Orientation Controlled Protein Nanocapsules by Enzymatic Removal

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Orientation Controlled Protein Nanocapsules by Enzymatic Removal of a Polymer Template Chaeyeon Lee, Aran Hwang, Leeja Jose, Ji Hyun Park, Jaekwang Song, KyuHwan Shim, Seong Soo A An, and Hyun-jong Paik Biomacromolecules, Just Accepted Manuscript • DOI: 10.1021/acs.biomac.8b00965 • Publication Date (Web): 28 Sep 2018 Downloaded from http://pubs.acs.org on September 28, 2018

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Biomacromolecules

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Orientation Controlled Protein Nanocapsules by

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Enzymatic Removal of a Polymer Template

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Chaeyeon Lee,† Aran Hwang,† Leeja Jose,† Ji Hyun Park,‡ Jae Kwang Song‡, KyuHwan Shim, §

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Seong Soo A.An, § and Hyun-jong Paik†,*

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AUTHOR ADDRESS †

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Department of Polymer Science and Engineering, Pusan National University, Busan, Korea

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46241 ‡

Research Center for Bio-Based Chemistry, Korea Research Institute of Chemical Technology (KRICT), Daejeon, Korea 34114

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§

Department of Bionano Technology, Gachon University, Sungnam, Korea 13120

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KEYWORDS.

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Nitrilotriacetic acid (NTA), biodegradable polymer, non-covalently connected micelle,

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enzymatic core degradation, protein nanocapsule, orientation control of protein

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ABSTRACT. Protein nanocapsules are potentially useful as functional nanocarriers because of

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their hollow structure and high biocompatibility, and the intrinsic activity of their protein

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constituents. However, the development of a facile method for the preparation of oriented

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Biomacromolecules 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

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nanocapsules that retain their protein activity has been challenging. Here we describe the

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preparation of protein nanocapsules through the enzymatic removal of polymer templates.

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Nickel(II) nitrilotriacetic acid-end-functionalized poly(lactic acid) (Ni2+-NTA-PLA) was

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introduced as a polymeric template to immobilize hexa-histidine-tagged green fluorescence

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protein (His6-GFP) with consistent orientation. Following protein cross-linking and core-

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degradation, various measurements as a function of degradation time indicated the formation of

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hollow structures. We also demonstrated orientational control and activity preservation of the

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protein after capsule preparation. Protein nanocapsules prepared by this method can act as

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functional containers, taking advantage of the intrinsic function of their constituent proteins

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without additional modification.

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INTRODUCTION

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Protein nanocapsules have been developed as hollow, biocompatible vehicles.1-3 These structures

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are also broadly applicable since their constituent proteins can be of many different classes,

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including enzymes, antibodies, and antigens. One of the key factors that allow individual

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proteins in such structures to maintain their intrinsic function are the relative exposure and

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orientation of the active site.4-6 In this manner, various approaches have been reported for

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controlling the orientation of proteins immobilized on a substrate.4, 7-10 However, most methods

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used to prepare protein capsules use synthetic support systems based on non-specific adsorption

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and so do not allow for orientational control.2, 11-19 This study, which resulted in a new method

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for the preparation of hollow protein nanocapsules, focuses on 1) mild preparation conditions

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that do not destroy protein activity, and 2) the control of protein orientation.


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Biomacromolecules

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There have been two main approaches to forming hollow, protein-based structures. The first is

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to induce the self-assembly of protein subunits. This method is a traditional biological approach

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that results in monodisperse and consistently oriented protein capsules.3, 20-24 However, this

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method is only applicable to a few types of protein and requires delicate protein engineering to

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assign functionalities such as antigenicity, ligand-receptor interactions, and catalysis.24 The

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second approach employs a synthetic support, such as inorganic particles or an oil emulsion,

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which acts as a template that is removed following formation of the protein shell. This approach

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enables control of the size of the protein capsules by changing the size of the colloidal or

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inorganic particles.2, 15, 16, 18 However, many proteins denature if exposed to organic solvents

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during emulsion formation or harsh chemicals during the removal of an inorganic template.2, 17, 25,

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26

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generally immobilized on the template by non-specific adsorption. Each of the aforementioned

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approaches has strengths and weaknesses.

Also, controlling protein orientation with this method is difficult because proteins are

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Scheme 1. Preparation of protein nano capsule consisting of GFP (GFP-NC) through shell

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crosslinking and core degradation.


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The method described herein combines the advantages of these two approaches to efficiently

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manufacture hollow protein structures with a defined orientation. This is the first report detailing

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the use of a synthetic support to create an orientationally controlled nanocapsule system with

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preserved protein function. We synthesized nickel(II) nitrilotriacetic acid-end-functionalized

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poly(lactic acid) (Ni2+-NTA-PLA) and prepared PLA particles coated with hexa-histidine-tagged

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green fluorescence protein (His6-GFP). The polymeric core of these particles can be

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subsequently dissolved under mild conditions. The resulting hollow nanocapsules, composed

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solely of His6-GFP, were obtained following cross-linking of the protein shell and removal of the

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PLA core by enzymatic hydrolysis. This procedure is illustrated in Scheme 1. We demonstrated

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the potential of this system by fabricating protein nanocapsules with high intrinsic protein

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activity and consistent orientation.

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EXPERIMENTAL SECTION

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Materials. N,N-Bis[(tert-butyloxycarbonyl)methyl]-L-lysine tert-butyl ester (t-boc-NTA-NH2)

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was synthesized in accordance with a previously reported procedure.27 His6-GFP (six histidine-

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tagged green fluorescent protein) was expressed and purified according to previously published

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procedures.28 Acid-terminated poly(D,L-lactic acid) (PLA) (Mw 10,000–18,000), N-

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hydroxysuccinimide (NHS) (98%), N-(3-dimethylaminopropyl)-N′-ethylcarbodiimide

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hydrochloride (EDC) (commercial grade), trifluoroacetic acid (TFA) (99%), nickel(II) chloride

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(NiCl2) (98%), glutaraldehyde solution (grade I, 25% in H2O), sulforhodamine 101 (SR101),

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rhodamine B isothiocyanate-dextran (average mol wt ~ 10,000 and 70,000) (RITC-dex), and

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trypsin from bovine pancreas (type I, ~10,000 BAEE units/mg protein) were purchased from


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Biomacromolecules

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Aldrich (St. Louis, MO, USA). Sodium tetrahydroborate (NaBH4) (98%) was purchased from

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JUNSEI (Tokyo, Japan). Lipozyme TL 100L (100 KLU/g) was purchased from Novozymes

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(Bagsværd, Denmark).

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Measurements. Molecular weights (Mn) and molecular weight distributions (Mw/Mn) were

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determined using size exclusion chromatography against poly(methyl methacrylate) standards

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(SEC; Agilent, Santa Clara, CA, USA). The SEC was equipped with an Agilent 1100 pump, RID

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detector and PSS SDV (5 µm, 105, 103, 102 Å 8.0 mm × 300.0 mm) columns. Tetrahydrofuran

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(THF) was used as a mobile phase at a flow rate of 1.0 mL/min and a column temperature of 313

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K. Preparative high-performance liquid chromatography (HPLC) was performed with a YMC K-

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50 HPLC pump and YMC-GPC T30000 and T2000 columns were used to purify the resulting

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polymer. THF was used as a mobile phase at a flow rate of 10 mL/min. Matrix-assisted laser

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desorption/ionization time of flight mass spectrometry (MALDI-TOF MS) was performed with a

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an Autoflex speed mass spectrometer (Bruker, Billerica, MA, USA) equipped with a 2-kHz

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smartbeam-II laser. An accelerating voltage of 20 kV was applied in positive mode. Mass

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calibration was performed using homemade polystyrene (PS) standards. Each polymer sample

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was dissolved in THF to a concentration of 5 mg/mL. The cationization agent used was sodium

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trifluoroacetate dissolved in THF to a concentration of 2 mg/mL. The matrix trans-2-(3-4-tert-

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butylphenyl)-2-methyl-2-propenylidene)malononitrile (DCTB) was dissolved in THF to a

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concentration of 30 mg/mL. Stock solutions were mixed in a 10/1/1 ratio (matrix/analyte/cation)

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and deposited onto a MALDI target plate. Proton nuclear magnetic resonance (1H NMR) spectra

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were obtained on a Unity Inova 500 spectrometer (500 MHz; Varian, Palo Alto, CA, USA) and a

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Varian Unity Plus 400 spectrometer (400 MHz) at room temperature (RT) using CDCl3 and D2O.

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(Varian Deutschland Gmbh, Darmstadt, Germany). Dynamic light scattering (DLS) was


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Page 6 of 29

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performed with a 90 plus Particle Size Analyzer (Brookhaven Instruments Corporation, New

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York, NY, USA). Atomic force microscopy (AFM) was performed with an n-Tracer SPM

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(NanoFocus, Oberhausen, Germany). The cantilever was composed of silicon and had a

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resonance frequency of approximately 320 kHz and a nominal radius of curvature of less than 8

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nm. For AFM analyses, solutions of protein nanocapsules were spin-coated onto glass coverslips

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that had been washed with Piranha solution and dried under vacuum.29 AFM images were

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obtained in air at RT. Transmission electron microscopy (TEM) was performed on a Hitachi H-

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7600 instrument (Hitachi High-Technologies, Tokyo, Japan) at 80 kV. For TEM measurements,

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nanocapsule solution was dropped onto a carbon-coated copper grid, followed by negative

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staining with uranyl acetate solution. Photoluminescence (PL) spectra were obtained at an

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excitation wavelength of 470 nm or 583 nm (HR4000CG composite-grating spectrophotometer;

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Ocean Optics Inc., Dunedin, FL, USA). Protein nanocapsules were imaged on a Leica TCS SP8

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inverted microscope (Wetzlar, Germany). The resulting micrographs were analyzed with Leica

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software (LAS X) and visualized with an HCX PL APO 100× objective lens (numerical aperture,

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1.40) using a 458-nm argon laser and an HyD detector (462–520 nm) for GFP excitation and

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emission, and a 580-nm laser and HyD detector (591-670 nm) for guest molecule excitation and

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emission.

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Synthesis of t-boc-NTA-PLA (1). t-boc-NTA-PLA (1) was prepared by conjugating t-boc-

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NTA-NH2 with acid-terminated PLA through an EDC-coupling reaction. PLA (300 mg), NHS

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(5.30 mg, 0.0462 mmol), and EDC (7.20 mg, 0.0462 mmol) were added to a Schlenk flask

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containing 10 ml of dried dichloromethane (MC). The mixture was cooled to 0°C and stirred for

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30 min. After the NHS activation process, a solution of t-boc-NTA-NH2 in 2 ml of dried MC was

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added dropwise into the mixture and a flask was put into an oil bath at 25°C for 24 h. The


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Biomacromolecules

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resulting solution was washed with distilled water (3 × 15 mL). After the removal of MC, the

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obtained product was dissolved into THF. Then, t-boc-NTA-PLA (1) was isolated by preparative

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HPLC (mobile phase: THF) using the preparative column at a flow rate of 10 mL/min. Proton

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nuclear magnetic resonance (1H NMR) (CDCl3) showed the following: δ 1.10–1.11 (d, 3H); 1.44

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(s, 18H); 1.46 (s, 9H); 1.51–1.60 (m, 234H); 3.25 (m, 2H); 3.29 (t, 1H); 3.45 (dd, 4H); 4.36 (m,

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1H); 5.10–5.30 (m, 76H). The molecular weight of the obtained polymer was determined by SEC

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(Mn,SEC = 9,940 g/mol, Mw,SEC/Mn,SEC =1.41) and 1H NMR (Mn,NMR = 6,050 g/mol).

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Synthesis of NTA-PLA (2). The protecting group of NTA was removed as reported previously.28 t-boc-NTA-PLA (240 mg) was dissolved in 10 ml of dried MC in a flask. TFA (110

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µL, 1.44 mmol) was then slowly added to the flask and the solution was stirred at RT for 24 h.

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After evaporating TFA under reduced pressure, the product (2) was re-dissolved in 3 mL of MC

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and precipitated in 100 mL of isopropyl ether. The resulting precipitate, obtained by

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centrifugation (12,000 rpm for 30 min × 3), was dried in vacuo at 30°C for 24 h. The yield was

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200 mg. 1H NMR (CDCl3) showed the following: δ 1.10–1.11 (d, 3H); 1.51–1.60 (m, 231H);

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3.00–3.60 (m, 7H); 4.36 (m, 1H); 5.10–5.30 (m, 75H). The molecular weight of the obtained

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polymer was determined by SEC (Mn,SEC = 10,800 g/mol, Mw,SEC/Mn,SEC = 1.51) and 1H NMR

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(Mn,NMR = 5,806 g/mol).

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Synthesis of Ni2+-NTA-PLA (3). Finally Ni2+-NTA-PLA (3) was prepared with the addition

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of NiCl2 (156 mg, 1.2 mmol) to the solution of NTA-PLA (2) (200 mg) in 10 mL

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dimethylformamide (DMF). The mixture was stirred at RT for 24 h and precipitated in 300 mL

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of methanol. The resulting precipitate, obtained by centrifugation (12,000 rpm for 30 min × 3),

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was dried in vacuo at 30°C for 24 h. The yield was 170 mg.


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Preparation of His6-GFP-coated PLA nanoparticles (GFP/PLA). Ni2+-NTA-PLA (3) (2

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mg) was dissolved in dried DMF (1 mL). An aliquot (200 µL) of the Ni2+-NTA-PLA solution

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was added to 20 ml of phosphate buffer solution (PBS) (10mM, pH 7.5) containing His6-GFP

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(27 kDa, 2.1 mg) at a rate of 0.08 mL/h using a syringe pump under rapid stirring at 25°C. After

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the addition of Ni2+-NTA-PLA, the solution was stirred continuously to stabilize the protein-

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polymer hybrids. Residual protein that was not conjugated to Ni2+-NTA-PLA in the GFP/PLA

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solution was removed by filtering through a Ni2+-NTA agarose resin. The diameter of the

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resulting structure was measured by DLS. The binding efficiency of His6-GFP to the polymer

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was calculated by comparing the fluorescence intensity before and after resin filtration.

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Protein shell cross-linking of GFP/PLA. His6-GFP shells were cross-linked by reactions

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between amine and aldehyde groups. GFP/PLA solution (20 mL, GFP concentration: 74.5

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µg/mL) was added slowly to 240 µL of glutaraldehyde solution (5% in H2O) with stirring. The

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resulting solution was stirred constantly at RT for 1 h. After the reaction, 1.44 mL of NaBH4

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stock solution (10 mg/mL in H2O) were added to the GFP/PLA solution to reduce the imine and

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any residual aldehyde groups. After 1 h, the cross-linked GFP/PLA (C-GFP/PLA) was washed

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with 60 mL of PBS by centrifugal filtration (2,500 rpm for 30 min × 4) through an Amicon

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Ultra-15 (Mn cut-off 10 KDa; Amicon, Lexington, MA, USA) filter.

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Verification of enzymatic PLA degradation by 1H NMR analysis. C-GFP/PLA (10 mL)

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was washed repeatedly with deuterated PBS (10 mM, pH 7.5). Centrifugal filtration was used to

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concentrate the C-GFP/PLA solution 10-fold (2,500 rpm for 30 min × 3). Lipozyme TL 100L (6

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µL) was added to an NMR tube containing 1 mL of the concentrated C-GFP/PLA. The NMR

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tube was sealed and the solution was maintained at 25°C. PLA degradation was monitored by


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Biomacromolecules

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observing the relative peak area of the lactic acid (LA) methyl proton peaks over 20 days. Peak

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areas were normalized to the D2O peak area.

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Preparation of cross-linked GFP nanocapsules (C-GFP-NC) through core degradation.

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C-GFP/PLA was used without further treatment. Lipozyme TL 100L (3 µL, lipase) was added to

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a vial containing 5 mL of C-GFP/PLA. C-GFP/PLA with or without lipase was kept at 25°C and

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stirred for 28 days. After 28 days, 84.7 mg of imidazole was added to the solution of C-

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GFP/PLA with lipase to dissociate any NTA-Ni2+/His interactions. Centrifugal filtration

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(Amicon Ultra-15; Mn cut-off 100 KDa) was used to repeatedly wash the solution with PBS, with

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and without 250 mM imidazole, to remove lipase and residual polymer. During washing, any

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GFP that was not cross-linked with circumferential GFPs was also removed. The yield of C-

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GFP-NC was calculated by comparing the fluorescence intensity before and after imidazole

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washing. Changes in the height and diameter of C-GFP/PLA nanocapsules after lipase treatment

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as a function of degradation time were observed by DLS and AFM measurements. The

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maintenance of green fluorescence and spherical structures after C-GFP-NC preparation was

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measured by PL, super resolution confocal micrographs (SRCM), and TEM.

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Preparation of C-GFP-NC of different sizes. NTA-PLA (3) was dissolved in dried DMF to

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a concentration of 1 or 3 mg/mL. Each GFP/PLA particle solution was prepared and purified

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using the same conditions and methods described above for GFP/PLA preparation. The

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diameters of resulting structures were measured by DLS. The GFP/PLA particles were converted

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to C-GFP-NC using the same methods and cross-linking and lipase treatments described above.

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The final C-GFP-NCs were characterized by DLS, TEM and AFM measurements.


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Binding of His6-GFP and GFP/NC to Ni2+-NTA resin. A His6-GFP solution (3 mL, 31.5

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µg/mL) was prepared as a control. Solutions of C-GFP-NC (1.5 mL) and His6-GFP were added

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to a column packed with 150 µL of Ni2+-NTA agarose resin (binding capacity: 5–10 mg His-Tag

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fusion protein per mL resin; Merck Millipore, Billerica, MA, USA). The solutions were slowly

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eluted from the column. Changes in the intensity of green fluorescence before and after Ni2+-

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NTA resin filtration were measured by PL.

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Penetration of hydrophilic guest molecules into C-GFP-NC as a function of molecular

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weight. The final C-GFP-NC solution (3 mL, GFP concentration: 55.6 µg/mL) was concentrated

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to 417 µL (GFP concentration: 0.4 mg/mL) by centrifugal filtration (2,500 rpm for 30 min,

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Amicon Ultra-15; Mn cut-off, 10 KDa). Stock solutions (0.5 mg/mL) of RITC-dex-10K (MW =

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10,000 g/mol) and RITC-dex-70K (MW = 70,000 g/mol) were prepared in PBS. A 30-µL aliquot

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of concentrated C-GFP-NC was mixed with each stock solution (2 µL) and incubated for 3 days

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at RT. The penetration of hydrophilic guest molecules was confirmed by SRCM measurements.

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Each mixed solution was diluted (1:10) with PBS just prior to analysis by SRCM.

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Encapsulation efficiency (EE) and loading capacity of C-GFP-NC. SR101 (25 mg) was

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dissolved in 1 mL PBS. The resulting SR101 solution (2 µL of 25 mg/mL) was added to the

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concentrated C-GFP-NC solution (15 µL of 0.8 mg/mL) and stirred constantly at RT for 3 days.

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A chromatographic column (1.5 × 12 polypropylene Econo-Pac column) packed with 15 mL of

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Sephadex G-10 solution was used to separate C-GFP-NC loaded with dye from any un-loaded

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dye. Briefly, the mixture of dye and C-GFP-NC (10 µL) was loaded onto the column and eluted

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with PBS (pH 7.5, 10 mM). The eluate was collected in 10 fractions. The fluorescence intensity

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of each fraction was measured by PL spectroscopy. The EE, i.e., the amount of loaded


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Biomacromolecules

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dye/combined amount of loaded and un-loaded dye, and LC, i.e., the amount of loaded

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dye/amount of scaffold, were calculated from measurements of the relative fluorescence

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intensities of loaded and un-loaded dye.

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Biocompatibility of C-GFP-NC. Cell viability was estimated in the presence of C-GFP-NC,

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C-GFP-NC and free GFP. Culture media prepared with different concentrations of each GFP

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derivative (0.5, 1.0, 2.5, 5.0, and 1.0 µg/mL) were inoculated with HEK293 cells

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(1.0 × 104 cells/well) in 96-well plates. The plates were then cultured for 24 h in a humidified

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incubator with 5% CO2. After removing the supernatant, each solution was loaded into the wells

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and incubated for 24 h. Cell viability was measured in duplicate using a Celltiter-Glo assay kit

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(Promega, Madison, WI, USA) according to the manufacturer’s instructions. To confirm the

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possibility of C-GFP-NC degradation in a biological environment, a trypsin solution (20 µL of 1

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mg/mL) was added to a solution of C-GFP-NC (2 mL, GFP concentration: 55.6 µg/mL) and

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incubated for 6 h. Changes in the hydrodynamic radii and fluorescence intensity of the particles

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were measured by DLS and PL.

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RESULTS AND DISCUSSION


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Scheme 2. Synthesis of Ni2+-nitrilotriacetic acid-end-functionalized poly(lactic acid) (Ni2+-NTA-

3

PLA).

4 5

This research builds on the polymer-templated protein nanoball (PTPNB) system described in

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previous reports.29-31 The PTPNB system is a size-controllable method of preparing polymer-

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protein core-shell nanoparticles. This system is based on the specific interactions of NTA-Ni2+-

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His with polymer chain ends and multi-histidine tags on proteins. Therefore, proteins

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immobilized on these polymer particles maintain a consistent orientation even after dissociating

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from the polymer particles. Our previous studies introduced PTPNB systems based on Ni2+-NTA

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PS, a non-degradable hydrophobic polymer. In this paper, PLA containing Ni2+NTA chain ends

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(3) were synthesized as shown in Scheme 2 to construct core-degradable PTPNB for preparing

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protein nanocapsules.

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Biomacromolecules

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Figure 1. Proton nuclear magnetic resonance (1H NMR) spectra of t-boc-NTA-PLA (1) and

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NTA-PLA (2). (a) The presence of the t-boc-NTA group is evident on PLA after the coupling

4

reaction. The spectrum in (b) shows that the t-boc group was removed via acidification.


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Acid-terminated PLA was reacted with EDC and NHS for activation, followed by t-boc-NTA-

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NH2 for amidation. The presence of NTA moieties in PLA (1) was verified by 1H NMR (Figure

4

1a). Peaks at 1.44–1.45 ppm (l, k, (CH3)3-), 3.45 ppm (j, -CH2-), 3.28 ppm (i, -CH-), and 3.24

5

ppm (e, -CH2-) were assigned to NTA moieties. Based on the integral ratio of peak j (3.38–3.53

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ppm) to the methyl proton of the PLA chain terminus (d, 1.10–1.11 ppm), the t-boc-NTA end-

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functionality was calculated to be 99%. The chain structure of the resulting polymer was

8

investigated by MALDI-MS. As shown in Figure S1, the m/z value matched that of t-boc-NTA-

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PLA (theoretical m/z of 39mer C145H206N2O88Na+: 3,408, observed: 3,408). The number average

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Mn of acid-terminated PLA and t-boc-NTA-PLA (1) was calculated using the integral ratio of the

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d peak (1.10–1.11 ppm) to methine peaks (b,b´, 5.10-5.30 ppm) in PLA (Mn, NMR=5,270 and

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6,050 g/mol).

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NTA-PLA (2) was obtained by removing the tert-butyl group of (1) with TFA in MC. The

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removal of t-butyl protons in (1) was evidenced by the disappearance of peaks at 1.44–1.45 ppm

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(l, k) in the 1H NMR spectrum (Figure 1b). Comparing SEC traces of acid-terminated PLA, (1)

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and (2) (Figure S2) are indicative of the slight influence the synthetic process had on PLA

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degradation. There were no significant changes in SEC traces before and after the synthetic

18

process. Lastly, NiCl2 was added to a solution of (2) to prepare Ni2+-NTA-PLA (3).

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GFP/PLAs were prepared in accordance with a previously published procedure.29, 30 The

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polymer solution (2 mg/mL) was added to a solution of His6-GFP in PBS. The polymer

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aggregated in water due to hydrophobic interactions and His6-GFP was conjugated with exposed

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Ni2+-NTA on the surface of the PLA particle via NTA-Ni2+/His interactions. The mean diameter


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Biomacromolecules

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of the GFP/PLA particles was 89.6 ± 20.1 nm, measured by DLS (Figure S3a). To confirm that

2

protein activity was not significantly influenced by the particle formation process, changes in

3

GFP fluorescence intensity were determined by comparing PL spectra before and after the

4

removal of free GFP (Figure S3b). The His6-GFP binding efficiency calculated by PL analysis

5

was approximately 71.1% (Figure S3c).

6

7 8

Figure 2. Particle size distribution of cross-linked green fluorescence protein/ poly(lactic acid)

9

(GFP/PLA) (C-GFP/PLA) measured by dynamic light scattering (DLS) and a corresponding

10

transmission electron micrograph (TEM) with negative staining.

11 12

To prevent the destruction of the GFP layer after core degradation, the GFP shell was cross-

13

linked by adding a solution of glutaraldehyde after removing any non-conjugated GFPs from the

14

GFP/PLA solution. Glutaraldehyde reacts with the amine of lysine to form an imine group. The

15

residual aldehyde group and the imine group can be reduced to alcohol and a stable alkyl amine


15


1

with NaBH4. To confirm that protein activity was not significantly influenced by the cross-

2

linking process, changes in GFP fluorescence intensity were determined by comparing by PL

3

spectra acquired before and after the cross-linking reaction (Figure S4). TEM and DLS

4

measurements of C-GFP/PLA showed well-defined spherical structures 89 nm in diameter

5

(Figure 2).

Page 16 of 29

6

7 8

Figure 3. (a) Real-time 1H NMR spectra of oligomeric lactic acid (OLA) (1.65-1.55 ppm) and

9

lactic acid (LA) (1.40-1.50 ppm) methyl protons following enzyme treatment; (b) relative

10

integrated peak area of LA methyl protons versus degradation time over 20 days.

11 12

Degradation of the PLA core for preparing C-GFP-NC was performed with the addition of

13

lipase (Lipozyme TL 100L) to a solution of C-GFP/PLA at 25°C. First, quantitative analyses of

14

PLA degradation from C-GFP/PLA were conducted by 1H NMR spectroscopy. A solution of

15

deuterated PBS was prepared with G-GFP/PLA in an NMR tube. Lipase was added to the tube

16

and 1H NMR spectra were monitored as a function of incubation time for 20 days (Figure 3a).

17

There were no signals corresponding to OLA (1.55–1.65) ppm) or LA (1.40–1.50 ppm) methyl


16

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Biomacromolecules

1

protons on day 0 since the aqueous media did not dissolve the hydrophobic core of C-GFP/PLA.

2

Peaks corresponding to OLA and LA methyl protons became apparent on prolonged incubation

3

with lipase. The LA peak area relative to that of the D2O peak (4.7–4.9 ppm) is shown as a

4

function of incubation time in Figure 3b. The relative area of the LA peak increased to a

5

saturation point at 12 days. This indicates that the PLA core was degraded by lipase regardless of

6

the barrier presented by the C-GFP shell.

7

8 9

Figure 4. Atomic force micrographs (AFM) show (a) changes in the height of C-GFP/PLA

10

features after lipase treatment as a function of degradation time over 28 days and (b) after 4

11

weeks incubation with or without lipase.

12 13

The formation of hollow spheres through core cavitation can be validated by the expansion of

14

hydrodynamic radius despite a decrease in dry state height measured from a substrate.32 To

15

monitor the progress of C-GFP-NC preparation, we used AFM and DLS to track changes in the

16

height and hydrodynamic radius of C-GFP/PLA particles, with or without lipase. Changes in the


17


Page 18 of 29

1

height of C-GFP/PLA structures after lipase treatment were measured by AFM and constitute

2

critical evidence for C-GFP-NC formation (Figure 4a). Prior to lipase treatment, the initial height

3

of C-GFP/PLA was 63.6 ± 8.78 nm. The height decreased as enzymatic hydrolysis progressed.

4

After 28 days, C-GFP/PLA structures that had been treated with lipase were 4.52 ± 1.05 nm tall

5

as a result of core cavitation. In contrast, the height of C-GFP/PLA structures that had not been

6

incubated with lipase was 19.5 ± 3.99 nm after 28 days (Figure 4b). While the average particle

7

height decreased as enzymatic hydrolysis progressed, the DLS data indicate that the average the

8

hydrodynamic radius increased, suggesting swelling of the G-GFP shell. This effect was

9

accelerated with particles that had been treated with lipase (Figure 5a). As swelling of the C-

10

GFP/PLA particles progressed, an unidentified feature, less than 70 nm in diameter, appeared in

11

the multimodal DLS distribution. The size of this feature decreased with degradation time and

12

we speculate that it was formed from assemblies of OLA that had escaped the C-GFP/PLA

13

particles. The concurrent increase in diameter and maintenance of spherical shape was also

14

validated by comparing TEM micrographs with negative staining before and after core

15

degradation (Figure 5b and c). The micrographs show that the core region became darker than

16

the shell region after core removal, suggesting the formation of hollow structures. Final C-GFP-

17

NCs were obtained following the addition of excess imidazole, which served to dissociate the

18

NTA-Ni2+-His interaction and remove residual lipase and hydrolysate (Figure S5a). During

19

washing, GFP that had not been cross-linked with circumferential GFPs was also removed.

20

Comparisons of the fluorescence intensity before and after imidazole washing gave a calculated

21

74.6% yield of C-GFP-NC (Figure S5b). TEM and DLS measurements of C-GFP-NC indicated

22

well-defined spherical structures with hydrodynamic radii over 100 nm (Figure S5c).


18

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Biomacromolecules

1

The size of protein capsules can be controlled by controlling the template size.17 The basic

2

principles of PTPNB structure formation are polymer aggregation via hydrophobic effects in

3

water, and rapid stabilization of PTPNBs by subsequent coating with hydrophilic proteins. The

4

size of a PTPNB can be controlled by adjusting the extent of aggregation.29, 30 Thus, the size of

5

GFP/PLAs can be controlled by adjusting the amount of polymer. Ni2+NTA-PLA solutions in

6

DMF were prepared at different concentrations (1 mg/mL and 3 mg/mL, previous concentration:

7

2 mg/mL). Using the same conditions as described previously for GFP/PLA preparation, each

8

polymer solution was added to a mixture of His6-GFP in PBS. As a result, the average diameter

9

of GFP/PLA particles was controlled to 62.1 ± 11.4 nm or 184 ± 19.4 nm (Figure S6). The

10

GFP/PLA particles were then converted to C-GFP-NCs using the same methods and cross-

11

linking and lipase treatments discussed above. The successful formation of C-GFP-NCs, either

12

smaller or larger than the previously fabricated C-GFP-NCs, was verified by an expansion in

13

hydrodynamic radius in the DLS data despite relatively low dry-state particle heights, as

14

determined by AFM (Figure S7). This result shows that the PTPNB system is suitable for the

15

preparation of protein nanocapsules of various sizes. C-GFP-NCs, prepared from GFP/PLA, with

16

an average diameter of 89.6 nm, were selected for further investigation.

17


19


Page 20 of 29

1 2

Figure 5. (a) Changes in the average hydrodynamic radius, measured by dynamic light scattering

3

(DLS) for C-GFP/PLA particles, with or without lipase as a function of degradation time. (b, c)

4

TEM micrographs of negative-stained C-GFP/PLA particles, (b) before and (c) after core

5

degradation.

6 7


20

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Biomacromolecules

1 2

Figure 6. (a) Changes in the fluorescence intensity of C-GFP/PLA after GFP-NC formation

3

through enzymatic hydrolysis measured by photoluminescence (PL) spectroscopy. (b) Super

4

resolution confocal microscope (SRCM) images of C-GFP-NCs.

5 6

Enzymatic hydrolysis is a soft method for template removal. To verify that protein activity

7

was preserved in our nanocapsules, the fluorescence intensities of C-GFP/PLA and C-GFP-NC

8

were compared after 28 days of lipase treatment. As shown in Figure 6a, lipase treatment did not

9

decrease fluorescence intensity, suggesting that enzymatic hydrolysis did not damage the GFP.

10

Furthermore, SRCM images of C-GFP-NC verified that the spherical nanostructures exhibited

11

the same green fluorescence (Figure 6b).


21


1

Page 22 of 29

The efficiency of protein activity is largely a function of the degree of exposure and

2

orientation of the active site. In this regard, protein assembly into cage-like structures is an

3

effective means of obtaining hollow structures with a consistent protein orientation. Biological

4

approaches often require complex interactions to assign specific intrinsic functions to caged

5

proteins. Despite the importance of protein orientation in nanostructures, prior to this report,

6

there were no synthetic methods that allowed control of protein orientation in nanocapsules.

7

Previous studies employed non-specific absorption of proteins on a template. The primary

8

advantage of using the PTPNB system for nanocapsule preparation is the consistent orientation

9

of proteins via specific NTA-Ni2+-His interactions.

10

11


22

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Biomacromolecules

1

Figure 7. (a) Schematic illustration showing the different capturing phenomena of the Ni2+-NTA

2

agarose resin with hexa-histidine-tagged (His6)-GFP and GFP-NC. (b, c) Fluorescence intensity

3

changes before and after resin filtration of (b) His6-GFP and (c) C-GFP-NC.

4 5

To validate the controlled orientation of GFP in C-GFP-NC, we conducted the capture tests of

6

C-GFP-NC, and free His6-GFP as a control group, with a Ni2+-NTA agarose resin (Figure 7).33

7

We hypothesized that there would be no loss of fluorescence after Ni2+-NTA resin filtration of

8

C-GFP-NC since GFPs in C-GFP-NC are oriented such that the His6-tag is prevented from

9

interacting with the resin (Figure 7a). As expected, free His6-GFP was captured on the Ni2+-NTA

10

resin and exhibited a rapid decrease in fluorescence intensity (Figure 7b). In contrast, C-GFP-NC

11

passed through the Ni2+-NTA resin column without any loss of fluorescence (Figure 7c). These

12

results indicate that His6-tag is located on the interior of C-GFP-NC and that protein orientation

13

inside the structure is controlled.st

14

To characterize the various features of our C-GFP-NCs, we estimated their permeability, EE,

15

and LC. Hydrophilic guest molecules were encapsulated in C-GFP-NCs by first swelling the C-

16

GFP-NCs in a solution containing a target guest molecule. C-GFP-NCs were incubated with

17

RITC-dex-10K or RITC-dex-70K to determine the size range of guest molecules that can be

18

accommodated. Penetration of guest molecules into the C-GFP-NC was confirmed by SRCM

19

after diluting 1/10 with PBS (Figure S8a). The relative fluorescence intensity of RITC-dex in the

20

particle compared to the background fluorescence (Iparticle/Ibackground) was measured using ImageJ

21

(NIH, Bethesda, MD, USA) to determine the relative degree of RITC-dex penetration (Figure

22

S8b). RITC-dex-10K penetrated deeply into the C-GFP-NCs while the degree of penetration of


23


Page 24 of 29

1

RITC-dex-70K was minimal. This suggests that the pore size of our C-GFP-NCs is below 70

2

kDa. EE and LC were calculated by incubating C-GFP-NCs in a dense SR101 solution for 3

3

days. The resulting mixture was separated by gel filtration chromatography and fractions were

4

collected as being SR101-loaded (fractions 2 and 3) or SR101-unloaded (fractions 5 and above)

5

(Figure S9a, b). The EE and LC were calculated as 5.34% and 22.2% by comparing the

6

fluorescence intensities of SR101-loaded and unloaded C-GFP-NCs (Figure S9c).

7

To assess their potential for use in biomedical applications, we examined the biocompatibility

8

and biodegradability of C-GFP-NCs. High concentrations of C-GFP-NC did not exhibit any

9

cytotoxic effects on HEK293 cells (Figure S10). Furthermore, since C-GFP-NCs are composed

10

entirely of protein, they are easily degraded in a biological environment. To confirm this, we

11

treated a solution of C-GFP-NCs with trypsin, which is a type protease, and observed the

12

disappearance of capsule structure and a rapid loss of GFP fluorescence (Figure S11).

13 14

CONCLUSION

15

In summary, protein nanocapsules with preserved activity and defined protein orientation were

16

prepared from the combination of PTPNB and enzymatic hydrolysis. Ni2+-NTA-PLA was

17

synthesized as the enzyme-degradable component of a polymeric template. PLA nanoparticles

18

with oriented His6-GFP were prepared using Ni2+-NTA-PLA. After cross-linking of the GFP

19

shell, C-GFP-NC was obtained through enzymatic hydrolysis of the PLA core.

20 21

Protein nanocapsules prepared using this method meet the general needs of drug delivery vehicles, including bio-compatibility, size-controllability, and high loading capacity. Thus, these


24

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Biomacromolecules

1

nanocapsules can serve as multi-functional containers, without additional modifications,

2

depending on the specific functions of their constituent proteins.

3 4

ASSOCIATED CONTENT

5

Supporting Information.

6

The following data are available free of charge via the Internet at http://pubs.acs.org. SEC traces

7

and MALDI-TOF mass spectra of synthesized polymer; GFP fluorescence changes after reaction

8

and purification; structural analyses of final C-GFP-NCs, including DLS data and TEM and

9

AFM micrographs; SRCM images after incubation with RITC-Dex; calculations of EE and LC;

10

cell viability of C-GFP-NC; changes in diameter and GFP fluorescence after trypsin treatment

11

(file type, PDF)

12

AUTHOR INFORMATION

13

Corresponding Author

14

*[email protected] (H.-j. Paik)

15

Author Contributions

16

The manuscript was written through contributions of all authors. All authors have given approval

17

to the final version of the manuscript.

18

ACKNOWLEDGMENT

19

We thank Mr. Junyong Ahn and Prof. Taihyun Chang in Pohang University of Science and

20

Technology for his help in MALDI-MS analysis of the synthesized polymer. This work was


25


Page 26 of 29

1

supported by Mid-career Researcher Program (2013R1A2A2A01068818) and the Active

2

Polymer Center for Pattern Integration (No. 2007-0056091) through the National Research

3

Foundation (NRF) grant funded by the Korean government (MSIP). This study was also

4

supported by a grant from KRIBB research initiative program. The authors also acknowledge the

5

Korea Basic Science Institute, Seoul center, for assistance with the super resolution confocal

6

microscope analysis.

7

REFERENCES

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from (nitrilotriacetic acid)-end-functionalized polystyrenes and His-tagged proteins. Polym. Chem. 2013, 4, (7), 2286-2292. 30. Lee, C.; Choi, J. E.; Park, G. Y.; Lee, T.; Kim, J.; An, S. S. A.; Song, J. K.; Paik, H.-j., Size-tunable protein–polymer hybrid carrier for cell internalization. React. Funct. Polym. 2018, 124, 72-76. 31. Lee, C.; Jeong, J.; Lee, T.; Zhang, W.; Xu, L.; Choi, J. E.; Park, J. H.; Song, J. K.; Jang, S.; Eom, C.-Y.; Shim, K.; Seong Soo, A. A.; Kang, Y.-S.; Kwak, M.; Jeon, H. J.; Go, J. S.; Suh, Y. D.; Jin, J.-O.; Paik, H.-j., Virus-mimetic polymer nanoparticles displaying hemagglutinin as an adjuvant-free influenza vaccine. Biomaterials 2018, 183, 234-242. 32. Zhang, Y.; Jiang, M.; Zhao, J.; Ren, X.; Chen, D.; Zhang, G., A novel route to thermosensitive polymeric core–shell aggregates and hollow spheres in aqueous media. Adv. Funct. Mater. 2005, 15, (4), 695-699. 33. Zhang, P.; Chen, Y.; Zeng, Y.; Shen, C.; Li, R.; Guo, Z.; Li, S.; Zheng, Q.; Chu, C.; Wang, Z., Virus-mimetic nanovesicles as a versatile antigen-delivery system. Proc. Natl. Acad. Sci. 2015, 112, (45), E6129-E6138.

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Biomacromolecules

Graphical Abstract 88x35mm (300 x 300 DPI)


Orientation Controlled Protein Nanocapsules by Enzymatic Removal

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