Possible Roles for His 208 in the Active-Site Region of Chloroplast Carbonic Anhydrase fromPisum sativum

Archives of Biochemistry and Biophysics Vol. 361, No. 1, January 1, pp. 17–24, 1999 Article ID abbi.1998.0961, available online at http://www.idealibrary.com on

Possible Roles for His 208 in the Active-Site Region of Chloroplast Carbonic Anhydrase from Pisum sativum Harry Bjo¨rkbacka, Inga-Maj Johansson*, and Cecilia Forsman 1 Department of Biochemistry, Umeå University, S-901 87 Umeå, Sweden; and *UKBF, 3B 2nd Floor, Umeå University Hospital, S-901 85 Umeå, Sweden

Received July 2, 1998, and in revised form September 29, 1998

His 208 of chloroplast pea carbonic anhydrase is conserved among the dicotyledonous carbonic anhydrases. This His was replaced by an Ala to test whether a histidine residue at this position, in analogy to His 64 of human carbonic anhydrase II, acts as an internal proton shuttle. Values of the kinetic parameters k cat and k cat/K m for the H208A mutant enzyme are high over the pH range 6 –9 and of the same magnitude as those for the wild-type enzyme, indicating that this residue is not crucial for the catalytic event. The pH dependence of k cat/K m, reflecting the Zn–H 2O ionization, is, however, simplified to that of a simple titration with pK a 5 7.1 6 0.1 in the absence of His 208. Interaction with the proton-accepting buffer molecule is impaired in the mutant, and apparent K m values for the buffer have increased up to 20 times. Furthermore, the H208A mutant is more easily inactivated by oxidation than the wild-type enzyme. The results indicate that the pK a for a redox-sensitive Cys residue is decreased by at least one pH unit in the mutant, and the histidyl side chain seems to have a function in stabilizing the reduced and active form of the enzyme. An interaction with the redox-sensitive cysteines at positions 269 and 272 is proposed. © 1999 Academic Press Key Words: carbonic anhydrase; chloroplast; enzyme kinetics; proton transfer; redox.

Carbonic anhydrase (CA; EC 4.2.1.1) 2 is a zinc-containing enzyme which catalyzes the reversible hydra1 To whom correspondence should be addressed. Fax: 146 90 7867661. E-mail: [email protected]. 2 Abbreviations used: CA, carbonic anhydrase; CD, circular dichroism; DTT, dithiothreitol; TCEP, tris(2-carboxyethyl)phosphine; Taps, 3-[tris(hydroxymethyl)methylamino]-2-hydroxypropanesulfonic acid; Mops, 4-morpholinepropanesulfonic acid; Mes, 4-morpholineethanesulfonic acid; Mopso, 3-(N-morpholino)-2-hydroxypropanesulfonic acid; 2-ME, 2-mercaptoethanol; pPCAs, plasmid containing cDNA encoding the mature subunit of pea carbonic anhydrase.

0003-9861/99 $30.00 Copyright © 1999 by Academic Press All rights of reproduction in any form reserved.

tion of CO 2. From sequence homologies, the CAs are found to belong to three genetically distinct families, designated a-, b-, and g-CA (1). The a-family is dominated by all known animal CAs, but it also contains CAs from the unicellular alga Chlamydomonas rheinhardtii and some prokaryotic CAs (2– 4). The most extensively studied CAs belong to this family. b-CAs have been shown to be present in both C 3 and C 4 monoand dicotyledonous plants (5). They are also found in other photosynthesising organisms (6 –9) and in certain nonphotosynthesizing prokaryotes (10, 11). g-CA is the most recently identified gene family, and the only isolated enzyme belonging to this family is from the archaeon Methanosarcina thermophila (12). According to primary structures, the b-CA family can be divided into three groups representing monocotyledons, dicotyledons, and prokaryotes, with a high degree of sequence homology within each group. The three groups differ in their quaternary structures. The enzyme from C 3 dicotyledonous plants has been shown to be a homooctamer (13, 14) with no evidence for covalent linkage between the subunits. CA from monocotyledonous plants has been suggested to be a dimer (15, 16), and the prokaryotic CA from Escherichia coli is reported to be a dimer or a tetramer, depending on experimental conditions (10). Most biochemical studies on b-CAs have been done on CAs from C 3 dicotyledons, where the enzyme is localized in the chloroplast stroma, directed to the plastid by an N-terminal chloroplast transit peptide (17). There is no three-dimensional structure known for a b-CA. However, structural studies of CA from pea and hybrid aspen using circular dichroism (CD) indicate that a-helix is the dominating secondary structure (14, 18). This is in contrast to both the a- and g-CAs, which are composed mainly of large b-sheets, according to their X-ray crystal structures (19, 20). Also, the binding of the zinc ion differs among the different CA families. In both a- and 17

¨ RKBACKA, JOHANSSON, AND FORSMAN BJO

18

g-CAs three His residues and a water molecule constitute the ligands, giving an almost tetrahedral coordination geometry. Extended X-ray absorption fine structure analysis of spinach CA suggests a Cys–His–Cys– H 2O ligand scheme (21, 22). This hypothesis is supported by the presence of two cysteines (Cys 159 and Cys 222 in pea CA) and one histidine (His 219 in pea CA) that are conserved in all b-CA sequences. Several of the CAs are very efficient enzymes, and the kinetic properties of human isozymes I, II, and III have been extensively investigated (see (23) for a review). Among plant CAs, kinetic studies have been done on CA from pea (24, 25), spinach (22, 26), and parsley (27). They all have a high catalytic efficiency with k cat values between 10 5 and 10 6 s 21 and k cat/K m values of 10 7–10 8 M 21 s 21 at high pH. The kinetic data can essentially be fit into the general zinc-hydroxide mechanism proposed for the high-activity a-CA isozymes (28). The catalytically active group in this mechanism model is the zinc-bound water, which ionizes to a hydroxide ion. In the CO 2-hydration reaction, the basic form of the enzyme is active, while the reverse reaction needs the protonated form. The pH profiles of k cat and k cat/K m for human CA II both follow a simple titration curve with a pK a close to 7. According to the proposed mechanism, the catalytic event can be divided into two stages. The first stage is the interconversion between CO 2 and HCO 32 (Eq. [1]), the rate of which is related to k cat/K m. H 2O

EZn 2 OH 2 1CO2 7 EZn 2 HCO 2 3 43 EZn 2 H2O 1 HCO 2 3

[1]

B

EZn 2 H2O 7 H 1 2 EZn2OH 2 43 EZn 2 OH 2

[2]

BH 1

The second stage (Eq. [2]) is the regeneration of the active form of the enzyme, involving the transfer of a proton between the zinc-bound water molecule and surrounding medium. In human CA II this occurs in two steps, in which the amino acid His 64 acts in shuttling the proton between the active site and bulk buffer. At high buffer concentrations the intramolecular step (the first part of Eq. [2]) is rate limiting, while at low buffer concentrations the proton transfer to the buffer molecule will be rate determining (second part of Eq. [2]). Consequently, Eq. [2] is always reflected by the kinetic parameter k cat. The buffer thus participates in the reaction as a second substrate, resulting in a ping-pong mechanism. This means that the first stage of the reaction (Eq. [1]) is independent of the buffer concentration. Most of the kinetic properties observed for higher plant CAs are consistent with this mechanism. How-

ever, there are also data that are not easily explained by the model, indicating that some parts of the mechanism might differ from that of a-CA II. For pea CA, solvent hydrogen isotope effects on k cat around 2 have been found at pH 6 –9 (25), suggesting that a step involving proton transfer is at least partly rate determining. If the mechanism outlined in Eqs. [1] and [2] holds for plant CA, the parameter k cat/K m would be unaffected by changing the solvent from H 2O to D 2O. This seems to be the case at high pH, but at pH 6 and 7 there is an isotope effect on k cat/K m of 2.5–3, indicating that part of the mechanism may vary with pH. Furthermore, high buffer concentrations are needed to obtain saturating conditions for pea CA. Treating the buffer as a substrate, apparent K m values for the buffer between 4 and 185 mM have been reported (24). Notably, the K m values showed a pH dependence with the highest values obtained at high pH (3-[tris(hydroxymethyl)methylamino]-2-hydroxypropanesulfonic acid (Taps buffer), and decreasing as the pH was lowered (4-morpholinepropanesulfonic acid (Mops) and 4-morpholineethanesulfonic acid (Mes) buffers). The results from these kinetic investigations are insufficient to conclude whether there is an internal proton transfer group in pea CA or if the proton is transferred directly between the zinc-bound water and a buffer molecule. All chloroplastic CAs from C 3 dicotyledons have a conserved His residue at position 208 (pea CA numbering starting from the initiator Met). In a study by Provart et al. on pea CA, this histidine was replaced by a glutamine and the CO 2-hydration activity was measured in crude E. coli lysate using a changing-pH method (29). They found the activity of this mutant to be significantly lower than that of wild-type enzyme in Hepes buffer, but not in the same concentrations of barbital or imidazole buffer. This result led the authors to suggest that His 208 has a similar function in proton shuttling as His 64 in human CA II. In a H64A mutant of human CA II, which is devoid of the internal proton shuttle, imidazole-type buffers (but not biological buffers) could replace His 64 and act in direct proton shuttling. To increase our knowledge of the kinetic mechanism of CAs from higher plants, and to further elucidate the role of His 208, we have replaced His 208 with Ala, a residue without hydrogen bonding abilities, and studied the kinetic properties of the isolated mutant enzyme. The results presented indicate that although His 208 is not necessary for a high turnover rate, it is most likely located in the vicinity of the active site. We find the side chain of His 208 affects the pH dependence of k cat/K m, resulting in a shift to a simple titration for the mutant. The interaction between the mutant enzyme and the proton-accepting buffer molecule is impaired, and extremely high buffer concentrations are needed to obtain saturating conditions. Furthermore, this mutant is found to be more easily oxidized,

STUDIES OF A HIS 208 MUTANT OF PEA CARBONIC ANHYDRASE

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and thereby inactivated, than the wild-type enzyme. The cysteines responsible for the redox sensitivity were previously reported to be at positions 269 and 272 (14). The results indicate an interaction between His 208 and these cysteines, where the histidyl side chain stabilizes the reduced state of the cysteines. MATERIALS AND METHODS Mutagenesis. The mutagenesis/expression plasmid containing cDNA encoding the mature subunit of pea carbonic anhydrase (pPCAs) under control of the T7 RNA polymerase promoter has been previously described (30). In vitro site-directed mutagenesis was performed essentially according to the method of Kunkel (31), and mutants were identified by directly sequencing the plasmid DNA. Plasmid containing the mutation was used to transform E. coli BL21(DE3) (32), which carries a chromosomal copy of the T7 RNA polymerase gene under the control of the lacUV5 promoter. The complete pea CA coding region was checked by plasmid sequencing before the clone was used for protein production. Protein production and purification. E. coli BL21(DE3) harboring wild-type or H208A mutant pPCAs plasmids was grown at 23°C in rich medium complemented with 100 mg/liter ampicillin to A 600 5 0.5, at which point pea CA synthesis was induced by the addition of 0.5 mM isopropyl-b-D thiogalactopyranoside. The cells were harvested after an additional 15 h and resuspended in 50 mM Tris– H 2SO 4 (pH 8.0) before mechanical disruption using a French press or Bead-Beater (BioSpec Products, USA). The enzyme was purified by affinity chromatography using p-aminomethylbenzenesulfonamide hydrochloride (Sigma) coupled to epoxy-activated Sepharose 6B (Pharmacia Biotech) as previously described (24) and eluted with 0.1 M NaN 3. The H208A mutant protein was isolated in the presence of 10 mM cysteine in all buffers, including the cell resuspension buffer. Collected enzyme was dialyzed against several changes of degassed 10 mM Bistris-H 2SO 4 (pH 6.8) with the addition of 10 –30 mM cysteine or 0.1– 0.2 mM tris(2-carboxyethyl)phosphine (TCEP; Pierce) as reducing agents. The purity of the preparations was verified using SDS–PAGE and Coomassie brilliant blue staining. Enzymes were stored in small aliquots under N 2(g) at 220 or 250°C. Protein concentrations were determined from the absorbance at 280 nm. Using the method of Gill and von Hippel (33), the molar absorption coefficient was calculated to 28,200 M 21 cm 21. The enzyme concentrations refer to the quantity of subunits, assuming a molecular mass of 24.2 kDa. Kinetic measurements. Kinetic parameters were measured in an Applied Photophysics DX-17MV sequential stopped-flow spectrofluorimeter, or a Hi-Tech stopped-flow spectrophotometer, at 25°C by the changing pH-indicator method (34, 35). The buffer/indicator pairs used were Taps or Tricine with m-cresol purple measured at 578 nm, Mops or 3-(N-morpholino)-2-hydroxypropanesulfonic acid (Mopso) with 4-nitrophenol measured at 400 nm, and Mes with chlorophenol red measured at 574 nm. To prevent oxidation, all buffers contained 10 mM EDTA and 1 mM dithiothreitol or 0.2–5 mM TCEP for the H208A mutant. The enzyme/buffer solutions were stored in a gastight syringe until use. Initial rates were calculated by fitting the first part of the curve to a first-order rate equation. Initial rate data were fitted by nonlinear regression to the Michaelis–Menten equation using the GraFit program (Erithacus Software Ltd., England). The pH dependence of the inhibition of the CO 2-hydration activity by SCN – was measured at a constant CO 2 concentration of 12.7 mM and varying NaSCN concentrations. All kinetic values were calculated on an enzyme subunit basis. For the redox-sensitivity studies, the CO 2-hydration activity was measured at 12°C by the colorimetric method according to Rickli et al. (36).

FIG. 1. Reactivation of oxidized H208A mutant by SH-reducing agents. The CO 2-hydration activity was measured according to (36). Oxidized enzyme was incubated with Cys, DTT, 2-mercaptoethanol (2-ME), and TCEP in 20 mM Bistris-H 2SO 4, pH 6.8, or with buffer only as a control (oxidized). Samples were withdrawn for activity measurements at various time points, and the figures given show the maximal activity obtained. Freshly thawed enzyme in the presence of 100 mM 2-ME (reduced) is defined as having 100% activity.

RESULTS

Enzyme isolation. The H208A mutant turned out to be very susceptible to air oxidation. The catalytic activity dropped rapidly when isolated according to the method used for the wild-type enzyme. However, enzyme with high activity could be isolated by including 10 mM cysteine in the cell resuspension buffer and dialyzing the purified enzyme against buffer containing 0.2 mM TCEP instead of cysteine. The resulting enzyme preparation was stable for more than 1 h in the presence of a reductant. The overall structure of the mutant was checked by CD spectroscopy, and the spectra are practically identical to those previously reported for the wild-type enzyme (24) (data not shown). Redox sensitivity. The mutant enzyme was oxidized by air exposure in 20 mM Bistris-H 2SO 4, pH 6.8. At this pH the process is comparatively slow, and it takes at least 48 h before the activity is reduced below 5%. In comparison, the wild-type enzyme does not undergo significant oxidation unless pH is increased. Such inactive enzyme is considered to be oxidized, and the catalytic activity can be regained by the addition of a reductant. In Fig. 1 the degree of reactivation of H208A obtained by different -SH agents is presented. Freshly thawed enzyme in 20 mM Bistris-H 2SO 4, pH 6.8, and 100 mM 2-mercaptoethanol is taken as reduced and having 100% activity. Oxidized enzyme, retaining 7% of the activity, was used for incubation with the reduc-

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¨ RKBACKA, JOHANSSON, AND FORSMAN BJO

FIG. 2. pH dependence of the inactivation of H208A by oxidation. Freshly thawed enzyme was incubated in 50 mM buffer with 10 mM EDTA for 10 –15 min at room temperature, and the CO 2-hydration activity was measured according to Rickli et al. (36). The enzyme concentration was 3–5 mM and the buffers used were acetate (pH 5–5.5), Bistris-H 2SO 4 (pH 6 –7), Tris-H 2SO 4 (pH 7–9.3), and 2-cyclohexylamino-l-propanesulfonic acid (pH 9.7–10.7). (E) H208A, (h) wild type. (Inset) The activity of H208A at pH 8.8 (50 mM TrisH 2SO 4, 10 mM EDTA) as a function of time after mixing with buffer (F). At 15 min 2 mM TCEP (h) or 100 mM 2-mercaptoethanol (E) was added.

ing agents. The activity was measured at various time intervals, and the activation came to a plateau after approximately 1 h and did not change over a period of 3 h thereafter. This value is given in the figure. It is striking that only 2-mercaptoethanol reactivates the mutant to a level close to that of the fully reduced state, while cysteine, DTT, and TCEP all give around 20% reactivation. However, we find all reductants to be efficient in maintaining the reduced, active state, despite the inefficiency in reactivating the enzyme. The sensitivity to oxidation is pH dependent for both wild-type and mutant enzymes, as illustrated in Fig. 2. Freshly thawed enzyme was mixed with 50 mM of buffer, and the activity was measured after 10 –15 min. All samples contained 10 mM EDTA to trap metal ions that can catalyze the oxygen oxidation of SH groups. We find the H208A mutant to be more readily inactivated as pH increases compared to the wild type. The midpoint for the activity drop is at pH 9 for the wild type, while it has decreased to near pH 7.5 for the H208A mutant. To confirm that the inactivation is due to the formation of a disulfide bond, the activity of H208A at pH 8.8 was measured at various time points. After 15 min the activity was reduced by 70%. The addition of a reducing agent, 2 mM TCEP or 100 mM 2-mercaptoethanol, at that time point resulted in a

complete recovery of the activity (shown as insert in Fig. 2). This also shows that the oxidation is a reversible process. Steady-state kinetics. The CO 2-hydration activity was measured in the pH range 6 –9. The ionic strength was allowed to vary, since we did not want to add additional salt that might inhibit the reaction. We have previously found for the wild-type enzyme that Na 2SO 4 added to a total ionic strength of 50 mM has no activating effect, and at low pH the k cat values are significantly decreased due to SO 422 inhibition (24). The pH profiles of k cat and k cat/K m are presented in Fig. 3, which shows that the activity of this mutant is only slightly lower than that of the wild-type enzyme. Still, we can note two interesting differences. The first is in the parameter k cat, which has stronger pH dependence in the mutant, the largest difference from the wild-type occurring at low pH values. This is probably a consequence of the altered buffer dependence observed for H208A (see below and Table I). The second difference is in the parameter k cat/K m, where the wild-type enzyme has a rather complex behavior, probably with more than one titratable group affecting the activity. In contrast, the data obtained for the mutant can be nicely fit

FIG. 3. pH dependence of the kinetic parameters k cat and k cat/K m for the CO 2 hydration catalyzed by H208A and wild-type pea CA. Measurements were made at 25°C in Taps (pH 8.7– 8.9), Tricine (pH 8.2), Mops (pH 7.8 –7.2), Mopso (pH 6.8), and Mes (pH 6.2– 6.5) buffers. The k cat data are for 50 mM buffer for the wild type and 75 mM buffer for the H208A mutant, and the k cat/K m data are for buffers in the range 25–100 mM. (F) H208A, (E) wild type. The k cat/K m data for the H208A mutant were fitted to a simple titration curve with a pK a value of 7.1 6 0.1.

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STUDIES OF A HIS 208 MUTANT OF PEA CARBONIC ANHYDRASE TABLE I

Parameters for Buffer Activation of the CO 2-Hydration Activity of the H208A Mutant and the Wild-Type Enzyme Measured at 25°C Wild type a

H208A

Buffer

pH

(k cat) max b (ms 21) d

K buffer c (mM) d

(k cat) max b (ms 21) d

K buffer c (mM) d

Taps Mops Mes

8.8 7.3 6.2

1360 6 380 280 6 20 50 6 9

350 6 130 310 6 35 100 6 40

740 6 40 310 6 25 (135) e

110 6 10 52 6 12 (4) e

a

Data are from (24). (k cat) max represents k cat value/subunit at infinite buffer concentration. c K buffer is the apparent K m with respect to total buffer concentration. d Numbers represent means followed by standard deviation. e Extrapolated from the highest substrate concentrations, due to curved Eadie–Hofstee plots. b

to a simple titration curve with a pK a of 7.1 6 0.1. According to the catalytic mechanism described by Eqs. [1] and [2], this should be the pK a for the Zn–H 2O. Thus, the side chain of His 208 affects the ionization of the catalytically active zinc-bound water through direct or indirect interactions. The kinetic study of H208A was extended with an investigation of the dependence of the CO 2 hydration rate on the buffer concentration. In the catalytic cycle for CA, the buffer molecule can be regarded as a second substrate, accepting the proton produced from hydration of CO 2 (Eq. [2]). At low, neutral, and high pH we find k cat to be strongly dependent on the buffer concentration, while k cat/K m is almost unaffected, as illustrated for pH 8.8 in Fig. 4. This pattern is the same as that previously reported for the wild-type enzyme and in agreement with a ping-pong mechanism (24). k cat values at infinite buffer concentration, (k cat) max, and apparent K m values with respect to the total buffer concentration, K buffer, were estimated from secondary

FIG. 4. Effects of the buffer concentration on the CO 2 hydration catalyzed by H208A. Measurements were taken at 25°C in Taps buffer, pH 8.8, at total concentrations of 150 mM (ƒ), 75 mM (h), 50 mM (‚), and 25 mM (E).

plots (Table I). The K buffer values for the H208A mutant are very high, and clearly the buffer concentration used to obtain the k cat data in Fig. 3 (75 mM) is far below saturating level. In comparison to the K buffer values obtained for the wild-type enzyme, the mutant has a much stronger buffer dependence, and this effect of the H208A mutation increases with decreasing pH. This gives us one explanation for the stronger pH dependence of k cat for the H208A enzyme seen in Fig. 3, since the wild-type enzyme, in contrast to the mutant, operates under close to saturating conditions with respect to buffer at pH below 7. The parameter k cat/K m is independent of the buffer concentration due to the ping-pong mechanism, and thus the differences in k cat/K m are not explained by the different K buffer values. Furthermore, it is interesting to note that the (k cat) max value at high pH is almost two times higher for the mutant. Inhibition. The inhibition of the CO 2-hydration activity was measured at 12°C using the well-known CA inhibitors, acetazolamide, SCN 2, Cl 2, and SO 422. Compared to the wild-type enzyme, the binding of the inhibitors to the H208A mutant is only moderately weakened by a factor of two for the sulfonamide as well as for the anions. With the H208A mutant we obtain I 50 values of 57 mM for acetazolamide, 50 mM for SCN 2, 70 mM for Cl 2 and 530 mM for SO 422. The corresponding values for the wild-type enzyme are 28 mM, 20 mM, 40 mM, and 190 mM, respectively. In human CA II, k cat/K m and K i both depend on the ionization state of the Zn–H 2O. To probe the ionization of the H208A mutant, we investigated the pH dependence of the anion binding to the enzyme. A key interest was whether or not it is simplified to that of a single titration, in analogy to the behavior of k cat/K m. The inhibition by SCN 2 of the CO 2-hydration activities of the mutant and the wild type at different pH values is shown in Fig. 5. The difference between the two en-

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¨ RKBACKA, JOHANSSON, AND FORSMAN BJO

FIG. 5. pH dependence of the inhibition by SCN 2 of the CO 2hydration activity of H208A and wild-type enzyme at 25°C. Initial rates were measured at a constant CO 2 concentration of 12.7 mM and varying SCN 2 concentrations. K i values were obtained by linear regression of Dixon plots of 1/(initial rate). The buffers used were 75 mM Mes (pH 6.1– 6.7), 100 mM Mops (pH 7.1–7.8), 100 mM Tricine, pH 8.2, and 100 mM Taps (8.5– 8.8). (F) H208A, (E) wild type. Data for the wild-type enzyme are from (25).

zymes is only marginal and found mainly at pH values above 7, where the dissociation constants for the mutant are higher. The pH dependence is far from that of a simple ionization, which implies that the SCN 2 is affected by additional groups other than the zincbound water in both the wild type and the mutant. Using SCN 2 we also investigated the anion-inhibition patterns for the H208A mutant. At pH 9.0 an uncompetitive inhibition pattern was obtained, shifting to noncompetitive at pH 6.5 (Figs. 6A and 6B). This agrees well with previous results on wild-type pea CA (25) and spinach CA (22, 37). DISCUSSION

Many features of the catalytic behavior of chloroplast CA from higher plants are compatible with the

zinc-hydroxide mechanism proposed for human CA II. However, there are several results that indicate differences in the step(s) involving proton transfer between the active site and surrounding medium, and the mechanism might change with pH (25). Results obtained for the H208A mutant supporting the hypothesis that the general mechanism is the same as for the wild-type enzyme include the ping-pong patterns obtained when the buffer is treated as the second substrate. Furthermore, the SCN 2-inhibition patterns are unaltered and consistent with the idea of a protonated enzyme form, with strong anion affinity, preceding the rate-limiting proton-transfer step. It has previously been suggested that His 208 in pea CA acts as an internal protontransfer group, analogous to His 64 in human CA II (29). The results presented here on a H208A mutant do not give clear support to this conclusion. The high catalytic activity of H208A means that, despite the absence of the histidyl side chain, the proton transfer step still occurs at a reasonably high rate. It also means that a bulk-water molecule is not acceptor of the proton from the Zn–H 2O, since that would limit k cat to 10 3–10 4 s 21 (38). Instead, a buffer molecule must act as the final proton acceptor, as proposed by the ping-pong mechanism. Even though the results presented do not rigorously show that His 208 acts as an internal proton shuttle, it is clear that the interaction with the buffer is impaired in the mutant. The K buffer values are very high over the pH range 6 –9, while the wild-type enzyme shows decreasing K buffer values with decreasing pH. The (k cat) max/K base ratio, where K base is the Michaelis constant for the basic buffer species, puts an upper limit for the rate of H 1 transfer from CA to bulk buffer (39, 40). In human CA II, the limiting maximum value is approximately 1 3 10 9 M 21 s 21, close to that of a diffusion-controlled process. The corresponding value for wild-type pea CA is 100 times lower and thus far

FIG. 6. SCN 2 inhibition of CO 2 hydration catalyzed by H208A at 25°C. (A) 100 mM Taps buffer at pH 9.0 in the presence of (E) no inhibitor, (‚) 0.1 mM NaSCN, (h) 1 mM NaSCN. (B) 75 mM Mes buffer at pH 6.5 in the presence of (E) no inhibitor, (‚) 0.05 mM NaSCN, (h) 0.1 mM NaSCN.

STUDIES OF A HIS 208 MUTANT OF PEA CARBONIC ANHYDRASE

from diffusion control. The high K buffer values for the H208A mutant at neutral and low pH result in (k cat) max/ K base values less than 10% of those of wild-type enzyme, while at high pH the difference is only marginal. Thus, at high pH and in the absence of the histidyl side chain, the proton transfer to a buffer molecule is largely unaffected, but when pH decreases the proton transfer becomes more difficult in the mutant. In this context we note that the solvent hydrogen isotope effect for pea CA was reported to be pH dependent with k cat/K m being almost unaffected by the solvent at high pH, while at neutral and low pH an effect of 2.3–3 was observed (25). The parameter k cat/K m is assumed to reflect the ionization state of the Zn–H 2O. The pH dependence in k cat/K m is simplified to that of a single titration with pK a 5 7.1 for the H208A mutant. Due to the more complex pH dependence of the wild-type enzyme, no pK a value has been calculated, but according to the data in Fig. 3 it should be higher than for the mutant. This is not what would normally be expected as the result of a mutation where a positive charge is removed from the vicinity of the titrating group, and the decreased pK a could therefore be caused by a secondary effect of the mutation. The lower affinity of SCN 2 to the mutant at pH values above seven fits with a decreased pK a, since the Zn–OH 2 form of the enzyme has weak anion binding. The H208A mutant is strikingly more easily oxidized than the wild-type enzyme. The activation study shows that if the oxidation is allowed to proceed for a long time, only approximately 20% of oxidized H208A can be reduced to an active enzyme form by the addition of DTT, cysteine, or TCEP. There is no difference in the degree of reactivation whether the reductant is a monothiol, dithiol, or a phosphine. Using these reducing agents, the main fraction of the mutant molecules seems to be irreversibly inactivated. This was also reported for the wild-type enzyme, where 30 and 50% activation was obtained in DTT and cysteine, respectively (24). However, if 2-mercaptoethanol is added to oxidized enzyme, 80% of the activity can be regained for the H208A mutant and 60% for the wild type. An explanation for this more efficient activation can perhaps be found in connection with the observation that 2-mercaptoethanol, besides activating pea CA by reduction, also acts as an inhibitor. The molecule, when acting as an inhibitor, binds to the active site, probably close to the zinc as it can displace the sulfonamide inhibitor p-aminomethylbensenesulfonamide (C. Forsman, unpublished data). One possibility is that this binding gives a high local concentration of reductant, thus increasing the reducing power. However, the most likely orientation of 2-mercaptoethanol is such that the sulfur interacts with the zinc, which would mean that it would be unable to participate in a redox reaction. A

23

more plausible explanation for the high degree of activation could be that the binding of 2-mercaptoethanol to the active site favors formation of the active conformation of the enzyme, thus promoting a conversion from the oxidized, inactive enzyme form to the reduced and active form. A second 2-mercaptoethanol molecule should then be responsible for the redox reaction. This argument implies that oxidation of the enzyme gives some conformational change beyond formation of the disulfide bridge. We have previously reported indications of such changes (14). These studies are currently being extended and will be published elsewhere. The degree of reactivation is higher for the wild type than for H208A in 2 mM DTT or 30 mM Cys. This could mean that the native structure of the active site is to some extent stabilized by His 208, resulting in a higher tendency to adopt the inactive conformation in the absence of the histidyl side chain. We also note that in the study shown in Fig. 2, where oxidation was for only 15 min, 100% of the activity was regained upon reduction. The difference in reactivation that is observed in Figs. 1 and 2 could be an effect of the extent of oxidation or an effect of a conformational change occurring on a slower time scale. The disulfide-bond formation is promoted by the thiolate anion, giving a pH dependence of the reaction. The pH dependence of the inactivation shown in Fig. 2 could reflect the deprotonation of a disulfide-forming cysteine, and the pK a is then decreased by at least one pH unit in the absence of His 208. This would give an explanation to the observed increased oxidation sensitivity for the H208A mutant. The oxidation of SH groups is greatly influenced by the distance and orientation of the individual cysteines. The neighboring groups are also of great interest when trying to explain the rate of oxidation. For instance electronegative groups have been shown to reduce the rate of oxidation of dithiols, presumably by increasing the Cys pK a value (41, 42). One explanation to the higher redox sensitivity of H208A compared to the wild-type could be that His 208 interacts, directly or indirectly, with one of the redox-active cysteines. The effect could be stabilization of the protonated SH group, thus increasing the pK a value or sterically preventing a proper orientation for disulfide bond formation. The cysteines responsible for the oxidation sensitivity have been shown previously to be Cys 269 and Cys 272 (14). Replacing either of these with Ala or Ser resulted in mutants with characteristics that are unaffected by oxidation, but with very low catalytic activities that could not be increased by the addition of extra reducing agents. It should be noted that Cys 269 is conserved among all higher plant CAs, while Cys 272 is found only in combination with His 208 in the chloroplastic CAs from C 3 dicotyledonous plants. We have made the double mutant H208A/C272A, and the char-

24

¨ RKBACKA, JOHANSSON, AND FORSMAN BJO

acteristics of this enzyme are very similar to those of the single cysteine mutants at positions 269 and 272 (14). However, an interesting effect of the double mutation at positions 208 and 272 is the formation of disulfide bridges between subunits, as seen from SDS– PAGE run in the absence of 2-mercaptoethanol (unpublished data). This was not observed for the C272A single mutant, or for any other mutant that we have studied so far. Thus, in the absence of His 208 the SH group of Cys 269 seems to have a higher susceptibility to oxidation, and a disulfide bridge might form between Cys 269 residues in neighboring subunits of the H208A/C272A mutant. In summary, His 208 seems to be located in the active site region, where it contributes to a structure important for the buffer-dependent proton transfer. This structure also affects the titration of the zincbound water molecule. The histidine residue is not crucial for the catalytic turnover, and the activity of the H208A mutant is almost as high as it is in the wildtype. Our results indicate that a possible function of His 208 is to prevent the cysteine pair at positions 269 and 272 from being oxidized and the His side chain seems to be positioned so that it stabilizes the reduced state of these cysteines. ACKNOWLEDGMENTS The authors thank Professor Sven Lindskog for valuable discussions and Eleonore Ska¨rfstad for skillful technical assistance. Financial support from Magn Bergvalls Stiftelse, Stiftelsen Lars Hiertas Minne, and J. C. Kempes Minnes Stipendiefond are gratefully acknowledged.

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