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Biomineralized Metal-Organic Framework Nanoparticles Enable Intracellular Delivery and Endo-Lysosomal Release of Native Active Proteins Ting-Ting Chen, Jin-Tao Yi, Yan-Yan Zhao, and Xia Chu J. Am. Chem. Soc., Just Accepted Manuscript • DOI: 10.1021/jacs.8b04457 • Publication Date (Web): 14 Jul 2018 Downloaded from http://pubs.acs.org on July 14, 2018

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Journal of the American Chemical Society

Biomineralized Metal-Organic Framework Nanoparticles Enable Intracellular Delivery and Endo-Lysosomal Release of Native Active Proteins Ting-Ting Chen, Jin-Tao Yi, Yan-Yan Zhao, and Xia Chu* State Key Laboratory of Chemo/Bio-Sensing and Chemometrics, College of Chemistry and Chemical Engineering, Hunan University, Changsha, 410082, P. R. China

ABSTRACT: Efficient delivery and endo-lysosomal release of active proteins in living cells remain a challenge in proteinbased theranostics. We report a novel protein delivery platform using protein-encapsulated biomineralized metalorganic framework (MOF) nanoparticles (NPs). This platform introduces an adapted biomimetic mineralization method for facile synthesis of MOF NPs with high protein encapsulation efficiency and a new polymer coating strategy to confer the NPs with long-term stability. In vitro results show that protein-encapsulating MOF NPs have the advantages of preserving protein activity for months and protecting proteins from enzyme-mediated degradation. Live cell studies reveal that MOF NPs enable rapid cellular uptake, efficient release and escape of proteins from endo-lysosomes, and preservation of protein activity in living cells. Moreover, the developed platform is demonstrated to enable easy encapsulation of multiple proteins in single MOF NPs for efficient protein co-delivery. To our knowledge, it is the first time that protein-encapsulating MOF NPs have been developed as a generally applicable strategy for intracellular delivery of native active proteins. The developed proteinencapsulated biomineralized MOF NPs can provide a valuable platform for protein-based theranostic applications.

INTRODUCTION Proteins are essential players of diverse cellular processes, and protein therapy, which delivers proteins into the cell to replace dysfunctional proteins, has provided a promising approach to drug development for various diseases, such as 1 cancer, inflammation, and lysosomal storage diseases. Intracellular delivery of proteins also represents useful strategies 2 3 for cellular imaging and diagnosis, genome engineering, 4 and synthetic biology. However, native proteins are mostly membrane impermeable, and are prone to undergo degrada5 tion by proteolytic enzymes in cells, especially in lysosomes. Carrier systems for efficient delivery of functional proteins into cells is crucial to advance protein-based theranostics. Current protein delivery systems typically rely on genetic protein fusions with membrane permeable tags such as pro6 7 tein transduction domains or supercharged proteins, and protein-encapsulating nanocarriers using such as cationic

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polymers and inorganic nanoparticles liposomes, 13,14 (NPs). However, the genetic fusion systems are not applicable to native, non-fused proteins, and exhibit varied efficiency for different protein types or susceptibility to lysosomal protein degradation. The nanocarrier systems used to require complicated synthesis or covalent protein modification, and be lacking of efficiency in protein loading, releasing 15 and endo-lysosomal escape. Development of novel platforms that enable efficient delivery and endo-lysosomal escape of native proteins in their active conformation to desired intracellular sites still present a great challenge. Metal-organic frameworks (MOFs), a class of highly crystalline inorganic−organic hybrids constructing by bridging 16 metal ions or clusters with organic ligands, have emerged as valuable materials for diverse applications, including gas 17 18 19 storage, catalysis, and separation. Because of their finely tunable chemical composition, pore shape and size, morphology and biodegradability, MOFs are able to afford a promising platform in encapsulation and delivery of imaging 20,21 22-24 agents and chemical drugs. MOFs have also demonstrated potential as nanocarrier systems for nucleic acid 25,26 27 therapeutics and RNA−protein CRISPR complex. Because of the strong interactions between nucleic acid backbones and metal ions, the MOFs based nucleic acid carriers 25,26 probably form surface-adsorbed complexes, leading to limited loading efficiency or possible degradation of cargos by exterior surroundings. MOFs have also been reported to 28-32 encapsulate proteins via a de novo approach or biomimet33 ic mineralization. This encapsulation structure can enhance the activities of proteins and improve their stability against 33 various denaturing conditions. However, to our knowledge, MOFs have not been explored as a general strategy for encapsulation and intracellular delivery of proteins. Many issues such as facile synthesis of nanoscale proteinencapsulating MOFs, stability of MOF NPs in cell media, and intracellular delivery properties of MOF NPs remain elusive. Here we report the development of biomineralized protein-encapsulated MOF NPs as a novel platform for efficient intracellular delivery and endo-lysosomal release of protein, as illustrated in Scheme 1. The key hypothesis to our design is that the recently reported biomimetic mineralization proce33 dure can be adapted for synthesis of nanoscale proteinencapsulating MOFs and the MOF NPs can be stable in cell

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Scheme 1. Illustration of biomineralized MOF NPs for protein delivery in living cells.

media, as nanocarriers are documented to have better effi20-26 ciency for cellular uptake. As a proof for our design, we choose a low-cytotoxicity and biodegradable MOF material, zeolitic imidazolate framework-8 (ZIF-8), which forms by 2+ 34coordination between Zn and 2-methylimidazole (MIM). 37 By finely controlling the concentration ratio of MIM to 2+ Zn , we introduce a facile one-pot biomimetic mineralization method for synthesizing nanoscale MOFs encapsulated with high-loading proteins. This method is generally applicable to encapsulating different proteins with very high efficiency in no need of preliminary protein modification, affording an advantageous platform over current protein delivery systems. Unlike current MOFs based 25,26 nanocarrier systems for nucleic acids or RNA−protein 27 complex with a probable surface-adsorbed structure, the MOF NPs are shown to carry the proteins via a definite encapsulation mechanism. Further-more, we develop a simple procedure by modifying the MOF NPs with a biocompatible polyvinylpyrrolidone (PVP) coat-ing that affords excellent stability to the NPs in cell media for months. To our knowledge, it is the first time that MOF NPs have been synthesized to confer long-term stability in cell media, providing new possibilities of using these MOF NPs for biomedical applications. The resulting MOF NPs are also shown to have the advantages of preserving protein activity and protecting proteins from enzyme-mediated degradation. Moreover, we demonstrate that MOF NPs enable rapid cellular uptake, efficient release and escape of proteins from endo-lysosomes, and preservation of protein activity in living cells. Additionally, the facile one-pot synthesis approach has shown the ability to encapsulate multiple proteins in single MOF NPs, affording a useful strategy for protein co-delivery. To our knowledge, it is the first time that proteinencapsulating MOF NPs have been demonstrated as a generally applicable strategy for protein delivery in living cells. Therefore, the protein-encapsulated biomineralized MOF NPs may represent a valuable platform for protein delivery and protein-based theranostics.

RESULTS AND DISCUSSION Synthesis and characterization of biomineralized MOF NPs. Previous studies revealed that proteins efficiently

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induced formation of microscale MOFs via biomimetic min33 eralization. We hypothesized that protein-mediated biomimetic mineralization process were dominated by coordi2+ nation reaction between MIM and Zn , while proteins merely had effect on forming prenucleated clusters with ligands and metal ions. Motivated by the hypothesis, we considered that the size of protein-mediated biomineralized ZIF-8 was dominantly controlled by the concentrations of MIM and 2+ Zn . Accordingly, by using a high concentration ratio of 2+ MIM to Zn , which were reported to generate nanoscale ZIF38 8, we could adapt the biomimetic mineralization method for synthesis of protein-encapsulated ZIF-8 NPs. We started the synthesis with a model protein, bovine serum albumin (BSA). In a typical biomimetic mineralization synthesis, BSA (0.5 mg) was incubated with 0.9 mL aqueous solution of 3.15 o mmol MIM for 10 min at 30 C followed by addition and incubation for 10 min with 0.1 mL aqueous solution of 0.045 mmol zinc nitrate. After centrifugation, washing and resuspension, the resulting biomineralized BSA@ZIF-8 particles were characterized using transmission electron microscope (TEM) and dynamic light scattering (DLS). As anticipated, nanoscale monodisperse BSA@ZIF-8 particles with an average size of 92±7.9 nm were obtained in the biomimetic mineralization synthesis (Figure 1a). In the absence of BSA, the pure ZIF-8 NPs merely had an average size of 53±3.1 nm (Figure 1b). A closer interrogation revealed that the size of BSA@ZIF-8 NPs was remarkably dependent upon the BSA concentration (Figure S1 in SI). With a fixed concentration 2+ ratio of MIM to Zn at 70:1, BSA@ZIF-8 NPs gave increased sizes of 53±3.1, 60±8.0, 92±7.9 and 138±13.5 nm for increasing amounts of BSA (0, 0.25, 0.50 and 0.75 mg). The increased sizes of BSA@ZIF-8 NPs with increasing BSA amounts might be attributed to a facilitated aggregative growth kinetics mediated by BSA-seeded clusters, which allowed the formation of large crystals. Moreover, the concentration ratio of MIM to 2+ Zn also had a crucial effect on the size of BSA@ZIF-8 NPs 2+ (Figure S2 in SI). For concentration ratios of MIM to Zn at 40:1 and 70:1, the average sizes of BSA@ZIF-8 NPs were 150±13.6 and 92±7.9 nm, respectively. A higher concentration 2+ ratio of MIM to Zn at 100:1 produced BSA@ZIF-8 NPs with obvious polydispersity and aggregation, which was attributed to the formation of too small NPs to generate aggregates. The decreased size for protein-encapsulating MOFs at a higher 2+ concentration ratio of MIM to Zn was attributed to faster nucleation at the higher concentration ratio such that a large number of nuclei were produced and crystal growth stage was shortened. Taken together, these results demonstrated 2+ that, by using a high concentration ratio of MIM to Zn , the biomimetic mineralization method enabled facile and consistent synthesis of nanoscale protein-encapsulated ZIF-8 particles. Because BSA@ZIF-8 NPs synthesized using 0.5 mg 2+ BSA with 70:1 concentration ratio of MIM to Zn had a size of 92±7.9 nm, which was ideal for intracellular delivery, it was then chosen for subsequent studies. To ascertain that the biomimetic mineralization synthesis indeed produced BSA-encapsulated NPs, the as-prepared BSA@ZIF-8 NPs were washed with SDS, which was known to 33 remove surface bound proteins from the NPs, followed by Fourier transform infrared spectroscopy (FTIR) analysis (Figure 1c). Absorption bands characteristic of BSA were still -1 observed in the ranges from ~1,640 to 1,660 cm and from


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~1,510 to 1,560 cm , ascribed to the amide I and II bands, respectively, for the BSA@ZIF-8 NPs. In contrast, FTIR spectrum from the control sample prepared by mixing preformed pure ZIF-8 crystals and BSA followed by wash using SDS did not gave these two characteristic bands. These data testified that BSA protein was indeed encapsulated in the BSA@ZIF-8 NPs instead of being adsorbed on the surface of NPs. To further confirm the encapsulation mechanism, we then utilized the biomimetic mineralization method to synthesize ZIF-8 NPs encapsulated with ferritin, a protein allowing direct visualization in TEM (Figure S3 in SI). As expected, ZIF-8 NPs with average sizes of 64±4.6 and 198±13.6 nm were obtained with 1.0 mg and 3.5 mg ferritin, respectively, and encapsulation of multiple ferritin molecules in the NPs were observed. Moreover, increased ferritin loadings were obtained with increasing ferritin amounts. This result gave clear evidence for protein encapsulation in the ZIF-8 NPs.

Figure 1. Characterization of biomineralized MOF NPs. (a) TEM image of BSA@ZIF-8; (b) TEM image of ZIF-8; (c) FTIR spectra of BSA, ZIF-8, BSA@ZIF-8, ZIF-8 with BSA absorbed (ZIF-8/BSA) and ZIF-8 with BSA absorbed washed with SDS (ZIF-8/BSA/SDS); (d) PXRD patterns of simulated ZIF-8, synthetic BSA@ZIF-8 and ZIF-8; (e) NAA isotherms of ZIF-8 and BSA@ZIF-8; (f) TGA of ZIF-8 and BSA@ZIF-8. Further characterization of BSA@ZIF-8 NPs were performed using powder X-ray diffraction (PXRD), nitrogen adsorption analysis (NAA), and thermal gravity analysis (TGA). The PXRD data confirmed that BSA@ZIF-8 NPs retained the same crystalline form as the pure ZIF-8 NPs (Figure 1d), suggesting that a minor portion of proteins did not alter the crystalline structure of ZIF-8. NAA isotherms was best described as Type 1 in shape and BET analysis of the 2 data gave a surface area of 728 m /g for BSA@ZIF-8 NPs, which was much smaller than that for pure ZIF-8 NPs (1442 2 m /g) (Figure 1e). This data was consistent with the proteinencapsulation structure for BSA@ZIF-8 NPs. The TGA data

gave an estimate of the loading capacity as ~52.2 μg/mg for encapsulated BSA protein (Figure 1f), i.e. ~52.2 μg BSA encapsulated in 1 mg BSA@ZIF-8 NPs. According to the total amount of BSA@ZIF-8 NPs (~8.7 mg) obtained, the protein encapsulation efficiency, the relative amount of protein encapsulated inside BSA@ZIF-8 NPs to the protein amount initially provided in the reaction mixture for synthesis of BSA@ZIF-8 NPs, was calculated to be ~91%. UV-vis absorption analysis also verified the high protein encapsulation efficiency (~93%) (Figure S4 in SI). This encapsulation efficiency was much higher than that reported for nonbiomineralized synthesis of MOF NPs carrying RNA−protein 27 complex (~17%), indicating distinct mechanisms in the two synthetic methods. This high encapsulation efficiency also implied a salient advantage of the biomineralized method in cases of encapsulating costly or hard-to-available proteins. The biomimetic mineralization synthesis of proteinencapsulating MOF NPs is primarily facilitated by affinity of protein molecules toward imidazole ligands due to intermolecular hydrogen bonding and hydrophobic 33 interactions. The affinity is generally applicable to different proteins with no need of preliminary protein modification, affording an intrinsic advantage over current protein delivery systems. Moreover, the same synthetic procedure for different protein offers the possibility of co-encapsulating multiple cargos in single ZIF-8 NPs for protein co-delivery. Hence, we explored the ability of the biomimetic mineralization approach for synthesizing ZIF-8 NPs encapsulated with different proteins as well as coencapsulated with multiple proteins (Figure S5 in SI). It was found that with different proteins, such as enhanced green fluorescence protein (EGFP), β-galactosidase (β-Gal), caspase 3/HSA, and BSA/red fluorescence protein (RFP)/β-Gal, ZIF-8 NPs were obtained with good monodispersity, only the size exhibiting dependency on the proteins. Presumably, prenucleated clusters around the proteins were different from varying proteins, which affected the corresponding aggregative growth kinetics and thus controlled the size of ZIF-8 NPs. Long-term stability, protein release and activity preservation enabled by PVP-coated ZIF-8 NPs. To enable the protein-encapsulated ZIF-8 NPs to be applicable in intracellular delivery, stability of the protein-encapsulated NPs in cell media is a major concern. In our initial studies, the as-prepared BSA@ZIF-8 NPs from the biomimetic mineralization synthesis exhibited poor stability, forming polydisperse aggregates in few hours. We ascribed aggregation of the BSA@ZIF-8 NPs to strong affinity between surfaceexposed ligands or metal ions on the MOFs NPs. We envisioned that a polymer coating isolating individual MOFs NPs had the potential to prevent aggregation of the NPs. Hence, we selected PVP for the coating because of its excellent bio28,29 compatibility and reported affinity to ZIF-8. The PVPcoated BSA@ZIF-8 NPs were synthesized readily by adding PVP of a given amount in the aqueous solution of BSA@ZIF8 NPs followed by incubation for 30 min. TEM images showed that a clear coating layer of 10±1.6 nm thickness formed on each individual BSA@ZIF-8 NPs, and the coating did not alter the size of the core BSA@ZIF-8 NPs (Figure 2a). DLS and zeta potential analysis also manifested formation of the PVP-coated BSA@ZIF-8 NPs (Figure S6 in SI). TGA data showed that the amount of PVP coating was ~127.6 μg in 1



mg PVP-coated BSA@ZIF-8 NPs. N2 adsorption analysis revealed that the pore size decreased from 5.0 nm for BSA@ZIF-8 NPs to 2.9 nm for PVP-coated BSA@ZIF-8, indicating a certain degree of blockage or penetration of PVP in the pores (Figure S6 in SI). Though the coating process is simple, long-term stability for the PVP-coated BSA@ZIF-8 NPs were observed over three-month storage in a buffer as well as a cell medium supplemented with 10% fetal bovine serum, no aggregation and no appreciable size change occurring for the BSA@ZIF-8 NPs (Figure S7 in SI). This result demonstrated that PVP coating afforded an effective approach to prevent aggregation and decomposition of ZIF-8 NPs. Because long-term stability conferred better control of efficiency and reproducibility for protein delivery, our polymer-coating design indeed provided new possibilities of using MOF NPs for biomedical applications. An essential requirement for protein delivery system is to specific release of protein cargos in cells. Nanocarrier systems used to enter cells via an endocytic pathway, which were destined to localize in acidic organelles such as endosomes or lysosomes. Therefore, pH-responsive release of protein from the PVP-coated BSA@ZIF-8 NPs was investigated. Western blot assay of the PVP-coated BSA@ZIF-8 NPs incubated in water and buffers of different pH values showed that there was no release of BSA at pH 7.4 and in water, while acidic environments (pH 5.5) induced substantial release of BSA within 0.5 h and complete release in 2 h (Figure S8 in SI). Furthermore, we used the PVP-coated ZIF-8 NPs encapsulated with EGFP to validate the release (Figure 2b). The time-dependent fluorescence profiles of EGFP released from the PVP-coated EGFP@ZIF-8 NPs showed that complete release of proteins was achieved in 2 h at pH 5.5, slower release up to 5 h appeared at pH 6.0, and no release occurred at pH 7.4. These results demonstrated that ZIF-8 NPs allowed efficient release of encapsulated protein cargos under physiological acidic environments, a desirable property for protein delivery systems.

Figure 2. (a) TEM image of PVP coated BSA@ZIF-8; (b) EGFP release from PVP-coated EGFP@ZIF-8; (c) Protection of ZIF-8 encapsulated proteins. EGFP (1), EGFP + proteases (2), EGFP@ZIF-8 + proteases (3), EGFP released from EGFP@ZIF-8 (4), FDG (5), FDG + β-Gal (6), FDG + β-Gal treated by proteases (7), FDG + β-Gal released from βGal@ZIF-8 treated by proteases (8); (d) Long-term activity of β-Gal and β-Gal in β-Gal@ZIF-8.

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A unique advantage of biomineralized ZIF-8 NPs is protection of encapsulated proteins against its external environment, as reported for other microscale protein28-33 To demonstrate the protective encapsulating MOFs. properties, we chose NPs encapsulated with EGFP and β-Gal in the study (Figure 2c). It was found that the fluorescence signal for EGFP-encapsulated ZIF-8 NPs incubated with a protease mixture of trypsin and α-chymostrypsin merely showed negligible decrease, while the unencapsulated protein gave much lower fluorescence. This finding verified the ability of encapsulation in ZIF-8 NPs to protect the proteins from protease-mediated degradation. After EGFP was released from ZIF-8 NPs via acidic decomposition, the fluorescence signal also did not show appreciable change, indicating that encapsulation and release did not alter the fluorescence properties of EGFP. Similar finding was obtained with β-Gal. After digestion with the protease mixture, the unencapsulated enzyme only showed slight activity in catalyzing hydrolysis of its fluorescence substrate, C12-fluorescein di-3-Dgalactopyranoside (FDG). In contrast, after β-Galencapsulated ZIF-8 NPs digested using the protease mixture followed by release from the ZIF-8 NPs, the enzyme retained nearly the same activity as that for the intact enzyme before encapsulation. Taken together, these results verified that ZIF-8 NPs could not only protect encapsulated proteins from protease-mediated degradation, but also retain their activity throughout the encapsulation and release. This finding supported the potential of the ZIF-8 NPs as a useful protein delivery system. An additional benefit of protein-encapsulated ZIF-8 NPs is long-term preservation of the protein activity, as shown in the activity assay of β-Gal encapsulated in ZIF-8 NPs (Figure o 2d). After three-month storage at 4 C, the unencapsulated β-Gal displayed a substantial loss of activity (~82% decrease in fluorescence signal from substrate hydrolysis). In contrast, the enzyme released from ZIF-8 NPs stored for the same period merely showed a slight loss of activity (~16% decrease in fluorescence signal from substrate hydrolysis). Combing with the properties of facile synthesis and long-term stability of protein-encapsulated ZIF-8 NPs, long-term activity preservation is another very desirable property for this protein delivery system, implying its promise for up-scalable biomedical applications. Next, we examined the toxicity of the PVP-coated protein-encapsulated ZIF-8 NPs to living cells using HeLa cell line. After incubating cells with BSA@ZIF-8 NPs of varying concentrations for different periods, we observed that 80 µg/mL BSA@ZIF-8 NPs only showed marginal toxicity, with the cell viability decreased by ~12% after 6 h incubation and by ~18% after 48 h incubation (Figure S9 in SI). Higher concentrations of BSA@ZIF-8 NPs caused substantial toxicity to the cells after 6 h or 48 h incubation. The data suggested high biocompatibility of the BSA@ZIF-8 NPs at a working concentration not higher than 80 µg/mL BSA@ZIF-8 NPs, indicating that this system afforded a loading of BSA as high as ~4.2 µg/mL for protein delivery applications. Intracellular delivery and endo-lysosomal escape of PVP-coated protein-encapsulated ZIF-8 NPs. To inspect the ability of PVP-coated ZIF-8 NPs for intracellular delivery, we prepared the NPs encapsulated with fluorescein (FITC)labeled BSA and incubated the resulting NPs (80 µg/mL)


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with HeLa cells in a culture medium supplemented with 10% fetal bovine serum. After 2 h incubation, the cells displayed bright green fluorescence in the confocal laser scanning microscopy (CLSM) images, implying that the PVP-coated ZIF8 NPs were rapidly internalized within 2 h into the cells (Figure 3a). No bright green fluorescence was obtained in a control experiment in which the FITC-labeled BSA was directly incubated with the cells (Figure 3b), implying very low cellular uptake efficiency for free protein. Flow cytometry assay for different cell lines incubated for 2 h with PVP-coated ZIF8 NPs encapsulated with FITC-labeled BSA further confirmed rapid internalization of the NPs in different cells (Figure S10 in SI). Moreover, PVP-coated ZIF-8 NPs encapsulated with EGFP afforded substantially improved delivery efficiency for different cell lines as compared to an established delivery protocol using a fusion protein of EGFP with cell penetrating peptide TAT (Figure S11 in SI). These results favorably demonstrated the potential of PVP-coated ZIF-8 NPs as a generally applicable for intracellular delivery of encapsulated proteins.

Figure 3. Delivery and endo-lysosomal escape in HeLa cells of FITC-labeled BSA in PVP-coated ZIF-8 NPs. (a) BSA@ZIF8; (b) BSA; (c) BSA@ZIF-8 and early endosome localized EER; (d) BSA@ZIF-8 and lysosome localized LyR. Scale bar: 20 μm. To ascertain the localization of the delivered proteins, we performed fluorescence staining of subcellular organelles using early endosomes-RFP fusion protein (EER) that localizes exclusively in early endosomes, lysosomes-RFP fusion protein (LyR) for lysosomes localization, and Hoechst 33342 for nuclei, respectively. After 2 h incubation of cells with ZIF-8 NPs encapsulated with FITC-labeled BSA, the CLSM images showed no overlap for the green fluorescence from delivered BSA protein with the nuclei-specific blue fluorescence (Figure 3a). In contrast, the green fluorescence exhibited moderate overlap with red fluorescence for early endosomes (Figure 3c), and to a less degree with red fluorescence for lysosomes (Figure 3d). The colocalization coefficients (Pearson’s correlation coefficient) were 0.65 and 0.32, respectively, for early endosomes and lysosomes. A relatively large colocalization coefficient for delivered proteins and early endosomes implied a portion of ZIF-8 NPs in early endosomes, while a smaller coefficient for delivered proteins and late endosomes/lysosomes suggested a less portion of ZIF-8 NPs re-

tained in late endosome/lysosome. This result indicated an endocytic pathway for cellular uptake of the ZIF-8 NPs with efficient escape of the delivered proteins from endolysosomes into the cytoplasm. Confocal images at higher resolution gave clearer evidence for efficient endo-lysosomal escape of the delivered FITC-labeled BSA (Figure S12 in SI). Further imaging experiments were performed at different sections through z-axis using Lyso@tracker specific for acidic organelles (lysosomes or late endosomes) and a cell membrane stain to outline the cells. The images also revealed that the delivered proteins exhibited obvious cytosolic localization and there was little overlapping between lysosomes or late endosomes and the delivered proteins (Figures S13 and S14 in SI). This result confirmed that PVP-coated ZIF-8 NPs enabled efficient escape of encapsulated proteins from endolysosomes. The efficient endo-lysosomal escape might be ascribed to proton sponge effect of the imidazole ligands, which induced an extensive inflow of ions and water into 39,40 endosomes and destroyed their membrane. Next, the cellular uptake mechanism for the PVP-coated protein-encapsulated ZIF-8 NPs was interrogated (Figure S15 in SI). Incubation of HeLa cells using ZIF-8 NPs encapsulated o with FITC-labeled BSA at 4 C or in the presence of NaN3 showed only dim green fluorescence, which were much o weaker than that for cells incubated at 37 C in the absence of NaN3. Flow cytometry assay further testified that cells o incubated at 4 C or in the presence of NaN3 both strongly suppressed cellular uptake of the protein-encapsulated ZIF-8 NPs. This result suggested an energy-dependent process for the cellular internalization pathway. Further CLSM analysis revealed that cells pretreated with chlorpromazine (CPZ, clathrin-mediated endocytosis inhibitor), amiloride (macropinocytosis mediated endocytosis inhibitor), and nystatin (caveolar mediated endocytosis inhibitor) delivered negligibly decreased fluorescence, but cells pretreated with methyl-β-cyclodextrin (Me-β-CD, lipid-raft mediated endocytosis inhibitor) exhibited substantially decreased fluorescence. The Me-β-CD mediated suppression of cellular uptake was also verified by the flow cytometry experiment. The data evidenced that the PVP-coated proteinencapsulated ZIF-8 NPs entered cells mainly through a lipidraft mediated endocytosis pathway. Combining these results, we inferred that the internalization of PVP-coated proteinencapsulated ZIF-8 NPs was mainly an energy-dependent, lipid-raft-mediated endocytosis mechanism. It was noteworthy that this mechanism was somewhat different 41 from that reported for ZIF-8 NPs, which might be attributed to the PVP coating or the size difference of the NPs. Activity preservation and multiple protein delivery enabled by PVP-coated ZIF-8 NPs. To evaluate the ability of PVP-coated ZIF-8 NPs to deliver proteins with preserved activity in the cell, we used ZIF-8 NPs encapsulated with βGal and examined intracellular enzymatic activity after delivery. Two cell lines, Hacat and skvo3, which are known 42 for low expression of β-Gal, were chosen in this study. When Hacat and skvo3 cells were incubated with free β-Gal followed by the substrate FDG, no green fluorescence was found in the cells, testifying that the large molecular weight protein β-Gal by itself could not enter Hacat and skvo3 cells. In contrast, the cells incubated with β-Gal encapsulated PVP-



coated ZIF-8 NPs followed by FDG displayed strong green fluorescence (Figure 4a and b). The substantial increase of fluorescence in protein-delivered cells gave a direct indicator for high intracellular enzyme activity. Additional evidences for the activity of β-Gal in Hacat and skvo3 cells were obtained using a β-Gal Reporter Gene Staining Kit (Figure 4c and d). The appearance of colored products in cells incubated with β-Gal encapsulated PVP-coated ZIF-8 NPs confirmed the activity of β-Gal delivered in the cells. These data verified that PVP-coated ZIF-8 NPs were able to preserve the activity of proteins throughout the delivery process.

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was specifically induced by the delivery of caspase 3 in the cells. Moreover, cell viability assays for cells incubated for 4 h with caspase 3/HSA co-encapsulated ZIF-8 NPs of different concentration was performed (Figure 5c). The data revealed a dose-dependent decrease in cell viability for caspase 3/HSA co-encapsulated ZIF-8 NPs, while no substantial toxicity was found for HSA encapsulated ZIF-8 NPs. This result gave confirmative evidences for the activity preservation property of the protein delivery system.

Figure 4. Enzymatic activity in cells pretreated with β-Gal and PVP-coated β-Gal@ZIF-8. Confocal images for fluorescent C12-FDG substrate in Hacat (a) and skvo3 (b) cells. Scale bar: 20 μm; light micrographs for colorimetric substrate in Hacat (c) and skvo3 (d) cells. Scale bar: 25 μm. To further validate activity preservation property of the protein delivery system, we used PVP-coated ZIF-8 NPs coencapsulated with two proteins, caspase 3 and HSA. This coencapsulation system could be utilized to reduce the amount of some costly proteins such as caspase 3 in the biomineralized synthesis, with adequate activity still achieved for their biomedical applications. To examine the activity for caspase 3 in cells, we evaluated the apoptosis-induced cell death in response to the protein delivery. It is known that caspase 3 activation induces phosphatidylserine externalization at the early stage and at last leads to death of cells with damaged 43 plasma membrane. Hence, the apoptosis-inducing activity of caspase 3 could be investigated using Alexa Fluor® 488 labeled annexin V, a fluorescent staining reagent specific for 44 phosphatidylserine externalization, and propidium iodide (PI), a fluorescent dye impermeable to the intact cellular membranes but brightly stain nucleic DNA of cells with 45 damaged membranes. As anticipated, cells incubated with caspase 3/HSA co-encapsulated ZIF-8 NPs for 2 h displayed bright green fluorescence on their membranes (Figure 5a), indicator for phosphatidylserine externalization at the early stage of apoptosis. After 4 h incubation, bright green fluorescence on their membranes and red fluorescence in nuclei were both observed, suggesting the damage of cell membrane and apoptosis-induced cell death. In control experiments for cells without treatment using ZIF-8 NPs or treated using HSA-encapsulated ZIF-8 NPs, neither green nor red fluorescence was observed. Fluorescence intensity analysis (Figure 5b) and flow cytometry experiments (Figure S16 in SI) also verified that phosphatidylserine externalization appeared after 2 h incubation and membrane damage occurred after 4 h incubation. These results validated that apoptosis

Figure 5. Activity of caspase 3 delivered in HeLa cells. (a) Fluorescent images of control cells or cells treated with HSA@ZIF-8, and caspase 3/HSA@ZIF-8 for 2 or 4 h. Scale bar: 20 μm; (b) Average fluorescence intensity of control cells (1) and cells treated with HSA@ZIF-8 (2), and caspase 3/HSA@ZIF-8 for 2 h (3) or 4 h (4); (c) Cell viability treated with HSA@ZIF-8 or caspase 3/HSA@ZIF-8 of different concentrations for 4 h. Moreover, we examined the preservation of antibody targeting activity through delivery of an antibody toward tubulin in living cells. In the assay, FITC-labeled anti-tubulin antibody was delivered using the PVP-coated ZIF-8 NPs into HeLa cells with expression of fusion protein of red fluorescence protein (mCherry) and tubulin. The confocal images showed that the green fluorescence of delivered anti-tubulin antibody was remarkably colocalized with the red fluorescence of tubulin (Figure S17 in SI). The colocalization correlation coefficient was 0.7, indicating that most of delivered anti-tubulin antibodies were able to target tubulin in the cytosol. This finding clearly confirmed that proteins delivered using PVP-coated ZIF-8 NPs were efficiently released from endo-lysosomes into the cytosol and the released antibodies still retained their activity to bind to the antigen. Co-delivery of two or more different proteins has largely


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unexplored because of the difficulty in accommodating the divergent physicochemical properties of different proteins. Biomimetic mineralization afforded the advantage of encapsulating multiple proteins in MOF NPs using the same onepot synthetic chemistry with no preliminary protein modification, implying its potential for co-delivery multiple proteins. To demonstrate the potential, we prepared ZIF-8 NPs encapsulated with fluorescein (FITC)-labeled BSA, red fluorescence protein (RFP) and NIR-641-labeled β-Gal. CLSM revealed that individual protein-encapsulated ZIF-8 NPs were shown as bright spots in three fluorescent images, and the merged image displayed perfect co-localization of the three-color fluorescence (Figure S18 in SI). A closer examination of fluorescence intensity profiles further verified colocalization of the three-color fluorescent spots. This finding confirmed co-encapsulation of the three proteins in each single ZIF-8 NPs, in which the separation between different proteins could not be resolved in CLSM images. Co-delivery of the three proteins with different colors in cells using the ZIF-8 NPs gave images with bright fluorescence in three channels (Figure 6a), indicating successful co-delivery of the three proteins in cells. Moreover, though three proteins all displayed remarkably cytosolic localization in the cells, there were localization discrepancies between different proteins, suggesting release and separation of the three proteins in the cells. The separated localization of three proteins as an indicator for protein release from the encapsulated ZIF-8 NPs was also confirmed by the fluorescence intensity profiles (Figure 6b). This result demonstrated that proteinencapsulated ZIF-8 NPs afforded a useful platform for intracellular multiple protein co-delivery.

Figure 6. ZIF-8 NPs for proteins co-delivery. (a) Fluorescent images of HeLa cells incubated with ZIF-8 NPs encapsulated with fluorescein (FITC)-labeled BSA, red fluorescence protein (RFP) and NIR-641-labeled β-Gal. Scale bar: 10 μm; (b) Fluorescence intensity profile of regions of interest (white line).

CONCLUSION We developed biomineralized protein-encapsulated ZIF-8 NPs as a novel platform for efficient intracellular delivery and endo-lysosomal release of protein. By finely controlling the 2+ concentration ratio of MIM to Zn , a recently reported biomimetic mineralization procedure was adapted for facile one-pot synthesis of nanoscale protein-encapsulated ZIF-8. This method is generally applicable to synthesizing ZIF-8 NPs encapsulating different proteins with high efficiency in no need of protein modification. Moreover, the ZIF-8 NPs were shown to carry proteins via a definite encapsulation mechanism. This encapsulation structure had the advantages

of preserving protein activity and protecting proteins from enzyme-mediated degradation. Furthermore, a simple procedure to afford long-term stability to the ZIF-8 NPs in cell media was introduced using a biocompatible PVP coating. The protein-encapsulated ZIF-8 NPs was demonstrated to enable efficient cellular uptake, rapid release and escape of proteins from endo-lysosomes, and preservation of the protein activity in living cells. Moreover, the MOF NPs based platform had the ability to encapsulate multiple proteins in single MOF NPs, affording a useful strategy for protein co-delivery. To our knowledge, it is the first time that protein-encapsulating MOF NPs have been demonstrated as a generally applicable protein delivery platform. Therefore, the protein-encapsulated biomineralized MOF NPs may represent a valuable platform for protein delivery and protein-based theranostics.

MATERIALS AND METHODS Materials. BSA, HSA, nystatin and protease mixture of trypsin with α-chymostrypsin were supplied by Sangon Biotech Co., Ltd. (Shanghai, China). EGFP, EGFP-TAT fusion protein and RFP plasmid pCMV-mCherry-Tubulin-C-18 were purchased from Miaolingbio Bioscience & Technology Co., Ltd. (Wuhan, China). Recombinant human caspase 3 was obtained from R&D systems (Minneapolis, USA). Ferritin, βGal (500 U/mg), FITC-labeled anti-α-tubulin antibody, PVP (wt 40,000), FDG substrate, β-Gal Reporter Gene Staining Kit, sodium azide, amiloride, CPZ, Me-β-CD, NIR-641 Nsuccinimidyl ester, Hoechast 33342, zinc nitrate hexahydrate, 2-methylimidazole (MIM), methanol (HPLC grade), and MTT assay kit were purchased from Sigma-Aldrich (St. Louis, USA). HeLa, MCF-7, HepG2, Hacat and skvo3 cell lines were supplied by the cell bank of Central Laboratory at Xiangya Hospital (Changsha, China). CellLight® Early Endosome-RFP (expresses RFP fused to Rab5a, a protein localized to early endosomes), CellLight® Lysosome-RFP (expresses RFP fused to LAMP1, a lysosomal associated membrane protein 1), Cellmask deep red plasma membrane stain, Alexa Fluor® 488 annexin V, PI Dead Cell Apoptosis Kit and cell culture media were obtained from Thermo Fisher Scientific Inc. (MA, USA). All other chemicals were of analytical grade and purchased from Sinopharm Chemical Reagent Co. Ltd. (Shanghai, China). All solutions were prepared using ultrapure water, which was obtained through a Millipore Milli-Q water purification system (Billerica, MA), with an electric resistance >18.3 MΩ. FITC-labeled BSA was prepared by mixing 10 mg/mL BSA in 0.1 M phosphate buffer (PB, pH 8.0) with 1 mg/mL FITC at 37 °C for 2 h. NIR-641-labeled β-Gal was prepared by mixing 2mg/mL β-Gal in 0.1 M PB (pH 7.5) with 1 mg/mL NIR-641 N-succinimidyl ester at 37 °C for 2 h. The fluorophore-conjugated proteins were purified by dialysis against ultrapure water for 48 h to remove excess o fluorescent dye and stored at 4 C for future use. Synthesis of Protein Encapsulated ZIF-8 NPs. In a typical synthesis of ZIF-8 NPs encapsulated with proteins, a given amount of the protein (0.25, 0.50 or 0.75 mg BSA, 1.0 or 3.5 mg ferritin, 0.1 mg EGFP, 0.25 mg HSA, 0.5 mg β-Gal, 0.5 mg FITC-labeled anti-α-tubulin antibody, three protein mixtures of 0.098 mg RFP, 0.125 mg FITC-labeled BSA and 0.75 mg NIR-641 labeled β-Gal, or two protein mixture of 0.23 mg HSA and 0.02 mg caspase 3) was firstly added in an aqueous



solution of MIM (3.15 mmol, 0.9 mL) under vigorous stirring and incubated at 30 °C for 10 min. Then, an aqueous solution of zinc nitrate hexahydrate (0.045 mmol, 0.1 mL) was mixed with the above mixture and stirred at 30 °C for 10 min. The resulting ZIF-8 NPs were centrifuged at 3500 rpm for 20 min, washed with ethanol twice, washed using 5% SDS (w/w) aqueous solution at 50 °C to remove free proteins on the surface, and finally mixed and stirred with 3% PVP (w/w) aqueous solution for 30 min. The as-prepared PVP-coated protein-encapsulated ZIF-8 NPs were stored at 4 °C for future use. The ZIF-8 NPs encapsulated with proteins were lyophilized and weighed to determine the amount of products obtained in a single synthesis. The product amounts were ~8.7 mg BSA@ZIF-8, ~9.0 mg β-Gal@ZIF-8, ~8.1 mg EGFP@ZIF-8, ~10.2 mg HSA/Caspase-3@ZIF-8, and ~12.1 mg ZIF-8 NPs encapsulated with three proteins, respectively. Cell Culture and Fluorescence Imaging. HeLa cells, Hacat cells, skvo3 cells, MCF-7 cells and HepG2 cells were cultured in a RPMI 1640 medium supplemented with 10% fetal bovine serum, 100 U/mL penicillin and 100 U/mL streptomycin. All o cell lines were maintained at 37 C in a 100% humidified atmosphere containing 5% CO2. Fluorescence imaging of cells was performed as follows: HeLa cells were seeded on a 35-mm Petri dish with 10-mm o well in 2 mL culture medium at 37 C for 24 h. Then, the cells were incubated with 1 mL culture medium containing 5 μg/mL FITC labeled BSA, 80 μg/mL FITC-labeled BSA@ZIF-8 NPs, or 120 μg/mL ZIF-8 NPs encapsulated with FITC-labeled BSA, red fluorescence protein (RFP) and NIR-641-labeled βo Gal at 37 C for 1 or 2 h. The cells were washed three times with cold PBS before imaging. Fluorescence imaging of intracellular localization for the delivered proteins was performed as follows: HeLa cells were firstly transfected according to the instruction of the reagents with CellLight® Lysosomes-RFP or CellLight® Early 5 Endosomes-RFP (10 μL stock solution for 10 cells in 1 mL o culture medium) at 37 C for 16 h. The transfected cells were incubated with 80 μg/mL FITC-labeled BSA@ZIF-8 NPs at 37 o C for 2 h, and washed three times with cold PBS before imaging. Fluorescence imaging of intracellular activity for the delivered enzymes was performed as follows: Hacat cells and skvo3 cells were incubated with 5 μg/mL β-Gal or 0.09 mg/mL β-Gal@ZIF-8 NPs (β-Gal concentration in the NPs o solution was ~5 μg/mL or ~2.5U/mL) at 37 C for 2 h. Then, the cells were incubated with 300 μM chloroquine diphosphate for 30 min followed by 33.3 μM C12-FDG for another 1 h. The cells were washed three times with cold PBS before imaging. HeLa cells were incubated with 100 μg/mL HSA@ZIF-8 (protein concentration in the NPs solution was ~2.5 μg/mL) or HSA/caspase 3@ZIF-8 (protein concentrations in the NPs solution were ~2.3 μg/mL for HSA o and ~0.2 μg/mL for caspase 3, respectively) at 37 C for a given time. Then, the cells were incubated with Alexa Fluor® 488 annexin V and PI dead cell apoptosis kit for 30 min, followed by washing three times with cold PBS before imaging. All fluorescence images were acquired using an oil dipping objective (100×, 1.25 NA) on a Nikon confocal laser scanning fluorescence microscope at a sampling speed of 6.2

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pixel/Dwell and a size of 1024. The transmissivity of FITC, HV, and offset values were set to 10, 150, and 0 for green channel. The transmissivity of TRITC, HV, and offset values were 10, 150, and 0 for red channel. The transmissivity of Cy5, HV, and offset values were 10, 150, and 0 for deep red channel. Colorimetric detection of intracellular activity for the delivered enzymes was performed as follows: Hacat cells and skvo3 cells were incubated with 5 μg/mL β-Gal or 0.09 mg/mL β-Gal@ZIF-8 NPs (protein concentration in the NPs o solution was ~5 μg/mL or ~2.5 U/mL) at 37 C for 2 h. Then, the cells were fixed in a 1× fixation buffer (included in βGalactosidase Reporter Gene Staining kit) for 10 min. The fixed cells were subsequently incubated with 5-bromo-4chloro-3-indolyl β-D-galactopyrano-side (1 mg/mL) in 1× PBS (included in the kit) containing 2 mM MgCl2, 4 mM potassium ferricyanide, and 4 mM potassium ferrocyanide at 37 °C for 2 h followed by washing with 1× PBS before imaging using XD-202 inverted biological microscope (40× objective).

ASSOCIATED CONTENT Supporting Information Experimental methods including biomineralized synthesis of MOF NPs, TEM, FTIR and UV-vis experiments, protein encapsulation efficiency determination, MTT assay, and flow cytometry assay as well as additional figures. These materials are available free of charge on the ACS Publications website.

AUTHOR INFORMATION Corresponding Author [email protected]

ORCID Xia Chu: 0000-0002-4120-6131

Notes The authors declare no competing financial interests.

ACKNOWLEDGMENT This work was supported by the National Natural Science Foundation of China (Grants 21525522 and 21705039), and the National Postdoctoral Program for Innovative Talents (BX201600049).

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