Synthesis and fluorescence study of 7-azaindole in DNA oligonucleotides replacing a purine base

Article abstract of DOI:10.1016/S1386-1425(02)00004-5

The fluorescence spectroscopy of 7-azaindole (^7aIn) incorporated in DNA oligonucleotides is investigated. Incorporation of ^7aIn into DNA oligonucleotides is accomplished through standard solid-phase phosphoramidite chemistry. Fluores

Full text of DOI:10.1016/S1386-1425(02)00004-5

Spectrochimica Acta Part A 58 (2002) 2595–2603
www.elsevier.com/locate/saa
Synthesis and fluorescence study of 7-azaindole in DNA
oligonucleotides replacing a purine base
Ke Wang a, Sandra Stringfellow b, Shiming Dong b, Yugao Jiao b,
Hongtao Yu b,*
a Polymer Research Institute, Sichuan Union Uni6ersity, Chengdu, China
b Department of Chemistry, Jackson State Uni6ersity, Jackson, MS 39217, USA
Received 15 November 2001; accepted 17 December 2001
Abstract
The fluorescence spectroscopy of 7-azaindole (7aIn) incorporated in DNA oligonucleotides is investigated. Incorpo-
ration of 7aIn into DNA oligonucleotides is accomplished through standard solid-phase phosphoramidite chemistry.
Fluorescence emission of the 7aIn chromophore shifts slightly to the red (from 386 nm to 388 nm) upon glycosylation
at the N−1 position, but its relative fluorescence quantum yield increases 23 times, from 0.023 to 0.53. Upon
incorporation into DNA, the fluorescence emission of 7aIn is greatly quenched with fluorescence quantum yields of
0.020 and 0.016 in single and double strand DNA, respectively. The fluorescence emission for 7aIn in DNA
oligonucleotides shifts to the blue with an emission maximum at 379 nm. Both the strong fluorescence quenching and
the blue shift of the emission spectrum signify that 7aIn is stacked with neighboring DNA bases in both single and
double strand DNA. As the duplex DNA melts due to temperature increase, the fluorescence of the 7aIn chromophore
increases, indicating the transition from the less fluorescent duplex DNA to the more fluorescent single strand DNA.
Since this fluorescent 7aIn is a structural analog of purine, its fluorescence property may be utilized as a probe for
studying nucleic acid structure and dynamics. © 2002 Elsevier Science B.V. All rights reserved.
Keywords: 7-Azaindole; DNA; Fluorescence; Synthesis
such as DNA-protein interaction and DNA struc-
ture and dynamics [1–27]. Research findings in
1. Introduction
this area have provided some important informa-
The use of a non-natural ‘base’ to replace one
tion toward understanding our living systems at
of the natural nucleic acid bases in DNA has been
the molecular level. There are mainly two groups
an important tool for studying biological events
of ‘bases’ synthesized and incorporated into DNA
oligomers: those made by minor modification of
the natural bases or the derivatives of cyclic aro-
matic compounds. While the former includes 2-
* Corresponding author. Tel.: +1-601-979-3727; fax: +1-
aminopurine [1–8], inosine [9] and isoinosine
[9–11], 3- and 7-deazaadenine [12–15], and 3- and
601-979-3674
E-mail address: yu@ccaix.jsums.edu (H. Yu).
1386-1425/02/$ - see front matter © 2002 Elsevier Science B.V. All rights reserved.
PII: S1386-1425(02)00004-5

2596
K. Wang et al. / Spectrochimica Acta Part A 58 (2002) 2595–2603
7-deazaguanine [14,16], the latter includes analogs
of pteridine [17–19] and indole [20–24], benzene
[23–26], naphthalene and pyrene derivatives [24].
The first group of compounds, when replacing a
natural base in duplex DNA, can still form hydro-
gen bonds with the base in the opposite strand,
while the second group of compounds cannot
form any hydrogen bonds.
Fluorescence has been one of the very useful
techniques used for studying DNA structure and
dynamics. The fluorescence properties of the
above base analogs have been utilized for study-
ing DNA structure and duplex stability [1,2,10,14,
with CaH2 and distilled under nitrogen atmo-
sphere. 7aIn, obtained from ICN Biochemicals,
was recrystallized from alcohol and dried in vacuo
before use. 1-(a)-Chloro-3,5-di-O-(p-toluoyl)-2-
deoxy-D-ribose was purchased from Berry and
Associates, Inc. (Ann Arbor, MI) and used di-
1
rectly. H- and 31P-NMR spectra were recorded
on a Bruker AM250 MHz NMR spectrometer at
ambient temperature. The NMR spectra were in
ppm with tetramethylsilane as internal standard.
High-resolution mass spectrometric analyses
(HRMS) were performed at Louisiana State Uni-
versity’s Mass Spectrometry Facility using the
Fast Atomic Bombardment (FAB) technique.
DNA oligonucleotides containing 7aIn in the place
of either an adenine or a guanine were synthesized
by Gemini Biotech (Alachua, FL) using the phos-
phoramidite 4. The 4-step synthesis of 4 is shown
in Fig. 1.
Fluorescence emission spectra were recorded
using a Fluoromax-2 spectrofluorometer from In-
struments SA Inc. (Edison, NJ). Sample solutions
were prepared in 1×TBE buffer in the presence
of 0.2 M sodium chloride. The quantum yields
were determined using indole as a standard. The
excitation wavelength was set at 300 nm for 7aIn.
The sample temperature was controlled by an
external water bath and the actual solution tem-
perature was determined using a thermometer
with a temperature probe.
15,22,26],
protein-DNA
interactions
[4–
8,13,18,25], or simply as universal base analogs
[20,27]. Some of the unusual ‘bases’ are used as
anti-HIV or anti-cancer drugs [28–30].
One interesting fluorescent molecule used for
studying protein structure and dynamics is 7-
azaindole (7aIn) [31–38]. 7aIn is weakly fluorescent
in water due to dimer formation [34–36]. The
excitation energy of 7aIn is lost due to double
proton transfer, and therefore quenches the
fluorescence. After methylation of the N−1 ni-
trogen to prevent dimer formation, 7aIn becomes
strongly fluorescent. The fluorescence property of
7aIn, as well as its 3-alanyl derivative, 7-azatryp-
tophan, is well understood. It has been incorpo-
rated into peptides to obtain peptide/protein
structural information [31,33,39]. Due to the
unique fluorescence properties of this molecule,
we wish to report the incorporation of 7aIn into
duplex DNA to examine its fluorescence proper-
ties in 2%-deoxyriboside, in single and double
strand DNA, and the fluorescence change during
DNA duplex thermomelting.
2.2. Synthesis of 7-azaindole phosphoramidite 4
2.2.1. Glycosylation of 7aIn with 1-(h)-chloro-3,5-
di-O-(p-toluoyl)-2-deoxy-D-ribose (modified
procedure from Refs. [11,20,40,41])
Sodium hydride (475 mg, 11.9 mmol, 60% in
oil) was added to 86 ml of anhydrous acetonitrile
containing 7aIn (1.30 g, 11.0 mmol). The mixture
was stirred for 2 h at room temperature under
nitrogen atmosphere. Then the mixture was
cooled to −10 °C using an ice salt bath before
2. Experimental
2.1. Materials and instruments
Chemicals and solvents were purchased from
either Aldrich Chemical Company or Fisher Sci-
entific. All solid compounds were dried in vacuo
over phosphorous pentoxide. Anhydrous solvents
from Aldrich were used without further treat-
ment, except for acetonitrile, which was dried
1-(a)-chloro-3,5-di-O-(p-toluoyl)-2 - deoxy-D-ri-
bose (5.13 g, 13.2 mmol) was added portionwise
while stirring. The solution was kept at −10 °C
under nitrogen atmosphere for 3 h before it was
filtered to remove a small amount of insoluble
material. The filtrate was evaporated to give an

K. Wang et al. / Spectrochimica Acta Part A 58 (2002) 2595–2603
2597
MH+: 471.1935. Calc. 471.1935 (M+1) for
C28H26N2O5.
oily crude product, which was purified by silica
gel chromatography (column: 33×345 mm2) elut-
ing with ethyl acetate/hexanes (50:50). After evap-
oration of the solvent, 3.42 g white, foamy
product, 1%-(7-azaindolyl)-3%,5%-di-O-(p-toluoyl)-2%-
2.2.2. Synthesis of 2 by the hydrolysis of 1
Product 1 (3.42 g, 7.26 mmol) was added to 40
ml of methanol containing sodium methoxide pre-
pared by dissolving 75 mg of sodium (3.3 mmol)
in anhydrous methanol prior to the addition. The
reaction mixture was stirred at room temperature
for 3 h while large amounts of product precipi-
tated. After completion of the reaction, solid
NH4Cl was added to the solution to adjust its pH
to basic (pH 8) before it was filtered to collect the
deoxy- -riboside (1, 66%), was obtained. Rf=
D
0.75 (ethyl acetate/hexanes 50:50). 1H-NMR
(DMSO-d6, ppm): 8.26 (1H, d, H-C6), 7.98 (1H,
d, H-C4), 7.72 (1H, d, H-C2), 7.13 (1H, q, H-C5),
6.70 (1H, dd, H1%), 6.52 (1H, d, H-C3), 5.25
(0.5H, OH), 5.04 (0.5H, OH), 4.42 (1H, m, H3%),
3.85 (1H, m, H 4%), 3.55 (2H, m, H5% and H5¦),
2.61 (1H, m, H 2%) and 2.20 (1H, m, H2¦). HRMS:
Fig. 1. Synthesis of 1%-(7-azaindolyl)-2%-deoxy-D-riboside and the incorporation into DNA via the phosphoramidite chemistry.

2598
K. Wang et al. / Spectrochimica Acta Part A 58 (2002) 2595–2603
majority of the product, 1%-(7-azaindolyl)-2%-de-
oxy- -riboside (2), as a white powder. A further
portion of the product was crystallized from the
mmol) and 1.2 ml of diisopropylethylamine (6.9
mmol) under nitrogen atmosphere. The reaction
mixture was allowed to stir for 1 h at room
temperature before it was poured into 30 ml of
5% NaHCO3 aqueous solution and extracted with
CH2Cl2 (3×20 ml). The combined organic layers
were dried over anhydrous Na2SO4 and concen-
trated to produce the crude product, which was
purified by silica gel chromatography (column:
20×245 mm) eluting with ethyl acetate/hexanes
(70:30). After evaporation of the solvent, the
phosphoramidite 4, 1%-(7-azaindolyl)-5%-O-(4,
D
filtrate and the combined yield was 75% (1.28 g,
1
5.45 mmol). H-NMR (CDCl3, ppm): 8.30 (1H, d,
H-C6), 7.89 (1H, d, H-C4), 7.38 (1H, d, H-C2),
7.32 (9H, m, Ph), 7.09 (1H, q, H-C5), 6.92 (1H, t,
H1%), 6.82 (4H, m, Ph), 6.46 (1H, d, H-C3), 4.65
(1H, m, H3%), 4.08 (1H, m, H4%), 3.80 (6H, d,
2×CH3O), 3.38 (2H, m, H5% and H5¦), 2.68 (1H,
m, H2%) and 2.48 (1H, m, H2¦). HRMS: MH+:
235.1100; Calc. 235.1005 (M+1) for C12H14N2O3.
4%-dimethoxytrityl)-2%-deoxy-D-riboside-3%-O-(2-
2.2.3. Synthesis of 3 by the
4,4%-dimethoxytritylation of 2
cyano - ethyl ) - N,N - diisopropylphosphoramidite,
was collected as a white foam in 77% yield (390
mg, 0.53 mmol). Rf=0.76 (ethyl acetate/hexanes/
triethylamine 7:3:1). It separated into two spots
with nearly equal intensity on TLC with a less
polar solvent: Rf=0.38 and 0.44 (ethylacetate:
hexanes: triethylamine 3:7:1). The broadband de-
coupled 31P-NMR (CDCl3) gives two signals at
149.67 and 149.49 ppm with about a 1:1 intensity
ratio. The 1H-NMR spectrum is complex and
shows a mixture of two co-existing diastereomers.
4,4-Dimethoxytrityl chloride (745 mg, 2.20
mmol) was added to 14 ml of anhydrous pyridine
containing 6 mg of 4-(N,N-dimethylamino)-pyri-
dine (DMAP) and 469 mg of 2 (2.00 mmol) which
was dried over P2O5 in vacuo overnight before
use. The mixture was then stirred for 2.5 h at
room temperature before 10 ml of methanol was
added to quench the reaction. The resulting mix-
ture was poured into 70 ml of 5% aqueous
NaHCO3 solution and the mixture emulsified. The
emulsion was extracted using CH2Cl2 (3×40 ml)
and the combined organic layers were dried over
anhydrous Na2SO4. After evaporation of the sol-
vent, an oily product was obtained and purified
by silica gel chromatography (column: 20×310
mm2) eluting with ethyl acetate/hexanes (70:30) to
2.3. Preparation of DNA oligonucleotides
containing 7aIn
Phosphoramidite 4 was used directly for auto-
mated DNA synthesis performed by scientists at
Gemini Biotech (Alachua, FL). The synthesis was
carried out on a 1.0 mmol scale. All of the synthe-
ses were optimized for trityl release and all
oligonucleotides were HPLC purified. The se-
quence of the synthetic DNA oligonucleotides
are: Oligo-1: 5%-GCGCGC7aInCGCGCGC-3%
(self-complimentary); Oligo-2: 5%-CGC7aInGAAT-
TC-3%; Oligo-3: 5%-CGG7aInAATTC-3%; (matching
strand: 5%-GAATTCCCG-3%). The concentration
of Oligos 1–3 was determined by their absorption
value at 260 nm using Beer’s Law: C=A260/ml,
where A260 is the absorbance of the oligomer at
260 nm; m is the molar absorption extinction co-
efficient at 260 nm and l is the cell path length (1
cm). The m was calculated based upon the follow-
ing equation: m=0.89 [(A×15480)+(C×
7340)+(G×11760)+(T×8850)], where A, C, G,
T=number of A, C, G, and T bases in the
produce
trityl)-2%-deoxy-
1%-(7-azaindolyl)-5%-O-(4,4%-dimethoxy-
-riboside (3) as a yellowish foam
D
in 75% yield (805 mg, 1.50 mmol). Rf=0.59 (ethyl
1
acetate/hexanes 70:30). H-NMR (CDCl3, ppm):
8.30 (1H, d, H-C6), 7.89 (1H, d, H-C4), 7.38 (1H,
d, H-C2), 7.32 (9H, m, Ph), 7.09 (1H, q, H-C5),
6.92 (1H, t, H1%), 6.82 (4H, m, Ph), 6.46 (1H, d,
H-C3), 4.65 (1H, m, H3%), 4.08 (1H, m, H4%), 3.80
(6H, d, 2×CH3O), 3.38 (2H, m, H5% and H5¦),
2.68 (1H, m, H2%) and 2.48 (1H, m, H2¦). HRMS
(FAB-MS): MH+: 537.2402; Calc. 537.2312
(M+1) for C33H32N2O5.
2.2.4. Synthesis of phosphoramidite 4
2-Cyanoethyl diisopropylaminochlorophospho-
ramidite (620 ml, 2.76 mmol) was added to 10 ml
of anhydrous CH2Cl2 containing 3 (370 mg, 0.69

K. Wang et al. / Spectrochimica Acta Part A 58 (2002) 2595–2603
2599
oligomer. The coefficient for 7aIn was determined
to be 4400 at 260 nm. Double stranded Oligo-3 was
prepared by matching equimolar amounts of the
two complimentary strands followed by annealing.
The buffer used for all DNA samples were the
1×TBE buffer (89 mM Tris base, 89 mM boric
acid, 0.1 mM EDTA, pH 8.2) with 200 mM NaCl.
compound 3: chlorophosphoramidite: diisopropy-
lethylamine under our conditions, compared to
1:2:4 by Seela and Chen [10] for isoinosine or 1:6:4
by Cosstick et al. [13] for 3-deazaadenosine. The
large amount (10-equivalent) of diisopropylethy-
1
lamine proved to be necessary in our case. H-
NMR spectrum of 4 is very complex because
compound 4 exists as a diastereomeric mixture
since the phosphorous atom is chiral and it adopts
both an R and S-configuration. Actually, the two
diastereomers can be isolated on TLC (Rf=0.38
and 0.44) when eluting with ethylacetate: hexanes:
triethylamine=3:7:1. The two spots were nearly
equal in intensity and size (data not shown),
indicating that both diastereomers exist in nearly
equal amount. The broadband decoupled 31P-
NMR spectrum (not shown) has two peaks at
149.67 and 149.49 ppm with a 1:1 intensity ratio for
the single phosphorous atom in the molecule.
3. Results and discussions
3.1. Synthesis of 1%-(7-azaindolyl)-2%-deoxy-D-
riboside and its con6ersion into phosphoramidite
The 4-step synthesis and the respective reaction
yield for each step are shown in Fig. 1. The
glycosylation of 7aIn with 1-(a)-chloro-3,5-di-O-(p-
toluoyl)-2-deoxy-D-ribose was accomplished at −
10 °C with an admirable yield (66%). Prior to this
procedure, several attempts at room temperature
(20 °C) yielded multiple by-products, although
most chemists were successful with the glycosyla-
tion of other bases at room temperature [11,20,40].
The by-products were not characterized. Lowering
the reaction temperature by 30 °C effectively elim-
inated the unwanted reactions to yield 1. 1H-NMR
spectrum of purified 1 indicated that the b-isomer
was the only product, there was no sign indicating
the presence of the a-isomer. The glycosylation is
at the N−1, not the N−7, as shown by Seela and
Gumbiowski [41]. Hydrolysis of 1 in methanol in
the presence of a catalytic amount of sodium
methoxide was completed in 3 h at room tempera-
ture. Compound 2 crystallized readily during the
reaction and was easily isolated and purified. Trity-
3.2. Incorporation of 7aIn into DNA
oligonucleotide
In order to test whether the phosphoramidite 4
is suitable to be used for automated DNA synthe-
sis, the following oligonucleotides containing 7aIn
were prepared: Oligo-1: 5%-GCGCGC7aIn-
CGCGCGC-3%; Oligo-2: 5%-CGC7aInGAATTC-3%;
Oligo-3: 5%-CGG7aInAATTC-3%. Although 7aIn was
put in different sequences, the coupling proceeded
as planned. The coupling efficiency for 7aIn was
about the same as that for the natural bases A, T,
G, C as revealed by monitoring the trityl release at
each coupling step (data not shown). After HPLC
purification, the overall isolated yield for all
oligonucleotides ranged from 6 to 22% including 10
other DNA oligonucleotides containing 7aIn pre-
pared in a separate synthesis (not shown). This
yield is comparable with the synthesis of DNA
oligonucleotides comprised of only natural bases.
lation of
2
with 1.5-equivalent of 4,4%-
dimethoxytrityl chloride partially protected both
the 5%- and 3%-hydroxyl groups of the sugar as
revealed by 1H-NMR spectrum of the isolated main
product (not shown), although such procedure
performed on other modified bases protected only
the 5%-hydroxyl group [10,15,16,20,22]. Therefore,
only 1.1-equivalent of 4,4%-dimethoxytrityl chloride
was used for the reaction and a 75% yield for 3 was
achieved after chromatography purification. The
reaction formula for the synthesis of phospho-
ramidite 4 from the literature also had to be
modified. The optimized mole ratio was 1:4:10 for
3.3. Fluorescence emission spectra for 7aIn, 7aIn
2-deoxyriboside (7aIn-r), 7aIn in single stranded
DNA (7aIn-ss), and 7aIn in double stranded DNA
(7aIn-ds)
The fluorescence emission spectra for 7aIn, 7aIn-

2600
K. Wang et al. / Spectrochimica Acta Part A 58 (2002) 2595–2603
Fig. 2. Fluorescence emission spectra (from top to bottom): (1) 7-azaindole 2%-deoxyriboside (7aIn-r) whose intensity is lowered
10-fold to fit to the scale, (2) 7-azaindole (7aIn), (3) 7-azaindole in single stranded DNA (7aIn-ss), and (4) in double stranded DNA
(
7aIn-ds). The excitation wavelength was 300 nm. The emission intensity for each derivative was normalized by dividing its
absorbance value at this wavelength. The solvent was 1×TBE buffer (89 mM TRIS, 89 mM boric acid, 0.1 mM EDTA) at pH 8.2.
r, 7aIn-ss, and 7aIn-ds were recorded and plotted
together after normalization for the absorbance
value at 300 nm (Fig. 2). In order to fit all the
normalized spectra on the same graph, the
fluorescence emission of 7aIn-r was lowered 10-
fold. The glyco-linkage causes the fluorescence
intensity of 7aIn to increase by about 23 times.
This was expected because similar increase was
observed if 7aIn was N−1 methylated [31–38].
However, incorporation of 7aIn into DNA
quenches its fluorescence significantly as the
fluorescence intensity of 7aIn-ss and 7aIn-ds are
significantly lower than that for 7aIn-r. The
quenching is probably due to base stacking in
DNA. This strong quenching effect was also seen
for 2-aminopurine upon incorporation into DNA,
in which the fluorescence quenching is attributed
to both base stacking and hydrogen bonding [3].
Since 7aIn cannot form hydrogen bonds with the
opposite base, the fluorescence quenching should
be due to base stacking. In comparison, the
fluorescence emission of both 7aIn-ss and 7aIn-ds

K. Wang et al. / Spectrochimica Acta Part A 58 (2002) 2595–2603
2601
are only slightly lower than the fluorescence for
7aIn itself (Fig. 2). Also, the fluorescence of 7aIn in
double stranded DNA is weaker than in single
stranded DNA as shown in Fig. 2. Using 0.023 as
the fluorescence quantum yield for 7aIn deter-
mined using indole as a standard, the quantum
yields for 7aIn-r, 7aIn-ss, and 7aIn-ds are 0.53,
0.020, 0.016, respectively. In summary, the
fluorescence of 7aIn-r is 96% quenched upon in-
corporation into single stranded DNA, and fur-
ther quenched when incorporated into double
stranded DNA.
The fluorescence emission maximum for 7aIn in
water is at 286 nm. When it is glycosylated, its
emission maximum shifts slightly to the red at 288
nm. Upon incorporation into DNA, it shifts to
the blue at about 279 nm in both single and
double strand DNA. This blue shift signifies that
Fig. 3. Change of fluorescence emission spectra for 7-azaindole due to duplex melting as temperature increases. The DNA sequence
was d(5%-CGG7aInAATTC-3%)/d(3%-GCCCTTAAG-5%). The excitation wavelength was 300 nm. The concentration of the DNA was
10.7 mM in DNA strands. The spectra and their corresponding temperatures are shown in the graph. Insert: Fluorescence intensity
change at 380 nm versus temperature plot for the duplex.

2602
K. Wang et al. / Spectrochimica Acta Part A 58 (2002) 2595–2603
7aIn is in a less polar environment in DNA due to
base stacking.
[7] O.V. Petrauskene, S. Schmidt, A.S. Karyagina, I.I.
Nikolskaya, E.S. Gromova, D. Cech, Nucleic Acids
Res. 23 (1995) 2192.
[8] K.D. Raney, L.C. Sowers, D.P. Millar, S.J. Benkovic,
Proc. Natl. Aca. Sci. USA 91 (1994) 6644.
3.4. Fluorescence change for the incorporated
[9] K.T. Hughes, P.C.W. Gaines, J.E. Karlinsey, R.
Vinayak, M.I. Simon, EMBO J. 11 (1992) 2695.
[10] F. Seela, Y. Chen, Nucleic Acids Res. 23 (1995) 2499.
[11] F. Seela, Y. Chen, U. Bindig, Z. Kazimierczuk, Helv.
Chim. Acta 77 (1994) 194.
[12] A. Ono, T. Ueda, Nucleic Acids Res. 15 (1987) 3059.
[13] R. Cosstick, X. Li, D.K. Tuli, D.M. Williams, B.A.
Connolly, P.C. Newman, Nucleic Acids Res. 18 (1990)
4771.
[14] T. Grein, S. Lampe, K. Mersmann, H. Rosemeyer, H.
Thomas, F. Seela, Bioorg. Med. Chem. Lett. 4 (1994)
971.
[15] F. Seela, T. Grein, Nucleic Acids Res. 20 (1992) 2297.
[16] F. Seela, S. Lampe, Helv. Chim. Acta 74 (1991) 1790.
[17] S.L. Driscoll, M.E. Hawkins, F.M. Balis, W. Pfeiderer,
W.R. Laws, Biophys. J. 73 (1997) 3277.
[18] M.E. Hawkins, W. Pfeiderer, A. Mazumder, Y.G. Pom-
mier, F.M. Balis, Nucleic Acids Res. 23 (1995) 2872.
[19] M.E. Hawkins, W. Pfeiderer, F.M. Balis, D. Porter,
J.R. Knutzen, Anal. Biochem. 244 (1997) 86.
[20] D. Loakes, D.M. Brown, Nucleic Acids Res. 22 (1994)
4039.
[21] J.P. Erzberger, D. Barsky, O.D. Schrarer, M.E. Colvin,
D.M.I. Wilson, Nucleic Acids Res. 26 (1998) 2771.
[22] B.A. Schweitzer, E.T. Kool, J. Am. Chem. Soc. 117
(1995) 1863.
7aIn during DNA thermomelting
Since the fluorescence intensity for 7aIn is
weaker in double stranded DNA than in single
stranded DNA, it can be utilized to monitor
DNA thermomelting. Fluorescence emission spec-
trum for Oligo-3 is recorded as a function of
temperature and shown in Fig. 3. At 2 °C, the
DNA is mostly double stranded where its fluores-
cence intensity is the lowest. As the temperature
increases to 13 °C, the fluorescence increases
15%, and the increase continues until about 30 °C
where the maximum fluorescence increase is 30%
(Fig. 3 insert). If temperature continues to in-
crease, a slight decrease of fluorescence is ob-
served due to temperature related quenching
processes. Since this fluorescent 7aIn is structurally
similar to a purine base, its fluorescence property
may be utilized as a probe for studying nucleic
acid structure and dynamics.
[23] B.A. Schweitzer, E.T. Kool, J. Org. Chem. 59 (1994)
7238.
[24] S. Moran, R.X.-F. Ren, C.J. Sheils, S.I. Rumney, E.T.
Kool, Nucleic Acids Res. 24 (1996) 2044.
[25] J.C. Morales, E.T. Kool, Nature Struct. Biol. 5 (1998)
950.
[26] K.M. Guckian, T.R. Krugh, E.T. Kool, Nature Struct.
Biol. 5 (1998) 954.
Acknowledgements
The authors wish to thank the National Insti-
tutes of Health for the NIH-MBRS grant cS06
GM08047 and NIH-RCMI 12RR13459.
[27] C.L. Smith, A.C. Simmonds, I.R. Felix, A.L. Hamilton,
S. Kumar, S. Nampolli, P. Loakes, F. Hill, D.M.
Brown, Nucleosides Nucleotides 17 (1998) 541.
[28] C.W. Flexner, J.E. Hildreth, R.W. Kuncl, D.B. Drach-
man, The Lancet 339 (1992) 438.
[29] C.H. Jurgensen, B.E. Huber, T.P. Zimmerman, G. Wol-
berg, J. Immunol. 144 (1990) 653.
[30] P.R. Andreassen, R.L. Margolis, Proc. Natl. Acad. Sci.
USA 89 (1992) 2272.
References
[1] L.W. McLaughlin, T. Leong, F. Benseler, N. Piel, Nu-
cleic Acids Res. 16 (1988) 5631.
[2] R. Eritja, B.E. Kaplan, D. Mhaskar, L.C. Sowers, J.
Petruska, M.F. Goodman, Nucleic Acids Res. 14 (1986)
5869.
[31] A.V. Smirnov, D.S. English, R.L. Rich, J. Lane, L.
Teyton, A.W. Schwabacher, S. Luo, R.W. Thornburg,
J.W. Petrich, J. Phys. Chem. B 101 (1997) 2758.
[32] R.L. Rich, Y. Chen, D. Neven, M. Negrerie, F. Gai,
J.W. Petrich, J. Phys. Chem. 97 (1993) 1781.
[33] D.S. English, R.L. Rich, J.W. Petrich, Photochem. Pho-
tobiol. 67 (1998) 76.
[3] R.A. Hochstrasser, T.E. Carver, L.C. Sowers, D.P. Mil-
lar, Biochemistry 33 (1994) 11971.
[4] L.B. Bloom, M.R. Otto, J.M. Beechem, M.F. Good-
man, Biochemistry 32 (1993) 11247.
[5] L.B. Bloom, M.R. Otto, R. Eritja, L.J. Reha-Krantz,
M.F. Goodman, J.M. Beechem, Biochemistry 33 (1994)
7576.
[6] M.W. Frey, L.C. Sowers, D.P. Millar, S.J. Bencovic,
Biochemistry 34 (1995) 9185.
[34] Y. Chen, R.L. Rich, F. Gai, J.W. Petrich, J. Phys.
Chem. 97 (1993) 1770.

K. Wang et al. / Spectrochimica Acta Part A 58 (2002) 2595–2603
[35] M.S. Gordon, J. Phys. Chem. 100 (1995) 3974. (1994) 735.
2603
[36] P.-T. Chou, C.-Y. Wei, C.-P. Chang, M.-S. Kuo, J. Phys.
Chem. 99 (1995) 11994.
[39] P. Cioni, L. Erijman, G.B. Strambini, Biochem. Biophys.
Res. Commun. 248 (1998) 347.
[37] M. Negrerie, F. Gai, Bellefeuille, J.W. Petrich, J. Phys.
Chem. 95 (1991) 8663.
[40] Z. Kazimierczuk, H.B. Cottam, G.R. Revankar, R.K.
Robbins, J. Am. Chem. Soc. 106 (1984) 6379.
[38] F. Gai, R.L. Rich, J.W. Petrich, J. Am. Chem. Soc. 116
[41] F. Seela, R. Gumbiowski, Heterocycles 29 (1989) 795.