Synthesis and characterization of conformationally preorganized, (R)-diethylene glycol-containing γ-peptide nucleic acids with superior hybridization properties and water solubility

Article abstract of DOI:10.1021/jo200482d

Developed in the early 1990s, peptide nucleic acid (PNA) has emerged as a promising class of nucleic acid mimic because of its strong binding affinity and sequence selectivity toward DNA and RNA and resistance to enzymatic degradation by proteases and nucleases; however, the main drawbacks, as compared to other classes of oligonucleotides, are water solubility and biocompatibility. Herein we show that installation of a relatively small, hydrophilic (R)-diethylene glycol ("miniPEG", R-MP) unit at the γ-backbone transforms a randomly folded PNA into a right-handed helix. Synthesis of optically pure ^R-MPγPNA monomers is described, which can be accomplished in a few simple steps from a commercially available and relatively cheap Boc-l-serine. Once synthesized, ^R-MPγPNA oligomers are preorganized into a right-handed helix, hybridize to DNA and RNA with greater affinity and sequence selectivity, and are more water soluble and less aggregating than the parental PNA oligomers. The results presented herein have important implications for the future design and application of PNA in biology, biotechnology, and medicine, as well as in other disciplines, including drug discovery and molecular engineering.

Full text of DOI:10.1021/jo200482d

ARTICLE
pubs.acs.org/joc
Synthesis and Characterization of Conformationally Preorganized,
(R)-Diethylene Glycol-Containing γ-Peptide Nucleic Acids with
Superior Hybridization Properties and Water Solubility
Bichismita Sahu, Iulia Sacui, Srinivas Rapireddy, Kimberly J. Zanotti, Raman Bahal, Bruce A. Armitage,
and Danith H. Ly*
Department of Chemistry and Center for Nucleic Acids Science and Technology (CNAST), Carnegie Mellon University,
4400 Fifth Avenue, Pittsburgh, Pennsylvania 15213, United States
S Supporting Information
b
ABSTRACT: Developed in the early 1990s, peptide nucleic acid (PNA) has
emerged as a promising class of nucleic acid mimic because of its strong binding
affinity and sequence selectivity toward DNA and RNA and resistance to
enzymatic degradation by proteases and nucleases; however, the main draw-
backs, as compared to other classes of oligonucleotides, are water solubility and
biocompatibility. Herein we show that installation of a relatively small, hydro-
philic (R)-diethylene glycol (“miniPEG”, R-MP) unit at the γ-backbone trans-
forms a randomly folded PNA into a right-handed helix. Synthesis of optically
pure R-MPγPNA monomers is described, which can be accomplished in a few
simple steps from a commercially available and relatively cheap Boc-L-serine.
Once synthesized, R-MPγPNA oligomers are preorganized into a right-handed
helix, hybridize to DNA and RNA with greater affinity and sequence selectivity, and are more water soluble and less aggregating than
the parental PNA oligomers. The results presented herein have important implications for the future design and application of PNA
in biology, biotechnology, and medicine, as well as in other disciplines, including drug discovery and molecular engineering.
’ INTRODUCTION
an attractive reagent for many applications in biology, biotech-
nology, and medicine.16,17
Peptide nucleic acid (PNA) is a promising class of nucleic acid
mimic developed in the past two decades in which the naturally
occurring sugar phosphodiester backbone is replaced with N-(2-
aminoethyl)glycine units (Chart 1).1 PNA can hybridize to cDNA
or RNA just as the natural counterpart, in accordance with the
WatsonꢀCrick base-pairing rules, but with higher affinity and
sequence selectivity.2 Furthermore, PNA can invade selected
sequences of double-stranded DNA (dsDNA).3ꢀ5 The improve-
ment in thermodynamic stability has been attributed, in part, to
the lack of electrostatic repulsion in the backbone.1 The other
contribution may come from counterion release upon hybridiza-
tion, as opposed to condensation taking place with DNA and
RNA, resulting in an increase in the overall entropy of the
system.6 The enhancement in sequence selectivity, however, is
less well understood.7 Structural studies suggested that hydration
may play a key role in rigidifying the backbone of PNA upon
hybridization to DNA or RNA (or PNA), making it less accom-
modating to structural mismatches. Structural or spinal water
molecules have been observed in the X-ray structures of PNAꢀ
DNA8 and PNAꢀPNA9ꢀ11 duplexes bridging the amide proton
in the backbone to the adjacent nucleobase. Besides hybridiza-
tion properties, another appealing aspect of PNA is enzymatic
stability. Because of its unnatural backbone, PNA is not easily
degraded by proteases or nucleases.12 These properties, along
with the ease13,14 and flexibility15 of synthesis, together make PNA
PNA has so far been employed in a number of applications,
from molecular tools for probing and manipulating nucleic acid
structures and functions18ꢀ21 and regulation of gene expression22,23
to the recognition code for tagging molecules in drug discovery,24,25
amplifying genetic information in molecular evolution,26,27
and organizing molecular self-assembly in materials science and
nanotechnology.28ꢀ32 In spite of its many appealing features,
however, PNA has one major setback as compared to other
classes of oligonucleotides. Because of the charge-neutral back-
bone, PNA is only moderately soluble in aqueous solution.
Furthermore, it has a tendency to aggregate and adhere to
surfaces and other macromolecules in a nonspecific manner.33,34
This inherent property posts a considerable technical challenge
for the handling and processing of PNA—for instance, in the
development of microarrays and related surface-bound applica-
tions, because of its propensity to aggregate and collapse onto the
surface, causing poor and nonspecific binding,35,36 and in biology
and medicine, in part because of the concerns for off-target
binding and cytotoxicity.34
In attempts to address this concern, several approaches have
been taken, including incorporation of charged amino acid
Received: March 18, 2011
Published: May 27, 2011
r
2011 American Chemical Society
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Chart 1. Chemical Structures of DNA (RNA), PNA, and
Scheme 1. Synthesis of R-MPγPNA Monomersa
MP-Containing γPNA Units
residues, such as lysine at the termini37 or in the interior part of
the oligomer,38,39 inclusion of polar groups in the backbone,40
carboxymethylene bridge,41 and nucleobases,25,42 replacement
of the original (aminoethyl)glycyl backbone skeleton with a nega-
tively charged scaffold,43,44 conjugation of high molecular weight
polyethylene glycol (PEG) to one of the termini,36,45 fusion of
PNA to DNA to generate a chimeric oligomer,46ꢀ50 and redesign
of the backbone architecture.51 These chemical modifications
have led to improvement in solubility, but it is often achieved at
the expense of binding affinity and/or sequence specificity, not to
mention the requirement of an elaborate synthetic scheme in
some cases. Incorporation of cationic residues can improve water
solubility, but it can also lead to nonspecific binding due to an
increase in chargeꢀcharge interaction upon hybridization to
DNA or RNA. Conjugation of PNA to DNA (or RNA) can im-
prove water solubility as well, but it can compromise binding
affinity due to an increase in chargeꢀcharge repulsion and
conformational heterogeneity in the backbone.46,47,49 Herein
we report the development of a new class of conformationally
preorganized, diethylene glycol-containing γPNAs that exhibit
superior hybridization properties and water solubility as com-
pared to the original PNA design or any other chiral γPNA that
has been developed to date.
a Reagents and conditions: (a) NaH, BrCH2CH2OCH2CH2OCH3,
DMF, 0 °C, 68%; (b) isobutyl chloroformate, NaBH4, NMM, DME,
0 °C f rt, 84%; (c) DIAD, [(2,4-dinitrophenyl)sulfonyl]glycine ethyl
ester, TPP, THF, 0 °C f rt, 56%; (d) n-propylamine, CH2Cl2, rt, 81%;
(e) carboxymethylene nucleobase, DCC, DhbtOH, DMF, 50 °C,
67ꢀ85%; (f) 2 M NaOH/THF (1:1), 0 °C, 85ꢀ98%.
mainly on synthesis74ꢀ77 and covalent attachment of PNA to
other functional groups or molecular entities.78ꢀ80
’ RESULTS AND DISCUSSION
The helical sense, i.e., whether it adopts a right-handed or left-
handed helix, is determined by the stereochemistry. γPNAs pre-
pared from L-amino acids adopt a right-handed helix, while those
prepared from D-amino acids adopt a left-handed helix; however,
only the right-handed helical γPNAs hybridize to DNA and RNA
with high affinity and sequence selectivity. Furthermore,
we showed that the fully modified, L-alanine-derived γPNAs
Design Rationale. Diethylene glycol, commonly referred to as
“miniPEG” or MP, was chosen as a chemical moiety for
incorporation in the backbone of PNA because of its relatively
small size and hydrophilic nature and the fact it has been shown
to be nontoxic and nonimmunogenic.52ꢀ54 MP and larger mol-
ecular weight PEG have been incorporated into a number of
macromolecular systems, including peptides and proteins,55
nucleic acids,45,56ꢀ60 carbohydrates,61 synthetic polymers,62
dendrimers,63 liposomes,64,65 nanoparticles,66,67 and self-assembled
materials,68 just to name a few, in attempts to improve their
aqueous solubility, enzymatic stability, bioavailability, and phar-
macokinetics as well as to minimize their immunogenicity.
Incorporation of MP into the backbone of PNA is expected to
confer similar beneficial effects, including improvements in water
solubility and biocompatibility, along with reduction in aggrega-
tion and nonspecific binding. Inspection of the PNA backbone
reveals three obvious sites for incorporation of this chemical
moiety—R, β, and γ (Chart 1). We selected the γ-position
because prior studies from our group69ꢀ73 have revealed that
installation of a chiral center at this position induces helical
organization in the oligomer, providing an additional means for
fine-tuning the thermodynamic stability of PNA. γPNAs have
been made by other groups before, but their emphasis has been
(
S-AlaγPNAs) can invade mixed-sequence double-helical B-form
DNA (B-DNA).81 Though they are promising as antisense
and antigene reagents, S-AlaγPNAs are poorly soluble in water
and have a tendency to aggregate, presumably due to the charge-
neutral backbone and hydrophobic character of the methyl group
at the γ-backbone position. We surmised that replacement of the
methylgroupwithMPmight ameliorate someofthese issues while
maintaining the superior hybridization properties of the original
γPNA design because of the helical preorganization in the back-
bone upon installation of MP at the γ-position (Chart 1).
Monomer Synthesis. Boc-protected R-MPγPNA monomers
containing all four natural nucleobases (A, C, G, T) were
synthesized according to the procedures outlined in Scheme 1.
Alkylation of Boc-protected L-serine (1) with 1-bromo-2-(2-
methoxyethoxy)ethane under an optimized reaction condition:
slow addition of 1 into a vigorously stirred, chilled solution of
DMF containing 2 equiv of sodium hydride, followed by addition
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Scheme 2. Chemical Derivatization for 19F NMR Analysis
of 1-bromo-2-(2-methoxyethoxy)ethane afforded compound 2with
high optical purity (see the section below). The stoichiometry
and order of addition were determined to be essential for ob-
taining optically pure product. Slow addition of sodium hydride
was necessary to ensure complete deprotonation of the carboxyl
group prior to removal of the hydroxyl proton. Formation of the
carboxylate anion reduces the acidity of the R-proton, making it
less susceptible to deprotonation by base. Esterification of the
alkylated product 2 followed by reduction with sodium borohy-
dride yielded serinol 3. This chemical transformation, conversion
of the carboxylic acid to alcohol, renders the R-proton inert to
deprotonation and racemization in the subsequent reaction
steps. Coupling 3 to N-[(2,4-dinitrophenyl)sulfonyl]glycine ethyl
acetate, which was prepared according to the published procedure,82
employing the Mitsunobu reaction followed by removal of the
(2,4-dinitrophenyl)sulfonyl group with a mild base yielded
compound 5. DCC-mediated coupling of 5 with the appropriate
carboxymethylene nucleobases (A,83 C,14 G,83 T13) and hydro-
lysis of the resulting esters gave the desired monomers 7aꢀd.
Determination of Optical Purities. The optical purities of
key intermediates and final monomers were determined by 19F
NMR following chemical derivatization (Scheme 2). Corradini
and co-workers84 have previously shown that the enantiomeric
excess (ee) of chiral R-PNA monomers and oligomers, with the
latter obtained following acid hydrolysis, can be determined by
GC/MS following chemical derivatization. We have discovered
that 19F NMR is a convenient and accurate alternative method
for determining the ee values for γPNA monomers and prior
intermediates following removal of the Boc protecting group and
subsequent coupling with Mosher’s reagent, (+)-1-methoxy-
1-(trifluoromethyl)phenylacetyl chloride (MTPA-Cl). Figure 1
shows representative spectra of MTPA-derivatized MP-L-serine
2 (compound 8), MP-L-serinol 3 (compound 9), and thymine
R-MPγPNA monomer 7a (compound 10). The corresponding dia-
stereomers were prepared the same way starting from Boc-D-
serine and are included for comparison. A close inspection of
these spectra reveals no trace quantity of the other diastereomers
for the alkylated serine 8, serinol 9, or final monomer 7a,
indicating that they were optically pure. If racemization were to
occur, we would expect an additional peak in the spectrum, as
indicated by the arrow, corresponding to the chemical shift of the
other diastereomer. The ee values for serine 8, serinol 9, and final
thymine monomer 7a were estimated to be at least 99%, within
the detection limit of 19F NMR.85 At this point it is not clear why
Figure 1. 19F NMR spectra of MTPA-derived (A) MP-L-serine 2
(compound 8), (B) MP-L-serinol 3 (compound 9), and (C) thymine
monomer 7a (compound 10). The corresponding diastereomers were
prepared under identical conditions starting from Boc-D-serine and are
included for comparison.
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Table 1. Sequence of PNA Oligomersa
oligomer
sequence
no. of MP units
PNA1
PNA2
PNA3
PNA4
PNA5
PNA6
PNA7
PNA8
PNA9
PNA10
PNA1X
PNA1Y
PNA4X
PNA4Y
H-GCATGTTTGA-NH2
0
1
H-GCATGTTTGA-NH2
H-GCATGTTTGA-NH2
3
H-GCATGTTTGA-NH2
5
H-GCATGTTTGA-NH2
10
0
H-ACGGGTAGAATAACAT-NH2
H-ACGGGTAGAATAACAT-NH2
H-ACGGGTAGAATAACAT-NH2
H-ACGGGTAGAATAACAT-NH2
H-ACGGGTAGAATAACAT-NH2
H-LOrn(X)-LLys-GCATGTTTGA-NH2
H-LLys-GCATGTTTGA-LOrn(Y)-NH2
H-LOrn(X)-LLys-GCATGTTTGA-NH2
H-LLys-GCATGTTTGA-LOrn(Y)-NH2
1
3
5
8
0
0
5
5
a Underlined letters indicate R-MP γ-backbone modification. X =
fluorescein (FITC), and Y = tetramethylrhodamine (TAMRA).
the thymine R-MPγPNA monomer showed two rotamers (Figure 1C),
at ꢀ68.80 and ꢀ68.95 ppm, while thymine S-MPγPNA showed
only one (Figure 1C, inset). (The existence of the two rotamers
was determined by multinuclear and multidimensional NMR
analyses as described in an earlier work;73 they differ from one
another in the rotation around the C7ꢀN4 tertiary amide bond.)
We attribute the high optical purities of the intermediates and
final monomers (the others not shown) to the optimized, initial
alkylation step and the stereoselective conversion of carboxylic
acid to alcohol in the second step, on the basis of the procedure
reported by Rodriguez and co-workers.86
Oligomer Design and Synthesis. Three sets of PNA oligo-
mers were prepared (Table 1). The first set, comprising PNA1ꢀ5,
was designed to test the effects of MP on the conformation and
hybridization properties of PNA. The second set, consisting of
PNA6ꢀ10, was designed to test the effect of MP on water
solubility. The hexadecameric sequence was chosen for this study
because it represents a statistical length that would be required to
target a unique site within the mammalian genome or transcrip-
tome. The third set, constituting PNA1 and PNA4, each sepa-
rately linked to fluorescein (FITC) at the N-terminus (PNA1X
and PNA4X) and tetramethylrhodamine (TAMRA) at the C-
terminus (PNA1Y and PNA4), was designed to test the effect of
MP on self-aggregation on the basis of F€orster resonance energy
transfer (FRET). Inclusion of a lysine residue at the C-terminus
was necessary, since prior attempts to synthesize FITC- and
TAMRA-containing PNA oligomers without the lysine residue
resulted in sticky and poorly water soluble materials that were
difficult to purify and characterize.
All PNA oligomers, those with and without MP side chains, were
synthesized on a solid support according to the published procedures
of Christensen and co-workers .87 Unlike modifications made at the
R-backbone, which require further optimization of the reaction
condition to minimize racemization during the on-resin coupling,88
no precaution was necessary for coupling of R-MPγPNA monomers
on the resin. Upon completion of the last monomer coupling, the
oligomers were cleaved from the resin and precipitated with ethyl
ether. After being air-dried, the crude pellets were dissolved in a
water/acetonitrile mixture (80:20), purified by reversed-phase
HPLC, and characterized by MALDI-TOF mass spectrometry.
Figure 2. CD spectra of (A) the unhybridized (single-stranded) PNA
oligomers and (B) the corresponding PNAꢀDNA and PNAꢀRNA
(inset) hybrid duplexes at 5 μM strand concentration each. (A) Inset:
UV absorption spectra of the individual PNA oligomers at 90 °C,
showing that they had the same concentration. This indicates that the
differences in the CD spectra, both the amplitude and profile, are the
intrinsic properties of the oligomer themselves and not the variations in
concentrations. The samples were prepared in 10 mM sodium phos-
phate buffer at pH 7.4, and the CD spectra were recorded at room
temperature. The cDNA strand used was 50-TCAAACATGC-30.
Conformational Analysis. To determine the effect of MP on
the conformation of PNA, we measured the CD spectra of
PNA1ꢀ5. Consistent with the earlier finding,69 we did not
observe noticeable CD signals in the nucleobase absorption re-
gions for PNA1 (Figure 2A). This indicates one of two possibi-
lities: either that the PNA does not adopt a helical conformation
or that it does but with an equal proportion of a right-handed and
left-handed helix. The latter was suggested by MD simulations,89
but ruled out on the basis of NMR analyses.69 However, in the
case of PNA2ꢀ5, we noticed distinct exciton coupling patterns,
with minima at 242 and 280 nm and maxima at 220 and 260 nm,
characteristic of a right-handed helix.90 The amplitude of the CD
signals remained fairly constant as more MP units were added,
but the profiles slightly shifted toward that of the PNAꢀDNA2
and PNAꢀRNA2 double helices (Figure 2B), especially at the
242 and 280 nm minima. A gradual dip at the 242 nm minimum
generally indicates a tightening in the helical pitch of the
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Figure 3. (A) CD spectra of PNA5 and PNA2 (inset) as a function of
temperature. (B) Melting transition (Tm) of PNA2ꢀ5 as determined by
CD, monitored at 260 nm. The oligomer concentration was 5 μM,
prepared in 10 mM sodium phosphate buffer at pH 7.4.
Figure 4. UV melting profiles of (A) PNAꢀDNA and (B) PNAꢀRNA
hybrid duplexes at 5 μM strand concentration each in 10 mM sodium
phosphate buffer at pH 7.4. Both the heating and cooling runs were
performed; they both had nearly identical profiles (only the heating runs
are shown).
oligomer,91 from one that resembles that of a PNAꢀPNA92
duplex with 18 base pairs per turn to one that resembles that of a
PNAꢀDNA93 duplex with 13 base pairs per turn. Overall, the
CD profiles of PNA2ꢀ5 are similar to those of the corresponding
PNAꢀDNA and PNAꢀRNA hybrid duplexes (Figure 2B). The
major difference however is in the amplitude, which is roughly
doubled for the duplex as compared to the individual strand. This
is because the concentration of bases in the hybrid duplex is twice
that of the individual PNA strand. Taken together, these results
show that a single (R)-MP unit installed at the γ-backbone is
sufficient to preorganize PNA into a right-handed helix. Incor-
poration of additional MP units did not further improve base-
stacking, apparent from the similarities in the CD amplitudes, but
it did help to tighten the helical pitch of the oligomers, making
them more rigid and compact. This is apparent from the
temperature-dependent CD measurements, which showed less
dramatic reduction in the signal amplitude with temperature for
PNA5 as compared to that of PNA2 (Figure 3A). Even at a
temperature as high as 80 °C, a distinct CD profile still remained,
indicating that base-stacking still occurred at this temperature—
whereas it was completely denatured in the case of PNA2.
Overall the stability of the oligomers increases linearly with the
number of MP units incorporated (Figure 3B). The fact that
PNA5 adopts a helical conformation most closely resembling
that of PNAꢀDNA and PNAꢀRNA duplexes suggests that it
would hybridize to DNA and RNA more effectively than the
other oligomers in this series.
Thermal Stability. UV melting experiments were performed
to determine the effect of MP on the thermal stability of PNA
upon hybridization to DNA and RNA. Our result shows that
incorporation of a single MP side chain stabilized a PNAꢀDNA
duplex by 4 °C (Figure 4A). The extent of stabilization gradually
increased with additional MP units added but tapered off to
∼2.3 °C per unit for the fully modified PNA5. A similar pattern
was observed for RNA, but the extent of stabilization was less
compared to that of DNA (Figure 4B). The enhancement was
only 3 °C for the first MP unit incorporated and 1.2 °C per unit
for PNA5, as compared to 2.3 °C per unit with DNA. It is
interesting to note that the unmodified PNA1 binds more tightly
to RNA than to DNA, with a differential Tm (ΔTm) of 10 °C.
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Table 2. Thermodynamic Parameters for PNAꢀDNA and PNAꢀRNA Duplexes
PNAꢀDNAa
PNAꢀRNAb
oligomer ꢀΔH° (kJ/mol) ꢀTΔS° (kJ/mol) ꢀΔG° (kJ/mol)
Kd
ꢀΔH° (kJ/mol) ꢀTΔS° (kJ/mol) ꢀΔG° (kJ/mol)
Kd
PNA1
PNA2
PNA3
PNA4
PNA5
273 ( 5
319 ( 18
316 ( 11
329 ( 14
372 ( 11
224 ( 5
263 ( 16
256 ( 11
265 ( 12
294 ( 10
49 ( 1c
54 ( 1
59 ( 1c
65 ( 1
78 ( 2
2.5 ꢁ 10ꢀ9
3.2 ꢁ 10ꢀ10
5.1 ꢁ 10ꢀ11
3.5 ꢁ 10ꢀ12
4.6 ꢁ 10ꢀ14
289
333
350
356
365
229
232
280
283
287
60
68
71
73
78
3.5 ꢁ 10ꢀ11
1.2 ꢁ 10ꢀ12
4.3 ꢁ 10ꢀ13
1.7 ꢁ 10ꢀ13
2.1 ꢁ 10ꢀ14
a The averages of three trials (two from concentration dependence measurements and one from UV melting curve fitting). b UV melting curve fitting.
c The standard deviation is less than 1 kJ/mol. The temperature was 298 K.
Figure 6. SPR sensorgrams (solid black lines) and fits (red dotted lines)
for hybridization of PNA probes to immobilized cDNA. Solutions
contained 30 nM PNA. (Sensorgrams and kinetic fits for all PNA
concentrations from 10 to 50 nM are provided in the Supporting
Information.) Error bars at t = 420 s illustrate standard deviations for
Figure 5. Correlation between the Gibbs binding free energy (ΔG°)
and number of MP units.
three separate trials.
However, fully modified PNA5 displayed thermal stability iden-
tical with that of RNA as well as that of DNA. The lack of a
preferential binding for PNA5 is not clearly understood at this
point, but it may reflect the rigidity of the backbone. Because
PNA5 is more rigid and tightly wound as compared to PNA1, it
may not have the flexibility or conformational freedom to ac-
commodate the DNA and RNA template strands. Under such
circumstances, the DNA and RNA strands would have to under-
go a conformational change of their own to accommodate
the R-MPγPNA helix and for hybridization to take place. This may
explain why S-AlaγPNAꢀDNA prefers a P-form helix,73 a struc-
ture that is intermediate between the A- and B-form DNA, an
indication that DNA and RNA accommodate the γPNA ex-
igencies rather than the other way around. The RNA strand is less
accommodating to the R-MPγPNA, when it is fully modified,
because the RNA backbone is more rigid as compared to that of
DNA due to the presence of the chiral OH group at the 20-
position.94
Thermodynamic Analysis. To gain greater insight into the
contribution of MP to the stability of the PNAꢀDNA duplex, we
used van’t Hoff analysis to determine the thermodynamic para-
meters for PNA1ꢀ5 upon hybridization to cDNA and RNA
strands.95 The data are tabulated in Table 2. Our results show
that the Gibbs binding free energy (ΔG°) increases (to a rough
approximation) linearly with the number of MP units for PNAꢀ
DNA and follows a sigmoidal growth for PNAꢀRNA (Figure 5).
Incorporation of a single MP unit resulted in a net gain in binding
free energy of ∼5 kJ/mol for PNAꢀDNA, whereas it is less for
PNAꢀRNA—especially at the higher end. A reduction in the
equilibrium dissociation constant (Kd) by nearly 5 orders of mag-
nitude for PNA5ꢀDNA and 3 orders of magnitude for PNA5ꢀ
RNA was observed, as compared to that of the respective
PNA1ꢀNA and PNA1ꢀRNA duplexes. The binding free energy
gain in this case appears to be predominantly enthalpically driven
for both PNAꢀDNA and PNAꢀRNA, apparent from the grad-
ual increase in the ΔH° term with the number of MP units con-
comitant with modest enthalpyꢀentropy compensation.96
Kinetics of Hybridization and Dissociation. From a kinetic
perspective, the higher apparent affinity of the R-MPγPNA
oligomers indicated by the UV melting curves, from which the
equilibrium dissociation constants were extracted, could be due
to faster on rates and/or slower off rates. We used surface plasmon
resonance (SPR) analysis to study the hybridization kinetics. In
most other examples of PNAꢀDNA hybridization studied by
SPR, the PNA probe was immobilized to the chip while the DNA
target was captured from solution.97ꢀ99 One exception was pre-
vious work from our group, in which a DNA guanine quadruplex
target was immobilized and a homologous PNA probe hybri-
dized to form a PNAꢀDNA heteroquadruplex.100,101 In the
current studies, a biotinylated version of the DNA target was
immobilized on a streptavidin-conjugated, carboxymethylated
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Table 3. Association Rate Constant (ka), Dissociation Rate
Constant (kd), and Equilibrium Dissociation Constant (Kd)
for Hybridization of PNA Probes with a Complementary
DNA Target
Table 4. Sequence Mismatch Discriminationa
Tm (°C)
Tm (°C)
XꢀT
PNA1ꢀDNAb PNA5ꢀDNA PNA1ꢀRNAb PNA5ꢀRNA
oligomer
ka (Mꢀ1sꢀ1
)
kd (sꢀ1
)
Kd (M)
AꢀT
45
68
55
68
PNA1
PNA2
PNA3
PNA4
PNA5
4.7 ꢁ 105
9.7 ꢁ 105
6.2 ꢁ 105
6.6 ꢁ 105
8.0 ꢁ 105
13.0 ꢁ 10ꢀ4
4.1 ꢁ 10ꢀ4
1.9 ꢁ 10ꢀ4
0.3 ꢁ 10ꢀ4 a
0.4 ꢁ 10ꢀ4 a
2.8 ꢁ 10ꢀ9
C<>T
31 (ꢀ14)c
31 (ꢀ14)
35 (ꢀ10)
47 (ꢀ21)
48 (ꢀ20)
51 (ꢀ17)
37 (ꢀ18)
44 (ꢀ11)
40 (ꢀ15)
48 (ꢀ20)
52 (ꢀ16)
48 (ꢀ20)
4.2 ꢁ 10ꢀ10
3.0 ꢁ 10ꢀ10
4.1 ꢁ 10ꢀ11 a
5.4 ꢁ 10ꢀ11 a
G<>T
T(U)<>T
a Sequences: PNA1, H-GCATGTTTGA-LLys-NH2; PNA5, H-GCA-
TGTTTGA-LLys-NH2; DNA, 30-CGTACAXACT-5, X = A, C, G, T;
RNA, 30-CGUACAXACU-5, X = A, C, G, U. b The data for PNA1ꢀ
DNA and PNA1ꢀRNA mismatched binding were taken from ref 69.
c The value in parentheses indicates ΔTm between the perfect match and
mismatch.
a Uncertainty due to the calculated value approaching the limits of
detection of the instrument.
dextran chip. A relatively low surface density (ca. 100 response
units) of DNA target was used to limit mass transport effects on
the association kinetics. Solutions containing 10ꢀ50 nM PNA
were allowed to flow over the chip for 420 s, at which point the
flow was switched to a PNA-free buffer to allow net dissociation
of the hybridized PNA.
Table 5. Saturated Concentrations of PNA Oligomers
oligomer
no. of MP units
satd concn (mM)
PNA6
PNA7
PNA8
PNA9
PNA10
0
1
3
5
8
39
76
Individual sensorgrams for the unmodified (PNA1) and
R-MPγ-modified (PNA2ꢀ5) oligomers at 30 nM concentration
are shown in Figure 6. Relatively minor variation is observed in
the association kinetics, although singly modified PNA2 appears
to bind approximately twice as fast as the unmodified PNA.
Fitting the data to a 1:1 binding model yields association rate
constants (ka) that range from 4.7 ꢁ 105 to 9.7 ꢁ 105 Mꢀ1 sꢀ1
(Table 3). In contrast, significantly larger effects are seen in the
dissociation phase of the experiment, with the dissociation rate
constant (kd) varying by at least a factor of 50. Equilibrium
dissociation constants (Kd) calculated from the ratio of the dis-
sociation and association rate constants are also given in Table 3.
Unmodified PNA1 and fully modified PNA5 have Kd = 2.8 nM
and 54 pM, respectively. Note that the Kd values for PNA1ꢀ3
determined by SPR (Table 3) are similar to those determined by
UV melting experiments (Table 2). However, increasing diver-
gences are observed for PNA4 and PNA5, with the SPR-derived
values being 12- and 1200-fold greater, respectively. We attribute
this discrepancy to the very small degrees of dissociation
observed within the time scale of the SPR experiment. This in-
troduces a large uncertainty into the fitting of the dissociation
phases of the sensorgrams, translating into questionable Kd values.
However, we also note that the free energy changes given in Table 2
are based on the assumption that the heat capacity change for
PNAꢀDNA melting is zero, allowing the parameters determined at
the high melting temperature to be extrapolated to lower tempera-
tures. Future calorimetric measurements will allow us to determine
whether there is a heat capacity change in these hybridization/
melting transitions. Nevertheless, the SPR results clearly demon-
strate that the enhanced affinity of the R-MPγPNAs is due almost
entirely to the significantly slower dissociation kinetics. Thus, the
helical preorganization of the modified PNA does not translate into
significantly faster hybridization. This could be due to the fact that
the cDNA strand is likely to require some structural reorganization
of its own to hybridize to the PNA, negating the preorganization of
the latter. While the origin of the slower dissociation kinetics is not
clear at this time, one possible contributor is the network of
structured water molecules in the minor groove of the hybrid duplex,
which would need to be disrupted to separate the two strands.
Sequence Selectivity. On the basis of the CD, NMR,69 and
X-ray73 data which showed that γPNAs derived from L-amino
108
350
>500
acids adopt a right-handed helix and that the helix becomes more
rigid as more γ-chiral units are added in the backbone, one would
expect a fully modified PNA5 to hybridize to DNA and RNA
targets with greater sequence selectivity than PNA1. To verify
this prediction, we determined the thermal stabilities of PNA5ꢀ
DNA and PNA5ꢀRNA duplexes containing perfectly matched
(PM) and single-base-mismatched (MM) targets and compared
them to those from an earlier study with PNA1ꢀDNA and
PNA1ꢀRNA.69 Our results showed that, despite the strong bind-
ing affinity, PNA5 is able to discriminate between closely related
sequences. ΔTm ranges from ꢀ17 to ꢀ21 °C for PNA5ꢀDNA
and from ꢀ16 to ꢀ20 °C for PNA5ꢀRNA containing a single-
base mismatch (X = C, G, T), as compared to ꢀ10 to ꢀ14 °C for
PNA1ꢀDNA and ꢀ11 to ꢀ18 for PNA1ꢀRNA (Table 4). The
level of sequence discrimination is greater for PNA5ꢀDNA than
for PNA1ꢀDNA, and similar, if not slightly better, for PNA5ꢀ
RNA as compared to PNA1ꢀRNA. This result is consistent with
PNA5 adopting a more rigid helical motif, which is less accom-
modating to structural mismatches as compared to PNA1.
Water Solubility. To determine whether inclusion of MP in
the backbone has an effect on water solubility, we determined the
saturating concentrations of PNA6ꢀ10 (Table 1) in aqueous
solution using UV spectroscopy. It is interesting to note that
incorporation of a single MP unit enhanced the solubility of
PNA6 by nearly 2-fold (Table 5). The solubility of the oligomers
is further improved, albeit to a smaller extent, with additional MP
units. For PNA8, which contained eight MP units, we could not
determine the saturating concentration of this oligomer because
addition of a minimum amount of water (1 μL) into a vial con-
taining 2.7 mg of fluffy material resulted in complete dissolution.
On the basis of this result, we estimated the saturating concen-
tration to be at least 500 mM—more than an order of magnitude
higher than that of the unmodified PNA6. Solvation of the MP
side chain likely contributes to the enhanced solubility. The other
contributing factor may come from helical organization. Stacking
the nucleobases on top of one another in a helical arrangement
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Figure 8. Photographs of (A) PNA1XꢀPNA1Y and (B) PNA4XꢀP-
NA4Y samples at different temperatures upon excitation with a short-
wavelength (254 nm), hand-held UV lamp. The sample indicated as
“90 °C” was incubated in a heating block at 90 °C for 5 min prior to
placement on the UV lamp, after which the photographs were
immediately taken.
complexes and precipitation of the aggregates out of the
solution.
Self-Aggregation. Next we performed a FRET study to deter-
mine whether incorporation of MP into the backbone of PNA
would also help reduce aggregation. Different concentrations
of unmodified PNA1XꢀPNA1Y and homologous γ-modified
PNA4XꢀPNA4Y pairs (Table 1) were prepared by mixing
equimolar ratios of the individual oligomers in sodium phosphate
buffer. The samples were excited at 475 nm, the λmax of FITC,
and the emission was recorded from 480 to 700 nm. Upon
aggregation, in which the oligomers bearing FITC and TAMRA
come into contact with one another, excitation at 475 nm would
lead to energy transfer from FITC to TAMRA because of the
proximity of the two chromophores. Comparison of the FRET
efficiencies of the two systems at different concentrations should
provide an assessment of the effect of MP on the intermolecular
interaction of PNA.
Inspection of Figure 7A reveals that, at a concentration as low
as 1 μM of each PNA oligomer, a small but noticeable emission
appeared at 580 nm, an indication of FRET and correspondingly
aggregation between PNA1X and PNA1Y. The extent of aggre-
gation was further intensified with increasing concentrations of
oligomers, apparent from the fluorescent intensity of TAMRA at
∼580 nm upon excitation of the FITC donor at 475 nm. In
contrast, at 20 μM, the point at which nearly 70% FRET
efficiency was observed for PNA1XꢀPNA1Y, only ca. 5% FRET
efficiency was observed for PNA4XꢀPNA4Y (Figure 7B). This
indicates that the pair of γ-modified PNAs do not interact with
each other as much as the unmodified PNAs. The distinction is
apparent from photographs of the samples illuminated with a
short-wavelength (254 nm), hand-held UV lamp (Figure 8). The
PNA1XꢀPNA1Y solution displayed a light orange emission at
Figure 7. Fluorescent spectra of (A) PNA1XꢀPNA1Y and (B)
PNA4XꢀPNA4Y pairs at different concentrations. The samples were
prepared by mixing equimolar ratios of the oligomers in 10 mM sodium
phosphate buffer (pH 7.4). The samples were excited at 475 nm (FITC
λmax), and the emissions were recorded from 480 to 700 nm. The spectra
were normalized with respect to the FITC emission.
would shield the hydrophobic core from exposure to solvent. As
such, only the heteroatoms at the periphery of nucleobases are
exposed to and interact with the water molecules. This may
explain the nonlinear relationship between the number of MP
units and the saturating concentrations of PNA oligomers.
This suggestion was corroborated by additional experiments
(Figure S1, Supporting Information) which showed that in-
corporation of a single methyl group instead of MP at the γ-
backbone of PNA resulted in nearly 1.2-fold improvement in
water solubility, while the reversed trend was observed with
additional methyl groups. This is because addition of the alkyl
group beyond the first unit does not help to preorganize PNA
any more than it already has but instead introduces additional
hydrophobic character to the backbone. This would exacerbate
self-aggregation, leading to formation of large molecular weight
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helps reduce nonspecific binding with other macromolecules as
well—in this case DNA.
’ CONCLUSION
In summary, we have shown that a randomly folded PNA can
be preorganized into a right-handed helix by installing an R-MP
side chain at the γ-backbone. The MP-containing γPNA mono-
mers can be prepared in a few simple steps starting from a
commercially available and relatively cheap Boc-L-serine. We
attribute the high optical purities of the backbone intermediates
and final monomers to the optimized alkylation condition in the
first step and stereoselectivity of the reduction of carboxylic acid
to alcohol in the second step. In contrast to the (aminoethyl)-
glycine backbone prepared via the reductive amination route,
which degrades over time (unpublished data), compound 4 can
be stored over a prolonged period without signs of deterioration.
Another appealing feature of γ-backbone modification is that it is
configurationally stable. After the reduction step, i.e., conversion
of the carboxylic acid to alcohol, the γ-proton is inert to de-
protonation by base. Thus, there is no need for concern for
racemization in the subsequent reaction steps. Compared to the
previous attempts to improve water solubility, which is often
achieved at the expense of binding affinity and/or sequence
selectivity, incorporation of MP in the γ-backbone enhances
the hybridization properties as well as the water solubility of
PNA. Improvements in thermodynamics are attributed to slower
dissociation, while enhanced sequence selectivity is the result of
backbone preorganization and helical base-stacking arrange-
ment. Improvement in water solubility is the result of solvation
of the MP side chains as well as helical nucleobase arrangement.
Helical induction presumably permits greater shielding of the
hydrophobic cores of nucleobases and better hydrogen-bonding
interaction between heteroatoms at the periphery of nucleobases
and solvent molecules.
The results reported herein have important implications for
the future design and utility of PNA in biology, biotechnology,
and medicine. The improvements in hybridization properties may
enable R-MPγPNAs to invade double-helical DNA and structured
RNA, which may not be permissible with other classes of
oligonucleotide mimics. The former has already been demon-
strated in a recent study.81 The enhancements in water solubility
will facilitate the handling and processing of PNA while lessening
the concerns for nonspecific binding and cytotoxic effects.
Improvements in these areas, along with the flexibility of synth-
esis whereby other chemical functionalities can be installed at the
γ-backbone with ease, will further expand the utility of PNA
into other burgeoning disciplines, including drug discovery and
nanotechnology.
Figure 9. Result of a gel-shift assay following incubation of a 171 bp,
linear double-stranded DNA with different concentrations of PNA6 and
PNA10 in 10 mM sodium phosphate buffer (pH 7.4) at 37 °C for 16 h,
followed by electrophoretic separation on nondenaturing gel and SYBR-
Gold staining. Note that this particular DNA fragment, which was PCR
amplified from a Psuper plasmid vector, does not have a sequence
complementary to that of PNA6 or PNA10 (both contained the same
nucleobase sequence). The concentration of the 171 bp DNA fragment
was 0.4 μM.
room temperature and yellow-green hue at 90 °C, an indication
of the aggregate dissociating upon heating. In contrast, the PNA4Xꢀ
PNA4Y solution displayed the same color, yellow-green, at room
temperature as well at 90 °C, indicating that the oligomers were
well dispersed even at room temperature. Thus, the R-MP γ-
modification not only imparts enhanced solubility to PNA, but
also suppresses aggregation.
Off-Target Binding. It has been documented that, at moder-
ate concentrations, PNA tends to aggregate and stick to surfaces
and other macromolecules in a nonspecific manner.33 Such inter-
actions could lead to off-target binding and cytotoxic effects when
employed in the cellular context. Among the macromolecules
that PNA is known to interact nonspecifically with are nucleic
acids and proteins.34 To assess the extent of off-target binding of
PNA and R-MPγPNA, we performed a gel-shift assay. In this case
a DNA fragment (PCR product), 171 bp in length, was incubated
with different concentrations of PNA6 and PNA10 (Table 1) in
10 mM sodium phosphate buffer at 37 °C for 16 h. The two olig-
omers contained identical nucleobase sequences but differed from
another at the γ-backbone; PNA6 was unmodified, whereas
PNA10 was modified at every other position with an R-MP γ side
chain. Following incubation, the samples were separated on
nondenaturing polyacrylamide gel and stained with SYBR-Gold.
Since the target did not contain a sequence complementary to
that of the oligomers, we did not expect binding to take place, in
which case the intensity of the DNA band should remain fairly
constant, independent of the PNA6 and PNA10 concentrations.
Instead, we observed a drastic reduction in the intensity of the
DNA band with increasing concentrations of PNA6 (Figure 9).
At 10 μM (corresponding to a PNA:DNA ratio of 25:1) or higher,
the DNA band completely disappeared from the gel. We ob-
served neither a shifted band, indicating formation of a stable
complex, nor staining in the wells, which would indicate forma-
tion of globular complexes too large to traverse through the
polyacrylamide gel. One possible explanation for the disappear-
ance of the DNA band could be that, once formed, the PNAꢀ
DNA complex precipitated from solution and/or adhered to the
walls of the eppendorf tubes in which the samples were incu-
bated. On the other hand, for γ-modified PNA10, the intensity of
the DNA bands remained fairly constant even at a concentration
as high as 20 μM (PNA:DNA ratio of 50:1). This result is
consistent with the solubility and FRET data, indicating that
incorporation of MP at the γ-backbone not only improves the
hybridization properties and water solubility of PNA but also
’ EXPERIMENTAL SECTION
Monomer Synthesis. Boc-(2-(2-methoxyethoxy)ethyl)-L-serine
(2). To a stirred, ice-cold (0 °C) solution of NaH (60% suspended in
mineral oil, 1.7 g, 42.6 mmol) in dry DMF (80 mL) under an inert
atmosphere was added Boc-L-Ser-OH (4.05 g, 19.6 mmol) in DMF
(30 mL) dropwise over a period of 3 h. To this mixture, while still in
the ice bath, was added 1-bromo-2-(2-methoxyethoxy)ethane (6.4 mL,
42.6 mmol) at once. The ice bath was then removed, and the reaction
was allowed to gradually warm to room temperature and stirred for
another 3 h. The reaction mixture was quenched with H2O (100 mL) at
0 οC. The solvents (DMF and water) were evaporated under reduced
pressure at room temperature. Water (20 mL) was added to the crude
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residue and acidified with 5% HCl to pH ≈ 3 at 0 °C. The aqueous layers
were extracted with ethyl acetate (5 ꢁ 100 mL) and dried over Na2SO4.
The solvent was evaporated under reduced pressure, and the crude
mixture was purified by column chromatography to afford a colorless
liquid (4.1 g, 13.3 mmol): yield 68%; 1H NMR (CDCl3, 300 MHz) δ
6.84 (br s, 1H), 5.57 (d, J = 7.4 Hz, 1H), 4.46ꢀ4.38 (m, 1H), 3.93 (dd,
J1 = 3.9 Hz, J2 = 3.5 Hz, 1H), 3.90ꢀ3.49 (m, 9H), 3.37 (s, 3H), 1.41 (s,
9H); 13C NMR (CDCl3, 75 MHz) δ 28.2, 53.8, 58.7, 70.2, 70.7, 70.9,
71.7, 80.0, 155.8, 173.4; HRMS (ESI/MSm/z) M calcd for C13H25NO7-
Na 330.1529, found 330.1546.
Boc-(2-(2-methoxyethoxy)ethyl)-L-serinol (3). To a stirred solution
of Boc-(2-(2-methoxyethoxy)ethyl)-L-serine (2) (4 g, 13.0 mmol) in
20 mL of DME in an ice bath was added NMM (1.43 mL, 13.0 mmol)
dropwise under an inert atmosphere, followed by isobutyl chloroformate
(1.76 mL, 13.0 mmol). The reaction mixture was stirred at the same
temperature for another 0.5 h. The cold solution was filtered, and the
precipitate was washed with DME (2 ꢁ 10 mL). To the combined
filtrate stirred in an ice bath was added NaBH4 (0.741 g, 19.5 mmol in
10 mL water) slowly. A strong effervescence occurred. The aqueous layer
was extracted three times with ethyl acetate, and the combined organic
layers were washed with brine and dried over Na2SO4. The solvent was
evaporated under reduced pressure, and the oily residue was purified by
column chromatography to give a colorless liquid (3.2 g, 10.9 mmol):
yield 84%; 1H NMR (CDCl3, 300 MHz) δ 5.26 (br s, 1H), 3.84ꢀ3.50
(m, 12 H), 3.36 (s, 3H), 2.93 (br s, 1H), 1.42 (s, 9H); 13C NMR
(CDCl3, 75 MHz) δ 28.3, 51.5, 58.9, 63.2, 70.3, 70.4, 70.5, 71.3, 71.8,
79.5, 156.0; HRMS (ESI/MSm/z) M calcd for C13H27NO6Na 316.1736,
found 316. 1719.
Boc-(2-(2-methoxyethoxy)ethyl)-L-serine-Ψ[CH2N(o,p-diNBS)]Gly-
OEt (4). To a stirred, cold solution of o,p-diNBS-Gly-OEt (3.53 g,
10.5 mmol), triphenylphosphine (2.73 g, 10.5 mmol), and compound 3
(3.1 g, 10.5 mmol) in dried THF (20 mL) under an inert atmosphere
was added DIAD (1.48 mL, 10.5 mmol) dropwise over a period of 0.5 h.
The reaction mixture was allowed to warm to room temperature and
then stirred overnight. The solvent was evaporated, and the oily residue
was purified by column chromatography to afford a yellow solid (3.6 g,
5.91 mmol): yield 56%; 1H NMR (CDCl3, 300 MHz) δ 8.51 (dd, J =
2.1 Hz, 1H), 8.41 (d, J = 2.1 Hz, 1H), 8.32 (d, J = 8.7 Hz, 1H), 5.16 (d, J =
8.6 Hz, 1H), 4.34 (dd, J = 18.6 Hz, 2H), 4.10 (2 ꢁ q, J = 3.6 Hz, 2H),
4.01ꢀ3.87 (m, 1H), 3.70ꢀ3.49 (m, 12H), 3.39 (s, 3H), 1.43 (s, 9H),
1.23 (t, J = 7.1 Hz, 3H); 13C NMR (CDCl3, 75 MHz) δ 14.0, 28.2, 48.3,
48.7, 49.6, 58.9, 61.6, 70.0, 70.3, 70.4, 70.8, 71.9, 79.7, 119.4, 126.0,
132.0, 138.4, 147.9, 149.5, 155.6, 168.6; HRMS (ESI/MSm/z) M calcd
for C23H36N4O13SNa 631.1898, found 631.1840.
was dissolved in ethyl acetate (100 mL) and washed with a saturated
solution of NaHCO3 (100 mL) followed by 10% KHSO4 (100 mL). The
organic layer was washed with brine (50 mL) and dried over Na2SO4.
The solvent was removed under reduced pressure. The crude product
was purified by column chromatography to afford a white solid (0.480 g,
0.88 mmol): yield 67%; 1H NMR (DMSO-d6, 300 MHz) δ 7.27ꢀ7.19
[2 ꢁ d (Rot1,2), J1 = J2 = 1.0 Hz, 1H], 6.86ꢀ6.57 [2 ꢁ d (Rot1,2) [two
rotamers were found, as determined by multinuclear and multidimen-
sional NMR experiments73], J1 = J2 = 8.6 Hz, 1H], 4.80ꢀ4.36 [2 ꢁ ABq
(Rot1,2), J1 = J2 = 16.8 Hz, 2H], 4.33ꢀ3.65 (m, 5H), 3.58ꢀ3.25 (m,
12H), 3.22ꢀ3.15 [2 ꢁ s(Rot1,2), 3H], 1.73 (s, 3H), 1.35 (br s, 9H),
1.27ꢀ1.11 [2 ꢁ t(Rot1,2), J1 = J2 = 7.1 Hz, 3H]; 13C NMRmajor rotamer
(DMSO-d6, 75 MHz) δ 12.3, 14.4, 28.6, 47.9, 48.1, 48.6, 49.5, 58.5, 60.9,
70.0, 70.3, 71.7, 78.6, 108.5, 142.4, 151.4, 155.8, 164.8, 168.0, 169.3;
HRMS (ESI/MSm/z) M calcd for C24H40N4O10Na 567.2642 found,
567.2651.
Boc-(2-(2-methoxyethoxy)ethyl)-L-serine Adenine(Cbz) Ethyl Ester
(6b). Adenine ester 6b was prepared, purified, and characterized the
same way as described for 6a: 1H NMR (DMSO-d6, 300 MHz) δ 10.56
(br s, 1H), 8.59ꢀ8.55 [2 ꢁ s (Rot1,2), 1H], 8.30ꢀ8.24 [2 ꢁ s (Rot1,2),
1H], 7.55ꢀ7.20 (m, 5H), 7.04ꢀ6.53 [2 ꢁ d (Rot1,2), J1 = J2 = 8.5 Hz,
1H ], 5.35 [ABq (Rot1), J = 17.2 Hz, Rot2 appears as a br s at 5.15 ppm,
2H], 5.20 (s, 2H), 4.54ꢀ3.87 (m, 5H), 3.81ꢀ3.31 (m, 12H), 3.23ꢀ3.10
[2 ꢁ s (Rot1,2), 3H], 1.36 (s, 9H), 1.30ꢀ1.10 [2 ꢁ t (Rot1,2), J1 = J2 =
7.1 Hz, 3H]; 13C NMRmajor rotamer (DMSO-d6, 75 MHz) δ 14.4, 28.6,
44.3, 48.7, 49.3, 49.9, 58.4, 60.9, 66.7, 70.0, 70.2, 70.4, 71.0, 71.6, 78.7,
123.4, 128.3, 128.4, 128.8, 136.8, 145.5, 149.9, 151.9, 152.7, 152.9, 155.8,
167.4, 169.2; HRMS (ESI/MSm/z) M calcd for C32H45N7O10Na
710.3126, found 710.3110.
Boc-(2-(2-methoxyethoxy)ethyl)-L-serine PNA Guanine(Cbz) Ethyl
Ester (6c). Guanine ester 6c was prepared, purified, and characterized
the same way as described for 6a: 1H NMR (DMSO-d6, 300 MHz) δ
7.78ꢀ7.74 [2 ꢁ s (Rot1,2), 1H], 7.45ꢀ7.28 (m, 5H), 6.98ꢀ6.59 [2 ꢁ d
(Rot1,2), J1 = J2 = 8.5 Hz, 1H], 5.23 (s, 2H), 5.20ꢀ4.84 [ABq (Rot1), J =
17.1 Hz, m (Rot2), 2H], 4.45ꢀ3.77 (m, 5H), 3.76ꢀ3.33 (m, 12H),
3.22ꢀ3.11[2 ꢁ s (Rot1,2), 3H], 1.38ꢀ1.28 [2 ꢁ s (Rot1,2), 9H], 1.28ꢀ
1.10 [2 ꢁ t (Rot1,2), J1 = J2= 7.2 Hz, 3H ]; 13C NMRmajor rotamer
(DMSO-d6, 75 MHz) δ 14.4, 28.5, 44.1, 48.8, 49.4, 49.8, 58.4, 61.0,
67.7, 70.0, 70.1, 70.2, 70.4, 71.6, 78.4, 119.6, 128.5, 128.6, 128.8, 128.9,
135.9, 140.8, 147.8, 149.9, 155.0, 155.6, 155.8, 167.4, 169.2; HRMS
(ESI/MSm/z) M calcd for C32H45N7O11Na 726.3075, found 726.3070.
Boc-(2-(2-methoxyethoxy)ethyl)-L-serine PNA Cytosine(Cbz) Ethyl
Ester (6d). Cytosine ester 6d was prepared, purified, and characterized
the same way as described for 6a: 1H NMR (DMSO-d6, 300 MHz) δ
7.84 (d, J = 7.3 Hz, 1H), 7.46ꢀ7.26 (m, 5H), 6.99 (d, J = 7.3 Hz, 1H),
6.87ꢀ6.57 [2 ꢁ d (Rot1,2), J1 = J2 = 8.4 Hz, 1H], 5.17 (s, 2H), 4.93ꢀ
4.52 [2 ꢁ ABq (Rot1,2), J1 = J2 = 15.9 Hz, 2H], 4.37ꢀ3.68 (m, 5H),
3.63ꢀ3.29 (m, 12H), 3.22ꢀ3.15 [2 ꢁ s (Rot1,2), 3H], 1.35 (s, 9H),
1.27ꢀ1.09 [2 ꢁ t (Rot1,2), J1= J2= 7.1 Hz, 3H]; 13C NMRmajor rotamer
(DMSO-d6, 75 MHz) δ 14.4, 28.6, 48.7, 49.9, 58.5, 60.9, 66.9, 70.0, 70.3,
71.7, 78.6, 94.3, 128.3, 128.6, 128.9, 136.4, 151.1, 153.6, 155.4, 155.7,
163.6, 167.9, 169.2; HRMS (ESI/MSm/z) M calcd for C31H45N5O11Na
686.3013, found 686.3006.
Boc-(2-(2-methoxyethoxy)ethyl)-L-serine Thymine Monomer (7a).
To a stirred, cold solution of 6a (0.480 g, 0.88 mmol) in THF (10 mL)
was added 2 N NaOH (10 mL) dropwise over a period of 15 min. The
resulting mixture was stirred at the same temperature for another 0.5 h.
Upon completion of the reaction, as confirmed by TLC, H2O (20 mL)
was added, and the resulting mixture was extracted with ethyl acetate
(2 ꢁ 25 mL). The aqueous layers were combined, acidified with 5% HCl
to pH ≈ 4 at 0 οC, and then extracted with ethyl acetate (4 ꢁ 25 mL) and
dried over Na2SO4. The solvent was evaporated in vacuo, and the crude
product was purified by column chromatography to afford a colorless
solid (0.400 g, 4.5 mmol): yield 87%; 1H NMR (DMSO-d6, 300 MHz) δ
Boc-(2-(2-methoxyethoxy)ethyl)-L-serine-Ψ[CH2N]Gly-OEt (5). To
a stirred solution of 4 (2.6 g, 4.2 mmol) in dichloromethane (20 mL) was
added n-propylamine (7.0 mL, 85.4 mmol) under an inert atmosphere.
The reaction mixture was stirred at room temperature for another 20 min.
The solvent was evaporated, and the crude mixture was purified by
column chromatography to afford 5 as a light yellow liquid (1.3 g,
3.4 mmol): yield 81%; 1H NMR (CDCl3, 300 MHz) δ 5.13 (br s, 1H),
4.12 (q, J = 7.0 Hz, 2H), 3.78ꢀ3.63 (m, 1H), 3.62ꢀ3.33 (m, 12H), 3.32
(s, 3H), 2.71 (2 ꢁ dd, J1 = J2 = 5.9 Hz, 2H), 1.80 (br s, 1H), 1.39 (s, 9H),
1.22 (t, J = 7.3 Hz, 3H); 13C NMR (CDCl3, 75 MHz) δ 14.2, 28.3, 49.9,
50.3, 51.0, 58.9, 60.5, 70.4, 70.6, 71.4, 71.9, 79.1, 155.6, 172.4; HRMS
(ESI/MSm/z) M calcd for C17H34N2O7Na 401.2264, found 401.2278.
Boc-(2-(2-methoxyethoxy)ethyl)-L-serine Thymine Ethyl Ester (6a).
To a stirred solution of thymine acetic acid (0.287 g, 1.56 mmol) in dried
DMF (15 mL) under an inert atmosphere were added DCC (0.324 g,
1.56 mmol) and DhbtOH (0.254 g, 1.56 mmol). The resulting mixture
was stirred at room temperature for 1 h. Compound 5 (0.5 g, 1.32 mmol)
was dissolved in dried DMF (10 mL) and added to the above reaction
ο
mixture, and the resulting mixture was stirred at 50 C for 24 h. The
solvent was removed under reduced pressure, and the remaining residue
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7.27ꢀ7.18 [2 ꢁ s (Rot1,2), 1H], 6.88ꢀ6.53 [ 2 ꢁ d (Rot1,2), J1 = J2 =
8.2 Hz, 1H], 4.75ꢀ4.33 [ABq (Rot1), J = 17.5 Hz, m (Rot2), 2H],
4.01ꢀ3.59 (m, 3H), 3.55ꢀ3.25 (m, 12H), 3.22ꢀ3.19 [2 ꢁ s (Rot1,2),
3H], 1.73 (s, 3H), 1.35 (s, 9H); 13C NMRmajor rotamer (DMSO-d6,
75 MHz) δ 12.3, 28.7, 48.1, 49.3, 51.9, 58.5, 70.0, 70.2, 70.3, 71.0, 71.7,
78.2, 78.5, 108.5, 142.6, 151.5, 155.7, 164.8, 167.5, 168.5; HRMS (ESI/
MSm/z) M calcd for C22H35N4O10Na2 561.2148, found 561.2115.
Data for Boc-(2-(2-methoxyethoxy)ethyl)-L-serine adenine(Cbz)
monomer (7b): 1H NMR (DMSO-d6, 300 MHz) δ 8.57ꢀ8.53 [2 ꢁ s
(Rot1,2), 1H], 8.30ꢀ8.16 [2 ꢁ s (Rot1,2), 1H], 7.47ꢀ7.24 (m, 5H),
7.04ꢀ6.50 [2 ꢁ d (Rot1,2), J1 = J2 = 8.7 Hz, 1H], 5.22 [ABq (Rot1), J =
16.4 Hz, Rot2 appears as a br s at 5.10 ppm, 2H], 5.20 (s, 2H), 4.50ꢀ3.65
(m, 5H), 3.63ꢀ3.43 (m, 5H), 3.41ꢀ3.21 (m, 5H), 3.20ꢀ3.14 [2 ꢁ s
(Rot1,2),3H], 1.41ꢀ1.27 [2 ꢁ s (Rot1,2),9H]; 13C NMRmajor rotamer
(DMSO-d6, 75 MHz) δ 28.6, 44.5, 49.3, 58.5, 63.4, 66.7, 70.0, 70.2, 70.4,
71.0, 71.7, 78.2, 78.6, 123.3, 126.8, 128.3, 128.4, 128.8, 136.8, 145.7,
149.7, 151.8, 152.9, 155.7, 167.8; HRMS (ESI/MSm/z) M calcd for
C30H40N7O10Na2 704.2632, found 704.2620.
Data for 3,3,3-trifluoro-N-{2-hydroxy-1-[[2-(2-methoxyethoxy)-
ethoxy]methyl]ethyl}-2-methoxy-2-phenylpropionamide (9): 1H NMR
(300 MHz, CDCl3) δ 7.55 (m, 2H), 7.44 (br s, 1H), 7.40 (m, 3H), 4.13
(m, 1H), 3.83 (m, 1H), 3.70ꢀ3.60 (m, 9H), 3.54 (m, 2H), 3.41(m, 3H),
3.37 (s, 3H), 3.07 (br s, 1H); 13C NMR (CDCl3, 75 MHz) δ 50.5,
54.3, 58.4, 62.2, 69.8, 69.89, 69.9, 70.2, 71.3, 72.1, 83.4, 121.4, 125.2,
127.2, 128.0, 128.9, 132.0, 165.9; HRMS (ESI/MSm/z) M calcd for
C18H26F3NO6Na 432.1610 found 432. 1595; 19F NMR (300 MHz,
CDCl3) δ ꢀ69.43 (s, L-stereomer), ꢀ69.33 (s, D-stereomer).
Data for {[3-[2-(2-methoxyethoxy)ethoxy]-2-[(3,3,3-trifluoro-2-
methoxy-2-phenylpropionyl)amino]propyl][2-thyminylacetyl]amino}-
acetic acid (10): 1H NMR (300 MHz, DMSO-d6) δ 11.22 (br s, 1H),
8.33 and 8.25 (2 ꢁ d, J = 8.5 Hz, 1H), 7.54ꢀ7.35 (m, 5H), 7.09 and 6.96
(2 ꢁ s,1H), 4.52ꢀ4.26 (ABq, J = 16.8 Hz, 2H), 4.26ꢀ4.16 (m, 1H)
3.6ꢀ3.34 (m, 16H), 3.21 and 3.20 (2 ꢁ s, 3H), 3.10 (ABq, J =
14.0 Hz, 2H), 1.70 and 1.68 (2 ꢁ s, 3H); 13C NMRmajorrotamer
(DMSO-d6, 75 MHz) δ 11.8, 47.5, 48.0, 48.4, 52.2, 54.8, 58.0, 69.2,
69.5, 69.6, 69.9, 71.2, 83.7, 107.9, 127.0, 127.2, 128.3, 129.3, 133.3, 142.0,
151.0, 164.3, 165.3, 168.2, 171.6; HRMS (ESI/MSm/z) M calcd for
C27H35F3N4O10Na 655.2305 found 655.2299; 19F NMR (300 MHz,
CDCl3) δ ꢀ68.80 and ꢀ68.95 (s, L-stereomer), ꢀ68.70 (s, D-stereomer).
Oligomer Synthesis. Unmodified, Boc-protected PNA monomers
were purchased from a commercial supplier (they are no longer
available). The oligomers were synthesized on MBHA resin according to
the published protocol.87 Upon completion ofthelast monomercoupling,
the oligomers were cleaved from the resin (and the protecting groups
were simultaneously removed) by immersing the resin in a cocktail
containing m-cresol/thioanisole/TFA/TFMSA (150/150/900/300 μL—
for 100 mg of resin) for 2 h. The crude mixture was eluted and
precipitated in ethyl ether, dissolved in a water/acetonitrile mixture
(80:20), purified by reversed-phase HPLC, and characterized by MAL-
DI-TOF mass spectrometry. A solution of R-cyano-4-hydroxycinnamic
acid (10 mg of R-cyano-4-hydroxycinnamic acid in 500 mL of water with
0.1% TFA and 500 mL of acetonitrile with 0.1% TFA) was used as the
matrix for MALDI-TOF analysis. Concentrations of the oligomers were
determined by UV absorption at 260 nm at 90 °C in water using the
following extinction coefficients: 13 700 Mꢀ1 cmꢀ1 (A), 11 700 Mꢀ1 cmꢀ1
(G), 6600 Mꢀ1 cmꢀ1 (C), and 8600 Mꢀ1 cmꢀ1 (T).
Circular Dichroism (CD). All CD data were recorded at room
temperature unless otherwise stated. All spectra represent an average of
at least 15 scans, recorded from 320 to 200 nm at the rate of 100 nm/min.
A 1 cm path length cuvette was used, and the temperature was main-
tained at 22 °C. All spectra were processed using Origin software,
baseline subtracted and, unless otherwise noted, smoothed using a five-
point adjacent averaging algorithm.
Melting Temperature (Tm). All hybridization experiments were
performed by measuring the change in the absorbance at 260 nm as a
function of temperature. All samples were prepared by mixing a
stoichiometric amount of each strand (5 μM) in a 10 mM sodium
phosphate at pH 7.4 and annealed prior to each measurement. The
recorded spectra were smoothed using a five-point adjacent averaging
algorithm unless otherwise specified. The melting transitions were
determined by taking the first derivative of the absorbanceꢀtemperature
profile.
Data for Boc-(2-(2-methoxyethoxy)ethyl)-L-serine guanine(Cbz)
1
monomer (7c): H NMR (DMSO-d6, 300 MHz) δ 7.78ꢀ7.72 [2 ꢁ s
(Rot1,2), 1H], 7.48ꢀ7.23 (m, 5H), 7.00ꢀ6.61 [2 ꢁ d (Rot1,2), J1 = J2 =
8.6 Hz, 1H], 5.22ꢀ5.19 [2 ꢁ s (Rot1,2), 2H ], 5.03ꢀ4.84 [ABq (Rot1),
J = 17.3 Hz, br s (Rot2), 2H], 3.98ꢀ3.62 (m, 4H), 3.59ꢀ3.40 (m, 7H),
3.41ꢀ3.23 (m, 4H), 3.22ꢀ3.12 [2 ꢁ s (Rot1,2), 3H], 1.35ꢀ1.31 [2 ꢁ s
(Rot1,2), 9H];13CNMRmajor rotamer (DMSO-d6, 75MHz) δ28.6, 44.4, 49.3,
52.7, 58.5, 67.4, 70.0, 70.1, 70.2, 70.4, 71.0, 71.7, 78.2, 119.4, 128.4, 128.5,
128.6, 128.9, 136.1, 140.8, 147.9, 150.2, 155.7, 166.7, 167.9, 171.9; HRMS
(ESI/MSm/z) M calcd for C30H40N7O11Na2 720.2581, found 720.2581.
Data for Boc-(2-(2-methoxyethoxy)ethyl)-L-serine cytosine(Cbz)
monomer (7d): 1H NMR (DMSO-d6, 300 MHz) δ 7.84 (d, J =
7.2 Hz, 1H), 7.43ꢀ7.24 (m, 5H), 7.01ꢀ6.93 [2 ꢁ d (Rot1,2), J1 =
J2 = 7.3 Hz, 1H], 6.87ꢀ6.58 [2 ꢁ d (Rot1,2), J1 = J2 = 8.73 Hz, 1H ], 5.17
(s, 2H), 4.9ꢀ4.5 [ABq (Rot1), J = 15.8 Hz, br s (Rot2), 2H], 4.01ꢀ3.61
(m, 3H), 3.58ꢀ3.43 (m, 7H), 3.43ꢀ3.23 (m, 5H), 3.22ꢀ3.16 [2 ꢁ s
(Rot1,2), 3H], 1.35 (s, 9H); 13C NMRmajor rotamer (DMSO-d6, 75 MHz)
δ 28.1, 48.6, 48.9, 49.7, 52.2, 58.0, 66.4, 69.5, 69.7, 69.8, 70.4, 71.2, 77.8,
94.0, 127.9, 128.1, 128.4, 135.9, 150.9, 153.1, 155.1, 155.2, 163.0, 167.8,
172.2; HRMS (ESI/MSm/z) M calcd for C29H40N5O11Na2 680.2520,
found 680.2546.
Determination of Optical Purities. A General Procedure for
Preparing MTPA Derivatives. Compound 8. To a stirred, cold solution
of 2 (100 mg) in DCM (5 mL) was added 5 mL of TFA/m-cresol
solution (95:5). The ice bath was removed, and the reaction mixture was
stirred at room temperature overnight. The solvent was evaporated
under reduced pressure and triturated with diethyl ether (3 ꢁ 5 mL).
The remaining residue was dried under high vacuum overnight and used
in the next coupling step without further purification. The crude residue
was dissolved in DCM (5 mL) and chilled at 0 °C. DIPEA (2.0 equiv)
and (S)-(+)-R-methoxy-R-(trifluoromethyl)phenylacetyl chloride (MTPA-
Cl; 1.1 equiv) were added, and the reaction mixture was stirred at room
temperature overnight. After completion, the solution was diluted with
DCM and washed with water (2 ꢁ 10 mL) and then brine solution. The
organic layer was dried on Na2SO4, concentrated under reduced
pressure, and purified by column chromatography. Compounds 9 and
10 were prepared the same way.
Data for 3-[2-(2-methoxyethoxy)ethoxy]-2-[(3,3,3-trifluoro-2-
methoxy-2-phenylpropionyl)amino]propionic acid (8): 1H NMR
(300 MHz, CDCl3) δ 7.78 (br s, 1H), 7.56 (m, 2H), 7.37 (m, 3H),
4.47 (m, 1H), 4.03ꢀ3.7 (m, 2H), 3.68ꢀ3.45 (m, 8H), 3.40 (s, 3H), 3.30
(s, 3H); 13C NMR (DMSO-d6, 75 MHz) δ 42.7, 52.1, 54.8, 55.6, 57.9,
60.1, 68.9, 69.5, 69.6, 71.2, 72.3, 83.8, 127.2, 127.5, 128.1, 129.4, 165.3,
174.7; HRMS (ESI/MSm/z) M calcd for C18H24F3NO7Na 446.1403,
found 446.1359; 19F NMR (300 MHz, CDCl3) δ ꢀ69.58 (s, L-stereomer),
ꢀ69.24 (s, D-stereomer).
Thermodynamic Analysis. Thermodynamic parameters were
determined from concentration-dependent Tm measurements following
van’t Hoff analysis.95 ΔH° and ΔS° were obtained from a plot of 1/Tm
(Kꢀ1) vs ln CT, where CT is the total strand concentration. ΔH° was
obtained from the slope of the plot, which should be a linear function,
using the relationship slope = (n ꢀ 1)R/ΔH, where n is the molecularity
of the association interaction (in this case, n = 2) and R is the gas
constant (R = 8.314 J/(mol K)). ΔS° was determined from the y
3
intercept (ln CT = 0), where for a non-self-complementary oligonucleo-
tide the following relationship stands: slope = [ΔS° ꢀ (n ꢀ 1)R ln 2n]/
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ARTICLE
ΔH°. ΔG° was calculated at 298.15 K using the equation ΔG° = ΔH° ꢀ
TΔS° and K using the equation ΔG° = ꢀRT ln K.
perfectly matched and single-base-mismatched targets, CD melt-
ing curves of PNA2ꢀ5, and SPR sensorgrams and kinetic fits for
PNA1ꢀ5. This material is available free of charge via the Internet
at http://pubs.acs.org.
SPR Analysis. DNA and PNA concentrations were calculated by
measuring sample absorbance at 260 nm and 90 °C. At this temperature
the bases are unstacked, resulting in the extinction coefficient of the
oligomer being equal to the sum of the extinction coefficients of its bases.
The DNA extinction coefficients were obtained from the literature, while
thePNA extinctioncoefficients were provided bythecommercialsupplier.
SPR experiments were conducted on a Biacore T100 with a four-
channel carboxymethylated dextran-coated gold sensor chip (CM5) at
25 °C. Approximately 6300 response units (RU) of streptavidin was
immobilized on the chip’s surface at a flow rate of 5 μL/min using
standard N-hydroxysuccinimide/1-ethyl-3-[3-(dimethylamino)propyl]-
carbodiimide hydrochloride (NHS/EDC) coupling followed by capping
with ethanolamine. Then 100 RU of 50-biotinylated DNA was attached to
the chip via noncovalent capture at a flow rate of 2 μL/min and a sample
concentration of 25 nM. HBS-EP (0.01 M HEPES, 0.15 M NaCl, 3 mM
EDTA, and 0.005% Surfactant P-20, pH 7.4) was used as the running
buffer for both the streptavidin and biotinylated DNA immobilizations.
Kinetic information was obtained from the SPR studies by flowing a
dilution series (10ꢀ50 nM in 10 nM increments) of PNA across the
chip. All samples were run in triplicate at a flow rate of 50 μL/min with a
running buffer consisting of 10 mM sodium phosphate, pH 7.4, 100 mM
NaCl, 0.1 mM EDTA, and 0.005% Surfactant P-20. The chip was
regenerated with two 30 s pulses of 10 mM NaOH/1 M NaCl. Buffer
injections were conducted before each sample injection to flush any
lingering regeneration solution. All samples were reference corrected to
account for nonspecific binding by subtracting off the signal resulting
from flowing sample over a streptavidin-coated (i.e., no DNA) flow cell.
Kinetic constants were calculated using a 1:1 binding global fit model in
Biacore T100 Evaluation Software.
’ AUTHOR INFORMATION
Corresponding Author
*E-mail: dly@andrew.cmu.edu.
’ ACKNOWLEDGMENT
Financial support for this work was provided by the National
Institutes of Health (Grant GM076251) and the National Science
Foundation (NSF; Grant CHE-1012467) to D.H.L. SPR instru-
mentation was purchased with support from NSF Major Re-
search Instrumentaion Award 0821296. K.J.Z. gratefully acknowl-
edges support from Department of Defense, Air Force Office of
Scientific Research, National Defense Science and Engineering
Graduate Fellowship 32 CFR168a.
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S
Supporting Information. UV absorption spectra of
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