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: jvieregg@uchicago.edu
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
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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
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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
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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.
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