Non-Nucleosidic Analogues of Polyaminonucleosides and Their Influence on Thermodynamic Properties of Derived Oligonucleotides

The rationale for the synthesis of cationic modified nucleosides is higher expected nuclease resistance and potentially better cellular uptake due to an overall reduced negative charge based on internal charge compensation. Due to the ideal distance between cationic groups, polyamines are perfect counterions for oligodeoxyribonucleotides. We have synthesized non-nucleosidic analogues built from units that carry different diol structures instead of sugar residues and functionalized with polyamines. The non-nucleosidic analogues were attached as internal or 5′-terminal modifications in oligodeoxyribonucleotide strands. The thermodynamic studies of these polyaminooligonucleotide analogues revealed stabilizing or destabilizing effects that depend on the linker or polyamine used.


Introduction
The polyanionic character of nucleic acids and their synthetic fragments is a barrier for their introduction into cells. Many laboratories try to improve nucleic acid delivery to cells by synthesis of cationic bioconjugates [1][2][3], bioconjugate analogues with reduced polyanionic character [4][5][6], or a great variety of polymeric transfection media [7][8][9][10][11][12]. On the other hand, there are studies to elucidate a OPEN ACCESS specific mode of small cationic molecular drugs interactions with biomolecules (e.g., nucleic acids, proteins) useful in developing their more active analogues [13]. Among this clinically important class of compounds are antibiotics carrying aminosugar residues [14].
Chemically modified oligonucleotides have been utilized as indispensable materials for DNA gene therapy [15,16], gene regulation [17,18], chip technology [19,20] and recent nanotechnology [21][22][23] because of their hybridization affinity for target DNA and/or RNA molecules [24]. The polyanionic character of antisense and siRNA oligonucleotides is a major cause of insufficient cellular uptake and side effects such as binding to serum proteins. The combination of nucleotides with aminoalkyl chains greatly enhances the variety of possible structures as well as their potential application [24]. Thus, oligonucleotides possessing cationic functionalities in addition to the anionic phosphate backbone have been shown to exhibit promising properties [25][26][27][28]. Polyamines, putrescine, spermine and spermidine, are involved in the regulation of gene function [29,30]. In vitro, they stabilize DNA and RNA duplexes [31,32], especially ones with imperfect base pairing [33]. The distance between amino groups of three and four carbon atoms is practically the same as the distance between phosphate anions in the backbone of DNA making polyamines the perfect compounds for creating "zwitterionic" oligonucleotides [27]. It is known that polyamines interact with DNA and RNA in different ways. The electrostatic binding is performed via water molecules, by hydrogen bonding with polar functional groups or with hydrophobic surfaces of nucleobases. There have been numerous studies aimed at determining these interactions using NMR imaging, circular dichroism, Raman spectroscopy, IR spectroscopy, X-ray crystallography and differential scanning calorimetry. In spite of these extensive studies, precise mechanisms for the interaction between polyamine and DNA is still not fully understood in vivo [34].
The non-nucleosidic analogs of polyaminonucleosides have not been examined so far. Therefore, our aim was to elaborate versatile procedures for synthesis of non-nucleosidic polyamine derivatives and learn their properties within oligonucleotide chains. The choice of carbon chain skeletons of analogues ( Figure 1) was based on the extent of their commercial availability. The obtained polyamine building blocks were incorporated at the 5′-end and internal positions within an oligodeoxyribonucleotide chain and the resulting conjugates were evaluated for their hybridization properties.

Chemistry
The synthesis of the non-nucleosidic polyamine derived phosphoramidite building blocks containing different linkers is shown in Scheme 1. We decided to use 2,2-bis(hydroxymethyl)-propionic acid (bis-MPA, 1) that was earlier applied in dendrimer synthesis [35,36].
Bis-MPA (1) was protected with benzaldehyde [37] as it is more lipophilic than a isopropylidene group preferred to ease the final workup procedure and increase yield when using an excess of polar polyamines [38,39]. The protection of hydroxyl groups in bis-MPA gives rise to formation of two stereoisomers in the ratio ca 2:1, however, further reactions were carried out without separation of isomers. Several approaches for activation of carboxyl function (2) were checked. Reactions with 1,4-butylamine as a model amine in the presence of carbonyldiimidazole or 2-chloro-4,6-dimethoxy-1,3,5-triazine gave the expected n-butylamide derivative (results not shown). The best results were obtained when activation was performed with thionyl chloride in the presence of pyridine and traces of DMF. Thus, reaction with molar excess of putrescine (1,4-diaminobutane) at ambient conditions led to N-(4-aminobutyl)-5-methyl-2-phenyl-1,3-dioxane-5-carboxyamide 2 in 60% yield (Scheme 1).
We also synthesized the analogs based on the enantiomers (7) and (7a) which were coupled with excess of spermine in dry MeOH. Due to the lack of a chromophore in synthesized compounds, a reaction control was performed using TLC plates stained with a solution of fluorescein (free acid) in acetonitrile. Spermine derivatives (8, 8a) were isolated by several extractions with large amounts of organic solvent and used in the next step without additional purification. The configuration of the branching point in these linkers is retained throughout the synthetic procedure. After protection of the amine function with trifluoroacetyl groups, 1,3-dioxolane was hydrolyzed to prepare polyamine non-nucleosidic derivatives for selective 4,4′-dimethoxytritylation. In order to avoid transacetylation of trifluoroacetyl groups during the isopropylidene cleavage of 9 we used p-TsOH instead acetic acid. Otherwise, amino groups would be at least in part protected with Ac instead of TFA and their deprotection with ammonia would not be possible under the conditions of the final deprotection of oligonucleotides. Commercially available precursor α-hydroxy-γ-butyrolactone (12) was used as a starting material to obtain a polyaminonucleoside analogue (17). This compound provides a three carbon distance between the phosphate groups after introduction into the oligodeoxyribonucleotide strand. It also contains a chiral carbon center with a secondary hydroxyl group and the carbonyl group corresponding to the 3′-hydroxyl and 2′-carbon of nucleosidic residues, respectively. The lactone-spermine conjugate was subsequently obtained after protection of the secondary hydroxyl group of the lactone (13) with lipophilic and a bulky t-butyldimethylsilyl (TBDMS) group. Thus, this eased the isolation of the product (14) resulting from the coupling reaction with spermine (Scheme 1). The reaction of protected lactone 13 and unprotected spermine was performed by microwave synthesis to achieve the desired product 14 with 64% yield, even when both reagents were used in equimolar ratio. To eliminate side reactions during the condensation step of DNA synthesis, the protection of putrescine and spermine amino functions with trifluoroacetyl groups was ensured (as previously described [40]. For all non-nucleosidic analogue yields of trifluoroacetylation ranged from 58% (spermine derivatives) up to 70% (putrescine derivative) (Scheme 1). Conversion to the corresponding phosphoramidite derivative was preceded by removal of protecting groups of hydroxyl function with HCl (4) and p-TsOH (9, 9a) acids or Et3N*3HF (15). The protection 5′-OH-groups with a 4,4′-dimethoxytrityl group was used in a last step of the 3′-OH reaction with 2-cyanoethyl-N,N,N′,N′-tetraisopropylaminophosphane and 5-(ethylthio)-1H-tetrazole as the activator. However, due to similar physical properties, the separation of the products from impurities was difficult by silica gel column chromatography. This was resolved by precipitation from hexane. The spermine derivatives of (R) and (S) enantiomers of glyceric acid gained an additional chiral center as phosphoramidites (11, 11a). In the case of the polyamine derivative of α-hydroxy-γ-butyrolactone, opening of the lactone ring 14 led to a mixture of enantiomers. Thus, after phosphitylation, compound 17 as a mixture of enantiomers was obtained. All amidites were lyophilized and were stable during long-term storage at −20 °C.

Oligonucleotides
The phosphoramidites (6, 11, 11a, 17) were used for synthesis of polyamine analogs of oligodeoxyribonucleotides ( Figure 2) using a 12-mer as a reference sequence ( Table 1). The coupling efficiencies of these phosphoramidites were 53%-70% as determined by measuring detritylation. The modified oligodeoxyribonucleotides were obtained after a standard deprotection procedure, and their structures were confirmed by the MALDI-TOF ( Table 1). The oligodeoxyribonucleotides ON5-ON6 and ON9 were used as pure enantiomers and ON4 and ON8 as mixtures of diastereoisomers.  Figure 2. Non-nucleosidic polyamino-nucleoside analogues units in oligodeoxyribonucleotide strands.

Stability of Duplexes with Modified Oligodeoxyribonucleotides
The influence of conjugated polyamines on the stability of the DNA duplexes was studied using UV melting spectra. It was shown previously that polyamine conjugation to oligodeoxyribonucleotides results in stabilizing of DNA duplexes and triplexes [2][3][4]38,39,[41][42][43]. Recently, we described the NMR structure of a DNA duplex carrying a single spermine modified deoxycytidine unit [34]. This modification moderately stabilizes the DNA duplex ( Figure 3, dC Sp vs. RF1) and does not perturb the DNA structure.  (Table 2); ON4-ON6 and ON8-ON9-non-nucleosidic spermine analogues ( Table 2).
The 5′-end dangling modifications in oligodeoxyribonucleotide duplexes usually increase duplex stability and do not show large sequence dependence [44,45]. Since the terminal unpaired nucleotides are not involved in base pairing, stacking, electrostatic, perhaps to some extent, hydrophobic interactions are responsible for the thermodynamic effects of the 5′-and 3′-dangling ends. The nearest-neighborhood model assumes the duplex region for a dangling end to have the same calculated thermodynamic stability as a matching blunt-end duplex [32,33].
Thus, the stability was increased when X1 and X3 (Scheme 1) were incorporated at the 5′ position ( Table 1, entry ON4 and ON6), but for incorporations of X0 and X2 (Scheme 1) a moderate destabilizing effect was observed (Table 1, entry ON3 and ON5). Despite lack of a nucleobase and higher conformational flexibility at the 5′-end of the strand the observed stabilities are rather typical for 5′-end dangling units as described in the literature. Thermodynamic data (Table 2, Figure 1) show that X3 results in ΔΔG (−0.3 kcal/mol) similar to unpaired nucleotide (−0.1 to −0.5 kcal/mol) at the 5'-end. The change of ΔG for X1 is even higher (ΔΔG = −0.64 kcal/mol). These results suggest that the lack of a nucleobase does not affect the thermal stability of the duplexes. This might indicate that increased duplex stability is provided by the three positive charges in the spermine chain. In the case of the putrescine residue, X0, which introduces one positive charge only, a small decrease of duplex stability was observed. (Figure 1, ON3). One can conclude that the small decrease in duplex stability observed for ON5 ( Figure 1) containing spermine attached to the glycerol isomer (X2) is caused by the structure and configuration of this particular linker. Next, we investigated the influence of polyamine derivatives X0, X1 and X2 on duplex stability, inserted as bulges in the middle of the sequence (Table 1, entry ON7-ON9). The changes of duplex stability caused by single nucleotide bulges differ and depend on the type of flanking bases [46,47]. Incorporation of X0-X2 resulted in a lowering of melting temperature independently of polyamine residue. However, the melting data suggest that the duplex formation occurs with the proper W-C base pairing despite of a slight increase in free energy for ON7-ON9 (Table 2, ΔΔG ca. 3-4.5 kcal/mol). Moreover, the effect in ON7 where the putrescine bulge (X0) is flanked by GC/CG pairs is practically the same as for ON8 and ON9 (spermine bulge, X1 and X2) flanked by AT/TA pairs. This can be attributed to the stronger stabilizing effect of a spermine residue when compared to putrescine-three positive charges vs. one. Yet, even three positive charges of a spermine residue do not neutralize the bulge effect as such. The observed changes of ΔG for the studied duplexes (ON7-ON9) are in the range observed for duplexes with a single bulge (ΔΔG, 2-6 kcal/mol) [46,47].
Linkers X1 and X2 differ in length by one carbon and this additional carbon in X1 makes this linker somehow less rigid, allowing for more favorable placement of the polyamine residue. Thus, the energy gain of 1.5 kcal/mol in ON4 when compared to ON5. Therefore, ON4 duplex is more stable than the unmodified duplex. When the same modified linkers (X1 and X2) are inserted as bulges in a more spatially "demanding environment", the difference of free energies ΔΔG°37 of ON8 (+3.63) and ON9 (+4.43) is smaller (0.8 kcal/mol). The influence of linker structure is less profound when the modification is inserted in the middle of chain.
Linkers X2 and X3 carrying a spermine residue differ only in the configuration of carbon (S and R respectively). Comparison of ΔΔG°37 of ON6 (−0.3 kcal/mol) and ON5 (+0.87 kcal/mol) suggests that X3 allows more favorable placement of polyamine residue This seems to indicate that in this case the spermine residue attached to a glycerol linker with R-configuration (X3) is closer to the duplex charged surface.
We would like to conclude that the overall effect of DNA modification with polyamine analogues seems to offer a convenient way to modify nucleic acids properties. An attachment of polyamine residues via various open chain linkers allows to maintain the general scheme of base pairing. Moreover, the structure and configuration of the linkers seems to have a higher influence on stability when placed at the 5′-end of DNA duplexes.

General Methods
All reagents were of analytical grade, obtained from commercial resources and used without further purification. For synthesis, solvents with quality pro analysis were used. Solvents were dried and distilled following standard methods and kept over molecular sieve. All reactions were carried at room temperature unless described otherwise. Column chromatography was performed with silica gel (Merck KGaA, Darmstadt, Germany, 200-630 mesh) and TLC was carried out on precoated plates (Merck silica gel 60, F254). All NMR spectra were recorded at 298 K on Bruker AVANCE II (400 MHz, 1 H, 13 C) and Varian Unity (300 MHz, 31 P) spectrometers (Bruker BioSpin GmbH, Rheinstetten, Germany and Varian, Inc., Palo Alto, CA, USA). Chemical shifts (δ) are reported in parts per million (ppm). J values are given in Hz. Mass spectra were recorded on the MicroTofQ mass spectrometer with electrospray ionization (ESI) sources (ESI source voltage of 3.2 kV, nebulization with nitrogen at 0.4 bar, dry gas flow of 4.0 L/min at temperature 220 °C) and Bruker Autoflex MALDI-TOF (Bruker Daltonik GmbH, Bremen, Germany). Microwave reactions was performed in domestic microwave oven (800 W, Amica, Wronki, Poland). Some NMR (Figures S1-S16) and MS (Figures S17-S22) spectra are available in Supplementary.

Synthesis of Monomers
Benzylidene-2,2-bis(oxymethyl)propionic acid (2) [37]. Benzaldehyde (4.2 mL, 40.7 mmol) was added to a well-stirred solution of 2,2-bis(hydroxymethyl)-propionic acid (1) (5 g, 37 mmol) in DMF (30 mL), followed by catalytic p-TsOH (0.355 g, 1.85 mmol) with stirring at room temperature for 4 days. The reaction was quenched with NH4OH/EtOH (1 mL, 1:1). The solvent was evaporated and the residue dissolved in DCM (100 mL) and washed with NaHCO3 (2 × 100 mL). Organic extracts were dried over anhydrous MgSO4 and filtered then evaporated under reduced pressure. The residue was purified by recrystallization from DCM to obtain product 2 as mixture of two isomers (77%). 1  N-(4-Aminobutyl)-5-methyl-2-phenyl-1,3-dioxane-5-carboxamide (3). Thionyl chloride (1.26 mL, 17.8 mmol) was added dropwise to ice-cooled solution of 2 (2 g, 8.9 mmol) in 45 mL DCM containing 10% of pyridine and 3 drops of DMF. The mixture was stirred for 2 h at room temperature, and the excess thionyl chloride was removed by several co-evaporations with a mixture of DCM and toluene. The brown residue was dissolved in DCM (46 mL) and the temperature was lowered to −10 °C. Putrescine (4.47 mL, 44.5 mmol) and triethylamine (1.4 mL) in DCM (2 mL) was added after 15 min. The mixture was stirred for 1h at room temperature, and the reaction was quenched with saturated aqueous NaHCO3 (3 mL). The mixture was extracted with DCM (2 × 100 mL). The combined organic extracts were dried over anhydrous MgSO4 and concentrated under vacuum and purified by chromatography with 4% MeOH in DCM as the eluent to give 3 (0.910 g, 60% yield) as a yellow oil.  (4) 3 (0.430 g, 1.47 mmol) was co-evaporated with pyridine (3 × 5 mL), dissolved in pyridine (15 mL), followed by the addition of N-methylimidazole (0.16 mL, 1.46 mmol). Then, trifluoroacetic anhydride (0.593 mL, 4.41 mmol) was added dropwise to the mixture cooled at 0 °C. The mixture was stirred for 30 min and then poured into saturated aqueous NaHCO3 (30 mL), and extracted with DCM (3 × 50 mL). The combined organic extracts were dried over anhydrous Na2SO4 and concentrated under vacuum. The residue was purified by chromatography using DCM as the eluent to give 4 (0.500 g, 52% yield) as an oil. 1 (5). 4 (0.240 g, 0.61 mmol) was dissolved in a mixture conc. aq HCl/EtOH (1:5, v/v) and refluxed for 72 h. The excess of HCl was removed by several co-evaporations with a mixture of methanol and toluene and finally a brown oil with was dried by co-evaporation with anhydrous pyridine (2 × 5 mL) and dissolved in anhydrous pyridine (2.4 mL). To this solution 4,4′-dimethoxytrityl chloride (0.243 g, 0.72 mmol) was added and the reaction was quenched after 3 h by adding saturated aqueous NaHCO3. The resulting solution was extracted with DCM and the combined organic extracts were washed with brine, dried over Na2SO4 and concentrated under vacuum. The residual yellow oil was purified by chromatography with 5% MeOH in DCM as the eluent to give 0.18 g (49%) 5 as a white foam.

Oligonucleotide Preparation
The DNA dodecamers (5′-CTC AAG CAA GCT-3′, 5′-AGC TTG CTT GAG-3′, 5′-CTC ACA TGC GCG-3′, 5′-CGC GCA TGT GAG-3′) were synthesized using DNA synthesizer Gene Assembler Plus from Pharmacia-LKB (Uppsala, Sweden) or K & A Laborgerate GbR DNA/RNA (Frankfurt am Main, Germany), using standard phosphoramidite chemistry. The non-nucleosidic analogues (6, 11, 11a, 17) were inserted at positions marked with an X as listed in the Table 1. For the modified phosphoramidites, two-fold excess of phosphoramidites (in comparison to the standard protocol) and a prolonged coupling step of 10 min were used. The oligomers were cleaved from the CPG-support with 32% aqueous ammonia (room temperature, 1 h). The deprotection under standard conditions using concentrated aqueous ammonia at 55 °C overnight allowed for removal of all protecting groups, including trifluoroacetyls [3,38]. The oligomers were purified by the TLC on Merck 60 F254 TLC plates with n-propanol/aqueous ammonia/water solution (55:35:10, by vol.) as an eluent. The product band (least mobile) was cut out, eluted with water and desalted with Waters Sep-pak C-18 cartridges. First, the solution containing the oligonucleotide was loaded onto the cartridge and the column was flushed with 10 mM ammonium acetate (10 mL). In the next step, the oligonucleotides were eluted by flushing the cartridge with 5 mL of 30% acetonitrile/water solution. The fraction with the product was evaporated to dryness and the purity of oligonucleotides was monitored using HPLC and confirmed (by MALDI-TOF spectrometry Autoflex, Bruker). Oligonucleotides were also purified in a subsequent reverse phase HPLC step (UFLC system with LC-20AD pump; C(18)2 100 Å column (15 cm × 4.6 mm); starting from 0.01 M triethylammonium acetate (pH 7.0) up to CH3CN:CH3COOEt3N (40%/60%). Identity was confirmed by mass spectroscopy on MALDI-TOF (Autoflex, Bruker Daltonik GmbH, Bremen, Germany). The isolated yields of modified oligonucleotides were at the range of 23%-57%: 26%-57% and 23%-26% for ON3-ON6 and ON7-ON9 respectively.

Thermodynamic Analysis
UV melting profiles of the DNA duplexes were obtained in a buffer containing 100 mM sodium chloride, 20 mM sodium cacodylate, 0.5 mM Na2EDTA, pH 7.0. Duplexes were used in the 10 −3 -10 −6 M concentration range. Single strand concentrations were calculated from absorbance above 80 °C with single strand extinction coefficients approximated by the nearest-neighbor model [51]. The temperature range, in a heating-cooling cycle, was 0-90 °C with a temperature gradient of 1 °C/min. Thermal-induced transitions of each mixture were monitored at 260 nm with a Beckman DU 650 spectrophotometer with a temperature controller. The thermodynamic parameters were determined from fits of data acc. to a two-state model with the MeltWin 3.5 software [52].

Conclusions
Nucleic acids that carry polycationic modifications have many therapeutic and biotechnological advantages. Our studies corroborate that non-nucleosidic analogues of polyaminooligonucleosides maintain affinity and easily form duplexes with complementary strands. In some cases, this may increase the stability of modified complexes of nucleic acids. These new properties based on the rationale of charge masking do not change the scheme of the secondary structure of nucleic acids. Thus, they can ease transferring of nucleic acids past cellular barriers.