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We describe the chemical synthesis and preliminary biophysical and biochemical characterization of a series of mRNA 5' end (cap) analogs designed as reagents for obtaining mRNA molecules with augmented translation efficiency and stability in vivo and as useful tools to study mRNA metabolism. The analogs share three structural features: (i) 5',5'- bridge elongated to tetraphosphate to increase their affinity to translation initiation factor eIF4E (ii) a single phosphorothioate modification at either the α, β, γ or δ-position of the tetraphosphate to decrease their susceptibility to enzymatic degradation and/or to modulate their interaction with specific proteins and (iii) a 2'-O-methyl group in the ribose of 7-methylguanosine, characteristic to Anti-Reverse Cap Analogs (ARCAs), which are incorporated into mRNA during in vitro transcription exclusively in the correct orientation. The dinucleotides bearing modified tetraphosphate bridge were synthesized by ZnCl2 mediated coupling between two mononucleotide subunits with isolated yields of 30–65%. The preliminary biochemical results show that mRNAs capped with new analogs are 2.5–4.5 more efficiently translated in a cell free system than m7GpppG-capped mRNAs, which makes them promising candidates for RNA-based therapeutic applications such as gene therapy and anti-cancer vaccines.
Dinucleoside 5',5'-polyphosphates fulfill numerous regulatory, signaling, or energetic functions in biological systems. A particular representative of this group of biologically important molecules is the m7G-containing cap present on the 5'-end of all eukaryotic messenger RNAs. Different types of cap structures exist in nature,1 but they all contain guanosine methylated at the N7-position connected by a 5', 5'-triphosphate bridge to the first nucleotide of the RNA transcript (Fig 1. A).
The cap plays a key role in a variety of cellular processes related to mRNA metabolism, including splicing, nucleo-cytoplasmic transport, intracellular targeting, translation, translational repression, and turnover.1,2 Therefore, synthetic analogs of the cap are widely employed as tools for studying these physiological processes as well as being useful in biotechnology. There are also promising potential clinical applications.
One application that has been intensively explored by our groups is the use of cap analogs as reagents for in vitro synthesis of capped mRNAs. Capped mRNAs may be employed for expression of proteins in either in vitro translation systems or in cultured cells. Recently, mRNA have emerged as an attractive alternative to classical DNA-vector based gene delivery due to several advantages; the use of mRNA avoids the risk of insertional mutagenesis, is effective in non-dividing cells, and does not require introduction into the nucleus because mRNA is translated in the cytoplasm.3
Synthesis of capped mRNA can be achieved by transcribing a DNA template in vitro with either a bacterial4 or bacteriophage5,6 RNA polymerase in the presence of all four NTPs and a dinucleotide cap analog such as m7GpppG. These polymerases normally initiate transcription with nucleophilic attack by the 3′-OH of GTP on the α-phosphate of the next nucleoside triphosphate specified by the DNA template. If a cap analog such as m7GpppG is present in the reaction mixture at a ~5:1 ratio to GTP, the transcription is initiated mainly by attack of 3′-OH of the cap dinucleotide rather than that of GTP, leading to formation of capped transcripts of the form m7GpppGpNp(Np)n. Unfortunately, attack can occur by the 3′-OH of either the Guo or m7Guo moieties of the cap dinucleotide, producing one third to one half transcripts capped in a reversed orientation, i.e., Gpppm7GpNp(Np)n.7 Such reverse-capped transcripts decreased the overall translational activity of mRNA. This problem was overcome several years ago by the introduction of anti-reverse cap analogs (ARCAs) bearing either 3′-O-methyl, 3’-H or 2′-O-methyl modifications in the m7Guo moiety, ensuring 100% correct orientation.8–10 ARCA-capped mRNAs were shown to have higher translational efficiency than m7GpppG capped mRNAs both in vitro8,10 and in cultured cells.11–13
One important advantage of the ARCA method over the post-transcriptional enzymatic capping is the option to introduce additional chemical modifications into the mRNA 5' end that can modify the behavior and properties of the mRNAs. Among dozens modified synthetic cap analogs examined, several have been identified that augment mRNA translational efficiency, both in vitro and in cultured cells.10,13 Two features of the modified cap structure have emerged that influence the overall yield of protein produced from the mRNA, particularly in cultured cells.11,13 The first one is the cap's intrinsic ability to compete with natural mRNAs for recruitment to the translational machinery, and the second is its susceptibility to enzymatic decapping.
During recruitment of mRNA to the translational machinery, the cap is specifically recognized by eukaryotic initiation factor 4E (eIF4E). mRNA recruitment to form the 48S pre-initiation complex is the rate-limiting step for initiation of translation under normal circumstances (absence of virus infection, cellular stress, etc.) and plays a central role in regulation of translation.14 The presence of cap greatly increases the rate of translation. Affinity of the cap for eIF4E, and hence for the translational machinery, can be increased by modifying the 5',5'-triphosphate bridge. One class of compounds with modifications of this type are cap analogs in which the 5',5'-bridge is extended from triphosphate to tetraphosphate, e.g. m7GppppG, m27,2'-OGppppG or m27,3'-OGppppG.10 Generally, these analogs have binding affinities to eIF4E that are ~9–10-fold higher than for the corresponding triphosphates and promote translation 20–30% more efficiently when introduced into mRNA. However, it should be mentioned that further increasing the affinity to eIF4E may exert opposite effect on mRNA translational efficiency. Surprisingly, it was found that extending the 5',5'-bridge to pentaphosphate increases the binding affinity of cap analogs to eIF4E by a factor ~3,5–5 compared to tetraphosphates, nonetheless, mRNAs capped with these analogs are translated with efficiency comparable or even lower than for corresponding triphosphates.10
A second important role of the mRNA cap is protecting mRNA against degradation.15 Exonucleolitic mRNA cleavage in the 5'→3' direction cannot take place until the cap is removed from mRNA 5' end by a decapping pyrophosphatase termed Dcp1/Dcp2 (a heterodimer consisting of a regulatory and catalytic subunit). The enzyme cleaves the cap, provided it is attached to an RNA of at least 20 nt, between the α and β phosphate moieties to release m7GDP and 5′-phosphorylated mRNA chain. This is then degraded by the exonuclease Xrn1.16,17 Both the 5'→3' and 3'→5' pathways play major roles in mRNA degradation.15 We have recently demonstrated that mRNAs capped with an ARCA analog containing a phosphorothioate moiety at the β position of the triphosphate bridge, m27,2’-OGppSpG (D2),18 were resistant to decapping by Dcp2 in vitro, resulting longer half-lives in cultured mouse mammary cells (257 min compared to 155 min for m27,2’-OGpppG-capped mRNA).13 An unexpected further benefit was that m27,2’-OGppSpG (D2)-capped mRNA was 2.4-more efficiently translated in these cells than m27,2’-OGpppG-capped mRNA.
In the present work, we aimed at obtaining analogs that would combine both features that can enhance the translational yield of mRNA: high affinity for eIF4E and resistance to cleavage by Dcp2 in the expectation that these two features would independently improve the translational properties of mRNA in living cells. We therefore synthesized a series of tetraphosphate ARCA analogs modified with phosphorothioate group at various positions of the 5',5'-tetraphosphate bridge (Fig. 1C).
The new tetraphosphate analogs could potentially be useful in other spheres of mRNA metabolism, based on our recent findings with triphosphate phosphorothioate ARCA analogs modified at different positions. Those modified at the γ-position were resistant to another decapping enzyme, DcpS, which acts on the products of 3′→5′ mRNA decay. DcpS cleaves only short capped 5'-oligonucleotides or m7GpppN dinucleotides released after processive degradation of mRNA by the exosome initiated from 3'-end. Cleavage is between the β and γ phosphates of the 5′,5′-triphosphate bridge, releasing m7GMP and a downstream (oligo)nucleotide.19 Resistance to DcpS is an important factor in design of long-lasting cap analogs that can inhibit translation.
Recently, a cap analog modified at the α-position, m27,2’-OGpppSG (D2), was shown to enhance microRNA-mediated translational repression by a factor of 2 with no change in translation efficiency, providing new insights into the mechanism of this recently discovered but poorly understood process.20
We designed and prepared a series of four cap analogs 1–4 bearing a single phosphorothioate moiety at either the α-, β-, γ-, or δ-positions of the tetraphosphate chain and 2′-O-methyl group (ARCA modification) in the m7Guo moiety. We refer to these as 4P-S-ARCAs (Fig. 1C). The synthetic routes provide these analogs with good yields and can also be applied to the synthesis of a variety of other biologically relevant dinucleoside polyphosphates. Due to the presence of a P-stereogenic center, each 4P-S-ARCA exists as two P-diastereomers, designated D1 and D2 based on their elution order during reverse phase (RP) HPLC. Two pairs of diastereomers were successfully resolved by RP HPLC, but we could not achieve this for the other two. Subsequently, the new analogs were subjected to biophysical and biochemical characterization. We determined their binding affinities for eIF4E, susceptibility to decapping by DcpS, and translational efficiencies in a rabbit reticulocyte lysate (RRL) system. Other biochemical properties of mRNAs capped with these analogs, including susceptibility to the human decapping enzyme Dcp2, translational efficiency in cultured mammalian cells, and half-life in mammalian cells, are currently under investigation. Preliminary results suggest that mRNA capped with a mixture of diastereomers (D1 plus D2) of m27.2'-OGpppSpG (compound 2) is resistant to in vitro decapping by hDcp2 but mRNA capped with m27.2'-OGppSppG (compound 3) is not.21
The key step in the preparation of each new analog (1–4) was formation of a thio-modified tetraphosphate bridge by ZnCl2-mediated coupling between a nucleoside 5'-O-(thiopolyphosphate) and a second nucleotide activated by conversion into the P-imidazolide. Generally, this approach offers the possibility of synthesizing each compound three ways,22 (see Discussion). For each analog we chose the most straightforward synthetic pathway (Scheme 1). Analogs 1 and 4, which are modified at the α- and δ-positions, were obtained by the "3+1*" strategy, i.e., by coupling a nucleoside 5'-O-(1-thiotriphosphate) with a nucleoside 5'-monophosphate P-imidazolide. Analogs 2 and 3, which are modified at the β- and γ-positions, were obtained by the "2 + 2*" strategy, i.e., by coupling a nucleoside 5'-(2-thiodiphosphate) with a nucleoside 5'-diphosphate imidazolide. An alternative "3 + 1*" route was also tested for analog 2.
The guanosine 5'-O-(1-thiotriphosphate) 5 was obtained by 5'-thiophosphorylation of guanosine in trimethylphosphate in the presence of a non-nucleophilic base, 2,6-lutidine,23 followed by addition of tributylammonium pyrophosphate. As monitored by RP HPLC (see Experimental Section), the thiophosphorylation proceeded smoothly, and more than 90% conversion of nucleoside was observed within 5 h. After addition of pyrophosphate solution in DMF and weakly-alkaline hydrolysis (pH 7.5–8.0) a mixture of products was obtained consisting mainly of the desired guanosine 5'-O-(1-thiotriphosphate) (~60%), guanosine 5'-O-thiophosphate (~10 %) and guanosine 5'-O-(1-thiopentaphosphate) (~15 %). The products were separated by DEAE Sephadex A-25 ion-exchange chromatography, after which 5 was isolated as 7:10 (D1:D2) diastereomeric mixture with 46% yield. The synthesis of 6 was similar to that of 5, but the reaction was less efficient for two reasons: (i) thiophosphorylation proceeded significantly slower, the final conversion (after 24h) not exceeding 80%, and (ii) the byproduct, 7,2'-O dimethylguanosine 5'-O-(1-thiopentaphosphate), was formed more efficiently (~25%). Moreover, 31P NMR analysis unfortunately revealed that the desired product co-eluted with unreacted pyrophosphate, probably due to the partial neutralization of the negative charge on the α-phosphate by the positive charge in the m7Gua moiety. Consequently, additional HPLC purification was necessary before the synthesis scheme could be continued. Compound 6 was finally isolated as 7:10 (D1:D2) diastereomeric mixture with 30% yield. Both 5 and 6 were used in the subsequent coupling reactions as diastereomeric mixtures.
The final couplings between 5 and 7 or 6 and 8 leading to 1 or 4, respectively, were mediated by ~16-fold ZnCl2 excess. The P-imidazolide derivatives, 7 and 8, synthesized using a dithiodipyridine/triphenylphosphine activating system,24 were used in 1.5–2-fold excess to maximize consumption of the second reactant. Both coupling reactions proceeded smoothly, and essentially complete conversion of 5 or 6 into tetraphosphate-bridged dinucleotides was observed by RP HPLC after several hours (Fig. 2A). The respective HPLC conversions and isolated yields achieved after ion-exchange purification are shown in Table 1.
The synthesis of analogs modified at internal positions of the tetraphosphate bridges, i.e., 2 and 3, involved formation of mixed phosphate-thiophosphate anhydride bonds in the final step (Scheme 1 B and C). This required the nucleoside 5'-(2-thiodiphosphates) 11 and 12 or 5'-(3-thiotriphosphate) 15, which were synthesized with 80–85% yields by a procedure reported previously by our group.25 In the case of mixed anhydrides, the formation of dinucleotides proceeded slower than for unmodified ones. The reaction progress usually stopped after 3 d despite the presence of some reactants remaining. Partial hydrolysis of 11, 12 or 15 also occurred due to extended reaction times; consequently, HPLC conversions and isolated yields were lower than in case of 1 and 4 but were still acceptable (Table 1, Fig. 2B). In the case of analog 2, both synthetic pathways proved to be equally efficient.
Analogs 1–4 were isolated after ion-exchange chromatography as diastereomeric mixtures. The diastereomers of 1 and 4, in which the stereogenic P-center is next to the nucleoside moiety, were well separated by either analytical or preparative RP HPLC (see Experimental Section). The differences between diastereomers of 2 and 3 were less distinct, perhaps because the P-stereogenic center is located in the more flexible part of tetraphosphate bridge and is more distant from the nucleoside stereogenic centers. Hence, we were unable to find conditions in which these diastereomers could be separated from each other on a preparative basis, although different mobile phases and two types of columns were tested. However, in case of 2 we managed to separate small amounts of compound sufficient to determine some biochemical properties (~0.5 mg). Thus analogs 1 and 4 were subjected to biological studies as diastereomerically pure samples (1a and 4a = D1, 1b and 4b = D2), but 3 was studied as a diastereomeric mixture. Compound 2 was studied both as a diastereomeric mixture (2) and as pure diastereomers (2a = D1, 2b = D2).26
All newly synthesized compounds were characterized by MS ESI (−), 1H and 31P NMR. Each pair of diastereomers showed some small but significant differences in their 1H chemical shifts, particularly for H8 of nucleobase and H1' of the ribose moiety of either guanosine or 7,2'-O-dimethylguanosine. For 3 distinctive differences were also observed for the m27,2'-OGuo H3' and both methyl groups. In case of all compounds modified with sulfur at a phosphate neighboring a nucleoside (both nucleoside triphosphates 5 and 6 and dinucleoside tetraphosphates 1 and 4) the H8 proton of the diastereomer with shorter RP HPLC retention time (D1) was more up-field shifted than that of D2, with ΔδH chemical shift differences being in the range of 0.2–0.7 ppm (see Experimental Section and Table S1). For ATPαS and its analogs, Major et al.27 showed that the RP HPLC-faster diastereomer with less shielded H8 proton has the SP configuration; their model is that the H8 shift depends on its distance from the S atom, which is different for the two diastereomers. The same correlation between the RP HPLC retention time, H8 proton shift and the absolute configuration was subsequently reported for other structurally related compounds, including guanosine nucleotides.28 Some minor differences between diastereomers were also noted in the 31P NMR spectra of 1–6. One observation was that the signal of the phosphorothioate moiety in compounds 1, 4, 5 (GTPαS) and 6 was 0.2–0.3 ppm up-field shifted for D1 compared to D2 (Table S1). The absolute configuration of GTPαS was first reported by Conolly et al.;29 the RP HPLC-faster diastereomer was assinged as SP. Later Ludwig and Eckstein reported that the α-phosphorus signals in GTPαS (and other NTPαSs) was ~0.2 ppm more up-field for D1 (SP) than for D2 (RP).30 The abovementioned similarities in the 1H and 31P spectra of compounds 1 and GTPαS (5) suggest that their D1 diastereomers have the same, i.e. SP, absolute configuration. This is further supported by the observation that 1a and 1b are formed in the same diastereomeric ratio as the 5a and 5b ratio before reaction. On the basis of analogous observations made for compounds 4 and 6, one can presume that also in their cases the isomer D1 is of SP configuration. However, this assumption is necessary to be further confirmed, e.g. by X-ray crystallography.
The binding affinities of new analogs for murine eIF4E were determined by fluorescence quenching (Fig. 3).31 Experimental curves obtained upon titration of eIF4E with increasing concentrations of cap analogs are depicted in Fig. 3A. The descending fragment of each curve is due to the quenching of intrinsic Trp fluorescence in eIF4E upon cap binding, whereas the increasing fluorescence signal after reaching the plateau originates from fluorescence emission of unbound cap analog.31 The KAS values and free energies of binding (ΔG°) for the new analogs and reference compounds are presented in Table 2 and Fig. 3B. As expected, all new analogs have binding affinities at least 10-fold higher than the unmodified cap structure, m7GpppG. The comparison to m27,2'-OGppppG reveals that the phosphorothioate moiety generally stabilizes complexes with eIF4E. Modifications in the δ-, γ-, and β-positions (analogs 4a, 4b, 3, 2a, 2b) give additional energetic gains which are similar to those reported for the corresponding modifications in the triphosphate series.32 By contrast, the phosphorothioate substitution in the α-position, which represent the 'extra' phosphate moiety (analogs 1a and 1b), has virtually no influence on the binding affinity.
The susceptibility of new analogs to enzymatic hydrolysis by the cap-specific human DcpS pyrophosphatase was tested by monitoring enzymatic reactions progress by RP HPLC, as described in Experimental Section (Fig. 4). In all experiments m7GpppG was used as a positive control. The amount of DcpS was adjusted to assure complete cleavage of m7GpppG to m7GMP and GDP within less than 15 min. Under these conditions, analogs bearing modification at the δ-position remained undigested after 24h of incubation with DcpS, regardless of the P-center absolute configuration, and hence, were considered to be resistant to hydrolysis. Analogs 1–3 were hydrolyzed more than 80% by hDcpS to m27,2'-OGMP and the corresponding nucleoside triphosphate in 2 h, which is comparable to hydrolysis of m27,2'-OGppppG. The identification of hydrolysis product was confirmed by co-injections with authentic samples of m27,2'-OGMP, GTP, and GTPαS SP or RP. The analysis of DcpS degradation products of analogs 1a and 1b adds support to our contention that their absolute configurations around P-stereogenic centers are the same as those of GTPαS (D1) and (D2), respectively, i.e., SP and RP (Fig. 4).
The new analogs were also subjected to hydrolysis by a non-specific enzyme, Snake Venom Phosphodiesterase I (SVPDE). SVPDE is known to hydrolyze nucleic acids, NTPs, and other structurally related compounds to release nucleoside 5'-monophosphates. The enzyme has been commonly used to determine absolute configurations of phosphorothioate diester bonds and nucleoside-1-thiotriphosphates, as it more readily cleaves phosphorothioate moieties of RP than SP configuration.33,34 By means of this enzyme we hoped to additionally confirm the configurations of analogs modified at the δ-position of tetraphosphate bridge (4a and 4b). We found that SVPDE was capable of hydrolyzing m7GpppG, and surprisingly, the initially (after 15 min) released products were majorly GMP and m7GDP rather than m7GMP and GDP (~1:3 ratio). During prolonged incubation with SVPDE, the initially formed small amounts of GDP were converted into GMP. Subsequently, a dephosphorylation of GMP was observed, whereas m7GDP remained intact, which suggests it is not a substrate for SVPDE. In the case of m27,2'-OGppppG this regioselectivity of cleavage was even more notable as initially almost exclusively GMP and m27,2'-OGTP were formed. A prolonged incubation resulted, however, in simultaneous dephosphorylation of GMP and cleavage of m27,2'-OGTP to m27,2'-OGMP. Surprisingly, when analogs modified at α-position (1a and 1b) were subjected to SVPDE digestion the regioselectivity, regardless the configuration of P-stereogenic center, was opposite to that observed for m7GpppG and m27,2'-OGppppG. The complete cleavage of both 1a and 1b occurred during 1.5 h and the only products observed were m27,2'-OGMP and GTPαS either D1 or D2. The reaction mixture remained practically unchanged after further incubation with enzyme, so unfortunately, under these conditions, we were not able to observe stereoselectivity of the phosphorothioate moieties cleavage. The analogs modified at the δ-position, 4a and 4b, were cleaved to release GMP and m27,2'-OGTPαS (D1) or (D2), respectively. Similarly as for 1a and 1b no further cleavage of NTPαS occurred. Nonetheless, this experiment adds support also to the assumption of 4a and 4b absolute configurations and indentifies SVPDE as a new tool for studying modified dinucleotide cap analogs since its specificity is complementary to that of DcpS.
Finally, the new analogs were investigated for their utility as reagents for increasing the translational activity of mRNA. Transcripts encoding firefly luciferase were synthesized by SP6 RNA polymerase in the presence of a DNA template, four NTPs, and a cap dinucleotide (1a, 1b, 2, 2b, 3, 4a, 4b). The assays included three reference compounds: m7GpppG, m27,3’-OGpppG, and m27,2'-OGppppG. In vitro-synthesized mRNAs were translated in micrococcal nuclease-treated RRL system and the activity of luciferase per µg of RNA was determined by luminometry after 60 min (Fig. 5A). Cap-dependent translational efficiencies were calculated by correcting the translation of each mRNA in particular experiment by subtracting the translation of mRNA capped with ApppG (a nonfunctional cap structure), which is considered to be cap-independent. The overall translational efficiencies and cap-dependent translational efficiencies (both normalized to the translation of m7GpppG-capped mRNA) are shown in Table 2.
In our experiment we could reproduce results for cap analogs that were previously tested.10 For instance, mRNA capped with m27,3’-OGpppG and m27,2’-OGppppG were translated 1.79 ± 0.33 and 2.33 ± 0.61 more efficiently than m7GpppG-capped mRNA, respectively, which is similar to the previously reported values of 1.88 ± 0.40 and 2.56 ± 0.18. Importantly, mRNAs capped with all of the new 4P-S-ARCAs were not translated less efficiently than mRNA capped with the parent compound m27,2’-OGppppG. Their relative translation efficiencies ranged from 2.5 to 4.5 compared to m7GpppG (Table 2, Fig. 5B), which makes them attractive candidates for further studies in living cells. mRNAs capped with two analogs, 3 and 2b, were translated more than 3-fold more efficiently than m7GpppG-capped mRNA.
Nucleoside and dinucleoside polyphosphates are biologically important molecules. Numerous methods for their synthesis have been developed, several of them providing synthetic routes to dinucleoside tri- or tetraphosphates modified at non-bridging positions in the phosphate group(s).29,35–40. However, most of these methods have drawbacks with regard to synthesis of mRNA cap analogs. These include limited structural variety of possible products available by a particular method (e.g., limitation to symmetrical compounds or those modified at the α-position with respect to nucleoside.) or requirement for protected nucleosides as starting materials. Due to the presence of positive charge in 7-methylguanine, m7Guo is unusually polar and thus poorly soluble in organic solvents. It is also susceptible to hydrolysis both in acidic and basic aqueous solutions,41 which are required for removal of the majority protecting groups. These facts make synthesis of m7Guo-containing nucleotides even more challenging.
As first noted by Kadokura et al.42 and then explored by our laboratory,8,10,32,43 and others,44 the formation of pyrophosphate bonds via coupling of nucleotides with nucleotide P-imidazolide derivatives in DMF is highly accelerated by the presence of excess of anhydrous zinc chloride. The rationale for ZnCl2 catalysis is at least two-fold. First, it increases solubility of nucleotides in DMF, probably through complexing negatively charged phosphates. Due to that, it is possible to obtain completely homogenous DMF solutions of nucleotide triethylammonium salts or even sodium salts in the case of P-imidazolide derivatives. Second, ZnCl2 is thought to act as Lewis acid to activate imidazole as a leaving group.
The synthesis of 4P-S-ARCAs has also been carried out taking advantage of this phenomenon. The formation of dinucleoside tetraphosphates modified at either α- or β-position with respect to the closest nucleoside can be generally achieved by any of the pathways depicted in Scheme 2. The choice of particular pathway should depend on the availability of the appropriate substrates as well as on the purification method to be applied.
For the analogs 1 and 4 pathways A, B and C were taken into consideration. Pathway B was excluded as nucleoside 5'-O-(1-thiodiphsophates) can be obtained by procedures similar to those for 5'-O-(1-thiotriphosphates), however, with lower yields.28,45 Moreover, nucleoside diphosphates are available from corresponding activated monophosphates used in pathway A. This, would finally elongate the synthesis giving no obvious advantage over pathway A. Pathway C, would involve coupling of nucleoside 5'-monothiophosphates, which are easily available and imidazolide derivatives of nucleoside triphosphates, which are possible to obtain,10 however due to decreased solubility in DMF with lower yields than those for mono- and diphosphates. Moreover, from our experience gained during the synthesis of thio-modified dinucleoside triphosphates results that formation of mixed phosphate-thiophosphate anhydride bond is much slower than for unmodified pyrophosphate bond, what would presumably enhance by-products formation hampering products purification. Hence, the pathway A appeared to us as a favorable, since it involves most available substrates and, additionally, the formation of tetraphosphate bridge involves coupling between two unmodified phosphate moieties. This route proved to be efficient as the coupling reactions leading to analogs 1 and 4, were relatively rapid (several h) with HPLC conversions exceeding 90%
In regard to the analogs 2 and 3 pathways D, E and F should be considered. Pathway D, although shares some advantages of pathway A, is less favorable since nucleoside 5'-(2-thiotriphosphates) are possible to obtain only through complicated procedures. Pathways E and F seemed to be both acceptable and both proved to be similarly efficient giving the HPLC conversions from 50 to 70 %.
It is worth emphasizing that the described synthetic method can be also applied for other biologically important dinucleoside polyphosphates.46 It would be particularly advantageous for the synthesis of unsymmetrical compounds for instance analogs of uridine adenosine tetraphosphate,47 or for compounds bearing highly polar nucleosides.
The determination of binding affinity to eIF4E by means of fluorescence quenching titration is a relatively simple yet very useful biophysical test which enables one to predict the functionality of any newly synthesized cap analog. A sufficiently high binding affinity for eIF4E, is essential for the recognition of mRNA by translational machinery and efficient competing of mRNAs possessing modified caps with natural mRNAs.
In one of our previous works,11 we identified a triphosphate ARCA analog modified with methylene group between α and β phosphates, m27,3'-OGppCH2pG, which protected mRNA against hydrolysis by Dcp2 decapping pyrophosphatase. Due to that such mRNA had prolonged half-life when transfected into cultured cells. However, the methylene modification coincidently decreased the binding affinity to eIF4E from 10.2 ± 0.3 (for m27,3'-OGpppG) to 4.41 ± 0.2 µM−1. This caused that mRNA capped with m27,3'-OGppCH2pG, despite being more stable in vivo, was translated even less efficiently than that capped with regular ARCA.11 Another example of the influence of the binding affinity to eIF4E on mRNA translation is the tetraphosphate ARCA analog, m27,2'-OGppppG, which due to the additional phosphate moiety has KAS much higher than corresponding triphosphate and due to that was shown to enhance mRNA translation efficiency in vitro.10 It is worth mentioning that the combination of methylene modification with the presence of the additional, fourth phosphate moiety has been shown recently to compensate the loss of binding affinity.43
The high binding affinity for eIF4E of new tetraphosphate S-ARCAs constituted thus an important premise that all those analogs should be at least as efficient in promoting translation as m27,2'-OGppppG. The particular KAS values were dependent both on the position of the phosphorothioate modification and on the absolute configuration of the P-stereogenic center. Their comparison to m27,2'-OGppppG and to S-ARCA triphosphate series reveals that the phosphorothioate moiety in the δ-, γ- and β-positions of tetraphosphate correspond to γ-, β- and α-modifications in the triphosphate bridge.32,48 This implies that eIF4E 'counts' the phosphates starting from m7Guo, which is in agreement with the well-established knowledge that m7Guo moiety is crucial for the mRNA 5' end recognition by eIF4E. For instance the determined ΔG° binding energy for 4a was, respectively, 0.40 kcal mol−1 and 0.54 kcal mol−1 lower than that of 4b and m27,2'-OGppppG. In the triphosphate series the ΔΔG° differences between m27,2'-OGpSppG (D1) and either its (D2) counterpart or m27,2'-OGpppG were 0.59 kcal mol−1 and 0.69 kcal mol−1, respectively. This observation indicates that the energetic gains arising from the presence of an additional phosphate and an oxygen-to-sulfur substitution are additive. For analog 3 we were only able to determine the binding affinity for the D1/D2 mixture. In the triphosphate S-ARCA series modification at the corresponding position (β) resulted in notably different KAS values for D1 and D2 (43.1 ± 1.4 versus 19.3 ± 2.2 µM−1, respectively). From the titration performed on the 1:1 mixture of 4a and 4b, which also differ in their KAS, arises that the KAS value determined for diastereomeric mixture is roughly a mean of KAS values for pure diastereomers. Interestingly, the KAS value for analog 3 (282 ± 12) is similar to that of pentaphosphate ARCA analog, m27,3'-OGp5G. As mentioned in the Introduction, this analog, despite its high KAS value (299 ± 20), was not effective in promoting translation when incorporated into mRNA 5' end. 10 The proposed explanation was that the extraordinarily high binding affinity interferes with release of the cap at the end of translation initiation, and thereby inhibits further translation events. However, the high translation efficiency of analog 3 suggests that some other factors may contribute to this decreased efficiency of mRNA capped with m27,3'-OGp5G, e.g. low capping percentage during in vitro transcription, or non-specific interactions of highly charged pentaphosphate chain with other proteins.
The results of in vitro translation of mRNAs capped with new 4P-S-ARCAs further support the initial assumption that combining the phosphorothioate modification with elongation of the 5',5'-bridge to tetraphosphate may produce analogs with beneficial properties in terms of preparation of translationally efficient mRNA. The results suggest that the largest increase in mRNA translation efficiency may be caused by modifications in the internal positions of the tetraphosphate chain (i.e. β and γ). The somehow unexpected difference between translation efficiency of 2 (diastereomeric mixture) and 2b (diastereomer D2) underlines the need for finding a method of its separation into pure diastereomers in order to explore this phenomenon.
The more efficient mRNA translation in vitro may benefit biotechnological in vitro protein production. The determination of translation efficiency in RRL is also a useful preliminary test enabling selection of most attractive analogs for expensive studies in cultured cells. However, conventionally used rabbit reticulocyte lysates (RRLs), are micrococcal nuclease pre-treated to deplete of any endogenous mRNAs present in the lysate. Therefore, this assay does rather not confer the cellular conditions since: (i) there is no competition of investigated mRNA with natural mRNAs (ii) the influence of its stability (half-life) on overall translation yield is rather marginal. Both of these factors have large impact on the overall protein synthesis in vivo.13 The results from our previous works suggest that in cellular conditions the differences in translation yield between mRNAs capped with modified and unmodified analogs should be more pronounced.
The ability to produce more stable and more efficiently translated in vivo mRNAs can be advantageous for applications in mRNA-based gene therapy. Several approaches have been exploited, to improve the stability and/or translation efficiency of mRNA vectors in vivo. This include extending the polyA length, alterations of mRNA 3' end, introducing into mRNA modified nucleosides, which retain coding properties of natural ones.12,49,50,3 Modification of mRNA cap structure is another possibility to facilitate mRNA-based gene delivery. Particularly attractive application of mRNA in therapy is transfection of patient’s own dendritic cells with mRNAs encoding tumor-associated antigens to evoke auto-immunization against cancer cells51.
We believe that the 4P-S-ARCAs will also enable some new insights into mRNA related physiological processes. The synthetic cap analogs have been commonly used to investigate mechanism of initiation of protein biosynthesis, and the discovery of the involvement of decapping enzymes in the mRNA degradation pathways, expanded further the scope of processes that have been studied by means of cap analogs. One of the still unsolved problems is the detailed mechanism of Dcp2 decapping enzyme, which belongs to the NuDiX family of pyrophosphatases.16 It is known that the enzyme utilizes cap structures on intact mRNAs and cleaves between α and β phosphates of the triphosphate bridge. The finding that the phosphorothioate group in the β position protected mRNA from decapping suggested that the β-phosphate is the one directly involved in the catalytic mechanism.13 However, since Dcp2 requires both cap structure and mRNA chain for its activity it is unclear whether the enzyme recognizes the phosphate groups 'counting' them from the m7Guo site or from the mRNA body. The susceptibility to Dcp2 of tetraphosphate S-ARCA modified in the β and γ-positions, which is currently under investigation, may address this issue.
On the other hand, the analogs modified in the δ-position proved to be resistant to DcpS decapping enzyme, which cleaves cap structures lacking mRNA chain. DcpS-resistant cap analogs with high affinity for eIF4E are important in the context of developing stable inhibitors of cap dependent translation which could counteract elevated levels of eIF4E in cancer cells.52 It is assumed that DcpS functions in the cytoplasm to prevent the accumulation of complexes between eIF4E and cap structures that would otherwise accumulate following 3'→5' mRNA decay,53 what implies that only DcpS resistant analogs could act as potent eIF4E inhibitors in cellular conditions. Moreover, it was also found that DcpS fulfils also some other biological functions, and constitutes an interesting object for investigation by itself. Its presence has been detected also in the nucleus, where it is implicated in the mRNA splicing, and recently, it has been identified as therapeutic target for spinal muscular atrophy.54,55 Non-hydrolysable cap analogs such as m27,2'-OGpSpppG may serve as invaluable tools to investigate these processes for instance by serving as inhibitors of DcpS activity.
Finally, it has been recently discovered that by modifying 5',5'-phosphate bridge in the mRNA cap structure one can observe enhancement in translation repression mediated by microRNAs.20 In the quoted work, two cap analogs have been identified, which in Drosophila melanogaster embryos extract evoke ~2-fold stronger miRNA mediated inhibition of translation initiation but are "neutral" towards general cap-dependent translation: the triphosphate ARCA modified at α-position, m27,2'-OGpppSG (D2), and the analog with 5',5'-bridge extended to hexaphosphate, m7Gp6G. Thus the 4P-S-ARCA modified at the α-position, which somehow combines the structural features of these two analogs may be an interesting tool for studying mechanism of microRNAs action.
A general methodology for the synthesis of unsymmetrical dinucleotides bearing a single phosphorothioate modification in either position of tetraphosphate chain has been developed. Four pairs of tetraphosphate S-ARCAs have been synthesized with good yields. Biophysical and biochemical characterization of new analogs indicates 4P-S-ARCAs as good candidates for reagents to produce mRNA transcripts with high translational activity. Stabilizing effects of the additional phosphate as well as the phosphorothioate substitutions on the cap-eIF4E complex are additive, which is reflected in KAS values much higher than for triphosphate cap analogs and notably higher than for unmodified tetraphosphate ARCA. This property of 4P-S-ARCAs should reflect in kinetics of translation initiation complex formation and make synthetic transcripts more competitive than endogenous mRNA in vivo.
It is also shown that phosphorothioate modification in the δ-position of tetraphosphate bridge protects dinucleotide cap analog against hydrolysis by DcpS. Similarly, the modification in the appropriate position of tetraphosphate bridge should decrease the susceptibility of 4P-S-ARCA containing mRNA transcripts to hydrolysis by Dcp2/Dcp1, which would affect their half-life and give additional contribution to translational activity in vivo.
The results of studies on translation in vitro show that mRNAs capped with new analogs are 2.5–4.5 more efficiently translated in a cell free system than m7GpppG capped mRNAs. On the basis of our previous studies one can predict the effect of novel cap on translational efficiency would be more pronounced when measured in cultured cells. These observations indicate 4P-S-ARCAs s promising candidates for studies on therapeutic applications of mRNA, which include mRNA-based gene delivery and antitumor vaccination. In addition to the biotechnological and potential therapeutic applications 4P-S-ARCAs appear as useful tools for elucidating mechanisms of various mRNA-related physiological processes.
Solvents and other reagents were purchased from Sigma-Aldrich and used without further treatment, unless otherwise stated. Reaction mixtures were analyzed by analytical reverse phase (RP) HPLC, which was performed on a Agilent Technologies Series 1200 apparatus using a Supelcosil LC-18-T RP column (4.6 × 250 mm, 5µm, flow rate 1.3 mL/min) developed with a 0–50% linear gradient of methanol in 0.05 M ammonium acetate buffer (pH 5.9) within 30 min, UV-detection at 260 nm and fluorescence detection (excitation at 280 nm and detection at 337 nm). Samples for HPLC analysis were prepared by dissolving 5 µL of a reaction mixture in 100 µL of either water or, in case of ZnCl2 containing reactions, aqueous solution of EDTA (10 mg/mL) and NaHCO3 (5 mg/mL).
Nucleotides were purified and isolated from reaction mixtures by ion-exchange chromatography on DEAE-Sephadex A-25 (HCO3− form) column using a linear gradient of triethylammonium bicarbonate (TEAB) in deionized water (1.6 L of each) and, after evaporation under reduced pressure with repeated additions of ethanol and drying in a vacuum dessicator over P2O5, isolated as triethylammonium (TEA) salts.
The final products were additionally purified (and separated) by semi-preparative RP HPLC, which was performed on either Waters 600E Multisolvent Delivery System apparatus using Discovery RP Amide C-16 HPLC column (25 cm × 21.2 mm, 5 µm, flow rate 5.0 mL/min) (or Ascentis C18 HPLC column (25 cm × 21.2 mm, 5 µm, flow rate 5.0 mL/min) developed with linear gradients of methanol in 0.05 M ammonium acetate (AAC) buffer (pH 5.9). UV-detection at 260 nm. After repeated freeze-drying, the products were isolated as ammonium salts. For compound 6 triethylammonium acetate (TEAAC) buffer (pH 7.0) was used in order to isolate compounds as TEA salts.
Yields were preferably calculated based on optical density miliunits at 260 nm (mODU260) of substrates and isolated products, measured in 0.1 M phosphate buffer of pH either 6 (for m7Guo mononucleotides) or pH=7 (for cap dinucleotides and Guo nucleotides). Optical density miliunits mODU260 are defined as absorption of compound solution in 0.1 M phosphate buffer of appropriate pH at 260 nm multiplied by volume of the solution in ml. Extinction coefficients taken for calculations were εpH=6= 11,400 M−1 cm−1 for m7Guo mononucleotides, εpH=7= 22,600 M−1 cm−1 for cap dinucleotides and εpH=7= 12,000 M−1 cm−1 for Guo mononucleotides.
The structures and homogeneities of synthesized nucleotides were confirmed by chromatography on analytical RP HPLC, mass spectrometry using negative electrospray ionization (MS ESI−) and both 1H and 31P NMR spectroscopy. 1H NMR and 31P NMR spectra were recorded in D2O at 25°C for samples at 1.5–3.0 mg mL−1 concentrations on a Varian UNITY-plus spectrometer at 399.94 MHz and 161.90 MHz, respectively. 1H NMR chemical shifts in ppm were reported to sodium 3-trimethylsilyl-[2,2,3,3-D4]-propionate (TSP) in D2O as an internal standard. 31P NMR chemical shifts in ppm were reported to 80% phosphorus acid in D2O as an external standard. J values are given in Hz. Mass spectra were recorded on a Micromass QToF 1 MS spectrometer.
2'-O-methylguanosine was synthesized as described by Kusmierek and Shugar.56 7,2'-O-dimethyl guanosine was obtained from 2'-O-methylguanosine by methylation with CH3I analogously as described earlier for 7-methylguanosine.57 Tributylammonium pyrophosphate was prepared according to Ludwig and Eckstein.30 Thiophosphate TEA salt was obtained from its trisodium salt as described earlier.25 Guanosine 5'-monophosphate disodium salt was purchased from Fluka and before use converted into its triethylammonium (TEA) salt by passing through Dowex 50 W × 8 (100–200 mesh), evaporating collected fractions to dryness and drying in vacuum over P2O5. Nucleotides: guanosine 5'-diphosphate (9), 7,2'-O-dimethylguanosine 5'-monophosphate and 7,2'-O-dimethylguanosine 5'-diphosphate (10) (all as TEA salts) as well as imidazolide derivatives 7, 8, 13 and 14 (all as sodium salts) were synthesized as described previously.10,43
To a suspension of guanosine (7710 mODU260 , 250 mg, 0.64 mmol; dried overnight in vacuum over P4O10) in trimethyl phosphate (3.75 ml) cooled to 0°C on ice/water bath were added 2,6-dimethylpyridine (460 µl, 4 mmol) and PSCl3 (270 µl, 2.65 mmol). The reaction progress was under HPLC control. After 6 h tributylammonium pyrophosphate solution (15.9 ml of 0.5M solution) in DMF and tributylamine (0.61 ml) were added. The reaction was maintained at RT for 15 min and then quenched by addition of 145 ml of water and a portion of 0.7 TEAB to pH 8 and extracted twice with ether. The product was separated by DEAE Sephadex chromatography using a linear gradient of 0–1.2 M TEAB to yield 5 (3550 mODU260, 310 mg, 0.30 mmol, 46%) as a 72:100 SP:RP (D1:D2, 5a:5b) diastereomers mixture. ESI MS (−) m/z 537.98 (Calc. for C10H15N5O13P3S: 537.96). 5a: Analytical RP HPLC Rt = 3.5 min δH 8.29 (1H, s, H(8) G); 5.95 (1H, d, J1',2' 6.3, H(1') G); 4.85 (1H, ~t, J1',2' 6.3 J2',3' 4.8, H(2')); 4.60 (1H, dd, J2',3' 4.8, J3',4' 3.2, H(3')); 4.39 (1H, m, H(4')); 4.30 (2H, m, H(5') H(5")). δP 43.12 (1P, d, JP1,P2 27.0, P1), −10.81 (1P, d, JP2,P3 19.6, P3), −23.98 (1P, dd, JP1,P2 27.0, JP2,P3 19.6, P2) 5b: Analytical RP HPLC Rt = 4.1 min δH 8.22 (1H, s, H(8)); 5.95 (1H, d, J1',2' 6.3, H(1')); 4.85 (1H, ~t, J1',2' 6.3 J2',3' 4.8, H(2')); 4.60 (1H, dd, J2',3' 4.8, J3',4' 3.2, H(3')); 4.39 (1H, m, H(4')); 4.30 (2H, m, H(5') H(5")), δP 43.64 (1P, d, JP1,P2 27.0, P1), −10.81 (1P, d, JP2,P3 19.6; P3), −24.02 (1P, dd, JP1,P2 27.0, JP2,P3 19.6; P2)
A suspension of 7,2’-O-dimethylguanosine (4500 mODU260, 150 mg, 0.39 mmol; dried overnight in vacuum over P4O10) in trimethyl phosphate (2.25 ml) was cooled to 0°C on ice/water bath were added 2,6-dimethylpyridine (340 µl, 2.9 mmol) and PSCl3 (150 µl, 1.4 mmol) were added. The reaction progress was under HPLC control. After 24 h to the reaction mixture were added tributyloammonium pyrophosphate solution (14 ml of 0.5M solution in DMF) and tributylamine (0.67 ml). The reaction was maintained at RT for 15 min and then quenched by addition of 120 ml of water and a portion of 0.7 TEAB to pH 8 and extracted twice with ether. The product was separated by DEAE Sephadex chromatography using a linear gradient of 0–1.1 M TEAB to yield 6 (1580 mODU260, 270 mg, 0.14 mmol, 25%) as a 7:10 SP:RP (D1:D2, 6a:6b) diastereomers mixture. The 31P NMR analysis revealed that the isolated product was contaminated with ~1.5 eq. of pyrophosphate. Consequently, the product was further purified by semi-preparative RP HPLC using a linear gradient of 0–50% MeOH in TEAAC (pH 7.0) within 45 min and the diastereomers were collected together yielding, after repeated freeze-drying, 6 (1350 mODU260, 120mg, 0.12 mmol, 30%) as a 7:10 diastereomeric mixture. Additionally, a small portion of 6 was purified using a linear gradient of 0–25% MeOH in AAC (pH 5.9) within 120 min, the diastereomers were collected separately (6a Rt= 41 min (SP, D1), 6b Rt= 45 min (RP, D2)) and after repeated freeze drying isolated as NH4+ salts. ESI MS (−) m/z 566.02 (calc. for C12H19N5O13P3S: 565.99). 6a: Analytical RP HPLC Rt = 8.5 min δH 9.33 (1H, s, H(8)); 6.15 (1H, d, J1',2' 2.6, H(1')); 4.70 (1H, ~t, J2',3' = J3',4' ~5.3, H(3')); 4.42-4.33 (4H, overlapped m, H(2'), H(4'), H(5'), H(5")); 4.17 (3H, s, NCH3); δP 43.54 (1P, d, JP1,P2 28.0, P1), −10.32 (1P, d, JP2,P3 20.0; P3), −23.73 (1P, dd, JP1,P2 28.0, JP2,P3 20.0, P2) 6b: Analytical RP HPLC Rt = 9.0 min, δH 9.30 (1H, s, H(8)); 6.14 (1H, d, J1',2' 2.1, H(1')); 4.66 (1H, ~t, J2',3' = J3',4' ~5.3, H(3')); 4.46-4.27 (4H, overlapped m, H(2'), H(4'), H(5'), H(5")); 4.16 (3H, s, NCH3); δP 43.33 (1P, d, JP1,P2 27.6, P1), −10.55 (1P, d, JP2,P3 19.8, P3), −23.73 (1P, dd, JP1,P2 27.6, JP2,P3 19.8, P2)
An appropriate nucleotide imidazolide derivative (1 eq.) was mixed with thiphosphate TEA salt (3–4 eq.) in DMF followed by immediate addition anhydrous ZnCl2 (at least 8 eq.) and vigorous shaking until homogenous solution was obtained. The reaction was quenched after 15–25 min by being diluted with ~20 volumes of aqueous solution of disodium EDTA dihydrate (in 1:1 ratio to ZnCl2) and NaHCO3 (a half of the amount of EDTA in mg). If necessary the pH was adjusted to 7–7.5 with additional portion of NaHCO3. The product was isolated by ion exchange chromatography. Product containing fractions were poured together and evaporated to dryness by repeated additions of 96% ethanol and, finally, absolute ethanol until white solid resulted.
Obtained starting from 7 (2110 mODU260, 100 mg, 0.18 mmol), thiophosphate TEA salt (obtained from 200 mg of trisodium salt hydrate, ~0.5 mmol) and ZnCl2 (235 mg, 1.73 mmol) in 3 ml of DMF. IE purification was accomplished with 0–0.9 M linear gradient of TEAB to yield 1790 mODU260 (130 mg, 0.16 mmol, 84 %) of 11 as a TEA salt. ESI MS (−) m/z 486.03 (Calc. for C12H18N5O10P2S: 486.02) δP 35.80 (1P, d, JP1,P2 30.7, P2), −11.66 (1P, d, JP1,P2 30.7, P1)
Obtained starting from 8 (3570 mODU260, 150 mg, 0.30 mmol), thiophosphate TEA salt (prepared from 400 mg of trisodium salt hydrate, ~0.75 mmol) and ZnCl2 (350 mg, 2.6 mmol) in 3 ml of DMF. IE purification was accomplished with 0–1.0 M linear gradient of TEAB to yield 2820 mODU260 (180 mg, 0.23 mmol, 79 %) of 12 as a TEA salt. ESI MS (−) m/z 457.89 (Calc. for C10H14N5O10P2S: 457.99) δP 34.69 (1P, d, JP1,P2 31.4, P2), −11.57 (1P, d, JP1,P2 31.4 Hz, P1)
Obtained starting from 14 (1600 mODU260, 100 mg, 0.14 mmol), thiophosphate TEA salt (obtained from 200 mg of trisodium salt hydrate, ~0.5 mmol) and ZnCl2 (240 mg, 1.76 mmol) in 3.5 ml of DMF. IE purification was accomplished with 0–1.1 M linear gradient of TEAB to yield 1290 mODU260 (110 mg, 0.11 mmol, 80 %) of 15 as a TEA salt. ESI MS (−) m/z 565.98 (Calc. for C12H19N5O13P3S: 565.99) δP 39.18 (1P, d, JP2,P3 28.7, P3), −11.24 (1P, d, JP1,P2 20.4, P1), −23.83 (1P, dd, JP2,P3 28.7, JP1,P2 20.4, P2)
To a suspension of 7 (1800 ODU260, 93 mg, 0.16 mmol) and 5 (1590 ODU260, 0.13 mmol) in DMF (5 ml) anhydrous ZnCl2 (244 mg, 2.54 mmol) was added and the mixture was shaken until reagents dissolved (1–2 min). The reaction was maintained at RT for 6 h and then quenched by being diluted with a solution of 940 mg (2.54 mmol) of disodium EDTA in 100 ml of water and adjusted to pH 6 with solid NaHCO3. The product was isolated on DEAE Sephadex using 0–1.3 M gradient of TEAB to yield 1800 ODU260 (0.080 mmol, 60 %) of 1 (TEA salt) as a 7:10 SP:RP diastereomers mixture. The diastereomers were separated using 0–25% gradient of MeOH in AAC (pH=5.9) within 120 min (1a: Rt= 43 min (SP, D1), 1b: Rt= 49 min (RP, D2)) to yield 484 ODU260 (19.4 mg, 0.02 mmol, 16%) of D1 (1a) and 761 ODU260 (31 mg, 0.033 mmol, 25%) of D2 (1b) as NH4+ salts. ESI MS (−) m/z 911.05 (Calc. for C22H31N10O20P4S1: 911.04). 1a: Analytical RP HPLC Rt = 6.8 min; δH 9.19 (1H, s, H(8)m7G); 8.06 (1H, s, H(8)G); 6.03 (1H, d, J1',2' 2.4, H(1')m7G); 5.83 (1H, d, J1',2' 6.0, H(1')G); 4.72 (1H, ~t, H(2')G); 4.61 (1H, ~t, H(3')m7G); 4.52 (1H, ~t, H(3')G); 4.45-4.42 (7H, overlapped m, H(4')m7G, H(4')G, H(2') m7G, H(5')G, H(5")G, H(5'), H(5")m7G); 4.10 (3H, s, N-CH3); 3.58 (3H, s, O-CH3), δP 43.70 (1P, d, JP3,P4 26.0; P4), −11.11 (1P, d, JP1,P2 18.0; P1), −23.80 (1P, ~t, JP1,P2 18.0, JP2,P3 17.2, P2), −24.90 (1P, dd, JP3,P4 26.0, JP2,P3 17.2; P3). 1b: Analytical RP HPLC Rt = 7.2 min; δH 9.16 (1H, s, H(8)m7G); 8.05 (1H, s, H(8)G); 5.99 (1H, d, J1',2' 2.4, H(1')m7G); 5.80 (1H, d, J1',2' 6.0, H(1')G); 4.75 (1H, ~t overlaped with HDO, H(2')G); 4.60-4.45 (2H, overlapped m , H(3')m7G; H(3')G); 4.40-4.30 (3H, overlapped m, H(4')G, H(5')G, H(5")G); 4.30–4.22 (3H, overlapped m, H(4') m7G, H(5'), H(5")m7G); 4.10 (3H, s, N-CH3);3.59 (3H, s, O-CH3),. δP 43.47 (1P, d, JP3,P4 26.3, P4), −11.27 (1P, d, JP1,P2 18.0; P1), −23.00 (1P, ~t, JP1,P2 18.0, JP2,P3 17.0; P2), −24.07 (1P, dd, JP3,P4 26.3, JP2,P3 17.0, P3)
To a suspension of 14 (Na salt, 102 mg; 1165 mODU260, 0.10 mmol) and 12 (TEA salt, 1730 mODU260, 0.14 mmol) in DMF (3 ml) anhydrous ZnCl2 (250 mg, 1.84 mmol) was added and the mixture was shaken until reagents dissolved (1–2 min). The reaction was maintained at RT for 3 d and then quenched by addition of 685 mg (1.84 mmol) of EDTA in 100 ml of water and adjusted to pH 6 with solid NaHCO3. The product was isolated on DEAE Sephadex using 0–1.3 M gradient of TEAB to yield 795 mODU260 (0.035 mmol, 35 %) of 2 as TEA salt. The product was additionally purified by semi-preparative RP HPLC using isocratic 10% MeOH and 90 % AAC (pH 5.9) yielding 640 mODU260 (31 mg, 0.031 mmol, 28 %) of 2 (NH4+ salt) as 8:10 (D1:D2, 2a:2b) diastereomers mixture.
To a suspension of 8 (Na salt, 107 mg; 2570 mODU260, 0.21 mmol) and 15 (TEA salt, 1290 mODU260, 110mg, 0.11 mmol) in DMF (2 ml) anhydrous ZnCl2 (775 mg, 5.69 mmol) was added and the mixture was shaken until reagents dissolved (1–2 min). The reaction was maintained at RT for 3 days and then quenched by addition of 2.11 g of EDTA (5.69 mmol) in 300 ml of water and neutralized with solid NaHCO3. The purification as in Method I yielded 1010 mODU260 (0.045 mmol, 39 %) of 2 as TEA salt as 9:10 (2a:2b, D1:D2) diastereomers mixture, and after additional HPLC purification 820 ODU260 (37 mg, 0.036 mmol, 31 %) of 2. ESI MS (−) m/z 911.05 (calc. for C22H31N10O20P4S1: 911.04). 2a: Analytical RP HPLC Rt = 7.5 min, δH 9.04 (1H, s, H(8)m7G); 8.10 (1H, s, H(8)G); 5.97 (1H, d, J1',2' 2.9, H(1')m7G); 5.80 (1H, d, J1',2' 5.9, H(1')G); 4.70 (1H, ~t, H(2')G); 4.56 (1H, ~t, H(3')m7G); 4.50 (1H, ~t, H(3')G); 4.41 (1H, m, H(5')G); 4.34 (2H, m, overlapped H(4')m7G, H(4')G); 4.27 (2H, m overlapped, H(2')m7G, H(5")G); 4.24 (2H, m, H(5'), H(5")m7G); 4.08 (3H, s, N-CH3); 3.59 (3H, s, O-CH3). δP 30.49 (1P, t, JP2,P3 = JP3,P4 24.7, P3), −11.14 (1P, d, JP1,P2 17.6, P1), −11.88 (1P, d, JP3,P4 24.7, P4), −23.84 (1P, dd, JP2,P3 24.7, JP1,P2 17.6, P2) 2b: Analytical RP HPLC Rt = 7.6 min, δH 9.00 (1H, s, H(8)m7G); 8.08 (1H, s, H(8)G); 5.96 (1H, d, J1',2' 2.9, H(1') m7G); 5.81 (1H, d, J1',2' 5.9, H(1') G); 4.70 (1H, ~t, H(2')G); 4.55(1H, ~t, H(3')m7G); 4.48 (1H, ~t, H(3')G); 4.40 (2H, overlapped m, H(4')m7G, H(5')G); 4.30 (3H, m (overlapped), H(2') m7G, H(4')G, H(5")G); 4.25 (2H, m, H(5'),H(5")m7G); 4.07 (3H, s, N-CH3), 3.62 (3H, s, O-CH3). δP 30.35 (1P, t, JP2,P3 = JP3,P4 24.7, P3), −11.14 (1P, d, JP1,P2 17.6, P1), −11.88 (1P, d, JP3,P4 24.7, P4), −23.86 (1P, dd, JP2,P3 24.7, JP1,P2 17.6, P2)
To a suspension of 13 (80 mg, 1450 mODU260, 0.12 mmol) and 12 (895 mODU260, 0.078 mmol) in DMF (5 ml) anhydrous ZnCl2 (172 mg, 1.27 mmol) was added and the mixture was shaken until reagents dissolved (1–2 min). The reaction was maintained at RT for 3 days and then quenched by addition of 480 mg (1.27 mmol) of disodium EDTA in 45 ml of water and neutralized with solid NaHCO3. The product was isolated on DEAE Sephadex using 0–1.2 M gradient of TEAB to yield 560 mODU260 (0.025 mmol, 32%) of 3. The product was additionally purified by semi-preparative RP HPLC using isocratic 10% MeOH and 90 % AAC (pH 5.9) yielding 435 ODU260 (20 mg, 0.019 mmol, 24 %) of 3 (NH4+ salt) as 7:10 (3a:3b, D1:D2) diastereomers mixture. ESI MS (−) m/z 911.04 (calc. for C22H31N10O20P4S1: 911.04) 3a: Analytical RP HPLC Rt = 7.5 min, δH 9.04 (1H, s, H8 m7G); 8.10 (1H, s, H(8)G); 5.97 (1H, d, J 2.9, H(1')m7G); 5.80 (1H, d, J 5.9, H(1')G); 4.70 (1H, ~t, H(2')G); 4.56 (1H, ~t, H(3')m7G); 4.50 (1H, ~t, H(3') G); 4.41 (1H, m, H(5')G); 4.34 (2H, m, overlapped H(4')m7G, H(4')G); 4.27 (2H, m overlapped, H(2')m7G, H(5")G); 4.24 (2H, m, H(5'), H(5")m7G); 4.08 (3H, s, N-CH3); 3.59 (3H, s, O-CH3), δP 30.21 (1P, t, JP1,P2 = JP2,P3 24.0; P2), −11.12 (1P, d, JP3,P4 18.5, P4), −11.96 (1P, d, JP1,P2 24.0, P1), −23.91 (1P, dd, JP2,P3 24.0, JP3,P4 18.5, P3) 3b: Analytical RP HPLC Rt = 7.6 min, δH 9.00 (1H, s, H(8)m7G), 8.08 (1H, s, H(8)G), 5.96 (1H, d, J 2.9, H(1')m7G), 5.81 (1H, d, J 5.9, H(1')G), 4.70 (1H, ~t, H(2')G); 4.55 (1H, ~t, H(3') m7G), 4.48 (1H, ~t, H(3')G), 4.40 (2H, overlapped m, H(4')m7G, H(5')G), 4.30 (3H, overlapped m, H(2')m7G, H(4')G, H(5")G), 4.25 (2H, m, H(5'), H(5")m7G), 4.07 (3H, s, N-CH3), 3.62 (3H, s, O-CH,), δP 30.21 (1P, t, JP1,P2 = JP2,P3 24.0, P2), −11.12 (1P, d, JP3,P4 18.5, P4), −11.96 (1P, d, JP1,P2 24.0, P1), −23.91 (1P, dd, JP2,P3 24.0, JP3,P4 18.5, P3)
To a suspension of 7 (Na salt, 65 mg; 1430 mODU260, 0.119 mmol) and 6 (TEA salt, 495 mODU260, 0.043 mmol) in DMF (6 ml) anhydrous ZnCl2 (246 mg, 1.81 mmol) was added and the mixture was shaken until reagents dissolved (1–2 min). The reaction was maintained at RT for 6.5 h and then quenched by addition of 673 mg (1.81 mmol) of disodium EDTA in 100 ml of water and neutralized with solid NaHCO3. The product was isolated on DEAE Sephadex using 0–1.3 M gradient of TEAB to yield 635 mODU260 (0.028 mmol, 65%) of 4 (TEA salt) as 7:10 (SP:RP, D1:D2, 4a:4b) diastereomers mixture. The diastereomers were separated by RP HPLC using 0–25% gradient of MeOH in AAC (pH=5.9) within 120 min (4a: Rt = 51 min (SP, D1), 4b: Rt = 52 min (RP, D2)) to yield 196 ODU260 (8.5 mg, 0.0087 mmol, 20%) of 4a and 308 ODU260 (13.4 mg, 0.013 mmol, 31%) of 4b (both as NH4+ salts). ESI MS (−) m/z 911.06 (calc. for C22H31N10O20P4S1: 911.04) 4a: Analytical RP HPLC Rt = 7.5 min, δH 9.04 (1H, s, H(8)m7G), 8.10 (1H, s, H(8)G), 5.97 (1H, d, J 2.9, H(1') m7G), 5.80 (1H, d, J 5.9, H(1') G), 4.70 (1H, ~t, H(2') G), 4.56 (1H, ~t, H(3') m7G), 4.50 (1H, ~t, H(3')G), 4.41 (1H, m, H(5')G), 4.34 (2H, overlapped m, H(4')m7G, H(4')G); 4.27 (2H, overlapped m, H(2')m7G, H(5")G), 4.24 (2H, m, H(5'), H(5")m7G), 4.08 (3H, s, N-CH3), 3.59 (3H, s, O-CH3). δP 43.92 (1P, d, JP1,P2 23.0, P1), −11.00 (1P, d, JP3,P4 16.6, P4), −22.71 (1P, ~t, JP3,P4 16.6, JP2,P3 16.5, P3), −23.63 (1P, dd, JP3,P4 23.0, JP2,P3 16.5, P2). 4b: Analytical RP HPLC Rt = 7.8 min : δH 9.00 (1H, s, H(8) m7G); 8.08 (1H, s, H(8)G); 5.96 (1H, d, J 2.9, H(1')m7G); 5.81 (1H, d, J 5.9, H(1')G); 4.70 (1H, ~t, H(2')G); 4.55 (1H, ~t, H(3')m7G); 4.48 (1H, ~t, H(3')G); 4.40 (2H, overlapped m, H(4')m7G, H(5')G); 4.30 (3H, overlapped m, H(2')m7G, H(4')G, H(5")G); 4.25 (2H, m, H(5'), H(5")m7G); 4.07 (3H, s, N-CH3), 3.62 (3H, s, O-CH3). δP 43.52 (1P, d, JP1,P2 22.5, P4), −11.06 (1P, d, JP3,P4 15.6, P4), −22.89 (1P, ~t, JP3,P4 15.6, JP2,P3 16.5, P3), −23.74 (1P, dd, JP1,P2 22.5, JP2,P3 16.5, P2)
Human DcpS was expressed in Escherichia coli according to the procedures described previously.58 The protein of 15 µM concentartion was stored at −80 °C in 20 mM Tris buffer, pH 7.5, containing 50 mM KCl, 0.2 mM EDTA, 1 mM DTT, 0.5 mM PMSF, and 20% glycerol. Snake Venom Phosphodiesterase I from Crotalus adamanteus (EC. 3. 1. 4. 1) was purchased from Sigma (0.14 u/mg solid) and before use dissolved at 1mg/mL concentration in 100 mM Tris buffer, pH 8.0, containing 100 mM NaCl, and 14 mM MgCl2 and 50% glycerol and stored in this form at −20 °C.
hDcpS assay: A cap analog at 40 µM concentration was treated with 3.0 µL of DcpS in 500 µl of 50 mM TRIS buffer, pH=7.9, containing 20 mM MgCl2 and 60 mM (NH4)2SO4 at 30 °C. After 15, 30, 75, 120 min and 24 h a 100 µl sample was collected from the reaction mixture and deactivated by incubation in 90 °C for 3 min.
SVPDE assay: A cap analog at 40 µM concentration was treated with 5.0 µL of SVPDE in 500 µl of 100 mM TRIS buffer, pH=8.0, containing 14 mM MgCl2 and 100 mM NaCl at 30 °C. After 15, 30, 90, 210, 330 min and 24 h a 50 µl sample was collected from the reaction mixture and deactivated by incubation in 90 °C for 3 min.
All samples were analyzed without further treatment by analytical RP HPLC using a 0–25% linear gradient of methanol in 0.1 M KH2PO4/K2HPO4 buffer (pH = 6.0) within 30 min.
Murine eukaryotic Initiation Factor eIF4E (residues 28–217) was expressed in BL21(DE3) Escherichia coli strain. The protein was purified from inclusion bodies then guanidinum-solubilized protein was refolded by a one-step dialysis and purified by ion-exchange chromatography on a HiTrapSP column without contact with cap analogs. The concentration of eIF4E was determined spectrophotometrically (ε280 = 53,400 cm−1M−1). Fluorescence titration measurements were carried out on an LS-55 spectrofluorometer (Perkin Elmer Co.) in 50 mM HEPES/ KOH (pH 7.2), 100 mM KCl, 0.5 mM EDTA, 1 mM DTT at 20.0 ± 0.2°C. 1 µL aliquots of cap analog solutions with increasing concentration were added to 1.4 mL of 0.1 µM protein solutions. Fluorescence intensities (excitation at 280 nm with 2.5 nm bandwidth and detection at 337 nm with 4-nm bandwidth and 290 nm cut off filter) were corrected taking sample dilution and the inner filter effect into account. Equilibrium association constants (KAS) were determined by fitting the theoretical dependence of fluorescence intensity on the total concentration of cap analog to the experimental data points, according to equation described previously.31 The concentration of protein was fitted as a free parameter of equilibrium equation showing amount of “active” protein. The final KAS was calculated as a weighted average of 3–4 independent titrations, with the weights taken as the reciprocals of the numerical standard deviations squared. Numerical nonlinear least-squares regression analysis was performed using ORGIN 6.0 (Microcal Software Inc.). The Gibbs free energy of binding was calculated from the KAS value according to the standard equation ΔG° = −RT ln KAS.
Capped, polyadenylated luciferase mRNAs were synthesized in vitro on a dsDNA template (amplified by PCR reaction) that contains: SP6 promoter sequence of DNA-dependent RNA polymerase, 5'UTR sequence of rabbit β-globin, the entire firefly luciferase ORF and a string of 31 adenosines. A typical in vitro transcription reaction mixture (40 µL) contained: SP6 transcription buffer (Fermentas), 0.7 µg of DNA template, 1U/µL RiboLock Ribonuclease Inhibitor (Fermentas), 0.5 mM ATP/CTP/UTP and 0.1 mM GTP and 0.5 mM dinucleotide cap analog (molar ratio cap analog:GTP 5:1). The reaction mixture was preincubated at 37°C for 5 minutes before addition of SP6 RNA polymerase (Fermentas) to final concentration 1U/µL and reaction was continued for 45 minutes at 37°C. After incubation, reaction mixtures were treated with DNase RQ1 (Promega) in transcription buffer, for 20 min at 37°C at concentration 1U per 1µg of template DNA. RNA transcripts were purified using NucAway Spin Columns (Ambion), integrity of transcripts was checked on a non-denaturating 1% agarose gel and concentrations were determined spectrophotometrically.
A translation reaction in RRL was performed in 10 µL volume for 60 minutes at 30°C, in conditions determined as optimal for cap-dependent translation. A typical reaction mixture contained: 40% RRL lysate (Flexi Rabbit Reticulocyte Lysate, Promega), mixture of amino acids (0.01 mM), MgCl2 (0.6 mM), potassium acetate (170 mM) and 5'-capped mRNA. Four different concentrations of each analyzed transcript were tested in an in vitro translation reaction. Activity of synthesized luciferase was measured with a luminometer.
We are indebted to Mike Kiledjian (Rutgers University) for providing the hDcpS encoding plasmid and Zbigniew M. Darzynkiewicz for expressing and purifying the protein, to the Laboratory of Biological NMR (Institute of Biochemistry and Biophysics of the Polish Academy of Sciences, IBB PAS) for access to the NMR apparatus and to Laboratory of Mass Spectrometry (IBB PAS) for recording MS spectra. Financial support from the Polish Ministry of Science and Higher Education (No. NN301 243 436), National Science Support Project 2008–2010 (No. PBZMNiSW-07/I/2007), Howard Hughes Medical Institute (to E.D.; No. 55005 604) and National Institute for General Medical Sciences (to R.E.R,; No. GM20818) is gratefully acknowledged.
§Electronic Supplementary Information (ESI) available: Table S1, HPLC profiles, 1H and/or 31P NMR spectra of compounds 1–6.