GUIDELINE for Toxoplasma gondii infection

Published: 01/01/2013
Last updated: 08/02/2024
Last reviewed: 11/06/2024

The Toxoplasma gondii infection in cats guidelines was first published by Hartmann et al. in J Feline Med Surg 2013; 15: 631-637. This update was authored by D. Addie and ABCD colleagues.

Key points

  • The cat is essential to the Toxoplasma gondii life cycle because Felidae are the only hosts in which T.  gondii can reproduce sexually.
  • Oocyst shedding normally happens only once in a cat’s lifetime, usually only from days four to eleven after ingestion of tissue cysts.
  • Cats mainly become infected by eating infected prey mammals or birds, or by being fed raw meat, and less commonly by ingesting oocyst-contaminated food or water. Transplacental and lactogenic transmission occur.
  • Oocysts are able to survive for a long time (over 100 days) in the environment, in soil and water.
  • Antibody prevalence studies show that around one third of cats becomes infected with T. gondii during their lifetime, yet few cats appear to develop clinical toxoplasmosis,
  • Toxoplasmosis should be suspected in any cat with uveitis, neurological signs, raised mesenteric lymph nodes, pneumonia / dyspnoea, muscular pain, lameness, myopathy.
  • A high, or rising, antibody titre and/or detection of IgM antibodies raises the suspicion of clinical signs being due to toxoplasmosis, and justifies treatment with clindamycin.
  • Definitive diagnosis of clinical toxoplasmosis is by detection of tachyzoites by cytology, immunohistochemistry, PCR, or cell culture.
  • T. gondii antibody positive cats no longer shed oocysts and neither are, nor will become, a risk for humans.
  • Toxoplasmosis is classified as a “major food borne pathogen” because the major source of infection for humans is eating meat containing tissue cysts or unwashed raw vegetables.
  • Oocysts are not infectious until they have sporulated 2 days after excretion in faeces, therefore cat faeces should always be removed daily and especially during pregnancy of the cat’s guardian.
  • There is no vaccine.

Agent properties

Toxoplasma (T.) gondii is an obligate intracellular coccidian parasite that can infect virtually all species of warm-blooded animals, including people, and birds. Domestic cats and wild felids (Matas Méndez et al., 2023) are the natural hosts – non-feline species serve only as intermediates (Dubey, 2005; Dubey and Lappin, 2006). T. gondii distinguishes the cat from other species (e.g. mouse) — and thus “knows” when to produce oocysts — by recognizing a single molecule, the fatty acid linoleic acid (Martorelli Di Genova et al., 2019). Felines are the only mammals that lack delta-6-desaturase activity in their intestines, which is required for linoleic acid metabolism, resulting in systemic excess of linoleic acid (Martorelli Di Genova et al., 2019).

Three infectious structures can be distinguished: sporozoites in oocysts, tachyzoites (the actively multiplying stage), and bradyzoites (the slowly multiplying stage) enclosed in tissue cysts.  In an experimental infection, oocysts were excreted in faeces between days 4 and 11 post-infection (Lin et al., 1992). Tachyzoites and bradyzoites are found in tissues and milk (Dubey, 2005; Dubey and Lappin, 2006). For the majority of its life cycle, T. gondii is haploid and reproduces asexually, the tachyzoite replicates rapidly, disseminates throughout the host, and is responsible for disease (English and Striepen, 2019).

Fig. 1. Toxoplasma life cycle. Cats and humans can be infected by transplacental or lactogenic transmission, or by ingestion of tachyzoites or bradyzoites in tissue (i.e. meat) cysts or oocysts. ©ABCD, Karin de Lange

Epidemiology

Prevalence

Antibody prevalence to T. gondii in cats varies geographically and with the age and lifestyle of the cats: the seroprevalence of T. gondii in cats in Thailand is low (4.8%): in contrast, cats living among animals farmed for meat tend to have a high seroprevalence (Dubey et al., 2020; also see the table below). Cats fed raw meat are more likely to be antibody-positive (Jokelainen et al., 2012).

Table 1. The prevalence of T. gondii in various European countries

Country What detected (method) Number of Cats Type of cats Prevalence % Reference
Belgium Antibodies

(IFA and ELISA)

567 healthy house cats 3m -7yrs old 25 De Craeye et al., 2008
Cyprus Antibodies

(ELISA)

123

32

pet

shelter or feral

37.1

40.6

Attipa et al., 2021
Finland Antibodies (MAT)* 445

45

445 purebred

45 shelter

49.7

35.6

Jokelainen et al., 2012
Finland DNA in faeces (PCR) 131 shelter 0.76 Jokelainen et al., 2012
Finland Toxoplasmosis (histopathology) 193 post mortem 3.1 Jokelainen et al., 2012
France DNA in faeces (PCR) 125 outdoor countryside cats 1.6 Poulle et al., 2017
France Antibodies (MAT) 130 from 5 farms 15 to 73 Simon et al., 2018
Germany DNA in faeces (PCR) 18,259 cats not defined 0.25 Herrmann et al., 2010
Germany Austria France Switzerland DNA in faeces

(PCR)

24,106 cats not defined 0.11 Schares et al., 2008
Greece Antibodies

(RIM)

805

748

pet cats

stray cats

20.6

23.1

Sioutas et al., 2022
Greece Antibodies

(IFAT)

75

260

24

0

95

457

pet cats

stray cats

catteries

pet shop

unknown

total

18.7

22.3

12.5

0

not given

20.8

Kokkinaki et al., 2023
Holland Antibodies

(ELISA)

450 pet cats 20.2 Opsteegh et al., 2012
Italy Antibodies (IFAT) 203 stray 30.5 Spada et al., 2012
Italy DNA in faeces (PCR) 146 healthy pet cats 10.3 Mancianti et al., 2015
Florence, Italy Antibodies (MAT) 50 colony cats 44 Mancianti et al., 2010
Florence, Italy Oocysts and DNA faeces (PCR) 50 colony cats 0

16

Mancianti et al., 2010
Latvia Antibodies

(ELISA)

216

26

pet

stray / shelter

50.5

61.5

Deksne et al., 2013
Latvia Antibodies

(ELISA)

163

73

indoor only

outdoor access

65.6

24.7

Deksne et al., 2013
Latvia Oocysts (flotation) 80 pet cats 2.5 Deksne et al., 2013
Madeira Antibodies

(MAT)

141 pet cats 30.5 Neves et al., 2020
Poland (South West) Antibodies

IFAT

208 “owned” 68.8 Sroka et al., 2018
Poland Oocysts and DNA in faeces (PCR) 41 “owned” 4.9

2.4

Sroka et al., 2018
Portugal Antibodies

(MAT)*

194 stray cats 24.2 Duarte et al., 2010
Lisbon, Portugal Antibodies

(MAT)*

423 stray cats 44.2 Waap et al., 2012
Lisbon, Portugal Isolation from brain homogenate 56 stray antibody-positive cats 23.8 Waap et al., 2012
Lisbon, Portugal Antibodies

(MAT)

215 pet cats 20.5 Esteves et al., 2014
Lisbon, Portugal DNA in faeces (PCR) 45 pet cats 35.6 Esteves et al., 2014
Portugal Antibodies

(ELISA)

183 pet cats 13.1 Pereira et al., 2023
Romania Antibodies

(MAT)

236 pet cats 47.0 Györke et al., 2011
Scotland Antibodies

(ELISA)

78 61 pet

17 shelter

19.2 Bennett et al., 2011
Barcelona, Spain Antibodies (MAT) 220 131 feral

89 pet

51.9

34.8

Gauss et al., 2003
Spain Antibodies

(MAT)*

316 stray cats 24.2 Montoya et al., 2018
Spain Oocysts (flotation) 459 stray cats 0 Montoya et al., 2018
Switzerland Oocysts (flotation then PCR) 252 171 pet, 43 stray, 38 with GIT signs 0.4 Berger-Schoch et al., 2011

* Recorded as DAT (direct agglutination test) but is same test that others called MAT; ELISA = enzyme linked immunosorbent assay; GIT = gastrointestinal tract; IFAT = indirect immunofluorescence antibody test; MAT = modified agglutination test; PCR = polymerase chain reaction; RIM = rapid immunochromatographic test.

Older cats are more likely to be antibody-positive to T. gondii (Gauss et al., 2003; Vollaire et al., 2005; Györke et al., 2011; Montoya et al., 2018; Xia et al., 2022; Pereira et al., 2023; Kokkinaki et al., 2023). In a study of 243 household cats in Romania, two peaks at around 10-months-old and 8-years-old were observed in the percentage of strongly positive blood samples (Györke et al., 2011). The antibody-prevalence was significantly higher in adult cats, cats fed meat or kitchen scraps, cats with outdoor access, and in cats from  rural areas (Györke et al., 2011; Deksne et al., 2013; Sioutas et al., 2022). Hunting was identified as the most significant risk factor for T. gondii infection in a Greek seroprevalence study (Sioutas et al., 2022). Being feral (Gauss et al., 2003) or stray was also a risk factor for T. gondii seropositivity in most (Wu et al., 2011; Xia et al., 2022; Pereira et al., 2023), but not all (Deksne et al., 2013) studies.

Birmans, Ocicats, Norwegian Forest Cats, and Persians are four to seven times more likely to be seropositive when compared with the Burmese and Korat cats (Jokelainen et al., 2012; Must et al., 2017). The differences in seroprevalence among cat breeds were most likely due to different lifestyles. A majority of the Korats, the breed with the lower seroprevalence, lived indoors only and received no raw meat (Jokelainen et al., 2012).

Cats of any age can shed T. gondii oocysts, although most shedding is observed in young cats (under one year of age) (Herrmann et al., 2010; Deksne et al., 2013; Simon et al., 2018). Oocyst shedding is rare even in antibody-positive cats: as can be seen from Table 1, less than 1% of faecal samples from Swiss, Spanish, and German cats (and also Korea) contained T. gondii oocysts (Schares et al., 2008; Herrmann et al., 2010; Berger-Schoch et al., 2011; Montoya et al., 2018; Ahn et al., 2019). However, in Syria, T. gondii-like oocysts were detected by flotation in the faeces of 26 of 68 (38.2%) of feral cats and 10 of 32 (31. 3%) pet cats, although as the authors say, without PCR, T. gondii oocysts are difficult to differentiate from those of Hammondia and Neospora which are similar in shape and size (Ismael and Al-Kafri, 2023), therefore the amount of T. gondii oocyst shedding was likely over-estimated.

Immunosuppression by pharmacological doses of immunosuppressive drugs (Lappin et al., 1991) and co-infection with feline immunodeficiency virus (FIV) (Lin et al., 1992; Lappin et al., 1996; Chi et al., 2021) or feline leukaemia virus (FeLV) do not cause re-excretion of oocysts in cats (Dubey et al., 2020)

There are no documented episodes of re-shedding of oocysts following natural infection of antibody positive cats, so far as we are aware, but occasional re-shedding of oocysts following experimental infection using a strain of T. gondii different from the original strain with which the cats were infected has been reported (Dubey et al., 1995; Malmasi et al., 2009; Zulpo et al., 2018). As shown in Table 1, in studies that look into oocyst shedding in the field, a very low percentage of cats sheds oocysts, indicating that re-shedding of oocysts is very unlikely in natural circumstances.

The annual burden of T. gondii oocysts in the environment is about 90 to 5,000 oocysts/square meter (Dabritz et al., 2007).  Oocysts can remain viable for over 200 days at temperatures under 30oC and for over 54 months at 4oC, but they are inactivated within one minute at 60oC (Dubey, 1998).

Oocyst shedding in the Northern hemisphere is more common between July and December than from January to June (Herrmann et al., 2010). The reason for this is unknown, but may be due to the availability of infected prey, such as rodents or birds, to cats (Herrmann et al., 2010).

Coprophagia with a subsequent intestinal passage by dogs plays a role in the dissemination of coccidian parasites for which cats are definitive hosts (Lindsay et al., 1997; Schares et al., 2005).

Transmission

The three major modes of transmission of T. gondii in all host species are congenital infection, ingestion of infected tissue and ingestion of oocyst-contaminated food or water (Dubey and Lappin, 2006; van Bree et al., 2018). Oocysts are able to survive for a long time in the environment, in soil and water.  In a laboratory experiment, 7.4% of T. gondii oocysts remained viable for over 100 days in dry conditions, and at 100 days 43.7% were viable in damp conditions (Lélu et al., 2012).  Less important routes of transmission are blood transfusions, organ transplants (Dubey, 2005; Dubey and Lappin, 2006) and tick bites (Sroka et al., 2003). T. gondii DNA was identified in two of 92 and four of 119 Ixodes ricinus, and 2.1% of 634 Dermacentor reticulatus, ticks in Poland (Sroka et al., 2003; Zajac et al., 2017;  Kocoń  et al., 2020). Whether or not ticks are able to transmit T. gondii infection to cats has not yet been established, but it seems possible given that toxoplasmosis associated with tick bites has been reported in humans (Sroka et al., 2003).

Parasitaemia during pregnancy of the host can cause placentitis and spread of tachyzoites to the foetus. Many kittens born to queens infected with T. gondii during gestation become infected transplacentally or when suckling. Lactogenic transmission is suspected because the organism has been detected in queens’ milk (Powell et al., 2001). Clinical signs are common in congenitally infected kittens, varying with the stage of gestation at the time of infection; some of these newborn kittens shed oocysts (Dubey, 2005; Dubey and Lappin, 2006).

There is no evidence for airborne transmission (Wadhawan et al., 2018).

Pathogenesis

The entero-epithelial (coccidian) life cycle

This cycle is found only in the feline host. Most cats are infected by ingesting intermediate hosts—typically rodents—infected with tissue cysts. Bradyzoites are released in the stomach and intestine from the tissue cysts when digestive enzymes dissolve their wall. They enter epithelial cells of the small intestine and give rise to schizonts, initiate five types of predetermined asexual stages, and merozoites released from the schizonts eventually form male and female gamonts. After fertilization, a wall is formed around the fertilized macrogamont to form an oocyst (Figs. 2, 3). Oocysts are round to oval, 10 x 12 μm in size, and are still unsporulated (not infectious) when passed in faeces. After exposure to air and moisture for one to five days, they sporulate to contain two sporocysts, each with four sporozoites (Dubey, 2005; Dubey and Lappin, 2006).

The entero-epithelial cycle is usually completed within three to ten days after ingestion of bradyzoite tissue cysts, which is the case in up to 97% of naive cats. Only after the rare event in which cats ingest oocysts or tachyzoites, is the formation of new oocysts delayed and shedding can occur up to 18 days (rarely more) after ingestion. However, only 20% of cats fed oocysts will shed oocysts (Dubey, 2005; Dubey and Lappin, 2006).

Fig. 2. T. gondii tissue cyst (unstained). Slowly-dividing bradyzoites can be seen inside. From the public domain, Wikipedia USA

Fig. 2. T. gondii tissue cyst (unstained). Slowly-dividing bradyzoites can be seen inside. From the public domain, Wikipedia USA

Fig. 3. Cysts develop in the tissues of many vertebrates, here in mouse brain; resting parasites (stained red) are enveloped by a thin cyst wall. Image is in the public domain, originally from the Agricultural Research Service, US Dept of Agriculture

Fig. 3. Cysts develop in the tissues of many vertebrates, here in mouse brain; resting parasites (stained red) are enveloped by a thin cyst wall. Image is in the public domain, originally from the Agricultural Research Service, US Dept of Agriculture

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. gondiiinfected 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. 5. Thoracic radiographs (latero-lateral view) of a cat with pulmonary toxoplasmosis (courtesy of Katrin Hartmann, Medizinische Kleintierklinik, Ludwig-Maximilians-Universitaet Muenchen, Germany).


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. 4. 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 hyperesthesia (courtesy of Katrin Hartmann, Medizinische Kleintierklinik, Ludwig-Maximilians-Universitaet Muenchen, Germany).


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.


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 hammondiBesnoitia 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. 6. T. gondii oocysts in fecal flotation. Source http://dpd.cdc.gov/dpdx/HTML/ImageLibrary/Toxoplasmosis_il.htm; US Center for Disease Control and Prevention


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 CT, 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

Drug ABCD recommendation Comment Evidence-based grade
Antiparasitic therapy
Clindamycin 10-12.5 mg/kg PO q12h for 28 days Treatment of choice for toxoplasmosis III
Trimethoprim sulfamethoxazole 15 mg/kg PO q12h Has been used as an adjunct in severe neurological cases (Wagner and Cooper, 2018) IV
Symptomatic topical therapy
Prednisolone acetate (1%) eye drops orTobramycin dexamethasone (Tobradex ophthalmic solution®, Alcon Comp, Egypt)and tropicamide1% eye drops (Mydria-cyl®, Alcon Comp, Egypt) 3-4 times/day. Use in addition to systemic clindamycin treatment
Apply topically to the eye q6-8h
For cats with toxoplasma-induced uveitis (to avoid secondary glaucoma and lens luxation) IV

 Evidence-based medicine (EBM) is a process of clinical decision-making that allows clinicians to find, appraise and integrate the current best evidence with individual clinical expertise, client wishes and patient needs. This article uses EBM ranking to grade the level of evidence of statements in relevant sections on diagnosis, disease management and control, as well as vaccination. Statements are graded on a scale of I to IV as follows:

✜ EBM grade I This is the best evidence, comprising data obtained from properly designed, randomised controlled clinical trials in the target species (in this context cats);

✜ EBM grade II Data obtained from properly designed, randomised controlled studies in the target species with spontaneous disease in an experimental setting;

✜ EBM grade III Data based on non-randomised clinical trials, multiple case series, other experimental studies, and dramatic results from uncontrolled studies;

✜ EBM grade IV Expert opinion, case reports, studies in other species, pathophysiological justification. If no grade is specified, the EBM level is grade IV.

Further reading: Roudebush P, Allen TA, Dodd CE, Novotny BJ. Application of evidence-based  medicine to veterinary clinical nutrition. J Am Vet Med Assoc 2004; 224: 1765–71.

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 64oC (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

Sources of infection for humans

“Contact with cats is not a risk factor for T. gondii infection” (Cook et al., 2000).

Most common routes of infection for humans

  • Ingestion of meat containing tissue cysts is the most common route of  infection (Kapperud et al., 1996; Cook et al., 2000; Tekay and Özbek, 2007; Belluco et al., 2016).
  • Ingestion of sporulated oocysts, either from the environment, e.g., through contact with contaminated soil, or from faeces of shedder cats is the second most common route. This can also happen when eating unwashed fruit or vegetables (Pinto-Ferreira et al., 2019).
  • Ingestion of sporulated oocysts through contact with contaminated water (Bell et al., 1995; Pinto-Ferreira et al., 2019) or ingesting fresh shellfish (Merks et al., 2023).

Less common routes of infection for humans 

  • Ingestion of tachyzoites in raw (unpasteurised) goat milk (Pinto-Ferreira et al., 2019).

Reducing risk of infection for humans who eat meat

  • Thorough cooking (to at least 67 oC) or freezing of meat (to minus 20 oC for at least 2 days) will inactivate tissue cysts (Dubey, 1988; Dubey et al., 1990; Lunden and Uggla, 1992; Dubey et al., 1998; Mirza Alizadeh et al., 2018).
  • Sporulated oocysts can be inactivated by freezing to minus 20 oC for at least 3 weeks (Kuticic and Wikerhauser, 1996, cited by Mirza Alizadeh et al., 2018).
  • Clean chopping boards etc. with boiling water (Mirza Alizadeh et al., 2018).

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

Recommendations to reduce the risk of parasite transmission from cat to human

  • Litter trays should be emptied daily so that oocysts do not have sufficient time (24 hours) to sporulate (Dubey et al., 2011).
  • Gloves should be worn when handling cat litter, and hands should be washed thoroughly after cleaning of litter trays.
  • Litter tray liners should be used if possible, and the tray cleaned regularly with detergent and scalding water.
  • Cat litter should be disposed in sealed plastic bags.
  • Children’s sandpits should be covered when not in use, to prevent cats from using them.
  • Only properly cooked food or commercial cat food should be fed.
  • Hands should be washed after contact with a cat (especially before eating).

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

For households with immunocompromised persons or pregnant women, the following additional advice is given:

  • Immunosuppressed persons and pregnant women should avoid contact with cat litter.
  • Cats should not be fed raw or partially cooked meat.
  • Cats should be discouraged from eating insects (e. g., cockroaches) (Wallace, 1972).
  • Cats should be tested for T. gondii antibodies; their presence indicates past infection. These cats will most likely not be a source of infection as they have completed their period of oocyst shedding.
  • Cats without antibody had not been infected earlier and, when newly infected, will shed oocysts in their faeces for a short time. If they are hunters then they should therefore be kept indoors during the phase of immunosuppression or pregnancy of the owner.

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).

  • Cats shedding oocysts in faeces are extremely rare (under 1%) (See Table 1 for references). In one study, only one of 250 cats shed T. gondii oocysts (Berger-Schoch et al., 2011).
  • Contact with cats has no influence on the probability of people developing antibodies to T. gondii, whereas consuming raw meat significantly increases the risk of acquiring the infection (Flegr et al., 1998; Cook et al., 2000; Tekay and Özbek, 2007; Belluco et al., 2016).
  • Veterinarians working with cats are not more likely to become infected with T. gondii or to suffer from toxoplasmosis than the general population, including people without cat contacts (Behymer et al., 1973; Sengbusch et al., 1976; Tizard and Caoili, 1976; DiGiacomo et al., 1990).
  • Stroking a cat will not spread the infection. Even when cats are shedding  in their faeces,  oocysts cannot be found on their coat (Dubey, 1995).
  • Cat ownership does not increase the risk of toxoplasmosis in persons with an HIV infection. Although toxoplasmosis is more common in HIV-infected persons  the disease results from reactivation of a previous infection rather than from acquiring a new infection.
  • Most people are infected with T. gondii through ingestion of undercooked meat, especially goat, mutton, and pork.
  • The risk of infection from cats is low, except for young children playing in soil contaminated with sporulated oocysts (Wallace et al., 1993).
  • Bites or scratches from an infected cat do not transmit the infection.
  • Infected cats under treatment with immunosuppressive drugs at standard doses do not start shedding oocysts in their faeces (Lappin et al., 1991).
  • Infected cats also do not re-shed oocysts in their faeces when they become immunosuppressed due to infection with FIV or FeLV (Lappin, 2001). Cats infected with FIV or FeLV that are subsequently infected with T. gondii do not shed oocysts for any longer or in any greater numbers than other cats (Lappin et al., 1996; Dubey and Lappin, 2006).
  • Newly identified strains of T. gondii are highly infectious for species other than cats; thus, cats might actually become less important in the spread of this infection.

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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