Disease Revisited (1-10-2001)
Stewart BSc (Hons)
Columnaris disease is a problem that has had a long and
often confusing history. What follows is broadly divided into three main
sections. Firstly, 'classic columnaris' is briefly outlined (see 1
for further information on columnaris disease; and 2
for further information on the Flavobacteriaceae). Secondly, recent
developments, specific to tropical fish are reviewed. Finally, what all
this may mean to the discus hobbyist is discussed - along with a 'real
life' example and potentially useful treatments. disease."
Columnaris disease is a bacterial disease of freshwater fish. It is the
second highest killer of farmed catfish in the USA 3,
and it is widely accepted that most species of fish are susceptible to
columnaris disease 4.
The aetiological agent of columnaris disease is the gram negative rod
bacteria, Flexibacter columnare (ex Herbert Spencer Davis 1922)
Bernardet and Grimont 1989 (Synonyms: "Bacillus columnaris;
Chondrococcus columnaris; Cytophaga columnaris; Flexibacter columnaris.
It is often referred to as "fin rot", "cotton wool
disease", "cotton mouth disease", or "saddleback
Outbreaks tend to occur following environmental stress and clinical
signs or symptoms may include yellowish brown or white lesions on the
gills, skin, or fins. The bacteria attach themselves to the fish where
they release protein and cartilage degrading enzymes. It may be argued
that the most important site of attachment is that of the gills. The
bacteria attach to the gills where they multiply, and eventually cover and
destroy the entire gill filament - if untreated, substantial damage can
occur, subsequently leading to the death of the fish 3.
In the early stages of the disease, the lesions may simply appear as an
area that is 'less glossy' than the surrounding scales. Advance lesions
may be round or oval in shape, and can if untreated, progress to an open
ulcer. In some fish the lesion may appear as a pale band, encircling the
body of the fish - hence the name saddleback disease 3.
Recent Developments in our understanding of
Prior to 1998, the majority of published works on columnaris disease
primarily focused on temperate farmed fish 5.
During 1998, Decostere, Haesebrouck and colleagues 4,
5, 6, 7, became the first to publicly correct this problem by
isolating and studying Flavobacterium columnare directly from tropical
aquarium fish. The fish species studied included black mollies Poecilia
sphenops, platies Xiphophorus maculatus, guppies Poecilia
reticulata, and tetras Cheirodon axelrod.
Decostere et al. 5
isolated four strains of F. columnare. Each of these strains were
compared to the F. columnare reference strain, NCMB 2248, and or
the 5 F. columnare strains that had previously been isolated from
temperate fish. Whilst there were similarities e.g. all were gram negative
'flexing' filamentous bacteria between 0.2-0.5µm x 5-8µm in size; all
had an optimum growth rate at between 25°C and 30°C - there were also
significant differences amongst the strains.
Importantly, the tropical strains continued to grow at 37°C; they
demonstrated a notable difference in virulence; and none were able to grow
in-vitro, when immersed in a 1% NaCl (Salt) solution. In tanks where
mortality occurred, the time to death following infection ranged from
between 8 hours and 6 days. It was also noted, that the strains were able
to enter the blood system and cause septicaemia.
The 4 'type strains', which were isolated, are listed in order of
- AJS1 was extracted from a fish with bleached and ulcerated skin, and
from a tank in which daily mortality occurred.
- AJS2 was extracted from a discoloured fish, which displayed fin rot,
especially in the caudal region; in addition, there was chronic
mortality in its aquarium.
- AJS3 was extracted from a fish that displayed small white patches
around mouth, opercula, and from tanks that had no significant
- AJS4 was extracted from a discoloured fish, with pale patches on the
skin and fins, especially in the dorsal region, and again from a tank
that had no significant mortality
In fish that were infected with F. columnare, the sequence of
visible physical symptoms included slight swellings at the site of
infection (infection was achieved by intramuscular injection of simple
contact infection), which subsequently developed into a white/blanched
patch. As the infection progressed, in the more virulent strains, the fish
began swimming at the water surface. Shortly before death, the fish lost
their ability to remain at the surface and became motionless, lying on
their sides on the floor of the aquarium. AJS1 was the most virulent
strain with acute mortality occurring 10 hours post-infection.
Further studies 5,
7 on tropical columnaris note that the development, progress, and
ultimate pathogenicity of the disease, is highly associated with the
ability of F. columnare to adhere to gill tissue. AJS-1 has a high
adherence ability and AJS-4 has a low ability. Therefore, as AJS-1 is the
more virulent and deadly strain, the mechanism and conditions that favour
attachment are of prime importance.
Factors that affect the gill adherence capability of F. columnare
include: temperature, over-crowding, excessive organic loads, ionic
composition of the water, excessive or poor handling, and slow movement of
water. The mechanisms hypothesised to explain these occurrences are not
fully understood - some suggestions include:
- The slow movement of water allows the bacteria to withstand the
water-flow, and turnover of the mucus cells/epithelium - thus allowing
the bacteria to remain in an area where there is a high concentration
of nutrients, that are efficiently obtained via 'slime layer'
localisation of dergradative enzymes.
- Ionic composition of water affects adhesion. MG++ and CA++ play a
part in adhesion by reducing surface potential and repulsive forces.
- High levels of nitrites and organic matter enhance adhesion; the
underlying mechanism involving nitrites is not known and it is
suspected that high levels of organic matter may result in debris
being trapped within the mucus layer at gills thus resulting in an
'ideal target site' for the bacteria.
- In regards to the cellular binding mechanism involved between gills
and the cells, a capsule incorporated, lectin-like carbohydrate has
been suggested 6.
Columnaris and the implications for
the discus hobbyist
There can be no doubt that columnaris disease can occur in virtually all
aquarium environments. It may be present even when there are no obvious
external signs. When external signs are present, they may take the form
of: yellowish brown lesions, 'cotton wool' tufts, the fish may simply look
'dull' or 'dark', or the fish may have small white patches on its fins or
body. In addition to these external signs of infection, it should be noted
that, one study found that in 40 % of all diagnosed cases of columnaris,
the internal organs were also affected (see 3
Stress is a major factor in columnaris disease and may involve any one,
or combination of stressors e.g. low oxygen levels, high nitrite levels,
comparatively high (or low) water temperatures, rough handling, mechanical
injury, overcrowding, water of inappropriate hardness etc (See 8,
9 for further details on stress, fish, its management and fish
In discus as with other tropical fish, many of theses stressors occur
during catching, bagging, transporting and subsequent reintroduction into
the new aquarium. The closed re-circulating system of the aquarium is an
ideal habitat for columnaris to spread and to result in high mortality
rates. In addition, it is not unusual for discus to be kept in overcrowded
tanks with inappropriate water parameters; or to be placed in tanks that
have immature filters.
For the discus hobbyist the external signs of columnaris to the naked eye
can closely resemble the signs presented by range of other parasites. It
is unlikely that the average hobbyist will have the necessary equipment,
time, experience and access to the drugs that are required to be able to
identify specific strains of F. columnare; however if they do, I
suggest they start with references 4,
5, 6, 7 and follow up their original methodology used by the authors
as a guide. However, gross identification of F. columnare is
certainly achievable using a reasonable microscope, a high powered lens,
appropriate accessories, and comparative slides, images, videos.
Rather than repeat information that is commonly available, I suggest
that, if you are new to microscopes and the gross identification of F.
columnare, that you pay a visit to the DPH
Articles Page - there, you will find a collection of excellent
information, images and videos.
I often meet people who moan that microscopes and vets are expensive and
a waste of time in regards tropical fish. In my experience an adequate 'scoping'
kit cost less than a decent pair of discus (certainly in the UK) and
with a little practice becomes a phenomenal diagnostic tool, and may
well aid the survival of your complete stock.
Secondly, if one takes along suffering stock to a vet/aqua culture
specialist who has the experience and equipment to identify specific
pathogenic bacteria - this again can cost less than the price of a
breeding pair. Be prepared to ask the individual if they have the
necessary equipment for accurate identification on the premises. In
saying this, I accept that you are not going to be able to use your
local dog and cat vet (unless you are very lucky) - Contact your local
fisheries advisory board; they often keep a list which can be issued to
the public of the laboratories that they use, e.g. I have used the
Ministry of Agriculture, Fisheries and Food MAFF; which are now the
Department for Environment, Food and Rural Affairs DEFRA to track down
the appropriate specialists in various regions of the UK for friends and
colleagues. Remember your stock is potentially worth thousands - it is
not good sense to rely on guess work and chance when confronted by any
potentially virulent disease.
The first step in managing columnaris is to prevent its occurrence. To do
this, it is important to minimise the amount of stress our fish are
subjected to e.g.:
- Proper handling - use soft nets and careful netting techniques to
avoid mechanical injury
- Ensure that there is adequate oxygen in shipping bags and tanks
- salt or other additives may be used to minimise the effect of
- Allow a sufficient run-in period for the maturation of new tanks
- coupled with regular partial water changes appropriate to
stocking density and feeding regime
- Do not overstock or keep discus in water of inappropriate hardness
- When receiving new fish - ACCEPT
that they have just undergone a journey that is likely to have placed
them under a phenomenal amount of stress - therefore, the risk of
columnaris (amongst other infections) is considerably higher than
- It is all to easy to blame the breeder or wholesaler when fish
arrive worse for the wear and an epidemic of columnaris breaks out
almost immediately the fish hits the tanks. In the main, breeders
or wholesalers will take every precaution possible to ship their
stock correctly - once the shipment leaves their premises though -
a lot can happen, from plunging pH levels, increasing carbon
dioxide levels/decreases in the available oxygen, chilling of
water, boxes being thrown through the air and dropped from
Hopefully the above is indicative of how important it is to quarantine
new stock correctly, and to take steps to reduce any population increases
of ecto- and endo-parasites, that may have occurred during the relocation
process. With this in mind I would like to draw attention to several
points re: quarantining discus.
- A quarantine tank (qt) is not necessarily a hospital tank (although
it may become one).
- It should contain a mature filter and conditioned water, prior to
the new fish being added. The qt tank's water should match the water
in which the discus will ultimately be placed.
- A record of observations are essential in preventing, diagnosing and
treating problems e.g. temp, pH, GH, KH, behaviour, physical
appearance, feeding practices, the appearance of waste etc. . . Don't
forget, during the qt period you are not only making observations for
visible signs of disease, you are also looking for signs of stress -
the most common pre-cursor to disease outbreaks.
- Many different quarantine periods have been suggested from 1 week to
6 weeks. In my opinion the minimum for discus is 4 weeks with the
preferred time being 6 weeks.
- If a 6 week quarantine period has been chosen - between weeks 4
& 5 add a discus from the main tank into the qt tank, in case the
newly acquired discus are unaffected carriers of, as yet, unidentified
Columnaris outbreak - an example
Whilst the best 'cure' for columnaris is prevention; F. columnare
is such a ubiquitous organism, that there is a high probability that at
some point in our fish keeping lives, our stock will suffer from an
outbreak. The example that follows is based on a factual occurrence and
the text was provided by Davis Gailitis.
'One day I looked into my tank and noticed that one of my discus
looked different' . . . 'she had a spot on her left side just below the
beginning of her dorsal fin. It looked like a scale had come up a bit. I
now know looking back that I should have looked closer at it. I just did
a casual glance and assumed that was what it was. I didn't think
anything of it, except to look at it again later that day'
'Things stayed the same for several days and I was not overly
'On the fourth day I noticed the scale that was turned up, now
looked like a small pimple, all white, around 3ml in diameter and about
the same in distance protruding from the body.'
'The next morning I looked at her when she was facing sideways to
me all of a sudden I could see, in her slime coat, this white opaque
haze, circular in form, about the size of an American dime.'
'Later that evening the opaque circle had grown t the size of an
American Quarter. Up until this point, my fish had behaved normally and
was eating with the rest. Tonight, she was at the back of the tank,
facing the corner, and getting progressively darker.'
Following discussion with Fred Goodall and further investigation -
several days of treatment were performed with the result of . . . 'She
is fine now and is horny as hell, she has also found a mate in the tank
and is on a four day cycle with laying of eggs!'
In my experience Davis' example is very common. It is important to
mention here that this is only one manifestation of columnaris. In many
cases columnaris has been known to effectively wipe-out several hundred
discus in a matter of weeks; with the first deaths occurring within hours
of a new shipment arriving at the retailers.
Should you be unfortunate enough to experience columnaris, the disease
must be brought under control as soon as practically possible.
In Davis' example, the treatment referred to, was the application of a
series of salt dips, coupled with in tank temperatures of 35°C, large
daily water changes using fresh conditioned water, and scrupulous tank
hygiene. Below, I list a couple of methods that I have used and that have
worked for me and others I have visited. If you choose to use the examples
- do so with great care - remember, the onus is on you to make judgements
regarding appropriate treatment and the current condition and status of
your fish and tanks - if you are unsure, nervous or need help, please ask,
or seek professional advice.
Discus Salt Dip Methodology
Type of salt to use:
The type of salt used should be non-iodized and contain no 'free flow' or
other additives (e.g. no iodine or sodium ferrocyanide etc. I have used
'Freshwater Aquarium Salt', rock salt and sea salt. If purchasing
non-aquarium salt please read the packaging carefully as current trends
show an increase in the use of additives even in natural products such as
rock salt (UK).
If new to salt dipping:
If new to dipping fish a good place to start would be with a 1.5-2% salt
solution; for more experienced users I would suggest you start with a 3%
solution immediately. The solution should be made up in a clean bucket or
spare (fishless) tank. Whilst it is preferable to weigh out the correct
amount of salt e.g. for a 2% solution one would use 20g of salt per litre
of water, the following approximate measure are given for the sake of
1 TABLESPOON of salt approximates to 15grams.
Therefore 1 TABLESPOON of salt per litre of water equates to a 1.5%
- If your bucket/tank contains 10 litres
of water you would add 13 tablespoons of
salt to get an
approximate 2% salt solution
- Or - If you place 3
US gallons of water in a 5 gallon bucket you would add 14½
tablespoons of salt - to get an approximate 2% solution.
And so on
- It is important to ensure that the salt is fully dissolved before
placing the fish in the bucket/tank and that the water temperature
matches the tank from which the fish are taken.
As you will need to multi-dip throughout the day, place a heater in the
tank, if you do not want to have to remake new salt solution each time;
personally I make a fresh solution for each dip.
Before you place the fish in the solution please remember that:
- The length of time that you can leave discus in the solution varies
greatly from a few seconds to 30 minutes
- 5 minutes would be a reasonable average
- The time they tolerate the dip DECREASES with the number of dips
performed in a 24 hour period
- The fish MUST NOT be left unattended
- During the treatment your fish may show some interesting
discolouration, do not worry this is short term.
Place the fish into the solution as quickly and as carefully as
possible - then observe closely.
Initially, the respiration of the fish will increase substantially, 120
gill beats per minute is not uncommon. At some point, the fish will
keel over on its side - and it is at
this point that the novice should remove the fish and
return it to its tank. If you are confident and experienced in dipping you
may want to extend the time that the fish is exposed to the saline
solution - I have found it effective to leave the fish in the solution
until the gill beats have slowed to around 20 bpm - irrespective of
whether the fish has keeled over or not.
Once the fish are returned to their tank they should within a few
minutes regain their composure. If they appear to be in difficulty, the
fish can be supported using your hands and then gently pulled backwards
through the water at a slow pace - so that water is forced over the gills.
I have dipped 5 times a day for up to 7 days and there has been no
lasting negative effect. It is important that the
dips continue until all visible signs of infection are gone.
In regards to salt dipping it may be argued that returning the dipped
fish to the tank from which it came is pointless - this is not the case.
Ideally we would want to return the fish to a tank which had been
sterilised and which contained 100% fresh (but conditioned) water, with
a matured filter. It is important to remember here, that all we are
doing is 'knocking back' the population of a ubiquitous organism and
promoting the production of copious amounts of slime layer in order that
the fish immune system can regain control and management of the problem.
If we must return the fish to the same tank etc. then large water
changes with conditioned water, scrupulous tank hygiene and the addition
of salt at a rate of 2 table spoons per 10 gallons (assuming that the
tank does not contain delicate plants or salt intolerant species) can
work wonders - in conjunction with correct salt dipping.
Whilst I personally prefer the use of salt - it should not be forgotten
that there are several alternatives. In relation to discus, various
combinations of: potassium permanganate, formalin, temperature, hydrogen
peroxide, acetic acid, large water changes, and so on, have been used to
help CONTROL the external infection - with the likelihood of success being
dependent on the virulence of the strain involved. For the sake of balance
I will summarise the use of potassium permanganate (as an insitu bath [not
1. Potassium permanganate and formalin must never be used in the same
water, nor should they be stored together. When combined, their vapours
are highly explosive.
2. I only use PP in BB tanks.
3. Wear rubber gloves, eye protection and old clothes - the aim is to
treat fish - not to become blind, to have sun-tanned hands or to stain
your best outfit. PP is a powerful oxidising agent and shows no
discrimination between human skin, fish gill filaments, mucous layers
etc - it will start with the top layer and continue to oxidise, until
exhausted - therefore, it must be handled and used with appropriate
respect. If you do get PP on your clothes or work surfaces, the
resulting brown stains of manganese dioxide can be removed by using a
solution composed of 1 parts 9% hydrogen peroxide, 4 parts white
vinegar, and 3 parts tap water. If you end up having to use this
solution to remove the stain from your skin, wash off the excess once
the stain is gone. Do not use: if the skin becomes red or
irritated, or on open wounds, or burns etc.
4. When working with PP in aquariums, it is probable that you will be
working with very small amounts e.g. 1g and dilutions of . . .
A 1g spoon can be purchased from some of the 'older' style chemists
which can be used for making up 2l of stock solution at a time; however,
many chemists will weigh out the necessary amount for you, if you tell
them what you are using the substance for. If you are new to the use of
PP, I strongly recommend you let the chemist weigh the necessary amounts
for you, so that you can see precisely what you are supposed to be
working with - you may be surprised at how small the amounts are.
5. Never expose discus that have open wounds to PP treatment.
Potassium Permanganate (KMnO4)
Potassium permanganate (PP) can be used to treat external columnaris. It
is added to the aquarium water, where it oxidises all organic matter,
until it reaches a point of exhaustion. Commonly, 2 mg/l of PP is added to
the aquarium water, and the tank is then monitored to ensure that the
water retains the resulting 'purple/red' for at least 4 hours. If the
colour begins to fade, then more PP will have to be added.
In my opinion, it is all to easy to overdose and kill you fish when you
simply rely on subtle changes in colour; therefore a 15 minute PP demand
test should be performed. By performing this test you will be able to
calculate the correct amount of PP to use, in respect to the organic
loading of the aquarium water in which the fish are to be treated.
Methodology to perform a 15 minute PP demand
test on aquaria and ponds that have a high organic load
If you have a bare bottom tank and perform regular water changes,
siphoning off any debris each day - you may find that even at 1mg/l the
water will retain a degree of pink colouration - if this is the case I
suggest that you use the standard treatment dose of 2mg/l PP and
maintain the colouration for four hours - Perform the test first, though
- Don't guess!
- Step 1
Prepare a 500mg/l water 'stock solution'. This is achieved by adding
0.5g of PP to 1 litre of distilled water.
- Step 2
Label four, 1 litre, transparent containers (e.g. 1, 2, 3, 4)
- Step 3
To each container add 500 ml of water taken from the tank that is to
- Step 4
Then add the following:
Container 1 - add 1ml of stock solution (results in a 1mg/l solution
Container 2 - " 3ml " " " " 3mg/l " "
Container 3 - " 5ml " " " " 5mg/l " "
Container 4 - " 7ml " " " " 7mg/l " "
- Step 5
After 15 minutes note the colour of the solution
- Step 6
Calculate the quantity of PP to use in the aquarium to be treated
A. You will find that the solution in one container will be
slightly pink, and in another the solution will be clear.
The concentration you use is the concentration that falls between
B. You then multiply the PP concentration selected, by a
factor of 2.5
This is the concentration of PP that you would use on the tank
containing the infected fish.
Container 2 (3 mg/l PP) has a slight pink colour
Container 1 (1 mg/l PP) is clear
Therefore you would use 2 mg/l x 2.5= a treatment dose of 5mg/l of PP
1. You are treating 200 litres of water
(see method below; it is important that you know precisely how much
water will be left in the tank after you have drained 50% or so off,
as it is this amount you will be treating
Length cm x Width cm x Height cm divided by 1000 = amount in
litres; don't forget to deduct the thickness of the glass if you are
measuring external edge to edge
2. You have calculated that the appropriate dose for
treatment is 2mg/l
3. You therefore need to add 200 x 2=400mg of
potassium permanganate to your 200 litre tank
a. This can be achieved be adding 800ml of the previously
created stock solution (remember it was made to a strength of
b. Or you could make new stock solution with 1 litre of
distilled water and add 1gram of PP to it.
i. You would then add 400ml of this solution to your 200
Once the final dose is calculated, there are two schools of thought, in
regards adding the PP to the aquarium. On the one hand, there are those
(myself included) that prefer to divide the total dose into 2mg/l portions
- adding each portion, only after the initial purple-red turns brownish
(tip: remove the cover glass and look into the tank from above to judge
colour most accurately). Others prefer to add the total amount of PP to
the tank as a single dose. There can be no doubt that the first scenario
is safer and less stressful, especially if you have severely infected
fish; therefore, it should be given some consideration. In addition, as
with the salt dip, discus can react in different ways to PP. In a tank of
6 - 5 may swim around normally and 1 may appear highly disorientated.
Now that the dose has been calculated, the next step is to siphon ½ of
the water out of the tank, cleaning up any waste material that you can.
This is followed by bypassing the filter system of your tank (PP will kill
you filter bacteria if you do not do this). If you use a canister filter,
you can use a spare 5 gallon bucket, filled with aquarium water and a
heater, and simply place the filter inlet and outlet into the bucket. An
air supply that can provide heavy aeration is then added to the tank. The
next step is to add the first portion of the total dose to the tank and
monitor the behaviour of the fish. Once the initial shade of purple/red
starts to turn brown, you should continue with the next portion of the
The most important points during this stage are:
1. Monitor the reaction of the fish; if they are severely stressed
a. you can either remove them, or
b. add hydrogen peroxide to neutralise the PP(dose given below in
2. Ensure that the purple-red colour is maintained for a minimum of 4
3. Remember that during this process you are stripping the slime
layer from the fish and that they will subsequently be 'vulnerable',
until a new slime layer develops; therefore, post treatment, tank
hygiene must be of the highest order.
Finally, once the 4 hour period is completed, you can neutralise the PP
and clear the tank by adding hydrogen peroxide to the water. The dosage I
use is ~ 1.3 ml of 9% hydrogen peroxide, for every 10 imperial gallons of
tank water, treated at a dose of 2mg/l of potassium permanganate. The
hydrogen peroxide and a 1ml syringe can be purchased from most chemists;
please check the strength of hydrogen peroxide you purchase as this varies
in strengths from 3% rising to 9%. The table below is provided for easy
conversion. If you require the molecular weights and conversion factors
used in creating table 1, please contact me.
||Potassium permanganate dosage used
||Amount of Hydrogen peroxide required
in ml @ strength 0.3%
||Amount of Hydrogen peroxide required
in ml @ strength 0.5%
||Amount of Hydrogen peroxide required
in ml @ strength 0.9%
Table illustrating the amount of hydrogen peroxide
required in ml, at various strengths, to neutralise a given amount of
Once the treatment is finished, I continue the aeration and add a
drip-feed of fresh conditioned water to fill the tank to its previous
level. Once the tank is full you can reconnect the filtration system. Keep
an extra eye on the ammonia and nitrite levels for the next couple of
weeks, and maintain a healthy regime of water changes; further treatments
may be necessary - if they are please remember that discus become less
tolerant to PP exposure.
Whilst salt and PP are can prove very effective in the management of
columnaris - there are times when additional methods may be necessary. For
example, in temperate and tropical aquaculture systems where the entire
stock of a breeder may be at risk; or where the disease has progressed to
the organs - antibiotics may be a reasonable option to pursue.
In my view, antibiotics are always a last recourse; and I only use them on
fish where there is a 'real' chance of survival. In other words, use
antibiotics only when the fish still has some 'vitality' to it and is not
1 step away from its final death throws.
In regards antibiotics two points should be remembered:
1. Breeding from antibiotic treated stock, too soon after the
treatment is irresponsible and ill-advised.
2. The effectiveness and toxicity of antibiotics varies with dose,
temperature, size and the development of fish; therefore, professional
veterinary advice should be sought - preferably for an aquaculture
specialist or similar.
et al. noted that in the tropical fish studied all F. columnare
strains reacted in a similar manner to antibiotics:
- High MIC values were found for: colistin (>128), sulfamethoxazole
(64) and neomycin (10±20)
- Low MIC values were found for kanamycin (2), nalidixic acid (2),
streptomycin (1±2), furazolidone (1±2), chloramphenicol (0.5±1),
oxytetracycline (0.5), lincomycin (0.125), erythromycin (0.031)
MIC values (Minimal Inhibitory Concentration) are defined as the lowest
concentration of drug, that inhibits more than 99% of the bacterial
population being investigated. Therefore, when treating F. columnare with
antibiotics, those with high MIC values should be avoided e.g. colistin,
sulfamethoxazole and neomycin. In regards the use of antibiotic feeds
developed for aquaculture - terramycin TM100 (contains 100 mg of active
ingredient oxytetracycline HCl) may be appropriate for larger
breeding/retail establishments; and medicated feeds such as Romet 30 and
Romet B (active ingredients sulfadimethoxine / ormetoprim) may not be an
optimum choice. Again, please remember that professional advice must be
undertaken before applying a drug regime.
In summary, columnaris disease is a deadly and common disease which can
have a catastrophic affect on fish populations; whether the fish are from
temperate water, tropical water, brackish water or sea water. The
ubiquitous organism can reach epidemic numbers when fish are subjected to
stressors e.g. the conditions that commonly occur during the handling and
transportation of fish. Currently, F. columnare is said to be the
organism responsible for columnaris disease; however there are a number of
strain permutations of this species. The most virulent and deadly strain
in tropical fish has been identified as AJS1 with the least deadly being
AJS4. These strains manifest themselves in slightly different ways;
therefore one fish keeper whose stock is suffering from AJS1 may loose his
or her entire stock overnight; whilst someone else's stock that have
contracted AJS4 may suffer a lengthy period of infection prior to
recovery. Within the range of strains discovered: all are tolerant of
higher temperatures than previously believed; all have a lower
reproductive rate when exposed to salt solutions from 1% upwards; all are
affected by antibiotics in a similar way. If visible symptoms are present,
the likelihood that internal damage is occurring is high. The more deadly
strains have a higher ability to attach themselves to the gills of their
hosts. It is likely that more strain variations will be found as research
continues, therefore the important thing to remember is to: keep an open
mind; note the symptoms; perform an accurate diagnosis of the gross
organism involved using a microscope, or an aquaculture expert; and
rapidly follow up with the preferred choice of treatment, whether it be a
salt dip, potassium permanganate bath or the use of antibiotics - coupled
with: scrupulous tank hygiene and appropriate stocking levels and feeding
regimes. Finally, if faced with an emergency and you suspect that your
fish may be suffering from a virulent form of columnaris disease, and you
neither have access to a microscope or appropriate expert, the 'broad
spectrum affect' of 2 days of repeated salt dips coupled with large
regular water changes may have wondrous results on a variety or microbes
Stewart BSc (Hons)
I wish to thank:
for his help in regards developing a salt dip methodology which I
consider to be specific to discus and for his time and patience reading
for the information and help he kindly supplied
for his generosity in allowing this information to be shared with discus
enthusiast from around the globe
for reinforcing the incentive to search out quality information in
regards Columnaris and all discus related topics
 Francis-Floyd, R., (1998), 'Columnaries Disease', FA-11:
Department of Fisheries and Aquatic Sciences, Florida, University of
 Jooste, P.J.; Hugo, C.J.; (1999), 'The taxonomy, ecology and
cultivation of bacterial genera belonging to the family Flavobacteriaceae',
International Journal of Food Microbiology, 53:81-94
 Durborow, R.M.; Thune R.L.; Hawke, J.P., et al.; (1998), 'Columnaris
Disease A Bacterial Infection Caused by Flavobacterium columnare', Southern
Regional Aquaculture Centre Publication, No. 479
 Decostere, A.; Haesebrouck, F.; Devriese, L. A.; (1998),
'Characterization of four Flavobacterium columnare (Flexibacter
columnaris) strains isolated from tropical fish', Veterinary
Microbiology, 62:1; 35 - 45
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of water quality and temperature on adhesion of high and low virulence
Flavobacterium columnare strains to isolated gill arches', Journal of
Fish Diseases, 22:1; 1 - 12
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'Characterization of the adhesion of Flavobacterium columnare (Flexibacter
columnaris) to gill tissue', Journal of Fish Diseases, 22:6;
465 - 474
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'The association of Flavobacterium columnare strains of high and
low virulence with gill tissue of black mollies (Poecilia sphenops)',
Veterinary Microbiology, 67:4 ; 287 - 298
 Francis-Floyd, R., (1997), 'Stress - Its Role in Fish Disease', Circular
919: Department of Fisheries and Aquatic Sciences, Florida,
University of Florida
 Rottmann, R.W.; Francis-Floyd, R.; Durborow, R.; (1992), 'The Role
of Stress in Fish Disease', Southern Regional Aquaculture Centre
Publication, Publication No. 474
 Francis-Floyd, R., (1995), 'The Use of Salt in Aquaculture', Fact
Sheet VM 86: Department of Fisheries and Aquatic Sciences, Florida,
University of Florida
 Francis-Floyd, R.; Klinger, R.E.; (1997), 'Use of Potassium
Permanganate to Control External Infections of Ornamental Fish', FA-37:
Department of Fisheries and Aquatic Sciences, Florida, University of