PURIFICATION, CHARACTERIZATION AND SITE-DIRECTED MUTAGENESIS OF A METHANOGEN RIBOFURANOSYLAMINOBENZENE 5â€™-PHOSPHATE SYNTHASE By MATTHEW EDWARD BECHARD A THESIS PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLOR IDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF MASTER OF SCIENCE UNIVERSITY OF FLORIDA 2003
Copyright 2003 by Matthew Edward Bechard
This work is dedicated to my parents Ka thryn and Edward Bechard. The love and support they provided throughout my life is what has enabled me to be where I am today, have what I have today, and be who I am toda y. None of this would be possible without them. This work is also dedicated to my grandfather â€œPepereâ€ who wanted with all his heart to see me get this fa r, but didnâ€™t get the chance.
ACKNOWLEDGMENTS I thank Dr. Madeline E. Rasche for her guidance, instruction, patience, and intellectual contributions. Her persistence, strong will, moral standing, and pure love of science have shaped me into a better scientist. I would also like to acknowledge Joseph Scott. His groundbreaking work laid the foundation for all of the ribofuranosylaminobenzene 5â€™-phosphate (RFAP) synthase studies in this work. Thanks go to Stephanie Wyles and Marco Caccamo, for their support and helpful discussions. Special thanks go to the undergraduate students in Dr. Rascheâ€™s Advanced Microbiology Laboratory. Their work on the alanine-screening mutations laid the foundation for our site-directed mutagenesis studies. I would especially like to thank Courtney Malone, Dina Greene, Rosemarie Garcia, and Sonya Chhatwal who worked on the initial characterization of the variants described in our study, as well as variants not described in this work. Finally the author would like to thank the rest of the committee (Dr. Nemat Keyhani and Dr. Julie Maupin-Furlow), for their endless helpful discussions and contributions to the interpretation of the work. iv
TABLE OF CONTENTS Page ACKNOWLEDGMENTS .................................................................................................iv LIST OF TABLES ............................................................................................................vii LIST OF FIGURES .........................................................................................................viii ABSTRACT .......................................................................................................................ix CHAPTER 1 INTRODUCTION........................................................................................................1 Overview.......................................................................................................................1 Methane Production......................................................................................................1 Methanogens.................................................................................................................3 Methanogenesis............................................................................................................6 Cofactors of Methanogenesis.....................................................................................11 Methanopterin and Folate...........................................................................................18 Biosynthesis of Methanopterin...................................................................................20 RFAP synthase............................................................................................................24 2 OVEREXPRESSION, PURIFICATION, CHARACTERIZATION, AND SITE-DIRECTED MUTAGENESIS OF A METHANOGEN RFAP SYNTHASE PRODUCED IN Escherichia coli..............................................................................29 Introduction.................................................................................................................29 Materials and Methods...............................................................................................31 Colorimetric Assay for RFAP Synthase.............................................................31 Heterologous Expression of MTH830 in E. coli................................................31 Purification of Recombinant RFAP Synthase.....................................................33 Determination of Optimal Enzymatic Conditions..............................................34 Kinetic Analyses of the Recombinant Methanogen RFAP Synthase.................35 Partial Purification and Analyses of M. thermautotrophicus RFAP Synthase...35 Partial Purification and Analyses of the M. thermophila RFAP Synthase.........36 Site-directed Mutagenesis of Recombinant RFAP Synthase..............................36 Partial Purification of RFAP Synthase Variants.................................................37 Kinetic Analyses of Recombinant RFAP Synthase Variants.............................38 v
Protein Quantitation and Gel Electrophoresis.....................................................38 Materials and Chemicals.....................................................................................38 Results.........................................................................................................................39 Overexpression of a Methanogen RFAP Synthase in E. coli.............................39 Purification of a Methanogen RFAP Synthase Produced in E. coli...................40 Determination of Optimal Enzymatic Conditions..............................................45 Kinetic Analyses of Recombinant RFAP Synthase............................................48 Bioinformatics of RFAP Synthase......................................................................52 Site-directed Mutagenesis...................................................................................53 Discussion...................................................................................................................54 3 CONCLUSIONS........................................................................................................63 Significance................................................................................................................63 Future Experimentation..............................................................................................65 LIST OF REFERENCES...................................................................................................67 BIOGRAPHICAL SKETCH.............................................................................................78 vi
LIST OF TABLES Table page 2-1. Primers used for site-directed mutagenesis................................................................37 2-2. Purification of RFAP synthase produced in E. coli....................................................43 2-3. Site-directed mutagenesis of the recombinant wild-type RFAP synthase..................54 vii
LIST OF FIGURES Figure page 1-1. Universal phylogenetic tree determined from rRNA sequence comparisons.............4 1-2. Autotrophic growth of hydrogenotrophic methanogens.............................................7 1-3. Methanogenesis from acetate (Aceticlastic pathway)...............................................12 1-4. Lactate oxidation pathway of A. fulgidus..................................................................14 1-5. Methanol utilization pathway of M. extorquens........................................................16 1-6. Structures of folate, methanopterin, and derivatives of methanopterin....................19 1-7. Proposed biosynthetic pathway for methanopterin...................................................22 2-1. Soluble versus insoluble fractions of RFAP synthase expression in KB1................41 2-2. Purification of recombinant RFAP synthase.............................................................42 2-3. Optimal temperature and pH of RFAP synthase.......................................................44 2-4. Effect of enzyme concentration on velocity of reaction...........................................46 2-5. Determination of pABA K m for recombinant RFAP synthase..................................47 2-6: Determination of pABA K m for native RFAP synthase............................................47 2-7: Determination of pABA K m for M. thermophila RFAP synthase.............................49 2-8. Phylogenetic relationship of RFAP synthases homologs...........................................50 2-9. Bioinformatic analysis of group 1 RFAP synthase genes.........................................51 viii
Abstract of Thesis Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Master of Science OVEREXPRESSION, PURIFCATION, CHARACTERIZATION, AND SITE-DIRECTED MUTAGENESIS OF A METHANOGEN RIBOFURANOSYLAMINOBENZENE 5â€™-PHOSPHATE SYNTHASE PRODUCED IN Escherichia coli By Matthew Edward Bechard December 2003 Chair: Madeline E. Rasche Major Department: Microbiology and Cell Science Methanogens and sulfate-reducing archaea are among a select group of microorganisms that use methanopterin as a one-carbon carrier instead of folate. The biosynthesis of methanopterin involves eighteen steps, the first of which is catalyzed by ribofuranosylaminobenzene 5â€™-phosphate (RFAP) synthase, a key enzyme proposed to be found only in methanopterin-containing organisms. This enzyme is interesting in that it catalyzes the decarboxylation of p-aminobenzoic acid (pABA) during the phosphoribosyl transfer from phosphoribosylpyrophosphate (PRPP) onto pABA to form RFAP. In our study, we have overexpressed, purified, and biochemically characterized the Methanothermobacter thermautotrophicus RFAP synthase recombinantly produced in Escherichia coli. Bioinformatic analysis of homologous methanogen RFAP synthase sequences reveal highly conserved blocks of amino acids, one of which resembles a region of dihydropteroate synthase proposed to be involved in pABA binding. The ix
residues D19 and R26, located in this conserved region of RFAP synthase, were altered by site-directed mutagenesis and then shown to be important for substrate binding. Because of the vital role of this enzyme in the synthesis of methanopterin and the importance of methanopterin in the generation of methane gas from methanogenic archaea, biochemical characterization and identification of the amino acids involved in the activity of a methanogen RFAP synthase would be beneficial. This work provides information about this essential methanopterin biosynthetic enzyme that could be key in reducing the problem of methane gas emitted from methanogenic archaea, and also in identifying other methanopterin-containing organisms. x
CHAPTER 1 INTRODUCTION Overview Methanogens are a select group of microorganisms that are classified under the third domain of life, the Archaea. In Methanothermobacter thermautotrophicus, a well studied methanogenic archaeon, methanopterin carries C 1 compounds as CO 2 is reduced to methane gas during methanogenesis, an essential process that is the only means of ATP production for these methanogenic organisms. As a result of the importance of methanopterin in the generation of methane, a potent greenhouse gas, the identification and characterization of the biosynthetic enzymes would be beneficial. The biosynthetic pathway for methanopterin involves eighteen steps, the first step of which is a key reaction catalyzed by the enzyme ribofuanosylaminobenzene 5â€™-phophaste (RFAP) synthase. The focus of our study is the recombinant production and biochemical characterization of RFAP synthase from M. thermautotrophicus. The knowledge gained from this project has potential to reduce the levels of methane released as a greenhouse gas from methanogenic organisms into the atmosphere. Methane Production Methane, the simplest of all hydrocarbons, is a greenhouse gas that has been implicated in the involvement of warming early earth, providing a temperature that was essential for the beginnings of life (Kasting 1997; Drake 2000; Pavlov et al. 2000). Greenhouse gases like methane trap solar energy in the atmosphere where it is then transmitted to the earth as heat. The methane released into the atmosphere is thought to 1
2 be removed by reacting with hydroxyl radicals present in the atmosphere, producing several byproducts that result in an indirect contribution to global warming as well (Drake 2000; Wuebbles and Hayhoe 2002). The lifespan of atmospheric methane is estimated to be between 8 and 11.8 years (Drake 2000). Over the years as the human population has increased, the production of methane from man-made and controlled sources, like sheep and cow populations, biomass burning, coal mining, landfills, and natural gas production, has also increased (Pimentel 1991; Drake 2000). As a direct result of this increased methane emission, the amount of reactive hydroxyl radicals is also on the decline thus increasing the lifespan of methane as well as the proposed problem of global warming (Drake 2000). There are many sources of atmospheric methane gas from both biogenic and nonbiogenic processes. Non-biogenic sources of methane include its release during coal and oil drilling, as well as during the incomplete combustion of fossil fuels (Johnson and Johnson 1995; Drake 2000). Methane gas is also present along with other hydrocarbons in natural gas. Biogenic sources of methane (which account for most of the total methane emitted) include landfills, wetlands, rice paddies, incomplete combustion of biomass, and the group of animals called ruminants (sheep, cows, and termites) (Khalil and Shearer 1993; Wuebbles and Hayhoe 2002). A significant amount of the biogenic methane, approximately 74%, comes from methanogenic organisms present in ruminants of livestock (Khalil and Shearer 1993). Methanogenic organisms are characteristically defined by their unique ability to use carbon dioxide, other one-carbon compounds, and acetate as growth substrates in the overall production of methane (Zinder 1993). Habitats for methanogenic organisms include environments such as freshwater and marine
3 sediments, the gastrointestinal tracts of animals, swamps, landfills, geothermal vents, and anaerobic waste digesters, where the anaerobic biodegradation of organic compounds provides methanogens with the proper substrates required for their growth (Zinder 1993; Thauer 1998). The generation of methane gas by methanogens present in ruminants is a significant source that produces 80 million tons of methane per year; and is second only to the processing and use of coal, oil, and natural gas (Khalil and Shearer 1993). As a result of the significant contribution of atmospheric methane by methanogens, inhibition of the essential metabolic pathways of methanogenesis is a strategy that is being used in reducing the generation of methane from these organisms (Wolin 1981). Nonspecific inhibitors of methanogenesis like 2,4-bis (trichoromethyl)-benzo[1,3]dioxins and the ionophore monensin have previously been developed with limited success (Thornton and Owens 1981; Davies et al. 1983). However these inhibitors also seem to negatively affect beneficial processes like acetogenesis, which provides nutrients used by the ruminant animals (Chen and Wolin 1979). Nonetheless an increase in knowledge of both methanogens and methanogenic processes has provided valuable information that allows for different, perhaps more successful, strategies for the inhibiting methanogenesis. One particular strategy involves inhibiting methanogenesis by developing an inhibitor specific for a methanopterin biosynthetic enzyme (Dumitru et al. 2003). Methanogens Methanogenic organisms are a group of microorganisms classified in the third domain of life, the Archaea (Woese 1987). Phylogenetic classification of organisms is based on a comparison of 16s rRNA and divides living organisms into three domains
4 Bacteria Archaea Eukarya Green non-sulfur bacteria Gram positives proteobacteria Entamoeba Slime molds Halophiles Methanosarcina Methanothermobacter Thermococcus Thermoproteus Pyrodictium Methanococcus Universal Ancestor Microsporidia Diplomonads Trichomonads Flagellates Ciliates Plants Fungi Animals Thermotogales Flavobacteria Cyanobacteria Figure 1-1. Universal phylogenetic tree determined from rRNA sequence comparisons (Adapted from Woese, C. R., O. Kandler, and M. L. Wheelis. 1990. Towards a natural system of organisms Proposal for the domains archaea, bacteria, and eucarya. Proc. Nat. Acad. Sci. USA 87:4576-4579).
5 (Bacteria, Archaea, and Eukarya) classifying Archaea closer to Eukarya (Figure 1-1) (Woese et al. 1990). Despite being separately classified the archaeal domain has characteristics that resemble both bacteria and eukarya (Woese 1987). Bacteria and Archaea are prokaryotic organisms that have similar morphologies as well as 30S and 50S ribosomal subunits with 16s rRNA and 23s rRNA (Jones et al. 1987; Howland 2000). However archaea, like eukaryotic organisms, use methionine as an initiator tRNA and multisubunit RNA polymerases (Jones et al. 1987). Unlike both Bacteria and Eukarya, however, archaea use ether-linked phospholipids in the makeup of their distinctly unique cell-wall structures (de Rosa et al. 1986). Archaea generally are divided into three phenotypes, methanogenic, extremely halophilic, and extremely thermophilic (Woese 1987). Archaea are also known to inhabit many extreme environmental conditions, including anaerobic, acidic, extremely thermophilic, and high salt (Howland 2000). Methanogenic organisms in particular are of great interest because of their ability to thrive under extreme anaerobic, reductive conditions, where they participate in the terminal reduction of one-carbon compounds to methane (Zeikus 1977). This ability makes methanogenic organisms an integral part of the biodegradation of biomass, where they compete with acetogenic organisms for the hydrogen and carbon dioxide produced as end products from other organisms (Mackie and Bryant 1994). Methanogens are further divided into five orders Methanobacteriales, Methanococcales, Methanomicrobiales, Methanosarcinales, and Methanopyrales (Boone et al. 1993). Two particular methanogenic organisms with diverse characteristics are Methanothermobacter thermautotrophicus (f. Methanobacterium thermoautotrophicum H) from the Methanobacteriales and Methanosarcina thermophila from the
6 Methanosarcinales. M. thermautotrophicus, a nonmotile rod shaped organism, grows hydrogenotrophically on H 2 /CO 2 at a pH range of 7.0 to 8.0 and a temperature range of 55 to 70C (Zeikus 1972). M. thermophila however, is slightly different in that it is a nonmotile, irregular shaped aggregate of cocci that uses the methanopterin derivative sarcinapterin as its one-carbon carrier (Zinder and Mah 1979). M. thermophila, unlike M. thermautotrophicus, also grows under more diverse conditions using acetate as well as methylated one carbon compounds like methanol and methylamines as growth substrates at a pH range of 5.5 â€“ 8.0 and temperatures ranging from 35 to 50C (Zinder and Mah 1979). Methanogenesis During the autotrophic growth of hydrogenotrophic methanogens like M. thermautotrophicus, hydrogen is used in the reduction of CO 2 to methane gas as seen in Figure 1-2 (Zeikus 1972). In contrast M. thermophila, which does not grow autotrophically on H 2 /CO 2 , uses acetate and other methylated one carbon compounds to generate methane and energy (Zinder and Mah 1979). The process of methanogenesis is the only means by which methanogenic organisms produce ATP (Thauer et al. 1993; Deppenmeier et al. 1996). In the initial step of methanogenesis by hydrogenotrophic methanogens, CO 2 is reduced by molecular hydrogen and attached to the coenzyme methanofuran (MFR) (Leigh et al. 1985). This initial step is an endergonic reaction catalyzed by the membrane bound hydrogenase, formyl-MFR dehydrogenase, and is proposed to involve several electron carriers driven by a transmembrane electrochemical ion potential (Bobik et al. 1990; Borner et al. 1991). The formyl group is transferred from MFR to the
7 CO 2 Formyl MFR 5-Formyl H 4 MPT Methenyl H 4 MPT Methylene H 4 MPT Methyl CoM Reductase Methyl H 4 MPT CoM â€“ S â€“ CH 3 CH 4 CoB â€“ SH CoB â€“ S â€“ S CoM e MethylH 4 MPT: HS-CoM Methyltransferase Na + Electron Transport Chain Na + /H + e Formyl-MF dehydrogenase e Hydrogenase 2 e Hydrogenase 1 Electron Transport chain Heterodisulfide reductase H 2 + F 420 Formyl-MF:H 4 MPT formyltransferase F 420 -reducing hydrogenase Methenyl-H 4 MPT Cyclohydrolase H 2 Methylene-H 4 MPT Dehydrogenase F 420 H 2 Methylene-H 4 MPT Reductase H 2 H + + CoM â€“ SH CoB â€“ SH Figure 1-2. Autotrophic growth of hydrogenotrophic methanogens. H 4 MPT, tetrahydromethanopterin; MFR, methanofuran; CoM â€“ S â€“ CH 3 , methylcoenzyme M; CoB â€“ SH, coenzyme B; CoB â€“ S â€“ S â€“ CoM, heterodisulfide; F 420 H 2 , coenzyme F 420 reduced form (Adapted from Deppenmeier, U., V. Muller, and G. Gottschalk. 1996. Pathways of energy conservation in methanogenic archaea. Arch. Microbiol. 165:149-163).
8 coenzyme tetrahydromethanopterin (H 4 MPT) by the enzyme formylmethanofuran: H 4 MPT formyltransferase (Donnelly and Wolfe 1986). The formyl group is reduced to the methenyl level by methenyl-H 4 MPT cyclohydrolase (Donnelly et al. 1985). Methenyl-H 4 MPT is further reduced to methylene-H 4 MPT by methylene-H 4 MPT dehydrogenase (Schworer and Thauer 1991). The electrons used in this reduction step are donated by the reduced form of coenzyme F 420 , which reduces these one carbon intermediates in the same manner as NAD (Jacobson and Walsh 1984; DiMarco et al. 1990). After donation of electrons from the reduced form of F 420 , the oxidized form is re-reduced by the F 420 -reducing hydrogenase (Thauer et al. 1993; Deppenmeier et al. 1996). The reduced F 420 is used in the reduction of methylene-H 4 MPT to methyl-H 4 MPT by methylene-H 4 MPT reductase (Te Brommelstroet et al. 1990). The next step in the methanogenic process proceeds in two steps. First, the methyl group is transferred from H 4 MPT to the corrinoid 5â€™-hydroxybenzimidaxolyl cobamide (Factor III) in the cob(I)alamin oxidation state (Kengen et al. 1990; Kengen et al. 1992). Next the methyl group is transferred to -mercaptoethanesulfonic acid or coenzyme M (Kengen et al. 1990; Kengen et al. 1992). This two step reaction is catalyzed by methyl-H 4 MPT:coenzyme M methyltransferase enzyme which is an integral membrane protein complex of several different subunits with factor III bound (Kengen et al. 1992). The transfer of the methyl group from H 4 MPT to coenzyme M is an exergonic reaction, with a free energy change of -29.7 kJ/mol, coupled to the translocation of sodium ions across the plasma membrane contributing to the sodium ion gradient used to generate ATP (Becher et al. 1992; Becher and Muller 1994; Deppenmeier et al. 1996). The terminal reduction of methyl-coenzyme M to methane is catalyzed by a nickel tetrapyrrol (F 430 ) bound
9 methyl CoM methylreductase (Ellermann et al. 1988; Ellermann et al. 1989; Hedderich et al. 1990). Electrons from 7-mercaptoheptanoyl-threonine phosphate or coenzyme B are used to reduce methyl-coenzyme M to a methyl group, released as methane (Ellermann et al. 1989; Ermler et al. 1997). The heterodisulfide of coenzyme M and coenzyme B (CoM-S-S-CoB), produced as a product of the reduction of methyl-coenzyme M to methane, is subsequently reduced to coenzyme M and coenzyme B by the FAD-containing heterodisulfide reductase (Hedderich et al. 1990). In the reduction of the heterodisulfide, electrons from molecular hydrogen are transported to CoM-S-S-CoB through an electron transport chain made up of a multisubunit protein complex that includes the heterodisulfide reductase (Hedderich et al. 1990). This reaction is also accompanied by the generation of a proton ion gradient (Hedderich et al. 1990; Becher and Muller 1994; Setzke et al. 1994). It is also thought that the exergonic reduction of the heterodisulfide also drives the endergonic reduction of CO 2 with H 2 to formyl-MFR (Kaesler and Schonheit 1989). The generation of sodium and proton motive forces during methanogenesis is used by Na + and H + -driven ATPases for the synthesis of ATP (Becher and Muller 1994; Deppenmeier et al. 1996). M. thermophila can not grow autotrophically on H 2 /CO 2 , instead this methanogen grows using either acetate or other methylated one-carbon compounds like methanol, and methylamines (Zinder and Mah 1979). The aceticlastic pathway, as seen in figure 1-3, is used during growth of M. thermophila on acetate (Ferry 1992; Thauer et al. 1993). In this pathway the acetate is first phosphorylated with ATP to acetyl phosphate by acetate kinase (Aceti and Ferry 1988). Acetyl-phosphate is then converted to acetyl-CoA by phosphotransacetylase (Latimer and Ferry 1993). Acetyl-CoA then interacts with carbon
10 monoxide dehydrogenase (CODH) which cleaves the C-C and C-S resulting in the attachment of the carbonyl group to the Ni/Fe-S component of CODH and attachment of the methyl group to factor III in the Co/Fe-S component (Abbanat and Ferry 1991; Raybuck et al. 1991). The methyl group is transferred to the H 4 MPT derivative tetrahydrosarcinapterin and reduced to methane as previously described for the CO 2 -reducing pathway (Hedderich et al. 1989; Jablonski et al. 1990; Grahame 1991; Jablonski and Ferry 1991; Schworer and Thauer 1991). The electrons needed for the reduction of the heterodisulfide come from oxidation of the carbonyl group to carbon dioxide (Hedderich et al. 1989; Schworer and Thauer 1991). Again, the generation of proton and sodium motive forces are coupled to the generation of ATP by ATPases (Peinemann et al. 1988; Deppenmeier et al. 1996). M. thermophila also has the ability to grow on other one-carbon compounds like methanol, producing methane and CO 2 as end products (Zinder and Mah 1979). Methanogenesis from methanol involves the disproportionated metabolism of methanol in an oxidative and reductive branch. The inability of these methanogens to use molecular hydrogen in the reduction of the one-carbon compounds forces the cells to oxidize one-fourth of the methanol to CO 2 in the oxidative branch of the pathway, to provide electrons for the reduction of the remaining three-fourths of the methanol to methane in the reductive branch (Zinder and Mah 1979; Deppenmeier et al. 1996). In the reductive branch of the pathway the methyl group of methanol is first bound to the corrinoid prosthetic group of methanol:5-hydroxybenzimidaxolyl methyltransferase (Van Der Meijden et al. 1983; Van Der Meijden et al. 1983). Subsequently the methyl group is transferred to coenzyme M by methyl-cobalamin:HS-CoM methyltransferase (Van Der
11 Meijden et al. 1983; Van Der Meijden et al. 1983). The terminal reduction of the methyl group to methane, catalyzed by methyl-coenzyme M reductase, occurs using coenzyme B as the reductant producing the heterodisulfide as a byproduct (Ellermann et al. 1989; Ermler et al. 1997). In the oxidative branch of the pathway it is thought that the methyl group is transferred from methyl-CoM to H 4 MPT by the Na + -translocating methyl-H 4 MPT:HS-CoM methyltransferase (Muller et al. 1988). From this point the methyl group is then oxidized to CO 2 in the reverse direction of the CO 2 reduction pathway described previously (Donnelly et al. 1985; Leigh et al. 1985; Donnelly and Wolfe 1986; Te Brommelstroet et al. 1990; Schworer and Thauer 1991). The reduction of the methyl group to CO 2 produces reducing equivalents (i.e. F 420 H 2 ) that are then used in the reduction of the heterodisulfide, which is coupled to the generation of a proton ion gradient used to generate ATP (Deppenmeier et al. 1990). Cofactors of Methanogenesis The study of methanogenic archaea led to the discovery of seven unusual coenzymes, coenzyme B, coenzyme M, F 420 , F 430 , factor III, methanopterin, and methanofuran (DiMarco et al. 1990; Maden 2000). However, as knowledge in this area increased it became apparent that a few of these cofactors, specifically, methanopterin, methanofuran, F 420 , and coenzyme M, are more widely distributed than previously thought. Most notably, the coenzyme methanopterin as well as a number of its derivatives has recently been found among both non-methanogenic archaea as well as methylotrophic bacteria (Mller-Zinkhan et al. 1989; Chistoserdova et al. 1998; Maden 2000). H 4 MPT was first found to have a role outside methanogenic archaea in the sulfate-reducer Archaeoglobus fulgidus, where it also serves as a one-carbon carrier (Mller-Zinkhan et al. 1989). A. fulgidus, although shown to use methanopterin,
12 Acetate ATP ADP + P i HS-CoA Acetyl-CoA CODH complex NiFeS component CoFeS component HS-CoA [CO] [CH 3 ] H 4 SPT CODH complex 2e CO 2 CH 3 -H 4 SPT H 4 SPT HS-CoM CH 3 -SCoM HS-CoB CoM-S-S-CoB CH 4 Figure 1-3. Methanogenesis from acetate (Aceticlastic pathway). CODH, carbon monoxide dehydrogenase; H 4 SPT, tetrahydrosarcinapterin; HS-CoM, coenzyme M; HS-CoB, coenzyme B; CoM-S-S-CoB, heterodisulfide; CH 3 -SCoM, methyl-coenzyme M; CH 3 -H 4 SPT, methyl-tetrahydrosarcinapterin. (Adapted from Ferry, J. G. 1992. Methane from acetate. J. Bacteriol. 174:5489-5495).
13 was also shown to use F 420 and methanofuran (Mller-Zinkhan et al. 1989). However, this non-methanogenic archaeon does not use the methylreductase system or any cofactors associated with it (Mller-Zinkhan et al. 1989). In A. fulgidus these methanogenic cofactors are utilized in a modified acetyl-CoA pathway used for the oxidation of lactate to CO 2 as shown in Figure 1-4 (Mller-Zinkhan et al. 1989). In this pathway lactate is first oxidized to pyruvate by lactate dehydrogenase and then further oxidized to acetyl-CoA by pyruvate dehydrogenase. From this point acetyl-CoA is then split by CODH, oxidizing the carbonyl group to CO 2 as previously described in the aceticlastic pathway. The methyl group is then oxidized to CO 2 by the hydrogenotrophic CO 2 reduction pathway in the reverse direction, using methanopterin dependent enzymes, F 420 -dependent hydrogenases, as well as methanofuran (Mller-Zinkhan et al. 1989). The presence of a dephospho derivative of the one-carbon carrier H 4 MPT, along with H 4 MPT dependent enzymes, in the methylotrophic bacteria Methylobacterium extorquens was an interesting discovery that showed a distribution of this cofactor beyond the Archaeal domain (Chistoserdova et al. 1998; Vorholt et al. 1999). It was shown by Chistoserdova et al. (1998) that M. extorquens has a 14-kb gene cluster containing 13 open reading frames, 10 of which were shown to be necessary for growth of the bacteria methylotrophically on one-carbon compounds. Of these 10 genes five were shown to have high sequence homology to enzymes in the H 4 MPT/MFR pathway in both methanogens and A. fulgidus. This was further supported by the detection of enzymatic activities and identification of genes encoding a methylene H 4 MPT dehydrogenase (MtdB), methenyl H 4 MPT cyclohydrolase (MchA), and formyl-methanofuran:H 4 MPT formyltransferase/hydrolase (Fhc)
14 Lactate P y ruvate 2 [H] Acet y l-Co A 2 [H] Meth y l H 4 MPT CO CO 2 2 [H] Methen y l H 4 MPT Meth y lene H 4 MPT 2 [H] 2 [H] Form y l MFR 2 [H] 5-Form y l H 4 MPT CO 2 Figure 1-4. Lactate oxidation pathway of A. fulgidus (Adapted from Mller-Zinkhan, D., G. Brner, and R. K. Thauer. 1989. Function of methanofuran, tetrahydromethanopterin, and coenzyme F420 in Archaeoglobus fulgidus. Arch. Microbiol. 152:362-368).
15 (Chistoserdova et al. 1998; Vorholt et al. 1998; Pomper et al. 1999; Hagemeier et al. 2000; Vorholt et al. 2000; Pomper et al. 2002). Of these enzymes methylene H 4 MPT dehydrogenase was shown to not have significant homology to its F 420 -dependent counterpart in methanogens, instead it was shown to have homology to another NADP-specific methylene H 4 MPT dehydrogenase that uses both H 4 MPT and H 4 F as substrates (Chistoserdova et al. 1998). Although, the formylmethanofuran: H 4 MPT formyltransferase/hydrolase in M. extorquens was shown to have homology to its counterpart in methanogens, the enzyme did not catalyze the oxidation of formyl:methanofuran to CO 2 as previously thought (Pomper et al. 2002). Instead the enzyme was shown to catalyze the hydrolysis of formyl:methanofuran to formate (Pomper et al. 2002). Using this information a methanopterin-dependent pathway was proposed for the oxidation of formaldehyde to CO 2 during the methylotrophic growth of M. extorquens on one-carbon compounds (Figure 1-5) (Vorholt et al. 1999; Pomper et al. 2002; Chistoserdova et al. 2003). Methylotrophic growth of M. extorquens on one-carbon compounds such as methanol, and methylamines results in the production of the toxic compound formaldehyde. Therefore, it was proposed that this pathway is used to not only detoxify the formaldehyde but also provide energy and cell carbon (Nunn and Lidstrom 1986; Vorholt et al. 1998; Chistoserdova et al. 2003). This methanol utilization pathway begins with the oxidation of methanol to formaldehyde by methanol dehydrogenase (Anthony 1982; Anderson and Lidstrom 1988). Formaldehyde at this point is attached to H 4 MPT by a novel formaldehyde activating enzyme, forming
16 Methanol HCOH Methylene H 4 MPT NAD(P)H Meth y lene H 4 F Methen y l H 4 F NADPH Methenyl H 4 MPT N 5 -Formyl H 4 MPT H 2 O Formyl Methanofuran H 2 O N 10 -Form y l H 4 F ATP Formate NADH CO 2 Figure 1-5. Methanol utilization pathway of M. extorquens. (Adapted from Chistoserdova, L., S. W. Chen, A. Lapidus, and M. E. Lidstrom. 2003. Methylotrophy in Methylobacterium extorquens AM1 from a genomic point of view. J. Bacteriol. 185:2980-2987)
17 methylene-H 4 MPT (Vorholt et al. 2000). Methylene-H 4 MPT is oxidized to formate and CO 2 by the corresponding aforementioned enzymes in a reversal of the CO 2 reduction pathway (Chistoserdova et al. 1998; Vorholt et al. 1998; Pomper et al. 1999; Hagemeier et al. 2000; Vorholt et al. 2000; Pomper et al. 2002). The formate produced is thought to feed into a H 4 F pathway, producing a pool of H 4 F bound intermediates utilized in the serine cycle as well as other biosynthetic reactions (Pomper et al. 2002). Coenzyme F 420 is a 5â€™-deazaflavin with a midpoint potential of -360 mV that serves as the central electron donor in methanogenic archaea (Jacobson and Walsh 1984). However, F 420 , like methanopterin, has also been found outside methanogens among the bacterial species, Streptococcus, Mycobacterium, and cyanobacteria where it serves as an electron donor (Mccormick and Morton 1982; Lin and White 1986; Bair et al. 2001). Coenzyme M (-mercaptoethanesulfonic acid) is a thiol cofactor that functions as a methyl carrier in the reduction of methyl-H 4 MPT to methane (Balch and Wolfe 1979). However, it has also been found to have a role outside of methanogens in the bacterial domain where it serves as a carrier and activator of alkyls in aliphatic epoxide carboxylation (Allen et al. 1999). The remaining three coenzymes, F 430 , coenzyme B, and factor III, have thus far not been found outside of methanogens (DiMarco et al. 1990; Maden 2000). F 430 is a non-fluorescent nickel tetrapyrrole with an absorbance max at 430 nm (Gunsalus and Wolfe 1978). The coenzyme is a prosthetic group non-covalently attached to methyl coenzyme M reductase carriers the methyl group during its reduction to methane (Thauer 1998). Coenzyme B (7-mercaptoheptanoylthreonine phosphate) is a low-molecular weight, heat-stable thiol cofactor that donates its electrons in the reduction of methyl-coenzyme M to
18 methane (Gunsalus and Wolfe 1980). 5â€™-hydroxybenzimidaxolyl cobamide or factor III is a derivative of the cobamide B 12 that has 5-hydroxybenzymidazole as its -ligand (Krautler et al. 1987). Methanopterin and Folate The transfer of one-carbon intermediates over different oxidation states involves the one-carbon carrier H 4 MPT and its derivatives, or H 4 F. The basic structure of methanopterin consists of a pterin ring attached to an arylamine, derived from pABA, with an extensive side chain attached to the arylamine, consisting of a ribitol residue attached to ribose 5â€™-phosphate with a hydroxyglutaric acid residue on the end (Figure 1-6) (Maden 2000; White 2001). The active form of methanopterin is the tetrahydro form in which the C7 and C11 positions are methylated (White 1996; Maden 2000; White 2001). The active sites of the coenzyme where the onecarbon intermediates are carried is at the N 5 and N 10 positions of the pterin moiety (White 1996; Maden 2000; White 2001). There have been several derivatives of methanopterin found in both the Archaeal and Bacterial domains all of which have the same basic structure of H 4 MPT with different side chains attached to the arylamine. Some of these methanopterin derivatives include dephospho-methanopterin, sarcinapterin, sulfopterin, and a modified form of methanopterin found in Pyrococcus furiosus, all of which are pictured in figure 1-6 (Lin and White 1988; Mller-Zinkhan et al. 1989; White 1991; White 1993; Chistoserdova et al. 1998; Vorholt et al. 1999). The structure of folate is very similar to that of methanopterin in that it also consists of a pterin ring attached to pABA however, with a less extensive side chain (Figure 1-6). The biologically active form of folate is also the fully reduced tetrahydro form with active sites at the N 5 and N 10 positions (Maden 2000). The two coenzymes, as previously described, are similar in that the basic structures
19 NNHNNOH2NCH3H2CCHCHHOOHCH2OCH2OPOHOOCCOOHCH2HH2CCOOHCHOHHNCH3OOHOH NNHNNOH2NNHCONHCHCOOHCH2COOH NNHNNOH2NH2CHNCHCHHOOHCHOHCH2OH NNHNNOH2NCH3H2CCHCHHOOHCH2OCH2OHCHOHHNCH3OOHOH NNHNNOH2NCH3H2CCHCHHOOHCH2OCH2OPOHOOCCCH2HCH2COOHCHOHHNCH3OHOOHONHCHCOOHCH2CH2COOH Sarcinapterin Dephospho methanopterin Sulfopterin Folate Methanopterin Figure 1-6. Structures of folate, methanopterin, and derivatives of methanopterin.
20 are a pterin ring attached to pABA. However, when the structure of these two coenzymes are compared there are a number of differences that distinguish these one-carbon carriers both structurally and thermodynamically. The most important of these differences is the absence of a carbonyl group para to the N 10 position in methanopterin that is normally present in folate (Maden 2000; White 2001). This carbonyl group para to the N 10 position in folate has an electron withdrawing effect on one carbon units attached to the N 10 position, whereas the methylene group para to the N 10 in methanopterin has a slightly electron-donating effect (Thauer et al. 1996; Maden 2000; White 2001). This key difference affects the thermodynamic properties of bound one-carbon units by conferring more negative mid-point potentials on the one-carbon redox transitions in methanopterin than in folate (Thauer et al. 1996; Maden 2000). The overall result of this difference is that methanopterin binds the one-carbon units more tightly than folate. Other differences between these coenzymes are in the side chains attached to the arylamine. The side chain of methanopterin is a long and complex side chain consisting of a ribitol residue attached to ribose 5-phosphate followed by a hydroxyglutarate residue, whereas in folate the side chain only consists of hydroxyglutarate residues attached to pABA. The polyglutamation of the folate side is however, thought to be involved in enzyme specificity, with different enzymes having preferences for different number of glutamates (Shane 1989). Another difference is the methylation of methanopterin at the C7 and C11 positions, thought to restrict the rotation of the long side chain, which is not present in folate (Maden 2000; White 2001). Biosynthesis of Methanopterin The biosynthesis of methanopterin was investigated for the methanogenic archaeon Methanosarcina thermophila and has been proposed to involve eighteen
21 steps (White 1996; White 2001). The biosynthetic pathway begins with the formation of the pterin ring from GTP in a slightly different manner than in folate biosynthesis. The GTP cyclohydrolase, which produces 7,8-dihydroneopterin triphosphate in folate biosynthesis, results in the formation of triaminopyrimidine triphosphate a novel intermediate that seems to be unique to the synthesis of methanopterin in methanogenic archaea (White 2001; Graham et al. 2002). This novel intermediate is then converted to 7,8-dihydroneopterin 2â€™:3â€™-cyclic phosphate by unknown enzymes, and finally to 6-hydroxymethyl-7,8-dihydropterin pyrophosphate in reactions identical to the corresponding reactions in folate biosynthesis (White 2001). This portion of the pathway produces a pool of pterin pyrophosphate that is used in the remaining nine steps of the biosynthesis of methanopterin (H 4 MPT) (Figure 1-7). The first of these steps, catalyzed by RFAP synthase, involves the condensation of pABA with PRPP, forming inorganic pyrophosphate, CO 2 and -ribofuranosylaminobenzene 5â€™-phophate (RFAP) (White 1996; Rasche and White 1998). RFAP is combined with 6-hydroxymethyl-7,8-dihydropterin pyrophosphate to produce 7,8-H 2 pterin-6-ylmethyl-4-(-D-ribofuranosyl)aminobenzene 5â€™-phosphate (White 1996; Xu et al. 1999). The product of that reaction is reduced and dephosphorylated to produce 7,8-H 2 pterin-6-ylmethyl-1-(4-aminophenyl)-1-deoxy-D-ribitol (White 1996; White 2001). The next three steps involve reaction of this intermediate with PRPP, activation by ATP, and condensation with hydroxyglutaric acid, resulting in the formation of demethylated dihydromethanopterin (White 1996; White 2001). The synthesis of demethylated dihydrosarcinapterin in M. thermophila occurs in the same manner only with the addition of another -linked glutamic acid residue (White 1996). The remaining steps involve methylation of the
22 HNNNHNH2NH2CHNOH2CCHOHCHOHCHOHCH2OH CO 2 PPi HNNNHNH2NH2CHNOH2CCHOHCHOHCHOHH2CCH2OOHOH2COPOO-O-OHPRPP+pAB OOHH2COPOO-O-H2N-RFAP Figure 1-7. Proposed biosynthetic pathway for methanopterin. (Adapted from White, R. H. 2001. Biosynthesis of the methanogenic cofactors. Vitam. Horm. 61:299337). OH HNNNHNH2NH2COPOOOO-POO-O-HydroxymethylpterinPPHNNNHNH2NH2CHNOH2CCHOHCHOHCHOHH2COPOOOHNNNHNH2NH2CONHOOHOHH2COPOO-O+ PPi 2H++2eH2OPi PRPP AMPATP HNNNHNH2NH2CHNOH2CCHOHCHOHCHOHH2CCH2OOHOH2COPOO-OOHPOO-OPOO-OHNNNHNH2NH2CHNOH2CCHOHCHOHCHOHH2CCH2OOHOH2COPOOHOOHCHCOOHCH2CH2COOH (S)-2-hydroxyglutaric acidPPi7,8-H2pterin-6-ylmethyl-4-(-D-ribofuranosyl)aminobenzene 5â€™-phosphate7,8-H2pterin-6-ylmethyl-1-(4-aminophenyl)-1-deoxy-D-ribitol 5â€™-phosphate7,8-H2pterin-6-ylmethyl-1-(4-aminophenyl)-1-deoxy-D-ribitol7,8-H2pterin-6-ylmethyl-1-(4-aminophenyl)-1-deoxy-5-[1--D-ribofuranosyl 5-phosphate]-D-ribitolDemethylated methanopterin HNNNHNH2NH2CHNOH2CCHOHCHOHCHOHCH2OH HNNNHNH2NH2CHNOH2CCHOHCHOHCHOHCH2OH2 CO PPi PPi HNNNHNH2NH2CHNOH2CCHOHCHOHCHOHH2CCH2OOHOH2COPOO-O-OHPRPP+pAB OOHH2COPOO-O-H2N-RFAP OH OOHH2COPOO-O-H2N-RFAP OH HNNNHNH2NH2COPOOOO-POO-O-HydroxymethylpterinPP HNNNHNH2NH2COPOOOO-POO-O-HydroxymethylpterinPPHNNNHNH2NH2CHNOH2CCHOHCHOHCHOHH2COPOOO-HNNNHNH2NH2CHNOH2CCHOHCHOHCHOHH2COPOOOHNNNHNH2NH2CONHOOHOHH2COPOO-O+ PPi 2H++2eH2OPi H2OPi PRPP AMPATP HNNNHNH2NH2CHNOH2CCHOHCHOHCHOHH2CCH2OOHOH2COPOO-OOHPOO-OPOO-OHNNNHNH2NH2CHNOH2CCHOHCHOHCHOHH2CCH2OOHOH2COPOO-OOHPOO-OPOO-OHNNNHNH2NH2CHNOH2CCHOHCHOHCHOHH2CCH2OOHOH2COPOOHOOHCHCOOHCH2CH2COOH HNNNHNH2NH2CHNOH2CCHOHCHOHCHOHH2CCH2OOHOH2COPOOHOOHCHCOOHCH2CH2COOH (S)-2-hydroxyglutaric acidPPi7,8-H2pterin-6-ylmethyl-4-(-D-ribofuranosyl)aminobenzene 5â€™-phosphate7,8-H2pterin-6-ylmethyl-1-(4-aminophenyl)-1-deoxy-D-ribitol 5â€™-phosphate7,8-H2pterin-6-ylmethyl-1-(4-aminophenyl)-1-deoxy-D-ribitol7,8-H2pterin-6-ylmethyl-1-(4-aminophenyl)-1-deoxy-5-[1--D-ribofuranosyl 5-phosphate]-D-ribitolDemethylated methanopterin
23 coenzyme at the C7 and C11 positions and finally its reduction from the dihydroform to the tetrahydro-form (White 1990; Caccamo, M., C. S. Malone, and M. E. Rasche, personal communication). Although the biosynthetic pathway for methanopterin is known and has been demonstrated to occur in M. thermophila, only five of the eighteen putative biosynthetic enzymes have been identified (White 1996; Xu et al. 1999; Graupner et al. 2000; White 2001; Graham et al. 2002; Scott and Rasche 2002; Caccamo M., C. S. Malone, and M. E. Rasche, personal communication). The first of these is the GTP cyclohyrolase III, encoded by the Methanococcus jannaschii gene MJ0798, which catalyzes the formation of triaminopyrimidine triphosphate from GTP (Graham et al. 2002). The gene product of MJ0301, which has high tertiary structure homology to dihydropteroate synthase from folate biosynthesis, has been shown to catalyze the condensation of the pterin ring with RFAP (Xu et al. 1999). Another of the enzymes, encoded by MJ1425, has been shown to catalyze the reduction of -ketoglutarate to (S)-2-hydroxyglutaric acid, which is a component of the side chain of methanopterin (Graupner et al. 2000). The last enzyme in the pathway has been identified as a dihydromethanopterin reductase, which catalyzes the reduction of dihydromethanopterin to tetrahydromethanopterin, and is encoded by the dmrA gene from the methylotrophic bacteria M. extorquens (Marx et al. 2003; Caccamo M., C. S. Malone, and M. E. Rasche, personal communication). Lastly the enzyme RFAP synthase, which catalyzes the condensation of pABA with PRPP to form RFAP, has been purified from the methanogen M. thermophila and identified in A. fulgidus (AF2089), as well as the methylotrophic bacteria M. extorquens (orf4) (Scott and Rasche 2002; Rasche et al. in press).
24 In contrast to the biosynthesis of tetrahydromethanopterin, the synthesis of tetrahydrofolate only involves three steps (Maden 2000). The first step in the pathway, catalyzed by the enzyme dihydropteroate synthase, involves the condensation of pterin pyrophosphate with pABA to form dihydropteroate (Baca et al. 2000). Dihydropteroate is combined with glutamic acid resulting in the formation of dihydrofolate, which is reduced to the physiologically active tetrahydro form. The biosynthetic pathway of folate is also known to be inhibited by the pABA analogs sulfonamides (Brown 1962). However, it has been shown that the sulfonamides do not inhibit methanopterin biosynthesis (White 1996; Rasche and White 1998). RFAP synthase The enzyme RFAP synthase has been shown to catalyze the transfer of the phosphoribosyl group of PRPP onto pABA to form RFAP, CO 2 , and inorganic pyrophosphate (White 1996; Rasche and White 1998; Scott and Rasche 2002). This enzyme is key in the biosynthetic pathway of methanopterin and is responsible for the removal of the carbonyl group of pABA in methanopterin (White 1996; Rasche and White 1998; Maden 2000). This carbonyl group is normally present in folate, its absence in methanopterin gives the coenzyme distinguishing structural and thermodynamic differences that set it apart from folate (White 1996; Rasche and White 1998; Maden 2000). RFAP synthase, because of the unusual nature of the chemistry it catalyzes, is proposed to be unique only to the synthesis of methanopterin and methanopterin-like derivatives (Scott and Rasche 2002; Bechard et al. 2003). The ability of RFAP synthase to catalyze the decarboxylation of a substrate during a phosphoribosyl transfer also makes this enzyme unusual among phosphoribosyltransferases. The only other enzyme known to catalyze chemistry of this nature is quinolinate phosphoribosyltransferase
25 (QAPRTase), which is responsible for the decarboxylation of quinolinic acid during the transfer of the phosporibosyl group of PRPP forming nicotinate mononucleotide, CO 2 , and pyrophosphate (Cao et al. 2002). RFAP synthase has been partially purified from M. thermophila and was shown to have optimal activity at a pH of 4.8 and a temperature of 50C (Rasche and White 1998). Biochemical characterization of the enzyme showed an apparent K m for pABA of 58M, and 3.6 mM for PRPP (Rasche and White 1998). Enzymatic mechanisms for the enzyme were proposed to occur in a sequential manner, one of two ways (Rasche and White 1998). The first of the proposed mechanisms involves a direct transfer mechanism where an enzyme-stabilized C-1 anionic intermediate of pABA serves as a nucleophile that undergoes an S N 1 reaction with PRPP displacing the pyrophosphate (Rasche and White 1998). The second of the proposed mechanisms involves a two-step S N 1-like reaction where the pyrophosphate is separated from PRPP forming a carboxonium ion at the C-1 position of PRPP, a reaction known to also occur in type I PRTases (Tao et al. 1996; Rasche and White 1998). This carboxonium ion intermediate of PRPP, thought to be stabilized by carboxylate residues, is thought to participate in an electrophilic aromatic substitution at the C-1 position of pABA resulting in its decarboxylation and formation of RFAP (Rasche and White 1998). RFAP synthase has been purified from M. thermophila to homogeneity. The N-terminal sequence was determined and used to identify more then 16 homologs of the enzyme in both the Archaeal and Bacterial domains (Scott and Rasche 2002). Two of these homologs, orf4 from M. extorquens and AF2089 from A. fulgidus, have since been shown to encode active RFAP synthases (Scott and Rasche 2002; Rasche et al. in press).
26 There are two evolutionarily distinct types of phosphoribosyltransferases (PRTases) that have distinctly different active site architecture. The type I PRTases consist of hypoxanthine guanine phosphoribosyltransferase (HGPRTases), orotate phosphoribosyltransferase (OPRTases), glutamine PRPP amidotransferase, uracil phosphoribosyltransferase (UPRTase), as well as PRPP synthase (Eads et al. 1994; Scapin et al. 1995; Schumacher et al. 1998; Smith 1999; Eriksen et al. 2000). The active site of these enzymes is made up of an / core structure with a characteristic Rossmann fold (Eads et al. 1994; Smith 1999; Cao et al. 2002). This solvent exposed active site is flanked by loop and hood structures which close over the active site during catalysis (Eads et al. 1994; Smith 1999; Cao et al. 2002). The type II PRTases are solely represented by a single enzyme QAPRTase and consist of a seven-stranded / barrel structure that contains the active site and serves as a cap for the / barrel structure of the other subunit (Cao et al. 2002). This enzyme was labeled a type II PRTase not only because of its different structure but also because the enzyme catalyzes the decarboxylation of its substrate during a phosphoribosyltransfer event, chemistry not seen in the type I PRTases (Cao et al. 2002). When the known properties of RFAP synthase are compared to the properties of the two types of PRTases it can be noted that the enzyme has characteristics of both types. The chemistry of RFAP synthase is similar to that of the type II PRTase; however, one of the proposed mechanisms for RFAP synthase has similarities to that of the type I PRTases (White 1996; Rasche and White 1998; Scott and Rasche 2002). The lack of a crystal structure, as well as further biochemical knowledge of RFAP synthase prevents any definitive categorization of the enzyme as either a type I or a type II enzyme.
27 RFAP synthase catalyzes a chemical reaction that gives methanopterin key distinguishing features. The chemistry catalyzed by RFAP synthase not only makes the enzyme important for the biosynthesis of methanopterin, but it also makes it specific for the biosynthesis of methanopterin and its derivatives. If the biosynthesis of methanopterin should in some way be disrupted or inhibited it would result in the inhibition of methanogenesis, because of the importance of methanopterin in the methanogenic process. The unique chemistry and specificity that RFAP synthase has for the synthesis of methanopterin makes this enzyme a good target for the development of an inhibitor. This inhibitor could serve as an antibacterial agent specific for methanogenic organisms, in much the same way that sulfonamides are for bacteria (Dumitru et al. in press). This methanogen â€œantibioticâ€ could have useful applications in reducing the generation of methane by methanogenic archaea present in the ruminants of animals (Dumitru et al. in press). Inhibitors of this nature are currently being developed and have in fact been shown to inhibit RFAP synthase (Dumitru et al. in press). These inhibitors have also been shown to halt the methanogenic process, both in pure methanogen culture, as well as artificial rumen cultures, while having no detrimental affect on the growth of beneficial organisms like acetogens (Dumitru et al. in press). In order to contribute to this overall goal more information is needed on the overall catalytic properties and structure of a methanogen RFAP synthase. This enzyme has been purified from the methanogen M. thermophila, however, because of the low abundance of the enzyme further biochemical characterization was not feasible. It is the focus of this study to gain a better understanding of methanogen RFAP synthases. This will provide important info about the enzyme to be used in the development of more
28 potent inhibitors of methanogenesis. The current work, therefore, describes the identification, overproduction, and purification of a methanogen RFAP synthase from an E. coli overproduction system. Bioinformatics and site-directed mutagenesis was used to gain a better understanding of not only the structure of the active site, but binding sites of the substrates, as well as amino acids involved in binding of one of the substrates pABA.
CHAPTER 2 OVEREXPRESSION, PURIFICATION, CHARACTERIZATION, AND SITE-DIRECTED MUTAGENESIS OF A METHANOGEN RFAP SYNTHASE PRODUCED IN ESCHERICHIA COLI Introduction The coenzyme methanopterin and its derivatives have been found among the third domain of life, the Archaea in both methanogenic and non-methanogenic organisms, where they replace folate as the primary one-carbon carrier (Leigh 1983; Vanbeelen et al. 1984; Mller-Zinkhan et al. 1989; Zhou and White 1992; White 1993; Maden 2000). In methanogenic archaea, methanopterin acts as the primary one-carbon carrier as these one-carbon compounds are reduced to methane (DiMarco et al. 1990; Thauer et al. 1993; White 2000). However, this coenzyme has recently been identified in the bacterial domain among the methylotrophs, where it coexists with folate (Chistoserdova et al. 1998). In these methylotrophic bacteria, methanopterin is utilized to carry one-carbon compounds as they are oxidized to and released as CO 2 (Vorholt et al. 1998; Chistoserdova et al. 2003). Hydrogenotrophic methanogens are of great interest because of their ability to produce methane gas through the hydrogen-dependent reduction of CO 2 (Thauer et al. 1993; Deppenmeier et al. 1996). Methanogens inhabit many diverse anaerobic environments where they release methane gas, a potent greenhouse gas, into the atmosphere contributing to the growing problem of global warming (Zeikus 1977; Keltjens and Vogels 1988; Drake 2000). The generation of methane gas as a result of the methanogenic process is the only means by which methanogenic archaea produce ATP, 29
30 making the process vital to their survival (Becher et al. 1992; Becher and Muller 1994). Methanopterin is also known to be essential in four of the steps of methanogenesis, making the coenzyme not only essential to the production of methane but also essential for the survival of the methanogenic organisms (Donnelly et al. 1985; Donnelly and Wolfe 1986; Te Brommelstroet et al. 1990; Schworer and Thauer 1991). Therefore, the characterization of enzymes involved in the biosynthesis of the essential coenzyme could lead to valuable information that would be key in reducing the problem of methane emitted from methanogenic archaea. In the first step of methanopterin biosynthesis, the enzyme RFAP synthase catalyzes the transfer of the phosphoribosyl group from phosphoribosylpyrophosphate (PRPP) onto p-aminobenzoic acid (pABA) to form -ribofuranosylaminobenzene 5â€™-phosphate (RFAP), CO 2 , and inorganic pyrophosphate. This is an interesting enzyme because of its ability to decarboxylate a substrate during a phosphoribosyl transfer event in the overall formation of C-riboside product rather than an N-riboside, properties that are only found in one other PRTase enzyme, quinolinate phosphoribosyltransferase (QAPRTase) (Rasche and White 1998; Cao et al. 2002; Scott and Rasche 2002). The decarboxylation of pABA in the reaction catalyzed by RFAP synthase gives methanopterin a distinguishing chemical difference that sets this coenzyme apart from folate (White 1996). Due to these properties it has been proposed that organisms which have an active RFAP synthase also contain methanopterin or a derivative of it (Rasche and White 1998; Scott and Rasche 2002; Bechard et al. 2003). Previously, RFAP synthase was purified from Methanosarcina thermophila, and the N-terminal sequence used to identify homologs in methylotrophic bacteria and
31 archaea (Scott and Rasche 2002). The RFAP synthase homolog from M. thermautotrophicus was recombinantly produced in E. coli, identifying the gene MTH0830 as encoding RFAP synthase (Bechard et al. 2003). However, a purified recombinantly produced methanogen RFAP synthase has not been biochemically analyzed. In the current work we have purified and biochemically characterized the recombinantly produced the M. thermautotrophicus RFAP synthase and subsequently generated and characterized site-directed mutants of the enzyme to provide further information on amino acids that play an important role in the overall mechanism and activity of the enzyme. These studies provide a better understanding of how a methanogen RFAP synthase catalyzes one of the most important biosynthetic steps in the synthesis of the essential coenzyme, methanopterin. Materials and Methods Colorimetric Assay for RFAP Synthase The RFAP synthase assay (White 1996) is based on the conversion of the arylamine product, RFAP, to its colored azo-dye derivative using nitrite and N-naphthylethylene diamine (Aldrich Chemical Co., Inc., Milwaukee, WI). Because these reagents also react with the arylamine substrate p-aminobenzoic acid (pAB) (Aldrich Chemical), RFAP must first be separated from pAB using a C18 column (White 1996; Rasche and White 1998). For full detailed protocol of the assay used in the current work see Bechard et al. (2003). Heterologous Expression of MTH830 in E. coli The polymerase chain reaction (PCR) was used to amplify the RFAP synthase homolog MTH0830 from the M. thermautotrophicus genome using established protocols with Pfu DNA polymerase (Sambrook and Russell 2001). The primer sequence was
32 synthesized commercially (Sigma Genosys, St. Louis, MO) and designed to engineer in an NdeI site 5â€™ and a BamHI site 3â€™ of the gene. The PCR product and the vector pET41a(+) (Novagen, Inc., Madison, WI) were digested with restriction enzymes overnight at 37C, and the two pieces were ligated yielding the plasmid designated pED2. pED2 was transformed into electrocompetent E. coli DH5-alpha cells, and the gene sequence was verified by dideoxy sequencing (Sambrook and Russell 2001). When MTH0830 was expressed in E. coli at 37C, most of the protein aggregated as inactive inclusion bodies (data not shown). Therefore, to increase the chances of producing soluble, active M. thermautotrophicus RFAP synthase, different induction temperatures and times were tested. In addition, the MTH0830 gene was coexpressed with the genes for the chaperone GroEL/ES and trigger factor provided on plasmid pG-Tf2 (Nishihara et al. 2000). For recombinant production of RFAP synthase (MTH0830), pED2 was transformed into E. coli BL21(DE3) cells containing pG-Tf2 (kindly provided by Dr. Tsunetaka Ohta). The expression cell line with pED2 and pG-Tf2 was called KB1.To test the effect of variable induction times and temperatures in the absence of chaperone, KB1 cells were grown in 1 liter of Luria-Bertani medium with kanamycin (50 g/mL) and chloramphenicol (17 g/mL) at 37C to an optical density at 600 nm of 0.6 to 0.8. MTH0830 expression was induced with IPTG at 1 mM, and the cultures were incubated at 37C for 2 h, 30C for 6 h, or 20C for 16 h. After the appropriate incubation times, cells were harvested by centrifugation. For expression in the presence of the chaperone, KB1 cells were grown in the same media to an optical density of 0.4, and tetracycline was added (final concentration, 50 ng per mL) for induction of the chaperone (Nishihara et al. 2000). After a 30-min incubation, expression of MTH0830
33 was induced with IPTG. The cultures were then transferred to a 20C incubator, and cells were grown for 16 h before centrifugation. After centrifugation cells were then disrupted by passage through French pressure cell. The cell lysate was then centrifuged at 31,000 x g for 1 h to separate the cell free extract (CFE, supernatant) from the insoluble fraction (pellet). Purification of Recombinant RFAP Synthase All forthcoming purification steps were performed at ambient temperature in the presence of 2 mM DTT. CFE (50 mL) was heat-treated at 65C for 15 min, and the heated sample was centrifuged at 13,000 x g for 15 min to remove precipitated proteins. Heat-treated CFE (25 mL) was loaded onto a 40-mL ceramic hydroxyapatite column (11cm x 2.5 cm) equilibrated with 50 mM TES [N-tris(hydroxymethyl)methyl-2-aminoethanesulfonic acid], 2 mM DTT (dithiothreitol) pH 6.8, and the column washed with 80 mL of the same equilibration buffer. The proteins were eluted using a 400-mL linear gradient from 0 mM NaH 2 PO 4 to 125 mM NaH 2 PO 4 in equilibration buffer at pH 6.8 collecting 20 mL fractions. Active RFAP synthase was found in fractions eluting between 60 and 80 mM NaH 2 PO 4 . The most active hydroxyapatite fractions were loaded onto a Mono Q HR 5/5 anion exchange column (Amersham-Pharmacia Biotech-Piscataway, NJ). The column was equilibrated with 50 mM TES, 5 mM MgCl 2 , 2 mM DTT pH 6.8, and washed with 300 mM NaCl in the equilibration buffer, followed by a 20-mL linear gradient from 300 mM to 600 mM NaCl in equilibration buffer. Active RFAP synthase was only found in fractions eluting between 375 and 390 mM NaCl. These active Mono Q fractions were then combined, concentrated and loaded onto
34 a Superdex 75 gel filtration column equilibrated with 50 mM TES, 150 mM KCl, and 2 mM DTT. The active pure RFAP synthase enzyme eluted after 19 min corresponding to a molecular mass of 68.7 kDa. Determination of Optimal Enzymatic Conditions To determine the temperature optimum for RFAP synthase, the velocity of hydroxyapatite purified enzyme was measured using 35 mM pABA, 8.8 mM PRPP and 30 mM MgCl 2 in 50 mM MES 150 mM NaCl (pH 6.8) buffer, at temperatures ranging from 37C to 80C. To determine the pH optimum, the velocity of hydroxyapatite purified enzyme was measured using 35 mM pABA, 8.8 mM PRPP and 30 mM MgCl 2 over a range of pH values from 4.5 to 8.0, in 50 mM acetate (pH 4.5 to 5.5), 50 mM PIPES (pH 6.5 to 7.0), and 50 mM bicine (pH 7.5 to 8.0). The ionic strength of the buffers used to determine the pH and temperature optima was approximately 150 mM, using NaCl as the primary salt. To determine optimal ionic strength needed for maximal activity of RFAP synthase the velocity of hydroxyapatite purified enzyme was determined using 35 mM pABA, 8.8 mM PRPP, and 30 mM MgCl 2 at ionic strengths ranging from 50 mM to 500 mM, using NaCl as the primary salt. To determine the optimal MgCl 2 concentration needed for maximal activity of RFAP synthase the velocity of hydroxyapatite purified enzyme was determined using 35 mM pABA, 8.8 mM PRPP, at MgCl 2 concentrations ranging from 5 mM to 50 mM. The optimal MgCl 2 for RFAP synthase was also determined under the same conditions using 17.6 mM PRPP. To examine the effect of enzyme concentration on activity the velocity of Mono Q purified enzyme (approximately 95% pure) was determined under optimal conditions using 35 mM pABA and 8.8 mM PRPP with various amounts of enzyme ranging from
35 0.012 nmol to 0.200 nmol (1 to 25 g). From this point on the amount of enzyme used in activity assays is within this linear range, approximately 8 to 12 g. Kinetic Analyses of the Recombinant Methanogen RFAP Synthase For determination of the K m for pABA, the concentration of PRPP was kept constant at 8.8 mM with 17.6 mM MgCl 2 , while the pABA concentration was varied from 181 M to 35 mM. All assays were done under optimal enzymatic conditions in 70 mM PIPES (pH 6.8), 10 mM MgCl 2 , 2 mM DTT buffer at 70C for 1 h. Partial Purification and Analyses of M. thermautotrophicus RFAP Synthase For preparation of the cell extract approximately 5 g of M. thermautotrophicus cells were resuspended in 10 ml of 50 mM TES (pH 6.8) buffer supplemented with 2 mM DTT containing DNAase I, disrupted by passage through a French pressure cell, and centrifuged at 31,000 x g for 1 h resulting in 9 mL of CFE (supernatant) at a protein concentration of approximately 17 mg/mL. The CFE was loaded onto a 10-ml ceramic hydroxyapatite column equilibrated with 50 mM TES (pH 6.8) buffer supplemented with 2 mM DTT and washed with 20 ml of the same equilibration buffer. A 100 ml linear gradient was applied to the column from 0 mM to 400 mM NaH 2 PO 4 in equilibration buffer at pH 6.8 collecting 5 ml fractions. Active RFAP synthase was found in fractions eluting between 150 mM and 200 mM NaH 2 PO 4 . To determine the apparent K m for pABA for the M. thermautotrophicus RFAP synthase, the specific activity of the most active hydroxyapatite fraction was measured at 70C for 1 h at a constant PRPP concentration of 8.8 mM with 10 mM MgCl 2 while varying the pABA concentration from 0.363 mM to 35 mM.
36 Partial Purification and Analyses of the M. thermophila RFAP Synthase The cell extract was prepared from 5 g of M. thermophila TM-1 cells as described for M. thermautotrophicus. The CFE was loaded onto a 75 ml Q-Sepharose column equilibrated with 50 mM TES (pH 6.8), 5 mM MgCl 2 , 2 mM DTT buffer and washed with two column volumes of the equilibration buffer. A 400-mL linear gradient from 0 mM to 600 mM NaCl in equilibration buffer at pH 6.8 was applied to the column, and 20-ml fractions were collected. Using conditions previously described by Rasche and White (1998), the apparent K m for pABA was determined using a constant PRPP concentration of 8.8 mM with 10 mM MgCl 2 , while varying the pABA concentration from 11.3 M to 1.45 mM. Site-directed Mutagenesis of Recombinant RFAP Synthase Residues R26 and D19 of the M. thermautotrophicus RFAP synthase were mutated in a non-conservative and conservative manner by site-directed mutagenesis. Plasmid pED2, which contains the RFAP synthase gene (MTH0830), was used as the template for site-directed mutagenesis (Bechard et al. 2003). The plasmids were purified using the Spin MiniPrep plasmid isolation kit from Qiagen (Valencia, CA). Site-directed mutagenesis was performed using the QuickChange XL kit (Stratagene, La Jolla, CA) with the following thermocycling conditions: denaturation at 95C for 30 s, primer annealing at 55C for 60 s, and extension for 12 min at 68C through 16 cycles. The forward primer sequences containing the codon mutations used to create the site-directed mutants studied in these experiments are depicted in Table 2-1. Dideoxy sequencing was used to verify the mutations (Sambrook and Russell 2001). For recombinant production of the RFAP synthase variants altered by site-directed mutagenesis, the plasmid with the mutated gene was transformed as previously
37 Table 2-1. Primers used for site-directed mutagenesis Variant protein Forward sequence of the mutagenic primer (altered codon is underlined) R26A 5â€™CTCAACGGTGAGAGGGGC GCA CTTGACGGTGGAGTTGG R26K 5â€™CTCAACGGTGAGAGGGGC AAG CTTGACGGTGGAGTTGG D19A 5â€™CCACCTGACCCTCATA GCG CTCAACGGTGAGAGGG D19N 5â€™CCACCTGACCCTCATA AAC CTCAACGGTGAGAGGG done for the wild-type in BL21(DE3) with the chaperone expressing plasmid pG-Tf2. Cells were grown and the gene expressed as done for the recombinant wild-type. After harvesting and disruption of the cells by French pressure cell lysis the solubility of the variants were assessed by SDS-PAGE analysis of the soluble and insoluble fractions. Partial Purification of RFAP Synthase Variants Variants were partially purified from cell extract (approximately 500 mg) by hydroxyapatite chromatography as described for the recombinant wild-type. Active RFAP synthase variants were found in fractions eluting between 80 mM NaH 2 PO 4 and 100 mM NaH 2 PO 4 . The most active hydroxyapatite fractions were loaded onto a Mono Q HR 5/5 anion exchange column (Amersham-Pharmacia Biotech-Piscataway, NJ). The column was equilibrated with 50 mM TES, 5 mM MgCl 2 , 2 mM DTT pH 6.8, and washed with 130 mM NaCl in the equilibration buffer, followed by a 30-mL linear gradient from 130 mM NaCl to 330 mM NaCl. Active RFAP synthase variants were found in fractions eluting between 120 mM and 130 mM NaCl. For comparison with the variants altered by site-directed mutagenesis, the recombinant wild type enzyme was partially purified in the exact same manner as done
38 Kinetic Analyses of Recombinant RFAP Synthase Variants To determine the K m for pABA for the RFAP synthase variants the specific activity of the Mono Q purified variants were determined in the presence of 8.8 mM PRPP and 16 mM MgCl 2 , with a range of pABA concentrations from 181M to 67 mM for the D19N variant, and 3.8 mM to 130 mM for the R26K variant. Due to the inability of the variants to function under the same optimal conditions as the recombinant wild-type, the assays were done under optimal conditions, for these enzymes, in 70 mM PIPES (pH 6.8), 10 mM MgCl 2 , 2 mM DTT buffer at 50C for three hours. The kinetic properties of the wild-type enzyme, partially purified in the same manner as the variants were also examined at 50C for three hours. Protein Quantitation and Gel Electrophoresis The purity of the proteins was determined using sodium dodecyl sulfate polyacrylamide gel electrophoresis (Garfin 1990) with Coomaise Brilliant Blue R-250 as the stain (Biorad). Protein concentrations were determined using the Bradford protein assay (Biorad) (Bradford 1976) with bovine serum albumin as the standard. Materials and Chemicals Naphthylethylene diamine and p-aminobenzoic acid were purchased from Aldrich Chemical Co., Inc. (Milwaukee, WI). PRPP was from Sigma Chemical Corp. (St Louis, MO), and gases were obtained from Strate Welding (Gainesville, FL). All other chemical were of reagent grade and purchased from Fisher Scientific (Suwanee, Ga).Pfu DNA polymerase was purchased from Stratagene. Restriction enzymes and T4 DNA ligase were from New England Biolabs (Beverly, MA).
39 Results Overexpression of a Methanogen RFAP Synthase in E. coli. BLAST searches indicate that the gene MTH0830 from the methanogen M. thermautotrophicus is an RFAP synthase homolog (Rasche and White 1998). Initially, we attempted to express the gene by transforming E. coli BL21(DE3) cells with a plasmid (pED2) containing MTH0830. However, this procedure resulted in a complete lack of RFAP synthase activity even when the induction temperature was lowered to 30C (data not shown). This led us to create an expression strain (KB1) containing both pED2 and the plasmid pG-Tf2, which carries the genes for the GroEL/ES chaperone under the control of a tetracycline promoter (Nishihara et al. 2000). Figure 2-1 illustrates the effects of induction temperature and the presence of chaperone on the production of soluble versus insoluble M. thermautotrophicus RFAP synthase. In these experiments, KB1 cells were grown at 37C, and then MTH0830 expression was induced with IPTG at the indicated temperatures and times. when MTH0830 was induced and incubated at 37C for 2 h, in the absence of the chaperone, SDS-PAGE analysis showed that a large quantity of the overproduced protein was present in the insoluble fraction (Figure 2-1, lanes 1 and 2). The resulting soluble fraction had only a trace of RFAP synthase activity (0.0056 nmol of RFAP produced per min per mg of protein, at pH 7.0 and 70C), possibly due to leaky expression of the chaperone from the pG-Tf2 plasmid. When the induction temperature was lowered to 30C and the induction time increased to 6 h, the specific activity of the soluble fraction (0.033 nmol of product per min per mg of protein) increased 6-fold when compared to the enzyme produced at 37C, and a corresponding increase in soluble protein was observed (Figure 2-1, lanes 3 and 4). The specific activity was further increased when the induction temperature was lowered to 20C for 16 h
40 (0.074 nmol of product per min per mg of protein) and was accompanied by a corresponding increase in soluble enzyme (Figure 2-1, lanes 5 and 6). To investigate whether the induction of the chaperone would further aid in producing soluble, active enzyme, KB1 cells were grown at 37C, and chaperone gene expression was induced with tetracycline, prior to the induction of MTH0830. Induction of the chaperone, in conjunction with MTH0830 expression at 20C for 16 h, resulted in the highest production of soluble, active enzyme. The specific activity (0.100 nmol of product per min per mg of protein) was approximately 20-fold higher than the activity of enzyme produced at 37C, with a significant increase in soluble protein and a marked decrease in protein found in the insoluble fraction (Figure 2-1, lanes 7 and 8). The results presented here show that lower induction temperatures, longer induction times, and the use of a chaperone all aided in the successful production of active RFAP synthase from M. thermautotrophicus. Purification of a Methanogen RFAP Synthase Produced in E. coli . The successful development of a recombinant production system for this methanogen RFAP synthase in E. coli allowed purification of the recombinant form of the enzyme for further biochemical analysis. RFAP synthase was purified initially with a heat-step at 65C for 15 min. We were able to do this because of the thermophilic nature of the enzyme and its native organism (Zeikus 1972). Following heat treatment and removal of the precipitated proteins, RFAP synthase was purified by hydroxyapatite, Mono Q, and Superdex 75 column chromatography. Previously, it was estimated that the overproduced enzyme has a molecular mass of 38 kDa by SDS-PAGE analysis (Bechard et al. 2003). The gel filtration analysis of the enzyme gave a molecular mass of 68.7 kDa suggesting that the enzyme is a homodimer
41 1 2 3 5 6 7 8 4 97.4 kDa 66.2 kDa 45 kDa 31 kDa 21.5 kDa 14 kDa Figure 2-1. Soluble versus insoluble fractions of RFAP synthase expression in KB1 cells. KB1 cells were grown at 37C and then induced with IPTG at the indicated temperatures. Cells were disrupted with a French press and centrifuged, producing a soluble fraction (supernatant) and insoluble fraction (pellet). Induction at 37C for 2 h, soluble (lane 1) and insoluble (lane 2); induction at 30C for 6 h, soluble (lane 3) and insoluble (lane 4); induction at 20C for 16 h, soluble (lane 5) and insoluble (lane 6). Induction of chaperone, followed by induction of MTH0830 and incubation at 20C for 16 h, soluble (lane 7) and insoluble (lane 8). Each lane contained between 15 and 30 g of protein. The arrow indicates the position of the recombinant RFAP synthase from M. thermautotrophicus.
42 1 2 3 4 5 6 Figure 2-2. Purification of recombinant RFAP synthase. Protein samples were boiled in the presence of 7.5% 2-mercaptoethanol in SDS-PAGE sample buffer and loaded onto a 12% polyacrylamide gel. The gel was stained with Coomassie Brilliant Blue R-250 (BioRad). Lane 1, Molecular mass markers. Lane 2, CFE (8 g). Lane 3, heated CFE (10.2 g). Lane 4, hydroxyapatite fraction (17 g). Lane 5, Mono Q fraction (2 g). Lane 6 Superdex 75 fraction (2 g). Arrows indicate the positions of the following molecular mass markers: phosphorylase b (97.4 kDa), bovine serum albumin (66.2 kDa), ovalbumin (45 kDa), carbonic anhydrase (31 kDa), soybean trypsin inhibitor (21.5 kDa), and lysozyme (14.4 kDa).
43 Table 2-2. Purification of RFAP synthase produced in E. coli . Purification Total activity 1 Specific activity Yield Fold step (nmol per min) (nmol per min per mg protein) (%) purification Cell-free extract 690 0.13 100 1 Heat-treated 390 0.79 57 6 Hydroxyapatite 260 14 38 110 Mono Q 110 290 16 2200 Superdex 75 30 360 4.2 2800 1 RFAP synthase assays were performed at 70C at pH 6.8 for one hour as described in the Material and Methods.
44 A. 00.050.10.150.18.104.22.1680.440455055606570758085Temperature (C) Velocity (nmol/min ) B. 00.050.10.150.22.214.171.1240.40.4544.555.566.577.588.5pHVelocity (nmol/min ) Figure 2-3. Optimal temperature and pH of RFAP synthase. A. The activity of hydroxyapatite purified RFAP synthase was assayed at various temperatures for one hour at pH 6.8 B. The activity of hydroxyapatite purified RFAP synthase was assayed at 70C for one hour, varying the pH from 4.5 to 8.0 as described in the Methods and Material section.
45 The enzyme was purified 2800-fold to homogeneity with a specific activity of 365 nmol/min/mg of enzyme (Figure 2-2 and Table 2-2). A 2800-fold purification of a recombinantly produced enzyme seems to be unusually high and might be due to the presence of a chaperone in the overexpression system. The interaction between the chaperone and the soluble enzyme may have affected the specific activity of the enzyme detrimentally, causing the unusually large increase in both the specific activity and fold purification of the enzyme when the chaperone was purified away from RFAP synthase at the Mono Q step (Figure 2-2 and Table 2-2). Determination of Optimal Enzymatic Conditions . To determine the optimal enzymatic conditions for the recombinant methanogen RFAP synthase, the activity of hydroxyapatite purified RFAP synthase was tested at different temperatures, with optimal activity found at 70C (Figure 2-3A). The activity of hydroxyapatite purified enzyme was also tested at pH values ranging from 4.5 to 8.0, revealing optimal activity at a pH of 7.0 (Figure 2-3B). The time course of the activity for the enzyme revealed a constant rate of activity up to 90 min (data not shown). The optimal MgCl 2 concentration needed for maximal activity was determined to be approximately 1.5 to 3 fold higher then the PRPP concentration used (data not shown). The activity of hydroxyapatite purified RFAP synthase remained constant up to an ionic strength of 200 mM (data not shown). From this point on, activity of RFAP synthase was measured using these optimal conditions. Under optimal conditions the effect of increasing the amount of enzyme used in the assay from 0.012 nmol to 0.200 nmol (1 to 25 g) on the velocity of the reaction was determined. A non-linear concave relationship was observed between the amount of
46 A. 00.20.40.60.8126.96.36.199.800.050.10.150.20.25Velocity (nmol RFAP produced/min)Amount of enzyme (nmol) 00.20.40.60.8188.8.131.52.800.050.10.150.20.25Velocity (nmol RFAP produced/min)Amount of enzyme (nmol) B. 010203040506070809010000.050.10.150.184.108.40.2060.4Specific Activity (nmol/min/mg)Amount of enzyme (nmol) 010203040506070809010000.050.10.150.220.127.116.110.4Specific Activity (nmol/min/mg)Amount of enzyme (nmol) Figure 2-4. Effect of enzyme concentration on velocity of reaction. A. The velocity of Mono Q purified RFAP synthase was measured using 35 mM pABA, 10 mM MgCl 2 , and 8.8 mM PRPP, as described in the Methods and Material section, against various amounts of enzyme. B. The specific activity of Mono Q purified enzyme was measure against various enzyme concentrations.
47 0 10 20 30 40 0 100 200[pABA] mMSpecific Activity(nmol/min/mg) Lineweaver-BurkeR2 = 0.9996-0.0500.050.10.15-101231/[pABA] mM1/Specific activity (nmol/min/mg) -1 0 10 20 30 40 0 100 200[pABA] mMSpecific Activity(nmol/min/mg) Lineweaver-BurkeR2 = 0.9996-0.0500.050.10.15-101231/[pABA] mM1/Specific activity (nmol/min/mg) -1 Figure 2-5. Determination of pABA K m for recombinant RFAP synthase . RFAP synthase activity was measured at a constant PRPP concentration of 8.8mM while varying the pAB concentration from 181M to 36 mM. The inset picture shows the Lineweaver-Burke plot from the Michaelis-Menton plot. 0 10 20 30 40 0.0 0.1 0.2[pABA] mM y Figure 2-6. Determination of pABA K m for native RFAP synthase . Cell extract from M. thermautotrophicus cells was partially purified by hydroxyapatite chromatography. Activity was measured using the active RFAP synthase fractions as described previously at a PRPP concentration of 8.8mM while varying the pABA concentration from 181 M to 36 mM. Sp eci fi c Ac tiv it (nmol/min/mg) Lineweaver-BurkeR2 = 0.9771-20-10010203040506070-2-101231/[pAB] mM1/Specific Activity (nmol/min/mg) -1 0 10 20 30 40 0.0 0.1 0.2[pABA] mM(nmol/min/mg) y it tiv Sp eci fi c Ac Lineweaver-BurkeR2 = 0.9771-20-10010203040506070-2-101231/[pAB] mM1/Specific Activity (nmol/min/mg) -1
48 enzyme used and the velocity of the reaction until about 0.1 nmol (6.5 g) of enzyme, at which point the relationship became linear (Figure 2-4A). When the specific activity of the enzyme is measured against variable amounts of enzyme, an increase in specific activity is seen until approximately 0.1 nmol (8.6 g) of enzyme at which point the activity levels off (Figure 2-4B). These data not only show that the linear range of activity is between 0.1 and 0.2 nmol of enzyme, but along with the molecular mass determination from the also suggests a need for the enzyme to be in the dimer form to obtain proper activity. Kinetic Analyses of Recombinant RFAP Synthase With the purification of this enzyme from the recently developed E. coli expression system, we were able to analyze the K m for the substrate pABA. When the specific activity of the Mono Q purified enzyme was measured at 8.8 mM PRPP and 17.6 mM MgCl 2 , the K m for pABA appears to be approximately 4.5 mM (Figure 2-5). When the K m was measured with the homogenous gel filtration purified enzyme a similar K m (approximately 1-2 fold higher) was observed (data not shown). During the course of the purification and biochemical analysis of the recombinant enzyme, it was observed that purification of the enzyme beyond the Mono Q step resulted in loss of enzyme stability, possibly having an effect on the catalytic competency of the enzyme. The K m for pABA determined here is significantly higher than the apparent K m for pABA determined for the M. thermophila enzyme (58 M) (Rasche and White 1998). However, when RFAP synthase was partially purified from M. thermautotrophicus delta H cells and the apparent K m measure, a value of 3.9 mM was observed (Figure 2-6), similar to the K m for the recombinant M. thermautotrophicus enzyme. To verify the K m for RFAP synthase from M. thermophila, the enzyme was partially purified and the apparent K m determined.
49 0.0 2.5 5.0 7.5 10.0 12.5 0.0 2.5 5.0 7.5[pABA] mM Figure 2-7. Determination of pABA K m for M. thermophila RFAP synthase . Cell extract from M. thermophila cells was partially purified by Q-sepharose chromatography. Activity was measured using the active RFAP synthase fractions as described in methods and materials section at a PRPP concentration of 8.8mM while varying the pABA concentration from 11.3 M to 1.45 mM. Sp efi Av(nmol/min/mg) ity ci c cti Lineweaver-Burke R2 = 0.953-0.200.20.40.6-60-40-2002040601/pABA mM 1/Specific Activity(nmol/min/mg) -1 0.0 2.5 5.0 7.5 10.0 12.5 0.0 2.5 5.0 7.5[pABA] mMefi Av(nmol/min/mg) ity cti Lineweaver-Burke R2 = 0.953-0.200.20.40.6-60-40-2002040601/pABA mM 1/Specific Activity(nmol/min/mg) -1 c ci Sp
50 Group 1Group 2Group 3 Group 1Group 2Group 3 Figure 2-8. Phylogenetic relationship of RFAP synthases homologs. The amino acid sequences of putative RFAP synthases were aligned with ClustalX (Thompson, J.D., 1997), and then TREEVIEW (Page, R.D.M., 1996) was used to generate the unrooted tree. The gene abbreviations are as follows: Methanosarcina mazei Mm0906 and Mm0322, Methanosarcina acetivorans MA4006 and MA0339, Methanosarcina barkeri MB2615, Methanococcus jannaschii MJ1427, Archaeoglobus fulgidus AF2089, Methanothermobacter thermautotrophicus MTH0830, Methanopyrus kandleri MK0558, Sulfolobus sulfataricus SSo0370, Sulfolobus tokodaii ST1329, Aeropyrum pernix APE1512 and APE2425, Pyrococcus horokoshii PH1228 and PH0227, Pyrococcus furiosus PF0903, Pyrococcus abysii PAB1694 and PAB0441, Methylobacterium extorquens orf4.
51 A. B. MM1592 --------------------------------------------------------MRIQMNMINVVSPSRLHLTLIDLNAEIGR MM0322 --------------------------------------------------------------MINVVSPSRLHLTLIDLNAEIGR MA0339 --------------------------------------------------------MRIQMNMINVVSPSRLHLTLIDLNAEIGR MB2615 ------------------------------------------------------MQFWSLTGMIKITTPCRIHMTLIDMNAEIGR MA4006 -------------------------------------------------------MSRGCPRMIKITTPSRLHITLIDMNGEIGR MK0558 --------------------------------------------------------------MVRVRSVSRIHVTLIDLHGGLGR MJ1427 ---------------------------------------------------------------MIIQTPSRIHMGLIDLNGSIGR AF2089 --------------------------------------------------------------MLRLRTPSRIHITLIDLNGSIGR MTH830 -----------------------------------------------------------MVIELIINTPSRLHLTLIDLNGERGR ST1389 -----------------------------------------------------------MFCMIKIIGLSRIHITLIDLEGKFGR SSO0370 --------------------------------------------------------------MIKIVGLSRIHITLFDLEGKYGR MM1592 VDGGVGITLESPSLEISAS-EADTVEVVGSSLLAG------------RMLKAAKAVIPEG---KGIKIHIKRDLPDHVGLGSGTQ MM0322 VDGGVGITLESPSLEISAS-EADTVEVVGSSLLAG------------RMLKAAKAVIPEG---KGIKIHIKRDLPDHVGLGSGTQ MA0339 VDGGVGITLESPGLEISAT-EADAVEVVGDSLLAG------------RMLKAAKAVLPAG---KGIRIHIKRDLSDHVGLGSGTQ MB2615 VDGGAGLTLSSPNIKITAE-EADGVNIEGRQGFAD------------RMKRAAESLLPEG---KGIRINVQEVYPAHVGFGSGTQ MA4006 VDGGAGLTLSSPNVKVTAA-EADEIRIEGLQEFAD------------RMIKAAEALLPEG---KGVRIDVESLIPAHVGFGSGTQ MK0558 VDGSVGVTLEGPRIELEVEPTGEGVKVDGEGEIAE------------KAERAARKVLDLYGIEGGVRIEVVRRYPEHVGLGSGTQ MJ1427 VDGGIGLALEEPNIKIEGK-ESDDISIEFDKKLIEKYGEDYIKSVRDRVYNTAIKVLDVIGG-EGVDLKILSLFPAHSGLGSGTQ AF2089 VDGGVGLALEEPHIEIKAK-ESE--TFVLKGEPIN----------RERFEIAAAKMAEYCG--RGAEIEVVSDYDAHVGLGSGTQ MTH830 LDGGVGITLNEPELVVGLE-ASDDMGVEFTSHAEGKLREEY----RSKIMEAARRTLKHIGSDEKFHFTVRSMFPAHSGLGSGTQ ST1389 LDGGIGVALKYPRIVLRSG----------------------------NCIKPNITLPFKIP-----DYCIEEDFEEHIGLGHTTQ SSO0370 LDGGAGVALKYPRIVVRSG----------------------------NCFKPEVSLPFDVP-----KICIEEDFEQHIGLGHTTQ MM1592 AALSVAAAINEIYGLGK-SVRELAVAVGRGGTSGIGVAAFEYGGFILDGGHRFR---DKGAFSPSAAS-RMPPGPVLFRRAFP-D MM0322 AALSVAAAINEIYGLGK-SVRELAVAVGRGGTSGIGVAAFEYGGFILDGGHRFR---DKGAFSPSAAS-RMPPGPVLFRRAFP-D MA0339 AALSAAAAVNEIYGLGQ-SVRELAVAVGRGGTSGIGVAAFENGGFILDGGHRFR---DKGAFSPSAAS-HMPPGPVLFRRDFP-D MB2615 SSLAAAAAVNELYGLGK-SVRELALAVKRGGTSGIGVAAFEKGGFIVDGGHKFN---DKNGFMPSAAS-KVPPGPVLFREDFP-Q MA4006 AALAVAAAVNELYGLEK-DVRELAFAVKRGGTSGIGVTAFEKGGFIVDGGHRFK---DKGAFMPSAAS-RVPPGPVLFREDFP-D MK0558 ATLSAAVGTLEAHGVEHYDVRELADALGRGGTSGIGVAAFERGGFIVDGGHVFGP-GGKEEFKPSAASGEVPPAPVISRLEVPED MJ1427 LSLAVGKLISKIYNKEM-NAYNIAKITGRGGTSGIGIGAFEYGGFLIDGGHSFGKGKDKEDFRPSSASKGVKPAPIIFRHDF--D AF2089 ISLAVGRAFSELYGLNL-TTRQIAEIMGRGGTSGIGVAVFDHGGLVVDGGHST---KEKKSFLPSSASR-AKPAPMIARLDFP-D MTH830 LSLATARLVAEYHGMKF-TARELAHIVGRGGTSGIGVASFEDGGFIVDAGHSS---REKSDFLPSSASS-ASPPPVIARYDFPEE ST1389 FLLSVAKLGAEYN-LKNIDVVELAKLVKRGGTSGVGVYAFKHGGFIVDGGHSKK---IKKEILPSDYS-TVSPPPLIARYDFP-SSO0370 YLLSVAKLAAEYN-FKRLNSYELAKLVKRGGTSGVGVYAFEYGGFIVDGGHSTK---VKRELLPSDFS-SAPPPPLIARLNFPâ€” Phosphate binding motif DHPS MM1592 RLHLTLIDLNAEIGR MM0322 RLHLTLIDLNAEIGR MA0339 RLHLTLIDLNAEIGR MB2615 RIHMTLIDMNAEIGR MA4006 RLHITLIDMNGEIGR MK0558 RIHVTLIDLHGGLGR MJ1427 RIHMGLIDLNGSIGR AF2089 RIHITLIDLNGSIGR MTH830 RLHLTLIDLNGERGR ST1389 RIHITLIDLEGKFGR SSO0370 RIHITLFDLEGKYGR S.aureus DEGADIIDVGGVSTR E.coli NAGATIIDVGGESTR T.tengcong EEGADIIDVGGESTR T.maritima EEGADIIDVGGMSTR Phosphate binding motif RFAP synthase MM1592 WNIVLAIP---NTK-GVHDAEEVDIFKKACPIPLSEVQEISHVILMQMLPALIEEDIDSFGRAVNHFQTVG----FKKREVELQP MM0322 WNIVLAIP---NTK-GVHDAEEVDIFKKACPIPLSEVQEISHVILMQMLPALIEEDIDSFGRAVNHFQTVG----FKKREVELQP MA0339 WHIVLAIP---DTK-GAHDVEEVDIFKKVCPVPLREVQEVSHVILMQMLPAIIEEDLESFGKAINHVQTVG----FKKREVELQP MB2615 WDMVVAIP---NDK-GMHDQEEVETFKKFCPLPVEEVREISHVVLMQMMPAVIEEDIVNFGAAVNHVQTVG----FNKRESLIWP MA4006 WDIVIAVP---NDK-GMHDQQEIDVFQEFCPLPIEEVREVAHMVLMKMMPAVIEEDIESFGAAVNHVQTVG----FNKSESLIWP MK0558 WRFVLAIP---EVERGAHGDKEVNIFKRYCPVPAREVGEICRWILMVMMPAVVEDDPEDFGRAVDAIQDLG----FKRVEVGLQH MJ1427 WETILIIP---KGE-HVYGKKEVDIFKKYCPVPLNEVEKICHLVLMKMMPAVVEKNLDDFGEVINKLQYLG----FKKVELSLQS AF2089 WNVVLAIP---DLK-GFFGEREVNLFQKSCPVPLEDVREICHLILMKMLPAVVEADLDEFGKALKRIQELG----FKKAEVEQYG MTH830 WNIIIAIP---EIDRSVSGRREVNIFQEYCPLPLRDVERLSHIILMKMMPAILEGDIEAFGESVNEIQGTG----FKRIERELQD ST1389 WYIYVNIP---KEGRKIYGKSELEIFKN---AKVEGIDTLTRIIFMKLIPAVIENDLEEALEAIGLIQNLG----FKKLEVSLQT SSO0370 WYIYVNVV---RGGRRVFGKDEINAFKQ---AKLEGLDTLARVVLMELIPSVAERDLEGALNAISLIQDLG----FKKVEVFLQT MM1592 ETVLNVMKYMQDNGASGSGVSSFGPVVYGIVESSVEAGRLQKEVQRMLDESL-GGEVLLTKGRNRGADIFGGSD----------MM0322 ETVLNVMKYMQDNGASGSGVSSFGPVVYGIVESSVEAGRLQKEVQRMLDESL-GGEVLLTKGRNRGADIFGGSD----------MA0339 EPVLKAMKYMQDNGASGSGVSSFGPVVYGIVGSQSEGKKLQKEVQHMLDESL-GGEVMLTKAKNRGADIFGGSG----------MB2615 EFVKNIASFMRSR-SYGAGVSSFGPVVYSFVDNKSEGRQLQSEVQKMLDESV-GGITMMTRAKNSGAEISEI------------MA4006 DFVKSIASFMRSR-SYGAGVSSFGPVVYAFVDNREEGRQLQAEVQKMLDESV-GGITMMTRAKNSGAEISNV------------MK0558 PVVREMMEVARSAGAYGAGLSSFGPTVYAVCDSPSARDVAQELEMYMREEGI-GGEVSVSEPRNEGFEVTG-------------MJ1427 DIVKDLINELHKD--VYAGLSSFGPTIYAFGD-KKL--IVEKANDIFDKYGV-YGEIIITKANNVGHKIW--------------AF2089 ELIKGCFDLAD-----CIGMSSTGPTVYAITD-SNAGGIERSLRDYFAEKGY-ECRTIVTKARNRGVEIEV-------------MTH830 PLIDRIIDSMISAGAPGAGMSSFGPAVYSVTD-EKPGNVAGAVAEIMGP-----GRIIVTGGRNRGAFMIK-------------ST1389 EEVKMLMKQLYQKG-YYSGISSFGPAVYTFVRSRREG---EELVSYFGG--------FVTEPNNEGAKVLWLKD----------SSO0370 DEVKKLMNDMVKAG-FPAGLSSFGPAVYTFVSSKREG---DELISRFGG--------FVSEPNNEGAKVFWSTMSS--------Figure 2-9. Bioinformatic analysis of group 1 RFAP synthase genes. A. Multiple amino acid sequence alignment of Group I RFAP synthases using the alignment program GeneDoc (Nicholas 1997). Possible phosphate binding motifs are indicated by brackets. B. Multiple amino acid sequence alignment showing homology between a region of dihydropteroate synthase (DHPS) and the indicated conserved region of RFAP synthase.
52 This resulted in an apparent K m value of 30 M, within the same order of magnitude as that determined by Rasche and White, 1998 (Figure 2-7). Bioinformatics of RFAP Synthase A phylogenetic tree was previously constructed for RFAP synthase homologs (Scott and Rasche 2002). An updated phylogenetic tree incorporating the newly identified RFAP synthase homologs shows that the homologs can be divided into three distinct groups, in agreement with the previous findings (Figure 2-8) (Scott and Rasche 2002). One group is made up of homologs mostly from methanogenic archaea, along with homologs from Archaeoglobus fulgidus and two Sulfolobus species. The other two groups are made up of the bacterial homolog from M. extorquens (orf4) and paralogs from three nonmethanogenic archaea, Pyrococcus abyssi (PAB0441/PAB1694), Aeropyrum pernix (APE2425/APE1512), and Pyrococcus horokoshii (PH0227/PH1228). When the amino acid sequences of the homologs are aligned according to individual groups, the group 1 RFAP synthases appear to have a great deal of conserved regions present that are either less prominent or not seen in the remaining two groups (Figure 2-8). Further analyses of the conserved regions individually reveal three possible motifs common to the group 1 RFAP synthase sequences. Two of the conserved regions that were analyzed show significant similarity to glycine rich P-loop sequences known to bind phosphoribosyl containing substrates (Fu et al. 2002; Kleiger and Eisenberg 2002). One of these possible PRPP binding motifs found only in group 1 RFAP synthase sequences, as indicated in figure 2-9, is similar to the consensus sequence [V/I]XGX 1-2 GX 2 GX 3 [G/A] known to stabilize a -strand and -helix interaction in Rossmann fold containing proteins that bind nucleotides (Figure 2-9) (Kleiger and Eisenberg 2002). The other conserved glycine rich region has some
53 similarity to the phosphate binding loop present in mevalonate kinase (Figure 2-9) (Fu et al. 2002). A third conserved region was analyzed and showed 40% similarity to a region of the enzyme dihydropteroate synthase that forms part of a channel suspected to bind pABA (Figure 2-8B) (Baca et al. 2000). This conserved region has a number of charged residues, including D19 and R26, which are conserved among the RFAP synthase and dihydropteroate synthase sequences. This suggests that the conserved region plays a role in binding pABA. Site-directed Mutagenesis To investigate the possible roles of these charged amino acids, site-directed mutagenesis was used to replace the conserved residues D19, and R26 with alanines. This work was performed by students in the summer 2002 MCB4034L Advanced Microbiology Laboratory. The resulting variants were expressed in E. coli, as previously done for the recombinant wild-type. The proteins were partially purified and characterized by the students in the advanced lab revealing that the two variants were soluble, but lacked of RFAP synthase activity. The non-conservative replacement of R26 and D19 with alanine completely removes the functional group of these amino acids, possibly explaining the lack of activity in spite of the production of soluble protein. To investigate if this was the case, more conservative mutations of R26 and D19 were constructed by Dina Greene. R26 was replaced with a lysine, and D19 was replaced with an asparagine. Initial characterization of the R26K and D19N variants by Dina Greene and Rosemarie Garcia, showed that they were soluble and had low activity. The temperature optimum for the enzymes however, were lower then the wild-type, 50C for R26K and 60C for D19N
54 (Table 2-3). For the sake of consistency the variants were studied at 50C. At this temperature the wild-type enzyme purified through the Mono Q step, without the heat-step, had a K m for pABA of approximately 3.9 mM, slightly lower than that of the wild-type at 70C, with a Vmax of 1.45 nmol/min/mg (Table 2-3). The variants D19N and R26K were then partially purified by hydroxyapatite and Mono Q column chromatography. A heat-step was not used because of the instability of the variants at temperatures higher than 50C. The R26K variant had a K m of approximately 38 mM, 9-fold higher than the K m for the wild-type under similar conditions (Table 2-3). The Vmax for the R26K variant was approximately 5.0 nmol/min/mg which is 3-fold higher than that of the wild-type. These data are consistent with a role for R26 in the binding of pABA. The D19N variant had a K m of approximately 12 mM, 3-fold higher than the wild-type (Table 2-3). The Vmax for the D19N variant was approximately 0.45 nmol/min/mg (Table 2-3). This result suggests that the residue plays a role in the binding of pABA, but the role might be a less significant one in terms of substrate binding. The Vmax differences between the wild-type and variants under similar conditions could suggest that altering the amino acids also slightly affects the catalytic properties as well. Table 2-3. Site-directed mutagenesis of the recombinant wild-type RFAP synthase Conservative substitution Soluble Temperature optimum (C) K m pABA 1 (mM) Vmax 1 (nmol/min/mg) W/T + 70 3.9 1.5 R26K + 50 38 5.0 D19N + 60 12 0.5 1 Assays for K m determination were conducted at 50C for 3 h. Discussion The formation of RFAP, catalyzed by RFAP synthase, from pABA and PRPP is an important part of the eighteen step methanopterin biosynthetic pathway that gives
55 methanopterin distinguishing chemical and thermodynamic differences that set it apart from folate (White 1996; Maden 2000; White 2001). As a result of the low abundance of RFAP synthase present in methanogenic archaea it was necessary to develop a generally applicable recombinant expression system for putative RFAP synthase genes in order to be able to further analyze the biochemical properties of the enzyme (Scott and Rasche 2002; Bechard et al. 2003). Using this system the putative RFAP synthase homolog from M. thermautotrophicus, gene MTH0830, was recombinantly produced in E. coli, biochemically characterized and purified to homogeneity with a final specific activity of 360 nmol/min/mg (Figure 2-2 and Table 2-2). The purified recombinantly produced methanogen RFAP synthase was a dimer of identical subunits with a native molecular mass of 68.7 kDa. Further biochemical analysis revealed a K m for pABA of 4.5 mM, higher than previously found for the M. thermophila homolog (Figure 2-5) (Rasche and White 1998). The current work is the first report of the purification and biochemical characterization of a recombinantly produced methanogen RFAP synthase. Bioinformatic analysis of conserved regions among the RFAP synthase sequences revealed possible substrate binding regions for pABA and PRPP as well as possible structural differences between the RFAP synthase homologs (Figure 2-9). Site-directed mutagenesis was used to further analyze this putative pABA binding region, identifying two conserved charged amino acids, D19 and R26, as playing an important role in the activity and binding of pABA for RFAP synthase (Table 3, Figure 2-9). Previously, the RFAP synthase homolog from A. fulgidus, gene AF2089, was recombinantly produced in E. coli, purified and partially characterized as a dimer of identical subunits with a molecular mass of 57 kDa and a final specific activity of 240
56 nmol/min/mg; under optimal conditions at 70C pH 7.0 (Scott and Rasche 2002; Bechard et al. 2003). The M. thermophila RFAP synthase homolog, the only other methanogen RFAP synthase studied, was partially purified from the native organism and found to have a similar molecular mass of 65 kDa with an apparent K m for pABA of 58 M and an apparent K m for PRPP of 3.6 mM (Rasche and White 1998). The M. thermophila enzyme was later purified to homogeneity and used to identify the RFAP synthase homologs (Scott and Rasche 2002). When the previously studied RFAP synthases are compared to the M. thermautotrophicus enzyme of this study it can be seen that the RFAP synthases all catalyze the same Mg 2+ dependent reaction in a dimer form of comparable molecular masses (Rasche and White 1998; Scott and Rasche 2002; Bechard et al. 2003). Slight differences in the optimal enzymatic conditions of the RFAP synthase homologs were observed, with the A. fulgidus and M. thermautotrophicus homologs being more thermophilic with higher rates of activity (Figure 2-3) (Rasche and White 1998; Scott and Rasche 2002; Bechard et al. 2003). When the recombinantly produced M. thermautotrophicus RFAP synthase was biochemically analyzed, some interesting differences were brought to light. The enzyme has a K m for pABA of 4.5 mM, nearly two orders of magnitude higher than the apparent K m for pABA determined for the M. thermophila enzyme (Figure 2-5) (Rasche and White 1998). Although the K m value of 4.5 mM seems to be an unphysiological value, when the native enzyme was partially purified from M. thermautotrophicus, the apparent K m was 3.9 mM, similar to that of the recombinantly produced enzyme (Figure 2-6). These results indicate that the pABA K m for this RFAP synthase homolog is in fact very different from the previously studied methanogen RFAP synthase.
57 There are many possible explanations for this apparent difference. One explanation is that the partial purification of native RFAP synthase from M. thermautotrophicus could have removed a crucial cofactor or protein needed for the enzyme to function properly, which would also be absent in the artificial production of the enzyme in E. coli. Another possible explanation for this difference could be the presence of RFAP synthase paralogs among the Methanosarcina species, both Methanosarcina mazei, and Methanosarcina acetivorans (Figure 2-8). The wide growth substrate range of the Methanosarcina species could require more than one paralogous functional RFAP synthases (isoenzymes), with different biochemical properties, differentially expressed according to the growth conditions (White 2000). The strict hydrogenotrophic nature of M. thermautotrophicus however requires that the methanogen be grown under very strict conditions, with only H 2 /CO 2 as the carbon source. Thus, only one functional RFAP synthase is likely required to maintain proper coenzyme levels for the organism to thrive. The presence of isoenzymes differentially expressed under different growth conditions, has been previously documented (Harms and Thauer 1996; Vaupel and Thauer 1998; Pennings et al. 2000). Since the M. thermautotrophicus cells used in the current study were grown hydrogenotrophically on H 2 /CO 2 , and the Methanosarcina thermophila cells used in previous studies were grown on acetate, this would be a plausible explanation for the apparent K m difference. Bioinformatic analysis, sequence alignments and phylogenetic comparison of the known RFAP synthase homologs indicates three groups (Figure 2-8 and Figure 2-9). More detailed analysis of conserved regions among the methanogen RFAP synthase
58 homologs of group 1 revealed three possible substrate binding regions for pABA and PRPP (Figure 2-9). The first of the regions has high homology to the phosphate binding loop of mevalonate kinase (Figure 2-9A). In mevalonate kinase this phosphate binding site is usually composed of a loop connecting a -strand to an -helix, with the consensus sequence PX 3 GLGSSAA (Yang et al. 2002). This binding loop is also known to interact with the pyrophosphate tail of ATP for mevalonate kinase (Fu et al. 2002; Yang et al. 2002). The presence of a similar region in the RFAP synthase sequences could mean a role for this conserved region in the binding of the pyrophosphate moiety of PRPP. The second of the highly conserved regions indicated in figure 2-9A has significant homology to the consensus sequence known to stabilize -strand and -helix interactions for Rossmann fold containing proteins that bind FAD and NAD(P) (Figure 2-9) (Kleiger and Eisenberg 2002). In these proteins this -strand/-helix loop not only stabilizes the Rossmann fold but also interacts with the nucleotide near the phosphoribosyl portion of the nucleotide (Kleiger and Eisenberg 2002). The presence of this sequence in the RFAP synthase homologs provides evidence that this region likely binds to the phosphoribosyl portion of PRPP and possibly works together with the other putative phosphate binding loop to form an overall PRPP binding region within the active site of the enzyme. It is interesting to note however, that this particular conserved region is not present in the group 2 and 3 RFAP synthase homologs. The third of these conserved regions (Figure 2-9B) has similarity to a region in dihydropteroate synthase thought to be involved in the binding of pABA (Figure 2-9B) (Baca et al. 2000). The structure of dihydropteroate synthase is made up of a â€œTIM
59 barrelâ€ fold composed of eight helices surrounding eight central -strands with the 8 loops connecting the -helices and -strands playing critical roles in catalysis (Baca et al. 2000). For the Mycobacterium tuberculosis dihydropteroate synthase it is thought that one of these 8 loops, loop 2, aids in the overall formation of a pABA binding site within a solvent inaccessible channel (Baca et al. 2000). The similarity of this conserved region with a portion of the loop 2 region of dihydropteroate synthase, along with the presence of an arginine and aspartic acid residue conserved between both pABA binding enzymes, could indicate a similar role for this conserved region in RFAP synthase. To test this hypothesis, site-directed mutagenesis was used to mutate two conserved charged residues in this region, R12 and D19. The non-conservative replacement of these residues to alanine and the soluble inactive production of the resulting variants reveal that the functional groups of the residues play an important role in the activity of RFAP synthase. However, the inactivity of the variants does not allow for the determination of the specific role of these amino acids. A role for a conserved arginine, like R26, as previously been described as contributing its guanidium group for hydrogen bonding as well as its main chain for hydrophobic interactions during the binding of pABA for dihydropteroate synthase (Achari et al. 1997; Baca et al. 2000). To determine whether or not the functional group of R26 could have similar interactions with pABA for RFAP synthase the residue was replaced with a lysine, removing the guanidium group while leaving the main side chain. The conservative replacement resulted in the production of an active soluble variant of RFAP synthase (Table 2-2). When the variant was partially purified and biochemically analyzed it was determined that higher maximal activities were
60 reached as compared to the wild-type with a 9-fold higher K m for pABA at 38 mM (Table 2-3). The increase in the K m for pABA along with the previously proposed role for the corresponding arginine in dihydropteroate synthase suggests that the guanidium group, as well as the main side chain, of R26 likely plays a role in binding of pABA for RFAP synthase, perhaps similar to the corresponding residue in dihydropteroate synthase. The presence of a conserved carboxylate residue, like D19, surrounded by hydrophobic residues has previously been proposed to be involved in the stabilization of a carboxonium ion intermediate formed during one of the proposed mechanisms of RFAP synthase, as well as in other PRTase enzymes (Scapin et al. 1995; Rasche and White 1998). To determine if D19 could play a similar role in RFAP synthase the residue was replaced with an asparagine, removing the carboxyl group and replacing it with an amide. The replacement of D19 with an asparagine resulted in the soluble active production of an RFAP synthase variant. However, further analysis of the partially purified variant revealed slightly lower maximal activities, as compared to the wild-type and R26K variant, with only a three fold increase in the K m for pABA to 12 mM (Table 2-3). The smaller increase in the K m for pABA when the carboxylic acid side chain of the aspartic acid residue is replaced with a more basic side group possibly suggests less of a direct role for the amino acid in substrate binding and more of a role in the overall catalysis of the enzyme. Identification of these highly conserved amino acids that appear to play a role in the overall catalysis and substrate binding of RFAP synthase suggests that this highly conserved region could play a vital role in not only substrate binding and catalysis, but in the structure of the active site of RFAP synthase as well.
61 Distinct differences in the active site structures of PRTases have resulted in two evolutionarily distinct types of PRTases. The active site of type I PRTases are made up of an / core structure with a characteristic Rossmann fold and a solvent exposed active site that is flanked by hood and loop structures which close over the active site during catalysis (Eads et al. 1994; Smith 1999; Cao et al. 2002). The active structure of type II PRTases, solely represented by QAPRTase, is made up of a seven stranded / barrel open sandwich structure, which serves as a cap for the active site of the other subunit (Cao et al. 2002). The data presented here along with the identification of a characteristic Rossmann fold and an apparent phosphate binding loop suggests a model for the active site structure and catalysis of the methanogen RFAP synthase, similar to that of type I PRTases (Eads et al. 1994; Smith 1999; Cao et al. 2002). The absence of this characteristic Rossmann fold in either of the two remaining groups of RFAP synthase homologs suggests not only structural differences but also differences in substrate binding modes between the groups of RFAP synthases. This proposed model for the active site structure and catalytic mechanism of RFAP synthase, like other type I PRTases, would involve the formation of a slightly hydrophobic active site that would be inaccessible to bulk solvent water upon the sequential or random binding of the substrates (Eads et al. 1994; Smith 1999; Cao et al. 2002). The ability of the active site of RFAP synthase to form a solvent inaccessible active site would be crucial for the enzyme to catalyze the phosphoribosyl transfer event, as bulk solvent water can serve as a nucleophile hydrolyzing the activated substrate PRPP (Heroux et al. 1999; Shi et al. 1999; Baca et al. 2000). This model would also accommodate a previously proposed two-step S N 1-like mechanism for RFAP synthase, which involves the formation of a
62 carboxonium ion intermediate of PRPP thought to be stabilized by carboxylate residues (Rasche and White 1998). The work presented here provides valuable information that would not only be important in the overall understanding of RFAP synthase but also in the interpretation of a crystal structure when it becomes available. This work, along with a crystal structure, would provide many insights that could have multiple applications. One specific application, proposed by Dumitru et al. (in press), involves developing an inhibitor specific for RFAP synthase that would not only result in inhibition of the biosynthesis of methanopterin, but inhibit the methanogenic process as well (Dumitru et al. in press). Current inhibitors being developed for RFAP synthase have in fact resulted in inhibition of the synthesis of methanopterin as well as growth of methanogenic organisms, and production of methane through methanogenesis (Dumitru et al. in press).
CHAPTER 3 CONCLUSIONS Significance The current study describes our contribution to increasing the understanding of the structure and overall catalysis of RFAP synthase. We have recombinantly produced and characterized the methanogen RFAP synthase from M. thermautotrophicus (MTH0830), a close relative of ruminal methanogens. Recombinant production of a soluble active methanogen RFAP synthase in this manner has provided RFAP synthase in quantities needed for its purification and biochemical characterization. Subsequent bioinformatic analysis and site-directed mutagenesis of the enzyme has also led us to propose the aforementioned model for the overall catalysis and active site structure of RFAP synthase. Furthermore, we propose that the identified Rossmann fold and phosphate binding loop sequences interact to form the binding site for PRPP. The conserved region that has homology to the suspected pABA binding region of dihydropteroate synthase, as well as the identification of a conserved arginine in the region that appears to be important for pABA binding, has led us to also propose that this region contributes to the overall formation of a slightly hydrophobic region of the active site that would provide important contacts with pABA (Achari et al. 1997; Baca et al. 2000). This proposed model also accommodates previously described mechanisms for RFAP synthase, as well as supporting the hypothesis that a hydrophobic area of the active site of RFAP synthase would be involved in the binding of pABA (Rasche and White 1998) (Dumitru et al. in 63
64 press). The work described in this study is the first report of the purification and biochemical characterization of a recombinantly produced methanogen RFAP synthase. RFAP synthase catalyzes the key step in the synthesis of methanopterin, providing the one-carbon carrier with distinguishing characteristics that set it apart from folate. Methanopterin also plays an essential role in the methanogenic process used by methanogens, not only to generate methane but also to provide the organisms with their only source of energy. Therefore, the identification and biochemical characterization of a methanogen RFAP synthase in the manner currently described is significant because it not only provides a better understanding of the overall structure and catalytic properties of this enzyme, but it also provides a foundation of knowledge that is useful for future work involving this enzyme. This work is also significant in that the knowledge gained here has the potential to impact the levels of methane released by these methanogenic organisms. In the rumen of domesticated livestock these methanogenic organisms compete with other organisms, such as acetogenic bacteria, for the molecular hydrogen and carbon dioxide required in their production of methane (methanogenesis) (Le Van et al. 1998; Deppenmeier 2002). However, acetogenic bacteria use the molecular hydrogen and carbon dioxide to produce acetate (acetogenesis), which is a beneficial nutrient for livestock (Wolin 1981; Le Van et al. 1998). Therefore, the presence of methanogenic archaea negatively affects the production of these beneficial nutrients through nonproductive conversion of acetate or molecular hydrogen and carbon dioxide to methane in methanogenesis (Wolin 1981; Le Van et al. 1998). Methane is also a known potent greenhouse gas, and its production from this source has been suspected to
65 contribute to global warming (Khalil and Shearer 1993; Drake 2000). Inhibition of the ruminal methanogens by targeting the essential methanogenic process has therefore been a longstanding strategy (Wolin 1981). However, previous inhibitors of the methanogenic process such as monensin have not been specific enough to target only the methanogenic organisms (Chen and Wolin 1979). As a result of the importance and specificity that RFAP synthase has for the synthesis of methanopterin and the important role of the one-carbon carrier in methanogenesis, RFAP synthase has been targeted for the development of inhibitors that would specifically inhibit this methanopterin biosynthesis (Dumitru et al. in press). Inhibitors of this nature should result in the overall inhibition of methanogens by specifically targeting methanogenesis (Dumitru et al. in press). The idea of inhibiting a specific group of organisms by targeting the biosynthesis of an essential cofactor was first employed in the development of sulfonamides. Sulfonamides are a class of antibacterial agents that specifically inhibit the growth of bacteria through the inhibition of the biosynthetic pathway for folate (Brown 1962). These folate biosynthetic inhibitors are in fact pABA analog inhibitors of pABA that inhibit the biosynthesis of folate by targeting the folate biosynthetic enzyme dihydropteroate synthase. This enzyme is essential in folate biosynthesis and is responsible for the condensation of pterin pyrophosphate with pABA forming the backbone of folate (Shiota et al. 1969; Roland et al. 1979). Future Experimentation In order to verify the proposed model as well as the involvement of these conserved regions in the binding of the substrates and overall catalysis of RFAP synthase, further experimentation is needed. Future experimentation would include more extensive kinetic
66 studies characterizing the K m for PRPP for the wild-type enzyme as well as additional characterization of the binding order of the substrates. These kinetic studies would provide increased knowledge of the mechanistic action used by the enzyme to catalyze this unique chemistry. Additional site-directed mutagenesis studies would help to further characterize the proposed pABA binding region, as well as the Rossmann fold and P-loop regions, leading to a better understanding of specific amino acids involved in direct interactions with pABA and PRPP. These studies would greatly contribute to a foundation of knowledge that would be useful in interpreting a crystal structure for RFAP synthase when it becomes available. Overall the development of a crystal structure for RFAP synthase in combination with the aforementioned experiments would provide valuable information about RFAP synthase that would increase the basic understanding of this essential methanopterin biosynthetic enzyme, while also having an impact on the development of more effective and specific methanogenesis inhibitors.
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BIOGRAPHICAL SKETCH Matthew E. Bechard was born on December 22, 1978 in Lewiston, Maine to Kathryn and Edward Bechard. Matthewâ€™s father was in the Air Force, causing him and his family to move soon after his birth to Griffis AFB, NY. where he attended grades K through second. After second grade the Bechard family moved to Eglin A. F. B. where Matthew attended third grade at Oak Hill Elementary School. Up until this point in his life Matthew was not exceedingly proficient at school. After moving to a small town near Eglin A.F.B., called Crestview, Matthew attended the fourth grade at Southside Elementary School where he was under the guidance of Donna Roberts. Donna Roberts was a very influential and caring teacher who was responsible for â€œlighting the fireâ€ in Matthewâ€™s mind and life. Matthew achieved the honor roll for the first time in Donna Robertâ€™s fourth grade class and has rarely ever left it since. Through middle school at Richbourg Middle School and high school at Crestview High School, Matthew continued to excel in school, setting his sights on medical school. Matthew graduated high school class of 1997 with high honors in the top 10% of his class with a Florida Bright Futureâ€™s Scholarship. He then obtained his Associate of Arts degree at Okaloosa Walton Community College, after which he applied and attended the University of Florida (UF). He graduated with honors from UF in May of 1999, obtaining Bachelor of Science degree in microbiology and cell science. While attending UF, Matthew conducted undergraduate research in the laboratory of Dr. Madeline Rasche, studying the enzyme RFAP synthase from the methylotrophic bacterium Methylobacterium extorquens. 78
79 intrigued by research Matthew decided to pursue a masterâ€™s degree under the guidance of Dr. Madeline E. Rasche, studying the enzyme RFAP synthase from the methanogenic archaeon M. thermautotrophicus. The focus of his masterâ€™s project was the recombinant production, purification, characterization, and site-directed mutagenesis of RFAP synthase from this methanogenic archaea. Matthew is graduating with a Master of Science degree in microbiology and cell Science in December of 2003. He plans to take a much-needed break from classes while pursuing his next course of action, which will most likely be an M.D./Ph.D. program.