Laura Brandt, DVM, MS, DACVP (Clinical Pathology)
Ethos Diagnostic Science, Regional Clinical Laboratory Director – Denver & Clinical Pathologist
Posted on 2019-02-07 in Clinical pathology
Cytological examination of fine needle aspirate samples, impression smears and cavitary effusions can provide vital information that can often help make or break a case. Cytology is a popular diagnostic tool because it is typically less invasive than surgical biopsy. Additionally, cytological assessment is less expensive and can provide more rapid results than histopathology. With appropriate case selection, good sample acquisition, and correct sample preparation cytology often provides a definitive diagnosis or can narrow down the differential diagnoses.
Cytology can simultaneously be a frustrating endeavor for clinicians, pet owners and pathologists alike, however. An inconclusive diagnosis can occur when case selection is not ideal for cytology or sampling and preparation techniques are not ideal. Major pitfalls for procuring and preparing good cytological preparations are geographic miss of the lesion, poor exfoliation or cells (either due to technique or lesion type), excessive hemodilution, excessive thickness of the sample, cellular rupture during processing, damage/destruction of the sample after collection/preparation. Cytological examination is also inherently limited by the fact that tissue architecture cannot be examined. Even when limited by lack of tissue architecture, cytology can still be a useful screening tool that can complement histological examination of tissues.
Case selection and successful sample procurement are the first step in obtaining a good quality, diagnostic cytology specimen. Cytology may be performed on cavity effusion, fluid from mass lesions, swabs from draining tracts and ears, fine needle aspirate biopsy of mass lesions and organs, airway washes, impression smears of mass lesions and ulcerated surfaces, bone marrow and cerebral spinal fluid (CSF). Any sample that can be smeared onto a slide and stained can be examined under the microscope, however the clinician expectations should be tempered according to the sample. For example, nasal flushes and cytology of nasal mucus often reveal inflammatory cells and bacterial organisms but will only very rarely contain tumor cells or fungal organisms, even when neoplasia or fungal infection are the primary underlying disease.
Mass lesions smaller than 0.5 cm (5 mm) in diameter may be too small to seat the tip of a needle deep enough into the lesion to cover the needle bevel, making it difficult to engage suction. Fine needle aspiration biopsy (FNAB) may still be attempted on these very small lesions but be prepared for a potentially non-diagnostic sample.
Be certain that the plan for sample acquisition is the most appropriate for the site. It is tempting to simply make impression smears of ulcerated lesions, but these preparations typically only procure superficial contents – keratin, normal or reactive epithelial cells, inflammatory cells, possibly microorganisms (e.g. bacteria), which are often indicative of a secondary issue. If a palpable mass is associated with ulcerated lesions be sure to obtain an FNAB of the actual underlying mass. Fine needle aspiration bypasses the surface contaminants and can provide the pathologist with a purer sampling of the primary lesion.
When performing an FNAB the center of the mass is a good place to sample, however, be aware that necrosis can be localized centrally, especially in large masses. For masses larger than 2 cm try to get aspirations from both the center and the periphery of the mass. Ultrasound guidance can also help isolate lesions internally and peripherally. If a hypoechogenic center of a mass is detected on ultrasound, this would be an indication to sample the periphery of the lesion, as well as the center. If you do use ultrasound gel while evaluating masses or internal organs be but certain to fully wipe off the gel prior to FNAB. All lubricant-type gels (lidocaine gel, lubricant, ultrasound gel) preferentially absorb Romanowsky stains, preventing the cells in the sample from staining properly (no matter how many times the slides are run through the stain). Cytology samples can be rendered completely non-diagnostic when contaminated with ultrasound gel.
Fine needle aspiration biopsy and spreading of the procured sample is a technique that takes some time to perfect. Early attempts may result in minimally useful information, but do not be deterred. Mastering the technique is worth your time in the long run. Do not use needles larger than 20-gauges to avoid excessive blood contamination. A 22-gauge needle is usually the ideal size, with the length of the needle varying according to the accessibility of the lesion.
To perform a traditional FNAB attach the syringe size of your choice to your 22-gauge needle. Typically, a 3 cc syringe is the perfect choice for soft, peripheral masses and enlarged lymph nodes. The larger the syringe the more powerful suction you will apply to the lesion. To begin the procedure, first pull on the plunger of the syringe to break the seal, then express all air out of the syringe. Insert the needle into the region of the lesion you wish to sample and apply negative pressure by drawing back on the plunger (about ¾ of the total volume of the syringe). Repeat this ~2-3 times. You may opt to redirect within the mass during the aspiration procedure, being careful to keep the tip of the needle within the mass. Release the plunger while keeping the tip of the needle seated within the mass. If the tip of the needle comes out of the mass during the aspiration procedure, the sample will be sucked into the syringe and will be difficult, if not impossible, to expel from the syringe (do over!). Once the negative pressure is released, withdraw the needle with attached syringe from the lesion. Detach the needle and fill the syringe with air. Re-attach the needle, point the needle bevel down and depress the syringe plunger, expressing all the air out in one swift, fluid motion, depositing the sample onto a glass slide.
Alternatively, you may try a non-aspiration technique, also referred to as a coring technique, in which the needle is held by the hub without an attached syringe and inserted and removed from the lesion multiple times using a quick, “stabbing” motion (see image at top of page). As with the traditional technique, the same pathway can be sampled, or one can opt to redirect during sampling to obtain cells from various portions of the lesion. This technique should be especially utilized for vascular lesions as the coring technique can result in less blood contamination than the suction from traditional FNAB. You may also consider a smaller gauge needle if you know a lesion is particularly vascular (e.g. splenic mass), but, once again, the versatile 22-gauge needle will often be a good choice. If too small a needle is selected, it may be difficult to obtain cells.
The next pitfall for the practioner is correctly preparing the sample. It is important to spread the sample into a monolayer, so cells can be separated from blood and properly assessed microscopically. Care must be taken, however, not to rupture the cells by applying too much pressure to the spreader slide. This latter concern often results in the cautious practioner expelling the FNAB contents onto a glass slide without spreading the sample. This well-meaning choice results in a “splatter prep,” which is often about as useful as splattered roadkill. Cells remain aggregated in regions of thick blood and appear rounded up and contracted, making morphological assessment (and a diagnosis) difficult. All samples applied to a glass slide should be appropriately spread for the best chance at a diagnosis.
To properly spread a cytology sample, apply the sample to one end of the slide (preferably close to the frosted edge of your slide). Then, lay another glass slide on top of the sample. Let the weight of the glass slide spread the sample. Do not apply extra pressure – this could rupture cells. Pull the two slides away from each other in the same horizontal plane until the slides are no longer touching. Do not pull the top slide up or the bottom slide down prematurely, as this will also cause cellular rupture.
Once cytology slides have been prepared, dry them quickly to air fix them. Prolonged drying can cause contracture of the cells, making it difficult to differentiated nucleus from cytoplasm and to assess the finer nuances of cell morphology. Most well-spread samples will dry quickly on their own or with some gentle waving back and forth in the air. Thicker preparations, including joint fluid and mucoid-type samples should be manually dried with a fan or hair dryer set on cool to avoid slow drying artifact. Slides should be completely dry before enclosing them in plastic slide cases as this can create an artifactually humid environment in which cells will rot and rupture on the glass slide en route to the lab. Do not heat fix slides – this is unnecessary and may damage cells.
Increase the odds of success by taking multiple aspirates from the same lesion. In truth it only takes one good quality slide to make the diagnosis, but it may take prepping 3-4 slides to produce one of diagnostic quality. Veterinarians have the added complication when making cytology samples of being unsure if they have procured the necessary sample prior to submission. This is in contrast to tissue biopsy and submission of formalin fixed tissues, in which case the veterinarian is certain they have given the pathologist a piece of affected tissue. It is completely appropriate to select 1-2 of one’s cytology slides to stain in-hospital and do a quick examination for cellularity. If nucleated cells are not present or only low numbers of cells are seen, consider obtaining more aspirates. Pathologists have wonderful, high-quality microscopes, but no microscope can make cells appear from nothing. Always send any inhospital stained slides with the unstained, air-fixed slides for the most complete assessment of lesions.
Impression smears are another method of producing cytology samples. These can be made from ulcerated external lesions or from biopsied tissue. As mentioned above, impression smears of ulcerated lesions tend to provide information about the surface of the lesion, so be prepared to concurrently perform an FNAB or tissue biopsy for a definitive diagnosis. Additionally, after taking your initial impression smears of the ulcer, you may gently clean the top layer of the ulcer off and make impression smears of clean, deeper, now exposed tissue and/or use a scalpel blade to scrape off cellular material that can be spread onto a glass slide. These preparations tend to be very bloody and can be difficult to interpret, however.
Impression smears from biopsied tissue should be made from freshly cut surfaces. Gently blot the tissue on paper towel until no blood is left behind. Once this has been achieved, press the tissue firmly against a glass slide and lift directly up. This may be repeated several times on clean parts of the same glass slide. The impression left behind should look similar to a lip print on a drinking glass. If your impression smear looks like a bloody version of this, go back to blotting until a blood-free, filmy appearance is achieved on your impression smear. Smearing or dragging the fresh cut surface along the glass slide will not produce good results.
Swab cytology specimens are made from samples collected from mucus membranes (vagina), ears and material expressed from fistulous tracks. The surface being sampled must be moist in order for sample to cling to the cotton tip of the swab. Simply insert the sterile swab into the area/lesion to be sampled and gently wipe the cotton tip along the moist surface. Remove the swab and gently roll the cotton tip of the swab along the surface of the slide. Do not drag or smear sample on the glass surface. If the material on the slide is thick and wet, assist the drying process with a fan or hair dryer set on cool. Fluid cytology can be performed on cavitary effusions, airway washes, synovial fluid, fluid from mass lesions and urine. Fluid cytology, especially of cavitary effusions, can be essential to a complete work up and can often produce a definitive diagnosis. For all fluids submitted to the laboratory, submit the fluid in an EDTA tube. In addition to preventing clot formation in your fluid sample, EDTA is also an excellent cell preservative. If culture of the fluid is anticipated, concurrent submission of an aliquot of fluid in a non-additive red top tube (RTT) or other sterile container is recommended. EDTA is bacteriostatic and can impact culture results. Lastly, if you are unable to immediately submit your fluid sample to a diagnostic laboratory, make a few direct (non-concentrated) preparations of the fluid on glass slides, dry the slides and submit these along with the EDTA tube of sample. Cells in fluid will degrade over time, even in EDTA, and it can be helpful for the pathologist to have a baseline example of how cells looked immediately upon collection. If flocculent material or mucus goobers are procured from airway washes, making squash preparations or impression smears with this material can also be useful.
Cerebral spinal fluid is a special fluid that is handled and processed in a unique way. Unless the fluid is grossly bloody, requiring anticoagulant, CSF is typically collected into a RTT. The volumes, cell count and protein content are often so low, that EDTA can dilute the sample may alter cell counts. The low-protein environment of CSF in vitro can lead to rapid degradation of cells. There are many protocols available to allow for addition of patient serum, hetastarch or even formalin to aliquots of CSF for cell preservation during transit to the laboratory. Studies have shown, however, that even in very low protein samples cells can be reliably identified and classified by pathologist 24 hours after sample collection. Still, it is recommended to process CSF as soon as possible after sampling. Sample processing is best performed by diagnostic laboratory personnel.
The key components need for best pathologist interpretation are good sample yield, well-spread sample and adequate clinical information. By following the above recommendations and a little practice, the diagnostic yield of cytological specimens can be greatly enhanced.
About the author
Dr. Brandt completed veterinary school at the University of Tennessee College of Veterinary Medicine in 2006. She then made her way to the Chicago suburbs where she completed a private practice, small animal rotating internship, followed by a year of clinical practice as an emergency clinician. While she enjoyed the interesting cases and animal contact, the serenity of time spent at the microscope won out and Dr. Brandt completed a clinical pathology residency at Colorado State University in 2011. Simultaneously, she obtained a Master’s degree in microbiology. Upon completion of her residency, Dr. Brandt took a position with Gribbles Veterinary, a commercial diagnostic laboratory in New Zealand, for a few years. Most recently, Laura was employed as an instructor at Cornell University where she enjoyed a busy and diverse cytology and hematology case load. Outside of work, Dr. Brandt enjoys cooking dinner for friends, live music, hiking and traveling.