Synthesis and evaluation of new spacers for use as dsDNA end-Caps

Article abstract of DOI:10.1021/bc100202y

A series of aliphatic and aromatic spacer molecules designed to cap the ends of DNA duplexes have been synthesized. The spacers were converted into dimethoxytrityl-protected phosphoramidites as synthons for oligonucleotides synthesis. The effect of the spacers on the stability of short DNA duplexes was assessed by melting temperature studies. End-caps containing amide groups were found to be less stabilizing than the hexaethylene glycol spacer. End-caps containing either a terthiophene or a naphthalene tetracarboxylic acid diimide were found to be significantly more stabilizing. The former showed a preference for stacking above an A·T base pair. Spacers containing only methylene (-CH₂-) and amide (-CONH-) groups interact weakly with DNA and consequently may be optimal for applications that require minimal influence on DNA structure but require a way to hold the ends of double-stranded DNA together.

Full text of DOI:10.1021/bc100202y

Bioconjugate Chem. 2010, 21, 1545–1553
1545
Synthesis and Evaluation of New Spacers for Use as dsDNA End-Caps
Pei-Sze Ng,† Brian M. Laing,†,‡ Ganesan Balasundarum,† Maneesh Pingle,† Alan Friedman,§ and
Donald E. Bergstrom*,†,‡
Department of Medicinal Chemistry and Molecular Pharmacology, Department of Biological Sciences, and Bindley Bioscience
Center, Purdue University, West Lafayette, Indiana 47907. Received April 25, 2010; Revised Manuscript Received June 18, 2010
A series of aliphatic and aromatic spacer molecules designed to cap the ends of DNA duplexes have been
synthesized. The spacers were converted into dimethoxytrityl-protected phosphoramidites as synthons for
oligonucleotides synthesis. The effect of the spacers on the stability of short DNA duplexes was assessed by
melting temperature studies. End-caps containing amide groups were found to be less stabilizing than the
hexaethylene glycol spacer. End-caps containing either a terthiophene or a naphthalene tetracarboxylic acid diimide
were found to be significantly more stabilizing. The former showed a preference for stacking above an A•T base
pair. Spacers containing only methylene (-CH2-) and amide (-CONH-) groups interact weakly with DNA and
consequently may be optimal for applications that require minimal influence on DNA structure but require a way
to hold the ends of double-stranded DNA together.
studies because of its inherent disorder. A disadvantage of the
stilbene linkers is that they are photosensitive. In addition, an
X-ray structure of a duplex oligonucleotide capped by a stilbene
linker has been obtained, and it was found that stilbene has
considerable influence on the crystallization because of its
tendency to self-associate (13) and the perylene-based linkers
have been shown to self-associate to form dimerized hairpins
(14). With these considerations in mind, we designed a set of
very simple amide containing spacers, in which the total spacer
length was close to the optimal 16-17 Å phosphate-oxygen to
phosphate-oxygen distance required to span the ends of a duplex
without distortion. The amide groups were expected to confer
significantly more rigidity to the spacers than would be expected
for hexaethylene glycol. On the basis of X-ray crystallographic
studies of DNA polymerases, it is known that amide groups
interact with the top face of a duplex base pair (15, 16). This
interaction appears to involve both a CH-π interaction (acidic
R-H positioned in close proximity with the π-orbitals of a
nucleobase) and stacking of an amide group and may provide
part of the driving force for the conformation switch between
the open and closed forms of the DNA polymerase (16). In the
event that these interactions are significant, and realizing that
one could not reasonably design a spacer with sufficiently
precise orientation and positioning of the amide groups, we
chose to make a series of amide-containing spacers in which
the amide positions vary (Figure 1). At the same time, the scope
of the study was expanded by including hydrophobic spacers
in order to weigh the influence of hydrophobicity on duplex
stability in direct comparison to the more hydrophilic amide
containing spacers. For these comparative studies, we have
adopted a very simple model system, a four base pair stem, for
which Tm values fall in a range that can be readily measured,
but for which differences in capping ability would be easily
differentiated.
INTRODUCTION
Short double-stranded DNA segments are required as com-
ponents of DNA-protein complexes for crystallographic studies.
However, it is often difficult to co-crystallize a protein with an
oligonucleotide segment that constitutes the recognition/binding
site. Among strategies that merit investigation as potentiators
of co-crystallization is the use of short preorganized DNA
segments in crystallization experiments. One way to effectively
preorganize short oligonucleotides into a duplex structure is to
covalently link the two complementary strands (1). It is well-
established that covalently attached linkers spanning the ends
ofcDNAstrandssignificantlystabilizetheduplexstructure(2-10).
This enhanced stabilization could be useful for increasing the
preorganization of DNA duplex fragments that are commonly
involved in the recognition and binding of enzymes involved
in degradation, repair, and synthesis of nucleic acids. We have
previously shown that DNA “end-capped” at both ends by non-
nucleotide linkers are useful for stabilizing short duplex
sequences for studying the DNA footprint of T4 DNA ligase
(11).
Ideally, the linker should be a molecule that would not alter
the duplex in a way that would either hinder binding or prevent
readjustment of the structure as dictated by the binding protein.
A linker that allows the protein to dictate the crystallization
process would be preferable. Although there are a number of
well-established linkers, particularly hexaethylene glycol, which
is commercially available as a DMTr protected phosphoramidite,
the family of stilbene linkers originally developed by Letsinger
and co-workers (8, 9) and the naphthalene and perylene based
linkers reported by McLaughlin and co-workers (3, 12), we
thought it worthwhile to expand the potential repertoire of
linkers. We were concerned that the hexaetheylene glycol spacer
was neither sufficiently stabilizing nor useful for crystallographic
EXPERIMENTAL PROCEDURES
* Correspondence to Donald Bergstrom, Department of Medicinal
Chemistry and Molecular Pharmacology, Birck Nanotechnology Center,
Purdue University, 1205 W State St, West Lafayette, IN 47907-2057.
Telephone: (765) 494-6275; Fax: (765) 494-1414. E-mail: bergstrom@
purdue.edu.
General Information. Reagents were purchased from Aldrich
Chemical Co., Inc. (Milwaukee, WI) and were used as obtained
unless otherwise stated. Anhydrous solvents used for synthesis
were purchased in Sure-Seal containers or distilled from
appropriate drying reagents. 2-Cyanoethyl tetraisopropylphos-
phorodiamidite and tetrazole were purchased from ChemGenes
† Department of Medicinal Chemistry and Molecular Pharmacology.
§ Department of Biological Sciences.
‡ Bindley Bioscience Center.
10.1021/bc100202y  2010 American Chemical Society
Published on Web 07/16/2010

1546 Bioconjugate Chem., Vol. 21, No. 8, 2010
Ng et al.
Figure 1. Structure of end-capped DNA sequences. (A) Synthetic scheme for the incorporation of end-caps into oligonucleotides sequences. (B)
Structures of end-caps used in this study. (C) Tetramer oligonucleotides sequences.
Corporation (Waltham, MA). Standard phosphoramidite mono-
mers and other reagents for oligonucleotide synthesis were
purchased from Glen Research Corp. (Sterling, VA). Thin-layer
chromatography (TLC) was performed using Sigma-Aldrich
TLC plates, silica gel on aluminum 60F-254 plates. Purification
of compounds by silica gel chromatography was performed
using Aldrich 230-400 mesh, 60 Å silica gel. Nuclear magnetic
resonance spectra were obtained using 250, 300, or 500 MHz
phoramidite chemistry. Oligonucleotides were either purified
by 20% polyacrylamide gel (PAGE, 60 cm × 40 cm × 0.15
cm) (trityl off) with 7 M urea in 1× Tris-borate-EDTA (TBE)
buffer at a constant power of 60 W or by HPLC (trityl on) on
a Varian C18 Dynamax-100 60 Å column (7.9 mm × 250 mm)
at a flow rate of 4 mL/min. Mobile phase A was 0.1 M
triethylammonium acetate (pH 7.0)/5% acetonitrile and mobile
phase B was 90% acetonitrile. The solvent program was a
gradient starting at 100% A and linearly increasing to 45% B
in 60 min. For analytical HPLC, a Varian Microsorb-MV-100
Å (4.6 mm × 250 mm) column at a flow rate of 1 mL/min was
used. The diode array detector was set at 260 nm. The
oligonucleotides were desalted by 2-propanol precipitation or
1
Bruker spectrometers. H and 13C spectra were referenced to
TMS, while 31P spectra were referenced to an 85% phosphoric
acid external standard.
Oligonucleotides (Figure 1) were synthesized on an Applied
Biosystems 392 DNA/RNA synthesizer using standard phos-

New Spacers for Use as dsDNA End-Caps
Bioconjugate Chem., Vol. 21, No. 8, 2010 1547
Scheme 1. Synthetic Pathway for the Preparation of the
Phosphoramidite Derivative of End-Cap 2a
Scheme 2. Synthetic Pathway for the Preparation of the
Phosphoramidite Derivatives of End-Caps 8 and 14a
a Reagents and conditions: (i) BuLi, -78°C; (ii) BF3 - Et2O, oxetane;
(iii) DMTrCl, pyridine; (iv) 2-cyanoethyl-N,N-diisopropyl-chlorophos-
phoramidite, diisopropylethylamine.
by elution using CH3CN/H2O (1:1) from C18 Sep-pak cartridges
(Waters). The oligonucleotides were characterized by MALDI-
TOF MS (Supporting Information Table S4).
Spacer Synthesis. The synthetic transformations are sum-
marized in Schemes 1-4.
2-(tert-Butyldimethylsilyloxy)ethylamine (6). To a solution of
ethanolamine (5) (7.24 mL, 120 mmol, 1 equiv) in anhydrous
CH2Cl2 (100 mL) at 0 °C were added anhydrous triethylamine
(18.5 mL, 132 mmol, 1.1 equiv), tert-butyldimethylsilyl chloride
(20.10 g, 132 mmol, 1.1 equiv), and a catalytic amount of
4-(N,N-dimethylamino)pyridine (DMAP) (0.5 g). The reaction
mixture was stirred under argon overnight. The reaction was
quenched with water (50 mL) and the layers separated. The
organic layer was washed with water, brine, dried (Na2SO4),
filtered, and concentrated in vacuo to give colorless oil (17.96
a Reagents and conditions: (iii) DMTrCl, pyridine; (v) tBuMe2SiCl,
N,N-dimethylaminopyridine (DMAP), Et3N; (vi) diglycolyl chloride,
Et3N; (vii) HCl, EtOH; (viii) 2-cyanoethyl-N,N,N′,N′-tetraisopropy-
lphosphorodiamidite, tetrazole.
dimethoxytrityl chloride (3.23 g, 90.8 mmol, 1 equiv). The
reaction mixture was stirred under argon overnight. Upon
completion as judged by TLC, pyridine was removed in vacuo.
The crude brown product was dissolved in CH2Cl2 (40 mL),
washed with aqueous 5% NaHCO3 and brine, and dried
(Na2SO4). CH2Cl2 was removed in vacuo and the crude product
purified by flash chromatography (silica gel, 0-5% CH3OH/
1
g, 86%). H NMR (250 MHz, CDCl3, δ, ppm): 3.63 (t, 2H, J
) 5.5 Hz), 2.78 (t, 2H, J ) 6 Hz), 0.81 (s, 9H), 0.06 (s, 6H).
13C NMR (75 MHz, CDCl3, δ, ppm): 65.16, 44.27, 25.90, 18.30,
-5.33. HRMS (EI, M-CH3+): calcd. for C7H18NOSi 160.1158;
act. 160.1165.
N,N′-Bis[2-(tert-butyldimethylsilyloxyethyl]-2,2′-oxidiaceta-
mide (7). To a solution of 6 (16.26 g, 93 mmol, 2.2 equiv) in
anhydrous CH2Cl2 (40 mL) at 0 °C was added anhydrous
triethylamine (17.8 mL, 127 mmol, 3 equiv), and diglycolyl
chloride (5 mL, 42.5 mmol, 1 equiv) was added dropwise with
a syringe. The reaction mixture was stirred under argon
overnight. The reaction was quenched with water (40 mL) and
the layers separated. The organic layer was washed with 5%
acetic acid (2 × 40 mL), 5% aqueous NaHCO3 (2 × 40 mL),
and brine, and dried (Na2SO4) and concentrated in vacuo to give
1
1% Et3N/CH2Cl2) to give a brown foam (1.67 g, 35.2%). H
NMR (250 MHz, CD2Cl2, δ, ppm): 6.82-7.46 (m, 13H), 4.03
(s, 2H), 4.01 (s, 2H), 3.78 (s, 6H), 3.59 (m, 2H), 3.48 (m, 2H),
3.33 (m, 2H), 3.20 (m, 2H). 13C NMR (75 MHz, CD2Cl2, δ,
ppm): 169.81, 168.93, 158.98, 145.32, 136.31, 130.32, 128.34,
128.21, 127.22, 113.52, 86.53, 71.44, 62.52, 62.02, 55.61, 42.23,
39.54. HRMS (ESI, M+H+): calcd. for C29H35N2O7 523.2444;
act. 523.2445.
N-[3-O-(2-cyanoethyl-N,N-diisopropylphosphoramidite)ethyl]-
N′-[2-(4,4′-dimethoxytrityl)oxyethyl]-2,2′-oxidiacetamide (10).
To a solution of 9 (211.2 mg, 0.404 mmol, 1 equiv) in anhydrous
CH2Cl2 (5 mL) were added anhydrous diisopropylethylamine
(0.25 mL, 1.45 mmol, 3.6 equiv) and 2-cyanoethyldiisopropyl
chlorophosphoramidite (0.49 mL, 0.485 mmol, 1.2 equiv). The
reaction was stirred for two hours at room temperature under
argon. When the reaction was completed as judged by TLC,
the reaction mixture was washed with aqueous 5% NaHCO3
and water and dried (Na2SO4). The crude product was further
purified by flash chromatography (silica gel, 5:4:1 hexane/
CH2Cl2/Et3N) to give a yellow oil (106.7 mg, 53.5%). 31P NMR
(100.8 MHz, CD2Cl2, δ, ppm): 148.3 (referenced to external
85% H3PO4).
1
a brown oil (14.75 g, 78%). H NMR (250 MHz, CDCl3, δ,
ppm): 4.07, (s, 4H), 3.71 (t, 4H, J ) 5.6 Hz), 3.41-3.48 (t,
4H, J ) 5.6 Hz), 0.090 (s, 18H), 0.076 (s, 9H). 13C NMR (75
MHz, CDCl3, δ, ppm): 168.14, 71.16, 61.58, 40.99, 25.78,
18.19, -5.42. HRMS (EI, M+H+): calcd. for C20H45N2O5Si2
449.2867; act. 449.2849.
N,N′-Bis(2-hydroxyethyl)-2,2′-oxidiacetamide (8; Scheme 2).
To a solution of 7 (14.06 g, 31.3 mmol) in 95% ethanol (15
mL) was added 2.9% w/w of conc. HCl in 95% ethanol (15
mL). The reaction mixture was stirred under argon for 15 min
and solvent removed in vacuo to give an orange solid. The
orange solid was dissolved in methanol (20 mL), washed with
hexane (3 × 30 mL), dried (Na2SO4), and concentrated in vacuo
1
to give an orange solid (6.47 g, 94%). H NMR (250 MHz,
N,N′-Bis-(4-hydroxybutanoyl)-1,3-propanediamine (18; Scheme
3). 1,3-Diaminopropane (2.0 mL, 24 mmol), δ-butyrolactone
(17) (6.2 mL, 80 mmol), and DMAP (40 mg, 0.32 mmol) in
methanol (30 mL) were kept under reflux for overnight. The
solvent was removed by rotary evaporation, and CH2Cl2 (50
mL) was added to the residue with stirring on low heat to break
up the white-colored solid. The solid was filtered and mixed
with CH2Cl2 (50 mL) and again stirred at low heat and filtered.
CDCl3, δ, ppm): 4.06 (s, 4H), 3.63 (t, 4H, J ) 5.8 Hz),
3.34-3.39 (m, 4H). 13C NMR (75 MHz, CDCl3, δ, ppm):
172.00, 71.42, 61.44, 42.58. HRMS (EI, M-OH+): calcd. for
C8H15N2O4 203.1032; act. 203.1034.
N-[2-(4, 4′-Dimethoxytrityl)oxyethyl]-N′-(2-hydroxyethyl)-
2,2′-oxidiacetamide (9). To a solution of 8 (2.00 g, 90.8 mmol,
1 equiv) in anhydrous pyridine (35 mL) at 0 °C was added 4,4′-

1548 Bioconjugate Chem., Vol. 21, No. 8, 2010
Ng et al.
Scheme 3. Synthetic Pathway for the Preparation of the
Phosphoramidite Derivatives of End-Caps 18 and 22a
silica gel chromatography in a gradient of methanol (0% to 5%)
in CH2Cl2 containing 0.5% Et3N. The appropriate fractions were
combined, evaporated, and dried under vacuum to give phos-
phoramidite 20 as pale yellow product (0.4 g, 54%), Rf 0.90
(CH2C2-MeOH 9:1 v/v); HRMS (ESI, M+H+) calcd. for
C41H57N4O7P 749.4043, found 749.4050; 31P NMR (100.8 MHz,
CDCl3, ppm) δ 165.0 (referenced to external 0.0485 M
triphenylphosphate in CDCl3).
N,N′-Bis-(5-hydroxypentanoyl)-1,3-propanediamine (22). 1,3-
Diaminopropane (1.7 mL, 20 mmol), δ-valerolactone (7.5 mL,
80 mmol), and DMAP (40 mg, 0.32 mmol) in methanol (30
mL) were refluxed overnight. The solvent was removed by
rotary evaporation, and CH2Cl2 (50 mL) was added to the
residue with stirring on low heat to break up the white-colored
solid. The solid was filtered and mixed with CH2Cl2 (50 mL)
and again stirred at low heat and filtered. This procedure was
repeated once more, and the final white solid was dried under
vacuum (4.65 g, 85%). 1H NMR (250 MHz, DMSO-d6, δ, ppm)
1.31-1.54 (m, 10 H), 2.03 (t, J ) 9 Hz, 4H), 2.99 (t, J ) 6 Hz,
2H), 3.01 (t, J ) 6 Hz, 2H), 3.33-3.38 (m, 4H), 4.35 (s, 2H),
7.74 (t, J ) 3 Hz, 2H); 13C NMR (75 MHz, DMSO-d6, δ, ppm)
22.30, 29.75, 32.42, 35.63, 36.60, 60.79, 172.45. HRMS (ESI,
M+H+) calcd. for C13H26N2O4 275.1970, act. 275.1972.
N-{5-[4,4′-(Dimethoxytrityl)oxy]pentanoyl}-N′-(5-hydroxy-
pentanoyl]-1,3-propanediamine (23). Diol 22 (2.75 g, 10 mmol)
was co-evaporated with dry pyridine (3 × 20 mL) and then
dissolved in dry pyridine (30 mL), and after cooling to 0 °C in
an ice bath, 4,4′-dimethoxytrityl chloride (4.4 g, 13 mmol) in
CH2Cl2 (20 mL) was added dropwise slowly. The flask was
removed from ice, and the reaction mixture was stirred
overnight, quenched with methanol (1 mL), diluted with CH2Cl2
(200 mL), washed with 5% NaHCO3 (2 × 200 mL) and brine
(100 mL), and dried over anhydrous sodium sulfate. The solution
was concentrated in vacuo, and the crude product was purified
by silica gel chromatography in a gradient of methanol (0% to
5%) in CHCl3 containing 0.5% Et3N. The appropriate fractions
were combined, evaporated, and dried in vacuum to give
compound 23 as pale yellow foam (2.65 g, 46%), Rf 0.5
(CHCl3-MeOH 9:1 v/v); 1H NMR (250 MHz, CDCl3, δ, ppm)
1.54-1.72 (m, 10H), 2.12-2.24 (m, 4H), 3.06 (t, J ) 6 Hz,
2H), 3.19 (t, J ) 6 Hz, 2H), 3.21 (t, J ) 6 Hz, 2H), 3.58 (t, J
) 6 Hz, 2H), 3.75 (s, 6H), 6.81 (d, J ) 6 Hz, 4H), 7.17-7.32
(m, 7H), 7.40-7.43 (m, 2H); 13C NMR (75 MHz, CDCl3, δ,
ppm) 22.01, 22.75, 29.61, 29.63, 31.98, 35.80, 35.89, 36.10,
36.47, 55.18, 61.78, 62.93, 76.75, 77.18, 77.60, 85.77, 112.99,
126.62, 127.71, 128.13, 129.98, 136.49, 145.23, 158.30, 173.92,
173.98. HRMS (ESI, M+Na+) calcd. for C34H44N2O6 599.3097,
found 599.3106.
a Reagents and conditions: (iii) DMTrCl, pyridine; (iv) 2-cyanoethyl-
N,N-diisopropyl-chlorophosphoramidite, diisopropylethylamine; (ix)
1,3-diaminopropane, DMAP, methanol.
This procedure was repeated once more, and the final white
solid was dried under vacuum (4.6 g, 80%). 1H NMR (250 MHz,
DMSO-d6, δ, ppm) 1.43-1.65 (m, 6H), 2.07 (t, J ) 9 Hz, 4H),
2.99 (t, J) 6 Hz, 2H), 3.01 (t, J ) 6 Hz, 2H), 3.34 (t, J )6 Hz,
4H), 4.48 (s, 2H), 7.76 (t, J ) 6 Hz, 2H); 13C NMR (75 MHz,
CDCl3, δ, ppm) 29.01, 29.65, 32.52, 36.62, 60.67, 172.52.
HRMS (ESI, M + H+) calcd. for C11H22N2O4 247.1657, found
247.1657.
N-{4-[4,4′-(Dimethoxytrityl)oxy]butanoyl}-N′-4-hydroxybu-
tanoyl-1,3-propanediamine (19). Diol 18 (2.46 g, 10 mmol) was
co-evaporated with dry pyridine (3 × 20 mL) and dissolved in
dry pyridine (30 mL), and after cooling to 0 °C in an ice bath,
4,4′-dimethoxytrityl chloride (4.4 g, 13 mmol) in CH2Cl2 (20
mL) was added dropwise slowly. The flask was removed from
ice, and the reaction mixture was stirred overnight, quenched
with methanol (1 mL), diluted with CH2Cl2 (200 mL), washed
with 5% NaHCO3 (2 × 200 mL) and brine (100 mL), and dried
over anhydrous sodium sulfate. The solution was concentrated
in vacuo, and the crude product was purified by silica gel
chromatography in a gradient of methanol (0 to 5%) in CHCl3
containing 0.5% Et3N. The appropriate fractions were combined,
evaporated, and dried in vacuo to give compound 19 as pale
yellow foam (2.9 g, 53%), Rf 0.50 (CHCl3-MeOH 9:1 v/v).
1H NMR (250 MHz, CDCl3, δ, ppm) 1.49-1.53 (m, 2H),
1.81-1.94 (m, 4H), 2.31 (t, J ) 6 Hz, 2H), 2.33 (t, J ) 6 Hz,
2H), 3.08-3.20 (m, 6H), 3.63 (t, J ) 6 Hz, 2H), 3.76 (s, 6H),
6.82 (d, J ) 12.3, 4H), 7.18-7.32 (m, 7H), 7.39 (d, J ) 9.0
Hz, 2H); 13C NMR (62.5 MHz, CDCl3, δ, ppm) 26.04, 28.18,
29.42, 33.70, 33.83, 35.75, 35.81, 55.11, 61.96, 62.32, 85.83,
112.96, 126.62, 127.68, 127.99, 129.86, 136.17, 144.94, 158.28,
173.76, 173.95. HRMS (ESI, M + Na+) calcd. for C32H40N2O6
571.2784; act. 571.2785.
N-[4-O-(2-cyanoethyl-N,N-diisopropylphophoramidite)bu-
tanoyl]-N′-{4-[4,4′-(dimethoxytrityl)oxy]butanoyl}-1,3-propanedi-
amine (20). The mono-DMT-derivative 19 (0.55 g, 1 mmol)
was co-evaporated with 10% dry pyridine in CH2Cl2 (3 × 10
mL), and DIEA (0.63 mL, 3.6 mmol) was added with stirring
under nitrogen followed by the addition of 2-cyanoethyl
diisopropylchlorophosphoramidite (0.27 mL, 1.2 mmol). The
reaction was performed at room temperature for two hours with
TLC monitoring. After reaction completion, the mixture was
quenched with 4 drops of anhydrous methanol, and CH2Cl2 (20
mL) was added. The methylene chloride solution was washed
once with 5% NaHCO3 (20 mL), once with brine (10 mL), and
then dried over anhydrous sodium sulfate. The solution was
concentrated in vacuo, and the crude product was purified by
N-[5-O-(2-Cyanoethyl-N,N-diisopropylphosphoramidite)pen-
tanoyl]-N′-{5-[4,4′-(dimethoxytrityl)oxy]pentanoyl}-1,3-pro-
panediamine (24). The mono-DMT-derivative 23 (0.4 g, 0.7
mmol) was co-evaporated with 10% dry pyridine in CH2Cl2 (3
× 10 mL), and DIEA (0.43 mL, 2.5 mmol) was added with
stirring under nitrogen followed by the addition of 2-cyanoethyl
diisopropylchlorophosphoramidite (0.19 mL, 0.84 mmol). The
reaction was performed at room temperature for two hours with
TLC monitoring. After reaction completion, the mixture was
quenched with 4 drops of anhydrous methanol, CH2Cl2 (20 mL)
was added, and the mixture was washed with 5% NaHCO3 (2
× 20 mL) and brine (10 mL) and dried over anhydrous sodium
sulfate. The solution was concentrated in vacuo, and the crude
product was purified by silica gel chromatography in a gradient
of methanol (0% to 5%) in CH2Cl2 containing 0.5% Et3N. The
appropriate fractions were combined, evaporated, and dried in
vacuum to give phosphoramidite 24 as pale yellow product (0.3
g, 55%), Rf 0.92 (CH2Cl2-MeOH 9:1 v/v); HRMS (ESI,
M+Na+) calcd. for C43H61N4O7P 799.4175, found 799.4169;

New Spacers for Use as dsDNA End-Caps
Bioconjugate Chem., Vol. 21, No. 8, 2010 1549
31P NMR (100.8 MHz, CDCl3, δ, ppm) 164.5 (referenced to
external 0.0485 M triphenylpohosphate in CDCl3).
Scheme 4. Synthetic Pathway for the Preparation of the
Phosphoramidite Derivative of End-Cap 29a
Methyl 2-(2-(Benzyloxy)ethoxy)acetate (26). 2-Benzyloxy-
ethanol (25) (9.96 mL, 70 mmol) was added to a suspension of
NaH (3.4 g, 84 mmol) in anhydrous THF (100 mL) under argon.
The mixture was stirred for 1 h at r.t., then cooled to 0 °C.
Methyl bromoacetate (7.72 mL, 84 mmol) was added slowly,
and the mixture was allowed to react at room temperature for
16 h. The reaction was quenched with H2O (30 mL) and the
product extracted with ethyl acetate (5 × 50 mL). The ethyl
acetate fractions were combined and washed successively with
H2O (3 × 100 mL) and brine (3 × 100 mL). The organic
fraction was dried over anhyd MgSO4 and concentrated in
vacuo. The crude product was purified by silica gel chroma-
tography (ethyl acetate-hexanes 1:4). The appropriate fractions
were combined, evaporated, and dried under vacuum to give
26 (13.2 g, 84%) as yellow oil. Rf 0.67 (ethyl acetate-hexanes
1:1 v/v). 1H NMR (300 MHz, CDCl3, ppm): 7.28 (m, 5H), 4.56
(s, 2H), 4.18 (s, 2H), 3.80-3.68 (m, 7H). 13C NMR (75 MHz,
CDCl3,, ppm): 170.8, 138.1, 128.3, 127.7, 73.2, 71, 69.5, 68.7.
HRMS (EI, M+) calcd for C12H16O4 224.1049, act. 224.1045.
2-(2-(Benzyloxy)ethoxy)acetic Acid (27). 26 (7 g, 29.38 mmol)
was dissolved in 20 mL methanol and heated to ∼35 °C with
stirring. KOH (4.95 g, 88.13 mmol) was added. The mixture
was stirred with TLC monitoring (1:1 ethyl acetate/hexanes)
for 1 h. The reaction was quenched by adding 80 mL H2O. The
unreacted ester was extracted with ether (2 × 50 mL). The
aqueous portion was acidified to pH 2 with 6 N HCl, then
extracted with ether (3 × 50 mL); the ether layers were
combined, dried over Na2SO4, and dried under vacuum to yield
27 (5.83 g 95%) as a clear oil. 1H NMR (300 MHz, DMSO-d6,
ppm): 12.62 (s, 1H), 7.33 (m, 5H), 4.48 (s, 2H), 4.04 (s, 2H),
3.55-3.65 (m, 4H). 13C NMR (75 MHz, DMSO-d6, δ, ppm):
171.73, 138.48, 128.26, 127.56, 127.43, 72.07, 71.74, 69.89,
67.16, 60.31. HRMS (ESI, M + H+) calcd. for C11H15O4
211.0970, act. 211.0971.
N,N′-Bis-(2-(2-benzyloxyethoxy)ethanoyl)-1,3-propanedi-
amine (28). 27 (2.13 g, 10.12 mmol) was dissolved in dry DMF
(10 mL) under argon. TBTU (3.25 g, 10.12 mmol) and
N-methylmorpholine (1.02 g, 10.12 mmol) were added and the
mixture stirred for 15 min. 1,3-Diaminopropane (0.28 mL, 4.05
mmol) was dissolved in dry DMF (5 mL) and added slowly.
The mixture was stirred overnight at room temperature. DMF
was evaporated under reduced pressure, and the residue was
dissolved in ethyl acetate (100 mL), and washed with 5%
NaHCO3 (75 mL × 2), 10% CuSO4 (75 mL × 2), and brine
(75 mL). The organic layer was dried over anhyd magnesium
sulfate and concentrated under vacuum. The crude product was
purified by silica gel chromatography (0-5% MeOH/ethyl
acetate). The appropriate fractions were combined, evaporated,
and dried under vacuum to yield 28 (0.92 g, 50%) as a light
brown oil, Rf 0.68 (10% MeOH/ethyl acetate 9:1 v/v). 1H NMR
(300 MHz, DMSO-d6,, ppm): 7.71 (t, J ) 7 Hz, 2H), 7.30 (m,
10H), 4.49 (s, 4H), 3.86 (s, 4H), 3.61 (m, 8H), 3.03 (q, 4H, J
) 7 Hz), 1.44 (t, 2H, J ) 7 Hz). 13C NMR (75 MHz, DMSO-
d6, ppm): 169.17, 138.33, 128.23, 127.52, 72.11, 70.23, 70.01,
68.95, 35.50, 29.38. HRMS (ESI, M + H+) calcd. for
C25H35N2O6 459.2495, act. 459.2502.
a Reagents and conditions: (iii) DMTrCl, pyridine; (iv) 2-cyanoethyl-
N,N-diisopropyl-chlorophosphoramidite, diisopropylethylamine; (x)
methyl bromoactetate, NaH, THF reflux, r.t.; (xi) MeOH, KOH, 35
°C; (xii) 1,3-diaminopropane, TBTU, r.t.; (xiii) H2, Pd/C.
169.53, 72.91, 69.99, 60.13, 35.71, 29.44. HRMS (ESI, M +
Na+) calcd. for C11H22N2NaO6 301.1376, act. 301.1372.
N-(2-(2-(4,4′-Dimethoxytrityloxy)ethoxy)ethanoyl)-N′-(2-(2-
hydroxyethoxy)ethanoyl)-1,3-propanediamine (30). Diol 29 (0.56
g, 2 mmol) was co-evaporated with dry pyridine (3 × 10 mL),
dissolved in dry pyridine (15 mL), and after cooling to 0 °C,
4,4′-dimethoxytrityl chloride (0.81 g, 2.4 mmol) in CH2Cl2 was
added dropwise slowly. The flask was removed from the ice
and the reaction mixture stirred overnight, quenched with
methanol (0.5 mL), and diluted with CH2Cl2 (50 mL). The
mixture was washed with 5% NaHCO3 (50 mL × 2) and brine
(50 mL) and dried over anhydrous sodium sulfate. The solution
was concentrated in vacuo, and the crude product was purified
by silica gel chromatography (0-5% MeOH/CH2Cl2 containing
1% Et3N). The appropriate fractions were combined, evaporated,
and dried in vacuum to give 30 as a yellow foam (0.12 g, 10%),
1
Rf 0.60 (8% MeOH/CH2Cl2). H NMR (500 MHz, DMSO-d6,
ppm): 7.85 (t, 1H, J ) 6 Hz), 7.74 (t, 1H, J ) 6 Hz), 7.38-7.39
(m, 2H), 7.31 (t, 2H, J ) 7.5 Hz), 7.22-7.26 (m, 5H), 6.89 (d,
4H, J ) 9 Hz), 4.76 (t, 1H, J ) 5 Hz), 3.91 (s, 2H), 3.85 (s,
2H), 3.73 (s, 6H), 3.63 (t, 2H, J ) 5 Hz), 3.53 (t, 2H, J ) 5
Hz), 3.47 (t, 2H, J ) 5 Hz), 3.08-3.14 (m, 6H), 1.54 (p, 2H,
J ) 6.5 Hz). 13C NMR (125.76 MHz, DMSO-d6, ppm): 29.37,
35.52, 35.61, 54.96, 59.99, 62.55, 69.92, 70.15, 70.22, 72.78,
85.45, 112.70, 113.11, 123.79, 126.57, 127.63, 127.71, 128.82,
129.57, 135.64, 135.99, 144.82, 149.52, 158.01, 169.07, 169.22.
HRMS (ESI, M+Na+) calcd. for C32H40N2NaO8 603.2682, act.
603.2680.
N-(2-(2-O-(2-Cyanoethyl-N,N-diisopropylphosphoramidite)-
ethoxy)ethanoyl)-N′-(2-(2-(4,4′-dimethoxytrityloxy)ethoxy)etha-
noyl)-1,3-propanediamine (31). 30 (0.11 g, 0.19 mmol) was co-
evaporated with 10% anhydrous pyridine in CH2Cl2 (2 × 10
mL), then dissolved in anhydrous CH2Cl2 (10 mL). DIEA (0.1
mL, 0.76 mmol) was added with stirring under argon followed
by the addition of 2-cyanoethyl diisopropylchlorophosphora-
midite (0.11 mL, 0.47 mmol). The mixture was stirred at room
temperature for 2 h with TLC monitoring (10% MeOH/ethyl
acetate, 1% Et3N). The reaction was quenched with 4 drops
anhydrous methanol and 20 mL CH2Cl2 was added. The reaction
mixture was transferred to a dry 125 mL separatory funnel and
N,N′-Bis-(2-(2-hydroxyethoxy)ethanoyl)-1,3-propanediamine (29;
Scheme 4). 28 (2.73 mg, 5.9 mmol) was dissolved in 40 mL
EtOH. Pd/C (10 wt %, 800 mg) was added, and the mixture
was treated with H2 gas (55 psi) on a Parr apparatus for 12 H.
The mixture was filtered over Celite on a sintered glass funnel,
evaporated, and dried in vacuum to give compound 29 as a white
1
solid (1.548 g, 94%). H NMR (300 MHz, DMSO-d6, ppm):
7.91 (s, 2H), 4.84 (s, 2H), 3.92 (s, 4H), 3.48 (m, 8H), 3.12 (m,
4H), 1.562 (m, 2H). 13C NMR (75 MHz, DMSO-d6, ppm):

1550 Bioconjugate Chem., Vol. 21, No. 8, 2010
Ng et al.
washed quickly with 5% NaHCO3 (10 mL × 2) and brine (10
mL). The organic layer was dried over anhydrous sodium sulfate
and concentrated under reduced pressure. The residue was
purified by silica gel flash chromatography (0-5% MeOH/ethyl
acetate, 1% Et3N). The appropriate fractions were combined,
evaporated, and dried in vacuum to give product as a pale yellow
oil (0.12 g, 81%). 31P NMR (202.46 MHz, DMSO-d6, ppm):
148.68 (referenced to external 85% H3PO4).
For comparison to previously studied end-caps, this study
included hexaethylene glycol (25), the napthalenetetracarboxy-
late dimides 26-28, and a tetrathymidine loop. In addition, we
prepared the terthiophene spacer 2, which provided an op-
portunity to study and compare an exceptionally hydrophobic
end-cap. Several research groups had reported that hydrophobic,
aromaticmoleculessuchasnaphthalenediimides32-34(3,12,17),
N,N-dimethylstilbene dicarboxamide (8), trimethoxystilbenes
(18), and stilbene ether (9) stabilized DNA duplexes through
stacking interactions with the DNA bases. We expected that
end-cap 2, with a terthiophene core, would also have significant
stacking interactions with the base pairs in a DNA duplex. The
hydrophilic end-caps 8, 14, 18, and 22 more closely resemble
the hexaethylene glycol linker (25) in hydrophilicity.
The strategy for synthesis of each of the spacers as a
symmetrical diol is illustrated in Schemes 1-4. Diol 2 could
be obtained in good yield by reaction of the dilithium salt of
terthiophene with 2 equiv of oxetane (10). Diols 8 and 14 were
obtained by reaction of the O-t-butyldimethylsilyl ethers of
ethanolamine and propanolamine with diglcolyl chloride, fol-
lowed by removal of the silyl protecting group with ethanolic
HCl. Details for the synthesis of diol 14 have been reported
(10). Diols 18 and 22 were obtained more directly by the
reaction of either γ-butyrolactone or δ-valerolactone with 1,3-
propanediamine. Diol 29 was obtained by reaction of methyl
bromoacetate with 2-benzyloxyethanol to form the benzyl
protected methyl ester (26), which is converted to the carboxylic
acid derivative for coupling with 1,3-propanediamine to form
the benzyl protected diol 28. Deprotection by catalytic hydro-
genation with Pd/C catalyst gives diol 29 in good yield. The
synthesis of diols 32-34 followed the procedure described by
Pingle et al. (10) Each diol is symmetrical and could be
synthesized as a mono-DMTr derivative by reaction with 1 equiv
of DMTr chloride. Yields are generally in the 35-50% range,
which, although low, was not limiting because of the ease with
which one can produce large amounts of the diols. Separation
of the monotritylated products from starting material and bis-
tritylated side product by silica gel chromatorgraphy was facile
in each case. The tritylated spacers were easily transformed into
the corresponding phosphoramidites by conventional protocols
and the 4,4′-dimethoxytrityl protected phosphoramidite deriva-
tives incorporated into the oligonucleotide on an automated
DNA synthesizer in good yield as illustated in Figure 1A. The
sequences synthesized for this study are included in Tables 2
and 3.
Spacers 2, 14, 32, 33, and 34. The synthetic procedures for
these spacers have been published (10).
Thermal Melting Measurements. Absorbance vs tempera-
ture (melting and annealing curves) profiles of the oligonucle-
otides were determined in Teflon-capped 1 cm path length quartz
cells under a nitrogen atmosphere using a Cary 3 Bio UV-visible
spectrophotometer with a 6 × 6 multicell block Peltier and a
temperature controller. Oligonucleotides were dissolved in 1.0
M NaCl, 10 mM Na2HPO4, 0.1 mM Na2EDTA, pH 7 degassed
buffer, and were prepared in three concentrations: 1 µM, 5 µM,
and 15 µM. Samples were covered with a layer of silicone oil
and denatured by raising the temperature to 85 °C for 10 min
prior to an experiment. Melting and annealing experiments were
conducted in two stages by starting the experiment at 85 °C
and cooling to 5 °C and heating from 5 °C and ending at 85
°C. Absorbance was measured at 260 nm and the heating rate/
cooling rate as 0.3 °C/min with data collection at 0.2 °C
intervals. The absorbance vs temperature data obtained on the
Cary 3 spectrometer were exported in ASCII format to the
software application Microcal Origin. The data were normalized
to produce relative absorbance values, and sigmoidal curves
were obtained using Boltzmann logged data-fit function. First
derivative of the melting curve was obtained from the above
sigmoidal curve and then fitted with a Gaussian function to
identify the center of the peak, which is Tm or the melting
temperature.
RESULTS AND DISCUSSION
Spacer Design. A series of four spacers, each containing two
amide groups (shown as the parent diols 8, 14, 18, and 22 in
Figure 1B), was designed based on three criteria. First, the
spacers in fully extended conformations had terminal oxygen
atoms separated by 14.6 to 17.3 Å. The distances were chosen
to lie within the middle of the range observed in crystal and
NMR structures between the terminal 3′-O and 5′-O of B-DNA
duplexes. We checked these distances by downloading a series
of PDB format files of B-DNA NMR and X-ray structures from
the Protein Data Bank of the Research Collaboratory for
Structural Bioinformatics (RCSB). These were displayed in the
molecular modeling program Sybyl (Tripos Inc.), wherein the
distance between the terminal 3′-O and 5′-O of 17 B-DNA
molecules was found to vary between 13.8 and 18.8 Å. The
average distance between the 3′-O and 5′-O of a B-DNA duplex
is 16.2 Å. The number of atoms (shortest direct path of
connectivity) and the distance between the terminal oxygen
atoms is shown for each spacer in Figure 1. This distance
provides a rough guide for design; the actual distance would
be expected to vary in order for the end-cap to accommodate
the geometry of the terminal base pair. Second, the series of
spacers provides variation in the placement of the carboxamide
groups over the top of the terminal bases. Direct contact appears
to exist between the glycine carboxamide group and the template
base in the closed form of Taq DNA polymerase (15, 16), but
there are too many variables to allow us to design a spacer that
would provide exactly the observed contact geometry. Third,
the linkers and their DMT protected monophosphoramidites
were easy and inexpensive to synthesize and could be incor-
porated into synthetic oligonucleotides on automated synthesiz-
ers with conventional reagents and conditions.
Thermal Melting Studies. Thermal melting studies were
carried out on four-base-pair duplexes containing the nucleosides
dA, T, dG, and dC and end-caps 14 and 18 (Table 2) and 2, 8,
22, 25, 29, 32, 33, and 34 (Table 3). The thermal melting profile
of all end-capped DNA duplexes at varied concentrations (1-15
µM) was a sigmoidal curve indicative of the formation of a
hairpin and a concentration independent, unimolecular process.
If the oligonucleotides were dimerizing rather than forming
hairpin stuctures, one would expect to see an increase in Tm
values with an increase in concentration. In order to determine
the end-cap preferences for specific base pairs, we examined
one sequence, IV, in which the end-caps were stacked over an
A•T base-pair and a second sequence, V, in which the end-
caps were stacked over a C•G base-pair. With two of the end-
caps, 14 and 18, we included additional sequences in which
the end-caps were placed over T•A (I and III) and G•C base
pairs (II).
The Tm values of the control sequences i-v (Table 1, no end-
caps) were not determined. These sequences contain only two
C•G (G•C) base pairs and consequently would be expected to
melt well below room temperature. Complementary 4-mers
containing all C•G (G•C) base pairs have been reported to have

New Spacers for Use as dsDNA End-Caps
Bioconjugate Chem., Vol. 21, No. 8, 2010 1551
Table 1. Calculated Free Energy of Change for Association of
Complementary Unmodified Tetramersa
Table 3. Tm Values of Oligonucleotides IV and V Containing 2, 8,
22, 29, 32-34, and TTTTa
DNA sequences (5′f3′)
-∆G (kcal/mol)
end-cap Tm values (°C)
i. GCAT/ATGC
ii. TACG/CGTA
iii. CGAT/ATCG
iv. GCTA/TAGC
v. ATGC/GCAT
3.59
3.16
3.37
3.12
3.54
oligonucleotide
2
8
22 25 29 32 33 34 TTTT
IV
V
66 39 46 50 48 62 53 60
62 42 56 61 56 75 66 74
39
58
a All DNA sequences are written in 5′f3′ direction. X: end-cap.
a All DNA sequences are written in the 5′f3′ direction. The free
energy factor for end effects was added only for the end opposite that
covalently linked by end-caps (see oligonucleotides I-V.
distances of 14.5 Å in a minimized structure in which the two
terminal C-C bonds are gauche. Linker 8 (14.6 Å fully
extended) spans only 11.4 Å in a minimized gauche conforma-
tion, while the longest linker, 22, which is 19.5 Å fully extended,
falls to 17.9 Å. The X-ray crystal structure for bis(2-hydroxy-
ethyl)-stilbene-4,4′-diether linker capping a six-base-pair stem
(13) clearly shows that the terminal -OCH2CH2O- exist in the
gauche form. In modeling studies of the bridged duplexes, the
3′-O to 5′-O distances were found to be 11.3, 14.7, 17.1, and
19.9 Å, for 8, 14, 18, and 22, respectively. These models are
interesting because each linker shows its own particular pattern
of conformational preferences, but the gauche form is noticeably
absent from the carbon bonds adjacent to the phosphodiester
group. In each of the structures, the 3′-endo (characteristic of
B-DNA) conformation is maintained in all deoxyribose residues
including those linked to the end-cap.
Comparison to the hexaethylene glycol end-cap (25) is
instructive. This spacer is far longer (21.4 Å) but contains
-OCH2CH2O- bonds for which the gauche conformation is
favorable. The corresponding Tm values for oligonucleotides IV
and V are higher (50 and 61 °C) than for any of the amide
containing end-caps. As a consequence, it is not possible to say
whether or not the amide groups provide any degree of
stabilization through direct interactions with the face of the
terminal base pair. It may be that the penalty for adopting higher
energy conformations in order to fit between the 3′-O and 5′-O
base pairs negates any advantage of amide-heterocycle interac-
tions. Finally, we modified linker 22 by replacing two carbon
atoms by oxygen atoms to create an analogue in which the two
terminals now contain the -OCH2CH2O- group (linker 29).
Presumably, this analogue, like 25 could more readily adopt a
gauche conformation. In fact, the resulting Tm values (Table 3)
were almost identical to those measured for compound 22. It is
possible that the increased flexibility was compromised by too
short of a spacer distance with both terminal -OCH2CH2O-
groups in a gauche conformation.
Stabilization by end-caps is greatest when aromatic residues
capable of stacking interactions are included as part of the
structure (3, 8, 13). For comparison, we have included two types
of aromatic moiety containing end-caps in these studies. The
naphthalene tetracarboxylic acid diimide linker was studied
previously by Bevers et al. (3); however, they used a
-OCH2CH2OCH2CH2- spacer between the phosphodiester groups
and the imide nitrogen atoms, whereas we used -(CH2)3-,
-(CH2)4-, and -(CH2)5- spacers (32, 33, and 34, respectively) in
our studies. Both Bevers et al. with the naphthalendicarboxamide
linkers and Lewis and co-workers (9) in the case of the stilbene
linkers included -OCH2CH2O- spacers as part of the structure.
Interestingly, the lack of this spacer in our linkers does not
appear to have compromised stability in all cases. One of these
linkers (34) is nearly identical to the Bevers et al. linker. The
difference is that we used OCH2CH2CH2CH2CH2- in place of
-OCH2CH2OCH2CH2-. Also, we measured Tm values in much
shorter oligonucleotides sequences (ATGC-34-GCAT, Tm ) 74°
and GCTA-34-TAGC, Tm ) 60° versus TCTTTTCTT-Y-
AGAAAAGAA, Tm ) 72°). Despite the shorter oligonucle-
otides, we obtained a higher Tm value when the naphthalene
tetracarboxylate diimide was stacked over the C•G base pair.
Our observed Tm value is significantly lower when the naph-
Table 2. Tm Values of Oligonucleotides I-V Containing 14 and 18a
end-cap Tm values (°C)
oligonucleotide
14
18
I GCATXATGC
II TACGXCGTA
III CGATXATCG
IV GCTAXTAGC
V ATGCXGCAT
48
45
44
41
51
53
50
48
44
56
a All DNA sequences are written in 5′f3′ direction. X: end-cap.
Tm values ranging from 16.7 to 27.8 °C. For comparison of
relative stability, we have tabulated the predicted free energies
of association for complementary sequences i-v (Table 1),
based on the unified parameters reported by SantaLucia (19).
For purposes of comparison, the free energy contribution from
the end effect was included only for the corresponding uncapped
end (compare to sequences in Table 2). From the calculated
free energies of association, we might predict the relative order
of the Tm values of the DNA duplexes of Table 1 to be i g v
> iii > ii g iv. Deviation from this order in the corresponding
end-cap containing oligonucleotides would indicate base-specific
association effects with the end-cap.
The Tm values observed for sequences I-V containing end-
caps 14 or 18 (Table 2) do not completely correlate with the
predicted trend of the Tm values of unmodified DNA duplexes
based on the calculated ∆G values in Table 1. The most stable
duplex is V, in which the end-caps are positioned over a C•G
base pair. Oligonucleotide I would be predicted to be slightly
more stable based on the calculated free energies for association.
Furthermore, the Tm values for oligonucleotide II containing
either end-cap 14 or 18 are higher than the Tm values for
oligonucleotide III, which is contrary to prediction. These results
suggest that there is a greater stabilization effect when the end-
caps are adjacent to either a G•C or a C•G base pair. This
observation is concordant with results reported by Benight and
co-workers (20) from a study on hairpin DNA oligonucleotides.
Stems with C•G base pairs closing the loop show greater
stability than stems in which the loop closing base pair is A•T.
Because of the number of different end-caps, we wished to
investigate, other end-caps were incorporated only into se-
quences IV and V. This was sufficient to probe differences
between the end-caps for stabilization of closing A•T versus
C•G base pairs. As can be seen from the data, all of the end-
caps stabilize the four-base-pair duplex, but the differences in
stabilization are significant. Among the diamide aliphatic linkers,
the degree of stabilization is related to the length. The shortest
linker (8) is least stabilizing with Tm values for IV and V at 39
and 42 °C, while the longest (22) is most stabilizing, with Tm
values for IV and V at 46 and 56 °C. Although 14, 18, and 22,
when fully extended, are all longer than the average distance
of 16.2 Å observed for the 3′-O to 5′-O span between duplex
ends in B-DNA, if the carbon-carbon bonds adjacent to these
oxygens are in a gauche conformation rather than an anti-
periplanar conformation, the lengths shorten by 1.6 to 2.9 Å.
For example, linkers 14 and 18, which are both near the
optimum length at 16.8 and 17.3 Å, respectively, both show

1552 Bioconjugate Chem., Vol. 21, No. 8, 2010
Ng et al.
Figure 2. Oligonucleotides VII and VIII do not form a stable hybrid
duplex with oligonucleotide VI. Combination of the same nucleotide
sequences with end-cap 18 linking VIII and VII to VI gives duplex
IX, which can be cocrystallized with Endo V to give the structure shown
in Figure 3.
Figure 3. Structure of oligonucleotide IX bound to Endo V (protein
not shown). (A) Complete structure. (B) Details of end-cap 18.
thalene was stacked over the A•T base pair. In the Bevers
sequence, the naphthalene is adjacent to a T•A base pair. It is
likely that the Bevers linker would have caused a similar
increase in stabilization when placed over a G•C or C•G base
pair. Nevertheless, based on our high Tm values it does not
appear likely that the absence of an -OCH2CH2O-spacer in our
linker significantly compromised the stability. Furthermore, the
C3 spacer (in linker 32), which would presumably be far too
short based on our earlier analysis (16.0 Å fully extended), is
the most stabilizing.
The hydrophobic terthiophene end-cap 2 stabilized oligo-
nucleotide IV (Tm ) 66 °C) to a greater extent than oligonucle-
otide V (Tm ) 62 °C). This difference is particulary profound
because nonbridged duplex V is considerably more stable than
duplex IV (Table 1). This presumably results from the greater
hydrophobicity of the A•T base pair versus the C•G base pair.
nucleotide. Details of the Endo V structure and its interaction
with sequence IX will be reported in a separate manuscript.
ACKNOWLEDGMENT
This work was supported by NASA under award NCC
2-1363, a grant from the National Institutes of Health
(1R21EB006308), the National Science Foundation (0554990),
the Indiana Elks Foundation, and Walther Cancer Institute.
Assistance from the National Cancer Institute Grant (P30
CA23168) awarded to Purdue University is also gratefully
acknowledged. A Purdue Doctoral Fellowship provided support
for Brian Laing.
Supporting Information Available: 1H, 13C, and 31P NMR
spectra of synthesized compounds; oligonucleotide MALDI-
TOFMS characterization data. This material is available free
of charge via the Internet at http://pubs.acs.org.
CONCLUSIONS
This study confirms that end-caps containing conjugated
aromatic π-systems, capable of base stacking, stabilize a short
oligonucleotide duplex significantly more than end-caps con-
taining other types of functionality. Moreover, we have now
demonstrated that a highly hydrophobic end-cap (2) enhances
the stability of a duplex to a greater extent when positioned
over an A•T base pair than when stacked over a C•G base pair.
The more hydrophilic naphthalene tetracarboxylate diimide
stabilizes a duplex to a much greater extent when stacked over
a C•G base pair.
The significantly greater stabilization of the four-base-pair
stems from a hexaethylene glycol in comparison to the four
diamide containing aliphatic end-caps included in this study
suggests that greater spacer flexibility may allow the terminal
base pair to achieve optimal conformation. Our results further
support the contribution of π-stacking interactions in duplex
stabilization by end-caps, as well as the importance of spacer
structure linking the aromatic residue to the oligonucleotides
phosphodiester groups.
An important goal of this research was to find spacers that
would not alter the duplex in a way that would either hinder
binding or prevent readjustment of the structure as dictated by
a protein binding to the dsDNA. On the basis of the Tm studies
and taking into account spacer length, spacer 18 was incorpo-
rated into sequence IX (Figure 2). The spacer made it possible
to create a single sequence (IX) that would take the place of
three separate oligonucleotides (GCTAACC (VI), GGTIA (VII),
and GC (VIII) required for the co-crystallization of the enzyme
Endo V with its substrate, a nicked DNA. Sequences VII and
VIII do not hybridize to the heptamer VI, because VIII is too
short (dimer) and VII contains an inosine mismatch. However,
as illustrated in Figure 3, the sequence is able to assume a duplex
structure that in the presence of Endo V can flip out the inosine
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