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Immunostaining Protocols

Immunostaining Workflow
Immunostaining Workflow

Immunostaining Protocols

Protocols for Cryostat Sections

Fresh Frozen (then Fixed) Tissue Sections

  1. Snap-freeze small tissue blocks (5x5x3 mm) in liquid nitrogen
  2. Transfer to cryostat and cut thin (5-30 m) sections.
  3. Collect specimens on clean poly-L-lysine-coated glass slides and dry at room temperature overnight (if you want to stain the same day let air-dry for 1-2 hrs. until completely dry). Thorough drying is essential for good adhesion to the slides.
  4. Fix sections in acetone or absolute ethanol at 4°C for 15 min. Use fresh ethanol or acetone for every 10-15 slides for best results.The organic solvents absorb moisture from the air and tissue, as they do so, they lose their ability to fix the tissue effectively.
  5. Thoroughly air-dry at room temperature or on mild heat (30-37°C). It is during this stage that much of the chemical fixation is being finalized; improper air-drying will lead to "soft" sections and likely loss of proper reactivity.
  6. Proceed with immunostaining or freeze.

Fixed, Frozen Tissue Sections

  1. Fix tissue either by perfusion with fixative or by immersion in fixative for a set time period. Most commonly, 4% Paraformaldehyde (PFA) solutions are used.
  2. Fixed tissue is then prepared for cryoprotection by submerging the target tissue in a hydrostabilizing solution. The cryoprotection is complete when the target tissue no longer floats in the stabilizing solution. Because it works well and is relatively inexpensive, PBS+sucrose solutions ranging from 10% (less protection), to 30% (w/v) sucrose (greater protection) are often used.
  3. Once stabilized, tissues can be removed from the protectant solution and frozen at -70°C until sectioned.
  4. Sectioned via cyrostat (5-40 μm*), where sections can be collected directly onto slides, or floated onto slides via a PBS/waterbath. Usually up to 3 sections per slide can be placed; each spaced well apart. The spacing prevents reagent mixing between samples.
  5. Sections-on slides are thoroughly air/warmed dried on a slide warmer, usually overnight or at least 2-3 hrs. at 40-50°C.
  6. Prepared slides can be stored dry at -70°C until stained. Equilibrate to room temperature and briefly redry prior to rehydration and staining.
*Individual skill and tissue type will determine the thickness of the sections. Sections between 10-15 m provide the best results for clarity and integrity. Sections between 6-9 m tend to tear during cutting, resulting in rough edges that can increase.

Protocol for Paraffin-embedded Sections

  1. Conventional deparaffinization and dehydration sequence
    1. Incubate sections in Xylene: 2 to 3 changes, 5 min. each.
    2. 100% absolute ethanol: 2 changes, 3 min. each
    3. 95% ethanol: 2 changes, 3 min. each
    4. 80% ethanol: 3 min.
    5. 50% ethanol: 3 min.
    6. Rinse with distilled water, PBS, or Tris buffer: 2 changes, 3 min. each.
    7. Note: Once sections have been rehydrated, do not allow them to dry.
  2. Place slides in prewarmed (37°C) 0.1% trypsin in PBS for 5-60 min. or 0.4% pepsin in 0.01N HCl for 30 min. to one hour. Follow by rinsing with distilled water
  3. If peroxidase conjugate is used, endogenous peroxidase should be blocked at this stage. Peroxidase activity results in the decomposition of hydrogen peroxide (H2O2). It is a common property of all hemoproteins such as hemoglobin, myoglobin, cytochrome and catalases. Suppression of endogenous peroxidase activity in formalin-fixed tissue entails the incubation of sections in 3% H2O2 for 8-10 min. Methanolic H2O2 treatment (1 part 3% H2O2 plus 4 parts absolute methanol) for 20 min. can also be used, but it is not recommended for specimens where cell surface markers are to be stained. Methanolic treatment may also detach frozen sections from their carrier glass.
  4. Wash twice with PBS.
  5. Proceed with immunostaining procedure (see Antibody Staining section).

General Protocol for Immunohistochemical Staining

The following general protocol is intended for use as a guideline in developing antibody-specific procedures. Different antibodies and tissues may require changes to this procedure. Review of individual product datasheets and relevant literature references may be helpful in customizing this procedure for specific applications.

Single-protein detection (specificity) on western blots is an absolute requirement. It is strongly advised that the researcher test each antibody on a western blot of the tissue they intend to use, to control for differences between tissues. Once the western blot is verified, the antibody can be tested on tissue sections, with negative controls consisting of no primary antibody (to see if there is direct staining by the secondary antibody), and a no secondary antibody control (to see if the primary antibody contributes endogenous peroxidase activity, or autofluorescence).

Since the protocols below are general, it is highly recommended to review the methodology and variations in IHC protocols; Immunocytochemical Methods and Protocols (second edition), edited by Lorette C. Javois, from Methods in Molecular Medicine, volume 115, Humana Press, 1999 (ISBN 0-89603-570-0).

  1. Gently rinse slide containing sections with distilled water or buffer from a wash bottle. Place slide in room temperature buffer bath for 5 minutes to rehydrate sections.
  2. Using a Kimwipe®, gently remove excess liquid from around the specimen. Avoid touching the tissue directly.
  3. Apply 4-6 drops of normal serum, (normal serum from the host of the secondary antibody), diluted 1:5-1:30 (final conc. 3%-20%). Incubate for 20-30 minutes at 37°C.
  4. Tap off serum and wipe away excess. Do not rinse.
  5. Perform any antigen retrieval if necessary.
  6. Apply 25-50 L of rabbit (mouse) primary antibody, diluted appropriately, per tissue section. Antibody should cover sections completely. Incubate for desired time (see above for suggested parameters and temperatures). If optimal antibody dilution is unknown, perform a series of antibody dilutions in the range of 1:20-1:1,000 to obtain initial results.
  7. Note: Antibody diluent is often very important for consistent reactivity. Simple solutions are easier to troubleshoot then complex ones, thus antibodies diluted only with simple buffers (PBS or TBS) are usually recommended.
  8. Rinse slide gently with distilled water or buffer from a wash bottle, and incubate in a buffer bath for 3x5 minutes (changing buffer in between washes).
  9. Note: For all procedures it is important to see that each step is adequately buffered, and that non-reacted solutions are washed away after each step.
  10. Apply 25-50 L of enzyme-conjugated antibody directed against rabbit (mouse) immunoglobulins, diluted appropriately. Incubate 45-60 minutes.
  11. Rinse slide gently with distilled water or buffer from a wash bottle, and incubate in a buffer bath for 3x5 minutes (changing buffer in between washes).
  12. Apply substrate-chromogen solution and incubate until desired color intensity has developed.
  13. Rinse gently with distilled water from wash bottle.
  14. Counterstain and coverslip.

IHC Select; Heat Induced Epitope Retrieval (HIER) Technique for Paraffin Sections

  1. Cut and mount sections on slides coated with "apes" {APES (3-aminopropyltriethoxysilane).
  2. To avoid sections becoming detached, sections should be mounted on "APES" covered slides, then dried at 37°C overnight followed by drying at 60°C for 60 minutes.
  3. Deparafinize sections through alcohols and rehydrate with distilled water; sections are now ready for antigen retrieval.

Steam/Pressure Antigen Retrieval: Suggested Protocol

  1. Rice Steamer: Black and Decker HS2000 Type 1 Using one of the following target retrieval solutions
    • 10mM Citrate Buffer, pH 6.0
    • 1.0mM EDTA, pH 8.0
    • 10mM Tris EDTA (T.E.) Buffer, pH 9.0
    • Note: If an enzymatic digestion is required after the high temperature unmasking technique, it is performed after the initial antigen retrieval step in most cases.
  2. Add 1 liter of room temperature distilled water to Hi Fill mark.
  3. Turn on the steamer's timer when you reach your last alcohol, thus allowing the water to come to boiling (8-15 minutes).
  4. Place your slides in a staining bucket containing the target retrieval solution of your choice and incubate for 2 min.
  5. Add 10 milliliters of fresh target retrieval solution to a reagent tray and using a capillary gap technique; place your tissue slide and a blank clean slide into the tray face to face allowing the reagent to cover the tissue (about 250 ul). Alternative applications methods can be used such as Coplin jars but more reagents will be used.
  6. Place the reagent tray with your slides into the steamer's vegetable tray and placed it on the already boiling water; cover steamer with its lid and incubate.
  7. Incubate for 20 minutes. Add one more minute to your time for a total of 21 min. This allows the target retrieval solution to equilibrate to the same temperature of the distilled water. (If using larger quantities of liquid, timings must be increased).
  8. After timer goes off, remove vegetable tray and allow your slides to cool down for about 10 to 15 minutes. Rinse with room temperature PBS before proceeding with staining protocol.

Immunocytochemistry General Protocol

Immunocytochemistry (ICC), by definition is the demonstration of a tissue constituent in situ by detecting specific antibody-antigen interactions where the antibody has been tagged with a visible label. The visual marker may be a fluorescent dye, colloidal metal, hapten, radioactive marker or the more commonly light microscopy an enzyme. Experimental samples ranging from frozen sections, cell culture/suspension, to whole tissue samples have been used. Ideally, maximal signal strength along with minimal background or non-specific staining are required to give optimal antigen demonstration. It is recommended to review the methodology and variations in protocols from; Immunocytochemical Methods and Protocols (second edition), edited by Lorette C. Javois, from Methods in Molecular Medicine, Humana Press, 1999 (ISBN 0-89603-570-0).

Procedure

  1. Place a previously autoclaved, 13 mm circular glass coverslip in the well of a 24 well plate.
  2. Plate approximately 1mL of cell suspension into each well. Incubate 24 hours in a 37°C CO2 incubator.
  3. Aspirate media from wells
  4. Add appropriate fix and incubate for the appropriate time at room temp.
  5. Wash the cells with PBS, twice, for 15 minutes. Do not shake.
  6. Cover cells with 400 l of 1% BSA in PBS and incubate for 1 hour at room temp.
  7. Wash the cells with PBS for 15 minutes.
  8. Incubate the cells with primary antibody of interest in 1% BSA in PBS and incubate for 2 hours at room temperature.
  9. Wash the cells twice with PBS for 5 minutes.
  10. Incubate the cells with a dilution of IgG fluorescein conjugated secondary antibody of choice in 1% BSA in PBS for 1 hour at room temperature.
  11. Wash the cells three times with PBS.
  12. Aspirate well dry.
  13. Clean a glass slide with Alconox™ and water, follow with a rinse in 70% ETOH, dry using a Kimwipe.
  14. Place a drop of Aqua Poly-Mount mounting media on cleaned slide.
  15. Using forceps and/or a 26 gauge needle with the tip bent, retrieve the glass coverslip from the well and place it cell side down on top of the drop of mounting media.
  16. Let dry at room temperature, seal edge if desired and examine the cells under a fluorescent microscope.

출처: http://www.millipore.com