LIFE HISTORY TYPES AMONG PROTOZOA

There is a wide variety of life history strategies used by parasitic protozoa to be successful. The major ones of importance to wildlife are presented in this section (Table 6.1).

Direct (Homoxenous) Life Cycles

No Cyst Stage

TRICHOMONAS CALLINAE

CAUSATIVE AGENT (CLASSIFICATION, morphology) The genus Trichomonas and other trichomonads are classified among the Parabasilia of the Supergroup Excavata (App. 1: Table 5) (Adl et al. 2005). The genus Trichomonas includes a number of species infecting birds and mammals, of which one, Trichomonas gallinae, is a wildlife pathogen that infects the upper respiratory tract of birds (Cole 1999b). Another, T. foetus, is a parasite of the urogenital system of cattle (BonDurant and Honigberg 1994), and T. vaginalis infects the urogenital system of humans (Leber and Novak 1999).

host range and distribution Tricho­monas gallinae has a worldwide distribution, with the domestic pigeon (Columba livia) serv­ing as primary host (BonDurant and Honigberg 1994). In North America, it is speculated that T. gallinae generally occurs wherever domes­tic pigeons and mourning doves (Zenaida macroura) are found (Stabler 1954, Cole 1999b). This parasite also infects other columbiforms, as well as falcons and hawks, owls, passer­ine birds, upland birds, waterfowl, and gulls (BonDurant and Honigberg 1994, Cole 1999b).

life cycle and variations Trichomo- nads have the simplest kind of protozoan life cycle, in which the organism occurs only as a trophozoite, with no cyst. These parasites mul­tiply by binary fission (Leber and Novak 1999).

reservoirs and transmission The common pigeon (Columba livia) is believed to be the ultimate source of T. gallinae infec­tion throughout the world (BonDurant and Honigberg 1994). The parasite was introduced to the United States with the pigeons and doves brought by European settlers (Stabler 1954).

Because there is no resistant cyst stage, trans­mission from host to host must be relatively direct. In pigeons, the parasite is spread dur­ing courtship behavior and infection of young through the process of regurgitive feeding of partially digested foodstuffs (“pigeon's milk”) produced in the crops of the adults (BonDurant and Honigberg 1994). Because T. gallinae is not found in the intestinal tract past the muscu­lar stomach, it cannot be transmitted in fecal droppings; rather, it is transmitted only from the mouth, and secondarily the nares and eyes (BonDurant and Honigberg 1994). Transmis­sion to species other than pigeons is through ingestion of parasites from water contaminated by infected pigeons, or through ingestion of infected hosts (Stabler 1954, Pokras et al. 1993, Cole 1999b).

clinical effects and diagnosis Although trichomoniasis is a general term for an infection, the disease also is termed “canker” in columbiform birds and “frounce” in raptors (Pokras et al. 1993). For columbiforms, most T. gallinae strains are nonpathogenic or only moderately pathogenic for their avian hosts; however, there also are a number of virulent strains (BonDurant and Honigberg 1994). Virulent strains kill young birds, and thus often are selected out of a population. Benign strains may provide immunity against the more virulent strains.

The normal sites of infections in colum- biforms are the mouth, pharynx, esophagus, and crop. Virulent strains may cause lesions in the upper digestive tract and spread to other parts of the body through these ulcerations (BonDurant and Honigberg 1994). Infected birds often appear listless, with ruffled feath­ers, and with yellowish caseous lesions around the beak or eyes.

Trichomonas gallinae can be diagnosed in wet smears from the mouth and oropharyngeal cavity of pigeons or by cultivation of swabs taken from those areas (BonDurant and Honigberg 1994). Stabler argued that examining samples directly in saline solution was better than trying to culture organisms. He described the following technique:

Sit, place the pigeon on its back on the examiner’s left leg; place the floor of the thumb against the tip of the lower mandible and pull back gently, thus opening the bird’s mouth; with a pair of curved forceps, stroke the roof of the mouth back of the palatal flaps in an effort to remove saliva for examination (the curve ofthe forceps should stroke and lift the rear of the flaps); dab saliva in a drop of saline and examine on ’scope. This procedure, when mastered, is quick, easy, and much more efficient than all the swabbing and culturing! (Stabler 1975)

We found this technique to work very well. More recently, the InPouch TF culture system (BioMed Diagnostics, San Jose, California) has been very effective compared with other tech­niques, including wet mount and direct micro­scopic examinations, for detecting T. gallinae (Cover et al. 1994, Bunbury et al. 2005).

population effects Trichomonas gallinae is a very common parasite of pigeons and some­times can cause mortality involving thousands to tens of thousands of birds (Cole 1999b, Hofle et al. 2004). Trichomoniasis is considered by many avian disease specialists to be the most important disease of mourning doves (Zenaida macroura) in North America (Conti 1993). Band-tailed pigeons (Columbafasciata) also have experienced considerable losses (Cole 1999b). The role of T.

Trichomoniasis in raptors (frounce) has caused mortality in falcons, and less commonly in hawks and owls, for hundreds of years (Bert 1619, Stabler 1954, Cooper 2002), and may have serious impacts on some raptors populations (Cooper and Petty 1988, Pepler and Oettle 1992). Trichomoniasis among wild raptors is linked to their feeding on urban pigeons. With Cooper’s hawks (Accipiter cooperii), prevalence of T. gallinae was significantly higher among urban nestlings (85%) than nestlings in undeveloped natural areas (exurban sites) (9%) (Boal et al. 1998); nestling mortality also was higher in urban than in exurban nests (Boal and Mannan 1999). In contrast, very few adults were found infected (Boal et al. 1998). In a study among northern gos­hawks (Accipiter gentilis), overall nestling preva­lence among urban birds was 65% and tended to increase with age of nestlings and brood size (Krone et al. 2005). However, prevalences of T. gallinae in raptors can vary considerably by site (Rosenfield et al. 2002), and the overall impact of this parasite on raptors is not well understood.

special problems Although the parasite has a very broad distribution, introduction of T. gallinae to new sites has been of some concern. Domestic pigeons were introduced to the Galapa­gos Islands in 1972 or 1973 (Harmon et al. 1987). A high prevalence later was found among these introduced birds, along with T. gallinae infec­tions among endemic Galapagos doves (Zenaida galapagoensis); doves examined on pigeon-free islands were not infected (Harmon et al. 1987).

control For virulent T. gallinae strains, removing infected individuals and any other source of infection is an effective strategy to pro­tect susceptible birds (BonDurant and Honigberg 1994). Preventing concentrations of doves at bird feeders and artificial watering sites can help reduce transmission among wild populations (Cole 1999b).

Most common antibiotics are ineffec­tive against T. gallinae infections and are not practical for wild populations (BonDurant and Honigberg 1994, Cole 1999b). Several nitroimidazole compounds are effective when administered in drinking water, but this group of drugs cannot be used for any food-producing animals in the United States (McDougald 1991, BonDurant and Honigberg 1994).

Cyst Stage Present

EIMERIA spp.

causative agent (classification, morphology) Members of the genus Eimeria fall within the Supergroup Chromoalveolata (App. ι: Table 5) (Adl et al. 2005). As Apicom- plexa, all are parasitic (Lindsay and Todd 1993). Most Eimeria spp. are parasites of the gastroin­testinal tract, although some also invade other tissues (Lindsay and Todd 1993).

host range and distribution Members of the genus Eimeria have a worldwide distri­bution, with many species affecting a wide variety of hosts among all classes of vertebrates (Gylstorff and Grimm 1987, Cole 1999a, Friend and Franson 1999, Bush et al. 2001, Duszyn­ski and Upton 2001). Well over 800 species of Eimeria are reported from mammals alone, and this may be only a fraction of the total in mam­mals (Duszynski and Upton 2001).

While at least 60 avian species of Eimeria have been reported (Todd and Hammond 1971), many of these species are not well understood. Much of the information on avian coccidiosis is through work on domestic birds. There cur­rently are seven recognized species of Eimeria in chickens, at least four species in domestic turkeys, 10 species in pheasants, two species in guinea fowl, at least six from partridge and quail, and about 15 species in geese (Long 1993). At least six species have been described from raptors (Cawthorn 1993).

life cycle and variations All members of the genus Eimeria have a direct life cycle (Fig.

reservoirs and transmission It is not clear whether most species of Eimeria have only a single vertebrate host (Lindsay and Todd 1993, Bush et al. 2001) or if they regularly can infect more than one species (Duszynski and Upton 2001). Among wild mammals, members of the Orders Rodentia and Artiodactyla have the largest numbers of known Eimeria species, followed by Lagomorpha, Carnivora, Diprodon- tia, Insectivora, and Chiroptera (Duszynski and Upton 2001). Eimeria spp. infections are reported among wild birds of the Orders Anseriformes, Galliformes, and Passeriformes (Todd and Hammond 1971). Transmission typically is by ingestion of a mature oocyst containing infective sporozoites (Duszynski and Upton 2001).

clinical effects and diagnosis Enteritis, associated with diarrhea, is a common clinical syndrome. Some Eimeria spp. also may invade the liver, leading to hepatitis. However, pathoge­nicity of coccidian parasites in wild mammals may not be of great significance, and appears to be a disease due primarily to human intervention (Duszynski and Upton 2001).

Among geese, there is a renal and an intesti­nal form of coccidiosis (Long 1993). Renal coc- cidiosis, caused by E. truncata, develops only in the kidneys and the cloaca near its junction with the ureters; the route to infecting the kid­neys is unknown (Long 1993). Most other avian species of Eimeria are not considered severely pathogenic (Long 1993).

When a host is not killed, diagnosis of a coc- cidian infection typically depends on finding oocysts in the feces and observing them com­pletely sporulate (Duszynski and Upton 2001). The polymerase chain reaction and random amplified polymorphic DNA tests also are used to detect and identify coccidia (Comes et al. 1996, Barta et al. 1997, Duszynski and Upton 2001).

population effects Among mammals, Eimeria stiedae, for example, may be a limit­ing factor on young wild rabbits (Oryctolagus cuniculus) of Australia in normal and wet years, but not in dry years (Dunsmore 1971). Most wild birds pass small numbers of oocysts in their feces without apparent ill effects, but coccidiosis becomes important as a disease when animals live under crowded or dense populations that allow buildup of infective oocysts in the envi­ronment and ingestion by susceptible hosts (Long 1993). Among domestic birds, it is specu­lated that the poultry industry is dependent on continuous medication in the feed (Long 1993).

Renal coccidiosis, caused by E. truncata, has caused losses as high as 87% among infected, presumably domestic, flocks of birds in Iowa (Long 1993). This parasite also has been asso­ciated with mortality among Canada geese in North America (Critcher 1950, Farr 1954, Hanson et al. 1957)

special problems No ongoing special problems are noted for Eimeria spp.

control Among domestic birds, there are a wide variety of anti-coccidial chemo­therapies used to combat Eimeria infections (Long 1993). There also have been some efforts to immunize domestic birds by use of attenuated strains of Eimeria, as well as vac­cinating with specific antigens (Long 1993). Immunity does not play a significant role; among domestic birds immunity lasts only 4 to 6 weeks in birds recovering from infection (Long 1993).

Indirect (Heteroxenous) Life Cycles

Use Vertebrates as Both Intermediate and Definitive Hosts

TOXOPLASMA CONDll

CAUSATIVE agent (CLASSIFICATION, morphology) Toxoplasma gondii is classi­fied as a coccidian parasite, with Eimeria and Sarcocystis, among the Apicomplexa in the Chromoalveolata (Dubey and Odening 2001). The name Toxoplasma (toxon: arc, plasma: form) is derived from the crescent shape of the tachyzoite stage (Hill et al. 2005). Toxo­plasma gondii is the sole member of the genus (Hill et al. 2005); currently, three dominant genotypes (I, II, III) are recognized in the species (Grigg et al. 2001, Su et al. 2003, Volkman and Hartl 2003). Recently, another type (X) has been observed in marine mam­mals (Miller et al. 2004).

host range and distribution Although it can only complete its life cycle in felids (Dubey and Odening 2001), T. gondii can infect nearly all mammal and bird species and has a worldwide distribution (Lehmann et al. 2006). In recent years, it has been described and become widely reported among marine mammals (Dubey 2004). Toxoplasma gondii is postulated to originally have been a parasite of South American felids that underwent a rela­tively recent worldwide expansion from the effects of both migratory birds and the expansion of maritime travel; in particular, the transatlantic slave trade may have played a considerable role in this expansion (Lehmann et al. 2006).

life cycle and variations Wild and domestic felids are the only known definitive hosts; oocysts that successfully sporulate in the environment and infect susceptible hosts are formed only in cats (Fig. 6.9: (Dubey

et al. 1970). In contrast to other coccidia, sev­eral life history stages of T. gondii can infect both intermediate hosts and definitive hosts. The three infectious stages of T. gondii are spo­rozoites, tachyzoites (trophozoites), and brady- zoites, developing in meront-like tissue cysts called pseudocysts) (Dubey 2004).

Most natural infections probably are acquired by ingesting sporulated oocysts with sporozoites in food or water contaminated with cat feces, or by ingesting pseudocysts in infected meat (Hill et al. 2005). Oocysts transform into tachyzoites shortly after ingestion by intermedi­ate hosts (Fig. 6.9: Γa). Tachyzoites rapidly pro­liferate in pseudocysts by binary fission (Bush et al. 2001) and destroy infected cells during acute infection. Tachyzoites localize in neural and muscle tissue and develop into bradyzoites within pseudocysts (Fig. 6.9: Γ^). Bradyzoites multiply slowly in tissue cysts (Bush et al. 2001). Cats become infected by consuming intermedi­ate hosts with these tissue cysts (Fig. 6.9: ^ι,Λ).

Toxoplasma gondii also may spread by infected macrophages to mesenteric lymph nodes and then to distant organs by invasion of the lymphatic system and blood (Hill et al. 2005). The host cells die and release tachyzo­ites, which invade adjacent cells and continue the process (Wilson and McAuley 1999, Hill et al. 2005). Tachyzoites are pressured by host immune responses to transform into bradyzoites and to form meront-like pseudo­cysts (Wilson and McAuley 1999). Although

FIGURE 6.9 Life cycle of Toxoplasmagondii. Three sources of infection (a, b, c) occur for both intermediate and definitive hosts. Bradyzoites in pseudocysts will become tachyzoites if ingested by a new host, and then form bradyzoites in that intermediate host; or they will go on to form gametes and oocysts if they are in a felid definitive host (Courtesy of Centers for Disease Control and Prevention's Division of Parasitic Diseases and Malaria, www.dpd.cdc.gov/dpdx).

pseudocysts containing bradyzoites may develop in visceral organs, including lungs, liver, and kidneys, they are more prevalent in muscular and neural tissues, including the brain, eye, skeletal, and cardiac muscle (Hill et al. 2005). Infected intermediate hosts may contain infec­tive cysts for their lifetime and shed tachyzo- ites, but they cannot complete the life cycle (Hill et al. 2005).

Generally, cats are more readily infected by ingesting bradyzoites than oocysts (Dubey 2004); ingesting as few as one bradyzoite can lead to a cat shedding millions of oocysts (Dubey 2001). Virtually all cats ingesting pseudocysts containing bradyzoites will shed oocysts, whereas less than half of the cats ingesting tachyzoites or oocysts will complete the life cycle (Dubey and Frenkel 1972).

When felids ingest pseudocysts from infected prey, the released bradyzoites penetrate epithelial cells of the small intestine (lamina propria) (Dubey 2004). In the epithelial cells, T. gondii multiplies (enteroepithelial cycle) and forms meronts (also called schizonts), which, in turn, continue developing to merozoites, gametes, zygotes, and oocysts (Dubey and Frenkel 1972, Hill et al. 2005). Formed oocysts are discharged into the intestinal tract by rup­ture of the epithelial cells and sporulate about 1-5 days after passing into the environment (Hill et al. 2005).

As this enteroepithelial cycle progresses, bradyzoites also penetrate the feline intestine and multiply as tachyzoites, eventually dissemi­nating to other tissues in the cat. Both intesti­nal and extraintestinal stages of T. gondii may occur for several months, or even remain for the life time of the cat (Hill et al. 2005).

Often, wild or domestic animals can become infected (Fig. 6.9: ^ι^). Humans become infected by eating these animals, as well as by eating food or water contaminated by cat feces, including from cat litter boxes (Fig. 6.9: l6). In a more seri­ous problem, developing fetuses may be infected transplacentally, sometimes with fetal mortality or severe neurological consequences after birth.

RESERVOIRS AND TRANSMISSION One collective population estimate of domestic and feral cats in the United States ranges from about 93 to 133 million cats [Introduced Species Sum­mary Project: Domestic Cat (Felis catus), www.columbia.edu/itc/cerc/danoff-burg/invasion_bio /inv_spp_summ/Felis_catus.html]. Up to 2% of all cats are estimated to be shedding T. gondii oocysts at any given time, with occasional higher prevalences reported (Dubey 2004).

Three known routes of transmission for T. gondii include transmission through oocysts shed in the feces of wild and domestic felids, ingestion of tissue cysts from meat, and con­genital transmission (Fayer et al. 2004). The oocysts are the only stage to enter the general environment, and they can remain infectious for months to years under moist temperate con­ditions while being widely dispersed in large numbers (Yilmaz and Hopkins 1972, Frenkel et al. 1975); thus, oocysts are viewed as the most likely means of transmission to non-carnivores (Fayer et al. 2004). Land-based surface runoff is a likely source of T. gondii oocysts for marine mammals (Miller et al. 2002a).

Immunity does not eradicate infection, and T. gondii tissue cysts persist several years after acute infections (Hill et al. 2005). It is not clear how T. gondii is destroyed in immune cells (Renold et al. 1992). Extracellular forms of the parasite are directly affected by antibody, but intracellular forms are not; it is believed that T-lymphocytes and lymphokines are more important than humoral factors in immune destruction of T. gondii (Renold et al. 1992).

clinical effects and diagnosis Pathoge­nicity of T. gondii is determined by many factors, including the susceptibility of the host species, virulence of the parasitic strain, and the life cycle stage ingested; oocyst-induced infections are the most severe clinically in intermediate hosts, independent of dose (Dubey 2004). Although cats of any age can die of toxoplasmosis, kittens and those with depressed immunity are the most susceptible (Dubey and Carpenter 1993a, 1993b). Toxoplasmosis can be a severe disease in domes­tic animals, and may cause embryonic and fetal death, abortion, stillbirth, and neonatal death (Dubey 2004).

Clinical signs of toxoplasmosis are nonspe­cific and mimic those of several other diseases (Dubey and Odening 2001). Trophozoites can invade many tissues, leading to fever, spleno­megaly, myocarditis, pneumonitis, and enceph­alitis. The pathology described among marine mammals includes meningoencephalitis, with lesser observations of hepatitis, myocarditis, and lymphadenitis, as well as disseminated and congenital infections (Dubey et al. 2003, Kreuder et al. 2003, Dubey et al. 2004, Miller et al. 2004).

Diagnosis of toxoplasmosis is made by bio­logic, serologic, or histologic methods, or by their combination (Hill et al. 2005). Oocysts in infected cats are diagnosed by concentra­tion methods such as flotation in high-density sucrose solutions (Ruiz and Frenkel 1980). Isolation of T. gondii from animals by inocula­tion of laboratory animals and tissue cultures can lead to definitive diagnoses (Dubey and Odening 2001). However, for epizootiologi­cal surveys, serological prevalence is a better measure of T. gondii prevalence (Dubey 2004). Serological tests include the dye test, indirect fluorescent antibody test (Miller et al. 2002b), modified agglutination test (MAT), ELISA, and immunosorbent agglutination assay test (Dubey and Beattie 1988). Some work with polymerase chain reaction is promising (Hill et al. 2005).

population effects Generally, the para­site causes few problems among wildlife popu­lations. One exception may be with sea otters (Enhydra lutris), and this is addressed in the next section.

special problems Toxoplasma gondii has been detected in a wide variety of geographic areas from seals (Lambourn et al. 2001, Dubey et al. 2003, Measures et al. 2004, Honnold et al. 2005), dolphins (Dubey et al. 2003, Cabezon et al. 2004, Dubey et al. 2005), sea otters (Cole et al. 2000, Lindsay et al. 2001b, Miller et al. 2002b, Dubey et al. 2003), and other marine mammals (Cruikshank et al. 1990, Dubey et al. 2003, Philippa et al. 2004). Pathology and mortality have been described from dolphins (Inskeep et al. 1990, Jardine and Dubey 2002, Resendes et al. 2002), a seal (Dubey et al. 2004), and sea otters (Kreuder et al. 2003, Miller et al. 2004). In an experi­mental infection, Toxoplasma established patent infections in gray seals (Halichoerus grypus), but did not cause pathology (Gajadhar et al. 2004).

The southern sea otter (Enhydra lutris nereis) is listed as threatened under the Endan­gered Species Act and is considered a keystone species strongly influencing the abundance and diversity of other species within the kelp forest ecosystem (Jessup et al. 2004). There has been a recent decline as a consequence of high mortality rather than low recruitment, with considerable concern among biologists and managers (Estes et al. 2003, Gerber et al. 2004). Current evidence is that toxoplas­mosis is making a significant contribution to this mortality (Kreuder et al. 2003, Hill et al. 2005). Of the T. gondii evaluated from sea otters in one study, 40% were Type II and 60% were Type X; no Types I or III were observed (Miller et al. 2004). Subsequent work has an estimate of 72% Type X in sea otters, with the observation that the same Type X has been isolated from other marine mammals as well (Conrad et al. 2005).

Of the potential sources of infection, ingestion of sporulated oocysts seems most likely; sea otters do not prey on warm-blooded animals, which are the usual intermedi­ate hosts of T. gondii (Conrad et al. 2005). There is evidence that Toxoplasma gondii has invaded the marine ecosystem through runoff from terrestrial sources (Miller et al. 2002a, Fayer et al. 2004). Sea otters in areas of high freshwater runoff are more likely to be exposed to T. gondii than otters from low or medium freshwater runoff (Miller et al. 2002a). Increased risk of sea otter mortal­ity has been linked to increased exposure to T. gondii (Miller et al. 2002a, Kreuder et al. 2003, Conrad et al. 2005).

Collectively, there is good evidence that Toxoplasma oocysts are spread from ter­restrial runoff into the marine ecosystems, where they are concentrated in the resident shellfish; shellfish are a major food source of the California sea otters and serve as a means of transmission (Mitchell and Sinai 2006), or as paratenic hosts (Arkush et al. 2003). Eastern oysters (Crassostrea virginica) are able to remove T. gondii oocysts from sea­water readily and retain their infectivity for at least 85 days (Lindsay et al. 2001a, 2004). Similarly, another bivalve mussel, Mytilus galloprovincialis, also is able to take up and retain infective T. gondii oocysts (Arkush et al. 2003). It also has been observed that anchovies (Family Engraulidae), a small common saltwater fish, can be infected with T. gondii after exposure (Massie and Black 2008); these fish are common prey items of marine mammals. Thus far, natural infec­tions have not been reported in these species, however, and their actual role in the epizooti­ology of the disease is unclear (Conrad et al. 2005). Interestingly, the oocysts also sporu- late readily and remain viable in seawater for at least 6 months (Lindsay et al. 2003).

Reasons for the unusual morbidity and mor­tality from toxoplasmosis in sea otters, com­pared to the subclinical or mild infections seen in most immunocompetent terrestrial animals, are unclear (Conrad et al. 2005).

control Prevention of Toxoplasma among humans and domestic animals generally is by maintaining cleanliness and reducing inges­tion of contaminated food or water (Hill et al. 2005). There are no specific recommendations to prevent transmission of T. gondii by drink­ing water because the level of contamination is unknown and because it can be transmitted by several modes (Dubey 2004). At present there is no vaccine to control toxoplasmosis in any species (Dubey and Odening 2001).

Arthropod-borne Apicomplexa

PLASMODIUM RELICTUM (AVIAN MALARIA)

causative agent As Apicomplexa (Adl et al. 2005), all members of the genus Plasmo­dium are parasitic. There are about 34 recog­nized species of Plasmodium described from birds (Bennett et al. 1993). Over 25% of all avian host species are parasitized by members of the genus Plasmodium (Bennett et al. 1982, 1994).

host range and distribution Plasmo­dium relictum has a worldwide distribution (Bennett et al. 1993, Peirce 2005, Kimura et al. 2006); it is the most common avian Plas­modium species and has been reported from over 300 species and at least 70 families of birds (Bennett et al. 1993, Valkiunas 2005).

life cycle and variations Using the general malarial life cycle model (Fig. 6.5), P. relictum sporozoites are introduced to suscep­tible birds by the bites of infected culicine mos­quitoes. Sporozoites invade a variety of cells, including the Malpighian body, and later the lungs, brain, and spleen (van Riper et al. 1994). After three or more generations of merogony in these tissues, the merozoites invade eryth­rocytes, where they also undergo merogony; in older literature this was referred to as exoeryth- rocytic (out of erythrocyte) schizogony and erythrocytic schizogony. The resulting merozo- ites then develop into gametocytes (van Riper et al. 1994).

Gametocytes ingested by Culex spp. mosqui­toes form gametes that fuse into zygotes that, in turn, become motile (ookinetes); ookinetes penetrate midgut epithelial cells and form an oocyst on the hemocoel side of the gut. Following sporogony, oocysts rupture and sporozoites are released into the hemocoel of the mosquito; from here they migrate to the salivary glands to be injected into the new avian host during a blood meal by the mosquito (van Riper et al. 1994).

reservoirs and transmission The avian malaria cycle requires the avian vertebrate host and culicine mosquitoes as the inverte­brate hosts (van Riper et al. 1994). Although P. relictum completes its development in at least 26 species of mosquitoes (Huff 1965, Garnham 1966), culicine mosquitoes are judged to be the most likely transmitters under natural condi­tions (van Riper et al. 1994).

clinical effects and diagnosis Clini­cally ill birds may appear listless, have ruffled feathers, lose their appetite, and have difficulty breathing; pathologic changes often involve anemia as well as enlargement of the spleen and liver (van Riper et al. 1994). Some infected birds may show few if any symptoms.

population effects The parasite P. relictum Capistranoae has been associated with severe population declines of native birds in Hawaii, including extinction of many native species of Hawaiian honeycreepers (Drepanidinae). These events followed introduction of the vector, the house mosquito, Culex quinquefasciatus, in approximately 1826 and introduction of the parasite through migratory birds (Warner 1968, van Riper et al. 1986).

special problems Endemic Hawaiian forest birds are highly susceptible to infection with Plasmodium relictum capistranoae, with some having mortality rates of 65-90% after being bitten by a single infective mosquito (Atkinson et al. 1995, 2000, 2001b). High susceptibility of native Hawaiian honeycreep­ers (Family Fringillidae, Subfamily Drepanid- inae) to avian malaria is believed to be one of the primary factors responsible for the disap­pearance of these birds from lowland habitats on the main Hawaiian islands (Atkinson et al. 2001a). There is compelling evidence that the near absence of native forest birds below elevations of 900 m in Hawaii was due to the Culex spp. vector and parasite transmission (van Riper et al. 1986). With further work, it was established that malaria transmission was greatest in mid-elevation forests where the highly susceptible native forest birds over­lapped in distribution with mosquitoes and where these native birds themselves served as the primary reservoirs of infection (van Riper et al. 1986, Atkinson et al. 2001a, Woodworth et al. 2005). The native birds were almost entirely absent from low-elevation study areas because of high mosquito densities; malaria prevalence in the primarily exotic, disease­resistant lowland avifauna was low (van Riper et al. 1986, Woodworth et al. 2005).

There is evidence that at least one native Hawaiian forest bird, the amakihi (Hemignathus virens), has endured the parasite, has mounted an immune response (Atkinson et al. 2001a), and is even recolonizing low-elevation habitats that contain higher densities of the culicine vec­tor (Woodworth et al. 2005).

control No specific measures are taken to control avian malaria in wild birds.

TRYPANOSOMA SPP. (NAGANA)

CAUSATIVE AGENT (CLASSIFICATION, morphology) The nagana disease complex involves three species of trypanosomes (T. vivax, T. congolense, T. brucei brucei) (Logan-Henfrey et al. 1992); they are classified in the Kinetoplastea among the Euglenozoa in the Excavata Super­group (Adl et al. 2005). These are long, spindle­shaped cells (about 2 ? 20 mμ) with a single flagellum (Logan-Henfrey et al. 1992). All three are salivarian parasites typically passed through the bites of tsetse flies.

host range and distribution Distribu­tion of this group of parasites is between 14°N and 29°S on the African continent (Wells and Lumsden 1969). Distribution is closely linked to the distribution of tsetse flies (Glossina spp.), which infest about 11 million km² of Africa, or about one-half of the available arable land. A wide variety of African ungulates are infected with this complex of organisms (Ashcroft 1959, Baker et al. 1967, Geigy et al. 1967, Baker 1969). Most of these wild ungulates are resis­tant to the trypanosomes, but occasional pathol­ogy and disease has been reported among them (Molyneux 1982).

LIFE cycle and variations The trypomas- tigote and the epimastigote (Fig. 6.8) stages are found among hosts of the African salivar- ian trypanosomes (Logan-Henfrey et al. 1992). Infected tsetse flies inoculate metacyclic trypo- mastigotes into the skin of animals, where the trypanosomes grow for several days and cause local swellings. They enter the lymph nodes and then the bloodstream, where they divide rapidly by binary fission (Fraser and Mays 1986).

After the tsetse fly feeds on a host infected with trypanosomes, the bloodstream trypomas- tigotes develop into procyclic trypomastigotes in the midgut, then into epimastigotes, and then into metacyclic trypomastigotes (Logan- Henfrey et al. 1992). There also are a number of variations in the timing of the life cycle, the various stages developed in the hosts, and the host sites at which the parasites concentrate among the trypanosome species (Fig. 6.10) (Logan-Henfrey et al. 1992).

reservoirs and transmission These African trypanosomes typically are passed by the bite of an infected tsetse fly. There are potentially 23 tsetse fly species involved, classi­fied into three taxonomic groups according to their preferred ecological habitat. The Glossina morsitans group typically is found in savanna, the G. palpalis group prefers areas around riv­ers and lakes, and the G. fusca group lives in high forest areas (Logan-Henfrey et al. 1992). The limits of tsetse infestation are determined primarily by climate and secondarily by vegeta­tion (Ford 1971, Logan-Henfrey et al. 1992).

In addition to the typical transmission by hematophagous arthropods, there also is evidence that transmission of trypanosomes to predators such as lions (Panthera leo) and hyenas (Crocuta crocuta) can occur by ingestion of infected ungulates (Baker et al. 1967, Baker 1969, Geigy and Kauffman 1973). This insight is supported by laboratory studies (Heisch 1963, Moloo et al. 1973).

CLINICAL EFFECTS AND DIAGNOSIS The severity of disease varies with the species of animal and the trypanosome species involved (Molyneux 1982). There are individual case reports of wild ungulate species in which lesions were attributed to trypanosome infec­tions, but generally African ungulates have an innate tolerance of trypanosome infections (Molyneux 1982). For cattle and other suscep­tible species, the primary clinical sign is ane­mia, with an associated stunting and wasting

FIGURE 6.10 Life Cycle of a Salivaria (Trypanosoma brucei gambiense) (Courtesy of Centers for Disease Control and Prevention’s Division of Parasitic Diseases and Malaria, www.dpd.cdc.gov/dpdx).

(Molyneux 1982). Most infected cattle have a chronic disease course with high mortality (Fraser and Mays 1986); mortality usually is associated with circulatory disorder, damage to the heart, and the anemia (Molyneux 1982). There are few consistent clinical signs or inter­nal lesions specific to the disease, and other dis­eases in Africa (e.g., anaplasmosis, babesiosis, theileriosis) can be confused with trypanoso­miasis (Logan-Henfrey et al. 1992).

Trypanosome infections may be detected directly by microscopic observation of the para­sites, by immunoassays for trypanosome anti­gens, or by biochemical techniques that detect trypanosomal DNA. Identifying trypanosome infections also is based on serological detec­tion of anti-trypanosomal antibodies (Logan- Henfrey et al. 1992).

population effects There is little overall impact on indigenous wild ungulate species. However, much of Africa between 14°N and 29°S was excluded for cattle due to the severe impacts of trypanosomiasis. There is virtually no overlap between the cattle-raising regions of Africa and the range of tsetse flies and the trypanosomes (Donelson and Turner 1985).

special problems Cattle have been denied vast areas of Africa by the occurrence of trypano­somiasis. Trypanosomiasis is regarded economi­cally as the most important livestock disease in Africa (Jawara 1990), with an estimated annual loss to African farmers in meat production alone estimated at US$5 billion in the 1990s (Gyening 1990, Logan-Henfrey et al. 1992).

control Historically, risk of trypanosomi­asis was minimized by avoiding tsetse-infected areas. More recently, control of trypanosomia­sis in livestock relies on the treatment of the infected host and on vector control (Logan- Henfrey et al. 1992).

In the early 1900s, it was observed that the mortality among wild ruminants from rinderpest was so high that the tsetse fly, espe­cially Glossina morsitans, was severely reduced in some areas such as Kruger National Park, from lack of suitable hosts (Stevenson-Hamilton 1911, Duke 1919, Carmichael 1933). At that time, “game elimination” became a technique used to limit the sources of blood meals available to the tsetse flies, and to eliminate trypanosome reservoirs (Logan-Henfrey et al. 1992); this in turn was intended to reduce tsetse fly popula­tions, specifically Glossina morsitans, and to enhanced success of cattle in the region; there was some evidence for success with this approach (Clarke 1964, Ford 1971). Between the 1920s and the 1960s, it was estimated that 1.3 million game animals were killed in tsetse control programs (Matthiessen and Douthwaite 1985). However, the public outcry against wildlife hunting as a means of tsetse control (Cockbill 1967) and the developing evidence for its overall inefficiency (Glover 1965), along with the advent of insec­ticides, largely ended the practice by the early 1970s (Matthiessen and Douthwaite 1985). At that time there also was an increasing apprecia­tion of the game animals as valuable in them­selves and as important national resources (Molyneux 1982).

Vector control historically involved bush clearing in an effort to destroy tsetse fly habitat, to limit the sources of blood meals available to the tsetse flies, and to eliminate trypanosome reservoirs (Logan-Henfrey et al. 1992). Pastoral­ists and arable farmers tend to clear brush as part of their land use, making the land unsuitable for the flies (Matthiessen and Douthwaite 1985). Some savanna woodland clearing was done to destroy G. morsitans; while there is evidence that this has been effective in some regions, there is no overall assessment of its effects, but there is speculation that continued deforestation and other habitat changes changed and reduced the occurrence of tsetse flies, and trypanosomiasis, in parts of Africa (Molyneux 1982).

Game destruction and bush clearing were replaced by insecticides for tsetse control (Jordan 1974, Allsopp 1984). A variety of insec­ticides have been used, but generally have had limited effectiveness and often have had signifi­cant side effects on nontarget species (Du Toit 1954, Graham 1964, Wilson 1972, Molyneux 1982, Matthiessen 1985, Matthiessen and Douthwaite 1985).

In recent years, use of traps and screens impregnated with pyrethrum has been effec­tive in reducing tsetse flies of the G. palpalis and

G. fusca groups; also, a number of attractants are effective for capture of the G. morsitans group of flies (Logan-Henfrey et al. 1992). There has been considerable interest in biologi­cal control, including use of sterilized males, but no effective techniques have emerged (Logan-Henfrey et al. 1992). There also is exploration of developing more trypanotoler- ant animals, but these animals represent only a small portion of the cattle in Africa (Logan- Henfrey et al. 1992).

Treatment of individually affected animals can be accomplished with a number of drugs, including dimazene aceturate, quinapuramine sulfate, and homidium bromide, among oth­ers (Logan-Henfrey et al. 1992). There is little likelihood of an effective vaccine because of the unique ability of trypanosomes to evade the immune system while in the blood stream by switching on new genes to encode new surface antigens (Donelson and Turner 1985). Trypano - somes may possess up to 1,000 variable surface glycoprotein (VSG) genes and, from these, they select certain ones to express when they are at the metacyclic stage of development, others as trypomastigotes in the early stages of infection in the blood, and still others during the later stages of chronic (Boothroyd 1985, Borst 1986, Van der Ploeg 1987, Pays and Steinert 1988).

Helminth-borne

H∣sτoMθNAs Meleacridis

CAUSATIVE AGENT (CLASSIFICATION, morphology) Histomonas meleagridis is included among the Trichomonadida of the Parabasalia, in the Supergroup Excavata (Adl et al. 2005). Histomonas meleagridis is one of two species in this genus and the only one of patho­genic significance. Two forms of H. meleagridis have been described, a lumen form with one or two flagellae, and a tissue form without flagellae; both range from 6 to 20 mμ in size (McDougald and Reid 1978). There is also some evidence for a third form parasitizing gonads of larvae of the cecal nematode Heterakis gallinae; a 3 μm form has been observed in cecal worm eggs (BonDurant and Wakenell 1994).

cost RANGE AND DISTRIBUTION Most, if not all, gallinaceous birds are suscep­tible hosts (Cole and Friend 1999), including chukar (Alectoris chukar), pheasant (Phasianus colchicus), quail (Order Galliformes), grouse (Family Phasianidae), wild turkeys (Meleagris gallopavo), jungle fowl (Gallus gallus), guinea fowl (Family Numididae), and peafowl (Pavo spp.) (BonDurant and Wakenell 1994). Histomonas meleagridis essentially has a world­wide distribution (Cole and Friend 1999).

life cycle and variations Histomonads multiply by binary fission. The H. meleagridis life cycle is unique in that it is one of the rare examples of a helminth-borne life history pat­tern; in addition, earthworms typically are used as paratenic hosts (Cole and Friend 1999). Histomonas meleagridis uses the cecal nematode Heterakis gallinae as a vector for entry into the avian hosts (BonDurant and Wakenell 1994). Although histomonads can be passed directly under optimal circumstances, H. meleagridis quickly loses viability upon cooling outside the host or inside a dead host; thus, development of some resting stage inside the nematode eggs appears essential for survival of H. gallinae, and this nematode is considered critical to the natu­ral transmission of H. meleagridis (BonDurant and Wakenell 1994).

While the histomonads are carried inside the nematode egg (Tyzzer 1934), it still is not clear how the parasite becomes incorporated into the egg and the nature of the form in which it may exist for several years (BonDurant and Wakenell 1994). Both the male and female

H. gallinae become infected with Histomonas; the histomonads then become incorpo­rated into the eggs of the female (Lee 1970, McDougald 1991). There is some evidence that the male Heterakis may become infected first and transfer the infection to the female during copulation (Springer et al. 1969, McDougald 1991). Unembryonated Heterakis eggs are not infective; also, the processes of hatching or physical destruction of the egg seem important in release of infective histomonads (BonDurant and Wakenell 1994).

reservoirs and transmission Heterakis gallinae, the nematode involved, is an ascarid with a direct life cycle, but one that also com­monly uses earthworms as paratenic hosts. After the histomonad eggs are shed into the environment, the embryos develop to second stage larvae (L ). On being ingested, the nema­tode larvae develop to adulthood in the ceca of their avian hosts. The histomonads accompany­ing the egg also infect the birds and may cause pathology before infecting the new generation of helminth ova produced by infecting H. gallinae.

In addition, earthworms often ingest fecally contaminated soil containing H. gallinae eggs infected with histomonads (Kemp and Franson 1975, BonDurant and Wakenell 1994). Thus earthworms concentrate the nematode eggs and enhance transmission; earthworms also take the eggs to new sites as they migrate in the soil and help preserve the eggs from fungi and predation by other soil invertebrates (Lund et al. 1966). After earthworms ingest Heterakis gallinarum eggs, L larvae hatch out and invade the earthworm tissues; thus the earthworm concentrates the H. gallinarum larvae as well as the histomonads present (Lund et al. 1966). Susceptible birds consuming these earth­worms become infected by both the Heterakis gallinarum nematode and Histomonas meleagri- dis. Thus, while earthworms are not required for the nematode or histomonad life cycle, they are considered to play a critical role in enhanc­ing natural transmission of each (BonDurant and Wakenell 1994).

clinical effects and diagnosis The dis­ease histomoniasis also is called blackhead because infections often cause cyanosis, a blue or black appearance on the skin of the head on turkeys and some other birds due to an excess of reduced hemoglobin in the blood (Cole and Friend 1999). There are no clinical signs specific to histomoniasis. Infected wild turkeys often are listless, amd have ruffled feathers and drooped wings. Feces are often sulfur-yellow in color and, combined with other field signs, are highly sug­gestive of histomoniasis (Cole and Friend 1999). The term enterohepatitis also is applied, reflect­ing their invasion of the cecum and, in more serious cases, causing multifocal necrosis in the liver (BonDurant and Wakenell 1994)

Turkeys, grouse, and partridge develop severe disease and have mortality rates that can exceed 75% of the birds infected (Cole and Friend 1999). In contrast, histomoniasis is less severe in gray partridge (Perdix perdix) and northern bobwhites (Colinus virginianus) (Cole and Friend 1999).

Pathologic effects of Histomonas appear to involve an interaction with fecal bacteria. Patho­logic effects are fully present only with the full complement of fecal bacteria and are reduced in the presence of single species or limited com­binations of fecal bacteria (Franker and Doll 1964, Springer et al. 1970). Histomonas does not colonize germ-free animals (Franker and Doll 1964, Kemp 1974).

A presumptive diagnosis can be based on the presence of characteristic gross pathology and history. Laboratory confirmation generally involves microscopic examination for living flagellates with one or two flagellae, an axostyle- like projection, and a 6-20 mμ diameter size; these organisms also can be cultured in Dwyer's medium (BonDurant and Wakenell 1994).

population effects Turkeys are highly sus­ceptible to Histomonas meleagridis infections (his- tomoniasis, blackhead). The parasite can cause catastrophic losses in both wild and domestic turkey populations (Cole and Friend 1999).

special problems One interesting case study involving Histomonas in wildlife occurred in the southeastern United States; it illustrates the complexity of parasitic interactions with their hosts. In the 1960s, there was evidence that wild turkey populations in the southeastern United States were declining; it was proposed that the turkeys were suffering from histomoniasis transmitted from bobwhites (Colinus virginianus) infected with Histomonas-infected Heterakis gal­linarum eggs (Kellogg and Reid 1970). But the bobwhites carried Heterakis bonasae rather than H. gallinarum, and only H. gallinarum is known to carry Histomonas (Kellogg and Reid 1970). Thus bobwhites seemed relatively unimportant in contaminating soil with Histomonas-bearing Heterakis eggs (Lund and Chute 1971). However, Indian red jungle fowl (Gallus gallus murghi) previously released in the range of eastern wild turkey (Meleagris gallopavo silvestris) carried Histomonas spp. and were considered a more likely reservoir host for histomoniasis (Kellogg et al. 1972).

control A variety of anthelminthics are used in commercial operations (BonDurant and Wakenell 1994). Because of the high suscepti­bility of turkeys compared to chickens, wild and domestic turkeys should not have exposure to habitat frequented by chickens; the develop­ment in the turkey industry of confinement rearing has reduced exposure to histomonads (BonDurant and Wakenell 1994).

Opportunistic Soil and Water Organisms

ACANTHAMOEBA SPP.

causative agent (classification, morphology) Acanthamoeba is a genus of amebae that generally are free living, but include several species of that can be pathogenic to wild­life, domestic animals, and humans (Visvesvara 1999, John 2001). The genus derives its name from the distinctive feature of its acanthopodia (Gr. acanth: spine or thorn), tapering spike-like pseudopodia, and is classified in the Amoebo- zoa (Adl et al. 2005). Trophozoites average about 24 to 56 μm in length (John 1993). Species often have been named on the basis of variation in cyst morphology among Acanthamoeba (John 1993), but are not easily distinguished (Costas and Griffiths 1986, John 1993).

host range and distribution Acan- thamoeba are distributed worldwide in soil and water (John 1993). Although generally free-living amebae, they are able to cause dis­ease and even death in numerous animals, including humans (John 2001).

LIFE CYCLE AND VARIATIONS The life cycle is simple and includes a motile feeding tropho­zoite (ameba stage) undergoing binary fission and a resting cyst stage (John 2001).

reservoirs and transmission Under most conditions, Acanthamoeba spp. are free- living phagotrophs in freshwater ponds, rivers, lakes, soil, sewage, and sludge, feeding mainly on bacteria (Ma et al. 1990, Visvesvara and Stehr-Green 1990, John 2001). They occasion­ally also have been isolated from brackish water and seawater (Sawyer et al. 1976). There is no evidence for an animal reservoir or carrier host for Acanthamoeba (John 1993).

Infections result from environmental exposure; animal-to-animal transmission is unknown (John 2001). Some infections are associated with aquatic exposure (John 2001). In other cases, infections may occur through the lower respiratory tract or through ulcers of the skin or mucosa, or direct contact with the eye (John 1993).

CLINICAL EFFECTS AND DIAGNOSIS Acan- thamoeba infections can involve the central nervous system, eye, and other organs; invasion of the central nervous system appears to be by the circulation, with the initial focus of ame­bic infection occurring elsewhere in the body (John 2001). Syndromes can include an amebic meningoencephalitis, granulomatous amebic encephalitis, and keratitis (John 1993). Both the trophozoite and cyst stages can occur in tissues of infected animals (John 2001).

Primary amebic meningoencephalitis usually is fatal in humans (John 1993), and generally is diagnosed by microscopic detection of living or stained amebae in cerebrospinal fluid or by cul­ture (John 2001). Granulomatous amebic enceph­alitis is diagnosed by microscopic detection of trophozoites in cerebrospinal fluid or detection of trophozoites or cysts in brain tissue. Corneal infections can be diagnosed by microscopic detec­tion of amebae from corneal scrapings or histo­logic examination of corneal tissue (John 2001).

Acanthamoeba can be difficult to culture from infected animals (John 2001). Species of Acanthamoeba are diagnosed by analysis of isoenzyme patterns with electrophoresis and isoelectric focusing (Daggett et al. 1985), restriction enzyme analysis of mitochondrial DNA (Byers et al. 1983, Costas et al. 1983), as well as restriction endonuclease digestion of whole-cell DNA (McLaughlin et al. 1988).

In general, there is not a good correlation between pathogenicity and species of Acan- thamoeba (John 1993). However, there are genetic markers to help distinguish pathogenic from nonpathogenic strains of Acanthamoeba (Howe et al. 1997).

population effects Although numer­ous wild and domestic animals are susceptible to infection, there are no reported population effects of any significance (John 2001).

special problems Acanthamoeba spp. can serve as host for a number of pathogenic agents, and enhance their survival in soil or water. The capacity for some bacteria not normally consid­ered soil and water organisms to survive in the environment for extended periods is influenced by their ability to successfully infect free-living Acanthamoeba. After being ingested by amebae, the bacteria grow, multiply, and eventually lyse the amebae and are released back into the envi­ronment. In addition to serving as a host for the soil saprophyte Listeria monocytogenes (Ly and Muller 1990), this unique relationship has been linked with extended environmental survival for Francisella tularensis (Abd et al. 2003), Salmonella enterica (Tezcan-Merdol et al. 2004), Chlamydia pneumoniae (Essig et al. 1997), Pasteurella multo- cida (Hundt and Ruffolo 2005), Mycobacterium avium (Cirillo et al. 1997, Steinert et al. 1998), and Mycobacterium leprae (Jadin 1975), among others. Thus, their extended capacity to survive in the environment by parasitizing amebae com­plicates the development of strategies to control these bacteria. There also is evidence that Acan- thamoeba can serve as suitable hosts for echovi­ruses and polioviruses (Danes and Cerva 1981).

control All work with prevention and control has been directed toward human cases.

There is no known satisfactory treatment for primary or granulomatous amebic meningo­encephalitis (John 2001). There are several treatments for corneal infections, including topical treatments, epithelial debridement, and systemic ketoconazole (John 2001), as well as protective treatment of contact lenses (Hughes et al. 2003). No work has been reported with wildlife or domestic animals.

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