skin and gill scrapes,
an essential diagnostic procedure
Written by Frank Prince-Iles
Taking a skin scrape
Taking and preparing a skin scrape involves using a blunt scraper, such
as a wooden spatula, to gently take a sample of mucus from either
immediately behind the gill cover, alongside the dorsal fin or the base of
The scraper is held at approximately 45 degrees to the body and drawn
backwards towards the tail in a smooth movement, lifting off a small
amount of mucus from the sample site. The mucus sample is then smeared
onto a clean microscope slide along with a drop of pond water. Never use
tap water as any residual chlorine could kill any parasites that are
The sample is then covered with a cover-slip and examined under the
microscope, usually low-power, for the presence and number of parasites. I
would normally take at least two samples, from different sites, from each
fish being examined.
Taking a skin scrape and gill biopsy
|Taking a skin scrape using a wooden spatula to gently
lift off a mucus sample from just behind the operculum
|The mucus sample is put onto a glass microscope
slide. A drop of pond water is added and a cover slip put on top
|Taking a small gill biopsy using a fine pair of
There are certain considerations if the examination is to yield useful
results and avoid causing damage to the fish being examined. The most
important concern is to avoid injury to the fish during what should be a
simple, safe procedure.
If the fish does have parasites they will be found in the mucous layer
(though some will also penetrate the epidermis, e.g. 'Ich'), so it is the
mucous layer we need to sample for the wet-slide preparation. It is
important to realise that damage caused to the epidermis while taking a
mucus sample will be detrimental to the fish, so only gentle pressure with
a blunt scraper should be applied - never use anything sharp that might
damage the epidermis.
To sedate or not sedate - that is the
Another consideration is whether the fish should be sedated while a
scrape is taken. This is a subject that most books and magazines seem to
avoid so I will put my neck on the block and give my thoughts on the
In the first instance we want the fish to be still enough to do the
scrape properly, while at the same time avoiding damage to the epidermis.
With two people, one holding the fish and the other taking the scrape, it
is possible to sample smaller fish and docile larger ones without the need
for sedation. However, if there is likely to be a lot of flapping around,
there is a real chance that mucus may be stripped off by the constant
handling, thus giving an inaccurate result or, alternatively, a danger
that the fish may be damaged.
On the other hand, if fish are routinely sedated prior to taking a
scrape, there is the slight added risk that the fish could die from an
overdose of anaesthetic. There is also the consideration as to whether the
anaesthetic will affect the parasites and give a false result- although
this has not been my experience if the scrape is taken and prepared
quickly. In short, it seems to be a question of experience in deciding
whether the procedure can be carried out effectively and safely without
anaesthetic or whether the additional risk involved in sedating the fish
is the lesser of two evils.
I must say that I have come across some rather inaccurate conclusions
when people have tried to take scrapes from large, unsedated, lively fish!
When dealing with larger koi on my own I invariably find MS222 anaesthetic
a help. Your thoughts on this subject would be welcome!
If a parasitic infestation of the gill is suspected it is possible to
sample mucus from the gill. It goes without saying that this procedure is
potentially dangerous and must be carried out with extreme care and only
on sedated fish.
A mucus sample can be taken from the gills by gently inserting a
cotton-bud under the operculum and rolling it over the gill filaments.
Under no circumstances should any pressure be exerted that may damage this
vital and delicate structure. The mucus is then spread onto the microscope
slide. For accurate results it is important that the sample is prepared
and examined as quickly as possible.
With care it is also possible to take a small sample of gill using fine
scissors to take a small biopsy from the lamellae tips. I urge that this
procedure is not carried out unless one is totally confident about the
procedure as the tiniest slip could cause considerable damage to the fish.
How many is too many?
Generally one or two observed parasites per slide should not be a
reason for concern, whereas ten or more per field of view would be.
Usually, when there is a serious infestation it is quite obvious as the
slide often seems alive with parasites!
There are two potential problems that may be encountered when examining
a skin or gill scrape. All of the smaller parasites are virtually
transparent and unless the slide is scanned slowly and methodically there
is always the possibility that some parasites may be missed, especially if
they are not moving. If there seems to be a problem it is worth lowering
the microscope condenser to improve contrast or, better still, if the
option exists, try viewing the slide under dark-field.
If the mucus scrape is particularly thick it may be necessary to view
at different levels, starting from the bottom and working progressively to
the top. Otherwise, there is the chance that smaller parasites may be
missed because they are hidden in the mucus. This becomes more likely at
Don't jump to conclusions
The second, more common, potential problem is simply jumping to
conclusions. It is just too easy to spot the most obvious parasites, for
example skin and gill flukes, and conclude that these are the problem.
Even when you have spotted parasites it is vital that the whole slide is
still methodically examined to make sure that nothing important is missed
and so you build up to the correct conclusion. My own record is: five
different species of parasite on one slide - but
it took a full 15 minutes to find them all!
If a fish is found to be heavily infested it is worth taking a scrape
from one or two others - to determine whether there is a general problem
in the pond or if this is an isolated instance. It is often the case that
rather than the whole pond requiring treatment, just the one fish is ill
and the parasite explosion has resulted as a secondary infection for that
fish, not the whole collection.
Practice make perfect
Although the microscope is a fairly simple instrument to set up and
use, interpreting what you see can sometimes be difficult. It really
requires practice until such times as you know what is normal and what is
As a piece of advice I would suggest familiarizing yourself with
healthy or non-urgent fish mucus samples - rather than waiting until a
serious case arises. By getting used to taking and examining mucus samples
you will be better prepared to make a judgment when a serious problem
What to look for
I am assuming that initially the microscope is going to be used as as
simple diagnostic aid to carry out procedures such as skin and gill
examinations. It can of course be used for more advanced studies, such as
plant and animal cell structure. You will probably have seen photos of
slides on the site showing details of various fish organs such as gills.
This is histology and involves special preparation and staining of the
At a basic level, microscopy would be used as part of a routine
examination to check mainly for external parasites. This involves taking a
skin or gill scrape / sample and making a simple wet mount as previously
Initially it is very easy to get confused by what you see on the slide
- particularly if small non-parasitic aquatic animals happen to be
sampled, leading to fears of some new, frightening parasite or disease!
The first stage is to recognize what normal mucus looks like and
disregard any "debris" or unimportant "introductions".
These could be things like air bubbles trapped under the cover slide which
appear as circular, dark-rimmed "grommets". Or perhaps normal
cellular debris which appears as stationary, irregular-shaped - often dark
patches. You are also likely to see trapped algae of all shapes and sizes.
This is why it is important to get as much practice as possible before
using your microscope in earnest.
Normal mucus in a wet mount at 100x
|Normal mucus which is seen a lumpy,
transparent, light to grey substance. Also shown is a trapped
air-bubble which is often mistaken for a strange parasite, and
some typical cellular debris. Active parasites should be easily
At the risk of oversimplifying what can be a complex issue, in the vast
majority of cases (>90%), the usual findings would be a common parasite
problem. In freshwater fish such as koi and goldfish these usually involve
just five different types namely, flukes, Costia, Trichodina, Chilodonella
or white-spot. So, simply being able to recognize these common beasties
will make you a reasonably competent microscopist. There are, as already
suggested, less common things you might come across, but once you are
familiar with these basics, it is fairly easy to focus on any abnormal
findings and either look them up in a good book or ask for advice.
In most slide preparations the parasites will be "alive and
kicking" and so they are easy to spot and recognize. However, it is
important to scan the entire slide, slowly and methodically looking for
parasites that are still. These can be more difficult to spot as all of
these parasites are transparent and therefore tend to blend in with the
mucus. However, with the exception of Costia, which is virtually
impossible to spot unless it is moving, they can be seen once you have the
experience and have "got your eye in".
This is why it is important to examine slides as soon as they are
prepared as many parasites die if left for any period.
There are many good books available which describe the common
parasites. There are further details and photos on the relevant pages of
this site. The movies of parasites will also give some idea of the way
each parasite moves; an important diagnostic feature. I have included a
short summary of each of the parasites below, but you will need to check
the relevant pages for more detail;
- Trichodina: Medium=sized round
parasites with a series of inner, concentric rings. These zoom around
like flying saucers. Go to page
- Costia: The smallest parasite
that is often easy to miss. A fast-moving parasite recognized by its
flashing and twinkling as it moves in and out of focus Go to page
- Chilodonella: A medium-sized
oval-shaped parasite that turns and glides. Go to page
Article and pictures placed here with permission from the author,