Plasmid Extraction

Reagents and Materials
  • LB Broth supplemented with Ampicillin (100 µg/mL)
  • Glycerol stock or freshly transformed E. coli colonies
  • Qiagen Plasmid Mini Kit (buffers P1, P2, N3, PE, and spin columns)
  • 1.5 mL microcentrifuge tubes (EP tubes)
  • Nuclease-free water (pre-warmed to ~55 °C)
  • Centrifuge (capable of 13,000 rpm)
  • Shaking incubator (37 °C, 220 rpm)
  • Spectrophotometer (e.g., NanoDrop)
  • Pipettes and sterile tips
Procedure
Bacterial Culture
  1. Inoculate 45–50 mL Amp+ LB broth (100 µg/mL) with 1–2 µL glycerol stock or a single transformed colony.
  2. Incubate overnight (>12 h) at 37 °C, 220 rpm.
    Tip: Use three 50 mL Falcon tubes, each containing ~15 mL of medium, to improve aeration.
Cell Harvesting
  1. Pellet the cultures at 8250 rpm for 5 min at room temperature (RT).
  2. Discard the supernatant completely.
Resuspension
  1. Resuspend the pellet in 1–2 mL Buffer P1 (cold).
  2. Aliquot 250 µL of the resuspension into 1.5 mL microcentrifuge tubes.
    Note: Subsequent steps refer to individual aliquots.
Cell Lysis
  1. Add 250 µL Buffer P2, gently invert 6–10 times to mix (do not vortex).
  2. Incubate at RT for 2–3 min to ensure complete lysis.
Neutralization
  1. Add 350 µL Buffer N3, invert 6–10 times to mix.
  2. A white precipitate should form, indicating successful neutralization. If no precipitate appears, the lysis step may have failed.
Clarification
  1. Centrifuge at 13,000 rpm for 10 min at RT to pellet cell debris.
  2. Carefully transfer the clear supernatant to a fresh tube.
DNA Binding
  1. Load up to 800 µL of the supernatant onto a spin column.
  2. Centrifuge at 13,000 rpm for 1 min. Avoid transferring debris; load in 100 µL portions if needed.
Column Washing
  1. Discard flowthrough.
  2. Add 500 µL Buffer PE, centrifuge at 13,000 rpm for 1 min.
  3. Repeat once more for a total of two washes.
  4. Discard flowthrough and centrifuge again (dry spin) at 13,000 rpm for 1 min to remove residual ethanol.
Elution
  1. Air-dry the column for 1–2 min.
  2. Transfer the column to a clean 1.5 mL tube.
  3. Add 50 µL pre-warmed nuclease-free water, let stand for 1 min.
  4. Centrifuge at 13,000 rpm for 1 min to elute plasmid DNA.
    Important: The flowthrough contains your plasmid DNA—do not discard it.
Quantification & Storage
  1. Measure DNA concentration and purity using 1.5–2 µL on a spectrophotometer (e.g., NanoDrop). For multiple preparations of the same plasmid, pool high-purity samples for larger stocks.
  2. Label tubes clearly with plasmid name and concentration. Store plasmid DNA at −20 °C.

Cell Culture Handling

Reagents and Materials
  • Pre-warmed complete culture medium (DMEM/F12, 10% BSA, 1% PenStrep)
  • Sterile PBS (phosphate-buffered saline)
  • Trypsin-EDTA solution (room temperature)
  • Falcon tubes (5 or 15 mL)
  • Hemocytometer and coverslip
  • KimWipes and 70% ethanol
  • Micropipettes and sterile tips
  • Centrifuge capable of 500 × g
  • Inverted microscope
1. Media Change

When to Perform: Every 2–3 days, or if a high proportion of dead cells is observed.

  1. Aspirate spent medium carefully using a vacuum pump with a 1000 µL tip, tilting the dish slightly and avoiding contact with the bottom or sides.
  2. Add half the well volume of sterile PBS gently down the side of the dish. Swirl to wash away serum and debris.
  3. Aspirate PBS and repeat for a total of two washes.
  4. Add the full well volume of pre-warmed complete medium, pipetting gently to avoid mechanical stress.
  5. Inspect cells under a microscope to confirm they remain adherent and undamaged.
2. Cell Seeding and Passage

When to Perform: Once cells reach >80% confluence or if the medium is highly spent.

  1. Perform media aspiration and two PBS washes as described in the Media Change section.
  2. Add 1/4 well volume of Trypsin-EDTA dropwise and swirl gently to coat evenly.
  3. Incubate at 37 °C for 1–2 min until cells detach (refractile particles visible under microscope).
  4. Quench Trypsin by adding half the well volume of complete medium.
  5. Transfer the cell suspension to a sterile Falcon tube. Rinse the dish with ~1 mL medium to collect remaining cells and combine in the same tube.
  6. Centrifuge at 500 × g for 5–10 min to pellet cells.
  7. Aspirate supernatant carefully using progressively smaller tips (1000 µL then 100 µL) to avoid disturbing the pellet.
  8. Resuspend pellet in 1–2 mL fresh complete medium depending on pellet size.
  9. Calculate volume needed for seeding; add fresh medium to new dishes first, then add cell suspension dropwise. Swirl gently in a figure-eight motion to ensure even distribution.
  10. Confirm even adhesion under a microscope; check again after ~12 hours.
3. Cell Counting

When to Perform: To determine cell concentration or seed at precise density.

  1. Clean hemocytometer and coverslip with ethanol and KimWipe.
  2. Place coverslip over counting chamber.
  3. Dilute 10 µL cell suspension into 1 mL medium (adjust if necessary).
  4. Load diluted suspension under the coverslip using the groove along the edge.
  5. Count cells in the four large corner squares under a microscope.
  6. Calculate concentration: Cells/mL = Average count × 104 × dilution factor.
  7. Repeat with a lower dilution if counts are too low for accuracy.

Western Blot

Reagents and Materials
  • Protein samples (quantified by BCA assay)
  • 4× SDS loading dye
  • Nuclease-free water (ddH₂O)
  • 4–12% gradient SDS-PAGE gel (15-well)
  • 1× MOPS running buffer
  • Protein ladder/marker
  • PVDF or nitrocellulose (NC) membrane
  • eBlot Transfer System (or equivalent)
  • 1× TBST (Tris-buffered saline + 0.1% Tween-20)
  • 5% nonfat milk & 5% BSA (in 1× TBST)
  • Primary antibodies: Anti-Rabbit GAPDH (1:8000), Anti-Mouse FLAG-Tag (1:5000)
  • Secondary antibodies: Goat Anti-Rabbit HRP (1:2000), Goat Anti-Mouse HRP (1:2000)
  • Chemiluminescent substrate
  • UV or chemiluminescence imaging system
Procedure
  1. Sample Preparation: Equalize protein concentrations. Prepare 20 µL total per sample (5 µL 4× SDS dye + protein + ddH₂O). Heat-denature at 95 °C for 10 min.
  2. Gel Electrophoresis: Assemble gel in tank with 1× MOPS buffer. Load 15 µL of each sample and ladder. Run at 120 V for ~60 min.
  3. Membrane Transfer: Assemble transfer stack and run on standard eBlot protocol (~15 min).
  4. Blocking: Incubate membrane in 5% milk in 1× TBST for 1 h at room temperature (RT).
  5. Primary Antibody Incubation: Incubate membranes overnight at 4 °C in 5% milk with primary antibodies (GAPDH: 1:8000, FLAG-Tag: 1:5000).
  6. Washing: Wash membrane 3× for 5 min each with 1× TBST.
  7. Secondary Antibody Incubation: Incubate for 1 h at RT in 5% BSA with HRP-conjugated secondary antibodies (1:2000).
  8. Detection: Apply chemiluminescent substrate and visualize bands using an imaging system.

LNP Synthesis

Reagents and Materials
  • Cholesterol, stock solution 5 mM in chloroform
  • DOTAP, stock solution 5 mM in chloroform
  • Phosphate-buffered saline (PBS), ice-cold
  • Round-bottom flask, Rotary evaporator, Sonicator (probe and bath)
  • Syringe and 0.22 µm polyethersulfone (PE) filter
Procedure
  1. Stock Preparation: Prepare 5 mM solutions of Cholesterol and DOTAP in chloroform. Store at −20 °C.
  2. Lipid Mixing: In a round-bottom flask, combine 0.75 mL Cholesterol and 0.25 mL DOTAP. Add 1 mL chloroform and vortex.
  3. Solvent Evaporation: Remove chloroform using a rotary evaporator for 10–15 min until a white lipid film forms.
  4. Lipid Rehydration: Add 2 mL ice-cold PBS. Sonicate in a bath sonicator at 45 Hz for 60 sec.
  5. Film Disruption: Perform alternating cycles of vortexing (60 sec) and probe sonicating (30 Hz for 30 sec) on ice until the film is rehydrated.
  6. Filtration: Pass the lipid suspension through a 0.22 µm PE filter 2–3 times.
  7. Storage: Store the LNP solution at 4 °C for up to 4 weeks.