Protein Nanoparticle Formation Using a Circularly Permuted α-Helix

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Protein nanoparticle formation using a circularly permuted ##-helix-rich trimeric protein Norifumi Kawakami, Hiroki Kondo, Masayuki Muramatsu, and Kenji Miyamoto Bioconjugate Chem., Just Accepted Manuscript • DOI: 10.1021/acs.bioconjchem.6b00735 • Publication Date (Web): 16 Jan 2017 Downloaded from http://pubs.acs.org on January 17, 2017

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Protein nanoparticle formation using a circularly permuted α-helixrich trimeric protein Norifumi Kawakami*, Hiroki Kondo, Masayuki Muramatsu and Kenji Miyamoto* Department of Bioscience and Informatics, Faculty of Science and Technology, Keio University, 3-14-1 Hiyoshi, Kohokuku, Yokohama, Kanagawa 223-8522, Japan.

ABSTRACT: We here report the production of highly spherical protein nanoparticles based on the domain-swapping oligomerization of a circularly permuted trimeric protein, major histocompatibility complex (MHC) class II-associated chaperonin. The size distribution of the nanoparticles can be adjusted to between 40-100 nm in diameter and thus these particles are suitable as drug carriers following purification under basic conditions. Our approach involves no harsh treatments and could provide an alternative approach for protein nanoparticle formation.

Protein nano-cages and nanoparticles are promising molecular materials for drug delivery systems owing to their biocompatibility and biodegradability.1, 2 Protein cages such as ferritin, a 24-mer iron storage protein, and viral capsids have been extensively studied as molecular capsules incorporating drugs in their inner space.3-8 In parallel with these studies, artificially designed protein supramolecules also provide caged proteins with novel architectures and could be used as drug carriers.9-12 However, disassembly and reassembly of these caged proteins is often required for incorporation of small molecules including the drug into the cages, reducing the yield of capsule formation13-16. Protein nanoparticles produced by random protein association with or without denaturation of the template protein provide alternative drug carriers.17-19 Such protein nanoparticles can entrap drug molecules during particle formation. The process of forming nanoparticles uses safer and milder conditions than previously reported methods, but still requires harsh treatments such as heat and organic solvent treatments for purification or destabilization of the template protein to initiate aggregation.20-23 High temperature and organic solvents can spontaneously degrade the drug molecule and result in unwanted incorporation of solvent in the particles, respectively. Heat and solvent treatments might appear to be critical, depending on the nanoparticle formation mechanism. However, we anticipated that a protein designed to have a “destabilized structure” might interact with identical protein molecules for stabilization and formation of nanoparticles, without the need for any destabilization treatment. To examine this possibility, we focused on a domain swapping mechanism that can produce oligomeric proteins from a monomeric protein.24, 25 Domain-swapped oligomers can be produced experimentally by placing the protein under specific conditions, such as by acidification of the sample or by the addition of organic solvents.26-29 Circular permutation (CP) is another approach for inducing artificial domain swapping. CP, found in nature, now become a gene manipulation technique

that connects the original N- and C-termini of a protein with an arbitrary linker peptide and produces new N- and C-termini by cutting at a desired position in the sequence.30, 31 CP can thus rearrange the relative domain positions of the template protein. Most proteins subjected to CP maintain the original structure but some monomeric proteins change to dimeric or trimeric proteins, suggesting that CP destabilizes the monomeric state, resulting in formation of oligomeric proteins.32 We hypothesized that if we subjected natural oligomeric proteins to CP, the destabilized protomers might interact with multiple protomers to form large aggregates resembling nanoparticles. To test this hypothesis, we explored template proteins fulfilling the following criteria: i) The 3D structure is known. ii) The template protein forms a trimeric structure, which is the smallest odd number oligomer. We avoided oligomeric proteins with a even-fold symmetry axis, such as dimers and tetramers, which easily produce stable domainswapped structures similar to the structure of the original oligomeric state. iii) The protein has a long flexible random coil structure at the original N- or C-terminus that can be altered by CP into a long loop structure to avoid formation of the original structure, thus simultaneously facilitating domainswapping between multiple protein molecules. We selected the trimeric domain of major histocompatibility complex (MHC) class II-associated chaperonin (trimeric domain, PDB: 1IIE).33 The protein monomer is composed of one long and two short α-helices and has a long flexible random coil structure at the C-terminus. Here we report the effect of CP on the aggregation of the template protein. Figure 1A and 1B shows the original protein structure and the circularly permuted protein model structure, respectively. We first purified native trimeric domain to confirm the oligomeric state of the wild-type protein. The protein was purified by anion exchange chromatography, followed by Niaffinity chromatography. The purified protein was concentrated to more than 2 mg/mL and subjected to Blue Native-PAGE (BN-PAGE) to separate oligomeric proteins without dissocia-

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tion. A smeared band was obtained corresponding to a molecular weight of less than 66 kDa (Figure 2A). We also tested the effect of organic solvent on protein aggregation by adding 5 vol% ethanol to the purified protein prior to lyophilization, followed by analysis of the lyophilized protein by BN-PAGE. No molecular weight shift was observed, indicating that the trimer structure was stable under these conditions (Figure S1).

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change between proteins. Indeed, the protein expressed in Escherichia coli cells was found in the insoluble fraction, indicating accelerated protein-protein interaction. Insoluble CPH6G4 was solubilized using 6 M urea pH 8.0 and purified by DEAE-Sepharose and Ni-Sepharose column chromatography.

Figure 1. The structure of template protein A) from PDB 1IIE, and a manually drawn model of the protein after being subjected to CP B). The positions of the new N- and C-termini were determined to be residue numbers 159 and 158 (scissors in right panel of A)), respectively, and the original N- and Ctermini were connected by a H6G4 peptide linker (circle in right panel of B)).

Figure 2. Electrophoretic analyses of the wild-type protein and CP-H6G4. A): BN-PAGE result of the wild-type protein. B): BN-PAGE result of CP-H6G4 purified at pH 8.0, and two dimensional SDS-PAGE analysis of the CP-H6G4 sample after separation by BN-PAGE. The long arrow shows the orientation of the gel placed on the second acrylamide gel. Next, the template protein was subjected to CP. The positions of the new N- and C-termini were residue numbers 159 and 158, respectively, corresponding to a position between two short α-helices (Figure 1B). Peptide sequences containing 6 to 11 amino acids (H6G0-5) were used as a linker to connect the original N- and C- termini. Proteins with four glycine residues in the linker showed the highest expression level and thus H6G4 peptide was used as the linker sequence. The designed protein (CP-H6G4) has a greater distance between the two short helices than the wild-type protein and thus should exhibit a higher frequency of helix ex-

Figure 3. TEM image of CP-H6G4 purified at pH 8.0. The sample was stained with 2% uranyl acetate. Panels A) and B) show images of the non-aggregated and aggregated parts of the sample, respectively. Panel C) shows size distributions of the spherical structures calculated based on the TEM images (N = 1325). The urea was removed before eluting the proteins bound on the Ni-Sepharose column. Dialysis could not be used to remove urea because a visible precipitate formed. The eluted

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Figure 4. Protein purified at pH 9.8 was analyzed by BN-PAGE and TEM. A): Time course of the aggregation of CP-H6G4 purified at pH 9.8 and analyzed by BN-PAGE. The numbers above each lane show the incubation time after dissolving the lyophilized protein. B) and C): TEM images of the protein after incubation for 24 h. Panels B) and C) show the non-aggregated and aggregated parts of the sample, respectively. Inset in the panel B) shows a magnified image of one particle. D): The size distribution of the spherical structures, calculated based on the TEM images (N = 1036). The particles were slightly larger than particles receiving the protein was immediately concentrated to more than 2 mg/mL by ultrafiltration, followed by separation using BN-PAGE most attention as drug carriers (less than 100 nm). Making the (Figure 2B). In contrast to the wild-type protein, the bands for pH basic prior to treatment with organic solvent decreased the CP-H6G4 appeared at positions corresponding to molecular size of protein nanoparticles when human serum albumin was weights of 200 kDa and 400 kDa. A smeared band extending used as the template protein21. We therefore purified CP-H6G4 from 500 kDa to over 1000 kDa was also observed. Twounder basic pH conditions. CP-H6G4 was purified in one step dimensional SDS-PAGE of these bands showed that all conusing a Ni-column at pH 9.8 in the presence of 6 M urea. After sisted of CP-H6G4 (Figure 2B). This strongly indicated that the removal of the urea, the protein was lyophilized to concentrate application of CP to the trimeric domain sequence facilitated the sample for BN-PAGE analysis. Since protein aggregation protein aggregation. Note that no aggregated band was obis dependent on the incubation time at neutral pH conditions, served when the wild-type protein was refolded using 6 M urea we expected that aggregation would increase with time at basic (Figure S2A). The bands at 200 kDa and 400 kDa suggested pH conditions. The lyophilized protein samples were incubated that this oligomerization process was not random. These bands for various times after dissolving in MilliQ water at room temdisappeared after incubation for 24 h at room temperature, perature and then separated by BN-PAGE. Protein separated indicating that protein aggregation is dependent on incubation immediately after dissolving in water provided smeared bands time. The sample after incubation for 24 h was visualized by with a molecular weight corresponding to less than 150 kDa, negatively stained transmission electron microscopy (TEM) whereas the sample incubated for 24 h showed a band with a using uranyl acetate. Interestingly, large numbers of highly high molecular weight distribution at around 1000 kDa (Figure 4A). Interestingly, no 200 or 400 kDa bands observed in samspherical protein aggregates were observed (Figure 3A and B). ples treated at neutral pH conditions were observed for any The average flattening ratio of the particles was 0.07 ± 0.06. incubation time, indicating that different intermediates were The distribution of the major axis length ranged from 60 to formed by protein-protein association under basic conditions. 400 nm, as directly measured from the TEM image (Figure Extending the incubation time to 72 h gave a visible precipi3C).

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tate, but diluting the protein concentration to below 0.02 mg/mL at 24 h incubation almost completely prevented further aggregation even after incubation for 72 h (Figure S3). Despite differences in the band patterns between the samples exposed to neutral and basic conditions, the TEM image of the latter sample stained with uranyl acetate showed spherical structures similar to that observed under neutral pH conditions (Figure 4B and C) but with a lower size distribution (from 40-100 nm; Figure 4D). Note that we also purified wild-type protein at pH 9.8 and confirmed the effect of urea, but no band at 1000 kDa position was observed (Figure S2B and C). The spherical shape of protein aggregates at two different pH conditions likely reflects a structural feature of the template protein. Ogihara et al. reported that the topological rearrangement of the α-helices in a coiled-coil protein consisting of three α-helices results in formation of a protein fibril by domain swapping.34 The template was the monomeric protein and therefore the rearranged monomer simply provides one αhelix to the second monomer, and the second monomer provides an α-helix to the third monomer to form a fibril structure (a one-dimensional extension). In contrast, our template protein was a trimeric protein. Subjecting this trimeric protein to CP would induce domain swapping at multiple positions, allowing extension to a spherical shape (three dimensional extension). The decreased size of the spherical structures at basic pH suggests that an increase in the negatively charged area of the particle surface might prevent further aggregation. We found that the addition of cationic metal ions to the protein after 24 h incubation at basic pH resulted in protein precipitation detected as the remained band at the entrance of the gel by BNPAGE analysis (Figure S4), providing further support that the protein sphere has a negatively charged surface. The results also implied that the aggregated parts of the protein observed in TEM analysis (Figure 4C) would be experimental artifact because no detectable band at the entrance of the gel is observed in the sample without metal ion treatment (Figure S4). In conclusion, we found that subjecting trimeric MHC class II chaperonin to CP resulted in the production of spherical particles, indicating that CP facilitated interaction between multiple protein molecules due to their flexible loop sequence which prevented adoption of the original trimeric structure. The shape of the aggregates remained unchanged at basic and neutral pH but the size and size distribution become smaller and narrower, respectively, at basic pH. In contrast to conventional methods for generating protein nanoparticles, our method does not require any harsh treatment at either the purification or particle formation step. Furthermore, the spherical aggregates remained stable for at least 3 days after purification. We, therefore believe that this approach may be an alternative method for creating protein nanoparticles.

ASSOCIATED CONTENT Supporting Information The Supporting Information is available free of charge on the ACS Publications website. (1) Procedures of protein purification, electrophoresis, mutagenesis, and TEM analysis. (2) Primer list. (3) BN-PAGE data confirming the effect of ethanol on aggregations. (4) BN-PAGE data checking the long-term stability of the spherical aggregates. (PDF)

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AUTHOR INFORMATION Corresponding Author * E-mail: [email protected] * E-mail: [email protected]

Notes The authors declare no competing financial interest.

ACKNOWLEDGMENT The authors wish to acknowledge Division for Medical Research Engineering, Nagoya University Graduate School of Medicine, for technical support of TEM observation. This work was supported by Keio Gijuku Academic Development Funds to N. K. from Keio University.

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