Acute respiratory distress in wild caught round goby (Neogobius melanostomus)

Heindrich N. Snyman, Michelle Wodzak, Kerri Nielsen, Deborah Pakes

Animal Health Laboratory, University of Guelph, Kemptville, ON (Snyman); University of Toronto, Scarborough, ON (Pakes, Wodzak, Nielsen)

AHL Newsletter 2019;23(4):11-12.

In mid-July, 30 adult wild round goby (Neogobius melanostomus) were collected from Lake Erie along the shore of Kingsville, ON. These fish were destined to establish a research colony at a local University. At the research facility, fish were housed in groups of 6-7 individuals in re-circulating 20 gallon tanks with aerators, rocky substrate, and PVC tubes for hides. Water temperatures were maintained at 20-24°C and fish were exposed to a ~ 1% salt bath upon intake. Shortly after introduction to the tanks, fish started to exhibit rapid breathing, irregular darting behavior, and clustering around the intake flow pipe. This was followed by progressive worsening of the respiratory distress and a decrease in appetite ultimately culminating in the mortality of 5 fish ~ 1 week after initial introduction. Water quality parameters were within normal limits (ammonia levels were 0-0.25 ppm, nitrite levels <1ppm, and water pH 7.8-8.2). Three fish were fixed whole in formalin and submitted to the Animal Health Laboratory for analysis.

Histological evaluation revealed widespread and marked hyperplasia of the lamellar gill epithelium as well as the epithelium lining branchial and oral cavity in all three fish with extensive blunting and fusion of secondary lamellae. Scattered throughout the affected foci were numerous irregular, round to oval, 50 to 200 µm diameter, subepithelial ciliated protozoal cysts that contained prominent 1-2 µm thick pale outer eosinophilic hyaline walls. Cysts contained a single large 20 × 120 µm crescent-shaped, deeply basophilic eccentric macronucleus and abundant finely granular to vacuolated basophilic cytoplasm with large amounts of phagocytized erythrocytes and cellular debris. The size, histomorphology, and particularly the subepithelial localization are highly characteristic of Ichthyophthirius multifiliis.

I. multifiliis is a well-known ciliated protozoal fish pathogen and causes the characteristic fish disease referred to as “ich” or “white spot disease”.  It is most common in ornamental tropical freshwater fish but can affect any freshwater fish species. Along with temperature, stress plays a major role in epidemics and the associated stress of handling, capture, and shipping likely played a significant role in the initiation of disease in this colony. Mortality rates are highest in warmer water (15 - 25°C) but significant disease can still occur in temperatures as low as 10°C.

Treatment of this pathogen is often very problematic as it exists as both free-living (tomont) and parasitic (trophont and theront) life stages. Briefly, the ich trophont (feeding stage) feeds in a capsule formed in the skin or gill epithelium which forms the characteristic “white spots” that lends this disease its name. After feeding, the parasite breaks through the epithelium and is free in the water column during which time it forms a capsule and divides (tomont stage). The tomont capsule is very sticky and can adhere to nets, substrate, plants, etc. Tomites subsequently break through the capsule and form motile, infective theronts which then reinfect the gill epithelium. Disease is further complicated by a particularly rapid and prolific rate of replication with the whole life-cycle being completed in as little as 3-8 days at optimum temperatures of 23-24°C and with a single tissue trophont being able to produce up to 2000 re-infecting theronts.

Figure 1. A Histology of gills with widespread lamellar epithelial hyperplasia with lamellar fusion (asterisks). B Sub-epithelial ciliated protozoal cysts (arrows) contain a characteristic crescent-shaped macronucleus (asterisks) with abundant phagocytized erythrocytic and cellular debris.


1. Dickerson HW. Protozoan and Metazoan Infections: Ichthyophthirius multifiliis and Cryptocaryon irritans (Phylum   

      Cilliophora). In: Woo, PTK, ed. Fish Diseases and Disorders. Wallingford, Oxfordshire, UK: CAB International, 2006.

2. Frasca S et al. In: Terio KA et al, ed. Pathology of Wildlife and Zoo Animals. London, UK: Elsevier, 2018. 

Diagnosing enteric disease in backyard chickens

Emily Martin, Marina Brash, Emily Brouwer and Kate Todd

Animal Health Laboratory, University of Guelph, ON

AHL Newsletter 2019;23(4):12-14.

Veterinarians that treat backyard flocks can encounter cases of enteric disease due to a number of different etiologies. Due to the length of the intestinal tract, enteric signs can range from gasping, distended crop (crop stasis, crop impaction, pendulous crop), diarrhea, pasty vents, and emaciation. Morbidity and mortality will be variable. How do you approach diagnosing these flocks?

On physical examination, examine the oral cavity for plaques and the crop for distention and consistency (empty/full, doughy/firm). Examine the vent for evidence of fecal or urate concretions, hemorrhage and trauma (Fig. 1A). Birds have two types of normal droppings: brown and well-formed feces with a cap of white urate material or loose brown cecal droppings. Loose cecal droppings need to be differentiated from diarrhea. Knowing the normal appearance of chicken droppings will help detect diarrhea. The colour of droppings can provide clues to disease; for example: green due to increased bile related to anorexia, brilliant yellow due to histomoniasis, orange associated with sloughing of the intestinal mucosa, and white due to excessive urates from the kidneys. Other changes to look for include the presence of undigested feed (malabsorption), blood or abnormal smell.

If there is no mortality but there are changes in the droppings, consider collecting fecal material for parasitology and bacterial culture. If there are oral plaques, swabs (viral transport media) of the plaques and choanal slit could also be collected to perform PCR tests for specific diseases (e.g. poxvirus, Trichomonas). You can watch a video on how to collect samples with these swabs at: (must be registered and signed in to to view this page)

If there is mortality present, this is an indication for postmortem examination and more intensive sampling. Postmortem examination could be done at a clinic or birds could be shipped to the Animal Health Laboratory. If birds are shipped, please refer to the AHL website for a PM submission form ( and packaging instructions (

Clinic postmortem: Cut the commissure of the beak and open the oral cavity to look for oral plaques. Rule outs include wet pox, candidiasis, trichomoniasis, impacted fine feed and vitamin A deficiency. Collect pieces of the oral mucosa, including plaques, for PCR and histology. 

Cut down the esophagus into the crop and examine the crop wall and content. Collect pieces of crop for histology. Open the bird and use the gizzard to reflect organs and cut across the esophagus-proventriculus junction to remove the intestines. Open the proventriculus and gizzard and pull out the content for examination. Birds allowed outside will often eat fibrous material that can impact the gizzard and crop (Fig. 1B) or may ingest toxic plants or foreign items (Fig. 1C) that can block or perforate the intestines. Examine the external surface of the small intestines for petechial hemorrhages (e.g. coccidiosis) then cut open most of the length of the small intestines to examine for the presence of worms and irregular mucosal surfaces; the latter may indicate necrotic enteritis (Fig. 1D). If there is increased fluid or irregular mucosal surfaces, tie off pieces of the intestines and submit for culture. For histology, collect pieces of duodenum (with pancreas), jejunum (mid small intestine), and ceca (cut transversely through the ileum and both ceca). Open the colon and cloaca to examine content. In younger birds, examine the bursa to evaluate size and collect pieces for histology and PCR.

Table 1: List of chicken enteric disease tests available at AHL.

DISEASE (by lab section)





Parasitology Lab





Intestinal parasites (coccidia, nematodes, tapeworms




fecal float, histology

Trichomonas gallinae





Clinical pathology





Crop wash




cytology, histology

Virology Lab





Poxvirus (nodules, crusts)





Newcastle/APMV-1 (Avian Paramyxovirus 1)





Avian Influenza/Influenza A





Infectious Bursal Disease (IBD)





Bacteriology Lab





Bacterial culture (feces, intestines)





Mycology/Fungal culture










Mycotoxin (feed analysis)





Sampling Summary:

NOTE: If screening a flock for disease, up to 5 swabs or tissues can be pooled for PCR testing. PCRs are generally the preferred test as they target specific diseases and can be done quickly.

Live Bird Diagnostics:

  • Choanal slit/tracheal swab (viral transport medium) is the optimal sample for PCR (larger number of tests available).
  • Photos of affected birds can be submitted to AHL along with the case history to

Dead Bird Diagnostics:

NOTE: If a postmortem is refused by the owner, fecal material and swabs from the choanal slit could still be collected for PCR.

  • Postmortem is the key test for lesion identification and appropriate sample collection.
  • Histology is an option to screen tissues for lesions or determine the etiology of specific lesions such as nodular structures (bacterial, fungal, neoplasm). Collect a wide variety of tissues to place in formalin, including all major organs: lung, liver, spleen, and kidney in addition to intestine.
  • PCR tests can be performed on swabs or tissues. If you are sending tissues, package each tissue type separately and label. Do not mix intestinal tissue with organ tissue.
  • Bacterial/fungal culture can be performed on gel swabs or tissues.

For more information on enteric diseases in chickens, please refer to the OAHN website:

Reportable diseases in Canada include Salmonella Pullorum, Salmonella Gallinarum, Avian Influenza and Newcastle Disease. If high mortality and/or clinical signs lead you to suspect Avian Influenza or Newcastle Disease – quarantine the flock and phone CFIA!

                    A Vent trauma. B Gizzard impaction with fibrous material. C Foreign body (nail) perforating the gizzard. D Necrotic enteritis (Turkish towel mucosa).           

Figure 1. A Vent trauma. B Gizzard impaction with fibrous material. C Foreign body (nail) perforating the gizzard. D Necrotic enteritis (Turkish towel mucosa).

Incidental cysticercosis in a pet rabbit

Emily Brouwer, Marina Brash

Animal Health Laboratory, University of Guelph, ON

AHL Newsletter 2019;23(4):15.

A four-year-old, male castrated, Dutch cross domestic rabbit presented to the submitting veterinarian for cystotomy to remove uroliths. On presentation, the rabbit was in good body condition with no significant findings on preanesthetic physical examination or blood tests. The rabbit was routinely anesthetized, and the abdomen surgically approached via a 3 cm long caudal ventral midline incision. Upon entering the abdomen, three abnormal cystic structures were identified. All cystic structures were free-floating in the abdomen and had no visible vascular attachments. The three cysts were removed, and a biopsy sample of the bladder wall was collected for histology (Fig. 1A). Grossly, these cysts were thin-walled, contained clear fluid, and each had a thickened white asymmetrical linear focus in the wall. Each cyst was fixed in formalin, and routinely processed for histologic examination. Histologically, the cysts contained flocculent eosinophilic fluid surrounded by a thick, fibrous capsule. Corresponding to the white area noted grossly are cestode larvae with convoluted hyaline cuticle overlying palisading epithelial cells and loose mesenchyme containing faint basophilic crystalline concretions (calcareous corpuscles). Scolices had prominent refractile hooks (Fig. 1B).

These cysts were determined to be histologically characteristic of encysted tapeworm larvae. In rabbits, this is most likely Cysticercus pisiformis, the larval stage of Taenia pisiformis. Rabbits are the intermediate host for a variety of carnivore tapeworm species, but the most likely species (T. pisiformis) has the dog as the definitive host. Dogs become infected with the adult tapeworms and release ova into the environment through the feces, which are then ingested by the rabbit. It is unknown if this rabbit had access to an outdoor space that was shared with canids, or if the household had dogs as well. Typically, these infections are incidental, where cysts are identified during abdominal surgery or postmortem. Less frequently, there can be clinical signs related to mass-effect if there is a heavy burden. Lethal cysticercosis is rare in rabbits, but has been described in cases of extensive hepatic parenchymal involvement.

A Free-floating cysts identified in the abdomen (right). The tissue on the left is a biopsy of the urinary bladder. B. Encysted tapeworm larva with prominent hooks (H&E 2X magnification).

Figure 1. A Free-floating cysts identified in the abdomen (right). The tissue on the left is a biopsy of the urinary bladder. B. Encysted tapeworm larva with prominent hooks (H&E 2X magnification).


1. Barthold SW et al. Parasitic diseases. In: Barthold SW et al, eds: Pathology of Laboratory Rabbits and Rodents, 4th ed. Chichester,UK: Wiley Blackwell, 2016.

2. Graham-Brown J et al. Lethal cysticercosis in a pet rabbit. Vet Rec Case Rep, 2018; 6e000634.doi:10.1136/vetreccr-2018- 000634.