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Bioorganic Molecules in the Cosmos and the Origins of Darwinian Molecular Systems

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BIOORGANIC MOLECULES IN THE COSMOS AND THE ORIGIN OF DARWINIAN MOLECULAR SYSTEMS By ALONSO RICARDO A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLOR IDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY UNIVERSITY OF FLORIDA 2004

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This work is dedicated to my family.

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ACKNOWLEDGMENTS I will like to thank Dr. Steven Benner for the research opportunities and guidance that he has offered me. I also thank Dr. Matthew Carrigan for his help, friendship and for being an excellent lab partner. I am grateful to the people in the mass spectrometry laboratory at the University of Florida and to Dr. Maurice Swanson for providing laboratory space and collaboration I also thank Fabianne for her unconditional love and support. Finally, I am forever grateful to my family for their constant support, care, and consideration that helped me to made this work possible. iii

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TABLE OF CONTENTS page ACKNOWLEDGMENTS .................................................................................................iii LIST OF TABLES ...........................................................................................................viii LIST OF FIGURES ...........................................................................................................ix ABSTRACT .....................................................................................................................xiii CHAPTER 1 ORIGINS....................................................................................................................1 Background and Significance.....................................................................................1 Prebiotic Chemistry.............................................................................................1 Sugars in the Prebiotic Environment..................................................................3 RNA World and the Problem of Ribose Accumulation......................................6 The Role of Minerals on Ribose Formation and Accumulation.........................8 2 PREBIOTIC SYNTHESIS OF SUGARS..................................................................9 Introduction................................................................................................................9 Boron Chemistry and Complexation Mechanism in Polyols............................13 pH dependence on the stability of boric acid esters and borate esters.......16 Differential coordination of borate to polyols...........................................17 Interaction of boron with carbohydrates : aldoses and ketoses..................19 Stabilization of pentoses towards decomposition in borate.......................23 Materials and Methods.............................................................................................23 Chemicals..........................................................................................................23 Enzymes............................................................................................................24 Analytic Instrumentation...................................................................................24 Ultraviolet analysis (UV)...........................................................................24 Gas chromatography (GC).........................................................................24 Mass spectrometry (MS)............................................................................24 NMR Spectroscopy....................................................................................25 Synthetic Preparations.......................................................................................25 Synthesis of colemanite.............................................................................25 Synthesis of deuterated Colemanite...........................................................25 Synthesis of pentoses in the presence of colemanite.................................25 iv

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Synthesis of pentoses in the presence of calcium hydroxide.....................26 Derivatization of pentoses for gas chromatography analysis....................26 Sugars Degradation Experiments......................................................................27 Sugar decomposition in the presence of calcium deuteroxide...................27 Sugar decomposition in calcium deuteroxide and colemanite...................27 Enzymatic Assays.............................................................................................28 Ribitol dehydrogenase assay......................................................................28 Cysteine-carbazole test..............................................................................29 DIOS Analysis..................................................................................................29 Preparation of PSi surfaces........................................................................29 Competition Experiments.................................................................................30 1, 4-Anhydroerythritol (AET) vs Pentoses................................................30 13 C-Ribose vs Pentoses..............................................................................30 Results......................................................................................................................30 Synthetic Preparations.......................................................................................30 Synthesis of pentoses in the presence of colemanite.................................30 Synthesis of pentoses in the presence of calcium hydroxide.....................32 Sugar Degradation Experiments.......................................................................33 Sugar decomposition in the presence of Calcium Deuteroxide.................33 Sugar decomposition in the presence of Colemanite.................................34 Enzymatic Assays.............................................................................................35 DIOS Analysis: Competition Experiments.......................................................36 1, 4-Anhydroerythritol (AET) vs Pentoses................................................36 13 C-Ribose vs Pentoses..............................................................................36 Discussion................................................................................................................39 3 CATALYSIS AND THE RNA WORLD................................................................43 Introduction..............................................................................................................43 Materials and Methods.............................................................................................45 Preparation of Precursor DNAzymes via PCR (Maniatis et al, 1982)..............45 Preparation of single-stranded DNAzymes.......................................................47 5-End Labeling of DNA..................................................................................48 DNAzyme Kinetic Assays................................................................................49 Cloning and Sequencing DNAzymes................................................................49 In vitro Selection...............................................................................................50 Results......................................................................................................................54 In vitro Selection...............................................................................................54 Cleavage of 614 does not go Completion.........................................................56 Inhibition by Incompletely Removed Complementary Strand.........................57 An approach to Chemical Equilibrium does not Account for the Plateau........59 Testing if the Cleavage Products are Acting as Catalysts or Inhibitors............60 Improperly Folded ribose-614 Accounts for Part of the Plateau......................61 Mutations Introduced into 614 during Cloning and Sequencing......................63 Ribose-614 catalysis is not Mg++-dependent...................................................66 Ribose-614 cleaves in trans..............................................................................66 v

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Various ribose-containing substrates are cleaved by deoxyribose-614.............69 Competition Studies of Ribose-614 Cleavage..................................................72 Saturation kinetics in trans cleavage by deoxyribose-614................................73 Compound deoxyribose-614 cleaves with multiple-turnovers..........................74 Catalytic power in trans is unaffected by annealing protocol...........................74 The commitment step for deoxyribose-614 cleavage........................................75 Dependence on temperature of deoxyribose-614 cleavage...............................77 Predictions of the energetically favored structure.............................................79 Discussion................................................................................................................81 4 DETECTING ORGANIC MOLECULES ON MARS............................................90 Introduction..............................................................................................................90 Oxidation of Alkanes Under Martian Conditions.............................................93 Oxidation of Alkylbenzenes Under Martian Conditions..................................96 Oxidation of PAHs under Martian Conditions.................................................96 Oxidation of Kerogen under Martian Conditions.............................................97 Oxidation of Amino and Hydroxyacids under Martian Conditions..................99 The Amounts and Fates of Organic Carboxylic Acids.....................................99 Failure of Viking 1976 to detect Organic Carboxylic Acids..........................102 The Infrared Spectra of the Martian Surface...................................................104 Detecting the Missing Organics on Mars........................................................107 Materials and Methods...........................................................................................108 Chemicals........................................................................................................108 Analytic Instrumentation.................................................................................108 Fluorescence analysis...............................................................................108 Infrared analysis.......................................................................................108 High performance Liquid chromatography (HPLC-MS)........................108 Synthetic Preparations.....................................................................................109 Synthesis of Mellitic Acid Salts......................................................................109 Synthesis of manganous mellitate (1)......................................................109 Synthesis of zinc mellitate (2)..................................................................109 Synthesis of cupric mellitate (3)..............................................................109 Synthesis of nickel mellitate (4)..............................................................109 Synthesis of cobalt mellitate (5)..............................................................109 Synthesis of magnesium mellitate (6)......................................................110 Synthesis of calcium mellitate (7)............................................................110 Synthesis of iron mellitate (8)..................................................................110 Synthesis of fluoresceins: Fiegls test......................................................110 Results....................................................................................................................111 Synthesis of Mellitic Acid Salts......................................................................111 Fluorescence Spectra Analysis........................................................................111 Discussion..............................................................................................................116 vi

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APPENDIX A NMR DEGRADATION EXPERIMENTS............................................................119 B DIOS COMPETITION EXPERIMENTS..............................................................124 LIST OF REFERENCES.................................................................................................130 BIOGRAPHICAL SKETCH...........................................................................................137 vii

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LIST OF TABLES Table page 2.1. Borate complexes of aldopentoses, aldohexoses and ketohexoses.............................21 2.2. Retention times of trimethylsilyl derivative of pentoses............................................32 2.3. Half life of pentoses under alkaline conditions determined by 1 H NMR...................35 3.1. Name, sequence, and description of oligonucleotides................................................52 3.2. Data from plot ln[S] t versus time for ribose-614 cleavage.........................................68 4.1. Expected metastable products from organic substances.............................................93 4.2 Fluorescence analysis of the fluoresceines of benzenecarboxylates..........................111 viii

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LIST OF FIGURES Figure page 1.1. Proposed autocatalytic glycolaldehyde regeneration....................................................5 2.1. Some organic compounds detected in the ISM...........................................................11 2.2. D-ribose structure........................................................................................................12 2.3. Lewis structure representation of boric acid and borate.............................................14 2.4. Mechanism of boronic acid complexation by acidic ligands......................................15 2.5. Borate complexation by non-acidic ligands................................................................16 2.6. Free energy diagram of threo and erythro diols..........................................................18 2.7. Free energy diagram of syn-, and anti-,-diols.....................................................18 2.8. Structures of the B-L 2 spirane complex......................................................................22 2.9. Psi surface preparation...............................................................................................29 2.10. HPLC-MS analysis of reaction mixture containing colemanite...............................31 2.11. Detection of ribose-borate comples by ESI (-) ion mode.........................................31 2.12. GC trace of the reaction mixture containing colemanite..........................................32 2.13. GC trace of the reaction mixture containing Ca(OH) 2 ..............................................33 2.14. Incubation of ribose in Ca(OH) 2 solution.................................................................34 2.15. Incubation of ribose in the presence of Ca(OH) 2 + colemanite................................35 2.16. Anhydroethrythritol (AET)-pentose borate ions detected by DIOS.........................36 2.17. DIOS spectra of competition experiment D-arabinose vs AET................................37 2.18. Competition experiments between the different pentoses and AET.........................37 2.19. 13 C, 12 C-D-ribose borate ions detected by DIOS.......................................................38 ix

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2.20. DIOS spectra of competition experiment 13 C ribose vs 12 C ribose...........................38 2.21. Ratio of borate complexes of 13 C-ribose vs pentoses...............................................39 2.22. Suggested mechanism for pentose formation...........................................................41 3.1. In vitro selection experiment representation...............................................................45 3.2. Sequence of the initial library and DNAzymes.........................................................46 3.3. Ribose-614 cleavage..................................................................................................57 3.4. Cleavage products do not affect ribose-614 cleavage...............................................61 3.5 Gel-purification of ribose-614 at cleavage plateau.....................................................62 3.6. Reheating ribose-614 results in additional cleavage.................................................64 3.7. Sequence alignment of cleaved and uncleaved cloned 614.......................................65 3.8. Initial rate of ribose-614 cleavage as a function of [ribose-614]...............................67 3.9. Ribose-614 cleavage rate is concentration dependent...............................................68 3.10. Both deoxyribose-614 (left panel) and ribose-614 (right panel)............................70 3.11. Cat+ribose competes with ribose-614 for cleavage................................................70 3.12. Compound deoxyribose-614 can cleave various substrates......................................71 3.13. Various substrates can compete with ribose-614 for self-cleavage.........................72 3.14. Cleavage of various substrates by 614 is reduced...................................................76 3.15. Ribose-614 rate of self-cleavage in trans.................................................................78 3.16. Burst kinetics...........................................................................................................79 3.17. A univariate statistical distribution...........................................................................82 4.1 Oxidative degradation of the generic alkane...............................................................95 4.2.Oxidative degradation of naphthalene to phthalic acid................................................98 4.3. Martian surface IR spectrum.....................................................................................105 4.4. Mars dust, labradorite standard and labradorita........................................................105 4.5. Mars dust and magnesita-labradorite mixture spectra..............................................106 x

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4.6. Phthalic acid yield fluorescein..................................................................................108 4.7. Infrared spectra of manganous mellitate (KBr)........................................................112 4.8. Infrared spectra of zinc mellitate (KBr)....................................................................112 4.9. Infrared spectra of cupric mellitate (KBr)................................................................113 4.10. Infrared spectra of nickel mellitate (KBr)...............................................................113 4.11. Infrared spectra of cobalt mellitate (KBr),..............................................................114 4.12. Infrared spectra of magnesium mellitate (KBr)......................................................114 4.13. Infrared spectra of calcium mellitate (KBr)............................................................115 4.14. Infrared spectra of iron mellitate (KBr)..................................................................115 4.15. Infrared spectra of aluminium mellitate (KBr).......................................................116 4.16. Infrared spectra of the mellitate salts......................................................................117 4.17. Structures of the fluorescein derivatives of pyromellitic acid................................118 A1.1. D-arabinose incubation in the presence of calcium hydroxide, pD:12..................120 A1.2. D-arabinose incubation in the presence of calcium-hydroxyde + borate...............120 A1.3. D-lyxose incubation in the presence of calcium hydroxide, pD:12.......................121 A1.4. D-lyxose incubation in the presence of calcium hydroxide + borate pD:12........121 A1.5. D-ribose incubation in the presence of calcium hydroxide, pD:12........................122 A1.6. D-ribose incubation in the presence of calcium hydroxide + borate, pD:12.........122 A1.7. L-xylose incubation in the presence of calcium hydroxide, pD:12.......................123 A1.8. L-xylose incubation in the presence of calcium hydroxide + borate, pD:12.........123 A2.1. DIOS spectra of 1,4-Anhydroerythritol vs arabinose............................................124 A2.2. DIOS spectra of arabinose vs 13 C-ribose...............................................................124 A2.3. DIOS spectra of 1,4-Anhydroeythritol vs lyxose...................................................125 A2.4. DIOS spectra of lyxose vs 13 C-ribose....................................................................125 A2.5. DIOS spectra of 1,4-Anhydroerythritol vs ribose..................................................126 xi

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A2.6. DIOS spectra of 12 C-ribose vs 13 Cribose.............................................................126 A2.7. DIOS spectra of 1,4-Anhydroerythritol vs Xylose................................................127 A2.8. DIOS spectra of xylose vs 13 C-ribose....................................................................127 A2.9. DIOS spectra of 1,4-anhyroerythritol vs ribulose..................................................128 A2.10. DIOS spectra of ribulose vs 13 C-ribose................................................................128 A2.11. DIOS spectra of Xylulose vs 13 C-ribose..............................................................129 xii

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Abstract of Dissertation Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy BIOORGANIC MOLECULES IN THE COSMOS AND THE ORIGIN OF DARWINIAN MOLECULAR SYSTEMS By Alonso Ricardo May, 2004 Chair: Steven Benner Major Department: Chemistry Two critical unsolved issues in the origins of life field are the prebiotic formation of the molecular building blocks of life and from these, the appearance of a self-replicating molecule that undergoes Darwinian evolution. In the present dissertation, both issues were addressed by an experimental approach from which the following findings are reported. A plausible prebiotic route for the synthesis of sugar pentoses starting from materials known in the interstellar matrix was achieved. Ribose, one of the pentoses and constituents of ribonucleic acid, was generated from a reaction mixture containing boron minerals. The role of boron in this process was found to be dual. Boron coordinates to glyceraldehyde blocking the enolization process and binds to the pentose sugar preventing decomposition. The formation of ribose appears to be the natural consequence of the intrinsic chemical reactivity of compounds available from the interstellar medium under alkaline, calciferous conditions. As these conditions are not excluded from the xiii

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early Earth, it is also not possible to exclude the availability of pentoses at the time when life originated. In vitro selections performed in the presence of Mg++ generated DNA sequences capable of cleaving an internal ribonucleoside linkage. Several of these, surprisingly, displayed intermolecular catalysis and catalysis independent of Mg++, features that the selection protocol was not explicitly designed to select. A detailed physical organic analysis was applied to one of these DNAzymes, termed 614. The DNAzyme 614 is more active in trans than in cis, and more active at temperatures below the selection temperature than at the selection temperature. Many of these properties are unreported in similar systems, and these results expand the phenomenology known for this class of DNA-based catalysts. A brief survey of other catalysts arising from this selection found other Mg++-independent DNAzymes, and provided a preliminary view of the ruggedness of the landscape relating function to structure in sequence space. Finally, in the last chapter of this dissertation a method was designed, that allows the evaluation and detection of potential organic molecules on Mars. The method was tested with synthetic salts of mellitic acid, that are likely to be formed under Martian conditions. The presence of these molecules in the martian soil, was evaluated by direct comparison with the recently published Infrared spectra of the Mars surface xiv

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CHAPTER 1 ORIGINS Background and Significance Prebiotic Chemistry Experimental prebiotic chemistry as a modern approach to study the origins of life was born just over 50 years ago, in the work of Stanley Miller (1953). Miller demonstrated that applying electric discharges to a mixture of reduced gases in the presence of water generates a brown solution containing amino acids. Neither sugars nor nucleobases were produced under these conditions. Millers experiment, which at the time was considered to reproduce conditions on early Earth atmospheres, was a modern example of the abiotic origin of biological molecules. Since then, the approach in prebiotic experiments remains the same. A mixture containing molecules believed to be present in a prebiotic environment is exposed to a source of energy. If any amount of the desired product is detected, it is then claimed that, if there was plausibly a historical moment during which the Earth contained the starting materials, it is then possible to assume that the mentioned reaction took place, allowing the compound to accumulate over long periods of time. This logic seems intuitively valid. The methodology, however, can be flawed at different stages: The retrosynthetic analysis used to determine the most likely starting materials, may ignore the geochemical constraints necessary to make the reaction relevant. 1

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2 Starting materials can be chosen mainly based on reactivity rather than prebiotic relevance. Water is included/excluded in the model by convenience; no logical reasons other than the instability of reactants and products in the chosen solvent are addressed. In some cases concentrations of reactants are controlled at stoichiometric ratios to bias the reaction towards one desired product. This is a common practice in organic chemistry. When applied to prebiotic chemistry, however, this practice fuels arguments that origin required intelligent design. It is worth mentioning that limiting reagents are not prohibited as long as their abundance can be explained through geochemical constraints. The statements expressed above do not necessarily apply to prebiotic chemistry elsewhere in the universe. For example, an inventory of interstellar compounds detected by radioastronomical methods and the inventory of organic compounds found in meteorites suggest that the non-terrean chemical repertoire is rich in molecules that have a short half life under terrean conditions, but nonetheless exist in the cosmos. These molecules constitute valuable starting materials for terrean chemistry. Biologically relevant compounds may eventually arise when interstellar material delivered to the Earth (by meteor or comet impact) interacts with terrean molecules, volcanic emissions, water, or surface minerals. Therefore, for a prebiotic experiment to be meaningful, chemistry and geology must be linked; only through a concise analysis of the chemical evolution (organic and inorganic) of our planet will we be able to explore the possible outcomes of an experiment within natural constrains.

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3 In the end, this methodology will not conclude how in fact life began, or what chemistry indeed happened, but at the very least, will offer evidence of a plausible mechanism (able to suffer scientific scrutiny) for the formation of key molecules present in modern life. Sugars in the Prebiotic Environment Sugars, and pentoses in particular, are one of the building blocks of nucleic acids. Other sugars, including hexoses, are key throughout metabolism and structural biochemistry. Sugars therefore are logical targets for prebiotic chemistry experiments. Early attempts to synthesize sugars from simpler molecules were not done explicitly to reproduce prebiotic events. In 1861 Butlerow reported the formation of a brown, sweet tasting compound resulting from the reaction of an aqueous formaldehyde solution in the presence of calcium hydroxide (Ca(OH) 2 ). The product had a molecular formula (calculated by elemental analysis) corresponding to C 7 H 14 O 6 and was named methylenitan. The term formose to describe the same product composition was introduced by Loew in 1886 while reacting gaseous formaldehyde and calcium hydroxide. Further characterization of the reaction products by derivatization showed that instead of being a single product, the formose sugar was a heterogeneous mixture of monosaccharides. This suggested a complex reaction mechanism with many possible outcomes. The first mechanistic studies on the formose reaction focused on the isolation of intermediates. From this work, it was found that glycolaldehyde, glyceraldehyde and dihydroxyketone were early addition products of formaldehyde condensation (Henry, 1895; Neuberg, 1902). These compounds can then react with either additional

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4 formaldehyde, or cross-react to produce a mixture containing tetroses, pentoses, hexoses, and branched sugars, in both aldoand ketoforms. A kinetic analysis of the formose reaction shows an initial period during which products are not made (the induction period). This is followed by a period in which all the compounds are formed relatively quickly. Breslow suggested a mechanism to explain the formose reaction in 1959 (Figure 1.1). According to Breslow, if any direct joining of formaldehyde molecules exists, it must be very slow. In this way, he explained the induction period. The novelty in Breslows work was to introduce the concept of an autocatalytic reaction during which the first addition product, glycolaldehyde, is regenerated. Because the autocatalytic process does not require condensations between two electrophiles (as in the condensation of formaldehyde), the autocatalytic formation of glycolaldehyde proceeds at a fast rate in alkaline conditions. Several authors tested this mechanism by starting the reaction from the intermediates suggested by Breslow (Pfeil & Ruckert, 1960; Ruckel, Pfeil & Scharf, 1965). These authors obtained similarly complex products. Further, the addition of glycolaldehyde at the beginning of the reaction was found to reduce significantly the induction period. With the invention of chromatography, formose sugar composition was better characterized and found to consist of 10% C 4 30% C 5 55% C 6 5 % > C 6 (detectable by GC-MS analysis) sugars when formaldehyde is ca. 99% consumed (Weiss et a1. 1970; Decker et al. 1982).

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5 CC OH O H H H -OH CC OH O H H Ca++ CH O H HO H OC C C H OH H OH H H -OH CC O C H H HO OH H H -OH CC O C H OH HO H H HC H O HO H OHC H H C O C OH C H OH H H OC H C OH C O C H OH H H H H OC C OH H H H +O OH H H -OH GlycolaldehydeFormaldehydeGlyceraldehydedihydroxyacetoneFormaldehyde2-tetrulosaaldotetrosaGlycolaldehydeGlycolaldehyde enediol Isomerization Figure 1.1. Proposed autocatalytic glycolaldehyde regeneration in the formose reaction. Parallel to the discovery of the formose reaction, studies on the chemical composition of nucleic acids were also yielding some important results. In a series of papers between 1891-1894 Albrecht Kossel deduced the structure of the heterocyclic bases that constitute nucleic acids. Levene and Jacobs (1909) completed the structure of the nucleoside of the RNA (known at the time as nucleic acid of plant origin) by finding that the carbohydrate portion was made of the sugar D-ribose. The elusive structure of the carbohydrate in DNA (named nucleic acid of animal origin) remained unsolved until in 1929, when Levene and London concluded that deoxyribose was indeed the sugar component.

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6 With the advent of prebiotic chemistry in the 1960s, it was not long before the formose synthesis was reexamined from a prebiotic context as a way of obtaining the sugars necessary for biology. With the realization that ribose is the backbone of RNA, and having detected this sugar as a product in the formose reaction (although ribose overall yield and long term stability were low in the experiment), prebiotic chemists were satisfied, with the idea of having found that ribose was prebiotically available. RNA World and the Problem of Ribose Accumulation In 1962 Alexander Rich hypothesized that RNA might have played both catalytic and genetic roles in early forms of life. This idea was furthered by comments from Woese (1967), Orgel (1968) and Crick (1968), who remarked that transfer RNA (tRNA) appeared to be an RNA molecule attempting to fold like an enzyme. The discovery of catalytic RNA by Cech, Altman, Usher, and others supported this notion (Kruger et al. 1982; Guerrier-Takada et al. 1983; Usher & McHale, 1976). Gilbert proposed in 1986 that organisms in an RNA world may have been a precursor to the contemporary protein-DNA-RNA organisms that dominate life on Earth today. In the RNA world, life used RNA as the sole genetically encoded biological catalysts. However, the plausibility of the RNA world hypothesis is obviously conditioned to the success of generating RNA from a plausible prebiotic soup. The difficulties of generating RNA without the assistance of a pre-existing living system have been noted by many authors and can be divided into two categories: (a) those directly related to the activity of water, and (b) those that are not. In the first category, RNA is thermodynamically unstable in water with respect to hydrolysis. Further, many RNA nucleobases are hydrolytically unstable. Cytidine, for

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7 example, deaminates with a half life of ca. 10 2 years to give uridine (Frick, Mac Neela & Wofelden, 1987). Adenosine deaminates to give inosine at a slower rate, while guanosine deaminates to give xanthosine. These reaction modes, favored at high pH, are matched by depurination and depyrimidinylation reactions at low pH, which are also favored thermodynamically in water. A series of criticisms of the RNA origins of life hypothesis are not directly related to water, however, but rather concern ribose itself. These focus on difficulties of creating the necessary amounts of ribose to support a RNA world in the early Earth, and the instability of ribose under prebiotic conditions where it might be generated. This problem was addressed by Robert Shapiro in 1988, who, focusing on the formose reaction, concluded that the synthesis and accumulation of ribose in any significant amount under prebiotic conditions were very unlikely events. Shapiros comments are correct; the overall yield of ribose in formose is less than 1% after an arbitrary length of time, and less if the incubation is allowed to continue indefinitely. Ribose itself contains both an electrophilic center (carbon-1) and a nucleophilic center (carbons 1 and 2 of the enediolate) (Figure 2.2). This makes ribose unstable under basic conditions with respect to further reactions with formaldehyde, glycolaldehyde, or itself, or other nucleophiles and electrophiles that are emerging under formose conditions. Not surprisingly, in the presence of Ca(OH) 2 ribose is converted to higher condensation products, branched chain sugars, and (ultimately) a brown, largely intractable polymer of undefined composition. Browning of the mixture is pronounced within an hour at room temperature and within minutes at 60 C. Thus, the extent of

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8 accumulation of ribose as a product of formaldehyde and glycolaldehyde at steady state (formation minus destruction) is quite low. Given this, Stanley Miller and coworkers, commented that the rate of this decomposition reaction is so high that it suggested that the backbone of the first genetic material could not have contained ribose or other sugars because of their instability (Larralde, Robertson & Miller, 1995). The Role of Minerals on Ribose Formation and Accumulation While the scientific literature contains many reports concerning the use of minerals in prebiotic experiments, few of these publications include a geochemical explanation of how and why the mineral was available on the early Earth. Ponnamperuma reported the use of the clay mineral kaolin (aluminium hydroxide silicate (Al 2 (OH) 4 .Si 2 O 5 ) a weathering product of feldspar), as a catalyst in the formose reaction at low concentrations of formaldehyde (Gabel & Ponnamperuma, 1967). Kaolin was shown to facilitate the condensation reaction of formaldehyde at a pH lower than that of calcium hydroxide solutions. Because of this, the sugar products obtained were stable over longer periods of time. Still, the problem of selective formation of ribose was not solved; the overall yield was later calculated by Miller to be approximately 3.8% (Miller, 1984), again after arbitrary time under arbitrary conditions. Recently, Zubay (1998) reported the use of a combination of lead and magnesium salts in the presence of formaldehyde to generate aldopentoses that constituted 30% of the total product. Here, the lower basicity and solubility of lead and magnesium hydroxide was exploited to moderate the formose decomposition processes. The discussion on the geochemistry of lead was only limited to a list of lead containing minerals.

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CHAPTER 2 PREBIOTIC SYNTHESIS OF SUGARS Introduction We do not know what organic molecules were present on the early Earth. We may, however, look at compounds in the interstellar medium (ISM), within meteorites, in comets, and in other solar system bodies, to provide a clue. No direct information indicates what fraction of interstellar and cometary compounds would be delivered to early Earth in unaltered form. It is likely that some would be transformed in icy bodies, especially by high energy particles and photons (Bernstein et al. 2002). It is also known that Earth-based chemistry would influence the composition of material. Hydrogen cyanide (HCN), is generated by example, upon comet impact, by an unknown mechanism. At this point in our development of knowledge of the chemistry of the solar system, it is pragmatic to assume a set of compounds such as those shown in Figure 2.1 as our starting point. How might the molecules in Figure 2.1 be transformed on Earth to give ribose in a stable form ? It is clear that the prebiotic soup would be exposed to rocks and minerals that have some solubility in water at atmospheric pressure (these are found on modern Earth as evaporates). In this light, we re-examined the formose reaction, recognizing that both formaldehyde and glycolaldehyde are found in the ISM. In its native form, glycolaldehyde can act as an electrophile (the carbon of the C=O group) and as a nucleophile (the alpha carbon, once the 2-position proton is abstracted). As the enolate, however, glycolaldehyde can act only as a nucleophile. To facilitate the 9

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10 enolization, a cationic species that coordinates the two oxygens of the enediol is needed. The O-C=C bond angle is 120 This places the two oxygens at some distance, implying the need for a large metal ion to bridge the long O-O distance. The large Ca ++ serves this role. Indeed Ca ++ was the cationic species originally used by Butlerow in 1861, and is the most common catalyst used at present for the formose reaction. Formaldehyde can act only as an electrophile. The calcium-stabilized enolate of glycolaldehyde can react only a a nucleophile. Therefore, the reaction of formaldehyde and the calcium-stabilized enolate of glycolaldehyde is constrained to give glyceraldehyde as a pair of enantiomers. Glyceraldehyde has a 1,2 diol unit. Glyceraldehyde, however, can act intrinsically both as an electrophile (the carbon of the C=O group) and as a nucleophile (the alpha carbon, once the 2-position proton is abstracted). The ability of glyceraldehyde to act as both a nucleophile and an electrophile means that it can cross-react to form compounds that resemble tar. For example, reaction of glyceraldehyde as a nucleophile with formaldehyde gives a branched sugar lacking the 1,2-diol moiety. Tar formation is, of course, the standard outcome of the formose reaction, which has been criticized for its prebiological relevance, as noted above. When glyceraldehyde acts as an electrophile with the calcium-stabilized enolate of glycolaldehyde acting as a nucleophile, however, a pentose is the only product. Four enantiomeric pairs of diastereomeric pentoses exist: ribose, arabinose, xylose and lyxose. As it is drawn in the open chain form, it appears as if ribose can also act both as an electrophile and as a nucleophile (Figure 2.2). This would also permit it to form undesired further products.

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11 Figure 2.1. Some organic compounds detected in the ISM.

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12 It is clear, however, that these pentoses cannot do so in their ring closed form. Thus, ribose closes to give either a six-membered ring (a pyranose, both in the alpha and beta anomeric forms, about 75% of the total) or a five membered ring (furanose, both in the alpha and beta anomeric forms, about 25% of the total). We then asked: what mineral components might stabilize the glyceraldehyde against undesired reaction, while directing it towards the reaction that creates ribose? Figure 2.2. D-ribose structure. The open form of ribose contains electrophilic and nucleophilic centers. Here, we do not seek a large complexing species that can bridge the distant oxygens on an enediol, but rather a small complexing species that can bridge the short oxygen-oxygen distance on a 1,2 diol. Here the O-C-C bond angle is only 109. The most obvious complexing species for this purpose is small borate. Borate is well known to form a complex with diols, with a micromolar dissociation constant (Beseken, 1949). As a borate complex, glyceraldehyde can act as an electrophile. The C=O group is not affected. The borate complex of glyceraldehyde is not expected to enolize easily, however, under alkaline conditions. Abstraction of the 2-proton by base is discouraged by the negative charge already on boron.

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13 Interestingly, borate is expected to stabilize the cyclic form of ribose as well (Figure 2.2). The cyclic form presents two hydroxyl groups in a cis configuration. It is well known that borate complexes with 1,2-diol and 2,3-diol are specially stable. These observations generated the hypothesis that is tested in this chapter: Perhaps borate, if it were present under formose conditions, would manage the reactivity of glyceraldehyde, and stabilize ribose, the desired product. Boron is known in carbonaceous chondrites, where it is almost certainly present as borate (Zhai & Shaw, 1994). Boron is relatively scarce, relative to carbon and other light elements, however, due to the inefficiency of its synthesis in nuclear reactions. Borate is, however, excluded from many silicate minerals. For this reason, it appears in the residual melts as lava cools. Here, it is found in tourmalines, minerals that are found in many forms, including colorful forms used as gemstones. Tourmaline weathers from rocks as they are exposed on the surface to generate borate salts, which are generally modestly soluble in water. For example, colemanite is soluble in water to the extent of 0.82 g/L. As a consequence, colemanite and other borate-containing minerals are found in deserts and other dry environments, often under alkaline conditions. Here, they are known as evaporites, as they are crystallized from water as it evaporates. These evaporites form under conditions that are close to the conditions that generate pentose. Boron Chemistry and Complexation Mechanism in Polyols A clear understanding of the chemistry of boron is necessary for the evaluation of our hypothesis. Boron has five electrons, which can be assigned to an electronic configuration 1s 2 2s 2 2p x 1 In the hybrid orbital state, one electron from the 2s orbital is promoted to the 2p orbital (2s 1 2p x 1 2p y 1 ) to make a an sp 2 hybrid orbital in which each of the three electrons is located in an orbit and able to accept one electron from another

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14 element to form a covalent complex with the boron atom (i.e.: B(OH) 3 boric acid). The additional 2p electron orbit is able two hold a pair of electrons from another element, (which completes the octet around boron) this property explains the Lewis acid character of trigonal boron complexes. Bxxx...OO....O...........+++HHHBxxx...OO....O...........+++HHHO.......+H -OHBoric AcidHydroxyl ionBorate ion pKa =9.1 Figure 2.3. Lewis structure representation of boric acid and borate. Boric acid has a trigonal planar structure in which the B-O bond length is 1.37 In the tetrahedral borate ion, the B-O bond length is 1.48 which makes the hydroxyl group a better proton acceptor (more basic) and therefore a better leaving group when compared to the hydroxyl group in trigonal boron (Pizer & Tihal, 1992). The reactive form of boron (trigonal vs tetrahedral) towards complexation is dependent on the pH of the solution under study. Is generally assumed that in reactions carried out at the pK a of boric acid (and above), the reactive species is borate, while the trigonal boron is responsible for the reactivity at lower pH values. Van Duin et al. (1984) suggest that aqueous boric acid exists as an adduct with a water molecule, to give a species whose geometry is tetrahedral, but still neutral. This suggestion makes sense when explaining the reactivity of boric acid towards esterification reactions, in this way a loosely bound water molecule is easily substituted by a hydroxyl group).

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15 The complexation of boric acids and borate with dicarboxylic acids, -hydroxycarboxylic acids, diols and polyols, has been studied in detail for many years (Mazurek & Perlin, 1963; Pizer & Kustin, 1968; Davis & Mott, 1979; Van duin et al. 1984; Van duin et al. 1985; Verchere & Hlaibi, 1986; Pizer & Tihal, 1996; Ito et al. 2003). In the case of acidic ligands such as dicarboxylic acids, -hydroxycarboxylic acids and 1,2 dihydroxybenzenes, the mechanism of boronic acid complexation (at pH values lower than the pK a of the boronic acid under study) is believed to occur through the nucleophilic attack of a hydroxyl group of the ligand (1) to the trigonal boron (2) generating an associative transition state in which a proton from the entering ligand is transferred to a leaving hydroxide originally coordinate to boron (Figure 2.4). (Kustin & Pizer, 1968; Pizer & Tihal, 1992, Pizer & Tihal, 1996) BHO R O H O H C C O O H BOH R OH +HOC C HO O BHO R O O C C O +H3O+ 123 Figure 2.4. Mechanism of boronic acid complexation by acidic ligands. In boric acid (R= OH), the resulting borate monoester (3), has the potential to form a bis-substituted complex by reacting with an additional ligand molecule. In non-acidic ligands, such as diols and polyols, the predominant complexation reaction occurs with the tetrahedral borate (Figure 2.5). The mechanism of complexation to borate its not yet well understood. But it has been suggested (Pizer & Tihal, 1992) that complexation occurs in two steps, is probably associative in character, and also involves proton transfer. An associative mechanism does not necessarily imply an increase in the coordination number on boron. But even if this is the case, hypervalent pentacoordinated

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16 boron complexes are known in the literature (Yamashita et al. 2000; Lee & Martin, 1984). Therefore, a pentacoordinated transition state in borate complexation remains as a possibility. BHO R OH OH +CC H HO HO H H H BR HO O OH BR HO O O C C H H H H +BO O O O C C H H H H C C H H H + 2 H2O C H H C H OH H + H2OBOH OH O O C C H H H H + H2OA.B.H CC H HO HO H H H rdsfast Figure 2.5. Borate complexation by non-acidic ligands. A) Attack of the hydroxyl moiety of the diol to borate is rate limiting. The cyclization reaction to make the five member borate ester is fast. B) In the case of esters of boric acid at high pH, bis-substituted complexes are form by reaction with an additional diol molecule. pH dependence on the stability of boric acid esters and borate esters. 11 B NMR experiments allowed comparison of esters of boric and borate formed by diols, -hydroxycarboxylic acids and dicarboxylic acids (Van Duin et al, 1984). In general it was found that boric acid esters of -hydroxycarboxylic acids proved to be more stable than those of diols or dicarboxylic acids. Also, it was concluded that the optimal pH stability of boric acid and borate esters, can be predicted by using the charge rule: Esters of boric acid and borate in aqueous medium show the highest stability at that pH where the sum of the charges of the free esterifying species is equal to the charge of the ester.

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17 This rule is better illustrated as: Esters of boric acid are most stable at low pH where dissociation of B(OH) 3 and ionization of the bidentate ligand (L) hardly occurs (for carboxylic acids and alpha-hydroxyacids). B(OH)3 + L B(OH)L + 2H2O Dissociation of boric acid favors formation of borate esters of 1,2-diols. A pH pK a of boric acid is necessary (in other words, diols only form borate complexes at an alkaline pH). B-(OH)4 + nL B-Ln + 2n H2O In the case of -hydroxycarboxylic acids and dicarboxylic acids, pH dependent optima are involved. The borate diester of an -hydroxycarboxylic acid shows maximal stability at pH = pK a (L) : B(OH)3 + L + LB-L2 + 3 H2O The borate monoester, however, occurs preferentially at pK a (L) < pH < pK a boric acid. A similar situation applies to dicarboxylic acids at pH = (pK a1 + pK a2 )/2 B(OH)3 + LB-(OH)2L + H2O The previous rules agree with the expected behavior for a reaction with an associative transition state in which proton transfer is involved postulated by Pizer & Tihal in 1992. Differential coordination of borate to polyols. The preferentiality of borate binding to different diols and polyols was studied by measuring the association constants of different borate complexes (Van Duin et al, 1985). Experiments were done by 11 B NMR at a high pH allowing only borate ions to be the

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18 reactive species. The results of the relative stabilities of complexes obtained by these experiments can be summarized as: Tridentate borate > bidentate borate > monodentate borate complexes Increasing the number of hydroxyl groups increases the stability of borate esters. Diols in a threo-, diol conformation > terminal diol > erythro-, diol (explained by the steric interactions of the R groups (Figure 2.6), in the case of terminal diols a loss of enthropy is responsible for the difference) OH R' R OH threo-diolO O B R R' OHOH erythro-diol OH R' HO R OH R' OH R O O B R' R OHOH GTGBTGBT GE-TS GEGBEGBEWhere GE-TS GT + 1 kand GBE GBT +1G= kcal/mol Figure 2.6. Free energy diagram of threo and erythro diols and their borate complexes. Coulumbic repulsion decreases stability (introduction of carboxylate residues in the diols, creates repulsion with the negatively charged borate) syn-, diols > anti-,-diols (Figure 2.7). OH R' OH R OH R' R OH OB O OH OH R' R OB O OH OH R R' GSGBSGAGBSGBAGBAWhere GA GS-1 and GBAGBS+1.5G: kcal/molsyndiolanti-diol Figure 2.7. Free energy diagram of syn-, and anti-,-diols and their borate complexes.

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19 Interaction of boron with carbohydrates : aldoses and ketoses The interaction of boron with carbohydrate molecules has been studied in the past with special interest due to the potential applications in stereochemistry and structure determination. Also in the case of carbohydrate mixtures, chromatographic separations may be improved by having boron preferentially coordinated to specific molecules within a family of structurally related sugars. Several methods have been published to measure the association constants (K asso ) between boron and carbohydrates: Potentiometry, pH titration, ( 11 B, 13 C, 1 H) NMR, calorimetry. However, reported values in the literature differ dramatically depending of the technique used, reasons for these differences other than experimental error include: Uncertainty of the anomeric and conformational composition of carbohydrates-borate complexes at equilibrium. Stoichiometric ratio of borate(boric)-carbohydrate species is assumed to be homogeneous at a certain pH value and either consistent of borate monoester or borate diester complexes ( work by Van Bekkums group has shown that borate monoester and diesters coexist at certain pH values) Reproducible values of association constants have been obtained by using a combination of techniques. Mazurek & Perlin (1963) used thermometric measurements of vapor pressure equilibria and 1 H-NMR to determine the different complexes of borate with D-glucose, D-threose and cis-3,4-dihydroxytetrahydrofuran (Table 2.1). The most important conclusion of their work is the finding that the complexation of D-glucose to borate proceeds via a pyranose to furanose interconversion. This finding is important, since the calculation of association constants by other groups, assume that the ratio of

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20 sugar conformers in the presence of borate is the same as in the aqueous sugar without borate. Mazurek also suggested that pyranose cis-diols do not make strong borate complexes, and that a sugar-borate ratio 2:1 yields spirane complexes of the sugars under study. Verchere & Hlaibi (1987) presented the first comprehensive analysis that considers the effect of borate in the conformational equilibrium of carbohydrates and later these authors include the contribution of this effect in the calculation of the association constants. By using a combination of potentiometric titration and 11 B NMR spectroscopy, Verchere confirmed that 1:1 and 2:1 carbohydrate-borate complexes were obtained for every sugar and then calculated the stability constant for each of those complexes (Table 1.1) (Chapelle & Verchere, 1988; ibid, 1989). Structural information of the different borate complexes was also deduced from NMR studies (Figure 2.8), allowing a direct interpretation of the stability constants for the monoester and the diester General conclusions from data in Table 2.1 and Figure 2.8 included: While sugars in aqueous solution mainly adopt the 6 member ring pyranose conformation, in the presence of borate, this configuration is forced into the five member borate-furanose form. At limited amounts of borate, formation of spirane complexes is preferred over the monochelate complex. The trend in stability of spirane comples in the monosaccharides shown is : ketohexoses > aldopentoses > aldohexoses

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21 Table 2.1. Borate complexes of aldopentoses, aldohexoses and ketohexoses. n.d: not determined. Only the strong furanose-borate esters are shown. Sugar (furanose form) Donor sites B L 2 (borate ester) Stability constant(log ) B (OH) 2 L Stability constant (log ) B L 2 % Complexed as B L 2 1,4-anhydroerythritol 2,3 n.d n.d n.d -D-threose 1,2 n.d n.d n.d -D-ribose (70%) -D-ribose (30%) 1,2 2,3 2.26 4.28 95 % -D-arabinose 1,2 2.14 2.99 30 % -D-xylose 1,2 1.95 3.74 80 % -D-lyxose (60%) -D-lyxose (40%) 2,3 1,2 2.15 3.39 50 % -D-allose -D-allose 1,2 2,3 n.d 3.94.4 85 % -D-glucose 1,2 1.80 3.05 55 % -D-mannose 2,3 2.01 2.74 20 % -L-galactose 1,2 1.99 2.56 20 % -D-psicose 2,3 n.d 6 > 99 % -D-fructose 2,3 2.82 4.97 n.d -D-sorbose 2,3 < 3.5 5.75 n.d -D-tagatose -D-tagatose 3,4 2,3 n.d n.d n.d The anomeric hydroxyl posses a high reactivity towards the formation of borate complexes. In those cases where the anomeric hydroxyl is not involved in complexing, is found in a trans position to the vicinal borate-diol ring system. Within the aldopentoses, the trend in stability of spirane borate complexes follows: ribose > xylose > lyxose > arabinose This difference in stability can be explained by considering steric effect between the R 1 group: CH 2 OH and the borate ring. In 1,2-cis coordinated ribose and xylose

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22 O R2 H R1 H O HO HOB R2 O H R1 HO H H OOB O R2 H R1 HO O H HOB R2 OH H R1 O O H HOB OH R2 H R1 H H O OOB R2 O H R1 H H HO OOB 1,2--D-xylofuranose R1 = CH2OH ; R2 = H1,2--D-glucofuranose R1 = CHOH-CH2OH ; R2 = H2,3--D-sorbofuranose R1 = R2 = CH2OH1,2--D-arabinofuranose R1= CH2OH; R2 = H1,2--L-galactofuranose R1= CHOH-CH2OH ; R2 = H2,3--D-fructofuranose R1 = R2 = CH2OH1,2--D-ribofuranose R1 = CH2OH ; R2 = H1,2--D-allofuranose R1 = CHOH-CH2OH; R2 = H2,3--D-psicofuranose R1 = R2 = CH2OH2,3--D-ribofuranose R1 = CH2OH ; R2 = H2,3--D-allofuranose R1 = CHOH-CH2OH; R2 = H2,3--D-lyxofuranose R1 = CH2OH ; R2 = H2,3--D-mannofuranose R1 = CHOH-CH2OH; R2 = H3,4--D-tagatofuranose R1 = R2 = CH2OH1,2--D-Lyxofuranose R1 = CH2OH ; R2 = H1,2--D-mannofuranose R1 = CHOH-CH2OH; R2 = H2,3--D-tagatofuranose R1 = R2 = CH2OH Figure 2.8. Structures of the B-L 2 spirane complex formed by monosacharides and borate. The stability constants for these compounds are tabulated in table 1.1. the borate ring is trans to the R 1 group preventing steric interactions. With xylose, the 3-OH and the R 1 group are cis destabilizing the furanose ring. In 1,2 cis coordinated lyxose and arabinose, the R 1 group is in cis position to the borate ring, creating steric interference. The arabinose complex has the 3-OH in a trans position to both R 1 and borate ring, in 1,2-coordinated lyxose the 3 groups are cis decreasing the stability of the complex. However lyxose has the ability of

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23 coordinating through hydroxyls 2,3 to borate, increasing the overall concentration of the complex. Stabilization of pentoses towards decomposition in the presence of borate Borate complexation with carbohydrates should change the reactivity of the sugars towards enolization. As mentioned before in the case of the pentoses, borate is expected to lock the pentose in a closed form, rendering the complexed sugar largely unreactive. A literature review shows that indeed this is the case. Borate protects monosacharides towards alkaline degradation and the extent of protection is proportional to the stability of the complex. (Bruijn, Kieboom & Van Bekkum, 1986). However the stabilization of aldopentoses had not been reported. Mendecino (1960) reports the isomerization D-xylose at alkaline pH in the presence of borate to yield D-xylulose (2-ketopentose). Once formed, the ketopentose was found to be stabilized by borate towards degradation at high values of pH and temperature. In the present chapter we will explore the role of borate and its minerals in the prebiotic synthesis and stabilization of the aldo-pentoses and in particular ribose. Materials and Methods Chemicals All reagents used for synthesis or as standards were purchased from Sigma-Aldrich Co., and were of their highest quality, if not mentioned otherwise. Glycolaldehyde was obtained as the dimer from ICN Biomedicals. Sodium deuteroxide was purchased from Cambridge Isotope Laboratories. Pyridine was purchased from Fluka in anhydrous

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24 quality. Calcium deuteroxide was prepared in situ from metallic calcium and deuterium oxide. 13 C fully isotopically labeled D-ribose was purchased from Omicron Biochemicals Inc. Silicon chips for desorption ionization on silicon (DIOS) analysis were obtained from Silicon Sense Inc. Enzymes Aerobacter aerogenes Type I lyophilized cells as a crude source of ribitol dehydrogenase were obtained from Sigma-Aldrich. Analytic Instrumentation Ultraviolet analysis (UV) UV analysis was performed in a Cary Varian spectrophotometer interfaced to a MS windows based computer. Gas chromatography (GC) Gas Chromatography Analysis was perform in a Perkin Elmer 1500 GC equipped with a flame ionization detector and a DB-5 capillary column. Mass spectrometry (MS) Gas ChromatographyMass spectrometry analysis was performed in a Finnigan LCQ Ion Trap GC-MS equipped with a DB-5 column. Injector temperature : 300 C, the temperature program used for the analysis was: 60 C250 C at 3.5 C/min, 250 C 300 C at 20 C/min, final temperature was held for 20 minutes. Analysis were done in collaboration with Dr. Lidia Nikole Matveeva, HPLC-ESI analysis were performed by Dr. Jodie Johnson at the University of Florida.

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25 Desorption Ionization On Silicon (DIOS-MS) analysis was performed on a Bruker Daltonics (Billerica, MA) Reflex II MALDI-TOF mass spectrometer in the negative reflectron mode in collaboration with Qian Li. NMR Spectroscopy Mercury 300 MHz. 1 H NMR spectra, referenced to the respective solvent (D 2 O). Synthetic Preparations Synthesis of colemanite Calcium hydroxide (0.74 g ; 10 mmole) and boric acid (1.84 g; 30 mmole) were added to 30 mL of Milli-Q water. The resulting suspension was stirred for an hour, and then the pH was adjusted to a final value of 12 by dropwise addition of sodium hydroxide (10 M) under stirring. Clear off-white crystals of colemanite (Ca 2 B 6 O 11 .5H 2 O) precipitated from the slurry after one hour. Synthesis of deuterated Colemanite Calcium hydroxide (74 mg ; 1 mmole) and boric acid (184 mg; 3 mmole) were added to a Fisher brand conical tube (10 mL, polypropylene) containing deuterium oxide (10 mL). The resulting solution was immersed in liquid nitrogen and lyophilized. The resulting white solid of deuterated colemanite was stored in a desiccator. Synthesis of pentoses in the presence of colemanite Glycolaldehyde dimer (6 mg, 0.05 mmol) and D,L-glyceraldehyde (15 mg, 0.16 mmol were dissolved together in an aqueous alkaline colemanite slurry (15 mL, pH: 12). The mixture was heated at 45 C for 60 minutes. The reaction was quenched by addition of Dowex W-50 resin H + form until a pH of 5.0 was obtained. The resulting clear solution was quickly filtered through a Nalgene 0.2 m filter, into a Fisher brand centrifuge tube (50 mL) and immediately immersed into liquid nitrogen until complete

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26 solidification of the solution. Lyophilization yielded a white solid powder which was dissolved in anhydrous methanol (5 mL) and subjected to rotary evaporation under high vacuum (repeated 3 times) yielding a white solid (18 mg, 88 % mass recovery of the total carbon input). Synthesis of pentoses in the presence of calcium hydroxide Glycolaldehyde dimer (6 mg, 0.05 mmol) and D,L-glyceraldehyde (15 mg, 0.16 mmol were mixed and dissolved into an aqueous alkaline calcium hydroxide slurry (0.74 g, 15 mL, pH: 12). The mixture was heated at 45 C for 60 minutes (the solution turned clear brown after 20 minutes). The reaction was quenched by addition of Dowex W-50 resin H + form until a pH of 5.0 was obtained. The resulting clear brown solution was quickly filtered through a Nalgene 0.2 m filter, into a Fisher brand centrifuge tube (50 mL) and immediately immersed into liquid nitrogen until complete solidification of the solution. Lyophilization yielded a dark brown syrup with an strong odor reminiscent of caramel. The syrup was dissolved in anhydrous methanol (5 mL) and subjected to rotary evaporation under high vacuum (repeated 3 times) the syrup remained unchanged after this treatment and it was not possible to obtain a solid even after high vacuum exposure (19 mg, 91% mass recovery of total carbon input). Derivatization of pentoses for gas chromatography analysis Derivatization of pure samples of pentose was usually achieved by adding anhydrous pyridine (400 L) to the corresponding solid pentose (5 mg, 33 nmol) followed by addition of N,O-bis(trimethylsilyl)trifluoroacetamide (100 L) under Ar atmosphere, the derivatized samples were used as standards for GC analysis.

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27 Pentoses resulting from synthetic procedures, were derivatized by addition of anhydrous pyridine (800 L) and of N,O-bis(trimethylsilyl)trifluoroacetamide (200 L ) under Ar atmosphere. Sugars Degradation Experiments Sugar decomposition in the presence of calcium deuteroxide Calcium deuteroxide (16.5 mg) and sodium benzoate (6.1 mg) were added to a 2.5 mL Eppendorf tube containing 1.5 mL of deuterium oxide. This solution was vortexed and then equilibrated at room temperature for 30 min (pD: 12). After equilibration, the tube was immersed into a 4 C ice-water bath. An aqueous solution of the corresponding pentose (D-arabinose, D-lyxose, D-ribose, D-xylose and D-ribulose; 500 L, 1.0 M) was transferred by pipette to the ice-cold solution and vortexed for a period of 1 min. An aliquot (900 L) of the resulting solution was then removed and used as the sample for the 1 H NMR degradation experiment. Pentose degradation was measured by calculating the ratio of the integral for the sharpest signal corresponding to the anomeric proton (hydrogen in C-1) over the value of the internal standard integral (sodium benzoate). Half-life for the decomposition of each pentose was calculated by plotting the mentioned ratio versus time. Sugar decomposition in the presence of calcium deuteroxide and synthetic colemanite Calcium deuteroxide (16.5 mg), deuterated colemanite powder (83 mg), and sodium benzoate (6.1 mg) were added to a 2.5 mL eppendorf tube containing 1.5 mL of deuterium oxide. This solution was vortexed and then equilibrated at room temperature for 30 min (pD adjusted to 12 by sodium deuteroxide addition when necessary). After equilibration, the tube was immersed into a 4 C ice-water bath. An aqueous solution of

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28 the corresponding pentose (D-arabinose, D-lyxose, D-ribose, D-xylose and D-ribulose, 500 L, 1.0 M) was transferred by pipette to the ice-cold solution and vortexed for a period of 1 min. An aliquot (900 L) of the resulting solution was then removed and used as the sample for the 1 H NMR degradation experiment. Pentose degradation was measured by calculating the ratio of the integral for the sharpest signal corresponding to the anomeric proton (hydrogen on C-1) over the value of the internal standard integral (sodium benzoate). Half life for the decomposition of each pentose was calculated by plotting the mentioned ratio versus time. Enzymatic Assays Ribitol dehydrogenase assay An aliquot (1 mL) of the reaction mixture between glyceraldehyde + glycolaldehyde (with/without boron) was removed after 60 min and treated with a sodium borohydride solution (100 L, 0.06g/mL in cold water) followed by incubation at room temperature for 30 min. This solution was then acidified to a final pH of 5.0 by addition of aqueous HCl (200 L, 3 M) and passed through a short column containing Dowex-50 resin H + form. An aliquot (650 L) of the eluent was lyophilized and the resulting solid was dissolved in anhydrous methanol (650 L) and subjected to rotary evaporation under high vacuum (repeated 2 times). The white solid obtained after methanol evaporation was dissolved in potassium phosphate buffer (200 L, 1.0 M, pH: 10) and then aqueous sodium borate ( 10 L, 19 mM) was added. The enzymatic reaction was initiated by addition of nicotinamide adenine dinucleotide (NAD + 15 L, 100 mM in water), and of Aerobacter aerogenes Type I lyophilized cells suspension (50 L 0.02 g/mL in 1 M potassium phosphate buffer) and incubated for 60 min at 30 C. After incubation, the reaction was placed on an

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29 ice bath for 5 minutes, and then centrifugated to remove cell debris. The presence of 2-pentulose in the resulting solution was confirmed by the cysteine-carbazole test. Cysteine-carbazole test An aliquot (100 L) of solution from the ribitol dehydrogenase assay, was transferred to a quartz cuvette containing a cysteine solution (20 L, 1.5% in water). sulfuric acid (600 L, 15.2 M) and carbazole (20 L, 0.12% in ethanol) were then added and mixed thoroughly. The reaction was kept at room temperature for a period of 60 min. Presence of ribulose in the mixture was confirmed by the appearance of a strong violet color (UV/vis : max = 535 nm). DIOS Analysis Preparation of PSi surfaces PSi surfaces were manufactured with HOME (H 2 O 2 -Metal) etching method: a silicon chip of 2 cm was cut from an n-doped crystalline silicon wafer and sputter coated with a thin layer of Au at an argon pressure of 40 Torr and a current of 10 mA. For the production of an array PSi chip, an aluminum mask with 4 holes (0.8 mm in diameter) was used in Au coating. The Au coated silicon chip was then immersed in an etching solution (49% HF: 30% H 2 O 2 : Ethanol = 1: 1: 1, v: v: v) for 10-20 s (Figure 2.9). Samples of sugars (0.5 L) were directly pipetted from the mother solution into the PSi surfaces. a b c Figure 2.9. Psi surface p r eparation : (a) An alum inum m a sk was placed on top of a crysta lline s i licon ch ip, ( b ) Only exp o se d areas were coated w ith Au, (c) P S i spots were produced on Au-coated regions

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30 Competition Experiments 1, 4-Anhydroerythritol (AET) vs Pentoses 1,4-Anhydroerythritol (100 L, 0.1 M, in water) and the corresponding pentose (100 L, 0.1 M) in water were mixed in a Eppendorf tube (1.5 mL) containing water (700 L) and vortexed thoroughly. An aqueous sodium borate solution (100 L, 0.025 M) was then added, and the resulting mixture was vortexed and equilibrated for 2 h at room temperature before DIOS analysis. 13 C-Ribose vs Pentoses 13 C-Ribose (10 L, 0.1 M, in water) and the corresponding pentose (10 L, 0.1 M in water) were mixed into a Eppendorf tube (500 L) containing water (70 L) and vortexed thoroughly. An aqueous sodium borate solution (70 L, 0.025 M) was then added and the resulting mixture was vortexed and equilibrated for 2 h at room temperature before DIOS analysis. Results Synthetic Preparations Synthesis of pentoses in the presence of colemanite The presence of pentoses in the reaction products after 60 minutes of incubation under formose conditions was directly confirmed by HPLC/ESI (-)-MS analysis. The dipentose-borate complex was detected as an m/z: 307 molecular ion (Figure 2.10). Analysis of a standard of di-ribose-borate complex is shown for comparison in Figure 2.11.

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31 Figure 2.10. HPLC-MS analysis of reaction mixture containing colemanite. Analysis of the products from the reaction between glyceraldehyde an glycolaldehyde in the presence of synthetic colemanite (Ca 2 B 6 O 11 .5H 2 O) at pH:12, 45 C. (top) ESI-MS indicates the presence of the pentose-borate dimeric complex ion MW: 307. (middle) MS/MS fragmentation pattern of the 307 ion/peak. Fragmentation is consistent with that observed from the standard. (bottom) HPLC-MS trace of reaction mixture showing only various isomeric pentoses. Figure 2.11. Detection of ribose-borate comples by ESI (-) ion mode.(electrospray ionization negative ion mode). Gas chromatography analysis of the trimethylsilyl derivatives of the reaction products is shown in Figure 2.12. Peaks were assigned by coinjection of authentic standards, using retention time values (Table 2.2).

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32 Figure 2.12. GC trace of the reaction mixture containing colemanite. Analysis of the silylated products (after 60 min) from the reaction between glyceraldehyde and glycolaldehyde in the presence of synthetic colemanite(Ca 2 B 6 O 11 .5H 2 O) at pH:12, 45 C. Peak assignation: (1) Lyxose/Arabinose, (2) Lyxose/Arabinose, (3) Ribose, (4)Xylose, (5) Xylose. Anomeric forms of the sugars are not assigned. Notice the absence of peaks corresponding to compounds eluting after 5. Table 2.2. Retention times of trimethylsilyl derivative of pentoses by GC analysis. Pentose Retention Time (min) Intensity ratios Arabinose 28.34; 29.49 10:1 Lyxose 28.17; 29.81 1:1 Ribose 30.04 1 Xylose 28.79; 31.97; 34.22 1:2:5 Synthesis of pentoses in the presence of calcium hydroxide A large portion of insoluble material for this reaction remained insoluble in pyridine after trimethylsilylation at 60 min. Gas chromatography analysis showed an

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33 heterogeneous composition with a low signal intensity (data not shown). Analysis of the reaction products after just 20 minutes is shown in Figure 2.13. Peaks were assigned by coinjection of authentic standards and retention time values. Figure 2.13. GC trace of the reaction mixture containing Ca(OH) 2 Analysis of the silylated products (after 20 min) from the reaction between glyceraldehyde and glycolaldehyde in the presence of Ca(OH) 2 at pH:12, 45 C. Peak assignation: (1) Lyxose/Arabinose, (2) Lyxose/Arabinose, (3) Ribose, (4) Xylose, (5) Xylose + other. Anomeric forms of the sugars are not assigned. Compounds to the right, eluting after peak, 5 include tar. Sugar Degradation Experiments Sugar decomposition in the presence of Calcium Deuteroxide Alkaline degradation of sugars in the presence of calcium deuteroxide occurred at a high rate. After a few minutes of reaction, the sugar solutions turned clear brown, and after 1 h some precipitate formed in the NMR tube. The calculated rates of

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34 decomposition (Table 2.3) were obtained by measuring the value of the integral of the signal associated with the anomeric proton and comparing this value with that of the internal standard (Figure 2.14). Figure 2.14. Incubation of ribose in Ca(OH) 2 solution. Decomposition was monitored by 1 H NMR at 25 C. Note loss of signals at (ppm): 4.85, 4.95, 5.4, corresponding to the signal of H-1 from various anomeric forms of ribose. Sugar decomposition in the presence of Colemanite Addition of colemanite had a dramatic effect in the rate of alkaline degradation. All of the sugars were stabilized towards decomposition (when compared to the samples without colemanite). Sugar solutions remained clear after several hours and in the case of ribose it remained unchanged for several days (Figure 2.15). The calculated rates of decomposition are shown in Table 2.3.

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35 Figure 2.15. Incubation of ribose in the presence of Ca(OH) 2 + colemanite. Degradation was monitored by 1 H NMR at 25 C. Note that signals corresponding to H-1 of various anomeric forms of ribose at (ppm): 5.3, 5.5, 5.7, remain unchanged even under incubation for 5 days at high pH. Table 2.3. Half life of pentoses under alkaline conditions determined by 1 H NMR estimated by loss of selected NMR signals (see appendix 1). Pentose Half life (min): Ca(OD) 2 Half life (min): Ca(OD) 2 + Colemanite D-ribose 291 2700 D-arabinose 144 259; 716; 331 L-arabinose 124 380; 274 D-lyxose 69; 177 1382 L-xylose 66 572; 258 L-Lyxose 184 1201 ribulose 14 2028 Enzymatic Assays The presence of ribulose after ribitol dehydrogenase treatment was confirmed by the cysteine-carbazole test. A dark violet color was observed when samples of the reaction between glyceraldehyde and glycolaldehyde were analyzed by this assay.

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36 DIOS analysis: Competition Experiments. 1, 4-Anhydroerythritol (AET) vs Pentoses The different complexes detected by DIOS are depicted in Figure 2.16. A typical spectra obtained by DIOS analysis is shown in Figure 2.17. O OOO OOB O OOO OOB HO OH O OOO OOB HO OH OH HO m/z : 215m/z : 261m/z : 307 Figure 2.16. Anhydroethrythritol (AET)-pentose borate ions detected by DIOS. Pentose borate complex is depicted through C2 and C3 (cis diol). However, binding through C1-C2 (cis diol) is also known to occur when possible. Each pentose was analyzed independently, in competition with 1,4-anhydroerythritol (AET). The relativies intensities reported for each detected ion (Figure 2.18) were calculated using equation (1) : % Relative intensity = 100 x Intensity of individual ion : equation (1) Intensity of all detected ions Error bars represent the standard deviation calculated from a set of 10 different measurements (for complete set of spectra see appendix 2). 13 C-Ribose vs Pentoses Fully isotopically labeled ribose (13C-ribose, 98.8% labeled) was used for competitions experiments because the difference in mass allows resolution of peaks with non-isotopically labeled pentoses. The different compounds detected in this experiment

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37 are shown in Figure 2.19. A typical DIOS spectra of this experiment is shown in Figure 2.20. Figure 2.17. DIOS spectra of competition experiment D-arabinose vs AET. Boron (10 mM) was set as the limiting reagent to favor spirane formation. Pentose vs 1,4 -Anhydroerythritol0.0%10.0%20.0%30.0%40.0%50.0%60.0%70.0%m/z 215 m/z 261 m/z 307Percentage of relative intensity xylose arabinose lyxose ribose riblulose Figure 2.18. Competition experiments between the different pentoses and AET. Relativive intensities represent the molar fraction of the different borate complexes in solution.

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38 O13C13C13C13C OOO13C13C13C13C OOB 13C HO OH 13C OH HO m/z : 317O13C13C13C13C OOO OOB 13C HO OH OH HO m/z : 307O OOO OOB HO OH OH HO m/z : 312 Figure 2.19. 13 C, 12 C-D-ribose borate ions detected by DIOS. Pentose borate complexes detected in the competition experiment using isotopically labeled ribose. Figure 2.20. DIOS spectra of competition experiment 13 C ribose vs 12 C ribose. Boron (10 mM) was set as the limiting reagent to force the formation of spirane compounds. Each pentose was analyzed independently in competition for boron binding against 13 C-ribose. Figure 2.21 shows the ratio of 13 C-ribose/pentose in equilibrium detected by DIOS. Error bars represent the standard deviation calculated from a set of 10 different measurements.

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39 13C-d-ribose vs pentose 0.001.002.003.004.005.006.007.008.00peak intensity ratio of m/z 307 : m/z 317 ribose lyxose arabinose xylose xylulose ribulose Figure 2.21. Ratio of borate complexes of 13 C-ribose vs pentoses. 12 C-D-ribose was included in the analysis to test the isotope effect in ionization-vaporization efficiency, however, no such effect was observed during the experiment. The slight deviation from the value of 1.0 in D-ribose is due to the presence of 1.1% unlabeled carbon in the 13 C-D-ribose used for the competition experiment. Discussion Reaction intermediates of the formose reaction were used as starting materials for the synthesis of pentoses under alkaline conditions. When the reaction was made in the presence of calcium hydroxide, a simple visual inspection gave some clue of the composition. The solution which was originally transparent, turned clear brown after 20 minutes and after one hour dark brown, which, in the case of sugars, is an indicator of decomposition with branching (know as caramelization or browning). Gas chromatography analysis showed that pentoses were indeed present at the early stages of the reaction in addition to other uncharacterized carbonaceous material as seen in Figure

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40 2.13 (probably tetroses, hexoses and branched sugars). However, further incubation of the alkaline mixture, showed a decrease in the total amount of detectable carbon caused by degradation of pentoses to higher molecular weight material that it is not volatile and therefore undetectable by the GC analysis. These results agree with the expected reactivity of the formose mixture. When boron was introduced as the mineral colemanite into the reaction mixture, visual inspection indicated that the decomposition process was either absent or slow. Browning was not observed even under long incubation periods. GC analysis of the reaction after a period of one hour shows the presence of pentoses as the major products, which indicates that boron is playing an active role in the control of the regiochemistry, and also stabilizing the pentoses product once these are formed. Because an equimolar amount of the fourth aldopentoses was detected, indicates that there was not stereoselectivity in the condensation reaction in the presence of borate. A proposed mechanism that will account for these observations is depicted in Figure 2.22. The mechanism depicted above, explains the regiochemistry obtained from the reaction between glyceraldehyde and glycolaldehyde. But it does not address the stabilization of the pentoses by boron, once formed, towards alkaline degradation. Degradation experiments in the presence of colemanite, showed that all the aldo-pentoses are stabilized by boron, following the trend: ribose > lyxose > arabinose > xylose. Shifts in the NMR signal corresponding to the anomeric proton of the monosaccharides are consistent with previous literature reports indicating that boron coordinates to pentoses, by favoring the formation of a closed furanose conformation.

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41 CC H OH C O H OH H H CC H O C O H O H H CC H O C O H O H H B electrophilic centernucleophiliccenteracidichydrogenstill electrophilicNo longer acidic due toanionic borateNo longer nucleophilicH O H HO H Ca(OH)2O HO H H Ca++ GlyceraldehydeGlycolaldehyde Borate electrophilic center H HO OH H OH H O H OH H H HO OH H O H H HO H HO OH H O H OH H OH H OH H O H H OH H H OH H H OH H OH H H D-riboseD-arabinoseD-xyloseD-lyxose Figure 2.22. Suggested mechanism for pentose formation. The aldol condensation reaction between glyceraldehyde, and glycolaldehyde in the presence of borate is shown. (Pentose products in the Fischer representation are shown only in the D-form for simplicity). Because the rate of degradation of these sugars is proportional to the amount of pentose in the open form, the half life should be directly proportional to the amount of pentose-borate ester in each case. Boron coordinates pentoses in different fashions to make spirane complexes; this is of course an obvious consequence of the spatial orientation of the different cis-hydroxyl groups. DIOS experiments were focused in understanding these binding preferences. A qualitative description of the preferential binding of borate for the different pentoses shows that the preferential order for binding is: ribose > lyxose > arabinose > xylose. This information confirms that the stabilization of pentoses (increased half life)

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42 under alkaline conditions is a direct consequence of borate-pentose complex formation. This synthesis of pentoses in the presence of the mineral colemanite is therefore plausibly prebiotic. Indeed, in the presence of borate, and given that ribose is the first compound in the formose product progression that offers a non-aldehydic cyclic form with unhindered cis-diols, the formation of ribose appears to be the natural consequence of the intrinsic chemical reactivity of compounds available from the interstellar medium under alkaline, calciferous conditions. As these conditions are not excluded from the early Earth, it is also not possible to exclude the availability of pentoses at the time when life originated. This example of how minerals can productively control organic reactivity reminds us of the fact that prebiotic chemistry is occurring on a planet, in the context of a larger geology. Minerals must be considered as we constrain models for the origin of life.

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CHAPTER 3 CATALYSIS AND THE RNA WORLD Introduction Ten years ago, Szostak, Joyce, Ellington, and others applied in vitro selection (IVS) to libraries of nucleic acids to extract nucleic acids that catalyze simple reactions, such as RNA ligation (Bartel & Szostak, 1993) and RNA cleavage (Breaker & Joyce, 1994). This work opened the possibility of using in vitro selection to ask quantitative questions about the performance of these catalysts. This includes questions concerning the mechanism of specific nucleic acid enzymes, as well as broader questions, such as how functional behavior is distributed in nucleic acid sequence space, and whether adding chemical functionality to nucleic acids, either by modifying the nucleobases or by adding cofactors, can enhance the catalytic potential of a nucleic acid library (Breaker, 2000; Perrin et al, 2001). Before broader questions can be addressed using in vitro selection, it is necessary to explore some of the specific features displayed by many nucleic acid catalysts that have emerged from selection experiments. For example: (a) Partial conversion. Many studies of individual nucleic acid catalysts report that their reaction goes only partially to completion. Hammerhead ribozymes, for example, are frequently reported to cleave only 40 60% at plateau (Stage-Zimmermann & Uhlenbeck, 1998) (Kore et al, 2000). DNAzymes with ribonuclease activity similar to the ones studied here are also frequently reported with cleavage plateaus of 25 65% (Perrin 43

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44 et al, 2001; Geyer & Sen, 1997; Faulhammer & Famulok, 1996; ibid, 1997). The same is seen for DNAzymes with DNase activity (Carmi et al, 1996). (b) Intramolecular versus intermolecular reactions. Many reactions catalyzed by nucleic acid catalysts are selected to be intramolecular, making the term catalyst technically incorrect; the catalyst is not regenerated. Many of the intramolecular reactions have analogous intermolecular processes that are truly catalytic, however, and these are often accessible both during the selection and in the subsequent kinetic analysis. (c) Loss of the most active catalysts during the set-up. In many selection schemes, a library of catalysts must be synthesized and folded before the selection step begins. This leads to the possibility that active catalysts in a pool will be lost before the selection system can extract them. (d) Michaelis-Menten kinetic behavior. Many nucleic acid catalysts bind their substrate in a reversible step prior to the step where chemical bonds are made or broken. This should generate saturation kinetics similar to those seen in protein enzymes. It is, of course, impossible to address these questions for the general reaction catalyzed by all nucleic acid catalysts. Rather, these questions must be addressed for individual nucleic acid catalysts working on specific reactions. Such studies follow the tradition in physical organic chemistry, where many detailed studies of specific reactions eventually generate a body of literature that addresses broader issues in catalysis. For this work, we began with work of Breaker and Joyce, who selected for DNA enzymes that catalyze the cleavage of a ribonucleotidyl-3->5-deoxyribonucleotide linkage in an oligodeoxyribonucleotide (Breaker & Joyce, 1995). The reaction almost certainly proceeds with the attack on the phosphorus electrophilic center via the

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45 ribonucleotide 2-hydroxyl group (Figure 3.1). An adaptation of the Breaker-Joyce selection procedure led to several new catalysts for this reaction (Figure 3.2). One of these DNAzymes, designated 614, was studied in detail to address some of the issues outlined above. NN NN NH2 O O O P OO O O-H 5'-constant region Biotin StreptavidinsupportNN NN NH2 O O O P OO O-5' constant region Biotin Streptavidinsupport 3'-constant region 3'-constant region HO Figure 3.1. In vitro selection experiment representation. A library of DNA oligonucleotides containing an internal adenine riboside and a 40-nt random region (represented in red) is attached to a solid support. The DNA-catalyzed ribonuclease reaction proceeds with the attack on the phosphorus electrophilic center by the ribonucleotide 2-hydroxyl group, cleaving the phosphodiester backbone and releasing the catalytic portion of the DNAzyme from the solid support. Materials and Methods Preparation of Precursor DNAzymes via PCR (Maniatis et al, 1982) DNAzymes were prepared by PCR amplification of the template (synthesized by Integrated DNA Technologies (Coralville IA), or from a clone) using a catalytic strand primer (cat+ribose or cat+deoxyribose, see Table 3.1) and a complementary strand primer (compl, compl+5P, or compl+tail). For trans cleaving assays, DNAzymes were generated without the internal ribose using the cat+deoxyribose primer, identical to the

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46 Figure 3.2. Sequence of the initial library and DNAzymes isolated from the in vitro selection. (A) The initial library was based on the sequence used by Breaker and Joyce, with an internal adenine riboside incorporated at position 28 to provide a cleavable linker. Two nucleotide substitutions were introduced to eliminate one of the clamps that were designed to hold the substrate and catalyst portions of the molecules in the library together. The original two nucleotides in the Breaker Joyce sequence are shown in bold above and below the sequence used in this study. Base pairing that could form binding clamps are highlighted in grey. (B) DNAzymes isolated from multiple rounds of IVS. Sequences 614, 62/615, 616, and 625 represent the major sequences classes initially cloned from the seventh round of the initial IVS. Sequences variants of these major classes were isolated from additional rounds of selection and are grouped according to sequence class (only variations from the prototype sequence are shown). Colored boxes are shown to highlight common motifs. Sequences isolated more than once are shown with the number of isolates identified in parenthesis following the name.

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47 cat+ribose primer except that the single ribo-adenosine in the cat+ribose was replaced with a 2-deoxyribose-adenosine. To obtain single stranded catalysts, DNAzymes were produced using a catalytic strand primer and either a 5-phosphorylated complementary strand primer (compl+5P, for use with lambda exonuclease) or a complementary strand primer with an 15-nt poly-deoxyriboadenosine tail appended to the 5 via an 18-atom hexaethyleneglycol-based linker (compl+tail, for use in asymmetric PCR). This linker prevents polymerase read-through. Templates all had the same 5-constant region and 3-constant region to which complementary and catalytic strand primers bind, separated by 40 nucleotides. Ribose-614, deoxyribose-614, and the library templates were synthesized and PAGE purified by IDT DNA technologies. Typical conditions for a PCR contained up to 1 ng template, 100 nM catalytic strand primer, 100 nM complementary strand primer, 100 M dNTPS, 10 mM KCl, 20 mM Tris-HCl (pH 8.8), 10 mM (NH4)2SO4, 2 mM MgSO4, 0.1% Triton X-100, 3-4 units polymerase (Taq or Vent exo-), and alpha-32P-CTP (10 Ci, for internally labeled samples), for a total volume of 100 L. The PCR amplification cycle consisted of an initial incubation (3 min, 96C) followed by 20 PCR cycles of (45 sec at 96C, 45 sec at 50C, and 2 min at 72C). PCR for in vitro selections (IVS) used Vent exopolymerase (3-4 units) and excess catalytic strand primer (400 nM), with up to 40 cycles of PCR in the early rounds. Preparation of Single-stranded DNAzymes Double stranded DNAzymes generated via PCR with a 5-phosphorylated complementary strand primer (compl+5P) were converted to single-stranded DNA by digestion of the complementary strand using lambda exonuclease (which is specific for

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48 the 5-phosphorylated strand of double stranded DNA). The DNA was recovered by EtOH precipitation, resuspended in exonuclease solution (25 L, 5 units lambda exonuclease, 67 mM glycine-KOH (pH 9.4), 2.5 mM MgCl2, 50 g/mL BSA, for a 100 L PCR). Samples were mixed and incubated (37C 30 min). Reactions were terminated with formamide stop dye (1 mg/mL xylene cyanol, 1 mg/mL bromophenol blue, 10 mM EDTA, in 98% formamide) and heating (80C 10 min). The single-stranded products were resolved by 8% PAGE/urea, and full-length ssDNA products excised. Gel slices of individual samples were crushed with individual disposable mortars and were eluted in buffer (350 L, 500 mM NH4OAc, 0.1 mM EDTA, 0.1% SDS, pH 7) overnight. Gel-purified samples were extracted with phenol/CHCl3/isoamyl alcohol (25:24:1), then with CHCl3/isoamyl alcohol (24:1), and the resulting DNA was precipitated in NH4OAc and EtOH. Single stranded DNAzymes were also purified via asymmetric PCR using a complementary strand primer with a 15-nt poly-deoxyriboadenosine tail connected to the 5-end of the primer by a C18 linker (compl+tail), at which Vent and Taq polymerases terminate. Complementary strand molecules generated by extension of a complementary strand primer containing the 5-tail are longer than the full-length catalytic strand, and were separated from the catalytic strand by PAGE/urea (8% acrylamide). Gel-purified samples were excised, extracted and recovered as before. 5-End Labeling of DNA Single-stranded DNA (20 mM) was 5-labeled with gamma-32P-ATP (20 Ci) using T4 polynucleotide kinase (10 units) in Tris-HCl buffer (70 mM, pH 7.6, 10 mM MgCl2, 5 mM dithiothreitol, final volume 10 L, 30 min 37C). An equal volume TE

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49 was then added, and the mixture heated (20 min 70C). End-labeled DNA was separated from unincorporated nucleotides by spinning through a G-25 column (600 g, 3 min). DNAzyme Kinetic Assays DNAzymes and substrates were isolated via EtOH precipitation and re-suspended in HEPES buffer (50 mM, pH 7). Their concentration was estimated by Cherenkov counting. Samples were diluted to twice the desired final concentration with additional HEPES buffer. For trans assays, enzyme and substrates were typically mixed (unless otherwise noted) and diluted to twice the desired final concentration. Samples were then mixed with equal volume 2X reaction buffer (typically 2M NaCl, 2 mM MgCl2, 50 mM HEPES pH 7). Mixtures were then heated (96C, 3 min), and slowly cooled to 23C (over 10 min) collectively termed slow cooling. The initial time zero point was the time at which the sample completed the slow cooling. Unless otherwise noted, reactions were run at 25C and terminated at various times by diluting an aliquot of the reaction into the formamide stop dye, followed by freezing (-20C). Samples were resolved using 8% PAGE/urea and the product cleaved quantified using a Bio-Rad phosphorimager. Data was analyzed using the GraphPad Prism 3.0a software package. Cloning and Sequencing DNAzymes Single-stranded DNAzymes were converted to duplex DNA by PCR amplification, usually with only the complementary strand primer compl (96C for 3 min; followed by 3 cycles of 45 sec at 96C, four min ramp cool to 50C plus 45 sec at 50C, and 1 min at 72C; followed by 7 min at 72C). When cloning cleaved and uncleaved 614, the desired single-stranded DNA (either cleaved product or uncleaved reactant) was first isolated via PAGE/urea. PCR of the unpurified 614 reaction mixture containing both cleaved and uncleaved 614 with only compl primer yielded clones only of uncleaved 614

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50 molecules; cleaved 614 molecules were therefore cloned by PCR amplifying gel purified cleaved fragments using the compl primer with the 5-truncated catalytic primer cat.nt29-45. Fresh double stranded PCR products were cloned using the TOPO TA Cloning System (InVitrogen) and plated on agarose plates containing ampicillin. Transformed cells were given only 15-30 min to recover in antibiotic-free media prior to plating on antibiotic containing plates (to prevent recently transformed clones from doubling prior to plating, favoring isolation of only unique species from the original pool). Clones were transformed into the TOPO TA Cloning Vector and DNA was prepared from individual clones using the alkaline lysis protocol. Clones were sequenced using the 1224 plasmid sequencing primer (3.2 pM, 5CGCCAGGGTTTTCCCAGTCACGAC) with 300-500 ng plasmid DNA template, 2X final concentration Big Dye reaction buffer and 2 L Big Dye Terminator Sequencing mix in a final volume of 10 L. Samples were overlaid with mineral oil and amplified by PCR (25 cycles, 96C for 30 sec, 50C for 15 sec, and 60C for 4 min). Sequencing samples were analyzed on an Applied Biosystems Prism 310 Genetic Analyzer. Sequencing results were confirmed by examining the chromatograms manually using the Sequencher software package. In vitro Selection DNAzyme libraries for the first round of selection were prepared by a single cycle of run-off PCR using the library template (5-GTGCCAAGCTTACCGTCAC-N40-GAGATGTCGCCATCTCTTCC (where N indicates equal molar concentrations of A, T, G, and C) and cat+ribose primer. Run-off reactions volumes ranged from 3 to 20 mL, each containing 1 ng/L library template, 100 nM catalytic strand primer, 100 M

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51 dNTPs, 10 mM KCl, 20 mM Tris-HCl (pH 8.8), 10 mM (NH4) 2SO4, 2 mM Mg2SO4, 0.1% Triton X-100, 20 units/mL Vent exo-, and 4.3 Ci/mL alpha-32P-CTP. An aliquot (1 mL) of each sample mixture (minus polymerase) was placed in Eppendorf tubes (1.5 mL), heated (96C, 8 min, then slowly cooled to 55C over 30 min). Polymerase was added, and the samples were incubated (72C, 15 min). DNA was recovered by EtOH precipitation with NH4OAc and glycogen as a carrier by storing overnight at -80C and then centrifuging in Corex tubes (16,000 g, 40 min, 4C). The EtOH was removed and the pellet re-suspended in water (200 L). DNA was recovered by precipitation a second time with EtOH (NH4OAc) and centrifugation (16,000 g, 20 min, 4C). After removing the EtOH, the pellet was resuspended in 1X binding buffer (1 M NaCl, 1 mM EDTA, 50 mM HEPES pH 7) and bound to a streptavidin column. The unbound material was removed by flushing the column with wash buffer (50 mM HEPES pH 7). The complementary strand was removed by washing quickly with 0.2M NaOH, followed again by wash buffer. The cleavage reaction was initiated by eluting the wash buffer and replacing it with reaction buffer (1M NaCl, 1 mM MgCl2, 50 mM HEPES pH 7). The cleaved products were eluted from the column after two hours incubation. The eluted material was used in a PCR with the cat+ribose and compl+5P primers to generate material for the next round of selection.

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52 Table 3.1. Name, sequence, and description of oligonucleotides used throughout this chapter. Names used in text Description ribose-614 5 CTGCAGAATTCTAATACGACTCACTATrAGGAAGACATGGCGACTCTCACATCATGCGAGCACACGCAATAGCCTGATAAGGTTGGTAGTGACGGTAAGCTTGGCAC A DNAzyme isolated from the in vitro selection in this study. It catalyzes the cleavage of the ribo-adenosine embedded within the cat+ribose primer, either as a part of its own sequence, or in trans. ribose-614C72T A mutant of ribose-614 in which the cytosine at position 72 is replaced by a thymidine, resulting in a 30-fold lower rate of cleavage at 100 nM. library template 5' GTGCCAAGCTTACCGTCAC(n40)GAGATGTCGCCATCTCTTCC The complementary strand template containing a region of 40 nucleotide randomized region flanked on both sides with constant regions, one complementary to cat+ribose, and the other identical to compl. ribose-library A random library generated by PCR amplification of the library template with cat+ribose and compl primers. Without selection, this library is predominately inactive. Selection of this library gave rise to DNAzyme 614, and others. All sequences contain the cat+ribose primer and therefore have the potential to function as substrates for 614 cleavage. ribose-lib62 An individual clone isolated from a the ribose-library. This sequence has no intrinsic ribonuclease activity of its own, but contains the cat+ribose primer and therefore can function as a substrate for 614 cleavage. deoxyribose-614, deoxyribose-lib62, deoxyribose-614C72T The same sequences as ribose-614, ribose-lib62, and ribose-614C72T but with the ribo-adenosine at position 28 replaced by a deoxyribose-adenosine (via PCR amplification with cat+deoxyribose primer). deoxyribose-614 is still capable of cleaving the ribo-adenosine embedded within the cat+ribose primer, but only in trans.

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53 Table 3.1 Continued Names used in text Description compl-614 The complementary strand to deoxyribose-614. cat+ribose 5CTGCAGAATTCTAATACGACTCACTATrAGGAAGACATGGCGACTCTC Primer used to generate full length molecules from template, thus incorporating a ribo-adenosine (rA) at position 28. This oligo has no catalytic activity of its own and was therefore used as a substrate in trans cleavage assays. cat+deoxyribose The same oligonucleotide sequence as cat+ribose but with a deoxyribonucleotide replacing the rA. This primer was used to generate full length DNAzymes without the ribo-adenosine moiety. DNAzymes incorporating cat+deoxyribose can not be cleaved and therefore act as a true enzyme. compl 5GTGCCAAGCTTACCGTCA This primer is complementary to the 3-end of the catalytic strand of the full length DNAzymes used in this study, and is used to generate double stranded DNAzymes via PCR. compl+5P This primer is the same sequence as the compl primer but with a phosphate attached to the 5-position. This primer is used to generate double stranded DNAzyme via PCR, followed by degradation of the complementary strand by lambda exonuclease (an enzyme that specifically degrades 5-phosphorylated DNA). compl+tail This primer is the same sequence as the compl primer but with a 15-nt poly-adenosine attached to the 5-position via an 18-atom hexaethyleneglycol-based linker. Polymerase can not efficiently read through this linker, so PCR using this primer genearates a double stranded DNAzymes with a complementary strand that is 15-nt longer than the catalytic strand, thus allowing purification of the catalytic strand via PAGE/urea. chase The cat+deoxyribose oligo was used as the chase substrate for cleavage assays with 614 functioning as the enzyme.

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54 Table 3.1 continued Names used in text Description 28-nt product The 28-nt product generated from cleavage of ribose-614 (106-nt). 78-nt product The 78-nt product generated from cleavage of ribose-614 (106-nt). cat.nt29-45 5-GGAAGACATGGCGACTCTC A 5-truncated version of cat+deoxyribose (containing nucleotides 29 45) used to generate dsDNA of the 78-nt ribose-614 cleavage product for subsequent cloning. Results In vitro Selection An in vitro selection experiment to select for a DNAzyme having ribonuclease activity was conducted using a procedure adapted from Breaker and Joyce (Breaker & Joyce, 1995). A library was constructed containing a 5-primer constant region and a substrate segment, which contained an internal ribo-adenosine at position 28. This was connected to a 3-primer constant region by a segment, 40 nucleotides in length, having random sequence. The 5-constant region used by Breaker and Joyce was altered to minimize a base paired clamp engineered to favor association between the substrate portion and random portion (see Figure 3.2a) of the DNAzyme. Table 3.1 presents a summary of the many oligonucleotides discussed in this chapter. A cycle of selection and amplification was applied for 7 rounds to the library. A sampling of the DNA molecules recovered was then cloned and sequenced. Several DNA molecules having distinct sequences were isolated. When inspected for catalytic activity, the molecules cleaved themselves with apparent first order rate constants ranging

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55 from 0.015 to 0.049 hr -1 (Figure. 3.2b). These rate constants are low in comparison to optimized DNAzymes obtained in the presence of divalent cation (ca. 60 hr-1 )(Santoro & Joyce, 1997; 1998), but comparable to Mg++-independent catalysts reported by Geyer and Sen (0.17 hr -1 ) and by Faulhammer and Famulok (0.006-0.024 hr -1 ). Additional rounds of selection isolated variants of the sequences initially identified in round 7. The behavior of one catalyst, termed 614, isolated from the seventh round of selection was examined. DNAzyme 614 was chemically synthesized and purified by polyacrylamide gel electrophoresis (PAGE) by Integrated DNA Technologies (Coralville IA). Large quantities of internally radiolabeled 614 were generated by PCR using this 614 template. The catalytic strand of the double stranded PCR products, termed ribose-614, was separated from its complement by two methods. In some cases, ribose-614 was generated with a 5-phosphorylated complementary strand primer (compl+5P). Following PCR, the complement was digested with lambda exonuclease. Alternatively, ribose-614 was generated using a complementary strand primer with a 15-nt poly-adenosine tail attached to its 5 end via a PEG linker (compl+tail). By increasing the length of the complementary strand, the tail allowed PAGE separation of the catalytic strand from its complement. Purified ribose-614 was resuspended in HEPES buffer, and cleavage reactions were initiated by adding equal volume of 2X reaction buffer, immediate heating to 96C for 3 min, and slow cooling to 23C over 10 min (the set-up). The reaction was then followed by gel electrophoresis. Ribose-614 (100 nM) cleaves itself at a rate of 0.015 hr -1 slow in comparison to previously isolated DNazymes, but still significantly faster than the

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56 uncatalyzed reaction (the uncatalyzed cleavage rate is negligible in 300 hours, estimated from the cleavage of ribose-library shown in Figure 3.5) Cleavage of 614 does not go completion A progress curve for ribose-614 cleavage shows that the product formation ceased after ca. 65% conversion; this is a cleavage plateau (Figure 3.3a). Such plateaus are frequently seen in selected DNAzymes and RNAzymes, but are rarely explained. We considered six possible explanations for the failure of the cleavage reaction to go to completion: 1. The complementary strand may have been incompletely removed in the set-up. This strand may inhibit the reaction, or may erroneously increase the estimate of the uncleaved product if the radiolabeled complementary strand is assigned to the reactant; 2. Part of the substrate may be missing the adenine riboside that offers the cleavable site, due to failure in the synthesis of the cat+ribose primer; 3. The reaction might be reversible, with the 65% cleavage representing the achievement of the equilibrium between uncleaved substrate and cleaved products; 4. The products of the reaction may inhibit 614; 5. Part of 614 may adopt a conformation that is inactive; 6. Part of the 614 might no longer have the correct sequence, having incurred mutation during the synthesis of the template or amplification. We examined each possibility.

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57 Figure 3.3. Ribose-614 cleavage is unaffected by the addition of 10% complementary strand. Addition of equimolar amounts of the complement greatly lowers cleavage rate and plateau. (a) Cleavage profile for ribose-614 (100 nM)(triangles), or ribose-614 (115 nM) plus complementary strand (11.5 nM)(squares). (b) Cleavage profile for ribose-614 (222 nM)(triangles), or ribose-614 (200 nM) plus equimolar complementary strand (circles). Inhibition by Incompletely Removed Complementary Strand does not Account for the Plateau The complement of 614 (termed compl-614) was added in small amounts to ribose-614 ([ribose-614] = 115 nM, [compl-614] = 11.5 nM). The initial cleavage rate and plateau were unchanged (Figure 3.3a). Similar results were seen at a lower concentration of ribose-614 (11.5 nM) plus compl-614 (1.15 nM). Experiments with equal amounts of ribose-614 (200 nM) and compl-614 (200 nM) found that the plateau was dramatically lower ( Figure 3.3b), even though the initial cleavage rate was unchanged. Similar results were seen with ribose-614 at 50 nM plus compl-614 at 50 nM. This showed that the complementary strand inhibited cleavage by 614, and the importance of removing the complementary strand prior to measurement of DNAzyme kinetics. We then asked whether the presence of complementary strand could explain the incomplete cleavage of ribose-614. Two independent procedures (exonuclease digestion

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58 and asymmetric PCR) were used to generate ribose-614 lacking its complement. Experiments with each yielded approximately the same cleavage plateau. This suggested that the plateau is not the consequence of incomplete removal of compl-614, as it is unlikely that the amounts of complement remaining following the two procedures are the same. In fact, it is difficult to imagine that any compl-614 remains following the purification using the asymmetric PCR procedure, as the complementary strand is 15-nt longer than the catalytic strand, and would almost certainly be removed by PAGE purification. To assess the amount of complementary strand remaining after exonuclease treatment, we treated ribose-614 generated via the exonuclease method with base (0.5 M NaOH, 80C, 1 h). This cleaves all of the substrate at the adenine riboside site. The amount of cleaved products was similar to that with the 5-32P labeled cat+ribose primer (85-95% cleaved for ribose-614 vs. 90-97% cleaved for cat+ribose primer following treatment with base). The incomplete cleavage of cat+ribose primer by base suggest a lack of chemical susceptibility of the primer (discussed below). Since contaminating complementary strand cannot be present in the cat+ribose primer, the ca. 5% difference between base cleavage of the cat+ribose primer and the full length ribose-614 is an estimate of the upper limit of the amount of complementary strand that might remain from incomplete degradation by exonuclease. The efficiency with which lambda exonuclease removes the complementary strand was also tested using a 5-32P labeled cat+ribose primer to synthesize full-length, double stranded ribose-614 without 5-phosphate on the complementary strand primer. The duplex PCR product was divided into two aliquots. One was subjected to standard

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59 exonuclease treatment, while the other sample was untreated. The samples were resolved by PAGE-urea to determine the amount of full-length product. Less than 10% of the original phosphorylated strand (compared to untreated sample) remained following exonuclease digest. These results rule out contamination by residual compl-614 as a major cause of incomplete 614 cleavage An Approach to Chemical Equilibrium does not account for the Plateau The failure of a substrate to be completely transformed to product may result from an approach to chemical equilibrium between substrates and products in a reversible reaction. To test for this, ribose-614 (200 nM) was incubated for 144 hours. At this time, plateau had been reached, and the reaction mixture was divided into three aliquots. One aliquot was reserved. The second was diluted 25-fold with reaction buffer. Diluting is expected to drive the equilibrium towards the cleaved product. An equal amount of a variant of ribose-614, deoxyribose-614, was added to the third aliquot. The compound deoxyribose-614 is the same sequence as ribose-614, but with an uncleavable uncleaveable adenine 2'-deoxyriboside replacing the cleavable adenine riboside at position 28. As discussed below, deoxyribose-614 is a true catalyst, acting in trans. Its addition in excess to ribose-614 ensured that catalytic activity was sufficient to see cleavage at the plateau if it was occurring. Neither treatment significantly altered the plateau, excluding reversibility as its explanation. In a second experiment, the 78-nt product of ribose-614 cleavage was isolated via gel purification. This product was radiolabeled and, following gel purification, incubated (in excess) with full-length ribose-614 (unlabeled). As the unlabeled ribose-614 becomes cleaved, and if the reverse reaction (ligation) occurs, excess 78-nt ensures that it may capture by ligation the 28-nt product, converting 78-nt to full-length ribose-614 over

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60 time. The reaction was monitored for 400 hours. No conversion of 78-nt to full-length material was observed. This also excludes reversibility as an explanation for the plateau. Testing if the Cleavage Products are Acting as Catalysts or Inhibitors By incorporating a short self-complementary segment that encourages a hairpin to clamp together the substrate and catalytic portions of the DNAzyme, Breaker and Joyce hoped to increase the chance that their library would contain molecules that self-cleave rapidly in cis. Further, cis-cleavage might be expected to predominate over trans cleavage, because the substrate is covalently bonded to the catalyst. The possibility that either cleaved product continues to act as a ribonuclease in trans or as an inhibitor cannot be ruled out. This is especially true for 614, where the clamp that might favor a hairpin, and therefore cis-cleavage, is not present. To test this, the 28and 78-nt products were gel purified. These products were added in equal amounts to a sample of uncleaved ribose-614. Cleavage of ribose-614 occurred at the same rate in the presence or absence of the 28and 78-nt fragments (Figure 3.4). This suggests that 28and 78-nt do not act as inhibitors or catalysts, at least when both are present in stoichiometric amounts (approximately the highest concentration they reach under normal 614 cleavage conditions). The 28and 78-nt products were also added to a sample of the ribose-library used in the selection. The ribose-library contains the cat+ribose primer followed by a segment of random sequence. Thus, it contains substrates that are (for the most part) not catalysts, and serves as the opposite of deoxyribose-614, which is a catalyst but not a substrate. The ribose-library alone does self-cleave to a detectable extent, as expected given that an unselected, random library has very few active catalysts.

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61 0 50 100 150 200 250 300 0 10 20 30 40 50 60 70 80ribose-614 ribose-614 + 28-nt product ribose-614 + 78-nt product ribose-614+ 28 & 78-nt prod. ribose-library + 28-nt product ribose-library + 78-nt product ribose-library + 28 & 78-nt prod. ribose-library hours Figure 3.4. Cleavage products do not affect ribose-614 cleavage, and only minimally alter ribose-library cleavage. Ribose-614 or ribose-library were incubated either alone (400 nM), or with the addition of the 28-nt product of 614 cleavage (400 nM), the 78-nt product of 614 cleavage (400 nM), or both (400 nM each). Data are fit to a single exponential curve. Errors in percent cleaved are < percentage points. The 28-nt product alone did not cleave ribose-library, showing that this product is not a catalyst. The 78-nt product alone did cleave ribose-library, although with a very low rate constant (kobs = 0.0006 hr-1, 35-fold lower than ribose-614 under similar conditions). The 28-nt product with the 78-nt fragment reduced cleavage of the ribose-library below that observed with the 78-nt fragment alone. This inhibition is presumably because 28-nt product competes with the ribose-library substrate in binding to the 78-nt catalyst. Inhibition of 614 by 28-nt is not noticeable presumably because 28-nt does not compete as effectively for 614 as a second molecule of 614 does. Improperly Folded ribose-614 Accounts for Part of the Plateau We then asked whether ribose-614 folds into active and inactive conformations, with the inactive form contributing to the cleavage plateau. Predictions of the conformation of ribose-614 using Mfold (Zuker, 2003) showed several potential structures. Close inspection of ribose-614 on a non-denaturing gel suggested two bands

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62 present in the starting material, one converting to a third band upon incubation under reaction conditions, and the other remaining unchanged. This is consistent with the hypothesis that ribose-614 adopts active and inactive conformations. To test this, ribose-614 was incubated until the plateau was reached (140 h). The mixture then was diluted into 2 volumes of formamide and heated (90C, 2 min). The uncleaved ribose-614 was purified by PAGE/urea, resuspended in buffer, re-folded using the slow cooling protocol, and subjected to cleavage conditions a second time. An additional 25% of the ribose-614 sample was cleaved in the 300 hours following gel purification. This additional cleavage following gel purification suggests that some of the initially uncleaved ribose-614 was in an inactive conformation. It is notable, however, that the gel-purified ribose-614 reached a plateau of ca. 25%, significantly lower than the cleavage plateau of the original sample (Figure 3.5). 0 50 100 150 200 250 300 350 0 10 20 30 40 50 60 70 80gel purified ribose-614 ribose-614 hours Figure 3.5 Gel-purification of ribose-614 at cleavage plateau results in additional cleaveage, indicating a fraction of 614 is folded into an inactive conformation. Ribose-614 (222 nM) was allowed to self-cleave for 140 hours, nearing cleavage plateau (triangles, top curve), at which point half the sample was resolved with denaturing PAGE. The fraction of ribose-614 remaining uncleaved at 140 hours was gel-purified, resuspended in reaction buffer (to a final concentration of 100 nM) and incubated for additional time (diamonds, bottom curve; percentage cleaved as a fraction of the label in the gel purified product, not of initial substrate). Errors in percent cleaved are < percentage points

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63 If misfolding were the only cause of incomplete cleavage, a plateau of approximately 65% would be expected in the second round of cleavage. The fact that the gel-purified sample reaches a lower plateau indicates that misfolding may account for only ca. 9% of total uncleaved fraction in the original sample (as an additional 25% of the sample cleaved following gel purification, and ca. 35% of the original sample remains uncleaved, the fraction of the initial sample that is misfolded is 25% of 35%, or 9%). We considered the possibility that gel-purification removed an inhibitor of cleavage. Ribose-614 was incubated to reach the plateau (141 h), and an aliquot was heated and slowly cooled. Any molecule in an inactive conformation was thus given another chance to adopt an active conformation (Figure 3.6). As in the gel-purification experiments, denaturing and refolding via this procedure increases the amount of material cleaved. The cleavage plateau over the first 150 hours following re-heating is about 10% higher than the untreated sample (in agreement with the estimate of 9% above), but still well below the ca. 90% cleavage when treated by strong base. When untreated samples are incubated beyond 300 hours, the cleavage levels approach those seen when samples are reheated at 140 hours, suggesting that re-folding into active conformations may occur slowly at room temperature. Thus, alternative conformations account for some, but not all, of the plateau seen between 150 and 300 h. Mutations Introduced into 614 during Cloning and Sequencing Account for part of the Plateau The ability of base to cleave ca. 94% of the cat+ribose primer suggests that about 6% of the cat+ribose primer, and therefore any full length DNAzyme made from the cat+ribose primer, may be missing the adenine riboside unit that is the site for cleavage.

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64 0 50 100 150 200 250 0 10 20 30 40 50 60 70 80ribose-614 ribose-614denatured andrefolded at 141 hours hours Figure 3.6. Reheating ribose-614 results in additional cleavage, indicating a fraction of ribose-614 is folded into an inactive conformation. Ribose-614 (100 nM) was allowed to react until it reached cleavage plateau (141 h), at which point half of the sample was denatured by heating to 96C for 3 min, and slowly cooled to 23C over 10 min. Errors in percent cleaved are < percentage points Other sequence variants may be present throughout ribose-614 as well, either as a result of mutations introduced during the synthesis of the primers and template, or mutations introduced by the polymerase during PCR. These mutations may reduce or eliminate the catalytic power of a fraction of the ribose-614 DNAzyme pool. This possibility was examined by cloning and sequencing DNA molecules from the cleaved and uncleaved fractions of ribose-614 after the plateau had been reached (Figure 3.7). Of the 29 sequenced clones of the 78-nt cleaved product of ribose-614, only four (14%) were found to be mutants. The mutations, at three sites, were in the N40 region between the primers. In contrast, of the 66 clones from the portion of ribose-614 that remained uncleaved at the plateau, only 64% had at least one mutation (42 individuals). Some 18% had a mutation in the N40 region, 5% had a mutation in the complementary strand primer, and 48% had a mutation in the catalytic strand primer (including 20% of the uncleaved ribose-614 molecules sampled that were missing the adenine riboside cleavable site).

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65 Excluding molecules missing the adenine riboside cleavable site, 44% of the uncleaved molecules possessed a mutation. This corresponds to 15% of the total initial population (44% of ca. 35% of total initial population remaining uncleaved at plateau). The extent of misfolding in 614 was estimated by assuming that all unmutated sequences from the uncleaved pool at the plateau remained uncleaved because they had adopted an inactive conformation. Given this, 13% of the total population is calculated to be misfolded (36% of the unreacted material is not mutated, and the unreacted material at plateau is ca. 35% of the total). k obs(hr-1)clone name: Sequence at 100 nM 614wt(x25) --------------------------------GGAAGACATGGCGACTCTCACATCATGCGAGCACAC-G-CAATAGCCTGATAAGGTTGGTAGTGACGGTAAGCTTGGCAC 0.015 614mut#1 --------------------------------GGAAGACATGGCGACTCTCACATCAT ACGAGCACAC-G-CAATAGCCTGATAAGGTTGGTAGTGACGGTAAGCTTGGCAC 0.0005614mut#2 --------------------------------GGAAGACATGGCGACTCTCACATCATGCGAGCACAC-G-CAATAG ACTGATAAGGTTGGTAGTGACGGTAAGCTTGGCAC 0.0064614mut#3(x2) --------------------------------GGAAGACATGGCGACTCTCACATCATGCGAGCACAC-G-CAATAGCCTGATAAGGT -GGTAGTGACGGTAAGCTTGGCAC 0.0040614wt(x21) CT-GCAGAATTCTAATACG-ACTCA-CTA-TAGGAAGACATGGCGACTCTCACATCATGCGAGCACAC-G-CAATAGCCTGATAAGGTTGGTAGTGACGGTAAGCTTGGCAC 0.015 614mut#31 CT-GCAGAATTCTAATACG-ACTCA-CTA-TAGGAAGACATGG NGA NT NTCACATCATGCGAGCACAC-G-CAATAGCCTGATAAGGTTGGTAGTGACGGTAAGCTTGGCAC614mut#33(2) CT-GCAGAATTCTAATACG-ACTCA-CTA-TAGGAAGACATGGCGACT NTCACATCATGCGAGCACAC-G-CAATAGCCTGATAAGGTTGGTAGTGACGGTAAGCTTGGCAC614mut#5(x6) CT-GCAGAATTCTAATACG-ACTCA-CTA-T -GGAAGACATGGCGACTCTCACATCATGCGAGCACAC-G-CAATAGCCTGATAAGGTTGGTAGTGACGGTAAGCTTGGCAC614mut#11 CT GCAGAATTCTAATACG-ACTCA-CTA-T -GGAAGACATGGCGACTCTCACATCATGCGAGCACACA-CAATAGCCTGATAAGGTTGGTAGTGACGGTAAGCTTGGCAC 0.0013614mut#12 CT-GCAGAATTCTAATACG-ACTC --CTA-T -GGAAGACATGGCGACTCTCACATCATGCGAGCACAC-G-CAATAGCCTGATAAGGTTGGTAGTGACGGTAAGCTTGGCAC614mut#13 CT-GCAGAATTCTAATACG-ACTCA-TA-T -GGA -GACATGGCGACTCTCACATCATGCGAGCACAC-G-CAATAGCCTGATAAGGTTGGTAGTGACGGTAAGCTTGGCAC614mut#14(x2)CT-GCAGAATTCTAATACG-ACTCA-CTA--GGAAGACATGGCGACTCTCACATCATGCGAGCACAC-G-CAATAGCCTGATAAGGTTGGTAGTGACGGTAAGCTTGGCAC614mut#16 CT-GCAGAATT NTAATACG-ACTCA-CTA-T -GGAAGACATGGCGACTCTCACATCATGCGAGCACAC-G-CAATAGCCTGATAAGGTTGGTAGTG -CGGTAAGCTTGGC -C614mut#17 CT-GCAGAATTCTAATACG-ACTCA-CTA-T -GGAAGACATGGCGACTCTCACATCATGCGAGCACAC-G-CAATAGCCTGATAAGGTTGGTAGTGACGG NAAGCTTGGCAC 614mut#18 CT-GCAGAATTCTAATACG-ACTCA-CT --AGGAAGACATGGCGACTCTCACATCATGCGAGCACAC-G-CAATAGCCTGATAAGGTTGGTAGTGACGGTAAGCTTGGCAC614mut#19(x3)CT-GCAGAATTCTAATACG-ACTCA-CT --TAGGAAGACATGGCGACTCTCACATCATGCGAGCACAC-G-CAATAGCCTGATAAGGTTGGTAGTGACGGTAAGCTTGGCAC614mut#22(x2)CT-GCAGAATTCTAATACG-ACTCA-CTA ATAGGAAGACATGGCGACTCTCACATCATGCGAGCACAC-G-CAATAGCCTGATAAGGTTGGTAGTGACGGTAAGCTTGGCAC614mut#24 CT-GCAGAATTCTAATACG-ACTCA-CTA-AGGAAGACATGGCGACTCTCACATCATGCGAGCACAC-G-CAATAGCCTGATAAGGTTGGTAGTGACGGTAAGCTTGGCAC 614mut#25 CT-GCAGAATTCTAATACG-ACTCA-CTA-TAGGAAGACATGGC AACTCTCACATCATGCGAGCACAC-G-CAATAGCCTGATAAGGTTGGTAGTGACGGTAAGCTTGGCAC 0.0005614mut#26 CT-GCAGAATTCTA -TACG-ACTCA-CTA-TAGGAAGACATGGCGACTCTCACATCATGCGAGCACAC-G-CAATAGCCTGATAAGGTTGGTAGTGACGGTAAGCTTGGCAC614mut#27 CT-GCAGAATTCTA CTACG-ACTCA-CTA-TAGGAAGACATGGCGACTCTCACATCATGCGAGCACAC-G-CAATAGCCTGATAAGGTTGGTAGTGACGGTAAGCTTGGCAC614mut#28 CT-G GAGAATTCTAATACG-ACTCA-CTA-TAGGAAGACATGGCGACTCTCACATCATGCGAGCACAC-G-CAATAGCCTGATAAGGTTGGTAGTGACGGTAAGCTTGGCAC614mut#29 CT TG CAGAATTCTAATAC C-AC NCA GCTA-TAG NAAGACATGGCGACTCTCACATCATGCGAGCACAC-G-CAATAGCCTGATAAGGTTGGTAGTGACGGTAAGCTTGGC NC614mut#30 CT-GCAGAATTCTAATACG-ACTCA-CTA-TAGGAAGACAT AGCGACTCTCACATCATGCGAGCACAC-G-CAATAGCCTGATAAGGTTGGTAGTGACGGTAAGCTTGGCAC 0.0003614mut#32 CT-GCAGAATTCTAATACG-ACTCA-CTA-TAGGAAG TCATGGCGACTCTCACATCATGCGAGCACAC-G-CAATAGCCTGATAAGGTTGGTAGTGACGGTAAGCTTGGCAC614mut#34 CT-GCAGAATTCTAATACG-ACTCA-CTA-TAGGAAGACATGGCGAC -CTCACATCATGCGAGCACAC-G-CAATAGCCTGATAAGGTTGGTAGTGACGGTAAGCTTGGCAC 0.0003614mut#35 CT-GCAGAATTCTAATACG-ACTCA-CTA-TAGGAAGACATGGCGACTCTCACATCATGCGAGCACAC-G-CAATAGCCTGATAAGGTTGGTAGTGACGG AAAGCTTGGCAC614mut#37 CT-GCAGAATTCTAATACG-ACTCA-CTA-TAGGAAGACATGGCGACTCTCACATCATGCGAGCACAC-G-CAATAGCCTGATAAGGTTGGTAGTG -CGGTAAGCTTGGCAC614mut#38 CT TGCAGAATTCTAATAC C-AC NCA GCTA-TAG NAAGACATGGCGACTCTCACATCATGCGAGCACAC-G-CAATAGCCTGATAAGGTTGGTAGTGACGGTAAGCTTGGC NC 614mut#39 CT-GCAGAATTCTAATACG-ACTCA-CTA-TAGGAAGACATGGCGACTCTCACATCA CGCGAGCACAC-G-CAATAGCCTGATAAGGTTGGTAGTGACGGTAAGCTTGGCAC 0.0181614mut#40 CT-GCAGAATTCTAATACG-ACTCA-CTA-TAGGAAGACATGGCGACTCTCACATCATGCGAGCACAC AG-CAATAG TCTGATAAGGTTGGTAGTGACGGTAAGCTTGGCAC 0.0003614mut#41 CT-GCAGAATTCTAATACG-ACTCA-CTA-TAGGAAGACATGGCGACTCTCACATCATGCGAGCACAC-G-CAATA TCCTGATAAGGTTGGTAGTGACGGTAAGCTTGGCAC614mut#42 CT-GCAGAATTCTAATACG-ACTCA-CTA-TAGGAAGACATGGCGACTCTCACATCATGCGA TCACAC-G TCAATAGCCTGATAAG CTTGGTAGTGACGGTAAGCTTGGCAC614mut#43 CT-GCAGAATTCTAATACG-ACTCA-CTA-TAGGAAGACATGGCGACTCTCACATCATGCGA TCACAC-G-CAATAGCCTGATAAG CTTGGTAGTGACGGTAAGCTTGGCAC614mut#44 CT-GCAGAATTCTAATACG-ACTCA-CTA-TAGGAAGACATGGCGACTCTCACATCATGCGAGCACAC-G-CAATAGCCTGATAAGGTTG -TAGTGACGGTAAGCTTGGCAC 0.0001614mut#45 CT-GCAGAATTCTAATACG-ACTCA-CTA-TAGGAAGACATGGCGACTCTCACATCATGCGAGCACAC-G-CA TTAGCCTGATAAGGTTGGTAGTGACGGTAAGCTTGGCAC614mut#46 CT-GCAGAATTCTAATACG GACTCA-CTA-TAGGAAGACATGGCGACTCTCACATCATGCGAGC TCAC-G-CAATAGCCTGATAAGGTTGGTAGTGACGGTAAGCTTGGCAC614mut#47 CT-GCAGAATTCTAATACG-ACTCA-CTA-TAGGAAGACATGGCGACTCTCACATCATGCGAGCACAC-G-CAATAG TCTGATAAGGTTGGTAGTGACGGTAAGCTTGGCAC 0.0005614mut#48 CT-GCAGAATTCTAATACG-ACTCA-CTA-TAGGAAGACATGGCGACTCTCACATCATGCGAGCACA T-G-CAATAGCCTGATAAGGTTGGTAGTGACGGTAAGCTTGGCAC614mut#49 CT-GCAGAATTCTAATACG-ACTCA-CTA-TAGGAAGACATGGCGACTCTCACATCATGCGAGCACAC-G-CAAT TGCCTGATAAGGTTGGTAGTGACGGTAAGCTTGGCAC 0.0014614mut#49 CT-GCAGAAT1CTAATACG-A2TCA-CTA-TAG3AAGACATGG4GACTCTCAC5TCATGCGAG6ACAC-G-CAAT7GCCTGATAA8GTTGGTAGT9ACGGTAAGC0TGGCAC 0 Figure 3.7. Sequence alignment of cleaved and uncleaved cloned 614 sequence variants. Mutations of 614 are highlighted in yellow; N40 region between primers is bold in each sequence and underlined in the original (no mutation) 614 sequence. Apparent first order catalytic rate constants (at 100 nM, in units of hr-1) are shown to right for a sample of mutant-614 DNAzymes. This is consistent with the estimate above based on the outcome of a cycle of unfolding and refolding.

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66 Several of the 614 mutants were tested to see if the mutations in fact reduced the rate of self-cleavage. Eleven of the twelve 614 mutants that were tested showed cleavage rates reduced by 10 to 100 fold compared to ribose-614. Ribose-614 Catalysis is not Mg++-dependent The mono and divalent cation requirements for ribose-614 activity were then examined. Reducing the concentration of NaCl from 1 M to 0.1 M eliminated cleavage. Only a modest change in the rate of cleavage of ribose-614 (100 nM) was observed when the reaction was run in the absence of MgCl2 (with and without 1 mM EDTA, initial rate 1.32 x 10-2 hr-1 and 1.37 x 10-2 hr-1, respectively) or in the presence of MgCl2 (1 mM, 1.53 x 10-2 hr-1; 10 mM, 1.70 x 10-2 hr-1; or 100 mM, 2.52 x 10-2 hr-1). Catalysis by 614 therefore does not require Mg++ in significant amounts even though 614 was selected in the presence of MgCl2 (1 mM). Requirement of a trace of Mg ++ cannot, of course, be excluded. The rates of two other catalysts isolated from this selection (ribose-62 and ribose-616) were also largely unchanged when the MgCl2 (1 mM) of the reaction buffer was replaced with EDTA (1 mM). For ribose-625, another molecule isolated from the selection, cleavage was eliminated by replacement of MgCl2 by EDTA at 25C. Ribose-614 Cleaves in trans A series of experiments was then performed to examine the rate of cleavage as a function of concentration. Self-cleavage is expected to be kinetically first order. Cleavage in trans is expected to be kinetically second order, meaning that an apparent first order rate constant kobs will not be independent of [DNAzyme], and fall to zero as [DNAzyme] falls to zero. A convenient way to separate simultaneous intra and intermolecular

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67 processes is to plot the apparent kobs as a function of [DNAzyme]. The y-intercept is kuni for the unimolecular reaction, while the slope of the line, reflects a second order process. Figure 3.8 shows a plot of the apparent kobs versus [ribose-614]. At low [ribose-614], the rate is independent of [ribose-614]; primary cleavage profiles are shown in Figure 3.9. At higher concentrations, however, the apparent kobs increases with increasing [ribose-614]. These results are consistent with a model that includes both unimolecular and bimolecular processes. Figure 3.8. Initial rate of ribose-614 cleavage as a function of [ribose-614]. At higher concentrations, a second order process is apparent. The rate constant for the apparent first order process (uncorrected for the cleavage plateau), from the y intercept, is ca. 0.006 hr-1. A best-fit line extrapolated to infinite dilution for [ribose-614] 3.5 nM and below (the region where the process is mostly unimolecular) gives an intercept corresponding to a unimolecular rate constant of 0.006 hr-1. This is an underestimate because the rate constant, based on initial rates, is uncorrected for the cleavage plateau (see below). An alternative way to obtain a rate constant for a first order process plots the log of uncleaved [ribose-614] (substrate remaining) versus time. Here, the progress of cleavage at low [ribose-614] fit a linear model well, while the progress at higher concentrations did not (Table 3.2). This suggests that below 20 nM, a unimolecular rate process

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68 0 50 100 150 200 250 0 25 50 75430.0 nM 108.0 nM 19.4 nM 3.5 nM 1.1 nM 0.3 nM K exp fit (h -1 ) =0.0610.0310.0230.0200.0110.011A [ribose614 ] hours 0 2 4 6 8 10 12 14 0.0 2.5 5.0 7.5 10.0 12.5 15.0430.0 nM 108.0 nM 19.4 nM 3.5 nM 1.1 nM 0.3 nM m (h -1 ) =0.032330.016270.009590.008990.006950.00610B [ribose614 ]hours Figure 3.9. Ribose-614 cleavage rate is concentration dependent. (A) Complete time course for ribose-614 at various concentrations. Rates are estimated based on a fit to a single first-order exponential equation. (B) Linear initial phase of time course. Rates are estimated based on the slope of best fit line. dominates, while a bimolecular rate process contributes above 20 nM. The rate constant estimated in this manner for [ribose-614] at 0.3 nM was 0.0042 hr-1 (uncorrected for the plateau), in good agreement with the estimate above. Table 3.2. Data from plot ln[S] t versus time for ribose-614 cleavage at various concentrations fit to a linear equation. [ribose-614] nM 430 108 19.4 3.5 1.1 0.3 (slope of ln[S]t)) 0.0032 0.0035 0.0030 0.0031 0.0030 0.0042 R2 0.78 0.89 0.92 0.90 0.97 0.97 We then asked if the progress curve for cleavage at high and low [ribose-614] fit best to a single exponential (in which both the rate and plateau can vary) or the sum of two exponential equations (in which both the rate and plateau can vary for each of two

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69 equations). At ribose-614 concentrations < 3.5 nM, progress curves fit best to a single exponential. At higher concentrations, the progress curve is fit best by the sum of two independent exponentials. This suggests that a single process dominates transformation of ribose-614 at low concentrations. If multiple active conformers exist at low [ribose-614], they are either in rapid equilibrium, or self-cleave with comparable rate constants. The first order constant was then corrected given information collected above about the plateau. After correction, a single exponential curve fit to the progress curve for 0.3 nM ribose-614 cleavage under unimolecular conditions. A rate constant 0.011x10-2 hr-1 was estimated (with a plateau of 61%). Various ribose-containing Substrates are Cleaved by deoxyribose-614 To establish trans cleavage in this system, internally labeled ribose-614 (which yields the 78-nt fragment as the only labeled product) was incubated with 5-32P-labeled cat+ribose primer (100 nM). The cat+ribose contains the substrate domain of ribose-614, but is not catalytically active. Here, cat+ribose is cleaved, as is ribose-614 (Figure 3.10, right panel). This establishes trans cleavage by ribose-614. The cleavage of ribose-614 (100 nM) in the presence of cat+ribose primer (100 nM) was lower than ribose-614 incubated alone (at 100 nM), suggesting that the cat+ribose primer competes with ribose-614 for trans cleavage by ribose-614 (Figure 3.11). Trans cleavage was also tested by challenging deoxyribose-614 (a modified form of ribose-614 in which the adenine riboside was replaced by a non-cleavable deoxyadenosine) with various substrates, each containing the cat+ribose sequence. These included (a) the cat+ribose primer, (b) a pool of molecules incorporating the cat+ribose primer followed by a random region and a 18-nt constant region (ribose-library), which

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70 collectively have no detectable catalytic activity, (c) a single clone from the ribose-library, (ribose-lib62), which also has no detectable catalytic activity, and (d) a mutant of 614 (614C72T) that has ca. 30 fold reduced activity compared to 614, at 100 nM. Figure 3.10. Both deoxyribose-614 (left panel) and ribose-614 (right panel) cleave cat+ribose primer in trans. Internally labeled deoxyribose-614 (100 nM) or ribose-614 (100 nM) were mixed with 5-end labeled cat+ribose (100 nM) and incubated under reaction conditions. The deoxyribose-614 does not contain the ribo-adenosine and therefore does not cleave itself. Unless otherwise stated, all kinetics were performed under standard reaction conditions, namely 1M NaCl, 1 mM MgCl2, 50 mM HEPES, 25C, and errors in percent cleaved are < percentage points. 0 50 100 150 200 250 0 10 20 30 40 50 60cleavage of ribose-614 cleavage ofcat+riboseby ribose-614 cleavage of ribose-614withcat+ribose competitor substrate hours Figure 3.11. Cat+ribose competes with ribose-614 for cleavage. Ribose-614 (100 nM) was incubated with and without cat+ribose (100 nM). Cat+ribose is not catalytic and when incubated alone is not cleaved. Data are fit to a single first-order exponential equation.

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71 Each substrate was cleaved by deoxyribose-614 (Figure. 3.12) although with different efficiencies. Ribose-614C72T (which should fold like ribose-614) is cleaved by deoxyribose-614 more slowly (suggesting that ribose-614C72T may fold so as to make the substrate domain inaccessible to catalysts), while the cat+ribose primer is cleaved faster. The fact that the small cat+ribose primer is better than full length substrates is consistent with the view that folding in longer substrates inhibits trans cleavage by deoxyribose-614. The unimolecular and bimolecular mechanisms by which 614 can cleave are summarized in Scheme 3.1. Figure 3.12. Compound deoxyribose-614 can cleave various substrates in trans. Unlabeled deoxyribose-614 (270 nM) was incubated with various radiolabeled substrates (cat+ribose, closed squares; ribose-lib6lo2, closed triangles; ribose-library, open diamonds; or ribose-614C72T, open circles) each at 7.5 nM). Each substrate possessed the same 5-primer sequence containing the adenine riboside. Compound deoxyribose-614 does not cleave itself, and percentage cleaved is the fraction of the substrate converted to product. Data are fit to a first order exponential curve. Data points being accurate to < percentage. Errors in the measurement of time are minute.

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72 Unimolecular Kinetic Scheme:614unfolded k1 (uni)k-1 (uni)614folded kcat (uni)ProductsBimolecular Kinetic Scheme:E + S k1 (bi)k -1 (bi)E.S k cat (bi)E.PE = ribose-614 or deoxyribose-614S = ribose-614, ribose-lib62 or cat+ribose Scheme 3.1. 614 can cleave via a unimolecular and bimolecular mechanism. Competition Studies of Ribose-614 Cleavage We then tested the ability of three of the above substrates, each with their ribose replaced by 2-deoxyribose (cat+deoxyribose primer, deoxyribose-614C72T, and deoxyribose-lib62), to compete with ribose-614 for self-cleavage. These competitors were added in 9-fold excess over ribose-614. For comparison, 9-fold excess of deoxyribose-614 was added to ribose-614. The addition of deoxyribose-614 (270 nM) to ribose-614 (30 nM) increased the rate of ribose-614 cleavage to the same level as seen for ribose-614 at 300 nM (Figure 3.13). 0 50 100 150 200 250 300 0 10 20 30 40 50 60 70 80d-614 cat-ribose d-lib62 d-614C72T no competitor Competitor (270 nM) added to ribose614hours Figure 3.13. Various substrates can compete with ribose-614 for self-cleavage. A nine fold excess of unlabeled deoxyribose-lib62, deoxyribose-614, deoxyribose-614C72T, or cat+deoxyribose was added to radiolabeled ribose-614 (33 nM) at time zero. Errors in percent cleaved are < percentage points.

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73 The cleavage of ribose-614, however, was reduced by excess cat+deoxyribose or deoxyribose-lib62. This showed that these molecules compete for ribose-614 self-cleavage. Curiously, addition of a 9-fold excess of deoxyribose-614C72T to ribose-614 did not reduce the rate of ribose-614 cleavage, but rather increased it. The ability of deoxyribose-614C72T to cleave ribose-614 may indicate that the mutation alters the ability of deoxyribose-614C72T to function as a substrate without inactivating its catalytic domain, consistent with the observation above that ribose-614C72T is itself a poor substrate for deoxyribose-614 cleavage. The competition study above was done at high [ribose-614] to favor trans cleavage. A parallel study at low concentrations of ribose-614 (favoring unimolecular cleavage) showed no inhibition of ribose-614 self-cleavage by deoxyribose-lib62 competitor. Saturation Kinetics in trans cleavage by deoxyribose-614 Two mechanisms for the trans bimolecular reaction were considered. In the first, deoxyribose-614 cleaves its substrate on every encounter. This gives a linear slope in a plot of an apparent rate constant vs. [catalyst] or [substrate] over the entire concentration range. An alternative second order process is possible, however. Here, substrate and catalyst form a complex that can dissociate before it reacts. This gives saturation Michaelis-Menten kinetics. To search for saturation kinetics, we exploited deoxyribose-614 as a DNAzyme that cannot act upon itself, but must act in trans. Concentrations of deoxyribose-614 were scanned to find a range where the apparent second order rate constant reaches a plateau reflecting saturation. Various substrates were used, including cat+ribose, ribose-lib62,

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74 and ribose-614, at low amounts relative to deoxyribose-614. Cleavage rates did not increase (2000 to 6000 nM deoxyribose-614), suggesting saturation of the bimolecular process. Assuming that unbinding is fast compared to reaction, a calculated fit gave a dissociation constant (Kd) of 29.0, 37.3, and 25.5 nM for cat+ribose primer, ribose-614 and ribose-lib62 substrates, respectively. The kcat(bi), determined from the maximum rate of cleavage with saturating enzyme, was 0.056, 0.048, and 0.049 hr-1 for cat+ribose, ribose-614 and ribose-lib62 substrates, respectively. Compound deoxyribose-614 Cleaves with Multiple-turnovers To show that deoxyribose-614 catalyzes cleavage with multiple turnovers, either cat+ribose or ribose-lib62 substrate were incubated in 4-fold excess over deoxyribose-614 (133:33 and 400:100 nM substrate:enzyme). With 400 nM enzyme and 100 nM substrate, two turnovers of substrate were observed at 100 hours for cat+ribose substrate and 200 hours for the ribose-lib62 substrate. As seen under single-turnover conditions, the cat+ribose substrate is cleaved faster under multiple-turnover conditions than the full-length ribose-lib62 substrate (at equivalent concentrations). This may be attributable to a better ability of the cat+ribose substrate to bind with or dissociate from the enzyme, or a lower proclivity for forming nonproductive interactions. Catalytic Power in trans is Unaffected by Annealing Protocol The impact of the slow cooling protocol on trans cleavage was tested by mixing deoxyribose-614 with substrate (ribose-lib62 or cat+ribose) either before or immediately

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75 after the slow cooling. This change in the annealing procedure did not alter noticeably the cleavage profile. At high concentrations (200 nM), the rate of cleavage of ribose-614 was unchanged by omitting the slow cooling. This also shows that intermolecular folding at the start of the reaction is not significantly effected by the annealing protocol, even though denaturation by heating allows molecules in inactive conformations to refold into active conformations. Similar experiments at low [ribose-614] favoring cis-cleavage showed a slight decrease in initial rate with the omission of slow cooling. The Commitment Step for deoxyribose-614 Cleavage If the rate limiting step is dissociation of the product-enzyme complex, a burst is expected in the initial phase of multiple turnover kinetics. To seek a burst, the concentration of deoxyribose-614 as a catalyst was held constant at 20 nM while the concentration of cat+ribose as a substrate was varied from 100 to 2000 nM. Turnover of substrate to product remained linear well beyond the initial turnover for all substrate concentrations, revealing no burst phase and suggesting that the overall rate is not limited by DNAzymeProduct dissociation. To estimate the relative magnitudes of kcat(bi) and k-1(bi), a chase was performed with unlabeled cat+deoxyribose added after four hours. The cat+deoxyribose chase is a substrate analog that cannot be cleaved, and therefore should compete with the labeled substrate. If all of the substrate-catalyst complex proceeds to product (which is the case if kcat(bi) >> k-1(bi)), then addition of the unlabeled chase will not influence the subsequent rate of appearance of labeled products. If, however, the rate of substrate dissociation from catalyst is faster than cleavage (kcat(bi) << k-1(bi)), then unlabeled chase molecules

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76 lacking a cleavable site should consume the newly disassociated catalyst, and the production of labeled products should largely cease. In the chase, deoxyribose-614 enzyme (2 M, saturating) acted in trans on small amounts (4 nM) of ribose-614 substrate. When the cat+deoxyribose chase was present at 5-fold excess over enzyme (10 M added at t = 4 h), a decrease in the rate of cleavage of ribose-614 after addition of the chase was observed (Figure 3.14a). The cleavage levels of ribose-614 following the addition of chase is reduced below that seen for a predominately cis-cleavage reaction of 1.1 nM ribose-614. This suggests that kcat(bi) << k-1(bi). Figure 3.14. Cleavage of various substrates by 614 is reduced by the addition of unlabeled chase.This indicates that the rate of ES dissociation is faster than the chemical step of cleavage. (A) Saturating amounts of deoxyribose-614 (2 M) cleaving ribose-614 (4 nM), with (closed squares) and without (open circles) chase (10 M) added at 4 h. (B) Saturating amounts of deoxyribose-614 (2 M) cleaving ribose-lib62 (4 nM), without chase (open circles) and with chase (5 M, closed triangles or 10 M, closed squares) added at 4 h. Similar results were seen following the addition of chase to ribose-614 under trans-cleavage conditions. Cleavage of ribose-614, however, was not completely eliminated by the chase. This suggested several explanations, including the persistence of cis cleavage

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77 in the presence of chase, or insufficient chase. Chase experiments conducted with ribose-614 cleaving in cis (2 nM) showed no significant change in cleavage following the addition of 30 nM chase (data not shown). If kcat(bi)<
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78 temperature (25C). Similarly, the rate for cis cleavage of ribose-614 is essentially the same at 4C and 25C (data not shown). The rate of the chemical step is expected (as an approximation) to increase two-fold for every 10C increase in temperature (Laidler & Peterman, 1979). Thus, it appears as though the overall rate of catalysis is largely affected by the folding/association step and not the chemical step. Figure 3.15. Ribose-614 rate of self-cleavage in trans is increased at lower temperatures. Ribose-614 (200 nM) was incubated at various temperatures from 4C to 42C. Initial rates were calculated and plotted vs. temperature. The rate of cleavage of ribose-lib62 in trans by deoxyribose-614 in single-turnover experiments is comparable at 15 and 25C, however. Similar experiments performed under multiple-turnover conditions reveal a rate enhancement at lower temperatures during cleavage of the first substrate, followed by cleavage of additional substrates at a slower rate (Fig. 3.16). This burst phase seen under multiple turnover conditions at 15C is not seen at 25C. This indicates that lower temperatures increase the initial rate by

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79 stabilizing intermolecular association between enzyme and substrate or, perhaps, through greater stability of the folded form of the enzyme. This stabilization presumably also slows the dissociation of product from enzyme to within the range of k -1(bi) and therefore reduces the rate for multiple-turnover subsequent to the first turnover. Figure 3.16. Burst kinetics (a higher cleavage rate during the first turnover compared to subsequent turnovers) is enhanced at lower temperatures. Ribose-lib62 substrate (400 nM) was incubated under multiple turnover conditions with deoxyribose-614 (100 nM) at 15C (closed squares) and 25C (open circles). The number of substrate turnovers was calculated by multiplying the percent of substrate cleaved by initial substrate concentration and dividing by [deoxyribose-614]. Predictions of the Energetically Favored Structure are not Supported by Experimental Data The ability of 614 to cleave in cis is not particularly surprising, because this is the function for which the DNAzyme was selected. It is therefore interesting that 614 is able to cleave various substrates in trans, and that the apparent first order rate constant at saturation for trans cleavage is 4-fold higher than the first order rate constant for cis-cleavage. The ability of 614 to cleave the cat+ribose primer, as well as a library of

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80 molecules containing the cat+ribose primer, suggests that 614 has a binding motif that pairs with a part of the cat+ribose primer common to all substrates, thus positioning a separate catalytic motif near the adenine riboside cleavage site. Chemical modifications specific for single-stranded DNA were used to probe secondary structure Deoxyribose-614 (300 nM) was folded overnight with and without excess unlabeled cat+deoxyribose substrate (10 M). The mixtures were then treated with either potassium permanganate (which modifies thymine) or dimethyl sulfate (which, at high salt concentrations, modifies guanine) for 2 and 5 min. Recovered samples were resolved by PAGE-urea next to a 10-bp ladder. All thymine and guanine nucleotides were somewhat sensitive to their respective reagents, indicating that either some 614 is not folded at all, or adopts multiple conformations. Nonetheless, T44, T46, T74, G33, G41, G65, G71, and G75 all demonstrated some degree of differential protection (numbers refer to the sequence of 614 shown in Figure. 3.7). The pattern of protected and sensitive positions observed under chemical modifications is not simultaneously compatible with any single structure predicted for 614 folded in cis or trans. This suggested that either a mixture of conformers exists, or none of the predicted structures accurately reflect the single true structure of 614. Likewise, each of the eight lowest free energy structure predictions made by Mfold was examined by generating mutations that would disrupt critical helixes. In no case could a loss of function arising from a nucleotide replacement be rescued by a compensatory mutation predicted by Mfold. This suggested that none of the predicted conformations dominated (data not shown).

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81 It is interesting to note that many of the structures reported for in vitro selected DNAzymes are based on unverified predictions using programs such as Mfold. Although such programs have been useful for predicting structures for many RNA and DNA molecules, this does not appear to be the case with 614. Discussion In vitro selection experiments offer the possibility of learning how molecular behavior is distributed within a sequence space defined by the building blocks of a biopolymer, in this case DNA. Structures within DNA sequence space are countable. For standard DNA, 4n sequences exist, where n is the length of the biopolymer. This distribution is relevant to issues as diverse as the origin of life and biomedicine. For the first, we wish to know how large a hypothetical prebiotic pool of random nucleic acid sequences must have been to contain the physical and catalytic requirements for life. For the second, we may want therapeutic or diagnostic DNAzymes, and need to know the likelihood of obtaining these from selection experiments. In both, we may ask whether added compounds (including divalent metal ions or other cofactors) increase the likelihood of obtaining catalysts. The answers to such questions might exploit the language of univariate statistical analysis (Johnson et al, 1994; Nelson, 1982). A distribution function P(kobs) relates the probability of finding a molecule with a particular catalytic power, kobs, to kobs itself. Because good catalysts are presumed to be scarce in the initial library relative to poor catalysts, and excellent catalysts are presumed to be scarce relative to good ones, this distribution is expected to be a decreasing function of kobs. Using such analyses, a better

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82 pool is defined as one having a distribution shifted to the right (Fig. 3.17). Adding a useful cofactor to the pool should shift the distribution to the right as well. To address such "big" questions, catalysts being examined must be sufficiently well behaved that their behavior can be quantitatively analyzed. Many DNAzymes are not, including (upon first inspection) 614. In particular, the reaction of 614 did not go to completion. Ca. 35% of the starting material is unreacted even after prolonged incubation. This makes quantitative analysis difficult. Figure 3.17. A univariate statistical distribution plotting the probability of encountering a catalyst with a particular kobs as a function of kobs, for two catalysts, one with a better pool (open circles, exponential function with b = 2), the other with a poorer (closed squares, exponential function with b = 5). In each case, the area under the line sums to unity. Because good catalysts are scarce in the initial library relative to poor catalysts, and excellent catalysts are scarce relative to good catalysts, this distribution is a decreasing function of kobs. The distribution shown is exponential; the true nature of the distribution in any particular space is, of course, unknown. Key questions in in vitro selection ask whether this curve is exponential, zipfian, or is best captured by mathematical approximations having multiple parameters.

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83 Similar behaviors are well known in the literature of nucleic acid catalysts. Hammerhead ribozymes, for example, are frequently reported to cleave to only 40 60% at the plateau. DNAzymes with ribonuclease activity and deoxyribonuclease activity are also frequently reported with self-cleavage plateaus of 25 65%. This behavior must be understood before any quantitative analysis can begin. This work accounts for the incomplete cleavage (the plateau) displayed by DNAzyme 614. Approximately 6-7% of the total ribose-614 sample does not cleave because it is missing the adenine ribonucleoside linkage, which is the site of cleavage. Approximately 9-13% of the total sample appears to fold into an unreactive conformation, which repopulates the active conformation via a cycle of denaturation and renaturation. As much as 15% of the unreacted material at the plateau appears to be sequences containing mutations introduced during amplification or the synthesis of template. These numbers sum to 30-35%, approximately the amount of DNAzyme left uncleaved at the plateau. Less than 5% of the plateau in our experiments can be attributed to the presence of DNA complementary to the catalyst. Nevertheless, these results show the importance of efficient removal of the complementary strand. Contamination by 10% of the complementary strand has little effect on initial cleavage rate or cleavage plateau, while greater contamination can greatly reduce the cleavage plateau and therefore estimates for cleavage rates. For this reason, we believe both asymmetric PCR and exonuclease degradation are superior to column purification (under strong base) as a way of eliminating the complementary strand.

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84 Given this understanding of the progress curve of the reaction undergone by 614, we asked whether the features of this DNAzyme reflect the environment under which the DNAzyme was selected. We expected that the presence of Mg++ in the selection would lead to catalytic molecules that used Mg++ as a cofactor. In part, this expectation arose because Breaker and Joyce themselves reported a Mg++-dependent DNAzyme emerging from their selection. Indeed, the Breaker-Joyce IVS protocol, by using Mg++ as the trigger to begin the selection process, seems to require that the DNAzyme be active only with Mg++. In addition, this expectation is based on a general view that the catalytic potential of DNA is poor. DNA lacks a range of functional groups, in particular, many of those present in proteins (Benner et al, 1987). In this view, the probability of finding a good catalyst that exploits a useful cofactor (such as Mg++) is greater than the probability of finding a good catalyst that does not. In terms of the distributions shown in Figure 3.17, adding Mg++ should shift the distribution to the right. For both reasons, we were surprised to discover that ribose-614 is as active in the absence of Mg++ as it is in its presence. This behavior was not unique to ribose-614 in this selection. It was also observed for two other deoxyribozymes isolated from the same selection, termed ribose-62 and ribose-616. A fourth DNAzyme generated in this work, ribose-625, was found to be Mg++-dependent at the temperature used for the selection, but evidently not at 4 C. While this sample is far from statistical, it suggests that the distribution of catalysts in the pool examined does not greatly favor Mg++-dependent catalysts over Mg++-independent catalysts. This, in turn, implies that adding Mg++ to the library does not shift the distribution in Figure 3.17 greatly to the right.

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85 The emergence from a selection of catalysts that fail to take advantage of available cofactors is not unprecedented. For example, Faulhammer and Famulok report a selection performed in the presence of histidine (20 mM) at a low concentration of Mg++ (0.5 mM). These authors hoped to select a DNAzyme that exploited histidine as a cofactor. They isolated, however, a DNAzyme that was highly active with Ca++ in the absence of histidine. This behavior was selected despite the fact that Ca++ was excluded from the selection. Mg++, present in the selection, was a poor substitute for Ca++. At the same time, Roth and Breaker, also seeking a histidine-dependent nucleic acid enzyme, found one (Roth & Breaker, 1998). Our results differ from those reported by Breaker and Joyce, who analyzed a single, Mg++-dependent catalyst that emerged from an analogous selection. Breaker and Joyce did not survey their catalysts to determine the ratio of Mg++-independent and Mg++-dependent catalysts. Thus, it is conceivable that their study of a Mg++-dependent catalyst differs from our study of a Mg++-independent catalyst because of the stochastic nature of the selection experiment. Other explanations are conceivable, however. Although the concentrations of Mg++ were the same in our selection and the Breaker-Joyce selection, the sequences used by Breaker and Joyce possessed two pairs of self-complementary sequences (each referred to as a clamp). These clamps were introduced by design into the library so as to anchor the substrate and enzyme regions of the molecule together with a four base pair clamp on one side of the cleavage site, and a six base pair clamp on the other. The combined length of these clamps increased to 15 base pairs following selection.

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86 In the selection experiments reported here, a binding clamp of this length was not introduced by design. This difference, which was not considered to be significant at the outset, may have had consequences. The absence of extensive clamps may transfer selective pressure onto the folding/association step, and away from the catalytic step. Thus, selection within a library lacking clamps may not generate as strongly catalysts that exploit cofactors as selection within a library having clamps. This may explain why the catalysts that we examined are Mg++-independent, while the catalyst examined by Breaker and Joyce is Mg++-dependent. A curious parallel is found when comparing the Faulhammer/Famulok selection experiment cited above, and the analogous experiment by Roth and Breaker. The former failed to isolate histidine-dependent DNAzymes; the latter generated several histidine-dependent DNAzymes. Two differences in their selection protocols appear relevant. The Faulhammer/Famulok protocol included low amounts of MgCl2 (0.5 mM) in addition to the histidine cofactor during selection; the Roth/Breaker selection did not. Further, the Roth/Breaker libraries included engineered binding clamps, analogous to the clamps in the Breaker-Joyce selection for ribonucleases. The Faulhammer/Famulok libraries did not. It may be coincidental that selections with clamped libraries in two cases generated catalysts that exploited an external cofactor, while analogous selections with libraries lacking the clamp generated catalysts that did not. These results suggest, however, that the presence of base pairing between the substrate and enzyme portions of the nucleic acid enzyme may be important to the experimental outcome.

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87 Including clamps between the substrate and enzyme motifs in the DNAzyme appears also to give faster catalysts. The catalyst that Breaker and Joyce examined in detail has a rate constant of 0.12 hr-1 before reselection; reselection improved this to 1.2 4.6 hr-1. Likewise, the catalyst reported by Santoro and Joyce had rate constants of 6 hr-1 and 600 hr-1 before and after reselection. The metal-cofactor independent catalyst reported by Sen and Geyer had rate constants of 0.17 hr-1 and 0.4 hr-1 before and after reselection. In contrast, the rate constants for the metal-cofactor independent catalysts selected here are considerably smaller (0.015 to 0.05 hr-1, both before reselection). This work suggests that if one wishes to obtain a fast catalyst, one should engineer clamps into the sequence. Indeed, Santoro and Joyce found that reducing the length of the clamps (for a DNAzyme selected with clamps) to six base pairs or fewer (on both sides of the clamp) reduced kcat 10 fold for an intermolecular reaction and increases KM 100 fold. On the other hand, to understand catalytic potential in a truly random pool, one is advised not to use engineered clamps. The information in an engineered clamp containing a 14 base pair duplex is substantial, even considering that the exact bases paired in not important. The specification of 14 nucleotides in a 4 letter alphabet is 414 108. This is significant relative to the size of a typical library (1013 molecules). What do these experiments tell us about sequence landscape and evolvability within DNA libraries, where the reaction sought is the cleavage of a ribonucleotide linkage? Many sequence variants of 614 were isolated that differed at only one or two positions. Most of these exhibited cleavage rates that were more than an order of magnitude slower than that for 614 itself. In contrast, variants of 614 changed at one or two sites were isolated following additional rounds of selection. These generally had

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88 rates that were an order of magnitude faster than those for the original 614. The fact that a 100-fold variation in rates arising from only minor changes in sequence suggests a degree of ruggedness to the landscape relating sequence space to catalysis in this system (Yomo, 2003). What do these experiments tell us about the origin of life? The hypothesis that life emerged as a nucleic acid capable of self-replication (Rich, 1962) remains disputed (Shapiro, 1988). Our recent discovery of a plausibly prebiotic route to ribose is encouraging the re-examination of this hypothesis (Ricardo et al, 2004). In no case, however, does DNA appear to be a candidate for the first living biopolymer. To the extent that conformation and catalysis in RNA and DNA is analogous, however, we may offer a few observations. First, binding clamps of sufficient length are unlikely to occur frequently in a prebiotic soup. If binding clamps of 14 base pairs are required (the optimal length estimated by Santoro and Joyce), it would increase the required size of a prebiotic library by a factor of 108. Without extensive clamps, it appears (if this work can be taken as a model) that catalysis is limited by folding. Folding, in the absence of extensive clamps, occurs more favorably at low temperatures. To the extent that the behavior of 614 is representative, cold temperatures would appear to be more favorable for a nucleic acid origins of life than high temperatures. Last, trans cleavage in this system is incidental to selection. It is, however, a likely result of the promiscuous nature of nucleic acid binding. Under the conditions in which a nucleic acid catalyst was evolving, selection likely acted upon a small set of nucleic acid catalysts isolated within a compartment. Significant trans interactions would be possible

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89 within the compartment. If trans reaction is generally associated with a nucleic acid catalyst that is capable of a cis reaction, this may be relevant to the origin of life.

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CHAPTER 4 DETECTING ORGANIC MOLECULES ON MARS Introduction Mars has become a prime target in human space exploration during the last decade. This renewed interest is in part due to the nascent field of astrobiology and the suggestion that a vestige of ancient Martian life was found in the interior of the Allan Hills meteorite (Mc Kay, et al. 1996). Several landing missions have since been directed to the red planet but about half of these were unable to descend safely to the Martian surface. Unfortunately, many of the instruments directly relevant to the fields of chemistry and astrobiology were lost in the unsuccessful landings. Therefore, the knowledge of the chemistry of the Martian surface remains limited to the experimental results obtained by the Viking missions nearly 30 years ago and the Pathfinder rover in 1999. As this dissertation is being completed, new data is also emerging from the opportunity rover. The Viking 1976 missions to Mars performed several experiments designed to assess the potential for life on the planet. The results were puzzling. Samples of Martian soil taken from the top 10 cm of the Martian surface released dioxygen when exposed to humidity (Oyama & Berdahl, 1977). At least one compound in a set of radiolabeled organic compounds (formate, D,L-lactate, glycollate, glycine and D,L-alanine) released radiolabeled carbon dioxide when placed in aqueous solution on the Martian surface, evidently via oxidative processes (Levin & Straat, 1979). Last, a gas chromatography-mass spectrometry (GC-MS) experiment looking for volatile products from a sample of 90

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91 soil heated for 30 seconds at 500 C did not detect any organic molecules (Biemann et al, 1977). The failure to detect organic molecules by GC-MS was especially surprising since some 2.4 x 108 g of unaltered carbon comes to Mars each year via meteor (Table 4.1)(Flynn, 1996; Hayatsu & Anders, 1981; Mullie & Reisse, 1987). Many meteoritic organic compounds are volatile and should have been detected by GC-MS (Sephton, Pillinger, & Gilmour, 1998). Pyrolysis should have generated volatile products from many of the non-volatile compounds, including the polymeric organic substance known as "kerogen", which accounts for the majority of organic material coming to Mars via meteorite, and as much as 1-3% of the weight of some meteorites (Hayes & Biemann, 1968). These too should have been detected by Viking, but were not. These results have been interpreted as evidence that the Martian surface contains no organic molecules of any kind, presumably because the Martian regolith carries an oxidant powerful enough to convert all organics to carbon dioxide. Coupled with the absence of liquid water on the surface of Mars, and the irradiation of the surface by ultraviolet light, the failure to detect organic substances led many to conclude that one must dig deeply (and perhaps very deeply) below the Martian surface to have a chance of encountering any organic molecules that may have arisen from life on Mars (perhaps present several billion years ago, when the surface of Mars was more like the surface of Earth at that time), or organic molecules that may have been delivered to Mars via meteorite (McKay et al, 1998; Kieffer et al, 1992). As this conclusion was influencing the design of missions to Mars, Benner et al, 1999 re-examined it in light of what is known about the oxidation of organic compounds,

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92 the nature of organic substances likely to come to Mars, and the features of the Viking 1976 analysis that limited the kinds of organic molecules that it could have detected. The examination suggested that organic compounds that arrive to Mars via meteorite are most likely to be converted to carboxylic acid derivatives that would not be easily detected by GC-MS. Organic molecules generated on Mars itself by non-biological (Hubbard et al, 1973; Chyba & Sagan, 1992; Hubbard, Hardy, & Horowitz, 1971; Horowitz & Hobby, 1977) or (entirely hypothetical) biological synthesis (Levin, 1977) should suffer similar fates at or near the surface. The hypothesis of Benner et al, started with the fact that Mars is exposed to ultraviolet radiation with sufficient energy to cleave water to give H and HO radicals. Some of the H radicals must recombine to give dihydrogen (H2), which escapes into space (Hunten, 1974; ibid, 1979), leaving behind the HO radical, which could react directly with organic substances, dimerize to give H2O2,( McDonald, de Vanssay, & Buckley, 1998) or generate peroxides or other species through combination with minerals in the Martian soil. The HO radical reacts directly with most organic molecules. In aqueous solution under terrestrial conditions, the second order rate constants range from 107 to 1010 Lmol-1sec-1 for reactions that include hydrogen atom abstractions and additions to double bonds. The concentration of HO on Mars is 1-2 x 105 cm-3, a number similar to the concentration of HO radical in the atmosphere at the surface of Earth (Hunten, 1979). Benner et al. considered five types of organic compounds (Table 4.1) known to come to Mars via meteorites: alkanes, alkylbenzenes, naphthalene and higher polycyclic

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93 aromatic hydrocarbons, kerogen, and amino and hydroxy acids. They then formulated a hypothetical mechanism of how HO and H2O2 might transform these compounds in generic oxidation pathways, and whether metastable intermediates in these pathways might accumulate. Table 4.1. Expected metastable products from organic substances in the Murchison meteorite (Hayatsu & Anders, 1981; Mullie & Reisse, 1987) Substance Concentration Metastable (parts per million) Products Acid insoluble kerogen 14500 benzenecarboxylic acids Aliphatic hydrocarbons 12-35 acetate Aromatic hydrocarbons 15-28 benzenecarboxylic acids Monocarboxylic acids ~330 acetate/oxalate 2-Hydroxycarboxylic acids 14.6 acetate/carbonate Alcohols (primary) 11 acetate Aldehydes 11 acetate Ketones 16 acetate, benzenecarboxylic acids Amines 10.7 acetate Urea 25 carbonate Heterocycles 12 carbonate, other products Oxidation of Alkanes under Martian conditions Alkanes react generically with the HO radical via abstraction of a hydrogen radical (H) at a tertiary center preferentially, then a secondary center, and last a primary center. This relative reactivity reflects the relative stability of the radical products, and is displayed under a wide range of conditions. Thus, a straight chain alkane would lose an internal hydrogen in the generic mechanism (Figure 4.1a). The resulting secondary radical is extremely reactive, and will be trapped by almost anything available. It will react with another HO radical to yield a secondary alcohol. It can transfer an electron to (for example) Fe+3, generating a carbocation that can be

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94 trapped by water (for example, from a hydrated mineral), also generating the secondary alcohol. Other products are possible, but a secondary alcohol is the generic intermediate in the oxidative degradation of n-alkanes (Figure 4.1a). HO abstracts an H from the carbon attached to the alcohol oxygen more readily than it abstracts H from the parent alkene. Thus, the secondary alcohol (under generic conditions) is expected to react faster than the parent. It will therefore not accumulate, but yield a ketone. The ketone, in turn, should undergo further oxidation to generate an ester, which will be cleaved to give a carboxylic acid and a primary alcohol, which will be oxidized directly to another carboxylic acid. Alternatively, the ketone might enolize, suffer oxidation, and then lead to a fragmentation to generate two carboxylic acids (Figure 4.1a). By these steps, the generic oxidation pathway for alkanes leads to carboxylic acids. These are, of course, subject to further oxidation. The abstraction of an H from the carbon attached to the COOH group is expected to be an important mode of oxidation involving HO. This will ultimately generate the next shorter carboxylic acid. An alternative mode involves the one electron oxidation of the carboxylate anion to give the carboxylate radical, which would lose carbon dioxide to give the radical of the shorter-by-one-carbon alkane. Depending on the trap, the product would be an alkane (and the process would resume), or another more easily oxidized derivative. In this cascade of intermediates, the carboxylic acid is the first that is slower to degrade than to be formed. Under typical Fenton conditions, for example, acetic acid reacts with the HO radical 100 times more slowly than does ethanol (Walling, 1999). Carboxylic acids are therefore likely to accumulate. Further, acetate is more stable to

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95 further reaction under generic conditions than propanoic acid and longer alkylcarboxylic acids. Thus, acetic acid accumulates especially effectively. Exemplifying the "generic oxidation" pathway are some "brand name" oxidations. In a Kuhn-Roth oxidation, for example, an alkane is refluxed in a solution of concentrated chromic acid (Kirsten & Stenhagen, 1952). Insignificant amounts of ketone or alcohol products can be isolated as intermediates in the oxidation cascade that follows; these are too unstable with respect to further oxidation. Organic alkanecarboxylic acids (butanoic acid and propanoic acid, for example) can be isolated as metastable intermediates, however. Upon incubation for longer times, these are further degraded to acetic acid. Acetic acid too can be oxidized, to give carbon dioxide. Nevertheless, acetate is more stable than longer chain alkanecarboxylic acids, and accumulates. This makes the Kuhn-Roth oxidation useful for elucidating the structure of natural products. The amount of acetate produced from a known amount of alkane corresponds to the number of methyl groups in the alkane. Figure 4.1 Oxidative degradation of the generic alkane (represented here by pentane) to acetic acid (a), toluene to benzoic acid (b), and kerogen to (mellitic acid) (c).

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96 Oxidation of Alkylbenzenes Under Martian Conditions The HO radical abstracts a benzylic H from the alkyl group of alkylbenzenes (such as toluene) to give a rather stable benzyl radical (Figure 4.1b). This may trap HO, or lose an electron to Fe3+ and then trap water, in each case forming benzyl alcohol. Benzyl alcohol is more reactive than toluene under generic conditions. It is not expected to accumulate, but rather be converted to benzaldehyde. Benzaldehyde is also unstable with respect to further oxidation, and also should not accumulate in the generic process. Rather, it should be converted to benzoic acid. Benzoic acid no longer has a benzylic hydrogen to lose to a radical oxidant. It is still thermodynamically unstable in the presence of oxidants to conversion to carbon dioxide. But it is metastable, resistant to further oxidation, and accumulates. Because benzoic acid has no hydrogen on the carbon adjacent to the COOH group, it also lacks a path available to alkanecarboxylic acids for further oxidative degradation. Further generic oxidative degradation involves a one electron oxidation of the benzoate anion, which decarboxylates to yield the phenyl radical, which then can be converted to benzene or phenol. This generic pathway can be illustrated by a specific oxidative process with commercial importance. Benzoic acid is synthesized on ton scales via the oxidation of toluene. The stability of benzoic acid under these oxidizing conditions is sufficient to allow benzoic acid to accumulate in the industrial process (Heberger et al, 1994). Oxidation of Polyaromatic Hydrocarbons (PAHs) Under Martian Conditions The generic oxidation of polycyclic aromatic hydrocarbons involves the addition of the HO radical to give a hydroxycyclohexadienyl radical. This suffers further oxidation to give eventually a single core aromatic rings to which carboxylic acids are attached

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97 wherever a second ring was fused. Thus, naphthalene, phenanthrene and anthracene all give phthalic acid in the generic oxidation process (Barbas, Sigman & Dabestani, 1996; Theurich et al, 1997). Higher PAHs give benzenetricarboxylic, tetracarboxylic, pentacarboxylic, and hexacarboxylic acids (Figure 4.1c)(Juettner, 1937). The generic pathway can be exemplified with laboratory reactions of naphthalene, which is 1-6 ppm in some carbonaceous chondrites (Pering & Ponnamperuma, 1971). The pseudo first-order rate constant for the first step in the reaction between naphthalene and the HO radical (Figure 4.2) is 0.035 min-1 (Bunce et al, 1997). The rate constants for further oxidation 1and 2-naphthol are higher (0.88 and 0.27 min-1). This implies that neither 1nor 2-naphthol will accumulate. The metastable end products are phthaldehyde and phthalic acid. This is true for a variety of other conditions (oxidation catalyzed by TiO2, by SiO2, (Barbas et al, 1993) and by Fe2O3 (Guillard et al, 1993). This uniformity in outcome argues that the oxidation of naphthalene will generically yield phthalic acid as a metastable intermediate (Lane et al, 1996). The metastability of phthalic acid to further oxidation has commercial significance. An important industrial synthesis of phthalic acid begins with the oxidation of naphthalene (Lowenheim et al, 1975). Phthalic acid is also produced from naphthalene under simulated Martian conditions (Or & Holzer, 1979). Oxidation of Kerogen Under Martian Conditions Polymeric organic material ("kerogen") has no defined structure. On Earth, kerogen (coal, for example) comes via metamorphosis of biological matter. Under generic oxidation

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98 Figure 4.2. Oxidative degradation of naphthalene to phthalic acid. Solid arrows indicate reactions documented in the literature with citations. Dotted arrows indicate transformations presumed to occur but without documentation in the cited literature. conditions, the aromatic portions of kerogen generate benzenecarboxylic acids, with one carboxylic acid group for every position on the core benzene ring that was attached to a carbon in the parent structure. These are metastable, accumulate, and are isolated and quantitated when defining the structure of kerogens. For example, treating coal with alkaline permanganate oxidized its carbon to carbonic acid (H2CO3, 42%), acetic acid (CH3COOH, 2%), oxalic acid (HOOC-COOH, 7%) and benzenecarboxylic acids (48%), with a trace of succinic acid (HOOC-CH2-CH2-COOH) (Bone, Horton & Ward, 1930). Kerogen is the most abundant organic substance in meteorites. As with terrestrial

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99 kerogen, the kerogen from the Murchison meteorite gives benzenecarboxylic acid products when oxidized (Hayatsu et al, 1977; Hayatsu et al, 1980). These are stable in refluxing nitric acid for 27 hours. Oxidation of Amino and Hydroxyacids Under Martian Conditions Polyfunctional molecules are easier to oxidize than unfunctionalized carboxylic acids. Thus, hydrogen peroxide (a mild oxidant) will, in the presence of iron salts, catalyze the oxidative decarboxylation of alpha-hydroxyacids to give carbon dioxide and the shorter aldehyde. This reaction, well known in sugar chemistry, has a brand name (the "Ruff degradation")( Wieland & Franke, 1927). Because of their multiple sites of reactivity, polyfunctionalized compounds are not expected to survive the generic oxidation conditions; they are the most likely to be converted entirely to carbon dioxide on the surface of Mars. One intermediate in the oxidative degradation of polyfunctional organic compounds is oxalic acid, HOOC-COOH. A report from 1928 suggests that the iron salt of oxalic acid is stable to further oxidation (Walton & Graham, 1928). Thus, oxalic acid may be a metastable intermediate in the generic oxidation pathway on Mars, where iron is abundant. The Amounts and Fates of Organic Carboxylic Acids This discussion makes the case that aromatic and aliphatic carboxylic acids are the metastable products of generic oxidation of meteoritic organic compounds. The generic oxidation pathway is exemplified by so many specific (admittedly terrestrial) reactions, and is so well supported by Organic Structure Theory, that it seems plausible that these products are found on Mars as well. Assuming that meteorites bring 2.4 x 108 g/year of organic carbon to Mars (Flynn, 1996), that kerogen in the Murchison meteorite is representative of this organic carbon,

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100 that Mars-bound kerogen generates the same proportion of benzenecarboxylic acids as the Murchison kerogen, and that the mass yield of these is 10%, then ca. 2 x 1017 grams of benzenecarboxylic acids should have been generated on Mars since its surface dried three billion years ago. This corresponds to 2 kg of benzenecarboxylic acids per square meter of the Martian surface (the surface area of Mars is ca. 1014 m2). It is possible that these compounds were diluted by wind and impact into the Martian regolith. If mixed in the regolith to a depth of one meter, 2 kg of benzenecarboxylates would contribute ca. 500 ppm by weight of the first meter of surface of Mars (the density of Mars is ca. 4 gm/cm3). If gardening mixes the material to a depth of 1 km, benzenecarboxylates will be present at a concentration of 500 ppb. The Viking MS had a sensitive to the ppb level, and should have detected these had it had access to them. Other processes might have removed organic carboxylic acids from the immediate surface. Carboxylic acids react with metal oxides to form salts (Martell & Smith, 1977; Avdeef, 1993). These often display some solubility in water. If the Martian surface has been exposed episodically to water, organic salts may have been removed from the surface by leaching, and concentrated in sub-surface environments. Examples on Earth include highly soluble salts (e.g., halite) and poorly soluble salts (e.g. gypsum). This process is almost certainly less important than gardening in the recent past, as surface water on Mars has been scarce for billions of years, and iron salts of benzenecarboxylic acids are poorly soluble (Giammar & Dzombak, 1998; Wu et al, 1996). Most important, of course, are chemical reactions that would degrade the carbon skeleton. The Fenton reaction serves as a model, even recognizing that it is best known as

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101 an aqueous process (Walling, 1997). The Fenton reaction is believed to involve HO radical generated from H2O2 (Chen et al, 1998). Hematite and goethite (both iron oxides believed to exist on the Martian surface) are effective catalysts (Lin & Gurol, 1998; Watts et al, 1997). The Fenton reaction is known to degrade organic molecules ranging from 2-methylnaphthalene (an aromatic compound) to n-hexadecane (an aliphatic compound) entirely to carbon dioxide, if given sufficient time. Even relatively resistant molecules can be degraded. For example, trinitrotoluene (TNT) is converted by H2O2 in water to trinitrobenzoic acid, from there to trinitrobenzene, and from there to oxalic acid as the primary organic end product. The oxalic acid is removed only when the mixture is exposed to light (Li, Comfort & Shea, 1997). Fenton chemistry converts benzoic acid into hydroxybenzoic acid and guanosine into 8-oxoguanosine (Sandstrom et al, 1997). A UV-accelerated Fenton reaction is also known in aqueous solution, and is proposed to generate Fe2+ by photoreduction (Sun & Pignatello, 1993). The efficiency of the Fenton reaction depends on the ligands around iron (Pignatello & Baehr, 1994; Dean, & Nicholson, 1994). For example, ferric oxalate initiates the destruction of other molecules (Safarzadeh-Amiri Bolton & Cater, 1997; ibid, 1996). Benzoic acid inhibits the Fenton reaction in certain terrestrial experiments (Zakharov & Kumpan, 1996). Aromatics are protected against degradation by more easily oxidized species (Owen et al, 1996). Thus, it is difficult to predict the consequences of Fenton chemistry on Mars, even if we assume that the process is analogous to the aqueous process known in the laboratory. An alternative path for the oxidative degradation of carboxylic acids involves the one electron oxidation of their anions to give the corresponding carboxylate radicals

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102 (Lamrini, 1998). These will lose carbon dioxide to generate an organic radical, which will then be trapped as part of the oxidative cascade. Photons can accelerate this process, and are likely to be important on the UV-irradiated surface of Mars (Or & Holzer, 1979; Bullock et al, 1994). Alkanecarboxylic acids are particularly susceptible to photochemical degradation. Many benzenecarboxylic acids are quite stable to photochemical degradation, however (Jeevarajan, & Fessenden, 1992). Phthalic acid derivatives, for example, yield phthalic anhydride under prolonged irradiation (Balabanovich, & Schnabel, 1998; Balabanovich, Denizligil & Schnabel, 1997), but no further degradation of the phthalic acid core. Failure of Viking 1976 to Detect Organic Carboxylic Acids If laboratory reactions are taken as examples of the generic oxidation pathway, the rates for the destruction of benzenecarboxylates are at least 103 to 106 fold slower than their rates of formation. Depending on the tempo of chemistry overall on Mars (and remembering that the billion years available for the accumulation of meteoritic organics is also available for the destruction of the derived benzenecarboxylates), substantial amounts of the kilogram of benzenecarboxylates expected to be generated per square meter should have survived. Their concentration would fall below the nominal sensitivity of the Viking 1976 mass spectrometer only if more than 99% of these were destroyed and if gardening diluted these to an average depth of 1 km or greater. To gain access to the Viking mass spectrometer, however, the organic molecule first pass through a gas chromatograph. Only volatile molecules can do so. Salts of organic carboxylic acids are not volatile. Thus, the salts of benzenecarboxylic acids, oxalic acid, and acetic acid would not be directly detectable by the Viking GC-MS experiments, even if they had been present.

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103 The ability of the Viking experiments to detect organic carboxylates therefore depends critically on the ability of these carboxylates to generate volatile products in the sample preparation (pyrolysis for 30 seconds at 200, 350, and 500C). The three generic oxidation products, benzenecarboxylic acids, acetic acid, and oxalic acid, could be detected only with difficulty by the Viking GC-MS. Oxalic acid generates carbon dioxide, carbon monoxide and water under pyrolysis. These were in fact detected, but are all also components of the Martian atmosphere. Higher benzenecarboxylic acids also do not easily yield volatile pyrolysis products. Benzenehexacarboxylate, a non-volatile compound, will eventually release carbon dioxide upon pyrolysis and become benzenepentacarboxylate (Manion, McMillen & Malhotra, 1996) and then benzenetetracarboxylate. The salts of these, however, are also not volatile. Acetic acid and its salts may be pyrolyzed to give volatile products. At high concentrations, acetone is formed (Davis, & Schultz 1962). However, the iron (II) acetate and iron (II) propionate salts are reported to be "amazingly stable up to 400-500C" (Granito & Schultz, 1963). For these reasons, the Viking experiments do not exclude the possibility that the soil being tested contained organic carboxylic acids, especially benzenecarboxylic acids in substantial amounts. To examine experimentally this conclusion, Benner et al 2000, synthesized the iron (III) salts of phthalate, mellitate and benzene-1, 2, 4-tricarboxylate. These were then subjected to thermolysis-mass spectrometry following heating from 25 to 400 in 30 seconds. The phthalate, mellitate and oxalate salts were separately heated to 400 in a quartz capillary in a direct insertion probe. Under these conditions, which give

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104 the mass spectrometer better access to the probe than the Viking MS had, no signal was observed from the iron (III) salts of mellitic acid and benzene-1,2,4-tricarboxylic acid. Iron (III) phthalate yielded a peak that might have been detected by the Viking GC-MS, corresponding to phthalic anhydride. Iron (II) acetate generated acetone, acetic acid and acetic anhydride in addition to water and carbon dioxide. Iron (III) oxalate releases carbon monoxide as well as carbon dioxide and water. This suggests that the Viking experiments can rule out substantial amounts of acetate on the surface (the top 10 cm), and modest amounts of phthalate, but not higher benzenecarboxylates, which are the principal products of the generic oxidative degradation of organic materials arriving via meteorite. The Infrared Spectra of the Martian Surface The infrared spectra of the Martian surface (Figure 4.3) was recently obtained by using data obtained with the Termical Emission Spectrometer (TES) instrument on board of the Mars Global Surveyor (MGS) (Bandfield, et al, 2003). The assignment of the signals on the spectra by Banfield et al, was done with the assumption that carbonate minerals are present in the surface of Mars (since not organic molecules had been detected on the planet). Several carbonate minerals mixtures were prepared in different proportions until the infrared spectra of the prepared sample closely resemble the most distinct features of the Martian surface spectra (Figure 4.4). From this comparison, the author concluded that magnesite (MgCO3) was the mineral which adsorption lines fit better the Martian IR spectra (Figure 4.5). The carbonate mineral was found to be distributed uniformly around the planet with not indication of a concentrated source and calculated to have an abundance in the surface of

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105 ~2-5% weight on Mars surface. The authors however recognize that many other combinations of minerals will probably give a better match to the spectra. Figure 4.3. Martian surface IR spectrum. The top spectra corresponds to the highalbedo surface dust spectrum, dashed lines represent standard deviation. The spectra at the bottom corresponds to terrestrial particulate basalt, shown to illustrate the resemblance in spectral features between 300-1200 cm-1. Figure 4.4. Mars dust, labradorite standard and labradorita + minerals mixture spectra. (see absorptions in the regions of 1350 cm-1 to 1580 cm-1 and a carbonate absorption at ~880 cm-1)

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106 Figure 4.5. Mars dust and magnesita-labradorite mixture spectra. The predominant abundance of magnesite over siderite (FeCO 3 ) or calcite (CaCO 3 ) was not explained in the paper, also details in the 1380-1520 cm -1 region were leave unexplained (the elemental abundance in the martian soil for the metals Mg, Fe and Ca are: magnesium~5%, iron ~13%, calcium ~4% ; Rieder et al, 1997). In the present chapter salts of benzenecarboxylates will be synthesized by the general method described by Galwey (1965), and their infrared spectra will be compared to that of the martian soil to evaluate if these salts could account for the unexplained features in the spectra.

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107 Detecting the Missing Organics on Mars Any attempt to perform qualitative and quantitative chemistry on Mars using a robotical lander, will be conditioned to the non-chemical aspects associated with space exploration (i.e: payload, complexity of the task, etc.). With this in mind, an experimental scheme for the detection of benzenecarboxylates is presented and evaluated on this chapter. Fluorescence derivatization of the different benzenecarboxylates will be attempted by using a one step reaction known as the Fiegls test (Feigl, 1961) which has been traditionally used in classical organic chemistry for the qualitative identification of 1,2-dicarboxylic acids. Fiegls test requires the addition of an excess 1,3-benzenediol (resorcinol) and catalytic amounts of acid (sulfuric acid) to the dicarboxylic acid containing molecule under analysis. The reaction occurs in a few minutes if the mixture is heated at 120 C and the product formed contains the xanthene ring structure (Figure 4.6). Fluorescein is the reaction product of phthalic acid (or anhydride) under Fiegls conditions and is of widely use in chemical and biological fluorescence sensing systems due to the high quantum yield and chemical stability that allows its detection at the single molecule level (Figure 4.6). The addition of carboxylic units to the phtalic acid core does not appear to affect the fluorescent ability of the fluorescein derivatives. Carboxy-fluorescein the product of resorcinol condensation with 1,2,4 benzene tricarboxylic acid is also a common fluorophore in molecular biology applications due to its enhanced solubility in water as a result of the additional carboxylic residue.

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108 O OH HO O O COOH COOH + HO OH H2SO4Phthalic AcidResorcinolFluorescein Figure 4.6. Phthalic acid yield fluorescein when heated with resorcinol in the presence of acid. The reactivity of the iron salt of benzenecarboxylates hypothesized to be present on Mars surface, will be also tested towards fiegls reaction. Materials and Methods Chemicals All reagents and materials were purchased from Sigma-Aldrich Co. in their highest quality and used without any further purification. Analytic Instrumentation Fluorescence analysis Flourescence spectra (emission and absorption) were recorded in a SpectroFluorometer Fluorolog 3. Courtesy of Professor Weihong Tan at the University of Florida. Infrared analysis Infrared spectra (KBr pellets) were recorded in a Perkin Elmer 1600 series FT-IR. Courtesy of Professor Kirk Schanze at the University of Florida. High performance Liquid chromatographymass Spectrometry (HPLC-MS) HPLC-MS (ESI) analysis were carried out by Dr. Jodie Johnson at the University of Florida.

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109 Synthetic Preparations Synthesis of Mellitic Acid Salts Crystalline aluminium mellitate (mellite) was obtained as a gift from Dr. Steven Benner mineral collection. Synthesis of manganous mellitate (1) Manganous carbonate (MnCO 3 ; 0.55 g, 4.3 mmol) and mellitic acid (0.5 g, 1.4 mmol) were mixed in water (10 mL) and boiled under stirring to complete evolution of gas, the solution was cooled, filtered and the precipitate was washed thoroughly with Milli-Q water. The resulting solid was dried in air at 80 C. Synthesis of zinc mellitate (2) The procedure for the preparation of this salt was the same as the one used for the synthesis of compound (1). The amounts used were: zinc carbonate-hydroxide hydrate (ZnCO 3 .2Zn(OH) 2 .H 2 O; 0.68 g, 2.1 mmol) and mellitic acid (0.25 g, 0.7 mmol). Synthesis of cupric mellitate (3) The procedure for the preparation of this salt was the same as the one used for the synthesis of compound (1). The amounts used were: basic copper carbonate (CuCO 3 .Cu(OH) 2 ; 0.96 g, 4.3 mmol) and mellitic acid (0.5 g, 1.4 mmol). Synthesis of nickel mellitate (4) Basic nickel carbonate (2NiCO 3 .3Ni(OH) 2 .4H 2 O; 0.61 g, 1.05 mmol) and mellitic acid (0,25 g, 0.7 mmol) were mixed in water (5 mL) and heated for 30 min at 90 C. The product was filtered off, washed with water, ethanol and ethyl ether, and dried at 60 C. Synthesis of cobalt mellitate (5) The procedure for the preparation of this salt was the same as the one used for the synthesis of compound (4). The amounts used in the experiment were: cobalt carbonate

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110 (CoCO 3 ; 0.52 g, 4.3 mmol) and mellitic acid (0.5 g, 1.4 mmol). Synthesis of magnesium mellitate (6) The procedure for the preparation of this salt was the same as the one used for the synthesis of compound (5). The amounts used in the experiment were: magnesium carbonate hydroxide pentahydrate (4MgCO 3 .Mg(OH) 2 .5H 2 O; 0.25 g, 0.5mmol) and mellitic acid (0.25 g, 0.7 mmol). Synthesis of calcium mellitate (7) The procedure for the preparation of this salt was the same as the one used for the synthesis of compound (5). The amounts used in the experiment were: calcium carbonate (CaCO 3 ; 0.42 g, 4.3 mmol) and mellitic acid (0.5 g, 1.4 mmol) Synthesis of iron mellitate (8) Ferric hydroxide, was precipitated by the addition of an excess concentrated ammonium hydroxide (1,5 mL) to a aqueous solution of ferric chloride (FeCl 3 5 mL, 1 M). The resulting mixture was boiled for 30 minutes, and the precipitated solid washed with hot water and removed by decantation. Mellitic acid (0.56g, 1.6 mmol) and water (5 mL) were then added to the solid ferric hydroxide and this solution was heated for 30 minutes at 90 C. The product was washed by decantation, filtered and dried in air at 60 C. Synthesis of fluoresceins: Fiegls test In a typical reaction, the poly-benzecarboxylic acid (phtalic acid, benzene tricarboxylic acid, pyrometllitic acid and mellitic acid) (1 eq) was mixed with resorcinol (2 eq) and heated by 5 min at 120 C at which time sulfuric acid (0.1 mL) was added. The resulting brown mixture was heated by additional 5 min, then cooled and made alkaline by addition of a concentrated solution of sodium hydroxide (5 M).

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111 Results Synthesis of mellitic acid salts The infrared spectra of each of the mellitic acid salts are presented below: Manganous mellitate (Figure 4.7), zinc mellitate (Figure 4.8), cupric mellitate (Figure 4.9), nickel mellitate (Figure 4.10), cobalt mellitate (Figure 4.11), magnesium mellitate (Figure 4.12), calcium mellitate (Figure 4.13), iron mellitate (Figure 4.14) and aluminium mellitate (Figure 4.15). Fluorescence spectra Analysis The Fiegls test derivatives of the different benzenecaboxylate showed strong fluorescence in alkaline medium. Table 4.2 shows the different excitation and emission signals obtained during the analysis. Table 4.2 Fluorescence analysis of the fluoresceines of benzenecarboxylates. Acid Excitation (nm) Emission (nm) Blank 475 ---1,2 benzene dicarboxylic acid (phtalic acid) 475 512 1,2,4 benzenetricarboxylic acid 475 517 1,2,4,5 benzene tetracarboxylic acid (pyromellitic acid) 475 517 Benzene hexacarboxylic acid (mellitic acid) 475 512 Iron phtalate 475 512

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112 Figure 4.7. Infrared spectra of manganous mellitate (KBr). Figure 4.8. Infrared spectra of zinc mellitate (KBr).

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113 Figure 4.9. Infrared spectra of cupric mellitate (KBr). igure 4.10. Infrared spectra of nickel mellitate (KBr). F

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114 Figure 4.11. Infrared spectra of cobalt mellitate (KBr), Figure 4.12. Infrared spectra of magnesium mellitate (KB r)

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115 Figure 4.13. Infrared spectra of calcium mellitate (KBr) Figure 4.14. Infrared spectra of iron mellitate (KBr)

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116 KBr) Discussion be en Figure 4.15. Infrared spectra of aluminium mellitate ( A comparison of the Infrared spectra of the different synthetic mellitic carboxylates salts, indicates that all the salts show absorption bands in the region of 1350-1600. A superimposed IR spectrum of the mellitates of the most abundant elements on Mars, with the IR of the surface of Mars is shown in Figure 4.16. Not conclusive arguments candone about the presence of mellitates in the martian surface in the absence of Martian samples. At the very least, we can say that the resemblance in the IR spectra keeps opthe possibility for contemplating the presence of benzene carboxylates on Mars and therefore detection systems that target these compounds should be included when designing future missions to the red planet.

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117 Figure 4.16. Infrared spectra of the mellitate salts of alum inium, iron, magnesium and calcium. Note the appearance of signals in the 1350-1500 region unexplained in previous reports by Banfield et al. The Fiegls test analysis shows that all the benzenecarboxylic acids studied were able to produce fluorescent derivatives, allowing identification even when coordinate to metals as in the carboxylates. The structure of the different fluoresceines was confirmed by mass-spectrometry and is consistent with the addition of two resorcinol molecules to the dicarboxylic acid moiety in benzenecarboxylates. When resorcinol was used in large excess over the 1,2,4,5 tetracarboxylic acid, two major products were observed after HPLC-ESI analysis. These products correspond to the expected mono and di-fluorescein derivatives of the tetracarboxylic acid (Figure 4.17). In general all the fluorescein derivatives show a similar spectrum The intensity of the emissio n decreases as successive carboxyl substituents are added to the benzene core

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118 O OH HO O O COOH HOOC O OH HO O O O OH HO O O 1,2-pyromellitic fluoresceinpyromellitic di-fluorescein Figure 4.17. Structures of the fluorescein derivatives of pyromellitic acid (1,2,4,5-benzenetetracarboxylate) detected by HPLC-MS (ESI). The absorption and emission spectra for the different derivatives (Table 4.2) show that the specific maximum emision spectral line for each compound makes it possible to distinguish and identify each individual fluorescein. The sensitivity of the method allows it to be a powerful tool for the search of oxidized organic molecules rising from kerogens. This approach is suitable for both Earthbound research on SNC meteorites and in situ studies on Mars.

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APPENDIX A NMR DEGRADATION EXPERIMENTS Half life of the different isomeric aldopentoses was calculated by measuring the ratio of anomeric proton/ internal standard (integral value) at different time points. In some cases several half life times are reported for the same aldopentose; this is caused by different rates of decay of the different anomeric proton signals. These phenomena could be explained by considering preferential protection of the different borate complexes possible for each aldopentose. Because the rate of decomposition in each case is proportional to the amount of pentose in the aldehydic form, those boron-pentose complexes that stabilize the closed ring furanose form (locking it) will be more stable towards decomposition. Another possible explanation is that the aldopentoses are interconverting (into another pentose or the corresponding ketose) and this alkaline isomerization is catalyzed by borate. 1 H NMR spectra were superimposed using the software package MestRe-C 2.3a obtained free of charge at: http://www.mestrec.com/. 119

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120 Figure A1.1. D-arabinose incubation in the presence of calcium hydroxide, pD:12. F igure A1.2. D-arabinose incubation in the presence of calcium-hydroxyde and borate, pD:12

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121 Figure A1.3. D-lyxose incubation in the presence of calcium hydroxide, pD:12. igure A1.4. D-lyxose incubation in the presence of calcium hydroxide + borate pD:12. F

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122 Figure A1.5. D-ribose incubation in the presence of calcium hydroxide, pD:12. Figure A1.6. D-ribose incubation in the presence of calcium hydroxide + borate, pD:12.

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123 Figure A1.7. L-xylose incubation in the presence of calcium hydroxide, pD:12. Figure A1.8. L-xylose incubation in the presence of calcium hydroxide + borate, pD:12.

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APPENDIX B DIOS COMPETITION EXPERIMENTS Figure A2.1. DIOS spectra of 1,4-Anhydroerythritol vs arabinose. F igure A2.2. DIOS spectra of arabinose vs 13C-ribose. 124

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125 Figure A2.3. DIOS spectra of 1,4-Anhydroeythritol vs lyxose Figure A2.4. DIOS spectra of lyxose vs 13C-ribose

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126 Figure A2.5. DIOS spectra of 1,4-Anhydroerythritol vs ribose F igure A2.6. DIOS spectra of 12C-ribose vs 13Cribose

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127 Figure A2.7. DIOS spectra of 1,4-Anhydroerythritol vs Xylose Figure A2.8. DIOS spectra of xylose vs 13 C-ribose

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128 Figure A2.9. DIOS spectra of 1,4-anhyroerythritol vs ribulose Figure A2.10. DIOS spectra of ribulose vs 13 C-ribos e

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129 13 Figure A2.11. DIOS spectra of Xylulose vs C-ribose.

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Alonso Ricardo was born in Santiago de Cali, Colombia, in 1977. He received a B.S in chemistry in 2000 from the Universidad del Valle while working with Dr. Luz Marina Jaramillo. After completing his B.S he joined the chemistry graduate program at the University of Florida working under the supervision of Dr. Steven A. Benner to BIOGRAPHICAL SKETCH pursue a doctorate. 137


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BIOORGANIC MOLECULES INT THE COSMOS AND THE
ORIGINT OF DARWINIAN MOLECULAR SYSTEMS














By

ALONSO RICARDO


A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL
OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT
OF THE REQUIREMENTS FOR THE DEGREE OF
DOCTOR OF PHILOSOPHY

UNIVERSITY OF FLORIDA


2004

































This work is dedicated to my family.
















ACKNOWLEDGMENTS

I will like to thank Dr. Steven Benner for the research opportunities and guidance

that he has offered me. I also thank Dr. Matthew Carrigan for his help, friendship and for

being an excellent lab partner. I am grateful to the people in the mass spectrometry

laboratory at the University of Florida and to Dr. Maurice Swanson for providing

laboratory space and collaboration I also thank Fabianne for her unconditional love and

support. Finally, I am forever grateful to my family for their constant support, care, and

consideration that helped me to made this work possible.




















TABLE OF CONTENTS


page

ACKNOWLEDGMENT S ................. ................. iii........ ....


LI ST OF T ABLE S ................. ................. viii............


LI ST OF FIGURE S .............. .................... ix


AB S TRAC T ......_ ................. ..........._..._ xiii..


CHAPTER


1 ORIGIN S ................. ...............1.......... ......


Background and Significance ................. ...............1............ ....
Prebiotic Chemistry............... ... .............
Sugars in the Prebiotic Environment .............. ...... ...............3.
RNA World and the Problem of Ribose Accumulation ................. .........._..._..6
The Role of Minerals on Ribose Formation and Accumulation .........._............8


2 PREBIOTIC SYNTHESIS OF SUGARS .....__.....___ ........... .............


Introduction .............. .. ...... .. .... ..... ... ........
Boron Chemistry and Complexation Mechanism in Polyols ..........................13
pH dependence on the stability of boric acid esters and borate esters.......16
Differential coordination of borate to polyols. ................. ...................1
Interaction of boron with carbohydrates : aldoses and ketoses. ................. 19
Stabilization of pentoses towards decomposition in borate ................... ....23
M materials and M ethods ................. ...............23....... ......
Chemicals............... ...............2

Enzym es .............. ..... ...............24....... ......
Analytic Instrumentation................ ...........2
Ultraviolet analysis s (UV) ................. ........_.. ........24__ ....
Gas chromatography (GC) ................. ....._._ ......... ............2
Mass spectrometry (MS)............... ...............24..
N MR Spectroscopy ................. ......... ...............25.......
Synthetic Preparations............... ..............2
Synthesis of colemanite ................ ...............25...
Synthesis of deuterated Colemanite ................. ........... ................ ..25
Synthesis of pentoses in the presence of colemanite ................ ...............25












Synthesis of pentoses in the presence of calcium hydroxide...................26
Derivatization of pentoses for gas chromatography analysis ....................26
Sugars Degradation Experiments ............... ... .......... .. ....... .... ...........2
Sugar decomposition in the presence of calcium deuteroxide...................27
Sugar decomposition in calcium deuteroxide and colemanite ................... 27
Enzymatic As says .............. ...............28....
Ribitol dehydrogenase assay ................................._ ............2
Cysteine-carbazole test .............. ...............29....
DIOS Analysis .............. .. ...............29...
Preparation of PSi surfaces .............. ...............29....
Competition Experiments .............. ......... ...............3
1, 4-Anhydroerythritol (AET) vs Pentoses .....__ ............. ...... ........._30
13C-Ribose vs Pentoses .............. ...............30....
Results .............. ..... ...............30.
Synthetic Preparations............... .... ... .......3
Synthesis of pentoses in the presence of colemanite ................ ...............30
Synthesi s of pentoses in the presence of calcium hydroxide ................... ..3 2
Sugar Degradation Experiments .............. .............. ... .. .......3
Sugar decomposition in the presence of Calcium Deuteroxide .................33
Sugar decomposition in the presence of Colemanite ................. ...............34
Enzymatic As says .............. .......... ...............3
DIOS Analysis: Competition Experiments. ................... ..............3
1, 4-Anhydroerythritol (AET) vs Pentoses .....__ ............. ...... ........._36
13C-Ribose vs Pentoses .............. ...............36....
D discussion .............. ...............39....


3 CATALYSIS AND THE RNA WORLD .............. ...............43....


Introduction ............... ...............43....
M materials and M ethods ............ _...... .._ ............ .... ...... ............4
Preparation of Precursor DNAzymes via PCR (Maniatis et al, 1982)............. .45
Preparation of single-stranded DNAzymes............... ...............47
5' -End Labeling of DNA ................. ......... ...............48. ..
DNAzyme Kinetic Assays .............. ...............49....
Cloning and Sequencing DNAzymes............... ...............49
In vitro Selection ........._._._..... ..... ...............50....
R results .............. .. ...............54...
In vitro Selection .........._.... .... ...._ ... ...............54....
Cleavage of 614 does not go Completion ................. ... ......... ................ 56
Inhibition by Incompletely Removed Complementary Strand .........................57
An approach to Chemical Equilibrium does not Account for the Plateau........59
Testing if the Cleavage Products are Acting as Catalysts or Inhibitors ............60
Improperly Folded ribose-614 Accounts for Part of the Plateau ................... ...61
Mutations Introduced into 614 during Cloning and Sequencing ......................63

Ribose-614 catalysis is not Mg +-dependent ................ ................. ......66
Ribose-614 cleaves in trans ........._._.. ...... ...............66.











Various ribose-containing substrates are cleaved by deoxyribose-614............_69
Competition Studies of Ribose-614 Cleavage ............... .. ...................7
Saturation kinetics in trans cleavage by deoxyribose-614 .........._..... ..............73
Compound deoxyribose-614 cleaves with multiple-turnovers.............._.._. .......74
Catalytic power in transrt~t~rt~t~rt~t~rt~ is unaffected by annealing protocol...........................74
The commitment step for deoxyribose-614 cleavage. ............ .. ......._.._.....75
Dependence on temperature of deoxyribose-614 cleavage .........._..... ..............77
Predictions of the energetically favored structure ................. ............. .......79
D discussion .............. ...............8 1....


4 DETECTINTG ORGANIC MOLECULES ON MARS .............. ....................9

Introduction .............. ... .......... .......... .. .. .......9
Oxidation of Alkanes Under Martian Conditions ................ ............. .......93
Oxidation of Alkylbenzenes Under Martian Conditions ................. ...............96
Oxidation of PAHs under Martian Conditions .............. ....................9
Oxidation of Kerogen under Martian Conditions .............. ........ ...............9
Oxidation of Amino and Hydroxyacids under Martian Conditions ................. .99
The Amounts and Fates of Organic Carboxylic Acids .............. ...................99
Failure of Viking 1976 to detect Organic Carboxylic Acids ................... .......102
The Infrared Spectra of the Martian Surface ................. ................ ...._.104
Detecting the Missing Organics on Mars ......_. .............. .................1 07
M materials and M ethods ................. ...............108....... ......
Chem icals................... .............10
Analyti c Instrumentati on ...._.._.._ ........... ...............108..
Fluorescence analysis............... ...............10
Infrared analysis................... ... ... .. ..................10
High performance Liquid chromatography (HPLC-MS) ........................108
Synthetic Preparations. ............ ........... ......____ .......... 10
Synthesis of Mellitic Acid Salts ....._____ ............. ....___ ............0
Synthesis of manganous mellitate (1) ....._._.__ ........_._ ........._......109
Synthesis of zinc mellitate (2) ................. ................................109
Synthesis of cupric mellitate (3) ................. .............. ......... .....109
Synthesis of nickel mellitate (4) ................. ........................... ...109
Synthesis of cobalt mellitate (5) .............. .....................109
Synthesis of magnesium mellitate (6) ................. .......... ...............1 10
Synthesi s of calcium mellitate (7) ................. ............... ......... ...1 10
Synthesis of iron mellitate (8) ................. ...............110........... ...
Synthesis of fluoresceins: Fiegl's test ....._____ .........__ ................110
Results ..........._............... ..._ ...........__ ............11
Synthesis of Mellitic Acid Salts ....__ ......_____ .......___ ............1
Fluorescence Spectra Analysis ................. ...............111................
Discussion ................. ...............116......... ......











APPENDIX

A NMR DEGRADATION EXPERIMENTS ................. ............... ......... ...119


B DIOS COMPETITION EXPERIMENTS ................ .............. ....__ .....124

LIST OF REFERENCE S ................. ...............130................


BIOGRAPHICAL SKETCH ................. ...............137......... ......

















LIST OF TABLES


Table pg

2.1i. Berate complexes of aldopentoses, aldohexoses and ketohexoses. ................... .........21

2.2. Retention times of trimethylsilyl derivative of pentoses. ................ ............... .....32

2.3. Half life of pentoses under alkaline conditions determined by 1H NMR. .................35

3.1. Name, sequence, and description of oligonucleotides ................. .........__..........52

3.2. Data from plot In[S]t versus time for ribose-614 cleavage. ................ ................. .68

4.1i. Expected metastable products from organic substances ................. ............. .......93

4.2 Fluore science analy si s of the fluoresceines of b enzenecarb oxyl ates ................... .......11 1


















LIST OF FIGURES


Figure pg

1.1. Proposed autocatalytic glycolaldehyde regeneration. ..........._... ..._... .............5

2. 1. Some organic compounds detected in the ISM. ........._.._.. ...._... ................11

2.2. D-ribose structure................. ..............1

2.3. Lewis structure representation of boric acid and borate. ............. ......................1

2.4. Mechanism of boronic acid complexation by acidic ligands. .............. ..................15

2.5. Berate complexation by non-acidic ligands............... ...............16

2.6. Free energy diagram of threo and erythro diols. ....._._._ ... ...._. ........._.....18

2.7. Free energy diagram of syn-a,y and anti-a,y-diols. ............. ......................18

2.8. Structures of the B-L2 Spirane complex. .............. ...............22....

2.9. Psi surface preparation............... ..............2

2. 10. HPLC-MS analysis of reaction mixture containing colemanite.. ........._... ..............3 1

2. 11. Detection of ribose-borate comples by ESI (-) ion mode ................. ................ ..3 1

2. 12. GC trace of the reaction mixture containing colemanite. ................ ................. ..32

2. 13. GC trace of the reaction mixture containing Ca(OH)2 .........__. ...... .._._...........3 3

2. 14. Incubation of ribose in Ca(OH)2 Solution. ....._.__._ .... ... .__. ......_._.........3

2. 15. Incubation of ribose in the presence of Ca(OH)2 COlemanite. .............. ..............35

2.16. Anhydroethrythritol (AET)-pentose borate ions detected by DIOS.. .......................36

2. 17. DIOS spectra of competition experiment D-arabinose vs AET............... ................37

2. 18. Competition experiments between the different pentoses and AET .......................37

2. 19. 13 ,12C-D-ribose borate ions detected by DIOS. ................ .......... ...............38











2.20. DIO S spectra of competition experiment 13C ribose VS 12C ribose. ................... .......3 8

2.21. Ratio of borate complexes of 13C-ribose vs pentoses. ............. ......................3

2.22. Suggested mechanism for pentose formation.. ............ ...............41.....

3.1. In vitro selection experiment representation ................. ...............45........... .

3.2. Sequence of the initial library and DNAzymes. ............. ...............46.....

3.3. Ribose-614 cleavage .............. ...............57....

3.4. Cleavage products do not affect ribose-614 cleavage .............. .....................6

3.5 Gel-purification of ribose-614 at cleavage plateau ........................... ...............62

3.6. Reheating ribose-614 results in additional cleavage .............. ....................6

3.7. Sequence alignment of cleaved and uncleaved cloned 614. .................. ...............65

3.8. Initial rate of ribose-614 cleavage as a function of [ribose-614] ............... .... ...........67

3.9. Ribose-614 cleavage rate is concentration dependent .............. ....................6

3.10. Both deoxyribose-614 (left panel) and ribose-614 (right panel) ............................70

3.11. Cat ribose competes with ribose-614 for cleavage .................... ...............7

3.12. Compound deoxyribose-614 can cleave various sub states ................. .................7 1

3.1 3. Various sub states can compete with ribose-614 for self-cleavage............._.._........72

3.14. Cleavage of various substrates by 614 is reduced. ............. .....................7

3.15. Ribose-614 rate of self-cleavage in transrt~rtrt~rt~r~rt~rt ........._._. ._...... ._ __ ................7

3.16. Burst kinetics. ............. ...............79.....

3.17. A univariate statistical distribution. ............. ...............82.....

4.1 Oxidative degradation of the generic alkane. ............. ...............95.....

4.2.Oxidative degradation of naphthalene to phthalic acid. .........._...._ ........_.. .........98

4.3. Martian surface IR spectrmm................ ..............10

4.4. Mars dust, labradorite standard and labradorita............... ..............10

4.5. Mars dust and magnesita-labradorite mixture spectra. ............. ......................0











4.6. Phthalic acid yield fluorescein. .............. ...............108....

4.7. Infrared spectra of manganous mellitate (KBr). ................ ......... ................1 12

4.8. Infrared spectra of zinc mellitate (KBr) ................. ...............112.............

4.9. Infrared spectra of cupric mellitate (KBr). ................ ...............113........... ..

4. 10. Infrared spectra of nickel mellitate (KBr) ................. ...............113........... .

4. 11. Infrared spectra of cobalt mellitate (KBr), ................. ...............114............

4. 12. Infrared spectra of magnesium mellitate (KBr) ................ .......... ...............1 14

4. 13. Infrared spectra of calcium mellitate (KBr) ................. ...............115............

4. 14. Infrared spectra of iron mellitate (KBr) ................ ...............115.............

4. 15. Infrared spectra of aluminium mellitate (KBr) ................ ......... ................1 16

4. 16. Infrared spectra of the mellitate salts ................. ...............117........... .

4. 17. Structures of the fluorescein derivatives of pyromellitic acid. ............. ..... ...........1 18

A1.1. D-arabinose incubation in the presence of calcium hydroxide, pD: 12. ...............120

Al.2. D-arabinose incubation in the presence of calcium-hydroxyde + borate............_...120

Al.3. D-lyxose incubation in the presence of calcium hydroxide, pD: 12. ................... ...121

A1.4. D-lyxose incubation in the presence of calcium hydroxide + borate pD:12........121

Al.5. D-ribose incubation in the presence of calcium hydroxide, pD: 12........................ 122

A1.6. D-ribose incubation in the presence of calcium hydroxide + borate, pD: 12. ........122

Al.7. L-xylose incubation in the presence of calcium hydroxide, pD: 12. ...................... 123

Al.8. L-xylose incubation in the presence of calcium hydroxide + borate, pD: 12. ........123

A2.1. DIOS spectra of 1,4-Anhydroerythritol vs arabinose. ............. .....................2

A2.2. DIOS spectra of arabinose VS 13C-ribose. ............. .....................124

A2.3. DIOS spectra of 1,4-Anhydroeythritol vs lyxose................. ...............12

A2.4. DIOS spectra of lyxose vs 13C-ribose ................ ................ ........ ...._..125

A2.5. DIOS spectra of 1 ,4-Anhydroerythritol vs ribose ................. ................ ...._.126










A2.6. DIOS spectra of 12C-ribose VS 13C- ribose ................ ...............126.............

A2.7. DIOS spectra of 1,4-Anhydroerythritol vs Xylose .............. .....................2

A2.8. DIOS spectra of xylose vs 13C-ribose ................ ...............127.............

A2.9. DIOS spectra of 1 ,4-anhyroerythritol vs ribulose ................. ........................128

A2. 10. DIOS spectra of ribulose VS 13C-ribose ................. ...............128........... .

A2. 11. DIOS spectra of Xylulose vs 13C-ribose. ................ ..............................129















Abstract of Dissertation Presented to the Graduate School
of the University of Florida in Partial Fulfillment of the
Requirements for the Degree of Doctor of Philosophy

BIOORGANIC MOLECULES INT THE COSMOS AND
THE ORIGIN OF DARWINIAN MOLECULAR SYSTEMS

By

Alonso Ricardo

May, 2004

Chair: Steven Benner
Major Department: Chemistry

Two critical unsolved issues in the origins of life field are the prebiotic formation

of the molecular building blocks of life and from these, the appearance of a self-

replicating molecule that undergoes Darwinian evolution. In the present dissertation, both

issues were addressed by an experimental approach from which the following findings

are reported.

A plausible prebiotic route for the synthesis of sugar pentoses starting from

materials known in the interstellar matrix was achieved. Ribose, one of the pentoses and

constituents of ribonucleic acid, was generated from a reaction mixture containing boron

minerals. The role of boron in this process was found to be dual. Boron coordinates to

glyceraldehyde blocking the enolization process and binds to the pentose sugar

preventing decomposition. The formation of ribose appears to be the natural consequence

of the intrinsic chemical reactivity of compounds available from the interstellar medium

under alkaline, calciferous conditions. As these conditions are not excluded from the









early Earth, it is also not possible to exclude the availability of pentoses at the time when

life originated.

In vitro selections performed in the presence of Mg++ generated DNA sequences

capable of cleaving an internal ribonucleoside linkage. Several of these, surprisingly,

displayed intermolecular catalysis and catalysis independent of Mg +, features that the

selection protocol was not explicitly designed to select. A detailed physical organic

analysis was applied to one of these DNAzymes, termed 614. The DNAzyme 614 is more

active in trans than in cis, and more active at temperatures below the selection

temperature than at the selection temperature. Many of these properties are unreported

in similar systems, and these results expand the phenomenology known for this class of

DNA-based catalysts. A brief survey of other catalysts arising from this selection found

other Mg++-independent DNAzymes, and provided a preliminary view of the ruggedness

of the landscape relating function to structure in sequence space.

Finally, in the last chapter of this dissertation a method was designed, that allows

the evaluation and detection of potential organic molecules on Mars. The method was

tested with synthetic salts of mellitic acid, that are likely to be formed under Martian

conditions. The presence of these molecules in the martian soil, was evaluated by direct

comparison with the recently published Infrared spectra of the Mars surface















CHAPTER 1
ORIGINS

Background and Significance

Prebiotic Chemistry

Experimental prebiotic chemistry as a modern approach to study the origins of life

was born just over 50 years ago, in the work of Stanley Miller (1953). Miller

demonstrated that applying electric discharges to a mixture of reduced gases in the

presence of water generates a brown solution containing amino acids. Neither sugars nor

nucleobases were produced under these conditions. Miller's experiment, which at the

time was considered to reproduce conditions on early Earth atmosphere's, was a modern

example of the abiotic origin of biological molecules.

Since then, the approach in prebiotic experiments remains the same. A mixture

containing molecules believed to be present in a prebiotic environment is exposed to a

source of energy. If any amount of the desired product is detected, it is then claimed that,

if there was plausibly a historical moment during which the Earth contained the starting

materials, it is then possible to assume that the mentioned reaction took place, allowing

the compound to accumulate over long periods of time.

This logic seems intuitively valid. The methodology, however, can be flawed at

different stages:

*The retrosynthetic analysis used to determine the most likely starting materials,

may ignore the geochemical constraints necessary to make the reaction relevant.










Starting materials can be chosen mainly based on reactivity rather than prebiotic

relevance.

* Water is included/excluded in the model by convenience; no logical reasons other

than the instability of reactants and products in the chosen solvent are addressed.

* In some cases concentrations of reactants are controlled at stoichiometric ratios to

bias the reaction towards one desired product. This is a common practice in organic

chemistry. When applied to prebiotic chemistry, however, this practice fuels

arguments that origin required intelligent design. It is worth mentioning that

limiting reagents are not prohibited as long as their abundance can be explained

through geochemical constraints.

The statements expressed above do not necessarily apply to prebiotic chemistry

elsewhere in the universe. For example, an inventory of interstellar compounds detected

by radioastronomical methods and the inventory of organic compounds found in

meteorites suggest that the non-terrean chemical repertoire is rich in molecules that have

a short half life under terrean conditions, but nonetheless exist in the cosmos. These

molecules constitute valuable starting materials for terrean chemistry.

Biologically relevant compounds may eventually arise when interstellar material

delivered to the Earth (by meteor or comet impact) interacts with terrean molecules,

volcanic emissions, water, or surface minerals. Therefore, for a prebiotic experiment to

be meaningful, chemistry and geology must be linked; only through a concise analysis of

the chemical evolution (organic and inorganic) of our planet will we be able to explore

the possible outcomes of an experiment within natural constrains.










In the end, this methodology will not conclude how in fact life began, or what

chemistry indeed happened, but at the very least, will offer evidence of a plausible

mechanism (able to suffer scientific scrutiny) for the formation of key molecules present

in modern life.

Sugars in the Prebiotic Environment

Sugars, and pentoses in particular, are one of the building blocks of nucleic acids.

Other sugars, including hexoses, are key throughout metabolism and structural

biochemistry. Sugars therefore are logical targets for prebiotic chemistry experiments.

Early attempts to synthesize sugars from simpler molecules were not done

explicitly to reproduce prebiotic events. In 1861 Butlerow reported the formation of a

brown, sweet tasting compound resulting from the reaction of an aqueous formaldehyde

solution in the presence of calcium hydroxide (Ca(OH)2). The product had a molecular

formula (calculated by elemental analysis) corresponding to C7H1406, and was named

"methylenitan." The term "formose" to describe the same product composition was

introduced by Loew in 1886 while reacting gaseous formaldehyde and calcium

hydroxide. Further characterization of the reaction products by derivatization showed that

instead of being a single product, the formose sugar was a heterogeneous mixture of

monosaccharides. This suggested a complex reaction mechanism with many possible

outcomes.

The first mechanistic studies on the formose reaction focused on the isolation of

intermediates. From this work, it was found that glycolaldehyde, glyceraldehyde and

dihydroxyketone were early addition products of formaldehyde condensation (Henry,

1895; Neuberg, 1902). These compounds can then react with either additional









formaldehyde, or cross-react to produce a mixture containing tetroses, pentoses, hexoses,

and branched sugars, in both aldo- and keto- forms.

A kinetic analysis of the formose reaction shows an initial period during which

products are not made (the induction period). This is followed by a period in which all

the compounds are formed relatively quickly.

Breslow suggested a mechanism to explain the formose reaction in 1959 (Figure

1.1). According to Breslow, if any direct j oining of formaldehyde molecules exists, it

must be very slow. In this way, he explained the induction period. The novelty in

Breslow' s work was to introduce the concept of an autocatalytic reaction during which

the first addition product, glycolaldehyde, is regenerated. Because the autocatalytic

process does not require condensations between two electrophiles (as in the condensation

of formaldehyde), the autocatalytic formation of glycolaldehyde proceeds at a fast rate in

alkaline conditions.

Several authors tested this mechanism by starting the reaction from the

intermediates suggested by Breslow (Pfeil & Ruckert, 1960; Ruckel, Pfeil & Scharf,

1965). These authors obtained similarly complex products. Further, the addition of

glycolaldehyde at the beginning of the reaction was found to reduce significantly the

induction period.

With the invention of chromatography, formose sugar composition was better

characterized and found to consist of 10% C4, 30% Cg, 55% C6, 5 % > C6 (detectable by

GC-MS analysis) sugars when formaldehyde is ca. 99% consumed (Weiss et al. 1970;

Decker et al. 1982).











ca

H C O O OH O O H
C--OH C-C H H HC- -C H)
~OH HH C 77 HJ H
Glycolaldehyde H O
Formaldehyde Glyceraldehyde


O OHFormaldehyde
HO OHH


H/ H H HH
HJ H C
~OH H
dihydroxyacetoneH

Glycolaldehyde


OH OI OH H OH O) OHP~ C--C H
Isomerization OH
H- C- C-C---- H H

H3 H H H H HO OH

~O2-tetrulosa loers

Glycolaldehyde
enediol

Figure 1.1. Proposed autocatalytic glycolaldehyde regeneration in the formose reaction.

Parallel to the discovery of the formose reaction, studies on the chemical


composition of nucleic acids were also yielding some important results. In a series of


papers between 1891-1894 Albrecht Kossel deduced the structure of the heterocyclic


bases that constitute nucleic acids. Levene and Jacobs (1909) completed the structure of


the nucleoside of the RNA (known at the time as nucleicc acid of plant origin") by


finding that the carbohydrate portion was made of the sugar D-ribose. The elusive


structure of the carbohydrate in DNA (named nucleicc acid of animal origin") remained


unsolved until in 1929, when Levene and London concluded that deoxyribose was indeed


the sugar component.









With the advent of prebiotic chemistry in the 1960's, it was not long before the

formose synthesis was reexamined from a prebiotic context as a way of obtaining the

sugars necessary for biology. With the realization that ribose is the backbone of RNA,

and having detected this sugar as a product in the formose reaction (although ribose

overall yield and long term stability were low in the experiment), prebiotic chemists were

satisfied, with the idea of having found that ribose was prebiotically available.

RNA World and the Problem of Ribose Accumulation


In 1962 Alexander Rich hypothesized that RNA might have played both catalytic

and genetic roles in early forms of life. This idea was furthered by comments from Woese

(1967), Orgel (1968) and Crick (1968), who remarked that transfer RNA (tRNA)

appeared to be an RNA molecule attempting to fold like an enzyme. The discovery of

catalytic RNA by Cech, Altman, Usher, and others supported this notion (Kruger et al.

1982; Guerrier-Takada et al. 1983; Usher & McHale, 1976). Gilbert proposed in 1986

that organisms in an "RNA world" may have been a precursor to the contemporary

protein-DNA-RNA organisms that dominate life on Earth today. In the RNA world, life

used RNA as the sole genetically encoded biological catalysts. However, the plausibility

of the RNA world hypothesis is obviously conditioned to the success of generating RNA

from a plausible prebiotic soup.

The difficulties of generating RNA without the assistance of a pre-existing living

system have been noted by many authors and can be divided into two categories: (a)

those directly related to the activity of water, and (b) those that are not.

In the first category, RNA is thermodynamically unstable in water with respect to

hydrolysis. Further, many RNA nucleobases are hydrolytically unstable. Cytidine, for










example, deaminates with a half life of ca. 102 years to give uridine (Frick, Mac Neela &

Wofelden, 1987). Adenosine deaminates to give inosine at a slower rate, while guanosine

deaminates to give xanthosine. These reaction modes, favored at high pH, are matched by

depurination and depyrimidinylation reactions at low pH, which are also favored

thermodynamically in water.

A series of criticisms of the "RNA origins of life" hypothesis are not directly

related to water, however, but rather concern ribose itself. These focus on difficulties of

creating the necessary amounts of ribose to support a RNA world in the early Earth, and

the instability of ribose under prebiotic conditions where it might be generated. This

problem was addressed by Robert Shapiro in 1988, who, focusing on the formose

reaction, concluded that the synthesis and accumulation of ribose in any significant

amount under prebiotic conditions were very unlikely events.

Shapiro's comments are correct; the overall yield of ribose in formose is less than

1% after an arbitrary length of time, and less if the incubation is allowed to continue

indefinitely. Ribose itself contains both an electrophilic center (carbon-1) and a

nucleophilic center (carbons 1 and 2 of the enediolate) (Figure 2.2). This makes ribose

unstable under basic conditions with respect to further reactions with formaldehyde,

glycolaldehyde, or itself, or other nucleophiles and electrophiles that are emerging under

formose conditions.

Not surprisingly, in the presence of Ca(OH)2, ribose is converted to higher

condensation products, branched chain sugars, and (ultimately) a brown, largely

intractable polymer of undefined composition. "Browning" of the mixture is pronounced

within an hour at room temperature and within minutes at 60 oC. Thus, the extent of









accumulation of ribose as a product of formaldehyde and glycolaldehyde at steady state

(formation minus destruction) is quite low. Given this, Stanley Miller and coworkers,

commented that the rate of this decomposition reaction is so high that it suggested that

"the backbone of the first genetic material could not have contained ribose or other sugars

because of their instability" (Larralde, Robertson & Miller, 1995).

The Role of Minerals on Ribose Formation and Accumulation

While the scientific literature contains many reports concerning the use of minerals

in prebiotic experiments, few of these publications include a geochemical explanation of

how and why the mineral was available on the early Earth.

Ponnamperuma reported the use of the clay mineral kaolin (aluminium hydroxide

silicate (Al2(OH)4. Si205) a weathering product of feldspar), as a catalyst in the formose

reaction at low concentrations of formaldehyde (Gabel & Ponnamperuma, 1967). Kaolin

was shown to facilitate the condensation reaction of formaldehyde at a pH lower than that

of calcium hydroxide solutions. Because of this, the sugar products obtained were stable

over longer periods of time. Still, the problem of selective formation of ribose was not

solved; the overall yield was later calculated by Miller to be approximately 3.8% (Miller,

1984), again after arbitrary time under arbitrary conditions.

Recently, Zubay (1998) reported the use of a combination of lead and magnesium

salts in the presence of formaldehyde to generate aldopentoses that constituted 30% of

the total product. Here, the lower basicity and solubility of lead and magnesium

hydroxide was exploited to moderate the formose decomposition processes. The

discussion on the geochemistry of lead was only limited to a list of lead containing

minerals.















CHAPTER 2
PREBIOTIC SYNTHESIS OF SUGARS

Introduction

We do not know what organic molecules were present on the early Earth. We may,

however, look at compounds in the interstellar medium (ISM), within meteorites, in

comets, and in other solar system bodies, to provide a clue.

No direct information indicates what fraction of interstellar and cometary

compounds would be delivered to early Earth in unaltered form. It is likely that some

would be transformed in icy bodies, especially by high energy particles and photons

(Bernstein et al. 2002). It is also known that Earth-based chemistry would influence the

composition of material. Hydrogen cyanide (HCN), is generated by example, upon comet

impact, by an unknown mechanism. At this point in our development of knowledge of the

chemistry of the solar system, it is pragmatic to assume a set of compounds such as those

shown in Figure 2.1 as our starting point.

How might the molecules in Figure 2.1 be transformed on Earth to give ribose in a

stable form ? It is clear that the prebiotic soup would be exposed to rocks and minerals

that have some solubility in water at atmospheric pressure (these are found on modern

Earth as evaporates). In this light, we re-examined the formose reaction, recognizing that

both formaldehyde and glycolaldehyde are found in the ISM.

In its native form, glycolaldehyde can act as an electrophile (the carbon of the C=0

group) and as a nucleophile (the alpha carbon, once the 2-position proton is abstracted).

As the enolate, however, glycolaldehyde can act only as a nucleophile. To facilitate the









enolization, a cationic species that coordinates the two oxygens of the enediol is needed.

The O-C=C bond angle is 120 o. This places the two oxygens at some distance, implying

the need for a large metal ion to bridge the long O-O distance. The large Ca++ serves this

role. Indeed Ca" was the cationic species originally used by Butlerow in 1861, and is the

most common catalyst used at present for the formose reaction.

Formaldehyde can act only as an electrophile. The calcium-stabilized enolate of

glycolaldehyde can react only a a nucleophile. Therefore, the reaction of formaldehyde

and the calcium-stabilized enolate of glycolaldehyde is constrained to give

glyceraldehyde as a pair of enantiomers. Glyceraldehyde has a 1,2 diol unit.

Glyceraldehyde, however, can act intrinsically both as an electrophile (the carbon

of the C=0 group) and as a nucleophile (the alpha carbon, once the 2-position proton is

abstracted). The ability of glyceraldehyde to act as both a nucleophile and an electrophile

means that it can cross-react to form compounds that resemble tar. For example, reaction

of glyceraldehyde as a nucleophile with formaldehyde gives a branched sugar lacking the

1,2-diol moiety. Tar formation is, of course, the standard outcome of the formose

reaction, which has been criticized for its prebiological relevance, as noted above.

When glyceraldehyde acts as an electrophile with the calcium-stabilized enolate of

glycolaldehyde acting as a nucleophile, however, a pentose is the only product. Four

enantiomeric pairs of diastereomeric pentoses exist: ribose, arabinose, xylose and lyxose.

As it is drawn in the open chain form, it appears as if ribose can also act both as an

electrophile and as a nucleophile (Figure 2.2). This would also permit it to form

undesired further products.










H/O'H


O=C=S


H--CEN O=C=0


O
I I
H/C'H

NH
I I
H'C


H

H'N'H

O
I I
HOZ,CH


S
I I
H/C'H


H-CEC--H

H--N=C=0


H--N=C=S

H--CEC--N EC
H--C-C--C-C--H
H-CEC--CEN


N
III
C
C
H I'H
H


I-
N
C
H I'H
H


O

I I

H


O
H--C-C--H
H
H H


H H
.C=C
H H


H
,N--CEN
H


H--CECC c
1H


H
C=C=0
H


H H
O S
I I
H CH H C'H
H H


H O
H--C ~OC


H I


H/ '',
I\
HH


H-C-C--C-C--C-N
H H
H--CEC--CEC--CEC-H N

HC--C C--C C-H H I'H
HI H

\ N
C--C C--C N ^//
/I H /C
H C

H--CC-C_C--CC--CN /C
H H
H-CEC-CEC-CEC-CEC-CEN

H-C-C-C-C-C-C-C-C-C-C-C-N


H H

H--C CG,C
H I


H H
H--C C
/ 'O' l' H
H H


H C\CO
H 11
H H


C ,C9N
HIC
H 11
H H
Figure 2.1. Some organic compounds detected in the ISM.










It is clear, however, that these pentoses cannot do so in their ring closed form.

Thus, ribose closes to give either a six-membered ring (a pyranose, both in the alpha and

beta anomeric forms, about 75% of the total) or a five membered ring (furanose, both in

the alpha and beta anomeric forms, about 25% of the total).

We then asked: what mineral components might stabilize the glyceraldehyde

against undesired reaction, while directing it towards the reaction that creates ribose?

n ce p iielectrophili c CO HO OH
center~ ~o

center
| O~O
acidiC H C OH B
hydrogen
H-C OH O O


H COH OH

D-ribose (open diribose-borate complex
form with reactive (the cyclic form, stabilized
C=0) by borate,is largely unreactive)
Figure 2.2. D-ribose structure. The open form of ribose contains electrophilic and
nucleophilic centers.

Here, we do not seek a large completing species that can bridge the distant

oxygens on an enediol, but rather a small completing species that can bridge the short

oxygen-oxygen distance on a 1,2 diol. Here the O-C-C bond angle is only 1090. The most

obvious completing species for this purpose is small borate. Berate is well known to

form a complex with diols, with a micromolar dissociation constant (Boieseken, 1949).

As a borate complex, glyceraldehyde can act as an electrophile. The C=0 group is

not affected. The borate complex of glyceraldehyde is not expected to enolize easily,

however, under alkaline conditions. Abstraction of the 2-proton by base is discouraged by

the negative charge already on boron.









Interestingly, borate is expected to stabilize the cyclic form of ribose as well

(Figure 2.2). The cyclic form presents two hydroxyl groups in a cis configuration. It is

well known that borate complexes with 1,2-diol and 2,3-diol are specially stable. These

observations generated the hypothesis that is tested in this chapter: Perhaps borate, if it

were present under formose conditions, would manage the reactivity of glyceraldehyde,

and stabilize ribose, the desired product.

Boron is known in carbonaceous chondrites, where it is almost certainly present as

borate (Zhai & Shaw, 1994). Boron is relatively scarce, relative to carbon and other light

elements, however, due to the inefficiency of its synthesis in nuclear reactions.

Berate is, however, excluded from many silicate minerals. For this reason, it

appears in the residual melts as lava cools. Here, it is found in tourmalines, minerals that

are found in many forms, including colorful forms used as gemstones. Tourmaline

weathers from rocks as they are exposed on the surface to generate borate salts, which are

generally modestly soluble in water. For example, colemanite is soluble in water to the

extent of 0.82 g/L. As a consequence, colemanite and other borate-containing minerals

are found in deserts and other dry environments, often under alkaline conditions. Here,

they are known as evaporites, as they are crystallized from water as it evaporates. These

evaporites form under conditions that are close to the conditions that generate pentose.

Boron Chemistry and Complexation Mechanism in Polyols

A clear understanding of the chemistry of boron is necessary for the evaluation of

our hypothesis. Boron has five electrons, which can be assigned to an electronic

configuration I s2 2s2 2px In the hybrid orbital state, one electron from the 2s orbital is

promoted to the 2p orbital (2sl 2pxl 2pf ) to make a an sp2 hybrid orbital in which each of

the three electrons is located in an orbit and able to accept one electron from another










element to form a covalent complex with the boron atom (i.e.: B(OH)3 boric acid). The

additional 2p electron orbit is able two hold a pair of electrons from another element,

(which completes the octet around boron) this property explains the Lewis acid character

of trigonal boron complexes.



Hydroxvl ion : O
H;O'B01;H -OH **
.x : H:0B.O:H
.. pKa=-9. 1 :O:
H *+
Boric Acid
Borate lon
Figure 2.3. Lewis structure representation of boric acid and borate.

Boric acid has a trigonal planar structure in which the B-O bond length is 1.37 1.

In the tetrahedral borate ion, the B-O bond length is 1.48 a, which makes the hydroxyl

group a better proton acceptor (more basic) and therefore a better leaving group when

compared to the hydroxyl group in trigonal boron (Pizer & Tihal, 1992).

The reactive form of boron trigonall vs tetrahedral) towards complexation is

dependent on the pH of the solution under study. Is generally assumed that in reactions

carried out at the pKa of boric acid (and above), the reactive species is borate, while the

trigonal boron is responsible for the reactivity at lower pH values.

Van Duin et al. (1984) suggest that aqueous boric acid exists as an adduct with a

water molecule, to give a species whose geometry is tetrahedral, but still neutral. This

suggestion makes sense when explaining the reactivity of boric acid towards

esterification reactions, in this way a loosely bound water molecule is easily substituted

by a hydroxyl group).









The complexation of boric acids and borate with dicarboxylic acids, a-

hydroxycarboxylic acids, diols and polyols, has been studied in detail for many years

(Mazurek & Perlin, 1963; Pizer & Kustin, 1968; Davis & Mott, 1979; Van duin et al.

1984; Van duin et al. 1985; Verchere & Hlaibi, 1986; Pizer & Tihal, 1996; Ito et al.

2003).

In the case of acidic ligands such as dicarboxylic acids, a-hydroxycarboxylic acids

and 1,2 dihydroxybenzenes, the mechanism of boronic acid complexation (at pH values

lower than the pKa of the boronic acid under study) is believed to occur through the

nucleophilic attack of a hydroxyl group of the ligand (1) to the trigonal boron (2)

generating an associative transition state in which a proton from the entering ligand is

transferred to a leaving hydroxide originally coordinate to boron (Figure 2.4). (Kustin &

Pizer, 1968; Pizer & Tihal, 1992, Pizer & Tihal, 1996)




| + HO-C O------O -C R
,a O H-C= 0 Ro B--------p--OCO HOBOC +H0~
1 2 HO


Figure 2.4. Mechanism of boronic acid complexation by acidic ligands. In boric acid (R=
OH), the resulting borate monoester (3), has the potential to form a bis-
substituted complex by reacting with an additional ligand molecule.

In non-acidic ligands, such as diols and polyols, the predominant complexation

reaction occurs with the tetrahedral borate (Figure 2.5). The mechanism of complexation

to borate its not yet well understood. But it has been suggested (Pizer & Tihal, 1992) that

complexation occurs in two steps, is probably associative in character, and also involves

proton transfer. An associative mechanism does not necessarily imply an increase in the

coordination number on boron. But even if this is the case, hypervalent pentacoordinated










boron complexes are known in the literature (Yamashita et al. 2000; Lee & Martin,

1984). Therefore, a pentacoordinated transition state in borate complexation remains as a

possibility.

A.


HOn OH HOC rds O- -C --c- atHO OC-
"*B HO,- + H20 B + H20
OHHO--C-H RB HO ,--
R HO OH H O
B.
H HHH

HC..0 OH HOH H -~- H C',.-O OC- H
H-- ~ + |OH |OC- B-~/~ + 2 H20

H H H H

Figure 2.5. Berate complexation by non-acidic ligands. A) Attack of the hydroxyl moiety
of the diol to borate is rate limiting. The cyclization reaction to make the five
member borate ester is fast. B) In the case of esters of boric acid at high pH,
bis-substituted complexes are form by reaction with an additional diol
molecule.

pH dependence on the stability of boric acid esters and borate esters.

11B NMR experiments allowed comparison of esters of boric and borate formed by

diols, a-hydroxycarboxylic acids and dicarboxylic acids (Van Duin et al, 1984). In

general it was found that boric acid esters of a-hydroxycarboxylic acids proved to be

more stable than those of diols or dicarboxylic acids. Also, it was concluded that the

optimal pH stability of boric acid and borate esters, can be predicted by using the "charge

rule" :

"Esters of boric acid and borate in aqueous medium show the highest stability at

that pH where the sum of the charges of the free esterifying species is equal to the charge

of the ester."









This rule is better illustrated as:

* Esters of boric acid are most stable at low pH where dissociation of B(OH)3 and

ionization of the bidentate ligand (L) hardly occurs (for carboxylic acids and

alpha-hydroxyacids).

B(OH)3 + L B(OH)L + 2H2,0
* Dissociation of boric acid favors formation of borate esters of 1,2-diols. A pH >

pKa of boric acid is necessary (in other words, diols only form borate complexes at

an alkaline pH).

B-(OH)4 + nL B-Ln + 2n H120
* In the case of a-hydroxycarboxylic acids and dicarboxylic acids, pH dependent

optima are involved. The borate diester of an a-hydroxycarboxylic acid shows

maximal stability at pH = pKa (L) :


B(OH)3 + L + L- BL, + 3H,O
* The borate monoester, however, occurs preferentially at pKa (L) < pH < pKa boric

acid. A similar situation applies to dicarboxylic acids at pH = (pKai + pKa2)/2

B(OH)3 + L- B-(OH)2L + H120
The previous rules agree with the expected behavior for a reaction with an

associative transition state in which proton transfer is involved postulated by Pizer &

Tihal in 1992.

Differential coordination of borate to polyols.

The preferentiality of borate binding to different diols and polyols was studied by

measuring the association constants of different borate complexes (Van Duin et al, 1985).

Experiments were done by 11B NMR at a high pH allowing only borate ions to be the











reactive species. The results of the relative stabilities of complexes obtained by these

experiments can be summarized as:

* Tridentate borate > bidentate borate > monodentate borate complexes

* Increasing the number of hydroxyl groups increases the stability of borate esters.

* Diols in a threo-aP diol conformation > terminal diol > erythro-a,P diol (explained


by the steric interactions of the R' groups (Figure 2.6), in the case of terminal diols

a loss of enthropy is responsible for the difference)

threo- n,P-diol erythro- n,P-diol


OH
R'


GBT


R OOHGrBE RII" O B bOH

RIl~nn O OH Where GE-Ts GT + 1 kand GBE GBT +
G= kcal/mol

Figure 2.6. Free energy diagram of threo and erythro diols and their borate complexes.

*Coulumbic repulsion decreases stability (introduction of carboxylate residues in the

diols, creates repulsion with the negatively charged borate)


*syn-a,y diols > anti-a,y-diols (Figure 2.7).

syn-a,y &o anti-a,y &o

4OH





AG~s OH AGBA




Whee G~o G,-1 and GBDG,,+1 5
G kcal/mol
Figure 2.7. Free energy diagram of syn-a,y and anti-a,y-diols and their borate complexes.









Interaction of boron with carbohydrates : aldoses and ketoses

The interaction of boron with carbohydrate molecules has been studied in the past

with special interest due to the potential applications in stereochemistry and structure

determination. Also in the case of carbohydrate mixtures, chromatographic separations

may be improved by having boron preferentially coordinated to specific molecules within

a family of structurally related sugars.

Several methods have been published to measure the association constants (Kasso)

between boron and carbohydrates: Potentiometry, pH titration, (11B, 13C 1H) NMR,

calorimetry. However, reported values in the literature differ dramatically depending of

the technique used, reasons for these differences other than experimental error include:

* Uncertainty of the anomeric and conformational composition of carbohydrates-

borate complexes at equilibrium.

* Stoichiometric ratio of borate(boric)-carbohydrate species is assumed to be

homogeneous at a certain pH value and either consistent of borate monoester or

borate diester complexes ( work by Van Bekkum's group has shown that borate

monoester and diesters coexist at certain pH values)

Reproducible values of association constants have been obtained by using a

combination of techniques. Mazurek & Perlin (1963) used thermometric measurements

of vapor pressure equilibria and 1H-NMR to determine the different complexes of borate

with D-glucose, D-threose and cis-3,4-dihydroxytetrahydrofuran (Table 2.1). The most

important conclusion of their work is the finding that the complexation of D-glucose to

borate proceeds via a pyranose to furanose interconversion. This finding is important,

since the calculation of association constants by other groups, assume that the ratio of










sugar conformers in the presence of borate is the same as in the aqueous sugar without

borate. Mazurek also suggested that pyranose cis-diols do not make strong borate

complexes, and that a sugar-borate ratio 2: 1 yields spirane complexes of the sugars under

study .

Verchere & Hlaibi (1987) presented the first comprehensive analysis that considers

the effect of borate in the conformational equilibrium of carbohydrates and later these

authors include the contribution of this effect in the calculation of the association

constants. By using a combination of potentiometric titration and 1B NMR spectroscopy,

Verchere confirmed that 1:1 and 2: 1 carbohydrate-borate complexes were obtained for

every sugar and then calculated the stability constant for each of those complexes (Table

1.1) (Chapelle & Verchere, 1988; ibid, 1989). Structural information of the different

borate complexes was also deduced from NMR studies (Figure 2.8), allowing a direct

interpretation of the stability constants for the monoester P; and the diester p2 -

General conclusions from data in Table 2.1 and Figure 2.8 included:

* While sugars in aqueous solution mainly adopt the 6 member ring pyranose

conformation, in the presence of borate, this configuration is forced into the Hyve

member borate-furanose form.


* At limited amounts of borate, formation of spirane complexes is preferred over the

monochelate complex.


* The trend in stability of spirane comples in the monosaccharides shown is :


ketohexoses > aldopentoses > aldohexoses










Table 2. 1. Berate complexes of aldopentoses, aldohexoses and ketohexoses. n.d: not
determined. Only the strong furanose-borate esters are shown.
Sugar Donor sites B La Stability Stability % Complexed
(furanose form) (borate ester) constant(log Pr) constant (log as B-L2
B (OH)2L Pt) B-L2
1,4- 2,3 n.d n.d n.d
anhydroerythritol
P-D-threose 1,2 n.d n.d n.d
a-D-ribose (70%) 1,2

P-D-ribose (30%) 2,3 22 .89
P-D-arabinose 1,2 2.14 2.99 30 %
a-D-xdlose 1,2 1.95 3.74 80 %
a-D-lyxose (60%) 2,3

P-D-lyxose (40%) 1,2 21 .95
a-D-allose 1,2
p-D-llos 2,3n.d 3.9- 4.4 85 %

a-D-glucose 1,2 1.80 3.05 55 %
a-D-mannose 2,3 2.01 2.74 20 %

a-L-galactose 1,2 1.99 2.56 20 %
a-D-psicose 2,3 n.d 2 6 > 99 %
p-D-fructose 2,3 2.82 4.97 n.d
a-D-sorbose 2,3 < 3.5 5.75 n.d
a-D-tagatose 3,4

P-D-tagatose 2,3 n.d n.d n.d



*The anomeric hydroxyl posses a high reactivity towards the formation of borate

complexes. In those cases where the anomeric hydroxyl is not involved in

completing, is found in a trans position to the vicinal borate-diol ring system.


*Within the aldopentoses, the trend in stability of spirane borate complexes follows:


ribose > xylose > lyxose > arabinose

This difference in stability can be explained by considering steric effect between

the R1 group: CH20H and the borate ring. In 1,2-cis coordinated ribose and xylose













R1B-O0
O,



H R2
HOH

1,2-P-D-arabinofuranose R = CHOH: R,= H
1,2-ac-L-galactofuranose R = CHOH-CH2OH: R = H
2,3-P-D-fructofuranose R = R,= CHOH



R1 O O)H
H H

H R2





2,3-P-D-ribofuranose R = CHOH : R, = H
2,3-P-D-allofuranose R = CHOH-CH2OH: R = H


1,2-ac-D-xylofuranose R1= CH2OH R, = H
1,2-ac-D-glucofuranose R,= CHOH-CHOH: R, = H
2,3-ac-D-sorbofuranose R1= R2=CH OH



R1 O R2
H H


H~ O




1,2-ac-D-ribofuranose R = CH OH; R, = H
1,2-ac-D-allofuranose R = CHOH-CH2OH: R2 = H
2,3-ac-D-psicofuranose R = R = CH2OH


B R1 B-O0



H RLF2
H OHH H
HHH

2,3-ac-D-lyxofuranose R = CHOH :R, =H 1,2-P-D-Lyxofuranose R = CH2OH :R2 -H
2,3-ac-D-mannofuranose R1= CHOH-CH2OH R2= H 1,2-P-D-mannofuranose R = CHOH-CH2OH: R = H
3,4-ac-D-tagatofuranose R = R = CH2OH 2,3-P-D-tagatofuranose R = R = CH2OH

Figure 2.8. Structures of the B-L2 Spirane complex formed by monosacharides and
borate. The stability constants for these compounds are tabulated in table 1.1.

the borate ring is transrt~t~rt~t~rt~t~rt~ to the R1 group preventing steric interactions. With xylose,


the 3'-OH and the R1 group are cis destabilizing the furanose ring.


*In 1,2 cis coordinated lyxose and arabinose, the R1 group is in cis position to the


borate ring, creating steric interference. The arabinose complex has the 3'-OH in a


transrt~t~rt~t~rt~t~rt~ position to both R1 and borate ring, in 1,2-coordinated lyxose the 3 groups are


cis decreasing the stability of the complex. However lyxose has the ability of










coordinating through hydroxyls 2,3 to borate, increasing the overall concentration

of the complex.


Stabilization of pentoses towards decomposition in the presence of borate

Berate complexation with carbohydrates should change the reactivity of the sugars

towards enolization. As mentioned before in the case of the pentoses, borate is expected

to lock the pentose in a closed form, rendering the completed sugar largely unreactive. A

literature review shows that indeed this is the case. Berate protects monosacharides

towards alkaline degradation and the extent of protection is proportional to the stability of

the complex. (Bruijn, Kieboom & Van Bekkum, 1986). However the stabilization of

aldopentoses had not been reported.

Mendecino (1960) reports the isomerization D-xylose at alkaline pH in the

presence of borate to yield D-xylulose (2-ketopentose). Once formed, the ketopentose

was found to be stabilized by borate towards degradation at high values of pH and

temperature.

In the present chapter we will explore the role of borate and its minerals in the

prebiotic synthesis and stabilization of the aldo-pentoses and in particular ribose.


Materials and Methods

Chemicals

All reagents used for synthesis or as standards were purchased from Sigma-Aldrich

Co., and were of their highest quality, if not mentioned otherwise. Glycolaldehyde was

obtained as the dimer from ICN Biomedicals. Sodium deuteroxide was purchased from

Cambridge Isotope Laboratories. Pyridine was purchased from Fluka in anhydrous










quality. Calcium deuteroxide was prepared in situ from metallic calcium and deuterium

oxide.

13C fully isotopically labeled D-ribose was purchased from Omicron Biochemicals

Inc. Silicon chips for desorption ionization on silicon (DIOS) analysis were obtained

from Silicon Sense Inc.

Enzymes

Aerobacter aerogenes Type I lyophilized cells as a crude source of ribitol

dehydrogenase were obtained from Sigma-Aldrich.

Analytic Instrumentation

Ultraviolet analysis (UV)

UV analysis was performed in a Cary Varian spectrophotometer interfaced to a MS

windows based computer.

Gas chromatography (GC)

Gas Chromatography Analysis was perform in a Perkin Elmer 1500 GC equipped

with a flame ionization detector and a DB-5 capillary column.

Mass spectrometry (MS)

Gas Chromatography- Mass spectrometry analysis was performed in a Finnigan

LCQ lon Trap GC-MS equipped with a DB-5 column. Inj ector temperature : 300 oC,

the temperature program used for the analysis was: 60 oC- 250 oC at 3.5 oC/min, 250 oC -

300 oC at 20 oC/min, final temperature was held for 20 minutes. Analysis were done in

collaboration with Dr. Lidia Nikole Matveeva, HPLC-ESI analysis were performed by

Dr. Jodie Johnson at the University of Florida.









Desorption Ionization On Silicon (DIOS-MS) analysis was performed on a Bruker

Daltonics (Billerica, MA) Reflex II MALDI-TOF mass spectrometer in the negative

reflectron mode in collaboration with Qian Li.

NMR Spectroscopy

Mercury 300 MHz. 1H NMR spectra, referenced to the respective solvent (D20).

Synthetic Preparations

Synthesis of colemanite

Calcium hydroxide (0.74 g ; 10 mmole) and boric acid (1.84 g; 30 mmole) were

added to 30 mL of Milli-Q water. The resulting suspension was stirred for an hour, and

then the pH was adjusted to a final value of 12 by dropwise addition of sodium hydroxide

(10 M) under stirring. Clear off-white crystals of colemanite (Ca2B6011.5H20)

precipitated from the slurry after one hour.

Synthesis of deuterated Colemanite

Calcium hydroxide (74 mg ; 1 mmole) and boric acid (184 mg; 3 mmole) were

added to a Fisher brand conical tube (10 mL, polypropylene) containing deuterium oxide

(10 mL). The resulting solution was immersed in liquid nitrogen and lyophilized. The

resulting white solid of deuterated colemanite was stored in a desiccator.

Synthesis of pentoses in the presence of colemanite

Glycolaldehyde dimer (6 mg, 0.05 mmol) and D,L-glyceraldehyde (15 mg, 0.16

mmol were dissolved together in an aqueous alkaline colemanite slurry (15 mL, pH: 12).

The mixture was heated at 45 oC for 60 minutes. The reaction was quenched by addition

of Dowex W-50 resin H' form until a pH of 5.0 was obtained. The resulting clear

solution was quickly filtered through a Nalgene 0.2 Cpm filter, into a Fisher brand

centrifuge tube (50 mL) and immediately immersed into liquid nitrogen until complete









solidification of the solution. Lyophilization yielded a white solid powder which was

dissolved in anhydrous methanol (5 mL) and subj ected to rotary evaporation under high

vacuum (repeated 3 times) yielding a white solid (18 mg, 88 % mass recovery of the total

carbon input).

Synthesis of pentoses in the presence of calcium hydroxide

Glycolaldehyde dimer (6 mg, 0.05 mmol) and D,L-glyceraldehyde (15 mg, 0.16

mmol were mixed and dissolved into an aqueous alkaline calcium hydroxide slurry (0.74

g, 15 mL, pH: 12). The mixture was heated at 45 oC for 60 minutes (the solution turned

clear brown after 20 minutes). The reaction was quenched by addition of Dowex W-50

resin H' form until a pH of 5.0 was obtained. The resulting clear brown solution was

quickly filtered through a Nalgene 0.2 Cpm filter, into a Fisher brand centrifuge tube (50

mL) and immediately immersed into liquid nitrogen until complete solidification of the

solution. Lyophilization yielded a dark brown syrup with an strong odor reminiscent of

caramel. The syrup was dissolved in anhydrous methanol (5 mL) and subj ected to rotary

evaporation under high vacuum (repeated 3 times) the syrup remained unchanged after

this treatment and it was not possible to obtain a solid even after high vacuum exposure

(19 mg, 91% mass recovery of total carbon input).

Derivatization of pentoses for gas chromatography analysis

Derivatization of pure samples of pentose was usually achieved by adding

anhydrous pyridine (400 CLL) to the corresponding solid pentose (5 mg, 33 nmol) ,

followed by addition of N,O-bis(trimethylsilyl)trifluoroacetamide (100 CLL) under Ar

atmosphere, the derivatized samples were used as standards for GC analysis.









Pentoses resulting from synthetic procedures, were derivatized by addition of

anhydrous pyridine (800 CLL) and of N,0-bis(trimethyl silyl)trifluoroacetamide (200 CL )

under Ar atmosphere.

Sugars Degradation Experiments

Sugar decomposition in the presence of calcium deuteroxide

Calcium deuteroxide (16.5 mg) and sodium benzoate (6.1 mg) were added to a 2.5

mL Eppendorf tube containing 1.5 mL of deuterium oxide. This solution was vortexed

and then equilibrated at room temperature for 30 min (pD: 12). After equilibration, the

tube was immersed into a 4 oC ice-water bath. An aqueous solution of the corresponding

pentose (D-arabinose, D-lyxose, D-ribose, D-xylose and D-ribulose; 500 CLL, 1.0 M)

was transferred by pipette to the ice-cold solution and vortexed for a period of 1 min. An

aliquot (900 CLL) of the resulting solution was then removed and used as the sample for

the 1H NMR degradation experiment.

Pentose degradation was measured by calculating the ratio of the integral for the

sharpest signal corresponding to the anomeric proton (hydrogen in C-1) over the value of

the internal standard integral (sodium benzoate). Half-life for the decomposition of each

pentose was calculated by plotting the mentioned ratio versus time.

Sugar decomposition in the presence of calcium deuteroxide and synthetic
colemanite

Calcium deuteroxide (16.5 mg), deuterated colemanite powder (83 mg), and

sodium benzoate (6.1 mg) were added to a 2.5 mL eppendorf tube containing 1.5 mL of

deuterium oxide. This solution was vortexed and then equilibrated at room temperature

for 30 min (pD adjusted to 12 by sodium deuteroxide addition when necessary). After

equilibration, the tube was immersed into a 4 oC ice-water bath. An aqueous solution of









the corresponding pentose (D-arabinose, D-lyxose, D-ribose, D-xylose and D-ribulose,

500 CLL, 1.0 M) was transferred by pipette to the ice-cold solution and vortexed for a

period of 1 min. An aliquot (900 CLL) of the resulting solution was then removed and used

as the sample for the 1H NMR degradation experiment.

Pentose degradation was measured by calculating the ratio of the integral for the

sharpest signal corresponding to the anomeric proton (hydrogen on C-1) over the value of

the internal standard integral (sodium benzoate). Half life for the decomposition of each

pentose was calculated by plotting the mentioned ratio versus time.

Enzymatic Assays

Ribitol dehydrogenase assay

An aliquot (1 mL) of the reaction mixture between glyceraldehyde +

glycolaldehyde (with/without boron) was removed after 60 min and treated with a sodium

borohydride solution (100 CLL, 0.06g/mL in cold water) followed by incubation at room

temperature for 30 min. This solution was then acidified to a final pH of 5.0 by addition

of aqueous HCI (200 pIL, 3 M) and passed through a short column containing Dowex-50

resin H+ form. An aliquot (650 pIL) of the eluent was lyophilized and the resulting solid

was dissolved in anhydrous methanol (650 CLL) and subj ected to rotary evaporation under

high vacuum (repeated 2 times).

The white solid obtained after methanol evaporation was dissolved in potassium

phosphate buffer (200 CLL, 1.0 M, pH: 10) and then aqueous sodium borate ( 10 CLL, 19

mM) was added. The enzymatic reaction was initiated by addition of nicotinamide

adenine dinucleotide (NAD 15 CLL, 100 mM in water), and of Aerobacter aerogenes

Type I lyophilized cells suspension (50 CLL 0.02 g/mL in 1 M potassium phosphate

buffer) and incubated for 60 min at 30 oC. After incubation, the reaction was placed on an









ice bath for 5 minutes, and then centrifugated to remove cell debris. The presence of 2-

pentulose in the resulting solution was confirmed by the cysteine-carbazole test.

Cysteine-carbazole test

An aliquot (100 CLL) of solution from the ribitol dehydrogenase assay, was

transferred to a quartz cuvette containing a cysteine solution (20 CLL, 1.5% in water).

sulfuric acid (600 CLL, 15.2 M) and carbazole (20 CLL, 0.12% in ethanol) were then added

and mixed thoroughly. The reaction was kept at room temperature for a period of 60 min.

Presence of ribulose in the mixture was confirmed by the appearance of a strong violet

color (UV/vis : 3Lax = 535 nm).

DIOS Analysis

Preparation of PSi surfaces

PSi surfaces were manufactured with HOME (H202-Metal) etching method: a

silicon chip of 2x2 cm was cut from an n-doped crystalline silicon wafer and sputter

coated with a thin layer of Au at an argon pressure of 40 Torr and a current of 10 mA.

For the production of an array PSi chip, an aluminum mask with 4x4 holes (0.8 mm in

diameter) was used in Au coating. The Au coated silicon chip was then immersed in an

etching solution (49% HF: 30% H202: Ethanol = 1: 1: 1, v: v: v) for 10-20 s (Figure 2.9).

Samples of sugars (0.5 CLL) were directly pipetted from the mother solution into the

PSi surfaces.




~1a b c6--J7,~ee

Figure 2.9. Psi surface preparation: (a) An aluminum mask was placed on top of a
crystalline silicon chip, (b) Only exposed areas were coated with Au, (c) PSi
spots were produced on Au-coated regions










Competition Experiments

1, 4-Anhydroerythritol (AET) vs Pentoses

1,4-Anhydroerythritol (100 CLL, 0.1 M, in water) and the corresponding pentose

(100 CLL, 0.1 M) in water were mixed in a Eppendorf tube (1.5 mL) containing water

(700 CLL) and vortexed thoroughly. An aqueous sodium borate solution (100 CIL, 0.025

M) was then added, and the resulting mixture was vortexed and equilibrated for 2 h at

room temperature before DIOS analysis.


13C-Ribose vs Pentoses

13C-Ribose (10 CLL, 0.1 M, in water) and the corresponding pentose (10 CIL, 0.1 M

in water) were mixed into a Eppendorf tube (500 CLL) containing water (70 CIL) and

vortexed thoroughly. An aqueous sodium borate solution (70 CLL, 0.025 M) was then

added and the resulting mixture was vortexed and equilibrated for 2 h at room

temperature before DIOS analysis.



Results

Synthetic Preparations

Synthesis of pentoses in the presence of colemanite

The presence of pentoses in the reaction products after 60 minutes of incubation

under formose conditions was directly confirmed by HPLC/ESI (-)-MS analysis. The

dipentose-borate complex was detected as an m/z: 307 molecular ion (Figure 2. 10).

Analysis of a standard of di-ribose-borate complex is shown for comparison in Figure

2.11.














16e 20 uL


















247 1

277 5
2451


EUSpecW-Dauleq-200144 0891144~B307343P YI4054,whee~na
H To-RP.015,asorrareDG 51 o~s
SEQI-203144#B349 RT.279-297 AIV.4 S 1 262-267 NL 361EA
F 6lE5Asid=200Flillrn [145001001000
100- 239 1



60 n 209 3 73 3 4
408




SEO203144sB547 RT 276240I AY3 SB 262-257 NL 54rE4
F c aEbl idi2 00 Funmn2 307 00@3 0010 0010 3500
21711


40 i


20_


Y*0111~1
~i~ll


Figure 2. 10. HPLC-MS analysis of reaction mixture containing colemanite. Analysis of

the products from the reaction between glyceraldehyde an glycolaldehyde in

the presence of synthetic colemanite (Ca2B6011.5H20) at pH: 12, 45 oC. (top)
ESI-MS indicates the presence of the pentose-borate dimeric complex ion

MW: 307. (middle) MS/MS fragmentation pattern of the 307 ion/peak.

Fragmentation is consistent with that observed from the standard. (bottom)
HPLC-MS trace of reaction mixture showing only various isomeric pentoses.


E W-spmorsetWDt 4202s 41
HgredPrr 2,100140-6)~ > s{60]-pEsI
SEQI-202941s47 RT2?.18 AV- s 8,150(17-19 NL.GAS
F: CE1al0g2200 Fullms [12500-100000]


08~444 02 55 12 PM AR-(14$-, Ribose borat:20 uL


"-on


8 so- MnW : 307
6 '






mb


Figure 2. 11. Detection of ribose-borate comples by ESI (-) ion mode.(electrospray
ionization negative ion mode).


Gas chromatography analysis of the trimethylsilyl derivatives of the reaction



products is shown in Figure 2. 12. Peaks were assigned by coinj section of authentic


standards, using retention time values (Table 2.2).










chromatogram Plot C:\GCPODATA\2169El Date: LO1078 /03 16 :04 : 26
Comment: El AR5--4 60/O-250/3. 5-300/20:,250, 300 ; !uL
Scan No: 1827 Retention Time: 35 :30 RIC: 203653 Mass Range: 50 746
Plotted: 1 to 3653 Range: 1 to 3653 100% = 12484930
100 % 2











TOT4












600 1200 1800 2400 3000 3600
15:03 25:03 35:03 45:04 55:04

Figure 2. 12. GC trace of the reaction mixture containing colemanite. Analysis of the
silylated products (after 60 min) from the reaction between glyceraldehyde
and glycolaldehyde in the presence of synthetic colemanite(Ca2B6011.5H20)
at pH: 12, 45 oC. Peak assignation: (1) Lyxose/Arabinose, (2)
Lyxose/Arabinose, (3) Ribose, (4)Xylose, (5) Xylose. Anomeric forms of the
sugars are not assigned. Notice the absence of peaks corresponding to
compounds eluting after 5.

Table 2.2. Retention times of trimethylsilyl derivative of pentoses by GC analysis.
Pentose Retention Time (min) Intensity ratios

Arabinose 28.34; 29.49 10:1

Lyxose 28.17; 29.81 1:1

Ribose 30.04 1

Xylose 28.79; 31.97; 34.22 1:2:5


Synthesis of pentoses in the presence of calcium hydroxide

A large portion of insoluble material for this reaction remained insoluble in


pyridine after trimethylsilylation at 60 min. Gas chromatography analysis showed an










heterogeneous composition with a low signal intensity (data not shown). Analysis of the

reaction products after just 20 minutes is shown in Figure 2. 13. Peaks were assigned by

coinj section of authentic standards and retention time values.

Chromratogram Plot C:\GCQ\DATAl2162El D~ate: 10/09 03 OB :45 :30
Comment: El AR5027-b20 60/0-250/3.5-300/20;r250. 300 ; !uL
Scan No: 1930 Retention Time: 35 :33 RIC: 612445 Mass Range: 50 745
Plotted: 1 to 3660 Range: 1 to 3660 100% s 4784370
100 %


600
15:03


1
~-J

?---~ i


3000
55:04


1200
25:03


1800
35:03


2400
415:04


Figure 2. 13. GC trace of the reaction mixture containing Ca(OH)2. Analysis of the
silylated products (after 20 min) from the reaction between glyceraldehyde
and glycolaldehyde in the presence of Ca(OH)2 at pH: 12, 45 oC. Peak
assignation: (1) Lyxose/Arabinose, (2) Lyxose/Arabinose, (3) Ribose, (4)
Xylose, (5) Xylose + other. Anomeric forms of the sugars are not assigned.
Compounds to the right, eluting after peak, 5 include tar.

Sugar Degradation Experiments

Sugar decomposition in the presence of Calcium Deuteroxide

Alkaline degradation of sugars in the presence of calcium deuteroxide occurred at a

high rate. After a few minutes of reaction, the sugar solutions turned clear brown, and

after 1 h some precipitate formed in the NMR tube. The calculated rates of


I;"I :ii.


3600









decomposition (Table 2.3) were obtained by measuring the value of the integral of the

signal associated with the anomeric proton and comparing this value with that of the

internal standard (Figure 2.14).


300 MHz Proton NMR
Solvent : Deuterium Oxiide (D20)
Internal Standar: Sodiurn Benzoate j
Ribose + Calciurn Hidroxide















Fiur 214 Icuatonofrio e i aOH)ouin Dcmoitio wa monitoe by
H ~~~I NM t2 C oelsso inl t6(pm:48,49,54








of the suar were sbtailize towad decomosi nC(H2sltion. (wencomparedtion the smpltoes





without colemanite). Sugar solutions remained clear after several hours and in the case of

ribose it remained unchanged for several days (Figure 2.15). The calculated rates of

decomposition are shown in Table 2.3.











300 MHz Proton NMR
Solvent : Deuterium Oxide
Internal Standar : Soaium Benzoate
Rlbose + Calcium H-idroxide + Colemanite


















unhage eve under inuaioo 5 days j, at -r high pH

Tablre 2.31. H ualf ife of pentoses undte r alalne odtonsa(H) determined e by N Rad



estimated by loss of selected NMR signals (see appendix 1).
Pentose Half life (min): Half life (min):
Ca(OD) Ca(OD) + COlemanite
D-ribose 291 2700
D-arabinose 144 259; 716; 331
L-arabinose 124 380; 274
D-lyxose 69; 177 1382
L-xylose 66 572; 258
L-Lyxose 184 1201
ribulose 14 2028


Enzymatic Assays

The presence of ribulose after ribitol dehydrogenase treatment was confirmed by

the cysteine-carbazole test. A dark violet color was observed when samples of the

reaction between glyceraldehyde and glycolaldehyde were analyzed by this assay.









DIOS analysis: Competition Experiments.

1, 4-Anhydroerythritol (AET) vs Pentoses

The different complexes detected by DIOS are depicted in Figure 2.16. A typical

spectra obtained by DIOS analysis is shown in Figure 2. 17.






00 o ooq





m/z :215 m/z :261 m/z :307
Figure 2.16. Anhydroethrythritol (AET)-pentose borate ions detected by DIOS. Pentose
borate complex is depicted through C2 and C3 (cis diol). However, binding
through C1-C2 (cis diol) is also known to occur when possible.

Each pentose was analyzed independently, in competition with 1,4-

anhydroerythritol (AET). The relatives intensities reported for each detected ion (Figure

2. 18) were calculated using equation (1) :

% Relative intensity = 100 x Intensity of individual ion : equation (1)

SIntensity of all detected ions


Error bars represent the standard deviation calculated from a set of 10 different

measurements (for complete set of spectra see appendix 2).

13C-Ribose vs Pentoses

Fully isotopically labeled ribose (13C-ribose, 98.8% labeled) was used for

competitions experiments because the difference in mass allows resolution of peaks with

non-isotopically labeled pentoses. The different compounds detected in this experiment



















3000



2000

a.1

1000


1IIIII


are shown in Figure 2. 19. A typical DIOS spectra of this experiment is shown in Figure

2.20.


0 50 100 150 200 250 300 350
lexport /home/data/Li~ian/1 0_17_03/12_2Lin/pdata/1 unknown


400 450 mn/2
Fri Oct 17 12:35:17 2003


Figure 2. 17. DIOS spectra of competition experiment D-arabinose vs AET. Boron (10
mM) was set as the limiting reagent to favor spirane formation.



Pentose vs 1,4 -Anhydroerythritol


70.0%

60.0%

~50.0% -1 -I O xylose

~40.0% m1 T arabinose
I ~ ~ ~ I t lyxose
30.0%
o ribose
20.0% -1 aI I riblulose

S10.0%

0.0%

m/z 215 m/z 261 mlz 307


Figure 2. 18. Competition experiments between the different pentoses and AET.
Relativive intensities represent the molar fraction of the different borate
complexes in solution.


-LJ-ILI~~L~w..JIl...i








38


HO-1'3C
HOHO-13C"
OH 13CHOHO
'3C~nn130 -136







10130


OH O OH 3C--OH

m/z : 307 m/z : 312 m/z : 317

Figure 2.19. 13 1'2C-D-ribose borate ions detected by DIOS. Pentose borate complexes
detected in the competition experiment using isotopically labeled ribose.







3000












2 9030 1 3033 /

lexpore/hame/data/Ligian/02 ~ ~ ~ ~ ~ ~ ~ I CNG47efpaa1ukow h e 2 93:520
Fiue220. DIO setaocmpiinexrimn13rioeV12rbs.Brn(0
mM) was se stelmtn raett oc h frainoprn
compounds.
Eahpnoewsaalzdidpnetyincmeiinfrboo idn gis


13C-iboe. Fgur 2.1 shws he rtioof 3C-rbos/penosein quilbrim deectd b

DIOS. Erro bar rersn th stnaddvaincluaedfo e f1ifrn


measurements.











1C-d-ribose vs pentose


8.00

S7.00-

S6.00-

a 5.00-

o 4.00-

S3.00-

S2.00-




O ribose a lyxose arabinose o xylose a xylulose a ribulose


Figure 2.21. Ratio of borate complexes of 13C-ribose vs pentoses.

12C-D-ribose was included in the analysis to test the isotope effect in ionization-

vaporization efficiency, however, no such effect was observed during the experiment.

The slight deviation from the value of 1.0 in D-ribose is due to the presence of 1.1%

unlabeled carbon in the 13C-D-ribose used for the competition experiment.

Discussion

Reaction intermediates of the formose reaction were used as starting materials for

the synthesis of pentoses under alkaline conditions. When the reaction was made in the

presence of calcium hydroxide, a simple visual inspection gave some clue of the

composition. The solution which was originally transparent, turned clear brown after 20

minutes and after one hour dark brown, which, in the case of sugars, is an indicator of

decomposition with branching (know as caramelization or browning). Gas

chromatography analysis showed that pentoses were indeed present at the early stages of

the reaction in addition to other uncharacterized carbonaceous material as seen in Figure









2.13 (probably tetroses, hexoses and branched sugars). However, further incubation of

the alkaline mixture, showed a decrease in the total amount of detectable carbon caused

by degradation of pentoses to higher molecular weight material that it is not volatile and

therefore undetectable by the GC analysis. These results agree with the expected

reactivity of the formose mixture.

When boron was introduced as the mineral colemanite into the reaction mixture,

visual inspection indicated that the decomposition process was either absent or slow.

Browning was not observed even under long incubation periods. GC analysis of the

reaction after a period of one hour shows the presence of pentoses as the maj or products,

which indicates that boron is playing an active role in the control of the regiochemistry,

and also stabilizing the pentoses product once these are formed. Because an equimolar

amount of the fourth aldopentoses was detected, indicates that there was not

stereoselectivity in the condensation reaction in the presence of borate.

A proposed mechanism that will account for these observations is depicted in

Figure 2.22.

The mechanism depicted above, explains the regiochemistry obtained from the

reaction between glyceraldehyde and glycolaldehyde. But it does not address the

stabilization of the pentoses by boron, once formed, towards alkaline degradation.

Degradation experiments in the presence of colemanite, showed that all the aldo-

pentoses are stabilized by boron, following the trend: ribose > lyxose > arabinose >

xylose. Shifts in the NMR signal corresponding to the anomeric proton of the

monosaccharides are consistent with previous literature reports indicating that boron

coordinates to pentoses, by favoring the formation of a closed furanose conformation.










Glyceraldehyde

electrophilic 11il
center elctrophilic
nucleophilicNolne
acidic cete nulohlc
hydrogen 'C H'C H

H-C-OH H-- -CHH
H--C--OHBorate H-C~-O/ \O-C--H Ca+


H ~~c dli due toH CH
anionic borate


H, O H, O H ,O H, O

H--~OH HO--H H--OH HO--H

H-- OH H--~OH HO--H HO--H
H-- OH H--OH H--OH H--OH
H-- OH H--OH H--OH H--OH

HH H H
D-ribose D-arabinose D-xylose D-lyxose
Figure 2.22. Suggested mechanism for pentose formation. The aldol condensation
reaction between glyceraldehyde, and glycolaldehyde in the presence of
borate is shown. (Pentose products in the Fischer representation are shown
only in the D-form for simplicity).

Because the rate of degradation of these sugars is proportional to the amount of

pentose in the open form, the half life should be directly proportional to the amount of

pentose-borate ester in each case. Boron coordinates pentoses in different fashions to

make spirane complexes; this is of course an obvious consequence of the spatial

orientation of the different cis-hydroxyl groups.

DIOS experiments were focused in understanding these binding preferences. A

qualitative description of the preferential binding of borate for the different pentoses

shows that the preferential order for binding is: ribose > lyxose > arabinose > xylose.

This information confirms that the stabilization of pentoses (increased half life)


Glycolaldehyde

electrophilic


O H

C a(OH)2 H









under alkaline conditions is a direct consequence of borate-pentose complex formation.

This synthesis of pentoses in the presence of the mineral colemanite is therefore

plausibly prebiotic. Indeed, in the presence ofborate, and given that ribose is the first

compound in the formose product progression that offers a non-aldehydic cyclic form

with unhindered cis-diols, the formation of ribose appears to be the natural consequence

of the intrinsic chemical reactivity of compounds available from the interstellar medium

under alkaline, calciferous conditions. As these conditions are not excluded from the

early Earth, it is also not possible to exclude the availability of pentoses at the time when

life originated.

This example of how minerals can productively control organic reactivity reminds

us of the fact that prebiotic chemistry is occurring on a planet, in the context of a larger

geology. Minerals must be considered as we constrain models for the origin of life.















CHAPTER 3
CATALYSIS AND THE RNA WORLD

Introduction

Ten years ago, Szostak, Joyce, Ellington, and others applied in vitro selection (IVS)

to libraries of nucleic acids to extract nucleic acids that catalyze simple reactions, such as

RNA ligation (Bartel & Szostak, 1993) and RNA cleavage (Breaker & Joyce, 1994). This

work opened the possibility of using in vitro selection to ask quantitative questions about

the performance of these catalysts. This includes questions concerning the mechanism of

specific nucleic acid enzymes, as well as broader questions, such as how functional

behavior is distributed in nucleic acid sequence "space", and whether adding chemical

functionality to nucleic acids, either by modifying the nucleobases or by adding

cofactors, can enhance the catalytic potential of a nucleic acid library (Breaker, 2000;

Perrin et al, 2001).

Before broader questions can be addressed using in vitro selection, it is necessary

to explore some of the specific features displayed by many nucleic acid catalysts that

have emerged from selection experiments. For example:

(a) Partial conversion. Many studies of individual nucleic acid catalysts report that

their reaction goes only partially to completion. Hammerhead ribozymes, for example,

are frequently reported to cleave only 40 60% at plateau (Stage-Zimmermann &

Uhlenbeck, 1998) (Kore et al, 2000). DNAzymes with ribonuclease activity similar to the

ones studied here are also frequently reported with cleavage plateaus of 25 65% (Perrin









et al, 2001; Geyer & Sen, 1997; Faulhammer & Famulok, 1996; ibid, 1997). The same is

seen for DNAzymes with DNase activity (Carmi et al, 1996).

(b) Intramolecular versus intermolecular reactions. Many reactions catalyzed by

nucleic acid catalysts are selected to be intramolecular, making the term "catalyst"

technically incorrect; the catalyst is not regenerated. Many of the intramolecular reactions

have analogous intermolecular processes that are truly catalytic, however, and these are

often accessible both during the selection and in the subsequent kinetic analysis.

(c) Loss of the most active catalysts during the set-up. In many selection schemes, a

library of catalysts must be synthesized and folded before the selection step begins. This

leads to the possibility that active catalysts in a pool will be lost before the selection

system can extract them.

(d) Michaelis-Menten kinetic behavior. Many nucleic acid catalysts bind their

substrate in a reversible step prior to the step where chemical bonds are made or broken.

This should generate saturation kinetics similar to those seen in protein enzymes.

It is, of course, impossible to address these questions for the general reaction

catalyzed by all nucleic acid catalysts. Rather, these questions must be addressed for

individual nucleic acid catalysts working on specific reactions. Such studies follow the

tradition in physical organic chemistry, where many detailed studies of specific reactions

eventually generate a body of literature that addresses broader issues in catalysis.

For this work, we began with work of Breaker and Joyce, who selected for DNA

enzymes that catalyze the cleavage of a ribonucleotidyl-3'->5'-deoxyribonucleotid

linkage in an oligodeoxyribonucleotide (Breaker & Joyce, 1995). The reaction almost

certainly proceeds with the attack on the phosphorus electrophilic center via the










ribonucleotide 2'-hydroxyl group (Figure 3.1). An adaptation of the Breaker-Joyce

selection procedure led to several new catalysts for this reaction (Figure 3.2). One of

these DNAzymes, designated 614, was studied in detail to address some of the issues

outlined above.




NH2 [3'-constant region NH2`
[3'-constant region] N N
B0n- i 5'-constant region ]-0~ Blotn-[5cosatrgn]-
O O-H O ,O
Streptavidin O=,P O- Streptavidin d 'O-
support (~support

HO






Figure 3.1. In vitro selection experiment representation. A library of DNA
oligonucleotides containing an internal adenine riboside and a 40-nt random
region (represented in red) is attached to a solid support. The DNA-catalyzed
ribonuclease reaction proceeds with the attack on the phosphorus electrophilic
center by the ribonucleotide 2'-hydroxyl group, cleaving the phosphodiester
backbone and releasing the catalytic portion of the DNAzyme from the solid
support.

Materials and Methods


Preparation of Precursor DNAzymes via PCR (Maniatis et al, 1982)

DNAzymes were prepared by PCR amplification of the template (synthesized by

Integrated DNA Technologies (Coralville IA), or from a clone) using a catalytic strand

primer (cat ribose or cat deoxyribose, see Table 3.1) and a complementary strand

primer (compl, compl+ 5'P, or compl tail). For transrt~t~rt~t~rt~t~rt~ cleaving assays, DNAzymes were

generated without the internal ribose using the cat deoxyribose primer, identical to the











A

5' -Biotin-CTGCAGAATTCTAATACGC-ITCACTAT GAAAATC
SI Ij I G
3' -CACGGTTCGAATGGCAGTG- -(N40) --CTCT-CA


B


614 re$plncaaa BI~TPAGG'TTGC;TAG 5.015
r#10.3
r#11.131
r#11.27 (

r1.5r#12.14Br1.541 1I
r#12.159

r#15.161 0.006
r#15.58 0.128
r#15.162 0.161
r#15.53 0.163
62 TATT AG 0.049
615
r#11.137
r#12.147
r#15.169 0.017
r#15. (9)


r#12. (2)
r#12.146 I
625 TGTGCTAGGTGTTCTCTGAGCCAGACGTTAGTGTAGTTAAG 0.045

Figure 3.2. Sequence of the initial library and DNAzymes isolated from the in vitro
selection. (A) The initial library was based on the sequence used by Breaker
and Joyce, with an internal adenine riboside incorporated at position 28 to
provide a cleavable linker. Two nucleotide substitutions were introduced to
eliminate one of the clamps that were designed to hold the substrate and
catalyst portions of the molecules in the library together. The original two
nucleotides in the Breaker Joyce sequence are shown in bold above and below
the sequence used in this study. Base pairing that could form binding clamps
are highlighted in grey. (B) DNAzymes isolated from multiple rounds of IVS.
Sequences 614, 62/615, 616, and 625 represent the maj or sequences classes
initially cloned from the seventh round of the initial IVS. Sequences variants
of these maj or classes were isolated from additional rounds of selection and
are grouped according to sequence class (only variations from the prototype
sequence are shown). Colored boxes are shown to highlight common motifs.
Sequences isolated more than once are shown with the number of isolates
identified in parenthesis following the name.









cat ribose primer except that the single ribo-adenosine in the cat ribose was replaced

with a 2' -deoxyribose-adenosine. To obtain single stranded catalysts, DNAzymes were

produced using a catalytic strand primer and either a 5'-phosphorylated complementary

strand primer (compl+ 5'P, for use with lambda exonuclease) or a complementary strand

primer with an 15-nt poly-deoxyriboadenosine tail appended to the 5' via an 18-atom

hexaethyleneglycol-based linker (compl tail, for use in asymmetric PCR). This linker

prevents polymerase read-through. Templates all had the same 5'-constant region and 3'-

constant region to which complementary and catalytic strand primers bind, separated by

40 nucleotides. Ribose-614, deoxyribose-614, and the library templates were

synthesized and PAGE purified by IDT DNA technologies.

Typical conditions for a PCR contained up to 1 ng template, 100 nM catalytic

strand primer, 100 nM complementary strand primer, 100 CIM dNTPS, 10 mM KC1, 20

mM Tris-HCI (pH 8.8), 10 mM (NH4)2SO4, 2 mM MgSO4, 0.1% Triton X-100, 3-4 units

polymerase (Taq or Vent exo-), and alpha-32P-CTP (10 CICi, for internally labeled

samples), for a total volume of 100 CIL. The PCR amplification cycle consisted of an

initial incubation (3 min, 960C) followed by 20 PCR cycles of (45 sec at 960C, 45 sec at

500C, and 2 min at 720C).

PCR for in vitro selections (IVS) used Vent exo- polymerase (3-4 units) and excess

catalytic strand primer (400 nM), with up to 40 cycles of PCR in the early rounds.

Preparation of Single-stranded DNAzymes

Double stranded DNAzymes generated via PCR with a 5'-phosphorylated

complementary strand primer (compl+ 5'P) were converted to single-stranded DNA by

digestion of the complementary strand using lambda exonuclease (which is specific for









the 5'-phosphorylated strand of double stranded DNA). The DNA was recovered by

EtOH precipitation, resuspended in exonuclease solution (25 CIL, 5 units lambda

exonuclease, 67 mM glycine-KOH (pH 9.4), 2.5 mM MgCl2, 50 Clg/mL BSA, for a 100

CIL PCR). Samples were mixed and incubated (370C 30 min). Reactions were

terminated with formamide stop dye (1 mg/mL xylene cyanol, 1 mg/mL bromophenol

blue, 10 mM EDTA, in 98% formamide) and heating (800C 10 min). The single-stranded

products were resolved by 8% PAGE/urea, and full-length ssDNA products excised. Gel

slices of individual samples were crushed with individual disposable mortars and were

eluted in buffer (350 CIL, 500 mM NH40Ac, 0.1 mM EDTA, 0.1% SDS, pH 7) overnight.

Gel-purified samples were extracted with phenol/CHCl3/isoamyl alcohol (25:24:1), then

with CHCl3/isoamyl alcohol (24:1), and the resulting DNA was precipitated in NH40Ac

and EtOH.

Single stranded DNAzymes were also purified via asymmetric PCR using a

complementary strand primer with a 15-nt poly-deoxyriboadenosine tail connected to the

5'-end of the primer by a C18 linker (compl+tail), at which Vent and Taq polymerases

terminate. Complementary strand molecules generated by extension of a complementary

strand primer containing the 5'-tail are longer than the full-length catalytic strand, and

were separated from the catalytic strand by PAGE/urea (8% acrylamide). Gel-purified

samples were excised, extracted and recovered as before.

5'-End Labeling of DNA

Single-stranded DNA (20 mM) was 5'-labeled with gamma-32P-ATP (20 CICi)

using T4 polynucleotide kinase (10 units) in Tris-HCI buffer (70 mM, pH 7.6, 10 mM

MgCl2, 5 mM dithiothreitol, final volume 10 CIL, 30 min 370C). An equal volume TE









was then added, and the mixture heated (20 min 700C). End-labeled DNA was separated

from unincorporated nucleotides by spinning through a G-25 column (600 g, 3 min).

DNAzyme Kinetic Assays

DNAzymes and substrates were isolated via EtOH precipitation and re-suspended

in HEPES buffer (50 mM, pH 7). Their concentration was estimated by Cherenkov

counting. Samples were diluted to twice the desired Einal concentration with additional

HEPES buffer. For transrt~t~rt~t~rt~t~rt~ assays, enzyme and substrates were typically mixed (unless

otherwise noted) and diluted to twice the desired Einal concentration. Samples were then

mixed with equal volume 2X reaction buffer (typically 2M NaC1, 2 mM MgCl2, 50 mM

HEPES pH 7). Mixtures were then heated (960C, 3 min), and slowly cooled to 230C

(over 10 min) collectively termed "slow cooling". The initial "time zero" point was

the time at which the sample completed the slow cooling. Unless otherwise noted,

reactions were run at 250C and terminated at various times by diluting an aliquot of the

reaction into the formamide stop dye, followed by freezing (-200C). Samples were

resolved using 8% PAGE/urea and the product cleaved quantified using a Bio-Rad

phosphorimager. Data was analyzed using the GraphPad Prism 3.0a software package.

Cloning and Sequencing DNAzymes

Single-stranded DNAzymes were converted to duplex DNA by PCR amplification,

usually with only the complementary strand primer compl (960C for 3 min; followed by 3

cycles of 45 sec at 960C, four min ramp cool to 500C plus 45 sec at 500C, and 1 min at

720C; followed by 7 min at 720C). When cloning cleaved and uncleaved 614, the

desired single-stranded DNA (either cleaved product or uncleaved reactant) was first

isolated via PAGE/urea. PCR of the unpurified 614 reaction mixture containing both

cleaved and uncleaved 614 with only compl primer yielded clones only of uncleaved 614









molecules; cleaved 614 molecules were therefore cloned by PCR amplifying gel purified

cleaved fragments using the compl primer with the 5'-truncated catalytic primer cat.nt29-

45. Fresh double stranded PCR products were cloned using the TOPO TA Cloning

System (InVitrogen) and plated on agarose plates containing ampicillin. Transformed

cells were given only 15-30 min to recover in antibiotic-free media prior to plating on

antibiotic containing plates (to prevent recently transformed clones from doubling prior

to plating, favoring isolation of only unique species from the original pool).

Clones were transformed into the TOPO TA Cloning Vector and DNA was

prepared from individual clones using the alkaline lysis protocol.

Clones were sequenced using the 1224 plasmid sequencing primer (3.2 pM, 5'-

CGC CAGGGTTTTCC CAGTCAC GAC) with 3 00-5 00 ng pl asmi d DNA template, 2X

final concentration Big Dye reaction buffer and 2 CIL Big Dye Terminator Sequencing

mix in a final volume of 10 CIL. Samples were overlaid with mineral oil and amplified by

PCR (25 cycles, 960C for 30 sec, 500C for 15 sec, and 600C for 4 min). Sequencing

samples were analyzed on an Applied Biosystems Prism 310 Genetic Analyzer.

Sequencing results were confirmed by examining the chromatograms manually using the

"Sequencher" software package.

Inz vitro Selection

DNAzyme libraries for the first round of selection were prepared by a single cycle

of run-off P CR u sing the lib rary template (5'- GTGC CAAGC TTAC CGTC AC -N40 -

GAGATGTCGCCATCTCTTCC (where N indicates equal molar concentrations of A, T,

G, and C) and cat+ribose primer. Run-off reactions volumes ranged from 3 to 20 mL,

each containing 1 ng/CIL library template, 100 nM catalytic strand primer, 100 CIM









dNTPs, 10 mM KC1, 20 mM Tris-HCI (pH 8.8), 10 mM (NH4) 2SO4, 2 mM Mg2SO4,

0.1% Triton X-100, 20 units/mL Vent exo-, and 4.3 CICi/mL alpha-32P-CTP. An aliquot

(1 mL) of each sample mixture (minus polymerase) was placed in Eppendorf tubes (1.5

mL), heated (960C, 8 min, then slowly cooled to 550C over 30 min). Polymerase was

added, and the samples were incubated (720C, 15 min).

DNA was recovered by EtOH precipitation with NH40Ac and glycogen as a carrier

by storing overnight at -800C and then centrifuging in Corex tubes (16,000 g, 40 min,

40C). The EtOH was removed and the pellet re-suspended in water (200 CL). DNA was

recovered by precipitation a second time with EtOH (NH40Ac) and centrifugation

(16,000 g, 20 min, 40C). After removing the EtOH, the pellet was resuspended in lX

binding buffer (1 M NaC1, 1 mM EDTA, 50 mM HEPES pH 7) and bound to a

streptavidin column. The unbound material was removed by flushing the column with

wash buffer (50 mM HEPES pH 7). The complementary strand was removed by washing

quickly with 0.2M NaOH, followed again by wash buffer.

The cleavage reaction was initiated by eluting the wash buffer and replacing it with

reaction buffer (lM NaC1, 1 mM MgCl2, 50 mM HEPES pH 7). The cleaved products

were eluted from the column after two hours incubation. The eluted material was used in

a PCR with the cat+ribose and compl+5 'P primers to generate material for the next

round of selection.










Table 3.1. Name, sequence,
chapter.
Names used in text


ribose-614


and description of oligonucleotides used throughout this

Description


5'-
CTGCAGAATTCTAATACGACTCACTATrAGGAAGA
CATGGCGACTCTCACATCATGCGAGCACACGCAAT
AGCCTGATAAGGTTGGTAGTGACGGTAAGCTTGGC
AC

A DNAzyme isolated from the in vitro selection in this
study. It catalyzes the cleavage of the ribo-adenosine
embedded within the cat+ribose primer, either as a part of
its own sequence, or in trans.

A mutant of ribose-614 in which the cytosine at position 72
is replaced by a thymidine, resulting in a 30-fold lower rate
of cleavage at 100 nM.

5'-
GTGC CAAGC TTACC GTCAC(n40)GAGATGTCGCCAT
CTCTTCC

The complementary strand template containing a region of
40 nucleotide randomized region flanked on both sides with
constant regions, one complementary to cat+ribose, and the
other identical to compl.

A random library generated by PCR amplification of the
library template with cat+ribose and compl primers.
Without selection, this library is predominately inactive.
Selection of this library gave rise to DNAzyme 614, and
others. All sequences contain the cat+ribose primer and
therefore have the potential to function as substrates for 614
cleavage.

An individual clone isolated from a the ribose-library.
This sequence has no intrinsic ribonuclease activity of its
own, but contains the cat+ribose primer and therefore can
function as a substrate for 614 cleavage.

The same sequences as ribose-614, ribose-lib62, and
ribose-614AC72T but with the ribo-adenosine at position
28 replaced by a de oxyribose-adeno si ne (via PCR
amplification with cat+deoxyribose primer). deoxyribose-
614 is still capable of cleaving the ribo-adenosine embedded
within the cat+ribose primer, but only in trans.t~t~rt~t~rt~t~rt~


ribose-614AC72T



library template


ribose-library








ribose-lib62




deoxyribose-614,
deoxyribose-lib62,
deoxyribose-
614AC72T










Table 3.1 Continued

Names used in text


Description


conpl-614

cat+ribose


The complementary strand to deoxyribose-614.


C TGC AGAATT CTAATAC GAC TCAC TATrAGGAAGA
CATGGCGACTCTC

Primer used to generate full length molecules from
template, thus incorporating a ribo-adenosine (rA) at
position 28. This oligo has no catalytic activity of its own
and was therefore used as a substrate in trans cleavage
assays.

The same oligonucleotide sequence as cat+ribose but with
a deoxyribonucleotide replacing the rA. This primer was
used to generate full length DNAzymes without the ribo-
adenosine moiety. DNAzymes incorporating
cat+deoxyribose can not be cleaved and therefore act as a
true enzyme.


cat+deoxyribose


conpl


5'-GTGCCAAGCTTACCGTCA


This primer is complementary to the 3'-end of the catalytic
strand of the full length DNAzymes used in this study, and
is used to generate double stranded DNAzymes via PCR.

This primer is the same sequence as the conpl primer but
with a phosphate attached to the 5'-position. This primer is
used to generate double stranded DNAzyme via PCR,
followed by degradation of the complementary strand by
lambda exonuclease (an enzyme that specifically degrades
5'-phosphorylated DNA).

This primer is the same sequence as the conpl primer but
with a 15-nt poly-adenosine attached to the 5'-position via
an 18-atom hexaethyleneglycol-based linker. Polymerase
can not efficiently read through this linker, so PCR using
this primer genearates a double stranded DNAzymes with a
complementary strand that is 15-nt longer than the catalytic
strand, thus allowing purification of the catalytic strand via
PAGE/urea.

The cat+deoxyribose oligo was used as the chase substrate
for cleavage assays with 614 functioning as the enzyme.


compl+5 'P







conpl+tail









chase









Table 3.1 continued

Names used in text Description


28-nt product The 28-nt product generated from cleavage of ribose-614
(106-nt).

78-nt product The 78-nt product generated from cleavage of ribose-614
(106-nt).

cat. nt29-45 5 '-GGAAGACAT GGC GAC T CTC

A 5'-truncated version of cat deoxyribose (containing
nucleotides 29 45) used to generate dsDNA of the 78-nt
ribose-614 cleavage product for subsequent cloning.



Results

In vitro Selection

An in vitro selection experiment to select for a DNAzyme having ribonuclease

activity was conducted using a procedure adapted from Breaker and Joyce (Breaker &

Joyce, 1995). A library was constructed containing a 5'-primer constant region and a

substrate segment, which contained an internal ribo-adenosine at position 28. This was

connected to a 3'-primer constant region by a segment, 40 nucleotides in length, having

random sequence. The 5'-constant region used by Breaker and Joyce was altered to

minimize a base paired "clamp" engineered to favor association between the substrate

portion and random portion (see Figure 3.2a) of the DNAzyme. Table 3.1 presents a

summary of the many oligonucleotides discussed in this chapter.

A cycle of selection and amplification was applied for 7 rounds to the library. A

sampling of the DNA molecules recovered was then cloned and sequenced. Several

DNA molecules having distinct sequences were isolated. When inspected for catalytic

activity, the molecules cleaved themselves with apparent first order rate constants ranging









from 0.01_5 to 0.049 hrl (Figure. 3.2b). These rate constants are low in comparison to

optimized DNAzymes obtained in the presence of divalent cation (ca. 60 hr )(Santoro &

Joyce, 1997; 1998), but comparable to Mg++-independent catalysts reported by Geyer

and Sen (0.17 hr ) and by Faulhammer and Famulok (0.006-0.024 hrl ). Additional

rounds of selection isolated variants of the sequences initially identified in round 7.

The behavior of one catalyst, termed 614, isolated from the seventh round of

selection was examined. DNAzyme 614 was chemically synthesized and purified by

polyacrylamide gel electrophoresis (PAGE) by Integrated DNA Technologies (Coralville

IA). Large quantities of internally radiolabeled 614 were generated by PCR using this

614 template. The catalytic strand of the double stranded PCR products, termed ribose-

614, was separated from its complement by two methods. In some cases, ribose-614 was

generated with a 5'-phosphorylated complementary strand primer (conpl 5 'P).

Following PCR, the complement was digested with lambda exonuclease. Alternatively,

ribose-614 was generated using a complementary strand primer with a 15-nt poly-

adenosine tail attached to its 5' end via a PEG linker (conpl tail). By increasing the

length of the complementary strand, the tail allowed PAGE separation of the catalytic

strand from its complement.

Purified ribose-614 was resuspended in HEPES buffer, and cleavage reactions were

initiated by adding equal volume of 2X reaction buffer, immediate heating to 960C for 3

min, and slow cooling to 230C over 10 min (the set-up). The reaction was then followed

by gel electrophoresis. Ribose-614 (100 nM) cleaves itself at a rate of 0.015 hr- slow in

comparison to previously isolated DNazymes, but still significantly faster than the









uncatalyzed reaction (the uncatalyzed cleavage rate is negligible in 300 hours, estimated

from the cleavage of ribose-library shown in Figure 3.5)

Cleavage of 614 does not go completion

A progress curve for ribose-614 cleavage shows that the product formation ceased

after ca. 65% conversion; this is a "cleavage plateau" (Figure 3.3a). Such plateaus are

frequently seen in selected DNAzymes and RNAzymes, but are rarely explained. We

considered six possible explanations for the failure of the cleavage reaction to go to

completion:

1. The complementary strand may have been incompletely removed in the set-up.

This strand may inhibit the reaction, or may erroneously increase the estimate of

the uncleaved product if the radiolabeled complementary strand is assigned to the

reactant;

2. Part of the substrate may be missing the adenine riboside that offers the cleavable

site, due to failure in the synthesis of the cat ribose primer;

3. The reaction might be reversible, with the 65% cleavage representing the

achievement of the equilibrium between uncleaved substrate and cleaved products;

4. The products of the reaction may inhibit 614;

5. Part of 614 may adopt a conformation that is inactive;

6. Part of the 614 might no longer have the correct sequence, having incurred

mutation during the synthesis of the template or amplification. We examined each

possibility.










A -.:, .B -..









Figure 3.3 Rioe-1 claaei uafce b hdiiof1%copeetr








Sribos-614(0 n)pu eqioa complementaryin~161 stran~d (circles).

Iniiione .3 Rby Incompletely i Remov eed b ComleentaryStrand does n ot Acount fry
th lteau d diino qioa mut o h opeetgetylwr
Tecomlemvaenrte ofd 614termed(a cnl-61) ase addfied insal munst ribose-6

614 ([riose-614]= 115 n, [on plo-614] =11.5 nM). h initia cmlemnavag sratand 1.


of rbose614(11ars.5 nM ) p laae flus cool61 (1.15-1 nM). Eprm nts writeqanle) aounso
~~ribose-614 (200 nM) an opl-64 20 nquM)l foundtha teplteauy wtasd dramatcall

loer( igue33) vn huhteiitial b nopee Rmv cmleavnage rate asd uncaned. Simla results


were seenen wit rioe614 at50 M plu conpl-614) wat 50e inM Thsm showed tha thoe-

61 [ioe64 1 M complementar strandM).T inhibite cleavage b61,adteiprncofraemovng h


o ioe6 ( M l comple64(.5nM.Eprmentar stran prior to masurmen ofDNzmkiecs







We then asked whether the presence of complementary strand could explain the

incomplete cleavage of ribose-614. Two independent procedures (exonuclease digestion









and asymmetric PCR) were used to generate ribose-614 lacking its complement.

Experiments with each yielded approximately the same cleavage plateau. This suggested

that the plateau is not the consequence of incomplete removal of conpl-614, as it is

unlikely that the amounts of complement remaining following the two procedures are the

same. In fact, it is difficult to imagine that any conpl-614 remains following the

purification using the asymmetric PCR procedure, as the complementary strand is 15-nt

longer than the catalytic strand, and would almost certainly be removed by PAGE

purification.

To assess the amount of complementary strand remaining after exonuclease

treatment, we treated ribose-614 generated via the exonuclease method with base (0.5 M

NaOH, 800C, 1 h). This cleaves all of the substrate at the adenine riboside site. The

amount of cleaved products was similar to that with the 5'-E2P labeled cat ribose primer

(85-95% cleaved for ribose-614 vs. 90-97% cleaved for cat ribose primer following

treatment with base). The incomplete cleavage of cat ribose primer by base suggest a

lack of chemical susceptibility of the primer (discussed below). Since contaminating

complementary strand cannot be present in the cat ribose primer, the ca. 5% difference

between base cleavage of the cat ribose primer and the full length ribose-614 is an

estimate of the upper limit of the amount of complementary strand that might remain

from incomplete degradation by exonuclease.

The efficiency with which lambda exonuclease removes the complementary strand

was also tested using a 5'-E2P labeled cat ribose primer to synthesize full-length, double

stranded ribose-614 without 5'-phosphate on the complementary strand primer. The

duplex PCR product was divided into two aliquots. One was subj ected to standard










exonuclease treatment, while the other sample was untreated. The samples were resolved

by PAGE-urea to determine the amount of full-length product. Less than 10% of the

original phosphorylated strand (compared to untreated sample) remained following

exonuclease digest. These results rule out contamination by residual compl-614 as a

maj or cause of incomplete 614 cleavage .

An Approach to Chemical Equilibrium does not account for the Plateau

The failure of a substrate to be completely transformed to product may result from

an approach to chemical equilibrium between substrates and products in a reversible

reaction. To test for this, ribose-614 (200 nM) was incubated for 144 hours. At this time,

plateau had been reached, and the reaction mixture was divided into three aliquots. One

aliquot was reserved. The second was diluted 25-fold with reaction buffer. Diluting is

expected to drive the equilibrium towards the cleaved product. An equal amount of a

variant of ribose-614, deoxyribose-614, was added to the third aliquot. The compound

deoxyribose-614 is the same sequence as ribose-614, but with an uncleavable

uncleaveable adenine 2'-deoxyriboside replacing the cleavable adenine riboside at

position 28. As discussed below, deoxyribose-614 is a true catalyst, acting in trans. Its

addition in excess to ribose-614 ensured that catalytic activity was sufficient to see

cleavage at the plateau if it was occurring. Neither treatment significantly altered the

plateau, excluding reversibility as its explanation.

In a second experiment, the 78-nt product of ribose-614 cleavage was isolated via

gel purification. This product was radiolabeled and, following gel purification, incubated

(in excess) with full-length ribose-614 (unlabeled). As the unlabeled ribose-614 becomes

cleaved, and if the reverse reaction (ligation) occurs, excess 78-nt ensures that it may

capture by ligation the 28-nt product, converting 78-nt to full-length ribose-614 over










time. The reaction was monitored for 400 hours. No conversion of 78-nt to full-length

material was observed. This also excludes reversibility as an explanation for the plateau.

Testing if the Cleavage Products are Acting as Catalysts or Inhibitors

By incorporating a short self-complementary segment that encourages a hairpin to

clamp together the substrate and catalytic portions of the DNAzyme, Breaker and Joyce

hoped to increase the chance that their library would contain molecules that self-cleave

rapidly in cis. Further, cis-cleavage might be expected to predominate over trans

cleavage, because the substrate is covalently bonded to the catalyst. The possibility that

either cleaved product continues to act as a ribonuclease in transrt~t~rt~t~rt~t~rt~ or as an inhibitor cannot

be ruled out. This is especially true for 614, where the clamp that might favor a hairpin,

and therefore cis-cleavage, is not present.

To test this, the 28- and 78-nt products were gel purified. These products were

added in equal amounts to a sample of uncleaved ribose-614. Cleavage of ribose-614

occurred at the same rate in the presence or absence of the 28- and 78-nt fragments

(Figure 3.4). This suggests that 28- and 78-nt do not act as inhibitors or catalysts, at least

when both are present in stoichiometric amounts (approximately the highest

concentration they reach under normal 614 cleavage conditions).

The 28- and 78-nt products were also added to a sample of the ribose-library used

in the selection. The ribose-library contains the cat+ribose primer followed by a

segment of random sequence. Thus, it contains substrates that are (for the most part) not

catalysts, and serves as the opposite of deoxyribose-614, which is a catalyst but not a

substrate. The ribose-library alone does self-cleave to a detectable extent, as expected

given that an unselected, random library has very few active catalysts.





















0 50 1DD 150 200 250 300
hours
nbose-614 )( nbose-library
A nbose-614 +28-nt product nbose-library + 28-nt product
r nbose-614 +78-nt product o nbose-library + 78-nt product
nbose-614 + 28 & 78-nt prod a nbose-library + 28 & 78-nt prod

Figure 3.4. Cleavage products do not affect ribose-614 cleavage, and only minimally
alter ribose-library cleavage. Ribose-614 or ribose-library were incubated
either alone (400 nM), or with the addition of the 28-nt product of 614
cleavage (400 nM), the 78-nt product of 614 cleavage (400 nM), or both (400
nM each). Data are fit to a single exponential curve. Errors in percent cleaved
are < +2 percentage points.

The 28-nt product alone did not cleave ribose-library, showing that this product is

not a catalyst. The 78-nt product alone did cleave ribose-library, although with a very

low rate constant (kobs = 0.0006 hr-l, 35-fold lower than ribose-614 under similar

conditions). The 28-nt product with the 78-nt fragment reduced cleavage of the ribose-

library below that observed with the 78-nt fragment alone. This inhibition is presumably

because 28-nt product competes with the ribose-library substrate in binding to the 78-nt

catalyst. Inhibition of 614 by 28-nt is not noticeable presumably because 28-nt does not

compete as effectively for 614 as a second molecule of 614 does.

Improperly Folded ribose-614 Accounts for Part of the Plateau

We then asked whether ribose-614 folds into active and inactive conformations,

with the inactive form contributing to the cleavage plateau. Predictions of the

conformation of ribose-614 using M~fold (Zuker, 2003) showed several potential

structures. Close inspection of ribose-614 on a non-denaturing gel suggested two bands










present in the starting material, one converting to a third band upon incubation under

reaction conditions, and the other remaining unchanged. This is consistent with the

hypothesis that ribose-614 adopts active and inactive conformations.

To test this, ribose-614 was incubated until the plateau was reached (140 h). The

mixture then was diluted into 2 volumes of formamide and heated (900C, 2 min). The

uncleaved ribose-614 was purified by PAGE/urea, resuspended in buffer, re-folded using

the slow cooling protocol, and subjected to cleavage conditions a second time. An

additional 25% of the ribose-614 sample was cleaved in the 300 hours following gel

purification. This additional cleavage following gel purification suggests that some of

the initially uncleaved ribose-614 was in an inactive conformation. It is notable,

however, that the gel-purified ribose-614 reached a plateau of ca. 25%, significantly

lower than the cleavage plateau of the original sample (Figure 3.5).





S50-
S40-
S30-



0 50 100 150 200 250 300 350
hours
A nbose-614 gelpunfled nbose-614

Figure 3.5 Gel-purification ofribose-614 at cleavage plateau results in additional
cleaveage, indicating a fraction of 614 is folded into an inactive conformation.
Ribose-614 (222 nM) was allowed to self-cleave for 140 hours, nearing
cleavage plateau (triangles, top curve), at which point half the sample was
resolved with denaturing PAGE. The fraction of ribose-614 remaining
uncleaved at 140 hours was gel-purified, resuspended in reaction buffer (to a
final concentration of 100 nM) and incubated for additional time (diamonds,
bottom curve; percentage cleaved as a fraction of the label in the gel purified
product, not of initial substrate). Errors in percent cleaved are < +2 percentage
points









If misfolding were the only cause of incomplete cleavage, a plateau of

approximately 65% would be expected in the second round of cleavage. The fact that the

gel-purified sample reaches a lower plateau indicates that misfolding may account for

only ca. 9% of total uncleaved fraction in the original sample (as an additional 25% of the

sample cleaved following gel purification, and ca. 35% of the original sample remains

uncleaved, the fraction of the initial sample that is misfolded is 25% of 3 5%, or 9%).

We considered the possibility that gel-purification removed an inhibitor of

cleavage. Ribose-614 was incubated to reach the plateau (141 h), and an aliquot was

heated and slowly cooled. Any molecule in an inactive conformation was thus given

another chance to adopt an active conformation (Figure 3.6). As in the gel-purification

experiments, denaturing and refolding via this procedure increases the amount of material

cleaved. The cleavage plateau over the first 150 hours following re-heating is about 10%

higher than the untreated sample (in agreement with the estimate of 9% above), but still

well below the ca. 90% cleavage when treated by strong base. When untreated samples

are incubated beyond 300 hours, the cleavage levels approach those seen when samples

are reheated at 140 hours, suggesting that re-folding into active conformations may occur

slowly at room temperature. Thus, alternative conformations account for some, but not

all, of the plateau seen between 150 and 300 h.

Mutations Introduced into 614 during Cloning and Sequencing Account for part of
the Plateau

The ability of base to cleave ca. 94% of the cat ribose primer suggests that about

6% of the cat ribose primer, and therefore any full length DNAzyme made from the

cat ribose primer, may be missing the adenine riboside unit that is the site for cleavage.












70-




S30-



U 50 100 150 200 250
hours
Snbose-614 A nbose-614 denatured and
refolded at 141 hours

Figure 3.6. Reheating ribose-614 results in additional cleavage, indicating a fraction of
ribose-614 is folded into an inactive conformation. Ribose-614 (100 nM) was
allowed to react until it reached cleavage plateau (141 h), at which point half
of the sample was denatured by heating to 960C for 3 min, and slowly cooled
to 230C over 10 min. Errors in percent cleaved are < +2 percentage points

Other sequence variants may be present throughout ribose-614 as well, either as a

result of mutations introduced during the synthesis of the primers and template, or

mutations introduced by the polymerase during PCR. These mutations may reduce or

eliminate the catalytic power of a fraction of the ribose-614 DNAzyme pool.

This possibility was examined by cloning and sequencing DNA molecules from the

cleaved and uncleaved fractions of ribose-614 after the plateau had been reached (Figure

3.7). Of the 29 sequenced clones of the 78-nt cleaved product of ribose-614, only four

(14%) were found to be mutants. The mutations, at three sites, were in the N40 region

between the primers.

In contrast, of the 66 clones from the portion of ribose-614 that remained uncleaved

at the plateau, only 64% had at least one mutation (42 individuals). Some 18% had a

mutation in the N40 region, 5% had a mutation in the complementary strand primer, and

48% had a mutation in the catalytic strand primer (including 20% of the uncleaved

ribose-614 molecules sampled that were missing the adenine riboside cleavable site).















Excluding molecules missing the adenine riboside cleavable site, 44% of the uncleaved



molecules possessed a mutation. This corresponds to 15% of the total initial population



(44% of ca. 3 5% of total initial population remaining uncleaved at plateau).



The extent of misfolding in 614 was estimated by assuming that all unmutated



sequences from the uncleaved pool at the plateau remained uncleaved because they had



adopted an inactive conformation. Given this, 13% of the total population is calculated to



be misfolded (36% of the unreacted material is not mutated, and the unreacted material at



plateau is ca. 35% of the total).



clone name: Sequence 1 n
614wt (x25) -------------------GACATGGCCTCTCAC ATCATGCGAGCACAC-G-CAATAGCCT~ATAAGGTITGGTACGTACTGCC .1
614mut#1 -------------------GACATGGCGCTCTCAC ATCATACGAGCACAC-G-CAATAGCCT~GATAAGGTITG;GTACGPACTGAC00
614mut#2 ----------------GC4AAGACATGGCGCTCTCAC ATCATGCGAGCACAC-G-CAATAG ACT~ATAAGGTTU;AGGPACGPCTGA 00r
614mut#3(x2) -------------------GACATGGCGCTCTCAC ATCATGCGAGCACAC-G-CAATAGCCT~ATAAGGT -U;ATAGGTPGPAGA 00
614wt (x21) CT-MAGPTTCTAATACG-ACTCA-CTA-TAGGMGACATUCCTTCC ATCATGCGAGCACAC-G-CAATAGCCT~ATAAGGTITGGPA CGTACTGCC .1
614mut#31 CT-MAGPTTCTAATACG-ACTCA-CTA-TAGGMGACATMG NGANTNTCACATCATGCGAGCACA C-G-CAATAGCCTATA GGI TACGPATGGCAA
614mut#33(2) CT-MAGPTTCTAATACG-ACTCA-CTA-TAGGMGACATGCMC NTCACATCATGCGAGCACAC-G-CAATAGCCT~ATAAGGITGGPAPGCTGCA
614mut#5 (x6) CT-MAGPTTCTAATACG-ACTCA-CTA-T -MAGACATGGCGCTCTCAC ATCATGCGAGCACAC-G-CAATAGCCT~ATAAGGTITGGPA CGTAGTGGA
614mut#11 CT MCAGPTTCTAATACG-ACTCA-CTA-T -MAGACATGGCGCTCTCAC ATCATGCGAGCACAC- A-CAATAGCCT~GATAAGGTI~UTGGTGPA PACTGAC00
614mut#12 CT-MAGPTTCTAATACG-ACTC -CTA-T -MGACATGGCGCTCTCAC ATCATGCGAGCACAC-G-CAATAGCCT~ATAAGGTITGGPA CGTAGTGGA
614mut#13 CT-MAGPTTCTAATACG-ACTCA- -TA-T-M-CATGGCGCTCTCAC ATCATGCGAGCACAC-G-CAATAGCCT~ATAAGGTITGGPA CGTAGTGGA
614mut#14 (x2) CT-MAGPTTCTAATACG-ACTCA-CTA- --MGACATGGCGCTCTCAC ATCATGCGAGCACAC-G-CAATAGCCT~ATAAGGTITGGPA CGTAGTGGA
614mut#16 CT-MAGPTT NTAATACG-ACTCA-CTA-T -MAGACATGGCGCTCTCAC ATCATGCGAGCACAC-G-CAATAGCCT~ATAAGGTITGGPA CGTAGTT
614mut#17 CT-MAGPTTCTAATACG-ACTCA-CTA-T -MAGACATGGCGCTCTCAC ATCATGCGAGCACAC-G-CAATAGCCT~ATAAGGTITGGPA CGNAGTGGA

614mut#18 CT-MAGPTTCTAATACG-ACTCA-CT --AGGAAGACATGGURCTCTCAC ATCATGCGAGCACAC-G-CAATAGCCT~ATAAGGTITGGPA CGTAGTGGA
614mut#19(x3) CT-MAGPTTCTAATACG-ACTCA-CT -TAGGMGACATGGCGCTCTCAC ATCATGCGAGCACAC-G-CAATAGCCT~ATAAGGTITGGPA CGTAGTGGA
614mut #2 2(x2) CT-MAGPTTCTAATACG-ACTCA-CTA ATAGGAAGACATGGURCTCTCAC ATCATGCGAGCACAC-G-CAATAGCCT~ATAAGGTITGGPA CGTAGTGGA
614mut #2 4 CT-MAGPTTCTAATACG-ACTCA-CTA- -AGGAAGACATGGURCTCTCAC ATCATGCGAGCACAC-G-CAATAGCCT~ATAAGGTITGGPA CGTAGTGGA
614mut #25 CT-MAGPTTCTAATACG-ACTCA-CTA-TAGGMGACATGGC AACTCTCAC ATCATGCGAGCACAC-G-CAATAGCCT~ATAAGGTITGGPA CGTACTGCA .0
614mut#26 CT-MAGPTTCTA -TACG-ACTCA-CTA-TAGGMGACATGGCGCTCTCAC ATCATGCGAGCACAC-G-CAATAGCCT~ATAAGGTITGGPA CGTAGTGGA
614mut #27 CT-MAGPTTCTA CTACG-ACTCA-CTA-TAGGMGACATGGCGCTCTCAC ATCATGCGAGCACAC-G-CAATAGCCT~ATAAGGTITGGPA CGTAGTGGA
614mut #2 8 CT-GGAGATTCTAATACG-ACTCA-CTA-TAGGMGACAT GGTCCC ATCATGCGAGCACAC-G-CAATAGCCT~ATAAGGTITGGPA CGTAGTGGA
614mut #2 9 CT TGCAGAATTCTAATAC C-ACNCAGCTA-TAG NAAGACATGGCGCTCTCAC ATCATGCGAGCACAC-G-CAATAGCCT~ATAAGGTITGGPA CGTPGTTGC
614mut#30 CT-MAGPTTCTAATACG-ACTCA-CTA-TAGGMGACAT AGCGACTCTCAC ATCATGCGAGCACAC-G-CAATAGCCT~ATAAGGTITGGPA CGTACTGCA .0
614mut #32 CT-MAGPTTCTAATACG-ACTCA-CTA-TAGGMG TCATGGCGCTCTCAC ATCATGCGAGCACAC-G-CAATAGCCT~ATAAGGTITGGPA CGTAGTGGA
614mut #34 CT-MAGPTTCTAATACG-ACTCA-CTA-TAGGMGACATUCMC -CTCAC ATCATGCGAGCACAC-G-CAATAGCCT~ATAAGGITGGPAACTGGAC.00
614mut #35 CT-MAGPTTCTAATACG-ACTCA-CTA-TAGGMGACATUCCTTCC ATCATGCGAGCACAC-G-CAATAGCCT~ATAAGGTITGGPA CGAAGTGGA
614mut#37 CT-MAGPTTCTAATACG-ACTCA-CTA-TAGGMGACATUCCTTCC ATCATGCGAGCACAC-G-CAATAGCCT~ATAAGGTITGGPA CGAACTGG
614mut #38 CT TGCAGTTCTAATAC C-ACNCAGCTA-TAG NAAGACATGGCGCTCTCAC ATCATGCGAGCACAC-G-CAATAGCCT~ATAAGGTITGGPA CGTPGTTGC

614mut#39 CT-MAGPTTCTAATACG-ACTCA-CTA-TAGGMGACATUCCTTCC ATCACGCGAGCACAC-G-CAATAGCCT~ATAAGGPTGGPA TAGTPGTTGA 00
614mut #4 0 CT-MAGPTTCTAATACG-ACTCA-CTA-TAGGMGACATUCCTTCC ATCATGCGAGCACAC AG-CAATAG TCT~ATAAGGPU;A TGG ACGPCTGA 003
614mut#41 CT-MAGPTTCTAATACG-ACTCA-CTA-TAGGMGACATUCCTTCC ATCATGCGAGCACAC-G-CAATA TCCTGATAAGGT~UTTGGPACGTACTGA
614mut #42 CT-MAGPTTCTAATACG-ACTCA-CTA-TAGGMGACATUCCTTCC ATCATGCG TCACAC-G TCAATAGCCT~ATAAG CTIUTATCGG TPGPAGA
614mut#43 CT-MAGPTTCTAATACG-ACTCA-CTA-TAGGMGACATUCCTTCC ATCATGCG TCACAC-G-CAATAGCCT~ATAAG CTIUTATCGG TPGPAGA
614mut#44 CT-MAGPTTCTAATACG-ACTCA-CTA-TAGGMGACATUCCTTCC ATCATGCGAGCACAC-G-CAATAGCCT~ATAAGGTITG -TAGTAGTACTGA000
614mut #45 CT-MAGPTTCTAATACG-ACTCA-CTA-TAGGMGACATUCCTTCC ATCATGCGAGCACAC-G-CA TTAGCCT~ATAAGGTI~UT GGPA GTAGTGG
614mut#46 CT-MAGPTTCTAATACG GACTCA-CTA-TAGGMGACATGGCGCTCTCAC ATCATGCGAGC TCAC-G-CAATAGCCT~ATAAGGTI~UTGGTGPA PGCTGA
614mut#47 CT-MAGPTTCTAATACG-ACTCA-CTA-TAGGMGACATUCCTTCC ATCATGCGAGCACAC-G-CAATAG TCT~ATAAGGPU;A TGG ACGPCTGA 00F
614mut#48 CT-MAGPTTCTAATACG-ACTCA-CTA-TAGGMGACATUCCTTCC ATCATGCGAGCACA T-G-CAATAGCCT~ATAAGGPTUTG GTGPA PGCTG
614mut#49 CT-MAGPTTCTAATACG-ACTCA-CTA-TAGGMGACATUCCTTCC ATCATGCGAGCACAC-G-CAAT TGCCT~ATAAGGITU;AGGPACGPCTGA 001
S2




Figure 3.7. Sequence alignment of cleaved and uncleaved cloned 614 sequence variants.

Mutations of 614 are highlighted in yellow; "N40" region between primers is

bold in each sequence and underlined in the original (no mutation) 614

sequence. Apparent first order catalytic rate constants (at 100 nM, in units of

hr-l) are shown to right for a sample of mutant-614 DNAzymes.


This is consistent with the estimate above based on the outcome of a cycle of



unfolding and refolding.









Several of the 614 mutants were tested to see if the mutations in fact reduced the

rate of self-cleavage. Eleven of the twelve 614 mutants that were tested showed cleavage

rates reduced by 10 to 100 fold compared to ribose-614.

Ribose-614 Catalysis is not Mg++-dependent

The mono and divalent cation requirements for ribose-614 activity were then

examined. Reducing the concentration of NaCl from 1 M to 0.1 M eliminated cleavage.

Only a modest change in the rate of cleavage of ribose-614 (100 nM) was observed when

the reaction was run in the absence of MgCl2 (with and without 1 mM EDTA, initial rate

1.32 x 10-2 hr-l and 1.37 x 10-2 hr-l, respectively) or in the presence of MgCl2 (1 mM,

1.53 x 10-2 hr-l; 10 mM, 1.70 x 10-2 hr-l; or 100 mM, 2.52 x 10-2 hr-l). Catalysis by 614

therefore does not require Mg++ in significant amounts even though 614 was selected in

the presence of MgCl2 (1 mM). Requirement of a trace of Mg" cannot, of course, be

excluded. The rates of two other catalysts isolated from this selection (ribose-62 and

ribose-616) were also largely unchanged when the MgCl2 (1 mM) of the reaction buffer

was replaced with EDTA (1 mM). For ribose-625, another molecule isolated from the

selection, cleavage was eliminated by replacement of MgCl2 by EDTA at 250C.

Ribose-614 Cleaves in trans

A series of experiments was then performed to examine the rate of cleavage as a

function of concentration. Self-cleavage is expected to be kinetically first order. Cleavage

in trans is expected to be kinetically second order, meaning that an apparent first order

rate constant kobs will not be independent of [DNAzyme], and fall to zero as [DNAzyme]

falls to zero. A convenient way to separate simultaneous intra and intermolecular











processes is to plot the apparent kobs as a function of [DNAzyme]. The y-intercept is kuni

for the unimolecular reaction, while the slope of the line, reflects a second order process.

Figure 3.8 shows a plot of the apparent kobs versus [ribose-614]. At low [ribose-

614], the rate is independent of [ribose-614]; primary cleavage profiles are shown in

Figure 3.9. At higher concentrations, however, the apparent kobs increases with

increasing [ribose-614]. These results are consistent with a model that includes both

unimolecular and bimolecular processes.

O 035-


~0025-



0i 0010- a
10005-

D 1 1 10 100 1000
[ribose-614] nM

Figure 3.8. Initial rate of ribose-614 cleavage as a function of [ribose-614]. At higher
concentrations, a second order process is apparent. The rate constant for the
apparent first order process (uncorrected for the cleavage plateau), from the y
intercept, is ca. 0.006 hr-l.

A best-fit line extrapolated to infinite dilution for [ribose-614] 3.5 nM and below

(the region where the process is mostly unimolecular) gives an intercept corresponding to

a unimolecular rate constant of 0.006 hr-l. This is an underestimate because the rate

constant, based on initial rates, is uncorrected for the cleavage plateau (see below).

An alternative way to obtain a rate constant for a first order process plots the log of

uncleaved [ribose-614] (substrate remaining) versus time. Here, the progress of cleavage

at low [ribose-614] fit a linear model well, while the progress at higher concentrations

did not (Table 3.2). This suggests that below 20 nM, a unimolecular rate process












A vs-
K exp fR
[nhose-614] (td)-i
m m ~430 0nM 0 061
g 50-1 A-~~ 1080nM 0031
7 194nM 0023
35nM 0020
25- 1 1nM 0 011
S 0 3nM 0 011

U50 100 150 200 250
hours
B
150-

125- y-" [nm-l] at)
S430 0nM 0 03233
~100-( j A I 1080nM 001627
75-1 / 7/ 194nM 000959

50- 35nM 000899
1nM 000695
25- 03nM 000610

0 2 6 8 10 12 14
hours


Figure 3.9. Ribose-614 cleavage rate is concentration dependent. (A) Complete time
course for ribose-614 at various concentrations. Rates are estimated based on
a fit to a single first-order exponential equation. (B) Linear initial phase of
time course. Rates are estimated based on the slope of best fit line.

dominates, while a bimolecular rate process contributes above 20 nM. The rate constant


estimated in this manner for [ribose-614] at 0.3 nM was 0.0042 hr-l (uncorrected for the


plateau), in good agreement with the estimate above.


Table 3.2. Data from plot In[S]t versus time for ribose-614 cleavage at various
concentrations fit to a linear equation.
[ribose-614] nM 430 108 19.4 3.5 1.1 0.3

(slope oflIn[S],,) 0.0032 0.0035 0.0030 0.0031 0.0030 0.0042

R2 0.78 0.89 0.92 0.90 0.97 0.97




We then asked if the progress curve for cleavage at high and low [ribose-614] fit


best to a single exponential (in which both the rate and plateau can vary) or the sum of


two exponential equations (in which both the rate and plateau can vary for each of two









equations). At ribose-614 concentrations <3.5 nM, progress curves fit best to a single

exponential. At higher concentrations, the progress curve is fit best by the sum of two

independent exponentials. This suggests that a single process dominates transformation

of ribose-614 at low concentrations. If multiple active conformers exist at low [ribose-

614], they are either in rapid equilibrium, or self-cleave with comparable rate constants.

The first order constant was then corrected given information collected above about the

plateau. After correction, a single exponential curve fit to the progress curve for 0.3 nM

ribose-614 cleavage under unimolecular conditions. A rate constant 0.011x10-2 hr' was

estimated (with a plateau of 61%).

Various ribose-containing Substrates are Cleaved by deoxyribose-614

To establish trans cleavage in this system, internally labeled ribose-614 (which

yields the 78-nt fragment as the only labeled product) was incubated with 5'-32P-labeled

cat ribose primer (100 nM). The cat ribose contains the substrate domain of ribose-

614, but is not catalytically active. Here, cat ribose is cleaved, as is ribose-614 (Figure

3.10, right panel). This establishes transrt~t~rt~t~rt~t~rt~ cleavage by ribose-614. The cleavage of ribose-

614 (100 nM) in the presence of cat ribose primer (100 nM) was lower than ribose-614

incubated alone (at 100 nM), suggesting that the cat ribose primer competes with

ribose-614 for transrt~t~rt~t~rt~t~rt~ cleavage by ribose-614 (Figure 3.11).

Trans cleavage was also tested by challenging deoxyribose-614 (a modified form of

ribose-614 in which the adenine riboside was replaced by a non-cleavable

deoxyadenosine) with various substrates, each containing the cat ribose sequence. These

included (a) the cat ribose primer, (b) a pool of molecules incorporating the cat ribose

primer followed by a random region and a 18-nt constant region (ribose-library), which







70


collectively have no detectable catalytic activity, (c) a single clone from the ribose-

library, (ribose-lib62), which also has no detectable catalytic activity, and (d) a mutant

of 614 (614AC72T) that has ca. 30 fold reduced activity compared to 614, at 100 nM.


Time (hours)


0 3 9 16 23 4 69 91

< d-614 r61> W gg


O 2 4 7 10 24I 32 48B 72 95

r n *


cleaved r-614 >


- a a ag)


< cat-ri~se z 11,1)1~


, I* *


< cleaved cat-ribose >


* e *


Figure 3.10. Both deoxyribose-614 (left panel) and ribose-614 (right panel) cleave
cat+ribose primer in trans. Internally labeled deoxyribose-614 (100 nM) or
ribose-614 (100 nM) were mixed with 5'-end labeled cat+ribose (100 nM) and
incubated under reaction conditions. The deoxyribose-614 does not contain
the ribo-adenosine and therefore does not cleave itself. Unless otherwise
stated, all kinetics were performed under standard reaction conditions, namely
IM NaC1, 1 mM MgCl2, 50 mM HEPES, 250C, and errors in percent cleaved
are < +2 percentage points.


hours
cleavage of ribose- 614
V cleavage ofribose-6814 with cat+nbose competitor substrate
A cleavage of catmnbose by rbose-614

Figure 3.11. Cat ribose competes with ribose-614 for cleavage. Ribose-614 (100 nM)
was incubated with and without cat ribose (100 nM). Cat ribose is not
catalytic and when incubated alone is not cleaved. Data are fit to a single
first-order exponential equation.










Each substrate was cleaved by deoxyribose-614 (Figure. 3.12) although with

different efficiencies. Ribose-614AC72T (which should fold like ribose-614) is cleaved

by deoxyribose-614 more slowly (suggesting that ribose-614AC72T may fold so as to

make the substrate domain inaccessible to catalysts), while the cat ribose primer is

cleaved faster. The fact that the small cat ribose primer is better than full length

substrates is consistent with the view that folding in longer substrates inhibits transrt~t~rt~t~rt~t~rt~

cleavage by deoxyribose-614. The unimolecular and bimolecular mechanisms by which

614 can cleave are summarized in Scheme 3.1.




70-

60-






20-

10-


0 10 20 30 40 50 60 70 80 90 100
hours
Substrate:
Scat+ribose O ribose-library
A ribose-lib62 o ribose-614AC72T


Figure 3.12. Compound deoxyribose-614 can cleave various substrates in trans.
Unlabeled deoxyribose-614 (270 nM) was incubated with various radiolabeled
substrates (cat ribose, closed squares; ribose-lib61o2, closed triangles;
ribose-library, open diamonds; or ribose-614AC72T, open circles) each at
7.5 nM). Each substrate possessed the same 5'-primer sequence containing
the adenine riboside. Compound deoxyribose-614 does not cleave itself, and
percentage cleaved is the fraction of the substrate converted to product. Data
are fit to a first order exponential curve. Data points being accurate to < +2
percentage. Errors in the measurement of time are 11 minute.







72


Unimolecular Kinetic Scheme:
k1(uni) kcat (uni)
614unfolded = 614folded Products


Bimolecular Kinetic Scheme:


k1 (bi) k cat (bi) E = ribose-614 or deoxyribose-614
E +S E-S a EP S = ribose-614, ribose-lib62 or cat ribose
k-1 (bi)
Scheme 3.1. 614 can cleave via a unimolecular and bimolecular mechanism.


Competition Studies of Ribose-614 Cleavage

We then tested the ability of three of the above substrates, each with their ribose

replaced by 2'-deoxyribose (cat deoxyribose primer, deoxyribose-61 4AC72T, and

deoxyribose-lib62), to compete with ribose-614 for self-cleavage. These competitors

were added in 9-fold excess over ribose-614. For comparison, 9-fold excess of

deoxyribose-614 was added to ribose-614.

The addition of deoxyribose-614 (270 nM) to ribose-614 (30 nM) increased the rate

of ribose-614 cleavage to the same level as seen for ribose-614 at 300 nM (Figure 3.13).


80-1
70- 4Competitor (270 nM)
60- A added to rihose-814
A o A d-614

15 40- d-614AC72T
A 0 no competitor
30 +.o d-lib62
20- e~
V V cat-ribose


O 50 100 150 200 250 300
hours


Figure 3.13. Various substrates can compete with ribose-614 for self-cleavage. A nine
fold excess of unlabeled deoxyribose-lib62, deoxyribose-614, deoxyribose-
614AC72T, or cat deoxyribose was added to radiolabeled ribose-614 (33
nM) at time zero. Errors in percent cleaved are < +2 percentage points.










The cleavage of ribose-614, however, was reduced by excess cat deoxyribose or

deoxyribose-lib62. This showed that these molecules compete for ribose-614 self-

cleavage.

Curiously, addition of a 9-fold excess of deoxyribose-614AC72T to ribose-614 did

not reduce the rate of ribose-614 cleavage, but rather increased it. The ability of

deoxyribose-614AC72T to cleave ribose-614 may indicate that the mutation alters the

ability of deoxyribose-614AC72T to function as a substrate without inactivating its

catalytic domain, consistent with the observation above that ribose-614AC72T is itself a

poor substrate for deoxyribose-614 cleavage.

The competition study above was done at high [ribose-614] to favor transrt~t~rt~t~rt~t~rt~ cleavage.

A parallel study at low concentrations of ribose-614 (favoring unimolecular cleavage)

showed no inhibition of ribose-614 self-cleavage by deoxyribose-lib62 competitor.

Saturation Kinetics in trans cleavage by deoxyribose-614

Two mechanisms for the trans bimolecular reaction were considered. In the first,

deoxyribose-614 cleaves its substrate on every encounter. This gives a linear slope in a

plot of an apparent rate constant vs. [catalyst] or [substrate] over the entire concentration

range.

An alternative second order process is possible, however. Here, substrate and

catalyst form a complex that can dissociate before it reacts. This gives saturation

Michaelis-Menten kinetics.

To search for saturation kinetics, we exploited deoxyribose-614 as a DNAzyme that

cannot act upon itself, but must act in trans. Concentrations of deoxyribose-614 were

scanned to find a range where the apparent second order rate constant reaches a plateau

reflecting saturation. Various substrates were used, including cat ribose, ribose-lib62,










and ribose-614, at low amounts relative to deoxyribose-614. Cleavage rates did not

increase (2000 to 6000 nM deoxyribose-614), suggesting saturation of the bimolecular

process.

Assuming that unbinding is fast compared to reaction, a calculated fit gave a

dissociation constant (Kd) Of 29.0, 37.3, and 25.5 nM for cat ribose primer, ribose-614

and ribose-lib62 substrates, respectively. The kcat(bi), determined from the maximum

rate of cleavage with saturating enzyme, was 0.056, 0.048, and 0.049 hr-l for cat ribose,

ribose-614 and ribose-lib62 substrates, respectively.

Compound deoxyribose-614 Cleaves with Multiple-turnovers

To show that deoxyribose-614 catalyzes cleavage with multiple turnovers, either

cat ribose or ribose-lib62 substrate were incubated in 4-fold excess over deoxyribose-

614 (133:33 and 400: 100 nM substrate:enzyme). With 400 nM enzyme and 100 nM

substrate, two turnovers of substrate were observed at 100 hours for cat ribose substrate

and 200 hours for the ribose-lib62 substrate.

As seen under single-turnover conditions, the cat ribose substrate is cleaved faster

under multiple-turnover conditions than the full-length ribose-lib62 substrate (at

equivalent concentrations). This may be attributable to a better ability of the cat ribose

substrate to bind with or dissociate from the enzyme, or a lower proclivity for forming

nonproductive interactions.

Catalytic Power in trans is Unaffected by Annealing Protocol

The impact of the slow cooling protocol on transrt~t~rt~t~rt~t~rt~ cleavage was tested by mixing

deoxyribose-614 with substrate (ribose-lib62 or cat ribose) either before or immediately









after the slow cooling. This change in the annealing procedure did not alter noticeably the

cleavage profile.

At high concentrations (200 nM), the rate of cleavage of ribose-614 was unchanged

by omitting the slow cooling. This also shows that intermolecular folding at the start of

the reaction is not significantly effected by the annealing protocol, even though

denaturation by heating allows molecules in inactive conformations to refold into active

conformations. Similar experiments at low [ribose-614] favoring cis-cleavage showed a

slight decrease in initial rate with the omission of slow cooling.

The Commitment Step for deoxyribose-614 Cleavage

If the rate limiting step is dissociation of the product-enzyme complex, a burst is

expected in the initial phase of multiple turnover kinetics. To seek a burst, the

concentration of deoxyribose-614 as a catalyst was held constant at 20 nM while the

concentration of cat ribose as a substrate was varied from 100 to 2000 nM. Turnover of

substrate to product remained linear well beyond the initial turnover for all substrate

concentrations, revealing no burst phase and suggesting that the overall rate is not limited

by DNAzyme*Product dissociation.

To estimate the relative magnitudes of kcat(bi) and k-1(bi), a chase was performed

with unlabeled cat deoxyribose added after four hours. The cat deoxyribose chase is a

substrate analog that cannot be cleaved, and therefore should compete with the labeled

substrate. If all of the substrate-catalyst complex proceeds to product (which is the case if

kcat(bi) >> k-1(bi)), then addition of the unlabeled chase will not influence the subsequent

rate of appearance of labeled products. If, however, the rate of substrate dissociation from

catalyst is faster than cleavage (kcat(bi) << k-1(bi)), then unlabeled chase molecules










lacking a cleavable site should consume the newly disassociated catalyst, and the

production of labeled products should largely cease.

In the chase, deoxyribose-614 enzyme (2 gM, saturating) acted in trans on small

amounts (4 nM) of ribose-614 substrate. When the cat deoxyribose chase was present at

5-fold excess over enzyme (10 pM added at t = 4 h), a decrease in the rate of cleavage of

ribose-614 after addition of the chase was observed (Figure 3.14a). The cleavage levels

of ribose-614 following the addition of chase is reduced below that seen for a

predominately cis-cleavage reaction of 1.1 nM ribose-614. This suggests that kcat(bi) <

k-1(bi)


A 8o- B o




S50- g






chase added att = 4 h chase added at t= 4 h
U 5 10 15 20 25 30 35 40 45 50 0 5 10 15 20 25 30 35 40 45 50
hours hours

Figure 3.14. Cleavage of various substrates by 614 is reduced by the addition of
unlabeled chase.This indicates that the rate of E* S dissociation is faster than
the chemical step of cleavage. (A) Saturating amounts of deoxyribose-614 (2
pLM) cleaving ribose-614 (4 nM), with (closed squares) and without (open
circles) chase (10 pLM) added at 4 h. (B) Saturating amounts of deoxyribose-
614 (2 pLM) cleaving ribose-lib62 (4 nM), without chase (open circles) and
with chase (5 pLM, closed triangles or 10 pLM, closed squares) added at 4 h.

Similar results were seen following the addition of chase to ribose-614 under trans-

cleavage conditions. Cleavage of ribose-614, however, was not completely eliminated by

the chase. This suggested several explanations, including the persistence of cis cleavage









in the presence of chase, or insufficient chase. Chase experiments conducted with ribose-

614 cleaving in cis (2 nM) showed no significant change in cleavage following the

addition of 30 nM chase (data not shown).

If kcat bi)<
more rapidly than it will form products. If association is also relatively rapid, however,

with insufficient chase, the enzyme may bind and release a number of chase molecules

until it finds a labeled substrate. The reduction in cleavage should therefore depend on

the relative amounts of chase, enzyme, and substrate molecules.

To test this directly and without self-cleavage, chase experiments were performed

with saturating levels of deoxyribose-614 enzyme (2 pM) cleaving cat ribose substrate

(4 nM) and chase in either 5 or 10 gM. Cleavage is greatly reduced, but not eliminated,

following the addition of the chase (Figure 3.14b). The reduction of cleavage was

slightly greater with chase in 5-fold excess over enzyme as compared to only a 2.5-fold

excess. Together, these experiments indicate that the rate of E*S dissociation is greater

than the rate of chemical step (kcat bi) << k-1 bi)).

Dependence on Temperature of deoxyribose-614 Cleavage

The temperature dependence of initial rates for ribose-614 cleavage under transrt~t~rt~t~rt~t~rt~

cleavage conditions is shown in Fig. 3.15. The rate is lower by a factor of 2.5 at 250C

than at 150C, and falls dramatically at higher temperatures. Thus, as true for many

biological catalysts, it appears as if the catalyst-substrate complex required for trans

cleavage unfolds (or dissociates) at higher temperatures.

The rate of cleavage of ribose-614 under trans cleavage conditions is essentially

the same between 40C and 150C, and 2.5 fold greater than the rate at the selection










temperature (250C). Similarly, the rate for cis cleavage of ribose-614 is essentially the

same at 40C and 250C (data not shown). The rate of the chemical step is expected (as an

approximation) to increase two-fold for every 100C increase in temperature (Laidler &

Peterman, 1979). Thus, it appears as though the overall rate of catalysis is largely

affected by the folding/association step and not the chemical step.




0.075-




S0.050-




0.025-




0.000 I I I I I I I" I
0 5 10 15 20 25 30 35 40 45
te mpe rature (c.C)


Figure 3.15. Ribose-614 rate of self-cleavage in trans is increased at lower temperatures.
Ribose-614 (200 nM) was incubated at various temperatures from 40C to
420C. Initial rates were calculated and plotted vs. temperature.

The rate of cleavage of ribose-lib62 in transrt~t~rt~t~rt~t~rt~ by deoxyribose-614 in single-turnover

experiments is comparable at 15 and 250C, however. Similar experiments performed

under multiple-turnover conditions reveal a rate enhancement at lower temperatures

during cleavage of the first substrate, followed by cleavage of additional substrates at a

slower rate (Fig. 3.16). This "burst phase" seen under multiple turnover conditions at

15oC is not seen at 25oC. This indicates that lower temperatures increase the initial rate by







79


stabilizing intermolecular association between enzyme and substrate or, perhaps, through

greater stability of the folded form of the enzyme. This stabilization presumably also

slows the dissociation of product from enzyme to within the range of k-1(bi), and therefore

reduces the rate for multiple-turnover subsequent to the first turnover.




1.50-

1.25-1
I o
o 1.00-
e =0

S0.75_1 o

E 0.50-1
co a 15"C
0.25-
o 25"C
0.00 I
O 10 20 30 40 50
hour


Figure 3.16. Burst kinetics (a higher cleavage rate during the first turnover compared to
subsequent turnovers) is enhanced at lower temperatures. Ribose-lib62
substrate (400 nM) was incubated under multiple turnover conditions with
deoxyribose-614 (100 nM) at 150C (closed squares) and 250C (open circles).
The number of substrate turnovers was calculated by multiplying the percent
of substrate cleaved by initial substrate concentration and dividing by
[ deoxyribose-614].

Predictions of the Energetically Favored Structure are not Supported by
Experimental Data

The ability of 614 to cleave in cis is not particularly surprising, because this is the

function for which the DNAzyme was selected. It is therefore interesting that 614 is able

to cleave various substrates in transt~t~rt~t~rt~t~rt~ and that the apparent first order rate constant at

saturation for trans cleavage is 4-fold higher than the first order rate constant for cis-

cleavage. The ability of 614 to cleave the cat ribose primer, as well as a library of









molecules containing the cat+ribose primer, suggests that 614 has a binding motif that

pairs with a part of the cat+ribose primer common to all substrates, thus positioning a

separate catalytic motif near the adenine riboside cleavage site.

Chemical modifications specific for single-stranded DNA were used to probe

secondary structure Deoxyribose-614 (300 nM) was folded overnight with and without

excess unlabeled cat+deoxyribose substrate (10 CIM). The mixtures were then treated

with either potassium permanganate (which modifies thymine) or dimethyl sulfate

(which, at high salt concentrations, modifies guanine) for 2 and 5 min. Recovered

samples were resolved by PAGE-urea next to a 10-bp ladder.

All thymine and guanine nucleotides were somewhat sensitive to their respective

reagents, indicating that either some 614 is not folded at all, or adopts multiple

conformations. Nonetheless, T44, T46, T74, G33, G41, G65, G71, and G75 all

demonstrated some degree of differential protection (numbers refer to the sequence of

614 shown in Figure. 3.7). The pattern of protected and sensitive positions observed

under chemical modifications is not simultaneously compatible with any single structure

predicted for 614 folded in cis or trains. This suggested that either a mixture of

conformers exists, or none of the predicted structures accurately reflect the single true

structure of 614.

Likewise, each of the eight lowest free energy structure predictions made by M~fold

was examined by generating mutations that would disrupt critical helixes. In no case

could a loss of function arising from a nucleotide replacement be rescued by a

compensatory mutation predicted by M~fold. This suggested that none of the predicted

conformations dominated (data not shown).









It is interesting to note that many of the structures reported for in vitro selected

DNAzymes are based on unverified predictions using programs such as M~fold. Although

such programs have been useful for predicting structures for many RNA and DNA

molecules, this does not appear to be the case with 614.

Discussion

In vitro selection experiments offer the possibility of learning how molecular

behavior is distributed within a sequence space defined by the building blocks of a

biopolymer, in this case DNA. Structures within DNA sequence space are countable. For

standard DNA, 4n sequences exist, where n is the length of the biopolymer. This

distribution is relevant to issues as diverse as the origin of life and biomedicine. For the

first, we wish to know how large a hypothetical prebiotic pool of random nucleic acid

sequences must have been to contain the physical and catalytic requirements for life. For

the second, we may want therapeutic or diagnostic DNAzymes, and need to know the

likelihood of obtaining these from selection experiments. In both, we may ask whether

added compounds (including divalent metal ions or other cofactors) increase the

likelihood of obtaining catalysts.

The answers to such questions might exploit the language of univariate statistical

analysis (Johnson et al, 1994; Nelson, 1982). A distribution function P(kobs) relates the

probability of finding a molecule with a particular catalytic power, kobs, to kobs itself.

Because good catalysts are presumed to be scarce in the initial library relative to poor

catalysts, and excellent catalysts are presumed to be scarce relative to good ones, this

distribution is expected to be a decreasing function of kobs. Using such analyses, a better










pool is defined as one having a distribution shifted to the right (Fig. 3.17). Adding a

useful cofactor to the pool should shift the distribution to the right as well.

To address such "big" questions, catalysts being examined must be sufficiently well

behaved that their behavior can be quantitatively analyzed. Many DNAzymes are not,

including (upon first inspection) 614. In particular, the reaction of 614 did not go to

completion. Ca. 3 5% of the starting material is unreacted even after prolonged

incubation. This makes quantitative analysis difficult.


P= be(-bkcat)

0.25-
-- b = 2

0.20 -* b= 5







0.05-


0.00
0.0 0.2 0.4 0.6 0_8 1.0

kat

Figure 3.17. A univariate statistical distribution plotting the probability of encountering a
catalyst with a particular kobs as a function of kobs, for two catalysts, one
with a better pool (open circles, exponential function with b = 2), the other
with a poorer (closed squares, exponential function with b = 5). In each case,
the area under the line sums to unity. Because good catalysts are scarce in the
initial library relative to poor catalysts, and excellent catalysts are scarce
relative to good catalysts, this distribution is a decreasing function of kobs.
The distribution shown is exponential; the true nature of the distribution in
any particular space is, of course, unknown. Key questions in in vitro
selection ask whether this curve is exponential, zipflan, or is best captured by
mathematical approximations having multiple parameters.









Similar behaviors are well known in the literature of nucleic acid catalysts.

Hammerhead ribozymes, for example, are frequently reported to cleave to only 40 60%

at the plateau. DNAzymes with ribonuclease activity and deoxyribonuclease activity are

also frequently reported with self-cleavage plateaus of 25 65%. This behavior must be

understood before any quantitative analysis can begin.

This work accounts for the incomplete cleavage (the plateau) displayed by

DNAzyme 614. Approximately 6-7% of the total ribose-614 sample does not cleave

because it is missing the adenine ribonucleoside linkage, which is the site of cleavage.

Approximately 9-13% of the total sample appears to fold into an unreactive

conformation, which repopulates the active conformation via a cycle of denaturation and

renaturation. As much as 15% of the unreacted material at the plateau appears to be

sequences containing mutations introduced during amplification or the synthesis of

template. These numbers sum to 30-3 5%, approximately the amount of DNAzyme left

uncleaved at the plateau.

Less than 5% of the plateau in our experiments can be attributed to the presence of

DNA complementary to the catalyst. Nevertheless, these results show the importance of

efficient removal of the complementary strand. Contamination by 10% of the

complementary strand has little effect on initial cleavage rate or cleavage plateau, while

greater contamination can greatly reduce the cleavage plateau and therefore estimates for

cleavage rates. For this reason, we believe both asymmetric PCR and exonuclease

degradation are superior to column purification (under strong base) as a way of

eliminating the complementary strand.









Given this understanding of the progress curve of the reaction undergone by 614,

we asked whether the features of this DNAzyme reflect the environment under which the

DNAzyme was selected. We expected that the presence of Mg++ in the selection would

lead to catalytic molecules that used Mg++ as a cofactor. In part, this expectation arose

because Breaker and Joyce themselves reported a Mg++-dependent DNAzyme emerging

from their selection. Indeed, the Breaker-Joyce IVS protocol, by using Mg++ as the

trigger to begin the selection process, seems to require that the DNAzyme be active only

with Mg++

In addition, this expectation is based on a general view that the catalytic potential

of DNA is poor. DNA lacks a range of functional groups, in particular, many of those

present in proteins (Benner et al, 1987). In this view, the probability of finding a good

catalyst that exploits a useful cofactor (such as Mg++) is greater than the probability of

finding a good catalyst that does not. In terms of the distributions shown in Figure 3.17,

adding Mg++ should shift the distribution to the right.

For both reasons, we were surprised to discover that ribose-614 is as active in the

absence of Mg++ as it is in its presence. This behavior was not unique to ribose-614 in

this selection. It was also observed for two other deoxyribozymes isolated from the same

selection, termed ribose-62 and ribose-616. A fourth DNAzyme generated in this work,

ribose-625, was found to be Mg++-dependent at the temperature used for the selection,

but evidently not at 4 OC. While this sample is far from statistical, it suggests that the

distribution of catalysts in the pool examined does not greatly favor Mg++-dependent

catalysts over Mg++-independent catalysts. This, in turn, implies that adding Mg++ to the

library does not shift the distribution in Figure 3.17 greatly to the right.









The emergence from a selection of catalysts that fail to take advantage of available

cofactors is not unprecedented. For example, Faulhammer and Famulok report a selection

performed in the presence of histidine (20 mM) at a low concentration of Mg++ (0.5

mM). These authors hoped to select a DNAzyme that exploited histidine as a cofactor.

They isolated, however, a DNAzyme that was highly active with Ca++ in the absence of

histidine. This behavior was selected despite the fact that Ca++ was excluded from the

selection. Mg++, present in the selection, was a poor substitute for Ca++. At the same

time, Roth and Breaker, also seeking a histidine-dependent nucleic acid enzyme, found

one (Roth & Breaker, 1998).

Our results differ from those reported by Breaker and Joyce, who analyzed a single,

Mg++-dependent catalyst that emerged from an analogous selection. Breaker and Joyce

did not survey their catalysts to determine the ratio of Mg++-independent and Mg++-

dependent catalysts. Thus, it is conceivable that their study of a Mg++-dependent catalyst

differs from our study of a Mg++-independent catalyst because of the stochastic nature of

the selection experiment.

Other explanations are conceivable, however. Although the concentrations of Mg++

were the same in our selection and the Breaker-Joyce selection, the sequences used by

Breaker and Joyce possessed two pairs of self-complementary sequences (each referred

to as a "clamp"). These clamps were introduced by design into the library so as to anchor

the substrate and enzyme regions of the molecule together with a four base pair clamp on

one side of the cleavage site, and a six base pair clamp on the other. The combined length

of these clamps increased to 15 base pairs following selection.










In the selection experiments reported here, a binding clamp of this length was not

introduced by design. This difference, which was not considered to be significant at the

outset, may have had consequences. The absence of extensive clamps may transfer

selective pressure onto the folding/association step, and away from the catalytic step.

Thus, selection within a library lacking clamps may not generate as strongly catalysts that

exploit cofactors as selection within a library having clamps. This may explain why the

catalysts that we examined are Mg++-independent, while the catalyst examined by

Breaker and Joyce is Mg++-dependent.

A curious parallel is found when comparing the Faulhammer/Famulok selection

experiment cited above, and the analogous experiment by Roth and Breaker. The former

failed to isolate histidine-dependent DNAzymes; the latter generated several histidine-

dependent DNAzymes. Two differences in their selection protocols appear relevant. The

Faulhammer/Famulok protocol included low amounts of MgCl2 (0.5 mM) in addition to

the histidine cofactor during selection; the Roth/Breaker selection did not. Further, the

Roth/Breaker libraries included engineered binding clamps, analogous to the clamps in

the Breaker-Joyce selection for ribonucleases. The Faulhammer/Famulok libraries did

not.

It may be coincidental that selections with clamped libraries in two cases generated

catalysts that exploited an external cofactor, while analogous selections with libraries

lacking the clamp generated catalysts that did not. These results suggest, however, that

the presence of base pairing between the substrate and enzyme portions of the nucleic

acid enzyme may be important to the experimental outcome.