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Protocols

 

IHC/IF Fixation

Fixation and permeabilization for IHC and ICC

Fixation

Fixation should immobilize antigens while retaining cellular and subcellular structure. It should also allow for access of antibodies to all cells and subcellular compartments. The fixation and permeablization method used will depend on the sensitivity of the epitope and antibody themselves, and may require some optimization.

Fixation can be done using crosslinking reagents, such as paraformaldehyde. These are better at preserving cell structure, but may reduce the antigenicity of some cell components as the crosslinking will obstruct antibody binding. For this reason, antigen retrieval techniques may be required, particularly if there is a long fixation incubation or if a high percentage of crosslinking fixative is used. Another option is to use organic solvents. These remove lipids while dehydrating the cells. They also precipitate proteins on the cellular architecture.

4% Paraformaldehyde

  1. Add 4% paraformaldehye to slides for 10 min only

  2. Rinse with PBS or PBS 1% BSA

Fixing in paraformaldehyde for more than 10-15 minutes will cross link the proteins to the point where antigen retrieval may be required to ensure the antibody has free access to bind and detect the proteins of interest.

Ethanol

  1. Add 100-200 μl per slide of cooled 95% ethanol, 5% glacial acetic acid for 5-10 min. Wash with PBS or PBS 1% BSA

Methanol

  1. Add 100-200 μl per slide of ice cold methanol

  2. Place at -20°C for 10 min

  3. Wash with PBS or PBS 1% BSA

Methanol will also permeabilize, but not in all cases as some epitopes are very sensitive to this. Can try acetone instead for permeabilization if required.

Acetone

  1. Add 100-200 μl per slide ice cold acetone. Place at -20°C for 5-10 min

  2. Wash with PBS or PBS 1% BSA

Acetone will also permeabilize, no permeabilization step required.

Permeabilization should only be required for intracellular epitopes when the antibody required access to the inside of the cell to detect the protein. However, it will also be required for detection of transmembrane membrane proteins if the epitope is in the cytoplasmic region.

Solvents

  1. Acetone fixation will also permeabilize

  2. Methanol fixation can be used to permeablize but is not always suitable.

These reagents can be used to fix and permebilize, or can be used after fixation with a crosslinking agent such as paraformaldehyde to permeabilize the cells.

Detergents

Triton x-100

  1. Use 0.1 to 0.2% in PBS, 10 min only

  2. These will also partially dissolve the nuclear membrane and are therefore very suitable for nuclear antigen staining

As these are harsh detergents, they will disrupt proteins if they are used at higher concentrations or for longer amounts of time which will affect staining results.

Tween 20, Saponin, Digitonin and Leucoperm

  1. Use 0.2 to 0.5% for 10-30 min.

These are much milder membrane solubilizers. They will give large enough pores for antibodies to go through without dissolving plasma membrane. Suitable for antigens in the cytoplasm or the cytoplasmic face of the plasma membrane. Also suitable for soluble nuclear antigens.

Special recommendations

Cytoskeletal, viral and some enzyme antigens usually give optimal results when fixed with acetone, ethanol or formaldehyde (high conc.).

Antigens in cytoplasmic organelles and granules will require a fixation and permeabilization method depending on the antigen. The epitope needs to remain accessible.

Histology

Tissue for paraffin embedding

For the best preservation of tissue morphology, the thickness of the tissue should be between 0.2 and 0.5 cm.

  • For further haematoxylin and eosin (H&E) staining, we recommend that you use a fresh solution of 10% neutral buffered formalin with a volume of at least 10 times that of the specimen

  • For further immunoassaying, we recommend that you use 70% ethanol with a volume of at least 10 times that of the specimen

Tissue for frozen section

For the best results, do not fix tissue for frozen section histology in a chemical fixative. Place tissue in a mould and surround by a cryoprotectant (e.g., OCT) before being snap frozen.

  • For help with freezing technique, please contact the histology lab

  • Tissue should be frozen as soon as possible to prevent the damaging effects of autolysis.

Fluorescence

Immuofluorescence Protocol

General procedure

  1. Grow cells on glass coverslips or prepare cytospin or smear preparation

  2. Rinse briefly in phosphate-buffered saline (PBS)

Fixation

  1. Fix the samples either in ice-cold methanol, acetone (1-10 min) or in 3-4% paraformaldehyde in PBS pH 7.4 for 15 min at room temperature

  2. Wash the samples twice with ice cold PBS. Permeabilization: If the target protein is expressed intracellularly, it is very important to permeabilize the cells. Note: acetone fixed samples do not require permeabilization

  3. Incubate the samples for 10 min with PBS containing 0.25% Triton X-100 (or 100 μM digitonin or 0.5% saponin). Triton X-100 is the most popular detergent for improving the penetration of the antibody. However, it is not appropriate for the use of membrane-associated antigens since it destroys membranes

  4. Wash cells in PBS three times for 5 min

Blocking and incubation

  1. Incubate cells with 1% BSA in PBST for 30 min to block unspecific binding of the antibodies (alternative blocking solutions are 1% gelatine or 10% serum from the species that the secondary antibody was raised in)

  2. Incubate cells in the diluted antibody in 1% BSA in PBST in a humidified chamber for 1 hr at room temperature or overnight at 4°C

  3. Decant the solution and wash the cells three times in PBS, 5 min each wash

  4. Incubate cells with the secondary antibody in 1% BSA for 1 hr at room temperature in dark

  5. Decant the secondary antibody solution and wash three times with PBS for 5 min each in dark

Counter staining

  1. Incubate cells on 0.1-1 μg/ml DAPI (DNA stain) for 1 min

  2. Rinse with PBS

Mounting

  1. Mount coverslip with a drop of mounting medium

  2. Seal coverslip with nail polish to prevent drying and movement under microscope

  3. Store in dark at 4°C

TEM

E.M. PROCESSING SCHEDULE for EPOXY RESIN

  1. Fix tissue in 2.5% glutaraldehyde in 0.1M sodium cacodylate buffer at 4oC, for a minimum of 4 hours. Tissue should be cut into approx. 1mm cubes for fixing. This may be done in a drop of fix on a sheet of dental wax, using a razor blade. Place in glass processing vials and close with plastic caps. (Tissue may be stored at this stage.)

  2. Wash in 0.1M buffer - 1 hour x 2. (Or overnight at 4oC.)

  3. Post fix in 1% osmium tetroxide in distilled water - 1 hour. (Mix equal quantities of 2% aqueous OsO4and distilled water and use immediately.)

  4. Rinse in distilled water - 5 mins. x 2

  5. Dehydrate in 70% ethanol - 20 mins. x 2. (May be stored overnight at this stage if absolutely necessary.)

  6. Dehydrate in 90% ethanol - 10 mins. x 2

  7. Dehydrate in 100% ethanol - 20 mins. x 2

  8. Epoxy resin mixture (50/50) - 1 hour

  9. Epoxy resin - overnight - with caps removed from vials

  10. Embed in labelled capsules with freshly prepared resin

  11. Polymerise at 60oC - 48 hours

Notes

  • All steps must be performed in a fume cupboard and gloves should be worn throughout

  • Osmium tetroxide and resin waste should be collected in bottles for safe disposal

  • For steps 2 to 10 the processing vials should be on a rotating mixer

  • It is often beneficial to increase the processing times for specimens with a high connective tissue content such as skin, tendon, cartilage, cornea, etc. This should be done for all steps and especially for the 50/50 mixture at step 9 (which can even be extended to an overnight treatment). Failure to increase the times may result in a poorly infiltrated block and therefore weak sections which will fail under the electron beam, or even be impossible to cut in the first place


E.M. PROCESSING SCHEDULE - ACRYLIC RESIN

  1. Fix tissue in 2.5% glutaraldehyde in 0.1M sodium cacodylate buffer at 4oC, for a minimum of 4 hours. Tissue should be cut into approx. 1mm cubes for fixing. This may be done in a drop of fix on a sheet of dental wax, using a razor blade. Place in glass processing vials and close with plastic caps. (Tissue may be stored at this stage.)

  2. Wash in 0.1M buffer - 1 hour x 2. (Or overnight at 4oC.)

  3. Post fix in 1% osmium tetroxide in 0.2M buffer - 1 hour. (Mix equal quantities of 2% aqueous OsO4and distilled water and use immediately.)

  4. Rinse in 0.1M buffer - 5 mins. x 2

  5. Dehydrate in 70% ethanol - 20 mins. x 2. (May be stored overnight at this stage if absolutely necessary.)

  6. Dehydrate in 90% ethanol - 10 mins. x 2

  7. Dehydrate in 100% ethanol - 20 mins. x 2

  8. LR White resin - overnight.

  9. LR White resin - 1 hour

  10. LR White resin - 1 hour

  11. Embed in closed labelled gelatine capsules filled with fresh resin

  12. Polymerise at 60oC - 24 hours. (Polymerise at 50oC for immunocytochemistry.)

Notes

  • All steps must be performed in a fume cupboard and gloves should be worn throughout

  • Osmium tetroxide/buffer waste should be collected in bottles for safe disposal. Resin waste may be polymerised

  • Steps 2 to 10 - processing vials should be on a rotating mixer

  • If results are urgently required steps 8-10 may be shortened by performing 4-6 changes of resin at 60oC over 3 hours

  • Be aware of the recommendations for processing 'difficult' tissues in the notes for epoxy resin processing. The same will apply when using acrylic resin

  • If polymerisation using the accelerator is necessary step 3, osmium tetroxide, must be omitted or artefact will occur due to overheating. Step 3 should also be omitted if immunocytochemistry is to follow. Resin which is nearing its "use by" date should not be used with OsO4 as some polymerisation may occur during step 8


AGAR/RESIN EMBEDDING OF CELL SAMPLES

Cell samples suspended in fluid may produce a pellet which is cohesive enough to process after spinning at 5,000 rpm for 5 mins. But if not they should be embedded in high strength agar gel.

Samples are best put straight into fixative upon collection. If they arrive at the laboratory in any other medium spin them in 1.5ml Eppendorf tubes at 5,000 rpm for 5 mins., take off the supernatant, replace it with 2.5% glutaraldehyde in 0.1M sodium cacodylate buffer and leave for a minimum of 4 hours at 4oC.

  1. Decant the fixative and replace with 0.1M sodium cacodylate buffer

  2. Re-suspend the sample and leave for 2 hours (or overnight) then re-spin

  3. Prepare a 1% solution of high strength agar in distilled water by bringing to the boil whilst stirring

  4. Decant the buffer from the sample tubes and take them and the agar solution to the centrifuge

  5. When the agar solution has cooled to approximately 60oquickly fill each tube with it, resuspend the samples and spin them at full speed for 30 secs - 1 min. (Maximum of 4 samples at a time or the agar will set before the sample can be spun to the bottom of the tube)

  6. Cool the tubes in a beaker of cold water to set the agar

  7. Remove the agar plug with a mounted needle and cut off the end containing the sample

  8. Cut up the sample in agar (1mm cubes) and place in 0.1M sodium cacodylate buffer

  9. Continue with the E.M. processing schedule from step 3

SEM

SEM tissue processing using HMDS

  • Fix cells sample in 2.5% glut in 0.1M phosphate buffer pH7.2 for 12 hours at 4 C

  • Buffer wash 2 * 5 mins

  • 1% osmium in distilled water 1 hour

  • Buffer wash 3 * 5 mins

  • 70%, 80%, 90% ethanol 10mins each

  • 100% ethanol 3 * 10 mins

  • HMDS 3 mins

  • Samples then removed and placed in desiccator for 25 mins to avoid water contamination

Useful links

Here are some useful links to help you with your microscopy and histology work:

Publications

https://abdn.pure.elsevier.com/en/equipments/microscopy-and-histology