Category Archives: Protocol

Transfection of Mouse N2a Cells using Lipofectamine/plus

1.  Quick thaw the cells in 37 C water bath and add to 10 ml of growth medium. Spin 5 min at 800 rpm to remove DMSO from freezing medium.  DMSO is toxic to growing cells.  Resuspend in 10 ml growth medium in 75 cm2 TC flask and place in 5% CO2/37 C incubator.

2. Plate approximately 5 x 105 cells into 60mm petri dishes. Wait until the cells reach 80% confluent.

3.  Dilute 2-4 ug DNA of each construct to be transfected into 250 ul of OptiMEM or DMEM medium. Add 8 ul of Plus Reagent (Invitrogen # 11514-015) to the DNA/MEM mix.  Let sit 15 min at room temperature.

4. Separately, mix 12 ul of Lipofectamine (Invitrogen # 18324-111) with 250 ul of OptiMEM or DMEM.

5. Mix DNA/MEM/Plus solution with Lipofectamine/MEM.  Let sit 15 min at room temperature.

6. Remove growth medium from 60mm cell cultures.  Rinse once with 2 ml OptiMEM or DMEM, replace with another 2 ml MEM to remove serum from the plate.

7. Add preincubated DNA/Plus/Lipofectamine/MEM solution to the cells.  Mix gently.

8. Grow cells 3 hr – overnight.  Add 2.5 ml N2a medium and continue culture.

If developing stable transfectants, add G418 (Gibco/Life Tech # 1181-031) to the growth medium  to select for positive cells.  Use at 400-800 ug/ml for selection of clones, and 200-400 ug/ml for maintenance.

9. 48 hours post transfection, harvest the cells for the future assay.

Growth medium:

DMEM, high glucose, w/L-gluatmine       Gibco # 11965-092     250ml

Fetal bovine serum, qualified           Gibco # 26140-079      50ml

OptiMEM 1 medium                    Gibco # 31985-070          250ml

Penicillin-Streptomycin            Gibco # 15140-122             5ml

Filter to 0.2 um.  Stable for several months at 4 C.


1. Solutions and reagents

1.1 Xylene

1.2. Ethanol, anhydrous denatured, histological grade (100%, 95%, 70%, 50%)

1.3. Washing buffer/TBST: 1X TBS/0.1% Tween-20, pH to 7.6.

1.4. Distilled water (dH 2O)

1.5. Antigen Retrieval Solution: 0.01M Sodium Citrate Buffer, pH 6.0

To prepare Antigen Retrieval stock solutions:

- 10X Stock: Dissolve 29.4 g sodium citrate trisodium salt dihydrate (C 6H 5Na 3O 7 2H 2O)
in 1 liter of dH2O. Add 5mL Tween-20.

- 1X Working Solution: Mix 200mL 10X stock with 1800mL dH2O; pH to 6.0

1.6. 3% Hydrogen Peroxide

1.7. Blocking Buffer: 10% serum in PBS (serum origin depends on the host of the secondary antibody)

1.8. Primary Antibody Diluent: 5% serum in PBS (serum origin depends on the host of the secondary antibody)

1.9. Hematoxylin

1.10. Permanent Mounting Medium

2. IHC Protocol

2.1. Deparaffinization/Rehydration

2.1.1. Heat slides in an oven at 65 °C for 1 hour.

2.1.2. De-paraffinize/hydrate using the following series of washes: two Xylene washes (3 min each), followed by two 100% ethanol rinses (3 min each), followed by 95% ethanol, 70% ethanol, 50% ethanol, 30% ethanol, followed by TBST wash for 3 min on a shaker.

2.2. Antigen Retrieval

This is recommended Heat Induced Epitope Retrieval (HIER) using Decloaking Chamber/Pressure Cooker. Hot water bath or Microwave with temperature sensor can be also used (protocol would vary depending on the method used).

2.2.1. Add 500 ml of dH 2O to Decloaker/Pressure Cooker.

2.2.2. Immerse slides into staining dish containing Antigen Retrieval Solution. Place staining dish into decloaking chamber.

2.2.3. Program to run for 30 seconds at 125° C, followed by 10 seconds at 90° C.

2.2.4. Let it cool down to room temperature (10 – 20 minutes).

2.2.5. Removes slides and rinse in TBST.

2.2.6. Proceed to Staining step.
2.3. Staining

2.3.1. Wash slides with TBST for 3 min on a shaker.

2.3.2. Inactivate endogenous peroxidase by covering tissue with 3% hydrogen peroxide for 5 min.

2.3.3. Wash slides three times with TBST (3 min each on a shaker).

2.3.4. Block slides with the blocking solution for 1 hour.

2.3.5. Dilute primary antibody in primary antibody diluent per recommendation on data sheet.

2.3.6. Apply primary antibody to each section and incubate overnight in the humidified
chamber (4 °C).

2.3.7. Wash slides three times with TBST (3 min each on a shaker).

2.3.8. Apply to each section secondary HRP-conjugated anti-rabbit antibody diluted in the blocking solution per manufacturer’s recommendation; incubate for 30 min at room temperature.

2.3.9. Wash slides three times with TBST (5 min each on a shaker).

2.3.10. Add freshly prepared DAB substrate to the sections and incubate until stain develops (generally 1 min).

2.3.11. Rinse sections with water.

2.3.12. Counterstain with Hematoxylin (generally 10 seconds).

2.3.13. Rinse sections with water.

2.3.14. Dehydrate samples using two washes with 100% Ethanol (3 min each), followed by two rinses with Xylene (3 min each).

2.3.15. Mount coverslips on slides using permanent mounting medium


kill curve and stable transfection using puromycin

1.  Do a kill-curve to find out the lowest dose to kill your cells (4-7 days for puromycin). You can do that any time. You just need to know what concentration to use to select for your stable transfected clones.

Split your cells at 1:10 the day before the selection. Change the medium with different concentration of puromycine every 2 to 3 days. The normal conc. used for Puromycin is from 0.1 ug/ml to 10 ug/ml. So you can pick up a series of conc. within this range. For example, you can use 6 well plates, with each well containing a different conc. of puromycin (0, 0.1, 0.5, 1, 2, 10).  You can pick more point if you want, so that you can get more accurate data. Check your cells every day, after 1 week, find the lowest dose that kill all your cells (100%). That’s the concentration you want to use for selection.

2. Perform a transfection with the plasmid containing Puro resistant gene.  Twenty-four hrs post-transfection, passage the cells at 1:10 into fresh growth medium containing puromycin at the concentration you found from step 1.  A mock transfection should be performed in parallel as a control. Grow and passage the cells as necessary (usually 2-3 days), maintaining selection pressure by keeping puromycin in the growth medium. After 1-2 weeks, a large number of the cells will be killed; the cells that remain growing in the selective medium have retained the expression plasmid, which stably integrates into the genome of the targeted cells. Monitor the mock control to ensure the cells are dying, no cells attached at the bottom.

3. If you pool all the colonies, that is your stable pool. If you isolate individual colony, that is your individual clones. For isolation of individual colony, there are several commercial tools to do that. You can also keep on diluting your stable transfected cells to the point that you can assume every well only get one colony. Continue growing these cells in selection medium for 1-2 additional passages.  At this time,  each well contains a clonal population of stably transfected cells, which can be maintained in normal growth medium without the selection pressure of puromycin (although you may wish to grow the cells under “light pressure”, lower concentration of puromycin).  These populations can be used for      experiments or stored under liquid nitrogen in growth medium with 10% DMSO and 20% FBS for future use.

Western Blot Protocols

Western blot analysis
  1. Run SDS-PAGE gel, and then Western transfer the protein samples to nitrocellulose (NC) membrane for immunoblot analysis.
  2. After transfer, transfer the membrane to western-blot tray, briefly wash the NC membrane with distilled water.
  3. (Optional) Visualize the proteins on the membrane by Ponceau’s staining.
  4. Wash off the red stain with distilled water.
  5. Block the membrane with 5-10ml blocking buffer (made by 5% non-fat milk in 1xPBST) for 30 minutes at R/T.
  6. Dilute the primary antibody with blocking buffer according to the suggested dilution factor on datasheet (In case of anti-DDK mouse monoclonal antibody (TA100011, do 1:4000 dilution).
  7. Remove the blocking buffer and add enough diluted primary antibody to cover the membrane.
  8. Incubate the membrane with primary antibody for 1hr at R/T. (Note: Or you can do overnight incubation at 4C, make sure you cover the western-blot tray to prevent excessive evaporation). To prevent uneven coverage, the western-blot tray can be rocked on a rocker platform.
  9. Collect the primary antibody and store them at 4C for up to two weeks. (If you would like to store them longer, you can freeze the diluted antibody at –20C. Remember frequent freezing and thawing will gradually decrease the antibody titer.)
  10. Briefly wash the membrane with 1xPBST once to remove any excessive primary antibody.
  11. Add enough 1xPBST to cover the membrane and leave the Western-blotting tray on a rocker platform.
  12. Wash the membrane for 15 minutes. (Note: If the background is high, repeat this step for two to three times.), turn on the developer during the wash time.
  13. Dilute HRP-conjugated secondary antibody with blocking buffer (1:5000 or higher dilution is usually good for Goat anti-mouse-HRP; TA100015).
  14. Incubate the membrane with secondary antibody for 30 minutes to 1hr.
  15. Wash the membrane with 1xPBST for 15 minutes, and then 3 times (5 min/time).
  16. Prepare the chemiluminescence development substrate mixture by mixing equal amount of solution 1 and 2 (TA100016; Normally 1ml will be enough for one membrane).
  17. Prepare a plastic saran film, lay the film on a flat surface, and dispense 1ml of substrate mixture for one membrane on the plastic saran film.
  18. Use a forceps to take washed the blot from the western-blotting tray, flip it, lay on the substrate mixture, and then incubate for 1 to 5 minutes. (Note: To avoid air bubbles, always lay the blot by touching one edge first.)
  19. Remove excess Chemiluminescence Reagent and wrap the membrane in plastic. Place inside X-ray cassette.
  20. Expose to film and develop
Buffer preparation
1xPBS: This buffer is made by dissolving 8g of NaCl, 0.2g of KCl, 1.44g of Na2HPO4 and 0.24g of KH2PO4 into 800ml of distilled water. Then adjust the pH to 7.4 with HCl, and add H2O to 1 liter.
1xPBST: 0.05% Tween 20 in 1xPBS

Flow Cytometry for Intracellular Staining

  • Solutions and Reagents
    1. 1X Phosphate Buffered Saline (PBS): Dissolve 8g NaCl, 0.2g KCl, 1.15g Na2HPO4 and 0.2g KH2PO4 in 800mL distilled water (dH2O). Adjust the pH to 7.4 with HCl and the volume to 1 liter. Store at room temperature.
    2. Fixation buffer: 2% paraformaldehyde in 1xPBS
    3. Permeabilization buffer : 0.1% Triton X-100 in 1xPBS
    4. FACS buffer: 0.5% BSA , 0.05% Azide in 1xPBS
    5. Fluorescent dye conjugated secondary antibody.
  • Fixation
    1. Collect cells by centrifugation and aspirate supernatant.
    2. Fix the cell by 125μl cold fixation buffer, vortex briefly.
    3. Incubate at room temperature for at least 30 min or for 1hr 40C.
    4. Centrifuge for 5min at 300g,remove the supernatant.
  • Permeabilization
    1. Add 1ml permeabilization buffer to each tube.
    2. Centrifuge briefly, and aspirate supernatant.
    3. Resuspend cells in 125μl of permeabilization buffer and incubate at room temperature for 5min.
  • Staining
    1. Aliquot 1-2×106 cells into each tube.
    2. Add 1 ml FACS buffer to each tube, centrifuge to pellet the cells.
    3. Resuspend cell pellet with 125μl FACS buffer containing diluted primary antibody, vortex and incubate on ice for 30min.
    4. Rinse as before in FACS buffer by centrifugation.
    5. Resuspend cells in fluorescent dye conjugated secondary antibody, diluted in FACS buffer per manufacturer’s recommendations.
    6. Incubate for 30 minutes on ice.
    7. Rinse the cells as before in FACS Buffer by centrifugation.
    8. Resuspend cells in 0.5 ml PBS and analyze on flow cytometer

Immunoprecipitation (IP)

  1. Solutions and ReagentsLysis buffer:
    1. Typically use RIPA buffer (25 mM Tris-HCl pH 7.6, 150mM NaCl, 1% NP-40, 1% sodium deoxycholate, 0.1% SDS). Proteinase inhibitor cocktail should be added fresh before each use.
    2. The usage of SDS depends on the nature of the cells. For some cell lines, 0.1% SDS will release DNA and thus make it hard to extract proteins out. In those cases, omit SDS
  2. Preparation of cell lysates
    1. Add ice cold lysis buffer (1ml per 100mm-dish or 107 cells, or adjust based on your specific requirements). Scrape off cells (for adherent cells still on plate) and resuspend cells. Collect cells in a centrifuge tube and agitate for 30 min at 4°C.
    2. Spin cells at 4°C for 20 min at 12000 rpm.
    3. Save the supernatant which is the cell lysates.
  3. Pre-clearing
    1. Add normal serum or irrelevant antibody from the same species and isotypes as the IP antibody you will use. The amount should be at least 5-fold more than the amount you will use for IP. Incubate for 1 hr at 4°C.
    2. For 1 ml lysate, add 100 ul of proteins A or protein G beads slurry (50 ul solid bed volume), and incubate at 4°C for 30 min on a rotator.
    3. Spin down beads at 14000g for 5 min at 4°C.
    4. Save the supernatant which is the pre-cleared lysates.
  4. Immunoprecipitation (IP)
    1. Add IP antibody to the pre-cleared lysates. You will need to determine the best amount of antibody to use. As a starting point, you may use 1 ug antibody for every ml of lysates.
    2. Incubate for a certain amount of time (from 1 hr to overnight, depending on your specific conditions) at 4°C.
    3. Add 100 ul of protein A or protein G slurry (50 ul solid bed volume) to 1 ml lysate and incubate for 3 hr at 4°C on a rotator.
    4. Spin down beads, and remove supernatant.
    5. Wash beads 3 times with lysis buffer.
    6. Add SDS-PAGE sample buffer to beads. Boil and run gel.