All-Aqueous-Phase Microfluidics for Cell Encapsulation - ACS Applied

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All-Aqueous Phase Microfluidics for Cell Encapsulation Kaixuan Zhu, Yunru Yu, Yue Cheng, Conghui Tian, Gang Zhao, and Yuanjin Zhao ACS Appl. Mater. Interfaces, Just Accepted Manuscript • DOI: 10.1021/acsami.8b19234 • Publication Date (Web): 16 Jan 2019 Downloaded from http://pubs.acs.org on January 16, 2019

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ACS Applied Materials & Interfaces

All-Aqueous Phase Microfluidics for Cell Encapsulation Kaixuan Zhua,c,†, Yunru Yub,†, Yue Chenga,†, Conghui Tiana, Gang Zhaoa,*, Yuanjin Zhaob,*

a

Department of Electronic Science and Technology, University of Science and

Technology of China, Hefei, 230027, China b

State Key Laboratory of Bioelectronics, School of Biological Science and Medical

Engineering, Southeast University, Nanjing, 210096, China c

School of Electrical and Information Engineering, Suzhou Institute of Technology,

Jiangsu University of Science and Technology, Zhangjiagang, 215600, China. * Author to whom correspondence should be addressed. †

These authors contributed equally.

E-mail: [email protected] (G.Z.); [email protected] (Y.Z.)

KEYWORDS: microfluidics; cell encapsulation; microcapsule; emulsion; hydrogel

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ABSTRACT: Cell-laden hydrogel microcarriers are widely used in diverse biomedical applications like 3D cell culture, cellular therapy and tissue engineering, where microcarriers were generally produced by using oil, which is the common but not optimal choice, as oil may cause cytotoxicity or protein denaturation. Here, an allaqueous phase microfluidics is presented to achieve oil-free emulsification of cell-laden microcapsules and 3D cell culture. Aqueous solutions with different concentration gradients are used as immiscible continuous phase and dispersed phase, and oscillation from a solenoid valve facilitates the formation of microcapsules in the water-water interface. By adjusting aqueous phases flow rates and oscillating frequency, core-shell microcapsules with controllable structure can be stably and continuously generated. In further 3D cell culture, encapsulated cells maintained good viabilities and aggregated together. These features show that the oil-free microfluidic method may has broad prospects in many biomedical applications.

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1. INTRODUCTION In recent years, hydrogel microcarriers with biocompatibility and semipermeable properties have attracted broad attention in the fields of materials chemistry and biomedical engineering.1-8 Hydrogels are generally made into micron-sized droplets or fibers through multiple methods like microfluidics, electrostatic spraying and ultrasonic crushing, which provide a mild microenvironment for scaffold-based 3D cell culture, drug delivery and other applications.9-17 The porous structure of hydrogel ensures the substances exchange between cells and the external environment, and it also protects the cells from mechanical damage and immune rejection.18-20 Among the diverse hydrogel microcarriers forms (capsule, fiber, rod-shaped, etc.), spherical microcapsules with core-shell structures have been widely-used biological carriers due to their easy fabrication process and universality.21,

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Core-shell microcapsules generally

consist of liquid cores containing cells or other biological samples and hydrogel shells which split samples into individual spatial units. Micron-sized core-shell microcapsules are widely used scaffolds and carriers for cell culture and delivery.2326

Their inner cores can be extracellular matrix (ECM), collagen and other

biomaterials while the outer shells can be changed to natural polysaccharides or synthetic polymers as needed.2,

27, 28

However, when using droplet-based

microfluidics to produce such hydrogel capsule, oil is often inevitably introduced as a continuous phase to help form droplets, which is undesirable in some biomedical cases(Some proteins and cells are sensitive to organic solutions), and 3



subsequent washing process may bring about pollution and time waste.29-31 Therefore, the oil-free method for stable and controllable fabrication of microcapsules becomes a new strategy. Unlike the immiscibility between oil and water, specific aqueous solutions can also become immiscible when their concentrations exceed a certain range. This feature was utilized for the preparation of water-water emulsifier.32-34 The waterwater emulsifier avoid direct contact between cells and oil phase, providing a mild biological microenvironment.35,

36

Therefore, water-water emulsification method

have been widely used in many biomedical applications like bio-macromolecule separation and cell enrichment.37-39 Water-water emulsifier can be used for the generation single aqueous phase droplet. Based on that, the generation of double aqueous phase core-shell capsules requires for a water-water-water emulsification system, in which three aqueous solutions are incompatible with each other. The allaqueous phase emulsification systems have been utilized to generate multi-layered extracellular matrix and porous scaffolds as microcarriers for cells, proteins and DNA.40-42 However, since the interfacial tension between the aqueous phase solutions is relatively small, it is difficult to generate water-water-water emulsification capsules with controllable size and uniformity by conventional droplet-based microfluidic devices, which has significantly hampered the promotion of all-aqueous phase emulsification method.43, 44 Therefore, stable and controllable fabrication of core-shell microcapsules using all-aqueous phase system remains a challenge. 4


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Here, we present a novel all-aqueous phase microfluidics for controllable generation of core-shell microcapsules. To facilitate the emulsification process, an external interference was introduced based on an oscillating solenoid valve on the channel of shell phase to control the injection of the fluid at different frequencies, then the microcapsules could then be generated stably and continuously. The diameter of the core and the thickness of the shell can also be adjusted by changing the flow rates of each phase and oscillating frequency. By exploring their performance in cell encapsulation and 3D cell culture, this all-aqueous phase emulsification method was proved to show potentials in more biomedical applications like cell transplantation, drug delivery and tissue engineering. 2. EXPERIMENTAL SECTION 2.1. Materials and Reagents. Sodium carboxymethylcellulose (C4888, Medium viscosity) were purchased from Sigma (USA).Polyethylene glycol (Mw.10000, Reagent Grade) and sodium alginate (A602116-0100, Reagent Grade) was obtained from BBI LIFE SCIENCE CORPORATION (China). Calcium chloride (analytically pure) were bought from Shanghai Hushi Laboratorial Equipment Corporation (China). Solenoid valve (PS-1615NO) was bought from TAKASAGO ELECTRONIC CORPORATION (China). The information of fluorescent dye would be explained when mentioned 2.2. Capillary microfluidic design. Three different sized capillaries were coaxially aligned to form the tube-in-tube microfluidic devices (Figure 1a). The tip diameters of the capillaries for core phase and shell phase were 180 and 400 μm, respectively. The 5



outermost capillary for continuous phase had an inner diameter of 1000 μm. Capillaries were connected with syringe pumps using connector and hoses with 4 mm inner diameter. The solenoid valve was mounted on the hose connecting with the shell phase. When the solenoid valve was closed, shell phase can flow normally; when the solenoid valve was opened, the clamp on the valve would block the flow. The outlet of the device was connected via a hose to a petri dish containing calcium chloride solution. Due to the fragility of the capillary, the entire device was glued to an acrylic plate for easy handling. 2.3. Cell culture. Porcine adipose-derived stem cells (pADSCs) were chosen for experiments and were obtained as a gift from Prof. Zhang’s laboratory.45 Cells were cultured with DMEM/F12 containing 10% FBS, 50 µg mL−1 vitamin C (Sigma, USA), 10 ng mL−1 basic fibroblast growth factor (Pepro Tech, USA) and 0.002 mol L−1 GlutaMAXTM-100× (Life Technologies, USA) at 37 °C in a 5% CO2 humidified incubator. Culture medium was changed every 2-3 d till cells reached 80–90% intensity. Cells were digested with 0.25% w/v trypsin-EDTA (Sigma, USA) for 3 min, and centrifuged at 1000rpm for 3 min. Then, they were resuspended in sodium carboxymethylcellulose solution for further use. 2.4. Generation of cell-laden microcapsules. Prior to the experiment, the microfluidic device was cleaned with alcohol (75%) and UV sterilized for 30 minutes, and all used solutions were aseptically processed. The syringe pumps were connected to the device through hoses, and sterile water was firstly introduced to the device for humidifying the channel. In a typical experiment, polyethylene glycol (30%, w/v) was 6


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pumped as the continuous phase with the flow rate of 100 µl/min; mixture of sodium alginate (1%, w/v) and dextran (15%, w/v) was pumped as shell phase with the flow rate of 1.5µl/min and cell-suspended sodium carboxymethylcellulose (1%, w/v) was pumped as core phase with the flow rate of 1.5µl/min. The oscillating frequency was 2Hz and could be further adjusted. When pumps started, generated capsules flow into the calcium chloride solution. The outer sodium alginate shell of capsules cross-linked with calcium ions, forming the final cell-laden hydrogel microcapsules. 2.5. 3D cell culture and immunofluorescence staining. Cell-laden microcapsules were collected from calcium chloride solution and transferred to culture medium. During one week of 3D culture, microcapsules were sampled at day 1, 3, 5 and 7 to test cell viabilities by using an AO/EB staining kit (KeyGen BioTECH Co., Ltd, China). 2 μL of prepared staining solution was added to the cell-laden microcapsules (about 5-10 capsules) and incubated for 3 min at room temperature. The sample was then placed under a fluorescence microscope for observation: the green fluorescence represented live cells while the red represented dead cells. Two

membrane

protein

markers

CD44

and

CD29

were

used

for

immunofluorescence staining. After one week of 3D culture, the cell aggregates were released from microcapsules by using sodium citrate solution (75mM). Alginate hydrogel shell of capsules would gradually degrade within 3 min with the addition of sodium citrate solution, and cell aggregates could be collected for further staining after being washed twice with PBS and transferred to a glass slide. Immunol Staining Fix Solution (Beyotime, Haimen, China) was added to the samples and incubated at 4 °C 7



overnight. The fixed cell aggregates were then washed with PBS for three times to remove extra fix solution. Afterwards, Immunol Staining Blocking Buffer (Beyotime, Haimen, China) was added and incubate for 1 h, followed by staining with the primary antibody (1:10 dilution) of mouse antihuman CD44 monoclonal antibody (Proteintech, Wuhan, China) and purified mouse antipig CD29 (BD Pharmingen, USA) at 4 °C overnight. The samples were then washed with PBS and incubated in secondary antibody coupled to Alexa Fluor 488 (1:50 dilution, Thermo Fisher Scientific, USA) for 1 h in the dark. Next, washed the sample with PBS, cell nuclei were stained by using DAPI (Beyotime, Haimen, China) for 10 min, followed by washing with PBS. Finally, samples were imaged by a laser confocal microscopy. 3. RESULTS AND DISCUSSION 3.1. The all-aqueous phase microfluidic system and core-shell microcapsules fabrication. The all-aqueous phase microfluidic system and a typical capsule generation process were shown in Figure 1. The microfluidic device had three inlets for three aqueous phase respectively: I1 for continuous phase, I2 for shell phase and I3 for core phase. All liquid was pumped into the device through syringe pumps. A solenoid valve was mounted on the hose that connects the shell phases to shut the channel periodically. In order to establish the all-aqueous phase emulsification system, polyethylene glycol (30%, w/v) was used as continuous phase, mixture of sodium alginate (1%, w/v) and dextran (15%, w/v) was used as shell phase, sodium carboxymethylcellulose (1%, w/v) was used as the core phase. When syringe pumps started, three aqueous phases flowed into the device at different flow rates, forming 8


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microcapsules with the help of shear force. The outlet of the device was connected to a calcium chloride solution filled container, and the sodium alginate layer of the capsule rapidly cross-linked with the help of calcium ions to form solidified hydrogel capsules. However, since the interfacial tension between aqueous phase solutions was small, it was difficult to control the droplet generation with the help of flow shearing force. The disperse phase (core phase and shell phase) preferred to flow in a “threading” mode, where the disperse phase flowed stably within a long distance before finally splitting into droplets. To generate droplets quickly and controllably, we introduced a solenoid valve that directly interferes with the shearing process of the droplet to divide the continuous phase into separate droplets (Figure 1b). When the valve was off, the shell phase liquid flowed normally, and when the valve was on, the shell phase channel was instantly shut and squeezed the fluids to form droplet, as recorded in Movie S1. After crosslinking in CaCl2 solution, solidified core-shell microcapsules were collected for further observation and statistical analyses (Figure 2). In a typical experiment, flow rates of core phase, shell phase and continuous phase were 1.5, 1.5, and 100 µl/min respectively, and the frequency of the oscillating valve was 2Hz. The resultant capsules showed great uniformity, and the capsule/core diameters were conformed to the standard normal distribution (Figure 2a-b). By calculating the core-shell diameters of 117 microcapsules, we obtained the capsule diameter of 360.1 ± 16.3 μm and core diameter of 250.5 ± 18.8 μm. The C.V. value of the capsule diameter shown good monodispersity of the capsules (C.V. value indicates the deviation of the capsule diameter to average value). By mixing with the nano9



fluorescent microbeads, the capsules with fluorescent cores could also be generated (Figure 2c-d). It could be found that the core and shell of capsules had clear boundaries, and no diffusion occurred between the two phases. 3.2. Effect of flow rate and oscillating frequency on microcapsule size. In our system, uniform core-shell capsules with varied diameters could be continuously produced by simply adjusting the flow rate of each phase and oscillating frequency (Figure 3). With the increase of inner solution flow rate, the core diameter and the entire capsule diameter increased while the shell thickness decreased (Figure 3a-c); by increasing the flow rate of shell phase, the capsule diameter and the shell thickness increased, but there is almost no effect on the core diameter (Figure S1, Supporting Information). The effect of the oscillating frequency on the size of the capsule was also tested at 1Hz, 3Hz and 4Hz, keeping the flow rate unchanged (Figure 3d-f). It could be demonstrated that the increase in oscillating frequency caused the decrease of capsule diameter, and both core diameter and shell thickness became smaller. To further study the effect of different factors on capsule size, the actual capsule changes under different flow rates and calculated the actual generation efficiency (defined as “output flow rate”) (Figure S2, Supporting Information). It can be found that the output core phase flow rate kept a similar trend with input core phase flow rate, which meant changing the core phase flow rate can effectively adjust the core diameter of capsules, thereby changing the relative shell thickness. Similarly, comparison of the input shell phase flow rate and output shell phase flow rate showed that with the increase of shell phase flow rate, the difference between the output flow rate and the 10


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input flow rate became larger (Figure S3, Supporting Information). This meant that changing the shell phase flow rate did not effectively change the thickness of the shell. The effect of oscillating frequency on the output flow rates of core phase and shell phase was also tested when the input flow rate of core and shell phase were both 1.5 µl/min (Figure S4, Supporting Information). The experiment results showed that with the increase of oscillating frequency, the output flow rate and input flow rate maintain good agreement. However, the increase in frequency resulted in an increase in generated capsules amount within the same period. Therefore, the capsule diameter was reduced as the total volume kept unchanged. 3.3. Cell encapsulation and 3D culture. Without introducing any oil phases, this simple all-aqueous system could offer good biocompatibility in producing cellencapsulated capsules and avoid the washing process. By mixing the cell suspension with core phase, cell-laden microcapsules could be generated and tested for the cell viabilities. Cell viabilities was calculated based on above 200 cells in each experiments and repeat at least for 3 times in the same conditions. The final data is the mean and standard deviation of 3 viabilities. The cell viability of fresh cells after encapsulation and release were observed and analyzed. The cell-laden microcapsule images in bright field and fluorescent field showed a great cell viability after encapsulation and following release, (Figure 4a-d), and there was no significant difference between fresh cells and encapsulated cells (Figure 4e). These suggested that our all-aqueous system had virtually no effect on cell viability and was highly biocompatible. Moreover, we compared the effects of the oil-water method and our water-water method on cell 11



activity, adherence and proliferation. Figure S5 showed that there was no significant difference in the effect of the water-water and oil-water encapsulation method on cell viability (98% to 94%), but the use of oil-water encapsulation methods requires several washes before subsequent 3D culture, which may result in a slight decrease in cell viability. Our water-water method does not require washing of the capsules and can be taken directly for 3D culture, which is more convenient. Cell attachment and proliferation of released cells after encapsulation by the two method were also tested (Figure S6 & S7). Similarly, there was no significant difference between fresh group, oil-water group and water-water group, indicating that our all aqueous method has competitive performance in cell encapsulation to traditional method. Due to the mild generation process and the 3D micro-structure, the generated microcapsules could provide a biomimetic microenvironment for cell proliferation and differentiation, as proved by the results of 3D culture of the cell-laden capsules (Figure 5). Unlike 2D cell culture, encapsulated cells grew in a spatial microenvironment and tended to form cell aggregate. During the one-week culture, the capsules were sampled every two days to tested cell activities, and the green florescence represented live cells. It could be shown that cells gradually aggregated into a cluster and maintained high activity (Figure 5a). In addition to the cell viabilities, we also detected protein markers on the surface of cell aggregates after one week of culture (Figure 5b). CD-44 and CD29 were selected as the marker for immunofluorescence staining because they were two common cell membrane proteins, which played an important role in cell migration and cell adhesion. Blue fluorescence showed nucleus (DAPI) and red fluorescence showed 12


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two marker proteins (CD-44, CD-29), demonstrating that the two markers were still positively expressed on the cell surface during 3D culture. 4. CONCLUSION In conclusion, motivated by the limitation of existing oil-involved droplet microfluidics, we have developed a novel all-aqueous phase microfluidic method for highly controllable generation of core-shell microcapsules by introducing an external oscillating valve. Core-shell microcapsules with uniform and controlled morphologies could be generated by simply adjusting the flow rated of each phase and oscillating frequency. Furthermore, cell encapsulation and 3D culture were also accomplished to show the great biocompatibility of the method. Our all aqueous phase microfluidic system provides with a controllable encapsulation method without using oil phase. As we can envision, this method can be further employed in organoid construction, drug delivery and other significant applications in biomedical engineering.

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ASSOCIATED CONTENT Supporting Information is available free of charge on the ACS Publications website at DOI: XXX. Movie S1: A typical generation process of core-shell micarocapsules. Figure S1: Effect of shell phase flow rate on capsule size; Figure S2: Comparison of the input core phase flow rate and the output core phase flow rate; Figure S3: Comparison of the input shell phase flow rate and the output shell phase flow rate; Figure S4: Effect of oscillating frequency on the output flow rates of core phase and shell phase. Figure S5: Effect of two encapsulation methods (oil-water and water-water) on cell viability. (a) Cell viability of encapsulated cells after releasing from capsules. (b) Florescent staining of cells after encapsulation and release from the capsules. Figure S6. Attachment of encapsulated cells after releasing from capsules generated by two methods (oil-water and water-water) compared with fresh cells. Figure S7. Proliferation of encapsulated cells after releasing from capsules generated by two methods (oil-water and water-water) compared with fresh cells. (a) Cell proliferation times of released cells in 72 hours. (b) Morphology of the cell proliferation in 72 hours.

AUTHOR INFORMATION Corresponding Author *Email: [email protected] (G.Z.); [email protected] (Y.Z.) Author Contributions

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K.X.Z., Y.R.Y. and Y.C. contributed equally to this work. The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript. Funding Sources This research was supported by the National Natural Science Foundation of China (Grant Nos. 51476160 and 51522302) Notes The authors declare no competing financial interest.

ACKNOWLEDGMENT This research was partially performed at the USTC Center for Micro- and Nanoscale Research and Fabrication.

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Figure captions

Figure 1. The all-aqueous phase microfluidic system. a) Schematic of the all-aqueous phase microfluidic system for the fabrication of core-shell capsules. b) Actual fabrication process of core-shell capsules with the help of solenoid valve.

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Figure 2. Size distribution and morphology of produced microcapsules. a-b) Size distribution of a) microcapsules and b) microcapsule cores. c-d) Microcapsules encapsulated with florescent nanoparticles in c) optic field and d) florescent field. The scale bar is 200 μm.

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Figure 3. Effect of flow rate and oscillation frequency on capsule size. a-c) Morphology of the capsules at different core phase flow rates. The flow rates of shell phase and continuous phase were 1.5 µl/min, 100 µl/min respectively, and the frequency of the oscillating valve was 1Hz. d-f) Morphology of the capsules at different oscillating frequency. The flow rates of core phase, shell phase and continuous phase were 1.5 µl/min, 1.5 µl/min and 100 µl/min respectively. g) Effect of core phase flow rate on capsule size, n=90. h) Effect of oscillating frequency on capsule size, n=88. The scale bar is 200 μm. 25



Figure 4. Effect of encapsulation on cell viability. a-b) Morphology of cell-laden microcapsules in optic and fluorescent fields. c-d) Morphology of cells after releasing from microcapsules in optic and fluorescent fields. e) Histogram of cell viability before and after encapsulation. The scale bar is 200 μm.

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Figure 5. 3D culture of cell-laden microcapsules and immunofluorescence staining of cell aggregates. a) Morphology of cell-laden microcapsules during one week of 3D culture. b) Immunofluorescence staining of cell aggregates after one week of 3D culture. Cell nucleus (blue) and two protein markers (red) were stained. The scale bar is 50 μm.

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Table of Contents (TOC)

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All-Aqueous-Phase Microfluidics for Cell Encapsulation - ACS Applied

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