Study population
This study was conducted from January to December of 2018 in central Kentucky, USA, a warm temperate, fully humid, hot summer climate (Cfa), based on the Köppen-Geiger climate classification system. Horses from the University of Kentucky’s anthelmintic naïve parasitology herd were evaluated under the University of Kentucky’s Institutional Animal Care and Use Committee protocol 2012–1046. This closed herd had not received anthelmintics and has been located on the same grazing pasture since 1979 [30]. Horses were provided free access to pasture, hay, and mineral blocks, and were fed a ration balanced grain supplement over the course of the study. Two age groups were defined within this herd: a mature horse population with ages of 7–18 years (n = 19, 18 mares and one stallion) and foals born into the herd in 2018 and followed until necropsy at 4–8 months of age. All mares were bred in the field via live cover in 2017. Foals (n = 14; 9 fillies, 5 colts) were born between April and July 2018, and 13 of these were euthanized either at 4–6 months of age (n = 9) or at 6–8 months of age (n = 4).
Fecal and serum sample collection
Samples were collected biweekly from all mature horses and weekly/biweekly from the foals, depending on the behavior of the individual animals. Fecal samples were collected from fresh droppings or manually from the rectum. Serum samples were collected by venipuncture of the jugular vein(s) of each animal. Fecal samples were immediately packaged, labeled, and stored at 4 °C for further laboratory processing, and serum samples were stored at − 80 °C until processing.
Fecal egg counts
FECs were determined in triplicate using the Mini-FLOTAC method as described previously [31], using a saturated glucose-NaCl flotation medium (specific gravity of 1.25) and with a multiplication factor of five eggs per gram (EPG). Specimens were evaluated for strongylid, Parascaris spp., and S. westeri type eggs. However, due to an abundance of S. westeri in the foal samples, S. westeri egg counting was discontinued after a threshold of 5000 EPG was reached.
Coproculture and larval strongylid differentiation
Individual coproculture was carried out for all mature horses upon each collection, using 10 g of fecal matter and an equal amount of vermiculite to promote water retention and aeration, as described previously [32]. Samples were incubated at room temperature for 14 days and then placed in a Baermann apparatus for an additional 2 days. The entire sediment was harvested, and larvae were stored at 4 °C until identification. For identification, larvae were transferred to nematode counting slides and subsequently heat-inactivated by placing the nematode counting slide on a warming plate at 60 °C for 5 min, or until small bubbles formed. Samples were then examined microscopically at ×10 magnification, and third-stage larvae were counted up to 500. Larvae were identified to genus and/or species using a published key [33].
Strongylus vulgaris qPCR
Eggs were isolated from fecal samples via egg isolation methods described previously [34]. Briefly, 10 g of feces was mixed with 50 ml of tap water. The mixture was strained through a two-ply cheesecloth (18 × 36 in., grade 10 mesh, Fisher Scientific, Waltham, MA, USA) and centrifuged for 10 min at 300×g. The pellet was then suspended in 50 ml of glucose-NaCl flotation solution (SG = 1.25) and centrifuged again at 300×g for 10 min. The supernatant containing the eggs was then subjected to straining through a series of filters arranged by filter size (largest to smallest: 200 μm, 100 μm, and 27 μm) (pluriSelect Life Science, Leipzig, Germany). Isolated eggs were then retrieved from the 27 μm filter and stored in 100% EtOH at −20 °C until DNA isolation. Ethanol was removed from egg samples prior to DNA extraction via evaporation. The Quick DNA Fecal/Soil Microbe kit (Zymo Research, Irvine, CA, USA) was used to isolate DNA from egg samples according to the protocol provided by the manufacturer. Real-time polymerase chain reaction (qPCR) assay was used to identify S. vulgaris DNA in isolated eggs as described previously [35]. A mixture of Brilliant II QPCR Master Mix (Agilent Technologies, Santa Clara, CA, USA) and 4 μl of extracted DNA was used, and the temperature and cycles were set as described previously [35]. Primers and probes were obtained from Integrated DNA Technologies (Skokie, IL, USA), and concentrations were set as described previously [35]. Positive controls for the assay were obtained from an adult morphologically identified S. vulgaris specimen. For statistical analysis, all negative PCR reactions were recorded as a cycle threshold (Ct) of 100.
Strongylus vulgaris ELISA
Serum samples were measured for concentration of antibodies specific to the SvSXP antigen produced by migrating S. vulgaris larvae using an enzyme-linked immunosorbent assay (ELISA) as described previously [36]. All samples were diluted 1:50 with phosphate-buffered saline solution with Tween (PBST) (1:10 dilution), and a secondary antibody of horseradish peroxidase conjugated to goat anti-horse IgG(T) (Bethyl Laboratories, Inc., Montgomery, TX, USA) at a dilution of 1:40,000 was added to each well. Results were normalized as a percentage of the positive control to reduce inter-assay variability [36]. The positive control sample was obtained from a horse known to be infected with S. vulgaris.
Necropsy procedures
The foals were divided into two groups based on a previously reported biphasic appearance of ascarid worm burdens and egg shedding [37, 38], with nine foals (7 fillies, 2 colts) euthanized at the presumed peak ascarid burden age (4–6 months) and four foals (1 filly, 3 colts) euthanized at 6–8 months of age. Necropsy procedures followed previously published principles [39], and the following parasite species and stages were collected and enumerated. Attempts were not made to obtain specimens of S. westeri from the small intestines, as egg shedding patterns had indicated that populations of S. westeri were eliminated prior to euthanasia.
Cyathostomins
Encysted larval stages were enumerated for each of the large intestinal sections—cecum, ventral colon, and dorsal colon—using a mucosal digestion technique as described previously [40], and larvae identified to either early third stage (EL3) or developing stages [late third-stage (LL3) and mucosal fourth-stage larvae (L4)]. A multiplication factor of 500 per organ was used to estimate the total mucosal burden of each organ.
Luminal cyathostomin stages were enumerated for each large intestinal organ by examining a subsample representing a 1% aliquot of the total content volume and subsequently multiplying the count by 100 to estimate the total luminal cyathostomin burden for each foal.
Strongylus spp.
The cranial mesenteric (CMA) and celiac arteries were dissected, and migrating stages of S. vulgaris were collected and morphologically identified to stage (L4 and L5) and sex (L5 only). Similarly, the ventral abdominal walls and peri-renal fat tissues were inspected and dissected for the presence of migrating S. edentatus larvae, which were identified to stage (L4 and L5) and sex (L5 only) as well. Intestinal Strongylus spp. specimens were collected from the cecum, ventral colon, and dorsal by visual inspection of mucosal walls for attached parasites and macroscopical examination of the entire intestinal content.
Other parasites
Parascaris spp. specimens were collected by visual inspection of the entire small intestinal tract and by examining the intestinal contents from the large intestines. Specimens were identified to L4 or L5 (male and female), where appropriate. Ascarids in this documented herd have been karyotyped previously and identified as P. univalens [41], but karyotyping was not carried out in this study, so findings are reported as Parascaris spp. herein. Anoplocephala perfoliata were collected by inspection of the cecal mucosal walls and examination of the intestinal contents from the large intestines.
Statistical analyses
McNemar’s test was run comparing the qualitative results obtained with the coproculture and qPCR using online software (www.graphpad.com). All statistical models were run with SAS University edition software (Cary, NC, USA), and were analyzed with generalized mixed linear models using the glimmix procedure, with a Gaussian distribution assumed. Akaike information criterion and Akaike information criterion corrected were used to assess model fit. JMP Pro 14 software (Cary, NC, USA) was used to create figures and estimate confidence intervals and correlations. Associations of all measured parameters for each analysis was evaluated using traditional backward and forward elimination of variables. All variables with P-values < 0.25 were kept in the model. When variables were significant, a least-square means for Tukey’s pairwise comparison, odds ratio, and estimate were all performed, and interpretation of results carried out at a significance level α = 0.05.
Mature horses
Mature horses were categorized based on age, sex, egg shedding category, and parturition status. At each collection time point, horses were assigned to the following egg shedding categories: low shedders (0–99 EPG), moderate shedders (100–499 EPG), and high shedders (> 500 EPG). Furthermore, the horses were assigned to one of two age categories: 7–11 years (n = 7) and 12–18 years (n = 12). Each sample collection time point was assigned a corresponding collection number (1–24) and season: winter (December–February), spring (March–May), summer (June–August), and autumn (September–November). At each time point, mares were assigned a parturition status of either 0 (no foal) or 1 (foal). Models were constructed with FEC, ELISA, PCR, and coproculture results (% S. vulgaris and total S. vulgaris larvae counted) as outcome variables and with age, season, collection date, and parturition as input variables. “Horse_ID” was kept as a random effect, and repeated measures used with collection date, where appropriate.
Foals
In foals, S. vulgaris ELISA, FEC (strongylid, ascarid, S. westeri), and worm counts (S. vulgaris, S. edentatus, cyathostomins, Parascaris spp.) were analyzed as output variables. In addition to the ELISA values determined at each time point, a foal/mare ELISA ratio variable was created for each foal at each time point as the ratio between the ELISA value in the foal at the time of determination divided by the ELISA value of the corresponding dam at the time of parturition. Models were constructed analyzing these output variables with age, sex, or time point evaluated as input variables. “Horse_ID” was kept as a random effect, and repeated measures used with collection date, where appropriate.