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THhSIS UNIVERS TYL IIIIIIIIIIIIIIIIIIIIIII IIIII IIIIIIIIIIII 3 1293 01405 7693 This is to certify that the thesis entitled Biocatalytic Production of Arcmatics from D-Glucose presented by Michael Anthony Farabaugh has been accepted towards fulfillment of the requirements for M.S. degree in _Chem1$:l'.n¥__' firm Major professor Date [4/3/96 / I 0-7639 MS U is an Affirmative Action/Equal Opportunity Institution LIBRARY Michigan State University PLACE IN RETURN BOX to remove this checkout from your record. TO AVOID FINES return on or before date duo. DATE DUE DATE DUE DATE DUE MSU I. An Affirmative Adlai/Equal Opportunity Instituion BIOCATALYTIC PRODUCTION OF AROMATICS FROM D-GLUCOSE By Michael Anthony Farabaugh A THESIS Submitted to Michigan State University in partial fulfillment of the requirements for the degree of MASTER OF SCIENCE Department of Chemistry 1996 ABSTRACT BIOCATALYTIC PRODUCTION OF AROMATICS FROM D-GLUCOSE By Michael Anthony Farabaugh Transaldolase catalysis had a notable impact on the yield of aromatics produced from D-glucose. This effect is attributed to the fact that transaldolase catalyzes the formation of D-erythrose 4-phosphate and can increase the in vivo availability of this metabolite. The carbon flow entering the common pathway of aromatic amino acid biosynthesis was quantitated by measuring the concentration of 3-dehydroshikimate (DHS) in the culture supernatant of Escherichia coli aroE strains. Under certain conditions, overexpression of transaldolase raised the yield of DHS to near theoretical maximum. The Citrobacterfreundii gene encoding tyrosine phenol-lyase (tpl) was introduced into an E. coli construct that synthesizes catechol in order to achieve a biocatalytic synthesis of L-3,4-dihydroxyphenylalanine (L-DOPA) from D-glucose. The synthesis was impeded by an unfavorable reaction equilibrium. In the presence of catechol, cells expressing tyrosine phenol-lyase synthesized L-DOPA in 12% yield. To my wife Karen, for her constant love and support. ACKNOWLEDGMENTS I would like to thank Professor John Frost for his enthusiasm and encouragement. I would also like to acknowledge all the members of the Frost group who have given me their advice, support, and friendship. I wish them the very best in their endeavors. iv TABLE OF CONTENTS LIST OF TABLES .................................................................................. vii LIST OF FIGURES ............................................................................... viii LIST OF ABBREVIATIONS ...................................................................... ix CHAPTER 1 INTRODUCTION ................................................................................... 1 The Common Pathway ......................................................................... 1 Synthesis of Aromatics with E. coli. ......................................................... 2 Increasing In Vivo Availability of D-Erythrose 4-Phosphate .............................. 4 Increasing In Vivo Availability of Phosphoenolpyruvate .................................. 6 Rate-Limiting Enzymes of the Common Pathway .......................................... 8 Assembly of a Heterologous Microbial Biocatalyst ......................................... 9 References ...................................................................................... 14 CHAPTER 2 IMPACT OF TRANSALDOLASE ON THE BIOCATALYTIC PRODUCTION OF AROMATICS ......................................... 17 Background ..................................................................................... 17 Overexpression of Transaldolase in E. coli. ................................................ 19 Impact of Transaldolase on Aromatic Production .......................................... 22 Discussion ...................................................................................... 37 References ...................................................................................... 39 CHAPTER 3 PRODUCTION OF L-DOPA WITH TYROSINE PHENOL-LYASE ........................ 41 Background ..................................................................................... 41 Overexpression of Tyrosine Phenol-Lyase in E. coli ...................................... 43 Exploiting Tyrosine Phenol-Lyase for the Synthesis of L-DOPA ........................ 46 Discussion ...................................................................................... 51 References ...................................................................................... 53 CHAPTER 4 EXPERIMENTAL .................................................................................. 55 General Methods ............................................................................... 55 Chromatography .......................................................................... 55 Spectroscopic Measurements ............................................................ 55 Bacterial Strains ........................................................................... 56 Storage of Bacterial Strains and Plasmids ............................................. 56 Culture Medium ........................................................................... 56 Culture Conditions ....................................................................... 57 V 1H NMR Analysis of Culture Supernatant ............................................ 57 General Genetic Manipulations ......................................................... 58 Large Scale Purification of Plasmid DNA ........................................ 58 Small Scale Purification of Plasmid DNA ........................................ 6O Restriction Enzyme Digestion of DNA ........................................... 61 Agarose Gel Electrophoresis ....................................................... 61 Isolation of DNA from Agarose ................................................... 61 Treatment of Vector DNA with Calf Intestinal Alkaline Phosphatase ......... 62 Ligation of DNA ..................................................................... 62 Preparation and Transformation of Competent Cells ............................ 62 Purification of Genomic DNA ..................................................... 63 Enzyme Assays ........................................................................... 65 Chapter 2 ........................................................................................ 65 D-Erythrose 4-Phosphate ................................................................ 65 Assays ..................................................................................... 66 DAH(P) ............................................................................... 66 DAHP synthase ...................................................................... 66 Transaldolase ........................................................................ 67 Transketolase ........................................................................ 67 PCR Amplification of talB, tktA, and aroF ............................................ 68 Plasmid Constructions ................................................................... 7O pMF61A .............................................................................. 7O pMF6OA .............................................................................. 7O pMF63A .............................................................................. 71 pMF65A .............................................................................. 71 pMF66A .............................................................................. 71 pMF67 ................................................................................ 71 Chapter 3 ........................................................................................ 71 Tyrosine Phenol-Lyase Assay .......................................................... 71 PCR Amplification of tpl. ............................................................... 72 Construction of Plasmid pMF43A ...................................................... 73 Culture Conditions for Converting D-Glucose into L-DOPA ........................ 74 Culture Conditions for Converting Catechol into L-DOPA .......................... 75 HPLC Analysis of Reaction Mixtures .................................................. 75 Determination of KM Values for Tyrosine Phenol-Lyase ............................ 75 References ...................................................................................... 77 vi LIST OF TABLES Table 1 - Plasmids Constructed for DHS Accumulation Experiments ........................ 22 vii LIST OF FIGURES Figure 1 - The Common Pathway of Aromatic Amino Acid Biosynthesis .................... 2 Figure 2 - Reactions Catalyzed by Transketolase (A) and Transaldolase (B) ................. 5 Figure 3 - Biocatalytic Synthesis of Quinic Acid from D—Glucose ............................. 9 Figure 4 - Biocatalytic Synthesis of Catechol from D-Glucose ................................ 10 Figure 5 - Industrial Catechol Manufacture ...................................................... 11 Figure 6 - Biocatalytic Synthesis of Adipic Acid from D-Glucose ............................ 12 Figure 7 - Industrial Adipic Acid Manufacture ................................................... 13 Figure 8 - Transaldolase Involves the Transfer of a Dihydroxyacetone Unit ................ 18 Figure 9 - Mechanism of Transaldolase .......................................................... 18 Figure 10 - Preparation of Plasmid pMF52A .................................................... 21 Figure 11 - Cloning the 1.25 kb aroF Fragment: Preparation of pMF58A .................. 24 Figure 12 - Cloning the 2.2 kb tktA Fragment: Preparation of pMF51A .................... 25 Figure 13 - Preparation of Plasmid pMF61A .................................................... 26 Figure 14 - Preparation of Plasmids pMF57A and pMF6OA .................................. 27 Figure 15 - Preparation of Plasmids pMF63A and pMF65A .................................. 28 Figure 16 - DHS Production in Different E. coli aroE Strains ................................. 29 Figure 17 - Preparation of Plasmid pMF66A .................................................... 31 Figure 18 - Effect of Plasmid pMF66A on DHS Production .................................. 32 Figure 19 - Preparation of Plasmid pMF67 ...................................................... 34 Figure 20 - Effect of Plasmid pMF67 on DHS Production .................................... 35 Figure 21 - Effect of Plasmid pKL1.87A on DHS Production ................................ 36 Figure 22 - Proposed Synthesis of L-DOPA from D-Glucose ................................. 42 Figure 23 - Reactions Catalde by Tyrosine Phenol-Lyase ................................... 42 Figure 24 - Proposed Mechanism of Tyrosine Phenol-Lyase .................................. 44 Figure 25 - Preparation of Plasmid pMF38A .................................................... 45 Figure 26 - Nucleotide Sequence of the C itrobacter freundii tpl Gene ........................ 47 Figure 27 - Preparation of Plasmid pMF42A .................................................... 48 Figure 28 - Preparation of Plasmid pMF43A .................................................... 49 viii Ap bp Cm DAH DAHP DHQ DHS E4P EPSP HPLC IPTG kb NMR PEP PCR phe Tc TSP tyr LIST OF ABBREVIATIONS ampicillin base pair chloramphenicol 3deoxy-D-arabino—hepnilosonic acid 3-deoxy~D-arabin0-heptulosonic acid 7-phosphate 3-dehydroquinate 3-dehydroshikimate D—erythrose 4—phosphate 5-enolpyruvoylshikimate 3-phosphate hour high pressure liquid chromatography isopropyl B-D-thiogalactopyranoside kilobase minute nuclear magnetic resonance spectroscopy phosphoenolpyruvate polymerase chain reaction L—phenylalanine tetracycline L-tryptophan sodium 3-(trimethylsilyl)propionate-2,2,3,3-d4 L-tyrosine CHAPTER 1 INTRODUCTION The Common Pathway The common pathway of aromatic amino acid biosynthesis is found in plants, bacteria, and fungi and is named for the fact that chorismic acid is the common branch point from which the aromatic amino acids and related secondary metabolites are synthesized.1 The intermediates of the seven enzymatic reactions that convert phosphoenolpyruvate (PEP) and D-erythrose 4-phosphate (E4P) into chorismic acid (Figure 1) were completely identified by the early 19605 from studies of bacterial auxotrophs of Escherichia coli and Klebsiella aerogenes. The first committed step of aromatic amino acid biosynthesis involves the condensation of PEP and E4P to form 3-deoxy-D-arabino-heptulosonic acid 7- phosphate (DAHP), catalyzed by DAHP synthase.2 Three isozymes of DAHP synthase exist in E. coli, each of which is sensitive to feedback inhibition by one of the three aromatic amino acids. The genes aroF, aroG, or aroH encode for DAHP synthase (tyr), DAHP synthase (phe), and DAHP synthase (trp), respectively.l DAHP is converted into 3-dehydroquinate (DHQ) via DHQ synthase in a reaction that requires NAD. DHQ dehydratase catalyzes the formation of 3-dehydroshikimate (DHS) which contains the first double bond of the aromatic ring. The reduction of DHS by shikimate dehydrogenase in the presence of NADPH affords shikimate. Shikimate kinase produces shikimate 3- phosphate from shikimate and ATP. The reversible condensation of shikimate 3—phosphate and PEP to form 5-enolpyruvoylshikimate 3-phosphate (EPSP) is catalyzed by EPSP synthase. Chorismate synthase catalyzes the elimination of inorganic phosphate from 0P03H2 OH OH , CO2H A HO“ COZH HOI. C(32H O I' —.> PEP (OO‘HB PiOQ‘OH ——> _._ OH H203PO OH ; ; OH D-glucose H8 0 DAHP DHQ E4P COZH COZH COZH :33 £1 ° 6) 7% O > o _ OH /\ Ho" ; OH H203PO" ; OH H20 0” NADPH NADP 0” ATP ADP 0“ DHS shikimate shikimate 3-phosphate COZH COZH F (5‘0 IL ©OIL —> L-phenylalanine ' 6: "—"’ L-tyrosine I ; HHOPO' 002H COZH L-t to han PEP Pi F»: W p EPSP Chorismate (A) DAHP synthase (aroF aroG aroH); (B) DHQ synthase (aroB); (C) DHQ dehydratase (aroD); (D) shikimate dehydrogenase (aroE); (E) shikimate kinase (aroK aroL); (F) EPSP synthase (aroA); (G) Chorismate synthase (aroC). Figure 1 - The Common Pathway of Aromatic Amino Acid Biosynthesis EPSP to give Chorismate, which contains the second double bond of the aromatic ring. Three terminal pathways lead from Chorismate to L-phenylalanine, L-tryptophan, and L- tyrosine. Chorismate also serves as the precursor for other essential aromatic metabolites such as ubiquinone, enterochelin, and folic acid, which are involved in electron transport, iron uptake, and coenzyme biosynthesis, respectively.1 5 I . I I . w' I E I' Microorganisms possess a variety of biosynthetic enzymes, in addition to those of the common pathway, which can be utilized for the synthesis of aromatics from D-glucose. The advantage of using D-glucose is that it is inexpensive, nontoxic, and derived from renewable resources such as corn starch.3 This contrasts with traditional chemical synthesis of aromatics, which relies on petroleum-derived resources and typically involves carcinogenic starting materials such as benzene. Using Escherichia coli as the microbial host for aromatic production is derived from the fact that a tremendous amount of information has been accumulated on the biochemistry and molecular biology of this organism.1 It has become the archetype for extensive studies on protein expression, genetic engineering, and biocatalysis. One of the essential goals in the development of a microbial catalyst for aromatic synthesis is to direct a high percentage of the carbon consumed by the microbe into the common pathway of aromatic amino acid biosynthesis. Typically this has been accomplished by increasing the specific activity of the first enzyme of the pathway, DAHP synthase. This can be done by localizing a gene that encodes for one of the three isozymes of DAHP synthase on an extrachromosomal plasmid4 or by eliminating transcriptional control of one of the DAHP synthase genes.5 The result is an increased intracellular concentration of DAHP synthase verified by elevated enzyme activity. A complementary strategy is to introduce a mutation in one of the DAHP synthase genes resulting in a protein that is resistant to feedback inhibition by the aromatic amino acids.6 Increasing DAHP synthase activity does increase the percentage of carbon (D- glucose equivalents) committed to aromatic amino acid biosynthesis. This has been measured using E. coli aroB strains that possess an inactive DHQ synthase.7 Because these strains are incapable of converting DAHP into DHQ, the DAHP that accumulates intracellularly is exported into the culture medium, along with the dephosphorylated product 3-deoxy-D—arabino-heptulosonic acid (DAH).4 The amount of DAH(P) present in the culture supernatant can be directly quantified8 to provide a measure of the carbon flow directed into the common pathway. Experiments with E. coli aroB strains that contain a feedback-resistant copy of aroG-encoded DAHP synthase localized on an extrachromosomal plasmid demonstrated that increased DAHP synthase activity and 4 improved DAHP production are observed.7b However, a point was reached where further increases in DAHP synthase activity had no impact on the delivery of carbon flow into the common pathway. Increasing In Vivo Availability of Q-Egthrose 4-Phosphate At this point it becomes necessary to focus on the in vivo availability of the substrates of DAHP synthase: D-erythrose 4-phosphate (E4P) and phosphoenolpyruvate (PEP). If the intracellular concentration of either E4P or PEP is limiting, DAHP formation will be impeded, even in the presence of overexpressed DAHP synthase. Initial attention centered on E4P as the limiting substrate, for several reasons. D-Erythrose 4-phosphate has never been convincingly detected in a living system.9 The absence of intracellular E4P accumulation could be attributed to its rapid utilization subsequent to its formation inside the cell.9b’c Considering that this aldose phosphate is rather unstable and can easily undergo dimerization,10 it may be that biological systems have evolved to maintain very low steady-state concentrations of E4P. Nevertheless, when the demand for aromatic amino acid biosynthesis is high, limiting E4P levels may hinder the rate of formation of DAHP. Investigation into increasing the in vivo levels of E4P led to the enzymes of the pentose phosphate pathway. The pentose phosphate pathway forms a bridge between glycolysis and several biosynthetic pathways. It consists of an oxidative and a non-oxidative branch. The oxidative branch converts D-glucose 6-phosphate into D-ribose 5-phosphate and carbon dioxide, with concomitant production of NADPH. The non-oxidative branch converts D- fructose 6-phosphate (a D-glucose equivalent) into a variety of C—3 through C—7 monophosphate sugars. Two enzymes of the nonoxidative branch,ll transketolase and transaldolase, catalyze reactions that lead to the formation of D-ribose 5-phosphate, D- sedoheptulose 7-phosphate, and D-erythrose 4-phosphate (Figure 2). These sugars are required for the biosynthesis of nucleotides, lipopolysaccharides, and aromatic amino _ + - H ‘— H + - 6H OH OH 6H 5H 0 6H 0 D-fructose D-glyceraldehyde D-xylulose 6-phosphate 3-phosphate 5-phosphate H203PO OH 0 H203PO OH 0 A Hzoapo 0H H203PO OH 9H CH _ + - _ H H + - - 5H OH OH 6H 6H 3H 0 6H 6H 0 D-fructose D-ribose D-sedoheptulose 6-phosphate 5-phosphate 7-phosphate H203PO OH C_)H OH H203PO o B Hzoapo OH HZOSPO OH 0 _ - - + - H ‘ H + _ 6H 6H 0 6H 5H 0 6H OH OH D-sedoheptulose D-giyceraldehyde D-erythrose D-fructose 7-phosphate 3-phosphate 4-phosphate 6—phosphate (E4P) Figure 2 - Reactions Catalyzed by Transketolase (A) and Transaldolase (B) acids, respectively. The only source of D-erythrose 4-phosphate in bacterial biosynthesis is derived from reactions catalyzed by transketolase and transaldolase. Although both transketolase and transaldolase are responsible for E4P formation, efforts were focused on increasing transketolase activity for the following reasons. Transketolase catalyzes the reversible transfer of a two carbon ketol group between various aldose acceptors. The equilibrium constant for transketolase-catalyzed conversion of D- fructose 6-phosphate into E4P favors D-fructose 6-phosphate.12 The unwieldy chemical characteristics of free E4P might be avoided by maintaining an intracellular supply of D- fructose 6-phosphate that serves as a precursor to E4P.10a Furthermore, E. coli mutants with extremely low levels of transketolase are unable to grow in the absence of aromatic amino acid supplementation, presumably due to limitations in E4P availability. 13 A gene encoding transketolase was isolated from an E. coli genomic library by complementation of a mutant possessing reduced transketolase levels.7b The insert was 6 subcloned to give a 5 kb fragment that contained the tkt gene. Transketolase was purified to homogeneity from both wild-type E. coli and an overexpressing strain containing the 5 kb insert localized on an extrachromosomal plasmid. The protein is a homodimer with a subunit size of 72 kDa,7b which agrees closely with nucleotide sequence information.14 Recently a second gene for transketolase (designated tktB) was identified and sequenced. 15 However this isozyme of transketolase accounts for a small percentage of wild-type transketolase activity. 15 The isolation of the transketolase gene (now designated tktA) and successful overexpression in E. coli led to experiments that measured the impact of transketolase on the delivery of D-glucose equivalents into the common pathway of aromatic amino acid biosynthesis. This was illustrated by comparison of DAHP formation in two strains that possessed comparable levels of DAHP synthase, AB2847aroB/pRW5 and AB2847aroB/pRW5tkt.7b The only difference between these two strains is that AB2847aroB/pRW5tkt carries an extrachromosomal copy of tktA resulting in elevated levels of transketolase activity. Under the same culturing conditions AB2847aroB/pRW5tkt synthesized twice as much DAHP relative to AB2847aroB/pRW5, which confirmed that increasing transketolase activity leads to higher in vivo availability of E4P and increased carbon flow into the common pathway. Although it has been demonstrated that amplification of transketolase can increase the yield of DAHP, the effect of increasing the specific activity of transaldolase has not been explored. One of the goals of this thesis is to determine the impact of transaldolase on directing carbon flow into the common pathway. Increasing In Vivg Availability 9f Phosphoenglpyruvate In addition to D-erythrose 4-phosphate, phosphoenolpyruvate (PEP) has been proposed to limit the percentage of D-glucose committed to the common pathway.16 Stoichiometric analysis of the pathways for DAHP production in E. coli suggests that the 7 theoretical yield of DAHP production from D-glucose is limited by PEP due to the phosphotransferase system (PT S) for sugar uptake. During glucose transport PEP serves as a phosphate donor and is converted to pyruvate. In wild-type E. coli, this pyruvate is not likely to be recycled back to PEP because of the high energy cost. As a result, carbon flow directed through pyruvate goes on to produce organic acids, carbon dioxide, or cell mass. The problem of limited PEP availability has been addressed with several strategies. One plan to alleviate the burden of limited PEP supply is to increase the specific activity of PEP synthase, which catalyzes the conversion of pyruvate into PEP.15 Theoretical analyses predict that the maximum yield of DAHP produced from D-glucose can be increased twofold (from 43 mol % to 86 mol %) if pyruvate is recycled back to PEP. 16 The overexpression of PEP synthase was accomplished by localization of the pps gene on an extrachromosomal plasmid. When DAHP synthase, transketolase, and PEP synthase were overexpressed in an E. coli aroB strain in the presence of D-glucose, the final concentration and yield of DAHP approached the theoretical maximum. ‘6 Notably, the effect of PEP synthase was not observed without the overproduction of transketolase, suggesting that E4P is the first limiting metabolite. However, transketolase alone cannot increase the yield of DAHP from glucose to the theoretical maximum value because of the stoichiometric limitation of PEP. Other strategies to compensate for the limitations in PEP availability include elimination of enzymatic pathways that compete for intracellular PEP. Studies have been done with mutants lacking PEP carboxylase activity163v17 (converting PEP to oxaloacetate) and with mutants lacking pyruvate kinase activity18 (converting PEP to pyruvate). These approaches have been met with limited success. Changing the carbon source from D- glucose to D-xylose, a sugar whose uptake does not involve the phosphotransferase system, has been explored. 16" The theoretical yield of DAHP from D—xylose is not limited by PEP and can reach 71 mol % without pyruvate recycling.16b Rate—Limiting Enzymes of thg Cgmmon Pathway An increased surge of carbon flow into the common pathway creates a metabolic situation where individual common pathway enzymes become rate-limiting. These rate- limiting enzymes cannot convert substrate to product at a sufficient rate to avoid accumulation of substrate inside the cell. Export of the accumulating substrate into the culture supernatant occurs, resulting in lower percent conversions and reduced purity of desired aromatic products. Using 1H NMR analysis, common pathway substrates and related metabolites were identified from the culture supernatant of an E. coli auxotroph that had been designed to overexpress DAHP synthase and transketolase.19 The genes for common pathway enzymes whose substrates were present in the culture supernatant were introduced on extrachromosomal plasmids into the E. coli auxotroph. When a metabolite no longer accumulated as a result of elevated levels of a common pathway enzyme, the rate- limiting character of that enzyme was said to be removed. Progress toward decreasing the number of accumulating common pathway enzyme substrates and increased synthesis of aromatics has been achieved. In this fashion, 3-dehydroquinate (DHQ) synthase, shikimate kinase, 5-enolpyruvoylshikimate 3-phosphate (EPSP) synthase, and Chorismate synthase were identified as rate-limiting enzymes.19 Other methods that have been used to remove the rate-limiting character of common pathway enzymes include inactivation of the TyrR protein in E. coli. The TyrR protein, in combination with tyrosine or tryptophan, inhibits the transcription of aroL, which encodes shikimate kinase. 1330 The introduction of a mutant allele of tyrR into an E. coli strain that produces phenylalanine resulted in an increased specific activity of shikimate kinase, a decrease in the accumulation of shikimate, and a higher yield of aromatic end products.21 Recently, chromosomal insertion of a synthetic multi-gene cassette carrying aroB (encoding DHQ synthase), aroA (encoding EPSP synthase), and aroC (encoding Chorismate synthase) into an aromatic-synthesizing E. coli strain was accomplished in an effort to 9 avoid potential problems associated with plasmid instability and unnecessary overexpression of plasmid-encoded, common pathway enzymes.21 Assembly of a Heterolgggus Migrobial Biocatalyst Intermediates of the common pathway of aromatic amino acid biosynthesis can be transformed into a variety of aromatic-derived chemicals.22 Often this requires incorporation of enzyme activities not possessed by E. coli. Examples of this process are illustrated in the biocatalytic production of quinic acid,23 catechol,24 and adipic acid25 from D-glucose. Quinic acid is an important chiral starting material used widely in multistep chemical synthesis whose isolation from plant sources is expensive. The biocatalytic synthesis of quinic acid (Figure 3) has been reported using inexpensive, abundant D- glucose as starting material.23 OH DHQ quinate HO H ”OH " 002 synthase HO" 002” dehydrogenase “OI. C02H o _, o I _OH ;OH O_OH HO'_OH HO OH H203PO OH 5H 6H D-glucose DAHP DHQ quinic acid DHQ dehydratase COZH L-phenylalanine ——-> L-tyrosine O L-tryptophan - OH 5H DHS Figure 3 - Biocatalytic Synthesis of Quinic Acid from D-Glucose Quinic acid synthesis in E. coli requires the conversion of D-glucose into 3- dehydroquinate (DHQ) via the common pathway. DHQ can be converted into quinic acid by quinic acid dehydrogenase,26 an enzyme not found in E. coli. The quinic acid 10 dehydrogenase gene (qad) was isolated from the microorganism Klebsiella pneumoniae, which shares a close evolutionary relationship with E. coli.27 This relationship is evidenced by the fact that K. pneumoniae genes have been successfully expressed from their native promoters in E. coli.28 Although K. pneumoniae utilizes quinic acid dehydrogenase for the catabolism of quinic acid to DHQ, the reduction of DHQ to quinic acid is the thermodynamically preferred direction of reaction.26 The overexpression of quinate dehydrogenase, DAHP synthase, DHQ synthase, and transketolase on extrachromosomal plasmids was achieved in an E. coli aroD strain incapable of converting DHQ into DHS. The result was the creation of a heterologous microbe that converts D- glucose into quinic acid in 31 mol % yield.23 OH COZH COZH 0 "OH dehggitase decafbgéylase __> i - OH 0 _ OH OH OH Ho 6H 6H OH on D-glucose DHS PCA catechol shikimate dehydrogenase COZH , L-phenylalanine HO“ ' OH _—> :IyroIonIian 5H WP P shikimate Figure 4 - Biocatalytic Synthesis of Catechol from D-Glucose The biocatalytic synthesis of catechol from D-glucose also involved the recruitment of genes from K. pneumoniae.24 The microbial biocatalyst was created by utilizing enzymes from the common pathway of aromatic amino acid biosynthesis and from hydroaromatic catabolism29 (Figure 4). D-Glucose is converted to 3-dehydroshikimate 11 (DHS) through the common pathway. Catechol is formed from DHS by the action of two enzymes found in K. pneumoniae. DHS dehydratase3O (encoded by aroZ) catalyzes the formation of protocatechuic acid (PCA) from DHS, and PCA decarboxylase31 (encoded by aroY) converts PCA into catechol. Plasmid-based overexpression of DAHP synthase, DHQ synthase, and transketolase in an E. coli aroE strain that cannot convert DHS into shikimic acid results in the production of DHS from D-glucose.32 Isolation of the K. pneumoniae genes aroZ and aroY was followed by amplification of DHS dehydratase and PCA decarboxylase in the DHS-producing construct. This heterologous biocatalyst synthesized catechol in 33 mol % yield from D-glucose.24 OH OH A B C o —» 4 <5 —. + OH benzene cumene 0 phenol OH OH A catechol hydroquinone acetone (A) propylene, solid H3PO4 catalyst, ZOO-260°C, 400-600 psi. (B) 02, 80-1300C then S02, 60-1000C. (C) 70% H202, EDTA, Fe+2 or C0”, 70-800C. Figure 5 - Industrial Catechol Manufacture This environmentally compatible synthesis contrasts sharply with the dominant method33 of industrial catechol manufacture, which starts from petroleum-derived benzene (Figure 5). Benzene undergoes Friedel-Crafts alkylation to give cumene, which is subsequently converted into acetone and phenol in a Hock-type, air oxidation. Phenol is then oxidized with 70% hydrogen peroxide either in the presence of transition metal catalysts or in formic acid solution where the actual oxidant is performic acid. The resulting mixture of catechol and hydroquinone is separated into its pure components through successive distillations. The biocatalytic synthesis of catechol from D-glucose 12 simultaneously avoids the use of carcinogenic, petroleum-based starting materials and highly corrosive hydrogen peroxide. Catechol can be transformed into cis,cis-muconic acid with catechol 1,2- dioxygenase encoded by cat/1.29a This gene is not possessed by E. coli, but has been isolated from Acinetobacter calcoaceticus.34 Expression of catA in a microbe designed to produce catechol resulted in a new heterologous biocatalyst that converted D-glucose into cis,cis-muconic acid (Figure 6) in 30 mol % yield.25 Catalytic hydrogenation of the cis,cis-muconic acid from the culture supernatant at 50 psi for 3 h at room temperature afforded adipic acid in 90% yield.25 catechol H020 HOZC (051:: ©Lm‘1,—2—dioxygenase> I H2. Pt ——> I Ho OH 0021-1 COzH D-glucose cateHchol cis,cis-muconic acid adipic acid Figure 6 - Biocatalytic Synthesis of Adipic Acid from D-Glucose As with the synthesis of catechol, industrial synthesis of adipic acid33a'bv35 (Figure 7) poses several threats to the environment. Benzene is hydrogenated to produce cyclohexane, which is then oxidized in air to give a mixture of cyclohexanol and cyclohexanone. Nitric acid oxidation affords adipic acid. The byproduct of this last reaction, nitrous oxide}6 is involved in depletion of the ozone layer and the greenhouse effect.37 Some 10% of the annual increase in atmospheric nitrous oxide levels is related to adipic acid manufacture.36 The industrial process requires forcing reaction conditions, with temperatures as high as 250°C and pressures reaching 800 psi.33’i"‘”35 However, biocatalytic synthesis of adipic acid involves mild temperatures and pressures, avoids 13 OH i) cyclohexanol C H02C A B Q _+ O _. . _. o benzene cyclohexane COQH adipic acid cyclohexanone (A) Ni-Ale3, 370-800 psi, ISO-250°C. (B) Co, Oz, 120-140 psi, ISO-160°C. (C) Cu, NH4VO3, 60% HNO3, 60-800C. Figure 7 - Industrial Adipic Acid Manufacture generation of nitrous oxide, and starts with nontoxic D-glucose derived from abundant, renewable resources. Design of microbial catalysts often involves recruitment of enzyme activities not found in E. coli. This has been illustrated in the biocatalytic syntheses of quinic acid, catechol, and adipic acid. A similar strategy was proposed for creating a biocatalyst capable of converting D-glucose into L-3,4-dihydroxyphenylalanine (L-DOPA), a compound that is primarily used for treatment of Parkinson's disease.38 This required the activity of tyrosine phenol-lyase,39 an enzyme not found in E. coli, which converts catechol, pyruvate, and ammonia into L-DOPA. In addition to the transaldolase study mentioned previously, this thesis will describe the isolation of the tpl gene encoding tyrosine phenol-lyase from Citrobacter freundii and the expression of tpl in a catechol- producing biocatalyst. 10. ll. 12. 13. 14. 15. 16. 14 References (a) Pittard, A. J. In Escherichia coli and Salmonella typhimurium; Neidhardt, F. C., Ed.; American Society for Microbiology: Washington, DC, 1987; Vol. 1, pp 368-394. (b) Haslam, E. Shikimic Acid Metabolism and Metabolites; Wiley: New York, 1993. (c) Herrmann, K. M. In Amino Acids: Biosynthesis and Genetic Regulation; Herrmann, K. M., Somerville, R. L., Eds.; Addison-Wesley: Reading, 1983; p 301. Srinivasan, P. R.; Sprinson, D. B. J. Biol. Chem. 1959, 234, 716. Lee, H. In Industrial Uses of Agricultural Materials; Commodity Economics Division, Economic Research service, U. S. Department of Agriculture: 1993 (June); IUS-l. Frost, J. W.; Knowles, J. R. Biochemistry 1984, 23, 4465. (a) Cobbett, C. S.; Morrison, S.; Pittard, J. J. Bacterial. 1984, 15 7, 303. (b) Garner, C. C.; Herrmann, K. M. J. Biol. Chem. 1985, 260, 3820. (a) Ogino, T.; Garner, C.; Markley, J. L.; Herrmann, K. M. Proc. Natl. Acad. Sci. U.S.A. 1982, 79, 5828. (b) Weaver, L. M.; Herrmann, K. M. J. Bacteriol. 1990, 172, 6581. (a) Draths, K. M.; Frost, J. W. J. Am. Chem. Soc. 1990, 112, 1657. (b) Draths, K. M.; Pompliano, D. L.; Conley, D. L.; Frost, J. W.; Berry, A.; Disbrow, G. L.; Staversky, R. J.; Lievense, J. C. J. Am. Chem. Soc. 1992, 114, 3956. Gollub, E.; Zalkin, H.; Sprinson, D. B. Methods Enzymal. 1971, 17A, 349. (a) Paoletti, F.; Williams, J. F.; Horecker, B. L. Anal. Biochem. 1979, 95, 250. (b) Williams, J. F.; Blackmore, P. F.; Duke, C. C.; MacLeod, J. F. Int. J. Biochem. 1980, 12, 339. (c) Williams, J. F.; Arora, K. K.; Longenecker, J. P. Int. J. Biochem. 1987, 19, 749. (a) Blackmore, P. F.; Williams, J. F.; MacLeod, J. F. FEBS Lett. 1976, 64, 222. (b) Dike, C. C.; MacLeod, J. F.; Williams, J. F. Carbohydr. Res. 1981, 95, l. Horecker, B. L. In Comprehensive Biochemistry; Florkin, M., Stotz, E. H., Eds; Elsevier: Amsterdam, 1964; Vol. 15. Racker, E. In The Enzymes; Boyer, P. D., Ed.; Academic Press: New York, 1961; Vol. V, Chapter 24A. Josephson, B. L.; Fraenkel, D. G. J. Bacteriol. 1974, 118, 1082. Sprenger, G. A. Biochim. Biophys. Acta 1993, 1216, 307. Iida, A.; Teshiba, S.; Mizobuchi, K. J. Bacterial. 1993, 175, 5375. (a) Patnaik, R.; Liao, J. C. Appl. Environ. Microbial. 1994, 60, 3903. (b) Patnaik, R.; Spitzer, R. G.; Liao, J. C. Biotechnol. Bioeng. 1995, 46, 361. 17. 18. 19. 20. 21. 22. 23. 24. 25. 26. 27. 28. 29. 30. 31. 32. 33. 15 Miller, J. E.; Backman, K. C.; O'Connor, J. M.; Hatch, T. R. J. Ind. Microbial. 1987, 2, 143. Gubler, M.; Jetten, M.; Lee, S. H.; Sinskey, A. J. Appl. Environ. Microbial. 1994, 60, 2494. (a) Dell, K. A.; Frost, J. W. J. Am. Chem. Soc. 1993, 115, 11581. (b) Dell, K. A. PhD. Thesis, Purdue University, December, 1993. Ely, B.; Pittard, J. J. Bacteriol. 1979, I38, 933. Snell, K. D.; Draths, K. M.; Frost, J. W. J. Am. Chem. Soc, in press. Frost, J. W.; Draths, K. M. Chem. Brit. 1995, 31, 206. Draths, K. M.; Ward, T. L.; Frost, J. W. J. Am. Chem. Soc. 1992, 114, 9725. Draths, K. M.; Frost, J. W. J. Am. Chem. Soc. 1995, 117, 2395. (a) Draths, K. M.; Frost, J. W. J. Am. Chem. Soc. 1994, 116, 399. (b) Draths, K. M.; Frost, J. W. In ACS Synpasium Series 577: Benign by Design; Anastas, P. T., Farris, C. A., Eds.; American Chemical Society: Washington, DC, 1994; p 32. Davis, B. D.; Gilvarg, C.; Mitsuhashi, S. Methods Enzymal. 1955, 2, 307. Jensen, R. A. Mol. Biol. Eval. 1985, 2, 92. (a) Peng, H.-L.; Wang, P.-Y.; Wu, C.-M.; Hwang, D.-C.; Chang, H.-Y. Gene 1992, 117, 125. (b) Feldmann, S. D.; Sahm, H.; Sprenger, G. A. Mal. Gen. Genet. 1992, 234, 201. (c) Hama, H.; Wilson, T. H. J. Biol. Chem. 1992, 267, 18371. (a) Ornston, L. N .; Neidle, E. L. In The Biology of Acinetobacter; Towner, K. J ., Bergogne-Bérézin, E., Fewson, C. A., Eds.; Plenum: New York, 1991; pp 201- 237. (b) Giles, N. H.; Case, M. E.; Baum, 1.; Geever, R.; Huiet, L.; Patel, V.; Tyler, B. Microbial. Rev. 1985, 49, 338. (a) Gross, S. R. J. Biol. Chem. 1958, 233, 1146. (b) Tatum, E. L.; Gross, S. R. J. Biol. Chem. 1956, 219, 797. (c) Strmman, P.; Reinert, W. R.; Giles, N. H. J. Biol. Chem. 1978, 253, 4593. (d) Schweizer, M.; Case, M. E.; Dykstra, C. C.; Giles, N. H.; Kushner, S. R. Gene 1981, 14, 23. (a) Patel, J. C.; Grant, D. J. W. Antonie van Leeuwenhoek 1969, 35, 53. (b) Grant, D. J. W. Antonie van Leeuwenhoek 1970, 36, 161. Draths, K. M.; Frost, J. W. J. Am. Chem. Soc. 1990, 112, 9630. (a) Framck, H.-G; Stadelhofer, J. W. Industrial Aromatic Chemistry; Springer- Verlag: New York, 1988. (b) Szmant, H. H. Organic Building Blocks of the Chemical Industry; Wiley: New York, 1989. (c) Varagnat, J. In Kirk-0thmer Encyclopedia of Chemical Technology, 3rd ed.; Grayson, M, Ed.; Wiley: New York, 1981; Vol. 13. PP 39-69. 34. 35. 36. 37. 38. 39. 16 Neidle, E. L.; Omston, L. N. J. Bacterial. 1986, 168, 815. Davis, D. D.; Kemp, D. R. In Kirk-0thmer Encyclopedia of Chemical Technology, 4th ed.; Kroschwitz, J. 1., Howe-Grant, M., Eds.; Wiley: New York, 1991; Vol. 1, pp 466-493. Thiemens, M. H.; Trogler, W. C. Science 1991, 251, 932. Dickinson, R. E.; Cicerone, R. J. Nature 1986, 319, 109. Cotzias, G. C.; Papavasiliou, P. S.; Gellene, R. New Eng. J. Med. 1969, 280, 337. (a) Yamada, H.; Kumagai, H.; Matsui, H.; Ohgishi, H.; Ogata, K. Biochem. Biophys. Res. Commun. 1968, 33, 10. (b) Kumagai, H.; Yamada, H.; Matsui, H.; Ohkishi, H.; Ogata, K. J. Biol. Chem. 1970, 245, 1767. (c) Kumagai, H.; Yamada, H.; Matsui, H.; Ohkishi, H.; Ogata, K. J. Biol. Chem. 1970, 245, 1773. (d) Kumagai, H.; Kashima, N .; Torii, H.; Yamada, H.; Enei, H.; Okumura, S. Agr. Biol. Chem. 1972, 36, 472. (e) Carman, G. M.; Levin, R. E. Appl. Environ. Microbiol. 1977, 33, 192. CHAPTER 2 IMPACT OF TRANSALDOLASE ON THE BIOCATALYTIC PRODUCTION OF AROMATICS Background Biocatalytic synthesis of aromatics requires that a large percentage of the carbon source consumed by the microbe be committed to the common pathway of aromatic amino acid biosynthesis. This can be accomplished by increasing the specific activity of DAHP synthase, the first enzyme of the common pathway.1 However, a point has been reached where the flow of carbon into the common pathway is not limited by the catalytic activity of DAHP synthase, but rather by the in vivo availability of the substrates D-erythrose 4- phosphate (E4P) and phosphoenolpyruvate (PEP). Transketolase and transaldolase are the two enzymes of the pentose phosphate pathway that catalyze reactions leading to E4P (Figure 2, Chapter 1). Transketolase has been successfully overexpressed in E. coli and has been used to increase the delivery of carbon (D-glucose equivalents) into the common pathway.2 However, the effect of transaldolase on the biosynthesis of aromatics has not been explored. Transaldolase was first described by Horecker and Smymiotis3 in 1953. Present in animal, plant, and microbial cells, transaldolase is named for the fact that it catalyzes the transfer of aldol linkages from one sugar to another. Specifically, transaldolase transfers a three carbon dihydroxyacetone unit from a phosphorylated ketose donor to an aldose acceptor4 (Figure 8). The mechanism of the reaction43 involves the initial formation of a Schiff base intermediate between the carbonyl group of the ketose substrate and the 8- amino group of a lysine residue at the active site of transaldolase (Figure 9). The proton of 17 18 H203P0 0H 9H OH OH Hzoapo OH OH 9H 0H ' + H transaldolase; H + - H6 H5 0 H203PO o ‘ 3H 0 H203PO H6 0 D-sedoheptulose D-glyceraldehyde D-erythrose D-fructose 7-phosphate 3—phosphate 4-phosphate 6-phosphate Figure 8 - Transaldolase Involves the Transfer of a Dihydroxyacetone Unit H203P0 OH QH - u OH ”HEY H203P0 OH QH/ H203“) OH 9H HO (cl) ONH HO HO O H0 H0 ONH :N H20 \é IL \ N H O 0 H203PO OH HN \ HN' \ H20\i N kII H H Oil I O I l I M 2 OH 9H OH H203PO H6 0 Figure 9 - Mechanism of Transaldolase 19 the C-4 hydroxyl group is abstracted by the imidazole of a histidine residue, promoting release of the aldose product. The Schiff base carbanion can then react with a suitable aldose acceptor that produces a ketose product upon hydrolysis. The first step toward overexpression of transaldolase (or any enzyme) in E. coli is to isolate the gene that encodes for it. Once this has been achieved, the gene can be localized on an extrachromosomal plasmid vector.5 Because genes that are plasmid- encoded are present in multiple copies, they are often expressed at higher levels than chromosomal genes. This leads to increased enzyme activity for the corresponding gene products. Technical advances in molecular biology have greatly facilitated the use of plasmids. It is generally easier and faster to increase gene expression by introducing plasmids into the cell rather than performing manipulations directly on the bacterial chromosome. Overexpression of Traaaaldolase in E. coli Isolation of the gene encoding transketolase was facilitated with an E. coli auxotrophic mutant.5b This strain possessed extremely low levels of transketolase and therefore was unable to grow on pentoses such as D-ribose, D-xylose, and D-arabinose. Complementation of this mutant with an E. coli genomic library led to the isolation of the tktA gene. Unfortunately, this strategy could not be applied to isolation of the transaldolase gene, because no E. coli transaldolase mutants are known. Perhaps E. coli possesses more than one enzyme (or a transaldolase isozyme) that can catalyze the reactions of transaldolase. In this case, deletion of transaldolase activity in E. coli might not result in an easily identifiable phenotype. Two different regions of the E. coli chromosome have been identified which share homology with the transaldolase protein6 from Saccharomyces cerevisiae. One is located at 53 min and was identified as an open reading frame upstream of tktB, a gene encoding the minor isozyme of transketolase.7 The size of the open reading frame, designated as talA, 20 was not specified, and only 200 bp of the 3' end of the gene sequence was reported.7 The second putative tal locus is at 0 min, and was discovered as a pair of open reading frames during the sequencing of the 0-2.4 min region of the E. coli genome.8 Since the entire sequence of this region was available, it was amplified using the polymerase chain reaction (PCR). PCR is an in vitro method for amplifying DNA fragments.9 From the published sequence information, two oligonucleotide primers were designed to flank the locus at 0 min. The DNA that was amplified by PCR included the sequence that exhibited homology with yeast transaldolase6 (954 bp) plus an additional 322 bp outside this region. This was done to ensure that the entire gene and its native promoter would be amplified. Genomic DNA from wild-type E. coli strain RB79110 served as the template for PCR. The size of the major product from PCR was 1.3 kb, as expected. The next step was to determine if this 1.3 kb DNA fragment (designated talB) encoded for transaldolase. This required localization of the fragment onto a plasmid vector. For cloning purposes, the 5'-ends of each PCR primer had been designed to contain the recognition site for endonuclease NcaI. The PCR-amplified DNA fragment was digested with NcaI and cloned into the NcaI site of pBR325ll (Figure 10). The resulting plasmid, pMF52A, was transformed into wild-type E. coli strain DHSOI.12 Transaldolase activity was quantified from crude extracts of DHSOt and DHSOI/pMF52A. This is a coupled enzyme assay13 that measures the change in absorbance at 340 nm resulting from the disappearance of NADH. The specific activity of transaldolase from DH50I/pMF52A (8.6 units mg‘l) was approximately 6-fold higher than that of DH50t (1.5 units mg‘l), which verified that this 1.3 kb DNA fragment (talB) encodes for transaldolase. Escherichia coli genomic DNA t PCR Ncol Ncol 1.3 kb talB l1) Ncoi digest 21 Ncoi QR pBR325 6.0 kb { 1) Neal digest 2) CIAP treatment T4 Ligase Figure 10 - Preparation of Plasmid pMF52A 22 Table 1 - Plasmids Constructed for DHS Accumulation Experiments Plasmid Genes Encoded pMF61A aroF aroB pM F60A aroF aroB BIB pMF63A aroF aroB tktA pM F65A amF aroB taIB tktA Impact of Transaldolase on Aromatig Production Successful overexpression of transaldolase in E. coli led to experiments that measured the impact of transaldolase on delivery of carbon flow into the common pathway. Previous experiments with transketolase had been performed with E. coli aroB strains that possess an inactive DHQ synthase.2 The DAHP that accumulates intracellularly is exported into the culture supernatant and can be quantified using a colorimetric assay.14 The concentration of DAHP produced by the strain is a direct measure of the percentage of D- glucose committed to aromatic amino acid biosynthesis. However, results with transaldolase were attained using an E. coli araE mutant for the following reasons. An araE mutant blocked at shikimate dehydrogenase results in the accumulation of 3- dehydroshikimate (DHS) in the culture supernatant. Quantitation of DHS can be done with 1H NMR analysis by comparison of the integrated resonances corresponding to DHS to the integrated resonance of an internal standard. This has advantages over the colorimetric assay, in that it is more reliable and reproducible. In addition, 1H NMR facilitates detection and identification of any metabolite that may be accumulating in the culture medium. For this study four plasmids were constructed (Table 1). Each plasmid contains aroF (encoding DAHP synthase) and aroB (encoding DHQ synthase). Amplified expression of DAHP synthase increases the amount of carbon flow directed into the 23 common pathway.1 DHQ synthase is overexpressed to overcome the rate-limiting character of this enzyme.15 The genes for transketolase (tktA) and transaldolase (talB) were then added in all possible combinations. The aroF gene had previously been localized on a 2.4 kb EcaRI fragment.2a~C However, the aroF open reading frame is only 1068 bp.16 In order to eliminate the extraneous DNA sequence, the aroF gene including its native promoter was amplified by PCR to give a 1.25 kb EcoRI fragment (Figure 11). This shortened aroF gene was then cloned into pBR325 to form plasmid pMF58A (Figure 11). Furthermore, the gene for transketolase had been originally isolated on a 5 kb BamHI fragment,2 but the size of the open reading frame is only 1992 bp.17 The tktA gene was also amplified by PCR to eliminate nonessential DNA sequence, giving a 2.2 kb BamHI fragment containing tktA with its native promoter (Figure 12). This 2.2 kb tktA fragment was cloned into pBR325 to give pMF51A (Figure 12). The araB gene had been previously cloned as a 1.65 kb SphI fragment into plasmid pKD136.15a This 1.65 kb DNA fragment was not shortened further by PCR. Instead, the aroB gene was simply isolated from SphI digestion of pKD136. The assembly of the four plasmids listed in Table 1 was carried out as follows. Plasmid pMF61A encodes aroF and aroB and was prepared by cloning the 1.65 kb aroB fragment into plasmid pMF58A (Figure 13). Plasmid pMF60A containing aroF, aroB, and talB was constructed in two steps. The aroB fragment was first cloned into pMF52A to give pMF57A (Figure 14). Then the 1.25 kb aroF fragment was ligated into pMF57A, forming pMF60A (Figure 14). Plasmid pMF63A containing aroF, aroB, and tktA was constructed by cloning tktA into pMF61A. (Figure 15). Finally, talB was cloned into pMF63A to produce pMF65A (Figure 15). Each of these four plasmids (pMF60A, pMF61A, pMF63A, and pMF65A) was transformed separately into E. coli AB2834 araE.18 Because all plasmids were derived from pBR325,ll they contain the gene that confers resistance to the antibiotic ampicillin. 24 EcoRl EcoRl 2.5 kb aroFiragment aroF ‘— 2.5 kb—-> EcoRl EcoRi 1.25 kb aroF PCR product aroF *1 .25 kb" Plasmid pKD136 EooFtI PCR pBR325 ECORI EWRI 6.0 kb I 1.25 kb I K aroF EcoRI digest 1) EcoRl digest 2) CIAP treatment T4 Ligase Eoo aroF EooFII Figure 11 - Cloning the 1.25 kb aroF Fragment: Preparation of pMF58A 25 5.0 kb tktA fragment 2.2 kb tktA PCR product BamHI BamHI BamHl BamHl tktA ‘— 2.2 kb —’ Plasmid pKD136 i PCR BamHl BamHi 2.2 kb tktA l BamHl digest Figure 12 - Cloning the 2.2 kb tktA Fragment: BamHl O‘n pBR325 6.0 kb K 1) BamHl digest 2) CIAP treatment I T4 Ligase BamHl pMF51A KAI) 8.2 kb Preparation of pMF51A 26 pKD136 aroF EooR \Ap 15.15kb E RI aroEL- BamHl Go “I Sphl Sphl 1) Sphl digest . 2) CIAP treatment l Sphl dlgest Sphl Sphl 1.65 kb . ...........,.,,,,WW_ I aroB T4 Ligase Sphl Figure 13 - Preparation of Plasmid pMF61A 27 pKD136 \Ap 15.15kb aroB Sphl 1) Sphl digest . 2) CIAP treatment l Sphl dlgest Sphl Sphl .. 1.65 kb aroB T4 Ligase Sphl aroB Eco R NCO pMF57A Sphl aroF pMF58A 7.25 kb EooRi A p \ EooRl JD 1) EcoRi digest E RI d' t 2) CIAP treatment l 00 Iges ECORI E009] 1.25 kb aroF T4 Ligase Sphl Ncol EooFIl Figure 14 - Preparation of Plasmids pMF57A and pMF60A 28 1) BamHl (119953t BamHi digest 2) CIAP treatment BamHl BamHl 2.2 kb tktA T4 Ligase Ncol BamHl Ncol R pMF52A 7.3 kb taIB Ncol AP \-7 Sphl _ Ncol digest 1) Neal dlgest 2) CIAP treatment Ncol Ncol 1.3 kb I taIB T4 Ligase NCOI BamHl Sphl Figure 15 - Preparation of Plasmids pMF63A and pMF6SA 29 "10' % yield DHS Production DHS (mM) 7 1 4 0 54 ..................................................................................... 30 [:3 24h I 36h 36 “20 18 10 A82884/ A82834/ A82884/ A2834/ pMF61A pMF60A pMF63A pMF65A Figure 16 - DHS Production in Different E. coli araE Strains Addition of ampicillin to the culture medium provides selective pressure for the strain to maintain the plasmid. These strains were initially grown in rich medium, harvested, and then cultured in minimal medium that contained D-glucose. The culture supematants were analyzed by 1H NMR. The results from each strain are summarized in Figure 16. 1H NMR analysis indicated that overexpression of transaldolase (AB2834/pMF6OA compared to AB2834/pMF61A) improved the yield of DHS slightly, from 39 mol % to 44 mol %. Transketolase overexpression (AB2834/pMF63A) was more effective, producing DHS in 60 mol % yield. The strain that overexpressed both transketolase and transaldolase (AB2834/pMF65A) did not lead to further improvements in DHS production. The yield of DHS produced by AB2834/pMF65A was actually lower compared to AB2834/pMF63A, in which only transketolase was overexpressed. At this point it became necessary to determine if a metabolic burden was being imposed on AB2834/pMF65A resulting from 30 the overexpression of four proteins. The specific activity of DAHP synthase might be reduced to a point where the flow of carbon into the common pathway is limited. In order to compensate for the potential limitations of DAHP synthase activity, aroF was localized on a second plasmid to increase expression levels further. To ensure plasmid compatibility and plasmid maintenance, two vectors must have compatible replication origins and encode resistance to different antibiotics. Plasmid pSUl8'9 was chosen by virtue of its p15A replicon that is compatible with the pMBl replicon of pBR325-derived plasmids. The resistance to chloramphenicol encoded by pSU18 distinguishes it from plasmids pMF60A, pMF61A, pMF63A, and pMF65A, which encode resistance to ampicillin. The stability of two-plasmid constructs was maintained in culture medium containing both antibiotics. The aroF gene was cloned into pSU18, generating pMF66A (Figure 17). Transformation of each of the strains listed in Figure 16 with pMF66A was followed by 1H NMR analysis to determine the impact on DHS production. The specific activity of DAHP synthase was also measured20 from crude extracts of all eight strains. The results are summarized in Figure 18. The addition of aroF-encoding pMF66A increased the specific activity of DAHP synthase at least tenfold. However, this increase in enzyme activity did not significantly improve the DHS production, since the maximum yield of DHS was still 60 mol %. Furthermore, transaldolase and transketolase seemed to have little or no effect on improving the carbon flow into the common pathway in the presence of high levels of DAHP synthase. This suggested that the in vivo availability of D-erythrose 4-phosphate (E4P) was no longer rate-limiting under these conditions. Thus it became necessary to focus on increasing the intracellular supply of phosphoenolpyruvate (PEP). Techniques have been developed that increase the PEP available for aromatic biosynthesis. It has been demonstrated that overexpression of PEP synthase increases the production of DAHP from D-glucose.21 The PEP synthase effect is not observed without amplified expression of transketolase, which suggests that E4P is the first limiting 31 EcoFll Pstl Hindlll aroF EooRl 1) EcoFll digest I _ t 2) CIAP treatment £5009 dlges EooRl EooRI l 1.25 kb I I I aroF T4 Ligase EcoRl Pstl Hindlll pMF66A Cm 3.55 kb Figure 17 - Preparation of Plasmid pMF66A 32 mol % yield DHS Production DHS (mM) 7 1 4 0 54 I3 24h I 36h 36‘ DAHP Synthase Specific Activity (units mg'1) 5 1 2 3 4 5 6 7 8 1) A82834/pMF61A 2) A82884/pMF60A 3) AB2834/pMF63A 4) A82834/pMF65A 5) A32834/pMF61A/pMF66A 6) A32834/pMF60A/pMF66A 7) ABZ834/pMF63A/pMF66A 8) A82834/pMF85A/pMF68A Figure 18 - Effect of Plasmid pMF66A on DHS Production 33 metabolite.21a Overexpression of PEP synthase (encoded by pps) was carried out in this laboratory by Kai Li. A 3.1 kb Pstl fragment containing the pps gene was amplified by PCR and cloned into pSU19 to create pKLl.87A. The only difference between pSU19 and pSU18 is the orientation of its multiple cloning site relative to a lac promoter. The aroF gene was cloned into pKLl.87A to give pMF67 (Figure 19). Transformation of each of the strains listed in Figure 16 with pMF67 was followed by analysis of the DHS production in each new construct. The results are listed in Figure 20. AB2834/pMF61A/pMF67, which expresses elevated levels of DAHP synthase, DHQ synthase, and PEP synthase, synthesized DHS in 55 mol % yield from D-glucose. Additional overexpression of transaldolase in AB2834/pMF60A/pMF67 improved the yield to 66 mol %. Transketolase overexpression in AB2834/pMF63A/pMF67 resulted in a 79 mol % yield of DHS. There was not a significant increase in the yield of DHS resulting from overexpression of both transketolase and transaldolase under these conditions, as AB2834/pMF65A/pMF67 synthesized DHS in 81 mol % yield. At this point the effect of PEP synthase had only been observed under conditions where DAHP synthase activity was quite high. Maximizing the yield of DHS might not necessarily require such elevated DAHP synthase levels in the presence of overexpressed PEP synthase. Consequently each of the strains listed in Figure 16 were transformed with pps-encoding pKLl.87A. The DHS production from each new construct is reported in Figure 21. The results from these experiments illustrate the role of transaldolase in directing carbon flow into the common pathway. Comparing AB2834/pMF60A/pKL1.87A to AB2834/pMF6lA/pKL1.87A, it appears that the overexpression of transaldolase had no impact on the production of DHS. However, comparison of AB2834/pMF65A/pKL1.87A to AB2834/pMF63A/pKL1.87A is quite different. An increase in the yield of DHS from 69 mol % to 80 mol % was obtained by increasing the catalytic activity of transaldolase. 34 Hindlll Sphl Pstl Plac pKL1.87A 5.4 kb pps EcaFll digest Pstl 1) Ecol-1| digest 2) CIAP treatment I EooRl EooRI I aroF T4 Ligase Hindlll Sphl Pstl ‘; pps Figure 19 - Preparation of Plasmid pMF67 35 mol % yield DHS Production DHS (mM) 8 6 4 8 <— theoretical maximum 7 1 4 0 I 5 7 g -- 3 2 4 3 -- 2 4 24 h I 36 h _. grammar»: 3456 1) A82834/pMF61A 2) AB2864/pMF60A 3) A82834/pMF63A 4) A82834/pMF65A 5) A82834/pMF61A/pMF66A 6) A82834/pMF60A/pMF66A 7) A82834/pMF63A/pMF66A 8) A82834/pMF65A/pMF66A 9) ABZ834/pMF61A/pMF67 10) AB2834/pMF60A/pMF67 1 1) AB2834/pMF68A/pMF67 12) A82884/pMF65A/pMF67 I I1o1112° Figure 20 - Effect of Plasmid pMF67 on DHS Production 36 mol % yield DHS Production DHS
  • L-tyrosine HO' _ OH 6H L-tryptophan shikimate Figure 22 - Proposed Synthesis of L-DOPA from D-Glucose coz' tyrosine NH; 0 phenol-lyase_ /lL . + NH3 + ‘ 002 R H OH OH pyruvate ammonia R: H: phenol R: H: L-tyrosine R=OH: catechol R=OH: L-DOPA Figure 23 - Reactions Catalyzed by Tyrosine Phenol-Lyase 43 of L-tyrosine to phenol, pyruvate, and ammonia (Figure 23) and requires pyridoxal 5'- phosphate as a cofactor.8 This 0t,B-elimination reaction is readily reversible at high concentrations of pyruvate and ammonia, leading to the formation of L-tyrosine from phenol.9'10 When catechol is substituted for phenol, L—DOPA is formed9’ll (Figure 23). Spectrophotometric studies of tyrosine phenol-lyase indicated that ammonia is the first substrate that interacts with bound pyridoxal 5'-phosphate.10b Kinetic results showed that pyruvate is the second substrate bound, thus phenol (or catechol) is the third. 10b The proposed mechanism10b for this reversible reaction is represented in Figure 24. Enzyme- bound pyridoxal 5'-phosphate interacts with ammonia, then pyruvate, resulting in the formation of an enzyme-bound a-aminoacrylate intermediate. Phenol is then added, which leads to the release of the L-tyrosine product. The reverse of this reaction scheme would most likely occur in the degradation of L-tyrosine. Overexpression of Tyrosine Phenol-Lyase in E. coli Tyrosine phenol-lyase has been studied extensively in Escherichia intermedia,6a'C Erwinia herbicola,6d and C itrobacter freundii.6'3 The gene encoding this enzyme (tpl) has been cloned and expressed in E. coli, and the DNA sequence has been published. 12 PCR amplification of the tpl gene was performed using genomic DNA isolated from C. freundii (ATCC 29063). Initially, a 2.1 kb KpnI fragment of DNA was obtained from PCR that included the complete tpl gene. This DNA fragment was cloned into pSU18 to give pW38A (Figure 25). Transformation of wild-type E. coli strain DHSa with pMF38A was followed by an enzyme assay6e for tyrosine phenol-lyase. The specific activity of tyrosine phenol—lyase from the crude extract of DHSOt/pMF38A was 0.008 units mg'l. Tyrosine phenol-lyase activity in DHSOt/pMF38A was increased to a maximum value (0.033 units mg‘l) by addition of 0.1% L-tyrosine to the culture medium. This reflects the fact that expression of tyrosine phenol-lyase is normally induced by L-tyrosine.9’13 JVIV‘ Aliv‘ N HN NH2 ’0 POH 0 OH ““3 'o POH 0 OH 3 2 / I 3 2 / I N CH3 N CH3 l OH CH2 +H,N*co.- VVIV‘ CHzR NH, NICO; 031301120 / I OH N CH3 ll 6 OH OH H JVIV‘ CH2 wlv CH2 'oaporizc OH '03POHZC OH I l -—.—"-— l N CH N CH H 3 H 3 CH3 0 “1‘” 1 _ _ HN N co2 002 _ __._ 03POH2C OH N CH3 1 ’1” CH2 H - HN NJLco2 031301-120 /| OH N CH3 1 <5 WA 03POHZC /| OH ‘9 N CH3 H Figure 24 - Proposed Mechanism of Tyrosine Phenol-Lyase 45 EcoRl KpnlifindHl Citrobacter freundii genomic DNA i PCR Kpnl Kpnl I 2.1kb ll tpl lenl digest l 1) Kpnl digest 2) CIAP treatment T4 Ligase EcoRl Kpnl Hindlll KP” Figure 25 - Preparation of Plasmid pMF38A 46 Because the presence of L-tyrosine in the culture medium would be problematic, PCR of the tpl gene was repeated with a new set of oligonucleotide primers. This time the DNA sequence that was amplified by PCR did not include the region of DNA presumed to control induction by L-tyrosine12a (Figure 26). The product of this PCR reaction, a 1.5 kb Kpnl fragment, was cloned into pSU18 to form pMF42A (Figure 27). The specific activity of tyrosine phenol-lyase from crude extracts of DHSa/pMF42A was 0.065 units mg‘l. When 0.1% L-tyrosine was added to the culture medium, the tyrosine phenol-lyase activity was unchanged (0.068 units mg'l), which verified that the induction by L-tyrosine had been eliminated. Ex 1 itin T r sine Phe l-L ase rthe nthe i f -D A Successful overexpression of tyrosine phenol-lyase in E. coli was followed by incorporation of tpl into the catechol-producing construct AB2834/pKD136/pKD9.069A.l AB2834 is an E. coli aroE mutant that cannot convert DHS into shikimate. Plasmid pKD13614 carries the genes aroF, tktA, and aroB encoding for DAHP synthase, transketolase, and DHQ synthase respectively. Amplified expression of DAHP synthase and transketolase increases the carbon flow directed into DAHP synthesis. Elevated levels of DHQ synthase ensure that DAHP is completely converted into DHS. Plasmid pKD9.069A contains the genes for DHS dehydratase2 (mol) and PCA decarboxylase3 (aroY). AB2834/pKDl36/pKD9.069A synthesizes catechol in 33 mol % yield from D- glucose.1 In order to create an L-DOPA-producing construct, the 1.5 kb tpl fragment from pMF42A was cloned into pKD9.069A to give pMF43A (Figure 28). AB2834/pKD136 was transformed with pMF43A. Despite the fact that AB2834/pKD136/pMF43A possesses all the genes required for the synthesis of L-DOPA from D-glucose, only catechol was observed as an end product. This required closer examination of the literature regarding the use of tyrosine phenol-lyase to synthesize L-DOPA from catechol. 47 5 ' —WCCAGATGTAAATAACATGAAATAAGGGATTAGCAGGTTTCGGAATAAATA 70 5'-primer (2.1 kb product) GAAATCCTCTCTATCCCAATAAACATAGTATTATTCACCGTATAAATTCATTCCTGTCCCATCCTCCTGA 140 TACCTGCCATTTCCAGAAAATAACGCCCATGTGTATCAGGCCAAAAATGAGATCTAACTCACTGAAGCAA 210 ATAGCAAACTCTGAAACAACGCCGTTTTGTACACTTTGCTTTACACTTTTAGTGATGTGAATCACAAATA 280 TAAATCCGTGAAGTGATCCGACTCTCACAAAATGGGGTGTACTCATTCAGCAATACAGTATGAGCACAGA 350 CTGGAAAGTTAAGTTTTCAGTATTTCCCCTCTCCACGGGGGCACCATCGCAAATGGCAAATCAACACGCA 420 AAAAAAAACTTGTTGAATATGAACGGGTAAAAAAATGACGTGTGATTTGCATCACCTACATTTACAGCTT 490 TTTAAATTATTGCCAGTGATTAATGTTGACTAGGCTATTTCCATAATCAATTAAATCTTGCATAGTGCCT 560 CCACATTATTTCTQQQQQTQAQZEAQEAQQCGAATAEIIAIAIIIQAIQAQAQIIIATTGAIGAAQQAGG 630 L-tyrosine induction 5'-primer (1.5 kb product) -35 TATGCTTTACTTCACWCGTACATGACCACTGTTACTQQAQAAACAAAAIQAATTATCCGGCAGA 700 -10 SD start ACCCTTCCGTATTAAAAGCGTTGAAACTGTATCTATGATCCCGCGTGATGAACGCCTTAAGAAAATGCAG 770 GAAGCGGGATACAATACTTTCCTGTTAAATTCGAAAGATATTTATATTGACCTGCTGACAGACAGTGGCA 840 CTAACGCAATGAGCGACAAGCAGTGGGCCGGCATGATGATGGGTGATGAAGCCTACGCGGGCAGCGAAAA 910 CTTCTATCATCTGGAAAGAACCGTGCAGGAACTGTTTGGCTTTAAACATATTGTTCCTACTCACCAGGGG 980 CGCGGCGCAGAAAACCTGTTATCGCAGCTGGCAATTAAACCGGGGCAATATGTTGCCGGGAATATGTATT 1050 TCACTACTACCCGTTATCACCAGGAAAAAAATGGTGCGGTGTTTGTCGATATCGTTCGTGACGAAGCGCA 1120 CGATGCCGGTCTGAATATTGCTTTTAAAGGTGATATCGATCTTAAAAAATTACAAAAACTGATTGATGAA 1190 AAAGGCGCCGAGAATATTGCCTATATTTGCCTGGCAGTCACGGTTAACCTCGCAGGCGGGCAGCCGGTTT 1260 CCATGGCTAACATGCGCGCGGTGCGTGAACTGACTGCAGCACATGGCATTAAAGTGTTCTACGACGCTAC 1330 CCGCTGCGTAGAAAACGCCTACTTTATCAAAGAGCAAGAGCAGGGCTTTGAGAACAAGAGCATCGCAGAG 1400 ATCGTGCATGAGATGTTCAGCTACGCCGACGGTTGTACCATGAGTGGTAAAAAAGACTGTCTGGTGAATA 1470 TCGGCGGCTTCCTGTGCATGAACGATGACGAAATGTTCTCTTCTGCCAAAGAGTTAGTCGTTGTCTACGA 1540 AGGCATGCCATCTTACGGCGGCCTGGCCGGACGCGACATGGAAGCCATGGCGATTGGTCTGCGCGAAGCC 1610 ATGCAGTATGAGTACATCGAGCACCGCGTGAAGCAGGTTCGCTATCTGGGCGACAAGCTGAAAGCCGCTG 1680 GTGTACCGATTGTTGAACCGGTGGGCGGTCATGCGGTATTCCTCGATGCGCGTCCGTTCTGTGAGCATCT 1750 GACGCAGGACGAGTTCCCGGCGCAAAGCCTGGCTGCCAGTATCTATGTGGAAACCGGCGTACGTAGTATG 1820 GAGCGCGGAATTATCTCTGCGGGCCGTAATAACGTGACTGGTGAACACCACAGGCCGAAACTGGAAACCG 1890 TGCGTCTGACTATTCCACGCCGCGTTTATACTTACGCGCATATGGATGTAGTGGCTGACGGTATTATTAA 1960 ACTTTACCAGCACAAAGAAGATATTCGCGGGCTGAAGTTTATTTACGAGCCGAAGCAGCTCCGTTTCTTT 2030 ACTGCACGCTTTGACTATATQIAAATAATAATTATGGCCCCATCTCAGGATCGGTCCTTTTTTGATTTCT 2100 stop TTTSQAIQAAQAGQAAQIQQIII-3' 3'-primer The putative ribosomal binding site (SD), promoter sequences (-35, -10), start codon, and stop codon are underlined. The region that is responsible for induction by L-tyrosine is indicated by arrows. Two different S'-primers were used for PCR. The size of the PCR product associated with each 5'-primer is indicated in parentheses. Figure 26 - Nucleotide Sequence of the Citrobacterfreundii tpl Gene 48 EcoRl Kpnl Hindlll Citrobacter freundii genomic DNA l PCR Kpnl Kpnl l1.5kb.| tpl lenl digest l 1) Kpnl digest 2) CIAP treatment T4 Ligase EcoRl Kpnl Plac pMF42A m Kpnl Hindlll Figure 27 - Preparation of Plasmid pMF42A 49 EcoFil Kpnl Hindlll Plac pMF42A tp, Kpnl Hindlll 1) Kpnl digest Kpnl digest 2) CIAP treatment Kpnl Kpnl I 1.5 kb . I tpl T4 Ligase Hindlll Hindlll BamHl Figure 28 - Preparation of Plasmid pMF43A 50 When whole cells expressing elevated levels of tyrosine phenol-lyase are used to synthesize L-DOPA from catechol, the temperature of the reaction is typically 15-25 0C, and the pH is maintained at 80-85911lb The concentration of catechol is very high11b (230 mM) or maintained at a steady concentration9 (about 65 mM) by frequent additions to the reaction mixture. These conditions contrast with the method in this laboratory for producing catechol from D-glucose. Biocatalysts such as AB2834/pKDl36/pKD9.069A are incubated at 37 0C in a medium containing glucose buffered to near pH 7.0. In order to achieve the conversion of D-glucose to L-DOPA, variations in culture conditions were explored. Biocatalytic conversion of D-glucose into catechol was extremely sensitive to the temperature at which the cells were incubated. As the temperature was lowered from 37 0C to room temperature, the purity of catechol produced by AB2834/pKD136/pMF43A was compromised by increasing levels of protocatechuic acid (PCA) in the culture supernatant. Decreased temperature also led to slower rates of D-glucose consumption, reducing the combined yield of end products. The effect of pH adjustment was also explored. As the pH of the culture medium was increased from 7.0 to 8.0, the amount of PCA synthesized by AB2834/pKDl36/pMF43A increased at the expense of catechol. In an effort to drive the reaction equilibrium toward L-DOPA formation, pyruvate and ammonia were added to the culture medium. This decreased the consumption of D-glucose dramatically, which may have been the result of catabolic inhibition by pyruvate. Whole cells of AB2834/pKDl36/pMF43A and DH5a/pMF42A, which both overexpress tyrosine phenol- lyase, were used to synthesize L-DOPA from catechol (230 mM), pyruvate (330 mM), and ammonia (330 mM) using a modified procedure described by Foor.11b Formation of L- DOPA was monitored by HPLC. The yield of L—DOPA produced from both strains was 12 mol % based on catechol. However, no experimental conditions were identified for converting D-glucose into L-DOPA with AB2834/pKD136/pMF43A. 51 Further information about the characteristics of tyrosine phenol-lyase was attained by determining the Km values for catechol and L-tyrosine using Lineweaver-Burk analysis.15 In the synthetic direction from catechol to L-DOPA, the Km for catechol was 8 mM. In the degradation of L-tyrosine to phenol, however, the Km for L-tyrosine was 0.4 mM. This suggests that the affinity of tyrosine phenol-lyase for L-tyrosine is roughly 20 times greater than for catechol. Therefore it is likely that the conversion of D-glucose to L- DOPA with AB2834/pKD136/pMF43A was hampered by an unfavorable reaction equilibrium. The kinetic data is also consistent with the fact that L-DOPA synthesis with tyrosine phenol-lyase requires high concentrations of catechol, pyruvate, and ammonia, presumably to drive the reaction forward. my; At this point it was suggested that the addition of tyrosine phenol-lyase activity to an L-tyrosine-producing construct might lead to the synthesis of phenol. This would require that carbon flow be directed through the common pathway and through the terminal pathway leading from Chorismate to L-tyrosine. The most critical consideration for such a process is percent conversion. The theoretical maximum percent conversion of glucose into L-tyrosine is reduced (from 43 mol %) to 30 mol % because of the additional PEP required for EPSP synthesis. 16 Irnpediments to the biocatalytic synthesis of L-tyrosine that are caused by rate-limiting, common pathway enzymes would need to be removed. Selection of an appropriate host organism and elimination of pathways leading to undesirable by-products would also be essential. Although construction of a microbial biocatalyst that can convert D-glucose into phenol seems plausible, development of a biocatalytic route to phenol that can compete with traditional synthetic methods represents a more challenging task. An alternate approach which might be developed for producing L-DOPA from D- glucose involves the catalytic action of tyrosine hydroxylase. In mammals, tyrosine 52 hydroxylase catalyzes the conversion of L-tyrosine into L-DOPA. The gene encoding this enzyme has been isolated from rat and mouse cDNA libraries and successfully expressed in E. coli.17 The addition of tyrosine hydroxylase activity to an L-tyrosine-producing construct might lead to the synthesis of L—DOPA. However, while in vitro catalytic activity of tyrosine hydroxylase expressed in E. coli has been clearly demonstrated, achieving in vivo catalytic activity is problematic. Tyrosine hydroxylase requires 5,6,7,8- tetrahydrobiopterin as a cofactor. Dihydropteridine reductase is needed to regenerate the cofactor during the course of the reaction. E. coli does not biosynthesize biopterin nor does it possess a reductase system for regenerating it. The reductase could possibly be cloned and expressed in E. coli, or a different microbial host might be used which does synthesize and catalytically recycle biopterin. The production of L-DOPA from D-glucose might eventually be realized, provided that solutions are found for all of the impediments to tyrosine hydroxylase-catalyzed conversion of L-tyrosine to L-DOPA. A number of techniques have been developed that increase the percentage of carbon flow directed into the common pathway of aromatic amino acid biosynthesis,13'21 as well as deliver the carbon flow from DAHP to Chorismate.22 Recruitment of foreign genes into E. coli has been used to create microbial constructs that can synthesize aromatic chemicals from abundant, inexpensive D-glucose. The results of this chapter illustrate that the construction of E. coli strains for the production of aromatics is not as straightforward as it appears. Nevertheless, the experiments described in this thesis provide information that can be applied toward the development of future biocatalysts. Optimization of current processes and the engineering of new biosynthetic pathways in microorganisms represent fundamental challenges in biocatalysis. However, the effort required to surmount these challenges is justified by the benefits of environmentally compatible synthesis. 10. 11. 12. 53 Wes Draths, K. M.; Frost, J. W. J. Am. Chem. Soc. 1995, 117, 2395. (a) Gross, S. R. J. Biol. Chem. 1958, 233, 1146. (b) Tatum, E. L.; Gross, S. R. J. Biol. Chem. 1956, 219, 797. (c) Strflman, P.; Reinert, W. R.; Giles, N. H. J. Biol. Chem. 1978, 253, 4593. (d) Schweizer, M.; Case, M. E.; Dykstra, C. C.; Giles, N. H.; Kushner, S. R. Gene 1981, 14, 23. (a) Patel, J. C.; Grant, D. J. W. Antonie van Leeuwenhoek 1969, 35, 53. (b) Grant, D. J. W. Antonie van Leeuwenhoek 1970, 36, 161. (a) Framck, H.-G; Stadelhofer, J. W. Industrial Aromatic Chemistry; Springer- Verlag: New York, 1988. (b) Szmant, H. H. Organic Building Blocks of the Chemical Industry; Wiley: New York, 1989. (c) Varagnat, J. In Kirk-0thmer Encyclopedia of Chemical Technology, 3rd ed.; Grayson, M, Ed.; Wiley: New York, 1981; Vol. 13, pp 39-69. Cotzias, G. C.; Papavasiliou, P. S.; Gellene, R. New Eng. J. Med. 1969, 280, 337. (a) Yamada, H.; Kumagai, H.; Matsui, H.; Ohgishi, H.; Ogata, K. Biochem. Biophys. Res. Commun. 1968, 33, 10. (b) Kumagai, H.; Yamada, H.; Matsui, H.; Ohkishi, H.; Ogata, K. J. Biol. Chem. 1970, 245, 1767. (c) Kumagai, H.; Yamada, H.; Matsui, H.; Ohkishi, H.; Ogata, K. J. Biol. Chem. 1970, 245, 1773. (d) Kumagai, H.; Kashima, N.; Torii, H.; Yamada, H.; Enei, H.; Okumura, S. Agr. Biol. Chem. 1972, 36, 472. (e) Carman, G. M.; Levin, R. E. Appl. Environ. Microbiol. 1977, 33, 192. Kakihara, Y.; Ichihara, K. Med. J. Osaka Univ. 1953, 3, 497. Yoshimatsu, H. Med. J. Osaka Univ. 1957, 9, 727. Yamada, H.; Kumagai, H. Adv. Appl. Microbiol. 1975, I9, 249. (a) Yamada, H.; Kumagai, H.; Kashima, N.; Torii, H.; Enei, H.; Okumura, S. Biochem. Biophys. Res. Commun. 1972, 46, 370. (b) Nagasawa, T.; Utagawa, T.; Goto, J.; Kim, C.-J.; Tani, Y.; Kumagai, H.; Yamada, H. Eur. J. Biochem. 1981, 117, 33. (c) Enei, H.; Nakazawa, H.; Okumura, S.; Yamada, H. Agr. Biol. Chem. 1973, 37, 725. (a) Enei, H.; Yamada, H. In Progress in Industrial Microbiology; Aida, K., Chibata, I., Nakayama, K., Takinami, K., Yamada, H., Eds.; Elsevier: Amsterdam, 1986, Vol. 24, pp 280-285. (b) Foor, F.; Morin, N.; Bostian, K. A. Appl. Environ. Microbiol. 1993, 59, 3070. (a) Iwamori, S.; Yoshino, S.; Ishiwata, K.-I.; Makiguchi, N. J. Ferment. Bioeng. 1991, 72, 147. (b) Antson, A. A.; Demidkina, T. V.; Gollnick, P.; Dauter, 2.; Von Tersch, R. L.; Long, J .; Berezhnoy, S. N.; Phillips, R. S.; Harutyunyan, E. H.; Wilson, K. S. Biochemistry 1993, 32, 4195. (c) Polak, J.; Brzeski, H.; Biotechnol. Lett. 1990, 12, 805. (d) Suzuki, H.; Nishihara, K.; Usui, N.; Matsui, H.; Kumagai, H. J. Ferment. Bioeng. 1993, 75, 145. (e) Iwamori, S.; Oikawa, T.; Ishiwata, K.-I.; Makiguchi, N. Biotechnol. Appl. Biochem. 1992, 13. 14. 15. l6. 17. 18. 19. 20. 21. 22. 54 16, 77. (f) Kurusu, Y.; Fukushima, M.; Kohama, K.; Kobayashi, M.; Terasawa, M.; Kumagai, H.; Yukawa, H. Biotechnol. Lett. 1991, 13, 769. Enei, H.; Yamashita, K.; Okumura, S.; Yamada, H. Agr. Biol. Chem. 1973, 37, 485. Draths, K. M.; Frost, J. W. J. Am. Chem. Soc. 1990, 112, 9630. Lineweaver, H.; Burk, D. J. Am. Chem. Soc. 1934, 56, 658. Frost, J. W.; Lievense, J. New J. Chem. 1994, 18, 341. (a) Wang, Y.; Citron, B. A.; Ribeiro, P.; Kaufman, S. Proc. Natl. Acad. Sci. U.S.A. 1991, 88, 8779. (b) Ichikawa, S.; Nasrin, S.; Nagatsu, T. Biochem. Biophys. Res. Commun. 1991, 178, 664. (a) Cobbett, C. S.; Morrison, S.; Pittard, J. J. Bacterial. 1984, 1 5 7, 303. (b) Garner, C. C.; Herrmann, K. M. J. Biol. Chem. 1985, 260, 3820. (c) Cobbett, C. S.; Delbridge, M. L. J. Bacteriol. 1987, 169, 2500. (d) Ogino, T.; Garner, C.; Markley, J. L.; Herrmann, K. M. Proc. Natl. Acad. Sci. U.S.A. 1982, 79, 5828. (e) Weaver, L. M.; Herrmann, K. M. J. Bacteriol. 1990, 172, 6581. (a) Draths, K. M.; Frost, J. W. J. Am. Chem. Soc. 1990, 112, 1657. (b) Draths, K. M.; Pompliano, D. L.; Conley, D. L.; Frost, J. W.; Berry, A.; Disbrow, G. L.; Staversky, R. J.; Lievense, J. C. J. Am. Chem. Soc. 1992, 114, 3956. (c) Frost, J. W. US. Patent 5 168 056, 1992. (d) Draths, K. M. Ph.D. Thesis, Stanford University, June 1991. (a) Patnaik, R.; Liao, J. C. Appl. Environ. Microbial. 1994, 60, 3903. (b) Patnaik, R.; Spitzer, R. G.; Liao, J. C. Biotechnol. Bioeng. 1995, 46, 361. (a) Miller, J. E.; Backman, K. C.; O'Connor, M. J.; Hatch, R. T. J. Ind. Microbiol. 1987, 2, 143. (b) Backman, K. C. US. Patent 5 169 768, 1992. (a) Dell, K. A.; Frost, J. W. J. Am. Chem. Soc. 1993, 115, 11581. (b) Dell, K. A. PhD. Thesis, Purdue University, December, 1993. (c) Snell, K. D.; Draths, K. M.; Frost, J. W. J. Am. Chem. Soc., in press. CHAPTER 4 EXPERIMENTAL General Me hods Chromatography HPLC analysis utilized a Rainin HPLC system with a Rheodyne injector, HPXL pumps, and a model UV—l detector. A Rainin C18 Microsorb-MV analytical column (4.6 mm x 25 cm) was used for separation and identification of components of reaction mixtures. Dowex 50 (H+ form, 100-200 mesh) was purchased from Sigma. Dowex 50 was cleaned before use as follows. An aqueous suspension of resin was adjusted to pH 14 with solid potassium hydroxide and allowed to cool to room temperature. Bromine (approximately 1 mL per 100 mL of resin) was added until the supernatant was clear yellow, and the suspension was left at room temperature for at least 3 h with occasional agitation. Dowex 50 resin was collected by filtration and washed extensively with water followed by 6 N HCl. The resin was poured into a glass column and washed with 6 N HCl (5 column volumes) followed by exhaustive rinsing with water (at least 5 column volumes). Spectroscopic Measurements 1H NMR spectra were recorded on a Varian Gemini-300 spectrometer at 300 MHz. Chemical shifts were reported in parts per million (ppm) downfield from internal sodium 3- 55 56 (trimethylsilyl)propionate-Z,2,3,3-d4 (TSP, 5 = 0.00) with DzO as solvent. TSP was purchased from Lancaster. UV and visible measurements were recorded on a Perkin-Elmer Lambda 3b UV-vis spectrophotometer connected to an R100A chart recorder. Additional UV spectra were measured on a Hewlett Packard 8452A Diode Array Spectrophotometer equipped with HP 89532A UV-Visible Operating Software. Bacterial Strains E. coli DHSOt1 [F ' endAI hst17(r'Km+K) supE44 thi-I recAI gyrA relAI ¢80lacZAM15 A(lacZYA-argF)U169] and RB7912 (W3110 lacL8Iq) were obtained previously by this laboratory. AB28343 [tsx-352 supE42 2' aroE353 mol/1352 (1')]was obtained from the E. coli Genetic Stock Center at Yale University. Citrobacter freundii (ATCC 29063)4 was obtained from the American Type Culture Collection. Storage of Bacterial Strains and Plasmids All bacterial strains were stored at -78 0C in glycerol. Plasmids were transformed into DHSOt for permanent storage. Glycerol samples were prepared by adding 0.75 mL of an overnight culture to a sterile vial containing 0.25 mL of 80% (v/v) glycerol. The solution was mixed, left at room temperature for 2 h, and then stored at -70 0C. Culture Medium All solutions were prepared in distilled, deionized water. LB medium5 contained (per liter) tryptone (10 g), yeast extract (5 g), and NaCl (10 g). M9 medium5 contained (per liter) NazHPO4 (6 g), KH2PO4 (3 g), NH4C1 (1 g), NaCl (0.5 g), MgSO4 (0.12 g), thiamine (1 mg), and D-glucose (10 g). TB medium contained (per liter) tryptone (10 g), NaCl (5 g), and MgSO4 (1.2 g). Isopropyl B-D-thiogalactopyranoside (IPTG) (0.2 mM)was added to the culture medium for strains possessing plasmids derived from pSU18 and pSU19. Ampicillin (50 ug mL'l) and chloramphenicol (20 ug mL'l) were added to 57 appropriate cultures. Stock solutions of antibiotics were prepared in water with the exception of chloramphenicol (100% ethanol). Solutions of inorganic salts, magnesium salts, and carbon sources were autoclaved separately and then mixed. Antibiotics, thiamine, and IPT G were sterilized through 0.2 um membranes prior to addition to the culture medium. Solid medium was prepared by addition of 1.5 % (w/v) Difco agar to LB and 1.5 % (w/v) agarose (Sigma, type 11: medium EEO) to M9 medium. Culture Conditions For analysis of product accumulation, bacterial strains were cultured as follows. One liter of LB (4 L Erlenmeyer flask) containing the appropriate antibiotics and [PT G was inoculated with 10 mL of an overnight culture. Cultures were grown at 37 0C in a gyratory shaker at 250 rpm for 10 h. Cells were collected by centrifugation (4000 x g, 5 min) and resuspended in 1 L of M9 medium (4 L Erlenmeyer flask) containing the appropriate antibiotics and IPT G. Cultures were then returned to the shaker (37 0C, 250 rpm). 1H NMR Analysis of Culture Supernatant Samples (25 mL) of the culture were taken at timed intervals, and the cells were removed by centrifugation (4000 x g, 5 min). A portion (2 mL) of the culture supernatant was concentrated to dryness under reduced pressure. The residue was redissolved in D20 and concentrated to dryness (2 times). The residue was then redissolved in D20 containing a known concentration of sodium 3-(trimethylsilyl)propionate-2,2,3,3-d4 (TSP, 8 = 0.00). Concentrations of cellular metabolites in the supernatant were determined by comparison of the integrals of known metabolite resonances to the resonance corresponding to TSP in the 1H NMR. Cultures were grown in triplicate to establish mean values and standard deviations. 58 General Genetic Manipulations Recombinant DNA manipulations generally followed methods described in Sambrook et al.6 All restriction enzymes were purchased from Gibco BRL or New England Biolabs. T4 DNA ligase was obtained from Gibco BRL. Calf intestinal alkaline phosphatase was obtained from Boehringer Mannheim. Agarose (electrophoresis grade) was obtained from Gibco BRL. Phenol was prepared by addition of 0.1 % (w/v) 8- hydroxyquinoline to distilled, liquefied phenol. Extraction with an equal volume of 1 M Tris-HCl pH 8.0 (two times) was followed by extraction with 0.1 M Tris-HCl pH 8.0 until the pH of the aqueous layer was greater than 7.6. Phenol was stored at 4 0C under an equal volume of 0.1 M Tris-HCl pH 8.0. SEVAG was a mixture of chloroform and isoamyl alcohol (24:1 v/v). TE buffer contained 10 mM Tris-HCl pH 8.0 and 1 mM EDTA pH 8.0. Endostop solution (10X concentration) contained 50% glycerol (v/v), 0.1 M NazEDTA, pH 7.5, 1% sodium dodecyl sulfate (SDS) (w/v), 0.1% bromophenol blue (w/v), and 0.1% xylene cyanole FF (w/v). DNAse-free RNAse (10 mg mL") was prepared according to Sambrook et al.,6 and 0.12 mL RNAse was added to 1 mL of 10X endostop. Large Scale Purification of Plasmid DNA Plasmid DNA was purified on a large scale (0.5 mg) using a modified alkaline lysis method described by Sambrook et al.6 LB (500 mL in a 2 L Erlenmeyer flask) was inoculated with a strain that contained the plasmid. The culture was incubated in a gyratory shaker (250 rpm) for 16 h at 37 0C. Cells were harvested by centrifugation (4000 x g, 5 min, 4 0C) and then resuspended in 10 mL of cold GETL solution [50 mM glucose, 20 mM Tris-HCl (pH 8.0), 10 mM NazEDTA (pH 8.0) into which lysozyme (5 mg mL'l, Sigma) had been added immediately before use]. The suspension was stored on ice for 5 min. Addition of 20 mL of 1% (w/v) sodium dodecyl sulfate in 0.2 N NaOH was followed by gentle mixing and storage on ice for 15 min. To the sample was added 15 mL 59 of cold KOAc solution (3 M K+, 5 M acetate, prepared by combining 60 mL of 5 M potassium acetate, 11.5 mL of glacial acetic acid, and 28.5 mL of H20). Vigorous shaking resulted in formation of a white precipitate. After the suspension was stored on ice for 10 min, the cellular debris was removed by centrifugation (50000 x g, 20 min, 4 0C). The supernatant was transferred to two centrifuge bottles and isopropanol (0.6 volumes) was added to precipitate the DNA. After the samples were left at room temperature for 15 min, the DNA was recovered by centrifugation (20000 x g, 20 min, 4 0C). The DNA pellet was then rinsed with 70% ethanol and dried under vacuum (10 min). Further purification of the DNA sample involved precipitation with polyethylene glycol (PEG). The DNA was dissolved in TE (3 mL) and transferred to a Corex tube. Cold 5 M LiCl (3 mL) was added, and the solution was gently mixed. The sample was then centrifuged (12000 x g, 10 min, 4 0C) to remove high molecular weight RNA. The supernatant was transferred to another Corex tube and isopropanol (6 mL) was added, followed by gentle mixing. The precipitated DNA was collected by centrifugation (12000 x g, 10 min, 4 0C). The DNA was then rinsed with 70% ethanol and dried. After redissolving the DNA in 1 mL of TE containing 20 ug of RNAse, the solution was transferred to a 1.5 mL microcentrifuge tube and stored at room temperature for 30 min. To this sample was added 500 11L of 1.6 M NaCl containing 13% PEG-8000 (w/v) (Sigma). The solution was mixed and centrifuged (microcentrifuge, 5 min, 4 0C) to recover the precipitated DNA. The supernatant was completely removed, and the DNA pellet was then redissolved in 400 1.1L of TE. The sample was extracted sequentially with phenol (400 1.1L), phenol and SEVAG (400 1.1L each), and finally SEVAG (400 11L). Ammonium acetate (10 M, 100 uL) was added to the aqueous DNA solution. After thorough mixing, 95% ethanol ( 1 mL) was added to precipitate the DNA. The sample was left at room temperature for 5 min and then centrifuged (microcentrifuge, 5 min, 4 0C). The DNA was rinsed with 70% ethanol, dried, and then redissolved in SOOuL of TE. 60 The concentration of DNA in the sample was determined as follows. An aliquot (20 uL) of the DNA was diluted to 1 mL in TE, and the absorbance at 260 nm was measured relative to the absorbance of TE. The concentration of DNA was calculated based on the fact that the absorbance at 260 nm of a 50 ug mL'1 sample of plasmid DNA is 1.0. The purity of the DNA sample was estimated by calculating the ratio of the absorbance at 260 nm to the absorbance at 280 nm. If this ratio was greater than 1.8, the purity of the DNA was sufficient for cloning purposes. Small Scale Purification of Plasmid DNA Plasmid DNA was purified on a small scale (25 1.1g) using a modified alkaline lysis method described by Sambrook et al.6 An aliquot (1.5 mL) from an overnight culture (5 mL) of the strain containing the plasmid was transferred to a microcentrifuge tube. The cells were collected by centrifugation (1 min, room temperature) and the culture supernatant discarded. The cell pellet was liquefied by vortexing (30 sec) and then resuspended in 0.1 mL of cold GETL solution. The solution was stored on ice for 10 min. Addition of 0.2 mL of 1% (w/v) sodium dodecyl sulfate in 0.2 N NaOH was followed by gentle mixing and storage on ice for 10 min. To the sample was added 0.15 mL of cold KOAc solution (3 M K“, 5 M acetate). The solution was shaken vigorously and stored on ice for 5 min before centrifugation (15 min, 4 0C). The supernatant was transferred to another microcentrifuge tube and extracted with phenol (0.2 mL) and SEVAG (0.2 mL). DNA was precipitated by the addition of 95% ethanol (1 mL). The sample was left at room temperature for 5 min before centrifugation (15 min, room temperature) to collect the DNA. The DNA pellet was rinsed with 70% ethanol, dried, and redissolved in 100 uL TE. The DNA isolated from this method was used for routine cloning and restriction enzyme analysis, and the concentration was not quantified. 61 Restriction Enzyme Digestion of DNA Restriction enzyme digests were performed using buffer solutions supplied by BRL or New England Biolabs. A typical digest contained approximately 1 ug of DNA in 8 11L TE, 1 1.1L of restriction enzyme buffer (10X concentration), and l 11L of restriction enzyme (10 units). The reaction was incubated at 37 0C for 2 h. The digest was completed by addition of 1.1 1.1L of endostop solution (10X concentration) followed by agarose gel electrophoresis. When gel electrophoresis was not required, the reaction was stopped by addition of l uL of 0.5 M EDTA (pH 8.0) followed by precipitation of the DNA. DNA was precipitated by addition of 0.1 volume of 3 M NaOAc (pH 5.2), thorough mixing, and addition of 3 volumes of 95% ethanol. Samples were stored for at least 2 h at -78 0C. Precipitated DNA was recovered by centrifugation (15 min, 4 0C). To the DNA pellet was added 70% ethanol (100 uL), and the sample was centrifuged again (15 min, 4 0C). The DNA was dried and redissolved in TE. Agarose Gel Electrophoresis Agarose gels were run in TAE buffer containing 40 mM Tris-acetate and 2 mM EDTA (pH 8.0). The concentration of the gels was typically 0.7% agarose (w/v) in TAE buffer. Higher concentrations of agarose (1%) were used to resolve DNA fragments less than 1 kb. Ethidium bromide (0.5 ug mL'l) was included in the agarose to visualize the DNA fragments with a UV lamp. The size of the DNA was determined by using two sets of DNA standards: 7» DNA digested with Hindlll or A DNA digested with EcoRI and HindIII. Isolation of DNA from Agarose The band of agarose containing the DNA of interest was excised from the gel and chopped into smaller pieces with a spatula in a 1.5 mL microcentrifuge tube. Phenol (400 11L) was added to the agarose, and the sample was vortexed for several minutes. The 62 sample was placed in a dry ice/ethanol bath for 15 min and then centrifuged (15 min, 4 OC). The aqueous layer was transferred to another 1.5 mL microcentrifuge tube and then extracted sequentially with phenol (400 11L), phenol and SEVAG (400 uL each), and finally SEVAG (400 uL). The DNA was precipitated with 3 M NaOAc and 95% ethanol as previously described and redissolved in TE. Treatment of Vector DNA with Calf Intestinal Alkaline Phosphatase Plasmid vectors digested with a single restriction enzyme were dephosphorylated to prevent self-ligation. Digested vector DNA was dissolved in TE (88 uL). To this sample was added 10 11L of dephosphorylation buffer (10X concentration, Boehringer Mannheim) and 2 1.1L of calf intestinal alkaline phosphatase (2 units). The reaction was incubated at 37 0C for 1 h. The phosphatase was inactivated by addition of 1 1.1L of 0.5 M EDTA (pH 8.0) followed by heat treatment (65 0C, 20 min). The sample was extracted with phenol and SEVAG (100 uL each) to remove the phosphatase protein, and the DNA was precipitated as previously described and redissolved in TE. Ligation of DNA DNA ligations were designed so that the molar ratio of insert to vector was at least 3 to l. A typical ligation reaction contained 0.1 to 0.5 ug of vector and 0.5 to 1 ug of insert in a total volume of 14 11L. To this sample was added 4 11L of ligation buffer (5X concentration, BRL) and 2 uL of T4 DNA ligase (2 units). The reaction was incubated at 16 0C for at least 4 h and then used to transform competent cells. Preparation and Transformation of Competent Cells Competent cells were prepared using a procedure modified from Sambrook et al.6 An aliquot (1 mL) from an overnight culture (5 mL) was used to inoculate 100 mL of LB (500 mL Erlenmeyer flask) containing the appropriate antibiotics. The cells were cultured EE—t' ' 63 in a gyratory shaker (37 0C, 250 rpm) until they reached the mid-log phase of growth (judged from the absorbance at 600 nm reaching 0.6). The culture was poured into a large centrifuge bottle that had been previously sterilized with bleach and rinsed with autoclaved water. The cells were collected by centrifugation (4000 x g, 5 min, 4 0C) and the culture medium discarded. At this point all further manipulations were carried out on ice. The cell pellet was washed with 100 mL of cold 0.9% (w/v) NaCl and then resuspended in 50 mL of cold 100 mM CaClz. The suspension was stored on ice for at least 30 min and then centrifuged (4000 x g, 5 min, 4 0C). The cell pellet was resuspended in 4 ml. of cold 100 mM CaClz containing 15% glycerol (v/v). Aliquots (0.25 mL) were dispensed into 1.5 mL microcentrifuge tubes and immediately frozen in liquid nitrogen. The competent cells were stored at -78 0C with no significant decrease in transformation efficiency over a period of several months. Frozen competent cells were thawed on ice for 5 min before transformation. A small aliquot (l to 10 11L) of plasmid DNA or a ligation reaction was added to the thawed competent cells (0.25 mL). The solution was gently mixed and stored on ice for 30 min. The cells were then beat shocked at 42 0C for 2 min and placed on ice briefly (30 s). LB (1 mL, no antibiotics) was added to the cells, and the sample was incubated at 37 0C (no agitation) for l h. Cells were collected in a microcentrifuge (30 s), resuspended in a small volume of LB (0.1 mL), and then spread onto LB plates that contained the appropriate antibiotics. A sample of competent cells with no DNA added was also carried through the transformation procedure as a control. These cells were used to check the viability of the competent cells and to verify the absence of growth on selective medium. Purification of Genomic DNA Genomic DNA was purified using a modified method described by Silhavy.7 A single colony of the strain was inoculated into 100 mL of TB medium (500 mL Erlenmeyer flask). The cells were cultured in a gyratory shaker (37 0C, 250 rpm) for 12 h. 64 Centrifugation (4000 x g, 5 min, 4 0C) of the culture was followed by resuspension of the cell pellet in 5 mL of buffer [50 mM Tris-HCl (pH 8.0), 50 mM EDTA (pH 8.0)] and storage at ~20 0C for 20 min to freeze the suspension. To the frozen cells was added 0.5 mL of 0.25 M Tris-HCl (pH 8.0) that contained 5 mg of lysozyme. The suspension was thawed at room temperature in a water bath with gentle mixing and then stored on ice for 45 min. The sample was then transferred to a Corex tube. After addition of 1 mL of STEP solution [25 mM Tris-HCl (pH 7.4), 200 mM EDTA (pH 8.0), 0.5% SDS (w/v), and proteinase K (1 mg mL'l, Sigma), prepared just before use], the mixture was incubated at 50 0C for at least 1 h with gentle, periodic mixing. The solution was then divided into two Corex tubes, and the contents of each tube were extracted with phenol (4 mL). The organic and aqueous layers were separated by centrifugation (1000 x g, 15 min, room temperature), and the aqueous layer was transferred to another Corex tube. All transfers of the aqueous layer were carried out using wide bore pipette tips to minimize shearing of the genomic DNA. The contents of each tube were extracted again with a mixture of phenol (3 mL) and SEVAG (3 mL). Extractions with phenol/SEVAG were repeated (6 to 7 times) until the aqueous layer was clear. Genomic DNA was precipitated by addition of 0.1 volume of 3 M NaOAc (pH 5.2), gentle mixing, and addition of 2 volumes of 95% ethanol. Threads of DNA were spooled onto a sealed Pasteur pipette and transferred to a Corex tube that contained 5 mL of 50 mM Tris-HCl (pH 7.5), 1 mM EDTA (pH 8.0), and 1 mg of RNAse. The mixture was stored at 4 0C overnight to allow the DNA to dissolve completely. The solution was then extracted with SEVAG (5 mL) and centrifuged (1000 x g, 15 min, room temperature). The aqueous layer was transferred to another Corex tube and the genomic DNA was precipitated as before with 3 M NaOAc and 95% ethanol. The threads of DNA were spooled onto a Pasteur pipette and redissolved in 2 mL of 50 mM Tris-HCl (pH 7.5) and 1 mM EDTA (pH 8.0). Genomic DNA was stored at 4 oC. . .. ' ‘1.- .h' it“! 65 Enzyme Assays Cells were grown in LB medium as described under Culture Conditions. Centrifugation of the culture (4000 x g, 5 min, 4 0C) was followed by resuspension of the cell pellet in a buffer appropriate for the enzyme assay. The volume of resuspension buffer (mL) was usually twice the wet weight (g) of the cells. The cells were disrupted by two passages through a French pressure cell (SLM Aminco) at 18000 psi. Cellular debris was removed from the lysate by centrifugation (48000 x g, 20 min, 4 0C). Protein was quantified using the Bradford dye-binding procedure.8 A standard curve was prepared using bovine serum albumin. The protein assay solution (5X concentration) was purchased from Bio-Rad. Chapter 2 D-Erythrose 4—Phosphate D-Erythrose 4-Phosphate (E4P) was synthesized by oxidation of D-glucose 6- phosphate with lead tetraacetate.9 D-Glucose 6-phosphate (0.56 g, 2 mmol) was dissolved in water (4 mL) and added to 500 mL of glacial acetic acid in a 1 L three-necked round bottom flask. A solution of lead tetraacetate (1.51 g, 3.4 mmol) in 80 mL of acetic acid containing 1.2 mL of 6 N H2SO4 was added dropwise over a 2 h period under a nitrogen atmosphere with stirring. A fine white precipitate formed over the course of the reaction, which was removed upon filtration of the reaction mixture through Celite. The filtrate was concentrated by rotary evaporation to 80 mL and excess acetic acid was removed by azeotropic distillation with water (at least 5 times). The solution (80 mL) was loaded onto a column of Dowex 50 (H+ form, 90 mL) and eluted with 300 mL of water. The concentration of E4P in the eluent was determined by using an aliquot of the solution as a substrate in the transaldolase assay,10 which measures the change in absorbance at 340 nm resulting from a decrease in NADH (e = 6220 L mol'1 cm'l). Dilute E4P solutions (less 66 than 4 mM) were stored in a dark bottle at room temperature. The yield of E4P was 75% based on D-glucose 6-phosphate. Assays DAH(P) The combined concentration of DAH and DAHP was determined by thiobarbituric acid visualization of the periodate cleavage products.11 An aliquot (0.1 mL) of solution containing DAH/DAHP was added to 0.2 M NaIO4 in 8.2 M H3PO4 (0.1 mL) and incubated at 37 0C for 5 min. The periodate oxidation was quenched by adding 0.5 mL of a solution containing 0.8 M NaAst and 0.5 M NaZSO4 in 0.1 M H2804. Thorough mixing of this sample was followed by addition of 3 mL of 0.04 M thiobarbituric acid in 0.5 M NaZSO4 (pH 7.0). The mixture was incubated at 100 0C for 15 min and the pink chromophore was extracted into 4 mL of distilled cyclohexanone. The organic and aqueous layers were separated by centrifugation (2000 x g, 15 min, room temperature). The absorbance of the organic layer was measured at 549 nm (8 = 68000 L mol‘1 cm‘l). DAHP synthase DAHP synthase activity was measured according to the procedure described by Schoner.12 Harvested cells were resuspended in 50 mM potassium phosphate (pH 6.5) that contained 10 mM PEP and 0.05 mM CoClz. The cells were then disrupted as previously described. The lysate was diluted in potassium phosphate (50 mM), PEP (0.5 mM), and 1,3-propanediol (250 mM), pH 7.0. A dilute solution of E4P was concentrated to 12 mM by rotary evaporation and neutralized with 5 N KOH. A solution was then prepared that contained E4P (6 mM), PEP (12 mM), ovalbumin (1 mg mL'l), and potassium phosphate (25 mM), pH 7.0. An aliquot (0.5 mL) of the diluted lysate was combined with 1 mL of the E4P/PEP solution. After these solutions were mixed (time = 0), the reaction was incubated at 37 0C for 3 min. Aliquots (0.15 mL) were removed at T] arm mm: “a“ n J J i 67 timed intervals and quenched with 0.1 mL of 10% trichloroacetic acid (w/v). Precipitated protein was removed by microcentrifugation, and the DAH(P) in each sample was quantified by the thiobarbituric acid assay. One unit of DAHP synthase activity was defined as the formation of 1 umol of DAHP per min. Transaldolase Transaldolase was assayed according to the method described by Tsolas.10 The transaldolase-catalyzed conversion of D-fructose 6-phosphate and E4P into D- sedoheptulose 7-phosphate and D-glyceraldehyde 3-phosphate (GAP) was monitored. For the determination of transaldolase activity the GAP formed per unit time was related to the loss of NADH by coupling enzymes triosephosphate isomerase (TIM) and glycerophosphate dehydrogenase (GDH). Cells were resuspended in 50 mM potassium phosphate (pH 7.6) and disrupted in a French pressure cell as previously described. The assay solution contained D-fructose 6-phosphate (2.7 mM), E4P (0.2 mM), NADH (0.1 mM), triethanolamine (91 mM), pH 7.6, EDTA (91 mM), TIM (14 units), and GDH (2 units) in a total volume of 2.7 mL. The absorbance at 340 nm was monitored at room temperature until a stable baseline was obtained. An aliquot (0.05 mL) of diluted lysate was added and the absorbance at 340 nm monitored for 10 min. One unit of transaldolase activity was defined as the loss of one umol of N ADH (8 = 6220 L mol‘l cm'l) per min. Transketolase Transketolase activity was quantified using a procedure described by Paoletti.l3 The transketolase-catalyzed conversion of E4P and B—hydroxypyruvate into D-fructose 6- phosphate and C02 was assayed. Formation of D-fructose 6-phosphate was related to NADPH formation through coupling enzymes phosphoglucoisomerase (PGI) and glucose 6-phosphate dehydrogenase (G6PDH). Cells were resuspended in buffer containing potassium phosphate (50 mM), pH 7.4, MgC12 (1 mM), and dithiothreitol (0.2 mM). 68 Cells were disrupted as previously described. The assay solution (0.98 mL) contained triethanolamine (125 mM), pH 7.6, MgC12(5 mM), thiamine pyrophosphate (0.1 mM), [3- hydroxypyruvate (0.4 mM), NADP (0.4 mM), E4P (0.1 mM), G6PDH (3 units), and PGI (10 units). The solution was incubated at room temperature for several minutes until the absorbance at 340 nm was constant. An aliquot (0.02 mL) of diluted lysate was then added and the reaction was monitored at 340 nm for 20 min. One unit of transketolase activity was defined as the formation of one umol of NADPH (8 = 6220 L mol'1 cm‘l) per min. PCR Amplification of talB, tktA, and aroF PCR reactions were performed in a single-block thermocycler purchased from Ericomp. VentR® DNA polymerase was purchased from New England Biolabs. Oligonucleotide primers were synthesized by the Macromolecular Structure Facility in the Department of Biochemistry at Michigan State University. Stock solutions of 2'- deoxynucleoside 5'-triphosphates were purchased from Pharmacia. Plasmids used as templates for PCR were purified by PEG precipitation. Genomic DNA used as templates for PCR was isolated as previously described. The concentration of DNA from solutions of oligonucleotide primers was determined by measuring the absorbance at 260 nm. The extinction coefficient for each primer was calculated from the sum of the extinction coefficients for each dNTP14 (A = 15400 L mol‘l cm‘l; C = 7400 L mol'1 cm'1;G = 11700 L mol‘1 cm'l; T = 8700 L mol‘l cm'l). Primers were dissolved in water or TE and diluted to 10 uM. A typical PCR reaction contained the following: 10 1.1L of 2 mM dNTPs, 20 uL of each primer (10 11M), 10 1.1L of VentR® polymerase reaction buffer (10X concentration), VentR® polymerase (2 units) and template DNA in a total volume of 100 11L. The amount of template DNA was usually 1 ug for genomic DNA and 0.05 1.1g for plasmid DNA. PCR reactions were can'ied out in 0.6 mL microcentrifuge tubes. Mineral oil (100 uL) was added to each reaction mixture immediately before starting PCR to minimize evaporation of the sample. After completion of the temperature cycling program, the PCR reaction PF: 69 mixture was removed from the mineral oil. A small aliquot of the sample was analyzed by agarose gel electrophoresis. The remainder of the sample was extracted with a mixture of phenol and SEVAG (0.1 mL each). DNA was precipitated by addition of 0.1 volume of 3 M NaOAc (pH 5.2), thorough mixing, and addition of 3 volumes of 95% ethanol. A 1.3 kb fragment containing the talB gene was amplified from E. coli RB791 genomic DNA. Oligonucleotide primers used were as follows: JWF46 (5'-GTACGTACCCATGGTI'I‘AAGAAGTATATACGCTA—3') JWF47 (5'-CATGCATGCCATGG'I'I‘AAACAGTCTCGTTAAACA-3') The following temperature cycling program was used: 1 cycle of 94 0C (4 min); 30 cycles of 94 0C (1 min), 60 0C (1 min), 72 0C (1.3 min). The size of the product was confirmed on a 0.7% agarose gel. The PCR product was digested with Ncol and purified by agarose gel electrophoresis. Vector pBR32515 was digested with Neal and treated with calf intestinal alkaline phosphatase (CIAP). Ligation of the talB fragment into pBR325 resulted in the formation of plasmid pMF52A (7.3 kb). The orientation of the insert with respect to the host vector was determined by analysis of the DNA fragments that resulted from digestion with BamHI. A 2.2 kb tktA fragment was amplified from plasmid pKD130A.l6 The template DNA was prepared by linearizing pKD130A with Sphl before use. The following primers were used: JWF38 (5'-CGGGATCCTGGTCCGCAAACGGACATTA-3') JWF39 (5'-CGGGATCCAGAGATITCTGAAGC-3') The program for PCR was as follows: 1 cycle of 94 0C (4 min); 25 cycles of 55 0C (1 min), 72 0C (2.2 min), 94 0C (1 min). The size of the fragment was verified on a 0.7% agarose gel. The tktA fragment was digested with BamHI and gel purified. The insert was then ligated to pBR325 that had been previously digested with BamHI and treated with CIAP, resulting in the formation of pMF51A (8.2 kb). The orientation of the insert with 70 respect to the host vector was determined by analysis of the DNA fragments that resulted from digestion with AvaI. The aroF gene was amplified as a 1.25 kb fragment from plasmid pKD130A that had been previously linearized by Sphl. The primers used for PCR were as follows: JWF l 9 (5'-GGAA'ITC'I‘TAAGCCACGCGAGCCGT-3') JWF22 (5'-GGAATTCAAAGGGAGTGTAAATI‘TAC-3') The following program was used for PCR: 1 cycle of 94 OC (4 min); 30 cycles of 94 oC (1 min), 55 0C (1 min), 72 0C (1.3 min). The size of the PCR product was confirmed by running a 0.7% agarose gel. The DNA fragment was digested with EcoRI and isolated from an agarose gel. After digestion of pBR325 and treatment with CIAP, the vector was ligated to the aroF fragment to give pMF58A (7.25 kb). The orientation of the insert with respect to the host vector was determined by analysis of the DNA fragments that resulted from digestion with PvuII. Plasmid Constructions Plasmid pMF61A Plasmid pMF58A was digested with Sphl and treated with CIAP. The 1.65 kb aroB fragment was isolated from an agarose gel following Sphl digestion of plasmid pKDl36. Ligation of these two fragments gave plasmid pMF61A (8.9 kb). The orientation of the insert with respect to the host vector was determined by analysis of the DNA fragments that resulted from digestion with Add]. Plasmid pMF60A Plasmid pMF52A was digested with Sphl and treated with CIAP. The 1.65 aroB fragment was ligated to pMF52A to give pMF57A (8.95 kb). Digestion of pMF57A with EcoRI was followed by CIAP treatment. The 1.25 kb aroF fragment was purified by 71 agarose gel electrophoresis from an EcoRI digest of pMF58A. Ligation of the aroF fragment to pMF57A yielded pMF60A (10.2 kb). Plasmid pMF63A Plasmid pMF61A was digested with BamHI and treated with CIAP. The 2.2 kb tktA fragment was isolated from an agarose gel following BamHI digestion of pMF51A. Ligation of tktA into pMF61A resulted in plasmid pMF63A (11.1 kb). Plasmid pMF65A Plasmid pMF63A was digested with NcaI followed by CIAP treatment. Isolation of the 1.3 kb talB fragment from an NcoI digest of pMF52A was followed by ligation to pMF63A to give pMF65A (12.4 kb). Plasmid pMF66A Vector pSU18l7 was digested with EcoRI and treated with CIAP. Ligation of the 1.25 kb aroF fragment to pSU18 yielded pMF66A (3.55 kb). Plasmid pMF67 Plasmid pKLl.87A18 contains a 3.1 kb pps fragment in pSU19. Digestion of pKLl.87A with EcoRI and CIAP treatment was followed by ligation to the 1.25 kb aroF fragment. This resulted in the formation of plasmid pMF67 (6.65 kb). M13 Tyrosine Phenol-Lyase Assay Tyrosine phenol-lyase activity was assayed according to the method described by Kumagai, which measures the amount of pyruvate formed from L-tyrosine.l9 Cells were 72 resuspended in 50 mM potassium phosphate (pH 8.0) and disrupted with a French pressure cell as previously described. The assay solution (3.9 mL) contained L-tyrosine (1.25 mM) and pyridoxal 5'-phosphate (0.025 mM) in 50 mM potassium phosphate buffer (pH 8.0). Lysate (0.1 mL) was added (time = 0), and the mixture was incubated at 30 0C in a water bath with gentle shaking for 20 min. Aliquots (0.5 mL) were removed at timed intervals and quenched with 10% trichloroacetic acid (3 mL). Precipitated protein was removed by centrifugation (2000 x g, 15 min, room temperature). The pyruvate in each sample was quantified as the 2,4-dinitrophenylhydrazone derivative by the method of Friedemann.20 A 2,4-dinitrophenylhydrazine (2,4-DNP) reagent solution was prepared as follows. Solid 2,4—DNP was ground in a mortar with addition of small amounts of 2 N HCl until 100 mL was added, followed by gravity filtration of the solution. The 2,4-DNP reagent was stored at 4 OC and was prepared freshly each week. An aliquot (3 mL) of the deproteinized supernatant from the tyrosine phenol-lyase assay was added to 1 mL of 2,4-DNP reagent in a large test tube. The mixture was vortexed (15 s), and the hydrazone derivative was extracted into benzene (3 mL). The aqueous layer was discarded, and to the organic layer was added 10% NazCO3 (6 mL). After this solution was vortexed (30 s), an aliquot (5 mL) of the aqueous layer was withdrawn and added to 1.5 M N aOH (5 mL) in another test tube. This solution was then mixed briefly and left at room temperature for 10 min. The absorbance of this solution at 520 nm was measured. The concentration of pyruvate in the sample was determined from a standard curve that had been prepared with pyruvate. One unit of tyrosine phenol- lyase activity was defined as the formation of l umol of pyruvate per min. PCR Amplification of tpl The PCR conditions were identical to those previously described under Chapter 2 with the exception that oligonucleotide primers were synthesized by National Biosciences. 73 Initially a 2.1 kb fragment containing the tpl gene was amplified from Citrobacterfreundii (ATCC 29063) genomic DNA. Oligonucleotide primers used were as follows: Primer A (5'-'ITGGTACCCCCAGTAATTGGCGGGAAGT-3') Primer B (5'-ACGGTACCAAAGGAC'ITCCTGTTCATGG-3') The following temperature cycling program was used: 1 cycle of 94 0C (5 min); 25 cycles of 94 0C (l min), 55 0C (1 min), 72 0C (2.1 min). The size of the product was confirmed on a 0.7% agarose gel. The PCR product was digested with Kpnl and purified by agarose gel electrophoresis. Vector pSU18 was digested with Kpnl and treated with CIAP. Ligation of the 2.1 kb tpl fragment into pSU18 gave plasmid pMF38A (4.4 kb). The orientation of the insert with respect to the host vector was determined by analysis of the DNA fragments that resulted from digestion with Sphl. Iwamori identified a palindromic region of DNA located approximately 100 bp upstream from the translational start site for tpl.21 It was presumed that this region was concerned with the induction of L-tyrosine.21 In order to generate a tpl fragment that did not require L-tyrosine induction, a new PCR primer was designed to anneal to the 5’-end of the tpl gene immediately downstream from the aforementioned palindromic sequence. New Primer A (5'-GGGGTACCG'ITATATITCATCAGACTTT-3') With New Primer A and Primer B, a 1.5 kb tpl fragment was amplified from C. freundii genomic DNA. The PCR program was as follows: 1 cycle of 94 0C (5 min); 25 cycles of 94 0C (l min), 55 0C (1 min), 72 0C (1.5 min). The size of the PCR product was verified on a 0.7% agarose gel. The 1.5 kb tpl fragment was digested with Kpnl, gel purified, and ligated to pSU18 that had been digested with Kpnl and treated with CIAP. This resulted in the formation of plasmid pMF42A (3.8 kb). Construction of Plasmid pMF43A Plasmid pKD9.069A22 contains a 3.5 kb aroZ fragment and a 2.4 kb aroY fragment localized in pSU19. Digestion of pKD9.069A with Kpnl was followed by CIAP 74 treatment. Isolation of the 1.5 kb tpl fragment from a Kpnl digest of pMF42A was followed by ligation of this insert to pKD9.069A to give pMF43A (9.7 kb) Culture Conditions for Converting D-Glucose into L—DOPA The culture conditions described previously under General Methods were tried initially with AB2834/pKDl36/pMF43A, but no production of L-DOPA was observed. The effect of pH on the reaction was explored as follows. Six samples of LB (50 mL in a 250 mL Erlenmeyer flask) were each inoculated with 0.25 mL of an overnight culture of AB2834/pKD136/pMF43A. The cells were grown in a gyratory shaker (37 0C, 250 rpm) for 12 h. The cells were collected by centrifugation (4000 x g, 5 min) in bleach-sterilized bottles and washed with 10 mL of 50 mM potassium phosphate (pH 8.0). Each of the six cell pellets was resuspended in M9 medium (25 mL in a 125 mL Erlenmeyer flask) that had been adjusted to a different pH value (7.0, 7.2, 7.4, 7.6, 7.8, and 8.0). The cultures were then returned to the shaker (37 0C, 250 rpm). An aliquot (3 mL) of each culture was removed at 24h and 48h and the culture supernatant was analyzed by 1H NMR. The effect of temperature on the reaction was explored as follows. The procedure for growing the cells was the same as for the pH study, except that only three cultures were used. Each culture was incubated in M9 (pH 7.0) at a different temperature (37 0C, 30 0C, and 25 0C). An aliquot (3 mL) of each culture was removed at 24h and 48h and the culture supernatant was analyzed by 1H NMR. Addition of excess pyruvate and ammonia to the M9 culture was performed as follows. AB2834/pKD136/pMF43A was grown exactly as for the pH study. Six cultures were used. The M9 media was adjusted in the following manner. Pyruvate (330 mM) was added to two of the M9 cultures and the pH adjusted to 7.5 and 8.0. Ammonium chloride (330 mM) was added to two of the M9 cultures and the pH adjusted to 7.5 and 8.0. Both pyruvate (330 mM) and NH4C1 (330 mM) were added to two of the M9 cultures and the pH adjusted to 7.5 and 8.0. The six cell pellets were each resuspended in one of the M9 75 cultures and returned to the shaker (37 0C, 250 rpm). An aliquot (3 mL) of each culture was removed at 24h and 48h and the culture supernatant was analyzed by 1H NMR. Culture Conditions for Converting Catechol into L-DOPA The synthesis of L-DOPA from catechol was carried out using a modified procedure described by Foor.23 LB (50 mL) was inoculated with 0.25 mL of an overnight culture from a strain that overexpressed tyrosine phenol-1yase. The cells were harvested (4000 x g, 5 min) and washed with 10 mL of 50 mM potassium phosphate (pH 8.0). The cells were then resuspended in a 25 mL of a solution that contained catechol (230 mM), pyruvate (330 mM), NH4C1 (330 mM), B(OH)3 (240 mM), NaZSO3 (30 mM), and EDTA (7 mM) (adjusted to pH 8.5 with 5 N NH4OH). The cell suspension was incubated at room temperature for 24 h. An aliquot of the culture was removed and the culture supernatant analyzed by HPLC. The yield of L-DOPA was 12 mol % based on catechol. HPLC Analysis of Reaction Mixtures For the HPLC analysis of catechol and L-DOPA, an aliquot of the supernatant from the reaction was diluted in 25 mM potassium phosphate (pH 3.0). Diluted samples (20 to 50 uL) were analyzed on a Rainin HPLC using a C18 analytical column (4.6 mm x 25 cm). The HPLC program was as follows. For the first 5 min, the mobile phase was 5% (v/v) methanol in 25 mM potassium phosphate (pH 3.0). Then from 5 min to 20 min, methanol was increased over a linear gradient to 60 %. The absorbance at 280 nm was monitored and the flow rate was 1 mL min'l. The retention time for L-DOPA and catechol under these conditions was about 5.6 min and about 15.6 min, respectively. Determination of KM Values for Tyrosine Phenol-Lyase For the determination of the KM for catechol (8 mM), the assay solution (4 mL) contained 50 mM potassium phosphate (pH 8.0), 200 mM NH4Cl, 120 mM pyruvate, [CT 76 0.025 mM pyridoxal 5'-phosphate, 1 unit of tyrosine phenol-lyase, and catechol concentrations of 5, 10, 15, 20, 25, and 30 mM. The samples were incubated in a water bath at 30 0C with gentle shaking. Aliquots (0.5 mL) were removed at timed intervals and quenched with concentrated H2SO4 (5 1.1L). The precipitated protein was removed by microcentrifugation and the concentration of L-DOPA in the sample determined by HPLC analysis. A Lineweaver-Burk plot24 of V'1 versus [catechol]'l was generated, where V is the velocity of the reaction (umol L-DOPA formed per min) and [catechol] is the concentration of catechol (M). The x-intercept of this plot is equal to -KM'1. The KM for L-tyrosine (0.4 mM) was determined as follows. The normal tyrosine phenol-lyase assay (as previously described) was performed. The reaction contained 0.2 units of enzyme and varying concentrations of L-tyrosine (0.05, 0.1, 0.25, 0.5, l, and 2 mM). A plot of V'1 versus [L-tyrosine]'l was generated, where V is the velocity of the reaction (umol pyruvate formed per min) and [L-tyrosine] is the concentration of L-tyrosine (M). The x-intercept of this plot is equal to -KM'1. 10. ll. 12. 13. 14. 15. 16. 17. 18. 19. 20. 21. 77 References Current Protocols in Molecular Biology; Ausubel, F. M., Brent, R., Kingston, R. E., Moore, R. E., Seidman, J. G., Smith, J. S., Struhl, K, Eds.; Wiley: New York, 1987. Frost, J. W.; Bender, J. L.; Kadonaga, J. T.; Knowles, J. R. Biochemistry 1984, 23, 4470. Pittard, J .; Wallace, B. J. J. Bacteriol. 1966, 91, 1494. Carman, G. M.; Levin, R. E. Appl. Environ. Microbiol. 1977, 33, 192. Miller, J. H. Experiments in Molecular Genetics; Cold Spring Harbor Laboratory: Plainview, NY, 1972. Sambrook, J.; Fritsch, E. F.; Maniatis, T. Molecular Cloning: A Laboratory Manual; Cold Spring Harbor Laboratory: Plainview, NY, 1990. Silhavy, T. J .; Berrnan, M. L.; Enquist, L. W. Experiments with Gene Fusions; Cold Spring Harbor Laboratory: Plainview, NY, 1984. Bradford, M. M. Anal. Biochem. 1976, 72, 248. Sieben, A. S.; Perlin, A. S.; Simpson, F. J. Can. J. Biochem. 1966, 44, 663. (a) Tsolas, O.; Horecker, B. L. Arch. Biochem. Biophys. 1970, 136, 287. (b) Brand, K. In Methods of Enzymatic Analysis, 3rd ed.; Bergmeyer, H. U., Ed.; Verlag Chemie: Weinheim, 1983, Vol. 4, pp 334-340. Gollub, E.; Zalkin, H.; Sprinson, D. B. Methods Enzymal. 1971, 17A, 349. Schoner, R.; Herrmann, K. M. J. Biol. Chem. 1976, 251, 5440. Paoletti, F.; Williams, J. F.; Horecker, B. L. Anal. Biochem. 1979, 95, 470. Kiessling, L. L.; Griffin, L. C.; Dervan, P. B. Biochemistry 1992, 31, 2829. (a) Bolivar, F. Gene 1978, 4, 121. (b) Prentki, P.; Karch, F.; Iida, S.; Meyer, J. Gene 1981, I4, 289. Draths, K. M.; Frost, J. W. J. Am. Chem. Soc. 1990, 112, 1657. Martinez, E.; Bartolomé, B.; de la Cruz, F. Gene 1988, 68, 159. Li, K.; Frost, J. W., unpublished results. Kumagai, H.; Yamada, H.; Matsui, H.; Ohkishi, H.; Ogata, K. J. Biol. Chem. 1970, 245, 1767. Friedemann, T. E.; Haugen, G. E. J. Biol. Chem. 1943, I47, 415. Iwamori, S.; Yoshino, S.; Ishiwata, K.-I.; Makiguchi, N. J. Ferment. Bioeng. 1991, 72, 147. 78 22. Draths, K. M.; Frost, J. W. J. Am. Chem. Soc. 1995, 117, 2395. 23. Poor, F.; Morin, N.; Bastian, K. A. Appl. Environ. Microbiol. 1993, 59, 3070. 24. Lineweaver, H.; Burk, D. J. Am. Chem. Soc. 1934, 56, 658. Hrcnu STATE UN I It | IV~ fiIBRfiRIEs llllllllllllllllIllllllllIlllllllllllllllllllllllllllllllllll 31293014057693 l'lIC