JBC

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

Originally published In Press as doi:10.1074/jbc.M504537200 on July 21, 2005

J. Biol. Chem., Vol. 280, Issue 42, 35658-35669, October 21, 2005
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Supplemental Data
Right arrow All Versions of this Article:
280/42/35658    most recent
M504537200v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow reprints & permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Saladino, R.
Right arrow Articles by Di Mauro, E.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Saladino, R.
Right arrow Articles by Di Mauro, E.
Social Bookmarking
Add to CiteULike   Add to Complore   Add to Connotea   Add to Del.icio.us   Add to Digg   Add to Reddit   Add to Technorati  
What's this?

Origin of Informational Polymers

DIFFERENTIAL STABILITY OF 3'- AND 5'-PHOSPHOESTER BONDS IN DEOXY MONOMERS AND OLIGOMERS*{boxs}

Raffaele Saladino{ddagger}, Claudia Crestini§, Vincenzo Busiello¶, Fabiana Ciciriello||, Giovanna Costanzo¶, and Ernesto Di Mauro||1

From the {ddagger}Dipartimento AgroBiologia and AgroChimica, Università della Tuscia, Via San Camillo De Lellis, Viterbo 01100, Italy, §Dipartimento di Scienze e Tecnologie Chimiche, Università Tor Vergata, Rome 00133, Italy, Istituto di Biologia e Patologia Molecolari, Consiglio Nazionale delle Ricerche, Piazzale Aldo Moro, 5, Rome 00185, Italy, and ||Fondazione Istituto Pasteur-Fondazione Cenci-Bolognetti, Dipartimento di Genetica e Biologia Molecolare, Università La Sapienza di Roma, Piazzale Aldo Moro, 5, Rome 00185, Italy

Received for publication, April 26, 2005 , and in revised form, July 13, 2005.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 Methods
 RESULTS
 DISCUSSION
 REFERENCES
 
To survive, an informational macromolecule must solve the major problem set by its very polymeric nature: instability. This is especially true in prebiotic terms because of the presumed initial absence of protective structures (proteins, lipids, etc.). We have analyzed the stability of the {beta}-glycosidic and of the 3'- and 5'-phosphoester bonds in both deoxy monomers and deoxy oligomers under a large set of conditions. The results show a strong dependence of the relative stability of these bonds on the physico-chemical environment. A set of conditions has been identified in which the stability of polymers becomes comparable with that of the precursor monomers. In certain instances the stability of the 5'-phosphoester bond is even higher in the polymer than in the mononucleotide.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 Methods
 RESULTS
 DISCUSSION
 REFERENCES
 
If life on this planet had an early start as it seems (1-8), several conditions had to be satisfied at the same time: abundance of the starting biogenic materials, formation of precursors based on simple chemical processes, and simultaneous (or quasi so) presence of the building blocks to be used for the assembling of informational molecules. We have observed (Refs. 9-11 and references therein) that the one-carbon compound formamide is an highly versatile building block for the synthetic processes yielding all the necessary precursor nucleic bases. In terms of its possible role in allowing or favoring the formation of informational nucleic polymers, formamide chemistry also provides a possible solution to the lack of reactivity between ribose/deoxyribose with bases through a formose-like reaction yielding acyclonucleosides (10). The reported activity of formamide as activator of transphosphorylation (see "Discussion") provides additional possible relevance to its role in prebiotic reactions leading to pre-genetic informational polymers. The plausibility of its prebiotic role is related to (i) its presence in interstellar medium, in comets and asteroids (12), hence on early Earth; (ii) the ease of its formation by hydrolysis of HCN and its stability in liquid form over a wide range of temperatures (2.5-210 °C), thus favoring its concentration from dilute aqueous solutions; and (iii) the above mentioned catalyst-induced wealth of nucleic precursors.

The question arises as to whether the physico-chemical conditions (moderately high temperature and varying concentration of formamide in water) allowing active synthesis and potentially leading to polymerization favor or impair the survival of the polymerized materials. In reconstructing the passage from monomers to the information-bearing polymers that we know at present, two major pieces of the mosaic are markedly missing: the knowledge of the mechanisms leading to the formation of the {beta}-glycosidic and of the 3'-5' phosphodiester bonds and, once the polymer had been formed, the physico-chemical reason(s) leading to its very survival as a polymer. In other words, which is the Darwinian selective advantage that overcame the intrinsically higher instability of the polymeric form?

The analysis presented here focuses on the second question and is centered on two specific aspects: (i) we limit our analysis to DNA polymers and (ii) we explore the possibility that in formamide-containing solutions conditions exist in which the 3'- and the 5'-phosphoester bonds in the polymer are more stable than the corresponding ones in the monomer.

The stability of the phosphoester bonds of nucleic acids was analyzed in the 1960s and 1970s under various physico-chemical conditions. For comprehensive reviews of initial studies, see Chapter 10 in Ref. 13. These studies determined interesting differences, for instance, that the rate of cleavage of glycosidic bonds of free deoxynucleosides (14, 15) is 10-50 times higher relative to that in single-stranded DNA (16), that hydrolysis of glycosidic bonds in deoxynucleosides is >deoxynucleotides >DNA (17-19), and that the depurination is 4-fold in single-versus double-stranded DNA (rate constant single-stranded DNA = 4 x 10-9 sec-1, 70 °C, pH 7.4) (20). At apurinic sites in DNA, the deoxyribose residue occurs in equilibrium between the free aldehyde and the furanose form, leading to the cleavage of the phosphodiester bonds at both 5'- and 3'-positions (Ref. 21 and references therein).

However, a systematic and comprehensive comparison of the stability of the phosphoester bonds in the precursor monomers (both ribo and 2'-deoxyribo) versus the stability of the same bonds when present in DNA and RNA was not performed. A set of clear conclusions allowing a definitive picture of the relative stabilities of the phosphoester bonds in DNA and in RNA under various conditions is lacking. We have undertaken this analysis. We report here two first sets of data on DNA: we have defined (i) the stability of the 3'- and the 5'-phosphoester bonds in a model deoxyribo oligonucleotide as a function of temperature and of the water/formamide w/w ratios and (ii) the stability of the 3'- and the 5'-phosphoester bonds in monophosphate deoxynucleosides under a corresponding set of conditions. The comparison of these two sets of data provides information relevant to the monomer-versus-polymer stability bias.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
Methods
 RESULTS
 DISCUSSION
 REFERENCES
 
Materials
Adenine, 2'-deoxyribose, 2'-deoxyribose 5'-monophosphate, 2'-deoxyadenosine, 2'-deoxyadenosine 5'-monophosphate, and 2'-deoxyadenosine 3'-monophosphate were from Sigma, analytical grade.

Oligonucleotides—The degradation of oligonucleotides by formamide was studied on homogeneous polymers embedded in mixed sequence stretches. The overall approach consisted of the analysis of the degradation products of two synthetic 2'-deoxyoligonucleotides, each made of two short mixed sequence tails (10 and 6 bases, respectively) and of a central 30-base-long homogeneous stretch of As or Ts.

The oligonucleotides used were Oli1, 5'-ACCTAACCGG(A)30CCGGTT-3', and Oli2, 5'-CCCGAACCGG(T)30CCGGTT-3'. These oligonucleotides were designed such as to be complementary. Upon annealing, 4-nucleotide-long 5'-protruding tails remain at both extremities that can be used for selective labeling at 3'. In this study only the poly(dA) oligonucleotide (Oli1) was labeled and analyzed, as described (9).

For 3'-labeling, 2 µg of each oligonucleotide were annealed with the same amount of the complementary oligo and labeled with [{alpha}-32P]dCTP. Labeling was performed using T7 Sequenase (USBC; Amersham Biosciences). The labeled oligo was purified on a 16% denaturing acrylamide (19:1 acrylamide/bisacrylamide) gel, and the polyacrylamide was removed by a NuncTrap Probe purification column (Stratagene). 2 pmol (typically 30,000 cpm) of DNA were processed for each sample.

For 5'-labeling, 2 µg of poly(dA) oligonucleotide (Oli1) were labeled with [{gamma}-32P]ATP. Labeling was performed using polynucleotide kinase (Roche Applied Science), and the labeled oligo was purified and processed as above.


   Methods
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 Methods
RESULTS
 DISCUSSION
 REFERENCES
 
HPLC Analysis—Samples were resuspended at a final concentration of 1 mg/ml in water or in the appropriate formamide reaction mixtures (usually in 0.5-1.0 ml). Temperatures and incubation times are indicated. 10-µl aliquots of the reaction mixtures were diluted to a final concentration of 50% formamide in a final volume of 20 µl and injected into a SupelcosilTM LC-18-S 5-µm HPLC column (Supelco) 15 cm x 4.6 mm. Elution was performed at a flow rate of 2 ml/min at room temperature with methanol:30 mM ammonium phosphate, pH 5.3 (2.5:97.5), UV = 254 nm, pressure = 1.5 atm, on a HPLC Beckman System Gold instrument. Identification of the peaks was performed by comparison with real samples.

Oligonucleotide Degradation Protocols and Analyses—3'- and 5'-labeled oligonucleotides were treated under the time, temperature, and solution conditions indicated where appropriate. To stop the reaction a solution of 5 x 10-4 M (final concentration) of tetrasodium pyrophosphate (Sigma) dissolved in water was added to a final volume of 40 µl. The samples were vortexed for 1 min and then centrifuged at 13,000 rpm for 20 min. This procedure was performed twice. The wash was ethanol precipitated, resuspended in 5 µl of formamide buffer, heated for 2 min at 95 °C, and loaded on a 16% denaturing polyacrylamide gel (19:1 acrylamide/bisacrylamide). For these methods, see also Ref. 9.

Half-lives of the Bonds of the 3'- and 5'-Phosphoester Bonds in Oligonucleotides—For oligomers, the half-lives of the phosphodiester bonds were determined with standard procedure from plots similar to those reported in Figs. 2 and 3. Lines were drawn tangential to the initial part of the curves (i.e. Fig. 2), and the was graphically determined.

For the 3'-bonds, the half-life was calculated for the bonds located in three different positions from the label: the 7th, 20th, and 35th bonds, counting from the labeled extremity. We assumed that each bond in the poly(dA) stretch is cleaved with the same efficiency as the other bonds, and we have not taken into consideration poly(dA) internal nearest neighbor, sequence context, and cleavage-induced extremity effects, for which we have not observed any evidence. The homogeneity of cleavage in the poly(dA) stretch contrasts with the sequence dependence of formamide-induced cleavage rates. We have previously reported that the rate of cleavage in formamide is G ≥ A > C >> T (22, 23). The focus of the present analysis is the determination of the stability of the 3'- and 5'-phosphoester bonds related to adenine. Based on these considerations, we assumed that the actual half-life of, say, the 7th bond is given by the experimentally determined value (that is, the % value of the band under consideration relative to the total signal of the lane) multiplied by 7 and divided by 2; that of, say, the 20th bond by the observed value multiplied by 20 and divided by 2, etc. This procedure was adopted to average out experimental imprecision (deriving from gel electrophoresis, image scanning, background subtractions, tangential lane drawing, etc.) and to verify the linearity of the analytical range. When cleaving a terminally labeled oligomer, this type of kinetic analysis is meaningfully performed only in the less than one cut/molecule range (typically, cleaved molecules <30%). Above this % value, multiple hits prevent reliable analysis. This procedure provides a quantitative evaluation of the linearity of the analyzed range: under ideal conditions the of the 7th x 7 must correspond to that of the 20th x 20 and to that of the 35th x 35. To average out deviations from such correspondence, the value is calculated as the average of one central position and of two positions close to the two opposite extremities. The data are reported "as the sample average value" µ ± the "sample S.D. value" {sigma} of the values obtained by this procedure in the three different positions. These are computed as shown in Equation 1.


(Eq. 1)

The positions were selected as indicative of a centrally located bond (the 20th) and of two oppositely located bonds proximal to the extremities (the 7th to the 3'- and the 35th to the 5'-extremity, respectively).

For 5', the same procedure was followed, analyzing the positions 11th and 25th from the 5'-labeled extremity (Fig. 3). The smearing of the upper part of the gel, typical for 5'-labeled oligos (9), prevented analysis of more label-distal positions. For monomers, the values were graphically calculated from the data reported in Figs. 6, 7, 8.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 Methods
 RESULTS
DISCUSSION
 REFERENCES
 
The Half-life of the Phosphoester Bonds in a Deoxyribo Oligonucleotide
The 3'-Bond—The 3'-labeled 46-mer deoxyribo oligonucleotide shown in the lower part of Fig. 1 (see "Experimental Procedures") was treated at 90 °C for the indicated times in water (panel A) or in the presence of different amounts of formamide in aqueous solution (3, 10, 25, 33, 66, 100%). Panels B and C show two additional instances of the profiles obtained (25%, panel B; 100%, panel C). The treatments were always performed for two ranges of times, short (0-20 min) and long (20 min-12 h).



View larger version (51K):
[in this window]
[in a new window]
  		FIGURE 1.
Degradation of a 3'-labeled 46-base-long oligonucleotide containing a 30-bp-long dA stretch and mixed sequence extremities. The oligo was incubated at 90 °C for the time indicated on top of each lane in H2O(A), in 25% (B), or 100% formamide (C) and processed as described (see Oligonucleotide Degradation Protocols and Analyses under "Methods"). The a, b, and c letters point to the positions whose cleavage kinetics was determined, as detailed in Fig. 2.

 
The fast hydrolytic degradation of DNA and the regularity of the cleavage ladders are evident (panel A). Depurination is known to be faster than depyrimidination (11, 13). The degradations reported in the accompanying panels (B and C) performed in the presence of increasing amounts of formamide show that as a general trend formamide appreciably protects DNA from the hydrolytic process. The variations from this trend, namely at low formamide concentration and at high temperatures, are reported and discussed below.

Fig. 2 shows the quantitative evaluation of this type of degradation experiment. Panel A shows the cleavage kinetics at three different positions of the oligomer (namely, the 7th, 20th, and 35th positions) in H2O. Considering the number of 3'-phosphoester bonds in this 46-mer oligonucleotide (= 45) and the slower hydrolysis of the bonds proximal to the 9 pyrimidine residues (22, 23), each phosphoester bond would account upon complete cleavage for ~2.5% of the overall radioactive signal. Fig. 2A shows that this maximal value is rapidly reached for the 35th position (6.3 min); position 20 takes 14.7 min and position 7 48.4 min. The half-life is given by the projection on the abscissa of the intersection of the tangent with the ideal 1.25% line. In panel A the decrease of signal due to "multiple hits" becomes evident immediately after the maximum has been reached, as shown by the sloping down of curve a starting at 10 min and of curve b at 20 min. The multiple hit events being proportionally less frequent for the bond proximal to the labeled extremity (the 7th), no sloping effect is observed for curve c. The average of the 3'-phosphoester bond under these conditions (90 °C, H2O) is 142 ± 24 min. Panel B describes the effect of increasing formamide on the 3'-phosphoester cleavage reported above: enhancement by the low concentrations (i.e. 3%, see below) and protection by the higher ones.

The 5'-Bond—The same analysis was performed on the same oligo, labeled at the 5'-extremity. The resulting data are shown in Fig. 3. The average half-life of the 5'-phosphoester bond is 230 min ± 37 (90 °C, H2O). Formamide (panel B) has an overall protective effect.

Two Different Reaction Pathways
A detailed analysis was performed of the effect of temperature and formamide on the stability of the DNA chain. At the fixed time of 8 h, the kinetics of degradation was analyzed for the temperatures of 40, 50, 60, 70, 75, 80, and 90 °C at formamide concentrations of 0, 10, 25, 33, 66, and 100%. Fig. 4 shows two instances of this analysis on a 3'-labeled oligonucleotide. Panel A describes the increasing hydrolytic degradation as a function of increasing temperature in water; Panel B shows the effect of formamide on this degradation reaction. The data on panel B reveal that in the presence of formamide the degradation first increases as a function of the increasing temperature from 40 to 75 °C, then decreases at 80 °C (as shown by the arrow), and then increases again at 90 °C. This was taken as an indication of two different degradative reaction pathways, each dependent on a specific set of conditions (namely, dependent on the defined combination of temperature and water/formamide ratio). This observation was extended and confirmed by an extensive analysis of the effects of the physico-chemical variables (Fig. 5).

Fig. 5 shows a series of scanning profiles obtained from electrophoretic analyses similar to those shown in Fig. 4, presented as a function of increasing temperature (vertical) and increasing formamide % (horizontal). The first vertical row from the left (H2O, 0% formamide) describes the effect of increasing temperature in enhancing the rate of hydrolysis. The diagnostic feature inside each panel is the displacement of the degradation profile from the origin of the scanned lane on the left (the leftmost peak corresponding to the uncleaved sample) toward the bottom of the lane (the right side of the plot).

The results are interpreted according to the two following different degradation mechanisms. At temperatures ~100 °C or above and in the absence of water, formamide reacts with both purine and pyrimidine nucleobases by nucleophilic addition at position C (8) and position C (4) and C (6), respectively (Ref. 21 for purine, Ref. 22 for pyrimidines), causing the opening of the imidazole or pyrimidine rings and the ensuing cleavage of the {beta}-glycosidic bond. The {beta}-eliminations of both the 3'- and the 5'-phosphoester bonds and the consequent cleavage of the polymer follow (21, 22). The cleavage at 3' is faster than that at 5' (21-23). This reaction pathway can be referred to as Hydrolysis following Nucleobase Degradation (HND).2

In water the hydrolytic cleavages of the oligonucleotide occur following the direct cleavage of the {beta}-glycosidic bond (mainly leading to the removal of the non-degraded base). This mechanism can be referred to as Hydrolysis following Nucleobase Substitution (HNS). Both hydrolytic pathways are detailed under "Discussion" and in Fig. 11A, pathways A and B.



View larger version (19K):
[in this window]
[in a new window]
  		FIGURE 2.
The cleavage kinetics of the 46-bp-long 3'-labeled oligo shown in Fig. 1. Details in Half-lives of the Bonds under "Methods." A, the % cleavage of the analyzed bands (corresponding to the 7th, 20th and 35th 3'-phosphoester bonds from the label) is reported in ordinate and plotted as a function of the reaction time (abscissa). B, the % cleavage of the 7th bond (ordinate) as a function of time in various formamide concentrations (0, 3, 25, 100%, as indicated).

 



View larger version (30K):
[in this window]
[in a new window]
  		FIGURE 3.
As in Fig. 2, for the same oligo labeled at the 5'-extremity. The positions analyzed are the 11th and the 25th, as indicated. Filled symbols refer to a long-range kinetics (up to 4 h); open symbols refer to an experiment focusing on the initial part of the degradation reaction.

 
Inspection of the profiles (Fig. 5, three leftmost panels) of the samples treated at increasing concentrations of formamide revealed its increasingly protective effect against HNS hydrolysis, as evident in the upper horizontal rows of panels (40 and 50 °C, increasing formamide from left to right). The last vertical row on the right (100% formamide, increasing temperatures from top to bottom) shows patterns caused by {beta}-elimination cleavage due to the HND pathway. This latter reaction is markedly sequence-dependent as indicated in the "80 °C-100%" panel. The order of cleavage is G ≥A > C >> T (21-23).



View larger version (79K):
[in this window]
[in a new window]
  		FIGURE 4.
Degradation of the 3'-labeled poly(dA) as a function of the temperatures reported on top of each lane. 8-h treatment in H2O (A) or 33% formamide (B). The arrow (B) indicates the condition marking the passage between the two degradative reaction pathways (see Two Different Reaction Pathways under "Results").

 



View larger version (47K):
[in this window]
[in a new window]
  		FIGURE 5.
Scanning profiles of electrophoresis analyses similar to those shown in Fig. 4. The reactions were run at the formamide concentrations indicated on top and at the temperatures indicated on the left.

 
In conclusion, the left panels (both upper and bottom) show the products of hydrolysis following nucleobase substitution (HNS), whereas the panels on the right show the products of hydrolysis following nucleobase degradation (HND). HNS hydrolysis decreases from left to right and, in central panels, from top to bottom, as shown by the intermediate behavior observed at intermediate conditions, where the two mechanisms coexist. The chemical rational of these two reaction pathways is described under "Discussion."

The Stability of the {beta}-Glycosidic and 3'- and 5'-Phosphoester Bonds in Monophosphate Nucleosides
The {beta}-Glycosidic Bond—The stability of the {beta}-glycosidic bond in nucleosides in water was determined in pioneering studies under several conditions (24). We describe here the effect of formamide on the stability of this bond. Measurements were performed as a function of formamide concentration in water and of temperature. In addition to its interest per se, determination of the stability of this bond is a prerequisite for analysis of the stability of phosphoester bonds in nucleotides and in polymers thereof.

Fig. 6 shows the HPCL analysis of the degradation of deoxyadenosine (%, ordinates) as a function of time (abscissa) and of the concentration of formamide in the aqueous medium (from 0 to 100%, as indicated in each panel). The upper right panel side provides the interpretation key. The red letters indicate the starting compound, in this case deoxyribose (dR) bound to the nucleic base (B), in this case adenine. The cleavage (arrow) of the glycosidic bond (red) yields a compound (deoxyribose; black) that is not detected under our conditions and the base (B, green). The same color code is used in the plots of the following figures.

The kinetic disappearance of the deoxyadenosine (red line) is accompanied by the appearance of the base (green line). The two curves cross at 50% value (within minor experimental error). Thus, no other cleavage than that of the {beta}-glycosidic bond occurs in this range of conditions. The half-life of this bond at 90 °C in water is 8.5 x 103 min. Increasing concentrations of formamide first decrease and then (>25%) markedly increase the stability of this bond. Fig. 9A summarizes this double effect exerted by increasing formamide on the stability of the {beta}-glycosidic bond, observed both in the deoxyadenosine and in the deoxyadenosine monophosphate (these latter data are from Fig. 8).

The 3'-Phosphoester Bond—In addition, the degradation rates of the 3'-monophosphate form of deoxyadenosine were analyzed as a function of both temperature and formamide concentrations (Fig. 7). The disappearance of the 3'-dAMP (blue) in water (upper panel) is matched by the appearance of the nucleoside (red), thus being primarily caused by the cleavage of the 3'-phosphoester bond. The {beta}-glycosidic bond is cleaved with a slower kinetics, and the corresponding product (the base, green) only appears at a later time. Increasing formamide decreases the half-life of the 3'-phosphoester bond (see the 25% (middle) and the 100% (lower) panels) as shown by the faster sloping of the blue line in formamide, causing an earlier appearance of the nucleoside (red). Interestingly, the presence of the phosphate group in 3' stabilizes the {beta}-glycosidic bond, as shown by the plateauing of the red line in the 100% sample and by the delayed appearance of the base (see "Discussion").



View larger version (23K):
[in this window]
[in a new window]
  		FIGURE 6.
HPLC analysis of the degradation kinetics of deoxyadenosine as a function of the formamide concentration indicated in each panel. Red, deoxyadenosine; green, adenine. The deoxyribose is not detected in the HPLC analysis and is indicated in black in the scheme. Best fit curves were calculated with Microsoft Excel.

 



View larger version (15K):
[in this window]
[in a new window]
  		FIGURE 7.
HPLC analysis of the degradation kinetics of 3'-monophosphate deoxyadenosine. Color code as in Fig. 6. The whole 3'-dAMP molecule is indicated in blue.

 
The 5'-Phosphoester Bond—A detailed analysis of the numerous effects of formamide on the 5'-phosphoester bond of 5'-dAMP is shown in Fig. 8. Formamide strongly stimulates the disappearance of the 5'-dAMP (blue line) due to the cleavage of the 5'-phosphoester bond, resulting in the increasingly earlier formation of the nucleoside (red). An additional effect is the delayed appearance of the base (green) due to the slower cleavage of the {beta}-glycosidic bond. The fact that formamide strongly protects the {beta}-glycosidic bond is shown by the increasing distance between the red and green curves as a function of the increasing concentration of formamide.



View larger version (27K):
[in this window]
[in a new window]
  		FIGURE 8.
HPCL analyses of the degradation of 5'-monophosphate deoxyadenosine, as in Figs. 6 and 7.

 
The half-lives of the {beta}-glycosidic bond and of both the 3'- and 5'-phosphoester bonds as a function of increasing formamide are summarized in Fig. 9, A and B, respectively. The {beta}-glycosidic bond (Fig. 9A, red) is cleaved faster in the presence of low concentrations of formamide and then is markedly protected. This effect occurs for the {beta}-glycosidic bond in both adenosine and on its phosphate forms. The half-life values in Fig. 9 for the 5'-dAMP plot are taken from the degradations reported in Fig. 8. The overall trend is respected in both cases (as expected) with one important notation, the half-lives of the {beta}-glycosidic bond are constantly longer in the phosphate form. In water the effect is quite strong.

The 3'- and the 5'-phosphoester bonds (Fig. 9B) are cleaved faster in the presence of formamide, the effect being quite strong for the 5'-phosphoester bond, less so for the 3' one. The stability in water of the 5'-phosphoester bond is 10-fold higher ( min) than that at 3' ( min).

Summarizing the numerous effects exerted by water/formamide and temperature on the stability of the {beta}-glycosidic and phosphate bonds in the nucleoside and in its 3'- and 5'-phosphate forms at 90 °C, we observed for {beta}-glycosidic (i) intermediate stability in water (~ 10·103 min) for the nucleoside, (ii) enhanced cleavage by low formamide/water ratios (<33%) versus protection by higher ones (>33%) (Fig. 9A), and (iii) enhanced stability exerted by the presence of a phosphate group, especially in water (Fig. 9A). For the phosphoester we observed (i) a marked difference in the stability of water between the 5'- and the 3'-phosphoester bonds: high (>25·103 min) for the 5', an order of magnitude lower (~2.5·103 min) for the 3', and (ii) destabilization of both 5'- and 3'-phosphoester bonds by formamide (Fig. 9B).

The Differential Effects of Formamide on the Stability of the 3'- and the 5'-Phosphoester Bonds in Monomers and Polymers
The data reported above on the stability of the {beta}-glycosidic and phosphoester bonds in the monomers and polymers reveal strong and differential sensitivity. These bonds are highly sensitive to variations of the environmental conditions. The chemical rationales presented under "Discussion" provide an explanation for a large part of these differential stabilities based on state-of-the-art chemical knowledge and on previous well established observations.

Independent of the detailed chemical mechanisms involved, the plot of the half-life values of the 5'-phosphoester bonds in the monomer 5'-dAMP and in the oligomer as a function of formamide concentration (Fig. 10A) reveals a relevant property: formamide decreases the half-life of the 5'-phosphoester bond in the monomer and increases that of the same bond in the polymer. The effect is quite strong: from a of 28.5·103 min for the dAMP and of 0.23·103 for the polymer (ratio = 125), the situation is reversed starting at ~75% formamide. In formamide (100%) the same bond becomes more stable in DNA than in the precursor monomer. For the 3'-phosphoester bond the phenomenon and the trend are similar, although somewhat less marked: 2.55·103 versus 0.14·103 (ratio = 18.3) (Fig. 10B).


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 Methods
 RESULTS
 DISCUSSION
REFERENCES
 
We have compared the stabilities of the phosphoester bonds in water and found them to be quite different in the monomers relative to those in the DNA polymer. The difference is ~two orders of magnitude for the 5'-bond (Fig. 10A) and more than one order of magnitude for the 3'-bond (Fig. 10B). Given similar environmental conditions, the same 5'- and 3'-bonds would dissolve, respectively, 125 and 18.3 times faster in DNA than in its precursor monomer. In this physico-chemical frame, polymerization would obviously be forbidden. Formamide reverses these stability parameters, potentially allowing the survival of the polymeric form.

The Known Mechanisms of the Phosphoester Bond Cleavage—Different mechanisms for the cleavage of the phosphodiester bond for DNA and RNA have been reported in water. Most reactions leading to cleavage of the phosphodiester bond in RNA result from the direct rupture of the P-O bond by nucleophilic addition/elimination at the phosphorus atom. In the case of DNA the reaction requires a multistep process starting from the removal of the nucleobases. Under alkaline conditions both HND and HNS pathways are effective processes due to the high reactivity of hydroxide ions (Fig. 11A, pathways A and B) (25-28). Irrespective of the mechanism of removal of the nucleobase, the cleavage of the phosphodiester bond at C(3')-O and C(5')-O positions of the DNA backbone proceeds further by ring opening of the 2'-deoxyribose moiety and two successive {beta}-eliminations (Fig. 10A) (13). A quantitative evaluation is still lacking of the contribution of the HND and HNS pathways on the rate of hydrolysis of DNA in water. At high temperature (110 °C), neat formamide reacts with DNA through the nucleobase degradation pathway in a very selective manner. On the contrary, at lower temperature it shows a limited reactivity interacting strongly with DNA by noncovalent bonds (29, 30). These interactions are mainly due to selective molecular recognition processes with both purine and pyrimidine nucleobases involving the formation of multiple hydrogen bonds (31, 32). Moreover, formamide forms with water molecules stable 1:1 complexes in which a cyclic double hydrogen-bonded structure has been observed (33). In formamide/water mixtures and at low concentration of formamide (3-10% w/w), the rate of hydrolysis of 2'-deoxy polynucleotides increases because of the effect of a general alkaline catalysis. The protective effect exerted by formamide at concentrations higher than 25% and temperatures lower than 70 °C is probably due to the formation of an extensive and highly organized hydrogen bond network shielding the removal of nucleobases by reaction of hydroxide ions. At high concentration of formamide and at temperatures higher than 70 °C the base-selective degradation of nucleobases by formamide becomes the operatively predominant process (21-23). Thus, the transition between the two different types of hydrolytic pathways shown in titration experiments in Figs. 4 and 5 (HND plus HND with hydroxide ions versus HNS with formamide) presumably reflects the higher selectivity of formamide relative to hydroxide ions on the degradation of purine and pyrimidine nucleobases.



View larger version (14K):
[in this window]
[in a new window]
  		FIGURE 9.
The stability of the {beta}-glycosidic (A) and the 3'- and 5' phosphoester (B) bonds as a function of formamide concentration. A, the half-lives (min·103) of deoxyadenosine (dAdenosine) and of 5'-dAMP are calculated from the experiments reported in Figs. 6 and 8, respectively. B, half-lives are shown for the 3'- and 5'-phosphoester bonds. Data are from Figs. 7 and 8, respectively.

 



View larger version (15K):
[in this window]
[in a new window]
  		FIGURE 10.
Summary comparison of the stability of the phosphoester bonds in the polymer and in the monomer. A, 5'-phosphoester bonds. B, 3'-phosphoester bonds. The values of the phosphoester bonds of the 5'-dAMP (A) and of the 3'-dAMP (B) are those already shown in Fig. 9B. The values of the same bonds when embedded in the oligonucleotide are taken from digestion experiments similar to those reported in Fig. 1 (for the 3'-phosphoester bond) and from similar experiments (not shown) for the 5'-phosphoester bond.

 



View larger version (27K):
[in this window]
[in a new window]
  		FIGURE 11.
A, mechanism of DNA degradation. Pathway A, formamide or hydroxide nucleophilic addition on the C-8 position of the adenine ring followed by nucleobase degradation/elimination and two {beta}-elimination reactions with C(3')-O and C(5')-O bond cleavages. Pathway B, direct hydroxide nucleophilic substitution of adenine followed by two {beta}-elimination reactions with C(3')-O and C(5')-O bond cleavages. B, equation 1, mechanism of degradation of 2'-deoxyadenosine. Equation 2, mechanism of degradation of 2'-deoxyadenosine 5'-monophosphate.

 
The protective effect of formamide on the nucleobase removal was further confirmed on 2'-deoxyadenosine. In this case, the quantitative recovery of the free base adenine in the reaction mixture suggests that the HNS pathway was the main hydrolytic process in water and in weakly alkaline solutions (Fig. 11B, equation 1). As reported in Fig. 6 and summarized in Fig. 9, the rate of hydrolysis of the {beta}-glycosidic bond increases with increasing formamide concentration, reaching its highest value at 25% w/w formamide. Again, formamide concentrations higher than 33% w/w depress the hydrolysis, probably as a consequence of the shielding effect of the newly formed formamide hydrogen-bonding network against the hydroxide ions approaching the nucleobase.

In the case of adenine nucleotides, the cleavage of two reactive bonds, the phosphoester and the {beta}-glycosidic bonds, should be considered at the same time. In all the experimental conditions analyzed, cleavage of the phosphoester bond increases with increasing formamide concentration, reaching its highest value at 100% w/w formamide (Fig. 11B, equation 2). The hydrolysis of the {beta}-glycosidic bond is on the contrary markedly inhibited in increasing formamide as revealed by the presence of 2'-deoxyadenosine even for long reaction times (see for example the "100%" panel in Fig. 7). It is noteworthy that the stabilization of the {beta}-glycosidic bond was more efficient in the nucleotide relative to the nucleoside, suggesting a plausible participation of the phosphate moiety to the hypothesized hydrogen bond network.

Accordingly, direct and water-mediated hydrogen bonds between the polypeptide amide backbone and the phosphate of DNA play a significant role in the formation and stabilization of protein-DNA complexes (34-36). Because of the high directionality of the hydrogen bond, the position of the phosphate moiety on the nucleotide (5'-phosphate versus 3'-phosphate) is also a relevant stereo electronic parameter to be considered, 2'-deoxyribo-nucleoside 5'-monophosphate being the most stable derivatives (Fig. 9B).

Why Formamide?—Why should formamide be relevant in any scenario leading to the formation and survival of genetically relevant polymers?

First, its reactions at temperatures between 90 and 160 °C result in the production in acceptable yields of essentially all the nucleic base precursors (9-11, 37). We have observed the synthesis of adenine, purine, hypoxanthine, N9-formylpurine and other acyclonucleosides, N,N-diformyladenine, cytosine, hydroxypyrimidine, pyrimidinone, uracil, dihydrouracil, hydroxymethyluracil, thymine, AICA (5-aminoimida-zole-4-carboxamide), and fAICA (5-formamidoimidazole-4-carboxamide). Interestingly, the absence of guanine is compensated in this otherwise complete set of precursor bases by AICA and fAICA, known intermediates in the late steps of the biosynthesis of inosine monophosphate (Ref. 9 and references therein).

Second, acyclonucleosides may be of great relevance in the prebiotic synthesis of nucleoside derivatives because of their masked glycosidic bond. The formation of the glycosidic bond between nucleobases and sugars is a most difficult synthetic step; demonstration of this condensation under prebiotic experimental conditions has met difficulty (38, 39). The presence of the glycosidic bond in acyclonucleosides provides an interesting alternative: it shows that sugars have the chemical possibility of growing onto the nucleic bases (10) following a "formose reaction" pathway (40, 41), thus avoiding the difficult joining of preformed bases and sugars.

Third, the other relevant step in which formamide might have played an important role is the formation of phosphoester bonds. The question "where do the phosphates of nucleic acids come from?" cannot be answered until the chemistry of the formation of the phosphoester bond has been understood, possibly based on easily available inorganic phosphates. Evidence that formamide does favor phosphoester bond formation in abiotic conditions has been reported (42). Formamide also allows full solubility of the otherwise weakly soluble deoxyadenosine (43).

The synthesis products listed above are obtained in rich mixtures whose variety and quantitative composition depend on the catalyst present. Remarkably, the catalysts that were found to be active are among the most common components of the Earth's crust: clay (9), CaCO3, zeolites, silica, alumina, kaolin (37), titanium oxides (10), and olivines (11). The broad availability of the appropriate catalysts concurs with the required robustness of early abiotic polymerization events.

In conclusion, formamide chemistry provides on the one hand the frame for the production of a complete panel of precursor bases. On the other, it offers the testable possibility of closing the two gaps still open in understanding the plausible routes leading to pre-genetic polymers, the formation of the {beta}-glycosidic and phosphodiester bonds.

Why Study Stability?—It has been pointed out repeatedly and correctly that stability of the precursor molecules is a major concern in understanding the origin of informational polymers (44-46, 57). This consideration has supported the hypothesis of a cold origin of life, and a series of long-term experiments showed that purines can be obtained in decades of time in frozen solutions made of NH4CN or of the products obtained by electric discharge of a CH4, N2, NH3, and H2O mixture, among the most important components of the primitive Earth atmosphere (47, 48).

The rationale of this approach is that the degradation rates of precursor nucleic bases are relatively rapid (44-46) and that conditions should be found that might favor synthesis over degradation. The very fact of obtaining the bases after years in the frozen state indicates that those products accumulate and that synthesis is favored over degradation.

We observed the synthesis of the complete set of nucleobases at rather high temperatures, between 90 and 160 °C (9-11, 37). Therefore, to allow the possibility of polymerization and the survival of the products, two alternatives exists: either the synthesized molecules are quickly removed from the hot reactive site where synthesis has taken place and are transferred to a cold surrounding environment, as hypothesized in the "vents" system (49, 60), or conditions and reactions exist allowing the polymerization process at the same high temperatures.

The two scenarios are obviously not mutually exclusive. A general consensus is forming concerning the conditions of the early Earth upon which life appeared, as recently critically reviewed (50-53). The planet on which life first flourished was hot (51), was subject to impacts from extraterrestrial bodies, and was volcanically and hydrothermally active, anoxic, and highly UV bathed. In particular, temperatures on the early Earth were probably higher than they are today, both in the mantle and at the surface (51, 54). The average ocean temperature is estimated to have been >50 °C, possibly as high as 70-80 °C (50, 51, 54). Any form of life arising at this temperature was supposedly thermophilic or hyperthermophilic (55-56). The thermophilic roots of the evolutionary tree have been established (Ref. 58 and references therein).

The very fact that formamide yields the full panel of nucleobases shows the possibility of their accumulation in heat. In terms of the ability to evolve informational macromolecules, the intrinsic instability of the bases (44-46) and their prompt synthesis (Refs. 9-11, 37, reviewed in Ref. 59) does not constitute a disadvantage. To the contrary, the synthesis/degradation/resynthesis cycle offers the possibility of forming a dynamically equilibrating pool of precursors whose composition depends on the synthesis/degradation rate of each molecular species and on the catalysts present (59). The advantage provided by a flexible and adjustable pool of precursors is an important evolutionary property. This "equilibration-of-the-pool" hypothesis is presented in Ref. 59 and is discussed in detail in Ref. 61.

The way out from a futile cycle of syntheses/degradations of precursor nucleobases may be provided by polymerization itself. For this reason the study of the stability of the important bonds in the polymer relative to the stability of the same bonds in the monomers is the key point in understanding the basics of the thermodynamics and the kinetics of polymerization. It is probably quite relevant that formamide itself (whose functions in providing precursor nucleobases, acyclonucleosides, and in stimulating phosphoesterification are mentioned above) also provides the key to stabilizing bonds into polymers.

It has been pointed out that informational polymers are not limited to DNA and that other possibilities exist (38), as peptide nucleic acid (62) or threose nucleic acid (63). The central role of RNA as ur-molecule was hypothesized based on the intrinsic ability of RNA to act as a catalyst (64-68). This has stimulated the search for comparable properties in DNA leading to the discovery of DNA enzymatic activity (69-72).

It has been pointed out that in RNA the {beta}-glycosidic bonds are more stable than in DNA, whereas the contrary occurs for the 5'-phosphoester bond (Refs. 38 and references therein). This means, in principle, that in terms of keeping and/or evolving genetic information DNA could be a better suited molecule for both a depository function (more stable backbone) and for evolving information (more dynamically allowing changes of the base sequence). Thus, knowing the relative stabilities of the relevant bonds in DNA versus RNA is instrumental in drawing a more detailed scenario for the origin of informational macromolecules or at least for the passage from the RNA to the DNA worlds (73).

Quantitative analyses of the evolutionary effect of these differential stabilities are lacking. For the sake of comparison and deeper insight, measurement of bond stabilities in RNA and RNA precursors is in progress in our laboratories.

In conclusion, a limited but well defined ensemble of conditions was identified in which the 5'-phosphoester bond is more stable in DNA than in dAMP: >75% formamide, 90 °C. The polymer, or even a short oligomer, would not survive for long in these conditions because of its repetitive nature. However, our measurements and findings provide the basic assay for the evaluation of the conditions, of the catalysts, of the shielding surfaces, and of the physico-chemical environment that could sufficiently further enhance the stability of the polymers relative to that of the monomers enough to allow their survival, replication, and evolution.


   FOOTNOTES
 
* This work was supported by the Italian Space Agency, Genomica Funzionale Consiglio Nazionale delle Ricerche, Centro di Eccellenza di Biologia e Medicina Molecolare, Fondo per gli Investimenti della Ricerca di Base, and by Ministero Instruzione Università e Ricerca Cofinanziamento 2003 "La catalisi dei metalli di transizione nello sviluppo di strategie sintetiche innovative di eterocicli". The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

{boxs} The on-line version of this article (available at http://www.jbc.org) contains supplemental Fig. S1. Back

1 To whom correspondence should be addressed. Tel.: 39-06-49912880; Fax: 39-06-49912500; E-mail: Ernesto.dimauro{at}uniroma1.it.

2 The abbreviations used are: HND, hydrolysis following nucleobase degradation; HNS, hydrolysis following nucleobase substitution; HPLC, high pressure liquid chromatography. Back



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 Methods
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Dunlop, J. S., Muir, M. D., Milne, V. A., and Groves, D. I. (1978) Nature 274, 676-678[CrossRef][Medline] [Order article via Infotrieve]
  2. Walsh, M. M. (1992) Precambrian Res. 54, 271-293
  3. Lazcano, A., and Miller, S. L. (1994) J. Mol. Evol. 29, 546-554[CrossRef]
  4. Mojzsis, S. J., Arrhenius, G., McKeegan, K. D., Harrison, T. M., Nutman, A. P., and Friend, C. R. (1996) Nature 384, 55-59[CrossRef][Medline] [Order article via Infotrieve]
  5. Hofmann, H. J., Grey, Hickman, A. H., and Thorpe, R. I. (1999) Geol. Soc. Am. Bull. 111, 1256-1262[Abstract/Free Full Text]
  6. Westall, F., De Wit, M. J., Dann, J., Van Der Gaast, S., De Ronde, C., and Gerneke, D. (2001) Precambrian Res. 106, 93-116
  7. van Zuilen, M. A., Lepland, A., and Arrhenius, G. (2002) Nature 418, 627-630[CrossRef][Medline] [Order article via Infotrieve]
  8. Westall, F., Hofmann, B., and Brack, B. (2004) in Proceedings of the Third European Workshop on Exo/Astrobiology, Madrid, Spain November 18-20, 2003 (Vazquez, L., Brack, A., Amils, R., Cockell, C., Fernandez Remolar, D., Colangeli, L., Gomez-Elvira, J., Ehrenfreund, P., Lazaro, E., Hofmann, B., Martinez Frias, J., Penny, A., Perez-Mercader, J., Rodriguez-Manfredi, J. A., and Selsis, F., eds) pp. 37-40, ESA Special Publication 545, European Space Agency Publication Division, Noordwijk, The Netherlands
  9. Saladino, R., Crestini, C., Ciambecchini, U., Ciciriello, F. Costanzo, G., and Di Mauro, E. (2004) ChemBioChem. 5, 1558-1566[CrossRef][Medline] [Order article via Infotrieve]
  10. Saladino, R., Ciambecchini, U., Crestini, C., Costanzo, G., Negri R., and Di Mauro, E. (2003) ChemBioChem. 4, 514-521[CrossRef][Medline] [Order article via Infotrieve]
  11. Saladino, R., Crestini, C., Neri, V., Brucato, J. R., Colangeli, L., Ciciriello, F., Di Mauro, E., and Costanzo, G. (2005) ChemBioChem. 6, 1368-1374[CrossRef][Medline] [Order article via Infotrieve]
  12. Millar, T. J. (2004) in Astrobiology: Future Perspectives (Ehrenfreund, P., Irvine, W., Owen, T., Becker, L., Blank, J., Brucato, J., Colangeli, L., Derenne, S., Dutrey, A., Despois, D., Lazcano, A., and Robert, F., eds) pp. 17-31, Kluwer Academic Publishers, Boston
  13. Kochetov, N. K., Budovskii, E. I., Sverdlov, E. D., Simukova, N. A., Turchinskii, M. F., and Shibaev, V. N. (1972) in Organic Chemistry of Nucleic Acids (Kochetov, N. K., and Budovskii, E. I., eds) pp. 477-532, Plenum Press, London and New York
  14. Shapiro, R., and Kang, S. (1969) Biochemistry 8, 1806-1810[CrossRef][Medline] [Order article via Infotrieve]
  15. Garrett, E. R., and Mehta, P. J. (1972) J. Am. Chem. Soc. 94, 8542-8547[Medline] [Order article via Infotrieve]
  16. Lindahl, T., and Karlstrom, O. (1973) Biochemistry 12, 5151-5154[CrossRef][Medline] [Order article via Infotrieve]
  17. Shapiro, H. S., and Chargaff, E. (1957) Biochim. Biophys. Acta, 26, 596-608[Medline] [Order article via Infotrieve]
  18. Venner, H. (1966) Hoppe Seyler's Z. Physiol. Chem. 344, 189-196[Medline] [Order article via Infotrieve]
  19. Shapiro, R., and Danzig, M. (1973) Biochim. Biophys. Acta, 319, 5-10[Medline] [Order article via Infotrieve]
  20. Lindahl, T., and Nyberg, B. (1972) Biochemistry 11, 3610-3618[CrossRef][Medline] [Order article via Infotrieve]
  21. Saladino, R., Mincione, E., Crestini, C., Negri, R., Di Mauro, E., and Costanzo, G. (1996) J. Am. Chem. Soc. 118, 5615-5619[CrossRef]
  22. Saladino, R., Crestini, C., Mincione, E., Costanzo, G., Di Mauro, E., and Negri, R. (1997) Bioorg. Med. Chem. 5, 2041-2048[Medline] [Order article via Infotrieve]
  23. Negri, R., Costanzo, G., Saladino, R., and Di Mauro, E. (1996)) BioTechniques 21, 910-917[Medline] [Order article via Infotrieve]
  24. Kochetov, N. K., Budovskii, E. I., Sverdlov, E. D., Simukova, N. A., Turchinskii, M. F., and Shibaev, V. N. (1972) in Organic Chemistry of Nucleic Acids (Kochetov, N. K., and Budovskii, E. I. eds) pp. 425-448, Plenum Press, London and New York
  25. Jones, A. S., Mian, A. M., and Walker, R. T. (1968) J. Chem. Soc. 2042-2044
  26. Townsend, L. B, and Robins, R. K. (1963)) J. Am. Chem. Soc. 85, 242-243
  27. Zakharyan, R. A. Venkstern, T. V., and Baev, A. A. (1967) Biokhimiya 32, 1068-1078
  28. Lawley, P. D., and Brookes, P. (1964) Biochem. J. 92, 19C-20C[Medline] [Order article via Infotrieve]
  29. Bhattacharyya, A. J., and Feingold, M. (2001) Talanta 55, 943-949[CrossRef]
  30. Cornish, E. C., Beckham, S. A, and Maddox, J. F. (1998) BioTechniques 25, 948-954[Medline] [Order article via Infotrieve]
  31. Rozas, I., Alkorta, J., and Elguero, J. (2004) J. Phys. Chem. B 108, 3335-3341[CrossRef]
  32. Sponer, J., and Hobza, P. (2000) J. Phys. Chem. A 104, 4592-4597[CrossRef]
  33. Fu, A., Du, D., and Zhou, Z. (2003) THEOCHEM 623, 315-325[CrossRef]
  34. Harrison, S. C. (1991) Nature 353, 715-719[CrossRef][Medline] [Order article via Infotrieve]
  35. Pabo, C. O., and Sauer, R. T. (1992) Annu. Rev. Biochem. 61, 1053-1095[CrossRef][Medline] [Order article via Infotrieve]
  36. Travers, A. A. (1993) DNA-Protein Interactions, pp. 28-51, Chapman and Hall, London
  37. Saladino, R., Crestini, C., Costanzo, G., Negri, R., and Di Mauro, E. (2001) Bioorg. Med. Chem. 9, 1249-1253[CrossRef][Medline] [Order article via Infotrieve]
  38. Orgel, L. E. (1998) Trends Biochem. Sci. 23, 491-495[CrossRef][Medline] [Order article via Infotrieve]
  39. Pitsch, S., Eschenmoser, A., Gedulin, B., Hui, S., and Arrhenius, G. (1995) Orig. Life Evol. Biosph. 25, 297-334[CrossRef][Medline] [Order article via Infotrieve]
  40. Butlerow, A. (1861) Liebig's Ann. Chem. 120, 295-298
  41. Decker, P., Schweer, H., and Pohlmann, R. (1982) J. Chromatogr. 244, 281-291[CrossRef]
  42. Schoffstall, A. M, Barto, R. J., and Ramos, D. L. (1982) Orig. Life 12, 143-151[CrossRef][Medline] [Order article via Infotrieve]
  43. Schoffstall, A. M. (1976) Orig. Life 7, 399-412[CrossRef][Medline] [Order article via Infotrieve]
  44. Levy, M., and Miller, S. C. (1998) Proc. Natl. Acad. Sci. U. S. A. 95, 7933-7938[Abstract/Free Full Text]
  45. Bada, J. L., and Lazcano, A. (2002) Science 296, 1982-1983[Abstract/Free Full Text]
  46. Miller, S. L., and Lazcano, A. (1995) J. Mol. Evol. 41, 689-692[CrossRef][Medline] [Order article via Infotrieve]
  47. Levy, M., Miller, S. L., Brinton, K., and Bada, J. L. (2000) Icarus 145, 609-613[CrossRef][Medline] [Order article via Infotrieve]
  48. Miyakawa, S., Cleaves, H. J., and Miller, S. L. (2002) Orig. Life Evol. Biosph. 32, 209-218[CrossRef][Medline] [Order article via Infotrieve]
  49. Imai, E.-I., Honda, H., Hatori, K., Brack, A., and Matsuno, K. (1999) Science 283, 831-833[Abstract/Free Full Text]
  50. Westall, F. (2004) in Astrobiology: Future Perspectives (Ehrenfreund, P., Irvine, W., Owen, T., Becker, L., Blank, J., Brucato, J., Colangeli, L., Derenne, S., Dutrey, A., Despois, D., Lazcano, A., and Robert, F., eds) pp. 287-316, Kluwer Academic Publishers, Boston
  51. Knauth, L. P., and Lowe, D. R. (2003) Geol. Soc. Am. Bull. 115, 566-580[Abstract/Free Full Text]
  52. Lowe, D. R., Byerly, G. R., Kyte, F. T., Shukulyukov, A., Asaro, F., and Krull, A. (2003) Astrobiology 3, 7-48[CrossRef][Medline] [Order article via Infotrieve]
  53. Kyte, F. T., Shukolyukov, A., Lugmaor, G. W., Lowe, D. R., and Byerly, G. R. (2003) Geology 31, 283-286[Abstract/Free Full Text]
  54. Arndt, N. T. (1994) in Archean Crustal Evolution (Condie, K. C., ed) pp. 11-44, Elsevier, Amsterdam
  55. Wiegel, J., and Adams, M. W. (1998) in Thermophiles: The Keys to Molecular Evolution and the Origin of Life? (Adams, M. W. W., and Wiegel, J., eds) Taylor and Francis, Inc., Philadelphia, PA
  56. Forterre, P., Confalonieri, F., Charbonnier, F., and Duguet, M. (1995) Orig. Life Evol. Biosph. 25, 235-249[CrossRef][Medline]