Background

Hammondia hammondi, a coccidian parasite, is closely related to the zoonotic protozoan Toxoplasma gondii [1, 2]. Similar to T. gondii, H. hammondi uses felids such as the domestic cat as its definitive hosts [3]. Oocysts of T. gondii and H. hammondi are morphologically indistinguishable [3, 4]. Thus, faecal examinations solely based on microscopy cannot be used to estimate the prevalence of T. gondii or H. hammondi oocysts in feline hosts [5,6,7].

Laboratory mice, rats, hamsters, guinea pigs, wild rodents, rabbits, goats and dogs are susceptible to infection with H. hammondi oocysts [3, 6, 8,9,10,11,12,13,14]. While monkeys [15] are also experimental intermediate hosts of H. hammondi, no H. hammondi infection has so far been demonstrated in humans. No avian species, including chickens [16], quails [13] and pigeons [10, 13], could be infected with H. hammondi, which suggests avians are not its intermediate hosts [7]. In contrast to T. gondii, H. hammondi infections are basically asymptomatic in both definitive and intermediate hosts [3].

While several polymerase chain reactions (PCRs) are available for the diagnosis of T. gondii infections, only a few diagnostic PCRs for H. hammondi have been reported so far. Those for T. gondii are based on a variety of targets, including the B1 gene [17], the internal transcribed spacer (ITS)-1 region of ribosomal DNA (rDNA) [18] and a 529-base pair (bp) repetitive element, also known as TgREP-529 (GenBank AF146527) [19, 20]. The ones reported for H. hammondi are end-point PCRs, which use as targets ITS-1 rDNA [21] or a 529-bp repeat, which is similar to the T. gondii repetitive element and is designated here as HhamREP-529 [6]. To the best of our knowledge, no real-time PCR for the diagnosis of H. hammondi infections has so far been published. The availability of a real-time PCR will allow simplification of H. hammondi prevalence estimates. It will also allow the detection and quantification of oocysts of this parasite in feline faeces and in tissues of natural and experimental intermediate hosts.

Since TgREP-529 of T. gondii exists up to about 200 times in the genome of a single organism, as recently shown by third-generation sequencing [22], and sequence information on a homologue of this repetitive element in H. hammondi was readily available from prior studies [6], we decided to establish a HhamREP-529-based real-time PCR for H. hammondi. Unexpectedly, we faced several problems when we tried to establish a sensitive primer and probe combination, probably due to sequence variations between individual repeats, as observed earlier [6].

Methods

Parasite strains to generate reference DNA

Field isolates of H. hammondi oocysts were made available by IDEXX Laboratories, Ludwigsburg, Germany, and the University of Veterinary and Pharmaceutical Sciences, Brno, Czech Republic, in the frame of projects that are aimed at obtaining European T. gondii isolates (Additional file 1: Table S1). Faecal samples were tested by flotation and examined microscopically as described previously [7]. If T. gondii-like oocysts were detected, purified oocysts were genotyped at the species level by PCR using DNA extracted from oocysts. Occasionally, gamma interferon gene knockout (GKO) mice [C.129S7(B6)-Ifngtm1Ts/J; The Jackson Laboratory, Bar Harbor, ME] were in addition inoculated with sporulated oocysts via oral gavage; parasites were recovered as described previously [23] and genotyped. One H. hammondi oocyst isolate from Iran had been kindly provided by Dr. Morteza Hosseininejad, Faculty of Veterinary Medicine, Shahr-e Kord, Iran, in 2010. Oocysts of the isolate H.H.34 were from the USA, and whole genome sequences (GenBank: AHJH01004981.1) were recently obtained for these. H.H.34 oocysts were used to infect GKO mice to obtain tissue stages for DNA extraction, and DNA was also extracted from the oocysts (Additional file 1: Table S1; Additional file 2: Table S2).

To confirm the specificity of the Hham-qPCR1, we used DNA from three T. gondii strains (RH, ME49, NED), Neospora caninum (Nc-1), Besnoitia besnoiti (Evora), Hammondia heydorni, Eimeria bovis, Cystoisospora felis, Cystoisospora rivolta, Cystoisospora burrowsi, Cystoisospora canis, Cryptosporidium parvum, Sarcocystis hirsuta, Sarcocystis bovifelis, Sarcocystis hominis, Sarcocystis cruzi, Giardia spp. and Tritrichomonas foetus, in addition to DNA isolated from Hammondia hammondi. T. gondii, N. caninum and B. besnoiti were obtained by in vitro cultivation of the parasites, which yielded reference samples consisting of 100 ng/µl concentrated pure DNA. For the remaining oocyst reference DNA samples, DNA was extracted from field isolates (H. heydorni, Cystoisospora spp., Sarcocystis spp.; 104–105 oocysts or sporocysts) yielding 100 µl DNA for each parasite. Eimeria bovis oocysts were kindly provided by Dr. Christian Bauer, Institute for Parasitology, Giessen, Germany. Oocysts of C. parvum (Germany) were kindly provided by Prof. Dr. A. Daugschies, Institute of Parasitology Leipzig, Germany. Giardia spp. and T. foetus DNA were kindly provided by Dr. Christian Klotz, Robert Koch-Institut, Berlin, Germany and Dr. Klaus Henning, Friedrich-Loeffler-Institut, Jena, Germany.

Experimental infections of mice

GKO mice (n = 28) were orally inoculated with different isolates and varying doses, depending on the availability of field isolates, in experiments to isolate T. gondii or to trigger the permanent growth of H. hammondi in mice and eventually in cell culture (Additional file 2: Table S2). Sporulated oocysts were counted using a Neubauer chamber and mice infected via oral gavage as described [23].

DNA extraction

Oocysts were isolated from faeces using combined sedimentation and flotation by adding 13 ml concentrated sucrose (specific gravity of 1.3) to 1 ml faecal sediment as described previously [24]. Floating oocysts were collected, washed three times by centrifugation (1100× g, 7 min, without brake) and a five- to tenfold volume of phosphate-buffered saline. DNA was extracted from the final pellet using a phenol/chloroform purification method as previously described [23] or using the NucleoSpin Soil kit (Macherey and Nagel, Düren, Germany) following the manufacturer’s recommendations.

Tissue samples (25 mg) were extracted by using the NucleoSpin Tissue kit (Macherey and Nagel) following the manufacturer’s recommendations. Because it was not the aim of the present study to estimate parasite DNA concentration per infected cell but per type of tissue, DNA samples from tissues were not normalized but used as extracted.

Conventional endpoint PCRs

To confirm the presence of H. hammondi in oocyst samples, a previously published end-point PCR was applied as described to examine oocyst DNA [4].

To test DNA extracted from oocyst samples for coccidian DNA, a PCR was performed using the common apicomplexan small subunit ribosomal DNA (SSU-rDNA) primers COC-1 and COC-2 [25]. Primers were used at a final concentration of 0.5 mM and deoxynucleotide triphosphates at a final concentration of 250 mM each (Stratec Molecular, Berlin, Germany). Taq polymerase (Stratec Molecular) was used at 1U/25 μl with the buffer system supplied with the enzyme. The PCR cycling conditions were the same as previously reported [26]. To confirm the presence of Sarcocystis spp., primers SarcoFint and SarcoRint specific for Sarcocystis spp. SSU-rDNA were used as previously described [27]. To test for Giardia spp. DNA, the published primers AS1 GiardiaF and AS2 GiardiaR [28] were used, and for T. foetus DNA, we used the primer pair TFR1/TFR2 [29]. All reagents, except for primers, were the same as described above. They were used in the same concentrations as described for the coccidian PCR. The PCR cycling conditions were 94 °C for 5 min, followed by 35 cycles of 60 °C for 1 min, 72 °C for 1 min and 94 °C for 1 min. The PCR ended with a final extension at 72 °C for 10 min.

Identification of primer pairs suitable for amplifying H. hammondi DNA

To identify optimal primer pairs, a SybrGreen real-time PCR was performed using DNA (D9494) from a H. hammondi isolate (VB919008), diluted to resemble the DNA of approximately 5 oocysts/µl DNA. All possible combinations of forward and reverse primers (Additional file 3: Table S3) were checked in a SybrGreen real-time PCR using the iTaq Universal SYBR Green Super mix (Bio-Rad Laboratories, Munich, Germany) to test the H. hammondi template in comparison to a no-template control (DNA-grade water). Each primer pair was evaluated twice in two independent SybrGreen real-time PCRs and the results were recorded as mean Δ threshold cycle (Ct)pos-neg, i.e. the ΔCt between results (Ct values) for the H. hammondi template and the no-template control (Additional file 4: Table S4).

Novel real-time PCR to detect H. hammondi DNA

The quantitative TaqMan real-time PCR was performed as described previously [26], including the integration of an internal control (IC) system [26, 30]. The novel real-time PCR employing the optimal primer-probe combination was designated Hham-qPCR1. To monitor inhibition in real-time PCR, a heterologous plasmid DNA resembling the enhanced green fluorescent protein (EGFP) gene [30] was added to the reaction mix including the primers EGFP1-F, EGFP2-R and the probe EGFP1 [26]. A 712-bp fragment of the EGFP gene was amplified as the EGFP template and cloned into the pGEMTeasy standard cloning vector (Promega, Walldorf, Germany) in reverse orientation to obtain IC-2 DNA (pGEM-EGFP2-rev). The amount of IC-2 DNA added to each reaction was adjusted in such a way that it resulted in a Ct value of about 32 in the real-time PCR. Reactions were performed in a final volume of 20 μl using a commercial master mix (PerfeCTa MultiPlex qPCR ToughMix; Quantabio, VWR International, Darmstadt, Germany) and a CFX96 instrument (Bio-Rad Laboratories). Primers and probes (Additional file 3: Table S3) were purchased from MWG-Biotech (Ebersberg, Germany).

Standard concentrations for primers (500 nM; Additional file 3: Table S3) and probes (100 nM for H. hammondi probes, Cy5 labelled; Additional file 3: Table S3; EGFP1, HEX labelled [26]) were applied. These standard concentrations were also applied to the finally selected primer-probe combination (Hham275F, Hham81R, Hham222P). The cycling conditions in the Hham-qPCR1 were 95.0 °C (5 min, initial denaturation), followed by 45 cycles, during which the samples were first incubated at 95.0 °C for 10 s and then at 60.0 °C for 30 s. After each cycle, light emission by the fluorophore was measured. Real-time PCR results were analysed using the CFX manager software version 1.6 (Bio-Rad Laboratories).

Sensitivity and specificity of Hham-qPCR1

The analytical sensitivity was determined using a H. hammondi oocyst isolate with a high sporulation rate (P18/2900; sporulation rate 84%). Tenfold dilutions of three DNA samples isolated from 105 oocysts were tested by the Hham-qPCR1 in twofold repetition including the EGFP IC.

To confirm that the novel PCR recognized a broad range of H. hammondi specimens, oocyst isolates from Germany and other countries were tested by Hham-qPCR1. These DNA samples included oocyst DNA isolated from 19 H. hammondi samples of different origin from Germany, four samples from Denmark, two from Austria, and one each from France, the Czech Republic and the USA, i.e. n = 28 samples in total (Additional file 1: Table S1). The presence of H. hammondi had been confirmed by our previously published end-point PCR [6]. All these samples had tested negative in a Tg-qPCR [31, 32].

The analytical specificity was confirmed using DNA samples of parasites closely related to H. hammondi, which are mentioned in the “Parasite strains to generate reference DNA” section. The presence of DNA in these samples was confirmed by the SSU-rDNA end-point PCR as detailed in the “Conventional endpoint PCRs” section.

Statistical analysis

Calculations of medians and SDs were performed using EXCEL 2019, Microsoft Office (Microsoft, Seattle, WA).

To determine the relatedness of real-time PCR results and oocyst concentrations in samples, linear regression was performed using the lm command in R version 3.5.3 (R Core Team 2017). For the analysis, oocyst concentrations (number of oocysts per millilitre) were log 10 transformed.

Figures were assembled using R versions 3.5.3 or 4.0.0 (packages ggplot2, reshape and scales). Box plots displaying real-time PCR data of infected mice show the median, 25th and 75th percentiles, 1.5 interquartile ranges and outliers; PCR results were considered negative at a cut-off Ct value of 45.

Results

Location of primers

Potentially suitable primers and probes (Additional file 3: Table S3) for two 5’-nuclease quantitative real-time PCR assays were selected by in silico methods (Fig. 1). To this end, published sequences (n = 9) of HhamREP-529 were downloaded from GenBank (KC223619, JX477424, EU493279, EU493280, EU493281, EU493282, EU493283, EU493284, EU493285) and aligned with Geneious 10.0.9 (Multiple Geneious Alignment, Global alignment with free end gaps, Gap open penalty 12, Gap extension penalty 3, refinement iterations 2). Primers were selected to amplify a fragment with a maximum size of about 150–160 bp.

Fig. 1
figure 1

Workflow for establishing a quantitative real-time polymerase chain reaction for Hammondia hammondi (Hham–qPCR1) in oocysts and intermediate host tissues using a sensitive and specific primer-probe combination and an enhanced green fluorescent protein (EGFP)-based internal control (IC)

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Identification of optimal primer pairs to amplify H. hammondi DNA

In total, 39 primer pairs were evaluated with a SybrGreen assay and H. hammondi reference DNA D9494. Out of 51 primer pairs tested, n = 22 revealed ΔCtpos-neg values of > 4. Five of these 22 pairs revealed a large difference between two independent tests and in 17, the coefficient of variation of ΔCtpos-neg (Additional file 4: Table S4) was less than 0.4. These 17 primer pairs were further validated with up to four of the probes Hham55P, Hham75P, Hham110P or Hham222P (Additional file 3: Table S3) depending on the location of primers and probes (Fig. 2). Eleven of the 28 primer-probe combinations were excluded due to cross-reaction with T. gondii DNA (1000 ng). Four of the remaining primer-probe combinations showed mean relative fluorescence unit values > 1000 and mean Ct values of 33.6 to 35.3 in from three to five independent tests. The finally selected primer-probe combination consisted of the primers Hham275F and Hham81R and the probe Hham222P (Fig. 2; Additional file 3: Table S3). The analytical sensitivity and specificity were assessed for this primer-probe combination in a multiplex real-time PCR, in which EGFP-specific reagents were included as an IC to assess PCR inhibition, in addition to H. hammondi-specific reagents. This assay was designated as the Hham-qPCR1.

Fig. 2
figure 2

Locations of primers and probes on a fragment of the 529-base pair (bp) repeat sequence of Hammondia hammondi and Toxoplasma gondii. Consensus sequences of H. hammondi have been deposited in GenBank (KC223619, JX477424, EU493279, EU493280, EU493281, EU493282, EU493283, EU493284, EU493285); the RH529-1 sequence has been previously published [19]. Further details are provided in Additional file 3: Table S3. There are several sequence variants of H. hammondi; lowercase letters indicate polymorphic sites. Blue indicates a probe, green indicates a forward primer, red indicates a reverse primer, orange indicates a sequence difference, arrow-headed end indicates a 5′-end

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Analytical sensitivity and efficiency of the Hham-qPCR1

The analytical sensitivity of the Hham-qPCR1 was determined using a German H. hammondi oocyst isolate, which showed a high sporulation rate. Tenfold dilutions of three DNA samples isolated separately from 105 oocysts each were twice examined by the Hham-qPCR1 including the EGFP IC. A PCR efficiency (E) = 103.9%, a coefficient of determination (R2) = 99.7% and a slope of the regression line of 3.231 were recorded (Fig. 3).

Fig. 3a, b
figure 3

Characteristics of the Hham-qPCR1 with an EGFP IC. a DNA equivalents of 0.1 Hammondia hammondi oocyst reacted positively in the PCR [colours represent different DNA equivalents: 104 oocysts (red); 103 oocysts (orange); 102 oocysts (yellow), 10 oocysts (green); 1 oocyst (blue); 0.1 oocyst (black)]. b The Hham-qPCR1 was characterized by a PCR efficiency (E) of 103.9%, and linear regression using DNA equivalents of 104–10−1 oocysts yielded a coefficient of determination (R2) of 99.7%. RFUs Relative fluorescence units, Cq threshold cycle (Ct); for other abbreviations, see Fig. 1

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To confirm that the Hham-qPCR1 recognizes a broader range of H. hammondi isolates, we analysed DNA of oocysts collected from cat faeces. These DNA samples included 19 samples from Germany, four from Denmark, two from Austria and one from France, the Czech Republic and the USA each, i.e. a total of 28 samples. In the new Hham-qPCR1 these samples yielded Ct values between 15.9 and 28.3, which is equivalent to the DNA of 102–103 oocysts per microlitre (Additional file 1: Table S1). There was a linear relationship between the logarithm (log 10) of the number of isolated oocysts per millilitre and the observed Ct values, which was characterized by an R2 of 17.5%.

Analytical specificity

The analytical specificity of Hham-qPCR1 was assessed using DNA samples of parasites closely related to H. hammondi, i.e. Toxoplasma gondii (strains RH, ME49, NED), N. caninum (Nc-1), B. besnoiti (Evora), H. heydorni, E. bovis, C. felis, C. rivolta, C. burrowsi, C. canis, C. parvum, S. hirsuta, S. bovifelis, S. hominis, S. cruzi and DNA of parasites frequently observed in cat faeces like Giardia spp. and Tritrichomonas foetus. Presence of DNA in these samples was confirmed by end-point PCR (Fig. 4). In the Hham-qPCR1, no amplification with any of these DNA samples was observed.

Fig. 4
figure 4

Confirmation that samples used to determine the analytical specificity of Hham-qPCR1 contained sufficient parasite DNA. Lanes 1–19 Hammondia hammondi, Toxoplasma gondii (RH), Toxoplasma gondii (ME49), Toxoplasma gondii (NED), Neospora caninum (Nc-1), Besnoitia besnoiti (Evora), Hammondia heydorni, Eimeria bovis, Cystoisospora felis, Cystoisospora rivolta, Cystoisospora burrowsi, Cystoisospora canis, Cystoisospora parvum, Sarcocystis hirsuta, Sarcocystis bovifelis, Sarcocystis hominis, Sarcocystis cruzi, Giardia spp., Tritrichomonas foetus. DNA in lanes 1–13 had been amplified with Coccidia-specific primers COC-1 and COC-2, in lanes 14–17 with the Sarcocystis spp.-specific primers SarcoFint and SarcoRint, in lane 18 by the Giardia spp.-specific primers RH11_RH4 and RH11_RH5, and in lane 19 by the Trichomonas spp.-specific primers TRF1 and TRF2. M Marker (100 bp), C control (no template)

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Examination of tissue stages of H. hammondi

Tissues of 28 GKO mice infected with H. hammondi for varying time periods were available, from which DNA was extracted and tested by Hham-qPCR1 (Fig. 5a, b). From each of 28 mice, seven tissues (brain, heart, lung, liver, kidney, spleen and the proximal hind limb musculature) had been collected. In the first days post-infection (DPI; DPI 3, DPI 6), the median Ct values in Hham-qPCR1 ranged between 22.3 and 31.3 for organs including mostly muscle tissue (except for the distal hind limb and tongue, 39.2 or 38.9, respectively); by contrast, median Ct values in brain tissue ranged only from 36.5 to 39.9 (Fig. 5a). On DPI 42, skeletal muscle (i.e. proximal hind limb musculature) showed the lowest median Ct values (14.2) of all tissues in this comparison, followed by heart (17.3) and lung (20.4). Of the remaining organs, brain and spleen showed similar median Ct values (24.9 and 25.4). Liver and kidney also yielded Ct values in the same range (27.4 and 29.4). Later during infection (DPI 74–186 and 272), heart and skeletal muscle had the lowest median Ct values followed by lung, brain, kidney and spleen. However, Ct values at DPI 272 were considerably lower than during DPI 74–186 in mice (Fig. 5a).

Fig. 5a, b
figure 5

Hham-qPCR1 Ct values in various tissues of gamma interferon gene knockout mice inoculated with Hammondia hammondi. a For 28 mice necropsied 3, 6, 42, 74–186, or 272 days post-infection (DPI), seven tissues, brain (Bra), heart (Hrt), lung (Lun), liver (Liv), kidney (Kid), spleen (Spl) and the proximal hind limb musculature (HLp), were available. b For some of the mice, samples of the HLp, tongue (Ton), buccal musculature (Buc), proximal forelimb (FLp), distal forelimb (FLd), distal hind limb (HLd) and diaphragm (Dia) were also available and tested. Box plots show the median (line), 25th and 75th percentiles (box), 1.5 interquartile range (whiskers) and outliers (dots)

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Moreover, muscle tissues obtained from different locations on DPI 3, 6 and 42 were comparatively examined (Fig. 5b). On DPI 3 and 6, heart showed the lowest Ct values of all muscle tissues (median Ct 27.3 and 23.8), while heart and tongue had the highest Ct values on DPI 42 (Fig. 5b). The remaining muscle tissues showed very similar Ct values, i.e. medians ranged from 14.7 to 13.0. In all skeletal muscle tissues, the Ct values had dropped considerably between DPI 6 (Ct 27.0–25.3) and DPI 42 (Ct 10.2–13.3). In brain tissue, a similar decrease in Ct values was observed between DPI 6 and DPI 42. By contrast, for heart, a ΔCt of only 6.5 was observed between the median Ct values of DPI 6 and DPI 42 (Additional file 5: Table S5).

Discussion

The present study reports on the development and characterization of a 5’-hydrolysis real-time PCR (TaqMan PCR) for the quantitative detection of H. hammondi. To the best of our knowledge, this is the first real-time PCR for the detection of H. hammondi DNA in feline faecal samples or intermediate host tissues.

There are numerous end-point and real-time PCRs for T. gondii that target an up to about 200-fold repeated element in the genome of the parasite (TgREP-529). We identified a similar 529-bp repeat in H. hammondi and established the Hham34F/Hham3R end-point PCR to diagnose H. hammondi infection several years ago [6]. This PCR was subsequently applied to test feline faecal samples [5, 7], rodent tissues [33] and used in cell biological studies to elucidate differences between the life cycles of T. gondii and H. hammondi [34]. Although the use of HhamREP-529 seemed to be ideal for establishing a quantitative real-time PCR, and enough sequence data were available, no real-time PCR had previously been established for H. hammondi.

We found that only a small set of the primers that we had predicted in silico revealed enough specificity and sensitivity for the amplification of H. hammondi DNA. The identification of a suitable probe was also more difficult than expected. In a previous study, we cloned and sequenced several PCR amplification products of HhamREP-529 and observed considerable sequence variation [6]. It seems possible that these sequence polymorphisms prevented many of the in vitro-selected primers and probes from binding sufficiently to DNA.

The HhamREP-529 and the TgREP-529 sequences are similar, but not identical (identity ranging from 93.7 to 97.5%; [6]), and the H. hammondi primers and probes that were finally selected showed a considerable number of mismatches with respect to the T. gondii sequence. To confirm the analytical specificity relative to T. gondii, we used DNA samples from the three major European and North American clonal lineages of T. gondii. In addition, we confirmed the specificity with a variety of DNA samples from parasite species or genera related to H. hammondi or T. gondii, including parasitic protozoa frequently observed in cat faeces like Giardia spp. or T. foetus. Our results indicate that Hham-qPCR1 has an excellent analytical specificity.

When we tested DNA samples of more than 20 H. hammondi isolates from Austria, Denmark, France, Germany and the USA (oocysts), the Czech Republic (oocysts, mouse tissue) and Iran (mouse tissue), there was no indication that our Hham-qPCR1 lacks sensitivity for particular isolates. Due to limited knowledge on the global population structure of H. hammondi, we cannot exclude, however, the possibility that yet unidentified lineages of H. hammondi exist, which may be genetically diverse. If there are further H. hammondi lineages, the possibility remains that our primer-probe combination could show limited or no binding to the DNA of such lineages. However, experience with T. gondii, which has several genetically diverse haplogroups, suggests that TgREP-529 is relatively conserved. In general, only a few failures of TgREP-529-based PCRs for the detection of T. gondii have been reported [35,36,37,38,39,40]. However, there are some cases in which other PCR assays either based on ITS-1 rDNA [41] or on B1 as a target [42] detected a considerable number of additional T. gondii-positive DNA samples compared to the number of positive results obtained by TgREP-525-based PCRs.

An earlier study observed only a few differences among various strains of the canonical clonal lineages regarding the number of TgREP-529 repeat units per organism [43], and a very recent study reports similar observations [22]. It has been discussed, but so far not investigated, to what extent sequence polymorphisms between the tested canonical clonal lineages contributed to these findings [43]. Future studies need to assess if there are differences in the number of HhamREP-529 repeats in strains of H. hammondi.

We validated the analytical sensitivity of the Hham-qPCR1 with oocyst DNA. Similar to the results of our previous study using an end-point PCR [6], the analytical sensitivity of the Hham-qPCR1 was equivalent to the DNA content of 0.1 oocyst. Considering that the 200 µg faeces, that we usually used, yield a 100 µl DNA solution, the limit of detection comes close to 5 oocysts/g faeces, which is comparable to the analytical sensitivity reported for a T. gondii copro-PCR [44, 45] and a Besnoitia darlingi real-time PCR [26]. For comparison, the microscopical evaluation of faecal flotation by sucrose density centrifugation showed a limit of detection of 250 oocysts/g faeces [46].

To demonstrate the practical value of the novel Hham-qPCR1 in examining infections of intermediate hosts, we used tissue samples of GKO mice inoculated with H. hammondi oocysts. In contrast to infection with T. gondii, H. hammondi infections generally become chronic in GKO mice without causing disease (own unpublished data). Even when clinical signs were mentioned in previous studies, they were not described in detail [3, 5]. There are indications, however, that high doses of 106 oocysts cause serious illness in < 10% of inoculated GKO mice [5]. The reason why H. hammondi is not as virulent as T. gondii seems to be that it replicates only a few times during the tachyzoite stage and eventually differentiates into bradyzoites [34] that no longer cause cell death or harm to the host. As evidenced by laborious and time-consuming histological examinations, cyst-producing (i.e. BAG-1 positive) stages appeared as early as 7 DPI in mice, suggesting that, at least until then, H. hammondi multiplied in the tachyzoite stage [3]. Tachyzoite subculture and in vitro tissue cyst experiments suggest that there is significant tachyzoite multiplication in H. hammondi until about 15–18 DPI [34]. Thereafter, BAG-1-positive tissue cysts were microscopically visible in heart and skeletal muscles and, until 22 DPI, also in mesenteric lymph nodes, liver, lung, kidney and brain. After that time, tissue cysts were microscopically only observed in the lung, heart and skeletal muscles [3]. Our real-time PCR results corroborate these findings. In chronically infected mice (i.e. after 6 weeks of infection), the lowest Ct values were for skeletal muscle, heart and lung tissue (often Ct < 25, and especially in skeletal muscle Ct < 15); however, infections were also frequently observed in the other organs, but high Ct values dominated (often Ct > 30), if these organs were positive at all. This shows that H. hammondi can also persist in tissues other than those of the lung, heart or skeletal muscle. The dramatically decreasing Ct values in various skeletal muscles, characterized by ΔCt values of 10.2–13.3 between DPI 6 and DPI 42, are remarkable. They suggest a 103- to 104-fold increase in parasite loads in these tissues during this period of time. In the case of heart tissue, a ΔCt of only 6.5 was observed between DPI 6 and DPI 42, i.e. a 102-fold increase in parasite load. This finding may indicate that striated muscle supports multiplication and persistence of H. hammondi better than smooth muscle. This is in contrast to T. gondii, for which heart tissue was identified as a site of predilection for chronic T. gondii infections in many animal species [47]. As H. hammondi and T. gondii partially use the same animal species as intermediate hosts, differences in host tissue tropism may prevent, to some extent, interspecies competition in these very closely related parasites.

An increase in the H. hammondi parasitic load was also observed in brain tissue, which was characterized by a ΔCt of 11.7, i.e. a 104-fold increase in parasite load, between DPI 6 and DPI 42. However, the parasite load was 103 times lower than in skeletal muscle, which may suggest that H. hammondi tachyzoites only reach the brain late during infection. An increase in parasite load may be mainly due to limited tachyzoite and subsequent bradyzoite multiplication in immunologically isolated tissue cysts. This is in accord with earlier observations that there is no cerebral disease in H. hammondi-inoculated mice [3].

In mice sacrificed after 42–271 DPI, the results suggested that the parasitic load gradually decreased in all tissues over time. These findings are new, as in the only long-term infection study published so far, a single GKO mouse was used for each day from 22 until 127 DPI, and tissue cysts were observed in the lung and heart and skeletal muscle until 127 DPI, but were not enumerated [3].

Due to the similar, almost identical morphology of tissue cysts of T. gondii and H. hammondi, chronic H. hammondi infections may represent an interesting model for the study of the intra-cyst activity of chemical compounds that might be suitable for the treatment of chronic toxoplasmosis. As the definitive discrimination of H. hammondi from T. gondii is only possible at the encysted bradyzoite stage, and as re-activation of cysts does not seem to occur in this parasite, H. hammondi might be used as a model that could provide information on the true effect of compounds on encysted H. hammondi as a proxy for encysted T. gondii. The Hham-qPCR1 described here may thus help to quantify the effect of treatments. Moreover, Hham-qPCR1, a novel qPCR, may become a valuable tool for the further elucidation of cell biological differences between the usually pathogenic T. gondii and generally non-pathogenic H. hammondi, i.e. between two parasites which are genetically very closely related.

Conclusions

The newly established Hham-qPCR1 assay represents a novel diagnostic tool for epidemiological and cell biological studies on H. hammondi in cats as well as in intermediate hosts.