GUIDELINE for Toxoplasma gondii infection

The extra-intestinal life cycle

The extra-intestinal development of T. gondii is the same for all hosts, including cats, dogs, and people, whether tissue cysts or oocysts have been ingested. After the ingestion of oocysts, sporozoites hatch in the lumen of the small intestine and enter intestinal cells, including those in the lamina propria. Sporozoites divide into two by an asexual process known as endodyogeny, thereby becoming tachyzoites. These are lunate (falciform) in shape, approximately 6 x 2 μm, and multiply in almost any cell of the body. When the cell ruptures, releasing the tachyzoites, these infect new cells. Otherwise, tachyzoites multiply intracellularly for an undetermined period, and eventually encyst. Tissue cysts vary in size from 15 to 60 μm and usually conform to the shape of the parasitized cell. Tissue cysts are formed mainly in the CNS, muscles, and visceral organs, and probably persist for the life of the host. They can be reactivated after immunosuppression, which can then lead to clinical signs (Last et al., 2004; Dubey, 2005; Barrs et al., 2006; Dubey and Lappin, 2006; Lo Piccolo et al., 2019; Moore et al., 2022).

T. gondii affects the neurological system of the host

T. gondii blocks the innate aversion of rats for cat urine, instead making them attracted by the feline pheromone (House et al., 2011), which could increase the likelihood of a cat capturing an infected rat. This reflects adaptive, “behavioural manipulation” by T. gondii in optimizing the chances for completing the parasite’s life cycle: it reproduces sexually only in the feline intestine. T. gondii also alters olfactory preferences in humans; infected men rate cat urine (but not tiger urine) as pleasant while uninfected men do not, but T. gondii infected women found it less pleasant (Flegr et al., 2011).

The behavioural manipulation hypothesis postulates that a parasite will specifically manipulate host conduct essential for its transmission. However, the neural circuits for innate fear, anxiety, and acquired fright all overlap, raising the possibility that T. gondii could disrupt all of these non-specifically (Vyas et al., 2007).

Other evidence supporting the behavioural manipulation hypothesis includes that T. gondii–infected chimpanzees (Pan troglodytes troglodytes) lose their innate aversion towards the urine of leopards (Panthera pardus), their only natural predators. In contrast, no clear difference was found in the response of infected and uninfected animals towards urine collected from other definitive feline hosts that chimpanzees do not encounter in nature (Poirotte et al., 2016).

Immunity

Immunity to T. gondii in the cat is poorly understood. In the mouse and in humans, it is highly dependent on cell-mediated effector responses (Sanchez et al., 2010).

All infected cats develop IgG and IgA antibodies, about 80% also have IgM antibodies. IgG antibodies can take four to six weeks to appear, and maximal antibody titres are achieved within two to three weeks after first appearance (Dubey and Lappin, 2006).

In an experiment on 13 laboratory cats, following challenge with different strains of T. gondii immune levels which prevented re-excretion of T. gondii oocysts waned progressively from 1 to 3 years (Zulpo et al., 2018). Protection against oocyst re-excretion was present in 90%, 25%, and 33.4% of cats after 12, 24, and 36 months from the initial infection, respectively (Zulpo et al., 2018).

Clinical signs

Most T. gondii infections of cats are subclinical (Jokelainen et al., 2012; Dubey and Prowell, 2013), but when clinical signs do occur, they tend to be serious and life endangering. It is difficult to ascertain the prevalence of clinical toxoplasmosis, because the diagnosis can be missed in vivo (Cohen et al., 2016).  In a Finnish study, 3.1% of 193 cats undergoing post-mortem examination had toxoplasmosis: all six cats had an acute history of illness lasting approximately one week and they were young cats, up to two years of age (Jokelainen et al., 2012). However, cats of any age can develop toxoplasmosis.

Cats are generally asymptomatic while shedding toxoplasma oocysts (Dubey and Prowell, 2013), although diarrhoea has been  reported in some cases (Dubey and Prowell, 2013; Deksne et al., 2013) and experimentally infected cats (Lin et al., 1992). T. gondii oocysts were detected by PCR in 1% of 1,088 and 1.2% of 289 faecal samples from diarrhoeic cats (Paris et al., 2014; Paul and Stayt, 2019). Lymphocytic-plasmacytic enteritis was due to toxoplasmosis in two cats (Peterson et al., 1991). Toxoplasmosis was suspected to cause eosinophilic fibrosing gastritis in one case report (McConnell et al., 2007). Diarrhoea was reported in 4 cats with clinical toxoplasmosis (Bastan and Bas, 2018).

Clinical signs develop due to inflammation and tissue necrosis caused by dissemination and intracellular replication of tachyzoites (Dubey and Lappin, 2006). It usually originates from reactivation of a latent infection rather than after a newly acquired infection. If an infected cat is immunosuppressed, bradyzoites in tissue cysts replicate rapidly and disseminate again as tachyzoites.

Clinical toxoplasmosis has been documented in cats infected with FIV or FeLV (Davidson et al., 1993; Pena et al., 2017). Commonly used doses of glucocorticoids do not predispose to reactivation, but high doses (e.g. 10 to 80 mg/kg/day prednisolone) can reactivate latent infection with potentially fatal consequences (Lappin et al., 1992; Barrs et al., 2006; Lo Piccolo et al., 2019). Administration of cyclosporin to cats with renal transplants, immune-mediated disease or dermatologic disease has also been associated with clinical manifestations (Beatty and Barrs, 2003; Last et al., 2004; Barrs et al., 2006; Evans et al., 2017; Lo Piccolo et al., 2019; Salant et al., 2021). Immunosuppression by oclacitinib to suppress atopic skin disease led to fatal toxoplasmosis (Moore et al., 2022).   Immunosuppression by chemotherapy has also led to systemic toxoplasmosis (Murakami et al., 2018).

Although any organ can be involved, pneumonitis is the most common finding and it can be rapidly fatal (Jokelainen et al., 2012; Dubey et al., 2020;). At post-mortem examination lungs have marked pulmonary oedema (Jokelainen et al., 2012). One case presented with a pulmonary mass which was successfully treated (McKenna et al., 2021). One cat diagnosed by computed tomography as suffering from idiopathic pulmonary fibrosis recovered after clindamycin treatment was instigated in response to a high antibody titre (Stavri et al., 2021).

Following lungs (Fig. 4), the tissues most commonly affected by T. gondii are the liver (Jokelainen et al., 2012), CNS (Bastan and Bas, 2018), eyes (Cucoş et al., 2015; Bastan and Bas, 2018) (Fig. 5), mesenteric lymph nodes (Jokelainen et al., 2012; Cohen et al., 2016) and muscles (Fig. 6).

Cats with toxoplasmosis present with fever, depression, anorexia, neurologic signs (Jokelainen et al., 2012; Cucoş et al., 2015; Mari et al., 2016; Klang et al., 2018) (e.g., behavioural change, seizures, ataxia, polyneuropathy), uveitis (Cucoş et al., 2015; Jinks et al., 2016) (Fig. 5), icterus, lameness, muscle hyperaesthesia, myopathy, dyspnoea, diarrhoea, and weight loss (Jokelainen et al., 2012; Bastan and Bas, 2018; Butts and Langley-Hobbs, 2020; Güven and Ceylan, 2020), Cholecystitis (Lo Piccolo et al., 2019), neuropathy of a single cranial nerve (Wagner and Cooper, 2018) and spinal granuloma (Tyroller et al., 2023) have each been described in single case reports. Cutaneous ulcers and hyperaemic nodules have also been reported as clinical signs of toxoplasmosis (Anfray et al., 2005; Kul et al., 2011).

Toxoplasma antibodies were found in 60 (57%) of 105 cats with clinical signs of intra-ocular disease: 53 of the 60 cats responded completely or partially to clindamycin and topical tobramycin dexamethasone (Ali et al., 2021).

In a series of 22 cats with ocular toxoplasmosis, anterior and posterior uveitis were the most common ophthalmological signs; lens luxation, absent pupillary light reflex (PLR) and chorioretinitis were also encountered (Cucoş et al., 2015).

Toxoplasmosis presented as acute myocarditis in three cats: clinical, electrocardiographic, radiographic and echocardiographic abnormalities resolved following clindamycin treatment  (Simpson et al., 2005; Romito et al., 2022) and elevated troponin levels returned to within normal limits (Romito et al., 2022).

Although antibody positivity to T. gondii was associated with chronic kidney disease in humans (Babekir et al., 2022), no such association was made in a single study in cats (Hsu et al., 2011).

Congenital infection tends to be more serious than infection of the adult cat (Dubey and Lappin, 2006). Transplacentally or lactogenically infected kittens develop more severe signs and frequently die of pulmonary or hepatic disease (Dubey et al., 1995; Atmaca et al., 2013). Toxoplasmosis was identified as the cause of death of two 8 week old kittens which faded and died (Crouch et al., 2019).

Immune complex formation (Lappin et al., 1993) and deposition in tissues as well as delayed hypersensitivity reactions might be involved in the development of chronic forms of toxoplasmosis. Since T. gondii is not cleared from the body, neither naturally nor through treatment, toxoplasmosis can recur.

Fig. 4. Thoracic radiographs (latero-lateral view) of a cat with pulmonary toxoplasmosis (courtesy of Katrin Hartmann, Medizinische Kleintierklinik, LMU Munich, Germany).

Fig. 5. This cat had toxoplasmosis uveitis of his left eye four years prior to this photograph, treated successfully with clindamycin. However, anisocoria and iris discoloration remained, and some months following this photograph his lens luxated, necessitating enucleation. The cat survived 10 years after his toxoplasmosis (Photograph courtesy D. Addie, www.catvirus.com).

Fig. 6. Cat with toxoplasmosis suffering from myositis caused by T. gondii cysts. The cat presented in lateral recumbency, was unable to get up, and showed severe muscle hyperaesthesia (courtesy of Katrin Hartmann, Medizinische Kleintierklinik, LMU Munich, Germany).

Diagnosis

Definitive diagnosis of clinical toxoplasmosis is only confirmed when the organism is found in body fluids or tissue, but in practice many cases are diagnosed “empirically” by correlating presence of antibodies, clinical signs, and response to clindamycin therapy (Lappin et al., 1989). Around one third of cases have concurrent immunosuppressive conditions (Dubey et al., 2020).

Toxoplasma lesions in the lungs can appear like tumours/masses on imaging (Murakami et al., 2018), therefore, if safe to do so, it is important to take a fine needle aspirate of the lesions to confirm diagnosis.

During acute illness, tachyzoites can be detected in tissues and body fluids by cytology, PCR or bioassay in mice or cell culture (Dubey et al., 2020). Tachyzoites are rarely found in blood, but occasionally in CSF, aqueous humour (Powell et al., 2010), fine-needle aspirates of organs (e.g., lymph nodes), bile, and transtracheal or bronchoalveolar washings, and are common in the peritoneal and thoracic fluids of animals developing thoracic effusions or ascites.

While the gold standard for diagnosis is histopathology, if suitable samples cannot be taken for histopathology, then detection of tachyzoites by PCR, isolation by bioassay or cell culture confirms the diagnosis. A tentative diagnosis is sometimes based on rising IgG or IgM antibody titres (see antibody testing section below), exclusion of other causes for the clinical signs, and a favourable clinical response to anti-T. gondii drugs (Lappin et al., 1989; Dubey, 2005; Dubey and Lappin, 2006).

Laboratory changes

Little is published about biochemical and haematological findings in clinical toxoplasmosis.  The main abnormalities detected on blood biochemistry were elevated liver enzyme alanine aminotransferase (Lappin et al., 1989; Jokelainen et al., 2012; Bastan and Bas, 2018; Güven and Ceylan, 2020). Cats had hypoalbuminaemia in two reports (Bastan and Bas, 2018; Zandonà et al., 2018) while albumin was raised in another study (Lappin et al., 1989). In cases of myositis, one would expect creatine kinase to be raised.

Following experimental infection, neutrophil and lymphocyte counts reduced transiently (Lappin et al., 1996).

Detection of the infectious agent

Direct detection

Detection of tachyzoites

Ante-mortem diagnosis of clinical toxoplasmosis is ideally based on the detection of tachyzoites by cytology, immunohistochemistry (Jokelainen et al., 2012; Künzel et al., 2017; Zandonà et al., 2018), PCR, or cell culture. Detection of tachyzoites results in a definitive diagnosis: tachyzoites can be detected in various tissues and body fluids during manifestation of illness (Fig. 7). They are rarely found in blood, but occasionally in CSF or aqueous humour, fine-needle aspirates of organs (e.g. lymph nodes), and transtracheal or bronchoalveolar washings. Alternatively, a PCR can be performed using CSF, aqueous humour (Powell et al., 2010), bile (Lo Piccolo et al., 2019) or bronchoalveolar lavage fluid.

Fig. 7. Cytology of a fine needle aspirate of a cat with pulmonary toxoplasmosis and lung consolidation with numerous intracellular and extracellular T. gondii tachyzoites and cysts (arrows). (Courtesy of George Reppas, Vetnostics, Australia).

Detection of oocysts in faeces

Oocyst shedding can be detected by microscopy of faecal samples, with confirmation by PCR testing (Herrmann et al., 2010) and even genotyping.

Flotation methods for oocyst detection have the advantage of being rapid and able to be performed within the veterinary clinic, but the disadvantages of being relatively insensitive, requiring over 1000 oocysts per gram of faeces for detection (Györke et al., 2011). Also the technique is non-specific: T. gondii oocysts are morphologically indistinguishable from those of Hammondia hammondi, Besnoitia orcytofelisi,  Besnoitia darling and Neospora species (Dubey and Lappin, 2006; Monteiro et al., 2008; Herrmann et al., 2010; Lucio-Forster and Bowman, 2011; Ismael and Al-Kafri, 2023). In one study, oocysts were detected by flotation in 105 of 18,259 feline faecal samples, but only 46 of them were confirmed to be T. gondii oocysts (34 were H. hammondi oocysts) (Herrmann et al., 2010), but exact species can be differentiated by PCR testing.

Flotation solutions can be bought from veterinary suppliers or made in house by adding a measured amount of a salt or sugar to a specific amount of water to produce a solution with the desired specific gravity (SG) (Dryden et al., 2005). The standard flotation method for detecting T. gondii oocysts uses 262 mg/ml zinc chloride (ZnCl2) and 275 mg/ml sodium chloride (NaCl) (Herrmann et al., 2010; Mancianti et al., 2015), i.e. 262 grams of ZnCl2 and 275 grams NaCl dissolved in one litre of warm distilled or tap water. Zinc sulphate (371g ZnSO4 in 1,000 ml water (Pouillevet et al., 2017), saturated salt (350g NaCl in one litre of water, SG 1.18-1.20; Dryden et al., 2005) and modified Sheather’s sugar solutions have also been used (454 g granulated sugar, 355 ml tap water with a specific gravity of 1.27 g/ml). To prevent the growth of mould approximately 2-6 ml of 37% formaldehyde can be added. It always should be checked that the correct SG is achieved using a refractometer (hydrometer) which has a range of 1.000–1.400. The SG has to be optimised in order to allow parasite eggs to float and faecal debris to sink.

For PCR purified oocysts then the sugar solution is preferable to a zinc solution because the latter can interfere with the PCR (Berger-Schoch et al., 2011).

A small amount of faeces (2-5g) is mixed with 10 mls of the flotation solution in a test tube or centrifuge tube, and flotation solution is added until the tube is nearly full. At this stage, the sample could (optionally) be centrifuged at 1,200 rpm (280 ×g) for 5 minutes. The preparation is then allowed to stand for at least 10 minutes (up to 24 hours: if left too long the eggs will distort) until the eggs float to the top (Dryden et al., 2005; Schares et al., 2005). A sample is removed from the top to a McMaster or microscope slide using a tool such as a wire loop, straw, needle hub, or glass rod and transferred onto a slide with a coverslip. It is then examined by light microscopy using a magnification of at least 100 x (100-400x). T. gondii oocysts have a diameter of about 9–15μm (Schares et al., 2008; Herrmann et al., 2010; Nabi et al., 2018) (Fig. 8).

A cesium chloride method for purification of T. gondii oocysts from faeces of infected cats has also been described (Staggs et al., 2009).

Fig. 8. T. gondii oocysts in faecal flotation. (Source http://dpd.cdc.gov/dpdx/HTML/ImageLibrary/Toxoplasmosis_il.htm; US Center for Disease Control and Prevention).

PCR

PCR of faeces is useful for differentiating T. gondii oocysts from those of other protozoan species which appear similar (Herrmann et al., 2010). However, caution must be used when interpreting T. gondii  quantitative PCR results for samples of cat faeces: researchers fed a T. gondii infected mouse to an antibody positive cat and T. gondii DNA was detected for three days in the faeces following the mouse meal (albeit at high C_T, indicating a low level of DNA) (Poulle et al., 2016). Therefore, to use PCR to detect T. gondii oocyst shedding one must be certain that the cat did not eat an infected prey animal in at least the three days prior to the test.Faecal samples should be sent to the veterinary laboratory in a plain faecal pot and there is no need to send them on ice. For PCR purified oocysts then the sugar solution described above is preferable to a zinc solution because the latter can interfere with the PCR (Berger-Schoch et al., 2011).

Isolation and culture

Isolation of living organisms is not likely to be available to many practicing veterinary surgeons. Living viable T. gondii can be demonstrated in bioassays (usually using mice) or in Vero cell cultures. T. gondii was isolated from 15 of 56 brain and 10 of 15 muscle homogenates of stray cats from Lisbon in Vero cell cultures (Waap et al., 2012).

Indirect detection

Antibody tests to diagnose a sick cat

Anti-T. gondii antibodies are commonly found in both healthy (Table 1) and sick cats: in the USA, 31.6% of 12,628 clinically ill cats were seropositive for toxoplasma antibodies (Vollaire et al., 2005). Thus, their presence does not prove clinical toxoplasmosis, but does indicate that toxoplasmosis should be on the list of differential diagnoses in a cat who is sick with clinical signs suggestive of toxoplasmosis (Lappin et al., 1989). Antibodies of the IgM class indicate recent infection and can be occasionally detected in healthy cats, but again are not diagnostic proof of clinical toxoplasmosis. However, very high or rising T gondii-specific IgG or IgM titres raises the suspicion of clinical toxoplasmosis.

Antibody tests to assess risk to pregnant or immunosuppressed humans

For assessing human health risks, antibody test results from cats are useful. An antibody-negative cat could be shedding oocysts (early after infection, before antibodies have developed) or will likely shed oocysts if exposed for the first time.

An antibody-positive cat is extremely unlikely to shed oocysts because antibodies need two to three weeks to develop, and by that time, the initial infection has already been controlled and the cat has stopped shedding. In addition, oocyst shedding usually occurs only once in the cat’s lifetime. An antibody-positive cat is also unlikely to shed oocysts when re-exposed or immunosuppressed (Dubey, 2005; Dubey et al., 2020). In one study, cats inoculated with T gondii tissue cysts were orally re-challenged 77 months later, and a few of them did shed oocysts after this second challenge—although only low amounts and over a short time (Dubey, 1995). However, oocyst re-shedding has never been shown to occur in naturally infected cats and the very low oocyst shedding prevalence studies cited in Table 1 support a conclusion that the risk of shedding by an antibody-positive cat is extremely low.

Antibody tests available

A variety of toxoplasma antibody tests exist, including a commercial agglutination test (Toxo-Screen DA, bioMerieux, Marcy-l Etoile, France) referred to as modified or direct agglutination test: MAT (Györke et al., 2011) or DAT (Waap et al., 2012), respectively. Various laboratories have developed their own enzyme-linked immunosorbent assays  (ELISA), and there are commercial ELISA tests: ID Screen Toxoplasmosis Indirect Multi-species (ID.VET Innovative Diagnostics, France), Toxoplasma IgM, IgG (MyBioSource, Inc. USA) (Ali et al., 2021), and the point of care solid phase dot ELISA (Immunocomb, Biogal-Galed Laboratories, Israel) (Györke et al., 2011).

A commercially-available point of care rapid immunomigration  test  (FASTest TOXOPLASMAg,  MEGACOR Diagnostik, Hörbranz, Austria) showed 98.63% sensitivity, 100% specificity, and 99.32% accuracy when evaluated against a bank of samples from  different groups of cats, including 121 healthy seronegative cats, 146 seropositive cats with variable anti-Toxoplasma antibodies, and 25 cats positive for antibodies against other pathogens (Villanueva-Saz et al., 2023).

Indirect immunofluorescence tests (IFAT) can sometimes differentiate antibodies of the IgM, IgG (Sroka et al., 2018) and IgA isotypes.

One comparison of six T. gondii antibody tests is available, published by Györke et al. (2011) and concluded that the commercial ELISA ID.Vet was the best test available.

We recommend using a test which reports an antibody titre as opposed to reporting just  positive or negative.

Treatment

Clindamycin is the treatment of choice (Davidson, 2000; Cucoş et al., 2015) and should be administered at 10–12.5 mg/kg orally q12 h for four weeks (Table 2). Cats with systemic disease and uveitis should be treated with clindamycin in combination with topical glucocorticoids, to avoid secondary glaucoma and lens luxation (Lappin et al., 1989). Prednisolone acetate (1% solution) applied topically to the eye three to four times daily is generally sufficient.

Table 2. Treatment of Toxoplasmosis

Clinical signs not involving the eyes or the CNS usually begin to resolve within the first two to three days of clindamycin administration. CNS and ocular toxoplasmosis tend to respond more slowly. In cases of pulmonary toxoplasmosis, radiographic abnormalities might not resolve for several weeks. Prognosis is usually poor in pulmonary or hepatic disease, particularly in immunocompromised animals (Dubey et al., 2009).

Clindamycin inhibits oocyst shedding (Malmasi et al., 2009).

Vaccination

No toxoplasma vaccine exists commercially, although experimental vaccines have succeeded in reducing or preventing oocyst excretion (Zulpo et al., 2017; Ramakrishnan et al., 2019; Dubey et al., 2020), but not systemic infection (Ramakrishnan et al., 2019).

Prevention

Prevention of infection

Preventing toxoplasmosis in cats involves measures intended to reduce the incidence of infections and the shedding of oocysts into the environment. In order to avoid T. gondii infection, cats should preferably be fed commercially available, processed food, since heating destroys the parasite. The prevalence of feline T. gondii infection is usually higher in countries where cats are fed raw meat, although a study in Greece found no difference in antibody prevalence between cats which were fed raw meat and those which were not: the lack of difference was attributed to the habit of storing meat in the freezer in Greece (Sioutas et al., 2022). Freezing or irradiation can kill tissue cysts without affecting meat quality. If meat is fed to cats, it should be thoroughly cooked at temperatures of at least 64^oC (Rani and Pradhan, 2021), even if previously frozen.

Pets should be prevented from hunting and eating intermediate hosts (rodents) or mechanical transport hosts, such as cockroaches (Wallace, 1972) and earthworms. Cats should be prevented from entering buildings where food-producing animals are housed or where feed storage areas are located (Dubey, 2005).

Litter tray hygiene is important for preventing oocyst transmission to cats and humans especially in multicat environments: cat litter should be changed daily since sporulation takes 2-3 days (Dubey et al., 2011). Viable oocysts have been detected in used cat litter up to 14 days (Dubey et al., 2011). Unfortunately, so far, no cat litter has been found which inactivates toxoplasma oocysts (Dubey et al., 2011).

Prevention of toxoplasmosis in subclinically infected cats

The antibody status of cats to T. gondii should be determined prior to and during immunosuppressive therapy (Last et al., 2004; Barrs et al., 2006). If a cat has anti-T. gondii antibodies, there is a risk of re-activating dormant cysts and causing iatrogenic clinical toxoplasmosis, thus caution should be used prior to immunosuppressive treatments (Barrs et al., 2006). If the cat is antibody-negative, then avoidance of infection is important. If the cat has toxoplasma antibodies, his or her guardian should be apprised of clinical signs of toxoplasmosis to be on the watch for, and preventative clindamycin might be considered.

Zoonotic risk

T. gondii infection of humans is important because of risks to the unborn foetus, for immunocompromised persons (e.g. on chemotherapy), and because research links psychological and cognitive disorders (e.g. lower guilt proneness (Flegr and Havlícek, 1999); reduced IQ (Flegr et al., 2003); schizophrenia (Guimarães et al., 2022) and obsessive-compulsive disorder (Flegr and Horáček, 2017) to T. gondii infection. The life-long presence of Toxoplasma cysts in neural and muscular tissues, leads to prolongation of reaction times: latently infected people are at higher risk for having a road traffic accident (Havlicek et al., 2001; Flegr et al., 2002). Antibody prevalence in human beings is relatively high.

In the human medical literature toxoplasmosis is classified as a “major foodborne pathogen” (López Ureña et al., 2022) rather than a zoonotic infection. Consumption of contaminated food, especially undercooked or raw meat contaminated with T. gondii tissue cysts, is likely to be the major source of T. gondii infection for humans (Kapperud et al., 1996; Cook et al., 2000; Tekay and Özbek, 2007; Belluco et al., 2016; Pinto-Ferreira et al., 2019) (Box 1). People who knew to use separate chopping boards for raw and cooked food were significantly less likely to have been infected with T. gondii (Yan-Li et al., 2017). Exposure from oocyst-contaminated soil (Staggs et al., 2009; Egorov et al., 2018) or water has been reported. Water-borne outbreaks of toxoplasmosis have been reported worldwide and support the theory that exposure to environmental oocysts poses a health risk (Staggs et al., 2009).

A survey amongst obstetrician-gynaecologists in the USA to determine their knowledge and practices about toxoplasmosis prevention and testing found that most overestimated the risk of cat ownership vs environmental risk factors (Jones et al., 2010). A systematic review of risk factors for pregnant women is available (Cook et al., 2000); it stated, “Contact with cats was not a risk factor.”

In addition, studies of 269 (Minbaeva et al., 2013) and 673 (Jung et al., 2017) people with frequent contact with cats found that their antibody prevalence for toxoplasma was actually lower than in a “low-risk” groups who did not have frequent contact with cats.

Box 1: Sources of infection for humans

Veterinarians commonly get questions from immunocompromised or pregnant clients whether or not to get rid of their cat. If hygiene recommendations are followed (Box 2, 3), the risk of transmission is low (Box 4).

Box 2: Recommendations to reduce the risk of parasite transmission from cat to human

Box 3: Additional advice for households with immunocompromised persons or pregnant women

Box 4: Contact with cats is not a risk factor for T. gondii infection (Cook et al., 2000; Elmore et al., 2010; Minbaeva et al., 2013; Jung et al., 2017).

Acknowledgement

ABCD Europe gratefully acknowledges the support of Boehringer Ingelheim (the founding sponsor of the ABCD), Virbac and MSD Animal Health.

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