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. 2021 Nov 17;6(47):31595–31604. doi: 10.1021/acsomega.1c04074

Changes of C≡C Triple Bond Vibration that Disclosed Non-Canonical Cytosine Protonation in i-Motif-Forming Oligodeoxynucleotides

Ryota Itaya †, Wakana Idei †, Takashi Nakamura ‡, Tatsuya Nishihara †, Ryohsuke Kurihara §, Akimitsu Okamoto ∥, Kazuhito Tanabe †,*
PMCID: PMC8637604  PMID: 34869984

Abstract

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Non-canonical protonation at cytosine (C) in DNA is related to a formation of second order DNA structures such as i-motif, which has a role in gene regulation. Although the detailed structural information is indispensable for comprehension of their functions in cells, the protonation status of C in complicated environments is still elusive. To provide a reporter system of non-canonical protonation, we focused on the molecular vibration that could be monitored using the Raman spectroscopy. We prepared a cytosine derivative (PC) with an acetylene unit as a Raman tag, and found that the Raman signal of acetylene in PC in oligodeoxynucleotides (ODNs) changed due to protonation at the cytosine ring which shortened an acetylene bond. The signal change in i-motif-forming ODNs was also observed in crowded environments with polyethylene glycol, evidencing protonation in i-motif DNA in complicated environments. This system would be one of tracking tools for protonation in DNA structures.

Introduction

Among the four nucleobases in DNA, cytosine can adopt specific structures with protonation that are related with formation of higher order structures. For example, C+–G–C triplets with one protonated cytosine that interact with a G–C base pair formed triplex, and cytosine·hemiprotonated cytosine base pairs ([C–H–C]+) formed in DNA i-motifs. These non-canonical base pairing outside of Watson and Crick’s rule featuring hemiprotonated cytosine nucleobases can mediate complicated biological functions such as transfer of genetic information, gene manipulation, and regulation.1−7 One of the most attractive second order structures bearing non-canonical protonation is i-motif.8−22 An intercalating hemiprotonated [C–H–C]+ base pair that retains the quadruplex structure even at neutral pH is formed in this structure,23−26 which has a functional role in the gene regulation.

Detailed structural information of specific nucleobases in intracellular nucleic acids and their response to surrounding environments are important to design functional molecules for oligonucleotide therapeutics and comprehension of their functions. However, the protonation state of cytosine in DNA in complicated and messy environments remains elusive. Establishment of procedures to obtain the direct evidence for non-canonical protonation of cytosine under these complicated environments is still an issue, although much is known about the macroscopic characteristics of i-motif folding. Conventionally, spectroscopic approaches using absorption spectra, fluorescence emission of DNA-binding molecules and fluorophore-labeled ODNs, and nuclear magnetic resonance (NMR)8,9,27−29 have been applied to track protonation of cytosine;4 however, these methods have some drawbacks. Absorption spectra provide a sum of absorption bands of all protonated cytosine nucleobases in the strand, but localized protonation states cannot be identified. Modified cytosine with fluorescence properties was reported by Luedtke and co-workers to monitor the protonation.9 Although the emission of the modified cytosine was enhanced upon protonation, the quantum yields of its fluorescence are low and the chromophore should be excited by photoirradiation with short wavelength, and thus, it seems to be difficult to apply them to detect protonation under complicated conditions. In addition, a proton on cytosine was observed as an imino proton by NMR spectroscopic analysis, but the sensitivity of NMR is generally low. In light of these research contexts, we attempted to provide a reporting system of non-canonical protonation at specified cytosine under complicated conditions.

As an alternative implement to identify non-canonical protonation, we focused on molecular vibration as a characteristic that could be monitored using the Raman spectroscopy. Raman spectroscopy is a spectroscopic technique that is widely applied for the determination of vibrational modes of molecules.30 This spectroscopic method is used to obtain a structural fingerprint of the target molecules;31−33 therefore, we expected that protonation, including non-canonical protonation, would affect the vibrational modes of the target nucleobase in ODNs, and that Raman spectroscopy could become a key approach for reliable detection of the protonation state. Indeed, evidence that a specific substituent provides a characteristic band even in composite material also encouraged us to investigate the application of Raman spectroscopy.34−37

Herein, we prepared a modified cytosine derivative (PC, Figure 1) with a phenylacetylene unit as a reporter tag that exhibited a band around 2200 cm–1 in the Raman spectra. Initially, we assessed canonical protonation of PC and confirmed that a pH change resulted in a band shift of the signal from the acetylene unit (ca. 10 cm–1) upon protonation at the conjugated pyrimidine ring. We then applied the system to the identification of non-canonical protonation of PC in i-motif structures. Interestingly, the band shift of the acetylene unit signal of PC in i-motif-forming ODNs occurred around pH 4 and 5, where PC mononucleoside was not protonated. By the measurement of CD spectra, we attributed this signal shift of PC in ODNs to non-canonical protonation at PC. We also obtained evidence that molecular crowding of the cosolutes led to the protonation of PC in the i-motif structure even at circumneutral pH conditions according to the raised pKa. These results suggest that the non-canonical protonation at the nucleobase could be tracked by means of the Raman spectroscopy.

Figure 1.

Figure 1

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Outline of an identification of non-canonical protonation at Raman-tagged cytosine nucleobase in ODNs by Raman spectra. (A) Protonation of PC in ODNs. (B) Conceptual drawing of the signal shift in Raman spectra upon protonation. Non-protonated (black line) or protonated ODNs (blue line). (C) Non-canonical protonation of C–C+ base pair leads to i-motif formation of ODNs bearing PC in the strand under circumneutral or molecular crowding conditions.

Results and Discussion

Initially, we employed a PC mononucleoside (Figure 2) and assessed its molecular vibration.38 We measured the Raman spectra of PC with 532 nm excitation. As shown in Figure 3, PC showed a typical band for the acetylene group around 2220 cm–1 under neutral conditions. When the pH was decreased from 7.2 to 2.1, the characteristic band shift of the acetylene unit signal was observed. While the shift of the band was negligible between a pH of 7.2 and 3.5, the band changed significantly from 2220 to 2230 cm–1 under pH conditions below pH 3.0 (Figure 3C). As a control, we conducted similar experiments using uridine mononucleoside derivative with a phenylacetylene substituent (PU), which did not protonate even under acidic conditions. The PU also showed the typical band at 2226 cm–1, but the band did not change even under acidic conditions. To clarify the reason for the band shift of PC, we measured absorption spectra. Specific absorption bands that changed upon variation of pH were clearly observed, as shown in Figure 4A. At pH 1.6, the protonated form of PC with maximum absorption at 325 nm was observed, whereas only the neutral species with a maximum absorption at 310 nm was present at pH 5.5. The pKa value of PC was estimated as 2.8, which is smaller than that of the native cytosine nucleobase (pKa ca. 4.2),3,21,22,27 probably due to the electron-withdrawing nature of the phenylacetylene substituent. Thus, the protonation characteristics of PC obtained from the change of the absorption spectra are consistent with that obtained from its Raman spectra. We also evaluated the reversibility of the change of the signal of PC. Accordant with the pH effect on the Raman band of PC, the Raman signal of the acetylene unit was observed at 2229 cm–1 in the presence of 50 mM HCl, but was shifted to 2219 cm–1 upon addition of 50 mM NaOH to neutralize the sample solution (Figure 3D). This band shift was repeated upon the re-addition of HCl and NaOH. From these results, we assumed that the band shift of PC in the Raman spectra was attributed to the protonation of PC.

Figure 2.

Figure 2

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Chemical structures of modified nucleobases and sequences of ODNs used in this study. The i-motif-forming sites in ODNs are shown in bold.

Figure 3.

Figure 3

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Raman spectra of PC and PU with excitation at 532 nm. (A) Raman spectra of PC (50 mM) measured under different pH conditions (pH 7.2: black line, pH 5.2: blue line, pH 2.1: red line). (B) Raman spectra of PU (50 mM) measured under different pH conditions (pH 7.2: black line, pH 5.2: blue line, pH 2.1: red line). (C) Signal shift of PC (circle) and PU (triangle) as decreasing pH conditions. Results are shown with the mean ± SD (n = 3). (D) Reversible responses of Raman signal of PC (50 mM) to alternating changes of pH conditions (A: acidic conditions, N: neutralized conditions). The first sample was measured in the presence of 50 mM HCl. Then, 50 mM NaOH was added to neutralize the sample. To repeat the alteration of pH conditions, addition of HCl and NaOH was repeated.

Figure 4.

Figure 4

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(A) Absorption spectra of PC (100 μM) at pH 6.3 (deep red), 5.5 (red), 4.7 (orange), 3.9 (yellow), 2.9 (light green), 2.7 (green), 2.5 (light blue), 2.2 (blue), 2.1 (dark blue) and 1.6 (purple). (B) Concentration profiles of protonated or unprotonated PC. The data were obtained from absorbance at 301 nm in Figure 4A (pKa = 2.8). (C) Optimized structures of H+–PC from DFT calculations and calculated triple bond length of H+–PC and PC. Atom coloring-carbon (gray), hydrogen (white), oxygen (red) and nitrogen (blue). The sugar unit was replaced by methyl group. DFT calculations of H+–PC and PC were conducted at B3LYP/6-31G(d,p) and B3LYP/cccpVDZ level.

We then conducted density functional theory (DFT) calculations on PC at the B3LYP/6-31G(d,p) level to verify the effect of protonation on the band in the Raman spectra. This calculation method was selected on the basis of previous calculations that predicted a Raman spectral shift for protonation.39 The optimized form of protonated PC (H+–PC) is depicted in Figure 4C. Comparing the structures of H+–PC and non-protonated PC (Figure S1) revealed that although both structures had planer phenyl and pyrimidine rings, remarkable differences are observed in the acetylene group. The acetylene triple bond distance was shortened in H+–PC (H+–PC: 1.2162 Å, PC: 1.2181 Å), indicating that protonation affected the structure of the acetylene triple bond. Calculations conducted at a different level (B3LYP/ccpVDZ) gave similar structural results. Thus, it is likely that the protonation at the conjugated cytosine unit strengthened the acetylene bond and caused the shift of the vibrational frequencies of the acetylene unit in PC. From these results, we conclude that the protonation of PC could be tracked by monitoring the band shift in the Raman spectra.

We next attempted to introduce PC into ODNs and to monitor its protonation in the strand. In order to evaluate the protonation at cytosine located at various sequence and location, we prepared several types of ODNs with PC. The structures and sequences of the synthesized ODNs (Scheme S1) are summarized in Figure 2. We found that strand bearing PC formed their second-order structures (i-motif) at ambient temperature under moderately acidic conditions (pH 4.5) (e.g., Theating = 60.0 and 33.0 °C for ODN 6 and ODN 5. See Figure S2), although some destabilization probably due to the steric hinderance of phenylacetylene unit was observed for ODNs with short consecutive sequence of cytosine (Theating = 59.5 and 40.9 °C for corresponding ODNs of ODN 6 and ODN 5 without modification. See Figure S2). The absorption spectra revealed that PC in the strand protonated around pH 3.0 (Figure S3). We also introduced 5-acetylene-substituted uridine (AU) into ODNs as an internal standard for the measurement of the Raman spectra because the wavenumber of the acetylene unit on AU was negligibly affected by pH change (Figure S4). Initially, we measured the Raman spectra of ODN 1 under neutral conditions. In the Raman spectra, two bands were observed at 2220 and 2116 cm–1, which were assigned to acetylene units in PC and AU, respectively. Under acidic conditions (pH 2.0), the band of PC shifted to 2229 cm–1, while that of AU was not changed (Figure 5A, see also Figure S4). A gradual decrease of the pH revealed that an inflection point for change of wavenumber of PC occurred around pH 2.7 (Figure 5B), which is similar to the pKa value of PC. These results strongly indicate that the band shift in the Raman spectra is attributed to the protonation of PC in the ODNs. We also measured the Raman spectra of ODN 1 in the presence of its complementary strand (ODN 2) to form a duplex. We confirmed that the band of PC was observed at 2220 cm–1, indicating that canonical base paring and stacking interactions with marginal base pairs did not affect the vibration of acetylene in PC in the duplex. The PC in the duplex was also protonated under strongly acidic conditions (Figure S5).

Figure 5.

Figure 5

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Raman spectra (excitation at 532 nm) and CD spectra of ODNs. (A) Raman spectra of ODN 1 (1 mM) in 25 mM sodium phosphate buffer under different pH conditions (pH 7.0: black line, pH 5.2: green line, pH 4.2: blue line, pH 3.0: orange line, pH 2.0: red line). (B) Signal shift of ODN 1 as decreasing pH conditions. Results are shown with the mean ± SD (n = 3). (C) CD spectra of ODN 3 (10 μM) at varying pH values (pH 7.0: black line, pH 6.0: green line, pH 5.0: purple line, pH 4.5: dark blue line, pH 4.0: blue line, pH 3.0: red line). (D) Respective plots of the pH vs molar ellipticity at 283 nm obtained from CD spectra (C). (E) Raman spectra of ODN 3 (1 mM) in 25 mM sodium phosphate buffer under different pH conditions (pH 7.0: black line, pH 4.5: blue line, pH 2.0: red line). (F) Signal shift of ODN 3 as decreasing pH conditions. Results are shown with the mean ± SD (n = 3).

For a better understanding of the function of PC in the strand, we next applied the present system to the tracking of DNA second-order structure. We employed the i-motif structure, which formed via intercalating [C–H–C]+ base pairs in a C-rich sequence.8,40 Because i-motif is a typical structure with non-canonical cytosine protonation, we attempted to identify the protonation of cytosine directly by monitoring the vibrational mode of PC in i-motif structure. We prepared the 23-mer ODN 3, bearing AU and PC in the strand, and characterized its behavior. The i-motif formation of ODN 3 in aqueous solution was confirmed by the measurement of CD spectra (Figure 5C,D). The positive band at ca. 285 nm, characteristic of the i-motif, was observed under moderately acidic conditions around pH 4.0 and 5.0. We confirmed that an ODN with the same sequence for i-motif formation but without modification (ODN 4) exhibited similar CD spectra, indicating that even in the presence of the phenylacetylene unit on the nucleobase, ODNs formed an i-motif structure (Figure S6). Interestingly, the Raman spectra of ODN 3 showed that the band of the acetylene unit shifted from 2220 to 2223 cm–1 as the pH was decreased from 7.0 to 4.5 (Figure 5E,F), where PC in ODN 1 forming a random coil did not show any change in the spectra (see Figure 5A,B). Further decrease of pH to 2.0 resulted in an increase of wavenumber to 2228 cm–1. The band shift of the acetylene unit of ODN 3 below pH 3.0 was quite similar to that of the single-stranded ODN 1 and PC mononucleoside under strongly acidic conditions (Figures 5B and 3C). We also observed similar behavior, when the Raman spectra of another i-motif-forming ODN 5 was measured (Figure S7), indicating that the signal shift around pH 4 and 5 was a characteristic behavior for the i-motif structure. Given that the acidic conditions led to dissociation of second-order structures of ODNs into a random coil,41 we assumed that the protonation at PC occurred under pH conditions ranging between pH 2.0 and 3.0 in the random-coiled ODN 3. On the other hand, for the band shift of PC in ODN 3 around pH 4 and 5, evidence of i-motif formation at identical conditions, as proved by CD measurement, led to the following presumption. First, the specific environments in i-motif structure might accelerate protonation at PC even under moderately acidic conditions, in which PC in a random-coiled strand could not protonate based on its pKa (2.8). Second, the specific environments within the i-motif structure directly changed the vibrational mode of the acetylene unit in PC. To verify the band shift of i-motif-forming ODNs around pH 4 and 5, we compared the Raman spectra of ODN 6 and 7. We used ODNs without an internal standard, PU, for measurement of the Raman spectra because ODN 5 gave identical spectra when we conducted calibration using SiO2 as an external standard for measurements of the Raman spectra. Both ODN 6 and ODN 7 formed i-motif structures under moderately acidic conditions, as confirmed by CD spectra (Figure 6C,D). ODN 6 had a sequence that differed from that of ODN 3, and ODN 7 possessed PU in place of PC in ODN 6. As shown in Figure 6B, the change of the Raman band of PU in ODN 7 was negligible despite i-motif formation under the moderately acidic conditions, indicating that the direct effect of i-motif formation on the vibrational mode of the acetylene unit was limited (see Figure S8B). By contrast, the Raman band of PC in ODN 6 responded to pH change in a manner similar to that of ODN 3 and ODN 5. The signal of the acetylene unit of PC in ODN 6 shifted around pH 4 and 5. Previous reports speculated that i-motif formation raised the apparent pKa of protonated nucleobases due to global conformational changes,9 leading to the non-canonical protonation under moderately acidic conditions. Thus, these experimental results suggest that Raman signal changes at moderately acidic conditions were attributed to protonation at PC around pH 4 and 5 in the i-motif structure.

Figure 6.

Figure 6

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(A) Illustration of i-motif-forming ODNs bearing PC or PU in the strand. (B) Raman signal shift of ODN 6 (circle) or ODN 7 (triangle) in 25 mM sodium phosphate buffer under different pH conditions. The Raman signals of ODNs (1 mM) were obtained by using excitation at 532 nm. (C) CD spectra of ODN 6 (10 μM) at varying pH values (pH 7.0: black line, pH 4.0: blue line). (D) CD spectra of ODN 7 (10 μM) at varying pH values (pH 7.0: black line, pH 4.0: blue line).

Further attempts were made to identify protonation of cytosine in i-motif-forming ODNs in crowded environments. We employed molecular crowding environments42−44 and characterized protonation of ODNs. The conditions in cells are inherently molecularly crowded with biomolecules, which can significantly affect many aspects of cellular functions; thus, molecular crowding conditions using highly concentrated cosolutes are recognized as a mimic of the intracellular environments. Sugimoto and co-workers reported that molecular crowding accelerates the formation of higher order-stranded structures.43 In general, an i-motif structure was formed under moderately acidic conditions, while molecular crowding led to the formation of the i-motif structure even under neutral or circumneutral conditions. They predicted that the pKa of cytosine was further raised in such a microenvironment, but its protonation under these conditions was not identified directly. Thus, to track the protonation of cytosine in i-motif-forming ODN under molecular crowding conditions, we measured the Raman spectra of ODNs in the presence of polyethylene glycol (PEG) as a cosolute. First, we measured CD spectra of ODN 6 at pH 6.0 in the presence or absence of PEG. We confirmed that ODN 6 did not form an i-motif structure in the absence of PEG at pH 6.0, whereas addition of PEG up to 20% led to the formation of an i-motif structure even at pH 6.0 (Figure 7A). The Raman spectra of ODN 6 showed that addition of PEG to ODN 6 resulted in a signal shift from 2218.7 to 2222.1 cm–1 even at pH 6.0, indicating that PC in i-motif-forming ODN 6 was protonated at pH 6.0 under molecular crowding conditions. Control experiments using ODN 7 bearing PU instead of PC showed a negligible signal shift in the presence of PEG. In addition, the Raman spectra of ODN 8, which did not form an i-motif structure, but possessed PC in the strand, revealed that there was no change in the spectra at pH 6.0 in the presence or absence of PEG. Thus, PEG did not affect the protonation of PC in random-coiled strands, but induced its protonation in i-motif structures. From these experimental results, we conclude that the protonation of PC in the i-motif structure, even in complicated environments, could be tracked by monitoring changes in the Raman spectra.

Figure 7.

Figure 7

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(A) CD spectra of ODN 6 (25 μM) in aqueous solution containing sodium phosphate buffer (10 mM, pH 6.0) and NaCl (20 mM) in the presence (20%, blue line) or absence (black line) of PEG 300. (B–D) Raman spectra of ODNs (1 mM) in aqueous solution containing sodium phosphate buffer (10 mM, pH 6.0) and NaCl (20 mM) in the presence (20%, blue line) or absence (black line) of PEG 300. Results are shown with the mean ± SD (n = 3). (B) ODN 6 (C) ODN 7 (D) ODN 8.

Conclusions

We tracked the protonation of nucleobase by monitoring its vibrational mode. We prepared a cytosine derivative substituted with a phenylacetylene group as a Raman tag (PC) and monitored its Raman spectra. When the Raman spectra of the PC mononucleoside were measured, the band around 2200 cm–1, which was attributed to the acetylene group, shifted upon decreasing the pH to below 3.0. DFT calculations of PC suggested that the shortening of the acetylene bond due to protonation at the cytosine unit led to the band shift in the Raman spectra. We also confirmed that PC in ODN showed a band shift under acidic conditions due to its protonation. Thus, we tracked the protonation of nucleobases by monitoring changes in the Raman spectra. The most important study is that the Raman spectroscopy was applied to monitor the non-canonical protonation at PC, which is necessary for i-motif formation. We synthesized i-motif-forming ODNs with cytosine-rich sequences, in which one cytosine nucleobase was replaced with PC. The Raman spectra of the ODNs showed a band shift from 2220 to 2223 cm–1 around pH 4 and 5. Given that PC mononucleoside and PC in a random-coiled strand did not show any change under these conditions, the signal shift in i-motif-forming ODNs was attributed to the non-canonical protonation. The evidence of the i-motif formation around pH 4–5 as confirmed by the measurement of CD spectra also supported that the Raman signal shift around these pH conditions was induced by non-canonical protonation in i-motif structure. The Raman shift and protonation behavior was not affected by the sequences and location of PC in the strand. In addition, we explored the protonation of nucleobases in ODNs under molecular crowding conditions. Addition of PEG into the solution of i-motif-forming ODNs led to a signal shift in the Raman spectra even at pH 6.0 conditions, where the PC could not protonate in the absence of PEG because ODNs could not form i-motif structure. Thus, we concluded that non-canonical protonation on nucleobases even under complicated conditions could be monitored.

Although the present method to identify protonation of a nucleobase using the Raman spectroscopy would be a useful tool for analyzing a variety of structures and reactions of DNA strands, the present system needs improvements. First, the pKa of PC was smaller than that of cytosine mononucleoside due to the introduction of phenylacetylene unit at 5-position, and protonation of PC occurred under more acidic conditions than that of cytosine. Second, the small signal shift for protonation at PC and its small pKa may lead to misunderstandings. Third, the destabilization of duplex by phenylacetylene unit is also a task that should be improved. Thus, at present, PC with a Raman tag could not be a perfect alternative nucleobase of cytosine. However, present results suggest that the change of chemical structure in atomic level could be observed even in the complicated environments. In addition, preliminary experiments revealed that the signal of ODNs with PC was observed from the solution containing cell lysate (Figure S9), and thus, the system is expected to work in the cells. The improvement of the basic properties of PC to increase its pKa and eliminate of steric hinderance by modification of substituent on acetylene group and applications to monitoring of higher order structures of DNA in the cells are in progress.

Materials and Methods

General

Reagents were purchased from Wako pure chemical industries, Tokyo chemical industries, Sigma-Aldrich, nacalai tesque, Glen Research and Eurofins Genomics. The course of reactions was monitored by thin-layer chromatography on silica gel plates (Silica Gel 60 F254). Wakogel 60 was used for silica gel column chromatography. NMR spectra were recorded on a JNM-ECX500II (1H: 500 MHz, 13C: 500 MHz)spectrometer (JEOL) and chemical shifts are expressed in ppm downfield from tetramethylsilane, using residual protons in the solvents as an internal standard (chloroform: δ 7.26 in 1H NMR, δ 77.0 in 13C NMR dimethyl sulfoxide: δ 2.49 in 1H NMR, δ 39.5 in 13C NMR). FAB mass spectrometry was performed with a JMS-700A mass spectrometer (JEOL), using nitrobenzyl alcohol as a matrix. Matrix-assisted laser desorption ionization time-of-flight (MALDI-TOF) mass spectrometry were recorded on an AXIMER-LNR spectrometer (Shimadzu). The Raman spectrum measurement was carried out on a Renishaw inVia Raman microscope. UV–visible spectra were obtained at 250–600 nm using a JASCO V-630 Uv/vis spectrophotometer. DNA synthesis was performed with a 3400 DNA synthesizer (Applied Biosystems). Analytical and preparative high-performance liquid chromatographies were carried out with a D2000 HPLC system (Hitachi) and L7000 HPLC system (Hitachi) with a reversed phase column (Inertsil ODS-3, GL Science Inc., ϕ 10 mm × 250 mm, ϕ 4.6 mm × 250 mm). The column eluents were monitored by the UV absorbance at a flow rate of 0.6 (analysis) or 3.0 (preparative) mL min–1 with a 0–30% (over 60 min), 30–100% (over another 30 min) gradient of ACN/TEAA buffer (100 mM, pH 7.0). CD spectra were recorded using a JASCO J-805 circular dichroism spectrometer. PC,45PU,46AU,47 compound 6,48 and compound 7(49) were synthesized by the reported procedure.

Synthesis of Compound 2

A solution of 1 (584.2 mg, 1.00 mmol) in 4 mL of anhydrous dimethylformamide (DMF) and triethylamine was added to phenylacetylene (82 μL, 0.746 mmol), tetrakis(triphenylphosphine)palladium (28.3 mg, 0.0247 mmol), copper iodide (5.2 mg, 0.0273 mmol) and then stirred for 0.5 h at room temperature. The resulting mixture was extracted with ethyl acetate, washed with water then brine, dried over anhydrous magnesium sulfate, and concentrated. The crude product was purified by silica gel column chromatography (chloroform/methanol = 30/1) to give 2 (359 mg, 58%) as a yellow solid. 1H NMR (CDCl3): δ 8.21 (s, 1H), 7.44–7.32 (m, 5H), 6.28 (t, J = 6.5 Hz, 1H), 4.37 (m, 1H), 3.96–3.92 (m, 2H), 3.76 (dd, J = 11.0, 1.5 Hz, 1H), 2.48 (ddd, J = 13.0, 7.5, 2.5 Hz, 1H), 2.06 (ddd, J = 13.0, 6.5, 6.0 Hz, 1H), 0.89 (s, 9H), 0.88 (s, 9H), 0.13 (s, 3H), 0.11 (s, 3H), 0.06 (s, 3H), 0.05 (s, 3H); 13C NMR (CDCl3): δ 163.9, 154.3, 144.1, 131.4, 129.0, 128.8, 122.3, 95.3, 90.6, 88.2, 86.8, 80.8, 72.2, 62.9, 42.6, 26.2, 26.0, 18.8, 18.1, −4.5, −4.8, −5.0, −5.1; MS (FAB) m/z 556 [(M + H)+]; HRMS (FAB) calcd for C29H45N3O4Si2 [(M + H)+], 556.3027; found, 556.3025; mp 176.5–177.2 °C.

Synthesis of Compound 3

A solution of 2 (107.8 mg, 0.193 mmol) in 1 mL of anhydrous DMF was added to DMF-diethyl-acetal (1 mL) and then stirred for 14 h at room temperature. The resulting mixture was extracted with ethyl acetate, washed with water then brine, dried over anhydrous magnesium sulfate, and concentrated under vacuum. The crude product was purified by silica gel column chromatography (chloroform/methanol = 30/1) to give 3 (115.2 mg, 94%) as yellow oil. 1H NMR (CDCl3): δ 8.81 (s, 1H), 8.25 (s, 1H), 7.44–7.26 (m, 5H), 6.37 (t, J = 6.2 Hz, 1H), 4.37 (m, 1H), 3.95–3.91 (m, 2H), 3.77 (dd, J = 11.5, 1.4 Hz, 1H), 3.19 (s, 3H), 3.13 (s, 3H), 2.49 (ddd, J = 13.3, 4.4, 4.0 Hz, 1H), 2.04 (ddd, J = 13.3, 5.8, 5.8 Hz, 1H), 0.87 (s, 9H) 0.86 (s, 9H), 0.12 (s, 3H), 0.10 (s, 3H), 0.04 (s, 3H), 0.03 (s, 3H); 13C NMR (CDCl3): δ 170.5, 158.5, 155.2, 144.4, 131.8, 128.3, 128.0, 123.8, 99.0, 92.9, 88.2, 87.0, 83.9, 72.1, 62.6, 42.6, 41.4, 35.4, 26.3, 26.2, 18.7, 18.2, −4.4, −4.2, −4.8, −5.1; MS (FAB) m/z 611 [(M + H)+]; HRMS (FAB) calcd for C32H50N4O4Si2 [(M + H)+], 611.3449; found, 611.3464.

Synthesis of Compound 4

A solution of 3 (249.1 mg, 0.407 mmol) in 1.5 mL of anhydrous tetrahydrofuran was added to acetic acid (70 μL, 1.22 mmol) and tetrabutylammonium fluoride (1 N) and then stirred for 1 h at room temperature. The resulting mixture was neutralized with saturated aqueous sodium hydrogen carbonate, extracted with ethyl acetate, washed with water then brine, dried over anhydrous magnesium sulfate, and concentrated. The crude product was purified by silica gel column chromatography (chloroform/methanol = 30/1) to give 4 (128.4 mg, 82%) as white solid. 1H NMR (DMSO-d6): δ 8.61 (s, 1H), 8.41 (s, 1H), 7.42–7.32 (m, 5H), 6.10 (t, J = 6.5 Hz, 1H), 5.20 (dd, J = 4.5, 1.5 Hz, 1H), 5.11 (t, J = 4.5 Hz, 1H), 4.21 (m, 1H), 3.79 (m, 1H), 3.68–3.61 (m, 1H), 3.59–3.52 (m, 1H), 3.17 (s, 3H), 3.11 (s, 3H), 2.23–2.16 (m, 1H), 2.08–1.98 (m, 1H); 13C NMR (CDCl3): δ 170.6, 158.6, 155.0, 146.0, 131.6, 128.5, 128.3, 123.5, 99.6, 93.0, 88.7, 87.5, 83.1, 70.6, 61.8, 41.5, 40.9, 35.5; MS (FAB) m/z 383 [M + H]+; HRMS (FAB) calcd for C20H22N4O4 [M + H]+, 383.1720; found, 383.1718; mp 110.2–111.7 °C.

Synthesis of Compound 5

A solution of 4 (782.5 mg, 2.046 mmol) in anhydrous pyridine was added to 4,4′-dimethoxytrityl chloride (470.3 mg, 1.388 mmol) and then stirred for 3 h at room temperature. After concentration, the crude product was purified by silica gel column chromatography (chloroform/methanol/trimethylamine = 4003/4/1) to give 5 (237.6 mg, 18%) as white solid. 1H NMR (CDCl3): δ 8.83 (s, 1H), 8.28 (s, 1H), 7.45 (d, J = 8.0 Hz, 2H), 7.35 (m, 4H), 7.28–7.08 (m, 8H), 6.78 (m, 4H), 6.36 (t, J = 6.5 Hz, 1H), 4.49 (br s, 1H), 4.13 (m, 1H), 3.67 (s, 3H), 3.67 (s, 3H), 3.43 (dd, J = 13.4, 3.5 Hz, 1H), 3.34 (dd, J = 13.4, 4.1 Hz, 1H), 3.22 (s, 3H), 3.17 (s, 3H), 2.68 (m, 1H), 2.27 (m, 1H); 13C NMR (CDCl3): δ 170.6, 158.6, 158.6, 154.9, 144.6, 144.3, 135.8, 135.7, 131.4, 130.1, 130.0, 128.1, 128.0, 127.8, 127.0, 123.6, 113.4, 99.5, 92.9, 87.0, 86.3, 83.2, 72.4, 63.6, 55.2, 46.3, 42.4, 41.5; MS (FAB) m/z 685 [M + H]+; HRMS (FAB) calcd for C41H40N4O6 [M + H]+, 685.3087; found, 685.3026; mp 139.1–141.2 °C.

Synthesis of ODN 6 (General Procedure for the Synthesis of ODNs)

N,N-diisopropylethylamine (77 μL, 0.442 mmol) and 2-cyanoethyldiisoproryl-chlorophosphor-amidite (55 μL, 0.247 mmol) were added to compound (5, 65.5 mg, 0.0957 mmol) in anhydrous tetrahydrofuran (950 μL) and stirred for 4 h at room temperature. After the reaction, the mixture was filtered and placed on a DNA synthesizer. After automated DNA synthesis, ODN 6 was purified by reversed phase HPLC. The purity and concentration of the oligomers were determined by complete digestion with AP, P1, and phosphodiesterase I at 37 °C for 12 h. Identities of synthesized oligomers were identified by MALDI-TOF mass spectrometry (ODN 1: [M – H]− calcd 7634.2; found, 7634.3, ODN 3: [M – H]− calcd 6942.6; found, 6942.0, ODN 5: [M – H]− calcd 5460.8; found, 5460.1, ODN 6: [M – H]− calcd 6300.2; found, 6299.5, ODN 7: [M – H]− calcd 6301.2; found, 6300.4, ODN 8: [M – H]− calcd 7623.2; found, 7622.7).

Measurement of the Raman Spectra

The samples of ODNs were prepared in a sodium phosphate buffer and the sample solution was dropped on a hole glass slide (14–15 ϕ × 0.6 mm), then a cover glass (0.12–0.17 mm) was placed on the hole. The Raman spectra were measured by means of the excitation at 532 nm. The laser output was focused into the sample by 100 × objective lens (Leica N Plan Epi 100×/0.85). The slid width of the spectrograph was 20 μm. The laser power was 50 mW and exposure time for a point was 80–100 s. The Raman spectra were acquired with a confocal unit Renishaw inVia Raman microscope (Renishaw plc). The acquired images were analyzed and processed for multicomponent analysis with the software WiRE 2.0 (Renishaw plc, UK).

Measurement of Absorption Spectra

The samples were prepared and the absorption spectra were recorded on a JASCO-V630 spectrophotometer. UV spectra were obtained by monitoring the UV absorbance at 240–400 nm.

Measurement of CD Spectra

CD spectra were recorded using a JASCO J-805 circular dichroism spectrometer. The scans were gathered over the wavelength ranging from 200 to 350 nm in a 0.1 cm path length cell at the standard sensitivity, data pitch 0.5 nm, continuous scanning mode, scanning speed 50 nm min–1, response 8 s, and bandwidth 1 nm.

DFT Calculation

Structural data of PC and H+–PC were obtained from quantum chemical calculations performed with the GAUSSIAN 09 program (rev. E.01, Gaussian Inc.).50 First, structure of deoxycytidine calculated at B3LYP/6-31G(d) level was obtained from a Spartan Molecular database in a modeling software Spartan’18 (Wavefunction, Inc.). Then, the sugar unit was replaced by a methyl group and a phenyl acetylene unit was added to the C5 position of the methylated cytosine to construct pC using GaussView (ver 5.0). Geometry optimization and vibrational frequencies of the ground states of pC and H+–PC were calculated at B3LYP/6-31G(d) and B3LYP/ccpVDZ. The effects of interaction with the solvent (water, ε = 78.39) on the geometry and frequencies were taken into account by the integral equation formalism of the polarizable continuum model. The initial structure of H+–PC was constructed by adding hydrogen atom to N3 of the cytosine in the geometry optimized pC.

Acknowledgments

This work was supported in part by fund for research unit in Aoyama Gakuin University Research Institute (for K.T.) and Grant-in-Aid for Scientific Research (for K.T. grant number 20H02863).

Supporting Information Available

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acsomega.1c04074.

  • Detailed experimental procedure, synthetic scheme of ODNs and Raman spectra that could not be provided in the manuscript (PDF)

The authors declare no competing financial interest.

Supplementary Material

ao1c04074_si_001.pdf (1.2MB, pdf)

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