Cell activation and immunostaining

Materials needed

Extruder set (Avanti Polar Lipids Cat #610000)

0.1 μm polycarbonate membranes (Avanti Polar Lipids cat. #610005)

Ibidi silicone 8-well chamber (Ibidi cat. #80841) 

24 x 60 mm No 1.5 Coverglass

Lipids (see table below)

Streptavidin (Invitrogen cat. #43-4303)

Biotinylated pMHC monomers (collaborator)

His tagged ICAM (Sino Biological cat. #10346-H08H)

Chemicals: 1M KOH, Ethanol, acetone, PBS, 1M NiCl₂, 1M CaCl₂, 5% BSA in PBS

Bilayer preparation

All phospholipids were purchased from Avanti Polar Lipids.

Phospholipids are dissolved in chloroform and stored in glass vials with Teflon Seal/cap at -80°C.

Making Liposomes

1. In a glass vial mix stock phospholipids in choloroform to a molar ratio of 2% DOGS-NTA, 1% biotin DOPE, 97% DOPC, 0.5% DOPE-PEG5000 and a total amount of 1 mg phospholipid mixture.

2. In a fume hood evaporate the chloroform off using a nitrogen gas stream (dried lipids at the bottom of the tube have a white waxy appearance).

3. Leave the nitrogen on gently and dry for further 30 min to completely remove chloroform, alternatively leave under vacuum overnight.

4. Resuspend in 1 mL ultrapure water.

5. Vortex briefly and leave to hydrate for 0.5-1 hour

6. Vortex again to achieve a mixture of cloudy looking lipids 

7. Clean lipid extruder.

Critical - Ensure that the extruder is clean. Pump through EtOH 3x and then ultrapure water 3x. Ensure that no extra water remains in the syringe chambers. Remove plunges to dry if necessary.

Place one white cylinder inside the metal chamber. Then load a filter support dipped in ultrapure water. Then load the 0.1 μm membrane (shiny side up) into the chamber on top of the filter support. This is easiest if the membrane is kept dry. It will stick nicely to the wet filter support upon contact. Dip another filter support into milliQ and stick it to the second white cylinder, then slide both into the metal chamber on top of the membrane.

Critical - You must be aware of the orientation of the extrusion membrane. There is a shiny side and a dull side. We typically load the nut and bolt so that the shiny side is facing the nut side. Therefore, this is the side that you should first load first. After 15 extrusions, the liposomes will be on the ‘bolt’ side of the extruder, corresponding to the dull side of the membrane.

8. Remove air from the centre of the chamber.

Fill the first syringe (nearest the metal nut) with 1 ml ultrapure water and gently and smoothly extrude the water into the second syringe once. This will remove the air from within the chamber into the second syringe (notice the air bubble). Discard the water from the second syringe and re-attach. Reload the first syringe with the liposome mixture.

9. Make liposomes by extrusion through 0.1 μm filter (extrude 15 times). Push gently and smoothly throughout, avoid jerky movements and bubbles.

10. Store them in glass tube purged with nitrogen at 4°C until use (they should last for a week).

Preparing glass coverslip

1. Clean a 24 mm x 60 mm No. 1.5 coverglass by rinsing in ultrapure water

2. Using an ultrasonic bath, sonicate in 1M KOH for 20 min

3. Wash 3x in ultrapure water

4. Sonicate in ultrapure water for 15 min

5. Sonicate in 100% ethanol for 15 min

6. Sonicate in acetone for 15 min.

7. Dry cover glass in 70°C oven for ~3 hr or overnight.

8. Using expanded tabletop plasma cleaner (Harrick Plasma) plasm clean cloverslip for 3-5 min. Plasma cleaning leaves the surface of the glass highly hydrophilic and wettable which is critical for the charged interaction between phospholipid headgroups and the coverslip surface and hence for proper supported lipid bilayer formation. Phospholipids should be added within 15-20 min of plasma cleaning to ensure best results.

9. Attach a sticky Ibidi silicone 8-well chamber (Ibidi cat #80841) with base removed to the coverslip.

Preparing supported lipid bilayers

1. Dilute the liposome stock 1 in 5 (final lipid concentration is 0.2 mg/mL) in ultrapure water

2. Add calcium chloride to 10 mM (10 ul from 1M stock) 

3. Quickly invert twice and add 250 uL into each well of the plasma cleaned coverslip (see previous section) and incubate for 30 min at room temp.

4. Wash the well with 10 ml of ultrapure water using a continuous stream and a 10 ml serological pipette. Do this by:

(i) Filling each chamber to the top gently with ultrapure water before you add the 10 ml pipette tip. This will help prevent air bubbles which will destroy the membrane.

(ii) Hold the chamber over a glass beaker to catch milliQ as it overflows the well and runs off.

(iii) Use the pipette gun on low speed to prevent bubble formation.

(iv) Do not use the final ml of wash - use it as a back stop to ensure that a bubble of air is not blown into the well.

(v) Do not touch the bottom of the well with the pipette.

5. Remove the water from the well using a pipette, leaving 100 μl behind.

Note: never expose supported lipid bilayers to air and always leave ~100­­–150 μl of liquid in the well in all steps.

6. Wash gently in 1 ml of 1 mM EDTA pH 8.0.

7. Washing gently in 1ml of ultrapure water.

8. Add 250 μl 1 mM NiCl₂ in ultrapure water and incubate for 10 min at room temp.

9. Wash the bilayer gently in 1ml ultrapure water. As above.

10. Add 250 μl of streptavidin (1 μg/ml) in PBS with 1% BSA

11. Wash gently 3x with 1ml PBS. As above.

12. Add 250 μl of biotinylated protein (1 ug/ml biotinylated pMHC per well) and His tagged ICAM-1 (1 ug/ml per well) in PBS with 1% BSA and allow to bind for 30 mins at room temp.

13. Wash the bilayer gently 3x with 1 ml PBS

Note: The bilayers should be used straight away, as leaving them over night can affect lipid mobility.

Cell Activation and staining

Protocol is written for cell sample preparation in 8-well Ibidi chambers (Ibidi cat. # 80841) with supported lipid bilayers coated with cell ligands. Cells used were Jurkat cells.

Place culture media and fixative into a water bath

1.  Add cell culture media supplemented with 1 mM HEPES pH 7.4 to the well and warm in 37°C incubator for 15 min

2.  Count cells and resuspend in media at 6 x 10⁶ per ml

3.  Remove media from incubated wells leaving 100 μl and pipette in 50 μl of cell suspension containing 3 x 10⁵ cells 

Note: for supported lipid bilayers, never expose them to air and always leave ~100­­–150 μl of liquid in the well in all steps, even after fixation and during sample preparation. Failing to do so can result it high non-specific background staining during imaging.

4.  Incubate at 37°C for desired activation time (4 mins was used in this work)

5.  Fix cells with a final concentration of 4% paraformaldehyde (PFA) per well by adding 50 μl of 16 % PFA per well

6.  Incubate at 37°C for 15 mins

7.  Wash well x4 with 300 μl PBS followed by 2x with with 300 μl 5% BSA in PBS, leaving behind ~100 μl each time to ensure the supported lipid bilayer is not exposed to air. Leave to block surface for 1 hr.

8.  Permeabilize cells by adding 0.1 % saponin, 1% BSA in PBS and incubating at room temp for 15 min. Remove buffer leaving behind 100 μl. (Cells can be pre-stained with antibodies at this point)

9. Just prior to imaging on the microscope, add protein PAINT probe diluted to a concentration of 1 nM in PBS with 0.1 % saponin and 1% BSA.

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