Oil-free Acoustofluidic Droplet Generation for Multicellular Tumor

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Oil-free Acoustofluidic Droplet Generation for Multicellular Tumor Spheroid Culture Jacqueline A. De Lora, Frank A. Fencl, Aidira D. Y. Macias Gonzalez, Alireza Bandegi, Reza Foudazi, Gabriel P Lopez, Andrew P. Shreve, and Nick J. Carroll ACS Appl. Bio Mater., Just Accepted Manuscript • DOI: 10.1021/acsabm.9b00617 • Publication Date (Web): 09 Aug 2019 Downloaded from pubs.acs.org on August 12, 2019

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ACS Applied Bio Materials

Oil-free Acoustofluidic Droplet Generation for Multicellular Tumor Spheroid Culture Jacqueline A. De Lora1#, Frank A. Fencl1#, Aidira D.Y. Macias Gonzalez1#, Alireza Bandegi2, Reza Foudazi2, Gabriel P. Lopez1, Andrew P. Shreve1,*, Nick J. Carroll1,* 1Department of Chemical and Biological Engineering and Center for Biomedical Engineering, University of New Mexico, Albuquerque, NM, USA 2Department of Chemical and Materials Engineering, New Mexico State University, Las Cruces, NM, USA *Equal Contributors Corresponding Authors *E-mail: [email protected] (A.P.S.) *E-mail: [email protected] (N.J.C.)

Keywords: ATPS, cancer, spheroid, acoustics, microfluidics, hydrogels, capillary

Abstract We present an easy-to-assemble microfluidic system for synthesizing cell-loaded dextran/alginate (DEX/ALG) hydrogel spheres using an aqueous two-phase system (ATPS) for templated fabrication of multicellular tumor spheroids (MTSs). An audio speaker driven by an amplified output of a waveform generator or smartphone provides acoustic modulation to drive the breakup of an ATPS into MTS template droplets within microcapillary fluidic devices. We apply extensions of Plateau-Rayleigh theory to help define flow and frequency parameter space necessary for acoustofluidic ATPS droplet formation in these devices. This method provides a simple droplet microfluidic approach using off-the-shelf acoustic components for quickly initiating MTSs and subsequent 3D cell culture.

Introduction Multicellular tumor spheroids (MTSs) are 3D clusters of cancer cells that act as model systems to recapitulate avascular tumor environments. However their use to replace 2D cell culture approaches, which are widely recognized to lack critical aspects in accurately modeling in vivo tumor environments1, has not been widely implemented. MTS culture techniques face difficulties with production of consistently sized spheroids, ease of culture, maintenance, and analysis of cellular responses.2 Furthermore, conventional MTS initiation methods can take up to

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two weeks to produce populations amenable to handling. 3 These technical limitations hinder widespread use of MTS for in vitro biological assays. A method to reduce the preparation time required for MTS formation is to encapsulate and crowd multiple cancer cells in microdroplets or within hydrogel microspheres.4 Aqueous two-phase systems (ATPSs) of immiscible polymer solutions—an aqueous droplet or gel solution encapsulating cancer cells, surrounded by a waterbased continuous fluid—are suitable for MTS templates. Because ATPS microfluidic droplet technology is oil-free, it significantly simplifies culture preparation, control over cell aggregate formation, maintenance, delivery of reagents, as well as optical and biochemical analysis of MTSs.5-7 Various acoustofluidic devices have been implemented to load single cells into aqueous drops8 or to form MTS assemblies within microchannels 9. Previous work also exploited robotic microprinting technology to deposit one MTS that was formed using ATPSs in each well of a 384 microwell plate.10 Robotic microprinting is well-suited for high-throughput drug compound screening, but may not be easily accessible to researchers seeking an entry-level method for laboratory scale ATPS-based 3D cell culture. We therefore aim to develop a microfluidic platform using simple components that are easy to assemble for generating tens of thousands of ATPS templated MTSs per hour. The low interfacial tension of ATPS11-13 requires precise control of low flow rates within a narrow range for “passive”, spontaneous droplet formation to occur.14-19 By contrast, flow stream perturbations at specific frequencies induced by acoustic or mechanical energy can be used to form ATPS droplets within “active” microfluidic systems, enabling a more diverse set of flow conditions where droplet formation can occur.20-27 Droplet formation frequency in these active ATPS microfluidic devices typically ranges from 5-50 Hz, which overlaps the lower audible frequency range.17 Based on this observation, our goal is to use off-the-shelf acoustic components to assemble an easily constructed active microfluidics system for ATPS droplet generation. For ease of implementation, we choose glass capillaries for device construction and provide driving frequencies to the audio components using a smartphone function generator freeware app. Our design is guided by a Plateau-Rayleigh (PR) instability analysis, which predicts spontaneous (passive) droplet formation at various flow rates as a function of natural perturbation frequency, solution viscosities, and interfacial tension. Although spontaneous droplet formation is not observed in the absence of an acoustic perturbation (an observation in agreement with another study23), we do find a strong correlation between the experimental driving frequencies imposed by the audio speaker and the predicted P-R perturbation frequency where spontaneous droplet formation occurs. We use this fluidic apparatus to form template MTS drops (achieving droplet diameter of ≈100 µm) containing


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dextran, gel-forming alginate and cancer cells, dispersed in a poly(ethylene glycol)-containing aqueous carrier fluid. From these drops we form alginate hydrogel microspheres using a calcium crosslinker and subsequently observe mouse carcinoma cell proliferation with development of 3D cell-cell contacts representative of MTSs. The versatility and ease of use of this platform will make ATPS droplet and MTS technologies readily accessible to a variety of labs, enabling biomedical applications of 3D cell culture such as drug discovery, recapitulation of tumor environments, and organoid formation.2,28-30 Experimental Section ATPS Chemicals, Phase Enrichment, and Refractometry The process of preparing the ATPS for cell encapsulation and culture is illustrated in Figure S1. We prepare solutions (all components purchased from Sigma Aldrich) for ATPS acoustofluidic droplet generation using a physiological buffer to enhance cell viability during the droplet formation process. The buffer, referred to as HEPES buffer, is composed of Millipore water, 25 mM HEPES, and 150 mM NaCl, pH adjusted to 7.4. Dissolving 5% w/w dextran (MW ≈ 550000 Da) and 17% w/w poly(ethylene glycol) (MW ≈ 8000 Da) into 100 mL aliquots each of HEPES buffer produces the initial components of the ATPS. To assure complete dissolution of the polymers, the solutions are vigorously stirred on a magnetic plate for approximately 3 hours. The dextran and poly(ethylene glycol) phases are then added to a glass separatory funnel, mixed into a turbid suspension, and clamped in an upright position for 48 hours at room temperature. Within the 48 hours, the dextran and poly(ethylene glycol) solutions reach thermodynamic equilibrium and a distinct interface is observed between two phases, one enriched in dextran and one enriched in poly(ethylene glycol). For brevity, these equilibrated enriched solutions are referred to as DEX and PEG, respectively, and have an approximate 1:9 volume ratio. The DEX and PEG are carefully collected from the separatory funnel while maintaining purity of the two phases by discarding ~2 mL of the ATPS obtained near the interface. Because the final total volume fractions shift from the initial total volume fractions, we verify that the bottom enriched phase is primarily concentrated dextran and the top phase is enriched in poly(ethylene glycol). The DEX composition is estimated by measuring the index of refraction of the bottom phase and comparing to a standard curve. The solutions for the standard curve are prepared by mixing 5%, 10%, 20%, and 30% w/w dextran into HEPES buffer as described above. The refractive indices of the standard curve solutions and the enriched DEX are measured (n=5) using a Kruss, DR6200-TF Refractometer (Figure S2).



The DEX is further processed to achieve a cytocompatible solution that contains a gelation agent for encapsulation of cells into hydrogels within the device. We dissolve sodium alginate (ALG) into DEX to a final concentration of 5 mg/mL and vigorously stir for 24 hours. The gelation bath is composed of HEPES buffer with 1M CaCl2, which serves as the source of Ca2+ to ionically crosslink the alginate polymer31-34 within the DEX droplets. The DEX/ALG, PEG, and gelation bath solutions are sterilized for cell culture by filtering using a 0.2 µm Nalgene bottle top vacuum filter.

Preparing Cell Suspension in DEX/ALG for Acoustofluidic Droplet Generation EMT6 mouse mammary carcinoma cells are maintained using standard sterile cell culture conditions (37C, 5%CO2) in 150 mm tissue culture treated polystyrene dishes (Corning) with 20 mL of Minimal Essential Media supplemented with 10% Cosmic Calf Serum (Hyclone) and 1% Penicillin/Streptomycin (Sigma-Aldrich) to expand the population for encapsulation. In general, if the desired cell/droplet occupancy is 10 cells/100 µm diameter droplet then 2x10 7 cells/mL are required for the encapsulation protocol. Cells at ~80% confluence in 150 mm dishes are washed with 3 mL of 0.5% Trypsin +EDTA (Sigma-Aldrich) followed by incubation in 5 mL of 0.5% Trypsin +EDTA for up to 20 minutes to detach cells. The cells are then washed off the dish with gentle pipetting by adding 5 mL of media to the 5 mL of trypsin and cells. This 10 mL cell suspension is transferred into a 50 mL centrifuge tube and combined with cells recovered from additional 150 mm dishes, the cell sample is counted using a Z Series Coulter Counter, and centrifuged at 1500 RPM for 10 minutes to pellet. The supernatant is aspirated and the cell pellet is suspended into 20 mL of HEPES buffer with an additional 1 mM EDTA to sequester free Ca2+. The suspension volume corresponding to the desired final cell concentration is transferred to a fresh 50 mL centrifuge tube. This suspension volume is then centrifuged at 1500 RPM for 10 minutes to wash and pellet. The supernatant is aspirated and the cell pellet is resuspended into 500 µL of sterile DEX/ALG by gentle pipetting using a 1000 µL pipette. The 500 µL of DEX/ALG cell suspension is drawn up into an 18G needle/10 mL syringe (BD) and added dropwise to 2.5 mL of DEX/ALG in a 50 mL centrifuge tube while vortexing. The 3 mL cell suspension is passed through a 40 µm cell strainer test-tube filter insert (BD Falcon) into a fresh 50 mL centrifuge tube to produce the final single cell suspension in DEX/ALG. A sample is taken from this solution to count the final cell concentration as there is a small percentage of cell loss during the protocol due to the highly-viscous DEX/ALG and filtering out of any cell clumps. This dropwise-vortex mixing protocol and filtering of the DEX/ALG/cells promotes a single cell suspension, which reduces clogging in the microfluidics device. The DEX/ALG/cells


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suspension is gently drawn up into a 10 mL syringe for delivery to the droplet-generating stream in the acoustofluidics system.

Acoustofluidic Device Fabrication and Droplet Generation The design of the acoustofluidics device emphasizes simple fabrication and operation of reusable devices. As detailed in the following description, the device is designed in a coflowing fluidic geometry using round capillaries with defined nozzles, square capillaries, glass slides, needle injection ports, epoxy, and laboratory tubing. The defined nozzles are fabricated by using a Sutter Instrument Co. P-97 Micropipette puller to pull two, round borosilicate glass capillaries (World Precision Instruments, 1.0 mm OD, 0.58 mm ID). The two tapered capillaries are then filed with fine grain sand paper to produce either the injection nozzle (100 µm ID, 4.5 cm length) or collection nozzle (200 µm ID, 8.5 cm length), respectively. The device is constructed on the foundation of two glass slides (25 mm x 75 mm x 1 mm, Swiss plain microscope slide, precleaned, Fisherbrand) that are scored, broken, and joined with epoxy to achieve a final length of 11.3 cm. Although the microcapillaries used to assemble the microfluidics devices are fabricated in house as just described, we note that commercially available tapered capillaries with a defined nozzle diameter could be used, further simplifying device construction. The injection and collection nozzles are nested into a borosilicate square capillary (Harvard, 1.5 mm OD, 1.05 mm ID, 8.5 cm length) with the distance between the injection and collection nozzles being ~1.5x the ID of the injection capillary. The capillary fluidic channels are attached to the foundation with 5 Minute Epoxy (Deycon Epoxy) and nozzles are concentrically aligned using a Zeiss Axio microscope at 10X magnification before the epoxy cures. Blunt end needles (Probe Needles, M919) are fixed at the foundation with the two ends of the square capillary by cutting notches into the plastic needle housing to snugly cover the capillaries. These injection/collection ports are sealed by coating with epoxy. The port downstream of the nozzles is connected to a 10 mL plastic syringe (Becton Dickinson) to close the system and prime the device with Millipore water. The collection capillary is extended by attaching the device to another capillary using 1.5 cm length of Tygon tubing (Tygon E-3606, 2.38 mm OD, 0.79 mm ID). The additional 15.3 cm long capillary has a 90 elbow bent into it at ~13 cm by heating the capillary over an open flame and manually bending the glass. This bend enables the end of the collection capillary to submerge into an upright scintillation vial containing 20 mL of the gelation buffer. The DEX/ALG and PEG are injected using two syringes (Becton Dickinson, 10 mL or 60 mL plastic syringes, respectively) with blunt end needles connected to PE tubing (Scientific Commodities, Inc. Micro Medical, LDPE 1.32 mm OD, 0.86 mm ID). The fluids are injected using two syringe



pumps (Harvard Apparatus PhD 200), 74-80 cm from the microscope stage. PEG is injected through the needle port established at the entry to the square capillary, and DEX/ALG is injected by attaching the tube from the syringe pump directly to the injection circular capillary. The acoustic components of the design include an audio speaker (removed from the plastic housing of a computer speaker to expose the surface of the speaker as well as the leads), an amplifier (Fosi Audio, 50W 4 ohms, 20 Hz - 20 kHz, 0.04% THD Stereo Audio Amplifier Mini Hi-Fi Professional Amp for Home Speakers), and a sinusoidal waveform source (either an Agilent Technologies 33250A Function/Arbitrary Waveform Generator or a smartphone with a free function generating application such as f Generator V4.4.0 by EE Toolkit PRO). The output of the waveform generator is connected to the amplifier by exposing the leads of a BNC cable and splicing them into the signal input on the amplifier. In general, from the waveform generator we use a sinusoidal waveform between 10-60 Hz at a 10 Vp-p amplitude setting. In the smartphone version of our setup, an audio jack cable is stripped to expose the leads and is similarly spliced into the signal input on the amplifier. In each case, the signal output on the amplifier is then connected directly to the audio speaker. The volume dial on the amplifier is set to ~half of the maximum allowable, as required to observe droplet generation. Under these conditions, the amplified waveform measured at the speaker is not clipped or otherwise distorted and we estimate that the power input to the speaker is a few tenths of a Watt for these typical settings. For acoustofluidic droplet generation, the DEX/ALG injection tubing is attached to the speaker using laboratory tape approximately 10 cm from the microscope stage. Droplet generation is initiated by setting the flow rates of the DEX/ALG (100-400 µL/Hr) and PEG (5500 µL/Hr) on the syringe pumps and commencing fluid flow into the device. A DEX/ALG jet flows out of the injection nozzle and into the collection nozzle surrounded by PEG outer fluid in a core/annular fluid profile. We then turn on the speaker and use frequencies stepped in 5 Hz intervals from 10 Hz-60 Hz for each DEX/ALG flow rate. We image the formed droplets 5 cm downstream of the nozzles using an Andor Technology camera (Model No. DL-658M-TIL). Microscopy images are collected across the parameter space described above and analyzed using ImageJ for dispersed phase configuration (e.g., mono/polydisperse droplet, no droplet formation, etc.) as well as droplet diameter. We investigate three different independently constructed devices without cells and one device with cells (Figure S3 using waveform generator actuation and Figure S4 using smartphone actuation).


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Plateau-Rayleigh Analysis Analysis of stability of the co-flowing DEX inner stream contained within the PEG outer stream was carried out using previously published methods. 35 Briefly, in the exit channel of the fluidic device, the inner (outer) stream is flowing at a volumetric rate of 𝑄𝑖 (𝑄𝑒 ), the viscosity of the inner stream is 𝜂𝑖 and that of the outer stream is 𝜂𝑒 . A summary of parameters (e.g. solution viscosities, interfacial tension, flow rates, and device/flow stream dimensions) for device operation and the modeling that relates these parameters is reported in Table 1. The rheological measurements to obtain viscosities are reported in Figure S7b. Following Guillot et al.35, we calculate:

𝑥=

𝑟𝑖0 𝛼−1 =√ 𝑅𝑐 (1/𝜆) + 𝛼 − 1

(Eq 1)

where 𝑟𝑖0 is the radius of the inner stream, 𝑅𝑐 is the capillary radius, 𝜆 = 𝜂𝑖 /𝜂𝑒 , and

𝛼 = √1 + (1/𝜆)(𝑄𝑖 /𝑄𝑒 )

(Eq 2).

Under these conditions the exponential growth rate of a perturbation in the radius of the inner stream is35:

𝜔=

Γ 𝐹(𝑥, 𝜆)(𝑘 2 − 𝑘 4 ) 16 𝜂𝑒 𝑅𝑐 𝑥 9 (1 − (1/𝜆)) − 𝑥 5

(Eq 3).

Here 𝜔 is the growth rate, Γ is the interfacial tension, 𝐹(𝑥, 𝜆) = 𝑥 4 [4 − 𝜆−1 + 4 ln(𝑥)] + 𝑥 6 [−8 + 4 𝜆−1 ] + 𝑥 8 [4 − 3 𝜆−1 − (4 − 4 𝜆−1 ) ln(𝑥)]

and


(Eq 4)


𝑘=

2𝜋𝑓 0 𝑟 𝑈𝑖 𝑖

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(Eq 5).

𝑘 is the dimensionless wavenumber of the perturbation, where 𝑓 is the corresponding frequency and 𝑈𝑖 = 2𝑄𝑒 /𝜋𝑅𝑐 2 is the linear velocity of the inner stream. Parameters for our devices are: 𝑅𝑐 = 290 µm; 𝑄𝑒 = 5500 µL/h; 𝜂𝑖 = 0.9 Pa-s; 𝜂𝑒 = 0.012 Pa-s; Γ = 0.2 mN/m; 𝑄𝑖 = 100 to 400 µL/h; 𝑓 = 5 to 70 Hz. Within the parameter space of 𝑄𝑖 and 𝑓 explored, only positive values of 𝜔 are shown, as parameters leading to positive 𝜔 are predicted to lead to spontaneous droplet formation.

3D Suspension Culture About once/hour over three individual experiments, crosslinked droplets are allowed to settle in the 1 M CaCl2 gelation buffer so that the excess Ca2+ may be removed. These cell encapsulating hydrogels are transferred by gentle pipetting into a 125 mL spinner flask bioreactor (Corning) with 125 mL of complete media at 37C, 25 mM HEPES, and adjusted to pH 7.4. For the initial t0 time point, a 20 µL sample of hydrogels is pipetted onto a microscope slide (VWR) and placed onto a brightfield microscope for imaging using a Nikon camera and SPOT Basic software. The cell encapsulating hydrogel population is transferred into a spinner flask bioreactor, the head space is backfilled with 5% CO2, and the bioreactor is placed onto a slow stir plate in a warm room maintained at 37C for the duration of the cell culture time course (9 days). At days 3, 5, 7, and 9 samples are taken from the culture for imaging as described above and the culture media is refreshed. To refresh the media, the bioreactor is removed from the stir plate, allowing the droplets to settle to the bottom of the flask. The media is aspirated, leaving about 20 mL at the bottom of the bioreactor to avoid aspirating hydrogels, and then refilled to a final volume of 125 mL with complete media at 37C and backfilled with CO2 as previously described. The cells can be recovered from the hydrogel samples at each day by a typical EDTA/trypsin washing protocol for various downstream biological assays, but is not reported here.

Flow cytometry Cell viability is evaluated to measure cell death that could have developed in the population during handling and transfer to the DEX/ALG solution. W e use a LIVE/DEAD flow cytometric assay for apoptosis by Annexin V (AV) and necrosis by propidium iodide (PI) staining. For this assay, LIVE control cells are processed by trypsinization into a single cell suspension


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and remain in complete media on ice for the same amount of time as a droplet generation experiment. The DEAD control cells are treated by suspension in 50% ethanol for 60 minutes at 37C and 5% CO2 in the incubator. We keep the DEX/ALG cell suspension in ambient conditions to represent the duration of a droplet generation experiment (~2 hours). The suspension is then transferred to a 15 mL centrifuge tube and diluted with HEPES buffer to reduce the viscosity of the solution. The suspension is centrifuged at 3000 RPM for 10 minutes. In this first wash, the cells concentrate near the bottom of the test tube but have not yet been fully pelleted and so ~5 mL of the diluted DEX/ALG is removed by aspiration and 5 mL of HEPES buffer is added and vortexed. We repeat centrifugation at 3000 RPM for 10 minutes. This wash/centrifugation protocol is repeated once more to pellet the cells and remove residual DEX/ALG. The DEX/ALG/HEPES buffer supernatant is aspirated and the cells are washed in 10 mL of HEPES buffer followed by aliquoting into four 15 mL test tubes and centrifugation at 2000 RPM for 10 minutes. The staining protocol for the LIVE and DEAD control groups and the DEX/ALG population is as described in Alexa Fluor 488 AV/Dead Cell Apoptosis Kit with Alexa Fluor 488 AV and PI for flow cytometry from Invitrogen. The samples are measured using a rate of 100,000 events/group using an Attune NxT flow cytometer with the 488 nm laser and 530/30 emission filter (BL1 channel) for AV and a 590/40 emission filter (BL2 channel with yellow laser present) for PI. Analysis of the samples is performed using FCS Express with appropriate gating (Figure 4) and a compensation matrix (Figure S5).

Rheology The rheological properties of DEX/ALG without and with cells present is an important consideration for characterizing the acoustofluidic device behavior and for measuring the viscosity of the solutions for application of theory. A Discovery Hybrid Rheometer HR-3 with a 60 mm 0.5 cone plate and Peltier plate is used to measure the ATPS components of DEX/ALG without cells and with low or high cell concentrations, the HEPES buffer with similar cell concentrations for comparison, and PEG, as follows: (1) DEX/ALG without cells, with 1x107 cells/mL, and 1x108 cells/mL; (2) HEPES buffer without cells, with 1x107 cells/mL, and 1x108 cells/mL; and (3) PEG. Each sample is measured in duplicates with three repeats. The rheological characteristics of the samples is measured using a flow ramp test (shear rate from 0.01 to 1000/s) and the viscosity for each solution is calculated (Figures S6 and S7).



Results and Discussion The ATPS used here is composed of water-based, immiscible polymers that have initial concentrations of 5% w/w 550 kDa dextran and 17% w/w 8 kDa poly(ethylene glycol). Mixing of these dextran and poly(ethylene glycol) solutions produces a turbid suspension. Thermodynamic equilibrium is achieved resulting in dextran and poly(ethylene glycol) rich solutions that are partitioned into two distinct phases with different compositions. The final concentrations of the enriched polymer in each of the two phases, calculated using refractive index and volume change measurements (Figure S2), are ≈25% w/w DEX and ≈9% w/w PEG. Each enriched phase also has low concentrations of the co-solutes present. Added alginate in the enriched dextran phase at a final concentration of 5 mg/mL provides a hydrogel-forming material for 3D cell culture. Microfluidic droplet generation is initiated by acoustically activated breakup of a DEX/ALG/cells jet within a coflowing PEG outer fluid. Tubing that feeds the DEX/ALG/cell suspension into the microfluidic device is attached to an audio speaker using laboratory tape. The speaker provides acoustic actuation and is driven by the amplified output of a waveform generator or audio-jack of a smartphone. Without this acoustic actuation, we do not observe breakup of the coflowing jet into droplets within the device (Figure S3 e and f) in agreement with previously observed low interfacial tension ATPSs.23,36,37 The resultant DEX/ALG/cell droplets are crosslinked by collection in a Ca2+-containing gelation buffer. These hydrogels are collected and suspended in mammalian cell culture environments for durations up to 9 days. In addition to using readily available acoustofluidic components, this method integrates ATPS acoustofluidics, cytocompatible gelation, and long-term suspension culture to form multicellular tumor spheroid structures with 3D cell-cell interactions (Figure 1). To identify regimes that result in the formation of well-defined droplets in our system, we investigate a combinatorial range of DEX/ALG flow rates (100-400 µL/hr) and acoustic modulation frequencies (10-60 Hz) at a constant PEG flow rate of 5500 µL/hr. Three independently constructed devices without cell-loading were studied. In addition, one device was used to characterize flow regimes with cells present. Using brightfield microscopy and image analysis, we identify parameters that reproducibly generate monodisperse or polydisperse droplets versus cases with no jet breakup into droplets (Figure 2). To better understand the nature of experimentally observed monodisperse droplet formation within the parameter space that corresponds to droplet size distributions from 100-250 µm in diameter, we applied a mass conservation constraint (see Table 1, Note 1) and also calculated the Plateau-Rayleigh-based instability behavior for spontaneous droplet production in the co-flowing ATPS (Figure S6 and


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Figure 3 a-d).35 The mass conservation constraint (black curve) is in agreement with experimentally measured droplet diameters (colored data points) across the range of DEX/ALG flow rates investigated. Spontaneous production of droplets in the DEX/ALG stream is predicted by the Plateau-Rayleigh instability analysis, but in agreement with other studies,23,36,37 no droplet formation is observed without acoustic input. Interestingly, we do observe a strong correlation between flow and frequency parameter regions where spontaneous droplet formation is predicted in a non-driven system and where the relatively weak acoustic input used here produces monodisperse droplets (gray overlay). In one instance, at the 400 L/hr DEX/ALG flow rate (Figure 3d), we observe disagreement between the measured and predicted droplet diameters at acoustic driving frequencies above 35 Hz. Since droplet size is fully determined by flow rate and frequency of droplet production, this discrepancy suggests that the frequency of droplet production is not matched to the driving frequency. Indeed, the droplet size in these cases is consistent with a droplet production frequency that maps into the region of predicted spontaneous droplet production. This observation, also seen in isolated cases at other flow rates, as well as occasional observation of polydisperse droplet sizes, may indicate that droplet production involves a competition between relatively weak acoustic driving force and the growth rate of spontaneous perturbations at different frequencies. To further simplify device design, for selected flow and frequency combinations guided by the model, we also verify that replacing the waveform generator with a smartphone provides similar performance (Figure S4). The EMT6 mouse mammary carcinoma cell line is well characterized to grow in conventional multicellular spheroids, providing an excellent cancer-relevant model cell line for hydrogel-templated multicellular spheroid formation using our new approach.38 We use standard cell culturing conditions to expand the number of cells needed for encapsulation in DEX/ALG hydrogels and develop a protocol for preparing single cell suspensions. After adding the cells to the DEX/ALG phase, we operate the system at various flow rate and frequency combinations to determine the effect of cells on droplet generation. We do observe that the droplet formation parameter space is slightly modified in comparison to the absence of cells (Figure 2, points second from the bottom at each condition). Rheological measurements indicate that the presence of cells in the DEX/ALG, at both low and high cell concentrations, does not alter the viscosity (Figure S6). We speculate that differences in the monodisperse droplet generation regime may be related to small amounts of cell clumping observed within the DEX/ALG over the duration of a droplet generation experiment. Such aggregation of cells over time is a natural phenomenon that has been exploited for conventional multicellular spheroid formation but, in our case, may also somewhat influence droplet generation.39



A well-known challenge with the interface of microfluidic droplet generation and cell culture is the necessity for low volumetric flow rates, requiring that cells be out of optimal culture conditions for up to hours at a time. We demonstrate that the method of 3D cell encapsulation reported here is biocompatible, even given the time needed for device operation. We measure the fraction of live to necrotic cells recovered from a DEX/ALG sample that is exposed to the same device operating conditions (up to two hours under ambient temperature) using an adapted Annexin V/Propidium iodide flow cytometric assay (Figure 4 and Figure S5). We find that the majority of the cell population is viable when recovered and evaluated in comparison to the control groups. Furthermore, the viability of cells within these hydrogel environments proved exceptional after 9 days of suspension culture in spinner flask bioreactors. We demonstrate reproducibility and streamlined integration of ATPS acoustofluidic droplet generation with 3D cell culture by forming EMT6 cell-loaded DEX/ALG droplets over 3 individual experiments and cultures with varying starting cell concentrations, each with 3 independently constructed devices. The cellular proliferation in each experiment is qualitatively tracked immediately after encapsulation and up to 9 days later by phase contrast microscopy and imaging (Figure 5). The size distribution of the crosslinked hydrogels is measured for a representative culture and plotted by histogram in Figure S8. Here, the inset micrograph shows the hydrogel population with a mean diameter of 124 µm and CV of 18.3%. Although there is a slight variation in the size distribution of the formed hydrogels, this CV represents an improved dispersity when compared to the size distribution of mature MTSs formed using common solution-based methods, indicating that the size distribution obtained here is fully compatible with potential applications in cancer research.3 There is an apparent lag phase in the proliferation rate of the cells from initial encapsulation to day 3, which is likely caused by a low cell encapsulation concentration and individual cells being isolated from one another. This lag phase is a small trade-off in comparison to traditional MTS initiation approaches, which can take up to weeks before handling is possible. In contrast, the cellencapsulating hydrogels are easily handled immediately after formation, and are quickly transferred into suspension culture conditions for further maturation. Once cell-cell contact is established (e.g., day 5 and later), the growth rate becomes exponential and 3D cellular aggregation, as in mature MTSs, is observed.

Conclusion

The acoustofluidic ATPS hydrogel cell culturing system reported here is readily accessible to any standard biology or chemistry laboratory and provides a straightforward method to template


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the formation of MTS assemblies. Although this approach is designed with simple components, the system allows for modularity to include parallelization for increasing droplet production throughput and also has the potential to be automated for easy operation. We find a well characterized parameter space for monodisperse droplet generation of sizes appropriate for cell culture and demonstrate fidelity across multiple individually constructed devices. Analysis using Plateau-Rayleigh theory helps guide the choice of acoustic actuation parameters to drive the generation of monodisperse droplets, and can be adapted to other capillary dimensions or solution viscosities. We also demonstrate use of an amplified smartphone to apply appropriate acoustic driving fields. In addition, we validate this acoustofluidic ATPS hydrogel system as a biocompatible encapsulation method for the culture of cells in suspension over 9 days. Overall, the 3D cell culture strategy presented in this work meets many of the characteristics for ideal ATPS performance and holds great promise for: (1) long-term culture durations (up to months) to produce fully developed and uniform multicellular spheroids; (2) use of a range of cell concentrations to control the initiation and maturation of individual spheroids; (3) integration of multiple cell types to establish advanced cocultures that develop tissue organizations; and (4) encapsulation of stem cells together with appropriate suspension culture conditions including growth factors to enable the investigation and improved understanding of organoid development. This method could also be used to template the formation of membrane-less organelles for cellfree applications as well as for drug discovery.

Supporting Information Contents include details on: ATPS preparation and single cell suspension (Figure S1), index of refraction measurements to determine DEX concentration after enrichment (Figure S2), microscopy of regimes observed within devices operated by waveform generator (Figure S3) and smartphone actuation (Figure S4), flow cytometry compensation matrix used in LIVE/DEAD assay (Figure S5), rheology results (Figures S6 and S7), and hydrogel size distribution (Figure S8)

Acknowledgments We acknowledge James P. Freyer, PhD for his training in conventional spheroid formation techniques and the University of New Mexico for partially funding this work. A.P.S. and J.A.D. were, in part, supported for work on hydrogel droplet formation by the Photosynthetic Antenna Research Center (PARC), an Energy Frontier Research Center funded by the U.S. Department of Energy, Office of Science, Office of Basic Energy Sciences under Award Number DESC0001035. J.A.D. was supported for the cell culture and encapsulation research reported in this



publication by the National Cancer Institute of the National Institutes of Health under award number 5 F31 CA189682. The flow cytometry research reported in this publication was supported in part by the National Institute of General Medical Sciences of the National Institutes of Health under award number P30 GM110907. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health. Some graphics were created with icons from Biorender.com.

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Figure 1. Acoustofluidic device workflow for forming spherical hydrogels with encapsulated cells. The well-known ATPS of PEG and DEX enriched phases, the latter containing the gelforming agent ALG, is pumped into the glass capillary microfluidic device. Amplified acoustic flow stream perturbations are applied to the inner fluid with frequency control by a waveform generator (WFG) or smartphone. The droplets are crosslinked in a calcium bath to form hydrogels and transferred to suspension culture conditions where we observe the formation of MTS assemblies.



Figure 2. Acoustofluidics system flow rate/frequency parameter space. Flowing ATPS jet outcomes are defined as monodisperse (green circles) or polydisperse (red crosses) droplets or lacking drop formation (black squares). Each set of points represents four individual experiments, run using three individually constructed devices and an instance with cells in one case, completed by varying applied frequency and flow rates (Device 1 bottom point, Device 1 with cells 2nd from bottom, Device 2 3rd from bottom, and Device 3 top point). Each experiment uses a constant PEG flow rate of 5500 µl/hr, varying DEX/ALG flow rates (100 µL/hr, 200 µL/hr, 300 µL/hr and 400 µL/hr), and images are taken 5.5 cm downstream from the injection nozzle. These results from three independently constructed devices and prepared solutions emphasize the reliability of the overall method.


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Figure 3. Droplet size distribution is plotted against a theoretical mass conservation curve (black line) as a function of applied frequency and four flow rates (a-d) for three different microfluidic devices. The gray underlay presents the results of Plateau-Rayleigh analysis of a non-driven system (following the method of Guillot et al.35) where the exponential growth rate of a spontaneous perturbation is plotted along the x-axis (for this plot, full scale in each panel is 60 per second) versus the frequency of the perturbation on the y-axis.



Figure 4. The results from a LIVE/DEAD, apoptosis and necrosis flow cytometric assay indicate that the polymer solutions and the method to encapsulate the cells are biocompatible. The dead control population (a) is treated with ethanol, the live control population (b) is processed in parallel with the DEX/ALG population (c) and demonstrates that, importantly, a large majority of cells retain viability.


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Figure 5. Encapsulating EMT6 mouse mammary carcinoma cells in DEX/ALG droplets produces a reliable platform for observing cellular proliferation and eventual formation into MTS structures with cell-cell interactions. The cells are encapsulated on three separate occasions using three individually constructed devices with different and increasing initial cell encapsulation concentrations (vertical purple gradient, rows, Cultures 1-3) and observed to proliferate up to the end of a culture time course of 9 days (scale bars 50 µm).



Page 22 of 30

Table 1. Summary of Parameters for Device Operation and Modeling.

Capillary radius (µm)

290

Outer stream viscosity, 𝜂𝑒 (Pa-s)

0.012

Inner stream viscosity, 𝜂𝑖 (Pa-s)

0.9

Interfacial tension, Γ (mN/m)

0.2

Outer stream flow rate, 𝑄𝑒 (µL/h)

5500

Inner stream flow rate, 𝑄𝑖 (µL/h) (Note 1) 100 200 300 400

Inner stream radius (µm) (Note 2) 27.5 38.8 47.2 54.3

Notes: 1. From the inner stream flow rate and modulation frequency 𝑓, mass conservation between the unperturbed stream (flow rate 𝑄𝑖 ) and the stream broken up into 𝑓 droplets per second, 3 each droplet having volume of (4/3)𝜋𝑅𝑑𝑟𝑜𝑝 , yields a droplet diameter 𝑑 = 2𝑅𝑑𝑟𝑜𝑝 =

(3𝑄𝑖 /4𝜋𝑓)1/3. For 𝑄𝑖 in µL/h and 𝑓 in 1/s, 𝑑 in µm is (130.5 (3𝑄𝑖 /4𝜋𝑓)1/3 ). This quantity is plotted in Figure 3 as the mass conservation curve for each inner stream flow rate (solid lines). 2. Inner stream radius is calculated using Equations 1 and 2.


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Graphical Abstract


ACS Applied Bio Materials 1 2 3 4 5 6


Page 24 of 30

Page 25 of 30 1 2 3 4 5 6 7 8 9 10 11 12 13 14



Page 26 of 30

400


300

Monodisperse Polydisperse

200

No droplets

100

Flow Rate (µl/hr)

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23

10

20

30 ACS Paragon40Plus Environment 50 Frequency (Hz)

60

100 µl/hr

b

200ACS µl/hr Applied Bio Materials c

300 µl/hr

d

400 µl/hr

70

Page 27 ofa 30

Device 2 Device 3

Mass balance Plateau−Rayleigh

20

30

40

50

60

Device 1 Device 1 (cells)

10

Frequency (Hz)

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28

ACS Paragon Plus Environment 100

150

200

250 100

150

200

250 100

150

Droplet diameter (µm)

200

250 100

150

200

250

y


Page 28 of 30

4

10

a

3

PI

10

2

10

2

-10

Live Control Dead

3

-10

3

2

-10

3

-10

4

10

AV

4

10

5

10

10

6

10

b

PI

744 3 10

558 2

Count

10

2

-10 372

Live

DEX-Alginate Experiment 186 -10 3

3

2

-10

-10

3

10

4

10 0

c

1

-10

2

PI

4

AV

5

10

3

10

10

6

10

4

10

10

PI

87 3 10

65 1

Count

Necrosis

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

10

2

-10 44

DEX/ALG

22 3 -10 3 -10 0

2

-10

3

10

4

AV

5

10

10

ACS Paragon Plus Environment 2

-10

2

10

Apoptosis 10 3

4

10

6

10

Day 0

Day 3

Day 5


Culture 1

Page 29 of 30

[cells]/hydrogel

Culture 3

Culture 2

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28


Cell Proliferation/Time

Day 7

Day 9

ACSOperation Applied Bio Table 1. Summary of Parameters for Device andMaterials Modeling. 1 Capillary 2radius (µm) 3 4 5 290 6

Outer stream viscosity, 𝜂 " (Pa-s)

0.012

Inner stream viscosity, 𝜂 # (Pa-s)

Interfacial tension, Γ (mN/m)

Outer stream flow rate, 𝑄" (µL/h)


0.9

0.2

5500

Page 30 of 30 Inner stream flow rate, 𝑄# (µL/h) (Note 1) 100 200 300 400

Inner stream radius (µm) (Note 2) 27.5 38.8 47.2 54.3

Oil-free Acoustofluidic Droplet Generation for Multicellular Tumor

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