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Enzyme-Loaded Polyion Complex Vesicles as in Vivo Nanoreactors Working Sustainably under the Blood Circulation: Characterization and Functional Evaluation Daiki Sueyoshi, Yasutaka Anraku, Toru Komatsu, Yasuteru Urano, and Kazunori Kataoka Biomacromolecules, Just Accepted Manuscript • DOI: 10.1021/acs.biomac.6b01870 • Publication Date (Web): 24 Feb 2017 Downloaded from http://pubs.acs.org on February 25, 2017

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Biomacromolecules is published by the American Chemical Society. 1155 Sixteenth Street N.W., Washington, DC 20036 Published by American Chemical Society. Copyright © American Chemical Society. However, no copyright claim is made to original U.S. Government works, or works produced by employees of any Commonwealth realm Crown government in the course of their duties.

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Enzyme-Loaded Polyion Complex Vesicles as in Vivo Nanoreactors Working Sustainably under the Blood Circulation: Characterization and Functional Evaluation Daiki Sueyoshia,e, Yasutaka Anrakua,e, Toru Komatsub, Yasuteru Uranob,c, and Kazunori Kataokaa,c,d,e* a

Graduate School of Engineering, The University of Tokyo, 7-3-1 Hongo, Bunkyo-ku, Tokyo

113-8656, Japan b

Graduate School of Pharmaceutical Sciences, The University of Tokyo, 7-3-1 Hongo, Bunkyo-

ku, Tokyo 113-0033, Japan c

Graduate School of Medicine, The University of Tokyo, 7-3-1 Hongo, Bunkyo-ku, Tokyo 113-

0033, Japan d

Policy Alternatives Research Institute, The University of Tokyo, 7-3-1 Hongo, Bunkyo-ku,

Tokyo 113-1709 e

Innovation Center of Nanomedicine, Kawasaki Institute of Industrial Promotion, 3-25-14

Tonomachi, Kawasaki-ku, Kawasaki 210-0821, Japan

KEY WORDS: polyion complexes, polymersomes, nanoreactors, enzymes

ACS Paragon Plus Environment

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ABSTRACT Enzyme-loaded synthetic vesicles have attracted great attention for their feasibility to exert the efficient and prolonged functionality of loaded enzymes in harsh environments, such as in vivo. However, several issues remain regarding the optimization of their structures toward practical application. Herein, we fabricated polyion complex vesicles (PICsomes) loaded with Lasparaginase (ASNase@PICsomes), and conducted detailed characterization to ensure their utility as nanoreactors functioning under the harsh in vivo environment of the bloodstream. ASNase@PICsomes showed 100 nm-sized monodispersed vesicular structures. Fluorescence cross-correlation spectroscopy revealed essentially no empty PICsome fraction in the product, indicating the quantitative formation of ASNase@PICsomes. Furthermore, fluorescence anisotropy measurement showed that the loaded enzymes were located essentially in the inner aqueous phase of PICsomes, being successfully segregated from external environment. ASNase@PICsomes exhibited significantly prolonged enzymatic reaction compared with free ASNase after systemic injection into mice, corroborating their functionality as in vivo nanoreactors working under the blood circulation.

INTRODUCTION The compartmentalization of fragile functional substances into synthetic vesicular structures has attracted much attention due to its potential to preserve or even enhance the functionality of such substances.1-5 The compartments prevent interference from external molecules or environments, thus, protecting the substances to retain their inherent performance. A typical example of such encapsulated substances is catalysts, including enzymes, thus forming mesoscopic vesicular reactors. Compartmentalization is also a fundamental facet of living cells or subcellular


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organelles in eukaryotic cells, which are composed of a phospholipid bilayer with amphiphilic nature. Inspired by these vesicles composed of natural amphiphilic molecules, synthetic vesicles consisting of amphiphilic block copolymers, termed polymersomes, have been intensively studied for their relatively thick and robust membrane structures.6-8 In this regard, polymersomes enclosing catalytic species such as enzymes are prospective candidates for nano-/microreactors or artificial organelles working under the harsh biological conditions, such as in the blood stream or in the cellular interior.9-11 However, the vesicular walls of polymersomes have limited permeability,12 leading to the restricted flux of substances inward and outward, which is an essential function for vesicular reactors. Therefore, various strategies have been utilized in the design of vesicles with suitable permeability, typically by installing pore-forming components in the vesicle wall by embedding channel proteins13-15 or integrating stimuli-responsive polymers to gate the wall in response to external stimuli.16-18 An alternative approach to construct vesicular nanoreactors with the appropriate molecular exchange capability is the design of a vesicular membrane with intrinsic semipermeability. Worth noting in this regard is, polyion complex (PIC) vesicles (PICsomes) formed through electrostatic interaction-mediated self-assembly in aqueous media between oppositely charged block ionomers with polyethylene glycol (PEG) and homo ionomers.19,20 The PICsome membrane, consisting of a PIC layer sandwiched by two PEG layers, intrinsically exhibits semipermeability for hydrophilic molecules. Moreover, the encapsulation process of guest molecules into PICsomes is facilitated by utilizing a unique dynamic property of PICsomes in an aqueous milieu, i.e., a reversible dissociation / re-association behavior in response to mechanical stimuli exerted by vortex stirring.21 This process is advantageous for encapsulating molecules with various properties after hollow PICsome formation, leading to the successful formation of


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enzyme-loaded PICsomes as nanoreactors with preserved enzymatic activities in in vitro / in vivo environments.22,23

However, there are several issues remaining regarding the loading capacity and location of enzymes in PICsomes. It is important to adequately characterize the states of the loaded enzymes to ensure their functionality as nanoreactors. Herein, we conduct a detailed examination of the location of enzymes in the vesicle by a spectroscopic methodology. Moreover, a further investigation was implemented to gain insight into the loading capacity and the activity of the loaded enzymes.

We adopted L-asparaginase (ASNase) as the model enzyme to load into PICsomes. ASNase is a therapeutically relevant enzyme that exhibits anti-proliferative effects on asparagine synthetasedeficient cancer cells, such as leukemic cells, by hydrolyzing L-asparagine in the circulation.24 As the ultimate site in which this enzyme performs its function is the blood compartment, it can be a suitable model to demonstrate the functionality of enzyme-loaded PICsomes in such a harsh environment as the bloodstream. Therefore, the catalytic performance of intravenously injected ASNase-loaded PICsomes (ASNase@PICsomes) was examined by analyzing the L-asparagine level in the circulation, demonstrating the utility of ASNase-loaded PICsomes as therapeutically relevant in vivo nanoreactors.

EXPERIMENTAL Materials. β-benzyl-L-aspartate N-carboxy-anhydride (BLA-NCA) was purchased from Chuo Kaseihin Co. Inc. (Tokyo, Japan). α-methoxy-ω-amino poly(ethylene glycol) (MeO-PEG-NH2)


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(Mn = 2,000; Mw/Mn = 1.05) was purchased from Nippon Oil and Fats Co. Ltd. (Tokyo, Japan). 1,5-Diaminopentane (DAP) was purchased from Tokyo Chemical Industry Co. Ltd. (Tokyo, Japan). N-methyl-2-pyrrolidone (NMP) was purchased from Wako Pure Chemical Industries, Ltd. (Osaka, Japan). Dichloromethane (CH2Cl2) was purchased from Kanto Chemical Co, Inc. (Tokyo, Japan). DAP, NMP and CH2Cl2 were used after conventional distillation. DyLight488 NHS ester was purchased from Thermo Fisher Scientific Inc. (Waltham, MA, USA). Cy5monofunctional reactive dye pack was purchased from GE Healthcare (Waukesha, WI, USA). Dimethyl sulfoxide (DMSO) was purchased from Nacalai Tesque, Inc. (Kyoto, Japan). PEGpoly(α,β-aspartic acid) (PEG-b-P(Asp); Mn of PEG: 2000, DP of P(Asp): 75), poly([5aminopentyl]-α,β-asoartamide) (P(Asp-AP); DP: 82), and fluorescence-labeled PEG-b-P(Asp) were prepared according to a previously reported procedure.19,20,25 ASNase (Mn ~ 141,000) was purchased

from

Kyowa

Hakko

Kirin

Co.,

Ltd.

(Tokyo,

Japan).

1-Ethyl-3-(3-

dimethylaminopropyl)carbodiimide hydrochloride (EDC) was purchased from Tokyo Chemical Industry Co. Ltd. (Tokyo, Japan). Glycerol and Dulbecco’s phosphate-buffered saline (PBS) were purchased from Wako Pure Chemical Industries, Ltd. (Osaka, Japan). L-Aspartic acid β-(7amido-4-methylcoumarin) (Asp-AMC) and 7-amino-4-methylcoumarin were purchased from Sigma-Aldrich Co. LLC. (St. Louis, MO, USA). Balb/c mice (female, 5 weeks old) were purchased from Charles Liver Laboratories Japan, Inc. (Kanagawa, Japan). All animal experiments were conducted according to the guidelines for animal experiments at the University of Tokyo. Ammonium sulfate was purchased from Sigma-Aldrich Co. LLC. (St. Louis, MO, USA). L-Asparagine monohydrate was purchased from Wako Pure Chemical Industries, Ltd. (Osaka, Japan). L-Serine-2,3,3-d3 (d3-serine) was purchased from C/D/N Isotopes Inc. (PointClaire, Quebec, Canada). Acetonitrile was purchased from Nacalai Tesque, Inc. (Kyoto, Japan).


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Dansyl chloride and methylamine hydrochloride were purchased from Tokyo Chemical Industry Co. Ltd. (Tokyo, Japan). Preparation of fluorescence-labeled ASNase. DyLight488-NHS ester or Cy5 monoreactive dye pack dissolved in DMSO was mixed with 10 mM phosphate buffer (PB; pH 7.4) solution of ASNase (MW = 141,000, pI = 4.7 ± 1) in a volume ratio of 1:10, and reacted at 25˚C for 4 h. The resulting solution was purified by centrifugal ultrafiltration using a poly(ether sulfone) ultrafiltration membrane with a molecular weight cut-off (MWCO) of 10,000, with substituting the solvent to 10 mM PB (pH 7.4). The concentration of enzymes and fluorescent dyes conjugated to the enzymes in the purified solution was determined, respectively, using a Spectrophotometer (NanoDrop 1000, Thermo Fisher Scientific Inc., Waltham, MA, USA) by measuring the absorbance at 280 nm and the excitation wavelength of each fluorescent dye. Accordingly, two to three fluorescent dye molecules were estimated to be conjugated per enzyme molecule. Preparation of ASNase@PICsomes. PEG-b-P(Asp) and P(Asp-AP) were dissolved, respectively, in 10 mM PB (pH 7.4) to obtain a 1 mg/mL polymer solution. The solution was mixed to obtain an equal charge ratio of -COO- and -NH3+ units in the side chains of each ionomer and then vortexed to obtain PICsomes.20 To this solution, ASNase (or, in specified experiments, DyLight488-/Cy5-labeled ASNase) in 10 mM PB (pH 7.4) was added with the concentration of 2 mg/mL, followed by the vortex mixing to obtain PICsome loaded with ASNase (ASNase@PICsome). Subsequently, for cross-linking the PIC membrane, 10 mg/mL EDC in 10 mM PB (pH 7.4) (10 eq. to -COOH groups in the block aniomer PEG-b-P(Asp)) was added and reacted at 4˚C overnight. The resulting solution was purified by ultrafiltration using a membrane with MWCO of 300,000 to remove the residual EDC and ASNase, obtaining purified


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ASNase@PICsomes. During the ultrafiltration, the solvent was substituted with PBS (for in vivo studies), ultrapure water (for TEM and fluorescence anisotropy measurements), or 10 mM PB (pH 7.4; for other studies). Fluorescence-labeled PICsomes (Dy488-PICsomes) were prepared using 1 mg/mL PEG-b-P(Asp)-DyLight488 in 10 mM PB (pH 7.4) mixed with PEG-b-P(Asp) in the same buffer (1:39 v/v for FCCS measurements and 1:4 for fluorescence anisotropy measurements, respectively). Dynamic light scattering (DLS) measurements. The size and corresponding distribution of the PICsomes were evaluated by dynamic light scattering (DLS). DLS measurements were conducted at 25˚C using a Nano-ZS instrument (Malvern Instruments, Malvern, Worcestershire, UK) equipped with a diode-pumped solid state laser (532 nm) or a He-Ne laser (633 nm). The intensity-averaged hydrodynamic diameter and PDI were derived according to the cumulant method as described previously.20 Transmission electron microscopy (TEM). The morphology of ASNase@PICsomes was observed using transmission electron microscopy (TEM). First, the surface of copper grids (400 mesh) was subjected to hydrophilic treatment by plasma irradiation in vacuum. Then, 2 µL of the sample solution in ultrapure water was placed onto the grid, subsequently stained by dropping 2 µL of 50% v/v ethanol solution containing 2% wt uranyl acetate, followed by drying under room temperature. TEM measurements were conducted using a transmission electron microscope JEM-1400 (JEOL Ltd., Tokyo, Japan) operating at the acceleration voltage of 120 kV. Cryo-TEM observation. Cryo-TEM observation was performed to clearly verify the vesicular structure of ASNase@PICsomes with hollow inner spaces.

A 3 µL aliquot of

ASNase@PICsomes in ulrapurewater was placed onto a copper grid. The grid containing the


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sample solution was blotted onto a paper to remove excess sample solution in a Leica EM GP plunge freezer (Leica microsystems, Wetzlar, Germany). The resulting grid was immediately shock-frozen by prompt immersion into liquid ethane near its solidifying point. The grid bearing the frozen sample was placed into liquid ethane near its solidifying point. The grid bearing the frozen sample was placed into a TEM system (JEM-2100F, JEOL, Tokyo, Japan) using a cryotransfer holder (G-914, Gatan, Pleasanton, CA, USA) with a temperature maintained below 170˚C throughout the measurement. Fluorescence correlation spectroscopy (FCS). DyLight488-labeled ASNase (Dy488ASNase)-loaded PICsomes were analyzed by the fluorescence correlation spectroscopy (FCS). The FCS measurements were done using a confocal laser scanning microscopy (LSM 510 META/Confocor 3, Carl Zeiss AG, Jena, Germany) equipped with an Ar laser (488 nm) and a He/Ne laser (633 nm). The sample solution was placed in an eight-well Lab-Tech chambered cover glass (Nunc, Rochester, NY, USA), which was placed on a water-immersion objective (Zeiss C/Apochromat 40×). With Zeiss Confocor3 software, the fluctuation of the fluorescence signal was analyzed based on the autocorrelation function, and the diffusion time (average residence time in the confocal volume) of the fluorescent molecules was obtained by fitting the one-component three-dimensional free diffusion model. Additionally, the counts per molecule were obtained from the value of count rate and the number of fluorescent molecules for estimation of the number of loaded enzyme molecules per vesicle. Fluorescence anisotropy measurements of loaded enzymes. To obtain insight into the location of enzymes, fluorescence anisotropy measurements were performed, which reflect the rotational Brownian motion of fluorescent molecules. An aqueous solution of Dy488ASNase@PICsomes was placed into a 10 mm × 10 mm quartz cell. Fluorescence anisotropy r


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was measured at Ex/Em = 493/518 nm, using a fluorescence spectrophotometer FP8600 (JEOL Ltd., Tokyo, Japan), installed with autopolarizing filter units. The r value is the ratio of the intensity of the polarized fluorescence to that of the whole fluorescence, which can be obtained from equation (1):

r=

IVV − GIVH IVV + 2GIVH

G=

••• (1)

I HV I HH

where I indicates fluorescence intensity, with inferiors V and H representing vertical and horizontal placement of the excitation and emission polarizers, respectively. G is the instrumental function

which denotes relative instrumental sensitivity to emitted lights which

are vertically and horizontally polarized. In vitro enzyme activity assay of ASNase@PICsomes. L-Aspartic acid β-(7-amido-4methylcoumarin) (Asp-AMC) is used as a substrate of ASNase to liberate the fluorescent product of 7-amino-4-methylcoumarin (AMC) (Ex/Em = 350/450).26 Asp-AMC dissolved in PBS was mixed with a PBS solution of ASNase@PICsome at 37˚C, and the fluorescence intensity was measured for a defined time using a multiplate reader (Infinite M1000 PRO, Tecan Group Ltd., Männedorf, Switzerland). Here, by using a standard curve of AMC, the fluorescence intensity measured at defined time points was converted to the produced AMC concentration to obtain the reaction rate V, which was then plotted against various substrate concentrations [S] (MichaelisMenten plot). Additionally, the reciprocal of V was plotted against the corresponding reciprocal of [S] (Lineweaver-Burk plot), followed by fitting equation (2) to obtain the Michaelis-Menten constant Km:


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1 Km 1 1 = ⋅ + V Vmax [ S ] Vmax

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••• (2)

where Vmax is the maximum reaction rate for a specified [S]. Furthermore, using the same substrate, Asp-AMC, the changes in enzymatic units per certain volume of solution of ASNase@PICsomes over time were evaluated at 37˚C in the presence of 10% FBS. Here, one unit denotes the amount of enzyme that liberates 1 µmol of substrate at 37˚C per min. Evaluation of blood circulating property of ASNase@PICsomes. Cy5-ASNase@PICsomes and free Cy5-ASNase in PBS were intravenously injected into mice (Balb/c, female, 5 weeks old) and blood was collected from the postcaval vein using a heparinized syringe at defined time points (n = 5). The obtained blood was centrifuged at 4˚C at 15,000 rpm for 2 min and supernatant was collected. The fluorescence intensity of the obtained plasma was measured using a multiplate reader (Ex/Em = 650/670) and the residual amount of the labeled enzymes was evaluated. The half-life in blood was estimated according to equation (3)27: Half life = -0.693 × (t2 - t1) / [ln(c(t2)) - ln(c(t1))]

••• (3)

where t1 represents time 0, t2 represents 6 h for free ASNase and 24 h for ASNase@PICsome, respectively (because these were the maximum time points in the examined range at which remaining portion of injected samples was well above the detection limit), c(t1) and c(t2) denote remaining fraction of the labeled enzyme in the plasma at time 0 and t2, respectively. Evaluation of in vivo enzymatic reactions of ASNase@PICsomes in the circulation. ASNase@PICsome or free ASNase in PBS was intravenously injected into mice (Balb/c, female, 5 weeks old) with the dose of 100 unit/kg body weight (units against L-asparagine; adjusted using the units against Asp-AMC), which corresponded to approximately 60 mg polymer/kg


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body weight for ASNase@PICsome. After a defined time, blood was collected from the postcaval vein using a heparinized syringe (n = 3). The collected blood was centrifuged, and supernatant was collected for quantification of plasma component levels as described in the following procedure. Quantification of the plasma ammonia level. The plasma ammonia-nitrogen level was measured using a high throughput automatic clinical chemistry analyzer (Fuji Dri-Chem 7000i, Fujifilm, Tokyo, Japan), based on the coloration reaction of bromophenol blue appended in the Dri-Chem Slide (NH3-P II, Fujifilm, Tokyo, Japan) with ammonia. The obtained plasma was mixed with PBS (3:1 v/v), and 10 µL of the mixture was placed on a Dri-Chem Slide. The measured ammonia-nitrogen level was calibrated using a standard curve made by mixture of mouse plasma and standard ammonium sulfate in PBS (3:1 v/v). Quantification of the plasma amino acid level. The plasma L-asparagine (Asn) level was quantified by liquid chromatography/mass spectrometry (LC/MS) analysis, with using d3-serine (d3-Ser) as an internal standard. The obtained plasma was deproteinized using acetonitrile with 1% formic acid and Sirocco protein precipitation plate (Waters Co., Milford, MA, USA) as follows. First, d3-Ser and Asn were dissolved in a mixture of PBS and ultrapure water (1:9 v/v) to prepare two types of standard solution (Sol. 1: d3-Ser 600 µM, Sol. 2: d3-Ser, Asn 600 µM each). Then, 225 µL of acetonitrile with 1% formic acid was placed into a well of Sirocco protein precipitation plate and subsequently mixed with 25 µL of Sol. 1 or Sol. 2, respectively. Then, 50 µL of the plasma was added to each well, and vortexed for 1 min for precipitation of the plasma proteins, with a plastic lid fixed tightly on the plate. The resulting mixture in the well was filtered into a collection plate set in a manifold with the plate, by employing a vacuum pump for 3 min. A 50 µL aliquot of the resulting filtrate was added to 200 µL of 0.75 M sodium borate


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buffer (pH 9.5) and subsequently mixed with 50 µL of 5 mM dansyl chloride in acetonitrile and reacted at room temperature for 30 min for the dansylation of amino acids. The reaction was terminated by adding 25 µL of 1% methylamine hydrochloride. To the resulting mixture was added 10% formic acid in acetonitrile (1:1 v/v), which was placed into LC/MS for analysis. The LC/MS measurements were conducted using an octadecylsilyl silica column (InertSustain C18, 2.1 × 250 mm, GL Sciences Inc., Tokyo, Japan) and a LC/MS system (LC: 1260 Infinity, MS: 6130 Quadrupole LC/MS; Agilent Technologies, Inc, Santa Clara, CA, USA), under a neutral pH eluent condition with an increasing gradient of acetonitrile (eluent: 0.01 M ammonium formate in ultrapure water, and mixture of acetonitrile and water (4:1 v/v) with 0.01 M ammonium formate). The plasma Asn level CAsn (µM) was calculated using the formula (4):

CAsn =

AAsn,1 ⋅ DF ( AAsn,2 − AAsn,1 ) 50

AAsn,1 =

AAsn,1 Ad3 −Ser,1

AAsn,2 =

AAsn,2 Ad3 −Ser,2

••• (4)

where Ai,1 and Ai,2 (i = Asn, d3-Ser) are areas of the peak attributed to the dansylated amino acid i in the treated plasma sample mixed with Sol. 1 or Sol. 2, respectively, and DF is the dilution factor of the plasma sample due to the mixing of acetonitrile and Sol. 1 or Sol. 2.

RESULTS AND DISCUSSION Preparation and characterization of ASNase-loaded PICsomes (ASNase@PICsomes). ASNase@PICsomes were fabricated using the characteristic dynamic properties of PICsomes,


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i.e., the reversible fragmentation / reassembly behavior in response to mechanical stress imparted by vortex stirring in the solution.21 Briefly, solutions of PEG-b-P(Asp) (Mn of PEG: 2000, DP of P(Asp): 75) and P(Asp-AP) (DP of P(Asp-AP): 82) were mixed at charge-stoichiometric condition and vortexed to form hollow PICsomes via electrostatic interaction-mediated selfassembly. Then, ASNase was added to the solution and loaded in PICsomes through their reversible fragmentation / reassembly processes synchronized with on-off switching of vortex stirring. PICsomes undergo fragmentation by vortex stirring to form unit PICs, mainly single pairs of a PIC whose charge balance is neutral, which can subsequently reassemble into vesicles after the removal of the mechanical stress from vortex stirring. ASNase in the solution can be spontaneously loaded into PICsomes through this reassembly process of uPIC. Then, 10 mol equivalence of EDC to -COOH groups of the PEG-b-P(Asp) was added and incubated overnight for sufficient cross-linking of the PIC membrane, endowing PICsomes with high stability under physiological conditions. Note that as reported in our previous paper23, the EDC treatment seems to have negligible influence on the catalytic ability of encapsulated enzymes in PICsomes, presumably PIC membrane may act as the protection layer for inner enzyme from the possible attack by EDC. Residual EDC and unencapsulated ASNase were removed from the solution by ultrafiltration to obtain purified ASNase@PICsome sample. Removal of the unencapsulated ASNase was confirmed by size exclusion chromatography with UV detector set at 220 nm to monitor ASNase and PIC constituent polymers (Figure S1). Notably, the size of the PICsomes can be modulated just by changing the initial concentration of constituent polymers20 Here, the condition to form 100 nm-sized PICsomes, i.e., polymer solutions of 1 mg/mL each, was adopted due to their relatively low tissue disposition as well as longevity in blood circulation after systemic administration.25 Actually, narrowly-distributed (polydispersity index (PDI) = 0.01)


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particles with approximately 100 nm in size were obtained even in the presence of ASNase as determined by DLS measurements (Figure 1A). Furthermore, TEM and Cryo-TEM measurements revealed that the particles have characteristics of vesicular structures with hollow inner spaces (Figure 1B and Figure 1C, respectively), indicating the successful formation of enzyme-loaded PICsomes.

Figure 1. (A) A size histogram of ASNase@PICsomes derived from a DLS measurement. (B) A TEM image of ASNase@PICsomes stained with uranyl acetate. (C) A Cryo-TEM image of ASNase@PICsomes.

To confirm the loading of ASNase in PICsomes, fluorescence correlation spectroscopy (FCS) measurements were applied to DyLight488-labeled ASNase (Dy488-ASNase) and Dy488ASNase-loaded PICsomes (Dy488-ASNase@PICsomes) solutions. FCS is a technique used to examine the translational diffusivity and the number of fluorescent molecules, analyzing the fluctuation of fluorescence signals originating from fluorescent molecules going in and out of the optical confocal volume via an autocorrelation function.28 The normalized autocorrelation curve of fluorescent molecules loaded into mesoscopic-sized vehicles, such as PICsome, exhibits a


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shift to a longer decay time region from that of free fluorescent molecules due to an increase in the diffusion time of the fluorescent molecules after loading.20,29 Actually, Dy488ASNase@PICsomes exhibited an apparent shift in the autocorrelation curve, which reflects a significant increase in the diffusion time relative to free Dy488-ASNase (Figure 2, Table 1). This result indicates that the diffusivity of the free enzyme was lowered after the loading procedure, confirming that the enzymes were successfully loaded in larger structures, i.e., PICsomes. Another feature of the FCS method is that the number of loaded proteins per vesicle can be estimated by comparing derived counts per molecule (which correspond to fluorescence intensity per fluorescent molecule) of free proteins to those loaded in the vesicles.29 Following this methodology, the number of Dy488-ASNase loaded per PICsome was estimated. Derived counts per molecule were 99.4 kHz for Dy488-ASNase@PICsome and 49.2 kHz for free Dy488ASNase, suggesting that approximately 2 molecules of enzymes were loaded in a single PICsome.

Figure 2. FCS normalized autocorrelation curves of (i) free Dy488-ASNase (dashed line) and (ii) Dy488-ASNase@PICsome (solid line).


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Table 1. FCS fitting results of free Dy488-ASNase and Dy488-ASNase@PICsomes.

Sample

Diffusion Time / µs

Counts per molecule / kHz

free Dy488-ASNase

(2.29 ± 0.01) × 102

49.2

Dy488-ASNase @PICsome

(7.97 ± 0.05) × 103

99.4

This estimated number of enzymes per vesicle is based on the assumption that all the PICsomes are loaded with at least one enzyme molecule, but it is not excluded the possibility that some fraction of the PICsomes was loaded with no enzymes. To investigate this possibility, fluorescence cross-correlation spectroscopy (FCCS) measurements were performed. In FCCS, two laser lines simultaneously excite dyes with corresponding wavelengths in overlapped confocal volumes, and concurrent fluctuation of the fluorescence signal in separate channel is analyzed using a cross-correlation function.30,31 Here, the degree to which either of the two types of dyes is associated with the other type of the dye can be represented as relative crosscorrelation amplitude (RCA)32, denoting the fraction of those associated dyes normalized by either type of the dye [Supporting information]. FCCS measurements were applied to Cy5labeled ASNase (Cy5-ASNase) loaded in PICsomes prepared in part with DyLight488-labeled PEG-b-P(Asp) (Dy488-PICsomes). As a result, the RCA of Cy5-ASNase@Dy488-PICsomes was derived as 〜0.91 (Figure S2). Given that confocal volumes of the two excitation lights are spatially identical, this result indicates that 91% of the Dy488-PICsomes were loaded with at least one molecule of Cy5-ASNase, confirming only limited formation of empty PICsomes. Note that a significantly low RCA (~0.019) was observed for a mixing solution of Dy488-PICsome


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and free Cy5-ASNase, suggesting that false positive cross-correlation due to cross-talk in the two types of fluorescent channels in the FCCS measurement was negligibly low (Figure S3). Investigation on the location of loaded enzymes in PICsome. Although the FCS results indicated an association of ASNase with PICsomes, the possibility cannot be excluded that the enzymes are embedded in the PIC membrane, not loaded in the inner aqueous phase of PICsomes. To investigate this issue, fluorescence anisotropy (FA) measurements were employed, which reflect the rotational diffusion of fluorescent molecules. FA measurement is a technique used to examine the rotational behavior of fluorescent molecules by exciting them with polarized excitation light and detecting the degree of polarization of the emitted fluorescence.33,34 FA observed by steady excitation light ( r ) is in the theoretical range of 0 ~ 0.4, exhibiting a lower value as the rotational Brownian motion of the fluorescent molecules is vigorous. Applying an equivalent hard sphere model to the fluorescent molecule in the Perrin equation (5), equation (6) is obtained:

1 1 τ = 1+  r r0  θ  =

1 τ T 1+ kB ⋅ ⋅  r0  v η

••• (5)

••• (6)

where r0 is the initial fluorescence anisotropy, τ is the time constant of the fluorescence decay, θ is the time constant of the fluorescence anisotropy decay, kB is the Boltzmann constant, v is the volume of the equivalent hard sphere, T is the absolute temperature, and η is the solvent viscosity. Herein, the FA measurement was applied to free Dy488-ASNase and Dy488ASNase@PICsome. It can be expected that a relatively low r is observed if the enzymes are floated in the inner aqueous phase, whereas a significantly high r is observed if the enzymes are


17

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embedded in the PIC membrane (Scheme 1). Cross-linked PICsome containing Dy488-labeled PEG-b-P(Asp) in their membrane structure (Dy488-PICsome) was utilized as a control, resembling the situation in which enzymes are embedded in the PIC membrane (Scheme 1). Note that in the FA measurement, the effect of light scattering due to particles in the sample solution should be considered, which may affect the observed value of r . To examine this effect, free Dy488-ASNase solution was mixed with a hollow non-labeled PICsome solution. As a result, the effect of light scattering from PICsomes on the value of r was negligibly small in the concentration range of PICsomes employed in this study (Figure S4).

Scheme 1. Schematic illustration of assumed thermal rotational behaviors of (i) free Dy488ASNase, (ii) Dy488-ASNase@PICsome (floating in the inner aqueous phase), (iii) Dy488ASNase@PICsome (embedded in the membrane), and (iv) Dy488-PICsome as a control resembling the fluorophore microenvironment of (iii).

In pure water as the solvent, Dy488-ASNase loaded in PICsome (Dy488-ASNase@PICsome) exhibited r = 0.198, whereas Dy488-PICsome (PICsome with Dy488 labeling in their crosslinked membrane structure) exhibited r = 0.342 (Figure 3). This result indicates that the enzymes loaded in PICsomes have relatively free rotational motion compared with PEG-b-


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P(Asp) in cross-linked PIC membranes, suggesting that the enzymes maintain their rotational diffusion even after their loading into the PICsomes. However, the observed r of Dy488ASNase@PICsome ( r = 0.198) was slightly higher than that of free Dy488-ASNase ( r = 0.122). This result raises two possibilities: (i) enzymes are present both in the inner aqueous phase and in the membrane where rotational motion is highly restricted; and (ii) loaded enzymes are indeed floating in the inner aqueous phase, yet are in part susceptible to local viscosity increase in the inner cavity due to the presence of inner PEG layer of PICsome. To get insight into this issue, changes in r of samples was examined after increasing the solvent viscosity of inner aqueous space by adding glycerol (MW = 92.09) in the solution. Note that MW of glycerol is low enough to freely penetrate the PICsome membrane. According to equation (6), r increases with η. Thus, it can be reasonably assumed that r of the enzyme molecules floating in the inner aqueous cavity may increase with an elevation in the local viscosity, yet r of those embedded in the PICsome membrane may be insensitive to a viscosity change. In the 90% glycerol condition at 25˚C with viscosity of 163.6 mPa•s,35 r of Dy488-ASNase@PICsome revealed significant increase to a level comparable to that of free ASNase in the same condition (Figure 3). This is consistent with the possibility (ii) that ASNase molecules float in the inner aqueous phase, undergoing a restricted motion due to the increased local viscosity in the presence of glycerol. If some fraction of enzyme molecules is embedded in the PIC membrane as in the case with the possibility (i), Dy488-ASNase@PICsome should have higher r than free Dy488-ASNase even in 90% glycerol because the control Dy488-PICsome has significantly higher r value as seen in Figure 3. However, as aforementioned, this is not the case: they have comparable r value, consequently excluding the possibility of enzyme immobilization in the


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membrane structure. Eventually, it is safe to conclude that the loaded Dy488-ASNase is essentially present in the inner aqueous phase of PICsomes.

Figure 3. Steady-state fluorescence anisotropy ( r ) of (i) free Dy488-ASNase, (ii) Dy488ASNase@PICsome, and (iii) Dy488-PICsome with or without the presence of 90% glycerol (Ex/Em = 493/518 nm).

This complete segregation of enzymes into the inner cavity of PICsomes is worthwhile in terms of retaining their activity and avoiding immune responses in harsh in vivo conditions. It should also be noted that to our knowledge, there have been no reports clearly demonstrating the locations of enzymes loaded in mesoscopic-scaled synthetic vesicles. Thus, the present methodology based on FA measurements can be a useful characterization technique to clarify the location of loaded payloads in various nanocompartment systems. Enzymatic activity assay of ASNase@PICsomes. Enzymatic activity was investigated to verify functional aspects of ASNase@PICsomes. Here, as a substrate to examine enzymatic activity of ASNase, L-aspartic acid β-(7-amido-4-methylcoumarin) (Asp-AMC) was utilized.


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Asp-AMC is hydrolyzed by ASNase to liberate 7-amino-4-methylcoumarine (AMC), which emits fluorescence (Ex/Em = 350/450).26 A kinetic assay was conducted to evaluate the reaction between Asp-AMC and ASNase@PICsome in PBS at the physiological temperature of 37˚C. Eventually, the Michaelis-Menten constant (Km) to this substrate was derived as 194 µM for ASNase@PICsomes (Figure 4), which was comparable to 186 µM for free enzyme (Figure S5). This result indicates that the enzymatic activity of ASNase is well preserved even after the loading into PICsomes.

Figure 4. (A) A Michaelis-Menten plot and (B) a Lineweaver-Burk plot for hydrolysis of AspAMC by ASNase@PICsome at 37˚C.


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Many of enzymes are known to lose their activity under the physiological temperature of 37˚C due to gradual thermal denaturation.36-38 Furthermore, in the case of vesicular nanocarriers for enzymes, protein adsorption from biological milieu often results in the aggregation of nanocarriers, leading to deterioration of their biomedical functions. Therefore, the stability and the enzymatic activity of ASNase@PICsomes under a pseudo-physiological condition were examined for their feasibility as in vivo nanoreactors. ASNase@PICsomes were observed to retain their enzymatic units per unit volume of solution for 24 h even under 10% fetal bovine serum (FBS) at 37˚C (Figure S6). Additionally, the size and PDI obtained via DLS measurement were almost constant for ASNase@PICsome (Figure S6) over 24 h, indicating no vigorous interaction with serum proteins to undergo aggregation. In vivo blood circulation profiles of ASNase@PICsomes. To explore the utility of ASNase@PICsome as in vivo nanoreactor working in the blood compartment, their blood circulating ability was examined. Free Cy5-ASNase and Cy5-ASNase@PICsome were respectively injected into the tail vein of mice, and blood was collected at defined time points. Fluorescence intensity (Ex/Em = 650/670 nm) of the obtained plasma sample from the collected blood was measured to determine the residual amount of Cy5-ASNase in the sample. Eventually, almost all of the free Cy5-ASNase disappeared 24 h after intravenous injection into mice, whereas ca. 35% of Cy5-ASNase@PICsomes still remained (Figure 5). The half-life in blood estimated from the semi-logarithmic plot was 16.0 h for Cy5-ASNase@PICsomes and 2.53 h for free Cy5-ASNase, respectively. The half-life of free Cy5-ASNase obtained here is comparable to the literature value so far reported (~2 h).39 Furthermore, the half-life of systemically injected hollow PICsomes with the size of 102 nm was reported to be 16.6 h for tumor-inoculated mice.25


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Thus, blood circulation profiles of PICsomes are not impaired by the loading of ASNase, supporting the result that ASNase molecules are segregated in the inner aqueous phase of PICsomes.

Figure 5. Blood circulation profiles of (i) free Cy5-ASNase (open circles) and (ii) Cy5ASNase@PICsome. Each sample was injected into tail vein of BALB/c mice (female, 5 weeks old). At defined time points, blood was collected from their postcaval veins with heparinized syringe, followed by the centrifugation to obtain plasma sample. Relative residual amount of Cy5-ASNase in each formulation was determined by measuring fluorescence intensity of the obtained plasma samples (Ex/Em = 650/670 nm).

Evaluation of in vivo enzymatic reaction of ASNase@PICsomes under circulating blood. Both retained enzymatic activity and longevity in blood circulation of ASNase@PICsomes support their feasibility as in vivo nanoreactor working under circulating blood. To reveal this feasibility, change in the plasma level of reactant (L-asparagine; Asn) and reaction product


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(ammonia)

was

determined

after

intravenous

administration

Page 24 of 31

of

free

ASNase

and

ASNase@PICsome. Their dose was set at 100 unit/kg body weight for Asn. Blood was collected at defined time points, and obtained plasma was analyzed to quantify ammonia-nitrogen and Asn level. Ammonia-nitrogen was quantified by using a clinical dry-chemistry analyzer. The ammonianitrogen level 24 h after administration in mice treated with free ASNase was comparable to that of non-treated mice, whereas mice treated with ASNase@PICsomes showed significantly elevated ammonia-nitrogen level (Figure 6). This result is consistent with sustained enzymatic reaction by long-circulating ASNase@PICsomes in blood compartment. Then, the plasma level of Asn, the substrate for ASNase, was determined. The plasma Asn level was determined by quantitative LC/MS, after deproteinization of the obtained plasma and derivatization with dansyl chloride. As shown in Figure 7, Asn level returned to the level of non-treatment control for the mice treated with free ASNase 12 h after administration. In contrast, mice treated with ASNase@PICsome maintained significantly lowered plasma Asn level compared to the control even 24 h after administration (Figure 7).


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Figure 6. Ammonia-nitrogen concentration in plasma of the mice, 24 h after receiving intravenous administration of (i) free ASNase or (ii) ASNase@PICsome with the dose of 100 unit/kg body weight.

Figure 7. Time profiling of L-asparagine (Asn) concentration in plasma of the mice receiving intravenous administration of (i) free ASNase or (ii) ASNase@PICsome with the dose of 100 unit/kg body weight.

These results indicate that ASNase@PICsomes exhibit prolonged enzymatic reaction based on their longevity in blood circulation compared with free ASNase. Thus, the present study is the first to demonstrate the functionality of enzyme-loaded vesicles as nanoreactors in the circulating blood in a sustainable manner.

CONCLUSIONS Our results highlight the functionality of ASNase@PICsomes, compartmentalized by PIC membranes with semipermeable properties for water-soluble molecules, as sustainable enzymatic


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nanoreactors under harsh intravital conditions such as the circulating blood in vivo. ASNase was successfully encapsulated in PICsomes with high efficiency in a way that most of the formed PICsomes were loaded with enzyme molecules. The loaded enzymes were located in their inner aqueous phase, that is, the inside of the compartmentalized space, as examined by fluorescence anisotropy measurements. ASNase@PICsomes retained their enzymatic activity and exhibited sustained enzymatic reaction after systemic injection into mice due to their prolonged blood circulation compared with free ASNase. This system of PICsome-based enzymatic nanoreactors can address the issues of therapeutic exogenous proteins upon their use in intravital environments, such as rapid clearance, induction of immunological responses, and deterioration in enzymatic activity, opening a new avenue in enzyme therapeutics for various diseases requiring long-term efficacy and reduced side effects.

ASSOCIATED CONTENT Supporting Information Figures showing the purity of ASNase@PICsomes; FCCS results; the effect of light scattering in fluorescence anisotropy measurements; kinetic analysis of hydrolysis reaction of Asp-AMC by free ANase; the retention of enzymatic activity of ASNase@PICsomes under pseudophysiological condition. (PDF)

AUTHOR INFORMATION Corresponding Author *E-mail: [email protected]. Phone: +81-44-589-5700.


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Author Contributions The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript. Notes The authors declare no competing financial interest.

ACKNOWLEDGMENTS This work was supported in part by a Grant-in-Aid for Scientific Research (No. 15K12536 to Y.A.) from the Ministry of Education, Culture, Sports, Science, and Technology (MEXT) of Japan, Grant-in-Aid for Specially Promoted Research (No. 25000006 to K.K.) of the Japan society for the Promotion of Science (JSPS), and the Center of Innovation (COI) Program, Japan Science and Technology Agency (JST), Japan. D.S. thanks the Research Fellowships of JSPS and Graduate Program for Leaders in Life Innovation (GPLLI), the University of Tokyo. TEM measurements were conducted at the Research Hub for Advanced Nano Characterization at the University of Tokyo, with valuable help from Mr. H. Hoshi. We thank Dr. T. Shimada, Ms. S. Ogura, and Ms. A. Miyoshi for their assistance in animal experiments.

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