How to Recognize and Interpret Clinical Signs of Fish Illnesses
World Small Animal Veterinary Association World Congress Proceedings, 2015
N. Saint-Erne, DVM, CertAqV
Technical Service Veterinarian, PetSmart, Inc., Phoenix, AZ, USA

Learning Objectives

Learn how to examine a fish and perform clinical tests to diagnose commonly encountered fish diseases. Learn what treatments are used for specific diseases of ornamental fish.

When working with fish patients, just like with any other animal, collecting a good history is as important as the physical examination of the fish. Water quality parameters must also be tested, as poor water quality often is a significant factor in fish illness. Then, a live moribund fish should be examined when possible, or a freshly dead specimen. If the fish has been dead for more than a few hours, even if refrigerated, the chance of accurate diagnosis is diminished. Always wet the hands first before handling fish, or wear smooth latex gloves, to prevent damage to the slime coating of the skin of the fish.

After initial physical examination, biopsy samples should be taken. If fish anesthetic is available, add it to fresh dechlorinated water to anesthetize the fish prior to the biopsy. Have another container of dechlorinated water without anesthetic available in which to awaken the fish. Place the fish on a wet towel or chamois cloth. Be sure to keep the fish wet and handle it gently. Using a blunt blade or spatula, or even a plastic microscope coverslip, gently scrape a small sample of mucus off the body of the fish.

Place this in a drop of water on a microscope slide and cover it with the coverslip. If there are skin or fin lesions, take a sample from the margin of the lesion and prepare it the same way. Next, snip a small section of the caudal or other fin rays using small, sharp scissors, such as iris or suture scissors. Also take a sample of a few gill filament tips. Place these on a slide and prepare as with the skin scraping. Examine these samples microscopically at 40–400x magnifications.

Parasite infestation is a common cause of morbidity and mortality in aquarium fish. There are many parasites that affect fish, with protozoa being the most frequently encountered. Some of the species of protozoa that can cause significant fish loss include ciliated protozoa: Chilodonella, Tetrahymena, Epistylis, Ichthyophthirius, Trichodina; and flagellated protozoa: Ichthyobodo (Costia) and Spironucleus (an intestinal parasite). These protozoa parasites often cause similar signs, so accurate diagnosis requires microscopic examination of biopsy samples.

Monogenean trematodes are one-host flatworms that can affect the skin and gills of fish. The commonly found species are Dactylogyrus and Gyrodactylus. Dactylogyrus (gill flukes) are found on the tips of the gills and occasionally on the skin of fish. They cause gill filament hyperplasia resulting in hypoxia. Signs include rapid respiratory movements, fins held against the body, and flashing (scraping body on rocks or other objects in the aquarium). Gill tip biopsy will reveal the flukes upon microscopic examination. Gyrodactylus (skin flukes) occur mainly on the skin and fins, but occasionally are found on the gills.

Macroparasites of tropical aquarium fish include intestinal roundworms (Camallanus, Capillaria), tapeworms, leeches, anchor worms (Lernaea) and fish lice (Argulus). The female anchor worm develops an anchor-shaped head, which it embeds into the skin of the host fish, leaving its worm-like body extending from the skin. This causes a reddened sore, which may become secondarily infected with bacteria. Fish lice have a stylet mouthpart that repeatedly pierces the skin of the fish and releases toxic secretions. They feed on the tissue fluids released by this trauma.

Samples for bacterial culturing are taken from lesions or organs using a culture swab. Use sterile technique to avoid contamination from environmental or water microorganisms. Bacteria are grown in culture media at 20–25°C for 3–5 days. Growth is identified to bacterial species and then tested for antibiotic sensitivities. Many species of aquatic bacteria are resistant to commonly used antibiotics, so sensitivity testing is very important in selecting appropriate therapy. Treatments for pathogens such as bacteria and fungus also require different medications than for protozoa and helminthes, so accurate disease diagnosis is important to selecting the correct therapeutic agents.

Drawing blood samples from larger fish, especially koi, can be done from the caudal vein below the spine in the caudal peduncle. Use a 1-ml tuberculin syringe with a 22 or 23-gauge needle of appropriate length. A butterfly catheter can be attached to the syringe to facilitate handling of the needle separately from the syringe. Fill the hub of the needle with a drop of lithium heparin to prevent the blood from clotting. This is preferable to ammonium heparin or sodium heparin, but they can also be used for hematology testing. The ammonium and sodium heparins will affect those blood values if used in samples for serology or electrolyte testing. Ethylenediaminetetraacetic acid (EDTA) is not recommended to be used to prevent blood clotting in fish blood samples, as it may cause erythrocyte lysis.

To collect a blood sample from an anesthetized fish, the needle is inserted at an angle pointing craniodorsally from the ventral midline of the caudal peduncle until it hits the vertebrae. Withdraw the needle slightly, and it should be in the caudal vein. Light aspiration on the syringe plunger should be applied to collect the blood. The total volume of blood in most fish makes up about 5% of its body weight (50 ml/kg). Up to 20% of the total blood volume can safely be removed in fish that are not excessively ill. This allows for 0.1 ml of blood to be removed from a 10-gram fish (20% x 50 ml/kg x 10 g = 0.1 ml), 0.5 ml of blood removed from a 50-g fish, or 1 ml from a 100-g fish. Larger blood samples are usually not necessary, even though more blood could be drawn from a large fish such as a koi.

Diagnosing diseases in fish, especially internal problems, can be made easier with the use of radiology, ultrasound, and endoscopy equipment already present in the veterinary clinic. These advanced diagnostic techniques can be used with fish as easily as with other pet patients. Techniques such as CT and MRI, where available through specialty practices or veterinary colleges, can also be of great assistance in diagnosing abnormalities of the cardiovascular system, intestinal tract, gas bladder, and abdominal organs in larger fish, such as a koi with abdominal distension. These techniques, along with microscopic analysis of fluid and tissue biopsies, are very helpful in the diagnosis of fish diseases.

Radiographs of fish can be taken successfully with standard veterinary radiology equipment. The radiographs can be taken without anesthesia by briefly restraining the fish in a sealed plastic bag with a small volume of water. The fish in the plastic bag can be placed directly onto the film plate or digital sensor, and the bag taped down if necessary to hold the fish in the correct position. Fish can also be anesthetized and then taken out of the water and positioned for radiographs. Gas bladder (swim bladder) abnormalities, spinal deformities, abdominal masses, and occasionally ingested foreign objects can be evaluated using radiology. Abdominal viscera are not easily distinguishable in a radiograph without a contrast medium. A flexible rubber catheter can be inserted orally to place barium or iodinated contrast medium into the stomach or intestines to perform contrast studies on the intestinal tract. The dosage for barium is 5 to 10 mL/kg body weight, and the iodinated medium is dosed at 1 to 2 mL/kg. Be careful not to leak barium into the oral cavity and onto the gills, which could impair oxygen diffusion through the gills.

Ultrasound imaging can be performed on fish confined in a small container of water, as the water serves to couple the transducer to the fish's body, eliminating the need for ultrasound gel. Transducers of 5 to 10 MHz work well for visualization of internal organs at depths up to 13 to 20 cm into the body, with lower frequency transducers producing images at greater depths of tissue penetration. If not waterproof, the transducer can be placed inside a plastic cover (e.g., plastic bag, examination glove, or condom) for protection. The transducer can be held several centimeters away from the fish if it is in the water, and the transducer repositioned until the desired image is obtained. Motion imaging can be used for guided tissue biopsy collection, abdominocentesis (coeliocentesis), or pneumocystocentesis.


1.  Miller S, Mitchell M. Ornamental fish. In: Manual of Exotic Pet Practice. St. Louis, MO: Saunders-Elsevier; 2009.

2.  Roberts H. Physical examination in fish. In: Fundamentals of Ornamental Fish Health. Ames, IA: Wiley-Blackwell; 2010.

3.  Saint-Erne N. Advanced Koi Care. 2nd ed. Glendale, AZ: Erne Enterprises; 2010.


Speaker Information
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Nick Saint-Erne, DVM, CertAqV
PetSmart, Inc.
Phoenix, AZ, USA

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