Structures and Protonation States of HydrophilicâCationic Diblock

Article Cite This: J. Phys. Chem. B XXXX, XXX, XXX−XXX

pubs.acs.org/JPCB

Structures and Protonation States of Hydrophilic−Cationic Diblock Copolymers and Their Binding with Plasmid DNA 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: Complexation between plasmid DNA (pDNA) and a set of diblock copolymers, each with one cationic block and one hydrophilic, charge-neutral block, is examined. A range of hydrophilic block structures are explored, whereas the cationic block is fixed as poly(N-(2-aminoethyl) methacrylamide) (PAEMA) with a degree of polymerization of 60 ± 3. The hydrophilic blocks include poly(ethylene glycol) (PEG45), poly(oligo(ethylene glycol) methyl ether methacrylate) (P(OEGMA)51), and poly(2-deoxy-2-methacrylamido glucopyranose) (PMAG52). The numbers represent the degrees of polymerization and are chosen such that the diblock contour lengths are similar (37 ± 2 nm). The three diblock copolymers and a homopolycation control, PAEMA59, are compared with respect to their state of dissolution in aqueous environments, as well as their complexation with pDNA. The diblock copolymers are found to partially aggregate as pH increases above 6, whereas each separate block generally dissolves well over a wide pH range. The hydrophilic block proves to be a critical parameter in determining the structures of pDNA−diblock complexes. When the molar ratio of polycation amines to pDNA phosphates (i.e., N/P) is less than 1, a bulkier hydrophilic block leads to larger resulting complexes. As more polycations are added to the system (N/P > 1.5), colloidal stability becomes an important factor, making more water-soluble systems stabilize at smaller sizes. Further, the charge density effect on the binding thermodynamics is elucidated via calorimetric measurements. P(OEGMA)51-b-PAEMA60 exhibits a greater amount of endothermic pDNA binding per charged amine at higher pH, implying that lower cationic charge density promotes more phosphate pairing per amine on average. Also, the colloidal stability and the circular dichroism spectral evolution of the pDNA− PAEMA59 complexes are dependent on pH, showing noticeable differences between pH = 6.0 vs 7.4. To summarize, controlling the solution pH may be crucial in pDNA−polycation complexation, as it impacts polycation solubility, binding characteristics, and the final complex properties. The findings reported herein should aid researchers in drawing more rigorous structure− function correlations in the field of polymeric gene delivery.

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INTRODUCTION A pair of oppositely charged polyelectrolytes can spontaneously bind with each other in the solution state, resulting in complexes of various forms, such as layered films,1 coacervate phases,2 nanoparticles,3,4 or vesicles.5 In the 1980s, nucleic acid delivery emerged as an application of inter-polyelectrolyte binding,3 taking advantage of the ability of polycations to package nucleic acids into positively charged inter-polyelectrolyte complexes (termed “polyplexes”). Numerous polycations have since been developed, to improve not only gene delivery efficacy6,7 but also tissue-targeting ability,8−10 safety,11,12 and polyplex shelf life,13,14 depending on the priority specific to each application. Hydrophilic−cationic diblock copolymers (referred to as “diblocks” from this point forward) offer an especially versatile yet simple polycation structure. In these diblocks, the cationic block can complex with nucleic © XXXX American Chemical Society

acids, whereas the charge-neutral, hydrophilic block can sterically protect polyplexes from aggregation or precipitation. Besides providing colloidal stability, carbohydrate-containing hydrophilic blocks can additionally allow the polyplexes to target certain tissues,10,15,16 confirming the versatility of the diblock architecture. In most polyplex studies where diblocks and plasmid deoxyribonucleic acid (pDNA) are complexed, two assumptions are commonly accepted, although seldom noted explicitly: (i) the diblocks are highly water soluble in neutral-pH environments because both blocks are water soluble, and (ii) the hydrophilic blocks are rather “added-on” components that Received: August 8, 2017 Revised: January 15, 2018

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DOI: 10.1021/acs.jpcb.7b07902 J. Phys. Chem. B XXXX, XXX, XXX−XXX

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The Journal of Physical Chemistry B

Figure 1. Chemical structures and schematic drawings of the polycations used in this study: (a) PAEMA homopolymer, (b) PEG-b-PAEMA, (c) P(OEGMA)-b-PAEMA, and (d) PMAG-b-PAEMA. The PAEMA is colored orange, and the hydrophilic blocks are in blue. Note that the drawings are not to scale; the actual degree of polymerization of each block ranges from 45 to 64 (rather than 8).

simply provide colloidal stability or targeting ability, as opposed to representing a critical factor in the polyplex formation process. In general, the term “water soluble” is used somewhat loosely, meaning that some may say a polymer is water soluble when it dissolves in water and forms a clear solution, whereas others may say so only when the polymer fully dissolves as single molecules. To avoid confusion, it may be important to confirm and report the specific solution state of the polymer. In many schematic drawings of polyplex formation provided in the literature, the starting materials (pDNA and diblocks) are depicted to be single-molecularly dissolved and the polyplexes are depicted to be stable and micellelike, with an interpolyelectrolyte core and a discrete corona composed of the hydrophilic block.17−20 Although this perspective, along with the above-mentioned assumptions, can be valid for many systems, it may be useful and potentially critical to experimentally confirm and report the solution behavior of the starting materials. In fact, some studies21,22 have reported that hydrophilic−cationic diblocks may aggregate in water at neutral pH and that the presence of a hydrophilic block can alter the extent of pDNA−cationic block binding. Specifically, in our recent work,22 a higher hydrophilic-to-cationic block length ratio was shown to induce less changes in pDNA secondary structure upon binding, implying weaker binding strength, and to allow fewer cationic units to effectively access pDNA. The main goals of this work are to further elucidate diblock aggregation behavior and to examine the impact that the hydrophilic block architecture can have on polyplex properties. Diblocks containing different hydrophilic blocks and a common cationic block of poly(N-(2-aminoethyl) methacrylamide) (PAEMA) were systematically compared. Three distinctly structured hydrophilic blocks were utilized: poly(ethylene glycol) (PEG), representing the least bulky architecture, poly(2-deoxy-2-methacrylamido glucopyranose) (PMAG), containing carbohydrate pendants that introduce intermediate bulkiness, and poly(oligo(ethylene glycol) methyl ether methacrylate) (P(OEGMA)), representing the most bulky counterpart. A PAEMA homopolycation of similar length

was also utilized as a control. Figure 1 summarizes the four polycation structures studied herein. PEG-b-PAEMA and PMAG-b-PAEMA have previously been complexed with pDNA and have shown efficacy for gene delivery in vitro,10,23 where both displayed comparable or lower cellular toxicity than that of a commercial transfection agent. Although P(OEGMA)b-PAEMA has not been tested in DNA delivery studies to our knowledge, P(OEGMA) is chosen in this study as an architectural variation of PEG. PEG and P(OEGMA) are chemically similar but distinct in their physical structure, where P(OEGMA) is comb-structured and thus bulkier and stiffer at a given backbone length. In fact, a comparison between linear PEG and P(OEGMA) as hydrophilic blocks has been reported by Venkataraman et al.;24 with a fixed cationic block of poly(2dimethylaminoethyl methacrylate), polyplexes containing P(OEGMA) hydrophilic blocks displayed poorer in vitro delivery of a reporter gene (luciferase) than those containing PEG hydrophilic blocks. Also, a higher hydrophilic-to-cationic content ratio has been shown to lead to a larger aspect ratio of polyplexes, which correlated to better blood retention.25,26 Altogether, a comparison between linear PEG and its bulkier counterpart P(OEGMA) is useful, as the differences in their polyplex formation process are less known relative to those in their biological performance. This study carefully evaluates the solution states of hydrophilic−cationic diblocks and elucidates the role that hydrophilic block architecture and pH can play in pDNA− diblock interactions. Starting with a fixed contour length of the PAEMA block (≈20 nm), the three hydrophilic (PEG, P(OEGMA), and PMAG) blocks were synthesized to achieve similar contour lengths of the final diblocks (37 ± 2 nm). This was to compare hydrophilic block bulkiness in pDNA−diblock complexation. First tested was the hypothesis that the diblocks (Figure 1) may aggregate in aqueous solution under physiologically relevant pH conditions. The results shed light on the issue of diblock water solubility by showing that complete molecular dissolution may not occur even when the solution is visibly clear and both blocks are individually water soluble. As pH increases above 6.0, the diblock solubility B


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approximately 10 mL. The PEG macroCTA was then precipitated with excess diethyl ether three times and dried under vacuum at room temperature. The final product (2.12 g; yield = 91%) was a yellow solid and was stored in a dark, sealed container at 4 °C. The P(OEGMA) macroCTA was prepared by RAFT polymerization of OEGMA (Mn = 500). Briefly, OEGMA (4.9 g, 9.0 mmol), CEP (47 mg, 0.18 mmol), and 4,4′-azobis(4cyanopentanoic acid) (V-501) (3.0 mg, 0.011 mmol) were dissolved in a mixture of 1 M acetate buffer (pH = 5) and ethanol (4:1). After purging with nitrogen, polymerization was performed at 70 °C for 8 h. The reaction was terminated by exposure to air and simultaneous cooling. P(OEGMA) macroCTA was purified by dialysis against ultrapure deionized water. To polymerize AEMA from the PEG and P(OEGMA) macroCTAs, the procedure reported for PMAG-b-PAEMA synthesis23,29 was followed. Briefly, AEMA·HCl (200 mg, 1.2 mmol), V-501 (0.55 mg, 0.0020 mmol), and either of the macroCTAs (achieving a [AEMA]/[macroCTA] ratio of 62 as the target degree of polymerization) were dissolved in 0.1 M acetate buffer (pH = 5). After purging with nitrogen, polymerization was performed at 70 °C and terminated by exposure to air and cooling. PMAG-b-PAEMA and PAEMA were also prepared via RAFT polymerization, following previously reported procedures.22,23,29 All polycations (Figure 1) were dialyzed against ultrapure water and lyophilized before use. MacroCTA and Polycation Characterization. For PAEMA, P(OEGMA) macroCTA, PMAG macroCTA, P(OEGMA)-bPAEMA, and PMAG-b-PAEMA, the number-average molecular weights (Mn) and dispersities (Đ) were determined by size exclusion chromatography (SEC) in conjunction with multiangle light scattering. An Agilent 1260 Infinity system (Agilent Technologies, Santa Clara, CA) equipped with Eprogen CATSEC100, CATSEC300, and CATSEC1000 columns (Promigen Life Sciences, Downers Grove, IL), a Wyatt DAWN HELEOS II light scattering detector, and a Wyatt Optilab T-rEX refractometer (Wyatt Technology Corporation, Santa Barbara, CA) was utilized. The refractive index increment (∂n/∂c) of each polymer was independently measured using a Wyatt Optilab T-rEX refractometer. The SEC eluent consisted of a 1 wt% aqueous solution of acetic acid with 0.1 M sodium sulfate (pH = 3). The block lengths in P(OEGMA)-b-PAEMA and PMAG-b-PAEMA were obtained by comparing the Mn values (obtained from multiangle light scattering SEC as described above) of P(OEGMA) or PMAG macroCTA with the corresponding diblock (Table 1). The PEG macroCTA was also characterized via both electrospray ionization mass spectrometry using a Bruker BioTOF II (Bruker Corporation; Billerica, MA) and 1H NMR spectroscopy using a Bruker Avance III, from which Mn values were obtained. PEG-bPAEMA was characterized by NMR, from which the PEG-toPAEMA block length ratio was determined. Characterization results of all polycations are summarized in Table 1. Note that the total contour lengths of the diblocks are estimated to differ by less than 10%. pKa Measurement. Each polycation was titrated with an aqueous solution of NaOH, during which the solution pH was recorded using a Metrohm 719 S Titrino (Herisau, Switzerland). Specifically, the polycation was dissolved in 20 mL of 20 mM HCl (aq) to reach the AEMA moiety molarity of 1.4 mM and was titrated using 20 mM NaOH (aq). The derivative of

decreases and aggregates appear (even when the homopolymer of each block molecularly dissolves). Second, it was hypothesized that the hydrophilic block architecture plays a role in pDNA−polycation complexation and polyplex size. This was tested by tracking the polyplex size and size dispersity via dynamic light scattering (DLS) during the titration of pDNA with each polycation. With pDNA in excess, a positive correlation was found between hydrophilic block bulkiness (in the order of P(OEGMA) > PMAG > PEG) and the polyplex size. Isothermal titration calorimetry (ITC) was utilized in parallel and demonstrated that the enthalpic effects of pDNA−diblock binding are actually not a strong function of the hydrophilic block architecture. The third hypothesis tested was that the solution pH and thus the PAEMA block charge density impact the thermodynamics of pDNA−diblock binding. The amount of endothermic binding between pDNA and P(OEGMA)-b-PAEMA was measured as a function of pH, revealing its correlation with pH and elucidating how the binding tendency of each charged moiety is influenced by charge density. Collectively, both the hydrophilic block architecture and the PAEMA protonation state are important parameters in polyplex formation, contributing to the resulting polyplex size, dispersity, and colloidal stability, as well as the complexation thermodynamics. Our findings provide fundamental insights to controlling polyplex physical properties, which are important for nonviral gene carrier development and corresponding biological applications.

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EXPERIMENTAL METHODS Materials. pCMV-LacZ (PF462) was purchased from PlasmidFactory GmbH & Co. KG (Bielefeld, Germany). Oligo(ethylene glycol) methyl ether methacrylate (OEGMA; Mn = 500) was purchased from Sigma-Aldrich Co. LLC (St. Louis, MO) and was purified via alumina column filtration before use. N-(2-Aminoethyl) methacrylamide hydrochloride (AEMA·HCl) was purchased from Polysciences Inc. (Warrington, PA). 2-Deoxy-2-methacrylamido glucopyranose was synthesized as previously reported. 2 7 4-Cyano-4[(ethylsulfanylthiocarbonyl)sulfanyl]pentanoic acid (CEP) was purchased from Strem Chemicals Inc. (Newburyport, MA), and 4-cyano-4-[(propylsulfanylthiocarbonyl)sulfanyl]pentanoic acid was synthesized as previously reported.28 Sodium phosphate (monobasic, monohydrate) was purchased from Mallinckrodt Chemicals (Phillipsburg, NJ). All other chemicals were purchased from Sigma-Aldrich and used without further purification. Methods. Polymer Syntheses. For PEG-b-PAEMA and P(OEGMA)-b-PAEMA, PEG and P(OEGMA) macro chain transfer agents (macroCTAs) were prepared and then AEMA was polymerized via reversible addition-fragmentation chain transfer (RAFT) polymerization. First, to prepare the PEG macroCTA with a contour length of 19 nm, carbodiimide coupling between PEG monomethyl ether (mPEG; Mn = 2000) and CEP was performed via a modified literature procedure.28 mPEG (2.06 g, 1.03 mmol), CEP (543 mg, 2.06 mmol), and 4(dimethylamino)pyridine (126 mg, 1.03 mmol) were dissolved in 20 mL of dichloromethane (DCM). Into this mixture, N,N′dicyclohexylcarbodiimide (389 mg, 2.37 mmol) dissolved in 5.6 mL of DCM was added dropwise, while stirring at 0 °C. The reaction mixture was stirred at 0 °C for 12 h and then at room temperature for an additional 24 h. Dicyclohexylurea was removed via filtration, and the resulting clear solution was concentrated using a rotary evaporator until the volume was C


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The Journal of Physical Chemistry B Table 1. Polycation Characterization Data degree of polymerization

a

polycation

hydrophilic block

PAEMA block

Mn (kg/mol)

Đ

contour length (nm)

PAEMA59 PEG45-b-PAEMA59 P(OEGMA)51-b-PAEMA60 PMAG52-b-PAEMA63

0 45 51 52

59 59 60 63

7.7 12 40 21

1.05 N/Aa 1.34 1.10

20 39 36 37

NMR analysis was used for Mn determination, and thus Mw was not obtained.

Figure 2. Dissolved states of (a) PAEMA59 (pKa = 6.96), (b) PEG45-b-PAEMA59 (pKa = 7.14), (c) P(OEGMA)51-b-PAEMA60 (pKa = 6.97), and (d) PMAG52-b-PAEMA63 (pKa = 7.36), as functions of pH. The hydrodynamic size (Rh) distributions shown were obtained utilizing the regularized positive exponential sum (REPES) inversion program,28 and the y-axis is the relative intensity-weighted population. AEMA moiety molarity is 3.6 mM in all cases.

150 mM or higher salt concentrations.23 In all experiments, [N]0 (starting concentration of AEMA moieties in polycation solution) was 3.6 mM and [P]0 (starting concentration of phosphate groups in pDNA solution) was 0.12 mM. For polyplex formation, a polycation solution (filtered through a 0.45 μm syringe filter) was injected into a pDNA solution (filtered through a 0.45 μm syringe filter) stepwise, until the desired N/P ratio, the number ratio between the AEMA moieties and pDNA phosphate groups, was reached. The N/P ratio increment (ΔN/P) at each injection point was between 0.2 and 2 and will be specified for each technique described below. In every titration, the polycation and pDNA solutions were prepared from the identical phosphate buffer. Dynamic Light Scattering (DLS). Particle diffusion coefficients (D) were measured at 25 °C using a Brookhaven BI200SM instrument equipped with a programmable goniometer, a Mini-L30 laser (wavelength = 637 nm), and a BI-9000AT autocorrelator (Brookhaven Instruments Corporation, Holtsville, NY). For the polymer-only samples (polymers dissolved in 5 mM phosphate buffer), intensity autocorrelation functions

each titration curve (pH vs volume of added NaOH (aq)) was taken and fit to a double Gaussian distribution model, and the average pH value between the two Gaussian peak positions was assigned as the pKa (Figure S1 in the Supporting Information). Polyplex Formation. All pDNA and polycation solutions were prepared with phosphate buffers. The buffers consisted of 5 mM aqueous solutions of sodium phosphate monobasic monohydrate, and the desired pH (between 6.0 and 7.7) was achieved using NaOH. The buffer concentration of 5 mM was chosen to guarantee the buffering capacity to be higher than the polyelectrolyte concentration (in terms of amine or phosphate molarity), and the corresponding final ionic strength was 11 (± 4) mM. These low-ionic strength environments were chosen on the basis of the fact that in the literature, DNA and polycations are often initially mixed in pure water or nonphysiological buffers, even when final delivery is performed under physiological conditions. For example, in a previous work utilizing PAEMA and PMAG-b-PAEMA, polyplexes were formed in a low-salt environment (pure water with no salt added), whereas the final delivery was performed in media with D


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Figure 3. Circular dichroism (CD) spectra of the polymers of interest at (a) pH = 6.0 and (b) 7.4. The y-axis is the solution ellipticity normalized by [AEMA] (black) or [hydrophilic block repeat unit] (the rest). *The data for PAEMA, PMAG, and PMAG-b-PAEMA in (b) were obtained and reported previously22 and are replotted here for comparison.

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RESULTS AND DISCUSSION Water Solubility of Polycations. Each polycation (Table 1) was dissolved in 5 mM phosphate buffer with varying pH, and the solution state was explored by DLS. Figure 2 summarizes the results, elucidating the pH range over which each polycation is well dissolved (Rh ≤ 6 nm) vs partially aggregated. Although the PAEMA59 homopolymer exists as single molecules over a wide pH range, all three diblocks form a population of aggregates (Rh > 20 nm) as pH increases. In acidic environments (pH ≤ 6.0) where the PAEMA blocks are more than 90% charged, none of the polycations form large aggregates (see the caption of Figure 2 for the pKa values). There is no clear correlation between the pKa and the hydrophilic block structure, which may partially be due to the fact that at pH values near the nominal pKa, the diblocks form aggregates of various sizes rather than remaining well dissolved. It should be noted that Rh measured via DLS is weighted by scattering intensity,31 meaning that the molecularly dissolved population is still dominant, number-wise, at pH = 7.4 in Figures 2c and 2d. In the field of polymeric nucleic acid delivery, hydrophilic− cationic diblocks are often presumed to be well-dissolved molecules under nonbasic conditions, whereas it is rare to find rigorous experimental proof for the fully dissolved state. Figure 2 demonstrates that this is not always guaranteed, meaning that the solution state of a diblock may change throughout the polyplex formation process if pH is not controlled. The aggregation of diblocks comprising two water-soluble blocks has previously been discussed and attributed to a hydrophilicity difference between the two blocks.32,33 More specifically, aggregation of hydrophilic−cationic diblocks has been reported to be pH-responsive, allowing reversible transitions between the molecularly dissolved state and a supramolecular structure.34 As pH increases from 6.0 to 7.4, the PAEMA block degree of protonation decreases from over 90% to 30 − 50%, exposing the PAEMA hydrophobicity. The diblock behavior shown in Figures 2b−d can therefore be understood as aggregation in a nonselective solvent, driven by a solubility discrepancy between the two blocks. The solubility test for homopolymers of the hydrophilic blocks (Figure S2 in the Supporting Information) confirms that PEG45 and P(OEGMA)51 are, as expected, highly water soluble. Realizing that PAEMA, PEG, and P(OEGMA) homopolymers themselves do not aggregate even at pH = 7.4, the aggregation behavior of PEG45-b-PAEMA59 and

were obtained at a 90° scattering angle. For the other samples containing pDNA, scattering data were typically obtained from at least four different scattering angles per sample. D (and consequently hydrodynamic size via the Stokes−Einstein relation) distributions were obtained via regularized positive exponential sum (REPES) inversion30 of the autocorrelation functions. For monomodally distributed samples, the autocorrelation fitting analysis described previously22 was performed to determine the mean hydrodynamic radii (Rh). For bimodally distributed samples, autocorrelation functions were fit to a double exponential decay model to determine the Rh of each population. In polyplex preparation for DLS measurements, the initial pDNA solution volume (at N/P = 0) was always 0.24 mL and ΔN/P was equal to N/P ratio difference between the two adjacent measurement points. Isothermal Titration Calorimetry (ITC). Enthalpy changes during polyplex formation were measured using a MicroCal PEAQ-ITC Automated Instrument (Malvern, Westborough, MA) instrument at 25 °C. The solvents were 5 mM phosphate buffers (6.0 ≤ pH ≤ 7.4) prepared as above. Each ITC experiment consisted of two isothermal titrations: a main titration, where a polycation solution ([N]0 = 3.6 mM) was injected into a pDNA solution ([P]0 = 0.12 mM; initial volume = 0.20 mL) prepared with the matching buffer, and a background titration, where a polycation solution ([N]0 = 3.6 mM) was injected into the matching buffer with no pDNA. The N/P increment (ΔN/P) was 0.22 in all cases. During each titration, heat flow was recorded and integrated with respect to time (using MicroCal ITC-ORIGIN Analysis Software), providing the total instantaneous enthalpy change at each injection point. This conversion from heat flow into enthalpy is allowed because the system is at constant pressure with no noncompression work involved. The enthalpy change due to pDNA−polycation interactions (ΔHint) was determined by subtracting the background titration heat from the main titration heat, point by point. All ITC data presented in this article are in their raw form (ΔHint at each injection point vs N/ P), and data fitting using equilibration-assuming models was avoided. Polyplex formation is typically kinetically controlled, meaning that upon each injection of polycation the system will form a new kinetically accessible state (rather than reaching an equilibrium state). Therefore, even though the enthalpy measured at each step during pDNA−polycation mixing can be discussed at face value, fitting models that assume equilibration are not readily applicable. E


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Figure 4. Evolution of scattering intensity-weighted mean hydrodynamic size (Rh; left y-axis) and size dispersity (right y-axis) of complexes during the titrations of pDNA with polycations. pDNA ([P]0 = 0.12 mM) was titrated with (a) PAEMA59, (b) PEG45-b-PAEMA59, (c) P(OEGMA)51-bPAEMA60, and (d) PMAG52-b-PAEMA63 via stepwise injections. [N]0 = 3.6 mM and pH = 6.0 in all cases. The polyplexes formed with PEG45-bPAEMA59 at N/P = 1.5 (black arrow in (b)) precipitated after a week.

study, the cause of diblock aggregation seems to vary depending on the hydrophilic block chemistry. To summarize, aggregation of PEG45-b-PAEMA59 and P(OEGMA)51-bPAEMA60 at high pH is attributed to lower PAEMA charge density and consequently increased hydrophilicity difference between the two connected blocks. The bulkiest hydrophilic block, P(OEGMA), is more effective than PEG at limiting the extent of diblock aggregation. PMAG52-b-PAEMAE63, although showing comparable pH-dependent solubility to P(OEGMA)51-b-PAEMA60, is unique in that the PMAG itself has a tendency to partially aggregate (potentially via intermolecular hydrogen bonding). Complexation between pDNA and Highly Charged Polycations. At pH = 6.0, all four polycations are fully soluble in the 5 mM phosphate buffer (Figure 2) and are more than 90% protonated, representing densely charged systems. We determined that the effects of a hydrophilic block structure on polyplex formation would be more fairly evaluated using welldissolved polycations, rather than with partially aggregated systems. Thus, the sizes and dispersities of polyplexes were measured via DLS during the titration of pDNA with each polycation at pH = 6.0 (Figure 4). The primary hypothesis tested in this experiment was that a hydrophilic block, even though it does not directly interact with pDNA, impacts the polyplex size and dispersity. More specifically, we sought to determine whether a bulkier hydrophilic block would result in formation of a larger polyplex at given N/P ratios and concentrations. This prediction originally stems from the previously reported finding that a higher hydrophilic-to-cationic block length ratio leads to larger polyplexes.22 For the initial titration stages (N/P ≤ 1), the abovementioned hypothesis is found to hold; the evolution in polyplex size and dispersity varies with hydrophilic block architecture, and the average Rh is larger with bulkier

P(OEGMA)51-b-PAEMA60 must stem solely from two watersoluble but chemically different blocks being connected. This can be explained by solvation effects at the molecular level. For the PAEMA homopolymer, even when only sparsely charged, the advantage of aggregation (i.e., decreasing the area where deprotonated AEMA units and water meet) is smaller than the disadvantage (i.e., depleting water around protonated, charged AEMA units). However, the disadvantage may be diminished when a hydrophilic block (PEG or P(OEGMA)) is present due to the lowered density of charged AEMA moieties within the aggregates. In other words, when PAEMA is sparsely charged, adding a hydrophilic block can (counterintuitively) lower the polycation water solubility. In the case of PMAG-b-PAEMA, there is another driving force for aggregation. PMAG itself exhibits pH-dependent solubility (Figure S2c in the Supporting Information). This may stem from the impact of pH on the hydrogen bonding within and between PMAG molecules. Not only are most polysaccharides capable of hydrogen bonding, leading to intra- and intermolecular assembly, but Gress et al.35 have also shown that amide-glucopyranose hydrogen bonding can induce directional polymer self-assembly. Also, the circular dichroism (CD) spectra of PMAG (Figure 3) resemble those of helical macromolecules (e.g., a strong negative band near 215 nm)36−39 and all polymers except PMAG and PMAG-bPAEMA display negligible CD signals in the wavelength range between 185 and 250 nm. PMAG secondary structure is speculated to be helical and to contribute to the relatively low water solubility of PMAG, realizing that the PMAG homopolymer exhibits stronger ellipticity than PMAG-bPAEMA, particularly at pH = 6.0 (Figure 3a) where the diblock is well dissolved. Although the connection between the pH-dependent solubility and the secondary structure is a subject of ongoing F


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Figure 5. Evolution of ΔHint during the titrations of pDNA with (a) PEG45-b-PAEMA59, (b) P(OEGMA)51-b-PAEMA60, and (c) PMAG52-bPAEMA63. [N]0 = 3.6 mM, [P]0 = 0.12 mM, and pH = 6.0 in all cases.

are slightly smaller than those formed with P(OEGMA)51-bPAEMA60, the difference is minor compared to the difference seen in the initial stages (N/P ≤ 1). PMAG52-b-PAEMAE63 is distinct in that polyplexes continue to show signs of aggregation, with higher Rh and dispersity for N/P ≥ 1.5. We tentatively attribute this behavior to the secondary structure (helix) formation of PMAG, resulting in lower solubility compared to that of PEG- or P(OEGMA)-based polyplexes. In summary, the effect of the hydrophilic block is most evident in the low-N/P regime, where polymer aggregation is not a critical factor, whereas the hydrophilic block solubility (and thus the polyplex colloidal stability) becomes important in the high-N/P regime. It is important to realize that DLS provides the mean Rh and dispersity, but information about the actual polyplex shape is extremely difficult to extract, especially when there is polyplex shape heterogeneity. Even with a low dispersity (< 0.2), only a relatively narrow distribution of polyplex diffusion rates can be guaranteed, but not a well-defined shape or aspect ratio. Transmission electron microscopy is the most (if not the only) direct and accurate method of evaluating shapes for heterogeneous populations, and it should be noted that all discussion of DLS data in this article are limited to the Rh and dispersity values calculated on the basis of the distribution of diffusion rates. To evaluate the heat of binding between pDNA and welldissolved polycations, ΔHint was measured during pDNA− polycation titrations at pH = 6.0. The goals were to verify whether the complexation is an entropy-driven process and to observe the effects of varying the hydrophilic block on the binding enthalpy. Figure 5 presents the results, where in all cases, the complexation is endothermic, confirming that it is entropy-driven, and is completed by N/P ≈ 1.5. The maximum ΔHint is observed around N/P ≈ 1 and is comparable to thermal energy (≈ 600 cal/mol). The rapid decrease in ΔHint between N/P ratios of 1 and 1.5 indicates that most of the amine−phosphate pairing occurs at N/P < 1.5; this is consistent with the observation that the polyplex structures remain constant after N/P passes 1.5. Despite its pDNAcompacting ability (Figure 4), PEG45-b-PAEMA59 reveals the lowest ΔHint upon binding pDNA. This again suggests that the endothermic interactions are correlated with amine−phosphate pairing, rather than with pDNA structural changes. Here, one should note that ΔHint refers to the enthalpy (heat) of formation of a kinetically trapped polyplex state and is therefore potentially mixing route dependent. As shown in Table S1 in the Supporting Information, different polyplex structures are

hydrophilic block architecture. PEG45-b-PAEMA59 forms bimodally distributed polyplexes at N/P ≤ 1, with one population being similarly sized to the pDNA alone (∼80 nm) and one significantly more compact (∼25 nm). Even though bimodal distributions of PEG-based polyplexes have previously been predicted and observed,17,40,41 polyplex properties in the low-N/P regime have rarely been discussed. P(OEGMA)51-b-PAEMA60 forms the largest (∼150 nm) polyplexes in this initial regime, consistent with the prediction. PMAG52-b-PAEMA63 forms monomodal polyplexes (dispersity ≈ 0.2) that are similarly sized to pDNA. A more compact structure is achieved compared to P(OEGMA)51-b-PAEMA60, but there is no indication of a population smaller than that of pDNA (as observed with PEG45-b-PAEMA59). In the middle stage of the titrations (1 ≤ N/P ≤ 2), all four curves exhibit distinct increases in Rh. With the PAEMA59 control (Figure 4a), a monotonic increase in Rh is observed throughout the relatively small range of N/P ratios, except for the outlier at N/P ≈ 0.2. As N/P exceeds 1, the lack of a hydrophilic block leads to poor colloidal stability, immediately resulting in large aggregates (ca. 300 nm). With PEG45-bPAEMA59, the polyplexes formed at N/P ≈ 1.5 initially have Rh ≈ 150 nm but over time (a week) severely aggregate and eventually precipitate. Polyplexes formed with the other two diblocks (Figures 4c and 4d) do not precipitate even after a month, but they still show clear signs of aggregation; the dispersity values are high near N/P ≈ 1.5, and the average Rh values (≈ 250 nm) are significantly larger than those at N/P ≤ 1. Note that on the basis of the polycation pKa values, the overall polyelectrolyte charge of the system switches from negative to positive as the N/P ratio increases from 1 to 1.5. The charge switch corresponds to the aggregation point, suggesting that global charge neutrality can result in poor colloidal stability of polyplexes, even when the polycation is equipped with a hydrophilic block. In the terminal stages of the titrations (N/P ≥ 2), no further substantial changes in either average Rh or dispersity are observed with any of the diblocks, suggesting that stable, constant structures have been reached. In this regime, the hypothesis of the bulkiness effect no longer holds; the stable Rh does not have a clear correlation with the hydrophilic block bulkiness. We attribute this to the colloidal stability coming into play. As N/P increases above 2, both the PEG45-b-PAEMA59 and P(OEGMA)51-b-PAEMA60 systems overcome the tendency to aggregate near N/P ≈ 1.5 and exhibit dispersity below 0.3. Although the polyplexes formed with PEG45-b-PAEMA59 G


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Figure 6. Evolution of Rh (left y-axis) and size dispersity (right y-axis) of complexes during the titrations of pDNA with (a) PEG45-b-PAEMA59, (b) P(OEGMA)51-b-PAEMA60, and (c) PMAG52-b-PAEMA63. [P]0 = 0.12 mM, [N]0 = 3.6 mM and pH = 7.4 in all cases. Note: the data in (c) were obtained from our previous report.22

Figure 7. Evolution of ΔHint (after background subtraction) during the titrations of pDNA with P(OEGMA)51-b-PAEMA60 at (a) pH = 6.6, (b) 7.0, and (c) 7.4. [N]0 = 3.6 mM and [P]0 = 0.12 mM in all cases. (d) Total enthalpy change, i.e., summation of ΔHint recorded at every injection point, normalized by the PAEMA block degree of protonation.

impact of the hydrophilic block architecture on binding is also supported by CD measurements (Figure S3 in the Supporting Information) and is important, considering that various hydrophilic blocks are under development to advance beyond the traditional PEG, and thereby avoids reduction of interactions with cell membranes43 or accelerated clearance upon repeated dosing.44 Meanwhile, rather noticeable structural differences among the polyplexes were observed by DLS (Figure 4). We thus conclude that the endothermic interactions between pDNA and a cationic block do not strongly depend on the hydrophilic block chemistry and architecture but the polyplex structure and stability do. Effects of Polycation Charge Density. Studying low-pH systems is advantageous in that polycations are well dissolved (Figure 2) and thus no effect from aggregation is present. However, neutral-pH conditions are more often used for polyplex formation, especially in biological studies.18,45 To

achieved via different routes of pDNA−polycation mixing. It is, therefore, important to realize that the binding enthalpy presented in Figure 5 is specific to the corresponding stepwise mixing route. Meanwhile, PEG45-b-PAEMA59 is distinct in that an initial increase in ΔHint is observed (N/P < 1), suggesting binding cooperativity, which has previously been observed by Kim et al.42 in a similar system. Although the physical basis of this apparent cooperativity will be the subject of a future study, it can be at least partially attributed to the need for complexation initiation. At low N/P ratios, the probability for “binding sites” (phosphates and amines) to encounter each other may be positively correlated with how much binding has already happened. Altogether, however, there is no drastic difference in pDNA binding thermodynamics among the diblocks (Figure 5), considering the magnitudes of ΔHint or the end points (the N/P ratios at which ΔHint reaches zero). Such (minimal) H


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charged amines to pair with the phosphates, resulting in higher normalized Σ(ΔH). At high charge densities, because the polycation chain is stiffer and also because the pDNA double helix cannot bend easily, it may be difficult for all positive charges to bind with phosphates. Effects of Charge Density on Interactions between pDNA and PAEMA59. When pH is varied in the diblock systems, not only the charge density but also the dissolved state of the diblock is altered (Figure 2). Comparison between Figures 4 and 6 indicates that the PEG-based polyplex formation and properties are influenced by pH more noticeably than either of the P(OEGMA)- or PMAG-based counterparts, but one should recall that aggregated diblock populations are present at pH = 7.4. This introduces difficulty in differentiating the effects due to charge density from those due to polycation aggregation. Thus, to exclude the aggregation effect, the titration of pDNA with PAEMA59 homopolycation was studied under various conditions of pH ≤ 7.7; PAEMA59 does not aggregate over this pH range, as shown in Figure 2a. Very similar properties are observed for the pDNA− PAEMA59 polyplexes formed at pH = 7.7 (Figure 8) vs 7.4

elucidate the differences caused by altering pH and therefore PAEMA block charge density, polyplex sizes and dispersities were recorded during polyplex formation at pH = 7.4, where pDNA solutions were titrated with the diblocks of interest (Figure 6). With PEG45-b-PAEMA59 (Figure 6a), distinct differences are observed compared with the same experiment at pH = 6.0 (Figure 4b). In the initial stage (N/P ≤ 1), the bimodal size distributions are absent, although the polyplexes still remain more compact than those formed with P(OEGMA)51-b-PAEMA60. Possibly, being sparsely protonated, the PAEMA block no longer can effectively compact pDNA. At N/P ≈ 1.5, a large population (> 200 nm) emerges and persists throughout the rest of the titration. Lastly, the high-N/P polyplex size is significantly larger compared to that in the more highly charged case, even though the size dispersity remains below 0.3. On the other hand, with P(OEGMA)51-b-PAEMA60 (Figure 6b), the impact of pH is relatively minor. Except for suppression of aggregation at N/P ≈ 1.5, the trends in both size and dispersity are virtually the same as those observed with the same diblock at pH = 6.0 (Figure 4c). In the case of PMAG52b-PAEMAE63 (Figure 6c) as well, the size and dispersity evolution show a modest dependence on pH; polyplexes with Rh below 100 nm are formed at N/P ≤ 1, which aggregate at N/P ≥ 2. We attribute the small pH effect to the fact that unlike the PEG block, the P(OEGMA) and PMAG blocks inhibit the formation of compact polyplexes even at pH = 6.0. When the pDNA-compacting ability of a diblock is absent even with a high charge density, the resulting polyplex size may be less sensitive to polycation charge density. In summary, although the Rh increase during the middle stage of titrations (N/P ≈ 1.5) persists regardless of diblock structure and charge density, the charge density plays a noticeable role in the case of PEG45-b-PAEMA59, in both initial and terminal regimes of polyplex formation. To systematically study the effects of charge density, a single diblock, P(OEGMA)51-b-PAEMA60, was chosen and ITC profiles were obtained under various pH conditions. Figure 7 summarizes the results, where the evolution of ΔHint proves to be dependent on the PAEMA charge density. As suggested in the discussion of Figure 5, the endothermic effect is considered to stem mainly from pairing between polycation amines and pDNA phosphates. Figures 7a−c, along with Figure 5b, imply that a higher pH leads to lower total amine−phosphate pairing in the initial stages of the titrations (N/P ≤ 1). This is intuitively reasonable, as there are simply fewer protonated amines available for binding with phosphates. At the same time, however, the endothermic complexation tends to be slightly protracted to higher N/P ratios at higher pH. Consequently, the total binding enthalpy change, Σ(ΔH), is not a simple function of pH. This, in fact, gives rise to a pH-dependence of normalized Σ(ΔH) (Figure 7d). The amount of endothermic binding normalized by the amount of protonated amines has a positive correlation with pH (i.e., a negative correlation with the degree of protonation). More sparsely charged PAEMA blocks result in either stronger amine−phosphate pairing or larger portion of the charged amines participating in binding. We propose an explanation for this (somewhat counterintuitive) phenomenon, on the basis of the flexibility difference between pDNA and PAEMA. Figure S4 in the Supporting Information describes the flexibility effects on amine− phosphate binding; with low charge density, the flexible polycation may take conformations that allow most of the

Figure 8. Evolution of Rh (left y-axis) and size dispersity (right y-axis) of complexes during the titration of pDNA with PAEMA59 at pH = 7.7. [P]0 = 0.12 mM and [N]0 = 3.6 mM. The Rh data points that are off scale indicate precipitation.

(Figure S5 in the Supporting Information). In both cases, complete precipitation occurs as the N/P ratio exceeds a threshold value (1 or 1.2). Such behavior seems to be a characteristic of sparsely charged systems because at pH = 6.0 (Figure 4a), no precipitation is observed even when the N/P ratio reaches 2. This leads to the hypothesis that the pDNA− PAEMA59 binding mode (i.e., how the polycation binds to and alters the pDNA double helix) may be different at pH of 6.0 versus 7.4 or 7.7; the location and manner of the amine− phosphate interactions may be contingent on the PAEMA charge density. To test this, CD spectra of pDNA were recorded during titration with PAEMA59 at various pH values. The CD signals mainly originate from nucleobases stacked within pDNA helix,46,47 and pDNA−polycation binding can alter the stacking pattern, leading to changes in CD spectra. In 1974, Zama48 reported the effects of DNA concentration and polycation type (polylysine vs polyarginine) on the CD spectra, where both of the polycations reduced the positive CD signals at 280 nm under dilute conditions. Further, even negative signals were induced at 280 nm in some cases, described as an “anomaly”, which was attributed to structural regularity in aggregation (ordered aggregation).48 This is important to note for homopolycation-based polyplex studies because such I


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Figure 9. Evolution of CD spectra of pDNA during the titration with PAEMA59 at (a) pH = 6.0, (b) 7.4, and (c) 7.7. The y-axis is the solution ellipticity divided by the phosphate group molarity. [N]0 = 3.6 mM and [P]0 = 0.12 mM in all cases. Note: (b) has been adapted and reprinted with permission from ref 22. Copyright 2017 American Chemical Society.22

when DNA becomes single-stranded. We thus speculate that densely charged PAEMA59 may partially fray or melt the double helix due to stronger binding. Regardless of charge density (Figures 9a vs 9b and 9c), PAEMA59 binds to pDNA strongly enough to alter the π → π* transitions within the nucleobases. However, we observe two notable differences between the densely vs sparsely charged cases. First, when densely charged, the resulting polyplexes do not precipitate even at N/P = 2 (Figure 4a), whereas the sparsely charged counterparts show poorer colloidal stability and precipitate (Figure 8) in the buffered conditions. In fact, this relates to Figure S4 in the Supporting Information, which illustrates the effect of pH on polyplex formation due to the backbone flexibility difference between DNA and PAEMA. In the densely charged case, not all charged amines are able to pair with phosphates due to pDNA stiffness; this implies local overcharging (+/− > 1) in the complexation of pDNA and PAEMA, which may contribute to the colloidal stability of the polyplexes. In the sparsely charged case, because PAEMA is flexible, more complete amine−phosphate pairing is feasible, leading to charge neutrality and thus poorer stability. It should be noted that although the stiffness of unbound PAEMA likely increases slightly with increasing charge density (i.e., decreasing pH), it will surely remain more flexible than DNA, considering that most single-chain polyelectrolytes have significantly shorter persistence lengths than those of double-stranded DNA.55−58 This flexibility difference holds even at the ionic strengths used in this study (∼11 mM) because the persistence length of a polyelectrolyte with a flexible backbone displays only a logarithmic dependence on ionic strength and its variation is thus limited to about a factor of three when the ionic strength decreases from a physiological value (∼102 mM) to 11 mM.57,59−61 Considering that the effect of charge screening on flexibility resembles that of a decrease in charge density, any pH-dependence of PAEMA flexibility is assumed to have at most a modest impact on the discussion regarding Figure S4, where the incomplete cationic charge neutralization results in overcharged (+/− > 1) polyplexes. Second, via CD spectroscopy, the pDNA secondary structure is found to evolve differently during polyplex formations at different pH, implying that the pDNA−polycation binding mode also depends on the PAEMA charge density (Figure 9). It should be realized that changes in pDNA secondary structure, measured via CD, do not necessarily correlate with changes in the overall polyplex structure. Wilson and Bloomfield have previously observed that

systems tend to aggregate easily and are often delivered into cells in an aggregated state, without an understanding of the effect of aggregation on DNA structure. For the PAEMA case, however, there is no sign of an anomaly, and there is no reason to expect ordered aggregation (Figure 9). Choosakoonkriang et al.49 have previously reported the effects of homopolycation architecture on the DNA CD signals, where linear poly(ethylene imine) induced a greater reduction in the positive signal (280 nm) than its branched counterpart. Altogether, there are numerous factors contributing to polyplex CD spectra but an overall redshift and a decrease in positive signals are two of the most commonly observed features in polyplex formation. Although these features hold for the PAEMA system as well (Figure 9), the CD evolution recorded at pH = 6.0 is distinct from those at pH = 7.4 or 7.7, supporting the above hypothesis. In Figure 9a, the entire spectrum redshifts, showing a noticeable increase in the negative signals (250 nm), as the N/ P ratio increases. In Figures 9b and 9c, however, both the positive (275 nm) and the negative (250 nm) CD signals decrease throughout the titration and the positive signals essentially disappear when excess PAEMA59 is added (N/P > 1.2). Considering that the pDNA phosphate groups are fully charged within the pH range explored here, the way that the pDNA helical structure is altered during polyplex formation seems to be dictated by the PAEMA charge density. In the sparsely charged systems (Figures 9b and 9c), the decrease in the positive signals (275 nm) resembles that observed in the classical B → C transition in which the DNA axial pitch per base pair and the double helix diameter decrease.50,51 Changes in the negative signals around 250 nm, on the other hand, are often less systematic, and therefore most studies lack a clear explanation for them.49,52 The sparsely charged PAEMA systems, for example, exhibit a slight increase and then a strong decrease in this wavelength region with increasing N/P (Figures 9b and 9c). Although a physical interpretation of such a trend is difficult, it has previously been claimed that the CD signal around 250 nm is correlated with the helicity or handedness of the double helix.53,54 We thus suspect that the later decrease in the signal (at N/P > 1.2) indicates weakening of the pDNA helicity via strong binding. For the densely charged system (Figure 9a), it is unclear what the strong signal increase (doubling) at 245−250 nm represents in a physical sense. It is, though, worthwhile to mention Johnson and Tinoco’s calculation,46 which showed that the negative signals are larger than the positive signals J


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densely charged (pH = 6.0), a gradual shift in the CD spectrum was observed, suggesting potential melting of the double helix. The findings reported herein demonstrate the effects of hydrophilic block architecture and solution pH on polyplex formation or, more generally, on polyion−diblock interactions. The insights into various physical parameters in pDNA− polycation complexation gained from this study should aid in advancing polymeric nucleic acid delivery.

even a significant collapse of a DNA molecule into a compact polyplex can be accompanied by no change in the secondary structure.62 They suggested that the hydration state may be more directly related with the secondary structure, which could be another reason why the higher charge density and the potential overcharging in the low-pH system (Figure 9a) lead to a different CD response from the sparsely charged counterparts.

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CONCLUSIONS PEG-b-PAEMA, P(OEGMA)-b-PAEMA, and PMAG-bPAEMA diblocks were evaluated regarding their solution behavior as well as their pDNA complexation. It was discovered that hydrophilic−cationic diblocks are not guaranteed to be molecularly dissolved in water under physiological pH conditions (near 7). Although most of the hydrophilic homopolymers themselves were highly water soluble, their diblock counterparts aggregated as pH increased from 6.0 to 7.4. Diblock aggregation is attributed to the solubility difference between the hydrophilic and cationic blocks, rather than to inherent hydrophobicity of PAEMA alone, because the PAEMA homopolymer dissolved well at 6.0 ≤ pH ≤ 7.4. This finding underscores that careful understanding of the materials and the fundamental physical properties in the field of polyplexes may be valuable, as diblock solution behavior can be difficult to predict. The diblocks resulted in distinct polyplexes as well. At low N/P ratios (< 1), the hydrophilic block bulkiness was positively correlated to the mean Rh of polyplexes. A large portion of the polyplexes formed with PEG45-b-PAEMA59 were compact (less than 50% size of the bare pDNA), whereas those formed with P(OEGMA)51-b-PAEMA60 were larger than bare pDNA. At high N/P ratios (> 1.5), stable structures were reached with all diblocks, whose Rh values were determined mainly by the polyplex hydrophilicity rather than the hydrophilic block architecture. Meanwhile, enthalpic effects of pDNA−diblock complexation were found to be similar among all three diblocks, being weakly endothermic (ΔHint ∼ 102 cal/mol). Even though PEG45-b-PAEMA59 was unique in that it showed apparent binding cooperativity in the initial stage of titration, we conclude that the impact of hydrophilic block bulkiness on the pDNA−PAEMA59 block interactions is minimal. Charge density effects were also elucidated. With P(OEGMA)51-b-PAEMA60, the amount of endothermic binding per charged amine increased with increasing pH. In other words, lowering charge density enhanced the binding tendency of each charged amine, which can be explained by flexibility differences between pDNA and PAEMA. The homopolycation control (PAEMA) system further demonstrated other critical effects of charge density via both DLS and CD experiments. The polyplexes formed with PAEMA59 experienced more severe aggregation at lower charge density (pH ≥ 7.4), potentially due to more effective charge neutralization. No precipitation was observed at higher charge density (pH = 6.0), although the colloidal stability was still poorer than with the diblocks. We hypothesize that the pDNA−PAEMA59 binding mode (i.e., alterations in pDNA secondary structure upon binding) is different at low vs high charge densities, which is supported by the pDNA secondary structure changes tracked using CD spectroscopy. When sparsely charged (pH ≥ 7.4), PAEMA59 induced reductions of both positive and negative CD signals, implying a decrease in the pDNA double helix axial pitch. When PAEMA59 was

ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.jpcb.7b07902. Polycation pKa determination; solution states of PEG, P(OEGMA), and PMAG homopolymers at various pH values; schematic description of amine−phosphate pairing at different cationic charge densities, size, and dispersity evolution of polyplexes formed with PAEMA59 at pH = 7.4; and the properties of polyplexes formed via one-step mixing in unbuffered water (PDF)

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AUTHOR INFORMATION

Corresponding Authors

*E-mail: [email protected] (T.P.L.). *E-mail: [email protected] (T.M.R.). ORCID

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

The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript. 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. Isothermal titration calorimetry was carried out using a MicroCal PEAQ-ITC microcalorimeter, funded by the NIH Shared Instrumentation Grant S10-OD017982. We acknowledge Dr. Quanxuan Zhang, Dr. Yaoying Wu, and Anatolii Purchel for their help with polycation synthesis and characterization.

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REFERENCES

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