Sample Preparation for In Situ Cryotomography of Mammalian Cells

1. Grid preparation

NOTE: In the experimental design, plan for a maximum of 8-12 grids total per plunge-freezing session and 4-5 grids per well. More than that will lead to a very long plunge-freezing session, which may cause increased cell stress, ice contamination, and user errors.

1. Glow discharge
   1. Load an appropriate number of grids with a perforated carbon support film to be glow discharged at 15 mA for 45 s at 0.39 mBar atmosphere in a glow discharge device.
   2. Make sure the carbon side of the grid is facing up. When finished, transfer the grids to a clean container lined with filter paper (Figure 1A1).
      NOTE: Other grids may be used as long as the material is not toxic to cells (which excludes any copper grid). Finder grids are useful for correlative studies. Grids made of other materials are also possible. Many labs have had good results growing cells on top of SiO₂ support grids^8,9.
2. Adherent treatment
   1. Transfer the grids, fibronectin, PBS, plates, and tweezers to a biosafety cabinet. Treat both sides of the grids with 20 µg/mL bovine fibronectin by micropipetting two generously sized dots of adherent solution on a sterile surface, such as the lid of a 6 well plate. Ensure that the dots are large enough to envelop the entire grid (around 100 µL is recommended).
      NOTE: Prior to treatment with the selected adherent solution, there is the option of photo-micropatterning the grid here to optimize the attachment of cells on the center of grid squares. Reducing the ability of cells to attach to the bars will increase the number of imageable cells. This is ideal for experiments involving cryoFIB milling.
   2. Use 5/15 tweezers to manipulate the grids. Be sure to treat both sides of the grid in solution.
      ​NOTE: Other adhesive solutions may be used (fibrinogen, collagen, other extracellular matrix components, etc.). Fibronectin is used here, as it gives good results with a wide variety of cell lines. (U2OS, HeLa, Vero, Calu-3, Tzm-bl). The 5/15 tweezer is useful as its geometry allows improved maneuverability around the 6-well plates, 35 mm dishes, and micro slides. This results in less mechanical stress on the grids and a more intact carbon film. Make sure all required materials are placed in the biosafety cabinet prior to beginning, as this reduces the possibility for grid contamination.
3. Attaching grids
   1. Once treated with bovine fibronectin, the grid will become sticky. Using 5/15 tweezers, gently touch the non-carbon surface of the grid to the bottom of an empty dish/plate and open the tweezers. The grid should easily attach to the new surface (Figure 1A2).
      NOTE: Try to avoid placing grids in the direct center or the edges of the dish/plate. When adding cells, there is a tendency for cell density to accumulate in the center of the plate. This can result in grids with excessive cellular density. Alternatively, 3D-printed grid holders may be used here instead of attaching cells directly to the surface of the dish/plate¹².
4. Incubation
   1. Incubate the grids for 30 min in the fibronectin adherent solution (20 µg/mL). To ensure that the grids do not become dry during this process, micropipette 1-2 drops of more adherent solution directly on top of grids every 15 min (Figure 1A3).
      NOTE: The incubation period will vary depending on the adherent solution used. For bovine fibronectin, the recommended time is 30 min.
5. Washing
   1. Following the incubation period, remove excess adherent solution via micropipetting. Wash the grids by micropipetting drops of PBS directly on top of the grids. Repeat this 2-3 times. (Figure 1A4).
6. Sterilization
   1. Use UV light to sterilize the grids in the biosafety cabinet for 1 h. Place the grids as close to the UV source as possible to maximize sterilization (Figure 1A5). To ensure that the grids do not become dry during this process, micropipette 1-2 drops of PBS directly on top of grids every 10 min.
      ​NOTE : Steps 1.4-1.6 can be performed concurrently.

2. Seeding grids

1. Count cells:
   1. Detach and count the cells using mechanical or enzymatic methods. Determine the optimum number of cells to use for seeding prior to counting. For U2OS seeded in a 6-well plate, use anywhere between 6 x 10⁴ to 1.6 x 10⁵ cells per well (Figure 1B1).
      NOTE: The number of cells seeded will vary according to the cell type, the surface area of the well, dish or microslide used, the time between seeding and plunge freezing, and the experimental design. Test a range of cell numbers to identify conditions that will result in 0.25-1 cells per grid square at the time of plunge freezing. It is important to note that high cell density reduces the speed of heat transfer and may impact the vitrification process. To combat this, some labs have found success using cell strainers, which act to prevent cell clumps from forming¹³. Finally, this protocol results in the random attachment of cells to the carbon layer. If a targeted attachment is necessary, photo-micropatterning may be used (see step 1.3)¹⁴.
2. Adding cells and growth medium : Add cells via micropipette to the wells, avoiding bubbles (Figure 1B2). In a 6-well plate, a total well volume of 1.5-2.0 mL is recommended.
   NOTE: It is recommended to wet around the grids carefully before adding the entire volume. This will help prevent grids from detaching and floating in the solution. Gently move the plate side-to-side to facilitate homogeneous cell distribution on the well. Do not swirl the plate in a circular motion, as this will result in excessive cell density allocating in the center of the well.
3. Incubation: Incubate the slides at 37 °C for an appropriate time according to the experimental design, which will depend on downstream applications. For this example (transfection of HIV molecular clones), incubate between 16-24 h (Figure 1B3).
   ​NOTE: Longer incubation times will result in more cell division cycles. Therefore, adjust the starting number of cells for seeding accordingly.

3. Transfection

1. Preparing transfection mixture:
   1. Prepare an appropriate cationic liposome reagent for plasmid transfection into the selected cell line.
   2. In this transfection example, use 1 µg of total plasmid DNA per well (in a 6 well plate) to be transfected at a 1:3 ratio of HIViΔEnv plasmid to psPAX2 plasmid. For each well, dilute 1 µg of DNA into 50 µL of serum-free DMEM. Homogenize the mixture via repeatedly micropipetting or gentle swirling of the plate.
   3. For each well, dilute 3 µL of transfection reagent in serum-free DMEM, then homogenize the mixture again by micropipetting or gentle swirling. Add the entire diluted transfection reagent mixture to the diluted DNA mixture (step 3.1.2) and incubate at room temperature (RT) for 15 min (Figure 1C1).
      NOTE: While this mixture is incubating, replace the media in the wells with 1.5 mL of fresh DMEM. Be careful not to dislodge the grids from the bottom of the well.
2. To apply the transfection mixture, pipette 100 µL of the mixture from step 3.1.3 dropwise to each well, concentrating the drops over the grids to ensure contact (Figure 1C2). Change the growth medium 16-24 h after transfection (Figure 1C3).

4. Plunge freezing

NOTE: Keep cells at 37 °C until needed for blotting. Additionally, be sure to make a note of the carbon side of the grids throughout the process.

1. Cell density
   1. Check the grids in an inverted optical microscope and note cell densities on each grid.
      NOTE: Grids with less than 0.25 cells per grid square (or one cell every 4 grid squares), are considered 'empty'. Grids with 0.25-1 cells per grid square are considered 'normal'. Grids with more than 1 cell per grid square are considered 'full'.
   2. Take note of the cell density on each grid to allow the dialing of blotting times later during plunge-freezing. Add 2-3 mL of PBS to an empty plate/dish and warm to 37 °C.
2. Preparing fiducials
   1. To prepare gold fiducials for downstream cryotomography applications, pipette 50 µL of 10 nm colloidal gold bead solution into a clean 1.5 mL tube. Add 2 µL of 1 mg/mL BSA to the tube and homogenize by micropipetting repeatedly.
   2. Centrifuge the tube at 15,000-20,000 x g for 15 min. Micropipette to remove as much of the supernatant as possible without disturbing the pellet.
      NOTE: The pellet will be very loose.
   3. Add 50 µL of PBS to the pellet and resuspend. Centrifuge the tube again at 15,000-20,000 x g for 15 min.
   4. Using a 10 µL tip, directly aspirate 4 µL of the pellet, and transfer it to a clean 1.5 mL tube (Figure 1D1). Use 1 µL of the washed and concentrated gold per grid.
      NOTE: The type of fiducials used depends on downstream applications. For TEM tomography, use 10 nm gold beads. For soft X-ray tomography, use 200 nm gold beads. For correlation microscopy, use fluorescent latex beads. Fiducials are normally not necessary for experiments involving cryoFIB milling, as the milling process will destroy them.
3. Blotting
   1. Set up the environmental chamber to 95% humidity and 30°C (Figure 1D1). Select a grid using the 5/15 tweezers.
   2. Wash the grids with PBS and transfer them to plunging tweezers (Figure 1D2). Once secure, remove the 5/15 tweezers and slide the clamp on the plunging tweezers.
   3. Insert the grid into the plunge freezer, keeping the clamp secure (Figure 1D3). Add 1 µL of gold fiducials to the back side of the grid (Figure 1D4). Add 2-3 µL of PBS to the carbon side of the grid. Use automated blotting to blot the back side of the grid and plunge freeze into liquid ethane (Figure 1D5).
      NOTE: It is recommended to have between 3-4 µL of volume per grid for optimal blotting. Grids may retain a variable amount of PBS after washing. Adjust the added volume of PBS to account for pre-existing liquid retention on the grid. It is recommended that the gold fiducials be added to the back of the grid. Addition to the front can result in pools of gold fiducials near the point of insertion, whereas addition from the back results in a more homogenized solution. Blotting duration depends on the cell dentistry of the grid. 2 s for empty, 3 s for normal, and 4 s for full grids, respectively. The type of plunge freezer available may determine how blotting is approached: The Leica GP2 plunge freezer is the one used in this protocol and is capable of automated single-sided blotting by default. Thermo Fisher Vitrobot does automated double-sided blotting, though this often damages the cells attached to the carbon side. Manual blotting in the Vitrobot is also possible, as well as manual gravity plungers.