MTX (mitoxantrone) is perhaps the most promising drug used in the treatment of various malignancies. Comprehensive literature on the therapeutics has indicated it to be the least toxic in its class, although its mechanism of action is still not well defined. In the present study, we have evaluated the associated binding interactions of MTX with naked DNA. The mechanism of MTX binding with DNA was elucidated by steady-state fluorescence and a static-type quenching mechanism is suggested for this interaction. Thermodynamic parameters from van 't Hoff plots showed that the interaction of these drugs with DNA is an entropically driven phenomenon. The binding mode was expounded by attenuance measurements and competitive binding of a known intercalator. Sequence specificity of these drug–DNA complexes was analysed by FTIR (Fourier-transform infrared) spectroscopy and molecular modelling studies. CD spectroscopy and the plasmid nicking assay showed that the binding of this drug with DNA results in structural and conformational perturbations. EMSA (electrophoretic mobility-shift assay) results showed that these drug–DNA complexes prevent the binding of octamer TF (transcription factor) to DNA. In summary, the study implicates MTX-induced conformational instability and transcription inhibition on DNA binding.
The ability of small molecules to interfere with transcription and DNA replication makes them a major target for drug interaction studies. Many natural or synthetic drugs serve as analogues of interacting entities in the research of protein–nucleic acid recognition and provide site-specific reagents for molecular biology. Therefore the analysis of drug–DNA interaction is imperative for understanding the molecular mechanisms of the drug action and designing specific DNA-targeted drugs . Apart from interacting with proteins that bind to DNA or through DNA–RNA hybrids, small aromatic ligand molecules may bind to DNA double helical structures. Interactions may occur by: (i) intercalating between stacked base-pairs, thereby distorting the DNA backbone conformation and interfering with DNA–protein interaction, or (ii) as minor groove binders.
The anthracyclines are one of the widely studied types of such drugs , owing to their notable clinical efficacy against a wide range of malignancies . Among various anthracycline antibiotics, MTX (mitoxantrone) is regarded as the most promising anticancer drug available. It has major clinical value over other members of this group as MTX is an aglyconic anthracycline (Figure 1) reported to have less cardiotoxic effects when compared with the naturally occurring anthracyclines, namely doxorubicin and daunorubicin . In multiple sclerosis, MTX has also shown potential therapeutic efficacy that is worthy of approval by the FDA (Food and Drug Administration) . Although the interaction of these drugs with a range of cellular constituents is well known, the specific cellular mechanisms involved and the ultimate cause of tumour cell death are still to be elucidated. The drug has a high affinity for DNA, thus providing the driving force for further nuclear uptake. It is well known that intercalation is the most established form of interaction for naturally occurring anthracyclines and there is an extensive body of evidence to show that one of the first cellular responses is the impairment of topoisomerase II activity [6,7]. However, on reductive activation, the drug can also bind covalently to DNA . Some covalently binding agents have also been shown previously to disrupt the binding of TFs (transcription factors) to their specific consensus sequences . If these adducts prevent binding of TFs to DNA in tumour cells, then the sequence selectivity of the particular drug will determine on which gene TF assembly will be inhibited. In the present study, we have evaluated the temperature-dependent binding of these drugs and the thermodynamic parameters for this interaction. The effect of binding on the conformation of the DNA was also assessed. The sequence specificity of these drugs and drug–DNA adduct-induced transcription inhibition was also analysed.
Structure of MTX, an anthracycline antibiotic
MATERIALS AND METHODS
Highly polymerized type I calf thymus DNA sodium salt (7% Na content) and Hepes buffer salt were purchased from Sigma Chemical Co. and was deproteinated by the addition of chloroform and 3-methylbutan-1-ol to an NaCl solution. MTX was procured from Sigma–Aldrich Chemicals and the stock solution was prepared in water. Other chemicals were of reagent grade and used without further purification.
Preparation of stock solutions
Calf thymus DNA was dissolved to 0.5% (w/w) in 0.1 M Hepes buffer (pH 7.4) at 298 K for 24 h with occasional stirring to ensure the formation of a homogeneous solution. The purity of the DNA solution was checked from the absorbance ratio A260/A280. Since the attenuance ratio of the above purified DNA lies in the range 1.8<A260/A280<1.9, no further purification was needed. The concentration of DNA was spectroscopically determined using a molar absorption coefficient (ε) of 13200 M−1·cm−1 at 260 nm  and the molarity of the DNA solution was derived with reference to base-pairs. The stock solution of the drug with 10 mM concentration was also prepared.
Fluorescence measurements were made on a Shimadzu spectrofluorimeter, namely model RF-5301PC (Shimadzu), equipped with a 150 W xenon lamp. The fluorescence quenching of the drug at increasing molar ratios of DNA was recorded in the wavelength range 640–740 nm after exciting the MTX solution at 610 nm, using 3 nm/3 nm as slit widths. The drug concentration was fixed at 50 μM unless otherwise indicated, and the DNA concentration was varied from 5 to 55 μM. To evaluate the effect of temperature on drug–DNA interaction, fluorescence was recorded at four different temperatures, i.e. 298, 300, 306 and 310 K. The instrument was thermostatically controlled by a Neslab RTE-110 circulating water bath.
The stoichiometry of the interaction of the drug with DNA was determined by the method of continuous variations . The fluorescence change (ΔF=Ffluor–Ffluor+DNA) of a series of protein–drug mixtures was done, keeping the molarity of the mixture constant while varying the mole fraction of each.
The UV measurements of calf thymus DNA were recorded on a Shimadzu double beam spectrophotometer (model UV1700) by using a cuvette of 1 cm path length. The absorbance values of DNA in the absence and presence of complex were recorded in the range of 240–300 nm. The DNA concentration was fixed at 0.1 mM, while the compound was added in increasing concentrations.
FTIR (Fourier-transform infrared)
Infrared spectra of the nucleic acid solution were recorded on a Nicolet Magna 750FT-IR spectrophotometer (DTGS detector, Ni-chrome source and KBr beam splitter) with a resolution of 4 cm−1 and 60 scans.
Spectra processing procedures
Spectra of the sample solution and the buffer solution were collected using the same condition. Then the absorbance of the buffer solution was subtracted from the spectra of the sample solution to get the FTIR spectra of DNA.
The crystal structure of B-DNA used for docking was extracted from the structure deposited in the PDB (PDB ID: 453D) . The three-dimensional structure of the drug was generated in Sybyl 6 (Tripos) and its energy-minimized conformation was obtained with the help of the CHARMM (Chemistry at HARvard Macromolecular Mechanics) program . Water molecules were removed from the DNA PDB file. Polar hydrogen atoms and Gasteiger charges were added to prepare the DNA molecule for docking. Rotatable bonds in the ligands were assigned with the Auto Dock Tools in AutoDock [13a]. Ligand docking was carried out with AutoDock v4.0, Lamarckian GA (genetic algorithm). DNA was enclosed in the grid defined by Auto Grid having 0.375 Å spacing. Other miscellaneous parameters were assigned the default values given by the AutoDock program.
CD measurements of DNA in the presence and absence of the drug was made in the range of 220–320 nm on a Jasco-J820 spectropolarimeter coupled to a microcomputer using a quartz cell of 0.1 cm path length. All of the spectra were recorded at 298 K and the temperature was maintained by a thermostatically controlled Neslab RTE-110 circulating water bath. A stock solution of 150 μM DNA was prepared in 0.1 M Hepes buffer. The molar ratio of DNA concentration to drug concentration was 1:0, 1:2 and 1:4 for CD measurements.
Plasmid nicking assay
CCC (covalently closed circular) plasmid pBR322 DNA (0.25 μg) in a final volume of 30 μl was treated with varying concentrations of the drug and exposed to white light for 2 h. To this, 6 ml of 5× tracking dye [40 mM EDTA, 0.05% Bromophenol Blue and 50% (v/v) glycerol] was added and loaded on to 0.8% agarose gels. The gel was run at 50 mA for 3 h and stained with ethidium bromide (EtBr; 0.5 μg/ml) for 30 min at 4 °C.
Nuclear extract preparation
Extracts were prepared as described by Schreiber et al. . HEK-293T cells [HEK-293 (human embryonic kidney) cells expressing the large T-antigen of SV40 (simian virus 40); 5×106] at 70% confluency were washed twice with ice-cold PBS and resuspended in 10 mM Hepes (pH 7.4), 10 mM KCl, 0.1 mM EDTA, 0.1 mM EGTA, 1 mM DTT (dithiothreitol), 1 mM PMSF, 2 mM pepstatin A, 0.6 mM leupeptin, 1 mg/ml aprotinin and 0.6% Nonidet P40. The nuclei pellet was recovered after centrifugation at 1200 g for 10 min at 4 °C and resuspended in 20 mM Hepes (pH 7.9), 0.4 M NaCl, 1 mM EDTA and 1 mM EGTA. Aliquots were then incubated at 277 K for 30 min and centrifuged at 21000 g for 5 min at 4 °C and the supernatant containing nuclear proteins was removed. Protein concentration was determined by the Bradford method .
EMSA (electrophoretic mobility-shift assay)
The EMSA was performed by using the double-stranded synthetic oligonucleotides harbouring the octamer consensus motif ATGCAAAT (GATCCTAGCCCCCTCTATGCAAATGAGAAGCATTCCTTT). The synthetic oligonucleotides were end-labelled using 20 μCi of [γ-32P]dATP and 10 units of T4 polynucleotide kinase (MBI Fermentas). Drug–DNA reactions were set up with 0.1–6.0 μM of the tested drug and 10 nM of 32P-labelled oligonucleotides in a buffer containing 20 mM Hepes, 5 mM DTT, 7.5 mM MgCl2 and 5% glycerol. Reactions were incubated for up to 9 h at 0 °C. Then nuclear extracts were added to the reaction mixture along with the non-specific competitor poly(dI-dC)·(dI-dC)  and the binding reaction was further incubated for 20 min at room temperature (25 °C); then the samples were electrophoresed at constant voltage (200 V) under low ionic strength conditions (0.25 M Tris/acetate/EDTA buffer) on 6% (v/v) polyacrylamide gels. Gels were then dried and subjected to a phosphoimager (Typhoon 8600; Amersham Biosciences) for sensitive detection.
RESULTS AND DISCUSSION
Binding of MTX with DNA
The interaction of MTX with DNA was examined by monitoring the changes in the intrinsic fluorescence of the drug at varying concentrations of the DNA. The fluorescent emission spectra of the drugs as well as the effect of varying DNA concentrations on the fluorescence emission spectra were illustrated in Figure 2(A). MTX exhibited an emission maximum at 610 nm (Figure 2A, curve a). The fluorescence emission was gradually decreased with increasing concentrations of DNA, showing that the drug fluorescence was efficiently quenched on binding to DNA. Since the highest change in the drug fluorescence spectra on addition of DNA occurred at a wavelength of 675 nm, the fluorescence measurements at this wavelength yielded an accurate determination of the bound fraction. When the concentration of DNA was up to 25×10−6 M, the fluorescence intensity of MTX was almost completely quenched and a slight shift in its peak maxima was observed, indicating the saturation in binding. The red shift indicates that on binding to DNA the ring substituents on the fluorophore could slide into the base-pairs, illustrating that they were in an environment in which they were unable to hydrogen-bond with the solvent water molecules. The red shift observed at lower concentrations of DNA also indicated the higher affinity in the DNA–MTX complex. The Stern–Volmer quenching plot from the fluorescence titration results is shown in Figure 2(B). The fluorescence quenching constant (KSV) was evaluated using the Stern–Volmer equation :
where F0 and F are the fluorescence intensities in the absence and presence of DNA respectively and KSV is the Stern–Volmer quenching constant, which is a measure of the efficiency of quenching by DNA. The titration results were used to construct a plot of F0/F against [DNA]. KSV obtained from the slope of the linear line was 0.45×105 litres/mol. The Stern–Volmer plot is linear, indicating that only one type of quenching process occurs, either static or dynamic quenching . The procedure of quenching was further confirmed from the values of the bimolecular quenching rate constants, Kq, which are evaluated using the equation:
where τ0 is the lifetime of the protein without the quencher. Since fluorescence lifetimes are typically near 10−8 s, the bimolecular quenching constant (Kq) was calculated from the above equation and was found to be of the order of 1013 mol−1·s−1. By considering the equivalency of the bimolecular quenching constants, it can be seen that the latter is greater than the largest possible value (1×1010 mol−1·s−1)  in aqueous medium. Thus the fluorescence quenching is not initiated by a dynamic process but is rather due to a static process with ground state complex formation. The quenching procedure was further confirmed by the temperature dependence of quenching. Studying the temperature dependence pattern of quenching parameters can differentiate between dynamic and static quenching. The KSV values decrease with an increase in temperature for static quenching and the reverse will be observed for dynamic quenching. The trend in the present study (Table 1) indicates that the probable quenching mechanism of MTX fluorescence by DNA is a static type. Our results also corroborate with the earlier work on anthraquinone analogues .
Binding analysis of MTX with DNA
|T (K) .||KSV (×104) (litre/mol) .||Kq (×1012) (litre/mol per s) .||K (×105) (mol−1) .||N .||r2 .|
|T (K) .||KSV (×104) (litre/mol) .||Kq (×1012) (litre/mol per s) .||K (×105) (mol−1) .||N .||r2 .|
When ligand molecules bind independently to a set of equivalent sites on a macromolecule, the equilibrium between free and bound molecules is given by the equation :
where K and n are the binding constant and the number of binding sites respectively. Thus a plot of log(F0–F)/F against log[Q] (Figure 2C) can be used to determine K as well as n. The values of K and n at 298, 300, 306 and 310 K (Table 1) suggest the destabilization of the MTX–DNA complex with an increase in temperature.
Determination of the stoichiometry of the DNA–drug complexes was done using the Job's plot or the mole-ratio method. In this method, DNA and drug solutions in equimolar concentrations were mixed at different amounts such that the total number of moles of reactants in each mixture was kept constant. The molar ratio of reactants was varied across the set of mixtures. The fluorescence of each solution was then measured at the emission maxima of the drug, which was plotted against the mole fraction of one of the reactants (the drug in this case). The breakpoint in the plot corresponds to the mole fraction of the drug in the DNA–drug complex and thereby demonstrates the binding stoichiometry. The analysis and interpretation are simpler if, as in the present case, the DNA molecule has no fluorophore in the range of wavelengths scanned here. The relative fluorescence observed is directly proportional to the drug concentration present in the complex. As evident from the Job's plot (Figure 2D), a clear break-even point implies that the stoichiometric ratio of MTX/DNA at 298 K and pH 7.4 is 1:1.
Determination of thermodynamic parameters
The thermodynamic parameters ΔH and ΔS were evaluated from the following equation:
where R and T are the universal gas constant and absolute temperature respectively. The K values were determined at four different temperatures and a van 't Hoff plot of lnK against 1/T was plotted. The ΔH and ΔS values were obtained from the slope (–ΔH/R) and intercept (ΔS/R) of the van 't Hoff plot. Knowing these two values, ΔG° was calculated from the following standard equation :
For the MTX–DNA complex, ΔG° values were calculated at all four temperatures. Table 2 shows the thermodynamic parameters of the MTX–DNA interaction. The negative ΔG° values showing favourable binding reaction and temperature-independent ΔH° and ΔS° have been reported by earlier drug–ligand studies [19,20]. However, for the association, the binding enthalpy and ΔS° for MTX–DNA show that the binding reaction is entropically driven. For entropically driven drug–DNA interactions, the solvent molecules and the counter ions are expelled from the bound surface of both the interacting partners when they bind to each other, leading to entropic gain. As reported, the positive values of both ΔH° and ΔS° implies a typical hydrophobic interaction, whereas specific electrostatic interaction between ionic species in an aqueous solution is characterized by positive ΔS° and negative ΔH° values. Moreover, negative values of both ΔH° and ΔS° arise from van der Waals forces and hydrogen-bonding formation in low dielectic medium . Hence, the negative values of ΔH° and ΔS° in the present case showed that both van der Waals forces and hydrogen bonding play a major role in the binding of MTX to DNA.
|T (K) .||ΔG° (kJ/mol) .||ΔH° (kJ/mol) .||ΔS° (J/mol per K) .|
|T (K) .||ΔG° (kJ/mol) .||ΔH° (kJ/mol) .||ΔS° (J/mol per K) .|
UV–visible absorption studies were performed to further ascertain the interaction of DNA with MTX. Figure 3(A) shows the difference absorption spectra of DNA in the absence and presence of different concentrations of the drug. The maximum absorption of DNA was located at approx. 260 nm. It is well established that the absorption peak of the double helical DNA occurs at this wavelength. The UV attenuance showed a decrease in absorption bands with the increase in drug concentration (Figure 3A). The decrease in the difference spectra of the DNA on drug titration is indicative of complex formation between DNA and the drug. The increase of MTX concentration in the DNA at 260 nm exhibited hypochromism of 59% and bathochromism of 3 nm. This hypochromic effect is thought to be due to the interaction between the electronic states of the intercalating chromophore and those of the DNA bases increased . It is expected that the strength of this electronic interaction would decrease as the cube of the distance of separation between the chromophore and the DNA bases . So, obviously the large hypochromism observed in our experiments suggested the close proximity of the drug chromophore to the DNA bases. In addition, at a high concentration of DNA, a red shift in absorption maxima by 20 nm was observed. After interaction with the base-pairs of DNA, the π–π* orbital of the bound ligand can couple with the π orbital of the base-pairs, due to the decreased π–π* transition energy, which results in a bathochromic shift . This phenomenon indicated that upon binding to DNA the ring substituents on the chromophore could slide into the base-pairs; as a result they would be in an environment in which they were unable to form hydrogen bonds with the solvent water molecules. These changes are suggestive of the intercalative mode of binding.
Binding mode of MTX with DNA
Competitive binding of the drugs and EtBr with DNA
In another set of experiments, we further authenticate the intercalation of the drug by planning a competitive displacement experiment. EtBr, a typical intercalator, was used to compete with MTX. EtBr is an intercalator that approaches the DNA backbone via the minor groove . Figure 3(B) shows three fluorescence emission spectra of EtBr with different treatments. The first spectrum is 50 μM DNA with saturating EtBr. The second spectrum is 50 μM DNA+25 μM drug to which same concentration of EtBr was added as in the case of the first spectrum after 1 h incubation with the drug. The third spectrum is 50 μM DNA to which an identical concentration of EtBr is added as for the second spectrum followed by the addition of 25 μM of the drug. It is evident there are significant changes in the fluorescence spectra when the drug was added to DNA either before or after the addition of EtBr (Figure 3B). This shows that EtBr can be removed by the drug to a larger extent which indicates that the drug strongly competes with EtBr to bind to the DNA backbone. Putting together the results obtained from the UV absorption study and that of competitive binding, one can predict a possible intercalative mode of binding of MTX with the DNA.
Sequence specificity of drug–DNA binding
The spectral changes (intensity and shifting) of several prominent DNA signals in-plane vibrations at 1717 cm−1 (G, T), 1663 cm−1 (T, G, A and C), 1609 cm−1 (A, C), 1492 cm−1 (C, G) and 1222 cm−1 (PO2 asymmetric stretch) [25–27], were monitored at different drug concentrations by employing FTIR spectroscopy. Our results revealed an intercalation mode of binding as the minor intensity of DNA increases in-plane vibrations at 1717 (G), 1663 (T), 1609 (A) and 1222 cm−1 (PO2 asymmetric stretch) on MTX interaction (Figure 4A). The major shifting of the band at 1717 cm−1 (G) to 1698 and 1726 cm−1 on MTX treatment is indicative of drug intercalation mainly into the G–C base-pairs (Figure 4A). However, the thymine band at 1663 cm−1 exhibited no shifting on drug complexation. The adenine band at 1609 cm−1 showed an increase in absorbance with almost no shift in the peak because of which it was difficult to draw a certain conclusion on the nature of drug interaction with A–T bases (Figure 4A). At high drug concentrations, a major increase in the intensity of the guanine band at 1717 cm−1 was observed accompanied by the shifting of this vibration, which is indicative of MTX binding to guanine bases (Figure 4A). This illustrates the higher affinity of these drugs with G–C bases. Furthermore, the shifting of the PO2 asymmetric band at 1222–1224 cm−1 with an increase in intensity of this vibration illustrates the interaction of this drug with the backbone PO2 groups, implicating its binding near the phosphate backbone. The major increase in the guanine and cytosine vibrations and their respective shifts illustrates the higher affinity of the drug to interact with them.
Sequence specificity on DNA–MTX intraction
Modelling of MTX binding to DNA
Docking studies provide some insight into the interactions between the macromolecule and the ligand, which can corroborate the experimental results. Although the crystal structure of the complex can represent specific details of the interactions, general observations may be obtained from docking studies. The ligand has been made flexible to attain different conformations in order to predict the best fit orientation, and the best energy docked structure was analysed. The docked structure as shown in Figure 4(B) suggests that MTX could bind DNA by interacting with the bases and the phosphate backbone. The molecular modelling-predicted lowest energy conformation was the binding of MTX at the major groove. This could be explained as follows: in the major groove of double-stranded DNA, the guanine- and adenine-N-7 atoms are the more reactive sites, because they are free from strong steric hindrances and are not involved in the Watson–Crick hydrogen bonds. The binding site of the drug is predicted to be three base-pairs long. Figure 4(B) also illustrates the interaction of the aromatic ring and the side chains of the drug moiety with the base-pairs and the phosphate backbone of DNA, which helps to provide better ‘anchoring’ and stabilization of the drug–DNA adduct. The analysis of the docking results further revealed the binding of the drug at the G–C-rich region. This corroborates with our previous results of higher G–C affinity of the drug.
A CD spectrum is a sensitive reporter of any alteration in the DNA backbone. We have therefore used CD spectroscopy to identify the backbone distortions in DNA obtained on binding of the drug. Figure 5(A) shows the CD spectrum of 35 μM DNA alone (curve a) and with two different concentrations of MTX (curves b and c). As the concentration of the drug was increased, there was a very distinct change in the CD spectrum of the B-form DNA. It is important to note that the prominent changes reported in the present study were obtained at intermediate DNA or drug concentrations. At this drug concentration the maximum proportion of the DNA population is that of the bound DNA or vice versa, which is not the case with either high DNA or drug concentrations. On increasing the concentration of DNA or the drug, the CD contribution coming from free or unbound components masks the changes obtained in the CD spectra on the DNA–drug complex formation. Hence, the CD results presented exhibit substantial changes in the ellipticity values, which clearly suggested structural alterations in the DNA. Thus, in addition to surface binding of this drug on DNA, the intercalation of MTX may induce substantial conformational changes in DNA structure. As the drug alone does not exhibit any CD signal in this range, it was easier to interpret the results.
Structural alteration in DNA on MTX binding
Plasmid nicking assay
Figure 5(B) shows the agarose-gel electrophoretic pattern of MTX-treated plasmid DNA (pBR322). Treatment of supercoiled DNA with the drug resulted in complete transformation to the relaxed open form. A noticeable reduction in the intensity of the band corresponding to the CCC form of plasmid DNA occurred with a concomitant increase in the amount of nicked species. These transformations of the CCC structure into nicked or relaxed species with increasing drug concentration clearly suggest strand scission of DNA, resulting in structural perturbation of the nucleic acids.
The above two experiments suggest structural and conformational perturbation of DNA on interaction with MTX. These changes in DNA structure can be translated as a hindrance in the specific structure–activity relationship, which is a prerequisite for many vital processes.
Effect of drugs on transcription complex formation
The ability of this drug to inhibit TF–DNA complex formation was analysed by using EMSA. The octamer-binding element is important for promoter activation mediated through the generally expressed Oct-1 protein, such as for the histone H2B and snRNA (small nuclear RNA) genes. The octamer motif is also a specific binding site for octamer factors  encoded by the Brn-2 POU domain gene. Hence, we chose to use an octamer consensus motif for EMSA. Initial studies were performed to establish whether MTX binding would prevent octamer protein binding to their consensus motif. The crystal structure of Oct-1 protein bound to ATGCAAT revealed major contacts of the specific domain of the protein directly to the GC site within the sequence . The predicted specificity of the drug from the above experiments, supported by the tetracyclic structure of this drug, suggests that binding of this drug at the GC site would pose a direct blockage to TF–DNA interaction. Figure 6 showed MTX induced concentration-dependent inhibition of octamer binding with the consensus motif. To avoid non-specific artefacts, the binding reaction was carried out in the presence of the non-specific competitor poly(dI-dC)·(dI-dC). Overall, the inhibition of binding of octamer proteins correlates well with the formation of characteristic anthracycline-induced adducts . Although these trends are quite clear and highly reproducible in quadruplicate, there was some variation in the absolute level of formation of octamer protein–DNA complexes each time, but that could be the result of the varying concentration of TFs present in individual nuclear extracts . The MTX GC cross-link site in the motif may play a crucial role in the binding mode of the octamer protein, as the arginine present in the protein domain will form a hydrogen bond with O-6 of guanine and will also make contact with O-6 and N-7 positions of guanine on the opposite strand. Overall, the results from the present study indicate that MTX binding to DNA does have a potential inhibitory role in vivo with respect to protein binding to sequences containing 5′-GC sites. Because the octamer-binding proteins are able to recognize a variety of motifs with varying affinities of binding, it is likely that selective drug-induced inhibition of some octamer proteins would also arise in vivo. It is questionable whether the extent of inhibition of octamer protein binding facilitated by MTX would be significant enough to lead to detectably altered regulation patterns of specific cellular proteins, but it is possible that the effects of adducts could be further magnified when the complex interactions of proteins required in transcription initiation are considered.
Gel retardation assay of inhibition of binding by octamer proteins with increasing concentrations of MTX
In summary, the present paper demonstrates the hitherto unknown interaction of MTX directly with DNA. The association occurs in DNA with GC specificity. It is now important to establish whether there is a correlation between MTX treatment and selective targeting of genes that are high in GC content. MTX binding led to conformational transitions in the DNA structure. Although MTX has been a major inclusion in chemotherapy regimes for years and appears to act as a topoisomerase II inhibitor, a major research effort is still required that is directed toward elucidating its basic modes of action in tumour tissue. Unlike other earlier spectroscopic studies, most of the present observations are in aqueous medium under physiological conditions. The observed significant binding of MTX to DNA at 105 M−1 has possible clinical implications due to the recommendation of a high dosage range for MTX  unlike other anthracyclines, which led to its high availability within the tissues. Furthermore, our results refer to the number of base-pairs; hence if we consider n number of G–C base-pairs, then it will amplify the effect; within this purview, the results appear to be significant, assuming the biological importance of the present study in the context of its action and pharmacokinetics of the drug.
This research received no specific grants from any funding agency in the public, commercial or not-for-profit sectors.