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. Author manuscript; available in PMC: 2023 Nov 20.
Published in final edited form as: Ticks Tick Borne Dis. 2021 Oct 10;13(1):101847. doi: 10.1016/j.ttbdis.2021.101847

Direct injection of Amblyomma americanum ticks with Cytauxzoon felis

Tzushan S Yang a, Mason V Reichard b, Henry S Marr a, Leah A Cohn c, Laura Nafe b,c, Nathan Whitehurst d, Adam J Birkenheuer a,*
PMCID: PMC10658644  NIHMSID: NIHMS1944880  PMID: 34673404

Abstract

Cytauxzoon felis is a tick-borne hemoprotozoan parasite that causes life-threatening disease in domestic cats in the United States. Currently, the platforms for C. felis research are limited to natural or experimental infection of domestic cats. This study aims to develop an alternative model by infecting Amblyomma americanum ticks with C. felis via direct injection. Amblyomma americanum adults were injected with C. felis-infected feline erythrocytes through two routes: directly into the digestive tract through the anal pore (IA injection), or percutaneously into the tick hemocoel (IH injection). RNAscope® in situ hybridization (ISH) was used to visualize the parasites within the ticks at different time points after injection. Four months after injection, ticks were divided into 3 infestation groups based on injection methods and inoculum type and fed on 3 naïve cats to assess the ticks’ ability to transmit C. felis. Prior to the transmission challenge, selected ticks from each infestation group were tested for C. felis RNA via reverse transcription-PCR (RT-PCR). In both IA- and IH-injected ticks, ISH signals were observed in ticks up to 3 weeks after injection. The number of hybridization signals notably decreased over time, and no signals were detected by 4 months after injection. Prior to the transmission challenge, 37–57% of the sampled ticks were positive for C. felis RNA via RT-PCR. While the majority of injected ticks successfully attached and fed to repletion on all 3 cats during the transmission challenge, none of the cats became infected with C. felis. These results suggest that injected C. felis remained alive in ticks but was unable to progress to infective sporozoites after injection. It is unclear why this infection technique had been successful for other closely related tick-borne hemoprotozoa and not for C. felis. This outcome may be associated with uncharacterized differences in the C. felis life cycle, the lack of the feeding or molting in our model or absence of gametocytes in the inoculum. Nonetheless, our study demonstrated the potential of using ticks as an alternative model to study C. felis. Future improvement of a tick model for C. felis should consider other tick species for the injection model or utilize infection methods that more closely emulate the natural infection process.

Keywords: Cytauxzoon felis, Direct injection, Tick, In situ hybridization

1. Introduction

Cytauxzoon felis is an important tick-borne apicomplexan hemoparasite that can cause fatal disease in domestic cats in the United States (Birkenheuer et al., 2006; Lewis et al., 2012; Sherrill and Cohn, 2015; Wagner, 1976). Currently, even with the best medical care the mortality rate for acute cytauxzoonosis remains high (Cohn et al., 2011; Wikander et al., 2020). Additionally, the distribution of C. felis appears to be spreading and is no longer limited to the southeast and southcentral U.S. (Cohn and Birkenheuer, 2012; MacNeill et al., 2015). This shift is likely due to the expanding geographic range of the main tick vector, Amblyomma americanum (Raghavan et al., 2019). No vaccine is available for C. felis, and tick control is currently the only means to prevent infection in domestic cats (Reichard et al., 2013, 2019). The greatest barrier facing Cytauxzoon research, including the development of a vaccine, has been the inability to culture the parasite. Therefore, most research has relied on experimental or natural infection of domestic cats (Allen et al., 2019; Joyner et al., 2007; Reichard et al., 2010). There is a critical need to establish cost-effective alternative methods to study C. felis that minimize the use of live animals.

To date, ticks have mainly been thought of as “just a vector” rather than a host that can be used to study C. felis. Ticks typically acquire infective organisms through a blood meal while feeding on live animals. To minimize the usage of live animals for this process, several artificial infection methods have been established and utilized for other tick-borne pathogens (Bonnet and Liu, 2012). One of these methods is direct injection, which is a technique that has been used to experimentally infect ticks with several tick-borne pathogens (Goddard, 2003; Kariu et al., 2010; Kocan et al., 1986; Taank et al., 2020). Importantly, these include piroplasmids that are closely related to C. felis, Theileria and Babesia (Battsetseg et al., 2007; Jongejan et al., 1980; Walker et al., 1979).

The main objective of this study was to develop a model to infect ticks via direct injection with C. felis that eliminates the need for acquisition feeding on cats and subsequent molting. To achieve this goal, we injected A. americanum adults with C. felis, and assessed the viability of the parasites in the ticks as well as the ability of the injected ticks to transmit the infection to naïve cats.

2. Materials and methods

2.1. Tick maintenance

Pathogen-free adult A. americanum ticks were purchased from Oklahoma State University tick rearing facility (Stillwater, OK). Ticks were maintained in conditions recommended by the tick rearing facility with minor modifications. Specifically, ticks were stored in paper cartons in a humidity chamber with 90–99% relative humidity at 20–23°C with a 12 h light/12 h dark photoperiod. Ticks were checked every 1–3 days for mortality and fungal contamination. Paper cartons were changed every 3–4 weeks.

2.2. Inoculum

All blood samples were collected in accordance with North Carolina State University Institutional Animal Care and Use Committee (IACUC) approved protocol 20–254 or were diagnostic samples that were scheduled to be discarded. Blood was collected from privately-owned domestic shorthair cats diagnosed with acute cytauxzoonosis. Approximately 1to 2 ml of infected blood was collected into EDTA-containing tubes and shipped on ice to the author’s laboratory overnight. Upon arrival, thin blood smears were stained with Hema 3 stain (Fisher Scientific, Hampton, NH, USA) and examined via light microscopy for the presence of intraerythrocytic C. felis. The percent parasitemia was determined via light microscopy by counting the number of intraerythrocytic parasites per 1000 erythrocytes. Total DNA was extracted from an aliquot of each sample using a DNeasy Blood and Tissue kit (Qiagen, Valencia, CA, USA) following manufacturer’s instructions. Eluted DNA was tested for C. felis using a species-specific qPCR assay targeting the cox3 gene of C. felis as previously described (Schreeg et al., 2016). Only samples that were positive for both light microscopy (parasitemia 5–8%) and qPCR were used for injection.

Prior to injection, C. felis-infected blood was washed twice with equal volumes of either phosphate-buffered saline (PBS) (Gibco, Gaithersburg, MD, USA) or Alsever’s solution (Alfa Aesar, Haverhill, MA, USA). Packed cell volume was adjusted to 50–60%. PBS-washed inoculum was used immediately for injection. Alsever’s-washed inoculum was either used immediately or stored at 4 °C for up to 10 days until injection.

2.3. Tick injection

One hundred and fifty female and 150 male A. americanum adult ticks were used for injection. Males and females were divided into two groups for two different routes of injection: (1) Intra-anal pore (IA) injection into the tick midgut to mimic the natural route of infection; or (2) intra-hemocoel (IH) injection to bypass the midgut. Capillary needles for injection were prepared using a micropipette puller (Narishige, Amityville, NY). The tips were broken under a dissecting microscope and adjusted to match the size of the injection site. Approximately 5–10 μl of inoculum was then loaded into the capillary needles via microloader pipettes (Eppendorf, Hamburg, Germany).

Prior to injection, all ticks were surface sterilized as previously described (Mani et al., 2012). The ticks were then placed ventral side up on double-sided tape fixed on a clean glass slide, and the anterior 1/3 of the body secured by cellophane tape. For the IA group, the inoculum was injected directly into the tick’s anal pore using a microinjector (pneumatic picopump, WPI) until leakage of the inoculum was observed from the injection site (females ~5 μl, males ~2 μl). For the IH group, the cuticle of the ventral surface of the tick was nicked adjacent to the anal pore with the tip of a tuberculin syringe (27 gauge) and the capillary needle was inserted through the incision for injection until leakage of the inoculum was observed from the injection site. Excess inoculum around the injection site was cleaned with Kim wipes and 70% ethanol. Ticks were then returned to the humidity chamber.

2.4. RNAscope® in situ hybridization

For visualization of C. felis, IA- and IH-injected ticks were randomly selected for histopathology and in situ hybridization (ISH) at 24 h (n = 2), 1 week (n = 8), and 3 weeks (n = 8) post injection. At 4 months post-injection, ticks that tested positive for C. felis via RT-PCR (see Section 2.5.1) were used for ISH (n = 18).

A proprietary antisense probe (Cf-18S-rRNA) designed to target a 77 bp fragment of C. felis 18S rRNA (position 666–742 of GenBank Accession AF399930.1) was developed. RNAscope® assays are optimized for RNA detection and performed under conditions to minimize nuclear penetrance and therefore do not favor denaturation and hybridization of double-stranded chromosomal DNA (Advanced Cell Diagnostics; Newark, CA) (Wang et al., 2012). To validate the specificity of this probe, we used positive controls consisting of formalin-fixed paraffin-embedded tissues from cats diagnosed with acute cytauxzoonosis, and negative controls consisting of ticks injected with feline blood that tested negative for the presence of C. felis.

To prepare ticks for histopathology, ticks were first placed in 70% ethanol for 6–12 h, and then transferred to Bouin’s solution (StatLab, Columbia, MD, USA) and placed in a pressurized vacuum for one hour to facilitate fixative penetration. After fixation, ticks were washed in 70% ethanol for 5 min, and embedded ventral side up in molten Histogel™ until solidification. Ticks were then placed in tissue cassettes for paraffin embedding and sectioning (5 μm thick). Three sections per tick were stained with hematoxylin and eosin (H&E) and 3 unstained sections were used for ISH. ISH was performed using RNAscope® 2.5 HD red assay kits according to manufacturer’s instructions after optimizing pretreatment times for tick samples (10 min for target retrieval and 30 min for protease plus treatment).

2.5. Cytauxzoon felis detection of injected ticks

An overview of the transmission challenge is depicted in Fig. 1. Injected ticks were divided into 3 infestation groups based on injection methods (IA vs. IH) and inoculum type (washed with PBS vs. Alsever’s solution).

Fig. 1.

Fig. 1.

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Experimental design of tick C. felis transmission challenge. IA: Intra-anal pore injection; IH: Intra-hemocoel injection; PBS: Phosphate-buffered saline; RT-PCR: Reverse-transcriptase polymerase chain reaction.

2.5.1. RT-PCR prior to transmission challenge

Four months after injection, ticks were randomly sampled from each infestation group (group 1, n = 8; group 2, n = 14; group 3, n = 14) for C. felis two-step RT-PCR. Briefly, each tick was surface sterilized (Mani et al., 2012) and cut in half with a new scalpel blade on a clean glass slide. Total RNA was extracted from one half of the tick using an RNeasy Plus Micro kit (Qiagen, Valencia, CA, USA) according to the manufacturer’s instructions. Ten μl of eluted RNA was used to synthesize complementary DNA (cDNA) with Taqman Reverse Transcription Reagents (Applied Biosystems, Foster City, CA) following manufacturer’s instructions. One μl of cDNA was then used as template in a C. felis-specific real-time PCR assay targeting a 284 bp fragment of the C. felis 18S rRNA (Schreeg et al., 2016). This assay was selected as 18S rRNA is likely to be transcribed throughout all C. felis life stages. DNA from C. felis-positive cats were used as positive control for each set of reactions. Negative controls included cDNA from sham-injected ticks, RNA samples (no RT controls) and water (no template controls). All reactions were performed in CFX96™ Real-Time PCR Detection System combined with C1000™ or MiniOpticon Thermal Cycler (Bio-Rad, Hercules, CA 94,547, USA). Amplification and melt curves were analyzed via CFX96™ Manager software. Amplified products were randomly selected for sequencing (Genewiz, South Plainfield, NJ, USA) and analyzed bi-directionally with Geneious Prime (San Diego, CA, USA).

2.5.2. qPCR after transmission challenge

All ticks that were attached were removed at day 21 post-infestation. Genomic DNA was extracted from these ticks using a QIAamp DNA mini kit (Qiagen) according to manufacturer’s tissue extraction protocol with slight modifications. Briefly, male and non-engorged female ticks were cut in half with a clean scalpel blade, frozen in liquid nitrogen for 5–10 min and ground into fine powder using a pellet pestle (Fisher Scientific) within a 1.5 ml conical bottom microcentrifuge tube (Eppendorf). For digestion, 250 μl of ATL lysis buffer and 25 μl proteinase K were used at 56 °C overnight (12–18 h). Engorged females were similarly cut in half. These half ticks were homogenized using Bead Mill 4 (Fisher Scientific) for 90 s at 5 m/s after the addition of 500 μl of sterile PBS and five 3.5 mm steel beads (Biospec Inc., OK). This homogenate was digested overnight (12–18 h) at 56 °C in 500 μl of ATL buffer and 50 μl of proteinase K. For males and non-engorged females, 250 μl of AL buffer was added to each tick lysate. For engorged females, 500 μl of AL buffer was added to each tick lysate. Total DNA was then extracted from these lysates according to manufacturer’s instructions. DNA samples were then tested using a C. felis-specific cox3 qPCR assay (Schreeg et al., 2016). Plasmids containing C. felis cox3 inserts were used as positive controls for each set of reactions. DNA extracted from a non-injected unfed A. americanum female tick and water were used as negative controls. Amplification and melt curves were analyzed via CFX96™ Manager software. Amplicons were visualized in an Agarose gel after electrophoresis to confirm the correct amplicon size. Fourteen representative amplicons were submitted for bidirectional sequencing (Genewiz) and analyzed with Geneious Prime.

2.5.3. Comparison of C. felis detection pre- and post-transmission challenge

Prevalence of C. felis in injected ticks was estimated according to Bush et al. (1997) Ninety-five percent confidence intervals were calculated using QuickCalcs (2017). Proportions of C. felis infected ticks were compared using a chi-square tests or Fisher’s Exact tests (Sokal and Rohlf, 1995). A Fisher’s exact test was used if 20% of the expected values in contingency tables were less than 5. If there was not statistical difference between pre- and post-infestation of C. felis in ticks, data were combined for each group to compare injection routes. Significance level was set at 0.05.

2.6. Tick transmission challenge

Approximately 30 male and 30 female ticks were randomly selected from each infestation group and shipped to Oklahoma State University for the transmission challenge. Transmission feeding on naïve cats was performed as previously described (Reichard et al., 2010, 2009; Thomas et al., 2018) using procedures approved through Oklahoma State University IACUC. Tick attachment was estimated on day 3 and day 7 post-infestation, and unattached ticks were removed. All attached ticks were fed to repletion or removed by day 21 post-infestation and stored in 70% ethanol. Cats were monitored daily for signs of acute cytauxzoonosis, i.e. fever, lethargy, depression, inappetence and hyporexia.

2.7. C. felis PCR of naïve cats during transmission challenge

Whole blood was collected into EDTA tubes on days 0, 3, 7, 13, 17, 21, and 30 post infestation from each cat. Thin blood smears were stained with Hema 3 stain and screened for parasites via light microscopy. Total DNA was extracted as described above in Section 2.2. Each sample was then tested in ten replicate reactions using a qPCR assay targeting C. felis cox3 mitochondrial DNA. This assay was selected as it is more sensitive for the detection of early C. felis infection compared to 18S ribosomal DNA (Schreeg et al., 2016).

3. Results

3.1. Detection of C. felis ISH signals over time in ticks after injection

The ISH probe (Cf-18S-rRNA) is specific for C. felis and does not cross react with ticks injected with C. felis-negative feline erythrocytes (Fig. 2).

Fig. 2.

Fig. 2.

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Cytauxzoon felis 18S ISH probe is specific to infected feline cells with no cross-reactivity with negative feline erythrocytes or tick cells. (A) Positive control: H&E stained feline lung section with intravascular C. felis schizont-laden leukocytes (arrow) adhered to vascular endothelium. 40X. (B) Serial section of 2A probed with Cf-18S-rRNA showing robust hybridization signals (red) of C. felis schizonts. (C) Negative control: H&E stained tick injected with C. felis-negative feline erythrocytes through the anal pore into the midgut (MG) and rectal sac (RS). MG and RS are filled with feline erythrocytes. 20X. (D) Serial section of 2C probed with Cf-18S-rRNA showing no detectable hybridization signals in the injected feline erythrocytes or the surrounding tick cells.

Cytauxzoon felis hybridization signals were detected in ticks 24 h after injection with C. felis-positive inoculum. The number of hybridization signals gradually decreased over time after injection for both IA-and IH-injected ticks (Figs. 3 and 4) until they were undetectable at 4 months. Subjectively, signal intensity did not differ between inoculum types. Positive hybridization signals frequently correlated with areas where feline erythrocytes were present and C. felis signals were never detected within tick cells (Fig. 5).

Fig. 3.

Fig. 3.

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RNAscope® ISH assay in ticks after IA injection with C. felis: hybridization signals decreased over time within the ticks after injection. Thin arrows point to representative hybridization signals. (A,B) 24 h post-injection: numerous C. felis hybridization signals (red) were seen within the midgut lumen and scattered in the hemocoel (C,D) 1 week post-injection: the number of hybridization signals are no longer detected within the midgut lumen and were decreased in numbers in the hemocoel. (E,F) 3 weeks post-injection: hybridization signals (arrow) were only seen sporadically in the rectal sac and hemocoel. (G,H) 4 months post-injection (immediately prior to transmission challenge): no hybridization signals were detectable. PI: post-injection; MG: midgut; SG: salivary gland; T: tracheae; RS: rectal sac.

Fig. 4.

Fig. 4.

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RNAscope® ISH assay in ticks after IH injection with C. felis: hybridization signals decreased over time within the ticks after injection. Thin arrows point to representative hybridization signals. (A,B) 24 h post-injection: numerous C. felis hybridization signals (red) were seen within the hemocoel. (C,D) 1 week post-injection: the number of hybridization signals decreased in numbers in the hemocoel. (E,F) 3 weeks post-injection: hybridization signals (arrow) were only seen regionally in the hemocoel. (G,H) 4 months post-injection (immediately prior to transmission challenge): no hybridization signals were detectable. PI: post-injection; MG: midgut; SG: salivary gland; T: tracheae.

Fig. 5.

Fig. 5.

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Cytauxzoon felis hybridization signals within the ticks correspond with areas where injected feline erythrocytes are present. (A) H&E stained tick 1 week after IH injection with C. felis-infected feline erythrocytes. Numerous feline erythrocytes maintained its silhouette within the hemocoel (inset: higher magnification of asterisk area) (40X) (B) Serial section of 5A when C. felis 18S ISH probe is applied. Hybridization signals within the hemocoel correlated with the presence of feline erythrocytes (inset: higher magnification of asterisk area). MG: Midgut; IH: Intra-hemocoel injection; ISH: in situ hybridization.

3.2. Detection of C. felis by RT-PCR and PCR in ticks after injection

Four months after injection and prior to the transmission challenge, 36 ticks were tested for C. felis RNA: 3 of 8 ticks (37.5%; 13.5–69.6%) from cat infestation group 1 tested positive; 7 of 14 ticks (50%; 26.8–73.2%) from cat infestation group 2 tested positive; and 8 of 14 ticks (57.1%; 32.6–78.7%) from cat infestation group 3 tested positive. All positive samples matched the melting temperature of the positive control (85 ± 0.5 °C). C. felis was not detected in any negative control reactions, including the no-RT controls.

Tick attachment rates were adequate to excellent in all 3 infestation groups, ranging from 60 to 96% by day 7. At day 21 after tick infestation, 128 attached ticks were removed and tested for C. felis DNA by PCR: 10 of 33 ticks (30.3%; 17.3–47.5%) removed from cat 1 tested positive; 23 of 42 ticks (54.8%; 39.9–68.8%) removed from cat 2 tested positive; and 24 of 53 ticks (45.3%; 32.7–58.6%) removed from cat 3 tested positive. All positive samples had a melting temperature and amplicon size that matched the positive control. Representative amplicon sequences (n = 14) were confirmed to be C. felis (100% identity, GenBank accession KC207821). Cytauxzoon felis was not detected in any negative control reactions.

The proportion of the positive ticks detected between pre- and post-infestation did not significantly differ in any group (group 1: p-value = 0.69 [Fisher’s Exact test]; group 2: X2 = 0.0000, df = 1, p = 1.000; group 3: X2 = 0.239, df = 1, p = 0.625). Similarly, the proportion of positive ticks among groups did not differ statistically (X2 = 4.496, df = 2, p = 0.106).

3.3. Cytauxzoon felis transmission challenge

Cytauxzooon felis infection was not documented in any of the naïve cats throughout the transmission challenge. All 3 cats tested negative for C. felis by PCR prior to the challenge. The cat exposed to infestation group 1 ticks developed no signs of cytauxzoonosis. Fever was the only sign observed in the other 2 cats. For the cat exposed to infestation group 2 ticks, a fever (39.3 °C, 102.7 °F) was noted on day 13 post-infestation. The cat exposed to infestation group 3 ticks, fever was noted on both days 13 (39.6 °C, 103.3 °F) and 14 (39.7 °C, 103.4 °F) post-infestation. The fevers in both cats resolved without medical intervention. Cytauxzoon felis was not detected by PCR or microscopy in any blood samples collected throughout the transmission challenge.

4. Discussion

Since its discovery over 40 years ago in the United States, Cytauxzoon felis continues to be a significant cause of life-threatening illness in domestic cats living in enzootic areas (Cohn and Birkenheuer, 2012; Wagner, 1976; Wikander et al., 2020). The lack of an in vitro culture system for C. felis significantly limits our ability to study the parasite. Current experimental models require both acquisition and transmission feeding on domestic cats. Direct injection is an established technique that has resulted in successful infection of ticks for other pathogens and eliminates the need for acquisition feeding (Bonnet and Liu, 2012; Goddard, 2003; Kariu et al., 2010; Taank et al., 2020). Most notably, ticks that were injected with Theileria or Babesia, the two closest relatives of C. felis, were successfully infected and were able to transmit the infections to naïve animals (Battsetseg et al., 2007; Jongejan et al., 1980; Walker et al., 1979). Our goal was to infect A. americanum ticks with C. felis via direct injection.

In this study, A. americanum adults were injected with C. felis but failed to transmit this infection to naïve cats. It remains unclear whether or not these ticks actually became infected with C. felis after direct injection. Our ability to detect C. felis RNA transcripts in these ticks 4 months after injection suggests the presence of living C. felis. However, we were unable to determine whether these RNA transcripts indicated infection of these ticks with C. felis or just the persistence of erythrocytic life stages from the initial injection. The erythrocytic life stages of C. felis are presumed to be long-lived and can be detected in chronically infected cats for years (Allen et al., 2019; Kier et al., 1982). While the exact lifespan of these C. felis erythrocytic life stages has not been studied, the erythrocytic life stages of other closely related hemoprotozoa, like Babesia and Plasmodium sp., are resilient and able to survive up to 60 days in vitro (Cursino-Santos et al., 2014; Gebru et al., 2017). In this study, C. felis ISH signals were often detected within areas where erythrocytes were present and were never observed within tick cells. Therefore, it is possible that the ticks never became infected and C. felis organisms simply survived for months after injection. Other potential explanations for the failure of parasite development within these ticks include decreased infectivity of the inoculum for ticks or the absence of feeding and molting in the model leading to subsequent lack of stimuli for C. felis to infect tick cells and advance to the next life stages.

It is possible that the shipping and storage conditions (4 °C) may have diminished the number of viable parasites within the inoculums or affected the parasites’ ability to develop within ticks. While the effects of standard blood storage on C. felis viability have not been studied extensively, the authors have used C. felis-infected blood samples that were shipped and stored in similar conditions to successfully infect naïve cats via intravenous inoculation. In a T. parva infection study, inoculum was stored at 4 °C for up to 3 days prior to injection and was still able to successfully infect ticks (Jongejan et al., 1980). Also, Babesia spp. and Plasmodium spp. remain viable and infectious to vertebrate hosts via intravenous inoculation after 30 days in cold storage at 4°C (Cursino--Santos et al., 2014; Goheen et al., 2016). Unfortunately, the viability of our inoculums was not assessed prior to tick injection. C. felis RNA transcripts were not assessed by RT-PCR as these inoculums were not stored in an RNA-stabilizing solution. We also did not attempt to infect naïve cats with these inoculums directly via intravenous injection. As mentioned above, the detection of RNA transcripts in C. felis organisms in ticks after injection is strongly suggestive of live parasites in the inoculums. Based on our experience and previous experiments with related organisms, we believe it is highly likely that our inoculums contained viable parasites, but they either failed to infect the ticks or were unable to complete their life cycle.

Another potential explanation for the lack of parasite development after injection may be the absence of gametocytes in the inoculum. In other closely related piroplasmids, only a small proportion of intraerythrocytic merozoites transform into gametocytes and undergo gametogenesis in the tick midgut lumen once they are ingested (Jalovecka et al., 2018; Mehlhorn and Schein, 1984). For Babesia and Theileria, intraerythrocytic gametocytes are usually indistinguishable from other asexual erythrocytic stages without electron microscopy (Mehlhorn et al., 1980; Mehlhorn and Schein, 1984; Rudzinska et al., 1983). Unlike these parasites, gametocytes have not been described in C. felis. In a cat that is acutely infected with C. felis, merozoites enter erythrocytes after being released from schizont-laden leukocytes. These newly released merozoites may not have had adequate time to become capable of transforming into gametocytes. The duration of time necessary for this process to occur in C. felis has never been studied. For Theileria spp., experimentally infected calves with acute clinical disease (10–19 days after tick infestation or sporozoite inoculation) and high parasitemia have been used to successfully infect ticks (Walker et al., 1979; Warneckel et al., 1980; Watt et al., 2001; Watt and Walker, 2000). Plasmodium falciparum can take up to 12 days to fully mature from a merozoite into a gametocyte (Sinden, 2009). All inoculums used in this study were collected from cats with acute cytaxuzoonosis, however we were unable to determine the exact time lapse between tick infestation and onset of clinical signs as these were naturally occurring infections. In experimental tick transmission studies for C. felis, it takes 8–14 days after tick infestation for infected cats to develop clinical signs and intraerythrocytic stages were first detected via microscopy 16–18 days after infestation (Allen et al., 2019; Reichard et al., 2009; Thomas et al., 2018). Based on these data and our clinical experience, our assumption is that the cats we collected inoculums from were likely to have been infected via tick bite approximately 18–25 days prior to their clinical presentation. Despite our documentation of many intraerythrocytic parasites in our inoculums, it is uncertain whether any of these were gametocytes that were capable of infecting the ticks.

It is also possible that tick feeding, which is absent in our injection model, is required for the development of C. felis. Feeding may trigger critical signals for C. felis to continue its development within the tick’s midgut. Such triggers may include xanthurenic acid (XA), decreased oxygen tension, and a lowered ambient temperature, all of which have been used to induce gametogenesis in vitro for other piroplasmids (Jalovecka et al., 2016; Maeda et al., 2017; Mosqueda et al., 2004). In the current study, we presumed the injected organisms were exposed to decreased oxygen tension in the hemocoele or midgut after injection. Also after injection the ticks were housed at lowered ambient temperature (20–23 °C). While the role of XA in C. felis gametogenesis has not been studied, it could be an important element lacking in our model. XA is a molecule that is crucial in facilitating gametogenesis for Plasmodium spp. (Billker et al., 1998; Garcia et al., 1998). Importantly, increased XA concentrations are directly related to feeding as it is primarily produced by the salivary glands and secreted in the saliva which is ingested into the mosquitoes midgut during a blood meal (Matsuoka et al., 2007; Okech et al., 2006). If XA or other tick-derived molecules play a similar role in the process of gametogenesis for C. felis, infection studies that include feeding or the addition of XA to the inoculum prior to injection should be considered in future studies.

Additionally, it is possible that the molting process is essential for C. felis to continue its life cycle within the ticks, and the use of adult ticks in our model prohibited parasite development after injection. Successful injection studies for T. parva included molting. Jongejan et al. (1980) and Walker et al. (1979) injected engorged Rhipicephalus appendiculatus nymphs with infected bovine erythrocytes percutaneouly into the hemocoel or midgut. These T. parva-injected nymphs were then allowed to molt into adults that were able to successfully transmit the infection to naïve calves. However, the molting process may not be a necessary step for all piroplasmids. In another injection study for Babesia gibsoni and Theileria equi, a soft tick (Ornithodoros moubata) that is not believed to be a natural vector for either organism was selected for injection. Both nymphs and adult stages of O. moubata were directly injected in the hemocoel with B. gibsoni- or T. equi-infected erythrocytes (collected from experimentally infected dogs and in vitro culture, respectively). Both parasites were then able to successfully develop after injection without the process of molting and were detected in the ovaries and eggs. Notably, B. gibsoni was able to develop into infective sporozoites within the salivary glands and these ticks successfully transmitted the infection to naïve dogs (Battsetseg et al., 2007). Future injection studies could include either the molting process or utilize alternative tick species, such as soft ticks or Dermacentor variabilis.

An additional limitation in this study is the small number of cats subjected to the transmission challenge. In previous C. felis transmission studies using similar numbers of acquisition-fed ticks, 85–100% of naïve cats became infected after transmission challenge (Allen et al., 2019; Reichard et al., 2010; Thomas et al., 2018). In these experiments, C. felis was detected in 4–20% of the acquisition-fed ticks (Reichard et al., 2013, 2010) (Reichard, personal communication). Comparatively, we detected C. felis in 30–57% of the ticks we injected. One potential explanation for our higher detection level is the difference in sensitivity of molecular assays used. In our study, we used RT-PCR and qPCR assays targeting 18S rRNA and cox3 DNA of C. felis, while the previous studies utilized a conventional nested PCR targeting 18S rDNA (Reichard et al., 2013, 2010). Head-to-head comparisons of these assays have not been performed. Despite the higher detection levels in the current study, it is still possible that the transmission rate of injected ticks is lower than that of acquisition-fed ticks, and the number of cats used in this study was not sufficient.

Another potential factor affecting the outcome of our study is the interval between tick infection and transmission. The optimal infection-to-transmission interval has never been evaluated for C. felis and A. americanum. In previous experiments using acquisition-fed ticks, transmission challenges on naïve cats occurred 6–18 weeks after acquisition feeding (i.e. 2–12 weeks after molting) (Reichard, personal communication). Our transmission challenge occurred in a similar timeframe (16 weeks after injection). The time required for C. felis to develop within the ticks after injection is unknown, and it remains unclear whether the 16-week interval influenced the ticks’ ability to transmit the infection. To our knowledge, the only other injection study where adult ticks were used, the investigators injected adult O. moubata with B. gibsoni and the time lapse between injection and transmission challenge was not clearly specified (Battsetseg et al., 2007).

Even though this experiment was unsuccessful in transmitting C. felis to cats, we demonstrated the potential to study C. felis using a tick model. Our ability to detect C. felis in whole tick sections using ISH illustrates this as a promising platform to investigate the life cycle of C. felis in ticks. Additionally, as the injected parasites were able to remain alive for an extended period within the tick, this model could potentially serve as a source to initiate in vitro cultivation. To further refine and improve a tick infection model for C. felis, forthcoming studies should characterize the C. felis life cycle within the vertebrate and tick hosts, including the duration of time needed for merozoites to transform into gametocytes and when those parasites would become infectious to the tick host. These studies could assist with the timing and preparation of blood inoculums. Additionally, models could more closely emulate the process of natural infection (feeding and molting), attempt transmission with a larger number of cats, and determine the optimal interval between tick infection and transmission.

Acknowledgments

The authors would like to thank the technicians in the NCSU histology laboratory and Vector-borne Disease Diagnostic Laboratory for their technical support. We would also like to thank all the veterinarians who volunteered to participate in the study and supplied clinical blood samples.

Funding

This research is partially funded by Morris Animal Foundation (Grant ID: D21FE-805, tick injection, in situ hybridization, and molecular reagents) and a charitable organization which wishes to remain anonymous. Stipend support for T.S. Yang was provided by the NIH T32OD011130.

Footnotes

CRediT authorship contribution statement

Tzushan S. Yang: Writing – original draft, Writing – review & editing, Investigation, Project administration. Mason V. Reichard: Investigation, Resources, Writing – review & editing. Henry S. Marr: Investigation. Leah A. Cohn: Resources. Laura Nafe: Resources. Nathan Whitehurst: Methodology. Adam J. Birkenheuer: Supervision, Writing – review & editing.

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