Micropatterning Transmission Electron Microscopy Grids to Direct Cell Positioning within Whole-Cell Cryo-Electron Tomography Workflows

The protocol described here is a compilation of the cell culture, micropatterning, and imaging methods used by the Wright lab and the Cryo-EM Research Center at the University of Wisconsin, Madison. The workflow is presented in Figure 1. Additional training and instructional materials are available at the following sites: https://cryoem.wisc.edu or https://wrightlab.wisc.edu

1. Preparation of grids for patterning

1. Transfer the TEM grids onto a clean glass slide, carbon side up (the standard thickness of the carbon foil is 12 nm). Using a carbon evaporator, ACE600, evaporate 5-8 nm of additional carbon onto the grids to increase overall carbon film durability.
   NOTE: This step is not necessary for SiO₂ grids. This step may also be done in advance; store the coated grids in a low humidity environment such as a vacuum desiccator.
2. Transfer the grids to a grid prep holder and glow discharge the grids carbon side up. Using a glow discharge system, glow discharge the grids for 60 s at 10 mA with an 80 mm working distance and vacuum pressure of 1.0 x 10^-3 mbar. Do this within 15-30 min of the next step.
   NOTE: Grid prep holders can be commercially bought or homemade with a piece of filter paper on a small Petri dish.

2. Application of the anti-fouling layer

NOTE: Proper sterile technique should be used when handling the grids, and all solutions should be sterile and/or filter sterilized.

1. Transfer the grids (carbon side up) to a clean glass slide or coverslip with at least 1 cm of separation between the grids. Pipette 10 µL of 0.05% poly-L-lysine (PLL) onto each grid. Incubate the grids in a humid chamber, such as an enclosed plastic box with moist paper towels, for at least 30 min.
   NOTE: This step can be extended to overnight. Be sure the humidity level in the chamber is sufficient to prevent the grids from drying out.
2. Wash each grid three times with 15 µL of 0.1 M HEPES pH 8.5. For each wash, remove most of the liquid from the grid with a pipette without letting the grid dry. Add 15 µL of fresh buffer, incubate for at least 30 s and repeat. Leave each grid in 15 µL of 0.1 M HEPES after the final wash.
   NOTE: In this step and future steps, it is important to keep the grid wet and to avoid contact between the pipette and the grid.
3. Prepare 10 µL of 100 mg/mL polyethylene glycol-succinimidyl valerate (PEG-SVA) in 0.1 M HEPES pH 8.5 for each grid. The PEG-SVA will dissolve quickly with gentle mixing resulting in a clear solution.
   NOTE: Do not prepare the PEG-SVA solution in advance. PEG-SVA has a half-life of 10 min at pH 8.5. Avoid exposing the PEG-SVA stock to excessive moisture by storing it in a desiccator or dry environment at -20 °C and warming to room temperature before opening.
4. Immediately after preparing the PEG-SVA solution, remove the 15 µL drop of HEPES pH 8.5 from each grid (taking care not to dry the grid) and add a 10 µL drop of the PEG-SVA solution. Incubate the grids in a humid chamber for at least 1 h.
   NOTE: This step can be extended to overnight. Be sure the humidity in the chamber is sufficient to prevent the grids from drying out.
5. Wash each grid three times with 15 µL of sterile water. For each wash, remove most of the liquid from the grid with a pipette without letting the grid dry, add 15 µL of fresh water, incubate for at least 30 s and repeat. Leave each grid in 15 µL of water after the final wash.

3. Applying PLPP gel

1. Prepare a clean microscope coverslip for each grid. Complete the following steps for each grid, one grid at a time, to minimize the chance of the grid drying out.
2. Place a 1.0 µL drop of water on the center of the coverslip to help with placing the grid on the coverslip and keeping the grid wet. Carefully transfer the grid from the 15 µL water drop to the 1.0 µL water drop on the coverslip. Be sure to place the grid carbon side up.
3. Carefully place a polydimethylsiloxane (PDMS) stencil over the grid, taking care to keep the grid centered and to minimize stencil contact with the carbon foil of the grid.
4. Add 1.0 µL of 4-benzoylbenzyl-trimethylammonium chloride (PLPP) gel onto the grid. Pipette gently to mix (do not touch the grid with the pipette tip).
5. Move the coverslip with the grid to a dark location to dry. The gel will dry in approximately 15-30 min.

4. Calibration and design of the micropattern

1. Color one side of a glass coverslip with a highlighter. Add black lines from a fine-tipped permanent marker to make focusing easier. Place the coverslip on the microscope such that the colored side faces the objective lens. Using brightfield mode, focus on the highlighter.
2. Ensure the microscope and the micropatterning system is powered on, and the correct light path is set. Open Micromanager and the Leonardo software (Plugins > Leonardo) on the microscope computer.
3. Select calibrate and follow the on-screen instructions. Adjust the microscope focus so that the image projected onto the slide is in focus. The exposure time may need to be decreased. After calibration, select Pattern Now.
4. Record the micrometer/pixel (µm/px) ratio reported under calibration data in the top left window of the program (Figure 2, area 1). Use this ratio to determine the number of pixels to use per micrometer when designing a pattern.
5. After calibration, ensure that the software is now open with a live brightfield view from the microscope. Load a prepared grid on a coverslip (section 3) onto the stage with the grid facing the objective lens. Position the stage and adjust the focus so that the grid is visible in the software window.
6. Measure the size of the grid squares and grid bars in micrometers. The software includes a ruler activated by the button near the bottom left corner to measure the grid (Figure 2, area 2). For example, the patterns used here for a 200 mesh grid correspond to ~87 × 87 µm grid squares and ~36 µm grid bars.
   NOTE: The software offers flexibility in resizing patterns on the fly, so minor inaccuracies in measurement can be tolerated.
7. Based on the measurements and ratios above, create pattern(s) with any image creation software. The minimum feature size with a 20× objective is 1.2 µm. Patterns should be saved as uncompressed 8-bit .tiff files.
   1. Ensure the software does not rescale images to a different pixel size when saving. The pattern should fit within an 800 × 800 pixel box, sufficient to cover four grid squares.
      ​NOTE: Pixels with a value of 255 (white) will be patterned at the highest intensity (total dose of the laser) and pixels with a value of zero (black) will not be patterned. Any pixels with an intermediate value will be patterned with a dose of approximately (X/255)*total dose. In Figure 3A, pixel values of 255 and 129 were used for the greyscale patterns. Once the pattern is designed it can be saved and reused without modification.

5. Micropatterning

1. After calibration, ensure that the software is now open with a live brightfield view from the microscope. Load a prepared grid on a coverslip (section 3) onto the stage with the grid facing the objective lens. Position the stage and adjust the focus to see the grid in the software.
2. For an initial run, design a new template. In the software, select Add ROI (not shown, in the location of Figure 2 area 3) and choose a 3,000 µm circle. Position the circle ROI over the grid using the brightfield image on the screen as a guide. Press lock to secure the ROI.
3. After locking the ROI in place, select Add Pattern (not shown, in the location of Figure 2 area 3). Choose the pattern designed in section 4. Divide the grid into six regions to allow independent focusing and positioning in each region to factor for uneven grids. An 8 × 8 grid square region for each corner of the grid and a 2 × 8 grid square region on each side of the center, leaving the center four grid squares unpatterned (Figure 2, center image).
4. Use the replication options (Figure 2, area 4) to generate copies of the initial pattern to reach the desired number of total copies of the pattern. Adjust the spacing between copies to match the spacing between grid squares if necessary.
5. Set the Total Dose for the pattern. 30 mJ/mm² is a good starting point. See the discussion section for more details.
6. Under Expert Options (Figure 2, area 4) adjust the angle of the region to match that of the grid squares. Regions can be repositioned using the mouse. The ratio (size) of the patterns can also be adjusted. Iterate through adjusting the angle, position, space between, and ratio of the pattern until the patterns line up with the grid squares. Move the microscope stage to change the region of the grid in the live brightfield display.
7. Press Lock to save the changes made to the region.
8. To copy a region, click the Duplicate button (Figure 2, area 5, two sheets of paper icon) next to its name in the Actions panel on the left. To reposition, rename, or edit the copy, click on its name in the Action panel.
9. Repeat steps 5.4-5.9 as necessary to fill all of the desired regions.
10. Once the complete template is designed and positioned, Save the template file within the software (Figure 2, area 6, bar with up arrow icon in top toolbar).
11. When loading a previously saved template (bar with down arrow icon) center the ROI over the grid and press Lock. Click on each region in the Action Panel to change the angle, position, dose, and/or pattern file.
12. Once the template and patterns are positioned, uncheck all but one of the regions in the Action Panel in the software.
13. Use the microscope stage to navigate to that region and focus on the carbon foil. Clicking the Eyeball icon in the Action panel (Figure 2, area 5) will toggle the display of the pattern overlay on or off.
14. Once the grid is in focus, close the brightfield shutter and press the Play icon in the bottom right corner of the software to begin the patterning process, which can be monitored live.
15. In the action panel, select the box for the next region. Open the brightfield shutter so that the grid is visible and center that region using the microscope stage. Repeat steps 5.13-5.14 for each region in the Action Panel.
16. Remove the coverslip with the grid from the microscope, and immediately pipette 10 µL of sterile phosphate-buffered saline (PBS) onto the grid.
17. After 10 min, remove the stencil with tweezers, then wash the grid 3x with 15 µL of PBS. After the final wash, place each grid in 15 µL of PBS and move the grids to a dark location.

6. Deposition of ECM proteins

1. For cultured cells, follow steps 6.2-6.5; for primary Drosophila neurons, follow steps 6.6-6.10.
2. Prepare at least 15 µL of ECM for each grid. For BEAS-2B cells, prepare a final concentration of 0.01 mg/mL bovine fibronectin and 0.01 mg/mL fluorophore-conjugated fibrinogen in sterile PBS. For HeLa cells, prepare 0.01 mg/mL bovine collagen I and 0.1 mg/mL fluorophore-conjugated fibrinogen in sterile PBS.
3. Remove most of the PBS from each grid and apply 15 µL of the ECM. Incubate the grid in a humid chamber at room temperature for at least 1 h.
   NOTE: This step may be extended to overnight at 4 °C.
4. After incubation in ECM, wash each grid 5x with sterile PBS. For each wash, remove most of the liquid with a pipette without letting the grid dry, add 15 µL of fresh PBS, incubate for at least 30 s, and repeat. Leave each grid in PBS after the final wash.
   NOTE: Grids can be stored for up to a week in PBS at 4 °C with no observed deterioration in quality.
5. Use a fluorescence microscope to detect the fluorophore in the ECM to confirm patterning and that the carbon foil remained intact. A few broken squares are generally tolerable.
6. For primary Drosophila neurons, move the patterned grids to a 30 mm glass bottom dish containing sterile PBS.
7. Aspirate the PBS from the dish and apply 2 mL of 0.5 mg/mL fluorophore-conjugated concanavalin A. Incubate overnight at 25 °C in a sterile environment.
8. Remove the concanavalin A solution from the dish (without drying the grids) and wash the grids 3x with PBS. For each wash, add and remove 2 mL PBS from the dish.
9. Use a fluorescence microscope to detect the fluorophore in the ECM to confirm patterning and that the carbon foil remained intact. A few broken squares are generally tolerable.
10. After the final wash, remove the PBS from the glass bottom dish and add 2 mL of freshly-prepared, sterile-filtered supplemented Schneider's Drosophila media²¹, containing 20% heat-inactivated fetal bovine serum (FBS), 5 µg/mL insulin, 100 µg/mL penicillin, 100 µg/mL streptomycin, and 10 µg/mL tetracycline. Incubate at 25 °C in a sterile environment until the neurons are ready to be plated.

7. Preparation of primary Drosophila cells prior to seeding

1. Sterilize a 55 mm dissection dish with 70% EtOH, and then add to the dish 2-3 mL of sterile-filtered 1× dissection saline (9.9 mM HEPES pH 7.5, 137 mM NaCl, 5.4 mM KCl, 0.17 mM NaH₂PO₄, 0.22 mM KH₂PO₄, 3.3 mM glucose, 43.8 mM sucrose)²¹.
2. Pick 30-40 3^rd instar larvae gently from the food using a pair of tweezers.
3. Place the larvae into the tube with 1× PBS, then transfer them into the second tube with 1× PBS to wash the larvae.
4. Transfer the larvae into the tube with 70% EtOH to wash off the PBS, then transfer them into the second tube with 70% EtOH. Leave the larvae in the second tube for 2-3 min to sterilize the larvae.
5. Transfer the larvae into a tube with 1× dissection saline, then immediately transfer them into the second tube with 1× dissection saline.
6. Transfer individual larvae onto the dissecting dish containing 1× dissection saline. With a pair of forceps and a dissection microscope, quickly tear each larvae to extract the brain and transfer it to the third tube with 1× dissection saline. Repeat until all brains are extracted.
7. Centrifuge the tube containing the brains at 300 x g for 1 min.
8. Discard the supernatant, and wash with 1 mL of 1× dissection saline and centrifuge the tube at 300 x g for 1 min. Repeat this step one more time.
9. Discard the supernatant until 200-250 µL is left in the tube and add 20 µL of 2.5 mg/mL Liberase in 1x dissection saline.
10. Rotate the tube on a rotator for 1 h at room temperature; during this hour, pipette the solution 25-30 times every 10 min. By the end, the solution should be slightly opaque.
11. Centrifuge the cells at 300 × g for 5 min.
12. Discard the supernatant, then add 1 mL of supplemented Schneider's media. Pipette the solution 30 times to mix.
13. Centrifuge the cells at 300 × g for 5 min.
14. Discard the supernatant and wash the cell pellet by adding 1 mL of supplemented Schneider's media. Pipette the solution 30 times to mix.
15. Centrifuge the cells at 300 × g for 5 min.
16. Discard the supernatant, then resuspend the cell pellet with 300 µL of supplemented Schneider's media. Pipette the solution 30-40 times to mix.

8. Culture and RSV infection of BEAS-2B and HeLa cells

1. Maintain HeLa cells and BEAS-2B cells in T75 flasks at 37 °C and 5 % CO₂. Passage cells every 3-4 days once reaching approximately 80% confluency. Maintain HeLa cells in DMEM + 10% FBS + 1× Antibiotic-Antimycotic. Maintain BEAS-2B in RPMI + 10% FBS + 1× Antibiotic-Antimycotic^6,22,23.
2. For seeding of uninfected cells, skip to section 9. BEAS-2B and HeLa cells are susceptible to RSV infection; BEAS-2B cells were used for all experiments involving RSV shown here.
   NOTE: Perform all BSL-2 steps in compliance with institutional protocols using an appropriate biosafety cabinet (BSC) and personal protective equipment (PPE).
3. Prior to RSV infection of cells, passage 5 × 10⁴ cells per well into a 6-well plate (surface area ~9.6 cm²) with 2 mL of growth media and incubate overnight.
4. Trypsinize and count one well of cells. To trypsinize, aspirate media from one well and wash with 2 mL of sterile PBS without Mg²⁺ and Ca²⁺ to remove residual media. Add 500 µL of 0.25% trypsin solution. Incubate at 37 °C for 5-10 min. Periodically check the cells to see if they are released from the surface. Once the cells are released, add 1.5 mL of culture media.
5. Mix 100 µL of trypsinized cells with 100 µL of trypan blue. Pipette 10 µL of diluted cell mix into a hemocytometer. Count the cells and calculate the number of cells per well. Use this number to calculate MOI below.
6. Prepare a dilution of RSV-A2mK+²⁴ in growth media to achieve an MOI of 10 per-well in 750 µL of media. The MOI of RSV-A2mK+ can be calculated from fluorescent focus units (FFU) titers of the stock (For example: for 1.0 × 10⁵ cells per well and an RSV stock of 1.0 × 10⁸ FFU/mL, dilute the viral stock 1:75 to 1 × 10⁶ FFU/750 µL or 1.33 × 10⁶ FFU/mL).
7. Aspirate the media from the cells in the 6-well dish and add 750 µL of the viral solution from above to each well.
8. Rock the plate at room temperature for 1 h.
9. After 1 h, bring the total volume per well up to 2 mL with growth media pre-warmed to 37 °C and place the plate in an incubator set to 37 °C with 5% CO₂ for 6 h.
10. Trypsinize the cells to release them and proceed to seeding as described below. After seeding, incubate the grids for an additional 18 h before plunge freezing (for a total 24 h post-infection).

9. Cell seeding onto micropatterned grids

1. For cultured cells, follow steps 9.2-9.8; for primary Drosophila neurons, follow 9.9-9.11.
2. Trypsinize the cells to release them (see step 4 in section 8 above). To reduce cell aggregation, trypsinize the cells at 60 % or less confluency.
3. Mix 100 µL of trypsinized cells with 100 µL of trypan blue. Pipette 10 µL of the diluted cell mix into a hemocytometer and count the cells.
4. Dilute the cells in media to 2 × 10⁴ cells/mL.
5. Add 1 µL of media to the center of a 30 mm glass bottom dish to assist in placing the grid and prevent it from drying. Transfer the grid from the PBS on the coverslip to the center of the glass bottom dish. Add 10 µL of cell solution to the grid.
6. Using a brightfield microscope, observe cell adhesion to the grid after 5 min. If a majority of the patterns remain unoccupied, add an additional 10 µL drop of the cell solution. Keep the grids and cell solution at 37 °C during the incubations.
7. Repeat step 9.6 until most patterns are occupied or many occupied patterns begin to have multiple cells. Incubate the grid for 2 h in the incubator (37 °C, 5% CO₂).
8. Flood the dish with 2 mL of pre-warmed media and incubate overnight (37 °C, 5% CO₂).
9. For primary Drosophila neurons, remove the media from the grid-containing dish and plate the cells onto the dish.
10. Wait 30-60 min for the cells to attach, then add 2 mL of supplemented Schneider's media.
11. Culture the neurons in a 25 °C incubator for a minimum of 2-3 days before plunge-freezing.

10. Imaging and vitrification of patterned grids

1. Place the glass bottom dish containing the patterned grid and cultured cells on the fluorescence microscope.
2. Acquire images of the grid using brightfield and the appropriate fluorescent channels to detect the pattern and any other labeling in the cells. Ensure the cell density and positioning is suitable for downstream imaging and analysis.
   NOTE: Brightfield and fluorescent images were processed in the FIJI software package²⁵.
3. Prepare a cryo-plunge freezer; the type of freezing device will depend on availability, cost, and features that are most suitable for the sample.
   NOTE: Primary Drosophila neurons were prepared on an automated plunge-freezer, and the BEAS-2B cells were prepared using a semi-automated plunge-freezer.
4. Apply gold fiducials to the samples for proper alignment of the tilt series. Blot samples to remove excess media, then plunge-freeze the samples into a cryogen, such as liquid ethane cooled by liquid nitrogen. For primary Drosophila neurons, blot for 4 s from the backside. For HeLa and BEAS-2B cells, blot from both sides for 4-6 s. The frozen grids can then be stored in liquid nitrogen until further use.
5. Image vitrified cells in a cryo-electron microscope, operated at 300 kV with a direct electron detector camera. Set up tilt-series collection for each region of interest with software such as SerialEM²⁶ for cryo-EM/cryo-ET data collection.
   NOTE: Tilt-series of primary Drosophila neurons were collected on a direct electron detector from -60° to 60° bidirectionally at 2° increments at -8 µm defocus with a pixel size of 4.628 Å for a total dose of 70-75 e^–/Ų. Tilt-series of RSV-infected BEAS-2B were collected on a direct electron detector with an energy filter (20 eV slit) at -5 µm defocus with a pixel size of 4.603 Å and a total dose of ~80 e^–/Ų.
6. Process the tilt series to reconstruct tomograms.
   NOTE: Tomograms presented here were reconstructed using the IMOD package²⁷; lowpass filtering was done using the EMAN2 software package²⁸.