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Gill And Body Scrapes/samples


I've had a couple of people PM me about gill scrapes and how to conduct them latley, so I thought a general thread on how you conduct body samples could be of use to everyone, since I'm very interested to hear of alternate methods that may work better for goldfish in particular (I take samples the same way I do on koi, so there may be a more goldfish friendly way of conducting samples) I'm going to be using photos off fishdoc as I do not have someone to take photos while I work, so image credit to them.

Gill biopsy

When I need a tissue sample from the gills, and not just a mucous sample, I take a small biopsy from the from the lamellae tips in the gills. The fish is sedated for this procedure (because no matter how used to handling, if I went to take a tiny bit of your lung you would fight) and lifted so one gill is clear from the water. Sterile surgical steel scissors are then used to remove a tiny amount of tissue from the lamellae tips. This tissue is then prepared and mounted on a slide for investigation. I biopsy both gills. The fish is treated in salt to prevent any infections after taking samples.


On gill biopsies- Not recommended unless you absolutely NEED a tissue sample. It is very easy to do damage to a fish's gills via this method, and unless you have been instructed in how to take a biopsy or have a very thorough understanding of a fish's gills, it's best to leave this one alone.

Mucous scrape - Body

If large and hard to handle, sedate and lift the fish out of the water, place on clean towelling. If tame/small enough, just half lift them out of the water. There is usually no need to sedate a fish for a body scrape unless they are extremely large, impossible to restrain safely, or pose a threat to the handler (ie, piranhas) Using a blunt sterile scraper (I prefer metal ones, less likley to contaminate samples then wood or other porous materials), gently take a sample of mucus from either immediately behind the gill cover, alongside the dorsal fin or the base of the tail. Hold the scraper at 45 degrees to the body and draw it backwards towards the tail smoothly .5cm or so depending on size of fish. On larger fish you can take a larger sample if needed, but less is best for the fish. The scrape lifts a small amount of mucus from the sample site. Mount the mucus on a slide and investigate. Treat fish with salt to prevent infection of the scraped site.


On body scrapes- Very quick, safe and easy to conduct, even for an amateur as long as they have common sense. Some fish are so well handled you can lift them and scrape them with ease.

Mucous scrape- gills

If the fish is small, tame and used to having you touch around it's gills, you should not have to sedate for this procedure. if the fish is touchy, very large, impossible to restrain safely, or pose a threat to the handler (ie, piranhas) it's in the interest of safety to sedate them. Restrain the fish carefully, samples can then be taken from the gills by gently inserting a sterile cotton-bud under the operculum and rolling it over the gill filaments gently. DO NOT apply pressure. It is very easy to damage the gill filaments. We want the mucous on top, which will soak into the bud naturally, you do not need to apply pressure lest you push through the mucous to the filaments. Wipe the cotton bud onto the slide and mount it quickly. Treat the fish with salt while it recovers to prevent infections.

On gill scrapes- Not as easy as a body scrape, but not as hard as a biopsy to carry out correctly. However never carry out any scrape or biopsy without knowing exactly what you are doing lest you damage something.

With any scrape/biopsy

- Always wear gloves. Not only for the fish's safety, but for yours! Some diseases can be transmitted to humans.

- Always use clean sterile equipment.

- Salt fish afterwards to prevent infection. The combination of stress and taking samples (and sedations in some cases) can let secondary problems in while the immune system is down

- Never do a biopsy without first being shown how to conduct one in person by an experienced person. It's too easy to mess up

- Never Take a gill scrape if you are unsure of what you are doing. It's always best to be shown in person.

- Remember You are your own vet. When a fish gets ill it is very rare that we have the luxury of taking them into a clinic for treatment as we would a cat or dog. We are usually on our own to discover and treat issues. Take advantage of the collective knowledge of the forum for help

- If you are ever unsure don't do it. Especially if it concerns the gills. Its better to do broad treatments for issues then risk taking a biopsy or scrape and doing irreversible damage

I'm assuming everyone knows how to correctly mount a slide and use a microscope? If not I can add a guide for that later. Never use untreated water for a live slide, you will kill any organisms/parasites. Use distilled pure water, or even tank water if you have to, though it will contaminate the slide it's better then killing what you are looking for.

I'm also planning on posting images of normal mucus, as well as a parasite guide to follow this up with. And I'm very interested to see others methods for taking samples :)

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Normal mucous is grey and almost bubbly looking under the scope. debris and air bubbles are easily mistaken for parasites, so you should be ready to identify what is and what isn't normal in a healthy scrape before looking at your own.


Here's some healthy mucous with the usual things that trick people into thinking there's a problem, natural debris (usually microscopic food particles) and air bubbles. These are normal in a slide, bubbles are just from a bad mount (the better your mounting skills the less of these you will see) and the debris is just that, naturally occurring debris. No cause for concern here.

Not to oversimplify, but the 5 most common things I see on slides are flukes, costia, trichodina, chilodonella and white-spot. I'd estimate 80+% of the time that's what I end up seeing. Some strains of these are immune to what may have already be used to treat them (I've had prazi resistant flukes before, not fun at all) so don't skip looking for flukes jut because you've prazied recently. Learning to spot the 'common 5 issues' makes you more qualified them most "fish vets" I've encountered, so it's useful to have some knowledge, even if just theoretical of what they look like.

- White-spot are dark circles, slowly rotating, of varying sizes. Often with a lighter, horseshoe-shape visible inside. Quite large, hard to miss parasites.


White spot

- Flukes are long and worm-shaped and move in a looping action. Usually clearly visible hooks at one end.


Skin fluke


Skin fluke with developing embryo

- Trichodina are medium sized and round with a series of inner, concentric rings. These move fast like little UFOs




Trichondia in biopsy

- Costia is small and easy to miss. It's fast-moving though, and can be recognised by its flashing and twinkling as it moves in and out of focus


Costia in a histological section (cannot find a suitable scrape image)

- Chilodonella are medium, oval shaped parasite that turns and glide


I've tried to pick images closest to what you will see on a slide prepared by yourself. Gradually increase magnification too look for different problems when looking at a slide. Some very small parasites like costia will only show up at x400 magnification. Up the contrast if you can to get a better look at things.

Photo credit to fishdoc

Edited by Amber
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How to mount slides

Scrapes like this with living organisms should be wet mount. The following in purple is taken from the Museum of Science as it's a better description then my own.

Tools & Materials

* microscope

* flat slides

* cover slips

* eyedropper

* water

* toothpick

* paper towel

* sample


To make a wet mount place a sample on the slide. Using an eyedropper put a drop of water on the sample. Place one end of the cover slip on the slide and slowly lower the other end using the end of a toothpick. This will help to prevent air bubbles from getting trapped under the cover slip.

The water should just fill the space between the cover slip and the slide. If there is too much water and the cover slip is floating around, remove some water by holding the edge of a paper towel next to the edge of the cover slip. If there is too little water and some of the space under the cover slip is still dry, add more water by placing a drop right next to the cover slip. A little practice will help you learn how much water to add.

Once you have successfully identified your what is on your slide, if you have treatments on hand you can irrigate them through the slide by putting a drop of the treatment on one side then paper towel on the other to draw it through the specimen, and observing it to see what kills it. This way you can try treatments (especially useful if you have a resistant strain) on the slide instead on your fish until you find something that works.

Wet slides generally dry out within half an hour, so work fast, but thoroughly. Always use pure water, never untreated water as it will kill the parasites. If your really hard pressed use tank water, but be aware it may contaminate your slide.

it is normal to find afew parasites on a fish at any given time. All fish (and all creatures) carry a parasite load no matter how clean or well cared for. Even you have a parasite load of some sort! It is when this parasite load gets high that problems begin.

Edited by Amber
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