Article pubs.acs.org/Biomac
Hemocompatible Poly(NIPAm-MBA-AMPS) Colloidal Nanoparticles as Carriers of Anti-inflammatory Cell Penetrating Peptides Rush L Bartlett II, Matthew R. Medow, Alyssa Panitch,* and Brandon Seal Weldon School of Biomedical Engineering, Purdue University, 206 South Martin Jischke Drive, West Lafayette, Indiana, 47907 ABSTRACT: Anionic copolymer systems containing sulfated monomers have great potential for delivery of cationic therapeutics, but N-isopropylacrylamide (NIPAm) 2-acrylamido-2-methyl-1-propanesulfonic acid (AMPS) copolymer nanoparticles have seen limited characterization to date with regard to physical properties relevant to loading and release of therapeutics. Characterization of polymeric nanoparticles incorporating AMPS showed an increased size and decreased thermodynamic swelling ratios of AMPS containing particles as compared to NIPAm nanoparticles lacking AMPS. Particles with increasing AMPS addition showed an increased propensity for uniformity, intraparticle colloidal stability, and drug loading capacity. Peptide encapsulated in particles was shielded from peptide degradation in serum. Particles were shown not impede blood coagulation or to cause hemolysis. This study has demonstrated that AMPS incorporation into traditional NIPAm nanoparticles presents a tunable parameter for changing particle LCST, size, swelling ratio, ζ potential, and cationic peptide loading potential. This one-pot synthesis results in a thermosensitive anionic nanoparticle system that is a potentially useful platform to deliver cationic cell penetrating peptides.
■
fonic acid (AMPS).18 The few nanoparticle papers that have previously studied poly(NIPAm-MBA-AMPS) nanoparticles serve as a starting point for characterization of the physical properties, but lack assessment of this nanoparticle system for use in biomedical applications such as a drug delivery vehicle.19,20 As a result, additional characterization of poly(NIPAm-MBA-AMPS) nanoparticle formulations are warranted to further evaluate the unique properties of poly(NIPAm-MBA-AMPS) nanoparticles for future applications in drug delivery. Our lab’s interest in sulfate-containing pNIPAm nanoparticles is based on a long-term goal of delivering cellpenetrating peptides (CPPs).21,22 Prior work has demonstrated that the arginine residues within the primary sequence of our lab’s CPPs bind well to sulfated glycosaminoglycans through electrostatic interactions.23 However, there is currently no wellstudied synthetic drug delivery nanoparticle that contains high concentrations of sulfate. In this study of sulfate containing thermosensitive poly(NIPAm-MBA-AMPS) nanoparticles we not only exploit the nanoparticle swelling at low temperatures to facilitate peptide loading, but we characterize the hemocompatible nanoparticles according to varying AMPS concentration that directly tunes particle size, LCST, and charge-dependent drug loading of CPPs.
INTRODUCTION In an effort to provide targeted, localized methods for drug delivery and biomedical imaging, many research groups have developed polymeric nanoparticle systems. Several polymer types have been used including poly(ethylene glycol), poly(methylmethacrylate), chitosan, methacrylated hyaluronic acid, and poly(lactic-co-glycolic acid), to name a few. These systems have been previously reviewed by Oh et al.1 Most of these polymer systems depend on encapsulation techniques to load drugs that are sometimes too harsh for more delicate biological therapies.2 In these instances, diffusion loading under mild conditions is preferred. One extensively studied copolymer system involves a thermosensitive poly(N-isopropylacrylamide) (pNIPAm) backbone which reversibly shrinks above the 31−33 °C lower critical solution temperature (LCST).3−5 Due to this reversible shrinking, efficient diffusion loading of NIPAm nanoparticles is possible. Studies of various formulations of poly(NIPAm) nanoparticles have yielded advances for biomedical applications such as core shell systems,6,7 ion recognizable particles,8 tinted gels,9,10 and targeted drug delivery.6,11,12 Poly(NIPAm) nanoparticles investigated for biomedical applications are traditionally copolymerized with carboxylic acid comonomers like itaconic acid,13 or most commonly, acrylic acid (AAc).14,15 Often, these carboxylic comonomers serve not only as a colloidal stabilizer, but also as a means to facilitate further chemical reactions to incorporate molecules for improved targeting11 or imaging.16,17 Although many monomers with carboxylic acid functionalities have been copolymerized with N-isopropylacrylamide, relatively little characterization has been performed with sulfated monomers like 2-acrylamido-2-methyl-1-propanesul© 2012 American Chemical Society
■
EXPERIMENTAL SECTION
Materials and Methods. Chemicals. N-Isopropylacrylamide (NIPAm) was purchased from Polysciences Inc. (Warrington, Received: January 31, 2012 Revised: March 5, 2012 Published: March 27, 2012 1204
dx.doi.org/10.1021/bm300173x | Biomacromolecules 2012, 13, 1204−1211
Biomacromolecules
Article
Table 1. Molar Reactant Percentages Reaction % AMPS NIPAm AMPS
0% 838.7 mg 0 mg
2.5% 816.4 mg 38.3 mg
5% 794.1 mg 76.5 mg
7.5% 773.2 mg 114.8 mg
10% 752.3 mg 153.1 mg
15% 710.4 mg 190.9 mg
Hemolysis. Hemolysis assays were performed following ASTM F756 with a BioSpec-1601 spectrophotometer (Shimadzu, Columbia, MD, USA).26 Determination of total blood hemoglobin was accomplished by using Drabkin’s reagent and measuring the absorbance of free hemoglobin in solution at 540 nm. The total blood hemoglobin concentration was used to adjust the hemoglobin content of the blood sample to 10 ± 1 mg by adding blood and an equivalent amount of 1× PBS solution. Percent hemolysis was normalized by dividing total hemolysis determined by a 0.01% v/v of Triton X-100, subtracting the absorbance of blood only, and setting absorbance of PBS to zero. Blood, 1 mL, was incubated with 100 μL of a 1.5 mg/mL solution of the nanoparticles in PBS at 37 °C for 3 h under gentle shaking. After 3 h, samples were centrifuged at 750 g for 15 min. Supernatant was removed, added 1:1 to equivalent volume of Drabkins reagent and incubated for 15 min before measurement. Absorbance measurements at 540 nm were then recorded using a spectrophotometer. Peptide Synthesis and Purification. Therapeutic peptides for use in drug release studies were synthesized using standard 9-fluorenylmethyloxycarbonyl (FMOC) chemistry on Knorr-amine resin (Synbiosci Corp, Livermore, CA, U.S.A.). Two amino acid coupling steps were used to attach amino acids (Synbiosci Corp, Livermore, CA, U.S.A.). For the first coupling step, N-hydroxybenzotriazole (HoBt) and N,N′diisopropylcarbodiimide (DIC) were incubated with amino acid and resin for 30 min. A second 30 min coupling used 2-(1H-benzotriazole1-yl)-1,1,3,3-tetramethyluronium hexafluorophosphate (HBTU), lutidine, and amino acid to ensure high yields of pure product. After synthesis, peptides were cleaved with a cocktail of trifluoroacetic acid (Sigma-Aldrich, St. Louis, MO, U.S.A.), triisopropyl silane (TCI America, Boston, MA, U.S.A.), ethane dithiol (Alfa Asara, Ward Hill, Massachusetts, U.S.A.), and Milli-Q water. Peptide was immediately precipitated in ether, recovered by centrifugation, solubilized in MilliQ water, and lyophilized. Peptides were purified on a FPLC AKTA Explorer (GE Healthcare, Pittsburgh, PA, U.S.A.) with a 22/250 C18 prep-scale column (Grace Davidson, Deerfield, Illinois, U.S.A.) and an acetonitrile gradient with 0.1% trifluoroacetic acid. Peptide molecular weight was confirmed by time matrix-assisted laser desorption ionization time-of-flight (MALDI TOF) mass spectrometry with a 4800 Plus MALDI TOF/TOF Analyzer (Applied Biosystems, Foster City, CA, U.S.A.). Drug Loading and Release. To ensure proper drug loading, purified peptide was first dissolved in Milli-Q water to create a 1 mg/ mL loading solution. Then this solution was added to lyophilized nanoparticles of 0, 5, and 10% AMPS such that the final particle concentration in solution was 2 mg/mL. Then, the drug−nanoparticle loading solution complex was allowed to incubate for 24 h at 4 °C, to ensure particle swelling and facilitate peptide uptake in the particles, prior to centrifugation at 55000 rpm in an Optima L-90k Ultracentrifuge (Beckman Coulter, Indianapolis, IN, U.S.A.). Then diluent was isolated from the pellet. Both nanoparticle pellet and diluent were lyophilized for later analysis. Loaded nanoparticles were suspended in either Milli-Q water or sterile PBS at pH 7.4 (Invitrogen, Grand Island, NY, U.S.A.) at a concentration of 0.5 mg/mL loaded particles. Measurement of free peptide released into the solution was conducted using fluorescence analysis in a Costar 96-well plate by adding 20 μL of a sample solution to 180 μL fluoraldehyde ophthalaldehyde (OPA) solution (Thermo Scientific, Waltham, MA, U.S.A.). Fluorescent measurements of drug release were taken every 15 min for 3 h, then every 6 h until day two, and every 12 h until day three. After three days, samples were taken every 24−48 h until day 13. Then a final point was taken at 21 days. To ensure that only free peptide, and not particles, was present in the measurement sample, each sample was run through a 100000 MW cutoff membrane microcentrifuge tube (Omega, Norcross, GA, U.S.A.) prior to
PA, U.S.A.). N,N′-Methylenebisacrylamide (MBA), sodium dodecyl sulfate (SDS; 10% w/v in water), 2-acrylamido-2methyl-1-propanesulfonic acid (AMPSA), and potassium persulfate were acquired from Sigma-Aldrich (St. Louis, MO, U.S.A.). NIPAm, MBA, and AMPSA were stored under nitrogen at 4 °C. All water used in synthesis, dialysis, and testing was treated by a Milli-Q system (Millipore, Billerica, MA, U.S.A.; 18.2 MΩ·cm resistivity). Nanogel Synthesis. NIPAm-containing nanogels were synthesized using standard precipitation polymerization.24,25 Briefly, the nanogel compositions described in Table 1 were formed by dissolving varying amounts of NIPAm and AMPSA in 30 mL of degassed Milli-Q water in a three neck round-bottom flask. Then, 28.5 mg of MBA was dissolved in 10 mL of Milli-Q water and 164 μL of a 10% SDS in Milli-Q water solution were added, and the mixture was heated to 80 °C under nitrogen. After 30 min, 33.7 mg potassium persulfate, predissolved in 10 mL of degassed water, was added to the mixture to initiate polymerization. After 4 h, the reaction was cooled to room temperature. Then, the mixture was dialyzed against Milli-Q water for 7 days using a 15000 MWCO membrane. Post-dialysis concentrations varied between 6 and 15 mg/mL and were diluted or concentrated as necessary through lyophilzation and resuspension. To lyophilize nanoparticles, solutions were frozen to −80 °C for 12 h and then placed into a lyophilizer until the liquid was completely removed. Nanogel Characterization. Nanoparticles from three different reactions were characterized to validate repeatability of the synthesis for size and zeta (ζ) potential measurements. The hydrodynamic diameter of nanoparticles generated under each reaction condition was measured with dynamic light scattering (DLS) using a Nano-ZS90 Zetasizer (Malvern, Westborough, MA, U.S.A.) that was calibrated with polystyrene beads. Samples in disposable polystyrene cuvettes underwent 12 measurements per sample. For static temperature measurements, samples were equilibrated at the desired temperature for 5 min. Temperature sweep samples were equilibrated for 2 min for each half-degree temperature change. Sample zeta (ζ) potentials were measured by a Nano-ZS90 Zetasizer in folded capillary cells after DLS in Milli-Q water. Titration was used to determine the amount of AMPS incorporated by slowly adding 0.1 M NaOH. The amount of acid present was subtracted from residual potassium persulfate in 0% AMPS particles and then normalized to % reaction yield to give a final molar % AMPS in particles. This work will reference particles according to their feed amount of AMPS. TEM was conducted at the Purdue University Life Science Microscope facility on a FEI/Philips CM-100 Transmission Electron Microscope at 100 Kv using an uranyl acetate stain (UA) at pH 4.5. Discharged TEM sample grids were placed onto the top of a droplet of sample for 2 min. Then UA stain was added and samples were dried briefly before imaging at room temperature. Activated Partial Thromboplastin Time. Activated partial thromboplastin time (aPTT) was measured using a Hemochron Response whole blood coagulation system. A total of 2 mL of citrated bovine whole blood and 200 μL of nanoparticles in phosphate buffered saline (PBS) pH 7.4 (1.5 mg/mL final concentration in blood) were added to Hemochron tubes containing colloidal kaolin activating agent and 0.02% thimerosal preservative agent. PBS was calcium- and magnesium-free. The tubes were then measured and aPTT was recorded. The aPTT for each sample was measured in triplicate. Controls consisted of PBS without nanoparticles, a 6 mg/mL purified bovine collagen solution (Advanced Biomatrix, San Diego, CA, U.S.A.), and 0.01 mg/mL (final concentration) heparin (Sigma Aldrich, St. Louis, MO, U.S.A.). Measurements with heparin were stopped after 1000 s. 1205
dx.doi.org/10.1021/bm300173x | Biomacromolecules 2012, 13, 1204−1211
Biomacromolecules
Article
fluorescent analysis. Testing also revealed no free peptide stuck to this size exclusion membrane and that the membrane completely separated out the nanoparticles from the solution (data not shown). Images of particles containing fluorescein isothiocyanate (FITC) labeled peptide were taken with an Olympus FV1000 confocal microscope. Serum Studies. Samples tested include: peptide alone, serum alone, peptide in serum, and KAFAK-loaded 5% AMPS nanoparticles in serum. Each sample volume also contained 0.5× PBS except when stated. Samples containing peptide initially contained 0.5 mg of peptide alone or 1.12 mg of loaded 5% AMPS particles with an equivalent amount of 0.5 mg peptide. Samples with serum were 0.5× serum and 0.5× PBS. Samples without serum were 0.5× PBS and 0.5× Milli-Q water. Peptide alone, serum alone, peptide in serum, loaded nanoparticles alone, and loaded nanoparticles in serum were incubated for 12 h at 37 °C on a shaker. After the 12 h incubation, samples not containing nanoparticles were placed in the fridge at 4 °C for 24 h before evaluation. Samples containing nanoparticles were centrifuged for 2 h at room temperature at 30000 rpm in an Optima L-90k Ultracentrifuge (Beckman Coulter, Indianapolis, IN, U.S.A.). The nanoparticle pellet was briefly washed with Milli-Q water and then resuspended in a solution of 0.5× PBS and 0.5× Milli-Q water. Samples were then placed in the fridge for 24 h. Before the 24 h nanoparticle sample was placed in the fridge to incubate, but after the pellet was dissolved, an additional amount of 0.257 molar concentration of NaCl was added to induce peptide release. This brought the final NaCl concentration to 0.342 molar or twice normal salt concentration in PBS. Additional NaCl was added to facilitate total release by introducing additional counterions. Samples were evaluated by reverse phase chromotography on a FPLC (AKTA Explorer, GE Healthcare, Pittsburgh, PA, U.S.A.) with a 22/250 C18 prep-scale column (Grace Davidson, Deerfield, IL, U.S.A.) and an acetonitrile gradient with 0.1% trifluoroacetic acid. Peptide was quantified by integrating the area under the peak of the FPLC. Peptide molecular weight was confirmed by time-of-flight MALDI mass spectrometry with a 4800 Plus MALDI TOF/TOF Analyzer (Applied Biosystems, Foster City, CA, U.S.A.).
knowledge gained from studies pioneered by Garcia-Salinas et al., we varied AMPS addition to determine if this may be a useful parameter to control charge dependent release of therapeutic peptides from these nanoparticles. We observed a trend of increasing hydrodynamic diameter with increasing starting monomer percent of AMPS for measurements at 37 °C. At 23 °C, particle size did not significantly vary with AMPS. To determine the final composition of nanoparticles, titration was used to determine AMPS content normalized with respect to reaction yield and potassium persulfate incorporation in particles not containing AMPS. On average it was found that only 81 ± 3.1% of AMPS fed into the reaction was incorporated into the nanoparticle complex after purification. Table 2 indicates that there is no upper limit of AMPS incorporation up to 10% AMPS feed ratio evaluated, but we Table 2. Percent AMPS Monomer Incorporation by Titration Method mole % AMPS in rxn feed 0 2.5 5 7.5 10
normalized avg AMPS incorporated in nanoparticles (%) NA 1.95 3.98 6.23 8.48
± ± ± ±
0.10 0.21 0.33 0.90
observed a lack of particle formation at 15% according to TEM and light scattering. Accordingly, no reliable measurements of particles formed with 15% initial AMPS monomer were recorded. However, a small amount of lyophilizate was observed from a 15% formulation. This lyophilizate was likely long chain copolymer rather than nanoparticles. Particles did not degrade or aggregate when stored at 4 °C for over 18 months in Milli-Q water. The ability of poly(NIPAm-MBA-AMPS) nanoparticles to respond to temperature was maintained with the incorporation of AMPS. However, as indicated by the relative differences in particle size in Figure 1, the thermodynamic swelling ratio was shown to decrease in magnitude with increasing AMPS incorporation, presumably due to an increase in overall hydrophilicity of the NIPAm polymer. This is consistent with research demonstrating that, as hydrophilic content is added to a hydrophobic polymer backbone, the effect is an increase in overall system solubility.15,29,30 As seen in studies using NIPAm as the primary constituent of a bulk polymer, the phase transition temperature, or lower critical solution temperature (LCST), shifts to higher temperatures as the AMPS content of the polymer increases (Figure 2).29,31 The change in hydrodynamic diameter over a temperature sweep of 0.5 °C per 5 min from 25 to 40 °C is shown in Figure 2. Nanoparticles synthesized from MBA-NIPAm were used as a control and were shown to have a LCST consistent with literature around 31−33 °C.7 Increasing the addition of AMPS increased the LCST and the solvation energy of the copolymer leading to a reduction in swelling ratio. Specifically additions of AMPS above 5% led to swelling ratios of 1.25 ± 0.04 as compared to a swelling ratio of 2.3 ± 0.25 for 0% AMPS particles. Zeta Potential Measurements. AMPS incorporation and colloidal stability were indirectly measured by zeta (ζ) potential in Milli-Q water. Figure 3, as a general trend, shows that zeta potential increased in magnitude with the addition of up to 10%
■
RESULTS Effect of AMPS Addition on Particle Size. The addition of AMPS has a profound impact on the size and thermodynamic swelling of typical NIPAm systems. This allows AMPS to be used to tune the physical properties of the system. Dynamic light scattering (DLS) size measurements of nanoparticles as a function of increasing monomer reaction ratio of AMPS to NIPAm are shown in Figure 1. This data demonstrates a general trend of increased size with increased incorporation of AMP as both 23 and 37 °C. Consistent with studies by Garcia-Salinas et al., we observed stable particle formation with the addition of 5% molar AMPS into the poly(NIPAm-MBA) reaction.19,27,28 To expand the
Figure 1. Dynamic light scattering (DLS) hydrodynamic diameter measurements of poly(NIPAm-MBA-AMPS) nanoparticles at 23 and 37 °C; n = number of separate reactions measured. 1206
dx.doi.org/10.1021/bm300173x | Biomacromolecules 2012, 13, 1204−1211
Biomacromolecules
Article
an increase in the standard deviation of nanoparticle size with increasing AMPS. Lastly, due to a lack of particle formation above 10% it is likely that as hydrophilic content and charge density increases the reaction has a greater propensity for polymer chain formation instead of creation of nanoparticles. Together, these four inferences suggest the presence of an upper limit of AMPS monomer incorporation between 7.5 and 10% under the reaction conditions studied and that the composition of nanoparticles formed with 7.5 and 10% AMPS in the monomer feed ratio are substantially similar. Increasing AMPS further, above 10%, inhibited particle formation. Transmission Electron Microscopy (TEM). TEM images were taken at the Purdue Life Science Microscopy Facility. Figure 4 shows TEM images of nanoparticles from reactions of increasing concentrations of AMPS (from 0 to 10%). TEM images show that particles containing AMPS are more uniform than those without AMPS. Also, zeta potential measurements, an indication of colloidal stability, size uniformity, and uniformity of spacing, were confirmed as particles with AMPS were more uniform and evenly spaced than particles without AMPS in Figure 4A. The decreased particle diameter found using TEM as compared to DLS is due to DLS measurements being conducted in Milli-Q water as compared to TEM images taken with particles stained by UA on a dried TEM plate. Also, particles incorporating 2.5 and 5% AMPS seem to have a slight ring of lighter material that may indicate that AMPS is more concentrated at the surface at these lower reaction percentages, an idea consistent with hydrophobic groups of the NIPAm monomer preferentially segregating to the inner core of the polymer. However, it is also possible that this ring is an artifact of the negative staining procedure with UA as it was not present for all images or even on all of the particles in each image.
Figure 2. DLS temperature sweep from 23 to 40 °C of poly(NIPAmMBA-AMPS) nanoparticles.
Figure 3. Zeta potential (ζ) measurements of poly(NIPAm-MBAAMPS) nanoparticles; n = number of separate reactions measured.
AMPS. However there is no statistical difference between the 7.5% and 10% zeta measurements. Similarly, the sizing data from Figure 1 shows that there is no statistically relevant size difference between 7.5 and 10% particles. Also, Figure 1 shows
Figure 4. TEM images of poly(NIPAm-MBA-AMPS) nanoparticles from 0% AMPS to 10% AMPS: 0%, A; 2.5%, B; 5%, C; 7.5%, D; 10%, E. Images are shown at 52k magnification with a scale bar of 200 nm. 1207
dx.doi.org/10.1021/bm300173x | Biomacromolecules 2012, 13, 1204−1211
Biomacromolecules
Article
Blood Interaction Assays. As part of a preliminary assessment of utilizing poly(NIPAm-MBA-AMPS) as a blood contacting drug delivery vehicle, whole blood studies were done to determine the hemocompatibility of this system. Table 3
into the nanoparticle cores while they are expanded below the LCST, while above the LCST at physiological conditions they remain protected from proteases. Nanoparticle loading efficiencies, as a percent of final particle weight composed of KAFAK are shown in Table 4.
Table 3. Blood−Nanoparticle Interaction
blood PBS 0.0% 2.5% 5.0% 7.5% 10.0% collagen 6 mg/mL heparinI Triton X-100II
Table 4. Drug Loading Efficiency as Percent Drug Mass Per Loaded Particle
whole blood coagulation time (s)
normalized whole blood % hemolysis
avg (n = 3)
std dev
avg (n = 3)
std dev
236 235 238 234 237 227 227 26 >1000
14.3 17.3 15.7 11.8 14.9 7.3 7.2 7.7 NA
particle composition
mass % drug in each loaded particle after 24 h of loading
std dev (%)
0 −0.07 0.44 1.51 0.00 0.69
0.11 0.10 0.75 1.40 0.07 0.20
0% AMPS 5% AMPS 10% AMPS
17.8 45.3 60.5
7.1 9.5 6.2
100
4.29
Table 4 indicates that the addition of only 5% AMPS yields a roughly a 3× increase in the amount of drug loaded into the particles utilizing a passive diffusion loading method in Milli-Q water over a 24- hour incubation at 4 °C. Further addition of AMPS to 10% results in roughly a 4× increase over the amount of drug loaded without AMPS. To test the electrostatic mechanism of drug binding, release studies of KAFAK were conducted in both PBS and Milli-Q water to determine how particles would release bound KAFAK peptide. As mentioned in the introduction KAFAK has been previously shown to bind electrostatically to sulfated polymers. Figure 5 demonstrates that electrostatic interactions do indeed
I
Heparin concentration was 0.001 mg/mL; P > .05 as compared to Heparin. IITriton X-100 was 0.01% by volume; P > .05 as compared to Triton X-100.
shows coagulation times of various treatments of AMPS nanoparticles, 100 μL/ml blood to a final concentration of 1.5 mg/mL. No difference in coagulation time between treatment and control (PBS and collagen) were observed, P > .05, showing that the nanoparticles did not affect coagulation. A negative control of a 6 mg/mL bovine collagen solution was shown to induce clotting. As a positive control, heparin was seen to inhibit coagulation. After 1000 s, experiments in the presence of heparin were stopped due to lack of clotting. Then, hemolytic assay ASTM standard protocol F756 was used to check if hemolysis occurred upon the addition of poly(NIPAm-MBAAMPS) to the blood. Table 3 shows no statistically relevant hemolysis (P > .05) to be present for blood incubated with the 1.5 mg/mL concentration of the nanoparticles at 37 °C for 3 h. The data in Table 3 further indicates that poly(NIPAm-BISAMPS) nanoparticles are hemocompatible. This suggests that nanoparticles may be useful for a blood contacting in vivo drug release. Drug Loading and Release. Our lab has demonstrated previously that positively charged MAPKAP Kinase 2 (MK2) inhibiting peptide KAFAKLAARLYRKALARQLGVAA (abbreviated KAFAK), binds strongly to sulfated glycosaminoglycans through electrostatic interactions.23 KAFAK, a strong inhibitor of MK2, shows potential as a therapy for chronic inflammation conditions associated with MK2 mediated increases in cytokine activity such as rheumatoid arthritis.21,22,32 However, because of the highly nonspecific nature of the CPP associated with KAFAK and the susceptibility to enzymatic degradation, delivery of these serum sensitive therapeutic peptides is difficult in vivo without a site specific injection into an arthritic joint.32,33 As a result, a controlled release vehicle with a high loading capacity is needed to maintain the therapeutic activity of KAFAK over long time scales without repeated injections. Due to the strong affinity of KAFAK for sulfated moieties, AMPS doped poly(NIPAm) nanoparticles provide a tunable loading capacity based on the amount of incorporated AMPS. NIPAm was used as the polymer backbone to ensure the high molecular weight drug KAFAK was able to efficiently diffuse
Figure 5. Drug release profiles of KAFAK from 0, 5, and 10% AMPS containing particles in Milli-Q water and PBS over a 21 day period at 37 °C.
have a profound effect on the drug release kinetics because particles with AMPS release much less KAFAK. Also, in every case the amount of KAFAK released in Milli-Q water was significantly less than the amount of KAFAK released in a PBS environment. After 1 week 0% AMPS particles released near 100% of their drug in PBS, therefore it is reasonable to assume that the KAFAK peptide is not prohibitively encumbered by the shrunken polymer network over long time scales. However, the movement of KAFAK is retarded in the shrunken network because the amount of KAFAK loaded overnight at 4 °C requires 1 week at 37 °C to fully release. Taken together these observations demonstrate that the loading and release mechanism is driven by both physical hindrance and electrostatic interactions between KAFAK and the copolymer. After 1208
dx.doi.org/10.1021/bm300173x | Biomacromolecules 2012, 13, 1204−1211
Biomacromolecules
Article
Figure 6. Confocal image of FITC labled KAFAK inside 5% AMPS poly(NIPAm-MBA-AMPS) nanoparticle. (A) Particle at time zero. (B) Particle after releasing for 21 days at 37 °C in PBS. (C) Particle after incubation in serum at 37 °C for 12 h and incubation in 2× NaCl PBS for 24 h at 4 °C. (D) Overlay of fluorescence and light image at time zero. (E) Light image at time zero. (F) Zoomed out image after releasing for 21 days at 37 °C in PBS.
serum to determine if the peptide could be protected from degradation while within the particles in body fluids. Information from Table 5 indicates that KAFAK peptide
release KAFAK integrity was confirmed by MALDI TOF mass spectroscopy. Also, although actual mass of drug released were comparable between the 10% AMPS particles and the 0% AMPS particles in the PBS environment, the actual percent of initially loaded KAFAK released from the 10% was only 25% compared to nearly 100% by the 0% AMPS particles. 10% particles also released ∼3−7% more KAFAK than 5% particles. This is most likely due to the reduced shrinking ability of the 10% particles at 37 °C (19.4%) as compared to the 5% particles (29.3%). Enhanced particle contraction at 37 °C causes a reduction in void space between polymer strands that, combined with a high negative charge, further hindered KAFAK escape from the particle cores, as shown in Figure 6. Figure 6 indicates that KAFAK has released from the layer close to the surface of the poly(NIPAm-MBA-AMPS) nanoparticle after 21 days in a PBS solution but that KAFAK remained entrapped within the particle cores. Although data shown in Figure 5 demonstrates that a significant amount of KAFAK is released from the nanoparticles containing AMPS there is still a significant amount of KAFAK locked within the particles likely due to the combined effect of strong electrostatic interactions and a tightly intertwined polymer mesh while the poly(NIPAm-MBA-AMPS) is above the LCST. To better determine the ability of the poly(NIPAM-MBAAMPS) nanoparticles to deliver KAFAK in the physiological environment we conducted an enzymatic degradability assay in
Table 5. KAFAK Degradability Assay in Serum sample (A) serum control 12 h at 37 °C and 24 h at 4 °C (B) KAFAK peptide control 12 h at 37 °C and 24 h at 4 °C (C) serum KAFAK control 12 h at 37 °C and 24 h at 4 °C (D) serum and loaded nanoparticles 12 h at 37 °C, then purify and release 24 h at 4 °C in 2× salt (E) loaded nanoparticles control 12 h at 37 °C, then purify and release 24 h at 4 °C in 2× salt
integral of KAFAK peak between 30 and 31% acetonitrile
% recovery
0 mAU·mL
0
593.3 mAU·mL
100
0 mAU·mL
0
93.8 mAU·mL
15.8
238.1 mAU·mL
40.1
degrades within 12 h without a carrier to prevent degradation. Poly(NIPAM-MBA-AMPS) nanoparticles help protect KAFAK from degradation in serum by hindering the mobility of large proteases into the core of the drug-nanoparticle complex. This results in a large amount of KAFAK continuing to be present in the nanoparticle even after 12 h of incubation at 37 °C in serum. 1209
dx.doi.org/10.1021/bm300173x | Biomacromolecules 2012, 13, 1204−1211
Biomacromolecules
Article
of the bulk hydrogel as a mimic of the extracellular matrix.36 A second strategy employed by Lyon et al. and others utilizes a pH-sensitive hydrolytically degradable cross-linking system for use in the one-pot synthesis of poly(NIPAm) nanoparticles.37 Both strategies present unique opportunities for future studies with AMPS to facilitate a targeted release of therapeutic peptides from an AMPS containing nanoparticle system. Incorporating one or both into future research with AMPS containing nanoparticles is currently underway by our laboratory as the ability of a nanoparticle system to strongly sequester peptide and then release in a more specific manner could greatly suppress collateral damage due to nonspecific systemic release.
Figure 5 and Table 5 both demonstrate the difficulty of separating KAFAK from the AMPS containing nanoparticles after loading has occurred. Confocal images in Figure 6c indicate that even after incubation with serum for 12 h and further incubation at low temperature with 2× salt concentration some KAFAK still remains bound within the core of the nanoparticle. The mechanisms for the decrease in fluorescence observed in Figure 6c are likely manifold, including a some serum degradation of released material, peptide competitive binding/release with serum proteins, ionic destabilization of peptide binding in the presence of NaCl, and the swollen state of the nanoparticles at 4 °C for 12 h, which allowed for the release of fluorescent KAFAK. Similarly, releasing KAFAK from nanoparticles without serum yielded only a 40.13% recovery after 24 h at 4 °C in 2× salt. The difference in the control recovery and the 15.8% recovery from the sample containing serum and nanoparticles is likely due to either KAFAK degradation taking place at the surface of the particle or a competitive binding of serum proteins causing increased release of surface bound KAFAK during the 12 h incubation. Regardless of either outcome, a significant portion of KAFAK remains protected from serum degradation when loaded into poly(NIPAM-MBA-AMPS) nanoparticles. KAFAK integrity was confirmed by MALDI TOF MS.
■
CONCLUSIONS We have successfully synthesized a copolymer nanoparticle system comprised of poly(NIPAm-MBA-AMPS). The data demonstrates that approximately 7.5% AMPS in the synthesis feed gives maximal AMPS incorporation into the particles before system instability occurs. Particles containing AMPS are shown by TEM analysis to be uniform in size and to maintain better intraparticle spacing as compared with poly(NIPAm) alone. Biological characterization shows the particles to be hemocompatible. Most importantly, with only a short incubation below the particle LCST a very high loading capacity for cationic CPP KAFAK can be achieved. However, above the LCST the combination of electrostatic affinity and a tightly constricted polymer network makes it difficult to release all of the loaded KAFAK material. The range of reaction conditions studied show that addition of AMPS as a sulfated monomer can be used as a tool to tune, thermodynamic swelling, colloidal stability, drug loading, and release of peptide from the particles.
■
DISCUSSION Previously our laboratory demonstrated that controlled release of heparin-binding peptides from bulk hydrogels containing heparin has been previously demonstrated as a method to create a bulk scaffold for controlled release.34 Our current work demonstrates that synthetic nanoparticles containing sulfated moieties can mimic this heparin-mediated binding and release in a way that does not affect the hemoregulatory system and provides some protection from serum proteases. Loading of poly(NIPAm-MBA-AMPS) nanoparticles at 4 °C occurs in only a few hours, but at 37 °C, the nanoparticles can slowly release peptide for a week because the nanoparticles are in a collapsed state above the LCST. This system could be a useful delivery vehicle for intravenous injection or for direct injection into other body fluids, including intra-articular injection into the synovial fluid.35 However, due to the extremely high bonding affinity of KAFAK to nanoparticles containing AMPS, resulting in release of less than half of the loaded peptide, future experiments will need to investigate schemes that enable nanoparticles to degrade to liberate a higher percentage of KAFAK over time. Increasing electrostatic affinity by increasing the sulfate content in the nanoparticles also has a dual effect of reducing the particle’s ability to shrink above the LCST. This reduction in shrinking could enable increased release of uncharged molecules above the LCST, but the increase in electrostatic attraction between AMPS and cationic species will dampen the release of these molecules. Some optimization of the system could be employed to balance the reduction in swelling ratio with the added electrostatic attraction in order to optimize release, but future studies could also implement strategies for release by selective degradation of the particles. Several strategies could be employed to enable selective release including degradable cross-links. One strategy, demonstrated by West et al., has demonstrated the feasibility of incorporating collagenase sensitive peptides as cross-linkers into bulk hydrogels. These peptide cross-linkers are shown to degrade in the presence of collagenase for selective degradation
■
AUTHOR INFORMATION
Corresponding Author
*Fax: +1 765-496-1459. Phone: +1 765 496-1313. Notes
Moarae Matrix, Inc. has a world Wide exclusive license to the MK2 inhibitor peptide. A. Panitch owns greater than 5% of Moerae Matrix, Inc.
■
ACKNOWLEDGMENTS We would like to thank Debra Sherman for assistance with the TEM and imaging of the particles at the Purdue TEM Facility. The authors would also like to thank Yoon Yeo and Peisheng Xu for assistance with the nanosizer instrument.
■
REFERENCES
(1) Oh, J. K.; Drumright, R.; Siegwart, D. J.; Matyjaszewski, K. Prog. Polym. Sci. 2008, 33 (4), 448−477. (2) Liechty, W. B.; Peppas, N. A. Eur. J. Pharm. Biopharm. 2012, 80, 241−246. (3) Elaissari, A. Smart Colloid. Mater. 2006, 133 (9−14), 184. (4) Dai, S.; Ravi, P.; Tam, K. C. Soft Matter 2009, 5 (13), 2513− 2533. (5) Lyon, L. A.; Meng, Z. Y.; Singh, N.; Sorrell, C. D.; John, A. S. Chem. Soc. Rev. 2009, 38 (4), 865−874. (6) Blackburn, W. H.; Lyon, L. A. Colloid Polym. Sci. 2008, 286 (5), 563−569. (7) Gan, D.; Lyon, L. A. J. Am. Chem. Soc. 2001, 123 (31), 7511−7. (8) Ju, X.-J.; L., L.; Catherine Hui Niu, R. X.; Chu, L.-Y. Polymer 2009, 50, 922−929. 1210
dx.doi.org/10.1021/bm300173x | Biomacromolecules 2012, 13, 1204−1211
Biomacromolecules
Article
(9) Hu, Z.; Huang, G. Angew. Chem., Int. Ed. 2003, 42 (39), 4799− 802. (10) Ryojiro Akashi, H. T.; Komura, A. Adv. Mater. 2002, 14 (24), 1808−1811. (11) Weissleder, R.; Kelly, K.; Sun, E. Y.; Shtatland, T.; Josephson, L. Nat. Biotechnol. 2005, 23 (11), 1418−23. (12) Blackburn, W. H.; Dickerson, E. B.; Smith, M. H.; McDonald, J. F.; Lyon, L. A. Bioconjugate Chem. 2009, 20 (5), 960−8. (13) Krusic, M. K.; Ilic, M.; Filipovic, J. Polym. Bull. 2009, 63 (2), 197−211. (14) Gu, J. X.; Xia, F.; Wu, Y.; Qu, X. Z.; Yang, Z. Z.; Jiang, L. J. Controlled Release 2007, 117 (3), 396−402. (15) Meng, Z. Y.; Cho, J. K.; Debord, S.; Breedveld, V.; Lyon, L. A. J. Phys. Chem. B 2007, 111 (25), 6992−6997. (16) Hu, J. M.; Li, C. H.; Liu, S. Y. Langmuir 2010, 26 (2), 724−729. (17) Karg, M.; Hellweg, T. Curr. Opin. Colloid Interface Sci. 2009, 14 (6), 438−450. (18) Nowakowska, M.; Szczubialka, K.; Grebosz, M. J. Colloid Interface Sci. 2003, 265 (1), 214−9. (19) Garcia-Salinas, M. J.; Romero-Cano, M. S.; de las Nieves, F. J. J. Colloid Interface Sci. 2002, 248 (1), 54−61. (20) Szczubialka, K.; Jankowska, M.; Nowakowska, M. J. Mater. Sci.: Mater. Med. 2003, 14 (8), 699−703. (21) Lopes, L. B.; Furnish, E.; Komalavilas, P.; Seal, B. L.; Panitch, A.; Bentley, M. V. L. B.; Brophy, C. M. Eur. J. Pharm. Biopharm. 2008, 68 (2), 441−445. (22) Lopes, L. B.; Flynn, C.; Komalavilas, P.; Panitch, A.; Brophy, C. M.; Seal, B. L. Biochem. Biophys. Res. Commun. 2009, 382 (3), 535− 539. (23) Ishwar, A. R.; Jeong, K. J.; Panitch, A.; Akkus, O. Appl. Spectrosc. 2009, 63 (6), 636−641. (24) Gan, D. J.; Lyon, L. A. J. Am. Chem. Soc. 2001, 123 (31), 7511− 7517. (25) Hu, Z. B.; Huang, G. Angew. Chem., Int. Ed. 2003, 42 (39), 4799−4802. (26) Bailey, S. J., et al., Eds., F756; ASTM International: Baltimore, 2003; Vol. 13.01, pp 309−313. (27) Garcia-Salinas, M. J.; Romero-Cano, M. S.; de las Nieves, F. J. J. Colloid Interface Sci. 2001, 241 (1), 280−285. (28) Garcia-Salinas, M. J.; Donald, A. M. J. Colloid Interface Sci. 2010, 342 (2), 629−635. (29) Weng, Y.; Ding, Y.; Zhang, G. J. Phys. Chem. B 2006, 110 (24), 11813−7. (30) Gou, M.; Li, X.; Dai, M.; Gong, C.; Wang, X.; Xie, Y.; Deng, H.; Chen, L.; Zhao, X.; Qian, Z.; Wei, Y. Int. J. Pharm. 2008, 359 (1−2), 228−33. (31) Robb, S. A.; Lee, B. H.; McLemore, R.; Vernon, B. L. Biomacromolecules 2007, 8 (7), 2294−300. (32) Ward, B.; Seal, B. L.; Brophy, C. M.; Panitch, A. J. Pept. Sci. 2009, 15 (10), 668−674. (33) Lim, K. S.; Won, Y. W.; Park, Y. S.; Kim, Y. H. Biochimie 2010, 92 (8), 964−970. (34) Seal, B. L.; Panitch, A. Biomacromolecules 2003, 4 (6), 1572− 1582. (35) Lavelle, W.; Lavelle, E. D.; Lavelle, L. Med. Clin. North Am. 2007, 91 (2), 241. (36) Patel, P. N.; Gobin, A. S.; West, J. L.; Patrick, C. W. Jr. Tissue Eng. 2005, 11 (9−10), 1498−505. (37) South, A. B.; Lyon, L. A. Chem. Mater. 2010, 22 (10), 3300− 3306.
1211
dx.doi.org/10.1021/bm300173x | Biomacromolecules 2012, 13, 1204−1211
