Protocols

Immunohistochemical Staining and Analysis

Prepare frozen tissue sections

1. Place a freshly dissected tissue block (<5mm thickness) onto a pre-labeled tissue base mold.

2. Cover the entire tissue block with cryo-embedding media (e.g OCT)

3. Place the base molds containing the tissue block into liquid nitrogen and freeze completely.

4. Store the frozen tissue block at -80 οC until ready for sectioning.

5. Transfer the frozen tissue block to a cryotome cryostat at -20 οC and allow the temperature to equilibrate to the temperature of the cryotome.

6. Section frozen tissue block into a desired thickness (typically 5 μm) using the cryotome and place the tissue sections onto superfrost glass slides.

7. Dry the sections at room temperature overnight or store in -80οC for later use.

Immunofluorescence staining of frozen tissue sections

8. Fix the tissue sections in 300 ml of 4% PFA taken in coplin jar for 5 min.

9. Rinse the slides in phosphate-buffered saline at neutral pH for 3 changes in 3 different coplin jars, 5 mins each.

10. Incubate the slides in 1% BSA in Dulbecco’s Phosphate-Buffered Saline (DPBS) containing 0.1% Triton-X-100 for 30 mins, at room temperature.

11. Drain off the blocking buffer and Make liquid blocker around the tissue and allow drying for 15 sec.

12. Apply 100μl of an appropriately diluted primary antibody in 1% BSA in DPBS containing 0.1% Triton-X-100 to the sections on the slides and incubate the slides in humidified chamber for 1h at room temperature or overnight at 4οC.

13. Rinse the slides in 300 ml DPBS for 3 changes in 3 different coplin jars, 5 mins each.

14. Apply 100μl of an appropriately diluted secondary antibody in 1% BSA in DPBS containing 0.1% Triton-X-100 to the sections on the slides and incubate the slides in humidified chamber for 1h at room temperature.

15. Rinse the slides in 300 ml DPBS for 3 changes in 3 different coplin jars, 5 mins each.

16. Mount the coverslips gently onto the slides using 20μl Vectashield Mounting Medium containing DAPI for nuclear staining.

 

Flow Cytometry

Staining Cell Surface Antigens

1. Aliquot 100μL of cells per tube (1x106 cells total). Add 20 μL blocking reagent containing 2.4 G2 (anti- Fc receptor (0.5-1μg) for mouse cells. Incubate 20 mins on ice.

2. Add appropriately diluted unconjugated primary antibodies to each tube, vortex and incubate for 60 mins at 4ο C. Avoid exposure to light.

3. Add 200μL of 1x PBS to each tube to wash off excess antibody. Centrifuge for 5 mins at 250xg. Aspirate the supernatant, being careful not to disturb the pellet.

4. Add 100μL of 1X PBS to each tube. Add appropriately diluted fluorochrome-conjugated secondary antibodies to tubes. Vortex and incubate for 30 mins at 4οC. Avoid exposure to light.

5. Add 2 ml of 1x PBS to each tube to wash off excess antibody. Centrifuge for 5 mins at 250xg. Aspirate the supernatant, being careful not to disturb the pellet.

6. [Optional] Add a viability dye; either 7-AAD or Propidium iodide (5 μl/sample) to exclude dead cells from analysis.

Staining Intracellular Antigens

7. Stain cell surface antigens as described above (step 1 to 6)

8. After the last wash, discard the supernatant and pulse-vortex the sample to completely dissociate the pellet.

Fixation and Permeabilization

Fixation in 0.01% formaldehyde. Permeabilization in Triton-X-100 or NP-40 (0.5% in PBS)

9. Add 100 μL of fixative and incubate for 10 mins at room temperature.

10. Add 100 μL detergent based permeabilizing agent and incubate at room temperature for 15 minutes.

11. Centrifuge the cells at 300xg for 5 mins, discard the supernatant and re-suspend the pellet in remaining volume.

12. Follow antibody staining protocol as indicated in our “direct” and “indirect” protocols.

Note: If combining both direct and indirect staining together, the sequence of staining will be:

Primary antibody followed by fluorochrome-conjugated secondary antibody and then stain directly (fluorochrome-conjugated primary antibody).

 

Lysate Preparation

1. Weigh the tissue sample in a 50 mL tube.

2. Keep the sample on ice and wash with ice-cold 1x PBS and aspirate off the PBS.

3. Repeat until wash buffer appears clear.

4. Add sufficient volume of cold lysis buffer with 1X PIC, enough to cover the sample. Note: RNase/ Dnase (0.02 mg/ml) may be added to the samples to facilitate RNA and DNA digestion.

5. Grind/homogenise tissue in tube and incubate on ice for upto 60 mins.

6. Transfer mixture to microcentrifuge tubes and spin at 14, 00 rpm for 40 mins at 4οC.

7. Poke through lipid layer and remove supernatant. Discard the cellular debris and lipids.

8. If necessary, re-spin supernatant at 14, 000 rpm and repeat step 7 to obtain clean lysate free of debris and lipids.

9. Determine the protein concentration using Bradford Protein Assay.

10. Combine equal volumes of 2X SDS sample buffer and cell lysate supernatant. Pipet up and down or vortex several times to mix. Aliquots can be stored at -80οC for further use. Prior to use, heat samples at 95-100οC for 3-5 mins and then load immediately on SDS-PAGE gel. Avoid freeze/thawing lysate as much as possible.

1x Lysis Buffer Composition:

-10ml\A Tris pH 8.0 – 1.6 mg/ml Benzamidine HCL

-130 ml\A NaCL -1.0mg/ml Phenanthroline

-1% Triton-X-100 – 1.0 mg/mL Aprotonin

-10 ml\A NaF – 1.0 mg/ml Leupeptin

-10 ml\A NaPi pH 7.5 (sodium phosphate)- 1.0 mg/ml Pepstatin A

 


 

 

 

Body Fat Analysis in Rodents

1. Collect background blood sample (~75 μL; to have atleast 12 μL of plasma) from the mice into tubes containing EDTA.

2. Inject 3 μg deuterium oxide/g body weight intraperitoneally into the mice.

3. After 1 hour, collect a blood sample (~75 μL; to have atleast 12 μL of plasma) from the mice into tubes containing EDTA.

4. Centrifuge the blood samples to obtain plasma

5. Store at -80ο C

Note: Enrichment of approximately 0.2% to 0.4% is achieved at 3 μg deuterium oxide/g body weight.