Introduction

Immunocytochemistry is a technique used to assess the presence of a specific protein or antigen in cells (cultured cells, cell suspensions) by use of a specific antibody, which binds to it, thereby allowing visualization and examination under a microscope. It is a valuable tool for the determination of cellular contents from individual cells. Samples that can be analyzed include blood smears, aspirates, swabs, cultured cells, cell suspensions, and cytospin. Each sample is treated differently, yet all the methods are interchangeable. There is no one way to prepare these types of cell samples for immunocytochemical analysis.

Cells to be stained can be attached to a solid support to allow easy handling in subsequent procedures. This can be achieved by several methods: adherent cells may be grown on microscope slides, coverslips, or an optically suitable plastic support. Suspension cells can be centrifuged onto glass slides (cytospin), bound to solid support using chemical linkers, or in some cases handled in suspension.

Concentrated cellular suspensions that exist in a low-viscosity medium make good candidates for smear preparations. Dilute cell suspensions existing in a dilute medium are best suited for the preparation of cytospins through cytocentrifugation. Cell suspensions that exist in a high-viscosity medium, are best suited to be tested as swab preparations. The constant among these preparations is that the whole cell is present on the slide surface. For any intercellular reaction to take place, immunoglobulin must first traverse the cell membrane that is intact in these preparations. Reactions taking place in the nucleus can be more difficult, and the extracellular fluids can create unique obstacles in the performance of immunocytochemistry. In this situation, permeablizing cells using detergent (Triton X-100 or Tween-20) or choosing organic fixatives (acetone, methanol, or ethanol) becomes necessary.

HistoGel preparation will overcome the penetration difficulty into cellular nucleus since the cells cover with histogel will be treated as regular tissues for paraffin embedding and sectioning. Therefore, there is a possibility that cell nucleus can be presented for antibody binding.

Antibodies are an important tool for demonstrating both the presence and the subcellular localization of an antigen. Cell staining is a very versatile technique and, if the antigen is highly localized, can detect as few as a thousand antigen molecules in a cell. In some circumstances, cell staining may also be used to determine the approximate concentration of an antigen, especially by an image analyzer.

Cell Preparation

Preparation of Coated Slides

1.1 Position clean glass slides in a staining rack

1.2 Immerse the slides for 30 min in a large staining dish containing a 1:10 dilution of 0.1% poly-L-lysine solution in deionized water.

1.3 Remove the slides and oven dry for 1 hr at 60 C.

Cell Film Slide Preparation for Blood and Cell Suspension

2.1 Add a drop of cells (blood, cell suspension, etc.) to the end of the coated glass slide.

2.2 Hold the beveled edge slide at a 45° angle to the plane of the coated slide and gently touching the surface of the slide, back the edge over the drop of cells so they spread within the 45° angle, the width of the slide

2.3 Slide the beveled edge slide toward the other end of the preparation slide (in the direction of the 135° angle) with a rapid uniform motion. The cells will spread out over the surface of the slide and form a film with a feathered edge, if done properly.

2.4 Allow to air dry for 30 minutes.

2.5 Immerse in cooled acetone or other fixatives to fix the cells.

2.6 Rinse slides 3x5 min in PBS.

2.7 Incubate the slides with 0.25-0.5% Triton X-100 in PBS for 10 minutes to permeabilize the membranes (Note: there is no need for a permeabilization step following acetone or methanol fixation).

2.8 Rinse 3x5 min in PBS.

2.9 Blocking endogenous peroxidase by incubating in 3% H2O2 in PBS for 10-30 minutes (Do this step only if a peroxidase marker is to be used. Omit this step if a fluorescent or AP marker is to be used).

2.10 Rinse in 3x5 min in PBS.

2.11 Blocking with 2-5% normal serum in PBS for 1 hour (Normal serum should be the same species as the secondary antibody is raised).

2.12 Incubate with primary antibody and proceed to routine immunocytochemistry procedure (see below)

Swabbed Slide Preparation

3.1 Obtain sample on sterile swab.

3.2 Smear the sample onto the glass slide using the majority of the surface area to distribute the specimen.

3.3 Spray fix the material with Cyto Prep.

3.4 Allow to air dry.

3.5 Postfix the slides with 10% neutral buffered formalin for 30 seconds.

3.6 Rinse 3x5 min with PBS.

3.7 Incubate the slides with 0.25-0.5% Triton X-100 in PBS for 10 minutes to permeabilize the membranes (Note: there is no need for a permeabilization step following acetone or methanol fixation).

3.8 Rinse 3x5min in PBS to remove detergents.

3.9 Blocking endogenous peroxidase by incubating in 3% H2O2 in PBS for 10-30 minutes (Do this step only if a peroxidase marker is to be used. Omit this step if a fluorescent or AP marker is to be used).

3.10 Rinse in 3x5 min in PBS.

3.11 Blocking with 2-5% normal serum in PBS for 1 hour (Normal serum should be the same species as the secondary antibody is raised).

3.12 Incubate with primary antibody and proceed to routine immunocytochemistry procedure.

Cell Culture Slide Preparation

4.1 Transfer 200 ul of cell culture to the wells of a chamber slide or slides of your choice (Choice of slide design is often dictated by the experiment. Some slides have four wells, some have eight, some are glass, and some are plastic. Glass is recommended simply because the slide becomes more versatile, acetone can be used as a fixative, and finished slides can be dehydrated in ethanol and cleared in xylene)

4.2 Allow cells to grow to confluence with the addition of fresh media.

4.3 Wash the cells thoroughly 5x2 min in PBS.

4.4 Fix cells with 95% ethanol, 5% glacial acetic acid for 3-5 minutes.

4.5 Rinse 5x2 min in PBS.

4.6 Incubate the slides with 0.25-0.5% Triton X-100 in PBS for 10 minutes to permeabilize the membranes (Note: there is no need for a permeabilization step following acetone or methanol fixation).

4.7 Rinse 3x5 min in PBS.

4.8 Blocking endogenous peroxidase by incubating in 3% H2O2 in PBS for 10-30 minutes (Do this step only if a peroxidase marker is to be used. Omit this step if a fluorescent or AP marker is to be used. 

4.9 Rinse in 3x5min in PBS.

4.10 Blocking with 2-5% normal serum in PBS for 1 hour (Normal serum should be the same species as the secondary antibody is raised).

4.11 Incubate with primary antibody and proceed to routine immunocytochemistry procedure.

Preparation of Cell Culture Block for Paraffin Embedding Using HistoGel

5.1 Microwave HistoGel (Richard Allan Scientific, Cat# HG-4000-012) on low for 5-15 seconds. Make sure to loosen the cap before heating a tube of HistoGel to prevent rupturing of the tube (An alternatively you can also place HistoGel into a boiling water bath for 3-10 minutes.

5.2 After HistoGel is liquefied, the temperature may be lowered to 50 C ± 5 and it will remain in the liquid state. A lower temperature will allow the gel to solidify more quickly after it is dispensed onto a specimen.

5.3 Centrifuge ethanol-fixed cell suspension.

5.4 Remove the supernatants from the centrifuge tube.

5.5 Add 4-6 drops of liquefied HistoGel with a pipette to cell pellet at bottom of centrifuge tube.

5.6 Either vortex specimen for several seconds to adequately and thoroughly mix cells and HistoGel together, OR allow HistoGel to settle to the bottom of tube.

5.7 Allow HistoGel to solidify by cooling to near room temperature (<20 C). This can be achieved by use a cooling plate, ice tubes, freeze pack, or allowing to cool naturally.

5.8 Remove HistoGel pellet containing the specimen and place inside a tissue cassette.

5.9 Histologically process HistoGel “button” containing the cell pellet as a standard histology specimen.

5.10 Paraffin embedding and sectioning, and then following standard immunohisotchemical staining procedure.

Cytospin Slide Preparation

6.1 Position slides in slide holders with filter cards and sample chambers and attach to cytocentrifuge rotor.

6.2 Prepare cell suspension of 500 cells/mm3 (ul) with PBS (Preparations that are too dilute for cytocentrifugation may be dropped with a pipet using the location of the filter card as a guide, and allowed to dry).

6.3 Add 0.1 ml cell suspension to chamber and centrifuge at 1000 rpm for 5 minutes (It is important to get just the right amount of speed for cytocentrifugation, as too much speed will flatten the cell, and too little will not allow the cells to adequately bind to the slide).

6.4 Remove slide and immediately dip in 95% ethanol, 5% glacial acetic acid fixative for 2 minutes.

6.5 Rinse 3x5 min with PBS to remove fixative.

6.6 Incubate the slides with 0.25-0.5% Triton X-100 in PBS for 10 minutes to permeabilize the membranes (Note: there is no need for a permeabilization step following acetone or methanol fixation).

6.7 Rinse 3x5min in PBS to remove detergents.

6.8 Blocking endogenous peroxidase by incubating in 3% H2O2 in PBS for 10-30 minutes (Do this step only if a peroxidase marker is to be used. Omit this step if a fluorescent or AP marker is to be used).

6.9 Rinse 3x5 min in PBS.

6.10 Blocking with 2-5% normal serum in PBS for 1 hour (Normal serum should be the same species as the secondary antibody is raised).

6.11 Incubate with primary antibody and proceed to routine immunocytochemistry procedure.

Touch Prep Slide Preparation

7.1 Take the refrigerated acetone and add to a Coplin jar.

7.2 Position the excised tissue directly above the slides, best side facing the slide, using forceps (The purpose for the preparation of these types of slide is to examine the cells of a tissue quickly, with no need for tissue preparation and cutting. It is important to work swiftly, as the tissue will start to deteriorate the moment it is obtained. Therefore, it is important to have all the preparations ready for the rapid handling of the excised tissue. As soon as a surface of the tissue is decided on, it should immediately be touched to a waiting slide, oriented properly and ready to be fixed).

7.3 Align the slide horizontally on flat surface.

7.4 Gently touch the selected exposed tissue surface down onto the slide (Excess tissue fluid may be absorbed with a little gauze, but avoid touching any wet cells. Also, gentle pressure is required, but if too forceful, some cells may be destroyed).

7.5 Apply slight pressure then remove the tissue after a few seconds.

7.6 With a minimum of disturbance, immerse slide into the cold acetone solution for 5-10 minutes (For touch preps, the slide should be immersed in the acetone as soon as possible, but the cells need a moment to adhere to the plane of the glass. Slowly dip the slide into the acetone, as a violent action at this point could wash off some of the cells. As is the case with frozen sections, fixation is a matter of choice. In this instance, the use of acetone is preferred because the cells are still whole, and the membranes require disruption in order for the contents to be accessible for later analysis. However, an ethanol/acid fixative - 95% ethanol, 5% glacial acetic acid, is perfectly acceptable if desired).

7.7 Allowed to air dry.

7.8 Rinse the slides 3x5 min in PBS.

7.9 Incubate the slides with 0.25-0.5% Triton X-100 in PBS for 10 minutes to permeabilize the membranes (Note: there is no need for a permeabilization step following acetone or methanol fixation).

7.10 Rinse 3x5min in PBS to remove detergents.

7.11 Blocking endogenous peroxidase by incubating in 3% H2O2 in PBS for 10-30 minutes (Do this step only if a peroxidase marker is to be used. Omit this step if a fluorescent or AP marker is to be used. 

7.12 Rinse in 3x5 min in PBS.

7.13 Blocking with 5% normal serum in PBS for 1 hour (Normal serum should be the same species as the secondary antibody is raised).

7.14 Incubate with primary antibody and proceed to routine immunocytochemistry procedure.

Fixation

To ensure free access of the antibody to its antigen, the cells must be fixed and permeabilized. In general, fixation strengths and times are considerably shorter for cells than on the thicker, structurally complex tissue sections. For immunocytochemistry, sample preparation essentially entails fixing the target cells to the slide. Perfect fixation would immobilize the antigens, while retaining authentic cellular and subcellular architecture and permitting unhindered access of antibodies to all cells and subcellular compartments. Wide ranges of fixatives are commonly used, and the correct choice of method will depend on the nature of the antigen being examined and on the properties of the antibody used. Fixation methods fall generally into two classes: organic solvents and cross-linking reagents. Organic solvents such as alcohols and acetone remove lipids and dehydrate the cells, while precipitating the proteins on the cellular architecture. Cross-linking reagents (such as paraformaldehyde) form intermolecular bridges, normally through free amino groups, thus creating a network of linked antigens. Cross-linkers preserve cell structure better than organic solvents, but may reduce the antigenicity of some cell components, and require the addition of a permeabilization step, to allow access of the antibody to the specimen. Fixation with both methods may denature protein antigens, and for this reason, antibodies prepared against denatured proteins may be more useful for cell staining. Different fixation methods are described. The appropriate fixation method should be chosen according to the relevant application.

1.   Acetone Fixation

• Fix cells in -20°C acetone for 5-10 minutes.
• No permeabilization step needed following acetone fixation.

2.   Methanol Fixation

• Fix cells in -20°C methanol for 5-10 minutes.
• No permeabilization step needed following methanol fixation.

3.   Ethanol Fixation

• Fix cells in cooled 95% ethanol, 5% glacial acetic acid for 5-10 minutes.

4.   Methanol-Acetone Fixation

• Fix in cooled methanol, 10 minutes at –20 °C.
• Remove excess methanol.
• Permeabilize with cooled acetone for 1 minute at –20 °C.

5.   Methanol-Acetone Mix Fixation

• 1:1 methanol and acetone mixture.
• Make the mixture fresh and fix cells at -20 C for 5-10 minutes.

6.   Methanol-Ethanol Mix Fixation

• 1:1 methanol and ethanol mixture.
• Make the mixture fresh and fix cells at -20 C for 5-10 minutes.

7.   Formalin Fixation

• Fix cells in 10% neutral buffered formalin for 5-10 minutes.
• Rinse briefly with PBS.
• Permeabilize with 0.5% Triton X-100 for 10 minutes.

8.   Paraformaldehyde-Triton Fixation

• Fix in 3-4% paraformaldehyde for 10-20 minutes.
• Rinse briefly with PBS.
• Permeabilize with 0.5% Triton X-100 for 10 minutes.

9.   Paraformaldehyde-Methanol Fixation

• Fix in 4% paraformaldehyde for 10-20 minutes.
• Rinse briefly with PBS.
• Permeabilize with cooled methanol for 5-10 minutes at –20 °C.

Antibody Staining

The third step of cell staining involves incubation of cell preparations with antibody. Unbound antibody is removed by washing, and the bound antibody is detected either directly (if the primary antibody is labeled) or, indirectly using a enzyme-labeled or fluorochrome-labeled secondary reagent.

1. Direct Labeling of a Surface Antigen on Unfixed Cells in Suspension

1.1 Add 1 ml of the washed cell suspension to a 15 ml conical centrifuge tube and pellet the cells by centrifuging for 5 min at 200g.

1.2 Decant the supernatant, and add 50-100 ul of primary antibody diluted in PBS (Note: antibody concentration should be determined by titration of the stock solution and testing on a known positive specimen. Usually, working concentrations are in the range of 10-20 ug/ml. However, depending on the source this concentration could vary significantly).

1.3 Gently re-suspend the cells in this small volume by flicking the bottom of the tube with a finger.

1.4 Incubate on ice for 30 minutes.

1.5 Wash the cells three times in PBS, centrifuge for 5 min at 200g, and decanting the supernatant.

1.6 Add 50-100 ul of fluorescence (FITC, Taxes Red) conjugated secondary antibody and incubate on ice for 30 minutes.

1.7 Wash the cells three times in PBS, centrifuge for 5 min at 200g, and decanting the supernatant. After the last centrifugation, re-suspend the cells in two drops of PBS (Note: it is important to keep the cells concentrated at this stage to facilitate examination of an adequate number in the next step).

1.8 Mount the cells on a slide with a coverslip, and examine on the fluorescent microscope with the appropriate filters for fluorescein excitation and emission (Note: antibody molecules will not penetrate the cell membrane of living cells, so any cell showing fluorescence throughout its interior has died. If the cells are not kept cold or maintained with 0.1% sodium azide, capping will occur; antibody-antigen complexes will clustered into one region of the cell membrane resulting in a crescent-shaped fluorescent labeling pattern).

2. Fluorescence Labeling of Surface Antigens of Attached or Suspended Tissue-Culture Cells

2.1 Blocking: Wash cultured cells attached to tissue culture dishes in PBS, then incubate in a blocking buffer consisting of BSA-PBS for 5 min, and cool to 4 C. Cooling prevents subsequent endocytosis of any added antibody reagents, as well as minimizing lateral mobility of bound antibody in the plane of the plasma membrane.

2.2 Primary Antibody: Add the primary antibody to the living cells at 4 C in PBS, and incubate for 30 min. Do not pipet directly onto the cells, but add antibody solutions at the edge of the dish, and add wash solutions from a wide-mouth bottle or beaker to minimize the potential of removing cells by too vigorous a fluid steam. Do not allow the cells to dry at any step. Rock the dish back and forth to maintain coverage over the cells if the antibody solution volume is too small (Note: the most sensitive approach to surface antigen labeling is to incubate cells at 4 C while alive with antibodies. This prevents internal labeling background and also minimizes artifacts seen with some fixation. However, there are situations in which living cells are not available or the antigen of interest fails to react without some form of fixation. In these settings, formaldehyde fixation or other forms of fixation can be performed prior to incubation with antibodies).

2.3 Washing: Pipet out the primary antibody solution from the dish. Wash the dish again 3x5 min with PBS at 4 C.

2.4 Secondary Antibody: Add a fluorescent conjugated secondary antibody in PBS and incubate the cells at 4 C for 30-60 minutes.

2.5 Pipet out the secondary antibody and wash the dish 3x5 min in PBS at 4 C, followed by washing briefly in PBS at room temperature.

2.6 Fixation: Fix the cells attached to the dish in 3.7% formaldehyde-PBS at room temperature for 15 min (Note: the final formaldehyde fixation step links all of the antibody steps in place, preventing their dissociation. Immediate viewing of cells may not require this step, but after fixation, mounted cells can be kept at 4 C for several weeks with little loss of signal. Drying at any stage will severely affect the label, either causing it to b3e displaced or drastically altering morphology of the surface).

2.7 Mounting and Coverslipping: Wash the dish 3x5 min in PBS at room temperature, cover the cells with anti-fade mounting medium, and overlay with a coverslip.

3. Fluorescence Labeling of Intracellular Antigens of Attached or Suspended Tissue-Culture Cells

3.1 Fixation: Cultured cells attached to tissue-culture dishes are washed in PBS, then fixed in 3.7% formaldehyde in PBS for 10 min at room temperature. The dishes are then washed 3x5 min in PBS (Note: coverslips or glass slides can also be placed in the bottom of individual tissue-culture dishes for cell growth when a slide format is desired. Formaldehyde works well as a primary fixative in this setting because it preserves cell morphology well and the time of exposure to the fixative is short. Formaldehyde fixation at room temperature is very effective, but at 4 C, it is very poor fixative. Another fixative approach is the use of organic solvents, such as ethanol, methanol, and acetone. These precipitating fixatives also produce membrane permeability and generally yield a poorer quality of preservation, although with a high degree of permeability).

3.2 Blocking and Permeabilizing: Incubate dishes with blocking solution containing 0.5% Triton X-100 for 30 min at room temperature.

3.3 Primary Antibody: The primary antibody diluted with antibody dilution buffer or PBS is then added to the fixed cells and incubated for 1-2 hours at room temperature (Note: do not pipet directly on to the cells, but add antibody solutions at the edge of the dish, and add wash solutions gently at the edge of the dish to minimize cell detachment.

3.4 Washing: Decant the primary antibody solution, and immediately wash the dish 3x5 min with PBS. Do not let the cells dry at any step. Especially during washing, handle each dish individually, since leaving a washed dish without medium for even a few seconds can allow drying in the center of the dish.

3.5 Secondary Antibody: Add the fluorescent (FITC, Texas Red) conjugated secondary antibody in PBS and incubate for 30-60 minutes at room temperature.

3.6 Decant the secondary antibody and wash the cells 3x5 min in PBS.

3.7 Post-Fix: Fix the cells again using 3.7% formaldehyde freshly made as performed in the initial fixation. The purpose of the second fixation is to crosslink the antibodies in place and to prevent subsequent diffusion of label. If not postfixed in this way, the localization may not be stable for more than a few hours.

3.8 Wash the cells 3x5 min in PBS.

3.9 Mounting and Coverslipping: Mount the cells with anti-fading medium with coverslip.

Observation

1. Examine the cells under the microscope.
2. Record the results and taking pictures of the labeled cells.

Note: It is advisable to run the appropriate negative controls. Negative controls establish background fluorescence and non-specific staining of the primary and secondary antibodies. The ideal negative control reagent is a fluorochrome conjugated mouse monoclonal or myeloma protein. It should be isotype-matched, not specific for cells of the species being studied and of the same concentration as the test antibody. The degree of autofluorescence or negative control reagent fluorescence will vary with the type of cells under study and the sensitivity of the instrument used. For fluorescent analysis of cells with Fc receptors, the use of isotype-matched negative controls is mandatory.