Simple, Affordable, and Modular Patterning of Cells using DNA

1. Design experiment

1. Plan out the desired experiment, considering feature size, feature spacing, number of cell types involved, and the arrangement of cells with respect to one another. Refer to Supplemental File 1, a guide for experimental design, and Supplemental File 2, which contains example oligo sequences.
2. Design photomask using computer-aided design software. An example photomask is provided in Supplemental File 3.
   1. Draw a rectangle of the dimensions of a standard microscope slide (25 mm x 75 mm).
   2. Draw four rectangular regions 10 mm wide and 10 mm long, distributed evenly across the slide.
   3. Within each region, draw features that are the desired size, shape, and spacing for the experiment. Cells will adhere only to these features in the experiment.
   4. To create aligned photomasks for multiple cell types, create a master drawing with all sets of features, then save versions that correspond to each cell type.
   5. Order a high-resolution (at least 20,000 dots per inch) transparency photomask from this CAD drawing with the features drawn in 1.2.3 transparent and the larger regions black.

2. Photopattern DNA onto aldehyde-functionalized slides (protocol adapted from Scheideler et al.²⁹ )

1. If patterning multiple cell types, fabricate fiducial markers on the aldehyde-functionalized slide before any DNA patterning to facilitate alignment of features. Alternative methods for creating fiducial markers are suggested in Supplemental File 1.
   1. To create metal fiducial markers, apply S1813 positive photoresist as described in steps 2.3 – 2.11. Use a photomask that contains large features that will be easy to align later. Incorporate these features into the design of the photomasks that will be used for DNA patterning.
   2. Deposit a thin film (100 Angstroms) of titanium onto the slide using electron-gun evaporation²⁹. Remove excess metal and photoresist using acetone, and then proceed to the DNA photopatterning.
2. Prepare a 20 µM solution of a 5'-amine-modified oligo in DNA buffer (50 mM of sodium phosphate in water, pH = 8.5). See Supplemental File 2 for suggested oligo sequences.
   NOTE: It is possible to use as little as 5 µM of amine-modified oligo for some patterns and applications, so surface DNA concentration may need to be optimized.
3. Pre-heat a hot plate to 100 °C.
4. Use double-sided tape or a vacuum to attach an aldehyde-functionalized glass slide to the rotor of a spin coater.
   CAUTION: Slide detachment during spin-coating is a safety risk. Always use the spin coater in an enclosed container with a lid, such as an acrylic box.
   NOTE: Label a corner of the slide by using a diamond scribe or similar implement to scratch the glass. This helps with slide identification and orientation after the photoresist has been washed away.
5. Use a disposable pipette to drop the positive photoresist onto the aldehyde slide. For even coatings, add small drops of the photoresist across the slide, instead of one large drop in the middle (Supplemental Figure 1A).
6. Using the spin coater, spin the slide at 3000 rpm for 30 s.
7. Place the slide on 100 °C hotplate for 1.5 min (soft bake) to crosslink photoresist.
8. Remove the slide from hotplate. Place a photomask with the features desired for this experiment on top of the slide and weigh the photomask down with a piece of glass (Supplemental Figure 1B,C). Cover the entire setup in an opaque box (Supplemental Figure 1D). Expose with a UV lamp (365 nm wavelength, 360 mW, 5 inches from slide, total radiant energy density 100 mJ/cm²) for 2 min.
   NOTE: UV light will break the polymer bonds in the photoresist underneath transparent regions of the photomask, creating regions where DNA will later be able to adhere.
9. Develop the slide by immersing in developer solution for 3-5 min (Supplemental Figure 1E).
10. Rinse away excess developer solution with water. Dry under a stream of air or nitrogen. (Supplemental Figure 1F).
11. Confirm that the photolithography was successful by looking at the slide under the microscope. Because the photoresist is UV-light sensitive, do this step quickly and then store the slide in the dark while preparing other slides (if applicable).
    NOTE: A successfully patterned slide should have sharply defined edges for each feature, no cracking, and no feature distortion at the edges. Examples of correct and incorrect photolithography are provided in Supplemental Figure 2A. See Table 1 for troubleshooting suggestions if photolithography does not provide the desired feature quality.
12. Add a droplet of the 20 µM amine-modified oligo solution (Step 2.1) onto each photopatterned region of the slide. Use a pipette tip to gently spread the droplet across the entire region, being careful not to scratch the slide. (Supplemental Figure 1G).
13. Bake the slide in a 65-70 °C oven until the DNA solution has fully dried onto the slide surface (about 1 h).
14. Perform reductive amination by placing the patterned, baked slides in a 15 cm cell-culture dish and place in a fume hood on top of a shaker. Weigh out 100 mg of sodium borohydride. In a fume hood, add 40 mL of phosphate-buffered saline (PBS), gently mix, and add to the dish containing the patterned slides. Let the reaction proceed for 15 min with gentle shaking.
    NOTE: The amine on the oligo first forms a Schiff base with the aldehydes on the slide surface. This is a reversible covalent bond that must be converted to an irreversible bond prior to use in DPAC. Addition of a reducing agent (sodium borohydride) converts the Schiff base to a secondary amine by reductive amination.
    CAUTION: The reaction of sodium borohydride with water creates hydrogen gas and will continue to do so for hours or days after the reaction begins. Perform the reductive amination step in a fume hood and keep all sodium borohydride solution waste in an open or loosely capped container in the fume hood for at least 24 h.
15. Remove unreacted DNA by washing twice with 0.1% sodium dodecyl sulfate (SDS) in water, then three times with distilled water. Dry the slide under a stream of nitrogen or air.
16. Rinse the slide with acetone to remove the remaining photoresist.
    NOTE: At this point, the DNA has been irreversibly and covalently attached to the slide and all unreacted aldehyde functional groups have been converted to alcohols. The photoresist is no longer needed.
17. If multiple oligos will be patterned, return to step 2.4, align the photomask with fiducial marks, and repeat.
    ​NOTE: The experiment can be paused here. Store slides in a vacuum desiccator. Under dry conditions, the slides can be stored for up to 3 months without a loss of quality.

3. Make slide hydrophobic (optional) (protocol adapted from Todhunter et al.²⁴ )

NOTE: It is advantageous, but not required, to modify the slide's surface chemistry to render it more inert and hydrophobic. Non-specific cell attachment is reduced on these surfaces³³, thereby alleviating non-specific binding of cells to un-patterned areas of the slide. Additionally, if the patterned cells will ultimately be embedded within a hydrogel and transferred off the slide, the surface treatment is essential for reliable movement of the cell-laden hydrogel across the slide without distortion or tearing. Silanizing with (tridecafluoro-1,1,2,2-tetrahydrooctyl) dimethylchlorosilane results in the presence of hydrophobic fluoroalkyl groups on the slide surface.

CAUTION: Perform all steps from 3.1 onward in a chemical fume hood to prevent exposure to acetic acid and methylene chloride fumes.

1. Rinse slide with 10% acetic acid and then dry under an air stream.
2. In a glass Coplin jar, prepare a solution of 60 mL methylene chloride (dichloromethane), 0.6 mL of triethylamine, and 0.6 mL of (tridecafluoro-1,1,2,2-tetrahydrooctyl) dimethylchlorosilane. Stir with a metal spatula to mix.
   NOTE: These reagents are sensitive to water. They should be stored under dry conditions and used as fresh as possible.
3. Add the slide to the Coplin jar containing the silane solution. Place Coplin jar on an orbital shaker (set to 60-80 rpm) and allow the reaction of the silane and the slide to progress for 15 min.
4. Use metal forceps to remove the slide from the silane solution. Immerse slide in a Coplin jar containing methylene chloride for 1 minute to remove excess silane from the slide.
5. Immerse the slide in a 50 mL conical tube containing ethanol. Agitate. Immerse the slide in a 50 mL conical tube containing distilled water. Agitate.
   NOTE: Methylene chloride and water are not miscible, so an ethanol rinse is needed to remove excess methylene chloride before the final water rinse.
6. Remove the slide from the water and inspect it. The slide should be fairly dry, with any water droplets having a contact angle of greater than 90°. Allow slides to dry fully and store in vacuum desiccator until use.
   ​NOTE: The experiment can be paused here. Store the slide under dry conditions.

4. Prepare PDMS flow cells and slide for experiment

NOTE: Rectangular PDMS flow cells are used to concentrate the cells over the patterned regions of the slide. For experiments cultured in 3D, the flow cells form a mold for the hydrogel.

1. Make SU-8 master to use as mold for PDMS flow cells.
   1. Pre-heat hotplate to 95 °C.
   2. Add 5 mL of SU-8 2075 to a silicon wafer.
   3. Spin coat the SU-8 on the wafer at 500 rpm for 10s, followed by 1,000 rpm for 30s. This should create features up to 240 µm in height³⁴.
   4. Soft bake the wafer on the hotplate for at least 45 min.
   5. Remove the wafer from the hotplate. Put the photomask (see Supplemental File 4) (emulsion side down) on top of the wafer and weigh it down with a glass disc to ensure contact between the photomask and the slide.
   6. Expose with UV light (365 nm) for a radiant energy density of 350 mJ/cm².
   7. Bake wafer on the hotplate for 12-15 min.
   8. Place wafer in wide glass container. Cover wafer with SU-8 developer solution. Place on a shaker and develop while agitating for at least 15 min.
   9. Use forceps to remove the wafer from the developer solution. Rinse for 5 s by spraying more developer solution from a squirt bottle. Spray with isopropyl alcohol to rinse. If a white precipitate appears, return the wafer to the developer solution and develop for longer.
   10. Dry wafer under a stream of air or nitrogen.
   11. Bake slide for 5 min.
       ​NOTE: Once the master wafer has been created, it can be reused indefinitely as long as the features remain intact.
2. Prepare PDMS.
   1. In a weigh boat, add polydimethylsiloxane elastomer and crosslinker in a 10:1 ratio (by mass). Stir vigorously to ensure even mixing.
   2. De-gas the PDMS in a vacuum desiccator for 15-30 min until no more bubbles are visible.
   3. Place the master wafer in a 15 cm tissue culture dish. Pour PDMS over the wafer. If bubbles appear, de-gas in a vacuum desiccator for a few minutes.
   4. Bake in 60 °C oven for 3 h.
      ​NOTE: After baking, PDMS flow cells can be stored on the benchtop indefinitely.
3. Prepare PDMS flow cells for the experiment.
   1. Shortly before starting a CMO-DPAC experiment, cut out the required number of PDMS flow cells from the master wafer. Plasma oxidize with 10 cc/min room air for 90 s to render the surface hydrophilic.
   2. Cut out each individual flow cell so that there is 1-2 mm of PDMS remaining on each side, then cut open the top and bottom of the flow cell to create an inlet and outlet.
   3. Retrieve patterned slide created in Steps 2 and 3. Align on top of photomask.
   4. Using the photomask as reference, place the PDMS flow cells on the slide in the location of each patterned region.
   5. Add 50 µL of phosphate buffered saline (PBS) + 1% bovine serum albumin (BSA) to the inlet of each flow cell, as shown in Supplemental Figure 1H. Confirm that the flow cell is completely filled by the PBS + 1% BSA and that there are no large bubbles. Proceed immediately to Steps 5 and 6.
      ​NOTE: Blocking with BSA minimizes non-specific cell adhesion to the slide surface.

5. Lift and label cells with cholesterol-modified DNA

1. Prepare the cholesterol-modified DNA solutions.
   1. For each set of cells in the experiment, mix together 3 µL of a 100 µM stock solution of the cholesterol-modified Universal Anchor Strand with 3 µL of a 100 µM stock solution of an Adapter Strand. Incubate for 1 minute. This will pre-hybridize the oligos. Add 69 µL of phosphate-buffered saline (PBS) to create a 4 µM Universal Anchor + Adapter solution.
   2. For each set of cells in the experiment, add 3 µL of a 100 µM Universal cholesterol-modified Co-Anchor Strand stock solution to 12 µL of PBS, creating a 20 µM solution.
2. Prepare the single-cell suspension(s).
   1. For adherent cells, use trypsin or other dissociation agent to remove the cells from the culture flask. Add culture media to neutralize the trypsin and centrifuge to pellet the cells. For non-adherent cells, collect the cell suspension and centrifuge to pellet the cells.
   2. Resuspend the cell pellet in 1 mL of ice-cold PBS or serum-free media. Transfer 1-3 million cells to a 1.5 mL microcentrifuge tube. Centrifuge at 160 x g for 4 min.
      NOTE: If the cell type being used is prone to clumping/aggregation, use PBS without calcium and magnesium ions for all wash steps to reduce unwanted cell aggregation. If viability is a particular concern for the cell type being used, use serum-free media instead of PBS. Media containing fetal bovine serum is not recommended for cell labeling as it can hinder incorporation of lipid-modified oligos.³⁵
3. Label the cells with cholesterol-modified oligos.
   1. Resuspend the cell pellet in 75 µL of ice-cold PBS or serum-free media. Keep the cells in an ice bucket throughout the labeling and washing process to maximize cell viability and minimize loss of the cholesterol-modified oligos from the cell surface.
      ​NOTE: Resuspending the cells before adding the DNA ensures that the distribution of DNA is uniform across the cell population.
   2. Add the 75 µL of the 4 µM Universal Anchor + Adapter solution created in Step 5.1.1 to the microcentrifuge tube containing the cell suspension. Mix thoroughly by pipetting. Incubate for 5 min on ice.
   3. Add 15 µL of the Universal Co-Anchor Solution to the microcentrifuge tube. Mix thoroughly by pipetting. Incubate for 5 min on ice.
   4. Remove excess oligos from the cell suspension. Add 1 mL of ice-cold PBS or serum-free media to the microcentrifuge tube. Mix with a P1000 pipette. Centrifuge at 160 x g for 4 min at 4 °C. Discard the supernatant. Repeat two more times.
      NOTE: If cells are prone to clumping, pass the cell suspension through a 40 µm filter before the final wash. If cells are prone to adsorption onto the side of the microcentrifuge tube, consider pre-blocking the tube with casein.

6. Pattern the DNA-labeled cells

1. Resuspend the cells in ice-cold PBS or serum-free media to create a cell-dense solution of at least 25 million cells/mL.
   NOTE: For one slide using four of the 10 mm x 15 mm x 200 µm PDMS flow cells described in Step 4, about 100 µL of this dense cell suspension is required. Although most of these cells will not adhere to the pattern and will ultimately be discarded, having an extremely concentrated solution of cells over the pattern dramatically improves the efficiency of cell patterning.
2. Pick up slide and tilt it slightly. Add 25 µL of cell suspension to the inlet of each flow cell on the patterned slide. Remove the PBS + 1% BSA solution from the outlet, allowing the cell suspension to fill the PDMS flow cell. Incubate on ice or at room temperature for 30 s.
   NOTE: At this point, looking at the flow cell under a microscope should show densely packed cells with little to no gaps visible between cells. See Supplemental Figure 2B.
3. Aspirate 5 µL of cell suspension from the outlet of the slide and add it back into the inlet. Repeat 10 times per flow cell.
   NOTE: The adhesion of CMO-labeled cells to the DNA-patterned slide is nearly instantaneous. Flowing the cells over the pattern multiple times increases the probability that a cell will flow over a given DNA spot and be captured.
4. Gently pipette PBS or serum-free media into the inlet of each flow cell to wash out excess cells. Collect the cell suspension from the outlet. Repeat 2-4 times or until a visual inspection of the slide under the microscope confirms that there are no excess cells remaining.
   NOTE: It can be advantageous to save the excess cells from the first wash. If the patterning efficiency is unsatisfactory, the excess cells can be centrifuged and resuspended in a lower volume of PBS to create a more cell-dense solution, and then the process can be repeated from Step 6.2.
5. Repeat Steps 6.1-6.4 for each set of cells in the pattern. For patterns in which multiple cell types are directly patterned by the surface template, start with the least abundant cell type of the pattern and finish with the most abundant cell type.
   ​NOTE: It is advisable to do each round of cellular assembly sequentially instead of pooling the cells, even in conditions where the cells are all labeled with orthogonal DNA sequences. Pooling the cells effectively dilutes each cell population and reduces patterning efficiency.
6. After the final round of cell assembly is complete, the next steps will vary based on the specific experiment. If the cells are intended to remain on the glass, add media to a Petri dish containing the slide, and then gently use forceps to nudge the PDMS flow cells off of the slide. If the cells will be embedded into a hydrogel and cultured in 3D, proceed to Step 7.

7. Transfer into hydrogel for 3D culture (optional)

1. Prepare a hydrogel precursor solution containing 2% DNase.
   ​NOTE: The composition of the solution will vary based on experimental setup. Matrigel and mixtures of Matrigel and collagen I work well in this protocol, but other hydrogels are also possible.
2. Add 50 µL of hydrogel solution containing 2% DNase to the inlet of each flow cell. Aspirate the excess fluid from the outlet, driving the hydrogel solution into the flow cell. For viscous hydrogel precursors, tilting the slide slightly may be required to help the hydrogel flow into the flow cell.
3. Incubate the slide at 37 °C for 30-45 min (depending on hydrogel gelation kinetics) to allow the hydrogel to set and to cleave the DNA-based adhesion between the cells and the surface.
4. Remove each flow cell from the slide and place on top of hydrogel precursor solution.
   1. Add 50 µL of hydrogel precursor to a well of a 2-well chamber slide or a 6-well plate.
   2. Pipette 10 µL of PBS on either side of each flow cell.
   3. Use a razor blade or fine-point tweezers to distribute the PBS along the full length of the flow cell, then gently lift the sides of the flow cell so that the PBS rushes underneath the hydrogel.
      NOTE: This will "float" the hydrogel across the slide, allowing for transfer without distortion or tearing.
   4. Use a razor blade to gently move the flow cell to the edge of the glass slide.
   5. Invert the slide. With the razor blade, nudge the flow cell off the slide so that it lands on top of the razor blade.
   6. Pick the flow cell off the razor blade using curved forceps. Invert the flow cell so that the cells are on the bottom, and then place on top of the droplet of hydrogel precursor solution.
   7. Repeat Steps 7.4.1 – 7.4.6 for each flow cell.
5. Incubate for at least 30 min so that the hydrogel containing the patterned cells can bind to the hydrogel underlay, resulting in the full embedding of the patterned cells.
6. Remove the PDMS flow cell.
   1. Add enough media to immerse the PDMS flow cell.
      ​NOTE: The influx of media will loosen the adhesion between the hydrogel and the PDMS flow cell.
   2. Use curved forceps, oriented along the long axis of the flow cell, to gently nudge the flow cell until it pops off and floats into the media. Collect the flow cell with forceps and discard.
      ​NOTE: For optimal results, spread the curved forceps and apply gentle pressure to the walls of the PDMS flow cell. Apply force in the direction of the long axis of the flow cell.

8. Confirm successful labeling of cells with CMO (optional, for troubleshooting)

1. Order a fluorescently modified (FAM or AF647) oligonucleotide that is complementary to the surface adhesion sequence of the Adapter Strand being used in the experiment.
2. Label cells with CMO DNA and wash out excess DNA as described in Step 5. Resuspend in 200 µL of ice-cold PBS.
3. Make up a 4 µM solution of the fluorescently labeled complementary oligonucleotide in PBS. Add 200 µL of this solution to the cell suspension. Incubate on ice for 5 min.
4. Add 1 mL of ice-cold PBS. Mix. Centrifuge the cells to pellet them. Remove supernatant. Repeat this process two more times to wash out any DNA that has not hybridized.
5. Perform analytical flow cytometry to quantify the presence of DNA on the cell surface.
   1. On a flow cytometer, analyze control cells that have not been labeled with DNA. Set up gates based on this population.
   2. Analyze CMO-labeled cells that have been treated with a fluorescently labeled complementary oligonucleotide.
   3. Calculate mean fluorescence intensity.