©1996 by The American Society for Biochemistry and Molecular Biology, Inc.
Efficient Expression of the Gene for Spinach Phosphoribulokinase in Pichia pastoris and Utilization of the Recombinant Enzyme to Explore the Role of Regulatory Cysteinyl Residues by Site-directed Mutagenesis (*)

(Received for publication, November 10, 1995; and in revised form, January 2, 1996)

Hillel K. Brandes (2) Fred C. Hartman (1)(§) Tse-Yuan S. Lu (1) Frank W. Larimer (1)

From the  (1)Protein Engineering Program, Biology Division, Oak Ridge National Laboratory and the (2)University of Tennessee-Oak Ridge Graduate School of Biomedical Sciences, Oak Ridge, Tennessee 37831

ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
FOOTNOTES
REFERENCES

ABSTRACT

Phosphoribulokinase (PRK), unique to photosynthetic organisms, is regulated in higher plants by thioredoxin-mediated thiol-disulfide exchange in a light-dependent manner. Prior attempts to overexpress the higher plant PRK gene in Escherichia coli for structure-function studies have been hampered by sensitivity of the recombinant protein to proteolysis as well as toxic effects of the protein on the host. To overcome these impediments, we have spliced the spinach PRK coding sequence immediately downstream from the AOX1 (alcohol oxidase) promoter of Pichia pastoris, displacing the chromosomal AOX1 gene. The PRK gene is now expressed, in response to methanol, at 4-6% of total soluble protein, without significant in vivo degradation of the recombinant enzyme. This recombinant spinach PRK is purified to homogeneity by successive anion-exchange and dye-affinity chromatography and is shown to be electrophoretically and kinetically indistinguishable from the authentic spinach counterpart. Site-specific replacement of all of PRK's cysteinyl residues (both individually and in combination) demonstrates a modest catalytically facilitative role for Cys-55 (one of the regulatory residues) and the lack of any catalytic role for Cys-16 (the other regulatory residue), Cys-244, or Cys-250. Mutants with seryl substitutions at position 55 display non-hyperbolic kinetics relative to the concentration of ribulose 5-phosphate. Sulfate restores hyperbolic kinetics and enhances kinase activity, presumably reflecting conformational differences between the position 55 mutants and wild-type enzyme. Catalytic competence of the C16S-C55S double mutant proves that mere loss of free sulfhydryl groups by oxidative regulation cannot account entirely for the accompanying total inactivation.


INTRODUCTION

Phosphoribulokinase (EC 2.7.1.19) catalyzes the final step in the regeneration of ribulose 1,5-bisphosphate, the acceptor for CO(2) in photosynthetic carbon assimilation, from 3-phospho-D-glycerate. This kinase reaction, in which the -phosphoryl of ATP is transferred to the C-1 hydroxyl of Ru-5-P, (^1)provides the only avenue by which Calvin cycle intermediates can be rechanneled for utilization in CO(2) fixation. As a means of coupling photosynthetic carbon reduction with photosynthetic electron transport, the activity of PRK is modulated by photon flux. In the case of prokaryotic photosynthetic organisms, regulation of PRK (typically an octamer of 32-kDa subunits) is mediated by the allosteric activator NADH, the intracellular concentration of which is light-dependent (for a review, see (1) ). By contrast, regulation of eukaryotic PRK (typically a dimer of 40-kDa subunits) (2, 3, 4) is mediated by the redox active protein thioredoxin, which participates in sulfhydryl/disulfide exchange reactions(5, 6) . In the light, thioredoxin is maintained in its reduced form by the electron carrier ferredoxin. Reduced thioredoxin then reduces the regulatory disulfide of the inactive form of PRK to free sulfhydryls, thereby generating the active form of PRK.

In addition to the striking differences in molecular architecture and mode of regulation between prokaryotic and eukaryotic PRK, their catalytic parameters are also quite distinct. As an example, the k of plant PRK is about 10-fold higher than that of the bacterial counterpart(1) . Hence, the two classes of PRK must be considered as separate entities, each deserving of independent scrutiny. Structure-function studies of the bacterial (Rhodobacter sphaeroides) PRK have been greatly facilitated by the recent redesign of an expression plasmid that overproduces the recombinant enzyme in Escherichia coli to the extent of about 25% of the total soluble protein(7, 8) . Although the gene for spinach PRK has also been cloned (9, 10) and expressed in E. coli(11, 12) , the yield of recombinant enzyme is very low (leq0.5% of total soluble protein) due to the dual problems of susceptibility to proteolysis and inherent toxicity of spinach PRK to the host.

To circumvent these impediments, we have explored utilization of the yeast Pichia pastoris as an expression host for spinach PRK. In this paper, we report our success in this endeavor, including efficient purification of the recombinant enzyme and its characterization to validate authenticity to PRK isolated from spinach. We also describe construction, purification, and characterization of site-directed mutants of spinach PRK, in which the regulatory cysteinyl residues are replaced both individually and in combination so as to gain understanding of the molecular basis of oxidative deactivation.


EXPERIMENTAL PROCEDURES

Chemicals and Biologicals

Sequenase version 2.0 T7 DNA polymerase, Sequenase sequencing reagents, and DeltaTaq cycle sequencing system were obtained from U. S. Biochemical Corp. 5`-[alpha-S]dATP (1000 Ci/mmol) was purchased from DuPont NEN. Yeast nitrogen base without amino acids was obtained from Difco. MES, Hepes, Bicine, and DTT were purchased from Research Organics, Inc. Restriction endonucleases, calf intestine alkaline phosphatase, Klenow fragment of DNA polymerase I, and T4 DNA ligase were obtained from New England Biolabs. 4-(2-Aminoethyl)benzenesulfonylfluoride was obtained from Calbiochem, while additional buffer and assay components, sorbitol, DTNB, and Red-Agarose 3000-CL were purchased from Sigma. Oligonucleotides were synthesized on an Applied Biosystems model 392 DNA/RNA synthesizer using phosphoramidite chemistry. SuperQ-Toyopearl was obtained from TosoHaas. Spinach PRK was purified as described earlier(13, 14) . Ru-5-P was prepared from ribose 5-phosphate by the action of phosphoribose-isomerase, and its concentration was based on the reported equilibrium constant(15) .

Strains, Plasmids, and Phage

E. coli strain MV1190 Delta(lac-proAB) thi supE Delta(srl-recA)306::Tn10(tet^R) (F`: traD36 proABlacI^qZDeltaM15) was used for the propagation of plasmids and phage. Single-stranded dU-substituted template was produced in the E. coli host strain CJ236 dut ung thi relA (pCJ105 Cm^r), subsequent to its infection with M13KO7 helper phage. The P. pastoris expression system, purchased from Invitrogen Corp., includes the P. pastoris host strain GS115 his4, the AOX1 (methanol oxidase gene) intracellular expression vector pHIL-D2, 5` PCR/sequencing primer GACTGGTTCCAATTGACAAGCT, and 3` PCR/sequencing primer GCAAATGGCATTCTGACATCC.

Construction of the PRK Expression Cassette

Vector pHIL-D2 has a single EcoRI cloning site adjacent to the AOX1 promoter. This site was cleaved by EcoRI digestion, and the resulting 5`-overhangs were filled in by use of the Klenow fragment of DNA polymerase I plus dNTPs. An oligonucleotide linker containing a BglII site (pGAAGATCTTC) was ligated to the blunt-ended vector. The resulting vector (pHIL-D2B) was cleaved with BglII and dephosphorylated with calf intestinal alkaline phosphatase. The spinach PRK gene was excised from vector pSM100 (11) by BamHI digestion and was isolated by agarose gel electrophoresis. The PRK gene was ligated into pHIL-D2B to form the vector pFL451. The sequences of the BamHI/BglII junctions were confirmed by dideoxy-terminator cycle sequencing (according to instructions provided by the U. S. Biochemical Corp.) by use of the 5` and 3` primers supplied by Invitrogen.

Construction of PRK Mutations

Site-directed mutagenesis of the PRK gene was accomplished by the Kunkel procedure(16) . The primers used to construct C16S, C55S, C244S, and C250S have been described(11) . Each mutation was confirmed by dideoxy-terminator sequencing (17) in conjunction with the primers AACGACAACTTGAGGAAGATC for the Cys-Cys region and GAATCCAGAAAGCCA for the Cys-Cys region.

Integration of the PRK Expression Cassette into P. pastoris

Late log-phase cells for electroporation were concentrated 10-fold into 1 M sorbitol, 20 mM Hepes-NaOH (pH 7.5)(18) . Ten µg of NotI-cleaved pFL451, or the appropriate mutant derivative, was mixed with 100 µl of P. pastoris GS115 suspension (about 2 times 10^8 cells) and electroporated at 5000 V/cm (25-microfarad capacitance), with a 200-ohm parallel resistor. The time constant was typically 6 ms. Approximately 100 transformants/µg of DNA were obtained. Histidine-independent transformants were selected and subsequently screened for methanol utilization. The his methanol isolates were induced with methanol and screened for PRK production by Western blotting of cell extracts. Displacement of the AOX1 gene by the PRK cassette (see Fig. 1A) was confirmed by examination of Southern hybridizations of BglII-digested genomic DNA of P. pastoris probed with either labeled AOX1 DNA or PRK DNA (Fig. 1, B and C) and by PCR amplification of the AOX1 region from genomic DNA (Fig. 1D). The presence of the planned substitutions for Cys codons was confirmed by dideoxy-terminator cycle sequencing using PCR-amplified PRK template from genomic DNA of transformed P. pastoris.


Figure 1: Integration of the PRK expression cassette into P. pastoris chromosomal DNA. A, schematic representation of the displacement of the AOX1 sequence of GS115 by the expression cassette excised from pFL451 by NotI digestion. The displacement product, denoted Deltaaox1::PRK, lacks AOX1 and contains PRK-HIS4. The sequences flanking the PRK-HIS4 segment are homologous to the target sequences flanking AOX1. The 5` and 3` arrows denote the locations of sequencing/PCR primers. B and C, Southern transfer hybridization of NotI-BglII-digested plasmid and BglII-digested genomic DNAs. In B, AOX1 is used as the probe. The coding sequence of AOX2 is nearly identical to AOX1 and is thus also visualized. In C, PRK is used as the probe. D, ethidium bromide-stained agarose gel of PCR-amplified intact plasmid and bulk genomic DNAs. The identity of the amplified DNA was confirmed by dideoxy-terminator cycle sequencing (not shown). Note in B and D the absence of AOX1 in Deltaaox1::PRK and in C that the entire disruption cassette is incorporated into chromosome Deltaaox1::PRK.



Production of PRK from P. pastoris

Cultures (500 ml in 2-liter baffled flasks) were routinely grown in minimal medium (13.4 g/liter yeast nitrogen base without amino acids, 400 µg/liter biotin, 0.1 M MES-NaOH at pH 6.5) containing 1% (v/v) glycerol at 28 °C with vigorous shaking (250 rpm). After the cultures (originally inoculated at about 10^7 cells/liter) reached saturation (48-72 h), they were harvested by centrifugation and resuspended in 0.5 volume of minimal medium containing 0.5% (v/v) methanol. The incubation was continued for another 72 h, during which time additional methanol (5 ml/liter) was added at 24 and 48 h. Cells were harvested by centrifugation, washed once with ice-cold distilled water, and stored at -80 °C; the yield of wet, packed cells was 40-60 g/liter.

Assay of PRK Activity

Kinase activity was determined at 25 °C by a spectrophotometric assay in which ADP, generated by the kinase reaction, is coupled to NADH oxidation via phosphoenolpyruvate, pyruvate kinase, and lactate dehydrogenase(4, 19) . Assay mixtures (1 ml) at pH 8.0 routinely contained Bicine (50 mM), KCl (40 mM), MgCl(2) (10 mM), NADH (0.4 mM), ATP (1 mM), Ru-5-P (2.5 mM), phosphoenolpyruvate (3 mM), DTT (1 mM), pyruvate kinase (5 units), lactate dehydrogenase (6 units), and PRK (<0.2 units); reactions were initiated by the addition of ATP or the kinase. When determining the K(m) for Ru-5-P, ATP was held constant at 1 mM. When determining the K(m) for ATP, Ru-5-P was held constant at either 1 mM with wild-type and C16S mutant PRK or 2.5 mM with C55S, C16S-C55S, and C16S-C55S-C244S-C250S mutants.

Purification of Wild-type Recombinant PRK

Extractions and all steps of the purification were carried out at 4 °C. P. pastoris cell paste (50 g) was resuspended in 50 ml of pH 8.0 lysis buffer (100 mM Bicine, 5 mM EDTA, 10 µM leupeptin, 100 µM phenylmethylsulfonyl fluoride, 0.2 mM aminoethylbenzenesulfonyl fluoride, 22 µM pepstatin, 10 mM DTT) by vigorous vortexing. Cells were lysed by physical disruption with 0.5-mm glass beads in a BeadBeater (Biospec). Cell lysis was achieved during a 2-min agitation period (successive bursts of 10 s interspersed with pauses of 50 s). The cell lysis vessel and glass beads were rinsed twice with minimal volumes of lysis buffer. The combined rinses and suspension of ruptured cells were centrifuged at 10,000 times g for 10 min; the collected supernatant was recentrifuged at 120,000 times g for 60 min. The supernatant was brought to 20% (v/v) glycerol and frozen at -80 °C for overnight storage. The thawed supernatant was diluted with an equal volume of pH 8.0 buffer (50 mM Bicine, 1 mM EDTA, 0.2 mM aminoethylbenzenesulfonyl fluoride, 10 mM DTT), filtered (Gelman Acrodisc pore size of 0.8 µm), and applied at 5 ml/min to a 60-ml SuperQ-Toyopearl 650-M column (2.6 cm times 11.3 cm) equilibrated with the same buffer. Subsequent to complete elution of nonbound components, a 600-ml salt gradient was applied (0 to 0.5 M KOAc) at 10 ml/min while collecting 10-ml fractions. Fractions containing the bulk of the PRK activity were pooled, brought to 20% (v/v) glycerol, divided into two equal portions, and frozen at -80 °C. A thawed portion was dialyzed (Spectra/Por 2 membrane; 12,000-14,000 molecular weight cut-off) against 4 liters of pH 8.0 buffer (50 mM Bicine, 1 mM EDTA, 10 mM DTT) for 2 h, and the dialysis was repeated once. After the second dialysis, the sample was brought to pH 6.5 by the addition of solid Bicine and immediately applied at 1.5 ml/min to a 60-ml Red-Agarose 3000-CL column (2.2 cm times 15.8 cm) equilibrated with 10 mM Bicine (pH 6.5). After washing the column successively with 600 ml of 10 mM Bicine (pH 6.5), 10 mM DTT followed by 600 ml of 10 mM potassium phosphate (pH 6.8), 10 mM DTT, PRK was eluted with 10 mM potassium phosphate (pH 7.2), 10 mM DTT containing 5 mM ATP. Fractions containing the recovered PRK activity were pooled and concentrated to about 2 ml by ultrafiltration with a YM30 (Amicon) membrane. The concentrated sample (>10 mg/ml) was then dialyzed twice against 1 liter of 50 mM Bicine (pH 8.0), 1 mM EDTA, 5 mM potassium phosphate, 10 mM DTT, with inclusion of glycerol at a final concentration of 20% (v/v) in the buffer for the second dialysis. Finally, the sample was dialyzed once again in 50 mM Bicine (pH 8.0), 1 mM EDTA, 10 mM DTT, 20% (v/v) glycerol and subsequently stored at -80 °C.

All mutant PRKs were purified by the same protocol and displayed chromatographic behavior indistinguishable from that of wild-type.

The concentration of PRK was determined at 280 nm based on an extinction coefficient of 7.17 for a 1% (10 mg/ml) solution in a 1-cm light path(13) .

Polyacrylamide Gel Electrophoresis and Immunoblotting

Purified PRKs were subjected to electrophoresis under nondenaturing conditions without any sample manipulation other than appropriate dilution. For electrophoresis under denaturing conditions, the samples were diluted, brought to final concentrations of 0.5% (w/v) SDS, 1 mM 2-mercaptoethanol, and then heated at 100 °C for 1 min. In all cases, electrophoresis was carried out at 15 °C on 8-25% polyacrylamide PhastGel medium with a PhastGel apparatus (Pharmacia Biotech Inc.). Gels were stained with either silver or Coomassie Blue according to manufacturer's protocols. Western immunoblotting was carried out by standard procedures(20, 21) as previously applied to PRK(22) , using a Novex Xcell apparatus (Novex, San Diego, CA). Proteins from unstained gels (Novex, 14%) were transferred to nitrocellulose sheets (Novex) using the Novex blot module. The nitrocellulose membrane was impregnated with rabbit polyclonal anti-PRK IgG(22) . Following exposure of the membrane to secondary antibody (goat anti-rabbit IgG conjugated with horseradish peroxidase; Bio-Rad) and thorough washing, the immunoreactive proteins were detected by the color produced upon reaction of the horseradish peroxidase with H(2)O(2) and 3,3`-diaminobenzidine in 50 mM Tris-HCl (pH 7.2).

Kinetic Analyses

Kinetic data were analyzed by a Marquardt-Levenberg algorithm of nonlinear least squares fitting (SigmaPlot®, Jandel Scientific) to either v = V bullet S/(K + S) or v = V bullet S/(K + S + S^2/A), where A is the substrate inhibition constant.


RESULTS

P. pastoris as an Expression Host for PRK

After transformation of P. pastoris with the PRK expression cassette, only 10-15% of the his colonies were unable to grow on methanol. The slow-growth phenotype due to constitutive expression of AOX2 (the other methanol oxidase gene) introduced uncertainty in the scoring of replica plates. Effective distinction was achieved by replicating onto minimal plates lacking any carbon source, but including 100-200 µl of methanol in the lids of the plates, and incubating the plates in an inverted orientation. The AOX1 replicas grow well with methanol provided by evaporation, whereas aox1 replicas do not exhibit growth during 2 days of incubation.

For each construction, crude extracts were prepared from 10-12 methanol-induced his methanol isolates. These extracts were examined for PRK production by Western blotting with a polyclonal anti-PRK antiserum, and the most prolific producer of PRK was selected. These PRK-positive transformants were further characterized by Southern transfer hybridization, PCR amplification of the AOX1 region and DNA sequencing of the amplified segment (Fig. 1). Isolates chosen for production of PRK were devoid of AOX1 sequences and exhibited strong PRK signals. In the methanol-induced cultures, PRK reached 4-6% of the total soluble protein at 60-72 h post-induction. Longer term incubation did not increase the specific activity, and total protein levels declined. SDS-polyacrylamide gel electrophoresis and Western blot analysis of a large scale culture of the wild-type transformant is shown in Fig. 2. Note the absence of significant degradation of the PRK polypeptide chain. The diffuse leading edge of the PRK band detected by Western blotting is apparent even in the enzyme isolated from spinach and hence does not reflect an anomaly unique to the recombinant kinase.


Figure 2: Synthesis of PRK following methanol-induction of transformed P. pastoris. In both panels, lane identifications are as follows: lane 1, purified spinach PRK (specific activity of 500 units/mg); lane 2, extract from untransformed GS115 (specific activity of 0 units/mg); lane 3, uninduced Deltaaox1::PRK extract (specific activity of 0.7 unit/mg); lane 4, Deltaaox1::PRK extract 24 h post-induction (specific activity of 11 units/mg); lane 5, 48 h post-induction (specific activity of 12 units/mg); lane 6, 72 h post-induction (specific activity of 19 units/mg). A, SDS-polyacrylamide gel electropherogram stained with Coomassie Blue; lane 1, 500 ng of purified spinach PRK; lanes 2-6, 10 µg of total protein. B, Western transfer of an electropherogram probed with anti-PRK; lane 1, 50 ng of purified spinach PRK; lanes 2-6, 1 µg of total protein.



Isolation and Authenticity of Recombinant Spinach PRK from Transformed P. pastoris

The protocol described under ``Experimental Procedures'' offers recombinant PRK of high purity in good overall yields (50%) (Table 1). The initial chromatographic step, which utilizes a strong anion-exchange resin, effectively removes 80% of the extraneous proteins, while providing virtually full recovery of kinase activity. As with PRK extracted from spinach, the recombinant enzyme is selectively adsorbed by reactive red agarose, an affinity matrix for nucleotide binding domains, and efficiently eluted by ATP. Based on polyacrylamide gel electrophoresis under both nondenaturing and denaturing conditions, the size and charge of the recombinant enzyme and the PRK isolated from spinach are indistinguishable (Fig. 3). Catalytic equivalence of the two preparations is indicated by their closely matched V(max) and K(m) values (Table 2). The redox characteristic of spinach PRK, as reflected in its extreme sensitivity to DTNB (which oxidizes the regulatory sulfhydryls to an intrasubunit disulfide)(14) , is also embodied by the recombinant enzyme (Fig. 4); inactivation is completely reversed by DTT (data not shown). Thus, diverse structural and functional criteria suggest that the recombinant enzyme is identical to authentic spinach PRK.




Figure 3: Polyacrylamide gel electrophoresis of PRKs. A, Coomassie-Blue-stained nondenaturing gel with all samples at 1 µg/lane. B, silver-stained denaturing gel with all samples at 50 ng/lane. Lanes 1 and 8, molecular markers (Novex, Mark 12); lane 2, authentic spinach PRK; lane 3, wild-type recombinant; lane 4, C16S; lane 5, C55S; lane 6, C16S-C55S; lane 7, C16S-C55S-C244S-C250S.






Figure 4: Inactivation of recombinant PRKs by DTNB. Reaction mixtures contained PRK (10 µM subunits) in 50 mM Bicine (pH 8.0), 20% (v/v) glycerol, and inactivations were initiated by addition of DTNB to a final concentration of 50 µM. Periodically, aliquots were removed and assayed as described under ``Experimental Procedures.'' Samples depicted are as follows: wild-type (bullet), C55S (), C16S (circle), C16S-C55S (box), C16S-C55S-C244S-C250S (up triangle), controls of wild-type and mutant proteins lacking DTNB ().



Characterizations of Cys Ser Mutants of PRK

Spinach PRK contains four cysteinyl residues, which are located at positions 16, 55, 244, and 250(9, 10) . In earlier studies(11, 12) , Cys-16 and Cys-55 (the sites for redox regulation) were replaced individually by Ser via site-directed mutagenesis of E. coli transformation vectors. However, a double mutant was not constructed; thus, the potential synergistic effects from simultaneous loss of both sulfhydryls as occurs during oxidative regulation have not been evaluated. To avoid the inherent risks of comparing mutant enzymes derived from different hosts, at different times, we have again constructed the C16S and C55S single mutants, but by use of the Pichia system concurrently with original constructions of the C16S-C55S double mutant and the C16S-C55S-C244S-C250S quadruple mutant.

The electrophoretic mobilities of the four purified mutant proteins are similar to those of wild-type PRK (Fig. 3). However, each of the three mutants carrying a seryl substitution at position 16 migrates as a poorly resolved doublet under nondenaturing conditions. This microheterogeneity does not appear to be due to contamination by another protein, because the specific activity of C16S approximates that of the wild-type enzyme. Kinetic analyses of the mutant proteins are summarized in Table 2. In reasonable agreement with our earlier report(11) , particularly in view of crude preparations used at that time, C16S is almost fully active, and C55S displays only about 20% of wild-type activity. The activity of the C16S-C55S double mutant is decreased another 2-fold (10% of wild-type), but the quadruple mutant altogether lacking cysteinyl residues is not further impaired. Although none of the cysteinyl substitutions dramatically impact the K(m) for ATP (e.g. the same value was obtained for the quadruple mutant and wild-type enzyme), all three mutants with replacements for Cys-55 have substantially elevated K(m) values for Ru-5-P. These three mutants, in contrast to C16S and wild-type enzyme, are subject to inhibition by ATP at high concentrations in a manner adequately modeled by uncompetitive substrate inhibition.

One set of V(max) and K(m) values for C55S, C16S-C55S, and C16S-C55S-C244S-C250S presented in Table 2are based on velocity measurements at concentrations of Ru-5-P not exceeding 4 mM and fitting the data to a hyperbolic response. However, distinct deviation from hyperbolic kinetics is observed at higher concentrations of Ru-5-P, and saturation is not achieved even at 17 mM Ru-5-P (Fig. 5). Inclusion of sulfate (35 mM) in the assay mixture results in clear-cut hyperbolic kinetics and apparent enhancement of kinase activity at any given concentration of Ru-5-P. Hence, the second set of kinetic parameters presented in Table 2are those determined in the presence of sulfate. The stimulatory effect of sulfate on the three mutants lacking Cys-55 appears to be ion-selective, as chloride is ineffective and phosphate is inhibitory (Fig. 6). Sulfate does not enhance the activity of either the wild-type enzyme or the C16S mutant.


Figure 5: Effect of sulfate on the Ru-5-P-concentration dependence of kinase activity of position 55 mutants. Initial velocities in the presence and absence of 35 mM sodium sulfate are shown. As the relative responses for C55S, C16S-C55S, and C16S-C55S-C244S-C250S were the same, single curves are displayed for the two conditions. See ``Experimental Procedures'' for further details of assays.




Figure 6: Effect of anions on the relative activity of position 55 mutants. In all cases, the Ru-5-P and ATP concentrations were each 1 mM. As the relative responses for C55S, C16S-C55S, and C16S-C55S-C244S-C250S were the same, single curves are displayed for each anion. Error bars denote standard deviation based on averages of data derived from all three position 55 mutants. See ``Experimental Procedures'' for further details of assays.



The double and quadruple mutants (each of which lacks both regulatory sulfhydryls) are impervious to DTNB, while C55S and C16S (each of which retain one regulatory sulfhydryl) display wild-type and diminished sensitivity to DTNB, respectively (Fig. 4). Whereas 2 molar eq of 5-thio-2-nitrobenzoate are formed/mol of wild-type PRK subunit inactivated by DTNB (reflective of intrasubunit disulfide bond formation)(14) , only 1 molar eq of the chromophore is formed during inactivation of the C55S and C16S mutants (reflective of derivatization of the available sulfhydryl group by the reagent moiety) (data not shown). As in the case of the wild-type enzyme, the inactivations of C55S and C16S by DTNB are readily reversed by DTT (data not shown).


DISCUSSION

PRK is one of several chloroplast enzymes to be regulated by thioredoxin (for reviews, see (6) and (23) ). Others include NADP-dependent malate dehydrogenase, fructose-1,6-bisphosphatase, sedoheptulose-1,7-bisphosphatase, and glyceraldehyde-3-phosphate dehydrogenase. Among these enzymes, PRK is the only one in which the regulatory sulfhydryls (Cys-16 and Cys-55) are located at the active site(6, 14) . Hence, the question naturally arises as to whether the complete loss of kinase activity that accompanies disulfide bond formation between Cys-16 and Cys-55 is due to removal of catalytically important sulfhydryl groups or to a conformational change. Prior studies have indicated that both factors contribute to the oxidation-associated deactivation. Analysis of site-directed mutants with single amino acid substitutions for either of the regulatory cysteinyl residues showed that Cys-55 enhances k by 10-fold and that Cys-16 is inconsequential to catalysis(11, 12) . Localized conformational differences between the oxidized and reduced form of the enzyme were invoked by the observed efficient intrasubunit cross-linking of the regulatory sulfhydryls by bifunctional reagents spanning distances as short as 3.5 Å and as long as 9 Å(24) . Thus, a part of the deactivation concomitant with oxidation could be attributed to restricted conformational flexibility imposed by the disulfide bond.

We wished to evaluate further the basis of oxidative deactivation of PRK, because the catalytic consequences of simultaneous replacement of both regulatory sulfhydryls were not explored in earlier mutagenesis studies. Given the paucity of recombinant spinach PRK elaborated by reported E. coli expression systems, we sought a superior host not only to advance the present inquiry but to also facilitate future studies of the structure and function of the kinase.

Our original expression cassette for spinach PRK relied on a tac promoter in concert with a ribosomal binding site to maximize production of the mature form of the enzyme as found in the chloroplast(9, 11) . This promoter-ribosomal binding site unit had been used by our laboratory to express recombinant ribulose-bisphosphate carboxylase/oxygenase in E. coli at levels approaching 20% of the total soluble protein(25) . Levels of PRK expression, however, rarely surpassed 0.5% and frequently were only 0.2% of extractable protein. Even these meager levels of PRK production required cultures to be grown at or below 30 °C in the presence of arabinose, which suppressed the susceptibility of the recombinant enzyme to proteolysis. The arabinose effect could be duplicated in an arabinose-constitutive host without the addition of arabinose to the culture, suggesting that an inducible protein, rather than the sugar or its metabolites, was responsible for the protection. Western blots of crude extracts indicated some degradation, even under ``optimal'' conditions. Mutant PRKs, severely deficient in catalytic activity, were also poorly expressed, suggesting that the recombinant protein itself was eliciting a stress response. Invariably, induction of transcription of the wild-type or mutant genes encoding PRK led to cessation of cell division followed by cell death.

The PRK coding sequence, like many of the spinach genome, utilizes AGA or AGG codons for arginine predominantly (for 13 out of the 14 total arginyl residues), whereas the preferred arginine codons in E. coli are CGN. Arg-23 and Arg-24 of PRK are encoded by AGG; this presence of a tandem of rare codons early in the transcript may lead to ribosome pausing that is deleterious to high levels of expression and may further compromise the host by depleting charged pools of the rare tRNA(26) . Furthermore, the sequence AGGAGG resembles a ribosomal binding site and may lead to unproductive internal initiations on the transcript(27) . In efforts to overcome these potential problems, a clone of the arginyl-tRNA gene, dnaY, was co-expressed with PRK, and codons 23 and 24 were mutated to CGC codons. Neither of these measures, individually or in combination, enhanced PRK expression.

Aside from the tac promoter, several other highly regulated promoters were tried in alternative PRK expression cassettes. Neither the promoter of the trp operon (inducible with indole-acrylic acid), the P(L) promoter (inducible by heat shock in a cI857 host), nor a T7 phage promoter (controlled by induction of T7 RNA polymerase) supported PRK production above 0.2-0.3% of soluble protein.

In view of our inability to overcome these impediments, we turned our attention to eukaryotic host organisms. One system investigated was the GAL1 promoter of Saccharomyces cerevisiae, which is highly induced by galactose. A GAL1 promoter PRK expression cassette was constructed and introduced into the host yeast on a high copy episomal vector. Upon induction with galactose, the level of PRK produced was still only 0.2-0.3% of soluble protein. We were encouraged, however, to note very little degradation of the recombinant enzyme based on Western blots of the crude extract and were thus prompted to evaluate the alcohol oxidase promoter (AOX1) of the methylotrophic yeast P. pastoris as an avenue for obtaining recombinant PRK. Recently, a number of proteins that had proven difficult to produce in standard hosts have been obtained in high yields by use of this system(28) . Similarly, when induced with methanol, the transformant of P. pastoris described herein synthesizes large amounts of PRK (about 5% of total soluble protein), and the synthesis is not compromised by proteolytic degradation. Efficient expression, coupled with facile isolation, enables us to recover about 25 mg of purified PRK (either wild-type or mutants) from 1 liter of cell culture.

In corroboration of our earlier findings, Cys-16 of PRK does not play any role in catalysis. The V(max) of the C16S mutant equals that of the wild-type enzyme, and the very slight increase of the K(m) for ATP with the C16S mutant is not surprising, given the location of Cys-16 within the binding domain for ATP(12, 13, 29, 30) . In both our prior study (11) and an independent one(12) , the C55S mutant was reported to have a V(max) of 10% of the wild-type value. We now observe a value about twice this large with the same mutant constructed in the P. pastoris system. Greater credence can be placed in the present kinetic analyses, because the use of purified proteins eliminates the margin of error introduced by reliance on Western blotting to quantify kinase concentrations in the impure preparations analyzed before. Thus, Cys-55 would still be classified as catalytically facilitative, but its contribution to rate enhancement is maximally 5-fold. This contribution drops to only 2.5-fold if the stimulatory effect of sulfate is taken into account. The substantially increased K(m) for Ru-5-P, with the C55S mutant relative to the wild-type enzyme, is consistent with the assignment of Cys-55 to the binding domain for Ru-5-P(31) .

The C16S-C55S double mutant serves as a better gauge of the combined catalytic contribution of the two regulatory sulfhydryls than the single mutants assessed individually. Barring synergistic effects, insertion of a seryl substitution at position 16 of the C55S mutant would be catalytically negligible. However, the double mutant is only 50% (in the absence of sulfate) or 70% (in the presence of sulfate) as active as C55S, thereby revealing synergism between the two sites of substitution. Even so, the complete inactivation of PRK concomitant with oxidation to the disulfide form cannot be accounted for merely by the absence of free sulfhydryls. The significant catalytic competency retained by the double mutant is consistent with our prior contention that conformational constraints imposed by the disulfide bond contribute to oxidative deactivation of PRK(24) . Retention of activity by the quadruple mutant, devoid of cysteinyl residues, underscores the modest collective contribution of sulfhydryl groups to catalytic turnover.

We assume that the substantial stimulation of kinase activity of the position 55 mutants by sulfate reflects conformational differences between these three mutants and the wild-type enzyme and that sulfate favors the wild-type conformation. Although we have not characterized this phenomenon in detail, we note that sulfate activation of phosphoglycerate kinase has been thoroughly analyzed. In that case, activation is clearly mediated by a sulfate-induced conformational change(32, 33, 34, 35) .

In summary, we have designed the first efficient expression system for a thioredoxin-regulated PRK. We have proceeded to exploit this system via site-directed mutagenesis in order to clarify both the relevance of sulfhydryl groups to kinase activity and the basis of oxidation-associated, regulatory deactivation of PRK.


FOOTNOTES

*
This research was supported jointly by the United States Department of Agriculture under Grant 94-37306-0337 from the National Research Initiative Competitive Grants Program and by the Office of Health and Environmental Research, United States Department of Energy under Contract DE-AC05-84OR21400 with Lockheed Martin Energy Systems, Inc. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore by hereby marked ``advertisement'' in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

§
To whom correspondence and proofs should be addressed: Biology Div., Oak Ridge National Laboratory, P. O. Box 2009, Oak Ridge, TN 37831-8077. Tel.: 423-574-0212; Fax: 423-574-9297; hartmanfc{at}ornl.gov.

(^1)
The abbreviations used are: Ru-5-P, D-ribulose 5-phosphate; PRK, phosphoribulokinase; DTT, dithiothreitol; DTNB, 5,5`-dithiobis(2-nitrobenzoic acid); Bicine, N, N`-bis(2-hydroxyethyl)glycine; PCR, polymerase chain reaction; AOX, alcohol oxidase; MES, 2-(N-morpholino)ethanesulfonic acid.


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