Lipisome and Lipid-Coated Beads

General Notes

• All buffers should be filtered through a 0.2um filter to remove any particulates before use
• Avoid plastics until beads/lipisomes are generated, as lipids will stick to these
• Clean all equipment toughly with chloroform or 100% EtOH to avoid cross-contamination with lipids

Protocol

Updated Protocol for PtdSer Beads (As per Austin Lam and Brandon Dickson)

1. Using a glass syringe, add 30 μL PtdChol (10 mg/mL stock), 8 μL PtdSer (10 mg/mL stock), and 8 μL of biotinylated PtdEth (10 mg/mL stock) to a small amber vial.
   • Ensure that you flush the syringe completely with chloroform with chloroform every time before switching vials. Multiple flushes will be required.
   • If using rhodamine-PtdEth, use this last to avoid contaminating the other stocks with red lipid.
   • Flush vials containing the stock solutions with nitrogen gas, then cap and seal with parafilm before returning to freezer.
2. Evaporate the lipid mixture in the amber vial with a light stream of nitrogen gas, rotating the vial with your hands.
   • Continue for a few minutes until the vial is completely dry.
3. Add 400 μL of PBS into the vial. Vortex vial at max intensity for 1 minute to suspend completely.
4. Vortex the small container containing the silica beads. Transfer 100 μL of silica beads into a 1.5 mL tube. Wash with 1 mL ddH₂O at 5000 x g for 1 minute. Carefully remove supernatant. Repeat for a total of three washes.
   • Remove as much water as possible after the last wash.
5. Resuspend the beads in 400 μL of the lipid solution. Mix by pipetting.
6. Rotate the bead-lipid mix at room temperature for 20 minutes using the rotisserie-style spinner.
   • Cover tube with tin foil if using rhodamine-PtdEth to avoid photobleaching
7. Spin the beads down at 5000 x g for 30 seconds. Remove supernatant carefully.
   • This step removes unincorporated liposomes
8. Resuspend beads in 400 μL of PBS. Briefly purge the tube with nitrogen gas before capping. Store at 4°C for up to 4 days.

Lipid Preparation:

1. Using the lipid calculation spreadsheet, calculate the amount of lipids required.
   • If using dansylated lipids, add at 2-4% total molar amount. Higher concentrations will self-quench.
     
   • Generally speaking, 4μmol of lipids will produce enough for 1 weeks work
     
   • Once made, lipids are good for 3-4 days at 4oC
     

• In a hood:

2. Clean glass syringes 5x with choloform.
3. Using the syringes, transfer the desired amount of lipids from the stock vials into small glass tubes. Do not use plastic for any step.
   
4. Between each lipid, clean syringe by washing 5x with choloroform.
   
5. Before sealing each stock tube, fill tube with N2 or CO2 to prevent lipid oxidation
   
6. Dry the lipids to the lower ¼ of the tube by placing the tube at an ~45o angle, and rotating the tube while gently flowing N2 or CO2 into the tube.
   
7. Once the tubes are dried, place tubes beneath tube drier, and dry for an additional 2hrs-over night using N2 of CO2. Protect tubes from light during this time.
   
8. During step 7, clean syringes 5-10x in chloroform and return to storage area.
   

Lipisome Preparation:

1. To the dried lipids add 400μl of desired buffer (i.e. 20mM HEPES, pH 7.2)
   
2. Vortex each tube until as much white precipitate goes into solution as possible. Some lipids will not resuspend well (i.e. PC, PE)
   
3. Sonicate tubes 2-3x, 30sec, in ice-cold water, using an immersion sonicator at 30-35% maximum power.
   
4. Clean syringes for lipisome extruder in 100% EtOH and assemble.
   
5. Remove o-rings and mesh from extruders. Using syringes, pass 100% EtOH through each half of the extruder 5x, then the desired buffer 1x.
   
6. Wash o-rings in ddH2O and place back into extruder. Using a vacuum, remove any excess fluid from o-rings and ports.
   
7. Place 2 100nm-pore filters onto one o-ring. Filters should be centered on extruder, with an equal amount extending over o-rings on each side.
   
8. Insert into metal casing, followed by second half of extruder. Screw caps on tightly; while tightening watch filters through observation port to ensure they are sealed properly.
   
9. Place a long needle (i.e. 1.5” 18g) onto one glass syringe. Suck up lipid into syringe, making sure no air bubbles are trapped. If necessary lipids can be transferred into plastic eppindorfs if glass tubes are too long for needles.
   
10. Insert sample-containing syringe onto the short port of the extruder. Attach the empty syringe to the long-port. Press solution through extruder 30 times; if the solution remains turbid at this point pass through additional times until solution becomes clear. Collect purified lipisomes into eppindorf tubes, using the syringe on the long port to ensure maximum recovery and prevent contamination with unprocessed sample.
    
11. Disassemble extruder, throw away 100nm filters, and clean the extruder as described in steps 4-6 between each sample.
    
12. When complete clean extruder with EtOH as described above. When finished, reassemble extruder, with o-rings, and pass 100% EtOH through system 5 times. Disassemble completely and place in carrying case.
    
13. Liposomes can be stored at 4oC for 3-4 days.
    

Lipid-Coated Bead Preparation:

1. Dissassemble HPLC column and collect nucleosil beads into a lipid-free container. Store at -20oC until needed. For each lipid sample measure out 2mg beads. Dilute into 100μl chloroform per 2mg beads and sonicate.
2. Before step 6 of the lipid preparation (first drying of the aliquotted lipids) add 100μl of the bead/chloroform mixture. Once added, dry the lipid/bead preparation protocol as per usual.
3. Resuspend dried lipids in 1.5ml of desired buffer. Vortex and sonicate as described in lipisome section to suspend.
4. Transfer to 1.5ml eppidorf tubes by inverting glass tube overtop of eppidorf. Vortex in minifuge, 4000rpm, 4min.
5. Remove supernatant, resuspend in 1ml buffer, and spin at 6000 RPM for 4 min. Repeat 2-3x
6. Put an 8μl spot on a slide and cover with cover slip. View under microscope; if beads are clumped further processing is required; otherwise go to step 7.
7. If sample is still clumped, spin 5min at 6000 RPM
   
   • Remove supernatant
     
   • Gently profuse buffer across pellet to wash
     
   • Remove wash and resuspend
     
   • Sonciate in immersion sonicator, 30sec at 30% maximum power
     
   • Repeat until clumps are gone
8. Spin down cleaned sample at 4500RPM, 4min, resuspend in 1ml clean buffer, and count beads on scope using a hematocytometer and 100x magnification. Normalize bead concentration and store beads at 4oC until needed. Beads will last 3-4 days at 4oC.