11.1.1 Candida sp. Infection (Crop Mycosis)

Infection with an overgrowth of Candida sp. (synonyms candidiasis, crop mold, thrush) is common and affects the upper digestive tract in a wide variety of birds, especially young birds. The most common species is Candida albicans, although a variety of other species (C. tropicalis, C. glabrata, C. parapsilosis, C. krusei, C. lusitaniae) have been identified and these all have similar growth characteristics in culture (Sabouraud dextrose agar, 37 °C for 3–5 days) [2, 3]. Candida infection of the upper digestive tract has been well characterized in commercial chickens and turkeys, and described in pheasants, quail, guineafowl, peafowl, and partridges [3–6]. Although candidiasis is common, it is generally not a major clinical problem and usually occurs secondary to another underlying disease or condition; in fact, it is often caused by an extended use of oral antibiotics administered in the feed or drinking water.

Risk factors for Candida overgrowth include prolonged administration of antibiotics, malnutirion, poor flock hygiene, and immunosuppression from stress [2]. Most antibiotics in feed or drinking water have a label to use for less than 4–6 days. The infection most often affects the oral cavity, esophagus, and crop.

Candida is a ubiquitous yeast organism and can be part of the normal microflora; overgrowth is usually controlled by the normal bacterial microflora. When other diseases, such as those of the respiratory or digestive tract, are treated long term with antibiotics, the normal inhibitory microflora of upper digestive tract can be reduced to allow for yeast overgrowth. The organism can also be transmitted in contaminated drinking water, and infected adults that feed their precocial young can directly transmit the organisms [3].

Clinical signs are nonspecific because birds are often ill from some unrelated condition. Birds may appear unthrifty or have other more significant diseases being treated with oral antibiotics. Lesions appear grossly as a fine, white “pseudomembrane” or lining in the crop, upper esophagus or oral cavity (Figure 11.1). In the digestive tract, Candida growth starts as budding yeasts (blastospores) on the surface of the mucosa and progresses as branching, septate chains or pseudohyphae that extend into the deeper tissue (Figure 11.2). The condition may not be recognized until affected birds are examined by necropsy. The white membrane can be scraped or peeled off the mucosa. Gross and histopathologic lesions will confirm the diagnosis, although fungal culture can also be considered if speciation is necessary. The crop lesions when observed in gamebirds should be differentiated from Capillaria infection (crop worms).

The condition is often reversible by eliminating the primary disease condition, removing antibiotics and improving husbandry, but specific treatments such as copper sulfate solution (0.05%) in drinking water for several days [7] or addition of nystatin (100 g/ton) to feed [2] have been described. Prevention of this condition focuses on good husbandry, sanitary conditions, and reduction of primary stressors and infectious diseases in the flock. Judicious use of antibiotics is warranted, especially in young birds.

11.1.2 Capillariasis (Crop Worms, Threadworm)

There are multiple species of nematodes (family Capillariidae) that infect the digestive tract of pheasant, quail, partridge, and guineafowl (Table 11.1). Crop and small intestine are most often affected depending on the species of Capillaria, which are speciated based on the (i) morphology of the genitalia of the adult nematodes and (ii) location of infection in the host [14]. The nematode ova are distinctly bioperculate.

The most common Capillaria in gamebirds are Capillaria (Eucoleus) annulatus and Capillaria (Eucoleus) contortus, which infect the crop. These nematodes are long and slender and less than 60 mm long. Capillaria are shorter and thinner than Heterakis and Ascaridia (cecal worms and roundworms). There is generally a 30‐day life cycle from ova to adult. Adults embedded in the crop mucosa or small intestine produce ova that are shed in the feces. The ova of most Capillaria species require an arthropod or earthworm intermediate host for transmission, but Chaetoceros contortus can also be transmitted directly through consumption of the ova [31]. C. annulatus and C. contortus are embedded in the crop mucosa in large numbers, and the affected birds can be depressed, weak and emaciated, often dying without premonitory signs. Birds might occasionally gasp if agitated because of respiratory difficulty. When pressing on the crop of an infected live bird, one might express white to gray opaque crop fluid into the oral cavity [15].

In mild infections the crop mucosa can be covered with a thin, white film, while in heavy infections the entire crop wall can be thickened and fluid‐filled, with a rough, irregular mucosa covered by a thick white, fibrinonecrotic exudate (Figure 11.3). The infection can extend beyond the crop into adjacent regions of the esophagus. In species affecting the small intestine and cecum, such as C. caudinflata and C. phasianina, respectively, the affected organ can be distended with mucus and have a tan pseudomembrane or pointpoint ulcers on the mucosa [15].

Diagnosis can be made by examining mucosal scrapings or washings under a microscope to view the nematodes and distinctive bioperculate ova (Figure 11.4) or by histopathology of the affected segment of the digestive tract. Capillaria nematodes have distinctive histologic morphologic features (coelomyarian‐polymarian musculature and bacillary bands in hypodermis), including the distinctive bioperculate ova in the reproductive tract. Capillarid adults, depending on the species of nematode, burrow into the mucosa of the crop or deep into the glands of the small intestine to induce necrosis and heterophilic to granulomatous inflammation [13, 15].

Fenbendazole, mebendazole, and tramisole (levamisole) as off‐label medications are effective treatment for Capillaria in poultry [32–34], including peafowl [35]. Infections are most severe in birds raised on soil or floor because of the greater opportunity for fecal–oral transmission. Raising smaller gamebirds (quail, partridge) on wire at early ages can decrease the buildup of Capillaria ova. Rotating the use of pens or moving the flight pens intermittently will decrease the buildup of ova in the soil. Keep the soil dry and well drained to decrease the number of earthworms. Intermittent tilling of the soil may help cover manure to reduce exposure to nematode ova.

11.1.3 Trichomoniasis

Members of the protozoal family Trichomonadidae, mainly Trichomonas gallinae and Tetratrichomonas gallinarum, are digestive tract parasites in birds; infections have been described worldwide, most commonly affecting passerines, pigeons, raptors, chickens, and turkeys, among others. The condition is known as “canker” in pigeons and doves and as “frounce” in raptors and passerine birds.

Trichomonad infections have been described in pheasants, partridge, quail, and guineafowl. First, T. gallinae is more pathogenic than Tetratrichomonas gallinarum [36]. Infections are often worse and more extensive in birds that have concomitant infections and are raised under poor conditions. Fecal–oral spread and consumption of contaminated water and feed are the likely routes of transmission, but some authors have suggested retrograde infection of colon as an alternative route [37], particularly with Te. gallinarum. Infected birds excrete live parasites as soon as 2 days post infection, as demonstrated by experimental infection [38]. Field surveys of wild bobwhite quail failed to find T. gallinae, suggesting that either the infection is more common in farm‐raised birds or the route of transmission is hindered in nature [39].

Clinical signs associated with avian trichomoniasis are loss of appetite, vomiting, ruffled feathers, diarrhea, dysphagia, dyspnea, weight loss, increased thirst, inability to stand or to maintain balance, and a pendulous crop [38]. T. gallinae has been well described in Columbidae (pigeons) with necrotizing lesions in the upper digestive tract. Te. gallinarum can also cause typhlohepatitis in galliform and anseriform birds [38].

Trichomonads have been described in a variety of gamebirds. Trichomonas phasiani has long been associated with mortality in young pheasants, with yellow, fluid‐filled ceca containing the protozoa [40]. Wichmann and Bankowski described young chukar partridges with fatal Te. gallinarum infection, as confirmed by wet‐mount preparations collected from multifocal necrotic lesions in the cecum and liver [37]. Other trichomonas infections have been described in quail. Tritrichomonas gigantica caused mortality in coturnix quail [41], and a unique Tritrichomonas species has been described in a single coturnix quail [42].

Trichomonads colonize specific regions of the digestive tract after infection. T. gallinae occurs in the mouth, pharynx, esophagus, and crop, with the parasite rarely found posterior to the proventriculus. T. gallinae causes inflammation of the mucosa and accumulation of caseous exudate, which can block the esophagus and subsequently kill the host through starvation (Figure 11.5). Trichomonads can survive in a moist environment for 4–5 days [39]. Lesions associated with Te. gallinarum occur most often in the cecum and colon and can be variable in extent, often manifested as dilated fluid‐filled intestines. Te. gallinarum has caused fatal typhlocolitis in the red‐legged partridge with histopathologic evidence of necrotic foci containing intralesional protozoa in the cecum, liver, and spleen [36]. It appears that in many instances, Te. gallinarum can be present without clinical signs or gross lesions, but the protozoa can contribute to disease in birds with other enteric infections, including Salmonella, Spironucleus, and Blastocystis sp.

Diagnosis of trichomoniasis is based on gross lesions and observation of the protozoa through histology or cytologic wet mounts. Histopathology of Te. gallinarum can be based on protozoa situated in the mucosal glands or there can be moderate diffuse lymphocytic infiltration of the cecal or colonic mucosa associated with a large number of protozoa within the lumen. Live trichomonads can be observed on direct microscopic observation of motile protozoa via wet‐mount preparation (mucosal scraping placed on a wet glass slide) from a live or recently euthanized bird. Sample material can be obtained via swabbing the cloacae for Te. gallinarum [43] or the oral cavity for T. gallinae [44]. Trichomonads appear as elongated, oval shapes, which move briskly. The wet‐mount sample smeared on a glass slide can be stained with Wright‐Giemsa to demonstrate protozoal detail [45]. The oval to pear‐shaped protozoa are 4–6 μm by 8–14 μm. They contain a single axostyle, a 1–2 μmdiameter parabasal body and multiple anterior flagella (Figure 11.6).

Treatment can be challenging because of the lack of approved antiprotozoal drugs. Various nitroimidazoles, including metronidazole, dimetridazole, ronidazole, and carnidazole, have been considered the standard treatment for avian trichomoniasis [38, 46, 47], but these products are no longer approved for meat‐type birds. Prevention should be the major focus. Trichomonads have limited survival in a dry environment. Because trichomonads can be harbored in wild birds, strict biosecurity should prevent entry of wild carrier birds into pens and buildings, particularly those harboring gamebird chicks. Routine cleaning and disinfection of brooding pens between grow‐outs are essential.

11.2.1 Dispharynx Nasuta (Proventricular and Gizzard Worm)

Dispharynx nasuta, a spirurid nematode, has been recognized in wild passerine and galliform birds, including upland gamebirds, for many years. It appears to be present most often in wild‐caught upland gamebirds, but the risk for domestic birds remains. Dispharynx has been reported in pheasants [21], partridges [48], quail [49], peafowl [50, 51] and guineafowl [52]. D. nasuta adults reside in the proventriculus, and less often the esophagus or gizzard, of infected birds. The life cycle is often indirect with an invertebrate intermediate host. Adult nematodes produce eggs that are shed in host feces.

Birds become infected by consuming an isopod intermediate host that has eaten the nematode ova. The most common intermediate hosts involved with the indirect life cycle are isopod crustaceans – pillbugs (Armadillidium vulgare) and sowbugs (Porcellio scaber). Larvae develop to the infective stage in the intermediate host within 26 days and are viable in this host for up to 6 months. When the isopod is ingested by a bird host, the infective larvae then escape and migrate/attach to the proventriculus or gizzard lining where they develop into adults within 27 days [53].

The parasitic infections are often subclinical, but if infection is severe with a high parasite load, the birds can become emaciated, weak, and anemic from blood loss in the proventriculus. The proventriculus is distended with mucus and the proventricular walls are thickened. Numerous 5–8 mm long, white, coiled nematodes can be observed adhered to the mucosal lining (Figure 11.7) [21]. In light infections, inflammation and hypertrophy of the mucosa occur. In heavy infections, the adult worms penetrate the mucosa, creating deep ulcers, glandular hyperplasia, and thickening of the mucosal layer.

Numerous classic broad‐spectrum anthelmintics are effective against Dyspharynx nematodes, including several benzimidazoles (albendazole, fenbendazole, flubendazole, mebendazole, oxfendazole, etc.), levamisole, and macrocyclic lactones (e.g., ivermectin) [31].

11.2.2 Tetrameridosis (Tetrameres sp.)

This condition encompasses nematode parasitism associated with three genera of the family tetrameridae: Tetrameres, Microtetrameres, and Geopetitia. Adult nematodes are often embedded in the proventricular glands, with or without associated clinical signs. The condition is characterized in both aquatic and terrestrial birds, including pigeons [54]. The nematode infection has been reported in gamebirds, including ring‐necked pheasants [21], partridges [17], quail [55, 56], guineafowl [57–59] and peafowl [60]. In the literature, the species of Tetrameres is not always determined, but T. americana and T. pattersoni have been reported in bobwhite quail [55]. Tetrameres fissipina was reported in ring‐necked pheasants [21] and T. numida is described in helmeted guineafowl [59].

Tetramerid nematodes show sexual dimorphism. Both the female and smaller male nematodes are often in copulation within the proventricular glands. In the life cycle involving terrestrial bird hosts, gravid female nematodes residing in the proventricular glands lay eggs that are shed in the feces and ultimately consumed by intermediate hosts, such as grasshoppers, cockroaches or earthworms, depending on the genus or species of nematode [54]. The cycle is completed following consumption of the intermediate host.

Because this parasite is relatively rare in gamebirds, clincal signs are poorly described; however, in the authors’ experience, these nematode parasites are usually a secondary, subclinical finding. Fecal flotation is generally not useful for diagnosis and necropsy/histopathology is suggested. Gross lesions of tetrameridosis are usually observed when the birds are necropsied for other primary conditions. The female nematodes appear as dark, red 2–4 mm diameter nodules within the wall of the proventriculus (Figure 11.8). Histopathology is confirmatory and reveals cross‐sections of female and male nematodes within dilated proventricular glands with or without glandular atrophy and minimal inflammation. The need for treatment is unlikely, but control should be based on reducing exposure to the intermediate host in the soil by maintaining dry soil and practicing insect control.

11.2.3 Miscellaneous Gizzard Conditions

Factors associated with gizzard erosion and ulceration in domestic poultry include bacterial infection, histamine, gizzerosine, mycotoxins, and vitamin deficiencies [61, 62]. Affected gizzards usually have a thickened, dark tan to gray lining with multiple linear erosions and ulcerations (Figure 11.9). Additionally, koilin erosion and ulceration is commonly observed in young gamebirds with paratyphoid Salmonella infection or pica/litter consumption, resulting in litter accumulation in gizzard. In these instances, the damage to the koilin layer is caused by bacterial overgrowth and direct invasion of koilin by bacterial colonies.

Gizzard koilin erosions associated with infection with fowl adenovirus 1 (FAdV‐1) have been reported in coturnix quail [63] and bobwhite quail [64]. In the latter report, the 2‐week‐old bobwhite quail were depressed with loose droppings and increased mortality. The only lesions were observed in gizzards. Histologically, gizzards are inflamed with multifocal koilin degeneration and fragmentation, epithelial degeneration and necrosis, and infiltration of inflammatory cells. Necrotic epithelial cells contained large, basophilic intranuclear inclusions, and FAdV‐1 was isolated from affected gizzards. The lesions resembled those reported in broiler chickens infected with FAdV‐1. Both vertical and horizontal transmission were reported as important routes for the spread of FAdVs [65].

Gizzard “impaction,” more accurately referred to as litter eating, is common in young, commercial gallinaceous birds that are brooded on litter, such as wood shavings, straw, or peanut hulls. Young, curious birds raised on sawdust, sand or small particle size straw may consume the litter to excess, resulting in accumulation of the coarse material in the gizzard (Figure 11.10). Litter consumption may not affect the health of the birds, but it can reduce feed efficiency and promote bacterial enteritis. Alternatively, litter consumption is often a necropsy finding in birds that have been ill with a digestive tract disorder or have no access to feed because of lameness or malfunction of the feeder system. In these cases, the consumption of litter is not the primary issue, but it is a symptom of a larger management issue.

11.3.1 Salmonellosis

A variety of bacteria from the Enterobacteriaceae family can affect the digestive tract of birds, but Salmonella is recognized as the most important aerobic bacterial pathogen of the gamebird digestive tract. Salmonella sp. are Gram‐negative bacterial rods that can infect all types of birds and mammals. In gamebird chicks, Salmonella most often infects the digestive tract and yolk sac. It can also be harbored in the intestine of adult birds with no clinical signs. Salmonella infection in gamebirds is primarily a problem of young birds, although the adults (breeders) can still be involved as carriers and periodic shedders of bacteria. The veterinary literature distinguishes between avian Salmonellae that are largely avian‐adapted serovars – Salmonella pullorum and S. gallinarum, the causes of pullorum disease and fowl typhoid, respectively – and 200 other serovars (paratyphoid species) of Salmonella that are not avian adapted and can occur in a variety of species. These avian‐adapted and paratyphoid Salmonella serovars will be addressed separately.

Fowl typhoid and pullorum disease are caused by two different biovars of S. enterica subsp. enterica serovar gallinarum, a Gram‐negative bacterial rod in Salmonella serogroup D (nonflagellated organisms with O antigens 1, 9, and 12) in the family Enterobacteriaceae. S. enterica subsp. enterica serovar gallinarum biovar gallinarum, which causes fowl typhoid, is usually abbreviated as S. gallinarum, and S. enterica subsp. enterica ser. fallinarum biovar pullorum as S. pullorum [66]. Recently, S. pullorum (pullorum disease) and S. gallinarum (fowl typhoid) were taxonomically assigned to a single serovar (S. pullorum‐gallinarum), but because the microorganisms are both biochemically and genetically distinguishable, they are often referred to as separate species and conditions.

Salmonella pullorum and S. gallinarum are nonmotile and tend to be specific for birds, while the motile paratyphoid serotypes can infect a wide variety of birds and mammals. These have been described in most upland gamebirds.

Salmonella pullorum and S. gallinarum (S. pullorum‐gallinarum) are relatively nonmotile and are generally considered to be species specific for birds. Disease with typhoid has been described in pheasants, partridges, quail, peafowl, and guinea hens, as well as chickens and turkeys [66–69]. S. pullorum‐gallinarum can be harbored in adult birds and cause little to no clinical signs, but these microorganisms can be vertically transmitted from the ovary of the hen into the egg and then to hatchling chicks. Chicks can appear normal at hatch, but they can transmit Salmonella laterally and experience heavy mortality during the next 3 weeks, with death in the early brooding stages. For reasons previously stated, both S. pullorum and S. gallinarum are regularly monitored in US commercial poultry breeder flocks that participate in the National Poultry Improvement Plan (NPIP) [66].

Clinical signs of pullorum disease, apart from sudden mortality of hatchlings, are nonspecific. In one study, bobwhite quail infected with S. pullorum were depressed with closed eyes, ruffled feathers, profuse white diarrhea, and 75% mortality from hatch to 16–17 days of age [69]. There is delayed resorption of the yolk sac in chicks, with inflammatory foci, sometimes forming large nodules, in liver, heart, kidney, spleen, pancreas, intestine, and articular joints [66]. Fowl typhoid affects both young and old birds. As with pullorum disease, chicks with fowl typhoid may be found dead soon after hatching while depression, decreased appetite, weight loss, dehydration, ruffled feathers, and watery to mucoid yellowish diarrhea can be observed in live birds. In some instances, breeders infected with S. gallinarum will have decreased egg production as well. In one study, Japanese quail chicks infected with S. gallinarum had enlarged livers and spleens with pale, necrotic foci as well as similar foci in kidney and heart. Histologic evidence of sepsis with necrotic foci and intralesional bacteria is observed in a variety of internal organs, particularly liver and spleen [70].

Diagnosis of pullorum disease can be confirmed by assessment of clinical signs, gross and histologic lesions and bacterial culture. The whole‐blood plate test is routinely used to assess breeder flocks for the presence of antibodies (reactors) to S. pullorum or S. gallinarum, which are antigenically similar. Salmonellae can be isolated by recommended procedures [71]. Veterinarians or diagnostic labs that suspect a flock is infected with pullorum disease or fowl typhoid should contact their NPIP office or state veterinarian immediately. There is no antibiotic treatment for pullorum or fowl typhoid as the entire exposed flock will face euthanasia in an eradication effort as determined by the state veterinarian.

There are over 2000 serovars of paratyphoid Salmonella and about 10% of these have been described in poultry. A variety of paratyphoid Salmonella has been described in gamebirds, guineafowl, and peafowl. For example, there are reports of Salmonella derby and S. anatum in chukar partridge chicks [72], S. typhimurium in bobwhite quail [73], S. typhimurium, S. kentucky, and S. enteritidis in golden pheasants in a zoological exhibit [74]. Gross lesions of paratyphoid Salmonella in gamebirds are most prominent in chicks and include dehydration, distended abdomens, enlarged liver and spleen, dark or hemorrhagic yolk sacs and increased mucus and frothy fluid in the intestine (Figure 11.11) [73]. Gamebird chicks can pile in the corners of the pen. Dead chicks often have enlarged livers and spleens with caseous, white to tan exudate in the cecum (Figure 11.12). Birds can have a variety of histologic lesions indicative of sepsis, including interstitial pneumonia, necrotizing hepatitis and splenitis, and fibrinonecrotic typhlitis.

Diagnosis of paratyphoid Salmonella infections is readily made on gross and microscopic lesions and culture of the causative agent. Antibiotic treatment should be based on culture and sensitivity, but few antibiotics are approved for use in poultry. Paratyphoid infections can be treated with antibiotics in feed or drinking water, but affected chicks will continue to die because they are usually not eating or drinking (“starve‐outs”). Walking the house or running the feeder may stimulate chick activity. Antibiotics can help the birds in the pen that are least affected and most susceptible to contracting the infection. Because of a diminishing supply of approved antibiotics, some operations are looking at alternative therapies such as essential oils, probiotics, berry extracts, and organic acids [75].

Strong biosecurity is key in preventing the infection from getting into the farms and flocks. Several species of paratyphoid Salmonellae, such as S. enterica serovar enteritidis [76], can enter the egg before hatch while others infect the chick in the pen through shell contamination or lateral transmission in feces. Employees who have contact with infected birds can introduce the infection into a clean flock. A comprehensive biosecurity program should cover all potential sources of poultry farm contamination. Sources of Salmonella contamination include the following [72, 77, 78].

1. 1. Infected breeder flock
   
2. 2. Soiled or manure‐contaminated hatching eggs
   
3. 3. Contaminated byproducts in feed
   
4. 4. Feces of other infected chicks
   
5. 5. Hatchery contamination
   
6. 6. Vectors: mice, rats, insects, and wild birds
   
7. 7. Dirty litter or poorly disinfected flooring, feeders, drinkers
   
8. 8. Young and old birds in close proximity
   
9. 9. Vehicle traffic (delivery and feeder trucks) on the farm
   
10. 10. Human foot traffic

11.3.2 Ulcerative Enteritis (Quail Disease)

Ulcerative enteritis (UE) or “quail disease” is a severe bacterial disease produced by Clostridium colinum that is highly contagious and produces high mortality in bobwhite quail (Colinus virginianus) [79, 80]. C. colinum is a Gram‐positive, spore‐forming, anaerobic bacterial rod and, based on 16S rRNA sequencing, it is closely related to C. piliforme, the causative agent for Tyzzer disease [81]. The organism is hardy in the environment. More recently, other Clostridia, such as C. perfringens type A and C. sordellii, have been determined to cause similar lesions in the intestines of quail [82, 83]. UE is diagnosed most often in bobwhite quail, and has been reported in other upland gamebirds (pheasants, grouse, partridges, California quail), as well as young turkeys and white leghorn pullets. UE appears typically between 4 and 20 weeks of age with a mortality rate of up to 50% or more in untreated flocks [84–86]. The lesions are characterized by multifocal discoid ulcers in the small intestine and multifocal hepatic necrosis.

The microorganism spreads rapidly from bird to bird via the fecal–oral route. It is shed in the feces of infected birds and is hardy, persisting in the soil or litter for several months. UE is rare in quail raised on wire. The infection can be spread in contaminated feces by flies.

Quail are usually 4–10 weeks of age at sudden death associated with white, watery diarrhea. Birds that do not die suddenly will be depressed with closed eyes, ruffled feathers, and bloody diarrhea. Infected birds are thirsty and will huddle around the drinkers. The course of disease is about two weeks and can result in nearly 100% mortality in bobwhite quail [87].

Necropsy findings are consistent. Crops can be distended with fluid. Affected birds have deep, punctate to discoid, tan to yellow ulcers along the small intestine (Figures 11.13 and 11.14). The ulcers often penetrate the entire wall of the intestine to result in peritonitis and adherence of intestinal loops. The liver may or may not have pale foci of necrosis on the capsule and on cut surfaces. Histologic lesions are further supportive of the diagnosis and in the intestine consist of severe multifocal mucosal to transmural necrosis with numerous inflammatory cells and large Gram‐positive bacterial rods. Peritonitis with serosal fibrosis can occur adjacent to the transmural lesions. The liver has acute, moderate to marked, multifocal hepatocellular necrosis with fibrin exudation and varying numbers of intralesional rod‐shaped Gram‐positive bacteria [86]. The flock history along with the gross and histologic lesions are usually diagnostic.

The microorganism can be cultured from liver and intestine, but culture of the agent from typical lesions can be challenging because of fastidious growth requirements [80, 88]. Culture can be important if one is differentiating UE (C. colinum) from necrotic enteritis (C. perfringens) [89]. Polymerase chain reaction (PCR) testing for C. colinum has been reported, but the assay is not readily available for laboratory diagnosis [90].

Clostridium colinum is sensitive to a wide range of antimicrobials including penicillin, oxytetracycline, monensin, and bacitracin [88]. Bacitracin is generally the antibiotic of choice at 100–200 g/ton of feed for 7–10 days, or in the water at 0.25–0.50 g per gallon of water for 7–10 days. Antibiotics may not be successful if birds are not eating or drinking; workers should walk the pens and run the feeders and drinkers to encourage bird activity. UE rarely occurs in quail raised on wire, although the microorganism can be spread by flies. Rotation of pens on a regular basis can reduce exposure to Clostridium spores.

11.3.3 Enteric Viruses

The role that viruses play in gamebird enteric disease is poorly characterized. In the authors’ diagnostic experience, the most common viruses detected by electron microscopy of the feces of pheasants, partridges, and quail are rotavirus, reovirus, and adenovirus. Calicivirus has also been associated with enteritis in gamebirds and will be described below.

Because commercial gamebirds are grown in a concentrated confinement setting prior to release, one would expect them to have enteric viral diseases that are similar to those of other intensively farmed poultry; however, studies on the viral agents producing digestive disease in gamebirds are limited. Additionally, in the field, these various enteric viruses are often found in association with other infectious agents, such as coccidia and Salmonella sp. Therefore, the role of these viruses may be additive at best.

To characterize the effect of enteric viruses in a diagnostic setting, it is essential to receive freshly dead or euthanized birds to reduce autolysis artifact in microscopic sections of digestive tract. Bacterial culture, virus isolation, electron microscopy on negatively stained feces, and, for some viruses, PCR analysis of intestine or feces are part of a thorough enteric virus workup.

11.3.3.1 Enteric Rotavirus

Rotavirus is a nonenveloped, icosahedral, double‐stranded RNA virus of the reovirus family. The virus replicates in and destroys intestinal enterocytes to cause villus blunting, resulting in malabsorption and diarrhea. The nucleocapsid (genetic core) contains 11 segments of double‐stranded RNA. Each gene segment encodes a different viral protein to create the entire virus [91]. The virus must invade intestinal epithelial cells to survive in vivo.

Rotavirus was first identified in the diarrheic feces of turkey poults [92], and different strains of rotaviruses have been identified in turkeys, pigeons, chickens, guineafowl, lovebirds, partridges, and pheasants. In gamebirds, the virus typically causes diarrhea in pheasant and partridge chicks [93]. Avian rotaviruses can be characterized into different groups by serum neutralization, enzyme‐linked immunosorbent assay (ELISA) and by migration patterns on polyacrylamide gel electrophoresis [94]. Type A, D, F, and G rotaviruses have been detected in birds, and type A and D have been characterized in pheasants. By genetic analysis, pheasant rotaviruses are only distantly related to other poultry rotaviruses [95]. Most human rotaviruses are type A, but there is no evidence that avian rotaviruses can be transmitted to people; these viruses are antigenically and genetically dissimilar [96]. Avian rotaviruses have not been detected in humans and human rotaviruses have not been detected in birds.

Large numbers of rotaviruses are shed in the feces of infected birds. The virus is then contracted by swallowing contaminated manure which may be on the soil, feeder, drinker or equipment, shoes and clothing of workers. There is no evidence that rotavirus can be transmitted inside eggs. The virus must multiply in mature enterocytes at the tips (apices) of intestinal villi rather than the deeper crypts [97]. Damage to enterocytes with blunting of the villi will impair the absorption of fluid and nutrients, resulting in diarrhea with increased fluid in the intestinal lumen. Virus can be observed by electron microscopy of feces or intestinal contents [98, 99].

Clinical signs of rotavirus infection in pheasants and partridges usually start as early as 4–14 days of age, but signs can be observed in birds as old as 6 weeks. Clinical signs include death without premonitory signs, depression, huddling, piling in corners, decreased feed consumption, discomfort as evidenced by chirping and consumption of litter, and diarrhea or wet litter. Rotavirus infection in gamebird chicks is often complicated by Salmonella or Escherichia coli infection, and the virus can be apathogenic as a solitary infection [91]. Gross necropsy reveals dehydrated carcasses, fecal pasting of the vent feathers, pale viscera and small intestine, and ceca dilated with yellow frothy fluid (Figure 11.15) [94]. Gizzards can contain litter.

11.3.3.3 Other Enteric Viruses

Coronaviruses are enveloped, positive sense, single‐stranded RNA viruses in the subfamily Orthocoronavirinae, family Coronaviridae. These viruses have a wide range of hosts and have been associated with enteritis in quail and guineafowl. In one report, farm‐reared coturnix quail showed depression, diarrhea, and reduced growth rate, with the most severe mortality in the youngest birds (3 weeks). The prominent lesion in the quail was enteritis, with coronavirus observed in intestinal contents by electron microscopy [113]. Sequence analysis of the virus revealed genetic similarities to turkey coronavirus. Similarly, Japanese quail with high mortality had histologic lesions of decreased villous length, increased crypt depth, and lymphoplasmacytic infiltrates in the lamina propria of small intestine [114]. Metagenomic analysis of intestinal contents from the quail revealed predominantly coronavirus with a lesser percentage of picornavirus. The coronavirus shared genetic similarity to turkey enteric coronavirus and infectious bronchitis virus [114].

In France, there are reports of a novel avian coronavirus implicated as one of the causes of an enteric condition of guineafowl known as “fulminating disease.” Clinical signs of the fulminating disease of guineafowl were prostration, decreased water and feed consumption, with mortality reaching as high as 20% per day. Lesions of enteritis and an enlarged, white pancreas (pancreatic degeneration) were observed. In most instances, a coronavirus was identified by electron microscopy of intestinal contents, immunohistochemistry, PCR or whole‐genome sequencing [115, 116]. This novel virus has been sequenced and renamed as guineafowl coronavirus, a gammacoronavirus [117].

Caliciviruses are in the family Caliciviridae and are small, nonenveloped, single‐stranded RNA viruses with distinct cup‐shaped surface structures [118]. Gough et al. described calicivirus in the feces of 3–4‐week‐old pheasant chicks with mild enteritis, inappetence, and weight loss with eventual loss of 25% of the flock [119]

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