Phosphorothiolate Dinucleotide Cap Analogues - ACS Publications

Article Cite This: J. Am. Chem. Soc. 2018, 140, 5987−5999

pubs.acs.org/JACS

5′-Phosphorothiolate Dinucleotide Cap Analogues: Reagents for Messenger RNA Modification and Potent Small-Molecular Inhibitors of Decapping Enzymes Blazej A. Wojtczak,†,⊥ Pawel J. Sikorski,†,⊥ Kaja Fac-Dabrowska,† Anna Nowicka,‡ Marcin Warminski,‡ Dorota Kubacka,‡ Elzbieta Nowak,§ Marcin Nowotny,§ Joanna Kowalska,*,‡ and Jacek Jemielity*,† †

Centre of New Technologies, University of Warsaw, Banacha 2c Street, 02-097 Warsaw, Poland Division of Biophysics, Institute of Experimental Physics, Faculty of Physics, University of Warsaw, Pasteura 5 Street, 02-093 Warsaw, Poland § International Institute of Molecular and Cell Biology in Warsaw, 4 Ks. Trojdena Street, 02-109 Warsaw, Poland ‡

S Supporting Information *

ABSTRACT: The 5′ cap consists of 7-methylguanosine (m7G) linked by a 5′−5′-triphosphate bridge to messenger RNA (mRNA) and acts as the master regulator of mRNA turnover and translation initiation in eukaryotes. Cap analogues that influence mRNA translation and turnover (either as small molecules or as part of an RNA transcript) are valuable tools for studying gene expression, which is often also of therapeutic relevance. Here, we synthesized a series of 15 dinucleotide cap (m7GpppG) analogues containing a 5′phosphorothiolate (5′-PSL) moiety (i.e., an O-to-S substitution within the 5′-phosphoester) and studied their biological properties in the context of three major cap-binding proteins: translation initiation factor 4E (eIF4E) and two decapping enzymes, DcpS and Dcp2. While the 5′-PSL moiety was neutral or slightly stabilizing for cap interactions with eIF4E, it significantly influenced susceptibility to decapping. Replacing the γ-phosphoester with the 5′-PSL moiety (γ-PSL) prevented β-γpyrophosphate bond cleavage by DcpS and conferred strong inhibitory properties. Combining the γ-PSL moiety with α-PSL and β-phosphorothioate (PS) moiety afforded first cap-derived hDcpS inhibitor with low nanomolar potency. Susceptibility to Dcp2 and translational properties were studied after incorporation of the new analogues into mRNA transcripts by RNA polymerase. Transcripts containing the γ-PSL moiety were resistant to cleavage by Dcp2. Surprisingly, superior translational properties were observed for mRNAs containing the α-PSL moiety, which were Dcp2-susceptible. The overall protein expression measured in HeLa cells for this mRNA was comparable to mRNA capped with the translation augmenting β-PS analogue reported previously. Overall, our study highlights 5′-PSL as a synthetically accessible cap modification, which, depending on the substitution site, can either reduce susceptibility to decapping or confer superior translational properties on the mRNA. The 5′-PSL-analogues may find application as reagents for the preparation of efficiently expressed mRNA or for investigation of the role of decapping enzymes in mRNA processing or neuromuscular disorders associated with decapping.

■

INTRODUCTION

Phosphohydrolytic reactions play an important role in RNA biology. Indeed, cleavage of the phosphoanhydride bonds within the mRNA (mRNA) 5′ cap structure, known as decapping, is a key step in the regulation of mRNA stability and turnover.3,4 The 5′ cap, composed of 7-methylguanosine (m7G) linked by a 5′− 5′-triphosphate bridge to the first transcribed nucleotide, is the hallmark of the 5′ end of an mRNA molecule and is recognized by several specialized proteins involved in mRNA processing, transport, translation, and degradation (Figure 1).5−7 There are two major decapping enzymes engaged in bulk mRNA degradation, DcpS and Dcp1−Dcp2. Interestingly, while these two enzymes differ in substrate specificity, participate in different

Phosphate and oligophosphate mono- and diesters are key structural elements of many biomolecules, including nucleotides and nucleic acids. The enzymes governing phosphoester and phosphoanhydride bond cleavage within DNA and RNA participate in gene expression, cellular signaling, and metabolism.1,2 Consequently, phosphate ester and anhydride analogues have been invaluable tools in deciphering the mechanisms of biophosphate-related enzymatic transformations.2 Unfortunately, many of the analogues used to study these enzymes are complex, difficult to synthesize, or have biocompatibility issues, thus restricting their use and limiting our understanding of enzymatic function during various processes, including RNA processing. © 2018 American Chemical Society

Received: March 7, 2018 Published: April 20, 2018 5987

DOI: 10.1021/jacs.8b02597 J. Am. Chem. Soc. 2018, 140, 5987−5999

Article

Journal of the American Chemical Society

Figure 1. Structure, function, and applications of the mRNA 5′ cap and its chemically modified analogues. (A) Schematic structure of mRNA and its 5′ end. Sites for triphosphate bridge hydrolysis by major decapping enzymes are denoted with scissors. (B) The involvement of cap-recognizing proteins, eIF4E, DcpS, and Dcp2, in mRNA translation and degradation. (C) Structure of β-S-ARCA (m27,2′‑OGppSpG), a previously reported phosphorothioate cap analogue existing as two P-diastereomers, both conferring superior translational properties to mRNA. (D) Structure of the 5′-phosphorothiolate moiety, a non-diastereogenic modification incorporated into the mRNA cap structure in this study. (E) The concept of immune system programming relying on transfection of immature dendritic cells (iDCs) with synthetic 5′-capped mRNAs followed by DC maturation.

carrying bridging β-γ modifications (O-to-CH2, O-to-NH) or non-bridging modifications (γ-O-to-S, γ-O-to-BH3, β-O-toBH3).16−19 All these compounds inhibited DcpS in vitro, although at concentrations at least 20-fold higher than RG3039; therefore, their therapeutic potential is rather limited.20 The other major decapping enzyme, Dcp2 belongs to the nucleoside diphosphate linked to another moiety X (Nudix) hydrolase superfamily and is a catalytic subunit of the heterodimeric Dcp1−Dcp2 complex. Dcp2 cleaves 5′ capped mRNA between the α and β phosphates, releasing 7-methylguanosine diphosphate (m7GDP) and a 5′-monophosphorylated mRNA that is exposed to 5′-exonucleolytic degradation (Figure 1A,B).21−24 mRNA decapping by Dcp2 is the first irreversible step in 5′-to-3′ mRNA decay, thereby contributing to the regulation of mRNA fate. Consequently, Dcp2-resistant cap structures prolong mRNA half-life in vivo,25−27 and, if also characterized by high affinity for translation initiation factor 4E (eIF4E), confer superior translational properties to the mRNA.18,27 One of the Dcp2-resistant cap analogues with the most promising properties is β-S-ARCA (Figure 1C). In β-SARCA, a single O-to-S substitution at the β-phosphate induces a longer cellular mRNA half-life and higher translation efficiency.27 Such mRNAs with enhanced properties can benefit therapeutic applications relying on inducing protein expression in vivo, such as delivery of antigens eliciting anticancer immune response (currently under clinical trials; Figure 1E).28,29 Beyond its role in

mRNA decay pathways, and cleave the mRNA cap triphosphate bridge at different sites, they ultimately lead to a similar outcome: products of decapping and degradation of the mRNA (Figure 1A). DcpS, a member of the histidine triad (HIT) superfamily of pyrophosphatases, acts as a scavenger, degrading 7-methylguanine nucleotides arising from 3′-to-5′ mRNA degradation by the exosome, presumably to prevent their undesired accumulation in the cell.8 DcpS cleaves the cap between the γ and β phosphates, releasing 7-methylguanosine monophosphate (m7 GMP) and a nucleoside or oligoribonucleotide 5′diphosphate (Figure 1A,B). Moreover, DcpS has also been suggested to play a more general role in gene expression, particularly in regulation of mRNA splicing, transcript-specific degradation, and microRNA stability.9,10 DcpS has also been identified as molecular target for therapy of spinal muscular atrophy (SMA),11 a neuromuscular disorder caused by mutations in the survival motor neuron (SMN 1) gene.12 Notably, a linear correlation was found between the effectiveness of quinazoline derivatives to inhibit DcpS and their ability to improve motor function in a mouse SMA model.13−15 One of the most potent DcpS inhibitorsquinazoline derivative RG3039recently entered clinical trials, unfortunately without success. Phosphate-modified cap analogues are another class of DcpS inhibitors with applications in research and medicine. Among previously reported, DcpS-resistant cap analogues were those 5988


Article


modifications (none, phosphorothioate, or methylenebisphosphonate). Two compounds were additionally modified with a 2′-O-methyl group within the m7G, making them antireverse cap analogues (ARCAs) (i.e., analogues that can be incorporated into mRNA by in vitro transcription only in the forward orientation).32−34 After their synthesis, we thoroughly investigated the properties of the new analogues in the context of mRNA cap-related biological processes, including eIF4Emediated translation initiation and decapping by both DcpS and Dcp1−Dcp2. To our knowledge, this is the first report outlining the synthesis of multiple novel cap analogues using 5′PSL in addition to evaluating their structural and biochemical properties, including their resistance to enzymatic degradation as well as their ability to inhibit decapping or enhance mRNA translational potential.

bulk mRNA degradation, Dcp2 activity has also been recently linked to a reduced type I interferon antiviral immune response12 and found to act as a quality control enzyme during small nuclear RNA assembly.30 These findings open new directions for Dcp2related research. Despite the significant number of mRNA cap analogues developed to study cap-dependent processes, our understanding of the multifaceted functions of DcpS and Dcp2 is still limited. Furthermore, new decapping enzymes are also being continuously discovered, creating the space for new molecules to be designed to study them.4,31 Of particular interest in this context, are analogues that combine unique, beneficial biological properties with structural simplicity and synthetic availability, making them effective as well as easy to manufacture. In this study, we explored a new type of chemical modification in the mRNA cap, namely a substitution of the 5′-O atom to an S atom in the triphosphate chain, resulting in the formal replacement of the 5′-phosphoester with a phosphorothiolate (5′-PSL) moiety (Figure 1D and Chart 1). We synthesized and

■

RESULTS

Synthesis and Physiological Stability of 5′-PSL Cap Analogues. Synthesis of P−S−C-5′ linkages in nucleotide and oligonucleotide analogues is usually achieved by either SN2 Salkylation of an appropriate phosphorothioate-containing compound with a 5′-iodonucleoside (or -nucleotide) derivative or by taking advantage of phosphorus(III) chemistry.35−38 As the S-alkylation-based approaches do not require nucleoside protection, these were our first choice for the synthesis of the 5′-PSL cap analogues. We tested two strategies, both based on a combination of S-alkylation and phosphorimidazolide chemistries (Schemes 1−3). The first approach involves the multistep synthesis of a mononucleotide analogue containing a terminal phosphorothioate (PS) moiety, followed by S-alkylation with a 5′-deoxy-5′-iodonucleoside in the final step yielding the target dinucleotide. The second approach, which was especially effective for the synthesis of the heavily modified cap analogues (such as 9 or 14), involves the convergent synthesis of two mononucleotide subunits, followed by the transformation of one of them into an imidazolide derivative, and finally, coupling the subunits in a ZnCl2-mediated pyrophosphate bond formation reaction.39 The 5′-deoxy-5′-iodoguanosine derivatives 18 and 19 were obtained from unprotected nucleosides 16, 17 that were treated with I2 and triphenylphosphine in DMF.40,41 The resulting compounds were then converted into the N7-methylated derivatives 20 and 21 using methyl iodide in DMSO (Scheme 1). These reactions resulted in 77% and 44% isolated yields, respectively. Then, 5′-iodonucleosides 18 and 20 were reacted with trisodium thiophosphate in a water-DMF solution to form the corresponding 5′-phosphorothiolates 22, 25 (Scheme 1).42 After ion exchange (IE) purification, the yields of these compounds ranged from 53 to 65%. Compounds 22 and 25 were then converted into the P-imidazolides 23 and 26, respectively, using a dithiodipyridine/triphenylphosphine oxidation−reduction procedure,43 followed by coupling with triethylammonium phosphate or thiophosphate (Scheme 1). These coupling reactions afforded compounds 24, 27 or 28 in good yields (70−80%). Cap analogues 1−6 were synthesized by reacting 18, 20, or 21 with the specified nucleoside 5′-γ-thiotriphosphate18,44,45 (29− 33) in DMSO in the presence of DBU (Scheme 2). After depletion of the starting material, which was determined by RPHPLC analysis (Figure S1), the crude products 1−6 were purified by IE chromatography followed by semipreparative RPHPLC. This purification resulted in isolated yields of 5−51%.

Chart 1. Structures of the Cap Analogues Synthesized in This Study

biochemically characterized a set of 15 m7GpppG-derived, dinucleotide cap analogues containing a 5′-PSL moiety (compounds 1−15, Chart 1). The analogues primarily differ in the position of the PSL moiety (next to the m7G, next to guanosine (G), or at both sites), the number of phosphate groups (di- or triphosphate), and the presence of additional phosphate 5989


Article


Scheme 1. Synthetic Routes to Guanine and N7-Methylguanine Mononucleotide Analogues Containing a 5′-PSL Moietya

Conditions: (i) I2, imidazole, PPh3, NMP; (ii) Na3PSO3·12H2O, DMF; (iii) imidazole, PPh3, TEA, DTDP, DMF; (iv) H2PO4−·Et3NH+, DMF; (v) H2PSO3−·Et3NH+, DMF; (vi) MeI, DMF.

a

Scheme 2. Synthesis of Dinucleotides 1−6a

a

Conditions: (i) DBU, DMSO.

evaluating the biochemical properties of dinucleotides 1−15, we tested their chemical stability in aqueous buffers pH 3−9. After incubating for 24 h at 37 °C, an RP-HPLC analysis did not reveal any traces of analogue decomposition in the acidic and neutral buffers (Figure S3). At pH 9, slow 7-methylguanine imidazole ring opening was observeda reaction typical to all 7methylguanosine nucleotides47,48 (Figure S3, 4 open and 15 open). Next, we evaluated the general enzymatic susceptibility of selected cap analogues in fetal bovine serum (FBS). FBS contains various nucleases capable of hydrolyzing pyrophosphate and phosphodiester bonds and has been previously employed to study the general stability of modified nucleotides and oligonucleotides under physiological conditions.49 Cap analogues 2, 4, 5, and 8−15 were incubated in 10% FBS at 37 °C and analyzed by RP-HPLC at different time points (Figure S4A). The percentage of cap analogue remaining in the solution was plotted against time, and an exponential decay model was fitted to the experimental data to determine the half-life values (Figure S4B and Table S1). The majority of the selected cap analogues had half-lives similar to the reference analogue, m7GpppG (t1/2 = 26.9

Analogues 7−9 (Scheme 3) were obtained from imidazolides 23 and 26 via coupling with 5′-S-GDP (24), m7GpCH2p (34),16 or m7,2′‑OGDP (35)34 in the presence of excess ZnCl2. The isolated yields after IE and RP-HPLC purification varied from 6 to 40%. The synthesis of cap analogues 10−15 (Scheme 3), which bear two or three modifications (α-PSL, γ-PSL, or β-PS),27 was more demanding and required the use of freshly prepared 5′-Sm7GDP-βS (28) and m7GDP-βS (29), which due to their instability upon storage had to be reacted immediately after purification. These reactions involved coupling of 28 and 29 with 23 or 36, yielding cap analogues 10−15, which contain both PS and PSL moieties and are synthesized in the form of two Pdiastereoisomers designated D1 and D2 according to their elution order during RP-HPLC. (Scheme 3 and Figure S2A). The crude products were purified by IE chromatography followed by semipreparative RP-HPLC, resulting in product yields ranging from 28 to 56% (Figure S2B). Notably, nucleoside 5′-phosphorothiolates are susceptible to hydrolytic cleavage under acidic conditions.46 Therefore, before 5990


Article

Journal of the American Chemical Society Scheme 3. Synthesis of Dinucleotides 7−15a

a

Conditions: (i) ZnCl2, DMF.

± 1.3 min), suggesting that the 5′-PSL modification has little influence on analogue stability in FBS (Table S1). The Interaction of 5′-PSL Cap Analogues with eIF4E Closely Mimics That with Unmodified Caps. The interaction between the mRNA 5′ cap and eIF4E is crucial for canonical initiation of protein biosynthesis.50,51 In vitro transcribed mRNAs carrying cap analogues with a lower affinity for eIF4E than the natural cap show decreased translatability.25 Thus, high affinity for eIF4E is a desirable feature of synthetic cap analogues designed for mRNA engineering applications. To assess the influence of the 5′-PSL moiety on the eIF4E−cap interaction, we determined the binding affinities of analogues 1− 15 for murine eIF4E using time-synchronized fluorescence quenching titration (FQT, Figure 2A).52 The equilibrium association constants (KAS) calculated from replicate FQT experiments are summarized in Figure 2B and Table S2. Our results indicate that the 5′-PSL modification generally does not destabilize the complex with eIF4E, and in some cases, even stabilizes the interaction (Figure 2B). For example, compound 4 carrying a γ-5′-PSL moiety (i.e., a 5′-O-to-S substitution next to the m7G), had 1.6-fold higher binding affinity for eIF4E than m7GpppG. Similarly, a corresponding 2′-Omethylated derivative of 4 (compound 6) had a KAS that was 1.7fold higher compared to that of the parent compound,

Figure 2. The presence of the 5′-PSL moiety does not significantly alter cap-binding by translation initiation factor eIF4E. (A) Representative time-synchronized fluorescence quenching titration (FQT) curves of meIF4E with compounds 4, 15, and reference cap analogue (m7GpppG). (B) KAS values for cap-eIF4E complexes for analogues 1−15 determined by FQT (mean values from triplicates). (C) Structure of cap analogue 9 in complex with meIF4E (colored sticks) overlaid with the structure of m7GpppG−eIF4E (from PDB ID: 1L8B) (gray sticks, only the ligand is shown for clarity).

m27,2′‑OGpppG. Interestingly, the presence of an α-PSL modification (i.e., a substitution next to the G; compounds 2 and 7) or combination of the α-PSL and γ-PSL moieties (compound 9) minimally influenced eIF4E affinity. Analysis of the binding affinities for compounds 1, 3, 5, 8, and 10−15, which 5991


Article


Figure 3. Compound 15 potently inhibits hDcpS. (A) Susceptibility of m7GpppG (upper) and analogue 15 (lower) to hDcpS analyzed using RP-HPLC. m7GpppG is completely hydrolyzed within 15 min, while compound 15 is resistant. Data for all compounds are shown in Table 1. (B) Enzymatic assays used for hDcpS inhibitor evaluation: HTS assay based on artificial substrate20 used for initial evaluation of compounds 1−15; in vitro HPLC assay using unmodified m7GpppG as a substrate; and HeLa cell extract assay using fluorescein-labeled cap analogue as a substrate. (C−E) Dose response curves (mean values from triplicates) for selected cap analogues and RG3039 obtained in the three assays. (F) X-ray structure of cap analogue 15 in complex with hDcpS (residues 38−337). Left: Overall view of hDcpS-compound 15 complex. Color coding: teal, C-terminal domains; orange, N-terminal domains; purple, hinge regions; yellow sticks, histidine triads; gray sticks, overlaid m7GpppG ligand from the previously reported complex with H277NhDcpS (PDB id: 1ST0).56 Inset: Close-up view of the catalytic center in the closed state with both ligands. Additional amino acid contacts observed only for cap analogue 15 are marked in yellow, while the direction of the nucleophilic attack by His277 is marked as a red dashed line. (G) Comparison of ligand conformations presented in two orientations, showing different conformations and positions of triphosphate chains and guanosine moieties.

combine one or two 5′-PSL moieties with other previously reported triphosphate modifications, also confirmed the general compatibility of the 5′-PSL moiety with cap−eIF4E interaction. Combination of the 5′-PSL moiety with the triphosphate bridge shortened to diphosphate (compounds 1 and 3) or an

additional CH2 modification between the phosphates (compounds 5 and 8) weakened the interaction with eIF4E compared to m7GpppG (Table S2), as expected from previous studies.25,52 However, the beneficial effect of the γ-PSL moiety was still observed for these analogues, as the γ-PSL-containing com5992


Article

Journal of the American Chemical Society Table 1. Susceptibility to hDcpS Hydrolysis and hDcpS Inhibition Potency of 5′-Phosphorothiolate Cap Analogues compound

susceptibility to DcpS hydrolysisa

% inhibb

RG3039

NA

108 ± 1

m7Gpp m7GppSG (1) m7GpppSG (2) m7GSppG (3) m7GSpppG (4) m7GSppCH2pG (5) m27,2′‑OGSpppG (6) m27,2′‑OGpppSG (7) m7GpCH2ppSG (8) m7GSpppSG (9) m7GppSpSG D1 (10) m7GppSpSG D2 (11) m7GSppSpG D1 (12) m7GSppSpG D2 (13) m7GSppSpSG D1 (14) m7GSppSpSG D2 (15)

resistant hydrolyzed hydrolyzed resistant resistant resistant resistant hydrolyzed resistant resistant hydrolyzed hydrolyzed resistant resistant resistant resistant

69.5 ± 1.4 37.4 ± 9.5 38.8 ± 6.8 75.0 ± 4.8 93.5 ± 3.4 67.7 ± 1.8 24.1 ± 5.1 28.6 ± 2.3 49.1 ± 3.7 106 ± 3 25.4 ± 12.9 33.4 ± 4.6 107 ± 1 107 ± 1 107 ± 2 109 ± 1

m7GSp (25) m7GSpp (27)

resistant resistant

13.3 ± 12.5 84.8 ± 3.9

IC50 [μM]c 0.0478 ± 0.0106 (0.0707 ± 0.0164)d 5.2 ± 1.2 ND ND 2.93 ± 0.58 0.87 ± 0.17 6.2 ± 1.5 22.6 ± 6.3 72 ± 17 5.8 ± 1.3 0.285 ± 0.065 ND ND 0.178 ± 0.042 0.159 ± 0.027 0.214 ± 0.040 0.0430 ± 0.0080 (0.0447 ± 0.0089)d ND 1.52 ± 0.36

a Determined by HPLC. Conditions: 20 μM cap analogue, 10 nM hDcpS, 50 mM Tris/HCl buffer containing 200 mM KCl, 0.5 mM EDTA (pH 7.6), 20 °C. Cap analogues assigned “resistant” remained unhydrolyzed after 60 min. bFrom fluoride-release HTS assay: 60 μM m7GMPF (substrate), 5 μM inhibitor (1−15), 50 nM hDcpS, 30 °C, 50 mM Tris/HCl, 200 mM KCl, 0.5 mM EDTA, 0.75 mg/mL BSA (pH 7.6). Values were calculated as described in experimental; %inhib values above 100% result from fluctuations in background fluorescence levels. cFrom fluoride-release HTS assay: conditions as in b, except a 10-point half-log dilution series of inhibitor starting from 158 or 50 μM was tested. dValues in brackets are from the HPLC assay: conditions as in b, except 60 μM m7GpppG was used as a substrate.

hydrogen bonds with the triphosphate bridge), as well as several ordered water molecules buried in the binding pocket. The only differences resulting from the 5′-O-to-S substitutions included subtle changes in the length of the hydrogen bonds involving the γ-phosphate moiety, but the overall alignment of the triphosphate bridge remained unchanged. γ-PSL Modification Confers Resistance to Hydrolysis by hDcpS. We next investigated the susceptibility of the PSL cap analogues to hydrolysis by the human DcpS enzyme (Figure 3A and Figure S5). The cap analogues (1−15) or reference analogue m7GpppG (20 μM) were incubated with 10 nM hDcpS enzyme for 15, 30, or 60 min, followed by RP-HPLC analysis to estimate the percentage of remaining substrate. The results are summarized in Table 1, and representative RP-HPLC profiles are shown in Figure S5A−D. In this assay, 48% of the m7GpppG was hydrolyzed by hDcpS within 60 min. In contrast, cap analogues 3−6, 8, 9, and 12−15 were completely resistant to hydrolysis by hDcpS under the same conditions, indicating that 5′-O-to-S substitution next to the m7G effectively protects the cap structure from degradation. These findings are similar to those previously reported for O-to-CH2 substitution at the β−γ bridging position (exemplified here by compound 8) and several other modifications within the γ- or β-phosphate.17,19 In contrast, the PSL modification next to the G moiety (compounds 1, 2, 7, 10, or 11) minimally impacted cap susceptibility to hydrolysis by hDcpS. The susceptibility of cap analogues 3−6, 8, 9, and 12−15 was also studied under a higher enzyme:ligand ratio (200 nM hDcpS and 10 μM cap analogue), but this change did not affect the results (Figure 3A and Figure S5E,F). 5′-PSL Cap Analogues Resistant to hDcpS Hydrolysis Potently Inhibit Its Activity. The reduced hDcpS susceptibility observed for some of the synthesized analogues prompted

pounds 3 and 5 were observed to have higher eIF4E affinity than the corresponding compounds with α-PSL (compounds 1 and 8, respectively). In contrast, the combination of the 5′-PSL-moiety with a β-phosphorothioate moiety (β-PS; compounds 10−15) yielded analogues that had higher affinities for eIF4E than m7GpppG, with the D1 diastereomers having a higher affinity than the corresponding D2 isomers (Table S2). These data are in good agreement with previously reported properties of β-PS cap analogues.18 There was no additional effect provided by α-PSL in combination with β-PS, since analogues 10 and 11 had binding affinities very similar to those for corresponding diastereomers of parent compounds, m7GppS pG D1 or D2 (Table S2). Intriguingly, the combination γ-PSL and β-PS (compounds 12 and 13) or the combination of all three modifications (compounds 14 and 15) yielded analogues with KAS values lower than those for m7GppSpG D1 or D2, respectively, suggesting a slightly destabilizing effect of γ-PSL in these analogues. The influence of the 5′-PSL moiety on the cap−eIF4E interaction was further studied by X-ray crystallography. The cocrystal structure of analogue 9 (which contains both α-PSL and γ-PSL modifications and had an affinity for eIF4E comparable to m7GpppG) was determined to 1.92 Å resolution (PDB ID: 5OSX). Comparison of this structure with that of eIF4E-m7GpppG complex (PDB ID: 1L8B, Figure 2C),52 revealed very similar protein and ligand conformations, and protein−ligand contacts in both structures. This indicates that analogue 9 closely mimics m7GpppG in the eIF4E cap binding pocket, which is also consistent with our FQT data (Table S2). Indeed, both ligands were tightly bound by Trp56 and Trp102 (cation−π stacking with m7G), Glu103 (hydrogen bonds to N1 and N2 of m7G), Arg157 and Lys162 (ionic contacts and 5993


Article


fold more potent than RG3039 in this cell extract assay (EC50 = 0.70 ± 0.09 and 6.82 ± 1.08 nM, respectively). Finally, since analogue 15 appears to interact with both DcpS and eIF4E (Tables S2 and S3), we further investigated the selectivity of this inhibitor. Cap analogues targeting eIF4E have been shown to inhibit cap-dependent translation in cell-free systems.54 Therefore, we tested the effect of analogue 15 on the translation of ARCA-capped mRNA encoding firefly luciferase in HeLa cell extracts at concentrations that are inhibitory to hDcpS. We found that translation was inhibited at compound concentrations at least 100-fold higher than those required for inhibition of hDcpS (Figure S9), indicating that the concentration of compound 15 required for hDcpS inhibition would not disrupt normal mRNA translation. Co-crystallization of hDcpS with Analogue 15 Provides Insights into the Molecular Basis of Enzymatic Resistance and Inhibition. To elucidate the molecular basis of hDcpS resistance and to better understand the role of the three O-to-S substitutions in analogue 15, we determined the X-ray crystal structure of 15 in complex with hDcpS (residues 38−337) at 2.06 Å resolution (PDB ID: 5OSY, Figure 3F). hDcpS is a homodimeric enzyme with two active sites. Structural studies have revealed that in the apo state, both sites are symmetric and open.55 In the previously reported crystal structures of hDcpS in complex with either m7GpppG (PDB ID: 1ST0)56 or m7GDP (PDB ID: 1XMM),57 two distinct binding sites are observed and are associated with the open and closed states.57 The conformation change involves the movement of the entire Nterminal domain relative to the C-terminal domain.55,58,59 In our hDcpS-15 complex, the overall protein structure was similar to that previously described. However, significant differences were observed in the conformation of several amino acid side chains in the enzyme active site in both the closed and open states (Figure S10A). The closed state mimics the substrate-bound complex and involves numerous contacts formed between the protein and cap molecule, whereas in the open state, the ligand is less tightly bound and mimics the product-bound complex.56,57 Electron density maps (Fo − Fc maps) were clearly defined for inhibitor molecules bound to both active sites. In chain A (closed state), we were able to model the whole molecule of analogue 15, while in chain B (open state), the electron density was only visible for the m7G portion (Figure S10B). After refinement, a discontinuity in the Fo − Fc map for the γphosphate of the ligand bound in the closed state was observed, indicating an excess of electron density in our model. This phenomenon is likely due to a partial P−S bond break. Therefore, we also modeled two fragments of the initial molecule: 5′-thio-7-methylguanosine and 5′-thioguanosine-5′(2-thiodiphosphate), assuming that the γ-phosphate is disordered or absent in the cleaved product. These two alternate ligand set-ups (Figure S10B) were refined to group occupancies of 0.59 and 0.41 for the cleaved and non-cleaved molecules, respectively. The occurrence of partial 5′-S−Pγ bond cleavage under crystallization conditions was confirmed by RP-HPLC (Figure S11). Despite this 5′-S−Pγ bond breakage, the resulting fragments, with the exception of the γ-phosphate moiety, remained very well ordered inside the active site and retained the conformation of the initial, non-hydrolyzed inhibitor. Surprisingly, cap analogue 15 adopted a very compact conformation (Figure 3G), with an S-shaped triphosphate chain forming the core of the molecule. This chain appears to be stabilized by intramolecular hydrogen bonds between the 2′and 3′-hydroxyl groups of the ribose rings and the two

us to evaluate their inhibitory properties toward this enzyme. This was performed using a fluorescence-based high-throughput screening (HTS) assay (Figure 3B, top panel; Figures S6 and S7).20 The assay is based on the use of the unnatural hDcpS substrate 7-methylguanosine 5′-fluoromonophosphate (m7GMPF). Upon hydrolysis by hDcpS, m7GMPF releases fluoride ions, which can then be quantified by reacting them with a fluorogenic probe (bis(tert-butyldimethylsilylfluorescein)), thereby generating a fluorescent signal that is directly proportional to F− concentration. In the presence of an hDcpS inhibitor, fluorescence signal would be attenuated. Our initial screening was carried out in the presence of 60 μM substrate (m7GMPF), 5 μM inhibitor, and 50 nM hDcpS (monomer) and included 5′-PSL-containing cap analogues 1− 15 and mononucleotides 25 and 27 along with known hDcpS inhibitors (RG3039,14 m7GDP,53 and m7GpSppG (D1/D2)18) as references. The percentage of inhibition (% inhib) for each compound (Figure S6) was calculated as the ratio of signal intensity lost in the presence of inhibitor to the maximum response in the absence of an inhibitor (Table 1). This screening revealed that a group of cap analogues (compounds 4, 9, and 12− 15) strongly inhibit hDcpS (Figure S6). For these analogues, the IC50 parameters were also determined using the same HTS assay (Figure 3C, Figure S7, and Table 1). Notably, compound 4 (i.e., the m7GpppG analogue containing a single γ-PSL) appears to be a potent hDcpS inhibitor, with an IC50 of ∼0.87 μM, which is comparable to the most potent cap-derived hDcpS inhibitor reported in the literature (m7GppBH3pG D1, IC50 ≈ 1 μM under the same conditions).20 Structural alterations of analogue 4, such as shortening the triphosphate to diphosphate (compound 3), introduction of an O-to-CH2 substitution at the α-β bridging position (compound 5), or the addition of a 2′-O-methyl group to m7G (compound 6), all decreased the inhibition potency (by 3.4-, 7-, and 26-fold, respectively). Interestingly, combining the γPSL and α-PSL moieties in one compound increased inhibition by 3-fold (compound 9, IC50 = 0.285 μM). The inhibitory properties were also enhanced when the γ-PSL and β-PS moieties were combined, regardless of the configuration of the βP-stereocenter (compounds 12 and 13; 4.9- and 5.5-fold increased potency, respectively). The combination of all three modifications (γ-PSL, α-PSL, and β-PS) produced the strongest inhibitors among the tested compounds (compounds 14 and 15; Table 1) with compound 15 (IC50 = 43 nM) showing the highest level of inhibition, which was comparable to that of RG3039 in the same assay. To verify these results, analogue 15 was further analyzed with two other independent assays (Figure 3B, middle panel). An RPHPLC inhibition assay using m7GpppG as a substrate confirmed that the inhibition potency of 15 toward recombinant hDcpS is at nanomolar level and similar to RG3039 (Figure 3D). Finally, a fluorescence-based HPLC assay using cytoplasmic HeLa cell extracts was applied to verify the inhibitory properties of 15 against endogenous hDcpS activity. In this experiment, we monitored the degradation of fluorescein-labeled cap analogue (m7GpppA-FAM) in the extract (Figure 3B, bottom panel). Notably, concentration-dependent inhibition of probe cleavage was observed in the extracts in the presence of either compound 15, RG3039, or m7GpCH2ppG − a previously reported hDcpSresistant cap analogue with moderate inhibitory properties16,20 (Figure S8). As expected, both 15 and RG3039 were significantly more potent than m7GpCH2ppG (Figure 3E). Interestingly, in contrast to the other in vitro assays, compound 15 was about 105994


Article


9; DS = 0.44). The combination of γ-PSL with an O-to-CH2 substitution at the α-β-position resulted in similar DS (compound 5, DS = 0.14) compared to compounds containing only the γ-PSL. In contrast, combination of α-PSL with an O-toCH2 substitution at the β-γ-position, did not confer resistance (compound 8, DS = 0.69). Surprisingly, the combinations of PSL and β-PS modifications (as in compounds 10−15) increased resistance to decapping by SpDcp1−Dcp2 only in the case of compounds 10 (α-PSL and β-PS combined, DS = 0.11) and 14 (α-PSL, γ-PSL, and β-PS combined, DS = 0.30). Next, we tested the influence of the γ-PSL modification on RNA susceptibility to decapping by hDcp2. To this end, transcripts capped with ARCA analogues 6 and 7 were incubated with recombinant hDcp2 for up to 60 min and analyzed electrophoretically (Figure 4). The hDcp2 decapping assay

nucleobases located at the opposite sites. The arrangement of the m7G was similar to that observed in the m7GpppG and m7GDP complexes, with the m7G base stacked between Trp175 and Leu206 and the ribose in a 2′-endo puckering and C4′−C5′-sc (trans−gauche) conformation. One notable difference was the slightly elongated 5′-S−Pγ bond pointing in a different direction compared to the 5′-O−Pγ bond in m7GpppG, placing the γphosphate of 15 farther away from the HIT motif, particularly the His277 residue responsible for the nucleophilic attack during catalysis (Figure 3F).56 The negatively charged β-PS sulfur atom was sticking out toward a hinged α-helix and formed a strong ionic contact with the positively charged amine group of Lys142 and a hydrogen bond with a hydroxyl group of Ser272. One of the oxygen atoms of the adjacent α-phosphate also interacted with the hinge region residues (Lys142 and Tyr143), while the second oxygen atom was hydrogen bonded to the phenolic group of Tyr273, which plays a crucial role in closing the active site upon cap binding and stabilizing the closed state.56 RNAs Capped with Analogues Carrying the γ-PSL Moiety Are Resistant to Hydrolysis by hDcp2. In vitro resistance to the mRNA decapping enzyme Dcp2 is an important predictor of cap analogue-mediated mRNA stabilization. Unlike DcpS, Dcp2 acts at the beginning of the mRNA decay pathway, cleaving the cap between the α and β phosphates to release m7GDP and exposing the 5′-phosphorylated RNA to further degradation. For efficient catalysis, Dcp2 requires the presence of the RNA body as well as divalent metal ions. The effect of 5′-PSL on the susceptibility of the cap to Schizosaccharomyces pombe (Sp) Dcp1−Dcp2 and human Dcp2 was tested on short (25 nt) RNAs carrying 5′-PSL cap analogues.60 SpDcp1−Dcp2 susceptibility was tested for RNAs capped with analogues 2 and 4−15 (the diphosphate caps 1 and 3 were excluded from the analysis due to their unfavorable interactions with eIF4E). The caps were incorporated into the RNA by in vitro transcription, followed by trimming of the 3′ ends of the transcripts with DNAzyme 10−23 to reduce 3′-end heterogeneity and facilitate analysis.61 The transcripts were incubated with the SpDcp1−Dcp2 complex for various periods of time and resolved in high resolution polyacrylamide gel to separate capped and uncapped RNAs and estimate the progress of decapping over time (Figure S12A). The initial fractions of capped RNA versus total RNA (at time point 0) were used to estimate the capping efficiencies (Table S2). For all tested analogues capping efficiencies were above 70% indicating efficient incorporation into RNA by SP6 polymerase. To compare decapping susceptibilities of different caps, the fraction of capped RNA present in each sample was plotted as a function of time (Figure S12B). The greatest differences were observed at the 15 min time point; therefore, this time point was used for quantification (Table S2). Transcripts capped with the m7GpppG, ARCA, and β-S ARCA D2 reference compounds had decapping susceptibilities (DS, or the fraction of decapped RNA after 15 min) of 0.69, 0.52, and 0.43, respectively. Further, RNA capped with compounds 2 and 4 had DSs of 0.68 and 0.18, respectively, indicating that the presence of a 5′-O-to-S substitution next to the m7G (γ-PSL), but not next to the G (α-PSL), notably decreased the susceptibility of the RNA to SpDcp1−Dcp2. Consistent with this observation, the ARCA versions of caps 2 and 4 (i.e., compounds 7 and 6, ensuring their correct orientation) had analogous properties (DS = 0.52 and 0.07, respectively). Interestingly, the combination of two 5′-PSL modifications in one compound appeared to diminish the stabilizing effect compared to a single γ-PSL moiety (compound

Figure 4. A 5′-PSL moiety next to the m7G (γ-PSL) protects RNA from decapping by Dcp2. (A) Schematic representation of the hDcp2 susceptibility assay. (B) Example results from the assay (RNAs resolved by denaturing polyacrylamide gel electrophoresis (PAGE) and stained with SYBR Gold). (C) Susceptibility of 25-nt transcripts capped with various cap analogues to hDcp2. Fraction of capped RNA remaining in the reaction mixture is plotted as a function of time. Data are reported as the average values from two biological replicates ± SD. All data were normalized to time 0.

confirmed that the 5′-PSL modification stabilized RNA against decapping, but only if it was located at the γ-position (Figure 4 and Table 2). The DSs for RNAs capped with compounds 6 and Table 2. Biochemical Properties of Compounds 6 and 7 compound m7GpppG m27,3′‑OGpppG (ARCA) m27,2′‑OGppSpG D2 (β-S ARCA D2) m27,2′‑OGSpppG (6) m27,2′‑OGpppSG (7)

decapping susceptibilitya

normalized relative protein expressionb

mRNA halflife (h)c

0.69 ± 0.03 0.55 ± 0.04

1 1.90 ± 0.27

1.35 ± 0.34 1.28 ± 0.29

0.14 ± 0.03

2.70 ± 0.30

n.d.

0.18 ± 0.05 0.60 ± 0.19

2.07 ± 0.19 2.75 ± 0.29

1.54 ± 0.40 1.34 ± 0.23

a

Fraction of decapped RNA after a 15 min incubation with hDcp2. Data shown are the average values from duplicates ± SD. bTotal protein expression in cell lysates of firefly luciferase capped with given analogue normalized to the expression of m7GpppG-capped mRNA. The average values from three biological replicates ± SD are reported. c mRNA half-life of firefly luciferase transcript capped with given analogue in HeLa cells. The average values from three (two for m7GpppG) biological replicates ± SEM are reported. n.d. = not determined. 5995


Article

Journal of the American Chemical Society 7 were 0.18 ± 0.05 and 0.60 ± 0.19, respectively, compared to 0.69 ± 0.03 and 0.55 ± 0.04 for the reference compounds m7GpppG and ARCA. Interestingly, in contrast to the results from the SpDcp1−Dcp2 assay, the susceptibility of transcripts capped with compound 6 was comparable to that of RNA capped with β-S ARCA D2 (DS = 0.14 ± 0.03). α-PSL Enhances Cap-Dependent Translation. After biophysically and biochemically characterizing all of the novel cap analogues in the context of translation and decapping machinery, we investigated the properties of the mRNA molecules capped with these compounds. The influence of selected 5′-PSL cap analogues on mRNA expression was investigated in a cell-free translation system (rabbit reticulocyte lysate, RRL) and in cultured HeLa cells. First, a set of mRNAs encoding Renilla luciferase with modified 5′ ends were synthesized in vitro with SP6 RNA polymerase and translated in RRL. The experiments were performed under conditions optimized for cap-dependent translation as previously described.62 The cap-dependent nature of the system was confirmed by the low expression of negative control mRNA carrying an unmethylated (non-functional) 5′-cap analogue (GpppG) (Table S2). The results obtained for the mRNAs capped with analogues 1−15 and functional reference cap structures [m7GpppG, ARCA (m27,3′‑OGpppG), and β-S ARCA

luciferase was plotted as a function of time (Figure 5A), and total protein expression was calculated for each mRNA based on the

Figure 5. mRNAs capped with 5′-PSL cap analogues 6 and 7 are efficiently translated in living cells. (A) Representative time course of firefly luciferase mRNA expression in HeLa cells for a single biological replicate when the specified 5′-cap analogues were incorporated. (B) Total protein expressions in HeLa cells for various capped mRNAs. Bars represent integers under the expression curves shown in (A) normalized to m7GpppG. Data are reported as the average values from three biological replicates ± SD.

activity curve (Figure 5B and Table 2). Consistent with the translation efficiencies obtained in our RRL experiments, total protein expression was higher for mRNAs capped with analogue 7 compared to those capped with analogue 6 or unmodified ARCA (2.07 ± 0.19, 2.75 ± 0.29, and 1.90 ± 0.27, respectively). Notably, transcripts capped with compound 7 yielded similar amounts of protein as mRNA capped with β-S ARCA D2 (total protein expression = 2.70 ± 0.30) (Figure 5B and Table 2). Interestingly, total protein expression for mRNAs capped with analogue 6 was comparable to values obtained for ARCA capped mRNAs, despite that RNAs with compound 6 at their 5′ ends were more stable against hDcp2 than RNAs capped with ARCA or analogue 7 in an in vitro assay (Figure 4). This may suggest that resistance to hDcp2 hydrolysis may not translate into increased stability in vivo. Therefore, stability of mRNA capped with compound 6 and 7 in HeLa cells was investigated. To this end, HeLa cells were transfected with differently capped mRNAs, cells were lysed at various time points, and levels of firefly and GAPDH mRNA were measured by qRT-PCR. We found that the presence of cap analogue 6, 7, or ARCA at 5′ end of firefly mRNA does not significantly differentiate mRNA stability in HeLa cells (Table 2 and Figure S14).

D2 (m27,2′‑OGppSpG D2)] are summarized in Table S2, whereas Figure S13 shows representative data from a single biological replicate. mRNAs carrying the 5′-γ-PSL moiety (compound 4) or 5′-α-PSL moiety (compound 2), or a combination of these two modifications (compound 9) in non-ARCA analogues all had translation efficiencies that were comparable to that of m7GpppG-mRNA, indicating that these changes do not significantly influence in vitro translation. Alternatively, compounds containing the PSL modification and an additional methylenebisphosphonate moiety (α-β (compound 5) or β-γ (compound 8)) had decreased translation efficiency compared to m7GpppG-mRNA, whereas those with an additional β-PS moiety (cap analogues 2, 4, and 9) produced mRNAs that were translated more efficiently. These results are consistent with previous reports for compounds carrying these modifications. As expected, the 5′-PSL-ARCAs (analogues 6 and 7) also augmented translation efficiency (relative translation efficiencies = 1.73 ± 0.24 and 2.23 ± 0.31, respectively) compared to their non-ARCA versions (compounds 4 and 2). Interestingly, these values were between those obtained for the reference ARCAcapped mRNA (m27,3′‑OGpppG; translation efficiency = 1.56 ± 0.14) and mRNAs capped with both β-S-ARCA diastereomers (translation efficiencies = 2.81 ± 0.37 and 3.45 ± 0.42 for m7,2′‑OGppSpG D1 and D2, respectively), suggesting that the 5′PSL moiety in the ARCA-type compounds increases mRNA translation efficiency. The translation properties of transcripts capped with ARCAs 6 and 7 were additionally assessed in HeLa cells. In vitro transcribed mRNAs encoding firefly luciferase capped with compounds 6, 7, or with reference analogues m7GpppG, ARCA (m27,3′‑OGpppG), or β-S ARCA D2 (m27,2′‑OGppSpG D2) were transfected into HeLa cells. To account for variation in the transfection efficiency between samples, mRNA encoding Renilla luciferase capped with ARCA was used as an internal control for each transfection.63 The firefly and Renilla luciferase activities were measured using a dual reporter assay at different time points post-transfection. Firefly luciferase activity normalized to Renilla

■

DISCUSSION 5′-PSL is a sister modification to PS; however, in 5′-PSL, a sulfur occupies the bridging position within the modified phosphate moiety rather than the non-bridging position. Therefore, unlike PS, 5′-PSL does not introduce a new P-stereogenic center into the molecule, simplifying synthesis, purification, and biological evaluation of the resulting compounds. These characteristics make them ideal for the synthesis of mRNA cap analogues and the subsequent study of decapping enzymes, such as DcpS and Dcp2. In the present study, we explored the 5′-PSL linkage as an mRNA 5′ cap modification in 15 novel dinucleotide m7GpppGderived cap analogues. Notably, the present study is the first to explore the synthetic accessibility and biological properties of these particular compounds. The synthetic pathways used to synthesize the 5′-PSL cap analogues, which were based on a combination of SN2 Salkylation for 5′-PSL insertion and P-imidazolide chemistry for 5996


Article


electronic effects introduced by the β-PS moiety. These strong cap−enzyme contacts involving both the C- and N-hDcpS terminal domains may stabilize the closed enzyme state, a feature required for potent inhibition.11 Notably, the potency of analogue 15 in two independent in vitro assays was comparable to that of RG3039, a quinazoline-derived compound that alleviates SMA symptoms in a mouse model, but failed in clinical trials. Furthermore, in a HeLa cell lysate inhibition assay, 15 showed 10-fold greater potency, indicating its potentially higher selectivity. This assay also demonstrated that compound 15 is stable enough under physiological conditions to potently inhibit endogenous hDcpS. Importantly, we established the concentrations of compound 15 that provide complete hDcpS inhibition without any effect on translation. Another advantage of compound 15 is that unlike other small molecule inhibitors it can be easily incorporated into the 5′ end of the RNA by in vitro transcription. Thus, analogue 15 may facilitate studies of hDcpS activity, including the molecular mechanisms involved in long and short capped RNA discrimination as well as definition of the links between decapping, splicing modulation, and therapeutic relevance in SMA. Notably, other cellular and in vivo studies could also be performed provided that the compound is additionally modified or complexed with some carriers to enhance cell permeability.64−68 While additional work is necessary to elucidate the full use of compound 15, the current study firmly places it in the category of potent DcpS inhibitors. Following incorporation into RNA, we also studied the susceptibility of 5′-PSL cap analogues to Dcp2. All novel cap analogues were efficiently incorporated into short RNAs during in vitro transcription, consistent with previously reported findings that nucleoside 5′-phosphorothiolates can serve as RNA polymerase transcription initiators.40,69 RNAs capped with analogues containing a γ-PSL moiety, but not an α-PSL moiety, had increased resistance to both SpDcp1−Dcp2 heterodimer or hDcp2 monomer. Interestingly, this is the same position of modification that also conferred resistance to DcpS. This is unexpected considering that the two enzymes belong to different protein families, utilize different length RNA substrates, and cleave the cap at different positions via different mechanisms. Although we cannot yet explain Dcp2 resistance at the molecular level, it likely results from shifting of the β-phosphate away from the catalytic site in the Nudix domain caused by 5′-C−S−P bond elongation. This suggests that even subtle changes in the cap’s triphosphate chain geometry may cause significant changes in the biochemical properties of the resulting RNA molecules. Finally, we inspected the translational properties of mRNAs capped with 5′-PSL cap analogues in cell-free and cultured cell models. Generally, the 5′-PSL modification did not interfere with translation and in some cases increased the translational potential of the mRNA, especially in the case of analogues carrying the αPSL moiety.70 However, a more detailed analysis of this was not possible because most of the analogues could be incorporated into the mRNA in either the forward or reverse orientations70,71 and these two forms could not be differentiated nor quantified. This issue was circumvented using analogues 6 and 7, which are ARCAs that contain the γ- and α-PSL moieties, respectively. Their translation efficiencies in HeLa cells were compared to those of known cap analogues m7GpppG, m27,3′‑OGpppG (a

pyrophosphate bond formation, turned out to be moderately efficient, but thanks to the low number of synthetic steps appeared to be scalable and compatible with the practical applications of these compounds. Additionally, the chemical stability of these analogues in aqueous solutions of varying pH makes them compatible with biological applications. Although nucleoside 5′-phosphorothiolates undergo acid-catalyzed P−S bond cleavage in aqueous solutions,46 the 5′-PSL cap analogues synthesized here were chemically stable at pH 3−9, indicating that dinucleotides containing the 5′-PSL moiety are more stable than the corresponding mononucleotides. The stability of the cap analogues in blood serum (FBS) was also essentially unaffected by the 5′-PSL moiety. After evaluating the basic biocompatibility of the analogues, we evaluated the biochemical properties of the compounds using a set of assays involving three major cytoplasmic cap-recognizing proteins important for mRNA expression and turnover: eIF4E, DcpS, and Dcp1−Dcp2 complex. Our results show that the presence of the 5′-PSL modification had differential effects on these proteins. For eIF4E, the 5′-PSL moiety did not affect or even slightly stabilized the mRNA−enzyme interaction, indicating that the longer bonds are generally well accommodated in the cap-binding pocket. However, the single β-O-toS substituted cap analogues (e.g., β-S-ARCA D1) had higher affinity than corresponding analogues with multiple substitutions (e.g., analogues 10, 12, and 14), indicating that additional O-to-S substitutions may disrupt the optimal conformation for ligands carrying β-PS moiety. Dual 5′-PSL moieties in compound 9 did not impact eIF4E binding affinity. This corresponded with the eIF4E co-crystal structure, wherein triphosphate chain alignments in compound 9 and parent cap analogue (m7GpppG) were very similar. This is likely because structural perturbations introduced by longer C−S and S−P bonds compared to C−O and P−S bonds are accommodated by C5′−S−P versus C5′− O−P angle differences. The effects of the 5′-PSL moiety in cap analogue−eIF4E interactions was also indirectly supported by the high translational activity observed for the mRNAs capped with 5′-PSL analogues. Conversely, the 5′-PSL moiety does significantly influence the susceptibility of the cap analogues to decapping. Indeed, all of the analogues containing the γ-PSL moiety were resistant to hDcpS, whereas α-PSL had no influence. Moreover, these DcpS-resistant compounds were also potent hDcpS inhibitors. Compound 4, for example, had an IC50 < 1 μM, suggesting that it is more potent than known hDcpS-resistant analogues containing γ-PS, β-γimidodiphosphate, or β-γ-methylenebisphosphonate moieties. Additional O-to-S substitutions further improved these inhibitory properties, leading to the most potent inhibitorthe triplemodified analogue 15 (m7GSppSpSG D2)with low nanomolar potency. The hDcpS−15 complex crystal structure suggests that resistance to decapping and increased affinity for hDcpS likely arises from the accumulation of multiple steric and electronic effects. The primary factor involved in resistance is the altered γphosphate geometry caused by 5′-C−S−P bond elongation, which pushes the γ-phosphorus about 4.1 Å away from the site occupied by the unmodified cap, making it unavailable to the catalytic histidine triad. The additional α-PSL moiety likely ensures higher conformational flexibility enabling the ligand to adopt a more favorable conformation and form new ionic contacts with Lys 142 and Tyr 143, which were not observed in the previously reported structure of catalytically inactive mutant in complex with m7GpppG.52 Finally, it is probable that the interaction with Lys 142 is further stabilized by favorable

standard, commercially available ARCA) and m27,2′‑OGppSpG D2 (β-S-ARCA D2, optimized for efficient translation in vitro and in vivo).70 Notably, the mRNAs capped with 7 were translated more efficiently than mRNAs capped with 6 (despite similar half5997



■

lives) and were comparable to β-S-ARCA D2-terminated mRNAs. Therefore, in our model, it appears that the susceptibility of the 5′-PSL cap analogues to Dcp2 is not the primary determinant of translational activity. Furthermore, the high translational activity of compound 7 combined with its relatively simple synthesis and lack of P-stereoisomerism makes it an appealing alternative for the previously developed β-SARCA.

ACKNOWLEDGMENTS This work was supported by the Foundation for Polish Science (TEAM/2016−2/13), the National Centre of Research and Development in Poland (LIDER/001/003/L-5/13/NCBR/ 2014), the Ministry of Science and Higher Education (Poland, DI2012 024842), and the National Science Centre (ETIUDA 2017/24/T/NZ1/00345). We thank Mike Kiledjian (Rutgers University) for the hDcpS encoding plasmid, Christopher D. Lima (Sloan-Kettering Institute) for the hDcp2 encoding plasmid, John D. Gross (University of California, San Francisco) for plasmids encoding the Dcp1−Dcp2 complex from S. pombe, Stephen R. Ikeda (The National Institute on Alcohol Abuse and Alcoholism) for providing the hRLuc-pRNA2(A)128 plasmid, and Marek R. Baranowski (University of Warsaw) for chemical reagents used in HTS assay. Diffraction data have been collected on BL14.1 at the BESSY II electron storage ring operated by the Helmholtz-Zentrum Berlin.72

■

CONCLUSIONS In conclusion, we employed a wide range of biochemical assays to evaluate a set of 15 chemically synthesized cap analogues containing a 5′-PSL moiety. We found that 5′-PSL stabilizes interaction of the cap structure with eIF4E, can enhance mRNA translation, and in some cases, significantly reduces mRNA cap susceptibility to decapping. These findings provide new structural insights into specific protein−mRNA cap interactions, leading to the selection of novel cap analogues with superior biological properties. These superior analogues include: (i) compound 6 (m27,2′‑OGSpppG), which shows a notable decrease in susceptibility to hDcp2, (ii) compound 7 (m27,2′‑OGpppSG), which enhances mRNA translational potential in a manner that is comparable to β-S-ARCA D2 (m27,2′‑OGppSpG D2), and (iii) compound 15 (m7GSppSpSG D2), which inhibits hDcpS more potently than clinically tested RG3039. Taken together, our study demonstrates that the 5′-PSL moiety confers useful biological properties to mRNA caps, enabling modulation of therapeutically relevant cap-dependent processes. While still some additional work is necessary, this study highlights the 5′PSL moiety as a currently underexplored modification and provides insight into how it can be used to modulate the activity of dinucleoside 5′,5′-oligophosphates in normal biological processes as well as in disease.

■

■

REFERENCES

(1) Bowler, M. W.; Cliff, M. J.; Waltho, J. P.; Blackburn, G. M. New J. Chem. 2010, 34 (5), 784−794. (2) Korhonen, H. J.; Conway, L. P.; Hodgson, D. R. W. Curr. Opin. Chem. Biol. 2014, 21, 63−72. (3) Coller, J.; Parker, R. Annu. Rev. Biochem. 2004, 73, 861−890. (4) Grudzien-Nogalska, E.; Kiledjian, M. Wiley Interdiscip. Rev.-RNA 2017, 8 (1), e1379. (5) Ramanathan, A.; Robb, G.; Chan, S. Nucleic Acids Res. 2016, 44 (16), 7511−7526. (6) Furuichi, Y.; Shatkin, A. J. Adv. Virus Res. 2000, 55, 135−184. (7) Warminski, M.; Sikorski, P. J.; Kowalska, J.; Jemielity, J. Top. Curr. Chem. 2017, 375 (16), 211. (8) Liu, H.; Rodgers, N. D.; Jiao, X.; Kiledjian, M. EMBO J. 2002, 21 (17), 4699−4708. (9) Bail, S.; Kiledjian, M. RNA Biol. 2008, 5 (4), 216−219. (10) Song, M.-G.; Bail, S.; Kiledjian, M. RNA 2013, 19 (3), 390−399. (11) Singh, J.; Salcius, M.; Liu, S. W.; Staker, B. L.; Mishra, R.; Thurmond, J.; Michaud, G.; Mattoon, D. R.; Printen, J.; Christensen, J.; Bjornsson, J. M.; Pollok, B. A.; Kiledjian, M.; Stewart, L.; Jarecki, J.; Gurney, M. E. ACS Chem. Biol. 2008, 3 (11), 711−722. (12) Li, Y.; Dai, J.; Song, M.; Fitzgerald-Bocarsly, P.; Kiledjian, M. Mol. Cell. Biol. 2012, 32 (6), 1164−1172. (13) Gogliotti, R. G.; Cardona, H.; Singh, J.; Bail, S.; Emery, C.; Kuntz, N.; Jorgensen, M.; Durens, M.; Xia, B.; Barlow, C.; Heier, C. R.; Plasterer, H. L.; Jacques, V.; Kiledjian, M.; Jarecki, J.; Rusche, J.; DiDonato, C. J. Hum. Mol. Genet. 2013, 22 (20), 4084−4101. (14) Van Meerbeke, J. P.; Gibbs, R. M.; Plasterer, H. L.; Miao, W.; Feng, Z.; Lin, M. Y.; Rucki, A. A.; Wee, C. D.; Xia, B.; Sharma, S.; Jacques, V.; Li, D. K.; Pellizzoni, L.; Rusche, J. R.; Ko, C. P.; Sumner, C. J. Hum. Mol. Genet. 2013, 22 (20), 4074−4083. (15) Gopalsamy, A.; Narayanan, A.; Liu, S.; Parikh, M.; Kyne, R.; Fadeyi, O.; Tones, M.; Cherry, J.; Nabhan, J.; LaRosa, G.; Petersen, D.; Menard, C.; Foley, T.; Noell, S.; Ren, Y.; Loria, P.; Maglich-Goodwin, J.; Rong, H.; Jones, L. J. Med. Chem. 2017, 60 (7), 3094−3108. (16) Kalek, M.; Jemielity, J.; Darzynkiewicz, Z. M.; Bojarska, E.; Stepinski, J.; Stolarski, R.; Davis, R. E.; Darzynkiewicz, E. Bioorg. Med. Chem. 2006, 14 (9), 3223−3230. (17) Rydzik, A. M.; Kulis, M.; Lukaszewicz, M.; Kowalska, J.; Zuberek, J.; Darzynkiewicz, Z. M.; Darzynkiewicz, E.; Jemielity, J. Bioorg. Med. Chem. 2012, 20 (5), 1699−1710. (18) Kowalska, J.; Lewdorowicz, M.; Zuberek, J.; Grudzien-Nogalska, E.; Bojarska, E.; Stepinski, J.; Rhoads, R. E.; Darzynkiewicz, E.; Davis, R. E.; Jemielity, J. RNA 2008, 14 (6), 1119−1131. (19) Kowalska, J.; Wypijewska del Nogal, A.; Darzynkiewicz, Z. M.; Buck, J.; Nicola, C.; Kuhn, A. N.; Lukaszewicz, M.; Zuberek, J.; Strenkowska, M.; Ziemniak, M.; Maciejczyk, M.; Bojarska, E.; Rhoads, R. E.; Darzynkiewicz, E.; Sahin, U.; Jemielity, J. Nucleic Acids Res. 2014, 42 (16), 10245−10264.

ASSOCIATED CONTENT

* Supporting Information S

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/jacs.8b02597. The atomic coordinates and structure factors files of eIF4E−9 and hDcpS−15 complexes were deposited in the Protein Data Bank under accession codes 5OSX and 5OSY, respectively.

■

Article

Supporting Tables S1−S3, supporting Figures S1−S14, and detailed experimental procedures (PDF) 1 H, 31P NMR, and HRMS spectra for all novel compounds (PDF)

AUTHOR INFORMATION

Corresponding Authors

*[email protected] *[email protected] ORCID

Joanna Kowalska: 0000-0002-9174-7999 Jacek Jemielity: 0000-0001-7633-788X Author Contributions ⊥

B.A.W. and P.J.S. contributed equally.

Notes

The authors declare no competing financial interest. 5998


Article

Journal of the American Chemical Society (20) Baranowski, M. R.; Nowicka, A.; Jemielity, J.; Kowalska, J. Org. Biomol. Chem. 2016, 14 (20), 4595−4604. (21) She, M.; Decker, C. J.; Chen, N.; Tumati, S.; Parker, R.; Song, H. Nat. Struct. Mol. Biol. 2006, 13 (1), 63−70. (22) van Dijk, E.; Cougot, N.; Meyer, S.; Babajko, S.; Wahle, E.; Seraphin, B. EMBO J. 2002, 21 (24), 6915−6924. (23) Wang, Z.; Jiao, X.; Carr-Schmid, A.; Kiledjian, M. Proc. Natl. Acad. Sci. U. S. A. 2002, 99 (20), 12663−12668. (24) Valkov, E.; Jonas, S.; Weichenrieder, O. Curr. Opin. Struct. Biol. 2017, 47, 40−51. (25) Grudzien, E.; Kalek, M.; Jemielity, J.; Darzynkiewicz, E.; Rhoads, R. E. J. Biol. Chem. 2006, 281 (4), 1857−1867. (26) Su, W.; Slepenkov, S.; Grudzien-Nogalska, E.; Kowalska, J.; Kulis, M.; Zuberek, J.; Lukaszewicz, M.; Darzynkiewicz, E.; Jemielity, J.; Rhoads, R. E. RNA 2011, 17 (5), 978−988. (27) Grudzien-Nogalska, E.; Jemielity, J.; Kowalska, J.; Darzynkiewicz, E.; Rhoads, R. E. RNA 2007, 13 (10), 1745−1755. (28) Sahin, U.; Derhovanessian, E.; Miller, M.; Kloke, B.-P.; Simon, P.; Löwer, M.; Bukur, V.; Tadmor, A. D.; Luxemburger, U.; Schrörs, B.; Omokoko, T.; Vormehr, M.; Albrecht, C.; Paruzynski, A.; Kuhn, A. N.; Buck, J.; Heesch, S.; Schreeb, K. H.; Müller, F.; Ortseifer, I.; Vogler, I.; Godehardt, E.; Attig, S.; Rae, R.; Breitkreuz, A.; Tolliver, C.; Suchan, M.; Martic, G.; Hohberger, A.; Sorn, P.; Diekmann, J.; Ciesla, J.; Waksmann, O.; Brück, A.-K.; Witt, M.; Zillgen, M.; Rothermel, A.; Kasemann, B.; Langer, D.; Bolte, S.; Diken, M.; Kreiter, S.; Nemecek, R.; Gebhardt, C.; Grabbe, S.; Höller, C.; Utikal, J.; Huber, C.; Loquai, C.; Türeci, Ö . Nature 2017, 547 (7662), 222−226. (29) Kuhn, A. N.; Diken, M.; Kreiter, S.; Selmi, A.; Kowalska, J.; Jemielity, J.; Darzynkiewicz, E.; Huber, C.; Türeci, O.; Sahin, U. Gene Ther. 2010, 17 (8), 961−971. (30) Shukla, S.; Parker, R. Proc. Natl. Acad. Sci. U. S. A. 2014, 111 (32), E3277−E3286. (31) Taverniti, V.; Séraphin, B. Nucleic Acids Res. 2015, 43 (1), 482− 492. (32) Jemielity, J.; Fowler, T.; Zuberek, J.; Stepinski, J.; Lewdorowicz, M.; Niedzwiecka, A.; Stolarski, R.; Darzynkiewicz, E.; Rhoads, R. E. RNA 2003, 9 (9), 1108−1122. (33) Peng, Z.-H.; Sharma, V.; Singleton, S. F.; Gershon, P. D. Org. Lett. 2002, 4 (2), 161−164. (34) Stepinski, J.; Waddell, C.; Stolarski, R.; Darzynkiewicz, E.; Rhoads, R. E. RNA 2001, 7 (10), 1486−1495. (35) Li, N. S.; Frederiksen, J. K.; Piccirilli, J. A. Acc. Chem. Res. 2011, 44 (12), 1257−1269. (36) Eguaogie, O.; Cooke, L. A.; Martin, P. M.; Ravalico, F.; Conway, L. P.; Hodgson, D. R.; Law, C. J.; Vyle, J. S. Org. Biomol. Chem. 2016, 14 (4), 1201−1205. (37) Patel, B. K.; Eckstein, F. Tetrahedron Lett. 1997, 38 (6), 1021− 1024. (38) Shah, R.; Maize, K. M.; Zhou, X.; Finzel, B. C.; Wagner, C. R. Biochemistry 2017, 56 (28), 3559−3570. (39) Kadokura, M.; Wada, T.; Urashima, C.; Sekine, M. Tetrahedron Lett. 1997, 38 (48), 8359−8362. (40) Kim, I.-H.; Shin, S.; Jeong, Y.-J.; Hah, S. S. Tetrahedron Lett. 2010, 51 (26), 3446−3448. (41) McGee, D. P. C.; Martin, J. C. Can. J. Chem. 1986, 64 (9), 1885− 1889. (42) Zhang, B.; Cui, Z.; Sun, L. Org. Lett. 2001, 3 (2), 275−278. (43) Mukaiyama, T.; Hashimoto, M. Bull. Chem. Soc. Jpn. 1971, 44 (8), 2284−2284. (44) Strenkowska, M.; Wanat, P.; Ziemniak, M.; Jemielity, J.; Kowalska, J. Org. Lett. 2012, 14 (18), 4782−4785. (45) Kowalska, J.; Lewdorowicz, M.; Darzynkiewicz, E.; Jemielity, J. Tetrahedron Lett. 2007, 48 (31), 5475−5479. (46) Brear, P.; Freeman, G. R.; Shankey, M. C.; Trmcic, M.; Hodgson, D. R. W. Chem. Commun. 2009, 33, 4980−4981. (47) Darzynkiewicz, E.; Stepinski, J.; Tahara, S. M.; Stolarski, R.; Ekiel, I.; Haber, D.; Neuvonen, K.; Lehikoinen, P.; Labadi, I.; Lönnberg, H. Nucleosides Nucleotides 1990, 9 (4), 599−618.

(48) Mikkola, S.; Salomaki, S.; Zhang, Z.; Maki, E.; Lonnberg, H. Curr. Org. Chem. 2005, 9 (10), 999−1022. (49) Conway, J. W.; McLaughlin, C. K.; Castor, K. J.; Sleiman, H. Chem. Commun. 2013, 49 (12), 1172−1174. (50) Jackson, R.; Hellen, C.; Pestova, T. Nat. Rev. Mol. Cell Biol. 2010, 11 (2), 113−127. (51) Culjkovic, B.; Topisirovic, I.; Borden, K. Cell Cycle 2007, 6 (1), 65−69. (52) Niedzwiecka, A.; Marcotrigiano, J.; Stepinski, J.; JankowskaAnyszka, M.; Wyslouch-Cieszynska, A.; Dadlez, M.; Gingras, A.-C.; Mak, P.; Darzynkiewicz, E.; Sonenberg, N.; Burley, S. K.; Stolarski, R. J. Mol. Biol. 2002, 319 (3), 615−635. (53) Wypijewska, A.; Bojarska, E.; Lukaszewicz, M.; Stepinski, J.; Jemielity, J.; Davis, R. E.; Darzynkiewicz, E. Biochemistry 2012, 51 (40), 8003−8013. (54) Cai, A.; Jankowska-Anyszka, M.; Centers, A.; Chlebicka, L.; Stepinski, J.; Stolarski, R.; Darzynkiewicz, E.; Rhoads, R. E. Biochemistry 1999, 38 (26), 8538−8547. (55) Liu, S. W.; Rajagopal, V.; Patel, S. S.; Kiledjian, M. J. Biol. Chem. 2008, 283 (24), 16427−16436. (56) Gu, M. G.; Fabrega, C.; Liu, S. W.; Liu, H. D.; Kiledjian, M.; Lima, C. D. Mol. Cell 2004, 14 (1), 67−80. (57) Chen, N.; Walsh, M. A.; Liu, Y.; Parker, R.; Song, H. J. Mol. Biol. 2005, 347 (4), 707−718. (58) Neu, A.; Neu, U.; Fuchs, A. L.; Schlager, B.; Sprangers, R. Nat. Chem. Biol. 2015, 11 (9), 697−704. (59) Pentikainen, U.; Pentikainen, O. T.; Mulholland, A. J. Proteins: Struct., Funct., Genet. 2008, 70 (2), 498−508. (60) Steiger, M.; Carr-Schmid, A.; Schwartz, D. C.; Kiledjian, M.; Parker, R. RNA 2003, 9 (2), 231−238. (61) Coleman, T. M.; Wang, G.; Huang, F. Nucleic Acids Res. 2004, 32 (1), e14. (62) Rydzik, A. M.; Lukaszewicz, M.; Zuberek, J.; Kowalska, J.; Darzynkiewicz, Z. M.; Darzynkiewicz, E.; Jemielity, J. Org. Biomol. Chem. 2009, 7 (22), 4763−4776. (63) Warminski, M.; Sikorski, P. J.; Warminska, Z.; Lukaszewicz, M.; Kropiwnicka, A.; Zuberek, J.; Darzynkiewicz, E.; Kowalska, J.; Jemielity, J. Bioconjugate Chem. 2017, 28 (7), 1978−1992. (64) Zochowska, M.; Piguet, A. C.; Jemielity, J.; Kowalska, J.; Szolajska, E.; Dufour, J. F.; Chroboczek, J. Nanomedicine 2015, 11 (1), 67−76. (65) Neamtu, I.; Rusu, A. G.; Diaconu, A.; Nita, L. E.; Chiriac, A. P. Drug Delivery 2017, 24 (1), 539−557. (66) Gerson, T.; Makarov, E.; Senanayake, T. H.; Gorantla, S.; Poluektova, L. Y.; Vinogradov, S. V. Nanomedicine 2014, 10 (1), 177− 185. (67) Meier, C. Antiviral Chem. Chemother. 2017, 25 (3), 69−82. (68) Gollnest, T.; de Oliveira, T. D.; Schols, D.; Balzarini, J.; Meier, C. Nat. Commun. 2015, 6, 8716. (69) Zhang, L.; Sun, L.; Cui, Z.; Gottlieb, R. L.; Zhang, B. Bioconjugate Chem. 2001, 12 (6), 939−948. (70) Grudzien-Nogalska, E.; Stepinski, J.; Jemielity, J.; Zuberek, J.; Stolarski, R.; Rhoads, R. E.; Darzynkiewicz, E., Synthesis of anti-reverse cap analogs (ARCAs) and their applications in mRNA translation and stability. In Translation Initiation: Cell Biology, High-Throughput Methods, and Chemical-Based Approaches; Lorsch, J., Ed.; Academic Press, 2007; Vol. 431, pp 203−227. (71) Pasquinelli, A. E.; Dahlberg, J. E.; Lund, E. RNA 1995, 1 (9), 957− 967. (72) Mueller, U.; Förster, R.; Hellmig, M.; Huschmann, F. U.; Kastner, A.; Malecki, P.; Pühringer, S.; Röwer, M.; Sparta, K.; Steffien, M.; Ü hlein, M.; Wilk, P.; Weiss, M. S. Eur. Phys. J. Plus 2015, 130, 141.

5999


Phosphorothiolate Dinucleotide Cap Analogues - ACS Publications

Recommend Documents