Advertisement

Generation of Long RNA Chains in Water*

  • Giovanna Costanzo
    Footnotes
    Affiliations
    Istituto di Biologia e Patologia Molecolari, CNR, Università La Sapienza di Roma, P. le Aldo Moro, 5, Rome 00185, Italy
    Search for articles by this author
  • Samanta Pino
    Footnotes
    Affiliations
    Dipartimento di Genetica e Biologia Molecolare, Università La Sapienza di Roma, Università La Sapienza di Roma, P. le Aldo Moro, 5, Rome 00185, Italy
    Search for articles by this author
  • Fabiana Ciciriello
    Affiliations
    Dipartimento di Genetica e Biologia Molecolare, Università La Sapienza di Roma, Università La Sapienza di Roma, P. le Aldo Moro, 5, Rome 00185, Italy
    Search for articles by this author
  • Ernesto Di Mauro
    Correspondence
    To whom correspondence should be addressed. Tel.: 39-06-4991-2880; Fax: 39-06-4991-2500
    Affiliations
    Fondazione Istituto Pasteur-Fondazione Cenci-Bolognetti, c/o Dipartimento di Genetica e Biologia Molecolare, Università La Sapienza di Roma, P. le Aldo Moro, 5, Rome 00185, Italy
    Search for articles by this author
  • Author Footnotes
    * This work was supported by the Italian Space Agency, the MoMa project, and Agenzia Spaziale Italiana-Istituto Nazionale di Astrofisica (Grant I/015/07/0, Esplorazione del Sistema Solare) Italy.
    1 Both authors contributed equally to this work.
Open AccessPublished:October 02, 2009DOI:https://doi.org/10.1074/jbc.M109.041905
      The synthesis of RNA chains from 3′,5′-cAMP and 3′,5′-cGMP was observed. The RNA chains formed in water, at moderate temperatures (40–90 °C), in the absence of enzymes or inorganic catalysts. As determined by RNase analyses, the bonds formed were canonical 3′,5′-phosphodiester bonds. The polymerizations are based on two reactions not previously described: 1) oligomerization of 3′, 5′-cGMP to ∼25-nucleotide-long RNA molecules, and of 3′,5′-cAMP to 4- to 8-nucleotide-long molecules. Oligonucleotide A molecules were further extended by reciprocal terminal ligation to yield RNA molecules up to >120 nucleotides long and 2) chain extension by terminal ligation of newly polymerized products of 3′,5′-cGMP on preformed oligonucleotides. The enzyme- and template-independent synthesis of long oligomers in water from prebiotically affordable precursors approaches the concept of spontaneous generation of (pre)genetic information.

      INTRODUCTION

      The origin of informational polymers is not understood. The RNA polymerization process has been studied for five decades, the results showing that from preactivated precursors polymers of several tens can be obtained, as reviewed previously (
      • Orgel L.E.
      ). These pioneering studies provide the proof-of-principle that RNA precursors can self-assemble yielding linear polymers. However, the prebiotic validity of a process based on complex preactivation procedures is limited (
      • Orgel L.E.
      ,
      • Orgel L.E.
      ), and the problem of defining a prebiotically plausible chemical and thermodynamic scenario for the synthesis and accumulation of informational polymers remains open. The core of the problem is the standard state Gibbs free energy change (
      • van Holde K.
      The Origins of Life and Evolution.
      ,
      • Alberty R.A.
      ) stating that condensation reactions are very inefficient in water. Given that extant polymerizations occur in water, this is a major difficulty, only partially solved by the fact that these processes at present occur inside the active site of enzymes where water activity may be drastically reduced. The other part of the extant solution, fruit of evolution, is the use of biologically highly preactivated triphosphate nucleotides (
      • van Holde K.
      The Origins of Life and Evolution.
      ). In primordia, RNA molecules had no enzymes to catalyze their chain-wise growth, and highly activated precursors can be considered as prebiotic only with difficulty.
      We reasoned that for a pre-enzymatic polymerization to occur the solution must have relied on a simple and robust process. Ideally, such a process should have been based on compounds that were reactive yet relatively stable, chemically not too elaborate to allow their efficient production, and not too dissimilar from the products of their polymerization to minimize the chemical cost of the process.
      It was observed that phosphorylation of nucleosides occurs in formamide simply in the presence of a source of organic or inorganic phosphate at temperatures at which both the reactants and the products are stable (
      • Costanzo G.
      • Saladino R.
      • Crestini C.
      • Ciciriello F.
      • Di Mauro E.
      ). Phosphorylation occurs in every possible position of the nucleoside sugar moiety resulting, both for purine and pyrimidine nucleosides, in the production of 2′-, 3′-, 5′-, 2′,3′-cyclic, and 3′,5′-cyclic XMPs
      The abbreviation used is: XMP
      any one of AMP, GMP, CMP, or UMP.
      (
      • Costanzo G.
      • Saladino R.
      • Crestini C.
      • Ciciriello F.
      • Di Mauro E.
      ). The phosphorylation reaction is faster for the open than for the cyclic forms, whereas higher stability of the cyclic forms at higher temperature favors their accumulation.
      Coupled with the facile synthesis of all the nucleic bases from formamide (
      • Saladino R.
      • Crestini C.
      • Ciciriello F.
      • Costanzo G.
      • Di Mauro E.
      ) and with the formation of acyclonucleosides by TiO2-catalyzed formamide photochemistry (
      • Saladino R.
      • Ciambecchini U.
      • Crestini C.
      • Costanzo G.
      • Negri R.
      • Di Mauro E.
      ), the nonenzymatic phosphorylation of nucleosides (
      • Costanzo G.
      • Saladino R.
      • Crestini C.
      • Ciciriello F.
      • Di Mauro E.
      ) shows that the formation of cyclic monophosphate nucleosides is chemically simple and prebiotically plausible. The formation of both 2′,3′- and 3′,5′-cyclic XMPs in water starting from nucleosides and an inorganic source was also observed (
      • Saladino R.
      • Crestini C.
      • Ciciriello F.
      • Pino S.
      • Costanzo G.
      • Di Mauro E.
      ).
      The unsophisticated chemistry required for the formation of both open and cyclic nucleotides prompted us to investigate the possibility of their spontaneous polymerization. If so, nonenzymatic (pre)genetic polymerization could have taken place in warm little pond conditions, close to those imagined by Darwin (
      • Darwin F.
      ).

      EXPERIMENTAL PROCEDURES

      Materials

      Adenine, adenosine, adenosine 2′-monophosphate (2′-AMP), adenosine 3′-monophosphate (3′-AMP), adenosine 5′-monophosphate (5′-AMP), adenosine 2′,3′-cyclic monophosphate (2′,3′-cAMP), adenosine 3′,5′-cyclic monophosphate (3′,5′-cAMP), adenosine 5′-diphosphate (ADP), adenosine 5′-triphosphate (ATP), guanosine 3′,5′-cyclic monophosphate (3′,5′-cGMP), cytosine 3′,5′-cyclic monophosphate (3′,5′-cCMP), uridine 3′,5′-cyclic monophosphate (3′,5′-cUMP) were from Sigma-Aldrich and were analytical grade.

      Oligonucleotides

      The oligonucleotides 5′A243′, 5′C243′, 5′A12C123′, 5′A12U123′, 5′U243′, and 5′G243′ were purchased from Dharmacon and were provided unphosphorylated, at both the 5′ and 3′ extremities.

      Methods

      Polymerization Protocols and Analysis

      Concentrated solutions of the appropriate nucleotide (2′-AMP, 3′-AMP, 5′-AMP, 2′,3′-cAMP, 3′,5′-cAMP, 3′,5′-cGMP, 3′,5′-cUMP, and 3′,5′-cCMP) were diluted in water to the desired final concentration. Concentrations between 1 μm and 0.1 m were analyzed. Temperatures between 25 and 90 °C and pH values 3.2, 3.7, 5.0, 5.4, 6.1, 8.0, 8.2, and 8.4, obtained by Tris-HCl buffering of bidistilled deionized MilliQ water, were tested. Other variables are discussed where appropriate. After terminal labeling (see below) the samples were analyzed by gel electrophoresis.

      Acrylamide Gel Electrophoresis

      Standard methodologies were used, with the following specifications: 1) 12% polyacrylamide was used in analyses encompassing the whole product of the polymerization reaction, from the 32P-labeled monomer to the highest molecular weight fragments (>100 units), or 2) longer runs on 16% polyacrylamide gels were used for the analysis of low molecular weight polymers. With sequences allowing good resolution, the average chain length (Navg) of the oligomers was determined by the equation Navg = ΣiniNiini, where ni is the number of chain (in %) and Ni is the length of RNA chains in nucleotides.

      Reference Ladders

      The nucleotide ladders used as standard in the gel-electrophoretic analyses of the polymerization products consisted of partially hydrolyzed 24-mer Poly(G) or Poly(A) (Dharmacon), as appropriate. Products of combinatorial ligation of preformed oligonucleotides were also used as markers, obtained as detailed in a previous study (
      • Pino S.
      • Ciciriello F.
      • Costanzo G.
      • Di Mauro E.
      ). In practice, labeled 5′A243′ was mixed with unlabeled 5′A153′ yielding fragments of 39, 48, 69, 78, and 96 nucleotides in length.
      For details, handling and analysis of the RNA hydrolytic products see Ref.
      • Saladino R.
      • Crestini C.
      • Ciciriello F.
      • Di Mauro E.
      • Costanzo G.
      . In brief, terminally labeled RNA oligonucleotides were hydrolyzed in water at 90 °C for different time periods (between 0 and 24 h) and pre-analyzed on polyacrylamide gel.

      Terminal Labeling of the Material Polymerized from Unlabeled Cyclic Nucleotides

      The products of the polymerization reactions from cyclic nucleotides were ethanol-precipitated and dissolved in 44 μl of water. For de-phosphorylation, 1 μl of shrimp alkaline phosphatase (1 unit/μl, MBI Fermentas) was added along with 5 μl of 10× shrimp alkaline phosphatase buffer, and the reaction was incubated at 37 °C for 30 min, followed by phenol extraction and ethanol precipitation. Glycogen (1 μl of stock 20 mg/ml) was added to facilitate precipitation. RNA was pelleted by centrifugation, then dissolved in 16 μl of water and labeled at the 5′ termini with 32P. Phosphorylation was carried out by adding 1 μl of T4 polynucleotide kinase (T4 PNK, 10 units/μl, New England Biolabs), 2 μl of 10× PNK buffer and 0.5 μl of [γ-32P]ATP, followed by incubation at 37 °C for 30 min. For gel electrophoresis, 10-μl aliquots of the RNA samples were resuspended in 100% formamide and separated by electrophoresis on 12 or 16% polyacrylamide gels containing 7 m urea, along with the indicated markers.

      RNase Analyses

      Phosphodiesterase I from Crotalus adamanteus venom (International Union of Biochemistry 3.1.4.1., snake venom phosphodiesterase I (SVPD I)) from Sigma (in vials ≥ 0.4 unit, purified, catalog number P3243) is a 5′-exonuclease that hydrolyzes 5′-mononucleotides from 3′-hydroxy-terminated ribo-oligonucleotides. It cleaves both 2′,5′- and 3′,5′-phosphodiester linkages, and it was here typically used at 1 milliunit/assay in 40 mm Tris-HCl, pH 8.4, and 10 mm MgCl2 in 20-μl assays. One unit hydrolyzes 1.0 μmol of bis-(p-nitrophenyl)phosphate per minute at pH 8.8 at 37 °C.
      Nuclease P1 from Penicillium citrinum (International Union of Biochemistry 3.1.30.1) is from Sigma (Cat N8630), specific activity of 200 units/mg of protein. It catalyzes the sequence nonspecific endonucleolytic cleavage of single-stranded RNA to yield nucleoside 5′-phosphates and 5′-phospho-oligonucleotides. Specific for 3′,5′-phosphodiester linkages, it is here typically used at 20 units/sample in 40 mm Tris-HCl, pH 5.4, 5 mm NaCl, 0.5 mm MgCl2, in 20-μl assays. One unit liberates 1.0 μmol of acid-soluble nucleotides from RNA per minute at pH 5.3 at 37 °C.
      T1 from Aspergillus oryzae (EC 3.1.27.3) is a 3′-5′-specific ribonuclease. It cleaves with high preference at the 3′-end of G residues but at high concentration or at longer times will cleave also at other residues (
      • Steyaert J.
      ). One unit produces acid-soluble oligonucleotides equivalent to a ΔA260 of 1.0 in 15 min at pH 7.5 at 37 °C in a reaction volume of 1 ml.

      RESULTS

      The oligomerization capacity of the cyclic forms (2′-3′ or 3′-5′) of the four monophosphate nucleosides, guanosine, adenosine, cytidine, and uridine, was tested. The open nucleotides 5′-AMP, 3′-AMP, and 2′-AMP were also tested in water at temperatures between 40 and 90 °C. A number of additional variables were analyzed: concentration, time, addition of formamide (from 0 to 100%), presence of several minerals known to catalyze phosphorylation (
      • Costanzo G.
      • Saladino R.
      • Crestini C.
      • Ciciriello F.
      • Di Mauro E.
      ) or to increase the half-life of nucleic polymers (
      • Saladino R.
      • Crestini C.
      • Ciciriello F.
      • Di Mauro E.
      • Costanzo G.
      ,
      • Ciciriello F.
      • Costanzo G.
      • Crestini C.
      • Saladino R.
      • Di Mauro E.
      ,
      • Saladino R.
      • Crestini C.
      • Busiello V.
      • Ciciriello F.
      • Costanzo G.
      • Di Mauro E.
      ,
      • Saladino R.
      • Crestini C.
      • Neri V.
      • Ciciriello F.
      • Costanzo G.
      • Di Mauro E.
      ), addition of Na4P2O7 or Na5P3O10, and combinations thereof. Of all the conditions tested, the simplest proved to be the best: water between 40 and 90 °C. Several pH values (3.2, 3.7, 5.0, 5.4, 6.1, 8.0, 8.2, and 8.4) were tested. The results observed were marginally different. The afforded polymers were 5′-terminally labeled with [γ-32P]ATP by T4 polynucleotide kinase, and the products were characterized by gel electrophoresis, allowing detailed evaluation of the lower sized oligomers.

      Syntheses from Open Nucleotides

      No product of polymerization was observed upon incubation of 2′-AMP or 3′-AMP in water (nor in any of the reaction variants listed above) at temperatures encompassed between 40 and 90 °C for periods up to 400 h. Only degradation of the input nucleotides was observed (data not shown). 5′-AMP afforded only traces of oligomerized compounds whose total did not exceed 0.5% of the input (data not shown). The short half-life of 5′-AMP at 90 °C (35 h) (
      • Saladino R.
      • Crestini C.
      • Ciciriello F.
      • Di Mauro E.
      • Costanzo G.
      ) is not compatible with the possibility of accumulating oligomers.

      Syntheses from Cyclic Nucleotides

      3′,5′-cGMP

      Fig. 1 shows the products of polymerization obtained by treating 3′,5′-cGMP in water. The formation of oligomers is evident. 3′,5′-cGMP polymerized into RNA chains that reached a size of at least 25 nucleotides, the predominant oligomer being 8-mer. Panel A reports the synthesis obtained at 85 °C as a function of the 3′,5′-cGMP concentration, showing that, above the optimal concentration of 1 mm, chain elongation is impaired and the preferentially formed 8-mer accumulates. Panels B and C show the syntheses obtained at the optimal 1 mm and at the highest possible (before aggregation) 100 mm concentration as a function of the temperature. In both cases the highest temperature tested was the most favorable for chain extension. Below 60 °C the reaction rate dropped rapidly (data not shown).
      Figure thumbnail gr1
      FIGURE 1Nonenzymatic polymerization of 3′,5′-cGMP in water. A, 3′,5′-cGMP reacted at 0.1 (lane 1), 1 (lane 2), 10 (lane 3), or 100 (lane 4) mm concentration at 85 °C for 1 h in Tris HCl-buffered water, pH 8.2. Navg 10.32, 12.43, 10.19, and 8.88, respectively. marker: hydrolyzed 24-mer polyG, 16% polyacrylamide electrophoresis. B, nonenzymatic polymerization of 3′,5′-cGMP at 60 (lane 1), 75 (lane 2), or 85 °C (lane 3) for 1 h in Tris-buffered water, pH 8.2, 1 mm 3′,5′-cGMP. The Navg after 1 h was 10.10, 11.08, and 11.84, respectively. The more represented species is the 8-mer. C, 100 mm 3′,5′-cGMP. The Navg of the polymers obtained were 8.72, 9.02, and 9.34, respectively.
      The oligomers shown are the products of synthetic reactions lasting 1 h. In kinetic analyses it was observed that at the optimal concentration (1 mm) synthesis was fast, an Navg of 11.8 being reached during handling time (<1 min), followed by slow stepwise further growth. The kinetic constant of this further growth was determined by measuring the Navg of the oligonucleotide G chains formed as a function of time at 85 °C with 1 mm 3′,5′-cGMP and was 0.4 × h−1.

      3′,5′-cAMP

      Under the same conditions of the 3′,5′-cGMP polymerization, 3′,5′-cAMP polymerized by a two-step mechanism. Fig. 2 shows the two steps observed in a 3′,5′-cAMP-fed growth experiment. First, a family of short oligomers was synthesized rapidly. The steady-state Navg of 5.32 (Fig. 2, lane 1) was reached by 60 min (50% of molecules formed in 20 min). The kinetic constant of the reaction leading to the formation of the short oligonucleotide A molecules (Navg 5.32) was determined at 85 °C and was 2 × h−1. The short oligomers did not continue growing by slow ladder-wise addition, as for 3′,5′-cGMP, but extended their size forming a heterogeneous population (Fig. 2, lane 3) in which a rapidly formed 16-mer was prominent. Sequence extension lasted 200 h, forming molecules >100 nucleotides long (Fig. 2, lane 4). The distribution of the products of oligomerization beyond 28 nucleotides in length was size-discontinuous (see the numbering at the side of lane 4), comprising a complex series of fragments. Such heterogeneous numerical distribution is best interpreted as the result of ligation of shorter pieces. A model study (
      • Pino S.
      • Ciciriello F.
      • Costanzo G.
      • Di Mauro E.
      ) showed that mixing a limited number of different RNA oligomers in water yields a complex population of differently sized RNA fragments by nonenzymatic ligation. This second reaction, presumably based on ligation of the components of a heterogeneous population, is too complex to allow calculation of kinetic constants. By contrast, 2′,3′-cAMP yielded only short oligomers, up to tetramers (data not shown). Polymerization of 3′,5′-cUMP and 3′,5′-cCMP yielded only short fragments (Navg 5.49 and 5.45, respectively) at 85 °C, which did not grow further.
      Figure thumbnail gr2
      FIGURE 2Nonenzymatic polymerization of 3′,5′-cAMP in water. 3′,5′-cAMP reacted in water (85 °C) for 30 min (lane 2) or 3 h (lane 3). Lane 1: under-exposure of part of lane 2. marker: hydrolyzed 24mer poly(A). Lane 4, blow-up analysis of the population of fragments encompassed between 20 and >120 nucleotides, 2 × 102 h of reaction, and 12% acrylamide. RNA fragments obtained by ligation (
      • Pino S.
      • Ciciriello F.
      • Costanzo G.
      • Di Mauro E.
      ) were used as markers for lane 4 (not shown).

      The Bonds Formed, as Determined by RNase Analyses

      The type of phosphate bond formed in the polymers derived from 3′,5′-cGMP and 3′,5′-cAMP was analyzed by enzymatic digestion with SVPD I (EC 3.1.4.1, a 5′-exonuclease cleaving 3′-5′ and 2′-5′ phosphodiester bonds from the 3′-extremity in a nonprocessive manner) and with P1 endonuclease (EC 3.1.30.1, a 3′-5′-specific ribonuclease). Treatment of the products of polymerization with 1 milliunit of SVPD I or of P1 for 20 min at 37 °C completely converted the oligonucleotides into monomers, showing that the bonds formed are canonical 3′-5′ phosphodiester bonds (data not shown). For details of these RNase assays, see Ref.
      • Pino S.
      • Ciciriello F.
      • Costanzo G.
      • Di Mauro E.
      . The type of phosphate bond formed in oligonucleotide G was further analyzed, as described below, confirming the formation of 3′-5′ bonds.

      On the Mechanism of Polymerization

      Although detailed mechanistic aspects of the observed polymerization of cyclic nucleotides are beyond the aim of the present communication, the following facts elucidate the basics of the reaction: (i) the RNase digestion assays mentioned above show that the bonds formed by polymerization of 3′,5′-cyclic nucleosides are standard 3′-5′ phosphodiester bonds. Given that the starting monomers are 3′,5′-cyclic phosphates, this is not unexpected. The combined SVPD I and P1 RNase analyses rule out the formation of 2′-5′ bonds, of pyrophosphate bonds, or more complex alternatives. (ii) 3′,5′-cyclic nucleoside monophosphates hydrolyze in water yielding (in the temperature and pH conditions in which polymerization occurs) a mixture of 5′ and 3′ monophosphates, as verified by high performance liquid chromatography (data not shown) and as originally reported (
      • Smith M.
      • Drummond G.I.
      • Khorana H.G.
      ).
      Thus, the polymerization could occur according to two different alternative models. Model A consists of the reactive species that is a 5′-XMP afforded by the opening of the 3′ phosphodiester bond of the cyclic nucleotide. In this case, polymerization would occur via the 5′-phosphate reacting with the 3′-OH of another 5′-XMP, as indicated by the spark symbol in Fig. 3. The reactive species is a 3′-XMP, and the polymerization occurs via the 3′-phosphate reacting with the unphosphorylated 5′-extremity of another 3′-XMP molecule. Model A would lead to the phosphate group being on the top sugar molecule (as shown in Fig. 3), rather than on the lower sugar molecule (Model B, not shown).
      Figure thumbnail gr3
      FIGURE 3A simple model for the polymerization of 3′,5′-cGMP. The cyclic bond preferentially opens affording a 5′-GMP (
      • Smith M.
      • Drummond G.I.
      • Khorana H.G.
      ). The guanine moieties of two of these molecules are supposedly held in position by stacking (
      • Olson W.K.
      ,
      • Norberg J.
      • Nilsson L.
      ,
      • Norberg J.
      • Nilsson L.
      ). Transfer of the bond is favored at moderately high temperature. The 3′-phosphate bond is more stable in the polymer than in the monomer (
      • Saladino R.
      • Crestini C.
      • Ciciriello F.
      • Di Mauro E.
      • Costanzo G.
      ) thus justifying polymer prevalence in time.
      The bias would be solved in favor of Model A if neo-formed oligonucleotide G, obtained as described in Fig. 1, would ligate to the 3′ non-phosphorylated extremity of an acceptor oligonucleotide through 3′-5′ phosphodiester bonds (as schematically described in Fig. 5). The experiments reported below (FIGURE 4, FIGURE 5, FIGURE 6) show that this is the case: the neo-formed oligonucleotide G ligated with 3′-5′ bonds to the 3′-OH extremity of a 5′C243′ and of a 5′A12C123′ oligomer. Thus, Model A applies, as shown in Fig. 3.
      Figure thumbnail gr5
      FIGURE 5The 5′A12C123′ oligonucleotide (schematically shown in its 3′-extremity (left)) ligates to a neo-synthesized base-paired oligonucleotide G segment (right) through a standard 3′-5′ phosphodiester bond.
      In summary, in the presence of the thermodynamic driving force provided by stacking interaction, an isoenergetic phosphodiester exchange reaction is favored, affording the observed products. The possibility that the reaction occurs by general acid-base catalysis is disfavored by the observation that neither the 3′,5′-cAMP nor the 3′,5′-cGMP polymerizations are pH-dependent (between pH 3.2 and 8.4, data not shown).
      The fact that the order of the stacking potentials of the bases correlates with the corresponding polymerization rates (see below) establishes the relevance of stacking interactions in this reaction.

      RNA Chain Extension

      Nonenzymatic Ligation of Nonenzymatically Polymerized Oligonucleotide G to the 3′-Extremity of Preformed Oligonucleotide Cs

      Do cyclic nucleotides polymerize in the presence of preformed oligonucleotides? If so, is this condition interactive? The answer is positive, as described below. The following oligonucleotides were tested: 5′A243′, 5′C243′, 5′A12C123′, 5′A12U123′, 5′U243′, and 5′G243′. Each one of these oligonucleotides was reacted with 3′,5′-cAMP, 3′,5′-cGMP, 3′,5′-cCMP, and 3′,5′-cUMP.
      Fig. 4 Panel A shows the results of the reaction of 5′-labeled 5′C243′ with different concentrations of unlabeled 3′,5′-cAMP, 3′,5′-cGMP, 3′,5′-cCMP, and 3′,5′-cUMP, as indicated. The key observation is that 3′,5′-cGMP actively reacted with the preformed oligonucleotide, affording longer fragments. In particular, a group of molecules with a number average (Navg) of 42 formed in the presence of 3′,5′-cGMP (lanes 6–8), that grew up to an observed length of >50 nucleotides in the presence of the higher concentration of cyclic nucleotide (as counted in the right corner inset, showing a lower exposure of the relevant gel position). A slower migration band is also observed in the upper part of the lanes 6–8 (asterisk), probably representing a dimeric form of the extended sequence.
      Figure thumbnail gr4
      FIGURE 4RNA-chain extension by 3′,5′-cGMP-fed polymerization. A, 5′-labeled 5′C243′ reacted with 3′,5′-cAMP (lanes 3–5), 3′,5′-cGMP (lanes 6–8), 3′,5′-cCMP (lanes 9–11), or 3′,5′-cUMP (lanes 12–14). Each group of three lanes contained 0.1, 1.0, or 10 mm nucleotide, respectively. The reaction was in Tris-HCl-buffered water (pH 5.4) at 60 °C for 6 h. Lane 1: U, untreated; lane 2: no nucleotide. Inset: one-third autoradiographic exposure of the corresponding part of the gel. The model interprets the structure of the polymer in denaturing (left) and water condition (right). The polymer indicated by the asterisk is formed only in the presence of 3′,5′-cGMP. Its size is 84 ± 3 nucleotides (as determined by band counting in the appropriate autoradiogram exposure and plot graphical extrapolation). This multimer is interpreted as a dimeric form of the extended monomer, possibly caused by oligonucleotide G-oligonucleotide G ligation. Oligonucleotide C-oligonucleotide C dimerization did not occur in the absence of G-based cofactors or G extensions. The band corresponding to the 23-mer is missing. For an explanation of the reduced hydrolysis of the last phosphodiester bond at the 3′-extremity (see Ref.
      • Pino S.
      • Ciciriello F.
      • Costanzo G.
      • Di Mauro E.
      ). The 23-mer is produced in enzymatic degradations (see ). B, RNA-chain extension by 3′,5′-cGMP as a function of concentration. The reaction was performed as above in the presence of the indicated concentration of 3′,5′-cGMP. C, 5′-labeled 5′A12C123′ reacted with 3′,5′-cGMP. Lane 1: untreated; lane 2: 6 h, Tris-HCl-buffered water, pH 5.4, 60 °C. The C12 segment undergoes hydrolysis faster than the A12 segment (as detailed in Ref.
      • Ciciriello F.
      • Costanzo G.
      • Pino S.
      • Crestini C.
      • Saladino R.
      • Di Mauro E.
      ). Lanes 3–5: as in lane 2, in the presence of 0.1, 0.4, and 2 mm 3′,5′-cGMP, respectively. The fragment sizes of the G-encompassing fragments were determined by top-down band counting starting from the 48-mer dimer (24 × 2) in overexposed gel images. D, chain extension of 5′C243′ and of a 5′A12C123′ as a function of the 3′,5′-cGMP concentration. The data points show the % of full sized monomer molecules (filled symbols: ▴, 5′C243′; ●, 5′A12C123′) and of the extended molecules (open symbols, ▵ and ○) as a function of the 3′,5′-cGMP concentration indicated on the abscissa. Data are from experiments reported in B (and not shown).
      The Navg was calculated from graphical extrapolation of gel positions in the appropriate autoradiographic exposures. The band-compression effect characteristic of the C residues prevents a better resolution of high molecular weight oligomers and a more precise evaluation of fragment lengths. The system was explored with higher precision in 5′A12C123′polymers (see below).
      All the 5′C243′ fragments covalently reacted with oligonucleotide G oligonucleotides (lanes 7 and 8) and formed a new population reaching an average length of 42. This entails that in the solution in which the reaction takes place oligonucleotide Cs and oligonucleotide Gs interact, presumably by base-pairing, to form a double strand. Double strands withstand hydrolysis more than single strands. If this occurs also in our conditions and sequence set-up, a footprint of ∼18 bases in length should be produced, which is actually observed (Fig. 4A, dots in lane 7; scheme on the right side). The open dots at the bottom of the lane indicate where the footprint, is not observed, showing that the chain extension does not occur from the 5′-extremity.
      The following are also noted: 1) The C stretch is highly sensitive to hydrolytic degradation (as already reported (
      • Ciciriello F.
      • Costanzo G.
      • Pino S.
      • Crestini C.
      • Saladino R.
      • Di Mauro E.
      )); 2) 3′,5′-cAMP does not support polymerization growing on the 3′-extremity nor supports multimerization by ligation (as observed for 5′A243′ oligonucleotides (
      • Pino S.
      • Ciciriello F.
      • Costanzo G.
      • Di Mauro E.
      )). Starting at 10 mm concentration, 3′,5′-cAMP enhances the hydrolytic degradation of the 5′C243′ oligonucleotide (lane 5). The same behavior was observed on Poly(A)23U, Poly(A)24, and on Poly(G)24 (data not shown); 3) 3′,5′-cCMP and 3′,5′-cUMP are inert. Thus, only the reaction of oligonucleotide C with 3′,5′-cGMP was explored further.
      Fig. 4B shows the RNA-chain extension of 5′C243′ by 3′,5′-cGMP as a function of cyclic nucleotide concentration. Panel C shows selected examples of the same reaction on 5′A12C123′.
      Consistent with the calculated Navg of the oligonucleotide G polymerized from 3′,5′-cGMP reported in Fig. 1 (in synthesis reactions in which the 8-mer was prevailing), the family of oligonucleotide Gs that polymerized from 3′,5′-cGMP in the presence of the 5′A12C123′ 24-mer and that ligated to its 3′ C-extremity had an Navg of 8.75 (Fig. 4C). This Navg value was determined from the Navg calculated from the fragment sizes observed in the gel migration ladder (Navg = 32.75) subtracting 24 (that is, the size of the acceptor 24-mer oligonucleotide).
      The following is also noted: the footprint on the C12 moiety is shorter relative to the one on the C24 oligonucleotide, similar to the chains produced (Navg = 32.75, corresponding to an extension of 8.75 on the 24-mer and to a footprint ≥8 residues, as indicated by dots) and as predicted in a model based on the Poly(C)-Poly(G) base-pairing in water. 3′,5′-cAMP, 3′,5′-cCMP, and 3′,5′cGMP did not support chain extension on the 5′A12C123′ (nor on the 5′C243′; data not shown).
      The pre-synthesized oligonucleotide G did not bind to (nor did 3′,5′-cGMP-fed polymerization occur on) pre-synthesized Poly(A) oligonucleotides (data not shown), thus excluding that the 5′A-extremity of the 5′A12C123′ molecule supported RNA-chain extension on the Poly(C) oligonucleotides.
      The fact that a footprint is observed, starting from the position in which sequence extension begins (i.e. the 3′-extremity) and is oriented in the specular direction, provides an assay for the presence of newly formed complementary sequences. No footprint is observed on the 5′-extremity, indicating that sequence extension only occurs on the 3′-OH extremity based on the 5′ P-group from the incoming molecule, and not vice versa.
      A quantitative evaluation of the RNA-chain extension occurring on 5′C243′ and on 5′A12C123′ as a function of the cyclic nucleotide concentration is reported in Fig. 4D. The plot shows that the growth of short segments occurring on 5′A12C123′ (Navg = 8.75) levels off at lower concentration of 3′,5′-cGMP, relative to the growth on 5′C243′ (Navg = 18).
      The kinetic constant of the reactions leading to the formation of the extended monomers could not be determined, because the reaction was too fast even at the lowest concentration tested (200 nm 5′C243′ and 1 μm 3′,5′cGMP), at 40 °C. Reaction rates are given in Table 1.
      TABLE 1Quantitative analysis of chain extension and terminal ligation test-systems results
      PolymerSubstrate/cofactorChain extension rate
      The chain extension rates were determined based upon densitometry measurements of autoradiograms of gel electrophoretic analysis of extension reactions (i.e., as in Fig. 4) and have been normalized with respect to the extension rate of 5′A12C123′ in the best observed conditions (6 h, 60 °C, pH 6.2, 1 mm, 3′,5′-cGMP), which has been scaled to 10,000.
      Half max
      “Half max” indicates the concentration of cyclic nucleotide at which the rate of product yield is one-half of the maximum extension or ligation rate.
      Terminal Ligation rate
      The terminal ligation rates were determined with the methodology described in footnote a, relative to the ligation rate of 5′A243′, which has been scaled to 10,000 (see Ref. 8).
      Half max
      C243′,5′-cAMP0NA
      NA, not applicable.
      0NA
      3′,5′-cGMP4.600 ± 900260 mmNot determined
      Oligonucleotide C does not dimerize. In the presence of 3′,5′-cGMP it forms a multimeric form due to a more complex phenomenon (as described in the legend to Fig. 4A).
      NA
      A12C123′,5′-cAMP0NA27 ± 102 mm
      3′,5′-cGMP10.000 ± 1.500120 mm500 ± 1002 mm
      A243′,5′-cAMP0NA10,000 ± 1,5002 mm
      3′,5′-cGMP0NA1,200 ± 2002 mm
      a The chain extension rates were determined based upon densitometry measurements of autoradiograms of gel electrophoretic analysis of extension reactions (i.e., as in Fig. 4) and have been normalized with respect to the extension rate of 5′A12C123′ in the best observed conditions (6 h, 60 °C, pH 6.2, 1 mm, 3′,5′-cGMP), which has been scaled to 10,000.
      b “Half max” indicates the concentration of cyclic nucleotide at which the rate of product yield is one-half of the maximum extension or ligation rate.
      c The terminal ligation rates were determined with the methodology described in footnote a, relative to the ligation rate of 5′A243′, which has been scaled to 10,000 (see Ref.
      • Saladino R.
      • Crestini C.
      • Ciciriello F.
      • Pino S.
      • Costanzo G.
      • Di Mauro E.
      ).
      d NA, not applicable.
      e Oligonucleotide C does not dimerize. In the presence of 3′,5′-cGMP it forms a multimeric form due to a more complex phenomenon (as described in the legend to Fig. 4A).
      In conclusion, 3′,5′-cGMP efficiently polymerizes in the presence of Poly(C) and is covalently bound to its 3′-extremity. Given that the 3′-extremity of the 5′A12C123′ oligonucleotide bears no phosphate but ends by OH in 3′, and given that the ligation occurred via 3′-5′ phosphodiester bonds (see below, section on Characterization of the Bond Formation), the observed chain-extension necessarily occurred by ligation through the 5′-phosphate group carried by the neo-polymerized oligonucleotide G, as shown in Fig. 5.

      The Rate-limiting Step

      Is the rate-limiting step of the polymerization reaction the dinucleotide formation step or the extension reaction step? In the 3′,5′-cGMP system, to answer this question we tried to measure the kinetic constant of the dinucleotide formation by lowering the concentration of 3′,5′-cGMP down to the detection limit of the assay. The results, reported in Fig. 6, show that the shortest observed measurable chain is the G8 oligomer (Navg 8.75) and that its formation is immediate. The experiment also shows that the amount of elongated polymer formed depends on the concentration of 3′,5′-cGMP, not on a kinetically limiting step. Given that the kinetic constant of the elongation reaction, as determined in the same optimal conditions (85 °C, 1 mm 3′,5′-cGMP), is relatively low (0.4 × h−1) the limiting step is chain elongation.
      Figure thumbnail gr6
      FIGURE 6The chain extension reaction by 3′,5′-cGMP on oligonucleotide C is concentration-dependent. The reaction was performed in the same conditions described in , in the presence of 10, 30, 60, or 90 μm 3′,5′-cGMP for 0, 1, 10, or 60 min, as indicated. For each group of time points the amount of elongated chain does not vary.
      As for the 3′,5′-cAMP system, the kinetic constant for chain elongation is 2 × 10−1 h. The kinetics of formation of the dimer was followed by high performance liquid chromatography analysis of the polymerized products. At no time point was the dimer observed to accumulate relative to the trimer, tetramer, etc. showing that in this system the limiting step is the dimer formation.

      Characterization of the Bond Formed upon Ligation of the 5′A12C123′ Oligonucleotide with the Neo-synthetized Oligonucleotide G

      The 5′A12C123′ oligonucleotide was reacted with 3′,5′-cGMP (60 °C, 6 h, 400 μm 3′,5′-cGMP), then treated with T1 or SVPD I ribonucleases. Fig. 7 shows that the 5′A12C12 G8.753′ is sensitive to the two nucleases, thus confirming the 3′-5′ nature of the phosphodiester bonds formed, both in the oligonucleotide G and between the oligonucleotide G and the 5′A12C123′ oligomer.
      Figure thumbnail gr7
      FIGURE 7T1 and SVPD I ribonuclease treatment of the 5′A12C12G8.753′ RNA, synthesized as in C. A, treatment of the 5′-labeled RNA with 40 units of T1 for the indicated times in Tris HCl-buffered water, pH 7.2, 2 mm EDTA, in the presence of 1 mm 3′,5′-cGMP. B, 400 units. Units are defined as in Ref.
      • Steyaert J.
      . The presence of 3′,5′-cGMP markedly decreases the ribonuclease activity, as reported (
      • Eun H.M.
      ). C, SVPD I treatment as indicated in the figure and as detailed in a previous study (
      • Pino S.
      • Ciciriello F.
      • Costanzo G.
      • Di Mauro E.
      ).
      A vast literature has accumulated on the preferential formation of the 3′-5′ over the 2′-5′ phosphodiester linkages or (more often and contrary to) in oligomerizations entailing nucleoside-5′-phosphorimidazolides and related phosphoramidates or in carbodiimide-mediated ligations (reviewed in Ref.
      • Orgel L.E.
      ). The syntheses may preferentially form one type of linkage (i.e. the 2′-3′ linkage) (
      • Lutay A.V.
      • Chernolovskaya E.L.
      • Zenkova M.A.
      • Vlassov V.V.
      ) or the other (
      • Rohatgi R.
      • Bartel D.P.
      • Szostak J.W.
      ) or both (
      • Sawai H.
      • Wada M.
      • Kouda T.
      • Nakamura
      • Ozaki A.
      ). For an in-depth review of this topic see (
      • Sawai H.
      • Wada M.
      • Kouda T.
      • Nakamura
      • Ozaki A.
      ,
      • Sawai H.
      • Totsuka S.
      • Yamamoto K.
      • Ozaki H.
      ). In summary of our RNases analyses: the linkage in the oligomers formed from 3′,5′-cGMP and 3′,5′-cAMP is 3′-5′. The discrepancy with the fact that the 2′,3′ is the most commonly observed linkage in abiotic polymerizations from preactivated compounds may be explained simply by the fact that none of the previously reported syntheses was performed with 3′,5′-cyclic nucleotides in water, as in our case.

      Increased Stability of RNA oligonucleotides in Water Is Caused by the Presence of Cyclic Monophosphate Nucleosides

      The half-life of RNA oligonucleotides in water has been a matter of detailed analyses (
      • Saladino R.
      • Crestini C.
      • Ciciriello F.
      • Di Mauro E.
      • Costanzo G.
      ,
      • Ciciriello F.
      • Costanzo G.
      • Pino S.
      • Crestini C.
      • Saladino R.
      • Di Mauro E.
      ). As expected, and based on a large body of previous studies, the observed half-life of RNA molecules depends on sequence composition, temperature, pH, and concentration. In the present analysis, the products of polymerization from 3′,5′-cGMP and 3′,5′-cAMP showed unexpectedly high t½ values. It was found that the increased life span of the oligonucleotides in water is induced by the presence of the free cyclic nucleotide, presumably due to interference with the hydrolytic degradation process by stacking interaction (Fig. 8).
      Figure thumbnail gr8
      FIGURE 8The half-life of an 5′A243′ oligomer in the absence (−, left) or in the presence (+, right) of 20 mm 3′,5′-cAMP. The oligomer was terminally labeled at 5′ and treated in water (90 °C and pH 5.4) for the time indicated on the top of each lane. Further details are available elsewhere (
      • Pino S.
      • Ciciriello F.
      • Costanzo G.
      • Di Mauro E.
      ,
      • Ciciriello F.
      • Costanzo G.
      • Pino S.
      • Crestini C.
      • Saladino R.
      • Di Mauro E.
      ). The t½ values observed were 36 h in the absence and 184 h in the presence of the cyclic nucleotide. Higher concentrations of the cyclic nucleotide further increased the t½ values. U, untreated.

      DISCUSSION

      How Was RNA Polymerization Started?

      A key step missing in the reconstruction of the origin of living systems is an abiotically plausible synthesis of RNA. To fill this gap, for the robust synthesis and the simultaneous presence of all the necessary nucleic acid precursors (which is possible in principle (
      • Saladino R.
      • Neri V.
      • Crestini C.
      • Costanzo G.
      • Graciotti M.
      • Di Mauro E.
      )), an abiotic procedure for their activation and a thermodynamically sound polymerization mechanism are needed.
      Using this logic we have analyzed nucleotide oligomerization in the conceivably simplest solvent and environment: water at temperatures between 40 and 90 °C. Despite the limits set in principle by the standard-state Gibbs free energy change problem (
      • van Holde K.
      The Origins of Life and Evolution.
      ,
      • Alberty R.A.
      ), we observed that the process does actually take place in water and report the nonenzymatic formation of RNA chains in water from 3′,5′-cyclic nucleotides.
      We describe three mechanisms for nonenzymatic RNA generation: RNA polymerization from monomers, RNA ligation, RNA extension by polymerization on pre-existing oligomers, and ligation. RNA ligation was recently reported in a model study performed on Poly(A) oligomers (
      • Pino S.
      • Ciciriello F.
      • Costanzo G.
      • Di Mauro E.
      ).
      We observe that 3′,5′-cGMP polymerized into RNA chains at least 25 nucleotides long (Fig. 1), the predominant oligomer being the 8-mer. At the optimal 1 mm concentration, synthesis was fast, a Navg of 11.8 being reached within 1 min, followed by slow stepwise further growth. Canonical 3′,5′-phosphodiester bonds were formed, as determined by RNase sensitivity. 3′,5′-cAMP polymerized more slowly to oligomers that reached an Navg of 5.32 within 1 h. These oligomers expanded their size by inter-fragments ligation for a period of at least 200 h, yielding molecules >100 nucleotides long.

      The Plausibility of 3′,5′ Cyclic Nucleotides as Precursors in Nonenzymatic Polymerizations

      Nonenzymatic polymerizations require preactivated monomers (
      • van Holde K.
      The Origins of Life and Evolution.
      ,
      • Alberty R.A.
      ). The results obtained with the phosphoramidated nucleotides commonly used (
      • Mansy S.S.
      • Schrum J.P.
      • Krishnamurthy M.
      • Tobé S.
      • Treco D.A.
      • Szostak J.W.
      ,
      • Lohrmann R.
      ,
      • Ferris J.P.
      • Ertem G.
      ,
      • Kawamura K.
      • Ferris J.P.
      ,
      • Kanavarioti A.
      • Monnard P.A.
      • Deamer D.W.
      ,
      • Monnard P.A.
      • Kanavarioti A.
      • Deamer D.W.
      ,
      • Huang W.
      • Ferris J.P.
      ,
      • Ferris J.P.
      • Joshi P.C.
      • Wang K.J.
      • Miyakawa S.
      • Huang W.
      ) show that the accumulation of polymerized forms is possible once suitable activated monomers are available. Although these studies provide useful data on the formation and properties of RNA oligomers formed by chemical synthesis, their prebiotic relevance was questioned (
      • Orgel L.E.
      ,
      • Orgel L.E.
      ). The action of several organic agents (
      • Miller S.L.
      • Parris M.
      ,
      • Steinman G.
      • Lemmon R.M.
      • Calvin M.
      ,
      • Beck A.
      • Orgel L.E.
      ) and of inorganic polyphosphates (
      • Schwartz A.
      • Ponnamperuma C.
      ) on polymerization in aqueous solution was reported.
      An innovative nonenzymatic polymerization system was recently reported describing the lipid-assisted synthesis of RNA-like polymers from mononucleotides (
      • Rajamani S.
      • Vlassov A.
      • Benner S.
      • Coombs A.
      • Olasagasti F.
      • Deamer D.
      ). Chemical activation of the mononucleotides was not required. Instead, synthesis of phosphodiester bonds was driven by the chemical potential of fluctuating anhydrous and hydrated conditions, with heat providing the activation energy. Chemical complexity prevented the full analysis of the RNA-like products of this otherwise promising system.
      Cyclic nucleoside monophosphates were suggested as possible prebiotic compounds (
      • Usher D.A.
      • Mc Hale A.H.
      ,
      • Renz M.
      • Lohrmann R.
      • Orgel L.E.
      ), the driving force for polymerization being their high reactivity and the large negative standard enthalpy of hydrolysis. The prebiotic relevance of these polymerizations was questioned, because efficient synthesis was observed with 2′,3′- but not with 3′,5′-cyclic forms.
      In the possibly simplest activation system so far described, the phosphorylation of nucleosides by free phosphates or phosphate minerals in formamide was observed (
      • Costanzo G.
      • Saladino R.
      • Crestini C.
      • Ciciriello F.
      • Di Mauro E.
      ). The system affords 2′-AMP, 3′-AMP, 5′-AMP, 2′,3′-cAMP, and 3′,5′-AMP providing prebiotically plausible precursors to polymerization.
      Nucleoside phosphorylation also occurs in water (
      • Saladino R.
      • Crestini C.
      • Ciciriello F.
      • Pino S.
      • Costanzo G.
      • Di Mauro E.
      ). Treatment of adenosine in water with 1 m KH2PO4 afforded the five phosphorylated forms. A high concentration of phosphate donor is necessary and in optimized conditions (16 h, 1 m KH2PO4, 90 °C, pH 6.1) the total amount of phosphorylated products reaches only the 7.3% of the input adenosine. In these conditions the half-lives of the open phosphorylated forms 2′-AMP, 3′-AMP, and 5′-AMP are 15, 23, and 35 h, respectively, whereas the 2′,3′- and 3′,5′-cAMP cyclic forms have half-lives of 165 and 450 h, respectively (
      • Saladino R.
      • Crestini C.
      • Ciciriello F.
      • Di Mauro E.
      • Costanzo G.
      ). Adenosine half-life in the same environments is 450 h. Thus, the formation of cyclic nucleotides also occurs in water, although not efficiently and at high temperature. Cyclic monophosphate nucleosides can be synthesized abiotically by a two-stage nucleobase assembly process on a sugar-phosphate scaffold, as shown for cytidine-2′,3′-cyclic phosphate (
      • Powner M.W.
      • Gerland B.
      • Sutherland J.D.
      ).
      The stability of cyclic monophosphate nucleosides and of their precursor is of concern when one attempts to retrace the route followed by initial nascent ribopolymers. A possible solution is provided by the observation that in monophosphate ribonucleotides the 3′-phosphate bond, the weakest bond in water, is stabilized upon polymerization (
      • Saladino R.
      • Crestini C.
      • Ciciriello F.
      • Di Mauro E.
      • Costanzo G.
      ). This property may endow the polymer with an evolutionary edge over the monomer, allowing accumulation of complex chemical information. Protective conditions, like inclusion in micelles, interaction with mineral surface (
      • Ciciriello F.
      • Costanzo G.
      • Crestini C.
      • Saladino R.
      • Di Mauro E.
      ) or inner strata (i.e. in clays (
      • Bernal J.D.
      ,
      • Cairns-Smith A.G.
      )), cycles of displacement into cooler surroundings, etc., might have played an important role in the formation and accumulation of activated precursors.

      On the Mechanism of Polymerization

      The observed polymerizations only occur with cyclic nucleotides and do not take place with noncyclic forms. Sizeable polymerization is observed only with 3′-5′ cyclic nucleotides whereas the 2′,3′ cyclic ones only afford very short chains (up to tetramers).
      These facts help to focus on the possible mechanism, based on the formation of the internucleotide bonds requiring the opening of the cyclic phosphate bridge. The nonenzymatic joining of oligoadenylates on a polyuridylic acid template was reported (
      • Renz M.
      • Lohrmann R.
      • Orgel L.E.
      ). In that case 3′,5′-linked hexa-adenylic acid with a 2′,3′-cyclic phosphate terminus was shown to couple on a polyuridylic acid template in the presence of ethylenediamine, most often yielding a dodecamer.
      Before that, syntheses of oligomers were obtained from 2′,3′-cAMP (
      • Usher D.A.
      • Mc Hale A.H.
      ) upon polymerization on a poly(U) or from 2′,3′-cAMP evaporated from solution in the presence of catalysts such as aliphatic diamines (
      • Verlander M.S.
      • Lohrmann R.
      • Orgel L.E.
      ). The self-polymerization afforded oligonucleotides of chain length up to at least 6. In both the reported reactions the opening of the phosphate cyclic bridge supposedly provided the necessary activation energy.
      Nonenzymatic template-directed ligation of terminally preactivated oligonucleotides was reported (Refs.
      • Rohatgi R.
      • Bartel D.P.
      • Szostak J.W.
      ,
      • Sawai H.
      • Wada M.
      • Kouda T.
      • Nakamura
      • Ozaki A.
      ,
      • Mansy S.S.
      • Schrum J.P.
      • Krishnamurthy M.
      • Tobé S.
      • Treco D.A.
      • Szostak J.W.
      ,
      • Gao K.
      • Orgel L.E.
      , and
      • Joyce G.F.
      and references therein). In these works the formation of the internal phosphodiester bond is attributed to the template-mediated proximity of the reactive groups. In contrast to these systems, the syntheses reported here require no special preactivation, no catalyst, and no dry chemistry, and polymerization spontaneously occurs in water.

      A Role for Stacking Interactions

      The observed polymerizations occur in solution. The question thus arises as to how nucleic bases interact, rapidly and not based on sequence complementarity, and pertains first to the conditions allowing stacking of nucleoside monophosphates in solution.
      Stacking free energy profiles for all 16 natural ribodinucleoside monophosphates in aqueous solution were reported (
      • Olson W.K.
      ,
      • Norberg J.
      • Nilsson L.
      ,
      • Norberg J.
      • Nilsson L.
      ). The potential of mean force calculations showed that the free energy profiles displayed the deepest minima and the highest barriers and, therefore, the highest stacking abilities, for the purine-purine dimers, especially for ApA and GpG. The free energy of stabilizing the stacked state were 2–6 kcal/mol higher for purine-purine dimers than for pyrimidine-pyrimidine dimers. Base combinations with different stacking potentials (ApA > GpG > UpU ≅ CpC) (
      • Norberg J.
      • Nilsson L.
      ) show a corresponding order of decreasing polymerization rate (A > G > U ≅ C), reinforcing the explanation that the formation of oligonucleotides in solution relies on stacking for the passage from monomer to short oligonucleotides to occur.
      The explanation for the formation of long sequences by terminal ligation (
      • Saladino R.
      • Crestini C.
      • Ciciriello F.
      • Pino S.
      • Costanzo G.
      • Di Mauro E.
      ) (Figs. 4 and 5) relies in the studies by Holcomb and Tinoco (
      • Holcomb D.N.
      • Tinoco Jr., I.
      ) and by Brahms et al. (
      • Brahms J.
      • Michelson A.M.
      • Van Holde K.E.
      ) who first described the double strand formation by ribo-Poly(A) and the relationship (
      • Brahms J.
      • Michelson A.M.
      • Van Holde K.E.
      ) between Poly(A) length and strand coupling. Poly(A) strands are held together by stacking (
      • Holcomb D.N.
      • Tinoco Jr., I.
      ,
      • Finch J.T.
      • Klug A.
      ), the double strands are parallel and ligate terminally (
      • Pino S.
      • Ciciriello F.
      • Costanzo G.
      • Di Mauro E.
      ) in the absence of enzymes, affording molecules with standard 3′,5′ bonds in their entire length (
      • Pino S.
      • Ciciriello F.
      • Costanzo G.
      • Di Mauro E.
      ). The stacking-unstacking process is considered to be dependent on temperature (
      • Norberg J.
      • Nilsson L.
      ), pH (
      • Norberg J.
      • Nilsson L.
      ,
      • Holcomb D.N.
      • Tinoco Jr., I.
      ), and fragment size (
      • Brahms J.
      • Michelson A.M.
      • Van Holde K.E.
      ) and, in general, favored by lower temperature and pH, and longer size. The study of the free energy profiles of stacking for all 16 natural ribonucleoside monophosphates based on potential of mean force calculations shows that many different conformations, with different degrees of stacking, are possible, revealing the gradual nature of the stacking phenomenon (
      • Norberg J.
      • Nilsson L.
      ). This explains the variation in equilibrium constants and fraction stacking of ribodinucleoside monophosphates reported (
      • Frechet D.
      • Ehrlich R.
      • Remy P.
      • Gabarro-Arpa J.
      ,
      • Ezra F.S.
      • Lee C.H.
      • Kondo N.S.
      • Danyluk S.S.
      • Sarma R.H.
      ,
      • Brahms J.
      • Maurizot J.C.
      • Michelson A.M.
      ,
      • Davis R.C.
      • Tinoco Jr., I.
      ,
      • Kang H.
      • Chou P.J.
      • Johnson Jr., W.C.
      • Weller D.
      • Huang S.B.
      • Summerton J.E.
      ) and predicts that various degrees of stacking may occur also in sub-optimal conditions, such as higher temperature.
      Hence, we hypothesize that the oligomerization reactions from 3′,5′-cGMP and from 3′,5′-cAMP described in Figs. 1 and 2 rely on the stacking interaction of the purine moieties of the cyclic nucleotides, followed by the opening of the phosphodiester cyclic bond and the consequent formation of the internucleotide phosphodiester bridge. This latter part of the reaction is favored by high temperature.
      The ligation process involved in the formation of the long A stretches has been described (
      • Pino S.
      • Ciciriello F.
      • Costanzo G.
      • Di Mauro E.
      ). The sequence extension due to the terminal ligation reaction of Poly(G) on Poly(C) described in Figs. 4 and 5 need not be different from this type of ligation. Nevertheless, while the Poly(A) ligation occurred on parallel-bound double strands of A residues held by stacking, the latter occurred on antiparallel hydrogen-bonded base-paired double strands. The versatility of the set of nonenzymatic polymerization reactions leading to longer sequences (Fig. 9) is possibly the most relevant property of these self-polymerizing systems.
      Figure thumbnail gr9
      FIGURE 9Abiotically formed cyclic precursors may actually have started their evolution toward complexity in a warm little pond, as first conceived by Darwin (
      • Darwin F.
      ).

      REFERENCES

        • Orgel L.E.
        Crit. Rev. Biochem. Mol. Biol. 2004; 39: 99-123
        • Orgel L.E.
        Trends Biochem. Sci. 1998; 23: 491-495
        • van Holde K.
        The Origins of Life and Evolution.
        in: Halvorson H.O. van Holde K.E. Alan R. Liss, Inc., New York1980: 31
        • Alberty R.A.
        Biophys. Chem. 2006; 121: 157-162
        • Costanzo G.
        • Saladino R.
        • Crestini C.
        • Ciciriello F.
        • Di Mauro E.
        J. Biol. Chem. 2007; 282: 16729-16735
        • Saladino R.
        • Crestini C.
        • Ciciriello F.
        • Costanzo G.
        • Di Mauro E.
        Chem. Biodiv. Helv. Chim. Acta. 2007; 4: 694-720
        • Saladino R.
        • Ciambecchini U.
        • Crestini C.
        • Costanzo G.
        • Negri R.
        • Di Mauro E.
        ChemBioChem. 2003; 4: 514-521
        • Saladino R.
        • Crestini C.
        • Ciciriello F.
        • Pino S.
        • Costanzo G.
        • Di Mauro E.
        Res. Microbiol. 2009;
        • Darwin F.
        The Life and Letters of Charles Darwin. Vol. 3. John Murray, London1888: 18 (letter to Joseph Hooker)
        • Pino S.
        • Ciciriello F.
        • Costanzo G.
        • Di Mauro E.
        J. Biol. Chem. 2008; 283: 36494-36503
        • Saladino R.
        • Crestini C.
        • Ciciriello F.
        • Di Mauro E.
        • Costanzo G.
        J. Biol. Chem. 2006; 281: 5790-5796
        • Steyaert J.
        Eur. J. Biochem. 1997; 247: 1-11
        • Ciciriello F.
        • Costanzo G.
        • Crestini C.
        • Saladino R.
        • Di Mauro E.
        Astrobiology. 2007; 7: 616-630
        • Saladino R.
        • Crestini C.
        • Busiello V.
        • Ciciriello F.
        • Costanzo G.
        • Di Mauro E.
        J. Biol. Chem. 2005; 280: 35658-35669
        • Saladino R.
        • Crestini C.
        • Neri V.
        • Ciciriello F.
        • Costanzo G.
        • Di Mauro E.
        ChemBioChem. 2006; 7: 1707-1714
        • Smith M.
        • Drummond G.I.
        • Khorana H.G.
        J. Am. Chem. Soc. 1960; 83: 698-706
        • Ciciriello F.
        • Costanzo G.
        • Pino S.
        • Crestini C.
        • Saladino R.
        • Di Mauro E.
        Biochemistry. 2008; 47: 2732-2742
        • Lutay A.V.
        • Chernolovskaya E.L.
        • Zenkova M.A.
        • Vlassov V.V.
        Biogeosci. Discuss. 2006; 3: 1-21
        • Rohatgi R.
        • Bartel D.P.
        • Szostak J.W.
        J. Am. Chem. Soc. 1996; 118: 3340-3344
        • Sawai H.
        • Wada M.
        • Kouda T.
        • Nakamura
        • Ozaki A.
        ChemBioChem. 2006; 7: 605-611
        • Sawai H.
        • Totsuka S.
        • Yamamoto K.
        • Ozaki H.
        Nucleic Acids Res. 1998; 26: 2995-3000
        • Saladino R.
        • Neri V.
        • Crestini C.
        • Costanzo G.
        • Graciotti M.
        • Di Mauro E.
        J. Am. Chem. Soc. 2008; 130: 15512-15518
        • Mansy S.S.
        • Schrum J.P.
        • Krishnamurthy M.
        • Tobé S.
        • Treco D.A.
        • Szostak J.W.
        Nature. 2008; 454: 122-125
        • Lohrmann R.
        J. Mol. Evol. 1977; 10: 137-154
        • Ferris J.P.
        • Ertem G.
        J. Am. Chem. Soc. 1993; 115: 12270-12275
        • Kawamura K.
        • Ferris J.P.
        J. Am. Chem. Soc. 1994; 116: 7564-7572
        • Kanavarioti A.
        • Monnard P.A.
        • Deamer D.W.
        Astrobiology. 2001; 1: 271-281
        • Monnard P.A.
        • Kanavarioti A.
        • Deamer D.W.
        J. Am. Chem. Soc. 2003; 125: 13734-13740
        • Huang W.
        • Ferris J.P.
        Chem. Commun. 2003; 12: 1458-1459
        • Ferris J.P.
        • Joshi P.C.
        • Wang K.J.
        • Miyakawa S.
        • Huang W.
        Adv. Space Res. 2004; 33: 100-105
        • Miller S.L.
        • Parris M.
        Nature. 1964; 204: 1248
        • Steinman G.
        • Lemmon R.M.
        • Calvin M.
        Proc. Natl. Acad. Sci. U.S.A. 1964; 52: 27-30
        • Beck A.
        • Orgel L.E.
        Proc. Natl. Acad. Sci. U.S.A. 1965; 54: 664-667
        • Schwartz A.
        • Ponnamperuma C.
        Nature. 1968; 218: 443
        • Rajamani S.
        • Vlassov A.
        • Benner S.
        • Coombs A.
        • Olasagasti F.
        • Deamer D.
        Origins Life Evol. Biosphere. 2008; 38: 57-74
        • Usher D.A.
        • Mc Hale A.H.
        Science. 1976; 192: 53-54
        • Renz M.
        • Lohrmann R.
        • Orgel L.E.
        Biochim. Biophys. Acta. 1971; 240: 240463-240471
        • Powner M.W.
        • Gerland B.
        • Sutherland J.D.
        Nature. 2009; 459: 239
        • Bernal J.D.
        Proc. Phys. Soc. Sect. A. 1949; 62: 537-558
        • Cairns-Smith A.G.
        J. Theor. Biol. 1966; 10: 53-88
        • Verlander M.S.
        • Lohrmann R.
        • Orgel L.E.
        J. Mol. Evol. 1973; 2: 303-316
        • Gao K.
        • Orgel L.E.
        Origins Life Evol. Biosphere. 2000; 30: 45-51
        • Joyce G.F.
        Annu. Rev. Biochem. 2004; 73: 791-836
        • Olson W.K.
        Nucleic Acids Res. 1975; 2: 2055-2068
        • Norberg J.
        • Nilsson L.
        J. Am. Chem. Soc. 1995; 117: 10832-10840
        • Norberg J.
        • Nilsson L.
        J. Phys. Chem. 1995; 99: 13056-13058
        • Holcomb D.N.
        • Tinoco Jr., I.
        Biopolymers. 1965; 3: 121-133
        • Brahms J.
        • Michelson A.M.
        • Van Holde K.E.
        J. Mol. Biol. 1966; 15: 467-488
        • Finch J.T.
        • Klug A.
        J. Mol. Biol. 1969; 46: 597-598
        • Frechet D.
        • Ehrlich R.
        • Remy P.
        • Gabarro-Arpa J.
        Nucleic Acids Res. 1979; 7: 1981-2001
        • Ezra F.S.
        • Lee C.H.
        • Kondo N.S.
        • Danyluk S.S.
        • Sarma R.H.
        Biochemistry. 1977; 16: 1977-1987
        • Brahms J.
        • Maurizot J.C.
        • Michelson A.M.
        J. Mol. Biol. 1967; 25: 481-495
        • Davis R.C.
        • Tinoco Jr., I.
        Biopolymers. 1968; 6: 223-242
        • Kang H.
        • Chou P.J.
        • Johnson Jr., W.C.
        • Weller D.
        • Huang S.B.
        • Summerton J.E.
        Biopolymers. 1992; 32: 1351-1363
        • Eun H.M.
        Enzymology Primer for Recombinant DNA Technology. Academic Press, Inc., New York1996: 647-673