Cytopreparatory Techniques

Wet Fixation

Wet fixation, i.e., submerge the cell sample immediately into the fixative solution. The cell sample remains in the fixative solution until it arrives in the laboratory, where it is accessioned and stained. The cells are not exposed to air at any time during this fixation method. The alcohols – 95% ethanol, isopropanol, and methanol – are all satisfactory fixatives to submit specimens for evaluation.

Wet Fixation with Air Drying

Wet fixation with subsequent air drying, i.e., submerge the cell sample immediately into the fixative solution, remove the sample from the fixative solution after a specified time, air dry the cell sample, and place it into a container for transport to the laboratory. On arrival in the laboratory, the slide is placed in 95% ethanol or its equivalent prior to staining to rehydrate the cell sample.

Spray Fixation

Spray fixation, i.e., immediate fixation of the wet cell sample with a commercially available spray fixative. The spray-fixed cell sample is allowed to air dry and is then placed in a container for transport to the laboratory. Spray fixatives usually consist of an alcohol base blended with a waxy substance (carbowax) that provides a thin protective coating for the cells. When received in the laboratory, the spray-fixed slides must have the waxy substance removed prior to staining. To remove the waxy coating, the slides are placed in two separate rinses of 95% ethanol or two separate rinses of tap water. Slides are placed in the first ethanol rinse for approximately 30 min. One may see opaque, waxy particles float to the surface of the ethanol solution. This first 95% ethanol rinse is discarded. The slides are then placed into the second 95% ethanol rinse for 10–15 min. The use of these two ethanol rinses is usually sufficient to dissolve the waxy substance. In very hot and humid environments, it may be necessary to add a third ethanol rinse. It should be noted that some physicians spray fix more heavily than others, and this factor may add to the difficulty of treating all specimens in the same fashion. Some manufacturers of spray fixatives recommend removal of carbowax with water. We have found 95% ethanol to be more effective and less time-consuming than water rinses. Additionally, after the ethanol rinses, the cells are better prepared for the uptake of dyes than with water rinses alone. However, if availability and cost are factors, water rinses will suffice. It should be noted that if the carbowax is not removed completely, small, waxy particles will be carried into the staining solutions. Nuclei will then appear foggy and lack chromatinic detail and the cytoplasm may exhibit a pale blue color ( Fig. 33-1 ). Alcohol-based, non-aerosol spray fixatives are preferred. One spray container can fix approximately 700–1000 cell samples. It is suggested that one use a commercially prepared fixative designed specifically for cytologic use instead of alcohol-based hairsprays, as has been done in the past. One of the reasons for this recommendation is that companies producing hairsprays may alter their formulas, replacing viable components with substitutes that may alter the quality of cell preservation. When using non-aerosol fixatives, the spray nozzle may sometimes become clogged. It is advisable at the beginning of each day to test the spray emitting from the nozzle to ensure the spray is evenly distributed. Non-aerosol sprays are more likely to produce an uneven spray than aerosol sprays. Non-aerosol spray fixatives may be held closer to the specimen than aerosol sprays. For some products, 3–6 inches is recommended, whereas, for others, 6–10 inches is the recommended distance. One should try to obtain an even spray across the entire slide and avoid spraying the labeled or frosted portion of the slide, where a heavy coating of waxy residue makes it difficult to number or label the slide. If the frosted area is sprayed, it may be necessary to scrape off or otherwise remove the waxy coating on the frosted end of the slide before numbering or applying a label to the slide.

Liquid-based Fixation for Papanicolaou Tests

Liquid-based fixation may be methanol or ethanol based, depending on the manufacturer. Two systems are currently in use, while other systems are still undergoing development. Both of the currently used systems collect cell samples into liquid fixative solutions. Human papillomavirus (HPV) DNA testing, immunocytochemistry, tumor classification, and other special studies ( Chlamydia and Gonorrhea ) can also be performed on these preparations directly from the vial. One liquid-based system which requires no special preparative devices was developed in the late 1990s. This technique utilizes an easily made metastable alcoholic gel. It is compatible with many fixation agents, including CytoRich Red, CytoLyt, and PreservCyt. The specimen is received into the laboratory and centrifuged. The supernatant is poured off and an equal volume of the gel is added. The mixture is vortexed into a cell slurry. The resultant sediment can be either placed onto glass slides and evaluated in its entirety or placed onto a slide using the Hettich cytocentrifuge. Because there is no filtering mechanism, cellular debris, mucus, and inflammation will still be part of the presentation. The resulting slides are comparable with the conventional Papanicolaou smear and other ThinPrep preparations.

Liquid-based technology further resolves the issue of cells being obscured due to the presence of inflammation and blood. Cellular preservation and sampling problems inherent with the conventional Papanicolaou smear are effectively eliminated with liquid-based Papanicolaou testing. This technology entails direct transfer of cells from the collection device into a fixative solution. Diagnostic confidence is enhanced when the liquid-based method is utilized to obtain a representative sample. Currently, two liquid-based preparation systems are in use that account for approximately 80% of all Papanicolaou collection methods in the USA. ThinPrep and SurePath utilize different processing techniques to achieve the same result: eliminate background milieu and provide a well-fixed concentration of representative epithelial cells ( Figs 33-2 , 33-3 ). ThinPrep technology utilizes a filter process, whereas the SurePath system incorporates a gravity sedimentation process. A ThinPrep specimen may be collected with a “broom,” spatula, or endocervical brush and is then rinsed immediately into a vial containing a methanol-based solution (PreservCyt). This eliminates smearing the sample on a slide and ensures that a greater proportion of cellular material will be captured for microscopic evaluation. The vial containing the specimen is placed in the ThinPrep processor. A plastic cylinder containing a polycarbonate filter membrane is placed onto a holder which is then put into the processor. The filter is immersed into the vial. As processing proceeds, the cylinder is spun gently to produce forces that reduce large cell clusters and break up mucus, blood, and background debris. A vacuum draws the specimen through the filter-trapping epithelial cells and organisms, while mucus, blood, and inflammation pass through the filter. When the pressure across the polycarbonate membrane is high, the processor determines that the filter is full. The vacuum is stopped and the filter is inverted and drained of fluid. Reverse pressure is applied to gently “blow” the cell sample off the filter and on to the glass slide. This produces a thin, monolayered circle of epithelial cells (20 mm) that is fixed immediately and ready for staining.

The SurePath system, as mentioned above, incorporates a gravity sedimentation process. After a specimen is collected, the head of the “broom” can be detached and submitted to the cytology laboratory in the collection vial. The vial contains an ethanol-based solution that fixes the sample immediately. Steps in the preparatory process include layering of the cell sample on to a liquid density gradient, vortexing, and centrifugation. During this process, vortexing breaks up large cell aggregates and mucus. Density gradient centrifugation separates useful cellular elements (filtrate) from obscuring inflammation and debris. The filtrate is placed in a chamber and applied to a glass slide by gravity sedimentation. The end result is an evenly layered circle of cells (13 mm) on the slide that may be stained immediately on the SurePath processor ( Table 33-1 ).

Lysing Fixation for Bloody Samples

For conventional gynecologic smears, H.W. Boschann (pers. comm. 1962) recommended placing a drop of acetic acid into the 95% ethanol fixative in a Coplin jar both to fix the cells and to lyse the red blood cells. Some laboratories use Carnoy's fixative, others prefer Clarke's fixative or a modification of either fixative. CytoRich Red and CytoLyt (methanol-based) may be used for lysing bloody non-gynecologic specimens for liquid-based preparations. PreservCyt has been granted Food and Drug Administration (FDA) approval for Chlamydia / Gonorrhea and HPV DNA testing. Blood-lysing protocols vary by liquid-based manufacturer. Operating instructions must be followed meticulously for best cytopreparatory results. Various methods for fixation and treatment of bloody cell samples have been suggested. Some recommend fixing the sample first and then placing the slide into a lysing solution, whereas others place the slide directly into the lysing medium. It is important with either approach to rinse the slide thoroughly in 95% ethanol following the lysing solution and before staining in order to stop the action of the lysing solution. Whichever fixation and lysing method is applied to bloody samples, the solutions should be discarded after each use.

The following solutions may be used to lyse red blood cells:

• Carnoy's (always prepare fresh): absolute ethanol, chloroform, glacial acetic acid (6 : 3 : 1)

• Modified Carnoy's: 95% ethanol, chloroform, glacial acetic acid (7 : 2.5 : 0.5); 95% ethanol, chloroform, glacial acetic acid (6 : 3 : 1); or 95% ethanol, glacial acetic acid (6 : 1). The shelf-life of Carnoy's solution is not known, but it does become hydrochloric acid at some point in time. This is the reason it is suggested that Carnoy's should be mixed fresh each time it is used and discarded after each use. Carnoy's fixative is an excellent nuclear fixative as well as a preservative of glycogen. Because cells may shrink or round up when lysing media are used, it is important to reduce the staining times in hematoxylin and eosin (EA), especially when using Carnoy's fixative

• Clarke's solution: absolute ethanol, glacial acetic acid (3 : 1)

• One drop of concentrated hydrochloric acid per 500 mL of 95% ethanol

• Ten percent glacial acetic acid (this is followed by placing the slide in 95% ethanol)

  • CytoLyt/Glacial acetic acid (9 : 1) – FDA approved for bloody gyn ThinPrep re-preps. (procedure 4)

  • CytoLyt is a commercial liquid medium used as a quasi-fixative and an agent to remove red blood cells from specimen samples. It has the ability to hold over specimens so deleterious effects to the cells will not occur. Cell samples prepared from CytoLyt must be fixed for an additional 20 min in PreservCyt before further processing.

• Commercially available fixatives such as CytoRich Red (supplied by Thermo-Fisher Scientific) may be used (procedure 5)

  • CytoRich Red reduces red blood cells and background and fixes cells completely. Removal of the red blood cells allows the diagnostic cells to be visualized, especially in fine-needle aspirates when a rapid interpretation is required.

  • The solution is said to be a useful fixative for small tissue biopsies and for immunocytochemistry. It works in cytocentrifuged cell samples as well as in liquid-based prepared cell samples.

Progressive Staining Method

In the progressive method, the nuclear chromatin is stained with hematoxylin to the intensity desired. The time the cells are exposed to the hematoxylin is not sensitive because the nuclear uptake is complete after a finite time span. The stain usually has a lower hematoxylin concentration and slowly and selectively stains nuclear chromatin. The blueing agent, following the hematoxylin, sets the nuclear dye in place. The cytoplasm is barely tinted. Lithium carbonate, Scott's tap water substitute (STWS), and some commercially available solutions are the most commonly used blueing agents. Tap water may serve as a blueing agent if the pH is higher than 8. Today, ammonium hydroxide is seldom used as a blueing agent because of its variability in formulations and day-to-day instability.

The progressive method is the most commonly used method in laboratories today. It is easier to control and provides more stability on a day-to-day basis. There are differences among laboratories that use the progressive method ( Table 33-2 ). In the University of Chicago section of cytopathology procedure (prior to 1992), the stains were made from scratch according to the formulas described later in this chapter ( Fig. 33-4 ).

The procedure used by the University of Rochester utilizes hematoxylin #2 from Richard Allan and Orange G (OG) and EA from Baxter Harleco. The Richard Allan clarifier #2 is a reagent useful in eliminating excessive background staining by hematoxylin without removing nuclear stain. It enhances cytoplasmic transparency and should be used only with the progressive staining technique.

The Richard Allan blueing reagent is a buffered blueing agent that enhances nuclear detail by changing the nuclear stain from a reddish blue to a crisp blue-purple.

The third procedure ( Table 33-2 ) is a variation of the Papanicolaou progressive staining method as used by the Johns Hopkins cytopathology laboratory (K. Plowden, pers. comm. 2007).

Johns Hopkins uses Gill #1 hematoxylin, Gill EA, and Gill OG from Harleco.

Gill recommends the following steps when staining manually. These steps are part of the staining procedure used by the Johns Hopkins cytopathology laboratory:

• Follow times and dips posted on the staining schedule.

• A “dip” requires about a second. A dip is gently raising the staining rack until it clears the solution, and without jarring it against the sides or bottom of the staining dish, and lowering the rack until it is totally submerged in the solution. Ten dips effectively replace one solution by another.

• Drain solutions well between baths but do not allow the preparations to dry.

• Change stains (hematoxylin every 3–4 weeks, OG and EA every other week).

• Record changes on the maintenance sheet.

• Filter stains through separatory funnels twice daily (see “ Cross-contamination method to avoid floaters ”, below).

• Filter xylene through course filter paper to eliminate water, cells, and debris.

Regressive Staining Method

Using the regressive method, one deliberately overstains the nucleus with a non-acidified hematoxylin such as Harris's. The solution has a high concentration of hematoxylin and rapidly stains the entire cell. The excess stain is removed with a diluted hydrochloric acid solution. The decolorizing acid is then removed by a bath of running tap water. Timing in the acid bath is essential for the final appearance of the nuclear pattern. More often than not, hypochromasia rather than hyperchromasia is the result. The cytoplasm as well as the nucleus is decolorized by the diluted hydrochloric acid. If the acid bath is inadequate, the contrast between the chromatin and parachromatin is less and uptake of the counterstains is lessened.

A procedure for the regressive staining method once used by the Papanicolaou laboratory (C.M. Street, pers. comm. 1959) is shown in Table 33-3 . The fixative is 95% ethanol, the same fixative recommended for the progressive staining procedure. The second regressive staining procedure is from the Thomas Jefferson University Hospital cytology laboratory. In place of non-acidic Harris hematoxylin, Gill #3 hematoxylin from Surgipath has been substituted. Please note that, prior to staining, the slides are placed in 95% ethanol for at least 20 min.