HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS		QUICK SEARCH:   [advanced] Author: Keyword(s): Year:  Vol:  Page:

J. Biol. Chem., Vol. 279, Issue 1, 540-546, January 2, 2004

This Article Abstract Full Text (PDF) All Versions of this Article: 279/1/540    most recent M305499200v1 Submit a Letter to Editor Alert me when this article is cited Alert me when eLetters are posted Alert me if a correction is posted Citation Map Services Email this article to a friend Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Request Permissions Citing Articles Citing Articles via HighWire Citing Articles via Google Scholar Google Scholar Articles by Ohuchi, S. J. Articles by Inoue, T. Search for Related Content PubMed PubMed Citation Articles by Ohuchi, S. J. Articles by Inoue, T. Social Bookmarking                      What's this?

Artificial Modules for Enhancing Rate Constants of a Group I Intron Ribozyme without a P4-P6 Core Element^*

Shoji J. Ohuchi, Yoshiya Ikawa, Hideaki Shiraishi, and Tan Inoue¶

From the Graduate School of Science and Graduate School of Biostudies, Kyoto University, Kyoto 606-8502, Japan

Received for publication, May 27, 2003 , and in revised form, October 6, 2003.

	ABSTRACT

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
In this paper we report newly selected artificial modules that enhance the k_cat values comparable with or higher than those of the wild-type ribozyme with broad substrate specificity. The elements required for the catalysis of Group I intron ribozymes are concentrated in the P3-P7 domain of their core region, which consists of two conserved helical domains, P4-P6 and P3-P7. Previously, we reported the in vitro selection of artificial modules residing at the peripheral region of a mutant Group I ribozyme lacking P4-P6. We found that derivatives of the ribozyme containing the modules performed the reversal of the first step of the self-splicing reaction efficiently by using their affinity to the substrate RNA, although their k_cat values and substrate specificity were uninfluenced and limited, respectively. The results show that it is possible to add a variety of new domains at the peripheral region that play a role comparable with that of the conserved P4-P6 domain.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Group I intron ribozymes catalyze two consecutive transesterification reactions to excise themselves from the precursor RNAs and ligate the flanking exons together (1). Their core region consists of two completely conserved helical domains, P3-P7 and P4-P6, which are connected via base-triples, which can be seen in Fig. 1A (2–5). Biochemical studies indicate that P7 and J8/7 regions in the P3-P7 domain are essential for the catalysis of those reactions mentioned above (6–12). A study employing the T4 td group I intron demonstrated that a mutant ribozyme lacking both the P4-P6 domain and the base triples (M1 mutant, Fig. 1B) can still perform the trans-esterification reactions with moderate activity that was 10³ times lower than that of the wild-type ribozyme at higher concentrations of magnesium (13).

View larger version (23K): [in this window] [in a new window]   FIG. 1. Secondary structure of the ribozymes. A, wild-type T4 td ribozyme. B, M1 mutant ribozyme. The insertion sites for random sequences (N40 or N70) are shown by gray boxes. An arrowhead indicates the 5' splice site. Gray lines indicate known tertiary interactions.

We report here newly selected modules that enhance the activity of the mutant ribozyme lacking P4-P6 domain with broad substrate specificity. After 12 rounds of in vitro selection followed by an additional modification and selection, the selected variants performed the reaction with second order rate constants (k_cat/K_m values) one order of magnitude larger than the wild-type ribozyme. Furthermore, their reaction rates (k_cat values) were comparable with that of the wild-type ribozyme.

	EXPERIMENTAL PROCEDURES

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Library Construction—We used the N40 library that was constructed as previously described with minor modifications (14). An oligodeoxynucleotide, td-R (5'-ATTATGTTCAGATAAGGTCGTTAATC-3'; all oligodeoxynucleotides were purchased from Hokkaido System Science, Japan), instead of td-Rv, was used as the reverse primer throughout the experiments. In this primer the -G portion was eliminated to inhibit side reactions (circularization). The N70 library was constructed with essentially the same procedure as above using an oligodeoxynucleotide, Ban-N70 (5'-CGTGGACAGGGTGAGCGAGGCACCN₇₀GAGCCCTGTC-CACGCTCGCTCAC-3'; where N stands for any nucleotide, and restriction sites are shown in italics), as the template for producing the RNA pool. About 1.0 x 10¹⁴ molecules of each sub-library DNA were transcribed by using T7 RNA polymerase to yield the RNA pools (20 copies each).

In Vitro Selection—We performed in vitro selection as previously described with minor modifications (14–16). The pool RNA was dissolved in 1.25x reaction buffer (1x reaction buffer contains 50 mM Hepes-KOH, pH 7.5, 2.5 mM spermidine) followed by denaturation by incubating at 65 °C for 10 min. The RNAs were folded by adding volume of 100 mM MgCl₂ (final concentration was 10 mM) followed by incubation at 37 °C for 10 min. The reaction was initiated by adding 1/9 volume of the substrate RNA (S-2, 5'-biotin-GUAGCACAUCAUCGCAUCACAAAUCAGGU-3', or S-3, 5'-biotin-AUCAUCGCAUCACCACGUAGAAAUC-AGGU-3'; Dharmacom Research). The reaction mix was incubated at 37 °C. We arrested the reaction by adding 1/10 volume of 200 mM EDTA. Table I shows that the concentration of the substrate RNA and the reaction time were progressively decreased. Biotin affinity purification, reverse transcription, and cDNA elution were performed as described (14), and selective PCR was carried out by using a selective primer (5'-GTAGCACATCATCGCATCACAAATCAGG-3' for the substrate S-2 and 5'-ATCATCGCATCACCACGTAGAAATCAGG-3' for the substrate S-3) and td-R. A nested PCR was performed as described.

A doped clone 22 library was constructed as follows. We amplified the fragments for insertion with an oligodeoxynucleotide, DPD22 (5'-CGTGGACAGGGTGAGCGAGGCACCCTCGTTTGTACTGAAAAGCAATCGGGGGAATATGGTAGTGAGCCCTGTCCACGCTCGCTCAC-3'; underlined nucleotides were 30% doped (70% of original nucleotide and 3 x 10% of the other nucleotides)), by employing ExTaq DNA polymerase (Takara Shuzo). Restriction sites are in italics. The PCR product was digested with BanI and BanII. This fragment was ligated with a 5' and 3' fragment of the BanI/BanII digest of clone 22 DNA by using T4 DNA ligase. The ligated DNAs (1.2 x 10¹³ molecules) purified by native PAGE were used for transcription by using T7 RNA polymerase (40 copies each). Two rounds of in vitro selection from the doped libraries were carried out as described above under the following conditions; reaction time was 15 s, and the substrates for round 1 and round 2 were 2 µM S-2 and 2 µM S-3, respectively.

Hydroxyl Radical Footprinting—We conducted hydroxyl radical footprinting as described in Celander (17). The RNAs (1–2 µg) were dissolved in a 2x folding buffer (1x folding buffer contains 50 mM MOPS,¹ pH 7.3, 100 mM NH₄Cl,5mM dithiothreitol) followed by denaturation by incubation at 65 °C for 10 min. The RNAs were folded in the presence of 10 or 50 mM MgCl₂ followed by incubation at 37 °C for 10 min. The cleavage reaction was initiated by adding free radical generator. The resulting mixture contains 5 mM EDTA, 2.5 mM Fe(NH₄)₂(SO₄)₂, 5 mM dithiothreitol, 5 mM ascorbic acid, and 0.05% H₂O₂. The cleavage reaction was carried out at 37 °C for 1–3 min and arrested by adding an equal volume of 1 M thiourea followed by ethanol precipitation. The cleavage sites were mapped by reverse transcription with a 5' end-labeled primer, 8-BanI-Rv (14), and ReverTra Ace (Toyobo) followed by 7% denaturing PAGE.

	RESULTS

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
In Vitro Selection—To select artificial modules that enhance the activity of the M1 mutant ribozyme lacking the P4-P6 domain together with the base triples, we constructed its derivatives, which possess random sequences for the selection. The random sequence consisting of 40 or 70 nucleotides was inserted into peripheral loop L7.1, L8, or L9 (boxes in Fig. 1B) of the M1 mutant. Each pool of RNA contained 1.0 x 10¹⁴ different molecules. Three pools containing either a 40- or 70-nucleotide random sequence were mixed to form the N40 and N70 libraries, respectively. Active ribozymes were selected from the libraries by attempting the reversal reaction of the first step of self-splicing (Fig. 2). The selection conditions were restricted gradually by reducing the reaction time. Two substrates, S-2 and S-3, with a different 5' tag sequence were employed alternately after the completion of each cycle to inhibit the enrichment of the clones specialized for accepting a particular substrate (Table I). The variants in Round 12 pools were cloned because the increased activity was observed for the two pools containing N40 or N70 libraries (Fig. 3) that can utilize both S-2 and S-3 as the substrate. Sequences of the clones are shown in Supplemental Figs. 1 and 2.

View larger version (14K): [in this window] [in a new window]   FIG. 2. Scheme for the in vitro selection. Circled B and the black and white region in the substrate RNA indicate the 5' biotin moiety and the sequence switched after each cycle (black) and the constant sequence (white), respectively. SA, streptavidin beads.

View larger version (25K): [in this window] [in a new window]   FIG. 3. Increase in the activity due to the in vitro selection. A, the reaction of the wild-type (lanes 1 and 2), M1 mutant (lanes 3 and 4), Round 12 pools of the N70 library (lanes 5 and 6), or Round 12 pools of the N40 library (lanes 7 and 8) were carried out with 10 µM substrate RNA (lanes 1, 3, 5, and 7; S-2, lanes 2, 4, 6, and 8; S-3) for 1 min. B, each pool was reacted with 10 µM substrate RNA (S-2 or S-3) for 2 min.

To determine whether each clone has broad substrate specificity, we attempted the reaction with a new substrate (S-4) with a different sequence to that of S-2 or S-3 except the P1 region (Fig. 4). The S-4 substrate served as the substrate for clone 10 from the N70 pool to the extent comparable with S-2 or S-3. The clones from the N40 pool reacted efficiently with S-4 without exception, and the results indicate that the ribozymes with broad substrate specificity were selected as anticipated. However, all clones from the N70 pool except clone 10 failed to react with S4. This may be attributed to a partial sequence of their insertion, GUGUGUG (178–184, Supplemental Fig. 1), which is partially complementary to S-2 (CACN₁₀CAC) and S-3 (CACCAC).

View larger version (16K): [in this window] [in a new window]   FIG. 4. Activities of clones from Round 12 pools. The activity was determined with 10 µM substrate RNA (S-2, S-3, or S-4). Clones with similar sequences are indicated by underlines.

The inserted region in clones 10 (including the restriction sites) and 22 were 21 and 30% doped to form doped 10 and 22 libraries, respectively. The pools with 1 x 10¹³ variants were constructed followed by two rounds of in vitro selection. Clones were isolated from two pools that were as active as the parental clones (clone 10 and clone 22) after the selection. We found that 27 of 28 clones and 27 of 29 clones from doped 10 and 22 pools, respectively, showed comparable activity to the corresponding parental clones (Supplemental Figs. 3 and 4).

The Mfold program predicted that the clones from the doped 10 library contained two conserved helices, two base pairs extension of P8 (G1:C77 and G2:C76, Fig. 5, A and C) and a stem consisting of seven base pairs (base pairs between A4-C10 and G32-U38, Fig. 5, A and C). It was also observed that the sequence of U56-U67 was highly conserved (Fig. 5, A and C, see also Supplemental Fig. 3). Fig. 5C also shows that clone 107 was predicted to possess an extra stem consisting of seven base pairs (base pairs between U12-A18 and U24-A30) that was not found in clone 10. The corresponding stems were also observed for the majority of the clones (green in Supplemental Fig. 3). The predicted secondary structure of clone 107 suggests that the base pairings may stabilize the conserved structure and/or inhibit the formation of alternative forms (Fig. 5C).

View larger version (19K): [in this window] [in a new window]   FIG. 5. Secondary structural models of clone 10 (A), clone 22 (B), clone 107 (C), and clone 204 (D). In C and D, the numbers on the nucleotides are set based on those in A and B, respectively. Red boxes indicate the conserved helix structures, blue letters indicate the conserved primary sequences, and red letters with gray arrows indicate the mutations. The insertion and deletion are indicated by (ins.) and , respectively. The nucleotides highlighted with gray circles are complementary to L1 (see "Directed Evolution of Clone 10 and Clone 22" under "Results").

View larger version (20K): [in this window] [in a new window]   FIG. 6. A, proposed base pairings (red) between the inserted module and L1 in clone 204. B, mutational analysis of the base pairings. Sequence substitutions are highlighted with red boxes. nt, nucleotides. The numbers at the left are kobs values of the reaction with 0.5 µM S-2.

The Structural Roles of the Inserted Modules—That clone 107 and 204 can efficiently carry out the reaction in the presence of relatively low concentrations of magnesium ions (10 mM) indicates that the inserted modules play a role in stabilizing the active structure. To determine whether the core regions of the clones are highly structured, as is the case for the wild-type ribozyme, we performed hydroxyl radical footprinting experiments (28). The core region, P3, P7, and flanking regions between P3 and P7 of clones 107 and 204 were highly protected (Fig. 7, B and C; see also Supplemental Fig. 5, B and C) at 10 mM magnesium ions, where the corresponding region of the wild-type ribozyme was also highly protected (Fig. 7A and Supplemental Fig. 5A). However, the core region of M1 mutant was not protected or moderately protected or in the presence of 10 or 50 mM magnesium ions, respectively (Fig. 7D and Supplemental Fig. 5D). The results indicate that the inserted modules stabilize the catalytic core directly by interacting with the core region and/or indirectly by stabilizing the overall structure.

View larger version (105K): [in this window] [in a new window]   FIG. 7. Hydroxyl radical footprinting of the wild-type ribozyme (A), clone 107 (B), clone 204 (C), and M1 mutant (D). Lanes 1, 2, 3, and 4 are sequence ladders of A, C, G, and U, respectively. Lanes 5 and 6 in A–C and lanes 5–7 in D are the reverse transcription products using template RNAs without the hydroxyl radical cleavage as the control. Quantified data are shown in Supplemental Fig. 5.

According to the recently refined model structure, the T4 td ribozyme has a shorter P1 and longer P2 compared with the that previously proposed (Fig. 8A) (20). The substrate RNAs used in the selection were designed based on the previous model (Fig. 8B). When a substrate RNA with complete Watson-Crick base pairs in the extended P1 was used for the cleavage reaction, the reaction of the wild-type ribozyme was biphasic (an initial fast and following slow phase) (20).

View larger version (13K): [in this window] [in a new window]   FIG. 8. Model of the P1-P2 structure. A, the refined model (wild-type) structure of P1-P2. B, a predicted structure of the modified P1-P2 with the P2 mutations based on the originally predicted model. The nucleotides involved in the rearrangement and the mutated sites are highlighted by blue boxes and gray circles, respectively. The consensus mutations are shown with red letters.

	DISCUSSION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
The essential elements required for catalysis in Group I introns are concentrated in P3-P7 domain of the conserved core region, which also contains the P4-P6 domain. Previously, we reported the in vitro selection of modules residing at L7.1, L8, or L9 of a mutant T4 td ribozyme (M1 mutant) lacking the P4-P6 domain that is known to assist the functioning of P3-P7. The variants containing the modules performed the reversal reaction of the first step of self-splicing efficiently. No enhancement, however, was observed for the primary rate constant (k_cat value). In this study our new method of in vitro selection by employing the pools at the peripheral region of the ribozyme successfully provided modules that contribute to giving broad substrate specificity and high k_cat values. The result indicates that it is possible to construct a variety of modules that play a role comparable with that of P4-P6 at the core region of a Group I intron ribozyme.

The variants were highly active at a concentration of 10 mM magnesium ions due to the structural stabilization of the inserted modules, as shown by hydroxyl radical footprinting. The higher k_cat values of the variants suggest that the modules enhance the catalysis, such as the P4-P6 domain, which moderately contributes to the catalysis by using its L4 loop (21, 22). We did not observe any structural resemblance between the inserted modules and the P4-P6 domain, suggesting that they function differently, although it can be envisaged that an arbitrary module packs into the position, where the P4-P6 domain is located, to function like the P4-P6 domain.

The variants so far examined could not perform the intact splicing reaction due to their inability to perform the second step reaction that is chemically equivalent to the reversal reaction of the first step (data not shown) (24, 25). This can be explained by postulating that the loss of the second step activity is due to the loss of the ability to align the substrate at the catalytic site and that the P4-P6 element somehow conducts a smooth transition from the first to second step (26). The selected modules may not assist or allow the orientation of the 3' end of the ribozyme to be set properly in the substrate helix, called P10, that is responsible for conducting the second step (2). For example, the base pairings between the inserted module in clone 204 and L1 should prevent P10 formation (Fig. 6). Furthermore, some variants, including clone 107, could not conduct the first step reaction presumably due to the weak affinity to the guanosine cofactor (data not shown). Consequently, the modules obtained by the selection method employing either first- or second-step reactions may not allow a smooth transition between the two steps; thus, a selection procedure based on the splicing activity may be required to select a module whose function is completely equivalent to that of the P4-P6 element.

These results may be clues for understanding the origin of Group I intron ribozymes. The ribozyme can perform the reaction in a similar way to the reversal reaction of the first step by using a dinucleotide, GpN, as a substrate (27). The repetition of the reaction that adds an N at the 3' end of the substrate on the guide sequence is a template-directed RNA polymerization. Thus, it has been proposed that the ancestral form of Group I intron ribozymes might be an RNA polymerase (27). The role of the P4-P6 element that switches the first and second step in the splicing is not required for conducting this reaction. Our results show that a variety of modules that enhance the activity of the ribozyme without the P4-P6 element can be obtained, suggesting that a primitive RNA containing only P3-P7 element and its fairly simple helper module may be sufficient to compose a primordial RNA polymerase. The P4-P6 element might have evolved from such a helper module for composing the highly sophisticated self-splicing intron RNA.

	FOOTNOTES

 
^* This work was supported by grants-in-aid for Scientific Research on Priority Areas (to T. I.), the Encouragement of Young Scientists (to Y. I.) from the Ministry of Education, Science, Sports, and Culture, Japan, and Takeda Science Foundation (to T. I.). 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.

The on-line version of this article (available at http://www.jbc.org) contains Supplemental Figs. 1-5.

^¶ To whom correspondence should be addressed. E-mail: tan{at}kuchem.kyoto-u.ac.jp.

¹ The abbreviations used are: MOPS, 4-morpholinepropanesulfonic acid; FE, final extent.

	ACKNOWLEDGMENTS

 
We thank members of the Inoue Laboratory for helpful advice and comments.

	REFERENCES

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 

1. Cech, T. R. (1990) Annu. Rev. Biochem. 59, 543-568[CrossRef][Medline] [Order article via Infotrieve]
2. Jaeger, L., Michel, F., and Westhof, E. (1996) in Catalytic RNA (Eckstein, F., and Lilley, D. M. J., eds) pp. 33-51, Springer-Verlag, Berlin
3. Cech, T. R., and Golden, B. L. (1998) in The RNA World (Gesteland, R. F., Cech, T. R., and Atkins, J. F., eds) Vol. 2, pp. 321-349, Cold Spring Harbor Laboratory, Cold Spring Harbor, NY
4. Michel, F., and Westhof, E. (1990) J. Mol. Biol. 216, 585-610[CrossRef][Medline] [Order article via Infotrieve]
5. Golden, B. L., Gooding, A. R., Podell, E. R., and Cech, T. R. (1998) Science 282, 259-263[Abstract/Free Full Text]
6. Waring, R. B. (1989) Nucleic Acids Res. 17, 10281-10293[Abstract/Free Full Text]
7. Christian, E. L., and Yarus, M. (1992) J. Mol. Biol. 228, 743-758[CrossRef][Medline] [Order article via Infotrieve]
8. Christian, E. L., and Yarus, M. (1993) Biochemistry 32, 4475-4480[CrossRef][Medline] [Order article via Infotrieve]
9. Streicher, B., Westhof, E., and Schroeder, R. (1996) EMBO J. 15, 2556-2564[Medline] [Order article via Infotrieve]
10. Berens, C., Streicher, B., Schroeder, R., and Hillen, W. (1998) Chem. Biol. (Lond.) 5, 163-175
11. Ortoleva-Donnelly, L., Szewczak, A. A., Gutell, R. R., and Strobel, S. A. (1998) RNA (N. Y.) 4, 498-519
12. Pichler, A., Berens, C., and Schroeder, R. (2000) in Ribozyme Biochemistry and Biotechnology (Krupp, G., and Gaur, R. K., eds) pp. 7-25, Eaton Publishing, Natick, MA
13. Ikawa, Y., Shiraishi, H., and Inoue, T. (2000) Nat. Struct. Biol. 7, 1032-1035[CrossRef][Medline] [Order article via Infotrieve]
14. Ohuchi, S. J., Ikawa, Y., Shiraishi, H., and Inoue, T. (2002) Nucleic Acids Res. 30, 3473-3480[Abstract/Free Full Text]
15. Green, R., and Szostak, J. W. (1992) Science 258, 1910-1915[Abstract/Free Full Text]
16. Green, R. Ellington, A. D., and Szostak, J. W. (1990) Nature 347, 406-408[CrossRef][Medline] [Order article via Infotrieve]
17. Celander, D. W. (2000) in Current Protocols in Nucleic Acid Chemistry (Beaucage, S. L., Bergstrom, D. E., Glick, G. D., Jones, R. A., eds) pp. 6.5.1-6.5.6, John Wiley & Sons, Inc., New York
18. Zuker, M., Mathews, D. H., and Turner, D. H. (1999) in RNA Biochemistry and Biotechnology (Barciszewski, J., and Clark, B. F. C., eds) pp. 11-43, Kluwer Academic Publishers, Norwell, MA
19. Mathews, D. H., Sabina, J., Zuker, M., and Turner, D. H. (1999) J. Mol. Biol. 288, 911-940[CrossRef][Medline] [Order article via Infotrieve]
20. Pichler, A., and Schroeder, R. (2002) J. Biol. Chem. 277, 17987-17993[Abstract/Free Full Text]
21. Strobel, S. A., and Ortoleva-Donnelly, L. (1999) Chem. Biol. (Lond.) 6, 153-165
22. Szewczak, A. A., Kosek, A. B., Piccirilli, J. A., and Strobel, S. A. (2002) Biochemistry 41, 516-525
23. Deleted in proof
24. Suh, E., and Waring, R. B. (1992) Nucleic Acids Res. 20, 6303-6309; Correction (1993) Nucleic Acids Res. 21, 1054[Abstract/Free Full Text]
25. Kay, P., and Inoue, T. (1987) Nature 327, 343-346[CrossRef][Medline] [Order article via Infotrieve]
26. Ikawa, Y., Yoshioka, W., Ohki, Y., Shiraishi, H., and Inoue, T. (2001) Genes Cells 6, 411-420[Abstract]
27. Doudona, J. A., and Szostak, J. W. (1989) Nature 339, 519-522[CrossRef][Medline] [Order article via Infotrieve]
28. Heuer, T. S., Chandry, P. S., Belfort, M., Celander, D. W., and Cech, T. R. (1991) Proc. Natl. Acad. Sci. U. S. A. 88, 11105-11109[Abstract/Free Full Text]

CiteULike

Complore

Connotea

Del.icio.us

Digg

Reddit

Technorati

This article has been cited by other articles:

				Y. Ikawa, T. Shiohara, S. Ohuchi, and T. Inoue Concerted Effects of Two Activator Modules on the Group I Ribozyme Reaction J. Biochem., April 1, 2009; 145(4): 429 - 435. [Abstract] [Full Text] [PDF]

				N. Kim, H. H. Gan, and T. Schlick A computational proposal for designing structured RNA pools for in vitro selection of RNAs RNA, April 1, 2007; 13(4): 478 - 492. [Abstract] [Full Text] [PDF]

				J. GEVERTZ, H. H. GAN, and T. SCHLICK In vitro RNA random pools are not structurally diverse: A computational analysis RNA, June 1, 2005; 11(6): 853 - 863. [Abstract] [Full Text] [PDF]

This Article Abstract Full Text (PDF) All Versions of this Article: 279/1/540    most recent M305499200v1 Submit a Letter to Editor Alert me when this article is cited Alert me when eLetters are posted Alert me if a correction is posted Citation Map Services Email this article to a friend Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Request Permissions Citing Articles Citing Articles via HighWire Citing Articles via Google Scholar Google Scholar Articles by Ohuchi, S. J. Articles by Inoue, T. Search for Related Content PubMed PubMed Citation Articles by Ohuchi, S. J. Articles by Inoue, T. Social Bookmarking                      What's this?