Introduction
Clinical signs of illness displayed by fish are often not specific enough to determine a definitive aetiological diagnosis. Common techniques a veterinarian needs to be conversant with include taking a history, examining tank/pond conditions, testing the water quality, checking for clinical signs of disease, microscopic examination of wet mounts of skin mucus, fin and gill biopsies, sampling for bacterial cultures, faecal examination, phlebotomy, exploratory laparotomy, necropsy and histopathology.
This presentation will familiarise the participants with the practical aspects of veterinary medicine, ranging from point-of-care diagnostics, through to sampling for laboratory testing.
Water Quality Analysis
Fish rely on the qualities of water for many biological processes, and their health depends on several key parameters that include water temperature, ammonia (NH3), nitrite (NO2-), nitrate (NO3-), pH, carbonate hardness (KH, also known as alkalinity), general hardness (GH, also known as permanent hardness), dissolved oxygen (DO) and salinity. There are more than 40 thousand species of fish, and each fish species has specific environmental requirements to live. Understanding these requirements is necessary to provide good diagnosis and management recommendations. Table 1 provides parameters for the common biotypes.
Table 1. Water quality parameters for popular groups of aquatic animals
| |
Units
|
Freshwater
tropical general
|
Freshwater
Amazonian
|
Freshwater
African
|
Koi & goldfish
|
Marine reef
tropical
|
|
Temperature
|
°C
|
24–28
|
24–28
|
24–28
|
20–24
|
24–25
|
|
Ammonia
|
mg/L
|
< 0.2
|
< 0.2
|
< 0.2
|
< 0.2
|
0
|
|
Nitrite
|
mg/L
|
< 0.2
|
< 0.2
|
< 0.2
|
< 0.2
|
0
|
|
Nitrate
|
mg/L
|
< 50
|
< 50
|
< 50
|
< 50
|
5–10
|
|
pH
|
Units
|
6.5–7.5
|
6.5–6.8
|
8.0–8.2
|
6.5–7.5
|
8.2–8.5
|
|
KH
|
mg/L
|
70–100
|
70–100
|
125–225
|
70–150
|
125–225
|
|
GH
|
mg/L
|
100–200
|
80–150
|
300–500
|
150–200
|
Calcium (380–450 mg/L) &
magnesium (1200–1300 mg/L)
tested separately
|
|
Salinity
|
g/L
|
0
|
0
|
0
|
0–5
|
30–35
|
Pre-Physical Examination
Taking a complete history is required, noting the species affected, speed of onset, morbidity, mortality, changes in husbandry, previous treatments, and whether there had been new introductions. During this time, the veterinarians should be observing the fish from afar, observing for clinical signs of illness displayed by fish, such as clamped fins, reduced activity, staying at the bottom, piping (gasping for air), flashing (scraping flanks against substrate or sides of tank).
Physical Examination
Aeration should be given as soon as the fish is brought to the clinic or laboratory. The right-sized container should be provided so the veterinarian can evaluate behaviour, swimming pattern and overall condition of the fish. External lesions might be visible at this point as well. Ulcers, masses, cloudiness of skin (mucus production), erosion, irregularities of the fish, exophthalmia (pop-eye), abdominal swelling, skeletal deformities, protruding scales (dropsy). Colour change may serve as indicator of fish health. Note that these are nonspecific gross and clinical signs, and diagnostic tests are vital.
Microscopic Examination of Wet Mounts
Parasites are best observed live, by preparing wet mounts and examined via light microscopy. It is very important that wet mounts are examined immediately (at 40 to 400x magnifications) so the parasites are still moving. Things to observe for under the microscope include motility (sessile versus free-swimming), pattern of motility (e.g., ciliated protozoa tend to move in a smooth gliding motion, whereas flagellated protozoa tend to be quick and erratic), morphology (e.g., Trichodina are circular, Ichthyobodo are pear-shaped) and size (e.g., Tetrahymena are relatively uniform in size, whereas the size of Ichthyophthirius tomites can be variable). The procedure for preparing wet mounts follows.
To sample skin mucus, using the edge of a coverslip or glass slide held at approximately 45° angle, gently scrape a small sample of mucus from the body in a caudal direction. Mucus should be collected just from behind the pectoral fin, at the base of the dorsal fin and at the caudal peduncle. These are areas that the fish cannot scrape (scratch) and dislodge the parasites. Place this in a drop of water on a microscope slide (wet mount) and cover it with a coverslip. In small fish, a coverslip may be used to collect skin mucus samples, and in larger fish a glass slide or the foil pack of a scalpel blade can be used.
Next, a gill biopsy of the tips of several primary lamellae should be examined for parasites, bacteria and fungi. Hold the fish in a net, then use the tip of the scissors to lever the operculum (gill cover), and hold the operculum open by sliding your thumb gently towards the operculum, wedging it in place. Hold the scissors blades perpendicular to the primary filaments and sample only the distal tips of the primary gill filaments (up to 20% of the length of primary filaments). Only a small sample is required (3–5 filaments). It is better to collect smaller gill biopsy samples from multiple fish rather than a larger amount from any one fish. Be careful not to induce iatrogenic haemorrhage. Fine, blunt-tipped scissors with a serrated cutting edge is recommended. Again, place these on a slide with a drop of water. Sedatives may be used in fish that are large, venomous, or are difficult to restrain.
A faecal sample can be collected from the fish or from the water from the transport container and microscopically examined for bacteria, protozoa, and helminth ova. Faecal samples can be stored in plastic bags or other sealed containers for up to three days without preservatives if air is excluded from the container. Preserve faecal samples in 10% formalin (the same concentration as for submitting histopathology samples). If coccidia are suspected, place the faeces in 2.5% potassium dichromate solution. For examination for intestinal protozoa other than coccidia (particularly Giardia spp. or amoebae), the sample should be fixed using sodium acetate - acetic acid - formalin (SAF) fixative. This fixative also preserves helminth eggs.
Blood Tests
Blood samples can be drawn from the caudal vein of large fish. It is located ventral to the spine and can be accessed from a ventral approach with an anaesthetised fish in dorsal recumbency. Use a 1- to 3-ml syringe with a 22-gauge needle of appropriate length that has been pre-treated with heparin anticoagulant. The needle is inserted at an angle, craniodorsally from the ventral midline of the caudal peduncle until it contacts the vertebrae. Draw back on the plunger to collect blood. A minimum of 0.5 ml is required for biochemistry testing; however, smaller amounts can be prepared for qualitative haematology analysis as blood smears and for blood culture (for bacteria).
Necropsy
The number of fish to submit for diagnosis depends on the size, age, client (farmer vs. pet owner). The key is to ensure you and your client(s) understand the risks in taking diagnostic samples if it is not a terminal fish (live vs. dead fish post-diagnosis). Note that dead fish are of limited diagnostic value. Fish may be anesthetised for diagnostic sampling if necessary.
All diagnostic utensils should be on hand and a comfortable and clean area for necropsy should be ready. Containers for subsamples (virology, bacteriology, molecular, histology, etc.). Upon exposure of internal organs, great care needs to be taken to avoid contamination (rupture of gastrointestinal tract is a common source). The general anatomy of fish is not dissimilar to other animals; however, there is some terminology the aquatic veterinarian needs to be familiar with for describing lesion locations and diagnostic sampling.
Samples for bacterial and viral culture are prioritised and sampled with aseptic technique. Samples typically include blood-filtering organs like spleen, kidney, liver and heart. For some other agents with tissue tropism, the eye and brain are also useful.
For histopathology, the gills and gut are prioritised since these organs autolysed rapidly. As a general rule, sample every organ for histopathology. If in doubt, sample it.
Bacteriology
Testing confirms presence of organism. It does not mean it is causing disease. Bacteriology sampling should always be taken first with proper aseptic technique to minimise contamination. Additionally, samples for bacteriology testing must be taken prior to treatment with antibiotics to be valid. The sites for sample collection may vary. Effectively, sample where the lesion is.
Samples for bacterial culture from skin wounds should be taken at the margin of an acute lesion. If enteritis is suspected, then take a sample from the gut. For cases of septicaemia, the best site for sampling is the posterior kidney. It is the target organ in many diseases due in part to the extensive blood supply, and it traps more than 70% of bacteria because of the large number of phagocytes lining the renal sinusoids. It is also the organ least likely to be iatrogenically contaminated when conducting the necropsy, because it is situated in the retroperitoneal region, abutting the ventral aspect of the spinal column and covered by the swim bladder. In dead specimens, the swim bladder is removed carefully to reveal the kidney.
In the laboratory, a pipette is inserted through the membrane and kidney material is aspirated. In the field, a transport swab can be used to obtain a sample. The best type of bacterial transport swab contains Amies media with charcoal. The charcoal helps with improved survival of aquatic organisms and fastidious ones. It is practical to have a small tip for more accurate aseptic sampling of organs in small fishes. Although the samples in the transport swabs can be stored for up to 2 weeks at room temperature, it is advisable to store them in the refrigerator and/or send them straight away. Delays can alter the proportion of bacteria isolated (e.g., skin swabs can be overgrown with Klebsiella spp.). Ideally, the swabs should be processed at the laboratory within 24–48 hours of collection.
If swabs are not used, kidney tissue can be collected aseptically. It may be appropriate to pool samples in lots of 5 individuals to reduce costs. The samples would be homogenised in the laboratory. Fresh kidney samples should be maintained at 0–4°C (not frozen), and they must be set onto media within 48 hours of collection (preferably within 24 hours).
In very small, dead fish, the skin is decontaminated with antiseptic, dried with a sterile gauze swab and the back of the fish is cut, taking care not to enter the coelomic cavity. A sample can be taken from the area of haemorrhage (the kidney is well vascularised).
Smears on glass slides are also useful for the bacteriologist as an aid to identifying the bacteria. Slides can be prepared by spreading a thin layer of macerated kidney tissue or of the lesion, and heat fix the slides. Special stains are then applied to the slides to determine the bacteria's staining properties.
Conclusion
It is hoped that the reader will agree that sampling protocols between widely different species are not that dissimilar. Aseptic techniques and prioritising sample collection are sometimes required. Field diagnostics and sampling for laboratory analysis are complementary and help in disease diagnosis and management in all situations.
References
1. Loh R. Fish Vetting Essentials. Perth, Australia: Richmond Loh Publishing; 2011.
2. Loh R. Fish vetting techniques and practical tips. Perth, Australia: Richmond Loh Publishing; 2014 (DVD).