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Western blot analysis protocol

Sample Preparation

1. Heat samples in Laemmli buffer at 70°C - 90°C for 10 min.

2. Load 20 - 30 μg/lane of membranes/lysates sample or 2-5 x 105 cell lysate/lane.

3. Perform SDS-PAGE according to standard protocols.

Transfer

1. Perform transfer according to the manufacturer’s instructions.

Note: High molecular weight proteins may need longer transfer times.

Western blot

1. Block membranes with blocking solution (3% BSA in PBS + 0.1% Tween-20 and 0.05% NaN3) for 2-5 hr at room temperature with gentle agitation.

2. Add primary antibody diluted in blocking solution at the appropriate dilution. Incubate for 2-3 hr at room temperature or overnight at 4°C with gentle agitation. (If control antigen is used, refer to the protocol describing the use of control antigen).

3. Discard primary antibody and wash membranes by incubating with washing buffer (PBS + 0.1% Tween-20) 3 times for 15 min at room temperature.

Note: Do not use solutions containing NaN3 from this point if you are using a secondary antibody conjugated to horseradish peroxidase (HRP).

4. Incubate secondary antibody at appropriate dilution in washing buffer for 1 hr at room temperature with gentle agitation.

5. Wash membranes with washing buffer 3 times for 15 min at room temperature.

6. Proceed to detection using an ECL system. If using a commercial kit, perform according to the manufacturer’s instructions.


Immunohistochemistry protocol

Reagent Preparation

1. Neutral-buffered Formalin, 10% (NBF), 1 liter

Double -distilled H2O 900 ml

Di-sodium hydrogen phosphate, anhydrous (Na2HPO4) 6.5 grams

Sodium di-hydrogen phosphate, monohydrate (NaH2PO4 . H2O) 4.0 grams

Formaldehyde, 37% solution 100 ml

The pH should be about 7.0. Adjust if necessary with 1N NaOH or 1M HCl. Store at 4°C.

Final concentrations:

Formaldehyde 3.7%

Na2HPO4 46 mM

NaH2PO4. H2O 29 mM

2. Sodium Citrate Buffer pH 6.0, 1 liter

Tri-sodium citrate (di-hydrate) 2.9 g

Double distilled water 1000 ml

Mix to dissolve sodium citrate and adjust pH to 6.0 with 1M HCl.

Add 0.5 ml Tween 20. Store at room temperature or at 4°C if storing for longer than 3 months.

Final concentrations:

Sodium citrate 10 mM

Tween 20 0.05%

3. TBST (Tris-Buffered Saline, 0.05% Tween20), 1 liter

10X TBS 100 ml

Double-distilled water 900 ml

Tween 20 0.5 ml

4. Blocking buffer

5% serum, milk or BSA (bovine serum albumin) in TBST

Protocol

1. Tissue Preparation:

Fix tissue in 10% neutral buffered formalin for at least 24 hours. Embed in paraffin wax according to embedding machine manufactures instructions. Formalin has a very slow diffusion coefficient so the tissue needs to be no more than 1 cm thick.

2. Tissue Sectioning:

Prepare 4 - 12 µm sections on the microtome and place on clean, positively - charged microscope slides. Heat in tissue-drying oven for 45 minutes at 60°C.

3. Deparaffinization:

Wash slides 3 times for 5 minutes in xylene.

4. Rehydration:

    1) Wash slides 3 times for 3 minutes in 100% alcohol.

    2) Wash slides 2 times for 3 minutes in 95% alcohol.

    3) Wash slides 2 times for 3 minutes in 80% alcohol.

    4) Rinse slides for 5 minutes in running distilled water.

5. Antigen retrieval:

There are several methods of antigen retrieval. The most common is heat-mediated retrieval in citrate buffer.

    1) Heat slides in 10 mM sodium citrate buffer, pH 6.0 at 95 -100°C for 20 minutes.

    2) Remove from heat and let stand at room temperature in buffer for 20 minutes.

    3) Rinse in TBST 1 minute.

There are other antigen retrieval procedures available and the type of retrieval and the incubation time for antigen retrieval may require some optimization.

6. Immunostaining:

(Do not allow tissues to dry at any time during the staining procedure).

    1) Add 100 µl per slide of blocking solution. Incubate 20 to 30 minutes at room temperature.

    2) Drain the blocking solution from slides. Apply 100 µl per slide of diluted primary antibody at recommended concentration. Incubate 45 minutes at room temperature or overnight at 4°C.

    3) Wash slides in 1X TBST 4 times for 5 minutes

    4) Apply a 100 µl per slide of diluted conjugated secondary antibody. Incubate for 30 minutes at room temperature.

    5) Wash slides in 1X TBST 4 times for 5 minutes.

    6) Apply color development (i.e. enzyme substrate) 30 minutes, or follow manufacturer’s instructions.

    7) Wash slides in 1X TBST 4 times for 5 minutes

    8) Wash slides in distilled water for 1 minute.

7. Dehydrate and mount slides:

This method should only be used if the chromogen substrate is alcohol insoluble.

    1) Wash slides in 2 changes of 80% alcohol, 1 minute each.

    2) Wash slides in 2 changes of 95% alcohol, 1 minute each.

    3) Wash slides in 3 changes of 100% alcohol, 1 minute each.

    4) Wash slides in 3 changes of xylene 1 minute each.

    5) Apply coverslip


Immunocytochemistry protocol for live cells

Cell preparation/labeling:

1. Plate cells in chosen chamber slides and grow 1-2 days in appropriate medium. Cells need to attach strongly to plate.

Important: Some cell lines will need special coating i.e. poly-lysine of the chamber slides to aid in cell attachment. The specific type of coating needs to be determined empirically as it varies between chamber types and/or cell lines.

2. Wash cells 2-3 times with ice-cold assay buffer (PBS + 2% BSA + 0.05% NaN3).

3. Add primary antibody at the appropriate dilution in ice-cold assay buffer. Incubate 1 hr at 4°C.

NOTE: If using fluorescent labeled antibody, skip to step 6.

4. Wash cells 2-3 times with ice-cold assay buffer.

5. Add fluorescently-labeled secondary antibody at the appropriate dilution in ice-cold assay buffer and incubate 1 hr at 4°C protected from light.

6. Wash 3-5 times with assay buffer and drain well. Add buffer to cover cells and proceed to the microscope.


Immunocytochemistry protocol for fixed cells

Cell preparation/labeling:

1. Plate cells in chosen chamber slides and grow cells 1-2 days in appropriate medium. Cells need to attach strongly to plate.

Important: some cell lines will need special coating i.e. poly-lysine of the chamber slides to aid in cell attachment. The specific type of coating needs to be determined empirically as it varies between chamber types and/or cell lines.

2. Wash cells 2-3 times with ice-cold PBS.

3. Fix cells by adding paraformaldehyde (PFA) 1- 4% in PBS 10 min at room temperature.

Note: the optimal PFA dilution depends on the cell type and should be established experimentally.

4. Wash cells 2-3 times with ice-cold PBS.

5. Permeabilize cell membranes by adding saponin assay buffer (PBS + 2% BSA + 0.1% saponin) 10 min at room temperature.

6. Block unspecific binding sites with 5% goat serum in saponin assay buffer for 15 min at room temperature.

7. Add primary antibody at the appropriate dilution in saponin assay buffer and incubate for 1-2 hr at 4°C.

NOTE: If using fluorescent labeled antibody, skip to step 10.

8. Wash cells 3 times with saponin assay buffer.

9. Add fluorescently-labeled secondary antibody at the appropriate dilution in saponin assay buffer and incubate 1 hr at 4°C protected from light.

10. Wash 3-5 times with saponin assay buffer and drain well. Add buffer to cover cells and proceed to the microscope.


Flow cytometry protocol

Cell preparation/labeling

1. Transfer 0.5-2 x 106 cells into a microtube. Centrifuge 5 min. at 300 x g. Discard supernatant.

2. Carefully resuspend the cell pellet with 20-100 μl ice-cold labeling buffer (PBS + 2% BSA + 0.05% NaN3).

3. Add primary antibody at the appropriate dilution in labeling buffer. Incubate on ice 30-60 min.

4. Wash the unbound antibody by filling the microtube with labeling buffer, centrifuge 5 min. at 300 x g and discard supernatant. Repeat twice.

5. Add fluorescently-labeled secondary antibody at the appropriate dilution in ice-cold labeling buffer. Incubate on ice protected from light 30-60 min.

6. Wash the unbound antibody by filling the microtube with labeling buffer, centrifuge 5 min. at 300 x g and discard supernatant. Repeat twice.

7. Resuspend the cells with ice-cold labeling buffer. Keep on ice protected from light until analyzed with a flow cytometer.

Note: If working with a fluorescently-labeled primary antibody skip steps 5 and 6 and proceed directly to step 7.

Antigen manipulation

Incubate the antibody, in parallel, with and without the antigen (the ratio antibody/antigen is available in the certificate of analysis delivered with the antibody) in a small volume (500 µl of 1% BSA in PBS or 3% skim milk) for 1 hr at room temperature with rotation. After the incubation time, dilute each vial to the desired working dilution with the buffer you use and apply it to the membranes for parallel experiments.

Compare the results of antibody alone versus antibody with antigen. The disappearance of the requested band will confirm the specificity of the antibody.


Tissue membrane preparation protocol

Membrane lysate preparation:

1. Remove tissue of interest from animal and flash freeze in liquid nitrogen. Store tissue at -80°C until further use.

2. Suspend frozen tissue in 5 volumes of ice-cold lysis buffer [4 mM HEPES pH 7, 320 mM Sucrose, 5 mM EDTA pH8, protease inhibitor cocktail (Complete, EDTA-free, Roche)].

3. Homogenize the tissue with a polytron homogenizer. Centrifuge homogenates10 min. at 2000 x g, 4°C. Discard large debris.

4. Transfer the supernatant to a clean tube and resuspend the pellet in 2 volumes lysis buffer and re-homogenize.

5. Centrifuge the homogenate10 min. at 2000 x g, 4°C. Combine supernatant with that of step 4.

6. Centrifuge the supernatant (from steps 4 and 5) 1 hr at 100,000 x g, 4°C.

7. Discard the supernatant. Resuspend the pellet (which contains tissue membranes) in 2 volumes lysis buffer and briefly homogenize with a polytron homogenizer.

8. Measure protein concentration using the Bradford method. Adjust protein concentration to 3 mg/ml with lysis buffer. Store protein samples at - 80°C until further use.


Tissue lysate preparation protocol

Lysate preparation:

1. Remove tissue/organ of interest from the animal and flash freeze in liquid nitrogen. Store the tissue/organ at -80°C until further use.

2. Suspend the frozen tissue in 5 volumes ice-cold lysis buffer [50 mM Tris pH 7.4, 5 mM EDTA pH 8, 1% Triton X-100 and protease inhibitor cocktail (Complete, EDTA-free, Roche)].

3. Homogenize the tissue with a polytron homogenizer. Centrifuge homogenates1 hr at 100,000 x g, 4°C.

4. Transfer the supernatant to a clean tube and measure protein concentration using the Bradford method. Adjust protein concentration to 3 mg/ml with lysis buffer. Store lysates at - 80°C until further use.


siRNA transfection protocol

Using the following procedure to transfect siRNA into mammalian cells in a 12-well dish. Adjust cell and reagent amounts proportionately for wells or dishes of different sizes. Optimization may be necessary.

1) Split the cells one day before transfection to reach 70 - 80 % confluence in the day of transfection.

2) For each transfection sample, prepare the transfection complexes as follows:

In a sterile 1.5 ml microtube, mix the following reagents (for 12-well dish):

50 µl        Serum-free DMEM (or any other serum free-medium, PBS, DPBS etc.)

40 pmol   siRNA (2 µl x 20 µM siRNA)

3) Pipetting up and down the mixture a few times. Spin briefly in a centrifuge. Leave the mixture at room temperature for 5 min.

4) Add the entire mixture directly to cells in the 12-well culture dish. Tilting the dish a few times to mix. Return the dish to a CO2 incubator. Serum concentration (0-20%) in the growth medium has no effect on the transfection efficiency.

5) Optional: At anytime between 5 to 24 hours after the transfection, replace the medium in the dish with 1 ml (or any volume you’d like) fresh growth medium. If toxicity is not observed, change of medium is not needed.

6) Gene knockdown should be assayed 24-72 hours post-transfection.

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