Submitted date: 12/12/2018 • Posted date: 13/12/2018 Licence: CC
BY-NC-ND 4.0 Citation information: Marras, Alexander; Vieregg,
Jeffrey; Ting, Jeffrey; Rubien, Jack D.; Tirrell, Matthew (2018):
Polyelectrolyte Complexation of Oligonucleotides by Charged
Hydrophobic – Neutral Hydrophilic Block Polymers. ChemRxiv.
Preprint.
Polyelectrolyte complex micelles (PCMs, core-shell nanoparticles
formed by complexation of a polyelectrolyte with a
polyelectrolyte-hydrophilic neutral block polymer) offer an
attractive solution to the critical problem of delivering
therapeutic nucleic acids, but few structure-property studies have
been carried out to date. We present data comparing oligonucleotide
PCMs formed with poly(vinylbenzyl trimethylammonium) as the
cationic block to those using poly(lysine), which is more commonly
used. Despite its higher charge density, increased hydrophobicity,
and permanent charge, pVBTMA appears to complex DNA more weakly
than does poly(lysine). Using small angle X-ray scattering and
electron microscopy, we find that, at physiological ionic strength,
PCMs formed from both cationic blocks exhibit very similar
structure-property relationships, with PCM radius determined by the
cationic block size and shape controlled by the hybridization state
of the oligonucleotides. These observations narrow the design space
for optimizing therapeutic PCMs and provide new insights into the
rich polymer physics of polyelectrolyte self-assembly.
File list (2)
Alexander E. Marras1, Jeffrey R. Vieregg1*, Jeffrey M.
Ting1,2,
Jack D. Rubien3, and Matthew V. Tirrell1,2
1 Institute for Molecular Engineering, University of Chicago,
Chicago, IL, 60605
2 Institute for Molecular Engineering at Argonne National
Laboratory, Lemont, IL, 60439 3 Departments of Biology and Physics,
Swarthmore College, Swarthmore, PA 19081
* Correspondence:
[email protected]
Abstract
Polyelectrolyte complex micelles (PCMs, core-shell nanoparticles
formed by complexation of a polyelectrolyte with a
polyelectrolyte-hydrophilic neutral block polymer) offer an
attractive solution to the critical problem of delivering
therapeutic nucleic acids. Nucleic acid PCMs with hydrophilic
cationic blocks have produced promising results in vitro and in
animal models. We recently published a set of structure- property
relationships for poly(lysine)-poly(ethylene glycol) –
oligonucleotide PCMs that enables rational design of micelles with
desired size and shape, but significant work remains to optimize
this promising technology. Few systematic studies have been
conducted on how polycation properties such as charge density,
hydrophobicity, and choice of charged group influence PCM
properties, despite evidence that these strongly influence the
complexation behavior of polyelectrolyte homopolymers. In this
article, we report a comparison of oligonucleotide PCMs and
polyelectrolyte complexes formed by poly(lysine) and
poly((vinylbenzyl) – trimethylammonium) (pVBTMA), a styrenic
polycation with comparatively higher charge density, increased
hydrophobicity, and a permanent positive charge due to its
quaternary amine. All of these differences have been individually
suggested to provide increased complex stability, but we find that
pVBTMA in fact complexes oligonucleotides more weakly than does
poly(lysine), measured by stability as a function of monovalent
salt. Using small angle X-ray scattering and electron microscopy,
we find that, at physiological ionic strength, PCMs formed from
both cationic blocks exhibit very similar structure-property
relationships, with PCM radius determined by the cationic block
size and shape controlled by the hybridization state of the
oligonucleotides. These observations narrow the design space for
optimizing therapeutic PCMs and provide new insights into the rich
polymer physics of polyelectrolyte self-assembly.
Introduction Developing effective non-viral methods for delivery of
nucleic acids and other macromolecular therapeutics is one of the
most pressing challenges for nanomedicine and polymer science
[1-4]. The potential power of engineered nucleic acids as
therapeutic agents is severely limited by the difficulty of
overcoming the physical and biological barriers to using them as
practical drugs. DNA and RNA molecules’ large size, hydrophilicity,
and negative charge largely prevent them from crossing cell
membranes and promote their rapid clearance from circulation.
Exogeneous nucleic acids are also readily degraded by cellular and
serum nucleases and are potent activators of the innate immune
system. As a result, therapeutic applications to date have required
extensive chemical modification and/or encapsulation of the nucleic
acids, most commonly by liposomes and other lipid nanoparticles
assembled by hydrophobic interactions [5-8]. These approaches have
demonstrated the effectiveness of nucleic acid therapeutics but
come with significant drawbacks, including toxicity,
immunogenicity, and most particularly, limited biodistribution. In
circulation, lipids are rapidly complexed by apolipoproteins and
routed to the liver for metabolism. As a result, nucleic acid drugs
to date have either been limited to liver targets or delivered
locally. This fundamental limitation suggests the need for
alternative strategies for nanoparticle self-assembly; one of the
most promising being polyelectrolyte complexation.
Polyelectrolyte complexation describes the preference for
oppositely-charged macroions to associate with each other in
aqueous solution rather than with small counterions, due to their
lower translational entropy per unit charge [5]. If the attraction
is strong enough, this leads to phase separation despite all
components (usually polymers, but also charged particles, as
studied by Paul Dubin and others [6]) being individually
solvophilic. The resulting polymer-rich phase can either be liquid
(complex coacervate) or a solid precipitate, and the factors that
determine which one is formed remain largely unknown despite many
years of study. We also lack a quantitative ability to predict how
molecular properties such as charge density, charge patterning,
chirality, hydrophobicity, and hydrogen bonding propensity
determine the boundaries of phase separation and the properties of
the resulting complex phase. Despite this, complex coacervates and
precipitates are widely used in industry and have gained increasing
attention as vehicles for drug delivery [5,7].
Nucleic acids are strongly-charged polyanions, and phase-separated
complexes have been observed when DNA molecules (ranging in length
from as long as entire chromosomes to as short as individual
nucleotides) are mixed with cationic polymers [8,9]. Complexation
neutralizes the nucleic acids’ charge, and the resulting complexes
(sometimes termed ‘polyplexes’) can be internalized by cells via
endocytosis. The cationic polymers poly(lysine) and
poly(ethyleneimine) are widely used for gene transfection in
vitro and are effective, although toxicity and immunogenicity can
become a problem as polymer length increases [10,11]. More
importantly, however, the resulting complexes lack colloidal
stability in circulation, largely limiting them to local
applications in vivo.
If the polycation is conjugated to a neutral hydrophilic polymer
such as poly(ethylene glycol) (PEG), nanoparticles are produced
instead of macrophase separation: the hydrophilic neutral block
forms a corona around a neutralized polyion core (Figure 1). This
is visually reminiscent of surfactant micellization, and the
resulting nanoparticles are referred to as polyelectrolyte complex
micelles (PCMs, also referred to as polyion complex micelles, block
ionomer complexes, and coacervate-core micelles), though the forces
driving self-assembly are ionic rather than hydrophobic [12].
Conceptually, PCMs assembled with nucleic acids as the polyanions
are attractive delivery vehicles: in addition to the charge
neutralization and steric protection from nucleases afforded by the
polyion core, the neutral corona provides colloidal stability and
size control to allow optimization of circulation properties, as
well as a platform for attaching targeting ligands to further
improve biodistribution [2]. Assembly of multiple oligonucleotides
in each PCM also increases the potency of each cell internalization
event, and PCM formation does not require extensive chemical
modification of the nucleic acids, preserving biological function.
Several promising results in vitro and in small animal models
confirm the potential of this strategy [2,13-15], but much work
remains to optimize PCMs as safe, efficacious nucleic acid delivery
vehicles, as well as to improve our understanding of the physics of
PCM self-assembly.
We recently investigated structure-property relationships for PCMs
formed from DNA oligonucleotides and poly(lysine)-PEG block
polymers, which are by far the most common choice for
oligonucleotide delivery [16]. Over a wide range of polymer and
block lengths, we found that the PCM core radius is determined
solely by the length of the charged block and is independent of
both the length and hybridization state (single- vs
double-stranded) of the oligonucleotides. Interestingly, however,
we found that oligonucleotide hybridization had a large effect on
the shape of the nanoparticles, with single-stranded
oligonucleotides forming spheroidal micelles and double-stranded
oligonucleotides forming long wormlike micelles, apparently via
coaxial stacking of the DNA helices. Small-angle X-ray scattering
(SAXS) also revealed parallel packing of DNA helices inside the PCM
cores that had previously only been observed for condensed
genomic-scale DNA. These results provide design rules for
constructing oligonucleotide PCMs of desired size and shape, with
exceptionally low polydispersity, but do not address the question
of whether poly(lysine) is an optimal cationic polymer to use for
oligonucleotide delivery.
Figure 1. Polyelectrolyte complexation and PCM nanoparticle
formation. DNA is a highly charged polyanion. When mixed with
polycations (right), polyelectrolyte complexes are formed
(macrophase separation). When mixed with cation- hydrophilic
neutral block polymers (left), microphase separation produces PCMs.
In both scenarios, the hybridization state of the nucleic acid
(single- vs double-stranded) determines the nature of the product
(liquid droplets vs solid precipitates for the complexes,
spheroidal vs cylindrical PCMs). This study compares the effect of
hydrophilic (poly(lysine), pLys) and hydrophobic
(poly((vinylbenzyl) trimethylammonium), pVBTMA) polycations in
determing the properties of the complexes and PCMs. This study
describes an experimental comparison of the styrenic polycation
poly((vinylbenzyl) trimethylammonium) and poly(lysine) (pVBTMA and
pLys, Scheme 1) as polycations for assembling PCMs and
polyelectrolyte complexes with DNA. pVBTMA and pVBTMA-PEG block
polymers are readily accessible via aqueous reversible
addition-fragmentation chain transfer (RAFT) polymerization [17]
and differ from pLys in several respects that might be expected to
influence their complexation behavior. pVBTMA has a higher linear
charge density (2 backbone atoms per repeat unit vs 3), and its
aliphatic backbone, aromatic side chain, and minimal propensity for
hydrogen bonding make it much more hydrophobic than pLys. pVBTMA’s
quaternary ammonium is also permanently charged compared to the
primary amine of pLys, which could potentially be deprotonated in
the dense environment of the complexes. These differences make for
a stringent test of the universality of the design rules we derived
for pLys-PEG oligonucleotide PCMs. Additionally, all of these
factors have been linked to increased complex stability, which
might be expected to improve nuclease resistance and
Single-stranded DNA
Double-stranded DNA
Micelles
Spheroidal
Complexes
Precipitates
circulation time in the therapeutic setting [18]. As in our
previous work, we utilize a multi-modal characterization strategy
in which light scattering, SAXS, and light and electron microscopy
are used together to provide a more complete picture of the
complexes and PCMs than could be obtained from any individual
technique.
Scheme 1. Chemical structures of homo- and block polymers used in
this study. pLys(A,B) is cationic and hydrophilic, where pVBTMA
(D,E) is also cationic but hydrophobic. PEG blocks (B,E) are
neutral and hydrophilic. These polymers complex with DNA
oligonucleotides (C), which are anionic and hydrophilic, to form
phase- separated assemblies.
Materials and Methods
RAFT Synthesis of pVBTMA and pVBTMA-PEG. pVBTMA homopolymers and
pVBTMA-PEG block polymers were synthesized by aqueous reversible
addition- fragmentation chain transfer (RAFT) polymerization as
described by Ting et al [17]. A detailed description is included in
Supplementary Material section S1. Briefly, pVBTMA homopolymers
were synthesized with 35 and 172 repeat units using (vinylbenzyl)
trimethylammonium chloride monomer,
4-cyano-4-(phenylcarbonothioylthio) pentanoic acid chain transfer
agent (CPhPA, Scheme 1D) and VA-044 thermal initiator in degassed
acetate buffer solution/ethanol (3:1 v/v). pVBTMA-PEG block
polymers were synthesized using trithiocarbonyl macro-CTAs
containing PEG blocks of 1k, 5k and 10k MW (Scheme 1E) using the
same initiator and buffer. Monomer conversion was assessed by 1H
NMR spectroscopy; methods and representative spectra are shown in
Figures S1 – S5. Polymers
(E) pVBTMA-PEG
SS C12H25
O
p
were extensively dialyzed against NaCl, then water, and lyophilized
to provide the chloride salt as a free-flowing powder. Lengths and
molecular weights for all the polymers are shown in Table S1.
Polymer characterization. The absolute molecular weight of the
pVBTMA homo- and block polymers was determined by size-exclusion
chromatography with multi-angle light scattering (SEC-MALS) using a
Waters SEC instrument equipped with three columns (Waters
Ultrahydrogel 500, 250, 120), a diode array detector (Waters 2998),
an Optilab T-rEX (Wyatt) refractive index detector, and a miniDAWN
TREOS II (Wyatt). For all the polymers except the
pVBTMA(10)-PEG(5k), the mobile phase was acetonitrile : water (39.9
: 60 v/v) with 0.1% trifluoroacetic acid and the flow rate was 1.0
mL/min. Due to its short length, pVBTMA(8)-PEG(5k) was run in
aqueous 0.1 M NaNO3 with 0.1% (v/v) NaN3. Polymer dn/dc values were
determined using an Abbe refractometer with a red light-emitting
diode at 25 °C; measured refractive index values at increasing
polymer concentrations were collected in triplicate and fitted by
linear regression (Figure S3).
DNA and poly(lysine) polymers. DNA oligonucleotide sequences (Table
S2) were designed for minimal self-complementarity and internal
structure formation using the NUPACK software tool [19].
Oligonucleotides were ordered from Integrated DNA Technologies and
used without further purification. Sheared salmon sperm DNA
(average length 2000 bp) was purchased from Invitrogen. DNA
solutions were resuspended in water at 20 mM charge concentration
(mols phosphate / L) prior to use. pLys and pLys-PEG polymers were
purchased from Alamanda Polymers as chloride salts and were
neutralized with NaOH and resuspended in water at 10 mM charge
concentration prior to use.
Micelle preparation. PCMs were prepared using the salt-annealing
method described by Lueckheide et al [16]. Briefly, the
polyelectrolytes (2mM final charge concentration) were mixed in PBS
buffer (pH 7.4), then concentrated NaCl solution was added to
obtain 1M final concentration (400 µL total solution volume) to
dissolve the complexes. The salt concentration was then slowly
reduced over 36 hours by step dialysis with a 2000 MWCO membrane
(Slide-a-lyzer G2, ThermoFisher) to a final working concentration
of 1x PBS (155mM NaCl, 1mM KH2PO4, 3mM Na2HPO4-7H2O, pH 7.4).
Small-Angle X-Ray Scattering. SAXS measurements were made at
beamline 12- ID-B of the Advanced Photon Source at Argonne National
Laboratory. Micelle samples were irradiated in a thin-wall glass
capillary flow cell with a photon energy of 14 keV. Data was
reduced and background was subtracted as described in Ref. [16].
Fitting was performed using the multi-level modeling macros
distributed with the Irena software package [20] for Igor Pro as
described in the same reference.
Electron Microscopy. Cryo Transmission Electron Microscopy (TEM)
samples were flash frozen onto lacey carbon film grids (LC200-CU,
Electron Microscopy Sciences)
and imaged on a FEI Talos TEM at an acceleration voltage of 200kV.
Negative-stained samples were deposited on carbon coated square
grids (CF200-Cu-UL, Electron Microscopy Sciences), dried, and
stained with 2% uranyl formate, and imaged on a FEI Tecnai G2
Spirit TEM at an acceleration voltage of 120 kV.
Micelle salt dependence. To assess the stability of the micelles vs
salt, 200 µL micelle samples were prepared as described above and
titrated with 5M NaCl. Light scattering intensity was measured at a
90-degree angle using a Brookhaven Instruments BI-200SM system with
a 637 nm laser at room temperature. Critical salt concentrations
were classified as the point where no structure is seen in the
autocorrelation function and scattering intensity drops below 5000
counts per second.
Homopolymer complex preparation. Polyelectrolyte complexes were
prepared at pH 7 and room temperature. Double-stranded DNA was
prepared by annealing complementary strands at 65 °C for 5 minutes
followed by slow cooling to RT. 18.2 M water, concentrated PBS, and
NaCl (when applicable) solutions were mixed, followed by addition
of the DNA and then the polycation for a final concentration of 2mM
charge concentration of polyelectrolytes and 1x PBS. Samples were
mixed thoroughly after addition of each polyelectrolyte. Aliquots
were prepared separately at the indicated NaCl
concentrations.
Optical Microscopy. Phase and morphology of the homopolymer
complexes were observed by bright field optical microscopy using a
Leica DMI-6000B inverted microscope with white light illumination
and 10-20X magnification. 100 µL aliquots of the complex
suspensions were placed in ultra-low attachment 96 well plates
(Costar, Corning). Images were taken shortly after mixing and then
again 4 hours later, with the latter used unless noted to the
contrary.
Results
Polyelectrolyte Complex Micelle Formation and Morphology. In order
to evaluate pVBTMA as a cationic polymer for nucleic acid delivery,
we first prepared PCMs using pVBTMA-PEG block polymers and
single-stranded DNA (ssDNA) oligonucleotides and compared them to
PCMs assembled from pLys block polymers of similar lengths using
SAXS and electron microscopy (EM). None of the polymers formed
detectable structures on their own (Figure S6). The block polymer
with the shortest pVBTMA polycation (pVBTMA(8)-PEG(5k) also did not
phase separate when mixed with DNA, and block polymers with the
shortest PEG lengths (1k MW) formed large aggregates rather than
nanoparticles. In all other cases, however, both cationic polymers
readily formed PCMs when mixed with DNA under these conditions,
with low polydispersity in radius. We were able to accurately fit
the SAXS scattering profiles with a combination of hard-surface
form factor, power-law, and diffraction peak models
(Figure S7). EM imaging (cryo and conventional) also confirms PCM
formation with both cationic polymers.
The low-q region of the SAXS scattering profiles provides
information on PCM size and shape, and numerous similarities are
observed for pLys and pVBTMA micelles. With both polymers,
spheroidal micelles are observed for PCMs containing single-
stranded oligonucleotides, as shown by the flat (q0) scattering
intensity in the low-q region of the SAXS data and corroborated by
the EM images (Figures 2A-B). Fitting shows that the micelle radii
are similar for both polymers (Figure 4). As previously observed
with pLys, we saw no dependence of PCM radius on oligonucleotide
length over a 9-fold range in the latter, but a marked increase in
radius with cationic block length (Figure 4). These results, which
extend over a larger range of block lengths than our previous work,
suggest that the principles governing PCM formation apply over a
wide range of polymer structures and chemical properties. Direct
modification of the pVBTMA chain end- groups in
otherwise-equivalent systems also impacts the micelle particle size
(Figure S8 and Table S3). Additional EM and SAXS data and fits are
available in Tables S4-S7 and Figures S9-S19.
For double-stranded DNA (dsDNA), we observe long, wormlike micelles
with the pLys-PEG block polymers, consistent with our previous
report (Figure 3A), but in this case we see a qualitative
difference for PCMs prepared with pVBTMA-PEG. The micelles exhibit
low polydispersity in radius, with fitted values very similar to
those found for pLys-PEG PCMs (Table 1), but the lengths are
shorter and highly variable between and within samples (Figure
3B,C). This leads to power-law dependencies of the SAXS scattering
intensity in the low-q region that are not consistent with
spheroidal (q0), rigid cylinder (q1), or flexible cylinder (q2)
form factor models (Table 1). Accordingly, we did not attempt to
fit the lengths of these micelles and used a sphere form factor
model with a low-q cutoff to determine the radius alone (Figure 3D,
S19). Just as with single-stranded DNA, we see minimal dependence
of micelle radius on DNA length (Figure 4), even for sheared salmon
sperm DNA (mean length ~2000 bp), which we included as an analog
for plasmids and other full-length gene constructs. Also, similarly
to the ssDNA case, we observe a marked increase in PCM radius with
charged block length (Table 1, Figure 4).
Figure 2. Characterization of single-stranded oligonucleotide
polyelectrolyte complex micelles. pLys-PEG and pVBTMA-PEG both form
spheroidal micelles with ssDNA in all cases tested. Examples here
show cryo electron micrographs of pLys(200)-PEG(10k) with 88 nt DNA
(A) and pVBTMA(72)-PEG(10k) with 88 nt DNA (B). Small-angle X-ray
scattering intensity profiles and fits provide further structural
information, as shown with four examples in (C) with fitting
results shown in the table. Fit results for all ssDNA PCMs are
shown in Tables S4-S5. Scattering curves are vertically offset for
clarity. Scale bars = 50 nm.
The higher-q region (q 0.08 -1) of the SAXS data provides
information about the internal structure of the micelles.
Consistent with our previous observations, the ssDNA PCMs showed,
with few exceptions, Porod exponents near 2 regardless of whether
pLys or pVBTMA was used as the cationic blocks. This indicates that
the polymers are behaving like ideal (i.e. uncharged) chains and is
consistent with idea of a liquid-like PCM core for these micelles.
For dsDNA PCMs formed with pLys-PEG, we previously
showed that a prominent diffraction peak at q ≈ 0.23 -1 resulted
from parallel, hexagonal packing of dsDNA helices within the PCM
cores very similar to those observed in toroidal condensates of
genomic DNA [16]. Though shifted to lower q and broadened (Figure
3D), a similar diffraction peak is observed for many pVBTMA-PEG
PCMs, implying similar, though perhaps less regular, ordering
within these micelles.
Figure 3. Characterization of double-stranded oligonucleotide
polyelectrolyte complex micelles. A discrepancy in length and shape
is seen with dsDNA PCMs. pLys-PEG forms long flexible cylinders
(A), while pVBTMA-PEG forms cylindrical or spheroidal PCMs with
polydisperse lengths (B, C). (D) shows representative SAXS data for
both systems with fitting results available in Table 1. Panel A is
pLys(50)-PEG(5k) + 88 bp DNA, (B) is pVBTMA(194)-PEG(10k) + 10 bp
DNA, and (C) is pVBTMA(24)-PEG(5k) + 88 bp. (A, C) negative-stain
conventional EM and (B) is a cryo-EM micrograph. Scale bars = 50 nm
unless noted. PCM Stability. The primary driving force for
polyelectrolyte complexation is entropy gain from counterion
release [21]. Increasing the background ionic strength decreases
this, eventually resulting in dissolution of the complexes. The
critical ionic strength required for dissolution therefore provides
a measure of the stability of a given complex [22]. To compare the
stability of pVBTMA-PEG vs. pLys-PEG PCMs, we titrated concentrated
NaCl into PCMs prepared at physiological ionic strength while
monitoring light scattering intensity. As shown in Figure 5, PCMs
containing pVBTMA-PEG are considerably less stable than those
containing pLys-PEG: pLys-PEG PCMs are stable up to 600-700 mM NaCl
compared to pVBTMA-PEG PCMs which no longer form micelles above
300-400 mM.
Table 1: Representative SAXS fit parameters for dsDNA PCMs. All
PCMs show low polydispersity in radius, but the pVBTMA-PEG PCMs
exhibit intermediate scaling laws at low-q indicating significant
polydispersity in length. pLys-PEG data are fit using flexible
cylinder form factors (plus power law and diffraction peak at
higher q) while pVBTMA-PEG data is fit using a sphere form factor
cut off at low-q. Fit results for all dsDNA PCMs are shown in
Tables S6-S7.
Sample Mean Radius (nm)
Low-q power law
pVBTMA(194)-PEG(10k) + ds10 31.3 0.02 No 0.65 pVBTMA(194)-PEG(10k)
+ ds22 30.3 0.04 Yes 1.59 pVBTMA(53)-PEG(5k) + ds22 12.1 0.09 Yes
1.92 pVBTMA(53)-PEG(5k) + ds88 11.0 0.05 Yes 1.13
pLys(200)-PEG(10k) + ds10 19.9 0.03 Yes pLys(200)-PEG(10k) + ds22
16.3 0.01 Yes pLys(50)-PEG(5k) + ds22 8.8 0.03 Yes pLys(50)-PEG(5k)
+ ds88 11.0 0.03 Yes
Figure 4. PCM radii from SAXS fits vs polyelectrolyte length. For
both cations, PCM radius is largely insensitive to nucleic acid
length but increases strongly with the length of the cation blocks.
Data tables are shown in Supplementary Materials.
Figure 5: pVBTMA PCMs are much less resistant to added salt
compared to pLys PCMs. Critical salt concentration (measured by
drop in light scattering intensity) is plotted vs. cation identity
and length.
Homopolymer complexation. In order to more fully understand the
complexation behavior of pVBTMA and DNA, we next compared complexes
formed by pVBTMA/pLys homopolymers (no neutral block) and DNA
oligonucleotides over a similar range of lengths as used in the PCM
studies. We have previously observed that, under these conditions,
pLys and DNA readily phase separate, forming either liquid droplets
(coacervates) or solid precipitates depending on whether the DNA is
single- or double- stranded [23]. As shown in Figure 6, pVBTMA
shows the same phase behavior when complexed with long DNA (88
nt/bp and above), but is significantly less effective at forming
phase-separated complexes with shorter oligonucleotides. (35)mer
pVBTMA does not phase separate at all with 22 nt ssDNA, and both
(35) and (174)mer pVBTMA form coacervates rather than precipitates
when mixed with shorter double-stranded oligonucleotides. Overall,
pVBTMA requires longer polyelectrolyte lengths for phase separation
to occur compared to pLys.
Finally, we measured the effect of increasing ionic strength on the
pVBTMA – DNA homopolymer complexes and compared this to our
previous results for pLys – DNA complexes[23]. Qualitatively, we
observe similar trends for both polycations (Figure 7). As salt
concentration increases, solid precipitates (for double-stranded
DNA complexes, representative data for pVBTMA(35) – dsDNA(22) at
150 mM NaCl shown in panel A) melt into coacervates (panel B, 400
mM NaCl), then dissociate into solution. Complexes formed from
longer polymers are more stable than those formed from shorter
ones. In each case, however, the phase transitions (solid – liquid,
liquid – soluble) occur at much lower salt concentrations for
pVBTMA than for pLys, with physiological ionic
strength (PBS buffer) sufficient to melt the pVBTMA(35) – dsDNA(22)
complexes into coacervates. The critical ionic strengths required
to dissolve the homopolymer complexes are also quite close to those
measured for the PCMs (Figure 5), implying that the geometric
constraints imposed by microphase separation do not significantly
decrease the stability of the polyelectrolyte complexes in the
core.
Figure 6: Phase diagrams for ssDNA (A,C) and dsDNA (B,D) complexing
with homopolymers pLys (A-B) and pVBTMA (C-D). pVBTMA requires
longer oligonucleotides for phase separation to occur, suggesting a
less stable interaction with DNA. E) Representative optical
micrograph of coacervate complexes (22-nt ssDNA + 50- aa pLys). F)
Representative optical micrograph of precipitate complexes (88-bp
dsDNA + 53mer pVBTMA).
Figure 7: Phase transitions of homopolymer complexes with
increasing salt concentration. As ionic strength increases,
precipitates (A, pVBTMA(35) – dsDNA(22)) melt into coacervate
droplets (B) before eventually dissolving. Scale bars are 50 µm.
Critical ionic strengths for melting (green to orange) and
dissolution are plotted in panel C for four different combinations
of polyelectrolyte lengths. At each length combination, phase
transitions occur at lower ionic strength for pVBTMA than for pLys.
Discussion
These results provide, to our knowledge, the first systematic
characterization of pVBTMA’s ability to complex nucleic acids;
previous studies had been limited to single- angle DLS measurements
of complexes made with highly-polydisperse sheared genomic DNA
[24]. This reflects a more general trend in the field: with only
few exceptions [25,26], investigations of polycations for nucleic
acid delivery have focused on hydrophilic polymers such as pLys,
poly(ethyleneimine), poly(amidoamine), and cationic polysaccharides
[10,12], many of which are analogs of natural products. This may
represent a missed opportunity, as synthetic polymers provide
access to diverse structural motifs, many of which are
bio-orthogonal, and allow tuning of intermolecular interactions
(hydrophobicity, hydrogen bonding, cation-pi, etc.) over a wider
range than is possible with naturally-inspired polycations. Given
the diverse range of interactions available with nucleic acids
(charge interactions through the phosphate anion, pi-pi and
cation-pi interactions with the nucleobases, hydrogen bonding from
the sugar and nucleobase heteroatoms, and more), it seems
reasonable to assume that synthetic polymers may provide more
optimal vehicles for therapeutic delivery. This diversity, however,
raises an important question: to what degree can lessons learned
with one polycation be transferred to another? This study was
designed to provide a stringent test of that proposition; as
discussed earlier, pVBTMA has a 50 percent larger linear
charge
density than pLys, has an aliphatic backbone and aromatic side
chains, is largely unable to form hydrogen bonds, and contains a
permanent positive charge rather than the ionizable primary
amine.
Remarkably, we find many more similarities than differences when
comparing PCMs formed by the two polycations. pLys-PEG and
pVBTMA-PEG both form micelles with low polydispersity in radius
when prepared via salt annealing, and the PCM radii are, in most
cases, quantitatively similar as well. (Figures 2-4; Table 1,
S4-7). We have previously reported that the radius of
oligonucleotide PCMs formed with pLys-PEG block polymers is
controlled by the length of the charged block but largely
insensitive to oligonucleotide length [16]. These results extend
this conclusion to larger block lengths (n ≈ 200) and show that the
same pattern is true for pVBTMA-PEG PCMs (Figure 4), suggesting
that this may be a universal property for PCMs.
We also observe that, below critical lengths for both the charged
and neutral blocks, the complexes aggregate rather than forming
nanoparticles. Similar results have been reported for PCMs formed
from synthetic acrylate-based polyelectrolytes and block polymers,
though those authors discussed it in terms of core-corona ratio
[27]. While this requirement is likely general, determining the
exact criteria for colloidal stability and whether a quantitative
analog to the ‘packing parameter’ that describes hydrophobic
micellization exists for PCMs remains an open question.
We have previously seen that the hybridization state of the nucleic
acid is a key parameter in determining the behavior of
polylectrolyte complexes and PCMs assembled using pLys as the
polycation [16,23]. Single-stranded oligonucleotides produce liquid
complexes and spheroidal micelles, while double-stranded nucleic
acids produce solid complexes and long cylindrical micelles with
substantial internal ordering in the form of hexagonally-packed and
coaxially-stacked DNA helices. With both SAXS and EM, we observe
similar, though not identical behavior when pVBTMA is used as the
polycation. Single-stranded pVBTMA-PEG PCMs are spheroidal, and
double-stranded PCMs form cylinders, but the lengths of the latter
are much shorter than seen with pLys and are highly polydisperse. A
similar pattern is found for the homopolymers: liquids are observed
for ssDNA complexes, and solids are only seen with dsDNA, but the
short- polymer region where no phase separation is observed is
larger with pVBTMA, and liquids are also observed for complexes
containing short (10 and 22 bp) double-stranded
oligonucleotides.
One hypothesis that explains these differences is that the
interactions between DNA and pVBTMA are simply weaker than those
with pLys; this is strongly supported by the NaCl stability data,
which show that PCMs and homopolymer complexes containing pVBTMA
are significantly easier to melt and/or dissolve with salt than
those containing pLys. In this interpretation, the polydispersity
in length observed for the
dsDNA + pVBTMA-PEG PCMs results from some combination of the more
favorable entropy associated with a larger number of shorter PCMs
and mechanical instability of the micelles during experimental
manipulations. At the molecular level, this length dispersity
implies less efficient coaxial stacking of DNA helices, while the
observed broadening and shift of the packing diffraction peak to
lower q when pLys-PEG is replaced by pVBTMA-PEG reflects less
effective helical packing through a decrease in order and an
increase in average inter-helix spacing.
While this hypothesis appears to be consistent with our data, it
may also seem counterintuitive, as many of the structural
differences between pVBTMA and pLys (higher charge density,
increased hydrophobicity, non-susceptibility to neutralization)
have been individually shown or suggested to increase
polyelectrolyte complex stability [28-31]. We have also observed
that pVBTMA and polystyrene sulfonate (PSS) form polyelectrolyte
complexes that are extremely stable with respect to salt [32]. A
closer look at the structures and potential intermolecular
interactions of the polyelectrolytes suggests several solutions to
this apparent conflict. One possibility with some experimental
evidence is that, despite their permanent charge, the steric bulk
of quaternary amines limits their effectiveness for polyelectrolyte
complexation by enforcing a larger separation between opposing
charges than is attainable with smaller cations and counterions.
Schlenoff and coworkers recently reported that poly(allylamine) and
poly(vinylamine), which have primary amines as charged groups,
formed stronger complexes than did three quaternary
amine-containing polycations, including pVBTMA [33]. Similarly,
Izumrudov and coworkers found that pLys and several other primary
amine-containing polycations formed complexes with sheared genomic
DNA that were more resistant to salt than those formed with
quaternary amine-containing polycations [34]. Hydrophobic
interactions offer additional opportunities for stabilization, but
the disparate structures of DNA and pVBTMA may hinder short-range
interactions such as pi stacking that are favored in symmetrical
polyelectrolytes such as pVBTMA and PSS. Conjugation of hydrophobic
moieties to one end of the polyelectrolytes has been shown to
increase the serum stability of nucleic acid PCMs [18,35], and the
smaller radii we observe in PCMs with pVBTMA-PEG block polymers
containing C12 tails (Figure S8) highlights the potential of RAFT
polymerization to access these more complex architectures. A final
possibility is that pLys-DNA complexes could also be stabilized by
hydrogen bonding interactions that are not available to
pVBTMA.
While representing only a single pair of polycations, the large
structural differences between pLys and pVBTMA suggest that the
similarities in PCM properties that we observe may be universal,
while the differences provide useful leads for further
investigation into the physics of polyelectrolyte self-assembly as
well as possible solutions to the pressing problem of nucleic acid
delivery. The diversity of structures accessible in synthetic
polymers, as well as the capability of modern polymer
synthesis
techniques such as RAFT to access these in a relatively modular
manner, offers exciting new possibilities on both fronts. In
particular, systematic variation of charged groups, side chain
structures, and backbone architectures should be able to unravel
the effects of hydrophobicity, charge density and type, and
hydrogen bonding and enable rational design of optimal delivery
vehicles for therapeutic nucleic acids.
Author Contributions: Conceptualization: A.E.M and J.R.V. Polymer
synthesis and characterization: J.M.T. Data collection: A.E.M. and
J.D.R. Data analysis: A.E.M. and J.R.V. Research direction and
funding: M.V.T. All authors participated in interpreting the data
and drafting and editing the manuscript. Funding: This work was
supported by the U.S. Department of Energy Office of Science,
Program in Basic Energy Sciences, Materials Sciences and
Engineering Division and used resources of the Advanced Photon
Source, a U.S. Department of Energy (DOE) Office of Science User
Facility operated for the DOE Office of Science by Argonne National
Laboratory under Contract No. DE-AC02-06CH11357. J.M.T.
acknowledges support from the NIST-CHiMaD Postdoctoral Fellowship,
supported by the U.S. Department of Commerce, National Institute of
Standards and Technology (NIST) through the Center for Hierarchical
Materials Design (CHiMaD) under financial assistance award
70NANB14H012. Acknowledgments: Parts of this work were carried out
at the Soft Matter Characterization Facility and the Advanced
Electron Microscopy Facility of the University of Chicago. The
authors thank Dr. Xiaobing Zuo for his assistance with the SAXS
data acquisition and Dr. Tera Lavoie for assistance with electron
microscopy. Conflicts of Interest: The authors declare no conflict
of interest. References [1] B. Pelaz, C. Alexiou, R.A.
Alvarez-Puebla, F. Alves, A.M. Andrews, S. Ashraf, et
al., Diverse Applications of Nanomedicine, ACS Nano. 11 (2017)
2313–2381. doi:10.1021/acsnano.6b06040.
[2] H. Cabral, K. Miyata, K. Osada, K. Kataoka, Block Copolymer
Micelles in Nanomedicine Applications, Chem Rev. 118 (2018)
6844–6892. doi:10.1021/acs.chemrev.8b00199.
[3] R.L. Juliano, The delivery of therapeutic oligonucleotides,
Nucleic Acids Res. 44 (2016) 6518–6548.
doi:10.1093/nar/gkw236.
[4] S.F. Dowdy, Overcoming cellular barriers for RNA therapeutics,
Nat Biotech. 35 (2017) 222–229. doi:10.1038/nbt.3802.
[5] J. van der Gucht, E. Spruijt, M. Lemmers, M.A. Cohen Stuart,
Polyelectrolyte complexes: bulk phases and colloidal systems,
Journal of Colloid and Interface Science. 361 (2011) 407–422.
doi:10.1016/j.jcis.2011.05.080.
[6] E. Kizilay, A.B. Kayitmazer, P.L. Dubin, Complexation and
coacervation of polyelectrolytes with oppositely charged colloids,
Adv Colloid Interface Sci. 167 (2011) 24–37.
doi:10.1016/j.cis.2011.06.006.
[7] C.G. de Kruif, F. Weinbreck, R. de Vries, Complex coacervation
of proteins and anionic polysaccharides, Current Opinion in Colloid
& Interface Science. 9 (2004) 340–349.
doi:10.1016/j.cocis.2004.09.006.
[8] V.A. Bloomfield, DNA condensation by multivalent cations,
Biopolymers. 44 (1997) 269–282.
doi:10.1002/(SICI)1097-0282(1997)44:3<269::AID-BIP6>3.0.CO;2-
T.
[9] J.R. Vieregg, T.Y.D. Tang, Polynucleotides in cellular mimics:
Coacervates and lipid vesicles, Current Opinion in Colloid &
Interface Science. 26 (2016) 50–57.
doi:10.1016/j.cocis.2016.09.004.
[10] U. Lächelt, E. Wagner, Nucleic Acid Therapeutics Using
Polyplexes: A Journey of 50 Years (and Beyond), Chem Rev. 115
(2015) 11043–11078. doi:10.1021/cr5006793.
[11] A. Hall, U. Lächelt, J. Bartek, E. Wagner, S.M. Moghimi,
Polyplex Evolution: Understanding Biology, Optimizing Performance,
Mol Ther. 25 (2017) 1476– 1490.
doi:10.1016/j.ymthe.2017.01.024.
[12] I.K. Voets, A. de Keizer, M.A. Cohen Stuart, Complex
coacervate core micelles, Adv Colloid Interface Sci. 147-148 (2009)
300–318. doi:10.1016/j.cis.2008.09.012.
[13] R.J. Christie, Y. Matsumoto, K. Miyata, T. Nomoto, S.
Fukushima, K. Osada, et al., Targeted Polymeric Micelles for siRNA
Treatment of Experimental Cancer by Intravenous Injection, ACS
Nano. 6 (2012) 5174–5189. doi:10.1021/nn300942b.
[14] Z. Ge, Q. Chen, K. Osada, X. Liu, T.A. Tockary, S. Uchida, et
al., Targeted gene delivery by polyplex micelles with crowded PEG
palisade and cRGD moiety for systemic treatment of pancreatic
tumors, Biomaterials. 35 (2014) 3416–3426.
doi:10.1016/j.biomaterials.2013.12.086.
[15] C.-H. Kuo, L. Leon, E.J. Chung, R.-T. Huang, T.J. Sontag, C.A.
Reardon, et al., Inhibition of atherosclerosis-promoting microRNAs
via targeted polyelectrolyte complex micelles, J Mater Chem B. 2
(2014) 8142–8153. doi:10.1039/c4tb00977k.
[16] M. Lueckheide, J.R. Vieregg, A.J. Bologna, L. Leon, M.V.
Tirrell, Structure- property relationships of oligonucleotide
polyelectrolyte complex micelles, Nano Lett. (2018)
acs.nanolett.8b03132. doi:10.1021/acs.nanolett.8b03132.
[17] J.M. Ting, H. Wu, A. Herzog-Arbeitman, S. Srivastava, M.V.
Tirrell, Synthesis and Assembly of Designer Styrenic Diblock
Polyelectrolytes, ACS Macro Lett. 7 (2018) 726–733.
doi:10.1021/acsmacrolett.8b00346.
[18] Y. Oe, R.J. Christie, M. Naito, S.A. Low, S. Fukushima, K.
Toh, et al., Actively- targeted polyion complex micelles stabilized
by cholesterol and disulfide cross- linking for systemic delivery
of siRNA to solid tumors, Biomaterials. 35 (2014) 7887–7895.
doi:10.1016/j.biomaterials.2014.05.041.
[19] J.N. Zadeh, C.D. Steenberg, J.S. Bois, B.R. Wolfe, M.B.
Pierce, A.R. Khan, et al., NUPACK: Analysis and design of nucleic
acid systems, J Comput Chem. 32 (2011) 170–173.
doi:10.1002/jcc.21596.
[20] J. Ilavsky, P.R. Jemian, Irena: tool suite for modeling and
analysis of small-angle scattering, J Appl Crystallogr. 42 (2009)
347–353. doi:10.1107/S0021889809002222.
[21] J. Fu, J.B. Schlenoff, Driving Forces for Oppositely Charged
Polyion Association in Aqueous Solutions: Enthalpic, Entropic, but
Not Electrostatic, J Am Chem Soc. 138 (2016) 980–990.
doi:10.1021/jacs.5b11878.
[22] S. Perry, Y. Li, D. Priftis, L. Leon, M. Tirrell, The Effect
of Salt on the Complex Coacervation of Vinyl Polyelectrolytes,
Polymers. 6 (2014) 1756–1772. doi:10.3390/polym6061756.
[23] J.R. Vieregg, M. Lueckheide, A.B. Marciel, L. Leon, A.J.
Bologna, J.R. Rivera, et al., Oligonucleotide-Peptide Complexes:
Phase Control by Hybridization, J Am Chem Soc. 140 (2018)
1632–1638. doi:10.1021/jacs.7b03567.
[24] E. Haladjova, G. Mountrichas, S. Pispas, S. Rangelov,
Poly(vinyl benzyl trimethylammonium chloride) Homo and Block
Copolymers Complexation with DNA, J. Phys. Chem. B. 120 (2016)
2586–2595. doi:10.1021/acs.jpcb.5b12477.
[25] K.A. Howard, P.R. Dash, M.L. Read, K. Ward, L.M. Tomkins, O.
Nazarova, et al., Influence of hydrophilicity of cationic polymers
on the biophysical properties of polyelectrolyte complexes formed
by self-assembly with DNA, BBA - General Subjects. 1475 (2000)
245–255. doi:10.1016/S0304-4165(00)00076-3.
[26] A.V. Kabanov, I.V. Astafieva, I.V. Maksimova, E.M. Lukanidin,
G.P. Georgiev, V.A. Kabanov, Efficient transformation of mammalian
cells using DNA interpolyelectrolyte complexes with carbon chain
polycations, Bioconjug. Chem. 4 (2002) 448–454.
doi:10.1021/bc00024a006.
[27] S. van der Burgh, A. de Keizer, M.A. Cohen Stuart, Complex
Coacervation Core Micelles. Colloidal Stability and Aggregation
Mechanism, Langmuir. 20 (2004) 1073–1084.
doi:10.1021/la035012n.
[28] K. Sadman, Q. Wang, Y. Chen, B. Keshavarz, Z. Jiang, K.R.
Shull, Influence of Hydrophobicity on Polyelectrolyte Complexation,
Macromolecules. 50 (2017) 9417–9426.
doi:10.1021/acs.macromol.7b02031.
[29] P. Jha, P. Desai, J. Li, R. Larson, pH and Salt Effects on the
Associative Phase Separation of Oppositely Charged
Polyelectrolytes, Polymers. 6 (2014) 1414– 1436.
doi:10.3390/polym6051414.
[30] V.S. Rathee, H. Sidky, B.J. Sikora, J.K. Whitmer, Role of
Associative Charging in the Entropy-Energy Balance of
Polyelectrolyte Complexes, J Am Chem Soc. 140 (2018) 15319–15328.
doi:10.1021/jacs.8b08649.
[31] J.B. Schlenoff, A.H. Rmaile, C.B. Bucur, Hydration
Contributions to Association in Polyelectrolyte Multilayers and
Complexes: Visualizing Hydrophobicity, J. Am. Chem. Soc. 130 (2008)
13589–13597. doi:10.1021/ja802054k.
[32] H. Wu, J.M. Ting, O. Werba, S. Meng, M.V. Tirrell,
Non-equilibrium phenomena and kinetic pathways in self-assembled
polyelectrolyte complexes, The Journal of Chemical Physics. 149
(2018) 163330–11. doi:10.1063/1.5039621.
[33] J. Fu, H.M. Fares, J.B. Schlenoff, Ion-Pairing Strength in
Polyelectrolyte Complexes, Macromolecules. 50 (2017) 1066–1074.
doi:10.1021/acs.macromol.6b02445.
[34] V.A. Izumrudov, M.V. Zhiryakova, S.E. Kudaibergenov,
Controllable stability of DNA-containing polyelectrolyte complexes
in water-salt solutions, Biopolymers. 52 (1999) 94–108.
doi:10.1002/1097-0282(1999)52:2<94::AID-BIP3>3.0.CO;2-O.
[35] H.J. Kim, K. Miyata, T. Nomoto, M. Zheng, A. Kim, X. Liu, et
al., siRNA delivery from triblock copolymer micelles with
spatially-ordered compartments of PEG shell, siRNA-loaded
intermediate layer, and hydrophobic core, Biomaterials. 35 (2014)
4548–4556. doi:10.1016/j.biomaterials.2014.02.016.
download fileview on ChemRxivMarras_chemArxiv.pdf (3.51 MiB)
Alexander E. Marras, Jeffrey R. Vieregg, Jeffrey M. Ting,
Jack D. Rubien, and Matthew V. Tirrell S1. Experimental Information
Materials. The following reagent grade materials were used as
received, unless otherwise specified:
4-cyano-4-(phenylcarbonothioylthio)pentanoic acid (CPhPA, Sigma),
poly(ethylene glycol) methyl ether (2-methyl-2-propionic acid
dodecyl trithiocarbonate) (PEG-C12, Sigma, Reported Mn 1100, 5000,
and 10,000 g/mol), poly(ethylene glycol) 4-
cyano-4-(phenylcarbonothioylthio)pentanoate (PEG-Sty, Sigma,
Reported Mn 10,000 g/mol), (vinylbenzyl)trimethylammonium chloride
(VBTMA, Sigma, 99%), 2,2'-azobis[2-
(2-imidazolin-2-yl)propane]dihydrochloride (VA-044, Wako Chemicals,
USA), 1,6- diphenyl-1,3,5-hexatriene (DPH), acetic acid (glacial,
Sigma, ≥99.85%), sodium acetate trihydrate (Sigma, ≥99%), hydrogen
peroxide (H2O2, Sigma, 30% w/w in H2O), ethanol (anhydrous, Decon
200 proof), and SnakeSkin dialysis tubing / Slide-A-Lyzer dialysis
cassettes (MWCO 3.5K, 22 mm, Thermo Scientific). Acetate buffer
solution was prepared using 0.1 M acetic acid and 0.1 M sodium
acetate trihydrate (0.1 M) (42/158, v/v), adjusted to pH 5.2 for
RAFT polymerizations. Unless otherwise stated, all water was used
from a Milli-Q water purification system at a resistivity of 18.2
MΩ-cm at 25 °C.
RAFT polymerization synthesis. A full description of the reversible
addition- fragmentation chain transfer (RAFT) synthesis can be
found in Ting et al.1 The following pVBTMA-PEG block
polyelectrolytes were prepared in a Carousel 12 Plus Reaction
Station (Radleys, Saffron Walden, UK): 53-5k, 105-5k, and 72-10k.
The 60-1k and 24-5k systems were synthesized in a round bottom
flask, while the 8-5k system was quenched at ~10% conversion of a
105-5k, anticipated from previously-conducted kinetics
experiments.1 pVBTMA(35) and pVBTMA(172) homopolymers were also
prepared in the carousel reactor. In general, to each glass
container, the chemical precursors (monomer, RAFT macromolecular
PEG chain transfer agent / CPhPA chain transfer agent, and VA-044
initiator) were combined in acetate buffer solution and sealed.
10:1 equivalence of RAFT chain transfer agent to initiator was
used. Carousel reactions were degassed simultaneously via three
freeze-pump-thaw cycles; a mixture of acetate buffer solution and
ethanol (3:1, v/v) was used for nitrogen-mediated bubbling to degas
the round bottom flask reactions. Reactions were run at 50 °C under
constant stirring for 21 h, with
2
the final monomer conversion confirmed by 1H NMR spectroscopy
before quenching by cooling to room temperature and opening the
reactors to air.
1H NMR Spectroscopy. 1H NMR experiments were conducted on a Bruker
AVANCE III HD 400 Mhz NanoBay spectrometer with 16 transients to
minimize signal-to-noise. 1H NMR spectra were processed and
analyzed using iNMR (Version 5.5.7). Figure S1 shows a
representative 1H NMR spectrum for the pVBTMA(172)
homopolymer.
1H NMR (400 MHz, D2O) : 1.0-2.5 ppm (alkyl backbone, 3H, –CH2-CH–),
2.5-3.1 ppm (9H, –N-(CH3)3), 4.1-4.5 ppm (2H, –CH2-N–), and 6.1-7.3
ppm (4H, ArH).
Figure S1. Representative 1H NMR spectrum of pVBTMA(172) in D2O.
Fig. S2 shows a representative 1H NMR spectrum for the
pVBTMA(60)-PEG(1k) block polymer. 1H NMR (400 MHz, D2O) : 1.1-2.5
ppm (alkyl backbone, 3H, –CH2-CH–), 2.6- 3.0 ppm (9H, –N-(CH3)3),
3.4-3.6 ppm (4H, –O-(CH2)2–), 4.0-4.5 ppm (2H, –CH2-N–), and
6.3-7.3 ppm (4H, ArH).
H2O
HO
O
NC
S
S
n
3
Figure S2. Representative 1H NMR spectrum of pVBTMA(60)-PEG(1k) in
D2O. dn/dc Measurements. Fig. S3 shows the data used to determine
the change in refractive index (dn/dc) values for pVBTMA(172). The
calculated coefficient of determination (R2 = 0.99) confirms the
linear relationship between refractive index and polymer
concentration. Previous dn/dc measurements of the pVBTMA-PEG
diblock polyelectrolytes can be found in the literature.1
Figure S3. Determination of dn/dc for pVBTMA(172) in the
size-exclusion chromatography mobile phase (60% water, 39.9%
acetonitrile, 0.1% trifluoroacetic acid), via (A) measurements
taken in triplicate and (B) a linear regression to the average of
the data points.
O O
4
Size Exclusion Chromatography. Fig. S4 shows a representative SEC
chromatogram using the refractive index (RI) detector for the
pVBTMA(60)-PEG(1k) system. All samples outlined in Table S1 below
demonstrated a unimodal peak.
Figure S4. Representative SEC RI chromatogram of the
pVBTMA(60)-PEG(1k) in a mobile phase of 59.9% water, 40%
acetonitrile, and 0.1% trifluoroacetic acid. RAFT end group
removal. The RAFT end-group removal reaction was carried out
following protocols reported by Jesson et al.2 H2O2 was used as a
mild oxidant to cleave thiocarbonylthio chain ends at a molar ratio
of 5:1 (H2O2 to chain transfer agent) at 70 °C for 8 h open to the
air in water, at 7.5% w/w. Visually, the solution changed from
light yellow to colorless. Previous reports have shown that PEG can
be susceptible to free radical-induced degradation in water when
exposed to UV/H2O2.3 Thus, as a precaution reactions were conducted
in a dark environment. As seen in Figure S5, the disappearance of
the polymer trithiocarbonate peak centered at 310 nm in water by
UV-vis spectroscopy confirms successful end-group removal for the
PVBTMA(72)-PEG(10k) diblock.
Figure S5. UV-vis spectroscopy of the pVBTMA(72)-PEG(10k) system in
water before (purple curve) and after (pink curve) introduction of
H2O2 to remove chain end-groups.
5
Summary of Polymers and DNA Oligonucleotides. Table S1 shows a
summary of the characterized polymers used in this study. Table S2
contains the DNA oligonucleotide sequences. Table S1. pVBTMA
Polymer Characterization.
Polymer Sample ID PEO Mn (kg/mol)
Charged Block DP a
pVBTMA PVB50 - 35 7.7 1.12 PVB100 - 172 36.7 1.09
pVBTMA-PEG
50-1k 1 60 14.2 1.17 10-5k 5 8 6.6 1.33 30-5k 5 24 10.0 1.28 50-5k
5 53 18.3 1.12 100-5k 5 105 30.9 1.08 100-10k 10 72 24.4 1.22
200-10k 10 194 47.7 1.11 a Experimentally determined degree of
polymerization for the charged blocks. b Experimentally-measured
absolute molecular weight (Mn), determined by SEC-MALS at 35 °C. c
Dispersity () = Mw / Mn. Table S2. DNA Oligonucleotide
Sequences.
Length (nt) Sequence (5’ – 3’) 10 TCAACATCAG 22
CTACCGTCGCATTCAGCATTCA
88 TCAACATCAGTCTGATAAGCTATGGATACTCGTCTGGACTACTTA
CTCACTCATTCATCACTATCTACCGTCGCATTCAGCATTCATG
S2. Scattering and Imaging
Figure S6. SAXS data for individual polyelectrolytes show no
structure formation. Both polyelectrolytes are at 2mM charge
concentration in 1xPBS to match experimental conditions. There is
no y-axis offset for this plot.
10-5
10-4
10-3
10-2
10-1
100
101
2 3 4 5 6 7 0.1
2 3 4
7
Figure S7. All fitting for SAXS data incorporated the following
models: 1) size distribution (purple, above), 2) unified level
(green), and 3) diffraction peak (orange), when applicable. Size
distribution fits include a shape factor for spheroids, rigid
cylinders, or flexible cylinders assuming a Schulz-Zimm
distribution and provide micelle shape and size information in mid
to low q ranges. A unified level fit is used at high q values (q ≥
~0.1 A-1). For data with a high q diffraction peak, this model was
added. Original data with error bars is shown in light blue and the
final model incorporating all three fits is in black.
10-5
10-4
10-3
10-2
10-1
100
101
2 3 4 5 6 7 0.1
2 3 4 5
q [A-1]
SAXS Data Flexible Cylinder Fit Unified Level Fit Diffraction Peak
Fit Final Fit
8
Figure S8. SAXS data to show the effect of pVBTMA end group on PCM
size. Styrene and OH end groups appear very similar to each other
but larger than the C12 PCMs. 22 nt DNA was used for each sample.
Curves are vertically offset for visual clarity. Table S3: SAXS
fitting results for Figure S19 showing the effect of pVBTMA end
group on PCM size.
Sample Mean
Porod exponent
10-3
10-2
10-1
100
101
102
103
104
105
2 3 4 5 6 7 8 9 0.1
2 3 4
9
Figure S9. Cryo TEM image of pVBTMA(72)-PEG(10k) + 88 nt ssDNA
showing spherical micelles. The elongated object in the bottom
right is the lacey carbon grid coating.
10
Figure S10. Cryo TEM image of pLys(200)-PEG(10k) + 88 nt ssDNA
showing spherical micelles.
11
Figure S11. Negative stained TEM image of pVBTMA(24)-PEG(5k) + 88
bp dsDNA showing cylinder and worm-like micelles with similar
diameters but varying lengths.
12
Figure S12. Negative stained TEM image of pVBTMA(105)-PEG(5k) + 10
bp dsDNA showing aggregation and a small population of spheroidal
micelles.
13
Figure S13. Negative stained TEM image of pVBTMA(72)-PEG(10k) +
salmon sperm DNA (~2000bp) showing worm-like micelles with similar
diameter and varying length.
14
Figure S14. Negative stained TEM image of pVBTMA(194)-PEG(10k) + 22
bp dsDNA showing a population of mainly spherical micelles.
15
Figure S15. Negative stained TEM images of pLys(50)-PEG(5k) + 88 bp
dsDNA showing long flexible worm-like micelles.
16
Table S4: Results of SAXS fitting for pLys-PEG + ssDNA
micelles
Sample Mean Radius
(nm) PDI (σ2/R2) Aspect Ratio Porod exponent
pLys(10)-PEG(5k) + ss22 4.2 0.345 1.0 2.9 pLys(10)-PEG(5k) + ss88
4.3 0.007 2.0 2.8 pLys(30)-PEG(5k) + ss10 8.8 0.099 1.6 1.8
pLys(30)-PEG(5k) + ss22 8.7 0.026 1.6 2.2 pLys(30)-PEG(5k) + ss88
7.9 0.027 1.6 2.3 pLys(50)-PEG(5k) + ss10 16.0 0.013 1.4 2.3
pLys(50)-PEG(5k) + ss22 16.2 0.010 1.4 1.9 pLys(50)-PEG(5k) + ss88
15.3 0.018 1.3 2.0 pLys(100)-PEG(5k) + ss10 25.6 0.017 1.2 1.9
pLys(100)-PEG(5k) + ss22 26.0 0.020 1.1 2.1 pLys(100)-PEG(5k) +
ss88 23.5 0.000 1.4 1.8 pLys(100)-PEG(10k) + ss10 17.7 0.008 1.5
1.8 pLys(100)-PEG(10k) + ss22 18.2 0.017 1.5 2.1 pLys(100)-PEG(10k)
+ ss88 16.6 0.014 1.4 1.8 pLys(200)-PEG(10k) + ss10 42.5 0.020 1.3
1.5 pLys(200)-PEG(10k) + ss22 42.9 0.024 1.1 2.1 pLys(200)-PEG(10k)
+ ss88 41.4 0.016 1.1 1.7 pLys(10)-PEG(20k) + ss10 3.1 0.991 1.3
2.1 pLys(10)-PEG(20k) + ss22 6.1 0.127 2.5 3.0 pLys(10)-PEG(20k) +
ss88 2.1 0.692 3.0 2.4 pLys(50)-PEG(20k) + ss10 13.0 0.007 1.9 2.6
pLys(50)-PEG(20k) + ss22 12.0 0.015 1.9 2.0 pLys(50)-PEG(20k) +
ss88 11.5 0.017 1.8 2.1 pLys(100)-PEG(20k) + ss10 19.1 0.039 2.3
2.2 pLys(100)-PEG(20k) + ss22 20.6 0.058 1.5 2.7 pLys(100)-PEG(20k)
+ ss88 19.3 0.013 1.6 2.2
17
Table S5: Results of SAXS fitting for pVBTMA-PEG + ssDNA
micelles
Sample Mean Radius
(nm) PDI (σ2/R2) Aspect Ratio Porod exponent
pVBTMA(24)-PEG(5k) + ss10 10.1 0.048 1.5 2.0 pVBTMA(24)-PEG(5k) +
ss22 8.7 0.051 1.6 2.7 pVBTMA(24)-PEG(5k) + ss88 8.8 0.031 1.7 1.5
pVBTMA(53)-PEG(5k) + ss10 13.1 0.052 3.3 1.7 pVBTMA(53)-PEG(5k) +
ss22 12.1 0.035 1.9 1.9 pVBTMA(53)-PEG(5k) + ss88 12.3 0.024 2.3
1.1 pVBTMA(72)-PEG(10k) + ss10 17.7 0.010 1.5 1.6
pVBTMA(72)-PEG(10k) + ss22 15.9 0.018 1.4 1.7 pVBTMA(72)-PEG(10k) +
ss88 14.7 0.020 1.5 1.7 pVBTMA(194)-PEG(10k) + ss10 30.0 0.020 1.5
1.6 pVBTMA(194)-PEG(10k) + ss22 28.1 0.101 2.0 1.8
pVBTMA(194)-PEG(10k) + ss88 22.7 0.015 2.2 3.2
18
Table S6: Results of SAXS fitting for pLys-PEG + dsDNA
micelles
Sample Mean Radius
(nm) PDI (σ2/R2) Packing
Peak Porod exponent pLys(10)-PEG(5k) + ds22 3.7 0.029 N 2.4
pLys(10)-PEG(5k) + ds88 4.0 0.031 Y 2.6 pLys(10)-PEG(5k) + salmon
4.6 0.104 Y 1.3 pLys(30)-PEG(5k) + ds10 3.8 0.625 Y 3.7
pLys(30)-PEG(5k) + ds22 1.8 1.735 Y 2.8 pLys(30)-PEG(5k) + ds88 6.4
0.039 Y 2.1 pLys(50)-PEG(5k) + ds10 8.6 0.018 Y 2.2
pLys(50)-PEG(5k) + ds22 8.8 0.034 Y 2.8 pLys(50)-PEG(5k) + ds88
11.0 0.029 Y 1.8 pLys(50)-PEG(5k) + salmon 11.4 0.053 Y 1.4
pLys(100)-PEG(5k) + ds10 13.7 0.009 Y 1.9 pLys(100)-PEG(5k) + ds22
13.5 0.021 Y 2.2 pLys(100)-PEG(5k) + ds88 13.9 0.027 Y 1.9
pLys(100)-PEG(10k) + ds10 15.1 0.006 Y 1.8 pLys(100)-PEG(10k) +
ds22 15.2 0.020 Y 2.2 pLys(100)-PEG(10k) + ds88 12.2 0.008 Y 1.9
pLys(100)-PEG(10k) + salmon 13.3 0.066 Y 2.2 pLys(200)-PEG(10k) +
ds10 19.9 0.027 Y 2.1 pLys(200)-PEG(10k) + ds22 16.3 0.012 Y 2.4
pLys(200)-PEG(10k) + ds88 18.1 0.046 Y 1.9 pLys(10)-PEG(20k) + ds10
7.0 0.043 N 2.2 pLys(10)-PEG(20k) + ds22 7.2 0.022 N 2.5
pLys(10)-PEG(20k) + ds88 2.5 0.270 Y 2.3 pLys(10)-PEG(20k) + salmon
4.7 0.050 Y 2.5 pLys(50)-PEG(20k) + ds10 11.0 0.000 Y 2.0
pLys(50)-PEG(20k) + ds22 10.8 0.002 Y 2.3 pLys(50)-PEG(20k) + ds88
10.5 0.029 Y 2.3 pLys(50)-PEG(20k) + salmon 8.8 0.112 Y 1.6
pLys(100)-PEG(20k) + ds10 18.3 0.024 Y 1.9 pLys(100)-PEG(20k) +
ds22 17.1 0.004 Y 1.9 pLys(100)-PEG(20k) + ds88 14.4 0.018 Y 2.0
pLys(100)-PEG(20k) + salmon 13.0 0.064 Y 2.8
19
Table S7: Results of SAXS fitting for pVBTMA-PEG + dsDNA micelles.
Low qx values from data, other parameters from fit.
Sample Mean Radius
exponent Low qx pVBTMA(24)-PEG(5k) + ds10 10.8 0.043 N 1.5
0.45
pVBTMA(24)-PEG(5k) + ds22 10.8 0.044 Y 1.6 0.46
pVBTMA(24)-PEG(5k) + ds88 8.6 0.037 Y 1.8 0.96
pVBTMA(24)-PEG(5k) + salmon 8.4 0.045 Y 2.2 1.90
pVBTMA(53)-PEG(5k) + ds10 14.5 0.060 N 1.3 1.45
pVBTMA(53)-PEG(5k) + ds22 12.1 0.088 Y 1.8 1.92
pVBTMA(53)-PEG(5k) + ds88 11.0 0.052 Y 1.5 1.13
pVBTMA(53)-PEG(5k) + salmon 11.7 0.055 Y 1.4 2.03
pVBTMA(105)-PEG(5k) + ds10 14.5 0.029 N 1.2 1.79
pVBTMA(105)-PEG(5k) + ds22 15.2 0.038 N 1.4 2.32
pVBTMA(72)-PEG(10k) + ds10 18.7 0.018 N 1.2 0.89
pVBTMA(72)-PEG(10k) + ds22 19.1 0.019 N 1.6 1.72
pVBTMA(72)-PEG(10k) + ds88 15.5 0.052 Y 1.7 1.15
pVBTMA(72)-PEG(10k) + salmon 12.7 0.077 Y 2.2 1.99
pVBTMA(194)-PEG(10k) + ds10 31.3 0.021 N 1.5 0.65
pVBTMA(194)-PEG(10k) + ds22 30.3 0.044 Y 2.0 1.59
pVBTMA(194)-PEG(10k) + salmon 17.3 0.046 Y 2.0 2.38
20
Figure S16. SAXS data and fits for all pLys-PEG and ssDNA micelles.
Data is offset in y- axis for clarity.
21
Figure S17. SAXS data and fits for all pLys-PEG and dsDNA micelles.
Data is offset in y- axis for clarity.
22
Figure S18. SAXS data and fits for all pVBTMA-PEG and ssDNA
micelles. Data is offset in y-axis for clarity.
23
Figure S19. SAXS data and fits for all pVBTMA-PEG and dsDNA
micelles. Data is offset in y-axis for clarity. Dotted black lines
indicate low q fitting limit for sphere fit. S3. SI References 1.
Ting, J. M.; Wu, H.; Herzog-Arbeitman, A.; Srivastava, S.; Tirrell,
M. V. Synthesis and
Assembly of Designer Styrenic Diblock Polyelectrolytes. ACS Macro
Lett. 2018, 7, 726– 733.
2. Jesson, C. P.; Pearce, C. M.; Simon, H.; Werner, A.; Cunningham,
V. J.; Lovett, J. R.; Smallridge, M. J.; Warren, N. J.; Armes, S.
P. H2O2 Enables Convenient Removal of RAFT End-Groups From Block
Copolymer Nano-Objects Prepared via Polymerization-Induced
Self-Assembly in Water. Macromolecules 2016, 50, 182–191.
download fileview on ChemRxivMarras_SI.pdf (5.78 MiB)