5328±5337
Nucleic Acids Research, 2002, Vol. 30 No. 23
ã 2002 Oxford University Press
Probing triplex formation by EPR spectroscopy
using a newly synthesized spin label for
oligonucleotides
Peter M. Gannett*, Eva Darian, Jeannine Powell, Edward M. Johnson Il,
Claudius Mundoma1, Nancy L. Greenbaum1, Chris M. Ramsey1, Naresh S. Dalal1 and
David E. Budil2
West Virginia University, Department of Basic Pharmaceutical Sciences, PO Box 9530, Morgantown, WV 26506,
USA, 1Florida State University, Department of Chemistry and Biochemistry, Tallahassee, FL 32306, USA and
2
Northeastern University, Department of Chemistry, Boston, MA 02115, USA
Received May 17, 2002; Revised August 21, 2002; Accepted October 1, 2002
ABSTRACT
Spin labels have been extensively used to study the
dynamics of oligonucleotides. Spin labels that
are more rigidly attached to a base in an oligonucleotide experience much larger changes in their
range of motion than those that are loosely tethered.
Thus, their electron paramagnetic resonance spectra show larger changes in response to differences
in the mobility of the oligonucleotides to which they
are attached. An example of this is 5-(2,2,5,5-tetramethyl-3-ethynylpyrrolidine-1-oxyl)-uridine (1). However, the synthesis of this modi®ed DNA base is
quite involved and, here, we report the synthesis of
a new spin-labeled DNA base, 5-(2,2,6,6-tetramethyl4-ethynylpiperidyl-3-ene-1-oxyl)-uridine (2). This spin
label is readily prepared in half the number of steps
required for 1, and yet behaves in a spectroscopically analogous manner to 1 in oligonucleotides.
Finally, it is shown here that both spin labels 1 and
2 can be used to detect the formation of both
double-stranded and triplex DNA.
INTRODUCTION
Spin labels have been used for many years to probe the
conformational mobility and other structural properties of
oligonucleotides (1±3). Their utility lies in the electron
paramagnetic resonance (EPR) signal that can be observed
and its sensitivity to motion. The EPR spectra of unconstrained
spin-labeled oligonucleotides are mainly isotropic. As the
spin-labeled oligonucleotide becomes more constrained the
corresponding EPR spectrum becomes anisotropic. Thus, by
monitoring the EPR spectrum of a spin-labeled oligonucleotide
it is possible to observe the formation of a variety of DNA
structures such as loops and double-stranded DNA (dsDNA)
(4±7). Note that the degree of constraint can be analyzed in
quantitative terms by computer simulation of the spectra and
extraction of the correlation time for the spin probe (7,8).
The structure of the spin-labeled oligonucleotide signi®cantly affects the extent to which the EPR spectrum changes
upon binding. Spin-labeled oligonucleotides that are attached
to DNA or RNA bases by a ¯exible tether (e.g. single bonds)
show relatively small differences between single- and doublestranded DNA states (4,5). Similarly, spin labels attached to
the backbone of DNA, such as by covalent attachment to a
phosphorothioate, also show small but measurable differences
between unbound and bound states (9). Spin labels that are
more rigidly attached to the oligonucleotide, show large
changes in their EPR spectra between the unbound and bound
forms, as demonstrated, for example, by Spaltstein et al.
(10,11). These latter workers synthesized a spin-labeled
thymidine analog phosphoramidite (1) (Fig. 1), and prepared
several oligonucleotides containing this spin label. In turn,
these oligonucleotides were used to examine the EPR spectra
of a variety of DNA structures. These workers obtained a
range of correlation times spanning ~1±20 ns, effectively
spreading out the time scale so that it was easier to discern
different environments or different DNA structures.
The spin label 1 has been shown to be very useful for
detecting and distinguishing between various DNA structures.
However, the synthesis of 1 requires 12 steps and similar, but
more synthetically accessible, probes would facilitate their
use. Therefore, we designed and synthesized a new spin label,
2, and its phosphoramidite, which can be prepared in six steps
from readily available starting materials (12). In addition, we
have measured the EPR spectra of single-, double-, and triplestranded oligonucleotides containing either 1 or 2, and show
that they are quite similar based on their EPR spectra, thermal
denaturation temperatures and circular dichroism (CD) spectra. Finally, we have found that oligonucleotides containing
either spin label can be used to detect triplex DNA (txDNA)
and may aid in determining txDNA rigidity and stability.
*To whom correspondence should be addressed. Tel: +1 304 293 1480; Fax: +1 304 293 2576; Email: [email protected]
Nucleic Acids Research, 2002, Vol. 30 No. 23
MATERIALS AND METHODS
General
Solvents and reagents were obtained from Aldrich
(Milwaukee, WI) and were used without puri®cation unless
otherwise noted. NMR spectra were obtained on a Varian
Gemini 300 broadband spectrometer. Methylene chloride was
dried by distillation from phosphorus pentoxide. Triethylamine (TEA), pyridine and THF were dried by distillation from lithium aluminium hydride. Dimethylformamide
(DMF) was puri®ed by distillation from barium oxide.
Samples for NMR were either dissolved in CDCl3 or
CD3CN and treated with 1.5 equivalents of phenylhydrazine
or dissolved in 1:1 D2O:acetone-d6 and treated with 1.5
equivalents of sodium dithionite to convert the nitroxide to the
hydroxylamine prior to data acquisition (13). Thus, the
reported proton and carbon spectra refer to the hydroxylamine
derived from the corresponding nitroxides (1±7). Unmodi®ed
phosphoramidite DNA bases and CPG resins were obtained
from Glen Research (Sterling, VA). Mass spectra (MS) were
recorded on an Agilent 5973N (low resolution) or a Finnigan
MAT 90 (high resolution).
Phosphoramidite synthesis
5-(2,2,6,6 - Tetramethyl-4 -ethynylpiperidyl -3-ene-1-oxyl)-uridine (7). Spin probe nitroxide 6 (0.41 g, 2.35 mmol) (12) was
added to a solution of 5-iodo-2¢-deoxyuridine (5-IdU) (1 g,
2.8 mmol) in DMF (17 ml). The mixture was placed in the
dry-ice/methanol bath and exposed to three freeze±thaw
cycles, applying a vacuum between freezing and thawing of
the reaction mixture, to deoxygenate. Copper iodide (0.67 g,
3.5 mmol) and tetrakis(triphenyl-phosphine)palladium(0)
(0.42 g, 0.36 mmol) were then added followed by a ®nal
freeze±thaw cycle. Finally, TEA (0.5 ml, 3.6 mmol) was
added to a mixture and the reaction was stirred for 12 h at 25°C
(11). The solvents were removed in vacuo, the residue was
suspended in 20% methanol±CH2Cl2, and ®ltered through a
plug of silica gel. Chromatography (silica gel, 10%
methanol±CH2Cl2) afforded the thymidine analog 7 (0.34 g,
70%) as a yellow gum. IR (CHCl3) cm±1: 3700, 3650, 3400
(b), 2400, 1740, 1700, 1610, 1510, 1250, 1035. 1H NMR
(CDCl3, phenylhydrazine) d (ppm): 1.19 (6H, s), 1.28 (6H, s),
2.16 (2H, m), 2.35 (2H, s), 3.748 (1H, m), 3.803 (1H, m), 3.99
(1H, m), 4.34 (1H, m), 6.212 (1H, t), 6. 256 (1H, s), 8.67 (1H,
s). UV (CHCl3) lmax (log e): 240 (3.86). HRMS calculated for
C20H26N3O6: 404.4437. Found: 404.4440.
5-(2,2,6,6 -Tetramethyl - 4-ethynylpiperidyl -3 -ene-1-oxyl)-5¢(4,4¢-dimethoxyltriphenyl)-uridine (8). The thymidine analog
7 (1.3 g, 3.2 mmol) was added to 4,4¢-dimethoxytriphenylmethyl chloride previously dissolved in pyridine (10 ml). The
reaction was stirred for 2 h under N2 at 25°C, quenched by the
addition of methanol (5 ml), concentrated in vacuo, and
puri®ed by column chromatography (silica gel, 5%
methanol±CH2Cl2) to give the monoprotected nucleoside 8
(1.95 g, 80%) as a yellow solid. IR (CCl4) cm±1: 3695, 3620,
3395 (b), 3020, 3010, 2990, 2402, 1715, 1705, 1610, 1510,
1455, 1249, 1035. 1H NMR (CDCl3, phenylhydrazine) d
(ppm): 1.60 (6H, d), 1.69 (6H, d), 2.66 (2H, s), 2.76 (1H, m),
5329
3.01 (1H, m); 3.719 (1H, d), 3.816 (1H, d), 3.95 (1H, s), 4. 065
(3H, s), 4.075 (3H, s), 4.47 (1H, m), 5.081 (1H, d), 5.781 (1H,
s), 6.511 (1H, s), 9.271 (1H, s). UV (CHCl3) lmax (log e): 257
(s, 4.03), 276 (3.91), 308 (4.00). HRMS calculated for
C41H44N3O8: 706.8023. Found: 706.8100.
5 -(2,2,6,6 -Tetramethyl-4 -ethynylpiperidyl -3-ene -1-oxyl)-5¢(4,4¢-dimethoxyltriphenyl)-uridine phosphoramidite (2). To a
solution of 8 (0.29 g, 0.41 mmol) in 2.5 ml of dry CH2Cl2 were
added TEA (143 ml, 1.026 mmol) and 2-cyanoethyl-diisopropylchloro-phosphoramidite (107 ml, 0.45 mmol). The
reaction was stirred for 1 h at 25°C and formation of the
product was monitored by TLC. A second portion of 2cyanoethyl-diisopropylchloro-phosphoramidite was then
added (39 ml, 0.17 mmol) and stirred for another hour. The
reaction mixture was concentrated in vacuo, THF:benzene
(1:4, 2 ml) added and stirred for 10 min. The precipitate was
removed by ®ltration, the ®ltrate was concentrated in vacuo,
and the residue co-evaporated twice with benzene (3 ml).
Chromatography on silica gel (petroleum-ether:EtOAc:TEA,
50:50:1) afforded phosphoramidite 2 (0.32 g, 86%) as yellow
powder. M.p. 158±160°C. IR (CHCl3) cm±1: 3695, 3610,
3020, 3005, 2985, 2450, 1740, 1580, 1506, 1240, 1190, 1175,
1030, 920. 1H NMR (CD3CN, phenylhydrazine) d (ppm):
1.15/1.17 (6H, s, (CH3)2), 1.33/1.35 (6H, s, (CH3)2), 1.250/
1.251 (12H, s, CH(CH3)2), 2.26 (2H, m, CH2), 2.26 (1H, m,
H-2¢), 2.56 (1H, m, H-2"), 2.63/2.73 (1H, t, J = 6 Hz, CH2CN),
3.56 (1H, m, H-5¢), 3.57 (1H, m, H-5"), 3.67/3.71 (1H, m,
CH(CH3)2), 3.74 (1H, m, POCH), 3.84 (6H, s, OCH3), 3.92
(1H, m, POCH), 4.23 (1H, m, H-4¢), 4.84 (1H, m, H-3¢), 5.72
(1H, bs, CH = C), 6.28 (1H, m, H-1¢), 6.85 (4H, d, J = 8, ArH3,3¢,5,5¢), 7.26 (4H, d, J = 8, ArH-2,2¢,6,6¢), 7.26±7.39 (4H, m,
ArH-2",3",5",6"), 7.42 (1H, dd, J = 2, 8.5 Hz, ArH-4"), 8.4
(1H, bs, NH), 8.78/8.81 (1H, s, H-6). UV (CHCl3) lmax (log e):
240 (3.86), 275 (3.53), 302 (3.51). HRMS calculated for
C50H62N5O9P: 907.0239. Found: 907.0135.
Oligodeoxyribonucleotide synthesis
Large-scale (10±20 mmol) synthesis was conducted on a
modi®ed ABI 430A protein synthesizer. The oligonucleotides
T15, A15, 5spT15 (T75spTT7) and 6spT15 (T76spTT7) were
synthesized using the solid-phase phosphoramidite protocol.
In all cases where the spin-labeled thymidine was a part of the
oligonucleotide, it was the eighth base. The oligonucleotides
were cleaved from the resin by treatment with concentrated
NH4OH (28±30%, 12 ml) at room temperature for 1 h and
®ltered through a 0.2 mm ®lter disk. The cleavage of protecting
groups for the A15 oligonucleotide was accomplished by
heating the ®ltrate at 55°C for 20 h and then dried down on a
SpeedVac. Final puri®cation of oligomers was achieved by
FPLC using a Bio-Rad TSK DEAE-5-PW column. T15 was
puri®ed under isocratic conditions (53% B; buffer A, 10 mM
NaOH, pH 11.8; buffer B, 10 mM NaOH, 1 mM NaCl, pH
11.8; ¯ow rate, 7 ml/min, detecting at l = 260 nm
wavelength). A15 was puri®ed with a gradient (20±26% B
over 45 min), as were 5spT15 and 6spT15 (40±55% B, 90 min).
The oligomers were desalted with reverse-phase Waters
Sep-Pak (C-18) cartridges, conditioned with methanol and
water and eluted with 60% aqueous methanol.
5330
Nucleic Acids Research, 2002, Vol. 30 No. 23
DNA sample preparation
dsDNAs A15:T15, A:5spT15, A15:6spT15 and txDNAs T15-A15:
T15, 5spT15-A15:T15, 6spT15-A15:T15 [Watson±Crick base pair
indicated by a colon (:) and Hoogsteen base-paired strand by a
hyphen (-)] were used in the studies presented here. All DNA
samples were made up in phosphate buffer (10 mM NaH2PO4,
pH 7.4) and sodium chloride (100 mM). Duplex formation was
achieved by mixing equivalent amounts of *T15 (*T15 = T15,
5spT or 6spT) and A [based on optical density (OD)] and, in
15
15
selected cases, MgCl2, heating to 90°C for 30 min and then
slowly cooling to room temperature. txDNAs are prepared by
®rst forming Watson±Crick duplex A15:T15 in the same
manner as described above, then adding the third strand, *T15
(*T15 is either 5spT15 or 6spT15) to the duplex and then stored at
4°C for 24 h.
UV melting pro®les
UV-monitored melting temperature experiments were conducted at 260 nm using a Cary 300 spectrometer on the
duplexes A15:T15, A:5spT15, A15:6spT15, and triplexes T15A15:T15, 5spT15-A15:T15, 6spT15-A15:T15, under the following
conditions: 10 mM NaPO4, 100 mM NaCl pH 7.4, and 10 or
50 mM MgCl2 (~0.5 OD). Spectra were recorded over a range
of 5±90°C at a rate of 0.25°C/min. Tm values were determined
from the maxima of the ®rst derivative of the thermal
denaturation curve. Enthalpies and entropies were calculated
from van't Hoff plots. All calculations were performed using
the vendor supplied software (Win UV Thermal, Ver. 2.00).
Error estimates of 10% for the thermodynamic parameters
were based on: (i) the range of values obtained from three
individual samples for each duplex studied and (ii) reproducibility of the values obtained by computer ®t of the data as
there is some dependence of the computed values on the range
of data selected.
Circular dichroism measurements
The CD spectra were recorded on an AVIV Model 62A CD
spectrometer. Solutions were ~20±30 mM in duplex and
triplex. Unless otherwise stated, solutions were prepared in
10 mM phosphate buffer, pH 7.4, and 100 mM NaCl.
Magnesium ion concentrations of either 10 or 50 mM were
achieved by the addition of aliquots of 1.75 M MgCl2 to the
solution of the duplex or triplex. The ®nal sample volume was
400 ml. Spectra were recorded as function of temperature
every 5°C from 30 to 75°C for dsDNAs and from 5 to 75°C for
txDNAs.
EPR measurements
Continuous wave (CW) EPR spectra were obtained on a
Brucker EMX X-band or Varian E-12 spectrometer. Samples
were dissolved in phosphate buffer and loaded either in glass
capillaries (10 ml) or in a ¯at cell at oligonucleotide
concentrations of ~2 mM in single-stranded (ssDNA),
dsDNA or txDNA. Spectra were acquired at either 0 or
25°C under the conditions noted in the ®gure captions.
EPR simulation
Computer simulation of EPR spectra were performed using
non-linear-least-squares ®tting routine with modi®ed
Levenberg±Marquardt algorithm in a slow-motion regime
Figure 1. Structures of the spin-labeled phosphoramidites 1 (5spT15) and
2 (6spT15).
(8). Average rotational diffusion rate, rotational anisotropy
and Gaussian inhomogeneous line broadening are varied to get
the best ®t. Goodness-of-®t was determined from the
chi-squared value, which represents the sum of weighted
residuals, and the correlation factor between the experimental
data and the calculated spectrum. Visual judgement of the ®t
was also taken into account. The program requires values for
the g-values (gx, gy and gz) and hyper®ne coupling constants
(ax, ay and az for nitrogen). Initial values were taken from
Hustedt et al. (14). Measurements on frozen samples of T15
and A15:*T15 at 95 GHz were made to con®rm these quantities.
As no signi®cant difference was observed, the values used by
Hustedt were used.
RESULTS AND DISCUSSION
Synthesis of phosphoramidites 1 and 2
The synthesis of phosphoramidite 1 (Fig. 1) has been
previously reported and the published procedures were
followed (11). The full synthesis of phosphoramidite 2 has
not been previously reported and is shown in Figure 2 (12).
The synthesis of this compound began with the commercially
available nitroxide 3. This compound was condensed with the
lithium salt of triisopropylsilylacetylene in THF to give
the acetylenic alcohol 4. The alcohol was then eliminated to
the enyne 5, and the triisoproylsilyl group then removed by
treatment with tetrabutylammonium ¯uoride in wet THF to
give 6. Next, 6 was coupled to 5-iodouridine to give the
nucleoside 7. Attachment of the 5¢ and 3¢ groups necessary for
automated DNA synthesis was accomplished under standard
conditions and yielded the phosphoramidite 2.
Oligodeoxynucleotide synthesis and puri®cation
Oligonucleotides were prepared using the phosphoramidite
methodology on CPG and the standard reaction cycle. The
coupling ef®ciencies were 98±99% for both unmodi®ed and
modi®ed oligonucleotides based on the ODs of the isolated
oligonucleotides. However, FPLC of the 5spT15- or 6spT15modi®ed oligonucleotides showed the presence of three
oligonucleotides. The major product was the oligonucleotide
with an intermediate retention time (42.5 min for 5spT15;
42.0 min for 6spT15). These products correspond to the 15mer
Nucleic Acids Research, 2002, Vol. 30 No. 23
5331
Figure 2. Synthetic scheme followed for the preparation of the new spin label phosphoramidite 2. Reaction conditions: (a) TIPS-CºC-Li, THF, ±78°C,
(b) SOCl2, pyridine, 0°C, (c) TBAF, THF, (d) Pd[P(C6H5)3)]4, CuI, TEA, DMF, (e) trityl-chloride, TEA, CH2Cl2, (f) Cl-P(OCH2CH2CN)(N(i-Pr)2), TEA,
CH2Cl2.
oligonucleotide containing the spin label and both gave an
ESR signal. The oligonucleotides isolated with the longer
retention times (43.7 min for 5spT15; 44.7 min for 6spT15) did
not give EPR signals directly, but did following the addition of
hydrogen peroxide. FPLC of these EPR samples showed that
the EPR silent species eluting at 43.7 min for 5spT15 or 44.7 min
for 6spT15 were converted to the EPR active species eluting at
42.5 min for 5spT15 or 42.0 min for 6spT15. Thus, the EPR silent
oligonucleotides with a longer retention time are the
hydroxylamine derivatives of 5spT15 or 6spT15.
The oligonucleotides with shorter retention times (40.4 min
for 5spT15; 40.8 min for 6spT15) were EPR silent and did not
give rise to an EPR signal upon the addition of hydrogen
peroxide. Mass spectral analysis of the latter oligonucleotides
revealed that they had a molecular weight of ~30 less than
expected for the spin-label-bearing 15mer. This suggests that
this oligonucleotide had lost NO which may occur during the
iodine oxidation step. It is known that nitroxides are sensitive
to oxidation by halogens and are converted to nitrones, which
can subsequently decompose with the loss of NO (15). This
point was further investigated by stopping the oligonucleotide
synthesis after the addition of the spin label, T7*T, after the
12th base, T7*TT4, and after the 15th base, T7*TT7 (*T = 5spT
or 6spT, *T always located at the eighth position), cleaving
from the resin, and examining by FPLC and EPR. FPLC of
T7*T showed mainly one oligonucleotide and it was EPR
active. As the number of synthesis cycles continued to 12 and
then 15, increasing amounts of the oligonucleotide which had
lost the NO group appeared, consistent with the hypothesis
that the iodine oxidation step is responsible for decomposing
the nitroxide. Thus, oxidants, other than iodine, may improve
the purity of spin-labeled oligonucleotides prepared by
automated DNA synthesis.
Thermal denaturation studies
Thermal denaturation curves of dsDNA and txDNA without
and with spin-label modi®cation were measured. These
studies were conducted for several reasons. First, thermal
denaturation data for unmodi®ed and spin-labeled oligonucleotides have been previously measured on CGCGAATTCGCG and CGCGAAT5spTCGCG duplexes (11).
Nearly identical Tm values were determined, indicating that
the spin-label modi®cation did not signi®cantly alter the
stability of duplex. Here, we wanted to determine if this was
also the case for the dsDNAs A15:5spT15 and the A15:6spT15 and
the triplexes 5spT15-A15:T15 and the 6spT15-A15:T15. In addition, it was necessary to measure the temperature at which the
spin-labeled duplexes and triplexes melted so that the
temperature at which the EPR studies should be conducted
could be determined.
The dsDNA thermal denaturation curves were obtained on
samples made up in 10 mM phosphate buffer (pH 7.4) and
100 mM sodium chloride to which magnesium chloride was
added. Representative curves are shown in Figure 3A.
Thermal denaturation and thermodynamic data for all magnesium chloride concentrations are shown in Table 1. There
5332
Nucleic Acids Research, 2002, Vol. 30 No. 23
Figure 3. Thermal denaturation curves for (A) dsDNAs A15:T15 (solid line),
A15:5spT15 (dashed line) and A15:6spT15 (dotted line) and (B) the txDNAs
5spT -A :T
6spT -A :T
15
15 15 (dashed line) and
15
15 15 (dotted line). All samples
were dissolved in 10 mM sodium phosphate buffer pH 7.4, 100 mM NaCl
and 50 mM MgCl2.
are no signi®cant differences in the Tm values measured for the
unmodi®ed and modi®ed dsDNAs though the shape of the
pro®les suggests some differences in the degree of cooperativity as the dsDNAs melt. Likewise, the thermodynamic
parameters for the unmodi®ed and modi®ed oligonucleotides
are all nearly identical. Though not statistically signi®cant, the
general trend suggests that the modi®ed oligonucleotides are
slightly more stable. The values reported here for Tm, DG°,
DH° and DS° are comparable to those obtained computationally from the program MELTING (16).
Thermal denaturation curves of the txDNAs (*T15-A15:T15;
*T is the third, triplex strand, A15:T15 is the Watson±Crick
base-paired dsDNA) were also measured. Two Tm values,
identi®ed by determining the maxima of the ®rst derivative of
the thermal denaturation curve, are observed for txDNAs. The
®rst value corresponds to the triplex melting to give a singlestranded oligonucleotide, *T15 (*T15 = T15, 5spT15 or 6spT15)
and a dsDNA (A15:T15) and the second Tm for the melting of
the duplex. The Tm for the triplex melting is not always
observed, in part due to the small difference in absorbance
between the triplex and the duplex in the presence of an
unassociated triplex strand (17). However, at relatively high
concentrations of magnesium chloride, conditions known to
stabilize txDNA (18), two transitions can be observed for all
three triplexes as shown in Figure 3B. In Table 2 are reported
the Tm values for the unmodi®ed and modi®ed txDNAs. As
was observed for the dsDNAs, they do not signi®cantly differ
from one another under a given set of conditions.
Thermodynamic parameters could not be extracted from the
data as there was not a suf®cient break between the triplex and
duplex portions of the melting curves.
Within experimental error, there is no signi®cant difference
in the thermal denaturation temperatures determined for the all
three duplexes or triplexes. This indicates that the replacement
of T with either 5spT or 6spT has little, if any, effect on the
overall stability of either the duplexes or triplexes of which
they are a part. This is further supported by the CD and EPR
studies (see below). In addition, the thermal denaturation data
allow the determination of which species (triplex, duplex or
single-stranded) are present at a given temperature aiding the
interpretation of both the CD and EPR data.
Circular dichroism
CD spectroscopy has been shown to be a useful tool to
distinguish between homopolymer A:T duplexes and T-A:T
triplexes. We have used CD spectra here for three main
purposes. First, we used CD to show that the A15:T15 duplex
and T15-A15:T15 triplex are present by comparison with
previously published data. Secondly, for the spin labels to be
useful for biological studies, it is important to demonstrate that
Table 1. UV melting temperatures for the unmodi®ed and modi®ed
5spT
and
6spT
duplexesa
Duplex
Tm (°C)
DG° (kcal/mol)b
DH° (kcal/mol)
DS° (cal/mol´K)
A15T15
24
35
45
25
36
46
24
36
45
±8.7
±13.5
±18.8
±8.7
±14.8
±19.1
±8.6
±14.8
±18.9
±100
±116
±126
±102
±118
±134
±100
±117
±127
±312
±345
±365
±319
±349
±393
±312
±353
±369
A15:5spT15c
A15:6spT15d
aDNA melting temperatures were measured in solutions of ~0.5 OD oligonucleotide in sodium phosphate
buffer (10 mM, pH 7.4) and sodium chloride (100 mM). Tm values were determined from the maxima of the
®rst derivative of the thermal denaturation curve.
bFree energies calculated at 293 K. Errors in the thermodynamic data are estimated to be ~10% (see Materials
and Methods).
cSamples also contained 10 mM MgCl .
2
dSamples also contained 50 mM MgCl .
2
Nucleic Acids Research, 2002, Vol. 30 No. 23
Table 2. UV melting temperatures for the unmodi®ed and modi®ed
and 6spT triplexesa
5333
5spT
txDNA
Tm (°C) 10 mM Mg2+
Tm (°C) 50 mM Mg2+
T15-A15:T15
5spT -A :T
15
15 15
6spT -A :T
15
15 15
20
19
19
26
26
26
aOnly
the melting temperature (Tm) of the triplex is reported. DNA melting
temperatures were measured on solutions of ~0.5 OD oligonucleotide
in sodium phosphate buffer (10 mM, pH 7.4) and sodium chloride
(100 mM).
they do not produce any signi®cant conformational changes
and thus give rise to CD spectra comparable with unmodi®ed
oligonucleotides. Finally, CD data can be used to detect the
presence of the triplex form, the duplex form, and the
temperature-dependent conversion of the former to the latter.
Thus, the methodology can be used as an alternative to, or
supplement to, the thermal denaturation data.
In Figure 4A are shown the CD curves for A15:T15,
A15:5spT15 and A15:6spT15. Overall, the curves are quite similar
to one another and show positive ellipticity at 218 and 282 nm
and negative ellipticity at 248 and 210 nm (19,20). There are
some noticeable differences between the unmodi®ed and spinlabeled duplexes. In particular, the spin-labeled duplexes
produce larger positive ellipticities at 218 and 282 nm and
more negative ellipticities at 210 and 248 nm than do the
unmodi®ed duplexes. While the source of these differences is
unknown, they may be due to differences in the lightabsorption properties of the spin label, to changes in secondary
structure, or a combination of both (21). Molecular dynamics
calculations on the unmodi®ed and modi®ed dsDNAs have
shown that both adopt a B-DNA-like structure. However, the
unmodi®ed duplex more closely resembles canonical B-DNA
while the modi®ed dsDNAs are beginning to resemble
A-DNA (22). The differences observed in the CD spectra
between the unmodi®ed and modi®ed DNAs support this as
A-DNAs show more positive ellipticities at ~280 nm and more
negative ellipticities at ~210 nm (21). Thus, the shape of the
CD curves and the observed minima and maxima indicate that
there are some secondary structural differences, though
probably small, and both unmodi®ed and modi®ed dsDNAs
adopt a B-DNA-like conformation.
As noted, CD spectra can be used to distinguish between
txDNA and dsDNA for homopolymers of T-A:T and A:T,
respectively (23). Here, we have used CD for this purpose and
have measured the CD spectra of samples of *T15-A15:T15
(*T15 = T15, 5spT15 and 6spT15). Figure 4B shows a series of CD
spectra recorded on a sample containing 6spT15-A15:T15 over
the temperature range of 5± 40°C. Among the key features to
note are that at 5°C, there are maxima at 230, 260 and 284 nm,
minima at 210, 248 and 267 nm, and an isoelliptic point at
263 nm. These spectral features are very similar to 5spT15A15:T15 (data not shown) and to those previously reported CD
spectra of T-A:T triplexes and strongly support the presence of
a txDNA in our samples.
As the sample is warmed, the maxima at 230 and 284 nm
increase while the maxima at 260 nm gradually diminishes in
amplitude. The maxima at 230 and 284 nm shift toward
shorter wavelengths although the shift for the 230 nm peak is
Figure 4. CD spectra of (A) A15:*T15, where *T15 is T15 (circles), 5spT15
(squares), and 6spT15 (triangles) at 25°C, (B) 6spT15-A15:T15 triplex recorded
from 5°C (triangles) to 40°C (squares), and (C) 6spT15-A15:T15 triplex
sample in (B) from 40°C (squares) to 75°C (circles). All samples were
dissolved in 10 mM phosphate buffer pH 7.4, 100 mM NaCl and 50 mM
MgCl2.
much larger than that for 284 nm. The minimum at 248 nm is
seen to decrease in amplitude but its position remains constant.
In total, these observations are completely analogous to those
reported for T12-A12:T12 (19,20). Moreover, the unmodi®ed
T15-A15:T15 and 5spT15-A15:T15 triplexes give CD spectra
(data not shown) that were nearly identical to those presented
in Figure 4B.
Upon raising the temperature from 40 to 75°C a new set of
spectral changes occur (Fig. 4C). First, several of the
characteristics of the spectrum at 40°C, where the dsDNA
form should be the main species present, are quite different
5334
Nucleic Acids Research, 2002, Vol. 30 No. 23
The spectra shown in Figure 4B and C are very similar to
the CD spectra for T15-A15:T15 and 5spT15-A15:T15 (data not
shown). The data suggest that neither the 5spT nor the 6spT spin
labels signi®cantly affect the structure or conformation of the
triplex of which they are a part. Likewise, the temperature
dependence of all three sets of spectra is very similar and
supports the position that the spin-label modi®ed thymidine is
nearly identical to a thymidine that it replaces.
EPR studies
Figure 5. EPR spectra of (A) 5spT15 at 0°C, (B) 5spT15 at 25°C, (C) 6spT15 at
0°C and (D) 6spT15 at 25°C. All spectra were recorded on samples that were
dissolved in 10 mM phosphate buffer pH 7.4, 100 mM NaCl and 50 mM
MgCl2. Spectrometer settings: receiver gain 6.32 3 103, sweep width 100 G,
modulation amplitude 1.00 G, modulation frequency 100 kHz, microwave
power 20 mW, time constant 5.12 ms. The solid line is the experimental
spectrum and the dashed line is the simulated spectrum.
from those observed at 5°C. The short wavelength maximum
is signi®cantly blue shifted (230 ® 218 nm) and the maximum
at 260 nm is no longer present. However, the maximum at
282 nm and the minimum at 248 nm are still present though
both have diminished in intensity. Raising the temperature
produces smaller changes in the two maxima, relative to those
seen for the triplexes. In contrast, the relative change seen for
the minima at 248 nm is larger on heating the duplex than that
observed for the triplex.
Table 3. Correlation times for ssDNA, dsDNA and txDNA containing
6spT
One species
0°C
t^
t||
N%b
25°C
3.03
4.056
2.111
100
1.07
6spT:A
6spT
The EPR spectrum of 5spT15, at 0°C, is shown in Figure 5A.
This spectrum shows slight anisotropic broadening in the high
®eld line as has been previously observed (11) for the 12mer
analog with the spin probe 1, located on the sixth base. Here,
the spectra for 5spT15 were ®tted using the program NLSL and
the computer ®t is also shown (dashed line) (8). The ®tting
procedure provides the average correlation time (tc), a
measure of mobility for the spin label, with shorter times
indicating greater mobility. These times are reported for both
0 and 25°C temperatures.
As spectra become more anisotropic, meaning that the
motion of the probe slows, it is more appropriate to calculate
the two components of correlation time, t^ and t||, since they
are related to the rotational rate constants R^ and R||. As the
motion slows, the spectrum becomes more sensitive to R|| and
less sensitive to R^. The correlation time obtained for the
ssDNA was ~3 ns, slightly longer than the correlation time
reported for the 12mer (1 ns). Warming the sample to room
temperature gives the spectrum shown in Figure 5B. The
spectrum is similar to that obtained at 0°C (Fig. 5A) though
spin labela
One species
Two species
Fast
Slow
167
38.57
74.00
100
10.7
10.5
62.89
8.615
47
0.33
157
109.5
245.2
53
25.7
6spT-A:T
One species
170
53.40
151.6
100
0.95
Two species
Fast
Slow
4.83
30.78
4.006
5
0.60
307
38.73
3570
95
18.3
aCorrelation times are expressed in nanoseconds, and they were determined by simulation of the experimental EPR data using the program NLSL, unless
otherwise noted.
bN% refers to the percentages of each species present.
Table 4. Correlation times for ssDNA, dsDNA and txDNA containing
5spT
One species
0°C
t^
t||
N%b
25°C
2.97
4.430
1.791
100
1.15
5spT:A
5spT
spin labela
One species
Two species
Fast
Slow
163
87.55
156.4
100
9.33
11.6
4.849
18.18
45
0.43
88.3
77.54
196.4
55
14.3
5spT-A:T
One species
163
24.65
548.3
100
1.03
Two species
Fast
Slow
3.25
3.534
0.724
7
0.68
303
32.32
3265
93
26.7
aCorrelation times are expressed in nanoseconds, and, unless otherwise noted, they were determined by simulation of the experimental EPR data using the
program NLSL.
bN% refers to the percentages of each species present.
Nucleic Acids Research, 2002, Vol. 30 No. 23
Figure 6. EPR spectra of (A) A15:5spT15 at 0°C, (B) A15:5spT15 at 25°C,
(C) A15:6spT15 0°C and (D) A15:6spT15 25°C. All spectra were recorded on
samples that were dissolved in 10 mM phosphate buffer (pH 7.4), 100 mM
NaCl and 50 mM MgCl2. Spectrometer settings were the same as those
used for the data in Figure 5. The solid line is the experimental spectrum
and the dashed line is the simulated spectrum.
the up®eld line has sharpened and the correlation time has
decreased to ~1 ns.
The EPR spectrum of the 6spT15 spin-labeled oligonucleotide is also shown (Fig. 5C). This spectrum is nearly
indistinguishable from that of 5spT15 and the correlation time
obtained by computer simulation of the spectrum con®rms this
(Tables 3 and 4). As was observed for the 5spT15 spin probe,
warming the sample causes the high ®eld line to sharpen.
Fitting the spectrum shows that the correlation time has
decreased by a factor of 3 to ~1 ns (Tables 3 and 4), as
observed for the 5spT15 spin probe.
Duplexes with A15 and 5spT15 or 6spT15 were prepared
(10 mM phosphate buffer pH 7.4, 100 mM NaCl) and the EPR
spectra of these samples recorded at 0 and 25°C (Fig. 6). The
spectra were simulated to calculate tc for both spin labels and
at both temperatures. It has been shown that duplex formation
signi®cantly reduces the mobility of attached spin labels and
the spectra become very anisotropic as can be seen by
comparison of the EPR spectra in Figures 5 and 6. The spectra
in Figure 6 were each ®t assuming either (i) that only the
duplex was present or (ii) that two species were present, the
duplex and, for example, some dissociated *T15. The former
assumption gives correlation times that are nearly identical for
the duplexes containing 5spT or 6spT, 163 and 167 ns,
respectively (Tables 3 and 4).
The two species ®t results in two correlation times for each
spectrum (5spT15 or 6spT15), one corresponding to a conformationally more mobile species (fast) and one that is more
constrained (slow, duplex). While the two species assumption
gave statistically improved ®ts for each spectrum, this is likely
an artifact of the process. There are several reasons for this
statement. First, the most likely candidate for the fast species
is the single-stranded oligonucleotide bearing the spin probe.
If so, then the computed correlation times for the fast species
should be the same as for the ssDNAs in the absence of A15.
This is not the case as the fast species have correlation times
approximately three times larger (Tables 3 and 4). The results
at room temperature are similar to those obtained at 0°C.
5335
Figure 7. EPR spectra of (A) 5spT15-A15:T15 at 0°C, (B) 5spT15-A15:T15 at
25°C, (C) 6spT15-A15:T15 at 0°C and (D) 6spT15-A15:T15 at 25°C. All spectra
were recorded on samples that were dissolved in 10 mM sodium phosphate
buffer (pH 7.4), 100 mM NaCl and 50 mM MgCl2. Spectrometer settings
were the same as those used for the data in Figure 5. The solid line is the
experimental spectrum and the dashed line is the simulated spectrum.
Finally, the sub-spectra that correspond to the fast species do
not resemble the single-stranded oligonucleotide (data not
shown).
Note, that while the correlation times for the single-stranded
oligonucleotides are quite similar to those reported for 5spT12,
those found here for the duplexes are larger, by a factor of 10,
than those reported for 5spT12A12. The discrepancy may be due
to the different methods used to ®t the data or to differences in
the conditions used. The correlation times for 5spT12:A12 were
obtained by treating the duplex as a cylinder to which the
probe was attached and analyzed using hydrodynamic theory.
Here we have used the MOMD method for simulation (24).
Which method is more appropriate for these systems is
debatable. However, the basic conclusions are not affected.
Spin labels have been used to study a variety of DNA
structures including dsDNA, loops (both stem and helical
regions), and base-pair mis-matches and bulges (4±7). To our
knowledge, they have not been used to detect txDNA
formation. Here, we show that spin labels can also be used
for this purpose. The duplex between A15 and T15 was
prepared (A15:T15) and to this duplex was added either 5spT15
or 6spT15. After annealing overnight at 4°C, under conditions
known to produce txDNA (5spT15-A15:T15 or 6spT15-A15:T15),
the EPR were recorded at 0°C and at room temperature. The
resulting EPR and spectral simulations are shown in Figure 7.
It is obvious that the spectra obtained at 0°C are anisotropic
and the mobility of the strand has been constrained. This, in
turn, implies that the third strand has become bound to its
target and thus the EPR spectra shown in Figure 7 are those of
txDNA. In addition, the thermal denaturation and CD data
support this claim.
The triplex EPR data were ®t as described for the duplexes,
®tting the data assuming either one or two species to be
present. As in the case of the duplexes, the ®ts were improved
for the two species ®ts and the calculated correlation times are
shown in Tables 3 and 4. However, unlike the duplexes,
the correlation times obtained for the fast species agree with
5336
Nucleic Acids Research, 2002, Vol. 30 No. 23
those obtained for the single-stranded oligonucleotides. This
suggests that, in this case, the two-species ®t is a better choice.
Moreover, the calculated sub-spectra for 6spT15-A15:T15 (0°C)
con®rm this and the calculated spectrum for the fast species is
essentially the same as the spectrum that is calculated for
6spT
15 at 0°C (Fig. 5C).
The triplex simulations that were based on two sites give
correlation times that are twice those for the duplexes (single
species ®t). This increase in correlation time suggests that
txDNA may be more rigid than dsDNA. Alternatively, the
difference may simply be due to the increased size of the
triplex relative to the duplex. It is dif®cult to distinguish
between these two possibilities and to do so will require
additional studies to determine which factor is responsible for
the difference. In addition, molecular modeling and molecular
dynamics calculations may further help to resolve this
question.
Finally, the temperature dependence of the triplex EPR data
addresses whether the spectra in Figure 7 are txDNA with the
spin-labeled strand as the Hoogsteen or the Watson±Crick
base-paired strand. There is some ambiguity in this regard as
the spin-labeled strand, with the exception of the spin label, is
identical with the T15 Watson±Crick base-paired strand.
Therefore, since the 5spT15 or the 6spT15 strand could have
exchanged with the T15 strand, the EPR spectra shown in
Figure 7 might correspond to T15-A15:5spT15 or T15-A15:6spT15
instead of 5spT15-A15:T15 or 6spT15-A15:T15. This seems
unlikely as samples were maintained at 4°C or below, except
to obtain the room temperature spectrum, conditions where the
duplex (A15:T15) is stable (Tm ~ 45°C). More important are the
EPR spectra. As the txDNA samples are warmed to room
temperature (Fig. 7B and D), the spectra change to the
spectrum of mainly 5spT15 or 6spT15. If strand exchange had
occurred then warming the txDNA samples to room temperature would have lead to the same spectra as obtained for
dsDNAs A15:5spT15 or A15:6spT15. Comparison of the spectra
shown in Figure 6B and D with those in Figure 7B and D
shows that this is clearly not the case.
CONCLUSIONS
The synthesis of a new spin label nitroxide in a six-membered
ring, which is rigidly attached to a uridine, has been achieved
and its preparation described. This new spin-labeled DNA
base, 2, is synthetically more simple to prepare than a
previously described, analogous spin label, 1, and, at the same
time, provides comparable yields during automated DNA
synthesis and similar EPR data. The effect of the spin label on
the thermal denaturation temperature for dsDNA and txDNA
is minimal and unmodi®ed and modi®ed DNAs have similar
Tm and thermodynamic properties suggesting that the spin
labels do not signi®cantly alter the stability of dsDNAs or
txDNAs that contain them. CD spectra of the unmodi®ed and
modi®ed dsDNAs are similar, though there are some differences that suggest the spin labels may perturb the secondary
structure and render it more A-DNA-like than is the
unmodi®ed dsDNA. The CD spectra also show that the
modi®ed DNAs can form txDNA. The EPR spectra show that
the new spin label can be used to monitor dsDNA formation
and, in addition, both spin labels can be used to detect txDNA
as well. The EPR data suggest that the spin label in txDNA is
slightly less mobile than dsDNA though this may be due to the
increase in size of the txDNA relative to the dsDNA.
ACKNOWLEDGEMENTS
We thank the Florida State University BASS laboratory for
their assistance in the preparation and puri®cation of the
oligonucleotides used in the course of this work. C.M. is a
UNCF-P®zer Biomedical Research Fellow. We thank the NIH
(GM 57630) and NSF-EPSCoR (9871948) for their ®nancial
support of this work.
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