7 Microtomy
Paraffin and frozen
Introduction
The basic principles of microtomy are applicable to both paraffin and frozen sections. This chapter will discuss the techniques necessary to provide quality microscopic slides for clinical and research histology.
Microtomy
Microtomy is the means by which tissue is sectioned and attached to a surface for further microscopic examination. Most microtomy is performed on paraffin wax embedded tissue blocks. The basic instrument used in microtomy is the microtome; an advancing mechanism moves the object (paraffin block) for a predetermined distance until it is in contact with the cutting tool (knife or blade). The specimen moves vertically past the cutting surface and a tissue section is produced. Good technique is achieved through ongoing practice.
Types of microtome
There are several types of microtome, each designed for a specific purpose, although many have multifunctional roles.
Rotary microtome
The rotary microtome is often referred to as the “Minot” after its inventor. The basic mechanism requires the rotation of a fine advance hand-wheel by 360° degrees, moving the specimen vertically past the cutting surface and returning it to the starting position. The rotary microtome may be manual (completely manipulated by the operator), semi-automated (one motor to advance either the fine or course hand-wheel), or fully automated (two motors that drive both the fine and the course advance hand-wheel). The mechanism for block advancement may be retracting or non-retracting. Its advantages include the ability to cut thin 2–3 µm sections and its easy adaption to all type of tissue (hard, fragile or fatty) sectioning. Technological advances in the automation of microtomy have improved section quality, increased productivity, and improved occupational safety for the technologist. Eliminating manual hand-wheel operation of the microtome reduces the incidence of repetitive motion disorders, a common occupational health problem in the histology laboratory.
Base sledge microtome
With the sledge microtome, the specimen is held stationary and the knife slides across the top of the specimen during sectioning. Used primarily for large blocks, hard tissues, or whole mounts, it is especially useful in neuropathology and ophthalmic pathology. Three micron sections are difficult to produce. Further information regarding section of undecalcifed bone is available in Chapter 16.
Rotary rocking microtome
Commonly used in cryostats, the retracting action moves the tissue block away from the knife on the upstroke, producing a flat face to the tissue block.
Sliding microtome
The knife or blade is stationary and the specimen slides under it during sectioning. This microtome was developed for use with celloidin-embedded tissue blocks.
Microtome knives
There are many shapes, sizes and materials for microtome knives. Knives were developed to fit specific types of microtome, and to cope with different degrees of hardness of tissues and embedding media. Most steel knives have been replaced with disposal blades, although exceptions include the tool-edge knives for resin, and steel knives for some cryostats.
Disposable blades
The introduction of disposable stainless steel blades has revolutionized microtomy in the laboratory. Disposable blades are used for routine microtomy and cryotomy, providing a sharp cutting edge that can produce almost flawless 2–4 µm sections. Disposable blade holders are incorporated into the microtome or an adaptor may be purchased. The blades may be purchased in dispensers, with or without a special polytetrafluoroethylene (PTFE) coating, allowing ribbons to be sectioned with ease, reducing resistance during microtomy:
• The clearance angle should be adjusted in small increments to eliminate problems that occur with the ribboning of the tissue.
• Over-tightening the disposable blade in the clamping device may cause cutting artifacts, such as thick and thin sections.
• The clamping device must be clean and free of defects. During sectioning the hand-wheel must be turned slowly.
• Extremely hard tissues may pose a problem for disposable blades.
Reliability of a constant sharp edge, ease of use, low or high profiles adaptable to a variety of tissue and paraffin types, and low cost relative to steel knife sharpening make these blades a mainstay in most laboratories.
Paraffin section cutting
Equipment required
Flotation (water) bath
A thermostatically controlled water bath is used for floating out tissue ribbons after sectioning. The temperature of the water in the bath should be 10°C below the melting point of the paraffin to be sectioned. Care should be taken to prevent water bubbles from being trapped under the section. This can be accomplished by using distilled water in the bath. Alcohol or a small drop of detergent may be added to the water to reduce the surface tension, allowing the section to flatten out with greater ease.
Drying oven or hot plate
Drying ovens incorporate fans that keep the warm air circulating around the slides. The temperature setting should be approximately that of the melting point of the paraffin. If the oven is too hot there may be distortion to the cells, causing dark pyknotic nuclei or nuclear bubbling; cells that are completely devoid of nuclear detail. Drying times vary depending on the type of tissue, the number of slides to be dried and size of the drying device. Many automated stainers have drying ovens as part of the instrument, so the time and temperature is easily regulated. Special care should be taken when drying delicate tissues or tissues from the central nervous system; a lower temperature is required to prevent splitting and cracking of the section; 37°C for 24 hours is recommended.
Brush and forceps
Forceps, brushes or teasing needles are helpful in the removal of folds, creases and bubbles that may form during the floating out of the section on the water bath. They are also helpful for manipulating the section as it passes across the edge of the blade.
Slides
For normal routine work, 76 × 25 mm slides are universally used. Although slides are available in a variety of thicknesses, those specified as 1.0–1.2 mm in thickness are preferred because they do not break as easily. Most slide racks are made to accommodate this slide size. Larger slides are available for use with specialty tissues such as eyes or brains. Unique identification numbers or codes, patient name or other information should be etched, embossed or written on each slide. Automated instruments that imprint the patient’s information on the glass slide are readily available. Chemical-resistant pens and pencils are routinely used to label the slide.
Slides that are positively charged or pre-treated with an adhesive resist detachment of the tissue from the slide during staining. Colored, frost-ended slides may be used to identify special handling (decal, special stains, immunohistochemistry, etc.).
Section adhesives
Provided clean slides are used and sections are adequately dried, the problem of sections detaching from the slide during staining should not occur. There are occasions when sections may detach from the slide:
• Exposure to strong alkali solutions during staining
• Cryostat sections for immunofluorescence, immunohistochemistry or intra-operative consultation
• Central nervous system (CNS) tissues
• Sections that are submitted to extreme temperatures
• Tissues containing blood and mucous
Adhesives may alleviate the problem of tissue loss. Protein adhesives such as albumen, gelatin and starch may be prone to bacterial growth or heavy staining; close monitoring will prevent these problems. Adhesives that may be used:
Poly-L-lysine (PLL)
Poly-L-lysine is bought as a 0.1% solution, which is further diluted for use, 1 in 10 with distilled water. Slides are coated with the diluted solution and allowed to dry. The effectiveness of the coating to adhere the tissue to the slide will diminish within a few days.
3-aminopropyltriethoxysilane (APES)
Slides are dipped in a 2% solution of APES in acetone, drained, dipped in acetone, and drained again; the process is complete when the slides are dipped in distilled water. Slides are placed upright in a rack to dry. These slides are useful for cytology, especially specimens that may be bloody or contain proteinaceous material.
Charged or plus slides
Laboratories often use slides that have been manufactured with a permanent positive charge. Placing a positive charge on the slides is accomplished by coating the slide with a basic polymer in which a chemical reaction occurs, leaving the amino groups linked by covalent bonds to the silicon atoms of the glass. These slides have proven to be superior in their resistance to cell and tissue loss during staining or pre-treatments such as enzyme and antigen retrieval.
Practical microtomy
The expertise that must be gained to become a competent microtomist cannot be achieved from textbooks. Practical experience under the guidance of a skilled tutor is the best way to gain the confidence and coordination necessary to manipulate the microtome and the sections produced. Techniques will be described, providing information and helpful hints for use during microscopy.
Setup of the microtome
Maintenance of the microtome is important to the production of quality slides for diagnosis. The manufacturer’s recommendation regarding the proper care of the instrument should be closely followed. A departmental policy should be implemented outlining daily, weekly, quarterly and yearly preventive maintenance procedures.
The water bath and the microtome should be ergonomically positioned to reduce stress and tension on the employee’s neck and shoulders. The water bath may be filled with distilled or tap water, and adjusted to the proper temperature of the paraffin. Care should be taken to reduce air bubbles that may distort the tissue section.
The blade should be sharp and defect free. The blade or knife holder should be adjusted to optimize the clearance angle, the distance between the lower facet angle and the surface of the block face. The recommended angle varies from 2–4° for paraffin to 5–7° for frozen sections. The correct angle reduces friction as the blade passes through the block, preventing compression of the section. Determining the exact angle is largely a matter of trial and error. Clamps and screws must be firmly tightened. If a disposable blade is to be used, care should be taken to ensure enough pressure is being exerted on the blade to provide support, but it should not be over-tightened, since this causes thick and thin sectioning.
Sectioning
Trimming the tissue blocks
The paraffin block may be faced or “rough cut” by setting the micrometer at 15–30 µm or by advancing the block using the coarse feed mechanism. Aggressive trimming will cause “moth holes” artifacts. Care must be taken to ensure that the block clamped in the chuck has been retracted so there is no contact with the blade on the initial down-stroke. It is possible to damage the tissue by gouging or scoring when trimming the block.
Cutting sections
Blocks should be arranged in numerical order on an ice tray, cooling both the tissue and the paraffin, giving them a consistent temperature. A small amount of water is absorbed into the tissue, causing slight swelling, and making sectioning easier. Over-soaking may cause expansion and distortion of the tissue section. Proper processing greatly reduces or eliminates the need to pre-soak blocks. Routine surgical material should be cut at 3–4 µm. The micrometer setting does not guarantee that each section will be that exact thickness. Thickness depends on many factors, including temperature, knife angle and cutting speed. Experience will determine the speed of the stroke, but in general, one should use a smooth, slow stroke. If there is difficulty cutting a smooth flat section, warming the block face with warm water, or gently exhaling breath onto the block surface during sectioning, may help. This has the effect of expanding the block, giving a slightly thicker section. Ideally, successive sections will stick edge-to-edge due to local pressure with each stroke, forming a ribbon. If the entire block is to be sectioned and retained, the ribbons are stored in a receptacle for future use. Ribbons of sections are the most convenient way of handling sections. When a ribbon of several sections has been cut, the first section is held by forceps, or teasing needle, and the last section eased from the knife edge with a small brush.
Floating out sections
The floating out of the ribbon must be smooth, with the trailing end of the ribbon making contact with the water first. The slight drag produced when the rest of the ribbon is laid on the water is sufficient to remove most, if not all, of the folds that occur. Sections are floated on the water bath, shiny side down. Folds in the section may be removed by simply teasing with the forceps. Approximately 30 seconds should be long enough for a ribbon to flatten, since prolonged time on the water causes excessive expansion, distorting the tissue. Individual sections or ribbons may be floated onto the slide. Circular structures such as eyes may be difficult to flatten. Various techniques are useful in these situations, such as placing the section on a slide which has been pre-flooded with 50% alcohol. The slide is gently immersed in the water bath and the section of eye will float on the surface. The presence of the alcohol will set up diffusion currents that help to flatten the tissue section. The water bath should be cleaned after each block is cut, removing debris and tissue fragments by dragging tissue paper across the surface. Cleanliness cannot be overemphasized; debris (“pick-up”) material from different blocks is a serious problem.
Drying sections
The small amount of water held under the section will allow further flattening to occur when heat is applied to dry the section. The temperature should be at the melting point of the paraffin. Automated stainers have drying ovens as part of the instrumentation. Slides may be attached to the stainer with individual slide holders or in racks that are designed for the instrument. It is important to eliminate over-heating during the slide drying stage, as cellular details may be compromised. Hot plates may cause localized overheating of the slide. When delicate tissues are to be dried, less distortion will occur if the temperature is reduced and the time prolonged. Overnight drying at 37°C is recommended for many tissues.
Cutting hard tissues
Since the introduction of disposable blades, cutting hard tissues is less problematic. The reason for cutting difficulties is more likely poor fixation or over-processing. Prolonged soaking of the block, or exposing the block surface to running tap water for 30 minutes, may often overcome many of the problems associated with cutting hard tissues. A slight reduction in the knife angle may also yield results. If these remedies fail, softening agents may be used on the surface of the block.
Surface decalcification
When small foci of calcium are present in the tissue section, cutting a quality section may be difficult. The block may be removed from the chuck after rough cutting the tissue and placed face down in a dish that contains a small amount of decalcification solution.
The time for exposure to the decal will vary depending on the tissue; closely monitor the progress of the decal. The block is rinsed well, blotted dry, chilled and returned to the microtome. An immediate section should be taken since the decalcification achieved will be limited. Diagnostic materials may be compromised if over-decalcification occurs. It must be noted that the staining properties of the tissue may be affected after this treatment and allowances must be made to achieve optimum results.
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