Versatile Route to Colloidal Stability and Surface Functionalization of

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A versatile route to colloidal stability and surface functionalization of hydrophobic nanomaterials Heidi Renee Culver, Stephanie D Steichen, Margarita Herrera-Alonso, and Nicholas A Peppas Langmuir, Just Accepted Manuscript • DOI: 10.1021/acs.langmuir.6b00929 • Publication Date (Web): 20 May 2016 Downloaded from http://pubs.acs.org on May 24, 2016

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A versatile route to colloidal stability and surface functionalization of hydrophobic nanomaterials Heidi R. Culver,a,b Stephanie D. Steichen,a,b Margarita Herrera-Alonso,c Nicholas A. Peppas*,a,b,d,e a

Institute for Biomaterials, Drug Delivery, and Regenerative Medicine, C0800, The University of

Texas at Austin, Austin, TX 78712, United States b

Department of Biomedical Engineering, C0800, The University of Texas at Austin, Austin, TX

78712, United States c

Department of Materials Science and Engineering, The Johns Hopkins University, Baltimore,

MD 21218, United States d

McKetta Department of Chemical Engineering, C0400, The University of Texas at Austin,

Austin, TX 78712, United States e

College of Pharmacy, A1900, The University of Texas At Austin, Austin, TX, 78712

ABSTRACT: We introduce a general method for stabilization and surface functionalization of hydrophobic nanoparticles using an amphiphilic copolymer, poly(maleic anhydride-alt-1octadecene)-poly(ethylene glycol) methacrylate (PMAO-PEGMA). Coating nanoparticles with PMAO-PEGMA results in colloidally stable nanoparticles decorated with reactive carboxylic

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acid and methacrylate functionalities, providing a versatile platform for chemical reactions. The versatility and ease of surface functionalization is demonstrated by varying both the core material and the chemistry used. Specifically, the carboxylic acid functionalities are used (1) to conjugate wheat germ agglutinin to conducting polymer nanoparticles via carbodiimidemediated coupling and the methacrylate groups are used (2) to link cysteamine to the surface of poly(ε-caprolactone) nanoparticles via thiol-ene click chemistry, and (3) to link temperature responsive polymer shells to the surface of gold nanoparticles via free radical polymerization.

1. INTRODUCTION Nanomaterials are useful for biomedical applications including controlled drug delivery, in vivo imaging, and biosensing due to their unique chemical or physical properties.1–3 Prior to use, nanomaterials are commonly surface functionalized to enhance their colloidal stability and resistance to non-specific protein adsorption. Additional surface functionalization enables the nanomaterials to intelligently interact with their environment and can greatly enhance their utility. For example, decorating the nanomaterial surface with affinity ligands such as proteins or aptamers enables active targeting.4 Furthermore, environmentally responsive polymer shells can be grown on surface modified nanomaterial cores for controlled or triggered drug release.5 Surface functionalization of certain nanomaterials can be achieved through coupling techniques that are specific to their surface composition. A notable example of this is silane coupling, in which covalent bonds are formed between certain metal oxides, such as silica, and organo-functional alkoxysilanes.4 Most other inorganic nanomaterials do not readily form new covalent bonds, so they are often coated with silica to improve their stability and enable functionalization via silane coupling.6 As another example, thiols readily self-assemble on the surface of inorganic nanomaterials including gold, silver, and quantum dots, enabling


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bioconjugation through the functional groups opposite the thiols.4,7 However, the dynamic nature of the thiol-metal interaction can limit shelf life, particularly for quantum dots.8 For organic nanomaterials, polymer synthesis can be tailored to include reactive groups for bioconjugation, but this is not always straightforward.9 These examples cover only a small subset of nanomaterials and for most others there is no prevalent method for surface functionalization. As an alternative to material-specific conjugation strategies, amphiphilic molecules can be used to stabilize a variety of hydrophobic nanomaterials.4 The amphiphile imparts colloidal stability to the nanomaterials and can also provide functional groups that serve as reactive handles for surface modification.10 One commonly used amphiphile is poly(maleic anhydridealt-1-octadecene)-grafted-poly(ethylene glycol) (PMAO-PEG). PMAO-PEG was first introduced by Colvin et al. as a low-cost, easy to make polymeric amphiphile that could be used to stabilize and prevent aggregation of quantum dots and magnetic nanoparticles.11 PMAO-PEG is still most commonly employed as a stabilizer for these two nanomaterials, but its use has been extended to other inorganic nanomaterials, including carbon nanotubes,12 metal oxide nanoparticles,13 and lanthanide doped upconverting nanoparticles (Figure S1).14 The utility of PMAO-PEG as a stabilizer stems from its “polysoap” architecture: the hydrophobic octadecene chains associate with the nanomaterials and the PEG chains associate with water, providing colloidal stability that can withstand physiological conditions.12 Colvin et al. also demonstrated that the carboxyl groups generated during maleic anhydride ring opening are useful for linking amine-containing biomolecules to the particles via carbodiimide mediated coupling techniques.11 To expand the reactive potential of PMAO-PEG, we designed a methacrylated derivative to enable additional chemical routes for surface modification, such as thiol-ene click chemistry or free radical polymerization. Furthermore, this


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work extends the use of PMAO-PEG based polymers to organic nanomaterials as well, serving a whole other realm of the nanotechnology field. Herein we explore the use of PMAO-PEGMA as a stabilizer for three hydrophobic materials and demonstrate its reactive versatility. Specifically, solvent displacement techniques were used to form PMAO-PEGMA stabilized nanoparticles of (1) poly(9,9-dioctylfluorene-alt-benzothiadiazole) (PFBT), a fluorescent, semiconducting polymer, (2) poly(ε-caprolactone (PCL), a biodegradable polymer, and (3) 1-dodecanethiol capped gold nanoparticles (AuNPs). Each particle was then functionalized using a different technique as summarized in Scheme 1.

Scheme 1. Surface functionalization strategies used to modify (A) Fluorescent conducting polymer nanoparticles (PFBT CPNs), (B) biodegradable poly(ε-caprolactone) nanoparticles (PCL NPs), and (C) hydrophobically modified gold nanoparticles (AuNPs).


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2. EXPERIMENTAL 2.1 Materials Chemicals and solvents were purchased from Sigma Aldrich or Fisher Scientific and used as received. Spectra/Por dialysis tubing was used for dialysis (12-14 kDa MWCO unless otherwise specified). Ultrapure water (final resistance = 18.2 MΩ) was obtained from a Barnstead GenPure purification system from Thermo Scientific. 2.2 Instrumentation FTIR spectroscopy was performed on a VERTEX 70 spectrometer (Bruker). 1H NMR spectroscopy was carried out on a VARIAN DirectDrive 400 MHz. Fluorescence spectroscopy for critical micelle concentration determination was performed on a Fluorolog3 Fluorimeter (HORIBA Scientific). Cytation 3 Cell Imaging Multi-Mode Plate Reader (BioTek Instruments) was used for fluorescamine assay, PFBT NP concentration determination, and absorption spectroscopy of AuNPs. Dynamic light scattering measurements were taken on a


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ZetaSizer Nano-ZS (Malvern). TEM images were obtained on a FEI Tecnai Transmission Electron Microscope operating at 80 kV. Confocal images were obtained on an FV10i-DOC microscope (Olympus). 2.2

Synthesis

of

PMAO-PEGMA

Poly(maleic

anhydride-alt-1-octadecene)-grafted-

poly(ethylene glycol) methacrylate (PMAO-PEGMA) was synthesized by adapting a published protocol for synthesizing PMAO-PEG (Scheme 2).15 Briefly, 1.75 g of PMAO (average Mn 30,000-50,000) was dissolved in chloroform (20 mL) and combined with 4.5 g of PEGMA (average Mn of PEG segment = 400) dissolved in chloroform (10 mL). The mixture was stirred for 6 hours before turning the temperature to 61°C. Once the temperature had reached 61°C, several drops of concentrated sulfuric acid were added. The reaction was allowed to reflux overnight night before adding 20 mL of 1N NaOH. The solution was left refluxing for 6 hours. Chloroform was removed in vacuo and the salts and any unreacted PEGMA were removed via dialysis against ultrapure water for 7 days with frequent water changes. The product was dried via lyophilization and obtained as a fluffy, cream-colored solid: 1H NMR (400 MHz, CDCl3) δ 6.12 (s, 1H), 5.57 (s, 1H), 4.29 (b 4H), 3.64 (m, ~24H), 1.94 (s, 3H), 1.25 (br, ~32H), 0.87 (t, 3H). Scheme 2. Synthesis of PMAO-PEGMA.

2.3 Determination of critical micelle concentration of PMAO-PEGMA The critical micelle concentration (CMC) of PMAO-PEGMA was determined using the pyrene fluorescence


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method.16 A 2.4 x 10-5 M pyrene solution in acetone was prepared and 50 µL of this solution was added to each of 11 scintillation vials that were left uncapped to evaporate the acetone. A range of PMAO-PEGMA solutions (0 – 1 mg/mL) in water were prepared via serial dilution. Each polymer solution (2 mL) was added to each of the scintillation vials and vortexed for 20 seconds. The emission at 390 nm and the ratio of excitation intensities at 338 and 332.5 nm were plotted against PMAO-PEGMA concentration on logarithmic scales and fit to a sigmoidal curve. In both cases, the CMC was taken as the intersect of the tangent to low concentration points and the tangent to the inflection point. 2.4 Preparation of PMAO-PEGMA stabilized NPs For fluorescent conducting polymer nanoparticles (CPNs), poly(9,9-dioctylfluorene-alt-benzothiadiazole) (PFBT) (average Mn 17,000-23,000) was dissolved in THF (1.33 mg/mL) and added dropwise (0.5 mL/minute) to a stirring solution (400 rpm) of PMAO-PEGMA in water (2.5 mg/mL) using a Harvard Apparatus syringe pump. THF was removed via dialysis against ultrapure water for 4 days with frequent water changes. For biodegradable poly(ε-caprolactone (PCL) nanoparticles (PCL NPs), PCL (average Mn 10,000) was dissolved in acetone (6.25 mg/mL) and added dropwise to a stirring solution (400 rpm) of PMAO-PEGMA in water (5 mg/mL) using a Harvard Apparatus syringe pump. A PCL-only control was prepared using the same procedure, but in the absence of PMAO-PEGMA in the water. Acetone was removed under reduced pressure followed by dialysis against ultrapure water for 4 days with frequent water changes. Both sets of polymer nanoparticles were stored at 4°C until use. Aliquots were lyophilized with 5% sucrose (w/w) to determine the effect of the drying process on particle size and stability. For gold nanoparticles (AuNPs), colloidal gold was prepared using the Turkevich method.17 Particles were modified with 1-docecanethiol and coated in PMAO-PEGMA based on previously


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described methods for stabilizing AuNPs with amphiphilic polymers.18 Briefly, AuNPs were diluted 1:5 (100 uL:500 uL) in water and then centrifuged at 3000 x g for 30 minutes. The supernatant was removed and the particles were re-suspended in water (1 mL) and then recentrifuged at 3000 x g for 30 minutes. The supernatant was again removed and this time each pellet was suspended in 1 mL of DMF. AuNPs in DMF (3.6 mL) were combined with PMAOPEGMA in DMF (400 µL, 6 mg/mL). Ultrapure water (1 mL) was injected at a rate of 8.3µL/min. Meanwhile, a 1% solution (v/v) 1-dodecanethiol/DMF solution was prepared. After the 1 mL of water had been added, 40 µL of the 1-dodecanethiol solution was injected, followed by another 12 mL of water at 33.3 µL/min. AuNPs were dialyzed against ultrawater to remove excess 1-dodecanethiol and then centrifuged at 6500 x g for 30 min to concentrate them and remove excess PMAO-PEGMA. Particles were stored in solution at room temperature until use. 2.5 Conjugation of WGA to PFBT CPNs PMAO-PEGMA coated PFBT CPNs were suspended in 0.1M MES buffer (pH 6.0) and the carboxyl groups of PMAO-PEGMA were activated

using

1-ethyl-3-(3-dimethylaminopropyl)carbodiimide

HCl

(EDC)

and

N-

hydroxysulfosuccinimide (sulfo-NHS). Briefly, EDC (2.5 mg) and sulfo-NHS (5.5 mg) were sequentially added to PFBT nanoparticles (0.8 mg) and then mixed at room temperature for 15 minutes. Activated particles were separated from excess EDC and sulfo-NHS via centrifugation. Particles were then re-suspended in 1X PBS (2 mL) and combined with WGA in 1X PBS (2 mL, 2 mg/mL). Conjugation was carried out for 2 hours, followed by extensive dialysis against 0.1X PBS to remove unconjugated WGA (50 kDa MWCO Float-A-Lyzer, Spectra/Por). The final concentration of particles was determined using a fluorescence calibration curve and the amount of WGA conjugated per mass of particles was determined using a BCA assay from Thermo Fisher. The resultant WGA-CPN conjugates were stored at 4°C until use.


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2.6 Cell culture and confocal microscopy L929 murine fibroblast cells were maintained in high-glucose Dulbecco’s Modified Eagle’s Media (DMEM) supplemented 10% fetal bovine serum, 1% L-glutamine, and 1% penicillin-streptomycin solution. Round glass coverslips (18 mm in diameter) were sterilized for cell seeding and imaging by subsequent incubations at 37°C in 1N HCl (8-10 hours), 25:75 ethanol:water (20 minutes), 50:50 ethanol:DI (20 minutes), and finally 75:25 ethanol:DI (10 minutes). Glass coverslips were rinsed with modified DMEM and then added to a 12-well plate. L929 cells at passage 15 were seeded at a concentration of 18,750 cells/well and allowed to grow to 70-80% confluency. After reaching the desired confluency, cells were rinsed twice with ice cold 1X PBS and then incubated in ice cold IC Fixation Buffer (Invitrogen) for an hour at room temperature. The fixation buffer was removed and then the cells were rinsed twice with room temperature 1X PBS. To the cells, either a 0.02 mg/mL WGA-AlexaFluor488, 0.02 mg/mL WGA-CPN, or a 0.02 mg/mL unconjugated CPN solution was added and allowed to incubate for either 10 minutes (for WGA alone) or 30 minutes (for WGA-CPNs and unconjugated CPNs). After incubation, the cells were rinsed with 1X PBS twice and then allowed to dry overnight at 4oC. The coverslips were then removed from the 12-well plate and affixed to microscope slides with ProLong Gold AntiFade reagent with DAPI (Molecular Probes by Life Technologies). The mounted coverslips and slides were dried for 24-48 hours before imaging. Images were obtained on an FV10i-DOC inverted laser-scanning confocal microscope (Olympus) using the built-in UPLSAPO 60X phase contrast oil immersion objective (NA=1.35) with the confocal aperture diameter set to 1 Airy disk unit and zoom of 1.7x. DAPI was excited using a 405 nm laser and detected using a 420-460 nm barrier filter setting. AlexaFluor488/PFBT CPNs were excited using a 473 nm laser and detected using a 490-590 nm barrier filter setting. The two fluorophores


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were detected sequentially with PMT detector sensitivity of 620V and a Kalman average of 4 lines for all images. Olympus Fluoview software was used to adjust brightness and contrast, keeping adjustments the same for compared images. ImageJ was used to quantify the intensity of cell membrane staining compared to background intensity. Large fluorescent aggregates were excluded from quantification of the background intensity. 2.7 Conjugation of cysteamine to PCL NPs Cysteamine hydrochloride (5 mL, 5 mg/mL) and ammonium persulfate (APS) (1 mL, 1 mg/mL) were combined in a small round bottom flask and placed in an ice bath. PCL NPs (2 mL, 4 mg/mL) were added dropwise to the cysteamine HCl/APS solution under rapid stirring. The reaction was left stirring on ice for 150 minutes. Excess APS and cysteamine HCl were removed via dialysis against ultrapure water for 4 days with frequent water changes. Aliquots (1 mL) were lyophilized and weighed to determine the mass concentration. The reaction was repeated in triplicate. The amount of cysteamine conjugated to the PMAO-PEGMA coated PCL NPs was determined using a fluorescamine assay. PCL NPs conjugated to cysteamine and unconjugated PCL NPs (negative control) solutions were prepared at concentrations ranging from 0-0.35 mg/mL in 1X PBS. Fluorescamine in acetone (0.3 mg/mL, 20 µL) was added to each particle concentration (1 mL) and mixed end-over-end for 15 minutes at room temperature in the dark. The fluorescence intensity at 460 nm (λex = 360 nm) was measured for each particle concentration. The amount of cysteamine/mass of particle was determined by comparing fluorescence intensity values to a calibration curve prepared using cysteamine HCl. 2.8 Growth of poly(NIPAM-co-MAA) on AuNPs PMAO-PEGMA coated AuNPs (4.5 mL, 0.78 nM) were combined with 5 mL of each monomer solution (methylene bisacrylamide, BIS, 1.24 mg/mL: 5 mol%), (N-isopropylacryamide, NIPAM, 13.6 mg/mL: 75 mol%), and


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(methacrylic acid, MAA, 2.75 mg/mL, 20 mol%). APS in water (0.5 mL, 60 mg/mL) was injected after heating the reaction to 70°C and nitrogen purging for 30 minutes. Nitrogen was bubbled through the reaction for another 10 minutes, and then the reaction was allowed to proceed for 4 hours. NPs were purified via dialysis against ultrapure water for 4 days with frequent water changes. 3. Results and discussion 3.1 PMAO-PEGMA characterization Successful ring opening of maleic anhydride and grafting of PEGMA to PMAO were confirmed using FTIR (Figure S2) and 1H NMR spectroscopy. By comparing the integrals of the 1H NMR peaks corresponding to the protons of the terminal methyl group of octadecene (δ 0.87) and the vinylic protons of PEGMA (δ 6.12 and 5.57), it was determined that PEGMA was grafted to 50% of the maleic anhydride groups (Figure S3). After confirming the chemical structure, the critical micelle concentration (CMC) of PMAO-PEGMA was determined to be 3.34 ± 2.58 µg/mL using the pyrene fluorescence method (Figure 1, Figure S4, Table S1).16 This value is much lower than for small molecule surfactants and is comparable to or lower than reported CMC values for other amphiphilic polymers.19,20 This low CMC suggests that NPs stabilized with PMAO-PEGMA will have excellent thermodynamic stability even in dilute conditions.21


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Figure 1. Determination of PMAO-PEGMA CMC. Representative data showing the ratio of pyrene excitation intensity at 338 nm to excitation intensity at 332.5 nm (I338/I332.5) (λem = 390 nm). The excitation I338/I332.5 ratio of pyrene is constant at concentrations before micelles begin to form (i.e., below the CMC), but then increase with increasing PMAO-PEGMA concentration as pyrene transitions from water into the less polar interior of the micelles. The CMC was measured for three independent PMAO-PEGMA batches and in each case was determined from the intersection of the tangent to low concentration points and the tangent to the inflection point. The average CMC determined based on the I338/I332.5 excitation ratio data (3.34 ± 2.58 µg/mL) was in close agreement to the CMC determined by fluorescence intensity (3.05 ± 3.18 µg/mL) (Figure S4 and Table S1). 3.2 NP-protein conjugation via carbodiimide chemistry In the first application, PFBT CPNs were coated by PMAO-PEGMA to form highly fluorescent CPNs. CPNs have extraordinary brightness and large two-photon absorption cross-sections, making them useful for imaging and sensing applications.22,23 After nanoprecipitation, the structural characteristics of the CPNs were characterized using TEM and DLS. TEM confirmed that the particles were spherical and had an


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average diameter of 93.1 ± 15.3 nm (Figure 2A). This dried state diameter was smaller than the hydrodynamic diameter found by DLS (174.1 ± 81.8 nm), likely due to hydration shells generated from interactions between the PEG chains and water molecules (Figure 2B).24 CPNs had a highly negative zeta potential of -52.4 ± 7.4 mV due to the presence of pendant carboxyl groups of PMAO-PEGMA, which were used to link wheat germ agglutinin (WGA) to the CPNs via carbodiimide chemistry (Scheme 1A).

Figure 2. Characterization of PMAO-PEGMA stabilized PFBT CPNs. (A) TEM image of PMAO-PEGMA stabilized PFBT CPNs stained with 2% uranyl acetate. Scale bar represents 200 nm. (B) DLS intensity size distribution of PMAO-PEGMA stabilized PFBT CPNs (Dh = 174.1 ± 81.8 nm, PDI = 0.221 ± 0.129, ZP = -52.4 ± 7.4 mV). (C) Fluorescence excitation and emission spectra of PFBT CPNs (Excitation spectra: λem = 535 nm, λex, max = 466 nm; emission spectra: λex = 425 nm, λex, max = 542 nm). WGA was used as a model affinity ligand because it selectively binds N-acetylglucosamine and N-acetylneuraminic acid, which are found on cell membranes.25 Using a BCA assay, the amount of WGA conjugated to the CPNS was found to be 105 nmol/mg particle (Figure S5). Based on the number of particles per milliliter (4.6 ± 0.3 x 1011 particles/mL) as determined by


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Nanoparticle Tracking Analysis (Malvern NanoSight), a CPN surface area of 9.52 x 104 nm2 (based on Dh = 174.1 nm), and a projected area of 24.63 nm2 per WGA molecule (assuming Dh = 5.6 nm26), this corresponds to 985 WGA molecules/particle or approximately 25.5% surface coverage. While there was no observable effect of WGA conjugation on CPN stability in the week following the coupling reaction (during which the experiments were performed), long-term storage in 0.1X PBS (6 months) led to aggregation, likely due to electrostatic interactions between WGA (pI ~ 8.5) and anionic CPNs as well as general instability of proteins in solution. To test the targeting ability of these CPN-WGA conjugates, they were used to stain cell membranes of fixed L929 fibroblasts. WGA-AlexaFluor488 conjugates were used as a positive control because AlexaFluor488 has similar excitation and emission spectra as PFBT CPNs (Figure 2C). Confocal microscopy showed that CPN-WGA conjugates provided high contrast staining of L929 fibroblast membranes (Figure 3A). However, staining with CPN-WGA conjugates was more diffuse than WGA-AlexaFluor488 conjugates, likely due to the large size of the CPN relative to the small molecule fluorophore (Figure 3B). Interestingly, CPNs not conjugated to WGA also localized on the cell membranes, but the cell membrane/background fluorescence ratio was 36% lower than that of the CPN-WGA conjugates and there was more aggregation of CPNs compared to CPN-WGA conjugates (Figure 3C). This example demonstrates how PMAO-PEGMA may be used to link amine-containing molecules to nanomaterials using EDC/NHS coupling and demonstrates the utility of CPNs for imaging applications.


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Figure 3. Confocal images of L929 fibroblasts. Plasma membranes were stained by (A) WGACPN conjugates, (B) WGA-AlexaFluor488 conjugates, or (C) unconjugated CPNs. In all cases, nuclei were stained with DAPI. Scale bar represents 20 µm. Image acquisition and processing settings were kept the same for comparing membrane stain intensity. 3.3 NP-thiol conjugation via thiol-ene reaction In the second application, PMAO-PEGMA was used to stabilize nanoparticles of PCL (PCL NPs). PCL is useful as a drug delivery carrier because of its biodegradability and in nanoparticulate form can host hydrophobic drugs or imaging agents.27 While PCL NPs can be made via solvent displacement without the addition of a surfactant (Dh = 198.4 ± 50.6 nm), they quickly aggregate and fall out of solution (Figure S6). Colloidal stability of PCL NPs can be significantly improved by coating them with an amphiphilic stabilizer during the solvent displacement process. Colloidal stability of PMAOPEMGA coated PCL NPs was demonstrated by measuring hydrodynamic diameter after varied storage conditions. Specifically, periodic DLS measurements over 40 days show minimal changes in size distribution and polydispersity over time, indicating NP stability when stored in water (Figure 4A). If PMAO-PEGMA had detached from the PCL NPs over this long-term storage, we would have expected the PCL to form large aggregates similar to the PCL-only control. PCL NPs were also stored in water, 1XPBS, or cell culture media for 12 hours to probe


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colloidal stability in more physiologically relevant conditions. The average hydrodynamic diameter of the PCL NPs decreased slightly in 1XPBS (-9.4 nm) and increased slightly in cell culture media (+11.8 nm) compared to PCL NPs in water (Figure 4B). In all cases the PDI remained below 0.204 and no large aggregates were observed, suggesting that PMAO-PEGMA does aid in NP stability. Lastly, when lyophilized without a cryoprotectant, large aggregates (around 3-5 µm) were detected by DLS when particles were resuspended in water, however when lyophilized with a common cryoprotectant (5% sucrose) no large aggregates were observed (Figure 4C). With sucrose, there was a slight increase in the hydrodynamic diameter of 10.0 nm and PDI increased from 0.104 to 0.134 after lyophilization. Like the CPNs, PCL NPs had a highly negative zeta potential of -64.6 ± 1.3 mV. TEM images reveal that PCL NPs are spherical with an average diameter of 227.5 ± 114.9 nm (Figure 4D), which was larger than the hydrodynamic diameter determined by DLS (156.4 ± 45.2 nm). This discrepancy may be due to coalescence of particles upon drying on the TEM grid.

Figure 4. Characterization of PMAO-PEGMA stabilized PCL NPs. DLS measurements were taken to test the stability of PCL NPs after (A) 40 days stored in water, (B) 12 hours incubated in water, 1XPBS, or cell culture media, or (C) lyophilization in 5% sucrose (w/w). (D) TEM image of PCL NPs stained with 2% uranyl acetate. Scale bar represents 500 nm.


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With these PCL NPs, we demonstrate the use of methacrylate groups to link thiol-containing molecules to PMAO-PEGMA stabilized nanoparticles via a free-radical thiol-ene reaction. Click reactions between thiols and enes can be performed under a variety of mild conditions, making them an attractive route for bioconjugation.28 It has been shown that the presence of oxygen, low reaction temperatures, and excess thiol can increase conversion in free radical thiol-ene reactions by diminishing side reactions such as thiol oxidation and reactions between methacrylate groups.29–31 In accordance with these findings, reactions were carried out in an ice bath in the presence of air and with an excess of cysteamine (Scheme 1B). Using a fluorescamine assay to determine the number of amine groups present, the amount of cysteamine was measured at 0.85 mg cysteamine/mg particle (Figure 5A, Figure S7). Successful surface modification was further confirmed by a 36.3 mV increase in the zeta potential of the PCL NPs after reaction with cysteamine. The hydrodynamic diameter of PCL NPs decreased by 10 nm after modification, possibly due to favorable interactions between the terminal amine groups from cysteamine and carboxyl groups of PMAO-PEGMA, causing collapse of the PEG corona (Figure 5B). The PDI did not increase greatly, suggesting that despite any change in conformation of the PEG chains, they still provided effective stabilization. Many thiol-modified biomolecules (e.g., aptamers or peptides) and thiols with additional reactive groups are commercially available and widely used for functionalization of gold or quantum dots.4 Now, through thiol-ene reactions, immobilization of these molecules could be achieved on any nanomaterial coated with PMAO-PEGMA.


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Figure 5. Characterization of PMAO-PEGMA coated PCL NPs after cysteamine modification. (A) Fluorescence intensity of fluorescamine at 460 nm (λex = 360 nm) as a function of PCL NP concentration before and after conjugation to cysteamine (n=3, mean ± SD). (B) DLS intensity size distributions showing slight decrease in average hydrodynamic diameter and increase in polydispersity after conjugation to cysteamine (Dh = 144.7 ± 63.2 nm, PDI = 0.200 ± 0.012). 3.4 Growth of responsive polymer shells on NPs via free-radical polymerization In the final application, PMAO-PEGMA was used to stabilize hydrophobically modified AuNPs. Gold nanomaterials are widely used in biosensing, imaging, and triggered release of therapeutics.32


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PMAO-PEGMA improved the stability of AuNPs in 1X PBS compared to their citrate capped predecessors, as determined by broadening of the absorbance peak of citrate capped AuNPs but not PMAO-PEGMA coated AuNPs (Figure 6A). PMAO-PEGMA coated AuNPs had a zeta potential of -23.4 ± 6.4 mV and uniform structure with diameter of 13.7 ± 0.8 nm (Figure 6B).

Figure 6. Characterization of PMAO-PEGMA coated AuNPs. (A) Broadening of the localized surface plasmon resonance (LSPR) peak of gold nanomaterials indicates instability. There was significant peak broadening of citrate capped AuNPs in 1X PBS (red) but not for PMAO-PEGMA coated AuNPs (black), indicating improved stability after coating with PMAOPEGMA. (B) TEM image of PMAO-PEGMA coated AuNPs. Scale bar represents 100 nm.


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Gold nanomaterials are useful for controlled release of therapeutics because of their large, tunable absorption cross-sections, which enable localized heating and collapse of Nisopropylacrylamide (NIPAM) based polymers. This thermally responsive behavior can be used to trigger drug release or localize particles at areas of irradiation.5,33 In this work, NIPAM was co-polymerized with methacrylic acid (MAA) and crosslinked with methylene bisacrylamide to generate polymer shells on PMAO-PEGMA coated AuNPs (Scheme 1C). Core-shell architecture was confirmed by TEM (Figure 7A), with each particle containing a single AuNP roughly centered in the polymer. This suggests two things: (1) PMAO-PEGMA prevented aggregation of the AuNPs during polymerization and (2) the methacrylate groups of PMAOPEGMA facilitated growth of polymer shells via radical capture of oligomers and monomers.34 Increasing the seed concentration of AuNPs could prevent the observed secondary nucleation (i.e., particles without AuNPs cores). The narrow distribution (PDI = 0.027 ± 0.016) and absence of large particles (>400 nm) detected by DLS suggest that the coalescence observed on TEM was due to the drying process rather than crosslinking or aggregation of the NPs in solution (Figure 7A-C). DLS was used to characterize the temperature responsive behavior of the core-shell particles. As expected, the hydrodynamic diameter decreased near the lower critical solution temperature (LCST) of NIPAM (32-34°C), shrinking by ~90 nm as temperature increased from 16-60°C (Figure 7D). This example demonstrates the utility of PMAO-PEGMA as a modifier for nanomaterials on which one wishes to grow environmentally responsive polymer shells via free-radical polymerization.


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Figure 7. Characterization of AuNP-poly(NIPAM-co-MAA) core-shell NPs. (A) High magnification TEM image showing core-shell architecture and (B) low magnification TEM image demonstrating coalescence of NPs observed on TEM grids (arrows). Particles were stained with 2% uranyl acetate. Scale bars represent 100 nm (A) and 1 µm (B). (C) DLS distribution of AuNP-poly(NIPAM-co-MAA) core-shell particles at 24°C. The dry state diameter (133.7 ± 16.3 nm) is approximately 58% of the swollen state diameter (230.5 ± 37.9 nm). (D) NP diameter as a function of temperature showing LCST behavior of poly(NIPAM-co-MAA) coated AuNPs. 4. Conclusion The use of PMAO-PEGMA as an amphiphilic stabilizer opens up new avenues for functionalization of hydrophobic nanomaterials. We demonstrate that, in addition to inorganic nanomaterials, polymeric NPs can be made via solvent displacement with PMAO-PEGMA as the stabilizer. By modifying PMAO-PEG to include a methacrylate group, free radical based thiol-ene reactions and polymerizations were possible. We believe that PMAO-PEGMA will be of great utility to other scientists, as it provides a more universal method for nanomaterial stabilization and functionalization. ASSOCIATED CONTENT


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Supporting Information. The supporting information is available free of charge: Distribution of nanomaterials previously modified with PMAO-PEG, polymer characterization (FTIR, 1H NMR, CMC), BCA assay for quantifying WGA conjugation to CPNs, photos demonstrating instability of uncoated PCL NPs, fluorescamine assay calibration curve (PDF) AUTHOR INFORMATION Corresponding Author *E-mail: [email protected] Notes The authors declare no competing financial interests. ACKNOWLEDGMENT We thank the Cockrell Family Regents Chair for their generous support. During this work, HRC was supported by a National Science Foundation Graduate Research Fellowship (DGE1610403). We would like to acknowledge the Institute for Cellular and Molecular Biology, particularly Dwight Romanovicz for TEM technical support. REFERENCES (1) (2) (3) (4)

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