Cytology for the General Practitioner (Part 1)

In-house cytology is an inexpensive diagnostic tool that can be utilized easily in just about any veterinary hospital – in theory. While I’ve never been to a hospital where they didn’t have Diff-Quik stain, slides, needles and a microscope, cytology can be a tricky skill to master because not everything is black and white (or pink and blue). It’s also difficult to justify the time spent contemplating which purple blobs you’re looking at while the next patient is waiting. Because of this, some vets are discouraged enough about cytology that they don’t even feel it’s worth laying their own eyes on the slides. Don’t get me wrong, if there is ever any doubt, a board-certified clinical pathologist should absolutely look at your slides. But wouldn’t it be nice to be able to give the owner some sort of idea what you think of it before sending them home? I do think we can get to this point without being pathologists ourselves. Also, taking a look at a slide in-house is important for making sure there are actual readable cells in your sample, if for nothing else. My goal in this blog is to give you enough of the basics of cytology to feel comfortable at least laying your eyes on the slide before sending it off, and maybe eventually feeling brave enough to take a stab at a tentative diagnosis (pun intended)!

Getting the sample

There are many things that can be sampled for cytology – impression smears, scrapings, swabs, tape preps, aspirates, and more. For the scope of this discussion, I will focus on aspirates of masses. So, how do you ensure the best chance at success when poking that unknown mass?

Aspiration technique

There is always some discussion about whether it’s better to use the suction technique or the “woodpecker” technique. In the suction technique, sample is aspirated with a needle attached to a syringe and gentle suction applied in 3-4 directions within the mass before removing from the tissue. The woodpecker technique involves using the needle without the syringe, and poking up and down in multiple directions, then attaching it to a syringe of air to expel onto the slide. I personally like to use the woodpecker method for most things, but either method can work fine in most cases – use whatever you’re comfortable with. I would say, though, that perhaps the woodpecker should be considered for sampling of highly vascular (i.e. thyroid) masses or samples where the cells are fragile (i.e. lymph nodes).

I like to use a 22-gauge needle for pretty much all FNAs. Using a smaller gauge can cause more cell breakage because it shoves more cells through a smaller space. However, if you get a sample that is too bloody, using a 25-gauge could potentially reduce that blood. On the other hand, using a 20-gauge needle could get you more cells in your sample. I always get at least two samples, making a total of 4 slides once I smear them. When you go to splatter the cells onto the slide, try to minimize the amount of “splatter” by holding the bevel of the needle as close to the slide as possible. I like to use 2ml of air as a standard. When you smear the slides together, the key to keeping the cells intact is to be very gentle with your top slide. I don’t recommend pushing much at all with the top slide. For a lymph node, for example, you may even want to just set the slide on top and slide it across without any pressure at all.

Staining

Everyone has their own favorite ritual for staining slides. I keep it very simple – 30 seconds in each stain. If it seems like a particularly wet slide, maybe leave it in the fixative (the first jar) for 60 seconds. If you’re a dipper, 12 to 20 dips each should work. If you are worried about losing the sample in the stain, it is usually okay to use a hair dryer on the low setting from a bit of a distance away to dry it first. Yes, it is possible to dry a lipoma enough to be able to look at it and make sure it’s not a liposarcoma – I’ve done it! If you’re ever in doubt, there can never be too much fixation, so more time in the fixative can never hurt.

Drying

I always rinse the slide by dribbling a small stream of water onto the frosted part of the slide and letting it fall off the rest of the slide. If you’re really good about keeping track of which side the sample is on (I default to the side with the frost), you can confidently wipe the back of the slide with your bibulous paper or paper towel, and then blot the side with the sample gently. It’s probably ideal to let most things air dry, but I’m impatient, so I don’t do much air drying unless I really do think the sample will wash away if I’m not careful (i.e. lipomas). It is pretty important to get the slide quite dry, though. Water artifact can look pretty weird under magnification and may lead you astray in your interpretation.

Taking a look

The point I stress the most about looking at a cytology slide is not to skip the part where you scan at low magnification. It’s a habit I’ve had to learn myself, but I always start at the 4x or 10x objective. Of course, if your goal is just to determine if there are readable cells before sending it out, you may only need to go as far down as the 10x objective. Otherwise, this step is very important because it gives you the opportunity to pick up on how cellular the sample is, whether the cells exfoliated with any structure, and if there are any patterns. Just examining these characteristics alone, you can choose what cytomorphologic category you think the mass falls under, which already narrows it down to a finite set of differentials. A mass that does not exfoliate well (despite good sampling technique) could be a different type of mass than one that exfoliates a lot of cells. Cells that clump together really tightly likely signify a completely different tumor than one with cells that spread out individually or in sheets. The formation of certain patterns by the cells can sometimes be a key clue to make you lean toward one type of tumor versus another, including a malignant tumor rather than a benign mass. In more rare cases, scanning at low magnification can prevent you being fooled into thinking there is only an inflammatory population and no neoplastic population. If you zoom in on the first cells you see, you could miss an entire population that happened to also be present. Similarly, low-mag scanning can also catch instances when there are two different neoplastic populations in the same mass, which is rare but definitely can happen!

I will briefly describe the differences you could see with each of the four categories, and in the next blog I will go deeper into each one, with some differentials and examples. Of note, there are plenty of inflammatory lesions we could discuss (and perhaps we can go back and do that in another blog), but for now I’ll talk about neoplastic lesions. Again, these general characteristics are things you can often pick up on before even touching the 100x objective.

• Epithelial neoplasms will generally exfoliate a decent number of cells, which will usually be large round to polygonal cells with distinct borders, that clump together tightly.
• Mesenchymal neoplasms often don’t exfoliate well, and if they do you will see spindle, stellate (star-like), round, or fusiform-shaped cells with indistinct (fading) borders that exfoliate in looser sheets or individually.
• Round cell neoplasms are moderately cellular, and will have round cells with distinct borders, which will spread out individually.
• Naked nuclei neoplasms are usually pretty cellular, but will look like free-floating nuclei on top of a background of ruptured cytoplasms, so they will generally spread out, but may cluster occasionally.

Here is a table to summarize these:

Stay tuned for the next blog in this series, and we’ll take a deeper dive into each of these categories and discuss some common differentials to keep in mind from each one!