M.P. McLeod • S. Choudhary • K. França Department of Dermatology and Cutaneous Surgery,

University of Miami Leonard M. Miller School of Medicine, Miami, FL, USA

K. Nouri (*)

Department of Dermatology and Cutaneous Surgery,

University of Miami Leonard M. Miller School of Medicine,

Miami, FL, USA

Sylvester Comprehensive Cancer Center,

University of Miami Hospital and Clinics, Miami, FL, USA e-mail: knouri@med.miami.edu

Summary: Tissue Transport

•The major goal of transporting tissue from the surgical site to the histology processing center is to maintain specimen orientation.

•The 12 o’clock position of the specimen should be oriented to line up with the 12 o’clock position of the transferring device.

•The specimen should be transferred with toothed forceps, and care must be taken to avoid damaging or dropping the specimen.

•After transferring the specimen and sufficient hemostasis is achieved, the Mohs’ map should be drawn.

•A standard system should be devised and strictly followed for designating specific details of a map.

12.1Tissue Transport

The major goal of transporting tissue from the surgical site to the histology processing center is to maintain specimen orientation [1]. There is ample opportunity for orientation error, and every precaution should be made to safeguard against it. A transferring device such as an index card or gauze on a plate with the 12 o’clock position marked is very useful for maintaining orientation and preventing specimen dropping [1]. Guards around the plate can prevent unexpected movement of air from changing the specimen’s orientation. The device should be held closely to the wound when transferring the specimen from the surgical site to the plate in order to avoid dropping the specimen or placing it in the incorrect orientation. The 12 o’clock position of the specimen should be oriented to line up with the 12 o’clock position of the transferring device. The specimen should be transferred with toothed forceps, and care must be taken to avoid damaging or dropping the specimen.

After transferring the specimen and sufficient hemostasis is achieved, the Mohs’ map should be drawn. Care should be taken to draw the precise shape of the specimen. It should be at least the same size of the specimen if not larger so that it can demonstrate important details of subsequent stages. A standard system should be devised and strictly followed for designating specific details of a map. For example, short radiating lines are commonly used to denote borders with epithelial margins. Wavy lines can be used to denote nonepithelized margins. Subsequent stages should also be drawn with reference to the previous stage so that a “pictorial story” can be followed until the end of the procedure [1].

Summary: Initial Processing

•The specimen should be divided into as few sections as possible because more sections can lead to orientation confusion.

•Tissue scores or relaxing incisions can also be implemented that allow the specimen to flatten for placement onto the microscope slide.

•Inks can also be very useful in keeping proper specimen orientation throughout histologic processing.

•The cryostat uses a refrigerated microtome to cut histological sections of tissue that are placed onto slides.

•The specimen is placed upon a microtome chuck, and optimum cutting temperature (OCT) is used to fix the tissue to the chuck.

•The slides should be labeled with the patient’s medical record number, stage of surgery, and procedure date prior to sectioning the specimen with the microtome to avoid confusion and ease the work flow.

•Staining of the slides must occur so that colorless cellular material can be visualized underneath the microscope.

•The most commonly used stain for Mohs micrographic surgery is hematoxylin and eosin (H&E).

•Following transport and initial processing of the slides, the specimen is now ready to be analyzed by the Mohs micrographic surgeon

12.2Initial Processing

The Mohs map and transferring device with specimen are then carefully transported to the histological processing center. If the specimen is too large to fit on the slides, it must be divided. The specimen should be divided into as few sections as possible because more sections can lead to orientation confusion [1]. There are a few different ways to divide the specimen. Depending on the specimen shape, it can be divided in half so that there is a top and bottom or left and right. If the specimen is very large, it can be divided into numerous sections. Tissue scores or relaxing incisions can also be implemented that allow for the specimen to flatten for placement onto the microscope slide. These incisions should be superficial enough to allow the tissue to flatten but should not extend deep enough to distort the histology [1].

Inks can also be very useful in keeping proper specimen orientation throughout histologic processing. The color coding scheme should be decided upon and strictly followed as any deviation could lead to orientation error. Colors that are clearly distinct from one another should be used in order to avoid confusion. Sections should be inked one at a time and placed back into the transfer device to avoid orientation error. In addition, the ink should be placed approximately 1–2 mm underneath the surface of each section [1].

The cryostat uses a refrigerated microtome to cut histological sections of tissue that are placed onto

slides. The specimen is placed upon a microtome chuck, and optimum cutting temperature (OCT), a type of tissue glue, is used to fix the tissue specimen to the chuck. There are a number of methods used to freeze the specimen. Each cryostat is equipped with a small workstation (the freeze bar) that is approximately 20–30°C below the temperature of the rest of the cryostat [1]. This lower temperature results in an area where tissue can be quickly frozen.

The basic method of tissue freezing and fixing to the chuck involves placing the chuck directly on the freeze bar and placing OCT on its surface. As soon as the OCT begins to turn opaque, the specimen is placed with the deep margin facing upwards and the superior margin directly touching the OCT. A heat extractor can be used to flatten the tissue as it lies upon the OCT so that the peripheral margins come into the same plane as the deep margins. Additional OCT is then placed over top of the specimen after it is frozen [1].

When processing very small specimens, a technique that uses a glass slide may be helpful. The specimen is placed with the superior margin touching the glass slide surface. The lateral edges are then gently pushed down on the slide so that they are in plane with the deep margin. OCT is then placed on top of the specimen with the slide on the freeze bar. The microtome chuck is also placed on the freeze bar, and OCT is placed on it as well. While the OCT on the microtome is in a solid state, the glass slide is inverted and placed on top of the chuck containing the OCT. The glass slide containing the specimen is kept on top of the OCT on the microtome until it freezes. After the OCT freezes, a small amount of heat is placed on top of the glass slide with a finger, and the glass slide is gently removed from the frozen OCT. Additional OCT is then placed over the specimen. Upon freezing of the OCT, the specimen is now ready to be positioned for cutting with the microtome [1].

Another technique is to directly place the specimen with its deep margin touching the freeze bar. Again, the lateral margins should be gently pushed downward to maintain the same plane. Next, OCT is poured directly over the specimen. The OCT covering the specimen should be allowed to freeze. The chuck is again placed on the freeze board, and OCT is poured onto it until it reaches a semisolid state at which time the specimen, frozen in OCT, should be inverted and placed in the semisolid OCT on the microtome chuck. Additional OCT is then added to the specimen on the chuck as required [1].

The specimen can also be prepared using a cryomold in combination with the microtome chuck. The specimen is placed with its superior margin against the surface of the chuck. OCT is poured into the mold on top of the specimen. After that, the mold along with the chuck is placed on the freeze bar and allowed to freeze making sure that the specimen does not change orientation. Once the specimen is frozen in place, additional OCT is poured into the cryomold so that the specimen is covered. When the entire specimen is frozen in OCT, the cryomold can be peeled away leaving the specimen for sectioning [1].

The slides should be labeled with the patient’s medical record number, stage of surgery, and procedure date prior to sectioning the specimen with the microtome to avoid confusion and ease the work flow. Electrostatically charged slides are highly recommended as they encourage the specimen to adhere to the slide when being cut with the microtome. In addition, different slide colors can be used to demarcate particular Mohs stages.

Once the specimen is prepared on the chuck and properly placed on the microtome, sectioning the tissue may commence. Whenever possible, the epidermis should be placed perpendicular to the microtome blade in order to prevent tissue folding. The cryostat temperature should be set at −20°C to −29°C [1]. A temperature that is too low can lead to specimen shattering when the microtome blade cuts a section. On the other hand, when the temperature is too high, the tissue can “aggregate” together instead of creating a thin line of OCT and specimen on the microtome stage. Sometimes, the digital temperature reported on the cryostat is not accurate. In order to ensure that the microtome blade and stage are at the desired temperature, a thermometer can be placed in close approximation to these structures and compared to the digital display. In addition to the temperature, the microtome blade should be monitored for dullness and buildup of OCT and/or specimen [1].

When first beginning to rotate the microtome blade, the outer layers of OCT have to be cut away until reaching the actual specimen. Upon reaching the specimen, it may require realignment to achieve as flat a specimen as possible to create uniform slides. The sections should be approximately 5–10 mm thick. At this point, the anti-roll plate should be engaged, and the slides should be appropriately labeled and ready for “picking up” the specimen from the microtome stage. The slides should be kept at room temperature, so that

when the slide is gently placed upon the microtome stage, the cold freshly cut specimen quickly adheres to the slide. The specimen should be consistently placed on the slides in the same manner so that orientation confusion is not introduced [1].

Staining of the slides must occur so that colorless cellular material can be visualized underneath the microscope. The most commonly used stain for Mohs micrographic surgery is hematoxylin and eosin (H&E) [1]. Newer stains including immunohistochemical stains are discussed in Chapter 15. Hematoxylin is derived from the logwood tree. When combined with a metal salt, hematoxylin becomes a cation known as hematein. Since hematein has a positive charge, it can bind to negatively charged molecules. In the case of molecular structures, this is most commonly RNA and DNA, which are negatively charged. On the other hand, eosin is a negatively charged molecule and stains other molecules such as proteins that have a positive charge. Hematoxylin stains structures blue, while eosin stains the structures that it binds pink. Slide staining can occur manually or using an automated device. The automated slide staining technique has developed widespread use because it uses fewer reagents and does not demand as much time from the histotechnician (Figure 12.1). For example, in the manual method, the histotechnician must continually change the waterrinse areas; however, the automated staining device continually fills these areas with fresh water [1].

There are many different protocols available for staining slides with H&E. It is important for a Mohs histology processing center to use a protocol that yields the highest quality slides. Protocols can be modified based upon experience and preferences.

First, the tissue can be fixed. If using frozen sections, fixation is not necessary, but 10% neutral buffered formalin can be used. Next, water can be used to hydrate the slides. It is recommended to use distilled water as regular tap water can exhibit different alkalinities. Next, the dye is added. Hematoxylin is most commonly used first as a regressive stain. Following the regressive stain, the “bluing” step uses ammonia or alkaline salts to enhance the nuclear structures stained by the hematoxylin. Eosin is used to counterstain the specimen and dye the cytoplasmic proteins different shades of pink. The last step is dehydration and clearing. This step uses 95% alcohol and is important for determining how much pink to red color remains on the slide. The alcohol removes the excess water from the slide. All the water must be removed before the clearing step is used to improve the transparency of slides. Traditionally, xylene has been used as the predominant clearing agent; however, it has been found to be toxic. Less toxic substitutes have been developed and now include aliphatic hydrocarbons and essential oils [1].

The last step in the preparation of standard H&E slides is applying the coverslip. When removing the