Upon completing this chapter, the student should be able to:
List equipment employing temperature control and state proper temperature for each instrument.
Describe types of microscopy and identify indications for their use:
Light (compound microscope)
Polarizing
Phase-contrast
Darkfield
Fluorescence
Electron (scanning and transmission)
State the most common objectives used in light microscopy and the approximate magnification of each.
Describe the method of determining total magnification.
Define:
Clearance angle
Bevel angle
Wedge angle
Resolution
Ocular
Objective
Micrometry
Microtomy
Microscopy
Achromatic
Apochromatic
Binocular
Parfocal
List 3 types of microtomes and identify the use of each.
Describe routine microtome maintenance.
Describe the different types of microtome blades.
Identify the various blade angles and the significance of each.
Identify at least 6 microtomy problems and the appropriate corrective action for each.
Identify at least 4 artifacts that may occur during section flotation.
Describe the mechanisms of closed tissue-processing systems.
Identify critical factors in processing tissue on a short cycle.
Describe the role of vacuum in processing tissue.
Correlate maintenance of all laboratory instruments with quality control and/or quality assurance.
Discuss the principle of pH and the use of pH meter.
Describe how to obtain an accurate measurement when using a balance.
List at least 3 section adhesives and state why the use of an adhesive may be necessary.
Describe 2 applications in which a microwave oven can be useful.
Describe the principle of microwave-generated heat.
Describe the types of automatic stainers used in histopathology.
Describe the relationship of temperature (blade, chamber, room) to cutting frozen sections.
Identify problems encountered when using the cryostat and the appropriate corrective action.
Describe the basic function and benefits of recycling reagents.
Describe micropipette techniques for consistency.
List 4 steps that might decrease tissue carryover in staining baths.
Microscopes
Microscope is crucial for quality control.
Used to monitor quality of routine and special stains.
Slides should be reviewed for processing, sectioning, and staining quality.
Reviewing slides is a good time to learn histologic characteristics of tissues.
A good histology textbook or atlas should be available for review and study.
Pathologist or staff technologist should review slides with the student.
Light Microscope
Magnifying glass (1 lens) is a simple microscope.
Light microscope combines 2 simple microscopes, therefore, it is a compound microscope.
The lens systems consist of objective and ocular (eyepiece) lenses.
Objectives provide image magnification and resolution.
Common objectives:
Scanning lens (x2.5 to ×4 magnification)
Intermediate lens (×10 to ×20 magnification)
High-powered dry lens (×40 to ×45 magnification)
Oil immersion lens (×90 to 100 magnification)
Magnification alone is not the aim of the finest microscopes.
Resolving power is the ability to reveal fine detail or discriminate between adjacent details.
Resolving power is measured as the least distance between 2 objects at which they can be discerned as 2 separate structures.
With the light microscope, objects must be separated by at least 0.2μm to be seen as 2 separate structures.
Microscopic magnification is the apparent ratio of the object as seen through the microscope (virtual image) to the size of the object as it appears to the unaided eye at a distance of 10 inches.
Linear magnification is expressed in units known as diameters (x).
Oculars commonly have a ×10 magnification, but x5 oculars and ×15 oculars are also used.
Total magnification is determined by multiplying the magnifications of the ocular and the objective (e.g., ×10 oculars and a ×45 objective would give a total magnification of x450).
The substage is found below the stage and can be moved up and down.
The substage consists of the condenser and iris diaphragm.
The condenser functions primarily to concentrate light on the tissue section.
The condenser must be centered accurately and focused on the same plane as the tissue section.
The amount of illumination on the section is regulated by the iris diaphragm.
The iris diaphragm should be adjusted so that peripheral light rays are blocked, and the light passing through the tissue should be limited so that it just fills the front lens of the objective.
The iris diaphragm must be readjusted as the objectives are changed.
Micrometry, or microscopic measurement, uses a micrometer scale located in 1 eyepiece and a stage micrometer.
The stage micrometer contains a millimeter scale engraved in 1/10 and 1/100 graduations.
The eyepiece has an engraved arbitrary scale; the value of each division of the eyepiece is calculated for each objective by matching the comparable number of divisions on the stage micrometer.
The separate stage micrometer is no longer needed once this value has been determined.
When white light enters a lens, it is split (refracted) into the colors of the visible light spectrum.
Because colors are refracted at different angles, each has a different point of focus; therefore, an uncorrected lens will give an image surrounded by color fringes.
This is known as chromatic aberration.
Most laboratory microscopes consist of achromatic objectives that are corrected for 2 colors, red and blue.
Apochromatic objectives, corrected for 3 colors and also for other lens aberrations, are more expensive and are not necessary for routine use, but ideally they should be used for photomicrography.
Objectives should be parfocal: all objectives then will have the focal point in the same plane, and magnification, or objectives, can be changed without the need to refocus.
Parfocality is accomplished through a series of adjustment in the microscope, but all oculars must be adjustable for simultaneous observers to have a parfocal view.
Microscope Maintenance
Keep the microscope covered when not in use.
Clean the lenses frequently with lens paper. Do not use other paper tissue.
Remove immersion oil immediately after use.
Use xylene on the objectives only as a last resort, and then use it sparingly and remove it immediately.
Do not dismantle the objectives.
When using immersion oil, be careful that the high-powered dry lens is not dragged through the oil.
Reduce the light to a minimum or turn it off when the microscope is not in use.
Remove the slides from the stage when the microscope is not in use.
Try to always focus upwards, never downwards, especially with the higher power objectives.
Do not touch the surface of the lens with fingers.
Polarizing Microscope
Used for the identification of crystals such as talc, silica, or urates.
Used to make the identification of amyloid stained with Congo red more specific.
Used to examine tissue for substances exhibiting double refraction, anisotropism, and birefringence.
Birefringence: transmitting light unequally in various directions.
Anisotropism: having unlike properties in multiple directions.
Double refraction: a single light ray has been split into 2 rays that emerge from the crystal at different points.
A crystal splits light rays because of its uneven optical density, and the rays are refracted, or bent, to differing degrees.
Polarized illumination is achieved by interposing a polarizing device (polarizer) between the light source and the specimen, and inserting a second polarizing filter (analyzer) between the specimen and the eye.
Natural light vibrates in many planes, whereas light emerging from the polarizer vibrates in only 1 plane.
If the light path of the analyzer is aligned, or parallel, then the light will pass and the field will appear bright.
If the analyzer is rotated so that the optical paths are crossed, the light rays are blocked and the field will appear dark.
When optical paths are crossed, material having the property of anisotropism or birefringence appears bright against this dark background.
A compound, or light, microscope may be converted easily to a polarizing microscope by placing 1 piece of polarizing film on top of the light source (polarizer) and another on top of the microscopic slide (analyzer).
Polarizing discs also may be installed in the microscope by installing the polarizer in the substage filter carrier and mounting the analyzer inside or on top of the eyepiece.
The eyepiece analyzer or the polarizer is then rotated, and the field varies between bright and dark, with any doubly refractive particles appearing bright against a dark background.
Usually the blue filter (if used) should be removed to maximize the birefringence.
Through the polarizing microscope, amyloid stained with Congo red will exhibit an apple green birefringence.
Some normal tissue components, such as collagen, exhibit intrinsic birefringence because of an asymmetric alignment of chemical bonds, ions, or molecules.
Collagen will appear silver with polarization of Congo red stains.
Foreign substances such as talc may also be identified using polarized light.
Talc, or starch, often is introduced into tissue sections from the gloves of surgeons, nurses, or pathologists and it presents the characteristic configuration of a Maltese cross when polarized.
Phase-Contrast Microscope
Used for the examination of unstained specimens, especially unstained living cells, and allows almost transparent objects to be seen clearly.
A standard binocular microscope can be converted to a phase-contrast microscope by replacing the condenser and objectives with special phase equipment, but the performance of converted light microscopes is not equal to that of the specially designed phase-contrast microscope.
This type of microscopy is not used in a routine histopathology laboratory.
Darkfield Microscope
Directly transmitted light is excluded, and only scattered or oblique light is used in darkfield microscopy.
Although most objects examined microscopically are transparent, they also have the property of reflecting light rays.
If oblique light that does not enter the objective is directed on these objects, the objects will appear self-luminous against a dark background.
In darkfield microscopy; objects appear much larger than they are because of their light-scattering properties, and frequently, fine structures are seen much more easily with darkfield than with light microscopy.
The preparation to be examined with the darkfield microscope must be thin and free of extraneous refractive material such as air bubbles, oil, or red blood cells.
This type of microscopy is used primarily for the study of unstained microorganisms and for silver grains in radioactive staining procedures, and is rarely used in routine histopathology.
Fluorescence Microscope
Fluorescence is an optical phenomenon in which light of 1 wavelength is absorbed by a substance and almost instantly reemitted as light of a longer wavelength.
In fluorescence microscopy, a substance is bombarded with short-wavelength light in the ultraviolet (UV), violet, or blue range, and visible light is emitted.
Short-wavelength light absorbed by an atom or molecule boosts the energy level of electrons to a higher orbit; the return of electrons to the original level of excitation results in the loss of energy, which is expressed as visible light.
Mercury or halogen lamps are the usual light source for critical work; an exciter filter placed between the light source and the specimen transmits light of the desired wavelength but obscures all visible light.
A second filter, the barrier filter, is placed in the eyepiece to absorb all UV rays and to allow only visible light rays to pass.
The barrier filter protects the eyes from the damaging effects of UV light and reduces nonspecific fluorescence so that the fluorescent object is seen as a bright object against a dark background.
Some compounds and some tissue components such as collagen fibers fluoresce naturally.
This property of natural fluorescence is known as primary or autofluorescence and may cause problems for the inexperienced microscopist because some tissue components give the same color autofluorescence as the fluorochrome; this is especially true when using blue light.
Substances that are not naturally fluorescent may become fluorescent by interacting with a fluorochrome, ie, a dye that fluoresces if excited by UV light.
The filter combinations must be defined for each fluorochrome so that maximum excitation energy will be provided at the optimal wavelength.
Fluorescein conjugates used in immunofluorescence work are maximally excited in the blue range, ~495 nm.
In immunofluorescence techniques, 1 of the components of the reaction (antibody, antigen, or complement) is labeled with a fluorescent dye such as fluorescein isothiocyanate (FITC).
In the direct immunofluorescence technique, labeled primary antibodies are applied directly to tissue sections to locate and combine with antibodies, complement, or even antigens deposited in tissues such as kidney or skin.
Indirect techniques first apply an unlabeled primary antibody to the section.
This is followed by an FITC-labeled antibody that is raised in a second animal species against the animal species producing the primary antibody.
The second antibody against immunoglobulin G is frequently raised in goats.
Indirect techniques are considered to be more sensitive than the direct techniques.
Complement immunofluorescence techniques also use >1 step.
Acid-fast bacilli and amyloid also can be demonstrated by staining with fluorescent dyes.
Auramine-rhodamine is used for the detection of acid-fast bacilli, thioflavin T for amyloid, and thioflavin S for plaques and tangles.
Immunofluorescent preparations are subject to fading, which can be controlled to some degree by careful selection of the mounting media and of the excitation wavelength; however, slides should be viewed as soon as possible after staining and photographs should be taken for a permanent record.
Thioflavin S dye is stable and allows the slide to be stored; the tissue may be reviewed years later.
Fluorescence microscopy requires special equipment, skill, and training.
Electron Microscope
Objects must be at least 0.2μm apart to be distinguished as separate structures with the ordinary light microscope.
Crystalline structures no more than 0.35nm, or 0.00035μm, apart can be resolved with the electron microscope, but biologic structures are difficult to resolve if they are not separated by at least 2.5nm.
The upper limit of useful magnification with the light microscope is ~1,000 diameters (x); enlargement of photomicrographs above this magnification yields no additional information (empty magnification).
The electron microscope has a magnification range from ~1,000-500,000 diameters, with useful information (resolution) obtained over the entire range.
Resolution is dependent upon the wavelength of the energy source used for illumination.
The electron microscope obtains extra resolving power by replacing the ordinary light source of a light microscope with an electron gun, that is, an electrified tungsten filament that emits electrons.
The extra resolving power results from the fact that an electron beam has a much shorter wavelength than visible light.
The electron gun (electron source), the electron beam, and the specimen are all maintained under a vacuum.
The electron beam is aimed at the specimen and focused by varying the strength of electromagnetic fields, equivalent to the glass lenses of light microscope.
The resulting image is visualized by projection onto a fluorescent screen.
There are 2 types of electron microscopy-transmission and scanning.
In transmission electron microscopy, the specimen (typically a very thin section) either transmits electrons (producing electron-lucent, or clear, areas in the image) or deflects electrons (producing electron-dense, or dark, areas in the image), much as light is either transmitted or blocked by a histologic section.
A 2-dimensional black and white image is seen on the fluorescent screen.
With transmission electron microscopy, one can appreciate not only the relationship between cells but the ultrastructure of the cell itself.
Transmission electron microscopy is very useful in the diagnosis of muscle and kidney disease.
In scanning electron microscopy, a dramatic 3-dimensional image results as the electron beam sweeps the surface of the specimen and releases secondary electrons.
The highest effective magnification with the scanning electron microscope is much less than that of the transmission electron microscope, but 1 of the great merits of the scanning electron microscope is the great depth of focus.
The scanning microscope is used to study surfaces of an object or specimen, and has been used to study cell surface membrane changes in the evolution of malignancy and other pathologic processes.
Electron microscopes are very expensive and require skilled operators with an extensive knowledge of microscopic anatomy.
Microtomes
Rotary, sliding, and clinical freezing microtomes are the types most frequently encountered in histopathology laboratories in the United States.
The ultramicrotome used for cutting 0.5μm plastic sections for light microscopic orientation and 90nm sections for electron microscopy is a retracting microtome.
Today, the rotary microtome is standard in laboratories where routine paraffin and frozen sections are the sole requirements.
Rotary Microtome
Operates with a screw feed or a computerized motor.
The block moves up and down, and either the blade holder or the block advances a preset number of micrometers with each revolution of the wheel.
This type of microtome is found in most cryostats and is the type most commonly used for sectioning glycol methacrylate and paraffin embedded material.
Routine maintenance:
Clean the microtome thoroughly at the end of each day or shift by carefully removing all accumulated paraffin with a soft brush or soft cloth moistened with xylene and then drying the microtome thoroughly, following the manufacturer's instructions for cleaning.
If the model requires, apply microtome oil or grease to all sliding parts as indicated by the manufacturer; lubricating should be done on a routine schedule as recommended by the manufacturer.
Document the service, repair, or routine preventive maintenance performed by someone skilled in microtome maintenance; the microtome should be in good working condition for consistency and operator safety.
Cover the microtome when it is not in use.
Microtome designs offer a broad range of features providing ergonomic benefits.
The wheel can be semi- or completely motorized, giving a smooth wheel rotation.
Sectioning speed and section thickness are easily controlled, and fewer thick-thin or sections with microchatter are obtained.
Some models have an additional device that assists with block alignment, reducing the need for retrimming a previously cut block.
Ensuring minimal tissue loss when recutting a previously sectioned block not only preserves precious tissue but saves time.
These design features improve both working conditions and section quality.
Sliding Microtome
The block is held stationary on the sliding microtome, and the blade is moved along a horizontal plane past the block face.
As the blade is returned to the starting position, it completes each section cycle and a screw feed causes the block to be raised toward the blade at a predetermined thickness.
This type of microtome is used for sectioning celloidin and large paraffin blocks; it is not used in routine histopathology.
The care of the sliding microtome is the same as that of the rotary microtome.
Clinical Freezing Microtome
Has been replaced to a great degree by the cryostat; however, free-floating sections required for some special stains are easier to obtain with this instrument than with the cryostat.
This microtome clamps to the table top and is relatively portable.
A chuck with an attached supply of carbon dioxide allows for the freezing of a tissue section placed in a horizontal plane.
The blade swings out over the chuck containing the frozen tissue specimen, and as the blade is returned to its starting position, the micrometer screw advances the block upward toward the blade.
Sections must be removed from the blade edge and floated in a dish of distilled water.
The sections may be mounted on slides from the distilled water and dried before staining, or they may be stained and then mounted at the end of the procedure.
This method of preparing frozen sections is not good for friable tissue.
Airborne disease transmission is also more likely to occur with this method, because human tissue may contain possible infection hazards that can be spread by the bursts of carbon dioxide used for freezing.
Micrometer Setting
The micrometer setting is very important.
This setting is only approximate and is not an exact determination of section thickness; the actual thickness is influenced by the condition of the microtome and the quality of the blade edge as well as the skill of the microtomist.
Microtomy problems will be discussed in the following section on microtome blades.
The blade is a major component of the microtome and as such will be addressed separately from the microtome.
Microtome Blades
A sharp microtome blade with an edge free of defects is essential for obtaining good sections.
Successful sectioning of poorly processed tissue is often possible with a sharp blade; conversely, nondiagnostic sections may be obtained with a dull blade, even when tissue is optimally processed.
Glass knives are used for cutting sections of plastic embedded material.
Disposable blades are standard in the laboratory.
The blades are made of austenitic stainless or carbon steel with or without various coatings.
There are 2 types of disposable blades, high profile and low profile.
The type of microtome blade holder will dictate which of the 2 types is used.
Because each microtome brand has a different recommendation for settings for the blade's angle, it is suggested that the technician review the manual for settings and guidelines for the blade.
As disposable blades have replaced the older steel knives in most laboratories, these knives will not be discussed.
The disposable blades are held securely and rigidly in a holder designed for this purpose, and they have edges superior to the older knives.
Section quality is greatly improved because of the introduction of these blades; however, it should be noted that not all disposable blades produce sections of equal quality.
Used disposable blades and glass knives should be discarded in a puncture-proof biohazard "sharps" container that is then incinerated because of the safety hazard and potential for contamination with human tissue.
Regardless of the type of material to be sectioned and the type of blade used, section quality is determined by the condition of the edge and the clearance angle more than by any other factors.
The clearance angle in relation to the block face is very important when cutting sections.
This angle routinely is ~3-8 degrees, but the microtome design can dictate the angle.
It is recommended that, upon acquisition of a new microtome, the operational angle be confirmed before use.
Errors in properly establishing the clearance angle can lead to many different sectioning problems.
The rate at which sections are cut influences quality, and undue speed invariably yields sections of poor quality.
Each type of tissue has an optimum cutting speed that is dependent on the nature of the material, the angle and cutting edge of the blade, and the thickness of the section desired.
A general guide for cutting good paraffin sections is that the microtome drive wheel should be rotated -1 revolution per second.
An automated microtome in which the rotation of the wheel is motorized improves section thickness consistency because of controlled speed through the tissue.
For most routine hematoxylin and eosin (H&E) staining, the sections should never be greater than 1 cell layer thick, and should be cut at 3-4 pm settings on the micrometer scale.
The thickness of a section can be determined microscopically by focusing up and down through the section.
If the section is 3-5 μm in thickness, then all nuclei will be in 1 plane of focus; if some nuclei go out of focus as others come into focus, then the section is too thick.
Tissues such as bone marrow and kidney biopsy specimens are routinely cut thinner (2-3 μm), allowing for better nuclear detail.
If the microtome is properly set, and thick sections are being produced, the microtome may require service.
Troubleshooting Microtomy
Many microtomy problems associated with sectioning paraffin, frozen, or celloidin tissue blocks are similar.
With each of the aforementioned techniques, irregular, skipped, or excessively thick and thin sections are usually the result of either too little or too much blade tilt (clearance angle).
The problems of irregular, skipped, or thick and thin sections usually can be corrected by adjusting the blade so that the clearance angle between blade and specimen is correct.
Grooved, scored, smeared, and deformed sections are frequently produced by a dull edge, and moving the existing blade to an unused area or replacing with a new blade will usually correct this microtomy problem.
Regular lengthwise or vertical scratches and splits in the sections usually are caused by a defect in the edge, calcium, bone, or another hard material present in the specimen also can cause this artifact.
If the defect remains in exactly the same area of the new sections, then the problem is in the specimen and not the blade edge.
Mushy sections result from insufficient dehydration or clearing.
While backing up these tissues to the dehydrating agent and then reprocessing may help, the sections rarely will be as good as those from an originally well processed tissue sample and additional testing such as molecular studies or immunohistochemistry results may be compromised.
Crooked Ribbons
Crooked ribbons result when the horizontal edges (top & bottom) of the block are not parallel.
They may also be caused if the lower block edge is not parallel to the blade edge when sectioning.
Crooked ribbons occur when the block is not evenly chilled or the hardness of the paraffin varies from 1 side of the block to the other.
Imperfections in the blade edge should also be considered as a cause.
This artifact can be prevented by:
ensuring that the upper and lower block edges are parallel
ensuring that the lower block edge is parallel to the cutting edge of the blade
ensuring that there are no problems with the blade edge
ensuring that the block is evenly chilled
Block Face Unevenly Sectioned
When the block holder is adjusted so that it is not parallel to the blade, 1 side of the block is exhausted while attempting to get a complete section of the block face.
This results in uneven sectioning of the block face.
When this occurs, it wastes tissue and may cause the loss of important areas of tissue.
Uneven sectioning can be prevented by ensuring at the beginning of sectioning that the block holder is adjusted so that the block face and the blade are perfectly parallel.
The use of microtome calibration tools to align the microtome chuck with the blade provides a quick and standardized method to avoid unevenly faced sections.
In addition, it is a good practice to align the block to the blade holder before each day or shift for microtomes that are heavily used.
Holes in the Section
Holes occur when a block is faced too aggressively.
Small flecks of tissue are removed from the block, leaving a hole; liver, brain, and lymph nodes are especially prone to this artifact.
Holes seen in sections that are a microtomy artifact will decrease in size as successive ribbons are cut and will finally disappear.
Brown [2009] reports that tissue can become excessively dehydrated and brittle during processing because of extended time in alcohol or dehydrating steps.
Brittle tissue, if not adequately rehydrated before sectioning, may tear or split, creating holes
Holes may appear in the ribbon if all of the air is not displaced from the tissue during infiltration.
This occurs most frequently with lung tissue; it is not a sectioning artifact, and these holes will not disappear with continued ribboning.
Holes in sections may be corrected or prevented by:
facing the block to expose the tissue, followed by soaking the block briefly in ice water or with a wet piece of cotton before sectioning
facing the block less aggressively, with smaller micrometer advances of the block for each section removed
facing large autopsy brain sections at 5 μm intervals using a smooth rotation until the tissue is fully exposed
knowing the type of tissue being sectioned and making modifications as necessary
if there is sufficient tissue in the block, cutting and discarding ribbons until the holes disappear; but this technique will potentially lose valuable tissue or the area of interest
Failure Of Ribbon to Form
The failure to obtain a ribbon is most commonly caused by a dull blade, but also may result from paraffin that is too sticky (not enough plastic) or too hard (too high a melting point); too much blade tilt; or a room temperature that is too high or too low.
The formation of a ribbon depends on enough heat being generated by the friction occurring as each section is cut to cause the sections to adhere to each other.
If the blade is too cold, it is difficult to generate enough heat for section adherence.
This problem may be prevented or corrected by:
choosing a paraffin with a lower melting point
decreasing the tilt of the blade (smaller clearance angle)
changing the room temperature
Lifting of the Section from the Blade as the Block is Raised
Lifting of sections from the blade frequently is caused by a dull blade or by too little blade tilt.
A warm room or paraffin that is too soft can also cause this artifact.
This may be prevented or corrected by:
increasing the tilt of the blade (greater clearance angle)
changing to a paraffin that is harder (slightly higher melting point)
If a ribbon cannot be obtained after checking and correcting for any identified problems, single sections usually can be picked up by touching the top or bottom edge of the section with a wet applicator stick held with the long surface of the stick parallel to the section.
Forceps may also be used to pick up the section.
Water Trapped Underneath Tissue During Section Pick-Up Onto Slide
The technique used to pick up a tissue section that has been laid out onto the flotation bath is important.
The objective is to reduce the amount of water collected between the tissue and slide when picking it up.
When using positive charged slides, a technique or motion referred to as pulling is preferred over scooping.
The technique of pulling is achieved when a clean slide is placed into the flotation bath just below the surface, positioned at an angle (~30 degrees).
Once the tissue section makes initial contact with the slide and is securely anchored, the tech performs a pulling motion that pushes the water out as the section is being picked up.
The opposite motion of scooping positions the slide under the flotation bath surface in a horizontal position and the tech picks up the tissue by raising the slide parallel with the tissue.
Scooping is undesirable because it traps water under the section and typically requires additional manipulation or wicking to release the water by breaking the paraffin edge.
If the water is not adequately removed from between the section and slide, artifacts of tissue folds, holes, washing, or uneven staining may result.
When using automated staining platforms that conduct the drying step onboard and/or stain the slide in a horizontal position, it is imperative that the microtomist use the pulling technique.
The pulling technique will reduce residual water presence and promote better drying and tissue slide adhesion.
Washboarding or Undulations in the Section
This artifact most commonly occurs in very hard tissue such as uterus, or in overfixed tissue.
It is a macroscopic type of chatter easily seen when the sections are on top of the water bath.
It may occur from worn microtome parts that allow too much tolerance in some of the moving parts, and from loose clamping of the blade or block.
This may be prevented by:
ensuring that the paraffin is filled on the top of the cassette to provide support for tissue when clamped in the block holder
ensuring that both the block and blade are tightly clamped in the microtome. A block or blade that is very loosely clamped can cause the blade to chop into the block, resulting in loss of some of the section or pieces of tissue
ensuring that the block holder shaft is not overextended. Microtome models in which the blade is stationary and the block advances or retracts can create washboarding when the shaft is overextended
ensuring that the microtome is in good working order and has routinely scheduled maintenance
decreasing the blade tilt (smaller clearance angle)
Chatter, or Microscopic Vibration, in the Section
Luna [1988] relates this artifact primarily to overdehydration or a lack of moisture in the tissue.
Microscopic chatter also can be caused by a dull blade; by too much blade tilt, which causes the section to be scraped rather than cut; and by cutting too rapidly.
This may be prevented or corrected by:
ensuring that the processing schedule does not cause overdehydration of tissues; process different tissues types on different schedules. Best practice is to process biopsy tissue separate from large or fatty tissue types.
restoring moisture to tissue by facing the block and soaking it briefly in ice water, or placing it face down on an ice tray. A wet laboratory grade tissue paper (Kimwipe, Kimberly-Clark Corp, Roswell, GA) covering the thumb can also be used to rub the block face with water from the water bath; this slightly warms the block face while rehydrating the tissue. Do not put an ungloved thumb into water bath as it will result in squamous cell contaminant.
decreasing the tilt of the blade (smaller clearance angle)
decreasing cutting speed; 1 revolution of the wheel per second is considered a reasonable cutting speed
Skipped or Varied Thickness of Sections (Thick & Thin Sections)
Thick and thin sections may be caused by too little blade tilt, so that the bottom, rather than the top, of the blade facet contacts the block.
Compression of the block results, but with continued advance the blade edge, does finally contact the block and a section is cut.
Alternating compression and sectioning of the block yields skipped and thick sections.
Alternating thick and thin sections can also be caused by loose or worn microtome parts.
This may be prevented or corrected by:
increasing the tilt of the blade (greater clearance angle)
ensuring that the microtome is in good working order and that routine maintenance is scheduled and documented
Compressed, Wrinkled, or Jammed Sections
A dull blade, a blade gummed with paraffin, paraffin sticking in the back side of the holder, too-little blade tilt, too-rapid cutting, or a too-warm room will all cause this microtomy artifact.
These problems can be prevented by:
keeping paraffin from building up on the blade back; the edge should be kept free of paraffin by wiping (up, never down) with gauze slightly dampened with xylene
ensuring that a sharp blade is in use; changing if necessary
increasing the tilt of the blade (greater clearance angle)
decreasing cutting speed; 1 revolution of the wheel per second is considered a reasonable cutting speed