Complexation between DNA and Hydrophilic ... - ACS Publications

Article pubs.acs.org/JPCB

Complexation between DNA and Hydrophilic-Cationic Diblock Copolymers Seyoung Jung,† Timothy P. Lodge,*,†,‡ and Theresa M. Reineke*,‡ †

Department of Chemical Engineering and Materials Science, University of Minnesota − Twin Cities, 421 Washington Avenue SE, Minneapolis, Minnesota 55455, United States ‡ Department of Chemistry, University of Minnesota − Twin Cities, 207 Pleasant Street SE, Minneapolis, Minnesota 55455, United States S Supporting Information *

ABSTRACT: We examine connections among polycation composition, DNA-polycation binding thermodynamics, binding strength, and resulting complex properties, for circular and linear DNA and hydrophilic diblock copolymers possessing cationic blocks. Two poly(2-deoxy-2-methacrylamido glucopyranose)block-poly(N-(2-aminoethyl) methacrylamide) (PMAG-bPAEMA), with block degrees of polymerization of PMAG56-bPAEMA30 and PMAG52-b-PAEMA63, are employed. DNA binding behavior of these diblocks is also compared with that of a PAEMA homopolymer, in order to evaluate the role of the hydrophilic, charge-neutral PMAG block. In addition, DNA structure was varied, utilizing both circular and linear DNA with the same contour length. The enthalpy change due to DNA-polycation interactions (ΔHint) is observed via isothermal titration calorimetry (ITC) during titrations of DNA with the polycations. With circular DNA, a higher cationic content is found to result in a completion of binding with a smaller amount of polycation, as well as a larger initial ΔHint. In contrast to the common understanding that a neutral block simply provides colloidal stability, the PMAG block turns out to significantly impact both the extent of the binding and the size and dispersity of the final complexes. With a lower cationic content, the complex is less compact, but both the size and dispersity are more stable. Changes in circular dichroism spectra of DNA are shown to be correlated with PMAG-to-PAEMA block length ratio. PMAG52-b-PAEMA63 leads to stronger binding with DNA, compared to PMAG56-b-PAEMA30. Better-defined polyplexes and more disruption in the DNA helices are observed when the PMAG-toPAEMA ratio is lower. All in all, while PMAG itself does not directly interact with DNA, the DNA-polycation binding turns out to be sensitive to the balance between the DNA-PAEMA attraction and PMAG solvation. In addition, it is confirmed that polyelectrolyte complexation is favored both entropically and enthalpically when the ionic strength of the solution is low. While only endothermic interactions occur in the buffered systems, exothermic initial interactions are observed in low-salt, unbuffered cases. Finally, complexes formed with linear DNA show clear bimodal size distributions, distinct from those formed with circular DNA. Collectively, these data provide insights into the controllable parameters in DNA-polycation complexation, which may advance the development of polymeric vehicles for large biomolecules such as nucleic acids.

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INTRODUCTION Polyelectrolytes are of great interest in many fields, due in large part to their spontaneous complexation behavior and controllable properties in aqueous environments. Applications of controlled polyelectrolyte complexation range from versatile layer-by-layer film fabrication1−3 and nanoreactor construction,4,5 to protein purification,6,7 drug encapsulation,3,8 and gene delivery.9,10 In particular, the field of nucleic acid delivery has extensively utilized synthetic polycations as an alternative to traditional viral delivery vehicles.10−12 Spontaneous binding between anionic nucleic acids and polycations can result in interpolyelectrolyte complexes termed polyplexes. Polyplexes can promote delivery of nucleic acid payloads, due to their net positive charge; the positive surface charge participates in nonspecific electrostatic interactions with cell surfaces, leading © 2017 American Chemical Society

to endocytosis via a variety of mechanisms, and ultimately delivery into the cell.13−18 Over the past three decades, a wide variety of polycations have been tested for their ability to deliver nucleic acids into a range of cell types.12,19,20 Diverse synthesis routes to polycations have also been employed, ranging from stepgrowth21,22 and group-transfer polymerizations23 to controlled radical polymerizations.24−26 While many polyplexes have been explored for delivery applications, detailed investigations into their formation process and physical properties are relatively few. Rigorous characterization of complexation between Received: November 12, 2016 Revised: January 17, 2017 Published: March 7, 2017 2230

DOI: 10.1021/acs.jpcb.6b11408 J. Phys. Chem. B 2017, 121, 2230−2243

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The Journal of Physical Chemistry B

Comparison between circular and linear DNA should elucidate the restrictions in polycation binding that stem from conformational constraints. In this study, two PMAG-b-PAEMA diblock copolymers with different PAEMA block lengths, and a PAEMA homopolymer control, have been synthesized and compared with respect to their binding behavior with DNA. Two different solution environments (unbuffered and buffered) are explored to assess the effects of ionic strength on the binding thermodynamics; also, circular and linear DNA are compared. In short, the effects of (a) the PMAG-to-PAEMA ratio, (b) the DNA morphology, and (c) solvent buffering capacity on polyplex formation are investigated. The enthalpy change due to the complexation is directly observed via isothermal titration calorimetry, and the evolution in complex size and dispersity, and in DNA helical structure, are observed via multiangle dynamic light scattering and circular dichroism spectroscopy, respectively. We find that the neutral block (PMAG)while it does not directly interact with DNAsignificantly alters the characteristics of the binding between DNA and PMAG-b-PAEMA. With circular DNA in particular, the complexation strength is shown to decrease with increasing PMAG-to-PAEMA block length ratio in the copolymer, resulting in less compact polyplex structures.

deoxyribonucleic acid (DNA) and polycations has been pursued via time-resolved light scattering,27 nuclear magnetic resonance spectroscopy,28 and electron microscopy.26,29 However, the correlations among polycation structure, DNApolycation binding thermodynamics, and the resulting polyplex physical properties have not yet been fully elucidated. Such understanding is important for predicting behavior of polyplexes in complex environments, e.g., interactions with proteins in blood, and will advance clinical use of polyplexes. The binding between DNA and hydrophilic diblock copolymers with cationic blocks has been much less explored, yet such copolymers provide a number of advantages over homopolymers.13,30,31 For example, a charge-neutral, hydrophilic block can endow polyplexes with colloidal stability, and can potentially promote cell-polyplex interactions. Herein, we aim to establish a quantitative comprehension of polyplex formation using diblock copolymers possessing one cationic block and one charge-neutral hydrophilic block, or cationicneutral diblock copolymers. The neutral block imparts hydrophilicity and often steric bulk, which help to stabilize the polyplexes against aggregation and precipitation under high ionic strength conditions. Although poly(ethylene glycol) (PEG) has been the most studied neutral blockas it is one of the few inexpensive, FDAapproved hydrophilic polymers32,33it can significantly reduce the interactions between the polyplex and the cell membrane,34 potentially activate the human immune system,35 and cause accelerated clearance upon repeated dosing.36 Methacrylate-,25 phosphorylcholine-,37 methacrylamide-,38 and carbohydratecontaining26 neutral blocks have been developed to overcome the limitations of PEG. In particular, carbohydrate-containing neutral blocks stabilize polyplexes while exhibiting little toxicity, and also have the ability to interact with specific cell membrane receptors via multivalent carbohydrate-lectin interactions.39−42 Accordingly, in this work poly(2-deoxy-2-methacrylamido glucopyranose) (PMAG) and poly(N-(2-aminoethyl) methacrylamide) (PAEMA) are used as model neutral and cationic blocks, respectively. A series of PMAG-b-PAEMA diblocks have previously been synthesized and assessed for gene delivery, in which both high cell viability in vitro and polyplex stabilization at high ionic strengths were achieved.26,43,44 Different solution environmentsboth unbuffered45−47 and buffered48−50have been utilized as the media for polyplex formation. DNA-polycation binding in a buffered environment is predicted (and found) to be different from that in unbuffered water where the ionic strength is minimal. Due to release of counterions, polyelectrolyte complexation is generally entropically favored regardless of ionic strength. However, the complexation is supposed to be exothermic only at very low ionic strengths, because a large Debye length is required for a substantial reduction of electrostatic energy upon the formation of polyelectrolyte pairs.51 Here, DNA-polycation binding is investigated in both buffered and unbuffered media to gain a more detailed understanding of the complexation process. DNA architecture is also hypothesized to impact binding with polycations. While circular plasmid DNA (pDNA) is much more prevalent than linear DNA in delivery studies, various morphologies of DNA are of therapeutic interest; thus, the effect of DNA morphology on polyplex formation is important from both fundamental and applied points of view. With a given contour length, linear DNA is suspected to exhibit a more effective initial binding with PMAG-b-PAEMA than circular DNA, because linear DNA is conformationally less restricted.

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EXPERIMENTAL SECTION Materials. β-Galactosidase-encoding, circular plasmid DNA with a cytomegalovirus promoter (pCMV-LacZ, PF462) was purchased from PlasmidFactory GmbH & Co. KG (Bielefeld, Germany). Linear DNA with the same number of base pairs (7164 bps) was obtained by digesting pCMV-LacZ with EcoRV restriction endonuclease, which performs an isolated double strand break, purchased from New England BioLabs Inc. (Ipswich, MA). N-(2-Aminoethyl) methacrylamide hydrochloride (AEMA·HCl) was purchased from Polysciences Inc. (Warrington, PA). Sodium phosphate (monobasic, monohydrate) was purchased from Mallinckrodt Chemicals (Phillipsburg, NJ). 4,4′-Azobis(4-cyanopentanoic acid) (V-501) was purchased from Sigma-Aldrich Co. LLC (St. Louis, MO) and recrystallized using methanol. 4-Cyano-4(propylsulfanylthiocarbonyl)sulfanylpentanoic acid (CPP)52 and 2-deoxy-2-methacrylamido glucopyranose (MAG) monomer53 were synthesized as previously reported. All other chemicals were purchased from Sigma-Aldrich and used without further purification. PMAG-b-PAEMA (Figure 1) and the corresponding homopolymers (PAEMA and PMAG) were synthesized via reversible addition−fragmentation chain trans-

Figure 1. Chemical structure of PMAG-b-PAEMA. x and y represent the degrees of polymerizaion of PMAG block and PAEMA block, respectively. 2231


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the phosphate groups in a DNA solution ([P]0) was 60−240 μM. For all experiments, the polycation and DNA solutions shared the same solvent. For the PMAG homopolymer, as a negative control with no positively charged moieties, the G/P rationumber ratio of the glucopyranose units in PMAG to the phosphate groups in DNAsubstituted for the N/P ratio. The premixing concentration of the glucopyranose units ([G]0) was 3.6 mM in all experiments. Dynamic Light Scattering (DLS). The mean hydrodynamic radii (Rh) of DNA and polyplexes were measured at 25 °C with a Brookhaven BI-200SM light scattering instrument equipped with a Mini-L30 laser (637 nm), an automated goniometer, and a BI-9000AT autocorrelator (Brookhaven Instruments Corporation, Holtsville, NY). Scattering angles of 30°, 50°, 70°, 90°, and 110° were used, and the data analysis methods are discussed in Supporting Information. Isothermal Titration Calorimetry (ITC). ITC experiments were performed on a MicroCal VP-ITC (Malvern Instruments, Westborough, MA) at 25 °C. Before each measurement, the sample cell and the injection syringe were thoroughly cleaned and rinsed with appropriate solvent. DNA and polymer solutions were then placed in the sample cell and the injection syringe, respectively. For a background titration, the solvent was placed into the sample cell. The injection volume was 5 or 10 μL, and the injection interval was adjusted between 300 and 600 s to achieve high baseline quality. Raw ITC data (heat flow rate vs. time) show a peak at each injection, and integrating each with respect to time produces the final ITC data as total heat absorption at each injection vs N/P (or G/P) ratio. In all ITC profiles in this report, the heat absorption (enthalpy change) solely from DNA-polymer interactions (ΔHint) was calculated by subtracting the enthalpy change measured in a background titration (ΔHbgd) from that measured in a main titration (ΔHmain) point by point. Circular Dichroism (CD). CD spectra were obtained using a Jasco J-815 CD Spectropolarimeter (Jasco Inc., Easton, MD) at room temperature. 240 μL of a DNA solution with [P]0 = 120 μM was placed in a Spectrosil quartz cuvette with a path length of 1 mm (VWR International, Radnor, PA). Wavelengths from 210 to 320 nm were scanned with a bandwidth of 1 nm, at a rate of 50 or 100 nm/min. A polymer solution with [N]0 (or [G]0) = 3.6 mM was added directly into the cuvette to increase the N/P ratio, and was immediately and vigorously mixed with the existing DNA solution in the cuvette. The increments in N/ P ratio varied between 0.2 and 2, and are specified with each CD spectrum. For measurements of CD spectra of polymeronly solutions, 240 μL of a polymer solution with [N]0 (or [G]0) = 0.72 mM was added into the cuvette. The background CD signal from solvent was subtracted from all spectra, and none of the spectra were smoothed or further processed.

fer (RAFT) polymerization, following the previously reported procedure.26,43 As listed in Table 1, two PMAG-b-PAEMA copolymers with similar PMAG block lengths but different PAEMA block Table 1. Polymer Characteristics degree of polymerization abbreviated name PMAG56-bPAEMA30 PMAG52-bPAEMA63 PAEMA59 PMAG54

PMAG block

PAEMA block

∂n/∂ca (mL/g)

Mnb (kg/mol)

Đb

56

30

0.175

18

1.13

52

63

0.181

21

1.10

0 54

59 0

0.193 0.168

7.7 13

1.05 1.16

a

Refractive index increments, determined by a refractometer equipped with a 660 nm light source. bNumber-averaged molecular weights and dispersities, determined by SEC equipped with a refractive index (RI) detector and a multiangle light scattering detector.

lengths, one PAEMA homopolymer, and one PMAG homopolymer were synthesized. The molecular weights were determined by size exclusion chromatography (SEC) equipped with multiangle light scattering detectors, and the refractive index increment (∂n/∂c) was measured for each polymer. Diblock copolymer compositions were calculated from the molecular weights measured before and after the second block polymerization. All polymers were dialyzed against ultrapure deionized water and lyophilized before use. Methods. Polymer Characterization. The refractive index increments (∂n/∂c) were measured on a Wyatt Optilab T-rEX refractometer (660 nm light source; Wyatt Technology Corporation, Santa Barbara, CA). The number-averaged molecular weights (Mn) and dispersities (Đ) were determined by SEC, using an Agilent 1260 Infinity system (Agilent Technologies, Santa Clara, CA). The SEC instrument was equipped with Eprogen columns CATSEC100, CATSEC300, and CATSEC1000 (Promigen Life Sciences, Downers Grove, IL) in tandem, a Wyatt DAWN HELEOS II light scattering detector (658 nm laser light source), and an Optilab T-rEX refractometer (660 nm). The eluent used for SEC was 1 wt% aqueous acetic acid solution in which sodium sulfate was dissolved (0.1 M) to achieve pH = 3.0. Polyplex Formation. All solutions and buffers were prepared using ultrapure deionized water from a Millipore Synergy filtration system (EMD Millipore Corporation, Darmstadt, Germany). Solvents used in this study were unbuffered water and phosphate buffers (ionic strength = 11 mM; 6.0 ≤ pH ≤ 7.4). Phosphate buffers were prepared by dissolving sodium phosphate monobasic monohydrate in ultrapure deionized water (5 mM) and adding sodium hydroxide until the desired pH was achieved. To prepare DNA solutions, concentrated stock solutions were dialyzed against the desired solvents extensively over 3 days. To prepare polymer solutions, the polymers in powder form were directly dissolved in the desired solvents. To form polyplexes, a filtered polycation solution was mixed into a filtered DNA solution in a stepwise manner, up to the desired N/P ratioi.e., the number ratio of the protonatable amine moieties in the polycation to the phosphate groups in DNA. The premixing concentration of the protonatable amine groups in a polycation solution ([N]0) was 3.6 mM in all experiments. The premixing concentration of

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RESULTS From this point forward, for convenience, “pDNA” will refer to the circular pDNA (pCMV-LacZ) that is not linearized. The digested, linearized form of pCMV-LacZ will be specifically called “linear DNA”. Effective pDNA-Polycation Binding Observed via ITC. Enthalpy changes due to pDNA-polymer interactions (ΔHint) were measured via ITC, during stepwise addition of each polymer to pDNA, to determine whether the complexation is enthalpically favorable and to track the extent of pDNApolymer interactions at different stages of titration. Figures 2a− d show the ITC profiles for the titrations of pDNA with 2232


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Figure 2. (a)−(d) The evolution of ΔHint (left y-axis) and scattering intensity-weighted mean hydrodynamic size (Rh; right y-axis) during the titrations of pDNA with the polymers in phosphate buffer (pH = 7.4; ionic strength = 11 mM). [N]0 (or [G]0) is 3.6 mM and [P]0 is 0.12 mM in all cases. (a) PMAG56-b-PAEMA30 injected into pDNA; (b) PMAG52-b-PAEMA63 injected into pDNA; (c) PAEMA59 injected into pDNA; (d) PMAG54 injected into pDNA. (e) Photo of the polyplex solutions made with PAEMA59 at N/P ratios of 0.8 (left) and 1.1 (right), which are before and after the aggregation point, respectively.

generally decreases in the initial stage of titration. It is important to realize that the dispersity values found in DLS need not accurately represent the structural dispersity of DNA. With large, charged molecules, both the diffusion and the internal motions contribute to the scattering correlation function. In fact, purely in terms of molar mass, DNA used in this study is narrowly distributed (Figure S3 in Supporting Information). Nonzero apparent dispersity is measured via DLS, mainly due to the architectural dispersity, the contribution from the internal motions, as well as the weak scattering at the relevant concentrations ([P]0 = 120−240 μM). Therefore, throughout this article, the high dispersity of pDNA refers not simply to its DLS result but to its conformational variety. With PMAG56-b-PAEMA30, the dispersity gradually decreases until the N/P ratio reaches the maximum ΔHint point (indicated as a vertical dotted line). With PMAG52-bPAEMA63, the dispersity drops sharply at an N/P ratio as low as 0.5, but increases slightly at higher N/P ratios. PAEMA59 forms well-defined polyplexes at low N/P ratios (≤ 1), although severe aggregation occurs after the N/P = 1 point. With PMAG54, the size dispersity remains constant

PMAG56-b-PAEMA30, PMAG52-b-PAEMA63, PAEMA59, and PMAG54, respectively, under the same buffered environment (pH = 7.4; ionic strength = 11 mM). The mean Rh evolution profilesobtained via DLSare overlaid on the ITC profiles as well. Polyplex formation is found to be endothermic in these particular systems. The maximum in ΔHint is reached after several steps of polycation addition, rather than at the first addition. With PAEMA59, severe aggregation of polyplexes is seen when N/P > 1, and the solution turns optically turbid (Figure 2e). Figure 2d shows the negative control result obtained with PMAG54, where Rh remains constant, and ΔHint is near zero throughout the titration. Dispersity Decrease Observed Using DLS. In addition to the mean Rh, the dispersity in Rh is also important in characterizing interpolyelectrolyte complexes. Figure 3a−c shows the evolution of dispersity (reduced second cumulant, see Supporting Information) during polyplex formation using the three polycations. Note that the Rh distribution obtained by the regularized positive exponential sum (REPES) inversion program54 is monomodal for every pDNA-based polyplex in this study (Figure 3d), and that the polyplex dispersity 2233


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Figure 3. (a)−(c) The evolution of size dispersity during the titrations of pDNA with the polymers in phosphate buffer (pH = 7.4, ionic strength = 11 mM). [N]0 = 3.6 mM in all cases. [P]0 used are the same as in Figure 2. (a) PMAG56-b-PAEMA30 injected into pDNA; (b) PMAG52-bPAEMA63 injected into pDNA; (c) PAEMA59 injected into pDNA. The dotted lines in (a) and (b) indicate the maximum ΔHint points observed in Figure 2. All dispersity values are averages of the values measured at 30°, 50°, 70°, 90°, and 110°. Although the measured values themselves depended on angle, the trends of dispersity evolution were consistent (Figure S4 in Supporting Information); thus, here, we report the average values. (d) The Rh distribution of polyplexes formed with pDNA and PMAG56-b-PAEMA30 as in (a). The scattering angle of 90° was used for REPES analyses. Note that the small populations at Rh < 10 nm are artifacts from the short relaxation times rather than actual particles.

formed in unbuffered water where the ionic strength is minimized. To understand the differences in polyplex formation in the presence and absence of the buffering salts, polyplex formation was investigated in unbuffered water. Figure 5a,b shows the results obtained with PMAG56-b-PAEMA30 and PMAG52-b-PAEMA63, respectively. With PMAG56-b-PAEMA30, a great increase in Rh occurs during the initial stage of titration, and then the size is maintained until high N/P ratios are reached. This is very similar to the result shown in Figure 2a. With PMAG52-bPAEMA63, the size does not change initially but increases around the N/P ratio of maximum ΔHint. Compared to the buffered case (Figure 2b), however, the size increase at high N/ P ratios is not as dramatic. This suggests the absence of aggregation, which is consistent with the fact that polyplex aggregation is more prone to occur at higher ionic strengths.43 Figure 6a−c shows the evolution of polyplex dispersity in unbuffered water. Again, all polyplexes show lower dispersity compared to pDNA alone, and PMAG56-b-PAEMA30 induces the most gradual decrease in dispersity. The size distributions of polyplexes remain monomodal as shown in Figure 6d. The effect of the PMAG block can be further evaluated by comparing the impact of PMAG52-b-PAEMA63 and PAEMA59 on the pDNA helical structure in unbuffered solution. Figure 7 shows the results, again supporting the hypothesis that, at low N/P ratios, a smaller PMAG-to-PAEMA block length ratio leads to stronger binding to the pDNA helix. PMAG56-b-PAEMA30 did not induce any noticeable change in

throughout the titration (see Figure S5b in Supporting Information). Correlations Between Evolution of CD Spectra and ITC Profiles. Changes in the CD spectrum of DNA indicate changes in the local helical structure, i.e., the secondary structure. Strong binding between DNA and a polycation is expected to alter the secondary structure, and thus CD has been used to monitor the interactions between DNA and polycations.55,56 Figure 4a−c compares the CD spectra of pDNA during stepwise introduction of the three polycations. Because the PMAG block displays strong negative CD signals at wavelengths shorter than 240 nm (Figure 4d), the discussion here only concerns the wavelength range where the CD signal from the PMAG block is absent, i.e., between 245 and 320 nm. The ignored portion of CD spectra is shaded gray. For PMAG56-b-PAEMA30, with the lowest cationic content, the pDNA secondary structure remains relatively constant throughout the titration (Figure 4a). Even with a longer cationic block (PMAG52-b-PAEMA63), the overall change is not dramatic, and observable changes occur only for N/P > 0.5 (Figure 4b). On the other hand, PAEMA59 induces significant changes in pDNA secondary structure, suggesting the strongest interactions with pDNA (Figure 4c). pDNA-Polycation Complexation in Unbuffered Water. In many gene delivery studies, polyplexes are formed in pure, unbuffered water, while most ITC studies have been done in buffered systems. For PMAG-b-PAEMA in particular, the previous gene delivery studies43,44 utilized the polyplexes 2234


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Figure 4. (a)−(c) The evolution of CD spectra of pDNA during the titrations with PMAG56-b-PAEMA30 (a), PMAG52-b-PAEMA63 (b), and PAEMA59 (c) in phosphate buffer (pH = 7.4; ionic strength = 11 mM). [N]0 = 3.6 mM and [P]0 = 0.12 mM in all cases. The y-axis is molar ellipticity, i.e., the solution ellipticity divided by the final phosphate group molarity. (d) CD spectra of the PMAG homopolymer (black), PMAG-bPAEMA copolymer (red), and PAEMA homopolymer (blue) in water. On the y-axis, the solution ellipticity is divided by the MAG repeat unit molarity (black and red) or AEMA repeat unit molarity (blue).

Figure 5. Evolution of ΔHint (left y-axis) and mean hydrodynamic size (right y-axis) during the titrations of pDNA with the diblock copolymers in unbuffered water with no salt added. [N]0 = 3.6 mM in all cases. (a) PMAG56-b-PAEMA30 injected into pDNA ([P]0 = 0.080 mM); (b) PMAG52b-PAEMA63 injected into pDNA ([P]0 = 0.12 mM).

the CD spectrum of pDNA (Figure S8b in Supporting Information), similar to the buffered case. In summary, while exothermic interactions emerge in unbuffered aqueous conditions (Figure 5), the complexation behavior is largely similar between buffered vs unbuffered systems, especially concerning the effects of the PMAG-toPAEMA ratio on both the polyplex dispersity and the pDNA secondary structure. This similarityobserved via DLS and CDis important because the effects of added ions on polyplex formation are rarely discussed, while various environments are chosen by different researchers in the field of nucleic

acid delivery. Polyplex formation in unbuffered environment (Figures 5−7) is discussed in detail in Supporting Information. Effects of pH on pDNA-Polycation Complexation. The binding between pDNA and PMAG56-b-PAEMA30 was studied in two buffers with the same ionic strength (11 mM) but different pH values (7.4 vs 6.8). This experiment was performed to examine if a pH-induced increase in the number of charged AEMA moieties (by decreasing pH) affects ΔHint similarly to increasing the PAEMA block length at a fixed pH. Figure 8 shows the results, where the x-axis has been converted from N/P ratio to charge ratio. The ITC profile obtained with PMAG56-b-PAEMA30 at pH 6.8 (open circles) is similar to 2235


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Figure 6. (a)−(c) The evolution of size dispersity during the titrations of pDNA with the polycations in unbuffered water. [N]0 = 3.6 mM in all cases. [P]0 used are the same as in Figure 5. (a) PMAG56-b-PAEMA30 injected into pDNA; (b) PMAG52-b-PAEMA63 injected into pDNA; (c) PAEMA59 injected into pDNA ([P]0 = 0.12 mM). The dotted lines in (a) and (b) indicate the points where ΔHint is at maximum, which was observed using ITC. All dispersity values are averages of the dispersities measured at 30°, 50°, 70°, 90°, and 110°. (d) The Rh distribution of polyplexes formed with PMAG56-b-PAEMA30 as in (a). The scattering angle of 90° was used for these REPES analyses.

that obtained with PMAG52-b-PAEMA63 at pH 7.4 (Figure 2b). Initial binding is more evident with a higher cationic content in polymer, which can be achieved with either a lower pH or a longer PAEMA block. The maximum ΔHint is observed at a similar charge ratio even when the cationic charge density is varied significantlyfor example, the ratio (number of deprotonated amine groups)/(number of protonated amine groups) decreases by four times (from 1.1 to 0.27) as the pH is lowered from 7.4 to 6.8. Complexation Behavior of Linear DNA. The studies with pDNA (Figures 2−8) suggest that the pDNA-polycation interactions are affected by the chemical composition of the polycation, particularly by the ratio between the numbers of cationic AEMA and neutral MAG units. We hypothesize that this is due to the constraints on pDNA structure, whereby polycations with bulky neutral blocks are prevented from easily accessing phosphate groups. If so, linear DNA is less conformationally confined, and polycations may have a better access to the DNA phosphate groups (the “binding sites”). To this end, the pDNA was cleaved with a restriction enzyme to linearize the structure, allowing us to examine and compare the binding behavior with the same polycations. Figure 9 presents the complex size evolution during the titrations using linear DNA. PMAG52-b-PAEMA63 and PAEMA59 are compared in particular, to investigate the influence of the PMAG block. In contrast to the circular pDNA case, the complexes at most N/P ratios show bimodal size distributions in both buffered and unbuffered conditions, with both polycations. The bimodal size distributions were also confirmed via transmission electron microscopy (TEM), as shown in Figure S9 in the Supporting Information.

Unfortunately, ITC is not readily applicable for studying the interactions between linear DNA and polycations, due to the formation of a bimodal distribution of aggregates. The heat of binding obtained using ITC would therefore be influenced not only by the molecule-to-molecule interactions between DNA and the polycation, but also by multiparticle aggregation. We therefore concluded that ITC profiles obtained with linear DNA would not be directly interpretable. Figure 10 shows the CD spectra of linear DNA upon the additions of PMAG52-b-PAEMA63 and PAEMA59. Since linear DNA has less confined morphology than pDNA, the polycationswith or without PMAG blockare hypothesized to bind to linear DNA even at the initial stage of titration. Indeed, in Figure 10, spectral shifting is found at the first injection points (note the black and dark olive lines in Figure 10a and c).

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FURTHER DISCUSSION pDNA-Polymer Interactions Under Buffered Conditions. From the observations reflected in Figures 2−4, we propose a schematic picture of polyplex formation in buffer (Figure 11). It should be noted that, while Figure 11 is one of the most reasonable explanations conforming to the data, other possibilities may exist. The open or partially open conformations of pDNA shown in Figure 11a, for instance, simply represent pDNA conformations that exhibit large values of Rh, but not all possible conformations. In Figure 2a, the increase in Rh at the first few injection points is observed, but no significant ΔHint is detected. For a similar system, Jiang et al.57 observed more open and extended pDNA conformations within polyplexes as the solvent became 2236


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Without a neutral block (Figure 11b), PAEMA59 is able to access the pDNA phosphate groups with more ease, and welldefined polyplexes are immediately formed even at low N/P ratios. However, as more PAEMA59 is added, the system approaches charge neutrality and the polyplexes precipitate. With PMAG52-b-PAEMA63 (Figure 11d), behavior intermediate to the two previous cases above is found. Relatively compact, narrowly distributed complexes are formed at low N/ P ratios. The low number density of both the polycations bound to the pDNA and PMAG blocks in the corona, however, results in limited colloidal stability. As the N/P ratio further increases (> 2), the observed Rh increases, although to a much lower extent than with PAEMA59. This size increase is due to limited aggregation behavior of the polyplexes, as proven by static light scattering (SLS), see Figure S10 in Supporting Information. The slight increase in dispersity is also consistent with this picture. At a given pH and N/P ratio, the polyplexes formed with PMAG52-b-PAEMA63 will have fewer neutral PMAG chains in the polyplex corona than with PMAG56-bPAEMA30, if we assumed that a similar number of AEMA units are bound to each pDNA molecule, regardless of the diblock composition. A low number density of PMAG blocks in the polyplex corona leads to low colloidal stability and aggregation. Thus, while a neutral block may make the pDNA-polycation binding weaker, sufficient density in the corona is necessary to facilitate colloidal stability of the resulting polyplexes. Meanwhile, Figure 2d (corresponding to the illustration Figure 11c) confirms that the PMAG block by itself does not interact with pDNA, even though its presence or absence in a diblock greatly impacts the complexation. In this specific study, the amounts of uncomplexed polycations at different N/P ratios are not a subject of detailed discussion. This is mainly because previous studies using different polycation systems have shown that over 75% of polycations participate in binding with pDNA when the N/P ratio is low (≤ 5),28 and that uncomplexed polycations do not significantly impact the physical structure of polyplexes.58 The majority of polycations are thus assumed to be bound to DNA in the low N/P ratio range (< 5) explored in this study. General Characteristics of the Binding Between pDNA and PMAG-b-PAEMA. In Figure 2, the maximum magnitude of ΔHint is observed after several injections of polycation rather than at the first injection point. This indicates an apparent cooperativity in the binding, i.e., the tendency of pDNA to bind increases with further injections of polycation, when the N/P ratio is low. Apparent positive cooperativity has also been observed with other pDNA-polycation systems.59−61 This cooperativity may be partly attributed to the fact that pDNA is a conformationally confined macromolecule. Structural restrictions may result in limited initial accessibility between the “binding sites” (phosphate groups and amine groups), and thus maximum interactions may not be possible at the beginning of a titration. In contrast, absence of cooperativity has been observed between a cationic-neutral diblock copolymer and an oppositely charged flexible polymer, showing Langmuir-like binding.62 Here, “Langmuir-like” refers to the systems showing an S-shaped ITC curve where the magnitude of ΔHint consistently decreases throughout the titration. Bindings between small cations and pDNA and between oppositely charged homopolymers have also often been observed to be Langmuir-like.63,64 These examples support the inference that the non-Langmuir binding observed in Figure 2 stems from the structural nature of pDNA. Double-

Figure 7. Evolution of CD spectrum of pDNA during the titrations with PMAG52-b-PAEMA63 (a) and PAEMA59 (b) in unbuffered water. [N]0 = 3.6 mM and [P]0 = 0.12 mM in all cases.

Figure 8. Evolutions of ΔHint during titrations of pDNA with PMAG56-b-PAEMA30 in two phosphate buffers (ionic strength = 11 mM for both cases; pH = 7.4, filled circles; pH = 6.8, open circles). [N] = 3.6 mM, [P] = 0.12 mM. In the legend, “amine” refers to the protonatable amines in the diblock.

more polar (i.e., became a better solvent for the neutral block): in the most polar solvent (water), conformations were consistently more open and extended than the ones observed in solvents that are less good for the neutral block (see Figure 2 in ref 57). Adapting this result to our system, a higher PMAGto-PAEMA block length ratio is expected to result in a more open structure of pDNA, leading to a larger Rh. This is because a neutral block in its good solvent prefers to be surrounded by solvent rather than to approach close to pDNA, driving the pDNA to change its conformation to be more accessible (Figure 11a). 2237


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Figure 9. (a) and (b) are the evolution of hydrodynamic radius of complexes during the titrations of linear DNA in phosphate buffer (pH = 7.4; ionic strength = 11 mM) with PMAG52-b-PAEMA63 (a) and PAEMA59 (b). (c) and (d) show the evolution of hydrodynamic size of complexes during the titrations of linear DNA in unbuffered water with PMAG52-b-PAEMA63 (c) and PAEMA59 (d). [N]0 = 3.6 mM and [P]0 = 0.12 mM in all cases. The REPES program54 was employed to extract the bimodal size distributions, using the scattering data obtained at 30°. With PAEMA59, the complexes formed micron-sized aggregates at N/P ratios higher than 1.2.

Figure 10. (a) and (b) are the evolution of CD spectra of linear DNA during the titrations in phosphate buffer (pH = 7.4; ionic strength = 11 mM) with PMAG52-b-PAEMA63 (a) and PAEMA59 (b). (c) and (d) are the evolution of CD spectra of linear DNA during the titrations in unbuffered water with PMAG52-b-PAEMA63 (c) and PAEMA59 (d). [N]0 = 3.6 mM and [P]0 = 0.12 mM in all cases. 2238


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Figure 11. Schematic depiction of the interactions between pDNA and the four model polymers PMAG56-b-PAEMA30 (a), PAEMA59 (b), PMAG54 (c), and PMAG52-b-PAEMA63 (d). Black double helical chains represent pDNA molecules. Orange and dark blue lines represent neutral (PMAG) and cationic (PAEMA) blocks, respectively. Note that it is not drawn to scale. In reality, the contour length of the pDNA is 2 orders of magnitude longer than those of the polymers. With the PMAG homopolymer, no DNA-polymer binding is observed. In (a) and the first part of (d), the N/P ratio around 1 is exemplified, although the structure in (a) would hold for most N/P ratios (as implied by Figure 2a). The second (far right) part of (d) represents N/P ≥ 2.

hence suggest that having a longer PAEMA block enhances the tendency of the diblock to access and bind to pDNA. This effect of PAEMA length is found to be consistent with the CD results, discussed subsequently. Second, the maximum ΔHint is observed at a lower N/P ratio, near unity. This again suggests that the accessibility of PMAG-b-PAEMA to pDNA increases with increasing PAEMA length. Third, ΔHint abruptly decreases to near zero after the maximum value is achieved, as opposed to PMAG56-b-PAEMA30, which shows prolonged endothermic interactions after the maximum ΔHint point (Figure 2a). This implies that a higher PMAG-to-PAEMA ratio results in a more gradual completion of the endothermic complexation. In the case of PAEMA59 (Figure 2c), aggregation and precipitation of polyplexes are seen around N/P = 1, along with a sharp spike in the ITC profile. This underscores a major advantage of using cationic-neutral diblocks, in that they impart colloidal stability. As shown in Figures 2a and 2b, the diblock copolymers do not result in severe aggregation at any N/P ratio although, in the case of PMAG52-b-PAEMA63, limited aggregation is seen at high N/P. Additionally, the polyplexes formed with PMAG56-b-PAEMA30 exhibit almost the same Rh even after multiple days (see Figure S11 in Supporting Information). This stability provided by the PMAG block shows the promise of hydrophilic carbohydrate-containing neutral blocks in nucleic acid delivery. Additionally, the higher charge density of PAEMA affects the pDNA-polycation interactions in a very similar way as a longer PAEMA block (Figure 8). Increasing the cationic content, either by decreasing pH or by increasing the PAEMA length, facilitates the initial endothermic binding. Meanwhile, the ITC profiles measured at pH 7.4 and 6.8 show a maximum at almost the same charge ratio (+/−), implying that there exists a charge ratio at which the most effective form of binding is initiated. However, this interpretation assumes no increase in the pKa of the amine groups during polyplex formation, while all ΔHint values reported in this article likely include an enthalpy change

stranded pDNA can adopt various conformations ranging from supercoiled to relaxed circular forms. Meanwhile, PMAG-bPAEMA possesses a charge-neutral block (PMAG) that has no thermodynamic driving force to bind or approach pDNA. In fact, a diblock copolymer with a longer neutral block has been shown to bind with DNA more weakly than with a shorter neutral block, via DNA quenching assay.59 Additionally, PMAG contains bulky glucopyranose units that may further deter PMAG-b-PAEMA from easily accessing the DNA phosphate groups, and thus some pDNA conformations are better than others at allowing PMAG-b-PAEMA to bind effectively. Focusing on the ITC profile in Figure 2a, ΔHint gradually increases and peaks around N/P = 2, which is the charge neutrality point. (The degree of protonation of the PAEMA block is 50% at pH = 7.4, and thus N/P = 2 is equivalent to the charge ratio +/− = 1.) The maximum ΔHint may seem smaller than the thermal energy (ca. 600 cal/mol), but one should again note that the PAEMA block is only 50% protonated. Since the number of amine groups that can participate in the binding is smaller than the total number, the heat absorption from pDNA-polycation interactions is in fact comparable to the molecular thermal energy. The ITC profile obtained with PMAG52-b-PAEMA63 (Figure 2b) displays three distinct features compared to PMAG56-b-PAEMA30. First, noticeable endothermic interactions with pDNA are detected even at the first injection point. This can be attributed to the fact that a longer cationic block results in a larger entropy increase upon the pDNApolycation binding. (Change in entropy does not directly affect ΔHint itself, but if more binding is promoted due to an overall decrease in free energy change (ΔG), ΔHint at each injection point should increase.) The more counterions released per polycation during complexation, the larger the entropic advantage. Nevertheless, the entropic contribution may not be sufficient to explain how doubling the PAEMA length increases ΔHint from near zero to hundreds of cal/mol. We 2239


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secondary structure of PMAG, e.g., helical structure. Yin et al. have previously reported helical polycations that showed similar CD spectra to PMAG, and studied their DNA delivery efficiencies in vitro.68 However, the polycations in the referenced work are polypeptide-based, being inherently distinct from methacrylamide-based, RAFT-polymerized polymers such as PMAG. To our knowledge, there has not been a report of nonpeptide polycations that show secondary structures and have the ability to deliver nucleic acids. The dispersity in Rh for pDNA itself is large, and for such large molecules, diffusivity is strongly dependent on conformation. Once well-defined polyplexes are formed, the dispersity decreases (Figures 3 and 6). This was also visualized by atomic force microscopy (AFM) (Figure S14 in Supporting Information) under the assumption that the size dispersity is conserved after solvent removal. Even though the pDNA molecules have various initial conformations, they eventually converge to a specific conformation upon complexation with a polycation. The fact that PMAG56-b-PAEMA30 induces the most gradual dispersity decrease also confirms that a higher PMAG-to-PAEMA ratio leads to a weaker tendency to bind with pDNA. Altogether, the information obtained from ITC, DLS, and CD is mutually consistent. Both the pDNA-polycation interactions and the formation of well-defined polyplexes are discouraged when the PMAG-to-PAEMA block length ratio in the polycation is high. In addition, initial pDNA-polycation binding is significant only when the cationic content in the polycation is large. Differences between pDNA and Linear DNA in Polycation Complexation. A pDNA molecule is intrinsically confined in terms of physical motion, due to its closed structure and also due to its superhelicity. These conformational restrictions can be removed simply by cutting the pDNA at one spot, making it linear. We therefore hypothesized that linear DNA would provide better accessibility to all three polycations, regardless of the PMAG-to-PAEMA ratio. The evolution of the CD spectrum of linear DNA (Figure 10) supports this hypothesis. Independent of the block composition, a shift is observed in the spectrum even at low N/P ratios. In fact, it was found that even PMAG56-b-PAEMA30 is able to bind and distort the secondary structure of linear DNA (Figures S8c and S8d in Supporting Information). Another reason why binding with linear DNA is less affected by the PMAG content may be the fact that linear DNA has open ends that are more susceptible to the helix disruption caused by polycations, compared to pDNA which is a closed loop. The spectral evolution patterns in Figures 10b and 10d are strikingly similar to Figures 4c and 8b, respectively. This suggests that a PAEMA homopolymer can effectively bind to both pDNA and linear DNA. It is important to note that PMAG-b-PAEMA still alters the DNA secondary structure much less than does PAEMA, even in the case of linear DNA. In buffered systems, PAEMA almost completely eliminates the negative (250 nm) and positive (275 nm) CD signals (Figure 10b), while PMAG-b-PAEMA is limited in how much it can disrupt the base stacking in DNA helix (Figure 10a). We hence conclude that the PMAG block significantly restricts the extent and strength of DNA-polycation binding, which conflicts with the common understanding that the only function of a neutral block is to form a stabilizing corona for the polyplex. As shown in Figure 9, with linear DNA, the presence or absence of the PMAG block does not significantly affect the

due to the pKa increase leading to proton uptake. For the buffered systems in this work, the contribution from proton uptake was assumed to be consistent regardless of polycation structure and of charge density, as a single buffer condition (phosphate-based, at ionic strength = 11 mM) and a single cationic block chemistry (PAEMA) were used throughout all experiments. CD Spectral Evolution and Connection with DLS and ITC. In Figure 4b, no observable change occurs in pDNA secondary structure until the N/P ratio is increased to 0.5. Then a clear red shift (e.g., a shift of the positive peak from 275 to 285 nm) occurs when the N/P ratio is increased to 1. Such red shifts are reported when pDNA complexes with polycations under buffered conditions.55,56 In terms of the overall shape of the CD spectra, no dramatic change is caused by PMAG52-bPAEMA63, and the change is even smaller with PMAG56-bPAEMA30 (Figure 4a). In a negative control experiment with PMAG54, no change is observed in the CD spectrum (Figure S12 in Supporting Information), again confirming no interactions between DNA and PMAG. This brings up the underlying assumption in this study: the potential hydrogen bonding between the hydroxyl groups in the PMAG block and the bases in the DNA chain either does not exist or minimally contributes in polyplex formation. Prevette et al. have previously suggested the presence of nonionic hydrogen bonding between hydroxyl groups of poly(glycoamidoamine)s (PGAAs) and DNA,65 and PMAG may seem to be capable of interacting with DNA via hydrogen bonding as well. However, such nonionic attraction is hypothesized to come into play only when the ionic long-range attraction has readily brought the hydroxyl groups close to the DNA chain, which may not be the case for diblocks. While PGAAs exhibit the charge-neutral hydroxyl groups and the cationic amines in an alternating fashion, PMAG-b-PAEMA is a diblock where the hydroxyl groups are segregated into a separate neutral block (PMAG) away from the DNA-PAEMA binding sites. In addition to this inherent difference between PGAAs and PMAG-b-PAEMA diblocks, the control experiments discussed so far (Figure 2d and Figures S5b and S12 in Supporting Information) indicate that PMAG alone does not interact with DNA in any way. We suggest that a bulky neutral block limits the binding to pDNA and thus how much the polycation can disrupt the secondary structure of pDNA. This conclusion is strongly supported by the result obtained with the PAEMA homopolymer; in Figure 4c, observable changes in the secondary structure are readily achieved even at an N/P ratio as low as 0.4, and the magnitudes of both the positive (280 nm) and negative (250 nm) bands continuously decrease throughout the titration. CD signals of double-stranded DNA stem from base−base interactions from which π → π* transitions arise.66,67 Therefore, the noticeable decrease in CD signals may be attributed to the disruption in base stacking and thus to “untilting” of the DNA double helix. In other words, PAEMA59 is able to progressively prevent π → π* transitions by strongly binding to the DNA double helix. This is consistent with the size and dispersity evolution, where PAEMA59 forms compact and well-defined polyplexes prior to the point of aggregation. ITC profiles also indicate that a polycation with a higher AEMA content has a stronger tendency to complex with pDNA. It is worth noting that PMAG-containing polymers display CD signals at wavelengths ≤ 240 nm (Figures 4a, b, and d). While a discussion about the source of PMAG optical activity is beyond the scope of this article, it implies the existence of 2240


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complications in designing polymeric delivery vehicles for nucleic acids, which is important for further biomedical and clinical development.

polyplex size distribution at low N/P. PAEMA59 results in colloidally unstable systems at N/P > 1.2, but other than that, the general trends in size evolution are similar in all cases. Both PMAG52-b-PAEMA63 and PAEMA59 result in bimodal size distributions, which were never observed with pDNA. The smaller particles consistently displayed Rh around 90−200 nm, and the larger particles had Rh larger than 300 nm. The smaller particles are suspected to be polyplexes containing a single DNA molecule, and the larger particles contain multiple DNA molecules. Compared to pDNA, which intrinsically has a more confined structure, linear DNA spans a longer distance and a larger volume in a solution. Linear DNA can thus extend more freely into space and interact with neighboring polymers. This conformational freedom consequently provides a higher chance to form large complexes. In summary, DLS and CD results confirm that linear DNA-polycation complexes form differently from the pDNA counterpart. Linear DNA enabled all model polycations to bind effectively at initial stage of complexation, while pDNA is sensitive to the existence of PMAG. Furthermore, linear DNA leads to bimodal particle distributions, while pDNA-polycation complexation only leads to a decrease in polyplex dispersity.

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ASSOCIATED CONTENT

* Supporting Information S

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.jpcb.6b11408. Measurement of pKa of the PAEMA block, analysis methods for multiangle DLS data, purity of DNA confirmed by gel electrophoresis, dispersity evolution measured at multiple scattering angles, evolution of particle size during titrations of pDNA with PAEMA59 and PMAG54 homopolymers, conservation of size dispersity upon additions of PMAG54, pH-dependence of solubility of the PMAG block, complexation between pDNA and PMAG52-b-PAEMA63 at pH = 6, evolution of pDNA secondary structure upon additions of PMAG56-b-PAEMA30, TEM images of the bimodal size distributions of complexes formed from linear DNA, SLS of polyplex aggregates, polyplex stability over time, evolution of pDNA secondary structure upon additions of PMAG54, further discussions about the ITC data based on ITC data fitting, and visualization of polyplex dispersity decrease via AFM (PDF)

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CONCLUSIONS In this study, a selective set of polycations and DNA were evaluated in their complexation process, and fundamental questions were addressed with regard to the effects of (a) the block length ratio, (b) the DNA architecture, and (c) solvent buffering capacity on polyplex formation. It was discovered that the complexation between PMAG-b-PAEMA and pDNA is profoundly impacted by the balance between the DNAPAEMA block binding and the PMAG block solvation effects. When the PAEMA block is short (in the case of PMAG56-bPAEMA30), the binding is weakespecially at low N/P ratiosand loosely structured polyplexes are formed. The colloidal stability, however, benefits from the high PMAG-toPAEMA ratio, as the dense coronas prevent aggregation. With PMAG52-b-PAEMA63, with a longer PAEMA block, even a modest quantity can effectively complex with pDNA and form well-defined polyplexes. The charge ratio at which the maximum interaction occurs is also lower with a longer PAEMA block. However, due to the low PMAG-to-PAEMA ratio, the colloidal stability of the final polyplexes is not as high as the case of PMAG56-b-PAEMA30. Altogether, although the cationic block is the one that actively interacts with pDNA chains, the bulky neutral block exhibits a substantial impact on the binding behavior and polyplex properties. In unbuffered water, where the ionic strength is minimal, polyplex formation consists of two distinct steps. The first step is enthalpically favorable; and the second step is endothermic, where the binding is associated with changes in pDNA secondary structure. Regardless of the existence of buffering agents, however, effective binding between pDNA and polycations is generally accompanied by a decrease in the dispersity of complex size. DNA morphology is a significant factor in polyplex formation. With the same contour length, linear DNA results in bimodal size distributions when complexed with PMAG-bPAEMA, whereas pDNA only forms a single population of polyplexes. This study emphasizes the impact of the unique architecture of both diblock-structured polycations and pDNA on their complexation and solution behavior. The results presented herein provide insight into the fundamental

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AUTHOR INFORMATION

Corresponding Authors

*E-mail: [email protected]. *E-mail: [email protected]. ORCID

Timothy P. Lodge: 0000-0001-5916-8834 Theresa M. Reineke: 0000-0001-7020-3450 Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS This work was supported primarily by the National Science Foundation through the University of Minnesota Materials Research Science and Engineering Center (MRSEC) under Award Number DMR-1420013. This work was partially funded by the National Institutes of Health (NIH) program under Award Number 1-DP2-OD00666901. AFM imaging reported in Supporting Information was carried out at the College of Science & Engineering Characterization Facility at University of Minnesota. We acknowledge Dr. Yaoying Wu and Dr. Zachary P. Tolstyka for their help with polycation synthesis and characterization, Dr. Lisa E. Prevette for helpful discussions on ITC experiments, and Peter Schmidt for TEM imaging presented in Supporting Information. We also thank Dr. Robert Geraghty at University of Minnesota for sharing the ITC instrument.

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