Protected 5-(hydroxymethyl)uracil nucleotides bearing visible-light photocleavable groups as building blocks for polymerase synthesis of photocaged DNA†

Soňa Boháčová ^a, Lucie Ludvíková ^b, Lenka Poštová Slavětínská ^a, Zuzana Vaníková ^ac, Petr Klán ^b and Michal Hocek *^ac
aInstitute of Organic Chemistry and Biochemistry, Czech Academy of Sciences, Flemingovo namesti 2, CZ-16610 Prague 6, Czech Republic. E-mail: hocek@uochb.cas.cz
^bDepartment of Chemistry and RECETOX, Faculty of Science, Masaryk University, Kamenice 5, CZ-625 00 Brno, Czech Republic
^cDepartment of Organic Chemistry, Faculty of Science, Charles University in Prague, Hlavova 8, Prague 2, CZ-12843, Czech Republic

First published on 2nd February 2018

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Introduction

Photocleavable protecting groups¹ are gaining growing attention due to their applications in organic synthesis, chemical biology and many other areas. In biomolecules, photocaging^1,2 and cleavage of the photoremovable protective groups by light, which is both biocompatible and bioorthogonal, is often used for triggering or switching biological processes. Photocaging of nucleic acids³ is a particularly active field. The photocaging on the nucleobase (mostly at N3 or exocyclic O⁴/N⁴ atoms of pyrimidines) is used⁴ to prevent the base-pairing of oligonucleotides which only after photodeprotection can hybridize with the complementary strand. On the other hand, photocaging of the phosphate backbone⁵ increases the resistance to hydrolysis and influences the interactions with other biomolecules. Photochemical deprotection can therefore either trigger or block biological processes, i.e., primer extension, transcription, antisense effect, RNA interference, translation, etc. It is important to mention that all these types of caged oligonucleotides and nucleic acids can only be synthesized chemically because the corresponding caged (deoxy)nucleoside triphosphates (NTPs or dNTPs) are not suitable substrates for polymerases.^3–5 On the other hand, photocaging in the major-groove site of nucleotides and nucleic acids has been studied less frequently. Photocaged 5-(aminopropargyl)-pyrimidine dNTPs were used for polymerase synthesis of DNA and photochemical release of the reactive amino functions in the major groove.⁶ Nitrobenzyl- (NB), and -phenylethyl-caged 5-(hydroxymethyl)pyrimidine or 7-hydroxymethyl-7-deazapurine 2′-deoxyribonucleoside triphosphates (dNTPs) were used as reversible terminators of primer extension in sequencing.⁷ On the other hand, 6-O-(2-nitrobenzyl)guanosine and 4-O-(2-nitrobenzyl)uridine triphosphates⁸ were not substrates for RNA polymerases until photochemical decaging.

5-(Hydroxymethyl)uracil (5hmU) is a rare DNA base previously found in bacteriophages,⁹ microorganisms,¹⁰ and also eukarytotic genomes.¹¹ The Schultz group has even evolved¹² a bacterium which replaced thymines with 5hmU in genome showing that this base is biocompatible. Unlike the related 5-(hydroxymethyl)cytosine (5hmC),¹³ which is a known epigenetic mark and intermediate in active demethylation of DNA,¹⁴ the biological role of 5hmU is not known so far.¹⁵ In order to create an artificial epigenetic system for the regulation or switching of gene expression, we have systematically studied¹⁶ the influence of chemical modifications in the major groove of DNA on bacterial transcription and found that some small modifications are tolerated by the bacterial RNA polymerase and transcription factor and the modified DNA still serves as a template for transcription. Obviously, any bulkier modifications in the major groove inhibited the transcription.¹⁶ Recently we found¹⁷ that the presence of 5hmU or 5hmC in template DNA can even enhance (up to 3.5 times) or inhibit bacterial transcription (depending on the promoter) and thus even hmU could be an epigenetic mark.

Previously, we have also studied the influence of major-groove modifications on recognition and cleavage of DNA by restriction endonucleases (REs) and found that some small modifications at T/U¹⁸ or A¹⁹ are tolerated (and the REs cleave the modified DNA), whereas all bulky modifications or any modifications at G²⁰ or C¹⁸ bases inhibit the cleavage. We also developed²¹ transient chemical protection of DNA from RE cleavage by polymerase incorporation of bulky 7-(triethylsilylethynyl)-7-deazaadenine which, after desilylation, released 7-ethynyl-7-deazaadenine-modified DNA being fully cleavable by several REs. In order to develop a more biocompatible and bioorthogonal switch, we recently reported²² the use of 5-[(2-nitrobenzyl)oxymethyl]uracil dNTP as a substrate for DNA polymerases leading to the enzymatic synthesis of DNA photocaged in the major-groove. The presence of the bulky NB group in the major groove blocked specific interactions and cleavage by restriction endonucleases (REs) but, after photodeprotection by irradiation using UV light (365 nm), the 5-hydroxymethyluracil-containing DNA was recognized and cut by REs. It was a good proof of the principle, but for any in cellulo or in vivo applications, the use of UV light is not desirable^1b because of its toxicity and limited penetration through tissues. Therefore, it is desirable to develop alternative photocaging groups for 5hmU which would be compatible with the synthesis of modified dNTPs, polymerase incorporation and would be cleavable by visible light. Major challenges in the design and development of new photocaging groups for nucleoside triphosphates are (i) limited chemical stability of some groups (e.g. carbonates) during the triphosphorylation steps and (ii) insufficient substrate activities of some very bulky dNTPs for polymerase incorporations to DNA (in particular by PCR). For example, the nitrophenethyl-caged dNTPs were incorporated by polymerases but terminated the primer extension.⁷ Here we report on the synthesis of 5hmU dNTPs protected with 6-nitropiperonyl (NP)²³ and anthryl-9-methyl (An)²⁴ groups and comparison of substrate activities and photocleavage reactions with parent NB-caged nucleotides.

Results and discussion

Synthesis

Although there were some reports on selective modification of hydroxymethyl groups in 5-hydroxymethyl-2′-deoxyuridine,²⁵ for the desired ether formation in the introduction of the photocaging groups, we chose an opposite approach based on nucleophilic substitution of a suitably protected 5-bromomethyl-2′-deoxyuridine derivative (Scheme 1). In analogy to the previous related studies,^7,22,26 we started from 3′,5′-bis-O-(TBDMS)-3-Boc-protected 5-bromomethyl-2′-deoxyuridine 1 and performed substitution reactions with the corresponding benzyl-type alcohol in the presence of 2,2,6,6-tetramethylpiperidine in CH₃CN at 50 °C. The addition of AgOTf was important for the precipitation of AgBr and thus shifting the equilibrium towards products. The desired fully protected caged nucleosides 2a and 2b were isolated in moderate yields (10 and 21%). The An-caged intermediate 2c could not be isolated in pure form and was used in the next step in the crude form. Subsequently, the BOC group was removed using sodium bicarbonate²⁷ in MeOH at 50 °C, whereas the common reagents such as CF₃COOH or TBAF failed. The TBDMS group was then removed using Et₃N·3HF²⁸ in THF. The desired photocaged nucleosides dU^NB and dU^NP were isolated in good yields of 64 and 57%, whereas the dU^An was isolated in low 9% overall yield.

Phosphorylation reactions were carried out with POCl₃ in trimethyl phosphate at 0 °C. Quenching of the reaction with 2 M TEAB gave the desired monophosphates dU^NBMP, dU^NPMP and dU^AnMP in moderate yields after isolation by HPLC. To form dNTPs, phosphorylation was carried out in the same way followed by the addition of pyrophosphate and tributyl amine in DMF. The targets dU^NBTP, dU^NPTP and dU^AnTP were obtained in moderate yields after isolation by HPLC. The uncaged 5-hydroxymethyl-2′-deoxyuridine triphosphate dU^hmTP was prepared in 27% yield by the photolysis of dU^NPTP (with 59% recovery of the starting material) in water using irradiation by UV LEDs at either 375 or 385 nm followed by the separation on HPLC (Scheme 2).

Photophysical properties of modified monophosphates and quantum yields of uncaging reactions

Initially, we recorded the absorption spectra of the model caged monophosphates dU^RMP in water and calculated the molar absorption coefficients for selected wavelengths in the range of 355–437 nm (Table 1 and Table S1 and Fig. S1–S7 in the ESI†). The parent dU^NBMP exerted an absorption maximum at 264 nm and showed a very low absorption above 300 nm. At 365 nm (under which the NB group is cleaved²²), the molar absorption coefficient is only 238 M⁻¹ cm⁻¹ (Table 1, Table S1 and Fig. S3 in the ESI†). On the other hand, the nitropiperonyl-caged nucleotide dU^NPMP has a strong absorption maximum at 358 nm and still a relatively significant absorption at 400 nm (1940 M⁻¹ cm⁻¹, Table 1 and Fig. S4 in the ESI†). dU^AnMP showed three absorption maxima at 349, 367 and 386 nm as expected for an anthracene moiety. The spectra of dU^NPMP and dU^AnMP (Table S1, Fig. S4 and S6 in the ESI†) indicate that photochemical deprotection can be possible using light of wavelengths up to 425 nm.

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Then we determined the quantum yields (QY) of uncaging. The measurements were carried out at 365 nm in water with a ferrioxalate actinometer²⁹ according to published procedures³⁰ giving an absolute value of QY. The QY of photocleavage of dU^NPMP was moderate with the value of 4.4 ± 0.8%, whereas the QY value for dU^AnMP was negligible (in the range of an experimental error). Our attempts to measure QYs of previously published NB-group-protected dU^NBMP failed because of two factors. (a) The molar absorption coefficient of dU^NBMP at 365 nm was too low and the measurements had to be carried out in too concentrated solutions, and (b) the nitrosobenzene photoproduct formed during the reaction strongly absorbs at 365 nm. The fluorescence quantum yields of modified monophosphates in water were also determined at the excitation wavelength of 365 nm using quinine sulfate dissolved in 0.5 M H₂SO₄ as a standard³¹ giving relative values of QYs. U^AnMP is moderately fluorescent with the quantum yield of 4.48 ± 0.15%; U^NBMP and U^NPMP are not fluorescent.

Incorporation of caged nucleotides into DNA

Caged triphosphates dU^NBTP, dU^NPTP and dU^AnTP as well as the uncaged dU^hmTP were tested as substrates for DNA polymerases.³² At first, the primer extension (PEX) reaction was tested using a 19-mer template (for sequences of ONs see Table S4 in the ESI†) encoding for the incorporation of one modified dU^R followed by three guanines. Fig. 1a shows the successful incorporation of all modified dU^R nucleotides using KOD XL (for successful PEX using Vent (exo-) polymerase, see Fig. S12 in the ESI†). The extended modified oligonucleotide (ON) products were confirmed by the MALDI-TOF analysis showing masses corresponding to U^hm-containing ONs (Fig. S13 in the ESI†) because the laser irradiation using in MALDI cleaves the photoremovable groups. In the case of An modification, both caged and uncaged products were detected (Fig. S14 in the ESI†). We also performed a simple kinetics of single nucleotide extension (Fig. S15 and S16 in the ESI†) showing that the incorporation of modified dU^RTPs was only slightly slower compared to that of TTP. Then, the PEX reaction was performed with a more demanding 31-mer template designed for the incorporation of 4 dU^R modifications (Fig. 1b). This PEX proceeded successfully with dU^hmTP, dU^NBTP and dU^NPTP giving a clean fully extended product. U^ANTP appeared to be a slightly worse substrate for DNA polymerase giving the mixture of extended products also containing truncated ONs. The worse outcome was probably due to the bulkiness and hydrophobicity of the anthracene group.

Next, we tested dU^hmTP, dU^NBTP, dU^NPTP and dU^AnTP in PCR using either a 98-mer or (a more demanding) 297-mer template. In agreement with our previous communications,^17,22 the PCR reactions with dU^hmTP and dU^NBTP, as well as with the new nitropiperonyl-caged dU^NPTP, proceeded but the products of amplification were rather weak. The agarose gel of the 98-mer PCR (Fig. 1c) shows the bands of the modified PCR products only after the concentration of the sample, whereas the longer PCR (Fig. 1d) showed only very weak bands with somewhat faster mobility even after the concentration of the sample. However, in all cases, the identity of the modified amplicons was confirmed by sequencing. The bulky and hydrophobic dU^AnTP did not give any amplification products in either of the PCR reaction.

Photochemical deprotection of caged DNA

In order to study the photocleavage of caged DNA, we used the previously described²² approach utilizing the cleavage of modified DNA by RsaI restriction endonuclease for monitoring the photochemical deprotection of DNA. The caged DNA containing bulky protecting groups is not cleaved by RE, whereas the DNA containing small hydroxymethyluracil modification is fully cleaved (Scheme 3). Therefore, we prepared 30-mer PEX products containing a dU^R modification in the recognition site for RsaI. We tested the cleavage of the DNA by RE before and after irradiation using different LED diodes. In our previous work, we showed²² that the NB-caged DNA can be efficiently liberated by irradiation at 355 nm. Now, we tested the photoremoval of all three caging groups with UV or visible light at different wavelengths (355, 375, 385, 400 and 425 nm). Uncaging experiments at 437 nm and higher wavelengths could not be tested because of the photo-bleaching of the 6-FAM used for labeling and visualization of ONs on gels (Fig. S17†). Considering the negligible absorbances of dU^RMPs at 437 nm (see Table S1 in the ESI† for molar absorption coefficients at 437 nm), we do not expect that liberation would proceed at 437 nm and higher wavelengths.

Fig. 2 shows the results of photochemical deprotection reactions of modified DNA at 375 and 425 nm (b), whereas Table 1 (and Fig. S22–S27 in the ESI†) gives complete data at all tested wavelengths. ImageJ software was used to quantify the ratios of uncaged DNA (Table 1). Because of different optical power of the different LEDs, the reaction times differed; we tested at least 3 different irradiation times to reach (almost) complete conversion when possible. In all cases, the observed photochemical cleavage of the An group was very inefficient (even at 355 nm) and the conversions were very low (up to 15% even with prolonged time). The cleavage of NB and NP groups proceeded with comparable efficiencies at lower wavelengths (355–385 nm) reaching good conversions (92–95%) in reasonable reaction times (15–120 min). Only at higher wavelengths (400 and 425 nm), the nitropiperonyl group was cleaved significantly faster. At the highest wavelength of 425 nm, the cleavage of the nitrobenzyl group did not reach full conversion, whereas the cleavage of the NP group was still almost quantitative. Taking into account the almost negligible absorbance of the nitrobenzyl group at 400 and 425 nm, it was rather surprising that the NB group was still at least partly cleavable. However, as expected, uncaging of NPs was much faster and more efficient at these wavelengths, and we conclude that this group can be removed with visible light (Table 2).

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Conclusions

We prepared three types of photocaged derivatives of 2′-deoxy-5-(hydroxymethyl)uridine nucleoside, mono- and triphosphates protected by 2-nitrobenzyl-, 6-nitropiperonyl- and anthryl-9-methyl groups, and tested their enzymatic incorporations to DNA and photochemical deprotections. The NB- and NP-caged dNTPs (dU^NBTP and dU^NPTP) were good substrates for DNA polymerases and worked in both PEX and PCR reactions. The An-linked dU^AnTP was a less efficient substrate but it was still possible to incorporate it to at least some sequences using PEX. The liberation of photocaged DNA was tested by monitoring the cleavage of DNA with REs. We show that both NB- and NP-caged DNA can be efficiently deprotected by UV irradiation, whereas the nitropiperonyl group is suitable for uncaging with visible light at 400 or 425 nm. Therefore, the NP-caging has a better potential for in cellulo or in vivo photocaging of 5-(hydroxymethyl)uracil in DNA. The An group does not apparently have much potential in DNA photocaging but still the An-substituted dNTP can find some use in polymerase synthesis of DNA bearing a fluorescent extended aromatic system for applications in imaging or selection of hydrophobic aptamers.

Experimental

Full experimental part is given in the ESI.† Selected typical experiments are given below.

3′,5′-Bis-O-(tert-butyldimethylsilyl)-3-tert-butoxycarbonyl-5-(6-nitropiperonyloxy)methyl-2′-deoxyuridine (2b)

6-Nitropiperonyl alcohol (240 mg, 1.22 mmol) and silver triflate (308 mg, 1.20 mmol) were suspended in dry acetonitrile and 2,2,6,6-tetramethylpiperidine (205 μL, 1.21 mmol) was added. The mixture was stirred for 10 min at 50 °C. Then solution of 1 in acetonitrile was added and the reaction was stirred for another 2 h at 50 °C. The solvent was removed under reduced pressure and the mixture was separated by chromatography (hexane:ethyl acetate 5:1) to give yellow oil (97 mg, 21%). ¹H NMR (500 MHz, DMSO-d₆): 0.03, 0.04, 0.078 and 0.081 (4 × s, 4 × 3H, CH₃Si); 0.84 and 0.87 (2 × s, 2 × 9H, (CH₃)₃C); 1.51 (s, 9H, (CH₃)₃COCO); 2.18 (ddd, 1H, J_gem = 13.5 Hz, J_2′a,1′ = 6.2 Hz, J_2′a,3′ = 3.2 Hz, H-2′a); 2.31 (ddd, 1H, J_gem = 13.5 Hz, J_2′b,1′ = 7.3 Hz, J_2′b,3′ = 5.9 Hz, H-2′b); 3.70 (dd, 1H, J_gem = 11.4 Hz, J_5′a,4′ = 4.4 Hz, H-5′a); 3.74 (dd, 1H, J_gem = 11.4 Hz, J_5′b,4′ = 4.6 Hz, H-5′b); 3.84 (td, 1H, J_4′,5′a = J_4′,5′b = 4.5 Hz, J_4′,3′ = 3.0 Hz, H-4′); 4.26 (s, 2H, B-CH₂O); 4.35 (dt, 1H, J_3′,2′b = 5.8 Hz, J_3′,4′ = J_3′,2′a = 3.1 Hz, H-3′); 4.80 (s, 2H, OCH_2⁻Ph); 6.10 (dd, 1H, J_1′,2′b = 7.3 Hz, J_1′,2′a = 6.2 Hz, H-1′); 6.24 (s, 2H, OCH₂O); 7.26 (s, 1H, H-6′′); 7.70 (s, 1H, H-3′′); 7.79 (s, 1H, H-6). ¹³C NMR (125.7 MHz, DMSO-d₆): −5.31, −5.28, −4.73 and −4.56 (CH₃Si); 17.94 and 18.23 ((CH₃)₃C); 25.89 and 25.98 ((CH₃)₃C); 27.24 ((CH₃)₃COCO); 39.5 (CH₂-2′); 62.78 (CH₂-5′); 65.02 (B-CH₂O); 68.79 (OCH_2⁻Ph); 72.11 (CH-3′); 85.56 (CH-1′); 86.71 ((CH₃)₃COCO); 87.46 (CH-4′); 103.69 (OCH₂O); 105.51 (CH-3′′); 107.36 (CH-6′′); 110.03 (C-5); 132.28 (CH-1′′); 139.86 (CH-6); 141.05 (C-2′′); 147.05 (C-4′′); 147.82 ((CH₃)₃COCO); 148.06 (C-2); 152.49 (C-5′′); 159.98 (C-4). MS (ESI): m/z (%):789 (50)[M + Na + H]²⁺, 788 (100) [M + Na]⁺; HRMS (ESI): m/z calcd for C₃₅H₅₅N₃O₁₂NaSi₂: 788.32195 [M + Na]⁺; found: 788.32165.

3′,5′-Bis-O-(tert-butyldimethylsilyl)-5-(6-nitropiperonyloxy)methyl-2′-deoxyuridine (3b)

Compound 2b (131 mg, 0.17 mmol) was dissolved in methanol, NaHCO₃ (144 mg, 1.70 mmol) was added and the suspension was stirred at 56 °C overnight. The solvent was removed under vacuum and the residue was extracted with ethyl acetate and purified by column chromatography (hexane:EtOAc 3:2) to give 3b as yellow powder (81 mg, 71%). ¹H NMR (500 MHz, DMSO-d₆): 0.039, 0.044, 0.080 and 0.082 (4 × s, 4 × 3H, CH₃Si); 0.85 and 0.87 (2 × s, 2 × 9H, (CH₃)₃C); 2.12 (ddd, 1H, J_gem = 13.4 Hz, J_2′a,1′ = 6.2 Hz, J_2′a,3′ = 3.1 Hz, H-2′a); 2.21 (ddd, 1H, J_gem = 13.4 Hz, J_2′b,1′ = 7.8 Hz, J_2′b,3′ = 6.0 Hz, H-2′b); 3.69 (dd, 1H, J_gem = 11.3 Hz, J_5′a,4′ = 4.3 Hz, H-5′a); 3.74 (dd, 1H, J_gem = 11.3 Hz, J_5′b,4′ = 4.6 Hz, H-5′b); 3.80 (td, 1H, J_4′,5′a = J_4′,5′b = 4.5 Hz, J_4′,3′ = 2.9 Hz, H-4′); 4.20 and 4.23 (2 × dd, 2 × 1H, J_gem = 11.6 Hz, J_CH2,CH2 = 0.7 Hz, B-CH₂O); 4.35 (dt, 1H, J_3′,2′b = 5.9 Hz, J_3′,4′ = J_3′,2′a = 3.0 Hz, H-3′); 4.78 (d, 2H, J_CH2,CH2 = 0.7 Hz, OCH_2⁻Ph); 6.14 (dd, 1H, J_1′,2′b = 7.8 Hz, J_1′,2′a = 6.2 Hz, H-1′); 6.23 (s, 2H, OCH₂O); 7.28 (s, 1H, H-6′′); 7.65 (s, 1H, H-6); 7.68 (s, 1H, H-3′′); 11.50 (bs, 1H, NH). ¹³C NMR (125.7 MHz, DMSO-d₆): −5.35, −5.34, −4.75 and −4.59 (CH₃Si); 17.91 and 18.18 ((CH₃)₃C); 25.86 and 25.94 ((CH₃)₃C); 39.42 (CH₂-2′); 62.90 (CH₂-5′); 65.15 (B-CH₂O); 68.57 (OCH_2⁻Ph); 72.25 (CH-3′); 84.38 (CH-1′); 87.03 (CH-4′); 103.59 (OCH₂O); 105.39 (CH-3′′); 107.41 (CH-6′′); 110.55 (C-5); 132.43 (CH-1′′); 139.32 (CH-6); 141.02 (C-2′′); 146.95 (C-4′′); 150.61 (C-2); 152.40 (C-5′′); 163.13 (C-4). MS (ESI): m/z (%): 688 (100) [M + Na]⁺, 689 (42)[M + Na + 2H]³⁺, 1353 (22) [2M + Na]⁺; HRMS (ESI): m/z calcd for C₃₀H₄₈N₃O₁₀Si₂: 666.28748 [M + H]⁺; found: 666.28727.

5-(6-Nitropiperonyloxy)methyl-2′-deoxyuridine (dU^NP)

Compound 3a (80 mg, 0.12 mmol) was dissolved in dry THF, triethylamine trihydrofluoride (171 μL, 1.05 mmol) was added and the reaction was stirred overnight at rt. The solvent was removed under reduced pressure and the mixture was separated by chromatography (DCM:MeOH 7:1) to give dU^NP as yellow powder (42 mg, 80%). ¹H NMR (500 MHz, DMSO-d₆): 2.05–2.15 (m, 2H, H-2′); 3.55 (ddd, 1H, J_gem = 11.8 Hz, J_5′a,OH = 5.1 Hz, J_5′a,4′ = 3.9 Hz, H-5′a); 3.59 (ddd, 1H, J_gem = 11.9 Hz, J_5′b,OH = 5.4 Hz, J_5′b,4′ = 3.9 Hz, H-5′b); 3.79 (td, 1H, J_4′,5′a = J_4′,5′b = 3.9 Hz, J_4′,3′ = 2.9 Hz, H-4′); 4.20 and 4.23 (2 × dd, 2 × 1H, J_gem = 11.6 Hz, J_CH2,CH2 = 0.8 Hz, B-CH₂O); 4.24 (m, 1H, H-3′); 4.79 (d, 2H, J_CH2,CH2 = 0.8 Hz, OCH_2⁻Ph); 5.02 (t, 1H, J_OH,5′a = J_OH,5′b = 5.3 Hz, OH-5′); 5.25 (d, 1H, J_OH,3′ = 4.3 Hz, OH-3′); 6.16 (t, 1H, J_1′,2′b = J_1′,2′a = 6.8 Hz, H-1′); 6.23 (s, 2H, OCH₂O); 7.33 (s, 1H, H-6′′); 7.68 (s, 1H, H-3′′); 7.99 (s, 1H, H-6); 11.44 (bs, 1H, NH). ¹³C NMR (125.7 MHz, DMSO-d₆): 39.85 (CH₂-2′); 61.49 (CH₂-5′); 65.28 (B-CH₂O); 68.52 (OCH_2⁻Ph); 70.64 (CH-3′); 84.41 (CH-1′); 87.67 (CH-4′); 103.57 (OCH₂O); 105.32 (CH-3′′); 107.41 (CH-6′′); 110.34 (C-5); 132.75 (CH-1′′); 139.82 (CH-6); 140.92 (C-2′′); 146.90 (C-4′′); 150.49 (C-2); 152.50 (C-5′′); 163.01 (C-4). MS (ESI): m/z (%): 460 (100) [M + Na]⁺, 461 (21)[M + Na + H]²⁺, 897 (39)[2M + Na]⁺, 898 (16) [2M + Na + H]²⁺; HRMS (ESI): m/z calcd for C₁₈H₂₀N₃O₁₀: 438.11432 [M + Na]⁺; found: 438.11433.

5-(6-Nitropiperonyloxy)methyl-2′-deoxyuridine monophosphate (dU^NPMP)

Nucleoside dU^NP (22 mg, 0.05 mmol) and proton sponge (22 mg, 0.10 mmol) were suspended in trimethyl phosphate (150 μL) in an argon purged flask and the suspension was cooled to 0 °C. Then, redistilled POCl₃ (5 μL, 0.06 mmol) was added. The reaction mixture was stirred at 0 °C for 45 min and then aqueous solution of TEAB (2 M, 2 mL) was added and the mixture was evaporated under reduced pressure. The product was purified by HPLC chromatography (0.1 M aq. TEAB 0 → 50% methanol), co-evaporated several times with water and converted to sodium salt on Dowex. Monophosphate dU^NPMP (6 mg, 21%) was obtained as a yellowish lyophilizate (water). ¹H NMR (500 MHz, D₂O): 2.35 (ddd, 1H, J_gem = 14.1 Hz, J_2′a,1′ = 6.5 Hz, J_2′a,3′ = 4.0 Hz, H-2′a); 2.39 (ddd, 1H, J_gem = 14.1 Hz, J_2′b,1′ = 7.3 Hz, J_2′b,3′ = 5.9 Hz, H-2′b); 3.91–3.97 (m, 2H, H-5′); 4.13 (m, 1H, H-4′); 4.44 (s, 2H, B-CH₂O); 4.56 (dt, 1H, J_3′,2′b = 5.8 Hz, J_3′,2′a = J_3′,4′ = 4.0 Hz, H-3′); 4.87 and 4.91 (2 × d, 2 × 1H, J_gem = 14.0 Hz, OCH_2⁻Ph); 6.15 (m, 2H, OCH₂O); 6.27 (bt, 1H, J_1′,2′b = J_1′,2′a = 6.9 Hz, H-1′); 7.15 (s, 1H, H-6′′); 7.60 (s, 1H, H-3′′); 8.01 (s, 1H, H-6). ¹³C NMR (125.7 MHz, D₂O): 39.13 (CH₂-2′); 64.21 (d, J_C,P = 4.6 Hz, CH₂-5′); 66.25 (B-CH₂O); 69.71 (OCH_2⁻Ph); 71.75 (CH-3′); 85.91 (CH-1′); 86.74 (d, J_C,P = 8.3 Hz, CH-4′); 104.08 (OCH₂O); 106.27 (CH-3′′); 109.67 (CH-6′′); 111.65 (C-5); 132.13 (CH-1′′); 142.10 (C-2′′); 142.33 (CH-6); 147.88 (C-4′′); 152.74 (C-2); 153.00 (C-5′′); 166.72 (C-4). ³¹P NMR (202.4 MHz, D₂O): 4.55 (s, 1P, P-5′). MS (ESI): m/z (%): 516 (100) [M]⁻, 517 (23)[M + H]; HRMS (ESI): m/z calcd for C₁₈H₁₉N₃O₁₃P: 516.06610 [M]⁻; found: 516.06531.

5-(6-Nitropiperonyloxy)methyl-2′-deoxyuridine triphosphate (dU^NPTP)

Nucleoside dU^NP (40 mg, 0.09 mmol) and proton sponge (40 mg, 0.18 mmol) were suspended in trimethyl phosphate (270 μL) in an argon purged flask and the suspension was cooled to 0 °C. Then, redistilled POCl₃ (9 μL, 0.10 mmol) was added. The reaction mixture was stirred at 0 °C for 45 min and then ice-cold solution of (NHBu₃)₂H₂P₂O₇ (208 mg, 0.46 mmol) and tributyl amine (90 μL, 0.38 mmol) in anhydrous DMF (4 mL) was added. The reaction mixture was stirred at 0 °C for another 1 h. Then aqueous solution of TEAB (2 M, 2 mL, 4 mmol) was added and the mixture was evaporated under reduced pressure. The residue was co-evaporated several times with water. The product was purified by HPLC chromatography (0.1 M aq. TEAB 0 → 50% methanol), co-evaporated several times with water and converted to sodium salt on Dowex. Triphosphate dU^NPTP (17 mg, 24%) was obtained as a yellowish lyophilizate (water). ¹H NMR (500 MHz, D₂O): 2.34–2.40 (m, 2H, H-2′); 4.18 (qd, 1H, J_4′,5′a = J_4′,5′b = J_4′,3′ = 3.5 Hz, J_4′,P = 2.0 Hz, H-4′); 4.22 (ddd, 1H, J_gem = 11.6 Hz, J_5′a,P = 5.1 Hz, J_r = 3.2 Hz, H-5′a); 4.26 (ddd, 1H, J_gem = 11.6 Hz, J_5′b,P = 6.1 Hz, J_5′b,4′ = 3.6 Hz, H-5′b); 4.40 and 4.47 (2 × d, 2 × 1H, J_gem = 12.1 Hz, B-CH₂O); 4.66 (td, 1H, J_3′,2′a = J_3′,2′b = 5.4 Hz, J_3′,4′ = 3.7 Hz, H-3′); 4.84 and 4.92 (2 × d, 2 × 1H, J_gem = 14.0 Hz, OCH_2⁻Ph); 6.15 (t, 2H, J_CH2,LR = 1.3 Hz, OCH₂O); 6.26 (t, 1H, J_1′,2′b = J_1′,2′a = 6.7 Hz, H-1′); 7.13 (s, 1H, H-6′′); 7.58 (s, 1H, H-3′′); 7.96 (s, 1H, H-6). ¹³C NMR (125.7 MHz, D₂O): 39.31 (CH₂-2′); 65.65 (d, J_C,P = 5.5 Hz, CH₂-5′); 66.29 (B-CH₂O); 69.77 (OCH_2⁻Ph); 70.73 (CH-3′); 85.91 (CH-1′); 86.20 (d, J_C,P = 9.1 Hz, CH-4′); 104.11 (OCH₂O); 106.27 (CH-3′′); 109.84 (CH-6′′); 111.55 (C-5); 132.11 (CH-1′′); 142.08 (CH-6); 142.15 (C-2′′); 147.90 (C-4′′); 152.47 (C-2); 152.94 (C-5′′); 166.06 (C-4). ³¹P NMR (202.4 MHz, D₂O): −20.95 (t, 1P, J_β,α = J_β,γ = 19.8 Hz, P_β); −10.57 (d, 1P, J_α,β = 19.5 Hz, P_α); −5.11 (bd, 1P, J_γ,_β = 20.0 Hz, P_γ). MS (ESI): m/z (%): 516 (41) [M + H-HP₂O₆]⁻, 596 (100) [M + 2H-HPO₃]⁻, 618 (60) [M + H + Na-HPO₃]⁻; HRMS (ESI): m/z calcd for C₁₈H₂₀N₃O₁₉P₃Na: 697.98070 [M + H + Na]⁻; found: 697.97937.

PEX – multiple modifications

The reaction mixture (20 μL) contained KOD XL DNA polymerase (2.5 U μL⁻¹, 0.02 μL), mixture of dATP, dCTP and dGTP (1 mM, 1 μL), dTTP (1 mM, 1 μL) or modified dU^RTP (1 mM, 2 μL), 6-FAM labelled primer 248sh (3 μM, 1 μL), 31-mer template TINA-prb4basII (3 μM, 1.5 μL) and 10× buffer for KOD XL DNA polymerase (2 μL) supplied by the manufacturer. Reaction mixtures were incubated for 30 min at 60 °C in a thermal cycler. After the reaction samples were denatured by the addition of stop solution (20 μL, 80% [v/v] formamide, 20 mM EDTA, 0.025%, [w/v] bromophenol blue, 0.025% [w/v] xylene cyanol, PCR water) heating was followed for 5 min at 95 °C. The reaction mixtures were separated using 12.5% denaturing PAGE.

Uncaging of DNA

The reaction mixture (20 μL) contained KOD XL DNA polymerase (2.5 U μL⁻¹, 0.2 μL), mixture of dATP, dCTP and dGTP (1 mM, 0.2 μL), dTTP or modified dUTP (1 mM, 0.3 μL) 6-FAM labelled primer 248sh (3 μM, 1 μL) and 30-mer template RsT (3 μM, 1.5 μL) and 10× buffer for KOD XL DNA polymerase (2 μL) supplied by the manufacturer. Reaction mixtures were incubated for 30 min at 60 °C in a thermal cycler.

After that, the reaction samples were either:

(a) denatured by the addition of the stop solution (23 μL, 80% [v/v] formamide, 20 mM EDTA, 0.025%, [w/v] bromophenol blue, 0.025% [w/v] xylene cyanol, PCR water) and water (3 μL) and heated at 95 °C for 5 min,

(b) incubated with RSaI (0.8 μL) in CutSmart buffer (2.2 μL) and denatured by addition of the stop solution (23 μL) and heated at 95 °C for 5 min, or

(c) irradiated with a UV LED followed by incubation with RSaI (0.8 μL) in CutSmart buffer (2.2 μL) and denatured by the addition of the stop solution (23 μL) and heating at 95 °C for 5 min.

Reaction mixtures were separated using 12.5% denaturing PAGE.

Conflicts of interest

There are no conflicts to declare.

Acknowledgements

This work was supported by the Czech Academy of Sciences (Praemium Academiae award to M. H.) and by the Czech Science Foundation (17-03419S to S. B., Z. V. and M. H.; GA13-25775S to P. K.). The RECETOX research infrastructure was supported by the projects of the Czech Ministry of Education (LO1214 and LM2011028; P. K.).

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Footnote

† Electronic supplementary information (ESI) available. See DOI: 10.1039/c8ob00160j

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