VIII Fish health and diseases

1 Disease in general*

1.1 Disease is defined as a disturbance in function or structure of any organ or part of the body. When applied to livestock, this definition can be enlarged to include detrimental behavioural changes e.g. in feeding response, within a herd or a community. The full potential economic impact of disease is not often realised. Thus, disease may result in:

1. deaths

2. stunting, unsightly changes etc

3. longer grow-out periods

4. poor feed conversion ratios

5. lower maximum stocking levels

6. opportunity cost of production, especially in a seedstock-scarce situation when stocks lost through disease cannot be replaced

1.2 Disease may arise for various reasons and it is common to find more than one cause of disease. For example, disease may be caused by one factor but is then masked by disease caused by another factor. In such a case, the second disease may be opportunistic i.e. it takes advantage of a weakened state brought about by the first disease. It is therefore very important to determine the underlying causes or origin of a disease. Causes of diseases are as follows:

1. stress e.g. importation mortalities

2. pathogens e.g. cryptocaryoniasis

3. adverse environmental conditions e.g. plankton blooms

4. toxic factors e.g. drug overdose

5. nutritional imbalance e.g. liver lipidosis

6. miscellaneous e.g. swim bladder syndrome

* Chong, Y.C. and Chao, T.M., 1986. Common Diseases of Marine Foodfish. 34pp. Primary Production Fisheries Handbook No. 2. Published by Director of Primary Production, Primary Production Department, Ministry of National Development.

1.3 Disease arising from different causes show different external signs typical of a particular disease. Examples of such signs are sudden death, changes in swimming behaviour, refusal of food, skin ulcers etc. These signs in combination with laboratory tests, enable us to diagnose (= identify) the disease. Consequently, appropriate measures can be taken to limit the spread of that disease, to cure sick fish and to prevent recurrence of disease. The control of fish diseases, as a whole, requires an understanding of all common diseases: why they arise, how to recognise them and how to deal with them. The following sections describe current knowledge on the diseases of local farmed marine foodfish.

2 Common diseases of marine food fish farmed in Singapore

2.1 Figures VIII/1–19 show the common diseases of marine food fish. They are:-

2.1.1 Ectoparasites

(Figures 1–11)

1. Ciliates (Figures 1–7)

   1. Cryptocarvon irritans immature trophont (1000X).

   2. Cryptocaryoniasis in estuarine grouper, artificial transmission, initial stage with white spots.

   3. Cryptocaryoniasis in seebass, normal above, diseased below with cogenitally missing dorsal fin.

   4. Cryptocaryoniasis in golden snapper, balding due to loss of skin from head region.

   5. Brooklyenella sp. showing parallel lines of cilia.

   6. Brooklynelliasis in estuarine grouper with extensive skin damage and subcutaneous bleeding.

   7. Trichodina sp. (1000X), ventral view showing ‘teeth’ and lateral cilia.

2. Others (Figures 8–10)

   8. Monogenean gill parasite Diplectanum sp. (100X).

   9. Isopod parasite Nerocila sp., adult female (2.6X)

   10. Isopod parasite Nerocila sp., male phase (16X).

2.1.2 Bacterial diseases (Figures 11–12)

11. Bacterial fin rot bacteria comprising Myxobacter sp, Vibrio sp, Gram-positive cocci etc. (Gram stain 1000X).

12. Bacterial fin rot in newly imported seabass.

2.1.3 Fungal diseases (Figures 13–15)

13. ‘Saprolegniasis’ aseptate fungus in lesion (1000X).

14. ‘Saprolegniasis’ of seabass.

15. Ichthyosporidosis of red grouper, showing ‘pothole’ lesion on head.

2.1.4 Viral disease (Figures 16–17)

16. Seabass experimentally infected with lymphocystis virus

17. Histological section of lymphocystis tumour, showing bubble-like lymphocysts (100X).

2.1.5 Diseases of unknown aetiology (Figures 18– 19)

18. Swin bladder syndrome in estuarine grouper breeder. Note also abnormal colour of liver.

19. Exophthalmus (Popeye).

3 Laboratory methods of identifying diseases

3.1 Ectoparasites

3.1.1 Organisms which live on the external parts of a living host (ie deriving nutrients from the host) and give nothing in return are known as ectoparasites. A parasite, whose demands on the host may be so great as to cause disease may be regarded as a pathogen.

3.1.2 Ectoparasites that are common in locally cultured marine food fishes are the protozoans, flatworms and isopods.

3.1.3 Examples of ectoparasites to be found are:

1. Protozoans, usually the ciliates e.g. Cryptocaryon irritans, Trichodina sp. and Brookynella sp. common on gills and body surface of grouper and seabass.

2. Monogenean trematodes eg Diplectanum sp., common on the gills of grouper (Epinephelus tauvina) and seabass (Lates calcarifer). Diplectanum sp. found on cultured Epinephelus tauvina was been described by Kabata in 1985.

3. Isopods e.g. Nerocila sp on gill, body surface and mouth of young seabass and marine red tilapia hybrid (Oreochromis).

4. The isolation of an ectoparasite for examination is covered in Practical VIII/1.

5. Staining of parasites is not usually practised in the laboratory because fresh specimens are good enough for temporary mounting with glycerine-alcohol (5%) to be identified. Specific stains and staining procedures are adopted for some parasites. The following procedure in Table VIII/1 is good for the staining of monogeneans:

Table VIII/1

Staining of monogeneans*

1. Place the specimens in water for 15 minutes to release eggs from their uteri.

2. Relax the worms in 70°C water.

3. Fix in AFA (alcohol-formalin-acetic-acid), 70% ethyl alcohol or Bouin's solution.

4. Downgrade to water and stain with Gower's Carmine.

5. Differentiate slowly in 0.5% HCl in 70% alcohol for 1–2 hours, depending on the size of specimen.

6. Gower's Carmine stains precisely, giving an unusual red colour with a bluish tinge.

* Kabata, 1985 and Roberts, 1978. This will be carried out if samples are available.

3.2 Bacterial pathogens

3.2.1 Bacterial pathogens are responsive for a very high proposition of mortalities in cultured marine food fishes, and occur in conjunction with other stress factors.

3.2.2 Most bacteria do not cause disease. However, as they multiply very fast and are opportunistic, that is, if a fish is stressed, wounded or heavily infested with ectoparasites, they are predisposed to pathogenic bacteria.

3.2.3 The most common bacterial pathogens of cultured marine food fishes are Vitrio and Mycobacterium, gram-negative and positive rods respectively (Vibrio Parahaemclyticus. V. algynolyticus and Myocobacterium sp.

3.2.4 The method of preparing tissue smears and staining procedures are shown in Tables VIII/2, 3 below:

Table VIII/2

Method of preparing of tissue smears for staining

1. Dissect the fish to expose the kidney which is to be found on the dorso-posterior wall of the body cavity.

2. Sample one loopful (using 3mm loop size) of tissue from the kidney and spread it evenly on a microscope slide.

3. Heat fix the smear by passing the underside of the slide over a flame.

4. Stain the smear with gram stain or Kinyoun acid-fast stain (see below):

4 Therapy of fish diseases

4.1 Definition

Therapy (also treatment) is the means employed in effecting the cure of the disease that fish have contracted. It is an interaction of three factors, namely, the virulent pathogen, the susceptible fish and the therapeutic agent or medication.

4.2 Principles of therapy

4.2.1 As in human diseases, the therapy of fish diseases with various medications and chemotherapeutic agents is only for the purpose of buying time, not for killing 100 percent of the disease organisms present. Medications are used to check the disease organism, retard its growth or even kill the pathogen but, in the end, it is the protective mechanism (immune system) of the fish that must overcome the disease-producing organism if the treatment or cure is to be successful. Fish farmers must keep this basic principle in mind every time a treatment is considered, if it is to be a success.

4.2.2 Prior to the treatment, we should ask the following questions so that a decision can be made on whether or not to proceed with the treatment:

1. What is the prognosis ie. the probability of a successful treatment?

2. Is it economically feasible to treat the fish?

3. Can the fish withstand the treatment considering their condition?

4. Does the loss rate and the disease present warrant treatment?

4.2.3 If treatment is warranted, the following four factors will first have to be considered before the treatment is carried out:

1. The water

   The volume of water in the treatment tank or rearing unit to be treated must be known before any treatment is contemplated. An overestimation of the water volume means too much of the therapeutant will be used. This could probably kill all fish or result in excessive cost. An underestimation of the volume means that insufficient therapeutant will be used and the pathogen will therefore not be controlled. Other water quality factors such as temperature and turbidity will affect the efficacy of the therapeutants because oxidation would take place or other bacteria may be present. Sufficient dissolved oxygen could be ensured by providing strong aeration. This is especially the case for formalin treatment. The estimated optimum fish biomass for treatments in tanks is 4 kg/m3 and 8 kg/m3 for treatments in canvas bags under field conditions. This would also depend on fish species and size and the size of tanks or bags (see Practical VIII/2, Mass treatment of fish under field conditions).

2. The fish

   Fish of different species, age and condition react differently to the same drug/chemical. Certain fish species or size are much more sensitive to a particular drug/chemical than are other species. Fish which have been weakened from stress may even succumb to treatment attempts before the pathogen eg. potassium permanganate at 10 ppm for 5–60 minutes is very effective in removing Lernaea in freshwater fish, while the same dosage and duration could be lethal to marine fish like grouper and seabass from experience.

   If a new chemical/drug is introduced, it is advisable to test the drug/chemical on a small number of fish before treating the entire lot.

3. Chemical/drug (therapeutant)

   The toxicity of the chemical/drug to a particular species of fish to be treated should be known. The effect of water chemistry on the toxicity of the chemical should also be understood. Some drugs break-down rapidly in the presence sunlight, while the mixing of chemicals may enhance or intensify the toxicity of the component chemicals (synergism) eg. a mixture of 0.45 ppm malachite green and 150 ppm formalin is more effective in treating Cryptocaryon irritans than 150 ppm formalin alone.

4. The disease

   Although this factor appears to be self-evident, it is one which is widely disregarded. Most chemicals used in the treatment of fish diseases are expensive and are generally effective only against certain groups of disease-causing organisms. Use of the wrong chemical or drug will usually mean that several days to a week may pass before one realises that the treatment was not effective, during which time large numbers of fish may be unnecessarily lost.

4.3 Methods of treatment

4.3.1 Various methods of treatment and drug application have been adopted in the control of fish diseases. There is no specific method that is superior to others, as this depends on the specific situation faced. However, treatment can be divided into three main types:

1. Addition of drugs/chemicals to water eg. nitrofurazone and formalin.

2. Addition of drugs/chemicals to feed eg. oxytetracycline, furazolidone.

3. Application of drugs/chemicals directly to fish.

1. Addition of drugs/chemicals to water

   1. Dip

      A strong solution of a chemical is used for a relatively short period of time. This method can be dangerous because the solutions used are concentrated. The difference between an effective dose and a killing one can be very narrow. However, when handling small numbers of fish, this is one of the most effective method of treatment. Eg. using a 300 ppm of formalin to treat cryptocaryoniasis in grouper. This high dosage may kill or at least damage the gill.

      Fish are usually placed in a net and dipped into a strong solution of the chemical for 15–45 seconds depending on the type of chemical, concentration and the species of fish being treated. In the culture of marine fishes in netcages, this method is used during transfer, and for sorting and culling of fingerlings for the purpose of sanitization or prophylaxis.

   2. Flush

      This method is fairly simple and consists of adding a stock solution of a chemical to the water supply of the unit to be treated, and then allowing it to flush through the unit. It is popular method in hatcheries and raceways. An adequate water flow must be available so that the chemical can be completely flushed through the unit or system within the predetermined period of time. In the field, canvas bags or floating tanks which are used to hold fish under treatment have seawater pumped to flush the medication by overflowing. However, this method is not to be encouraged since the excessive use of the chemical may pose a potential danger to the ecosystem.

   3. Prolonged treatment in water

      There are two types of prolonged treatment; a short term bath and indefinite prolonged treatment.

      Bath

      The required amount of chemical or drug is added directly to the water in the rearing or holding unit and left for a specific period of time. The drug/chemical is then quickly flushed with fresh seawater. Several precautions must be observed with this method of treatment to prevent serious losses. Although a treatment time of 1 hour may usually be recommended, the fish should always be observed throughout the treatment period, and at the first sign of distress such as gasping at the surface, fresh seawater should be added quickly.

      The bath treatment may reduce dissolved oxygen to the point where the fish are stressed and when losses actually occur. The more fish per unit volume of water the more likely this problem will develop. Aerators of same type must be installed in the treatment unit to ensure an adequate oxygen supply for the fish.

      Extreme caution must be used to ensure that the drug/chemical is evenly dissolved throughout the unit being treated to prevent the occurrence of undissolved lumps or particles or ‘hot spots’ of the drug/chemical. Fish may be killed or severely damaged by swimming through these ‘hot spots’. On the other hand, fish that avoid these ‘hot spots’ may not be exposed to a high enough concentration for the chemical to be effective. The problem of ‘hot spots’ is easily solved by dissolving the chemical well in a small bucket of water before distributing the solution with a scoop over the surface of the tank or canvas bag in which treatment is to take place.

      In treatments carried out at fish farms, the bath water can be tipped out from the canvas bag or pumped out. In both, the fish should be held in an inner netcage to facilitate water evacuation.

      Indefinite prolonged treatment

      This method is usually employed in treating fish in ponds or culture tanks where a low concentration of chemical is applied and allowed to dissipate naturally. This method is not practical for fish culture because water through-flow has to be restored fairly early. If tanks or canvas bags are used, frequent change of medicated seawater may be required, because seawater easily fouls if the treatment has to last for more than four hours. This makes the treatment an expensive and tedious one. For example, `vibriosis' in grouper requires prolonged treatment of nitrofurazone, usually until the red lesions healed.

2. Addition of drugs/chemicals to feed

   1. In the treatment of some diseases, the medication must be fed or in some way introduced into the system of the sick fish. This can be done by either incorporating the medication in compounded feed or by weighing out the correct amount of drug, putting it in a gelatin capsule and then introducing it into the fish by force feeding. In the case of large carnivorous fish like grouper and seabass, the gelatin capsule filled with drug can be stuffed into the mouth of whole trash fish before feeding it to them. Force-feeding is applicable to small and large fish alike.

   2. As oral application of therapeutics is based on body weight, a standard unit of treatment is given in terms of grammes of active drug per unit weight of fish per day or variations of this. Medicated feeds contain levels of medication calculated to deliver the desired dose expressed in terms of fish weight.

3. Application of medication directly to fish

   1. This is the simplest way to ensure that medication is introduced into the system. The exact dosage can be administered and the interference of the environment can be largely eliminated. However, this method requires the use of qualified and experienced personnel as the fish can be stressed from improper handling. In intensive culture like fish farming in netcages where large numbers of fish must be rapidly medicated, this direct method is of limited use. It is useful in the treatment of small and/or valuable stocks such as spawners and ornamental fishes. Medication is usually introduced into the fish by injection.

   Injection

   This technique of direct application requires that the fish be first anaestnetized. Generally, there are two routes of injection:

   Intraperitoneal injection (IP) is done by introducing medication into the body cavity. The IP injects the medication into the fish body cavity. The point of injection varies with the anatomy of the fish. For grouper, this is between the lateral line and anus. The needle is inserted until the tip penetrates the body wall. This is recognised by the sudden lack of pressure when inserting the needle. As soon as the tip of the needle is in the body cavity, the required amoung of medication is injected and the needle then withdrawn.

   Intramuscular injection (IM) effects the administration of medication through muscle tissue. IM injections are given in the muscular portion of the fish, lateral to the dorsal fin. The syringe and needle should be held on a line parallel to the long axis of the body and at about a 45° angle. The needle is inserted to a depth of about 1/4 to 1/2 inch and the medication slowly injected directly into the muscle tissue of the back. The injection must be done slowly to reduce back-pressure that would otherwise force the medication out of the muscle through the needle puncture.

   IP is more effective as it distributes the medication directly into the viscera.

   Topical application

   This is an external treatment. The medication is applied on to the affected area of the skin by swabbing with a cotton wool or painting with a brush. Swabbing with 0.1% potassium permanganate solution, tincture of iodine or its substitutes, or with malachite green can be made for fungal infections and focal skin inflammation caused by bacteria.

4.4 Treatment of fish at floating fish farms

Mass treatment of fish is covered in Practical VIII/2. This is done for prophylactic or therapeutic reasons, while sanitisation is done for imported fish fry/fingerlings.

5 Sanitisation guidelines for imported fish fingerlings

5.1 Need to sanitise imported fish fingerlings

1. In 1987, Singapore imported 30.6 million fish fingerlings and prawn fry for mariculture. The imports were valued at S$2.7 million* . The main fish species imported are the seabass, Lates calcarifer, (3.0 million fingerlings or 9.8% of the total import, valued at S$2.0 million, or 74% of the total value) and the grouper, Epinephelus tauvina, (0.9 million fingerlings or 3% of the total import, valued at S$0.3 million, or 11% of the total value). 26.7 million shrimp fry (87% of the total import) valued at S$0.3 million, or 11% of the total value). gives the breakdown.

2. Mortality of imported fish is considered to be the single most imported cause of loss to fish farmers. An estimated 75% of the stocks lost during farming occurs at the early stage, and may be attributed to stress from transportation. This loss occurs in the first three weeks after arrival and stocking in netcages. Fingerlings are presently air-freighted into Singapore. Except for the seabass which is available from commercial hatcheries, fish are caught from the wild and are therefore subject to capture and holding stress prior to export. They are further stress during transportation and adaptation to a new environment.

3. Some stress effects have been observed in newly imported fish. Severe irreversible shock is occasionally encountered during the release of fingerlings from their packings. This may result in mortalities of as high as 50%. Physiological failure probably accounts for the mortalities observed in the first few weeks. The conditions producing stress are mainly those of transport, high ammonia, nitrite and carbon dioxide levels which precipitate altered organ function. For example, the gills of imported fish often produce excessive musus which interferes with oxygen uptake, while dehydration often results from extensively damaged skin and causes breakdown in kidney function. Under such stress, the fish will refuse to eat. This weakens them further and consequently predisposes them to infection.

*1 S$2.1 = US$1.00

5.2 Sanitisation

1. Sanitisation improves immediate post-importation survival by reducing parasite and pathogen loads; restricts the entry of parasites and pathogens into local waters; accelerates the healing of wounds before shipment.

2. Sanitisation can involve the treatment of fish prior to shipment (pre-shipment sanitisation), during transportation (trans-shipment sanitisation) and after arrival at the farm (on-farm sanitisation), but the feasibility of this depends on the reliability of the handling agents concerned.

3. Sanitisation trials with grouper fingerlings imported from the Philippines have shown that mortality can be reduced through sanitisation, while unsanitised fish have been observed to be more susceptible to protozoal and bacterial attacks.

5.3 Methods of sanitisation

Table VIII/4 gives sanitisation procedures generally employed. On-farm or a combination of all 3 methods described have been found to be more effective.

Table VIII/4

Methods of sanitisation

1. Preshipment sanitisation

   This is done at source. The fish agent is asked to bathe the fish to be exported in 10 ppm acriflavin sea water for half an hour. The fish are held in 20-litre capacity styrofoam boxes each containing 18 litres of sea water, and at a biomass of 20 – 30 g per litre (150 × 3 g fish per box. The water is well-aerated during treatment and changed completely after the treatment.

2. Transhipment sanitisation

   Fingerlings, starved for 2 days are packed at 170 – 200 fish of 3 – 4g size fish per litre for seabass (biomass of 75 – 100 g/1) and 750 – 35 fish of 0.3 – 30 g size fish per litre for grouper (biomass of 38 – 132 g per litre) depending on size and transportation. Clear plastic bags (0.08 mm thick, 100 cm × 65 cm) are used, each containing 7 – 8 litres of filtered sea water into which is dissolved nitrofurazone at 10 ppm. Oxygen is bubbled into the packing water to inflate the bag at 50 – 70% by volume of the gas. The bag is secured by double rubber bands. An outer plastic bag of similar specifications is used to give double layers and secured with rubber bands. The fish consignment in this way is continually bathed in nitrofurazone during transportation. Inflated bags measure 50 cm in diameter and 15 – 20 cm in height. They are packed 1 to a styrofoam box (38 cm × 38 cm × 19 cm or 48 cm × 36 cm × 32.5 cm) with lid. The boxes are lagged with newspaper to minimise heat entry.

3. On-farm sanitisation

   On arrival at the farm, all bags of fish are floated in sea water to acclimate the fish to ambient temperature (28 – 30°C) before on-farm sanitisation in tanks. Fibreglass tanks (60 cm × 60 cm × 28 cm, 100-litre capacity) containing 75 litres of well-aerated, unfiltered sea water are used for on-farm fish sanitisation which comprises a 100 ppm formalin bath of 1 hour, followed by a 30 ppm nitrofurazone bath for 4 hours. Fish are stocked in the tank at 4 – 10 per litre (biomass of 10 g/litre). All water is discarded after the formalin bath and the same tanks replenished with new, unfiltered sea water for the nitrofurazone treatment.

5.4 Beneficial effects of fish sanitisation

1. Improved survival

   It has been observed that survival of unsanitised imported grouper fingerlings, two weeks after arrival, and in floating netcages is generally low, ranging from 18 – 55%. Sanitisation can improve survival from 1.1 – 2 times. The survival in seabass is better, ranging from 60 – 80% for unsanitised consignments, and improving by 1.2 times in sanitised consignments. These improvements in survival are significant to farmers as fish supply is seasonal and prices high.

2. Reduction of ectoparasites

   It has been observed that sanitised fish are generally cleaner than unsanitised ones having less ectoparasites and are sometimes even devoid of them.

5.5 Limitations in fish sanitisation

1. The effect of sanitisation depends on fish. initial condition and size. Fish which are initially stressed or are too small and weak for transportation will easily die with or without sanitisation. Stress from transportation and handling can negate the beneficial effects of sanitisation, and their handling during sanitisation itself can add on to this stress and adversely affect the consignment. Sanitisation should therefore be practised with care, involving minimal handling.

2. It is also important to realise that once-only sanitisation will not guarantee good survival, and that good farm management involving observation and quick decision making is important.

6 Quarantine

Quarantine is to ensure that the consignment is clear of transmittable diseases before its release into local waters. It involves the holding of fish under controlled conditions in tanks over an approved period and checks for diseases. Quarantine is not practised at present because fish fingerlings sourced from the wild or from hatcheries are not certified for health and would therefore all have to be quarantined. The frequency and volume of imported live fingerlings and fry are high and complete control cannot as yet be assured. The quarantine operation may further compromise fish survival since it involves handling and transfer from tanks to farms.

7 Diagnostic Methodology

Diagnosis is the art or the act of determining the nature of a disease. To diagnose a disease quickly and efficiently might seem a difficult and formidable task. It would be made simple if certain general guidelines and rules are followed. Below is a description of diagnostic methods commonly adopted by fish health workers.

7.1 Recording of case history

Fish farmers who report the disease case, are usually interviewed by the Fish Health personnel for the purpose of filling up the Fish Examination Report Form (of which Tables VII/5–7 are examples).

7.2 Post-mortem examination

7.2.1 Selection of specimens for examination

Systematic specimen selection is important for an accurate evaluation of the disease problem. Three categories of fish should be collected. They are the clinically asymptomatic fish (the ‘healthy’ fish), moribund fish and fresh mortalities within 30–60 minutes of death. The collection method of each category varies with the culture method used. In netcage culture conditions, the ‘healthy’ fish are normally not collected unless it is very necessary for comparison during necropsy. This is because they can only be properly sampled if the netcage is hauled up. However this might impose further stress and damage to the ailing stock. Collection of specimens from a raceway or tank is comparatively easier if the water therein is clear enough for the fish to be observed. Specimens can be taken out with a long-handled net.

7.2.2 Necropsy

Necropsy should be performed as soon as the specimens have been collected. ‘Healthy’ fish are to be examined first so as to recognize the normal tissue appearance under that particular culture condition. Fresh mortalities are then necropsied and any lesions should be taken note of. Moribund fish are to be examined last. At this juncture, a provisional diagnosis can usually be drawn. Additional specimens can be taken if specific or special examination is required.

b   Necropsy consists of three basic procedures:

1. External examination

2. Internal examination and

3. Record of findings

1. External examination

   This begins with close observation of the fish skin for evidence of external parasites or lesions such as ulcers, raised scales, reddened fins etc. Next, gross examination of the gills should be made for parasites and abnormal colouration. A wet mount of the gill tissue can be made by killing the fish, cutting out one gill arch and placing it in several drops of filtered seawater between a microscopic slide and cover slip. This fresh, unstained tissue can then be examined microscopically for lamellar changes and parasites.

2. Internal examination

   This begins by making a ventral midline incision from between the pectoral fins posterior to the anus. The view of the viscera can be expanded by removing the body wall on one side of the fish by means of an arching incision from immediately anterior to the anus, dorsally to the spinal column and then anteriorly and ventrally, to meet the ventral incision at the base of operculum. The viscera is thus exposed. The internal organs are observed in situ for abnormal colouration, positioning or enlargements. Abnormal fluid in the body cavity should also be noted at this time and examined for bacteria.

   Following in situ examination, the liver, spleen, pancreas stomach, intestines and gonads can be removed as a unit. This is accomplished by incising the oesophagus and hepatic ligament anteriorly and the intestine posteriorly. These organs should then be separated and examined individually for abnormalities.

   The removal of these organs exposes the swim bladder and kidney for examination. The swim bladder should be examined in situ, and then removed to allow complete access to the kidney.

   The only remaining organ to be examined is the brain. It can be removed intact by incising the epaxial muscles at their attachment to the skull and peeling the roof of the cranium away from the brain with a pair of small, heavily constructed forceps.

   Last of all, the skeletal muscle should be incised and examined.

3. Record of findings

   The final step in necropsy is the recording of all the observations made on the Fish Examination Report Form. Photographs of any significant gross lesions may also be helpful in the diagnosis of difficult cases in the future.

7.3 Histopathological examination.

7.3.1 Histopathological examination is needed in cases of infections, nutritional, neoplastic and endoparasitic diseases, as these are undetectable during post-mortem examination. Samples should be taken from tissues with lesions or suspected lesions and fixed in either 10% formal saline or Bouin's fixative. The tissue should be collected after gross internal examination. Each piece of tissue should not exceed 6 mm in thickness for adequate fixation, and the volume of fixative should be at least three times the tissue volume. Samples of tubular organs should be taken (with a sharp blade) as cross-sections before the organ is opened for internal examination. This minimizes distortion in the microscopic sections, allowing the pathologist to critically judge the microscopic anatomy.

7.3.2 Fry and smaller fish not easily necropsied can be reserved for histopathological examination by opening their body cavity, making two incisions into the epaxial (dorsal) musculature on both sides, and puncturing the swim bladder. The whole fish can then be submerged in fixative.

7.4 Bacteriological examination

7.4.1 Isolation of the causative agent is essential for accurate diagnosis and treatment of bacterial diseases. Therefore, bacteriological examination should accompany all cases in which the history and gross lesions suggest such a disease. Aseptic necropsy technique is required to allow good bacteriological isolation.

7.4.2 Bacterial isolation from gills and any external lesions cannot be done aseptically. Attempts should be made to find out the bacterial flora of these sites to compare with the normal bacterial flora of seawater. This is always a laborious process and the causative bacteria cannot be accurately identified most of the time.

7.4.3 Isolation attempts are usually made from the abdominal fluid or internal organs. The kidney, which is a haemopoietic tissue is the organ chosen for bacterial isolation because it is a natural filter of pathogens. The kidney is also well-protected by the vertebrae and bacteriological sampling can be done aseptically. To prevent contamination by bacteria from the environment, the descaled body surface where the incisions have to be made should be sterilized with 75% alcohol.

Screening tissue smears can be made from internal organs on microscopic slides. These slide preparations should be heat-fixed and stained with Gram's stain or Kinyoun's acid fast stain (Tables VIII/2, 3) a specific stain for Gram-positive bacteria. Bacterial septicemia can be provisionally diagnosed with this simple and fast technique.

7.5 Virological examination

Isolation of a suspected virus is necessary for an accurate diagnosis and for imposing quarantine measures to prevent a viral epizootic. Virological examination is usually attempted when all other examinations fail to yield results or when the clinical signs are indicative of a viral attack. Cell lines from local marine food fish species are required for the isolation of virus from suspected fish. Attempts are presently being made to establish cell lines from local marine food fish. In the mean time, cell lines of temperate fish eg. BF-2 from blue fin have been used for routine isolation of virus have not met with success.

8 Fish examination is covered in Practical VIII/1

Practical VIII/1
Fish examination

1 Objective(s)

1.1 To assess health condition of farmed fish through gross external examination and estimation of ectoparasite load.

1.2 To post-mortem farmed fish and make an internal examination of organs.

1.3 To make a bacteriological examination of fish organs (kidney inoculum).

2 Materials

2.1 Live or moribund fish from commercial floating fish farms.

2.2 Dissecting set, trays

2.3 Microscopes

2.4 Bacteria culture media

1. Trypticase/Tryptose/Tryptone soy agar with seawater (TSA/SW) prepared as an agar plate.

2. Thiosulphate citrate bile salt agar (TCBS) prepared in plates.

2.5 Routine bacteriology facilities like spirit lamp, and inoculation loops.

3 Procedure

3.1 Examination of fish on-farm, and assessment of farming conditions.

This is the initial stage of a case study, and is done at the farm. Records are made according to Table VIII/5.

3.2 Gross external examination of fish

3.2.1 Visual assessment

1. Examine the fish you have collected for external clinical signs of disease and injury.

2. Record your findings in Table VIII/6.

3. Make an interim assessment of fish health condition at this stage.

3.2.2 Ectoparasite examination

1. Preparation of skin scraping

   1. Place a drop of filtered seawater on a microscope slide.

   2. Remove a small portion of skin and mucus from the side of the fish body, using a small metal spatula or scalped.

   3. Transfer the sample in the drop of seawater on the slide.

   4. Place a cover slip over the sample.

   5. Examine for ectoparasites under the microscope. Distinguish the ciliate protozoans especially. Count the numbers of ectoparasite per skin scaping.

   6. Make several scrapings, count and record the numbers of ectoparasite each time. Classify the ectoparasite if possible.

2. Preparation of gill lamellae

   1. Place a drop of filtered seawater on a microscope slide.

   2. Kill the fish by cranial incision.

   3. Remove 5 – 7 complete gill lamellae from the fish, using a pair of scissors to cut them out from the base.

   4. Transfer the gill lamellae in the drop of seawater on the slide.

   5. Place a cover slip on the lamellae and gently press to spread them.

   6. Blot the excess water with some blotting/filter paper.

   7. Examine for and count ectoparasites as in [a (vii)].

   8. Examine several samples of 5 – 7 lamellae/sample, count and record as before.

3.3 Post-mortem examination

3.3.1 External appearance of fish

Observe for any abnormalities (eg frayed fins, scale-drop, clubbed gills) of external organs and record your findings in Table VIII/7.

3.3.2 Internal appearance of fish

1. Dissect the fish according to the following methods:

Method of fish dissection

1. Kill the fish with a cranial incision before dissection.

2. Scale the fish along the area for dissection, with subsequent disinfection using a paper towel soaked with 75% alcohol.

3. Make a scalpel incision at the abdomen just above the cloaca. This incision should be deep enough to open the abdominal cavity without puncturing the intestine.

4. Starting at the incision, make a scissor cut in a crescent to a point just at the top of the gill.

5. Make a second scissor cut from the incision along the midline to the bottom of the gill.

6. Reflect the cut sides of the abdomen to expose the internal organs.

2. Make a drawing of fish internal organs (consult your lecturers for recognition of fish parts). Observe for any abnormalities (eg bleeding or discoloured organs, especially the liver, and excessive fat) and record your findings in Table VIII/7.

3.3.3 Bacteriological examination (kidney inoculum)

1. The most common site for bacterial culture in fish is the hind kidney. This organ is usually located along the ventral side of the backbone and is protected from cavity contamination by a protective membrane.

2. Make a kidney inoculum on the TSA/SW and TCBS plates provided according to the following method:

Method of kidney sampling for bacteria

1. Pull the internal tissues forward towards the gills and cut free from the fish.

2. Reflect the swim bladder and visceral organs

3. The kidney is apparent along the vertebrae as a dark, reddish organ.

4. Take a loop of kidney tissue with a sterile inoculating loop.

5. Streak the kidney tissue on TSA/SW and TCBS plates.

6. Mark the agar plates for easy identification.

7. Incubate the plates with the inoculants at room temperature (air-conditioned, 23°C) for 24 – 48 hours.

8. Observe the gross morphology of the different colony types that are produced.

3. Record your findings in Table VIII/7.

3 Interim assessment of fish from gross external examination and ectoparasite occurrence.

3.1 What was the general condition of the gills, judging from its appearance?

3.2 Could the fish have been particularly stressed by ectoparasites?

Practical VIII/2
Mass treatment of fish under field conditions

1 Objective

To go through procedure for on-farm mass treatment*¹ of grow-out fish, using floating canvas bags.

2 Plan

Mass treatment of fish in canvas bags

3 Materials

3.1 600 seabass of 300g mean weight.

3.2 2 (2m × 2m × 2.5m) canvas bags (2m × 2m galvanised iron frame to assist in sinking the canvas bags -optional, 2 (2m × 2m × 2m) hapa (knotless) netcages of mesh size 9mm (5.16").

3.3 Small (0.5 HP) seawater pump, air stones, air valves, air blower, generator.

3.4 Dissolved oxygen meter with temperature sensor.

3.5 Scoop nets, containers, telecounters, weights for netcage and canvas bags.

3.6 Formalin and nitrofurazone.

3.7 Microscopes, dissecting set, trays

4 Procedure for the mass treatment of fish in floating canvas bags.

4.1 Fish exemination before and after treatments.

The procedures in Practical VIII/1 apply. Record data in Table VIII/6 attached.

4.2 Preparation of fish and facilities for treatment

1. Starve the seabass (300g) for 24 hours before treatment.

2. Set the two canvas bags in vacant raft units, if possible adjacent to the netcage containing the fish to be treated.

3. Make sure that the air pockets in the immersed canvas bag are expelled. This can be facilitated by weighting the bottom corners of the bag to allow it to stretch open.

4. Tie weights to the bottom corners of the hapa netcage and set it in the canvas bag.

5. Set up the generator and air supply system (blower, air tubing and stones) to supply air to the water in the canvas bag. This is necessary if:

   1. fish density is high

   2. treatment is prolonged

   3. the species is sensitive to low dissolved oxygen concentration

   4. the species uses up dissolved oxygen rapidly

   5. the chemical/drug used in treatment reduces oxygen in water eg. formalin

6. Take dissolved oxygen reading of the seawater in the canvas bags.

7. Estimate the mean weight of fish by bulk weighing, and transfer them from their netcage (about 300 fish) into the hapa netcage within the canvas bag, following the procedure detailed and covered in IV P/6 Netcage operation and maintenance. Note the number of fish transferred.

8. Calculate the volume of seawater in the canvas bag and the biomass of fish in this volume.

*¹ Formalin (100 ppm × 1/2 hour) and nitrofurazone (15 ppm × 4 hour).

4.2 Treatment of fish

1. Calculate the weight of formalin required for the volume of seawater in the canvas bag to give 100 ppm and nitrofurazone to give 15 ppm (see method of calculating dosages in VI p/8 Treatment trials and tolerance tests).

2. Formalin (100 ppm × 1/2 hour) treatment of fish is in canvas bag as follows :

   1. Mix the weighed out formalin well in a bucket of seawater.

   2. Spread the formalin solution evenly in the canvas bag.

   3. Bathe the fish in the 100 ppm formalin for 30 minutes ensuring adequate aeration.

   4. Monitor dissolved oxygen during treatment from time to time and observe fish behaviour.

   5. Seawater-formalin is released by lowering the canvas bag which is then filled as previously, or by pumping out. In the latter case, fresh seawater is pumped in to replace the vacated seawater.

3. Nitrofurazone (15 ppm × 4 hour) treatment of fish is done in canvas bags as follows :

   1. Mix the weighed out nitrofurazone well in a bucket of seawater and distribute the solution evenly in the canvas bag.

   2. Bathe the fish in the 15 ppm formalin for 4 hours.

   3. Monitor dissolved oxygen and fish behaviour as on the previous treatment.

   4. Since treatment is prolonged, it may be necessary to siphon out detritus*2 from the bottom of the canvas bag from time to time. However, this may not be necessary since fish have been starved and sick fish will usually not have fed.

   5. At the end of the treatment, pull the canvas bag to one side and re-set the original netcage in the same raft unit.

   6. Pull out the hapa netcage to transfer the treated fish into their original netcage.

   7. Remove the canvas bag, wash and rinse with sodium hypochlorite and dry before folding and storage.

   8. Re-tie the netcage containing the fish into position.

*2 Siphoning can be done by inserting a UPVC cylinder (0 10 cm × 1.5m long) which is closed at one end into the sea adjacent to the canvas bag. Detritus is siphoned into the container which will have lowered the seawater level outside the bag for siphoning to be effected.

5 Record fish examination data in Table VIII/5–7

+1 Record according to the following scale:
(+) few (1–10 nos/sample)
(++) moderate (10–35 nos/sample)
(+++) abundant (> 35 nos/sample)

D45/naca-8

Figure VIII/1

Cryptocaryon irritans immature trophont (1000X).

Figure VIII/2

Cryptocaryoniasis in estuarine grouper, artificial transmission, initial stage with white spots.

Figure VIII/3

Cryptocaryoniasis in seabass, normal above, diseased below with cogenitally missing dorsal fin.

Figure VIII/4

Cryptocaryoniasis in golden snapper, ‘balding’ due to loss of skin from head region.

Figure VIII/5

Brooklyenella sp. showing parallel lines of cilia.

Figure VIII/6

Brooklynelliasis in estuarine grouper with extensive skin damage and subcutaneous bleeding.

Figure VIII/7

Trichodina sp. (1000X), ventral view showing ‘teeth’ and lateral cilia.

Figure VIII/8

Monogenean gill parasite Diplectanum sp. (100X).

Figure VIII/9

Isopod parasite Nerocila sp., adult female (2.6X)

Figure VIII/10

Isopod parasite Nerocila sp., male phase (16X).

Figure VIII/11

Bacterial fin rot bacteria comprising Myxobacter sp, Vibrio sp, Gram-positive cocci etc. (Gram stain 1000X).

Figure VIII/12

Bacterial fin rot in newly imported seabass.

Figure VIII/13

‘Saprolegniasis’ aseptate fungus in lesion (1000X).

Figure VIII/14

‘Saprolegniasis’ of seabass.

Figure VIII/15

Ichthyosporidosis of red grouper, showing ‘pothole’ lesion on head.

Figure VIII/16

Seabass experimentally infected with lymphocystis virus

Figure VIII/17

Histological section of lymphocystis tumour, showing bubble-like lymphocysts (100X).

Figure VIII/18

Swim bladder syndrome in estuarine grouper breeder. Note also abnormal colour of liver.

Figure VIII/19

Exophthalmus (Popeye)