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Originally published In Press as doi:10.1074/jbc.M406053200 on July 28, 2004

J. Biol. Chem., Vol. 279, Issue 40, 42041-42054, October 1, 2004
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Structural Analysis of DNA Interactions with Biogenic Polyamines and Cobalt(III)hexamine Studied by Fourier Transform Infrared and Capillary Electrophoresis*

Amin Ahmed Ouameur and Heidar-Ali Tajmir-Riahi{ddagger}

From the Department of Chemistry-Biology, University of Québec at Trois-Rivières, C.P. 500, Trois-Rivières, Québec G9A 5H7, Canada

Received for publication, June 1, 2004 , and in revised form, July 26, 2004.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Biogenic polyamines, such as putrescine, spermidine, and spermine are small organic polycations involved in numerous diverse biological processes. These compounds play an important role in nucleic acid function due to their binding to DNA and RNA. It has been shown that biogenic polyamines cause DNA condensation and aggregation similar to that of inorganic cobalt(III)hexamine cation, which has the ability to induce DNA conformational changes. However, the nature of the polyamine·DNA binding at the molecular level is not clearly established and is the subject of much controversy. In the present study the effects of spermine, spermidine, putrescine, and cobalt(III)hexamine on the solution structure of calf-thymus DNA were investigated using affinity capillary electrophoresis, Fourier transform infrared, and circular dichroism spectroscopic methods. At low polycation concentrations, putrescine binds preferentially through the minor and major grooves of double strand DNA, whereas spermine, spermidine, and cobalt(III)hexamine bind to the major groove. At high polycation concentrations, putrescine interaction with the bases is weak, whereas strong base binding occurred for spermidine in the major and minor grooves of DNA duplex. However, major groove binding is preferred by spermine and cobalt(III)hexamine cations. Electrostatic attractions between polycation and the backbone phosphate group were also observed. No major alterations of B-DNA were observed for biogenic polyamines, whereas cobalt(III)hexamine induced a partial B -> A transition. DNA condensation was also observed for cobalt(III)hexamine cation, whereas organic polyamines induced duplex stabilization. The binding constants calculated for biogenic polyamines are KSpm = 2.3 x 105 M-1, KSpd = 1.4 x 105 M-1, and KPut = 1.02 x 105 M-1. Two binding constants have been found for cobalt(III)hexamine with K1 = 1.8 x 105 M-1 and K2 = 9.2 x 104 M-1. The Hill coefficients indicate a positive cooperativity binding for biogenic polyamines and a negative cooperativity for cobalt(III)hexamine.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
The biogenic polyamines putrescine [NH2(CH2)4NH2], spermidine [NH2(CH2)4NH(CH2)3NH2], and spermine [NH2(CH2)3NH(CH2)4NH(CH2)3NH2] (Structure 1) are the most prevalent polyamines in mammalian cells. They are small aliphatic polycations involved in numerous diverse biological processes, such as the ability to modulate gene expression and enzyme activities, activation of DNA synthesis, cell proliferation and differentiation, and others (1-8). Other functions of polyamines have been related to DNA protection from external agents (9, 10) and against radiation damage (11, 12). In general, spermidine and spermine are present in millimolar concentrations in vivo, whereas putrescine levels are slightly lower (13-15). However, the most important characteristic of polyamines within the cells is to bind nucleic acids and DNA in particular (1, 6).



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  		STRUCTURE 1.
Chemical structures of spermine, spermidine, putrescine, and cobalt(III)hexamine. Carbon, nitrogen, and hydrogen atoms are shown in cyan, blue, and white colors, respectively.

 
Polyamines also have the ability to induce DNA conformational transitions. Their bindings have been reported to promote the conversion of the right-handed to a left-handed ZDNA (16-19) or to an alternative form of right-handed helix, A-DNA (20-22). However, polyamines can also bind B-DNA, thus inducing no major biopolymer conformational changes (23-25).

Another consequence of polyamine binding is the condensation of DNA occurring with both naked DNA (26-30) and chromatin (31, 32). Immunocytochemical studies of spermidine and spermine have shown that these polyamines are associated with highly compacted mitotic chromosomes (33, 34), inducing more stabilizing than regulating effects on the chromatin structure during the cell cycle (35). On the other hand, it has been shown that cobalt(III)hexamine cation is five times more efficient as a condensing agent than spermidine, having the same positive charges (36). These studies indicate that polyamine-induced DNA condensation is important to the cellular functions in vivo. However, the question of how they bind to DNA has not been clearly established.

One of the early discoveries about polyamine·DNA interactions is the observation that polycation could stabilize double-stranded DNA (37). Putrescine, spermidine, and spermine can increase the melting temperature (Tm) of DNA in a concentration-dependent manner by as much as 40 °C in low salt buffer solution, compared with that in the absence of polyamines (37, 38). Investigating on the mechanisms of this effect provides support for both electrostatic interaction between the polyamine amino-protonated groups and the negatively charged backbone group (38-40), as well as a direct interaction with DNA bases (21, 41).

Both experimental and theoretical methods have been used to determine the binding positions of polyamines on B-form DNA. These studies have concentrated mainly on spermine cation, whereas a few structural details about the binding of the other polyamines to DNA are known. Rather contradictory views concerning the binding sites of spermine and other polyamines have emerged. X-ray diffraction from crystals of B-form dodecamer revealed a spermine bound into the major groove of DNA duplex (42). In photoaffinity cleavage experiments (43), the binding position of spermine in solution appeared to be in the minor groove of B-DNA. From the NMR study of polyamine with DNA dodecamer, it has been proposed that the spermine cation appeared to be mobile (44). This study suggested a rapid diffusion of the spermine along the DNA duplex with specific tight binding sites or delocalized binding with no discrete sites. Feuerstein et al. (45) found that spermine prefers the major groove of B-DNA followed by minor groove interaction with phosphate binding being the least favorable. Different theoretical studies (46, 47) agreed with this hypothesis of spermine cation interacting with the dsDNA bases across the major groove. However, recent molecular mechanics simulation studies (48-50) found that spermine occupies more varied sites, including binding along the backbone and bridging both the major and minor grooves. On the other hand, recent Raman study (24) found that putrescine and spermidine prefer the minor groove of B-DNA, whereas spermine binds to the major groove.

Thus, the binding positions of spermine and other polyamines on B-DNA have not been firmly resolved, and it is probable that both the base sequence and the environment have an important influence on polycation binding. In the present work, we have studied the interactions between calf-thymus DNA and biogenic polyamines as well as cobalt(III)hexamine cation using affinity capillary electrophoresis, Fourier transform infrared, and circular dichroism spectroscopic methods. Evidence for DNA condensation and helix stabilization is provided. Furthermore, the influence of polyamine charge and concentration on DNA structural changes has been discussed, and the presence of specific base-polyamine binding is reported.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Materials—Polyamines spermine, spermidine, putrescine, and cobalt(III)hexamine were purchased from Sigma. Highly polymerized type I calf-thymus DNA sodium salt (7% sodium content) was purchased from Sigma, and deproteinated by the addition of CHCl3 and isoamyl alcohol in NaCl solution. To check the protein content of DNA solution, the absorbance at 260 and 280 nm was recorded. The A260/A280 ratio was 1.85, showing that the DNA was sufficiently free of protein (51). Other chemicals were of reagent grade and used without further purification.

Preparation of Stock Solutions—Sodium-DNA (8.3 mg/ml) was dissolved in 50 mM NaCl (pH 7.20) at 5 °C for 24 h with occasional stirring to ensure the formation of a homogeneous solution. The final concentration of the stock calf-thymus DNA solution was determined spectrophotometrically at 260 nm using a molar extinction coefficient {epsilon}260 = 6600 cm-1 M-1 (expressed as the molarity of phosphate groups) (52, 53). The UV absorbance at 260 nm of a diluted solution (1/250) of calf-thymus DNA used in our experiments was 0.661 (path length was 1 cm), and the final concentration of the stock DNA solution was 25 mM in DNA phosphate. The average length of the DNA molecules, estimated by gel electrophoresis, was 9000 bp (molecular mass ~ 6 x 106 Da). The appropriate amount of polyamines (0.3-25 mM) was prepared in distilled water and added dropwise to the DNA solution, to attain the desired polyamine/DNA(P) molar ratios (r) of 1/80, 1/40, 1/20, 1/10, 1/4, 1/2, and 1 at a final DNA concentration of 12.5 mM (4.15 mg/ml) for infrared measurements. For capillary electrophoresis, the polyamine/DNA(P) ratios were 1/800, 1/400, 1/200, 1/100, 1/50, 1/25, 1/12.5, and 1/6.25 with a final DNA concentration of 1.25 mM. The pH of the solutions was adjusted at 7.0 ± 0.2 with a pH meter ORION model 210A, using NaOH solution.

FTIR Spectra—Infrared spectra were recorded with a FTIR spectrometer (Impact 420 model) equipped with deuterated triglycine sulfate detector and KBr beam splitter, using AgBr windows. Spectra were recorded after 2-h incubation of polyamine with the polynucleotide solution and measured in triplicate (three individual samples of the same polynucleotide and polyamine concentrations). Interferograms were accumulated over the spectral range 400-4000 cm-1 with a nominal resolution of 2 cm-1 and a minimum of 100 scans. The water subtraction was carried out using 0.1 M NaCl solution at pH 7.0 ± 0.2 as a reference (54). A good water subtraction is considered to be achieved if there is a flat baseline around 2200 cm-1, where the water combination mode is located. This method yields a rough estimate of the subtraction scaling factor, but it removes the spectral features of water in a satisfactory way (54). The infrared spectra of polyamine·DNA complexes with molar ratios higher than 1/4 for spermine, spermidine, and cobalt-hexamine and higher than 1 for putrescine could not be recorded as a homogenous solution, due to DNA precipitation and solid gel formation.

The difference spectra [(DNA solution + polyamine) - (DNA solution)] were obtained using a sharp DNA band at 968 cm-1 as an internal reference. This band, which is due to deoxyribose C-C and C-O stretching vibrations, exhibits no spectral changes (shifting or intensity variation) upon polyamine·DNA complexation and cancelled out upon spectral subtraction (55). The spectra presented here were smoothed with a Savitzky-Golay procedure (54).

The plots of the relative intensity (R) of several peaks of DNA in-plane vibrations related to A-T, G-C base pairs and the stretching vibrations such as 1717 (guanine), 1663 (thymine), 1609 (adenine), 1492 (cytosine), and 1222 cm-1 ( groups), versus the polyamine concentrations were obtained after peak normalization using,

(Eq. 1)

where Ii is the intensity of absorption peak for pure DNA and DNA in the complex with i concentration of polyamine, and I968 is the intensity of the 968 cm-1 peak (internal reference).

Circular Dichroism Measurements—Spectra were recorded with a Jasco J-720 spectropolarimeter. For measurements in the near-UV region, a quartz cell with a path length of 0.1 cm was used. Five scans were accumulated at a scan speed of 50 nm/min, with data being collected at every nanometer from 200 to 320 nm. Sample temperature was maintained at 25 °C using a Neslab RTE-111 circulating water bath connected to the water-jacketed quartz cuvettes. The concentration of the calf-thymus DNA solutions was kept at 1.25 mM (0.4 mg/ml) in 25 mM phosphate buffer, pH 7.0. CD spectra of DNA·cobalt(III)hexamine complexes were recorded with molar ratios in the range of 0, 1/15, 1/10, 1/8, 1/6, and 1/4. Spectra were corrected for buffer signal, and conversion to the molar ellipticity [{theta}] was performed with Jasco Standard Analysis software.

Capillary Electrophoresis—A P/ACE System MDQ (Beckman) with photodiode array detector was used to study polyamine·DNA interaction. An uncoated fused silica capillary of 75-µm inner diameter (total length of 57 cm) and effective length of 50 cm (to the detector) was used. The capillary was conditioned each day by rinsing with 1 N sodium hydroxide for 30 min, followed by a 15-min wash with 0.1 N sodium hydroxide. Then it was rinsed with running buffer for 30 min at high pressure (50 p.s.i.), followed by a baseline run for 20 min at the voltage used for the experiments (25 kV). Between runs, the capillary was rinsed with NaOH 1 N for 2 min, followed by rinsing with running buffer for 3 min at high pressure. The capillary was flushed with distilled deionized water for 30 min at the end for each day and filled with deionized water overnight. Samples were injected using a voltage injection at 10 kV for 5 s. Electrophoresis was carried out at a voltage of 25 kV for 10 min using normal polarity. All runs were carried out at 25 °C. High voltage gave fast separation, and low current (the current in the capillary was typically 30 µA) gave less Joule heating, allowing the complex to stay intact during electrophoresis. The capillary inlet and outlet vials were replenished after every five runs. The polyamine binding experiments were performed in a sample buffer containing 20 mM, Tris-HCl, pH 7.0 ± 0.2, using constant concentration of calf-thymus DNA (1.25 mM) and variable concentrations of polyamines. The stock solutions of polyamines (2.5 mM) and DNA (2.5 mM) were prepared in the sample buffer. The solutions were mixed to attain polyamine/DNA(P) molar ratios of 1/800 to 1/6.25 in the presence of 1.25 mM DNA. Each sample was allowed to equilibrate for 30 min and tested with two separate runs for the same stock solution. The electropherograms were monitored at 260 nm and were collected and analyzed with the Beckman P/ACE Windows controller software.

Data Analysis—Affinity capillary electrophoresis (ACE) was used to detect a shift in mobility when polyamine binds to DNA. The binding constants for the polyamine·DNA complexes can be determined by Scatchard analysis using mobility shift of DNA complexes (56, 57). The extent of saturation (Rf) of the DNA was determined from the changes in the migration time of DNA in the presence of various concentrations of polyamine using the following equation,

(Eq. 2)

where m is the migration time of DNA measured for any added polyamine concentration, whereas mo and ms correspond to the migration time of pure and polyamine-saturated DNA, respectively.

The binding constant Kb, given by,

(Eq. 3)

was determined by fitting the experimental values of Rf and polyamine concentration to Equation 3.

(Eq. 4)

Rearrangement of this gives a convenient form for Scatchard analysis (Equation 4).

(Eq. 5)

Using the affinity capillary electrophoresis (ACE) method, many DNA·protein and metal·DNA complexes have been successfully identified (57-61). Because the charge-to-mass ratio of DNA·ligand complex is generally different from that of the unbound DNA and ligand separately, the complex migrates separately from the free ligand (59). ACE provides both qualitative and quantitative information on molecular interaction. Its scope includes the detection of complex formation, the identification of an active component for binding in a multicomponent mixture, the identification of structural requirements for recognition, the analysis of the equilibrium constant and stoichiometry for binding reactions, and concentration measurement, based on immunochemical recognition (62).

The cooperativity of the binding can be analyzed using a Hill plot. Assuming one binding site for polyamine (Equation 5), the equation below (Hill equation) can be established,

(Eq. 6)

where, n (Hill coefficient) measures the degree of cooperativity, and Kd is the dissociation constant. The linear plot of log(Rf/(1 - Rf)) versus log[polyamine] has a slope of n and an intercept on the log[polyamine] axis of log Kd/n (63). The quantity n increases with the degree of cooperativity of a reaction and thereby provides a convenient and simplistic characterization of a ligand-binding reaction (63).


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
Polyamines-DNA Complexes Studied by FTIR Spectroscopy—The FTIR spectral features of DNA·polyamine complexes between 1800 and 600 cm-1 are presented in Figs. 1 and 2. To avoid precipitation, different polycation concentrations were selected for the biogenic polyamines and cobalt(III)hexamine (26). Molar ratios of 1/80 to 1/4 were studied for spermine (+4), spermidine (+3), and cobalt(III)hexamine (+3) due to their high positive charges. However, molar ratios of 1/80 to 1 were used for putrescine (+2) with no precipitation under our experimental conditions. These ranges are physiologically justified, because millimolar concentrations of polyamines have been found in the nucleus of eukaryotic cells (1, 6). The relevant wavenumbers and assignment of the main infrared bands of the calf thymus DNA are listed in Table I.



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  		FIG. 1.
FTIR spectra in the region of 1800-600 cm-1 for pure DNA, free polyamine, and spermine·DNA (A) and spermidine·DNA (B) adducts in aqueous solution at pH 7.0 ± 0.2 (top three spectra) and difference spectra for polyamine·DNA adducts obtained at various polyamine/DNA(P) molar ratios (bottom two spectra).

 



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  		FIG. 2.
FTIR spectra in the region of 1800-600 cm-1 for pure DNA, free polyamine, and putrescine·DNA (A) and cobalt(III)hexamine·DNA (B) adducts in aqueous solution at pH 7.0 ± 0.2 (top three spectra) and difference spectra for polyamine·DNA adducts obtained at various polyamine/DNA(P) molar ratios (bottom two spectra).

 


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  		TABLE I
Measured wavenumbers, relative intensities, and assignments for the main infrared bands of calf-thymus DNA in aqueous solution at pH 7 ± 0.2

 
At low polyamine concentrations (r = 1/80), these four polyamines bind preferably to DNA bases. Evidence for this comes from the changes in the intensity and shifting of the absorption bands in the region 1800-1550 cm-1, due to the in-plane DNA vibrational frequencies (54, 55, 64-66). A major increase in the intensity was observed mainly for a guanine band at 1717 cm-1 (30-15%), thymine band at 1663 cm-1 (40-20%), and adenine band at 1609 cm-1 (40-15%) (Fig. 3). These intensity changes observed in the region 1800-1500 cm-1 were associated also with band shifts upon polycation interaction. In the spectra of Spm·DNA1 complexes, the bands at 1717 and 1609 cm-1 shifted downward by 4 cm-1, whereas the band at 1663 cm-1 shifted by 1 cm-1 (Fig. 1A). Similarly, the bands at 1663 and 1609 cm-1 were shifted by 1-2 cm-1 in the spectra of Spd·DNA, Put·DNA, and Co(III)·DNA adducts (Figs. 1B and 2). A similar trend was observed for the DNA band at 1717 cm-1, in-plane stretching vibrations of double bond of guanine (C7=N) located at the major groove (64, 66). This band shifted downward by 4 cm-1 in the spectra of Spd·DNA and Co(III)·DNA complexes, whereas in the Put·DNA adducts it was displaced by 2 cm-1 (Figs. 1B and 2). These spectral changes are supported by the difference spectra presented in Figs. 1 and 2. The positive features observed at 1715, 1665, and 1600 cm-1 in the difference spectra of Spm·DNA adduct (Fig. 1A) are related to an increase in the intensity of the guanine, thymine, and adenine bands upon polycation interaction as a result of cation bindings to the guanine-N7, thymine-O2, and adenine-N7 reactive sites. With respect to both intensity increase and shifting of the bands, we suggest that spermine interacts mainly with the guanine-N7 and adenine-N7 reactive sites located in the major groove of dsDNA. In the difference spectrum of spermidine (Fig. 1B), the positive features at 1705 and 1666 cm-1 are due to cation binding to the guanine-N7 and the thymine-O2 reactive sites. With respect to the shifting of the guanine band at 1717 cm-1 (4 cm-1), spermidine interaction occurs mainly with the guanine-N7 site followed by a weak interaction with A-T base pair.



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  		FIG. 3.
Intensity ratio variations for several DNA in-plane vibrations as a function of polyamine concentration. A-D, intensity ratios for the DNA bands at 1717 (guanine), 1663 (thymine), 1609 (adenine), 1492 (cytosine), and 1222 cm-1 ( vibration) for spermine, spermidine, putrescine, and cobalt(II)hexamine, respectively.

 

In the spectra of Put·DNA complexes, the bands at 1717 (G), 1663 (T), and 1609 cm-1 (A) were shifted downward by 2 cm-1 (Fig. 2A). These spectral changes together with the presence of positive features at 1710, 1662, and 1607 cm-1 in the difference spectra of Put·DNA complexes have been assigned to putrescine interactions with the guanine-N7, thymine-O2, and adenine-N7 reactive sites. In the case of Co(III)·DNA complexes, the difference spectra showed three positive bands at 1710, 1663, and 1602 cm-1 (Fig. 2B). Because a major displacement was observed for the band at 1717 cm-1 (4 cm-1) followed by 2 cm-1 for the band at 1609 cm-1 (A), we conclude that the cobalt(III)hexamine binds strongly to the guanine-N7 atom and to a lesser extent with the adenine-N7 reactive site. In addition, the shifting of the band at 1578 cm-1, which involves in-plane C8=N7 stretching vibration of the purine ring (mainly guanine residues) (64, 65) is indicative of a major cobalt(III)hexamine complexation with the guanine-N7 atom.

Other DNA vibrational frequencies in the region 1550-1250 cm-1 showed minor spectral changes upon polyamine complexation. The band at 1492 cm-1, which is related largely to the cytosine residues (65, 66), exhibited no major shifting (1 cm-1), and its relative intensity did not change significantly at different polycation concentrations (Figs. 1, 2, 3). Thus, the possibility of an interaction between cytosine and the polyamines cannot be included. It is worth mentioning that the weak positive features centered at 1482 cm-1 in the difference spectra of Spm·DNA and Spd·DNA complexes originating from polyamine methylene scissoring vibrations (67, 68) are not attributable to DNA vibrations (Fig. 1, A and B). Therefore, our results support the existence of hydrophobic interactions between the polyamine cations and DNA duplex.

To establish a possible interaction of polycation with the backbone phosphate group, the infrared spectra of dsDNA in the region 1250-1000 cm-1 were examined. The two strong absorption bands located at 1222 and 1088 cm-1 are assigned mainly to the asymmetric and symmetric stretching vibrations of the groups, respectively (64-66, 69, 70). An increase (10-15%) in the relative intensity of the absorption band (1222 cm-1) was observed with the shift of this vibration to 1225 cm-1 in all four polyamine·DNA complexes (Fig. 3, r = 1/80). The difference spectra of the polyamines provide clear evidence for an interaction between the groups of DNA and the polyamines, as deduced from the positive features appearing at 1230 and 1080 cm-1 (spermine), 1240 and 1099 cm-1 (spermidine), 1225 and 1088 cm-1 (putrescine), and 1243 and 1098 cm-1 (cobalt-hexamine) (Figs. 1 and 2, r = 1/80). These spectral changes reflect the interaction of polyamines with the oxygen atoms of the backbone phosphate groups.

At r = 1/40, a minor interaction of cobalt(III)hexamine with A-T base pair was observed. Evidence for this comes from the further increase in the intensity of the thymine band at 1663 cm-1 and adenine band at 1609 cm-1 (Fig. 3D). However, these two bands did not appreciably shift (1 cm-1) upon cobalt cation complexation (r = 1/40 to 1/10). This indicates a weak interaction of cobalt(III)hexamine with AT bases. However, the guanine band at 1717 cm-1 and the phosphate band at 1222 cm-1 were shifted by 3-4 cm-1 (spectra not shown). This evidence supports the preferential binding to the guanine-N7 and oxygen atom of groups by cobalt(III)hexamine cation. As the cobalt(III)hexamine concentration increased (r = 1/20), the intensities of the DNA vibrations at 1717 and 1222 cm-1 were also enhanced (Fig. 3D). These changes were accompanied by the shift of the two bands by 4 and 3 cm-1, respectively (spectra not shown), supporting a continued cation interaction with guanine-N7 site and the backbone group. Our findings are further supported by the x-ray study of B-DNA decamer d(AGGCATGCCT) (71), which identified two different binding sites for ; one with the guanine-N7 site in the major groove of 5'-AGG bases and another with the oxygen atom of the phosphate group. Similarly, the capillary electrophoresis results showed two major bindings for cobalt(III)hexamine·DNA complexes with K1 = 1.8 x 105 M-1 and K2 = 9.2 x 104 M-1, that can be assigned to the cation interactions with N7 of AG bases located in the major groove and the backbone groups, respectively.

Spermine strongly interacts with the guanine and adenine-N7 reactive sites as supported by the shifts of the bands at 1717 and 1609 cm-1 by 4 cm-1 together with a major increase of their relative intensities (Fig. 3A, r = 1/40 to 1/10). This evidence supports the preferential binding of spermine to the major groove via guanine and adenine bases, which continued until r = 1/20 for the guanine and adenine bases and until r = 1/10 for the guanine bases. As for the spermidine, similar behaviors were observed for the bands at 1717 and 1609 cm-1. In the spectra of spermidine·DNA complexes (r = 1/40), these two bands shifted by 4 and 2 cm-1, respectively. The above indicates the preferential bindings to the guanine and adenine-N7 reactive sites, located in the major groove, at low spermidine concentrations (r = 1/80 to 1/40). The thymine-O2 atom is the favored reactive site at the minor groove, because it is not involved in the Watson-Crick network of base pairing. At r = 1/20 to 1/4, the band at 1663 cm-1 (T) shifted downward by several wavenumbers (3 cm-1) upon spermidine addition (Fig. 1B, r = 1/4), which supports preferential binding with the minor groove in addition to its binding to the major groove at higher concentrations.

In the case of putrescine, the three major bands at 1717 (G), 1663 (T), and 1609 cm-1 (A) were shifted downward by 2 cm-1 (r = 1/40 to 1/10). Similarly, the relative intensities of these bands increased significantly (Fig. 3C), and this gives rise to the putrescine preferential bindings to the major and minor groove of the DNA duplex. As the concentration of putrescine increased further (r = 1/4 to 1), a decrease in the intensity of DNA vibrations at 1717, 1663, and 1609 cm-1 was observed (Fig. 3C). This decline in the intensity is the consequence of helix stabilization caused by a partial putrescine-phosphate interaction. Such decrease in infrared intensities of DNA vibrations is also observed when cation-phosphate binding resulted in partial helix stabilization (66). In our opinion, putrescine is more fluid than other polyamines at low and high concentrations and seems to bind at different sites, including the binding to the major and minor grooves, as well as the backbone phosphate group stabilizing the DNA duplex at high cation content. These findings are consistent with our capillary electrophoresis results that showed higher Hill coefficient (n) for putrescine compared with other polyamines as a result of a high cooperativity binding of putrescine to duplex DNA.

DNA Structural Changes at High Polycation Concentrations—At high spermine, spermidine, and cobalt(III)hexamine content (r = 1/4), several spectral changes were observed for the base and phosphate vibrations. Indeed, we observed a decrease in intensity of the absorption bands at 1717 (guanine-N7), 1663 (thymine-O2), 1609 (adenine-N7), and 1222 (phosphate) cm-1 (Fig. 3, A, B, and D). The term that has been adopted for this decrease is infrared hypochromism (67) and can be provoked by the base staking and base pairing as a consequence of the cation interaction (19, 24). In the case of the spermine and spermidine, this hypochromism is not arising from DNA condensation. In our opinion, this is due to DNA stabilization, which occurs at high polyamine concentrations just before the precipitation threshold. Similarly, the bands at 1717 and 1609 cm-1 in Spm·DNA and Spd·DNA spectra were shifted by several cm-1 (Fig. 1, A and B). The band at 1663 cm-1, which is assigned mainly to the C2=O2 stretching modes of the thymine residues, was shifted to the lower wavenumbers (1660 cm-1) in the spectra of Spd·DNA adducts (Fig. 1B, r = 1/4), whereas no major shifting was observed for this band in the spectra of Spm·DNA complexes (Fig. 1A).

These results are supported by the infrared difference spectra (Fig. 1). For spermine, the positive bands at 1715 and 1697 cm-1 have been assigned to the interaction with guanine-N7 atom, whereas the positive band at 1600 cm-1 was assigned to its interaction with the adenine-N7 atom. The interaction of Spm with thymine residues is not considered, because the corresponding band (1663 cm-1) exhibited no shifting. In the difference spectra of Spd·DNA complexes, the bands at 1660 and 1600 cm-1 have been assigned to the polycation interaction with thymine-O2 and adenine-N7 reactive sites. The interaction of putrescine at high concentration (r = 1) with DNA bases is evidenced by the positive difference features at 1715, 1660, and 1612 cm-1, that have been assigned to the guanine-N7, thymine-O2, and adenine-N7 reactive sites, respectively (Fig. 2A, r = 1). However, the infrared spectra of putrescine·DNA complex showed small wavenumber shifts (1 cm-1) for the three base bands (Fig. 2A). This indicates a weaker interaction of putrescine with these bases as the result of duplex stabilization.

The hypochromism effect observed for the base and phosphate vibrations at high cobalt(III)hexamine concentration (Fig. 2B) can be attributed to the formation of an aggregated state in solution in which polyanions (dsDNA) and polycations (cobalt(III)hexamine) dispose alternatively in a highly packaged structure similarly to a liquid crystal (27). This result is supported by the infrared difference spectra of Co(III)·DNA complexes (Fig. 2B, r = 1/4). As a general trend, the spectra exhibited negative features for the bases and the phosphate vibrations. The negative features at 1710 (base), 1200 (), and 1075 cm-1 () that emerged in the difference spectra are assigned to a reduced intensity caused by the DNA condensation. Similarly, the infrared spectra of Co(III)·DNA complex exhibited wavenumber shifts for the purine bands at 1717 cm-1 (guanine-N7) and 1609 cm-1 (adenine-N7) by 5 and 3 cm-1, respectively. This result gives rise to the preferential binding of the cobalt(III)hexamine with the major groove of the AG bases and the groups, in agreement with the precedent discussion for the stretching modes of the bases and phosphate groups.

The interpretation of the absorption bands shift in the region between 1550 and 1250 cm-1 of DNA complexed with biogenic polyamines is informative at high polycation concentrations. In this region, the IR spectra of spermine, spermidine, and putrescine show a pair of bands around 1480 and 1319 cm-1. In the spectra of Spm·DNA and Spd·DNA complexes, the former band is overlapped by the band at 1491 cm-1 involving in-plane ring vibration of cytosine residues (65, 66). These two bands are assigned to the polyamine methylene scissoring vibrations (67, 68). The spectral changes observed for these vibrations support the existence of a hydrophobic interaction between the polyamine cation and DNA duplex. Similarly, positive features at 1480 and 1334 cm-1 in the difference spectra of spermine and spermidine indicate hydrophobic contact with the methyl groups of dsDNA. The IR spectrum of putrescine showed a maximum band at 1570 cm-1 related to the N-H bending vibration of the groups (67, 68). At higher putrescine concentration, this band shifted to a lower frequency (1565 cm-1) with a positive feature at 1561 cm-1 in the difference spectra of the polycation·DNA adducts (Fig. 2A), indicating an active participation of these groups in polyamine complexation (Fig. 2A). The two positive peaks at 1492 and 1329 cm-1 are originating from methylene vibrations of putrescine, which support the existence of polycation hydrophobic interaction with DNA duplex. Further evidence of Co(III)·DNA interaction with DNA bases can be obtained from the shift of the bands in the region 1550-1250 cm-1. In the pure DNA, the band at 1529 cm-1 is assigned to the in-plane vibrations of cytosine and guanine residues (65, 72). This band disappeared in the spectra of the complex at high cobalt(III)hexamine concentration and was replaced by two weak bands at 1560 and 1450 cm-1 (Fig. 2B). These observed spectral changes reflect the interaction of cobalt cation with the G-C base pair.

The IR difference spectra of polyamine·DNA complexes at high polycation concentrations (r = 1/4) provide clear evidence for further cation interaction with the negatively charged groups, as deduced from the positive features appearing at 1230 and 1081 cm-1 (spermine), 1240 and 1099 cm-1 (spermidine), 1225 and 1087 cm-1 (putrescine), and 1245 cm-1 (cobalthexamine) (Figs. 1 and 2). The negative features observed at 1200 and 1075 cm-1 in the difference spectra of cobalt(III)hexamine are due to DNA condensation at high cobalt(III)hexamine concentrations. Similarly, the band at 1222 cm-1 was shifted by 2-3 cm-1 in the spectra of all polycation·DNA complexes (Figs. 1 and 2). The IR spectrum of the pure DNA in the region 1050-700 cm-1 is dominated by two intense bands, namely at 1050 and 968 cm-1, assigned to the C-O and C-C stretching vibrations of deoxyribose (65, 69, 70, 73). The band at 1050 cm-1 exhibited no major shifting (1 cm-1), whereas the band at 968 cm-1 was observed at the same frequency, suggesting that no direct polycation-sugar interaction occurred in these polyamine·DNA complexes (Figs. 1 and 2).

DNA Conformation—The region 1000-700 cm-1 is of particular interest for DNA conformation. The DNA B-form exhibits an absorption band with medium intensity at 836 cm-1, which disappears at lower humidities (74). The 836 cm-1 band, which involves the sugar-phosphodiester mode, is considered as a major marker band for the B-DNA conformation (64, 65, 74). Minor spectral changes were observed for the marker bands in the spectra of the biogenic polyamine·DNA complexes. However, these minor changes do not lead to conformational transitions such as B {leftrightarrow} A {leftrightarrow} Z. The observed spectral changes are due to minor alterations of the sugar-phosphate geometry upon polycation-DNA interactions. However, at high cobalt(III)hexamine content a new weak band emerged at 861 cm-1 (Fig. 2B, r = 1/4), which is related to A-form DNA (65, 69, 74). The presence of this band can be due to the progressive dehydration of the B-DNA and a partial B -> A transition (74) upon addition of cobalt(III)hexamine cation.

To verify whether cobalt(III)hexamine cations induce DNA conformational changes, circular dichroism analyses were performed using constant DNA concentration (1.25 mM) and various Co(III)/DNA(P) molar ratios. The CD spectrum of pure DNA (Fig. 4, line a) shows two positive bands at 275 nm (large band) and 221 nm (weak band) and a negative one at 245 nm, typical of B-DNA conformation (75, 76). The DNA bands at 275 and 245 nm were not changed upon the addition of cobalt(III)hexamine suggesting that DNA remain as B-conformation. On the other hand, the intensity of the weak positive band at 221 nm decreased and shifted to a higher wavelength (224 cm-1) upon addition of cobalt(III)hexamine cation. This was followed by an increase in the molar ellipticity of the negative band around 212 nm (Fig. 4, lines b-f), characteristic of A-DNA conformation (76). Hence, the conformation of the DNA may be an intermediate between B-DNA and A-DNA. These findings suggest that the new weak band, which appeared at 861 cm-1 in the infrared spectra of the Co(III)·DNA complexes at higher cobalt(III)hexamine content (r = 1/10 to 1/4), is due to the partial B -> A transition.



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  		FIG. 4.
CD spectra of highly polymerized calf thymus DNA in 25 mM NaCl (pH ~ 7) at 25 °C and at various concentrations of cobalt(III)hexamine. a, pure DNA (1.25 mM); b, Co(III)/DNA(P) = 1/15; c, Co(III)/DNA(P) = 1/10; d, Co(III)/DNA(P) = 1/8; e, Co(III)/DNA(P) = 1/6; f, Co(III)/DNA(P) = 1/4.

 
When a complete B -> A transition occurs the DNA marker bands such as 836 cm-1 appears at about 820-810 cm-1, the stretching vibration at 1222 cm-1 shifts toward a higher frequency at 1230-1240 cm-1, and the guanine band at 1717 cm-1 appears at 1710-1700 cm-1 (64, 77). In the B -> Z transition, the guanine band at 1717 cm-1 shifts to 1690 cm-1 and the phosphate vibration at 1222 cm-1 appears at 1216 cm-1, whereas the sugar-phosphate band at 836 cm-1 displaces toward a lower frequency at 800-790 cm-1 (64). Because no such spectral changes occurred in biogenic polyamine·DNA adducts, DNA remains in B-conformation in these complexes.

Stability of Polyamines·DNA Adducts by Capillary Electrophoresis—The binding of the polyamine·DNA complexes was also studied by capillary electrophoresis. Mixtures containing constant amounts of dsDNA (1.25 mM) and various amounts of polyamine in molar ratios of 1/800, 1/400, 1/200, 1/100, 1/50, 1/25, 1/12.5, and 1/6.25 were prepared and subjected to electrophoresis using an uncoated fused silica capillary of 75-µm inner diameter (effective length of 50 cm) at 25 kV. The electropherograms were monitored at 260 nm and at 25 °C in a run buffer of 20 mM Tris-HCl (pH 7.0 ± 0.2). The experimental data related to the migration times of pure DNA and its complexes with different concentrations of biogenic polyamines and cobalt(III)hexamine are shown in Table II.


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  		TABLE II
Experimental values of the migration time of pure DNA and polyamine-DNA adducts following incubation of a constant concentration of DNA (1.25 mM) with various concentrations of polyamines

Incubations were carried out in the run buffer (20 mM Tris-HCl, pH 7.0 ± 0.2) at 25 °C for 30 min prior to capillary electrophoresis.

 
The saturated curve is shown in Fig. 5A. The migration time of polyamine·DNA adducts gradually increased as the polyamine concentration increased and nearly reached a plateau at low polyamine concentration (around 50 µM) (Fig. 5A). Based on these results, binding constants of the polycation·DNA complexes were determined by Scatchard analysis as described under "Experimental Procedures." Quantitative determination of the binding constant for polyamine·DNA complexes will help to clarify the molecular mechanism of DNA stabilization and condensation by polyamines. Scatchard analysis for the biogenic polyamine·DNA adducts showed one overall binding constant for each polyamine with KSpm = 2.3 x 105 M-1, KSpd = 1.4 x 105 M-1, and KPut = 1.02 x 105 M-1 (Fig. 5B). However, no significant differences were detected in their binding affinities that could explain that spermine, spermidine, and putrescine may have a common molecular mechanism for DNA binding. Similar Scatchard analysis for the cobalt(III)hexamine·DNA complexes showed two binding constants with K1 = 1.8 x 105 M-1 and K2 = 9.2 x 104 M-1 (Fig. 5C). These results suggest a high and a low affinity for DNA upon cobalt(III)hexamine complexation. Furthermore, it may be speculated that the stronger interaction is due to major groove binding involving the purine-N7 atom, whereas the weaker is due to its binding with the backbone group, in agreement with the infrared data. Direct contacts between cobalt(III)hexamine and nucleic acids were observed by x-ray study (71) in which one of the cobalt(III)hexamine cations was located in DNA major groove binding via hydrogen bonding to the guanine residue (N7 and O6 atoms). The second cobalt(III)hexamine cation was positioned between DNA molecules forming hydrogen bonds to the phosphate oxygen atoms from different strands.



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  		FIG. 5.
A, plot of increase in migration time (in minutes) of polyamine·DNA complexes from capillary electrophoresis following incubation of a constant concentration of DNA (1.25 mM) with various concentrations of polyamines. The increase in migration time of the polyamine·DNA complexes was determined by subtracting the migration time of pure DNA from that of each polyamine·DNA adducts. B and C, Scatchard plots for biogenic polyamine·DNA and cobalt(III)hexamine·DNA complexes, respectively. D, Hill plots for polyamine·DNA complexes.

 
The cooperativity of the binding was analyzed using a Hill plot (Fig. 5D). For biogenic polyamines spermine, spermidine, and putrescine, the Hill coefficients, n, were 1.24, 1.47, and 1.74, respectively. These Hill coefficients were all greater than unity, indicating a positive cooperativity binding of these three polyamines with the dsDNA (63). In our opinion, this positive cooperativity reflects the biological importance of the biogenic polyamines within the cell. Indeed, depletion of polyamines was shown to cause a minor destabilization of the chromatin structure (35). In contrast, the Co(III)-DNA interaction was characterized by a negative cooperativity with Hill coefficient (n) of <1 (Fig. 5D). This may be due to the nonspecific interaction between cobalt(III)hexamine and DNA.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
Effect of Biogenic Polyamines and Cobalt(III)hexamine on Conformation and Stability of Calf-thymus DNA
The purpose of this study was to examine the effects of different polyamines on DNA structure and to determine their preferential binding sites at various polycation concentrations. The physiological functions of the polyamines are gradually clarified at the molecular level (6, 15). Among the biological functions of polyamines, one notable feature is their ability to condense DNA (27). In our experiments, we found no evidence to suggest calf-thymus DNA condensation by biogenic polyamines. It was previously suggested that the polyamines do not regulate the chromatin condensation state during the cell cycle, although they might have some stabilizing effect on the chromatin structure (35). It seems polyamines alone can not induce DNA condensation in cellular systems, and several other factors are involved in the stabilization of the compact forms of DNA such as specific proteins (histones in eukaryotes or histone-like proteins in prokaryotes), macromolecular crowding or DNA supercoiling. It appears that these different factors contribute to the stabilization of the compact forms of DNA (78). On the other hand, high concentrations of polyamines are generally found in the G1 phase of the cell cycle (1). It has been suggested that these high concentrations are required in the cell preparation for DNA synthesis. In our experimental setting, DNA stabilization occurs at molar ratios of 1/4 for spermine and spermidine, and between 1/4 and 1 for putrescine, corresponding to 6.25 mM for spermine and spermidine, and 6.25 to 25 mM for putrescine, that are close to physiological concentrations (the polyamines intracellular concentrations are in the millimolar range).

In contrast, the infrared results showed DNA condensation by cobalt(III)hexamine as exhibited by negative features in the infrared difference spectra of Co(III)·DNA complexes at high Co(III)/DNA ratio (r = 1/4). Our findings agree with those of Widom and Baldwin (36), who showed that efficiently aggregates random sequence natural DNA without any CD changes. The condensation process would be consistent with the formation of two direct hydrogen bonds between the punine-N7 of the major groove and the phosphate group of DNA.

The infrared spectra also provided the evidence that biogenic polyamine·DNA interaction leads to no major biopolymer conformational changes with DNA remaining in the B-family structure. This finding agrees with recent Raman studies of different genomic DNA, including calf-thymus DNA (23, 24), that find no conformational changes upon polyamine complexation. Indeed, polyamine-bound genomic DNA maintains the B-form structure even upon precipitation (23). The capability of biogenic polyamines to bind and condense genomic DNA while conserving the native B-form secondary structure may have important biological implications. Of particular interest is the present finding that cobalt(III)hexamine induced a partial B -> A transition. The infrared spectrum of pure DNA (Figs. 1 and 2, top spectra) shows a clear band at 836 cm-1 and no band at 860 cm-1, which is the pattern for pure B-family structures (74). Co(III)·DNA complexes at high cobalt(III)hexamine concentration (r = 1/4) gives a very similar spectrum (Fig. 2B, r = 1/4), although a small band is observed near 860 cm-1 and a weak band near 836 cm-1 is reduced, which indicate some degree of A-DNA formation. We believe that the incomplete transition from B to A form is due to a partial dehydration of DNA upon cobalt(III)hexamine complexation. Indeed, pure DNA when hydrated at 75% relative humidity exhibits the same behavior with a major band at 860 cm-1 and a weak band at 836 cm-1 (74). From ultrasonic and densimetric measurements, it has been found that cobalt(III)hexamine reduce the hydration of the atomic group of DNA (79).

To clarify the conformational changes of B-DNA upon cobat(III)hexamine complexation, circular dichroism results are helpful. No noticeable variation in the molar ellipticity of the bands in the region 240-300 nm were observed (Fig. 4, a-f) suggesting that DNA remains in B-form. However, minor spectral changes observed in the region 200-240 nm (reduced intensity and shifting of the band 221 nm, followed by intensity increase of the band around 212 nm) are characteristics of A-conformation (76). It is possible that some sequences of DNA undergo A-transition, while the rest still remain in B-form. The contrasting differences between cobalt(III)hexamine and biogenic polyamine binding might relate to the different molecular mechanism for DNA binding. Thus, cobalt(III)hexamine cations bind to nitrogen base sites (including N7) and the backbone groups. Conversely, biogenic polyamines bind primarily to the phosphates of the DNA backbone increasing the base pairing and stacking of B-DNA. In fact, this is consistent with our infrared hypochromism effect observed at high biogenic polyamine concentrations.

Preferred Binding Models for Polyamine·DNA Complexes and Their Biological Implication
Our spectroscopic data are used to build models for each biogenic polyamine·DNA adducts. It should be noted that several models are proposed for polyamine·DNA complexes in recent theoretical and spectroscopic studies (22, 24, 25, 48). To present some interaction models, the intramolecular distances should be taken into account. It was seen that polyamines attached to B-DNA are in trans torsion angles (22, 48). As predicted by theoretical calculations (80), this is the lowest energy conformation for these polyamines. Intramolecular distances for dsDNA and polyamines can be approximately calculated using HyperChem software (version 7.0 (2002) Hypercube, Alberta, Canada). Thus, the distances between the outer primary amino group in the all-trans conformation are 6.23, 11.13, and 16.04 Å for putrescine, spermidine, and spermine, respectively. The distance between a primary amino group and a secondary amino group separated by a trimethylene chain is around 5 Å (Structure 1). In addition, spermine contains two inner secondary amino groups that are separated by 6.23 Å. The double-stranded DNA in B-form was constructed using HyperChem Nucleic Acids Databases. A model for the polyamine cation was built manually using drawing tools of the HyperChem Model Builder and was subjected to geometrical optimization.

Putrescine—Our infrared spectra of putrescine·DNA complexes showed that putrescine binds to the major and minor grooves of DNA duplex at both low and high polycation concentrations. In addition, electrostatic interaction with negatively charged backbone phosphate groups occurs for putrescine·DNA complexes.

In the major groove of double strand DNA, the guanine- and adenine-N7 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 distance between these atoms and the oxygen atom of the nearest phosphate group is around 6.36 and 6.47 Å, respectively, which correlates well with the putrescine N-N distance (6.23 Å). The primary amino groups of putrescine could make two hydrogen bonds with the oxygen atom of the group and either the guanine-N7 or adenine-N7 atoms of the same strand at the major groove (Fig. 6A). Intragroove interaction by two adjacent groups (6.34 Å) was also proposed (Fig. 6A). However, the molecular size of putrescine could not allow it to have interstrand binding. In a minor groove model, estimated lengths of 6.02 Å can be obtained for the thymine-O2 atom and group of the same strand, which also correlates with the putrescine N-N distance. We therefore propose a putrescine·DNA model where the polyamine groups are in contact with a phosphate group and a thymine-O2 atom in the minor groove as shown in Fig. 6B.



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  		FIG. 6.
Preferential binding model proposed for putrescine·DNA complexes. A, phosphate and purine-N7 and phosphate-phosphate intragroove from the same strand of the major groove. B, phosphate and thymine-O2 from the same strand of the minor groove.

 
Spermidine—Infrared data for spermidine·DNA complexes indicate preferential binding by the major groove at low cation concentration and by both major and minor grooves at higher concentration. The smaller molecular size of putrescine allowed it to interact with both minor and major grooves. The presence of a secondary amino group extends the binding possibility of spermidine (N-N distance in an all-trans spermidine ~ 11.2 Å), which allows it to bind two reactive sites located at different DNA strands. Considering the interatomic distances in the major groove, there are suitable values for the spermidine interaction involving three adjacent phosphate groups (~11.9 Å) from the same strand (Fig. 7A). The distances between oxygen atoms of these phosphate groups are O2P1-O2P2 ~ 6.3 Å and O2P2-O1P3 ~ 5.7 Å, which correlate well with the distances between the amino groups of spermidine (~6.23 Å and ~5 Å, respectively) (Fig. 7A). In the model in Fig. 7B, two contacts with DNA bases are shown in the major groove of dsDNA. In this model, the two outer groups are near the N7 atom of adenine and oxygen of phosphate group (11 Å), whereas the inner group is in contact with the N7 atom of the guanine. Relevant interatomic lengths for this model are ~7 Å (O2P-N7(G)) and ~4 Å (N7(G)-N7(A)), which correlate slightly to the N-N distance of spermidine (~6.23 Å and ~5 Å).



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  		FIG. 7.
Preferential binding model proposed for Spd·DNA complexes. A, three phosphates adjacent from the same strand of the major groove. B, intrastrand across the major groove. C, intrastrand across the minor groove.

 
Several models involving the binding of spermidine in the minor groove of dsDNA have been proposed recently (24, 25). In these models, the authors showed an interstrand contact between polyamine amino groups and T(O2), A(N3), and groups. In our model (Fig. 7C), we propose an intrastrand binding possibility in which the outer amino groups are near thymine-O2 and -O4 of sugar (~11.3 Å). In addition, this model allows for interaction between an adenine-N3 atom and the inner spermidine- group. Hydrophobic contacts with the methylene-C5' can also occur (Fig. 7C). It should be noted that some phosphate-phosphate interstrand distances at the dsDNA minor grooves could favor the attachment of spermidine molecules (model not showed), whereas it is not possible at the major groove.

Spermine—Based on infrared data, the major groove is the most favorable binding site on dsDNA for spermine. Spermine interaction with the bases at the major groove was demonstrated by x-ray studies performed on spermine-oligonucleotide complexes (21, 42). In addition, the interaction between spermine and DNA has been widely studied using theoretical calculations (45, 47). These studies indicated that the interaction at the major groove of alternating purine/pyrimidine sequences appears to be the most favorable of all models presented and is associated with significant bending of DNA. The influence of base composition (and sequence) on the process of interaction between spermine and synthetic polynucleotides has shown significant differences between A-T and G-C base pairs (31). Thus, diverse binding models could be considered for spermine, because the interactions with the bases are sequence-dependent. Recent Raman studies (24, 25) propose the interactions along and across the major grooves involving contacts between the inner amino groups and purine-N7 and thymine-O4 atoms, which also permit hydrophobic contact between CH2 group of thymine and methylene group of spermine. These models correlate well with our infrared data, because interactions with the purine-N7 atoms and hydrophobic contacts are observed. In our major groove model, the outer primary amino groups of spermine could bind with groups from different strands (~16.4 Å), which correlate well with the distances between the outer primary amino groups of spermine (~16.04 Å). This model allows for inner interactions with N7 atoms of guanine and adenine bases (Fig. 8). As mentioned above, spermidine molecules favor the phosphate-phosphate interstrand attachment in the minor groove, whereas spermine could favor the interstrand attachment at both the minor and major grooves. In our opinion this might justify DNA protection against radiation, oxidation, and thermal denaturation.



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  		FIG. 8.
Preferential binding model proposed for spermine·DNA complexes. Interstrand is shown along the major groove.

 
Polyamines play an essential role in cell growth and differentiation, and interaction with nucleic acids could explain some of their important biological functions (1, 6, 8). Several studies indicate that polyamines play a critical role in protecting DNA strand breaks induced by radiation and oxidative stress (9-12). In fact, such DNA damage decreases the survivability of polyamine-deficient Escherichia coli mutant cells (11). However, the mechanisms by which this occurs are unknown. Our proposed models involve both intra- and interstrand interactions between polyamines and DNA. In our opinion, the intrastrand interaction would justify the ability of these polyamines to protect DNA against strand breaks. However, DNA strand breaks were not prevented as much by putrescine as by spermidine and spermine (putrescine afforded a lower protection) (9-12). This difference could be explained by the difference in the charges and sizes of polyamines. On the other hand, the interstrand interaction would explain the ability of spermine and spermidine to offer higher protection.


   FOOTNOTES
 
* This work was supported by the Natural Sciences and Engineering Research Council of Canada and Fonds pour la formation de Chercheurs et l'Aide à la Recherche (Québec). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

{ddagger} To whom correspondence should be addressed. Tel.: 819-376-5052 (ext. 3310); Fax: 819-376-5084; E-mail: tajmirri{at}uqtr.ca.

1 The abbreviations used are: Spm, spermine; Spd, spermidine; Put, putrescine; Co(III), cobalt(III)hexamine; ACE, affinity capillary electrophoresis; FTIR, Fourier transform infrared; CD, circular dichroism; r, polyamine/DNA(P) molar ratio. Back


   ACKNOWLEDGMENTS
 
We highly appreciate the financial support of the Natural Sciences and Engineering Research Council of Canada and Fonds pour la formation de Chercheurs et l'Aide à la Recherche (Qué-bec) for this work.



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Tabor, H., and Tabor, C. W. (1984) Annu. Rev. Biochem. 53, 749-790[CrossRef][Medline] [Order article via Infotrieve]
  2. Pegg, A. E. (1988) Cancer Res. 48, 759-774[Abstract/Free Full Text]
  3. Heby, O. (1981) Differentiation 19, 1-20[CrossRef][Medline] [Order article via Infotrieve]
  4. Tkachenko, A., Nesterova, L., and Pshenichnov, M. (2001) Arch. Microbiol. 176, 155-157[CrossRef][Medline] [Order article via Infotrieve]
  5. Marton, L. J., and Pegg, A. E. (1995) Annu. Rev. Pharmacol. Toxicol. 35, 55-91[CrossRef][Medline] [Order article via Infotrieve]
  6. Cohen, S. S. (1998) A Guide to the Polyamines, Oxford University Press, New York
  7. Casero, R. A., and Woster, P. M. (2001) J. Med. Chem. 44, 1-26[CrossRef][Medline] [Order article via Infotrieve]
  8. Thomas, T., and Thomas, T. J. (2001) Cell. Mol. Life Sci. 58, 244-258[CrossRef][Medline] [Order article via Infotrieve]
  9. Ha, H. C., Sirisoma, N. S., Kuppusamy, P., Zweier, J. L., Woster, P. M., and Casero, R. A., Jr. (1998) Proc. Natl. Acad. Sci. U. S. A. 95, 11140-11145[Abstract/Free Full Text]
  10. Ha, H. C., Yager, J. D., Woster, P. A., and Casero, R. A., Jr. (1998) Biochem. Biophys. Res. Commun. 244, 298-303[CrossRef][Medline] [Order article via Infotrieve]
  11. Oh, T. J., and Kim, I. G. (1998) Biotech. Techniques 12, 755-758[CrossRef]
  12. Douki, T., and Bretonniere, Y. (2000) J. Cadet. Radiat. Res. 153, 29-35
  13. Morgan, D. M. L. (1990) Biochem. Soc. Trans. 18, 1080-1084[Medline] [Order article via Infotrieve]
  14. Watanabe, S., Kusama-Eguchi, K., Kobayashi, H., and Igarashi, K. (1991) J. Biol. Chem. 226, 20803-20809
  15. Igarashi, K., and Kashiwagi, K. (2000) Biochem. Biophys. Res. Commun. 271, 559-564[CrossRef][Medline] [Order article via Infotrieve]
  16. Thomas, T. J., Gunnia, U. B., and Thomas, T. (1991) J. Biol. Chem. 266, 6137-6141[Abstract/Free Full Text]
  17. Bancroft, D., Williams, L. D., Rich, A., and Egli, M. (1994) Biochemistry 33, 1073-1086[CrossRef][Medline] [Order article via Infotrieve]
  18. Hasan, R., Alam, M. K., and Ali, R. (1995) FEBS Lett. 368, 27-30[CrossRef][Medline] [Order article via Infotrieve]
  19. Ruiz-Chica, J., Medina, M. A., Sanchez-Jiménez, F., and Ramírez, F. J. (2001) Biochem. Biophys. Res. Commun. 285, 437-446[CrossRef][Medline] [Order article via Infotrieve]
  20. Minyat, E. E., Ivanov, V. I., Kritzyn, A. M., Minchenkova, L. E., and Schyolkina, A. K. (1978) J. Mol. Biol. 128, 379-409[CrossRef]
  21. Jain, S., Zon, G., and Sundaralingam, M. (1989) Biochemistry 28, 2360-2364[CrossRef][Medline] [Order article via Infotrieve]
  22. Real, A. N., and Greenall, R. J. (2004) J. Biomol. Struct. Dyn. 21, 469-488[Medline] [Order article via Infotrieve]
  23. Deng, H., Bloomfield, V. A., Benevides, J. M., and Thomas, G. J., Jr. (2000) Nucleic Acids Res. 28, 3379-3385[Abstract/Free Full Text]
  24. Ruiz-Chica, J., Medina, M. A., Sanchez-Jiménez, F., and Ramírez, F. J. (2001) Biophys. J. 80, 443-454[Medline] [Order article via Infotrieve]
  25. Ruiz-Chica, J., Medina, M. A., Sanchez-Jiménez, F., and Ramírez, F. J. (2003) Biochim. Biophys. Acta 1628, 11-21[Medline] [Order article via Infotrieve]
  26. Gosule, L. C., and Schellman, J. A. (1976) Nature 259, 333-335[CrossRef][Medline] [Order article via Infotrieve]
  27. Pelta, J., Livolant, F., and Sikorav, J.-L. (1996) J. Biol. Chem. 271, 5656-5662[Abstract/Free Full Text]
  28. Bloomfield, V. A. (1997) Biopolymers 44, 269-282[CrossRef][Medline] [Order article via Infotrieve]
  29. Lin, Z., Wang, C., Feng, X., Liu, M., Li, J., and Bai, C. (1998) Nucleic Acids Res. 26, 3228-3234[Abstract/Free Full Text]
  30. D'Agostino, L., and Di Luccia, A. (2002) Eur. J. Biochem. 269, 4317-4325[Medline] [Order article via Infotrieve]
  31. Marquet, R., Colson, P., and Houssier, C. (1986) J. Biomol. Struct. Dyn. 4, 205-218[Medline] [Order article via Infotrieve]
  32. Sen, D., and Crothers, D. M. (1986) Biochemistry 25, 1495-1503[CrossRef][Medline] [Order article via Infotrieve]
  33. Hougaard, D. M., Fujiwara, K., and Larsson, L. I. (1987) Histochem. J. 19, 643-650[CrossRef][Medline] [Order article via Infotrieve]
  34. Sauve, D. M., Anderson, H. J., Ray, J. M., James, W. M., and Roberge, M. (1999) J. Cell Biol. 145, 225-235[Abstract/Free Full Text]
  35. Laitinen, J., Stenius, K., Eloranta, T. O., and Hölttä, E. (1998) J. Cell. Biochem. 68, 200-212[CrossRef][Medline] [Order article via Infotrieve]
  36. Widom, J., and Baldwin, R. L. (1980) J. Mol. Biol. 144, 431-453[CrossRef][Medline] [Order article via Infotrieve]
  37. Tabor, H. (1962) Biochemistry 1, 496-501[CrossRef][Medline] [Order article via Infotrieve]
  38. Thomas, T. J., and Bloomfield, V. A. (1984) Biopolymers 23, 1295-1306[CrossRef][Medline] [Order article via Infotrieve]
  39. Tsuboi, M. (1964) Bull. Chem. Soc. Jpn. 37, 1514-1522[CrossRef]
  40. Liquori, A. M., Constantino, L., Crescenzi, V., Elia, V., Giglio, E., Puliti, R., De Santis-Savino, M., and Vitagliano, V. (1967) J. Mol. Biol. 24, 113-122[CrossRef]
  41. Williams, L. D., Frederick, C. A., Ughetto, G., and Rich, A. (1990) Nucleic Acids Res. 18, 5533-5541[Abstract/Free Full Text]
  42. Drew, H. R., and Dickerson, R. E. (1981) J. Mol. Biol. 151, 535-556[CrossRef][Medline] [Order article via Infotrieve]
  43. Schmid, N., and Behr, J.-P. (1991) Biochemistry 30, 4357-4361[CrossRef][Medline] [Order article via Infotrieve]
  44. Wemmer, D. E., Srivenugopol, K. S., Reid, B. R., and Morris, D. R. (1985) J. Mol. Biol. 185, 457-459[CrossRef][Medline] [Order article via Infotrieve]
  45. Feuerstein, B. G., Pattabiraman, N., and Marton, L. J. (1990) Nucleic Acids Res. 18, 1271-1282[Abstract/Free Full Text]
  46. Pattabiraman, N., Langridge, R., and Kollman, P. A. (1984) J. Biomol. Struct. Dyn. 1, 1525-1533[Medline] [Order article via Infotrieve]
  47. Feuerstein, B. G., Pattabiraman, N., and Marton, L. J. (1986) Proc. Natl. Acad. Sci. U. S. A. 83, 5948-5992[Abstract/Free Full Text]
  48. Bryson, K., and Greenall, R. J. (2000) J. Biomol. Struct. Dyn. 18, 393-412[Medline] [Order article via Infotrieve]
  49. Korolev, N., Lyubartsev, A. P., Nordenskiöld, L., and Laaksonen, A. (2001) J. Mol. Biol. 308, 907-917[CrossRef][Medline] [Order article via Infotrieve]
  50. Korolev, N., Lyubartsev, A. P., Laaksonen, A., and Nordenskiöld, L. (2002) Biophys. J. 82, 2860-2875[Medline] [Order article via Infotrieve]
  51. Marmur, J. (1961) J. Mol. Biol. 3, 208-218
  52. Reichmann, M. E., Rice, S. A., Thomas, C. A., and Doty, P. (1954) J. Am. Chem. Soc. 76, 3047-3053[CrossRef]
  53. Vijayalakshmi, R., Kanthimathi, M., and Subramanian, V. (2000) Biochem. Biophys. Res. Commun. 271, 731-734[CrossRef][Medline] [Order article via Infotrieve]
  54. Alex, S., and Dupuis, P. (1989) Inorg. Chim. Acta 157, 271-281[CrossRef]
  55. Ahmed Ouameur, A., Nafisi, S., Mohajerani, N., and Tajmir-Riahi, H. A. (2003) J. Biomol. Struct. Dyn. 20, 561-565[Medline] [Order article via Infotrieve]
  56. Klotz, M. I. (1982) Science 217, 1247-1249[Abstract/Free Full Text]
  57. Guszcynski, T., and Copeland, T. D. (1998) Anal. Biochem. 260, 212-217[CrossRef][Medline] [Order article via Infotrieve]
  58. Foulds, G. J., and Etzkorn, F. A. (1998) Nucleic Acids Res. 26, 4304-4305[Abstract/Free Full Text]
  59. Li, G., and Martin, M. L. (1998) Anal. Biochem. 263, 72-78[CrossRef][Medline] [Order article via Infotrieve]
  60. Xian, J., Harrington, M. G., and Davidson, E. H. (1996) Proc. Natl. Acad. Sci. U. S. A. 93, 86-90[Abstract/Free Full Text]
  61. Marty, R., Ahmed Ouameur, A., Neault, J.-F., Nafisi, S., and Tajmir-Riahi, H. A. (2004) DNA Cell Biol. 23, 135-140[CrossRef][Medline] [Order article via Infotrieve]
  62. Shimura, K., and Kasai, K.-I. (1997) Anal. Biochem. 251, 1-16[CrossRef][Medline] [Order article via Infotrieve]
  63. Jack, R. C. (1995) Basic Biochemical Laboratory Procedures and Computing, pp. 147-198, Oxford University Press, New York
  64. Taillandier, E., and Liquier, J. (1992) Methods Enzymol. 211, 307-335[Medline] [Order article via Infotrieve]
  65. Taillandier, E., Liquier, J., and Taboury, J. A. (1985) Adv. Infrared Raman Spectrosc. 12, 65-113
  66. Arakawa, H., Ahmed, R., Naoui, M., and Tajmir-Riahi, H. A. (2000) J. Biol. Chem. 275, 10150-10153[Abstract/Free Full Text]
  67. Pavia, D. L., Lampman, G. M., and Kriz, G. S. (1979) Introduction to Spectroscopy: A Guide for Students of Organic Chemistry, Saunders Golden Sun-burst Series, Toronto, Canada
  68. Günzler, H., and Gremlich, H.-U. (2002) IR Spectroscopy, An Introduction, Wiley-Verlag Chemie, Germany
  69. Taillandier, E., Liquier, J., Taboury, J. A., and Ghomi, M. (1984) in Spectroscopy of Biological Molecules (Sandorfy, C., and Theophanides, T., eds) pp. 171-189, Reidel Dordrecht, Amsterdam
  70. Taillandier, E., Taboury, J. A., Adam, S., and Liquier, J. (1984) Biochemistry 23, 5703-5706[CrossRef][Medline] [Order article via Infotrieve]
  71. Nunn, C. M., and Neidle, S. (1996) J. Mol. Biol. 256, 340-351[CrossRef][Medline] [Order article via Infotrieve]
  72. Tsuboi, M. (1969) Appl. Spectrosc. Rev. 3, 45-90
  73. Theophanides, T., and Tajmir-Riahi, H. A. (1985) J. Biomol. Struct. Dyn. 2, 995-1004[Medline] [Order article via Infotrieve]
  74. DiRico, D. E., Jr., Keller, P. B., and Hartman, K. A. (1985) Nucleic Acids Res. 13, 251-260[Abstract/Free Full Text]
  75. Hou, M.-H., Lin, S.-B., Yuann, J.-M. P., Lin, W.-C., Wang, A. H.-J., and Kan, L.-S. (2001) Nucleic Acids Res. 29, 5121-5128[Abstract/Free Full Text]
  76. Kypr, J., and Vorlíèková, M. (2002) Biopolymers (Biospectroscopy) 67, 275-277
  77. Loprete, D. M., and Hartman, K. A. (1993) Biochemistry 32, 4077-4082[CrossRef][Medline] [Order article via Infotrieve]
  78. Zimmerman, S. B., and Murphy, L. D. (1996) FEBS Lett. 390, 245-248[CrossRef][Medline] [Order article via Infotrieve]
  79. Kankia, B. I., Buckin, V., and Bloomfield, V. A. (2001) Nucleic Acids Res. 29, 2795-2801[Abstract/Free Full Text]
  80. Marques, M. P. M., and Batista de Carvalho, L. A. E. (2000) in Biologically Active Amines in Food (Morgan, D. M. L., White, A., Sánchez-Jiménez, F., and Bardócz, S., eds) Vol. 4, pp. 122-129, Office for Official Publications of the European Communities, Luxembourg

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