Bilingual Peptide Nucleic Acids: Encoding the Languages of Nucleic Acids and Proteins in a Single Self-Assembling Biopolymer

Article abstract of DOI:10.1021/jacs.9b09146

Nucleic acids and proteins are the fundamental biopolymers that support all life on Earth. Nucleic acids store large amounts of information in nucleobase sequences while peptides and proteins utilize diverse amino acid functional groups to adopt complex s

Full text of DOI:10.1021/jacs.9b09146

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Article
Bilingual Peptide Nucleic Acids: Encoding the Languages of
Nucleic Acids and Proteins in a Single Self-Assembling Biopolymer
Colin S. Swenson, Arventh Velusamy, Hector S. Argueta-Gonzalez, and Jennifer M. Heemstra
J. Am. Chem. Soc., Just Accepted Manuscript • DOI: 10.1021/jacs.9b09146 • Publication Date (Web): 11 Nov 2019
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Bilingual Peptide Nucleic Acids: Encoding the Languages of Nucleic Acids and Proteins
in a Single Self-Assembling Biopolymer
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Colin S. Swenson, Arventh Velusamy, Hector S. Argueta-Gonzalez, Jennifer M. Heemstra*
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Department of Chemistry, Emory University, Atlanta, Georgia 30322, USA
*Correspondence:
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jen.heemstra@emory.edu
Abstract:
Nucleic acids and proteins are the fundamental biopolymers that support all life on Earth.
Nucleic acids store large amounts of information in nucleobase sequences while peptides and
proteins utilize diverse amino acid functional groups to adopt complex structures and perform
wide-ranging activities. Although Nature has evolved machinery to read the nucleic acid code and
translate it into amino acid code, the extant biopolymers are restricted to encoding amino acid or
nucleotide sequences separately, limiting their potential applications in medicine and
biotechnology. Here we describe the design, synthesis, and stimuli-responsive assembly
behavior of a bilingual biopolymer that integrates both amino acid and nucleobase sequences into
a single peptide nucleic acid (PNA) scaffold to enable tunable storage and retrieval of tertiary
structural behavior and programmable molecular recognition capabilities. Incorporation of a
defined sequence of amino acid side-chains along the PNA backbone yields amphiphiles having
a “protein code” that directs self-assembly into micellar architectures in aqueous conditions.
However, these amphiphiles also carry a “nucleotide code” such that subsequent introduction of
a complementary RNA strand induces a sequence-specific disruption of assemblies through
hybridization. Together, these properties establish bilingual PNA as a powerful biopolymer that
combines two information systems to harness structural responsiveness and sequence
recognition. The PNA scaffold and our synthetic system are highly generalizable, enabling
fabrication of a wide array of user-defined peptide and nucleotide sequence combinations for
diverse future biomedical and nanotechnology applications.
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Introduction:
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Nature encodes information, structure, and function in two basic forms of biopolymers:
nucleic acids and proteins. Nucleotide sequences in DNA or RNA encode both genetic information
and complementary molecular recognition properties, while amino acid sequences in peptides
and proteins convey complex information for structure and function. The robust and adaptable
performance of both of these biopolymer structures has been a fundamental driving force for
evolution on Earth and explains their continued ubiquitous presence in Nature. Moreover, the
straightforward design principles and privileged physicochemical properties of both nucleic acids
and proteins position them as attractive materials to leverage and integrate in applications beyond
their canonical roles.1–3 For example, combining the high density of negative charge of DNA
molecules with hydrophobic lipids or organic polymers enables assembly and formation of
complex nanostructures including micelles, tubes, and vesicles.4–6 While these DNA-polymer
conjugates successfully integrate structural responsiveness and can perform sequence-specific
recognition, they are difficult to control in biological systems and are limited to anionic backbones.
Conversely, peptides are able to express myriad biophysical characteristics through
programmable amino acid side-chain functionalities but lack the vast information storage and
highly modular molecular recognition capabilities of nucleic acids. The ability to combine the
functional properties of nucleic acids and proteins into a single biomolecule would largely
overcome these material limitations. A “bilingual” biopolymer, able to simultaneously speak both
amino acid and base-pairing languages, would allow for both the information processing capability
of nucleobase sequence recognition as well as the structural and functional versatility afforded to
peptides and proteins, together creating a powerful tool for biomedical and nanotechnology
applications.
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Despite these clear advantages, uniting these two chemical functionalities presents a
significant logistical challenge. We recognized that peptide nucleic acid (PNA) offered an ideal
scaffold for the design of a bilingual biopolymer, as is it capable of storing sequence-specific
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nucleotide code along a neutral pseudopeptide backbone.7–9 This backbone imparts a number of
desirable properties including extremely high affinity for complementary nucleic acids, enhanced
stability in complex biological environments, and most significantly, the ability to incorporate
amino acid side-chains at sequence-defined positions.10–12 Researchers have used the latter
capability to impact properties such as solubility, cell permeability, enhanced base-pairing, and
bioconjugation.13–22 Additionally, PNA-polymer conjugates have been constructed and shown to
self-assemble.23–25 While these are capable of binding to complementary nucleic acids, the
hydrophobic portion and nucleic acid code exist in different blocks of the polymer and thus
hybridization cannot be used to control assembly. In contrast to these previous examples,
integrating a complex amino acid sequence in the PNA backbone to encode assembly or
conformational information has yet to be explored. We hypothesized that strategically placing
hydrophobic or hydrophilic amino acid side-chains would enable us to controllably induce self-
assembly of amphiphilic structures (Figure 1). We further speculated that addition of a
complementary DNA or RNA strand would result in duplex formation with the PNA strand and a
dramatic change in amphiphilicity, effectively harnessing the nucleic acid code to direct disruption
of the assemblies. Here we demonstrate the design, synthesis, and characterization of such an
amphiphilic PNA sequence and show assembly into micellar architectures under aqueous
conditions. We also show stimuli-responsive disassembly using miRNA-21, a disease-related
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Figure 1: Schematic representation of bilingual PNA biopolymer. Amino acid side-chains at the -position
direct assembly, and disassembly is triggered by recognition of the nucleobase code by a complementary
DNA or RNA strand.
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RNA target.26 Together, these results represent the first example of a bilingual biopolymer capable
of simultaneously encoding both amino acid and nucleotide information for utilization in controlled
assembly and sequence-specific recognition.
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Results and Discussion:
Design and Synthesis of Amphiphilic PNA
To demonstrate the ability of PNA to function as a bilingual biopolymer, we aimed to create
a strand having a nucleobase sequence capable of specific RNA recognition and an amino acid
sequence to impart amphiphilic behavior to drive assembly, analogous to folding or quaternary
assembly of proteins. As a biologically relevant target, we designed a 12-nucleotide sequence
complementary to miRNA-21, a well characterized oncomiR that is upregulated in almost all
cancer types.26 Conveniently, this complementary sequence exhibits low overall purine content
and contains a single guanine nucleobase, averting sequence motifs that are known to be
problematic in PNA synthesis and hybridization (Table 1, Figure 2A).27 To impart hydrophobic
properties in our amino acid sequence, we selected an alanine side-chain, as we previously
observed aggregation of PNA having three methyl side-chains. For the hydrophilic portion of the
sequence, we chose a positively-charged lysine side-chain, which afforded convenient synthetic
access and is known to impart favorable solubility properties.12,28 The side-chains were placed at
Table 1: Sequences of amphiphilic PNA-A, control PNA-C, and amino acid containing PNA-aa. ”D” (orange)
denotes the 4-DMN dye monomer. Subscripts denote the amino acid single letter code. Alanine monomers
are in red. Lysine monomers are in blue. Masses were confirmed by ESI-TOF mass spectrometry.
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Figure 2: (A) Hybridization site of PNA oligomers to the miRNA-21 target used in this study. (B)
PNA monomers synthesized in this study: -methyl monomers of cytosine and thymine, -lysyl
monomers of adenine and thymine, and a solvatochromic 4-DMN monomer.
the -position along the PNA backbone using L-amino acids, as this enables installation of the
side-chains using amino acid precursors, and this location and stereochemistry have been shown
to confer increased binding affinity of PNA with complementary nucleic acids (Figure 1).12,13
Lastly, we integrated a solvatochromic fluorophore, 4-dimethylamino-naphthalimide (4-DMN), at
the hydrophobic terminus of the PNA (Figure 2B). This dye exhibits an enhancement in
fluorescence intensity in increasingly hydrophobic environments,29 and we envisioned that such
an arrangement in the hydrophobic portion would allow us to visualize assembly formation by
fluorescence spectroscopy.
We first synthesized the necessary -modified PNA monomers according to previously
reported procedures.22,28,30–32 Starting from Fmoc-protected L-amino acids, PNA backbones
containing lysine and alanine side-chains were produced. The -modified backbones were
coupled to bis(Boc)-protected nucleobase acetic acids using HATU in the presence of N-N-
diisopropylethylamine (DIPEA) to produce -modified monomers of A, T, and C (Scheme 1).
Synthesis of the 4-DMN PNA monomer was performed using a protocol adapted from that of Saito
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Scheme 1: General coupling procedure for the synthesis of -modified PNA
monomers. (a) Nucleobase acetic acid, HATU, DIPEA, DMF; (b) H2, Pd/C, MeOH.
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Scheme 2: Synthesis of 4-DMN PNA monomer. (a) 3-
(dimethylamino)propionitrile, isoamyl alcohol, reflux, quant. (b) Glycine
tert-butyl ester hydrochloride, DIPEA, DMF, 78%. (c) H2, Pd/C, methanol,
94%. (d) Fmoc (aminoethyl)glycine benzyl ester, HATU, DIPEA, DMF,
72%. (e) H2, Pd/C, methanol, quant.
and coworkers.33 First, 4-dimethylamino-naphthalic anhydride was generated by reaction of 4-
bromo-1,8-naphthalic anhydride with 3-dimethylamino-propionitrile in isoamyl alcohol.34 The
imide was then produced by condensation with glycine tert-butyl ester hydrochloride in the
presence of DIPEA.35 Removal of the tert-butyl protecting group by hydrolysis afforded the 4-DMN
acetic acid. Unmodified (glycine) PNA backbone was synthesized according to previously
reported procedure,36 and the 4-DMN acetic acid was coupled to the unmodified backbone using
HATU/DIPEA conditions to afford the 4-DMN PNA monomer (Scheme 2).
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Amphiphilic PNA and control PNA oligomers were then produced using solid-phase
synthesis employing HATU/DIPEA coupling conditions on a semi-automatic synthesizer with
microwave assistance. Initially, we sought to place three hydrophobic side-chains and three
hydrophilic side-chains equally spaced along the backbone. However, we were unable to obtain
full-length product with this design. We hypothesized that this was due to the close proximity of
semi-consecutive -modified monomers, which could severely lower the efficiency of the coupling
reactions, resulting in little to no product formation. With this in mind, we reduced the number of
-modified monomers to two alanine and two lysine side-chains. This design allowed placement
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of at least two unmodified monomers between each -modified residue, thereby mitigating
otherwise poor coupling yields due to steric strain. Using this design, we produced an amphiphilic
PNA sequence, PNA-A, as well as an unmodified control strand, PNA-C, containing a single C-
terminal lysine, as is typically used in PNA strands to ensure solubility. In order to demonstrate
the importance of using -modified monomers to encode the amino acid sequence along the PNA
backbone, we also synthesized a control amphiphilic PNA, PNA-aa. This PNA has two alanine
amino acids and two lysine amino acids on the C- and N-termini, respectively, and thus represents
a sequence that is amphiphilic, but not bilingual, as the nucleotide and amino acid codes exist in
separate blocks of the polymer (Table 1). The strands were purified via reverse-phase HPLC and
characterized by ESI-TOF mass spectrometry to confirm identity (Supporting Information).
After synthesis and characterization of the PNA sequences, we performed UV melting
experiments to analyze the hybridization of each sequence to a complementary DNA or the
miRNA-21 target. The presence of side chains at the -position is known to increase the duplex
stability of PNA to complementary nucleic acids.13,14,21 Thus, we were not surprised to find that
the complementary DNA exhibits a melting temperature (Tm) of 69.5 C with PNA-A, compared to
63.0 C and 64.5 C for PNA-C and PNA-aa, respectively (Figure S1A). miRNA-21 also displays
a higher Tm of 68.6 C with PNA-A, compared to 65.8 C and 67.5 C for PNA-C and PNA-aa,
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respectively, demonstrating that our -modifications improve the binding stability to
complementary sequences (Figure S1B).
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Characterization of Amphiphilic PNA Assembly
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The ability of proteins to form tertiary and quaternary structures based on the information
encoded in their primary amino acid sequence is a powerful tool. We envisioned that we could
mimic this characteristic of proteins by encoding an amphiphilic amino acid sequence into the
PNA backbone. After synthesis and characterization of the PNA strands, we sought to analytically
confirm assembly formation and determine the critical micelle concentration (CMC) of PNA-A. To
achieve this, we leveraged the ability of the 4-DMN dye to respond to changes in solvation, and
we hypothesized that the dye’s placement at the hydrophobic portion of the amphiphile should
result in enhanced fluorescence upon assembly (Figure 3A).29 CMC values determined using
fluorescence-based methods often utilize a constant concentration of dye to visualize an inflection
point where fluorescence increases. However, in our case the 4-DMN dye acts as an internal
reporter, and thus changes in concentration as the concentration of amphiphile is varied. Given
this approach, we instead quantified the relative fluorescence of PNA-A compared to the non-
assembling dye monomer in solution. Indeed, we observed a concentration-dependent increase
in fluorescence of amphiphile PNA-A over that of the 4-DMN monomer alone, suggesting the
formation of assemblies having the dye sequestered in their hydrophobic core. Specifically we
observe a discontinuous change in slope with increasing concentration, as is typical of micelle
formation. In contrast, the control strand, PNA-C, displayed very little fluorescence enhancement
with increasing concentrations, as it does not have an amino acid code to direct assembly (Figure
S2). While PNA-C does show a small amount of fluorescence signal over the 4-DMN monomer
alone, we surmise that this is attributable to the solvation state of the dye when placed near a
hydrophobic nucleobase in the PNA strand, as opposed to the complete solvation of the free
monomer. In any case, by observing the interception of the horizontal lower concentration and
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Figure 3: Characterization of amphiphilic PNA assembly. (A)
Schematic representation of assembly and subsequent 4-DMN
fluorescence response upon encapsulation. (B) Plot of the
fluorescence ratio of PNA-A to the 4-DMN monomer as a function of
concentration. Error bars represent SEM (n=3). (C) Histogram of the
size distribution of particles as measured by ImageJ using TEM
images. (D) TEM images of PNA-A at 100 M. Scale bars = 2000
nm (top left), 1000 nm (top right), 500 nm (bottom left) and 50 nm
(bottom right).
vertical upper concentration portions, we were able to estimate a CMC of ~317 nM, which is
sufficiently low for a broad range of stimuli-responsive biological applications (Figure 3B).
We next aimed to visually confirm the formation of assemblies and characterize their
morphology using transmission electron microscopy (TEM). Amphiphile PNA-A was first dissolved
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in ultrapure water and diluted to a concentration of 100 M. This solution was spotted onto
formvar/carbon-coated copper grids and stained using a 1% uranyl acetate solution for TEM
visualization. As expected, we observed the presence of spherical assemblies having an average
diameter of 36.9  9.9 nm, consistent with the formation of micelles (Figure 3C-D). Conversely,
the PNA-C control strand displayed only a small number of amorphous assemblies, likely due to
uncontrolled aggregation of PNA at this high of concentration or drying effects during sample
preparation (Figure S3). The amino acid amphiphile, PNA-aa, displayed a heterogeneous mixture
of amorphous assemblies, some having a similar size as the amphiphile PNA-A, but others being
significantly larger, and all lacking discrete and uniform shapes (Figure S4). This may be
explained by the flexibility of PNA-aa compared to PNA-A, as installing amino acid side-chains
directly onto the PNA backbone at the -position results in the oligomer adopting a right-handed
helical secondary structure that can be visualized via circular dichroism,14 whereas including them
outside of the nucleobases retains the flexibility of the backbone (Figure S6). Irrespective of the
explanation, the ability of PNA-A, but not PNA-aa, to form well-defined assemblies demonstrates
the unique capabilities of our bilingual biopolymers compared to other amphiphilic PNA conjugate
designs.
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In order to gain a better understanding of the assembly properties in solution, we also
carried out analysis using dynamic light scattering (DLS). Samples of PNA were prepared at 500
M, as lower concentrations failed to provide sufficient signal intensity for validated
measurements. For the PNA-A assemblies, we observed a hydrodynamic diameter of 110.2 
31.9 nm according to the number distribution (Figure S7). This is larger than observed by TEM,
but this difference is not entirely unexpected, as DLS has been shown to overestimate the sizes
of particles in solution because it is a measure of the hydrodynamic diameter as opposed to actual
size.37–39 Additionally, in TEM the sample is dried onto a surface during preparation, which may
cause the assemblies to decrease in size. Finally, the high concentrations required for DLS may
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lead to the formation of some larger particles, increasing the average size observed. DLS analysis
of PNA-aa reveals a similar trend, showing assemblies having a hydrodynamic diameter of 344.7
 92.3 nm (Figure S7), which is larger than the assemblies observed by TEM. The non-
amphiphilic PNA-C shows two populations in DLS, with a significant population at 2.4  0.5 nm
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and a small population at 54.9  15.4 nm (Figure S7). The smaller sized population is consistent
with single-stranded PNA while the larger may represent uncontrolled aggregation. This is in
agreement with the TEM data, as only sparse and amorphous assemblies were visualized for the
control PNA-C. Together, these results demonstrate our ability to encode amino acid information
into a PNA sequence to direct the formation of organized nanostructures under aqueous
conditions.
Stimuli-Responsive Switching of Amphiphilic PNA Assembly
Assembly of our PNA amphiphiles is driven by the amino acid code, and we envisioned
that disassembly could be driven by the nucleotide code (Figure 4A). While we designed the
amphiphilic PNA sequence to be responsive to miRNA-21, we used a complementary 12-
nucleotide DNA strand in our initial disassembly studies owing to greater ease of use and lower
cost. First, we sought to confirm the ability of the complementary DNA to bind to the amphiphilic
PNA using circular dichroism (CD). Incorporating -modifications into PNA oligomers imparts
chirality and orients the strand into a helical secondary structure, which can be observed by CD
spectroscopy at concentrations relevant to assembly formation.14 In order to detect PNA:DNA
hybridization and subsequent disassembly, we first prepared PNA-A assemblies at 100 M in 1x
phosphate buffered saline (PBS), pH 7.4, as this concentration has previously displayed
assemblies by TEM (Figure 3D). A range of concentrations of complementary or scrambled DNA
was added to the samples and CD signatures were recorded. We observed signals recording
maxima near 263 nm and 222 nm, with minima occurring near 244 nm and 201 nm, which are
indicative of a right-handed PNA:DNA duplex (Figure 4B).14,40 As the concentration of
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complementary DNA in the samples was increased, the CD signal also increased while
maintaining isosbestic points, suggesting a continuous increase in PNA:DNA duplex formation.
Free DNA alone at 100 M displayed maxima at 282 nm and 217 nm with minima occurring at
256 nm, further indicating that the increasing signal is caused by increased PNA:DNA duplex
formation. As expected, we observed no increase in signal when PNA-A was exposed to the
scrambled DNA sequence (Figure 4D). In fact, we observed a slight decrease in CD signal for
PNA-A in the presence of scrambled DNA, which we speculate could be caused by minor
electrostatic interference between the positively charged assemblies and the negatively charged
DNA. We also confirmed that the control, PNA-C, and amino acid-containing amphiphile, PNA-
aa, are capable of binding to complementary DNA at this concentration (Figure S8). Having
established the ability of PNA-A to decode complementary DNA, we next aimed to explore
whether similar decoding could be achieved using the full miRNA-21 target. Samples for CD
spectroscopy were prepared as previously described at 100 M but using only a single equivalent
of RNA. We observed a significant change in CD signal, in particular a large increase in intensity
at the 202 nm minima over that of PNA or miRNA alone, strongly suggesting the ability of the full
target miRNA-21 to hybridize with PNA-A in water at concentrations relevant to assembly (Figure
4F).
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Since our amphiphile is equipped with an internal fluorogenic dye, we sought to visualize
disassembly via fluorescence response. Samples of PNA-A at 10 M were prepared and mixed
with increasing amounts of complementary and scrambled DNA. As expected, increasing
amounts of complementary DNA caused the fluorescent signal to decrease (Figure S9). The
change in fluorescence was not as significant as that observed via concentration-dependent
disassembly, likely because the dye can undergo base stacking with the hybridized DNA, still
creating a somewhat hydrophobic environment. However, these data still provide evidence for
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DNA-induced disassembly. Interestingly, addition of the scrambled DNA sequence leads to an
increase in fluorescence signal. We hypothesize that this may result from non-specific interactions
between the negatively charged DNA and the positively charged micelles, and that the resulting
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Figure 4: Binding and disassembly of amphiphilic structures by complementary nucleic acids. (A) Schematic
representation of disassembly by hybridization of a complementary DNA or RNA strand. (B) CD plot
confirming hybridization of complementary DNA to PNA-A in 1x PBS at 100 M. (C) TEM image showing
disappearance of spherical structures in the presence of complementary DNA. (D) CD plot showing inability
of a scrambled DNA sequence to hybridize to PNA-A in 1x PBS at 100 M. (E) TEM image showing retention
of spherical assemblies in the presence of scrambled DNA. (F) CD plot confirming binding of full target
miRNA-21 to PNA-A in water at 100 M. (G) TEM image showing disappearance of spherical structures in
the presence of complementary RNA. Scale bars in all TEM images = 500 nm.
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charge repulsion screening leads to compaction of the assemblies and thus a more hydrophobic
environment in the micelle core.
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While the fluorescence data are promising, we wanted to further validate and confirm
disassembly using TEM. In order to visualize stimuli-responsiveness to DNA, amphiphile PNA-A
assemblies were prepared at a concentration of 200 M in 1x PBS. To these samples was added
one equivalent of the complementary or scrambled DNA sequence to a final concentration of 100
M. The samples were spotted onto grids and stained with uranyl acetate for TEM analysis. As
expected, we observed the disappearance of spherical assemblies upon addition of the
complementary DNA sequence, indicating stimuli-responsive disruption of the micelles (Figure
4C). As a control, we observed retention of small spherical nanostructures in the presence of non-
targeting scrambled DNA, strongly indicating that sequence-specific binding is required for
disassembly (Figure 4E). These assemblies appear to be slightly smaller than PNA-A alone,
supporting our hypothesis for the fluorescence increase described above. We also detected some
dark aggregates in the presence of both DNA sequences, and speculate this is likely due to the
negative stain, uranyl acetate, which non-specifically interacts with phosphate ions present in the
DNA backbone and PBS buffer, leading to precipitation. In any case, after confirming binding and
stimuli-responsive disassembly by target DNA, we sought to also demonstrate responsiveness to
the miRNA target of interest. Based on our previous results showing that one equivalent of DNA
was sufficient for binding and disruption of assemblies, we challenged our PNA-A amphiphile
constructs with a similar amount of miRNA-21. Assemblies were prepared in water and miRNA-
21 was added to give final concentrations of 100 M for both PNA and RNA. The samples were
then spotted onto grids and stained with uranyl acetate for TEM visualization. As expected, we
observed the disappearance of spherical assemblies in the presence of miRNA-21 (Figure 4G).
Consistent with our DNA studies, we observed similar staining precipitation patterns with miRNA-
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21, further suggesting that these objects are the result of non-specific artifactual precipitation
between the uranyl acetate stain and phosphate-rich oligomers.
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We had intended to also characterize disassembly using DLS, as this technique was
successful for characterizing the assemblies in solution. However, as previously stated, this
technique requires high concentrations (≥ 500 M) of assemblies in order to obtain a reliable
signal intensity.37–39 At these concentrations, we observed the formation of a precipitate upon
addition of the complementary target or a scrambled sequence. This is likely due to nonspecific
electrostatic interactions which caused uncontrolled and catastrophic aggregation, resulting the
presence of larger assemblies rather than the expected disappearance of assemblies. While
lower concentrations do show the expected lack of assemblies, this may be attributable to
insufficient signal and cannot necessarily be attributed to disassembly.
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Together, the Tm, CD, fluorescence, and TEM results demonstrate that both DNA and
RNA can bind sequence-specifically to PNA-A and disrupt assembly. Thus, even in the
assembled state, the nucleotide code of our bilingual PNA constructs can be accessed and used
to decode nucleic acid sequence information into supramolecular conformational changes. This
property is similar to that shown in other PNA amphiphile systems.23–25 However, these systems
rely upon the addition of a hydrophobic alkyl chain and hydrophilic amino acids on opposite ends
of the PNA oligomer to impart amphiphilic properties. While those designs are advantageous for
ease of synthesis, our design is fundamentally different in that the entire portion of the amphiphile
is capable of nucleobase-specific recognition. This allows greater control over the assembly and
disassembly properties imparted by the amino acid and nucleobase codes, thus providing a truly
modular bilingual biopolymer.
Conclusion:
Nature has evolved two languages for encoding the instructions and functions needed for
life. While extant biopolymers are limited to encoding a single language, we introduce here a non-
natural biopolymer that is able to simultaneously encode both. We report the synthesis and
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characterization of a PNA strand having amino acid-like side-chains comprised of hydrophobic
alanine and hydrophilic lysine, which when arrayed in a specific pattern are able to direct
assembly in a manner analogous to peptide aggregation or quaternary protein assembly. The
PNA strand also carries a specific nucleotide code, which is able to be decoded by complementary
DNA or RNA to direct stimuli-responsive disassembly. These assembly and disassembly events
can be observed both spectroscopically and using TEM, enabling characterization of the CMC
and morphology of the assemblies and the sequence-specific nature of the disassembly process.
The PNA scaffold is highly generalizable, allowing for the programming of both nucleotide and
amino acid sequences to impart unique biological and physicochemical properties.41 Thus, the
ability of this information-rich bilingual biopolymer motif to undergo ordered self-assembly and
conformationally respond to a variety of nucleic acid targets offers significant potential for
applications in nanomedicine and biotechnology. While our initial exploration focused on micellar
architectures, we envision that the bilingual biopolymer motif can be harnessed to recapitulate
many of the structural motifs found in proteins, such as sheets and coils. Moreover, we anticipate
that the ability to simultaneously interpret both of the biological codes can be extended beyond
the creation of stimuli-responsive assemblies to also enable the construction of custom adaptors
able to mediate precise interactions between specific protein and nucleic acid targets.
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Experimental Procedures
Abbreviations: Fmoc, fluorenylmethyloxycarbonyl; DMF, dimethylformamide; HATU, 1-
[Bis(dimethylamino)methylene]-1H-1,2,3-triazolo[4,5-b]pyridinium 3-oxid hexafluorophosphate;
DIPEA, diisopropylethylamine; TFA, trifluoroacetic acid; DCM, dichloromethane; Boc, tert-
butyloxycarbonyl; NMP, N-methyl-2-pyrrolidone.
PNA monomer synthesis
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Unmodified PNA monomers were purchased from PolyOrg, Inc. Fmoc -lysyl and -methyl
modified PNA backbones and nucleobase acetic acids of adenosine, thymine, and cytosine were
prepared by previously published procedures.22,28,30–32 Fmoc-L-amino acids and nucleobase
starting materials were purchased from Chem-Impex International, Inc. All other reagents and
solvents were purchased from Chem-Impex, Sigma-Aldrich, or Fisher Scientific and used without
further purification unless otherwise stated. All dry solvents were distilled and collected from a J.
C. Meyer solvent dispensing system prior to use to ensure dryness. Small molecule synthesis
was monitored by thin layer chromatography (TLC) on Merck silica gel 60 F254 glass plates and
visualized using UV light and staining with a ninhydrin solution followed by heating. Flash
chromatography was performed using SiliaFlash F60 grade silica purchased from SiliCycle Inc.
Compounds were characterized by proton magnetic resonance (1H) and carbon nuclear magnetic
resonance (13C) NMR using a Varian Inova 400 MHz spectrometer containing a Bore Oxford
magnet. Spectra were analyzed using the MestReNova software. The mass of compounds was
confirmed by the use of an Agilent 6230 electrospray ionization time-of-flight (ESI-TOF) mass
spectrometer.
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General coupling procedure for PNA monomer synthesis
Nucleobase acetic acid (1 eq) was dissolved in dry DMF under N2 in a round bottom flask.
To the flask was added HATU (1 eq) in dry DMF and DIPEA (2 eq). The mixture was stirred at
room temperature for 10 min before addition of -modified PNA backbone (0.67 eq) in dry DMF.
The mixture was stirred overnight or until consumption of the backbone as confirmed by TLC
using ninhydrin staining. Upon completion, the reaction was quenched with H2O, extracted with
ethyl acetate and washed with 1M hydrochloric acid (HCl), saturated aqueous sodium bicarbonate
(NaHCO3), H2O, and brine. The organic layer was dried with anhydrous sodium sulfate (Na2SO4)
and concentrated with rotary evaporation. The crude solid was purified by flash column
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chromatography (hexane to ethyl acetate gradient) to give the protected PNA monomer. The
purified benzyl-protected monomer was re-dissolved in methanol under N2. To the solution was
added palladium on activated charcoal (7%/wt.) and the N2 was replaced with a balloon of H2.
The reaction was stirred until complete deprotection as confirmed by TLC. Upon completion, the
solution was filtered through celite and concentrated with rotary evaporation to produce the final
-modified PNA monomer.
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1
Fmoc Lysyl Adenine Monomer (1). (43% yield) H NMR (400 MHz, DMSO-d6, two rotamers) 
8.75 (rotamer one, s, 1H) 8.73 (rotamer two, s, 1H) 8.45 (rotamer one, s, 1H) 8.43 (rotamer two,
s, 1H) 7.88 (d, 2H) 7.69 (t, 2H) 7.26-7.43 (m, 4H) 6.81 (s, 1H) 6.76 (s, 1H) 5.40 (m, 2H) 5.18 (m,
2H) 4.18-4.36 (m, 3H) 3.81 (m, 1H) 3.49 (m, 2H) 2.87 (m, 2H) 1.00-1.60 (m, 6H) 1.38 (s, 9H) 1.36
(s, 18H). 13C NMR (400 MHz, DMSO-d6)  170.78, 170.26, 167.04, 166.58, 156.25, 156.03,
155.64, 153.53, 150.10, 150.02, 148.96, 144.04, 143.91, 140.80, 140.79, 127.80, 127.52, 127.29,
126.94, 125.30, 120.19, 120.14, 83.31, 77.39, 46.95, 29.44, 28.40, 28.22, 27.40, 27.22. HRMS
(ESI-TOF) m/z 887.3907 (calc’d [M + H]+ = 887.4298).
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Fmoc Lysyl Thymine Monomer (2). (49% yield) H NMR (400 MHz, DMSO-d6, two rotamers) 
11.30 (rotamer one, s, 1H) 11.28 (rotamer two, s, 1H) 7.88 (d, 2H) 7.68 (t, 2H) 7.41 (t, 2H) 7.32
(t, 2H) 7.21 (s, 1H) 6.77 (m, 1H) 4.63 (m, 2H) 4.45 (s, 1H) 4.30 (m, 2H) 4.21 (m, 2H) 3.92 (m, 2H)
3.35 (m, 2H) 2.88 (m, 2H) 1.72 (rotamer one, s, 3H) 1.69 (rotamer two, s, 3H) 1.19-1.44 (m, 6H)
1.36 (s, 9H). 13C NMR (400 MHz, DMSO-d6)  170.32, 167.81, 167.45, 164.43, 156.14, 156.00,
155.62, 151.01, 143.91, 143.83, 140.79, 140.76, 127.66, 127.11, 125.24, 120.17, 108.14, 77.38,
65.34, 47.75, 46.87, 38.29, 29.33, 28.30, 22.90, 22.82, 11.96. HRMS (ESI-TOF) m/z 678.3166
(calc’d [M + H]+ = 678.3134).
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Fmoc Methyl Cytosine Monomer (3). (43% yield) H NMR (400 MHz, DMSO-d6)  7.95 (d, 1H)
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7.87 (d, 2H) 7.68 (d, 2H) 7.40 (m, 3H) 7.33 (t, 2H) 6.79 (t, 1H) 4.84 (m, 1H) 4.67 (m, 1H) 4.26 (m,
4H) 3.93 (m, 2H) 3.48 (m, 2H) 1.48 (s, 18H) 1.01 (d, 3H). 13C NMR (400 MHz, DMSO-d6)  168.03,
162.30, 162.22, 155.98, 165.63, 162.11, 149.70, 144.32, 141.14, 128.02, 127.48, 125.65, 120.53,
95.61, 84.92, 64.92, 52.48, 50.41, 48.89, 47.15, 45.68, 27.66, 18.76 HRMS (ESI-TOF) m/z
706.3113 (calc’d [M + H]+ = 706.3083).
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Fmoc Methyl Thymine Monomer (4). (85% yield) 1H NMR (400 MHz, DMSO-d6)  11.25 (s, broad,
1H) 7.87 (d, 2H) 7.68 (d, 2H) 7.41 (m, 3H) 7.32 (t, 2H) 7.22 (s, 1H) 4.63 (m, 1H) 4.44 (m, 2H) 4.25
(m, 3H) 3.83 (s, 2H) 3.44 (m, 2H) 1.70 (s, 3H) 0.99 (d, 3H) 13C NMR (400 MHz, DMSO-d6) 
168.36, 167.50, 164.86, 155.97, 151.46, 144.40, 144.29, 142.63, 141.13, 128.03, 127.51, 125.68,
120.52, 110.00, 108.37, 65.71, 48.26, 47.17, 45.76, 18.87, 12.32. HRMS (ESI-TOF) m/z 521.2051
(calc’d [M + H]+ = 521.2031).
Synthesis of 4-(dimethylamino)naphthalimide monomer:
The 4-DMN monomer was synthesized according to an adapted procedure from Saito, et. al.33
4-dimethylamino-napthalic anhydride (5). 4-bromo-1,8-naphthalic anhydride (1 eq) dissolved in
isoamyl alcohol was brought to reflux in an oil bath. 3-Dimethylamino-proprionitrile (4 eq) was
added and the mixture was refluxed overnight. The reaction was cooled to room temperature and
the precipitate filtered. The precipitate was washed with water and cold hexane and dried under
reduced pressure to yield a brown solid in quantitative yield. 1HNMR (400 MHz, CDCl3)  8.50 (d,
1H) 8.46 (d, 1H) 8.37 (d, 1H) 7.65 (t, 1H) 7.06 (d, 1H) 3.18 (s, 6H). 13C NMR (400 MHz, CDCl3) 
161.76, 160.78, 157.98, 135.03, 133.20, 133.00, 125.02, 124.83, 119.19, 113.18, 109.30, 44.69.
HRMS (ESI-TOF) m/z 242.0834 (calc’d [M + H]+ = 242.0812).
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4-dimethylamino-naphthalimide tert-butyl acetate (6). To a solution of 4-dimethylamino-napthalic
anhydride 5 (36 mmol) and glycine tert-butyl ester (54 mmol) in 180 mL of dry DMF was added
DIPEA (108 mmol). The turbid solution was brought to 80 °C and stirred overnight or until full
consumption of the anhydride by TLC. Upon completion, the reaction was quenched by addition
of 250 mL water and extracted with ethyl acetate (3x250 mL). The organic layer was washed with
water and brine, dried with anhydrous sodium sulfate, and concentrated using rotary evaporation.
The crude solid was purified by flash column chromatrography (1:1 ethyl acetate:hexane) to yield
a yellow solid (78%). 1H NMR (400 MHz, CDCl3)  8.50 (d, 1H) 8.40 (d, 1H) 8.36 (d, 1H) 7.59 (t,
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1H) 7.02 (d, 1H) 4.80 (s, 2H) 3.05 (s, 6H) 1.46 (9H). 13C NMR (400 MHz, CDCl3)  167.40, 164.28,
163.63, 157.20, 133.01, 132.93, 130.45, 125.25, 125.17, 122.58, 114.28, 113.21, 113.11, 81.98,
44.72, 41.96, 28.15. HRMS (ESI-TOF) m/z 355.16595 (calc’d [M + H]+ = 355.1652).
4-dimethylamino-naphthalimide acetic acid (7). 4-dimethylamino-naphthalimide tert-butyl acetate
6 (18 mmol) was dissolved in a 50% mixture of TFA in DCM (90 mL). After stirring for 3 hours,
TLC indicated reaction completion. The reaction was concentrated using rotary evaporation and
azeotroped with toluene (3x). The solid was triturated with hexane/DCM, filtered, and the collected
solid was dried under reduced pressure to yield the final product as a yellow solid (94%). 1H NMR
(400 MHz, DMSO-d6)  13.02 (s, broad, 1H) 8.40 (d, 1H) 8.35 (d, 1H) 8.21 (d, 1H) 7.66 (t, 1 H)
7.08 (d, 1H) 4.67 (s, 2H) 3.04 (s, 6H). 13C NMR (400 MHz, DMSO-d6)  169.59, 163.31, 162.53,
156.76, 132.53, 131.96, 130.77, 129.66, 124.84, 123.98, 121.67, 112.76, 112.35, 44.34, 40.94.
HRMS (ESI-TOF) m/z 299.10318 (calc’d [M + H]+ = 299.1026).
Fmoc 4-dimethylamino-naphthalimide (aminoethyl)glycine benzyl ester (8). 4-dimethylamino-
naphthalimide acetic acid 7 (2.67 mmol) was dissolved in dry DMF (15 mL) under N2 in a round
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bottom flask. The flask was charged with HATU (2.67 mmol) and DIPEA (4.46 mmol). The reaction
was stirred for 15 min before addition of Fmoc(aminoethyl)glycine benzyl ester PNA backbone
(1.78 mmol) in dry DMF (5 mL). The reaction was stirred for 2.5 h, at which time TLC showed
complete consumption of the backbone. Upon completion, the reaction was quenched with H2O,
extracted with ethyl acetate and washed with 1M HCl, saturated NaHCO3, H2O and brine. The
organic layer was dried with anhydrous sodium sulfate and concentrated by rotary evaporation.
The crude yellow solid was purified by flash column chromatography (hexane to ethyl acetate
gradient) to provide the product as a yellow solid (72%). 1H NMR (400 MHz, CDCl3)  8.51 (d, 1H)
8.38 (m, 2H) 7.69 (m, 3H) 7.57 (m, 2H) 7.31 (m, 9H) 7.01 (d, 1H) 6.23 (t, 1H) 5.15 (s, 2H) 5.10 (s,
2H) 4.40 (d, 1H) 4.30 (t, 2H) 4.13 (s, 2H) 3.40-3.70 (m, 4H) 3.05 (s, 6H) 13C NMR (400 MHz,
CDCl3)  169.81, 168.08, 164.43, 163.83, 157.23, 157.09, 144.20, 141.29, 135.23, 133.12,
131.56, 131.46, 130.57, 128.81, 128.68, 128.46, 128.39, 127.64, 127.13, 125.45, 125.20, 124.80,
122.68, 119.90, 114.41, 113.22, 67.89, 67.34, 49.32, 49.11, 47.29, 44.79, 40.98, 39.66. HRMS
(ESI-TOF) m/z 711.2901 (calc’d [M + H]+ = 711.2813).
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Fmoc 4-dimethylamino-naphthalimide PNA monomer (9). Fmoc 4-dimethylamino-naphthalimide
(aminoethyl)glycine benzyl ester 8 (1.49 mmol) was dissolved in methanol (25 mL) under N2.
Palladium on activated charcoal (7%/wt.) was added to the solution. The N2 was replaced by H2
and the reaction was stirred overnight, after which TLC showed full conversion to product. The
reaction was filtered through celite and concentrated using rotary evaporation to provide the
product as a yellow solid in quantitative yield. 1H NMR (400 MHz, DMSO-d6, two rotamers)  8.52
(rotamer one, d, 1H) 8.46 (rotamer two, d, 1H) 8.42 (rotamer one, d, 1H) 8.37 (rotamer two, d,
1H) 8.31 (rotamer one, d, 1H) 8.24 (rotamer two, d, 1H) 7.87 (rotamer one, t, 3H with Fmoc) 7.76
(rotamer two, t, 1H) 7.70 (d, 2H) 7.66 (d, 2H) 7.54 (t, 1H) 7.20 (rotamer one, d, 1H) 7.14 (rotamer
two, d, 1H) 4.98 (s, 2H) 4.80 (s, 2H) 4.34 (d, 2H) 4.27 (t, 1H) 3.56 (t, 2H) 3.15 (m, 2H) 3.10
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(rotamer one, s, 6H) 3.07 (rotamer two, s, 6H). 13C NMR (400 MHz, DMSO-d6)  167.39, 166.82,
136.39, 162.67, 156.75, 156.46, 156.13, 143.94, 140.75, 132.41, 131.85, 130.70, 129.75, 127.62,
127.09, 125.20, 124.99, 124.19, 122.05, 120.13, 112.88, 65.64, 46.77, 46.39, 44.39, 44.36, 38.26.
HRMS (ESI-TOF) m/z 621.2399 (calc’d [M + H]+ = 621.2344).
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PNA oligomer synthesis
PNA was synthesized on a Biotage SP Wave semi-automatic peptide synthesizer.
Synthesis began by down-loading 50mg of a rink amide MBHA resin (0.52 mmol/g) with 5 µmols
of the first Fmoc PNA monomer or Fmoc-Lys(Boc)-OH using HATU (1.2 eq), DIPEA (1.2 eq), and
2,6-lutidine (1.2 eq) in 200 µL dry NMP for 1 hour at room temperature followed by 1 hour of
capping using a solution of 9% acetic anhydride/13% 2,6-lutidine in DMF. Successive couplings
were performed using microwave assistance at 75 °C for 6 min with Fmoc PNA monomer (5 eq),
HATU (5eq), DIPEA (5 eq) and 2,6-lutidine (5 eq) in 400 µL dry NMP. After coupling, capping
(2x5 min with 1mL capping solution), washing (3x1.1 mL DMF, 3x1.1 mL DCM, then 3x1.1 mL
DMF), deprotection (3x2 min with 1mL 25% piperidine/DMF), and washing (same as previous)
completed a coupling cycle. All steps were monitored for completion by Kaiser test. Monomer
coupling efficiency was monitored by absorbance at 301 nm of the dibenzofulvene-piperidine
adduct using a Nanodrop 2000 spectrophotometer. Upon completion of synthesis, cleavage was
performed twice using 500 µL of cleavage solution (95% TFA/2.5% triisopropylsilane/2.5% H2O)
for 1 h. The crude oligomer was collected by ether precipitation and purified by reverse-phase
HPLC using an Agilent Eclipse XDB-C18 5 µm, 9.4 x 250 mm column at 60°C with a flow rate of
2 mL/min, monitored at 260 nm using a linear gradient (10-40%) of 0.1% TFA/acetonitrile in 0.1%
TFA/water. Identity was confirmed by ESI-TOF mass spectrometry.
Melting Temperature Analysis of PNA Strands
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Samples containing 5 M PNA and 5 M complementary DNA or miRNA-21 were
prepared from stock solutions in 1x PBS. The samples were transferred to an 8-cell quartz
microcuvette with a 1 cm pathlength. A Shimadzu UV-1800 spectrophotometer equipped with a
temperature controller and Julabo CORIO CD water circulator was used for the measurement.
Absorbance was monitored at 260 nm from 25-95 C at a rate of 0.5 C/min. Melting temperatures
were calculated by the first derivative method using a total of three independent trials.
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Preparation of PNA amphiphile assemblies
In order to form uniform assemblies, samples of PNA dissolved in water or 1x PBS were
heated to 95 C for 30 min. The samples were then cooled to room temperature and incubated
for at least 1 h before analysis.
PNA amphiphile CMC determination using internal 4-DMN monomer
The critical micelle concentration (CMC) of PNA-A was determined by the fluorescence
response of the internal 4-DMN monomer. Samples of PNA-A, control PNA-C, and fully
deprotected 4-DMN monomer ranging in concentration from 125 µM–0.0.061 µM in
1%DMSO/H2O were prepared from stock solutions. DMSO was used to initially dissolve the
monomer, then stock solutions were prepared in H2O to a final DMSO content of 1% in each
sample. Each sample was treated for amphiphile assembly as previously stated. Samples were
then transferred to a 384-well clear-bottom plate for analysis on a Biotek Cytation5 plate reader
(higher concentrations) or to a 50 L quartz cuvette for analysis by a Horiba Scientific Dual-FL
fluorometer (lower concentrations). Fluorescence measurements at 550 nm were collected using
an excitation of 458 nm. Absorbance was also measured at 458 nm to correct for any changes in
concentration between PNA samples and the monomer. A plot of the fluorescence enhancement
of the PNA samples over the monomer versus concentration was created using GraphPad Prism
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software and used to estimate the CMC. A semi-logarithmic fit to the data provided equations for
the upper and lower portions of the curve, which were then compared to determine a CMC value
of 317 nM.
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Characterization of assemblies by TEM
Samples of PNA at 1mM and 100 µM were prepared in H2O from stock solutions.
Assemblies were formed as previously stated. 3.5 µL of sample was spotted onto a 200-mesh
formvar/carbon-coated copper grid for 2 min before wicking with filter paper. 3.5 µL of a fresh 1%
uranyl acetate stain solution was then spotted onto the grids for 30 sec before wicking with filter
paper. The grids were allowed to dry at room temperature for 30 min before imaging using a
Hitachi HT7700 transmission electron microscope.
Size distribution determination from TEM
Images were analyzed using the NIH ImageJ software. Particle diameters were measured
manually by setting the scale of the image to the scale bar. Lines were drawn and measured in
triplicate for each particle that had clear boundaries without overlap in order to accurately
determine an average diameter for each particle. A total of 128 particles were measured and a
histogram of the data using 12 bins for size was produced using the GraphPad Prism software.
Determination of size using dynamic light scattering
Samples of PNA were prepared in water to a concentration of 500 µM. The samples were
heated to 95C for 30 min, followed by sonication at 50C for 10 min and incubation at RT for 1
hour to create uniform assemblies. Sizes were measured using a Particulate Systems NanoPlus
DLS nano particle analyzer and automically generated by the NanoPlus software.
Confirmation of binding of complementary DNA to PNA-A by CD
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DNA and RNA samples were purchased from Integrated DNA Technologies. Circular
dichroism was used to confirm binding and therefore disruption of PNA amphiphile assemblies.
Samples of PNA were prepared at 200 µM in 1x PBS. Assemblies were formed as previously
stated. DNA stocks at concentrations of 100 µM, 200 µM, 300 µM, and 400 µM were prepared in
1x PBS. 30 µL of PNA was mixed with 30 µL of DNA stocks to a final concentration of 100 µM
PNA and DNA concentrations of 50 µM, 100 µM, 150 µM, and 200 µM in 1x PBS. Samples were
incubated for 1 h before being analyzed by a JASCO J-1500 circular dichroism spectrometer.
Data points were collected every 1 nm from 190-300 nm using a continuous scanning mode of
200 nm/min at 23.5C. Samples containing a scrambled DNA sequence were prepared in the
same manner and analyzed simultaneously.
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Confirmation of binding of miRNA-21 to PNA-A by CD
Samples were prepared similarly to analysis by DNA. PNA-A samples of 200 µM in water
were prepared as previously described. A 200 µM stock of miRNA-21 was added to the samples
in water to a final concentration of 100 µM PNA and 100 µM miRNA-21. The samples were
incubated for 1 h before being analyzed.
PNA amphiphile disassembly by 4-DMN fluorescence
Samples of PNA-A at 11.11 M in 1.1xPBS were prepared and the assemblies formed as
previously stated. Complementary and scrambled DNA stocks ranging from 25 M–200 M were
prepared and 5 L of each was added to the appropriate PNA sample to a final concentration of
10 M PNA and 2.5 M-20 M DNA in 1xPBS. After addition, samples were incubated for 1 hour
before being transferred to a 384-well plate and analyzed using a Biotek Cytation5 plate reader.
Fluorescence measurements at 550 nm were collected using an excitation of 458 nm.
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PNA amphiphile disassembly by complementary DNA on TEM
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Samples of 400 µM PNA-A in 1x PBS were prepared from stock solutions. Assemblies
were formed as previously stated. To the samples was added complementary DNA or a
scrambled DNA sequence in 1x PBS for a final concentration of 200 µM PNA-A and 200 µM DNA.
Samples were incubated at room temperature for 1 h before preparation for TEM using the same
procedure described above.
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PNA amphiphile disassembly by complementary RNA on TEM
Samples of 200 µM PNA-A in water were prepared from stock solutions. Assemblies were
formed as previously stated. To the samples was added a 200 µM stock of miRNA-21 in water to
a final concentration of 100 µM PNA-A and 100 µM miRNA-21. The samples were incubated at
room temperature for 1 h before preparation for TEM.
Supporting Information
Supporting information available: characterization of monomers and PNA sequences and
additional TEM images.
Acknowledgements
This work was supported by the National Science Foundation (DMR 1822262 J.M.H.). The
authors also acknowledge the Robert P. Apkarian Integrated Electron Microscopy Core and NMR
Research Center at Emory University for access to instruments and technical assistance, and Mr.
Steve Knutson for helpful input on the writing of this manuscript.
Declaration of Interests:
The authors declare no competing interests.
References:
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