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Trichomonas

  • Esther Collántes-Fernández
  • Marcelo C. Fort
  • Luis M. Ortega-Mora
  • Gereon ScharesEmail author
Chapter

Abstract

The most widely known trichomonad in veterinary medicine is Tritrichomonas foetus. It is the etiologic agent of bovine tritrichomonosis, a sexually transmitted disease in extensively managed herds throughout many geographic regions worldwide. The same trichomonad species is also regarded as the causative agent of chronic diarrhea in the domestic cat, although more recent studies observed molecular differences between bovine- and feline-derived T. foetus. Trichomonosis in cats has a worldwide distribution and is mainly present among cats from high-density housing environments. Other trichomonads are found as inhabitants of the gastrointestinal tract in birds, such as Trichomonas gallinae. Particularly, Columbiformes, Falconiformes, Strigiformes, and wild Passeriformes can be severely affected by avian trichomonads. Diagnosis of trichomonosis is often complicated by the fragility of the parasite. To ensure valid test results, it is essential to collect and handle specimens in the right way prior to analysis. Cultivation tests, the specific amplification of parasites, or a combination of both test methods is the most efficient and most commonly used way to diagnose trichomonosis in animals. Bovine tritrichomonosis is mainly controlled by the identification and withdrawal of infected animals from bovine herds. The control of feline and avian trichomonosis relies mainly on preventive measures.

Keywords

Tritrichomonas foetus Trichomonas gallinae Bovine tritrichomonosis Feline tritrichomonosis Avian trichomonosis 

14.1 Morphology, Life Cycle, and Host-Pathogen Interactions

In veterinary medicine the most widely known trichomonad is Tritrichomonas foetus. It is located in the urogenital tract of cattle and considered the etiologic agent of bovine tritrichomonosis, a sexually transmitted disease throughout many geographic regions worldwide (Bondurant 2005; Ondrak 2016). The same trichomonad species was initially described in 1999 and finally confirmed in 2003 to be the causative agent of chronic diarrhea in the domestic cat (Gookin et al. 1999; Levy et al. 2003). Trichomonosis in cats has a worldwide distribution and is mainly present among cats from high-density housing environments such as catteries, shelters, or breeding facilities (Yao and Köster 2015). Other trichomonads in animals, for example, Tritrichomonas suis in swine—presumably genetically identical to T. foetus of bovine origin—are commensals and rarely involved in disease. T. suis was once thought to cause atrophic rhinitis in pigs, further studies were not able to establish a causal relationship, and T. suis is now considered a harmless nasal and gastrointestinal commensal in swine (BonDurant and Honigberg 1994). Isolates of T. suis that reside in the stomach, caecum, and nasal cavity of pigs are not of clinical significance to their porcine hosts (Fitzgerald et al. 1958; Hibler et al. 1960; Pakandl 1994; Mostegl et al. 2011).

Trichomonads are occasionally observed in the feces from dogs with diarrhea (Gookin et al. 2005). The parasite was also detected by culture in the feces of 17.2% of puppies from French breeding kennels, which indicates that T. foetus may be a common parasite in dogs (Grellet et al. 2010). However, molecular identity of the enteric trichomonads observed in these dogs was not investigated, and the relevance for dogs is not clear. More studies are required to determine the prevalence and clinical significance of T. foetus infection in dogs, since the finding could be also attributed to opportunistic overgrowth of the commensal, Pentatrichomonas hominis.

In human medicine, the most studied and relevant is Trichomonas vaginalis that affects over 150 million people worldwide and is the most common non-viral sexually transmitted disease (Van der Pol 2007).

Other trichomonads are found as inhabitants of the gastrointestinal tract in birds such as Tetratrichomonas gallinarum and Trichomonas gallinae. Particularly, Columbiformes, Falconiformes, Strigiformes, and wild Passeriformes can be severely affected by avian trichomonads, whereas the majority of infections in Galliformes and Anatiformes are subclinical although severe infections are occasionally reported (Amin et al. 2014).

Other examples of trichomonads found as inhabitants of the gastrointestinal tract are Trichomonas muris of mice and Pentatrichomonas hominis of a variety of vertebrate species (BonDurant and Honigberg 1994).

14.1.1 Morphology

Trichomonads are taxonomically framed in the Parabasalia class and Trichomonadida order. This order includes protists with a parabasal apparatus and three to five anterior kinetosomes and one posterior kinetosome. They usually bear flagella and have a conspicuous pelta-axostyle complex, and the recurrent flagella are often associated with a lamellar undulating membrane underlain by a striated costal fiber (Adl et al. 2005). The number of free flagella characterizes each genus of the family Trichomonadidae. Thus, the genus Tritrichomonas is characterized by having three free flagella, whereas the genera Tetratrichomonas and Pentatrichomonas possess four and five flagella, respectively. Among the various species of trichomonads thus far identified, only a number of them are regarded as pathogens (BonDurant and Honigberg 1994).

14.1.1.1 Tritrichomonas foetus

Bovine T. foetus isolated from the urogenital tract of cattle and feline isolates found in the gastrointestinal tract of the domestic cat are morphologically indistinguishable. However, there appears to be no association between T. foetus infection in cats and reported exposure to cattle (Gookin et al. 2004). There is an ongoing debate whether T. foetus from cattle and cats should be placed into separate species. A molecular separation of feline and bovine isolates of T. foetus based on a number of gene loci—summarized by Yao and Köster (2015)—seems to be possible, but on the transcriptomic level, a separation remains difficult (Reinmann et al. 2012; Slapeta et al. 2010, 2012; Sun et al. 2012; Morin-Adeline et al. 2014, 2015b). There is evidence that T. foetus isolates from cats and cattle show differences in pH tolerance (Sect. 14.2.2), and T. foetus from cats are able to survive a passage through the alimentary tract of slugs (Morin-Adeline et al. 2015a; Van der Saag et al. 2011). Thus, some authors believe that feline T. foetus represents a different species, and a change in name—to T. blagburni—has been proposed (Walden et al. 2013). It has been hypothesized that feline T. foetus has extended its host range into the bovine reproductive tract (Morin-Adeline et al. 2015a). The picture becomes even more complicated because T. foetus isolated from cattle seems to be morphologically and genetically identical to Tritrichomonas suis, that is, a commensal observed in the nasal cavity, stomach, cecum, and colon of the domestic pig (Felleisen 1998; Hampl et al. 2001; Tachezy et al. 2002; Reinmann et al. 2012; Slapeta et al. 2012; Sun et al. 2012). Consequently, it was assumed that T. suis and T. foetus belong to the same species (Tachezy et al. 2002; Lun et al. 2005; Frey and Müller 2012; Yao and Köster 2015). However, more recent epidemiological studies suggest that cross-species transmission from pigs to cattle on the same farm—e.g., by exposure to T. foetus-contaminated pig feces—is unlikely to occur (Mueller et al. 2015).

T. foetus has a trophozoite stage, with a pyriform or ovoid appearance, and a size ranging from 8 to 18 μm in length and 4 to 9 μm in width (BonDurant and Honigberg 1994) (Fig. 14.1). The trophozoite has several structures with locomotor function such as flagella and the undulating membrane. The flagella originate from the basal bodies or kinetosomes—located in the apical pole of the cell. Three of the flagella are of similar length to each other and are directed forward, while the fourth flagellum, called the recurrent flagellum, is directed toward the posterior part of the body, associated with it by an undulating membrane, and continues as a free flagellum beyond the posterior end of the undulating membrane (Taylor et al. 1994; Benchimol 2004).
Fig. 14.1

Tritrichomonas foetus trophozoites (size, 8–18 × 4–9 μm): schematic and microscopic representation

The internal organelles of T. foetus are similar to those of other trichomonads. The cytoplasm contains a series of support elements, including the pelta-axostyle complex, the costal fiber bordering the recurrent flagellum, and the parabasal apparatus. These elements together with the flagellum make up the cytoskeleton (Benchimol 2005). The axostyle originates in the same area of the birth of the flagellum—surrounded at this point by a chromatin ring—and is directed toward the back of the parasite, making prominence at the posterior end. The pelta is a semilunar structure, very little developed, located in the anterior part of the axostyle. Both structures form the pelta-axostyle complex—composed of groups of connected microtubules—forming a kind of slit that houses the nucleus and parabasal bodies (Benchimol 2004, 2005). In T. foetus, the axostyle has two functions; it serves as a support organelle and participates in the processes of cell division, allowing the constriction of the nuclei during the karyokinesis (Ribeiro et al. 2002). The costa is a rigid structure that sits on the inner margin of the undulating membrane and serves as a support. The parabasal filaments are also filaments striped perpendicularly, and their mission seems to support the parabasal body—i.e., the Golgi complex. The parabasal filaments and the parabasal body constitute the parabasal apparatus, located in the anterior part of the cell (Benchimol 2004, 2005). T. foetus has a simple anterior nucleus and hydrogenosomes, which appear as electro-dense corpuscles that act as functional substitutes for mitochondria. Other cellular components that can be observed in the cytoplasm are free ribosomes, polysomes, glycogen granules, vesicles, and vacuoles related to processes of endocytosis, digestion, and transport (Benchimol 2004, 2005). Under unfavorable conditions, such as a low concentration of nutrients, the presence of certain drugs such as griseofulvin, or abrupt changes in temperature, trophozoites internalize the flagellum and acquire a form of pseudocyst, which is not surrounded by a manifest cell wall (Pereira-Neves and Benchimol 2009).

As regards nutrition, trichomonads lack a cytostome; they are able to capture food through the cell surface by means of pinocytosis and phagocytosis, with the resulting formation of food vacuoles of different size. Like other trichomonads that inhabit body cavities, T. foetus feeds mainly on bacteria, whose proliferation depends on the environment conditions where the parasite is based (Petrin et al. 1998). From the metabolic point of view, T. foetus is unable to de novo synthesize purine and pyrimidine nucleotides, as well as complex phosphoglycerides or cholesterol. The parasite obtains its energy through the anaerobic catabolism of carbohydrates, although the trichomonads are able to survive in the presence of oxygen. As already noted, trichomonads lack mitochondria but possess hydrogenosomes that produce molecular hydrogen in anaerobiosis and reduce oxygen. In this way, the parasite manages to keep the pH of its environment close to neutrality favoring its own development (Kleydman et al. 2004).

The reproduction of T. foetus—like that of all trichomonads—is asexual (Petrin et al. 1998). The parasite divides by longitudinal binary fission in which the nuclear membrane persists—a type of mitosis referred to as cryptopleuromitosis. In addition, when compared to the trophozoite form, pseudocysts present a different mitosis model, since they first divide the nuclei without dividing their cytoplasm, leading to the formation of multinucleated polymastigotes that persist if the cells are maintained under conditions of stress. When the environmental conditions are again favorable, flagella are externalized, and the new flagellated trophozoites emerge from the multinucleated cells (Pereira-Neves and Benchimol 2009).

14.1.1.2 Trichomonas gallinae and Tetratrichomonas gallinarum

T. gallinae is the only trichomonad species with a clear pathogenic potential for birds (BonDurant and Honigberg 1994; Amin et al. 2014). T. gallinarum is commonly found in the large intestine of gallinaceous and anseriform birds, yet its role in causing disease either in naturally infected chickens and turkeys or via experimental infection is under discussion (Amin et al. 2014). T. gallinae trophozoites have an ovoidal to pyriform shape with a size of about 7–11 μm. They are provided with four free anterior flagella and a fifth recurrent one, which does not become free at the posterior pole as it extends for only two-thirds of the body length (Tasca and De Carli 2003; Mehlhorn et al. 2009). Trophozoites of T. gallinarum appear mostly pear shaped and range in size from 6 to 15 μm (Clark et al. 2003). They also have four free anterior flagella and a fifth recurrent one, which becomes free at the posterior pole. Another difference to T. gallinae is the occurrence of a sphere of lacunes of the endoplasmic reticulum surrounding in a regular distance the nucleus with its typical perinuclear membranes. Furthermore, the food vacuoles appear to be very large (Mehlhorn et al. 2009).

14.1.2 Life Cycle

14.1.2.1 Tritrichomonas foetus in Cattle

Bovine T. foetus is located in the genital tract of its natural hosts, Bos taurus taurus and Bos taurus indicus cattle (Skirrow and BonDurant 1988; Bondurant 2005; Sager et al. 2007). The preferred location of the parasite in the bull is the preputial cavity—concentrating mainly in the penile mucosa and adjacent areas of the posterior preputial mucosa—specifically on the surface of the stratified squamous epithelium of the penis and the proximal foreskin in the fornix area (Clark et al. 1974; Parsonson et al. 1974; Parker et al. 1999). This epithelium undergoes numerous folds—resulting in a greater development of crypts—where T. foetus can develop properly by providing a suitable microenvironment for facultative or microaerophilic anaerobic microorganisms (Rhyan et al. 1999). Infection may persist for the life of the bull, in spite of the presence of a measurable humoral immune response in the preputial cavity (Rhyan et al. 1999; Campero et al. 1990; Flower et al. 1983).

In the female, once the infection has occurred, the parasite colonizes the surface of the entire genital system—vagina, cervix, endometrium, and oviduct—in a period of 2 weeks (Parsonson et al. 1976). As observed in natural infections, the parasite is preferentially concentrated in the folds of the cervix (BonDurant 1997). The infection is self-limiting, and the parasite disappears simultaneously from all areas of the genital tract after a period of at least 90–95 days (Parsonson et al. 1976; Rae et al. 2004; Bondurant 2005). In experimental infections of nonpregnant heifers, T. foetus infection is typically cleared from the uterus and vagina between weeks 6 and 12 following infection (Parsonson et al. 1976; Anderson et al. 1996; BonDurant et al. 1993; Skirrow and BonDurant 1990a). A very small proportion of cows in infected herds—a fraction less than 1%—have been shown to remain infected throughout pregnancy and into the following breeding season. Fortunately, such carrier cows are rare (Bondurant 2005).

Under natural conditions, T. foetus is transmitted directly from an infected animal to a healthy animal, almost exclusively through natural mating (Bondurant 2005). The bulls become infected during the mating of infected cows, remaining asymptomatic carriers (Fig. 14.2). Very rarely, however, the parasite can be transmitted by other routes, for example, mechanically during the practice of artificial insemination or vaginal examination, if contaminated material is used—e.g., using the same glass rod or insemination pipette for different cows or not properly disinfected specula (Murname 1959; Goodger and Skirrow 1986). Mechanical transmission seems to be possible through a healthy bull—i.e., from an infected cow to a receptive cow—if the time between two services does not exceed 20 min (Clark et al. 1977; Goodger and Skirrow 1986; Bondurant 2005; Ondrak 2016).
Fig. 14.2

Life cycle of bovine Tritrichomonas foetus (David Arranz Solís from SALUVET-UCM is acknowledged for providing this graph)

T. foetus was shown to be able to survive in cryopreserved semen and may be present in semen if it is contaminated with preputial fluid during manual collection (Blackshaw and Beattie 1955). Given the resistance of this parasite in fresh, pure, or diluted semen, refrigerated and even cryopreserved, there is the possibility of transmission through artificial insemination with contaminated semen (Bondurant 2005).

Because bulls tend to mount each other, feces in the preputial cavity is commonly found. This fecal material may contain non-T. foetus trichomonads, such as Pentatrichomonas hominis and any number of Tetratrichomonas species that have been shown to be nonpathogenic (Taylor et al. 1994; Campero et al. 2003; Hayes et al. 2003). The opportunity for transmission of T. foetus between males is regarded as very limited.

14.1.2.2 Tritrichomonas foetus in Cats

Feline T. foetus appears to be host adapted, i.e., adapted to the intestinal tract of cats. After experimental orogastric infection of kittens, feline T. foetus has been demonstrated to colonize the lumen of the ileum, caecum, colon, and rectum 203 days after infection (Gookin et al. 2001). In naturally infected cats, massive numbers of trichomonads can be observed at the surface of the colonic epithelium and within the colonic crypts (Yaeger and Gookin 2005). Although the presence of T. foetus in the uterus of a cat with pyometra has been described, it has been speculated that the parasite could have accidentally accessed the genital area through contact with contaminated feces (Dahlgren et al. 2007). However, colonization of the reproductive tract in both male and female cats from breeding grounds with a high prevalence of feline tritrichomonosis has not been observed (Gray et al. 2010). Feline T. foetus infection occurs by direct fecal-oral transmission. Infected cats are shedding trophozoites with their feces, and transmission occurs when two or more cats share the same litter box (Fig. 14.3). Trophozoites would adhere to the hair of the animals and could be ingested during grooming (Gookin et al. 2004; Tolbert and Gookin 2009). The viability of the parasite in the environment is limited though it can withstand several days at room temperature facilitating its transmission (Hale et al. 2009). T. foetus-contaminated food and less likely water may be also a relevant route for transmission. Further, shedding of viable T. foetus has been demonstrated in some slug species, which were fed cat food spiked with trophozoites of a feline T. foetus isolate, suggesting that invertebrates like slugs could play a role as mechanical vectors (Van der Saag et al. 2011).
Fig. 14.3

Life cycle of feline Tritrichomonas foetus (David Arranz Solís from SALUVET-UCM is acknowledged for providing this graph)

14.1.2.3 Trichomonas gallinae and Tetratrichomonas gallinarum

The rock pigeon—Columba livia—was regarded as the primary host of T. gallinae and has been considered responsible for the worldwide distribution of this protozoal infection (Stabler 1954; Harmon et al. 1987). Other species within the Columbiformes, Falconiformes, Strigiformes, and, most recently, different Passeriformes have been recognized as potential hosts (Forrester and Foster 2008; Robinson et al. 2010). However, only a few natural occurrences of trichomonosis have been reported in gallinaceous birds like turkeys and chickens (Levine and Brandly 1939). The preferred site for T. gallinae is the upper digestive tract including the mouth, pharynx, esophagus, and crop, with the parasite rarely found posterior to the proventriculus (Cauthen 1936). Transmission by direct contact seems to be the most efficient route to establish an infection—e.g., via the crop milk from infected parent birds to the nestlings during feeding (Stabler 1954). In adult pigeons, the infection can occur during courtship while raptors can be infected from prey animals carrying the parasite. The infection of turkeys and chickens happens mainly via drinking water contaminated by pigeons (BonDurant and Honigberg 1994). Trichomonas gallinae is unable to form true cysts, even though cyst-like stages—pseudocysts—have been reported (Tasca and De Carli 2003; Mehlhorn et al. 2009). These pseudocysts may provide another route of transmission and an environmentally resistant stage during unfavorable conditions.

Tetratrichomonas gallinarum locates in the intestinal tract of different poultry species including chickens, turkeys, guinea fowl, quails, ducks, and geese and can be transmitted via consumption of contaminated food (BonDurant and Honigberg 1994). Pseudocysts of T. gallinarum have been reported in vivo and in vitro possibly protecting the parasite during fecal-oral transmission (Mehlhorn et al. 2009).

14.1.3 Host-Pathogen Interactions

14.1.3.1 T. foetus Infection in Cattle

Once infected, the male acts as an asymptomatic carrier throughout his life (Clark et al. 1974; Parsonson et al. 1974; Parker et al. 1999). Minor histological changes are observed with increased accumulations of neutrophils followed by an infiltrate of lymphocytes and plasma cells penetrating into the intraepithelial area and coalescing in the subepithelium to form lymphoid nodules (Rhyan et al. 1999; Bondurant 2005).

In the female, 2 weeks after infection, T. foetus may have colonized the different parts of the genital tract (Parsonson et al. 1976). The preferred location is in the cervix and cervicovaginal mucus, but the number of parasites varies throughout the estrus cycle, being higher in the days prior to estrus. The establishment of the parasite in the genital tract of the female does not seem to interfere with the fertilization nor with the early development of the embryo (Bielanski et al. 2004). In heifers experimentally infected with T. foetus, conceptus deaths peaked at 50–70 days of gestation (Parsonson et al. 1976). Occasional abortions of fetuses older than 4-month gestational age are reported, but typically losses occur 2 months earlier (Bondurant 2005). The infection in the female is usually self-limiting, disappearing between 2 and 4 months after the loss of the conceptus. The immunity that develops is not permanent and usually lasts for about 6 months; after 6 months, the female is again susceptible to infection. Carrier cows—these are cows that remain infected for at least 10 months—seem to fail to develop a protective immune reaction against T. foetus. Notable lesions in the maternal endometrium and fetal envelopes have been described only at the time of fetal loss (Parsonson et al. 1976).

The mechanisms of pathogenic actions that underlie the loss of the embryo or fetus are not known with accuracy and may include (1) the direct mechanical action of the parasite, (2) the adverse effects of enzymes secreted by the parasite, and (3) the alteration of the intrauterine environment mainly by antiparasitic inflammatory reactions of an infected dam (reviewed by Bondurant (2005) and Campero and Cobo (2006)). The increase in the number of microorganisms in the female genital tract occurs slowly and probably does not produce any relevant damage until this number exceeds a certain threshold. This fact would explain the long period of time between infection and loss of the conceptus.

14.1.3.2 T. foetus Infection in Cats

Few studies have examined the interaction of feline T. foetus with intestinal epithelium (recently reviewed by Yao and Köster (2015) and Tolbert and Gookin (2016)). Recent studies examining T. foetus infection in a co-culture model with monolayers of porcine intestinal epithelial cells suggest that adhesion to the intestinal epithelium occurs by means of specific receptor-ligand interactions (Tolbert et al. 2013). Pathogenesis of T. foetus on the intestinal epithelial cells has been suggested to be both contact-dependent and contact-independent. In the former, a cytopathic effect is mainly exerted via apoptosis induced by cell-associated proteases, whereas extracellular proteases are the major players in contact-independent cytotoxicity. Extracellular proteases may also play a role in evading complement killing.

14.1.3.3 T. gallinae and T. gallinarum Infections in Birds

The severity of the disease depends on the susceptibility of the infected birds together with the pathogenic potential of the incriminated strain and the stage of infection (Cooper and Petty 1988; Cole and Friend 1999). The severity of pathologic lesions of T. gallinae in the upper digestive tract varies from a mild inflammation of the mucosa to caseous areas that block the esophageal lumen (Stabler 1954). Some virulent strains are able to create diphtheritic membranes—associated with fibrinous lesions in internal organs such as the liver, lungs, and peritoneum—resulting in high mortality (Narcisi et al. 1991). Strains of moderate virulence are often associated with caseous abscesses in the upper digestive tract and oropharyngeal region, whereas no appreciable lesions are produced by avirulent strains (Cole and Friend 1999). In vitro, T. gallinae proliferation has been associated with a disintegration of the cell monolayer, and genetically different T. gallinae isolates caused diverse magnitudes of cytopathic effects on different cell lines (Amin et al. 2012a). However, little is known concerning the mechanism by which T. gallinae causes pathological changes in its hosts. Proteolytic proteins secreted by the parasite have been identified as contributing to the detachment of a cell monolayer (Amin et al. 2012b).

Various studies investigated the pathogenicity of T. gallinarum either in naturally infected chickens and turkeys or via experimental infection, with contradicting outcomes as reviewed by Amin et al. (2014). Recent studies have shown that in vitro, T. gallinarum has no destructive effect on cells and, in vivo, did neither produce clinical signs nor macroscopic or microscopic lesions in turkeys and specified pathogen-free chickens (Amin et al. 2011).

14.2 Clinical Effects and Diagnosis

14.2.1 Clinical Effects

14.2.1.1 Cattle

The clinical effects produced by the disease occur only in female cattle, causing early abortion and temporary infertility (reviewed by BonDurant (1997, 2005, 2007), Yule et al. (1989a), Rae and Crews (2006)). In males, T. foetus infection is asymptomatic and affects neither semen quality nor sexual behavior, but bulls can shed the organism indefinitely (Parsonson et al. 1974; Rhyan et al. 1999).

The parasite multiplication causes inflammation of the endometrium, cervical, and vaginal mucous membranes in cows or heifers following the infection at breeding (Parsonson et al. 1976; Rhyan et al. 1988; Anderson et al. 1996). Consequently, signs of mild vaginitis, cervicitis, or endometritis, such as mucopurulent vaginal discharge, may be observed, although generally there are no overt signs. Conception apparently proceeds normally, but almost all conceptuses are lost at some time early in gestation with early fetal death and resorption but also abortion—with a peak loss at 70–90 days (Parsonson et al. 1976; Bielanski et al. 2004). Infection can result in fetal maceration and pyometra. The consequence is infertility (Parsonson et al. 1976; BonDurant 1985; Ball et al. 1987; Anderson et al. 1996). Abortions of fetuses typically occur around 2 months of gestational age. Abortions of fetuses older than 4 months of gestational age due to trichomonosis have been occasionally reported. If the affected cow undergoes early fetal loss, it may cycle regularly without showing any signs but a prolonged inter-estrous interval (BonDurant 1985). Pyometra occurs in less than 5% of infected cows and is followed—as the corpus luteum of pregnancy is maintained—by a large purulent response (Rhyan et al. 1988); pyometra is probably a result of bacterial contamination that occurs at the time of fetal loss, when the cervix is likely to relax sufficiently to admit contamination from outside the environment (Rhyan et al. 1995a). Cows that are infected with T. foetus typically clear the infection within a few months, i.e., after three cycles (Parsonson et al. 1976). Immunity, however, is not permanent, and the cow will be subject to reinfection and embryonic death in subsequent breeding periods, and, as mentioned earlier, some infected cows may carry infections into the next breeding season (Skirrow 1987; Mancebo et al. 1995).

In an infected herd, bovine tritrichomonosis is associated with lowered fertility. The usual signs in the herd include return to estrus 1–3 months after breeding. At pregnancy exam time, a number of early pregnancies and open cows are observed. The period of infertility may last for another 2–6 months as a result of the infection. Other clinical features of the disease in the herd include many services per conception, poor pregnancy rates, long calving intervals, and calf crop reduction. In addition, the calving season is spread out causing batches of calves of different ages with a wide variation in weaning weights (Clark et al. 1983a; McCool et al. 1988; Rae 1989; Collantes-Fernandez et al. 2014).

14.2.1.2 Cats

T. foetus is recognized as an important cause of diarrhea in domestic cats. Typical clinical signs in natural infections are chronic or intermittent large bowel diarrhea, which can vary from subclinical to intractable (reviewed by Gookin et al. (1999), Gookin et al. (2001), Foster et al. (2004), Manning (2010), Yao and Köster (2015)). The feces are described as yellow-green in color, gassy, and malodorous with typical signs of colitis including fresh blood, mucus, fecal incontinence, tenesmus, and flatulence. The consistency of the feces can vary from liquid to semi-formed or cow pat (Stockdale et al. 2009). Severe cases may be accompanied by marked inflammation of the anal region, fecal incontinence, and rectal prolapse (Gookin et al. 1999; Foster et al. 2004; Tolbert and Gookin 2009; Bell et al. 2010). In addition, some infected cats have been reported showing systemic signs including anorexia, depression, vomiting, and weight loss (Stockdale et al. 2009). Mortality is extremely rare and only reported in kittens and is presumably caused by endotoxic shock because of deep lesions in the colonic mucosa (Holliday et al. 2009). The majority of infected cats maintain good body condition and appetite without signs of systemic illness (Gookin et al. 1999, 2001; Tolbert and Gookin 2009). The long-term prognosis for T. foetus-infected cats is usually good and most will eventually overcome the infection. Remission could take between 4 months and 3 years, with irregular episodes of diarrhea of variable length (Gookin et al. 1999, 2004). No abnormalities are routinely noted on hematology and serum biochemistry profile of some cats, though they remain infected and continue shedding the organism despite clinical improvement, i.e. these cats represent asymptomatic carriers. (Manning 2010). Some positive cats were also infected with other pathogens, like Cryptosporidium spp., Giardia spp., coccidian, or feline immunodeficiency virus (Gookin et al. 1999; Stockdale et al. 2009); concurrent infections may contribute and increase susceptibility and vulnerability to intestinal disease in infected cats.

14.2.1.3 Other Animals

In birds the two trichomonad species T. gallinarum and T. gallinae are commonly found (reviewed by Amin et al. (2014)). T. gallinarum parasitizes the large intestine of gallinaceous and anseriform birds. T. gallinarum induces usually a latent infection in the absence of clinical signs and lesions, and it is not clear whether T. gallinarum should be regarded a primary pathogen (Amin et al. 2011; Friedhoff et al. 1991). However, the presence of T. gallinarum may aggravate primary diseases—e.g., caused by Histomonas meleagridis—and coinfections have been observed (Grabensteiner and Hess 2006). T. gallinae is of veterinary and economic importance, as it causes avian trichomonosis, a disease with important medical and commercial implications, which is known as pigeon canker, canker, roup, or Gelber Knopf (Amin et al. 2014). T. gallinae is located in the upper digestive tract of pigeons, causing lesions (BonDurant and Honigberg 1994). The disease is characterized by greenish fluid and caseous lesions—whitish-yellowish fibrinous material—on the oropharyngeal membranes that can block the lumen of the esophagus impairing drinking and feeding. Clinical signs associated with avian trichomonosis 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 (Narcisi et al. 1991). Death may occur within 3 weeks of infection. Infected birds can also remain asymptomatic due to the infection with avirulent strains of trichomonads or a lower susceptibility as seen in older birds. Avian trichomonosis may also affect domestic fowl; in earlier studies severe outbreaks have been recorded in chickens and turkeys, but we are not aware on recent cases (Hawn 1937).

14.2.2 Diagnosis

14.2.2.1 Diagnostic Techniques

In cattle, the prescribed test for international trade is the identification of T. foetus by culture or PCR—World Organization for Animal Health (OIE), OIE Terrestrial Manual. The OIE Terrestrial Manual provides protocols for sampling, sample transportation, transport medium, culture media, culture conditions, and how to read out the culture test. In addition, the OIE Terrestrial Manual also provides recommendations for PCR analyses, which can be applied in combination either with or after culture as an ancillary test or—more often—direct as the primary test to examine bovine samples—i.e., preputial material, uterine or vaginal secretions, or abomasal content of aborted fetuses. Protocols to diagnose bovine tritrichomonosis have been described in detail previously (Sager et al. 2007). To diagnose T. foetus infection in cattle, PCR tests may have a higher or at least the same sensitivity as culture tests but have several advantages, because parasites in the sample do not need to be viable and PCR results are rapidly available in contrast to results of culture tests. On the other hand, culture tests are superior to PCR because of their relative easiness (Yao 2013).

In cats, both cultivation and PCR tests are regarded as optimal methods for a sensitive and specific detection of T. foetus in fecal samples, although currently PCR is regarded as the gold standard assay for diagnosis of feline T. foetus infection since detection is independent from parasite viability (Gookin et al. 2002, 2004; Manning 2010; Yao and Köster 2015). Under optimized conditions, PCR is the method of choice when samples have to be shipped—e.g., from practitioner to a veterinary laboratory. Similar to T. foetus in cattle, the success of cultivation tests is largely dependent on the viability of T. foetus in the sample (Hale et al. 2009).

In birds infected by T. gallinae, an immediate diagnosis by direct microscopy of material collected via swabbing the oral cavity during clinical examination or necropsy is possible. Also, T. gallinarum can be observed in fecal material collected from birds—e.g., by swabbing cloacae. However, although the direct detection by light microscopy is fast and inexpensive, it is regarded as insensitive, and low numbers of parasites may not be detected (Amin et al. 2014). Also in birds the use of cultivation for the detection of trichomonads is clearly superior in sensitivity as compared to direct microscopy (Cooper and Petty 1988; Bunbury et al. 2005).

14.2.2.2 Direct Microscopic Examination

In fresh samples it is possible to diagnose the infection with trichomonads by a direct light microscopical examination. Optimal is a 200 to 400-fold magnification. It is advantageous to pre-warm slides, to retain motility of trichomonads. A drop of physiological saline is added to the slide, mixed with a nearly equal volume of material collected, and mounted with a coverslip.

It is possible to apply conventional light microscopy—either using a conventional up-light or an inverted microscope—, phase-contrast microscopy, or dark-field microscopy. In phase-contrast microscopy, it is easier to see flagella. In dark-field microscopy, trichomonads appear as small rolling luminescent footballs. In conventional light microscopy, trichomonads are identified by their characteristic movement, which is described as rolling and jerky. They are flashing, due to their rolling movements. The presence of multiple anterior flagella and the characteristic refractile undulating membrane can be observed. However, one disadvantage of direct microscopy is that at the resolution of a conventional laboratory microscope, the exact number of anterior flagella cannot be determined.

The specificity of direct microscopy is very limited. The identification of the trichomonad species observed by this method is not possible, and confirmatory PCR analyses are necessary. For inexperienced examiners it might be difficult to differentiate trichomonads from other intestinal parasites—e.g., Giardia spp. in cat feces (Yao and Köster 2015). Also the intestinal commensal trichomonad Pentatrichomonas hominis might be misinterpreted as T. foetus.

Another disadvantage of direct microscopy is its low sensitivity. A sample is only positive, if it contains a sufficient number of parasites per milliliter specimen. For example, in cats, the fecal examination by direct microscopy is reported to have a diagnostic sensitivity of only about 14%, and also in bovine tritrichomonosis, direct examination is estimated to be 25% less sensitive than culture diagnosis (Gookin et al. 2004; Sager et al. 2007).

The advantage of direct microscopy as diagnostic tool is its speed and the low cost of examination. This is the reason why direct microscopy is often used by practitioners for the examination of T. foetus in cats and T. gallinae infection in birds (Forrester and Foster 2008; Amin et al. 2014).

Staining of trichomonads is possible using a number of stains, including Giemsa, silver, iron hematoxylin, malachite green, methylene blue, Papanicolaou, and acridine orange (Amin et al. 2011). A fast and inexpensive staining protocol—Giemsa or Diff-Quick and iodine—has been reported (Lun and Gajadhar 1999). Single parasites might be easier to inspect; however, the chance to find small numbers of parasites in a sample might decrease because parasites can no longer be identified by their characteristic movement or flashing essential for parasite identification in low concentrated samples. In tissues, the use of hematoxylin-eosin (HE) and periodic acid-Schiff (PAS) stains was proven to be advantageous for identification of the flagellates, especially in organs that contained only a few protozoal cells (Amin et al. 2011, 2014). Immunohistochemical techniques and in situ hybridization have been successfully applied to demonstrate trichomonads in histological sections of cat or bird tissues, respectively (Rhyan et al. 1995b; Yaeger and Gookin 2005; Liebhart et al. 2006; Mostegl et al. 2012). Immunohistochemical detection using monoclonal antibodies against T. foetus has been shown to be a valuable tool (Hodgson et al. 1990). A protocol for immunohistochemical detection using the monoclonal antibody Mab 34.7C4.4 is provided in the OIE Terrestrial Manual (www.oie.int/international-standard-setting/terrestrial-manual). Staining is also applied to confirm positive cultures by morphological criteria.

14.2.2.3 Culture

In vitro culture can be performed by incubating the samples at 25–37 °C in a growth medium. If parasites are present, their numbers will multiply in the culture over time, increasing the likelihood of their detection. A large number of culture systems for trichomonads and especially T. foetus have been developed and published. Presumably the first culture system for an axenic cultivation of T. foetus isolated from an aborted fetus—i.e., cultivation without bacteria or other living organisms—was reported by a German microbiologist (Witte 1933). From that time on, numerous reports of further cultivation protocols have been published.

Until now diagnostic culturing is of outmost importance for sensitive diagnosis of bovine tritrichomonosis. Also in the diagnosis of tritrichomonosis of cats, cultivation has been widely used for epidemiological studies or diagnostic purposes (Tables 14.2 and 14.3).

In bovine tritrichomonosis cultivation became an important diagnostic tool, because parasite numbers in bovine samples—e.g., preputial smegma or cervico-vaginal mucus—are usually too low to be detected by direct microscopy and a multiplication of parasites after a few days of cultivation increases the chance to find infected bulls.

The number of organisms in preputial secretions has been estimated and ranges from less than 200/mL up to more than 80,000/mL (Skirrow and BonDurant 1988). In bovine tritrichomonosis, diagnostic sensitivity of a single culture test on infected bulls has been estimated to range between 70 and 100% (Skirrow et al. 1985; Schönmann et al. 1994; Parker et al. 1999, 2003a, b). In a large field study, including 2832 mature bulls from 124 beef herds in Argentina, Bayesian estimation revealed a diagnostic sensitivity and specificity of 72.0% (59–87%) and 95.4% (94–96%), respectively (Perez et al. 2006). A repeated testing of bulls—e.g., three times with intervals of several days—has been shown to increase the diagnostic sensitivity of the cell culture test close to 100%. Of 29 samples collected from 5 experimentally infected bulls with resting periods of 2–4 days between samplings, 24 (83%) were determined as positive (Mukhufhi et al. 2003). In another study, consecutive testing over a period of more than 7 months resulted in the determination of an infection rate of 100% in 15 bulls (Clark et al. 1971). For bulls from herds in which T. foetus is endemic, two to four tests per bull may be required to ensure that the bull is not infected (Parker et al. 1999). A sexual rest of bulls for a minimum of about 1–2 weeks prior to sampling increases sensitivity (Yule et al. 1989a).

Also for the analysis of females, i.e., after sampling of cervico-vaginal mucus, sensitivity of cultivation is superior to direct microscopic examination (Simmons and Laws 1957; Skirrow and BonDurant 1988). In female cattle diagnostic sensitivity of culture tests has been reported in the range of 56 and 95% (Kimsey et al. 1980; Goodger and Skirrow 1986; Skirrow and BonDurant 1988; Parsonson et al. 1976). The infection in females is usually cleared within 3 months, and it is often difficult to isolate organisms from female cattle in the late stage of their infection.

In cats, the culture method is reported to have a detection limit of about 2 × 102 trophozoites and a diagnostic sensitivity from 26.4 to 58.8% (Hale et al. 2009; Gookin et al. 2004).

To achieve optimal test sensitivity, it is essential to retain as long as possible viability of trichomonads after sampling. The number of viable organisms decreases progressively after sampling (Todorovic and McNutt 1967; Tedesco et al. 1979; Reece et al. 1983; Skirrow et al. 1985; Kittel et al. 1998; Bryan et al. 1999; Parker et al. 1999). Immediate cultivation is ideal but rarely possible. A 1-day delay is estimated to cause a loss of diagnostic sensitivity of 10% (Sager et al. 2007). Sampling the parasite into transportation media providing nutrients has been shown to be essential for the survival of trichomonads, especially if time between sampling and starting cultivation is exceeding 2 days (Kimsey et al. 1980; Hale et al. 2009). An earlier study showed that physiological saline with 5% fetal serum or lactate Ringer’s solution was effective (Kimsey et al. 1980). A thyoglycolate transport medium was also shown to be suitable; however, sensitivity of the subsequent cell culture test was slightly lower than after transport using InPouch TF medium (BioMed Diagnostics, White City, OR, USA). Today the medium used for later cultivation is often also used as transportation medium—i.e., InPouch TF or Diamond’s medium (Bryan et al. 1999).

An alternative is the direct sampling into a commercially available transport and culture kit—InPouch TF—containing a selective medium, a medium optimized for T. foetus, and a medium repressing the growth of the contaminating bacterial flora. This commercial transport and culture kit is recommended not only for sampling in cattle but also for sampling in cats or birds (Thomas et al. 1990; BonDurant 1997; Gookin et al. 2004; Hale et al. 2009; Yao and Köster 2015).

According to the OIE Terrestrial Manual (www.oie.int/international-standard-setting/terrestrial-manual), bovine samples—after being added to transport media—should be protected from exposure to daylight and extremes of temperature, which should remain above 5 °C and below 38 °C (Bryan et al. 1999). For cat fecal samples, a storage for 1 to 24 h at room temperature (23–25 °C) was superior to a 4 °C storage for the same period of time as shown in experiments performed with fecal samples spiked with different T. foetus concentrations (2 × 102–2 × 104 T. foetus per gram of feces) (Hale et al. 2009).

Several culture media have been found suitable for the cultivation of trichomonads. Overviews on media have been provided in the OIE Terrestrial Manual, www.oie.int/international-standard-setting/terrestrial-manual/, and in several reviews (Skirrow and BonDurant 1988; Sager et al. 2007).

Currently, the most widely used system is InPouch TF, a commercial transport and cultivation kit (Yao 2013). As noncommercial medium the so-called Diamond’s medium is widely used, also in epidemiological studies (Tables 14.2 and 14.3). Modified Diamond’s medium is a trypticase-yeast extract-maltose medium which in most studies was used modified by the addition of heat-inactivated serum—e.g., of 5% heat-inactivated horse or lamb serum (Diamond 1957; Skirrow and BonDurant 1988; Sager et al. 2007). Both the use of modified Diamond’s medium and the InPouch TF kit are recommended for diagnosis of bovine tritrichomonosis by the OIE Terrestrial Manual (www.oie.int/international-standard-setting/terrestrial-manual, accessed 22. Febr. 2017).

The InPouch TF kit seems superior to Diamond’s medium to detect T. foetus infection in bulls (Schönmann et al. 1994; Appell et al. 1993; Mendoza-Ibarra et al. 2012; Yao 2013). As regards the cultivating of cat samples, in one study, the InPouch TF kit was found to be superior to modified Diamond’s medium (Gookin et al. 2004); however, in a recent study, comparison of both systems revealed a higher sensitivity when modified Diamond’s medium was used—ATCC medium 719 (Hale et al. 2009). A retrospective analysis of data revealed no statistical significant differences between cultivation with modified Diamond’s medium and InPouch TF.

A modified Plastridge medium containing antibiotics and antifungal agents as well as heat-inactivated bovine serum was recommended for initial cultivation of trichomonads and can be applied combined with modified Diamond’s medium for subsequent procedures—e.g., sub-cultivation (Reece et al. 1983; Skirrow and BonDurant 1988; Sager et al. 2007). In studies conducted in Argentina, a commercially available modified Plastridge medium has been applied (Mardones et al. 2008).

The preparation of modified Diamond’s medium and test vials, as well as sample processing and reading the results of the culture test, is described in the OIE Terrestrial Manual for bovine samples; many of the recommendations also apply for processing cat and avian samples (www.oie.int/international-standard-setting/terrestrial-manual, accessed 22. Febr. 2017).

Samples collected via preputial scraping—e.g., vigorously by brush, insemination pipette, or aspiration, usually about 0.5–1 mL—can be inoculated directly on top of the medium of a test tube, into the transportation medium, or into the upper chamber of the InPouch TF kit. In contrast, samples collected by preputial washing need to be centrifuged and the supernatant discarded in order to reduce volume. Reading the cell culture tests is performed by microscopic detection of the trichomonads. To increase specificity of the culture test, it is recommended to confirm observed parasites by PCR (Campero et al. 2003).

For cats, voided feces sampled directly from the litter box, rectal swabs obtained from rectal mucous membranes, or feces collected by manual extraction with the aid of fecal loops or by a colon flush technique can be the starting point for trichomonads cultivation (Yao and Köster 2015; Manning 2010; Tolbert and Gookin 2009).

In case of T. gallinae-infected birds, a cotton-tipped applicator moistened with sterile saline is used to swab the oral cavity, and swabs are added to InPouch TF culture devices or other commercial or noncommercial culture media (Forrester and Foster 2008; Rogers et al. 2016; Girard et al. 2014; Krone et al. 2005)).

Microscopic detection of culture-growing organisms can be done by light microscopy, on a wet mount slide prepared directly from the culture or through the plastic wall of the InPouch TF kit using a plastic clip provided by the supplier. The motile organisms may be seen under a standard microscope using a 200-fold or higher magnification. An inverted microscope may be useful for examining culture flasks containing culture medium. Culture media should be inspected by microscopic examination at regular daily intervals—from day 1 to day 7 after inoculation (Bryan et al. 1999; Lun et al. 2000). During the first 4–72 h of culturing, there might be an initial increase of parasite numbers, subsequently followed by a decrease. Organisms may be identified on the basis of characteristic morphological features (OIE Terrestrial Manual, www.oie.int/international-standard-setting/terrestrial-manual, accessed 22. Febr. 2017).

It has been shown for cattle and cats that without test confirmation—e.g., by using specific PCR or sometimes by using staining or electron microscopy—the diagnostic specificity of the culture method remains well under 100%. Therefore, a subsequent PCR analysis of culture-positive samples has been recommended to avoid false-positive findings (Parker et al. 2001; Campero et al. 2003; Ceplecha et al. 2013). Intestinal trichomonads were observed in virgin bull samples submitted for confirmation of InPouch TF culture diagnosis or culture diagnosis using Sutherland medium (BonDurant et al. 1999; Michi et al. 2016). The medium of InPouch TF-Feline is thought to be highly specific to T. foetus, and the morphologically similar flagellates P. hominis and Giardia spp. should not survive longer in this medium than 24 h (Gookin et al. 2003). However, the InPouch TF-Feline medium seems to be not entirely selective as P. hominis could be successfully cultivated and identified after 3 days following inoculation of InPouch TF-Feline medium using cat feces (Ceplecha et al. 2013).

14.2.2.4 DNA Detection

DNA detection has become one of the most important methods for the diagnosis of infections with trichomonads in cattle and cats. The major advantage of DNA detection by PCR is that it is independent of parasite viability and contaminating microbes that may inhibit trichomonad cultivation. Correspondingly, sensitivity of PCR is often reported to be higher than cultivation and direct microscopical examination (recently reviewed by Yao (2013)). However, PCR analysis has also a number of disadvantages since due to its sensitivity it is prone to carry-over and cross-contamination. In addition, samples may contain inhibitory components that may reduce the sensitivity of PCR or even disable amplification. Each lab should validate the entire diagnostic process—including DNA extraction and PCR amplification—prior to carry out PCR detection as laboratory-specific conditions, equipment, or consumables may have an impact on the outcome of the diagnostic process (Hoorfar et al. 2004; Conraths and Schares 2006).

Preputial material for DNA extraction is generally collected by sheath scraping combined with aspiration or by sheath washing. Material obtained by scraping may contain blood or fecal contaminations from outside the preputium, and material collected by preputial washing may contain urine. Blood, fecal components, and urine may act as PCR inhibitors, and, therefore, it is necessary to minimize or avoid such contaminations. Analytical sensitivities of PCR protocols are high and usually sufficient to detect the DNA of a single parasite (Table 14.1). In field samples, however, analytical sensitivity might be much lower and has been reported to be around 100 organisms per sample (Mukhufhi et al. 2003). In an assessment of diagnostic sensitivity carried out with spiked cat feces, ten organisms per 200 mg of feces were detected in 90% of nested PCR tests, and 100 organisms per 200 mg of feces were detected in 100% of nested PCR tests (Gookin et al. 2002).
Table 14.1

Veterinary relevant diagnostic PCRs to detect trichomonads

Target

Type of PCR

Name of primer

Primer, 5′–3′

Name of probe (type of probe)

Probe (type of probe)

Reported specificity

Reported sensitivity

Remarks

Reference

ITS1/5.8S rDNA/ITS2

End-point

TFR1, TFR2

TGC TTC AGT TCA GCG GGT CTT CC, CGG TAG GTG AAC CTG CCG TTG G

NA

NA

Amplifies T. foetus, T. suis, T. mobilensis, T. vaginalis, T. gallinae, T. tenax, P. hominis (Felleisen et al. 1998) . No amplification of bacterial DNA orpurified bovine genomic DNA

One or a few protozoa

Also referred to as pan-trichomonad PCR

Felleisen (1997)

ITS1/5.8S rDNA/ITS2

End-point

TFR3, TFR4

CGG GTC TTC CTA TAT GAG ACA GAA CC, CCT GCC GTT GGA TCA GTT TCG TTA A

NA

NA

Amplifies T. foetus, T. suis, T. mobilensis

One or a few protozoa

Often referred to as T. foetus-specific PCR

Felleisen et al. (1998)

18S rDNA, ITS1, 5.8S rDNA

End-point

TF211A, TF211B

CCT GCC GTT GGA TCA GTT TCG TTA, GCG CAA TGT GCA TTC AAA GAT TCG

NA

NA

Does not amplify Mycoplasma bovigenitalium, Ureaplasma diversum, or bovine genomic DNA

1 pg T. foetus DNA

Reported to produce few unspecific DNA bands

Nickel et al. (2002)

ITS1-5.8S rDNA-ITS2

End-point

Tricho-F/Tricho-R

CGG TAG GTG AAC CTG CCG TT (truncated TRF2, (Felleisen 1997)), TGC TTC AGT TCA GCG GGT CT (truncated TRF1 (Felleisen 1997))

NA

NA

Amplifies T. foetus, T. suis, T. mobilensis based on in silico analyses; amplified Pentatrichomonas hominis

NA

Used in human samples and in a study on cats (Profizi et al. 2013)

Jongwutiwes et al. (2000) and Duboucher et al. (2006)

18S rDNA, ITS1 and 5.8S rDNA

End-point

Forward, reverse

GTA GGT GAA CCT GCC GTT G (5’FAM labeled), ATG CAA CGT TCT TCA TCG TG

NA

NA

Amplifies T. foetus but also trichomonad DNAfrom a variety of genera; T. foetus (157 bp), Tetratrichomonas spp. (170–175 bp), Pentatrichomonas hominis (142 bp)

Accurate typing is possible from both the 1.0 and 0.1 pgtemplates

Using diagnostic size variants from within the internal transcribed spacer 1 (ITS1) region. Incorporation of a fluorescently labeled primer enables high sensitivity and rapid assessment of results for species identification

Grahn et al. (2005)

18S rDNA, ITS1 and 5.8S rDNA

End-point

Forward, reverse 5.8S primer

Forward primer (Grahn et al. 2005); TTC AGT TCA GCG GGT CTT C

NA

NA

Amplified T. foetus (367 bp), Tetratrichomonas sp. (379 bp), Pentatrichomonas sp. (333 bp), T. gallinae (364 bp), and T. vaginalis (363 bp)

0.1 pg

Analysis in a 2% agarose gel and by using fluorescent-labeled primers and 6% polyacrylamide gels; disadvantage: too much template makes typing difficult or impossible; advantage: low costs

Frey et al. (2009)

ITS1/5.8S rDNA/ITS2

End-point nested

TFR3, TFR4 (external); TFITS-F, TFITS-R (internal)

TFR3, TFR4, primer sequences published (Felleisen et al. 1998); CTG CCG TTG GAT CAG TTT CG, GCA ATG TGC ATT CAA AGA TCG

NA

NA

T. foetus-specific

Sensitivity in PBS: 1 organism, 70% ; 10 organisms, 90%; 100 organisms, 100%; sensitivity in 200 mg of feces: 10 organisms, 90%; 100 organisms, 100%

Single-tube nested PCR

Gookin et al. (2002)

ITS1/5.8S rDNA/ITS2

Real-time

TFR3, TFR 4

TFR3, TFR4, primer sequences published (Felleisen et al. 1998)

NA

NA

T. foetus-specific

 

SYBR® qPCR

Mueller et al. (2015)

5.8S rDNA

Real-time

T.foeForward (TFF2), T.foeReverse (TFR2)

GCG GCT GGA TTA GCT TTC TTT, GGC GCG CAA TGT GCA T

T.foeProbe (5’FAM/3’MGB-NFQ)

ACA AGT TCG ATC TTT G

Amplifies T. foetus, T. suis, T. mobilensis

3 fg DNA, 0.1–1 cells per assay

5’ Taq nuclease assay using a 3’ minor groove binder-DNA probe; no need for post-amplification processing

McMillen and Lew (2006)

SSU rDNA

End-point nested

External: 16Sl, 16Sr; nternal: TN3, TN4

External published by Cepicka et al. (2005): TAC TTG GTT GAT CCT GCC, TCACCTACCGTTACCTTG; internal: ATA GGA CTG CAA AGC CGA GA,

TGA TTT CAC CGA GTC ATC CA

NA

NA

Amplifies Trichomonas sp.

NA

NA

Robinson et al. (2010)

SSU rDNA

End-point

Tgf, Tgr

GCA ATT GTT TCT CCA GAA GTG, GAT GGC TCT CTT TGA GCT TG

NA

NA

Amplifies T. gallinarum

One protozoon per assay

Cross-reactions with T. gallinae. No cross-reactions were also observed with samples from other protozoa (Toxoplasma gondii, Eimeria tenella, Cryptosporidium spp., E. invadens, and E. ranarum)

Grabensteiner and Hess (2006)

Not reported

End-point + southernblot by probe

TF1, TF2

CAT TAT CCC AAA TGG TAT AAC, GTC ATT AAG TAC ATA AAT TC

Probe for Southern blot

CAT CAT TAA TGC CTT TTG ATG GAT CAG GCA ACC ATT TAT A

Amplifies T. foetus

Ten or occasionally fewer protozoa

Southern blot necessary to identify specific band. A 400 bp product from bovine genomic DNA is amplified. Multiple amplification products from DNA from a related organism, T. vaginalis; Southern blot is negative for T. vaginalis

Ho et al. (1994)

NA not applicable

It has been shown that also in PCR diagnosis the likelihood to detect T. foetus decreases with time between sampling and analysis. In one study it was reported that diagnostic sensitivity in PCR detection declined from 90% when samples were stored for 6 h to 31% when they were stored for 5 days (Mukhufhi et al. 2003). It has been hypothesized that hydrolases secreted by trichomonads are responsible for this effect (Thomford et al. 1996; Sager et al. 2007). Adding the DNA-stabilizing agent guanidinium thiocyanate—GuSCN in a final concentration of 200 mmol/L—or a commercial lysis buffer for sample collection known to preserve DNA for several months at room temperature to the transport medium did not improve results (Mukhufhi et al. 2003; Mendoza-Ibarra et al. 2012). Fecal samples from cats should be submitted within 24 h after sampling at room temperature or 4 °C.

To prevent amplification of carry-over contaminants, protocols that incorporate dUTP instead of dTTP as a nucleotide and allow to apply the Uracil-DNA Glycosylase (UDG) system have been reported (Longo et al. 1990; Felleisen et al. 1998). Contaminating amplicons carried over from previous PCRs can be removed by this system before a PCR amplification. Possible disadvantages of the UDG system are that it might become necessary to optimize the reaction mixture used for PCR—e.g., the MgCl2 concentration—and a standard PCR buffer may not work (Felleisen et al. 1998).

It is essential to monitor the presence of a potential inhibitor in each individual sample since preputial material, cervico-vaginal mucus, and fecal samples contain PCR-inhibiting components. There are different possibilities available. The TFR3/TFR4 PCR protocol includes an artificial internal control DNA carrying TFR3 and TFR4 sequences—based on pBluescript KS+ DNA—and generates control amplicons, via PCR amplification and composite primers (Table 14.1) (Felleisen et al. 1998; Sager et al. 2007). Internal controls of unrelated DNAs have been integrated into a 5′ nuclease assay—real-time PCR assays with TaqMan probes. In this type of assay it is possible to incorporate an unrelated DNA into the sample prior to DNA purification, which is later amplified in a multiplex assay along with the parasite DNA but detected by a probe carrying a fluorophore different than that of the parasite-specific probe. One commercially available T. foetus real-time PCR includes such a control system (VetMAX™-Gold Trich Detection Kit, Life Technologies). This principle of inhibition control is becoming more and more popular, also for in-house assay.

There are many ways, by which DNA from preputial smegma, cervico-vaginal mucus, fecal samples, oropharyngeal swabs, culture material, and others types of samples can be extracted. Often commercial kits but also in-house methods have been applied. Also non-purified but heat-treated samples were used with success (McMillen and Lew 2006). However, the use of unpurified DNA is prone to inhibition and is not generally recommended. It has been shown that inhibiting components could be successfully removed from preputial smegma by 5% Chelex®-100 and 0.05% agar solution (Chen and Li 2001). However, in another study, the use of Chelex®-100 caused significantly lower detection rates (Mendoza-Ibarra et al. 2012).

The majority of published diagnostic PCRs for T. foetus are targeting rRNA-coding genes (rDNA) and their flanking regions (Table 14.1). These regions include the 18S rRNA gene, the internal transcribed spacer (ITS)1 region, the 5.8S-rRNA gene, the ITS2 region, and the 28S rRNA gene. One of the first diagnostic PCRs established—the TFR3/TFR4 PCR—is widely used. The TFR3 primer targets the 3’ end of the 18S rRNA gene and the TFR4 primer the 5′ end of the 28S rRNA gene (Felleisen et al. 1998). Although this PCR assay is often referred to as being specific for T. foetus, also DNA of T. mobilensis—an intestinal parasite of squirrel monkey—or T. suis is amplified (Table 14.1).

The rRNA gene sequences have been widely used for phylogenetic studies in Parabasalia to which trichomonads belong. Other genes coding for cysteine proteinases—CP1, CP2, and CP4–CP9—and cytosolic malate dehydrogenase 1 (MDH1) have been used to differentiate T. foetus isolates from cattle and cat or to characterize new strains of T. foetus (Kleina et al. 2004; Cepicka et al. 2005, 2006; Gaspar da Silva et al. 2007; Kolisko et al. 2008; Sun et al. 2012; Slapeta et al. 2012; Casteriano et al. 2016). In T. gallinae further genes were used to define lineages (recently reviewed in Amin et al. (2014)).

Because 18S rRNA genes show limited differences between trichomonads, end-point assays have been applied using primers capable to amplify DNA of several trichomonad species simultaneously (Felleisen 1997). In these PCRs, species diagnosis was achieved in a second step, either by PCR-RFLP, by determination of the precise size of amplification products, or by single-strand conformation polymorphism (SSCP) (Hayes et al. 2003; Huby-Chilton et al. 2009).

The TFR3/TFR4 PCR protocol has been modified by using the TFR3/TFR4 primer pair for external amplification followed by an internal newly designed primer pair in a single-tube nested PCR (Gookin et al. 2002). The TFR3/TFR4 PCR protocol has been also modified into a SYBR®-based real-time PCR assay (Mueller et al. 2015).

A 5’ nuclease assay—i.e., a real-time PCR applying a TaqMan probe—based on rRNA gene sequences has been established to detect T. foetus, T. suis, and T. mobilensis (McMillen and Lew 2006). In this assay a 57 bp region of the 5.8S rRNA gene region is amplified (Table 14.1). As mentioned earlier in this section, a commercialized 5′’ nuclease assay is available (VetMAX™-Gold Trich Detection Kit) which has been used in epidemiological studies on T. foetus of cattle in Southern Africa (Casteriano et al. 2016).

14.2.2.5 Serological Techniques

Serological and other antibody detection tests have been established. However, they are of no importance for the diagnosis of trichomonosis. These tests include:
  • Agglutination and hemolytic tests. In T. foetus-infected cows, antibodies appear in the cervicovaginal mucus about 6 weeks after infection and persist for several months (Pierce 1947). A mucus agglutination test detected 32% (57 of 178) of cows in naturally infected herds, and no cows from clean herds tested positive (Pierce 1949). The mucus agglutination test was shown to be specific as no cross-reactions with Campylobacter fetus or Brucella abortus has been observed. However, the reliability of the test was strongly influenced by the type of mucus. The mucus agglutination test was regarded as herd test (Pierce 1949).

    A serum agglutination tests, similar to the mucus agglutination test, did not show results that correlated well with the T. foetus infection status of cows (Kerr 1944).

    A hemolytic assay has been established which showed in female cattle a diagnostic specificity of 96% and a diagnostic sensitivity of 94% (BonDurant et al. 1996). However, only 43% of chronically infected bulls tested positive when this test was applied (BonDurant et al. 1996).

  • Indirect ELISAs to detect parasite-specific antibodies. An indirect ELISA (iELISA) based on the TF1.17 surface antigen of T. foetus showed promising results when tested with cervico-vaginal mucus of heifers (Ikeda et al. 1995). TF1.17 surface antigen-specific IgG1, IgA, and IgM antibodies in the smegma of bulls naturally infected with T. foetus—as determined and measured by ELISA—were observed concurrently with T. foetus-positive smegma cultures (Rhyan et al. 1999); to the best of our knowledge, this approach was not further elaborated. In vaccination studies, IgG1, IgG2, IgA, and IgE responses were monitored in the preputial secretions or in sera of bulls by using a whole T. foetus cell antigen preparation for ELISA (Cobo et al. 2009).

    An iELISA coated with whole T. foetus parasites and fixed with ethanol was used to determine an isotype-specific antibody response in the reproductive tract secretions and sera of T. foetus-infected heifers (Skirrow and BonDurant 1990a). In cervical and vaginal secretions, parasite-specific IgA and IgG1 antibodies predominated 7–12 weeks after infection, while in serum, parasite-specific IgG1 and IgG2 antibodies were detected. Interestingly, elevated antibody levels were observed after reinfection using this iELISA (Skirrow and BonDurant 1990a). A similar iELISA with immobilized whole T. foetus parasites was used to monitor the T. foetus-antibody response in immunized heifers naturally challenged by being bred with a naturally infected bull (Cobo et al. 2002). The presented serological tests have not been used or validated for routine diagnostic purposes.

An iELISA has been also used under experimental conditions to detect antibodies against T. gallinarum and T. gallinae in poultry (Amin et al. 2011). For this iELISA, the plates were coated with T. gallinarum parasites in carbonate buffer per well.

14.2.2.6 Intradermal Test

Analogous to the tuberculin test, a Tricin test has been developed to identify T. foetus-infected cattle or herds (Kerr 1944). The antigen used for an intradermal Tricin test has been prepared by fixation of cultured T. foetus parasites using trichloroacetic acid. Skin reactions—i.e., an increase in skin thickness—were read 30–60 min after the application of the antigen. This test was regarded to represent a herd test (Kerr 1944).

14.2.2.7 Antigen Detection

Attempts to develop a sensitive and specific diagnostic antigen test for detecting T. foetus antigen in cervico-vaginal mucus have failed (Yule et al. 1989b). However, immunohistochemical techniques have been successfully applied to demonstrate trichomonads in histological sections (Rhyan et al. 1995b; Yaeger and Gookin 2005). For immunohistochemical detection, monoclonal antibodies developed to characterize T. foetus antigens revealed to be valuable tools (Hodgson et al. 1990). A protocol for immunohistochemical detection by using the monoclonal antibody Mab 34.7C4.4 is provided in the OIE Terrestrial Manual (www.oie.int/international-standard-setting/terrestrial-manual, accessed, 22. Febr. 2017).

14.2.2.8 Diagnosis in Different Hosts

Diagnosis in Cattle

A tentative diagnosis of trichomonosis as a cause of reproductive failure in a herd is based on the clinical history, signs of early abortion, repeated returns to service, or irregular estrous cycles. Confirmation of infection depends on the demonstration of the organism in placental fluid, stomach contents of the aborted fetus, uterine washings, pyometra discharge, vaginal mucus, or preputial smegma. In infected herds, the most reliable material for diagnosis is preputial scrapings (Kittel et al. 1998; Mukhufhi et al. 2003; Parker et al. 1999; Schönmann et al. 1994).

Bulls are the main reservoir for the parasite. Control programs focus on identifying and culling infected bulls and nonpregnant cows carrying the parasite. Prevention of transmission of the disease through culling practices relies on the ability to identify infected animals accurately.

Advances in cell culture and polymerase chain reaction (PCR) have increased the ability to detect the disease in bulls. However, the collection of an adequate sample is of outmost importance, as this step affects sensitivity, specificity, and repeatability. The low repeatability observed with most sample collection techniques can cause false-negative results. The most efficient method of sampling vaginal and preputial secretions is insertion of an insemination/infusion pipette inside the vaginal fornix or preputial cavity and performing short strokes while concurrently aspirating secretions (Cobo et al. 2007). Vagina or preputial cavity can be washed with PBS to recover more organisms, although this usually dilutes the sample. Alternatively, preputial secretions can be collected by scraping with a plastic or metal brush, with no significant differences in culture sensitivity compared to using a pipette (Tedesco et al. 1979; Parker et al. 1999).

Diagnosis in Bulls
Routine herd diagnosis is optimally performed on bulls and not on females, because bulls remain permanently infected while in most female cattle infection is only transient. In addition, sampling of bulls reduces costs as preputial samples have to be taken only from a smaller number of bulls. Diagnosis in the bull involves collecting, transporting, and culturing the sample in special growth media and tentatively identifying the organism by microscopic examination. Finally a confirmation of detection using PCR has to be carried out to confirm a T. foetus infection (Fig. 14.4). If samples are processed by PCR, the use of pooled direct preputial samples is possible. However, this strategy required repeated sampling to optimize sensitivity (Garcia Guerra et al. 2013). Optimally, all bulls belonging to the same herd should be sampled. In those herds in which periodic sanitary control is carried out without previous history of venereal diseases and with good pregnancy rates, at least two consecutive samplings of all bulls should be performed, with a minimum of 1 or 2 weeks time between them (Skirrow et al. 1985; Kimsey et al. 1980). Consecutive testing is necessary due to the low diagnostic sensitivity of an individual test. If both samplings revealed negative results, the herd can be considered tritrichomonosis free.
Fig. 14.4

Collection of preputial smegma samples and diagnostic methods for bovine tritrichomonosis

Some recommendations have to be taken into account at the time of sampling:
  • It is generally recommended to allow a sexual rest of about at least 1–2 weeks before taking a preputial sample to allow a repopulation of microorganisms in the preputial cavity and increase diagnostic sensitivity.

  • It is important to have adequate facilities to perform preputial sampling particularly when sampling a large number of bulls to avoid accidents and injuries.

  • The bulls should remain in a pen, without access to water approximately 12 h prior to sampling. This will prevent urination during sampling.

  • The external preputial area must be cleaned with disposable paper towels without soap or disinfectants, and—if necessary—preputial hairs should be trimmed to prevent contamination.

The presence of T. foetus seems to be confined to the preputial cavity, and T. foetus is localized in the preputial secretions and does not invade the epithelium of the penis or prepuce (Parsonson et al. 1974). T. foetus could not be cultured from the epididymis, ampulla, seminal vesicle, pelvic urethra, or testis. Macroscopic and microscopic examination of the genital tracts of infected bulls did not reveal any lesions populated by T. foetus (Parsonson et al. 1974).

A number of techniques for collecting preputial samples from bulls have been described (INTA, 2014, Técnicas de muestreo para el diagnóstico de enfermedades venéreas en bovinos, https://www.youtube.com/watch?v=G-1StrWaHKA, accessed 21. Febr. 2017; Navajo Technical College, 2012, Veterinary Technicians perform Trich testing, https://www.youtube.com/watch?v=lp8fpDVDOCE, accessed 21. Febr. 2017). In all these protocols, it is important to avoid contaminations, as this may introduce intestinal protozoa or PCR-inhibiting contaminants. Samples can be collected from bulls by scraping the preputial and penile mucosa with an artificial insemination pipette or scraper, by preputial lavage, or by washing the artificial vagina after semen collection (Cobo et al. 2007; Tedesco et al. 1979; Parker et al. 1999). The latter technique is not recommended as its sensitivity may be lower (Ostrowsky et al. 1974; Parker et al. 1999).

The collection of the preputial smegma can be carried out by using different devices:
  • A Cassou insemination pipette (Cassou straws, IMV Technologies, L’Aigle, France) covered by a plastic sheath. Inside the preputial cavity, the pipette is pulled forward through the plastic sheath to expose the tip and moved back and forward in short strokes adjacent to the glans penis, especially near the fornix, while aspirating and massaging the glands penis to release greater amounts of smegma into the pipette. A new plastic sheath is used for each bull (Cobo et al. 2007).

  • A sterile, dry, plastic insemination/infusion pipette of 43 cm length with a 10–15 mL syringe attached to one end is placed into the preputial fornix. The pipette is scraped vigorously across the preputial epithelium without aspiration, and then negative pressure is applied with the syringe to collect preputial smegma. The negative pressure is released before removing the pipette from the sheath, to avoid aspiration of urine or other contaminants. After removing the pipette from the sheath, the sample is placed immediately into a transport or culture medium. A new syringe and pipette are used for each bull (Rodning et al. 2008).

  • A plastic or metal scraping instrument of approximately 70 cm in length, having at the anterior end a grooved surface of approximately 10 cm in length by 0.8 cm in diameter, which is used to perform scraping of the preputial mucosa. The scraper is introduced into the preputial mucosa by performing 20–30 movements back and forward. It is then carefully withdrawn from the preputial cavity, avoiding contamination with the external part of the foreskin (Terzolo et al. 1992). The aspiration method for the collection of preputial secretions for cultural examination continues to be used although a specially designed scraping instrument was reported to possess advantages (Bartlett et al. 1947; Clark et al. 1971; Stuka and Katai 1969).

  • A gauze sponge, which to our knowledge is rarely used. In this procedure, the penis is extended by electrostimulation with a rectal probe. Once extended, a 16-ply gauze sponge is used to wipe around the glans and down the penile shaft and exposed preputial mucosa two to three times. Recently, it has been demonstrated that sponge sampling of T. foetus from bulls is a valid method (Dewell et al. 2016). The method facilitates easier collection once the penis is extended and is potentially safer for the veterinarian and the bull. Additionally, the gauze sponge method might be slightly more sensitive than the pipette method (Dewell et al. 2016).

After collection the material has to be inoculated in corresponding transport or culture media and should arrive at the laboratory within 24–36 h; during transportation, the sample should be protected from exposure to daylight and extremes of temperature.

Diagnosis in Heifers and Cows

Initially, the uterus was regarded as the definitive and persistent site of infection while the vagina was considered to be a relatively unreliable source of T. foetus for diagnosis (Bartlett 1947). Subsequently, however, the persistence of T. foetus in the vagina and/or cervix for periods up to 95 days post-infection was shown, and it was established that samples collected from vagina and the uterine cervix allow a reliable and accurate diagnosis (Parsonson et al. 1976). More recent studies of naturally infected cows reconfirmed that the cervix belongs to the preferred site of parasite location (Skirrow and BonDurant 1990b). The numbers of parasites present in the cervical-vaginal mucus fluctuate during the estrus cycle, and the largest numbers are seen a few days before estrus.

The time a cow remains infected was significantly longer for cows experiencing their first, mean 20.3 weeks, than for those experiencing either their second, mean 9.8 weeks, or their third period of infection, mean 11 weeks. However, the rate of isolation of T. foetus from samples of vaginal mucus collected from cows remained similar—mean 83.5%—irrespective of the period of infection or whether the cows showed normal fertility, infertility, or abortion (Clark et al. 1986).

The efficiency of diagnosis in cows increased with temporal proximity between the initial infection and the time of sampling (Clark et al. 1986). It is important to note that the detection of positive females is of value for the initial detection of the infection in a herd. However, it is not useful for a subsequent disease control because cows usually clear their infection and generally become immune, at least for the actual breeding season (BonDurant 1997; Fitzgerald 1986). Sampling should be performed when females with conception failures are observed or at the time of rectal palpation in nonpregnant females. Culture sensitivity is lower in cows than in bulls. However, it is important to point out that the optimal period for sampling is near the end of the service period (Skirrow and BonDurant 1990b; Terzolo et al. 1992).

Uterine and vaginal secretions can be collected in cows that have aborted, in those that have not been pregnant, or heifers. To perform the extraction of cervico-vaginal mucus, a sterile, dry, plastic 43 cm insemination pipette with a 10 mL syringe attached to one end or a Cassou insemination pipette (44 cm long × 0.64 cm outer diameter × 0.32 cm inner diameter; Cassou straws, IMV Technologies, L’Aigle, France) can be used. Opening the lips of vulva without having to fix the cervix rectally, the pipette is inserted in a dorsal-cranial direction into the vagina. After the pipette has been inserted, slight anteroposterior and circular movements are executed, performing aspiration simultaneously. The vacuum generated is usually sufficient to extract the cervical-vaginal mucus, which will vary in quantity and consistency depending on the moment of the estrous cycle in which it is extracted. In a low percentage of animals, the volume of mucus extracted may be scarce. In this case it is possible to introduce 5 mL of phosphate-buffered saline solution by performing a wash with subsequent extraction.

The use of a “screwhead scraper rod” for collecting of samples from the cervico-vaginal mucosa proved to be a practical method and calls for further comparative evaluation with other standard methods in use (Abbitt and Ball 1978). Apart from other extraction techniques, also the Bartlett glass pipette procedure, which is rarely used because it is complicated and the equipment often inaccessible, has been described (Hammond and Bartlett 1943).

Fetal Diagnosis

When abortion occurs, T. fetus can be isolated from placental fluids or cotyledons. However, the high degree of contamination of this material limits its use. Isolations can be made from samples taken from the fetal mouth. Swabbing the mucosa of the tongue and roof of the mouth has been recommended (Case and Keefer 1938). Nevertheless, the place where T. foetus is most consistently isolated is the abomasal fluid. The sample can be taken with sterile syringe and needle. Once the abomasum content has been extracted, it can be sent to the laboratory in the same syringe or can be inoculated in transport or cultured medium.

Diagnosis in Cats

The diagnosis of feline trichomonosis has been reviewed comprehensively (Tolbert and Gookin 2009; Manning 2010; Yao and Köster 2015). T. foetus infection is suspected in cats with recent—less than 6 months lasting—clinical signs of chronic large bowel diarrhea, in young, purebred cats, from densely housed origin. Routine coprological methods, like flotation-sedimentation or sodium acetate-acetic acid-formalin concentration (SAF), destroy or fix trichomonads, respectively, and fixation causes the loss of their characteristic movement which makes them hard to be recognized.

Currently, the preferred diagnostic methods for feline trichomonosis include visualization of the organism in direct smears or culture or T. foetus DNA detection by PCR (Gookin et al. 2001, 2002, 2004; Levy et al. 2003; Foster et al. 2004). Histopathological examinations of colon, cecum, and ileum samples are not routinely used but can be helpful to diagnose a feline tritrichomonosis.

It is important to mention that parasite shedding in feces is erratic throughout the course of infection being occasionally so low that it cannot be detected by diagnostic techniques. Consequently, it is advisable to resample cats showing clinical signs that have been tested negative or to increase diagnostic sensitivity by using more than one test method—e.g., culture and PCR (Gookin et al. 2002). Cats should not receive any antibiotics within several days prior to or at the time of testing.

Samples consist of fresh voided feces taken directly from the litter box, rectal swabs, and manual collection with the aid of fecal loops or by a colon flush technique (revised in Manning (2010), Yao and Köster (2015), Tolbert and Gookin (2009)). Samples collected with a fecal loop or by the colon flush technique are preferable. The technique of colon flush is demonstrated in a video clip the North Carolina State University, College of Veterinary Medicine website (http://www.youtube.com/watch?v=JMfZ9M80V8E, accessed 22. Febr. 2017). Freshly voided or diarrheic feces are considered ideal for testing whereas samples obtained from normal or dry stools are believed to be less suitable (Tolbert and Gookin 2009).

A diagnostic approach based on direct fecal smears, which may reveal motile trophozoites, can be employed during examination at pet clinics. Samples are suspended in saline and examined immediately under a cover slip at 200 to 400-fold magnification using a light or, preferably, a phase-contrast microscope (Fig. 14.5). Although direct smears represent a cheap, quick, and readily available technique, the sensitivity of microscopic examination of a direct fecal smear is low. The diagnostic sensitivity of a direct smear using samples from naturally infected cats has been shown to be only 14% in one study, but sensitivity can be increased by analyzing multiple fecal smears (Gookin et al. 2004). Other challenge associated with this diagnostic procedure is the skill of the practitioner in identifying motile trichomonads. T. foetus cannot precisely be distinguished by microscopical examination from P. hominis and is often misdiagnosed as Giardia spp. (Tolbert and Gookin 2009; Manning 2010; Yao and Köster 2015). A video clip demonstrating the classic jerky motility has been provided by the North Carolina State University, College of Veterinary Medicine website (https://www.youtube.com/watch?v=aF06jlbcF8E; accessed 22. Febr. 2017). The presence of Giardia spp. can be confirmed by a fecal enzyme-linked immunosorbent assay for Giardia-specific antigen. Differentiation of Giardia spp. and T. foetus is crucial to avoid a useless initiation of tritrichomonosis therapy using potentially neurotoxic ronidazole.
Fig. 14.5

Microscopic detection of Tritrichomonas foetus trophozoites (arrows) in feline feces (200× magnification)

In vitro culture of T. foetus can be started by incubating the sample feces in a suitable growth medium (Gookin et al. 2001, 2003). Diagnostic kits—for example, the InPouch TF-Feline test kit—can be an attractive option for the clinical practice due to the commercial availability, a shelf life of a year, and the simplicity of use. Sample submission requirement for in vitro culture diagnostic includes the collection of fresh feces—roughly the size of a lima bean—and the use of transport-growth media, InPouch TF or modified Diamond’s media. It is recommended to dilute the fecal samples in PBS or physiological saline—to about 1 to 20 v/v—to create a homogenous mixture with a semiliquid consistency to be used as the inoculum for the culture medium, reducing the bacteria burden in the sample (Arranz-Solis et al. 2016). Furthermore, it is crucial for a reliable diagnosis of feline trichomonosis to keep the time delay from sample collection to processing as short as possible, given the environmental fragility of T. foetus trophozoites (Gookin et al. 2004). In order to maximize the diagnostic sensitivity, cat feces should be inoculated into the culture within a 6-h period after voiding and to be submitted at room temperature—23 to 25 °C—to a specialized diagnostic laboratory within 24 h (Hale et al. 2009). False-negative findings can be due to refrigeration or delayed processing of feces.

After culture initiation, samples are then incubated at room temperature or at 37 °C; incubation at room temperature made more robust and long-lived cultures (Gookin et al. 2003). Cultures are examined microscopically periodically up to 7 days—normally every 24–48 h—on a wet mount slide prepared directly from the culture or through the plastic wall of the InPouch TF kit. Most of the microscopically positive culture samples are detected 72 h post-incubation. It has to be mentioned that the diagnosis by culture is more difficult for cats than for bovines due to the nature of the feces sample. As feces are directly cultured in enriched media, bacterial and/or fungal contamination is more likely to arise. Bacterial contamination severely impairs the culture of T. foetus and is able to reduce the sensitivity of the culture method. Incubation at 37 °C shows a quicker positive result but also more bacterial overgrowth potentially inhibiting the growth of T. foetus. If the commercial InPouch TF-Feline test kit is used, an accumulation of gas within the pouches due to an overgrowth of fecal flora is a commonly observed problem. Moreover, T. foetus and related parasites are hard to distinguish with light microscopy because of the similar morphology. Culture-positive samples require confirmation by PCR because also other trichomonads like P. hominis are able to grow (Ceplecha et al. 2013).

Molecular diagnosis is becoming more widely available than culturing the organism. Under optimized conditions, a direct diagnosis by PCR—i.e., without a prior cell culture test—offers a highly specific and a sensitive diagnostic alternative also able to detect nonviable parasites. Moreover, PCR is the method of choice to confirm positive results by culture. It is important to monitor a potential PCR inhibition in each individual sample. The choice of an appropriate DNA extraction technique greatly influences the reliability and sensitivity of PCR (Stauffer et al. 2008; Hale et al. 2009). Several methods have been successful employed for feline trichomonosis diagnosis, such as a modified procedure using the commercial QIAamp DNA stool minikit (QIAGEN, Hilden, Germany), Boom’s method, or ZR Fecal DNA kit ZR (Zymo Research, Orange, CA, USA), which was able to detect 10 T. foetus organisms per 100 mg feces in 100% of PCR reactions (Stauffer et al. 2008). Diagnostic T. foetus PCR using primers TFR3/TFR4 and a single-tube nested PCR using primers TFITS-F/TFITS-R in combination with primers TFR3/TFR4 are widely used for feline trichomonosis diagnosis (Table 14.1) (Felleisen et al. 1998). PCR amplification of DNA extracted from feces samples appears to be more sensitive than the InPouch TF culture, and a high level of agreement has been described between the culture and PCR detection (Gookin et al. 2002; Hosein et al. 2013; Arranz-Solis et al. 2016).

Finally, colonic histopathology can be used as a diagnostic tool. Colon, cecum, and ileum samples are collected during necropsy, surgery, or endoscopy. On histologic examination, large numbers of teardrop- to crescent-shaped trichomonads can be found associated with the colonic mucosal surface and less commonly within colonic crypt lumens. Histologic features include mild to moderate lymphoplasmacytic colitis with crypt micro-abscesses, increased mitotic activity, loss of goblet cells, and attenuation of superficial colonic mucosa (Yaeger and Gookin 2005). The probability of diagnosing a T. foetus infection based on histopathology increases with the number of submitted samples—a minimum of six colon samples is required to have a larger than or equal to 95% confidence of detecting T. foetus in at least one sample (Yaeger and Gookin 2005). Recently, a fluorescence in situ hybridization assay (FISH) and a chromogenic in situ hybridization (CISH) technique have been described to allow the localization and identification of T. foetus in formalin-fixed and paraffin-embedded samples, respectively (Gookin et al. 2010b).

Diagnosis in Other Animals

The trichomonad T. gallinae of birds is of veterinary importance, while Tetratrichomonas gallinarum is a commensal that can become important under certain circumstances—e.g., concurrent infections with other pathogens (recently reviewed in Amin et al. (2014)). Diagnostic techniques described for T. foetus can be also applied to the diagnosis of bird trichomonosis, including direct microscopy, cultivation, and PCR detection.

For direct microscopy, sample material can be collected by swabbing the oral cavity, T. gallinae, or cloacae, T. gallinarum, and is mounted either diluted or undiluted on glass slides. Glass slides need to be examined immediately to observe motile protozoa (Forrester and Foster 2008; Amin et al. 2014). As mentioned, staining of trichomonads is possible and may facilitate detection (Amin et al. 2011). Similar to bovine and feline trichomonosis, the sensitivity of direct microscopy is low, and situations in which the parasite load is low may likely result in false-negative results.

The use of samples to initiate cultures allows the amplification of the number of parasites making this procedure more sensitive than the direct microscopic examination of samples. Correspondingly, in a study of pigeons, only about half of the samples positive in culture revealed a positive result in wet mount microscopy (Bunbury et al. 2005). In case of T. gallinae-infected birds, material collected via swabbing from the oral cavity of the animal is added to a InPouch TF culture device or other commercial or noncommercial culture media (Bunbury et al. 2007; Forrester and Foster 2008; Rogers et al. 2016; Girard et al. 2014; Krone et al. 2005). For cultivation, specimens are usually incubated at 37 °C for several days and examined microscopically every 24 h. Although birds have a higher body temperature, 37 °C seems to be the optimal temperature for the cultivation of T. gallinarum (De Carli and Tasca 2002; De Carli et al. 1996; Amin et al. 2010).

For PCR, a number of primer pairs have been reported that target the ITS1-5.8SrDNA-ITS2 genomic region but which are not species-specific and able to detect virtually all trichomonad species. Most commonly used are primers TFR1 and TFR2 (Felleisen 1997). A nested PCR, targeting the 18S rRNA gene, ITS1, and 5.8S rRNA gene, was established but was not species-specific, amplifying also T. gallinae (Frey et al. 2009; Grahn et al. 2005). Another nested PCR has been developed for the amplification of the ITS1-5.8S rDNA-ITS2 genomic region of trichomonads used to detect T. gallinae-specific DNA from esophageal lesions of finches sampled during an epidemic of finch mortality (Robinson et al. 2010; Cepicka et al. 2005). The analytic specificity of this PCR has not been reported. Amplification primers specifically designed for the identification of T. gallinarum yielded also a PCR product of specific DNA of T. gallinae of identical size (Grabensteiner and Hess 2006).

An indirect ELISA has only been used under experimental conditions to detect antibodies against T. gallinarum and T. gallinae in poultry (Amin et al. 2011). Because birds are often latent carriers of trichomonads, there is hope that serological analyses in bird populations might be able to identify carrier birds and may help to better understand persistence and spread of these parasites (Amin et al. 2014).

14.3 Epidemiology

14.3.1 Epidemiology of Tritrichomonosis in Cattle

Bovine tritrichomonosis is a major problem worldwide, affecting a large proportion of herds in North and South America, in parts of Europe, Africa, Asia, and Australia (de Oliveira et al. 2015; Yao 2013; Perez et al. 2006; Mendoza-Ibarra et al. 2012, 2013; Madoroba et al. 2011; Yang et al. 2012; Guven et al. 2013; McCool et al. 1988). Bovine tritrichomonosis is considered endemic in herds managed under extensive conditions and using natural service for breeding (Bondurant 2005). The economic impact of tritrichomonosis infection has to be regarded severe in particular regions of the world. The calf crop in affected beef and dairy herds can be reduced up to 50% in beef operations (Rae 1989). Economic losses are variable and depend on the percentage of bulls infected and the susceptibility of the cows in the herd. Further losses, in addition to calf crop losses, include an extended breeding season due to an increased number of repeat breeders. Due to later calving, the growing periods for calves might be shortened, and there are batches of calves of different ages with a wide variation in weaning weights. Summarizing these losses, it was estimated that tritrichomonosis in a herd caused a 35% decrease in economic return per cow in an infected herd (Rae 1989). Economic losses in a case of bovine tritrichomonosis in a large Californian dairy herd were calculated at 665 US$ per infected cow (Goodger and Skirrow 1986).

The number of tritrichomonosis reports has drastically reduced in regions or production systems in which artificial insemination is the predominant mode of breeding and in which comingling of herds is avoided—for example, in the European dairy industry. A recent survey conducted in Switzerland involving 1362 preputial samples from bulls and 60 abomasal fluid samples of aborted fetuses from beef and dairy herds revealed no T. foetus-positive finding (Bernasconi et al. 2014). However, also in areas assumed to be largely free of bovine tritrichomonosis, a reestablishment of the infection in herds, especially in beef herds farmed under extensive, pastoral systems, is possible as shown by findings from Spain (Mendoza-Ibarra et al. 2012; Mendoza-Ibarra et al. 2013). Knowledge on risk factors is important for the implementation of effective measures to control tritrichomonosis. Results from risk factor studies were used to model effects of vaccination against tritrichomonosis in beef herds. A number of potential herd-level risk factors were assessed, including “no. of cows,” ”no. of young bulls,” ”trichomonad testing yes/no,” ”no. of trichomonad tests,” ”shared grazing,” “previous diagnosis of trichomonosis,” or ”duration of breeding season” (Villarroel et al. 2004).

T. foetus isolated from cattle and cats and T. suis from pigs are genetically very similar or in case of T. foetus from cattle and T. suis even indistinguishable. Nevertheless, there is no evidence that there are links between life cycles of T. foetus in cattle and cats or between life cycles of feline or bovine T. foetus and porcine T. suis. Most likely these parasites have evolved separately, and despite their genetic similarity, T. foetus of bovine and feline origin and porcine T. suis show biological traits which differ considerably.

As stated in Sect. 14.1.2, tritrichomonosis is an almost exclusively venereal transmitted disease in cattle and affects predominantly adult animals (Sager et al. 2007; Ondrak 2016). T. foetus is transmitted during coitus, mainly from an infected bull to an uninfected dam or vice versa (Ondrak 2016). Single mating with an infected bull may result in a 95% infection rate among susceptible heifers (Parsonson et al. 1976), but in general a transmission rate of 30–70% is assumed (Bondurant 2005).

A mechanical transmission either by uninfected bulls or by contaminated equipment or cryopreserved semen seems to be possible, but the relative importance of these routes of transmission is minor (Ondrak 2016; Clark et al. 1977; Murname 1959; Goodger and Skirrow 1986; Blackshaw and Beattie 1955; Clark et al. 1971; Skirrow and BonDurant 1988).

The following risk factors for tritrichomonosis in individual animals have been identified:
  • Carrier state and age as a risk factor of infection: Infection in bulls is reported to persist for more than 3 years and may persist for life (Rhyan et al. 1999; Campero et al. 1990; Flower et al. 1983; Bondurant 2005). Several studies established that the likelihood of bulls being infected seems to increase with age (Skirrow et al. 1985; McCool et al. 1988; BonDurant et al. 1990; Rae et al. 1999, 2004; Mendoza-Ibarra et al. 2012) (Table 14.1). In one experiment using bulls—3 to 6 years of age—all bulls more than 4 years old became infected after three to six services while only one of two young bulls, 2–3 years of age, became infected after nine services (Clark et al. 1974). Bulls less than 4 years of age are rarely carriers of T. foetus (BonDurant 1985; Perez et al. 1992; Ondrak 2016; Kimsey et al. 1980; Skirrow et al. 1985; BonDurant et al. 1990). The reason for this finding is not completely understood. Old bulls may have had a higher number of sexual contacts than young bulls. An often mentioned other possible reason is the more pronounced invaginations in the penile and preputial epithelium of older bulls—i.e., the crypts of these epithelia are becoming deeper and increase in number with the age of the bull (BonDurant 1985; Skirrow et al. 1985; McCool et al. 1988; Perez et al. 1992; Ondrak 2016). However, the hypothesis that anatomical changes are the cause for older bulls found to be infected more often was recently questioned because no age-related statistically significant differences were observed in the surface architecture of the penile and preputial epithelium of bulls (Ondrak 2016; Strickland et al. 2014).

In female cattle, age seems not to be associated with the likelihood of infection; however, there is evidence that repeated exposure induces resistance to infection (Simmons and Laws 1957; Clark et al. 1986; Skirrow and BonDurant 1990b). While mature bulls seem to remain infected for life, most cows are able to clear the infection after a few months—usually after 1 to 3 months—rarely longer (Parsonson et al. 1974, 1976; Skirrow and BonDurant 1990b; Bondurant 2005). Several studies of infected cows indicate that the os cervix is the preferred site of multiplication and persistence (Skirrow and BonDurant 1990b). Initial multiplication of T. foetus after infection seems to be followed by a decline of parasite numbers until next estrus (Bartlett and Hammond 1945). The numbers of parasites present in the cervicovaginal mucus seem to fluctuate during the estrus cycle, and the largest numbers are seen a few days before estrus (Hammond and Bartlett 1945). In two infected heifers that have been followed over time after experimental infection, T. foetus was not always observed in vaginal mucus by microscopic examinations or culture isolation suggesting fluctuations in vaginal parasite concentration; however, only in one heifer, there was a coincidence between the detection of T. foetus and the time of estrus (Simmons and Laws 1957). Not all cows are able to clear infection. Carrier dams have been reported—e.g., two chronically infected dams were observed in one Australian study 16 and 22 months after initial infection (Alexander 1953). In a Californian dairy herd, infected cows were found positive 9 weeks after pregnancy or 63 days after parturition (Skirrow 1987; Goodger and Skirrow 1986). In a more recent study from Argentina, several heifers remained infected up to 300 days after breeding which underlines the importance of these carrier state heifers for persistence of infection in affected herds (Mancebo et al. 1995).
  • Herd management practices: The risk of bulls of being tested T. foetus positive increased when the number of bulls used per unit was higher than 10 or the bull-to-cow ratio per unit was lower than 1 to 25. The higher number of bulls and lower bull-to-cow ratios are typical management practices in large herds to increase conception rates (Rae et al. 2004). It was hypothesized that by these practices the number of potential sexual contacts per bull and the probability for an individual bull to become positive are increased (Rae et al. 2004).

  • Breed disposition: The possibility of a breed predisposition was discussed based on study results suggesting a higher prevalence of infection in particular breeds (BonDurant et al. 1990; Perez et al. 1992; Rae et al. 1999, 2004; Skirrow et al. 1985; Abbitt and Meyerholz 1979). A number of epidemiological studies addressed this question (Table 14.2). It had been hypothesized that in B. taurus indicus bulls, due to their longer preputial length, the likelihood of being infected might be higher (BonDurant et al. 1990). Statistically significant differences in prevalence were observed when B. taurus taurus and B. taurus indicus or B. taurus taurus/B. taurus indicus crosses were compared; highest prevalences of infection were observed in B. taurus taurus (BonDurant et al. 1990). Another study in Costa Rica also found a strong association between the risk of positive findings and the Bos taurus taurus breed—as compared to Bos taurus indicus pure or hybrid breeds (Perez et al. 1992); an additional study also showed an increased risk in Angus, Charolais, Hereford, or Simmental breeds relative to B. taurus indicus (Rae et al. 2004) (Table 14.2). However, findings suggesting breed disposition should be interpreted with care; studies might have been biased by uneven study designs or by not paying enough attention to the differences in the way herds of particular breeds were operated (Ondrak 2016). It was hypothesized that prevalence differences could be due to the increased number of matings accomplished by B. taurus taurus bulls as compared to B. taurus indicus bulls in the same period of time, thus increasing the risk of exposure to infection (Perez et al. 1992). However, there are also some studies that did not observe differences in prevalence of infection between B. taurus taurus and B. taurus indicus (Dennett et al. 1974).

Table 14.2

Risk factor studies in bovine tritrichomonosis

Country

Region

Type of herd or animal

No. of herds examined/no. of herds positive (%)

No. of animals examined/no. of animals positive (%)

Diagnosis

Herd-level risk factors

Individual-level risk factors

Type of study

Reference

Argentina

Province Buenos Aires

Beef

42/173 (24)

NA

C (modified Plastridge medium, Tricoazul), all bulls per herd, three sequential tests

Rearing herds vs. full-cycle herds, p = 0.052; pregnancy rate in cows ≤ 90%, p = 0.005a; shared livestock with others, p = 0.003a; rotation of bulls, p<0.05; abortion, p<0.05; T. foetus reported in previous year, p = 0.001a

NA

Case-control

Mardones et al. (2008)

Argentina

Province La Pampa

Beef

194/3766 (5.15)

309/29178 (1.06)

C with modified Diamond’s medium (MDM), all non-virgin bulls, twice a year

Seasonal effect: highest no. of positive findings in February (pre-breeding season), no statistics provided; spatial clustering in south of La Pampa, p = 0.008; coinfection, high-risk cluster cells for bovine genital campylobacteriosis were also high-risk cells for bovine tritrichomonosis, p = 0.0014

NA

Cross-sectional

Molina et al. (2013)

Australia

Victoria River District

NA

27/41 (65.5)

81/1008 (8.0)

C (modified Plastridge medium)

NA

Tritrichomonosis infection rates varied significantly with age (p<0.0001), that is, increasing with age (p<0.05); no evidence of an increased likelihood of coinfection with Campylobacter fetus

Cross-sectional

McCool et al. (1988)

Costa Rica

NA

Dairy, mainly

10/63 (15.9)

14/225 (6.2)

C (InPouch TF)

NA

Bull in service: no vs. yes, p = 0.02a; age > 3 years, p = 0.02a; breed, B. taurus taurus vs. B. taurus indicus, p = 0.02a

Cross -sectional (results of pilot study not included)

Perez et al. (1992)

Spain

Asturias de la Montana

Beef

27/65 (41.5)

33/103 (32)

C (InPouch TF or thyoglycollate transport medium (TFTM) with modified Diamond medium (MDM))+PCR (Felleisen et al. 1998)

Increased number in repeat breeder cows, p = 0.007

Age >3 years, p = 0.04

Cross-sectional

Mendoza-Ibarra et al. (2012)

Spain

Asturiana de los Valles

Beef

12/229 (5.2)

13/327 (4.0)

C (InPouch TF)+PCR (Felleisen et al. 1998)

No statistical significant predictors

Age >3 years, p<0.001 (univariable)

Cross-sectional

Mendoza-Ibarra et al. (2013)

USA

Florida

Beef

17 (40.4)

119/1984 (6)

C

Herd size ≥500, p = 0.004b; bull-to-cow ratio = 1:<25, p = 0.039b

Age ≥5 years, p = 0.022a; breed, Angus, Charolais, Hereford, Simmental vs. B. taurus indicus; p<0.031a; herd management, number of bulls per group ≥10, p = 0.002a, bull-to-cow ratio <1:25, p = 0.03a; no knowledge of farmer on tritrichomonosis, p = 0.003; geographical area, South Florida vs. North Florida, p = 0.001a

Cross-sectional

Rae et al. (2004)

USA

Idaho

Beef

65/159 (40.9)

NA

dM or C (InPouch TF, modified Diamond medium (MDM))

Total cattle grazed on FS (US Forest Service) allotment >844, p<0.05; commingling on BLM (Bureau of Lands Management) allotment, FS (US Forest Service) allotment, or on any public land allotment, p<0.05

NA

Case-control

Gay et al. (1996)

USA

Wyoming

Beef

8/303+8 (2.6)

NA

C or PCR (three consecutive cell cultures or one PCR by an accredited diagnostic laboratory); herd status based on findings in the past 3 years

Allotments neighboring a positive herd(s), p = 0.0003; allotment type, open/public vs. private, p = 0.003; mingling with neighboring herd(s), p = 0.026

NA

Cross-sectional

Jin et al. (2014)

USA

California

Beef

9/57 (15.8)

30/729 (4.1)

C, modified Diamond medium (MDM)

NA

Age >3 years, p<0.025; breed (B. taurus taurus vs. B. taurus indicus (Bi) or Bi-hybrids), p<0.001)

Cross-sectional

BonDurant et al. (1990)

USA

Texas

Mainly non-virgin bulls, tested prior to interstate or intrastate commerce

NA

NA/NA; 1154 positive results/31202 tests (3.7)

C + real-time PCR

Spatial cluster in southeastern Texas identified (p<0.001)

Proportion of positive findings was highest in August (5.5%), no statistics provided

Cross-sectional

Szonyi et al. (2012)

NA not applicable, C culture

aStatistically significant in multivariate logistic regression

Nevertheless, a breed disposition and genetically based predisposition should not be completely ruled out. Further studies are needed to elucidate the reason why some of the infected dams are becoming carrier of T. foetus—being still infected after pregnancy—while others immediately eliminate infection during the first 2 to 3 months after loss of the conceptus (Alexander 1953; Goodger and Skirrow 1986; Skirrow 1987; Mancebo et al. 1995). It is possible that like in other diseases there are genetic determinants influencing a predisposition to acquire infection and to develop immunity against the pathogen.
  • Other individual-level risk factors: One study observed that the probability of positive findings is lower in bulls in service—i.e., in sexually active bulls (Perez et al. 1992) (Table 14.2). This is in accord with previous findings, which suggested that a depletion of the preputial T. foetus population might occur because of intense sexual activity. It also supports recommendations of a sexual rest of at least 1 to 2 weeks before sampling bulls in order to improve the likelihood of accurately identifying T. foetus-positive bulls (Clark et al. 1983a; BonDurant 1985; Yule et al. 1989a; Ondrak 2016). Variations in sexual activity between different seasons may also explain fluctuating differences in the proportion of positive findings in bulls during a year (Molina et al. 2013; Szonyi et al. 2012).

In addition, there are a number of herd-level risk factor studies—case-control and cross-sectional studies—that have been carried out to elucidate in more detail management practices and other factors increasing the risk of herds to acquire tritrichomonosis (Table 14.2) (Mardones et al. 2008; Molina et al. 2013; McCool et al. 1988; Perez et al. 1992; Mendoza-Ibarra et al. 2012, 2013; Rae et al. 2004; Gay et al. 1996; Jin et al. 2014; BonDurant et al. 1990; Szonyi et al. 2012). Many of the identified explanatory variables are related to predictors or risk factors in favor to increase the likelihood of venereal transmission of T. foetus in herds and between herds:
  • Transmission between herds: Risk factors were identified which characterized the likelihood to acquire tritrichomonosis from other herds. “Allotments neighboring a positive herd(s)”; “allotment type, open/public vs. private”; and “mingling with neighboring herd(s)” were recently identified as risk factors in T. foetus-positive beef herds in Wyoming, USA (Jin et al. 2014). These findings confirm previous observations in beef herds from Idaho, which identified “commingling on BLM—Bureau of Lands Management—allotment; FS, US Forest Service, allotment; or on any public land allotment” as an important predictor of positive herds (Gay et al. 1996). Also in Argentina “shared livestock with others” was identified as a significant risk factor for herds testing positive, that is, having positive bulls (Mardones et al. 2008). In summary, these findings suggest that any type of mingling herds and also fence-line contact with other herds represent important risk factors and should be avoided by farmers (Jin et al. 2014).

  • Transmission within herds: Other predictors characterized the likelihood of transmission within a herd. Obviously, herd size plays a role. A study on beef herds in Idaho, USA, identified “total cattle grazed on FS—US Forest Service—allotment more than 844” as a risk factor, and a study from Florida, USA, observed that herds with more than or equal to 500 cows had a higher risk of being positive (Gay et al. 1996; Rae et al. 2004). The latter study also identified particular herd management practices increasing the individual risk of bulls to test positive; therefore, most likely, herd size is a confounder explained by management practices in large herds in favor for T. foetus transmission, like “no. of bulls used per unit larger than or equal to 10” or “bull-to-cow ratio per unit smaller than 1 to 25” as discussed earlier (Rae et al. 2004). The management practice “rotation of bulls within a herd” has been identified in an Argentinian study to most likely favor to perpetuate transmission of infection within a herd (Mardones et al. 2008). The same may apply to an observation that among “rearing beef herds”—i.e., herds that rear cattle until the weight of 150–250 kg—the prevalence of positive herds was higher as compared to “full-cycle herds,” herds with breeding, rearing, and fattening. This observation was explained by hypothesizing a higher proportion of reproductively active animals resulting in an increased spreading of disease in “rearing beef herds” compared to “full-cycle herds” (Mardones et al. 2008). The risk factors related to a potential perpetuation/acceleration of transmission inside herds cannot be easily used to give recommendations with respect to better management practices, because these risk factors are either unspecific or difficult to change.

Herd-level predictors, possibly related to effects of tritrichomonosis include infertility, early fetal death, and rarely, abortion. In epidemiological studies “pregnancy rate in cows higher than or equal to 90%” and reporting “abortion” were found associated with T. foetus-positive beef herds in Argentina (Mardones et al. 2008). An “increased number in repeat breeder cows” was identified as a predictor for positive herds in extensively managed beef cattle in Spain (Mendoza-Ibarra et al. 2012).

Additionally, in a few studies, spatial clustering of T. foetus-positive herds was observed, and in one of these studies, it was also observed that high-risk clusters for T. foetus correlated to high-risk clusters for bovine genital campylobacteriosis (Molina et al. 2013; Szonyi et al. 2012). The observed spatial clustering remained unexplained in these studies.

14.3.2 Epidemiology of Tritrichomonosis in Cats

Although the typical clinical sign in natural infection is chronic or intermittent diarrhea, for many cats, no diarrhea was reported in 6 months preceding diagnosis (Xenoulis et al. 2013; Kuehner et al. 2011). Chronic T. foetus-associated diarrhea in most cats is likely to resolve spontaneously within 2 years of onset, and chronic infection with T. foetus—without clinical signs—after resolution of diarrhea appears to be common (Foster et al. 2004). In experimentally infected kittens, T. foetus infections was long lasting and that, in later phases of the infection, the presence of T. foetus in feces was not always associated with clinical signs—such as abnormal consistency of feces (Gookin et al. 2001). Thus, in addition to disease-infected cats, cats with asymptomatic infection are able to transmit infection to other cats.

As mentioned in Sect. 14.1.1, feline T. foetus tolerates a broader pH range than T. foetus from cattle (Morin-Adeline et al. 2015a). Like for T. foetus from cattle, no encysted stages are known for T. foetus from cats. However, the formation of pseudocysts, which may support survival in the environment, has been reported for T. foetus (Pereira-Neves and Benchimol 2009). In addition, it has been shown that feline T. foetus is able to survive outside of its host for at least 30 min on dry cat food and 180 min in drinking water or urine (Rosypal et al. 2012). Other studies showed that T. foetus can survive in cat feces for several hours or even days or up to at least 5 days in wet cat food (Hale et al. 2009; Van der Saag et al. 2011). Even the survival of a passage through the alimentary tract of slugs (Limax maximus, Limacus flavus) was demonstrated, suggesting that slugs could transmit T. foetus over short distance (Van der Saag et al. 2011). The predominant mode of infection is the fecal-oral route, and most likely a close contact of cats favors the spread of transmission.

Several epidemiological studies are reported, providing data on putative risk factors for T. foetus infection in individual cats or catteries, cat shelters, and breeding centers (Table 14.3). Most of the studies were small-scale studies, including only low numbers of individual cats and catteries, shelters, or breeding centers. Some of the studies were restricted to cats from shows or pedigree cats. Some studies were restricted to diarrheic cats or cats with a history of chronic diarrhea. Therefore, studies listed in Table 14.3 were stratified into (1) studies including only healthy or healthy and diseased cats and (2) studies including almost only diseased cats—i.e., cats with diarrhea or a history of diarrhea. In addition to studies listed in Table 14.3, a large number of case reports and small-scale studies are available—recently reviewed by Yao and Köster (2015)—which are not mentioned in the following because these studies provided no statistical evidence on putative risk factors. As indicated in Table 14.3, some studies were included after performing own statistical analyses based on data extracted from the reports.
Table 14.3

Risk factor studies in feline tritrichomonosis

Sampled animals

Country

Region

Type of animal

No. of catteries, shelters, or breeding centers examined/ no. of catteries positive (%)

No. of animals examined/no. of animals positive (%)

Diagnosis

Cattery-level risk factors

Individual cat-level risk factors

Type of study

Remarks

Reference

Only healthy or healthy and diseased cats

           
 

Canada

Ontario

Cats sampled at a cat clinic (n=140), cat shows (n=55), and a humane society (n=46)

NA

14/241 (5.8)

C (InPouch TF), PCR for confirmation of culture positives

 

Attendance to cat shows, p<0.05 (23.6% from the show cats were positive); history of another cat in the house with diarrhea in the past 6 months, p<0.01; fed a raw food diet, p<0.01; >5 cats per house, p<0.01; purebred vs. mixed breed, p<0.01

Cross-sectional

No significant association between the presence of T. foetus and diarrhea at the time of sampling or having a history of diarrhea in the past 6 months; “attendance to cat shows” was the only significant variable in bivariable models

Hosein et al. (2013)

 

France

NA

140 cats cattery-housed pedigree cats, participating in international cat shows

18/117 (15.9)

20/140 (14.3)

C (InPouch TF) + Tricho-F/Tricho-R-PCR (TRICHO-F and antisense primer TRICHO-R targeting the ITS1-5.8S rDNA-ITS2 region as previously described (Jongwutiwes et al. 2000); (Duboucher et al. 2006) + Seq (amplicon, cloned)

 

Age < 1 year, p = 0.057

Cross-sectional

No significant associations: size of cattery, type of food, vicinity of dogs

Profizi et al. (2013)

 

Germany

NA

230 purebred cats

23/124 (15.7)

36/230 (18.5); C-positive: 29/230, PCR-positive: 28/230

C + PCR (Grahn et al. 2005) + Seq (Representative amplicons)

 

Abnormal fecal consistency, p<0.001; history of diarrhea in the past 6 months, p = 0.027; age ≤1 year old, p<0.034; Norwegian Forest (NFO) cats had a significantly higher prevalence of T. foetus infection than other breeds, p< 0.001; history of diarrhea in cattery in the past 6 months (p = 0.01)

Cross-sectional

Any

Kuehner et al. (2011)

 

Japan

Hokkaido, Saitama

147 samples, also from cats with chronic diarrhea

NA

13/147 (8.8)

Cultivation (98 samples) + PCR (all samples)

NA

Presence of chronic diarrhea, p<0.0035

Cross-sectional

No significant differences with respect to age, breed, and whether cats were maintained indoors/outdoors between infected and uninfected cats (Fisher exact test)

Doi et al. (2012)

 

Norway

NA

52 clinically healthy cats participating in 3 cat shows

NA

11/21 (21)

Direct microscopy (n=39) + C (n=39) + nPCR (n=52), Seq of TFR3/TFR4 amplicons

NA

Own previous history of diarrhea, p = 0.1 (indicated as trend)

Cross-sectional, pilot study

Four samples positive for Giardia spp., one TF-Cryptosporidium spp. coinfection

Tysnes et al. (2011)

 

USA

NA

173 cats with and without clinical signs of tritrichomonosis; 32 purebred (including 2 purebred-cross cats), 143 mixed breed cats

NA

17/173 (9.8)

C + confirmation by TFR3/TFR4-PCR + Seq (amplicons)

NA

No statistical analysis provided in reference. Retrospective statistical analysis of presented data shows that purebred cats have a higher risk than mixed bred cats, Fisher exact test, p<0.001

Cross-sectional, pilot study

All positive cats had diarrhea. Positive cats were between 6 weeks and 12 yrs old, Negative cats 4 weeks to 13 yrs old. Positive cats often (53%) had concurrent infections (Giardia spp., Cryptosporidium spp., Coccidia, FIP)

Stockdale et al. (2009)

 

USA

NA

117 cats from 89 catteries at an international cat show

28/89 (31.5)

36/117 (30.8); In detail, fecal smear (5/36), fecal culture in modified Diamond’s medium (9/36), fecal culture in InPouch TF (20/36), or nPCR (34/36)

Fecal smear examination, C in modified Diamond’s medium, C in InPouch TF medium, or nPCR amplification (Gookin et al. 2002)

Loose stools or diarrhea in any cats within the past 6 months, P<0.052; coinfection with coccidia, P<0.059; square feet of facility available per cat was low, P<0.056

NA

Cross-sectional

An association between T. foetus and Giardia spp. infection was not significant (p = 0.075). There were no differences in age or sex between uninfected cats and cats having T. foetus infection. The proximity of the cattery to within 0.5 miles of agricultural species, type of diet fed to cattery cats (commercial or home cooked), and source of water were not associated with risk of infection

Gookin et al. (2004)

 

USA

California

Diarrheic (n=219) and control cats (n=54)

NA

10/223 (4.5)

C and/or PCR

NA

Age: mean age of positive cats 0.7 ± 0.3 years, mean age of negative cats 5.4 ± 5.0 years, p<0.001

Case-control

Any

Queen et al. (2012)

Almost only diseased cats

           
 

Europe

NA

Cats submitted to a private veterinary laboratory due to diarrhea

NA

166/1840 (9.02)

TFR3/TFR4-PCR

NA

No statistical analysis provided in publication. Results of retrospective analysis: age ≤1 years (65/628 positive), 2–15 years (33/755 positive), statistically significant difference, chi-square, p<0.001

Cross-sectional

Any

Galián et al. (2011a, 2011b)

 

Germany and Austria

NA

31 cats (6 weeks–14 years old; 30 had diarrhea)

NA

6/31 (19.4%)

nPCR+Seq (Amplicon)

NA

Purebred cats, p = 0.0153

Cross-sectional, pilot study

Coinfection with Giardia (ELISA) likely in four of the T. foetus-positive cats, there was only one cat Giardia positive but TF negative

Steiner et al. (2007)

 

Spain

NA

Cats from densely housed origins with a history of chronic diarrhea: family cats (n=15), breeding center cats (n=28), cat shelter cats (n=50); healthy cats: family cats (n=20), cat shelter cats (n=22)

3/4 (75); 50 cats were from 1 breeding center and 3 shelters

36/93 (38.7) diarrheic cats from cattery or family cats; 0/20 (0) healthy cats

C in modified Diamond’s medium and/or TFR4/TFR3-PCR

NA

Age ≤1 years, p = 0.014

Cross-sectional study, cats from densely housed origins with a history of chronic diarrhea and healthy cats

No difference between purebred (Persian) and mixed breed; female vs. male, p = 0.085

Arranz-Solis et al. (2016)

 

UK

NA

111 naturally voided diarrheic feline fecal samples

NA

16/111 (14.4)

nPCR (Gookin et al. 2002)

NA

Age ≤ 1 years, p = 0.0026; pedigree cat, p = 0.018; Siamese and Bengal breed, p = 0.0077

Cross-sectional, pilot study

Any

Gunn-Moore et al. (2007)

 

UK

NA

Fecal samples from diarrheic cats submitted to a private veterinary laboratory

NA

205/1088 (18.8)

Real-time PCR (IDEXX 8-way PCR assay: 5.8S rDNA, AF339736)

 

Age: 6–12 months (29.4%) vs. >12 months (15.2%), p<0.001; pedigree cats (37.8%) vs. non-pedigree cats (6.0%), p<0.001; T. foetus-positive findings tended to be found in combinations with Coronavirus, Clostridium perfringens, and Giardia sp

Cross-sectional

<6-month-old cats had lower T. foetus prevalence than 6–12-month-old cats

Paris et al. (2014)

NA not applicable, C cultivation, nPCR nested PCR, Seq sequencing

Individual risk factors for tritrichomonosis in cats include:
  • Diarrhea, abnormal fecal consistency, and history of diarrhea. In a number of studies including only healthy or healthy and diseased cats, it was observed that positivity for T. foetus was associated with a “previous history of diarrhea,” “history of diarrhea in the past 6 months,” “presence of chronic diarrhea,” and “abnormal fecal consistency” (Table 14.3) (Kuehner et al. 2011; Tysnes et al. 2011; Doi et al. 2012). These findings confirm that T. foetus is an important cause of diarrhea in cats.

The likelihood of a cat to test T. foetus positive was also increasing if there was a “history of another cat in the house with diarrhea in the past 6 months” or a “history of diarrhea in cattery in the past 6 months,” which suggested that individual positive cats were only part of a larger problem in a cattery, cat shelter, or breeding center (Kuehner et al. 2011; Hosein et al. 2013).
  • Age. With a few exceptions, it is now accepted that T. foetus associated with large bowel diarrhea is mainly a disease of young cats. Several studies revealed that cats about 1 year old or younger are more often T. foetus positive than elder cats (Table 14.3). These observations were made in studies including only healthy or healthy and diseased cats as well as in studies including almost only diseased cats (Profizi et al. 2013; Kuehner et al. 2011; Queen et al. 2012; Galián et al. 2011a, b; Arranz-Solis et al. 2016; Gunn-Moore et al. 2007; Paris et al. 2014). These epidemiological findings confirm the results of follow-up studies of cats with T. foetus infections, which clearly showed that clinical signs spontaneously resolve within 2 years of onset and that it is difficult to detect T. foetus in cats after several weeks of infection (Gookin et al. 2001; Foster et al. 2004). Meta-analysis on available data confirmed this view (Yao and Köster 2015). The reasons why especially young cats are affected by T. foetus diarrhea are not sufficiently understood. Possible explanations are a more intense contact of kittens to their mothers or to larger numbers of other cats in breeding centers, private households, or cat shelters, which may favor transmission of T. foetus and may also influence the dose by which kittens become infected. Immunity against feline T. foetus is poorly understood; recently the possibility that parasite-specific secretory IgA mediates immunity has been discussed (Yao and Köster 2015). Histological examinations in infected cats revealed influx of lymphocytes, plasma cells, and neutrophils into the subepithelial lamina propria (Yaeger and Gookin 2005). The inflammatory response might not only be involved in pathogenesis but may also be involved in the control of infection (Tolbert and Gookin 2016).

  • Breed. A number of studies identified an increased risk of infection in purebred, or pedigree cats, or particular breeds (Hosein et al. 2013; Stockdale et al. 2009; Steiner et al. 2007; Gunn-Moore et al. 2007; Paris et al. 2014). These observations remain largely unexplained. Most likely the conditions under which purebred or pedigree cats are reared are responsible for the increased risk. Purebred cats from breeding centers represent densely housed populations, and due to the frequent, close, and direct contact, the risk of infection might be higher than in mixed breed cats reared in less dense populations (Yao and Köster 2015; Arranz-Solis et al. 2016).

  • Other risk factors. One other risk factor identified was “attendance to cat shows,” which might be a confounding predictor; the predictor could be related to the increased risk of purebred or pedigree cats (Hosein et al. 2013). There is only a single study report on food-related predictors. In a Canadian study, “fed a raw food diet” has been identified as a risk factor (Hosein et al. 2013).

  • Other concurrent infections and impaired immune system. In experimental T. foetus infection with cats which shed naturally Cryptosporidium parvum oocysts and Cryptosporidium non-infected cats those with C. parvum infection had an earlier onset, more severe diarrhea, and increased number of trichomonads on direct fecal examination, as compared to non-infected cats (Gookin et al. 2001). In epidemiological studies none of the concurrent infections examined—Giardia sp., Cryptosporidium sp., coccidia, Clostridium perfringens, feline infectious peritonitis, and Coronavirus—revealed a statistical association either to infection or disease on the individual animal level (Table 14.3). Nevertheless, it has been discussed whether T. foetus alone can cause clinical signs without an impaired or immature immune system, concurrent infection, or other factors resulting in alterations in the colonic microflora (Manning 2010).

In catteries, cat shelters, or breeding centers, T. foetus may cause outbreaks of persistent large bowel disease (Holliday et al. 2009). There is only a single study on risk factors associated with occurrence of trichomonosis in catteries (Table 14.3). Similar to the findings in individual cats, the analysis on the cattery level also identified an association between T. foetus and abnormal fecal consistency or diarrhea; “loose stools or diarrhea in any cats within the past 6 months” was significantly associated with T. foetus-positive findings (Gookin et al. 2004). A second putative risk factor identified in this study was related to the cat population density in catteries; if the “square feet of facility available per cat was low,” this was associated with positive findings (Gookin et al. 2004). The observation that “coinfection with coccidia” was associated with T. foetus-positive findings may suggest that there are common risk factors in favor of mixed infections of coccidian parasites with T. foetus (Gookin et al. 2004). In epidemiological studies none of the concurrent infections examined revealed a statistical association either to infection or disease on the individual animal level.

14.3.3 Epidemiology of T. gallinae

As outlined in Sect. 14.2.1, T. gallinae affects a large number of avian species where it mainly parasitizes the oropharyngeal membranes—sinuses, mouth, throat, and esophagus—causing a disease characterized by greenish fluid and caseous lesions, of whitish-yellowish fibrinous material, on the oropharyngeal membranes (reviewed by Amin et al. (2010)).

T. gallinae infection seems not to be host species-specific. However, studies on parasite diversity suggest that there are subtypes more commonly found in certain bird species (Gerhold et al. 2008; Grabensteiner et al. 2010; Sansano-Maestre et al. 2009). T. gallinae is most common among domestic pigeons and wild doves, and these species may represent a reservoir also for other bird species. Accordingly, rock pigeons—Columba livawere regarded as source for the worldwide distribution of T. gallinae (Stabler 1954; Harmon et al. 1987). Most pigeons harbor this protozoan but rarely show clinical disease (Stabler 1954). Hawks, falcons, and owls may become infected, most likely via predation of other infected birds (Rogers et al. 2016). In recent years, severe outbreaks of avian trichomonosis caused by T. gallinae have been recorded initially in wild finches, later in Passeriformes, canaries, and psittacines in Europe and North America since 2005 (reviewed by Amin et al. (2010)). Trichomonosis is also reported in several other bird species, including corvids in California, USA (Anderson et al. 2009). T. gallinae has a low tenacity in the environment and is regarded as unable to survive a gastric passage, and droppings of birds are regarded as free of T. gallinae (Stabler 1954). T. gallinae has no cyst stage. However, as mentioned for T. foetus (Sect. 14.1.1), the formation of pseudocysts has been reported (Tasca and De Carli 2003). The importance of pseudocysts in the transmission of T. gallinae—e.g., via contaminated drinking water—is not sufficiently understood. Survival times in tap water and in carcasses are regarded as limited; a survival time of 8–48 h in carcasses has been reported (Erwin et al. 2000). The following important facts about transmission of T. gallinae need to be taken into account:
  • Transmission by crop milk or direct contact: Prevalences in pigeons can be very high, and pigeons are regarded as endemically infected, often without clinical signs. Feeding on infectious crop milk is regarded as an important route of infection for nestlings (Stabler 1954). In other bird species, feeding nestlings by regurgitation might also be an important route of transmission. Close contact—e.g., during courtship—is also regarded as a way by which the infection is spread between adult birds (Stabler 1954). In Southeastern USA, trichomonosis was diagnosed more often in the spring and summer months than in autumn and winter months, which might be related to the times of courtship and feeding nestlings (Gerhold et al. 2007).

  • Transmission by drinking water: It is believed that infection of turkeys and chickens is caused by T. gallinae-contaminated drinking water (Stabler 1954). Drinking water is also a likely source of infection for other bird species.

  • Effect of weather events on outbreaks of avian trichomonosis: T. gallinae-associated finch mortality usually peaked in summer and autumn, but a correlation with climatic events has not been observed (Neimanis et al. 2010; Robinson et al. 2010; Lawson et al. 2011). A coincidence of the emergence of avian trichomonosis with high temperature and low rainfall has been reported (Simpson and Molenaar 2006; Bunbury et al. 2007). It has been hypothesized that as a consequence of the dry and hot weather, numbers and volumes of water sources decline, which may favor, due to higher densities of birds aggregating at limited water sources, the transmission of T. gallinae (Bunbury et al. 2007).

  • Transmission by predation: Pigeons and doves may also serve as a source of infection for raptors (Boal et al. 1998; Krone et al. 2005). Although T. gallinae is a labile protozoan which does not remain viable for a prolonged period and is rapidly killed by desiccation, survival time seems to be long enough to be transmitted also to the nestlings of raptors (Krone et al. 2005).

14.3.4 Epidemiology of T. gallinarum

In contrast to T. gallinae, T. gallinarum is mainly found as a commensal in the intestine of many domestic bird species, including chicken, turkey, guinea fowl, duck, and goose (Friedhoff 1982; BonDurant and Honigberg 1994). As a parasite of the intestine, T. gallinarum is transmitted via consumption of contaminated food or drinking water. Cloacal as well as oral experimental infection of chickens and turkeys is possible.

14.4 Prevention

14.4.1 Prevention in Cattle

Due to the widespread use of artificial insemination, bovine tritrichomonosis has almost disappeared in dairy cattle industries, like in Western European countries, in the USA, or in Canada. However, the disease is still present in many areas of the world were cattle are raised on pastures and natural mating is used. Prevention and control measures are based on the distinctive epidemiologic features of bovine tritrichomonosis, a sexually transmitted disease where infected bulls are asymptomatic carriers and represent a permanent source of infection while in heifers and cows infection is typically transient (Clark et al. 1971, 1974; Parsonson et al. 1974; Skirrow and BonDurant 1990b).

Bovine tritrichomonosis belongs to the OIE-listed diseases (http://www.oie.int/en/animal-health-in-the-world/oie-listed-diseases-2017; accessed 22. Febr. 2017). T. foetus as a causative agent of this venereal transmitted disease may be present in semen if the semen has been contaminated with preputial fluid during manual collection (Bondurant 2005). Recommendations for the importation of cattle and bulls for breeding can be found in the Terrestrial Animal Health Code. In particular, emphasis has been placed in the measures applied to bull semen donor health status in order to avoid dissemination and transmission of the disease (http://www.oie.int/index.php?id=169&L=0&htmfile=chapitre_trichomonosis.htm; accessed 22. Febr. 2017).

In dairy herds and in some beef herds, artificial insemination is a very useful measure to reduce and eliminate infection. For bulls destined for artificial insemination, quarantine and periodic testing are required. Also, in semen trade, it is of great value to know the country of origin, the reproductive history of the bull, and the tests performed by the artificial insemination center (Eaglesome and Garcia 1997). In some areas of the world, as in the EU, specific regulations are applied to bovine semen trade and to regulate the sanitary conditions of the collection center and the animals. Specifically, bulls selected for entry into artificial insemination should be tested in quarantine before admission to the center, and regular testing of animals in service is included as basic measures to avoid the presence and dissemination of bovine tritrichomonosis (European Directive 88/407/EEC of 14th June 1988 and European Directive 2003/43/EC of 26th May 2003).

In other countries, different policies have been established to control the infection. In the USA, state regulations have endeavored to control the endemic disease as to minimize economic losses by testing bulls. Only the importation of T. foetus-free bulls is permitted for reproductive purposes while positive bulls are culled (Yao 2013). In the province of La Pampa, Argentina, with a bovine census close to four million heads, more than 80% of bulls are tested twice each year, and positive bulls were culled. By this measure, a significant decrease in the number of infected herds and animals has been achieved in this region in the last 10 years (Fort et al. 2016). Recently, an online tool has been developed in the USA—Trich CONSULT; www.trichconsult.org, accessed 22. Febr. 2017—that uses a series of questions to assess the T. foetus status and management practices of a herd. Based on the responses to the questions, this page provides feedback to users allowing them to evaluate the importance of implementing suggested control strategies (Ondrak 2016).

At present, a consensus exists concerning the most relevant measures as to how to prevent and control bovine tritrichomonosis (reviewed in Ball et al. (1987), McCool et al. (1988), Bondurant (2005), Rae and Crews (2006), Campero and Gottstein (2007), Yao (2013), Ondrak (2016)).

In herds where it is not possible to introduce artificial insemination and where natural mating is the normal practice, as is the case in many regions where extensive beef cattle are raised, the following measures to prevent the entrance of the infection are recommended.
  • Quarantine and testing of replacement bulls. Replacement should be done by virgin animals or bulls acquired only from disease-free herds with records of excellence in reproductive performance. All the bulls must be tested during quarantine before entrance in the herd. If the bull comes from a tritrichomonosis-free herd, the analysis of two samples of preputial smegma with 1–2 weeks of interval during the quarantine is recommended. Three or more samples must be analyzed if bulls are provided from an area where tritrichomonosis is known to be endemic (Campero and Gottstein 2007; Yao 2013; Ondrak 2016). The measure includes the prohibition of the use of communal, shared, or rented bulls, unless their herd of origin and individual health status are known.

  • Avoid communal pastures and keep fences in good conditions. These measures will help to control some of the two most important risk factors influencing disease transmission (Gay et al. 1996; Mardones et al. 2008; Jin et al. 2014).

  • No introduction of cows or heifers of unknown health status during the breeding season. Similar to bulls, heifers and cows must be acquired only from disease-free herds with records of excellence in reproductive performance. In Argentina several heifers were found infected up to 300 days after breeding; such heifers may represent carrier cows able to introduce infection into naïve herds (Mancebo et al. 1995).

In addition to the preventive measures outlined in Sect. 14.3, the following control measures are recommended to reduce the impact and eliminate the disease in case that bovine tritrichomonosis has been diagnosed in a herd or the herd is located in a tritrichomonosis endemic area:
  • Testing of bulls before the breeding season and culling of infected bulls. Efforts to control the disease focus on using diagnostic tests with a high sensitivity, low cost, and time efficacy. It is recommended to sample the animal twice or three times before the breeding season and every time new bulls are introduced into herds (Bondurant 2005; Campero and Gottstein 2007; Yao 2013). Once new cases are not detected, annual testing is recommended to verify the non-infected status of the herd. However, testing and culling policies alone, although effective in improving reproductive efficiency, do not allow the elimination of the disease since other putative risk factors associated with the disease are usually present in the management of beef herds (Yao 2013; Collantes-Fernandez et al. 2014).

  • Average age reduction of bulls and replacement of all bulls after four breeding seasons. Replacement with negative-tested young bulls reduces the prevalence since 3-year-old bulls seem not to be as susceptible as older bulls in natural service (Clark et al. 1971; Christensen et al. 1977). A strong association of infection rate with age has been reported in several studies (Rae et al. 1999; Mendoza-Ibarra et al. 2012).

  • Pregnancy examination. It is mandatory to know the reproductive performance and the magnitude of the infertility problem in the herd. All open and aborted cows should be culled or segregated in high-risk sub-herds or groups (Campero and Gottstein 2007).

  • Use of commercial vaccines. In the presence of a high prevalence of infection in an area, vaccination of all heifers and cows is a good measure to improve reproductive efficiency especially when risk factors associated with infection cannot be avoided—e.g., the use of communal pastures. Commercially available vaccines in the Americas offer an improvement in reproductive efficacy (Kvasnicka et al. 1989, 1992).

  • Segregation of the herd in low- and high-risk sub-herds or groups. In the low-risk sub-herds, only dams that have recently calved are pregnant for more than 5 months, and virgin females must be included. These must be serviced preferably by virgin bulls or by bulls with two negative examinations and coming from negative herds. In order to follow the effect of the infection, the same group of females should be serviced with the same male until the disease is controlled (Campero and Gottstein 2007).

  • Limiting of breeding season. The breeding season should be restricted to less than 4 months to reduce the duration of the possible transmission period as much as possible. In addition, with a shortened breeding season, it becomes easier to monitor reproductive performance.

With respect to vaccination against bovine tritrichomonosis, the main objective is to eliminate a T. foetus infection from the reproductive tract before fetal loss occurs without necessarily avoiding parasite colonization of the epithelium during the first 40 days post-infection (BonDurant et al. 1993; Bondurant 2005). The mucosal immune response in the genital area seems to be the main protective mechanism which is characterized by a local response with the presence of IgA and IgG1 and a mild systemic response characterized by the presence of IgG2 and IgG1 (Skirrow and BonDurant 1990a; Anderson et al. 1996; Corbeil et al. 2008). As a rule, cows immunized with T. foetus have a humoral response pattern similar to that described for natural infections (Herr et al. 1991; BonDurant et al. 1993). However, genital IgA levels appear to depend on the type of antigen, adjuvant, and route of administration employed.

As to our knowledge, commercial vaccines against bovine tritrichomonosis exist only in the Americas but not in Europe. One available inactivated vaccine is prepared from whole organisms and can be acquired in a monovalent formulation—Trich Guard®—but also in a polyvalent formulation combined with Campylobacter foetus subspecies venerealis and Leptospira, Trich Guard V5-L®. In addition, in Argentina, an alternative inactivated vaccine—Tricovac, Tandil Biological Laboratory—containing an oily adjuvant with a concentration of 5 × 107 trophozoites of T. foetus is available.

In several studies on heifers using whole-parasite-based vaccines, a reduction in the number of infected females, a shorter time of genital infection, and a higher percentage of pregnant females in comparison with control animals have been reported (Kvasnicka et al. 1989, 1992; Herr et al. 1991; Gault et al. 1995; Anderson et al. 1996; Cobo et al. 2002, 2004). In addition, a lower number of services, 1.44 vs. 1.73 in non-vaccinated animals, p = 0.16; higher percentage of pregnant animals at the first service, 66.7 vs. 33.3% pregnancies, p < 0.05; and a reduction of embryo/fetal losses of 56.4% were observed (Kvasnicka et al. 1992; Hudson et al. 1993).

Subunit vaccines have also been developed (Clark et al. 1983b; BonDurant et al. 1993; Corbeil et al. 1998). A trial with an experimental vaccine formulated with membrane antigens of T. foetus was conducted; cows were immunized with this vaccine and subsequently challenged by the vaginal route (Campero et al. 1999). The vaccine used was able to generate a specific humoral immune response and shorten the period of infection in the vaccinated animals compared to the controls. A greater efficacy of a T. foetus membrane vaccine compared to a whole cell vaccine was observed; the animals had a shorter duration of infection, a higher pregnancy rate, and a lower rate of fetal mortality (Cobo et al. 2002). Additional work has been done to identify T. foetus surface antigens such as TF1.17 and TF190 (Voyich et al. 2001). The application of the former in experimentally challenged heifers evidenced a rapid shortening of infection and a specific IgA production in genital secretions (Anderson et al. 1996; Corbeil et al. 1998).

Vaccine-induced immunity to T. foetus has not been studied in depth in bulls and is only mentioned in some earlier studies (Clark et al. 1983b, 1984; Soto and Parma 1989; Campero et al. 1990; Herr et al. 1991). In bulls older than 5 years, the whole cell vaccine lacked a preventive or curative effect (Clark et al. 1983b). Therefore, commercial Trich Guard® vaccine is not indicated for bulls (Herr et al. 1991; BonDurant 1997).

14.4.2 Prevention in Other Animals

Prevention of feline trichomonosis is based on interrupting the fecal-oral route of transmission, particularly in catteries, shelters, and other dense cat populations. Since the viability of the parasite in the environment is limited, strict cleaning and disinfection measures are sensible measures to be implemented (Hale et al. 2009). Additionally, contamination of food and water by T. foetus is also possible; consequently, regular replacement of water and food and cleaning and disinfection of watering troughs and food bowls should be recommended. Due to the suggested role of some invertebrates to function as mechanical vectors, the avoidance of their presence in the litter box area and food and drinking areas may help to prevent infection transmission (Van der Saag et al. 2011). Finally, limiting contact between infected and non-infected cats will help to interrupt transmission of T. foetus in the population.

Measures to prevent T. gallinae outbreaks in wild as well as in captive birds are focused on actions to reduce sources of infection. One of the major aims would be to prevent attracting birds to feeding places. If feeding is necessary, such places should fulfill minimum requirements with regard to sanitary conditions, like regular changing of food and disinfection as suggested in a recent review (Amin et al. 2014). Since T. gallinae seems to require a wet environment to persist, drying of buildings and housing facilities following washing will support to control trichomonad infections. The prevention of the infection of prey birds, like pigeons nesting near urban areas, is necessary to prevent infection of predator birds. Due to the loss of habitat, predators have replaced their traditional prey mainly by urban pigeons (Boal et al. 1998; Höfle et al. 2000; Estes and Mannan 2003; Krone et al. 2005).

14.5 Treatment

14.5.1 In Cattle

In the past various imidazoles were used to treat bulls, but none was both safe and effective, and drug resistant strains were found (reviewed in Bondurant (2005), Rae and Crews (2006)). Specifically, ipronidazole is probably the most effective drug, but due to its low pH, it frequently causes sterile abscesses at the injection sites, and resistances have also been observed (Skirrow et al. 1985). A systemic treatment using drugs like metronidazole or dimetridazole produces adverse side effects and resistant populations of trichomonads (Campero et al. 1987). Currently, there is no approved treatment for cattle infected with T. foetus because of concerns regarding toxic residues in meat (BonDurant 1997).

14.5.2 In Cats

Therapies traditionally used for treatment of protozoa are not successful for feline trichomonosis (reviewed in Manning (2010), Yao and Köster (2015)). Currently, ronidazole has been the most effective drug used to date and is recommended at 20–30 mg/kg orally once daily for 14 days (Gookin et al. 2006). Relapse of diarrhea is common, but cats can continue to carry the organism, and resistant strains of T. foetus to ronidazole have also been documented (Gookin et al. 2010a). In addition, neurological toxicity in cats treated with ronidazole in the range of 30–50 mg/kg has been reported (Rosado et al. 2007). It is therefore important that owners are informed of the potential side effects. Ronidazole is not registered for veterinary use, and informed consent is necessary prior to its use in cats, and it should only be prescribed in confirmed cases.

14.5.3 In Other Animals

The drugs of choice for the treatment of avian trichomonosis are nitroimidazoles (metronidazole, dimetridazole, ronidazole, and carnidazole) (reviewed by Amin et al. (2014)). However, subtherapeutic dosing and prophylactic use of these drugs against trichomonosis have resulted in emergence of resistant strains of T. gallinae (Franssen and Lumeij 1992; Munoz et al. 1998). In wild birds, treatment is not a practical approach and generally not considered an option due to the way of application (Cole and Friend 1999). These drugs can be only used in non-food-producing birds by veterinary prescription.

References

  1. Abbitt B, Ball L. Diagnosis of Trichomoniasis in pregnant cows by culture of cervical-vaginal mucus. Theriogenology. 1978;9(3):267–70.  https://doi.org/10.1016/0093-691x(78)90034-1.CrossRefGoogle Scholar
  2. Abbitt B, Meyerholz GW. Trichomonas fetus infection of range bulls in South Florida. Vet Med Small Anim Clin. 1979;74(9):1339–42.PubMedGoogle Scholar
  3. Adl SM, Simpson AG, Farmer MA, Andersen RA, Anderson OR, Barta JR, Bowser SS, Brugerolle G, Fensome RA, Fredericq S, James TY, Karpov S, Kugrens P, Krug J, Lane CE, Lewis LA, Lodge J, Lynn DH, Mann DG, McCourt RM, Mendoza L, Moestrup O, Mozley-Standridge SE, Nerad TA, Shearer CA, Smirnov AV, Spiegel FW, Taylor MF. The new higher level classification of eukaryotes with emphasis on the taxonomy of protists. J Eukaryot Microbiol. 2005;52(5):399–451.  https://doi.org/10.1111/j.1550-7408.2005.00053.x.CrossRefPubMedPubMedCentralGoogle Scholar
  4. Alexander GI. An outbreak of bovine trichomoniasis in Queensland and its control. Aust Vet J. 1953;29:61–6.CrossRefGoogle Scholar
  5. Amin A, Neubauer C, Liebhart D, Grabensteiner E, Hess M. Axenization and optimization of in vitro growth of clonal cultures of Tetratrichomonas gallinarum and Trichomonas gallinae. Exp Parasitol. 2010;124(2):202–8.  https://doi.org/10.1016/j.exppara.2009.09.014.CrossRefPubMedGoogle Scholar
  6. Amin A, Liebhart D, Weissenbock H, Hess M. Experimental infection of turkeys and chickens with a clonal strain of Tetratrichomonas gallinarum induces a latent infection in the absence of clinical signs and lesions. J Comp Pathol. 2011;144(1):55–62.  https://doi.org/10.1016/j.jcpa.2010.06.002.CrossRefPubMedGoogle Scholar
  7. Amin A, Bilic I, Berger E, Hess M. Trichomonas gallinae, in comparison to Tetratrichomonas gallinarum, induces distinctive cytopathogenic effects in tissue cultures. Vet Parasitol. 2012a;186(3-4):196–206.  https://doi.org/10.1016/j.vetpar.2011.11.037.CrossRefPubMedGoogle Scholar
  8. Amin A, Nobauer K, Patzl M, Berger E, Hess M, Bilic I. Cysteine peptidases, secreted by Trichomonas gallinae, are involved in the cytopathogenic effects on a permanent chicken liver cell culture. PLoS One. 2012b;7(5):e37417.  https://doi.org/10.1371/journal.pone.0037417.CrossRefPubMedPubMedCentralGoogle Scholar
  9. Amin A, Bilic I, Liebhart D, Hess M. Trichomonads in birds—a review. Parasitology. 2014;141(6):733–47.  https://doi.org/10.1017/S0031182013002096.CrossRefPubMedGoogle Scholar
  10. Anderson ML, BonDurant RH, Corbeil RR, Corbeil LB. Immune and inflammatory responses to reproductive tract infection with Tritrichomonas foetus in immunized and control heifers. J Parasitol. 1996;82(4):594–600.PubMedCrossRefGoogle Scholar
  11. Anderson NL, Grahn RA, Van Hoosear K, Bondurant RH. Studies of trichomonad protozoa in free ranging songbirds: prevalence of Trichomonas gallinae in house finches (Carpodacus mexicanus) and corvids and a novel trichomonad in mockingbirds (Mimus polyglottos). Vet Parasitol. 2009;161(3-4):178–86.  https://doi.org/10.1016/j.vetpar.2009.01.023.CrossRefPubMedGoogle Scholar
  12. Appell LH, Mickelsen WD, Thomas MW, Harmon WM. A comparison of techniques used for the diagnosis of Tritrichomonas-fetus infections in beef bulls. Agri-Practice. 1993;14(2):30–4.Google Scholar
  13. Arranz-Solis D, Pedraza-Diaz S, Miro G, Rojo-Montejo S, Hernandez L, Ortega-Mora LM, Collantes-Fernandez E. Tritrichomonas foetus infection in cats with diarrhea from densely housed origins. Vet Parasitol. 2016;221:118–22.  https://doi.org/10.1016/j.vetpar.2016.03.019.CrossRefPubMedGoogle Scholar
  14. Ball L, Dargatz DA, Cheney JM, Mortimer RG. Control of venereal disease in infected herds. Vet Clin North Am Food Anim Pract. 1987;3(3):561–74.PubMedCrossRefGoogle Scholar
  15. Bartlett DE. Trichomonas foetus infection and bovine reproduction. Am J Vet Res. 1947;8(29):343–52.PubMedGoogle Scholar
  16. Bartlett DE, Hammond DM. Pattern of fluctuations in number of Trichomonas foetus occurring in the bovine vagina during initial infections. 2. Application in diagnosis. Am J Vet Res. 1945;6(19):91–5.Google Scholar
  17. Bartlett DE, Hasson EV, Teeter KG. Occurrence of Trichomonas foetus in preputial samples from infected bulls. J Am Vet Med Assoc. 1947;110(839):114–20.Google Scholar
  18. Bell ET, Gowan RA, Lingard AE, McCoy RJ, Slapeta J, Malik R. Naturally occurring Tritrichomonas foetus infections in Australian cats: 38 cases. J Feline Med Surg. 2010;12(12):889–98.  https://doi.org/10.1016/j.jfms.2010.06.003.CrossRefPubMedGoogle Scholar
  19. Benchimol M. Trichomonads under microscopy. Microsc Microanal. 2004;10(5):528–50.  https://doi.org/10.1017/S1431927604040905.CrossRefPubMedGoogle Scholar
  20. Benchimol M. New ultrastructural observations on the skeletal matrix of Tritrichomonas foetus. Parasitol Res. 2005;97(5):408–16.  https://doi.org/10.1007/s00436-005-1480-x.CrossRefPubMedGoogle Scholar
  21. Bernasconi C, Bodmer M, Doherr MG, Janett F, Thomann A, Spycher C, Iten C, Hentrich B, Gottstein B, Muller N, Frey CF. Tritrichomonas foetus: prevalence study in naturally mating bulls in Switzerland. Vet Parasitol. 2014;200(3-4):289–94.  https://doi.org/10.1016/j.vetpar.2013.12.029.CrossRefPubMedGoogle Scholar
  22. Bielanski A, Ghazi DF, Phipps-Toodd B. Observations on the fertilization and development of preimplantation bovine embryos in vitro in the presence of Tritrichomonas foetus. Theriogenology. 2004;61(5):821–9.PubMedCrossRefGoogle Scholar
  23. Blackshaw AW, Beattie HER. The preservation of Trichomonas foetus at -79°C. Aust Vet J. 1955;31:214–6.CrossRefGoogle Scholar
  24. Boal CW, Mannan RW, Hudelson KS. Trichomoniasis in Cooper’s hawks from Arizona. J Wildl Dis. 1998;34(3):590–3.  https://doi.org/10.7589/0090-3558-34.3.590.CrossRefPubMedGoogle Scholar
  25. BonDurant RH. Diagnosis, treatment, and control of bovine trichomoniasis. Comp Cont Educ Pract. 1985;7(3):S179–88.Google Scholar
  26. BonDurant RH. Pathogenesis, diagnosis, and management of trichomoniasis in cattle. Vet Clin North Am Food Anim Pract. 1997;13(2):345–61.PubMedCrossRefGoogle Scholar
  27. Bondurant RH. Venereal diseases of cattle: natural history, diagnosis, and the role of vaccines in their control. Vet Clin North Am Food Anim Pract. 2005;21(2):383–408.  https://doi.org/10.1016/j.cvfa.2005.03.002.CrossRefPubMedGoogle Scholar
  28. BonDurant RH. Selected diseases and conditions associated with bovine conceptus loss in the first trimester. Theriogenology. 2007;68(3):461–73.  https://doi.org/10.1016/j.theriogenology.2007.04.022.CrossRefPubMedGoogle Scholar
  29. BonDurant RH, Honigberg BM. Trichomonads of veterinary importance. In: Kreier JP, editor. Parasitic protozoa. New York, NY: Academic; 1994. p. 111–88.CrossRefGoogle Scholar
  30. BonDurant RH, Anderson ML, Blanchard P, Hird D, Danaye-Elmi C, Palmer C, Sischo WM, Suther D, Utterback W, Weigler BJ. Prevalence of trichomoniasis among California beef herds. J Am Vet Med Assoc. 1990;196(10):1590–3.PubMedGoogle Scholar
  31. BonDurant RH, Corbeil RR, Corbeil LB. Immunization of virgin cows with surface antigen TF1.17 of Tritrichomonas foetus. Infect Immun. 1993;61(4):1385–94.PubMedPubMedCentralGoogle Scholar
  32. BonDurant RH, van Hoosear KA, Corbeil LB, Bernoco D. Serological response to in vitro-shed antigen(s) of Tritrichomonas foetus in cattle. Clin Diagn Lab Immunol. 1996;3(4):432–7.PubMedPubMedCentralGoogle Scholar
  33. BonDurant RH, Gajadhar A, Campero CM. Preliminary characterization of a Tritrichomonas foetus-like protozoan isolated from preputial smegma of virgin bulls. The Bovine. Practitioner. 1999;33:124–7.Google Scholar
  34. Bryan LA, Campbell JR, Gajadhar AA. Effects of temperature on the survival of Tritrichomonas foetus in transport, Diamond’s and InPouch TF media. Vet Rec. 1999;144(9):227–32.PubMedCrossRefGoogle Scholar
  35. Bunbury N, Bell D, Jones C, Greenwood A, Hunter P. Comparison of the InPouch TF culture system and wet-mount microscopy for diagnosis of Trichomonas gallinae infections in the pink pigeon Columba mayeri. J Clin Microbiol. 2005;43(2):1005–6.  https://doi.org/10.1128/JCM.43.2.1005-1006.2005.CrossRefPubMedPubMedCentralGoogle Scholar
  36. Bunbury N, Jones CG, Greenwood AG, Bell DJ. Trichomonas gallinae in Mauritian columbids: implications for an endangered endemic. J Wildl Dis. 2007;43(3):399–407.  https://doi.org/10.7589/0090-3558-43.3.399.CrossRefPubMedGoogle Scholar
  37. Campero CM, Cobo ER. Tritrichomonas foetus: patogénesis de la mortalidad embrionaria/fetal, caracterización de antígenos vacunales y respuesta inmune inducida. Rev Med Vet. 2006;87:47–56.Google Scholar
  38. Campero C, Gottstein B. Control measures, tritrichomoniasis. In: Ortega-Mora LM, Gottstein B, Conraths FJ, Buxton D, editors. Protozoal abortion in farm ruminants. Wallingford, Oxfordshire: CAB International; 2007. p. 290–301.Google Scholar
  39. Campero CM, Ballabene NC, Cipolla AC, Zamora AS. Dual infection of bulls with campylobacteriosis and trichomoniasis: treatment with dimetridazole chlorhydrate. Aust Vet J. 1987;64(10):320–1.PubMedCrossRefGoogle Scholar
  40. Campero CM, Hirst RG, Ladds PW, Vaughan JA, Emery DL, Watson DL. Measurement of antibody in serum and genital fluids of bulls by ELISA after vaccination and challenge with Tritrichomonas foetus. Aust Vet J. 1990;67(5):175–8.PubMedCrossRefGoogle Scholar
  41. Campero CM, Rossetti O, Medina D, Bretschneider G, Roppel M. Inmunización en vaquillonas nediante vacuna de membrana de Tritrichomonas foetus. Vet Argent. 1999;154:250–62.Google Scholar
  42. Campero CM, Rodriguez Dubra C, Bolondi A, Cacciato C, Cobo E, Perez S, Odeon A, Cipolla A, BonDurant RH. Two-step (culture and PCR) diagnostic approach for differentiation of non-T. foetus trichomonads from genitalia of virgin beef bulls in Argentina. Vet Parasitol. 2003;112(3):167–75.PubMedCrossRefGoogle Scholar
  43. Case CH, Keefer WO. Some ways to detect and prevent the spread of trichomoniasis in cattle. J Am Vet Med Ass. 1938;93:239–40.Google Scholar
  44. Casteriano A, Molini U, Kandjumbwa K, Khaiseb S, Frey CF, Slapeta J. Novel genotype of Tritrichomonas foetus from cattle in Southern Africa. Parasitology. 2016:1–6.  https://doi.org/10.1017/S003118201600158X.
  45. Cauthen G. Studies on Trichomonas columbae, a flagellate parasitic in pigeons and doves. Am J Hyg. 1936;23:132–42.Google Scholar
  46. Cepicka I, Kutisova K, Tachezy J, Kulda J, Flegr J. Cryptic species within the Tetratrichomonas gallinarum species complex revealed by molecular polymorphism. Vet Parasitol. 2005;128(1-2):11–21.  https://doi.org/10.1016/j.vetpar.2004.11.003.CrossRefPubMedGoogle Scholar
  47. Cepicka I, Hampl V, Kulda J, Flegr J. New evolutionary lineages, unexpected diversity, and host specificity in the parabasalid genus Tetratrichomonas. Mol Phylogenet Evol. 2006;39(2):542–51.  https://doi.org/10.1016/j.ympev.2006.01.005.CrossRefPubMedGoogle Scholar
  48. Ceplecha V, Svoboda M, Cepicka I, Husnik R, Horackova K, Svobodova V. InPouch TF-Feline medium is not specific for Tritrichomonas foetus. Vet Parasitol. 2013;196(3-4):503–5.  https://doi.org/10.1016/j.vetpar.2013.04.015.CrossRefPubMedGoogle Scholar
  49. Chen XG, Li J. Increasing the sensitivity of PCR detection in bovine preputial smegma spiked with Tritrichomonas foetus by the addition of agar and resin. Parasitol Res. 2001;87(7):556–8.PubMedCrossRefGoogle Scholar
  50. Christensen HR, Clark BL, Parsonson IM. Incidence of Tritrichomonas foetus in young replacement bulls following introduction into an infected herd. Aust Vet J. 1977;53(3):132–4.PubMedCrossRefGoogle Scholar
  51. Clark BL, White MB, Banfield JC. Diagnosis of Trichomonas foetus infection in bulls. Aust Vet J. 1971;47(5):181–3.PubMedCrossRefGoogle Scholar
  52. Clark BL, Parsonson IM, Dufty JH. Letter: infection of bulls with Tritrichomonas foetus through mating with infected heifers. Aust Vet J. 1974;50(4):180.PubMedCrossRefGoogle Scholar
  53. Clark BL, Dufty JH, Parsonson IM. Studies on the transmission of Tritrichomonas foetus. Aust Vet J. 1977;53(4):170–2.PubMedCrossRefGoogle Scholar
  54. Clark BL, Dufty JH, Parsonson IM. The effect of Tritrichomonas foetus infection on calving rates in beef cattle. Aust Vet J. 1983a;60(3):71–4.PubMedCrossRefGoogle Scholar
  55. Clark BL, Dufty JH, Parsonson IM. Immunisation of bulls against trichomoniasis. Aust Vet J. 1983b;60(6):178–9.PubMedCrossRefGoogle Scholar
  56. Clark BL, Emery DL, Dufty JH. Therapeutic immunisation of bulls with the membranes and glycoproteins of Tritrichomonas foetus var. Brisbane. Aust Vet J. 1984;61(2):65–6.PubMedCrossRefGoogle Scholar
  57. Clark BL, Dufty JH, Parsonson IM. The frequency of infertility and abortion in cows infected with Tritrichomonas foetus var. brisbane. Aust Vet J. 1986;63(1):31–2.PubMedCrossRefGoogle Scholar
  58. Clark S, De Gussem K, Barnes J. Flagellated protozoan infections in turkeys. World Poultry Turkey Special. 2003;5:20–3.Google Scholar
  59. Cobo ER, Cano D, Rossetti O, Campero CM. Heifers immunized with whole-cell and membrane vaccines against Tritrichomonas foetus and naturally challenged with an infected bull. Vet Parasitol. 2002;109(3-4):169–84.PubMedCrossRefGoogle Scholar
  60. Cobo ER, Favetto PH, Lane VM, Friend A, VanHooser K, Mitchell J, BonDurant RH. Sensitivity and specificity of culture and PCR of smegma samples of bulls experimentally infected with Tritrichomonas foetus. Theriogenology. 2007;68(6):853–60.  https://doi.org/10.1016/j.theriogenology.2007.06.019.
  61. Cobo ER, Morsella C, Cano D, Cipolla A, Campero CM. Immunization in heifers with dual vaccines containing Tritrichomonas foetus and Campylobacter fetus antigens using systemic and mucosal routes. Theriogenology. 2004;62(8):1367–82.  https://doi.org/10.1016/j.theriogenology.2003.12.034.CrossRefPubMedGoogle Scholar
  62. Cobo ER, Corbeil LB, Gershwin LJ, BonDurant RH. Preputial cellular and antibody responses of bulls vaccinated and/or challenged with Tritrichomonas foetus. Vaccine. 2009;28(2):361–70.  https://doi.org/10.1016/j.vaccine.2009.10.039.CrossRefPubMedGoogle Scholar
  63. Cole R, Friend M. Trichomoniasis. In: Friend M, Franson JC, editors. Trichomoniasis. Washington, DC: USGS-National Wildlife Health Center; 1999. p. 201–6.Google Scholar
  64. Collantes-Fernandez E, Mendoza-Ibarra JA, Pedraza-Diaz S, Rojo-Montejo S, Navarro-Lozano V, Sanchez-Sanchez R, Ruiz-Santa-Quiteria JA, Ortega-Mora LM, Osoro K. Efficacy of a control program for bovine trichomonosis based on testing and culling infected bulls in beef cattle managed under mountain pastoral systems of Northern Spain. Vet J. 2014;200(1):140–5.  https://doi.org/10.1016/j.tvjl.2014.02.003.CrossRefPubMedGoogle Scholar
  65. Conraths FJ, Schares G. Validation of molecular-diagnostic techniques in the parasitological laboratory. Vet Parasitol. 2006;136(2):91–8.PubMedCrossRefGoogle Scholar
  66. Cooper JE, Petty SJ. Trichomoniasis in free-living goshawks (Accipiter gentilis gentilis) from Great Britain. J Wildl Dis. 1988;24(1):80–7.  https://doi.org/10.7589/0090-3558-24.1.80.CrossRefPubMedGoogle Scholar
  67. Corbeil LB, Anderson ML, Corbeil RR, Eddow JM, BonDurant RH. Female reproductive tract immunity in bovine trichomoniasis. Am J Reprod Immunol. 1998;39(3):189–98.PubMedCrossRefGoogle Scholar
  68. Corbeil LB, Campero CM, Van Hoosear K, Bondurant RH. Detection of trichomonad species in the reproductive tracts of breeding and virgin bulls. Vet Parasitol. 2008;154(3-4):226–32.  https://doi.org/10.1016/j.vetpar.2008.03.014.CrossRefPubMedGoogle Scholar
  69. Dahlgren SS, Gjerde B, Pettersen HY. First record of natural Tritrichomonas foetus infection of the feline uterus. J Small Anim Pract. 2007;48(11):654–7.  https://doi.org/10.1111/j.1748-5827.2007.00405.x.CrossRefPubMedGoogle Scholar
  70. De Carli GA, Tasca T. Trichomonas gallinae: a possible contact-dependent mechanism in the hemolytic activity. Vet Parasitol. 2002;106(4):277–83.PubMedCrossRefGoogle Scholar
  71. De Carli GA, da Silva AC, Wendorff A, Rott M. Lysis of erythrocytes by Trichomonas gallinae. Avian Dis. 1996;40:228–30.PubMedCrossRefGoogle Scholar
  72. de Oliveira JM, da Silva GM, Batista Filho AF, de Melo Borges J, de Oliveira PR, Brandespim DF, Mota RA, Pinheiro JW Jr. Prevalence and risk factors associated with bovine genital campylobacteriosis and bovine trichomonosis in the state of Pernambuco, Brazil. Trop Anim Health Prod. 2015;47(3):549–55.  https://doi.org/10.1007/s11250-015-0761-3.CrossRefPubMedGoogle Scholar
  73. Dennett DP, Reece RL, Barasa JO, Johnson RH. Observations on the incidence and distribution of serotypes of Tritrichomonas foetus in beef cattle in north-eastern Australia. Aust Vet J. 1974;50(10):427–31.PubMedCrossRefGoogle Scholar
  74. Dewell GA, Phillips PE, Dohlman TM, Harmon KM, Gauger PC. Validation of a gauze sponge sampling methodology to detect Tritrichomonas foetus by real-time PCR. J Vet Diagn Invest. 2016;28(5):595–8.  https://doi.org/10.1177/1040638716653637.PubMedCrossRefGoogle Scholar
  75. Diamond LS. The establishment of various trichomonads of animals and man in axenic cultures. J Parasitol. 1957;43(4):488–90.PubMedCrossRefGoogle Scholar
  76. Doi J, Hirota J, Morita A, Fukushima K, Kamijyo H, Ohta H, Yamasaki M, Takahashi T, Katakura K, Oku Y. Intestinal Tritrichomonas suis (=T. foetus) infection in Japanese cats. J Vet Med Sci. 2012;74(4):413–7.PubMedCrossRefGoogle Scholar
  77. Duboucher C, Caby S, Dufernez F, Chabe M, Gantois N, Delgado-Viscogliosi P, Billy C, Barre E, Torabi E, Capron M, Pierce RJ, Dei-Cas E, Viscogliosi E. Molecular identification of Tritrichomonas foetus-like organisms as coinfecting agents of human Pneumocystis pneumonia. J Clin Microbiol. 2006;44(3):1165–8.  https://doi.org/10.1128/JCM.44.3.1165-1168.2006.CrossRefPubMedPubMedCentralGoogle Scholar
  78. Eaglesome MD, Garcia MM. Disease risks to animal health from artificial insemination with bovine semen. Rev Sci Tech. 1997;16(1):215–25.PubMedCrossRefGoogle Scholar
  79. Erwin KG, Kloss C, Lyles J, Felderhoff J, Fedynich AM, Henke SE, Roberson JA. Survival of Trichomonas gallinae in white-winged dove carcasses. J Wildl Dis. 2000;36(3):551–4.  https://doi.org/10.7589/0090-3558-36.3.551.CrossRefPubMedGoogle Scholar
  80. Estes WA, Mannan RW. Feeding behavior of Cooper’s Hawks at urban and rural nests in south-eastern Arizona. Condor. 2003;105(1):107–16.  https://doi.org/10.1650/0010-5422(2003)105[107:Fbocha]2.0.Co;2.CrossRefGoogle Scholar
  81. Felleisen RS. Comparative sequence analysis of 5.8S rRNA genes and internal transcribed spacer (ITS) regions of trichomonadid protozoa. Parasitology. 1997;115(Pt 2):111–9.PubMedCrossRefGoogle Scholar
  82. Felleisen RS. Comparative genetic analysis of tritrichomonadid protozoa by the random amplified polymorphic DNA technique. Parasitol Res. 1998;84(2):153–6.PubMedCrossRefGoogle Scholar
  83. Felleisen RS, Lambelet N, Bachmann P, Nicolet J, Muller N, Gottstein B. Detection of Tritrichomonas foetus by PCR and DNA enzyme immunoassay based on rRNA gene unit sequences. J Clin Microbiol. 1998;36(2):513–9.PubMedPubMedCentralGoogle Scholar
  84. Fitzgerald PR. Bovine trichomoniasis. Vet Clin North Am Food Anim Pract. 1986;2:277–82.PubMedCrossRefGoogle Scholar
  85. Fitzgerald PR, Johnson AE, Hammond DM, Thorne JL, Hibler CP. Experimental infection of young pigs following intranasal inoculation with nasal, gastric, or cecal trichomonads from swine or with Trichomonas foetus. J Parasitol. 1958;44(6):597–602.PubMedCrossRefGoogle Scholar
  86. Flower PJ, Ladds PW, Thomas AD, Watson DL. An immunopathologic study of the bovine prepuce. Vet Pathol. 1983;20(2):189–202.PubMedCrossRefGoogle Scholar
  87. Forrester DJ, Foster GW. Trichomonosis. In: Atkinson CT, Thomas NJ, Hunter DB, editors. Parasitic diseases of wild birds. Ames, IA: Wiley-Blackwell; 2008. p. 120–53.Google Scholar
  88. Fort M, Dubié D, Sago A, Goyeneche P, Fernández E, Collantes-Fernández E, Sánchez Sánchez R, Moreno Gonzalo J, Ortega-Mora LM. Evaluation of the performance of bovine trichomonosis control program in La Pampa-Argentina. Paper presented at the XXI Inter Comgress ANEMBE, Santiago de Compostela, Spain; 2016.Google Scholar
  89. Foster DM, Gookin JL, Poore MF, Stebbins ME, Levy MG. Outcome of cats with diarrhea and Tritrichomonas foetus infection. J Am Vet Med Assoc. 2004;225(6):888–92.PubMedCrossRefGoogle Scholar
  90. Franssen FF, Lumeij JT. In vitro nitroimidazole resistance of Trichomonas gallinae and successful therapy with an increased dosage of ronidazole in racing pigeons (Columba livia domestica). J Vet Pharmacol Ther. 1992;15(4):409–15.PubMedCrossRefGoogle Scholar
  91. Frey CF, Müller N. Tritrichomonas—systematics of an enigmatic genus. Mol Cell Probes. 2012;26(3):132–6.PubMedCrossRefGoogle Scholar
  92. Frey CF, Schild M, Hemphill A, Stunzi P, Muller N, Gottstein B, Burgener IA. Intestinal Tritrichomonas foetus infection in cats in Switzerland detected by in vitro cultivation and PCR. Parasitol Res. 2009;104(4):783–8.  https://doi.org/10.1007/s00436-008-1255-2.CrossRefPubMedGoogle Scholar
  93. Friedhoff KT. Pathogene, intestinale Flagellaten bei Tauben, Sittichen und Papageien. Collegium Veterinarium. 1982;63:329–34.Google Scholar
  94. Friedhoff KT, Kuhnigk C, Muller I. Experimental infections in chickens with Chilomastix gallinarum, Tetratrichomonas gallinarum, and Tritrichomonas eberthi. Parasitol Res. 1991;77(4):329–34.PubMedCrossRefGoogle Scholar
  95. Galián M, Gentil M, Heusinger A, Müller E. Tritrichomonas foetus as a cause of diarrhoea in cats. Tierärztliche Umschau Parasiten-Spezial. 2011a;1:5–8.Google Scholar
  96. Galián M, Heusinger A, Gentil M, Müller E. Tritrichomonas foetus en el gato. Argos Informativo Veterinario. 2011b;134:44–5.Google Scholar
  97. Garcia Guerra A, Hill JE, Waldner CL, Campbell J, Hendrick S. Sensitivity of a real-time polymerase chain reaction for Tritrichomonas fetus in direct individual and pooled preputial samples. Theriogenology. 2013;80(9):1097–103.  https://doi.org/10.1016/j.theriogenology.2013.08.011.PubMedCrossRefGoogle Scholar
  98. Gaspar da Silva D, Barton E, Bunbury N, Lunness P, Bell DJ, Tyler KM. Molecular identity and heterogeneity of trichomonad parasites in a closed avian population. Infect Genet Evol. 2007;7(4):433–40.  https://doi.org/10.1016/j.meegid.2007.01.002.CrossRefPubMedGoogle Scholar
  99. Gault RA, Kvasnicka WG, Hanks D, Hanks M, Hall MR. Specific antibodies in serum and vaginal mucus of heifers inoculated with a vaccine containing Tritrichomonas foetus. Am J Vet Res. 1995;56(4):454–9.PubMedGoogle Scholar
  100. Gay JM, Ebel ED, Kearley WP. Commingled grazing as a risk factor for trichomonosis in beef herds. J Am Vet Med Assoc. 1996;209(3):643–6.PubMedGoogle Scholar
  101. Gerhold RW, Tate CM, Gibbs SE, Mead DG, Allison AB, Fischer JR. Necropsy findings and arbovirus surveillance in mourning doves from the southeastern United States. J Wildl Dis. 2007;43(1):129–35.  https://doi.org/10.7589/0090-3558-43.1.129.CrossRefPubMedGoogle Scholar
  102. Gerhold RW, Yabsley MJ, Smith AJ, Ostergaard E, Mannan W, Cann JD, Fischer JR. Molecular characterization of the Trichomonas gallinae morphologic complex in the United States. J Parasitol. 2008;94(6):1335–41.  https://doi.org/10.1645/GE-1585.1.CrossRefPubMedGoogle Scholar
  103. Girard YA, Rogers KH, Gerhold R, Land KM, Lenaghan SC, Woods LW, Haberkern N, Hopper M, Cann JD, Johnson CK. Trichomonas stableri n. sp., an agent of trichomonosis in Pacific Coast band-tailed pigeons (Patagioenas fasciata monilis). Int J Parasitol Parasites Wildl. 2014;3(1):32–40.  https://doi.org/10.1016/j.ijppaw.2013.12.002.CrossRefPubMedGoogle Scholar
  104. Goodger WJ, Skirrow SZ. Epidemiologic and economic analyses of an unusually long epizootic of trichomoniasis in a large California dairy herd. J Am Vet Med Assoc. 1986;189(7):772–6.PubMedGoogle Scholar
  105. Gookin JL, Breitschwerdt EB, Levy MG, Gager RB, Benrud JG. Diarrhea associated with trichomonosis in cats. J Am Vet Med Assoc. 1999;215(10):1450–4.PubMedGoogle Scholar
  106. Gookin JL, Levy MG, Law JM, Papich MG, Poore MF, Breitschwerdt EB. Experimental infection of cats with Tritrichomonas foetus. Am J Vet Res. 2001;62(11):1690–7.PubMedCrossRefGoogle Scholar
  107. Gookin JL, Birkenheuer AJ, Breitschwerdt EB, Levy MG. Single-tube nested PCR for detection of Tritrichomonas foetus in feline feces. J Clin Microbiol. 2002;40(11):4126–30.PubMedPubMedCentralCrossRefGoogle Scholar
  108. Gookin JL, Foster DM, Poore MF, Stebbins ME, Levy MG. Use of a commercially available culture system for diagnosis of Tritrichomonas foetus infection in cats. J Am Vet Med Assoc. 2003;222(10):1376–9.PubMedCrossRefGoogle Scholar
  109. Gookin JL, Stebbins ME, Hunt E, Burlone K, Fulton M, Hochel R, Talaat M, Poore M, Levy MG. Prevalence of and risk factors for feline Tritrichomonas foetus and Giardia infection. J Clin Microbiol. 2004;42(6):2707–10.  https://doi.org/10.1128/JCM.42.6.2707-2710.2004.CrossRefPubMedPubMedCentralGoogle Scholar
  110. Gookin JL, Birkenheuer AJ, St John V, Spector M, Levy MG. Molecular characterization of trichomonads from feces of dogs with diarrhea. J Parasitol. 2005;91(4):939–43.  https://doi.org/10.1645/GE-474R.1.CrossRefPubMedGoogle Scholar
  111. Gookin JL, Copple CN, Papich MG, Poore MF, Stauffer SH, Birkenheuer AJ, Twedt DC, Levy MG. Efficacy of ronidazole for treatment of feline Tritrichomonas foetus infection. J Vet Int Med. 2006;20(3):536–43.CrossRefGoogle Scholar
  112. Gookin JL, Stauffer SH, Dybas D, Cannon DH. Documentation of in vivo and in vitro aerobic resistance of feline Tritrichomonas foetus isolates to ronidazole. J Vet Int Med. 2010a;24(4):1003–7.  https://doi.org/10.1111/j.1939-1676.2010.0534.x.CrossRefGoogle Scholar
  113. Gookin JL, Stone MR, Yaeger MJ, Meyerholz DK, Moisan P. Fluorescence in situ hybridization for identification of Tritrichomonas foetus in formalin-fixed and paraffin-embedded histological specimens of intestinal trichomoniasis. Vet Parasitol. 2010b;172(1-2):139–43.  https://doi.org/10.1016/j.vetpar.2010.04.014.CrossRefPubMedGoogle Scholar
  114. Grabensteiner E, Hess M. PCR for the identification and differentiation of Histomonas meleagridis, Tetratrichomonas gallinarum and Blastocystis spp. Vet Parasitol. 2006;142(3-4):223–30.  https://doi.org/10.1016/j.vetpar.2006.07.011.CrossRefPubMedGoogle Scholar
  115. Grabensteiner E, Bilic I, Kolbe T, Hess M. Molecular analysis of clonal trichomonad isolates indicate the existence of heterogenic species present in different birds and within the same host. Vet Parasitol. 2010;172(1-2):53–64.  https://doi.org/10.1016/j.vetpar.2010.04.015.CrossRefPubMedGoogle Scholar
  116. Grahn RA, BonDurant RH, van Hoosear KA, Walker RL, Lyons LA. An improved molecular assay for Tritrichomonas foetus. Vet Parasitol. 2005;127(1):33–41.  https://doi.org/10.1016/j.vetpar.2004.08.018.CrossRefPubMedGoogle Scholar
  117. Gray SG, Hunter SA, Stone MR, Gookin JL. Assessment of reproductive tract disease in cats at risk for Tritrichomonas foetus infection. Am J Vet Res. 2010;71(1):76–81.  https://doi.org/10.2460/ajvr.71.1.76.CrossRefPubMedGoogle Scholar
  118. Grellet A, Bickel T, Polack B, Boogaerts C, Casseleux G, Biourge V, Grandjean D. Prevalence of Tritrichomonas foetus in puppies from French breeding kennels. J Vet Int Med. 2010;24:1572.Google Scholar
  119. Gunn-Moore DA, McCann TM, Reed N, Simpson KE, Tennant B. Prevalence of Tritrichomonas foetus infection in cats with diarrhoea in the UK. J Feline Med Surg. 2007;9(3):214–8.  https://doi.org/10.1016/j.jfms.2007.01.003.CrossRefPubMedGoogle Scholar
  120. Guven E, Bastem Z, Avcioglu H, Erdem H. Molecular determination of Tritrichomonas spp. in aborted bovine foetuses in Eastern Anatolian Region of Turkey. Vet Parasitol. 2013;196(3-4):278–82.  https://doi.org/10.1016/j.vetpar.2013.03.031.CrossRefPubMedGoogle Scholar
  121. Hale S, Norris JM, Slapeta J. Prolonged resilience of Tritrichomonas foetus in cat faeces at ambient temperature. Vet Parasitol. 2009;166(1-2):60–5.  https://doi.org/10.1016/j.vetpar.2009.07.032.CrossRefPubMedGoogle Scholar
  122. Hammond DM, Bartlett DE. The distribution of Tritrichomonas foetus in the preputial cavity of infected bulls. Am J Vet Res. 1943;4:143–9.Google Scholar
  123. Hammond DM, Bartlett DE. Pattern of fluctuations in numbers of Trichomonas foetus occurring in the bovine vagina during initial infections. 1. Correlation with time of exposure and with subsequent estrual cycles. Am J Vet Res. 1945;6(19):84–90.Google Scholar
  124. Hampl V, Pavlicek A, Flegr J. Construction and bootstrap analysis of DNA fingerprinting-based phylogenetic trees with the freeware program FreeTree: application to trichomonad parasites. Int J Syst Evol Microbiol. 2001;51(Pt 3):731–5.  https://doi.org/10.1099/00207713-51-3-731.CrossRefPubMedGoogle Scholar
  125. Harmon WM, Clark WA, Hawbecker AC, Stafford M. Trichomonas gallinae in columbiform birds from the Galapagos Islands. J Wildl Dis. 1987;23(3):492–4.PubMedCrossRefGoogle Scholar
  126. Hawn C. Trichomoniasis of turkeys. J Infect Dis. 1937;61:184–97.CrossRefGoogle Scholar
  127. Hayes DC, Anderson RR, Walker RL. Identification of trichomonadid protozoa from the bovine preputial cavity by polymerase chain reaction and restriction fragment length polymorphism typing. J Vet Diagn Invest. 2003;15(4):390–4.PubMedCrossRefGoogle Scholar
  128. Herr S, Ribeiro LM, Claassen E, Myburgh JG. A reduction in the duration of infection with Tritrichomonas foetus following vaccination in heifers and the failure to demonstrate a curative effect in infected bulls. Onderstepoort J Vet Res. 1991;58(1):41–5.PubMedGoogle Scholar
  129. Hibler CP, Hammond DM, Caskey FH, Johnson AE, Fitzgerald PR. The morphology and incidence of the trichomonads of swine, Tritrichomonas-suis (Gruby and Delafond), Tritrichomonas-rotunda, nsp and Trichomonas-buttreyi, nsp. J Protozool. 1960;7(2):159–71.  https://doi.org/10.1111/j.1550-7408.1960.tb00725.x.CrossRefGoogle Scholar
  130. Ho MS, Conrad PA, Conrad PJ, LeFebvre RB, Perez E, BonDurant RH. Detection of bovine trichomoniasis with a specific DNA probe and PCR amplification system. J Clin Microbiol. 1994;32(1):98–104.PubMedPubMedCentralGoogle Scholar
  131. Hodgson JL, Jones DW, Widders PR, Corbeil LB. Characterization of Tritrichomonas foetus antigens by use of monoclonal antibodies. Infect Immun. 1990;58(9):3078–83.PubMedPubMedCentralGoogle Scholar
  132. Höfle U, Blanco J, Palma L, Melo P. Trichomoniasis in Bonelli’s eagle nestlings in south-west Portugal. In: Redig PT, Cooper JE, Remple TD, editors. Raptor Biomedicine III. Minneapolis, MN: University of Minnesota Press; 2000. p. 45–51.Google Scholar
  133. Holliday M, Deni D, Gunn-Moore DA. Tritrichomonas foetus infection in cats with diarrhoea in a rescue colony in Italy. J Feline Med Surg. 2009;11(2):131–4.  https://doi.org/10.1016/j.jfms.2008.06.004.CrossRefPubMedGoogle Scholar
  134. Hoorfar J, Wolffs P, Radstrom P. Diagnostic PCR: validation and sample preparation are two sides of the same coin. APMIS. 2004;112(11-12):808–14.PubMedCrossRefGoogle Scholar
  135. Hosein A, Kruth SA, Pearl DL, Richardson D, Maggs JC, Peach HA, Peregrine AS. Isolation of Tritrichomonas foetus from cats sampled at a cat clinic, cat shows and a humane society in southern Ontario. J Feline Med Surg. 2013;15(8):706–11.  https://doi.org/10.1177/1098612X13475617.CrossRefPubMedGoogle Scholar
  136. Huby-Chilton F, Scandrett BW, Chilton NB, Gajadhar AA. Detection and identification of Tetratrichomonas in a preputial wash from a bull by PCR and SSCP. Vet Parasitol. 2009;166(3-4):199–204.  https://doi.org/10.1016/j.vetpar.2009.08.026.CrossRefPubMedGoogle Scholar
  137. Hudson DB, Ball L, Cheney JM, Mortimer RG, Bowen RA, Marsh DJ, Peetz RH. Development and testing of a bovine trichomoniasis vaccine. Theriogenology. 1993;39(4):929–35.PubMedCrossRefGoogle Scholar
  138. Ikeda JS, BonDurant RH, Corbeil LB. Bovine vaginal antibody responses to immunoaffinity-purified surface antigen of Tritrichomonas foetus. J Clin Microbiol. 1995;33(5):1158–63.PubMedPubMedCentralGoogle Scholar
  139. Jin Y, Schumaker B, Logan J, Yao C. Risk factors associated with bovine trichomoniasis in beef cattle identified by a questionnaire. J Med Microbiol. 2014;63(Pt 6):896–902.  https://doi.org/10.1099/jmm.0.074971-0.CrossRefPubMedGoogle Scholar
  140. Jongwutiwes S, Silachamroon U, Putaporntip C. Pentatrichomonas hominis in empyema thoracis. Trans R Soc Trop Med Hyg. 2000;94(2):185–6.PubMedCrossRefGoogle Scholar
  141. Kerr WR. The intradermal test in bovine trichomoniasis. Vet Rec. 1944;56:303–5.Google Scholar
  142. Kimsey PB, Darien BJ, Kendrick JW, Franti CE. Bovine trichomoniasis—diagnosis and treatment. J Am Vet Med Assoc. 1980;177(7):616–9.PubMedGoogle Scholar
  143. Kittel DR, Campero C, Van Hoosear KA, Rhyan JC, BonDurant RH. Comparison of diagnostic methods for detection of active infection with Tritrichomonas foetus in beef heifers. J Am Vet Med Assoc. 1998;213(4):519–22.PubMedGoogle Scholar
  144. Kleina P, Bettim-Bandinelli J, Bonatto SL, Benchimol M, Bogo MR. Molecular phylogeny of Trichomonadidae family inferred from ITS-1, 5.8S rRNA and ITS-2 sequences. Int J Parasitol. 2004;34(8):963–70.  https://doi.org/10.1016/j.ijpara.2004.04.004.CrossRefPubMedGoogle Scholar
  145. Kleydman Y, Yarlett N, Gorrell TE. Production of ammonia by Tritrichomonas foetus and Trichomonas vaginalis. Microbiology. 2004;150(Pt 5):1139–45.  https://doi.org/10.1099/mic.0.26939-0.CrossRefPubMedGoogle Scholar
  146. Kolisko M, Cepicka I, Hampl V, Leigh J, Roger AJ, Kulda J, Simpson AG, Flegr J. Molecular phylogeny of diplomonads and enteromonads based on SSU rRNA, alpha-tubulin and HSP90 genes: implications for the evolutionary history of the double karyomastigont of diplomonads. BMC Evol Biol. 2008;8:205.PubMedPubMedCentralCrossRefGoogle Scholar
  147. Krone O, Altenkamp R, Kenntner N. Prevalence of Trichomonas gallinae in northern goshawks from the Berlin area of northeastern Germany. J Wildl Dis. 2005;41(2):304–9.  https://doi.org/10.7589/0090-3558-41.2.304.CrossRefPubMedGoogle Scholar
  148. Kuehner KA, Marks SL, Kass PH, Sauter-Louis C, Grahn RA, Barutzki D, Hartmann K. Tritrichomonas foetus infection in purebred cats in Germany: prevalence of clinical signs and the role of co-infection with other enteroparasites. J Feline Med Surg. 2011;13(4):251–8.  https://doi.org/10.1016/j.jfms.2010.12.002.CrossRefPubMedGoogle Scholar
  149. Kvasnicka WG, Taylor RE, Huang JC, Hanks D, Tronstad RJ, Bosomworth A, Hall MR. Investigations of the incidence of bovine trichomoniasis in Nevada and of the efficacy of immunizing cattle with vaccines containing Tritrichomonas foetus. Theriogenology. 1989;31(5):963–71.PubMedCrossRefGoogle Scholar
  150. Kvasnicka WG, Hanks D, Huang JC, Hall MR, Sandblom D, Chu HJ, Chavez L, Acree WM. Clinical evaluation of the efficacy of inoculating cattle with a vaccine containing Tritrichomonas foetus. Am J Vet Res. 1992;53(11):2023–7.PubMedGoogle Scholar
  151. Lawson B, Robinson RA, Neimanis A, Handeland K, Isomursu M, Agren EO, Hamnes IS, Tyler KM, Chantrey J, Hughes LA, Pennycott TW, Simpson VR, John SK, Peck KM, Toms MP, Bennett M, Kirkwood JK, Cunningham AA. Evidence of spread of the emerging infectious disease, finch trichomonosis, by migrating birds. EcoHealth. 2011;8(2):143–53.  https://doi.org/10.1007/s10393-011-0696-8.CrossRefPubMedGoogle Scholar
  152. Levine D, Brandly A. A pathogenic trichomonas from the upper digestive tract of chickens. J Am Vet Med Assoc. 1939;95:77–8.Google Scholar
  153. Levy MG, Gookin JL, Poore M, Birkenheuer AJ, Dykstra MJ, Litaker RW. Tritrichomonas foetus and not Pentatrichomonas hominis is the etiologic agent of feline trichomonal diarrhea. J Parasitol. 2003;89(1):99–104.  https://doi.org/10.1645/0022-3395(2003)089[0099:TFANPH]2.0.CO;2.CrossRefPubMedGoogle Scholar
  154. Liebhart D, Weissenbock H, Hess M. In-situ hybridization for the detection and identification of Histomonas meleagridis in tissues. J Comp Pathol. 2006;135(4):237–42.  https://doi.org/10.1016/j.jcpa.2006.08.002.PubMedCrossRefGoogle Scholar
  155. Longo MC, Berninger MS, Hartley JL. Use of uracil DNA glycosylase to control carry-over contamination in polymerase chain reactions. Gene. 1990;93:125–8.PubMedCrossRefGoogle Scholar
  156. Lun ZR, Gajadhar AA. A simple and rapid method for staining Tritrichomonas foetus and Trichomonas vaginalis. J Vet Diagn Invest. 1999;11(5):471–4.PubMedCrossRefGoogle Scholar
  157. Lun Z, Parker S, Gajadhar AA. Comparison of growth rates of Tritrichomonas foetus isolates from various geographic regions using three different culture media. Vet Parasitol. 2000;89(3):199–208.PubMedCrossRefGoogle Scholar
  158. Lun ZR, Chen XG, Zhu XQ, Li XR, Xie MQ. Are Tritrichomonas foetus and Tritrichomonas suis synonyms? Trends Parasitol. 2005;21(3):122–5.  https://doi.org/10.1016/j.pt.2004.12.001.CrossRefPubMedGoogle Scholar
  159. Madoroba E, Gelaw A, Hlokwe T, Mnisi M. Prevalence of Campylobacter foetus and Trichomonas foetus among cattle from Southern Africa. Afr J Biotechnol. 2011;10(50):10311–4.CrossRefGoogle Scholar
  160. Mancebo OA, Russo AM, Carabajal LL, Monzon CM. Persistence of Tritrichomonas foetus in naturally infected cows and heifers in Argentina. Vet Parasitol. 1995;59(1):7–11.PubMedCrossRefGoogle Scholar
  161. Manning K. Update on the diagnosis and management of Tritrichomonas foetus infections in cats. Top Companion Anim Med. 2010;25(3):145–8.  https://doi.org/10.1053/j.tcam.2010.08.001.CrossRefPubMedGoogle Scholar
  162. Mardones FO, Perez AM, Martinez A, Carpenter TE. Risk factors associated with Tritrichomonas foetus infection in beef herds in the Province of Buenos Aires, Argentina. Vet Parasitol. 2008;153(3-4):231–7.  https://doi.org/10.1016/j.vetpar.2008.01.038.CrossRefPubMedGoogle Scholar
  163. McCool CJ, Townsend MP, Wolfe SG, Simpson MA, Olm TC, Jayawardhana GA, Carney JV. Prevalence of bovine venereal disease in the Victoria River District of the Northern Territory: likely economic effects and practicable control measures. Aust Vet J. 1988;65(5):153–6.PubMedCrossRefGoogle Scholar
  164. McMillen L, Lew AE. Improved detection of Tritrichomonas foetus in bovine diagnostic specimens using a novel probe-based real time PCR assay. Vet Parasitol. 2006;141(3-4):204–15.  https://doi.org/10.1016/j.vetpar.2006.06.012.CrossRefPubMedGoogle Scholar
  165. Mehlhorn H, Al-Quraishy S, Aziza A, Hess M. Fine structure of the bird parasites Trichomonas gallinae and Tetratrichomonas gallinarum from cultures. Parasitol Res. 2009;105(3):751–6.  https://doi.org/10.1007/s00436-009-1451-8.CrossRefPubMedGoogle Scholar
  166. Mendoza-Ibarra JA, Pedraza-Diaz S, Garcia-Pena FJ, Rojo-Montejo S, Ruiz-Santa-Quiteria JA, San Miguel-Ibanez E, Navarro-Lozano V, Ortega-Mora LM, Osoro K, Collantes-Fernandez E. High prevalence of Tritrichomonas foetus infection in Asturiana de la Montana beef cattle kept in extensive conditions in northern Spain. Vet J. 2012;193(1):146–51.  https://doi.org/10.1016/j.tvjl.2011.09.020.CrossRefPubMedGoogle Scholar
  167. Mendoza-Ibarra JA, Ortega-Mora LM, Pedraza-Diaz S, Rojo-Montejo S, Ruiz-Santa-Quiteria JA, Garcia-Pena FJ, Navarro-Lozano V, Cuevas-Martin Mdel C, Osoro K, Collantes-Fernandez E. Differences in the prevalence of Tritrichomonas foetus infection in beef cattle farmed under extensive conditions in northern Spain. Vet J. 2013;196(3):547–9.  https://doi.org/10.1016/j.tvjl.2012.10.026.CrossRefPubMedGoogle Scholar
  168. Michi AN, Favetto PH, Kastelic J, Cobo ER. A review of sexually transmitted bovine trichomoniasis and campylobacteriosis affecting cattle reproductive health. Theriogenology. 2016;85(5):781–91.  https://doi.org/10.1016/j.theriogenology.2015.10.037.CrossRefPubMedGoogle Scholar
  169. Molina L, Perea J, Meglia G, Angon E, Garcia A. Spatial and temporal epidemiology of bovine trichomoniasis and bovine genital campylobacteriosis in La Pampa province (Argentina). Prev Vet Med. 2013;110(3-4):388–94.  https://doi.org/10.1016/j.prevetmed.2013.02.019.CrossRefPubMedGoogle Scholar
  170. Morin-Adeline V, Lomas R, O'Meally D, Stack C, Conesa A, Slapeta J. Comparative transcriptomics reveals striking similarities between the bovine and feline isolates of Tritrichomonas foetus: consequences for in silico drug-target identification. BMC Genomics. 2014;15:955.  https://doi.org/10.1186/1471-2164-15-955.CrossRefPubMedPubMedCentralGoogle Scholar
  171. Morin-Adeline V, Fraser ST, Stack C, Slapeta J. Host origin determines pH tolerance of Tritrichomonas foetus isolates from the feline gastrointestinal and bovine urogenital tracts. Exp Parasitol. 2015a;157:68–77.  https://doi.org/10.1016/j.exppara.2015.06.017.CrossRefPubMedGoogle Scholar
  172. Morin-Adeline V, Mueller K, Conesa A, Slapeta J. Comparative RNA-seq analysis of the Tritrichomonas foetus PIG30/1 isolate from pigs reveals close association with Tritrichomonas foetus BP-4 isolate ‘bovine genotype’. Vet Parasitol. 2015b;212(3-4):111–7.  https://doi.org/10.1016/j.vetpar.2015.08.012.CrossRefPubMedGoogle Scholar
  173. Mostegl MM, Richter B, Nedorost N, Maderner A, Dinhopl N, Weissenbock H. Investigations on the prevalence and potential pathogenicity of intestinal trichomonads in pigs using in situ hybridization. Vet Parasitol. 2011;178(1-2):58–63.  https://doi.org/10.1016/j.vetpar.2010.12.022.CrossRefPubMedPubMedCentralGoogle Scholar
  174. Mostegl MM, Wetscher A, Richter B, Nedorost N, Dinhopl N, Weissenbock H. Detection of Tritrichomonas foetus and Pentatrichomonas hominis in intestinal tissue specimens of cats by chromogenic in situ hybridization. Vet Parasitol. 2012;183(3-4):209–14.  https://doi.org/10.1016/j.vetpar.2011.07.050.CrossRefPubMedPubMedCentralGoogle Scholar
  175. Mueller K, Morin-Adeline V, Gilchrist K, Brown G, Slapeta J. High prevalence of Tritrichomonas foetus ‘bovine genotype’ in faecal samples from domestic pigs at a farm where bovine trichomonosis has not been reported for over 30 years. Vet Parasitol. 2015;212(3-4):105–10.  https://doi.org/10.1016/j.vetpar.2015.08.010.CrossRefPubMedGoogle Scholar
  176. Mukhufhi N, Irons PC, Michel A, Peta F. Evaluation of a PCR test for the diagnosis of Tritrichomonas foetus infection in bulls: effects of sample collection method, storage and transport medium on the test. Theriogenology. 2003;60(7):1269–78.PubMedCrossRefGoogle Scholar
  177. Munoz E, Castella J, Gutierrez JF. In vivo and in vitro sensitivity of Trichomonas gallinae to some nitroimidazole drugs. Vet Parasitol. 1998;78(4):239–46.PubMedCrossRefGoogle Scholar
  178. Murname D. Field and laboratory observations on trichomoniasis of dairy cattle in Victoria. Austr Vet J. 1959;35:80–3.CrossRefGoogle Scholar
  179. Narcisi EM, Sevoian M, Honigberg BM. Pathologic changes in pigeons infected with a virulent Trichomonas gallinae strain (Eiberg). Avian Dis. 1991;35(1):55–61.PubMedCrossRefGoogle Scholar
  180. Neimanis AS, Handeland K, Isomursu M, Agren E, Mattsson R, Hamnes IS, Bergsjo B, Hirvela-Koski V. First report of epizootic trichomoniasis in wild finches (family Fringillidae) in southern Fennoscandia. Avian Dis. 2010;54(1):136–41.  https://doi.org/10.1637/8952-060509-Case.1.CrossRefPubMedGoogle Scholar
  181. Nickel DD, Olson ME, Schultz GA. An improved polymerase chain reaction assay for the detection of Tritrichomonas foetus in cattle. Can Vet J. 2002;43(3):213–6.PubMedPubMedCentralGoogle Scholar
  182. Ondrak JD. Tritrichomonas foetus prevention and control in cattle. Vet Clin North Am Food Anim Pract. 2016;32(2):411–23.  https://doi.org/10.1016/j.cvfa.2016.01.010.CrossRefPubMedGoogle Scholar
  183. Ostrowsky JE, Frene AJ, Rodriguez Dubra C, Rutter B. Obtención de muestras prepuciales para el diagnóstico de Tritrichomonas foetus porraspado de mucosas. Rev Med Vet. 1974;55:525–8.Google Scholar
  184. Pakandl M. The prevalence of intestinal protozoa in wild and domestic pigs. Vet Med (Praha). 1994;39(7):377–80.Google Scholar
  185. Paris JK, Wills S, Balzer HJ, Shaw DJ, Gunn-Moore DA. Enteropathogen co-infection in UK cats with diarrhoea. BMC Vet Res. 2014;10:13.  https://doi.org/10.1186/1746-6148-10-13.CrossRefPubMedPubMedCentralGoogle Scholar
  186. Parker S, Campbell J, Ribble C, Gajadhar A. Comparison of two sampling tools for diagnosis of Tritrichomonas foetus in bulls and clinical interpretation of culture results. J Am Vet Med Assoc. 1999;215(2):231–5.PubMedGoogle Scholar
  187. Parker S, Lun ZR, Gajadhar A. Application of a PCR assay to enhance the detection and identification of Tritrichomonas foetus in cultured preputial samples. J Vet Diagn Invest. 2001;13(6):508–13.PubMedCrossRefGoogle Scholar
  188. Parker S, Campbell J, Gajadhar A. Comparison of the diagnostic sensitivity of a commercially available culture kit and a diagnostic culture test using Diamond’s media for diagnosing Tritrichomonas foetus in bulls. J Vet Diagn Invest. 2003a;15(5):460–5.PubMedCrossRefGoogle Scholar
  189. Parker S, Campbell J, Ribble C, Gajadhar A. Sample collection factors affect the sensitivity of the diagnostic test for Tritrichomonas foetus in bulls. Can J Vet Res. 2003b;67(2):138–41.PubMedPubMedCentralGoogle Scholar
  190. Parsonson IM, Clark BL, Dufty J. The pathogenesis of Tritrichomonas foetus infection in the bull. Aust Vet J. 1974;50(10):421–3.PubMedCrossRefGoogle Scholar
  191. Parsonson IM, Clark BL, Dufty JH. Early pathogenesis and pathology of Tritrichomonas foetus infection in virgin heifers. J Comp Pathol. 1976;86(1):59–66.PubMedCrossRefGoogle Scholar
  192. Pereira-Neves A, Benchimol M. Tritrichomonas foetus: budding from multinucleated pseudocysts. Protist. 2009;160(4):536–51.  https://doi.org/10.1016/j.protis.2009.05.001.CrossRefPubMedGoogle Scholar
  193. Perez E, Conrad PA, Hird D, Ortuno A, Chacon J, Bondurant R, Noordhuizen J. Prevalence and risk-factors for Trichomonas-fetus infection in cattle in northeastern Costa-Rica. Prev Vet Med. 1992;14(3-4):155–65.  https://doi.org/10.1016/0167-5877(92)90013-6.CrossRefGoogle Scholar
  194. Perez A, Cobo E, Martinez A, Campero C, Spath E. Bayesian estimation of Tritrichomonas foetus diagnostic test sensitivity and specificity in range beef bulls. Vet Parasitol. 2006;142(1-2):159–62.  https://doi.org/10.1016/j.vetpar.2006.06.021.CrossRefPubMedGoogle Scholar
  195. Petrin D, Delgaty K, Bhatt R, Garber G. Clinical and microbiological aspects of Trichomonas vaginalis. Clin Microbiol Rev. 1998;11(2):300–17.PubMedPubMedCentralGoogle Scholar
  196. Pierce AE. The demonstration of an agglutinin to Trichomonas foetus in the vaginal discharge of infected heifers. J Comp Pathol Ther. 1947;57(2):84–97.PubMedCrossRefGoogle Scholar
  197. Pierce AE. The mucus agglutination test for the diagnosis of bovine trichomoniasis. Vet Rec. 1949;61:347–9.Google Scholar
  198. Profizi C, Cian A, Meloni D, Hugonnard M, Lambert V, Groud K, Gagnon AC, Viscogliosi E, Zenner L. Prevalence of Tritrichomonas foetus infections in French catteries. Vet Parasitol. 2013;196(1-2):50–5.  https://doi.org/10.1016/j.vetpar.2013.01.021.CrossRefPubMedGoogle Scholar
  199. Queen EV, Marks SL, Farver TB. Prevalence of selected bacterial and parasitic agents in feces from diarrheic and healthy control cats from northern California. J Vet Int Med. 2012;26(1):54–60.  https://doi.org/10.1111/j.1939-1676.2011.00843.x.CrossRefGoogle Scholar
  200. Rae DO. Impact of trichomoniasis on the cow-calf producer’s profitability. J Am Vet Med Assoc. 1989;194(6):771–5.PubMedGoogle Scholar
  201. Rae DO, Crews JE. Tritrichomonas foetus. Vet Clin North Am Food Anim Pract. 2006;22(3):595–611.  https://doi.org/10.1016/j.cvfa.2006.07.001.CrossRefPubMedGoogle Scholar
  202. Rae DO, Chenoweth PJ, Genho PC, McIntosh AD, Crosby CE, Moore SA. Prevalence of Tritrichomonas fetus in a bull population and effect on production in a large cow-calf enterprise. J Am Vet Med Assoc. 1999;214(7):1051–5.PubMedGoogle Scholar
  203. Rae DO, Crews JE, Greiner EC, Donovan GA. Epidemiology of Tritrichomonas foetus in beef bull populations in Florida. Theriogenology. 2004;61(4):605–18.PubMedCrossRefGoogle Scholar
  204. Reece RL, Dennett DP, Johnson RH. Some observations on cultural and transport conditions for Tritrichomonas foetus var. brisbane. Aust Vet J. 1983;60(2):62–3.PubMedCrossRefGoogle Scholar
  205. Reinmann K, Muller N, Kuhnert P, Campero CM, Leitsch D, Hess M, Henning K, Fort M, Muller J, Gottstein B, Frey CF. Tritrichomonas foetus isolates from cats and cattle show minor genetic differences in unrelated loci ITS-2 and EF-1alpha. Vet Parasitol. 2012;185(2-4):138–44.  https://doi.org/10.1016/j.vetpar.2011.09.032.CrossRefPubMedGoogle Scholar
  206. Rhyan JC, Stackhouse LL, Quinn WJ. Fetal and placental lesions in bovine abortion due to Tritrichomonas foetus. Vet Pathol. 1988;25(5):350–5.PubMedCrossRefGoogle Scholar
  207. Rhyan JC, Blanchard PC, Kvasnicka WG, Hall MR, Hanks D. Tissue-invasive Tritrichomonas foetus in four aborted bovine fetuses. J Vet Diagn Invest. 1995a;7(3):409–12.PubMedCrossRefGoogle Scholar
  208. Rhyan JC, Wilson KL, Burgess DE, Stackhouse LL, Quinn WJ. Immunohistochemical detection of Tritrichomonas foetus in formalin-fixed, paraffin-embedded sections of bovine placenta and fetal lung. J Vet Diagn Invest. 1995b;7(1):98–101.PubMedCrossRefGoogle Scholar
  209. Rhyan JC, Wilson KL, Wagner B, Anderson ML, BonDurant RH, Burgess DE, Mutwiri GK, Corbeil LB. Demonstration of Tritrichomonas foetus in the external genitalia and of specific antibodies in preputial secretions of naturally infected bulls. Vet Pathol. 1999;36(5):406–11.PubMedCrossRefGoogle Scholar
  210. Ribeiro KC, Mariante RM, Coutinho LL, Benchimol M. Nucleus behavior during the closed mitosis of Tritrichomonas foetus. Biol Cell. 2002;94(4-5):289–301.PubMedCrossRefGoogle Scholar
  211. Robinson RA, Lawson B, Toms MP, Peck KM, Kirkwood JK, Chantrey J, Clatworthy IR, Evans AD, Hughes LA, Hutchinson OC, John SK, Pennycott TW, Perkins MW, Rowley PS, Simpson VR, Tyler KM, Cunningham AA. Emerging infectious disease leads to rapid population declines of common British birds. PLoS One. 2010;5(8):e12215.  https://doi.org/10.1371/journal.pone.0012215.CrossRefPubMedPubMedCentralGoogle Scholar
  212. Rodning SP, Wolfe DF, Carson RL, Wright JC, Stockdale HD, Pacoli ME, Busby HC, Rowe SE. Prevalence of Tritrichomonas foetus in several subpopulations of Alabama beef bulls. Theriogenology. 2008;69(2):212–7.  https://doi.org/10.1016/j.theriogenology.2007.09.014.
  213. Rogers KH, Girard YA, Woods L, Johnson CK. Avian trichomonosis in spotted owls (Strix occidentalis): Indication of opportunistic spillover from prey. Int J Parasitol Parasites Wildl. 2016;5(3):305–11.  https://doi.org/10.1016/j.ijppaw.2016.10.002.CrossRefPubMedPubMedCentralGoogle Scholar
  214. Rosado TW, Specht A, Marks SL. Neurotoxicosis in 4 cats receiving ronidazole. J Vet Int Med. 2007;21(2):328–31.CrossRefGoogle Scholar
  215. Rosypal AC, Ripley A, Stockdale Walden HD, Blagburn BL, Grant DC, Lindsay DS. Survival of a feline isolate of Tritrichomonas foetus in water, cat urine, cat food and cat litter. Vet Parasitol. 2012;185(2-4):279–81.  https://doi.org/10.1016/j.vetpar.2011.11.003.CrossRefPubMedGoogle Scholar
  216. Sager H, Ferre I, Henning K, Ortega-Mora LM. Tritrichomoniasis. In: Ortega-Mora LM, Gottstein B, Conraths FJ, Buxton D, editors. Protozoal abortion in farm ruminants. Wallingford, Oxfordshire: CAB International; 2007. p. 232–62.Google Scholar
  217. Sansano-Maestre J, Garijo-Toledo MM, Gomez-Munoz MT. Prevalence and genotyping of Trichomonas gallinae in pigeons and birds of prey. Avian Pathol. 2009;38(3):201–7.  https://doi.org/10.1080/03079450902912135.CrossRefPubMedGoogle Scholar
  218. Schönmann MJ, BonDurant RH, Gardner IA, Van Hoosear K, Baltzer W, Kachulis C. Comparison of sampling and culture methods for the diagnosis of Tritrichomonas foetus infection in bulls. Vet Rec. 1994;134(24):620–2.PubMedCrossRefGoogle Scholar
  219. Simmons GC, Laws L. Observations on bovine tritrichomoniasis. Aust Vet J. 1957;33:249–53.CrossRefGoogle Scholar
  220. Simpson V, Molenaar F. Increase in trichomonosis in finches. Vet Rec. 2006;159(18):606.PubMedCrossRefGoogle Scholar
  221. Skirrow S. Identification of trichomonad-carrier cows. J Am Vet Med Assoc. 1987;191(5):553–4.PubMedGoogle Scholar
  222. Skirrow SZ, BonDurant RH. Bovine trichomoniasis. Vet Bull. 1988;58:591–603.Google Scholar
  223. Skirrow SZ, BonDurant RH. Immunoglobulin isotype of specific antibodies in reproductive tract secretions and sera in Tritrichomonas foetus-infected heifers. Am J Vet Res. 1990a;51(4):645–53.PubMedGoogle Scholar
  224. Skirrow SZ, BonDurant RH. Induced Tritrichomonas foetus infection in beef heifers. J Am Vet Med Assoc. 1990b;196(6):885–9.PubMedGoogle Scholar
  225. Skirrow S, BonDurant R, Farley J, Correa J. Efficacy of ipronidazole against trichomoniasis in beef bulls. J Am Vet Med Assoc. 1985;187(4):405–7.PubMedGoogle Scholar
  226. Slapeta J, Craig S, McDonell D, Emery D. Tritrichomonas foetus from domestic cats and cattle are genetically distinct. Exp Parasitol. 2010;126(2):209–13.  https://doi.org/10.1016/j.exppara.2010.04.024.CrossRefPubMedGoogle Scholar
  227. Slapeta J, Müller N, Stack CM, Walker G, Lew-Tabor A, Tachezy J, Frey CF. Comparative analysis of Tritrichomonas foetus (Riedmuller, 1928) cat genotype, T. foetus (Riedmuller, 1928) cattle genotype and Tritrichomonas suis (Davaine, 1875) at 10 DNA loci. Int J Parasitol. 2012;42(13–14):1143–9.  https://doi.org/10.1016/j.ijpara.2012.10.004.
  228. Soto P, Parma AE. The immune response in cattle infected with Tritrichomonas foetus. Vet Parasitol. 1989;33(3-4):343–8.PubMedCrossRefGoogle Scholar
  229. Stabler RM. Trichomonas gallinae: a review. Exp Parasitol. 1954;3(4):368–402.PubMedCrossRefGoogle Scholar
  230. Stauffer SH, Birkenheuer AJ, Levy MG, Marr H, Gookin JL. Evaluation of four DNA extraction methods for the detection of Tritrichomonas foetus in feline stool specimens by polymerase chain reaction. J Vet Diagn Invest. 2008;20(5):639–41.PubMedCrossRefGoogle Scholar
  231. Steiner JM, Xenoulis PG, Read SA, Suchodolski JS, Globokar M, Huisinga E, Thucre S. Identification of Tritrichomonas foetus DNA in feces from cats with diarrhea from Germany and Austria. J Vet Int Med. 2007;21(3):649.Google Scholar
  232. Stockdale HD, Givens MD, Dykstra CC, Blagburn BL. Tritrichomonas foetus infections in surveyed pet cats. Vet Parasitol. 2009;160(1-2):13–7.  https://doi.org/10.1016/j.vetpar.2008.10.091.CrossRefPubMedGoogle Scholar
  233. Strickland L, Edmondson M, Maxwell H, et al. Surface architectural anatomy of the penile and preputial epithelium of bulls. Clin Therio. 2014;6:445–51.Google Scholar
  234. Stuka P, Katai P. Rapid demonstration of bull trichomoniasis in unstained smear preparations from preputial scrapings. Acta Vet Hung. 1969;19:385–9.Google Scholar
  235. Sun Z, Stack C, Slapeta J. Sequence differences in the diagnostic region of the cysteine protease 8 gene of Tritrichomonas foetus parasites of cats and cattle. Vet Parasitol. 2012;186(3-4):445–9.  https://doi.org/10.1016/j.vetpar.2011.12.001.CrossRefPubMedGoogle Scholar
  236. Szonyi B, Srinath I, Schwartz A, Clavijo A, Ivanek R. Spatio-temporal epidemiology of Tritrichomonas foetus infection in Texas bulls based on state-wide diagnostic laboratory data. Vet Parasitol. 2012;186(3-4):450–5.  https://doi.org/10.1016/j.vetpar.2011.11.075.CrossRefPubMedGoogle Scholar
  237. Tachezy J, Tachezy R, Hampl V, Sedinova M, Vanacova S, Vrlik M, Van Ranst M, Flegr J, Kuldaa J. Cattle pathogen Tritrichomonas foetus (Riedmuller, 1928) and pig commensal Tritrichomonas suis (Gruby & Delafond, 1843) belong to the same species. J Eukaryot Microbiol. 2002;49(2):154–63.PubMedCrossRefGoogle Scholar
  238. Tasca T, De Carli GA. Scanning electron microscopy study of Trichomonas gallinae. Vet Parasitol. 2003;118(1-2):37–42.PubMedCrossRefGoogle Scholar
  239. Taylor MA, Marshall RN, Stack M. Morphological differentiation of Tritrichomonas foetus from other protozoa of the bovine reproductive tract. Br Vet J. 1994;150(1):73–80.  https://doi.org/10.1016/S0007-1935(05)80098-3.CrossRefPubMedGoogle Scholar
  240. Tedesco LF, Errico F, Del Baglivi LP. Diagnosis of Tritrichomonas foetus infection in bulls using two sampling methods and a transport medium. Aust Vet J. 1979;55(7):322–4.PubMedCrossRefGoogle Scholar
  241. Terzolo HR, Argento E, Catena MC, Cipolla AL, Martinez AH, Tejada G, Villa C, Betancor R, Campero CM, Cordeviola JM, Pasini MI. Procedimientos de laboratorio para el diagnóstico de la Campylobacteriosis y Tricomoniasis genital bovina. In: Comisión Científica Permanente de Enfermedades Venéreas de los Bovinos. Balcarce, Argentina: INTA; 1992. p. 1–33.Google Scholar
  242. Thomas MW, Harmon WM, White C. An improved method for the detection of Tritrichomonas-fetus Infection by culture in bulls. Agric Pract. 1990;11(1):13–7.Google Scholar
  243. Thomford JW, Talbot JA, Ikeda JS, Corbeil LB. Characterization of extracellular proteinases of Tritrichomonas foetus. J Parasitol. 1996;82(1):112–7.PubMedCrossRefGoogle Scholar
  244. Todorovic R, McNutt SH. Diagnosis of Trichomonas foetus infection in bulls. Am J Vet Res. 1967;28(126):1581–90.PubMedGoogle Scholar
  245. Tolbert MK, Gookin J. Tritrichomonas foetus: a new agent of feline diarrhea. Compend Contin Educ Vet. 2009;31(8):374–81. 390; quiz 381.PubMedGoogle Scholar
  246. Tolbert MK, Gookin JL. Mechanisms of Tritrichomonas foetus pathogenicity in cats with insights from venereal trichomonosis. J Vet Int Med. 2016;30(2):516–26.  https://doi.org/10.1111/jvim.13920.CrossRefGoogle Scholar
  247. Tolbert MK, Stauffer SH, Gookin JL. Feline Tritrichomonas foetus adhere to intestinal epithelium by receptor-ligand-dependent mechanisms. Vet Parasitol. 2013;192(1-3):75–82.  https://doi.org/10.1016/j.vetpar.2012.10.019.CrossRefPubMedGoogle Scholar
  248. Tysnes K, Gjerde B, Nodtvedt A, Skancke E. A cross-sectional study of Tritrichomonas foetus infection among healthy cats at shows in Norway. Acta Vet Scand. 2011;53:39.  https://doi.org/10.1186/1751-0147-53-39.CrossRefPubMedPubMedCentralGoogle Scholar
  249. Van der Pol B. Trichomonas vaginalis infection: the most prevalent nonviral sexually transmitted infection receives the least public health attention. Clin Infect Dis. 2007;44(1):23–5.  https://doi.org/10.1086/509934.CrossRefPubMedGoogle Scholar
  250. Van der Saag M, McDonell D, Slapeta J. Cat genotype Tritrichomonas foetus survives passage through the alimentary tract of two common slug species. Vet Parasitol. 2011;177(3-4):262–6.  https://doi.org/10.1016/j.vetpar.2010.11.054.CrossRefPubMedGoogle Scholar
  251. Villarroel A, Carpenter TE, BonDurant RH. Development of a simulation model to evaluate the effect of vaccination against Tritrichomonas foetus on reproductive efficiency in beef herds. Am J Vet Res. 2004;65(6):770–5.PubMedCrossRefGoogle Scholar
  252. Voyich JM, Ansotegui R, Swenson C, Bailey J, Burgess DE. Antibody responses of cattle immunized with the Tf190 adhesin of Tritrichomonas foetus. Clin Diagn Lab Immunol. 2001;8(6):1120–5.  https://doi.org/10.1128/CDLI.8.6.1120-1125.2001.CrossRefPubMedPubMedCentralGoogle Scholar
  253. Walden HS, Dykstra C, Dillon A, Rodning S, Givens D, Bird R, Newton J, Lindsay D. A new species of Tritrichomonas (Sarcomastigophora: Trichomonida) from the domestic cat (Felis catus). Parasitol Res. 2013;112(6):2227–35.  https://doi.org/10.1007/s00436-013-3381-8.CrossRefPubMedGoogle Scholar
  254. Witte J. Bakterienfreie Züchtung von Trichomonaden und dem Uterus des Rindes in einfachen Nährböden. Zentralblatt für Bakteriologie, Parasitenkunde, Infektionskrankheiten und Hygiene. 1933;128:188–95.Google Scholar
  255. Xenoulis PG, Lopinski DJ, Read SA, Suchodolski JS, Steiner JM. Intestinal Tritrichomonas foetus infection in cats: a retrospective study of 104 cases. J Feline Med Surg. 2013;15(12):1098–103.  https://doi.org/10.1177/1098612X13495024.CrossRefPubMedGoogle Scholar
  256. Yaeger MJ, Gookin JL. Histologic features associated with Tritrichomonas foetus-induced colitis in domestic cats. Vet Pathol. 2005;42(6):797–804.  https://doi.org/10.1354/vp.42-6-797.CrossRefPubMedGoogle Scholar
  257. Yang N, Cui X, Qian W, Yu S, Liu Q. Survey of nine abortifacient infectious agents in aborted bovine fetuses from dairy farms in Beijing, China, by PCR. Acta Vet Hung. 2012;60(1):83–92.  https://doi.org/10.1556/AVet.2012.007.CrossRefPubMedGoogle Scholar
  258. Yao C. Diagnosis of Tritrichomonas foetus-infected bulls, an ultimate approach to eradicate bovine trichomoniasis in US cattle? J Med Microbiol. 2013;62(Pt 1):1–9.  https://doi.org/10.1099/jmm.0.047365-0.CrossRefPubMedGoogle Scholar
  259. Yao C, Köster LS. Tritrichomonas foetus infection, a cause of chronic diarrhea in the domestic cat. Vet Res. 2015;46:35.  https://doi.org/10.1186/s13567-015-0169-0.CrossRefPubMedPubMedCentralGoogle Scholar
  260. Yule A, Skirrow SZ, Bondurant RH. Bovine trichomoniasis. Parasitol Today. 1989a;5(12):373–7.PubMedCrossRefGoogle Scholar
  261. Yule A, Skirrow SZ, Staats J, Bondurant RH. Development and preliminary assessment of a polyclonal antibody-based enzyme immunoassay for the detection of Tritrichomonas foetus antigen in breeding cattle. Vet Parasitol. 1989b;31(2):115–23.PubMedCrossRefGoogle Scholar

Copyright information

© Springer International Publishing AG, part of Springer Nature 2018

Authors and Affiliations

  • Esther Collántes-Fernández
    • 1
  • Marcelo C. Fort
    • 2
  • Luis M. Ortega-Mora
    • 1
  • Gereon Schares
    • 3
    Email author
  1. 1.Faculty of Veterinary SciencesUniversity Complutense of MadridMadridSpain
  2. 2.Veterinary Public Health GroupNational Institute of Agricultural Technology (INTA)AnguilArgentina
  3. 3.Friedrich-Loeffler-Institut, Federal Research Institute for Animal Health, Institute of EpidemiologyGreifswald-Insel RiemsGermany

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