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Lipid Stabilized Solid Drug Nanoparticles for Targeted Chemotherapy Zhipeng Zeng, Pengfei Zhao, Lixin Liu, Xiaohu Gao, Hai-Quan Mao, and Yongming Chen ACS Appl. Mater. Interfaces, Just Accepted Manuscript • DOI: 10.1021/acsami.8b07024 • Publication Date (Web): 19 Jul 2018 Downloaded from http://pubs.acs.org on July 19, 2018

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ACS Applied Materials & Interfaces

Lipid Stabilized Solid Drug Nanoparticles for Targeted Chemotherapy

Zhipeng Zeng,†, # Pengfei Zhao,†, # Lixin Liu,*,† Xiaohu Gao,*, ‡ Hai-Quan Mao, §,⊥ Yongming Chen*, † †

School of Materials Science and Engineering, Center of Functional Biomaterials, Key Laboratory of Polymeric Composite Materials and Functional Materials of Ministry of Education, GD Research Center for Functional Biomaterials Engineering and Technology, Sun Yat-sen University, Guangzhou 510275, China ‡ Department of Bioengineering, University of Washington, Seattle, WA98195, USA § Institute for NanoBioTechnology and Department of Materials Science and Engineering, Johns Hopkins University, Baltimore, MD 21218, USA ⊥

Department of Biomedical Engineering and Translational Tissue Engineering Center, Johns Hopkins University School of Medicine, Baltimore, MD 21287, USA

Keywords: Drug delivery; flash nanoprecipitation; nanoparticles; targeting; cancer therapy

ABSTRACT: Nanoparticle-based chemotherapeutics have gained widespread interest in medicine due to their tunable pharmacokinetics and pharmacodynamics. Various drug delivery vehicles have been developed including polymer, liposome nanoparticles, and some of them have already made clinical impacts. Despite these advances, drug payload of these formulations is limited (typically < 10%). Here, we report a general and scalable approach to prepare lipid-coated solid drug nanoparticles by combining flash nanoprecipitation and extrusion technique, which enables optimization of individual steps separately and flexibility in selection of nanoparticle surface functionalities. Using methotrexate as a model drug, the nanoparticles significantly outperformed free drug in tumor growth suppression.

Nanoparticle-based drug delivery systems are of significant interest in medicine over conventional forms of free drug molecules due to their unique advantages. By formulating drug molecules into nanoparticles, key parameters for efficient treatment such as drug solubility, stability, pharmacokinetics, interaction with the biological systems, side effects, and cell-specific targeting, are completely redefined, creating a new paradigm for therapeutic engineering. Indeed, biodegradable materials such as liposomes, polymers, and natural proteins are frequently used in drug formulation, and some of them are already in clinical uses.1-3 Despite these advantages, common limitations of these conventional drug delivery vehicles include low drug payload, in most cases falls below 10% (weight percentage),4-6 and poor reproducibility preventing downstream scale-ups. High drug loading capacity and large-scale reproducibility are highly desired for lowering cost, increasing drug bioavailability, reducing side effects, and clinical translation. In this context, a simple and scalable nanoparticle production platform, flash nanoprecipitation (FNP), has recently been reported by Prud’homme and co-workers.7,8 A typical FNP process uses a multi-inlet vortex mixer, where a water-miscible organic solvent containing drug and amphiphilic (e.g., a block copolymer) molecules is injected through one of the inlets and water is supplied through the others. Rapid ACS Paragon Plus Environment

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mixing of the streams in a confined microenvironment creates super-saturation of the drug molecules, leading to precipitation and formation of drug nanoparticles. It is believed that the hydrophobic domains of amphiphilic molecules co-precipitate with hydrophobic drugs forming the nanoparticle core, whereas the hydrophilic moieties of surfactants locate on the surface of the nanoparticle.9,10 Although the stabilizing agent is not always needed to produce uniform nanoparticles,11 they are important for long-term storage and drug delivery in vivo to provide colloidal stability and anchor point for targeting ligand conjugation. During the ultrafast mixing, however, it is technically difficult to promote nucleation of drug molecules in the core while confining the surfactants on the surface to form the ideal drug coresurfactant shell structure.12,13 Here, we report a new fabrication strategy by combining the powerful FNP technology with the wellestablished extrusion-based lipid coating chemistry that is rich in surface functionality. As shown in Scheme 1, solid drug nanoparticles are produced with FNP in the first step without a stabilizing reagent. In the second step, the nanoparticles with transient stabilities are immediately subject to membrane extrusion in the presence of lipid mixtures for surface coating and functionalization. By separating particle formation and surface treatment into individual steps, both processes can be optimized without compromise. Methotrexate (MTX) was used as a model drug to evaluate the two-step fabrication approach. In the current study, the concentration of MTX in N-methyl pyrrolidone and dimethylformamide mixture (v/v 3:7) was fixed at 4 mg/mL. At a flow rate of 6 mL/min, dynamic light scattering (DLS) showed that the particle size could be readily tuned by varying the volume ratio of the organic solvent phase and the water phase (Figure 1A). At the volume ratio of 1/9, the MTX nanoparticles (MTX NP) reached a compact size of 49.1 ± 2.1 nm. This is an excellent size range for both in vitro cell uptake and in vivo tissue penetration. Previous research has shown that particles of approximately 50 nm in diameter have the highest rate of endocytosis,14,15 whereas under in vivo uses, compact nanoparticles penetrate tissues significantly deeper than large nanoparticles (except for tissues that are hyper-permeable).16 It is worth mentioning that similar compact particle formation was also achieved using other drug molecules such as doxorubicin and chlorambucil to demonstrate broad applicability of this approach (Figure S1). We have reason to believe that the hydrophilic segments of these three naturally amphiphilic drugs endowed the drug NPs with transitory stability,17 which provided sufficient time for the lipid coated procedure. The resulting drug nanoparticles were subjected to lipid coating for colloidal stability via repeated extrusions through a porous membrane. The lipid-coated MTX nanoparticles (MTX NP@lipid) exhibited a size increase of approximately 10 nm and a zeta potential change from neutral to -10 mV due to the negatively charged lipid molecules (Figure 1B). Further confirmation of successful lipid coating can be

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visualized in the transmission electron microscopy (TEM) images (Figure 1D). As the nanoparticle sample solutions dried on TEM grids, some of them tend to cluster randomly. Without lipids on the surface, the solid drug nanoparticle clusters showed no gap between adjacent particles. In contrast, for MTX NP@lipid, a small gap was always observed for adjacent particles. This thin lipid layer is critical for colloidal stability over long-term storage (Figure 1E). DLS revealed that MTX NP@lipid showed negligible size change in buffers after 7 days. In contrast, MTX NP gradually grew into large agglomerates, likely due to aggregation and Ostwald ripening.18 For comparison, MTX and the same amphiphilic lipids were also used in the conventional FNP process where MTX and lipids were fed into the micromixer in a single step through two of the four inlets (Figure 1C). Stable nanoparticles were also obtained, with slightly increased particle size and size distribution (71.1 ± 10.4 nm). Major differences between the conventional single-step process and the sequential two-step process, however, are the drug loading efficiency and capacity. Under the sequential two-step process, the drug loading efficiency and payload improved from 39.2% to 84.5% and from 30.5% to 62.3%, respectively. This is understandable since it is challenging to completely synchronize the precipitation rates of the hydrophobic drug and coating rate of the lipids under rapid mixing to promote formation of core-shell structures. Next, we tested the drug release profile of MTX NP@lipid (Figure 1F). While an initial burst release was observed for both bare and lipid-coated MTX nanoparticles, the lipid-coated ones showed a sustained release curve over 24 h. Interestingly, the release profile was also pH-responsive. Reducing the pH from 7.4 to 5.5 accelerated the release rate by some extent. This pH-dependent behavior likely arises from phase reorganization of the lipid molecules and subsequent destabilization of the lipid shell as previously reported,19 and can be beneficial to drug delivery to tumors because tumor microenvironment often is slightly acidic.20-22 To characterize cell uptake and cytotoxicity, a human breast cancer cell line, MCF-7, was treated with free MTX, MTX NP@lipid, or FA-lipid coated MTX nanoparticles (MTX NP@lipid-FA, Figure 1B shows its size profile) at the same MTX concentration. For visualization, the MTX molecule was conjugated to a fluorescent dye, sulfo-Cyanine7 NHS ester (Cy7). Semi-quantitative fluorescence microscopy revealed a time- and dose-dependent cell uptake for all three treatment groups. At 30 min incubation time and the same MTX-Cy7 concentration determined by fluorescence intensity for example (Figure 2A, C), MTX NP@lipid-FA showed small but meaningfully higher uptake than MTX NP@lipid (1.1 fold), which can be attributed to enhanced endocytosis of compact nanoparticles14 and the ligand-receptor interaction. The MTX celluar uptake was further increased to 1.5 fold after 2 h incubation (Figure 2B, C). The internalized MTX competitively inhibits dihydrofolate reductase and ACS Paragon Plus Environment


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consequently inhibits DNA synthesis leading to cell apoptosis.23 Indeed, a similar trend of dosedependent antitumor effect with specific cell targeting was observed for all three treatment groups compared to the untreated control (Figure 2D, E), with MTX NP@lipid-FA being the most effective. For example, cells treated for 48 h at a MTX concentration of 120 µg/mL showed reductions of cell viability from 92.1% (untreated) to 57.2% (free MTX), 50.8% (MTX NP@lipid), and 30.8% (MTX NP@lipid-FA) based on flow cytometry with Annexin-V labeling (Figure 2F). The antitumor effects of the three MTX agents were further evaluated under in vivo conditions using mice implanted with MCF7 tumors. Mice were given saline, free MTX, MTX NP@lipid, or MTX NP@lipid-FA via tail vein injection every 3 days at a dose of 10 mg/kg. As shown in Figure 3A, for mice treated with free MTX, the average tumor size was 59% smaller (867 ± 320 mm3) than those in the saline treatment control group (2,139 ± 1,032 mm3). In comparison, the MTX NP@lipid, and MTX NP@lipid-FA were more effective in tumor growth inhibition, with average tumor sizes of 71% and 88% smaller, respectively, leading to substantially longer survival (Figure 3B). The tumor suppression effect was further confirmed with an ex vivo molecular assay, TUNEL, characterizing the degree of cell apoptosis (Figure 3C). Tumor sections from mice treated with MTX NP@lipid-FA (48 h after the first therapeutic administration) showed high-density apoptotic cells, in contrast to mice treated with free MTX, and MTX NP@lipid. Another key advantage of lipid-coated MTX NP over free drug is the reduced side effect. Throughout the treatment course, animal weight loss and behaviors were routinely monitored. Mice treated with MTX NP@lipid and MTX NP@lipid-FA showed persistent body weight, and undetectable of tissue damage (Figure S2 and Figure S3). In contrast, free MTX caused weight loss, hunchback, and drowsiness in mice. Histology study also indicated damage to the heart (tissue atrophy and disordered patterns of myoneme) and kidney (cell karyopyknosis and nephron atrophy). To understand the reasons for the excellent tumor growth suppression effect of MTX NP@lipid-FA, biodistribution and preliminary pharmacokinetics of two MTX nanoparticle formulations were examined. MTX was labeled with Cy7. Non-invasive imaging revealed systemic distribution of MTX NP@lipid and MTX NP@lipid-FA within the first 4 h post administration, with gradual accumulation of the nanoparticles in the tumors (Figure 4A). After 8 h post injection, MTX NP@lipid and MTX NP@lipid-FA produced the highest ratios of tumor accumulation over the systemic background. Major organs (heart, liver, spleen, lung, and kidney) and tumor were isolated 8 h post administration for ex vivo quantitative fluorescence imaging (Figure 4B, 4C). MTX NP@lipid-FA accumulation in the tumor was 2.1-fold higher than MTX NP@lipid, which explained the differential therapeutic effects. The plasma circulating half-life for free MTX is 0.59 h and those for MTX NP@lipid and MTX NP@lipidFA were prolonged to 2.66 and 2.78 h (Figure 4D), respectively. MTX NP@lipid-FA exhibited


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extended drug accumulation and retention in the tumor region compared to the non-targeted nanoparticles. In conclusion, we have developed an effective approach to prepare lipid-coated solid drug nanoparticles by combining FNP and the lipid extrusion technique in a sequential manner, which allows optimization of both steps separately. FNP enables scalable fabrication of compact solid drug nanoparticles, whereas the rich chemistry of lipids and lipid derivatives offers flexible selection of targeting ligands. Using chemotherapeutic drug, MTX, as a model, we showed that nanoparticles of various sizes could be made and stabilized with lipids. When applied to MCF-7 cells in vitro, moderate improvement in antitumor activity was observed for drug nanoparticles compared to free drug. Under in vivo conditions, significantly higher therapeutic effect for MTX NP@lipid-FA was found over the free drug due to improved pharmacokinetic and biodistribution profiles. With further development, we expect this dual-step drug nanoparticle preparation process can become a general approach for nanoparticle drug production and have immediate impact on clinical translation.

ASSOCIATED CONTENT Supporting Information This Supporting Information is available free of charge on the ACS Publication website. Materials, instruments, experimental sections, and supplemental figures.

AUTHOR INFORMATION Corresponding Author *Xiaohu Gao. E-mail: [email protected] *Yongming Chen. E-mail: [email protected] *Lixin Liu. E-mail: [email protected]

Author Contributions #

Zhipeng Zeng and Pengfei Zhao contributed equally to this work.

Notes The authors declare no competing financial interest.

ACKNOWLEDGMENT This work was supported by the Natural Science Foundation of China (No. 51533009), Guangdong Innovative and Entrepreneurial Research Team Program (No. 2013S086), Science Foundation of Guangdong Province (No. 2014A030312018), and China Postdoctoral Science Foundation Project (2017M612803) for financial supports.

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Scheme 1. Schematic illustration of the fabrication procedure of lipid stabilized solid drug nanoparticles.



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Figure 1. Characterization of MTX NP and MTX NP@lipid. (A) Size control of MTX NP through regulating organic phase (O) /water (W) volume rate. (B) Size and surface potential of MTX NP, MTX NP@lipid and MTX NP@lipid-FA. (C) Size distribution of MTX NP@lipid prepared by one-step and two-step method. (D) TEM images of MTX NP, MTX NP@lipid and MTX NP@lipid-FA (scale bar = 500 nm). (E) Size stability of MTX NP, MTX NP@lipid in different solutions. (F) Drug release of MTX NP@lipid at different pH values (5.5 and 7.4) at 37 °C, compared with free MTX and MTX NP.


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Figure 2. Cellular uptake and cytotoxicity of free MTX, MTX NP@lipid and MTX NP@lipid-FA. MTX NP@lipid and MTX NP@lipidFA internalized by MCF-7 cells after (A) 0.5 h and (B) 2 h incubation (cell nuclei were stained by DAPI; Green represented for intracellular lysosomes/endosomes; Red represented for Cy7 fluorescence). (C) Quantitative analyzing cell uptake of MTX NP@lipid and MTX NP@lipid-FA. MTX were labeled with by Cy7. **P < 0.01. Cell viability of (D) MCF-7 cells and (E) A549 cells after 48 h incubation of free MTX, MTX NP@lipid and MTX NP@lipid-FA. (F) Cell apoptosis of MCF-7 cells induced by 48 h incubation of free MTX, MTX NP@lipid and MTX NP@lipid-FA. Annexin-V+ and PI+ cells were defined as late apoptotic/necrotic cells; Annexin-V+ and PI- cells were defined as early apoptotic cells.


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Figure 3. In vivo antitumor effect of MTX NP@lipid-FA to breast tumor-bearing mice. (A) Mouse tumor growth after intravenous injection of saline, free MTX, MTX NP@lipid and MTX NP@lipid-FA. Drugs were injected through tail vein at day 0, 3, 6, 9, 12 and 15. *P < 0.05; **P < 0.01 (n = 6). (B) Mouse survival after different treatments. (C) Images of mouse tumors sections with H&E and TUNEL staining (Yellow arrowhead represented for cell lysis, and blue arrowhead represented for cell pyknosis; green fluorescence represented for apoptotic cells). Mouse tumors were collected 48 h after first-time injection of different drugs.


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Figure 4. In vivo biodistribution of MTX NP@lipid and MTX NP@lipid-FA in tumor-bearing mice after intravenous injection. (A) In vivo fluorescence images showing the distribution of saline, free MTX, MTX NP@lipid and MTX NP@lipid-FA at different time points post-injection. (B) Ex vivo images of mouse hearts (he), livers (li), spleens (sp), lungs (lu), kidneys (ki) and tumors (tu) 8 h after injection of (a) saline, (b) MTX NP@lipid and (c) MTX NP@lipid-FA. (C) Semiquantitative fluorescence intensity of collected organs and tumors after 8 h injection of free MTX, MTX NP@lipid and MTX NP@lipid-FA (n = 3). **P < 0.01, compared with MTX NP@lipid; ##P < 0.01, compared with free MTX. (D) Pharmacokinetics of free MTX, MTX NP@lipid and MTX NP@lipid-FA in mouse blood, detected by HPLC (n = 5).


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