Intercalation and groove binding of an acridine–spermine conjugate on

DNA sequences: an FT–Raman and UV–visible absorption study

L. Perez-Floresa, A.J. Ruiz-Chicaa, J.G. Delcrosb, F. Sanchez-Jimenezc, F.J. Ramıreza,*

aDepartamento de Quımica Fısica, Facultad de Ciencias, Universidad de Malaga, Malaga 29071, SpainbGroupe Cycle Cellulaire, CNRS UMR 6061, 1FR97, Faculte de Medecine, Universite Rennes 1,

2 Avenue du Professeur Leon Bernand, 35043 Rennes Cedex, FrancecDepartamento de Biologıa Molecular y Bioquımica, Facultad de Ciencias, Universidad de Malaga, Malaga 29071, Spain

Received 6 September 2004; accepted 14 October 2004

Available online 28 January 2005

Abstract

Acridine and acridine derivatives are known as powerful DNA intercalators. Interactions of the acridine–spermine conjugate N1-(Acridin-

9-yl)-1,5,9,14,18-pentaazaoctadecane on two 16-mer oligonucleotides containing either alternating guanine–cytosine or adenine–thymine

sequences were studied by optical spectroscopies. UV–visible absorption spectra of oligonucleotide/conjugate solutions at different molar

ratios were recorded. The conjugate bands in the 350–500 nm region showed strong hypochromism and slight red shift in the presence of the

oligonucleotides, thus indicating that the acridine moieties intercalate into adjacent base pairs of the oligonucleotides. These effects stopped

near the 1:1 molar ratio, indicating that each oligonucleotide chain can only host one conjugate molecule. Raman spectra of solutions 60 mM

(in phosphate) of the oligonucleotides and 3 mM of the conjugate were also recorded. Upon intercalation, the spectra showed relevant

wavenumber shifts for skeletal and base vibrations, which have been largely attributed to the interactions of the positively charged side chain

groups with the reactive sites of the base residues. Raman data suggested the existence of sequence selectivity induced by the spermine tail.

Intercalation together to spermine interaction by the major groove was favoured for the guanine–cytosine sequence, while no groove

preference was achieved for the adenine–thymine sequence.

q 2004 Elsevier B.V. All rights reserved.

Keywords: GC; AT; Acridine; Spermine; UV–vis; Raman

1. Introduction

Biogenic polyamines spermine and spermidine are

essential molecules for cell proliferation and differentiation

in all living organisms [1]. In cells, they bind to nucleic

acids and proteins, acting as modulators of their macromol-

ecular conformations, stabilities and functions. It has been

demonstrated that tumoural cells need greater polyamine

amounts than their normal counterparts [2]. This fact makes

polyamines useful tools in anti-cancer strategies, since they

can act as vectors to introduce into cells covalently attached

DNA interacting anti-tumoural drugs [3,4]. This potential

0022-2860/$ - see front matter q 2004 Elsevier B.V. All rights reserved.

doi:10.1016/j.molstruc.2004.10.086

* Corresponding author. Tel.: C34 5 2132 258; fax: C34 5 132 000.

E-mail addresses: delcros@univ-rennes1.fr (J.G. Delcros), kika@

uma.es (F. Sanchez-Jimenez), ramirez@uma.es (F.J. Ramırez).

application has encouraged researchers to conjugate poly-

amines to known DNA intercalators or groove binders

because they are potent cytotoxic agents [4]. Polyamine–

drug conjugates would combine the great affinity of

the polyamines for DNA and the anti-proliferative action

of the drugs, thus decreasing secondary effects on non-

tumoural cells.

Acridine and acridine derivatives are known to have

chemotherapeutic activity, as they are DNA intercalators

[5]. Intercalation of planar molecules between the base pairs

causes the DNA helix to extend and decreases its winding

angle [6]. As demonstrated for several acridine derivatives

[7,8], these effects can inhibit the activity of topoisome-

rases, which are the enzymes responsible for the DNA

folding into cells [9]. Topoisomerase inhibition originates

that many DNA-involving processes are disrupted, so

leading to cell death.

Journal of Molecular Structure 744–747 (2005) 699–704

www.elsevier.com/locate/molstruc

Fig. 1. Chemical structure of the acridine–spermine conjugate N1-

(Acridin-9-yl)-1,5,9,14,18-pentaazaoctadecane.

L. Perez-Flores et al. / Journal of Molecular Structure 744–747 (2005) 699–704700

The following facts have been demonstrated for a

polyamine tail covalently attached to acridine: (i) it does

not alter the interacalating capability of the drug, (ii) it

enhances the DNA affinity of the conjugate, since the

polyamine side chain allows for additional groove binding

modes, (iii) it preserves the topoisomerase inhibitory

activity of acridine [10,11]. In a recent study, the effect of

the spermine conjugation on the cytotoxic and transport

properties of acridine has been reported [12]. Several

acridine–spermine conjugates were assayed, all of them

showing a high affinity for both DNA and the polyamine

transport system. Nevertheless, the structures of these

DNA–conjugates complexes in solution are not completely

known.

In this work we present an UV–visible and FT–Raman

study of the interaction of an acridine–spermine conjugate

on two 16-mer double-stranded oligonucleotides with either

the adenine–thymine (A–T) or the guanine–cytosine (G–C)

alternating sequence. This conjugate was constituted by a

spermine molecule and an acridine ring bridged by a

trimethylene-amino group, Fig. 1. Changes in both elec-

tronic absorption and vibrational Raman features of the two

sequences in the presence of the conjugate were analysed

and interpreted in order to propose preferential binding

modes between these molecules.

2. Experimental

2.1. Samples

The single-stranded 16-mer oligonucleotides d(GC)8 and

d(AT)8 were synthesized by Pharmacia-Biotech (Sweden).

The double-stranded oligonucleotides ds(GC)8 and ds(AT)8

were obtained and tested as previously reported [13,14].

Details about synthesis of the acridine–spermine conjugate

N1-(Acridin-9-yl)-1,5,9,14,18-pentaazaoctadecane are

given elsewhere [12]. To preserve physiological conditions,

10 mM TRIS buffer, 100 mM sodium chloride, was always

used as the solvent. Final pH was adjusted to 7.5 by using

hydrogen chloride.

2.2. Absorption spectroscopy

Absorption spectra were recorded at the room tempera-

ture in an Agilent 8453 spectrophotometer supplied with a

diode array detector. We used standard quartz cells, of 1 cm

path length, where 2 ml of a 2 mM solution in the conjugate

was placed. Once the first absorption spectrum was

achieved, 1 ml of a stock solution 40 mM (in phosphate)

of the oligonucleotide, either ds(GC)8 or ds(AT)8, was

added, followed by a new spectral acquisition. This

procedure allowed us to increase the oligonucleotide

concentration by 20 mM after each addition without

changing appreciably the total volume, thus preserving the

conjugate concentration. Features obtained for its absorp-

tion bands over the whole series of spectra are, therefore,

comparable. The final range of oligonucleotide concen-

trations studied was from 20 to 120 mM.

2.3. Raman spectroscopy

Fourier transform Raman spectra were recorded at room

temperature by means of a Bruker Equinox 55 spectrometer,

purged with dry nitrogen. Solutions at 3 mM of conjugate

concentration and 60 mM of oligonucleotide concentration

prepared for both ds(GC)8 and ds(AT)8. FT-Raman spectra

were recorded in a Bruker Equinox 55 Fourier-transform

spectrometer supplied with a Raman module. Spectra were

obtained at a resolution better than 2 cmK1, using the

excitation line at 1064 nm from a Nd-YAG laser working at

500 mw. Three microlitres of each solution were introduced

in a 0.8 mm diameter capillary. The capillary was then

placed into a spherical cuvette of 10 mm diameter, made of

sapphire, suitable for recording good quality spectra from

very low volumes of liquid sample.

Backscattering collection of the Raman radiation was

performed, and a minimum of 2000 scans were always

accumulated. Individual scans were examined by the

recording routine, being automatically discarded when the

mean intensity deviations were greater than 10% over

the full interferogram length. This procedure prevented

artefacts in the resulting spectra prior averaging, and was

performed at least with two independently prepared

samples for each concentration. Spectral treatment was

performed by using the Bruker OPUSq spectroscopic

software.

3. Results and discussion

3.1. Absorption spectroscopy

The electronic absorption spectrum of the conjugate is

shown in Fig. 2. It is dominated by the acridine bands, which

have been assigned on the basis of linear dichroism studies

[15]. All the bands correspond to in-plane p/p* electronic

transitions, since the out-of-plane n/p* transitions are too

weak to be observed, when compared with the in-plane

ones. In the ultraviolet region from 200 to 300 nm the

conjugate has intense peaks at 220, 259 and 265 nm. This

also is the region in which the DNA bases present their

absorption bands [16], so that it is not suitable to follow the

conjugate–oligonucleotide interaction. In the visible region

Fig. 2. Electronic spectrum of the acridine–spermine conjugate. Fig. 4. Electronic spectra of the conjugate, at a concentration 2 mM, in the

presence of increased concentrations of ds(AT)8: 0, 20, 40, 60, 80, 100 and

120 mM.

L. Perez-Flores et al. / Journal of Molecular Structure 744–747 (2005) 699–704 701

between 300 and 500 nm the spectrum of the conjugate

shows two band systems: two very weak bands, measured at

310 and 327 nm, which were assigned to electronic

transitions polarized along the acridine long-axis, and

three weak bands, namely at 432, 409 and 391 nm, assigned

to mixed polarizations. This region is free of base residue

absorptions, so that it was selected to monitor structural

changes on the acridine moiety of the conjugate when

interacting with nucleotides.

The electronic absorption spectra of the conjugate

between 350 and 500 nm in the presence of different

concentrations of ds(GC)8 and ds(AT)8 are shown in Figs. 3

and 4, respectively. The addition of both oligonucleotides to

a solution of the conjugate-induced slight red shift (bath-

ochromism) and a noticeable intensity reduction (hypo-

chromism) of the conjugate bands. These changes evidence

the existence of a strong interaction between the conjugate

and the oligonucleotides. More specifically, the hypochro-

mic effect has been associated to a stacking interaction

between the acridine moieties and the aromatic rings of the

bases [17]. This is explained since the inner region of a

Fig. 3. Electronic spectra of the conjugate, at a concentration 2 mM, in the

presence of increased concentrations of ds(GC)8: 0, 20, 40, 60, 80, 100 and

120 mM.

double helix is hydrophobic. When the acridine ring

intercalates, it goes from a strongly polar to a very low

polar medium, which would explains the observed

hypochromism.

The intensity reduction of the acridine bands when

adding ds(GC)8 or ds(AT)8 on the coujugate stopped at an

oligonucleotide concentration slightly greater than 60 mM,

in phosphate residues. As observed in Figs. 3 and 4, further

oligonucleotide charges did not induce noticeable changes

on the conjugate bands. In addition, the interactional

equilibrium produces isosbestic points at 440 nm for

ds(AT)8 and 441 nm for ds(GC)8, which suggests that

only two species of the conjugate exist in solution when

increasing the oligonucleotide–conjugate molar ratio.

Taking into account the oligonucleotide chain length

(16 base pairs), it can be deduced that 60 mM in phosphate

is approximately equivalent to 2 mM in oligonucleotide

molecules. Since the conjugate concentration is 2 mM, the

present results suggest that one 16-mer oligonucleotide

chain could only host one conjugate molecule.

The interpretation of these data in terms of conjugate–

oligonucleotide interaction can be summarized in two

points: (i) at conjugate–oligonucleotide molar ratios higher

than 1:1, the preferred binding mode likely intercalative; it

would explain the stable minimum of absorbance at 60 mM

in phosphate; (ii) at molar ratios lower than 1:1, both

intercalation and groove binding could be present, thus

explaining the isosbestic point. As groove binding does not

produce noticeable changes on the polarity of the acridine

surroundings, it will not significantly contribute to the

observed hypochromism. The hypothesis of groove binding

is clearly favoured by the presence of a spermine moiety in

the tail, since the affinity of the polyamines by the groove

reactive sites of DNA has been widely supported [1,2].

Nevertheless, the role of the aromatic ring in this interaction

mode is not clear, because it is expected that a groove

interaction does not significantly change the polar environ-

ment of the acridine ring.

Fig. 5. Raman spectra of the oligonucleotide ds(GC)8 alone (60 mM in phosphate; bold, down), ds(GC)8 in the presence of the conjugate (60 mM in phosphate

and 3 mM, respectively; bold, up), and the conjugate alone (3 mM; grey).

L. Perez-Flores et al. / Journal of Molecular Structure 744–747 (2005) 699–704702

3.2. Raman spectroscopy

We recorded Raman spectra of aqueous solutions

containing the acridine–spermine conjugate, at a concen-

tration 3 mM, and either the oligonucleotides ds(GC)8 or

ds(AT)8 at a concentration 60 mM (in phosphate). They are

shown in Figs. 5 and 6, respectively. At the light of the

results discussed in the precedent section, the conjugate

Fig. 6. Raman spectra of the oligonucleotide ds(AT)8 alone (60 mM in phosphate;

and 3 mM, respectively; bold, up), and the conjugate alone (3 mM; grey).

molecules would intercalate with site exclusion into

the oligonucleotide molecules. Consequently, the molar

ratio used in Raman experiments ensure a conjugate

concentration high enough to assume an almost complete

intercalation. Assignments were based on previous studies

[18–21], and they have been extensively discussed else-

where [13,14]. As a general feature common to both

oligonucleotides, intensity reductions were observed for

bold, down), ds(AT)8 in the presence of the conjugate (60 mM in phosphate

L. Perez-Flores et al. / Journal of Molecular Structure 744–747 (2005) 699–704 703

the bands assigned to the in-plane vibrations of the bases.

This effect has been described as the main consequence of

intercalative bindings on Raman spectra [22]. In addition,

some wavenumber shifts on the oligonucleotide bands were

observed upon conjugate addition. They largely correspond

to reactive sites of the aromatic rings of the bases, able to

interact with the positively charged ammonium groups of

the spermine moiety.

The more relevant wavenumber shifts measured in the

Raman spectra of ds(GC)8, Fig. 5, were: 1488/1485,

1452/1461, 1435/1433, 1258/1255, 1177/1171,

825/831, 722/726 and 709/702 cmK1. The intense

band at 1488 cmK1 is considered as a groove binding maker

band. It corresponds to a stretching vibration of the purine

rings, being largely assigned to the N7 atoms at the

oligonucleotide major groove [18,19]. Taking into account

that most of the acridine rings are intercalated at the molar

ratio used for recording these Raman spectra, the observed

shift should be a consequence of the interaction of the tails

with the base N7 positions. The bands measured at 1258 and

1177 cmK1 were also assigned to in-plane stretching

vibrations of the bases, involving both guanine and cytosine.

They shifted downwards by 3 and 6 cmK1, respectively.

The bands appearing between 1400 and 1500 cmK1 have

been assigned to bending vibrations of methyl and

methylene groups of the sugar moieties [18,19]. Changes

of these bands are usually attributed to hydrophobic

interaction with aliphatic groups of the ligands. The band

at 825 cmK1 has been assigned to an oligonucleotide

skeletal vibration, largely the phosphodiester, O–P–O,

stretching one. Insertion of a plane ligand between adjacent

base pairs provoke helix extension and unwinding that

involve the phosphodiester bridges [7]. It justifies the

measured shift (6 cmK1) upon the conjugate addition.

Bands between 700 and 800 cmK1 were assigned to in-

plane bending vibrations of the bases. The observed shifts

can be therefore caused by both intercalation of the acridine

moiety and groove binding of the spermine side chain.

The Raman bands of ds(AT)8 that showed relevant

wavenumber shifts are: 1461/1457, 1435/1439, 1304/1302, 1205/1210 and 793/796 cmK1. As in the case of

ds(GC)8, they correspond to in-plane stretching and bending

vibrations of the base residues [20,21]. Changes on groove

binding marker bands were not measured, while the skeletal

phosphodiester stretching band at 840 cmK1 exhibited a

negligible shift upon interaction. These facts suggest that

the conjugate could interact differently with AT and GC

sequences. However, intercalation is supported by the

acridine bands at 1597 and 700 cmK1, which shifted to

1591 and 709 cmK1, respectively, in the presence of

ds(AT)8. A vibrational dynamics of acridine, based on ab

initio force field calculations [23], assigned these bands to

stretching and bending in-plane vibrations, respectively,

which justifies the observed opposite behaviour.

In summary, the Raman spectra indicate that the

presence of a covalently attached spermine molecule to an

acridine ring could induce sequence selectivity on the

intercalation of this aromatic group on a double helix.

Selectivity on the interaction with nucleotide chains has

been observed for several acridine derivatives, as

methylene blue [24] or 9-amino-6-chloro-2-methoxyacri-

dine [25]. Our data suggest that intercalation together to

spermine interaction in the major groove is favoured for

guanine–cytosine sequences. Preferential interaction of

this biogenic polyamine to the major groove has been

demonstrated for genomic DNA [26] and GC sequences

[13], thus supporting this result. The interaction with

adenine–thymine sequences does not seem to have clear

groove preferences. Previous studies on biogenic poly-

amines [14] proposed that putrescine and spermidine

interact by the minor groove on a 15-mer adenine–

thymine oligonucleotide, while spermine could interact by

both the minor and the major grooves. Relevant Raman

changes were, nevertheless, observed at polyamine

concentrations rather greater than those used in this

work. This possibility was not able for the acridine–

spermine conjugate because the intense acridine Raman

signal would hide the oligonucleotide bands.

Acknowledgements

This work was supported by the Spanish Ministry of

Science and Technology, grants BQU2003-4168 (J.R.C. and

F.J.R.) and SAF2002-2586 (M.A.M. and F.S.J.). We would

also like to thank P.A.I. for support to groups FQM-159 and

CVI-267.

References

[1] S.S. Cohen, A Guide to the Polyamines, Oxford University Press,

New York, 1998.

[2] U. Bachrach, N. Seiler, Cancer Res. 41 (1981) 1205.

[3] C. Wang, J.G. Delcros, J. Biggerstaff, O. Phanstiel, IV, J. Med. Chem.

46 (2003) 2663.

[4] M. Demeunynck, F. Charmantray, A. Martelli, Curr. Pharm. Des. 7

(2001) 1703.

[5] A. Albert, The Acridines, Edward Arnold Ltd, London, 1996.

[6] S. Neidle, M. Waring, Molecular Aspects of Anticancer Drug–DNA

Interactions, CRC Press, Boca Raton, FL, 1993.

[7] M.J. Waring, B.A.J. Ponder, The Search for New Anticancer Drugs,

Kluwer Academic Publishers, Dordrecht, 1992.

[8] L.A. McDonald, G.S. Eldredge, L.R. Barrows, C.M. Ireland, J. Med.

Chem. 37 (1994) 3819.

[9] D.M. Morgan, Biochem. Soc. Trans. 18 (1990) 1080.

[10] I.S. Blagbrough, S. Taylor, M.L. Carpenter, V. Novoselskiy,

T. Shamma, I.S. Haworth, J. Chem. Soc., Chem. Commun. (1998);

929.

[11] L. Wang, H.L. Price, J. Juusola, M. Kline, O.I. Phanstiel, J. Med.

Chem. 44 (2001) 5590.

[12] J.G. Delcros, S. Tomasi, S. Carrington, B. Martin, J. Renault,

I.S. Blagbrough, P. Uriac, J. Med. Chem. 45 (2002) 5098.

[13] J. Ruiz-Chica, M.A. Medina, F. Sanchez-Jimenez, F.J. Ramırez,

Biochem. Biophys. Res. Commun. 285 (2001) 437.

L. Perez-Flores et al. / Journal of Molecular Structure 744–747 (2005) 699–704704

[14] J. Ruiz-Chica, M.A. Medina, F. Sanchez-Jimenez, F.J. Ramırez,

Biochim. Biophys. Acta 11 (2003) 1628.

[15] D. Fornasiero, T. Kurusev, Chem. Phys. Lett. 117 (1985) 176.

[16] K.E. Van Holde, W.C. Johnson, P.S. Ho, Principles of Physical

Biochemistry, Prentice-Hall, Englewood Cliffs, NJ, 1998.

[17] M. Nastasi, J.M. Morris, D.M. Rayner, V.I. Seligy, A.G. Szabo,

D.F. Williams, R.E. Williams, R.W. Yip, J. Am. Chem. Soc. 98

(1976) 3979.

[18] J.M. Benevides, G.J. Thomas Jr., Nucleic Acids Res. 11 (1983) 5747.

[19] J.M. Benevides, A.H. Wang, G.A. Van der Marel, J.H. Van Boom,

Biochemistry 28 (1989) 304.

[20] M. Katahira, Y. Nishimura, M. Tsuboi, Y. Sato, Y. Mitsui, Y. Iitaka,

Biochim. Biophys. Acta 867 (1986) 256.

[21] I. Movileanu, L.M. Benevides, G.J. Thomas Jr., J. Raman Spectrosc.

30 (1999) 637.

[22] G.J. Thomas Jr., M. Tsuboi, Adv. Biophys. Chem. 3 (1993) 1.

[23] I. Bandyopadhayay, S. Manogaran, J. Mol. Struct. (Theochem) 507

(2000) 217.

[24] E. Tuite, B. Norden, J. Am. Chem. Soc. 116 (1994) 7548.

[25] K. Fukui, K. Tanaka, Nucleic Acid Res. 24 (1996) 3962.

[26] J. Ruiz-Chica, M.A. Medina, F. Sanchez-Jimenez, F.J. Ramırez,

Biophys. J. 80 (2001) 443.