Spectroscopic studies of the interaction of aspirin and its important metabolite, salicylate ion, with DNA, A·T and G·C rich sequences

Article abstract of DOI:10.1016/j.saa.2010.08.078

Among different biological effects of acetylsalicylic acid (ASA), its anticancer property is controversial. Since ASA hydrolyzes rapidly to salicylic acid (SA), especially in the blood, interaction of both ASA and SA (as the small molecules) with ctDNA, oligo(dA·dT)₁₅ and oligo(dG·dC)₁₅, as a possible mechanism of their action, is investigated here. The results show that the rate of ASA hydrolysis in the absence and presence of ctDNA is similar. The spectrophotometric results indicate that both ASA and SA cooperatively bind to ctDNA. The binding constants (K) are (1.7 ± 0.7) × 10³ M^-1 and (6.7 ± 0.2) × 10³ M^-1 for ASA and SA, respectively. Both ligands quench the fluorescence emission of ethidium bromide (Et)-ctDNA complex. The Scatchard plots indicate the non-displacement based quenching (non-intercalative binding). The circular dichroism (CD) spectra of ASA- or SA-ctDsNA complexes show the minor distortion of ctDNA structure, with no characteristic peaks for intercalation of ligands. T_m of ctDNA is decreased up to 3 °C upon ASA binding. The CD results also indicate more distortions on oligo(dG·dC)₁₅ structure due to the binding of both ASA and SA in comparison with oligo(dA·dT)₁₅. All data indicate the more affinity for SA binding with DNA minor groove in comparison with ASA which has more hydrophobic character.

Full text of DOI:10.1016/j.saa.2010.08.078

Spectrochimica Acta Part A 77 (2010) 1077–1083
Contents lists available at ScienceDirect
Spectrochimica Acta Part A: Molecular and
Biomolecular Spectroscopy
journal homepage: www.elsevier.com/locate/saa
Spectroscopic studies of the interaction of aspirin and its important metabolite,
salicylate ion, with DNA, A·T and G·C rich sequences
a,∗
a
a
b
S.Z. Bathaie , L. Nikfarjam , R. Rahmanpour , A.A. Moosavi-Movahedi
a
Department of Clinical Biochemistry, Faculty of Medical Sciences, Tarbiat Modares University, P.O. Box: 14115-111, Tehran, Iran
Institute of Biochemistry and Biophysics, University of Tehran, Tehran, Iran
b
a r t i c l e i n f o
a b s t r a c t
Article history:
Received 28 May 2010
Received in revised form 1 August 2010
Accepted 26 August 2010
Among different biological effects of acetylsalicylic acid (ASA), its anticancer property is controversial.
Since ASA hydrolyzes rapidly to salicylic acid (SA), especially in the blood, interaction of both ASA and SA
(
as the small molecules) with ctDNA, oligo(dA·dT)15 and oligo(dG·dC)15, as a possible mechanism of their
action, is investigated here. The results show that the rate of ASA hydrolysis in the absence and presence
of ctDNA is similar. The spectrophotometric results indicate that both ASA and SA cooperatively bind
to ctDNA. The binding constants (K) are (1.7 ± 0.7) × 10 M and (6.7 ± 0.2) × 10 M for ASA and SA,
respectively. Both ligands quench the fluorescence emission of ethidium bromide (Et)–ctDNA complex.
The Scatchard plots indicate the non-displacement based quenching (non-intercalative binding). The
circular dichroism (CD) spectra of ASA– or SA–ctDsNA complexes show the minor distortion of ctDNA
Keywords:
3
−1
3
−1
Small molecules
DNA–ligand interaction
DNA structural change
Aspirin
◦
Salicylate
structure, with no characteristic peaks for intercalation of ligands. Tm of ctDNA is decreased up to 3 C
Circular dichroism
upon ASA binding. The CD results also indicate more distortions on oligo(dG·dC)15 structure due to the
binding of both ASA and SA in comparison with oligo(dA·dT)15. All data indicate the more affinity for SA
binding with DNA minor groove in comparison with ASA which has more hydrophobic character.
©
2010 Elsevier B.V. All rights reserved.
1
. Introduction
of cancer were reviewed elsewhere [19,20]. In addition, the reduc-
tion in neuronal function recovery [21] and neurotic outgrowth by
aspirin [22] were shown. Aspirin also activates some carcinogens
such as N-hydroxyarylamines to produce carcinogen-DNA adducts
[23–24].
Using the powder extracted from the bark of willow to treat
pain and reduce fever can be traced back to the fifth century BC.
In 1829, salicin (the parent of the salicylate) was isolated from wil-
low bark. Sodium salicylate was developed along with salicylic acid
Aspirin hydrolyzes spontaneously to salicylic acid through the
intramolecular general base catalysis by a carboxylate group [25].
The kinetic parameters for ASA hydrolysis were reported in differ-
(
SA) in 1875 as a pain reliever. In 1897 Felix Hoffman made aspirin
(
acetylsalicylic acid or ASA, Fig. 1), with reduced effect of irritat-
◦
ing the stomach. Today aspirin is used not only as a painkiller, but
also as an inhibitor of prostaglandin release. It also affects differ-
ent pathways and has been known for various pharmacological
properties, including: increasing blood vessel elasticity, acting as
an anti-inflammatory drug, inhibition of platelet aggregation and
blood clotting [1–4]. Furthermore, its beneficial effects on treat-
ment of cataract [5], sickle cell [6] and type I diabetes mellitus [7]
were reported.
ent media. The half-life of aspirin in rat blood at 37 C is about 0.21 h
(12.6 min) and it is hydrolyzed rapidly by arylesterases [26].
In 1968, it was reported that aspirin at therapeutic concentra-
tions, acetylates a variety of body constituents, including plasma
proteins such as albumin, IgG, IgA, transferrin [27]. Then, acetyla-
tion of endoperoxide synthesis (in cylooxygenase, Cox-1 and Cox-2)
[28–31], thus inhibition of prostaglandines biosynthesis [32] was
shown. Acetylating effect of ASA on some other proteins were
reported in the next years [5,6,33,34].
The anticancer effect of aspirin in some cancerous cell lines
[
[
[
8–10], through induction of apoptosis [11–16], inhibition of DNA
8,16] or protein synthesis [8], inhibition of proteasome function
17] and at higher doses, the inhibition of angiogenesis [18] have
On the other hand, the effect of salicylate ions, as the most
important metabolite of ASA, on the inhibition of palmitate oxi-
dation [35], growth of tumor cell lines [8,36] and angiogenesis in
endothelial cells with the same potency as aspirin, through Cox-
independent mechanism [18] were shown.
been reported. However, the controversial results about its bene-
fit on cancer treatment or prevention in human and animal model
According to the above mentioned results, it seems that acety-
lation is not the only mechanism of aspirin action and salicylate
ions also play an important role. Since, the exact mechanism of ASA
action and the molecular target of both ASA and SA are not clear yet,
∗ Corresponding author. Tel.: +98 21 8288 3850/3851; fax: +98 21 8288 4555.
E-mail address: zbatha2000@yahoo.com (S.Z. Bathaie).
1
386-1425/$ – see front matter © 2010 Elsevier B.V. All rights reserved.
doi:10.1016/j.saa.2010.08.078

1
078
S.Z. Bathaie et al. / Spectrochimica Acta Part A 77 (2010) 1077–1083
were 10 nm, ꢁex = 525 nm and ꢁem = 584 nm and no filter used. The
procedures for fluorometric studies of ctDNA–ethidium bromide
Et) complex in the absence and presence of ASA or SA have been
performed according to the previously reported method [39].
(
2.4. Circular dichroism (CD)
CD measurements were made on
a Jasco Model J-810
◦
CD recorder at 25 C. Data reported as molar ellipticity, [ꢂ]
2
−1
(
deg cm dmol ), based on the average weight of nucleotide was
equal to 330. The molar ellipticity for ctDNA was determined as
[
ꢂ] = (ꢂ × 330)/cl, where c is the ctDNA concentration in mg/ml, l is
ꢁ
the light path length in centimeter, and ꢂ is the measured ellip-
ticity in degrees at a wavelength of ꢁ. The molar ellipticity for
oligonucleotides was determined by using their molecular weight.
Fig. 1. The structural formula of SA (up) and ASA (down).
we attempt to investigate the interaction of these compounds with
3
. Results and discussion
DNA and the preference of their binding to A·T or G·C sequences
(
oligo(dA·dT) or oligo(dG·dC) ), as a possible mechanism for
1
5
15
3.1. Spectrophotometry
some of their actions.
Because of the rapid hydrolysis of aspirin in solutions, the kinetic
2
. Experimental
of ASA hydrolysis in the buffer condition was first investigated.
Some spectra at different concentrations of aspirin and at differ-
ent days up to a week were plotted. Three different peaks were
observed at 208 nm, 230 nm and 295.5 nm, as well as two isosbestic
points at 245 nm and 268 nm in the spectra belong to each concen-
tration at different day (Fig. 2A). The same set of spectra was plotted
in the presence of ctDNA (Fig. 2B).
Overally, the decrement at 260 nm (valley production) and the
increment in the peak at 295.5 nm are indicative of aspirin hydrol-
ysis and salicylate production as a function of time [40]. The
isosbestic point in the closed system indicates the conversion of
the reactants to the products with a constant ratio and it is due to
the linear changes of the different components in the reaction [41].
In other words, in this reaction aspirin has been converted directly
to salicylic acid.
2.1. Materials
Calf thymus DNA (ctDNA) was extracted by the method
explained previously [37]. Fiftheen bases olignucleotides (dA, dT,
dG and dC) were purchased from Cinagen. Co., Iran, Aspirin and
salicylic acid were obtained from Sigma, tris was from Merck, and
all of the other materials used were of analytical grade.
The experiments were carried out at pH 7.4; using 0.05 M tris
buffer, at room temperature in the absence and presence of 1 M
NaCl.
The ctDNA concentrations were determined using an extinction
−1
−1
coefficient of 6600 M cm at 260 nm and expressed in terms of
base molarity (Mb) or milli molar base (mMb) [38].
The kinetic parameters, including half-life (t1/2) and pseudo-first
order rate constant (k) for ASA hydrolysis calculated according to
the incremental changes in absorbency at 295.5 nm versus the time
at different concentrations of ASA in the absence (Fig. 2C) and pres-
ence of ctDNA (Fig. 2D) based on the Eqs. (1) and (2) [41]:
2
.2. Ultraviolet spectrophotometry
The spectrophotometric measurements were made with Shi-
madzu Model-3100 Double-beam Spectrophotometer. Since, both
aspirin and DNA show the absorption maximum at 260 nm, dif-
ferential spectroscopy was selected as a method of choice. By this
method a small change in the absorption is detectable. Thus, two
sets of experiments (a and b) were designed as follows:
v = k[ASA]
(1)
(2)
t1/2 = 0.693/k
By comparison of the UV-spectra of ASA, up to a week, in the
absence and presence of ctDNA (Fig. 2A and B), some changes in the
peaks are observed. They are including: the decrease in the peaks
at 260 nm and 208 nm accompanying with a red shift at 208 nm.
These changes indicate the ctDNA–ASA interaction. However, there
are no significant changes at 295 nm in the presence or absence of
ctDNA (Table 1) that indicate ctDNA has no effect on the rate of ASA
hydrolysis.
(
a) Two cuvettes (sample and blank) containing 2.5 ml of ctDNA
solutions, with the desired concentration, were titrated by lig-
and (ASA/SA) and buffer (10 ␮l in each injection), respectively.
The absorbency (A ) at each wavelength is related to the both
1
free and bound ligand.
(
b) The similar set of titration was done, using the cuvettes con-
taining 2.5 ml of buffer only. The absorbency (A ) at each
2
wavelength is related to the free ligand.
Our experiments also indicate that if aspirin is prepared daily,
it remains stable during the course of the reaction with almost no
significant amounts of salicylate formation.
(
c) By assumption that the absorbency of the free ligand is similar
in both sets of experiments, absorbance of the bound ligand
(
ctDNA–ligand complex) was obtained as: ꢀA = A1 − A2.
All calculations performed by the data processing mode of the
spectrophotometer’s software.
Table 1
Pseudo-first order rate constant and the half-life of aspirin hydrolysis in tris buffer
0.05 M, pH 7.4 in the presence and absence of DNA.
2.3. Spectrofluorometry
Substance
k (h−1)
t½ (h)
ASA–DNA
ASA
0.0188 ± 0.0016
0.0195 ± 0.0023
36.99 ± 2.83
35.79 ± 3.39
The Hitachi MPF-4 spectrofluorimeter, which was computer-
ized in our lab, was used. The scan speed was 60 nm/min, both slits

S.Z. Bathaie et al. / Spectrochimica Acta Part A 77 (2010) 1077–1083
1079
Binding parameters of ctDNA–ASA interaction are calculated
according to the changes at 260 nm (Fig. 3A). One can fit the ꢀA
vs. [ligand] curve to the following equation:
h
h
ꢀ
A = ꢀAmax((K[ligand]) /(1 + (K[ligand]) )
(3)
where h is the Hill coefficient, K is the equilibrium associa-
tion constant, and ꢀAmax is the maximum perturbation in the
ctDNA absorbance. Analyzing the data of ASA binding is shown
3
−1
in Fig. 3A gives values of 1.8 ± 0.3 for h, (1.7 ± 0.7) × 10 M for
K, and 0.079 ± 0.033 for ꢀAmax. A similar analysis for SA bind-
3
−1
ing gives values of 7.2 ± 1.3 for h, (6.7 ± 0.2) × 10 M for K, and
0
.0042 ± 0.0002 for ꢀAmax. The Hill coefficients indicate more
cooperativity in the SA–ctDNA interaction in comparison with ASA
binding to ctDNA, may be due to the steric hindranceof the addi-
tional acetyl group on ASA. The binding constants seem to be low,
but they are comparable with that reported for other ligands such
as
The cooperative binding constant (KCo), which was first defined
by Sano et al. [43], shows the binding strength regardless of the
cooperativity parameter. It is obtained as:
KCo = K × q
(4)
0
where k is the intrinsic (or obtained) binding constant and q is the
0
cooperative interaction parameter (or Hill coefficient). The average
KCo for SA and ASA interaction with ctDNA in this study is equal
4
−1
to (2.5 ± 0.3) × 10 M . This value is very similar to that recently
obtained by us using electrochemical method to investigate the
binding of ASA/SA with the immobilized ctDNA on the nanofiber
polypyrrole electrode (unpublished data).
3.3. Spectrofluorimetry
Ethidium bromide (Et) fluorescence emission is increased upon
interaction with ctDNA [44]. Both ASA and SA quench the flu-
orescence emission of ctDNA–Et complex with no effect on the
emission of Et alone. The Scatchard analysis of the fluorescence
data of ctDNA–Et complex in the absence and presence of different
concentrations of ASA or SA are calculated as follows [39]:
Fig. 2. Absorption spectroscopy. (A) Absorption spectra of ASA (0.125 mM) at 0 h,
4 h, 48 h, 72 h, 121 h and 144 h after preparation, at room temperature. (B) The
2
same as before but in the presence of 25 ␮g/ml DNA. (C) Absorbance changes of
different concentrations of ASA (0.125 mM (ꢀ), 0.25 mM (ꢁ), 0.375 mM (ꢀ), 0.5 mM
(
×) and 0.625 mM (᭹) at 295.5 nm against time, which are used for determination
Iobs = I + If
(5)
b
of kinetic parameters. The plots for higher concentrations of ASA, 0.625–1.25 mM,
are not shown. (D) The similar plots as (C) but in the presence of 0.147 mMb DNA.
Arrows show the direction of changes in the spectra by time.
where the observed emission at any time of the experiment is Iobs,
I and I are the fluorescence emissions of free and bound ethidium
f
b
bromide. Ib and I are functions of the concentration of bound (c )
f
b
and the remaining Et (c ).
f
3.2. Differential spectroscopy
Iobs = k c + k (c
− cb)
(6)
b
b
f
total
All molecules in this study (ctDNA, ASA and SA) have the
The constants kb and k are determined experimentally. c is
total
controlled and cb is determined by monitoring of Iobs using Eq. (7).
absorbance peak at 260 nm, thus differential spectroscopy is used
as the method of choice for investigating the interaction of ASA or
SA with ctDNA. Thus as explained in the parts (a) and (b) of Section
f
Iobs − k C
f
total
2
, interaction of different concentrations of ASA or SA with ctDNA
Cb =
(7)
k − k
b
f
were studied. Then, the changes in the absorbency (ꢀA) at different
wavelengths were calculated and plotted using the data processing
software of the UV–vis Shimadzu Spectrophotometer Model 3100,
as explained in part (c) of Section 2. The obtained data are plot-
ted in the Fig. 3A–C. An increase in the absorbance at 260 nm after
complex formation between ASA/SA with ctDNA is seen in Fig. 3A.
Both plots have the sigmoidal shape. The changes (decrement) in
the peak at 208 nm as well as the amount of the red shift (ꢀꢁ) of
the peak at this wavelength, versus the ligand concentration are
shown in Fig. 3B and C, respectively. All plots in Fig. 3 are indica-
tive of ctDNA interactions with both ASA and SA. The more red
shift observed at 208 nm in the presence of ASA is due to its more
hydrophobic character in comparison with SA. The red shift due to
more hydrophobic ligands was also reported elsewhere [42].
Once the concentration of the bound dye is known, the binding
of the Et may be described in terms of the Scatchard equation:
ꢃ
= nK − ꢃ K
(8)
Cf
where ꢃ is the ratio of bound Et to ctDNA base pair; n is the maxi-
mum value of ꢃ and K is the intrinsic binding constant of the Et.
Fig. 4A and B show the Scatchard plots of Et binding to ctDNA in
the absence and presence of different concentrations of ASA or SA,
respectively. These Figures indicate the changes in both the slope
(K) and the intercept (n) of the Scatchard plot of Et–ctDNA complex
upon the addition of ASA or SA (as a second ligand). The determined
values of the K and n are shown in Table 2.

1
080
S.Z. Bathaie et al. / Spectrochimica Acta Part A 77 (2010) 1077–1083
Fig. 3. The plots of the absorbency changes obtained by differential spectroscopy (as explained in Section 2): (A) at 260 nm for DNA–ASA (♦) or DNA–SA (ꢂ) complexes; (B) at
2
08 nm for DNA–ASA (♦) or DNA–SA (ꢂ) complexes; and (C) the amount of the red shift of the peak (ꢀꢁ) upon complex formation against the ligand concentration (similar
marks as A & B were used). [DNA] = 25 ␮g/ml in tris buffer 0.05 M, NaCl 1 M, pH 7.4.
Fig. 4. The Scatchard plots for Et–DNA complexes in the absence and presence of ASA (left) or SA (right) in tris buffer 0.05 M, NaCl 1 M, pH 7.4. The Scatchard plot in left are
shown as follows: Et–DNA complexes alone (ꢃ), as well as 0.18 mM of ASA (ꢁ) and 0.37 mM of ASA (᭹). The plots in right are Et–DNA complexes alone (ꢃ), 0.11 mM of SA
(ꢁ) and 0.29 mM of SA (᭹). At concentrations more than 0.53 mM of the ligands, no more changes were observed.

S.Z. Bathaie et al. / Spectrochimica Acta Part A 77 (2010) 1077–1083
1081
Table 2
Binding parameters for Et–DNA complex in the absence or presence of different
concentration of ASA or SA.
K (M 1
−
)
n
Ligand
mM
5
Et (alone)
ASA + Et
1.528 × 10
0.239
5
5
5
0.18
1.485 × 10
1.447 × 10
1.437 × 10
0.171
0.153
0.146
0
0
.37
.53
5
5
5
SA + Et
0.11
1.146 × 10
1.128 × 10
1.103 × 10
0.180
0.166
0.153
0
0
.29
.53
Quenching of the emission of Et–ctDNA complex in the presence
of various ligands was reported previously [39,45–50], and differ-
ent Scatchard plots have been obtained for the observed behaviors
[
[
39,51,52]. According to the classification of Howe-Grant et al.
51] the changes in both K and n have named as type B or non-
competitive behavior. The proposed mechanism for such behavior
is energy transfer from excited ctDNA bases not only to an inter-
calated Et through direct contact but also to that bind in a minor
groove [46,48–50,53]. The obtained results are also compatible
with the results obtained by Neault et al. that indicated the binding
of ASA to the backbone phosphate, as well as A–T and G–C base
pairs of ctDNA through hydrogen bonding [54].
Fig. 6. The hyperchromicity of DNA due to the raising in the temperature in the
absence (a) and prescence of different concentrations (0.125 mM, 0.25 mM, 0.5 mM,
0.75 mM, 0.875 mM, which were named b to f) of ASA.
confirmed in this study. It is important to note that they solved ASA
in methanolic solution, which may affect the ctDNA conformation.
In addition, the concentration of both ctDNA and ASA was very high
at that study. Interaction of aspirin with RNA, through the G–C and
A–U base pairs and the backbone PO2 group, was also shown. Such
interactionconserves the RNA structureintheA-conformation [57].
3
.4. Circular dichroism (CD)
Fig. 5 shows the CD spectra of ctDNA in the absence and pres-
ence of Et, ASA or SA. As it is seen a significant peak at 308 nm
is observed for ctDNA–Et complex (curve 4), which is a charac-
teristic peak for the intercalation of Et between ctDNA base pairs
[
55,56]. In addition, the broad and positive bands at 275 nm as well
3
.5. Tm determination
as increasing in the negative band at 248 nm are other reasons for
Et binding through intercalation [55,56]. The mentioned peaks are
absent in the CD spectra of ASA– or SA–ctDNA complexes (curves
The hyperchromicity of ctDNA with increasing temperature in
the absence or presence of different concentrations of ASA is also
investigated (Fig. 6) and the Tm of ctDNA is calculated according to
the method of Li using Eqs. (9) and (10) [58]. With 3 C decrease in
the Tm of ctDNA, it reached to 77 C in the presence of ASA (com-
pared to its initial value of 80 C in the absence of ASA). As it is
reported previously, reduction in the Tm is a characteristic of all
2
and 3). The small changes in the peaks at 275 nm and 248 nm are
only due to the minor structural changes in the ctDNA upon ASA
or SA bindings. These results confirm the non-intercalative mech-
anism of binding obtained by fluorescence data. However, the B-
to A-ctDNA transition that was shown by Neault et al. [54] is not
◦
◦
◦
known reagents that can form hydrogen bonds with nucleotides
[
59]. This result also confirms the non-intercalative mechanism of
binding and induction of minor instability on ctDNA structure upon
interaction with ASA [54].
A(t) − A (t)
0
h(t) =
h(t)
× 100
(9)
A
0
∂
h(t + 1) − (t − 1)
=
(10)
∂t
2
where h is the hyperchromicity, A is the absorbance at each tem-
perature and A0 is the absorbance at beginning of the experiment.
3.6. Preferential binding
To study the preference of these ligands for different ctDNA
sequences, their interaction with oligo(dA·dT)15 and oligo(dG·dC)15
are investigated by CD, Fig. 7.
A slight decrease in the ellipticity at 275 nm, accompanying with
an increase in the negative peak at 248 nm is observed in the CD
spectra of oligo(dA·dT)15 after interactions with ASA or SA. The sim-
ilar changes are observe in the CD spectra of oligo(dG·dC)15,but at
higher ligand/ctDNA ratio. These changes were leveled off after lig-
and/ctDNA molar ratio of about 2 in A·T sequence, but it began at
higher molar ratio and continued to more than 5.5 in G·C sequence.
Fig. 5. The spectropolatimetry. The CD spectra of DNA (0.112 mMb) alone (curve 1);
DNA–ASA (curve 2, 1.4 mM ASA); DNA–SA (curve 3, 1.4 mM SA and DNA–Et (curve
4
, 8.4 ␮M Et).

1
082
S.Z. Bathaie et al. / Spectrochimica Acta Part A 77 (2010) 1077–1083
Fig. 7. The CD spectra of oligo(dG·dC)15 titrated by ASA (G·C–ASA) and SA (G·C–ASA); and oligo(dA·dT)15 titrated by ASA (A·T–ASA) and SA (A·T–SA). The oligonucleotide
concentrations were 0.112 mMb and titration was done by ligands between 0 and 2.5 mM. In the case of G·C sequences, maximum changes were observed up to 2.13 mM of
ASA and 2.28 mM of SA, but in the presence of A·T, after 1.47 mM of both ligands there were no more changes in the spectra >230 nm. In all cases by addition of more ligands
the peaks below 230 nm destroyed completely (the plots not shown). In all figures the double stranded oligonuleotide is shown as black line, in the presence of ligand at the
concentrations named above are shown as gray bold lines, and in the presence of lower ligand concentrations are shown as thin gray lines.
It indicates that interaction of ASA or SA with A·T sequences is hap-
pened at lower ligand concentration. These results are similar to
that reported previously about interaction of ASA with ctDNA [54].
In addition, as it is seen in the figures (Fig. 7), there are some
perturbations in the ellipiticity of each of the mentioned oligonu-
cleotides at wavelengths below 240 nm. These changes are induced
in the GC sequence at lower molar ratio of ligand/ctDNA than A·T
sequence (0.92 and 1.13, respectively) and their intensities are
more. The perturbations at lower wavelengths on the CD spectra
may indicate some distortions on the hydrogen bonds [60].
of the non-intercalative ligands. By comparing these two ligands
with each other, it shows that interaction of SA with ctDNA is
more cooperative than ASA, but the latter induces more changes
on ctDNA structure.
Both ligands interact with A·T oligonucleotide at lower concen-
trations in comparison with G·C sequence, and the changes induced
by each of them (ASA or SA) in the A·T sequence are less than
that induced on G·C sequence. It means that the observed changes
in the G·C sequence is more serious and appeared at higher lig-
and/oligonucleotide molar ratio. The observed changes in the CD
signals below 240 nm indicate that both ASA and SA affect the
hydrogen bonding in the ctDNA structure, which is more in G·C
oligonucleotide.
Finally, based on the previous suggestion [21] and the results of
this study, a more thorough investigation concerning the possible
side effects of aspirin especially for its daily usage prescriptions
[
61], especially in children and during the pregnancy is suggested.
References
4
. Conclusion
[
[
1] J. Morley, Proc. R. Soc. Med. 7 (70 Suppl.) (1977).
2] P.-O. Hagen, Z.-G. Wang, E.M. Mikat, D.B. Hakel, Ann. Surg. 195 (1982).
Both ASA and SA, as two known small molecules, interact with
ctDNA through the groove binding. The ctDNA has no effect on ASA
hydrolysis. Both ASA and SA induce little instability and minor dis-
tortion on ctDNA structure, which is confirmed by the CD spectrum
[3] S.G. Albert, B.I. Hasnain, D.G. Ritter, J.H. Joist, D. Arshag, Diabetes Res. Clin. Pract.
0 (2005) 195–199.
4] N. Zimmermann, E. Gams, T. Hohlfeld, Eur. J. Cardiothorac. Surg. 34 (2008)
3–108.
[5] W. Qin, J.B. Smith, D.L. Smith, Biochim. Biophys. Acta 1181 (1993) 103–110.
7
[
9
◦
and 3 C decreases in the Tm of ctDNA. These are the characteristics

S.Z. Bathaie et al. / Spectrochimica Acta Part A 77 (2010) 1077–1083
1083
[
[
6] A.S.L. Xu, Y. Ohba, L. Vida, R.J. Labatka, R.E. London, Biochem. Pharmacol. 60
2000) 917–922.
7] A. Jafarnejad, S.Z. Bathaie, M. Nakhjavani, M.Z. Hassan, J. Pharmacol. Exp. Ther.
24 (2008) 850–857.
[31] T. Shimokawa, W.L. Smith, J. Biol. Chem. 267 (1992) 12387–12392.
(
[32] W.L. Smith, W.E. Mc Lands, J. Biol. Chem. 246 (1971) 6700–6704.
[33] I.M. Klotz, J.W.O. Tam, Proc. Natl. Acad. Sci. U.S.A. 70 (1973) 1313–1315.
[34] J.P. Bohney, M.L. Fonda, R.C. Feldhoff, FEBS Lett. 298 (1992) 266–268.
3
[
[
8] T.C. Viljoen, C.H. van Aswegen, D.J. du Plessis, Oncology 52 (1995) 465–469.
9] S.J Shiff, M.I. Koutsos, L. Qiao, B. Rigas, Exp. Cell Res. 222 (1996) 179–188.
[35] J.F.T. Glasgow, B. Middleton, R. Moore, A. Gray, J. Hill, Biochem. Biophys. Acta
1454 (1999) 115–125.
[
10] P. Ricchi, S. Pignata, A di Popolo, A. Memoli, A. Apicella, R. Zarrilli, A.M. Acqua-
[36] D.J. Elder, A. Hague, D.J. Hicks, C. Paraskeva, Cancer Res. 56 (1996) 2273–2276.
[37] S.Z. Bathaie, A.A. Moosavi-Movahedi, A.A. Saboury, Nucleic Acids Res. 27 (1999)
1001–1005.
[38] S. Aktipis, A. Kindelis, Biochemistry 12 (1973) 1213–1221.
[39] K.G. Strothkamp, R.E. Strothkamp, J. Chem. Educ. 71 (1994) 77–79.
[40] G. Alibrandi, N. Micali, S. Trussoand, A. Villari, J. Pharm. Sci. 85 (1996)
1105–1108.
[41] J.W. Moore and R.G. Pearson, John Wiley & Sons, New York. (1981).
[42] N. Mudasir, Yoshioka, H. Inoue, J. Inorg. Biochem. 77 (1999) 239–247.
[43] T. Sano, T. Nakatsuka, H. Tsuruta, S. Harada, Bull. Chem. Soc. Jpn. 60 (1987)
759–760.
[44] J.B. LePecq, C. Paoletti, J. Mol. Biol. 27 (1967) 87–106.
[45] P. Fromherz, B. Rieger, J. Am. Chem. Soc. 108 (1986) 5361–5362.
[46] R.F. Pasternack, M. Caccam, B. Keogh, T.A. Stephenson, A.P. Williams, E.J. Gibbs,
J. Am. Chem. Soc. 113 (1991) 6835–6840.
[47] A.K. Paul, H. Mansuri-Torshizi, T.S. Srivastava, S.J. Chavan, M.P. Chitnis, J. Inorg.
Biochem. 50 (1993) 9–20.
[48] K.-M. Hyun, S.-D. Choi, S. Lee, S.K. Kim, Biochim. Biophys. Acta 1334 (1997)
312–316.
[49] A. Arabzadeh, S.Z. Bathaie, H. Farsam, M. Amanlou, A.A. Saboury, A. Shockravi,
A.A. Moosavi-Movahedi, Int. J. Pharm. 237 (2002) 47–55.
[50] S.Z. Bathaie, A. Bolhasani, R. Hoshyar, B. Ranjbar, F. Sabouni, A.A. Moosavi-
Movahedi, DNA Cell Biol. 26 (2007) 533–540.
[51] M. Howe-Grant, K.C. Wu, W.R. Baue, S.J. Lippard, Biochemistry 15 (1976)
4339–4346.
viva, Int. J. Cancer 73 (1997) 880–884.
[
[
11] C.J. Barnes, I.L. Cameron, W.E. Hardman, M. Lee, Br. J. Cancer 77 (1998) 573–580.
12] L. Qiao, R. Hanif, E. Sphicas, S.J. Shiff, B. Rigas, Biochem. Pharmacol. 55 (1998)
5
3–64.
[
[
[
13] C. Sotiriou, M. Lacroix, L. Langneaux, G. Berchem, J. Jacques Body, Anticancer
Res. 19 (1999) 2997–3006.
14] L. Deng, S. Hu, A.R. Baydoun, J. Chen, X. Chen, X. Cong, Cell Prolif. 42 (2009)
7
21–730.
15] Q. Gu, J.D. Wang, H.H.X. Xia, M.C.M. Lin, H. He, B. Zou, S. Ping Tu, Y. Yang, G.
Liu, X. Kum, S. Lam, W. Man Wong, A.O.O. Chan, M. Fung Yuen, H. Fu Kung, B.
Chun-Yu Wong, Carcinogenesis 26 (2005) 541–546.
[
[
[
[
[
[
[
[
[
16] E. Castano, M. Calmau, M. Marti, F. Berrocal, R. Bartrons, J. Gil, J. Pharmacol. Exp.
Ther. 280 (1997) 366–372.
17] P. Dikshit, M. Chatterjee, A. Goswami, A. Mishra, N.R. Jana, J. Biol. Chem. 281
(
2006) 29228–29235.
18] G.M. Borthwick, A.S. Johnson, M. Partington, J. Burn, R. Wilson, H.M. Arthur,
FASEB J. 20 (2006) 2009–2016.
19] C. Bosetti, S. Gallu, C. La Vecchia, Recent Results Cancer Res. 181 (2009)
31–251.
20] P.C. Elwood, A.M. Gallaghar, G.G. Duthie, L.A.J. Mur, G. Morgan, Lancet 373
2009) 1301–1309.
21] K.I. Maynard, P.M. Arango, D. Chen, C.S. Ogilvy, Neurosci. Lett. 249 (1998)
59–162.
22] F. Sabouni, M. Firouzi, M. Taghikhani, A.A. Ziaee, S. Semnanian, Biochem. Bio-
phys. Res. Commun. 248 (1998) 165–167.
23] R.F. Minchin, K.F. Ilett, C.H. Teitel, P.T. Reeves, F.F. Kadlubar, Carcinogenesis 13
1992) 663–667.
24] W.G. Humphreys, F.F. Kadlubar, F.P. Guengerich, Proc. Natl. Acad Sci. U.S.A. 89
1992) 8278–8282.
2
(
1
[52] B.C. Baguley, M. Le Bret, Biochemistry 23 (1984) 937–943.
[53] T.G. Gantchev, H. Ali, E. van Lier, Eur. J. Biochem. 217 (1993) 371–376.
[54] J.F. Neault, M. Naoui, M. Manfait, H.A. Tajmir-Riahi, FEBS Lett. 382 (1996)
26–30.
[55] S. Aktipis, W.W. Martz, A. Kindelis, Biochemistry 14 (1975) 326–331.
[56] M.J. Carvlin, E. Mark, R. Fiel, Nucleic Acids Res. 11 (1983) 6141–6154.
[57] J.F. Neault, H.A. Tajmir-Riahi, J. Phys. Chem. B 101 (1997) 114–116.
[58] H.J. Li, Methods Cell Biol. 18 (1978) 385–396.
(
(
[
[
[
[
25] A.R. Fersht, A.J. Kirby, J. Am. Chem. Soc. 89 (1967) 857–4863.
26] S.K. Bakar, S. Niazi, J. Pharm. Sci. 72 (1983) 1024–1025.
27] R.N. Pinckard, D. Hawkins, R.S. Farr, Nature 219 (1968) 68–69.
28] F.J. van der Ouderaa, M. Buytenhek, D.H. Nugteren, D.A. van Dro, Eur. J. Biochem.
[59] C.R. Cantor, in: W. Saenger (Ed.), Springer-Verlag, New York, 1983, pp. 126–149.
[60] D.M. Gray, R.L. Ratliff, M.R. Vaughan, in: D.M.J. Lilley, J.E. Dahlberg (Ed.). Aca-
demic Press, Inc. New York, 1992.
1
09 (1980) 1–8.
[
[
29] G.J. Roth, E.T. Machuga, J. Ozols, Biochemistry 22 (1983) 4672–4675.
30] C. Yokoyama, T. Takai, T. Tanabe, FEBS Lett. 231 (1988) 347–351.
[61] D.P.M. Symmons, Lancet 348 (1996) 1394–1395.