1722-1728
Nucleic Acids Research, 1995, Vol. 23, No. 10
1995 Oxford University Press
Spectroscopic studies of chimeric DNA-RNA and
RNA 29-base intramolecular triple helices
J. Liquier, E. Taillandier, R. KM nek1, E. Guittet1, C. Gouyette2 and T. Huynh-Dinh2*
Laboratoire CSSB, URA CNRS 1430, UFR de Medecine, University Paris Nord, 74 rue Marcel Cachin, 93012
Bobigny Cedex, France, 1lnstitut de Chimie des Substances Naturelles, CNRS, 98198 Gif sur Yvette, France
and 2 Unit6 de Chimie Organique, URA CNRS 487, Insitut Pasteur, 28 rue du Docteur Roux, 75724 Paris Cedex
15, France
Received February 8, 1995; Revised and Accepted April 6, 1995
ABSTRACT
Fourier transform infrared (FTIR), UV absorption and
exchangeable proton NMR spectroscopies have been
used to study the formation and stability of two
intramolecular pH-dependent triple helices composed
by a chimeric 29mer DNA-RNA (DNA double strand
and RNA third strand) or by the analogous 29mer RNA.
In both cases decrease of pH induces formation of a
triple helical structure containing either rUNJA.dT and
rC+*dG.dC or KTrA.rU and rC+*rG.rC triplets. FTIR
spectroscopy shows that exclusively N-type sugars
are present in the triple helix formed by the 29mer RNA
while both N- and S-type sugars are detected in the
case of the chimeric 29mer DNA-RNA triple helix.
Triple helix formation with the third strand RNA and the
duplex as DNA appears to be associated with the
conversion of the duplex part from a B-form secondary
structure to one which contains partly A-form sugars.
Thermal denaturatlon experiments followed by UV
spectroscopy show that a major stabilization occurs
upon formation of the triple helices. Monophasic
melting curves indicate a simultaneous disruption of
the Hoogsteen and Watson-Crick hydrogen bonds in
the intramolecular triplexes when the temperature is
increased. This is in agreement with imino proton NMR
spectra recorded as a function of temperature. Comparison with experiments concerning intermolecular
triplexes of Identical base and sugar composition
shows the Important role played by the two tetrameric
loops In the stabilization of the intramolecular triple
helices studied.
INTRODUCTION
Triple-stranded nucleic acid structures have been recently
extensively studied as potentially involved in several fundamental biological functions, such as transcriptional regulation
and recombination (1-3). Triple helices have been used as
powerful biological tools, for example, as artificial nucleases
(4,6) or as highly sequence-specific repressors to modulate
* To whom correspondence should be addressed
protein recognition of DNA (7,8). Physico-chemical studies of
triple helices such as X-ray fiber diffraction (9,10), NMR
(11-14), vibrational spectroscopy (15-21) and UV absorption
spectrophotometry (13,22-25) have provided a considerable
amount of structural data. Triple helices can be formed by binding
of a DNA or RNA oligomer to a double-stranded nucleic acid.
Another possibility to obtain a triple helix is a sequence which,
under well controlled circumstances, is able to fold allowing the
formation of base triplets (14,26). As different duplexes other
than DNA, such as double helical RNA, DNA-RNA hybrids,
RNA hairpins are also involved in biological processes, an
important question is, what is the influence of the sugar backbone
geometry in the formation of the triple helices, how can a third
strand fit into the major groove of a duplex and consequently will
there be any structural modifications in the targeted duplex
structure? Moreover, what are the relative stabilities of the
possible triple helices, in particular when both deoxy and
ribonucleotide sequences are involved? These questions have
been recently investigated by several authors in the case of
intermolecular triplexes (25,27-30). In the present work we have
considered two analogous oligomers which should be able to
form intramolecular triple helices containing pyr*pur.pyr base
triplet motifs where the third strand is parallel with respect to the
purine strand. In such a configuration the third pyrimidine strand
is known to Hoogsteen base pair with the purine strand (31-33).
In the notation pyr*pur.pyr, the (.) corresponds to Watson-Crick
base pairing of the initial duplex and the (*) corresponds to third
strand Hoogsteen base pairing. The sequences studied are:
a 29mer RNA r(GAGAGAA-CCCC-UUCUCUC-UUUUCUCUCUU) and the analogous chimeric 29mer DNA-RNA
d(GAGAGAA-CCCC-TTCTCrC-TTTT)-r(CUCUCUU). They
were chosen based on a 28mer DNA oligonucleotide studied by
Sklenar and Feigon, with one additional loop nucleotide (34).
They were designed to allow intramolecular folding at acidic pH
forming triple helices containing rO^rG.rC and rU*rA.rU base
triplets and r(CCCC) and r(UUUU) loops in the first case, and
rC+*dG.dC and rU*dA.dT base triplets and d(CCCQ and
d(lTlT) loops in the second (Fig. 1). The formation of the triple
helices has been followed by FTIR spectroscopy which also
provides structural data concerning the sugar conformations. The
Nucleic Acids Research, 1995, Vol. 23, No. 10 1723
with analytical HPLC, capillary electrophoresis and PAGE of 32 P
5' end-labeled sample. The oligonucleotide was completely
degraded by snake venom phosphodiesterase.
The mixed DNA-RNA sample was also prepared at the 10
u.mole scale with a double coupling procedure for each ribonucleoside and a single coupling for the deoxynucleoside (99.5%
average trityl yield). After desalting and HPLC purification, the
hybrid oligonucleotide Na+ form was obtained with a final yield
of 10% (9.4 mg).
Upi
5
i
Solution preparation and UV spectroscopy
T-C-T-C-T-C-T-T
'i9
18
17
18
15
14
'«
12*
p
—
_
Figure 1. Schematic representation of the studied triple helices. Top: 29mer
RNA: U*A.U and C^G.C base triplets, rC4 and rU4 loops. Bottom: 29mer
DNA-RNA: U'A.T and C+*G.C base triplets, dC4 and dT4 loops. Shaded
bases: ribonucleotides.
stability of the triplexes has been studied by UV absorption
spectrophotometry (thermal denaturation) and NMR spectroscopy. The results were compared in the case of the chimeric
DNA-RNA sequence with those obtained for the 18mer DNA
part of the molecule d(GAGAGAACCCCTTCTCTC).
MATERIALS AND METHODS
Oligonucleotide synthesis
Two analogous 29mers were synthesized: the first containing only
riboses itGAGAGAA-CCCC-UUCUCUC-UUUU-CUOJCUU),
the second formed by a 22mer DNA followed by a 7mer RNA:
d(GAGAGAA-CCCC-TTCTCTC-TTTT)-r(CUCUCUU). The
two 18mers (5' ends of the 29mers), r(GAGAGAA-CCCCUUCUCUC) and d(GAGAGAA-CCCC-TTCTCTC) as weU as
the 7mer (3' end of the 29mers) r(CUCUCUU) were also prepared.
These oligomers will be referred to in the text as 29mer RNA,
29mer DNA-RNA, 18mer DNA stem, 18mer RNA stem and 7mer
RNA.
The short oligonucleotides (7-18mer) in the DNA or RNA
series were synthesized at the 1 fj.mole scale with standard
procedures on an ABI 380 B.
The 29mer RNA was synthesized with a 10 (imole column,
with Milligen/Millipore phosphoramidites and a double coupling
procedure for each base addition (average coupling yield 99.5%
by trityl titration). Deprotection was carried out using NH3
saturated ethanol at 55°C for 24 h followed by 24 h at room
temperature. The 2'-TBDMS protecting group was removed with
TBAF/THF (1.1 M, 50 molar excess) for 40 h at room
temperature. After neutralization with a solution of 1 M TEAA,
the crude product was desalted twice on a Sephadex G-10
column, eluting with 0.1 M then 0.05 M TEAA. The eluted
compound was lyophilized and then purified using preparative
HPLC (Nucleosil 300 A 5-C18 1/2" x 25 cm; flow rate 5.5
ml/min of 0.01 M TEAA pH 7, with a 5-25% CH3CN gradient
in 20 min). The purified oligonucleotide was exchanged with a
Dowex 50 W-X8 Na+ column, then lyophilized to yield 14.6 mg
(15.5% total yield of purified product). Its purity was checked
All oligomers were dissolved in 20 mM NaCl. Samples were
heated at 90° C for 5 min then rapidly cooled and stored overnight
at 4°C. The pH of the solutions was then adjusted by addition of
small amounts of HO or NaOH and measured before and after the
denaturation experiments using a MI4152 microcombination
probe from Microelectrodes Inc.
Sample concentrations were determined spectrophotometrically using computed sequence-dependent extinction coefficients
given by a nearest neighbor model involving average nucleotide
and dinucleotide extinction coefficients (35): 29mer RNA 10
100/M/cm, 29mer DNA-RNA 88OO/M/cm, 18mer DNA stem
9200/M/cm. Oligomer concentration was usually 1.5 |iM strand
except for the experiments carried out with the 29mer RNA for
which the concentration was 3 (iM strand. Absorption spectra and
absorbance versus temperature profiles (measured at the maximum of the absorption band -262 nm) were recorded with a
Kontron Uvikon 941 spectrophotometer. The temperature of the
cell holder (sample and reference) was varied by circulating
liquid using a Huber water bath controlled by a Huber PD415
temperature programmer. The measurements were initiated ~8°C
and carried up to 85°C. The temperature increase rate was
0.1°C/min. Measurement of the temperature was performed
directly in the reference cell.
NMR spectroscopy
Imino proton spectra as a function of temperature were recorded
on a Bruker AMX 600 spectrometer using a 125 }is S-excitation
pulse (36) centered on the water resonance. The samples 29mer
RNA and 29mer DNA-RNA were 1.2 and 0.8 mM oligomer,
respectively, 20 mM NaCl, pH 4.8 in 95% H2O/5% D2O. Data
were processed using the GIFA software, developed in our
laboratory.
Infrared spectroscopy
For FTIR spectroscopy, samples were deposited between ZnSe
windows without spacer. The sample volume was 2 u.1 and the
usual concentration 3 mM strand. The pH was measured directly
in the sample using a MI4152 microelectrode. Deuterated
samples were obtained by drying the sample in H2O solution
under nitrogen and redissolving in the same volume of D2O. The
pH values given in the text for D2O solutions are not corrected and
correspond to the pH measured in H2O before hydrogendeuterium exchange.
FTIR spectra were recorded with a Perkin Elmer 1760
spectrophotometer coupled to a P.E.7700 minicomputer. Usually
25 scans were accumulated. Data treatment performed with the
Galaxy Spectracalc program included solvent substraction,
baseline correction and spectral normalization using the phos-
1724 Nucleic Acids Research, 1995, Vol. 23, No. 10
h)
8
1700
1700
1700
1600
1500
Figure 2. Founer transform infrared spectra recorded in D2O solutions in the region of the base in-plane double bond stretching vibrations: (a) 29mer RNA pH 7.4;
(b) 29mer RNA pH 4.8; (c) 29mer DNA-RNA pH 4.8; (d) 29mer DNA^UMA pH 8.9. (e) poly rU; (f) poly rA. poly rU; (g) poly rU*poly rA.poly rU; (h) poly rC;
(i) poly rG.poly rC; (j) poly rC"*poly rG.poly rC. [Marker of U*A-T or U*A-U triplex formation: decrease of therelativeintensity of the adenosine band {IIIIf). Marker
of C+'G-C triplex formation: emergence of a cytidine band (::::::).]
phate symmetric stretching vibration located around 1090/cm as
internal standard.
RESULTS AND DISCUSSION
Infrared spectroscopy: triple helix formation
Figure 2 presents the FTIR spectra recorded in D2O solutions of
the 29mer RNA (Fig. 2a and b) and of the 29mer DNA-RNA
(Fig. 2d and c) in basic and acidic pH conditions. The spectral
region shown contains the absorption bands of the base in-plane
double bond stretching vibrations. This region is extremely
sensitive to base pair formation in duplex and triplex nucleic acids
and contains well established marker bands reflecting the
formation of U*A.U, U*A.T and C+*G.C base triplets. In triple
helices formed by interstrand monobase oligo or polynucleotides
(for instance poly rU*poly rA.poly rU) rU*rA.rU base triplets are
evidenced by a decrease in the relative intensity of the adenine
absorption located around 1632/cm (16,37,38) (Fig. 2e, f and g).
The formation of the rO^rCrC base triplets, for example in poly
rC+*poly rG.poly rC, is reflected on the FTIR spectrum by the
emergence of a high wavenumber absorption assigned to a
C2=O2 carbonyl vibration of protonated cytosine involved in
Hoogsteen type base pairing with guanine in a rG.rC WatsonCrick duplex (17,19) (Fig. 2h, i and j). In the case of the 29mer
RNA studied here, when the pH was decreased, formation of
rU*rA.rU and rO^rCrC base triplets was expected. We observe
on the spectrum of the 29mer RNA recorded at pH 4.8 (Fig. 2b)
that the adenine band located at 1622/cm in the spectrum recorded
at pH 7.4 (Fig. 2a) is no longer present, which shows that the
rU*rA.rU base triplets were formed. Moreover, the spectrum of
the 29mer RNA recorded at pH 4.8 presents a strong new band at
1707/cm characteristic of the formation of rC¥*xG.xC base
triplets.
If we now consider the spectra of 29mer DNA-RNA recorded
in similar pH conditions we can observe that when the pH was
decreased, the adenine contribution located ~1622/cm at pH 8.9
(Fig. 2d) is no longer present (Fig. 2c)reflectingthe formation of
the rU*dA.dT triplets. The weak contribution found at 1633/cm
can be assigned to vibrations of the unpaired thymines in the loop
(this band is observed at 1633/cm in dT 12 or poly dT and is shifted
to 164 I/cm when the dA.dT base pair is formed for instance in
poly dA.poly dT). The emergence of a band at 1702/cm reflects
the formation of the rC+*dG.dC base triplets in the 29mer
DNA-RNA.
Formation of the triplexes was confirmed by the study of the
FTTR spectra of the 29mers recorded in H2O solution. Figure 3
presents the FTIR spectra recorded between 1520 and 1150/cm
of the 29mer DNA-RNA at pH 8.9 (Fig. 3b) and pH 4.8 (Fig. 3c),
as well as that of the 18mer DNA stem (Fig. 3a). The absorption
observed at 1495/cm in the spectra of the 18mer DNA stem and
of the chimeric 29mer DNA-RNA at pH 8.9 involves the bending
vibration of the N7C8H group of purine nucleotides (39). This
absorption is sensitive to interactions at the N7 of these bases. We
can observe that when the pH of the 29mer DNA-RNA sample
is decreased to 4.8, this absorption is shifted to 1489/cm,
reflecting the folding back of the molecule and the binding of the
third strand on the N7 sites of the purine strand of the duplex. A
similar result has been obtained in the case of the 29mer RNA. In
Nucleic Acids Research, 1995, Vol. 23, No. 10 1725
1400
1300
12OO
Figure 3. Fourier transform infrared spectra recorded in H2O solutions of
(a) I8mer DNA stem; (b) 29mer DNA-RNA pH 8.9; (c) 29mer DNA-RNA
pH 4.8; (////) absorption involving the motion of the guanosinc N7 site (:::::::::)
absorption characteristic of thymidines in B family helices.
this case the purine absorption involving the N7C8H bending
vibration is shifted still further from 1495 to 1478/cm (spectra not
shown).
This spectral region can give us other information concerning
the geometry of the molecules studied. Thus, in the case of the
29mer DNA-RNA, the thymidine nucleotides are in a B
family-form geometry as shown by the presence of the 128 I/cm
absorption band. For thymidines involved in A family-form
geometries this absorption is shifted to lower wave numbers
(~1275/cm) (40).
Infrared spectroscopy: sugar conformations
Figure 4 presents the FTIR spectra of the 29mer RNA and
DNA-RNA between 950 and 750/cm, region in which bands
characteristic of the sugar conformations are observed, at neutral
and acidic pH. Marker IR absorptions of the N-type sugars (C3'
endo/anti, A family-form geometry) are observed around 866 and
814/cm, while the S-type sugars (C2' endo/anti, B family-form
geometry) are detected by an absorption located ~834/cm (for
review see 40). RNA oligomers classically contain sugars in
N-type geometry (A family-type conformation). As expected, the
spectra of the 29mer RNA in double-stranded as well as in
triple-stranded configurations present exclusively N-type sugar
absorptions observed at 866 and 814/cm (Fig. 4a and b). When we
consider the spectrum of the 29mer DNA-RNA recorded at basic
pH (Fig. 3c) we observe a strong absorption at 834/cm and a weak
contribution at 868/cm. The former bandreflectsthe DNA duplex
B family-type structure of the stem with S-type sugars, while the
second contribution can be assigned to the ribose tail r(CUCUCUU) which is expected to contain N-type sugars. When the pH
830
BOO
700
Figure 4. Fourier transform infrared spectra recorded in D2O solutions in the
region of the vibrations characteristic of sugar conformations: (a) 29mer RNA
pH 7.4; (b) 29mer RNA pH 4.8; (c) 29mer DNA-RNA pH 8.9; (d) 29mer
DNA-RNA pH 4.8. (////) N-type sugars ( \ \ \ \ ) S-type sugars.
is decreased and the triple helix formed, we observe a major
modification of the spectrum in this region (Fig. 3d). A very
strong contribution is now detected at 865/cm, while only a weak
band is observed at 834/cm. The binding of the third ribose strand
to the purine deoxyribose strand of the duplex has induced a
repuckering of the sugars which are now mainly in N-type
conformation. The weak contribution at 834/cm may be due to the
deoxy sugars of the d(TTTT) loop, as we have seen above that the
thymidines remain in a B-type geometry. The triple helix
formation with the third strand RNA and the duplex as DNA
appears to be associated with a conversion of the duplex part from
a B-form secondary structure to a geometry containing mainly
N-type sugars.
UV spectroscopy: triple helix stability
We shall first present the results concerning the 29mer DNARNA and the corresponding 18mer DNA stem. In a second part
we shall compare these data with those obtained on the 29mer
RNA.
Figure 5c and d presents the melting profiles of the 29mer
DNA-RNA recorded at pH 7.4 and 4.8, respectively. At pH 7.4
the melting curve is monophasic with a Tm at 50.3°C. When the
pH is lowered to 4.8 still only one transition is observed but with
a Tm of 59.7 ° C. It seems thus reasonable to propose that at pH 4.8
a new structure is present.
To assign these transitions, we have studied the melting
behaviour of the DNA 18mer stem. Figure 5a presents the
absorbance versus temperature profile of the 18mer DNA stem
recorded at pH 7.4. Only one transition is detected with a
midpoint located at 50.1 °C. Several thermal denaturation studies
have been previously published concerning oligonucleotide
1726 Nucleic Acids Research, 1995, Vol. 23, No. 10
b)
20 3O 4O 5O 60 70
20 30 -+O 50 60 70
20 30 4O 30 60 70
Figure 5. UV spectroscopy: plots of the absorbance at the maximum -260 nm versus temperature of: (a) 18mer DNA stem d(GAGAGAA-CCCC-TTCTCTC) pH 7.4.
(b) 18mer DNA stem d(GAGAGAA-CCCC-TTCTCTC) pH 4.8. (c) 29mer DNA-RNA d(GAGAGAA-CCCC-TTCrCTC-TTTT)-r(CUCUCUU) pH 7.4. (d) 29mer
DNA-RNA d(GAGAGAA-CCCC-TTCTCTC-TTTT)-r(CUCUCUU) pH 4.8. (e) 29mer RNA r(GAGAGAA-CCCC-UUCUCUC-UUUU-CUCUCUU) pH 7.4.
(f) 29mer RNA r(GAGAGAA-CCCC-UUCUCUC-UUUU-CUCUCUU) pH 4.8. The ordinate scale is in arbitrary units: the curves are normalized considering
arbitrary values of optical density equal to 0 and 1 at the beginning and the end of the experiment.
folding in solution (42-44). Generally a two-step melting profile
was observed. The first transition occurring at low temperatures
is concentration-dependent and assigned to an intermolecular
duplex ++ hairpin conversion. The second one, independent of
oligomer concentration, was assigned to the hairpin ++ coil
conversion. In our case the possible duplex ++ hairpin transition
is not observed Two factors may explain this. First, the thermal
treatement of our sample, heated at 90° C for 5 min then quickly
cooled, to ensure a maximum formation of hairpin structure in
solution. Secondly, the low oligomer concentration which favors
the existence of intramolecular hairpins rather than intermolecular duplexes. We can see that the melting temperature of
the 18mer DNA stem at neutral pH (7"m = 50.1 °C) is very close
to that of the 29mer DNA-RNA (Tm = 50.3°C) in similar salt,
concentration and pH conditions. It seems thus reasonable to
consider that in these conditions the 29mer DNA-RNA is formed
by a Watson-Crick base paired DNA hairpin d(GAGAGAACCCCTTCTCTC) with a dangling llmer DNA-RNA
d(TTTT)-r(UUCUCUC) at the 3' end. Such a model has been
proposed for a 28mer DNA with a closely related sequence
d(GAGAGAA-CCCC-TTCTCTC-TTT-CTCTCTT) in similar
pH conditions (34).
When the pH of the 18mer DNA stem is decreased to 4.8 (Fig.
5b) we observe that the midpoint temperature is strongly shifted
down (AT = -10°C). A significant destabilisation of the stem part
of the hairpin is detected. The decrease of the pH induces
protonation of the cytosines in the DNA loop. An electrostatic
repulsion between the adjacent protonated cytosines may then
occur leading to partial opening of the double-stranded stem
adjacent to the loop. This is completely different from what was
observed for the 29mer DNA-RNA. In the latter case the decrease
of the pH of the solution down to 4.8 induced a shift of the melting
temperature to higher values (AT = 9.4°Q which can be
explained as follows. The protonation of the cytosines in the
dangling RNA part of the 29mer molecule now allows the
formation of rC+*dG.dC base triplets, which in alternation with
rU*dA.dT base triplets form a stable triple helix. A monophasic
melting profile is then detected with a Tm at 59.7°C. Such
monophasic melting has been previously reported in the case of
intra- or intermolecular DNA triple helices (26,45) For example
in the case of a 31mer DNA d(AGAGAGAA-CCCC-
TTCTCTCT-TTT-TCTCTCTT) (sequence very similar to that
studied here but exclusively containing deoxyriboses) the intramolecular triplex conformation studied at pH 5.5 predominates
up to at least 60°C, and melts in a single transition rather than
going through an intermediate partial duplex state (46). In our
case the complete formation of the DNA-RNA triple helix
requires a decrease of the pH to a lower value (pH = 4.8) which
is in agreement with the lower pK values of rC nucleotides when
compared to dC nucleotides. In the case of the chimeric 29mer
DNA-RNA the melting curves observed at pH values intermediate between 7.4 and 4.8 show two transitions and we can clearly
see in Table 1 that these two transitions are detected at -50.3 and
59.7°C, which correspond to the values of the monophasic
transitions obtained at pH 7.4 and 4.8, respectively. This should
reflect, at intermediate pH values, the coexistence of molecules
still in the original DNA hairpin conformation with a dangling
llmer at the 3' end (Tm at 5O.3°C) and of molecules in triple
helical geometry (7"m at 59.7°C).
Table 1. Half dissociation temperatures (7"m) for triple helices studied in 20
mMNaCI
TmCO
Sample
pH
18merDNA
7.4
50.1
4.8
40
7.4
50.3
5.8
50.8
57.1
5.4
50.4
59.8
29mer DNA-RNA
59.7
4.8
18mer DNA +7mer RNA
29merRNA
7.4
48.2
4.8
40.9
7.4
27.6
63.7
6.8
29.6
62.9
5.8
33.7
60.7
5.4
39.4
62.9
4.8
62.9
'
Nucleic Acids Research, 1995, Vol. 23, No. 10 1727
T
17
T
15
U
26
U
24G5G3
U 1 7 U , 5 U 2 4 G S G3
Figure 6. NMR spectra of the imino resonances of (a) 29mer DNA-RNA and (b) 29mer RNA at pH 4.8 as a function of temperature. Assignments are indicated for
the resonances at the highest temperature at which they were observed.
The NMR spectra of the imino protons of 29mer DNA-RNA
at pH 4.8 as a function of temperature are presented in Figure 6a.
The assignments shown were obtained from imino-imino and
imino-amino correlations in a NOES Y spectrum at 10°C and will
be described in detail elsewhere. Peaks, indicative of hydrogen
bonds stable on the NMR timescale, could be observed for all the
imino protons involved in base pairing with the exception of C+23
and U29. The disappearance of these signals with increasing
temperature, an effect of the imino proton's increased rate of
exchange with water protons, is directly correlated with the
melting behavior of the molecule. Although this method is not as
precise as thermal denaturation followed by UV spectrophotometry, it does show the melting behavior for individual base pairs.
In the case of 29mer DNA-RNA, the triplex appears to be
relatively stable up to 60 ° C; the disappearance of the G1 and U12
resonances indicate a fraying effect at the helix ends. Both
Watson-Crick and Hoogsteen base paired strands then melt
simultaneously between 60 and 70 °C, consistent with the
monophasic melting behavior observed by UV. We are currently
working towards the structure determination of the 29mer
DNA-RNA by NMR.
We have tried to form a triple helix by addition of a
stoichiometric amount of r(CUCUCUU) to a solution of the
18mer DNA stem. At pH 7.4 the melting profile (not shown), as
expected, was similar to those of the 18mer DNA stem alone and
of the 29mer DNA-RNA. However when the pH was decreased
to 4.8, no stabilization was found; on the contrary, a melting
temperature of 4O.9°C was obtained, comparable again with that
of the 18mer DNA stem, showing that the triple helix was not
formed, but instead a destabilization of the DNA stem had
occurred (Table 1). This shows the important role played in the
case of the 29mer DNA-RNA by the d(TTTT) loop which
bridges the stem and the potential third strand. In the absence of
such a bridge, protonation of the cytosines in the d(CCCC) loop
occurs prior to the formation of the triplex which is shown by the
destabilization of the stem.
The 29mer RNA denaturation has been studied in a pH range
between 7.4 and 4.8. The results are given in Table 1. Two clearly
resolved transitions are observed, one for the duplex (higher r m )
and one for the triplex (lower 7"™). The stability of the triplex
increases markedly as the pH decreases from 7.4 to 4.8 (increase
of the lower Tm value from 28 to 60 °C) whereas the Tm of the
duplex is constant over this pH range (~60°C). A possible
explanation for the biphasic transition at neutral pH (Fig. 5e) may
be that some of the rU*rA.rU base triplets are already formed.
The first transition (27.6°C at pH 7.4) would then correspond to
the disruption of the weakly bound third strand. When the pH is
decreased, cytosines of the third strand are progressively
protonated which allows the formation of some rCl"*rG.rC base
triplets stabilizing an incompletely formed triple helix, hence a
higher temperature for the disruption of the third strand (29.6°C
at pH 6.8,33.7°C at pH 5.8 and 39.4°C at pH 5.4). Finally at pH
4.8 (Fig. 5f) the triple helix is well formed and the melting occurs
at 62.9 °C simultaneously for the hydrogen bonds of the third
strand and the Watson-Crick bonds of the initial duplex stabilized
by the third RNA 'arm'. The same pH-dependence of the lower
Tm had been observed for the d((T4O-)3T4)*d((A4G)3A4)-d((T4C)T)4) triple helix (47). In that case also the oligomer has two
clearly resolved transitions, one for the duplex (higher T^ and
one for the triplex (lower T^. The latter increases with
decreasing pH and finally <pH 6 triplex and duplex melt at the
same temperature (which moreover is the melting temperature of
the duplex alone at any pH).
The NMR spectra of the imino protons of 29mer RNA as a
function of temperature are presented in Figure 6b. The
assignments of the imino protons based on the imino-imino and
imino-amino NOESY correlations are described elsewhere (48).
At 60°C imino resonances from triplets U24A2U17 and
U26A4U15 are still intense and sharp, indicating that these triplets
are not melted, in agreement with the data presented on the similar
intramolecular DNA triple helix (14). Our results obtained on the
772* Nucleic Acids Research, 1995, Vol. 23, No. JO
29mer RNA at pH 4.8 indicate, as in the case of the 28mer DNA
studied by Macaya et al. (14), that the intramolecular triplex
conformation predominates up to at least 60°C and that it melts
in a single transition rather than going through an intermediate
partial duplex state, which is consistent with the results of optical
melting experiments.
Comparison of the results obtained for the 29mer DNA-RNA
and the 29mer RNA triple helices shows that the stability of the
triple helix formed by three ribose strands is greater than that
obtained by binding of a third ribose strand to a deoxyribose
duplex. This influence of the chemical nature of sugars on triple
helix stability had been previously pointed out (25,27-28). Our
results are in agreement with the data presented by Roberts and
Crothers (27) obtained on triple helices in which the third strand
is identical in length to the duplex used. Moreover our work
concerns intramolecular triple helices, stabilized by two tetrameric
loops, which is also the case for the duplexes used by Roberts and
Crothers. A different order for the triplex stabilities (r*d-d triplex
more stable than r*r-r triplex) has been proposed by Escude et al.
(25) and Han and Dervan (28). In these two cases the results have
been obtained by targeting a long duplex (23 or 35mer) by a far
shorter oligomer (11 or 18 mer). In such a system the melting of
the duplex part is not considered and thus the simultaneous
disruption when the temperature is increased of the Hoogsteen
and Watson-Crick hydrogen bonds which occurs in the triplexes
formed by three strands of identical length not taken into account.
This may explain the difference in the observed stability order
between the two-types of experiments.
In summary, triple helix formation of d(GAGAGAA-CCCCTTCTCTC-TTTT) r(CUCUCUU) induced by folding of the
chimeric molecule by lowering of the solution pH has been
demonstrated. Binding of the third RNA strand to the duplex
DNA induces a conformational transition of the duplex part
initially in B-form geometry with S-type sugars to a structure
containing mainly N-type sugars.
REFERENCES
1 PostelJE-H., Flint,SJ., Kessler.DJ. and Hogan,M.F. (1991) Proc. Natl.
Acad. Sci. USA, 88, 8227-8231.
2 Maher, LJ., Ill (1992) Biochemistry, 31, 7587-7594. .
3 Gngoriev,M., PraseuthJ)., GuyesseAL., RobinJ5., Thuong,N.T.,
Hdlene.Cand Harel-BellanA (1993) C.R. Acad. Sci., Life Sci., 316,
492-495.
4 FrancoisJ.C, Saison-Behmoaras,T., Barbier.C, Chassignol,M., Thuong,
NT.
5 and Helene.C. (1989) Proc. Natl. Acad. Sci. USA, 86, 9702-9706.and
Helene,C. (1989) Proc. Nail. Acad. Sci. USA, 86, 9702-9706.
6 Strobel,S.A.and Dervan,P.B. (1991) Nature (London), 350, 172-174.
7 MaherXJ-, m, Wold.B. and Dervan,P.B. (1989) Science, 245, 725-730.
8 PanuewskiJ5., Kwinkowskijvi., WilkA and KlysikJ. (1990) Nucleic
Acids Res., 18,605-611.
9 Amott,S.and Seising J£. (1974) / Mol. Biol., 88, 509-521.
10 Arnott^., BOTKLPJ., Selsing.E. and Smith.PJ.C. (1976) Nucleic Acids Res.,
3, 2459-2470.
11 Rajagopal J>. and FeigonJ. (1989) Nature (London), 339, 637-640.
12 De los Santos,C, Rosen,M. and Patel.D. (1989) Biochemistry, 28,
7282-7289.
13 PilchJD.S., Levenson,C.and Shafer,R.H. (1990) Proc. NatL Acad. Sci.
USA, 87, 1942-1946.
14 MacayaJR.F., GilbertJ3.E., Malek,S., SinsheimerJ. and FeigonJ. (1991)
Science, 254, 270-274.
15 Thomas.G.A. and Peticolas.W.L. (1983) J. Am. Chem. Soc., 105, 986-992.
16 LiquierJ., CoffinierJ5., Firon,M. and Taillandier,E. (1991) / Biomol.
Struct. Dyn., 9, 437^45.
17 AkhebatA., Dagneaux.C, LiquierJ. and Taillandiei\E. (1992) J. Biomol.
Struct. Dyn., 10, 577-588.
18 Howard,F.B., Miles,H.T., Liu.K., FrazierJ., Raghunathan.G. and
Sasisekharan.V. (1992) Biochemistry, 31, 10671-10677.
19 Oualijvi., Letellier.R., Adnet,F, LiquierJ., SunJ.S., Lavery.R. and
Taillandier.E. (1993) Biochemistry, 32, 2098-2103.
20 Ouali,M., Letellierji, SunJ.S., AkhebatA, Adnet,F., LiquierJ. and
Taillandier,E. (1993) J. Am. Chem. Soc., 115, 4264-^270.
21 LjquierJ. Letellier.R-, Dagneaux.C, Ouali,M., Morvan,F, Raynier.B.,
ImbachJ.L. and Taillandier,E. (1993) Biochemistry, 32, 10591-10598.
22 Xodo,L.E., Manzini.G., QuadnfoglioJ1., Van der Marel.G. and Van
BoomJ.H. (1991) Nucleic Acids Res., 19, 5625-5631.
23 MergnyJ.L., CollierJ)., RougieAl-, Montenay-Garestier.T^nd Hilfene.C.
(1991) Nucleic Acids Res., 19, 1521-1526.
24 Roug£e,M, Faucon,B., MergnyJ.L., BarceloJ1'., Giovannangeli,C,
Garestier.T. and H^lfene.C. (1992) Biochemistry, 31, 9269-9278.
25 Escud£,C, Fran^oisJ.C, SunJ.S., Ott,G., Sprinzjvi., Garcstier.T. and
Hflfcne.C. (1993) Nucleic Acids Res., 21, 5547-5553.
26 Wang.E., Malek,S. and FeigonJ. (1992) Biochemistry, 31,4838-^846.
27 Roberts.R.W. and Crothers,D.M. (1992) Science, 258, 1463-1466.
28 Han.H. and Dervan,P.B. (1993) Proc. NatL Acad. Sci. USA, 90,
3806-3810.
29 Semerad.C.L. and MaherJ^J., Ill (1994) Nucleic Acids Res., 22,
5321-5325.
30 Han,H. and Dervan,P.B. (1994) Nucleic Acids Res., 22, 2837-2844.
31 Le Doan.T, PerrouaulO.., PraseuthJD., Habhoub,N., DecoutJL.,
ThuongJvl.T., LhommeJ. and Heiene.C. (1987) Nucleic Acids Res., 15,
7749-7760.
32 MoserJrI.E. and Dervan,P.B. (1987) Science, 238, 645-650.
33 Lyanuchev.V.I., Mirkin.S.M., Frank-Kamenitskii,D. and Cantor,C.R.(1988)
Nucleic Acids Res., 16, 2165-2178.
34 Sklenar.V and FeigonJ. (1990) Nature (London), 345, 836-838.
35 Fasman,G.D. (1975) in Handbook of Biochemistry and Molecular Biology,
vol. I,p597.
36 Smallcombe.S. (1993) /. Am. Chem. Soc., 115, 4776^785.
37 MitesJ-I.T. and FrazierJ. (1964) Biochem. Biophys. Res. Comm., 14,
21-28.
38 Ohms J. and Ackerman.T. (1990) Biochemistry, 29, 5237-5244.
39 Adam,S., BourtayreJ5., LiquierJ. and TaillandicrJE. (1986) Nucleic Acids
Res., 14, 3501-3513.
40 LiquierJ., AkhebatA., Taillandierji., Ceolin.F, Huynh-Dinh,T. and
IgolenJ. (1991) Spectrochimica Acta, 47a, 177-186.
41 TaillandierJE. and LiquierJ. (1992) in Methods in Enzymology, HJ.Lilley
and JJ.Dahlberg (eds) Acad. Press, 211, 307-335.
42 Wemmer,D.E., Hare,D.R.and Reid3R. (1985) Nucleic Acids Res., 13,
3755-3771.
43 Haasnoot,,C.A.G., Hilberts.C.W., van dcr Marel,G-A., van BoomJ.H.,
Singh.U., Pattabhiraman N.and Kollman^.A. (1986) / Biomol. Struct.
Dyn., 3, 843-857.
44 Xodo,L-E., Manzini,G., Duadrifoglio,F, van der Marel.G.A.and van
BoomJ.H. (1988) Biochemistry; 27, 6321-6326.
45 Kan.L.S., CallahanJJ.E., TrapaneJ.L. and MillerJ>.S. (1991) / Biomol.
Struct. Dyn., 8, 911-933.
46 Macaya,R., Wang,E., SchultzeJ3., Sklenar,V. and FeigonJ. (1992) J. Mol.
Biol., 225, 755-773.
47 Wilson,W.D., Hopkins,H.P., Mizan.S., Hamilton,D.D. and Zon,G. (1994) J.
Am. Chem. Soc., 116, 3607-3608.
48 Klinck.R., GuittetJE., LiquierJ., Taillandier.E., GouyeOe.C. and Huynh
Dinh.T. (1994) FEBS Lett., 355, 297-300.