Fluorescent Protein Capped Mesoporous ... - Wiley Online Library

DOI: 10.1002/chem.201302026

Fluorescent Protein Capped Mesoporous Nanoparticles for Intracellular Drug Delivery and Imaging Xinjian Yang ,[a, b] Zhenhua Li ,[a, b] Meng Li ,[a, b] Jinsong Ren,*[a] and Xiaogang Qu*[a] Abstract: A multifunctional system for intracellular drug delivery and simultaneous fluorescent imaging was constructed by using histidine-tagged, cyan fluorescent protein (CFP)-capped magnetic mesoporous silica nanoparticles (MMSNs). This protein-capped multifunctional nanostructure is highly biocompatible and does not affect cell viability or proliferation. The CFP acts

not only as a capping agent, but also as a fluorescent imaging agent. The nanoassembly was activated by histidinebased replacement, leading to release Keywords: drug delivery · fluorescent probes · imaging agents · mesoporous materials · nanostructures

Introduction Controlled-release systems have attracted great attention owing to their applicability in drug delivery. To date, diverse organic and inorganic carriers have been investigated for their utility as delivery systems.[1] Among these materials, mesoporous silicas have been recognized as attractive nanocarriers due to their stable mesoporous structures, large surface areas, tunable pore sizes and volumes, good biocompatibility, and facile surface functionalization.[2] In particular, mesoporous silica nanoparticles are useful scaffolds for equipping with “smart caps” on the pore outlets. Recent studies have demonstrated that inorganic nanoparticles,[3] polymers,[4] larger supramolecular assembles[5] and biomolecules[6] could be used as blocking caps to control the opening or closing of pore entrances of mesoporous silica. In response to different stimuli, such as pH,[7] light,[8] competitive binding,[9] enzymes,[10] and redox potential,[11] the well-designed gate-keepers could be opened according to previous on-command release systems. Despite these burgeoning achievements, many of the existing cap systems still present

[a] X. Yang , Z. Li , M. Li , Prof. J. Ren, Prof. X. Qu State Key Laboratory of Rare Earth Resource Utilization and Laboratory of Chemical Biology Changchun Institute of Applied Chemistry Chinese Academy of Sciences Changchun, 130022 (P.R. China) Fax: (+ 86) 0431-85262625 E-mail: [email protected] [email protected] [b] X. Yang , Z. Li , M. Li Graduate School of the Chinese Academy of Sciences Beijing, 100039 (P.R. China)

of drug molecules encapsulated in the nanopores into the bulk solution. The fluorescent imaging functionality would allow noninvasive tracking of the nanoparticles in the body. By combining the drug delivery with cell-imaging capability, these nanoparticles may provide valuable multifunctional nanoplatforms for biomedical applications.

challenges in terms of biocompatibility and the multifunctionality of the capping agents. Native biomacromolecules, such as fluorescent proteins, have several advantages over the existing caps for drug delivery, since they would enable high biocompatibility and act as imaging agents. Fluorescent proteins have been recognized as efficient tools for gene expression, cellular localization of proteins, and various dynamic intracellular events.[12] Histidine (His) is an important amino acid for humans and animals, and an abnormal level of histidine is an indicator for many diseases.[14] In this work, we report the construction of a multifunctional platform for simultaneous fluorescent imaging and bioresponsive (histidine) controlled drug delivery by using His-tagged, cyan fluorescent protein (HCFP)-capped magnetic mesoporous silica nanoparticles (MMSNs). As shown in Scheme 1 below, HCFP acts as a capping agent through the chemical interaction between Ni nitrilotriacetate (Ni-NTA) complex and His-tagged protein. This simple and highly selective interaction has been used extensively for purification, surface immobilization, and in vitro detection of recombinant His-tagged proteins.[13] As demonstrated in the literature, His-tagged protein could be removed from the Ni-NTA moiety by introduction of imidazole. We thus expected that the drug-release process would occur in the presence of histidine, which has an imidazole group. Magnetic nanoparticles were incorporated within the mesoporous silica framework due to their potential biomedical applications, such as in hyperthermia treatment of cancer, as contrast agents for magnetic resonance imaging, magnetic separation, and sorting of cells and proteins (biorecognition), and for targeted drug delivery.[15]

Supporting information for this article is available on the WWW under http://dx.doi.org/10.1002/chem.201302026.

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FULL PAPER NHS) in 2-(N-morpholino)ethanesulfonic acid (MES) buffer and subsequently treated with nitrilotriacetate (NTA) to obtain MMSN-NTA, which was used to immobilize nickel(II) ions. The surface functionalization of various MMSNs was monitored by FTIR spectroscopy (see Figure S4 in the Supporting Information). The emerging absorption band at around 1560 cm 1 for MMSNs-COOH can be assigned to the acylamide vibration of the attached succinic acid molecules. After the decoration of the surface of MMSNs with NTA molecule by means of an acylamide bond, the absorption at around 1560 cm 1 increased. The energy-dispersive X-ray spectrum (see Figure S5 in the Supporting Information) showed that Ni2 + was chelated on the Figure 1. a) SEM image of MMSNs. b) Nitrogen sorption isotherms of the MMSNs. Inset: pore distribution of surface of nanoparticles. The MMSNs obtained from desorption branch. TEM images of NTA-MMSNs (c) and HCFP-capped MMSNs (d). closure reaction was performed in phosphate-buffered saline (PBS) at room temperature by mixing MMSNs-NTA-Ni Results and Discussion with HCFP. After magnetic separation and washing steps, TEM images and dynamic light scattering (DLS) were used The MMSNs were prepared according to the modified to demonstrate the attachment of HCFP on the surface of method reported previously.[16] The structure of the resulting MMSNs. No clear difference in the shape of the proteinnanoparticles was characterized by low-angle X-ray diffraccapped MMSNs was observed in comparison with uncapped tion, SEM, and TEM (Figure 1 and Figures S1 and S2 in the Supporting Information). The mean diameter of the nanoparticles estimated from SEM and TEM images was about 55 nm, which is highly desirable, because small nanoparticles usually stay longer in blood circulation and can be taken up more easily by cancer cells. No hysteresis loop was apparent in their saturation magnetization curves (see Figure S3 in the Supporting Information), which indicates that the MMSNs are superparamagnetic. Thus, the MMSNs could be easily separated by applying a magnetic field after dispersal in aqueous solution. The N2 adsorption-desorption isotherms of the MMSN showed a typical Type IV curve with a specific surface area of 499 m2 g 1, average pore diameter of 2.8 nm, and a narrow pore distribution (Figure 1 b). The modification procedure is shown in Scheme 1 a. The surface of nanoparticles was first functionalized with amino groups by treatment with 3-aminopropyltriethoxysilane (APTES) to give MMSN-NH2. Silica particles functionalized with carboxyl groups (MMSN-COOH) were obtained by allowing MMSNNH2 to react with succinic anhydride in DMF. The resultant carboxyl group on the surface was activated with 1-[3-(dimethylamino)propyl]-3-ethylcarbodiimide hydrochloride Scheme 1. Synthetic route to HCFP-capped magnetic mesoporous silica (EDC) and N-hydroxysulfosuccinimide sodium salt (sulfo(a) and histidine-responsive guest molecule release (b).

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nanoparticles (Figure 1 c and d). However, a border appeared after anchoring the protein on the nanoparticles. This result was related to efficient grafting of HCFP to the nanoparticles and blocking of the pores of the nanoparticles. The DLS experiment showed that the hydrodynamic diameter of the observed particles was slightly increased after protein attachment (see Figure S6 in the Supporting Information). This result further demonstrated the highly efficient attachment of HCFP to NTA-modified nanoparticles. To demonstrate histidine-triggered uncapping of HCFP, HCFP-capped MMSNs were incubated with histidine solution in PBS at pH 7.4. As shown in Figure 2 a and Figure S7

Figure 2. Release profiles of His-CFP (a) and IBU (b) from HCFPcapped MMSNs in PBS in the presence of 50 mm of histidine.

in the Supporting Information, we observed a significant increase in the amount of HCFP released by measuring the fluorescence intensity of the supernatant after 5 h of incubation, which indicates that HCFP molecules were released from the pore outlet of MMSNs due to replacement by histidine. However, the increase in fluorescence intensity of the supernatant is negligible in the absence of histidine. In an additional control experiment, an increase in fluorescence also occurred on addition of imidazole to the solution of HCFP-capped MMSNs (see Figure S7 in the Supporting Information). The different uncapping efficiencies of histidine and imidazole is due to their difference in steric hindrance. To further investigate histidine-responsive release behavior of HCFP-capped MMSNs, ibuprofen (IBU) was loaded as a guest molecule by soaking MMSNs-NTA-Ni and IBU in hexane (loaded amount: 8.2 wt %, see Figure S8 in the Supporting Information). To examine the capping efficiency, the HCFP-capped MMSNs loaded with IBU were first dispersed in PBS solution without histidine molecules. The released amount of CFP or IBU of less than 5 % after 4 h indicated no pronounced leakage of the entrapped guest mole-

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cules. This result indicated that the capping strategy was successful with good efficiency. The triggered release of IBU was investigated by addition of histidine to the HCFPcapped MMSNs. In contrast to the experiment without histidine, up to about 40 % of IBU was released from HCFP capped-MMSNs within 8 h after introduction of histidine under otherwise identical conditions, which suggested a good response to histidine (Figure 2 b). This result shows that the linkage between HCFP and MMSNs-NGTA-Ni could be dissociated through a competitive displacement reaction in the presence of histidine, and the His-tagged proteins are uncapped from the MSN system. Similar results were obtained when the IBU release experiment was carried in real media (see Figure S9 in the Supporting Information). Nonspecific protein adsorption is a problem that restricts the use of many nanomaterials. In our system, the released amount decreased only slightly, that is, few protein molecules from the medium were adsorbed after HCFP was released from the surface of MMSNs. This result guarantees the use of these materials in physiological fluids. To assess the in vitro safety and biocompatibility of the HCFP-coated MMSNs, we analyzed mitochondrial activity of Hela cells following incubation with different concentrations of HCFP-coated MMSNs. No significant in vitro cytotoxicity was observed, even at a nanoparticle concentration of 1 mg mL 1 (see Figure S10 in the Supporting Information). Next, investigations of the controlled-release system were extended to an in vitro study for intracellular delivery by introducing a chemotherapeutic agent, namely, doxorubicin (DOX), which is the most utilized anticancer drug against a range of neoplasms, including acute lymphoblastic and myeloblastic leukemias, as well as malignant lymphomas. To confirm the toxicity of DOX, HepG2 cells were incubated with DOX in culture medium without fetal bovine serum (FBS) at 37 8C, 5 % CO2. After 2 h incubation, the unbound DOX molecules were removed, and fresh medium (10 % FBS) was added for further cell growth (48 h). The relative viability of cells with different treatments was determined by MTT assay. The results shown in Figure 3 a demonstrated that DOX at micromolar levels had high toxicity toward HepG2 cells and could greatly inhibit cell proliferation. However, the cells treated with DOX-loaded HCFPcapped nanoparticles (loaded amount: 7.7 wt %) showed low toxicity, which further confirmed the high capping efficiency. Having established that our drug-release system is histidine-responsive, we next tested the cytotoxicity of the chemotherapeutic agent utilized in this study to assess the tumoricidal potential of our controlled drug-delivery system. To this end, cells were incubated with 50 mm of histidine 2 h after adding DOX-CFP-MMSNs under the same condition for 48 h. Figure 3 a shows that DOX-loaded, HCFP-capped MMSNs greatly increased the death of HepG2 cells compared to DOX by itself. This was also confirmed by the morphological changes of nuclei (Figure 3 b–d). Cells in the presence of DOX-loaded CFP-MMSNs showed normal morphology of nuclei after 48 h of treatment, On the contrary, nuclear condensation and deformation were revealed in


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Figure 4. Fluorescent images of HepG2 cells incubated with HCFPMMSNs (a) and HCFP-MMSNs-DOX (b) on adding histidine.

Figure 3. a) Cytotoxicity of HepG2 cells exposed to free DOX, HCFPMMSNs-DOX and HCFP-MMSNs-DOX incubated with 50 mm of histidine for 48 h. The bottom panel shows the nuclei of HepG2 cells exposed to HCFP-MMSNs-DOX (b), free DOX (c), and HCFP-MMSNs-DOX in the presence of 50 mm of histidine (d) after 48 h of incubation.

most of the cells treated with free DOX and DOX-loaded CFP-MMSNs in the presence of histidine. Importantly, nuclear fragmentation occurred in the cells treated with DOXloaded CFP-MMSNs on adding histidine. Taken together, these results implied that cell death could be attributed to intracellular DOX release from HCFP-capped MMSNs triggered by histidine. Furthermore, the nanocomposites could also be used for cell imaging. Compared to fluorescent organic molecules and inorganic nanoparticles, such as semiconductor quantum dots, the HCFP-capped MMSNs capped system could provide the possibility of cell imaging due to their photoluminescence (PL) properties and nontoxicity. Although the Fe3O4 could quench the fluorescence of CFP, the silica shell and the high loading of CFP make this platform suitable for cell imaging (see Figure S11 in the Supporting Information). Based on this result, we evaluated the capability of the HCFP-capped MMSNs for bioimaging by incubating them with HepG2 cells under physiological conditions. As shown in Figure 4 a, the cells show strong cyan fluorescence on the entire cytoplasm after 4 h of incubation with HCFP-capped MMSNs. Furthermore, to visualize intracellular delivery of anticancer drugs by the HCFP-capped MMSNs system for intracellular therapeutic applications, HCFP-capped MMSNs-DOX were studied. Bright cyan fluorescence was also found in the entire cell cytoplasm after 4 h of incubation in the presence of histidine (Figure 4 b), which is consistent with the previous result that CFP was replaced slowly and suggests that this material could be effectively taken up by the cancer cells and is mainly localized in the cytoplasm

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instead of entering the nucleus. By contrast, red fluorescence throughout the cell nuclei indicated that the release of DOX molecules from the MMSNs was due to replacement of CFP by histidine, and thus confirmed the applicability of the HCFP-capped MMSNs system for intracellular delivery and release of drugs for the cancer therapy. Hence, the fluorescent imaging capacity of HCFP-NTA-MMSNs makes this platform an ideal candidate for tracking cancer cells.

Conclusion We have constructed a multifunctional system for intracellular drug delivery and simultaneous fluorescent imaging by using His-CFP-capped MMSN nanoparticles. This proteincapped multifunctional nanostructure is highly biocompatible and does not affect cell viability or proliferation. The CFP acts not only as capping agent, but also as a fluorescent imaging agent. The nanoassembly was activated by histidine-based replacement, which led to release of drug molecules encapsulated in the nanopores into bulk solution. The fluorescent imaging functionality would allow noninvasive tracking of the nanoparticles within the body. By combining drug delivery with the cell-imaging capability, these nanoparticles may provide valuable multifunctional nanoplatforms for biomedical applications.

Experimental Section Synthesis of iron oxide nanoparticles: FeCl3·6 H2O (2.43 g) and FeCl2·4 H2O (1.195 g) were dissolved in H2O (20 mL). The solution was stirred at 90 8C for 10 min. Then NH3·H2O (6 mL) and oleic acid (0.4 g) were added to the solution, followed by another 3 h of stirring. The mixture was washed with water and absolute ethanol several times. The synthesized iron–oleic acid complex was then dried under vacuum overnight. Mesoporous silica formation: The dried oleic acid capped iron oxide NPs were dissolved in chloroform. The large aggregates were removed by a magnet. Two milliliters (10–20 mg mL 1) of the NC solution was mixed with 0.4 g of cetyltrimethylammonium bromide (CTAB, Aldrich) and water (20 mL). The mixture was then sonicated and stirred vigorously, and the chloroform solvent was boiled off from the solution. Then distilled water (172 mL) and sodium hydroxide (1.4 mL, 2 m) were added and the mixture heated to 80 8C. After the temperature had stabilized,


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tetraethyl orthosilicate (0.5 mL) was added slowly to the aqueous solution. The solution was stirred for another 2 h. The synthesized materials were centrifuged and washed with methanol. The CTAB surfactant was removed from the mesopores by dispersing the as-synthesized materials in a solution of ammonium nitrate (160 mg) and 95 % ethanol (60 mL) and heating the mixture at 60 8C for 30 min. The materials were then centrifuged and washed with ethanol. Modification of MMSNs: The MMSNs (1.0 g) were treated with APTES (0.5 mL) in dry toluene (30 mL) for 12 h under N2 gas to give NH2MMSNs. After the reaction, the nanoparticles were collected by filtration and washed with methanol. The NH2-MMSNs (100 mg) were treated with succinic anhydride (1.00 g) in DMF solution (20 mL) under N2 gas for 8 h with continuous stirring. Thus, carboxyl groups were formed on the MMSN surface for conjugation of NTA. After a thorough water wash, the carboxylated nanoparticles (MMSN-COOH) were activated with EDC (10 mg mL 1, 15 mL) and sulfo-NHS (10 mg mL 1, 15 mL) in MES buffer (pH 6.0) for 15 min at room temperature with continuous stirring. The pH of the solution was adjusted to 8.0, followed by the addition of NTA (0.1 g mL 1) at room temperature with continuous stirring for 24 h. The resulting nanoparticles were washed with water three times and then mixed with 1 m NiSO4 to give MMSNs-NTA-Ni, followed by washing with water and ethanol. HCFP attachment and release: The closure reaction was performed in PBS at room temperature for 2 h by mixing MMSNs-NTA-Ni (2 mg mL 1) with His-tagged CFP (0.5 mg mL 1). Then the nanocomposites were magnetically separated, followed by washing twice with PBS. The release of HCFP was monitored at 474 nm with a fluorescent spectrophotometer. HCFP-capped nanoparticles (1 mg mL 1) were stirred slowly with histidine (50 mm). At selected time intervals, a sample (0.1 mL) was removed and immediately replaced with an equal volume of fresh PBS. Then nanoparticles in the removed solution were magnetically separated, and the concentrations monitored by measuring the fluorescence of the supernatant. Drug loading and release: IBU was dissolved in hexane at a concentration of 60 mg mL 1. MSN-NTA-Ni (100 mg) was then added to this solution at room temperature for 24 h. After magnetic separation, the IBU-loaded nanoparticles were washed three times with PBS. The closure reaction was performed in PBS at room temperature for 2 h by mixing MMSNsNTA-Ni with His-tagged CFP. The capping efficiency was monitored by FL spectroscopy. The loading of IBU was calculated from the difference in the concentration of initial and residual drug in the supernatant. The IBU loading in MMSNs was determined to 8.2 wt %. The release experiment was conducted in PBS with histidine (50 mm). Drug-loaded HCFP capped nanoparticles (1 mg mL 1) were stirred slowly with histidine (50 mm). At selected time intervals, a sample (0.1 mL) was removed and immediately replaced with an equal volume of fresh PBS. Then nanoparticles in the solution were removed by magnetic separation. Then the protein was removed by an ultracentrifugal filter device (10 000 NMWL) at 13 000 rpm for 20 min. IBU in the protein-free solution was monitored at 222 nm by using a UV/Vis spectrophotometer. MTT assay: HepG 2 cells were placed in a 96-well plate for 24 h. On the day of the experiment, cells were washed with prewarmed PBS (100 mL). Freshly prepared DOX-loaded nanocarriers (prepared in PBS, loading efficiency was 7.7 wt %) were added to the cell. Free DOX was used as a positive control. Untreated cells were used as a negative control. The cells were incubated for 6 h and then washed with PBS. The cells were further incubated in prewarmed growth medium for 42 h in a 5 % CO2 incubator at 37 8C. Afterwards, cells were incubated in media containing MTT (0.5 mg mL 1) for 3 h. The precipitated formazan violet crystals were dissolved in DMSO (150 mL) at 37 8C. The absorbance was measured at 490 nm by multidetection microplate reader.

Acknowledgements Financial support was provided by the National Basic Research Program of China (Grant 2012CB720602, 2011CB936004) and the National Natu-

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ral Science Foundation of China (Grants 21072182, 21210002, and 91213302).

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Received: May 27, 2013 Revised: July 29, 2013 Published online: September 20, 2013

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