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Cells as Factories for Humanized Encapsulation Zhengwei Mao,^,† Regis Cartier,^,† Anja Hohl,† Maura Farinacci,‡ Anca Dorhoi,‡ Tich-Lam Nguyen,§ Paul Mulvaney,§ John Ralston,|| Stefan H. E. Kaufmann,‡ Helmuth M€ohwald,† and Dayang Wang*,†,|| †
Max Planck Institute of Colloids and Interfaces, D-14424, Potsdam, Germany Max Planck Institute for Infection Biology, Department of Immunology, 10117, Berlin, Germany § School of Chemistry and Bio21 Institute, University of Melbourne, VIC 3010, Australia Ian Wark Research Institute, University of South Australia, Adelaide, SA 5095, Australia
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‡
bS Supporting Information ABSTRACT: Biocompatibility is of paramount importance for drug delivery, tumor labeling, and in vivo application of nanoscale bioprobes. Until now, biocompatible surface processing has typically relied on PEGylation and other surface coatings, which, however, cannot minimize clearance by macrophages or the renal system but may also increase the risk of chemical side effects. Cell membranes provide a generic and far more natural approach to the challenges of encapsulation and delivery in vivo. Here we harness for the first time living cells as “factories” to manufacture cell membrane capsules for encapsulation and delivery of drugs, nanoparticles, and other biolabels. Furthermore, we demonstrate that the built-in protein channels of the new capsules can be utilized for controlled release of encapsulated reagents. KEYWORDS: Drug delivery, encapsulation, nanoparticles, cell membranes, nanostructures ncapsulation is of paramount importance in various fields of medical applications to improve the biocompatibility of diagnostic and therapeutic substances, reduce their toxicity, and at the same time facilitate their efficient delivery to specific targets in the body in a large mass fraction.1 A great variety of biodegradable and/or biocompatible materials, such as phospholipid liposomes,2 amphiphilic molecular micelles,3 polyelectrolyte microcapsules4 or polymer microspheres,5 erythrocyte ghosts,6 and vesicles derived from bacteria,7 virus-like particles,8 to name a few, have been employed to produce nanometer- or micrometer-sized carriers for encapsulation. These carriers may be further rendered stealthlike after deliberate surface modification with, for example, poly(ethylene glycol) (PEG).9 However, all synthetic carriers developed thus far are rapidly reused and cleared, notably by phagocytes, possibly due to the non-natural surface characteristics in terms of chemistry, morphology, and mechanics. To circumvent this important challenge, we exploit living cells as factories to encapsulate drugs and nanoparticles (NPs) in “natural vesicles”, composed of various cell membrane lipids and proteins. To underline their immense promise in drug delivery and encapsulation, these natural vesicles here are termed cell membrane capsules (CMCs). These CMCs maintain all the membrane functions and cytosolic proteins of the parental cells. Here our pilot study investigating the macrophage response to drug or NP-laden CMCs has been carried out to assess the potential of CMCs to avoid attack by the immune system. We demonstrate that CMCs act as unprecedented delivery vehicles, in which encapsulated substances can be processed stepwise by
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cellular enzymes and then be selectively released through protein channels of the membrane, in a controlled and sustained manner. It is well-known that living cells can endocytose molecules or NPs from the surrounding environment into natural vesicles composed of cell membrane lipids and proteins—endosomes or lysosomes. Conversely, these molecules and NPs can be released from the cells via exocytosis to restore the membranes and cytoplasm of the cells or as a means to deliver important proteins into the extracellular environment.10 The integration of endocytosis and exocytosis provides a mechanism for harnessing living cells in order to produce CMCs for encapsulation. However, exocytosis is usually signal- or receptor-specific and difficult to manipulate. In 1972, Sanger and Holtzer reported the first observation of genesis of nonexocytic, natural vesicles from cells treated with cytochalasin B (CB), a drug that affects cytoskeleton membrane interactions.11After about 2 decades, Bousquet et al. and Constantin et al. have demonstrated that CB does not induce cell apoptosis and that the cytoskeleton of CB-treated cells can be recovered after removal of the CB.12,13 Pick et al. have most recently shown that the membrane functions of the parental cells are preserved in such CB-induced vesicles, comprising biologically active surface receptors and ion pumps.14 Inspired by these studies, we have achieved the first success in encapsulation of drugs and NPs in natural vesicles by using CB to treat cells Received: March 9, 2011 Revised: April 6, 2011 Published: April 12, 2011 2152
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Scheme 1. Graphic Depiction Showing the Use of Living Cells to Produce CMCs for Humanized Encapsulation of Drugs and Inorganic NPs
Figure 1. (a) CLSM image of the CMCs derived from HEK293 cells stained with carboxyfluorescein. Inset shows a high magnification CLSM image. (b) Release profiles of rhodamine 6G (9) and carboxyfluorescein (b), from CMCs vs time. The released fractions are obtained by dividing the amount of released dyes by the total amount of dyes encapsulated in CMCs.
laden with drugs and/or NPs (Scheme 1). Fluorescence microscopy revealed that upon CB treatment mouse fibroblasts (3T3 cells) with the cytoplasm stained by carboxyfluorescein shrank and released spherical capsules in the presence of trypsin under mechanical force (Figure S1a, Supporting Information). The strong green fluorescence of the resultant CMCs indicated the presence of the carboxyfluorescein and thus of cytosolic fluids of the parental cells (Figure S1b, Supporting Information). The hydrophobic membranes of the resulting CMCs were stained by 2-diphytanoyl-sn-glycero-3-phosphoethanolamine-N-(7-nitro-21,3-benzoxadiazol-4-yl) (NBD-PE) (Figure S1c, Supporting Information). Most CMCs (70%) were in the size range of 1 2 μm (Figure S1d, Supporting Information). A single 3T3 cell produced >30 CMCs. Given that hundreds of millions of cells can be harvested from a culture dish (9 cm in diameter), the present approach can be easily scaled up for production of billions of CMCs at relatively low cost. Similar CMCs were produced from human embryonic kidney cells (HEK293 cells) (data not shown). The CMCs were stable and well-dispersed in buffer solutions containing serum but tended to aggregate in serum-free buffer solution after 8 h (Figure S2, Supporting Information). This suggests that the proteins in the serum act as steric
stabilizers for the CMCs, which is usually expected for cell stabilization in biological medium.15 Carboxyfluorescein diacetate (cFDA) was used to assess cytoplasm function within the CMCs and membrane integrity. cFDA is a hydrophobic, nonfluorescent ester and can freely diffuse through the hydrophobic cell membrane. Following hydrolysis by esterase in the cytoplasm,16 cFDA is transformed into hydrophilic, fluorescent, carboxyfluorescein, which cannot penetrate the cell membrane. Figure 1a reveals that after incubation with cFDA, the CMCs (derived from HEK293 cells) became fluorescent. Thus, the esterase is enclosed within the CMCs and remains active. Furthermore, the capsule membrane remains intact; otherwise, leakage of hydrophilic substances into the aqueous media is expected. Only 8% of carboxyfluorescein was released out of the CMCs in 8 h, indicating a release rate of approximately 0.11 μg/(mL/h). No release was detected over the following 18 h period (Figure 1b). These observations demonstrate that the CMC membranes remain intact. In contrast, hydrophobic rhodamine 6G (inert to the esterase), encapsulated in CMCs, was released at a speed of 0.27 μg/(mL/h); 24.7% of encapsulated rhodamine 6G was released after 26 h. Similar release rates were observed for CMCs derived from 3T3 2153
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Figure 2. Cumulative release profiles of calcein (9, 0) and carboxyfluorescein (b, O) released from HEK 293-derived CMCs (a) and 3T3 cells (b). Dye release was implemented in Tris buffer (0, O) and Tris buffer bearing 10 mM ATP (9, b).
cells (Figure S3, Supporting Information). Hence, CMC membranes are permeable to hydrophobic, but not hydrophilic, molecules suggesting that the release rate is determined by the solubility of the dyes, their diffusion within the cytoplasm, and their ability to diffuse through the membranes. Cell membranes are composed of a variety of protein channels for transfer of ions and molecules across the membranes. For instance, tumor cells such as HEK293 cells usually have multidrug resistance proteins (MRP) overexpressed in their membrenes and form a channel to pump out a variety of anticancer drugs into the extracellular environement with the aid of adenosine triphosphate (ATP).17 In the current work, hydrophobic calcein acetoxymethyl ester (calcein-AM) was encapsulated in HEK293 cells, via diffusion across the cell membranes, induced by its hydrophobicity. The esterase in the cytoplasm hydrolyzed the ester to hydrophilic and fluorescent calcein. Calcein was therefore satisfactorily encapsulated in the CMCs derived from the cells. Calcein is a well-known probe to test MRP channels in cell membranes.18 Figure 2a shows the release profile of substances from the CMCs derived from HEK293 cells in Tris buffer at 37 C. Regardless of the presence of ATP, only 10% of the carboxyfluorescein, which did not interact with the MRD channels, was released from the CMCs in 26 h. In contrast, approximately 30% of calcein has been released in 26 h at a rate of 0.28 μg/(mL/h). The presence of 10 mM ATP causes a fast and profound release of calcein; approximately 80% of calcein has been released in the first 8 h at a rate of 1.8 μg/(mL/h) and the residual 20% in the following 18 h at a rate of 0.18 μg/(mL/h). This ATP-dependent calcein release could be also due to the fact that extracellular ATP at high concentration can result in an increase in cell membrane permeability to small molecules due to dissipation of the membrane potential.19 Accordingly, we have conducted release of calceins from the CMCs derived 3T3 cells, which have a much lower number of MRP channels compared with HEK293 cells. The presence of ATP noticeably increases the calcein release from the CMCs due to membrane permeability increase. But the release rate and the total release in a given time are much smaller than those from the CMCs derived from HEK293 cells (Figure 2b). This underlines the presence of MRP channels in CMCs derived from HEK293 cells and its significant role in the calcein release. The release of small amounts of calcein in the absence of ATP is attributed to the fact that other ATP sources were available in the cytoplasm of the CMCs. The present study should be the first report of harnessing the protein channels in cell membranes for controlled release of an encapsulated drug or molecular probe.
Inorganic NPs have been widely recognized as innovative diagnostic probes and/or drugs for biomedical applications due to their unique size-dependent electronic, photonic, magnetic, and surface chemical properties.20 23 However, renal clearance and macrophage clearance limit circulation kinetics for NPs in vivo.24 Here we exploit living cells for encapsulation of inorganic NPs. Two types of PEGylated, hydrophilic CdSe quantum dots (QDs), 5.3 and 5.9 nm in diameter, were prepared as described elsewhere;25 they exhibited photoluminescence emission bands centered at 522 and at 570 nm, respectively (Figure S4, Supporting Information). CdSe@PEG QDs were taken up by 3T3 cells via endocytosis and entrapped in the endolysosomes of the cells (Figures S5a and S6a, Supporting Information). The QDs egressed from the endosomes and lysosomes into the cytoplasm after treatment of the QD-loaded cells with chloroquine, a drug that induces significant swelling of endosomes and lysosomes26 (Figures S5b and S6b, Supporting Information). After CB treatment, the QDs dispersed in the cytoplasm were effectively encapsulated in the CMCs (Figure 3, panels a and b). When mixtures of differently sized QDs were used for cellular uptake, CMCs loaded with multiple QDs were obtained and their emission color was easily encoded by the molar ratios of the different QDs in the original aqueous dispersions (Figure 3c). Compared to the original QDs, the CMCs loaded with QDs showed little change in their emission bands and profiles (Figure 3d). Relying on the fluorescence spectra of the QD-loaded CMCs, we estimated that each CMC encapsulated about 30 QDs. Following a similar protocol, aqueous 14 nm gold NPs, stabilized by rhodamine-labeled poly(oligo(ethylene glycol) methacrylate),27 were also encapsulated in the CMCs with a yield of about 10 gold NPs per CMC (Figure S7, Supporting Information). Similarly, two kinds of CdSe QDs (with green and red color) and gold NPs were encapsulated in the CMCs derived from HEK293 cells (Figure S8, Supporting Information). Thus CMCs can be used to encapsulate a wide range of molecules and even nanoscale biolabels, and these can impart CMCs with multimodal functionality. Since the CMC membranes are almost identical to those of the parental cells and CB treatment does not cause apoptosis, it is useful to test whether CMCs act as “humanized” vehicles capable of evading immune attack. To this end, we studied the internalization of CMCs by the human macrophage cell line, human acute monocytic leukemia cell line (THP-1). Prior to this study, we had tested the viability of THP-1 macrophages in the presence of CMCs. As expected, the CMCs showed little cytotoxicity (Figure S9, Supporting Information). Uptake of 2154
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Figure 3. (a) CLSM fluorescence images (left panel) and overlay of CLSM transmission and fluorescence images (right panel) of 3T3 CMCs, loaded with 5.3 nm CdSe@PEG QDs. (b) CLSM fluorescence images (left panel) and overlay of CLSM transmission and fluorescence images (right panel) of 3T3 CMCs, loaded with 5.9 nm CdSe@PEG QDs. (c) CLSM fluorescence images of 3T3 CMCs, loaded with both 5.3 and 5.9 nm QDs. The fluorescence image in the left panel has been obtained via the green fluorescent channel to reveal the small QDs and the fluorescence image in the middle panel via the red fluorescence channel to reveal the large QDs. The overlay of these two images is shown in the right panel to reveal the coexistence of both small and large QDs. (d) The fluorescence spectra of original 5.3 nm (dashed, green curve) and 5.9 nm (dashed, red curve) CdSe QDs in PBS buffer and 3T3 CMCs loaded with 5.3 nm QDs (solid, green curve), 5.9 nm (solid, red curve), and both (solid, yellow curve) in PBS buffer. The excitation wavelength is 350 nm.
Figure 4. Macrophage internalization of CMCs loaded with PEGylated CdSe QDs (a and b) and PLGA particles loaded with CdSe@ODA QD (c). CMCs have been derived from HEK 293 (a) and 3T3 cells (b). PLGA particles and CMCs were incubated with human macrophage THP-1 for 1 h (upper panels) and 24 h (lower panels). Macrophage uptake was assessed by flow cytometry. Dead and living macrophages were differentiated by PI staining. (d) Histogram of fractions of fluorescence-positive THP-1 macrophages after incubation with CMCs and PLGA particles for 1 h (upper panel) and 24 h (low panel).
PEGylated CdSe QD-loaded CMCs derived from HEK293 and 3T3 cells by THP-1 macrophages is shown in panels a and b of Figure 4, respectively. A minute fraction of macrophages became fluorescent after 1 h of incubation with CMCs; 12.6% for the
CMCs derived from HEK293 cells and 17.3% for those derived from 3T3 cells (upper panel in Figure 4d). After 24 h of incubation, 30.9% of the macrophages became fluorescent for the CMCs derived from HEK293 cells and 38.3% for those 2155
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Nano Letters derived from 3T3 cells (lower panel in Figure 4d). Similar uptake behavior of macrophages was observed in CMCs loaded with fluorescent dyes. Poly(lactic-co-glycolic acid) (PLGA) particles, loaded with octadecylamine (ODA) capped CdSe QDs,28 were used as controls, representing conventional artificial carriers (Figure S10a, Supporting Information). In stark contrast to CMCs, PLGA particles were rapidly internalized by THP-1 macrophages (Figure 4c). 61.8% of the macrophages were fluorescent after 1 h of incubation with PLGA particles, and over 90% of macrophages became fluorescent after 24 h of incubation (Figure 4d). Lipid vesicles are another conventional class of commonly used artificial carrier, labeled here by NBD-PE (Figure S10b, Supporting Information). After 1 h of incubation with lipid vesicles labeled by NBD-PE, approximately 6% of the macrophages were fluorescent, which may be due to the fact that lipid vesicles are compositionally similar to the cell membranes (Figure S11a, Supporting Information). Nevertheless, 92% of the macrophages became fluorescent after 24 h of incubation (Figure S11b, Supporting Information). In summary, although CMCs cannot completely evade long-term macrophage surveillance, they are far more efficient than artificial carriers. The surfaces of CMCs were conjugated by Fc fragments to enhance macrophage recognition and internalization (Figure S12, Supporting Information). The resulting Fc-CMCs showed rapid macrophage internalization, that is, 45% of macrophages became fluorescent after 1 h of incubation with the capsules and close to 80% after 24 h of incubation (Figure S11, Supporting Information). In conclusion, we have developed an innovative encapsulation methodology—using living cells as factories to produce CMCs that can encapsulate a variety of drugs and NPs. The functions of the membranes and cytoplasms of parental cells are well preserved in CMCs and can be harnessed to provide an unprecedented mechanism for controlled loading and release of active substances, e.g., by using functional pumps such as MRP channels. CMCs are noncytotoxic and can effectively minimize recognition and internalization by macrophages, thus evading immune attack in the body. Hence the CMCs provide the first intrinsically biocompatible and functional drug delivery and release vehicles. Importantly, CMCs can be harnessed from a wide range of natural or recombinant cells, enabling them to be manufactured with tailored membrane and cytoplasm functionalities. On the other hand, CMCs are almost identical to cells without nuclei, but they are abiotic and intrinsically robust, allowing easy study of a number of biological processes occurring at the cellular level. Furthermore, they will also raise exciting new opportunities to modulate the global immune system and address generic immunological problems.29
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Supporting Information. Experimental details and additional characterization data. This material is available free of charge via the Internet at http://pubs.acs.org.
’ AUTHOR INFORMATION Corresponding Author
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’ ACKNOWLEDGMENT This work is financially supported by the Max Planck Society. D.W. and H.W. thank Deutsche Forschungsgemeinschaft (MO283/38-1) for financial support. T.L.N. thanks DEST for support through ISI Grant CG110036. Dr. Andreas Lankenau (Fraunhofer Institute for Biomaterials Engineering, Potsdam, Germany) is acknowledged for assistance with cell cultures. ’ REFERENCES (1) Hubbell, J. A. Science 2003, 300, 595–596. (2) Huang, S. L. Adv. Drug Delivery Rev. 2008, 60, 1167–1176. (3) Savic, R.; Luo, L. B.; Eisenberg, A. Science 2003, 300, 615–618. (4) Johnston, A. P. R.; Cortez, C.; Angelatos, A. S.; Caruso, F. Curr. Opin. Colloid Interface Sci. 2006, 11, 203–209. (5) Mundargi, R. C.; Babu, V. R.; Rangaswamy, V.; Patel, P.; Aminabhavi, T. M. J. Controlled Release 2008, 125, 193–209. (6) Kriebardis, A. G.; Antonelou, M. H.; Stamoulis, K. E.; Economou-Petersen, E.; Margaritis, L. H.; Papassideri, I. S. Transfusion 2008, 48, 1943–1953. (7) MacDiarmid, J. A.; Amaro-Mugridge, N. B; Madrid-Weiss, J.; Sedliarou, I.; Wetzel, S.; Kochar, K.; Brahmbhatt, V. N; Phillips, L.; Pattison, S. T; Petti, C.; Stillman, B.; Graham, R. M; Brahmbhatt, H. Nat. Biotechnol. 2009, 27, 643–651. (8) Noad, R.; Roy, P. Trends Microbiol. 2003, 11, 438–444. (9) Romberg, B.; Hennink, W. E.; Storm, G. Pharm. Res. 2008, 25, 55–71. (10) Cooper, G. M. The Cell: A Molecular Approach, 2nd ed.; Sinauer Associates Inc.: Sunderland, MA, 2000. (11) Sanger, J. W.; Holtzer, H. Proc. Natl. Acad. Sci. U.S.A. 1972, 69, 253–257. (12) Bousquet, P. F; Paulsen, L. A.; Fondy, C.; Lipski, K. M.; Loucy, K. J.; Fondy, T. P. Cancer Res. 1990, 50, 1431–1439. (13) Constantin, B.; Imbert, N.; Besse, C.; Cognard, C.; Raymond, G. Biol. Cell 1995, 85, 125–135. (14) Pick, H.; Schmid, E. L.; Tairi, A.; Llegems, E.; Hovius, R.; Vogel, H. J. Am. Chem. Soc. 2005, 127, 2908–2912. (15) Norde, W. In Bacterial Adhesion and Preventive Dentistry; Ten Cate, J. M., Leach, S. A., Arends, J., Eds.; IRL Press: Oxford, U.K., 1984; pp 1 17, (16) Morono, Y.; Takano, S.; Miyanaga, K.; Tanji, Y.; Unno, H.; Hori, K. Biotechnol. Lett. 2004, 26, 379–383. (17) Aller, S. G.; Yu, J.; Ward, A.; Weng, Y.; Chittaboina, S.; Zhuo, R.; Harrell, P. M.; Trinh, Y. T.; Zhang, Q.; Urbatsch, I. L.; Chang, G. Science 2009, 323, 1718–1722. (18) Feller, N.; Broxterman, H. J.; Wahrer, D. C. R.; Pinedo, H. M. FEBS Lett. 1995, 368, 385–388. (19) Arav, R.; Friedberg, I. FEBS Lett. 1996, 387, 149–151. (20) Alivasatos, P. Nat. Biotechnol. 2004, 22, 47–52. (21) Schultz, D. A. Curr. Opin. Biotechnol. 2004, 14, 13–22. (22) Medintz, I. L.; Tetsuo Uyeda, H.; Goldman, E. R; Mottoussi, H. Nat. Mater. 2005, 4, 435–446. (23) McCarthy, J. R.; Weissleder, R. Adv. Drug Delivery Rev. 2008, 60, 1241–1251. (24) Choi, H. S.; Liu, W. B.; Misra, P.; Tanaka, E.; Zimmer, J. P.; Ipe, B. I.; Bawendi, M. G.; Frangioni, J. V. Nat. Biotechnol. 2007, 25, 1165. (25) Lees, E. E.; Nguyen, T. L.; Clayton, A. H. A.; Mulvaney, P. ACS Nano 2009, 3, 1121–1128. (26) Mellman, I.; Fuchs, R.; Helenius, A. Annu. Rev. Biochem. 1986, 55, 663–700. (27) Edwards, E. W.; Chanana, M.; Wang, D. J. Phys. Chem. C 2008, 112, 15207–15219. (28) Jasieniak, J.; Mulvaney, P. J. Am. Chem. Soc. 2007, 129, 2841– 2848. (29) Kotov, N. Science 2010, 330, 188–189.
These authors contributed equally to this work. 2156
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