Distribution of Ionizable Groups in Polyampholyte Microgels Controls

Article Cite This: Biomacromolecules 2019, 20, 1578−1591

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Distribution of Ionizable Groups in Polyampholyte Microgels Controls Interactions with Captured Proteins: From Blockade and “Levitation” to Accelerated Release Wenjing Xu,†,§,‡ Andrey A. Rudov,⊥,†,‡ Ricarda Schroeder,†,§ Ivan V. Portnov,⊥ Walter Richtering,# Igor I. Potemkin,*,⊥,†,∥ and Andrij Pich*,†,§,¶ Biomacromolecules 2019.20:1578-1591. Downloaded from pubs.acs.org by UNIV AUTONOMA DE COAHUILA on 04/09/19. For personal use only.

†

DWI-Leibniz Institute for Interactive Materials e.V., Forckenbeckstraße 50, 52074 Aachen, Germany Functional and Interactive Polymers, Institute of Technical and Macromolecular Chemistry, RWTH Aachen University, Forckenbeckstraße 50, 52074 Aachen, Germany ⊥ Physics Department, Lomonosov Moscow State University, GSP-1, 1-2 Leninskiye Gory 119991 Moscow, Russian Federation # Institute of Physical Chemistry, RWTH Aachen University, Landoltweg 2 52056 Aachen, Germany ∥ National Research South Ural State University, Chelyabinsk 454080, Russian Federation ¶ Aachen Maastricht Institute for Biobased Materials (AMIBM), Maastricht University, Brightlands Chemelot Campus, Urmonderbaan22, 6167 RD Geleen, The Netherlands §

S Supporting Information *

ABSTRACT: A striking discovery in our work is that the distribution of ionizable groups in polyampholyte microgels (random and core−shell) controls the interactions with the captured proteins. Polyampholyte microgels are capable to switch reversibly their charges from positive to negative depending on pH. In this work, we synthesized differently structured polyampholyte microgels with controlled amounts and different distribution of acidic and basic moieties as colloidal carriers to study the loading and release of the model protein cytochrome c (cyt-c). Polyampholyte microgels were first loaded with cyt-c using the electrostatic attraction under pH 8 when the microgels were oppositely charged with respect to the protein. Then the protein release was investigated under different pH (3, 6, and 8) both with experimental methods and molecular dynamics simulations. For microgels with a random distribution of ionizable groups complete and accelerated (compared to polyelectrolyte counterpart) release of cyt-c was observed due to electrostatic repulsive interactions. For core− shell structured microgels with defined ionizable groups, it was possible to entrap the protein inside the neutral core through the formation of a positively charged shell, which acts as an electrostatic potential barrier. We postulate that this discovery allows the design of functional colloidal carriers with programmed release kinetics for applications in drug delivery, catalysis, and biomaterials.

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release of the payload from microgels.6,16−18 Since the pH in, e.g. cancer cells, differs from the surrounding pH, a controlled docking to specific cells (via functionalization with specific receptors) with subsequent controlled release (via the change in pH) can be realized.19 Based on their unique architecture, microgels enable capacious uptake and controlled release of architecturally complex macromolecular species.20 Microgels containing ionizable groups are well suited to operate as drug delivery systems because potential drugs can be bound via electrostatic interactions and easily be released through a change in pH or the addition of salt. The swelling behavior of microgels and thus their effective storage volume can easily be tuned by a variation in pH, ionic strength and the number of incorporated charges.21 For instance, Y. Li et al. synthesized oxidized, negatively charged starch microgel and

INTRODUCTION Extensive studies have been performed in the last years focusing on the use of microgels as delivery systems for drugs and proteins due to their facile functionalization and their high biocompatibility.1−7 Microgels can transport hydrophobic drugs that are otherwise insoluble,8 enhance their metabolic stability,9 offer protection against immunogenicity10 or degradation,11 and enhance the drug’s circulation time within the body.12,13 Through the right choice of monomers at the stage of microgel synthesis, the balance between hydrophilicity/hydrophobicity, the presence of charges at specific pH, the size and the response to external stimuli can be tuned. Drugs can be attached to the microgel polymer network either by chemical (covalent) bounds or physical (electrostatic) binding. Chemical binding is often used if diffusional leaking is undesired and the release of the drug is realized through the degradation of the whole polymer network (i.e., by the use of cleavable cross-linkers)14,15 Physical binding offers the opportunity to use pH as an external stimulus to trigger the © 2019 American Chemical Society

Received: December 19, 2018 Revised: February 28, 2019 Published: March 1, 2019 1578

DOI: 10.1021/acs.biomac.8b01775 Biomacromolecules 2019, 20, 1578−1591

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Biomacromolecules

Figure 1. Scheme of microgels used in present study. N, pure PNIPAm microgel; V, pure PVCL microgel; V−, anionic PVCL microgel; V±, PVCL polyampholyte microgel with random distribution of ionizable groups; V−N+, core−shell polyampholyte microgel consisting of an anionic PVCL core and a cationic PNIPAm shell.

studied the uptake and release of fluorescently labeled, positively charged lysozyme via confocal laser scanning microscopy.22 They showed that binding of proteins has an electrostatic nature. Increasing the salt concentration or changing the pH value triggers the release of the lysozyme. They also found out that the protein could bind to multiple binding sites inside the microgel at lower protein concentration leading to a decreased mobility of the protein. M. Smith and L. A. Lyon used negatively charged poly-N-isopropylacrylamideco-acrylic acid (PNIPAm-co-AAc) microgels to study the influence of charge density on the uptake of cyt-c.23 Multiangle light scattering (MALS) measurements demonstrated that the molecular weight MW of the loaded particles increased with increasing AAc content indicating that large amounts of cyt- c are bound by the polymer network. At the same time, the microgel radius decreased due to a release of counterions and electrostatically driven attraction of charged species.24−26 T. Hoare and R. Pelton, however, were the first to address the importance of the distribution of ionizable groups within the polymer network on the uptake and release of drugs.27 The authors synthesized PNIPAm microgels containing different acids such as acrylic acid, methacrylic acid, vinylacetic acid, and fumaric acid as comonomers. Due to differences in the reactivity parameters, microgels with various distribution of carboxylic groups were formed. They showed that different extents of loading could be achieved depending on the location of the ionizable groups. Though the presence of carboxylic groups enables the binding of oppositely charged protein via electrostatic interactions, a change to lower pH renders the microgel neutral, i.e. the release of protein is solely based on diffusion. A more pronounced and effective release, however, can be achieved via repulsion, i.e. microgel and protein have the same charges through a change in pH. This can be realized by using polyampholyte microgels that contain both acidic and basic moieties and exhibit an isoelectric point (IEP), i.e. they are positively charged at low pH while being negatively charged at pHs above the IEP. Conversely, there are reports on the interactions of polyelectrolyte microgels with proteins, but to the best of our knowledge, no detailed study on the polyampholyte microgels. Different from polyelectrolyte microgels, polyampholyte microgels are able to switch reversibly their charge from positive to negative as a function of pH.28−31 This unique property allows the coexistence of two important pH-triggered functions in polyampholyte microgels: (a) efficient binding of proteins by electrostatic attraction forces and (b) stimulation

of release of proteins accelerated by electrostatic repulsion forces. Under the hypothesis that the distribution of ionizable groups of opposite charges in microgels in random or core− shell fashion may control the pH-triggered binding and release of proteins, we studied the interactions of a series of polyampholyte microgels of known chemical composition and internal structure (Figure 1) with a model protein. In our study cytochrome c (cyt-c) was chosen as a model protein. Cyt-c has a small size of Mw ∼ 12 kDa, thus hypothetically could freely penetrate through the microgel pores and it has positive charges over a wide pH range due to its isoelectric point at pH 10.2. Microgels used in the present study are based on poly-N-vinylcaprolactam (PVCL) and poly-N-isopropylacylamide (PNIPAm) as main monomers. This not only gives the microgels temperature responsive characters but with the incorporation of itaconic acid (IA, pKa: 3.84 and 5.55) and 1vinylimidazole (VIm, pKa: 6.0) by copolymerization approach they also show pH sensitivity. Due to the presence of ionizable groups, they could undergo abrupt and reversible changes in volume in response to both temperature as well as pH changes. This multisensitivity was used to study the controlled release of cyt-c as a response to the pH. Hereby, polyampholyte microgels with different distribution (random or core−shell) of ionizable groups within the polymer network as well as reference microgels (polyelectrolyte or neutral analogues) were synthesized and tested as carrier systems. The polyampholyte microgels were characterized by means of light scattering measurements, electrophoresis, and transmission electron microscopy. The experimental uptake and release kinetics of cyt-c by polyampholyte microgels (random and core−shell) at different pH values of the outer surroundings was investigated by UV−vis. The surface of a protein is extremely complex, containing varying degrees of hydrophobic/hydrophilic residues, chemical moieties, and charges. The precise interpretation and prediction of protein absorption, localization, and orientation in the microgels, therefore, is difficult. Nevertheless, we have shown experimentally that both PVCL and PNIPAm microgels demonstrate low hydrophobic contribution to the protein uptake and release. The major role is played by the electrostatic interactions between cyt-c and the charged functional groups within the microgels. We demonstrate for the first time that core−shell distribution of the ionizable groups in the microgel can serve as an additional tool to entrap the protein inside the carriers. Namely, we show that similarly charged shell of the microgel could lock the proteins inside the 1579

DOI: 10.1021/acs.biomac.8b01775 Biomacromolecules 2019, 20, 1578−1591

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Biomacromolecules Table 1. Reagents Used for the Synthesis of the Reference Microgels VCL

NIPAm

IA

BIS

AMPA

Sample

g

mmol

g

mmol

g

mmol

g

mmol

g

mmol

ratio VCL/NIPAm:IA (mol %)

N V V20−

− 0.461 0.314

− 3.312 2.256

0.375 − −

3.314 − −

− − 0.070

− − 0.538

0.015 0.015 0.015

0.097 0.099 0.099

0.013 0.013 0.011

0.047 0.05 0.05

− − 80:20

Table 2. Reagents Used for the Synthesis of the Microgels with a Random Distribution of Ionizable Groups VCL Sample ±

V10 V20± N20±

BIS

IA

VIm

AMPA

g

mmol

g

mmol

g

mmol

g

mmol

g

mmol

ratio VCL:VIm:IA (mol %)

0.803 0.800 0.650

5.769 5.750 5.744

0.040 0.044 0.039

0.260 0.285 0.259

0.093 0.249 0.187

0.715 1.914 1.436

0.068 0.180 0.135

0.723 1.913 1.436

0.031 0.047 0.035

0.114 0.173 0.129

80:10:10 60:20:20 60:20:20

Table 3. Reagents Used for the Synthesis of Core Microgels VCL

BIS

IA

AMPA

Sample

g

mmol

g

mmol

g

mmol

g

mmol

Remarks

V10−

1.876

13.478

0.065

0.422

0.175

1.343

0.053

0.194

anionic core

Table 4. Reagents Used for the Synthesis of the Shell NIPAm

BIS

VIm

AMPA

Sample

g

mmol

g

mmol

g

mmol

g

mmol

ratio IA:VIm in core:shell (mol %)

N10+ 5N10+

1.572 3.790

13.892 33.492

0.062 0.189

0.402 1.223

0.126 0.315

1.339 3.347

0.056 0.152

0.208 0.559

1:1 1:5

Synthesis of Pure PVCL, PNIPAm, and Anionic PVCL Reference Microgels. Pure PVCL and PNIPAm microgels (V and N) were all synthesized via one-step free-radical precipitation polymerization.1 The appropriate amount of the monomer (VCL or NIPAm) and cross-linker (BIS) (Table 1) were first mixed in 30 mL of distilled water under N2 atmosphere at 70 °C in a round-bottom flask under constant stirring for 30 min. The reaction was afterward started by the addition of the initiator AMPA and allowed to polymerize for 2 h. For the anionic PVCL microgel (V20−), the appropriate amount monomers (PVCL and IA) and cross-linker (BIS) (Table 1) were mixed in 30 mL. The reaction mixture was regulated to pH 10 and heated up to 70 °C under N2 atmosphere for 1 h under continuous stirring followed by the addition of AMPA to start the reaction. The polymerization was allowed to react for 2 h. All microgels were dialyzed directly after the polymerization using a composite regenerated cellulose membrane from Millipore (NMWCO 12 000−14, 000) for 5 days. Synthesis of Microgels with a Random Distribution of Ionizable Groups. Microgels with a random distribution of ionizable groups were synthesized via one-step free-radical precipitation polymerization.28 Appropriate amounts of monomers (VCL/ NIPAm, VIm, IA) and cross-linker (BIS) (Table 2) were first mixed in 80 mL of distilled water under N2 atmosphere at 70 °C in a double-wall reactor under constant stirring for 1 h. The reaction was afterward started by the addition of the initiator AMPA and allowed to polymerize for 4 h. The synthesized microgel solution was then directly cleaned via dialysis using a composite regenerated cellulose membrane from Millipore (NMWCO 12 000−14 000) for 5 days. Synthesis of Microgels with Core−Shell Distribution of Ionizable Groups. Microgels with a core−shell distribution of ionizable groups were synthesized in a two-step approach employing free-radical precipitation polymerization in aqueous media wherein the first step core microgels were synthesized followed by the addition of the shell in the second step. For the core-microgel, appropriate amounts (Table 3) of VCL, IA, and BIS were dissolved in 150 mL of distilled water and heated up to 70 °C while purging with N2 in a stirred round-bottom flask. After 1 h, the initiator AMPA was added and the reaction was carried out for 2 h under constant stirring. After

neutral core via only electrostatic repulsion of the proteins from the shell. Such metastable state of the blocked proteins can exist very long time (at least months) due to the high potential barrier whose value is primarily controlled by the fraction of charged groups in the shell. In other words, local repulsion of the proteins from the similarly charged shell enforces them to be localized inside the microgel core rather than be expelled into the outer solvent which is thermodynamically more favorable but kinetically impossible. On the other hand, a random distribution of cationic and anionic groups leads to an accelerated release of the proteins (compared to the polyelectrolyte analogue) due to the repulsion between charged proteins and cations of the microgel. We also perform the molecular dynamic simulations to expand understanding of processes occurring in the system. We demonstrate the electrostatic nature of uptake and release of the proteins and their trapping by the core−shell microgels. We also analyzed the regimes of weak, medium, and strong hydrophobic interactions between the proteins and the microgels. Our model not only reproduces the experimental data but also extends our knowledge about the location of proteins within the microgels at different pH values.

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MATERIALS AND METHODS

Reagents. N-Vinylcaprolactam (VCL, 98%), itaconic acid (IA, ≥99%), 1-vinylimidazole (VIm, ≥99%), N-isoprylacrylamide (NIPAm, 97%), cytochrome c (cyt-c, from equine heart), 2,2′azobis[2-methylpropionamidine] dihydrochloride (AMPA, granular, 97%), and N,N′-methylene(bis)acrylamide (BIS, 99%) were purchased from Sigma-Aldrich. VCL and NIPAm were recrystallized from hexane and dried under vacuum before use. Water used in the experiments was purified using a Millipore water purification system with a minimum resistivity of 18 MΩ*cm. 1580

DOI: 10.1021/acs.biomac.8b01775 Biomacromolecules 2019, 20, 1578−1591

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Biomacromolecules the synthesis, the core microgels were remained in the round-bottom flask without cooling down or cleaning. Subsequently, the appropriate amounts of NIPAm, VIm, BIS, and AMPA (Table 4) for the shell synthesis were dissolved in preheated distilled water (70 °C, 100 mL) and quickly added to the core dispersion. The reaction was again allowed to continue at 70 °C for a further 2 h. The core−shell microgel dispersions were cleaned by dialyzing for 5 d against water using a composite regenerated cellulose membrane from Millipore (NMWCO 12 000−14 000). Protein Loading Experiments. 5 mg of microgel was redispersed in 3 mL of a buffer with pH 8 at room temperature. 0.89 mg of cyt-c (1:106 excess to the added microgels, even after taking into account that itaconic acid possesses two carboxylic groups) was added into the microgel solution, and the mixture was allowed to stir overnight to reach equilibrium. At this state, maximum loading of cyt-c into the microgel carriers should be achieved. Before the start of dialysis (5 L), UV−vis measurements from 300−800 nm using a UV−visible spectrophotometer (CARY 100 Bio from Agilent Technologies, USA) were conducted at 25 °C, the value of the characteristic peak intensity (I0) at 530 nm was obtained. The samples were afterward dialyzed in a composite regenerated cellulose membrane from Millipore (NMWCO 30 000) in a buffer at pH 8 in order to remove free swimming (not bound to the microgels) cyt-c for 24 h. UV−vis measurements were repeated again where a decrease in the peak intensity of the characteristic peak at 530 nm could be observed (Ix). The fraction of bounded cyt-c to the microgels was calculated by using the following formula: x% = (Ix/I0) × 100%. Protein Release Studies. The microgel samples preloaded with cyt-c in pH 8 at room temperature were dialyzed against dialysis water (5 L). During dialysis, UV−vis measurements were carried out. The following parameters were changed: the pH of the dialysis water was changed from pH 8 to pH 3 or pH 6. The measurements were all carried out at room temperature and salt concentration of the dialysis water was kept at 30 mM. Determination of Amounts of Ionizable Groups by FTIR Spectroscopy. The amount of itaconic acid and imidazole groups in microgels (Table S1) was determined by FTIR spectroscopy as reported in the previous work.30 Fourier transform infrared spectroscopy (FTIR) spectra (resolution 4 cm−1) were recorded using a Nicolet NEXUS 670 Fourier transform IR spectrometer. Samples were mixed with KBr powder and then pressed to form a transparent KBr pellet before measurement. Dynamic Light Scattering and Electrophoretic Mobility. The hydrodynamic radius RH and the electrophoretic mobility of the microgels were measured using a Zetasizer NanoZS (Malvern, UK). Unless otherwise noted, measurements were taken at 25 °C after equilibrating the samples for at least 15 min pH trends were measured from 3 to 10 in 0.5 steps using 0.1 M HCL and NaOH; respectively. Before all measurements, the samples were filtered with a 1.2 μm PTFE filter. Static Light Scattering. The molecular weight of the microgel particles was measured with static light scattering using a Fica goniometer (SLS Systemtechnik, Germany) with a laser of 543 nm. The samples were measured in an angle range of 25° to 145° with 5° intervals. For each sample, five dilutions were prepared with concentration from 0.1 to 2 mg/mL. Prior to SLS measurements, the changing refractive index increment (dn/dc, see Table S2) of polyampholyte microgels was determined using a refractometer (SLS Systemtechnik, Germany). Both measurements were performed at 25 °C. Transmission Electron Microscopy. Morphologies of polyampholyte microgels were detected by TEM using a Zeiss Libra 120 (Zeiss, Germany). For the sample preparation, one drop of the microgel dispersion was deposited at room temperature on a TEM copper grid (Formvar carbon film, 400 mesh, Plano GmbH, Germany) and dried overnight. TEM was operated at an acceleration voltage of 120 kV. The images were recorded using a CCD camera system (Ultra Scan 1000, Gatan, Germany). Ultraviolet−visible Spectroscopy. UV−vis spectra were taken at a UV−visible spectrophotometer using a CARY 100 Bio (Agilent

Technologies, USA). A given concentration of microgel solution in buffer was measured at room temperature in the range of 200−800 nm using 1.5 mL UV-cuvette semimicro (12.5 mm × 12.5 mm × 45 mm). Computer Simulations. Brownian molecular dynamics (MD) simulations within a standard coarse-grained model with implicit solvent were performed at the supercomputer JURECA, Jülich Supercomputing Centre.32 The LAMMPS package was used.33 Without loss of generality, the simulations were carried out using dimensionless units, where the fundamental quantities such as mass m, the diameter of the bead σ, and the Boltzmann constant kB are considered to be equal to 1. All beads of the microgels (including monomer units, cross-links, and charged groups), protein molecules and counterions were modeled as Lennard-Jones particles with the same diameter σ = 1 and mass m = 1. The equations were integrated with a time step Δt = 0.005τ, where τ = σ(m/ε)1/2 is the standard time unit for a Lennard-Jones fluid. We set T = 1.0ε. The calculations were carried out in NVT ensemble. The Ewald summation technique with 10−5 precision was used to compute the electrostatic interactions. The microgels were designed as described in details in refs 30, 31. In brief, fully stretched subchains of an ideal microgel (all subchains have equal length) were connected through tetrafunctional cross-links in such a way to repeat a unit cell of the diamond crystal lattice. Then cubic microgel template consisting of 15 × 15 × 15 modified unit cells mentioned above was constructed. The fraction of the cross-links in the template was approximately equal to 5% so that the number of beads in each subchain was 10. To provide spherical shape to the microgel particles, a sphere was inscribed into the template. Then all beads outside the sphere were “cut off”. The remaining beads have been assigned a type denoted as V (by analogy with experiment). In the case of the core−shell microgel, two concentric spheres were inscribed into the frame. All the beads located in the inner sphere forming the core of the microgel have been assigned a type V. The beads of the template located in-between the inner and outer spheres forming the shell of the microgel were denoted as N. The rest of the beads outside the outer sphere were “cut off”. As a result, we have designed a series of different microgels: homogeneous core microgel consisting of 3500 beads, V = (3500:0), symmetric core−shell microgel, VN = (3500:3500) and asymmetric core−shell microgels with more bulky shells, V3N = (3500:10500) and V7N = (3500:24500). The digit before N defines how many times the number of beads in the shell is greater than in the core. The microgels were modeled as pH-sensitive. The fraction of cationic and anionic groups depends on pH. Only cationic groups are “switched on” at pH 3 and only anionic at pH 8. . At the intermediate pH values, the fractions of the ions of both sorts change linearly (one increases and other decreases) with pH: the electrophoretic mobility is a linear function of pH.30,31 Maximum fraction of cationic (at pH = 3) and anionic groups (at pH = 8) in the microgel set to value 10% from the number of beads of the shell and the core, respectively (see Table 5). In the case of polyampholyte microgel with random distribution of ionizable groups, V±, both anionic and cationic groups are randomly (homogeneously) distributed throughout the whole gel. For polyampholyte core−shell microgels, V−N+, V−3N+, and V−7N+, the anionic and cationic groups are distributed either in the core or in the shell regions, respectively. Each charged group has an absolute

Table 5. Characterization of Microgels in Computer Simulations Amount of neutral beads Sample ±

V V−N+ V−3N+ V−7N+ 1581

Amount of ionizable beads

Fraction of ionizable beads

nV

nN

n+

n−

φ−

φ+

ratio n−:n+

3500 3500 3500 3500

0 3500 10500 24500

350 350 350 350

350 350 1050 2450

0.1 0.1 0.1 0.1

0.1 0.1 0.1 0.1

1:1 1:1 1:3 1:7

DOI: 10.1021/acs.biomac.8b01775 Biomacromolecules 2019, 20, 1578−1591

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Biomacromolecules

× 106), the saturated number of the proteins in the microgels corresponds to the equilibrium state. To avoid dependence of the number of bound proteins on their total number in the simulation box, the system was “dialyzed”: each microgel with absorbed proteins was transferred into empty box (pure solvent) with further annealing. After t = 106 simulation steps, some small number of the proteins escaped the microgel. To double check the equilibrium number of the captured molecules, we also considered an excess number of the proteins in the microgels as starting structures. The release of the proteins caused by pH switching from 8 to 6 or 3 was calculated as a number of remaining captured proteins vs time.

value of the elementary charge and the corresponding counterions provide macroscopic electric neutrality of the system. Cytochrome c is a small globular protein. The primary, secondary, and tertiary structures of cytochrome c are already well studied. The presence and distribution of hydrophobic, hydrophilic, and charged groups in the protein structure are established. We propose a simple model for this globular protein, reflecting its main physical properties. Namely, we consider soluble, spherical, monodisperse proteins having identical surface structure. We design them in the following way (Figure S13). Each protein consists of 123 beads of 3 types: single central bead, 114 neutral and 8 charged beads distributed on the sphere of radius 2.5σ. Both Lennard-Jones (L-J) and FENE potentials support the spherical shape of the proteins. Each surface bead is interconnected with 5 neighbor surface beads (stiffness coefficient K0 = 20ε/σ2 and maximum extension of the bond R0 = 2σ) and with the central bead (stiffness coefficient K1 = 30ε/σ2 and maximum extension of the bond R1 = 4σ). The interaction between the surface beads describes via L-J potential, ε1 = 0.01ε, rmin = 1σ. The interaction of the surface beads with the central bead is also described via L-J potential, ε2 = 0.01ε, rmin = 4σ. Such choice of the parameters provides stable nondeformable impenetrable spherical proteins of the constant diameter, D ∼ 4.5σ. Proteins did not aggregate with each other and could freely move within the microgel without any topological constraints. This model of the protein has several advantages including the ability to create ideal spherical globule with defined surface pattern and charge distribution, the acceleration of simulation time due to the neglection of inner inconsiderable beads. We consider 8 elementary cationic groups located on the surface of the nanoparticle. In the pH range of interest, the electrophoretic mobility of cyt-c has positive value monotonously decaying with increasing pH (Figure S7). The electrospray mass spectrum of native cytochrome c in water demonstrates that the most intense charge states of cyt-c in the positive ion mass spectrum are at +6 to +9 while the charge state distribution ranges from +4 to +13.34,35 For the purity of the computer experiments, the total charge of the protein in our model is considered to be constant and equal to +8 independently on pH. Monovalent mobile counterions provide electric neutrality of the proteins. The interactions between any pair of beads were described through the truncated-shifted Lennard-Jones potential. The dimensionless Lennard-Jones interaction parameters εV, εN, and εVN = (εV × εN)1/2, describe bead−bead interactions in the core (εV), shell (εN), and between the core and the shell (εVN). We confine ourselves by consideration of good solvent conditions for the microgels, εV= εN= εVN = 0.01ε, rmin = 1σ. We also consider that the proteins are slightly repelled from the core of the microgel and are not absorbed into the microgel in the absence of charges, εV cyt = 0.1ε, rmin = 1σ. To mimic hydrophobic (short-range attractive) interactions between the shell of the microgel and the proteins, we vary the value εN cyt enhancing attraction between them. We analyze the regimes of weak, medium, and strong hydrophobic interactions between the proteins and the microgel shell, εN cyt = 0.1ε, εN cyt = 0.3ε and εN cyt = 0.5ε, rmin = 1σ. At εN cyt = 0.1ε uptake of the proteins by the microgel is driven only by electrostatics. The total number of the proteins in the simulation box of the volume (150σ)3 is 150. The total charge of the proteins, Qtot = 150·(+8) = +1200 is bigger than the maximum charge of anionic groups in the core of the microgels, Qcore = −350. In other words, we add an excess of proteins which can be absorbed by the microgel in the solution. However, it should be mentioned that after the uptake process, many proteins are not bound to the microgels but are swimming freely in the system. Uptake of the proteins is simulated as follows. First, the microgels are annealed in a good solvent during t = 107 simulation steps. Then the proteins are placed around the microgels. Such placement does not give accurate data on the uptake kinetics, but it is more efficient for calculating the equilibrium number of bound molecules. The number was calculated via (i) analysis of correlation functions of bound and free molecules and via (ii) direct counting of the proteins which belonged to the microgels. Both ways of calculations gave similar results. We can anticipate that after long simulation time (t = 5

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RESULTS AND DISCUSSION Structure and Properties of Polyampholyte Microgels with a Random Distribution of Ionizable Groups. Polyampholyte microgels with random distribution of ionizable groups were successfully synthesized by free-radical precipitation polymerization.28,29,32 The molar ratio between the acidic and basic moieties within the microgels was kept 1:1, but the total amount of ionizable groups was varied with respect to the wanted microgel systems (Table 2). Microgels without ionizable groups and microgels with only acidic groups were used as reference samples. Quantitative FTIR analysis was done to determine the theoretical amounts of acidic and basic groups in the microgels (Note S2 and Table S1).28 The internal structure of polyampholyte microgels with random distribution of ionizable groups (V20±) was confirmed with TEM after staining the samples with uranyl acetate (U(Ac)3). Uranyl acetate was used for selective staining of carboxylic acid groups in polymers, thus increasing the electron density (dark areas) in TEM images. Accordingly, the TEM image in Figure 2A indicates that stained areas rich in carboxylic acid groups

Figure 2. (A) TEM image of randomly distributed polyampholyte microgels at pH ∼ 6 (V20±), the sample were stained with U(AC)3 to visualize the location of the carboxylic groups. (B) Electrophoretic mobility of the microgels. The gray dotted line at EM = 0 is a guide to better see the isoelectric point of the microgels. All measurements were conducted at T = 25 °C.

are randomly distributed through the microgels. Although the resolution of the TEM image is not high enough to observe single polymer chains, it enables the interpretation of the spatial distribution of functional groups within the microgels. For the polyampholyte microgels with random distribution of ionizable groups, we have an average diameter of 110 nm. If the distribution would not be random, localized accumulation of uranyl ions would be observed in the image, similar to the case of core−shell microgels (Figure 3A). The hydrodynamic radii of polyampholyte microgels with random distribution of ionizable groups and the reference microgels as a function of pH were determined by dynamic light scattering (DLS) (Figure S8). Microgels with a random 1582

DOI: 10.1021/acs.biomac.8b01775 Biomacromolecules 2019, 20, 1578−1591

Article

Biomacromolecules

to the microgels, cyt-c has its IEP at 10.2 and is positively charged over the whole pH range from 3 to 8 (Figure S7). Structure and Properties of Polyampholyte Microgels with Core−Shell Distribution of Ionizable Groups. Core−shell distributed polyampholyte microgels were successfully synthesized via two-step free-radical precipitation polymerization with variable molar ratios between acidic and basic moieties (Tables 3 and 4).30,31,43 Like polyampholyte microgels with random distribution of ionizable groups, the amounts of acidic and basic groups in the core−shell distributed polyampholyte microgels were determined by potentiometric titration and quantitative FTIR analysis (Note S2 and Table S1). The internal structure of the microgels was confirmed with TEM where the samples were stained with uranyl acetate (U(Ac)3) before (Figure 3A). Accordingly, the core−shell structure of the synthesized polyampholyte microgels with controlled distribution of ionizable groups could be clearly detected. It can be observed that carboxylic groups are mainly localized in the microgel core (darker part). Dark spots in the shell come from entangled U3+. With DLS, the pH dependency of the core−shell distributed polyampholyte microgels was determined. The hydrodynamic radii exhibit an asymmetric pH-dependent swelling behavior (Figure S9). Core−shell microgels swell significantly at low pH due to the positively charged shell. The shell is able to swell in a larger extent than a negatively charged core at high pH, for which the neutral shell can play a role of a corset restricting the core swelling.31 The EM of core−shell distributed polyampholyte microgels with increasing shell thickness is presented in Figure 3B. For microgel samples V10−N10+ and V10± (Figure 2B), the isoelectric point is at the same position reflecting the same total amount of ionizable groups within the microgels. For V10−5N10+ the isoelectric point is shifted slightly to higher pH for an increasing core−shell ratio. Moreover, the absolute value of the EM for V10−5N10+ is higher at low pH than at high pH due to the fact that negative charges are “hidden” in the microgel core contributing only slightly to the overall EM. The EM of the model protein cyt-c at 10.2 is given (Figure S7) to establish that the binding procedure for polyampholyte core−shell microgels can also be operated at pH 8 where the microgels and cyt-c are oppositely charged. Dynamic (DLS) and static (SLS) light scattering experiments were also performed for core−shell distributed polyampholyte microgels pH 3, 6.5, and 8 providing the hydrodynamic radius RH and the radius of gyration RG, respectively (Figure S1). Uptake and Release of cyt-c by Polyampholyte Microgels with Random Distribution of Ionizable Groups. Polyampholyte microgels with randomly distributed ionizable groups V10± and V20± were loaded with cyt-c at pH 8. The selection of pH was based on the fact that at pH = 8 cyt-c is still positively charged and almost 90% of carboxylic groups in microgels are deprotonated (Figure 2B). In this situation, we could achieve the highest loading of cyt-c into microgels due to electrostatic attraction. In our experiments, both cyt-c and microgel were mixed in a buffer at pH 8 and allowed to stir for 24 h to reach the saturation. Subsequently, the mixture was filled into dialysis tubes and dialyzed for time span of 24 h. After loading of cyt-c into microgels the solutions appear to be intensively red colored. The heme group of cyt-c has a distinctive red color that enables the monitoring of the release both with the naked eye and with UV−vis spectros-

Figure 3. (A) TEM image of core−shell distributed polyampholyte microgels (V10−5N10+) at pH ∼ 6; the sample was stained with U(AC)3 to visualize the location of the carboxylic groups. (B) Electrophoretic mobility of the microgels. The gray dotted line at EM = 0 is a guide to better see the isoelectric point of the microgels. All measurements were conducted at T = 25 °C.

distribution of ionizable groups show a V-shaped curve, i.e. they swell symmetrically at low pH (protonation of imidazole groups) and high pH (deprotonation of carboxylic acid groups). The swelling is slightly more pronounced with the increasing amount of carboxylic acid and imidazole groups (comparing samples V10±, V20±, and N20±). For polyelectrolyte microgels (V20−), the hydrodynamic radius increases linearly at low pH due to the deprotonation of carboxylic groups and reaches equilibrium at around pH 7 and thus no Vshaped curve could be observed here. As expected, the hydrodynamic radius for uncharged microgels (V and N) remained unchanged during pH variation. Both dynamic (DLS) and static (SLS) light scattering experiments were performed at pH 3, 6, and 8, providing the hydrodynamic radius RH and the radius of gyration RG, respectively (Figure S1). The ratio RG/RH gives information about the density distribution within a microgel and thus its internal structure.36,37 A RG/RH ratio of 0.775 is typical for hard spheres, implying a homogeneous density distribution.38 Microgels in the swollen state have loose chains on the particle surface that result in a large RH, but have no influence on RG.39,40 The RG/RH ratio of microgels without ionizable groups was 0.55 and not pH dependent. Those values are comparable to high generation star-like polymers with a very open structure.41,42 In contrast, for polyampholyte microgels considerably lower RG/RH ratios (0.40−0.45) were detected at pH 3 and 8, indicating open porous structure due to strong swelling. Polyampholyte microgels at their IEP (i.e., pH ∼ 6.5) have a RG/RH ratio similar to neutral microgels as a result of the charge compensation. Thus, light scattering results presented in Figure S8 indicate that polyampholyte microgels change their swelling degree and internal structure at different pH values. Furthermore, we measured the electrophoretic mobility (EM) of microgels with randomly distributed functional groups and polyelectrolyte microgel (Figure 2B). The two polyampholyte microgels (V10± and V20±) contain both equal ratios of acidic and basic groups but different total amounts of ionizable groups (Table S1). They possess an IEP at pH ∼ 6. Below the IEP, imidazole groups are positively charged, while above the IEP, the carboxylic groups are deprotonated and therefore negatively charged. Similar to the hydrodynamic radii, the EM values are a bit larger for microgels with a higher amount of ionizable groups. In contrast 1583

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Biomacromolecules Table 6. Amount of cyt-c Loaded to the Microgel Carriers in (% and mg/mg)a Sample

N

V

V20−

N20±

V10±

V20±

Uptake (%) Uptake (mg cyt-c in mg microgel)

23 0.041

21 0.037

55 0.098

57 0.101

46 0.082

53 0.094

a The fraction of bounded cyt-c to the microgels were calculated by the using the following formula: x% = (Ix/I0) × 100%. I0 is the value of the characteristic peak of cyt-c at 530 nm detected by UV−vis before washing with a buffer with pH 8 and Ix is the value detected after washing out the free cyt-c which are not bound to the microgels.

Figure 4. Change of amount of bound cyt-c with dialysis time after change of pH 8 to 3, 6, and 8 for polyampholyte microgels with (A) V10± and (B) V20±. Change of amount of bound cyt-c with dialysis time at pH 3 for polyampholyte microgels V10± and V20±, and the polyelectrolyte microgel V20− as a comparison (C). All measurements were conducted at T = 25 °C. 100% refers to the fraction of bounded cyt-c to the microgels at t = 0, while 0% corresponds to the complete release of cyt-c from the microgels.

microgels to the release procedure. The release of cyt-c from polyelectrolyte microgels was investigated as a reference experiment. After successful loading of the protein into different microgels, we studied the pH-triggered protein release. For each type of microgel carrier, we calculated both the absolute (mg/mg) and relative amounts (%) of released cyt-c. The relative released amounts were determined as a ratio the initial loaded cyt-c (Table 6) to the released quantity of cyt-c. Polyampholyte microgels with random distribution of ionizable groups loaded with cyt-c were kept at pH 8, 6, and 3, and protein release was determined by UV−vis spectroscopy (Figure 4A,B). At pH 8 (strong electrostatic attraction between negatively charged microgel and positively charged protein), approximately 44% (0.036 mg) and 54% (0.051 mg) of loaded cyt-c is released from V10± and V20±, respectively. The probable explanation for this phenomenon is that weakly bound cyt-c, located at the surface of the microgels is desorbing into the aqueous solution. If pH is decreased to 6 (IEP, charge compensation) cyt-c is released much faster from the microgel for both systems. At pH 3 we observed an accelerated release of cyt-c from microgels, which is complete after 6 h. Figure 4A,B indicates that the change in pH switches the interactions between protein and microgel from electrostatically attractive to electrostatically repulsive. The random distribution of ionizable groups in the microgels favors fast and complete release of protein under electrostatically repulsive conditions. For further confirmation of the electrostatic caused phenomena, we compared the release of cyt-c from polyampholyte microgels with the release from polyelectrolyte microgels (Figure 4C). For polyampholyte microgels cyt-c was fully released after ∼21 h while the release from polyelectrolyte microgels was significantly slower. After 7 days, ∼20% (0.0795 mg) of the initial loaded protein amount still remained inside the polyelectrolyte microgels. This result strongly indicates

copy. Though cyt-c has the highest absorbance at 402 and 416 nm (γ-band belonging to Fe(III) and Fe(II), respectively), the smaller peak at 530 nm (α-band) was used as reference peak since this peak is not changing with the oxidation state of the ion in the heme structure. Pure microgel samples show no peaks in this range (Figure S3). The loading experiments of cyt-c were also operated with reference microgels (N, V, V20−, and N20±)in the same way as described in the Materials and Methods for comparison. Table 6 shows the fraction/amount of loaded cyt-c within different microgels. For uncharged microgels systems around 20% (0.041 mg cyt-c bound to 1 mg N and 0.037 mg cyt-c bound to 1 mg V) of the initially added cyt-c were captured after the washing process. This effect can be easily explained by the hydrophobic interactions between the microgels and cyt-c.44 Higher loading of cyt-c was achieved by using charged microgels (V10±, V20−, N20±, and V20±) according to electrostatic attraction forces of two opposite charged systems. At pH 8, we achieved a loading of 46% (0.082 mg cyt-c) cyt-c in 1 mg V10±. For the charged microgels (V20− and N20± and V20±), more cyt-c could be loaded; however, the loading efficiency is not in linear dependency to the number of ionizable groups within the microgels as shown in Table 6. By comparing the polyampholyte microgels V10± and V20± (53% loading, 0.094 mg cyt-c in 1 mg V20±), only slight differences were visible. This phenomenon was already reported by T. Hoare and R. Pelton;27 they reported in their study that when positively charged protein molecules come into contact with a negatively charged microgel particle, the protein will bind to the carboxylic groups most easily accessible, i.e. on the particle surface. Binding of cyt-c leads to a local collapse of the polymer network, thus “blocking” the polymer network for further cyt-y to enter (Figure S5). Taking this into account, the release studies were continued with polyampholyte microgels (V10± and V20±) to investigate the influence of the amount of ionizable group within the 1584

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Biomacromolecules Table 7. Amount of cyt-c Loaded to the Microgel Carriers in (% and mg/mg)a Sample

N

V

V20−

V10−N10+

V10−5N10+

Uptake (%) Uptake (mg cyt-c in mg microgel)

23 0.041

21 0.037

55 0.098

52 0.093

71 0.126

a The fraction of bounded cyt-c to the microgels were calculated by the using the following formula: x% = (Ix/I0) × 100%. I0 is the value of the characteristic peak of cyt-c at 530 nm detected by UV−vis before washing with a buffer with pH 8 and Ix is the value detected after washing out the free swimming cyt-c which are not bound to the microgels.

Figure 5. Change of amount of bound cyt-c with dialysis time after change of pH 8 to 3, 6, and 8 for polyampholyte microgels with (A) V10−N10+ and (B) V10−5N10+ (C) Change of amount of bound cyt-c with dialysis time at pH 3 for polyampholyte core−shell microgels V10−N10+,V10−5N10+ and polyelectrolyte microgels V20− as comparison. All measurements were conducted at T = 25 °C. 100% refers to the fraction of bounded cyt-c to the microgels at t = 0, while x% corresponds to the fraction of released cyt-c from the microgels at t = x.

that the release of cyt-c from polyampholyte microgels with randomly distributed ionizable groups is driven by electrostatic repulsion. Uptake and Release of cyt-c by Polyampholyte Microgels with Core−Shell Distribution of Ionizable Groups. Uptake studies of cyt-c with core−shell distributed polyampholyte microgels were performed in the same way described above for polyampholyte microgels with randomly distributed ionizable groups (at pH 8 where microgels and cytc possess opposite charges). Since two different monomers (VCL and NIPAm) were applied during the synthesis of core− shell microgels, it is necessary to explore whether there is an influence of different monomers to the interaction between microgels and cyt-c. For this, uncharged microgels (V and N) were used to study the effect of the polymer structure on protein uptake. The uptake results from Table 7 show that approximately 20% (0.041 mg cyt-c in 1 mg N and 0.037 mg cyt-c in 1 mg V) of the initially added cyt-c after removing all the free cyt-c (cyt-c not bound to the microgel carriers) could be bound to the uncharged microgel systems. With this knowledge, we ensure that the interaction between cyt-c and the two monomers (VCL and NIPAm) are similar. Therefore, having both monomers within the same gel will not affect the further studies. As expected, the binding ability for the core− shell microgels V10−N10+ analogous the polyelectrolyte microgel V20−. The EMs for both microgel systems are very similar at pH 8 (Figure 2B and Figure 3B) and this probably leads to similar cyt-c binding amounts. By increasing the shell thickness, much more cyt-c is bound to the microgels although at pH 8 the EM value for V10−5N10+ is similar to V10−N10+ (Figure 3B) indicating that besides electrostatic attraction, forces hydrophobic interactions between hydrophobic amino acid residues of the N- and C- terminal α-helices of cyt-c and the hydrophobic backbone of the polymer chain in the microgels can occur.44

Furthermore, we compared the release of cyt-c from core− shell distributed polyampholyte microgels at various pH values of the aqueous solutions (Figure 5A,B). At pH 8, core−shell microgels exhibit a negatively charged core, while the shell is electrically neutral. In this situation, a small amount of protein could escape from core−shell microgels with a thin shell (V10−N10+) but by increasing the shell thickness the microgels are able to delay or completely prevent the cyt-c from release. By reducing the pH to 6 (IEP, charge compensation) ∼40% (0.062 mg) cyt-c is released from the microgel carriers V10− N10+. In contrast, for microgel V10−5N10+ there is no protein release observed indicating that cyt-c is safely bound in the microgel carrier due to the thick shell even after 160 h. This indicates that the hydrophobic forces within the microgel shell are not only responsible for the higher loading of the protein but also prevent its release. A change to pH 3 leaves the core electrically neutral, while the shell is now positively charged. It can be assumed that the protein that was formerly bound via hydrophobic interactions in the shell is now released due to the strong electrostatic repulsion explaining the release of ∼80% (0.074 mg) for V10−N10+ and 70% (0.084 mg) for V10−5N10+ of protein after 24 h. Cyt-c that was bound inside the microgel core via electrostatic attractions is now “trapped” within the microgel network by the positively charged shell. Though partial release of cyt-c was observed, the release itself is considerably slowed down. It has to be mentioned that the presence of long-range electrostatic interactions between the microgels and cyt-c and trapping of cyt-c in the metastable state does not allow analyzing the release process within the Fick’s laws of diffusion. Moreover, we compared the release of the cyt-c at pH 3 from polyelectrolyte microgel V20− with the release from polyampholyte core−shell microgel (Figure 5C). One can notice that after 7 days the core−shell system entraps more than 20% protein and the release is almost completed. Contrary, in the case of the V20− despite a similar amount of 1585

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Biomacromolecules Table 8. Absolute Amount of cyt-c Loaded to the Microgel in Computer Simulationsa Amount of cyt-c in the corea

Total amount of cyt-c in the gel

εN cyt

εN cyt

Sample (ratio Vim:IA)

0.1

0.3

0.5

V± V−N+ V−3N+ V−7N+

− 32 ± 2 33 ± 2 34 ± 2

− 21 ± 2 18 ± 1 20 ± 1

− 17 ± 3 0 0

0.1 43 43 42 40

± ± ± ±

1 1 1 1

0.3

0.5

− 52 ± 1 66 ± 1 75 ± 1

− 52 ± 1 70 ± 0 82 ± 0

a

Amount of central beads of nanoparticles in the sphere of the radius equal to the gyration radius of the core. Center of the sphere was set to the point of center mass of the microgel/protein complex.

subchains occurs. In the symmetric case, when the fraction of positively and negatively charged groups in the microgel is equal, microgels with random distribution of ionizable groups, V±, is characterized by symmetric V-shaped curve.28 The microgels have a nearly homogeneous distribution of polymer throughout the whole volume independently on the pH values. On the contrary, core−shell distributed polyampholyte microgel, V−N+, in the symmetric case is characterized by asymmetric V-shaped curve.29 Microgels with the biggest shell, V−7N+, at higher pH swells more because the absolute amount of the positively charged groups in the shell significantly exceeds the amount of negatively charged groups in the core at low pH values. That is why the isoelectric point of such microgels is shifted to higher pH. A distinctive feature of the core−shell polyampholyte microgel is the dense layer formation at the core−shell interface at intermediate pH values. Such elevation of the concentration is a result of the electrostatic complexation of oppositely charged subchains, which can interpenetrate in the interfacial region and form a dense interpolyelectrolyte complex. The depth of the interpenetration is determined by the core−shell composition of the microgel, length of the subchains and fraction of the charged units. On the basis of our theoretical knowledge with high accuracy, we could assume that experimentally synthesized microgels (Figures S8 and S9) are polyampholyte microgel with random and core−shell distributed ionizable groups and declared VIm:IA ratio. Computer Simulations: Uptake of the Proteins. All the neutral microgels V, VN, V3N, and V7N at εV= εN= 0.01ε, εN cyt = 0.1ε do not absorb proteins. Nevertheless, under the same conditions εV= εN= 0.01ε, εN cyt = 0.1ε at pH 8 polyampholyte microgels capture proteins (Table 8). The only reason, triggering uptake of the proteins in that case, is the electrostatic attraction between the oppositely charged systems. Keeping in mind that the total number of charged groups in the core was 350, around 350/8 ≈ 43 proteins is needed theoretically in order to neutralize them. One could see from Table 8 that the amount of bound cyt-c in the microgel, V±, and core−shell microgel with a thin shell, V−N+, is equal (43). This means that the net charge of the microgels created by the anions was fully compensated by cyt-c. Uptake of the proteins is accompanied by the collapse of the microgels (Figure S14) due to the substitution and release of small monovalent counterions by the proteins acting as multivalent macroions. Absorbed proteins tend to move in the direction of the core of the microgel. While the proteins are distributed homogeneously through the V± microgel in the equilibrium, the proteins in core−shell microgels are located either in the core or in its immediate vicinity. (Figure 6A). Increasing the

protein left in the microgel, the slow release is still continued for a few more weeks (data not shown). These results strongly suggest that the release of cyt-c from polyelectrolyte microgels is diffusion-driven rather than caused by electrostatic repulsion. In addition, we can conclude that the controlled incorporation of ionizable groups in microgels allows designing colloidal carriers with programmed release properties in aqueous solutions. Furthermore, we studied and compared the release kinetic of cyt-c from polyampholyte core−shell microgels V10+N10−, where the positive charges are located inside the core and negative charges in the shell with V10−N10+ (Figure S12). For V10+N10− at pH 8, a maximum loading of 48% (0.085 mg cytc/mg microgel) could be achieved which is similar to that of V10−N10+ (52%). By comparing the release kinetic of the two polyampholyte core−shell microgels with opposite distribution of charges, at pH 3, both microgels possess a fast release. After 48 h, only 20% cyt-c remained inside the microgel carriers. This phenomenon can be explained in the way that for V10+N10−, cyt-c is mostly loaded to the outer shell of the microgel via electrostatics at pH 8, and once the thin shell loses its charge at pH 3, cyt-c will be released very fast, similar to V10−N10+. Since no big difference was detected for the release kinetic at pH 3, no further comparison was performed at pH 8. The different release profiles are of special interest in the biomedical field. Microgels as drug or RNA carriers requires the control over the release profiles as some drugs are only effective in some parts of the body and a too early release will be either ineffective or harmful. These results show that the time frame for the release of proteins can be tuned by controlling the spatial distribution of ionizable groups within polyampholyte microgels. Computer Simulations: Characterization of Microgels. Using the same models and methodology, internal structure and properties of similar polyampholyte microgels with random and core−shell distribution of ionizable groups were investigated in our previous papers.30,31 As it was predicted, both microgels with random and core−shell distribution of ionizable groups reveal a nonmonotonous shape of the swelling curve. At both low and high pH values the microgels swell due to the electrostatic repulsion of similarly charged ionizable groups and the osmotic pressure of counterions. On the contrary, the microgels deswell at intermediate pH values. This behavior is caused by the reduction of the electrostatic repulsion between similarly charged groups on the subchains, attraction between oppositely charged groups and decrease of osmotic pressure of counterions due to their release from the microgels. The minimum of the swelling degree is achieved at the isoelectric point when complete complexation of oppositely charged 1586

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Figure 6. Computer simulation snapshots of V−N+ and V−7N+ core−shell polyampholyte microgels at pH 8 under different regimes: (A, E) weak hydrophobic interactions between the proteins and microgels shell, εN cyt = 0.1ε, (B, F) medium hydrophobic interactions between the proteins and microgels shell, εN cyt = 0.3ε, and (C, G) strong hydrophobic interactions between the proteins and microgels shell, εN cyt = 0.5ε. Snapshots represent the slice of the microgels of 10σ width through the center of mass of the microgel/proteins complex.

size of the microgel shell leads to a decrease in the number of bound proteins (Table 8, Figure 8 B1, C1) due to the steric barrier of the shell and weakening of the electrostatic interaction on large distances. In the regimes of medium and strong hydrophobic interactions between the proteins and the microgel shell, εN cyt = 0.3ε and εN cyt = 0.5ε, the absolute amount of bound proteins in the microgels increases, but the localization of the proteins varies (Figure 8). At εN cyt = 0.3ε, significant amount of proteins are localized out of the core (Table 8) restraining in the shell, which is more pronounced for the microgels with the thicker shell (Figure 6F). If we take a higher value of the interaction parameter εN cyt = 0.5ε, almost all proteins remain in the shell and do not reach the core (Figure 6C,G). Based on the experimental data presented in Tables 6 and 7, one could conclude that in experiments we deal with the regime in-between weak and medium hydrophobic interactions. Indeed, according to Table 7, as discussed in the Materials and Methods, 20% binding for neutral microgels were achieved, revealing that the process is taking place in the regime εN cyt ∼ 0.2ε. Computer Simulations: Release of the Proteins. First, we consider the release of the proteins from microgels with random distribution of ionizable groups. Switching the pH value provokes the release procedure of the proteins (Figure 7). Similar to the experiment, if pH is decreased to 6 (isoelectric point) all 44 proteins are released from the microgel. At pH 3, we also observed the complete and accelerated release of cyt-c from microgels due to the strong repulsion. The presence of cations changes the electric fields inside the microgel. The resulting force acting on the proteins at any point within the microgel is now leading them to the outside. Thus, we confirm that the presence of homogeneously distributed cations arising from pH switching is the main reason for the release of cyt-c.

Figure 7. Relative number of cytochrome c in the random V± microgels at different pH (simulation).

The explanation of the experimental release curves of the proteins from core−shell distributed microgels is more intricate and intriguing. For core−shell distributed microgels the charges are separated from each other. Moreover, the degree of hydrophobic interactions between the proteins and microgels shell plays a crucial role in the release process. The results of simulations for V−N+, V−3N+, and V−7N+ describing the uptake/release with the corresponding parameter in case of weak hydrophobic interactions, εN cyt = 0.1ε, are presented in Figure 8 A1, B1, and C1, respectively. As mentioned above, after uptake at εN cyt = 0.1ε, the proteins are localized in the core or in its immediate vicinity. Switching the pH from 8 to 6 is accompanied by a decrease in the core charge and the appearance of the opposite sign charge in the shell. One could see that for the microgel with a thin shell, V−N+, more than 80% of the proteins are released due to the weakening of electrostatic attraction between the core and the cyt-c. The rest proteins are situated in the core. The same 1587

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Biomacromolecules

Figure 8. Relative number of cytochrome c in the core−shell V−N+ (A), V−3N+ (B), V−7N+ (C) microgels at different pH (simulation). Numbers 1, 2, and 3−corresponds to the different value of interaction parameter between the shell and cytochrome c particles, εN cyt, 0.1ε (slight repulsion), 0.3ε (slight attraction), and 0.5ε (strong attraction), respectively.

Figure 9. (top) Snapshot of V−3N+ microgel with entrapped protein at pH 3 (neutral core and similarly charged shell). The case of weak hydrophobic interactions between the proteins and microgels shell, εN cyt = 0.1ε. The zoom demonstrates “levitation” of the protein molecules inside the core of the microgel. (bottom) Equipotential (thin closed) lines of homogeneously charged spherical shell (R1 and R2 being the inner and outer radii of the shell) (A) and of discrete point-like charges (red dots) evenly distributed on the circle (cross-section of the sphere) of the radius R2 (B). The value of the potential is encoded in the color: the brighter the color, the higher the potential. Small black arrows depict the direction and intensity of the electric field. The inserted parts correspond to the potential of the electric field as a function of radial coordinate r. In the case of the discrete charge, U(r) is plotted in the direction crossing the circle in-between of two neighbor charges.

phenomenon as “levitation”. Picturing the proteins float and retain in the center of the microgel without any physical reasons (at first sight), like the zoomed in part presented in Figure 9. Despite a confined geometry, proteins aggregate neither with each other nor with the subchains of the microgel. At εN cyt = 0.1ε, the proteins do not contain attractive (hydrophobic) domains and long-range Coulomb repulsion forces act between them. A natural question arises: what is the reason for the localization of charged proteins inside the

trend can be observed for the microgels with the thicker shell. However, the amount of trapped proteins increases which cannot be explained only by the steric restriction of the shell. At pH 3, when the core is neutral and the shell is similarly charged to cyt-c, microgel with a thin shell, V−N+, does not contain any proteins due to the repulsion. Surprisingly, For V−3N+ and V−7N+ at pH 3, 21% and 32% (relative to the number of absorbed proteins at pH 8) of the loaded proteins are captured inside the microgels (Figure 9). We consider this 1588

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Article

Biomacromolecules microgel with penetrable but similarly charged shell? At first glance, if we imagine that the molecules are confined by a spherical, homogeneously charged shell, we can argue that the shell is not able to keep them in the cavity. Indeed, in this case, the electric field exists only within and outside the shell. In the cavity, the field is equal to zero due to the spherical symmetry of the object and absence of the charge. A 2D picture of equipotential lines and electric field (black arrows) is shown in Figure 9A. The inserted part in Figure 9A corresponds to the electrostatic potential of the shell. It is constant in the cavity and gradually decays to zero at an infinite distance from the shell. Thus, if positive charges are added into the cavity, they will move to infinity decreasing the electrostatic energy (they move in the direction of lower electrostatic potential). However, in reality, the charged groups in the shell of the core−shell microgels have a discrete distribution and cannot be approximated by a homogeneously “smeared out” charge, since the distance between the groups is of the same order of magnitude as the size of the guest molecules rather than smaller. By plotting an electric field profile of point-like charges evenly distributed on a circle (cross section of the sphere), the field inside the cavity is not equal to zero (Figure 9B). It possesses a maximum value near the charges and decays away from them. If we plot the electrostatic potential along the radius crossing the circle equidistantly from the neighbor charges (the direction of the minimum possible potential), it has a local minimum at r = 0 and absolute minimum at r = ∞. These two minima are separated by a potential barrier whose height is controlled by the density of the point-like charges: the larger the density, the higher the barrier. Therefore, if a charged molecule is placed in the center of the sphere, it can reach the equilibrium state (at r = ∞) only in the case of overcoming the potential barrier. In other words, the energy of the thermal motion of the charged molecule has to be larger than the height of the barrier. Otherwise, the charge will be trapped in the metastable state within the sphere for an infinitely long time and the probability of its release will be exponentially small. As explained in the section “Characterization of microgels” in the case of amphoteric core−shell microgels with the same fraction of charged groups in the core and the shell, the thicker the shell the greater the absolute number of charged groups inside it for a given pH. In our case at pH 3 total charges of the shells are Qshell (V−N+) = +350, Qshell (V−3N+) = +1050 and Qshell (V−7N+) = +2450. Therefore, the physical reason for the trapping of cyt-c in the V−3N+ and V−7N+ core−shell microgels is the presence of a high potential barrier, which is produced by similarly charged groups of the microgel. The total shell charge for V−N+ is not enough to trap even a single protein. It is worth noting that the size and shape of loaded molecules affect the possibility of overcoming electrostatic barrier. Thus, for example, in simulations complete release of linear polyelectrolytes of approximately the same molecular weight and charge density as modeled protein under the same conditions occurs.45 It is likely that the stretching and unfolding of the polyelectrolytes give rise to their charge redistribution contributing to an increasing probability of chains penetration through the charged porous shell. Interestingly, a disturbance of spherical symmetry in the system may cause the release of proteins. Video S2 illustrates an adsorption of the V−7N+/cyt-c complex at solid substrate. Strong deformation of the microgel distorts the electrostatic

interaction, provoking a complete release of the proteins trapped in the metastable state. It should be noted that microgel/cyt-c complex must be subjected to long-term deformation; otherwise, microgel will have time to adjust its structure to avoid release (Video S1). In the case of medium hydrophobic interactions between the proteins and microgels shell, εN cyt = 0.3ε a majority of proteins are localized in the shell, especially in the microgels with the thicker shell (Figure 6). Switching the pH from 8 to 3, a vast release of the proteins from the microgel occurs (Figure 8 A2, B2 and C2). Similar to the polyampholyte microgels with randomly distributed ionizable groups. At εN cyt = 0.5ε, the short-range attraction between the shell and proteins is so strong, that changing the pH from 8 to 3 pH will not induce the release of proteins (Figure 8 A3, B3, and C3). The theoretical predictions are in good agreement with the experimental data. Considering the region between weak and medium hydrophobic interactions, the proteins that are formerly bound to the shell during uptake were released according to electrostatic repulsion. The left proteins are trapped with high probability in the metastable state in the core region.

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CONCLUSIONS In summary, we have demonstrated that polyampholyte microgels are efficient colloidal carriers for entrapment and release of proteins. The most striking finding of our work is that the distribution of ionizable groups in polyampholyte microgels (random and core−shell) controls the interactions with the captured proteins from entrapment and “levitation” to accelerated release. In this work, the pH-triggered release of cytochrome c (cyt-c) as a model protein from polyampholyte microgels with a different distribution of ionizable groups (random and core−shell) was studied experimentally and by computer simulations. Polyampholyte microgels with defined chemical structure (controlled amounts of acidic and basic groups) and morphology (controlled distribution of acidic and basic groups: random and core−shell) were synthesized and their properties in aqueous solutions were systematically investigated. Polyampholyte microgels exhibit pH-dependent swelling and can switch their charge from positive (low pH) to negative (high pH). Microgels were loaded with cyt-c and protein release was investigated at pH = 3, 6, and 8. For microgels with a random distribution of ionizable groups complete release of cyt-c was observed at pH 6 and pH 3. For core−shell distributed polyampholyte microgels the ionizable groups are located in predefined regions (negative charges in the core and positive charges in the shell). In this case, proteins are bound both in the core and shell due to different forces. This also enables the entrapment of cyt-c inside the core through the formation of a positively charged shell, which acts as a potential barrier. On the contrary, part of the proteins bound inside the shell could be released according to electrostatic repulsion induced by pH switching. All the experimental results are supported and explained by computer simulations. These findings demonstrate that controlling amount and distribution of ionizable groups in polyampholyte microgels allows the design of functional colloidal carriers with programmed release kinetics for applications in drug delivery, catalysis and biomaterials. 1589

DOI: 10.1021/acs.biomac.8b01775 Biomacromolecules 2019, 20, 1578−1591

Biomacromolecules

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ASSOCIATED CONTENT

ACKNOWLEDGMENTS The authors gratefully acknowledge the computing time granted by the John von Neumann Institute for Computing (NIC) and provided on the supercomputer JURECA32 at the Jülich Supercomputing Centre (JSC).

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.biomac.8b01775.

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Article

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RG/RH ratio as a function of pH for microgels with different distribution of ionizable groups, FTIR measurements to proof the incorporation of the monomers and the theoretical amount, time depending release UV−vis measurements, characteristic peaks of cytochrome c detected with UV−vis spectroscopy at different pH, proof of the electrostatic interactions between the microgels and cytochrome c, schematic illustration of protein absorption, TEM images of random distributed and core−shell distributed polyampholyte microgels, proof of the positive charges of cyt-c over a broad pH range, pH depending size measurements of polyampholyte microgels, release kinetic (relative released cyt-c amount in %) of cyt-c from polyampholyte microgels, release kinetic of cyt-c (relative released cyt-c amount in %) and comparison from opposite charged core−shell distributed polyampholyte microgels at pH 3, illustration of modeled protein structure, simulation curves of the absolute number of cytochrome c in the random, 1:1, 1:3, and 1:7 core−shell distributed microgels vs time during uptake and release at different value of interaction parameter between the shell and cytochrome c particles, simulation curves of the gyration radius of random, 1:1, 1:3, and 1:7 core−shell distributed microgels vs time during uptake and release at different value of interaction parameter between the shell and cytochrome c particles, binding ability data of different polyampholyte microgels (PDF) Video S1 illustrating short-term deformation of the microgel/cytochrome-c complexes (AVI) Video S2 illustrating long-term deformation of the microgel/cytochrome-c complexes (AVI)

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AUTHOR INFORMATION

Corresponding Authors

*A. Pich. E-mail: [email protected]. *I. I. Potemkin. E-mail: [email protected]. ORCID

Wenjing Xu: 0000-0003-4120-7609 Andrey A. Rudov: 0000-0001-7364-4286 Walter Richtering: 0000-0003-4592-8171 Igor I. Potemkin: 0000-0002-6687-7732 Andrij Pich: 0000-0003-1825-7798 Author Contributions ‡

W.X. and A.A.R. contributed equally.

Funding

The financial support of the Deutsche Forschungsgemeinschaft (DFG) within Collaborative Research Center SFB 985 “Functional Microgels and Microgel Systems”, the Russian Foundation for Basic Research, project # 19-03-00472, and the Government of the Russian Federation within Act 211, contract # 02.A03.21.0011, is gratefully acknowledged. A.P. thanks the VolkswagenStiftung for financial support. Notes

The authors declare no competing financial interest. 1590

DOI: 10.1021/acs.biomac.8b01775 Biomacromolecules 2019, 20, 1578−1591

Article

Biomacromolecules

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DOI: 10.1021/acs.biomac.8b01775 Biomacromolecules 2019, 20, 1578−1591