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. 2003 Feb 15;31(4):1331–1338. doi: 10.1093/nar/gkg203

Structural effect of the anticancer agent 6-thioguanine on duplex DNA

Jen Bohon 1, Carlos R de los Santos 1,a
PMCID: PMC150222  PMID: 12582253

Abstract

The incorporation of 6-thioguanine (S6G) into DNA is an essential step in the cytotoxic activity of thiopurines. However, the structural effects of this substitution on duplex DNA have not been fully characterized. Here, we present the solution structures of DNA duplexes containing S6G opposite thymine (S6G·T) and opposite cytosine (S6G·C), solved by high-resolution NMR spectroscopy and restrained molecular dynamics. The data indicate that both duplexes adopt right-handed helical conformations with all Watson–Crick hydrogen bonding in place. The S6G·T structures exhibit a wobble-type base pairing at the lesion site, with thymine shifted toward the major groove and S6G displaced toward the minor groove. Aside from the lesion site, the helices, including the flanking base pairs, are not highly perturbed by the presence of the lesion. Surprisingly, thermal dependence experiments suggest greater stability in the S6G-T mismatch than the S6G-C base pair.

INTRODUCTION

Thiopurines are used clinically to treat childhood acute lymphoblastic leukemia as well as other forms of leukemia. They are also used to treat a variety of other conditions, are used as immunosupressants in transplant surgery, and most recently have been shown to impair HIV replication (1,2). These therapeutic agents are transformed through cellular processing to 6-thioguanine triphosphate, the active metabolite, and then incorporated into the DNA duplex (1,3). It has been shown that 6-thioguanine (S6G) exhibits a delayed cytotoxic activity, halting the cell cycle in the G2 phase after one round of replication (4). Types of damage caused by the presence of S6G include single- and double-strand breaks, cross-linking to protein and DNA, sister chromatid exchange events, and large-scale chromosomal damage (4–10). Experiments in vitro show that DNA synthesis with S6G in the template strand can occur, but that it is quite slow (11). S6G also blocks the action of some restriction enzymes, greatly decreases the rate of ligation, and can inhibit sequence-specific binding of proteins to the substituted DNA (12). The presence of S6G in DNA also inhibits the ability of RNase H to hydrolyze RNA from hybrid duplexes (13). Additionally, S6G is methylated in vivo by S-adenosylmethionine, forming S6-methylguanine (S6meG), a mutagenic lesion that miscodes for thymine 50% of the time (14). Both S6meG-T and S6G-T pairs are bound by hMutSα (15,16), and cells deficient in the components of this repair complex are often resistant both to S6G treatment and methylating agents, and exhibit a mutator phenotype (17). Thus, several theories about the mechanism of cytotoxicity of S6G have focused on improper processing by the post-replicative mismatch repair system (14,18,19) with parallels to the cytotoxic mechanism of O6-methylguanine (14,20).

This single-atom substitution clearly has a substantial impact on cellular biology. However, the structural changes caused by its incorporation into DNA have not been well determined. Structural and thermodynamic research has been done on G-T mismatches (21–24) and on O6- and S6-methyl guanine hydrogen bonding (25), allowing interesting bases for comparison. Previous work relating to structural properties has indicated that the pairing of S6G with cytosine (26,27) is likely to be quite unstable. Modified neglect of differential overlap (MNDO) calculations have shown that the presence of the sulfur could significantly affect the hydrogen-bonding and general stability of duplex DNA (26). A recent study has suggested that S6G does not form Watson–Crick hydrogen bonds at all and that the structural repercussions of the presence of this lesion extend well beyond the lesion site (27). However, our findings clearly indicate that S6G does indeed form hydrogen bonds, both opposite cytosine and opposite thymine. Interestingly, thermal dependence experiments point toward a greater degree of stability in the S6G-T mismatch than the S6G-C pair. The chemical structure of S6G and the sequences of DNA used in this research are shown in Figure 1.

Figure 1.

Figure 1

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Chemical structure of 6-thioguanine and sequence of lesion containing duplexes (S represents 6-thioguanine and X represents either cytosine or thymine).

MATERIALS AND METHODS

Synthesis of deoxyribo-oligonucleotides

Protected 6-thioguanosine precursor for solid phase DNA synthesis was purchased from Glen Research, Inc., and was incorporated into the oligonucleotide sequence by standard phosphoramidite chemistry procedures. Eleven oligomer DNA d(CGTACSCATGC), d(GCATGTGTACG) and d(GCATGCGTACG) (where S represents 6-thiodeoxyguanosine) were synthesized at the SUNY Stony Brook DNA synthesis facility. Purification of modified and unmodified oligonucleotides was performed as described previously (28). Purity was >95%.

Duplex formation and sample preparation

A 1:1 stoichiometry was obtained for each duplex by addition of the appropriate amount of each strand as calculated by Generunner (1994, V.3.00, Hastings Software, Inc.). NMR experiments performed afterward did not reveal peaks that might suggest excess of either strand. The samples consisted of ∼300 OD260 U of duplex dissolved in 0.7 ml of a 25 mM phosphate buffer containing 50 mM NaCl and 0.5 mM EDTA at a pH of 6.78, for a DNA concentration of 4.3 mM. These samples were lyophilized and placed into either 99.96% D2O (D2O buffer) or 10% D2O/90% H2O (H2O buffer) solution at pH 6.78 for collection of NMR data.

NMR experiments

NMR experiments were conducted on Varian (INOVA) spectrometers at 500 and 600 MHz. Proton chemical shifts were referenced to 3-(trimethylsilyl)-propionate-2,2,3,3,-d4 at 0 p.p.m. Phase-sensitive NOESY, COSY, DQFCOSY, TOCSY and COSY45 data were taken with each duplex in D2O buffer at 25°C. NOESY mixing times collected for the S6G·T duplex were 50, 100, 150, 200 and 250 ms. For the S6G·C duplex, 50, 100, 200 and 300 ms mixing times were collected. TOCSY experiments were done with a mixing time of 120 ms. NOESY data with mixing times of 120 and 220 ms were taken for each duplex in H2O buffer at 3°C. Time-domain data sets consisted of 4096 by 300 complex data points in the first and second dimensions, respectively. For the acquisition of a COSY45 spectrum, the number of complex points in both dimensions was doubled. One-dimensional experiments following the thermal dependence of imino protons were performed in H2O buffer at: 3, 10, 15, 25, 39, 45 and 52°C for S6G·T; 3, 10, 17, 24, 31, 39, 45, 52 and 58°C for S6G·C; 3, 10, 15, 20, 25, 30, 35, 40, 45, 50, 55, 60, 65, 70 and 75°C for G·C; and 3, 10, 15, 20, 25, 30, 35, 40, 45, 50, 55 and 60°C for G·T. NMR data were processed using FELIX95 and FELIX98 (Accelrys, San Diego, CA) software running on Silicon Graphics workstations. Shifted-sinebell window functions were applied to all data before Fourier transformation. Exponential multiplication with a line broadening of 2 Hz and baseline correction were applied to the thermal dependence data. Noise running parallel to the first dimension that appeared in two-dimensional data sets was systematically removed by a subtraction macro in FELIX95.

Distance calculation and molecular dynamics simulation

Interproton distance calculation and molecular dynamics were carried out using XPLOR3.1 on Silicon Graphics workstations (29). Molecular dynamics simulations were performed in vacuo using a CHARMM-derived force field (30) and the dielectric constant was set to 4 (31). Partial charges for S6G were derived from the literature (32,33). Distances were calculated by directly inputting the NOE cross-peak volumes obtained from FELIX for all mixing times into a relaxation protocol in XPLOR. These volumes were normalized against CH5-H6 and H2′-2′′ NOEs, which correspond to fixed distances (34). Only a relaxation energy term related to the difference between the back-calculated and experimental NOE intensities was included during minimization. A grid search was performed to find the best-fit isotropic correlation time of 2.25 ns. The volumes were given error bounds of 30% initially and the structure was put through 300 steps of minimization. The bounds were tightened to 10%, and then 1%, with 300 steps of minimization between each narrowing. After final minimization, distances were extracted from the coordinates of the structure using Perl (35). A total of 430 distinct NOE distances for the S6G·T mismatch duplex and 336 for the S6GC duplex were restrained during molecular dynamics using square-well potential energy functions. Distance boundaries of ±0.6 Å were used on most distances acquired from single-peak assignments, but were increased to ±0.8 Å for overlapping peaks. Based on NMR data, Watson–Crick alignment for normal pairs was enforced by distance restraints with bounds of ±0.1 Å from crystallographically determined distances. In the case of the lesion-containing base pairs, hydrogen bonds across S6G-C were enforced using canonical distances with bounds of ±0.3 Å, while those of S6G-T were constrained between 2.6 and 3.2 Å (23). Sugar conformations were derived from COSY45 data and enforced using empirical dihedral angle potential energy functions (36,37). Similarly, backbone dihedral angles were restrained within a range encompassing A- and B-form DNA. Covalent bond lengths of all hydrogens were kept constant during the simulations through use of the SHAKE algorithm (38). Two structures (canonical A- and B-form DNA) were generated in InsightII (Accelrys) for each duplex and then minimized to generate the initial structures used in the molecular dynamics simulations. Five variations in initial temperature (50, 75, 100, 125 and 150 K) and three variations in the length of the high-temperature step (30, 35 and 40 ps) were used for each starting structure (A- and B-form DNA) of each duplex for a total of 30 structures for each duplex. The temperature was increased gradually from the starting value to 500 K over a period of 5 ps. During this time, the NOE force constant was also increased gradually from 2 to 150 kcal/mol-Å2. The dynamics were allowed to progress for 30–40 ps at 500 K and then the temperature was cooled to 300 K over a period of 10 ps. The simulation then continued for 160 ps at 300 K. After this point, the structures were further refined by 30 ps of molecular dynamics at 300 K restrained by the full relaxation matrix method (39,40). During this period, the NOE force constant was brought to zero over 5 ps as the scale of a potential function based on the intensity difference between experimental and back-calculated NOESY spectra was increased (see Supplementary Material, Fig. S1). A total of 1750 and 1110 experimental volumes were used with 30% boundaries for the S6G·T and S6G·C duplexes, respectively. The structures were then minimized to obtain the final NMR-refined structures for each duplex. Analysis of the structural parameters was performed using Curves (41).

RESULTS

Structural determination

Assignment of non-exchangeable protons was based on NOESY, COSY, DQFCOSY and TOCSY spectra using standard procedures (42–44). Intraresidue and sequential base to H1′, H2′, H2′′ and H3′ crosspeaks characteristic of right-handed duplex DNA were observed for both the S6G·C and S6G·T duplexes. These connectivities were seen throughout the duplexes, including the lesion site and surrounding residues (Fig. 2). Interstrand adenine H2 to H1′ sugar proton connectivities reinforced the observed regularity of the helices. The duplexes exhibited similar NMR spectra, with analogous chemical shifts for most crosspeaks. The chemical shift data for both duplexes measured at 25°C are available in the Supplementary Material (Tables S1 and S2).

Figure 2.

Figure 2

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Base-to-H1′ region of NOESY spectra of the S6G·C (left) and S6G·T (right) duplexes, recorded in 100% D2O buffer at 25°C. The spectra were taken at mixing times of 300 and 250 ms for the S6G·C and S6G·T duplexes, respectively. Peaks are labeled D1–D2.

Assignment of the exchangeable protons was straightforward (43). Imino proton crosspeaks due to interactions with adjacent bases establish base pair formation and normal base stacking throughout both duplexes (Figs 3 and 4). In the S6G·C duplex, the crosspeaks due to the interaction of S6GH1 with the C17 amino protons (Fig. 3, panel 2), as well as those due to stacking interactions with G16 and G18 (Fig. 3, panel 3, peaks A and B), are clearly present. The chemical shift and sharpness of the S6GH1 signal, as well as the chemical shift separation of C17 amino protons (1.98 p.p.m.), provide clear indications that the S6G-C base pair adopts Watson–Crick alignment. However, we note that the intensity of these crosspeaks is weak in comparison with all non-terminal base pairs in the duplex. The diagonal peak of S6GH1 (Fig. 3, panel 3) and the corresponding signal in the one-dimensional spectrum (Fig 3, panel 1) are also of low intensity. Taken together, these observations suggest a relatively rapid exchange of S6GH1 with the solvent. In the case of the S6G·T duplex, the S6GH1 and T17H3 protons appear as sharp signals in the one-dimensional spectrum (Fig. 4, panel 1) and the diagonal peaks for these protons are as intense as any other non-terminal imino proton in the duplex (Fig. 4, panel 3). There is also a very strong crosspeak between S6GH1 and T17H3, indicating that they are quite close spatially. S6GH1 and T17H3 clearly interact with G18H1 and G16H1 (Fig. 4, panel 3, peaks B–E), indicating that both mismatched bases are intrahelical and stack well with flanking base pairs. These observations, as well as the interaction of T17H6 with G16H8 but not G18H8, demonstrate that the S6G-T mismatch assumes a wobble pairing similar to that of G-T (22–24).

Figure 3.

Figure 3

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Panel 1: one-dimensional trace labeling imino protons. Panel 2: imino–amino region. Blue boxes enclose crosspeaks due to interaction with cytosine amino protons participating in hydrogen bonds. Red boxes indicate interactions with cytosine amino protons not participating in hydrogen bonds. Green boxes enclose crosspeaks due to interaction with adenine H2 protons. Panel 3: imino–imino region. Crosspeak labeling: (A) S6GH1–G18H1, (B) S6GH1–G16H1. Blue asterisks denote crosspeaks due to stacking interactions with imino protons of adjacent bases. Panels 2 and 3 are regions of a NOESY spectrum of S6GC, recorded in 10% D2O buffer at 3°C with a mixing time of 180 ms.

Figure 4.

Figure 4

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Panel 1: one-dimensional trace labeling imino protons. Panel 2: Imino–amino region. Blue boxes enclose crosspeaks due to interaction with cytosine amino protons participating in hydrogen bonds. Red boxes indicate interactions with cytosine amino protons not participating in hydrogen bonds. Green boxes enclose crosspeaks due to interaction with adenine H2 protons. Panel 3: Imino–imino region. Crosspeak labeling: (A) S6GH1–T17H3, (B) G18H1–T17H3, (C) G16H1–T17H3, (D) G18H1–S6GH1, (E) G16H1–S6GH1. Blue asterisks denote crosspeaks due to stacking interactions with imino protons of adjacent bases. Panels 2 and 3 are regions of a NOESY spectrum of S6G·T, recorded in 10% D2O buffer at 3°C with a mixing time of 220 ms.

Molecular dynamics simulations restrained by these NMR data yielded a family of structures for each duplex (see Supplementary Material, Fig. S2). An averaged structure (Fig. 5) was calculated and minimized for each family to yield the reported coordinates for S6GC (PDB no. 1KB1) and S6G·T (PDB no. 1KBM). Twenty-nine refined structures converged to an average RMSD of 0.64 Å from the averaged S6GT structure, with a maximum deviation of 1.06 Å. In the case of S6G·C, 25 refined structures converged to an average RMSD of 0.65 Å from the averaged structure, with a maximum deviation of 1.03 Å. The structures are regular in shape and generally conform to parameters (41) for normal B-form DNA.

Figure 5.

Figure 5

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Energy minimized averaged structures for S6G·C (left) and S6G·T (right). Blue, S6G; green, C or T opposite S6G.

The S6G-T mismatch (Fig. 6) assumes a wobble-type base pairing comparable to that of a previously reported G-T mismatch (21), but with increased hydrogen bond distances (Table 1). However, the planarity is not entirely conserved and a base pair buckle of –11° occurs between S6G and T17 (Fig. 5). The S6G-C pair displays a generally planar orientation within the duplex (Fig. 5), but the hydrogen-bonding distances are clearly different from those of a normal G-C pair. The distances surrounding the sulfur are increased an average of ∼10%, while the bond distance nearest the minor groove is actually decreased by 8.3% (Table 1). This causes a base pair opening of 7° toward the major groove. Despite these deviations from normal DNA, a closer look at the lesion site reveals the clear localization of perturbations (Figs 5–7). The flanking base pairs are basically undisturbed, retaining planarity and Watson–Crick alignment. This regularity is also evident for base pairs further from the lesion. Comparative structural parameters are listed in Table 1.

Figure 6.

Figure 6

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Base pair configurations. Top left, S6G-C; bottom left, canonical G-C (49); top right, S6G-T; bottom right, G-T (23). Hydrogen bonds are shown in yellow.

Table 1. Structural parameters.

Base pair Shear (Å) Buckle (°) Propeller twist (°) Opening (°) Hydrogen bond distances (Å)
S6G-C –1.0 –3 –4 7 GN1–CN3, 3.2; GS6–CN4, 3.1; GN2–CO2, 2.8
G-Ta –2.6 –1 –14 2 GO6–TN3, 2.7; GN1–TO2, 2.6
S6G-T –3.3 –11 –8 18 GS6–TN3, 3.3; GN1–TO2, 2.9
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Values for G-C were derived from Curves analysis of a canonical InsightII (Accelrys) B-form structure: shear, 0.0; buckle, 0; twist, 3; opening, –4. Hydrogen bond distances: GN1–CN3, 3.0; GO6–CN4, 2.9; GN2–CO2, 2.9.

aValues derived from Curves (41) analysis of previously reported structure (PDB no. 1BJD) (23).

Figure 7.

Figure 7

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Stacking interactions of S6G·C (left panel) and S6G·T (right panel). The top of each panel shows the interactions with the flanking bases above the lesion site (fifth and eighteenth bases), and the bottom shows the interactions with the flanking bases below the lesion (seventh and sixteenth bases).

Stability effects

NMR-based temperature dependence experiments following the broadening of imino peaks were utilized in order to investigate the relative thermal stability of both experimental and control duplexes. The control duplexes have sequences identical to the experimental duplexes except for the presence of guanine in place of S6G. Chemical shifts of lesion-containing duplexes as well as those of the control duplexes were similar enough to allow easy comparison of the spectra (Fig. 8). The imino protons in the S6G·T and G·T duplexes display similar patterns of temperature dependence. While the signals of the terminal pairs decrease quite quickly, those at the lesion site disappear at only a slightly lower temperature than normal base pairs in the duplexes. This effect is highly localized, as the base pairs flanking the lesion begin to broaden only as the entire structure collapses. The G-T and S6G-T mismatches thus do not appear to be much less stable than any other pair in these duplexes. This is in contrast to what is seen for the S6G·C duplex. It is immediately apparent upon inspection of this data that the imino proton signal of the S6G-C base pair disappears considerably faster than those of the G-C, G-T and S6G-T pairs. The broadening of the S6GH1 peak in this duplex in fact parallels that of the terminal peaks. The most striking comparison of the duplexes occurs at 35°C (Fig. 8), where the lesion site imino proton peaks are still quite sharp for S6G·T, but the S6G·C S6GH1 peak has clearly broadened. This implies a faster apparent opening rate for S6G-C than S6G-T, suggesting a decreased stability at the lesion site in S6G·C. Interestingly, even in the case of S6G·C, the flanking base pairs seem quite stable, and do not show significant instability until >55°C, when the entire imino region begins to ‘melt’. This indicates a general stability in the base pairing of the duplex that is not perturbed by the presence of this rather unstable S6G-C pair.

Figure 8.

Figure 8

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Temperature dependence experiments. Spectra (3, 15, 35 and 50°C) are depicted for each duplex as labeled. Experiments were carried out in 10% D2O/90% H2O buffer phosphate, pH 6.78. Base pairs in the sixth and seventeenth positions (lesion site) are marked with blue asterisks. Flanking guanine residues in the sixteenth and eighteenth positions are marked with red arrows. Terminal base pairs appear in close proximity and are marked with a single green arrow for each duplex.

DISCUSSION

The structures solved in this work indicate that the presence of S6G causes only a localized distortion of duplex DNA. The alignment at the lesion site is basically similar to that of canonical G-C or previously solved G-T structures. The hydrogen bonding distances at the lesion site are generally increased due to the large atomic radius of sulfur and lower electronegativity comparative to the normal oxygen in this position. The exception to this is the GN2–CO2 distance in the S6G-C pair, which is actually decreased to 2.76 Å from the normal value of 3.01 Å (Table 1). In addition, there is an opening toward the major groove of 7° in the S6G-C pair and 16° in the S6G-T mismatch. Energetic analysis indicates that these perturbations contribute to the reduced stability of base pairs containing the S6G lesion. However, they do not disturb the flanking base pairs significantly, nor the remainder of the base pairs in the duplexes. This is verified by both structural studies and thermal stability experiments. Our results are in direct conflict with a previous report, which suggests that S6G causes generalized distortion of the duplex and does not form a base pair (27). Several factors make it unlikely that the disparity in these findings is due to the differences between the sequences used in these studies. The previous study utilized a thioguanine-modified ‘Dickerson’ dodecamer duplex which contained two S6G-C pairs separated by four A-T base pairs. Although the experimental data shown in that study were limited, it is stated in the text that the A-T base pairs were present, including those flanking the lesion site. Because the center of the duplex is intact, it is unlikely that the lack of evidence for S6G-C base pair formation results from instability caused by the proximity of the two lesions. Also, recent base pair kinetics experiments have shown that the sequence of the ‘Dickerson’ duplex does not increase the opening rate of the fourth G-C pair (45,46), which is the site of S6G substitution (27). Thus, the failure of the previous study to observe a Watson–Crick S6G-C pair cannot be readily explained by the sequence composition. Based on these considerations and our results, the assertion that S6G does not participate in a base pair in the ‘Dickerson’ duplex deserves more investigation.

The apparent thermal instability of the S6G-C base pair relative to the S6G-T base pair (Fig. 8) is difficult to explain. The possibility that S6G-C undergoes imino proton exchange by a different mechanism than S6G-T, perhaps involving internal catalysis, is highly unlikely. Our current investigation of base pair dynamics indicates that the exchange rates of both S6G-C and S6G-T depend on the presence of an external catalyst. Based on the similarity of our experimentally determined solution structures, pKa values of S6G in both duplexes are likely to be nearly identical. Therefore, the increased exchange rate of the imino proton must be ascribed to an increased opening rate of the S6G-C base pair. Clearly, this increased rate cannot be attributed to hydrogen bonding, as S6G-C has all three hydrogen bonds in place, while S6G-T has only two (Table 1). Furthermore, the base stacking (Fig. 7) is quite similar for both duplexes and is unlikely to contribute to this effect. A possible explanation is a decreased entropic content at the lesion site in S6G·C. A full thermodynamic investigation is needed to address this issue.

Although the incorporation of S6G causes only minor distortions in the overall structure of duplex DNA, these changes create the environment for cytotoxicity. This could be due to differential recognition of the S6G sites by cellular enzymes such as those required for repair and replication. The thermal stability of the S6G-T mismatch is similar to that of G-T mismatches, and it has been shown that they are incised with equivalent frequency by normal cellular repair systems (17). This, coupled with the similarity in structure and hydrogen bonding to G-T mismatches revealed by this work, leads to the conclusion that it is not likely that the mispairing of S6G with thymine directly leads to cell death. However, these S6G-T lesions are ‘repaired’ to form S6G-C lesions, which are only minimally recognized by mismatch repair proteins (14), and are likely to persist in the genome. The modified hydrogen bonding distances have a significant effect on the stability of the S6G-C pair, increasing the base pair opening rate and making it more accessible to solvent and other nuclear components. This ‘loose’ pairing could facilitate endogenous methylation of the S6G residue to form S6meG, a known mutagenic lesion, and thus lead to cytotoxicity (14). This alone cannot completely explain the cytotoxicity of S6G, however, as it would take many cycles of replication for mutations to accumulate enough to cause cytotoxicity, and S6G is known to interrupt replication after one cycle. An explanation that better fits the time frame is the possibility that repetitive incorporation and excision events occur during replication due to the proofreading functions of replicative DNA polymerases. This has been termed ‘futile cycling’, and has recently been shown to occur in the case of O6meG (47). It is reasonable to assume that S6meG could be subject to the same effect. S6G itself is a poor template in vitro for polymerization and ligation reactions (11) and may also be subject, even unmethylated, to futile cycling.

After acceptance of this manuscript, an article reporting on the structure and dynamics of duplex DNA containing a single S6G-C base pair was published (48). Our observations are in complete agreement with this paper.

SUPPLEMENTARY MATERIAL

Supplementary Material is available at NAR Online.

[Supplementary Material]
nar_31_4_1331__index.html (1.1KB, html)

Acknowledgments

ACKNOWLEDGEMENTS

We thank Cecilia Torres for the synthesis and purification of oligodeoxynucleotides. This research was supported by NIH Grant CA47995 and CA77094.

PDS nos 1KB1 and 1KBM

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