Fabrication and Operation of Microfluidic Hanging-Drop Networks

Abstract

The hanging-drop network (HDN) is a technology platform based on a completely open microfluidic network at the bottom of an inverted, surface-patterned substrate. The platform is predominantly used for the formation, culturing, and interaction of self-assembled spherical microtissues (spheroids) under precisely controlled flow conditions. Here, we describe design, fabrication, and operation of microfluidic hanging-drop networks.

1. Introduction

The hanging-drop technique enables the formation of scaffold-free 3D spherical microtissues by seeding a defined number of cells into hanging drops of a specific culture medium. Cells sediment by gravity force, aggregate, and form a spherical microtissue at the liquid–air surface [1]. Hanging-drop networks (HDNs) expand the isolated hanging drops toward a fully interconnected network of hanging drops and enable controlled liquid flow between the hanging drops [2]. Adding perfusion functions allows for continuous medium exchange, for application of compound dosage protocols and for interaction of different microtissue types to realize multi-tissue or the so-called “body-on-a-chip” setups. The hanging-drop technology has also been tested for stem cell culturing [3]. Further, electrical impedance spectroscopy [4], biosensor readout methods [5], and on-chip peristaltic pumps [6] have been integrated.

Hanging-drop networks are designed as completely open microfluidic systems at the bottom of an inverted, surface-patterned substrate (Fig. 1). They inherently fully exploit the benefits of the liquid–air interface with respect to low cell adhesion and reduced compound adsorption. Further, their open nature ensures gas exchange, prevents bubble formation and gives access to the liquid phase and the microtissues at every position in the network. Finally, their fabrication is simple, and the design of the networks is very versatile.

Fig. 1.

Photograph of a line of hanging drops filled with green food dye. As a result of surface energy minimization, all five hanging drops have the same radius and, therefore, comprise the same liquid volume

The surface patterns underneath the inverted substrate guide the liquid by surface tension and capillary forces. Rim structures are used to distinguish wetted regions from dry regions and prevent the liquid from flowing over the whole surface in an uncontrolled way. The design of the rim structures defines where drops are formed. Circular patterns induce the formation of hanging drops, whereas extended narrow structures produce a channel-like structure. Through the variation of the feature dimensions many different arrangements and configurations of hanging-drop networks can be realized. Robust operation of hanging-drop networks depends on a few basic principles that have to be followed during the design and arrangement of the hanging drops and interconnection channels; the respective features will be explained in Subheading 3.1.

The hanging-drop network structures are made of PDMS casted from a micropatterned SU-8 mold. The stability of the chips can be increased through bonding of the PDMS substrate onto a glass slide with fluid access holes. The chips are placed in a custom-made chip holder and connected to conventional pumps via tubing. All fabrication and setup steps are described in detail in Subheadings 3.2, 3.3 and 3.4. The fabrication of the SU-8 mold is done in a cleanroom. All other steps can be performed in a conventional laboratory.

The open nature of the HDN-systems ensures bubble-free initial liquid filling of the microfluidic network. Further, the loading of cells and preformed microtissues is straightforward and easy to perform. Different approaches for loading cells and spheroids are presented in Subheading 3.5.

In comparison to the more common closed microfluidic systems, flow control is different in hanging-drop networks. For a stable drop size to be maintained over extended time, the liquid needs to be actively infused and actively withdrawn at both inlet and outlet, respectively, at a precision, which is not available in most commercial systems. We present two different methods that allow for robust perfusion over an extended culturing period. In the first method, described in Subheading 3.6, the outlet is conceived as a hanging drop and a needle is placed at the liquid–air interface. The position of the needle then defines the drop size in the whole network (we call it “needle-outlet” method, here). This method does not require a microscope and can be used for perfusion in a conventional incubator. In the second method, described in Subheading 3.7, we use microscopy to estimate the average drop height by (software) autofocusing the individual microtissues. The micro-tissues are always located at the bottom of each drop on the liquid–air interface and the z-position of the objective with the microtissue in focus can be used as measure for the drop height. Based on this value the flow rate is adjusted online via proportional–integral feedback control (“feedback” method). Finally, in Subheading 3.8 we describe, how spheroids can be retrieved from the microfluidic system for further downstream analysis.

2. Materials

2.1. SU-8 Mold

1. Transparency masks of the design printed at 50800 dpi resolution (Selba SA, Versoix, Switzerland).

2. 4-inch silicon wafer (525 μm thick, single-side polished).

3. Cleanroom equipped with mask aligner, spin coater, hot plate (see Note 1), ultrapure water bath, glass beakers, spiders, and covers.

4. Chemicals:

   • (a)
     Acetone (semiconductor grade)
   • (b)
     Isopropanol (semiconductor grade)
   • (c)
     SU-8 100 negative photoresist (Microchem Corp., Newton, MA, USA)
   • (d)
     mr-Dev 600 Developer (Micro Resist Technology GmbH, Berlin, Germany)
   • (e)
     Trichloro(1H,1H,2H,2H-perfluorooctyl)silane (Sigma-Aldrich, Buchs, Switzerland).

2.2. PDMS Chip

1. Polydimethylsiloxane (PDMS), Sylgard 184, (Dow Corning GmbH, Wiesbaden, Germany).

2. Vacuum desiccator.

3. Scotch tape or custom-built wafer holder for PDMS casting.

4. Hot plate (see Note 1).

5. Cutter blade.

6. Hollow punchers 0.75 mm and 2.00 mm (Harris Uni-Core, Ted Pella, Inc., Redding, CA, USA).

7. Microscopy glass slides, standard size (1 mm thick, 25 mm x 75 mm or 50 mm x 75 mm).

8. Diamond drill of 1.2 mm diameter.

9. O₂ plasma cleaner/sterilizer 50 W at 0.3 mbar (Diener Electronic GmbH & Co., Ebhausen, Germany).

10. (For needle-outlet) N-124S Nanoport connector and F-124S Standard Head Fitting (Idex Health & Science GmbH, Wertheim, Germany).

11. (For needle-outlet) Araldite rapid two-component glue (Huntsman Advanced Materials GmbH, Basel, Switzerland).

2.3. Setup

1. Custom-built chip holder, or PDMS spacer blocks.

2. One-well culture dish (Nunc OmniTray, Thermo Fisher Scientific, Rochester, NY, USA).

3. Humidifier pad (InSphero AG, Schlieren, Switzerland).

4. Tubing and connectors:

   • (a)
     Polytetrafluoroethylene (PTFE) tubing, ID 0.3 mm/0.5 mm, OD 0.6 mm/1 mm (Bola GmbH, Grünsfeld, Germany).
   • (b)
     Standard luer lock syringe-tubing connectors 22 GA.” Bent 90 deg. (APM Technica AG, Heerbrugg, Switzerland).
   • (c)
     Standard luer lock syringe-tubing connectors 25 + 32 GA straight (APM Technica AG).
   • (d)
     Silicon tubings Tygon LMT-55, ID 0.38 mm, Wall 0.91 mm (Idex Health & Science GmbH, Wertheim, Germany).
   • (e)
     Peristaltic tubing Tygon S3 E-LFL, ID 0.27 mm, 0.91 mm wall (Idex Health & Science GmbH).

5. neMESYS syringe pump base and dosing units (Cetoni GmbH, Korbussen, Germany).

6. Peristaltic pump (Ismatec, ISM935C, Idex Health & Science GmbH).

2.4. Microscopy

1. Automated inverted microscope (DMI6000B, Leica Microsystems).

2. Objectives (Leica Microsystems).

   • (a)
     5 x /0.12 HCX FL Plan.
   • (b)
     10 x /0.30 Ph1 HCX PL Floutar.

3. Camera (DFC340FX, Leica Microsystems).

4. (Optional) C-Mount 0.70x.

5. Environmental box for microscope (“The Box,” Life Imaging Services, Basel, Switzerland).

6. Temperature controller (“The Cube,” Life Imaging Services).

7. Stage-top incubator and automated gas mixer (“The Brick,” Life Imaging Services).

2.5. Microscope Control Software “YouScope”

1. Youscope version R2016 or higher, available at http://www.youscope.org/ for Windows 7 (32bit or 64bit) or higher.

2. The control software is necessary for the “feedback” method; optional for “needle-outlet” method (e.g., for imaging and/or switching flows).

3. Methods

3.1. Design of the Chips

1. Design the microfluidic hanging-drop networks by using 2D CAD software (e.g., Clewin, AutoCAD). A very basic hanging-drop network is presented as an example in Fig. 2a. The layout features one inlet, one outlet, and five drop-sites that are interconnected through short, semiopen connection channels. The drop structures have a diameter of 3.5 mm and are arranged in a row (see Note 2). The drop pitch is 4.5 mm, which corresponds to the 384-well-plate format, making it compatible with routine imaging and pipetting. A 200-μm-wide circular rim defines the drop sites and guides the liquid on the microfluidic chip. Cross sections 1, 2, and 3 in Fig. 2b illustrate the surface pattern of the inverted substrate. The depth of the fluidic structure is 500 μm. The groove around the rim is 250 μm deep and 800 μm wide. Inlet and outlet areas have a diameter of 1.5 mm. The small channels between the drops are 1 mm long, 200 μm wide, and 500 μm deep.

2. Printing of two high-resolution transparency masks is required for the fabrication process of the SU-8 mold. Figure 2c shows both layers for the hanging-drop line in Fig. 2a.

3. The reported dimensions for drops and channels have been optimized with regard to drop and network stability and media perfusion characteristics (see also ref. 2). Design custom hanging-drop network configurations (e.g., larger arrays, different interconnections, other inlet and outlet position) respecting the guidelines given above. Arrays larger than 8-by-8 drops and more than 8 inlets and outlets become rather difficult to realize. An example of a 4-by-4 array is presented in Fig. 3a. The array includes also a microfluidic gradient generator allowing for exposing spheroids to different compound concentrations [7]. Additional microfluidic features may be devised, but then have to be tested. The channel dimensions should, in general, include a width of 200 μm and a height of 500 μm.

4. The integration of capillary stop valves enables sequential filling of the microfluidic network and subsequent reconfiguration. An example is given in Fig. 3b. The 4-by-6-drop array has four inlets and four outlets. Each column can be loaded with a different liquid or cell suspension through the cell loading ports located at the top of the columns. After loading, the columns are connected by adding liquid through the connecting ports (see also Subheading 3.5). The optimized design and respective mask layers of the capillary stop valves are shown in Fig. 3c, d.

Fig. 2. Design of the hanging-drop network.

(a) Line of five hanging drops with single inlet and outlet. All important dimensions are indicated. (b) Schematic cross sections of drop, channel, and outlet site. (c) Masks required for fabrication of the negative-tone SU-8 mold of the hanging-drop line in (a)

Fig. 3. Examples of different hanging-drop networks.

(a) 4-by-4 interconnected hanging-drop array with preceding microfluidic gradient generator structure. (b) 4-by-6 hanging-drop array including the reconfiguration option, so that every column can be loaded independently. (c) Details of the valves indicated in (b). (d) Design and of the masks for fabricating the valves

3.2. Fabrication of the SU-8 Mold

1. Take a 4-in. polished silicon wafer and perform a dehydration bake for 5 min at 200 °C on a hot plate. Proceed right afterward with SU-8 spin coating.

2. Spin-coat the first layer of SU-8 100 (~250 μm height, Fig. 4a): Pour SU-8 100 directly from the 500-ml bottle onto the wafer (see Note 3). Use the following two-step spin-coating program: (1) Spreading step with a ramp (500 rpm/s) to 500 rpm held for 5 s, (2) spinning step with a ramp (500 rpm/s) to 1500 rpm held for 30 s. After coating, remove the wafer from the spin coater and clean the backside of the wafer by using a tissue and Acetone (see Note 4).

3. Soft-bake the wafer on a leveled hot plate using the following protocol: 15-min ramp from RT to 65 °C, hold for 30 min, 15-min ramp from 60 °C to 95 °C, hold for 90 min, slow cool down to RT in ~60 min. Slow ramps reduce internal stress and improve the adhesion of SU-8 to the silicon substrate.

4. UV-expose the first SU-8 layer on a mask aligner through the first mask (Fig. 4b). SU-8 is a negative-tone resist; exposed areas are cross-linked and remain on the wafer after development. Use soft-contact mode and a broadband exposure dose of 400 mJ/cm² (see Note 5).

5. After exposure, perform the postexposure bake on a hot plate using the following protocol: 15-min ramp from RT to 65 °C, hold for 5 min, 15-min ramp from 60 °C to 95 °C, hold for 30 min, slow cool down to RT in ~60 min (Fig. 4c).

6. Spin-coat the second layer of SU-8 100 (~250 μm height) onto the first SU-8 layer using the same procedure described in steps 2 and 3 (Fig. 4d).

7. Align the second mask with respect to the SU-8 structures on the wafer and UV-expose the second SU-8 layer using a broadband exposure dose of 400 mJ/cm² (Fig. 4e, f). After exposure, perform the postexposure bake using the same parameters described in step 5 (see Note 6).

8. Develop the unexposed SU-8 in a glass beaker by using mr-Dev 600 developer. Put the wafer upside down on a spider and fill the beaker with developer until the wafer is completely immersed. Cover the beaker. Develop for about 60 min and agitate the developer from time to time (Fig. 4g) (see Note 7). Once all unexposed resist has been dissolved, rinse the wafer with fresh developer followed by isopropanol and then rinse it thoroughly in an ultrapure water bath. Spin-dry the wafer for 60 s at 2500 rpm or let it dry in air.

9. Vapor silanization of the SU-8 mold: Put the clean wafer, together with 5 μl of trichloro(1H,1H,2H,2H–perfluorooctyl)silane applied on a glass slide, into a desiccator. Apply house vacuum or a pump vacuum for 2–3 min and close the valve. Leave the wafer in the silane atmosphere for 2 h. Purge the chamber with air and remove the wafer.

Fig. 4. Fabrication of the SU-8 mold.

(a) Spin coating of the first SU-8 layer. (b) Exposure through the first mask. (c) Postexposure bake and cross-linking. (d) Spin-coating of the second SU-8 layer. (e) Alignment and exposure through the second mask. (f) Postexposure bake and (g) development of un-exposed SU-8

3.3. Fabrication of the PDMS Hanging-Drop Network

1. Add Sylgard 184 silicone elastomer and Sylgard 184 silicone elastomer curing agent at a ratio of 10:1 (e.g., 44 g in total) into a dust-free plastic cup and mix thoroughly. Place the cup into a vacuum desiccator for ~30 min until no air bubbles are visible anymore.

2. Add tape as a wall around the wafer or place the wafer in a petri dish to create a containment for the PDMS with the wafer at the bottom (see Note 8). Custom-built wafer holders may also be used.

3. Pour 20 g of degassed PDMS on the wafer (3 mm thick, Fig. 5a). Pour 10 g of PDMS in an empty petri dish (1 mm thick), which is later used as a mask for the surface activation. Degas the PDMS under vacuum for additional 30 min to remove all bubbles.

4. Place wafer and petri dish on a leveled hot plate for 2 h at 75 °C to fully cure the PDMS (see Note 1).

5. After curing and cool down, carefully remove the structured PDMS from the mold and the PDMS layer out of the petri dish (Fig. 5b). Cut the PDMS into single-chip pieces using a cutter blade and punch holes at inlet and outlet using a 0.75-mm hollow puncher.

6. Prepare the PDMS mask by cutting the thin PDMS layer into slides fitting the hanging-drop network array by using a cutter blade. Punch holes at the corresponding drop positions using a 2-mm hollow puncher (Figs. 5c and 6).

7. Take a microscopy glass slide and drill holes with a diameter of 1.2 mm at the inlet and outlet positions by using a diamond drill (see Note 9).

8. Clean the PDMS bonding surface as well as the glass slide with soap and successively rinse it with DI water, acetone and isopropanol and dry the two parts with an air gun.

9. Place the PDMS chip and the glass slide into the plasma cleaner with the bonding surfaces facing upward. Activate the surfaces for 25–30 s using oxygen plasma at 50 W and 0.5 mbar.

10. Remove the parts from the plasma cleaner and place the PDMS chip onto a dust-free paper with the nonstructured surface at the top. Align the microscopy glass slide to the chip by eye and bring them into contact starting at one side so that no air bubbles are trapped. Slightly press them together (Fig. 5d).

11. (For needle-outlet) Glue an N-124S Nanoport connector on the glass slide at the outlet hole. Use a two-component glue (Araldite), which cures rapidly. Apply slight pressure on the connector from the top for several minutes to tightly fix it (Fig. 5e) (see Note 10).

12. (For needle-outlet) Remove the plastic part of a 32-GA and a 25-GA standard luer lock syringe-tubing connector. Glue the 32-GA needle tip inside the 25-GA needle tip by carefully adding glue to the thinner tip and sliding it into the thicker one. Make sure that approximately 5 mm of the inner needle stays uncovered. Glue the combined needle into a F-124S standard head fitting with the uncovered part in front (Fig. 5f).

Fig. 5. Fabrication of the PDMS chip.

(a) Molding of PDMS from the SU-8 structure. (b) PDMS chip with SU-8 replica. (c) PDMS mask for chip preparation. (d) PDMS chip bonded on a glass slide. (e, f) Application of the needle-outlet

Fig. 6.

Preparation of the chip. A PDMS mask with holes is used to selectively activate the inside of the circular hanging-drop areas and channels, while the rim is protected and remains hydrophobic

3.4. Chip Preparation

1. Carefully clean the PDMS chip and the PDMS mask with soap, water, acetone, isopropanol and dry the two parts with an air gun (see Note 11).

2. Align the holes in the PDMS mask with the circular hanging-drop areas on the PDMS chip before plasma activation and press them slightly together so that a good contact of the PDMS mask and the rim is ensured (Fig. 6) (see Note 12).

3. Put the chip into the plasma cleaner with the openings of the mask facing upward. Activate the chip for 45–60 s using an oxygen plasma at 50 W and 0.5 mbar. In this step, the inside of the circular areas and channels is turned hydrophilic, while the rim structures covered by the PDMS mask remain hydrophobic.

4. Remove the mask from the PDMS chip and affix it to a custom-built holder with the hanging-drop structures facing down. Alternative: Use prefabricated PDMS blocks to support the flipped chip (see Fig. 9) (see Note 13).

5. (For needle-outlet) Insert the prepared needle fitting into the Nanoport connector (Fig. 5f). Visually adjust the needle so that the tip is 0.5–1.0 mm below the rim structure.

6. Take a sterile one-well culture dish (OmniTray box) and place a humidifier pad inside. Soak it with sterile DI water. Place the PDMS chip and custom-built holder inside.

Fig. 9. Experimental setup on the microscope.

(a) The hanging-drop chip is placed on PDMS blocks inside an OmniTray box. The stage-top incubator and the incubator box ensure high humidity and a constant temperature to minimize evaporation. The inflow is provided by neMESYS syringe pumps. Flow rates into the microfluidic chip are adjusted through a feedback controller, which has been implemented in YouScope to keep the drop height constant. Outflow is generated by a peristaltic pump and is constant during the experiment. (b) Enlarged view on the outlet of the chip

3.5. Cell and Spheroid Loading

Option 1—Loading of a cell suspension

1. Perform the loading sequence within 15 min after oxygen plasma activation.

2. Prepare the cell suspension and adapt the cell concentration with respect to the final cell numbers required per hanging drop.

3. Calculate a volume of 8 μl per drop and load the total liquid volume that is required for filling of the selected drop (sub) array into a conventional pipet (200-μl tip).

4. Load liquid or the prepared cell suspension through one of the inlets (see Note 14). The liquid will automatically spread over the whole network, and hanging drops will simultaneously develop below all circular structures (Fig. 7a). For large networks, one or more additional inlets at drops inside the network may be used to achieve a more homogeneous cell distribution.

5. For hanging-drop networks that can be reconfigured through capillary stop valves, load the defined cell suspensions through the respective loading ports (Fig. 7b, c). Let the cells sediment for 5–10 min and then load a small amount (~5 μl) ofmedium through the connection port to connect the different subarrays.

Fig. 7. Cell and spheroid loading.

(a) Loading of liquid or cell suspension through the inlet and formation of hanging drops of uniform size. (b) Sequential loading of subarrays of hanging drops (here columns) and subsequent connection. (c) Photograph of the loading using food dye and details on the capillary stop valve function. (d) Loading of the network with drops facing upward. (e) Transfer of a spheroid into a standing drop

Option 2—Loading of preformed microtissues

1. Seal inlet and outlet with sticky tape.

2. Flip the chip so that the hanging-drop structures are facing upward.

3. Load medium into the network until standing drops are formed at all circular areas. The medium can be loaded at several positions in the network (Fig. 7d).

4. Pick up the selected spheroids from your dish or well plate into the tip of a 100/200 μl pipet tip together with ~20 μl of medium.

5. Observe and wait until the spheroid settles at the bottom of the tip at the liquid–air interface (Fig. 7e).

6. Hold the tip vertically and bring the tip into contact with the selected standing drop. The spheroid will transfer without any pipetting actuation.

7. Load all spheroids into the network. If the drops have been designed with the required pitch, a multichannel pipet may be used for parallel spheroid transfer (see Note 15).

8. Flip the chip back into hanging drop mode and place it onto the holder (see Note 16).

9. Remove the sticky tape at inlet and outlet.

3.6. Operation Using “Needle-Outlet” Method

1. Choose an appropriate incubator for your experiments with 5% CO₂ and >95% humidity. The incubator should have the possibility to insert tubing from the outside.

2. Place the syringe pump and the peristaltic pump as close to the incubator as possible.

3. Prepare the tubing connections from the syringe pumps to the PDMS chip with polytetrafluoroethylene (PTFE) tubing of appropriate length (ID 0.3 mm). Connecting pieces are prepared by removing plastic and glue from standard luer lock syringe-tubing connectors (22 GA Bent 90). They are then connected to the PTFE tubing by short flexible silicon tubing (ID 0.38 mm) (Fig. 9b).

4. Fill glass syringes with culture medium and mount the tubing to the syringe, which is then mounted onto the neMESYS syringe pumps. Prefill the tubing with medium to remove any air bubbles before connecting it to the hanging-drop chip.

5. Use special peristaltic tubing (ID 0.27 mm) for the peristaltic pump. A PTFE tubing of appropriate length is directly inserted in the peristaltic tubing. The other end of the PTFE tubing is inserted in a short silicon tubing adapter piece (ID 0.38 mm), which is later used to connect to the outlet needle (Fig. 8).

6. Put the one-well culture dish (OmniTray box), including the PDMS chip prepared in Subheading 3.4, into the incubator. Connect inlet and outlet tubing to the chip. Check the horizontal position of the PDMS chip (see Note 17).

7. Close the lid of the culture dish and the incubator. Make sure that none of the tubing is squeezed.

8. In this operation mode, the drop size is controlled by the position of the outlet needle. Liquid is removed from the chip if the drop size increases and the liquid–air interface moves below the needle tip. The liquid removal stops as soon as the liquid–air interface then again reaches the needle tip. This back and forth between liquid withdrawal and no withdrawal ensures a constant drop height (Fig. 8). As indicated in the figure, the outlet produces a segmented flow with alternating air and liquid plugs.

9. Start the perfusion of the chip. The programed withdrawal rate should always be set higher than the inlet rate for stable chip operation (1.5–2 times the inlet rate). A continuous flow rate of 0.5–10 μl/min has been successfully tested. As an alternative, pulsed inlet flow can be applied. The average input volume should, however, always compensate for the evaporation in the incubator and to guarantee a constant liquid volume in the chip and never exceed the continuous withdrawal rate.

10. (for needle-outlet) If the volume of the hanging drops is increasing too much, decrease the needle length by turning the needle fitting counter-clockwise. Turn the fitting in the opposite direction to increase the volume of the hanging drops.

11. (Optional) The withdrawn liquid can be sampled by simply guiding the tubing downstream of the peristaltic pump into a sampling device or container.

Fig. 8.

A back and forth between liquid withdrawal (a) and no withdrawal (b) is used to control the drop volume and the overall liquid volume in the chip

3.7. Operation Using “Feedback” Method

1. Install YouScope and Qmix SDK (first time). Configure You-Scope as described in [8] and add a “NemesysPump” device.

2. Switch on the microscope and set the temperature controller of the environmental box to 37 °C for 3–4 h before starting the experiment.

3. Select a 5 x or 10 x objective (Fig. 9).

4. Set the culturing conditions inside the stage-top incubator with an automated gas mixer to 5% CO₂ and >95% humidity and a gas flow rate of 10 l/h.

5. Prepare the tubing connections from the syringe pumps to the PDMS chip with polytetrafluoroethylene (PTFE) tubing (ID 0.3 mm, OD 0.6 mm) and metal connecting pieces. Connecting pieces are prepared by removing plastic and glue from standard luer lock syringe-tubing connectors (22 GA Bent 90). They are then connected to the PTFE tubing by short flexible silicon tubing (ID 0.38 mm).

6. For the inlet, fill glass syringes with culture medium and mount the tubing to the syringe with standard luer lock syringe-tubing connectors (22 GA), which are then affixed to the neMESYS syringe pumps. Prefill the tubing with medium. Remove any air bubbles.

7. Special peristaltic tubing (ID 0.27 mm) is used for the peristaltic pump. All the other part of the tubing remains the same as described in step 5. Prefill the peristaltic tube with medium. The outlet of the tube can be used for sampling of the medium.

8. Affix the one-well culture dish (OmniTray box) including the PDMS chip prepared in Subheading 3.4 to the microscopy stage. Connect all inlet and outlet tubes to the chip. Close the dish with a lid and put the stage-top incubator on top (see Note 18).

9. Check the horizontal position of the PDMS chip by checking the z-position on all four edges of the chip. Adjust if needed.

10. Check if all channels and capillary stop valves are filled with liquid.

11. Check the size of the hanging drops and adjust the drop height if needed by slowly infusing (~5 μl/min) additional medium through the syringe pumps (see Note 19).

12. Identify autofocus settings for feedback control in YouScope (first time): Manually focus on a microtissue inside a drop. Create a “simple measurement” with #executions = 1. Add an autofocus job using “exhaustive search,” with upper and lower bounds approximately plus/minus half the drop height. Before starting the measurement, double-click on “autofocus results”. Display a plot with x-column = “relative focus,” y-column = “focus score,” and “scatter plot,” and start the measurement. Adjust the imaging settings and the focal score algorithm until the focus-score plot becomes bell-shaped, with a single maximum around zero and a good signal-to-noise ratio (see Note 20).

13. For time-lapse imaging and drop-height control (Fig. 9), create a microplate measurement in YouScope. Choose the 384-well-plate format, or create a custom microplate if using a nonstandard chip format. Select microplate positions corresponding to the drop layout, and manually focus the microtissues in each drop. Add a “droplet-based microfluidic” job, and select all connected syringes. Use “Brent Optimization” for the autofocus, and all other settings as previously identified. Select “syringe-table” as the controller strategy, select the target flow rate (see Note 21), and allow the controller to deviate approximately half the target flow rate to adjust the drop height. Measure or estimate (based on the chip design) the first-order relationship between drop height and volume around the target height. Choose at least one syringe to generate the inflow (the target flow is distributed equally between all selected syringes). If using the peristaltic pump to generate the outflow, do not choose “outflow” for any syringes. By adding rows to the “syringe table,” one can switch during the measurement between syringes containing different media. To take additional images, e.g., for quantifying fluorescence, add the respective imaging jobs to the imaging protocol after the “droplet-based microfluidic” job.

14. Start the measurement and, at the same time, the withdrawal through the peristaltic pump with a flow rate equal the target flow rate of the controller (see Note 22). If necessary, stop the measurement and the flow to adjust the controller settings (see Note 21).

3.8. Retrieval

1. Collect a selected microtissue from a single drop using a conventional pipet.

2. (Alternative) Bring the hanging drop network into contact with the surface of a petri dish and collect the microtissues from there by using a pipet. The petri dish can be filled with medium. By using this alternative collection method the order and registration of the microtissues will be lost.

3. (Alternative) Transfer all microtissues in parallel to a special receiver plate. The receiver plate features hydrophilic spots at the locations of the hanging drops. In this way, the registration of the microtissues is maintained (for more details please refer to [3]).

4. Notes

1. Make sure to horizontally level the hot plate.

2. Hanging drops can be designed and operated under stable conditions if they feature diameters between 2.5 mm and 5 mm.

3. SU-8 is very viscous. The final height has ~10% variation. The amount poured onto the wafer needs to be the more or less the same for each layer.

4. If small air bubbles appear on the wafer, simply prick them with a small needle.

5. The resolution depends on the print quality of the mask. Use chromium masks on glass if critical features are smaller than 30 μm. For larger structures, such as presented here, 50800-dpi foil masks are sufficient.

6. Special alignment structures are placed at the sides of both mask layers and promote precise alignment.

7. When the developer becomes yellow, exchange it for fresh one.

8. Make sure that the PDMS is not going underneath the wafer.

9. For precise positioning of the holes, place the punched PDMS chip onto the microscopy glass slide and mark the positions of the access holes directly on the glass.

10. For precise alignment, use a 25-GA needle tip inside the Nanoport connector that protrudes into the punched PDMS hole during fixation so that the connector and outlet are cocentered.

11. A toothbrush can be used with soap to remove persistent dust or other debris in the channels and drop structures.

12. No contact of the mask will lead to plasma activation of the rim and potential failure of the chip during operation.

13. Height of the PDMS block should be ~8 mm, yielding a total distance of the drops from the OmniTray bottom of approximately 3–4 mm.

14. Loading flow rates of 300–400 μl/min can be applied. The total chip volume depends on the chip design. In general, one drop (without channel) with a diameter of 3.5 mm has a volume of ~8 μl. This yields a drop size/height of ~800 μm (measured from the rim structure).

15. Place the network on a heating mat, if the procedures take longer. Be quick to minimize evaporation.

16. Do the flipping fast and with a uniform rotation rate along the axis of the drop rows.

17. Due to the additional height of the Nanoport connector, a small window needs to be removed from the lid of the culture dish to provide access for the outlet tubing.

18. Properly tape the tubing on the microscope stage.

19. The drop height can be measured through the difference in the z-position of the rim structure of the chip and the specimen located at the bottom of the hanging drop.

20. Small improvements in autofocus quality can significantly improve long-term stability of the controller. Try to slightly over-expose the autofocus search images, clean the optics, use prefiltered medium, switch off lights, use Köhler illumination, and consider to decrease magnification. In our experience, autocorrelation based focus scores with lags between one and eight showed the best results.

21. If the average drop height and the target flow rate show long-lasting oscillations, increase the “mean droplet’s height learn speed” of the observer, and increase the time-constants for the proportional and integral parts of the controller. If individual drop height measurements are noisy, decrease the “individual droplet’s height learn speed” to avoid high frequency fluctuations in the flow. If only small flow adjustments are necessary, manually switch on the flow and let the droplets settle before configuring and starting the controller.

22. The maximal flow rate highly depends on the array design and the volume of the interconnecting channels, and, in general, on the flow resistance between inlet and outlet. Up to 20 μl/min for a single drop line can be applied without problems. In some cases, high flow rates can lead to substantially different drop sizes (smaller drops toward the outlet) as a result of the pressure drop over the interconnecting channels in the network (for more details see ref. 2).