2’-F-Arabinonucleic Acid (2’-F-ANA) Phosphoramidites

Arabinonucleosides are epimers of ribonucleosides with the chiral switch being at the 2’ position of the sugar residue.   2’-F-ANA adopts a more DNA-like B-type helix conformation, not through the typical C2’-endo conformation but, rather, through an unusual O4’-endo (east) pucker.  However, the presence of the electronegative fluorine leads to a still significant increase (ΔT_m1.2° C/mod) in melting temperature per modification.¹  2’-F-ANA-containing oligonucleotides exhibit very high binding specificity to their targets.  Indeed, a single mismatch in a 2’-F-ANA – RNA duplex leads to a ΔT_m of -7.2 °C and in a 2’-F-ANA - DNA duplex a ΔT_m of -3.9 °C.²   

The presence of fluorine at the 2’ position in 2’-F-ANA leads to increased stability to hydrolysis under basic conditions relative to RNA and even 2’-F-RNA.^1,3   The stability of 2’-F-ANA to nucleases also makes this a useful modification for enhancing the stability of oligonucleotides in biological environments.²  2’-F-ANA hybridizes strongly to target RNA and, unlike most 2’ modifications, induces cleavage of the target by RNase H.  Phosphorothioate (PS) 2’-F-ANA is routinely used in these applications due to its increased nuclease resistance.  Alternating 2’-F-ANA and DNA units provide among the highest potency RNase H-activating oligomers.  Both the “altimer” and “gapmer” strand architectures consistently outperform PS-DNA and DNA/RNA gapmers.⁴  

siRNA oligos were found to tolerate the presence of 2’-F-ANA linkages very well.  High potency gene silencing was demonstrated⁵ with siRNA chimeras containing 2’-F-RNA and/or LNA and 2’-F-ANA.  The high efficacy of these chimeras was attributed to the combination of the rigid RNA-like properties of 2’-F-RNA and LNA with the DNA-like properties of 2’-F-ANA.

References

1. E. Viazovkina, M.M. Mangos, M.I. Elzagheid, and M.J. Damha, Curr Protoc Nucleic Acid Chem, 2002, Chapter 4, Unit 4 15.

2. J.K. Watts, and M.J. Damha, Can. J. Chem., 2008, 86, 641-656.

3. J.K. Watts, A. Katolik, J. Viladoms, and M.J. Damha, Org Biomol Chem, 2009, 7, 1904-10.

4. A. Kalota, et al., Nucleic Acids Res., 2006, 34, 451. 

5. G.F. Deleavey, et al., Nucleic Acids Res., 2010, 38, 4547-4557, J.K. Watts, et al., Nucleic Acids Res., 2007, 35, 1441-1451, T. Dowler, et al., Nucleic Acids Res., 2006, 34, 1669-1675.