Nucleic Acid Triple Helices: Stability Effects of ...

Current Organic Chemistry, 2002, 6, 1333-1368

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Nucleic Acid Triple Helices: Stability Effects of Nucleobase Modifications Jordi Robles 1, Anna Grandas1, Enrique Pedroso1, F. Javier Luque2, Ramón Eritja3 and Modesto Orozco4* 1

Departament de Química Orgànica. Facultat de Química. Universitat de Barcelona. Martí i Franquès 1. Barcelona 08028. Spain 2

Departament de Fisicoquímica. Facultat de Farmàcia. Universitat de Barcelona. Avgda Diagonal sn. Barcelona 08028. Spain 3

Instituto de Biologia Molecular de Barcelona. CSIC. Jordi Girona 18-26. Barcelona 08034. Spain

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Departament de Bioquímica i Biologia Molecular. Facultat de Química. Universitat de Barcelona. Martí i Franquès 1. Barcelona 08028. Spain Abstract: In the last years DNA triplexes have been the subject of a very intense research effort, which has been summarized in a vast number of books and reviews (see for example refs. [1-16]). After a general introduction to DNA polymorphism and the structure of triple helices, we will examine here general approaches to the stabilization of triple helices. Methods based in external binding to DNA and backbone modification will be briefly summarized, and the reader will be addressed to more specific reviews. Finally, we will survey strategies for triplex stabilization based on modification of nucleobases, paying particular attention to recent advances in the field.

DNA: A POLYMORPHIC MACROMOLECULE DNA is highly flexible and can adopt a large variety of helical structures depending on the sequence, ionic environment, temperature, solvent, or the presence of ligands [15-28]. Some transitions between different DNA forms occur under physiological conditions and can largely influence the functionality of the DNA [15-28]. The most relevant biological form is the B-DNA duplex [29,30; see Figure 1], which is an antiparallel right-handed double helix with a 10 bp/turn periodicity similar to the structure early suggested by Watson and Crick [31] in the middle of the past century. The same general structure has been found in different DNA crystals [25,30,32-34] and in solution structures as determined by NMR techniques [3436] or by molecular dynamics simulations [38-41] in nearphysiological conditions. Since the sixties modified versions of the B-DNA helix like the D, T, or C forms have been reported in both synthetic and natural DNAs [15,26,42-45], and new modified forms are still being discovered [46]. The A-DNA duplex [see Figure 1] is an antiparallel righthanded helix with a 11 bp/turn periodicity [15,26,43,46-51]. It is the most stable conformation of RNA and DNA-RNA hybrids in physiological conditions [15,17-20,51,52]. This structure is adopted by the DNA only in non-physiological (low hydration) conditions, and typically in sequences rich in d(G·C) steps [15,17-20,26]. The transition between B and A

*Address correspondence to this author at the Molecular Modelling and Bioinformatics Unit, Institut de Recerca Biomedica, Parc Cientific de Barcelona, Baldiri i Reixac 1-5, Barcelona 08028, Spain; Tel: 93– 4037156; Fax: 93– 437157; E mail: [email protected] 1385-2728/02 $35.00+.00

forms of DNA has been found experimentally by changing the solvent composition [15,17-20,34], and theoretically by altering artificially the conformational preference of the 2’deoxyribose moiety from C3’-endo to C2’-endo [41]. Intermediate forms between A and B structures [the E-form) have been recently described in crystal structures of modified DNAs rich in d(G·C) steps [53,54], suggesting that there is a continuum ensemble of structures between the canonical A and B forms [53-55]. Other antiparallel helical DNA duplexes have been experimentally found. A left-handed helix (the Z-form) was characterized by CD, fiber diffraction, X-ray and NMR experiments [15,17,56-61] for sequences rich in d(G·C) steps, specially upon methylation of cytosines [15,17,56-61]. This left-handed helix and its transition to the B-DNA form have been examined by means of theoretical methods [6264]. More recently, other antiparallel DNA helical structures have been described for repetitive sequences based on nonWatson-Crick pairings (see Figure 2), like Hoogsteen or reverse Hoogsteen d(A-T) pairings [46,65], the latter involving adenosines in the syn conformation [65]. Parallel DNA duplexes (see Figure 3) are known since 1980 [66]. Though they were initially found in crystals of small pieces of intercalated DNA [66], both experimental and theoretical evidence have shown its existence in both hairpin and linear DNAs [67-83]. They involve either Hoogsteen or reverse Watson-Crick H-bond patterns (see Figure 2). The reverse Watson-Crick helix is preferred for d(A·T) sequences [66-72,78-81], while the Hoogsteen pairing is favored for d(G·C)-rich sequences, specially at acidic pHs, or in presence of 8-amino derivatives [74,82-88]. Though parallel helices are not fully characterized,

© 2002 Bentham Science Publishers Ltd.

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Fig. (1). Representation of B (left), A (middle) and Z (right) DNA structures.

Fig. (2). Schematic representation of the Watson-Crick and Hoogsteen (normal and reverse) pairings of A:T.

experimental and theoretical data [70,78-81] suggest that the general structure of a reverse Watson-Crick helix is not very different from that of the B-DNA. In constrat, the Hoogsteen helix seems to be more similar to the Hoogsteen subunit of a parallel DNA triplex (see above and refs. [82,85]). The DNA can also be found as multistranded structures like Holliday junctions [89], triplexes and quadruplexes [123]. Tetraplexes are formed following i) i-DNA and ii) G-

DNA motifs. The i-DNA (see Figure 4) is formed by polyd(C) sequences when two parallel d(C-C) duplexes are intercalated leading to “out of the plane” tetrameric structures [90-95]. The cytosine-cytosine pairing is stabilized by hemiprotonation of cytosines (see Figure 5), a type of interaction which also occurs in some poly(dC) or polyd(CT) duplexes [96]. The resulting tetraplex is rigid and stable even at neutral pH, as noted by both experimental data and theoretical studies [90-96]. The i-DNA might be a potential

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Fig. (3). Representation of poly d(A:T) parallel duplex in the reverse Watson-Crick (left) and Hoogsteen (right) conformations [see reference 81].

therapeutic target [97,98], because it can contribute to the stabilization of some telomeric sequences [90]. Because the G-DNA (see Figure 6) was found in G-rich sequences in telomeric regions at the 3’-end of chromosomes [99], it is an interesting target for the treatment of cancer and aging-related phatologies [97-102]. The G-DNA is defined by “in the plane” G-quartets, which are stabilized by double H-bonds (see Figure 7) and the interaction of the O6(G) [located in the interior of the helix) with small cations like Na+ or K+ [100-111]. The G-DNA can exist with parallel orientations of the four strands, as well as with different antiparallel arrangements [100-111]. Riboses are always in the C2’- endo region, and glycosidic bonds can be all in the anti conformation (parallel helix), or half in the anti and half in the syn conformation (antiparallel helix). In all cases the four-stranded helices are very stable provided counterions

are placed inside the central channel of the helix [100-111]. Interestingly, solution studies have shown that K+ is preferred over Na+ as stabilizing cation [102,104,105], but crystals of parallel G-DNA could be obtained only in presence of Na+ as counterion [111-113]. This suggests that the recognition properties of G-DNA can change depending on small structural features related to the helical topology [114].

TRIPLE HELICES The possibility that the DNA migth form triple helices was suggested in 1953 by Pauling and Core [115], and demonstrated four years later by Rich and coworkers [115]. Since then, triplexes have been largely investigated owing to their potential impact in biotechnological and biomedical

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Fig. (4). Representation of the crystal structure of the i-DNA (PDB code 1BQJ)

(not the pyrimidines) of a DNA duplex by H-bond interactions in the major groove of the duplex. Triplexes are traditionally classified in two main schemes derived from the orientation of the third strand with respect to the WatsonCrick duplex: i) parallel (H-DNA) and ii) antiparallel (H*DNA) triplexes. In the first case the third strand makes Hoogsten-type H-bonds with the Watson-Crick purine, while reverse Hoogsteen interactions occur in the antiparallel triplex (see Figure 8). Fig. (5). Schematic representation of an hemiprotonated cytosine:cytosine pair.

areas [1-16,117-129]. There is now a vast amount of theoretical and experimental information on the structure [116,130-149], stability [1-16,83,84,150-165], binding of drugs [1-16,146-149,166-169] or proteins [1-16,146149,170-173], or the possible formation of triplexes in the cell [1-16,129,174-177]. Intermolecular or intramolecular triplexes are formed when a third oligonucleotide strand interacts with the purines

Triplexes can be formed combining DNA, RNA or peptide nucleic acid (PNA) strands in almost all-possible combinations [23]. The stability of triplexes relative to the parent duplexes is determined by four main factors: i) the intrinsic stability of the nucleotides, ii) the repulsion between phosphates, iii) the stacking interactions, and iv) the Hoogsteen or reverse Hoogsteen H-bonds. Thus, triplexes are more stable when the building blocks (nucleotides) are not in unstable conformations, or minor tautomeric or ionic forms. Similarly, the stability of the triplex increases in an ionic environment able to screen the phosphate-phosphate



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Fig. (6). Representation of the crystal structure of the parallel G-DNA (PDB code 352D)

As noted by Mills et al [163], it is difficult to apply concepts of “stability” to structures whose relative folding free energies change with the solvent or the temperature. However, in general parallel triplexes are expected to be more stable [75,160] than the antiparallel ones, and are better known from a structural and functional point of view.

PARALLEL TRIPLEXES Parallel triplexes are characterized by a third strand rich in pyrimidines, Hoogsteen H-bonds, and parallel orientation of the third strand with respect to the central purine strand. Following Mills et al. [163], parallel triplexes can be classified depending on the nature of the triplet as: i) pure pyrimidine triplexes, ii) GT triplexes and iii) mixed TCG triplexes (see Figure 8). Fig. (7). Schematic representation of a G quartet.

repulsion between the duplex and the third strand. Finally, a correct pattern of H-bonds and stacking interactions is also necessary for the stabilization of the triplexes (see below).

The pure pyrimidine triplexes are based on the T-A·T and C+-G·C motifs (the first base refers to the Hoogsteen strand, and the symbols “dot“ and “dash -“ refers to Watson-Crick and non-Watson-Crick pairings, respectively). Two Hoogsteen H-bonds stabilize the interaction of T with A and

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Fig. (8). Different type of Hoogsteen (labelled in roman) and reverse Hoogsteen interactions (labelled in italics) in parallel and antiparallel triplexes. To avoid confusion between parallel and antiparallel trios of equal molecular composition antiparallel (reverse Hoogsteen) trios d(T-A·T) and d(G-G·C) trios are marked with “*”

C with G. The GT triplexes contain normal T-A·T triplets, but the C +-G·C motif is replaced by G-G·C triplets where the Hoogsteen guanine interacts through Hoogsteen H-bonds with the Watson-Crick guanine. Finally, the mixed TCG triplexes contain the three Hoogsteen motifs: T-A·T, C+-G·C and G-G·C triplets. Most of the structural information on parallel triplexes was obtained [1-16] for pure or almost-pure pyrimidine triplexes, which were the first ones to be discovered [115]. The first structural information on parallel triplexes was derived from fiber diffraction data by Arnott and coworkers [130]. Based on these data, the 3-D model for poly d(T-A·T) triplex suggest an A-type model, with a rise of 3.3 Å, a twist around 30 degrees, and all the sugars in the North C3’-endo conformation [130]. This structural model, which is still present in many textbooks, received some support from early NMR data [178], but later NMR and IR studies demonstrated that the A-model was not correct in aqueous solution [135,179,180]. A few high-resolution structures of triplexes coming from Patel’s, Feigon’s, and Lane’s groups are available in structural databases [23,132-141,156,181, 182-186]. Most of the data correspond to triplexes based in the pure pyrimidine motif, though some information on purine motifs and triplexes with non-homopurine tracks is available [181]. Since NMR structural parameters are sequence-dependent, it is difficult to obtain average parameters from a very scarce number of structures. For pyrimidine triplexes, however, though the general helical parameters derived from fiber diffraction data are not very far from the NMR values, it is clear that the shape of the helix predicted from fiber

diffraction and NMR experiments is different due to the different sugar puckerings. Thus, NMR data detect all the sugars in the South and South-East regions, as expected for a B-DNA [131-142]. A certain displacement to the South-East region is found in sugars of the Hoogsteen strand [23], but the North puckering were very minor, in disagreement with the fiber model of triplex DNA. Recently, state of the art molecular dynamics (MD) calculations showed that the B-type conformation (as defined from NMR and IR data in ref. [179] was more consistent than the A-type conformation for pyrimidine triplexes, at least in dilute aqueous solution [146-148]. Thus, MD simulations starting from A, B or P (the crystal structure of the PNA-DNA·PNA triplex [187]) trajectories converged quickly to an ensemble of structures very close to the Bform. The MD trajectories sampled structures very similar to those found in NMR experiments [see refs. 146-148 for discusion]. Thus, all sugars were found in the South and South-East regions [146], with a small displacement to the South East regions for the Hoogsteen pyrimidine strand, in agreement with NMR data [22,132-141,156,181,182-186]. A control simulation performed for a PNA-DNA·DNA triplex showed an excellent convergence to the crystal P-form [149], irrespective of the starting structure, confirming that the force-field and the protocol simulations were not biased to reproduce the NMR data. The poly d(T-A·T) triplex (see Figure 9) is a righthanded helix with the bases oriented perpendicular to the helical axis. The triplets are tightly hydrogen-bonded, as noted by NMR [23,132-141,156,181, 182-186], X-ray [131,187] and MD simulations [142-147,186-189], and there



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Fig. (9). Representation of poly d(T-A·T) triplex [from reference 146].

are excellent stacking interactions involving the Hoogsteen pyrimidines (see Figure 10). The third strand divides the major groove of the duplex in two asymmetric units, which following Shields et al. [146] are denoted as major part (MM) and minor part (mM) of the major groove (see Figure 11). The minor groove of the triplex is similar in size to that of a duplex, and MD simulations [146] show that there is a “spine of hydration” similar to that found in crystal structures of B-DNAs [25,32,191]. However, its shape is slightly modified from an “U” form to a “V” form (see Figure 12), which can explain why minor groove binders such as netropsin or dystamicin [167,168,178] bind to the triplex, but with less affinity than to a duplex. Though the mM groove is narrow, MD simulations [146] show that it can accommodate a water molecule or a small polar group, which agrees with water residence times detected from NMR experiments by Radhakrishnan and Patel [132]. This finding led to the suggestion that a small polar group attached to the position C8 of purines could stabilize the pyrimidine triplex [85,146,188-190].

Surprisingly, the third strand does not block the ability of the DNA to interact with large molecules [146,147]. In fact, the width of the MM groove is similar to that of a DNA duplex [NMR structures are typically too short for an accurate determination of the dimension of the MM groove), although the recognition properties have completely changed. Based on the large size and flexibility of the MM groove and the similarity between the minor groove in duplexes and triplexes, it was suggested [146] that DNA triplexes could interact with proteins that typically recognize DNA duplex. The GAGA binding protein, a Zn-finger protein which interacts with DNA duplex through minor and major groove contacts, was the first protein able to bind simultaneously and with sequence specificity to DNA duplex and triplex [170]. The major form of cytosine (neutral keto-amino) cannot interact with the guanine through double Hoogsteen-like Hbonds due to the lack of a donor at position N3. The stability of Hoogsteen G-C interactions can then be explained (see Figure 13) by [147] i) a wobble pair with the neutral keto-

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Fig. (10). Detail of stacking interactions in poly d(T-A·T) triplex.

than expected [148], the global interaction energy is similar to that of a Hoogsteen A-T pair in the gas phase [148,192]. Nevertheless, this pairing is handicapped by the fact that the imino tautomer is disfavored by 1.5 kcal/mol with respect to the amino form in the gas phase [194,195,196] and by 6-7 kcal/mol in aqueous solution [192-196].

Fig. (11). Groove definitions in parallel triplexes

amino cytosine, ii) a “Hoogsteen-like” interaction with the neutral-keto-imino tautomer of cytosine, and iii) a “Hoogsteen-like” interaction with the N3-protonated form of the keto-amino tautomer of cytosine. The wobble G-C pair implies an H-bond interaction between N4(C) and N7(G), and a favorable interaction between H8(G) and N3(C). In the gas phase its interaction energy is similar to that of a Hoogsteen A-T pair [149,150]. However, this pairing is not likely to occur due to bad stacking of the Hoogsteen cytosine, which has to be displaced to perform the wobble pairing [148]. Pairing with the imino cytosine tautomer implies two H-bonds, and even though the interaction between N4(C) and O6(G) is weaker

The last possibility, and the most likely one to occur, is the protonation of cytosine, leading to two very strong Hbonds in the gas phase [192]. Protonation of cytosine is disfavored in aqueous solution at neutral pH, since the pKa of cytosine is around 4.2 [1]. However, the negative charge of phosphates and the possibility to form a double H-bond with guanine lead to an apparent increase of the pKa of cytosine in the triplex [105,132-134,140,159,196]. NMR data and the pH-dependence of triplexes rich in d(C-G·C) trios support that the N3-protonated form of cytosine determines the stability of d(C-G·C) trios in purine strands of the type d(-GAGA-) or d(-AGGA-) [105,132134,140,159,196]. Theoretical studies indicate that for a d(CG·C) step surrounded by d(T-A·T) steps the presence of the protonated form of cytosine is favored by more than 2 kcal/mol with respect to that of the imino form [147]. Molecular dynamics calculations have shown that a triplex with a central purine d(-GAGA-)n is stable in aqueous solution if all the Hoogsteen cytosines are protonated, while it unfolds if they are in the imino form [147]. Similar results have been found for a triplex with a central purine track



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Fig. (12). Grooves in parallel triplexes compared with those of a standard B-DNA duplex. Left Figure shows the mM (TOP groove) and m (BOTTOM groove) grooves of a parallel triplex. Figure in the middle shows the major and minor grooves of a B-DNA duplex and Figure in the right illustrates the MM groove of a parallel triplex.

d(AGGA)n [189], confirming that the protonated form is responsible for the stabilization of d(C-G·C) steps in d(GAGA)-type triplexes. The role of Hoogsteen protonated cytosines is not so clear in triplexes containing three or more contiguous guanines in the same Watson-Crick strand. Molecular dynamics simulations have shown [147,148] that triplexes having three consecutive protonated cytosines in the Hoogsteen strand are unstable and unfold spontaneously at neutral pH and low ionic strength. This can be explained by the electrostatic repulsion between the Hoogsteen cytosines (C-C distance around 3.4 Å). This finding agrees with calorimetric and NMR studies [152,157] and suggest that in triplexes containing several contiguous guanines a small percentage of neutral Hoogsteen cytosines [probably in the imino form) should exist to screen the repulsion between protonated cytosines [147,148]. A fast interchange between neutral and protonated cytosine can easily occur due to the vicinity between the N3 and N4 atoms of two contiguous Hoogsteen cytosines [147]. The structure of triplexes containing d(C+-G·C) trios is not very different to that of a poly d(T-A·T) triplex [146148], as noted by the regularity along the sequence of d(GAGA)-type triplexes found in NMR studies [181]. The largest differences occur in the grooves, which are slightly narrower and less attractive for cations due to the presence of protonated cytosines. As noted above, no direct high-resolution structural data is available for GT and TCG triplexes. However, the fact that both triplexes contain a large percentage of pure pyrimidine trios, and that parallel triplexes can incorporate “anomalous” triads without dramatic distortions in the helix [197] suggest that their structural features are similar to those of pure pyrimidine triplexes. A high resolution crystal structure by

Fig. (13). Different H-bonding patterns for the stabilisation of Hoogsteen G-C interactions.

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Kenard’s group [131], containing isolated Hoogsteen-like d(G-G·C) trios confirms that the formation of this type of triplets is compatible with the general structure of a pure pyrimidine triplex.

ANTIPARALLEL TRIPLEXES Antiparallel triplexes (H*-DNA; 198-200) are characterized by a third strand rich in purines, reverse Hoogsteen H-bonds, and antiparallel orientation of the third strand with respect to the central purine strand. Following Mills et al. [163] triplexes can be classified depending on the nature of the triplets as: i) GA triplexes, ii) GT* triplexes, and iii) mixed GTA triplexes (see Figure 8). The GA triplexes are based on the G-G·C and A-A·T motifs, where two reverse Hoogsteen H-bonds stabilize the interactions between the purines. The GT* triplexes contains G-G·C and T-A·T triplets, like in a GT triplex (see above), but now stabilized by reverse Hoogsteen interactions. Finally, mixed GTA triplexes contain all type of reverse Hoogsteen triplets: G-G·C, A-A·T and T-A·T. Antiparallel triplexes are less stable than the corresponding parallel triplexes in near-physiological conditions [75,160,201,202]. They exhibit a large dependence on the counterion atmosphere, and typically high concentration of divalent ions are necessary for their stability [1,122,20,197-201,203,204]. This may reflect strong phosphate-phosphate repulsion [22] which are reduced by the screening effect of cations, or the need for specific N7ion contacts [204]. Interestingly, antiparallel triplexes have been more successful than parallel triplexes in anti-gene strategies [205], which has been related to their pHindependence. The amount of high-resolution experimental structural information on antiparallel triplexes is very limited. Only one structure solved by Patel’s group using NMR data [182] is found in the December-2001 release of PDB (pdb134d). The structure corresponds to a short intramolecular GT* triplex with reverse Hoogsteen d(G-G·C) and d(T-A·T) trios with sequence d(AGGAGGA) in the central purine strand. It is a regular right-handed helix with all the sugars in the South and South-East regions, with average rise and twist of 3.3 Å and 30 degrees. Other NMR data for a similar sequence [206] show helices in the B-family, but with very different twist, suggesting that the structure of antiparallel triplexes can largely depend on sequence or solvent composition. In fact, as noted by Lane and Jenkins [23], the different triplets that can be found in an antiparallel triplex are not fully isomorphous, which can also contribute to the reduced stability of these structures. The amount of theoretical data on antiparallel helices is also limited, and mostly correspond to the seminal work of Pettitt’s group on triplexes based on the reverse-Hoogsteen d(G-G·C) motif [144,145,207,208]. These studies, which provided interesting explanations on the role of solvent in the stabilization of the triplex later verified by NMR experiments [105], were however handicapped by the use of an old-generation force-field with a known A-philic tendency [105,207,208].

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STABILIZATION OF TRIPLE HELICES The biomedical use of triplexes is limited by several factors: i) the triplex-forming oligonucleotide (TFO) must pass through several biological membranes, ii) it must be resistant to degradation by cellular nucleases, iii) the triplex formation has to be very sequence-specific, and iv) the resulting triplex has to be stable, ideally even for nonhomopurine sequences. The strategies examined to date to increase the triplex stability rely on i) the addition of external compounds, and ii) the modification of the oligonucleotide. The final result should be a triplex stable under physiological conditions (low temperature, moderate ionic strength, neutral pH) irrespective of the sequence of the targeted duplex. Stabilization by Addition of External Compounds Several ligands bind to DNA triple helices and might then modulate their stability. They can be classified into three families: i) intercalators, ii) groove binders, and iii) crosslinkers. These molecules are often tethered to the TFO to increase the triplex stability, while keeping the sequence specificity [1,4]. Groove binders like netropsin, berenil, DAPI, Hoeschst 33258 or dystamicin [167,168,178,210] bind to the triplex, but with less affinity than to a duplex, leading to a net destabilization of the triplex and in a few cases to the inhibition of the triplex formation [211]. In other conditions, nevertheless, the triplex is stabilized. For instance, berenil, which destabilizes triplexes at 1 M NaCl, stabilizes them in the absence of salt [212,213], suggesting that different binding modes of the groove binder can affect the duplex triplex equilibrium. While berenil, netropsin or DAPI destabilize pure DNA triplexes in normal laboratory conditions, Pilch and Breslauer [210] have shown that they stabilize hybrid triplexes containing mixed DNA and RNA strands. Similar results have been recently obtained by the aminoglycoside drug neomycin [214,215]. Finally, as demonstrated by Robles and McLaughlin, when standard minor groove binders such as Hoescht 33258 or berenil are linked to the 5’-end of the TFO, the stability of the DNA triplex increases [216,217], which opens interesting possibilities for the triplex stabilization in antigene therapies. Intercalators are known to interact with triplexes since the sixties, when it was shown that ethidium bromide interacts with RNA triplexes [218]. Other studies [219] demonstrated that it also binds to poly dA-type triplexes, stabilizing them with respect to parent duplexes [219,220], a result also found for dye methylene blue [221]. Since then, many other intercalators have been studied, alone or tethered to TFOs, as triplex-stabilizing agents [1,4,210]. The benzopyridoindole familty developed by Héléne’s group [222-225] stabilizes specifically DNA triplexes. Similar results were found for benzopyridoquinoxaline [225] and other quinoline derivatives [226-228]. Sulfonamide derivatives of anthraquinones have interesting stabilizing properties for dA-type triplexes [229]. Related amidoanthraquinones were explored by Fox and coworkers [230,231], who found excellent stabilization of both parallel


and antiparallel triplexes. Hélène and coworkers showed that acridines tethered to TFO stabilize triplexes [232] by intercalation in the duplex/triplex junction, even though acridines alone do not show a dramatic stabilizing effect on triplexes [227]. Azobenzene [233] and naphtalene dimimide derivatives [234] also display excellent triplex-stabilizing properties when tethered to a TFO. Other aromatic systems such as quinoxalines, coraline, ellipticine and phenylimidazoles are able to intercalate into triplex DNA [4]. Despite the existence of a high resolution NMR structure of a triplex DNA bound to an intercalator [195], little is known about the physical nature of intercalation in triplexes. The excellent binding of intercalators might be related to the larger size of the π-system in triplexes compared to duplexes, which is expected to enlarge the favorable dispersion interactions with the drug. Apparently, in parallel triplexes most intercalators prefer adenine-rich to guaninerich sequences, in opposition to the behavior in duplexes, which is probably related to electrostatic repulsion generated by the protonated Hoogsteen cytosines in d(C-G·C) triads. Considering the impact of triplex intercalation in antigene therapy, a more intense theoretical and experimental effort seems necessary to understand the intercalation in triplexes. Finally, the use of crosslinking agents like psoralen tethered to TFOs have been analyzed to make permanent triplexes with antisense purposes [235-237], to design specific endonucleases [239,240], to introduce a mutation in a given position of the DNA [16,241-242], or to protect the DNA against UV-induced pyrimidine dimerization [124]. The possibility to photoactivate the crosslinking by irradiation with a suitable wavelength opens very important biomedical and biotechnological possibilities for these compounds.

Stabilization by Modification of the Oligonucleotides From a biomedical and biotechnological point of view, triplexes are useful to target endogenous (natural) nucleic acids. They can be formed at different levels: i) a single stranded TFO is added to target an endogenous DNA duplex, ii) a TFO is added to an endogenous single stranded RNA, and iii) a TFO is added to target a endogenous duplex DNA by means of strand-displacement mechanism. In all the cases, the TFO can be chemically-modified at either the backbone or nucleobase level in order to increase its affinity for the endogenous nucleic acid. Modifications in the backbone have been very popular, since they also permit to modulate the incorporation of the TFO to the cell and its resistance to nucleases [1,4,5,243]. Most backbone modifications were designed to stabilize DNA·RNA duplexes for antisense purposes (for a recent review see ref. [5], but a few of them have been applied to the triplex stabilization. Backbone modifications can be classified into three categories: i) sugar modification, ii) phosphate modification, and iii) full backbone replacement.


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Sugar Modification 2’-Deoxyribose has been extensively modified [1,4,5] i) to change the sugar conformation to a more triplex-like conformation, and ii) to reduce the entropic cost of TFO binding by restricting the sugar mobility. The first modification analyzed was to replace one or several of the RNA strand(s) by DNA [244-251]. The results are somehow confuse, and different well-known laboratories give different orders of relative stability for pure DNA and DNA-RNA hybrid triplexes [compare for instance results in refs. 244,248 and 250]. This indicates that, in addition to the type (parallel or antiparallel) of triplex and the sequence and length of the oligo, the relative stability of pure DNA and hybrid DNA-RNA triplexes also depend on subtle experimental conditions [245,251,252]. The structure of parallel triplexes containing RNA in the third strand (RNA-DNA·DNA triplexes) has been analyzed by experimental FTIR and NMR techniques [205,250,251] and molecular mechanics calculations [252]. FTIR experiments [205] suggested that the sugars of the purine DNA strand change from South to North puckerings. However, theoretical calculations [252] and NMR experiments [250,251] have shown that all the sugars of the DNA strands are in the South and South-East regions, and North puckerings are only found in the RNA strand. In fact, all NMR data suggests that the structure of the RNADNA·DNA triplex is in general very similar to that of a pure DNA triplex, except for some alterations in the grooves related to the presence of C3’-endo conformations [250,251]. A very successful modification of RNA strand for triplex stabilization was the methylation at O2’ [250,253,254]. Thus, triplexes having 2’-O-methylated DNA strands are very similar to normal DNA triplexes [250], but they are more stable than pure DNA and RNA-DNA·DNA triplexes [250,252]. Furthermore, 2’-O-methylated-TFOs are resistant to nucleases [4,254], which increases their potential biomedical and biotechnological possibilities. Recently, Torigoe and coworkers [255] have shown that very stable duplexes can be formed using 2’-4’ bridged nucleic acids (BNA; see Figure 14) in the TFO. The mechanisms of triplex stabilization by BNAs have not been fully clarified [255], but they are probably related to a rigidification of the TFO. A similar approach that eliminates the phosphate group is the riboacetal linkage (see Figure 14), where a methylene 2’-3’ linkage leads to an important stabilization of the triplex [256]. Phosphate Modifications Different changes in the phosphate moiety have enlarged the triplex stability and/or provided nuclease resistance. A very popular antisense strategy is to replace the phosphate group by phosphorothioate (Figure 14), which yields stable RNA·DNA duplexes and increased resistance to nucleases [257,258], while maintaining similar charge distribution. However, phosphorotioates are not very popular in antigene strategies since they lead to unstable antiparallel triplexes and avoid formation of parallel triplexes when present in the TFO [259-261]. Neutralization of the TFO phosphate

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Fig. (14). Different backbone subsitutions in the TFO

backbone has also been examined [256,262-265]. Thus, alkylphosphonate and methoxyethylamidate derivatives [262,263] were examined as neutral surrogates of the phosphate group (see Figure 14), but in general with only a moderate success [1,262,263], which indicates that the presence of a bulky group attached to the phosphorus atoms is in general unfavorable for triplex stability. More drastic substitutions of the entire phosphate by neutral groups, such as formacetal or thioformacetal (see Figure 14), were also analyzed [262,265], but without great success, which can be explained by the different geometry of these groups compared to that of the phosphate. Replacement of the O3’>P5’ phosphodiester linkage by a N3’->P5’ phosphoramidate linkage [see Figure 14 and refs. 266-269] was very successful, yielding very stable parallel triplexes in physiological conditions [266-269], probably due to a correct

preorganization of the TFO [264]. Bruice and coworkers [270-272] examined more drastic changes, such as the substitution of the entire phosphate group by cationic linkages like guanidinio or methylated thiorueas (see Figure 14). Instead of the usual repulsion, these changes revealed favorable backbone-backbone and stable triplexes [272,273]. Other authors [274-279] have analyzed aminoalkylfosforamidates substitutions, finding in some cases important stabilization of the triplexes [274-276]. Full backbone Replacement The replacement of the entire phosphoribose backbone by other polymerizing units has been largely analyzed for antisense purposes [5,126,280-285]. Morpholino derivatives (see Figure 14 and refs. [286-289]) were examined as



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Fig. (15). Neutral analogs that mimic protonated cytosine: (I) Pyrimidine analogs

antisense agents, but their profile as triplex-forming agents is not excellent [290,291]. Peptide nucleic acids (PNAs, see Figure 14) developed by Nielssen and cowokers were more successful. In this case the phosphoribose backbone is replaced by a peptide chain leading to extremely stable duplexes with both DNA and RNA [292-295], and have excellent antisense properties [285,295]. Furthermore, the PNA form very strong triplexes of different stoichiometry, it being able to displace one strand of stable DNA duplexes [296-302]. These features, in conjunction with the nuclease and protease resistance of PNAs, make this molecule to be an excellent candidate for antigene purposes [126,272-279]. A large amount of theoretical [149,303-306] and experimental work [302,307-309] has been focused on the structural characteristics of hybrids of peptide nucleic acids and DNAs, including triple helical structures [302,306]. The PNA backbone seems very flexible in some regions, while more rigid in others [306], inducing changes in the DNA backbone which makes all hybrid structures containing PNA more similar to A-type conformations than to B-type structures [149,302-309]. The lack of charge in the backbone and the special plasticity of PNA can be responsible for the high stability of complexes involving PNA. Some recent approaches to improve PNA-technology (limited often by solubility or poor cellular-uptake) rely on the use of DNAPNA chimeras [310], on the use of olefinic peptide nucleic acids (OPA, see Figure 14 and ref. [311], or oxy-PNAs [312], but none of these strategies has been able to improve the characteristics of normal PNAs.

[152,314,315]. The pKa value of the nucleoside [4.2] was described to be a little bit lower than that of the 3'- or 5'nucleotides [15], and to vary depending on the position of cytidine within oligonucleotide chains. pKa Values such as 4.5 [313], 5.5 [316], 7.0 [314], or even larger than 8.5 [152] have been described for the N3 of cytosine in polynucleotides. NMR experiments showed that the aparent pKa of the nucleobase may go up to about 9.5 when it's placed in an internal position, but down to 6-7 for terminal cytidines [196]. Not unexpectedly, cytosine protonation becomes more troublesome when the third strand has to interact with a duplex containing G tracts. As previously discussed in the first part of this review, these structures have been described to be less stable than those with an (A/G)-mixed purine central strand [315,317-320].

MODIFICATION OF THE NUCLEOBASES

Neutral analogs that Mimic Protonated Cytosine

1. Base Analogs Aimed at Stabilizing Parallel Triple Helices.

Neutral analogs that reproduce the double donor hydrogen bond pattern of protonated cytosine have been designed as derivatives of either the pyrimidine (Figure 15) or the purine rings (Figure 16). The first group of nucleobases should, in principle, give rise to the most stable triplexes, since they cause little or no distortion in the sugarphosphate backbone. In the second group of analogs, the presence of a purine system in the pyrimidine-rich Hoogsteen strand separates the anomeric carbon from the

As indicated above, the main problem associated with the formation of parallel triple helices (also referred to as the pyrimidine motif) is the protonation of C residues in the Hoogsteen strand. Many authors have stated that C+-G·C triplets are more stable than T-A·T triads [75,153,313], but, as expected, their stability varies depending on the pH

Consequently, with this scenario many of the efforts of the synthetic chemists have been addressed at obtaining either: 1) neutral C analogs mimicking protonated cytosine, with two hydrogen bond donor groups, in order to achieve pH-independent triplex formation; or 2) more basic C analogs, since greater ease of protonation should correlate with higher triplex stability. We will also review thymine and other cytosine modifications that yield more stable triplexes, either by hydrophobic effects, increased stacking or from additional interactions with the phosphates. 5Methylcytosine (MeC), which was one of the first bases shown to stabilize triplexes and often used as reference nucleobase, is included in this group.

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Fig. (16). Neutral analogs that mimic protonated cytosine: (II) Purine analogs

Watson-Crick duplex, breaking the isomorphism of the triple helix. Even though this results in structurally less homogeneous triplex systems, stabilizing effects have been observed.

strands to get information on the effect of positive charges (due only to the presence of the lysine residues) on the kinetics of the mechanism of strand displacement by positively-charged PNAs [325].

Pseudoisocytosine and Analogs

At pH=4.5, pseudoisocytosine formed triplets less stable than protonated cytosine [323]. The stability of ΨC-G·C triplets was shown to slightly increase as the pH decreased, which was related to the fact that at acidic pH the base is protonated, the tautomeric equilibrium no longer exists, and triplex formation is not penalized by the choice of the appropriate tautomeric form [323].

Pseudoisocytosine (ΨC, Figure 15a) was obtained from pseudouridine [321-323]. Several pseudoisocytosine derivatives have been prepared and introduced in oligonucleotide analogs: the 2'-deoxynucleoside [323], the ribo- and 2'-Omethylribonucleosides [321-323], and the PNA derivative [324]. Pseudoisocytosine replaced protonated cytosine either to recognize isolated G·C base pairs [321] or to form triplexes with (G·C)n segments [322]. UV and CD thermal denaturation studies indicated that, in all cases, a triple helix was formed, whereas no triple helix was detected using MeC at pH=7. The 2'-O-methyl-ΨC ribonucleoside formed more stable triplexes than the 2'-deoxy derivative [323]. In one of the thermal denaturation experiments [321], two transitions were observed at the typical triplex melting temperature, which were ascribed to the presence of the two possible pseudoisocytosine tautomers (N1-H or N3-H). The formation of (ΨC-containing PNA)-DNA·PNA triplexes has been shown to take place at pH values ranging from 5 to 9. A very small decrease in the melting temperature was found upon increasing the pH from 5 to 9 (∆Tm=-6.5 oC), as compared with the cytosine-containing PNA triplex (∆Tm=-30 oC) [324]. The stabilizing effect of pseudoisocytosine was high enough so as to allow for the formation of intramolecular triplexes with only two bases in the loops, whose stability was not dependent on the presence of magnesium [323]. ΨC-Containing PNAs linked to different numbers of lysines have been used as Hoogsteen

With the idea of having only the ΨC tautomer with the desired two hydrogen bond donor pattern, the 2'-Omethylribo pyrazine analog was obtained (Figure 15b) [326]. When introduced in a single position of the third strand sequence, this nucleobase analog was shown to form triplexes whose stability did not change with the pH, and more stable than those formed by C at pH=8. 6-Oxo-5-Alkylcytosines 6-Oxocytosines have been obtained either unsubstituted at the 5-position (C oxo) or substituted with methyl (CMeoxo) or allyl groups [327-331] (Figure 15c). Subsequent glycosilation reactions afforded the corresponding 2'-deoxy[327,329,331], ribo- [331] or 2'-O-methylribonucleosides [328,331], as well as a glycerol derivative [330]. The C oxo and CMeoxo-containing 2'-deoxyoligonucleotides [327,329,331] have been shown to give triple helices more stable at basic pH (pH>7) than those formed by oligonucleotides containing either C or MeC, with melting temperatures remaining essentially unchanged at pH values ranging between 6 and 8. The 5-allyl-6-oxocytosine analog did not form stable triplexes [331]. Triplets containing the

Nucleic Acid Triple Helices: Stability Effects of Nucleobase Modifications oxo

Meoxo

C analog were more stable than those formed by C [328], suggesting that the methyl group at the 5 position might cause unfavorable steric interactions.

Under neutral or acidic conditions, C +-G·C and MeC+-G·C triplets were shown to be more stable than those including the 6-oxo analogs [327-331]. It has been argued that, under these conditions, the π systems of C and MeC are more extended than those of Coxo or CMeoxo, allowing for more stable triple helices as a result of greater stacking interactions with the neighboring nucleobases. It is not excluded that the carbonyl group at the 6 position prevents the nucleoside from adopting the anti conformation required to form the triplet. However, in the presence of spermine, these analogs formed triple helices more stable than MeC [329], and this effect was more pronounced at high pH (pH=8.5). Oligonucleotides containing various CMeoxo were shown to be able to form triple helices with (G·C) n sequences [329] in the presence of spermine, which was not the case when the Hoogsteen strand contained MeC. More stable triplexes were obtained when the third strand contained CMeoxo and MeC at every other positions. CMeoxo was linked to glycerol (aCMeoxo) [330] to minimize the steric interactions between the carbonyl at position 6 of the nucleobase and the sugar, since the acyclic sugar analog is more flexible than the ribose system. A DNA duplex containing the (G·C)5 sequence was shown to form a triplex with a third chain containing five aCMeoxo at these pairing positions, under conditions in which neither C nor MeC were able to yield a triple helix. The stability of this triplex did not change with the pH. The most stable triple helices were formed when the third chain contained alternate MeC and aCMeoxo. Ribonucleotides containing these base analogs did not form stable triple helices [331]. With respect to the 2'-Omethyloligoribonucleotides, moderately stable triplexes were formed only when a single Coxo analog was present [328]. Oligonucleotides either containing more nucleobase analogs or formed by mixed deoxyribose and ribose backbones yielded fairly unstable triplexes, and in some cases triplex formation was not even observed [331]. Other Monocyclic Analogs Other five-membered heterocyclic systems have been designed as neutral analogs of protonated cytosine (Figure 15d) [332]. So far, only the preparation of the deoxyribonucleosides has been described. 8-Oxoadenine Different groups have proposed to use 8-oxoadenine (oxoA, Figure 16a) [333-335] or its N2-methyl derivative [336] to replace protonated cytosine in C+-G·C triads. The nucleosides were obtained by modification of 2'deoxyadenosine, and the phosphoramidite derivative is commercially available. The syn conformation was predicted for adenosine analogs substituted at the 8 position [337], but NMR experiments showed that the preferred conformation


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was anti when they formed Watson-Crick pairings, and that the nucleobase existed as the keto tautomer [338]. Both melting denaturation experiments [333] and footprinting techniques [336] showed that oxoA formed oxoAG·C triplets with no pH dependence, interacting either with isolated G·C pairs or with (G·C)n tracts. Triple helices containing oxoA were more stable than those containing MeC at pH>7.4, but the situation was reversed below this pH value [333], probably because of the higher isomorphism of Me C-G·C triads. The circular dichroism spectra of triplexes containing oxoA were identical to those containing MeC [333], thus showing that both have similar structures. Even though oxo A is an adenosine analog, triplexes containing oxoA-A·T or oxo A-T·A triads were not formed [333]. Triple helix formation with the oxoA analog could not always be assessed by UV melting denaturation studies, since triplex to duplex transitions were not clearly detected [334]. Gel shift experiments showed that oxoA-containing oligonucleotides formed stable triplexes even at pH slightly above 7.4. The affinity of the modified oligonucleotide for the duplex was highest at pH=5.9, and this effect was related to the acidity of the N7 proton (pKa=8.7). Different biological experiments support the use of oxoA to replace C+. On the one hand, oxoA-containing oligonucleotides inhibited the in vitro transcription of RNA Pol II [139]. On the other, ribo- and 2'-O-methylribo oxoAmodified oligonucleotides were able to prevent an endonuclease from being synthesized [339]. N7-guanine and Analogs Nucleosides with a C1'-N7 guanine linkage (7G) (Figure 16b) were prepared by glycosylation of N2isobutyrylguanosine [340-342]. This nucleoside analog can replace C + in parallel triple helices. Its affinity for G·C base pairs has been shown to be virtually the same at pH 7.0 or 7.5, but to be sequence-dependent [341,342]. The affinity of Me C and 7G for single G·C base pairs was essentially the same, but the affinity of (MeC/T)-containing oligonucleotides for duplexes with G·C base pairs at every other position was three times higher than that of (7G/T)-oligonucleotides. Conversely, (7G/T)-oligonucleotides hybridized to duplexes with G·C tracts with an affinity five times higher than (MeC/T)-oligonucleotides. This sequence-dependent affinity was related to the lack of isomorphism with T-A·T triplets. NMR studies on an intramolecular triplex [137] showed that the conformation of 7 G in 7G-G·C triplets was different from that of C in C+-G·C triplets, which caused a substantial distortion in the overall regularity of the triplex structure. Triplex formation with a 7G acyclic glycerol derivative, a7G, was also examined (Figure 16b) [343]. Using this derivative, the stereochemistry of the ribose system is maintained, but the flexibility must be higher. Five contiguous a7G residues binding to a (G·C)5 tract were found to yield more stable triplexes than MeC, whereas an isolated a7G-G·C triad was found to be less effective.

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thermodynamic parameters calculated from the melting curves suggested that the β7H-U·A triplet might be stabilized by an additional and relatively unusual H bond between the C5-H of U and the carbonyl group at the 6 position of β7H. Further semiempyrical calculations indicated that such a hydrogen bond would be only 1.12 kcal.mol-1 less stable than a "typical" H bond. In addition, since the β7H-G·C triplet was, at pH=7, virtually as stable as the MeC-G·C triplet, the authors inferred that the β7H-G·C triplet might be stabilized by the (N3-H)-CO hydrogen bond, as expected, and two CHCO hydrogen bonds (Figure 17). Fig. (17). Structure proposed for the 7H-G.C triad

The capacity of pyrazole analogs of 7G (0P1 and P2, Figures 16c and 16d, respectively) [344,345] to mimic C+ was also evaluated. Footprinting experiments [344,345] showed that their affinity for G·C pairs was very different. The affinity of P2 was smaller than that of P1, probably as a result of the distortion caused by the position of the sugarnucleobase linkage. P1 recognized its target with an affinity similar to that of C, but had the advantage of forming triplets within a larger pH range, that is, at pH>7. Footprints with P1-containing oligonucleotides were observed at higher pH values than in the case of MeC-modified oligonucleotides. MeC had a fourfold higher affinity for alternate G·C sequences than P1, whereas oligonucleotides with P1 interacted with (G·C)5 sequences with higher affinity. NMR Structural studies of intramolecular triplexes containing one P1-G·C triplet in the middle of the Hoogsteen strand showed that, even though the P1-G·C triplet is not isomorphous with T-A·T, the distortion caused by P1 is relatively small [346]. The α− and β-N7-hypoxantine deoxynucleosides (α7H and β7H, respectively), as well as the β-ribo- (β7HOH) and β2'-O-methylribonucleosides (Figure 16e) have also been evaluated as proton donors able to interact with G·C pairs [347,348]. The stability of the triplets, and the affinity and selectivity of these analogs were assessed from both footprinting and UV thermal denaturation experiments. The α7H analog yielded triplexes less stable than MeC [347]. The β7H analog gave rise to certainly more stable triple helices, and its capacity to recognize G·C pairs, either in alternate or in adjacent G·C sequences, was also better [348]. The β-ribo isomers afforded slightly worse results. From theoretical calculations of the triplet structure [348], it was inferred that the triad was stabilized by a single [N3-H]- N7 hydrogen bond. The comparison beween the analog β7H, 7G and P1 indicated that the additional amine group present in the latter might cause some steric hindrance, which would prevent the hydrogen bond from being a key factor in the stabilization of the triplets. Subsequent experiments [349] showed that β7H was also able to form β7H-U·A triplets, but not β7H-T·A (melting temperatures of the triplex to duplex transitions: 40 oC for β7H-G·C, 30 oC for β7H-U·A, and 18 oC for β7H-T·A). The

It is finally to be mentioned that hypoxantine has been proposed as "universal base" for triplex formation [350). The negative aspect of extensively using this promiscuous nucleobase analog was that triple helices decreased in stability.

Cytosine analogs with increased basicity Aminopyridines Different C-nucleosides with 2-aminopyridine rings (P) (Figure 18a) have been prepared and evaluated as cytosine analogs with a higher pKa and the same pattern of hydrogen donor groups [351-354]: i) 2'-deoxy-5-(2-aminopyridine) nucleosides, either in the α− or β−configuration (αP and βP, respectively), ii) the 5-(2-amino-3-methylpyridine) β−2'deoxynucleoside (MeP), and iii) the 2'-O-methylribo-5-(2aminopyridine) derivative (POMe). The reported pKa values for the α and β anomers of the P nucleosides were 6.2 and 5.9, respectively [351]. At pH=7, the affinity for duplexes with G·C base pairs at every other position was fairly similar for both of the αP and βP analogs [351], and the slightly higher affinity shown by αP-oligonucleotides was related to the greater basicity of the nucleoside. Under the same conditions, neither C nor MeC formed a triple helix. Both αP- and βP-containing oligonucleotides formed the most stable triplexes at pH=5.5, and no triplex was formed at pH>8 [351]. In addition, the half life of mixed (T/αP) sequences in serum was higher than that of those containing T and βP. Footprinting experiments showed that the affinity for duplexes containing the (G·C)4 sequence decreased in the order βP>αP>C [352]. From molecular dynamics studies [352] it was inferred that the αP isomers would not cause a significant distorsion in the triplex structure as compared with the βP derivatives. Two hydrogen bonds between guanine and the P ring could be established for both of the isomers. The structure of the triple helix was shown to be more A- than B-DNA-like both for the αP and βP analogs. Thermal denaturation studies and footprinting experiments showed that oligonucleotides containing βP and Me P at every other position formed triple helices more stable than MeC in the 6 to 8 pH range [353,354]. The stability of triplexes with (G·C)n tracts was higher when MeP replaced Me C in the third chain (pH=6-8). POMe oligonucleotides were basically unable to form triple helices at pH above 6.4.



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Fig. (18). Cytosine analogs with increased basicity

None of these analogs gave rise to duplexes more stable than those formed by the natural nucleobases, from which the authors concluded that these nucleobases tend to form Hoogsteen rather than Watson-Crick hydrogen bonds [353,354]. Other authors have also shown that 2-aminopyridine can suitably replace protonated cytosine in triple helices [161]. The 2-aminopyrimidine analog (C1'-C5 linkage as well) destabilized triplexes under the same experimental conditions, as expected from the relative pKa values of the nucleobases (2-aminopyridine: 6.8, cytosine: 4.2, 2aminopyrimidine: 3.3). It is to be noticed that 2aminopyrimidine was able to recognize C·G pairs. 6-Aminocytosine This analog (Figure 18b), more basic than cytosine (pKa=6.8), was transformed into the nucleoside by a glycosilation reaction [355]. Footprinting experiments showed that oligonucleotides containing 6-aminocytosines at different positions were not able to form triple helices. It was inferred that the 6-amino group might change the conformation of the glycosidic bond, hindering the anti conformation required for triplex formation. 4-substituted Cytosine Analogs The 4-guanidine analog of cytosine (Figure 18c, R=R1=R2=R3=H) was initially devised as a nucleobase able to recognize G·C or C·G pairs in a "switch" mode [14, 356], depending on whether it was linked to the ribose in the α or β configuration [357]. The synthesis of 4-guanidinecytosine and 4-guanidine-5-methylcytosine (Figure 18c, R=CH3,

R1=R2=R3=H) deoxynucleoside analogs were described [357]. 4-Guanidino-5-methylcytosine derivatives (Figure 18c, R=CH3) have also been prepared with the idea of reproducing the hydrogen bond pattern of C+ as well as getting protonated nucleobases at neutral pH. Preparation of the 2'-deoxyribonucleosides and their introduction in oligonucleotide chains was described [358]. The unsubstituted guanidine analog (R1=R2=R3=H) was relatively unstable to the synthesis and purification procedures, so only the methylsubstituted guanidine analogs were studied. Oligodeoxynucleotides containing one or three non contiguous 5-methylcytosine analogs with either methyl(R3=CH3, R1=R2=H) or dimethylguanidine (R1=R2=CH3, R3=H; R1=H, R2=R3=CH3) groups at the 4-position were examined as the Hoogsteen strand in parallel triplexes (pH=6.5, 7 and 7.5) [359]. No triplex formation was found by UV thermal denaturation studies, whereas triple helices were formed with the C- or MeC-containing oligonucleotides. CD spectra lead to the same conclusion: the CD of the (modified oligonucleotide)-duplex mixture (pH=6.5) corresponded to the sum of the spectra of the different components, while different bands were observed when unmodified oligonucleotide was used instead. Even though the 4-guanidine substituents should be able to form two hydrogen bonds with guanine, as shown by molecular modelling inspection, the (4-guanidino-5-methyl)-G·C triad is not isomorphous with C+-G·C or T-A·T. The distortion caused in the structure of the helix would only allow for weak stacking interactions, which might account for the

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Fig. (19). Pyrimidine analogs with other stabilizing effects: (I) 5-Methylcytosines and derivatives

inability of guanidine-modified oligonucleotides to form triplexes. Two additional neutral cytosine derivatives, closely resembling the 4-guanidinocytosine analog, were designed to have similar binding properties and will be discussed here. Nucleobases with either two hydrogen bond donor groups (X, Figure 18d), or one acceptor and one donor groups (Z, Figure 18e), were prepared and introduced in oligonucleotides [360,361]. The α anomers were expected to interact with the purines of the central strand, giving rise to Xα-G·C or Zα-A·T triplets. Conversely, the β nucleosides would interact with the purines of the opposite strand of the duplex, in Xβ-C·G or Zβ-T·A triplets. Footprinting experiments were carried out to assess whether X- and Zcontaining oligonucleotides were able to recognize duplexes, with contradictory results. The nuclease S1 treatment indicated that one of these modified oligonucleotides was able to interact with a duplex and protect it from being degraded, while DNase 1 digestion suggested that no triplex had been formed.

PYRIMIDINE ANALOGS WITH OTHER STABILIZING EFFECTS 5-Methylcytosine and Derivatives 5-Methylcytosine ( MeC, Figure 19a) has been substituted for cytosine in the third strand of triple helices for many years. More stable triplexes were found when either a single or a few cytosines were replaced by MeC [362-365]. On the contrary, UV and CD thermal denaturation studies showed that oligonucleotides containing MeC interacted inefficiently with duplexes having G·C tracts [364,365]. MeC-containing 2'-O-methyloligoribonucleotides formed triplexes less stable than C-containing oligodeoxynucleotides [367], and replacing C by MeC in PNA-DNA·PNA triplexes also had a destabilizing effect [368]. Many comparisons between MeC and C have been carried out [322,336,342,345], and, as stated before, the effectiveness of new nucleobase analogs is usually compared with that of MeC. The stabilizing effect of MeC has been related to an increase in entropy, upon binding to the duplex, due to desolvation of the helix [365], and to stronger stacking

interactions [369]. Even though the pKa of MeC (4.4) is only 0.2 units higher than that of C, the pH range of triplex formation with MeC is extended to pH values 0.5-1 units higher than with C-containing oligonucleotides [64]. The carbocyclic analog of MeC, cMeC (Figure 19a), was shown to be more basic (pKa: 4.8) than the 2'deoxyribonucleoside [370]. Nevertheless, replacement of C by cMeC was shown to destabilize parallel triple helices, as revealed by UV melting curves and footprinting experiments (∆Tm=-1.7 oC/substitution). The lower triplex forming efficiency of cMeC was attributed to changes in the cyclopentane conformation with respect to the ribose ring, since the X-ray structure of the nucleoside showed a C1'-exo pucker. This conformation, if maintained within the Hoogsteen strand, would be unfavorable for the formation of the triplex. The 5-methylcytosine ring has also been modified at the 4 position by addition of long, positively charged chains that may add favorable electrostatic interactions with the phosphate groups. The 4-amine group has been replaced by spermine (C sp) and other polyamines (Figure 19b) [371,372], or N-alkylated with an ω-deoxy-ω-aminotetraethyleneglycol chain (Cteg) (Figure 19b) [373]. The rationale for these modifications is that spermine and other biogenic polyamines are known to stabilize triple helices, at millimolar concentrations, even in the absence of Mg2+ and at neutral pH [374]. The higher the number of amine groups of the polyamine, the greater the stabilizing effect [spermine>spermidine>putrescine) [375]. Moreover, covalent linkage of spermine to the 5' end of an oligonucleotide has been shown to render more stable triplexes at pH=6.5 [376], making the addition of free polyamine or Mg2+ unnecessary for their formation. Oligonucleotides in which C was replaced by Csp formed parallel triple helices at pH=7.3 and in the absence of Mg2+, conditions in which neither C nor MeC formed the complex [371,372]. The effect of one spermine moiety at the 4position was similar to that of adding a 103-fold excess of free amine with respect to the oligonucleotide. The stability of C sp-containing triplexes was found to decrease as the pH decreased, which was ascribed to electrostatic repulsions between the protonated amine groups. No duplex stabilization has been found with Csp.



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2+

Fig. (20). Pyrimidine analogs with other stabilizing effects: (II) 5Halopyrimidines

The stability of triple helices diminished as the number of Csp increased, but it was still greater than when C was replaced by MeC [372]. Best results were obtained for oligonucleotides modified at the ends of the chain. Csp (pKa=3.7) is less basic than C, most probably as a result of the effect of several positive charges at the spermine chain, which lower the basicity of N3. In any case, the stabilizing effect of the positive charges of spermine compensate for the loss of one hydrogen bond in the formation of the triplets.

Mg , and melting profiles did not show the typical hysteresis curve [373]. The stabilizing effect of the tetraethyleneglycol chain, with a single positively charged group, was lower than that of spermine, and it was shown to be higher when Cteg was placed at the ends of the oligonucleotide sequence. Since Cteg was less charged than Csp, triplex stabilization could no longer be attributed, to the same extent, to the (protonated amine]-phosphate electrostatic interactions, but to other reasons such as hydrophobic effects related to desolvation of the grooves. It is to be mentioned that polyethyleneglycol had been shown to stabilize triple helices, the effect being greater when the molecular weight and the concentration of the additive increased [377]. Finally, for comparison purposes we would like to mention here that the α-anomer of the N4-spermine analog of cytosine has also been obtained [378]. The α-β·β triplexes were shown to be less stable when the α Hoogsteen chain contained the 4-modified cytosine derivative instead of C. 5-Halopyrimidines

The UV melting profile of triplexes with Csp at the third chain did not show the typical hysteresis curve [372]. This kind of profile is usually observed for triplex formation under conditions that strongly favor hybridization, such as high ionic force or the presence of polyamines or divalent cations, so the experimental data were interpreted as the result of an association process assisted by the phosphateamine electrostatic interactions.

The replacement of thymine or uracil by 5-bromouracil (BrU, Figure 20a) was shown to have the same positive outcome as the C by MeC substitution: triple helices were formed within a larger pH range [364]. Incorporation into the same chain of both BrU and MeC was described to have additive effects on the stability of the complex [364,379]. With respect to T or BrU, uracil analogs with iodine (IU) or amine groups at the 5 position had a negative effect on the stability of triple helices [380].

Many of the properties of the N4-(ω-deoxy-ωaminotetraethyleneglycol)-5-methylcytosine analog, Cteg, were common to Csp: duplex formation was destabilized, triplex formation took place at pH=7.3 in the absence of

Cytosine analogs with bromine ( BrC) or iodine ( IC) atoms at the 5 position (Figure 20b) have also been obtained. Triplex formation between a hairpin duplex and a third chain including these nucleobases was shown to be highly

Fig. (21). Pyrimidine analogs with other stabilizing effects: (III) 5-Propynylpyrimidines and other modifications at the 5 position

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Fig. (22). Triplex formation by cooperative binding of two oligonucleotides modified at the junction

unfavored [380]. These experimental results were in good agreement with those obtained by molecular dynamics calculations [381]. These studies suggested that replacement of C by BrC in the Hoogsteen strand had a destabilizing effect, while substitution of T by BrU in the duplex gave rise to more stable triplexes. This differential behavior was attributed to changes in stacking interactions. 5-Propynylpyrimidines and Analogs The methyl group of both thymine and 5-methylcytosine has been exchanged by a propynyl group (Figure 21a,b) [382]. The idea of introducing a propynyl group was to bring about hydrophobicity to the triplex complex, favoring both desolvation and stacking interactions. Positive results in parallel triplex formation were obtained when 5-propynyl-U replaced T (∆Tm=2.4 oC/modification), but not when 5propynyl-C replaced C (∆Tm=-3.4 oC/modification) [382]. 5-Propynyl-C was found to be less basic (pKa=3.30) than C, but triplexes formed with oligonucleotides containing 5propynyl-C were found to have the same pH sensitivity as the unmodified chains. The cooperative association of two short oligonucleotides with a DNA duplex, to yield a triplex, was certainly enhanced when 5-propynyl-U was substituted for T at the ends of both of the short oligonucleotides [383] (Figure 22). A positive but lower effect was observed when only the "3' chain" was modified. This was explained on the basis of favorable stacking interactions between the propynyl group and the terminal nucleobase of the adjacent chain end, which, in a right-handed helix, can only take place when the propynyl group is placed at the end of the "3' chain" [383]. NMR Studies of a parallel intramolecular triplex in which T was replaced by 5-propynyl-U showed that its structure closely resembled that of an unmodified triplex [186], with an A-DNA like conformation in some helical parameters (X displacement and inclination). Increased stacking between the nucleobases and higher hydrophobicity were ascribed to the presence of propynyl groups. The results from thermal denaturation experiments and gel shift assays have also been used to compare the effect of many different nucleobase and backbone modifications on oligonucleotides triplex forming ability [384]. The most stable parallel triple helices were those in which the third strand contained 5-propynyl-U or MeC. It is also worth mentioning here that the stabilizing effect that both 5-propynyl-U and 5-propynyl-C have on duplexes has been successfully exploited in antisense strategies.

Expression of the TAg gene in cell culture was inhibited by a phosphorothioate oligonucleotide containing these analogs [385]. The stability of triplexes containing 5-(aminopropargyl)uracil (Figure 21c) was shown to depend on the pH, as expected for nucleosides derivatized with amine groups [386]. The increase in stability was of about 12 oC per replaced T at the Hoogsteen strand. The contribution of the positively charged groups was ascertained by the fact that, at pH=9, when the aminopropargyl groups are no longer protonated, triple helices containing this analog were as stable as those containing 5-propynyl-U. Interestingly, triplex stability was shown not to decrease with Hoogsteen strands containing various aminopropargyl-uracils at adjacent positions, which was ascribed to favorable electrostatic interactions between the phosphates and the positively charged amines [386]. Quantitative footprinting experiments showed that the affinity for duplexes of (5-aminopropargyl-U)-modified oligonucleotides was considerably higher than that of the unmodified chains [386]. Triple helices with many C+-G·C triplets were even formed at pH=6 when T was replaced by 5-propargylamine-U in the third strand. Nucleosides with 2'-O-(2-aminoethyl)ribose (Figure 21c) have been shown to stabilize triple helices [387]. Hoogsteen strands containing doubly modified nucleosides, with 5aminopropargyl and 2'-aminoethoxy groups, strongly favored triplex formation [388]. With respect to unmodified Hoogsteen chains, the increase in Tm was of 25.8 oC for an oligonucleotide with two aminopropargyl-uracils, and of 48.4 oC for an oligonucleotide with two doubly-modified nucleosides, under the same experimental conditions. Additional stabilization of triplexes by the presence of two positive charges has also been searched for by preparing 5-modified uracil derivatives with other diaminoalkyl appendices (Figure 21c) [389]. No increase in stability was found in this case. Pyrimidines with other Modifications at the 5 Position Looking for the stabilizing effect of either positively charged groups or aromatic systems able to intercalate between base pairs, 5-carboxyuracil was modified by covalent linkage to diamines or anthraquinone derivatized systems (Figure 21d) [390]. Only the anthraquinonemodified bases, when placed at the 5' end of an oligonucleotide chain, showed a significant stabilizing effect on the stabilization of parallel triple helices at pH=7.



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Fig. (23). Polycyclic pyrimidine analogs: (I) Additional rings fused at positions 4 and 5 of pyrimidines

Fig. (24). Polycyclic pyrimidine analogs: (II) Additional rings fused at positions 5 and 6 of pyrimidines

Polycyclic Analogs Additional Rings Fused at Positions 4 and 5 of Pyrimidines The bicyclic system F (Figure 23a) has an extended π system which, in principle, should favor the stacking interactions with other nucleobases. F formed F-A·T triplets in parallel triple helices, and was able to recognize G and A in duplexes [391]. This behavior was explained on the basis of the tautomeric equilibrium between F1 and F2 (Figure 23a). F1 is the C-like tautomer, while F2 is T-like, as deduced from the hydrogen acceptor/donor pattern of each molecule. Footprinting analysis showed that the specificity of F for A·T pairs was the same as that of T, but its affinity was lower. Other authors have reached similar conclusions with Hoogsteen chains containing guanines and thymines [392]. It was also evaluated whether the cooperative union of two triplex forming oligonucleotides could be improved by replacing T by F at the neighboring 3' and 5' positions (Figure 22) [391], but the values of the association constant were of the same order in both cases. The PNA monomers of the bicyclic and tricyclic systems shown in Figures 23b-d (bT, tT and tC, respectively) were

prepared, and formation of the corresponding PNADNA·PNA triplexes was evaluated [393]. bT-Containing PNA Hoogsteen strands gave rise to triplexes more stable than PNA chains including T or ΨC (∆Tm=1.5 oC/modification], both when placed at neighboring and at every other positions. Third strands with (tT)T tracts destabilized the triplexes (∆Tm=-3.0 oC/modification), but a slightly stabilizing effect was observed for oligonucleotides with adjacent tT nucleobases. This effect might be related to increased stacking. Finally, replacement of C by tC was shown to destabilize triplexes at neutral pH. The effect was greater in sequences with tC at every other position (∆Tm=–2.7 oC/modification) than in sequences with contiguous tC (∆Tm=-1.0 oC/modification). The change in Tm upon varying the pH was greater than for cytosine. Even though the pKa value of the heterocycle was not determined, these results were correlated with a lower basicity of the nucleobase. Additional Rings Fused at Positions 5 and 6 of Pyrimidines Different 2,4-quinazolinedione systems (Q), with fused benzene-thymine rings [Figure 24a), have been examined as T analogs [394]. Both duplexes and triplexes were

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The fluorescence of 1 (Figure 24b) did not change with the pH within the range 5.0-8.0, but, upon protonation, the fluorescence of 3 decreased, and the emission maximum was shifted to longer wavelengths [398]. On the basis of this behavior, the authors proposed that fluorescence experiments could be used to probe the protonation state of 3-G·C triplets in triple helices.

2. BASE ANALOGS AIMED AT STABILIZING ANTIPARALLEL TRIPLE HELICES Fig. (25). Analogs of A aimed at stabilizing antiparallel triple helices

destabilized, slightly the former and significantly the latter. The analogs with halo- or nitro groups at the benzene ring (Figure 24a) did not afford better results [395]. The tricyclic 1 and 2 systems (Figure 24b and 24c, respectively) [396] destabilized duplexes (∆Tm≈-6 o C/modification). Parallel triplexes with one or two analogs, forming X-A·T triplets (X=1, 2), were slightly destabilized (∆Tm≈-2.5 oC/modification). This was interpreted as the result of increased stacking with the bases of the adjacent triplets, which might interfere with the regularity of the triple helix. The polycyclic system 1 renders fluorescent oligonucleotides, which allows parallel triplexes containing 1 at the Hoogsteen strand [or within the duplex) to be studied by fluorescence techniques. The analog 1 was also used to evaluate the affinity of a segment of the HIV Tat protein for a stem-loop segment of the TAR RNA [397]. The tricyclic system 3 (Figure 24c) is the fluorescent analog of cytosine (pKa=4) [398]. 2'-O-Methyloligoribonucleonucleotides containing 1 and 3 were hybridized with DNA·DNA or RNA·(2'-OMeRNA) duplexes. The resulting triplexes were found to be less stable than triplexes with (C/T)-containing Hoogsteen chains (∆Tm between -7 and 10 oC), and were not formed when 3 replaced C in the all RNA triplexes.

Adenine Analogs Several base analogs have been proposed to substitute adenine in triplex-forming oligonucleotides (TFO) to form antiparallel triplexes (Figure 25). Purine binds A·T base pair similarly to adenine but with one single hydrogen bond [399]. In addition purine was found to bind to C·G base pairs by formation of one H-bond to cytosine. 6-Alkyl-G was proposed as adenine analog but no data on triplex formation is available [400]. 2-Aminopurine was also proposed as substitute of adenine in antiparallel triplexes [401]. Recently, the ability to form antiparallel triplexes of four isomeric forms (7-, 9-, α− and β-) of 2-aminopurine 2’-deoxyriboside were analyzed [402]. Both the α-9- and the β-7-isomers bind to A·T base pairs similarly to a thymine in a GToligonucleotide. The α-9-isomer binds also to G·C base pair with as similar stability to A·T base pair [402]. Overcoming K+-Inhibition of Triplex formation via GQuartet Structures Triple helix formation involving guanine-rich oligonucleotides is inhibited by physiological ions, particularly K+, most likely due to oligonucleotide aggregation such as formation of guanine quartet structures. Several guanine analogs have been designed to prevent G-quartet structure (Figure 26). Oligonucleotides carrying 6-thioguanine were found to resist K + mediated inhibition but triplexes were less

Fig. (26). Analogs aimed at overcoming K+-inhibition of triplexes via G-quartet



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Fig. (27). Analogs aimed at interacting with both Watson-Crick bases

Fig. (28). N4-Cytosine derivatives aimed at interacting with both Watson-Crick bases

stable than triplexes with guanines [403-406]. The increased radius and decreased electronegativity of sulfur at the 6position of guanine destabilize potential guanine quartets. Substitution of thymines by 7-deazaxanthines in G,T-rich oligonucleotides has shown also triplex stabilization [403, 407, 408]. 9-Deazaguanine and 6-thio-7-deazaguanine have been also prepared to avoid K+-inhibition of triplex but a strong destabilization was found [409, 410]. Recently, 7chloro-7-deazaguanine was found to resist K+-inhibition and to form triplexes with similar or slightly better stability than triplexes having guanine [411]. 7-Deazaguanine was found to inhibit G-quartet structures but no data on triplexes were described [412]. Protonated backbones were found to form more stable triplex at high K+ concentration [413]. The introduction at the ends of the G-rich oligonucleotides of short duplex structures [414, 415] or small ligands [416] also destabilize G-quartet and enhance triplex formation.

3. HETEROCYCLIC COMPOUNDS AIMED AT INTERACTING WITH BOTH WATSON-CRICK BASES An innovative approach for the sequence-specific recognition of double-stranded DNA is the use of heterocyclic compounds which interact with base pairs in the major groove. The first molecule of this type was the 4-(3benzamidophenyl)imidazole, D3, (Figure 27a) described by the group of Dervan [417]. This compound was incorporated in a parallel triplex and it was show to bind specifically to direct TA and CG base pairs but not to inverse AT and GC base pairs. NMR experiments showed that the specific binding of the imidazole derivative is achieved by intercalation instead of forming sequence-specific hydrogen bonds [418]. The three aromatic rings of D3 stack on the bases of all three strands and mimics a triplet [418]. Also in a parallel triplex context, a series of N4-cytosine derivatives (Figure 28) have been studied [195, 419-422]. N4-(6-aminopyridinyl)-C (Figure 28a) and N4-(propyl)-C

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Fig. (29). Extension of targets accessible to triplex formation by alternate-strand.

(Figure 28c) form stable base pairs with inverse GC base pairs [195, 419, 421]. N4-(acetamidopropyl)-C (Figure 28b) form stable base pairs with direct CG base pairs [195]. N4(3-carboxypropyl)- and N4-(5-carboxytriazolyl)-C (Figure 28d) form stable base pairs with direct CG base pairs and inverse TA base pairs [420]. Recently, oligonucleotides carrying N4-anthraniloyl-, N4-ureidoand N4ureidocarbamoyl-C (Figure 28e) derivatives have been prepared [422]. Unfortunately, these last nucleosides do not form base triplets with any of the four base pairs in parallel triplexes. Other heterocyclic derivatives designed to interact with Watson-Crick bases are shown in Figure 27. The imidazole derivative b (Figure 27) was demonstrated by NMR to interact with CG base pair in chloroform at the nucleoside level [423]. The thiazol derivative c (Figure 27) show selective binding of inverted pyrimidine·purine base pair over direct purine·pyrimidine within a parallel triplex. But no descrimination between CG or TA base pair was found [424]. Recently, the derivative d (Figure 27) was found to form a stable parallel triplex with a pyrimidine interruption within the polypurine sequence, especially on an inverted AT base pair [425]. Several other heterocyclic compounds which may interact with Watson-Crick bases through the major groove have been described at the nucleoside level and their triplex binding properties are being evaluated [426428].

Linker molecules that have been used to introduce 3’-3’ linkages are 2-deoxy-1,4-anhydroribitol [429], 1,3propanediol, xylene [430], regular phosphodiester linkages [431] and lysine [432]. The design of the 5’-5’ linked oligonucleotides is difficult because the distance between the 5’-ends of the third strand is much longer than between the 3’ends. This has been achieved by using lineal molecules connected through the bases [433-435]. The second type of alternate-strand triple helix is when the two TFO to be linked are designed to form two different types of triplexes: one parallel and the other antiparallel (Figure 29b). Then TFO consists of both CT sequences (to form parallel triplex) and GA sequences [to form antiparallel triplexes). In this case both halves have the same polarity and TFO are easier to prepare [436-438]. In these mixed triplexes crossing the mayor groove is difficult due to the structure of the phosphate bonds. Strand recognition is easier at purine-pyrimidine junctions than at pyrimidine-purine junctions [436-438]. A third strategy consists in linking two TFO sequences having their triplex target sequences separated by several bases which are incompatible with triplex formation (Figure 29c). Two oligopurine sequences of 12 bases separated by 10 bases (roughly one helix turn) have been simultaneously bound by hybrid oligonucleotides carrying a flexible linker molecule longer than 20-25 rotatable bonds [439].

4. EXTENSION OF TARGETS ACCESSIBLE TO TRIPLEX FORMATION

Triplex with Interruptions Polypyrimidine Tracks

Alternate-Strand Recognition

An important limitation of triplex is the presence of interruptions in the polypurine-polypyrimidine tracks. This is the presence of a pyrimidine on a polypurine track or viceversa. Usually the base at Hoogsteen position binds to Watson-Crick purine. In the case of an interruption the base at the third strand has to bind to the pyrimidine site of duplex instead of the purine site. A large effort has been devoted in solving this problem [165, 440]. Some of the novel base analogs designed to bind both Watson-Crick bases are able to form stable triplexes in the presence of interruptions (see above). The ability of the natural bases to bind to the pyrimidine site was evaluated by affinity cleaving [441]. The best base to bind to a T·A interruption was G [441, 442].

When the DNA target is made of close or adjacent oligopurine-oligopyrimidine tracks it is possible to link the triplex-forming oligonucleotides with the appropriate spacer molecules (see Figure 29). Triplex formation is enhanced due to cooperative binding. This approach is known as alternate-strand triple helix formation. The most common mode of alternate-strand triple helix formation is when two parallel triplexes are adjacent. In this case pyrimidine triplexforming oligonucleotides are in opposite directions (Figure 29a). This requires joining the TFO sequences either with 3’3’ or 5’-5’ linkages to change the orientation at the junction.

in

the

Polypurine-



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Fig. (30). Base analogs aimed at overcoming interruptions at polypurine-polypyrimidine tracks.

Recently the structural characteristics of the G-T·A triad has been determined by NMR [443]. The best base to bind to a C·G interruption is T [441]. In order to avoid destabilization produced by unfavorable steric hindrance effects abasic analogs were studied. Good results were described for the (4aminobutyl) propane-1,3-diol (Figure 30g) probably due to interactions of protonated amino group and phosphates [444].

451] and 5-bromo-C [380] were found to destabilize triplex (see above).

Bases that were designed to bind all four natural bases in duplex structures (universal bases, [445]) were analyzed for their binding to interruptions on triplex [446-448]. Azole derivatives (Figure 30a-c) were shown to bind to both TA and CG inversion sites but more specifically to TA inversion sites [447]. 3-Nitropyrrole (Figure 30d) was found to bind specifically to CG inversion site and N6-methoxy-2,6diaminopurine (Figure 30f) binds to T·A inversion site [446, 448].

2’-Deoxyformycin A was found to bind with high affinity to CG inverted sites within target sites in antiparallel triplexes [453].

A systematic study of six uracil derivatives to bind at CG and TA inversion sites in a GT oligonucleotide to form antiparallel triplexes found thymine and 5-fluorouracil to have the best binding efficacy for CG inversion sites [449]. 5-Fluorouracil binds also to TA inversion sites [449]. Recently a study of eight base analogues opposite each direct (AT or GC) or inverted (TA or CG) base pairs in a parallel triplex context has been made [450]. None of the analogs studied were found to bind efficiently to TA inverted sites. But, 5-propynyl- (Figure 21a) and 5-bromo-U (Figure 20a) were found to bind better than T to CG inverted sites. Surprisingly, in this study, 5-propynyl-C (Figure 21b) and 5bromo-C (Figure 20b) were found to bind similar than C to direct GC base pair [450]. In a different sequence context but still the same parallel triplex context 5-propynyl-C [382,

The use of N-(2-hydroxyethyl)glycine residue having several small ligands was studied to bind to a TA inverted site in a parallel triplex context [452]. Only the monomer carrying an anthraquinone molecule (Figure 30i) gave a more stable triplex than the unmodified oligonucleotide.

5. BASE ANALOGS AT WATSON-CRICK POSITIONS Several base analogs have been incorporated at the Watson-Crick positions (Figure 31). In some cases new triads were found. Although the use of these analogs is limited due to the fact that the modification is located at the target site, in some cases it is possible to overcome this inconvenient and design oligonucleotides with enhanced binding properties. Pyrimidine Modifications Pseudouridine (Figure 31) is able to form two types of new triads. The T-pseudoU·T [454] and the A-pseudoU·A triad [455]. In these triads pseudouridine is located at the Watson-Crick position at the purine strand (replacing an A). Binding of the nucleoside at the Hoogsteen strand (T or A) is made through H-bonds with pseudouridine instead of the A in the normal T-A·T or A-A·T. For this reason, these triads are known as inverted motifs. A similar result was found if

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Fig. (31). Base analogs aimed at stabilizing triplexes when located at Watson-Crick positions

Fig. (32). Base analogs aimed at overcoming interruptions at polypurine-polypyrimidine tracks.

5-amino-U (Figure 31) was located at the central position [456, 457]. The amino group at position 5 of uracil is acting as H-donor group for Hoogsteen pairing as it does one of the N-H groups of pseudouridine. Moreover it has been shown that 5-aminouracil is able to bind to G, 2-aminopurine and T within an antiparallel context and C, T and A in parallel orientation [457]. Purine Modifications The introduction of an amino group at position 8 of the Watson-Crick purines (A, G and hypoxanthine, Figure 31) produces a high stabilization of parallel triplexes [188-190, 458, 459]. The stabilizing properties of these analogues were hypothesized long time ago [460-462] but experimental data was not available due to self-aggregation of homopolymers containing these bases. Recently, the preparation of oligonucleotides carrying 8-aminopurines was described and a high stabilization effect on triplex was found [188-190, 458, 459]. The triplex-stabilization properties of the amino group at position 8 is due to a combined effect of the gain of one Hoogsteen purine-pyrimidine H-bond (Figure 32) and the propensity of the amino group to be integrated into the ‘spine of hydration’ located in the minor-major groove of the

triplex. In order to use the triplex-stabilization properties of 8-aminopurines, hairpins formed by a polypyrimidine part linked to a polypurine sequence carrying 8-aminopurines were prepared [191]. These hairpins showed enhanced binding to polypyrimidine targets by triple helix formation. The presence of 8-aminopurines also stabilized Hoogsteen parallel-stranded duplex structure [191, 192]. The increased binding properties of hairpins carrying 8-aminopurines may be of special interest to the development of applications based on triple helix formation.

REFERENCES [1]

Soyfer, V.N.; Potaman, V.N. Triple-Helical Nucleic Acids. Springer-Verlag: New York. 1996.

[2]

Frank-Kamenetskii, M.D.; Mirkin, S.M. Annu. Rev. Biochem., 1995, 64, 65.

[3]

Luyten, I.; Herdewjn, P. Eur.J.Med.Chem., 1998, 33., 515.

[4]

Fox, K.R. Current Med. Chem., 2000, 7, 17.

[5]

Micklefield, J. Current Med. Chem., 2001, 8, 1157.


[6]

Vasquez, K.M.; Wilson,K.H. Trends in Biochem., 1993, 23, 4.

[7]

Chubb, J.M.; Hogan, M.E. Trends in Biotechnol., 1992, 10, 132.

[8]

Feigon, J.; Koshlap, K.M.; Smith, P.E.Methods Enzymol., 1995, 261, 225.

[9]

Mirkin, S.M.; Frank-Kamenetskii, M.D. Annu. Rev. Biomol. Struct. 1994, 23, 541.

[10]

Plum, G.E.; Pilch, D.S.; Singleton, S.F.; Breslauer, K.J. Annu. Rev. Biophys. Biomol. Struct., 1995, 24, 319.

[11]

Hélène, C.; Toulme, J.J. Biochim. Biophys. Acta, 1990, 1049, 99.

[12]

Strobel, S.A.; Dervan, P. Methods Enzymol., 1992, 216, 309.

[13]

Sun,J.S.; Garestier,T.; Hélène,C. Curr. Opin. Struct. Biol., 1996, 6, 327.

[14]

Thuong,, N.T.; Hélène,C. Angew. Chem. Int. Ed. Engl., 1993, 32, 666.

[15]

Saenger, W. Principles of Nucleic Acid Structure. SpringerVerlag: New York. 1984.

[16]

Malvy, C.; Harel-Bellan, A.; Pritchard, L.L. Triple helix forming oligonucleotides. Kluwer Academic. Dordrecht. 1999.

[17]

Bloomfield, V.A.; Crothers, D.M.; Tinoco, I (eds). Nucleic Acids: Structures, Properties and Functions. University Science Books: Sausalito. 2000.

1359

[30]

Arnott, S.; Hukins, D.W.L. J.Mol.Biol., 1973, 81, 93.

[31]

Watson, J.D.; Crick, F.H.C. Nature, 1953, 171, 737.

[32]

Drew, H.R.; Wing, R.M.; Tanako, T.; Broka, C.; Tanaka, S.; Itakura, K.; Dickerson, R. Proc. Natl. Acad. Sci. USA., 1981, 78, 2179.

[33]

Kennard, O.; Hunter, W.N. Q. Revs. Biophys., 1989, 22, 327.

[34]

171 structures are labelled as B-DNA in the Dec 21st 2001 update of PDB. 218 entries are labelled as A-DNA.

[35]

Pardi, A.; Hare, D.R.; Wang, C. Proc. Natl. Acad. Sci. USA, 1988, 85, 8785.

[36]

Van de Ven,F.J.M.; Hilbers,C.W. Eur. J. Biochem., 1988, 178, 1.

[37]

Chen, J.W.; Chou, S.H.; Salazar, M.; Reid, B.R. J. Mol. Biol., 1992, 228, 1037.

[38]

Cheatham, T.E.; Kollman, P.A. J. Am. Chem. Soc., 1995, 117, 4193.

[39]

Cheatham, T.E.; Kollman, P.A. J. Mol. Biol., 1996, 259, 434.

[40]

Jayaram, B.D.; Sprous, M.A.; Young, M.A.; Beveridge, D.A. J. Am. Chem. Soc., 1998, 120, 10629.

[41]

Soliva, R.; Luque, F.J.; Alhambra, C.; Orozco, M. J. Biomol. Struct. Dyn., 1999, 17, 89 [1999).

[42]

Arnott, S.; Chandrasekharan, R.; Hukins, D.W.L.; Smith, P.J.C.; Watts, L. J. Mol. Biol., 1974, 88, 523.

[43]

Davis, D.R.; Baldwin, R.L. J. Mol. Biol., 1963, 6, 251.

[44]

Marvin, D.A.; Spencer, M.; Wilkins, M.H.F.; Hamilton, L.D. J. Mol. Biol., 1961, 3, 547.

[18]

Calladine, C.R.; Drew, H.R. Understanding Academic Press: San Diego. 1997.

[19]

Dickerson,R.E. Nucleic Acid Res., 1998, 26, 1906.

[20]

Blackburn, G.M.; Gait, M.J. (Eds). Nucleic Acids in Chemistry and Biology. IRL Press. Oxford, 1990.

[45]

Mokul’skii, M.A.; Kapitanova, K.A.; Mokul’skaya, T.D. Mol. Biol., 1972, 6, 714.

[21]

Neidle, S. (Ed). Oxford Handbook of Nucleic Acid Structure. Oxford University Press. Oxford. 1999.

[46]

Premilar, S.; Albiser, G. Eur. Biophys. J., 2001, 30, 404.

[47]

[22]

Sinden, R.R. DNA Structure and Function. Academic Press, San Diego. 1994.

Fuller, W.; Wilkins, M.H.F.; Wilson, H.R.; Hamilton, L.D. J. Mol. Biol., 1965, 12, 60.

[48]

[23]

Lane, A.N.; Jenkins, T.C. Curr. Org. Chem., 2001, 5, 845.

Arnott, S.; Hukins, D.W.L. Biochem. Biophys. Res. Commun., 1972, 47, 1504.

[24]

Berman, H.M.; Olson, W.K.; Beveridge, E.L.; Westbrook, J.; Gelbin, A.; Demeny, T.; Hsieh, S.H.; Srinavasan, A.R.; Scheneider, B. Biophys.J., 1992, 63, 751.

[49]

Wang, A.H.J.; Fujii, S.; van Boom, J.H.; Rich, A. Proc. Natl. Acad. Sci. USA, 1982, 79, 3968.

[50]

[25]

Dickerson, R.E. Methods in Enzymol., 1992, 211, 67.

Shakked, Z.; Rabinovich, D.; Cruse, W.B.T.; Egert, E.; Kennard, O.; Sala, G.; Salisbury, S.A.; Viswamitra, M.A. Proc. Roy. Soc. London Ser. B., 1981, 231, 479.

[26]

Leslie, A.G.W.; Arnott, S.; Chandrasekharan, R.; Ratliff, R.L. J. Mol. Biol., 1980, 143, 49.

[51]

Arnott, S.; Hukins, D.W.L.; Dover, S.D.; Fuller, W.; Hodgsn, A.R. J. Mol. Biol., 1973, 81, 107.

Shaked, Z.; Guerstein-Guzikevich, G.; Eisenstein, M.; Frolow, F.; Rabinovich, D. Nature, 1989, 342, 456.

[52]

Milman, G.; Langridge, R.; Chamberlin, M.J. Proc. Natl. Acad. Sci. USA., 1967, 57, 1804.

Shaked, Z.; Rabinovich, D. Prog. Biophys. Molec. Biol., 1986, 47, 159.

[53]

Vargason, J.M.; Eichman, B.F.; Ho.; P.S. Nature Struct. Biol., 2000, 7, 758.

[27] [28] [29]

Arnott, S. Prog, Biophys. Mol. Biol., 1970, 21, 267.

DNA.


1360


Orozco et al.

[54]

Vargason, J.M.; Henderson, K.; Ho.; P.S. Proc. Natl. Acad. Sci. USA., 2001, 98, 7265.

[78]

Mohammadi, S.; Klement, R.; Shchyolkina, A. K.; Liquier, J.; Jovin, T. M.; Taillandier, E. Biochemistry, 1998, 37, 16529.

[55]

Dickerson, R.E.; Ng, H.-L. Proc. Natl. Acad.Sci. USA., 2001, 98, 6986.

[79]

Pattabiraman, N. Biopolymers, 1986, 25, 1603.

[56]

Pohl,F.M.; Jovin,T.M. J. Mol. Biol., 1972, 67, 375.

[80]

Zhou, N.; Germann, M. W.; van de Sande, J. H.; Pattabiraman, N.; Vogel, H. J. Biochemistry, 1993, 32, 646.

[57]

Davis, D.R.; Zimmerman. Nature, 1980, 283, 11. [81]

[58]

Wang,, A.H.J.; Quigley, F.J.; Kolpak,G.; van der Marel,G.; van Boom, J.H.; Rich, A. Science, 1980, 211, 171.

Cubero,E.; Luque,F.J.; Orozco,M. J. Am. Chem. Soc., 2002, 123, 12018

[82] [59]

Wang,, A.H.J.; Quigley, F.J.; Kolpak, G.; Crawford, J.L.; van Boom, J.H.; van der Marel, G.; Rich.A. Nature, 1979, 282, 680.

Cubero, E.; Aviño, A.; de la Torre, B. G.; Frieden, M.; Eritja, R.; Luque, F.J.; González, C.; Orozco, M. J. Am. Chem. Soc., In Press 2002.

[83] [60]

Drew, H.; Takano,T.; Tanaka,S.; Itakura,K., Dickerson,R.E. Nature, 1980, 286, 567.

Kandimalla, E. R.; Agrawal, S.; Venkataraman, G.; Sasisekharan, V. J. Am. Chem. Soc., 1995, 117, 6416.

51 structures are labelled as Z-DNA in the Dec 21st 2001 release of PDB.

[84]

[61]

Kandimalla, E. R.; Agrawal, S. Biochemistry, 1996, 35, 15332.

[85] [62]

Olson, W.K.; Srinavasan, A.R.; Marky, N.L.; Balaji, V.N. Cold Spring Harbor Symp. Quant. Biol., 1982, 47, 229.

[63]

Harvey, S.C. Nucleic Acids Res., 1983, 11, 4867.

Aviñó, A.; Morales, J. C.; Frieden, M.; de la Torre, B. G.; Güimil-García, R.; Cubero, E.; Luque, F. J.; Orozco, M.; Azorín, F.; Eritja. Bioorganic Med. Chem. Lett., In Press 2002.

[64]

Pearlman, D.A.; Kollman, P.A. Biopolymers, 1990, 29, 1193.

[86]

Hashem, G. M.; Wen, J. D.; Do, Q.; Gray, D. M. Nucl. Acids Res., 1999, 27, 3371.

[65]

Abrescia, N.G.A.; Thompson, A.; Dinh, T.H.; Subirana, J.A Proc. Natl. Acad. Sci. USA, In Press. 2002.

[87]

Escudé, C.; Mohammadi, S.; Sun, J. S.; Nguyen, C-H.; Bisagni, E.; Liquier, J.; Taillandier, E.; Garestier, T.; Hélène, C. Chemistry and Biology, 1996, 3, 57.

[66]

Westhof, E., Sundaralingam, M. Proc. Natl. Acad. Sci. USA.,1980 77, 1852.

[88]

Germann, M. W.; Kalisch, G. T.; van de Sande, J. H. Biochemistry, 1998, 37, 12962.

[67]

Rippe, K.; Jovin, T. M. Methods in Enzymology, 1992, 211, 199.

[89]

Ortiz-Lombardía,M.; González,A.; Eritja,R.; Aymamí,J.; Azorín,F.; Coll,M. Nature Structural Biology, 1999, 6, 1.

[68]

Sande, J. H.; Ramsing, N. B.; Germann, M. W.; Elhorst, W.; Kalisch, B. W.; Kitzing, E. V.; Pon, R. T.; Clegg, R. C.; Jovin, T. M. Science, 1988, 241, 551.

[90]

Gehring, K.; Leroy, J.L.; Guéron, M. Nature, 1993, 363, 561.

[69]

Ramsing, N. B.; Rippe, K.; Jovin, T. M. Biochemistry, 1989, 28, 9528.

[91]

Chen, L.; Cai, L.; Zhang, X.; Rich, A. Biochemistry, 1994, 33, 13540.

[70]

Germann, M. W.; Vogel, H. J.; Pon, R. T.; van de Sande, J. H. Biochemistry, 1989, 28, 6220.

[92]

Ahmed, S.; Kintanar, A.; Henderson, E. Nature Struct. Biol., 1994, 1, 83.

[71]

Rippe, K.; Ramsing, N. B.; Jovin, T. M. Biochemistry, 1989, 28, 9536.

[93]

Leroy, J.L.; Guéron, M.; Mergny, J.L.; Hélène, C. Nucleic Acids Res., 1994, 22, 1600.

[72]

Rentzeperis, D.; Rippe, K.; Jovin, T. M.; Marky, L. A. J. Am. Chem. Soc., 1992, 114, 5926.

[94]

Spackova, N.; Berger, I.; Egli, M.; Sponer, J. J. Am. Chem. Soc., 1998, 120, 6147.

[73]

Germann, M. W.; Kalisch, B. W.; Pon, R. T.; van de Sande, J. H. Biochemistry, 1990, 29, 9426.

[95]

Phan, A.T.; Guéron, M.; Leroy, J.L. J. Mol. Biol., 2000, 299, 123.

[74]

Lavelle, L.; Fresco, J. R. Nucl. Acids Res., 1995, 23, 2692.

[96]

Gray, D.M.; Vaughn, M.; Ratliff, R.L.; Hayes, F.N. Nucleic Acids Res., 1980, 8, 3695.

[75]

Scaria, P. V.; Shafer, R. H. Biochemistry, 1996, 35, 10985.

[97]

Jenkins, T.C. Curr. Med. Chem., 2000, 7, 99.

[76]

Bhaumik, S. R.; Kandala, V. R.; Govil, G.; Liu, K.; Miles, H. T. Nucl. Acids Res., 1995, 23, 4116.

[98]

Perry, P.J.; Jenkins, T.C.; Exp. Opin. Invest. Drugs., 1999, 8, 1981.

[77]

Liu, K.; Miles, T.; Frazier, Biochemistry, 1993, 32, 11802.

[99]

Williamson, J.R.; Raghuraman, M.K.; Cech, T. R. Cell, 1989, 59, 871.

J.;

Sasisekharan,

V.

[100] Rhodes, D.; Giraldo, R. Curr. Opin. Struct. Biol., 1995, 5, 311.


[101] Mergny, J.L.; Maillet, P.; Lavelle, F.; Riou, J.F.; Lauoi, A.; Hélène, C. Anti Cancer Drug Des., 1999, 14, 327.


1361

[126] Wang,, G.; Xu, X.; Pace, B.; Glazer, P.M.; Chan, P.; Goodman, S.R.; Shokolenko, O. Nucleic Acid Res., 1999, 27, 2806.

[102] Kervin, S.M. Curr. Pharm. Des., 2000, 6, 441. [103] Xu, X.; Hamhouria, F.; Thomas, S.D.; Burke, T.J.; Girvan, A.C.; McGregor, WG.; Trent, J.O.; Miller, D.M.; Bates, P.J. J. Biol.Chem., 2001, 276, 43221. [104] Hud, N.V.; Smith, F.W.; Anet, F.A.L.; Feigon, J. Biochemistry, 1996, 35, 15383.

[127] Barre, F.X.; Ait-Si-Ali, S.; Giovannangeli, C.; Luis, R.; Robin, P.; Pritchard, L.L.; Hélène, C.; Harel-Bellan, A. Proc. Natl. Acad. Sci. USA, 2000, 97, 3084. [128] Bernal-Méndez, E.; Leumann, C.J. J. Biol. Chem., 2001, 276, 35320. [129] Chan, P.P.; Glazer, P.M J. Mol. Med., 1997, 75, 267.

[105] Radhakrishnan, I.; Patel, D.J. Biochemistry, 1994, 33, 11405.

[130] Arnot, T.S.; Bond, P.J.; Selsing, E.; Smith, P.J.C. Nucleic Acids Res., 1976, 133, 1405.

[106] Deng, H.; Braulin, W. H. J. Mol. Biol., 1996, 255, 476. [107] Sundquist, W.I.; Klug., A. Nature, 1989, 342, 825.

[131] Van Meervelt, L.; Vlighe, D.; Dautant, A.; Gallois, B.; Précigoux, G., Kennard, O. Nature, 1995, 374, 742

[108] Scaria, P.V.; Shire, S.J.; Shafer, R.H. Proc. Natl. Acad. Sci. USA, 1992, 89, 10336.

[132] Radhakrishnan, I.; Patel, D.J. Structure, 1994, 2, 395. [133] Rajagopal, P.; Feigon, J. Nature, 1989, 22, 637.

[109] Spackova, N.; Berger, I.; Sponer, J. J. Am. Chem. Soc., 1999, 121, 5519.

[134] Rajagopal, P.; Feigon, J. Biochemistry, 1989, 28, 7859.

[110] Spackova, N.; Berger, I.; Sponer, J. J. Am. Chem. Soc., 2001, 113, 3295.

[135] Macaya, R.F.; Schultze, P.; Feigon, J. J. Am. Chem. Soc., 1992, 114, 781.

[111] Laughlan, G.; Murchie, A.I.H.; Norman, D.G.; Moore, M.H.; Moody, P.C.E.; Lilley, D.M.; Luisi, B. Science, 1994, 265, 520.

[136] Radhakrishnan, I.; Gao, X.; de los Santos, C.; Live, D.; Patel, D.J. Biochemistry, 1991, 30, 9022.

[112] Philips, K.; Dauter, Z.; Murchie, A.I.H.; Lilley, D.M.J.; Luisi, B. J. Mol. Biol., 2000, 297, 627. [113] Luisi, B. Private communication. [114] Soliva, R.; Luque, F.J.; Orozco, M. To be published. [115] Pauling, L.; Corey, R.B. Proc. Natl. Acad. Sci. USA., 1953, 39, 84. [116] Felsenfeld, G.; Davis, D.R.; Rich, A. J. Am. Chem. Soc., 1957, 79, 2023. [117] Giovannangeli, C.; Hélène, C. Nature Biotech., 2000, 18, 1245.

[137] Koshlap, K.M.; Schultze, P.; Brunar, H.; Dervan, P.D.; Feigon, J. Biochemistry, 1997, 26, 2659. [138] Gotfredsen, C.H.; Schultze, P.; Feigon, J. J. Am. Chem. Soc., 1998, 120, 4281. [139] Young, S.L.; Krawczyk, S.H.; Matteuci, M.D.; Toole, J.J. Proc. Natl. Acad. Sci. USA., 1991, 88, 10023. [140] Asensio, J.L.; Brown, T.; Lane, A.N. Nucleic Acids Res., 1998, 26, 3677. [141] Bartley, J.P.; Brown, T.; Lane, A.N. Biochemistry, 1997, 36, 14502. [142] Powell, S.W.; Jiang,, L.; Russu, I.M. Biochemistry, 2001, 40, 11065.

[118] Vasquez, K.M.; Narayanan, L.; Glazer, P.M. Science, 2000, 290, 530.

[143] Laughton, C.A.; Neidle, S. J. Mol. Biol., 1992, 223, 519.

[119] Rocher, C.; Dalibart, R.; Letellier, T.; Précigoux, G.; Lestienne, P. Nucleic Acids Res., 2001, 29, 3320.

[144] Mohan, V.; Smith, P.E.; Pettitt, M. J. Am. Chem. Soc., 1993, 115, 9297.

[120] Van Dongen, M.J.P.; Doreleijiers, J.F.; van der Marel, G.; van Boom, J.H.; Hilbers, C.W.; Wijmenga, S. Nature Struct. Biol., 1999, 6, 854.

[145] Weerasinghe, S.; Smith, P.E.; Mohan, V.; Cheng, Y.K.; Pettitt, B.M. J. Am. Chem. Soc., 1995, 117, 2147.

[121] Wang,, G.; Chen, Z.; Zhang,, S.; Wilson, G.L.; Jing, K. Nucleic Acids Res., 2001, 29, 1801. [122] Cooney, M.; Czernuszewicz, G.; Postel, E.H.; Flint, S.J.; Hogan, M.E. Science, 1988, 241, 456.

[146] Shields, G.C.; Laughton, C.A.; Orozco, M. J. Am. Chem. Soc., 1997, 119, 7463. [147] Soliva, R.; Laughton, C.A.; Luque, F.J.; Orozco, M. J. Am. Chem. Soc., 1998, 120, 1126.

[123] Vuyisich, M.; Beal, P.A. Nucleic Acid Res., 2000, 28, 2369.

[148] Soliva, R.; Luque, F.J.; Orozco, M. Nucleic Acids Res., 1999, 27, 2248.

[124] Lyamichew, V.I.; Frank-Kamenetskii, M.D.; Soyfer, V. Nature, 1990, 344, 568.

[149] Shields, R.; Laughton, C.A.; Orozco, M. J. Am. Chem. Soc., 1998, 120, 5895.

[125] Lin, F.L.; Majumdar, A.; Klotz, L.C.; Reszka, A.P.; Neidle, S.; Seidman, M.M. J. Biol. Chem., 2000, 275, 39117.

[150] Srinivasan, A.R.; Olson, W.K. J. Am. Chem. Soc., 1998, 120, 484.

1362


Orozco et al.

[151] Singleton, S.F.; Dervan, P.B. J. Am. Chem. Soc., 1994, 116, 10376.

[176] Lee, J.S.; Woodworth, M.L.; Latimer, L.J.; Morgan, A.R. Nucleic Acids Res., 1987, 15, 1047.

[152] Plum, G.E.; Breslauer, K.J. I. Mol. Biol., 1995, 248, 679.

[177] Kohwi, Y.; Malkhosyan, S.R.; Kohwi-Shigematsu, T. J. Mol. Biol., 1992, 223, 817.

[153] Roberts, R.W.; Crothers, D.M. Proc. Natl. Acad. Sci. USA, 1996, 93, 4320. [154] Mundt, A.A.; Crouch, G.J., Eaton, B.E. Biochemistry, 1997, 36, 13004. [155] Rodrigues-Hoyne, P.; Gacy, A.M.; McMurray, C.T.; Maher, L.J. Nucleic Acids Res., 2000, 28, 770. [156] Asensio, J.L.; Dosanjh, H.S.; Jenkins, T.C.; Lane, A.N. Biochemistry, 1999, 37, 1588. [157] Leitner, D.; Schröder, W.; Weisz. J. Am. Chem. Soc., 1998, 120, 7123. [158] Leitner, D.; Schröder, W.; Weisz. Biochemistry, 2000, 39, 5886.

[178] Umemoto, K.; Sarma, M.H.; Gupta, G.; Luo, J.; Sarma, R.H. Biochemistry, 1990, 112, 4539. [179] Raghunathan, G.; Miles, Biochemistry, 1993, 32, 455.

H.T.;

Sasisekharan,

V.

[180] Howard, F.B.; Miles, H.T.; Liu, K.; Frazier, J.; Raghunathan, G.; Sasiskharan, V. Biochemistry, 1992, 31, 10671. [181] PDB contains 18 entries labeled as DNA triplexes at Dec 21st 2001. Nine of these entries correspond to unique structures of DNA-DNA·DNA triplexes solved from NMR data. There is also one RNA-DNA·DNA structure [pdb1r3x), and two PNA-DNA·PNA triplexes [pdb1hzs and pdb1pnn).

[159] Sugimoto, N.; Wu, P.; Hara, H.; Kawamoto, Y. Biochemistry, 2001, 40, 9396.

[182] Radhakrishnan, I.; Patel, D.J. Structure, 1993, 1, 135.

[160] Chandler, S.P.; Fox, K.R. Biochemistry, 1996, 35, 15038.

[183] Schultze, P.; Koshlap, K.M.; Feigon, J. Biochemisty, 1997, 36, 2659.

[161] Chen, D.L.; McLaughlin, L.W. J. Org. Chem., 2000, 65, 7468.

[184] Radhakrishnan, I.; Patel, D.J. J. Mol. Biol., 1994, 241, 600.

[162] Goobes, R.; Minsky, A. J. Biol. Chem., 2001, 276, 16155.

[185] Tarkoy, M.; Phipps, A.K.; Schultze, P.; Feigon, J. Biochemistry, 1998, 37, 5810.

[163] Mills, M.; Arimondo, P.B.; Lacroix, L.; Garestier, T.; Hélène, C.; Klump, H.; Mergny, J.L. J. Mol. Biol., 1999, 291, 1035.

[186] Phipps, A.K.; Tarkoy, M.; Schultze, P.; Feigon, J. Biochemistry, 1998, 37, 5820.

[164] Debin, A.; Laboulais, C.; Ouali, M.; Malvy, C.; Le Bret, M.; Svinarchuk, F. Nucleic Acids Res., 1999, 27, 2699. [165] Gowers, D.M.; Fox, K.R. Nucleic Acids Res., 1999, 27, 1569. [166] Gryaznov, S.M.; Lloyd, D.H. Nucleic Acids Res., 1993, 21, 5909.

[187] Betts, L.; Josey, J.A.; Veal, J.M.; Jordan, S.R. Science, 1995, 270, 1838. [188] Güimil-Garcia, R.; Ferrer, E.; Macías, M.J.; Eritja, R.; Orozco, M. Nucleic Acids Res., 1999, 27, 1991. [189] Cubero, E.; Güimil-Garcia, R.; Luque, F.J.; Eritja, R.; Orozco, M. Nucleic Acids Res., 2001, 29, 2522.

[167] Durand, M.; Thuong,, N.T.; Maurizot, J.C. J. Biol. Chem., 1992, 267, 24394.

[190] Soliva, R.; Güimil-Garcia, R.; Blas, J.R.; Eritja, R.; Asensio, J.L.; González, C.; Luque, F.J.; Orozco, M. Nucleic Acids Res., 2000, 28, 4531.

[168] Park, Y.W.; Breslauer, K.J. Proc. Natl. Acad. Sci. USA, 1992, 89, 6653.

[191] Shui, X.; McFail-Isom, L.; Hu, G.G.; Williams, L.D. Biochemistry, 1998, 37, 8341.

[169] Wilson, W.D.; Mizan, S.; Tanious, F.A.; Yao, S.; Zon, G. J. Mol. Recogn., 1994, 7, 89.

[192] Colominas, C.; Luque, F.J.; Orozco, M. J. Am. Chem. Soc., 1996, 188, 6811.

[170] Jiménez, E.; Vaquero, A.; Espinás, M.L.; Soliva, R.; Orozo, M.; Bernués, J.; Azorín, F. J. Biol.Chem., 1998, 273, 24640.

[193] Szczesniak, M.; Szczepaniak, K.; Kwiatkowski, J.S.; BuBulat, K.; Person, W.B. J. Am. Chem. Soc., 1988, 110, 8319.

[171] Brosh, R.M.; Majumdar, A.; Desai, S.; Hickson, I.D.; Bohr, V.A.; Seidman, M.M. J. Biol. Chem., 2001, 276, 3024. [172] Guieysse, A-L.; Praseuth, D.; Hélène, C. J. Mol. Biol., 1997, 267, 289. [173] Musso, M.; Nelson, L.D.; van Dyke, M.W. Biochemistry, 1998, 37, 3086. [174] Rodrigues-Hoyne, P.; Edwards, L.M.; Viari, A.; Maher, L.J. I. Mol. Biol., 2000, 302, 797. [175] Bucher, P.; Yagil, G. DNA Sequence., 1991, 1, 157.

[194] Dreyfus, M.; Beusande, O.; Dodin, G.; Dubois, J.E. J. Am. Chem. Soc., 1976, 93, 6338. [195] Wang,, E.; Koshlap, K.M.; Gillespie, P.; Dervan, P.B.; Feigon, J. J. Mol. Biol., 1996, 257, 1052. [196] Asensio, J.L.; Deshai, J.; Bergqvist, S.; Brown, T.; Lane, A.N. J. Mol. Biol., 1998, 275, 811. [197] Radhakrishnan, I.; Patel, D.J. Structure, 1994, 2, 1.


[198] Bernués, J.; Beltran, R.; Casasnovas, J.M.; Azorin, F. EMBO J., 1989, 8, 2087. [199] Bernués, J.; Beltran, R.; Casasnovas, J.M.; Azorin, F. Nucleic Acids Res., 1990, 18, 4067. [200] Beltran, R.; Martínez-Bañbas, A.; Bernués, J.; Bowater, R.; Azorin, F. J. Mol. Biol., 1993, 230, 966.


1363

[223] Pilch, D.S.; Waring, M.J.; Sun, J.S.; Rougée, M.; Nguyen, C.H.; Bisagni, E.; Garestier, T.; Hélène, C. J. Mol. Biol., 1993, 232, 926. [224] Escudé, C.; Sun, J.S.; Nguyen, C.H.; Bisagni, E.; Garestier, T.; Hélène, C. Biochemistry, 1996, 35, 5735. [225] Silver, G.C.; Sun, J.S.; Nguyen, C.H.; Boutorine, A.S.; Bisagni, E.; Hélène, C. J. Am. Chem. Soc., 1997, 119, 263.

[201] Washbrook, E.; Fox, K.R. Biochem. J., 1994, 301, 569. [202] Cheng, Y.K.; Pettitt, B.M. J. Am. Chem. Soc., 1992, 114, 4465. [203] Malkov, A.; Shafer, R.H. Biochemistry, 1996, 35, 10985. [204] Debin, A.; Laboulais, C.; Ouali, M.; Malvy, C.; Le Bret, M.; Svinarchuk, F. Nucleic Acids Res., 1999, 27, 2699. [205] Dagneaux, C.; Liquier, J.; Taillander, E. Biochemistry, 1995, 34, 16618 [206] Ji, J.; Hogan, M.E.; Gao, X. Structure, 1996, 4, 425.

[226] Wilson, W.D.; Tanious, F.A.; Mizan, S.; Yao, S.; Kiselyov, A.S.; Zon, G.; Strekowski, L. Biochemistry, 1993, 32, 10614. [227] Cassidy, S.A.; Strekowski, L.; Wilson, W.D.; Fox, K.R. Biochemistry, 1994, 33, 15338. [228] Chandler, S.P.; Strekowski, L.; Wilson, W.D.; Fox, K.R. Biochemistry, 1995, 34, 15338. [229] Kan, Y.; Armitage, B.; Schuster, G.B. Biochemistry, 1997, 36, 1461.

[207] Cheng, Y.K.; Pettitt, B.M. Biopolymers, 1994, 35, 457.

[230] Fox, K.R.; Polucci, P.; Jenkins, T.C.; Neidle, S. Proc. Natl. Acad. Sci., USA, 1995, 92, 7887.

[208] Weerasinghe, S.; Smith, P.; Pettitt, B.M. Biochemistry, 1995, 34, 16269.

[231] Keppler, M.D.; Neidle, S.; Fox, K.R. Nucleic Acids Res., 2001, 29, 1935.

[209] Chalikian, T.V.; Plum, G.E.; Sarvazyan, A.P.; Breslauer, K.J. Biochemistry, 1994, 33, 8629.

[232] Sun, J.C.; Giovannangeli, C.; Francois, J.C.; Kurfust, K.; Monteny-Garestier, T.; Asseline, U.; Saison-Behmoaras, T.; Thuong, N.T.; Hélène, C. Proc. Natl. Acad. Sci. USA, 1991, 88, 6023.

[210] Pilch, D.S.; Breslauer, K.E. Proc. Natl. Acad. Sci. USA, 1994, 91, 9332. [211] Stonnehouse, T.J.; Fox, K.R. Biochim. Biophys. Acta, 1994, 1218, 322. [212] Durand, M.; Thuong, N.T.; Maurizot, J.C. Biochimie, 1994, 76, 181. [213] Durand, M.; Thuong, N.T.; Maurizot, J.C. J. Biomol. Struct. Dyn., 1994, 11, 1191. [214] Arya, D.P.; Coffee, R.L.; Willis, B.; Abramovitch, A.I. J. Am. Chem. Soc., 2001, 123, 5385. [215] Arya, D.P.; Coffee, R.L.; Charles, I. J. Am. Chem. Soc., 2001, 123, 11093 [216] Robles, J.; Rajur, S.B.; McLaughin, L.W. J. Am. Chem. Soc., 1996, 118, 5820. [217] Robles, J.; McLaughlin, L.W. J. Am. Chem. Soc., 1997, 119, 6014. [218] Le Pecq, J.B.; Paoletti, C. C.R. Acad. Sci., 1965, 260, 7033. [219] Morgan, A.R.; Lee, J.S.; Pulleyblak, D.E.; Murray, N.L.; Evans, D.E. Nucleic Acids Res., 1979, 7, 547. [220] Scaria, P. V.; Shafer, R. H. J. Biol. Chem., 1991, 266, 5417. [221] Tuite, E.; Norden, B. J. Chem. Soc. Chem. Commun., 1995, 53. [222] Mergny, J.L.; Duval-Valentin, G.; Nguyen, C.H.; Perrouault, L.; Faucon, B.; Rougée, M.; MontenayGarestier, T.; Nisagni, E.; Hélène, C. Science, 1992, 256, 1681.

[233] Asanuma, H.; Yoshida, T.; Liang, X.; Komiyama, M. Chem. Lett., 2000, 108. [234] Gianolio, D.A.; Segismundo, J.M.; McLaughlin, L.W. Nucleic Acids Res., 2000, 28, 2128. [235] LeDoan, T.; Perroualt, L.; Praseuth, D.; Habhoub, N.; Decout, J.L.; Thuong, N.T., Lhomme, J.; Hélène, C. Nucleic Acids Res., 1987, 15, 7749. [236] Praseuth, D.; Perroualt, L.; LeDoan, T.; Chassignol, M.; Thuong, N.; Hélène, C. Proc. Natl. Acad. Sci. USA, 1991, 88, 5602. [237] Takasugi, M.; Guendouz, A.; Chassignol, M.; Decout, J.L.; Lhomme, J.; Thuong,, N.T.; Hélène, C. Proc. Natl. Acad. Sci. USA, 1991, 88, 5602. [238] Grigoriev, M.; Praseuth, D.; Guyesse, A.L.; Robin, P.; Thuong, N.T.; Hélène, C.; Harrel-Bellan, A. Proc. Natl. Acad. Sci. USA, 1993, 90, 301. [239] Perroault, L.; Asseline, U.; Rivalle, C.; Thuong, N.T.; Bisagni, E.; Giovannangeli, C.; LeDoan, T.; Hélène, C. Nature, 1990, 344, 358. [240] Grant, K.B.; Dervan, P.B. Biochemistry, 1996, 35, 12313. [241] Wang, G.; Levy, D.D.; Seidman, M.M.; Glazer, P.M. Mol. Chell. Biol., 1995, 15, 1759. [242] Wang, F.; Seidman, M.M.; Glazer, P.M. Science, 1996, 271, 802. [243] Luyten, I.; Herdewijn, P. Eur. J. Med. Chem., 1998, 33, 515.

1364


[244] Roberts, R.W.; Crothers, D.M. Science, 1992, 258, 1473. [245] Rougée, M.; Faucon, B.; Mergny, J.L.; Barcelo, F.; Giovannangeli, C.; Garestier, T.; Hélène, C. Biochemistry, 1992, 31, 9269. [246] Singlenton, S.F.; Dervan, P.B. Biochemistry, 1992, 31, 10995. [247] Singlenton, S.F.; Dervan, P.B. J. Am. Chem. Soc., 1992, 114, 6957. [248] Han, H.; Dervan, P.B. Proc. Natl. Acad. Sci. USA, 1993, 90, 3806. [249] Semerad, C.L.; Maher, L.J. Nucleic Acid Res., 1994, 22, 5231. [250] Asensio, J.L.; Carr, R.; Brown, T.; Lane, A.N. J. Am. Chem. Soc., 1999, 121, 11063. [251] Gotfredsen, C.H.; Schultz, P.; Feigon, J. J. Am. Chem. Soc., 1998, 120, 4281. [252] Srinavasan, A.R.; Olson, W.K. J. Am. Chem. Soc., 1998, 120, 484.

Orozco et al.

[267] Zhou-Sun, B.W.; Sun, J.S.; Gryaznov, S.N.; Liquier, J.; Garestier, T.; Hélène, C.; Thailandier, E. Nucleic Acids Res., 1997, 25, 1782. [268] Giovannageli, C.; Perrouault, L.; Escudé, C.; Gryaznov, S.; Hélène, C. J. Mol. Biol., 1996, 261, 386. [269] Browne, K.A.; Dempcy, R.O.; Bruice, T.C. Proc. Natl. Acad. Sci. USA, 1995, 92, 7051. [270] Dempcy, R.O.; Browne, K.; Bruice, T.C. J. Am. Chem. Soc., 1995, 117, 6140. [271] Arya, D.P.; Bruice, T.C. J. Am. Chem. Soc., 1998, 120, 12419. [272] Arya, D.P.; Bruice, T.C. J. Am. Chem. Soc., 1999, 121, 10680. [273] Arya, D.P.; Bruice, T.C. Proc. Natl. Acad. Sci. USA, 1999, 96, 4384. [274] Letsinger, R.L.; Singman, C.N.; Histand, G.; Salunke, M. J. Am. Chem. Soc., 1988, 110, 4470. [275] Bailey, C.D.; Dagle, J.M.; Weeks, D.L. Nucleic Acids Res., 1998, 26, 4860.

[253] Wang, S.; Kool, E.T. Nucleic Acid Res., 1995, 23, 1157. [254] Vickers, T.A.; Wyatt, J.R.; Burckin, T.; Bennett, C.F.; Freier, S.M. Nucleic Acids Res., 2001, 29, 1293. [255] Torigoe, H.; Hari, Y.; Sekiguchi, M.; Obika, S.; Imanishi, T. J. Biol. Chem., 2001, 276, 2354. [256] Jones, R.J.; Swaminathan, S.; Milligan, J.F.; Wadwani, S.; Froehler, B.C.; Matteucci, M.D. J. Am. Chem. Soc., 1993, 115, 9816 [257] Stein, C.A.; Kreig, A.; [Eds). Applied Antisense Oligonucleotides Technology, John Wiley & Sons: New York, 1998.

[276] Laurent, A.; Naval, M.; Debart, F.; Vasseur, J.J.; Rayner, B. Nucleic Acids Res., 1999, 27, 4151. [277] Jung, P.M.; Histand, G.; Letsinger, R.L. Nucleosides & Nucleotides, 1994, 13, 1597. [278] Chatuverdi, S.; Horn, T.; Letsinger, R.L. Nucleic Acids Res., 1996, 24, 2318. [279] Robles, J.; Ibáñez, V.; Grandas, A.; Pedroso, E. Tetrahedron Lett., 1999, 40, 7131. [280] Nielsen, P.E.; Egholm, M. Peptide Nucleic Acids. Protocols and Applications. Horizon Scientific Press: Wymondham. 1999.

[258] Uhlmann, E.; Peyman, A. Chem. Rev., 1990, 90, 543. [281] Chopra, I. Exp. Opin. Invest. Drugs, 1999, 8, 1203. [259] Latimer, L.J.P.; Hampel, K.; Lee, J.S. Nucleic Acid Res., 1989, 17, 1549.

[282] Nielsen, P.E. Annu. Rev. Biophys. Biomol. Struct., 1995, 24, 167

[260] Hacia, J.G.; Wold, B.J.; Dervan, P.B. Biochemistry, 1994, 33, 5367.

[283] Hyrup, B.; Nielsen, P.E. Bioorg Med. Chem., 1996, 4, 5-23.

[261] Lacoste, J.; Francois, J.C.; Hélène, C. Nucleic Acid Res., 1997, 25, 1991.

[284] Uhlmann, E.; Peyman, A.; Breipohl, G.; Will, D.W. Angew. Chem. Int. Ed., 1988, 37, 2796.

[262] Matteucci, M.; Lin, K.Y.; Butcher, S.; Moulds, C. J. Am. Chem. Soc., 1991, 113, 7767. [263] Trapane, T.L.; Hogrefe, R.I.; Reynold, M.A.; Kan, L.S.; Ts’o, P.O.P. Biochemistry, 1996, 35, 5495.

[285] Hanvey, J.C.; Peffer, N.J.; Bisi, J.E.; Thomson, S.A.; Cadilla, R.; Josey, J.A.; Ricca, D.J.; Hassman, C.F.; Bonham, M.A.; Au, K.G.; Carter, S.G.; Bruckenstein, D.A.; Boyd, A.L.; Noble, S.A.; Babiss, L.E. Science, 1992, 258, 1481.

[264] Kiber-Herzog, L.; Kell, B.; Zon, G.; Shinozuka, K.; Mizan, S.; Wilson, W.D. Nucleic Acids Res., 1990, 18, 3534.

[286] Stirchak, E.P.; Summerton, J.E.; Weller, D.D. Nucleic Acid Res., 1989, 17, 6129.

[265] Jones, R.J., Swaminathan, S.; Milligan, J.F.; Wadwani, S.; Matteucci, M. J. Org. Chem., 1993, 58, 2983.

[287] Summerton, J.; Weller, D. Antisense Nucleic Acid Drug Dev., 1997, 7, 187.

[266] Escudé, C.; Giovannangeli, C.; Sun, J.S.; Lloyd, D.H.; Chen, J.K.; Gryaznov, S.M.; Garestier, T.; Hélène, C. Proc. Natl. Acad. Sci. USA., 1996, 93, 4365.

[288] Summerton, J.; Stein, D.; Huang, S.B.; Matthews, P.; Weller, D.; Partridge, M. Antisense Nucleic Acid Drug Dev., 1996, 6, 267.



1365

[289] Taylor, M.G.; Paulaskis, J.D.; Weller, D.D.; Kobzik, L. J. Biol. Chem., 1996, 271, 17445.

[312] Kuwahara, M.; Arimitsu, M.; Sisido, M. J. Am. Chem. Soc., 1999, 121, 256.

[290] Basye, J.; Trent, J.O.; Gao, D.; Ebbinghaus, S.W. Nucleic Acids Res., 2001, 29, 4873.

[313] Völker, J.; Klump, H.H. Biochemistry, 1994, 33, 13502.

[291] Lacroix, L.; Arimondo, P.B.; Takasugi, M.; Hélène, C.; Mergny, J.L. Biochem. Biophys. Res. Commun., 2000, 270, 363. [292] Nielsen, P.E.; Egholm, M.; Berg, R.H.; Buchardt, O. Science, 1991, 254, 1497-1500. [293] Egholm, M.; Buchardt, O.; Nielsen, P.E.; Berg, R.H. J. Am. Chem. Soc., 1992, 114, 1895. [294] Egholm, M.; Buchardt, O.; Christensen, L.; Behrens, C.; Freier, S.M.; Driver, D.A.; Berg, R.H.; Kim, S.K.; Norden, B.; Nielsen, P.E. Nature, 1993, 365, 566. [295] Doyle, D.F.; Braasch, D.A.; Simmons, C.G.; Janowski, B.A.; Corey, D.R. Biochemistry, 2001, 40, 53. [296] Wittung, P.; Nielsen, P.; Norden, B. J. Am. Chem. Soc., 1997, 119, 3189. [297] Wittung, P.; Nielsen, P.; Norden, B. Biochemistry., 1997, 36, 7973. [298] Nielsen, P.E.; Christensen, L. J. Am. Chem. Soc., 1996, 118, 2287. [299] Lohse, J.; Dahl, O.; Nielsen, P.E. Proc. Natl. Acad. Sci. USA, 1999, 96, 11804. [300] Ishihara, T.; Corey, D.R. J. Am. Chem. Soc., 1999, 121, 2012. [301] Kurakin, A.; Larsen, H.J., Nielsen, P.E. Chem. & Biol., 1988, 5, 81. [302] Betts, L.; Josey, J.A.; Veal, J.M.; Jordan, S.R. Science, 1995, 270, 1838. [303] Almarsson, O.; Bruice, T.; Kerr, J.; Zuckermann, R.N. Proc. Natl. Acad. Sci. USA, 1993, 90, 7518.

[314] Gaffney, B.L.; Kung, P.P.; Wang, C.; Jones, R.A. J. Am. Chem. Soc., 1995, 117, 12281. [315] Plum, E.G.; Park, Y.W.; Singleton, S.F.; Dervan, P.B.; Breslauer, K.J. Proc. Natl. Acad. Sci. USA, 1990, 87, 9436. [316] Singleton, S.F.; Dervan, P.B. Biochemistry, 1992, 31, 10995. [317] Moser, H.E.; Dervan, P.B. Science, 1987, 238, 645. [318] Maher, L.J.; Dervan, P.B. Biochemistry, 1990, 29, 8820. [319] Kiessling, L.L.; Griffin, L.C.; Dervan, P.B. Biochemistry, 1992, 31, 2829. [320] Sagamoto, N.; Wu, P.; Hara, H.; Kawamoto, Y. Biochemistry, 2001, 40, 9396. [321] Ono, A.; Ts'o, P.O.P.; Kan, L.S. J. Am. Chem. Soc., 1991, 113, 4132. [322] Ono, A.; Ts'o, P.O.P.; Kan, L.S. J. Org. Chem., 1992, 57, 3225. [323] Chin, T-M.; Lin S-B.; Lee, S-Y.; Chang, M-L.; Cheng, A.Y-Y.; Chang, F-C.; Pasternack, L.; Huang, D-H.; Kan, LS. Biochemistry, 2000, 39, 12457. [324] Egholm, M.; Christensen, L.; Dueholm, K.L.; Buchardt, O.; Coull, J.; Nielsen, P.E. Nucleic Acids Res., 1995, 23, 217. [325] Kuhn, H.; Demidov, M.V.; Frank-Kamenetskii, M.D.; Nielsen, P.E. Nucleic Acids Res., 1998, 26, 582. [326] Von Krosigk, U.; Benner, S.A. J. Am. Chem. Soc., 1995, 117, 5361. [327] Xiang, G.; Soussou, W.; McLaughlin, L.W. J. Am. Chem. Soc., 1994, 116, 11155.

[304] Sen, S.; Nilsson, L. J. Am. Chem. Soc., 1998, 120, 619.

[328] Berressem, R.; Engels, J.W. Nucleic Acids Res. 1995, 23, 3465.

[305] Topham, C.M.; Smith, J. J. Mol. Biol., 1999, 292, 10171038.

[329] Xiang, G.; Bogacki, R.; McLaughlin, L.W. Nucleic Acids Res., 1996, 24, 1963.

[306] Soliva, R.; Sherer, E.; Luque, F.J.; Laughton, C.A.; Orozco, M. J. Am. Chem. Soc., 2000, 122, 5997.

[330] Xiang, G.; McLaughlin, L.W. Tetrahedron, 1998, 54, 375.

[307] Brown, S.C.; Thomson, S.A.; Veal, J.M.; Davis, D.G. Science, 1994, 265, 777. [308] Erikson, M.; Nielsen, P.E. Nature Struct Biol., 1996, 3, 410. [309] Leijon, M.; Gräslund, A.; Nielsen, P.E.; Buchardt, O.; Norden, O.; Kristensen, S.M.; Eriksson, M. Bicohemistry, 1994, 33, 9820.

[331] Parsch, U.; Engels, J.W. Chem. Eur. J., 2000, 6. 2409. [332] Bédu, E.; Benhida, R.; Devys, M.; Fourney, J-L. Tetrahedron Lett., 1999, 40, 835. [333] Miller, P.S.; Bhan, P.; Cushman, C.D.; Trapane, T.L. Biochemistry, 1992, 31, 6788. [334] Jetter, M.C.; Hobbs, F.W. Biochemistry, 1993, 32, 3249.

[310] Uhlmann, E.; Peyman, A.; Breipohl, G.; Will, D.W. Angew. Chem. Int. Ed. 1998, 37, 2797.

[335] Miller, P.S.; Bi, G.; Kipp, S.A.; Fok, V.; Delong, R.K. Nucleic Acids Res., 1996, 24, 730.

[311] Schütz, R.; Cantin, M.; Roberts, C.; Greiner, B.; Uhlmann, E.; Leumann, C. Angew. Chem. Int. Ed., 2000, 39, 1250.

[336] Krawczyk, S.H.; Milligan, J.F.; Wadwani, S.; Moulds, C.; Froehler, B.C.; Matteucci, M.D. Proc. Natl. Acad. Sci. USA, 1992, 89, 3761.

1366


Orozco et al.

[337] Giessner-Prette, C.; Pullman, B.J. J. Theor. Biol. , 1977, 65, 189.

[360] Blanalt-Feidt, S.; Doronina, S.O.; Behr, J-P. Nucleosides & Nucleotides, 1999, 18, 605.

[338] Guschlbauer, W.; Duplaa, A.M.; Guy, A.; Téoule, R.; Fazakerley, G.V. Nucleic Acids Res., 1991, 19, 1753.

[361] Blanalt-Feidt, S.; Doronina, S.O.; Behr, J-P. Tetrahedron Lett., 1999, 40, 6229.

[339] Ushijima, K.; Ishibashi, T.; Yamakawa, H.; Tsukahara, S.; Takai, K.; Maruyama, T.; Takaku, H. Biochemistry, 1999, 38, 6570.

[362] Lee, J.S.; Johnson, D.A.; Morgan, A.R. Nucleic Acids Res. , 1979, 6, 3073.

[340] Rao, T.S.; Durland, R.H.; Revankar, G.R. J. Heterocycl. Chem., 1994, 31, 935. [341] Hunziker, J.; Priestley, E.S.; Brunar, H.; Dervan, P.B. J. Am. Chem. Soc., 1995, 117, 2661. [342] Brunar, H.; Dervan, P.B. Nucleic Acids Res., 1996, 24, 1987. [343] St. Clair, A.; Xiang, G.; McLaughlin, L.W. Nucleosides & Nucleotides, 1998, 17, 925. [344] Koh, J.S.; Dervan, P.B. J. Am. Chem. Soc., 1992, 114, 1470. [345] Priestly, E.S.; Dervan, P.B. J. Am. Chem. Soc., 1995, 117, 4761. [346] Radhakrishnan, J.; Patel, D.J.; Priestley, E.S.; Nash, H.M.; Dervan, P.B. Biochemistry, 1993, 32, 11228. [347] Marfurt, J.; Hunziker, J.; Leumann, C. Bioorg. Med. Chem. Lett., 1996, 6, 3021. [348] Marfurt, J.; Parel, S.P.; Leumann, C. Nucleic Acids Res., 1997, 25, 1875. [349] Marfurt, J.; Leumann, C. Angew. Chem. Int. Ed. Engl., 1998, 37, 175. [350] Mills, M.; Völker, J.; Klump, H.H. Biochemistry, 1996, 35, 13338. [351] Bates, P.J.; Laughton, C.A.; Jenkins, T.C.; Capaldi, D.C.; Roselt, P.D.; Reese, C.B.; Neidle, S. Nucleic Acids Res., 1996, 24, 4176. [352] Cassidy, S.A.; Slickers, P.; Trent, J.O.; Capaldi, D.C.; Roselt, P.D.; Reese, C.B.; Neidle, S. Nucleic Acids Res., 1997, 25, 4891. [353] Hildbrand, S.; Blaser, A.; Parel, S.P.; Leumann, C.J. J. Am. Chem. Soc., 1997, 119, 5499. [354] Hildbrand, S.; Leumann, C.J. Angew. Chem. Int. Ed. Engl., 1996, 35, 1968. [355] Pudlo, J.S.; Wadwani, S.; Milligan, J.F.; Matteucci, M.D. Bioorg. Med. Chem. Lett., 1994, 4, 1025. [356] Doronina, S.O.; Behr, J. P. Chem. Soc. Review, 1997, 26, 63.

[363] Lee, J.S.; Woodsworth, M.L.; Latimer, L.J.; Morgan, A.R. Nucleic Acids Res., 1984, 12, 6603. [364] Povsic, T.J.; Dervan, P.B. J. Am. Chem. Soc., 1989, 111, 3059. [365] Xodo, L.E.; Manzini, G.; Quadrifoglio, F.; van der Marel, G.A.; van Boom, J.H. Nucleic Acids Res., 1991, 19, 5625. [366] Lee, J.S.; Woodsworth, M.L.; Latimer, L.J.P. Biochemistry, 1984, 23, 3277. [367] Shimizu, M.; Koizumi, T.; Inoue, H.; Ohtsuka, E. Bioorg. Med. Chem. Lett., 1994, 4, 1029. [368] Ferrer, E.; Shevchenko, A.; Eritja, R. Bioorg. Med. Chem., 2000, 8, 291. [369] Singleton, S.F.; Dervan, P.B. Biochemistry, 1992, 31, 10995. [370] Froehler, B.C.; Ricca, D.J. J. Am. Chem. Soc., 1992, 114, 8320. [371] Barawkar, D.A.; Kumar, V.A.; Ganesh, K.N. Biochem. Biophys. Res. Commun., 1994, 205, 1665. [372] Barawkar, D.A.; Rajeev, K.G.; Kumar, V.A.; Ganesh, K.N. Nucleic Acids Res., 1996, 24, 1229. [373] Rajeev, K.G.; Jadhav, V.R.; Ganesh, K.N. Nucleic Acids Res., 1997, 25, 4187. [374] Hampel, K.J.; Crosson, P.; Lee, J.S. Biochemistry, 1991, 30, 4455. [375] Thomas, T.; Thomas, T.J. Biochemistry, 1993, 32, 14068. [376] Tung, C.H.; Breslauer, K.J.; Stein, S. Nucleic Acids Res., 1993, 21, 5489. [377] Spink, C.H.; Chaires, J.B.; J. Am. Chem. Soc., 1995, 117, 12887. [378] Meyer, A.; Morvan, F.; Rayner, B.; Imbach, J-L. Nucleosides & Nucleotides, 1999, 18, 1631. [379] Ferrer, E.; Fàbrega, C.; Güimil-García, R.; Azorín, F.; Eritja, R. Nucleosides & Nucleotides, 1996, 15, 907. [380] Ferrer, E.; Wiersma, M.; Kazimierzczak, B.; Müller, C.W.; Eritja, R. Bioconjugate Chem., 1997, 8, 757.

[357] Doronina, S.O.; Behr, J-P. Tetrahedron Lett., 1998, 39, 547.

[381] Yang, L.; Liu, M.; Deng, W.; Wang, C.; Bai, C.; Kan, L-S. Biophysical. Chem., 1999, 76, 25.

[358] Robles, J.; Grandas, A.; Pedroso, E. Tetrahedron, 2001, 57, 179.

[382] Froehler, B.C.; Wadwani, S.; Terhorst, T.J.; Gerrard, S.R. Tetrahedron Lett., 1992, 33, 5307.

[359] Robles, J.; Grandas, A.; Pedroso, E. unpublished results.

[383] Colocci, N.; Dervan. P.B. J. Am. Chem. Soc., 1994, 116, 785.



1367

[384] Mills, M.; Arimondo, P.B.; Lacroix, L.; Garestier, T.; Klump, H.; Mergny, J.L Biochemistry, 2002, 41, 357.

[407] Milligan, J.F.; Krawczyk, S.H.; Wadmani, S.; Matteucci, M.D. Nucleic Acids Res., 1993, 21, 327.

[385] Wagner, R.W.; Matteucci, M.D.; Lewis, J.G.; Gutiérrez, A.J.; Moulds, C.; Froehler, B.C. Science, 1993, 260, 1510.

[408] Faraqi, A.F.; Krawczyk, S.H.; Matteucci, M.D.; Glazer, P.M. Nucleic Acids Res., 1997, 25, 633.

[386] Bijapur, J.; Keppler, M.D.; Berqvist, S.; Brown, T.; Fox, K.R. Nucleic Acids Res., 1999, 27, 1802.

[409] Rao, T.S.; Lewis, A.F.; Durland, R.H.; Revankar, G.R. Tetrahedron Lett., 1993, 34, 6709.

[387] Cuenoud, B.; Casset, F.; Husken, D.; Natt, F.; Wolf, R.M.; Altmann, K-H.; Martin, P.; Moser, H.E. Angew. Chem. Int. Ed. Engl., 1998, 37, 1288.

[410] Rao, T.S.; Lewis, A.F.; Hill, T.S.; Revankar, G.R Nucleosides & Nucleotides, 1995, 14, 1.

[388] Sollogoub. M.; Domínguez, B.; Fox, K.R.; Brown, T. J. Chem. Soc. Chem. Commun., 2000, 2315. [389] Michel, J.; Moreau, S. Nucleosides & Nucleotides, 1999, 18, 1633. [390] Ono, A.; Haginoya, N.; Kiyokawa, M.; Minakawa, N.; Matsuda, A. Bioorg. Med. Chem. Lett., 1994, 4, 361. [391] Staubli, A.B.; Dervan, P.B. Nucleic Acids Res., 1994, 22, 2637. [392] Durland, R.H.; Rao, T.S.; Jayaraman, K.; Revankar, G.R. Bioconjugate Chem., 1995, 6, 278. [393] Eldrup, A.B.; Nielsen, B.B.; Haaima, G.; Rasmussen, H.; Kastrup, J.S.; Christensen, C.; Nielsen, P.E. Eur. J. Org. Chem., 2001, 1781. [394] Michel, J.; Toulmé, J-J.; Vercauteren, J.; Moreau, S. Nucleic Acids Res., 1996, 24, 1127. [395] Michel, J.; Gueguen, G.; Vercauteren, J.; Moreau, S. Tetrahedron, 1997, 53, 8457. [396] Godde, F.; Toulmé, J-J.; Moreau, S. Biochemistry, 1998, 37, 13765.

[411] Aubert, Y.; Perrouault, L.; Hélène, C.; Giovannangeli, C.; Asseline, U. Bioorg. Med. Chem., 2001, 9, 1617. [412] Raynaud, F.; Asseline, U.; Roig, V.; Thuong, N.T. Tetrahedron, 1996, 52, 2047. [413] Dagle, J.M.; Weeks, D.L. Nucleic Acids Res., 1996, 24, 2143. [414] Kandimalla, E.R.; Agrawal, S. Nucleosides & Nucleotides, 1995, 14, 991. [415] Gamper, H.B.; Kutyavin, I.V.; Rhinehart, R.L.; Lokhov, S.G.; Reed, M.W.; Meyer, R.B. Biochemistry, 1997, 36, 14816. [416] Svinarchuk, F.; Cherny, D.; Debin, A.; Delain, E.; Malvy, C. Nucleic Acids Res., 1996, 24, 3858. [417] Griffin, L.C.; Kiessling, L.L.; Beal, P.A.; Gillespie, P.; Dervan, P.B. J. Am. Chem. Soc., 1992, 114, 7976. [418] Wang, E.; Koshlap, K.M.; Gillespie, P.; Dervan, P.B.; Feigon, J. J. Mol. Biol., 1996, 257, 1052. [419] Huang, C.Y.; Miller, P.S. J. Am. Chem. Soc., 1993, 115, 10456. [420] Verma, S.; Miller, P.S. Bioconjugate Chem., 1996, 7, 600.

[397] Arzumanov, A.; Godde, F.; Moreau, S.; Tolumé, J-J.; Weeds, A.; Gait, M.J. Helv. Chim. Acta, 2000, 83, 1424. [398] Godde, F.; Toulmé, J-J.; Moreau, S. Nucleic Acids Res. 2000, 28, 2977.

[421] Huang, C.Y.; Bi, G.; Miller, P.S. Nucleic Acids Res., 1996, 24, 2606. [422] Guzzo-Pernell, N.; Tregear, G.W.; Haralambidis, J.; Lawlor, J.M. Nucleosides & Nucleotides, 1998, 17, 1191.

[399] Stilz, H.U.; Dervan, P.B. Biochemistry, 1993, 32, 2177. [400] Raynaud, F.; Asseline, U.; Roig, V.; Thuong, N.T. Tetrahedron, 1996, 52, 2047. [401] Roig, V.; Kurfust, R.; Thuong, N.T. Tetrahedron Lett., 1993, 34, 1601. [402] Parel, S.P.; Leumann, C.J. Nucleic Acids Res., 2001, 29, 2260. [403] Olivas, W.M.; Maher-III, L.J. Nucleic Acids Res., 1995, 23, 1936.

[423] Zimmermann, S.C.; Schmitt, P. J. Am. Chem. Soc., 1995, 117, 10769. [424] Lehmann, T.E.; Greenberg, W.A.; Liberles, D.A.; Wada, C.K.; Dervan, P.B. Helv. Chim. Acta, 1997, 80, 2002 [425] Guianvarc’h, D.; Benhida, R.; Fourrey, J.L.; Maurisse, R.; Sun, J.S. Chem. Comm., 2001, 1814. [426] Purwanto, M.G.M.; Lengeler, D.; Weisz, K. Tetrahedron Lett., 2002, 43, 61. [427] Lengeler, D.; Weisz, K. Tetrahedron Lett., 2001, 42, 1479.

[404] Marathias, V.M.; Sawicki, M.J.; Bolton, P.H. Nucleic Acids Res., 1999, 27, 2860.

[428] Saito, A.; Kuroda, R. Tetrahedron Lett., 1999, 40, 4837.

[405] Rao, T.S.; Durland, R.H.; Seth, D.M.; Myrick, M.A.; Bodepudi, V.; Revankar, G.R. Biochemistry, 1995, 34, 765.

[429] Horne, D.A.; Dervan, P.B. J. Am. Chem. Soc., 1990, 112, 2435.

[406] Gee, J.E.; Revankar, G.R.; Rao, T.S.; Hogan, M.E. Biochemistry, 1995, 34, 2042.

[430] McCurdy, S.; Moulds, C.; Froehler, B. Nucleosides & Nucleotides, 1991, 10, 287.

1368


[431] De Napoli, L.; Messere, A.; Montesarchio, D.; Pepe, A.; Piccialli, G.; Varra, M. J. Org. Chem., 1997, 62, 9024. [432] Barone, G.; De Napoli, L.; Di Fabio, G.; Giancola, C.; Messere, A.; Montesarchio, D.; Petraccone, L.; Piccialli, G. Bioorg. Med. Chem., 2001, 9, 2895. [433] Asseline, U.; Thuong, N.T. Tetrahedron Lett., 1994, 35, 5221. [434] Zhou, B.W.; Marchand, C.; Asseline, U.; Thuong, N.T.; Sun, J.S.; Garestier, T.; Hélène, C. Bioconjugate Chem., 1995, 6, 516. [435] Ueno, Y.; Ogawa, A.; Nakagawa, A.; Matsuda, A. Bioorg. Med. Chem. Lett., 1997, 6, 2817.

Orozco et al.

[447] Durland, R.H.; Rao, T.S.; Bodepudi, V.; Seth, D.M.; Jayaraman, K.; Revankar, G.R. Nucleic Acids Res., 1995, 23, 647. [448] Kukreti, S.; Sun, J.S.; Loakes, D.; Brown, D.; Nguyen, C.H.; Bisagni, E.; Garestier, T.; Hélène, C. Nucleic Acids Res., 1998, 26, 2179. [449] Durland, R.H.; Rao, T.S.; Revankar, G.R.; Tinsley, J.H.; Myrick, M.A.; Seth, D.M.; Rayford, J.; Singh, P.; Jayaraman, K. Nucleic Acids Res., 1994, 22, 3233. [450] Amasova, O.A.; Fresco, J.R. Nucleic Acids Res., 1999, 27, 4632. [451] Eritja, R.; Ferrer, E.; Güimil García, R.; Orozco, M. Nucleosides & Nucleotides, 1999, 18, 1619.

[436] Jayasena, S.D.; Johnston, B.H. Biochemistry, 1992, 31, 320. [437] Jayasena, S.D.; Johnston, B.H. Biochemistry, 1993, 32, 2800. [438] Jayasena, S.D.; Johnston, B.H. Nucleic Acids Res., 1992, 20, 5279. [439] Kessler, D.J.; Pettitt, B.G.; Cheng, Y.K.; Smith, S.R.; Jayaraman, K.; Vu, H.M.; Hogan, M.E. Nucleic Acids Res., 1993, 21, 4810.

[452] Mokhir, A.A.; Connors, W.H.; Richert, C. Nucleic Acids Res., 2001, 29, 3674. [453] Rao, T.S.; Hogan, M.E.; Revankar, G.R. Nucleosides & Nucleotides, 1994, 13, 95. [454] Trapane, T.L.; Christopherson, M.S.; Roby, C.D.; Ts’o, P.O.P.; Wang, D. J. Am. Chem. Soc., 1994, 116, 8412. [455] Bandaru, R.; Hashimoto, H.; Switzer, C. J. Am. Chem. Soc., 1995, 60, 786.

[440] Doromina, S.O.; Behr, J.P. Chem. Soc. Rev., 1997, 63. [441] Beal, P.A., Dervan, P.B. Nucleic Acids Res., 1992, 20, 2773. [442] Griffin, L.C.; Dervan, P.B. Science, 1989, 245, 967. [443] Jiang, L.; Russu, I.M. Nucleic Acids Res., 2001, 29, 4231. [444] Kandimalla, E.R.; Manning, A.N.; Venkataraman, G.; Sasisekharan, V.; Agrawal, S. Nucleic Acids. Res., 1995, 23, 4510. [445] Loakes, D. Nucleic Acids Res., 2001, 29, 2437. [446] Orson, F.M.; Klysik, J.; Bergstrom, D.E.; Ward, B.; Glass, G.A.; Hua, P.; Kinsey, B.M. Nucleic Acids Res., 1999, 27, 810.

[456] Rana, V.S.; Barawkar, D.A.; Ganesh, K.N. J. Org. Chem., 1996, 61, 3578. [457] Rana, V.S.; Ganesh, K.N. Nucleic Acids Res., 2000, 28, 1162. [458] Güimil-García, R.; Bachi, A.; Eritja, R.; Luque, F.J.; Orozco, M. Bioorg. Med. Chem. Lett., 1998, 8, 3011. [459] Kawai, K.; Saito, I.; Sugiyama, H. Tetrahedron Lett., 1998, 39, 5221. [460] Hattori, M.; Frazier, J.; Milles, H.T. Biochemistry, 1975, 18, 5033. [461] Hattori, M.; Frazier, J.; Milles, H.T. Biopolymers, 1976, 15, 523. [462] Kumar, R.K.; Gunjal, A.D.; Ganesh, K.N. Biochem. Biophys. Res. Comm., 1994, 204, 788.