A Comparison with Biogenic Polyamines - American Chemical Society

2712

Biomacromolecules 2008, 9, 2712–2718

DNA Interaction with Antitumor Polyamine Analogues: A Comparison with Biogenic Polyamines C. N. N’soukpoe´-Kossi,† A. Ahmed Ouameur,† T. Thomas,‡,§ A. Shirahata,| T. J. Thomas,§,⊥ and H. A. Tajmir-Riahi*,† De´partement de Chimie-Biologie, Universite´ du Que´bec a´ Trois-Rivie`res, C. P. 500, Trois-Rivie`res (Que´bec), G9A 5H7, Canada, Department of Environmental and Occupational Medicine, Environmental and Occupational Health Sciences Institute, Department of Medicine, and The Cancer Institute of New Jersey, University of Medicine and Dentistry of New Jersey, Robert Wood Johnson Medical School, New Brunswick, New Jersey 08903, and Department of Biochemistry and Cellular Physiology, Josai University, Saitama, Japan Received April 16, 2008; Revised Manuscript Received June 16, 2008

Biogenic polyamines, putrescine, spermidine, and spermine, are ubiquitous cellular cations and exert multiple biological functions. Polyamine analogues mimic biogenic polyamines at macromolecular level but are unable to substitute for natural polyamines and maintain cell proliferation, indicating biomedical applications. The mechanistic differences in DNA binding mode between natural and synthetic polyamines have not been explored. The aim of this study was to examine the interaction of calf thymus DNA with three polyamine analogues, 1,11-diamino4,8-diazaundecane (333), 3,7,11,15-tetrazaheptadecane · 4HCl (BE-333), and 3,7,11,15,19-pentazahenicosane · 5HCl (BE-3333), using FTIR, UV-visible, and CD spectroscopy. Polyamine analogues bind with guanine and backbone PO2 group as major targets in DNA, whereas biogenic polyamines bind to major and minor grooves as well as to phosphate groups. Weaker interaction with DNA was observed for analogues with respect to biogenic polyamines, with K333 ) 1.90 ((0.5) × 104 M-1, KBE-333 ) 6.4 ((1.7) × 104 M-1, KBE-3333 ) 4.7 ((1.4) × 104 M-1 compared to KSpm ) 2.3 ((1.1) × 105 M-1, KSpd ) 1.4 ((0.6) × 105 M-1, and KPut ) 1.02 ((0.5) × 105 M-1. A partial B- to A-DNA transition was also provoked by analogues. These data suggest distinct differences in the binding of natural and synthetic polyamines with DNA.

Introduction Polyamine analogues (Scheme 1) exert antitumor activity in multiple experimental model systems, including breast and lung cancer models, and are being used in clinical trials.1-12 Polyamine analogues can mimic some of the self-regulatory functions of biogenic polyamines but are unable to substitute for natural polyamines in their growth promoting role.13 Natural polyamines (putrescine, spermidine, and spermine) are ubiquitous cellular cations and are involved in cell growth and differentiation. They are capable of modulating gene expression and enzyme activities, activation of DNA synthesis, and facilitating protein-DNA interactions.13-20 Biogenic polyamines interact with DNA backbone phosphate groups via electrostatic interactions.21 This interaction has been shown to protect small DNA molecules from common damaging agents, such as ionization and reactive oxygen species.22 Natural polyamines also have the ability to induce DNA conformational transitions, B- to Z-DNA23-27 or B- to A-DNA 28-31 transitions in specific sequences. In addition, polyamines can bind with B-DNA without provoking any major biopolymer conformational changes.21,32 * To whom correspondence should be addressed. Tel.:819-376-5011 (ext. 3310). Fax: 819-376-5084. E-mail: [email protected]. † Universite´ du Que´bec a´ Trois-Rivie`res. ‡ Department of Environmental and Occupational Medicine, Environmental and Occupational Health Sciences Institute. § The Cancer Institute of New Jersey. | Josai University. ⊥ Department of Medicine.

Scheme 1. Chemical Structures of Polyamine Analogues

Biogenic polyamines induce DNA condensation in both isolated DNA33-36 and chromatin.37,38 In polyamine condensed DNA, the attractive component of the free energy is ∼2.3 ( 0.2 times larger than the repulsive component of the free energy between double helices.38 Several studies have shown that spermidine and spermine are associated with highly compacted mitotic chromosomes,39-41 thereby stabilizing the chromatin structure during cell cycle.40-42 These studies indicate that polyamine-induced DNA condensation is important to the cellular functions in vivo. Even though structural models for the binding of biogenic polyamines to duplex DNA have been presented,21,22,32 less is known about the binding of polyamine analogues with nucleic acids and the effect of such complex formation on DNA or RNA structure. Polyamine analogues bind oligonucleotides 15mer GC and 15mer AT via major and minor grooves, inducing B- to Z-DNA conformational changes for G-C base pairs and causing DNA aggregation.43 Analogues also induce aggregation and precipita-

10.1021/bm800412r CCC: $40.75  2008 American Chemical Society Published on Web 08/26/2008

Polyamine Analogue-DNA Interactions

tion of highly polymerized DNA.44 The effects of polyamine analogues on the aggregation, precipitation, and conformations of single-, double-, and triple-stranded DNA have been reported.45,46 DNA condensation by polyamine analogues is known and the effects of analogues on the stability of duplex structures, such as RNA-DNA, DNA-RNA, RNA-RNA, and DNA-DNA have been investigated.47,48 Analogue binding to protein was also recently reported.49 Even though most of these studies deal with the interaction of polyamine analogues with DNA, major information regarding analogue binding sites, the effects of such interaction on DNA stability, aggregation, precipitation, and conformation is missing. Furthermore, detailed comparison of the effects of polyamine analogues and biogenic polyamines with regard to the question of why natural polyamines are vital for DNA stability and function, while analogues can exhibit antitumor activity and cause DNA structural changes is not reported. In this report, we have studied the interactions between calfthymus DNA and polyamine analogues 333, BE-333, and BE3333 using Fourier transform infrared, circular dichroism, and UV-visible spectroscopic methods. Evidence for binding and stability of analogue-DNA complexation, DNA aggregation, and conformation is provided. In addition, a comparison is made with results obtained for biogenic polyamine-DNA complex formation. Our results suggest significant differences in the DNA binding modes of natural and synthetic polyamines.

Experimental Section Materials. Polyamine analogues, 333, BE-333, and BE-3333 were synthesized by A. Shirahata. Highly polymerized type I calf-thymus DNA sodium salt (7% Na content) was purchased from Sigma Chemical Co., 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 from protein.50 Other chemicals were of reagent grade and used without further purification. Preparation of Stock Solutions. Sodium-DNA (8.3 mg/mL) was dissolved in Tris-HCl (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 molar extinction coefficient of 260 ) 6600 cm-1 M-1 (expressed as molarity of phosphate groups).51,52 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 base pairs (molecular weight ∼ 6 × 106 Da). The appropriate amount of polyamines (0.25-1 mM) was prepared in distilled water and added dropwise to DNA solution, in order to attain the final polyamine concentrations of 0.125-0.50 mM at a final DNA concentration of 12.5 mM (4.15 mg/mL) for infrared measurements. FTIR Spectra. Infrared spectra were recorded with a FTIR spectrometer (Impact 420 model) equipped with DTGS (deuterated triglycine sulfate) detector and KBr beam splitter, using AgBr windows. Spectra were recorded after a 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 4000-400 cm-1 with a nominal resolution of 2 cm-1 and a minimum of 100 scans. The water subtraction was carried out using H2O solution at pH ) 7.0 ( 0.2 as a reference.53 A good water subtraction was considered to be achieved if there was a flat baseline around 2200 cm-1, where the water combination mode was located. This method yields a rough estimate of the subtraction scaling factor, but it removes the spectral features of water in a satisfactory way.53

Biomacromolecules, Vol. 9, No. 10, 2008

2713

The difference spectra [(DNA solution + polyamine analogue) (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 stretching vibrations, exhibits no spectral changes (shifting or intensity variation) upon polyamine analogue-DNA complexation and canceled out upon spectral subtraction. The spectra presented here were smoothed with a Savitzky-Golay procedure.53 The plots of the relative intensity (R) of several peaks of DNA inplane vibrations related to A-T, G-C base pairs, and the PO2- stretching vibrations, such as 1710 (guanine), 1663 (thymine), 1610 (adenine), 1491 (cytosine), and 1225 cm-1 (PO2- groups), versus the polyamine analogue concentrations were obtained after peak normalization using

Ri )

Ii I968

(1)

where Ii is the intensity of absorption peak for pure DNA and DNA in the complex with i concentration of polyamine analogue, and I968 is the intensity of the 968 cm-1 peak (internal reference).21 CD Spectroscopy. Spectra of DNA and its polyamine complexes were recorded at pH 7.2 with a Jasco J-720 spectropolarimeter. For measurements in the far-UV region (200-320 nm), a quartz cell with a path length of 0.01 cm was used under nitrogen atmosphere. Three scans were accumulated at a scan speed of 50 nm per minute, with data being collected at every nm 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. Spectra were corrected for buffer signal. The polyamine analogue concentrations used in our experiment were 125, 250, and 500 µM with a final DNA concentration of 2.5 mM. Absorption Spectroscopy. The absorption spectra were recorded on a Perkin-Elmer Lambda 40 spectrophotometer. Quartz cuvettes of 1 cm were used. The absorption spectra were recorded for free DNA (125 µM) and for its complexes with each polyamine analogue (1-60 µM). To calculate the polyamine analogue-DNA binding constant, the data were treated according to the following equations:

DNA + polyamine T DNA-polyamine complex

(2)

K ) [DNA-polyamine complex] ⁄ [DNA]uncomplexed [polyamine]uncomplexed (3) The values of binding constants K were obtained from the DNA absorption at 260 nm, according to methods published in the literature,54,55 where the bindings of various ligands to hemoglobin were described. For weak binding affinities, the data were treated using linear reciprocal plots based on the following equation:

1 1 1 1 · ) + A - A0 A∞ - A0 K(A∞ - A0) Cligand

(4)

where A0 is the absorbance of DNA at 260 nm in the absence of ligand, A∞ is the final absorbance of the ligated-DNA, and A is the recorded absorbance at different ligand concentrations. The double reciprocal plot of 1/(A - A0) versus 1/Cligand is linear and the binding constant (K) can be estimated from the ratio of the intercept to the slope.

Results Polyamine Analogue-DNA Complexes Studied by FTIR Spectroscopy. Analogue Base Binding. Evidence for polyamine analogue-base binding comes from the shift of the guanine band at 1710 cm-1 (guanine-CdO stretch) (Table 1)21,53,56-60 to a lower frequency at 1705 for 333-DNA and 1706 cm-1 for Be3333-DNA complexes, whereas it showed no shifting in the spectra of BE-333-DNA complex (Figures 1, 2 and 3). An increase in the intensity of the band at 1710 cm-1 was also observed with positive features at 1710 (333) and 1704 cm-1

2714


N’soukpoé-Kossi et al.

Table 1. Principal Infrared Absorption Bands, Relative Intensities, and Assignments for Calf-Thymus DNA in Aqueous Solutiona wavenumber intensityb (cm-1) 1710 1661 1610 1575 1527 1491 1225 1088 1053 968 892 834

vs vs s sh w m vs vs s s m m

assignment guanine (CdO stretching) thymine (C2dO stretching) adenine (C7dN stretching) purine stretching (N7) in-plane vibration of cytosine and guanine in-plane vibration of cytosine asymmetric PO2- stretch symmetric PO2- stretch C-O deoxyribose stretch C-C deoxyribose stretch deoxyribose, B-marker deoxyribose, B-marker

a At pH 7 ( 0.2.21 b Relative intensities: s ) strong, sh ) shoulder, vs ) very strong, m ) medium, w ) weak.

Figure 2. FTIR spectra in the region of 1800-600 cm-1 for free DNA, free polyamine, and BE-333-DNA adducts in aqueous solution at pH ) 7.0 ( 0.2 (top three spectra), and difference spectra for polyamine-DNA adducts obtained at various BE-333 concentrations (bottom two spectra).

Figure 1. FTIR spectra in the region of 1800-600 cm-1 for free DNA, free polyamine, and 333-DNA adducts in aqueous solution at pH ) 7.0 ( 0.2 (top three spectra), and difference spectra for polyamine-DNA adducts obtained at various 333 concentrations (bottom two spectra).

(BE-333 and BE-3333) upon DNA complexation (Figures 1-3, diff. 125 µM). The intensity of these positive features (1710-1704 cm-1) increased as polyamine analogue concentration increased (Figures 1-3, diff. 500 µM). The spectral changes observed for the band at 1710 cm-1 are due to major polyamine analogue-DNA interaction via guanine bases. Thymine band at 1661 cm-1 (thymine-O2) showed no major spectral shifting and intensity variations upon analogue-DNA interaction both at low and high polyamine contents (Figures 1-3). The negative and positive features centered at 1621-1648 cm-1 in the difference spectra of polyamine-DNA complexes is coming from H2O bending mode21,53 and not from thymine band at 1661 cm-1 (Figures 1-3), indicating no direct polyamine-thymine interaction. Similarly, the adenine band at 1610 cm-1 (adenineN7) exhibits no shifting and intensity changes due to a lack of polyamine analogue-adenine interaction (Figures 1-3). No major alterations of cytosine band at 1491 cm-1 (1 cm-1) can

Figure 3. FTIR spectra in the region of 1800-600 cm-1 for free DNA, free polyamine, and BE-3333-DNA adducts in aqueous solution at pH ) 7.0 ( 0.2 (top three spectra), and difference spectra for polyamine-DNA adducts obtained at various BE-3333 concentrations (bottom two spectra).

also be attributed to no direct polyamine-cytosine binding (Figures 1-3). It should be noted that infrared results showed that more base binding occurred for biogenic polyamines spermine, spermidine and putrescine21 than those of polyamine analogues studied herein. Biogenic polyamines bind DNA via guanine-N7, thymineO2, and adenine-N7 with no major interaction with cytosine


Figure 4. CD spectra of highly polymerized calf thymus DNA (pH ∼ 7.2) at 25 °C (2.5 mM) and spermine (A) and 333 (B), Be-333 (C), and BE-3333 (D) with 125, 250, and 500 µM polyamine concentrations.

bases,21 whereas analogue binding is limited to the guanine-N7 site and the backbone phosphate group. Polyamine analogue binding to guanine bases of oligunucloetides and polynucleotides is known by Raman spectroscopy.43,44 Analogue Phosphate Binding. Polyamine analogue binding to backbone phosphate group is characterized by the spectral alterations of the PO2 asymmetric and symmetric bands at 1225 and 1088 cm-1, respectively (Table 1). The PO2 asymmetric band at 1225 cm-1 and symmetric band at 1088 cm-1 shifted toward lower frequencies at 1223 and 1084 (333-DNA), 1224 and 1086 (BE-333-DNA), and 1224 and 1082 cm-1 (BE3333-DNA) upon analogue complexation (Figures 1-3, 500 µM). The observed spectral shifting was accompanied by intensity variations of the phosphate bands with positive features at 1225 cm-1 and 1088 cm-1 (333), 1027 cm-1 and 1091 cm-1 (BE-333), and 1225 cm-1 (BE-3333) on analogue-PO2 interaction (Figures 1-3, diff., 500 µM). The spectral changes observed are due to a major analogue-phosphate interaction, which contributes to the stabilization of polyamine-DNA complexation. Further evidence regarding analogue-PO2 interaction is also coming from the intensity ratio variations of symmetric and asymmetric PO2 bands at 1088/1225.53 The ratio of νs/νas was changed from 1.66 (free DNA) to 1.75 (333-DNA), 1.90 (BE-333-DNA), and 2.0 (BE-3333-DNA) upon polyamine analogue-PO2 interaction (Figure 1-3, 500 µM). CD Spectra and DNA Conformation. The CD spectra of free calf-thymus DNA and its complexes with polyamine analogues at different polyamine concentrations are shown in Figure 4. The CD of the free DNA composed of four major peaks at 210


2715

(negative), 221 (positive), 245 (negative), and 280 nm (positive; Figure 4). This is consistent with CD spectra of double helical DNA in B conformation.61-64When a complete B- to A-DNA transition occurred, the CD marker band at 210 nm gained negativity with minor shifting, 222 nm lost intensity, and shifted toward a higher wavenumber, the band at 245 nm lost negativity, while the positive band at 280 nm gained intensity and shifted to 270-267 nm.61,64 Upon addition of polyamine analogues (125, 250, and 500 µM), major spectral changes of CD bands occurred. The band at 210 nm gained negativity with minor shifting ((2 nm), the band at 222 nm lost intensity with no shifting, the band at 245 nm lost negativity, while the positive band at 280 nm lost intensity with no major shifting (Figure 4B-D). It should be noted that the loss of intensities for the bands at 280 and 222 nm were less pronounced for biogenic spermine than those of polyamine analogues, particularly for the band at 280 nm, which lost most of its intensity as the concentration of analogues increased (Figure 4A, 500 µM). At the first look, these spectral changes indicate that DNA remains in B conformation because the band at 280 nm lost intensity and exhibited no major shifting toward a lower wavelength. However, the increase in negativity of the band at 210 nm and increase in positivity of the band at 222 nm, with a decrease of negativity for the band at 245 nm in the CD spectra of analogue-DNA complexes can be attributed to a partial B- to A-DNA transition upon complex formation (Figure 4B-D). Additional evidence for a partial B- to A-DNA transition comes from the emergence of a new IR band at 862-861 cm-1 in the infrared spectra of 333-DNA (861 cm-1), BE-333-DNA (862 cm-1), and BE-3333-DNA (861 cm-1; Figures 1-3). Further support for a partial B- to A-DNA transition also comes from the spectral changes (intensity and shifting) of a sugar-phosphate band at 1053 cm-1. This band shifted to a higher frequency at 1060 cm-1 (333-DNA), 1056 cm-1 (BE333-DNA), and 1056 cm-1 (BE-3333-DNA), upon analogueDNA complexation (Figures 1-3). Similar CD and infrared spectral changes were observed for cobalt-hexamine complex of calf-thymus DNA, where a partial B to A transition was observed.21 The presence of a new band at 861 cm-1 in the IR spectra of Co(III)-DNA together with major intensity variations of the CD bands at 210, 221, and 245 nm were attributed to partial B- to A-DNA conformational changes.21 It should be noted that the infrared marker band at 861 cm-1 for A-DNA conformation56 was not observed in the infrared spectra of biogenic polyamines spermine-, spermidine-, and putrescineDNA complexes, where DNA remained in the B-DNA family structure.21 However, the major loss of intensity of the CD band at 280 nm is attributed to DNA aggregation as polyamine analogue content increased (Figure 4). The loss of intensity of the band at 280 nm was less pronounced in the case of spermine, where DNA aggregation occurred at a much higher biogenic polyamine/DNA(P) ratio (r ) 0.4)21 than those of the analogues studied herein (r ) 0.2; Figure 4A, 500 µM). The condensation and precipitation of DNA by polyamine analogues have been previously studied using Laser light scattering.48 Stability of Polyamine Analogue-DNA Complexes. UV spectroscopic results reveal several important aspects of polyamine analogue-DNA complexation (Figure 5). At very low polyamine analogue concentrations (1-35 µM), DNA band at 260 nm (dark line) gained intensity upon analogue complexation, which allowed us to calculate the overall binding constant (K) for polyamine-DNA complex formation (accord-

2716


N’soukpoé-Kossi et al. Table 2. Binding Constants for Polyamine Analogue-DNA Complexes Compared with Biogenic Polyamines polyamine analogues

binding constant (M-1) 1.9 × 104 6.4 × 104 4.7 × 104

333 BE-333 BE-3333 biogenic polyamines

binding constant (M-1) 2.3 × 105 1.4 × 105 1.0 × 105

spermine spermidine putrescine

Discussion

Figure 5. UV-visible results of calf-thymus DNA and its spermine (A), 333 (B), Be-333 (C), and BE-3333 (D) complexes: spectra of (a) free DNA (125 µM); (b) free polyamine (60 µM); (1-15) polyamine-DNA complexes 1 (2), 2 (4), 3 (6), 4 (8), 5 (10), 6 (12), 7 (15), 8 (20), 9 (25), 10 (30), 11 (35), 12 (40), 13 (45), 14 (50), and 15 (60 µM).

ing to the method described in Experimental Section; Figure 5). The low analogue concentrations used for the calculation of the binding constant were below DNA aggregation point. However, at higher polyamine content (40-60 µM), major decrease in intensity and shifting of the band at 260 nm occurred toward a higher wavenumber (Figure 5B-D, curves 13-15). The loss of intensity and shifting of the band at 260 nm were more pronounced in the case of BE-3333 than those of BE333- and 333-DNA complexes (Figure 5B-D). The major decrease in intensity of the band at 260 nm is due to DNA aggregation in the presence of polyamine analogues. However, DNA aggregation was not observed in the presence of spermine at similar polyamine concentrations (35-60 µM; Figure 5A). Previous IR spectroscopic data showed that DNA aggregation occurred in the presence of biogenic polyamine at higher polyamine/DNA molar ratios (r ) 0.5 for spermine, spermidine and r ) 1 for putrescine)21 than that of polyamine analogues used in the present study (r ) 0.3). The binding constants estimated for polyamine analogue-DNA complexes were K333 ) 1.90 × 104 M-1, KBE-333 ) 6.4 × 104 M-1, KBE-3333 ) 4.7 104 M-1. However, the binding constants reported for biogenic polyamine-DNA adducts were KSpm ) 2.3 × 105 M-1, KSpd ) 1.4 × 105 M-1, and KPut ) 1.02 × 105 M-1 (Table 2). The weaker binding affinity of polyamine analogues (Table 2) is due to a faster DNA aggregation in the presence of analogues, which establishes a major competition between analogue-DNA binding and DNA precipitation.

Major Differences between Polyamine Analogues and Biogenic Polyamines. Structural models based on spectroscopic studies showed biogenic polyamine binding to both major and minor grooves as well as to the backbone phosphate group of DNA duplex.21 Putrescine binds to the major and minor grooves of DNA duplex at both low and high polycation concentrations. Putrescine binding is with N7 guanine and adenine, thymineO2, as well as with the backbone PO2 groups.21 Spermidine showed preferential binding by the major groove at low cation concentration and by both major and minor grooves at higher concentration. The binding sites included N7 adenine and guanine, an interstrand contact between polyamine amino groups and thymine-O2, adenine-N3, and PO2- groups.21 Molecular models showed that the major groove of DNA duplex was the most favored binding site on DNA duplex at low polyamine concentration, while minor groove binding prevailed at high concentrations.21 These results are consistent with X-ray crystallographic studies of spermine-oligonucleotide complexes.65,66 Spermine binding sites included purine-N7, thymine-O2, and the backbone PO2 group.21 Theoretical calculations indicated that the interaction with the major groove of alternating purine/ pyrimidine sequences appeared to be the most favorable of all models presented, with a significant bending of the DNA duplex.67,68 It should be noted here that DNA remained in B-family structure upon its interaction with spermine, spermidine and putrescine.21 Based on our IR and CD spectroscopic results, polyamine analogues 333, BE-333, and BE-3333 bind DNA at guanine N7 site (major groove) and the backbone PO2 groups, both at low and high analogue concentrations. A partial B- to A-DNA transition occurs upon analogue-DNA complex formation, which is rather different from biogenic polyamineDNA complex formation, with no major alteration in the B-DNA conformation.21 Infrared data also show lower level of base binding for analogues compared to those of biogenic polyamine-DNA complexes.21 As a consequence, larger binding constants are estimated for biogenic polyamines than those of the analogue-DNA adducts (Table 2). The low stability of the analogue-DNA adduct can be due to a lower level of polyamine-base interactions and rapid aggregation of DNA in the presence of polyamine analogues. The order of stability of biogenic polyamine-DNA complexes is charge dependent: spermine > spermidine > putrescine.21 In the case of analogues, the binding constant of BE-333 is higher than that of BE-3333 (Table 2); however, the measurement of very low binding affinities is associated with large errors, and hence, binding constants of BE-333 and BE-3333 are within experimental errors. Several studies indicated that biogenic polyamines play a critical role in protecting DNA strand breaks induced by


radiation and oxidative stress.13,69,70 Our proposed models involve both intra- and interstrand interactions between natural polyamines and DNA.21 We believe that the intrastrand interaction would justify the ability of these polyamines to protect DNA against strand breaks. However, the low DNA binding affinity of analogues (in terms of the number of binding sites) and their ability to induce DNA aggregation and precipitation at a faster rate make them distinctly different from biogenic polyamines. The major differences observed herein for natural polyamine and analogues do not allow analogues to substitute for biogenic polyamine in their role as growth promoters, whereas analogues exert a growth inhibitory role and potential therapeutic effects on cancer cells. In conclusion, this is the first spectroscopic study on the binding mode of polyamine analogues with DNA, which explores the major differences between biogenic and synthetic polyamines in their ability to bind to specific sites on DNA. On the basis of our results, we can conclude that (a) due to rapid DNA aggregation and precipitation, the analogue-DNA binding sites are limited to the guanine N7 atom and the backbone phosphate group, while biogenic polyamines bind to the major and minor grooves as well as backbone phosphate group, (b) polyamine analogue-DNA complexes are less stable than natural polyamine-DNA complexes, and (c) A partial Bto A-DNA conformational transition occurrs due to the binding of polyamine analogues to DNA, whereas such a transition is not observed in biogenic polyamine-DNA complexes under the conditions of our experiment. These results might contribute to a better understanding of the mechanism of action of antitumor polyamine analogues.

Abbreviations 333: 1,1-diamino-4,8-diazaundecane · 4HCl BE-333: 3,7,11,15-tetrazaheptadecane · 4HCl BE-3333: 3,7,11,15,19-pentazahenicosane · 5HCl Put: putrescine Spd: spermidine Spm: spermine FTIR: Fourier transform infrared spectroscopy CD: circular dichroism Acknowledgment. We highly appreciate the financial supports of the Natural Sciences and Engineering Research Council of Canada (NSERC) for this work. The work in the Thomas laboratory was supported by grants from the National Institutes of Health through the National Cancer Institute (CA80163 and CA42439).

References and Notes (1) Huang, Y.; Keen, J. C.; Pledgie, A.; Marton, L. J.; Zhu, T.; Sukumar, S.; Park, B. H.; Blair, B.; Brenner, K.; Castro, R. A., Jr.; Davidson, N. E. J. Biol. Chem. 2006, 282, 19055–19063. (2) Huang, Y.; Keen, J. C.; Hager, E.; Smith, R.; Hacker, A.; Frydman, B.; Valasinas, A. L.; Reddy, V. K.; Marton, L. J.; Castro, R.A., Jr.; Davidson, N. E. Mol. Cancer Res. 2004, 2, 81–88. (3) Casero, R. A., Jr.; Woster, P. M. J. Med. Chem. 2001, 44, 1–26. (4) Gerner, E. W.; Meyskens, F. L., Jr Nat. ReV. Cancer 2004, 4, 781– 792. (5) Bergeron, R. J.; Neims, A. T.; McManis, J. S.; Hawthorne, T. R.; Vinson, J. R. T.; Bortell, R.; Ingeno, M. J. J. Med. Chem. 1988, 31, 1183–1190. (6) Bergeron, R. J.; McManis, J. S.; Liu, C. Z.; Feng, Y.; Weimar, W. R.; Luchetta, G. R.; Wu, Q.; Ortiz-Ocasio, J.; Vinson, J. R. T.; Kramer, D.; Porter, C. W. J. Med. Chem. 1994, 37, 3464–3474. (7) Davidson, N. E.; Mank, A. R.; Prestigiacomo, L. J.; Bergeron, J. R.; Casero, R. A., Jr Cancer Res. 1993, 53, 2071–2075.


2717

(8) Thomas, T.; Balabhadrapathruni, S.; Gallo, M. A.; Thomas, T. J. Oncol. Res. 2002, 13, 123–135. (9) Faaland, C. A.; Thomas, T. J.; Balabhadrapathruni, S.; Langer, T.; Mian, S.; Shirahata, A.; Gallo, M. A.; Thomas, T. Biochem. Cell Biol. 2000, 78, 415–426. (10) Thomas, T.; Thomas, T. J. J. Cell Mol. Med. 2003, 7, 113–126. (11) McCloskey, D. E.; Pegg, A. E. J. Biol. Chem. 2003, 278, 13881– 13887. (12) Ha, H. C.; Woster, P. A.; Yager, J. D.; Casero, R. A., Jr Proc. Natl. Acad. Sci. U.S.A. 1997, 94, 11557–11562. (13) Ha, H. C.; Sirisoma, N. S.; Kuppusamy, P.; Zweier, J. L.; Woster, P. M.; Casero, R. A., Jr Proc. Natl. Acad. Sci. U.S.A. 1998, 95, 11140– 11145. (14) Tabor, C. W.; Tabor, H. Annu. ReV. Biochem. 1984, 53, 749–790. (15) Pegg, A. E. Cancer Res. 1988, 48, 759–774. (16) Heby, O. Differentiation (Berlin) 1981, 19, 1–20. (17) Tkachenko, A. G.; Nesterova, L. Y. Biochemistry (Moscow) 2003, 68, 850–856. (18) Marton, L. J.; Pegg, A. E. Annu. ReV. Pharmacol. Toxicol. 1995, 35, 55–91. (19) Cohen, S. S. A Guide to the Polyamines; Oxford University Press: New York, 1988. (20) Thomas, T.; Thomas, T. J. Cell. Mol. Life Sci. 2001, 58, 244–258. (21) Ahmed Ouameur, A.; Tajmir-Riahi, H. A. J. Biol. Chem. 2004, 279, 42041–42054. (22) D′Agostino, L.; di Pietro, M.; Di Luccia, A. FEBS J. 2005, 272, 3777– 3787. (23) Ruiz-Chica, J.; Medina, M. A.; Sanchez-Jimeńez, F.; Ramı´rez, F. J. Biochim. Biophys. Acta 2003, 1628, 11–21. (24) Thomas, T. J.; Gunnia, U. B.; Thomas, T. J. Biol. Chem. 1991, 266, 6137–6141. (25) Bancroft, D.; Williams, L. D.; Rich, A.; Egli, M. Biochemistry 1994, 33, 1073–1086. (26) Thomas, T. J.; Messner, R. P. J. Mol. Biol. 1988, 201, 463–467. (27) Ruiz-Chica, J.; Medina, M. A.; Sanchez-Jimeńez, F.; Ramı´rez, F. J. Biochem. Biophys. Res. Commun. 2001, 285, 437–446. (28) Minyat, E. E.; Ivanov, V. I. I.; Kritzyn, A. M.; Minchenkova, L. E.; Schyolkina, A. K. J. Mol. Biol. 1978, 128, 379–409. (29) Jain, S.; Zon, G.; Sundaralingam, M. Biochemistry 1989, 28, 2360– 2364. (30) Real, A. N.; Greenall, R. J. J. Biomol. Struct. Dyn. 2004, 21, 469– 488. (31) Deng, H.; Bloomfield, V. A.; Benevides, J. M.; Thomas, G. J., Jr Nucleic Acids Res. 2000, 28, 3379–3385. (32) Ruiz-Chica, J.; Medina, M. A.; Sanchez-Jimeńez, F.; Ram; Ramı´rez, F. J. Biophys. J. 2001, 80, 443–454. (33) Gosule, L. C.; Schellman, J. A. Nature 1976, 259, 333–335. (34) Pelta, J., Jr.; Durand, D.; Doucet, J.; Livolant, F. Biophys. J. 1996, 71, 48–63. (35) Bloomfield, V. A. Biopolymers 1997, 44, 269–282. (36) Patel, M. M.; Anchoroquy, T. J. Biophys. J. 2005, 88, 2089–2103. (37) Flock, S.; Houssier, C. Biomol. Struct. Dyn. 1997, 15, 53–62. (38) Todd, B. A.; Parsegian, V. A.; Shirahata, A.; Thomas, T. J.; Rau, D. C. Biophys. J. 2008, 94, 4775–4782. (39) Feuerstein, B. G.; Williams, L. D.; Basu, H. S.; Marton, L. J. J. Cell. Biochem. 1991, 46, 37–47. (40) Sauve, D. M.; Anderson, H. J.; Ray, J. M.; James, W. M.; Roberge, M. J. Cell. Biol. 1999, 145, 225–235. (41) Laitinen, J.; Stenius, K.; Eloranta, T. O.; Ho¨ltta¨, E. J. Cell. Biochem. 1998, 68, 200–212. (42) Ordesson, S. M. Biochem. Soc. Trans. 2003, 31, 366–370. (43) Ruiz-Chica, A. J.; Medina, M. A.; Snachez-Jimenz, F.; Ramı´rez, F. J. Nucleic Acids Res. 2004, 32, 579–589. (44) Ramı´rez, F. J.; Thomas, T. J.; Antony, T.; Ruiz-Chica, J.; Thomas, T. Biopolymers 2002, 65, 148–157. (45) Saminathan, M.; Antony, T.; Shirahata, A.; Sigal, L. H.; Thomas, T.; Thomas, T. J. Biochemistry 1999, 38, 3821–3830. (46) Antony, T.; Thomas, T.; Shirahata, A.; Thomas, T. J. Biochemistry 1999, 38, 10775–10784. (47) Venkiteswaran, S.; Vijayanathan, V.; Shirahata, A.; Thomas, T.; Thomas, T. J. Biochemistry 2005, 44, 303–312. (48) Vijayanathan, V.; Thomas, T.; Shirahata, A.; Thomas, T. J. Biochemistry 2001, 40, 13644–13651. (49) Beauchemin, R.; N’soukpoe-Kossi, C. N.; Thomas, T.; Thomas, T. J.; Carpentier, R.; Tajmir-Riahi, H. A. Biomacromolecules 2007, 8, 3177– 3183. (50) Marmur, J. J. Mol. Biol. 1961, 3, 208–218. (51) Reichmann, M. E.; Rice, S. A.; Thomas, C. A.; Doty, P. J. Am. Chem. Soc. 1954, 76, 3047–3053.

2718


(52) Vijayalakshmi, R.; Kanthimathi, M.; Subramanian, V.; Balachandran, U. N. Biochem. Biophys. Res. Commun. 2000, 271, 731–734. (53) Alex, S.; Dupuis, P. Inorg. Chim. Acta 1989, 157, 271–281. (54) Stephanos, J. J. J. Inorg. Biochem. 1996, 62, 155–169. (55) Stephanos, J. J.; Farina, S. A.; Addison, A. W. Biochim. Biophys. Acta 1996, 1295, 209–221. (56) Taillandier, E.; Liquier, J. Methods Enzymol. 1992, 211, 307–335. (57) Di; Rico, D. E., Jr.; Keller, P. B.; Hartman, K. A. Nucleic Acids Res. 1985, 13, 251–260. (58) Loprete, D. M.; Hartman, K. A. Biochemistry 1993, 32, 4077–4082. (59) Andrushchenko,V, V.; Leonenko, Z.; van de Sande, H.; Wieser, H. Biopolymers 2000, 61, 243–260. (60) Dovbeshko, G. I.; Chegel, V. I.; Gridina, N. Y.; Shishov, O. P.; Tryndiak, Y. M.; Todor, V. P.; Solyanik, G. I. Biopolymers 2002, 67, 470–486. (61) Vorlickova, M. Biophys. J. 1995, 69, 2033–2043. (62) Kypr, J.; Vorlickova, M. Biopolymers 2002, 67, 275–277.

N’soukpoé-Kossi et al. (63) Kankia, B. I.; Bukin, V.; Bloomfiled, V. A. Nucleic Acids Res. 2001, 29, 2795–2801. (64) Nejedly, K.; Chladkova, J.; Vorlickova, M.; Hrabcova, I.; Kypr, J. Nucleic Acids Res. 2005, 33, 1–8. (65) Drew, H. R.; Dickerson, R. E. J. Mol. Biol. 1981, 151, 535–556. (66) Feuerstein, B. G.; Pattabiraman, N.; Marton, L. J. Nucleic Acids Res. 1990, 18, 1271–1282. (67) Feuerstein, B. G.; Pattabiraman, N.; Marton, L. J. Proc. Natl. Acad. Sci. U.S.A. 1986, 83, 5948–5952. (68) Bryson, K.; Greenall, R. J. J. Biomol. Struct. Dyn. 2000, 18, 393– 412. (69) Ha, H. C.; Yager, J. D.; Woster, P. A.; Casero, R. A., Jr Biochem. Biophys. Res. Commun. 1998, 244, 298–303. (70) Douki, T.; Bretonnier, Y.; Cadet, J. Radiat. Res. 2000, 153, 29–35.

BM800412R

A Comparison with Biogenic Polyamines - American Chemical Society

Recommend Documents