Eimeria macusaniensis, a large coccidian parasite of New World camelids, is one of the most damaging intestinal parasites in llamas and alpacas.1–8 It has been reported to also affect guanacos9,10 and is presumed to affect vicuñas as well. It has been theorized to be the same organism as Eimeria cameli, an important parasite of Old World camelids,11 although it is slightly different in appearance. Clinical signs of infection include lethargy, weight loss, diarrhea, a decrease in appetite, recumbency, seizures or other neurologic signs, colic, and dyspnea. In South America, E macusaniensis and Eimeria lamae are considered the most pathogenic coccidia, with coccidiosis counted among the leading causes of death in young camelids.2–4,8 In North America, E macusaniensis affects camelids of all ages, with the mortality rate equally high in adults and crias.1 On the basis of the literature,1–10 E macusaniensis infection currently is one of the major health problems of New World camelids.
One of the most difficult aspects concerning treatment of E macusaniensis infections is identifying infection in a timely manner. Clinical signs associated with infection by this parasite are common to many other, better known disorders and hence are frequently mis-identified. Colibacillosis and clostridial disease in particular have become major causes of death in crias, when in fact these bacterial infections often develop secondary to another intestinal process, such as coccidiosis.1,3,8 Additionally, the reported prepatent period for E macusaniensis is at least 32 days and may be as high as 58 days,12,13 far longer than the reported period of 15 to 16 days for E lamae,14 16 to 18 days for Eimeria alpacae,15 and 10 days for Eimeria punoensis.15 Prepatent disease appears to be common with E macusaniensis infection, with some alpacas dying of the disease within 20 days after presumptive ingestion of oocysts, which is as much as 2 weeks prior to appearance of oocysts in the feces of less severely affected herdmates.1 Affected camelids have also been reported in which clinical signs were evident for at least as long as the mean prepatent period and infection was confirmed at postmortem examination but no coccidial oocysts were detected during antemortem fecal flotation.7 Finally, the oocysts are large and dense and are poorly detected by many standard methods of fecal analysis.16
To address the frequency of prepatent disease and failure of standard fecal analytic techniques to identify E macusaniensis during the prepatent period, our research group developed a PCR assay to detect E macusaniensis DNA in feces. The assay detects the ITS-1 within the rDNA. Ribosomal DNA and its ITS regions often exist in thousands of copies and thus require small amounts of sample DNA for PCR amplification. The ITS-1 region has been successfully used in PCR assays to differentiate among Eimeria spp in feces collected from poultry17–19 and rodents.20
Fecal PCR assays for the detection of coccidia and other parasites have been used principally to identify specific species or to simplify the processing of large numbers of samples.17–21 Samples are processed in those situations to release DNA from the oocysts. We hypothesize that such assays could also prove to be useful in identifying prepatent infections. All stages of the parasite should have similar or identical DNA sequences, and it is possible that small amounts of this DNA may appear in feces before the oocysts appear in the feces. This would be especially useful for parasites such as E macusaniensis, which have long prepatent periods and cause prepatent disease. However, successful detection would also rely on free DNA surviving in the fecal environment, which is one of the least hospitable environments for DNA amplification.22,23 Techniques have been developed to improve free DNA recovery from feces and have been used successfully for procedures such as fecal genotyping of wildlife species,24 but their use in detecting free Eimeria DNA has not been reported to our knowledge.
The purpose of the study reported here was to evaluate a PCR assay for use in the detection of E macusaniensis in camelid feces at various stages of infection, including during the prepatent period after experimental inoculation. The goal was to enable early detection of infection, particularly in camelids with few or no oocysts in their feces, and hence allow more timely treatment and ultimately a greater survival rate for infected camelids.
Materials and Methods
Animals—Four adult alpacas (3 gelded males and 1 female; age, 4 to 9 years) from the Oregon State University research herd (all were herd members for at least 3 years) were screened via physical, fecal, and hemato-logic examinations prior to inclusion in the study. This herd was known to have exposure to E macusaniensis, E lamae, and strongyle-type worms on the basis of results of routine periodic fecal examination, but clinical disease attributable to endoparasitism had never been identified in herd members. The alpacas were housed in confinement in groups on concrete floors covered with straw. Fecal material was removed daily. All components of the study were conducted with the approval of the Oregon State University Institutional Animal Care and Use Committee.
Inoculation—Parasites of the family Eimeriidae undergo a self-limiting life cycle. Thus, the alpacas were confined for 62 days and determined to be free of detectable E macusaniensis by the use of fecal exami nation conducted on samples collected 55, 41, 28, 14, 10, and 3 days before inoculation. All fecal analyses were performed via the double-centrifugation sucrose-flotation technique,16 and PCR assays were conducted on the samples collected 55, 41, 14, and 3 days before inoculation. The 62-day period was chosen because it is longer than the longest reported prepatent period (58 days) for this parasite12,13; no E macusaniensis oocysts were found during the period before inoculation.
Each alpaca received an oral inoculum of 2 × 104 sporulated E macusaniensis (day 0). The inoculum was the minimum used in a previous study13 to create patent infection but not to induce clinical signs in naive crias. Oocysts for inoculation were isolated from feces of naturally infected alpacas and llamas by use of the double-centrifugation sucrose-flotation technique, followed by 5 washes in 1 × PBS solution.16 Separation of oocysts from fecal debris and the smaller Eimeria spp was achieved via size exclusion by passing the preparation through 104- and 40-μm mesh filters; E macusaniensis oocysts pass through the larger filter but not the smaller filter, whereas the smaller Eimeria spp pass through both filters. Oocysts were further purified by manual separation via light microscopy at 30 × magnification. Purified oocysts were sporulated by standard incubation in 2% (wt/vol) K2CrO7 for 21 days at 18°C, after which > 90% of the oocysts were confirmed by use of light microscopy to have sporulated.
The inoculation study lasted 7 weeks, which was approximately 1 week longer than the maximum reported prepatent period for E macusaniensis.12,13 After initiation of the study, alpacas were examined daily for signs of poor health. Feces were recovered from the rectum, and the double-centrifugation sucrose-flotation technique and PCR assay were conducted on samples collected weekly for 3 weeks, then twice a week for 4 weeks. Alpacas with signs of illness, including loss of body condition, inappetence, lethargy, severe anemia, or a severe decrease in blood protein concentration, were treated appropriately and removed from the study. At the conclusion of the study, all alpacas with detectable Eimeria spp in their feces were treated with ponazuril (20 mg/kg, PO, q 24 h for 5 days) and kept in confinement until feces were free of detectable oocysts.
DNA extraction—To obtain control DNA, E lamae and E macusaniensis oocysts were isolated from fresh feces obtained from naturally infected alpacas and llamas. Oocysts were isolated by use of the double-centrifugation, sucrose-flotation technique followed by 5 washes in 1× PBS solution.16 Oocysts were purified via size exclusion by use of 104- and 40-μm mesh filters to separate E macusaniensis from Eimeria spp and larger fecal debris. Purified oocysts were then allowed to incubate for 1 to 3 weeks at 18°C in 2% (wt/vol) K2CrO7 until sporulation was confirmed with microscopic examination. Sporulated oocysts were centrifuged and washed 5 times in 1× PBS solution to remove all K2CrO7. For E macusaniensis, the outer shell was mechanically disrupted by use of a ground-glass tissue grinder to release the sporozoites and unsporulated oocysts. Then, DNA was extracted via a commercial DNA extraction kita used in accordance with the manufacturer's protocol, except that the elution-buffer incubation time was increased from 1 minute to 5 minutes to increase DNA yield. The same protocol, without mechanical disruption because of the thinner wall of E lamae, was used for extraction of DNA from sporulated E lamae oocysts. Eimeria lamae were obtained from feces of naturally infected camelids and purified through double-centrifugation sucrose flotation and via size exclusion by washing the concentrated oocysts and separating them with 104- and 40-μm mesh filters to separate the smaller E lamae oocysts from the larger E macusaniensis oocysts. Final preparations were confirmed to be free of E macusaniensis oocysts by use of light microscopy. The DNA content was determined by use of a spectrophotometer,b and DNA was stored at −70°C until used.
For study samples, DNA from (mean ± SD) 0.24 ± 0.01 g of fresh feces was extracted via a commercial kitc used in accordance with the manufacturer's protocol. When possible, a scalpel blade was used to peel away the outer mucoid layer (2 to 3 mm) of fecal material from multiple fecal pellets to help reduce the presence of fecal inhibitors and degraded DNA.24 The resulting filtrate containing DNA was placed in aliquots and stored at −70°C until assayed.
Amplification and sequencing—The ITS-1 regions for E macusaniensis and E lamae were amplified with previously published primers (EF1 and ER1)17 at 50 pmol/50 μL of reaction by use of 1.25 U of Taq poly-merase in a master mixd and 0.1 μg of bovine serum albumin/mL. Reaction cycles consisted of an initial hot start at 95°C for 5 minutes; a denaturing step at 94°C for 2 minutes; then 35 cycles at 94°C for 30 seconds, 55°C for 30 seconds, and 72°C for 30 seconds; and a final extension cycle at 72°C for 5 minutes. Reactions were performed by use of a thermal cycler.e
Amplified products were separated electrophoreti-cally by use of 2% 89mM Tris-borate and 2mM EDTA agarose gels. Fragments were stained with ethidium bromide and visually inspected under UV light. Size of the DNA fragments was confirmed by use of a 50-bp ladder, and selected bands were excised and purified from the agarose gel via a commercial kitf used in accordance with the manufacturer's instructions. Purified products were sequenced at the Oregon State University Center for Genome Research and Biocomputing by use of a DNA sequencer machine.g,h Sequence analysis was performed by use of commercial software.i,j Comparisons of nucleotide sequences were performed by use of the Needleman-Wunsch global alignment algorithm. At least 3 isolates obtained from a minimum of 3 animals from 3 geographic locations within the states of Washington and Oregon were sequenced in both directions for each Eimeria spp, and the consensus sequences were submitted to GenBank.
Nested E macusaniensis–specific PCR assay—Primer sets for E macusaniensis- and E lamae–specific PCR assays were manually designed on the basis of sequences derived by use of conserved primers ER1 and EF1.17 The initial PCR reaction mix and conditions were as described previously, with DNA extracted from feces as a template for primary amplification via the conserved primers. Because amplification of samples by use of the conserved primers (EF1 and ER1) routinely did not yield a visible band, except for samples containing high numbers of oocysts, the conserved amplified product was then reamplified under the same conditions with species-specific primers for E macusaniensis. Primer sequences and expected product sizes were summarized (Appendix).
To test sensitivity and specificity, species-specific primer pairs for E macusaniensis (GenBank accession No. GU570449) and E lamae (GenBank accession No. GQ330537.1) were tested by use of a PCR assay against purified genomic target DNA from each species of coc-cidia. Purity was confirmed by the detection of amplified product bands at the appropriate DNA fragment size for each species when initially amplified by use of conserved primers and further confirmed by the presence or absence of appropriate product bands when further amplified by species-specific primers. The DNA fragment size was confirmed by use of a 50-bp ladder, and sequences were confirmed via forward and reverse sequencing.
The assay detected E macusaniensis DNA in 7 of 7 fecal samples spiked with 100 ng of E macusaniensis DNA and extractedc in accordance with the manufacturer's protocol. Three of 7 samples spiked with 10 ng of DNA and 1 of 9 samples spiked with 1 ng of DNA also had positive results, with unspiked fecal samples yielding no amplified product. Field samples with positive results by use of the double-centrifugation sucrose-flotation technique for E lamae, E alpacae, and E pu-noensis but negative results for E macusaniensis also failed to yield an amplification product when tested with E macusaniensis–specific primers. The sequences of recovered DNA from spiked samples were identical to those for the DNA used to spike the samples.
Fecal examination—Fecal samples were examined by use of the double-centrifugation sucrose-flotation technique.16 In brief, 2 g of feces was mixed in 98 mL of distilled water and allowed to soak overnight at 4°C to assist in liberating parasite ova from the fecal material. The following morning, the fecal slurry was mixed, and 10 mL of the slurry was immediately poured into a conical-bottom 15-mL centrifuge tube and centrifuged at 200 × g for 5 minutes in a swinging bucket centrifuge. Supernatant was decanted, and the pellet was resus-pended in 2 mL of sucrose solution (Sheather's solution without formalin; 3.74M; specific gravity, 1.27). An additional 10 mL of sucrose solution was added, and the sample was centrifuged again for 5 minutes at 200 × g. After the second centrifugation, the tube was filled with sucrose solution until a positive meniscus formed, then a 22 × 22-mm coverslip was placed on top. The coverslip was removed after 60 minutes and placed on a microscope slide.16 All E macusaniensis and small Eimeria oocysts were counted, and the total of each was multiplied by 5 to determine the number of oocysts of each type per gram of feces.
Results
ITS-1 sequences—The ITS-1 sequences for E macusaniensis and E lamae were amplified successfully, and the resulting sequences were submitted to GenBank. The ITS-1 region began at base position 82 and terminated 22 bp before the 5.8S rDNA. On the basis of results obtained by use of the EF1 and ER1 primers,17 the sequence length of ITS-1 was shorter in E macusaniensis than in E lamae (233 vs 267 bp, respectively), with 63% homology between the 2 species (Figure 1).
Schematic depicting the ITS-1 of Eimeria macusaniensis and Eimeria lamae. Conserved regions for primers (EF1 and ER1) are identified with a single underline; species-specific primers are double underlined. Identical bases at the same position for both organisms are indicated by a star. The ITS-1 region begins at base position 82 and ends 22 bp before the end of the sequence. There was 63% homology between the 2 species. Numbers at the right of each line indicate the last base position in that row.
Citation: American Journal of Veterinary Research 73, 1; 10.2460/ajvr.73.1.13
Schematic depicting the ITS-1 of Eimeria macusaniensis and Eimeria lamae. Conserved regions for primers (EF1 and ER1) are identified with a single underline; species-specific primers are double underlined. Identical bases at the same position for both organisms are indicated by a star. The ITS-1 region begins at base position 82 and ends 22 bp before the end of the sequence. There was 63% homology between the 2 species. Numbers at the right of each line indicate the last base position in that row.
Citation: American Journal of Veterinary Research 73, 1; 10.2460/ajvr.73.1.13
Schematic depicting the ITS-1 of Eimeria macusaniensis and Eimeria lamae. Conserved regions for primers (EF1 and ER1) are identified with a single underline; species-specific primers are double underlined. Identical bases at the same position for both organisms are indicated by a star. The ITS-1 region begins at base position 82 and ends 22 bp before the end of the sequence. There was 63% homology between the 2 species. Numbers at the right of each line indicate the last base position in that row.
Citation: American Journal of Veterinary Research 73, 1; 10.2460/ajvr.73.1.13
PCR primer specificity—Each species-specific primer was tested against genomic DNA from purified E macusaniensis and E lamae sporulated oocysts. Primer pairs amplified fragments of the predicted size from their species DNA without cross-species amplification or unexpected bands, which confirmed species specificity (Figure 2). These results were duplicated with test samples from the present study; these samples were confirmed positive for E macusaniensis oocysts but negative for smaller oocysts via fecal flotation. Conversely, a fecal sample from this study that was confirmed negative for E macusaniensis via PCR assay and fecal flotation but positive for E lamae resulted in an amplified product when E lamae primers were used, but no product was seen with the E macusaniensis–specific primer pair.
Agarose gels of species-specific PCR amplification of the ITS-1 region of E macusaniensis and E lamae DNA by use of primers developed for the study reported here after amplification with the genus-specific primers EF1 and ER1. Amplification products are indicated (arrows). Lanes are as follows: M = Molecular weight 50-bp ladder, 1 and 3 = an E macusaniensis isolate (164-bp product), and 2 and 4 = an E lamae isolate (236-bp product). Lanes 1 and 2 are results for E macusaniensis–specific primers, and lanes 3 and 4 are results for E lamae–specific primers. The > 300-bp product apparent in lane 4 is residual template carryover from the initial (conserved primers) amplification product.
Citation: American Journal of Veterinary Research 73, 1; 10.2460/ajvr.73.1.13
Agarose gels of species-specific PCR amplification of the ITS-1 region of E macusaniensis and E lamae DNA by use of primers developed for the study reported here after amplification with the genus-specific primers EF1 and ER1. Amplification products are indicated (arrows). Lanes are as follows: M = Molecular weight 50-bp ladder, 1 and 3 = an E macusaniensis isolate (164-bp product), and 2 and 4 = an E lamae isolate (236-bp product). Lanes 1 and 2 are results for E macusaniensis–specific primers, and lanes 3 and 4 are results for E lamae–specific primers. The > 300-bp product apparent in lane 4 is residual template carryover from the initial (conserved primers) amplification product.
Citation: American Journal of Veterinary Research 73, 1; 10.2460/ajvr.73.1.13
Agarose gels of species-specific PCR amplification of the ITS-1 region of E macusaniensis and E lamae DNA by use of primers developed for the study reported here after amplification with the genus-specific primers EF1 and ER1. Amplification products are indicated (arrows). Lanes are as follows: M = Molecular weight 50-bp ladder, 1 and 3 = an E macusaniensis isolate (164-bp product), and 2 and 4 = an E lamae isolate (236-bp product). Lanes 1 and 2 are results for E macusaniensis–specific primers, and lanes 3 and 4 are results for E lamae–specific primers. The > 300-bp product apparent in lane 4 is residual template carryover from the initial (conserved primers) amplification product.
Citation: American Journal of Veterinary Research 73, 1; 10.2460/ajvr.73.1.13
Experimentally inoculated alpacas—Infection was confirmed by the detection of E macusaniensis oocysts via fecal flotation in 1 alpaca at 31 days after inoculation and 3 alpacas at 35 days after inoculation. No fecal oocysts or other parasites were detected. One alpaca was removed from the study 30 days after inoculation because of clinical signs judged more likely to be related to peer interactions in confinement than to infection with E macusaniensis. None of the other infected alpacas developed clinical signs related to parasite infection.
The alpaca was removed from the study and treated because of the development of lethargy, anorexia, and weight loss. Despite an extensive evaluation, no definitive diagnosis was achieved. In retrospect, it was determined that these signs, particularly the weight loss, were evident prior to inoculation, and hence were not necessarily specifically attributable to the infection. Because of concerns that the treatments, including oral administration of barium sulfate for diagnostic purposes, might have interfered with the PCR assay by changing the fecal environment, posttreatment results were not recorded. This alpaca subsequently had a complete recovery.
None of the alpacas had positive results for the PCR assay prior to inoculation. Three of 4 alpacas developed brief strong positive reactions within 1 week after inoculation. The alpaca that was removed from the study on day 30 had a positive PCR result for feces collected on day 7 but negative PCR results for samples collected on days 14, 21, and 28.; E macusaniensis oocysts were detected via fecal flotation in that alpaca in samples collected on days 35 and 38. The second alpaca had positive PCR results for a fecal sample collected on day 7, negative PCR results for samples collected on days 14, 21, and 28, and positive PCR results for samples collected on days 31, 35, and 38; E macusaniensis oocysts were detected via fecal flotation in that alpaca in samples collected on days 35 and 38. The third alpaca had negative PCR results for fecal samples collected on days 7, 14, and 21 and positive PCR results for samples collected on days 28, 31, 35, and 38; E macusaniensis oocysts were detected via fecal flotation in that alpaca in samples collected on days 35 and 38. The fourth alpaca had positive PCR results for fecal samples collected on days 7 and 14, negative PCR results for samples collected on days 21 and 28, and positive PCR results for samples collected on days 31, 35, and 38; E macusaniensis oocysts were detected via fecal flotation in that alpaca in samples collected on days 31, 35, and 38. In all alpacas, sequences of amplified fragments were identical to the sequences for the E macusaniensis used for inoculation.
Three alpacas developed patent infections but remained in the study. They had positive PCR reactions at 28 or 31 days after inoculation, which was 3 to 7 days before oocysts were detected in feces. All results for PCR assays were positive once oocysts were detected in the feces. Oocyst counts ranged from 10 to 2,000 oocysts/g of feces. Results were repeatable by use of extracted DNA in samples stored at −70°C over a 12-month period. Small coccidian oocysts including E lamae were found in a number of the samples with negative results for E macusaniensis when tested by use of the PCR assay and fecal flotation. The source of these oocysts was unknown, but there was no evidence that they affected test results.
Discussion
On the basis of the findings for the study reported here, the PCR assay for E macusaniensis in fecal samples appeared to be as specific and more sensitive at some stages of infections than was the double-centrifugation sucrose-flotation technique, and the PCR assay was capable of detecting infection within the reported prepatent period. Specificity was established by lack of cross-reactivity with DNA isolated from E lamae or from samples with visible small coccidial oocysts and lack of positive results during the preinfection period. Although both diagnostic methods eventually identified all infected alpacas, the PCR assay clearly detected more infected alpacas during the prepatent period than did the flotation technique. The PCR assay yielded a positive result for all samples in which E macusaniensis oocysts were detected.
The PCR assay appeared to allow successful detection of infections within the reported prepatent period, which was the major goal of this study. Low-level preexisting infections could not be completely excluded, nor could we exclude the passage of inoculum oocysts immediately after inoculation or the early passage of oocysts 28 to 31 days after inoculation, but no oocysts were found via fecal flotation for any of these samples. The simultaneous increase in signal strength 7 to 14 days after inoculation and again before shedding strongly suggested that the PCR assay was detecting non–oocyst-associated DNA at those time points. This could include DNA from sporozoites, merozo-ites, microgametocytes, or other luminal stages of the parasites. These are all thin-walled forms that could potentially release DNA or be passed into the feces. Also, because the alpacas were adults from a herd with known exposure to E macusaniensis, their immune response may have impeded these intermediate stages of the parasite and contributed to their fecal passage. The gap between the early and late detection periods could have represented the period of sexual reproduction and macrogametocyte maturation, but this is speculation.
To our knowledge, a PCR assay has not previously been used to detect free coccidial DNA. The transient nature of the postinoculation signal could be interpreted to limit the clinical value of this test, but it is likely that camelids with infections resulting in clinical disease would be continuously ingesting infective parasites and would also be infected with a larger number of E macusaniensis than were used for the inoculum in the present study. Hence, clinically affected camelids might develop stronger, more continuous reactions.
The positive PCR reactions in samples collected 3 to 7 days before the onset of fecal shedding and before the anticipated minimum prepatent period for this parasite suggested that the PCR assay could yield clinically useful information that would not otherwise be available. Although some camelids infected with E macusaniensis die up to 2 weeks before their herdmates commence shedding oocysts, most clinical disease is evident in camelids shortly before or after the onset of shedding.1 Thus, identifying infection at the onset of shedding, when the PCR assay appeared reasonably sensitive, could justify timelier use of anticoccidial medications and potentially improve the clinical course of infected camelids. Additionally, detection of early infection could allow prophylactic administration of medication. A more extensive clinical trial is necessary to assess the value of the PCR assay for use in detecting natural infections.
One of the initial challenges in the present study was to overcome the effect of inherent PCR assay inhibitors, including humic acids, heme compounds, bilirubin, bile salts, and complex polysaccharides, present in feces.22,23 Although extensive research has been performed to compare various DNA extraction protocols for fecal material, many techniques are ill suited for use in a diagnostic setting because they are time-consuming, are tedious, or were found in preliminary trials to be ineffective for use with camelid feces. The commercial extraction protocol used in the present study involved a combination of chemical lysis and mechanical disruption techniques and was designed for samples with a high humic acid content; this combination appeared to work well for the disruption of the thick outer shell of E macusaniensis in pelleted feces of camelids.
ABBREVIATIONS
| ITS | Internal transcribed spacer |
DNeasy tissue kit, Qiagen, Valencia, Calif.
NanoDrop ND-1000 UV-Vis spectrophotometer, Nanodrop, Thermo Scientific, Waltham, Mass.
PowerSoil DNA isolation kit, MoBio Laboratories Inc, Carlsbad, Calif.
HotStarTaq Plus master mix, Qiagen, Valencia, Calif.
Bio-Rad PTC-200 DNA engine cycler, BioRad Laboratories, Hercules, Calif.
QIAquick gel extraction kit, Qiagen, Valencia, Calif.
BigDye terminator cycle sequencing kit, version 3.1, Applied Biosystems, Foster City, Calif.
3730 DNA analyzer, Applied Biosystems, Foster City, Calif.
ABI Prism 3730 data collection software, version 3.0, Applied Biosystems, Foster City, Calif.
ABI Prism DNA sequencing analysis software, version 5.2, Applied Biosystems, Foster City, Calif.
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Appendix
Conserved and species-specific primers, expected size of amplified product, and ITS-1 length for Eimeria macusaniensis and Eimeria lamae.
| Species | Primer sequence 5′-3′ (primer reference) | Expected product size (bp) | ITS-1 length (bp) | GenBank accession No. |
|---|---|---|---|---|
| Eimeria spp | Forward: aag ttg cgt aaa tag agc cct c (EF1)* | Variable | Variable | † |
| Reverse: aga cat cca ttg cgt aaa g (ER1)* | ||||
| E lamae | Forward: tgc ggc aac ttg cat tga tgc (ELF)† | 236 | 267 | GQ330537.1 |
| Reverse: cta cac caa gaa atg ctc ac (ELR)† | ||||
| E macusaniensis | Forward: ggc cgt ata tta acc aat cc (EmacF3)† | 164 | 233 | GU570449 |
| Reverse: taa tat gaa gat ggg tga ttc c (EmacR3)† |
Annealing temperature for all primers was 55°C.
Primers developed by investigators in another study.17
Primers developed in the study reported here.
= Not applicable.
