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• W2170585870 abstract "ADP-glucose pyrophosphorylase (AGPase) is a key regulatory enzyme of bacterial glycogen and plant starch synthesis as it controls carbon flux via its allosteric regulatory behavior. Unlike the bacterial enzyme that is composed of a single subunit type, the plant AGPase is a heterotetrameric enzyme (α2β2) with distinct roles for each subunit type. The large subunit (LS) is involved mainly in allosteric regulation through its interaction with the catalytic small subunit (SS). The LS modulates the catalytic activity of the SS by increasing the allosteric regulatory response of the hetero-oligomeric enzyme. To identify regions of the LS involved in binding of effector molecules, a reverse genetics approach was employed. A potato (Solanum tuberosum L.) AGPase LS down-regulatory mutant (E38A) was subjected to random mutagenesis using error-prone polymerase chain reaction and screened for the capacity to form an enzyme capable of restoring glycogen production in glgC−Escherichia coli. Dominant mutations were identified by their capacity to restore glycogen production when the LS containing only the second site mutations was co-expressed with the wild-type SS. Sequence analysis showed that most of the mutations were decidedly nonrandom and were clustered at conserved N- and C-terminal regions. Kinetic analysis of the dominant mutant enzymes indicated that theKm values for cofactor and substrates were comparable with the wild-type AGPase, whereas the affinities for activator and inhibitor were altered appreciably. These AGPase variants displayed increased resistance to Pi inhibition and/or greater sensitivity toward 3-phosphoglyceric acid activation. Further studies of Lys-197, Pro-261, and Lys-420, residues conserved in AGPase sequences, by site-directed mutagenesis suggested that the effectors 3-phosphoglyceric acid and Pi interact at two closely located binding sites. ADP-glucose pyrophosphorylase (AGPase) is a key regulatory enzyme of bacterial glycogen and plant starch synthesis as it controls carbon flux via its allosteric regulatory behavior. Unlike the bacterial enzyme that is composed of a single subunit type, the plant AGPase is a heterotetrameric enzyme (α2β2) with distinct roles for each subunit type. The large subunit (LS) is involved mainly in allosteric regulation through its interaction with the catalytic small subunit (SS). The LS modulates the catalytic activity of the SS by increasing the allosteric regulatory response of the hetero-oligomeric enzyme. To identify regions of the LS involved in binding of effector molecules, a reverse genetics approach was employed. A potato (Solanum tuberosum L.) AGPase LS down-regulatory mutant (E38A) was subjected to random mutagenesis using error-prone polymerase chain reaction and screened for the capacity to form an enzyme capable of restoring glycogen production in glgC−Escherichia coli. Dominant mutations were identified by their capacity to restore glycogen production when the LS containing only the second site mutations was co-expressed with the wild-type SS. Sequence analysis showed that most of the mutations were decidedly nonrandom and were clustered at conserved N- and C-terminal regions. Kinetic analysis of the dominant mutant enzymes indicated that theKm values for cofactor and substrates were comparable with the wild-type AGPase, whereas the affinities for activator and inhibitor were altered appreciably. These AGPase variants displayed increased resistance to Pi inhibition and/or greater sensitivity toward 3-phosphoglyceric acid activation. Further studies of Lys-197, Pro-261, and Lys-420, residues conserved in AGPase sequences, by site-directed mutagenesis suggested that the effectors 3-phosphoglyceric acid and Pi interact at two closely located binding sites. ADP-glucose pyrophosphorylase 3-phosphoglyceric acid large subunit small subunit polymerase chain reaction ADP-glucose pyrophosphorylase (AGPase)1 controls the synthesis of glycogen and starch in bacteria and plants, respectively. It catalyzes the formation of ADP-glucose from glucose 1-phosphate (Glc-1-P) and ATP with the concomitant release of pyrophosphate (PPi) (1Kavakli I.H. Slattery C.J. Hiroyuki I. Okita T.W. Aust. J. Plant Physiol. 2000; 27: 561-570Google Scholar, 2Slattery C.S. Kavakli I.H. Okita T.W. Trends Plant Sci. 2000; 5: 291-298Abstract Full Text Full Text PDF PubMed Scopus (146) Google Scholar, 3Preiss J. Sivak M. Zamski E. Schaffer A.A. Photoassimilate Distribution in Plants and Crops: Source-sink Relationships. Marcel Dekker, Inc., New York1996: 139-168Google Scholar). The product ADP-glucose then serves as the glucosyl donor for the formation of α-1,4-glucosyl chains by glycogen/starch synthase. Both bacterial and plant AGPases are allosterically regulated by small effector molecules whose nature reflects the primary carbon assimilatory pathway present in these organisms. Despite sharing sequence homology and similar catalytic and regulatory properties, the bacterial and plant AGPases have different quaternary structures. The bacterial AGPases are composed of four identical subunits with an approximate subunit molecular mass of 48 kDa (4Preiss J. Annu. Rev. Microbiol. 1984; 38: 419-458Crossref PubMed Scopus (282) Google Scholar). In contrast, the heterotetrameric plant enzyme is composed of a pair of large (LS) and small subunits (SS), encoded by different genes. The molecular mass of the plant LSs range from 51 to 60 kDa, whereas the SSs are from 50 to 54 kDa (5Nakata P.A. Greene T.W. Anderson J.M. Smith-White B.J. Okita T.W. Preiss J. Plant Mol. Biol. 1991; 17: 1089-1093Crossref PubMed Scopus (76) Google Scholar, 6Smith-White B.J. Preiss J. J. Mol. Evol. 1992; 34: 449-464Crossref PubMed Scopus (178) Google Scholar). Different approaches have been utilized in attempts to decipher the role of the two subunit types in higher plant AGPase function. Pyridoxal phosphate, a structural analogue of the activator 3-PGA, labels both the LS and SS of the spinach leaf isoform, suggesting that both subunits bind the activator (7Morell M.K. Bloom M. Knowles V. Preiss J. Plant Physiol. 1987; 85: 182-187Crossref PubMed Google Scholar). Further analysis showed that pyridoxal phosphate only labeled a single residue in the spinach leaf SS (which aligns with Lys-429 in the potato SS, see Fig.2), whereas three Lys residues (which align with Lys-124, Lys-414, and Lys-452 in the potato LS, Fig. 2) were labeled in the spinach leaf LS (8Ball K.L. Preiss J. J. Biol. Chem. 1994; 269: 24706-24711Abstract Full Text PDF PubMed Google Scholar, 9Preiss J. Stark D. Barry G.F. Guan H.P. Libal-Weksler Y. Sivak M.N. Okita T.W. Kishore G.M. Henry R. Ronalds J.A. Proceedings of the Improvement of Cereal Quality by Genetic Engineering. Plenum Publishing Corp., New York1994: 115-127Google Scholar). The multiple labeling patterns of the LS suggest that this subunit type plays a more dominant role in allosteric regulation than the small subunit, a view supported by genetic studies (10Preiss J. Miflin B.J. Oxford Surveys of Plant Molecular and Cellular Biology. Oxford University Press, Oxford1992: 59-114Google Scholar, 11Preiss J. Ball K. Smithwhite B. Iglesias A. Kakefuda G. Li L. Biochem. Soc. Trans. 1991; 19: 539-547Crossref PubMed Scopus (66) Google Scholar). A missense mutation in the LS gene of Arabidopsis caused only a partial reduction of AGPase activity and starch levels in leaves (12Lin T.-P. Caspar T. Somerville C.R. Preiss J. Plant Physiol. 1988; 88: 1175-1181Crossref PubMed Google Scholar). The isolated enzyme, which was found to be composed of only the SS, required much higher levels of 3-PGA for activation as compared with the wild-type (wild type) Arabidopsis leaf AGPase (13Li L. Preiss J. Carbohydr. Res. 1992; 227: 227-239Crossref Scopus (34) Google Scholar). Similarly, bacterial expression of the SS and LS inglgC− Escherichia coli AC70R1-504 (an AGPase deficient strain) indicated that the SS alone is able to form an active homotetrameric enzyme, although requiring much higher levels of 3-PGA for activation and lower levels of Pi for inhibition, as compared with the heterotetrameric wild-type enzyme (14Ballicora M.A. Fu Y. Wu M.-X. Sheng J. Nesbitt N.M. Preiss J. Nakamura Y. The 5th NIAR/COE International Symposium on the Regulation and Manipulation of Starch and Sucrose Metabolism in Plants. National Institute of Agriobiological Resources, Tsukuba, Japan1996: 5-11Google Scholar,15Salamone P.R. Greene T.W. Kavakli I.H. Okita T.W. FEBS Lett. 2000; 482: 113-118Crossref PubMed Scopus (33) Google Scholar). These results suggest that the SS is involved in both catalytic and regulatory functions, whereas the LS is involved primarily in regulation. The ability to produce an active plant enzyme in bacteria, capable of complementing a glgC (structural gene for AGPase) mutation and restoring glycogen production, enabled a biochemical-genetic approach to understand the role of the AGPase subunits in enzyme function. For example, the LS or SS cDNA of the potato AGPase was subjected to chemical mutagenesis with hydroxylamine and co-expressed with the wild-type counterpart subunit in aglgC−E. coli. With this technique, both substrate and effector binding mutants have been identified. Laughlin et al. (16Laughlin M.J. Chantler S.E. Okita T.W. Plant J. 1998; 14: 159-168Crossref PubMed Scopus (39) Google Scholar, 17Laughlin M.J. Payne J.W. Okita T.W. Phytochemistry. 1998; 47: 621-629Crossref PubMed Scopus (25) Google Scholar) isolated several potato SS mutants, which had defective substrate (ATP or Glc-1-P) binding properties. Additionally, random mutagenesis using the LS as a template yielded several up-regulated and down-regulated potato AGPase mutants (18Greene T.W. Chantler S.E. Kahn M.L. Barry G.F. Preiss J. Okita T.W. Proc. Natl. Acad. Sci. U. S. A. 1996; 93: 1509-1513Crossref PubMed Scopus (50) Google Scholar, 19Greene T.W. Woodbury R.L. Okita T.W. Plant Physiol. 1996; 112: 1315-1320Crossref PubMed Scopus (27) Google Scholar, 20Greene T.W. Kavakli I.H. Kahn M.L. Okita T.W. Proc. Natl. Acad. Sci. U. S. A. 1998; 95: 10322-10327Crossref PubMed Scopus (47) Google Scholar). Overall, results from these biochemical and genetic studies indicate that the two subunit types play different roles in enzyme function. The SS contains both regulatory and catalytic sites, whereas the LS contributes a regulatory function by modulating the catalytic activity of the SS by increasing the allosteric regulatory response. Because of the lack of structural information, the study of mutant enzymes generated by random mutagenesis is a plausible approach to understand enzyme function. However, this approach requires the ability to generate numerous mutations. Although hydroxylamine treatment has been employed to mutagenize LS, it is limited by the narrow spectrum (GC to AT transitions) of mutations generated. To map amino acids involved in effector binding in the potato LS, a reverse genetics approach was employed where a “loss of function” LS (E38A) was subjected to error-prone PCR and co-expressed with the wild-type SS. Revertants that stained darkly by iodine vapor were selected and further studied for the presence of dominant second-site mutations that conferred elevated glycogen accumulation than the wild-type condition. Sixteen dominant LS mutants, which fell into 11 classes, were identified with most of the second-site mutations clustered in two regions of the primary sequence. Kinetic analysis of these 11 AGPase classes showed that the dominant LS mutations conferred an up-regulatory phenotype, i.e. increased sensitivity to the activator 3-PGA and/or increased resistance to the inhibitor Pi. Random mutagenesis of the E38A LS cDNA was performed according to Cadwell and Joyce (21Cadwell R.C. Joyce G.F. PCR Methods Appl. 1992; 2: 28-33Crossref PubMed Scopus (834) Google Scholar) with modifications. 30 fmol of E38A LS in pML7 plasmid was amplified using the primers LSUampS and PLS-2 (Table I). The PCR mixture contained 1× mutagenic buffer (7 mm MgCl2, 0.5 mm MnCl2, 50 mm KCl, 10 mm Tris-HCl, pH 8.3, and 0.1% (w/v) gelatin), 0.2 mm dGTP, 0.2 mm dATP, 1 mm dCTP, 1 mm dTTP, 30 pmol of each primer, and 5 units of Taq polymerase. PCR was performed using a Promega Robo Cycler for 14 cycles at the following conditions: 40 s at 94 °C, 40 s at 61 °C, and 1.5 min 72 °C. PCR products were purified using a Qiagen PCR clean-up kit. The amplified products were digested with NcoI and HindIII, cloned into the corresponding sites of pML7 vector (22Ballicora M.A. Laughlin M.J. Fu Y. Okita T.W. Barry G.F. Preiss J. Plant Physiol. 1995; 109: 245-251Crossref PubMed Scopus (124) Google Scholar), and transformed intoglgC−E. coli, carrying the SS expression plasmid pML10. Revertants were identified by their ability to complement the glgC− phenotype thereby restoring glycogen production, which is readily scored by iodine staining of the bacterial colonies.Table IOligonucleotide primers used for amplification of the LS cDNA and generation of site-directed mutationsLSU ampS5′-GGGCCATGGCTTACTCTGTGATCACTA-3′PLS25′-GCTTATCATCGATAAGCTTCCTTCGG-3′A38E S5′-CTGGGAGGAGGAGAAGGGACCAAGTTATTC-3′A38E AS5′-GAATAACTTGGTCCCTTCTCCTCCTCCTCCCAG-3′K420A S5′-CGCAAAGATAGGAGCGAATGTTTCAATCATAAATAAAG-3′K420A AS5′-CTTTATTTATGATTGAAACATTCGCTCCTATCTTTGCG-3′K420E S5′-CGCAAAGATAGGAGAGAATGTTTCAATCATAAATAAAG-3′K420E AS5′-CTTTATTTATGATTGAAACATTCTCTCCTATCTTTGCG-3′M197I S5′-GATTTTGGGCTGGTCATAATTGACAGCAG-3′M197I AS5′-CTGCTGTCAATTATGACCAGCCCAAAATC-3′K197E S5′-GATTTTGGGCTGGTCGAGATTGACAGCAG-3′K197E AS5′-CTGCTGTCAATCTCGACCAGCCCAAAATC-3′P264E S5′-TTGAAATGGAGCTATGAAACTTCTAATGATTTTG-3′P264E AS5′-CAAAATCATTAGAAGTTTCATAGCTCCATTTCAA-3′P264G S5′-TTGAAATGGAGCTATGGGACTTCTAATGATTTTG-3′P264G AS5′-CAAAATCATTAGAAGTCCCATAGCTCCATTTCAA-3′P264K5′-TTGAAATGGAGCTATAAGACTTCTAATGATTTTG-3′P264K AS5′-CAAAATCATTAGAAGTTTCATAGCTCCATTTCAA-3′Underlined and bold nucleotides indicate the nucleotides used to replace the wild-type amino acid. S, sense; AS, antisense. Open table in a new tab Underlined and bold nucleotides indicate the nucleotides used to replace the wild-type amino acid. S, sense; AS, antisense. Site-directed mutagenesis was carried out using the Stratagene Quick-change Mutagenesis kit. The PCR reaction contained 30 fmol of DNA, 20 pmol of primers (Table I), 0.2 mm dNTPs, and 2.5 units of Pfu Turbo DNA polymerase. The PCR was carried out for 12 cycles under the following conditions: 40 s at 94 °C, 40 s at 55 °C, and 11 min at 68 °C. The PCR products were digested with DpnI to remove template plasmid DNA and transformed into E. coli DH5α. The presence of the site-directed mutations was confirmed by DNA sequencing through the Washington State University DNA sequencing facility. Glycogen was quantified using a glucose oxidase assay kit (Sigma). Cells were grown in 2% Kornberg's media (23Govons S. Vinopal R. Ingraham J. Preiss J. J. Bacteriol. 1969; 97: 970-972Crossref PubMed Google Scholar) containing the appropriate antibiotics for selection. Cells (500 mg) were harvested by centrifugation, resuspended in water, and then lysed by incubating at 100 °C. The samples were solubilized in 2 ml of Me2SO and 0.5 ml of HCl at 60 °C for 30 min, and then the pH was adjusted to 4.5 using NaOH in sodium citrate buffer. The samples were then treated with 3 units of amyloglucosidase at 55 °C for 40 min to hydrolyze glycogen into d-glucose. The amount of d-glucose was measured spectrophotometrically by measuring the formation of the quinoneimine dye using a coupled enzyme assay containing glucose oxidase and peroxidase. The activity of ADP-glucose pyrophosphorylase was determined in both the reverse (pyrophosphorylase: Assay A) and the forward (synthesis: Assay B) directions. Pyrophosphorylase assays were used to monitor purification of the enzyme. [32P]ATP formation was measured from [32P]PPi and ADP-glucose. The reaction mixture contained 50 mm HEPES, pH 7.5, 0.4 mg/ml bovine serum albumin, 5 mm MgCl2, 3 × 106 cpm/ml [32P]Pi, 1.5 mm NaPPi, 5 mm 3-PGA, 5 mm dithiothreitol, 1 mm ADP-glucose, 10 mm NaF, and 50–100 mg/ml protein in a final volume of 0.25 ml. Kinetic parameters were defined by the synthesis assay, which measured [14C]Glc-1-P incorporation into ADP-glucose. The reaction mixture contained 50 mm HEPES, pH 7.5, 0.4 mg/ml bovine serum albumin, 5 mmMgCl2, 5 mm 3-PGA, 5 mmdithiothreitol, 0.5 mm Glc-1-P, 0.2 units of inorganic pyrophosphatase, 1.5 mm ATP, 0.5 µCi of [14C]Glc-1-P in a final volume of 0.1 ml. Reactions were incubated at 37 °C for 10 min. Calf intestinal alkaline phosphatase (2 units/reaction in 1× CIAP buffer) was added to each reaction and incubated at 37 °C for 1 h. A reaction sample volume of 0.055 ml was blotted onto DEAE-81 paper and washed 3 times with distilled water. The filter was dried and product formation quantified by liquid scintillation spectrometry. Cells were grown in 1 liter of modified LB medium (23Govons S. Vinopal R. Ingraham J. Preiss J. J. Bacteriol. 1969; 97: 970-972Crossref PubMed Google Scholar) to anA600 of 1.2. AGPase expression was induced by the addition of 10 µg/ml nalidixic acid and 200 µm isopropyl-1-thio-β-d-galactopyranoside (final concentrations) and incubated with shaking at room temperature for 18 h. Cells were harvested by centrifugation at 6,000 ×g for 5 min, and the cell pellet was resuspended in lysis buffer (50 mm HEPES, pH 8.0, 5 mmMgCl2, 5 mm dithiothreitol, 1 mmEDTA, 10% glycerol) containing 500 µg/ml lysozyme, 0.5 mg/ml pepstatin, 0.5 mg/ml leupeptin, 0.5 mm benzamidine, and 1 mm phenylmethylsulfonyl fluoride. The sample was sonicated 3 times for 45 s and centrifuged at 30,000 × gfor 20 min at 4 °C. The supernatant was passed through a 0.2-µm filter, loaded onto an HQ-POROS anion exchange column equilibrated with buffer A (50 mm HEPES, pH 8.0, 5 mmMgCl2, 1 mm EDTA, 10% glycerol), and eluted from the column by a 200-ml linear gradient of buffer B (buffer A, pH 7.0, containing 1 m KCl). Active fractions were combined, and (NH4)2SO4 was added to a final concentration of 1 m. Samples were then loaded onto a C-4 hydrophobic interaction chromatography column, which was equilibrated with buffer C (50 mm HEPES, pH 7.0, 5 mmMgCl2, 1 mm EDTA, 1 m(NH4)2SO4, and 10% glycerol). Enzyme elution was carried out using a linear 200-ml linear gradient with buffer D (50 mm HEPES, pH 7.0, 5 mmMgCl2, 1 mm EDTA, 10% glycerol). Active fractions were pooled and dialyzed against buffer A overnight. Aliquots of concentrated enzyme were stored at −80 °C. Protein levels were determined by Bradford analysis (24Bradford M.M. Anal. Biochem. 1976; 72: 248-254Crossref PubMed Scopus (216377) Google Scholar) using the Bio-Rad reagent. Purity and integrity of partially purified enzymes were determined by analysis on 12% SDS-polyacrylamide gels followed by Coomassie Brilliant Blue staining and immunoblot analysis with antibodies specific for the potato AGPase LS and SS. Km and A0.5 values, corresponding to levels required for 50% maximal activity, were calculated using Lineweaver-Burk plots generated by the computer program Enzyme Kinetics 4.0 (Trinity software). Kmvalues for Mg2+ were determined using nonlinear regression plots in Excel (Microsoft). Inhibition kinetics (I0.5) corresponding to the inhibitor level required for 50% inhibition were calculated from the inhibition curve at specified allosteric activator levels using Cricket graph. Previous mutagenesis studies resulted in the identification of an LS mutant, which contained a single amino acid replacement, E38A. When co-expressed with the wild-type SS, the resulting AGPase required several hundred-fold higher levels of 3-PGA for activation than the wild-type enzyme (20Greene T.W. Kavakli I.H. Kahn M.L. Okita T.W. Proc. Natl. Acad. Sci. U. S. A. 1998; 95: 10322-10327Crossref PubMed Scopus (47) Google Scholar). Bacterial cells expressing this mutant LS with wild-type SS were null for glycogen accumulation when analyzed by iodine staining. To map the peptide regions that participate in allosteric effector binding, a second site genetic reversion approach was utilized. The LS E38A was subjected to a modified error-prone PCR protocol and co-expressed with the wild-type SS. Primary screening of putative allosteric mutants was achieved by iodine staining. Because cells co-expressing E38A LS and the wild-type SS are unable to accumulate glycogen and hence lack iodine staining, any colonies stained by iodine vapor were assumed to contain a second site mutation(s) in the LS that restored AGPase activity and, in turn, substantial quantities of glycogen. Approximately 12,000 colonies were examined by iodine staining, and of these, 30 colonies that accumulate high amounts of glycogen were identified. The various mutant LS cDNAs were then purified and subjected to site-directed mutagenesis to convert the primary mutation Ala-38 back to the corresponding wild-type residue (Glu) using the primers A38G sense and A38E antisense (Table I). The LS cDNAs containing only the secondary mutations were then co-expressed with the wild-type SS, and their activity was assessed by iodine staining. Iodine staining results indicated that 16 revertants contained dominant second-site mutations and accumulated more glycogen with respect to the wild-type LS based on increased iodine staining. Ten revertants stained as dark as wild type, whereas four revertants lost their iodine staining phenotype (data not shown). Glycogen amounts were measured enzymatically. Co-expression of the 16 LS dominant mutants with the wild-type SS in glgC−E. coli resulted in these bacterial cells accumulating more glycogen than cells co-expressing the wild-type LS and SS. DNA sequence analysis of the 16 dominant LS mutants indicated that they fell into 11 mutation classes (Table II). Some lines for example, 18, 20, and 23, have the same multiple amino acid changes, yet they are different independent mutant isolates because of the presence of different silent mutations in their coding regions (data not shown).Table IIAmino acid replacements in the 16 dominant mutantsCloneWild-type residuePositionReplacement residue2Lys420Arg11/30Lys197Met12Asp367AsnIle412Val13/22Asn175HisSer390ProGlu403Val14Pro261Leu15/16Asn145HisIle424Val17Phe193Tyr18/20/23Ser386Pro24Thr11SerSer73Cys25Ser390Pro26Ser188CysMet377ThrSeveral of the individually isolated mutants shared identical amino acid changes (e.g. 11 and 30). Open table in a new tab Several of the individually isolated mutants shared identical amino acid changes (e.g. 11 and 30). As shown in Fig. 1, the mutations were not randomly distributed, but instead 9 of the 11 mutations were clustered at two specific regions of the primary sequence, one located within the N-terminal half of the primary sequence and the other near the C terminus. The two exceptions were T11S, which contained one of the two mutations near the N terminus, and P261L, which is located near the middle (residue 261) of the polypeptide. Five of the mutations were clustered between residues 145 and 197. Even within this N-terminal region, three of the mutations occurred within 10 nucleotides that coded a region strongly conserved in the various plant AGPases. Interestingly, when mutations were mapped in the predicted secondary structure, almost all mutations were located in putative loop regions with the exception of K197M, which is located in a region having a β-sheet (Fig. 1). Likewise, mutations in the C-terminal half of the primary sequence were clustered between residues 367 and 425, with three changes occurring between residues 412 and 424 (Fig. 1). Overall, these data indicate that several loops contained within two regions of AGPase are important for the binding of allosteric effectors. The catalytic and allosteric regulatory properties of the 11 mutants AGPases were determined. Mutants were co-expressed with the wild-type SS in glgC−E. coli and partially purified using HQ POROS (anion exchange) and C-4 (hydrophobic interaction) chromatography with a final purity of ∼20%. Because degraded AGPase exhibits altered kinetic behavior (15Salamone P.R. Greene T.W. Kavakli I.H. Okita T.W. FEBS Lett. 2000; 482: 113-118Crossref PubMed Scopus (33) Google Scholar, 25Plaxton W.C. Preiss J. Plant Physiol. 1987; 83: 105-112Crossref PubMed Google Scholar, 26Kleczkowski L.A. Villand P. Luthi E. Olsen O.-A. Preiss J. Plant Physiol. 1993; 101: 179-186Crossref PubMed Scopus (137) Google Scholar), all mutant enzymes were subjected to SDS-polyacrylamide gel electrophoresis and subsequent analysis by immunoblot using potato anti-LS and anti-SS antibodies. No apparent degradation was evident for any of the mutants analyzed (results not shown). Km values for the substrates (ATP and Glc-1-P) and co-factor (Mg2+) of the various mutants were comparable with wild-type values (Table III). In contrast, changes in the affinity toward the allosteric activator 3-PGA and inhibitor Pi were apparent and varied depending on the mutant types. Two general classes of allosteric regulatory response mutants were evident. One class (mutants K420R, N145H/I424V, and S390P) showed similar 3-PGA activation responses (A0.5values) equivalent to the wild-type value (Table III) but required more than 3.3-fold Pi, respectively, to achieve 50% inhibition in the presence of 0.1 mm 3-PGA (Table III). The second class consisting of the remaining mutants showed alterations in both 3-PGA activation and Pi inhibition. They required ∼2-fold less 3-PGA to reach 50% maximal activation, and their I0.5values were 3.5-fold higher than the wild-type AGPase (TableIV). Among the second class, mutants D367N/I412V, F193Y, and N175H/S390P/E403V were the most sensitive to 3-PGA activation and the most resistant to Pi inhibition, requiring at least 5-fold less 3-PGA to achieve 50% activation and more than 6.5-fold greater levels of Pi for 50% inhibition than the wild-type enzyme (Table III).Table IIIKinetic parameters of the wild-type and mutant AGPasesWild-typeK420RK197MD367N/I412VN175H/S390P/E403VP261LN145H/I424VF193YS386PT11S/S73CS390PS188C/M377TATP (Km)0.30.350.30.250.290.190.310.190.280.180.180.3Glc-1-P (Km)0.260.250.230.180.350.240.250.170.260.230.230.27Mg2+(Km)2.53.13.32.12.22.52.52.12.51.92.43.23-PGA (A0.5)0.10.10.040.020.020.050.070.010.030.040.090.03Pi (0.1 mm3-PGA)0.090.70.190.50.60.20.60.60.20.30.30.24Pi (0.25 mm3-PGA)0.251.8ND3-aND, not determined.NDND0.85NDNDND0.8NDNDPi (1.0 mm3-PGA)NDNDND3.22.9ND3.55.92.9ND2.53.3The kinetic and regulatory properties of the mutants and wild-type are determined in this study. All values (in mm) were determined at least twice with S.D. less than 10% in all cases.3-a ND, not determined. Open table in a new tab Table IV3-PGA and Pi inhibition values of the wild-type and mutantsLSA0.5I0.5(mm)[3-PGA]/I0.5I0.5(mm)[3-PGA]/I0.5µmWild-type1000.09 at 0.1 mm 3-PGA1.110.26 at 0.25 mm3-PGA0.96K420R1000.68 at 0.1 mm3-PGA0.151.80 at 0.25 mm 3-PGA0.14K420A130.22 at 0.1 mm 3-PGA0.450.49 at 0.25 mm 3-PGA0.50K420E1150.26 at 0.1 mm 3-PGA0.380.75 at 0.50 mm3-PGA0.67K197M400.19 at 0.04 mm3-PGA0.21K197I320.50 at 0.1 mm3-PGA0.21.75 at 0.5 mm 3-PGA0.28K197E130.60 at 0.1 mm 3-PGA0.162.40 at 0.5 mm 3-PGA0.20P261L400.19 at 0.04 mm 3-PGA0.210.85 at 0.25 mm3-PGA0.29P261G15000.60 at 1.50 mm3-PGA2.50.90 at 3 mm 3-PGA3.34P261E6000.40 at 0.6 mm 3-PGA1.51.20 at 3 mm 3-PGA2.1P261K3000.16 at 0.30 mm 3-PGA1.924 (T11S/S73C)410.29 at 0.1 mm 3-PGA0.330.79 at 0.25 mm3-PGA0.3124 A (T11S)220.40 at 0.1 mm3-PGA0.251.90 at 0.50 mm 3-PGA0.25All LS mutants were co-expressed with the wild-type SS in glgC−E. coli and partially purified for kinetic analysis. All values are determined from synthesis assay data of at least two iterations, and the difference was <10% in all cases. The average values are shown below. The [3-PGA]/I0.5 ratio is an indicator of the general regulatory properties of the enzyme. It was obtained by measuring the I0.5 value (i.e. the amount of Pi that inhibits enzyme activity 50%) in the presence of a known quantity of 3-PGA. Open table in a new tab The kinetic and regulatory properties of the mutants and wild-type are determined in this study. All values (in mm) were determined at least twice with S.D. less than 10% in all cases. All LS mutants were co-expressed with the wild-type SS in glgC−E. coli and partially purified for kinetic analysis. All values are determined from synthesis assay data of at least two iterations, and the difference was <10% in all cases. The average values are shown below. The [3-PGA]/I0.5 ratio is an indicator of the general regulatory properties of the enzyme. It was obtained by measuring the I0.5 value (i.e. the amount of Pi that inhibits enzyme activity 50%) in the presence of a known quantity of 3-PGA. Several clones have multiple mutations. For example, clone 24 has two amino acid changes, T11S and S73C. To describe the effects of these two mutations independently, the wild-type pML7 and mutant 24 LS plasmid DNAs were digested with PstI/HindIII, and the 1.3-kilobase pair DNA fragments were exchanged between the two plasmids. Isolation of the T11S and S73C mutations on separate vectors (24A and 24B, respectively) were verified by DNA sequencing. Iodine staining results show that cells containing T11S (mutant 24A) LS accumulated higher amounts of glycogen than wild-type, whereas cells containing S73C (24B) LS did not accumulate glycogen, indicating that this mutation resulted in a defective enzyme. Kinetic analysis of 24A indicated that this single amino acid change conferred sensitivity to allosteric effectors, by requiring less 3-PGA for activation and more Pi for inhibition when compared with mutant 24 and wild-type AGPase (Table IV). To explore further the structure-function relati" @default.
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