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DIP per mole of protease (calculated Mi.=28,238). This diphosphorylated species represents at least 10% of the inactivated desMet-protease. In order to confine DIP incorporation to the active site of the protease, 200 |LIM protease was incubated for 2 hours with a series of DFP concentrations ranging from 0.3 to 2 mM. The deconvoluted ES mass spectra of the samples (Figures 2 and 3) and the activity data (Figure 4 and Table II) indicate a correlation between the appearance of the DIP-CMV species and protease inactivation, indicating that DIP-CMV protease has incorporated the diisopropylphosphoryl group at its active site serine.

FIGURE 4. Effect of DFP Treatment on Protease A c t i v i t y . Serial dilutions of CMV protease standard and DFP-treated protease samples (5-0.005 ^M) were assayed for their cleavage activity of the biotinylated peptide substrate for 2 hours, as described in the legend of Table 1. The amount of intact substrate remaining after quenching the reactions were monitored on the BIAcore.

A protease species with 2 DIP groups incorporated is apparent in samples incubated with 1.26, 1.63 and 2.07 DFP, although even in the sample incubated with 2.07 DFP, the diphosphorylated components represent less 5% of the inactivated protease. Based on Figure 3,

also mM mM than the

Inactivation of Human CMV by DFP

265

residual unmodified protease present in the samples incubated with 1.63 and 2.07 mM DFP is also less than 2%. Consequently, incubation of 200 jiM CMV protease at 22" C with 1.5 to 2 mM DFP was chosen as the optimum conditions for producing diisopropylphosphoryl-CMV protease suitable for crystallography. Table

DFP

II. Summary

Concentration

of DFP

Inhibition

of

CMV

Protease

mol DFP/mol

(mM) 0.33

1.70

69

0.62

3.10

36

1.26

6.30

14

1.63

8.50

6

2.07

10.4

4

4.00

22.0

0

Summary of CMV protease activity data presented in Figure 4 a n d the legend of Figure 2. The activities correlate well with t h e deconvoluted mass spectra presented in Figures 2 and 3.

IV. Conclusions CMV protease was inactivated by DFP under conditions optimum for its activity and stability (pH 7.8, 10% glycerol and 10 mM DTT). The results of electrospray mass spectrometry and protease activity measurements indicated that a 4 hour incubation of 180 |LIM CMV protease with 4 mM DFP completely inactivated the protease, but that within 3 hours nearly 10% of the protease had incorporated a second diisopropylphosphoryl (DIP) moiety. After performing 2 hour inactivations under the same conditions, but with lower DFP concentrations, it was found that 1.5 to 2 mM DFP was optimum for producing 98% DIP-CMV protease, which contained less than 5% protein with the second site labeled.

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Since the structure of the DFP-treated serine proteases resembles that of the tetrahedral transition-state intermediate, and the inactivated enzyme would not be susceptible to autoproteolysis, production of DIP-CMV protease would be useful for structure based drug design.

References 1.

Burck, P. J., Berg, D. H., Luk, T. P., Sassmannshausen, L. M., Wakulchik, M., Smith, D. P., Hsiung, H. M., Becker, E. W., Gibson, W., and Villarreal, E. C. (1994). J. Virol 68, 2937-2946. 2. Gibson, W. (1981). Virol 111, 516-537. 3. Irmiere, A., and Gibson, W. (1983). Virol 130, 118-133. 4. Stevens, J. T., Mapelli, C , Tsao, J., Hail, M., O'Boyle II, D., Weinheimer, S. P., and Diianni, C. L. (1994). Eur. J. Biochem. 2 2 6 , 361-367. 5. Holwerda, B. C , Wittwer, A. J., Duffin, K. L., Smith, C, Toth, M. V., Carr, L. S., Wiegand, R. C , and Bryant, M. L. (1994). /. Biol Chem. 269, 25911-25915. 6. O'Boyle II, D. R., Wager-Smith, K., Stevens III, J. T., a n d Weinheimer, S. P. (1995). /. Biol Chem. 270, 4753-4758. 7. Stroud, R. M., Kay, L. M., and Dickerson, R. E. (1974). /. Mol Biol 83, 185-208. 8. Fenn, J. B., Mann, M., Meng, C. K., Wong, S. F., and Whitehouse, C. M. (1990). Mass. Spectrom. Rev. 9, 37-70. 9. Covey, T. R., Bonner, R. P., Shushan, B. L, and Henion, J. D. (1988). Rapid Commun. Mass. Spectrom. 2, 249-256. 10. Cohen, J. A., Oosterbaan, R. A., and Berends, F. (1967). Meth. Enzymol 11, 686-702.

studies on the Status of Arginine Residues in Phospholipase A2 from Naja naja atra (Taiwan cobra) snake venom C. C. Yang, T. S. Yuo and C. Y. Chen Institute of Life Sciences, National Tsing Hua University, Hsinchu, Taiwan 30043, ROC

I . Introduction The enzyme phospholipase A2 (PLA2, EC 3.1.1.4) is widely distributed among various species in the animal kingdom, notably in the pancreatic tissues of mammals and venoms from snakes and bees. It specifically catalyzes the hydrolysis of the acyl-ester bond at the sn-2 position of l,2-diacyl-3-^/7phosphoglycerides in the presence of Ca^^ (Dennis, 1983). The enzyme activity of PLA2 is strongly dependent on the status of the substrate. Although PLA2 enzymes hydrolyze monomeric substrates, their activities increase by several orders of magnitude when the substrates are in aggregated form. This resuh has led to the hypothesis that there is a specific interfacial recognition site at the N-terminal region with specific affmity for lipidwater interfaces (Pieterson et al.^ 1974; van Dam-Mieras et al, 1975). Several studies have revealed that the N-terminal region is important for the activity of PLA2 enzymes (Dijkstra et al, 1981; van Eijk et al.^ 1984; Haruki et al.^ 1986; Oda et al, 1986). The X-ray crystallographic analysis of PLA2 enzymes (Dijkstra et al, 1983; Brunie et al, 1985; Scott et a/., 1990; White e/ a/., 1990) showed that a hydrogen bonding network connects the active site to the N-terminal region. Renetseder et al (1985) suggested that the hydrogen bonding network is an activation network which could stabilize the productive conformation of PLA2 enzymes for aggregated substrates. Yang and Chang TECHNIQUES IN PROTEIN CHEMISTRY VIII Copyright © 1997 by Academic Press All rights of reproduction in any form reserved.

267

C.C.Ymg etal.

268

(1988) suggested that the N-terminal region mediated the effects of lipid binding to the active site by the hydrogen bonding network, and causes architectural changes at the active site which result in an active conformation. The structural features of PLA2 showed by chemical modification to be important for enzymatic activity are histidine (Volwerk et al, 1974; Yang and King, 1980), aspartic acid (Fleer et al, 1981; Yang et al, 1983), tryptophan (Yang and Chang, 1984; Chang et al, 1993), tyrosine (Yang et al, 1985; Yang and Lee, 1986; Soons, et al, 1986), and lysine residues (Yang et al, 1982; Yang and Chang, 1989). Therefore, a study of the role of arginine residues might throw some light on the catalytic mechanism of PLA2 enzymes. PLA2 from N. naja atra (Taiwan cobra ) snake venom is an acidic single chain polypeptide consisting of 119 amino acid residues and contains five Arg residues at the position 16, 30, 42, 94 and 117. In this study, the Arg residues of A^. naja atra PLA2 were selectively modified with phenylglyoxal (PG), and the modified derivatives were separated by HPLC. Based on changes in physicochemical and biological properties afl;er Arg modification, the possible role played by Arg residues in A^. naja atra PLA2 is discussed.

II. Materials and Methods 1.

Materials

PLA2 from Naja naja atra (Taiwan cobra) venom was isolated and purified as previously described (Yang et al, 1981). Phenylglyoxal was purchased from Aldrich Chemical Co., 8-anilinonaphthalene sulfonate (ANS) was obtained from Pierce Chemical Co.. The SynChropak RP-18 column (4.6mm x 25cm) was obtained from Synchrom. All other reagents were of analytical grade. 2.

Chemical Modification of Arg Residues with Phenylglyoxal (PG)

PLA2 was modified with PG according to the procedure described by Takahashi (1968). One micromole of PLA2 in 1 ml of 0.1 M sodium borate buffer (pH 9.0) was incubated with 100-fold molar excess of PG. The reaction was allowed to proceed for 2 hr at 37°C, then quenched by the addition of a few drops of acetic acid. The modified proteins were immediately desalted by passing through a Sephadex G-25 column equilibrated with 0.1 M acetic acid and the protein fraction was lyophilized. The modified proteins were separated by HPLC on a SynChropak RP-18 column (4.6 mm x 25 cm), equilibrated with 0.1% TFA and eluted with a linear gradient of 28-35% acetonitrile for 35 min. Flow rate was 0.8 ml/min and the effluent was monitored at 280 nm. 3.

Identification of the Modified Arg Residues

To determine the positions of the Arg residues modified in the sequence of PLA2,

Status of Arg-Residues in Phospholipase A2

269

the PG derivatives were reduced and carboxymethylated (RCM) according to the procedure described by Crestfield et al. ( 1963 ). The RCM-proteins (1.4 mg) were dissolved in 0.2 M ammonium bicarbonate buffer at pH 7.8 and trypsin was add (30:1, w/w). Digestion was carried out at 37 C for 3 hr and the hydrolysates were lyophilized. The tryptic hydrolysates were separated by HPLC on a SynChropak RP-18 column (4.6 mm x 25 cm) equilibrated with 0.1% TFA and eluted by a linear gradient of 0-35 % acetonitrile for 105 min. 4.

Circular Dichroism (CD)

CD spectra of native and Arg-modified PLA2 were measured from 190 to 250 nm on a Jasco J-700 spectropolarimeter at a concentration of 0.285 mg/ml in 10 mM Tris buffer (pH 8.0) with a cell path-length of 0.1 cm. The estimated fraction [%] of the spectra were obtained by signal averaging five scans. 5.

Other Tests

Determination

of

PLA2

activity

and

antigenicity,

polyacrylamide-gel

PCM

0.6 PG-2

1 0 CO

<

0.4

• "

Native

1

0.2

iyA>'^--U 1

20 Time (min)

L

40

Figure. 1. Separation of Arg-modified PLA2 on a SynChropak RP-18 column. The column (4 6 mm x 25 cm) was equilibrated with 0.1% TFA and eluted with a linear gradient of 28-35% acetonitrile for 35 min. Flow rate was 0 8 ml/min and the effluent was monitored at 280 nm.

270

C.C.Yang era/.

electrophoresis, amino acid analysis and sequence determination, Ca^^-induced difference spectroscopy and fluorescence measurement were performed in essentially the same manner as previously described (Yang and Chang, 1984, 1988; Chang and Yang, 1988).

III. Results and Discussion A. Chemical Modification of PLAi by Phenylglyoxal (PG) PLA2 from N. naja atra snake venom was modified with 100-fold molar excess of PG at pH 9.0 for 2 hr. The modified proteins were separated by HPLC on a SynChropak RP-18 column and two major modified derivatives were isolated (Figure 1). Polyacrylamide gel electrophoresis revealed that both modified derivatives are homogeneous and move slower than the native PLA2. Amino acid analysis showed that only one Arg residue is modified in each of the PG-1 and PG-2 derivatives. In order to determine the positions of the Arg residues modified in the sequence of N. naja atra PLA2, PLA2 and the Arg-modified derivatives were digested with trypsin after reduction and carboxymethylation. The tryptic Table I. Amino Acid Compositions of Tryptic Peptides of RCM-PLA2 and Arg-modified PLA2"

Asp Glu Gly Ser His Arg Thr Ala Pro Tyr Val Met l/2Cys-Cys He Leu Phe Lys Corresponding peptide Arg-modified

A

B

C

D

5.6 (6) 1.0(1) 0.6(1)

0.9(1) 0.8(1)

5.5 (6) 0.8(1) 0.7(1)

0.8(1) 0.9(1)

2.1(2) 1.1 (1) 2.8 (3) 1.1(1) 1.6(2)

0.2(1) 0.9(1) 1.0(1)

1.0 (1) 0.9(1) 1.1(1) 0.7(1) 0.6(1) 0.3(1) 0.7(1)

0.8(1) 2.7 (3) 1.0(1) 1.5 (2)

0.2 (1) 0.8(1) 0.8(1)

1-2(1) 1.0(1) 0.7(1) 0.2 (1) 0.7(1)

0.9(1)

0.6(1) 101-119

2.3 (2)

7-19

101-119

7-19

Arg-117

Arg-16

^he peptides obtained from Figure. 2 were analyzed.

Status of Arg-Residues in Phospholipase Aj

271

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0.02

0.04|-

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40 60 Time (min) Figure. 2. HPLC profiles of the tryptic digests of native (1), and Arg-modified derivatives PG1 (2) and PG-2 (3). The tryptic hydrolysates were separated by HPLC on a SynChropak RP-18 column (4.6 x 25 cm) equilibrated with 0.1% TFA and eluted with a linear gradient of 0-35% acetonitrile for 105 min.

272

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NLYQFKNMIQCTVPSRSWWDFADYGCYCGRGGSGTPVDDL 60

80

DRCCQVHDNCYNEAEKISGCWPYFKTYSYECSQGTLTCKG

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GNNACAAAVCDCDRLAA

119

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Figure. 3. Amino acid sequence of .V. naja extra PLA2 (Tsai et ai, 1981; Pan et al, 1994) and tryptic peptides A, B, C and D.

hydrolysates were separated by HPLC on a SynChropak RP-18 coiumn and the elution profiles are shown in Figure. 2. The resuhs of amino acid analysis show that peaks A and B in the native PLA2, which represent the peptide fragments at positions 101-119 and 7-19 (Table I ), respectively, shifted to the respective peptides C and D on the chromatographic profiles of the modified derivatives PG-1 and PG-2 (Figure 2). Amino acid analysis and sequence determination of peaks C and D (Figure 3) revealed that they were derived from the peaks A and B in the native PLA2, but were missing an Arg residue at positions 117 and 16. These results unambiguously indicate that Arg-117 and Arg-16 are modified in derivatives PG-1 and PG-2, respectively. B. Properties of Arg-modified Derivatives As shown in Table II, a precipitous drop in enzymatic activity to 25.4% that of the native PL A? was observed when Arg-16 was modified. Modification of Arg-117 resulted in a decrease in enzymatic activity by only 12.1%. However, the antigenic activities of the both modified derivatives remained unchanged. The Arg-modified derivatives also enhanced the emission intensity of ANS dramatically, and the emission intensity of the ANS-enzyme complex increased in parallel with increasing concentration of CdC^ until the saturation level was reached. The dissociation constant of Ca^^ to ANS-enzyme complex (Kd) was Table II. Properties of the Native and Arg-modified PLA2 Enzymatic activity (%) Native PLA2

100

Antigenic activity(%)

Pl

100

5.2

Arg-117 modified

87.9

97

5.0

Arg-16 modified

25.4

95

5.0

273

Status of Arg-Residues in Phospholipase A2 Table HI. Physical Properties of Native and Arg-modified PLA2 ANS binding (Kd, liM) Ca^^ binding (Kd, mM)

with 10 mM Ca'"

without Ca'^

Native PLA2

0.53

85

180

Arg-117 modified

0.50

70

127

Arg-16 modified

0.71

58

98

calculated by Scatchard plots derived from the degree of saturation of the emission intensity (Table III). In order to find out whether there is specific interaction between ANS and the enzymes, Scatchard plots was obtained by titration of 5 //M protein with increasing dye concentration from 5 to 50 jwMin the presence or absence of Ca^^ (10 mM ). A plot of the intensity of fluorescence [F] vs. [F]/[ANS] gave lines with slopes corresponding to the respective dissociation constant of the complex. As listed in Table 3, the binding ability with ANS has been changed slightly afl;er Arg-modification. The fact that the Arg-modified derivatives are similar to native PLA2 in their affinity for C3^\ and the Scatchard plots revealed that there is only one kind of Ca^^ binding site indicates that the loss of the biological activity was not related to cofactor binding. CD spectra of Arg-modified derivatives are similar to that of PLA2, reflecting the fact that modification does not affect the secondary structure of PLA2. The antigenic activity of PLA2 remained unchanged afl:er Arg modification at position 16 or 117 Therefore, the observed loss of biological activity is more likely due to the disappearance of an electrostatic interaction between the PLA2 and substrates than the conformational change of the enzyme molecule. Scott et al. (1990) showed that the catalytic event occurred on a rigid internal surface that is well shielded from bulk solvent. In their discussion of the hydrophobic channel and the interficial recognition surface, they suggested that the phospholipid molecule undergoing hydrolysis leaves the aggregate and reaches this catalytic surface by facilitated diffusion through a hydrophobic channel whose opening is in the interfacial binding surface. Inspection of the tertiary structure of .¥. naja atra PLA2 shows that the residues at positions 2, 3, 5, 6, 9, 18, 30 and 63 constitute the hydrophobic channel (Figure 4) which are involved in the interaction of enzyme molecule with phospholipid substrates (White et al, 1990; Chang et al, 1993). Thus, it is conceivable that the modification of Arg-16 might directly distort the binding ability of PLA2 with substrate v/aj perturbs the phosphate portion of the substrate,

274

C. C Yang et al

Figure. 4. Tertiary structure of N. naja atra PLA2 showing the relative location of the Arg residues and the residues of which the "hydrophobic channel" is composed. The model is based on the result of White et al. (1990). The "hydrophobic channel" was formed by the residues at positions 2, 3, 5, 6, 9, 18, 30 and 63. Numbers in parentheses indicate the position of Arg residues at 16 and 117 in PLA2.

facilitates diffusion through the hydrophobic channel from interfacial surface, and causes a drastic loss in enzymatic activity. IV. Conclusions Phospholipase A2 (PLA2) from Naja naja atra (Taiwan cobra) snake venom was subjected to arginine modification with phenylglyoxal (PG), and two major derivatives were separated by HPLC. The results of amino acid analysis and sequence determination revealed that Arg-117 and Arg-16 were modified by PG. Modification of Arg-117 and Arg-16 resulted in a decrease in enzymatic activity of PLA2 by 12.1% and 74.6%, respectively. The Arg-modified derivatives were similar to native PLA2 in their affinity for Ca^^, and the Scatchard plots revealed that the loss of the biological activity is not related to cofactor binding. Modification did not significantly affect the secondary structure of the PLA2 molecule as revealed by CD spectra. The antigenic activity of PLA2 also remained unchanged after modification. The results indicate that Arg-16 is important for the biological activities of PLA2 and the modification of Arg-16

Status of Arg-Residues in Phospholipase Aj

275

might directly distort the binding ability of PLA2 with substrate vra. perturbs the phosphate portion of the substrate, facilitates diffusion through the hydrophobic channel from interfacial surface, and causes a drastic loss in enzymatic activity. Acknowledgments This work was supported by grant NSC 84-2311-B007-027 from the National Science Council, RepubUc of China. The authors would like to thank Miss F.S. Wu for her technical assistance. References Brunie, S., Bolin, J., Gewirth, D. and Sigler, P.B. (1985) J. Biol Chem. 260, 9742. Chang, L.S., Kuo, K.W. and Chang, C.C. (1993) Biochem. Biophys. Acta 1202, 216. Chang, L.S. and Yang, C.C (1988) J. Protein Chem. 7, 713. Crestfield, S.A.M., Moore, S. and Stein, W.H. ( 1 9 6 3 ) / Biol. Chem. 238, 622. Dennis. E.A. (1983) Phospholipases. In "The Enzymes" (Boyer, P.D.. ed). Vol. 16, p. 307. Academic Press, New York. Dijkstra, B.W., Kalk, K.H., Hoi, W.G.J, and Drenth, J. (1981) J. Mol. Biol. Ul, 97. Dijkstra. B.W., Renetseder, R., Kalk. K.H., Hoi. W.G.J, and Drenth, J. (1983) J. Mol. Biol. 168, 163. Fleer, E.A.M.. Verheij. H.M. and de Haas, G.H. (1981) Eur. J. Biochem. 113 , 283. Haruki, H.. Teshima, K., Samejima, Y., Kawauchi, S. and Ikeda. K. (1986) J. Biochem. 99, 99. Pan. P.M., Yeh, M.S., Chang. W.C, Hung, C.C and Chiou S.H. (1994) Biochim. Biophys. Res. Commun. 199, 969. Pieterson, W.A., Vidal J.C. Volwerk, J.J. and de Hass, G.H. (1974) Biochemistry 13, 1455. Renetseder, R., Brunie, S.. Dijkstra, B.W., Drenth. J. and Sigler. P.B. (1985) J. Biol. Chem. 260, 11627. Scott, D.L.. White, S.P.. Otwinowski. Z.. Yuan. W . Gelbs, M.H. and Sigler, P.B. (1990) Science 286, 1541. Soons, K.R., Condrea, E., Yang, C.C and Rosenberg. P. (1986) Toxicon 24, 679. Tsai, I.H.. Wu. S.H. andLo. T.B. (1981) Toxicon 19, 141. TakahashiK. (1968) J. Biol. Chem. 23. 6171. van Dam-Mieras, M.C.E., Slotboom, A, J., Pieterson. W.A. and de Haas, G.H. (1975) Biochemistry 14, 5387. van Eijk, J.H.. Verheij, H.M. and de Haas. G.H. (1984) Eur. J. Biochem. 140, 407. Volwerk, J.J.. Pieterson. W.A. and de Haas, G.H. (1974) Biochemistry 13, 1446. White. S.P., Scotte. D.L.. Otwinowski, Z.. Gelb, M.H. and Sigler, P.B. (1990) Science 286, 1560. Yang, C.C and Chang, L.S. (1984) J. Protein Chem. 3, 195. Yang, C.C and Chang, L.S. (1988) Toxicon 26, 721. Yang. C.C and Chang. L.S. (1989) Biochem. J. 262, 855. Yang. C.C, Chen, S.F. and Fan, Y.C (1983) Toxicon Suppl. 3, p. 509. Yang, C.C, Huang, C.S. and Lee, H.J. (1985)./. Protein Chem. 4, 87. Yang, C.C and Lee. H.J. (1986)./. Protein Chem. 5, 15. Yang, C.C. and King. K. (1980) Biochim. Biophys. Acta 614, 373. Yang. C.C, King, K. and Sun. T.P. (1981) Toxicon 19. 645. Yang, C.C, King, K., Sun. T.P. andHseu. W.S. (1982) The Snake 14. 110.

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Selective Reduction of the Intermolecular Disulfide Bridge in Human Glial Cell Line-Derived Neurotrophic Factor Using Tris-(2-Carboxyethyl)Phosphine

John O. Hui, John Le, Viswanatham Katta, Michael F. Rohde and Mitsuru Haniu Department of Protein Structure, 14-2-E Amgen Inc., Thousand Oaks, CA 91320

INTRODUCTION The essential roles played by the neurotrophic growth factors in the development of the nervous system have been well documented (1,2). Studies on how the different factors interact with their target cells and how such interactions eventually result in the generation of biological responses are actively being examined (3,4). Glial cell linederived neurotrophic factor (GDNF) is one of the more recently identified neurotrophic factors, first purified from the conditioned medium of a rat glial cell line (B49) by Lin and co-workers (5,6). Because of its ability in supporting the growth of mid-brain dopaminergic neurons in-vitro, GDNF has been implicated to have therapeutic potential in the treatment of Parkinson's disease (7). Mature GDNF is a single polypeptide with 134 amino acid residues containing 7 cysteines and functions as a glycosylated, disulfide linked dimer. Examination of the protein's primary structure suggests that GDNF is a distant member of the transforming growth factor-fi (TGF-fi) superfamily of growth factors (5). The recombinant protein has been expressed in Escherichia coli and is currently being developed as a candidate for human therapeutic use. Because the protein has to be refolded from inclusion bodies, it is important to identify the disulfide linkages of the final purified product. Based on the locations of the 7 cysteines with their relative positions that are conserved among all members of the TGF-6 family, it is expected that GDNF would share a similar disulfide structure as the other members TECHNIQUES IN PROTEIN CHEMISTRY VIII Copyright © 1997 by Academic Press All rights of reproduction in any form reserved.

277

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John O. Hui et al

of this class of growth factors. The crystal structure of TGF-62 has been solved to 1.8 A resolution by the study of Davies and associates (8) and the cystine linkages have been determined to exist in a "cystine knot", a threaded-ring conformation (9, 10). If the disulfides in GDNF do form the cystine knot, the following intramolecular cystine bridges would be expected to form: Cys^^ to Cys^^^; Cys^^ ^Q Cys^^^. Through this ring of 8 amino acid residues, the third intramolecular disulfide bond between Cys"*^ and Cys^02 would pass. The 2 monomers would then cross-link via a disulfide bond between Cys^^^. As can be expected from this complex structure, the protein was refractory to extensive proteolytic degradation under native conditions (11). In this investigation, we address the issue of disulfide assignment by subjecting GDNF to partial reduction using the chemical reagent tris-(2-carboxyethyl)phosphine (TCEP) in 0.17 M acetic acid at pH 2.5 and we report the structural characterization of the single reduced disulfide bond.

MATERIALS GDNF was expressed in Escherichia coli, refolded and purified to apparent homogeneity by the Process Development Department at Amgen Inc. The protein retains the initiation methionine and its biological activity is essentially identical to the native des-Met species (unpublished data). Tris-(2-carboxyethyl)phosphine (TCEP) and sequencing grade trypsin were obtained from Boehringer Mannheim. Dithiothreitol (DTT), iodoacetamide (lAM), N-ethylmaleimide (NEM), and 4-vinylpyridine (4-VP) were purchased from Sigma. Sequanal grade guanidine hydrochloride and urea, spectral grade trifluoroacetic acid (TEA) were products of Pierce. All other reagents were of the highest quality commercially available. HPLC grade water and acetonitrile from Burdick and Jackson were employed throughout.

METHODS Partial reduction of GDNF In a typical preparation, 2 |LI1 of 0.5 M TCEP was added to 500 |ig of GDNF (16.0 nmol) in 200 |Lil of 0.17 M acetic acid. Reduction was allowed to proceed at 45°C for 2 hours before the sample was chromatographed through a Vydac C4 column (0.46 x 25 cm) using a Hewlett Packard 1090 M liquid chromatograph. Solvent A was 0.1% TEA in water and solvent B was 0.1% TEA in 90% acetonitrile. The column was equilibrated in 10% solvent B and a flow rate of 0.7 ml per min was used. After sample application, the column was washed isocratically with 10% solvent B for 5 min. Protein elution was accomplished by a linear gradient in 2 steps: 10 to 30% solvent B over 5

Disulfide Bridge in Glial Cell Line-Derived Neurotrophic Factor

279

min, followed by 30 to 55% solvent B over 45 min. Protein elution was monitored by absorbance at 215 nm. The proteins were collected manually and concentrated under vacuum to a minimal volume prior to structural analysis. Alkylation of the partially reduced GDNF with N-ethylmaleimide To the partially reduced GDNF was added 100 |il of 0.2 M NEM followed by 100 |il of 6 M guanidine hydrochloride in 0.25 M Tris containing 1 mM EDTA at pH 8.5. The reaction was allowed to occur at room temperature for 30 min in the dark before it was quenched by adjusting the pH to 2.0 with 25% TFA. The modified protein was purified by reverse phase chromatography as described in the previous section. Complete reduction and pyridylethylation of the NEM modified GDNF To the lyophilized, NEM modified protein was added 50 |il of 6 M guanidine hydrochloride in 0.25 M Tris containing 1 mM EDTA at pH 8.5. Reduction was initiated by the addition of 5 |il of 0.1 M DTT. The sample was incubated at 45°C for 1 hour and the remaining free sulfhydryls were alkylated by 2 |il of 4-vinylpyridine. The sample was left at room temperature for 30 min before being purified by HPLC as described. Tryptic digestion of the modified GDNF To identify the residue that was modified by NEM, 100 |ig of the NEM treated, completely reduced and pyridylethylated GDNF was dissolved in 50 |Lil of 0.4 M ammonium bicarbonate containing 8 M urea. The solution was diluted with 150 |LI1 of water and 2 |ig of trypsin was added. Proteolysis was allowed to proceed at 37'C for overnight and the peptides generated were analyzed by chromatography through a Vydac C18 column (0.46 x 25 cm) at a flow rate of 0.7 ml per min. Solvents A and B were identical to those described above. The column was equilibrated in 2% solvent B. After sample injection, the column was washed with 2% B for 5 min before a linear gradient of 2 to 55% B over 1 hour was applied. Peptide elution was monitored by absorbance at both 215 nm and 254 nm. The peptides were manually collected and dried prior to structural characterization. Protein and peptide sequence analysis Automated Edman degradation of protein and peptide samples were performed using an Applied Biosystems sequencer (Model 470A or 477A) or a Hewlett Packard G1005A sequencer. Each sequencer was fitted with an on-line HPLC analyzer for the identification of phenylthiohydantoin derivatives.

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Mass spectrometry The molecular mass of protein and peptide samples was determined by electrospray ionization mass spectrometry using a Perkin-Elmer Sciex API 100 mass spectrometer. The sample was introduced either by infusion or by on-line liquid chromatography/mass spectrometry (LC/MS) using a splitter. The data were obtained by scanning from 450 to 2000 Da with a scan time of 5 s and a step size of 0.25 Da with 1.0 ms dwell time per mass step. The molecular mass of the sample was obtained using the software provided by the instrument manufacturer. SDS-PAGE Laemmli (12) gels (14%) were run in non-reducing conditions and stained with Coomassie Blue for detection of proteins as described (13).

RESULTS & DISCUSSION To show that the Escherichia coli expressed GDNF has been properly refolded, we have initiated a study to identify its disulfide bridges. However, establishing the disulfide bonds of a protein is often a labor intensive task. The protein of interest is usually subjected to extensive proteolytic or chemical degradation, peptides which are linked, ideally by a single cystine bridge, are purified and analyzed (1416). In many instances, the cysteines exist as a cluster as in human chorionic gonadotropin (17) or in a "knotted" structure found in GDNF; under such conditions the proteins are refractory to proteolytic degradation and data from partial digestion are difficult to interpret. We have subjected GDNF to chemical reduction and ask the question: Would any disulfide bond(s) be selectively reduced and thus identified by protein chemistry techniques? A major concern was of course the reactivity of the free sulfhydryl group. Once a cysteine is generated, disulfide scrambling will occur rapidly at neutral or alkaline pH and may generate misleading data (18). TCEP has been well documented to be active at acidic pH by the study of Gray (19), thus disulfide rearrangement would be minimized. Partial reduction of GDNF Reduction of GDNF with TCEP (the molar ratio of the number of cysteines in the protein to chemical reagent employed in the experiment was 1 to 4) at 45°C for 2 hours resulted in 3 major peaks (Figure 1). Examination of each peak by SDS-PAGE under nonreducing conditions showed that both peaks B (10% of the total protein) and C (30% of the total protein) migrated as monomer. Peak A was the remaining unmodified protein and peak C, because it migrated with

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the same retention time from reverse-phase HPLC as a sample of GDNF that had been reduced with DTT in the presence of 6 M guanidine hydrochloride (data not shown), indicating that it corresponded to the fully reduced protein. The identity of peak B was therefore of interest. Alkylation of the protein with NEM produced a predominant product with a molecular mass of 15319.0 Da (Figure 2). Based on the primary structure of GDNF and assuming all the cysteinyl residues were linked by disulfides, the dimeric protein should give a molecular mass of 30386.0 Da. If the dimeric GDNF was cross-linked by a single cystine bond, its specific reduction and alkylation with NEM should generate a component with a molecular mass of 15319.0 Da. Hence, our data are consistent with the model that the dimeric protein is cross-linked through a single disulfide bridge. 15319 7.0e5 -1 6.3e5 J 5.6e5 \

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Identification of the intermolecular disulfide bridge Because any nucleophilic amino acid side chain can react with NEM, it is crucial to establish the residue on GDNF that was modified by the reagent. The modified protein was therefore purified by HPLC, completely reduced with DTT in the presence of 6 M guanidine hydrochloride, and the remaining cysteines differentially alkylated with 4-vinylpyridine prior to tryptic digestion. The chromatographic behavior of the phenylthiohydantoin derivative of pyridylethyl Cys (PTH-pyridylethyl Cys) has been well characterized and can be identified readily (20). The tryptic map of a control GDNF sample where the protein had been completely reduced and pyridylethylated under denaturing condition is shown in Figure 3A. Peak 1 shows the co-purification of 2 peptides which were analyzed by both automated Edman degradation and mass spectrometry. They were identified to be L92vSDKVGQApeCpeCRPIAFDDDLSFLDDNLVYi20 (molecular mass 3443.8 Da; calculated = 3443.9 Da) and V97GQApeCpeCRPIAFDDDLSFLDDNLVYl20 (molecular mass = 2901.3 Da; calculated = 2901.3 Da), where peC is used to denote a pyridylethyl Cys throughout this Discussion. The 2 peptides contained the same 2 cysteinyl residues (residues 101 and 102) and were generated by incomplete digestion of the Lys^^-Val^^ bond. The cleavage after Tyr^^O suggests the presence of a contaminating chymotryptic-like activity in the commercial trypsin employed in this study. However, even highly purified preparations of trypsin had been shown to cleave certain aromatic bonds (21). Examination of the NEM modified sample indicated that peak 1 shifted to a later retention time (designated as peak 2 in Figure 3B). N-terminal analysis again gave 2 amino acid sequences: L92vSDKVGQAXpeCRPIAFDDDLSFLDDNLVYl20 (molecular mass = 3481.2 Da) and V97GQAXpeCRPIAFDDDLSFLDDNLVYl20 (molecular mass = 2938.7 Da), where X denotes the presence of a modified amino acid, which was eluted from the PTH-analyzer with a retention time of 6.8 min, approximately 1.5 min after PTH-Glu (Figure 4). If X corresponds to NEM-modified Cys, the expected mass of the 2 peptides would be 3463.8 Da and 2921.2 Da respectively. Therefore, our results showed that the 2 measured masses gave an increment of 18 Da, indicating the succinimide ring of NEM has been hydrolyzed to the succinamic acid derivative, which occurred during tryptic digestion. The retention time of the modified amino acid observed from the PTH-analyzer is also consistent with this hypothesis. Maleimide SH-reagents have been documented to be unstable at neutral and alkaline pH values (22). Sequence analysis of the remaining peptides showed that Cys ^^l ^v^as the only residue modified by NEM, demonstrating that GDNF dimerizes through a single disulfide bridge at Cys ^0^. The model based on the protein's homology with TGF-62 was hence confirmed. The

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partial reduction method described in this communication may be applicable to the other members of the TGF-C family and to the best of our knowledge, this is the first report on the chromatographic identification of the PTH-derivative of NEM modified Cys.

ACKNOWLEDGMENTS The authors are grateful to Drs Robert Rush, Tony Polverino and Scott Patterson for their critical comments on the manuscript.

REFERENCES 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20

Henderson, C.E. (1996) Current Opinion in Neurobiology 6, 6470. Thoenen, H. (1991) Trends Neurosci. 14, 165-170. Isackson, P.J. (1995) Current Opinion in Neurobiology 5, 350357. Curtis, R. and DiStefano, P.S. (1994) Trends Cell Biol 4, 383-386. Lin, L.-F. H., Doherty, D.H., Lile, J.D., Bektesh, S. & Collins, F. (1993) Science 260, 1130-1132. Lin, L.-F. H., Zhang, T.J., Collins, F. and Armes, L.G. (1994) /. Neurochem, 63, 758-768. Gash, D.M. et al, (1996) Nature 380, 252-255. Daopin, S., Li, M. and Davies, D.R. (1993) Proteins : Structure, Function, and Genetics 17, 176-192. McDonald, N. Q. and Hendrickson, W. A. (1993) Cell 73, 421424. Massague, J. (1996) Cell 85, 947-950 (1996). Haniu, M., et al. (1996) Biochemistry , manuscript submitted. Laemmli, U.K. (1970) Nature 227, 680-685. Hui, J.O., Tomasselli, A.G., Zurcher-Neely, H. and Heinrikson, R.L. (1990) /. Biol. Chem. 265, 21386-21389. Hui, J.O., Le, J., Viswanatham K., Rosenfeld, R., Rohde, M.F. and Haniu, M. (1996) /. Prot. Chem. 15, 351-358. Violand, B.N., SchUttler, M.R., Duffin, K.L. & Smith, C.E. (1995) /. Prot. Chem. 14, 341-347. Acklin, C , Stoney, K., Rosenfeld, R., Miller, J.A., Rohde, M.F. & Haniu, M. (1993) Int. ]. Peptide Protein Res. 41, 548-552. Mise, T. and Bahl, O.P. (1980) /. Biol. Chem. 255, 8516-8522. Creighton, T.E., Zapun, A. and Darby, N.J. (1995) Trends in Biotech. 13, 18-23. Gray, W.R. (1993) Protein Science 2, 1749-1755. Fullmer, C.S. (1984) Anal. Biochem. 142, 336-339.

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Maroux, S., Rovery, M., and Desnuelle, P. (1966) Biochim, Biophys. Acta 111, 147. Gregory, J.D. (1955) /. Am. Chem. Soc. 77, 3922-3923.

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EFFECTS OF SURFACE HYDROPHOBICITY ON THE STRUCTURAL PROPERTIES OF INSULIN Mark L. Brader^ Rohn L. Millican^ David N. Brems^*, Henry A. Havel^» Aidas Kriauciunas* and Victor J. Chen** Divisions of ^Pharmaceutical Sciences, %iopharmaceutical Development and ^Diabetes Research Lilly Research Laboratories, Indianapolis, IN 46285

Introduction A classical observation of protein structure is that the interiors of soluble globular proteins are composed mainly of hydrophobic amino acids. This structural arrangement has been compared to the interior of a micelle (Kauzmann 1959). The significance of internal hydrophobicity as a dominant factor in protein folding has recently been demonstrated in studies of the thermostability of T4lysozyme mutants, in which specific residues were replaced with amino acids bearing lipophilic side chains of varying sizes (Mendel et al., 1992; Eriksson et al., 1992). These studies showed that, in general, protein conformational stability correlated positively with the degree of internal hydrophobicity. The studies on the T4-lysozyme mutants prompted us to consider the converse, namely the effects of increasing the surface hydrophobicity of a globular protein. We chose to address this by comparing the conformational stability of a model protein in the presence and absence of a hydrophobic group attached covalently to a specific surface residue. Human insulin (HI) was selected as the model protein because it is a relatively small polypeptide (Mr = 5808) for which there is a wealth of structural, chemical and biological information. In addition, the insulin molecule possesses a rich conformational chemistry distinguished by thoroughly characterized structural transitions and ligand binding processes (Derewenda et al., 1989; Birnbaum et al., 1996; Bloom et al., 1995; Choi et al., 1993, Bryant et al., 1992). The derivative bearing a surface hydrophobic group that is the subject of this study is N^-palmitoylLys^29 human insulin (Pal-HI, Structure 1).

Authors to whom correspondence should be addressed * Current Address: Amgen Inc., Thousand Oaks, CA 91320 TECHNIQUES IN PROTEIN CHEMISTRY VIII Copyright © 1997 by Academic Press All rights of reproduction in any form reserved.

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Structure 1. N^-palmitoylLysB29 human insulin (Pal-HI)

Experimental Methods N-hydroxysuccinimidyl palmitate was prepared by dropwise addition of palmitoyl chloride to a rigorously stirred suspension comprising one equivalent each of N-hydroxysuccinimide and triethylamine in dry tetrahydrofuran at room temperature. After removal of solids by filtration, the solvent was evaporated and the product (yield -70%) was used without further treatment. Pal-HI was prepared by the addition of 1.2 equivalents of Nhydroxysuccinimidyl palmitate to a stirred solution of insulin in DMSO containing a 20-fold molar excess of tetramethylguanidine. After 30 minutes, the reaction was quenched with four volumes of O.IN HCl and the reaction mixture chromatographed on a Vydac C4-reversed phase column, eluted with a 0.1% trifluroacetic acid-acetonitrile gradient. Pal-HI (yield -40%) eluted at a solvent composition of about 40% acetonitrile. Static light scattering experiments were performed using a Brookhaven Instruments goniometer as described in Needham et al. (1995). A dn/dc value of 0.183 mL/g was employed. Absorption and circular dichroism spectroscopies were performed using Gary 3E and Aviv 62 DS instruments, respectively, under the conditions described in Brems et al. (1992). Guanidine-HCl induced denaturation experiments were conducted according to the method of Brems et al. (1990).

Results A biophysical study of Pal-HI was performed in order to assess the effects of palmitoylation on the structure and self-association of this derivative. Circular dichroism (CD) has been utilized as a convenient and sensitive technique for studying the conformational behavior and self-association of insulin. Insulin forms a hexamer that possesses two high affinity binding sites for certain divalent transition metal ions (Brader & Dunn, 1991). The far-UV CD spectra of HI and Pal-HI recorded under identical conditions are presented in Figure 1. In the absence (Figure 1, Panel A) and presence (Figure 1, Panel B) of Zn(II), the farUV CD spectra of HI and Pal-HI show only a slight difference indicating that palmitoylation has not greatly altered the secondary structure of the parent insulin molecule. The small difference is likely attributable to minor secondary structural

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differences between HI and Pal-HI and the fact that palmitoylation introduces an additional amide chromophore which absorbs in the far-UV region. In contrast, Figure 2 shows that the near-UV CD spectra have similar profiles, while the peak magnitude of that for Pal-HI is only about half that of HI. This change in the near-UV CD reflects a perturbation in the chromophoric environments of the tyrosine residues. The decreased CD magnitude may simply arise from a greater flexibility of the tyrosine residues in Pal-HI. Two alternative possibilities could account for this observation. Since the magnitude of the nearUV CD of insulin has been shown to correlate with hexamer assembly (Strickland & Mercola, 1976) the data of Figure 2 could indicate that palmitoylation reduces self association in the monomer-dimer-hexamer equilibrium. In addition, the different near-UV CD of Pal-HI may arise from a different conformation of aromatic side chains induced by the presence of the palmitoyl group in Pal-HI that is not directly correlated with self association. To investigate the self-association of Pal-HI in the presence and absence of Zn(II), a static light scattering study has been performed. The data in Figure 3 shows that metal-free Pal-HI undergoes a concentration dependent selfassociation process. In the presence of Zn(II), Pal-HI exhibits a different

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concentration-dependent association profile indicative of higher levels of aggregation. The data of Figure 3 may be compared qualitatively with published self-association data of HI as measured by light scattering (Hvidt, 1991) and analytical ultracentrifugation (Brems et al., 1992). Although the data of Figure 3 do not distinguish the extent of hexamer formation from other aggregation states, the increased association relative to insulin suggests that the reduction in magnitude of the near-UV CD spectrum of Pal-HI (Figure 2) probably comes from a perturbation in the conformation of the tyrosine residues as a result of palmitoylation of LysB29, as opposed to decreased self-association. This interpretation is supported by the spectra of Figure 2 which show that the nearUV CD of the Pal-HI Zn(II) hexamer is also considerably diminished. Another interesting example of the conformational behavior of the insulin molecule is exhibited by the transition-metal-substituted hexamer. Figure 4 shows that the near-UV CD signal for Pal-HI increases with increasing levels of Zn(II) and that this effect saturates at a value of two Zn(II) ions per hexamer, identical to the Zn(II)-binding of insulin. The structure of the zinc insulin hexamer has been characterized thoroughly by a series of X-ray crystal structures (Baker et al., 1988; Derewenda et al., 1989; Ciszak & Smith, 1994; Smith & Dodson, 1992a; Smith & Dodson, 1992b). The hexamer consists of three insulin dimers associated about a three-fold symmetry axis. Two high-affinity metal binding sites exist on the three-fold axis, each formed by three His^lO residues. These chelation sites may accommodate metal ions such as Zn(II), Co(II), Co(III), Cu(I) and Cu(II) (Brader & Dunn, 1991). The hexameric complex exists in two interconvertable conformational states in which the subunits are designated as either T or R according to their conformation. The R^ hexamer is stabilized by noncovalent interactions with phenolic compounds which bind to six hydrophobic pockets on the surface of the hexamer. With Co(II) ions coordinated at the HisBlO sites, the hexamer gives rise to characteristic Co(II) d-d transitions evident

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in the visible absorption spectra. The visible spectra of Co(II) hexamer complexes have been shown to be extremely sensitive to the nature of exogenous small molecule that can serve as ligand to the metal at the HisBio site. The visible absorption spectra of HI and Pal-HI recorded in the presence of Co(II) and small molecule ligands are presented in Figure 5. Under these conditions, HI exists as the Co(II)-R6-(Ligand) hexamer in which the visible absorption spectrum arises from the pseudotetrahedral Co(II)His3L chromophore, where L is a small molecule ligand that is coordinated to the Co(II) center (Brader & Dunn, 1991). The similarity of the Pal-HI and HI spectra recorded under the same conditions indicates that the Co(II) center in this hexamer must be very similar with respect to ligand identity and geometry. This result indicates that the Pal-HI molecule forms a Co(II) hexamer and that the structural features required for formation of dimer and hexamer remain largely unperturbed by palmitoylation. In addition, these results show that the palmitoylated derivative retains the capacity to undergo the ligand-promoted T6 to R6 conformational transition and the same general profile for the phenol-binding isotherm. The conformational stabilities of HI and Pal-HI have been compared by studying the respective guanidine-HCl induced denaturation profiles. Experiments were conducted utilizing a concentration of 0.1 mg/mL protein in pH 7.4 buffer containing 20% ethanol. Under these solution conditions, both HI and Pal-HI were determined by ultracentrifugation analysis (data not shown) to exist in the monomeric state. The denaturation profiles of HI and Pal-HI obtained by monitoring the circular dichroism at 224 nm as a function of guanidine-HCl concentration are shown in Figure 6. The denaturation transitions begin at about 2.5 M guanidine-HCl and were shown to be completely reversible. From these data, the free energies of unfolding were calculated to be 6.0 and 4.5 kcal/mol for Pal-HI and HI, respectively.

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Discussion The increased conformational stability of Pal-HI over HI is a surprising yet intriguing finding of this study. The introduction of a large hydrophobic group to the surface of a small protein like insulin could perturb its structure to the extent of altering the conformation. The present results show that the palmitoylation of HI has not significantly altered its basic structural properties. Pal-HI retains the ability to form Zn(II) and Co(II) hexamers, to form pseudotetrahedral Co(II)His3L centers, to undergo the T^ to R6 hexamer conformational transition, and for the metal-free monomer to adopt a secondary structure that is closely analogous to that of HI. Collectively, these results indicate that the Pal-HI molecule forms a hexamer that is highly analogous to that of HI with respect to structure, ligand binding properties and conformational flexibility. The static light scattering results suggest that the Zn(II)-Pal-HI hexamers have a greater tendency to aggregate. This behavior is consistent with a structural model of the Zn(II)Pal-HI hexamer in which the assembly of this hexamer is essentially identical to that of the Zn(II)-HI hexamer, but the palmitoyl groups are accommodated on the surface of the hexamer. In such a model, the palmitoyl groups would interact with hydrophobic residues on the hexamer surface. An arrangement of this type would be in accord with the perturbed near-UV CD observed for Zn(II)-Pal-HI and would explain the increased propensity for hexamer aggregation as being attributed to enhanced m/^r-hexamer hydrophobic interactions. The situation is less clear for the metal-free self-association process. Although the specific nature of the metal-independent self-association is not understood, it appears to be more complex than merely the apparent monomer-dimer-hexamer equilibrium in effect for the assembly of the hexamer. It is likely that the association process for PalHI involves new hydrophobic interactions in which the palmitoyl groups

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participate. The details of how the palmitoyl group affects the intra- and interhexamer contacts both in the absence and presence of metal ions remains to be elucidated. The polypeptide of Pal-HI is folded as for HI, therefore, it is of interest to determine how the hydrocarbon tail of the surface-residing palmitoyl moiety is accommodated in an environment that is extensively solvent exposed. The increase in conformational stability in Pal-HI, compared to HI, may be interpreted by proposing the existence of a surface hydrophobic pocket on the insulin molecule that is capable of intramolecularly binding the palmitoyl group attached to LysB29. As a result of this intramolecular interaction between the pocket and the fatty acyl chain, the structure of the polypeptide as a whole attains greater stability. The hydrophobic pocket is likely to comprise aromatic residues, which would be consistent with the observed perturbation of the near-UV CD spectrum. However, the purported structural alterations that have occurred to accommodate the palmitoyl group, evidently are local and do not greatly affect hexamerization, conformational flexibility or biological activity. It is interesting to note that in the mammalian cAMP dependent protein kinase, which is naturally N-terminal myristylated, the hydrocarbon tail of the myristyl moiety also folds back into the polypeptide and is bound in a cavity created by noncontiguous hydrophobic residues (Zheng et al., 1993). Furthermore, it was found that the myristylated protein showed increased thermostability when compared with its deacylated form (Yonemoto et al., 1993). An alternative explanation for the increased stability observed for Pal-HI is through a decreased entropy for the unfolded Pal-HI polypeptide. The acyl chain may interact with the hydrophobic side chains of the unfolded polypeptide thereby decreasing its entropy. An entropy reduction of the unfolded state would result in a decrease in the number of possible conformations thereby reducing the energy required to bring two chain elements together, with the overall effect of increasing the free energy of unfolding (Flory, 1956). For the present study of Pal-HI, LysB29 was chosen as the acylation site because of the simplicity of the conjugation chemistry. This residue resides in the C-terminus of the B-chain which is a region of the insulin molecule known to be extremely flexible (Weiss et al., 1989), and this region probably plays a minor role in the folding and unfolding process of unmodified insulin. A future direction of these studies is to investigate the effect of palmitoylation at a site corresponding to the more rigid and helical portion of the molecule with the aim of establishing how the hydrocarbon chain of the fatty acid would be accommodated and determining its impact on the overall structural integrity of the molecule.

Reference Baker, E. N., Blundell, T. L., Cutfield, J. F., Cutfield, S. M., Dodson, E. J., Dodson, G. G., Hodgkin, D. C , Hubbard, R. E., Isaacs, N. W., Reynaolds, C. D., Sakabe, K., Sakabe, N., & Vijayan, N. M. (1988) Phil Trans. Roy. Soc. Ser. B 319, 369-456. Birnbaum, D. T., Dodd, S. W., Saxberg, B. E. H., Varshavsky, A. D., & Beals, J. M. (1996) Biochemistry 35, 5366-5378. Bloom, C. R., Choi, W. E., Brzovic, P. S., Ha, J. J., Huang, S.-T., Kaarsholm, N. C, & Dunn, M. F. (1995) /. Mol. Biol. 245, 324-330. Brader, M. L., & Dunn, M. F. (1991) Trends Biochem. Sci. 16, 341-345. Brems, N. B., Alter, L. A., Beckage, M. J., Chance, R. E., DiMarchi, D. D., Green, L. K., Long, H. B., Pekar, A. H., Shields, J. E., & Frank, B. H. (1992) Protein Engineering 5, 527-533.

Effects of Surface Hydrophobicity on the Insulin

297

Brems, D. N., Brown, P.L., Heckenlaible, L. A., & Frank, B. H. (1990) Biochemistry 29, 9289-9293. Bryant, C, Strohl, M., Green, L. K., Long, H. B., Alter, L. A., Pekar, A. H., Chance, R. E., & Brems, D. N. (1992) Biochemisty 31, 5692-5698. Choi, W. E., Brader, M. L., Aguilar, V., Kaarsholm, N. C , & Dunn, M. F. (1993) Biochemistry 32, 11638-11645. Ciszak, E., & Smith, G. D. (1994) Biochemistry 33, 1512-1517. Derewenda, U., Derewenda, Z., Dodson, E. J., Dodson, G. G., Reynolds, C. D., Smith, G. D., Sparks, C, & Swenson, D. (1989) Nature 338, 594596. Eriksson, A. E., Basse, W. A., Zhang, X.-J., Heinz, D. W., Blaber, M., Baldwin, E. P., & Matthews, B. W. (1992) Science 255, 178-183. Flory, P.J. (1956) J. Am. Chem. Soc. 78, 5222-5232. Hvidt, S. (1991) Biophys. Chem. 39, 205-213. Kauzmann, W. (1959) Adv. Protein Chem. 14, 1-63. Mendel, D., Ellman, J. A., Chang, Z., Veenstra, D. L., Kollman, P. A., & Schultz, P. G. (1992) Science 256, 1798-1802. Needham, G. P., Pekar, A. H., Havel, H. A. (1995) J. Pharm. Sci. 84, 437442. Smith, G. D., & Dodson, G. G. (1992a) Proteins: Sruct. Func. Genet. 14, 401408. Smith, G. D., & Dodson, G. G. (1992b) Biopolymers 32, 1749-1756. Strickland, E. H., Mercola, D. (1976) Biochemistry 15, 3875-3884. Weiss, M. A., Nguyen, D. T., Khait, I., Inouye, K., Frank, B. H., Beckage, M., O'Shea, E., Shoelson, S. E., Karplus, M., & Neuringer, L. J. (1989) Biochemistry 28, 9855-9873. Yonemoto, W., McGlone, M. L., & Taylor, S. S. (1993) J. Biol. Chem. 268, 2348-2352. Zheng, J., Knighton, D. R., Xuong, N.-H., Taylor, S. S., Sowadski, J. M., & Ten Eyck, L. F. (1993) Protein Sci. 2, 1559-1573.

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The Effects of In Vitro Methionine Oxidation on the Bioactivity and Structure of Human Keratinocyte Growth Factor Christopher S. Spahr, Linda O. Narhi, James Speakman, Hsieng S. Lu, and Yueh-Rong Hsu Amgen Inc., Thousand Oaks, CA 91320

I. Introduction When the crystal structure of a protein is not available, other techniques can be employed to identify the amino acids that are involved in its structure and function. Commonly used techniques include chemical cross-linking, site-specific chemical modifications, and mutagenesis. Chemical modifications of Met residues using oxidizing agents such as hydrogen peroxide, t-butyl hydroperoxide, chloramine T, and sodium periodate have been useful in identifying structure and function relationships in many proteins (1-7). Keratinocyte growth factor (KGF) is a member of the fibroblast growth factor (FGF) family. The molecule is expressed by stromal fibroblasts and is involved in the proliferation and differentiation of epithelial cells in a paracrine mode (8). E.co/i-derived human KGF is biologically active (9) and may be clinically useful (10-12). In this study, hydrogen peroxide oxidation was performed on E.co/i-derived human KGF at pH 5.0 to understand the structure and function of the protein. Reverse-phase high performance liquid chromatography (RP-HPLC), peptide mapping, protein sequencing, and mass spectrometry were used to separate and identify the different methionine oxidized KGF species. Cation exchange HPLC was utilized to remove hydrogen peroxide from the different modified forms of the protein prior to mitogenic bioassay and circular dichroism (CD) analysis. Preferential oxidation of the methionine residues to methionine sulfoxide has enabled us to determine that Met 160 may play an important role in the biological function of KGF. TECHNIQUES IN PROTEIN CHEMISTRY VIII Copyright © 1997 by Academic Press All rights of reproduction in any form reserved.

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300

11. Materials and Methods Materials E.co//-derived human KGF contained a KGF polypeptide sequence (8) that started with Ser 24 as the N-terminus. The protein was produced and purified using methods similar to those described previously (9). Trifluoroacetic acid (TFA) was purchased from J.T. Baker. HPLC grade water and acetonitrile for RP-HPLC analysis were obtained from Burdick and Jackson. Urea was a product of Amresco. Sequencing grade trypsin was purchased from Boehringer Mannheim. Oxidation

of KGF with Hydrogen

Peroxide

KGF was incubated with hydrogen peroxide in 50 mM sodium acetate, pH 5.0 at a concentration of 0.4 mg/ml under various conditions to generate the oxidized KGF species. Detailed conditions were described in the legend to Figure 1. Reverse Phase HPLC RP-HPLC was performed using a Vydac C4 column (4.6 mm X 250 mm) connected to a Hewlett Packard 1090 HPLC system. The HPLC was equipped with a diode array detector and a PC based computer system for data processing. Solvent A was 0.1% TFA, while solvent B was composed of 0.1% TFA in 90% acetonitrile. The column was initially equilibrated at 3% B using a flow rate of 0.7 ml/min, with the absorbance monitored at 230 nm. The elution gradient consisted of isocratic conditions at 3% B for 5 minutes, followed by linear gradients to 40% B over 10 minutes, to 50% B over 20 minutes, and to 80% B over 5 minutes, and finally isocratic conditions at 80% B for 5 minutes. Tryptic Peptide

Mapping

The oxidized KGF species were vacuum lyophilized and then 100 \xl of 20 mM Tris-HCl, pH 7.0/2 M urea was added to each sample. About 25 |ig of protein was carboxyamidomethylated with iodoacetamide in the dark for 30 minutes at 25°C, using a 5:1 molar ratio of iodoacetamide to KGF. The carboxyamidomethylated KGF was digested with 0.625 [ig trypsin (40:1 ratio of KGF to trypsin) for 24 hours at 25°C. The proteolytic digests were quenched by the addition of 150 (il of 0.1% TFA, then injected onto a Vydac C8 column (4.6 mm X 250 mm) connected to a Hewlett Packard 1090 HPLC system. The

Methionine Oxidation of Human Keratinocyte Growth Factor

301

absorbance was monitored at 215 nm and a flow rate of 0.7 ml/min was used. The solvents used were identical to those described above. The digests were separated using isocratic conditions at 1% B for 5 minutes, then linear gradients to 36% B over 50 minutes, and to 70% B over 5 minutes, and finally isocratic conditions at 70% B for 5 minutes. The tryptic peptides collected from RP-HPLC were vacuum lyophilized, then resuspended in 0.1% TFA.

Protein Sequencing and Mass Spectrometry Peptides were sequenced using either an Applied Biosystems Model 470 or 477 or a Hewlett Packard GIOOOA protein sequencer, each equipped with narrow bore RP-HPLC for on-line analysis of the PTH-amino acids. Mass analysis of peptides was performed using matrix-assisted laser desorption/ionization mass spectrometry on a KRATOS MALDI III with a-cyano-4 hydroxy-cinnamic acid as the matrix.

Cation Exchange HPLC Cation exchange HPLC was performed using a TosoHaas TSK gel SP-5PW column (7.5 mm X 75 mm) connected to a Hewlett Packard 1050 liquid chromatograph. Buffer A was 20 mM sodium phosphate, pH 7.0, while buffer B was 20 mM sodium phosphate, pH 7.0/0.5 M sodium sulfate. The column was initially equilibrated with 30% buffer A at a flow rate of 1 ml/min. The elution was monitored by absorbance at 230 nm. The protein was eluted using isocratic conditions at 30% B for 20 minutes, followed by a linear gradient to 100% B over 1 minute, and finally isocratic conditions at 100% B for 20 minutes. The hydrogen peroxide eluted in the void volume. The KGF samples collected from cation exchange HPLC were buffer exchanged into phosphate-buffered saline. The protein concentrations were determined by UV absorption at 280 nm, assuming an extinction coefficient of 1.5 from a 0.1% protein solution. The concentrations were then adjusted to 1 mg/ml.

Circular Dichroism The far UV CD spectra and conformational stability, as determined by thermal denaturation, were compared using a Jasco J720 spectropolarimeter. The far UV CD spectra were determined using a cuvette with a 0.02 cm pathlength. Thermal stability was determined by continuously monitoring the change in the signal at 231 nm with increasing temperature in a thermal cuvette with a 0.1 cm pathlength and a Peltier JTC-345 thermal control unit.

Christopher S. Spahr et al

302

Balb/MK Cell Proliferation

Bioassay

The in vitro mitogenic bioassay used to determine the biological activity of KGF was similar to that described by Rubin, et al (13). The assay measured the incorporation of [^H]-thymidine by Balb/MK epidermal keratinocytes.

Kinetics of Methionine Oxidation of KGF The kinetics of oxidation of KGF were determined at room temperature with various time points. KGF was incubated with 0.5% hydrogen peroxide (v/v) in 50 mM sodium acetate, pH 5.0 at a concentration of 0.4 mg/ml. AUquots at various incubation times were injected onto a Vydac C4 column (4.6 mm X 250 mm) connected to a Hewlett Packard 1090 using the conditions described in the cation exchange HPLC section.

III. Results and Discussion Figure 1 shows RP-HPLC chromatograms of KGF samples oxidized by hydrogen peroxide under various conditions. KGF eluted as peak A in panel 1. Oxidized KGF eluted at earlier retention times (panels 2-5). As the temperature, the duration of incubation, or the 120 SO

120

fD&.rt&l

1

panel

2

painol

3

SO

i

g

°'

120

120-j

p a n e l -4-

soH

4.0

120-

panel

5

SO4.0 • 24

Tim»

25 (min.)

Figure 1. RP-HPLC chromatograms of the different species (A-F) generated by incubating KGF with hydrogen peroxide in 50 mM sodium acetate, pH 5.0 at a concentration of 0.4 mg/ml under various conditions. Panel 1- no hydrogen peroxide. Panel 2- 0.5% hydrogen peroxide, 1 hour at4°C. Panel 3- 0.125% hydrogen peroxide, 8 hours at 0°C. Panel 4- 0.5% hydrogen peroxide, 16 hours at 4°C. Panel 5- 2% hydrogen peroxide, 24 hours at room temperature.

Methionine Oxidation of Human Keratinocyte Growth Factor

303

40 Time (min.) Figure 2. Tryptic peptide maps of native KGF (A) and the individual oxidized KGF species (B-F) from Figure 1. Met containing peptides were labelled gl, hi, il, jl, kl, and 11, while the peptides containing the free sulfhydryl Cys 40 were labelled ml and nl. Oxidized forms of the same peptides were labelled g2, h2, i2, j2, k2,12, m2, and n2, respectively. Peptide i2 coelutes with the peptide T-V-A-V-G-I-V-A-I-K, while peptide 12 co-elutes with the peptide E-L-I-L-E-N-H-Y-N-T-Y-A-S-A-K.

Christopher S. Spahr et al

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hydrogen peroxide concentration was increased, the moditied protein eluted at earher retention times than native KGF. The individual oxidized KGF species observed on RP-HPLC were labelled from B-F. Sufficient quantities of KGF species A-F were generated using the conditions described in the legend to Figure 1. These forms were collected from RP-HPLC, then subjected to tryptic peptide mapping. The tryptic peptide maps of species A-F are shown in Figure 2. Peptides containing Met residues derived from native KGF (species A) were labelled as gl, hi, il, jl, kl, and 11. Peptides containing the free sulfhydryl Cys 40, now carboxyamidomethylated with iodoacetamide, were labelled as ml and nl. In the peptide map of species B, a large portion of peptide gl shifted to g2; in the map of species C, the majority of peptide hi shifted to h2; in the map of species D, peptides gl and hi shifted to g2 and h2 respectively; in the map of species E, peptides gl, hi, and il shifted to g2, h2, and 12 respectively; and in the map of species F, peptides gl, hi, il, jl, kl, 11, ml, and nl shifted to g2, h2, 12, j2, k2, 12, m2, and n2 respectively. The mass of peptides g2, h2,12, j2, k2, and 12 each increased by about 16 mass units as compared to the respective unoxidized peptides, consistent with the oxidation of the methionine residues to methionine sulfoxide (Table I). In summary, KGF species A was Table I. Sequence and mass spectrometry analysis of methionine containing peptides from the tryptic maps of the oxidized KGF species Peptide

Observed Sequence

Frag.

Obs. (MH+) Calc. (MH+) Mass

Mass

Mass

Difference

jl

W-T-H-N-G-G-E-M-F-V-A-L-N-Q-K

(125-139)

1732.1

1731.8

0.4 0.1 -0.2 0.3

kl

N-N-Y-N-I-M-E-I-R

(62-70)

1166.3

1166.6

-0.3 1

11

G-V-E-S-E-F-Y-L-A-M-N-K

(81-92)

1387.2

1387.7

-0.5

gl

G-T-Q-E-M-K

(56-61)

693.7

693.3

hi

S-Y-D-Y-M-E-G-G-D-I-R

(24-34)

1305.6

1305.5

il

T-A-H-F-L-P-M-A-I-T

(154-163)

1101.4

1101.6

1 ml

1 1 1 1

L-F-X-R (X= carboxyamidomethyl-Cys)

(38-41)

595.4

538.7

56.7

nl

R-L-F-X-R (X= carboxyamidomethyl-Cys)

(3741)

751.5

694.9

56.6

g2

G-T-Q-E-M-K

(56-61)

709.7

693.3

16.4 1

h2

S-Y-D-Y-M-E-G-G-D-I-R

(24-34)

1321.6

1305.5

16.1

1 ^ J2 1 ^ 12

1 m2 n2

T-A-H-F-L-P-M-A-I-T

(154-163)

1117.1

1101.6

15.5 1

W-T-H-N-G-G-E-M-F-V-A-L-N-Q-K

(125-139)

1747.9

1731.8

16.1

N-N-Y-N-I-M-E-I-R

(62-70)

1182.1

1166.6

15.5 1

G-V-E-S-E-F-Y-L-A-M-N-K

(81-92)

1403.4

1387.7

15.7

L-F-X-R (X= q^steic acid)

(38-41)

586.4

538.7

47.7 1

R-L-F-X-R (X= cysteic acid)

(3741)

742.6

694.9

47.7

Methionine Oxidation of Human Keratinocyte Growth Factor

305

unoxidized KGF; species B was identified to have most of Met 28 oxidized; species C had the majority of Met 60 oxidized; species D had Met 28 and 60 oxidized; and species E had Met 28, 60, and 160 oxidized. Methionine oxidation proceeded further, with no intermediate forms, from a species with Met 28, Met 60, and Met 160 oxidized (species E) to a species in which all six Met residues were oxidized (species F). The conditions required to oxidize all six Met residues also oxidized the free sulfhydryl Cys 40 to cysteic acid. Cys 40 has previously been suggested to reside in a solvent-inaccessible, buried environment (14). This data implies that the core of the protein opens up under these oxidizing conditions. As a result, the fully oxidized protein readily precipitated out of solution, therefore no further analysis was performed on it. In contrast, the conditions required to oxidize Met 28, Met 60, and Met 160 to methionine sulfoxide did not oxidize Cys 40 nor modify any other residues, as determined by peptide mapping. These oxidized KGF forms remained soluble and stable. The kinetics of oxidation for each Met residue was determined by RP-HPLC. The peak integration of the various oxidized forms taken at different time points was plotted as a function of time (Figure 3). After 60 minutes, about 90% of Met 28 and Met 60 were oxidized, while about 35% of Met 160 and essentially none of the Met 67, Met 90, and Met 132 residues had been converted to sulfoxide derivatives. The results indicate that Met 28 and Met 60 oxidize at a very rapid rate, while Met 160 was oxidized at a slightly slower rate. This data also suggests that Met 28 and Met 60 are located in an exposed environment on the surface of the protein and Met 160 may reside in a partially solvent-accessible environment. Met 67, Met 90, and Met 132 oxidize at substantially slower rates, suggesting that they reside at a buried environment that is relatively inaccessible to the oxidizing agent. Circular Dichroism and Mitogenic

Bioassay

Sufficient quantities of control KGF, KGF with Met 28 and Met 60 oxidized, and KGF with Met 28, Met 60, and Met 160 oxidized were generated as described in the legend to Figure 1. Cation exchange HPLC was utilized to rapidly remove the hydrogen peroxide from the protein samples prior to the Balb/MK mitogenic assay and CD analysis to avoid the RP-HPLC solvents that could potentially inactivate the protein. All samples were between 90-95% homogeneous, except for the KGF species with Met 28 and Met 60 oxidized which was approximately 85% homogeneous. From the analysis of these samples by CD, the far UV spectra of the KGF species with two methionine residues oxidized (Met 28 and 60) and the species with three methionine residues oxidized (Met 28, Met 60, and Met 160) were determined to be identical to that of native

Christopher S. Spahr et al

306

KGF with regard to the positive feature at 231 nm, as well as the psheet signal and the negative drift from the disulfide signals (Figure 4A). The oxidation of Met 28, Met 60, and Met 160 did not alter the secondary structure of KGF, nor the gross structure as monitored by the 231 nm peak in the far UV CD region. The stability of the two methionine oxidized species and three methionine oxidized species in solution was compared to that of native KGF using thermal denaturation (Figure 4B). Thermal denaturation of KGF is an irreversible reaction, as KGF precipitates following heat-induced denaturation, so stability was assessed by comparing the temperature at the onset of protein unfolding. The onset of melting occurred at 49°C in all samples. The identical melting points indicate that oxidation of Met 28, Met 60, and Met 160 did not result in a significant decrease in the rigidity or thermostability of the molecule. In the mitogenic bioassay, the KGF control had maximal activity around 34,000 cpm (Figure 5). To achieve 40% maximal activity in the bioassay (about 13,600 cpm), the KGF control required about 2 ng/ml and the KGF species with Met 28 and Met 60 oxidized required 6 ng/ml. However, to achieve this level of activiy in KGF with Met 28, Met 60, and Met 160 oxidized required about 2 |ig/ml. KGF with Met 28, Met 60, and Met 160 oxidized lost significant biological activity. Oxidation of Met 160 in KGF to methionine

73

.a X

O

.s o

-B

10

20

30

40

50

60

70

80

90

100110120

Time (min) Figure 3. The kinetics of oxidation for each Met residue of KGF. KGF was incubated with 0.5% hydrogen peroxide in 50 n\M sodium acetate, pH 5.0 at a concentration of 0.4 mg/ml at room temperature. D - Met 28 oxidized. A - Met 60 oxidized. • - Met 160 oxidized. O - Met 67, Met 90, and Met 132 oxidized.

307

Methionine Oxidation of Human Keratinocyte Growth Factor

CD
•o E. G

195 200

220

230

240

Wavelength (nm)

1

250

1

1

i

1

r

24 30 40 50 60 70 80 86

Temperature (°C)

Figure 4. (A) [left] Far UV CD spectra of the different KGF samples. 1 -KGF control without cation exchange HPLC step. 2 -KGF with Met 28 and Met 60 oxidized. 3 -KGF with Met 28, Met 60, and Met 160 oxidized. (B) [right] Thermal denaturation of the different KGF samples. The CD signal was monitored at 231 nm with increasing temperature. 1 -KGF control without cation exchange HPLC step. 2 -KGF with Met 28 and Met 60 oxidized. 3 -KGF with Met 28, Met 60, and Met 160 oxidized. 39500 37000 -

A

34500-

n3

32000 -

'J

29500-

6f)

27000 -

o M 1

s u > <

A D

D

A

24500 -

D

22000 1950017000-

d ft

14500120009500-

•

7000-

s

45002000-

Ql i f l ..(M .0 01

01

A

s •

m • A O

o

A D

•

A

•

D

•

•

•

O

O

o o

A

o

o o

O

0 11*1

.1

1

10

ng/ml

100

"•"I

1000

10000

100 000

Figure 5. Balb/MK nutogenic bioassay of the KGF samples, n -KGF control without cation exchange HPLC step. A -KGF control with cation exchange HPLC step. • -KGF with Met 28 and Met 60 oxidized. o -KGF with Met 28, Met 60, and Met 160 oxidized.

Christopher S. Spahr et al

308

sulfoxide did not appear to affect the secondary structure or the stabiUty of the molecule, yet it resulted in a severe loss in biological activity. These results imply that Met 160 may play an important role in the biological function of KGF.

IV. Conclusion In vitro hydrogen peroxide oxidation of the methionine residues of KGF to their sulfoxide derivatives demonstrated that Met 28, Met 60, and Met 160 are preferentially oxidized. Met 28 and Met 60 appear to be solvent-accessible and exposed on the surface of the protein, while Met 160 appears to be partially solvent-accessible. In contrast. Met 67, Met 90, and Met 132 oxidized much more slowly, which suggests they are located in a buried, less solvent accessible environment. Oxidation of these buried residues results in the generation of a highly unstable form of KGF that readily forms aggregates. Met 160 has been identified as a residue that may be critical to the biological function of KGF. A severe loss in biological activity was observed upon conversion of this residue to methionine sulfoxide, yet no change in secondary structure or thermostability was seen.

References 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14.

Hsu, Y-R., Narhi, L.O., Spahr, C.S., Langley, K.E., and Lu, H.S. (1996). Protein Scl 5:1165-1173. Keck, R.G. (1996). Anal Biochem. 236:56-62. Manning, M.C., Patel, K., and Borchardt, R.T. (1989). Pharmaceutical Res. 6:903-918. Anantharamaiah, G.M., Hughes, T.A., Iqbal, M., Gawish, A., Neame, P.J., Medley, M.F., and Segrest, J.P. (1988). ] Lipid Research. 29:309-318. Oda, T., and Tokushige, M. (1988). J Biochem. 104:178-183. Teh, L.C., Murphy, L.J., Huq, N.L., Sums, A.S., Friesen, H.G., Lazarus, L., and Chapman, G.E. (1987). ] Biol Chem. 262:6472-6477. De la Llosa, P., El Abed, A., and Roy, M. (1980). Can] Biochem. 58:745-748. Finch, P.W., Rubin, J.S., Miki, T., Ron, D., and Aaronson, S.A. (1989). Science. 245:752-755. Ron, D., Bottaro, D.P., Finch, P.W., Morris, D., Rubin, J.S., and Aaronson, S.A. (1993). ] Biol Chem. 268:2984-2988. Perkett, E.A. (1995). Current Opinion in Pediatrics. 7:242-249. Ulich, T.R., Yi, E.S., Cardiff, R., Yin, S., Bikhazi, N., Biltz, R., Morris, C.F., and Pierce, G.F. (1994). Amer J Pathol. 144:862-868. Staiano-Coico, L., Krueger, J.G., Rubin, J.S., D'limi, S., Vallat, V.P., Valentino, L., Fahey III, T., Hawes, A., Kingston, G., Madden, M.R., Mathwich, M., Gottlieb, G., and Aaronson, S.A. (1993). / Exp Med. 178:865-878. Rubin, J.S., Osada, H., Finch, P.W., Taylor, W.G., Rudikoff, S., and Aaronson, S.A. (1988). Proc. Natl. Acad. Sci. USA 86:802-806. Hsu, Y-R., Hsu, E. W-J., Katta, V., Brankow, D., Tseng, J., Hu, S., Morris, C.F., Kenney, C.W., and Lu, H.S. (1996). Biochem]. (in press).

SECTION IV Posttranslational and Other Modifications

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Effects of Enzyme Glycosylation on the Chemical Step of Catalysis, as Probed by Hydrogen Tunneling and Enthalpy of Activation Amnon Kohen, Thorlakur Jonssoii'* and Judith P. Klinman * Department of Chemistry, University of California, Berkeley, CA 94720

I. Introduction The effect of protein glycosylation on enzyme activity is poorly understood. Recent reports suggest that glycosylation does not change the enzyme conformation but does reduce the protein dynamic fluctuations (cf. Rudd et al., 1994). This effect may inhibit catalytic activity although it is not clear which kinetic steps will be predominantly affected. Most activity assays can not distinguish between effects on the chemical step and effects on internal enzymesubstrate rearrangements or product release. Hydrogen tunneling should be very sensitive to any change in the barrier crossing process for the chemical step. The contribution of quantum-mechanical tunneling to the hydrogen transfer step was demonstrated in the past with several enzymes (for review see Bahnson & Klinman, 1995) and most recently for lipoxygenase (Jonsson et al., 1996). In this study, we have investigated the effect of glycosylation on the tunneling contribution to the hydrogen transfer step in the glucose oxidase (G0)1 catalyzed reaction. Our results suggest that the degree of tunneling is affected by changes in the polysaccharide envelope on the surface of the protein. After establishing that the chemical step is largely rate limiting with [l-^H]-2-deoxyglucose as a substrate, enthalpies of activation have been measured with different glycoforms. A decrease in AH* is found to correlate with an increase in tunneling. Glucose oxidase (EC 1.1.3.4.) catalyzes the oxidation of glucose to gluconolactone and the subsequent reduction of oxygen to hydrogen peroxide in accordance with a ping-pong steady state kinetic mechanism (Pazur & Kleppe, 1964; Crueger & Crueger, 1990): ^.,

glucose oxidase

HO^^.L-0 HO

HO

FAD ^ H

^

FADH2

H.n. H2O2 > ^ ^ ^ ^ X ^ 0oo 2

HO^X-Ov HO

^O

* Current address: Kairos Scientific, Bldg. 62, 3350 Scott Boulevard, Santa Clara, CA 95054. Author to whom correspondence should be addressed. 1 Abbreviations: GO, glucose oxidase; KIE, kinetic isotope effect; TLC, thin-layer chromatography; kD, kilodalton; dd, doublet-doublet; ax, axial; eq, equatorial.; M.W., molecular weight. TECHNIQUES IN PROTEIN CHEMISTRY VIII Copyright © 1997 by Academic Press All rights of reproduction in any form reserved.

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The enzyme contains one very tightly, but noncovalently, bound FAD cofactor per monomer and is a homodimer with a molecular weight of 130-320 kD, depending on the extent of glycosylation. Previous reports suggest that the hydrogen transfer step is not fully rate limiting under steady state conditions for the oxidation of glucose (Bright & Gibson, 1967; Bright & Appleby, 1969; Kriechbaum et al., 1989). Consequently, anomerically protonated, deuterated and tritiated 2-deoxyglucose were used as substrates in the present study. Our results indicate that the protonated substrate is not fully rate limited by hydrogen transfer (see Results) but that deuterated and tritiated substrates can used as direct probes of the chemical step of the sugar oxidation. II. Materials and Methods A. Materials [l-3H]-2-Deoxyglucose (10 Ci/mmol), 2-deoxyglucose (grade III), glucose oxidase (EC 1.1.3.4.) fvom Aspergillus niger (VII-S, for synthesis only) and all salts and buffers (unless otherwise indicated) were from Sigma. [UL-14c]-2Deoxyglucose (255 mCi/mmol) was from American Radiolabeled Chemicals, Inc. Bis-tris propane was from Calbiochem. D2O (99.9% and 99.996%) was from Cambridge Isotope Laboratories or Aldrich. Sodium amalgam (5%) was from Anachima. Glucose oxidase from Aspergillus niger (grade I, 211 U/mg), hexokinase, endoglycosidase-H and a-mannosidase were from Boehringer Mannheim. The recombinant glucose oxidase from Aspergillus niger expressed in yeast (pSGO-2 plasmid - Frederick et al., 1990) was a generous gift of Dr. Steven Rosenberg from Chiron Co (Emeryville, CA). The pH of all buffers was adjusted at the experimental temperature. Synthesis of Anomerically Deuterated 2-Deoxyglucose [l-2H]-2-Deoxyglucose was synthesized from 2-deoxyglucose by oxidation to 2-deoxygluconolactone catalyzed by GO, followed by sodium amalgam reduction in D2O to yield the deuterated product. 2-Deoxyglucose (1 g; 3.78 mmol) was dissolved in 1 mL of 0.2 M potassium phosphate buffer pH 5.6. 155 mg of GO (25000 unit, type VII-S, containing - 1 % catalase) was added and the reaction was stirred gently under oxygen until all the reactant was converted to product (as detected by TLC [7:1 ethyl acetate:methanol - stained by "Yellow spray": 12g (NH4)6Mo7024-4H20 and 0.5g cerric ammonium nitrate in 300 ml 10% H2SO4, heated]). 25 mL of acetonitrile was added and the aqueous phase was re-extracted twice with acetonitrile. The organic phases were pooled and dried under reduced pressure and then lyophilized twice from 50 mL D2O (99.9%) to exchange all exchangeable hydrogens. The produced lactone, 0.894 g (3.38 mmol - 89.5% yield from previous step) was dissolved in 130 mL of D2O (99.996%) in a three-neck flask equipped with a mechanical stirrer and stirred under argon for 10 min. Reduction to [l-2H]-2-deoxyglucose was performed by a modification of the procedure of Isbell et al., (1962). The flask was cooled to -2 °C in a ice-salt bath and 10 g of NaHC204 (lyophilized twice from D2O) and 23 g of sodium amalgam (5%) were successively added. The reaction mixture was stirred under Ar and the temperature was raised to 4 °C. After all the lactone was converted to sugar (~ 1 h, by TLC detection), the pH was raised to 10 by addition of NaOD and 1 L of methanol was then added. The mercury and the precipitated salts were filtered through a fine sinter-glass and the volume was reduced to 5 mL. 500 mL Methanol was added and more salts were filtered out. The product was passed through a mixed ion-exchanger (BioRad, TMD-8; H+ZHCOs" form)

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until the conductivity was similar to that of distilled water (which served as eluent). The product was lyophilized and then crystallized twice from hot ethanol to yield 0.502 g (50.2% overall yield). No anomeric protons were detected by ^HNMR (assigned as dd peak at 5.23 ppm and dd peak at 4.78 ppm for the protonated sugar) which also revealed the loss of the vicinal anomeric /-coupling at the 2-deoxy protons (/i-2ax = 9.8Hz at 6 = 1.37 ppm, J\.2ax = 3.2Hz at 5 = 1.58, J\-2eq < IHz at 8 = 1.98 and /i.2eq = 3.2Hz at 6 = 2.11 ppm as measured for the protonated sugar). Electrospray mass-spectrometry (MNa+ = 188.1) detected no iH-contamination. Micro analysis found C 43.49% (calc. 43.68%) and H 7.54% (calc. 7.33%).

Synthesis of Anomerically Deuterated [UL-^'^C]-2-Deoxyglucose [UL-i^c, l-2H]-2-Deoxyglucose was synthesized from [UL-l4c]-2deoxyglucose according to the strategy described above. All the purification steps were performed by HPLC (Phenomenex CI8-NH2 column eluted with 87% acetonitrile in water). 65 jaCi [UL-l'^C]-2-Deoxyglucose (250 mCi/mmol) was "diluted" with unlabelled 2-deoxyglucose to a specific activity of 8.4 mCi/mmol and dissolved in 170 |iL of 0. IM potassium phosphate pH 7.2. 10 |iL of catalase (5000U) and 8 mg of GO (grade I) were added. The reaction was quenched after 14 min by adding 700 |iL of acetonitrile, and the precipitated protein and salts were spun down (the pellet contained 9 |iCi ^^C). Unreacted reactant and the lactone were separated by HPLC (Altech NH2 column, eluted with 87% acetonitrile, 13% H2O at 2 ml/min) and pure lactone (-30 |iCi) fractions were pooled and lyophilized. Exchangeable hydrogens were exchanged by lyophilizing twice from D2O (99.9%). The lactone was reduced in a similar apparatus to the one described above with 750 mg of NaHC204 (lyophilized twice from D2O) and 2.1g of sodium amalgam 5% in 10 mL of D2O (99.996%). After 45 min, 50 mL of Acetonitrile was added. The mercury and the precipitated salts were removed by filtration through a sinter-glass. The product was purified by HPLC (same conditions as above) yielding 10.8 |LiCi of [UL-l^C, l-^H]-2-deoxyglucose (17% overall yield). The deuterium content of radiolabelled product was not directly determined. In subsequent experiments no trend in D/T isotope effect values was detected with fractional conversion of substrate. The latter has been seen previously with incompletely deuterated substrates (Cha et al., 1989). Deglycosylation of Glucose Oxidase Both wild type (M.W. 155±10 kD) and recombinant (M.W. 260±50 kD) glucose oxidase were deglycosylated enzymatically to the same glycoform (M.W. 136±3 kD). Deglycosylation was conducted under non-denaturating conditions using a modification of the procedure described by Kalisz et al. (1990). Since the recombinant enzyme deglycosylation was much faster, and the deglycosylated enzyme from both sources had the same M.W. and activity parameters (data not shown), the recombinant enzyme was used to produce the deglycosylated GO used below. In a typical experiment, 50 mg of GO was incubated for 24 h in 60 mL of 30 mM potassium phosphate buffer pH 5.0 with 2.6 units of endoglycosidase H and 80 units of a-mannosidase. The reaction was monitored by SDS-PAGE until the GO showed a narrow (< 3 kD) band at 68 kD. The reaction mixture was concentrated by ultrafiltration (Amicon, YM30 membrane), and the buffer was replaced with 100 mM sodium acetate pH 4.5. The mixture was loaded on a cation-exchange column (Mono-S, Pharmacia) and the GO eluted

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with 100 mM sodium acetate pH 4.5. The GO was concentrated by ultrafiltration to 3.4 niL in 0.1 M potassium phosphate, pH 7. The enzyme activity was measured by a continuous spectrophotometric assay (see Methods), active site concentration was determined by FAD absorption at 452 nm (e = 12.83 mM-lcm-l) as described by Frederick et al. (1990) and the protein concentration was measured by the Bradford assay (BioRad reagent) using bovine serum albumin as standard, or by its absorption at 280 nm using a published factor of 1.67 O.D. per mg (Swoboda & Massey, 1965). The specific activity was 430 U/mg, and the overall yield of enzyme, based on active sites measurement (452 nm absorption), was about 40%. B, Methods 1. Competitive Kinetic Isotope Effect (KIE) Experiments In these experiments a tritiated substrate was mixed with I'^C-labeled protonated (for H/T KIE experiment) or deuterated (for D/T KIE experiment) substrate. The mixture was allowed to react in the presence of enzyme, under defined conditions of pH and temperature, and quenched at different fractional conversions. The quenched mixtures were analyzed by HPLC and liquid scintillation counting to determine the fractional conversion (f, determined from the ^^C counting) and tritium to ^^C ratio in the products ([^H/l^C]/ and [3H/1^C]OO for the ratio at the time point and the infinity point, respectively). The L/T KIE (also denoted by: T(V/K)L or ki^/kj) were calculated by equation 1 (Melander & Saunders, 1987): T(V/K)L=ln(l-f)/ln{ l-f[(3H/14C)//(3H/14C)oo]}

(1)

These experiments were carried out in 10 mM bis-tris propane (for pH 9.0) as it was found that at this pH the KIE is largest, suggesting that the hydrogen transfer step is closest to being rate-limiting. Prior to kinetic experiments, tritiated and I'^C labeled substrates were copurified by the same HPLC procedure described above (see Materials). Typically, each experiment contained 1.35 |LiCi ^H and 0.13 |iCi l^c (about 0.2 mM and 0.6 mM total substrate concentration for H/T and D/T KIE experiments, respectively). The reaction mixture (1.6 mL) was pre-equilibrated in a Neslab waterbath (±0.1 °C) and three zero time point (to) of 150 |iL each, were removed. The reaction was initiated by addition of enzyme and in general, seven time points (t^) were removed. After incubation overnight with more enzyme, an infinity time point (too) was taken. All samples were quenched with 1 |LiL of 70% HCIO4. The sample's pH was raised to 8.5 by addition of 2.5 |LiL of 6 M NaOH and all remaining substrate was converted to the 6-phospho derivative by addition of 4 mM MgCh, 0.5 mM ATP, 90 mM Tris/HCl pH 8.5 and 30 units of hexokinase followed by a one hour incubation.^ The coupling reaction was quenched with 0.4 mM HgCl2 and stored at -70 "C until HPLC analysis. Before analysis, each sample was thawed and tetrabutylammonium hydrogen phosphate was added to a final concentration of 5 mM, after which precipitated salts were spun down. The HPLC analysis was performed with a CI8 column (Phenomenex, UltraCarb) equilibrated with 5 mM tetrabutylammonium hydrogen phosphate pH 6.8 and the sample was eluted with 0 to 30% acetonitrile gradient. ^ Derivatization was found necessary for efficient HPLC separation. The quantitive conversion of the unreacted substrate was guaranteed (and tested) by a large excess of hexokinase over initial GO and of ATP over 2-deoxyglucose. The loss of GO activity following acid quenching and neutralization was assayed by an oxygen electrode.

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Tritiated water eluted in the dead-volume (2 min) followed by 2-deoxygluconolactone (5 min), 2-deoxygluconic acid (8 min), and 6-phospho-2deoxyglucose (15 min). Fractions were collected and analyzed using a liquid scintillation counter (Wallac). Since the substrate ^H/^^C ratio at to was equal to the ^H/l^c ratio in the products at too, an average of the three to was used as (3H/14C)OO in equation 1.

2. Initial Velocity Measurements Initial velocity measurements for enzyme activity (through the purification and before kinetic experiments) were performed by a continuous spectrophotometric assay as described by Lockridge et al. (1972) and modifed by Frederick et al. (1990) with a Hewlett-Packard 8452A diode array spectrophotometer equipped with a thermostatted cell holder. Initial velocity measurements in kinetic experiments were carried out by following the oxygen consumption with an oxygen electrode (Yellow Spring Instruments; 1 mL chamber). C. Data Analysis 1, KIE Analysis Analysis of the temperature dependence of KIE's was done by fitting them to equation 2 as described previously (Bahnson & Klinman, 1995 and other cited therein.). kL/kT = AL/Ar*exp[(ET-EL)/KT] (2) where kL/kx is L/T KIE, P^IA.j is the KIE on pre-exponential factors and EJ-EL is the isotope effect on the enthalpy of activation. The fitted parameters enable comparison of values for the AL/AT to theoretical limits from semiclassical models. The semiclassically calculated values range from 0.6 to 1.6 for AH/Ay and from 0.9 to 1.2 for AD/AT (Schneider & Stem, 1972; Bell, 1980; Melander & Saunders, 1987). For proper error analysis, curve fitting was carried out as a least root mean square fit exponential regression of KIE vs 1/T (using the software KaleidaGraph). Each experimental point (as shown in figure 1) was an average of two or three independent experiments with at least six time points and three zero time points per experiment. 2. Initial Velocity Data Analysis KM and kcat and their standard errors were determined by nonlinear least root mean square fit to equation 3 (using KaleidaGraph). v/[E]=kcat*[S]/(KM+[S])

(3)

where v is the initial velocity, [E] is the enzyme concentration and [S] the substrate concentration.

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III. RESULTS A. Competitive KIE Temperature Dependence The H/T and D/T KIE's for the three GO glycoforms were measured with the substrate 2-deoxyglucose (as described under Methods). The temperature was varied from 0 to 45°C and an example of representative results for the wild type enzyme are presented in Figure 1.

0.0033 0.0036 1/T [K-^] Figure 1. H/T (•) and D/T (A) KIE temperature dependence for the wild type GO. Data presented as KIE Arrhenius plots (see explanation and description under Methods). All the experiments were carried out in lOmM bis-tris propane buffer pH 9.0 as described under Methods.

The D/T KIE values were relatively larger than expected from the deuterium and tritium zero point energy differences. The H/T KIE values were large, but not as large as expected from semiclassical (no tunneling) calculations (Swain et al., 1958; Saunders, 1985). kn/kx = (fe/kT)^-26-3.34

(4)

where kn/kx and kp/kx are H/T and D/T KIE, respectively. The observed trend is kn/kx <(te/kx^-26-3.34

(5)

As discussed previously (Klinman 1995 and other cited therein), such a trend suggests that some kinetic complexity "masks" the intrinsic H/T KIE as discussed below in more details. Table I summarizes the KIE's on Arrhenius pre-exponential factor (AH/AX and AD/Ax) and their standard experimental error calculated from a nonlinear least root mean square fit (see data analysis under Methods) for the different GO glycoforms. Additionally, the ratio of ln(H/T) and ln(D/T) (the exponent of equation 4) at 25°C has been included. The different GO glycoforms are denoted by their molecular weight.

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Table I KIE's on the Arrhenius pre-exponential factor and the exponents relating H/T and D/T KIE at 25°C M.W. [kD]

AH/AX ^

AD/AT ^

exponent^

136

0.84 (±0.34)

1.47 (±0.09)

2.89 (±0.04)

155

1.28 (±0.18)

1.30 (±0.10)

2.93 (±0.05)

260

1.46 (±0.20)

0.89 (±0.04)

2.93 (±0.04)

a. The semiclassically calculated values range from 0.6 to 1.6 for A H / A J . b. The semiclassically calculated values range from 0.9 to 1.2 for AD/AX. c. See equation 4.

Values for AH/Ax are all within their semiclassical limits (see KIE analysis under Methods). On the other hand, the AQ/AX values for the three glycoforms differ significantly. The value for the heaviest GO glycoform (260 kD) is close to the lower semiclassical limit (0.9), while that for the wild type enzyme (155 kD) is shghtly above the upper semiclassical limit (1.22) and the factor for the lightest, deglycosylated form (136 kD) is clearly and significantly above this limit. This observation is quite rare but was predicted and observed in the past when substantial amount of tunneling was incorporated or expected. This issue is discussed in more detail elsewhere (Kohen, et al., 1996). B. Kinetic Complexity When more than one kinetic step is rate limiting in an enzymatic turnover, the reaction is said to be kinetically complex. The role of kinetic complexity in the glucose oxidase catalysis, was studied by calculating the ratio between the ln(H/T) and ln(D/T) KIE (Table I) which is the exponent term of equation 4. Through the temperature range studied, the distribution of these exponent values was rather narrow (2.82 to 2.98 and no more than 1.4% change for a single glycoform) suggesting a very small temperature dependence for the kinetic complexity. Moreover, the small trend in exponents with temperature can be shown to lead to H/T KIE's on the Arrhenius prefactors smaller than the intrinsic values for the lighter glycoforms and larger than the intrinsic value for the heavy one. This is a trend that further emphasizes the differences between them (Kohen et al., 1996). However, as the D/T KIE measurements are likely to be virtually free of kinetic complexity, the AQ/AX values reported in table I reflect the chemical step of the GO reaction. C. Enthalpy of Activation for [l'^H]-2-Deoxyglucose The enthalpy of activation for the oxidation of [l-2H]-2-deoxyglucose was determined with the three GO glycoforms. When using anomerically deuterated 2-deoxyglucose, the chemical step is rate limiting as discussed above. This substrate was synthesized on a gram scale (see Materials). Initial velocities for its oxidation catalyzed by the three GO glycoforms were determined at different temperatures by following the oxygen consumption using an oxygen electrode (see Methods). Determination of enthalpy of activation requires measurements of kcat under full substrate saturation over a significant temperature range. The initial velocity measurements were performed under pure oxygen atmosphere and with 0.5 M 2deoxyglucose (> 8*KM) in 10 mM bis-tris propane buffer pH 9 as described under

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Methods. At high temperatures, saturation of the enzyme with oxygen under atmospheric pressure could not be achieved. However, since GO exhibits a pingpong steady state kinetic mechanism (Swoboda & Massey, 1965; Bright & Gibson, 1967), the KM'S of 2-deoxyglucose and oxygen were determined at 25 °C and 45 °C and a simple Michaelis factor was applied to correct the Arrhenius slope: kcat = Vniax.obs. * (KM + [S]) / ([E] * [S]) (6) where Vmax.obs. is the observed Vmax^ [S], the oxygen concentration, KM is the measured Michaelis constant and [E] the enzyme concentration. The overall correction on the Arrhenius slope was less than a factor of 1.14 for all glycoforms and the trend among them was not affected. The enthalpies of activation thus determined are 8.1 (±0.36), 11.0 (±0.29) and 13.7 (±0.31) kcal/mole for the 136 kD, 155 kD and 260 kD glycoforms, respectively. IV. DISCUSSION The most significant results of this study are sunmiarized in Table II. Table II: D/T KIE's on the Arrhenius Pre-Exponential Factors and Enthalpies of Activation for [1-^H1-2-deoxyglucose with Glucose Oxidase Glycoforms^ M.W. (kD)

AD/AT^

AHD* (kcal/mole)

260

0.896 (±0.041)

13.64 (±0.31)

155

1.302 (±0.100)

11.06 (±0.29)

136

1.462 (±0.089)

8.18 (±0.39)

a. All the experiments were carried out under the same buffer and pH conditions (see Methods). b. The semiclassical calculated range is 0.9 to 1.22 (see text).

Two striking points arise from this table. First, the D/T KIE's on the Arrhenius pre-exponential factor (AD/AT) for the heavy GO glycoform is close to the lower limit of the semiclassical range. On the other hand, this factor is slightly above the upper semiclassical limit for the 155 kD glycoform and well above this limit for the light glycoform. Second, there is a correlation between a larger preexponential factor and a lower enthalpy of activation. Larger than unity KIE's on the Arrhenius pre-exponential factors are an indication of extensive tunneling and are expected to be accompanied by a lowered enthalpy of activation (Jonsson et al., 1996). Therefore, it appears that the degree of glycosylation on the surface of the protein has changed the nature of the chemical step of the enzyme catalysis. Apparently, more tunneling is involved in catalysis when the polysaccharide envelope size is reduced. These effects are discussed in detail elsewhere (Kohen et al., 1996). Acknowledgment We thank Dr. Steven Rosenberg (Chiron Co. Emeryville, CA) for the generous gift of recombinant glucose oxidase.

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References Bahnson, B. J., & Klinman, J. P. (1995) Meth. Enzymol 249, 373-397. Bell, R. P. (1980) The Tunneling Effect in Chemistry, Chapman & Hall, London & New York. Bright H. J., & Gibson, Q. H. (1967)7. Biol. Chem. 242, 994-1003. Bright, H. J., & Appleby, M. (1969) /. Biol. Chem. 244, 3625-3634. Cha, Y., Murray, C. J., & Klinman, J. P. (1989) Science 243, 1325-1330. Cruger, A., & Cruger, W. (1990) in Microbial Enzymes and Biotechnology (Rogarty, W. M., & Kelly, C. T. Ed.) Elsevier Applied Science, London, pp 177-227. Frederick, K. R., Tung, J., Emerick, R. S., Masiarz, E.F., Chamberlain, S. H., Vasavada, A., Rosenberg, S., Chakraborty, S., Schopter, L. M., & Massey, V. (1990) /. Biol. Chem. 265, 3793-3802. Gibson, Q. H., Swoboda, B. E. P., & Massey, V. (1964) /. Biol. Chem. 239, 3927-3934. Glickman, M. H., Wiseman, J. S., & Klinman, J. P. (1994) /. Am. Chem. Soc. 116, 793-794. Hecht, H. J., Kahsz, H. M., Hendle, J., Schmid, R. D., & Schomburg, D. (1993) /. Mol. Biol. 229, 153-172. Hwang, C. C , & Grissom, C. B. J. Am. Chem. Soc. 116, 795-796. Isbell, H. S., Holt, N. B., & Frush, H. L. (1962) in Methods in Carbohydrate Chemistry (Whistler, R. L., & Wolfarm, M. L. Ed.) Academic Press, New York, pp 276-280. Jonsson, T., Glickman, M., Sun, S., & Klinman, J.P. (1996) J. Am. Chem. Soc. in press. Kalisz, H. M., Hecht, H. J., Schomburg, D., & Schmid, R. D. (1990) J. Mol. Biol. 213, 207-209. KaUsz, H. M., Hecht, H. J., Schomburg, D., & Schmid, R. D. (1991) Biochim. Biophys. Acta 1080, 138-142. Kohen, A., Jonsson, T., & Khnman, J.P. (1996) submission to Biochemistry. Kriechbaum, M., Heilman H. J., Wientjes, F. J., Hahn, M., Jany, K. D., Gassen, H. G., Sharif, F. & Alaeddinoglu, G. (1989) FEBS Lett. 255, 63-66. Lockridge, O., Massey, V., & Sullivan, P. A. (1972) J. Biol. Chem. 247, 8097-8106. Melander, L., & Saunders, W. H. (1987) Reaction Rates oflsotopic Molecules, Krieger, R. E., Fl. Muller, D. (1928) Biochem. Z 199, 136-170. Pazur, J. H., & Kleppe, K. (1964) Biochemistry 3, 578-583. Rudd, P. M., Joao, H. C , Coghill, E., Fiten, P., Saunders, M. R., Opdenakker, G., & Dwek, R. A. (1994) Biochemistry 33, 17-22. Saunders, W. H. (1985)/. Am. Chem. Soc. 107, 164-169. Schneider, M. E., & Stern, M. J. (1972) J. Am. Chem. Soc. 94, 1517-1522. Swain, C. G., Stivers, E. C , Reuwer, J. F., & Schaad, L. J. (1958) /. Am. Chem. Soc. 80, 58855893. Swoboda, B. E. P., & Massey, V. (1965) / Biol. Chem. 240, 2209-2215.

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Profile Analysis of Oligosaccharides from Glycoproteins by PMP Labeling. Comparison of Chemical and Enzymatic Release Methods Using RP-HPLC and Mass Spectrometry Hanspeter Michel Yuemei Ma Barbara DeBarbieri, and Yu-Ching E. Pan Dept. of Analytical Research and Development Hoffmann-La Roche Inc. Nutley, NJ07110

I. Introduction Characterization of carbohydrates in glycoproteins has become an important task in the field of biotechnology, since a number of protein biopharmaceuticals are glycosylated. To gain detailed structural information, it would be advantageous to have a routine profiling method that can provide suitable samples for further analysis. Chemical or enzymatic methods can be employed to release intact oligosaccharides (1). Hydrazine is a commonly used reagent in the chemical release of both N- and 0-linked oligosaccharides (2,3). An instrument that performs automated hydrazinolysis, as well as purification and recovery of oligosaccharides, is commercially available. In the case of enzymatic treatment, peptide N-glycosidase F (PNGase F or N-glycanase) is the most commonly used enzyme; it can release virtually all known intact N-linked oligosaccharides. Among various electrophoretic and chromatographic methods that give reliable profile results of the released oligosaccharides, reversed-phase (rp) HPLC offers an advantage: since the system usually uses volatile or salt-free buffer, the recovered samples are suitable for further analysis without additional manipulation. Unfortunately, oligosaccharides lack a hydrophobic domain as well as a chromophore for sensitive detection. In order to circumvent TECHNIQUES IN PROTEIN CHEMISTRY VIII Copyright © 1997 by Academic Press All rights of reproduction in any form reserved.

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this disadvantage, Honda, et al. (4) have demonstrated that oligosaccharides labeled with l-phenyl-3-methyl-5-pyrazolone (PMP) are suitable for rp-HPLC analysis with conventional UV detection. And, as a result, two analysis kits are now available commercially to perform this method routinely (5,6). One kit performs enzymatic release of N-linked oligosaccharides followed by PMP labeling, and the second kit provides chromatographic conditions for separation of these labeled oligosaccharides. With the use of fetuin as a model system, a study was initiated to examine whether the PMP-labeling and rp-HPLC approach is also suitable for analyzing chemically released oligosaccharides. The recovered PMP-oligosaccharide samples from rp-HPLC were also analyzed by MALDI TOF and LC ESI MS.

II.

Materials and Methods

Bovine fetuin was from Sigma (St. Louis, MO). All chemicals and columns used for hydrazinolysis by automated chemical release were from Oxford GlycoSystems (Rosdale, NY). The N-linked Oligosaccharide Release and Labeling kit-PMP and Oligosaccharide HPLC kit-PMP were purchased from the Perkin-Elmer Corporation Applied Biosystem division (ABI, Foster City, CA). PNGase F was from New England BioLabs Inc. (Beverly, MA). Other chemicals and solvents were of the highest purity available. A.

Chemical release

Chemical release was performed with the use of an automated GlycoPrep 1000 instrument (Oxford GlycoSystems). Two release modes, namely N+0 and O, were performed using the programmed methods. Fetuin (250 \xg) was first dialyzed against 0.1% TFA, dried under vacuum and then subjected to hydrazinolysis. Fetuin was reacted with hydrazine under defined conditions of temperature and time: N+0 mode, which was performed at 95° C for 4 hr, released both N- and 0-linked oligosaccharides, and O- mode, which was performed at 60° C for 5 hr, released 0-linked oligosaccharides. The entire process, including solvent wash and column elution, took approximately 22 hr. B.

Enzymatic release

All the reagents included in the N-linked oligosaccharide release and labeling kitPMP were used for the treatment. The enzymatic reaction of 100 |ig of denatured fetuin sample was carried out at 37 °C for 2 hr using PNGase F from New England BioLabs. C.

PMP labeling of released oligosaccharides

PMP labeling was performed according to the instructions provided in the

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release and labeling kit-PMP. Briefly, the oligosaccharide samples released by either chemical or enzymatic treatments were reacted with PMP reagent in a sodium hydroxide solution at 70 °C for 120 min., acidified and extracted with ethyl acetate to remove excess PMP reagent. D.

RP-HPLC analysis of PMP-labeled oligosaccharides

The HPLC system used for the analysis of PMP-oligosaccharides included a Waters WISP autosampler and an ABI Model 120 HPLC system. ABFs oligosaccharide HPLC kit-PMP was used for the separation of PMPoligosaccharides. The kit included a C-18 reversed-phase column, buffer A (10% acetonitrile, 10% 1 M ammonium acetate, pH 5.5) and buffer B (25% acetonitrile, 10%) 1 M ammonium acetate, pH 5.5). Elution of PMPoligosaccharides was monitored at UV 245 nm with the use of a PE Nelson Turbochrom data system (Cupertino, CA) for data acquisition and analysis. E.

Mass spectrometric analysis

MALDI TOF (7) mass spectra were recorded on a Bruker Reflex instrument (Billerica, MA). The rp-HPLC fractions containing PMP labeled oligosaccharides were dried in a SpeedVac concentrator (Farmingdale, NY) and redissolved in water/acetonitrile (75/25, v/v). 2,5-Dihydroxybenzoic acid (DHB) was used as a matrix. Normally, 0.3 j^L of a half-saturated solution of DHB in water/acetonitrile/trifluoroacetic acid (70/30/0.1, v/v) was mixed on the sample dish with 0.3 |iL of the sample solution. Desorption and ionization was done with a nitrogen laser (337 nm) adjusted to minimum laser attenuation. Mass spectra (10 to 20) were accumulated in linear negative ionization mode with an acceleration voltage of 25 kV. Calibration was done externally with bovine insulin and angiotensin I. LC ESI (8) mass spectra were recorded on a Finnigan TSQ700 instrument (San Jose, CA) in positive ion mode. A sample aliquot was injected into a fused silica microcapillary column with an inside diameter of 100 f^m. The microcapillary was filled at the end with 10 cm of a C-18 reversed phase resin. PMP labeled oligosaccharides were eluted at ca. 1 |iL/min directly into the electrospray ionization source with a 10 min gradient of acetic acid in H2O (0.5 %), v/v) to 80 % acetonitrile. Determination of experimental molecular weights was done with the deconvolution software provided by the manufacturer.

III. Results and Discussion The strategy used to perform the PMP oligosaccharide profile analysis, with the use of bovine fetuin as a model system, is given in Scheme 1. Bovine fetuin, a major glycoprotein in fetal calf serum, has been widely used as a model for the study of glycoprotein structure. This glycoprotein contains both N- and 0linked oligosaccharides and detailed structural information is available (9-11).

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Glycoprotein

Chemical release by GlycoPrep 1000

Enzymatic release by PNGaseF

Oligosaccharides labeled with PMP

Reversed phase HPLC

Scheme 1. Strategy for PMP oligosaccharide profile analysis of glycoproteins.

Profile analysis: Typical profiles of PMP-oligosaccharides released from fetuin by the two chemical treatments, namely N+0 and O modes, and enzymatic treatment using PNGase F are shown in Fig 1. Comparable profile results were obtained from both chemically and enzymatically released oligosaccharides. The results indicate that PMP labeling can also be used for analyzing chemically released oligosaccharides. Five major oligosaccharide peaks were detected in these profiles. An eariier eluting peak corresponding to free PMP reagent was also detected in all of the profiles. It is believed to be due to insufficient solvent extraction of excess PMP reagent. The identity of peaks 1, 3 and 4 in the PNGase F profile has been reported previously (5). They are the three types of N-linked oligosaccharides in bovine fetuin: peak 1 - tetrasialyl triantennary, peak 3 - trisialyl triantennary, and peak 4 - bisialyl biantennary. Since the retention times of peaks 1,3 and 4 in the N+0 profile (Fig. lA) matched well with those detected in the PNGase F profile (Fig. IC), it is reasonable to assume that they are the same three N-linked oligosaccharides. To verify that this is indeed the case, mass spectrometric analysis was used to fijrther characterize the recovered PMP-oligosaccharide samples (see below). Peaks 2

Oligosaccharide Profile Analysis

325

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Hanspeter Michel et al

and 5, on the other hand, are likely to be the 0-linked oligosaccharides, because they are detected exclusively in the profiles of N+0, and O modes. They did not appear in the PNGase F profile. It was reported previously that there are three types of 0-linked oligosaccharides in fetuin: trisaccharide, tetrasaccharide and hexasaccharide (10,11). This information was usefijl for the identification of peaks 2 and 5 (see below).

Mass spectrometric analysis While MALDI TOP mass spectrometry is widely used for the analysis of unlabeled oligosaccharides (12), LC ESI mass spectrometry is seldom used, because of the lack of a suitable chromatographic system. On the other hand, PMP-oligosaccharides could be analyzed with MALDI TOP as well as with LC ESI mass spectrometry. Pig. 2 shows a comparison of mass spectra obtained from the PMP-oligosaccharide in HPLC peak 3. Within the accuracy of the method, MALDI TOP mass spectrometry showed no difference in the molecular weights between the chemically (Pig. 2A) and the enzymatically (Pig. 2B) released oligosaccharide. Identical molecular weights were also obtained with LC ESI mass spectrometry (Pig. 2C). The experimental average molecular weight of 3211 is in good agreement with the theoretical value of 3211.0. Molecular weights were also determined for the other two N-linked oligosaccharides (HPLC peaks 1 and 4), released either chemically or enzymatically. They also were in good agreement with the theoretical values (Table 1). Together with the intact PMP-oligosaccharide, a component at lower molecular weight was observed (marked in Pig. 2A and 2B with an asterisk). This component was formed through the loss of one PMP molecule (174) during sample storage in the HPLC buffer. Its appearance could be minimized by reducing the time between fraction collection and mass spectrometric analysis. This additional component was not seen in the LC ESI mass spectrum shown in Pig. 2C because the rp-HPLC separated it from the intact PMPoligosaccharide. Shown in Pig. 3 is the result from the analysis of HPLC peak 2 (Pig. 1) with LC ESI mass spectrometry. A major component with a molecular weight of 1004.5 was found. This observed average molecular weight is in good agreement with the theoretical value of the PMP labeled 0-linked trisaccharide (10, 11). The identity of this trisaccharide was fiarther confirmed with the presence of fragment peaks in the mass spectrum (marked in Pig. 3C with an asterisk). These fragments were produced through the ESI process and were a result of a loss of sialic acid and a hexose from the intact PMP-oligosaccharide. In the case of peak 5, the major component found in this sample has a molecular weight of 801.4 (data not shown). The origin of this oligosaccharide is not clear. However, the molecular weight indicated the presence of a component that consists of a PMP-labeled N-acetylneuraminic acid and a hexose. Such a structure has not been described to be present as a 0-linked oligosaccharide (10,11). Although preliminary collision induced dissociation (CID) analysis was

Oligosaccharide Profile Analysis

327

Relative Abundance 250 200 I

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Figure 2. Mass spectrometric analysis of the PMP-labeled N-linked oligosaccharide (HPLC peak 3). MALDI TOF mass spectrum of (A) chemically, (B) enzymatically released oligosaccharide, and (C) deconvoluted LC ESI mass spectrum of enzymatically released oligosaccharide. Asterisks denote fragments due to the loss of one PMP molecule.

Table 1. Mass spectrometric results obtained from the analysis of the five PMP-oligosaccharide peaks

Peak

Oligosaccharide + 2PMP *

Theoretical + 2PMP

1 2 3 4

tetrasialyl triantennary trisaccharide trisialyl triantennary bisialyl biantennary partially known

3502.2 1005.0 3211.0 2554.4 -

1 5

Average Molecular Weight | LC ESI MS 1 MALDI TOF MS chemical 1 PNGase F chemical PNGase F 1 ~ 3500 3501 3500 1 ~ 1004.5 ~ 3211 3210 3211 1 — 2555 2555 2554 1 801.4 -

* the molecular weights reported here are for PMP-oligosaccharides which contain two PMP molecules per oligosaccharide.

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in agreement, further experiments will be necessary to unambiguously establish the exact structure of this component.

IV.

Conclusion Remarks

With bovine fetuin as a model system, the use of PMP labeling in the routine rp-HPLC profile analysis of glycoproteins has been evaluated. Comparable profiles for oligosaccharides can be obtained regardless of whether they are released by automated chemical or manual enzymatic treatments. The PMP-oligosaccharide samples recovered from HPLC are suitable for MALDI TOF and LC ESI mass spectrometric analysis. The presence of all three N-Iinked and one of the 0-linked oligosaccharides was confirmed.

Oligosaccharide Profile Analysis

329

Acknowledgments We would like to thank Dr. H. Chokshi, K. Hollfelder and D. Ciolek for reviewing this manuscript.

References 1. 2. 3. 4. 5. 6. 7. 8.

9. 10. 11. 12.

Chaplin, M. F., and Kennedy (1994) Carbohydrate Analysis, A Practical Approach (IRL Press, Oxford). Takasaki, M., Mizuochi, T., andKobata, A. (1982) in Methods Enzymol. 83, 263-268. Patel, T., Bruce, J., Merry, A., Bigge, C, Wormald, M., Jaques, A., and Parekh, R. (1993) Biochem. 32, 679-693. Honda, S., Akao, E., Suzuki, S., Okuda, M., Kakehi, K., and Nakamura, J.(1989) Anal. Biochem. 180,351-357. Yuen, S. W., Fu, D., Zaidi, I, and O'Neill, R. (1994) ABI Poster Presentations in the Protein Society Annual Meeting. Fu, D. and O'Neill, R. A. (1995) Anal. Biochem, 227, 377-384. Hillenkamp, F., and Karas, M. (1990) Meth. Enzymol 193, 280-295. Hunt, D.F., Alexander, J.E., McCormack, A.L., Martino, P.A., Michel, H., Shabanowitz, J., Sherman, N., Moseley, M.A., Jorgenson, J.W., and Tomer, K.B. (1991) in Techniques in Protein Chemistry II (Villafranca, J.J., ed.), pp. 441-454. Academic Press, New York, USA. Green, E., Adelt, G., Baenziger, J. U. , Wilson, S., and Van Halbeek, H. (1988) J. Biol. Chem. 263, 18253-18268. Nilsson, B., Norden, N. E., and Svensson, S. (1979) J. Biol. Chem. 254, 4545-4553. Edge, A. S., and Sipro, R. G. (1987) J. Biol. Chem. 262, 16135-16141. Harvey, D. J. (1994) American Laboratory, December, 22-28.

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Positive Identification of Glycosylation Sites In Proteins And Peptides Using A Modified Beckman LF 3600 N-Terminal Protein Sequencer

^Xiaomei Lin, ^U Wulf Carson, ^Saber M. A. Khan, ^Clark F. Ford and ^Kristine M. Swiderek ^Beckman Research Institute of the City of Hope, Duarte, CA 91010. Beckman Instruments Inc., Fullerton, CA 92634 ^Department of Biochemistry and Biophysics Department of Food Science and Human Nutrition, Iowa State University, Ames, Iowa 50011

Introduction Posttranslational modifications such as the glycosylation of proteins and peptides can be easily overlooked during conventional Edman degradation. Usually, glycoamino acids appear as blank cycles during a sequencing run. Further sequence analysis is necessary after deglycosylation of the sample in order to assign the phenylthiohydantoin (PTH)- Ser or Thr oligosaccharide (O-linked Sac) derivatives and the PTH-Asn (N-linked Sac). Also, a consensus sequence of Asn-X-Ser (Thr) is an indication for PTH-Asn (N-linked Sac) derivatives (Elhammer et al., 1993). This kind of identification is a time and sample consuming task. Positive identification of the glycosylation sites during N-terminal sequence analysis could become a real advantage. The techniques developed for the LF 3600 N-terminal protein sequencer (Beckman Instr.) provide a fast and efficient way to positively identify Ser (0-Unked Sac), -Thr (0-Unked Sac) and Asn (N-linked Sac) carbohydrate structures during N-terminal sequence analysis (Gooley et al., 1995). Liquid phase anhydrous trifluoroacetic acid (TFA) is used to extract glycosylated, polar amino acid derivatives from the reaction cartridge. These amino acids are then converted into PTH derivatives which can be TECHNIQUES IN PROTEIN CHEMISTRY VIII Copyright © 1997 by Academic Press All rights of reproduction in any form reserved.

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positively identified during on-line chromatography. In addition, different forms of carbohydrate structures at a single amino acid site have different retention times and can be distinguished by further analysis (Gooley et al., 1995). After fraction collection of individual peaks during the chromatography of one sequencing cycle, the structure of oligosaccharides at one specific site can be further examined with subsequent analyses, such as mass spectral analysis, capillary electrophoresis (CE) or high pH anion exchange chromatography (HPAEC). Adding to the flexibiUty of the instrument, it can be switched back to regular mode for high sensitivity protein and peptide sequencing. This report will describe the use of a modified Beckman LF3600 microsequencing system for the positive identification of glycosylation sites of different protein and peptide samples. The analysis of glycopeptides from carcinoembryonic antigen (CEA) will be presented. CEA is a heavily glycosylated membrane bound glycoprotein which has been intensively characterized as an important tumor marker for colonic cancer. The example for a glycoprotein is the starch binding domain (SBD) of enzyme glucoamylase, which is an industrially important exoamylase used for saccharification of liquefied starch from various sources to produce glucose and finally fructose syrups (Belshaw et al., 1993). For both, CEA and SBD, the identification of the glycosylation sites is an important contribution to understand their functionaJity.

Materials and Methods Sample Source The protein asialoglycophorin (Sigma A 9791), which contains Olinked Ser and O-linked TTir carbohydrate structures, is used as a standard for glycoprotein sequencing. Glycophorin (Sigma G-9511) can also be used as standard, but desialation needs to be done prior to coupling in order to remove sialic acid. a-Lactalbumin (Sigma L-6385) is used as a standard for high sensitivity sequencing as well as glycosequencing to compare the two sequencing modes. The glycopeptides analyzed are HPLC fractions of carcinoembryonic antigen (CEA) after digestion with trypsin and chymotrypsin (Swiderek et al., 1993). CEA was isolated and purified from liver metastases of colon tumors as described elsewhere (Pritchard et al., 1976). The enzyme glucoamylase, containing engineered enterokinase recognition site, was expressed and purified from yeast. Following the digestion by enterokinase, the starch binding domain (SBD) which contains 109 amino acid residues is separated by gel filtration and desalted using a spin column (Khan et al., 1996).

Coupling

Reaction

One prerequisite of the glycosequencing technique is the covalent attachment of the glycoprotein to a solid support. For these studies

Glycosylation Sites Using N-Terminal Sequencer

333

Sequelon-AA membrane disks (Perceptive Biosystem GEN92(X)33) were used to attach the carboxyl terminal groups of protein or peptide samples. Sequelon-AA is a polyvinylidene difluoride (PVDF) membrane derivatized with aryl amine groups. To couple the sample, the membrane is prewet with 5 |Lil CH3CN and the same volume of sample of approximately 200 pmole proteins is applied to the disk and then dried. 1 mg of water soluble carbodiimide is dissolved in 100 |il of coupling buffer as coupling reagent (available in the Sequelon-AA membrane kit). After applying 5 |Lil of coupling reagent, the membrane is incubated for 30 minutes at room temperature. The disk is then washed twice with 50% methanol/H20 to remove the extra reagent. After coupling, the membrane is stored at 4°C in an eppendorf tube. When samples are attached via carboxyl groups, the protein or peptide must first be desialated prior to the coupling reaction. This is necessary to prevent the coupling of terminal carboxyl groups of sialated glycoamino acids to the support and to increase the yield of glyco amino acids extracted during the Edman reaction. The desialation procedure is carried out according to Gooley et al. (1995). A cap cut from a 1.5 ml microcentrifuge tube was used as a reaction chamber. After the protein is dissolved in 40 |il of 0.1 M TFA (Pierce), the sample is incubated at 80 XI for 40 minutes. The sample is then dried in a vacuum centrifuge to remove TFA and resuspended in 20% CH3CN/H2O.

Glycosequencing

Analysis

The samples are applied to a modified Beckman LF3600 DT protein sequencing system (Beckman Instruments Inc. #291106). Two cartridge block bottle positions are used for TFA delivery so either gas or liquid phase sequencing can be applied. Thus, the instrument can be easily switched to regular mode of high sensitivity protein sequence analysis. The delivery tubing and the programming of the sequencer were changed in order to deliver Hquid TFA through the valve block. Kel-F valves were added to the second TFA delivery reservoir to prevent corrosion. PTHderivatives of both amino acid and glycoamino acids are chromatographed using a Gold HPLC system equipped with a diode array detector at 268 nm range. The PTH derivatives of amino acids and glycoamino acids are separated by a custom made HPLC column (250 x 2.1 mm, Hypersil) at 50°C with a flow rate of 0.2 ml/min. A new buffer system is used, 25 mM ammonium formate, pH 4.0 as buffer A and 100 % acetonitrile (Burdick & Jackson) as buffer B to separate the different PTH-glycoamino acids as well as the PTH-amino acids. The stock solution for buffer A is made by adding 30% ammonium hydroxide (Aldrich #33,881-8) to 88% Formic Acid (Aldrich #39,938-8) with a final concentration of 250 mM pH 4.0 for ammonium formate and is stored refrigerated. A gradient for the separation of the PTH-derivatives is used according to the recommendation of the manufacturer.

Xiaomei Lin et al

334

Results and Discussions Cycle

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Time (min) Figure 1. (A) HPLC chromatogram of 20 pmole PTH-amino acids using the current glycosequencing method. 19 PTH-amino acids are separated by a modified reverse phase HPLC system, (B) Sequencing results of asialoglycophorin using the glycosequencing method. 200 pmole of sample were subjected to the analysis. The assigned PTH-amino acids of the 14 N-terminal residues of asialoglycophorin are LS*T*T*G/EVAMHT*T*T* S*S*S* (*representing the presence of a glycoamino acid).

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Figure lA shows the chromatogram of 20 pmole PTH-standard using the new chromatography system. The chromatogram represents the separation of a total of 19 PTH-amino acid derivatives except for cysteine. In addition, the new buffer system expands the front region of the gradient for the separation of PTH-glycoamino acids. Figure IB is a representative sequence analysis of the commercially available glycoprotein asialoglycophorin using the described glycosequencing method. 200 pmole of the sample are coupled to a A A-Sequelon membrane as described above prior to Edman sequencing. The first 14 cycles are displayed. Cycles 2, 3 and 4 show peaks of PTH derivatives of Ser (0-linked Sac) and Thr (Olinked Sac). The glyco amino acids are identified by a unique pair of peaks. For example, there are two resolved peaks of PTH-Ser (Sac) with retention times of 6.5 and 6.7 min in cycle 3 representing PTH-Ser (Sac) I and II. Two well separated peaks in cycle 4 represent PTH-Thr (Sac) I and II with retention times of 6.9 and 7.3 min. The peak pairs have been shown to represent stereo enantomers and can be present in different ratios (Gooley et al., 1995). In addition, different retention times of the peak pairs can indicate different saccharide structures. The positive identification of glycosylation sites is still possible after 10 cycles: Thr (Sac) in cycles 11, 12,13 and Ser (Sac) in cycles 14, 15, 16 are still well distinguishable. In contrast, no sequence assignments for cycles 2, 3 and 4 and 11-16 can be made using the conventional sequencing method (data not shown). A high background is observed in all cycles due to a high degree of impurity of the sample. SDS-PAGE analysis of this protein sample shows multiple bands on the gel (data not shown). To compare the glycosequencing procedure with the conventional high sensitivity sequence analysis, a-Lactalbumin is sequenced in both modes. Figure 2A and B show cycles 1-3 and cycles 12-14 from conventional and glycosequence analyses side by side. The features of each cycle appear to be similar for both runs, and the repetitive yields are approximately the same. However, for the conventional high sensitivity sequence analysis, the low level of sample amount required is about 10 pmole as demonstrated in Figure 2C. The lowest amount that can be subjected to glycosequencing is about 50 pmole (Figure 2B). Also, the lower limit of glycoamino acid detection depends on the efficiency of the coupling reaction which can vary for different protein and peptide samples. It should be noted that there is also an upper limit to couple samples to the membrane. For most of the samples, 2(X) pmole has proven to be an optimal amount for covalent coupling and further glycosequence analysis. Using the glycosequencing technique, we were able to identify the glycosylation sites of several samples submitted to us for analysis. Figure 3 shows the sequencing results of SBD domain of glucoamylase. Previous GC-MS studies had shown that only mannose is present in this protein (Khan et al., 1996). About 1 nmole of sample is coupled to the membrane disk. During subsequent glycosequence analysis, peaks with retention times of 6.5 and 7.5 in cycle 4 can be assigned to PTH-Thr (0-linked Sac). One of the retention time is different compared to those assigned for PTH-

Glycosylation Sites Using N-Terminal Sequencer

337

Thr (Sac) in asialoglycophorin, which indicates a different form of carbohydrate structure (Gooley et al., 1995). PTH-Thr (0-linked Sac) can also be identified in cycle 6. TTie carbohydrate structures linked to the Thr in this cycle are even more heterogeneous which is indicated by the presence of three peaks with retention times of 6.5, 6.9 and 7.3 min.

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Time (min) Figure 3. Sequence analysis of SBD domain using the glycosequencing method. Peaks in cycle 4 are PTH-Thr (Sac) with retention times of 6.95 and 7.40 min and those in cycle 6 are PTH-Thr (Sac) with retentiontimesof 6.45,6.95 and 7.25 min, respectively. The sequence of the first 15 amino acids in the peptiide is SCTT*PT*AVAVTFDLT.

338

Xiaomei Lin et al

Examples for peptides containing PTH-Asn (N-linked Sac) have been observed during the sequence analysis of fractions from the tryptic/chymotryptic digest of carcinoembryonic antigen (CEA). Figure 4A shows the sequence analysis of a CEA peptide. 'Rie peak with retention time of 7.0 min in cycle 2 is assigned as PTH-Asn (N-linked Sac). The sequence analysis of a different CEA peptide as an example for Asn (Nlinked Sac) appearing in later cycles is shown in Figure 4B. Asn (N-linked Sac) in cycle 9 with a retention time of 7.0 min is observed. The identification of these glycosylation sites in CEA peptides are consistent with previous mass spectral analyses and deglycosylation experiments (Swiderek et al., 1993).

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Time (min) Figure 4. Glycosequence analysis of different CEA peptides. (A) The analysis of a CEA peptide that contains Asn (Sac) N-linked carbohydrate structures in position 2 is shown: SN*NSKPVEDK. (B) The analysis of a CEA peptide that contains N-linked carbohydrate structures in position 9 is shown. Sequence: FTCEPEAQN^TTY.

Conclusions Several examples using sequencing techniques developed for die positive identification of glycosylation sites of proteins and peptides are

Glycosylation Sites Using N-Terminal Sequencer

339

presented. The analysis is straightforward and less time and sample consuming then previously described sequencing techniques. The technique requires extra steps such as the covalent coupling of the protein or peptide samples prior to the sequence analysis. However, the sequencer modification and operation is easily applied on a routine basis. The HPLC system can be easily adjusted resulting in good separations of PTH-amino acids and PTH-glycoamino acids. In addition, different carbohydrate structures attached to a single amino acid site can be distinguished. The different glycoforms collected from the HPLC can be further analyzed by LC/MS, GC/MS, high performance anion exchange chromatography (HPAEC) and capillary electrophoresis (CE) in order to obtain the information regarding the structures and heterogeneity of the oligosaccharides.

References Elhammer AP, Poorman RA, Brown E, Maggiora LL, Hoogerheide JG and Kezdy FJ (1993) / . Biol. Chem 268, 10029-10038. Gooley A A, Packer NH, Piasano A» Redmond JW, Williams KL (1995) Techniques in Protein Chemistry Vf, 83-90. Khan SMA & Ford CF (1996) in preparation. Pritchard DO & Todd CW (1976) Cancer Res. 36,4699-4701. Reilly PJ (1979) Appl. Biochem. Bioeng. 2, 185-206. Swiderek KM, Pearson CS and Shively JE (1993) Techniques in Protein Chemistry IV, 127-134.

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DEAMIDATION AND ISOASPARTATE FORMATION DURING in Vitro AGING OF A RECOMBINANT HEPATITIS E VACCINE CANDIDATE C. Patrick McAtee and Yifan Zhang Genelabs Technologies, Redwood City, CA 94063

I. Introduction Formation of isoaspartate via the deamidation of asparaginyl residues or isomerization of aspartyl residues constitutes a major source of instability in proteins and peptides (1). Isoaspartate arises through an intramolecular rearrangement that produces a succinimide (cyclic imide) intermediate (2). Spontaneous hydrolysis of the imide occurs with a half-life of several hours and generates an aspartyl residue linked to its C-flanking neighbor through the aspartate p-carbonyl (3). Isoaspartate-bearing proteins and peptides are specific substrates for a widely distributed protein methyltransferase; PIMT (EC 2.1.1.77) that uses S-adenosyl-L-methionine (Adomet) as its methyl donor (4). Several laboratories have become interested in monitoring the accumulation of isoaspartate in purified proteins during in vitro aging at physiological pH and temperature as deamidation and aspartate isomerization can lead to dramatic changes in the biological activity of proteins. Federal stability and purity regulations of protein based pharmaceutical products necessitates that the structure of parenteral molecules be clearly defined and consistent. Less well understood is the effect of deamidation on recombinant vaccine candidates. In evaluating a candidate vaccine for HEV (r62-kDa), it was found that the predicted amino acid sequence contained several potential deamidation sites (5, 6). II. Materials and Methods A.

Measurement of Isoaspartate Formation

Isoaspartate formation was determined by methylation of isoaspartyl residues by PIMT using [^H] S-adenosyl-L-methionine ([^H]SAM) as the methyl donor (Isoquant Kit (Promega)). Quantitation of the [^H] methanol released allowed a precise determination of the isoaspartic content in the protein. For quantitative measurements of isoaspartyl content, r62-kDa or its fragments were methylated for 30 minutes at 30°C and pH 6.8 in the presence of PIMT. Concurrently, a standard peptide (delta sleep inducing protein-DSIP) containing one isoaspartic group was incubated with PIMT under the same conditions. Following incubation of the protein and standard for varying time periods, the reaction was stopped by addition of 0.4M CAPS, pH 10; 5% SDS; 2.2% methanol; 0.01% m-cresol purple. PIMT activity was then quantitated by methanol diffusion into a scintillation cocktail followed by liquid scintillation counting. TECHNIQUES IN PROTEIN CHEMISTRY VIII Copyright © 1997 by Academic Press All rights of reproduction in any form reserved.

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C. Patrick McAtee and Yifan Zhang

Methylation reactions of trypsin digested peptide fragments were performed for 30 minutes at 30°C and were stopped by freezing at -70°C. The samples were thawed and immediately injected for RP-HPLC, followed by liquid scintillation counting of the collected fractions. B.

Tryptic Digestion

r62-kDa protein was adjusted to a concentration of 10 nmole in 50 |j.l in the following buffer: 8M urea in 50 mM Tris-HCl, pH 8.0, 0.5 mM DTT. The reduced protein was Scarboxymethylated, digested with modified porcine trypsin (Promega) at a trypsin to substrate ration of 1:100 (w/w), and incubated at 30°C for 2 hours. The reaction was stopped by the addition of 0.01 volumes of .IM PMSF. The digests were then stored at -70°C until HPLC analysis was performed.

C

HPLC Analysis and Mass Spectrometry

r62-kDa digests were chromatographed on a Vydac C,8 reverse phase column using an ABI Model 41 OB dual syringe pumping system. The flow-rate was maintained at 50 \x\l min and elution achieved using a linear gradient from 0.1% aqueous TFA to 0.1% TFA in acetonitrile. A Carlo Erba Phoenix 20 CU pump was used to deliver a mixture of methoxyethanol and isopropanol (1:1) (v/v) at 50 fil/min which was combined with the column eluent in a post column mixing chamber. An in line flow splitter was used to restrict flow to the mass spectrometer to approximately 10 \\\.l min. Detection was performed immediately following elution from the column at 214 nm using an ABI 759A variable wavelength detector. Mass spectrometric detection of tryptic digested r62-lcDa protein was achieved following post column solvent addition and flow splitting by a VG BioQ triple quadrupole mass spectrometer. Spectra were recorded in the positive ion mode using electrospray ionization. Peptides of interest were repurified off-line using a phosphate buffer system previously described (7). Repurified peptides were desalted and analyzed by LC-MS (ES) and fast atom bombardment analysis mass spectral analysis (FAB-MS). FAB-MS was carried out on repurified peptides using a VG analytical ZAB 2-SE high field mass spectrometer. Z).

Protein/peptide sequencing

Conventional sequencing was carried out on either an Applied Biosystem 477 that was equipped with on-line HPLC's for the identification of the resulting phenylthiohydantoin (PTH) amino acid derivatives. The instrument was operated based upon manufacturer's recommendations and 3 pmol PTH standards were routinely used.

Deamidation and Isoaspartate Formation in Hepatitis E Vaccine

343

III. Results and Discussion A.

Kinetics of Isoaspartate Formation During In Vitro Aging

To determine if r62-kDa accumulates isoaspartate during in vitro aging under mild conditions, a sample of the purified protein was incubated at 30°C for 5 days in a pH 7.5 Tris buffer. The aged r62-kDa was then assayed at a concentration of 120 pmoles for the methyl-accepting capacity in the presence of [methyl-^H] AdoMet and PIMT. A significant increase in methyl incorporation was detected in r62-kDa aged for as little as two days (Figure 1). Methylation of r62-kDa increased with time and reached a plateau of 1.4 moles of CH3/mol of r62-kDa under these conditions. Under identical conditions, control (not aged) r62-kDa could be methylated to only 0.04 mol/mol of r62-kDa.

0.4 0.6 0.8 1 pmoles IsoAsp/pmole r62-kDa

1.2

1.4

Figure 1. Kinetics of Isoaspartate Formation During In Vitro Aging To determine if r62-kDa accumulates isoaspartate during in vitro aging under mild conditions, a sample of the purified protein was incubated at 30°C for 5 days in a pH 7.5 Tris buffer.

344 B,

C. Patrick McAtee and Yifan Zhang Isoaspartate Containing Tryptic Peptides of Aged Recombinant 62-kDa

Samples of control and aged r62-kDa were reduced, alkylated, and then digested with trypsin as described above (Figure 2). Each digest was split into two portions. One portion was subjected to reverse phased HPLC at low pH with uV detection to generate a peptide elution profile (Figure 3, left panels). The second portion was enzymatically methylated with [methyl-^H] Adomet to radiolabel all of the isoaspartate bearing peptides. The labeled peptides were then chromatographed under the same conditions as the first portion (Figure 3, right panels). HPLC of the methylated aged sample indicated the presence of three major methyl-accepting peptides. The control sample yielded a methylation pattern qualitatively similar to the aged samples. As expected, however, the amount of methyl incorporation into the major peaks was considerably less in the control sample compared to the aged sample. 10

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190

200

ETSGVAEEEATSGLVMLCIHGSLVNSYTNTPYTGALGLLDFALELEFRNL 210

220

230

240

250

TPGNTNTRVSRYSSTARHRLRRGADGTAELTTTAATRFMKDLYFTSTNGV 260

270

280

290

300

GEIGRGIALTLFNLADTLLGGLLPTELISSAGGQLFYSRPVVSANGEPTV 310

320

330

340

350

KLYTSVENAQQDKGIAIPHDIDLGESRVVIQDYDNQHEQDRPTPSPAPSR 360

370

380

390

400

PFSVLRANDVLWLSLTAAEYDQSTYGSSTGPVYVSDSVTLVNVATGAQAV 410

420

430

440

450

ARSLDWTKVTLDGRPLDSTIQQYSKTFFVLPLRGKLSFWEAGTTKAGYPY 460

470

480

490

500

NYNTTASDQLLVENAAGHRVAISTYTTSLGAGPVSISAVAVLAPHSALAL 510

520

530

540

550

LEDTLDDYPARAHTFDDFCPECRPLGLQGCAFQSTVAELQRLKMKVGKTR EL Figure 2. Tryptic Peptides of r62-kDa Protein. Tryptic peptide sequences are indicated in tlie figure above the predicted sequence. A lower case a, b, or c following the peptide designation indicates that the peptide was obtained as a mixture of peptides. lUPAC nomenclature is used for amino acid abbreviation.

Deamidation and Isoaspartate Formation in Hepatitis E Vaccine

345

Control

Control

H ,M

°> T 1

pw \\U

J i|...^.4.C:',

['•

""

I"" v"''N'"'"

""T"'

11 Aged

JL^ Figure 3. Isoaspartate Containing Tryptic Peptides of Aged Recombinant 62-kDa Samples of control and aged r62-kDa were reduced, alkylated, and then digested with trypsin as described in the text. Each digest was split into two portions. One portion was subjected to reverse phased HPLC at low pH with uV detection to generate a peptide elution profile (left panels). The second portion was enzymatically methylated with [methyl-^H] Adomet to radiolabel all of the isoaspartate bearing peptides. The labeled peptides were then chromatographed under the same conditions as the first portion (right panels) and detected by radioactivity.

346

C.

C. Patrick McAtee and Tifan Zhang

Isolation and Characterization of Methyl-Accepting Sites

The tryptic peptides of aged r62-kDa were pooled and subjected to reversed-phase HPLC in a pH 6.0, sodium phosphate/acetonitrile solvent system as previously described ( 7 ). Compared to the control sample, the aged samples showed a significant elevation of peptide isoforms which were identical to the controls but did not sequence through asparagine residues when they were adjacent to a glycine, serine, or threonine residue (Table I). In contrast, there was also a decrease in aged samples of peptide isoforms which sequenced through the asparagine residues. These isoforms were also evaluated with the Isoquant assay for methyl-accepting capacity and it was determined that the peptides from the aged samples which did not allow complete sequencing by Edman degradation were readily methylated. Methyl accepting peptides which did not sequence efficiently also gained at least 1 atomic mass unit when examined by mass spectrometry. In a pH 6.0 solvent system, the individual peaks from the pH 2.0 solvent system were resolved into a series of peaks. The initial methyl-accepting site encompassing residues 199-208 resolved into 6 isoforms (a-f) when chromatographed at pH 6.0. These peaks were evaluated by Edman degradation and found to consist primarily of two sequences. Peak d did not sequence beyond residue 203 whereas peak e sequenced through to the end of the peptide. Likewise, methyl accepting peaks 2 and 3 were resolved on a pH 6.0 buffered system yielding similar results. The third methyl accepting peak was unusual in that the methyl accepting isoform did not sequence beyond residue 39. Unfortunately, sequencing yields were low enough in this region that the exact site(s) of deamidation could not be called with confidence. The aged peptide (A3a) gave rise to a methyl accepting species which also was 1.8 atomic mass units larger than that of the intact peptide. The deamidation at residue 60 would seem somewhat obvious as the asparagine residue is flanked by threonine on its carboxyl side. However, deamidation has also been known to occur when asparagine is flanked on its amino terminal side by serine, threonine, and to some extent, lysine (8). The asparagine at residue 41 is preceded by a threonine while another asparagine at residue 70 is preceded by a serine. This strongly suggests that the peptide encompassing residues 24-75 contains two potential deamidations. We have shown that in vitro aging of r62-kDa leads to considerable deamidation and isoaspartate formation primarily at Asn^^'*, and Asn^'*^ It is likely from mass spectrometry data that the asparagine at residue 206 may also be deamidated although it is difficult to verify that this residue is deamidated by sequence analysis. Significant deamidation also occurred at in residues 24-75 of the r62-kDa protein although it was difficult to assess the exact location of these deamidation sites. For all of these sites, the Asn residue is flanked on its carboxyl side by glycine, serine, or threonine, producing linkages that are highly susceptible to succinimide formation in small peptides. As mentioned above, deamidation involving amino terminal adjacent residues may also occur. It has been reported that a several deamidation sites in structured proteins do in fact occur at the same sequence pairs known to be the most labile in short flexible, model peptides (9). Highly flexible regions have also been indicated to play a role in antigenic determination. The three-dimensional structure of r62-kDa has not been determined so there is no detailed information on the local conformation or chain mobility at Asn^^'*, Asn ^^^, and

Deamidation and Isoaspartate Formation in Hepatitis E Vaccine

347

Asn^'*^ However, using the flexibility plot of Karplus and Schultz (10), of the ten most flexible peptides in the 62-kDa sequence, three of these peptides contained the three methyl accepting sites which were described in this work. Table I. Characterization of Methyl-Accepting Peptides* (Observed)

mass A

1087.8

1089.5

+ 1.7

No

1087.8

1087.8

0.0

241-255

Yes

1630.2

1631.4

+ 1.2

241-255

No

1630.2

1630.4

+ 0.2

24-75

Yes

5558.0

5559.8

+ 1.8

24-75

No

5558.0

5558.1

+ 0.1

HPLC Peak

Edman Sequence

Predicted Sequence

Aid

199-203

99-208

Yes

A1e

199-208

99-208

A2c

241-247

A2d

241-255

A3a

24-39

A3b

24-42*

Methyl Acceptor

mass

(Expected)

mass

*Sequencing yields were too low to give meaningful data beyond residue 42 *The methyl-accepting tryptic peptides of aged and control r62-kDa were pooled and repurified as described. The peptides were evaluated for isoaspartate formation by Edman degradation, methylation by PIMT, and FAB-MS. The nomenclature used to describe the HPLC peaks is as follows: "A" indicates the peptide was derived from an aged sample. The numbers 1 ,2, or 3 indicate whether the sample was derived from the first, second, or third methyl accepting HPLC peak using the pH 2.0 buffer system. The small case letters indicate the order of peaks resolved using the pH 6.0 buffer system

348

C. Patrick McAtee and Yifan Zhang

IV. Conclusion Deamidation has been shown to reduce the biological activity of several recombinant biopharmaceuticals. The predicted amino acid sequence of our r62-kDa vaccine candidate contained several potential deamidation "hotspots" in which an asparagine residue was flanked on its carboxyl side by a threonine, glycine, or serine residue or in one example the putative site(s) were flanked by a threonine or a serine on the amino terminal side. By methylation of isoaspartate residues generated through in vitro aging it was determined that three tryptic peptides contained notable isoaspartate accumulation. The presence of isoaspartate in these peptides was confirmed by sequence analysis and mass spectrometry. Whether deamidation plays a role in immune recognition has not been determined. However, alteration of a neutralizing epitope through deamidation at a crucial asparagine (or glutamine) residue could lead to either enhanced immune recognition or quite possibly infectious breakthrough in immunized populations. The 62kDa antigen has been previously shown to elicit protective immune responses in primates after heterologous challenge with live HEV (11). We are currently evaluating the effects of deamidation of the 62-kDa antigen to determine whether important parameters of immune response are affected in vivo. References 1.

Aswad, D. (1995) In Deamidation and Isoaspartate Formation in Peptides and Proteins {Asv/ad, D., Ed.) pp. 1-7, CRC Press.

2.

Bomstein, P. and Balian, G. (1977) Methods Enzymoi. 47, 132-145.

3.

Patel, K. and Borchardt, R.T. (1990) Pharm. Res. 7, 787-793.

4.

Johnson, B.A. and Aswad, D. (1990) In Protein Methylation (Paik, W.K. and Kim, S., Eds.) pp. 195210, CRC Press.

5.

McAtee, C. P., Zhang, Y., Yarbough, P.O., Bird, T., and Fuerst, T.R. (1996) Prat. Exp. Pur. (in press).

6.

McAtee, C.P., Zhang, Y., Yarbough, P.O., Fuerst, T.R., Stone, K.L., Samander, S., and Williams, K.R. (1996) J. Chromatog. B (in press).

7.

Paranandi, M. V., Guzzetta, A.W., Hancock, W.S., and Aswad, D. (1994) J. Biol. Chem. 269, 243253.

8.

Bischoff, R. and Kolbe, H. V.J. (1994) J. Chromatog. B 662,261-278.

9.

Wright, H. T. (1991) Protein Eng 4, 283-294.

10.

Karplus, P. A. and Schulz, G. E. (1985) Naturwissenschaf. 72, 212-213.

Deamidation and Isoaspartate Formation in Hepatitis E Vaccine 11.

349

Fuerst, T.R., Yarbough, P.O., Zhang, Y., McAtee, C. P., Tarn, A. W., McCaustland, K. A., Garcon, N., Spelbring, J., Carson, D., Myriam, F., Lifson, J.D., Slaoui, M., Prieels, J.-P., Margolis, H., and Krawczynski, K., (1996) In Enterically-Transmitted Hepatitis Viruses (Y. Buisson, P. Coursaget, and M. Kane, eds.) La Simarre, Joue-les-Tours (France) pp 384-392.

This Page Intentionally Left Blank

The Isolation and Characterization of Active Site Peptides in Lysyl Oxidase Sophie X. Wang and Judith P. Klinman* Departments of Chemistry and Molecular and Cell Biology University of California Berkeley, CA 94720 Katalin F. Medzihradszky and Alma L. Burlingame Department of Pharmaceutical Chemistry and the Liver Center University of California San Francisco, CA 94143

I. Introduction Lysyl oxidase (LO, EC 1.4.3.13) is an important extracellular, matrix-embedded protein. It catalyzes the oxidative deamination of the e-amino group of lysine side chains in elastin and collagen to form allysine ((DC-aminoadipic-y-semialdehyde) (1). The protein-bound aldehydic functional groups subsequently undergo either self-condensation or condensation with a second, unmodified lysine side chain to form inter- and intrachain crosslinks. These crosslinks confer both strength and altered solubility to the modified proteins. In the case of elastin, the presence of such crosslinks facilitates the return of protein fibers to their original size and shape after stretching (2-4). Since it has a central role in the biogenesis of connective tissues, the abnormal expression of enzyme activity is associated with a number of pathological conditions (5). Due to its important physiologic functions, the nature of the covalently bound cofactor in LO has been the subject of speculation for many years. Both redox cycling assays (6) and enzyme inhibition patterns (7, 8) indicate that LO contains a quinone-like structure at its active site. Resonance Raman studies of a C-terminal, cyanogen bromide cleavage product indicated spectral similarities to pyrroloquinoline quinone (PQQ), leading to the proposal of covalently bound PQQ (9). In Hght of the finding of topa (trihydroxyphenylalanine) quinone (TPQ) in a wide range of copper amine oxidases previously claimed to contain PQQ (10, 11), the presence of TPQ in LO appeared more feasible. However, the large size difference between LO and other amine oxidases, as well as the absence of the TPQ consensus sequence in LO (12), raised the possibility of a different cofactor existing in LO. * To whom correspondence should be addressed. TECHNIQUES IN PROTEIN CHEMISTRY VIII Copyright © 1997 by Academic Press All rights of reproduction in any form reserved.

351

Sophie X. Wang et al

352 PCX)H HCX)C

HIVJ-

HOOC

1:PQQ

In a recent paper, we have documented in detail the isolation and characterization of an active site, cofactor-containing peptide from bovine aorta LO (13). Investigation of this peptide by a combination of Edman sequencing, mass spectrometry and resonance Raman spectroscopy indicates a type of quinocofactor, previously undescribed, which is derived from the crosslinking of a modified tyrosine to the e-amino group of a lysyl side chain (13). This new cofactor is designated as lysine tyrosylquinone (LTQ, structure shown as 3). The present paper will be focused on the methodology of the isolation and mass spectroscopic characterization of the active site, cofactor-containing peptide from bovine aorta LO.

•J?

I

-N-(J;H-C-

HN ^ CH-CH2-CH2-CH2-CI-b-N>

CH2

C=0

I

3: LTQ

IL Materials and Methods A.

Protein Purification and Inactivation

LO was purified from two weeks-old bovine calf aorta by a modification of published procedures (14, 15). The aortas (ca. 600 g) were cleaned and finely ground prior to protein extraction. Extractions and all subsequent procedures were carried out at 4 °C. The ground aortas were extracted with 0.15 M NaCl-16 mM potassium phosphate (KPi) (saline buffer), 16 mM KPi, and 4 M urea-16 mM KPi buffer (pH 7.8), sequentially, in the presence of protease inhibitors PMSF (1 mM) and iodoacetamide (0.04% w/v). The saline buffer and KPi extracts contained negligible activity and were discarded. The urea buffer extracts were pooled (total of 4 to 5 L) and mixed with hydroxyapatite gel (100 g, preequilibrated in 4 M urea-16 mM KPi). After batch elution from hydroxyapatite, the urea-soluble enzyme solution was concentrated (ca. 700 ml) and dialyzed against 16 mM KPi (pH 7.8) buffer.

Active Site Peptides in Lysyl Oxidase

353

To the dialyzed, cloudy enzyme solution an equal volume of 1 M KPi (pH 7.75) was added to further precipitate the urea-soluble proteins. The suspension was centrifuged and the pellet was collected and redissolved in ca. 130 ml of 6 M urea, 0.03 M NaCl, 16 mM KPi (pH 7.8) buffer prior to being loaded on a Sephacryl S-200 column previously equilibrated with 6 M urea, 0.03 M NaCl, 16 mM KPi (pH 7.8). The column was eluted with the same buffer and fractions were collected. Those containing enzyme activity or having an enriched protein species of ~32 kDa as determined by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) (16) were pooled and concentrated. The concentrated eluant (20 ml) was applied to a Sephacryl S-100 column previously equilibrated with 6 M urea, 0.03 M NaCl, 16 mM KPi, pH 7.8. The column was eluted with the same buffer, fractions were collected and those containing only 32 kDa (LO) and 24 kDa species were pooled and concentrated. This is referred to as the "two-banded" protein, and was used without further purification (to reduce protein loss) for phenylhydrazine inactivation (see below). The two-banded enzyme (6.34 mg) has a specific activity of 0.019 U/mg when assayed at pH 7.2 with benzylamine as substrate (17, 18). It was dissolved in 3.43 M urea, 16 mM KPi (pH 8.0) and reacted with 240 nmoles of [I'^C]phenylhydrazine hydrochloride (universallv labeled, 5550 dpm/nmole) for 30 min. A chromophore was formed with a Imax of "^54 nm. The reaction was monitored on a Hewlett-Packard 8452A Diode Array Spectrophotometer by taking a scan every 3-5 min until the formation of the chromophore reached completion. Excess phenylhydrazine was removed by passing the reaction mixture through a Bio-Rad DG-10 desalting column, equilibrated and eluted with 2 M urea, 100 mM NH4HCO3 (pH 8.0). To determine the incorporation of I'^C-labeled phenylhydrazine in the LO protein, a sample of the desalted, radiolabeled two-banded material was analyzed by SDS-PAGE (16). The separating gel contained 10.5% acrylamide using N,N'-diallyltartardiamide as crosslinking reagent. The protein gel was stained with Coomassie Brilliant Blue G - Colloidal (Sigma) and subsequendy cut into 16 slices according to the position of the protein bands. The gel slices were then dissolved in 1 to 5 ml of 2% periodic acid solution, and the radioactivity in each gel slice was quantitated by liquid scintillation.

B.

The Isolation of the Active Site Peptide

The [l^C]-phenylhydrazine-labeled protein {ca. 6 mg, as described above) was lyophilized and redissolved in 3 ml of 6 M guanidine hydrochloride (19, 20). The resulting solution was capped, deoxygenated, flushed with argon and incubated for 30 min at 37 °C. Dithiothreitol was added carefully with a syringe to a 50-fold excess and the solution was flushed with argon and incubated at 37°C for an additional 4 hrs. The solution was then cooled on ice. To the cooled solution, iodoacetamide (44.4 mg) was added to give a slight excess over the estimated free SH groups. The mixture was left in the dark for 1 hr before the excess reagents were removed by desalting with a Bio-Rad DG-10 column, preequilibrated with 2 M urea, 100 mM NH4HCO3, pH 8.0. The reduced and carboxyamidomethylated LO was then lyophilized and redissolved in 100 mM NH4HCO3 (pH 8.0) containing 2 M urea at 37°C, with shaking. The digestion buffer also contained 5 mM CaCl2, which is an absolute requirement for thermolysin activity (21). Proteolytic digestion was initiated by the addition of thermolysin to 4% (w/w) of the total protein weight. A second

Sophie X. Wang et al

354

aliquot of protease was added after 24 hrs. The digestion was stopped after 49 hrs by placing the solution at -70°C (or by addition of EDTA, 2 mM). The enzymatic digest was subjected to high performance liquid chromatography (HPLC) purification to isolate the active site-containing peptides. After the preliminary HPLC purification, those thermolytic peptides which have absorption peaks at 438 nm were lyophilized and redissolved in 50 mM sodium phosphate (pH 8.0). Subdigestion was initiated by the addition of Asp-N endoproteinase to a final concentration of 2.5% (w/w) by weight of the substrate. The digestion mixture was incubated at 37°C with gentle shaking for 19 hrs and later stored at -70°C before it was purified by HPLC. The thermolysin digest was injected onto a Dynamax re versed-phase CI 8 column (5 |im, 300 A, 4.6 x 250 mm), equilibrated with solvent A (0.11% TFA, 5% CH3CN in H2O), on a Shimadzu HPLC system (see Fig. 2). Peptides were, eluted using a linear gradient of 20 to 30% solvent B (0.1% TFA, 80% CH3CN in H2O) over 65 min at a flow rate of 1 ml/min. Elution of peptides was monitored at 214 nm and 438 nm. The Asp-N subdigest of the cofactor-containing thermolytic peptide (e.g., the dominant peak in Fig. 2) was again separated on HPLC by injecting the mixture onto a Vydac reversed-phase C18 column (4.6 x 250 mm), preequilibrated with solvent A (0.3% triethyl-ammonium acetate or TEAA in H2O) and solvent B (0.3% TEAA, 60% CH3CN in H2O) with a ratio of solvent A : solvent B of 95 : 5. This was followed by a linear gradient elution of 5 to 50% solvent B over 60 min at a flow rate of 1 ml/min. Peptide elution was monitored at 220 and 438 nm. The cofactor containing peptides were collected and lyophilized and subsequently used for sequencing, mass spectrometry, and resonance Raman studies.

C.

Mass Spectrometric

Analyses

The electrospray mass spectrum of a LO active site peptide (sample 1) was acquired by an LC/ESIMS (electrospray ionization mass spectrometry) experiment, in which the sample was further purified by microbore reversedphase HPLC and the eluant was directly injected into an electrospray ionization mass spectrometer. An Applied Biosystems (ABI) microbore HPLC system was interfaced with a Micromass BioQ quadrupole mass spectrometer equipped with an electrospray source. The mass spectrometer was scanned in noncontinuum mode over a range of m/z of 350 to 2000 at 5 s/scan. The chromatographic separation was carried out on a C-18 column (ABI, 1x100 mm) equilibrated at 98% solvent A (0.1% TFA/H2O) and 2% solvent B (0.08% TFA/CH3CN). A linear gradient was started immediately after the injection, increasing solvent B concentration by 1% per min. The average molecular weight of the peptide was determinedfromthe doubly and the triply protonated ions detected. The accurate mass measurement was performed on a Micromass AutoSpec SE mass spectrometer using electrospray ionization on the triply charged ion of a cofactor-containing peptide (sample 2). Doubly charged ions of bradykinin (m/z 530.7885) and gramicidin S (m/z 571.3608) were used as calibration standards. Leu-enkephalin (MH+ at m/z 556.2771) was also included to verify the accuracy of the mass measurement. The monoisotopic molecular mass of this peptide was established by the mean [M+3H]3+ value obtained from four separate injections. Elemental compositions of the crosslinked residue were obtained by computer calculations.

Active Site Peptides in Lysyl Oxidase

355

Matrix-assisted laser desorption ionization (MALDI)-high energy collision-induced dissociation (CID) mass spectrum of an active site peptide (sample 4) was obtained on a Micromass AutoSpec SE orthogonal acceleration time-of-flight (TOP) tandem mass spectrometer. The ^^C isobar of the peptide MH"*" ion was selected as the precursor ion, at m/z 1597.6. The collision gas was Xe, and the collision energy was 800 eV. Fragments are labeled according to Biemann nomenclature (22). Peptide bond cleavage with charge retention at the N-terminus yields b fragments, and when the charge is retained at the C-terminus y ions are formed.

III. Results and Discussion A.

Peptide Isolations and Characterization

Unlike bovine serum amine oxidase, which is available in gram quantities and was used as the prototypic system for the establishment of TPQ, a typical preparation of LO yields about 5 mg (starting with about 500 grams of aorta tissue). In an effort to minimize protein loss, purification was stopped after the stage where LO (32 kDa) coelutes with a second protein (24 kDa) from Sephacryl S-200 gel filtration. The two-banded protein was labeled with [I'^CJphenylhydrazine, to yield the expected chromophore for a phenylhydrazone derivative of a quinone structure (Fig. 1). Analysis of the l^C-labeled protein by SDS gel electrophoresis showed almost exclusive incorporation of ^^C into the 32-kDa LO band (13). Therefore, the unreactive 24-kDa band would not affect subsequent procedures since selection of active site-derived peptides from LO was dependent on screening for l^^C in addition to monitoring the UV-Vis absorption of the newly formed chromophore. 0.45

O C i3

&

0.3 H 0.15H

800 Wavelength (nm) Figure 1. [^^Cjphenylhydrazine labeling of the LO. The two-banded LO enzyme (6.34 mg) was dissolved in 3.43 M urea-16 mM KPi buffer (pH 8.0) and reacted with 240 nmoles of [l^Q-phenylhydrazine hydrochloride for 30 minutes. A chromophore was formed at 454 nm. After labeling, the reaction mixture was loaded on a Bio-Rad DG-10 desalting column to remove the excess phenylhydrazine (see text for details).

Sophie X. Wang et al

356 2.56

20

30

40

50

60

Time (min)

Figure 2. HPLC purification of the thermolytic peptide. Thermolytic digest was injected onto a Dynamax reversed-phase C18 HPLC column pre-equilibrated with 0.11% TFA, 5% CH3CN in H2O. Peptides were eluted with an CH3CN gradient and monitored at both 214 nm and 438 nm (see text for details).

The HPLC elution profile of a thermolytic digest of the [I'^Clphenylhydrazine derivative of lysyl oxidase (Fig. 2) showed a large number of 214 nm peptide peaks (top panel), together with a single dominant 438 nm peak eluting at 36 to 37 min (bottom panel). Subsequent determination of radioactivity in the peptide fractions showed the coincidence of l^C radioactivity with the 438 nm peaks. Edman sequencing results showed that the thermolytic peptides were fairly long and that more than one peptide was present in the sample (13). In order to yield short peptides to facilitate the separation and analysis, thermolytic peptides (e.g., the dominant 438 nm peak in Fig. 2) were collected and further digested with Asp-N endoproteinase. The HPLC profile of this subdigestion is shown in Fig. 3. The dominant 438 nm peak (indicated by an arrow in Fig. 3) was collected and subjected to further analysis. Once again, Edman sequencing of the Asp-N digested peptide showed two amino acids in each round, indicating that two peptides were present in the sample (13). Both peptides can be located in the cDNA-derived LO protein sequence and are the same peptides as those observed in the thermolytic sample, except that they are shorter in length (13). This has led to the proposal of two crosslinked peptides, as shown below: Asp - Thr - (Tyr - derivative) - Asn - Ala - Asp Vai - Ala - Glu - Gly - His - (Lys - derivative) - (Ala - Ser)

Active Site Peptides in Lysyl Oxidase

357

0,32 n E c o

<

to

-e

o w n

<

20

30

40

60

Time (min)

Figure 3. HPLC purification of the Asp-N peptide. The Asp-N subdigest was loaded onto a Vydac reversed-phase C18 HPLC column pre-equilibrated with 0.3% TEAA, 3% CH3CN in H2O. Peptides were eluted with an CH3CN gradient and monitored at both 214 nm and 438 nm (see text for details).

Tyr and Lys residues were suggested to have been crosslinked to form the active site cofactor since they were not detectable at the expected recovery level during Edman sequencing. This was further supported by mutagenesis studies which are discussed in the full paper (13).

B. Mass Spectral Analysis of the Active Site Peptides The cofactor-containing peptide from the thermolysin/Asp-N digests (sample 1, Fig. 3) was analysed in an LC/ESIMS experiment and the mass spectrum obtained in this way showed doubly and triply protonated ions at m/z 720.9 and 480.9, respectively (Fig. 4). These values were used to establish the average molecular mass as 1439.71. The ion at m/z 747.4 in Fig. 4 is a double-charged Fe-adduct ion, which is believed to arise from the stainless steel capillary. Subtraction of the residue weights of the known amino acid residues (2Asp, Thr, Asn, 2Ala, Val, Glu, Gly, His) in the peptide, plus masses corresponding to two N-terminal protons and two C-terminal hyckoxyl groups, yielded a mass of 393.45 as the residue-weight for the unknown crosslinking cofactor structure labeled with phenylhydrazine. To calculate the elemental composition of an ion, its mass must be determined to three or four decimal places. The accurate mass measurement was carried out on another cofactor-containing peptide sample (sample 2) isolated

Sophie X. Wang et al

358

from different thermolytic and Asp-N digests. Leu-enkephalin (MH+ at m/z of 556.2771) was included to verify the accuracy of the mass measurement. In four separate injections of this compound, MH+ was measured as m/z 556.2760 (J = 0.0025), which shows an approximately 2 ppm deviation from the calculated mass. The mean [M+3H]3+ for sample 2 (from four separate injections) was m/z 533.2356 (J = 0.0026), which establishes the monoisotopic molecular mass of this peptide as 1596.6833. From the protein sequence, the mass of sample 1 and the CID data, sample 2 is the elongated version of peptide sample 1 with an AlaSer sequence at the C-terminus of the crosslinked Lys residue. This mass value shows an approximately 5 ppm deviation from the calculated m/z value for the corresponding peptide with the proposed structure. Again, subtraction of the residue weights of the known amino acid residues (2Asp, Thr, Asn, 3Ala, Ser, Val, Glu, Gly, His) in this peptide, plus masses corresponding to two N-terminal protons and two C-terminal hydroxyl groups, yielded a mass of 393.1830 as the residue-weight for the unknown crosslinking cofactor structure labeled with phenylhydrazine. A mass of 288.1377 for the crosslinked residue was obtained when the phenylhydrazine label was further subtracted. Within a 10 ppm deviation of this mass value, computer calculations permitted only two compositions which contained an odd number of N-atoms as required for this molecule (nitrogen-rule of mass spectroscopy) using composition limits of l^C 10/30; ^H 15/50; l^N 1/9; l^Q 2/10; ^^s 0/2 (where the numbers indicate the number of a particular atom required/allowed in the structure). The C12H22N3O3S composition shows only 1.7 ppm deviation from the measured mass value. However, there is no other indication of a sulfur atom present in the cofactor. Therefore, the C15H18N3O3 composition (C21H23N5O3 including the phenylhydrazine label), which exactly matches our proposed structure and shows a -10 ppm deviation from the measured mass value, is concluded to be the correct composition. 3+ 480.9

Vaverage = 1439.71 2+ 720.9

£1

<

747.4

illi,4i^.ll).li. ljii.i\l.fLi>l(ilj[Mil.4^.il).iu^iJ4.^l'M.Mi4 400

450

500

550

600

650

•" Iji'it' .in.,..i...i.u 700

750

800

650

900

050

m/z

Figure 4. Electrospray of the LO active site peptide (sample 1). The electrospray ionization mass spectrometer was scanned in noncontinuum mode over a range of m/z of 350 to 2000 at 5 s/scan (see text for details).

Active Site Peptides in Lysyl Oxidase

359

A reasonable structure for the phenylhydrazine derivative of this cofactor, which has the composition of C21H23N5O3, is given below, as the two possible tautomers (13). ''^'*'*'*^ H N—C H ~ C O^'*'^'^^

CHp

I

HN—CH

'

^ Azo form

Hydrazone form

The calculated mass for these tautomers is 393.1801 (monoisotopic mass) or 393.45 (average mass), which shows excellent agreement with the experimental mass of 393.1830 (monoisotopic mass from accurate mass measurement, sample 2) or 393.45 (average mass from LC/ESIMS experiment, sample 1) for the derivatized active site cofactor. The underivatized cofactor itself has a structure of the type shown as 3. This is consistent with the strong evidence for a quinonelike structure in LO (6, 8, 9, 23), and with the conclusion that the cofactor is comprised of a crosslink between a tyrosine derivative and a lysine residue. This structure could arise from an initial hydroxylation of Tyr to form dopa, followed by the oxidation of dopa to dopa quinone, and subsequent nucleophilic attack by the e-amino group of a lysine side chain to generate an aminoquinol (13). One of the peptides was subjected to high energy CID analysis (13). As documented therein, the CID data, together with results obtained with additional peptide samples (data not shown), provide independent evidence for the proposed crosslinked cofactor structure and the crosslinking site in the protein. The immonium ions, b type and y type ions observed in the fragmentation pattern have confirmed the presence of most amino acids in both peptides and supported the proposed cofactor structure (13). In particular, the fragment ion at m/z 902 is present due to cleavages between the a- and P-carbon of the modified Tyr residue (-590 Da) and a cleavage between the aromatic ring and the phenylhydrazine label (-105 Da), with charge retention on the N-terminal fragment. Unusual cleavages were also observed. The ion at m/z 884 seems to be formed via cleavages between the amino group and the a-carbon, and between the a-carbon and the carbonyl group of the Lys residue with charge retention on the crosslinked residue or peptide 1(13). The cleavage at both sides of the peptidyl a-carbon is a very rare event in mass spectral analysis. However, a novel cofactor structure analyzed by a new ionization technique could result in some unexpected fragmentation results. It should be noted that of the two possible nucleophiles in the second peptide. His and Lys, the His appears unmodified; therefore, establishing Lys as the most suitable residue to undergo crosslinking to a modified Tyr residue.

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Sophie X.Wang era/.

Conclusions

In order to confirm the structure for the LO cofactor, model compounds mimicking the proposed enzymic structure were synthesized (13, 24). The UVVis absorption spectrum of the model compound, with a Xmax of 504 nm which is red-shifted compared to other TPQ-containing amine oxidases, corresponded almost exactly to that of native LO (13). Under the same conditions, the resonance Raman spectrum of the phenylhydrazine derivative of the model compound was found to be superimposable with that of the isolated LO peptide labeled with phenylhydrazine, yet very different from the phenylhydrazine derivative of a TPQ model compound (13). As documented, a previously unknown cofactor has been identified in LO (13). It is formed by the crossslinking of two amino acid side-chains and catalyzes the redox reaction in the enzyme (13). This discovery extends the range of quino-structures now demonstrated to function as redox catalysts, provides insights into the biogenetic pathways leading to quinone production from amino acid precursors, and introduces the possibility of regulating LO activity through the design of cofactor-specific inhibitors.

References 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17. 18.

Pinnell, S. R. & Martin, G. R. (1968) Proc. Nad. Acad. Sci. U.S.A. 61, 708-716. Eyre, D. R., Paz, M. A. & Gallop, P. M. (1984) Ann. Rev. Biochem. 53, 717-748. Sanberg, L. B., Soskel, N. T. & Leslie, J. G. (1981) New Engl. J. Med. 304, 556579. Kagan, H. M. (1986) in Characterization and Regulation ofLysyl Oxidase (Academic Press, Orlando, FL), Vol. 1, pp. 321-389. Uitto, J. & Perejda, A. J. (1987) in Connective Tissue Disease (Marcel Dekker, New York). Paz, M. A., Fluckiger, R., Boak, A., Kagan, H. M. & Gallop, P. M. (1991) /. Biol. Chem. 266, 689-692. Buffoni, F., Ignesti, G. & Lodovici, M. (1981) Ital. J. Biochem. 30, 179-189. Gacheru, S. N., Trackman, P. C, Calaman, S. D., Greenaway, F. T. & Kagan, H. M. (1989) /. Biol. Chem. 264, 12963-12969. Williamson, P. R., Moog, R. S., Dooley, D. M. & Kagan, H. M. (1986) /. Biol. Chem. 261, 16302-16305. Janes, S. M., Mu. D., Wemmer, D., Smith, A. J., Kaur, S., Maltby, D. M., Burlingame, A. L. & Klinman, J. P. (1990) Science 248, 981-987. Klinman, J. P. & Mu, D. (1994) Annu. Rev. Biochem. 63, 299-344. Janes, S. M., Palcic, M. M., Seaman, C. H., Smith, A. J., Brown, D. E., Dooley, D. M., Mure, M. & Klinman, J. P. (1992) Biochem. 31, 12147-12154. Wang, S. X., Mure, M., Medzihradszky, K. F., Burlingame, A. L., Brown, D. E., Dooley, D. M., Smith, A. J., Kagan, H. M. & Klinman, J. P. (1996) Science 273, 1078-1084. Kagan, H. M., Sullivan, K. A., Olsson, T. A. & Cronlund, A. L. (1979) Biochem. J. Ill, 203-214. Williams, M. A. & Kagan, H. M. (1985) Anal. Biochem. 149, 430-437. Laemmli, U. K. (1970) Nature 111, 680-685. Trackman, P. C. & Kagan, H. M. (1979) /. Biol. Chem. 254, 7831-7836. Kagan, H. M. & Sullivan, K. A. (1982) Methods Enzymol. 82, 637-650.

Active Site Peptides in Lysyl Oxidase 19. 20. 21. 22. 23. 24.

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Fontana, A. & Gross, E. (1986) in Fragmentation of Polypeptides by Chemical Methods, ed. Darbre, A. (John Wiley and Sons, New York), pp. 67-120. Allen, G. (1989) in Sequencing of Proteins and Peptides (Elsevier Science Publishers B. v.. New York), pp. 58. Wilkinson, J. M. (1986) in Fragmentation of Polypeptides by Enzymic Methods, ed. Darbre, A. (John Wiley and Sons, New York), pp. 121-147. Biemann, K. (1990) Method. Enzymol. 193, 886-887. Williamson, P. R., Kittler, J. M., Thanassi, J. W. & Kagan, H. M. (1986) Biochem. J. 235, 597-605. Wang, S. X., Mure, M. & Klinman, J. P. (in preparation).

Acknowledgments The authors thank Dr. Herbert Kagan and co-workers for assistance leading to the enzyme preparations used for the studies described herein. This work was supported by NIH grants GM39296 to J.P.K. and NCRR BRTP P41 RR01614 to A.L.B., and supported by NSF Biol. Instru. Prog, grant DIR 8700766 to A.L.B.

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Complement activation in EDTA blood/plasma samples may be caused by coagulation proteases Philippe H. Pfeifer Tony E. Hugli Department of Immunology The Scripps Research Institute La Jolla, California 92037

Earl W. Davie Kazuo Fujikawa Department of Biochemistry University of Washington School of Medicine Seattle, Washington 98195-7350

L Introduction

C3a and C4a are important markers of alternative and classical pathway activation, respectively. Both pathways require divalent ions for activation, mainly Mg^^ for generation of the C3 convertase of the alternative pathway and Ca^^ for generation of the CI-complex in the classical pathway. It is known that EDTA, which complexes divalent ions, is a poor stabilizing agent to prevent ex vivo generation of C4a, whereas Futhan (nafamostat mesilate, a powerfiil serine protease inhibitor) is an excellent stabilizing agent in this regard (1-3). On the other hand, heparin, when combined with EDTA, significantly reduces ex vivo generation of C4a which may indicate an involvement of coagulation enzymes in C4 cleavage. Thrombin has been shown to be able to cleave complement factors (4). We hypothesized that other enzymes of the coagulation or fibrinolytic system were responsible for at least part of the ex vivo C4a generation observed in EDTA plasma. TECHNIQUES IN PROTEIN CHEMISTRY VIII Copyright © 1997 by Academic Press All rights of reproduction in any form reserved.

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Even though many coagulation factors require phosphohpids or Ca^^ for activation, we tested the hypothesis that some coagulation enzymes may be able to cleave C3 and/or C4 in a Ca^^- and phospholipid-free environment.

II. Materials and Methods Blood from healthy volunteers was drawn into standard green (heparin, 14.3U/ml) or lavender (lOmM EDTA) top tubes, or into syringes containing Futhan (nafamostat mesilate, Torii Pharmaceutical Co., Tokyo, Japan, 0.2mg/ml final concentration). Either whole blood or plasma, isolated by centrifugation at 4°C/1000xg for 15 minutes immediately after drawing, was kept at 4°C for up to 48 hours to assess generation of C3a and C4a. All of the plasma samples were then frozen at -70 °C until the C3a and C4a levels were measured by RJA (Amersham). Human C3 and C4 were obtained from Advanced Research Technologies (San Diego, CA). 20|ag of C3 or C4 were digested with the respective enzyme for 0 to 90 min in HEPES buffer (50mM HEPES and 150mM NaCl, pH 7.3). The reaction was stopped by adding Futhan at a final concentration of 0.5 mg/ml. The samples were then subjected to Tricine-PAGE (5), stained with Coomassie and analyzed by scanning the visualized bands (Personal Densitometer by Molecular Dynamics). Factor XI and plasma prekallikrein were prepared according to Tait and Fujikawa (6) while p-factor Xlla was prepared according to Fujikawa and McMullen (7). Prothrombin was isolated by the method of Mann (8). Factor X, factor IX and urokinase were the gifts of The Green Cross Co., Osaka, Japan. Factor XI and plasma prekallikrein were activated by trypsin at an enzyme to substrate ratio of 1 to 25 in lOOmM Tris-HCl at pH 8.0. The remaining trypsin was inactivated by aprotinin. Factor IX, factor X and prothrombin were activated by factor XIa, Russell's viper venom factor Xa activating enzyme and factor Xa (1 to 25-50 ratio), respectively, in lOOmM Tris-HCl, pH 8.5/5mM CaCl2. The resulting factor IXa, factor Xa and thrombin were purified by column chromatography using Waters DEAE 15HR resin. The individual samples were applied to the DEAE column that had been equilibrated with 50mM Tris-HCl at pH 8.0 and the proteins were eluted by a NaCl gradient (0 to 0.6M NaCl) in the same buffer. SDS-PAGE analysis demonstrated the complete conversion of the respective zymogens to their active forms. To assess the generation of antigenically active fragments, identical aliquots of C3 or C4 (40|ig/ml in HEPES buffer) were incubated with thrombin (9|i,g/ml), plasma kallikrein (35)Lig/ml), factor XIa (7ng/ml), factor Xa (20ng/ml), tissue-type plasminogen activator (t-PA) plus plasminogen (0.6+350^g/ml), or buffer alone. After 60 minutes the different aliquots were precipitated with the precipitating agent provided with the RIA kit to remove the precursor molecules and C3a and C4a were measured by RIA.

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III. Results Futhan stabilized complement activation in EDTA plasma for extended periods of time at 4°C, as measured by RIA. Background levels of C3a and C4a in stabilized plasma from whole blood drawn into Futhan+EDTA and stored at 4°C for up to 48 hours showed only minimal ex vivo activation (Table I). EDTA effectively inhibited ex vivo generation of C3a, whereas there was an ongoing production of C4a. Heparin also appeared to keep the C3a, but not the C4a, at a low level. Heparin + EDTA was no better in preventing ex vivo C3a generation than EDTA or heparin alone, but yielded a marked improvement in the stabilization of C4a. Indeed, C4a levels remained almost as low with this combination as with Futhan. In whole blood anticoagulated with the same reagents and stored at 4°C for up to 48 hours, a similar tendency as in plasma could be observed. Here too, heparin plus EDTA attenuated the generation of C3a and C4a while heparin alone stabilized C3a levels but allowed a 10- to 20-fold increase in C4a over baseline values. Again, EDTA+Futhan practically inhibited the ex vivo generation of C3a and C4a (not shown). Incubation of C3 and C4 for 60 minutes at 3TC with a number of enzymes involved in coagulation and fibrinolysis indicated that half of the enzymes tested were able to cleave C3 and C4 (Table II). Interestingly, factors IXa and Xlla (pXlla) did not seem to cleave C3 or C4, whereas factors XIa and Xa, thrombin, plasma kallikrein and plasmin clearly degraded both complement factors.

Table I. Generation of C3a and C4a inplasma at 4°C' 24 hours 0 hours 48 hours | 175 201 EDTA 173 201 200 235 C3a (ng/ml) EDTA + Futhan 123 228* heparin 183 171* 271* heparin + EDTA 229* 100 271 499 EDTA 106 131 90 C4a (ng/ml) EDTA + Futhan heparin 433 525 664* 109* heparin + EDTA 125* 134* ^Whole blood was anticoagulated with lOmM EDTA, heparin (14U/ml), heparin + 5mM EDTA or 0.2mg/ml Futhan + lOmM EDTA. Plasma was separated fi"om whole blood by centrifugation for 15 minutes at lOOOxg at 4''C. All samples were kept at 4° for the length of time indicated. Values are averages of two experiments except when marked with an asterisk for single values.

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Table 11. Qualitative differences in the C3 and C4 converting ability of some enzymes involved in coagulation and fibrinolysis^ 1 Enzyme C3 cleavage C4 cleavage Thrombin 4+ Vila + tissue factor + activated protein C Plasma callikrein + + activated XII (p-XIIa) XIa + + Xa + + IXa . . Urokinase t-PA + Plasminogen + + •

II

I

1

1 1 1 1

NV^ith the exception of factor Vila + tissue factor, all experiments were performed in Ca^^- and Mg^^-free buffers. (-: no cleavage, +: cleavage)

|i|BlHiii|if|'-JBiB^

W^iS^^^

|:N:!^^^^j^il;::f;3i|iii|^^Bii|*

i:BiHiHlB

/ lilililii:::;?

Figure 1: Kinetics of C3 degradation by factor Xa and plasmin. C3 was incubated at 200^g/ml for 0, 2, 30, 60 and 90 minutes at 3TC with 44ng/ml of factor Xa (lanes 1-5). Clearly visible are the C3 a- and p-chains having molecular weights of 115 and 75kD, respectively. The band of degraded material visible in lane three has an estimated M.W. of 105-1 lOkD and increases in intensity for 0 to 30 minutes. This pattern, where only the a-chain gets degraded, is similar to proteolysis seen with thrombin, plasma kallikrein and factor XIa (not shown). Lanes 6-10: degradation of C3 (200|Lig/ml) with tissue-type plasminogen activator (0.4^g/ml) and plasminogen (350^g/ml) under identical conditions. Clearly visible are the bands of C3 a-(l 15kD) and C3 p-chains (75kD) and plasminogen (90kD), but not of plasmin (78kD). Even though little degradation is apparent after 30 minutes of incubation, extensive cleavage of both the a- and p-chain occurs between 30 and 90 minutes. The larger C3 degradation product again has a M.W. of 105-1 lOkD whereas the smaller fragment of about 60kD indicates further degrading of the C3 molecule.

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Analysis by PAGE indicated that generally only the a-chain of C3 was cleaved, with the exception of plasmin which cleaved both the a- and the P-chains (Fig. 1). Similarly, only the a-chain of C4 was cleaved in most instances, the P-chain remaining intact and the y-chain possibly degraded only by kallikrein (Fig. 2). On a molar ratio the most active of the enzymes was factor XIa, which cleaved almost half of the C3a-chains at a 1:15,000 molar ratio. The highest degree of degradation of the C4a-chain was seen with thrombin at a 1:5 molar ratio (Table III). When comparing the fragment size of the native C3 or C4 a-chains with their degradation products, both showed a reduction in size of about lOkD, indicating that C3a or C4a-likefragmentscould have been generated. Indeed, after incubation of C3 or C4 with thrombin, kallikrein, factors XIa and Xa or t-PA plus plasminogen for one hour at 37°C, significant elevations in C3a, and particularly C4a, levels could be measured. This result clearly indicates that antigenically active fragments had been generated. However, no band in the lOkD region could be seen on the Coomassiestained gels.

Figure 2: Kinetics of C4 degradation by factor XIa (left) and kallikrein (right). C4 (200^g/ml) was incubated for 0, 2, 30, 60 and 90 minutes with factor XIa (9ng/ml) or kallikrein (45^g/ml) at 3TC in HEPES buffer. Clearly visible are the C4 a-, C4 P- and C4 y-chains of 93, 75 and 32kD, respectively. Both enzymes have similar activities, apparently cleaving the C4 a-chain at a moderate but constant rate over the whole 90 minutes. The degradation product of the C4 a-chain appears to be about 80-83kD. Gel analysis by scanning also revealed a possible slight degradation of the C4 y-chain by kallikrein, but not by any of the other active enzymes (i.e. factors Xa, XIa, thrombin or plasmin).

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Table III. Relative activity of various proteases in the cleavage of the a-chains of C3 and C4' molar enzyme : C3/C4 ratio % C3a cleaved 1 enzyme %C4a cleaved | 1 : 15,000 47 16 1 1 XIa 1 : 1,000 11 32 Xa thrombin 1:5 51 90 kallikrein 27 2:3 33 1 plasmin 4: 1 39* 4 1 ^C3 or C4 was incubated for 60 minutes at 37°C with the respective enzymes, subjected to tricine-PAGE and Coomassie staining and analyzed by scanning the different bands. The values were obtained after background subtraction and normalization to the intensity of the C3p and C4p bands, respectively, except where marked with an asterisk.

IV. Conclusions We found that EDTA-plasma shows little ex vivo generation of C3a at 4°C within 48 hours, whereas C4a levels increased significantly. This effect was even more pronounced in whole blood that had undergone the same treatment. Heparin by itself did not appear to be an effective stabilizing agent, but worked well when combined with EDTA. However, the best results were obtained with the serine protease inhibitor Futhan, indicating that enzymes of the complement activation and/or coagulation pathway possessed residual activity after the divalent ions in plasma had been chelated by EDTA. In our experiments factors Xa, XIa, thrombin and kallikrein of the coagulation system, as well as plasmin, were indeed able to cleave C3 and C4. For both molecules, the a-chain was preferentially attacked and initially diminished in size by about lOkD. This was paralleled by the appearance of antigenically active C3a- and C4a-like products, indicating that a C3a- or C4a-like fragment was generated. However, the fact that no band in the lOkD-area could be detected on the polyacrylamide gel probably means that the fragments generated from C3 and C4 are not necessarily identical with C3a and C4a. Since we used active coagulation enzymes that are not necessarily generated in EDTA plasma, we can not rule out that continued classical pathway activation in EDTA plasma accounts for the ex vivo conversion of C4.

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Bibliography 1. Fujii, S. and Y. Hitomi. 1981. New synthetic inhibitors of CI r, CI esterase, thrombin, plasmin, kallikrein and trypsin. Biochim. Biophys. Acta 661:342-345. 2. Watkins, J., G. Wild, and S. Smith. 1989. Nafamostat to stabihse plasma samples taken for complement measurements. Lancet 896-897. 3. Issekutz, A. C, D. M. Roland, and R. A. Patrick. 1990. The effect of FUT-175 (Nafamostat mesilate) on C3a, C4a and C5a generation in vitro and inflammatory reactions in vivo. Int. J. Immunopharmac. 12:1-9. 4. Hugh, T. E. 1977. Complement factors and inflammation: effects of a-thrombin on components C3 and C5. In Chemistry and biology of thrombin. R. L. Lundblad, J. W. Fenton, and K. G. Mann, editors. Ann Arbor Science, Arm Arbor, Mich. 345-360. 5. Schaegger, H. and G. von Jagow. 1987. Tricine-sodium dodecyl sulfatepolyacrylamide gel electrophoresis for the separation of proteins in the range from 1 to 100 kDa. Anal. Biochem. 166:368-379. 6. Tait, J. F. and K. Fujikawa. 1987. Primary structure requirements for the binding of human high molecular weight kininogen to plasma prekallikrein and factor XI. J. Biol. Chem. 262:11651-11656. 7. Fujikawa, K. and B. A. McMullen. 1983. Amino acid sequence of human p-factor Xlla. J. Biol Chem. 258:10924-10933. 8. Maim, K. G. 1976. Methods in Enzymology XLV. L. Lorand, editor. Academic Press, New York. 123-156.

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DISULFIDE-LINKED HUMAN STEM CELL FACTOR DIMER Method of IdentiHcation and Molecular Comparison to the Noncovalent Dimer

Hsieng S. Lu, Michael D. Jones, and Keith E. Langley Amgen Inc., Amgen Center, Thousand Oaks CA 91320

I. INTRODUCTION Stem cell factor (SCF), also termed "kit ligand" or "mast cell growth factor" (1-6), functions in the early stages of hematopoiesis, and is an important growth factor involved in the development and function of other cell lineages, including melanocytes and germ cells (7,8). A soluble SCF form of 165 amino acids is biologically functional and contains approximately 40% of N- and O-linked sugar moieties (9-11). Two soluble SCF^-l^^ forms recombinantly expressed in Escherichia coli in a non-glycosylated form (rhSCF) and by mammalian cells in a glycosylated form (9,12,13) contain native SCF structure and are biologically functional. SCF binds to its receptor, kit, to elicit its specific biological functions (1-3). The kit receptor belongs to the type III tyrosine kinase family whose members include receptors for macrophage colony-stimulating factor (M-CSF) and platelet derived growth factor (PDGF) (14-16). SCF, M-CSF and PDGF are all dimeric ligands that mediate receptor dimerization (12,13,17,18). In contrast with the M-CSF and PDGF dimers whose monomers are disulfidelinked (17-19), both glycosylated and nonglycosylated SCF dimers contain non-covalently linked monomers (10,12). The SCF noncovalently associated dimer was observed to undergo spontaneous dissociationreassociation of monomers in its native state (20). There are two intramolecular disulfide bonds present in each monomer of SCF molecule (Cys^-Cys^^ and Cys^^^-Cys^^^); and the production of active rhSCF from E. coli requires an oxidative folding procedure to recover its biological activity (13). Oxidation and folding of denatured and reduced rhSCF involves at least three major partially oxidized intermediates, I-l to /5, each containing a native-like or mis-paired disulfide bond (21). These forms appear to reach steady state equilibrium and are important folding intermediates. There are two off-pathway intermediates that are dimers linked by a single intermolecular disulfide bond (Cys43-Cys^^ and Cys^^Cys^^, respectively). These two intermediates exist during early folding time and disappear after folding. In the final folding mixture, the major folded TECHNIQUES IN PROTEIN CHEMISTRY VIII Copyright © 1997 by Academic Press All rights of reproduction in any form reserved.

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SCF is the noncovalently linked dimer (SDS-dissociable) and a small fraction is SDS-nondissociable dimer. In this report, we describe strategy and methodology leading to the verification of the structure of SDSnondissociable SCF dimer. The dimer is covalently linked by four intermolecular disulfide bonds involving all cysteinyl residues. The cysteines are paired as in the non-covalently associated dimer except that all pairings are intermolecular rather than intramolecular. Other structural models, involving interwining of intramolecular disulfide loops, are ruled out. The understanding in molecular properties of the noncovalently and covalently linked dimers provides some insights to the structure and function of SCF. Detailed purification and biological/biochemical characterization of the disulfide-linked dimer have been extensively described elsewhere (22). 11. MATERIAL AND METHODS Materials: Escherichia co//-derived rhSCF (SDS-dissociable dimer) was purified according to methods described previously (12,13). The recombinant molecule contains 165 amino acids plus an N-terminal methionine at position -1. lodoacetic acid was purchased from Sigma. HPLC solvents and water were purchased from Burdick and Jackson. Sequencing reagents and solvents were supplied by Applied Biosystems (Foster City, CA) and Hewlett Packard (Mountain View, CA). All other reagents were of the highest quality available. Isolation of SDS-nondissociable dimer: Recovery of rhSCF expressed in E, coli includes solubilization of rhSCF-containing inclusion bodies, oxidation and folding, and subsequent chromatographic steps (13). After cationic exchange chromatography using an S-Sepharose column, pooled SCF (approximately 1 L containing 600 mg rhSCF) was further subjected to C-4 reverse-phase chromatography performed with a BioCat liquid chromatographic system (Perceptive Inc., New Jersey) as described (22). Analytical HPLC analysis and structural characterization: SCF dimer hybridization studies were performed according a previously described cationic exchange chromatographic procedure (20). Reverse-phase (RP-) HPLC was performed using TFA-acetonitrile gradient elution. A Vydac C4 column (4.6 mm x 25 cm; 300 A) was equilibrated with 97% solvent A (0.1% TFA)/3% solvent B (0.1% TFA in 90% acetonitrile) with 215 and 280 nm UV detection at a flow rate of 0.7 ml/min. After samples were injected into the column, the following elution program was used: a linear gradient to 20% solvent B in 5 min and to 70% B in 60 min, then isocratic elution at 70% B for 20 min. N-terminal amino acid sequence analysis of peptides was performed on an automatic protein sequencer (Applied Biosystems Models 477A, 470) as described (10). Procedures used to sequence peptides recovered from gel bands electroblotted onto PVDF membranes were described in a previous report (23). Mass spectrometric analysis was performed in a Sciex API-Ill electrospray mass spectrometer by direct infusion of sample (0.1 mg/ml in 0.1% acetic acid) at 10 |i-l/min.

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Hydrogen peroxide oxidation of SCF dimer and CNBr cleavages: SDSnondissociable rhSCF dimer at 1 mg/ml in 10 mM sodium acetate, pH 5.0 was incubated with 0.5% (w/v) H2O2 at 25°C for 3 h (24). After reaction, the mixture was analyzed by analytical reverse-phase HPLC as described above. The conditions used were found to completely oxidize all Met residues except Met^S. Only a small fraction (about 10%) of Met^S was oxidized. A complete CNBr cleavage at the Met residues of H202-oxidized SCF dimer species was performed as follows. Vacuum-dried samples were redissolved in 70% formic acid (0.2 mg in 150 |Lil) and then incubated with freshly prepared CNBr (400 molar ratio to SCF) at 25°C for 24 h in the dark. All the cleaved samples were immediately vacuum dried for further analysis. Limited proteolysis by endoproteinase Lys-C: Sample was reconstituted in 20 mM Tris-HCl buffer, pH 7.5 (1 mg/ml) and digested with endoproteinase Lys-C (enzyme-to-substrate ratio= 1:100) at 25 C. At 15 min and 2 h, sample aliquots (100 |il each) were taken and digestion stopped by adding 5 |Lil of 20% TFA. Samples of 5-20 |ig were dried completely and subjected to SDS-PAGE as described below. Partial reduction of SCFs: One mg/ml solutions of SCF dimer species were incubated in the presence of 1.24 mg/ml dithiothreitol (DTT) in 0.1 M TrisHCl buffer (pH 8.5) containing 2.5 M urea, 60 mM NaCl, 2 mM EDTA. Aliquots of the reaction mixture were removed at selected time intervals and unreacted thiols were blocked by the addition of 1 M iodoacetic acid (10:1 molar ratio to the thiol) in 0.3 M Tris, pH 8.0 for 2 min at room temperature. Samples were then quickly frozen in a methanol/dry ice bath and subsequently analyzed by RP-HPLC using conditions described previously (21). Gel electrophoresis and electroblotting: Aliquots of dried samples (5-20 mg) were loaded onto individual lanes of precast 16% Laemmli polyacrylamide gels (10 wells; Novex Inc., San Diego, CA) and electrophoresed (25) under nonreducing and reducing conditions. After Coomassie blue staining and destaining, protein band intensity in each gel lane was measured using an image scanner (PDI Inc. New York). In separate analyses, gel bands were also electrophoretically transferred onto PVDF membrane and the Coomassie blue-stained bands were excised for N-terminal sequence analysis (23). III. RESULTS AND DISCUSSION Isolation of SDS-nondissociable SCF dimer: Expression of rhSCF in bacteria has resulted in the production of insoluble and inactive SCF accumulated in inclusion bodies. Solubilization and in vitro folding and oxidation are therefore necessary for the recovery and chromatographic purification of active SCF (12,13,21). The rhSCF isolated in this way is a noncovalently linked, SDS-dissociable dimer (12), like naturally occurring

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SCF (1). However, during the cationic exchange chromatography after folding and oxidation, we have noticed that rhSCF bands of 18 and 37 kDa co-elute, as analyzed by nonreducing SDS-PAGE (data not shown). The 37 kDa species was also detectable in the final folding mixture by HPLC (21).

[M+13H] 1556.3

2074.0

Nondissodble Dimer

—r— 60 Retention Time (min)

s^ (£

Nondissodable Dimer

1200

1400

[M+17H]] 2196.0 174.0

1600

U

1800

2000

2200

M/Z

Figure 1. A, SDS-PAGE analysis. From the left- standard (45, 31, 21 and 14 kDa, from the top), DTT-reduced rhSCF, DTT-reduced dimer, nonreduced rhSCF, and nonreduced dimer. B, RP-HPLC of wild type rhSCF and the SDS-nondissociable dimer. C. ESI-MS analysis of wild type rhSCF and the dimer. The multiply charged ions are indicated; and the molecular masses of each form obtained from the respective deconvoluted spectrum are 18,658.5±2.3 and 37,315.2+3.6, respectively. Preparative reversed-phase column chromatography resolved these two species from partially purified preparations obtained after cationic exchange chromatography. The isolated major form (80-85% of the total) has a molecular weight of 18 kDa under nonreducing SDS-PAGE, while the minor form (15-20%) has a molecular weight of 37 kDa (Fig. lA). Both forms migrates as 19 kDa bands on reducing SDS-PAGE as seen in Figure lA. The major form corresponds to the active non-covalently associated, SDSdissociable rhSCF dimer (17), while the minor form represents the SCF covalently linked, SDS-nondissociable dimer. Analytical RP-HPLC using TFA-acetonitrile gradient elution is shown in Fig. IB. This analysis provides a full resolution of the two species with the covalently linked dimer being eluted later (more hydrophobic). In electrospray mass spectrometric analysis shown in Fig. IC, the major form gave an average MH+ mass of

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18,658.5+2.3 (theoretical mass = 18,657.6), while the dimer gave an average mass of 37,315.6+3.6 (theoretical mass = 37,315.2). This data indicated that the covalent dimer contains two identical SCF monomer, like the dissociable dimer. The 37 kDa species is now referred as SDSnondissociable dimer to dintinguish from the normal dissociable dimer. Molecular comparison: Extensive comparison of the biological, biochemical and biophysical properties between the SDS-dissociable and nondissociable forms was reported (22). Many molecular properties are shared by both molecules, however, clear differences can also be observed. A brief summary of structural/functional similarity and difference is described in Table 1. An example of difference between the two forms can be demonstrated in a dimer dissociation-reassociation experiment using a cationic exchange HPLC method. The usual noncovalently associated dimer can undergo spontaneous rapid monomer dissociation-reassocaition (20). This was shown with the use of an NIOD variant of rhSCF which migrates differently from the wild type on ion-exchange HPLC as indicated in Figure 2. Upon mixing the NIOD and wild type molecules, the appearance of hybrid dimer could be monitored (Fig. 2, chromatogram 3). Not suprisingly, the SDS-nondissociable dimer did not undergo such dissociation-reassociation and subunit exchange (Fig. 2, chromatogram 6). Table L Comparison of the molecular properties between SDS-dissociable and nondissociable rhSCF dimers A. Similaiitv: -Profile in ion-exchange chromatography and gel filtration (simiUir charge and size). -In viU"o biologiciil activity to hematopoietic cells and receptor binding. Nondissociable dimer is tliiee-fold more active, but it binds to receptor with a half efficiency (22). -Identity in disulfide pairing, Cys^-Cys^^ and Cys^^-Cys^^^ (identical peptide map). -Identity in CD spectra, fluorescence spectra, and thermostability (simiUu* secondary and tertiaiy stiuctures and lociil environment). -Homodimer in solution. B. Difference: -The covalent dimer is not dissociated by SDS (linked by intennolecular disulfides). -The SDS-nondissociable dimer elutes later in reverse-phase cliiomatography (i.e., more hydrophobic). -The SDS-nondissociable dimer c^ui not undergo spontaneous monomer-dimer dissociation-reassociat ion.

Possible Structures for SPS-nondissociable rhSCF dimer: Since the disulfide bonds Cys^-Cys^^^ and Cys^^-Cys^^^ are present in both forms and the monomers of the nondissociable SCF dimer become dissociable in the presence of reducing agent (see Fig. 1 A), it follows that there are two types of structures to explain the lack of dissociation in SDS. These two types of models are inteiTnolecularly disulfide-linked and concatenated dimers (Fig.

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1

20

30

0

10

20

Retention Time (min)

Figure 2. Cationic exchange HPLC. Chromatograms 1-6: rhSCF, NIOD variant, rhSCF and NIOD variant mixture, nondissociable dimer, nondissociable dimer and rhSCF, and nondissociable dimer and NIOD variant. Incubation was in 10 mM NaOAc, pH 4.5, at 37 ^C for 20 h with concentration of each sample at 1 mg/ml; and 50 |il sample was injected. 3A and 3B, respectively). There are three disulfide-linked dimers, of which cysteines are involved in the formation of four intermolecular S-S linkages (structure Al), or two inter- and two intramolecular S-S bridges (structures A2 and A3). The five possible concatenated dimers would contain interlocked, but not covalently-linked, monomers (Fig. 3B). Structure Bl is a dimer concatenated by N-terminal disulfide loops of the two monomers, while structure B2 is interlocked by two C-terminal disulfide loops. Structures B3 and B4 are dimers with a respective N- and C-terminal disulfide loop of one monomer locked into the other monomer near a sequence region (between residues 44 and 88) shared by both N- and C-terminal disulfide loops. B5 is concatenated between the N-terminal disulfide loop of one monomer and the C-terminal loop of the other. In order to determine which structure(s) correspond(s) to the isolated SDSnondissociable dimer, the experiments described in the following were performed. Several strategies were followed according to the above models. As indicated in Fig. 3, Lys^^, Lys^^^ and Met^S are important sequence positions for the cleavages. Table 2 compares the expected cleavage results to the observed data for those particular structures shown in Fig. 3. Structural characterization of the SDS-nondissociable dimer: A. Limited endoproteinase Lys-C digestion: Figure 4A shows SDS-PAGE of digests generated by limited proteolysis (nonreducing conditions, for 15 min [lane 1] or 2 h [lane 2]) with Lys-C protease. The SCF polypeptide has

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Disulfide-Linked Human Stem Cell Factor Dimer

14 Lys residues along the polypeptide chain including Lys^^ and Lys^^^. After the limited proteolysis, bands are still apparent near the 36 kD position. When these bands were transferred to PVDF membrane and sequenced, two sequences, M"l-E-G-I-C... and S^^^-P-E-P-R..., were detected in equivalent yields, suggesting that there is complete cleavage after Lys^^^. Several small peptides were also isolated from the digest by reverse-phase HPLC, and shown by sequence and mass spectrometric analyses to be D^^-L-K, K^^^S-F-K, D149.S-R-V-S-V-T-K-P-F-M-L-R-P-V-A-A, and pl57.F-N-L-P-PV-A-A. The latter two are C-terminal peptides not in the disulfide loops. Identification of these small peptides indicates that there was also partial cleavages by Lys-C after Lys^^, Lys^^, Lys^^^, and Lys^^^. Verification of these complete and partial cleavages was provided by reducing SDS-PAGE (lanes 3 and 4, Figure 4A). Only three large peptides, 11, 10, and 7 kDa were seen; the 11 kDa and 10 kDa bands had the rhSCF N-terminal sequence M-1-E-G-I-C, and the 7 kDa peptide had the sequence S^O^.p-E-P-R... In this case, the key point is that cleavage after Lys^^^, i.e., within the Cys^^Cysl38 disulfide loop, still leaves material which migrates near 35 kDa on non-reducing SDS-PAGE. This finding is inconsistent with models B2, B4, and B5, but consistent with all of the other models (Table 2). Table 2, Assignment of SDS-nondissociable SCF dimer structure by specific cleavages Lys-C cleavage atLys99,103,148

Partial DTT reduction Cys"^, Cys^^ reduced

Expected^ Observed*^

Expected Observed

Expected Observed

Al A2 A3

monomer dimer dimer

monomer monomer monomer

dimer dimer dimer

dimer dimer dimer

dimer monomer dimer

dimer dimer dimer

81 B2 B3 B4 B5

dimer dimer monomer monomer dimer

monomer monomer monomer monomer monomer

dimer monomer dimer monomer monomer

dimer dimer dimer dimer dimer

monomer dimer monomer dimer monomer

dimer dimer dimer dimer dimer

Structures^

CNBr cleavage at Met^8

^Structural Al is the only model compatible with all experimental data (for details, see text). ^Expected result if the predicted structure is cleaved by specific cleavage methods used. ^Observed results were found in Fig. 4 and 5 (for details, see text).

B. CNBr cleavage of H202-oxidixed dimer at Met^^: Five methionines, i.e., Met"l, Met^^, Met^^, Met"*^, and Met^^^, are in the rhSCF sequence, with Met^^ and Met^^ in the N-terminal loop created by the Cys^-Cys^^ bond and Met"^^ in a sequence region shared by both disulfide loops. A complete

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CNBr cleavage at Met residues will open the N-terminal loop and cut in the area shared by both loops. However, a complete cleavage opens up all dimer structures (Fig. 3) and generates monomer forms under nonreducing SDSPAGE. In another approach, the SDS-nondissociable dimer was reacted with H2O2 under conditions which completely oxidize Met"^ Met^^, Met^^,and Met^^^, but only partially (about 10%, as indicated by peptide

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remains at the "dimer" position (about 35 kDa). The material at this band position has sequences corresponding to the rhSCF N-terminus and to the peptide beginning at Val^^, in a ratio of 2:1; this result is expected since 10% of the H202-treated material was oxidized at Met^^ and therefore uncleavable with CNBr. Models Al, B3, and B4 all allow for retention of "dimer" if only one chain and not the other is cleaved. C. Partial reduction ofdimeric SCFs: When native, SDS-dissociable rhSCF dimer was partially reduced with DTT followed by alkylation with iodoacetate and the resulting mixture was analyzed by reverse-phase HPLC and peptide mapping of the HPLC peaks, the Cys'^-Cys^^ bond was found to be preferentially reduced, with generation of an intermediate {1-2) containing only the Cys^^^-Cys^^^ bond (Fig. 5A, chromatogram 2); reduction of the Cys'^^-Cys^^^ bond follows at later times. When the SDS-nondissociable SCF dimer was similarly subjected to partial reduction and alkylation, no 7-2 was detected. Instead, two unique peaks, a and b, were resolved by HPLC at retention times later than that of the SDS-nondissociable SCF dimer (Fig. 5A, chromatogram 4). By sequence analysis, both species gave a clear PTH-Cys (Cm) signal at position 4; the signal for peak b was about half that for peak a. In addition, both peaks a and b migrate at the 36 kDa "dimer" position on non-reducing SDS-PAGE (Figure 5B, lanes 6 and 7). These A

B

kDa

-< 35

18 13

Figure 4. SDS-PAGE of peptide products of SDS-nondissociable dimer derived from chemical and proteolytic cleavages. A, endoproteinase Lys-C digestion. Lanes 1 and 2 (nonreducing), products at 40 and 10 |ig: lanes 3 and 4, as lanes 1 and 2, but reducing. B, CNBr cleavage of Met-oxidized dimer. Lane 1 (nonreducing), oxidized dimer; lane 2 (nonreducing), cleavage product: lanes 3 and 4, as lanes 1 and 2, but reducing.

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.

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Figure 5. Partial DTT reduction ofrhSCF dimers. A, RP-HPLC analysis. Chromatograms 1 and 2: SDS-dissociable dimer (N), untreated and DTTtreated (10 min), respectively; chromatograms 3 and 4, SDS-nondissociable dimer, untreated and DTT-treated (5 min), respectively. Each in 50 L | Lg was injected. 5 , Nonreducing SDS-PAGE of forms referred to in A. Lanes, from left, protein standards, R, N, SDS-nondissociable dimer, 1-2, a, and b. findings indicate that peak a is "dimeric" material in which both Cys^-Cys^^ disulfide bonds have been broken, and the Cys^^-Cys^^^ disulfide bonds are intact, while peak b is "dimeric" material in which only one of the Cys^Cys^^ disulfide bonds has been broken. Since there is no detectable "monomeric" material (as 7-2), we conclude that the data are inconsistent with models A2, Bl, B3, and B5, but consistent with the other models (Table 2). As summarized in Table 2, the only model for the SDS-nondissociable rhSCF dimer compatible with all the results of the last three experiments is Al, with four intermolecular disulfide bonds involving all four Cys residues of each monomer. Therefore, the SDS-nondissociable dimer is a disulfidelinked dimer, with no intramolecular disulfide bonds. None of the proposed concatenated dimers exist. Comparison and speculation of quaternary structure: As described in rhSCF folding studies (21), intermediate I-l with a Cys^-Cys^^ disulfide bond is the main intermediate form during rhSCF folding and oxidation. This and other intermediates lead to the non-covalently associated SCF dimer

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with intramolecular disulfides, but could also undergo disulfide rearrangement to form intermolecular disulfides. For such events to occur, the partially-oxidized rhSCF monomers would have to be associated prior to disulfide formation; we have shown that all of the intermediate forms that have been identified are in dimeric state (21). As described in Table 1, many of the biochemical and biophysical properties of the non-covalently associated dimer and the disulfide-linked dimer appear indistinguishable - including surface charge, molecular size, plus secondary and tertiary structure and local environments. The disulfide-linked dimer does behave differently than the non-covalently associated dimer on RPHPLC at low pH and in the monomer dissociation-reassociation experiments (Figs. 1 and 2). In each case the differences essentially reflect the covalent attachment of the disulfide-linked dimer. The biological properties of the covalent dimer are noteworthy. Its activity toward hematopoietic target cells is 3-fold higher than the activity of non-covalently associated dimer (Table 1). However, in c-kit receptor binding experiments, the disulfide-linked dimer if anything displayed slightly lower affinity for kit in comparison with the non-covalently associated dimer. This phenomenon may be due to the

B Figure 6. Proposed quaternary structures of rhSCF dimer and disulfidelinked dimer. A, SDS-dissociable rhSCF dimer with topology similar to MCSF. B, disulfide-linked dimer having all disulfides at the dimer interface. C, disulfide-linked dimer containing A and D helices swapped between subunits (distinguished as shaded and unshaded helices). The four helical structure (A-D helices) was derived from that proposed by Bazan (27).

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possibility that SCF dimer is necessary to mediate kit dimerization, or at least that SCF dimer may be more effective at doing so than SCF monomer, although monomeric SCF can mediate the dimerization and activation of kit receptor (26). Depending on the Ka for monomer association to dimer (20), it is possible that much of the noncovalently associated SCF dimer could be monomeric at the 0.05 -2 ng/ml range which is equivalent to the effective concentration for the biological assay, while the disulfide-linked dimer is dimeric at all concentrations. Our above observation implies that the overall quatemary structure, including interactions at the dimer interface, would be similar for the disulfide-linked and non-covalently associated dimers. In considering the structure of SCF, as pointed out by Bazan (27), there are many reasons to expect similarity to the structure of M-CSF, which is known (28). X-ray crystallographic structure of M-CSF dimer (28) includes the four-helix bundle for each monomer which had been proposed by Bazan for both M-CSF and SCF (27). The two monomers of M-CSF associate in head-to-head fashion, i.e., the top ends of the helix bundles associate leading to a flat and elongated overall shape. The SCF-equivalent intramolecular disulfide bonds (Cys^Cys^O and Cys^S-Cys^^Q) of M-CSF are at the ends of the helix bundles distal to the dimer interface. Given that the disulfide-linked SCF dimer described here is highly active, we suggest the following speculation as to how its structure may compare to the noncovalently associated SCF dimer. If the quatemary structure of noncovalently associated SCF (see the predicted four-helical bundle structure in Fig. 6, model A) is homologous to that of MCSF, the monomers of the disulfide-linked dimer would need to be inverted in order to accommodate the disulfide bond formation, without an adverse effect on activity (Fig. 6, model B). Alternatively, if the quaternary structure of the disulfide-linked SCF dimer (model B) and the noncovalently associated SCF dimer were similar to each other, both could be inverted relative to that of the M-CSF dimer. Thirdly, and perhaps most likely, the quatemary structures of the SCFs could be similar to each other and to that of M-CSF if, for the disulfide-Hnked SCF dimer, the proposed A and D heUces (27) were swapped between the monomers within the dimer as seen in model C (Fig. 6). Such swapping would be feasible within the constraints of the proposed SCF structure (i.e. similar to M-CSF structure), and could conceivably arise during the refolding of the E. C(9//-derived recombinant molecule. There is precedent for such swapping of helices or other domains between monomers within overall oligomeric structures, e.g., interleukin 5 (29) and many other proteins as well (30). ACKNOWLEDGMENTS We are indebted to technical help from microbial fermentation and recovery process development groups at Amgen Inc. in the expression and isolation of rhSCF.

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REFERENCES Zsebo, K.M., Wypych, J., et al (1990) Cell 63, 195-201. Martin, F.H., Suggs, S.V., et al (1990) Cell 63, 203-211. Zsebo, K.M., Williams, D.A., et al (1990) Cell 63, 213-224. Williams, D.E., Eisenman, J., et al (1990) Cell 63, 167-174. Copeland, N.G., Gilbert, D.J., et al (1990) Cell 63, 175-183. Huang, E., Nocka, K., et al (1990) Cell 63, 225-233. Russell, E.S. (1979) Adv. Genet. 20, 357-459. Silvers, W.K. (1979) The Coat Colors of Mice. A Model for Mammalian Gene Action and Interaction. Springer-Verlag, New York. 9. Lu, H.S., Clogston, C.L., et al (1992) Arch. Biochem. Biophys. 298, 150-158. 10. Lu, H.S., Clogston, C.L., et al (1991) J. Biol. Chem. 266, 81028107. 11. Langley, K.E., Bennett, L.G., et al (1993) Blood 81, 656-660. 12. Arakawa, T., Yphantis, D.A., et al (1991) /. Biol. Chem. 266, 1894218948. 13. Langley, K.E., Wypych, J., et al (1992) Arch. Biochem. Biophys. 295 21-28 14. Yarden, Y.,*Kuang, W.-J., et al (1987) EMBO J. 6, 3341-3351. 15. Ullrich, A. and Schlessinger, J. (1990) Cell 61, 203-212. 16. Miyajima, A., Kitamura, T., Harada, N., Yokota, T. and Aral, K.-I. (1992) Ann. Rev. Immunol. 10, 295-331. 17. Johnsson, A., Heldin, C.-H., Westermark, B. and Wasteson, A. (1982) Biochem. Biophys. Res. Commun. 104, 66-74. 18. Das, S.K. and Stanley, E.R. (1982) J. Biol. Chem. 257, 13679-13684. 19. Glocker, M.O., Arbogast, B., Schreurs, J. and Deinzer, M.L. (1993) Biochem. 'il, 482-488. 20. Lu, H.S., Chang, W.-C, et al (1995) Biochem. J. 305, 563-568. 21. Jones, M.D., Narhi, L.O., Chang, W.-C. and Lu, H.S. (1996) J. Biol. Chem., 271, 11301-11308. 22. Lu, H.S., Jones, M.D., et al (1996) J. Biol. Chem. Ill, 1130911316. 23. Fausset, P.R. and Lu, H.S. (1991) Electrophoresis 12, 22-27. 24. Hsu, Y.-R., Narhi, L.O., Spahr, C , Langley, K.E. and Lu, H.S. (1996) Protein Science 5, 1165-1173. 25. Laemmli, U.K. (1970) Nature 2276, 680-685. 26. Lev, D., Yarden, Y. and Givol, D. (1992) J. Biol. Chem. 267, 1597015977. 27. Bazan, J.F. (1991) CelUS, 9-10. 28. Pandit, J., Bohm, A., et al (1993) Science 258, 1358-1362. 33. Milburn, M.V., Hassell, A.M., et al (1993) Nature 363, 172-176 30. Bennett, M.J., Schlunegger, M.P., and Eisenberg, D. (1995) Protein Science 4: 2455-2468

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AUTOCATALYTIC REDUCTION OF A HUMANIZED ANTIBODY A. Ashok Kumar John Kimura Jennifer Running Deer ICOS Corporation Bothell, Washington 98021

I. INTRODUCTION Disulfide (-S-S-) bonds play an important role in the structure and function of proteins. IgG molecules are comprised of two heavy and two light chains linked by interchain disulfide bonds. In addition, intrachain disulfide bonds are also present in IgGs. Alkyl thiols have been used to demonstrate the structural and functional role played by disulfide bonds. Reduction of IgGs with alkyl thiols under denaturing conditions results in separation of light and heavy chains. Complimentarity determining region (CDR) grafting is a unique way of generating human antibodies with the same specificities as murine antibodies. We report here our work with a humanized antibody (hAB-1), its parent murine antibody (pMAB) and a control humanized antibody (hAB-2). Both pMAB and hAB-1 contain a cysteine residue in CDR-1 of the heavy chain. Our results demonstrate autocatalytic reduction of hAB-1 under denaturing conditions. We also report the identity of the multiple species formed due to autoreduction. N-ethylmaleimide (NEM), an alkylating reagent, reacts specifically with protein cysteines at neutral pH to form stable thioether bonds (1-4). Thiol specific reagents and in vitro mutagenesis experiments were used to confirm the involvement of a cysteine residue in the autoreduction of hAB-1. TECHNIQUES IN PROTEIN CHEMISTRY VIII Copyright © 1997 by Academic Press All rights of reproduction in any form reserved.

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II. MATERIALS AND METHODS Purification of Antibodies: Humanized antibodies (hAB-1 and hAB-2) were purified by the technique of Protein-A chromatography, followed by Phenyl Sepharose and Ion-exchange chromatographies. The murine antibody (pMAB) was purified by Protein-A chromatography. SDS-PAGE: Poly aery lamide gel electrophoresis of proteins was performed using NOVEX (San Diego, CA) pre-cast, 1 mm 12 % Trisglycine gels, by the method of Laemmli (5). Protein samples were boiled for 1-10 minutes in reducing or non-reducing NOVEX sample buffers (pH 6.8). For reducing gels, DTT was added to the sample to a final concentration of 50 mM. Samples were electrophoresed at a constant voltage of 200 volts for 70 minutes. Protein bands were visualized by staining with Coomassie Blue R-250. Aikylation of Protein Thiols: Free cysteines of pMAB and hAB-1 were blocked with NEM or iodoacetamide by incubating the proteins (50 \ig) with NEM (4 jag) or iodoacetamide (200 lag) in PBS at room temperature for 10 or 30 minutes. Matrix Assisted Laser Desorption lonization-Mass Spectrometry (MALDI-MS): Mass spectra of native and denatured antibodies were obtained with a PerSeptive Biosystems (Farmingham, MA.) Voyager Elite mass spectrometer operated in the linear mode with a Laser Sciences Inc., 337 nm nitrogen laser. hAB-1 was denatured by boiling the sample in 1.0 M guanidine-HCl, 50 mM Tris pH 7.5 buffer. Native and denatured samples were diluted with 20 mM Tris, 10 mM octylglucoside (Tris/OG) pH 6.8 buffer prior to MALDI-MS analysis. Proteins were spotted on the sample plate as a sandwich between two layers of the matrix. The bottom layer consisted of 100 mM sinapinic acid in acetonitrile and the top layer consisted of 50 mM sinapinic acid in 30% acetonitrile / 70% H2O / 0.07% TFA. The m/z scale of the instrument was calibrated using a HewlettPackard protein standard mixture. Cysteine to Serine Conversion: The heavy chain CDR-1 residue of hAB-1 was converted to a serine residue by standard in vitro mutagenesis techniques. The mutation was confirmed by sequencing the entire cDNA. The modified CS-hAB-1 was purified by Protein-A chromatography.

Autocatalytic Reduction of a Humanized Antibody

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III. RESULTS AND DISCUSSION A. Apparent Heterogeneity of Humanized Antibody hAB-1 Humanized antibodies hAB-1, hAB-2 and pMAB were examined by SDS-PAGE under non-reducing and reducing conditions (Figure 1). Nonreducing SDS-PAGE of hAB-1 revealed 10-15 protein bands in the molecular weight range of 25 to 200 kDa, while the hAB-2 protein had two minor bands (>95 kDa and <200 kDa) and a major band at 200 kDa. The parent murine antibody had two very closely spaced bands near 200 kDa. On reduction, all three proteins showed primarily two major bands corresponding to mass values of 50 kDa and 25 kDa. All three proteins contained a minor band just above the 25 kDa band. Occasionally, a very low molecular weight band, near the dye front, was seen with hAB-2 under reducing conditions. kDa MW 1 2 3 200.0 fp •— ,pi 116.3 97.4 . 'PI 66.3

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Figure 1. Apparent heterogeneity of humanized antibody hAB-1. Ten micrograms of each antibody were mixed with non-reducing or reducing SDS-PAGE sample buffer and boiled for ten minutes prior to electrophoresis. The NOVEX molecular weight markers (MW) are identified by their mass values. Samples: Lanes 1,5 pMAB, lanes 2,6 hAB-2 and lanes 3,7 hAB-1. HC and LC correspond to heavy and light chains of antibodies, respectively.

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B. Effect of Sample Preparation on Apparent Heterogeneity The possible role of sample preparation technique on the observed heterogeneity was examined by studying the effects of incubation temperature and duration on the appearance of multiple bands with hAB-1. Results from SDS-PAGE of hAB-1 and pMAB boiled for 1, 5, and 10 minutes in non-reducing buffer are shown in Figure 2. The intensity of the lower molecular weight bands increase with boiling time particularly for hAB-1. Figure 3 shows the effect of incubating the humanized antibody in non-reducing SDS-PAGE buffer at different temperatures. hAB-1 migrates as a doublet between the 116.3 kDa and 200 kDa markers when incubated at 5°C or 21°C. In addition, these samples also contain a minor band at 67 kDa. The 37°C samples showed the presence of >95 kDa band and 200 kDa band in addition to the bands seen with the 5°C or 21°C samples. Samples incubated at 60°C or boiling temperature had several low molecular weight bands.

MW kDa 200.0 — 116.3 _ 97.4 ""' 66.3 — 55.4 —

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3

4

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Figure 2. Influence of sample preparation time on apparent heterogeneity. pMAB and hAB-1 were mixed with non-reducing SDS-PAGE sample buffer and boiled for various times prior to electrophoresis. Boiling times: Lanes 1,2:1 minute, lanes 3,4:5 minutes and lanes 5,6:10 minutes. Samples: pMAB (lanes 1,3 and 5), hAB-1 (lanes 2,4 and 6).

Autocatalytic Reduction of a Humanized Antibody kDa 200.0 116.3 97.4 66.3 55.4

MW 1 2

3

4

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Figure 3. Influence of temperature on apparent heterogeneity. hAB-1 was incubated in non-reducing SDS-PAGE sample buffer for ten minutes at various temperatures prior to electrophoresis. Incubation temperatures: Lane 1: 5°C, lane 2: 21 °C, lane 3: 37°C, lane 4: 60'^C and lane 5: boiling. C. Molecular Integrity of hAB-1 Antibody Gel filtration HPLC of hAB-1 in 0.2 M sodium phosphate (pH 6.8) buffer showed that the majority of the protein (97%) was in the monomer form with retention time similar to the bovine gamma globulin (158 kDa) standard. MALDI-MS is a technique well suited for the examination of low and high molecular weight biomolecules (6,7). Figure 4 shows the mass spectra for hAB-1, hAB-2 and pMAB diluted in Tris/OG buffer. The spectra contain signals for (M+H)^^ (148 kDa), (M+2H)^^ (74 kDa), (M-f3H)^^ (49.5 kDa) and (M+4H)^^ (37.6 kDa) species in each case. In addition a minor signal at 23-24 kDa was observed.

A. Ashok Kumar et al.

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Figure 4. MALDI-MS of native hAB-1, hAB-2 and pMAB. Samples (2 pmols) in 20 mM Tris, 10 mM octylglucoside, pH 6.8 buffer were used for MS analysis. D. Autoreduction of hAB-1 Antibody The parent murine antibody (pMAB) and the humanized antibody (hAB1) both contain a cysteine residue in their heavy chain CDR-1. The role of this cysteine residue in the apparent heterogeneity of hAB-1 was examined by incubating the hAB-1 protein with thiol specific reagents iodoacetamide (lAA) and N-ethylmaleimide (NEM). Data in Figure 5 shows that preincubation with lAA or NEM can prevent the formation of low molecular weight species seen with untreated hAB-1. Treatment with thiol specific reagents suggests autoreduction as the cause for the apparent heterogeneity.

Autocatalytic Reduction of a Humanized Antibody

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21.5 lOMin.

30Min.

Figures. Role of a cysteine residue in autoreduction of hAB-1. hAB-1 (50 jig) was incubated with either NEM (4 lug) or lAA (200 ^ig) in PBS at room temperature for 10 or 30 minutes. At the end of the incubation period, treated and untreated samples (10 jiig) were boiled in non-reducing SDS-PAGE sample buffer and electrophoresed. Samples: Lanes 1,4: untreated, lanes 2,5: treated with NEM and lanes 3,6: treated with lAA. E. Autoreduction of hAB-1 and pMAB in Guanidine-HCl We have examined the mass spectra of all three proteins in 1.0 M guanidine-HCl with and without thermal denaturation. Samples incubated in guanidine-HCl at room temperature have very similar spectra as in Tris/OG buffer (data not shown). Mass spectra of samples boiled in guanidine-HCl (Figure 6) revealed that similar species are seen with hAB1 and pMAB and that both are subject to autoreduction under these conditions. Resuhs with hAB-2 were different compared to hAB-1 and pMAB. With all three samples, boiling in guanidine-HCl resulted in a significant loss of (M+H)^ signal for high molecular weight components (74.5-148 kDa). The loss of (M+H)^^ and (M+2H)^^ signals of IgG in all three cases could be due to precipitation of denatured high molecular weight components. In addition, guanidine-HCl could be interfering with desorption of the ionized high molecular weight components.

A. Ashok Kumar et al

392

hAB-1

i ^»w«n..

)ii I,J

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hAB-2

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13

N^wA.

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pMAB

20K 40K 60K 80K lOOK 120k 140K 160K m/z Figure 6. MS of hAB-l, hAB-2 and pMAB boiled in LO M guanidineHCl. Samples (1 mg/ml) in 50 mM Tris, 1.0 M guanidine-HCl pH 7.5 buffer were boiled for 5 minutes, diluted with Tris/OG buffer and analyzed by MALDI-MS. Table 1 lists the molecular mass values and possible species from the mass spectra of hAB-1 and pMAB boiled in guanidine-HCl. Observed species and probable identities include: the monomer (24 kDa), dimer (47.8 kDa), and trimer (71.9 kDa) of light chain, monomer (50 kDa) and dimer (100.5 kDa) of heavy chain, combinations of one heavy and one light chain (74.3 kDa), one heavy and two light chains (98.1 kDa), and two heavy chains and one light chain (124.5 kDa). The 74.3 kDa signal could be due to the (M+2H)"^^ species of IgG or the (M+H)"^^ species of a light chainheavy chain heterodimer. Also present were species with mass values of 172.5 kDa and 197 kDa, corresponding to an IgG complexed with an additional light chain or an additional heavy chain. Finally, we saw signals for several (M+2H)^^ species.

Autocatalytic Reduction of a Humanized Antibody

393

Table 1. Identity of Autoreduction Products of hAB-1 and pMAB Molecular Mass hAB-1 pMAR 197007

Possible Species IgG + HC

Charge Status (M+H)""

172513

172513

IgG + LC

(M+H)'"

148347

148018

IgG

(M+H)*'

124489

124209

2HC + LC

(M+H)^'

100475

100279

HC + HC

2(M+H)^'

98097

98186

HC + 2LC

(M+H)""'

74308

74118

HC + LC or IgG

(M+H)""' or (M+2H)"'^

71885

71929

3LC

3(M+H)*'

50404

50236

HC

(M+H)"

47848

48010

2LC

2(M+Hf'

37210

37158

IgG

(MMH)""*

25254

25149

HC

(M+2H)"^

24031

24027

LC

(M+nf

12039

12022

LC

(M+2H)"*'^

F. Identification of the Cysteine Residue Responsible for the Autoreduction of hAB-1 The heavy chain CDR-1 cysteine of hAB-1 was converted to a serine residue by in vitro mutagenesis. Non-reducing SDS-PAGE of the CShAB-1 antibody showed that the mutated protein was resistant to autoreduction (Figure 7), confirming the role played by this cysteine in the autoreduction during thermal denaturation.

394

MW

A. Ashok Kumar et al. 1

2

3

MW

1 2

3

4

5

w

Figure 7. Establishment of CDR-1 cysteine as the residue responsible for hAB-1 autoreduction. Ten micrograms of sample were boiled with or without reducing agent for ten minutes prior to electrophoresis. Samples: Lane 1: pMAB, lane 2: hAB-1, lane 3: NEM-treated hAB~l, lane 4: cysteine to serine mutant of hAB-1 and lane 5: hAB-2. IV. CONCLUSIONS The humanized antibody, hAB-1, was heterogeneous by non-reducing SDS-PAGE and MALDI-MS under denaturing conditions. NEM and iodoacetamide treatments established the role of a cysteine residue in generation of the heterogeneity. Reducing SDS-PAGE, HPLC gel filtration and MALDI-MS of native hAB-1 have shown that the native protein is intact and is similar to pMAB and hAB-2. Our results have shown that hAB-1, when denatured, is subject to autocatalytic reduction by a cysteine residue. This leads to the formation of low molecular weight species. The role played by the heavy chain CDR-1 cysteine of hAB-1 in this process was confirmed by in vitro mutagenesis of the cysteine to a serine residue. Our results have shown that a humanized antibody with a cysteine residue in its CDR is more susceptible to autoreduction compared to the

Autocatalytic Reduction of a Humanized Antibody

395

parent murine antibody. pMAB is not susceptible to autoreduction in nonreducing SDS-PAGE buffer, while thermal denaturation of pMAB in guanidine-HCl leads to autoreduction. hAB-1 is subject to autoreduction under both conditions. hAB-2 is not subject to reduction or breakdown under either conditions. Thus, reactivity of pMAB CDR-1 cysteine towards disulfide bonds is dependent on the denaturant employed to perturb the structure, while reactivity of hAB-1 CDR-1 cysteine is independent of the denaturant employed.

ACKNOWLEDGMENTS The authors would like to thank Dr. Leland Paul for critical reading of the manuscript and Eileen Jarvis for typing the manuscript.

REFERENCES 1. Heitz, J. R., Anderson, C. D., and Anderson, B. M. (1968). Arch. Biochem. Biophys. 127,'627-636. 2. Smyth, D. G., Blumenfeld, O. O., and Konigsberg, W. (1964). Biochem. J. 91, 589-595. 3. Gorin, G., Martic, P. A., and Doughty, G. (1966). Arch. Biochem. Biophys. 115, 593-597. 4. Partis, M. D., Griffiths, D. G., Roberts, G. C , and Beechey, R. B. (1983). J. Protein Chem. 2, 263-277. 5. Laemmli, U. K. (1970). Nature (London) 111, 680-685. 6. Zaluzec, E. J., Gage, D. A., and Watson, J. T. (1995). Prot. Expr. Purif. 6, 109-123. 7. Andersen, J. S., Svensson, B., and Roepstroff, P. (1996). Nature Biotech. 14, 449-457.

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SECTION V Interactions of Protein with Ligands

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Oxygen and Ascorbate Mediated Modification of a Recombinant Hemoglobin Bruce A. Kerwin, Edward Hess, Julie Lippincott, Ray Kaiser^ and Izydor Apostol Somatogen Inc., Boulder, Colorado 80301 ^ Eli Lilly and Co., Indianapolis, Indiana 46285

I. Introduction Recombinant hemoglobin, rHbl.l, is atrimeric protein composed of two p-globins, two genetically fused a-globins (di-a globin) and four hemes (see ribbon structure). In the reduced state the iron center of each heme group reversibly binds molecular oxygen. Upon binding oxygen the hemoglobin can autooxidize forming metHb which is incapable of binding oxygen: HbFe2+-02 ^ HbFe3+ + O2". The metHb can be reduced back to the ferrous state using ascorbate and reduced oxygen conditions (Vestling, 1941 and Gibson, 1943) as described by the mechanism Ascorbate2- + 2HbFe3+ -> 2HbFe2+ + dehydroascorbate (Al-Ayash and Wilson, 1979). In the presence of oxygen, however, ascorbate reacts with molecular oxygen to form dehydroascorbate and superoxide anion. The dehydroascorbate can undergo hydrolytic ring rupture to form 2,3-diketogulonic acid which in turn can react further with oxygen forming additional byproducts which may modify proteins (Washko et.al., 1992). One of the more prevalent protein modifications detected has been carboxymethylation of lysine groups to form Ns-(carboxymethyl)lysine (Dunn et al, 1990 and Ortwerth et al., 1992). Although it is known that the superoxide reacts with and oxidizes the hemoglobin little is known concerning the modification of the hemoglobin by dehydroascorbate and its byproducts. Studies were undertaken to determine how rHbl.l is modified in the presence of dehydroascorbate as well as in the presence of ascorbate and oxygen. TECHNIQUES IN PROTEIN CHEMISTRY VIII Copyright © 1997 by Academic Press All rights of reproduction in any form reserved.

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Bruce A. Kerwin et al

11. Materials and Methods A. Recombinant Hemoglobin Recombinant hemoglobin (rHbl.l) was produced at Somatogen as described by Looker et.al, 1992.

B. Ascorbic acid and Dehydroascorbate Ascorbic acid (sodium salt) and dehydroascorbate were both obtained from Sigma Chemical Co.

C Reverse Phase HPLC Reverse phase HPLC analysis was performed using a Zorbax C3 analytical column. The oven temperature was maintained at 40°C. Solvent A = H20/0.1% TFA and Solvent B = 100% acetonitrile/0.1% TFA. Flow rate = Iml/min. The column was equilibrated in 35% solvent B. Following sample injection the column was maintained for 5 minutes with 35% solvent B then ramped to 49% solvent B over 45 minutes. Samples were prepared by precipitation with icecold acid/acetone (Witkowska et al., 1993) and solubilization of the pellet in 0.1%) TFA/20%) acetonitrile.

D. Trypsin Mapping Tryptic mapping was performed on an HP 1090 HPLC modified with two independent switching valves for controlling flow to a Poroszyme immobilized trypsin Cartridge (Perseptive Biosystems) and an in-line Zorbax analytical CI8 column (Lippincott et al., manuscript in preparation).

E. Mass Spectrometry Mass spectrometry was performed using a Finnigan Mat LCQ.

F. Amino Acid Sequence Analysis Sequencing was performed on the isolated p-globin N-terminal tryptic peptide and the difference tryptic peptide using a Porton 2020 sequencer.

O2 and Ascorbate Mediated Modification of Recombinant Hb

401

n. Results and Discussion Studies were undertaken to determine how rHb 1.1 was modified by the oxygen mediated degradation of ascorbate. In the presence of ascorbate and ISOppm of oxygen the reverse phase HPLC profile of the hemoglobin was not significantly different from the control oxy-rHb without ascorbate (see Fig. 1). In contrast, when the hemoglobin was incubated with ascorbate and 15,000ppm of oxygen a decrease in the height of the P-globin peak was seen along with new peaks appearing on the lagging shoulder of the P-globin indicating a modification to the P-globin chain. Ascorbate is known to degrade in the presence of oxygen forming dehydroascorbate and superoxide anion which oxidizes the hemoglobin. In the sample containing only hemoglobin and 15,000ppm of oxygen the metrHb levels rose from 19% to approximately 40% while the met-rHb levels in the sample containing 15,000ppm oxygen and 5mM ascorbate demonstrated an overall decrease in the level of metHb to final concentration of 0.5% metHb (data not shown). This suggests that the superoxide anion produced during reaction of the ascorbate with oxygen alone was not responsible for the modification, rather the modification was due to a degradation product of the ascorbate. The initial product of ascorbate oxidation is dehydroascorbate. We incubated dehyrdroascorbate with the hemoglobin to determine if it could modify and produce similar reverse phase HPLC profiles as that seen with ascorbate and oxygen (see Fig. 1). Addition of dehydroascorbate to deoxy-rHb (middle panel Fig. 2) showed a reverse phase HPLC profile similar to that seen for ascorbate and oxygen (see Fig. 1). The height of the P-globin in the deoxy-rHb/ascorbate sample again decreased with new peaks appearing on the lagging shoulder of the P-globin. It appears that both the deoxy-rHb and oxy-rHb in the presence of dehydroascorbate are modified to the same degree (Fig. 2). However, we cannot exclude the possibility that residual oxygen leaking into the deoxy-rHb/ascorbate system through the septum was responsible for the modification. It is also possible that dehydroascorbate decomposition byproducts were already present prior to addition and caused the modification. The location of the modification was determined using trypsin mapping. Following dehydroascorbate modification of deoxy-rHb the main p-globin peak and the lagging shoulder of the P-globin were purified from reverse phase HPLC (see Fig. 2) and mapped with trypsin (see Fig. 3). The map of the unmodified Pglobin is shown in the upper panel and is not different from that of P-globin not exposed to ascorbate (data not shown). The map of the modified p-globin (lower panel Fig. 3) demonstrated a marked decrease in the N-terminal peptide (MHLTPEEK) of the p-globin eluting with a retention time of 51min. and the appearance of a difference peptide eluting with a retention time of 53.5min. This

Briice A. Kerwin et al

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025 20 DAD1 A. Sig=215.4

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min

5mM Ascorbate

600-

15,OOOppm Oxygen

900400300200100020

25

Jy 30

J 35

40

45

min

Figure 1: Revese phase HPLC analysis of hemoglobin modified by ascorbate and oxygen. Recombinant hemoglobin (50mg/ml) was equilibrated to the indicated oxygen tension followed by addition of ascorbate to a final concentration of 5mM or buffer (150mM NaCl, 5mM NaPi, pH7.4) as a control. Aliquots (0.5ml) were storred in stoppered 2ml vials for 15-16 days at 4°C then analyzed by reverse phase HPLC as described in materials and methods. The upper panel represents a typical chromatographic profile of rHbl.l. The peak at ~34.5min. represents the p-globin subunit and the peak at ~40min. represents the di-a-globin subunit. At 150ppm of oxygen no apparent modification of the protein primary structure was observed. However, at 15,000ppm of oxygen a distinct modification of the p-globin was observed.

O2 and Ascorbate Mediated Modification of Recombinant Hb

403

In contrast, when the hemoglobin was incubated with ascorbate and 15,000ppm of oxygen a decrease in the height of the (3-globin peak was seen along with new peaks appearing on the lagging shoulder of the P-globin indicating a modification to the P-globin chain. Ascorbate is known to degrade in the presence of oxygen forming dehydroascorbate and superoxide anion which oxidizes the hemoglobin. In the sample containing only hemoglobin and 15,000ppm of oxygen the metrHb levels rose from 19% to approximately 40% while the met-rHb levels in the sample containing 15,000ppm oxygen and 5mM ascorbate demonstrated an overall decrease in the level of metHb to final concentration of 0.5% metHb (data not shown). This suggests that the superoxide anion produced during reaction of the ascorbate with oxygen alone was not responsible for the modification, rather the modification was due to a degradation product of the ascorbate. The initial product of ascorbate oxidation is dehydroascorbate. We incubated dehyrdroascorbate with the hemoglobin to determine if it could modify and produce similar reverse phase HPLC profiles as that seen with ascorbate and oxygen (see Fig. 1). Addition of dehydroascorbate to deoxy-rHb (middle panel Fig. 2) showed a reverse phase HPLC profile similar to that seen for ascorbate and oxygen (see Fig. 1). The height of the p-globin in the deoxy-rHb/ascorbate sample again decreased with new peaks appearing on the lagging shoulder of the P-globin. It appears that both the deoxy-rHb and oxy-rHb in the presence of dehydroascorbate are modified to the same degree (Fig. 2). However, we cannot exclude the possibility that residual oxygen leaking into the deoxy-rHb/ascorbate system through the septum was responsible for the modification. It is also possible that dehydroascorbate decomposition byproducts were already present prior to addition and caused the modification. The location of the modification was determined using trypsin mapping. Following dehydroascorbate modification of deoxy-rHb the main p-globin peak and the lagging shoulder of the p-globin were purified from reverse phase HPLC (see Fig. 2) and mapped with trypsin (see Fig. 3). The map of the unmodified Pglobin is shown in the upper panel and is not different from that of P-globin not exposed to ascorbate (data not shown). The map of the modified P-globin (lower panel Fig. 3) demonstrated a marked decrease in the N-terminal peptide (MHLTPEEK) of the p-globin eluting with a retention time of 51min. and the appearance of a difference peptide eluting with a retention time of 53.5min. This

Bruce A. Kerwin et al

404

suggests that the dehydroascorbate modification of the deoxy-rHb is on the Nterminal tryptic peptide of the P-globin. DAD1 A, Sig=215.4 mAU-

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Figure 2: Reverse phase HPLC analysis of dehydroascorbate modified hemoglobin. Recombinant hemoglobin (50mg/ml in 150mM NaCl, 5mM NaPj, pH 7.4) was deoxygenated under a stream of humidified nitrogen, divided into two ahquots (3 ml each) in glass flasks stoppered with white rubber septa and stored at 4°C. Dehydroascorbate was prepared by evacuating and flushing a flask containing the dehydroascorbate solid followed by addition of deoxygenated water. Dehydroascorbate was added to each sample as an 8:1 molar ratio of dehydroascorbate: rHb 1.1. For the oxygenated sample one of the aliquots was reoxygenated by flushing its flask with oxygen prior to addition of the dehydroascorbate solution. Both samples were allowed to react overnight at 4°C prior to analysis. Samples were analyzed by reverse phase HPLC as described in materials and methods. The data indicate that modification by dehydroascorbate shows a similar reverse phase HPLC profile as seen for ascorbate and oxygen modification and that similar profiles are observed when modification is performed under both oxy and deoxy conditions.

O2 and Ascorbate Mediated Modification of Recombinant Hb

405

Figure 3: Tryptic mapping of the dehydroascorbate modified P-globin subunit. Dehydroascorbate modified deoxy recombinant hemoglobin was prepared as described in Fig. 2. The p-globin peak from 26-28 min. and the lagging shoulder of the p-globin peak from 2830min. were collected and mapped as described in materials and methods. The upper panel represents the tryptic map of the p-globin peak (26-28min.) and the lower panel represents the tryptic map of the lagging shoulder of the p-globin (28-3Omin.). The arrow in the upper panel indicates the position of the p-globin N-terminal peptide and the arrow in the lower panel indicates the position of the difference peptide present in the modified p-globin shoulder.

Modifications of other proteins by ascorbate have been reported on the NS-amino group of lysine to produce NS-(carboxymethyl)lysine which has a mass of 58amu (Dunn et al., 1990; Ortwerth et al., 1992). The mass of the difference peptide was determined using LC-MS (see Fig. 4 and Table I). The lower panel shows the spectrum of the modified peptide with a mass of 1055.2amu and xhe upper panel shows the spectrum of the P-globin N-terminal peptide with a mass of 983.5amu to produce a difference of 71.7amu.

Bruce A. Kerwin et al

406 (3-globin N-terminal peptide

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Figure 4: Mass spectrometry analysis of difference peptides from the dehydroascorbate modified p-globin subunit. Dehydroascorbate modified deoxy recombinant hemoglobin was prepared as described in Fig. 2. Trypsin digestion was performed as described in materials and methods and the p-globin N-terminal peptide (see upper panel Fig. 3) and the difference peptide (see lower panel Fig. 3) analyzed by LC-MS. The spectrum in the upper panel represents the p-globin N-terminal peptide and the spectrum in the lower panel represents the difference peptide. A summary of the data is presented in Table I.

This is different from that for dehydroascorbate which has a mass of 156amu and carboxymethylation which produces a mass increase of 58amu. Edman protein sequencing analysis of the difference peptide demonstrated a blocked Nterminus, suggesting that the N-terminal methionine was modified. Experiments are currently in progress to determine the exact structure of the modification.

O2 and Ascorbate Mediated Modification of Recombinant Hb

407

Table I: Summary of mass spectrometry and sequencing data from the unmodified and dehydroascorbate modified (3-globin peptides Sample

Sequence

Expected monoisotopic

Observed mass (amu)

N-terminal peptide

MHLTPEEK

983.5

983.5

Difference peptide

Blocked to Edman sequencing

p-globin

1055.2

A = 71.7

References Al-Ayash, A.I. and Wilson, M.T. (1979) Biochem. J. Ill

Ml.

Dunn, J.A., Ahmed, M.U., Murtiashaw, M.H., Richardson, J.M., Walla, M.D., Thorpe, S.R. andBaynes, J.W. (1990) Biochemistry 29:10964. Gibson, Q.H. (1943) Biochem. 1 37:615. Hoffman, S.J., Looker, D.L., Roehrich, J.M., Cozart, P.E., Durfee, S.L., Tedesco, J.L. and Stetler, G.L. (1990) Proc. Natl Acad. Sci. USA 87:8521. Looker, D., Abbott-Brown, D., Cozart, P., Durfee, S., Hoffman, S., Mathews, A.J., Miller-Roehrich, J., Shoemaker, S., Trimble, S., Fermi, G., Komiyama, N.H., Nagai, K. and Stetler, G. (1992) Nature 356:258. Ortwerth, B.J., Slight, S.H., Prabhakaram, M., Sun., Y. and Smith, J.B. (1992) Biochim. Biophys. Acta. 1117:207. Vestling, C.S. (1941) J. BiolChem. 143:439. Washko, P.W., Welch, R.W., Dhariwal, K.R., Wang, Y. and Levine, M. (1992) Anal. Biochem. 204:1. Witkowska, H.E., Bitsch, F. and Shackleton, C.H.L. (1993) Hemoglobin 17:227.

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Metal activation and regulation of E.coli RNase H James L. Keck and Susan Marqusee Dept. of Molecular and Cell Biology University of California, Berkeley Berkeley, CA 94720

Introduction: The ribonuclease H (RNase H) family of enzymes are ubiquitous nucleases that catalyze the hydrolysis of RNA in RNA»DNA hybrids (for review, see 1). In contrast to the well-studied ribonucleases A and Tl, RNase H does not employ the 2'-OH in RNA as a nucleophile but instead activates water as the nucleophile for hydrolysis in a metal-dependent reaction. The number and role(s) of divalent metal in the RNase H reaction mechanism are still unclear. Two RNase H mechanisms have been proposed based on wellcharacterized metal-dependent DNase activities ~ a one-metal mechanism (2,3,18), modeled after DNase I (4), and a two-metal mechanism (5,19), modeled after the exonuclease domain from Klenow fragment (6-8). The one-metal mechanism is supported by observation of a single Mg2+ binding to E.coli RNase HI via X-ray crystallography, NMR and isothermal titration calorimetry (9-11,18). Also, mutagenesis of conserved residues in RNase H shows that only three of these ten residues result in a complete loss of activity when mutated to alanine (12). Of these three acidic residues, two are found by x-ray crystallography to ligand a single Mg^^ (Asp 10 and Glu48) and the third (Asp70) is proposed to abstract a proton from the attacking nucleophilic water (3,9). This divalent metal is proposed to stabilize the r e a c t i o n ' s p e n t a c o v a l e n t p h o s p h o r e n e t r a n s i t i o n state intermediate. In contrast, the two-metal mechanism is supported by observation of two Mn^+ ions bound in the active-site in the crystal structure of HIV-1 RNase H domain (a d o m a i n of reverse transcriptase) (5). The Mn2+ ions are --4 A apart (as is seen in the Klenow fragment exonuclease domain (6,8)) and are bridged by a uranium heavy atom. It is thought that the uranium acts as an TECHNIQUES IN PROTEIN CHEMISTRY VIII Copyright © 1997 by Academic Press All rights of reproduction in any form reserved.

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James L. Keck and Susan Marqusee

artificial bridging ligand that would normally be satisfied by substrate. In this two-metal mechanism, one metal activates the hydroxyl nucleophile and the second metal stabilizes the p h o s p h o r e n e intermediate of the reaction. Both mechanistic hypotheses are daunted by the lack of structural information on RNase H bound to its nucleic acid substrate. An understanding of the RNase H mechanism is important for a number of reasons. First, the RNase H activity is essential to the lifecycle of HIV (for review, see 13). Mutations that merely reduce the reverse transcriptase RNase H activity are sufficient to completely inhibit virulence in the mutant in vivo (14). Its absolute requirement makes the RNase H activity a logical drug target for anti-HIV therapies. Development of knowledge-based inhibitors will require an understanding of the activity's mechanism. Second, a number of proteins with structures homologous to RNase H have been solved in the past two years, all of which are metal-requiring nucleic acid manipulating proteins (reviewed in 15-17). This superfamily of proteins, now termed "polynucleotide transferases", includes RNase H (5,18,19), resolvase (20), integrase (21,22) and Mu transposase (23). It is assumed that the enzymes share a common mechanism, so an understanding of RNase H mechanism will assist in clarifying the mechanism of other members of this superfamily. Clearly, a determination of the number and role(s) of metal in the E.coli RNase H active site will help establish the enzyme's mechanism. We have examined the metal dependence (Mn2+ and Mg2+) of E.coli RNase H activity. Mn^^-dependent activity requires much less metal for activation than does Mg2+-activity and is inhibited upon the further addition of Mn2+. Using electron paramagnetic resonance (EPR), we have measured two distinct Mn^^ binding constants, consistent with the concentration requirements for activation and inhibition in vitro. Our data are most consistent with a singledivalent metal catalyzed reaction which can be attenuated (inhibited) upon binding a second metal. We discuss a possible mechanism for metal activation and inhibition of RNase H in light of previous mutagenesis and structural studies.

Materials and Methods

Materials: pJK502 is a T7-overexpression vector that encodes for wild-type E.coli RNase HI. It was made by site-directed mutagenesis of pSMlOl (24), reverting three alanines (residues 13, 63 and 133) to cysteines, and then subcloning the resulting gene into a p E T l l a overexpression vector (details of plasmid sequence available upon request). Overexpression and purification of E.coli RNase HI were performed essentially as described in (24) for RNase H'*'. RNase H* is

Metal-Dependence of E. coli RNase H

411

a cysteine-free version of E.coli RNase HI and was a gift from Dung Vu. M e t h o d s : i^Nflse H activity assay. Production of R N A » D N A hybrid and RNase H activity assays were performed essentially as described in (25). MnCl2 stocks were made by serial dilution of a 1 M MnCl2 stock in 1% Nitric Acid. Final MnCl2 stocks were in 0.1% Nitric Acid. The p H of the final assay solutions were unchanged by the addition of the Nitric Acid as confirmed by measuring the p H of mock reactions containing buffer (Tris) and the appropriate volume of 0.1% Nitric Acid. Velocity values are specific activity (units/mg enzyme) measurements based on four time points in the linear range of enzymatic activity. Assays were performed in 50 mM Tris, p H 8.0/50 mM N a C l / 1 mM DTT/1.5 |LIM B S A / 1 |LIM (basepairs) RNA*DNA hybrid with 0.2 to 1.2 nM E.coli RNase HI at 37 °C. One unit is defined as the amount of enzyme needed to generate 1 |Limol of acid-soluble product in 15 minutes under our reaction conditions. Electron Paramagnetic Resonance (EPR) binding experiments: All EPR measurements were carried out on a Bruker ESP300E X-band spectrometer at ambient temperature. Lyophilized RNase H* was resuspended in 50 mM Hepes, p H 7.5, and diluted 1:1 with MnCl2 stocks made up in water ~ final concentrations were 25 mM Hepes, p H 7.5, 50 |LiM RNase H* and MnCl2 from 15 to 300 |iM. Free Mn2+ is the only component that gives an EPR spectrum, allowing calculation of [Mn2+]bound ([Mn2^]totai = [Mn2+]free + [Mn2+]bound). A Mn^^ peak (centered at 3629.6 G with a + / - 60 G sweep) was measured and then compared (height and peak shape) with Mn^^ standards in the same buffer conditions to determine the scalar difference. Using these scalar factors, [Mn^+lfree was calculated at various total Mn2 + concentrations. Dissociation constants were determined via Scatchard analysis of the data (26).

Results

Mn^-^-dependence of E.coli RNase HI: The Mn2+-dependence of E.coli RNase HI catalysis was determined using a soluble assay that monitors acid-solubility of radiolabeled RNA in an R N A ^ D N A hybrid (25). This Mn^^-dependence shows activation at low concentrations of Mn2+, followed by inhibition at higher Mn^^ concentrations (Figure 1). The optimum activity is achieved in 5 |iM M n C l 2 , and is 30 % of the maximum activity in MgCl2 (data not shown). Maximum inhibition at 1 mM MnCl2 is -'20-fold inhibited relative to activity at 5 |LiM MnCl2. Activation and inhibition of RNase H'*' was indistinguishable from RNase HI (data not shown).

James L. Keck and Susan Marqusee

412

Figure 1. Mn^'^-dependence of E.coli RNase HI activity Assays were performed as described in Materials and Methods. Data points are the average of two assays with standard deviations shown as error bars.

10

1000

100

[MnCl2],|iM

Mit^-^-binding by E.coli RNase H*: EPR spectrometry was used to determine the stoichiometry and affinity of Mn^^ binding to E.coli RNase H*. Scatchard analysis of the binding data show that E.coli RNase H* (a cysteine-free version of E.coli RNase HI) has multiple Mn2+ binding sites. Two binding sites were determined with dissociation constants (Kd) of -15 |xM and --60 |iM (Figure 2). Figure 2. Equilibrium Miri^"^-binding to E.coli RNase H*^ V represents the fraction of b o u n d M i f ^ e r total ^•^^" RNase H* as o 05 - \ described in §j Materials and rp*^ 0 04 ~ Methods. Dashed 'Splines indicate the Jg 0.03best fit two lines ^ representrag the o.02 data points. K ^ measurements o.Ol are the inverse of the fits' abscissa. Q" 0

Measured Dissociation Constants Kdi z 15 |iM Kd2=60|iM

•

\ \ \ §"^

\ 1

i

1

1

1

1

i

0.2

0.4

0.6

0.8

1

1.2

1.4

V

1.6 1

1

Metal-Dependence of E. coli RNase H

413

Figure 3. Comparison of Mn 2+-activation to simulated activation curves ?

^ 0.5-

t

both metals required for activation

^ 0.4O *X2

>^

^

first metal activates second metal inhibits

> 0.3-1

"S 0.2-1 0^

»-4

S 0.1 H 1

10

[MnCl2],|iM

T

100

1000

Comparison of metal binding to dependence of the RNase H catalysis: Using the determined Mn2+-binding constants to E.coli RNase H*, the relative populations of enzyme with either 0, 1 or 2 Mn2+ ions bound as a function of [Mn^+Jtotal were determined. Figure 3 shows two simulations of the Mn2+ dependence of E.coli RNase H activity; one for the case where both metals are required for activation and one for the case where one metal is activating and one inhibiting. A comparison of Mn^+ activation of E.coli RNase HI (from Figure 1) shows the similarity between the data and the later model. The overall shape similarity of the two plots is striking. The highest metal affinity binding correlates well to in vitro activation and binding of the lower affinity metal correlates approximately to inhibition (Figure 3). Relative RNase H activity is scaled to represent the observation that maximum Mn^+-dependent activity is -^-0.3 that for maximum Mg2+-dependent activity (27). Differences between the real and theoretical activation data may imply differences in metal binding in the presence of substrate. Discussion It has been known for over 20 years that either Mn2+ or Mg2+ can activate E.coli RNase HI. However, differences between metal requirements in Mn2+ and Mg2+ are only now beginning to be understood. Here, we have shown that the Mn2+ requirement for E.coli RNase HI activity is in the low micromolar range. This value can be contrasted to the relatively high (-0.1 to 1 mM) Mg^ + concentrations required for activity (10,27). Further, we have

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demonstrated that the Mn2+-dependent RNase H activity can be inhibited with higher Mn2+ concentrations (> 5 |iM). Mn^+ inhibition at higher metal concentrations could be due to a n u m b e r of factors, including: (1) metal-induced conformational changes in the RNA»DNA hybrid substrate or (2) metal binding to the enzyme that reduce it's activity. Similar inhibition has been documented for E.coli RNase H Mg^^-dependent reaction, with the inhibition attributed to substrate-metal association (28). This interpretation was based on the fact that E.coli RNase H binds Mg^+ with a 1:1 stoichiometry (10) and that the binding constant for Mg2+ to nucleic acid is similar to the inhibition constant (29,30). It is possible however, that binding studies reveal a second Mg2+ binding site on the enzyme only in the presence of substrate. With Mn2+, we can correlate metal binding to both activation and inhibition. We therefore, support the idea that Mn^^ inhibits as a result of binding an inhibitory site on the enzyme. We have determined here that E.coli RNase H can bind multiple (presumably two) Mn^+ ions with KdS of --15 |LIM and --60 |LIM. Upon comparison of metal-binding with our activation/inhibition data, the simplest model is that the tightest binding metal activates the enzyme while the second metal inhibits the activity (Figure 3). If both metals were required for activity, presumably there would be no inhibition at higher metal concentrations. Binding of the first metal in the absence of substrate correlates well with activation, but binding of the second metal appears weaker without substrate (i.e. the apparent inhibitory Kd is less than the measured Kd of the second metal binding). This discrepancy may indicate that substrate is involved in complete formation of the second metal binding site. Mechanism of E.coli RNase H: The metal-dependence of the RNase H reaction mechanism is not well understood. Currently there are two primary mechanisms that have been proposed: a onemetal mechanism and a two-metal activation mechanism. In light of the information presented in this paper, and in the context of information that has been gathered on the RNase H family of enzymes, we hypothesize that the RNase H mechanism is a singledivalent catalyzed reaction that can be attenuated by a second metal binding event. This hypothesis encompasses all of the seemingly contradictory information that has been presented to defend both the one and two-metal mechanisms. Figure 4 diagrams the basis of the proposed mechanism. In the absence of metal, E.coli RNase H is completely inactive. Upon addition of metal at activating concentrations (< 5 |LiM Mn^+), the tight metal binding site is filled and the enzyme is optimally active. We assume that the tightest binding metal binds in the single Mg2+ site observed crystallographically (9). Upon increasing the metal

Metal-Dependence of E. coli RNase H

415

Figure 4. Hypothetical metal-binding in the E.coli RNase H active site

Aspl34

AsplO r

Asp70

Glu48

Glu48

Inactive

Inhibited

concentration (> 5 |iM Mn^+), the second metal binding site (assumed from the co-crystal structure of the HIV RNase H domain with Mn2+ (5)) becomes occupied and the enzyme is inhibited. What is the mechanism of metal-inhibition? Our current hypothesis is based on E.coli RNase H active-site mutagenesis results coupled with the observation of two Mn^^-binding sites in the HIV RNase H domain structure. In the one-metal mechanism. Asp 70 abstracts a proton from the attacking nucleophilic water and then needs to deprotonate to reset the enzyme for the next hydrolysis (3). This deprotonation is believed to occur by shuffling the proton to His 124, since solvent is not accessible to Asp 70 and His 124 is nearby (within 4 A). Mutagenesis of His 124 to Ala results in a 100-fold reduction of kcat (3), presumably since Asp 70 must deprotonate through a less efficient mechanism. If His 124 is a liganding element for the second metal binding site, its pKa would be expected to shift down upon metal binding, making it more difficult to protonate. The effect of this pKa shift would be to inhibit the proton-transfer from Asp 70 to His 124, and thus slow the overall kcat for the reaction. We are currently testing this mechanism t h r o u g h mutagenesis and structural studies of E.coli RNase HI in Mn^^. Acknowledgments: We thank Mark Rabenstein and Yeon-kyun Shin for assistance with the EPR measurements. This work was supported by a grant from the N.I.H. (GM53321).

References

1. Hostomsky, Z., Hostomska, Z. and Matthews, D. A (1993) Ribonucleases H in Nucleases (ed. Linn, S. M., Lloyd, R. S. and Roberts, R. J.) 2nd Ed.,pp. 341-76, Cold Spring Harbor Laboratory, Cold Spring Harbor NY 2. Nakamura, H., Oda, Y., Iwai, S., Inoue, H., Ohtsuka, E., Kanaya, S., Kimura, S., Katsuda, C , Katayanagi, K., Morikawa, K., Miyashiro, H. and Ikehara, M. (1991) Proc. Natl Acad. Sci. U.S.A., 88, 11535-9

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3. Oda, Y., Yoshida, M., and Kanaya, S., (1993) /. Biol Chem. 268, 88-92 4. Suck, D. and Oefner, C. (1986) Nature 321, 620-5 5. Davies, J. F., Hostomska, Z., Hostomsky, S., Jordan, S. and Mathews, D. A. (1991) Science 252, 88-95 6. Beese, L. and Steitz, T. A. (1991) EMBO ]. 10, 25-33 7. Derbyshire, V., Grindley, N. D. F., and Joyce, C. M. (1991) EMBO J. 10,17-24 8. Freemont, P. S., Friedman, J. M., Beese, L. S., Sanderson, M. R. and Steitz, T. A. (1988) Proc. Natl Acad. Sci. U.S.A. 85, 8924-8 9. Katayanagi, K., Okumura, M., and Morikawa, K. (1993) Proteins 17, 337-46 10. Huang, H. W. and Cowan, J. A. (1994) Eur. J. Biochem 219, 253-60 11. Oda, Y., Nakamura, H., Kanaya, S. and Ikehara, M. (1991) /. Biomol. Nmr. 1, 247-55 12. Kanaya, S., Kohara, Y., Miura, Y., Sekiguchi, A., Iwai, S., Inoue, H., Otsuka, E. and Ikehara, M. (1990) /. Biol Chem.,265, 4615-21 13. Skalka, A.-M., and Goff, S. P. (eds) (1993) Reverse Transcriptase, Cold Spring Harbor Laboratory, Cold Spring Harbor, NY 14. Tisdale, M., Schulze, T., Larder, B. A. and Moelling, K. (1991) /. Gen. Virol 72, 59-66 15. Yang, W. and Steitz, T. A. (1995) Structure 3,131-4 16. Venclovas, C. and Siksnys, V. (1995) Nature Struct. Biol 2, 838-41 17. Rice, P., Cragie, R., and Davies, D. R. (1996) Curr. Op. Struct. Biol 6, 76-83 18. Katayanagi, K., Miyagawa, M., Matsushima, M., Ishikawa, M., Kanaya, S., Ikehara, M., Matsuzaki, T. and Morikawa, K. (1990) Nature 347, 306-9 19. Yang, W., Hendrickson, W. A., Crouch, R. J. and Satow, Y. (1990) Science 249, 1398-405 20. Ariyoshi, M., Vassylyev, D. C , Iwasaki, H., Nakamura, H., Shinagawa, H. and Morikawa, K. (1994) Cell 78, 1063-72 21. Dyda, F., Hickman, A. B., Jenkins, T. M., Engelman, A., Craigie, R. and Davies, D. R. (1994) Science 266, 1981-6 22. Bujacz, G., Jaskolski, M., Alexandratos, J. and Wlodawer, A. (1995) /. Mol Biol 253, 333-46 23. Rice, P. and Mizuuchi, K. (1995) Cell 82, 209-20 24. Dabora, J. M. and Marqusee, S. (1994) Protein ScL 3,1401-8 25. Keck, J. L. and Marqusee, S. (1995) Proc. Natl Acad. Scl U.S.A. 92, 2740-4 26. Scatchard, G. (1949) Ann. N. Y. Acad. Scl 51, 660-72 27. Keck, J. L. and Marqusee, S. (1996) /. Biol Chem. 271, 19883-7. 28. Black, C. B. and Cowan, J. A. (1994) Inorg. Chem. 33, 5805-8 29. Cowan, J. A. (1991) /. Am. Chem. Soc. 113, 6025-32 30. Black, C. B. and Cowan, J. A. (1994) /. Am. Chem. Soc. 116, 1174-8

Crystal structure of avian sarcoma virus integrase with bound essential cations Jerry Alexandratos\ Grzegorz Bujacz^'^, Mariusz Jaskolski^'^ and Alexander Wlodawer^*, ^Macromolecular Structure Laboratory, NCI-Frederick Cancer Research and Development Center, ABL-Basic Research Program, Frederick, Maryland ^Faculty of Food Chemistry and Biotechnology, Technical University of Lodz, Lodz, Poland ^Center for Biocrystallographic Research, Institute of Bioorganic Chemistry, Polish Academy of Sciences, Poznaii, Poland

George Merkel, Richard A. Katz and Anna Marie Skalka Institute for Cancer Research, Fox Chase Cancer Center, Philadelphia, Pennsylvania

I. Introduction Retroviral integrase (IN) is a virus-encoded enzyme that catalyzes nonspecific insertion of viral DNA into multiple sites on host DNA (1-3). Since DNA integration is an essential step in the retroviral replication cycle, this enzyme is an attractive target for inhibition of human immunodeficiency virus (HIV), the causative agent of acquired immunodeficiency syndrome (AIDS). Work over the last several years has resulted in a general understanding of the enzymatic mechanism, but more detailed analyses have been hampered by the lack of precise structural information. The situation changed when the crystal structures of the catalytic domains of both HIV-1 IN (4) and avian sarcoma virus (ASV) IN (5,6) became available. Precise data on the interaction of these enzymes and the essential ligands are necessary for understanding the structural basis of the reaction mechanism and for guiding rational drug design. Members of the structurally related superfamily of enzymes that include RNase H, RuvC resolvase, MuA transposase, and retroviral integrase contain at least three acidic residues in the active site and require divalent cations, such as Mg^"^ or Mn^"^, for their enzymatic activity. However, the precise placement of cations is reported in the X-ray crystal structures of only two of these proteins, E. coli RNase H and HIV-1 RNase H. Details of the location of metal ions in the active site of retroviral integrases can enhance our understanding of the catalytic mechanism of these enzymes and their relationship to that of other members of the superfamily. We present the structure of ASV IN catalytic domain with the essential cations Mg^"^ or Mn^"^ bound in the active site. In addition, we present the structure of an inactive complex of the catalytic domain of ASV IN with Zn^"^. TECHNIQUES IN PROTEIN CHEMISTRY VIII Copyright © 1997 by Academic Press All rights of reproduction in any form reserved.

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418

II. Methods The expression strategy, purification from E. coli, and activities of the purified ASV IN 52-207 fragment have been described previously (7). ASV IN protein crystals were produced from 20% polyethylene glycol (PEG) solution by the hanging drop vapor diffusion method, also described previously (5). All conditions yielded tetragonal crystals with approximate cell dimensions of a = b = 66 A, c = 81 A, space group of P432i2, with one molecule in the asymmetric unit. Crystals were soaked in metal chloride solutions in a synthetic mother liquor for at least 3 days each. Concentrations of 10 mM MnCl2,100 and 500 mM MgCl2, and 100 mM ZnCl2 solutions produced complete occupancy, whereas the occupancy was only partial in 20 mM MgCl2. X-ray diffraction data were collected at room temperature on a MAR 300 mm image plate detector, using a Rigaku RU200 rotating anode operated at 50 kV and 100 mA (Table I). Data were processed with DENZO and scaled with SCALE?ACK (8). Variations in the unit cell parameters for different crystals were less than 1.25 A, even between metal complexes and low temperature native structures. Electron density maps, calculated with the program PROTEIN (9), were interpreted using FRODO (10). The model underwent multiple cycles of restrained structure-factor least-squares refinement using PROLSQ (11). In addition to protein atoms and water molecules, the final models also include a well-ordered HEPES molecule that cocrystallized with the native protein. At least 60 water molecules were added to each structure during refinement.

III. Results and Discussion The central, catalytic domain of ASV IN is a five-stranded mixed P-sheet flanked by five a-helices. The active site is characterized by the presence of the D,D(35)E motif of three carboxylate-containing amino acids, the last two of which Table I. Summary of data collection and refmement Protein

Cell dimensions (A)

Resolution (A)

R-factor

R-free

Root mean square deviation from Mg structure

Low temperature selenomethionine

a,b= 65.40 c= 80.41

6.0-1.70

0.139

0.208

0.27

Mg (500 mM)

a,b= 66.05 c= 81.65

8.0-1.75

0.150

0.191

""

Mn(lOmM)

a,b= 66.24 c= 81.60

8.0-2.05

0.130

0.189

0.14

Zn(lOOmM)

a,b= 66.08 c=80.96

10.0-1.95

0.176

—-

0.17

Crystal Structure of Sarcoma Virus Integrase

419

are separated by 35 residues. The two aspartate residues are located on strand pi and the end of strand P4, with these strands being part of the stable P-sheet core of the protein. There is a long 10-residue loop between strand P5 and helix a4, which has higher B-factors and slightly different conformations in the different structures. This loop extends out of the compact shape of the molecule and appears quite flexible. The third catalytic residue appears near the end of helix a4 at one end of this flexible loop. The structure of ASV IN complexed with divalent cations (Mn^"^, Mg^"^, and Zn^"*") was solved at the resolution of 1.70 - 2.05 A. This enzyme is active in the presence of either Mn^"^ or Mg^"*", with the activity higher in the former than in the latter, and is inactive in the presence of Zn^"^. After refinement, the structures of Mn^"^ and Mg^"^ complexes were nearly identical in their overall architecture and in the metal binding scheme. A single ion of either metal interacts with the aspartate side chains of the D,D(35)E catalytic center and uses four water molecules to complete its octahedral coordination (Fig. la). The metal-ligand distance is within a 2.1 - 2.4 A range for the Mn^"^ ion and 2.05 - 2.15 A for the Mg^'^ion. Glu-157 does not take part in binding of these cations. Only small adjustments take place in the active site of ASV IN upon binding of a metal cofactor. The Asp-64 carboxylate shifts less than 0.4 A compared with the uncomplexed enzyme (5). A slight rotation of the Asp-121 carboxylate shifts the oxygen atom about 1 A. Only the non-metal-binding side chain of Glu-157 in the Mn^"*" structure moves a greater distance. This result agrees with previous studies, which show that even conservative mutations to these residues abolish protein activity. The soaking experiments were performed with modest metal salt concentrations. The 10 mM Mn^"^ concentration was only three times above that used in activity assays, and the 100 mM Mg concentration was similarly proportional to the concentration used in in vitro activity assays. We are certain that under the conditions of these soaking experiments only one of these catalytic metal cations occupies the active site at any one time. Because the final Mn^"^ Fo-Fc map showed a peak height of 6.5 o above background, we are very confident of its location. Metal binding with even one third of this occupancy would have been easily visible. Even the structure obtained using an extremely high 500 mM Mg^"^ concentration did not indicate a second metal-binding site. It is not known at this time whether one or two metals are required for the integration reaction to proceed, although the detailed modeling of reactions catalyzed by nucleotidyl tranferases appears to require two metal ions (12). Since we could find only one divalent cafion in the complex, we decided to examine whether other divalent cations could be bound with different stoichiometry. Even though it is known that Zn^"^ does not activate integrases, we used this cation because zinc chemistry is very similar to that of Mg (13). Unexpectedly, we observed a structure with two Zn^"^ ions coordinated by all three active site residues, and with one water ligand interacting with each metal ion (Fig. lb). As in the other structures, we observed only minimal conformational changes for both aspartate side chains. One of the Zn^"^ ions appears in essentially the same location as the Mn'^'^ and Mg^"*" ions, only 0.36 A from the position of the former and 0.29 A from the latter, coordinating with Asp-64 and Asp-121, respectively. The coordination of the second active site metal involves the other carboxylate oxygen

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Figure 1. Stereo views of active sites of ASV IN complexed with metals, la. Electron density map of Mg coordinated with four water molecules. Two active site carboxylate oxygens and four waters create the octahedral coordination for the metal cation, lb. Electron density map of two Zv?^ ions with coordinating two water molecules. Four active site carboxylate oxygens and two water molecules coordinate the metal cations.

Figure 2. Ribbon diagram of the ASV IN catalytic domam, with explicitly shown active site side chains for both Zv?^ (black) and Mn^"^ (grey) complexes. The metal locations and corresponding coordinated water molecules are indicated in the same colors.

Crystal Structure of Sarcoma Virus Integrase

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of Asp-64, as well as Glu-157. Not surprisingly, the side chain of Glu-157 was observed to rotate, as this residue did not previously point into the active site. The fact that only a side chain rotation, with no backbone displacement, was needed for the second cation to bind explains why even conservative mutations of these active site residues inactivate IN completely. The coordination also seems to extend between the metal ions themselves, since the distance between the Zn^"^ ions is only 3.5 A. Both Zn^"^ ions are coplanar with the two carboxylate oxygens from Asp-121 and Glu-157 and the liganded waters. Each Zn^^ is located in the center of a triangle formed by coordination with a carbonyl oxygen, a water molecule, and the other cation, with Asp-64 coordinating both Zn^"^ ions from below this plane. The distance between any oxygen Hgand and the metal ion is within 2.1-2.4 A. The number of waters coordinating the metal bound by IN may be crucial for catalytic activity. If one water is replaced by an incoming DNA phosphate ligand, then this active site-Zn^"^ arrangement will not have a water molecule available for the hydrolysis reaction (14). Another Zn^"*" ion was found bound in a distant part of the structure, with His-103 and three additional water molecules, forming a more typical tetrahedral coordination. Binding of the Mg^"^, Mn^"^, and Zn^"*" ions does not lead to significant structural modifications in the active site or the overall protein architecture when compared with native ASV IN (Table I). This result indicates that metal-binding sites are preformed in this IN structure. The observed configuration of the D,D(35)E residues may represent a catalytically-competent active site (Fig. 2). When one divalent cation is bound, the active site side chains remain in the positions seen in the PEG native structures. Binding of two Zn^"^ cations also causes no change to the IN backbone. The side chain of Glu-157 rotates with respect to the conformation seen in the protein complexed with the other metals. However, this is completely consistent with the side chain conformation seen in the native structure of IN crystals grown from ammonium sulfate. These minor differences between the active sites of ASV IN with different cofactors seem to reflect a tendency for structural flexibility in the active site of integrases. The two Zn^"^ ions are observed at a similar concentration as one Mg^"^ ion, with no overall changes to the protein, indicating a possible mode of binding for Mg^"^ under other conditions. This observation supports the hypothesis that a second metal-binding site exists for Mg^'^/Mn^'^, but forms in the presence of substrate and/or other domains of the protein.

A. Comparison with related enzymes Although many enzymes that are active in the processing of nucleic acids, such as nucleases, DNA polymerases, or reverse transcriptases, have acidic residues in the active sites and require divalent cations for activity, such cations have been reported only for a few published structures. One published structure of reverse transcriptase from Moloney murine leukemia virus, (MMLV RT) shows a single metal bound in the active site (15), whereas none of the available structures of HIV-1 RT show bound metals. In addition, the structures of MMLV RT and E. coli RNase H with bound metals have been solved at a lower resolution than the

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same proteins without metals. As our data show, the quaUty of the metalcontaining ASV IN structures are as good as that of the apoenzyme. Since the electron density maps are of excellent quality, we can describe the active site with high accuracy. We have compared compared the structure of the ASV IN active site with the active sites of the other members of this superfamily for which metal complexes have been described or inferred, namely both HIV-1 and E. coli RNases H, and E. coli RuvC resolvase. As reported by Yang and Steitz (16), the similarity of the cluster of acidic residues forming the active sites of the RNase H enzymes is striking. With alignment based on conserved secondary structure elements, we have found that the best agreement in the active site of these enzymes is with ASV IN Asp-64. The placement and direction of the analogous carboxylates are very similar in RNases H and in ASV IN (Fig. 3). ASV IN Asp-121 is close to its equivalent in HIV-1 RNase H (17) and E. coli RNase H (18), whereas the side chain of the equivalent residue in E. coli RuvC resolvase (19) is more distant (not shown). The third residue of the cluster, ASV IN Glu-157, is also in quite good agreement among these enzymes. The other acidic residues in this region do not have counterparts in ASV IN. Similar to what we have observed for ASV IN, the residues in the active site of £. coli RNase H are moved only slightly upon binding of the metal, shifting Ca atoms less than 0.4 A when comparing structures without (20) and with (18) divalent cations; the side chain acidic groups shift no more than 1.5 A. Interestingly, the two residues that coordinate the Mg^"^ ion move less than the other carboxylates, indicating that the part of the active site directly coordinating the metal ion has an invariant character. A similar case is noted with a comparison of the metal-bound and unbound active sites of MMLV RT (18,20) and HIV-1 RT (21), with less than a 1.5 A r.m.s. deviation among the three active site residues. The sole Mg^"*" ion reported for E. coli RNase H (22) is complexed by Asp10 (Asp-64) and by Glu-48 (no ASV IN equivalent). Although no precise data on the location of a divalent cation are available for RuvC resolvase, a Mn^"^-binding site apparently exists between Asp-7 (Asp-64) and Asp-141 (Glu-157) (19). ZnMn

D121 D98

^ 1 D64D43 m D64D.

^ E157E149

0153N145 Figure 3. Comparison of the active sites of the HIV-1 RNase H-Mn^"^ complex (16) (black) with the ASV EST-Zn^"^ complex (grey) shows the excellent alignment of catalytic residues and metal cations.

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Structural alignment of the crystallographically determined structures of ASV IN complexed with Zn^"^ and of the HIV-1 RNase H complexed with Mn^"^ was carried out using ALIGN (23). The general architecture of the ASV IN and RNase H monomers is significantly different, but it is possible to superimpose structurally conserved regions, three a-helices and one P-strand, which contain the active site and nearby structurally important residues. This alignment reveals a surprisingly good superposition of the catalytic residues, with the r.m.s. deviation of 1.3 A for the 28 atom pairs. The positions of the two Zn^"^ ions in ASV IN are very close to the two Mn^"^ ions in RNase H (17), which are directly coordinated by the carboxylates of Asp-43 (equivalent to Asp-64 in ASV IN) and Asp-98 (equivalent to Asp-121 in ASV IN), and between Asp-43 and Asp-149 (equivalent to Glu-157 in ASV IN). The distances between the two pairs of cations are 0.38 A for the Zn^"^ ions bound between the two aspartates and 0.48 A for the other Zn^"*" ion, less than the r.m.s. deviations between the protein atoms (Fig. 3). Although the three most highly conserved acidic residues are present in similar locations in all of these enzymes, the exact relationships between them are not strictly preserved. However, for the two (quite divergent) RNase H enzymes, the maximum differences in the positions of the carboxylates do not exceed 1.5 A, despite some disorder reported in the vicinity of the active site of the isolated HIV1 RNase H domain from HIV-1 RT (17). The minimal influence of the presence of the divalent cation on the disposition of residues in the active site of RNase H and ASV IN is mirrored in MMLV RT, where the differences observed in the positions of the three critical aspartates in the active site are not larger than 0.4 A when metal bound and unbound structures are compared. In the case of MMLV RT, however, the quality of the difference Fourier map is not sufficient to determine the details of the coordination of the metal. The similarities between the active sites of MMLV RT and ASV IN include the interaction of only two of the three carboxylates with a single Mn ion present in the active site. We have presented here the structure of the catalytic domain of ASV IN complexed with three different divalent cations. These results clearly show that the active site of this enzyme is preformed, in that only relatively small movements of side chains and no shifts of the main chain are needed in order to provide an environment suitable for cation binding. This is in contrast with the related core HIV-1 IN (4), in which no binding of the divalent cations could be shown by crystallographic means, and in which the constellation of active site residues differs significantly from their counterparts in ASV IN. However, antibody-binding experiments have shown that HIV-1 IN undergoes a conformational change when incubated with divalent cation cofactors (Asante-Appiah, E. and Skalka, A. M., personal communication). These results are consistent with the notion that the activity can be modulated by transitional order-disorder phenomena involving the active site, and that such conformational changes are be different for enzymes obtained from different sources. Although only a single divalent cation was observed upon soaking ASV IN in Mn^"^ and Mg^"^, the unexpected observation of two Zn^"^ ions binding to the active site of ASV IN could provide indirect proof of the hypothesis postulating the utilization of two cations for catalytic activity.

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Acknowledgements Research sponsored in part by the National Cancer Institute, DHHS, under contract with ABL. Other support includes National Institutes of Health grants CA47486 and CA06927, a grant for infectious disease research from Bristol-Myers Squibb Foundation, and an appropriation from the Commonwealth of Pennsylvania. The contents of this publication do not necessarily reflect the views or policies of the Department of Health and Human Services, nor does mention of trade names, commercial products, or organizations imply endorsement by the U.S. Government.

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8. 9. 10. 11. 12. 13. 14. 15.

16. 17.

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Katz, R. A., and Skalka, A. M. (1994). The retroviral enzymes. Annu. Rev. Biochem. 63, 133-173. Goff, S. P. (1992). Genetics of retroviral integration. Annu. Rev. Genet. 26, 527-544. Vink, C , Groeneger, O. A. M., and Plasterk, R. H. (1993). Identification of the catalytic and DNA-binding region of the human immunodeficiency virus type I integrase protein. Nucleic Acids Res. 1\, 1419-1425. Dyda, F., Hickman, A. B., Jenkins, T. M., Engelman, A., Craigie, R., and Davies, D. R. (1994). Crystal structure of the catalytic domain of HIV-1 integrase: Similarity to other polynucleotidyl transferases. Science 266, 1981-1986. Bujacz, G., Jaskolski, M., Alexandratos, J., Wlodawer, A., Merkel, G., Katz, R. A., and Skalka, A. M. (1995). High resolution structure of the catalytic domain of the avian sarcoma virus integrase. / Mol Biol. 253, 333-346. Bujacz, G., Jaskolski, M., Alexandratos, J., Wlodawer, A., Merkel, G., Katz, R. A., and Skalka, A. M. (1996). The catalytic domain of avian sarcoma vims integrase: conformation of the active-site residues in the presence of divalent cations. Structure 4, 89-96. Kulkosky, J., Katz, R. A., Merkel, G., and Skalka, A. M. (1995). Activities and substrate specificity of the evolutionarily conserved central domain of retroviral integrase. Virology 206, 448-456. Otwinowski, Z. (1992). An Oscillation Data Processing Suite for Macromolecular Crystallography, Yale University, New Haven. Sheriff, S. (1987). Addition of symmetry-related contact restraints to PROTIN and PROLSQ. J. Appl Crystallogr. 20, 55-57. Jones, T.A. (1985). Interactive computer graphics: FRODO. Methods Enzym. 115:157-171. Hendrickson, W. A. (1985). Stereochemically restrained refinement of macromolecular structures. Methods Enzymol. 115, 252-270. Steitz, T. A. (1993). DNA- and RNA-dependent DNA polymerases. Curr. Opin. Struct. 5/o/. 3,31-38. Cotton, F., and Wilkinson, K. (1988). Advanced Inorganic Chemistry (5th edition, Wiley-Interscience) Beese, L. S., and Steitz, T. A. (1991). Structural basis for the 3'-5' exonuclease activity of Escherichia coli DNA polymerase I: a two metal ion mechanism. EMBOJ. 1, 25-33. Georgiadis, M. M., Jessen, S. M., Ogata, C. M., Telesnitsky, A., Goff, S. P., and Hendrickson, W. A. (1995). Mechanistic implications fi*om the structure of a catalytic fragment of Moloney murine leukemia virus reverse transcriptase. Structure 3, 879-892. Yang, W., and Steitz, T. A. (1995). Recombining the structures of HTV integrase, RuvC and RNase H. Structure 3, 131-134. Davies, J. F.,II, Hostomska, Z., Hostomsky, Z., Jordan, S. R., and Matthews, D. A. (1991). Crystal structure of the ribonuclease H domain of HTV-1 reverse transcriptase. Science 252, 88-95. Yang, W., Hendrickson, W. A., Crouch, R. J., and Satow, Y. (1990). Structure of ribonuclease H phased at 2 A resolution by MAD analysis of the selenomethionyl protein. Science 249,139^-1405.

Crystal Structure of Sarcoma Virus Integrase 19.

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Ariyoshi, M., Vassylyev, D. G., Iwasaki, H., Nakamura, H., Shinagawa, H., and Morikawa, K. (1994). Atomic structure of the RuvC resolvase: A HoUiday jxmction-specific endonuclease from^. colL Celin, 1063-1072. Katayanagi, K., Miyagawa, M., Matsushima, M., Ishikawa, M., Kanaya, S., Nakamura, H., Ikehara, M., Matsuzaki, T., and Morikawa, K. (1992). Structural details of ribonuclease H from Escherichia coli as refined to an atomic resolution. J. Mol Biol. 11^, 1029-1052. Unge, T., Knight, S., Bhikhabhai, R., Lovgren, S., Dauter, Z., Wilson, K., and Strandberg, B. (1994). 2.2 A resolution structure of the amino-terminal half of HIV-1 reverse transcriptase (fingers and palm subdomains). Structure 2, 953-961. Katayanagi, K., Okumura, M., and Morikawa, K. (1993). Crystal structure of Escherichia coli Rnase HI in complex with Mg^"*" at 2.8 A resolution: proof for a single Mg^''"-binding site. Proteins 17: 337-346. Satow, Y., Cohen, G. H., Padlan, E. A., and Davies, D. R. (1986). Phosphocholine binding immunoglobulin Fab McPC603: An X-ray diffraction study at 2.7 A. J. Mol Biol 190, 593-604.

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Multidimensional NMR Studies of an Exchangeable Apolipoprotein and Its Interactions with Lipids

Jianjun Wang^, Daisy Sahoo^, Dean Schieve^, Stephane M. Gagne§, Brian D. Sykes§ and Robert O. Ryan^

^Lipid and Lipoprotein Research Group, »Protein Engineering Network Centres of Excellence, Department of Biochemistry, University of Alberta Edmonton, Alberta, Canada T6G 2S2

I. INTRODUCTION Exchangeable apolipoproteins are a class of functionally important proteins which play a key role in plasma lipoprotein metabolism. In this capacity they have been associated with several human disorders, including hyperlipidemia and cardiovascular disease (1,2). Apolipophorin-III (apoLp-III) is a model exchangeable apolipoprotein derived from the insect Manduca sexta (166 residues, Mr 18,380). ApoLp-III is a major hemolymph protein in the adult life stage and functions in lipid transport during sustained flight (3,4). Biophysical studies demonstrate that apoLp-III is a soluble monomeric protein at concentrations of 15 mg/ml (5). While the tertiary structure of M. sexta apoLp-III has not been solved. X-ray crystallography of apoLp-III from Locusta migratoria reveals a globular structure comprised of a bundle of five elongated amphipathic a-helices which are connected by short loops (6). A similar molecular architecture was also found for the 22 kDa N-terminal fragment of human apolipoprotein E (7). The crystal structure of L. migratoria apoLp-III was obtained for the protein in its lipid-free state. The lipid-bound structure of apoLp-III, however, is more interesting since it represents the active form of the protein. To date, no detailed structural reports for exchangeable apolipoproteins in complex with lipid have been reported. The crystal structure of lipid-free apoLp-III demonstrated that the five amphipathic heUces orient in such a way that their hydrophobic faces are directed toward each other to form a hydrophobic core while the hydrophilic faces of the helices are exposed to solvent. It has been hypothesized that, upon binding to a TECHNIQUES IN PROTEIN CHEMISTRY VIII Copyright © 1997 by Academic Press All rights of reproduction in any form reserved.

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lipid surface, the protein undergoes a major conformational change that results in opening of the helix bundle, with exposure of the hydrophobic surfaces of the helices which contact the Hpid (6). This putative conformational change is depicted in Figure 1:

Lipid Binding

Figure 1. Open conformation model of exchangeable apolipoproteins upon lipid-binding.

Since M. sexta apoLp-in is a well-behaved member of the exchangeable apolipoprotein family in terms of its physico-chemical properties, it represents a good candidate for investigation of the molecular details of the exchangeable apolipoproteins associated with lipid binding. A study of this system may reveal a general mechanism for exchangeable apolipoprotein-Hpid interaction. In a manner similar to all exchangeable apolipoproteins, apoLp-III resists crystallization when complexed with lipid. Thus, NMR is the only potentially useful high resolution technique to investigate structural changes of apoLp-III which accompany lipidbinding. To date, no NMR structures of exchangeable apolipoproteins have been reported. In order to carry out 3D and/or 4D-NMR experiments, however, the protein of interest must be either ^^N and/or ^^C-isotope labeled. Isotope labeling strategies require an efficient bacterial expression system and such a system has recendy been developed in this laboratory for apoLp-III from M. sexta (8). The present study describes the results of labeling experiments and presents a useful method to incorporate ^^N specifically and exclusively into peptide backbone amide nitrogens. 3D-NMR experiments have been performed on isotopically labeled apoLp-m and nearly complete assignment has been achieved. In addition.

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preliminary experiments provide direct experimental evidence in support of a significant conformational change in apoLp-III upon interaction with lipid. II. METHODS Materials. 1^NH4C1, ^^N-Leucine, ^^N-Glycine, ^^N-Valine and ^^N-Lysine were purchased from Cambridge Isotope Laboratories (Andover, MA). i^C-C^-Glucose and ^H-dodecylphosphocholine were obtained from Isotec. Inc. (Miamisburg, Ohio). Unlabeled amino acids were obtained from Sigma Chemical Co. (St. Louis, MO) Bacterial expression and isotope-labeling of recombinant apoLp-III. The coding sequence of M. sexta apoLp-III was cloned into the pET expression vector (Novagen Corp., Madison, WI) directly downstream from the pelB leader sequence cleavage site. Introduction of the plasmid vector into E. coli BL21(DE3) permits high level expression upon induction with 1 mM isopropyl 6-D thiogalactopyranoside (IPTG). Significant amounts of recombinant apoLp-III were secreted into the culture medium during expression and protein was isolated from the culture supernatant following five hours incubation at 30°C. Typically, a one liter cell culture produces 150 - 200 mg of pure apoLp-III (8). The purification procedure was essentially as described by Ryan et al. (8). Isotope-labeling of apoLp-III used M9 minimal media with l^NILtCl for ^^N-uniform labeling; 15NH4Cl/^^C6-glucose for l^N/l^C uniform labeling; I5]sj4eucine/1^NH4C1 for ^^N specific backbone nitrogen labeling; ^^N-amino acid of interest/19 other ^^N-amino acids for specific amino acid ^^N-labeling. NMR Spectroscopy. NMR experiments were carried out at 30 °C on a Varian Unity 600 spectrometer equipped with three channels, a pulse-field gradient triple resonance probe with an actively shielded z gradient and a gradient amplifier unit. NMR sample concentrations ranged from 0.5 to 1.1 mM, pH. 6.5±0.1, with 250 mM phosphate buffer and 0.5 mM NaN3. 2D ^H-^^N HSQC spectra were recorded using the enhanced sensitivity mode with 8 - 3 2 transients (9). Triple resonance HNCACB (10) and CBCA(CO)NNH (10) 3D-NMR spectra, recorded on an uniformly l^N/^^C-labeled H2O sample with 8 - 1 6 transients, correlates backbone amide protons of residue / with CA and CB atoms of residue / (HNCACB) and i-1 (HNCACB, CBCA(CO)NNH) for the backbone sequential assignment. l^N-edited NOESY (9) and l^N-edited TOCSY (9) were also acquked on an uniformly l^N-labeled H2O sample with 8-12 transients, for both backbone and sidechain assignments. A mixing time of 150 ms was used for ^^N-edited NOESY experiments and a mixing time of 59 ms was used for l^N-edited TOCSY

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experiments. Pulse field gradient HCCH-TOCSY (11) and simultaneous ^^N- and l^C-edited NOESY (12) were acquired for the sidechain assignment. A mixing time of 100 ms was used for simultaneous l^N- and ^^C-edited NOESY. Titrations of ^H-dodecylphosphocholine (DPC) to specific amino acid l^N-labeled apoLp-HI samples were monitored by 2D ^H-^^N HSQC spectra at pH 6.9 - 7.0 in order to investigate structural changes induced by lipid-binding. Electrospray ionization mass spectrometry. Molecular weight determinations for control and isotope enriched apoLp-IIIs were made using a VG quattro electrospray mass spectrometer (Fisons Instruments, Manchester, UK). Molecular weights were determined as the mean value calculated for several multiply charged ions within a coherent series. The instrument was calibrated using the series of ion peaks from horse heart myoglobin with a molecular mass of 16,951 daltons. Calculated masses were derived from the amino acid sequence using the program MacPro Mass (Terry Lee, City of Hope, Duarte CA). III. RESULTS and DISCUSSION ^^N Isotope-labeling Strategies In order to pursue heteronuclear multidimensional NMR experiments, a bacterial system for expression of apoLp-III has been developed which allows facile production of 150 - 200 mg/L l^N-labeled apoLp-III or 100 - 125 mg/L l5N/l3C-double labeled apoLp-III. Figure 2, panel A shows the iR-l^N HSQC spectrum of a 1.0 mM solution of lipid-free, uniformly ^^N-labeled apoLp-III. Panel A also indicates that, although the chemical shift dispersion in the ^Hdimension is rather small (6.5 ppm to 9.5 ppm), it is generally upfield shifted, consistent with the fact that the protein secondary structure is predominantly a-helix (13). The chemical shifts in the ^^N-dimension are well-dispersed which results in good separation of the overall crosspeaks. However, certain regions in the spectrum are still crowded as shown in Figure 2. The upper right comer of the HSQC spectrum of ^^N-labeled apoLp-III shown in Figure 2, panel A contains numerous doublet crosspeaks which are derived firom side chain amines of glutamine and asparagine residues. Since apoLp-III is rich in glutamine and asparagine (25 of the 166 amino acids), this region of the spectrum is

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crowded. In considering possible approaches to selectively label backbone amide nitrogens, we speculated that although bacteria efficientiy and readily transaminate nitrogen to a-amino nitrogens of other amino acids, they may prefer NH4CI as a precursor of Gin and Asn side chain amine nitrogens. Interestingly, when bacteria were grown in media containing ^^N-leucine and unlabeled NH4CI, specific labeling of backbone amide nitrogens was achieved (Panel B, Figure 2). The presence of unlabeled NH4CI is essential to this process because it provides an alternative, preferentially utilized, biosynthetic precursor of side chain nitrogen atoms. The isotope-labeling strategy shown in Panel B, Figure 2 is the first report, to our knowledge, which allows for specific isotope-labeling of the backbone amide nitrogen atoms of a protein. As a reference control, bacteria harboring the apoLpni/pET plasmid were cultured in medium containing ^^N-leucine as the sole nitrogen source. ApoLp-III obtained by this isotope-labeling method gave a HSQC spectrum which was indistinguishable from the spectrum shown in Figure 2, panel A (data not shown). This result confirms that bacteria are capable of redistributing nitrogen derived from ^^N-leucine into all backbone and side chain nitrogens in this protein. Importantly, when the uniform labeled spectrum is overlayed with the specifically backbone amide labeled spectrum, it is apparent that several backbone amide resonances are masked by the abundant side chain amine nitrogen resonances. Thus, this labeling strategy simphfies HSQC spectra, enabling identification of crosspeaks from backbone amides that otherwise overlap with crosspeaks from side chain amines. Spectroscopic methods (DEPT-HMQC and DEPT-SQC) have been developed which permit resolution of ^^NH and 15NH2 groups in proteins (14,15). However, the metabolic labeling strategy described herein offers an attractive altemative method, permitting specific enrichment of backbone amide nitrogens with l^N, effectively eliminating glutamine and asparagine side chain NH2 resonance, and thus simpHfying the spectrum. Mass spectrometric analysis of ^^Nbackbone amide nitrogen labeled apoLp-III indicated an isotope enrichment of 50%, demonstrating that unlabeled NH4CI competes with l^N-leucine derived nitrogens for incorporation into the backbone nitrogen atoms in the protein. By contrast, an isotope enrichment of > 95% was observed for uniformly l^N-labeled apoLp-III using l^NIL^Cl as the sole nitrogen source in M9 minimal media. It has been proposed that the amphipathic a-helix is the putative lipidassociating motif in exchangeable apolipoproteins (16,17). Further, it is generally accepted that binding of exchangeable apoHpoproteins to lipid surfaces involves contact of the apolar faces of the a-heUces, with the polar faces of the helices exposed to the solvent. Taken together with the known structures of L. migratoria apoLp-in and the 22 kDa N-terminal fragment of human apolipoprotein-E in the absence of lipid, it is apparent that a significant conformational change must

NMR Studies of an Exchangeable Apolipoprotein

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accompany lipid association. Hence, we are particularly interested in residues located in the apolar and/or polar faces of the helices. Leucine is a typical hydrophobic residue and lysine is a typical hydrophilic residue. In M, sexta apoLpni, there are 11 leucines and 23 lysines which are dispersed along the entire length of the protein sequence. These two residues are proposed to be located on the apolar and polar faces of the amphipathic helices, respectively. To accomplish specific labeling of the 11 leucine residues in apoLp-III, we cultured bacteria in M9 minimal media containing ^^N-leucine and the 19 other unlabeled amino acids. Figure 2, panel C reveals a spectrum consisting of 11 well-separated resonances, consistent with the conclusion that specific labeling of the 11 leucine residues in apoLp-ni has been achieved. For l^N/l^C-double labeling of apoLp-III, the amount of l^C6-glucose was optimized. Normally, minimal media contains 2.4 - 3.0 g glucose/L. However, it was found that efficient overexpression of ^^N/^^c double labeled apoLp-III can be achieved with as little as 1.2 g/L of ^3C6-glucose, significantiy reducing the cost of isotope-labeling. Strategy for Complete Assignment of ApoLp-III In order to completely assign the NMR spectra, heteronuclear 3D-NMR experiments have been performed. Experiments essential for the backbone assignment are the l^N-edited NOESY, HNCACB and CBCA(CO)NNH. These spectra allow for nearly complete assignment of the backbone atoms of apoLp-III. For an a-helical protein, strong/medium intensity of dNN(i>i±l) NOEs can usually be observed (18). The strategy for sequential assignment of apoLp-III took advantage of the dNN connectivities and used NH-NH walking based on the ^^Nedited NOESY spectrum. Figure 3 gives an example of data obtained from the NHNH walking strategy. This figure shows strip plots extracted from the 150 ms mixing time ^^N-edited NOESY spectrum of apoLp-III for residues 50 - 59. This strip plot clearly demonstrates the NH-NH walking strategy and, using this approach, we assigned 80% of the backbone N, HN atoms, as well as 80% Ha atoms with the help of a ^^N-edited TOCSY spectrum. Triple resonance HNCACB and CBCA(CO)NNH spectra correlate backbone amide protons of residue / with CA and CB atoms of residues / and /-I. Spectra obtained, which yielded crosspeaks correlating 8 5 - 9 5 % of apoLp-III residues, were used to confirm the assignment obtained by NH-NH walking, complete the remaining assignment and correct two mistakes. Using a combination of the above mentioned 3D-experiments, it was possible to assign nearly all of the backbone atoms of apoLp-III. Sidechain atom assignment was completed using 3D HCCH-TOCSY and simultaneous ^^N/^^C NOESY spectra (at 100 ms mixing time).

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The complete assignment of apoLp-III will be reported elsewhere. However, Figure 2, Panel C shows the assignment of 11 leucine residues of apoLp-IIL Lipid binding induced conformational changes in ApoLp-III In order to investigate structural changes in apoLp-III induced by lipid-binding, we prepared specifically ^^N-valine and l^N-lysine labeled protein using a labeling strategy similar to that described for l^N-ieucine above. Compared to leucine labeling, the efficiency of label incorporation was greater with l^^-lysine, consistent with the work of others (19). For l^N.yaline specific labeling, in addition to strong crosspeaks from direct incorporation of l^N-valine, several weaker crosspeaks were found in HSQC spectra. These data suggest scrambling of valine's a-amino nitrogen to other amino acids. This interpretation is consistent with the fact that, in bacteria, valine can donate its amino nitrogen direcdy to aketoglutarate to form glutamic acid. However, we can easily identify the crosspeaks from valine in l^N-^H HSQC spectra due to the much stronger intensity

NMR Studies of an Exchangeable Apolipoprotein

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of valine crosspeaks versus those arising from scrambling of the label. The assignment derived from 3D-NMR experiments also allows us to confirm the valine assignments. D38-dodecylphosphocholine (DPC) micelles were used to provide a lipid environment. This molecule, which possesses a phosphocholine head group and a single Ci2 hydrocarbon chain, mimics the phospholipid component of lipoprotein surface monolayers. About 40 DPC molecules form a micelle which has molecular weight of about 16 kDa. Thus, the apoLp-HI/DPC (1:1 protein/micelle) complex

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has a predicted molecular weight of about 34.4 kDa. In addition, a final molar ratio between apoLp-IH and DPC should be above 1:40 for the study of the lipid-induced conformational changes of apoLp-HI since under this condition, DPC concentration is much higher than its CMC, and DPC micelle concentration is also higher than the protein concentration. Hence, a 1:1 apoLp-IU/DPC complex will be obtained. For simplicity, specifically ^^N-valine and ^^N-lysine labeled apoLp-IIIs were used to study conformational changes of apoLp-HI upon lipid-binding. It is postulated that these two residues (and the helices they are associated with) will undergo significant repositioning when apoLp-EI binds to lipid. ^H-^^N HSQC spectra were used to monitor chemical shift changes of the crosspeaks arising from these two residues. Figure 4 shows the HSQC spectra of specifically ^^N-lysine labeled recombinant apoLp-HI in the absence and presence of ^H-DPC micelles. In this figure, the dark contour crosspeaks represent apoLp-III in the lipid-free helix bundle conformation whereas the light contour crosspeaks represent apoLp-III in the presence of DPC micelles. It is noteworthy that specifically ^^N-lysine labeled apoLp-in in its lipid-free state gives rise to only 16 distinct resonances, of which at least four crosspeaks represent more than one lysine residue due to resonance overlap (see the assignment in Figure 4). In addition, two lysine HSQC crosspeaks (K71, K136) were missing in Figure 4 due to the fast exchange of the amide protons of these two lysine residues under the experimental conditions we used for lipid titration (pH 6.9 - 7.0). The DPC:apoLp-ni mole ratio was 1:45 in Figure 4, which is well above the DPC critical micelle concentration of 1.3 mM. I^N-^H HSQC experiments were carried out on both samples to evaluate the effect of ^H-DPC titration. Our goal was to follow the chemical shift changes of each crosspeak shown in Figure 4 and, ultimately, obtain the assignment of lysine and valine residues in the lipid-bound state. Interestingly, these two samples behave differently upon DPC titration in terms of relative HSQC crosspeak chemical shift changes. While the valine crosspeaks are less sensitive to DPC titration, lysine crosspeaks are extremely sensitive. Small amounts of DPC, even less than its CMC (100 |j,g per 3 mg apoLp-III), cause dramatic changes in the chemical shift of lysine resonance, making it difficult to follow the changes in individual crosspeaks. For this reason, we do not show the assignment of apoLpIII lysine residues in the lipid-bound state (Figure 4). On the other hand, DPC titration-induced chemical shift changes in valine crosspeaks were more easily followed which allows us to obtain the assignment of valine residues of apoLp-III at its lipid-bound state. Since the DPC micelle concentration in Figure 4 is higher than the protein concentration, a 1:1 ratio of protein/DPC micelle complex is expected. In general, crosspeaks observed with lipid-bound apoLp-III are broader than those obtained with apoLp-III in the lipid-ft"ee state, consistent with association of the protein to the micellar surface. The dramatic differences in resonance

NMR Studies of an Exchangeable Apolipoprotein

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distribution between the lipid-free and lipid-bound apoLp-HI provides strong direct experimental support for the concept that lipid association is accompanied by a significant protein conformational change (5-6,20). Further detailed structural studies are currently in progress to characterize the molecular details of the lipidassociated conformation of apoLp-UI in terms of the structural model depicted in Figure 1. AKNOWLEDGEMENTS We thank Dr. Bill Bachovchin and David Corson for helpful discussions. ROR is a Senior Scholar of the Alberta Heritage Foundation for Medical Research and Medical Research Council of Canada Scientist. This work is supported by a grant from the Medical Research Council of Canada.

REFERENCES l.Weisgraber, K.H. (1994) Adv. Protein Chem.. 45, 249-302. 2.Weisgraber, K.H., Pitas, R.E. and Mahley, R.W. (1994) Curr. Opin. Struct. Biol, 4, 507-515. 3. Blacklock, B.J. and Ryan, R.O. (1994) Insect Biochem. Mol. Biol., 24, 855-873. 4. Ryan, R.O. (1994) Curr. Opin. Struct. Biol., 4, 499-506. 5. Kawooya J.K., Meredith, S.C, Wells, M.A., Kezdy, F.J. and Law, J.H. (1986) /. Biol Chem., 261, 13588-13591. 6. Breiter, D.R., Kanost, M.R., Benning, M.M., Wesenberg, G., Law, J.H., Wells, M.A., Rayment, I. and Holden, H.M. (1991) Biochemistry, 30, 603-608. 7. Wilson, C, Warden, M.R., Weisgraber, K.H., Mahley, R.W. and Agard, D.A. (1991) Science, ISl, 1817-1822. 8. Ryan, R.O., Schieve, D., Wientzek, M, Narayanaswami, V., Oikawa, K., Kay, CM. and Agellon, L.B. (1995) /. Lipid Res. 36, 1066-1072. 9. Zhang, O., Kay, L. E., Olivier, J. P. and Forman-Kay, J. D. (1994). J. Biomol-NMR 4, 845-858. 10. Muhandiram, D. R. and Kay, L. E. (1994). J. Magn. Reson. B103, 203-216. 11. Kay, L. E., Xu, G. Y., Singer, A. U., Muhandiram, D. R. and Forman-Kay, J. D. (1993). J. Magn. Reson. BlOl, 333-337. 12. Pascal, S. M., Muhandiram, D. R., Yamazaki, T., Forman-Kay, J. D. and Kay, L. E. (1994) /. Magn. Reson. B103, 197-201. 13. Ryan, R.O., Oikawa, K. and Kay, CM. (1993) /. Biol. Chem., 268, 1525-1530. 14. Kessler, H., Schmeider, P. and Kurz, M. (1989) /. Magn. Reson. 85, 400-405. 15. Tate, S.-L, Masui, Y. and Inagaki, F. (1991) /. Magn. Reson. 94, 625-630. 16. Segrest, J.P., Jackson, R.L., Morriseu, J.D., and Gotto, A.M.Jr. (1974) FEBS Lett., 38, 247-253.

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17. Segrest, J. P., Garber, D.W., Brouillette, C.G., Harvey, S.C. and Anantharamaiah, G.M. (1994).

Adv. Protein Chem. 45, 303-369.

18. Wuthrich. K. (1986) NMR of Proteins and Nucleic Acids. John Wiley & Sons, N. Y. 19. Muchmore, D.C., Mcintosh, L.P., Russell, C.B., Anderson, D.E. and Dahlquist, F.W. (1989). Methods EnzymoL, 177, 44-73. 20. Wientzek, M., Kay, CM., Oikawa, K. and Ryan, R.O. (1994) / . Biol. Chem., 269, 46054612.

NMR Methods for Analysis of CRALBP Retinoid Binding* Linda A. Luck"', Ronald A. Venters^, James T. KapronS, Karen E. Roth^, Seth A. Barrows"•, Sara G. Paradis'' and John W. Crabb3 ^Department of Biology, Clarkson University, Potsdam, NY 13699 Duke University NMR Center, Duke University, Durham, NC 27708 ^Protein Chemistry Facility, W Alton Jones Cell Center, Lake Placid, NY12946

I. Introduction Cellular retinaldehyde-binding protein (CRALBP) may play a key role in visual pigment regeneration as a substrate carrier/routing protein In the visual cycle, mediating the conversion of 11-c/s-retinol to 11-c/sretinaldehyde through interaction with an 11-c/s-retinol dehydrogenase in the retinal pigment epithelium (Saari et al., 1994). The protein exhibits retinoid stereoselectivity, only binding 11-c/s retinoids with high affinity and 9-c/s-retinaldehyde with lower affinity. 11-c/s-retinaldehyde bound to CRALBP is less susceptible to photoisomerization than when bound to rhodopsin (Saari and Bredberg, 1987). No evidence has been found for covalent linkage between retinoid and CRALBP. Toward identification of the CRALBP retinoid-binding pocket and definition of the structural properties of the protein that provide high ligand stereoselectivity and low photosensitivity, solution state NMR analysis has been initiated using human recombinant CRALBP labeled by biosynthetic isotope incorporation. A combination of heteronuclear gradient-enhanced ""SN N M R and one dimensional ^^F and "^^c NMR methods coupled with improved isotope incorporation methods and mass spectrometry, have proven to be complimentary approaches for characterizing CRALBP-ligand interactions. While these methods have been used separately elsewhere, usually as a primary approach to structural problems, here we emphasize the complimentarity of the *Thls work was supported In part by USPHS grant EY06603. TECHNIQUES IN PROTEIN CHEMISTRY VIII Copyright © 1997 by Academic Press All rights of reproduction in any form reserved.

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techniques and the advantages of combining the methods for studying protein-ligand interactions. Such protein biotechnology is suitable for characterizing a variety of protein-ligand interactions and is becoming more accessible through specialized biomolecular resource facilities.

II. Materials and Methods Human Recombinant CRALBP (rCRALBP). All human recombinant CRALBP used in this study was expressed as a fusion protein (339 residues, Mr=39,110) in bacteria [E c o / / s t r a i n BL21(DE3)LysS] with a His-tag N-terminal extension using the pET 19b vector, labeled with 11 -c/s-retinaldehyde in the crude cell lysate and purified by nickel affinity chromatography (Qiagen Ni-ATA resin) as previously described (Chen et al., 1994; Crabb et al., 1996). The presence of ligand was monitored by the characteristic UV absorption maximum at 425 nm. Prior to NMR analysis, sample purity was verifed by SDS-PAGE and Edman degradation, the sample quantified by amino acid analysis, and the solvent exchanged to 25 mM TrisCI, pH 7.5, 1 mM DTT-EDTA and the preparation concentrated using Amicon Centri prep concentrators (10,000 MW cut off). Because 11-c/sretinaldehyde is a light sensitive ligand, rCRALBP preparation was performed under dim red illumination to retain the ligand in the binding pocket and preserve the holoprotein conformation. Biosynthetic isotope incorporation was as described in section III. The ' ' ^ N ammonium chloride and "'SC-methyl-methionine were obtained from Cambridge Isotope Laboratories; 5-fluorotryptophan was from Sigma. Solution State NMR. " ' H - " ' 5 N gradient-enhanced sensitivityenhanced heteronuclear single quantum correlation (GESE-HSQC) experiments were carried out on a three-channel Varian Unity 600 spectrometer using a '^HI'^^CI'^^H triple-resonance probe equipped with an actively shielded Bz gradient coil (Farmer II and Venters, 1996). "•3c NMR spectra were acquired on Varian Unity 600 and General Electric GN300 instruments. '^^F NMR spectra were obtained at 564 MHz on a Varian Unity 600 spectrometer equipped with a 5mm ' ' H / " ' 9 F probe. 3-Fluorophenylalanine (-38.0 ppm) was used to calibrate the chemical shifts of fluorine resonances relative to trifluoroacetic acid at 0 ppm (Luck, 1995). All samples were adjusted to 10 % D2O (v/v) prior to NMR analysis. Spectra were acquired at 25 °C first under dark conditions then again after exposure of the protein samples to bleaching illumination. Mass Spectrometry and Other Analytical Procedures. Liquid chromatography electrospray mass spectrometry (LC-ESMS) was performed on approximately 1 jig protein samples with a Perkin Elmer Sciex API-300 triple quadrupole mass spectrometer fitted with an articulated ion spray source and set to scan over a range of 400-3000

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m/z at 7 s/scan in 0.25 Da steps using an orifice potential of 35 V. RPHPLC was performed with an Applied Blosystems Model 120 HPLC system (modified with a 75 |LII mixing chamber), a 5|a Vydac C18 column (1 X 250 mm), aqueous TFA/acetonitrile solvents and a flow rate of 50 |il/min. About 30% of the HPLC effluent was split to the mass spectrometer. Purified protein was quantified by phenylthiocarbamyl amino acid analysis (Applied Biosystems models 420H/130/920) and purity was evaluated by Edman degradation (Applied Biosystems models 470/120/900) as described elsewhere (Crabb et al., 1988). rCRALBP retinoid binding was monitored by ultraviolet spectral analysis and photoisomerization of bound retinoid achieved by exposure to bleaching illumination for 10-30 min at room temperature (Crabb etal., 1996).

III. Isotope Incorporation and Pre-NMR Analyses Recipes for defined media used to obtain uniform biosynthetic "•^N and 13c labeling of recombinant proteins expressed in bacteria have been published elsewhere (Venters et al., 1991). For uniform ""SN-labeling of rCRALBP, we utilized a modified minimal media containing 1 g/L [">5N,98%] ammonium chloride as the sole nitrogen source plus M9 salts, 2 mM MgS04, 1 jiM FeCIa, 100 |iM CaCl2, 50 |iM ZnS04, 10 lig/ml biotin, 10 |ig/ml folic acid, 0.1 |Lig/ml riboflavin, 5 |Lig/ml thiamine and 50 |ig/ml ampicillin. This media provided about a 2 fold improved yield of purified rCRALBP (5-6 mg/L) over that achieved with standard LB growth media. The level of ^^N isotope incorporation was evaluated by liquid chromatography electrospray mass spectrometry as shown in Fig. 1. For rCRALBP grown in the above media, the measured mass of the i^N labeled protein was 39,591 ±5 (compared with 39,594 for the calculated mass of the protein with 100% ''^N incorporation), indicating that essentially complete labeling was achieved. Several commercially available fluorinated analogs of aromatic amino acids are useful for "•Qp NMR studies, including 2-,3- and 4fluorophenylalanine, 4-,5- and 6-fluorotryptophan and 3-fluorotryrosine. High level incorporation of fluorinated amino acids can be accomplished by using a bacterial auxotroph strain for the amino acid of interest or by using glyphosate [N-(phosphonomethyl)-glycine] which inhibits the synthetic pathways of the aromatic amino acids (Kim et al, 1990). Because 5-fluorotryptophan is a protein synthesis inhibitor, protein production requires a balance with unfluorinated tryptophan. A 5:1 molar ratio of fluorinated to unfluorinated tryptophan yields up to 65% incorporation of the fluorinated amino acid in many bacterial auxotroph systems (Luck and Faike, 1991). Variable lower levels of fluorine incorporation are usually obtained without a bacterial auxotroph yet informative NMR signals are often still achievable by extending the NMR measurement time. We have obtained useful i^FTrp spectra from rCRALBP (which contains 2 mole Trp per mole protein) using 8-10 h NMR analysis periods. For these analyses,

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rCRALBP was produced using a minimal media (Luck and Faike, 1991) containing I\/I9 salts with 2x the normal phosphate concentration, 2% (w/v) case amino acids (Difco), 1% (v/v) glycerol, 10 |ig/ml thiamine, 100 iLig/ml ampicillin, 65 |ig/ml 5F-tryptophan (DL) and 8 |xg/ml tryptophan (L). LC-ESMS analysis of ""QP-labeled rCRALBP yielded the electrospray mass spectra shown in Fig. 1C. Multiple molecular species of the ^^F labeled protein are apparent in the deconvoluted spectra (inset) with measured masses of (a) 39,117±8, (b) 39,126±9 and (c) 39,146±8 which approximate the calculated masses for the unlabeled protein (39,110), the protein containing one equivalent of 19F-Trp (39,128) or two equivalents of ^^F-Trp (39,146). rCRALBP was labeled with "i^C-methyl methionine for NMR analysis with the same media used for "i^F labeling with the addition of 200 |ig/ml ''3C-methyl-methionine, 10 |xg/ml tryptophan and no 5fluorotryptophan. This formulation, which results in a 2:1 rato of "^^CMet to "'SC-Met, was used to keep isotope cost low while still providing useful "ISC NMR data with 10-12 h analysis times. Since ^3C-labeling has no adverse effect on protein synthesis or bacterial cell growth, higher amounts can be used for isotopic labeling to reduce NMR instrument time. The fusion rCRALBP contains 7 Met per mole protein and mass spectral analysis (Fig. IB) demonstrates significant "^^c incorporation was obtained based on a major molecular species with a measured mass of 39,121 ±5 (compared with a calculated mass of 39,117 for 100% incorporation). Compared with the uniformly labeled "•SN rCRALBP (Fig. 1A), the microheterogeneity of the ^^F and "^^C preparations from the presence of both labeled and unlabeled Trp and Met residues is readily apparent in the mass spectra (Fig. 1B,C).

IV. NMR Applications Gradient-enhanced sensitivity-enhanced heteronuclear single quantum correlation (GESE-HSQC) NMR is a rapid and sensitive highresolution, multidimensional methodology that requires advanced instrumentation and pure samples with uniform isotopic labeling in millimolar concentrations (Venters and Spicer, 1995), The experiment when applied to ""^N labeled protein correlates the ^mide proton with its directly bound ""^N nuclei. Concerted effort and significant associated costs in time and money can yield complete assignment of backbone resonances and protein three-dimensional solution structures; however, application of this methodolo'gy in a modest manner is affordable and appropriate for probing protein-ligand interactions. As an example, we present ""^N GESE-HSQC data that support localized conformational changes in rCRALBP when the ligand is removed from the retinoid binding pocket (Fig. 2). The GESE-HSQC pulsed field gradient experiment is depicted in Fig. 3. This type of NMR analysis is particularly useful for decerning whether ligand removal results in global or localized protein conformational change and, as in

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the case of rCRALBP, provides a rational foundation for pursuing identification of specific amino acid residues involved in the localized change. One dimensional "'^p NMR experiments offer advantages of low cost, sensitivity and simplicity. Fluorine provides a sensitive probe for monitoring protein conformational changes and ligand interactions in part because the chemical shift range is 100 fold larger than that of the proton due to the lone pair electrons (Gerig, 1994; Danielson and Faike, 1996; Sykes and Hull 1978). Most commonly, fluorine is substituted for hydrogen in the ring structure of aromatic amino acids such as tyrosine and tryptophan which enhances sensitivity due to the resonance electrons in the ring. Because of the usual low abundance of tryptophan in proteins and the low cost of 5-fluorotryptophan, tryptophan is generally the residue of choice for fluorine incorporation. A potential pitfall is that fluorine incorporation may cause protein denaturation and/or instability, depending on the site of incorporation and the nature of the protein. Partial apparent denaturation of 5fluorotryptophan labeled rCRALBP was observed by NMR, nevertheless, distinct "I^F resonances for each of the two labeled Trp were also observed (in NMR spectra collected over 8h from 18 mg/ml

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protein solutions before and after exposure to bleaching illumination). Apparent chemical shift differences of about 0.25 and 0.75 ppm for the two Trp were seen upon removal of ligand. The smaller shift suggests that this residue is experiencing a small conformational change whereas the larger shift suggests that the other Trp residue may be in more immediate contact with bound retinoid (data not shown). To determine which Trp residue in the CRALBP sequence is associated

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Figure 4. NMR Spectra of "^^C-Met labeled CRALBP with and without ligand. Solution state NMR spectra of CRALBP labeled with '•^c-Met (23 mg/ml) containing bound 11-c/s-retinaldehyde were recorded in the dark. The sample was then exposed to bleaching illumination and reanalyzed without bound ligand. Major "'^C chemical shift differences are apparent between the two spectra, suggesting that a Met residue may be in direct contact with bound retinoid and/or associated with the CRALBP retinoid binding pocket. NMR conditions include: pw90 (^^C) = 15 ^isec, sw (13Q) _ 13422 Hz, preacquisltion delay= 1 sec, "'H broadband decoupling during aquisition at a field strength of 1.85 KHz using MLEV-16.

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with the observed chemical shifts, site directed Trp to Phe mutants are being prepared for additional retinoid binding and ^^F NMR analyses. One dimensional "^^c NMR experiments are also relatively inexpensive and straightforward however natural abundance ""^C (0.0018 relative to ""H) does not exhibit the sensitivity in NMR of fluorine (0.8331 relative to 1H). Specific enrichment of sites, such as the methyl carbons of methionine can increase the sensitivity and ""^CMet has proven to be an effective tool for probing protein conformational changes (Beatty et al., 1996). The advantage of the innocuous substitution of carbon-13 for carbon-12 is counteracted by the smaller chemical shift dispersion of ""^C compared with ""^F. Residues of high abundance In the protein may cause overlap of resonances in the NMR. Another consideration is that while '•^cmethlonine Is less expensive than many isotopes, it remains more expensive than '•^F-Trp and "^^N ammonium chloride. NMR analysis of the ''^c-Met labeled rCRALBP (Fig. 4) was used to probe the possible interaction of any of the seven Met residues with retinoid. Preliminary one dimensional NMR results reveal a predominant set of ovelapping 13c resonances with minor chemical shift differences before and after bleaching (Fig. 4). However, a distinct and major ""^c chemical shift difference of about 1 ppm is also apparent after removal of ligand, suggesting that at least one Met residue may be in direct contact with bound retinoid. Site directed mutagenesis of the Met residues in CRALBP is underway and additional two dimensional HSQC NMR analyses will be used to assign specific Met residues to observed chemical shifts and ligand interactions.

V. Conclusions This study demonstrates the applicability of ""^N, ""^F and ^^c NMR methodology for studying ligand interactions in a light sensitive protein such as rCRALBP. Gradient enhanced sensitivity enhanced heteronuclear single quantum correlation "^^N NMR has provided evidence that rCRALBP undergoes a specific localized conformational change upon photoisomerization of 11-c/s-retinaldehyde and removal of the ligand from the binding pocket. The results from the multidimensional NMR measurements strongly support the likelihood that the ^^F Trp and i^c-Met NMR chemical shift differences observed for the protein with and without bound 11-c/s-retinaldehyde are associated with protein-ligand interactions. Site directed mutagenesis in conjunction with further NMR and ligand binding studies promises to identify components of the rCRALBP retinoid binding pocket. The effectiveness of these NMR experiments was greatly facilitated by careful quantification and protein characterization prior to NMR analysis, particularly by liquid chromatography electrospray mass spectrometry.

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References Beatty, E.J., Cox, M.C., Frenkiel, T.A., Tarn, B.M., Kubal, G., Mason, A.B., MacGillivray, R.T.A., Sadler, P.J. and Woodworth, R.C. (1996) J Amer Chem Soc. (in press). Chen, Y., Johnson, C, West, K., Goldflam, S., Bean, M.F., Huddleston, M.J., Carr, S.C., Gabriel, J.L and Crabb, J.W. (1994) In Techniques In Protein Chemistry V, J.W. Crabb, ed., pp 371-378, Academic Press, San Diego,CA. Crabb, J.W., Johnson, CM., Carr, S.A., Armes, LG. and Saari, J.C. (1988) J Biol Chem. 263, 18678-18687. Crabb, J.W., Chen, Y., Goldflam, S., West, K.A. and Kapron, J.T. (1996) In Techniques in Molecular Biology: Retinoids, Redfern, C, ed., Humana Press, NJ (in press). Danielson, M.A. and Faike, J.J. (1996) Annu. Rev. Biophys. Biomol. Struct 25: 163-195. Gerig JT (1994) Prog. Nucl. Magn. Reson. Spectrosc. 26:293370. Kim, H.W., Perez, J.A., Ferguson, S.J., Campbell, I.D. (1990) FEBS Letts. 272: 34-36. Luck, L.A. (1995) In Techniques In Protein Chemistry V I , J.W. Crabb, ed., pp 487-494, Academic Press, San Diego,CA. Luck, L.A. and FaIke, J.J. (1991) Biochemistry 30: 4248-4252. Sykes, B.D. and Hull, W.E. (1978) Meth Enzymol. 49: 270-295. Saari, J.C. and D.L. Bredberg (1987) J Biol Chem 262, 7618-7622. Saari, J.C, Bredburg, D.L. and Noy, N. (1994) Biochemistry 331: 3106-3112. Farmer II, B.T. and Venters, R.A. (1996) J BioMolecular NMR 7, 59-71. Venters, R.A., Calderone, T.L., Spicer, LD. and Fierke, CA. (1991) Biochemistry 30, 2291-4494. Venters, R.A. and Spicer, L.D. (1995) In Techniques In Protein Chemistry V I , J.W. Crabb, ed., pp 495-502, Academic Press, San Diego, CA

A Novel Method for Measuring the Binding Properties of the Site-Directed Mutants of the Proteins That Bind Hydrophobic Ligands: Application to Cellular Retinoic Acid Binding Proteins Honggao Yan, Lincong Wang and Yue Li

Department of Biochemistry Michigan State University East Lansing, Michigan

I. Introduction Many hydrophobic molecules such as vitamin A, vitamin D and steroid hormones play vital roles in a variety of cellular processes. Because of the low solubility of these molecules in water, it has been difficult to measure the binding properties of the site-directed mutants of the proteins that interact with these hydrophobic ligands such as cellular retinoic acid binding proteins (CRABPs) (Zhang et al. 1992; Chen et al. 1995). This has greatly hampered the studies of the quantitative structure-function relationships of these important proteins. Retinoic acid (RA), a hormonally active metabolite of vitamin A, has profound effects on cell growth, differentiation, and morphogenesis. Two types of proteins have been found to bind RA: nuclear retinoic acid receptors (RARs and RXRs) and CRABPs. RARs and RXRs are RA-activated transcriptional factors that regulate expression of target genes (Mangelsdorf et al., 1994). Although the physiological roles of CRABPs are not clear at present, they are thought to be involved in cellular transport and metabolism of RA (Ong et al., 1994). Two isoforms (CRABP-I and CRABP-H) have been characterized. Both CRABP-I and CRABP-H bind specifically to SiW-trans-TQimoic acid, but they differ in affinity for RA, expression pattern and regulation. It appears that the two isoforms may have distinct functions. The idea is supported by the fact that the sequence identity of human and mouse CRABP-I (99.3%) or human and TECHNIQUES IN PROTEIN CHEMISTRY VIII Copyright © 1997 by Academic Press All rights of reproduction in any form reserved.

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mouse CRABP-n (93.5%) is much higher than the sequence identity (13.1%) between the two isoforms from the same source. Four conserved residues (Arg-Ul, Leu-121, Arg-132 and Tyr-134 in CRABP-n) line at the bottom of the RA binding pockets of CRABPs and interact with the carboxyl group of RA (Kleywegt et al., 1994). Site-directed mutagenesis studies have shown that the two arginine residues are important for binding of RA (Zhang et al., 1992; Chen et al., 1995). However, the affinities of these mutants for RA have not been quantitatively determined because the current RA binding assays are inapplicable to mutants with greatly decreased affinity for RA. We have developed a novel competitive binding assay for measuring the dissociation constants of the site-directed mutants of CRABPs. We have used this novel method to evaluate the contribution of Leu-121 of CRABP-H to binding of RA in conjunction with site-directed mutagenesis and NMR. The results show that Leu-121 is also important for binding of RA and contributes to the binding energy by ~ 1.4 kcal/mol.

II. Experimental Procedures A. Site-Directed Mutagenesis The oligonucleotide for making L121A mutant was 5'-GGAACTGATCGCGACCATGACG-3'. The mutant was generated by the method of Kunkel (1985) and screened by DNA sequencing. In order to ensure that there were no unintended mutations in the mutant, the entire sequence of the mutated gene was determined. Both the wild-type and mutant proteins were purified by ion exchange chromatography using DEAE-cellulose DE53 followed by gel filtration using Sephadex G-50 (Wang et al., 1996).

B. Competitive Binding Assay The assays were designed to measure the affinity of a mutant for RA relative to that of the wild-type CRABP-H. The proteins were dissolved in a phosphate buffer (4 mM NaH2P04, 16 mM Na2HP04, 150 mM NaCl, pH 7.3). The concentrations of the protein stock solutions were measured by OD280 using the absorption coefficient 19,480 M-i cm-^ for CRABP-H. RA stock solutions were prepared in absolute ethanol. The concentrations of the RA stock solutions were determined by OD336 using the absorption coefficient of 45,000 M-i cm-i. The assays were carried out in equilibrium dialysis cells at room temperature. The two compartments of each dialysis cell were separated by a semipermeable membrane with a molecular weight cutoff of 6-8 kDa. One compartment was filled with the wild-type CRABP-H (1 ml), and the other with LI21 A. An equal amount of [^H] RA (100 nM) was added to each compartment. The proteins in both compartments were in large excess of RA

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(>20-fold). 100 |il of samples were taken from the two compartments after various times of incubation at room temperature, mixed with 5 ml of scintillation fluids and counted by a liquid scintillation counter. The equilibria in the two compartments that contain the wild-type CRABP-II and mutant proteins can be described by Eq. (1) and Eq. (2): J.

_ [WTIRA]

...

^

_ [MTIRA]

...

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J^d(MT) - [MT*RA]

^^ ^^

where WT, MT, WT«RA and MT#RA represent the wild-type CRABP-H, a CRABP-II mutant and their RA complexes, respectively. Therefore ^d{MT) _ [MTIWT^RA] WT\MT*RA] ^d{WT)

^3^

Since the concentrations of the proteins were much greater than their respective dissociation constants and the concentration of RA, [WT]»[\yr]^^^^p [MT] - [MT\,,i, [WT• RA\»[RA], and [MT• RA]»[RA]. Then the relative dissociation constant can be calculated by Eq. (4): ^dmT) '^diWT)

_

\.^T\otal^Wr L t^^ ]total ^MT

(4)

where C ^ and C^j are the measured radioactivities of the two compartments containing the wild-type CRABF-E and the mutant, respectively. It turned out that the system could not reach equilibrium in 2 days, presumably because of few free RA in solution to diffuse across the membrane. Since RA is not stable even in dark, the assay was redesigned to match the equilibrium conditions by varying the ratio of the protein concentrations of the wild-type and the mutant ([MT\^^^J[WT\^j^i). Thus the concentration of the mutant was varied while keeping the concentration of the wild-type at ~2 |iM. Initially the concentration of the mutant was increased in an exponential manner (e.g., 2, 20, 200 |iM). Then it was varied in a small range. Since an equal amount of RA was added to the two compartments of the dialysis cell, the two compartments should have the same RA concentration and radioactivity at the beginning of each assay. If [MT]j^^^i/[WT]j^t^i ^ i5:^(Mr)/^^(wr)' there would have a net transfer of RA across the semipermeable membrane separating the two compartments. Thus the radioactivity counts of the two compartments {C^ and C^^.) would differ after incubation for a certain period. When [MTl^^^J[WTl^^^i < K^^j^^/K^^^^, then C^-C^r >0. When [MTl^^J[WTl^^^, > K,,^r,/K,,^,, 0. When [MT],,J[WTl^,, = K,,^r)/K,,^,, then C^-C^= 0.

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C. NMR Spectroscopy NOESY was performed at 32 °C on a VXR-500 spectrometer operating at a proton frequency of 500 MHz. The protein was dissolved in 20 mM sodium phosphate, pH 7.5 (direct pH meter reading), 100 mM NaCl, 5 mM DTT in D2O. The protein concentration was ~2 mM. The data was acquired in the hypercomplex mode with a mixing time of 150 ms (Jeener et al., 1979; Macura & Ernst, 1980). The spectral width was 7200 Hz in both dimensions. 2048 complex points in the t2 dimension and 256 complex points in the tl dimension were acquired. 96 transients were collected for each FID. Data processing was performed on a Sun Sparc 10 station using VNMR software from Varian. The time domain data were zero-filled once and multiplied by shifted sinebell or Gaussian functions before Fourier transformation in both dimensions. Chemical shifts were referenced to internal sodium 3-(trimethylsilyl)-propionate-2,2,3,3d4.

III. Results and Discussions A. Competitive Binding Assay Two types of methods have been in general use for measuring binding of RA to CRABPs: fluorometry and radiometry. The radiometric method involves separation of bound from free RA by dextran-coated charcoal, gel filtration and other means. Substantial loss of bound ligand during the separation process makes the method unsuitable for measuring the dissociation constants of sitedirected mutants with greatly decreased affinity for RA. The very limited solubility of RA in water (-200 nM, Szuts & Harosi, 1991) also makes the fluorometric method inapplicable for determining the dissociation constants of these mutants. Studies of the quantitative structure-function relationships of CRABPs have been hampered by the lack of methods for measuring the affinities of site-directed mutants for RA (Zhang et al. 1992; Chen et al. 1995). We have developed a novel competitive binding assay for measuring the affinities of site-directed mutants for RA relative to that of the wild-type CRABP. The essence of the method is to monitor the competition between a mutant and the wild-type protein for binding of limited RA. Equilibrium dialysis cells are used for the assays. The two compartments of each dialysis cell are filled with the wild-type and mutant proteins respectively. The absolute concentration of RA is not important as long as the concentration of free RA is much smaller than that of bound RA. There is no need to separate bound from free RA. The transfer of RA from one compartment to the other is determined by measuring the radioactivities of the samples taken from the two compartments. The direction of the net transfer is dependent on the relative affinity of the proteins and the ratio of the protein concentrations of the two compartments. A representative result is shown in Figure 1. When the ratio of

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the concentrations of the two proteins ([L121A]/[WT]) is < 8, there is a net transfer of RA from the compartment containing L121A to the compartment containing the WT. When the ratio of the concentrations of the two proteins is > 12, there is a net transfer of RA from the compartment containing the WT to the compartment containing LI21 A. Since the relative K^ lies between the points with opposite net transfers, the K^ of L121A relative to that of the (^d(L121A/^d(WT)) ^^ 8-12.

Determination of the relative dissociation constant of a point mutant is sufficient for estimating the energetic contribution of the amino acid residue to ligand binding (AAG = RT\n{K^(^^j.^ I A:^(HT) ))• The method can also be used for measuring the relative dissociation constants of the mutants of other proteins that bind hydrophobic ligands such as RA receptors, vitamin D receptors and steroid hormone receptors.

i

-400

-600 H

[L121A]/[WT]

Figure 1. Competitive binding assays for measuring the dissociation constant of L121A mutant relative to that of the wild-type CRABP-II. The relative radioactivity is the radioactivity count of the compartment containing the wild-type protein minus that of the compartment containing L121A.

Honggao Yan et al

454

B. Conformational Characterization By NMR Since a decrease in the affinity of a mutant for RA may be caused by conformational changes, we compared the conformation of L121A with that of the wild-type protein by NMR. Parts of the NOESY spectrum of L121A are shown in Figure 2. We have recently made total sequential resonance assignment of the wild-type CRABP-E (Wang et al., in preparation). 18 interresidue NOEs between the aromatic protons in the wild-type protein have been identified and assigned. Among the 18 NOE cross peaks, 16 of them can be identified in the NOESY spectrum of LI21 A. The other two NOEs are rather weak in L121A. We have not assigned the NOEs between aromatic and aliphatic protons. Qualitatively, the aromatic-aliphatic NOE patterns of the wild-type and L121A are very similar. The results suggest that L121A mutant is properly folded and its conformation is highly similar to that of the wild-type protein. Thus the decrease in the affinity of L121A for RA is unlikely to be caused by conformational perturbations.

C. LeU'121 Is Important for Binding ofRA The results of the competitive binding assay and NMR characterization of L121A mutant suggest that Leu-121 is important for binding of RA. Leu-121 is located at the bottom of the RA binding pocket of CRABP-H. One of the methyl group of Leu-121 is in close contact with the carboxyl group of the bound RA (Kleywegt et al., 1994). The distance between the carbon of the methyl group and the oxygen of the carboxyl group is 3.26 A. The packing of the methyl group and the carboxyl group is very close to the optimal van der Waals interaction (Derewenda et al., 1995). On the basis of the relative dissociation constant, the van der Waals interaction between the methyl group of Leu-121 and the carboxyl group of RA contributes to the binding energy by ~1.4kcal/mol.

IV.

Conclusions

A novel competitive binding assay has been developed for measuring the relative dissociation constants of the site-directed mutants of CRABPs. Leu121 has been replaced with alanine by site-directed mutagenesis. The affinity of the mutant for RA is decreased by ~ 10-fold as measured by the competitive binding assay. NMR characterization indicates that the conformation of L121A mutant is very similar to that of the wild-type protein. The results taken together show that Leu-121 is important for binding retinoic acid and contribute to the binding energy by -1.4 kcal/mol.

Cellular Retinoic Acid Binding Proteins Fl

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Figure 2. Parts of the 500 MHz NOESY spectrum of L121A at 32 °C. The mixing time was 150 ms. Only the interresidue NOEs are labeled. The identities of the NOEs are: A, F65-2,6H ••• F71-2,6H; B, F65-2,6H ••• F71-3,5H; C, F65-4H • • W109-6H; D, F65-3,5 H ••• W109-6H; E, F50-2,6H ••• W87-7H; F, F71-4H • • W109-7H; G, F65-4H • • W109-7H; J, W87-5H •• F3-4H; K, F50-3,5H - F3-2,6H; L, F50-3,5H - W87-7H; M, F50-4H •. W87-7H; N, F50-4H ." F32,6H; O, F50-2,6H •• W87-6H; P, F50-2,6H •• F3-2,6H; Q, W87-5H ••• F3-2,6H; R, F3-4H •• W87-4H.

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Acknowledgments We are indebted to Dr. Anders Astrom for providing us the wild-type cDNA clone of human CRABP-II. This work was supported by funds from the REF Center of Protein Structure and Design and the Cancer Center at Michigan State University.

References Chen, L. X., Zhang, Z.-P., Scafonas, A., Cavalli, R. C , Gabriel, J. L., Soprano, K. J., & Soprano, D. R. (1995) J, Biol Chem. 270,4518-4525. Cogan, U., Kopelman, M., Mokady, S., & Shinitzky, M. (1976) Eur. J. Biochem. 65, 71-78. Derewenda, Z. S., Lee, L., & Derewenda, U. (1995) /. Mol. Biol 252, 248-262. Jenneer, J and Ernst, R. P. (1979) J. Chem. Phys. 71,4546-4553. Kleywegt, G. J., Bergfors, T., Senn, H., Le Motte, P., Gsell, B., Shudo, K., & Jones, T.'A. (1994) Structure 2, 1241-1258. Kunkel, T. A. (1985) Proc. Natl. Acad. Sci. U. S. A. 82,488-492. Macura, S and Ernst, R.P. (1980) Mol. Phys. 41, 95-117. Mangelsdorf, D. J., Umesono, K., & Evans, R. M. (1994) In The Retinoids: Biology, Chemistry, and Medicine (Spom, M. B., Rorberts, A. B., & Goodman, D. S., Eds.) pp 319-349, Raven, New York. Ong, D. E., Newcomer, M. E., & Chytil, F. (1994) In The Retinoids: Biology, Chemistry, and Medicine (Sporn, M. B., Rorberts, A. B., & Goodman, D. S., Eds.) pp 283-317, Raven, New York. Szuts, E. Z., & Harosi, F. I. (1991) Arch. Biochem. Biophys. 287, 297-304. Wang, L., Li, Y., & Yan, H. (1996) submitted to J. Biol. Chem.. Zhang, J., Liu, Z.-P., Jones, T. A., Gierasch, L. M., & Sambrook, J. F. (1992) Proteins: Struct, Funct., Genet. 13, 87-89.

A Strategy for Predicting the Ligand Binding Competence of Recombinant Orphan Nuclear Receptors using Biophysical Characterization Derril Willard^ Bruce Wisely^, Derek Parks^ Martin Rink^ William Holmes', Michael Milbum^, and Thomas Consler' Departments of'Molecular Sciences, ^Structural Chemistry, and ^Molecular Biochemistry. Glaxo Wellcome Research and Development, Research Triangle Park, NC, 27709 I. Introduction Nuclear receptors are a loosely related group of ligand dependent transcriptional regulators with varying degrees of sequence homology. These proteins have been historically associated with the steroid hormone receptors, e.g. estrogen and glucocorticoid receptors, by virtue of DNA binding domain sequence homology comprising two zinc finger motifs. Many of these are orphan receptors, having no defined ligand. The nuclear receptors present tempting targets in the pursuit of a systems based research approach since so many have now been cloned. However, when recombinant forms of a receptor are available before its cognate ligand has been identified, a problem arises. How does one determine if an orphan receptor is active for use in in vitro assays? Researchers now have access to unparalleled amounts of DNA sequence and genetic data. Families of homologous gene products can be studied with the intent of connecting specific proteins to various disease conditions. An obvious advantage to this wide scope of research is that once the mechanism of action has been elucidated for a few family members, this information can then be applied in a general sense to other homologues. Specifically, to apply this type of strategy to our studies, we approached the problem with two premises. First, we engineered recombinant constructs of orphan nuclear receptors to contain domains with hypothetical fiinctional homology to receptors with known activities and ligands. In particular, a great deal is known concerning retinoid X receptor a (RXRa) (1) and the domains necessary for DNA binding, retinoid binding, and selfi^hetero-association. For the purposes of this study, PPARa, PPAR5, PPARy, RXRa, and LXRa constructs were created to contain the putative ligand binding domains (LBD). The amino acid residues within this conserved contiguous region have been TECHNIQUES IN PROTEIN CHEMISTRY VIII Copyright © 1997 by Academic Press All rights of reproduction in any form reserved.

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shown to be both necessary and sufficient to demonstrate ligand binding competence for RXRa and other nuclear receptors. Structurally, the LBDs are composed primarily of multiple a-helicies (2,3). Second, we began to characterize each nuclear receptor using a variety of biophysical techniques. The object of this scrutiny was to compile a set of physical traits for each protein and to use these characteristics as a basis to compare the orphans with those receptors having defined ligand binding ability. Purified proteins were characterized by low resolution solution structure, thermal stability and propensity to self-associate or to aggregate. Observations of protein expression levels and solubility were also considered as qualitative estimates of native structure. Circular dichroism spectroscopy (CD) was employed to probe secondary structural features. CD monitored unfolding and differential scanning calorimetry (DSC) provided two separate measures of thermal stability. Static and dynamic light scattering (SLS and DLS, respectively) and analytical ultracentrifiigation were used to determine solution values for molecular size, association and aggregation state.

IL Materials and Methods A. Expression and purification Nuclear receptor LBDs were engineered to have an amino-terminal polyhistidinefiasiontag. Constructs were expressed in E. coli strain BL21 (DE3) using the T7 promoter. PPARa LBD was also expressed as a polyhistidine tagged recombinant in baculovirus infected T. ni. Protein purification in most cases was performed using a single nickel affinity step. Tagged protein was either initially purified by anion exchange chromatography then adsorbed onto a Pharmacia Chelating Sepharose Fast Flow column charged with nickel or adsorbed by nickel-chelating chromatography directly out of crude cell lysate. Proteins were eluted by 0-1 M linear gradient of imidazole in the lysis buffer.

B. Structure and stability Purified nuclear receptor proteins were buffer exchanged into PBS for CD spectral analysis using an Aviv model 62DS CD spectropolarimeter. The proteins were scanned repetitively in 0.1 cm quartz cuvettesfi^om197 to 300 nm in 1 nm wavelength increments. EUipticity was converted to molar ellipticity for comparisons. Thermal transitions were performed with the CD instrument above. Proteins were monitored at 222 nm over a temperature range of 5-80°C. Data were collected in 1°C increments with a slope of 10°C/min. Initially, data were fit to a simple sigmoidal mathematical relationship for comparison. The half-point of the thermal transition, T1/2, was determined by iterative fitting using the Boltzmann equation. Data were also fit to the following thermodynamic model:

Ligand Binding of Recombinant Orphan Nuclear Receptors

''

459

1 + exp"

where u = [{l/T-\/

T^'^AcJ^ - A//,,) + Ac^ ln(r/ T^)]/R

and where 0T is ellipticity at T (temperature in °K), 0N is the native protein ellipticity, 0D is the unfolded protein ellipticity, R is the gas constant, TD is the temperature at which the protein unfolding transition is half-complete, AHTD is the enthalpy change at TD, and Acp is the heat capacity change. DSC analyses were performed using a MicroCal MCS DSC unit. Data were analyzed to determine the midpoint of a two-state thermal transition (Tm) using the accompanying MicroCal Origin data analysis package. For each experiment approximately 1.5 mL of protein at concentrations ranging from 3-20 micromolar was analyzed during an increase in chamber temperature from 5-80 °C. The data were collected at a scan rate of 90 °C/hr and a filter period of 5 seconds. Scans were performed using PBS as the reference buffer.

C Association/Solution State Dynamic light scattering (DLS) measurements were performed on a DynaPro-801 instrument (Protein Solutions) and the data analyzed with the Autopro software package. All proteins were filtered through a 0.10 micron syringe filter and analyzed at 22°C. The translational difiiisional coefficient (DT) was obtained directly and the hydrodynamic radius (RH) is derived by a rearrangement of the following equation: DT

kT 67cr|RH

where k is the Boltzmann's constant, T is the absolute temperature and TJ is the solvent viscosity. Static light scattering (SLS) measurements were performed at 22°C on a Wyatt Technology Dawn DSP laser photometer (argon-ion laser 488 nm). Data were analyzed with the Astra software package. All proteins were filtered through a 0.10 micron syringe fiher and held at 30 psi during analysis. Each sample was run in duplicate and the refractive index increment was estimated to equal 0.180. Molecular weights are derived from a rearrangement of the following equation: K*c RiO)

1 + 2A2C M^P{e)

460

Derril Willard et al

where R(0) is the excess intensity of scattered light, c is sample concentration, A2 is the second viral coefficient, P(9) is the scattering function which depends on the molecular configuration and approaches 1 for proteins, and K* is an optical parameter. Sedimentation equilibrium analytical ultracentrifiigation was performed using a Beckman XL-A (Palo Alto, CA) centrifuge with two-channel or six-channel 12 nmi charcoal-filled epon centerpieces. Runs were performed at 20, 25, and 30 krpm at 4°C. Equilibrium was judged to be achieved by the absence of change between plots of several successive scans after approximately 20 hours. Solvent density was determined empirically at 4°C and 20°C using a Mettler DA-110 density/specific gravity meter calibrated against water. The partial specific volume of each protein was calculated using the method of Cohn and Edsall (4). Temperature differentials were incorporated using the appropriate equation (5) modified from values of each amino acid at 25°C (6). Raw data was analyzed by the Beckman/Microcal Origin non-linear regression software package using multiple iterations of the Marquardt-Levenberg algorithm (7) for parameter estimation.

D. Ligand Binding Binding constants were determined for PPARy and RXRa using ligands with known affinities. Each protein was incubated with the appropriate tritiated radioligand for two hours. Bound ligand was separated from free by gel filtration. The unbound radioligand was mixed with scintillation fluid and counted. For PPARa, PPAR5, and LXRa, similar assays were performed using compounds which had been implicated as nuclear receptor effectors in a separate cell-based assay.

III. Results Nuclear receptor LBDs were constructedfromthe homologous regions of the native sequences (Figure 1). Expression and purification of the nuclear receptors used in this study proceeded as detailed. N-terminal histidine fusion tags provided an easy and similar method of purification for each protein. In general expression yields were good (over 20 mg protein/fermentation liter) with the exception of LXRa which produced >5 mg protein/fermentation liter. Post nickel-chelating chromatography, the proteins were dialyzed into PBS for use in these studies, akhough long-term stability was found to vary from protein to protein over a range of buffer and storage conditions. In particular the histidine tag of PPARa produced in baculovirus infected T. ni cells was found to be processed away in all purification attempts shortly after elution from the nickelchelating chromatography step. Protein solubility as evidenced by ability to concentrate each nuclear receptor was found to be sufficient for the studies involved. The ultimate concentration attainable with LXRa was however

Ligand Binding of Recombinant Orphan Nuclear Receptors PPARa

461

. .6MSHNAIRFORMPRSEKAKLKAEILTCEHDIEDSETADLKSLAKRIYEAYLKNFN

PPARS

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PPARy LXRa RXRa

. .OMSHNAIRF6RNPQAEKEKLLAEISSDIDQLNPESADLRQALAKHLYDSYIKSFP ..QAHATSLPPRASS . .6NKREAVQEERQ_R6KDR ^NENEVESTSSANEDMPVERILEAELAVEP

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PPARS

MTKKKARSILT6KASHTAPFVIHDIETLWQAEKOLVWKQLVN6LPPYKEISVHVFYR

PPARY LXRa RXRa

LTKAKARAILT6KTTDKSPFVIYDMNSLNN6EDKIKFKHITPLQEQSKEVAIRIFQ6 PPQILPQLSPEQL6MIEKLVAAQQQCNRRSFSDRLRVTPWPMAPDPHSREARQQRFA KTETYVEANNOLNPS SPNDP VTN I

PPARa PPAR5 PPARY LXRa RXRa

CQCTSVETVTELTEFAKAIP6FANLDLNDQVTLLKYGVYEAIFAMLS SVNNKDGM CQCTTVETVRELTEFAKSIPSFSSLFLNI)QVTLLKYOVHEAIFAMLA__SIVNKDGL CQFRSVEAVQEITEYAKSIPGFVNLDLNDQVTLLKYGVHEIIYTMLA^SLMNKDGV HFTELAIVSVQIVDFAKQLP6FLQLSREDQIALLKTSAIEVMLLETS RRYNPGSE CQAADKQLFT_LVEWAKRIPHFSELPLDDQVILLRAGWNELLIASFSHRSIAVKDGI

PPARa PPAR5 PPARY LXRa RXRa

LVAYGNGFITREFLKSLRKPFCDINEPKFDFAMKFNALELDDSDISLFVAAIICCGD LVANGSGFVTREFLRSLRKPFSDI lEPKFEFAVKFNALELDDSDLALFI AAIILCGD LISEGQGFMTREFLKSLRKPFGDFMEPKFEFAVKFNALELDDSDLAIFIAVIILSGD SITFLKDFSYNREDFAKAGLQVEFINPIFEFSRANNELQLNDAEFALLIAISIFSAD LLATGLHVHRNSAHSAGVGAIFDRVLT ELVSKMRDMQMDKTELGCLRAIVLFNPD

PPARa PPAR5 PPARY LXRa RXRa

RPGLLNVGHIEKMQEGIVHVLRLHLQSWHPDDIFLFPKLLQKMADLRQL VTEHA RPGLMWVPRVEAIQDTILRALEFHLQANHPDAQYLFPKLLQKMADLRQL VTEHA RPGLLNVKPIEDIQDNLLQALELQLKLNHPESSQLFAKLLQKMTDLRQI ^VTEHV RPWVQDQLQVERLQHTYVEALHAYVSIHHPHDRLMFPRMLMKLVSLRTL SSVHS SKGLSNPAEVEALREKVYASLEAYCKHKYPEQPGRFAKLLLRLPALRSIGLKCLEHL

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QLVQIIKKTESDAALHPLLQEIYRDNY QMMQRIKKTETETSLHPLLQEIYKDMY QLLQVIKKTETDNSLHPLLQEIYKDLY EQVFALRLQDK KLPPLLSEIWDVHE FFFKLIGDTPIDTFLMEMLEAPHQMT

Figure 1. Primary sequence aligrunent of PPARa, PPARy, PPAR5, LXRa, and RXRa LBDs.

considerably lower than for the other constructs, resulting in precipitation of L X R a in the range of 0.5 mg/ml. P P A R a , PPARy, P P A R S , and R X R a all exhibited classic a-helical structure by C D spectroscopy (Figure 2). None of the four proteins had significant ellipticities in the aromatic region and all began downslopes near 240 nm. Minima at 222 n m and 208 n m were present in each scan with a crossover point at or near 200 nm. However, the spectrum for L X R a did not exhibit characteristic a-helical traits. The spectrum did not agree well with any major structural class and showed a considerable amount of scattering in the far U V . Thermal stability of the nuclear receptor L B D s was determined by both C D (Figure 3) and D S C melts. All of the proteins except L X R a showed secondary

Derril Willard et al

462

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Figure 2. CD spectra of A, PPARa; B, PPARy; C, PPAR8; D, RXRa; and E, LXRa LBDs. [0] denotes molar ellipticity in deg cm^/dmol, converted from raw ellipticity units. Data are averaged from multiple scans in each case and blank subtracted.

463

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'

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464

DerrilWillarderfl/.

Structural transitions over the temperature range 40-55°C. TD and Tm values were derived assuming a two-state model although in the case of RXRa there is some indication that a complex model should be applied. Melts were not reversible and in some cases resulted in precipitation of a portion of the sample. Ti/2 and Tm values derived by both techniques showed good agreement for each protein. CD unfolding data for LXRa appeared by visual inspection to exhibit an unfolding transition and T1/2 similar to those seen from the other nuclear receptors. However, attempts to fit the LXRa scans with the same program used to fit the other data failed to identify a TD in the above range. By DSC analysis, LXRa did not exhibit a transition as judged by the lack of any identifiable peak corresponding to AH over the experimental temperature range oflO-80°C. Association and aggregation states of the proteins were examined by SLS, DLS and sedimentation equilibrium analytical ultracentrifugation (Table 1). With the exception of SLS data for PPAR5, light scattering data observed some degree of aggregation in all cases (DLS data for PPARa was not determined). Analytical ultracentrifugation indicated that PPARa, PPARy and PPAR5 existed predominantly as monomers with molecular weights in good agreement with those calculated from the respective sequences. RXRa was shown to exist as a monomer-tetramer self-associating system in the absence of ligand. Analysis of data from LXRa runs indicated that the protein existed as a monomer but the molecular weight derived, 46.3 kD, did not agree with the calculated and apparent monomer molecular weight of 3L5 kD. Table L

S u m m a r y o f data f r o m biophysical techniques. CD Spectrum

TI«/TD(CD)'

Tm(DSC)'

MW*

SLS"

DLS"

PPARa

alpha helical

53.4/53.5

49

31.5

agg**

n.d.

31.7

Yes

PPARy

alpha helical

46.4/46.5

41.5

34.8

agg

agg

35.5

Yes

PPAR5

alpha helical

51/51

51

35.9

34

agg

36.8

Yes

RXRa

alpha helical

54/54

55

27.7

agg

agg

monomertetramer

Yes

LXRa

mixed beta/ undefined

49/*

no transition observed

31.5

agg

agg

46.3

No

Centrifugation" Binding'^!

" Values in units of °C, * Values in units of kilodaltons, *" gel filtration assay, ''aggregation, * no value obtained.

PPARy and RXRa LBDs exhibited the expected binding to their respective ligands. Novel ligands that had been implicated as PPARa and LXRa effectors in a cell-based assay were found to bind to PPARa and PPAR5 respectively in the gel filtration assay. No ligands were found to bind to LXRa in the gel filtration assay.

Ligand Binding of Recombinant Orphan Nuclear Receptors

465

IV. Discussion In our evolving strategy, we are attempting to apply low resolution structural, thermodynamic and solution state analyses in a systematic approach to develop a set of parameters indicative of native proteins for target families. Orphan nuclear receptor constructs were engineered to be homologues of constructs of verified functional nuclear receptors. Obvious initial check points in the strategy would be the lack of protein expression or the expression of only insoluble material. Four of the five constructs reported in this study provided amply expressed, soluble proteins. The low levels of both expression and solubility of LXRa provided initial indications that the construct might not exist in a native form. Qualitatively, CD spectral analysis indicated that LXRa was different from the other constructs, all of which exhibited classical a-helical structure. The LXRa spectrum clearly did not contain double minima at 208 and 222 nm but instead had a more narrow trough with a minimum around 220 nm. The spectrum is very similar to that taken from a PPARa sample which we progressed to 60°C and then rescanned (data not shown). This unfolded PPARa did not exhibit ligand binding in the gel fihration assay. Also, LXRa showed heavy scattering below 220 nm. Taken altogether, the CD data suggest that recombinant LXRa as expressed does not share a similar structure with the other four constructs. CD and DSC thermal stability studies are convenient techniques for comparing the thermodynamic characteristics of a group of proteins. While recognizing that homologous primary structure does not imply that the constructs would display similar thermodynamic properties, obvious outliers of Ti/2, TD, Tm, and AH calculations might be diagnostic for non-native proteins. Additionally, if significant misfolded populations were present, multiple transitions or less cooperative unfolding might be observed. All five constructs exhibited a range of 7°C for T1/2 as determined by CD. The range as determined by DSC for PPARa, PPARy, PPAR5, and RXRa was 13.5°C. LXRa did not produce a characteristic DSC melting profile over temperature change from 1080°C. This difference between the two techniques for LXRa is difficult to interpret. The CD transition is a measure of the difference of ellipticity at 222 nm while DSC is a measure of enthalpy of the system. The mean difference between CD Ti/2 and DSC Tm for each of the other constructs was less than 3°C. The solution behavior of proteins can ofl;en be good indicators of protein state. In this study both DLS and SLS were applied to probe for aggregation and solution molecular weight. Both techniques indicated that even constructs with verified ligand binding activity contained some proportion of aggregates. Light scattering in general is very sensitive to aggregates, even at levels representing only a small fraction of the protein populations. Analytical ultracentrifugation data indicated that RXRa existed as a monomer-tetramer (monomer-dimer with compound, data not shown) equilibrium. The other four nuclear receptors existed as monomers. Ultracentrifugation findings for the

Derril Willard et al

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monomeric constructs are consistent with small amounts of aggregate (as suggested by light scattering data) being spun down during the runs. These populations would be invisible to later data analysis. The good ideal species fit for LXRa which yielded an aberrant molecular weight is difficult to explain. If this construct is improperly folded, assumptions (e.g. regular hydration shell) inherent in the molecular weight calculation may have been inappropriate. This quality of LXRa may also be indicative of non-native structure. The primary research impetus behind this study is the desire to establish ligands and/or mechanisms of action for each nuclear receptor. Using a simple gel filtration assay which has been validated for characterizing RXRa and PPARy-binding ligands, we were able to test novel ligands which had been indicated as orphan effectors by a cell-based assay system (data not shown). Compounds that were identified as effectors of PPARa and LXRa were shown to bind PPARa and PPAR5, respectively, but not LXRa. Additionally, unfolded PPARa resembled LXRa in CD spectrum (see above) and did not retain ligand binding ability. No small molecules were found to bind the recombinant LXRa LBD. We have presented data which suggests that we can determine the usefiilness of a recombinant protein before we have an appropriate ligand in hand. These results are an encouraging start in our attempt to predict the binding competency of recombinant orphan nuclear receptors. Obviously, a considerable amount of work needs to be completed before a great deal of confidence can be placed in this type of predictive strategy. At present more than 20 human nuclear receptors have been identified and the number is certain to grow. Biophysical characterization of other nuclear receptors should aid in the construction and refinement of a database for use in further predictive studies.

Bibliography 1. Chen, Z.-P., Shemshedini, L., Durand, B., Noy, N., Chambon, P., and Gronemeyer, H. (1994) J. Biol Chem., 269 (41): 25770-25776. 2. Wurtz, J.-M., Bourguet, W., Renaud, J.-M., Vivat, V., Chambon, P., Moras, D., and Gronemeyer, H. (1996) Nature Struct. Biol, 3(1): 87-94. 3. Parker, M.G. and White, R. (1996) Nature Struct. Biol, 3(2): 113-115. 4. Cohn, E.J., and Edsall, J.T. (1943) Proteins, amino acids and peptides as ions and dipolar ions, Rheinhold, New York, p. 157. 5. Laue, T.M., Shah, B.D., Ridgeway, T.M., and Pelletier, S.L. {1991) Analytical ultracentrifugation in biochemistry and polymer science (Harding, S.E., Rowe, A.J., and Horton, J.C., eds). The Royal Society of Chemistry, Cambridge, p. 102. 6. Durschlag, H. (1986) Thermodynamic data for biochemistry and biotechnology (Hinz, H.J., ed). Springer-Verlag, New York, p. 45. 7. Marquardt, D.W. (1963)/. Soc. Ind Appl Math., 11:431-441.

SECTION VI Protein-Protein Interactions

This Page Intentionally Left Blank

Detection of /w/ra-Cellular Protein-Protein Interactions: Penicillin Interactive Proteins and Morphogene Proteins S. Bhardwaj R.A. Day Department of Chemistry University of Cincinnati Cincinnati, Ohio 45221-0172 I. INTRODUCTION Protein-protein interactions are recognized as central to understanding regulation of metabolic processes. There are, at present, no general methods for detection and measurement of protein-protein interactions in the intact cell. Current emphasis on this subject is the measurement of these interactions in vitro outside the cell (1). The use of recombinant DNA technology has been successful in many studies to demonstrate specific interactions; the "two-hybrid" system relies on transcriptional activity and relies on the modular nature of many particular sites (2). There are other "library based" methods which generally require varying degrees of genetic manipulation (1,2). We describe here a method that exploits the occurrence of salt-bridges formed by interacting proteins in their specific associations. The method applies to intra-cellular interactions in the intact cell including membrane proteins which normally can only be solubilized at the expense of their normal interactions (3). Salt bridges are ion pairs within hydrogen bonding distances and can be converted to covalent links by the gaseous reagent cyanogen (C2N2, N=C-C=N, ethanedinitrile). C2N2 has been demonstrated to be a salt-bridge specific reagent and in a carbodiimide-like mode to generate covalent bonds (4,5,6). This is depicted in Reaction I:

R^

/;^ NH2'^ III

O II

^

OC-R" + C2N2

^ \

•

R'"^

II

II

.N-C-R"+NH2C.C=N R'^

Reaction I The somewhat analogous carbodiimides will drive this mrm-molecular process and //z^^r-molecular condensations such as is exploited in peptide bond formation. C2N2 will only do the former (4,5), i.e., convert pre-existing saltbridged functional groups into a covalent link. An in vivo method must cause minimal perturbation of the cell at the critical point in the process from which TECHNIQUES IN PROTEIN CHEMISTRY VIII Copyright © 1997 by Academic Press All rights of reproduction in any form reserved.

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minimal perturbation of the cell at the critical point in the process from which the data will arise. The methodology must also provide identification of the proteins in question. The method described here satisfies the above criteria. Fluorescence/spectroscopic (F/S) labeled, active site directed substrate analogues mark the proteins in question. Cyanogen rapidly converts salt-bridges into covalent links (6). It is our working hypothesis borne out by the results to date that many, if not most, protein-protein associations involve specific salt-bridges. The modified proteins, in this study are the penicillin interactive proteins of Gram (+) and Gram (-) bacteria and the other morphogene proteins (MGPs) that interact with them. They are isolated by RPLC (7) following the F/S label. The proteins are unambiguously identified by computer based analysis of MALDI-TOF generated data from the tryptide and chymotryptide maps. The F/S labeled active site peptide is also identified.

n . EXPERIMENTAL MATERIALS AND METHODS A, Synthesis of active site directed fluorescence/spectroscopic (F/S) labeled filactams The synthesis of dansyl amino penicillanic acid (DNS-APA, I) was carried out according to the procedure outlined in Ref. (8). The e-dansyl monocyclic p-lactam of lysine (II) prepared essentially as described for another monocyclic p-lactam (9). JXH2

CH3OONH

II

B.

000'

^^<e"^^ '{CH2)4. NHSO2

©-N(CH3)2 (0>

Labeling of bacterial cells The labeling of Gram positive and Gram negative penicillin binding proteins (PBPs) is outlined in Scheme 1. Log phase bacterial cells were harvested by centrifuging them at 12,000 rcf for 10 minutes at 4''C. The cell pellet was washed twice with the 50 mM Tris HCl buffer (pH=7.2) and divided into three parts. One part of the pellet (control) was suspended in 500 ML of the Tris buffer and then added 25/uL of 15% SDS was added. It was left standing for 5 minutes. The other two equally divided pellets were then resuspended in 200 ML of buffer each and treated with 100-150 u^g dansyl (I)X APA or monocyclic p-lactam of N-€-dansyl lysine (II). The cell pellets were incubated for 20 minutes at 37°C. Treated cells were washed 3-4 times with buffer by centrifuging them for 5-10 minutes at 12,000 rcf and at 4"C so as to get rid of excess label as determined by visual inspection of the supernatant with UV irradiation. The cell pellets were resuspended in 200 uL of the Tris buffer. The

SCHEME 1: LABELING OF PBPS AND PBP-MORPHOGENE PRODUCT(MGP) COMPLEXES BY FLUORESCENCE SPECTROSCOPIC (F/S) PROBES BACTERIAL CELLS F/S probe 2. 1 Centrifuge

CONTROL CELLS

i LABELED CEUS | C2N2

CROSS-LINKEO PBP-MGP COMPLEXES

Centrifuge SOLUBLE UNLABELED PBP-S Reverse Phase Chromatography

Centrifuge Centrifuge SOLUBLE F/S LABELED PBP-MGP COMPLEXES

SOLUBLE F/S LABELED PBP COMPLEXES RPLC

CHROMATOGRAPHICALLY RESOLVED F/S LABELED PBP-MGP COMPLEXES

CHROMATOGRAPHICALLY RESOLVED F/S LABELED PBP COMPLEXES

SCHEME 2 : PROTEIN IDENTIFICATION CHROMATOGRAPHICALLY RESOLVED F/S LABELED PBF/FBP-MGP COMPLEXES

CHROMATOGRAPHIC FRACTION Enzymatic Digestion

PEPTIDE FRAGMENTS

MALDI-TOF/LSIMS Mass Spectrometry

Search the Protein Sequence database for muitipeptides of individual proteins using "MS-FIT' Program

Generation of Sequence fragments with matching Mol. wt.(s)

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of the Tris buffer. The label only pellet is treated with 25 /A. of 15% SDS and left standing for 5 minutes. C. Cyanogen Treatment Cyanogen may be obtained commercially or prepared readily in one step from AgCN(lO). Cyanogen is far less toxic than HCN, HjS or CO, but should be handled carefully (11). The headspace over the second F/S labeled pellet was swept out with -5 mL of cyanogen and allowed to stand at room temperature for 30 minutes. The color of cell pellet changes from cream to brown and the pellet disintegrates. This treatment was repeated three times with the same amount of CjNj. After the final CjNj treatment, the cell pellet was subjected to 25 yuL of 15% SDS for 5 minutes. The total volume was brought to 1 mL by adding Tris HCl buffer to each set and the cells were lysed by sonication. Cellular debris was removed by centrifugation at 12,000 rcf for 10 minutes at 4"C. The supernatant wasfilteredthrough 0.22 ^.m nylon membrane filter and diluted 5-10 fold before analysis by HPLC. The unused portion of the labeled supernatant was kept frozen at -80"C. D. Reverse phase chromatographic separation of penicillin binding proteins (PBPs) and cross-linked PBP-morphogene products (See Scheme 2) The chromatographic separation of hydrophobic membrane proteins, as well as hydrophilic "hybrid" proteins, was carried out using Microsorb-MV C18 reverse phase column (Rainin Instruments; 4.6x250 mm, 300 A pore size, 5 M particle size). The mobile phase for chromatography consisted of 99.9% acetonitrile with 0.1% trifluoroacetic acid as one solvent and 99.9% water with 0.1 % trifluoroacetic acid as the other solvent. The sample size was 50-80 yuL with lOOyuL loop attached to the injector. A gradient of 0-100% acetonitrile in 60 minutes was used with monitoring at 280 nm and 320 nm. A minimum of ten collections were made and the separated fractions were pooled. The column was cleaned periodically by injecting 100 ML of trifluroethanol (2-3 times) (12). The rechromatography of collected protein fractions was done using solvent gradients related to the retention time of that fraction in the original chromatographic profile. E. Enzymatic digestion of the RPLC purified penicillin binding proteins and "hybrid" proteins The enzymatic digestion of the purified proteins was carried out as described (13). The enzymes TPCK-treated pancreatic trypsin (EC 3.4.21.4; Type XIII) and TLCK-treated pancreatic chymotrypsin (EC 3.4.21; Type VII) werefromSigma Chemical Company. The amount of proteinase used was -2% of the concentration of the purified protein. The proteins were dissolved in 200 yuL of 0.1 N ammonium bicarbonate buffer (pH=7.0), followed by addition of the enzyme dissolved in deionized water. The enzymatic digestion was carried out by incubating the samples for -18 hours at 37° and the reaction was stopped by addition of 10% trifluoroacetic acid solution. The proteins were lyophilized.

Intra-CclMar Protein-Protein Interactions

473

E

Mass spectral analysis of digested proteins The samples were analyzed on the VG TofSpec-SE MALDI TOF mass spectrometer in the reflectron mode with positive ion detection. The samples were spotted on the sample plate in acetonitrile:water (60:40) or chloroform: methanol:TFA (1:1:0.1) mixture plus ammonium sulfate on alpha-C (a-cyano-4hydroxycinnamic acid) matrix. The ionization of the samples was carried out with Nd:YAG laser at 355 nm or nitrogen laser at 337 nm. Some of the fractions were analyzed by SIMS on a Kratos 890 mass spectrometer equipped with a Phrasor Scientific SIMS source. The mass spectral data were analyzed by the MSFIT program at the University of California, San Francisco. i n . RESULTS A,

F/S labeled PBPs The F/S labeled p-lactams were prepared as described. On the basis of their mode of action against bacteria I is termed "lytic" and II, "non-lytic." That is, diastereomers of II produce the microbiological Liesegang effect (MLEs)(14). I and n each produced characteristic chromatographic patterns in the F/S labeled penicillin interactive proteins, the penicillin binding proteins (PBPs), by RPLC (7). The hydrophobic PBPs elute much later than the cytosolic proteins. When monitored at 320 nm I-labeled PBPs showed a pattern of eight or nine major peaks (Fig. la) and >20 minor peaks. II-Labeled PBPs appeared essentially as one peak (Fig. lb). The major peaks were rechromatographed before further analysis of their proteinase digests by MALDI-TOF and/or SIMS. Two sets of peptides were generated from trypsin and chymotrypsin treatment. Characteristically each digest showed among the peptides only one F/S labeled peptide, the active site labeled peptide. The entire digest was needed for peptide mass mapping (15). The details of peptide mass-fmgerprinting of only one fraction out of several (Fig. la) analyzed is shown here. Digestion of fraction 1 (Fig. la) by the two proteinases was carried out. Shown are the results of MALDI-TOF analyses of the resultant tryptides (Fig. 2a) and chymotryptides (Fig. 2b). After eliminating peaks that were ambiguous either because (a) they could arise from more than one PBP or (b) could come from the proteinase itself, we were left with a set of peptide identities associated only with PBP IB. A typical result is shown in Tkble 1. In this case as with other reports the mass window used to examine the MALDI-TOF was less than ±3 amu as consistent with the range used in other studies (15-18). Peptide mass-fingerprinting of the I-labeled PBPs (Fig. la) revealed the eight known PBPs of E, coli (Tkble 2). Unlike SDS-PAGE analysis of labeled PBPs (19), the elution order by RPLC is not related directly to molecular weight but is dictated by hydrophobicity. II-Labeled PBPs presented only one peak (Fig. lb). Peptide-mass fingerprinting of the chymotryptic and tryptic digests revealed that it was a

S. Bhardwaj and R. A. Day

474

\2j4i6f

*320

'320

'320

'320

Figure 1. Chromatographic Traces of F/S Labeled Proteins from E. coli: (a) from i treated cells, (b) from i i treated cells, (c) from cells treated sequentially with i and C2N2, and (d) from cells treated sequentially with 11 and C2N2.

/ntra-Cellular Protein-Protein Interactions

400

600

800

1000

M/Z 1200

1400 M/Z 1800

475

1600

2200

2000

2600

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S. Bhardwaj and R. A. Day

complex containing the eight well-known PBft plus a candidate PBP, PHSE (Tkble 2). B.

Cyanogen linking ofPBPs to other morphogene proteins The I-treated E, coli cells were treated with C2N2 and again carried through the procedures up to and including peptide mass-fingerprinting. A new set of F/S labeled peaks intermediate in retention times between the early putative cytosolic proteins and the later eluting PBPs showed combinations of PBPs and known MGPs (20,21). The chromatographic profile of these "hybride" proteins is shown in Figure Ic. Space only allows us to show the identity of proteins found in fraction 1 (Tkble 3). The cyanogen trapped F/S labeled complexes reveal only PBPs and MGPs. Donachie (21) summarizes their functions. Chromatography of the E. coli proteins from the sequentially II and cyanogen treated cells (Fig. Id) showed F/S labeled components. The peptide massfingerprintingof proteolytic digests of these purified components showed only PBPs and MGPs (20). In Tkble 4 are shown the constituent proteins of one component. Found here are all the PBPs, save PBPS and PBP7, and seventeen MGPs. Not shown here are the results from a parallel analysis of the Gram (+) B, subtilis which gave entirely analogous results with I and II with and without C2N2 treatment (20). IV, A.

DISCUSSION

Specificity The most significant measurements of cellular processes must be done with no perturbation of the process. While zero perturbation is probably not attainable, nevertheless minimization of the perturbation is important. Cyanogen as a reagent readily diffuses into a cell, is itself apolar and noninteractive until it participates in a specific reaction. Its specificity is very high for hydration by bi-functional catalysis (6). When that catalysis is provided by a salt-bridge, it is accompanied by conversion of the salt-bridge to a covalent link. This study strongly supports a high specificity for pre-formed salt-bridges. The E, coli K-12 genome has almost completely been sequenced. It contains thousands of structural genes (3469 according to the data base Swiss Prot. r33). Of these, almost 70 have been identified as PBPs and MGPs (21). Wthout cyanogen treatment only PBPs were found in the F/S labeled proteins. With cyanogen only other MGPs were found covalently linked to the PBPs. That none of the other thousands of proteins became linked provides an extremely stringent internal control. It can be anticipated that this is a general property of protein-protein interactions and that any appropriately labeled protein could become a productive target for identification of proteins that may be interacting with it. It should be noted that in this study the C2N2 treatment

Intra-CellulsiT Protein-Protein Interactions

477

Table 1. Peptides Identified from Mass Spectral Data of Chymotryptides Shown in Fig. 2a Showing Identification of PBPIB. There were 7 matches P02919 penicillin-binding protein IB (PBP-IB), (94267.0Da) Mass Found

Mass Matched

Delta Da

Sequence Start

Peptide Sequence

End 540

(W)IADAPIAL(R)

1017.700

1016.556

1.144

415

422

(F)MQLVRQEL(Q)

1017.700

1019.451

-1.751

734

743

(L)YGASGAMSIY(Q)

1099.600

1100.610

-1.010

216

225

(F)VPRSGFPDLL(V)

1594.300

1592.811

1.489

202

215

(L)ITMISSPNGEQRLF(V)

2026.840

2027.997

-1.157

129

145

(L)EATQYRQVSKMTRPGEF(T)

2435.030

2437.040

-2.010

787

808

(W)TSDPQSLCQQSEMQQQPSGNPF(D) 1

Unmatched masses: 824.2 1126.7 1879.1 2515.1

Table 2. Identification of PBPs of E, coU K12 (ATCC 29079) Labeled with F/S labeled plactams. The intact log phase cells were treated with the F/S p-lactams for 10 minutes and then disrupted by sonication. After removal of the debris by centrifugation (10,000 rpm, 10 min.), ahquots of the supemate were separated chromatographically as described (39). The F/S labeled peaks were isolated and rechromatographed on a shallower gradient. The purified peaks were divkied and subjected to tiyptic and chymotryptic digestion respectively. The digests were analyzed by MALDI-TOF or by SIMS. The resultant mass spectral data were submitted for peptide mass fingerprinting at the UCSF Mass Spectrometry Facihty.

F/S p-Lactam

F/S Labeled Chromatographic Fraction

Protein

MoLWt.

PBPIA PBP4 PBPIB PBP7 PBP2 PBP6 PBP3 PBP5

93,636 51,798 94,266 34,245 70,856 43,639 63,877 44,444

PBPIA, PBPIB, PBP2, PBP3, PBP4, PBP5, PBP7, PHSE

was extreme, C2N2 driven reactions proceed at rates that vary by five orders of magnitude (4,6). For example, the subunits of hemoglobin have salt-bridges among them; it requires -60 seconds to covalently cross-link them (4). Thus, in this study, if there were any possibility of adventitious salt-bridge formation and conversion to covalently linked functionalities, it should have been seen.

|

478

S. Bhardwaj and R. A. Day

Table 3. Identification of PBPs and MGPs in Fraction I Isolated from the Sequential F/S p-Lactam DNS-APA (I) and CjN, IVeated E. coU Log Phase Cells. P^enthesis indicate possible but not confirmed by one or more peptides unique to the parenthetic protein. PBPS Protein PBPIA PBPIB PBP2 PBP3

Mol. Wt. 93,636 94,266 70,856 63,877

Protein (PBP4) (PBP5) PBP6 PBP7

Mol. Wt. 51,798 44,444 43,639 34,245

Other MorphogeneJProteins _ Protein Mol. Wt. Protein Mol. Wt. (FTSH) 70,708 ALRI 39,000 (SLT70) 73,369 (ENVC) 41.317 LON 87,438 MURA 44,817 SECA 101,909 FTSN 45,987 MUKB 176,935 FTSY 54,513

Table 4. Identification of PBPs and MGPs in Fraction I L Isolated from the Sequential F/S DNS-Monocyclic p-Lactam (II) and C^N^ Treated E, coli Log Phase Cells. Parenthesis indicate possible but not confirmed by one or more peptides unique to the parenthetic protein. Other Morphogene Proteins

PBPS Protein PBPIA PBPIB PBP2 PBPS

Mol. Wt 93,636 94,266 70,856 63,877

Protein Mol. Wt. (PBP4)r 51,798 (PBP5) 44,444 (PBP6) 43,639

Protein MUKB SECA (LON) (FTSH) SLT70 (MURE)

Mol. Wt. 176,935 101,909 87,438 70,708 78,969 53,212

Protein (FTSY) ENVC MURA (DDLA) ALRI MREB

Mol. Wt 54,513 41,317 44,817 39,315 39,060 39,952

B. Membrane proteins Membrane proteins are difficult to work with. Solubilization in a semifunctional form is possible with the aid of certain non-ionic detergents (3); the sheath of detergent molecules attached to the hydrophobic regions of these molecules compromise studies of interactions of membrane proteins. Thus, a method applicable to an intact cell or organelle may have some value. The system here involves membrane proteins (PBPs) and cytosolic proteins (most of the other MGPs) and thus demonstrates that such proteins and their interacting ligands can be accessed. C Covalently linked complex ofpenicillin interactive proteins The non-lytic II labeled a single peak in the chromatographic profile (Fig. lb). Since the denaturing conditions of isolation and chromatography separate the I-labeled PBPs (Fig. la), it is clear that non-lytic II prevents their separation from a covalently bonded complex. As with I-treatment, Il-treatment must be followed with C2N2 to show MGP association. More MGPs are associated in the latter case. The inference is that a covalently bonded network is normal and prevents perforation and lysis as had been observed (22); the typical lytic p-lactam causes this dissociation of the complex murasome situated

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479

in a pre-existing opening in the cell wall which is, then, a factor of importance in bacterial cell death. When the complex breaks up the cytosol is released. D. Sites of Salt Bridging There are two important consequences of the C2N2 treatment for the analysis. The first is that some peptides that contain cross-linked residues will be diminished and may not appear in the MSFIT analysis. The second is that such cross-linked residues identify the salt-bridged site(s); in the first phase of this study we identified only the interacting proteins. E. Limitations of the Method Thus far no data have been found which indicate that C2N2 produces zn/^r-molecular condensations between proteins that have no normal association to form complexes. Neither the published papers cited here nor in an extensive unpublished body of experiments from this laboratory where conditions had been optimized for z/z/^r-molecular amide bond formation had any detectable condensation been seen. The only important limitation arises from the peptide mass fragment analysis. It appears to be completely unambiguous for single proteins (Table 2). As the number of proteins in a mixture or in a complex increase the number of unambiguous M/Z values from the MALDI-TOF analysis decrease. An important observation is that when ambiguities arose in this study, only MGPs appeared in the C2N2 driven association. None of the other ~ 3400 proteins showed up. At this stage of development, it is clear that maximum complexity is limiting. For the ~70 MGP system there is a need to follow the time course of condensation not only to deal with complexity, but more importantly to collect information about rates and degree of association. V. CONCLUSION There is no way to know what fraction of protein-protein interactions involve salt-bridges; however, recent crystallographic studies of protein-protein complexes of cytokines and growth hormone with their receptors show multiple salt bridges at the interfaces (23). Those that do not have salt bridges or hydrogen-bonded carboxylate carbinol associations (24) must remain undetectable by this technique. However, these early results show a high specificity and a good fraction of the expected interactions. It seems likely in the case of PBP-MGP interactions that we have seen vegetative state complexes and can expect a different set for cell division. When the septasome complex(es) and any other specialized complex(es) are characterized, we feel that a different set of MGPs will be found. This technique also provides a means of identifying each covalently linked, inter-protein salt-bridged site through identification of the peptide component contributed by each protein of the linked pairs and/or multimers. This is an aspect of this ongoing project. Such data will provide more specific

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information on how the almost 70 MGP/PBPs mutually modulate each other and cell wall synthesis (20). ACKNOWLEDGEMENTS. We thank Professor Jayasimhulu for the SIMS analyses and Mr. J. E. Carlson for the MALDI-TOF work. We appreciate being given access to the MALDI-TOF unit by Professor J. Monaco. We appreciate having access to the MASS-FIT program at UCSF and assistance from K. Clauser. The cyanogen aspect of this work was developed in the past under NIH Grant GM42697. REFERENCES 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17. 18. 19. 20. 21.

22. 23. 24.

Phizicky, E.M. and Fields, S. (1995) Microbiol. Rev. 59, 94-123. M e n , J.R, Walber& M.W., Edwaixls, M.C., and EUedge, S.V. (1995) Trends Biochem. 5c?i., 20, 511-516. Kyte, J. (1995) Structure in Protein Chemistry, Garland Publishing (New York), p. 520 j ^ Day, R.A., Kirley, J., Tharp, R, Flicker, O., Strange, C. and Ghenbot, G. (1989) In (T.E. Hugli, Ed.) Techniques in Protein Chemistry, Academic Press, San Diego, pp. 517-525. Day, R.A., Hignite^ A. and Gooden, W.E. (1995) In (J.W. Crabb, Ed.) Techniques in Protein Chemistry VI, Academic Press, San Diego, pp. 435-442. Day, R.A., Tharp, R.L., Madis, M.E., Wallace, J.A., Silanee^ A.A., Hurt, P. and Mastruserio, N. (1990). Peptide Res. 3, 169-175. Day, R.A., Ahluwaha, R. and Du, Y. (1994) LC-GC12, 384-394. Cartwright, S.J., Tan, A.K. and Fink, A.L. (1989) Biochem. J., 263, 905-912. Ahluwalia, R., D ^ , R.A., and Nauss, J. (1995) Biochem. Biophys. Res. Common. 206, 577-583. Kirley, J.W., Day, R.A. and Kreishman, G.P. (1985) FEBS Lett. 193, 145. Fassett, D.W. (1983) In (F.A. I ^ y , Ed.) Industrial Hygiene and Toxicology, 2nd Ed., Interscience Publishers, NY, p. 2003. Bhardwaj, S. and Day, R.A. (1996) submitted to LC/GC. Lee, T.D. and Shively, J.E. (1990) Methods Enzymol. 193, 361-374. Day, R.A., Bhardwaj, S. and Bai, H. (1995) Miami Bio/Technology Short Reports 6, 88. Henzel, W.J., Billeci, T.M., Stults, J.T., Wong, S.C, Grimly, C and Watanabe, C. (1993) Proc. Natl. Acad. Sci. USA 90, 5011-5015. Pappin, D.J.C., Hojrup, P. and Bleaby, A.J. (1993) Current Biol. 3, 327-332. James, P., Quadron, M., Carafoh, E. and Gonnet, G. (1993) Biochem. Biophys. Res. Commun. 195, 58-64. Hynes, G., Sutton, C.W., U, S., and WiUison, K.R. (1996) EiSEB J. 10, 127-147. Waxman, D.J. and Strominger J.L. (1983) Ann. Rev. Biochem. 52, 825-869. Bhardwaj, S. (1996) Ph.D. Dissertation, University of Cincinnati. Donachie, W.D. (1993) In (M.A. de Pbdro, J.-V. Holtje and W. Loffelhaixlt, Eds.) Bacterial Growth and Lysis. Metabolism and Structure of the Bacterial Sacculus, Plenum, New York, pp. 409-18. Giesbrecht, P., Kersten, T., Madela, K., Grob, H., Bliimel, P. and Wecke, J. (1993) in de Pedro et al. op. cit. pp. 393-407. Ealick, S., Thiel, D , le Du, M., Walter, R., D'Arey, A., Chene, C , Fontoulabis, M., Garotta, G. and Winklet, F. (1996) Prot. Science 5 (Suppl. 1) 59. Karagozler, A.A., Ghenbot, G. and Day, R.A. (1993) Biopolymers 33, 687-692.

Use of Synthetic Peptides in Mapping the Binding Sites for hsp70 in a Mitochondrial Protein Antonio Artigues, Ana Iriarte, and Marino Martinez-Carrion Division of Molecular Biology and Biochemistry. School of Biological Sciences. University of Missouri-Kansas City, Kansas City, MO 64110

I. Introduction The members of the 70-kDa heat shock protein (hsp70) family perform functions that are essential for cell viability, both under normal and stress conditions. Constitutively expressed hsp70s are thought to be involved in the folding and assembly of newly synthesized proteins, disassembly of oligomeric proteins, protein degradation, and the transport of nascent peptide chains across membranes (l, 2). The structure of hsp70 consists of a variable C-terminal peptidebinding domain and a highly conserved N-terminal ATPase domain. The crystal structure of a peptide-C-terminal domain complex shows that the peptide substrate is bound in an extended conformation through numerous interactions from both side chains and backbone groups (3). As with all molecular chaperones, hsp70 shows a remarkable selectivity for unfolded structures, but low specificity towards the sequence of its potential peptide substrates. Among the few consensus features identified in peptides binding to hsp70 with high affinity is the presence of internal hydrophobic residues (3, 4), which agrees with the proposed role of this chaperone in binding to hydrophobic regions of unfolded proteins normally hidden in the native structure. However, a great variety of synthetic peptides, including organellar targeting sequences containing basic residues (5-7) also bind to hsp70. Perhaps the binding sites recognized by hsp70 depend on the specific role fulfilled by the chaperone with a given substrate. In any case, the precise characteristics of the targeting sequences that determine the recognition and binding of hsp70 to its multiple substrates remain largely obscure. The cytosolic (cAAT) and mitochondrial (mAAT) isozymes of aspartate aminotransferase share a significant degree of sequence homology TECHNIQUES IN PROTEIN CHEMISTRY VIII Copyright © 1997 by Academic Press All rights of reproduction in any form reserved.

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(63%), and almost identical crystallographic structures (8, 9). The mitochondrial isozyme is synthesized in the cytosol as a precursor protein (pmAAT) with a 29-residue presequence peptide that is required for targeting and import into mitochondria. Despite the broad substrate specificity of hsp70 mentioned before, this chaperone is able to discriminate between the two highly homologous AAT isozymes. Following either synthesis in cell-free extracts (lO) or refolding firom acid-denatured states the mitochondrial isozyme binds to hsp70, whereas its cytosolic counterpart does not interact with this molecular chaperone. Thus, these isozymes provide a particularly attractive system to delineate sequence elements that might be binding motifs for hsp70 in pmAAT, but are absent or hidden in cAAT. An initial screening with a series of tetradecameric peptides corresponding to the complete amino acid sequence of pmAAT has led to the identification of several binding regions scattered over the full length of the polypeptide chain.

II. Methods A. Protein

Purification

Purification of pmAAT and cAAT was carried out as previously described (11,12). After concentration by ultrafiltration (Amicon centricon, 30,000 molecular weight cutoff), the proteins were transferred to 2 mM Tris HCl, pH 7.5, a low ionic strength buffer suitable for the subsequent unfolding of the proteins at low pH (pH=2.0). Protein concentrations were estimated from the absorbance at 356 nm or 362 nm of the pyridoxal-5'-phosphate (PLP) cofactor bound to either pm- or cAAT, respectively (molar absorption coefficient of 8,500 M cm ), and Mr=46,597 for pmAAT or Mr=46,399 for cAAT. Hsp70 was purified following published procedures (13). Following the last ammonium sulfate precipitation, the protein was exhaustively dialyzed against 25 mM Tris HCl, pH 7.5, 20 mM NaCl, 10 mM p-mercaptoethanol and kept at 4 ^C until use. Hsp70 protein concentration was measured using a molar absorption coefficient at 280 nm of 47,800 M'^ cm"^ and Mr=70,000 (14).

B. Unfolding and Refolding of pmAAT and cAAT For the reversible acid unfolding of pmAAT, a stock solution of the enzyme in 2 mM Tris HCl, pH 7.5 was denatured by addition of diluted HCl to pH 2.0, followed by incubation for 90 min at room temperature (15). Refolding of pmAAT was performed by rapid dilution of the unfolded protein to 1.8 pM final concentration in refolding buffer (40 mM

Mapping Binding Sites for hsp70 in Mitochondrial Protein

483

Hepes, 0.1 mM EDTA, 1 mM DTT, 10 jaM PLP, pH 7.5) at 10 OC. When studying the effect of hsp70 on the refolding of the enzyme, hspTO ( 1.8 |LiM) was present in the refolding mixture before initiation of refolding by addition of the unfolded protein. For competition studies, hsp70 (1.8 |iM) was preincubated with a 70-fold molar excess of each peptide (120 |LiM) in refolding buffer for 2 to 16 h at 10 ^C before addition of aciddenatured pmAAT (1.8 |iM). The reaction mixture was incubated for an additional 2 h to allow complete refolding of the pmAAT molecules that were not complexed with hsp70. Binding of a peptide to hsp70 would result in an increase in the fraction of pmAAT molecules that are allowed to recover full catalytic activity. Therefore, peptide competition was examined by determining the recovery of pmAAT activity in the presence of hsp70 and in the presence or absence of peptide. The effect of the peptides on the spontaneous refolding of pmAAT was analyzed by measuring the yield of reactivation in samples containing 120 |LiM peptide but no chaperone. The yield of reactivation was determined by measuring the recovery of transaminase activity. Data are expressed as percentage relative to the activity of a control sample maintained under identical conditions.

C. Synthesis and Purification of Peptides The collection of 14-residue peptides spanning the complete amino acid sequence of pmAAT was a generous gift from Dr. B.M. Conti-Tronconi (University of Minnesota). The peptides were synthesized according to Houghten (16). The purity of the peptides was assessed by reverse phase HPLC using a C18 column (Vydac 218TP, 250 x 4.6 mm) and an acetonitrile/water gradient (5 to 70% over 30 min) containing 1% trifluoroacetic acid. The purity of the different peptide preparations ranged from 50-95%. Most of the contaminating peptides represented truncated peptides randomly missing amino acids from incomplete coupling. For screening purposes, these peptides were used without further purification. Selected peptides were further purified by HPLC reverse phase chromatography on a BioRad C18 Hi-Pore RP-318 semipreparative column (250 x 10 mm), using the same gradient as before. Major peaks were collected and the full-length peptide peak was identified by amino acid composition analysis. Peptide concentration was determined based on the molar composition obtained by amino acid analysis. The presequence peptide, MALLHSGRVLSGM-AAAFHPGLAAAASARA, was also synthesized as a single 29-mer peptide in an Applied Biosystems 433A peptide synthesizer at the Molecular Core Facility of the School of Biological Sciences and purified by reverse phase chromatography as described above.

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D. Enzymatic

Activities

The transaminase activity was measured at 37 ^C using L-aspartate and a-ketoglutarate as substrates, using a coupled assay with malate dehydrogenase, as described previously (17). To measure the ATPase activity of hsp70, the chaperone (0.5 |LIM) was incubated at 37 ^C in 40 mM Hepes, 45 mM KCl, 120 ^iM MgCl2, 60 ^iM ATP, pH 7.5, in the presence or absence of different synthetic peptides (120 |LIM). At different incubation times, a 20-|LI1 aliquot was withdrawn and assayed for ATP content on a Turner TD 15-e Luminometer, using the ATP bioluminescence assay kit from Sigma and following manufacturer's instructions. This assay measures the light emitted upon spontaneous decomposition of adenylate-luciferin produced by luciferase from ATP and luciferin substrates (18). When ATP is the limiting reagent, the light emitted is proportional to the ATP present in the sample, and the concentration of ATP can be calculated by reference to an ATP standard curve. The rate of spontaneous hydrolysis of ATP was estimated in samples incubated at 37 ^C in the absence of hsp70.

III. RESULTS AND DISCUSSION A. Effect of hsplO on the Refolding of pmAAT and cAAT In vitro refolding of the acid-unfolded isozymes results in the reconstitution of native-like proteins. Figure 1 shows the yield of reactivation of cAAT and pmAAT (1.8 |LiM) in the absence or presence of hsp70 (1.8 |uM). In the absence of hsp70, a significant recovery of activity (70-80%) can be achieved following spontaneous refolding of both proteins. However, when hsp70 is present, the yield of reactivation of pmAAT is reduced considerably, whereas the yield of reactivation of its cytosolic counterpart is not affected. Inhibition of pmAAT refolding by hsp70 results in the formation of insoluble aggregates of the hsp70-pmAAT complex. hsp70 does not affect the activity of the native protein. This effect of hsp70 is specific, since addition of high concentrations (1 mg/ml) of other unrelated proteins such as bovine serum albumin, aldehyde dehydrogenase, or malic dehydrogenase does not prevent pmAAT refolding and reactivation (data not shown).

Mapping Binding Sites for hsp70 in Mitochondrial Protein

i

100

80

K//X

•I 60 > O CO 0 V-

o

40

485

]

0

> 20

#

——

— \ — —\— ^

Hsp70 cAAT

pmAAT

Fig. 1. The effect of hsp70 on the refolding of cAAT and pmAAT. Refolding of acid unfolded cAAT or pmAAT was performed by rapid dilution of the denatured enzymes in the refolding buffer to a final protein concentration of 1.8 fiM. When present, hsp70 (1.8 ^iM) was added to the refolding buffer before initiation of the refolding reaction. After incubation for 120 min at 10 °C, the transaminase activity recovered was measured as indicated under Methods. Reactivation data are expressed relative to that of the native enzyme incubated under identical conditions.

B. Competition by pmAAT Peptides tact Unfolded pmAAT to hsp70

of Binding of In-

Taking advantage of the fact that hsp70 binds to unfolded pmAAT and markedly reduces the yield of reactivation (from 70% to 20%), we developed a competition assay to search for putative binding sites for hsp70 in the pmAAT polypeptide. In this assay, each peptide in a collection of 43 synthetic tetradecamers spanning the entire amino acid sequence of pmAAT was tested for its ability to compete with unfolded pmAAT for binding to hsp70. Since binding to hsp70 stops refolding of pmAAT, competition by a given synthetic peptide should result in an increase in the fraction of pmAAT activity recovered. Thus, the relative affinity of the different 14-mer peptides for binding to hsp70 was established by comparing the yield of pmAAT reactivation in the presence of hsp70 alone, or hsp70 that had been preincubated with a 70-fold molar excess

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Table I. Selective binding of pmAAT peptides to hsp70. Synthetic tetradecamer peptides corresponding to the amino acid sequence of rat liver pmAAT (p-1 to p-43 from the N-terminai to the C-terminal end) were tested for their ability to compete for the binding of unfolded pmAAT to hsp70 as described under Methods. The percentage of pmAAT activity recovered relative to that obtained in the presence of hsp70 alone (20%) represents an index of the magnitude of peptide competition of pmAAT binding to hsp70: <25% (-), 26-35% 0. 36-50%(+), 51-65% (++), >66 % (+++). The maximum yield of reactivation in the absence of hsp70 is 75 5 %.

Peptide sequence

Number

Activity recoveredCompetition a (+ hsp70, %)

none presequence MALLHSGRVLSGMA SGMAAAFHPGLAAA LAAAASARASSWWT SWWTHVEMGPPDPI PDPILGVTEAFKRD FKRDTNSKKMNLGV NLGVGAYRDDNGKP NGKPYVLPSVRKAE RKAEAQIAGKNLDK NLDKEYLPIGGLAD GLADFCKASAELAL ELALGENSEVLKSG LKSGRFVTVQTISG TISGTGALRVGASF GASFLQRFFKFSRD FSRDVFLPKPSWGN SWGNHTPIFRDAGM DAGMQLQGYRYYDP YYDPKTCGFDFSGA FSGALEDISKIPEQ IPEQSVLLLHACAH ACAHNPTGVDPRDE PRPEQWKEMAAVVK AVVKKKNLFAFFDM FFDMAYQGFASGDG SGDGDKDAWAVRHF VRHFIEQGINVCLC VCLCQSYAKNMGLY MGLYGERVGAFTW FTWCDKAEEAKRV AKRVESQLKILIRP LIRPLYSNPPLNGA LNGARIAATILTSP LTSPDLRKQWLQEV LQEVKGMADRIISM IISMRTQLVSNLKK NLKKEGSSHNWQHI WQHITDQIGMFCFT

20 p r e - p * b 58 p-1 C 45 p-2 C 47 p-3 c 50 p-4 d nd p-5 * 76 p-6 37 p-7 32 p-8 35 p-9 20 p-10 6 p-11 65 p-12 80 p-13 75 p-14 45 p-15 * 80 p-16 38 p-17 30 p-18 53 p-19 43 p-20 62 p-21 e nd p-22 42 p-23 * 80 p-24 52 p-25 11 p-26 39 p-27 e nd p-28 52 p-2 9 e nd p-30 69 p-31 * 80 p-32 45 p-33 52 p-34 34 p-35 46 p-36 44 p-37 33 p-38 e nd

+-f+ + + nd +4-+

+-h ++-h +++ + +-1--H + ++ + ++ nd + +++ + + nd + nd +-I-+ ++-H -f -I+ + nd

Mapping Binding Sites for hsp70 in Mitochondrial Protein FCFTGLKPEQVERL VERLTKEFSVYMTK YMTKDGRISVAGVT AGVTSGNVGYLAHA LAHAIHQVTK (a) (b) (c) (d) (e) (*)

p-39 p-40 p-41 p-42 p-43

* * * *

52 70 74 71 20

487 + +++ +++ +++ ^

Reactivation data are expressed relative to a sample of native pmAAT incubated under identical conditions. A 29-residue peptide corresponding to the entire presequence region of pniAAT. Tetradecameric peptides with 4-residue overlapping regions spanning the presequence and the first five residues of the mature sequence. Binding of peptide p-4 to hsp70 could not be analyzed by the competition assay due to its strong inhibition of pmAAT refolding, nd, not determined Peptides p-21, p-27, p-29 and p-38 show very low solubility in aqueous solutions. These peptides were selected for further characterization of their hsp70 binding by ATPase activity stimulation assays.

of the peptide. The results of this competition cissay are summarized in Table I. Addition of several synthetic peptides (p-5, p-12, p-13, p-15, p-23, p-30, p-31, p-40, p-41; p-42, rated as +++ in Table I) produced a complete reversal of the hsp70-induced reduction in pmAAT refolding, indicating that they bind stroncfly to the chaperone and thus prevent formation of a hsp70-pmAAT complex. Another group of peptides (labeled as ++ or + in Table I) induce only a partial recovery of pmAAT activity in the presence of hsp70, suggesting a lower affinity for binding to hsp70. Finally, equivalent concentrations of several peptides (those rated or in Table I) had very little or no effect on the interaction of pmAAT with hsp70, indicating that they do not bind to the chaperone. Obviously this competition assay would not be feasible if the synthetic peptides interfered with the spontaneous refolding of pmAAT. This was tested by monitoring the yield of reactivation in the presence of concentrations of peptide similar to those used in the competition experiments (120 M) but minus hsp70. Among the 43 tetradecamers tested, only p-4, whose sequence corresponds to the N-terminal peptide of the mature portion of pmAAT, had a marked effect on the recovery of pmAAT activity (Table I). In the presence of this peptide, the yield was reduced from about 7 0% to 8%, and there was extensive aggregation of the refolding polypeptide. Consequently, binding of this peptide to hsp70 could not be tested using the competition assay. Since in the native pmAAT dimer this N-terminal peptide interacts strongly with a hydrophobic pocket on the surface of the neighboring subunit (8, 9) the presence of an excess of the synthetic peptide may interfere with the dimerization step in the folding pathway. On the other hand four peptides (p-21, p-27, p-29 and p-38) have a very limited solubility in aqueous solutions and therefore could not be tested at concentrations similar to those used for the other peptides. When used at a lower concentration, they did not show any effect on either the spontaneous refolding of pmAAT or its binding to hsp70.

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The peptide sequences with highest affinity for binding to hsp70 are not clustered in a specific region of the polypeptide pmAAT chain, but rather are scattered over the entire amino acid sequence of the enzyme. The sequence of these regions shows several of the characteristics described for peptides with high binding afiBnity to hsp70 (6, 19), such as the presence of hydrophobic and positively charged residues. Moreover, with the exception of the presequence peptides, they are localized within regions of the enzyme that are normally hidden in the folded state of the protein. However, sequence homology analysis of the different high affinity peptides did not allow for the identification of a consensus sequence, which agrees with the known broad specificity of hsp70 for peptide substrates (l, 20, 21). In addition, the majority of the peptides with high binding affinity to hsp70 map to regions in the amino acid sequence of pmAAT having the lowest degree of homology with the corresponding position in the cytosolic homologue. In addition to the collection of tetradecamers having 4-residue overlapping ends (see Table I for sequences), we also tested the competition of a 29-residue peptide corresponding to the entire presequence peptide (pre-p in Table I). Interestingly, the effect of this peptide in the competition assay is more pronounced than that of each of the 14-mer peptides (p-1, p-2, and p-3) containing sequence elements from the same region (see first four entries in Table I). One possible explanation for this different behavior is that the targeting sequence recognized by hsp70 in the intact presequence peptide has been split in the three related shorter peptides. The effect of the presequence peptide is of particular interest since it is unique to the mitochondrial enzyme. The competition of the presequence peptide with pmAAT for binding to hsp70 is concentration dependent, with an apparent affinity constant of about 9.4 jiM (data not shown). Preincubation of hsp70 with saturating concentrations of the presequence peptide also stimulates the ATPase activity of hsp70 (see below. Table II). Binding of other mitochondrial presequences to hsp70 has been recently reported (5, 7).

C. stimulation of the ATPase Activity High Affinity Binding Peptides

of hsp70

by

Hsp70 has a weak ATPase activity, with turnover rates ranging from 0.0004 to 0.0012 s"^ (14). Peptides binding to the C-terminal domain of hsp70 induce a conformational change in the N-terminal domain (6, 23, 24), which leads to a discrete stimulation of the ATPase activity. Therefore, binding of substrates to hsp70 can also be tested by monitoring changes in its ATPase activity. For this reason, we next examined the effect of several of the pmAAT tetradecamers on the ATPase activity of hsp70 using a sensitive bioluminescence assay to monitor the decrease in ATP concentration with time. All of the peptides assayed were repu-

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Table II. The effect of peptides on the hsp70 ATPase activity. Hsp70 ATPase activity was measured by monitoring the disappearance of ATP substrate over time using a bioluminescence assay as described under Methods. The concentration of the various peptides in the assay mixture was 120 |j,M. Peptide

ATPase activity (nmole/min/mg)

Stimulation ^

none

0.92

1.00

pre-p p-5 p-15 p-23 p-31 p-40 p-41 p-42

1.70 1.82 1.75 1.34 1.63 1.50 1.30 1.30

1.85 1.98 1.90 1.46 1.77 1.63 1.41 1.41

p-43

1.00

1.09

^ Activity in the presence of peptide/basal activity in the absence of peptide.

rifled by RP-HPLC before use. Stimulation of the ATPase activity correlated well with peptide binding data obtained from competition experiments. The presequence peptide and several of the 14-mer peptides showing maximal competition with pmAAT for binding to hspTO induced a 1.5 to 2-fold ATPase stimulation (Table II). In contrast, p-43, the C-terminal peptide from pmAAT which did not bind to hspTO according to the competition assay, showed no stimulation of the chaperone ATPase activity. IV.

CONCLUSIONS

Possible hsp70 binding sites on the primary structure of the pmAAT polypeptide have been identifled by competition studies in which, previous to the initiation of pmAAT refolding, hsp70 had been preincubated with a series of synthetic tetradecameric peptides spanning the complete sequence of pmAAT. The rationale of this approach was based on two assumptions: i) hsp70 binds peptides in an extended, or at least flexible, conformation, and ii) sequence homology analysis and the use of peptides derived from a known sequence will allow the identiflcation of peptide motifs responsible for the differential interaction of hspTO with two homologous proteins, pmAAT and cAAT. The flrst assumption has recently been strengthened by the publication of the crystal structure of the hspTO peptide binding domain (3). The second has led to the mapping of putative binding sites of polypeptide regions that show maximum sequence divergence between the two isozymes.

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490

100

200

300

400

Position

Fig. 2. Structural comparison between cAAT and pmAAT. The average sequence homology between cAAT and pmAAT was calculated using the Plotsimilarity program included in the Wisconsin Package of the Genetics Computer Group suite of programs (version 8.0, 1984) with a window size of seven residues, after the proteins were aligned inserting gaps where necessary to maximize homology. A score of 1.5 corresponds to a region of perfect homology. The dotted line represents the overall average similarity between the two proteins. Horizontal bars indicate the position of peptides showing maximum competition with pmAAT for binding to hsp70. The peptides are identified by numbers as assigned in Table I.

Mechanistic studies on the structure-function of hsp70 have shown that upon binding of peptides there is a conformational change in hsp70 that results in a slight stimulation of hsp70 ATPase activity. Release of peptide substrates is expected to be a slow step and may require coupling to ATP hydrolysis and possibly the cooperation of other molecular chaperones. Consequently, in the absence of any other cytosolic factors, the binding of peptides to hspTO is basically irreversible. Considering these properties, several strategies have been used to identify substrate recognition features of hsp70. An initial screening of a battery of peptides derived from pmAAT for their ability to compete with pmAAT for the formation of a complex with hsp70 has allowed for a fast, easy, and accurate identification of protein sequences that efficiently bind to hsp70. Confirmation of the binding of selected peptides has been obtained by measuring the stimulation of the hsp70 ATPase activity as a consequence of the conformational change induced upon substrate binding.

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491

With the exception of the presequence-containing peptides, and in agreement with the generally accepted mechanism of hsp70 action, the peptides that bind with high affinity to hsp70 comprise sequences that are hidden in the native state of the protein. These peptides contain central hydrophobic and basic carboxyl terminal amino acids, but few acidic residues. More interestingly, a sequence homology comparison of the cytosolic and mitochondrial protein sequences shows that the mitochondrial peptides binding to hsp70 correspond to regions of major sequence dissimilarity between the two isozymes (Figure 2). This suggests that sequence divergences observed between the mitochondrial and cytosolic isozymes may have arisen as a consequence of biochemical specialization to ensure the different interaction of each enzyme with the cellular machinery responsible for protein folding and translocation in vivo, thus promoting efficient import into the organelle of pmAAT and rapid folding in the cytosol of cAAT. Detailed analyses of the binding properties of each peptide, including the accurate determination of the binding affinity of each region as well as the identification of the critical residues involved in the peptidehsp70 interaction, are in progress. Information gathered from these studies should contribute to a better characterization of putative recognition sites responsible for the distinct interaction of the two isozymes with hsp70.

Bibliography 1. Mckay, D. (1993) Advances in Protein Chemistry 44, 67-98. 2. Hendrick, J.D., and Hartl, F.U. (1993) Annu. Rev. Biochem. 62, 349-384 3. Zhu, X., Zhao, X., Burholder, W.F., Gragerov, A., Ogata, CM., Gottesman, M.E., and Hendrikson, W.A. (1996) Science 272, 1606-1614. 4. Flynn, G.C., Pohl, M.T. Flocco, M.T., and Rothmann, J.E. (1991) Nature 353, 726730. 5. Endo, T., Mitsui, S., Nakai, M., and Roise, D. (1996) J. Biol. Chem. 271, 41614167. 6. Takenaka, I.M., Leung, S.M., McAndrew, S.J., Brown, J.P., and Hightower, L.E. (1995) J. Biol. Chem. 270, 19839-19844. 7. Schmid, D., Baici, A., Gehring, H., and Cristen, P. (1994) Science 263, 971-973. 8. Malashkevich, V.N., Strokopytov, B.V., Borisov, V.V., Dauter, Z., Wilson, K.S., and Torchinsky, Y.M. (1995) J. Mol. Biol. 247, 111-124. 9. Jansonius, J.N., and Vincent, M.G. (1987) In Biological Macromolecules and Assemblies (Jurnak, F., and McPherson, A., Eds.) Vol. 3, pp. 187-285, John Wiley & Sons Inc., New York. 10. Lain, B., Iriarte, A., Mattingly, J.R. Jr., Moreno, J.I., and Martinez-Carrion, M. (1995) J. Biol. Chem. 42, 24732-2739. 11. Altieri, F., Mattingly, J.R. Jr., Rodriguez-Berrocal, F.J., Iriarte, A., Wu, T., and Martinez-Carrion, M. (1989) J. Biol. Chem. 264, 4782-4786. 12. Mattingly, J. R., Jr., Iriarte, A., and Martinez-Carrion, M. (1995) J. Biol. Chem. 270, 1138-1148.

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13. Welch, W.J., and Feramisco, J.R. (1985) Molecular and Cellular Biology 5,1494914959. 14. Palleros, D.R., Welch, W.J., and Fink, A.L. (1991) Proc. Natl Acad. Sci. U.S.A. 88, 5719-5723. 15. Artigues A., Iriarte, A., and Martinez-Carrion, M. (1994) J. Biol. Chem. 269, 21990-21999. 16. Houghten, R.A. (1985) Proc. Natl. Acad. Sci. U.S.A. 82, 5431- 5135. 17. Martinez-Carrion, M., Turano, C, Chiacone, E., Bossa, F., Giartosio, A., Riva, F., and Fasella, P. (1967) J. Biol. Chem. 242, 2397-2409. 18. Leach, F.R., and Webster, J.J. (1986) Methods in Enzymology 133, 51-70. 19. Fourie, A.M., Sambrook, J.F., and Gething, M.-J. (1994) J. Biol. Chem. 269, 30470 - 30478. 20. Gilk, B.S. (1995) Cell 80, 11-14. 21. Hightower, L.E., Sadis, S.E., and Takenaka, I.M. (1994). In The Biology of Heat Shock Proteins and Molecular Caperones, (Morimoto, R.I., Tissieres, A. and Georgopoulos, C, Eds.) pp. 179-207, Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York. 22. Mattingly, J. R., Jr., Iriarte, A., and Martinez-Carrion, M. (1993) J. Biol. Chem. 268, 26320-26327. 23. Takeda, S., and McKay, D.B. (1996) Biochemistry. 35, 4636-4644. 24. Park, K., Flynn, G.C., Rothman, J-E., and Fasman, G.D. (1993) Protein Science 2, 325-330.

Interfacing Biomolecular Interaction Analysis with Mass Spectrometry and the use of Bioreactive Mass Spectrometer Probe Tips in Protein Characterization Randall W. Nelson, Jennifer R. Krone, David Dogruel, Kemmons Tubbs Department of Chemistry and Biochemistry Arizona State University Tempe AZ 85287-1604 Russ Granzow and Osten Jansson Pharmacia Biosensor AB, S-751 82 Uppsala, Sweden OVERVIEW The past decade has seen the development of new and powerful technologies capable of the accurate characterization of biomolecules with extreme speed and sensitivity. Two of these techniques, Biomolecular Interaction Analysis (BIA) and matrix-assisted laser desorption/ionization time-of-flight mass spectrometry (MALDI-TOF), lend themselves particularly to such analyses; the former ideally suited for the real-time investigation of biomolecular interactions, the latter finding much use in the qualitative assessment of analytes. Although the two analytical approaches operate on mutually exclusive detection principles (either surface plasmon resonance detection of a refractive index change or the physical determination the molecular mass of a gas-phase ion), they can share a common denominator ~ the use of affinity interactions in selecting the analyte. Interfacing of the two thereby creates a unique approach for the investigation of the kinetic parameters of biomolecular interaction (using BIA), and the unambiguous confirmation of the presence of targeted affinity ligands by direct mass analysis (using MALDI-TOF). In other applications, MALDI-TOF analysis can be extended beyond the primary role of protein molecular weight determination by combination with analytical enzymologies. The simplest use of enzymes in combination with MALDI-TOF is digestion of analytes into smaller fragments using endoproteases. The masses of the fragments are then determined in order to confirm or deny the sequence of the protein (or the presence of a given variant of the analyte). Traditionally, digestions are performed with both the analyte and enzymes in solution. As a result, autolysis signals are frequently observed in the mass spectra. Enzyme autolysis can be eliminated by using proteases immobilized to chromatographic supports, but generally at the expense of speed and sensitivity in analysis. An alternative to using enzymatically active chromatographic supports is to covalently attach enzymes to the surface of the mass spectrometer sample introduction device (probe). The probe device thus serves a two-fold purpose: as the enzymatic agent used for modification of the analyte, and, as a sample introduction device into the mass spectrometer. Over the past few years we have been developing new mass spectrometric approaches for the rapid, sensitive, and, accurate characterization of proteins. Reported here are some of our findings on the interfacing of Biomolecular Interaction Analysis with mass spectrometry, and the use of enzymatically active - or bioreactive - mass spectrometer probe tips in the characterization of analytes. TECHNIQUES IN PROTEIN CHEMISTRY VIII Copyright © 1997 by Academic Press All rights of reproduction in any form reserved.

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494

INTERFACING BIA WITH MASS SPECTROMETRY I.

Introduction

Biomolecular Interaction Analysis (BIA) is an acronym given to a number of techniques used in the characterization of bio-specific interactions. A form of the technique is based on the non-destructive detection principle of surface plasmon resonance (SPR), and is capable of monitoring the binding of an analyte to a surface-immobilized binding partner in real-time [1]. Briefly, a biosensor surface (chip) comprised of an affinity ligand-derivatized carboxylated dextran layer coupled to a thin gold surface is monitored using SPR while the chip surface is exposed to the complementary affinant. Differences in surface concentration resulting from ligand-affinant interaction are detected as a change in the SPR signal, expressed in resonance units (RU), with 1000 RU corresponding to a surface concentration of- 1 ng/mml The resulting sensorgrams report the mass quantity of analyte bound to the chip surface as a function of time. Sensorgram data, as a function of analyte concentration, can then be used to determine kinetic parameters, and molar absorptivity constants, of the interaction [2]. i?

100 nm dextran u.

_JL_ 1

m/z

50 nm Gold

Polarized Light Source Resonance

Signal r-\_qsigr Surface Plasmon Resonance

Fig. 1 Biomolecular interaction analysis/mass spectrometry (BIA/MS). Biosensor chips are derivatized with affmant (or used with an affmant of streptavidin) and used in the BIA analysis of biological fluids. The chips are then introduced into a MALDI time-of-flight mass spectrometer and retained ligands analyzed by virtue of molecular weight.

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495

Although BIA is capable of providing pertinent information on ligand binding and kinetics, SPR detection is indirect. As a resuh the identity of the bound affinant(s) may not always be certain. This situation can hold particularly true in complex systems where there exists the possibility of binding muhiple, or unknown affmants, either non-specifically or in competition for the surface bound ligand. MALDI-TOF mass spectrometry differentiates between species by detection of analytes at precise mass-to-charge (m/z) values. When coupled with affinity isolation, this direct detection enables the unambiguous determination, or, possible identification, of the retained affinants. Interfacing of BIA with MALDI-TOF thus affords a powerful combination of techniques capable of real-time monitoring of biospecific interactions, and absolute determination of retained analytes. The coupling of BIA with MALDI-TOF mass spectrometry has therefore been explored [3,4,5]. An approach was taken in which BIA analyses were first performed; the retained analytes then mass analyzed directly from the sensor chips (see Fig. 1). II.

Materials and Methods

A.

Biomolecular Interaction Analysis

BIA analyses were performed on a rabbit anti-human IgG/human myoglobin system using a Pharmacia Biosensor BIAcore 2000 (Uppsala, Sweden). Individual flow cells of CMS (carboxylated dextran) sensor chips were derivatized with polyclonal rabbit anti-human IgG using an amine-coupling protocol described previously [6]. Cyano-stabilized human myoglobin (400 ng/mL) in the presence of human serum albumin (20 mg/mL) wasflowed(10 jiL/minute, 20 mM HEPES, 0.005% Tween 20, 150 mM NaCl, 5 mM EDTA, pH 7.4 (HBS)) over the antimyoglobin-derivatized flow cells for times ranging from 30 second to 3 minutes while monitoring the SPR signal. After incubation, the flow cell surfaces were (flow) rinsed with HBS for an additional 3 minutes before the chips were de-blocked from the instrument. Chips were dried and stored at ambient until mass spectrometric analysis. B.

MALDI Mass Spectrometry

Approximately 100 nL of a MALDI matrix, a-cyano-4-hydroxycinnamic acid, (-50 mM dissolved in 1:2, acetonitrile:1.4% TFA) was applied to each of the four flow cells (500|im x 2.0 mm) and allowed to air dry. The chips were next introduced into a prototype MALDI time-offlight mass spectrometer built specifically for analysis of the BIA chips. Briefly, the instrument consists of a linear translation stage/ion source capable of the precise targeting of each of the four flow cells under a focused laser spot (with a spatial resolution on the order of the diameter of the laser spot; ~ 200 |xm). Ions generated during a 4 ns laser pulse (355 nm; Q-switched frequencytripled Nd:YAG) were accelerated to a potential of 25 kV (in a continuous extraction mode) over a single-stage ion extraction source distance of ~ 1 cm before entering a 1.5 m field-free drift region. Ions signals were detected using a 2-stage hybrid (channel plate/discrete dynode) electron multiplier. Time-of-flight spectra were produced by signal averaging the individual spectra from 50 - 100 laser pulses (using a 500 Mhz; 500 MS/sec digital transient recorder). Custom software was used in acquisition and analysis of the mass spectra. Spectra were obtained in the positive ion mode and externally calibrated using equine cytochrome c (MW = 12360.7 Da) as a standard.

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III.

Results/Discussion

Sensorgrams of the antibody immobilization and myoglobin binding are shown in Fig. 2. Fig. 2A shows a sensorgram obtained for one of the flow cell during the amine-coupling of the anti-myoglobin IgG to the surface of the CMS sensor chip. Anti-human IgG (~ 2 mg/mL in HBS) was flow incubated over the chip surface for ~ 7 minutes before a ~ 2 minute rinse with HBS, followed by a ~ 7 minute blocking with ethanolamine. Thefinaldifference in the sensorgram reading of- 15,000 RU translates to ~ 15 ng of antibody covalently linked to the surface of the 1 mm^ area of the flow cell. Considering two binding sites per antibody molecule, a myoglobin binding capacity of 200 fmol is estimated for the flow cell. All four flow cells of the sensor chip were derivatized using identical conditions and resulted in virtually identical sensorgrams (i.e. < 1 % deviation in the amount of antibody bound). Fig. 2B shows sensorgrams obtained during the incubation of the anti-myoglobin-derivatized flow cells with human myoglobin. Sensorgrams for flow cells two and three are shown. A difference in the sensorgram signal of- 250 RU translates to approximately 20 fmoles of myoglobin retained in flow cell 2 (during the 2.5 minute incubation). The sensorgram signal for flow cell 3 indicates roughly half that amount (-10 fmole) retained during the shorter (1 minute) incubation time. Immobilization of anti-human myoglobin on CMS chip

45000 40000

1600-, 1400

Blocking

Human myoglobin bound on CMS chip

1200H S ^

1000 800

CO

§

600

i"

400 200 H 0 -200

100

200

300 400 Time (sec)

Fig. 2 Sensorgrams of CM5/anti-human myoglobin IgG/HSA; myoglobin system. (A) Covalent immobilization of IgG to flow cells. A sensorgram reading of 15,000 RU is indicated corresponding to an antibody binding capacity of- 200 fmol myoglobin. (B) Myoglobin retained by flow cells 2 (FC2), and 3 (FC3). Retention of 20 fmol, and 10 fmol, of myoglobin is indicated for flow cells 2, and 3, respectively.

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Fig. 3 shows the mass spectra obtained from the direct MALDI-TOF analysis of flow cells 2 and 3 of the anti-myoglobin-derivatized/myoglobin-incubated CMS sensor chip. Fig. 3 (lower) was one of ca. 5 mass spectra taken from the area within flow cell 2. Significant signal is observed for both the singly-and doubly-charged ion species of the myoglobin. A measured molecular mass of 17,150 + 15 Da was found for the myoglobin by averaging the centroided mass values of the 5 spectra acquired from flow cell 2. This molecular weight is significantly higher (~ 0.4 %) than that calculated for the mono-derivatized (cyano) myoglobin (MW = 17,080 Da). Considering that the myoglobin ion signals are fairly broad, the shift to higher mass is consistent with the attachment of multiple cyano groups to the myoglobin (creating a heterogeneous sample). Fig. 3 (upper) shows a mass spectrum obtained from within flow cell 3. From the sensorgram it was estimated that - 1 0 fmol of myoglobin was present within the area of the flow cell. Again ion signal is readily observed for the myoglobin. A measured mass of MW = 17160 + 15 Da was determined for the myoglobin using the average of ca. 5 mass spectra taken from within the area of flow cell 3.

FC3 FC2 20000 Fig. 3 BIA/MS of CM5/anti-human myoglobin IgG/HSA; myoglobin system flow cells 2 (FC2) and 3 (FC3). Ion signals are observed in both spectra for the singly- and doubly-charged myoglobin. Retention of species other than the myoglobin is observed in flow cell 3 (marked by *), possibly due to non-specific interactions or the specific retention of myoglobin fragments.

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A few issues of BIA/MS are worth noting. The first is the comparable sensitivities of the two techniques. BIA analyses registering above the ~ 100 RU level are generally considered significant. This sensorgram response translates to ~ 5 fmole of a 20 kDa protein retained over an area of ~ 1 mm^ (the area of a flow cell), an amount generally at the limit of detection of MALDITOF analysis (this is, of course, a general statement as the limits of detection observed during MALDI-TOF are highly dependent on acquisition factors, e.g., matrix and instrument, and the nature of the analyte). Furthermore, the overall sensitivity of the BIA/MS approach reported here (analysis of retained analytes directly from the sensor chip) is not compromised by sample losses associated with eluting the retained affinants and transfer to the mass spectrometer. In fact, there was no actual handling of the samples for mass spectrometry beyond the simple application of matrix solution to the sensor chip surface. While making no claims on the universality of the limits of detection, similar studies with other systems have demonstrated BIA/MS limits of detection of comparable to, or less than, those observed here [3,5]. A second aspect of the BIA/MS analysis is the observance of species in the mass spectra other than those targeted. Fig. 3 (upper) shows the presence of a number of lower molecular weight species retained along with the myoglobin. Blank analyses of flow cells derivatized with antibody and incubated with HBS/HS A buffer (no myoglobin) demonstrated the presence of a number of the lower molecular weight species, however, not all those observed in Fig. 3 (upper). A combination of both nonspecific retention of background species, and specific retention of myoglobin fragments (present in the starting solution) is suggested. Non-specific retention (while of obvious concern) can be compensated for during BIA analysis by blank substraction or saturation of the sensor chip surface. That is to say that the BIA analysis is concerned with the change in response, due to the biospecific interactions defined by the immobilized affinity ligand, after a baseline measurement is established. It is not easy, however, to compensate for the specific binding of non-targeted ligands while simultaneously analyzing for targeted ligands. By direct detection of retained species at defined molecular weights, and incorporation of quantitative methodologies [7,8], MALDI-TOF mass spectrometry has the potential to compensate for such competitive binding. BIOREACTIVE PROBE TIPS IN PROTEIN CHARACTERIZATION L

Introduction

A particular strength of MALDI-TOF mass spectrometry is the ability to analyze complex biological mixtures with little or no prior sample workup. This ability allows for a number of intricate analyses directed at the characterization - from primary to quaternary structure, and post-translational modifications - of proteins. Several such analyses involve the use of enzymes to modify a protein or peptide prior to analysis of the resultant using MALDI-TOF. More often than not, digestions are performed free in solution; a process which allows the possibility of the enzyme autolysis. The resulting autolysis products are recognized in the mass spectrum as interferences and pose a hinderance to the analysis through potential mis-interpretation, or masking of true analyte signals. A way to eliminate such interferences is to covalently immobilize the enzymes to a solid support, the complex then used as the enzymatic reagent. When considering the MALDI-TOF analysis, the support of choice is in fact the mass spectrometer probe device, which, when enzymatically-derivatized, serves to both digest the analyte, and to introduce the digestion mixture into the mass spectrometer [9,10]. There are several advantages to performing digestions using enzymatically-derivatized

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mass spectrometer probe tips. First is an overall increase in sensitivity as sample losses (in transfer and handling) are minimized. Lack of sample loss is critical in maintaining limits of detection throughout the process which are comparable to conventional MALDI analyses (elimination of sample losses is also a contributing factor to the number of proteolytic fragments observed in the mass spectrum during mass mapping). A second advantage is (as stated) the absence of interfering, or background signals due to autolytic digestion of the enzyme. The enzyme is covalently anchored to the probe surface preventing association into the MALDI matrix DSP/isopropanol ISmin.

Fig. 4 General approach of the bioreactive MALDI mass spectrometer probe tips. Gold plated probe tips are activated through the covalent attachment of enzymes (the general terminology of Au/enzyme is used to indicate the nature of the activated surfaces). The probe tips are then used for protein characterization by direct application of the analyte and time given for digestion. The digestions are stopped with the addition of a MALDI matrix, the reaction productmatrix mixture allowed to dry, and the probe tips are inserted into the mass spectrometer for MALDI-TOF analysis.

(negating desorption/ionization), and also prohibiting the freedom necessary for autolysis (which would also produce interferences). Third, digestions can be performed on a time scale equivalent to that required for the MALDI analysis (a few minutes). Covalent anchoring of the enzymes is again largely responsible for the ability to perform digestions rapidly because high effective enzyme concentrations can be used without introducing interferences. Digestion rates can be further increased by using the probe tips at elevated temperatures (accelerating diffusion limited processes and equilibrium kinetics). Lastly, use of enzymatically-derivatized probe devices is

Randall W. Nelson et al

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quite easy, requiring no more steps than those required for a normal MALDI analysis (application of analyte and matrix to the probe). Reported here is the use of bioreactive mass spectrometer probe tips to serially digest myoglobin. The object of the serial digestion was to simultaneously view the relative stability of molecule fragments of myoglobin (generated during an initial, limited digestion of the myoglobin under denaturing conditions using pepsin-active tips at low pH), by exposing the fragment set to extensive digestion (using trypsin tips) under re-naturing conditions. II.

Experimental

A graphic depiction of the experimental process is given in Fig. 4. Stainless steel probe tips were first sputter-coated with ~ 300 nm of gold, and then activated by treatment with dithiobis (succinimidyl propionate) (DSP)/isopropanol solution (for -- 30 minutes). Probe tips were then rinsed vigorously with isopropanol and either used directly (for amine linkage), or further derivatized (for carbodiimide mediated carboxylic acid linkage) by a 15 minute incubation with a solution of ethylene diamine (EDA):isopropanol: triethylamine (40:40:20%). Trypsin was linked through amine coupling by addition of the enzyme (0.1 mg/mL in 20 mM phosphate buffer; pH 8.0) directly to the DSP-derivatized probe tips. Pepsin was linked through carboxylate coupling by addition of the enzyme (0.1 mg/mL in acetate buffer; pH 4.5; 0.1 mg/mL l-ethyl-3-(3dimethylaminopropyl) carbodiimide) to DSP/EDA-derivatized gold tips. Tips were prepared in batches (20 - 40) with the reactions performed in 50 mL conical tubes, generally overnight at ~ 4°C, using volumes of enzyme solution equal to ~ 0.5 mL per probe tip. After incubation the tips were washed with liter volumes of ice-cold incubation buffer, dried, and stored at ambient until needed. For clarity, tips are termed as Au/enzyme to denote the gold surface and linked enzyme. Whale myoglobin (MW = 17,200.4 Da) was dissolved to 0.01 mg/mL (~ 0.6 |iM) in 20 mM ammonium acetate buffer, pH 2.7, and allowed to stand for ~ 30 min. A one minute pepsin digestion was performed by application of 3 |iL of the myoglobin solution directly to the surface of an Au/pepsin probe tip (maintained in a humidified environment at 60 °C). At the same time, 1.5 |aL of a 20 mM phosphate buffer (pH 10) was applied to the surface of an Au/trypsin tip (maintained in high humidity at 60 °C). After one minute, the tips were touched together, effectively transferring a portion of the peptic digest to the Au/trypsin tip (the combination of the two buffers resulted in a solution pH of- 7.5, as verified with pH paper). Immediately following, 1.5 fxL of a a-cyano-4-hydroxycinnamic acid solution (in 1:2; acetonitrile:1.5 % TFA (ACCA)) was applied to the ~ 2 fxL of the digest mixture remaining on the Au/pepsin tip. Trypsin digestion was then allowed to proceed for 5 minutes before termination by addition of 1.5 |LIL of the ACCA matrix. Samples were allowed to air dry prior to insertion of the probes into the mass spectrometer. MALDI-TOF mass spectrometry was performed using a Vestec LaserTec ResearcH linear time-of-flight mass spectrometer (Vestec Corp. Houston, TX), modified to accommodate the probe tips (see Fig. 4), and equipped with a two-stage gridded ion source operating at 30 kV. The rest of the instrument remained unchanged from that described previously [11]. Mass spectra were acquired in the positive ion mode with each spectrum the sum of 50 - 100 individual laser desorption/ionization events. Spectra were externally calibrated using horse heart cytochrome c (MW 12,360.7 Da) as a standard. Mass data was analyzed using protein analytical worksheet software (PAWS) [12].

Biomolecular Interaction Analysis with MS and MS Probe Tips

III.

501

Results/Discussion

A combination of enzymatically-active probe tips was used to investigate the regional stability of myoglobin. A set of molecular fragments representing different regions of the protein was first prepared by a limited pepsin digestion of the protein under denaturing conditions (pH ~ 3). The set was then exposed to further, more extensive, degradation (using trypsin) under native conditions (pH ~ 8). Regions of myoglobin that do not exhibit an intrinsic steric shielding by the tertiary structure of the molecule (the molecule being either the intact myoglobin or one of the fragments) are more susceptible to digestion by the trypsin, and therefore, signals representing these fragments are expected to be attenuated in the final mass spectrum. Regions of the myoglobin possessing a tighter tertiary structure (when folded under native conditions) will exhibit a higher degree of immunity to the trypsin digestion, and representative signals in the mass spectrum will be attenuated to a lesser extent.

c 0) 4

.> ''•3

JS

«

2H

20000

Fig. 5 One minute Au/pepsin digestion of whale myoglobin under denaturing conditions (pH 3, 60 °C) (A, grey). Peptic fragments digested for 5 minutes using an Au/trypsin probe tip (pH ~ 8,60 °C) (B). Select fragments have been completely digest indicating a relatively low degree of steric hinderance (to tryptic sites) in the final 46 residues of the protein. Ion signals are marked with residue numbers. Region indicated is shown in Fig. 7.

502

Randall W.Nelson era/. Residue Number N

20

40

60

80

100

120

140

^

^^^^"" " 30 153

110.153 • 13«-153

N

20

40

60

Residue Number 100 80

120

140

R

„

^

N

20

40

60

"

•

^

^

^

Residue Number 80 100

^

120

140

c • 13«-153

Fig. 6 Coverage maps derived from the Au/pepsin-Au/trypsin serial digestion of whale myoglobin. (A) Au/pepsin digest fragments. (B) Peptic fragments exhibiting a relatively high immunity to tryptic digestion. (C) Peptic fragments eliminated during trypsin digestion. Residue numbers are as indicated.

Fig. 5A shows the resuhs of myoglobin digested under denaturing conditions using an Au/pepsin tip. Strong ion signals representing fragments due to cleavage of the myoglobin at five sites, residues 29, 69, 106, 109, and, 137, are observed. All peptic fragments contain between one and nine trypsin cleavage sites. Upon exposure to an Au/trypsin tip (Fig. 5B), select fragments in the peptic mixture are observed to undergo complete digestion, whereas others exhibit a relative immunity to digestion. Fig. 6 shows mass coverage maps of the fragments from the pepsin digestion, the fragments surviving the Au/trypsin digestion, and those completely digested by the trypsin. In general, minimal stearic shielding of tryptic sites is observed in fragments comprised of the final two helices of the myoglobin. Fig. 7 shows an evolution of tryptic fragments derived from the original pepsin digest. Signals consistent with cleavage at three of the six possible trypsin sites present in the 107 - 153 region of the myoglobin are observed. There are no other strong ion signals in the Au/pepsin; Au/trypsin spectrum due to both pepsin and trypsin digestion of the myoglobin (other signals in the spectrum are consistent with cleavage at trypsin sites — confirmed by Au/trypsin digestion of myoglobin). This observation, and the survival of numerous fragments containing residues 1 - 106, is consistent with the steric inaccessibility of sites within the central, heme-coordinated region of the molecule (independent of the final 46 residues of the molecule).

503

Biomolecular Interaction Analysis with MS and MS Probe Tips

C 0)

I

0)

1000

2000

3000

4000

5000

6000

m/z

Fig. 7 Mass spectra showing the evolution of proteolytic fragments generated by the successive Au/pepsin Au/trypsin digestion of myoglobin. Ion signals representing peptic fragments of the myoglobin originating between residues 107-153 (A) are observed to undergo complete tryptic digestion (B) indicating relatively free access to trypsin cleavage sites. Ion signals are marked to indicate proteolytic fragments (by residue).

Obviously, more data is needed in order to make any broader statements on the relative degree of m^ra-molecular interaction of the myoglobin. However, serial digestions are possible in numerous combinations, and quite easy to perform using the bioreactive probe tips. Further, digestion of the myoglobin in the presence of denaturants (detergents, salts) is also possible to study the relative accessibility to proteolytic sites, yielding additional information on the overall structure of the molecule [13]. Finally, incorporation of quantitative MALDI-TOF techniques allows the tracking of digestions as a function of time, providing even further insight into the dynamics of digestion {i.e., determination of fragment pre-cursors and final products) and molecular stability [13, 14]. Currently, we are exploring such uses of the bioreactive probe tips, in combination with the defined and accurate mass spectrometric identification of proteolytic fragments, in the study of higher-order protein structure.

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FINAL REMARKS The rapid advancement of analytical technologies such as SPR-based biomolecular interaction analysis (BIA), and MALDI-TOF mass spectrometry, has allowed the routine characterization of biomolecules present in complex environments at physiological concentrations. Presented here has been the coupling of the two orthogonal techniques into a combined approach capable of observing real-time, solution-phase biospecific interactions (using BIA), and the rapid qualitative assessment of binding partners (using MALDI-TOF). The combined analysis is performed without compromise of the speed, sensitivity, or, accuracy of the constituent techniques, and therefore demonstrates the inception of a new bioanalytical approach: Biomolecular Interaction Analysis Mass Spectrometry (BIA/MS). An additional approach to biomolecular analysis, hioreactive mass spectrometry probe tips, has also been given. These are conceptually, and practically simple devices constructed to analytically modify biomolecules prior to mass spectrometric analysis. The bioreactive devices have proven quite convenient in use, and often necessary in maintaining high speed, sensitivity, and, accuracy in the mass spectrometric analysis of proteolytic mixtures. An obvious next step is the combination of the two techniques, BIA/MS with bioreactive mass spectrometry probe tips. Such an approach would thereby allow (all on a single surface), the real-time observance of affinity interaction followed by enzymatic modification and mass spectrometric characterization of retained ligands. REFERENCES 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14.

Szabo, A., Stoltz, L., and Granzow, R. (1995) Curr. Opinion Struc. Biol 5, 699- 705. Karlsson, R., Roos, H., Fargerstam, Persson, B., (1994) Methods: A Companion to Methods in Enzymology, 6 , 9 9 - 110. Krone, J.R., Nelson, R.W., Dogruel, D., Granzow, R., Williams, P., in Proceedings of the 5th Annual European BIAsymposium, Stockholm, Sweden, September 27-29, 1995, Ed. R. Millett. Page 173 - 179. Krone, J.R., Nelson, R.W., Dogruel, D., Williams, P., Granzow, R., (1996) BIAjournal 3, 16 - 17. Krone, J.R., Nelson, R.W., Dogruel, D., Williams, P., Granzow, R., (1996) Anal. Biochem. In press. BIAapplications Handbook (1994). Chapter 4. Nelson, R.W., McLean, M.A., Hutchens, T.W., (1994) Anal. Chem. 66, 1408 - 1415. Nelson, R.W., Krone, J.R., Bieber, A.L., Williams, P., (1995) Anal.Chem. 67, 1153 -1158. Dogruel, D., Williams, P., Nelson, R.W., (1995) Anal. Chem. 67, 4343 - 4348. Nelson, R.W., Dogruel, D., Krone, J.R., Williams, P., (1995) Rapid. Comm. Mass Spectrom. 9, 1380 1385. Vestec LaserTec ResearcH specification sheet, Vestec Corporation, Houston, TX, (1992). Beavis, R.C. Protein Analysis Worksheet Version 6.1.1, (1995). Patterson, D.H., Tarr, G.E., Hines, W.M., Vestal, M.L.,Proceeding of The 44*^ ASMS Conference on Mass Spectrometry and Allied Topics, Portland, Oregon, May 1996. In press. Lewis, J.K., Krone, J.R., Dogruel, D., Williams, P., Nelson, R.W., Proceeding of The 44* ASMS Conference on Mass Spectrometry and Allied Topics, Portland, Oregon, May 1996. In press.

Transition-State Theory and Secondary Forces in Antigen-Antibody Complexes Mark E. Mummert and Edward W. Voss, Jr. Dept. of Microbiology, University of Illinois at Urbana-Champaign, Urbana, IL 61801

I. Introduction Secondary forces, defined as those interactions exhibited outside of the classically defined antibody active site, have been demonstrated to modulate the conformation and free energy of binding of antifluorescein antibodies (1-3). Figure 1 defines and distinguishes primary from secondary interactive components. The ability of the epitopic environment to influence antibody binding has obvious immunological ramifications. Dissection of those interactions that influence the overall dynamic and thermodynamics of a given protein system is of general importance in understanding interfacial protein chemistry. The antifluorescein system is advantageous for evaluating and quantitating interfacial chemistry. Binding of fluorescein ligand in the antifluorescein active site results in bathochromic shifts of the ligand's absorption spectrum and a decrease in both the fluorescence quantum yield and lifetime. These properties allow sensitive spectral and kinetic measurements to be made (4). Changes in the spectral and kinetic properties of a given antifluorescein antibody upon interacting with fluorescein attached to a carrier molecule compared to fluorescein (devoid of carrier residues) thus provides important information about secondary force directed perturbations. Placement of the fluorescein moiety in various environments is easily achieved due to the availability of the highly reactive isothiocyanate derivative of fluorescein. Evaluations of secondary interactive components have been discussed (5-8). In general, the delineation between primary and secondary interactive components have been vague (9). An important advantage of the fluorescein system is that the ligand fills the active site (10-12) which has been conclusively demonstrated by X-ray crystallographic results for the monoclonal antifluorescein antibody (mAb) 4-4-20 (13-15). Thus, interactions with carrier residues associated with the ligand-carrier complex are by necessity outside of the primary interactions. An understanding of interfacial protein chemistry requires evaluation of the thermodynamics of the system under investigation as well as the energetic barriers responsible for the observed kinetics and affinity. Due to the kinetic methodology available for the antifluorescein system, the energetic barriers for complex decomposition TECHNIQUES IN PROTEIN CHEMISTRY VIII Copyright © 1997 by Academic Press All rights of reproduction in any form reserved.

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Mark E. Mummert and Edward W. Voss, Jr. Antibody variable domains

Carrier environment (highly charged protein or lipid membrane)

Figure 1. Schematic representation differentiating primary and secondary interactions. Secondary interactions are the result of interactions between regions surrounding the mouth of the active site and regions of the carrier environment surrounding the ligand. A highly charged protein or lipid interface represents an example of substrate exerting secondary effects.

can be evaluated (Figure 2). It is important to note that the kinetic measurements can be conducted in solution under near physiological conditions. Thus, the results obtained can be extrapolated to biological situations. In this report, we summarize the results of a study in which the energetic barriers of several protein/complex decompositions were analyzed utilizing transition-state theory. In essence, fluorescein 5-isothiocyanate was covalently linked to a variety of synthetic peptides and allowed to bind with the well defined high affinity 4-4-20 mAb. Differences in the rates of decomposition were measured at 275 K and 291 K and the height of energetic barriers calculated using classical transition-state analysis (16).

n. Methods and Materials A. Monoclonal anti-fluorescein antibody 4-4-20 mAb 4-4-20 was produced in ascitic fluid from pristane treated Balb/c mice and affinity purified using a fluorescein Sepharose 4B adsorbent (17,18). B. Peptide synthesis for use as carriers Peptides of different chemical composition acetylated in the amino-terminal position were synthesized using an Applied Biosystem model 430A peptide synthesizer at the University of Illinois Genetic Engineering Facility (Urbana, DL) employing solid-phase F-moc chemistry with standard amino acid protecting groups. The generic peptide design was as follows: Ac-NH-(X)6-K-(X)6-COOwhere Ac-NH denotes the acetylated a-amino group, X represents glutamate or arginine, and K is the central lysine residue available for FITC (I) derivatization. Peptides were desalted and purity verified by RP-HPLC. Purified peptides were analyzed by mass spectrometry to verify composition.

Transition-State Theory and Secondary Forces in Ag-AB Complexes

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Second Transition Stat*

Reaction Coordinate

Figure 2. Two dimensional reaction coordinate depicting the interaction of mAb 4-4-20 with homologous ligand. The x-axis is arbitrarily assigned reaction progression while the y-axis is the chemical potential. The height of the chemical potential barriers dictates the rate of the reaction. The encounter complex was included based on kinetic considerations (19). Monofluoresceinated peptides were synthesized by adding an equimolar concentration of FITC(I) to peptides. The reaction was adjusted to a pH of 10.3 with K2CO3 and incubated at ambient temperature overnight. The resulting reaction mixture was resolved over a P-2 column (Bio-Rad) equilibrated in 0.1 M phosphate, pH 8.0 to remove unreacted fluorescein from the peptides. Fluorescently labeled peptides were analyzed by thin layer chromatography with water saturated methyl ethyl ketone as the solvent system. C. Determination of unimolecular rate constants Ligand dissociation rates were determined at 275 K and 291 K utilizing the methodology and analysis as described in detail by (19). This technique provides an essentially unidirectional displacement of the fluorescein/antibody complex. D. Calculation of transition-state thermodynamic parameters All calculations have been described in detail elsewhere (3). Transition-state equations can be found in most elementary physical chemistry texts or in the classical work of Wynne-Jones and Eyring (16). III. Results A. Monofluoresceinated peptides Thin layer chromatographic analyses of monofluoresceinated peptides indicated a single fluorescent band for each of the labeled peptides. RF values were 0.90, 0.85, 0.83 and 0.76 for FDS, D12KF1, R6D6KF1 and R12KF1 respectively.

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Mark E. Mummert and Edward W. Voss, Jr.

Table L Comparative unimolecular rate constants at 275 K and 291 K for the interaction of FDS and monofluoresceinated peptides with mAb 4-4-20 Ligand

k.i^

k.^i,

^Ab^Asi

FDS

1.63(±0.02)xl0-4

1.92(iO.09)xl0-3

11.8

D12KF1

3.52(±O.62)xl0-3

1.06(±0.19)xl0-l

30.1

R6D6KF1

6.96(±1.02)xl0-3

I.15(dt0.42)xl0-1

16.5

R12KR

6.79(±0.25)xlO-3

6.08(±0.81)xl0"2

8.9

k.ia = unimolecular rate constant at 275 K k_n, = unimolecular rate constant at 291 K

B. Affinity of mAb 4-4-20 with various ligands In previous studies (2), the affinity constants (Ka) for the interaction of mAb 4-420 with fluorescein and monofluoresceinated peptides were measured at 275 K. The affinities of mAb 4-4-20 for FDS, D12KH, R12KF1 and R6D6KF1 were 3.14x10^° M'\ 1.49x10^ M"^ 7.49x10^ M^^ and 7.55x10^ M \ respectively. C. Unimolecular rate constants Unimolecular rate constants for decay of the mAb 4-4-20/fluorescein complex and mAb 4-4-20/monofluoresceinated peptide complexes were determined at 275 K and 291 K. The 16 K differential resulted in significant changes in the individual decay rates of the various complexes. The largest change with temperature was with the mAb 4-420/D12KF1 complex (30.1-fold), while the smallest change was with R12KF1 (8.9-fold). Importantly, the R6D6KF1 ligand resulted in an approximate average (16.5 -fold) of the poly anionic (D12KF1) and polycationic (R12KF1) environments. Table 1 summarizes these results. D. Relationship between enthalpy and entropy Table 2 summarizes the calculated transition state thermodynamic parameters (AH", AS" and AG"). The secondary effects that resulted from the carrier molecule caused an apparent enhancement in AH" and AS" relative to fluorescein devoid of carrier residues. The enhanced values of AS" offset the enhanced AH" with the net effect of lowering the overall energetic barriers (AG") of the 4-4-20/monofluoresceinated complexes relative to the 4-4-20/fluorescein complex (Table 3).

Transition-State Theory and Secondary Forces in A g - A B Complexes

509

Table IL Comparative thermodynamic transition-state parameters and transition-state equilibria for the interaction of mAb 4-4-20 with FDS and monofluoresceinated peptides at 275 K Ligand

AH^

AS^

AC"

K^

FDS

+23.96±0.06

+0.0110.00

+20.8210.07

2.84x10-^7

D12Kn

+33.28+1.95

+0.0510.00

+19.1511.95

6.03x10-^6

R6D6KF1

+27.3213.36

+0.0310.00

+18.7713.36

1.21x10-1^

R12KF1

N.A.

N.A.

N.A.

N.A.

AH^ = transition-state enthalpy (kcal/mol) AS"^ = transition-state entropy (kcal/mol/K) K"^ = transition-state equilibrium (dimensionless) N.A. = not applicable; does not conform to the theoretical assumptions of transition-state theory

E. K values Values for the transmission coefficient (K) at 275 K were 1.00, 1.02, 1.00 and 0.58 for FDS, D12KF1, R6D6KF1 and R12KF1, respectively. Transition-state theory assumes unity for K. Deviations of K from unity indicated poor approximation of the various transition-state thermodynamic parameters. Thus all complex decays were adequately described by transition-state theory, except for the R12KF1 peptide.

IV. Discussion Understanding those components that influence the interfacial binding properties in protein/protein and protein/ligand interactions is of basic importance in protein chemistry. In this report, we have defined a system that should allow the dissection of those chemical properties that influence primary interactions via an evaluation of the transition-state thermodynamic components. It is important to realize fundamental assumptions made in the calculations. At the temperatures utilized in these experiments (275 K and 291 K), it was assumed that complexes moved over energetic barriers with standard Arrhenius motion. Deviations from Ahrrenius motion (e.g., tunneling) usually result as a consequence of low temperature (20-22). It is also important to realize that the values calculated for AH^, AS"" and /SG^ are the upper limits of the system, since solvent was considered as a part of the system (23). This study suggested that secondary forces of the mAb 4-4-20 /monofluoresceinated peptide complexes modulated binding interactions via increased transition-state enthalpic and entropic contributions. The net result was a decreased energetic barrier that allowed modulation of the previously reported affinity constants of mAb 4-4-20 for the monofluoresceinated peptides due to variation of the unimolecular rate constant (2).

510

Mark E. Mummert and Edward W. Voss, Jr.

Table i n . Comparative differences in thermcxlynaniic transition-state parameters of monofluoresceinated peptides with respect to FDS at 275 K

Ligand

AAH^

AAS''

AAC

D12KF1

+9.32±1.95

•K).04±0.00

-1.67±1.95

R6D6KF1

+3.36±3.36

0.02±0.00

-2.05±3.36

R12KF1

N.A.

N.A.

N.A.

AAIT^ = change in transition-state enthalpy with respect to FDS (kcal/mol) AAS'^ = change in transition-state entropy with respect to FDS (kcal/mol/K) AAG^ = change in transition-state free energy with respect to FDS (kcal/mol) N.A. = not applicable

Increased values of AH" and AS" for the mAb 4-4-20/monofluoresceinated peptide complexes relative to the mAb 4-4-20/fluorescein complex decay were interpreted as resulting from inclusion of the carrier peptides. Increased enthalpic contributions may have resulted from actual binding interactions between the surface accessible complementarity determining regions (CDRs) surrounding the mouth of the antibody active site and the amino acids of the peptides. Whitlow et al. (15) reported that a significant percentage of the amino acids that compose the mAb 4-4-20 CDRs were solvent accessible when fluorescein was in the active site. The increased values for AH" also may have been due to differences in hydration of the antibody complexes. Enhanced AS" values for the antibody/peptide complexes may have been a result of the greater rotational, translational and vibrational degrees of freedom as the complexes decayed relative to the mAb 4-4-20/fluorescein complex. As in the AH" argument, hydration may also be an important factor to consider. Hydration has been shown to significantly influence the free energy of binding (14). We interpreted the inability of transition-state theory to predict the decay of the mAb 4-4-20/R12KF1 complex to be a result of differential conformational changes. Deviations of K from unity are a direct result of the inertial (solvent coupling) and diffusive (intramolecular dynamic) regimes (24-27). The frictional coefficient in both of these regimes dictates the value of K (24,25). Both inertial and diffusive regimes modulate K in proteins (27-29). We therefore proposed that the mAb 4-4-20/R12KF1 complex could not be evaluated by transition-state theory due to inertial and/or diffusive regimes. We conceived that the secondary forces dictated by R12KF1 resulted in greater perturbation of the antibody variable domains than the secondary forces dictated by either D12KF1 or R6D6KF1. It was postulated that the greater van der Waals volume for arginine (R~148 A^) as opposed to aspartic acid (D~91 A^) resulted in greater variable domain atomic coordinate displacement and thus enhanced frictional components. In conclusion, the antifluorescein system provides a reasonable model with which to evaluate interfacial interactions utilizing transition-state theory. Evaluations like those presented herein provide means to develop mechanistic models to describe interfacial interaction from an energetic barrier viewpoint.

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511

References Mummert, M.E. and Voss, E.W., Jr. (1995). Mol Immunol 32, 1225-1233. Mummert, M.E. and Voss, E.W., Jr. (1996) Mol Immunol in press. Mummert, M.E. and Voss, E.W., Jr. (1996) Biochemistry 35, 8187-8192. Voss, E.W., Jr. (1993) 7. Mol Recog. 6, 51-58. vanOss, C.J. and Absolom, D.R. (1984) In "Molecular Immunology" (Atassi, M.Z., vanOss,C.J. and Absolom, D.R., eds.) pp. 337-360. Marcel Dekker, New York. 6. vanOss, C.J., Good, R.J. and Chaudhuny, M.K. (1986) 7. Chromatog. 376, 111-119. 7. vanOss, C.J. (1992) In "Structure of Antigens" (Van Regenmortel, M.H.V., ed.) vol. 1, pp. 179-208. CRC Press, Inc., Boca Raton, FL. 8. vanOss, C.J. (1994) In "Immunochemistry" (vanOss, C.J. and Van Regenmortel, M.H.V., eds.) pp. 581-613, Marcel Dekker, New York. 9. vanOss, C.J. (1995) Mol Immunol 32, 199-211. 10. Voss, E.W., Jr., Eschenfeldt, W. and Root, R.T. (1976) Immunochemistry 12, 745749. 11. Omelyanenko, V.G., Jiskoot, W. and Herron, J.N. (1993) Biochemistry 32, 1042310429. 12. Carrero, J. and Voss, E.W., Jr. (1996) 7. Biol Chem. Ill, 5332-5337. 13. Herron, J.N., He, X-m., Mason, M.L., Voss, E.W., Jr. and Edmundson, A.B. (1989) Proteins: Struct., Funct., Genet. 5, 271-280. 14. Herron, J.N., Terry, A.H., Johnson, S., He, X-m., Gudday, L.W., Voss, E.W., Jr. and Edmundson, A.B. (1994) Biophys. 7. 67, 2167-2183. 15. Whitlow, M., Howard, A.J., Wood, J.F., Voss, E.W., Jr. and Hardman, K.D. (1995) Prot.Eng.^,lA9-16\. 16. Wynne-Jones, W.F.K. and Eyring, H. (1935) 7. Chem. Phys. 3, 492-502. 17. Kranz, D.M. and Voss, E.W., Jr. (1981) 7. Biol Chem. 257, 6987-6995. 18. Weidner, K.M., Denzin, L.K., Kim, M.L., Mallender, W.D., Miklasz, S.D. and Voss, E.W., Jr. (1993) Mol Immunol 30, 1003-1011. 19. Herron, J.N. (1984) In "Fluorescein Hapten: An Immunological Probe" (Voss, E.W., Jr., ed.) pp. 50-75. CRC Press, Inc., Boca Raton, FL. 20. Frauenfelder, H., Nienhaus, G.U. and Johnson, J.B. (1991) Ber. Bunsenges. Phys. Chem. 95, 272-278. 21. Wolynes, P. (1987) In "Protein Structure: Molecular and Electronic Reactivity" (Austin, R., Buhks, E., Chance, B., DeVault, D., Dutton, P.L., Frauenfelder, H. and Gol'daskii, V.I., eds) pp. 201-209, Springer-Verlag, Inc., New York. 22. Frauenfelder, H. (1979) In "Tunneling in Biological Systems" (Chance, B., DeVault, D.C., Frauenfelder, H., Marcus, R.A., Schriefer, J.R. and Sutin, N., eds.) pp. 627649. Academic Press, Inc., New York. 23. Beece, D., Eisenstein, L., Frauenfelder, H., Good, D., Marden,M.C., Reinisch, L., Reynolds, A.H., Sorensen, L.B. and Yue, K.T. (1980) Biochemistry 19, 51575157. 24. Chandler, D. (1978) 7 Chem. Phys. 68, 2959-2970. 25. Northrup, S. and Hynes, J.T. (1978) 7 Chem. Phys. 69, 5246-5260. 26. Hasha, D.L., Eguchi, T. and Jonas, J. (1982) 7. Am. Chem. Soc. 104, 2290-2297.

1. 2. 3. 4. 5.

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27. Doster, W. (1983) Biophys. Chem. 17, 97-103. 28. Karplus, M.A. and McCammon, J.A. (1981) FEES Lett. 131, 34-36. 29. McCammon, J.A. and Karplus, M. (1979) Proc. Natl Acad. Sci. U.S.A. 76, 35853589.

Thermodynamic Investigation of Enzyme and Inhibitor Interactions with High Affinityi Yudu Cheng, Jacek Slon-Usakiewicz, Jing Wang, Enrico O. Purisima and Yasuo Konishi National Research Council of Canada, Biotechnology Research Institute Montreal, Quebec, Canada

I. Introduction The enzyme and inhibitor binding interactions may be elucidated by the thermodynamic functions such as the free energy (AG), enthalpy (AH), entropy(TAS) and heat capacity(ACp). These thermodynamic functions are related through the following equation: AG°(T) = AH°(T) - TAS°(T) = [AH°(T°) - TAS°(T°)] + ACp[(T - T°) - Tin (T/T°)]

(1)

In the above equation, AG°, AH°, AS° and ACp are the thermodynamic functions relative to a standard state(1.0 mol/L for all chemical species and 25 °C), T° is the reference temperature (298.15 K in this work). The thermodynamic study plays a major role in accessing the molecular basis of enzyme and inhibitor interactions because the thermodynamic functions convey extensive information from the binding affinity to the conformational change. In general, AG is the affinity between enzyme and inhibitor. AH is the binding energy arisen from the van der Waals interactions, hydrogen bonding interaction, dehydration and other effects (e.g. deprotonation, ion-bridge etc.). AS measures the loss or gain in the rotational, translational and/or vibrational degrees of freedom in the conformational change and consists of both solvent and conformational contributions. ACp measures the temperature dependence of AH and AS. ACp may also be temperature dependent. In eq 1, ACp is assumed to be temperature independent for simplicity. We have conducted thermodynamic studies on the thrombin and its bivalent inhibitors' interactions in which the binding affinity ranges from Kj = 10-9 to Kj = 10-12 M (Ki is the inhibition constant)(l,2). Thrombin is a key enzyme regulating thrombosis in cardiovascular disease. The synthetic bivalent thrombin inhibitors possess an active-site binding segment, a linker and a fibrinogen recognition exosite (FRE) binding segment which is based on the C-terminal sequence of hirudin, AspH55-PheH56-GluH57_GluH58_IleH59_ProH60_GluH61-GluH62 -TyrH63_LeuH64_

1NRC publication No. 39931 TECHNIQUES IN PROTEIN CHEMISTRY VIII Copyright © Government of Canada All rights of reproduction in any form reserved.

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Yudu Cheng et al

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GlnH65-OH(HirudinH55-H65^ H stands for hirudin). Hirudin is a 65 amino acid protein and naturally occurring thrombin inhibitor with a Ki value of 2.2 x 10-14 M(3). The crystal structure of thrombin-hirudin complex(4) indicates that, besides those distinct electrostatic interactions of hirudin and thrombin, the complementary fit of the nonpolar residues seems to be of particular importance. The site-directed mutagenesis(5) and Gly substitution(6) studies of five nonpolar residues PheH56^ IleH59^ ProH60, TyrH63 and LeuH64 in the FRE binding segment showed that the residues PheH56 and IleH59 are crucial to the binding at the FRE. In order to understand the molecular details of the nonpolar residue and thrombin interactions at the FRE, the complete thermodynamic profiles (AG°, AH°, TAS° and ACp) of the analogs, in which the five nonpolar residues PheH56, IleH59, ProH60, TyrH63 and LeuH64 of a thrombin inhibitor P552(7) are consecutively substituted by Gly, were measured and analyzed in conjunction with the structural features obtained from the molecular modelling for these substitutions. The results show that the change in the binding free energy (AAG°) due to the Gly substitution has a linear correlation with the change in the molecular surface area (AAA) around the Gly substitution site, evidencing the structural basis of the free energy. Meanwhile, the components of AAG°, AAH° and TAAS°, appear no correlation with AAA because of the linear compensation of these two quantities, but are specific to the conformational effects (e.g the movement of the inhibitor's backbone and neighboring water molecules) due to the Gly substitution. In this article, we describe the technique procedures employed by us to measure and analyze the thermodynamic functions in eq 1 for the system of thrombin and inhibitor interactions.

II. Experimental method A. Materials Human a-thrombin and the fluorogenic substrate (Tos-Gly-Pro-Arg-AMCHCl) were purchased from Sigma. Fmoc derivatives of amino acids were purchased from Advanced ChemTech and Novabiochem. N-a-Fmoc-N-y-trityl-L-Gln-Wang resin was purchased from Applied Biosystem Inc. The solvents used in peptide synthesis were obtained from Anachemia Chemical Inc. and Applied Biosystems Inc. B, Peptide synthesis and

purification

The thrombin inhibitors are synthesized on a 396 Multiple Peptide Synthesizer (Advanced ChemTech) by using a conventional Fmoc strategy of solid phase peptide synthesis. Double couplings are performed throughout the synthesis. The peptides are purified on a preparative HPLC using a linear gradient of 20 to 50% acetonitrile in 0.1% TEA ( 0.5%/min gradient, 33 mL/min flow rate). The purified products with >98% purity estimated by an analytical HPLC are lyophilized. The final peptides are identified using a Beckman 6300 amino acid analyzer and a

Thermodynamics of Enzyme and Inhibitor Interactions

515

SCIEX API III mass spectrometer. C

Enzymatic

assay

The inhibition of the amidolytic activity of human a-thrombin is measured using Tos-Gly-Pro-Arg-AMC as a fluorogenic substrate in 50 mM Tris HCl buffer (pH 7.80) containing 0.1 M NaCl and 0.1% poly(ethylene glycol) 8000 at various temperatures from 10 to 45 °C. Human a-thrombin is stable in this temperature range in the presence of poly(ethylene glycol)(8). The temperature dependence of Km and V^ax is measured at 1—40 |iM and 30 pM of the substrate and thrombin, respectively. The Ki is measured under various temperatures at 40|LiM, 30pM and varying concentrations (0.3 - 100-fold of the Ki values in the temperature range of 10-45°C) of the substrate, thrombin and inhibitor, respectively. The steady-state velocity of the enzyme reaction is measured under the condition of XQ^ =383 nm and ?Lem =455 nm in a Hitachi F2000 spectrophotometer and a Perkin Elmer LS50B luminescence spectrometer. The running solutions is preincubated at the temperature of assay for 15 min. The temperature is controlled and monitored by using a HAAK^E Circulator and a YSI Series 400 Probe (±0.1 °C). The reaction starts with adding thrombin and the progressive curve is traced for 5-15 min. D, Molecular

modelling

Energy minimization. The crystal structure of the thrombin-P500 complex(9) is used as the starting structures for molecular modelling. P500 is a bivalent thrombin inhibitor with the sequence of dansyl-Arg-(D-Pip)-)LlAdod-Gly-HirudinH55-H65. Because the residues interested (PheH56^ IleH59, ProH60^ TyrH63 and LeuH64) locate only in the PRE binding segment, the bound state of the inhibitors is modelled using the sequence of Ac-HirudinH55-H65-NHCH3. The complex which includes the ERE segment Ac-HirudinH55-H65-NHCH3 and thrombin residues and water molecules within 6 A from any atom of the ERE segment is re-energy-minimized. This refined structure is the starting point for the structure modelling of the thrombin and analogous inhibitor complexation. The energy minimization for the analogs is conducted only for the atoms in the static substructure set which includes the residues within 4 A from the residue substituted. The AMBER force-field(lO) as implemented in SYBYL 6.1 (Tripos Inc.) is used with a non-bonded cutoff of 8 A, a dielectric constant of 80 and a gradient convergence tolerance of 0.005 kcal/molA. Conformational search, Monte Carlo sampling and energy minimization. In the case of the Gly substitution for the residue PheH56^ the considerably different thermodynamic profiles (small decrease in AH°, but large decrease in T°AS° and ACp) from other P552 analogs were observed, an alternative procedure, which includes the systematic conformational search, Monte Carlo sampling and energy-

Yudu Cheng et al

516

minimization(2), is applied for further study. The rotatable bonds of the backbone and the side chains are varied in 15 degree increment for (PheH56 or GlyH56) and GluH57 and 30 and 45 degree increment for AspH55 in the complex with thrombin in order to generate a database of sterically feasible conformations. The water molecules are not included in the conformational search but are included in the energy minimization. The conformers in the database are sampled and energyminimized. This is then followed by a clustering step where all of the energyminimized conformers are grouped into several clusters. Each cluster contains the conformers with similar energies and structures. The further energy minimization is conducted only for the conformer with the lowest energy in each cluster. Molecular surface area calculation. The molecular surface area is an envelope of a molecule from which the solvent is excluded(ll). The molecular surface area is estimated using the GEPOL algorithm(12) with the van der Waals radii used in the AMBER force-field(lO) and a solvent probe radius of 1.4 A. The polar molecular surface area is composed of oxygen, nitrogen and polar hydrogens (e.g., NH and OH), and nonpolar molecular surface area is composed of all other atoms. The molecular surface area of the bound state is calculated using the energy-minimized complex structures.The molecular surface area of the free state is calculated using the geometry of a tripeptide Gly-Xaa-Gly, where Xaa stands for the residues studied. The backbone conformation of the tripeptide is set as \|/ = 140° and (j) =140°. The side-chain conformations and their populations are determined based on the statistical survey of the side-chain conformations in 100 refined protein structures(13). The change in the molecular surface area(AAA) is estimated by using a thermodynamic circle shown in Scheme I, where E, I and V stand for the free enzyme, inhibitor and analog, respectively, and EI and EI' stand for the complex of thrombin and wild-type and mutated inhibitors, respectively. The circle satisfies that AAX = AX2 - AX] = AX4 - AX3 and enables that the relative thermodynamic function measured (AAG°, AAH°, T°AAS°) can be analyzed by using the structural properties predicted (conformations and AAA).

Scheme II

Scheme I E+I

AX| ^^

^1 AX3

AX4

E + r ^;;:

AX2

M

EI

Er

+

^2

+

^3

E

r

P + E (Ila)

ES ^5

EI - ^ r ^ EI k6

(Hb)

Thermodynamics of Enzyme and Inhibitor Interactions

517

III. Data analysis A. Kinetic data transformation For the system studied, the reaction between enzyme and substrate, enzyme and inhibitor may be described by Scheme 11. Conforming to Scheme Ila which represents the reaction of enzyme and substrate in the absence of inhibitor, the Michaehs constant (Km) and maximal velocity (Vmax) are given by Kn, = [E][S]/[E-S] =(k2+kp)/ki

(2)

Vn,ax = kp[E]

(3)

and

respectively. [E], [S] and [ES] are the concentrations of enzyme, substrate and enzyme-substrate complex, respectively. The enzymatic parameters, K^ and Vmax. are estimated at each temperature by using the equation: V = Vmax[S]/(Km+[S])

(4)

where v is the velocity of the enzyme and substrate reaction. Conforming to the Scheme lib which represents a slow-binding inhibition, the progressive curves of the enzymatic assay in the presence of a competitive inhibitor are analyzed using the following equation(14): P = Vst + (vo-Vs)(l-e-kt)/k

(5)

where P is the fluorescence intensity, Vs is the steady-state velocity, t is time, VQ is the initial velocity and k is a parameter relevant to the kinetic mechanism(15). The variation of steady-state velocity (Vs) with inhibitor concentration ([I]) obtained by using eq 5 is then used to determine the inhibition constant (Kj) through the following equation(16) Vs=V„ax[S]/{K„(l+[l]/Ki)+[S]}+Ve

(6)

where Ki = k4k6/k3(k5+k6) and represents the overall inhibition constant and Vc is a parameter used to account the deviation from the linearity (Vc > 0). Temperature dependence ofK^ and Vmax- Since both Km and Vmax are encountered in the calculation of the inhibition constant (Ki) at various temperatures, the temperature dependence of Km and Vmax should be a priori determined. Figure 1 shows the temperature dependence of Michaelis constant (Km)

Yudu Cheng et al

518

and maximal velocity (Vmax) in the range of 10-45 °C. The temperature dependence of Km and Vmax is analyzed using van't Hoff equation: InKm - InKd = AG°(T)/RT = [AH°(T) - TAS°(T)]/RT = {[AH°(T°) - TAS°(T°)] + ACp[(T-T°)-Tln(T/T°)]}/RT

(7)

Vmax = kp[E]T=A[E]T^(-E/RT)

(8)

and

respectively. The temperature dependence of Km is fairly weak at low temperature(< 25 °C), but becomes strong at high temperatures. Vmax is rapidly increased with temperature. The parameters in eqs 7 and 8 estimated are AH° =12.3 ± 0.5 kcal/mol, T°AS° = -5.1 ± 0.5 kcal/mol, ACp° = -0.80 ±0.09 kcal/mol-K, A = 9.65 X 1011 s-i and E = 10.4 kcal/mol. The values of AH° and T°AS° are in good agreement with those previously published for the same system(17). Temperature dependence of K^. Prior to determining the temperature dependence of Ki, the progressive curve of the enzymatic assay is analyzed using eq 5 in order to obtain the steady-state velocity. Figure 2 shows the assay data and the fitting results for the thrombin and substrate reaction inhibited by an inhibitor with varying concentrations at 25 °C. It is readily seen that steady-state velocity becomes more evident with increased inhibitor concentration ([I]) because the inhibitor slows down the decrease of the substrate concentration. Figure 3 shows 1/Vs vs. [I] for the same system in the inhibitor concentration range of 0 - 0.269 nM at 25 °C. The parameters of eq 6 for Figure 3 are Vmax = 28.8 |LiM/s, Km = 5.3 |LiM, Ki = 0.011 nM and Vc = 0.0000 l|lM/s. Similar to the temperature dependence of Km, the variation of inhibition constant with temperature may be analyzed by using van't Hoff equation (eq 6). Figure 4 shows InKi vs. T(K) for P552 and its analogs 12000.0 10000.0

:

[\]=o/i\]=^ ^<x^]=0.082

8000.0 ,^A-*^=0A2i

j _ 6000.0 4000.0 2000.0 0.0 r, iV: 1

1111

: :

^ -13.0

5,*^-*1i^^

^ j

^

•-***tn=0.26{

:

^

280 285 290 295 300 305 310 315 320

T (K)

Figure 1. Temperature dependence of Michaelis constant (K ) and maximal velocity (V ). The solid and dashed lines are the fitting results by using eqs 7 and 8 in the text, respectively.

0.00

100

200

300

400

t (sec.)

500

600

700

Figure 2. Progressive curves of the reaction of thrombin, substrate and inhibitor(P552) at 25 °C. The solid lines are the fitting results by using eq 5 in the text.

Thermodynamics of Enzyme and Inhibitor Interactions P552(F56G) and P552(I59G). In general, InKi vs. T (K) is nonlinear. This states that the binding enthalpy and entropy should be considered as temperature dependent. In fact, the fitting values of the heat capacity are largely negative, ranging from -644 to -193 cal/mol-K for the inhibitors listed in Table 1. It is noteworthy that the negative heat capacity is characteristic of the binding interactions.

280 285 290 295 300 305 310 315 320

[I] (nM)

Figure 3, Reciprocal of the steady-state velocity (1/v^) vs. the inhibitor concentration ([I]) for P552 at 25 °C. The data is fitted by using eq 6 in the text.

280 285 290 295 300 305 310 315 320

T(K)

519

T(K)

280 285 290 295 300 305 310 315 320

T(K)

Figure 4. Variation of the inhibition constant (Kj) with temperature: (A)P552, (B)P552(F56G) and (C)P552(I59G). The soHd Hnes are the fitting results by using eq 7 where K^ is replaced by Kj.

Thermodynamic

data analysis

Table 1 lists the thermodynamic functions of some inhibitors studied previously. The free energy change, AG°, is directly determined from the logarithm of the inhibition constant (i.e. RTlnK,), the enthalpy, entropy and heat capacity are the fitting results by using eq 7 where K^ is replaced by Kj. Based on Scheme I, by using the relative thermodynamic functions (AAG°, AAH°, T°AAS°) given by AAX = AXn, - AX^

(9)

where "m" and "w" stand for the "mutated" and "wild-type" inhibitors, respectively, we are able to analyze the thermodynamic changes due to the mutation in conjunction with the structural features predicted from the molecular modelling. It is interestingly noticed that, for most analogs, the decrease in the binding affinity is attributed to the less favorable enthalpy whereas, for P552(F56G), the decrease in the binding affinity is attributed to the unfavorable entropy instead enthalpy (see Table 1). Molecular modeling for P552(F56G) suggests that the backbone located in AspH55-GlyH56-GluH57 has a large movement, in which the C" of GlyH56 shifts to C^ position of the original residue PheH56, Meanwhile, the water molecules with

520

Yudu Cheng era/. Table I Thermodynamic functions of thrombin and inhibitor interactions(25°C)*

Inhibitor

P552

Ki

AG°

AH°

T°AS°

0.011±0.0 -14.97+0.02 -13.66±1.12

ACn

1.31±1.13 -0.644±0.211

P552(F56G)

5.20±0.32 -11.30±0.03 -12.74±0.69 -1.44±0.70 -0.193±0.131

P552(I59G)

1.94±0.03 -11.88±0.01

-8.08±1.23

3.80+1.24 -0.439±0.232

P552(P60G)

0.39±0.08 -12.84±0.10 -10.73±1.62

2.11±1.64 -0.573±0.305

P552(Y63G)

0.21±0.03 -13.19±0.11

-6.07±1.57

7.12+1.58 -0.623±0.294

P552(L64G)

0.32±0.02 -12.95±0.04 -10.22±1.04

2.73±1.05 -0.532±0.197

*Data is adapted from reference (1). Units: Ki-nM; AG°, AH°, T°AS°-kcal/mol; ACp-kcal/molK.

hydrogen bonds to AspH55 and GluH57 move towards thrombin and form some more hydrogen bonds. It is then beheved that the movement of both the backbone and water molecule towards thrombin must compensate the loss in the molecular interactions due to the removal of the phenyl ring of PheH56^ and reduce the configurational entropy of the inhibitor around this area. For the analogs other than P552(F56G), no such conformational 8000.0 V6:^( change is predicted. Correspondingly, the 6000.0 I59G0 / ^ \ enthalpy and entropy of these analogs "O 4000.0 L64G ^J^ exhibit regular changes. Moreover, the : S 2000.0 \ • 'y^ O linear enthalpy-entropy compensation are ca 0.0 F56G ^ o observed for the congeneric series of _ -2000.0 thrombin bivalent inhibitors. Figure 5 < -4000.0 \ shows the relative enthalpy (AAH°) vs. the -6000.0 , relative entropy(AAS°). The relative free -^°^ - , o ,-20.0 -10.0 0.00 10.0 AAS° (cal/mol-K) energy(AAG°) and the overall change in the r

P60G5^

"^^^G

m o l e c u l a r s u r f a c e a r e a a r e f o u n d t o b e Figure S. Enthalpy-emropy compensation for the congeneric ,. , A . i r j.t ^ U.-A. A.r^u series of bivalent thrombin inhibitors. The filled circles are data I m e a r l y c o r r e l a t e d t o r t h e s u b s t i t u t i o n o r t h e from reference ( D and the open drcles are data from reference

nonpolar residues at the FRE:

(17).

AAG° = 0.0154 AAAnet + 1-34 (kcal/mol) (r=1.00)

(10)

where AAAnet = AAAnpi - AAApoi ("npl" and "pol" stand for the nonpolar and polar molecular surface area, respectively) and are calculated from the inhibitors in their bound and free states by using Scheme I. It is important to notice that the enthalpyentropy compensation is an intrinsic property of solute-solute interaction but largely

Thermodynamics of Enzyme and Inhibitor Interactions

521

enhanced by the solvent participation, and this feature of the enthalpy-entropy compensation is essential for the linear correlation of the relative free energy and the molecular surface area change due to the mutation(18).

IV. Summary The method presented in this article is applicable to the enzyme and inhibitor interactions and is particularly useful for those systems with high binding affinity. Analysis of the thermodynamic functions in conjunction with the structural features predicable from the molecular modeling may access the structural origin of the enzyme and inhibitor interactions upon the point mutation or substitution. The method may be utilized to develop the strategy of the rational inhibitor/drug design and protein/enzyme engineering.

References 1. Cheng, Y., Slon-Usakiewicz, J., Wang, J., Purisima, E., & Konishi, Y. (1996) Biochemistry 35, in press. 2. Wang, J., Szewczuk, Z., Yue, S. -Y., Tsuda, Y., Konishi, Y., & Purisima, E. O. (1995) J. Mol. Biol. 253, 473-492. 3. Stone, S. R., & Hofsteenge, J. (1986) Biochemistry 25, 4622-4628. 4. Rydel, T. J., Ravichandran, K. G., Tulinsky, A., Bode, W., Huber, R., Roitsch, C , & Fenton, J. W., II. (1990) Sciences 249, 277-280. 5. Betz, A., Hofsteenge, J., & Stone, S. R.(1991) Biochemistry 30, 9848-9853. 6. Yue, S. -Y., DiMaio, J., Szewczuk, Z., Purisima, E. O., Ni, F., & Konishi, Y. (1992) Protein Eng. 5, 77-85. 7. Yuko, T., Cygler, M., Gibbs, B. F., Pedyczak, A., Fethiere, J., Yue. S. -Y., & Konishi, Y. (1994) Biochemistry 33, 14443-14451. 8. Borgne, S. L., & Graber, M. {\99A)Appl. Biochem. Biotech. 48, 125-135. 9. Fethiere, J., Tsuda, Y., Coulombe, R., Konishi, Y., & Cygler, M. (1996) Protein Sci. 5, 1174-1183. 10. Weiner, S. J., Kollman, P. A., Nguyen, D. T., & Case, D. A. (1986) J. Comp. Chem. 7, 230-252. 11. Richards, F. M. {1911) Annu. Rev. Biophys. Bioeng. 6, 151-176. 12. Pascual-Ahuir, J. L., Silla, E., & Tuiionm I. (1994) J. Comp. Chem. 15, 1127-1138. 13. Blaber, M., Zhang, X., Lindstrom, J.D., Pepiot, S. D., Baase, W. A., & Matthews, B. W. (1994) /. Mol Biol. 235, 600-624. 14. Segel, I. H. (1975) Enzyme Kinetics: Behavior and Analysis of Rapid Equilibrium and SteadyState Enzyme Systems pp 100-160, John Wiley & Sons. 15. Morrison, J. F., & Stone, S. R. (1985) Comments Mol. Cell Biophys. 2, 347-368. 16. Morrison, J. (1988) Adv. Enzymol. 61, 201-301. 17. Di Cera, E., De Cristofaro, R., Albright, D. J., & Fenton, J. W., II (1991) Biochemistry 30, 7913-7924. 18. Cheng, Y., Wang, J., Slon-Usakiewicz, J., Purisima, E. O., and Konishi, Y., manuscript in preparation.

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Development and characterization of a Fab fragment as a surrogate for the IL-1 receptor Y, Cong, A. S. McColl, T. R. Hynes, R. C. Meckel, P. S. Mezes, C. L. Lane, S. E. Lee, D. J. Wasilko, K. F. Geoghegan, I. G. Otterness and G. O. Daumy Central Research Division, Pfizer Inc., Eastern Point Road, Groton, Connecticut 06340 L Introduction Interactions between proteins account for a substantial number of biological signaling events. These include classical hormone-receptor interactions at the cell surface, as well as the great number of intracellular interactions revealed by analyses of signal transduction pathways. They represent an attractive but difficult set of potential targets for pharmaceutical intervention. Most successfiil drugs are compounds of relative mass < 500 Da that are bioavailable when taken orally. Experience has shown that such small compounds, while capable of being bound tightly to pocket-like sites that accommodate natural ligands of similar structure and size, usually cannot achieve binding with sufficiently high affinity to the molecular surfaces of proteins recognized by other proteins (1,2). As a result, it remains to be determined how drugs that disrupt protein-protein interactions can be developed. In these circumstances, it appears worthwhile to create model systems that allow aspects of this problem to be analyzed. We have elected to use a monoclonal antibody that binds to human interleukin-lp (IL-IP) by recognizing amino acid residues that are also recognized by the IL-lp receptor (IL-IR). A monoclonal antibody that recognizes the receptor-binding residues of a cytokine can be considered a surrogate for the cytokine's natural receptor. Such a reagent might be valuable in assessing the structural basis of cytokine-receptor affinity, and could furnish a starting point from which to attempt the design of smaller competitive agents (3, 4). Certain precautions need to be observed at the outset of this effort. To be an appropriate subject for downsizing, an antibody must achieve critical interactions with at least part of the receptor-binding surface of IL-1 p. This is to exclude selection of an antibody that blocks access of IL-lp to the receptor merely by steric overlap of its molecular bulk with the space occupied by bound receptor (5). Such an antibody, on downsizing, would lead to a compound that fails to compete with IL-ip binding. Here we describe the selection and characterization of an antibody, and its Fab fragment, that provides a suitable starting point for this endeavor. n . Methods A. Cloning and expression oflLlp and its mutant derivatives A clone of the human IL-lp gene, modified to reflect the preferred codon usage of £. coli, was obtained from R&D Systems (Minneapolis, MN) and subcloned into the expression vector pET22b (Novagen, Madison, WI). Site-directed mutagenesis was performed to TECHNIQUES IN PROTEIN CHEMISTRY VIII Copyright © 1997 by Academic Press All rights of reproduction in any form reserved.

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Y.Cong era/.

generate the desired mutations of IL-ip (6). The pelB leader encoded by the vector was eliminated by Ndel digestion and religation, resulting in the final expression construct. The mutations chosen for this study were a cysteine substitution of residue K138 and alanine substitutions of residues R4 and L6. While the K138C mutation does not affect receptor binding (7, 8), the R4A and L6A changes were expected to produce proteins defective in binding to the IL-IR Type I (9-11). Two mutant forms of IL-ip were constructed. One harbored only the single K138C substitution, and was termed "native" to denote the unaltered condition of its receptor-binding surface. The second, termed "mutant #1", harbored the R4A and L6A substitutions in addition to the K138C replacement. The mutations were verified by DNA sequencing. B. Preparation of wild type rhlL-ip and rhlL-ip mutants Recombinant wild type hIL-lp, the K138C mutant and the K138C, R4A, L6A triple mutant (mutant #1) were isolated from the soluble fraction ofE. coli lysates by ammonium sulfatefi*actionationand hydrophobic interaction chromatography. The purified proteins were characterized by SDS-PAGE, western blots, N-terminal sequence, size exclusion chromatography (SEC), isoelectric focusing (lEF), matrix assisted laser desorption ionization mass spectrometry (MALDI-MS), and electrospray mass spectrometry (ESMS). C. Biotinylation of K138C IL-lp mutants The K138C mutants (-1.5 mg in 0.5 mL of PBS) were treated with 50 mM DTT (removed by gel filtration) and then biotinylated using biotin-maleimide (Sigma) (2:1 molar ratio). The biotinylated product was purified by gel filtration on Superdex 75. Evidence of biotinylation was routinely obtained by western blots probed with an avidinHRP conjugate (Pierce) and by binding of the proteins to streptavidin-coated BIAcore chips (Pharmacia SA5). D. Isolation of monoclonal antibody (mAb) and preparation of Fab fragments BALB/c mice immunized with recombinant human interleukin-lp (rhIL-lp) were the source of splenocytes for the production of mAbs. The X63-Ag8.653 myeloma cell line was used as the fiision partner (12). The mAbs were selected by binding to biotinylated K138C IL-ip immobilized in streptavidin-coated plates. A clone (F18/1E3) that produced an anti-IL-ip (IgGi) with the slowest off-rate when tested on K138C IL-ip immobilized on BIAcore chips was scaled-up in two 1 L spinners and the mAb (1E3) was isolated by protein G affinity chromatography. Fab fi-agments were prepared by digestion with immobilized papain and purified by size exclusion chromatography on Superdex 75 (Pharmacia) after undigested IgG and Fc fragments were first removed by affinity chromatography on protein A. E. Determination of kinetic constants Biotinylated K138C and mutant #1 were immobilized onto streptavidin-derivatized biosensor chips (Pharmacia SA5) by direct injection. Kinetic analysis of the binding of soluble IL-IR to the immobilized forms of IL-ip was carried out at low density, i.e. below 100 refi*active units (RU). Each binding cycle, with either soluble rhIL-lR (Genzyme) or

Fab Fragment as Surrogate for IL-1 Receptor

525

Fab fragments as analytes, was performed at a constant flow of 5 |LiL/min in PBS containing 0.02 % Tween. After the binding cycle, regeneration of the chip to its RU base line was achieved with either 5 mM or 2.5 mM NaOH, depending on the amount of analyte bound (as gauged by RU). Rate constants of dissociation (koff) were determined from analyte-saturated chips in order to minimize rebinding. Kinject, as described in the BIAcore manual, was used to confirm that koff was being measured under conditions free of rebinding. Rate constants of association (kon) were determined at different analyte concentrations and averaged (ave kon). The ratio of kog/ave kon was used to determine the dissociation constant (Ka) of the soluble IL-1 receptor and Fab fragments for immobilized K138C and mutant #1. All the data analysis was carried out using the BIAevaluation software (Pharmacia). For competition experiments, Fab was chemically coupled to CMS chips activated by treatment with l-ethyl-3-(3-dimethylaminopropyl)carbodiimide hydrochloride and N-hydroxysuccinimide, and the binding of wild type IL-ip (29 nM) was measured at different concentrations of soluble IL-1 receptor (0-167 nM). m . Results rhIL-lp and its mutant and biotinylated mutant derivatives were characterized biochemically before use. All the proteins appeared homogeneous by SDS-PAGE and SEC, but microheterogeneity was observed by ESMS, MALDI-MS, DBF and N-terminal sequencing. This was largely accounted for by N-terminal variation. Although all of the recombinant forms of IL-1 P originally had an N-terminus of Met-Ala-Pro, products with N-termini of Met, Ala and Pro were obtained. Kinetics of the binding of soluble IL-IR to immobilized IL-lp K138C and mutant #1 were measured using a BIAcore system (Table I). Soluble IL-IR exhibited about 8fold higher affinity for the K138C mutant than for mutant #1. The higher Kd exhibited for mutant #1 resulted from both a 4-fold lower association constant (kon) and a 2-fold higher dissociation constant (kog). This experiment confirmed that mutant #1, with the R4A and L6A mutations, was defective in binding the receptor. Table I. Kinetics of IL-IR binding to biotinylated IL-lfi K138C and ''mutant m** IL-1 form Mutation Kd(nM) kofif (sec'^) kon (M^ sec"^) "native"

K138C

0.0037

8.7x10^

4.3

mutant #1

K138C, R4A, L6A

0.0087

2.4x10^

35.5

In a similar fashion, the binding of Fab fragment 103095 to the same two ligands was also assessed (Table II). The Fab fragment bound to IL-ip K138C with Kd ~ 200 nM, i.e. about 50-fold more weakly than the IL-IR exhibited for this ligand. The higher Kd resulted primarily from a 20-fold lower kon. However, the Fab expressed the same relative preference as the IL-IR between the two forms of IL-lp; it recognized the K138C mutant, but failed to recognize the mutant defective in receptor binding.

Y. Cong etal.

526

Table n . Kinetics of Fab binding to hiotinylated IL-lp K138C and ''mutant #i'' Kd(nM) Mutation IL-1 form kon (IVr^ sec'^) koflF (sec'^) "native" mutant #1

K138C K138C, R4A, L6A

0.0069

3.5x10*

197

no binding

For competition studies, the Fab fragment was chemically coupled to the BIAcore chip and wild type rhIL-ip was used as analyte. Figure 1 presents the BIAcore sensorgrams obtained when different concentrations of wild type rhIL-ip (0.6-29 nM) were passed sequentially over the immobilized Fab chip. Soluble IL-IR was able to inhibit the binding of wild type rhIL-lp to immobilized Fab (Figure 2). Inhibition increased with increasing levels of soluble receptor until about 75% of the binding signal was suppressed. A residual level of apparent binding was attributed to nonspecific interaction by soluble proteins with the chip.

120 TIME (sec) Figure 1. Sensorgram traces for binding experiments in which different concentrations ofrhlL-lp were allowed to bind to anti-IL-1 Fab immobilized on a biosensor chip. The arrow indicates the time at which analyte was injected

Fab Fragment as Surrogate for IL-1 Receptor

527

80' ^^ 3

£60 O 0)

c o

8-40

o

20 1

20

1

-

1

40 60 IL-1 receptor (nM)

_

1

n

1

80

Figure 2. Competitive inhibition by soluble IL-l receptor of the binding of wild type rhlL-ip to immobilized Fab.

IV. Discussion The receptor-binding site of IL-ip has been mapped extensively by site-directed mutagenesis (9-11). These studies have revealed that two regions located about 25 A apart are important for receptor interactions (Figure 3, see color insert). One region (Patch A) is formed by the loop between p-strands 3 and 4, and is located on the side of the pbarrel. It is comprised of H30 and Q32. The other (Patch B) is located at the open end of the p-barrel, and encompasses a discontinuous set of residues including R4, L6, F46,156, K93, K103 and E105. The spatial separation of the two regions makes it unlikely that they could be bridged by a designed small molecule or even by an antibody, since antibody epitopes typically encompass four to nine amino acids (13-15). Nevertheless, anti IL-ip neutralizing antibodies have been shown to block the IL-lp:receptor interaction, and were also shown to bind at different but overlapping regions where the receptor binds (5). Thus, an antibody binding interaction to one of the two domains was strong enough to block the IL-lp:receptor interaction. We sought such an antibody as a first step toward designing a small molecular weight IL-ip antagonist. The traditional method of generating a blocking antibody has been to elicit a series of antibodies against the ligand and then determine which of the antibodies are neutralizing. Instead, we set out to generate a series of antibodies against one of the critical binding patches in the IL-1P:IL-1R interaction. Our criterion for antibody selection was that an antibody should bind well with wild type IL-lp, but poorly against a

528

Y. Cong etaL

mutant IL-ip in which prominent residues of the patch B binding region had been substituted by alanine. We also sought an efficient way to select for antibody that recognized the patch B binding region. It has previously been shown (5, 8) that the site-selectively monobiotinylated IL-lp mutant K138C retains its binding to IL-IR on a streptavidin surface, presumably because biotinylation occurs on a surface residue remote from the two binding regions (Figure 3). We bound monobiotinylated, oriented K138C IL-lp to a BIAcore streptavidin-coated chip and carried out binding studies with IL-IR in the presence of different concentrations of either K138C or wild type IL-lp as competitor. No appreciable differences in binding constants were apparent between K138C and wild type IL-ip, confirming that K138C was fully active in receptor binding using the E14 murine cell line and in the traditional IL-l/LAF bioassay (data not shown). Therefore, use of biotinylated K138C allows coating of a functional, orientated IL-1 on a streptavidincoated microtiter plate. In a standard antibody selection protocol, IL-ip would have been applied to the wells of microtiter plates so that binding could occur in a random fashion. With much of the antigen denatured by interaction with the plastic, antibodies selected by the screen would include many capable of recognizing only denatured IL-1 p. These would lack the ability to block receptor binding by the native cytokine. To focus monoclonal antibody selection on the native receptor-recognizing regions, only antibodies that bound to biotinylated K138C orientated on streptavidin plates were selected for further study. As a result, instead of the relatively large number of positive clones that might have been expected (potentially >100 in this case), only six antibodies were obtained, and all six of these bound to biologically active IL-1 p. This result appeared to validate the strategy of using a strategically oriented IL-ip as the antigen in the screening step, although no systematic comparison with a less specific method was performed to confirm this. Since the rate constant of dissociation (kofif), rather than the rate constant for association, is the primary determinant of differences in the Kd, we determined the apparent koff for each of the antibodies. The antibody (1E3) with the lowest apparent koff, and therefore, presumably the lowest Kd, was chosen for further study with the triple mutant K138C, R4A, L6A. The triple mutant K138C, R4A, L6A was prepared and its binding to IL-IR was compared to that of K138C. The results confirmed the importance of R4 and L6 for ILIR binding. A 10-fold increase in Kd was found in the triple mutant compared to K138C alone. To minimize the effect of steric hindrance and divalent binding of the IgG-lE3, a Fabfi-agmentwas prepared and its binding to the triple mutant was compared with its binding to biotinylated K138C. Fab-1E3 failed to bind to the triple mutant. This result demonstrated the successful selection of an antibody to the receptor-binding surface of the IL-lp molecule. It also demonstrated a fundamental difference between the ILlp:antibody and the IL-lp:IL-lR binding interfaces. The BL-IR protein:protein interaction interface contains at least two spatially separated binding domains. Diminished binding due to mutation at one domain raises the Kd, but need not abolish binding, because residues elsewhere can still support a lower affinity interaction. By contrast, an antibody

Fab Fragment as Surrogate for IL-1 Receptor

529

binding domain encompasses a limited number of spatially contiguous residues. Changes in those spatially close critical residues more readily abolish antibody binding. Finally, although by the criterion of non-binding to the triple mutant, Fab-1E3 appeared likely to be a receptor antagonist, it was important to confirm this. Consistent with the direct involvement of R4 and L6 in receptor binding, IL-IR binding to wild type IL-ip decreased the binding of Fab-1E3, and conversely, the binding of Fab-1E3 to wild type IL-ip decreased the binding of IL-1 R. The techniques developed during these studies are broadly applicable to selecting surrogate receptor (or ligand) antibodies toward other protein ligand:receptor pairs. First, the use of a biologically active, oriented ligand can result in a much more efficient first selection for blocking antibodies. Second, negative selection using an appropriate mutant will directly provide a blocking antibody that will also be a surrogate receptor (or ligand). We used K138C for the first selection and the triple mutant K138C, R4, L6 for the second selection, and found the blocking antibody Fab-1E3. Replacing negative selection using an appropriate mutant with a traditional positive selection scheme based on blocking activity will, of course, provide blocking antibodies, but such a selection scheme will detect blocking antibodies that are not receptor surrogates and thus are poor candidates for downsizing. Fab-1E3 fits the criteria that it is a receptor surrogate and therefore should be suitable for downsizing. References 1. 2. 3. 4. 5. 6. 7. 8. 9.

10. 11. 12. 13. 14. 15. 16.

Braisted, A. C. and Wells, J. A. (1996) Proc. Natl. Acad. Sci. USA 93, 5688-5692. DeGrado, W. F. and Sosnick, T.R. (1996) Proc. Natl. Acad. Sci. USA 93, 5680-5681. Smythe, M. L. and von Itzstein, M. (1994) J. Am. Chem. Soc. 116, 2725-2733. Saragovi, H. U., Fitzpatrick, D., Raktabutr, A., Nakanishi, H., Kahn, M. and Greene, M. I. (1991) Science 253, 792-795. Simon, P. L., Kumar, V., Lillquist, J. S., Bhatnagar, P., Einstein, R., Lee, J., Porter, T., Green, D., Sathe, G. and Young, P. R. (1993) J. Biol. Chem. 268, 9771-9779. Kunkel, T. A., Roberts, J. D. and Zakour, R. A. (1987) Methods Enzymol. 154, 367-382. Wingfield, P., Graber, P., Shaw, A. R., Gronenborn, A. M., Clore, G. M. and MacDonald, H. R. {\9%9)Eur. J. Biochem. 179, 565-571. Chollet, A., Bomiefoy, J.-Y. and Odermatt, N. (1990) J. Immunol. Methods 127, 179-185. Labriola-Tomkins, E., Chandran, C, Kaffka, K. L., Biondi, D., Graves, B. J., Hatada, M., Madison, V. S., Karas, J., Kilian, P. L. and Ju, G. (1991) Proc. Natl. Acad Sci. USA 88, 1118211186. Grutter, M. G., van Oostrum, J., Priestle, J. P., Edelmann, E., Joss, U., Feige, U., Vosbeck, K. and Schmitz A. (1994) Prot. Eng. 7, 663-671. Evans, R. J., Bray, J., Childs, J. D., Vigers, G. P. A., Brandhuber, B. J., Skalicky, J. J., Thompson, R. C. andEisenberg, S. P. (1995) J. Biol. Chem. ll^S, 11477-11483. Kearney, J. F., Radbruch, A., Liesegang, B. and Rajewsky, K. (1979) J. Immunol. 123, 1548-1550. Kabat, E. {1910) Ann. N. Y. Acad Sci. 169, 43-54. Schecter, I. {1911) Ann. N Y. Acad Sci. 190, 394-419. Hodges, R. S., Heaton, R. J., Parker, J. M. R., Molday, L. and Molday, R. S. (1988) J. Biol. Chem. 263, 11768-11775. Priestle, J. P., Schaer, H. P. and Gruetter, M. G. (1989) Proc. Natl. Acad Sci. USA 86, 9667-9671.

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SECTION VII Macromolecular Assemblies

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Topology of Membrane Proteins in Native Membranes Using Matrix-assisted Laser Desorption lonization/Mass Spectrometry Kamala Tyagarajanl, John G. Forte ^ and R.Reid Townsend^ iDept. of Molecular and Cell Biology, University of California, Berkeley, CA 94720-3200 and ^Dept. of Pharmaceutical Chemistry, University of California, San Francisco, CA 941430446

I.

Introduction

Knowledge of the topological orientation of membrane proteins within native membranes is fundamental to establishing structure-function relationships. In particular, information on topology is important for understanding the structural basis underlying the translocation function of cation pumps like the Na,KATPase, Ca-ATPase or H,K-ATPase. Previous efforts to define topology have used both theoretical and experimental approaches, such as hydropathy plots, proteolysis of vesicles, binding of regio-specific antibodies, or labeling with group-specific, membrane-sided reagents followed by identifying the modified sites (1, 2). Proteolysis of sided vesicles followed by analysis of peptide products has been one of the most common approaches to determine exposed peptide sequences. Conversely, remaining membrane-associated peptides can be analyzed after exhaustive protease digestion. Many analyses have utilized SDSPAGE to separate proteolytic fragments followed by Edman sequencing of peptides or identification using regio-specific antibodies (3, 4). However these approaches are not useful for identifying small peptides (< 5 kDa) from proteolysis. Alternatively, HPLC separation of peptides followed by Edman sequencing is possible but time-consuming and the coelution of multiple peptides makes identification by Edman sequencing difficult. More recently, mass spectrometry has been used in the identification of peptides and glycopeptides, in topological studies (5-8). In this study, we used matrix-assisted laser desorption ionization /Mass Spectrometry (MALDI/MS) to identify the peptides released from gastric parietal cell microsomes. MALDI, because of its sensitivity and relative tolerance to the presence of salts and buffers was examined for the analysis of unfractionated proteolytic digests (9, 10). MALDI with post-source decay (PSD) analysis was used to obtain sequence information on peptides even in crude digestion mixtures. Our strategy (Figure 1) consisted of proteolysis of intact vesicles, centrifugation at high speeds to separate membrane bound and soluble fractions and analysis of the mixture of released peptides by MALDI/MS. In addition, to increase the TECHNIQUES IN PROTEIN CHEMISTRY VIII Copyright © 1997 by Academic Press All rights of reproduction in any form reserved.

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Kamala Tyagarajan et al

534

Protein in Vesicles Proteolysis ; Centrifugation / \ Pellet Supernatant Peptides V^

MALDI/MS

I

\ . RP-HPLC separation

HPLC fractions

;

MALDI/MS with PSD analysis Topological Models—Sequencing of peptides Figure 1. Methodology used to determine the topology of membrane proteins.

sensitivity and breadth of analysis, supernatant peptides were separated by reverse-phase HPLC and individual fractions were analyzed by MALDI/MS. PSD-analysis was also performed to obtain partial sequence information and identify peptides (11). On basis of the released peptide products a topological map for a major portion of the H,K-ATPase in gastric parietal cell tubulovesicles is proposed. We focused on the gastric H,K-ATPase as a test protein because i) purified gastric microsomal vesicles are highly enriched in the enzyme (> 85-90% purity), ii) the vesicles are oriented with a common asymmetry i.e. cytoplasmic side out (12), iii) the vesicles are sealed allowing selective cytoplasmic digestion, and iv) there is a pool of existing topological data from other methods (13, 14, 15& 16).

subunit beta-subunit

Cytoplasm

Figure 2. The gastric H,K-ATPase in gastric microsomal vesicles. The H,K-ATPase is a heterodimer composed of an a-subunit and a glycoprotein p-subunit, which are asymmetrically oriented.

Topology of Membrane Proteins Using MALDI/MS

535

Thus, the H,K-ATPase in microsomes is a useful model to develop new methods to determine protein topology. The cartoon in Figure 2 illustrates that the H,KATPase is a heterodimer composed of two subunit proteins: an a-subunit of 1035 amino acids, traversing the membrane either 8 or 10 times (13), with most of its mass cytoplasmically disposed (and therefore outside the vesicles); and a glycosylated (i-subunit of 300 amino acids, traversing the membrane once and, except for a short cytoplasmic tail, with most of its mass on the extracellular side (inside the vesicles).

II. Materials and Methods Materials. Trypsin, Lys C, chymotrypsin and adrenocorticotropic hormone fragment (18-39) were purchased from Sigma (St. Louis, Mo). Tris(hydroxymethyl)aminomethane (Tris), sucrose, acetonitrile, HPLC grade water and acetic acid were purchased from Fisher Scientific (Pittsburgh, PA). Matrix (a-cyano 4-hydroxy-cinnamic acid) was purchased from Hewlett Packard (Palo Alto, CA). The low-molecular weight calibration standard was purchased from Bio-Rad (Richmond, CA). Preparation of H,K-ATPase enriched microsomal vesicles. H,K-ATPasecontaining gastric microsomal vesicles were isolated from rabbit stomach as previously described (12). Crude microsomes were harvested from homogenized mucosa of unstimulated rabbit stomach (H2 receptor-blocked) as the membrane pellet sedimenting between 10 min at 13,000 x g and 1 hr at 100,000 x g. The pellet was resuspended in 10% sucrose, brought to 40% sucrose (9 ml), and overlaid with successive layers of 30% sucrose (11 ml), 10% sucrose (16 ml) [300 mM sucrose, 5 mM tris(hydroxymethyl)aminomethane (Tris), and 0.2 mM EDTA, pH 7.4] in a 37 ml tube. After centrifugation at 80,000 x g for 4 hr, the purified gastric microsomal vesicles were collected from the interface between 10% and 30 % sucrose and stored at 4° C until use. Trypsinization of H,K-ATPase-enriched gastric microsomal vesicles. Tubulovesicles (-100 jxg of protein) were treated with trypsin (5 jig) in Tris.HCl (20 mM, pH 7.5) at 37°C for 30 min. The vesicles were next centrifuged at 100,000 X g on a TLIOO table top centrifuge for 1 hr at 4°C. The supernatant was carefully separated from the pellet. The supernatant was next boiled for 5 min and stored at -20°C until further analysis. Reverse Phase-HPLC separation of tryptic digest.. The tryptic digest (60%) was separated on an Aquapore OD-300 (Applied Biosystems Inc) C18 reverse phase column (7 |i and 1 x 250 mm) using a Michrom UMA Model 600 HPLC system with eluant monitoring at 214 nm. The first 5 min of the gradient was isocratic at 5% eluant B (98% CH3CN, 0.1% TFA) and 95 % eluant A (2% CH3CN, 0.1% TFA). This was followed by a linear gradient of 5-15 % B in 15 min, 15-50% B at 75 min and 50-75 % B at 90 min. The flow rate was 50.0 |il/min. Individual fractions were collected and stored for subsequent use. MALDI/MS analysis. Supernatant (1 jxl) was diluted 1:2 with 50% CH3CN in water and this mixture was mixed with 2 |il of a-cyano 4-hydroxy-cinnamic acid, vortexed and centrifuged. One |il was spotted onto the target. MALDI/MS of samples was carried out on a TofSpec SE from Micromass (Manchester, UK), equipped with a reflectron and using a nitrogen laser (337 nm). Samples were

Kamala lyagarajan et al

536

initially examined in the linear mode to determine whether signals > 5 kDa were present. An accelerating potential of 25 kV, a reflectron voltage of 28.5 kV and an extraction voltage of 10 kV in the reflectron-ion mode were typically used. Thirty shots were usually averaged. The instrument was calibrated with peptides from a low molecular weight peptide set from Biorad (Richmond, CA). Molecular ions of bombesin and the 18-39 amino acid clip of adrenocorticotropic hormone fragment were used as calibration standards.

III.

Results and Discussion

The H,K-ATPase-enriched vesicles were trypsinized for 30 min using a trypsinrprotein ratio of 1:25 and then centrifuged to separate the pellet from the supematant fractions. An aliquot of the supernatant was analyzed by MALDI/MS as shown in Figure 3. The observed signals {m/z 600-4400) were assigned to masses (±2 Da) of the predicted tryptic digestion products for the gastric H,KATPase a-subunit as shown in Table 1. Since, the exposure to trypsin was for a limited period of time (30 min) incompletely cleaved tryptic peptides were also observed, so it was important to include the possibility of these incompletely cleaved peptides in the search through the molecular mass signals.

I

l^^i;^^,)!^^^ 800

1200

1600

2000 2400 m/z

2800

3200

3600

4000

Figure 3. MALDI mass spectrum of the total supernatant from the tryptic digest of the H,KATPase-enriched tubulovesicles. The H,K-ATPase was digested with trypsin and the vesicles centrifuged to separate supernatant from the pellet. An aliquot of the supernatant was analyzed by MALDI/MS in the reflectron ion mode using a-cyano 4-hydroxy cinnamic acid as a matrix. The signals are denoted by numbers and were assigned to a-subunit peptides (Table 1).

537

Topology of Membrane Proteins Using MALDI/MS

Table 1. Assignment of signals obtained by MALDI/MS of a tryptic digest supernatant from H,K-ATPase-enriched vesicles. The observed masses of the numbered signals shown in Figure 3 were assigned to the masses of a-subunit peptides. Signal

No. Observed MH+

Calculated MH+

a-subunit peptide Asp483-Lys487

1 2

663

662.4

819

818.5

Val435_Arg44l

3

899

898.6

Leu'78_Arg85

4

1043

1044.6

Ala32-Lys42

5

1047

1046.6

Leu7lO-Arg7l8

6

1056

1055.6

Gly206.Arg215

7

1076

1076.6

Ala673-Lys682

8

1088

1087.5

Thr695-Arg703

9

1093

1092.6

Asp86-Arg95

10

1196

1195.7

Leu659-Arg668

11

1239

1238.7

Asnl'74.Argl84

12

1283

1282.7

Tyr66.Arg'7'7

13

1324

1323.8

Leu659-Lys669 or Ala838.Arg848

14

1370

1370.7

Gly536_Arg546

15

1375

1374.7

Asp683_Arg694

16

1458

1458.7

Phe470-Arg482

17

1485

1485.7

Ser239-Arg251 or Arg456-Lys469

18

1619

1619.8

Val224.Arg238 or Asp5ll-Arg524

19

1679

1678.9

Ala43l-Lys445

20

1712

1711.9

Glu547-Arg562

21

1824

1823.9

Phe499-Arg513

22

2013

2013

23

2159

2159.1

Glu49-Lys65 Ala637.Arg658

24

2448

2449.2

Asn755-Arg777

25

2543

2544.2

Asn252_Arg275

26

2736

2737.4

Asn37l-Arg396

27

2793

2792.3

Glu49-Lys72

28

3553

3553.9

Asnl74.Lys205

29

4004

4005

Lys738.Arg'777

Based on mass analysis by MALDI/MS of the unfractionated tryptic digest, 29 tryptic peptides from the a-subunit were tentatively identified, but only two signals corresponded to (3-subunit peptides. These latter signals at m/z 791 and 1485 corresponded to peptides from the short cytoplasmic tail of the p-subunit, and included Met^-Lys'^ and Lys^-Lys^^ from the N-terminus of the sequence. No signals that corresponded to masses of the extracellular domain of the (3subunit peptides were observed, consistent with the vesicles being oriented with their cytoplasmic side-out and preservation of vesicular integrity during

538

Kamala Tyagarajan et al

proteolysis by trypsin. The a-subunit peptides tentatively identified by MALDyMS are listed in Table 1, including: Ala32-Lys42, Glu49-Lys65, Glu^^Lys'72, Tyr66-Arg'7'7, Leu'^^-Arg^^ and Asp^^-Arg^^ from the N-terminus (before membrane segment Ml); peptides Asn^'^^-Argi^^^ ^sn^'^^-Lys^o^^ Q\y206. Arg2i5^ Yai224_Arg238 Ser239.Arg25i and Asn252-Arg275 in the cytosolic loop between membrane segments M2 and M3; peptides in the large cytosolic loop between M4 and M5 which included Asn37i.Arg396, Ala^3i.Lys445 Arg^^e. Lys469, Phe470-Arg482, Asp483.Lys487, Phe499.Arg5i3, Gly536-Arg546, Glu547Arg562, Ala637_Arg658, Leu659.Lys669 , Ala673.Lys682, Asp683-Arg694, Thr695. Arg703, Leu7io_Arg7i8^ Lys738_Arg777 and Asn755-Arg777; and a peptide Ala838Arg848 from the cytosolic loop between membrane segments M6 and M7. All of these regions have previously been deduced to be cytoplasmic (15, 16). No peptides corresponding to any of the intramembrane segments or intravesicular (extracellular) regions of the a-subunit were observed. Thus the topological prediction obtained by analysis of the MALDI mass spectrum of the entire tryptic supernatant was consistent with the currently accepted topological model of H,KATPase (16). An assignment of the identified peptides to putative extracellular regions of the H,K-ATPase is schematically shown in Figure 4. Although analysis of peptide masses in the total supernatant allows a tentative identification, mass overlap at this resolution may lead to erroneous assignments. For example, signal suppression can lead to low intensity or abolition of certain peptide signals. Since the peptides are tentatively identified on the basis of mass alone, it is prudent to perform PSD analysis to obtain sequence information and confirm the identity. PSD analysis could be performed on some peptides in the total mixture; however, it was difficult to obtain sequence information on low Luminal solution

f\ Apical I ? plasma ^Jmembrane

Cytoplasm Figure 4. Topological model of the gastric H,K-ATPase. The topological model shown is adapted from a proposal by Besancon et al. (14). The model depicts the a-subunit having ten tranmsmembrane segments, denoted as Ml-MlO. Amino acid numbers are shown for the cytoplasmic ends of segments M1-M8. The glycoprotein p-subunit traverses the membrane once and has most of its mass luminally oriented. The darkened regions indicate peptides of the asubunit that were identified by MALDI/MS analysis of the total tryptic digest supernatant of H,K-ATPase-enriched vesicles (Figure 3 and Table 1).

Topology of Membrane Proteins Using MALDI/MS

539

40 Time (min) Figure 5. Reverse-phase HPLC of the supernatant from a tryptic digest of H,K-ATPaseenriched vesicles. Peak fractions were collected up to 60 min using the gradient described in "Methods".

intensity peptides and peptides that were separated by less than 14 Da. In order to obtain a series of purified peptides, we subjected the tryptic digest supernatant to RP-HPLC as described in "Methods". The RP-HPLC trace of the digest is shown in Figure 5. We collected 30 individual fractions and an aliquot of each was subjected to MALDI/MS. The MALDI/MS of each HPLC fraction showed the presence of several peptides which had sufficient mass-differences for successful PSD-analyses. Figure 6A shows the MALDI mass spectrum of a representative fraction, Fraction 13, from the HPLC preparation. Signals were observed at m/z 730, 1047, 1327, 1371, 1394, 1798, and 2141. The assignment of these signals to peptides of the a-subunit is summarized in Table 2. Although we had noted peptides at 1047 and 1371 in the total supernatant material, signals at m/z 730, 1327, 1394, 1798 and 2141 were apparent only after HPLC fractionation. The sequence and identity of the peptides was confirmed by PSD-analyses. The PSD spectrum of the signal at m/z 1798 is shown in Figure 6B. As an example, the PSD spectrum of m/z 1798 gave a series of y ions ranging from ya-yi? and the b ions from b3-b6 confirming the amino acid sequence to be identical to peptide '719LGAIVAVTGDGVNDSPALK737 of the a-subunit. Interestingly, the presence of the series of y ions from ys-yi? demonstrated that the sequon, '731 Asn-Asp-Ser'733, exists in a non-glycosylated form. It has been suggested that one of the Asn residues within the cytoplasmic domain of the a-subunit is glycosylated (17).

Kamala lyagarajan et al

540

700

m/z

900

Figure 6. MALDI mass spectrum of fraction 13 from RP-HPLC. The H,K-ATPase-enriched vesicles were trypsinized and centrifuged to separate supernatant from pellet. The supernatant was subjected to RP/HPLC and individual fractions collected and subjected to MALDI/MS. The MALDI mass spectrum (reflectron-ion mode) was obtained using a-cyano-4-hydroxy cinnamic acid as a matrix (Panel A). The signals were assigned to a-subunit peptides (Table 2). The signal at m/z 1798, indicated by an arrow was next subjected to PSD-analysis. The PSDspectrum of MH"*" 1798.4 is shown in Panel B. Only the peaks for the b and y fragment ions are labeled. The deduced amino acid sequence is shown at the top of the panel.

Topology of Membrane Proteins Using MALDI/MS

541

Table 2. Tryptic peptides of a-subunit in Fraction 13. The H,K-ATPase-enriched microsomes were trypsinized and centrifuged to separate the supernatant from the pellet. The supernatant was subjected to RP/HPLC and individual fractions were collected and analyzed by MALDI/MS. The MALDI mass spectrum of fraction 13 is shown in Figure 6A. The signals seen were assigned to a-subunit tryptic peptides, as shown below. Observed MH+

Calculated MH+

1047

1046.6

a-subunit peptide ^lOLVIVESCQR^ls

1327

1329.8

457IVIGDASETALLK469

1371

1370.7

536GQELPLDEQWR546

1396.8

661VPVDQVNRKDAR672

1798

1797.0

'719LGAIVAAVTGDGNDSPALK737

2141

2141.1

48KEMEINDHQLSVAELEQK65

1394

Use of alternative proteases Proteases other than trypsin may be used to increase the coverage of the protein sequence or resolve ambiguities from mass overlap. For example, Lys C for topological analysis gave results that were complementary to trypsin (data not shown). From a Lys C digest it was determined that several peptides from regions Ala^-Lys^^^, Seri^4-Lys223^ Arg^^^-Lys'^^^ and Asp^^s.LygSSi ^vere cytoplasmic. Again, a signal at m/z 2824 corresponded to the mass of peptide '7iOLeu-Lys'737 (2825 Da) of the a-subunit which includes Asn'731. These data were again consistent with the accepted topological model of the H,K-ATPase (Figure 4). Treatment of vesicles with chymotrypsin using similar conditions as for trypsin (1:20, chymoptrypsin:protein) and MALDI-MS analysis of the supernatant after centrifugation of the digest gave some interesting results. Signals at m/z 996, 1015, 1298, 1460 and 1678 were observed which corresponded to the masses of p-subunit peptides (Tyr2i9-Leu227, Seri5i-Leui59, Leu25i-Leu262, Cys58-Tyr69 and Arg^^-Tyr^^, respectively) and are known to have an intra-vesicular orientation. Further investigation including PSD analyses will be performed to confirm the identity of these peptides.

IV.

Conclusions

We have demonstrated the utility of MALDI/MS in combination with proteolysis to investigate the topology of a heterodimeric membrane glycoprotein, the gastric H,K-ATPase within its native microsomal membrane. MALDI/MS proved to be a rapid and sensitive method for topological analysis of membrane proteins in native membranes. The high sensitivity, and relative tolerance of MALDI/MS to buffers and some detergents, allowed rapid assessment of topology by examination of unfractionated supernatants from vesicular digests. The above approach may also be usefully employed to assess the reconstitution of proteins into vesicles and vesicular integrity. Analysis of HPLC fractions by MALDI with PSD analysis allowed the determination of partial peptide sequence and may prove suitable for identifying post-translational modifications of

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Kamala Tyagarajan et al

extravesiculj peptides. Finally, this approach should provide a convenient, extravesicular sensitive anc and rigorous assessment of protein topology in artificial and native membranes.

Acknowledgments This project was supported in part by NIH grant DK38792. The mass spectra were obtained at the UCSF Mass Spectrometry Facility supported by the Biomedical Research Technology Program of the National Center for Research Resources (NIH NCRR BRTP RR01614 and RR08282). The VG TofSpec SE was partially supported by Micromass, Beverley, MA.

References 1. Modyanov, N., Lutsenko, S., Chertova, E., Efremov, R. and Gulyaev, D. (1992) Acta Physiol. Scand. Supplementum, 607, 49-58. 2. Loo, T.W. and Clarke, D.M. (1995) J. Biol. Chem. 270, 843-848. 3. Serrano, R., Monk, B.C., Villalba, J.M., Montesinos, C. and Weiler EW. (1993) Eur. J. Biochem., 212, 737-744. 4. Ban, W.J. Jr, Abbott, A., Sun, Y. and Malik, B. (1992) Ann. New York Acad. Sci., 671, 436-439. 5. le Maire, M., Deschamps, S., Moller, J.V., La Caer, J.P. and Rossier, J. (1993) Anal. Biochem., 214, 50-57. 6. Mel, S.F., Falick, A.M., Burlingame, A.L. and Stroud, R.M. (1993) Biochemsitry, 32, 9473-9479. 7. Moore, C.R., Yates, J.R., Griffin, P.R., Shabnowitz, J., Martino, P.A., Hunt, D.F. and Cafiso, D.S. (1989) Biochemistry, 28, 9184-9191. 8. Poulter, L., Earnest, J.P., Stroud, R.M. and Burlingame, A.L. (1989) Proc. Natl. Acad. Sci., 86, 6645-6649. 9. Tsarbopoulos, A., Karas, M., Strupat, K., Pramanik, B.N., Nagabushan, T.L. and Hillenkamp, F. (1994) Anal. Chem., 66, 2062-2070. 10. Billeci, T. M., and Stults, J.T. (1993) Anal. Chem., 65, 1709-1716. 11. Spengler, B., Kirsch, D., Kaufmann, R. and Jaeger, E. (1992) Rapid Commun. Mass Spectrom., 6, 105-108. 12. Reenstra, W.W. and Forte, J.G. (1990) Meth. in Enzymol., 192, 151-165. 13. Bamberg, K. and Sachs, G. (1994) J. Biol. Chem., 269, 16909-16919. 14. Asano, S., Arakawa, S., Hirasawa, M., Sakai, H., Ohta, M., Ohta, K.and Takeguchi N. (1994) Biochem. J., 299, 59-64. 15. Sachs, G., Besancon, M., Shin, J.M., Mercier, F., Munson, K. and Hersey S. (1992) J. Bioenerg. Biomem., 24, 301-308. 16. Besancon, M., Shin, J.M., Mercier, F., Munson, K., Miller, M., Hersey, S. and Sachs, G. (1993) Biochemistry, 32, 2345-2355. 17. Tai, M.M, Im, W.B., Davis, J.P., Blakeman, D.P., Zurcher-Neely, H.A. and Heinrikson, R.L. (1989) Biochemistry, 28, 3183-3187.

Role of D-Ser*^ in the P-type Calcium Channel Blocker, co-Agatoxin-TK Tomohiro Watanabe, Manabu Kuwada, Kumiko Y. Kumagaye*, Kiichiro Nakajima*, Yukio Nishizawa and Naoki Asakawa Eisai Tsukuba Research Laboratories, 5-1-3 Tokodai, Tsukuba, Ibaraki 300-26, Japan and *Peptide Institute Inc., Protein Research Foundation, Osaka 562, Japan

I. Introduction Multiple types of voltage-dependent calcium channels in mammalian neurons play important roles in controlling various nervous functions such as synaptic transmission, gene expression, neuronal development and differentiation. There are at least four subtypes of the calcium channels, namely T-type, L-type, N-type, and P-type channels, classified on the basis of their electrophysiological and pharmacological properties. Among them, the P-type calcium channels have been reported to be primarily associated with neuronal transmission through regulating the release of excitatory amino acids and catecholamines (1-4). We have previously isolated a 48-amino-acid peptide, named co-agatoxin-TK (co-Aga-TK), from the venom of the funnel web spider, Agelenopsis aperta. It was found to be a potent blocker of the P-type calcium channels in rat cerebellar Purkinje neurons, but TECHNIQUES IN PROTEIN CHEMISTRY VIII

543

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Tomohiro Watanabe et al

had no activity against T-type, L-type, or N-type channels in brain neurons. The peptide has a unique structural profile including a high-density disulfide core structure with four disulfide bonds and a D-form amino acid, D-Ser, at position 46 (Fig. 1). Interestingly, coAga-TK contains two serine residues at positions 28 and 46 of which only Ser^^ is in the D-form (©-[o-Ser^^JAga-TK) (5,6). W e have also found in the spider venom a related peptide with the same amino acid sequence and disulfide pairings as those of CO-[DSej-46]Aga-TK except for the L-configuration of Ser^^ (co-[L-Ser46]AgaTK), though the L-Ser^^ toxin is about six times less abundant than the D-Ser^^ toxin (7). These findings raise the questions of why only Sef*^ of the two serine residues is in the D-form and why the two co-Aga-TKs containing opposite configuration at Ser^^ are both present in the Agelenopsis aperta venom. Heck et al. (8) have reported the presence in the venom of a novel peptide isomerase that specifically converts L-Ser^^ to D-Ser^^ residue of co-Aga-TK. We have recently reported the complete primary structure of the peptide isomerase, which is a 29-kDa glycoprotein consisting of a 243-residue heavy chain and an 18-residue light chain cormected by a single disulfide bond (9). This was the first report to assign the structure of a peptide isomerase from an eukaryotic organism that converts the chirality of amino acid residues. co-[D-Ser4^]Aga-TK has very low solubility under neutral conditions, which precluded detailed studies of its tertiary structure by NMR spectroscopy. However, Adams et al (10) and Yu et al (11) reported two-dimensional NMR analyses of co-[D-Ser4^]Aga-TK in acidic solution; they concluded that the cystine-rich region consists of a triple-stranded antiparallel p-sheet with four loops formed by four disulfides (Cys^-Pro^^), but the carboxyl-terminal tail was very poorly defined since the carboxyl-terminal ten residues containing the D-Ser'^^ residue (Arg^^-Ala^^) adopt a disordered structure. Our structure-function relationship studies of co-[D-Ser4^]Aga-TK demonstrated that co-[L-Ser^6]Aga-TK has 80- to 90-fold less potency towards the P-type calcium channels compared with CO-[DSer46]Aga-TK. Two proteolytic fragments of co-[D-Ser46]Aga-TK, namely co-Aga-TK (1-43) and a carboxyl-terminal peptide fragment, co-Aga-TK (44-48), did not exert any significant inhibition of P-type calcium channels or interfere with the blockade of the channels elicited by native co-Aga-TK (12). Furthermore, molecular dynamics calculations showed that the carboxyl-terminal sixamino-acid peptide of co-Aga-TK containing D-Ser^^ assumes a different conformation from that containing L-Ser^^. These data suggested that the specific conformation of the carboxyl-terminal

Role of D-Ser46 in w-Agatoxin-TK

545

tail generated by the D-Ser^^ residue, together with the triplestranded antiparallel p-sheet, might be essential for the blockade of the P-type calcium channels. loop-4

OOH Figure 1. Schematic diagram of the high-density disulfide core and carboxylterminal tail containing D-Ser^^ in Q)-[D-Ser^^]Aga-TK. The disulfide core structures are represented en the basis of the coordinates determined by NMR spectroscopy (11). Amino acid residues of the peptide are represented by single-letter abbreviations in the circles.

546

Tomohiro Watanabe et al

In this study, the conformations of co-[D-Ser4^]Aga-TK and co[L-Ser46]Aga-TK were investigated by the combination of sizeexclusion chromatography, circular dichroism (CD) measurement, and fluorescence spectroscopy in order to elucidate the structural and functional effects of the configuration of the Ser^^ residue in coAga-TK. We have found that co-[D-Ser4^]Aga-TK has a particularly compact molecular shape involving p-sheet structure, whereas co[L-Ser46]Aga-TK has a relatively unfolded or extended structure at physiological pH and ionic strength. These data are discussed in terms of the possible role of the configuration of the Ser^^ residue in determining the molecular conformation of ©-Aga-TK.

11. Experimental Procedures A. Peptides and Reagents co-[L-Ser46]Aga-TK and co-[D-Ser^^]Aga-TK were synthesized by Drs. K. Y. Kumagaye and K. Nakajima of Peptide Institute Inc. using a Applied Biosystems type 430A peptide synthesizer as described previously (5). Synthetic co-[D-Ser46]Aga-TK is commercially available from the company. High-purity guanidine hydrochloride was obtained ICN Biomedicals, Inc. (Aurora, OH). The phosphate-buffered saline, pH 7.4, was prepared by dissolving Dulbecco's PBS Powder (Nissui Pharmaceutical Co., Ltd., Tokyo) in Milli-Q water, and consists of 8.10 mM Na2HP04,1.47 mM KH2PO4, 2.68 mM KCl, and 137 mM NaCl. Other chemicals and reagents used were of reagent grade.

B. Size-Exclusion Chromatography The apparent molecular masses of co-[D-Ser4^]Aga-TK and CO-[LSer^6]Aga-TK were determined by size-exclusion chromatography (LKB GTI HPLC Systems) with a Pharmacia Superdex 75HR column (10 x 300 mm) or a TSK G3000SWXL column (10 x 300 m m ) equilibrated with Dulbecco's phosphate-buffered saline, pH 7.4, with or without 5.2 M guanidine hydrochloride. The peptides were eluted from the columns with the buffer at the flow rate of 0.5 ml/min at 25 °C and elution profiles were monitored by measuring the absorbance at 280 nm or 220 nm. The column was calibrated using a Pharmacia low-molecular-weight marker kit (blue dextran, bovine serum albumin, ovalbumin, chymotrypsinogen, and

Role of D-SeH6 in co-Agatoxin-TK

ribonuclease A), aprotinin Institute Inc.).

547

(Sigma), and substance P (Peptide

C. Spectroscopic analysis For CD and fluorescence spectroscopic analyses of CO-[LSer^^]Aga-TK and co-[D-Ser^^]Aga-TK, peptide samples were prepared by freshly dissolving the lyophilized peptides at a concentration of 150 |Lig/ml in Dulbecco's phosphate-buffered saline, pH 7.4, in the presence or absence of guanidine hydrochloride. CD spectra were recorded with a Jasco J-720WI spectropolarimeter at room temperature using a 0.1 cm path-length cell. In all cases, the buffer base-line spectrum was subtracted, and the results were expressed in terms of the mean residue ellipticity {0) in units of degrees cm^ dmol'^. Fluorescence spectra were determined with a Hitachi F4500 spectrofluorometer using a 1 cm path-length cell at 25 °C.

III. Results And Discussion A. Different molecular shapes of co-lD-Ser^^lAga-TK and co[L-Ser^^lAga-TK During the isolation and characterization of biologically active peptides from Agelenopsis aperta venom, we found that two stereoisomers of the P-type calcium channel blocker, CO-[DSer4^]Aga-TK and co-[L-Ser46]Aga-TK, were eluted in distinct fractions on size-exclusion chromatography (13). This finding was confirmed with synthetic standards of the two toxins on a Superdex HR75 column equilibrated with phosphate-buffered saline, pH 7.4, as shown in Fig. 2. co-[D-Ser4^]Aga-TK was found to be eluted from the column significantly later than co-[L-Ser^^]Aga-TK (the D-Ser toxin, 39.9 min; the L-Ser toxin, 31.5 min), in spite of their identical molecular mass of 5273 Da. The D-Ser toxin was eluted in close proximity to an 11-residue peptide, substance-P (molecular mass of 1348 Da), but the L-form toxin was eluted at a similar position to aprotinin (molecular mass of 6512 Da). Under the conditions used, each toxin was eluted as a single peak at the same position at a loading concentration from 2 ^M to 200 ^M, whereas aggregated or oligomeric forms were observed at the concentration of 2 mM.

Tomohiro Watanabe et al

548

O GO

^

co-[L-Ser^^]Aga-TK

0)

u

a o

67kDa I

43kDa 25kDa

14kDa TkDa I

(0-[i>Ser46]Aga-TK

10

20

I

30

40

50

60

Time (min) Figure 2. Size-exclusion chromatography of (o-[D-Ser^^]Aga-TK and co-[LSer46]Aga-TK. The two toxins (200 |iM) were analyzed en a Superdex HR75 column equilibrated with phosphate-buffered saline, pH 7.4, as described under Experimental Procedures. The molecular masses and elution positions of bovine serum albumin (67 kDa), ovalbumin (43 kDa), chymotrypsinogen (25 kDa), ribonuclease A (14 kDa), and aprotinin (7 kDa), used as calibration standards, are shown.

Role of D-Ser46 in co-Agatoxin-TK

549

The calibration of the size-exclusion column with standard proteins demonstrated that the L-Ser toxin has an apparent molecular mass of 6 kDa, which is close to the real molecular mass of the toxin. The apparent molecular mass of the D-Ser toxin was too small to evaluate accurately from the calibration data. These results indicated that both ©-[D-Ser^^] Aga-TK and co-[L-Ser^6] Aga-TK take monomeric form at physiological pH and ionic strength, but the two toxins are significantly different in apparent molecular mass. The apparent molecular mass of D-Ser toxin was dramatically increased by the addition of guanidine hydrochloride to the elution buffer, although that of the L-Ser toxin was not altered by the denaturing reagent. In the presence of 5.2 M guanidine hydrochloride, the D-form toxin was eluted at the same position as the L-form toxin and the apparent molecular masses of the two toxins were estimated as 6 kDa based on calibration with the standard proteins. CD and fluorescence spectroscopic analyses revealed that the two toxins were unfolded and lost their secondary and tertiary structure in 5.2 M guanidine hydrochloride at pH 7.4, as described below. It, therefore, appears that the D-Ser toxin forms a compact folded structure, whereas the L-Ser toxin has a relatively unfolded or extended structure. In order to see whether or not the elution behavior of the two toxins depends on the specificity of the separation support used, the two toxins were also analyzed on a TSK GSOOOSWXL column under the same elution conditions as those of the Superdex column. Similar results were obtained, i.e., the D-form toxin was eluted later than the L-form toxin with phosphate-buffered saline. These results confirm that the different elution behavior of the two toxins was caused by the distinct molecular shapes of the two toxins.

JB. Conformational analyses of o}-[D-Ser^^]Aga-TK and co[L'Ser^^lAga-TK We examined the CD spectra of co-[D-Ser^^]Aga-TK and CO-[LSer46]Aga-TK in phosphate-buffered saline, pH 7.4, to compare the secondary structures of the two toxins. As illustrated in figure 3, the spectrum of the D-Ser toxin showed a negative peak at 208 n m , while the spectrum of the L-Ser toxin had both a negative peak at 200 nm and broad positive ellipticity centered near 220 nm.

Tomohiro Watanabe et al

550

2000

o

S

-5000

-9000

220 Wavelength (nm)

250

Figure 3. CD spectra of co-[D-Ser'*^]Aga-TK and co-[L-Ser'*^]Aga-TK in phosphatebuffered saline, pH 7.4, in the presence or absence of guanidine hydrochloride. 1, co-[D-Ser^^]Aga-TK in phosphate-buffered saline; 2, co-[L-Ser^^]Aga-TK in phosphate-buffered saline; 3, co-[L-Ser^^]Aga-TK in phosphate-buffered saline containing 5.2 mM guanidine hydrochloride; 4, ©-[D-Ser'^^JAga-TK in phosphatebuffered saline containing 5.2 mM guanidine hydrochloride. CD spectra were recorded between 210 and 250 nm or 195 and 250 nm in the presence or absence of guanidine hydrochloride, respectively.

Role of D-Ser46 in w-Agatoxin-TK

551

These features are characteristic of peptide random coil and p-sheet structures, and the magnitude of the positive ellipticity band revealed a significant difference in p-sheet contents between the two toxins. The secondary structures of the two toxins were found to be disrupted by the addition of 5.2 M guanidine hydrochloride at pH 7.4, since the spectra changed to a pattern typical of predominantly random coil structure. It was concluded that CO-[DSer46]Aga-TK has a significantly higher p-sheet content than CO-[LSer46]Aga-TK under neutral conditions. Intrinsic fluorescence of co-[D-Ser46]Aga-TK and co-[L-Ser46]AgaTK was determined to compare the tertiary structures around the Trp and Tyr residues between the two toxins. The two toxins have a single residue each of Trp and Tyr in the disulfide-rich region, at positions 14 and 9, respectively. As shown in figure 4, tryptophan fluorescence with an emission maximum near 345 nm was strongly quenched in the D-Ser toxin, but not in the L-Ser toxin, whereas tyrosine fluorescence of the two toxins showed almost the same intensity at the emission maximum of 310 nm. Further, the intensity of tryptophan fluorescence of the D-form toxin, but not that of the L-form toxin, increased concomitantly with the increase of the concentration of guanidine hydrochloride at pH 7.4. These results clearly indicate that the Trp^^ residue in the L-form toxin is exposed to the solvent, but this residue of the D-form toxin is in a relatively hydrophobic environment. Previously, Yu et al. reported the solution structure of ©-[o-Ser^^JAga-TK at pH 4.0, showing that the indole side chain of the Trp^^ residue packs against the sulfur atoms of Cys^^-Cys^ and may serve to stabilize the loop formed by the disulfide bond (11). It is, therefore, suggested that the tryptophan fluorescence of the D-Ser toxin may be quenched by the sulfur atoms of the disulfide bond, whereas the indole chromophore of the L-Ser toxin may not be affected by the sulfur atoms due to the greater distance between the two groups. In conclusion, we have investigated the conformation of CO-[DSer46]Aga-TK and co-[L-Ser4^]Aga-TK at physiological pH and ionic strength using size-exclusion chromatography and spectroscopic methods. We have found that the apparent molecular mass of co[D-Ser^^]Aga-TK is significantly smaller than that of co-[L-Ser^^]AgaTK as determined by size-exclusion chromatography. CD spectra of the two toxins also revealed that co-[D-Ser4^]Aga-TK has a higher psheet content than co-[L-Ser46]Aga-TK. Furthermore, the intrinsic fluorescence of ©-[o-Ser^^JAga-TK showed that Trp^^ of CO-[DSer^6]Aga-TK is in a relatively hydrophobic environment compared with that of ©-[L-Ser^^JAga-TK. These data imply that

Tomohiro Watanabe et al

552 3727

30001 2000-1

1000-1 0.000 u O

250.0

300.0

350.0 (Emission)

400.0

300.0

350.0 (Emission)

400.0

3727

0.000 250.0

450.0

450.0

Wavelength (nm) Figure 4. Intrinsic fluorescence spectra of co-[D-Ser^6]Aga-TK and co-[L-Ser^^]AgaTK in phosphate-buffered saline, pH 7.4. Emission spectra were recorded between the wavelengths of 250 and 450 nm at t h e excitation wavelength of 280 nm.

Role of D-Ser46 in w-Agatoxin-TK

553

the D-Ser^^ residue of co-[D-Ser^^]Aga-TK may be involved in the formation of additional intramolecular p-sheet structure in the carboxyl-terminal region or between the disulfide core and carboxyl-terminal tail, which contributes to the compact folding of co-[D-Ser4^]Aga-TK. It is also likely that the additional p-sheet causes a change in the tertiary environment around the Trp^^ residue of co-[D-Ser4^]Aga-TK. Additional experiments to assess the biological importance of the carboxyl-terminal tail seem worthwhile. For instance, it would be interesting to examine the effects of sequential truncation of the carboxyl-terminal region of co-[D-Ser4^]Aga-TK on the blockade of the P-type calcium channels. Studies are in progress to characterize further the carboxyl-terminal conformation of co-[D-Ser46] Aga-TK.

Acknowledgments We thank Dr. Kozaki for helpful discussions and Dr. Takakuwa (Jasco), for measuring CD.

References 1. 2.

3. 4. 5.

6.

7.

8.

Olivera B.M., Miljanich G.P., and Ramachandran J. (1994) Annu. Rev. Biochem. 63, 823-8671. Niidome, T., Teramoto, T., Murata, Y., Tanaka, I., Seto, T., Sawada, K., Mori, Y., and Katayama, K. (1994) Biochem. Biophys. Res. Comtnun. 203, 1821-1827 Kimura, M., Yamanishi, Y., Hanada, T., Kagaya, T., Kuwada, M., Watanabe, T., Katayama, K., and Nishizawa, Y. (1995) Neuroscience 66, 609-615 Teramoto, T., Niidome, T., Miyagawa, T., Nishizawa, Y., Katayama, K., and Sawada, K. (1995) NeuroReport 6, 1684-1688 Kuwada, M., Teramoto, T., Kumagaye, K. Y., Nakajima, K., Watanabe, T., Kawai, T., Kawakami, Y., Niidome, T., Sawada, K., Nishizawa, Y., and Katayama, K. (1994) Mol Pharmacol. 46, 587-593 Kozaki, T., Kuwada, M., Narukawa, M., Nagai, Y., and Asakawa, N. (1996) in "Peptide Chemistry 1995" (Nishi, N., ed) Protein Research Foimdation, Osaka, 245-248 Watanabe, T., Teramoto, T., Kuwada, M., Shikata, Y., Niidome, T., Kawakami, Y., Sawada, K., Nishizawa, Y., and Katayama, K. (1995) in "Peptide Chemistry 1994" (Ohno, M., ed) Protein Research Foundation, Osaka, 253-256 Heck, S. D., Siok, C. J., Krapcho, K. J., Kelbaugh, P. R., Thadeio, P. P., Welch, M. J., Williams, R. D., Ganong, A. H., Kelly, M. E., Lanzetti, A. J., Gray, W. R., Phillips, D., Parks, T. N., Jackson, H., Ahlijanian, M. K., Saccomano, N. A., and Volkmann, R. A. (1994) Science 266,1065-1068

554 9. 10. 11. 12. 13.

Tomohiro Watanabe et al Shikata, Y., Watanabe, T., Teramoto, T., Inoue, A., Kawakami, Y., Nishizawa, Y., Katayama, and K., Kuwada, M. (1995) /. Biol Chem. 270, 16719-16723 Adams, M. E., Mintz, I. M., Reily, M. D., Thanabal, V., and Bean, B. P. (1993) Mol Pharmacol 44, 681-688 Yu, H., Rosen, M. K., Saccomano, N. A., Phillips, D., Volkmann, R. A., and Schreiber, S. L. (1993) Biochemistry 32, 13123-13129 Teramoto, T., Kuwada, M., Niidome, T., Sawada, K., Nishizawa, Y., and Katayama, K. (1993) Biochem. Biophys. Res. Commun. 196, 134-140 Watanabe, T., Shikata, Y., Oda, Y., Nishizawa, Y., Kuwada, M., and Asakawa N. The two dimensional HPLC purification of biologically active polypeptides and polyamines in funnel web spider venom, manuscript in preparation

Involvement of Basic Amphiphilic a-helical Domain in the Reversible Membrane Interaction of Amphitropic Proteins: Structural Studies by Mass Spectrometry, Circular Dichroism, and Nuclear Magnetic Resonance Nobuhiro Hayashi, Mamoru Matsubara, Koiti Titani, and Hisaaki Taniguchi Division of Biomedical Polymer Science, Institute for Comprehensive Medical Science, Fujita Health University, Toyoake, Aichi 470-11, Japan

I.

Introduction

A growing number of proteins have been shown to belong to the socalled "amphitropic" proteins which are neither "pure" membrane proteins nor soluble proteins (1). Interestingly, many of them are involved in the signal transduction, and their stimulation-dependent translocation plays important roles in the transmission of signals between the plasma membrane and the nucleus (2). They usually lack any apparent hydrophobic membrane-binding domain, but the importance of highly basic domains in the Src family proteins (2) and that of the basic amphiphilic domain in MARCKS (3) in the membrane association have been well established. In the latter case, the direct phosphorylation of the domain by protein kinase C regulates the reversible membrane association of MARCKS (3, 4). It is of interest to note that some of these amphitropic proteins are fatty acylated, and the modification is also involved in the membrane interaction (2, 5). One of the major phosphoproteins in neuronal growth cone, GAP-43 (growth-associated protein-43, also known as B50, Fl, P56, or neuromodulin), which is found associated with membrane cytoskeletal fractions (6), is very hydrophilic and lacks any apparent hydrophobic membrane-binding domain (7). Palmitoylation of two cysteine residues near the N-terminus has been assumed to be involved in the interaction with membranes (8, 9). However, we have recently shown that GAP-43 isolated from the membrane fractions is notpalmitoylated at all but still retains the ability to bind phospholipid membranes in vitro (10, 11). GAP-43 belongs to the MARCKS family of acidic hydrophilic membrane-associated proteins (12) and has a similar basic amphiphilic domain which serves as the calmodulin-binding domain and the phosphorylation domain by PKC. The involvement of the domain in the membrane-anchoring of GAP-43 has, in fact, been suggested (13, 14). In the present study, we first show a detailed mass spectrometric TECHNIQUES IN PROTEIN CHEMISTRY VIII Copyright © 1997 by Academic Press All rights of reproduction in any form reserved.

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analysis on the posttranslational modifications of GAP-43, which provides the basis for the understanding of the structures of molecules involved. The interaction of GAP-43 and that of the basic amphiphilic domain with membrane phospholipids are then studied using circular dichroism (CD) and nuclear magnetic resonance (NMR) to understand the underlying structural mechanisms in the interaction.

II. Materials and Methods A.

Materials

GAP-43 (10) and PKC (15) were purified from bovine brain as described previously. A peptide (QASFRGHITRKKLKGEK) corresponding to the calmodulin-binding domain of GAP-43, named GAP peptide, was synthesized using conventional tBoc chemistry in an ABI 430A peptide synthesizer (Applied Biosystems), and purified over a C18 reversedphase column (Vydac 218TP1010, The Separations Group) using a linear HjOacetonitrile gradient in the presence of 0.1% trifluoroacetic acid. Lipids purchased from Avanti Polar Lipids were suspended in 5 mM phosphate buffer (pH 7.5), and sonicated in a BRANSON SONIHER 250 sonicator for 30 min. The supernatant obtained after centrifugation in a tabletop centrifuge for 20 min was used as unilamellar liposomes. B.

Preparation

of Phosphorylated

GAP'43 and GAP

Peptide

Phosphorylation of the intact GAP-43 and GAP peptide by PKC was carried out in the reaction buffer (25 mM Tris-HCl buffer (pH7.5), 10 mM MgClj, 100 mM CaCl2, 80 |ig/ml phosphatidylserine, 8 M-g/ml dioleoyl glycerol, ImM ATP) at35°C for 90 min, and was stopped by adding 0.1% final concentration of trifluoroacetic acid. The extent of the phosphorylation was analyzed by mass spectrometry as described previously (10, 16). The phosphorylated GAP peptide was purified over a reversed-phase HPLC column. The phosphorylated GAP-43 protein was purified by ion exchange chromatography on a mono Q column (HR 5/5) using a linear gradient of NaCl (0 - 0.5M) in 20 mM Tris-HCl buffer (pH 7.5) containing 1 mM EDTA and 1 mM dithiothreitol. C.

Mass Spectrometrie

Analysis

Electrospray mass spectra were recorded in a PE Sciex API-Ill mass spectrometer as described previously (10, 16). A capillary HPLC was connected on-line to the electrospray interface of the mass spectrometer. D.

Circular Dichroism

CD spectra

were

(CD)

recorded

Spectrometry at

25°C

in

a JASCO

J-720

CD

Membrane Structure of GAP-43 Peptide

557

spectropolarimeter using a 0.1 cm cell. Concentration of the peptide was 20 }iM in 5mM phosphate buffer (pH7.3). The contents of secondary structures were calculated from the CD spectra using a CONTIN program (17) modified by Dr. F. Arisaka, Tokyo Institute of Technology.

E. NMR Spectrometric Analysis 500MHz proton NMR spectra were recorded on a Bruker DMX-500 spectrometer. Chemical shifts were measured relative to the methyl resonance of an internal reference, 4,4-dimethyl-4-silapentane-lsulfonate. GAP peptide (5mM) was dissolved in 90% H ^ - 10% D p , 99.98% D p , 50% H P -10% D p - 40% trifluoroethanol (TFE)-d3, or 60% D2O - 40% TFE-d3. The pH of the samples was 4.0 (direct meter reading). By using standard procedures for 2D proton NMR of proteins (18), the sequence-specific assignment of resonances was obtained from two-dimensional TOCSY (19), NOESY (20, 21), DQF-COSY with phase cycling (22) or with pulsed field gradient (23, 24), and TQF-COSY with pulsed field gradient (23, 24) spectra. All spectra were acquired at 25°C in the phase-sensitive mode using the time proportional phase increment technique. WATERGATE (25, 26) or presaturation was used for the water suppression. A total of 512 measurements with increasing t^ values were made, and 64 transients were accumulated for each measurement. For tj 2048 data points were taken, and the spectral widths along ^2 ^ ^ ^ 5000 Hz. The data were zero filled once in the f^ dimension. A cosine window function and a Gaussian function were used in f^ and fj dimension before Fourier transformation, respectively. For the NOESY spectra, the time-domain data were multiplied by Gaussian functions in both dimensions. All spectra were processed using Bruker XWIN-NMR or MSI Felix95.0 software packages. III.

Results and Discussion

A. Mass Spectrometric Analysis on the in Vivo Posttranslational Modifications Soft ionization techniques such as electrospray ionization and matrix assisted laser desorption are now routinely used to determine the mass of large hydrophilic polymers like proteins (27). However, as is usual for the ionization process, the presence of salts and detergents, which is common for biological samples, can affect the process significantiy. The use of the on-line capillary reversed-phase HPLC in combination of the electrospray mass spectrometer (LC/MS) has made it possible to analyze such samples directly (10,16, 28). When GAP-43 isolated from the membrane fractions of bovine brain was analyzed, a single major peak with a minor peak corresponding to a phosphorylated species was observed (Fig. la). To study the posttranslational modifications in detail, the protein was digested with specific proteases such as lysyl

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25145.0

>^

24,600

25,000

a

25,400

25,800

Mass (Da)

750

800

m/z

850

900

Fig. 1. Mass spectrometric analysis of GAP-43 purified from membrane fractions of bovine brain, (a) A deconvoluted mass spectrum of GAP-43. A deconvoluted mass spectrum of the N-terminal peptide before reduction (b) and after reduction (c). Peaks formed by oxidation of Met were also observed.

endoprotease and trypsin, and the resulting mixtures were directly analyzed with the same LC/MS apparatus. Since the cDNA sequence has been known, most of the peptides detected could be assigned solely from their masses, and the two peptides containing phosphorylation and a peptide corresponding to the N-terminal peptide were observed (10). Interestingly, the mass of the latter (796.3 Da) was slightly but significantly lower than the theoretical mass of 798.3 Da. Since the peptide contained two successive Cys residues, the peptide was treated with dithiothreitol, and directly analyzed with the LC/MS apparatus. As shown in Fig. lb, c, the mass of the peptide increased by 2 Da after the dithiothreitol treatment, suggesting that the two Cys residues form an intrachain disulfide bridge. Since no palmitoylated N-terminal peptide was detected to significant extent, we conclude that the isolated GAP-43 is not palmitoylated at the two Cys near the N-terminus. B. Conformational Change of GAP-^3 Phospholipid Binding

and GAP Peptide

upon

GAP-43 purified from bovine brain showed a CD spectrum with a single

Membrane Structure of GAP-43 Peptide

559

negative peak at around 197 nm in aqueous solution, which is typical for a random structure. At most 10% of the whole molecule seems to assume a-helix. Upon addition of acidic phospholipids such as phosphatidylglycerol (PG), however, a broad negative peak between 220 and 230 nm due to the increase in the a helix content was observed (Fig. 2a). All the acidic phospholipids tested but not neutral phospholipid such as phosphatidylcholine affected the CD spectrum in a similar way. A peptide corresponding to the calmodulin-binding domain of GAP-43 (GAP peptide) showed a similar random coil to a-helix conformational change upon phospholipid binding (Fig. 2b). The extents of the change in the CD spectra of the intact protein and the peptide are comparable, suggesting that only the domain interacts with the lipids and undergoes a conformational change to a-helix. This is reasonable, since the whole molecule of GAP-43 except for the calmodulin binding domain is hydrophilic and acidic without any hydrophobic amino acids. When ionic strength of the buffer was increased, the apparent affinity between the GAP peptide and the phospholipids decreased, suggesting that the interaction between the GAP peptide and the phospholipids involves electrostatic interaction (29, 30). The addition of TFE, a membrane

O

260

X3

o^ q

-4.0'

200

220

240

Wavelength (nm)

260

F^. 2. Effects of phospholipids on CD spectra of GAP-43 and GAP peptide. CD spectra of GAP-43 (a) and GAP peptide (b) were measured in the absence (O) and in the presence (•) of phosphatidylglycerol or phosphatidylcholine (A).

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mimicking reagent, caused a concentration dependent induction of the CD spectrum component typical for an a-helix. The a-helical content reached almost 100% in the presence of 40% TFE.

C.

Structural Analysis by Nuclear Magnetic

Resonance

The structural characteristics of the domain was further studied in detail by NMR techniques. Compared to the CD spectrometry, the NMR method gave more accurate and residue-specific information on the conformation. Large portion of the synthetic peptide formed a regular a-helix in the presence of TFE, as was evidenced by the consecutive NOE connectivities (Fig. 3a) (18). Fig. 3a shows that rather strong medium range ^H-^H NOE's of both ap(i, i+3) and aN(i, i+3) are detected in the region from Phe"^ to Lys^^ Furthermore, compared to the chemical shifts of a protons observed in GAP peptide with those obtained in random structure peptide (31), the characteristic upfield shifts of the a protons of GAP peptide except for those of two residues near the C terminus were observed (Fig. 3b). This feature is observable with a helical structure regions (32,33). These results indicate that the region (Phe^-Lys^^) forms a "regular" a-helix in the presence of TFE.

aN(i,i+1)

Q1A2 S3 F 4 R 5 G 6 H 7 |8 T9 R i ( K i i K i l i 3 K i 4 G i t i 6 K i 7

-0.60

-0.40

-0.20

0.00

Membrane Structure of GAP-43 Peptide

561

In the absence of TFE, GAP peptide showed a typical CD spectrum for a random structure (Fig. 2b). Due to resonance overlaps, many peaks in the NMR spectra could not be uniquely assigned except for several peaks. However, as is shown in Fig. 3c, the signals of a protons generally showed characteristic upfield shifts again, although the degrees were not so large as those obtained in the presence of TFE. Because Ala^ Ile^ Thr^ and Leu^^ each occurs only once in GAP peptide, and their methyl group signals give well-resolved signals in higher magnetic field region, it was possible to assign these residues. Interestingly, the a proton chemical shifts of all the assigned residues showed intermediate values between those typical for random coil and those for a helix obtained as above (Fig. 3b). Since the resonance overlaps observed is characteristic

Gly in random coil

CO

c^ Q_

Signals observed in a helix

X

00

Lys,Arg,Gln,Ser,His,Phe in random coil 8.7

8.4 8.1 F2 (ppm)

7.8

Fig. 3. NMR analysis of GAP peptide, (a) NOE connectivities of specified proton pairs observed in theNOESY spectra of GAP peptide in the presence of 40 % TFE are marked with a bar (aN(i,i+l)), open (aP(i,i+3)) and/or shaded (aN(i,i+3)) boxes, (b) Deviation in the chemical shifts of some of the a protons in the presence of TFE (open bars) and in the absence of TFE (shaded bars) from those observed with typical random coil (31) are indicated, (c) ocH (F1)/NH (F2) region of 500 MHs DQF-COSY spectrum of GAP peptide in 90 % H2O -10 % DjO. The regions, in which Lys, Arg, Gin, Ser, His, Phe, and Gly in random coil are observed, are indicated.

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for a random cx3il, and chemical sifts of a protons showed intermediatevalues between those of random coil and those of a helical structure, the GAP peptide in aqueous solution assumes an intermediate state between a random coil and a regular a helix. Such a "nascent" helical structure may deviate from ideal geometry, and/or the ends of the a-helix can fray (34, 35). The interaction of GAP peptide with phospholipids seemed to stabilize the conformation to induce an a helix, as is often the case of "nascent" a-helical structures which are usually induced or further stabilized by addition of the a-helix promoting solvent TFE (36,37).

IV.

Conclusions

GAP-43 lacks any hydrophobic region found in usual membrane proteins and the pal mi toy la ti on which has been implicated in the membrane anchoring is not present in the purified protein. However, the effector domain of basic amphiphilic nature has the ability to bind acidic phospholipids. The domain adopts an a helical conformation when put into hydrophobic environments as shown by the CD and NMR analyses. A growing body of evidence suggests that the basic amphiphilic a-helical domain, which has been initially found as a calmodulin binding motif, serves as a reversible membrane-association signal.

References 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13.

Burn, P. (1988) Trends Biochem. Sci. 13, 79-83. Resh, M. D. (1994) Cell 6,411-413. Taniguchi, H., and Manenti, S. (1993) /. Biol. Chem. 268,99609963. Kim, J., Shishido, T., Jiang, X., Aderem, A., and McLaughlin, S. (1994) /. Biol. Chem. 269, 28214-22821. Peitzsch, R. M., and McLaughlin, S. (1993) Biochemistry 32, 10436-10443. Meiri, K. P., and Gordon-Weeks, P. R. (1990) /. Neurosci. 10, 256-266. LaBate, M. E., and Skene, J. H. P. (1989) Neuron 3, 299-310. Zuber, M. X., Strittmatter, S. M., and Fishman, M. C (1989) Nature 341,345- 348. Skene, J. H. P., and Virag, I. (1989) /. Cell Biol. 108,613-624. Taniguchi, H., Suzuki, M., Manenti, S., and Titani, K. (1994) /. Biol. Chem. 269, 22481-22484. Hayashi, N., Matsubara, M., Titani, K., and Taniguchi, H. (1996) in preparation. Blackshear, P. J. (1993) /. Biol. Chem. 268,1501-1504. Houbre, D., Duportail, G., Deloulme, J. C, and Baudier, J.

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(1991) /. Biol Chem. 266, 7121-7123. Kim, J., Blackshear, P. J., Johnson,}. D., and McLaughlin, S. (1994) Biophys. ]. 67, 227-237. Manenti, S., Sorokine, O., Van Dorsselaer, A., and Taniguchi, H. (1992) /. Biol Chem. 267,22310-22315. Taniguchi, H., Manenti, S., Suzuki, M., and Titani, K. (1994) /. Biol Chem. 269,18299-18302. Provencher, S. W., and Glockner, J. (1981) Biochemistry 20,3337. Wiithrich, K. (1986) NMR of Proteins and Nucleic Acids, J. Wiley, New York. Bax, A., and Davies, D. G. (1985) /. Magn. Reson. 65,393-402. Jeener, J., Meier, B. H., Bachman, P., and Ernst, R. R. (1979) /. Chem. Phys. 71,4546-4553. Macura, S., Hyang, Y., Suter, D., and Ernst, R. R. (1981) /. Magn. i^eson. 43,259-281. Ranee, M., Sorensen, O. W., Bodenhausen, G., Wagner, G., Ernst, R. R., and Wiithrich, K. (1983) Biochem. Biophys. Res.Commun. 177,479-485. Baker, P., and Freeman, R. (1985) /. Magn. Reson. 64,334-338. Hurd, R. E. (1990) /. Magn. Reson. 87,422-428. Piotto, M., Saudek, V., and Sklenar, V. (1992) /. Biomol NMR 2,661-665. Sklenar, V., Piotto, M., Leppik, R., and Saudek, V. (1993) /. Magn. Reson. 102 (Ser. A), 241-245. Biemann, K. (1992) Annu Rev Biochem 61, 977-1010. Taniguchi, H. (1996) /. Mass Spectrm. Soc. Japan 44,443-457. McLaughlin, S. (1977) Curr. Top. Membr. Transp. 9,1-144. McLaughlin, S. (1989) Annu. Rev. Biophys. Biophys. Chem. 18, 13-136. Bundi, A., and Wuthrich, K. (1979) Biopolymers 18, 285-298. Pastore, A., and Saude, V. (1990) /. Magn. Reson. 90,165-176. Wishart, D., Sykes, B., and Richards, F. (1991) /. Mol Biol 222, 311-333. Dyson, H. J., Merutka, J., Waltho, J. P., Lerner, R. A., and Wright, P. E. (1992) /. Mol Biol 226, 795-817. Manning, M. C , Illangasekare, M., and Woody, R. W. (1988) Biophys. Chem. 31, 77-86. Munier, H., Blanco, F. ]., Precheur, B., Diesis, E., Nieto, J. L., Craescu, C. T., and Barzu, O. (1993) /. Biol Chem. 268,16951701. Shang, M., and Vogel, H. J. (1994) /. Biol Chem. 269, 981-985.

Acknowledgements We thank Mr. M. Suzuki for technical assistance. This work was supported in part by Grants-in-Aid from the Fujita Health University, Science Research Promotion Fund from the Japan Private School

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Promotion Foundation, Research Grant from the Naito Foundation for Medical Research, Grant-in-Aid for Scientific Research (C) (06680773) and Grants-in-Aid for Scientific Research on Priority Areas (06253218, 06276218, 07268221,07279242, 08249240 and 08260220) from the Ministry of Education, Science and Culture, Japan. M.M is a Research Fellow of the Japan Society of the Promotion of Science.

One-Dimensional Diffusion of a Protein along a Single-Stranded Nucleic Acid Bradley R. Kelemen Ronald T. Raines Department of Biochemistry University of Wisconsin Madison, WI 53706-1569

I. Introduction One-dimensional diffusion can accelerate the formation of site-specific interactions within biopolymers by up to lO^-fold (Berg et aL, 1981). Such facilitated diffusion is used by transcription factors and restriction endonucleases to locate specific sites on double-stranded DNA (von Hippel and Berg, 1989). The backbone of RNA, like that of DNA, could allow for the facilitated diffusion of proteins. Yet, the facilitated diffusion of a protein along RNA (or any single-stranded nucleic acid) has not been demonstrated previously. Bovine pancreatic ribonuclease A (RNase A; RNA depolymerase; EC 3.1.27.5) is a distributive endoribonuclease that catalyzes the cleavage of the P-O5' bond of RNA on the 3' side of pyrimidine residues. RNase A binds to polymeric substrates (Imura et aL, 1965; Trie et al., 1984; Moussaoui et al., 1995), but the mechanism by which RNase A locates a pyrimidine residue within a polymeric substrate is not known. Binding to phosphoryl groups is important for the one-dimensional diffusion of proteins along DNA (Winter et al, 1981), and may likewise provide nonspecific interactions necessary to generate one-dimensional diffusion by RNase A. RNase A has three defined phosphoryl group binding subsites, PO, PI, and P2, as well as three base binding subsites, Bl, B2, and B3 (Pares et al, 1991). The subsite interactions in the RNase A