Effect of the nematophagous fungus Pochonia chlamydosporia on soil content of ascarid eggs and infection levels in exposed hens

Effect of the nematophagous fungus Pochonia chlamydosporia on soil content of ascarid eggs and... Background: The nematophagous fungus Pochonia chlamydosporia can degrade ascarid (e.g. Ascaridia galli) eggs in agar and soil in vitro. However, it has not been investigated how this translates to reduced infection levels in naturally exposed chickens. We thus tested the infectivity of soil artificially contaminated with A. galli (and a few Heterakis gallinarum) eggs and treated with P. chlamydosporia. Sterilised and non-sterilised soils were used to examine any influence of natural soil biota. Methods: Unembryonated eggs were mixed with sterilised (S)/non-sterilised (N) soil, either treated with the fungus (F) or left as untreated controls (C) and incubated (22 °C, 35 days) to allow eggs to embryonate and fungus to grow. Egg number in soil was estimated on days 0 and 35 post-incubation. Hens were exposed to the soil (SC/SF/ NC/NF) four times over 12 days by mixing soil into the feed. On day 42 post-first-exposure (p.f.e.), the hens were euthanized and parasites were recovered. Serum A. galli IgY level and ascarid eggs per gram of faeces (EPG) were examined on days -1 and 36 (IgY) or 40 p.f.e. (EPG). Results: Egg recovery in SF soil was substantially lower than in SC soil, but recovery was not significantly different between NF and NC soils. SF hens had a mean worm count of 76 whereas the other groups had means of 355– 453. Early mature/mature A. galli were recovered from SF hens whereas hens in the other groups harboured mainly immature A. galli. Heterakis gallinarum counts were low overall, especially in SF. The SF post-exposure IgY response was significantly lower while EPG was significantly higher compared to the other groups. Conclusions: Pochonia chlamydosporia was very effective in reducing ascarid egg numbers in sterilised soil and thus worm burdens in the exposed hens. However, reduced exposure of hens shifted A. galli populations toward a higher proportion of mature worms and resulted in a higher faecal egg excretion within the study period. This highlights a fundamental problem in ascarid control: if not all eggs in the farm environment are inactivated, the resulting low level infections may result in higher contamination levels with associated negative long-term consequences. Keywords: Ascaridia galli, Heterakis gallinarum, Fungus, Biological control * Correspondence: sundar@sund.ku.dk Section for Parasitology and Aquatic Pathobiology, Department of Veterinary and Animal Sciences, Faculty of Health and Medical Sciences, University of Copenhagen, Dyrlægevej 100, 1870 Frederiksberg C, Denmark Full list of author information is available at the end of the article © The Author(s). 2018 Open Access This article is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated. Thapa et al. Parasites & Vectors (2018) 11:319 Page 2 of 11 Background respectively, of A. galli eggs in laboratory agar assays. Ascaridia galli and Heterakis spp., collectively known as However, the effect in a soil based assay was relatively ascarids, are economically important intestinal nema- lower (~45%) [42]. In the latter study, the fungal effective- todes of chickens worldwide. Ascaridia galli can impair ness was evaluated based on eggs recovered from soil the health [1–3], productivity [4–7] and welfare of before and after the fungal treatment. However, it is un- chickens [8]. Moreover, A. galli can reduce the vaccine known if the recovered eggs judged visually as viable are efficacy against Newcastle disease [9, 10] and increase indeed infective and whether the reduced contamination the susceptiblilty of chickens to other infectious diseases in the soil assay translates to lower worm burdens in such as fowl cholera [11]. On rare occasions, A. galli can chickens exposed to the fungus-treated soil. In addition, leave the host’s intestine, migrate up the oviduct and literature indicates that population composition of A. galli become enclosed inside one of the hen’s eggs, which is in chickens can be dose-dependent as shown by a reduced of aesthetic concern to the consumers [12, 13]. Com- number of inhibited larvae in the host’s intestine and a pared to A. galli, Heterakis spp. are less pathogenic, but shorter prepatent period in lightly infected compared to they can act as a vector for in ovo transmission of the heavily infected chickens [43, 44]. It is thus important to protozoan Histomonas meleagridis to turkeys and chick- use an in vivo infection model to assess how changes in ens [14]. Histomonas meleagridis is pathogenic [15, 16] exposure level as a result of fungal treatment of soil may and re-emerging in layer flocks in many European coun- modulate worm population dynamics within the host as tries, mainly after the ban of the prophylactic use of che- this may in turn alter on-farm transmission dynamics. motherapeutics in the European Union (EU) member The overall aim of this study was to evaluate the in countries [17–20]. vivo infectivity of soil experimentally contaminated with Recent studies have shown that A. galli and Heterakis ascarid eggs and treated with P. chlamydosporia. Both spp. are highly prevalent in European organic laying hen sterilised and non-sterilised soils were used to include flocks [21–23]. Both nematodes have a simple life-cycle any effect inherent to the natural soil biota and thus to that involves a pre-parasitic development phase evaluate the potential of P. chlamydosporia as an (i.e. free-living nematode eggs) in the environment such on-farm biocontrol agent. as litter and soil and a parasitic phase in the chicken’s intestine following ingestion of infective eggs [24, 25]. Methods Ascarid eggs have thick shells [26, 27] and they can Experimental design survive in the outdoor environment for up to 2–4 Unembryonated ascarid eggs were added to Petri dishes years [28, 29], but no effective means of inactivating with sterilised (S) or non-sterilised (N) soil, half of which eggs during or after embryonation in the farm yards were treated with spores of the fungus P. chlamydos- and pastures are currently available. At present, con- poria (F) while the other half were untreated (C). The trol of ascarid infections by farmers therefore solely Petri dishes were incubated at 22 °C for 35 days to allow relies on flock treatment with commercial anthelmin- the fungus to grow and the eggs to reach infectivity. tics. Of the anthelmintics, only flubendazole and After incubation, three subgroups of 10 hens per soil fenbendazole are available for use in layers in the EU treatment were exposed in-feed four times to the soil [30, 31]. As hens are rapidly reinfected due to con- from one of the four treatments (SC, SF, NF and NC). tinuous exposure to eggs present in the surroundings The hens were euthanized on day 42 post-first-exposure and do not appear to acquire protective immunity (p.f.e.) and examined for A. galli and Heterakis spp. To [32–34], repeated treatments are thus necessary. estimate the number of eggs in the Petri dishes, recovery However, overuse of these drugs may over time en- of eggs was tested before [i.e. day 0 post-incubation hance the risk of selecting for anthelmintic resistance. (p.i.)] and after incubation (day 35 p.i.) for each of the To be able to also combat the parasites in the environ- four soil treatments. ment, there is an increasing interest in using naturally occurring soil microfungi as is done to control agricul- Origin and isolation of ascarid eggs tural nematode pests [35]. An isolate of Pochonia Before collecting faeces, the infection status of A. galli chlamydosporia (syn. Verticillium chlamydosporium) and Heterakis spp. in a Danish organic layer farm with a (Ascomycota: Hypocreales), a microfungus of global oc- flock size of 3000 hens was examined through necropsy currence [36–38], that can mechanically and enzymati- of 18 randomly selected hens. The prevalence was 89% cally degrade the egg shell components (protein and for A. galli and 100% for Heterakis spp., and mean ± S.E. chitin) has already been developed as a biocontrol agent worm burdens were 40 ± 9 and 80 ± 21 worms, respect- against plant-parasitic nematode eggs [39]. The same iso- ively. Ascarid eggs (A. galli and Heterakis spp.) were iso- late [40] and two other isolates [41]of P. chlamydosporia lated from fresh hen faeces collected from the ground have subsequently been shown to kill ~70% and > 80%, using only the top part of the faeces as described by Thapa et al. Parasites & Vectors (2018) 11:319 Page 3 of 11 Thapa et al. [29]. The eggs were stored in sterile demi- (i.e. pre-incubation) soil moisture level per dish (23– neralised water at 5 °C for 6 days. Before use, a subsample 24% of the total soil weight) was estimated (105 °C, 24 of eggs was embryonated in 0.1 N H SO at 25 °C for 15 h). Five random dishes per treatment were used to esti- 2 4 days to assess percentage embryonation (i.e. ability to mate day 0 p.i. egg recovery, while the remaining dishes develop larva) of the egg batch [42] which was 95 ± 1% were incubated at 22 °C for 35 days in darkness. The (mean ± SE). weight of each incubated dish was recorded at days 0 and 35 p.i. to determine soil moisture loss (%). On day Preparation of fungal inoculum 35 p.i., the dishes were opened and 4 ml sterile water Parboiled rice initially soaked for 1 h in demineralised was added to each dish, re-sealed with parafilm and water and autoclaved (121 °C, 15 min) inside a polypropyl- stored at 10 °C for up to 18 days. On day 7 post-storage ene bag (Labsolute®, 300 g rice per bag) was inoculated at 10 °C (i.e. after incubation was terminated), egg with 10 ml P. chlamydosporia Biotype 10 spore suspen- recovery (i.e. exposure level) was estimated in five 7 4 sion [5.5 × 10 conidia and 1.6 × 10 chlamydospores in random dishes per treatment (see section on recovery 0.05% Triton® X-100 (Merck KGaA, Darmstadt, Germany) of eggs from soil). Soil from one random dish was harvested from 5-week-old culture in Sabouraud’sdex- selected and administered in the feed to a correspond- trose agar]. After incubating the rice at 25 °C for 25 days ing subgroup of hens (see section on animal exposure in darkness, 30 ml 0.05% Triton® X-100 was added to each to parasites). This exposure was repeated on days 11, 10 g of rice granules in centrifuge tubes and shaken gently 15 and 19 after incubation was terminated. to separate spores from the rice. The mixture was filtered through a 900 μm sieve to remove rice particles and cen- Recovery of eggs from soil trifuged (1831× g, 3 min) three times after re-suspension Fifty millilitres of 0.5 M NaOH was added to each of the in 0.05% Triton® X-100. Spore concentration was adjusted 40 dishes (n = 5 per treatment on day 0 and 35 p.i.) that 8 5 to 4 × 10 conidia and 1.5 × 10 chlamydospores per ml was then stored at 5 °C for 16 h. The soil was washed suspension. Spore germination was determined [42]tobe through 212 and 20 μm sieves and the material on the 96% and 91% for the conidia and chlamydsopores, latter was divided into four 50 ml tubes, centrifuged at respectively. 253× g for 7 min and the eggs were recovered as described by Thapa et al. [29]. For each dish, the egg Preparation of soil quantity and development stage (unembryonated, In April 2016, 15 kg sandy loam soil (pH 6.8) was pre-larvated, larvated or degenerated) was examined in a collected from a Danish experimental plot that was 20% subsample at 100× magnification [29]. established in 2002 and treated anually (2003–2015) with source separated organic household waste compost (CH) and sown with spring cereals [45, 46]. After Experimental animals and housing removing plant material and stones, the soil was sieved One hundred thirty pullets (ISA Warren, 18-weeks-old), (3 mm) and thoroughly homogenised. Three kilograms of raised indoors without previous anthelmintic treatment, soil was sterilised by autoclaving (121 °C, 30 min) inside a were obtained from a commercial breeder. On arrival polypropylene bag (Labsolute®, 200 g soil per bag, (day -15 p.f.e.), 10 randomly selected pullets were eutha- treatment S) while another 3 kg soil was kept without nized and examined for ascarid infections of the breeder autoclaving (treatment N). farm-origin (see section on recovery of worms). All necropsied pullets were found positive for tissue phase Fungal treatment of soil and eggs A. galli larvae (~0.5 mm long) with an overall mean ± For both soils (S, N), 44 replicate Petri dishes (14.5 × 2 cm) SE worm burden of 194 ± 97 A. galli, while only two each containing 46 g soil and 2 ml ascarid egg suspension birds harboured luminal Heterakis spp. giving an with approximately 8000 ± 260 eggs (mean ± SE) in sterile overall burden of 1 ± 1 worm per hen. The remaining water were prepared. Both soil types were randomised into pullets (n = 120), after random allocation into 12 control (C) and fungus treatment (F). To all SF and NF indoor pens (c.2.8 m ,10pullets perpen), were dishes, 2 ml fungal suspension containing approxi- therefore treated with flubendazole (Verminator®, 1.43 8 5 mately 8 × 10 conidia and 3 × 10 chlamydospores of mg flubendazole per kg live weight daily) in the feed P. chlamydosporia in 0.05% Triton® X-100 was added from days -13 to -6 p.f.e. The individual body weight whereas all SC and NC dishes received 2 ml 0.05% of all birds was measured on days -1 and 36 p.f.e. Triton® X-100 without fungus. The SC and SF dishes The birds were given pelleted feed (17.5% crude received an additional 635 μl of sterile water to balance protein, 4.5% crude fat) in two meals (110 g feed per the total moisture level between the S and N soils. The bird per day) and water ad libitum. Crushed oyster dishes were sealed with Parafilm ‘M’® and the initial shells were offered daily. Wood-chips and straw were Thapa et al. Parasites & Vectors (2018) 11:319 Page 4 of 11 used as bedding material. Pens were enriched with a antigens and one replicate serum sample per animal per perch and nests, and cleaned thoroughly once weekly. sampling day. A dilution series of a highly positive serum was used as standard and the highest concentration was Animal exposure to parasites set at the relative value 2. The 12 pens were allocated to the four treatment groups (SC, SF, NC and NF) in triplets (i.e. three subgroups per Statistical analyses treatment group). The hens were exposed to ascarid All statistical analyses were performed using SAS 9.4 contaminated soil on days 0, 4, 8 and 12 p.f.e. to mimic a (Cary, NC, USA). The main and interaction effects of moderate trickle infection. On each exposure, entire soil soil sterility (S, N), fungal treatment (C, F) and incuba- from one Petri dish was transferred to a 500 ml container tion time (days 0, 35 p.i.) on egg recovery from soil were with 150 g feed of the morning meal and 50 ml tapwater, analysed using a generalised linear model fitted with mixed thoroughly and spread in a tray (58 × 21 × 3 cm) in negative binomial distribution of errors (NBD) (proced- each pen. The feed was eaten within 10–15 min and the ure GENMOD). Soil moisture loss during incubation remainder of the meal was then given in the same tray. was analysed with a linear model (procedure GLM) with percent moisture loss as the outcome and soil sterility Recovery of worms (S, N), fungal treatment (C, F) and their interaction as The hens were euthanized by stunning and cervical predictors. Body weight at day 0 p.f.e. and weight gain dislocation on day 42 p.f.e. The A. galli worms in the (days 0 to 36 p.f.e.) in relation to soil sterility (S, N) and small intestinal lumen were isolated using an agar-gel fungal treatment (C, F) were analysed separately with a method [47] and collected using a 20 μm sieve. The linear-mixed model (procedure MIXED) with subgroup tissue phase larvae of A. galli (day -15 and 42 p.f.e.) and (i.e. pen) as a random effect. Worm burden (total, A. Heterakis spp. (day -15 p.f.e.) were isolated from the galli, H. gallinarum), proportion (%) of A. galli in the in- intestinal/caecal tissue by pepsin (1:3000 IU)-HCl (30%) testinal tissue, proportion (%) of A. galli in each length digestion [47] and collected on a 20 μm sieve. To category (< 0.5, 0.5–1.5, 1.5–3.0, 3.0–5.0, 5.0–8.0 cm) recover luminal Heterakis spp. (day -15 and day 42 and at day 40 p.f.e. EPG was analysed with a generalised p.f.e.), the caeca were opened and stored in tap water at linear mixed model (procedure GLIMMIX, NBD) that 5 °C. After 48 h, the caeca and contents were washed on included soil sterility (S, N), fungal treatment (C, F) and a20 μm sieve. All worm samples were stored in 70% their interaction as fixed effects and subgroup as a ran- ethanol and examined using a dissection microscope dom effect. The log-transformed IgY titre was analysed (30–40× magnification). All A. galli worms were cate- with a linear-mixed model (procedure MIXED) with soil gorised as < 0.5, 0.5–1.5, 1.5–3.0, 3.0–5.0 or 5.0–8.0 cm, sterility (S, N), fungal treatment (C, F), sampling time whereas Heterakis spp. were categorised as < 0.5 or ≥ 0.5 (days -1, 36 p.f.e.) and their interaction as fixed effects, cm. Moreover, Heterakis species were determined based subgroup as a random effect and individual bird as a on the length of the spicules [48, 49] of 50 randomly repeated measurement. At group level, the linear selected male worms (1 worm per hen and representing relationships between worm burden (total ascarid or A. all experimental groups) after exposing each worm to a galli) and the IgY titre difference between pre- and drop of 10% lactic acid in water (weight/weight). post-exposures were examined using a Spearman method (procedure CORR). The goodness of fit of each Faecal egg counts GENMOD and GLIMMIX model was assessed with the Individual faecal samples from all birds were collected ratio of Pearson’s χ and corresponding degress of free- on days -1 and 40 p.f.e. Ascarid eggs per gram faeces dom. The normality of residuals of each GLM and (EPG) was determined by a concentration McMaster MIXED model was examined by a q-q plot and a histo- technique (minimum detection limit: 20 EPG) using a gram, and homogeneity of residual variance assessed by flotation fluid of 500 g glucose monohydrate per litre of residual plots. For each model, the post-hoc significant saturated NaCl solution (specific gravity: 1.27) [50]. differences were determined with the differences of least squares means (Tukey-Kramer’s adjustment for multiple Ascaridia galli antibody (IgY) levels comparisons, P < 0.05). To determine the systemic antibody response as an indirect assessment of parasite exposure, individual Results blood samples from all birds were collected on days -1 Recovery of eggs from soil and 36 p.f.e. from a wing vein. Serum was separated by On day 0 p.i., the mean number of eggs recovered from centrifugation at 1000× g for 15 min and stored at -20 °C. the SC, SF, NC and NF soils were 8702–9673 with no sig- The A. galli IgY level was determined by ELISA according nificant differences between the treatments (P >0.9950 in to Norup et al. [51] using crude adult A. galli somatic all cases) (Fig. 1). Irrespective of treatment, > 97% of the Thapa et al. Parasites & Vectors (2018) 11:319 Page 5 of 11 Worm burdens The overall mean worm burdens of A. galli and H. galli- narum in hens in the four groups are shown in Fig. 2a and b, respectively. All 118 hens were A. galli positive, while 115 birds were H. gallinarum positive. The left and right spicules of H. gallinarum males had a mean ± SE length of 2086 ± 26 μm (range: 1554–2417 μm) and 723 ± 8 μm (range: 402–850 μm), respectively. The interaction between the soil sterility and fungal treatment strongly in- fluenced the total ascarid worm burden (F = 100.38, (1, 106 ) P < 0.0001) and the individual worm burdens of both A. galli (F = 96.85, P < 0.0001) and H. galli- (1, 106) narum (F = 10.07, P = 0.0020). Group SF hens (1, 106) Fig. 1 Mean (+ SE) number of ascarid eggs recovered from soil on thus had significantly lower worm burdens of both A. galli days 0 and 35 post-incubation at 22 °C. Approximately 8000 (P < 0.0001 in all cases) and H. gallinarum (P ≤ 0.0001 in unembryonated eggs were added to soil given four different all cases) compared to the three other groups that all had treatments (n =5) (Abbreviations: SC, sterilised control; SF, sterilised comparable A. galli (P > 0.3120 in all cases) and H. galli- with the fungus Pochonia chlamydosporia Biotype 10; NC, non- sterilised control; NF, non-sterilised with fungus). Different letters narum worm burdens (P > 0.9989 in all cases) (Fig. 2). above the bars indicate significant differences (P < 0.05, Heterakis gallinarum represented 6% of the total ascarids Tukey-Kramer’s adjustment for multiple comparisons) recovered eggs were unembryonated. On day 35 p.i., the mean egg number in the SC, SF, NC and NF soils was 5535 (36% reduction), 521 (94% reduction), 4176 (57% reduction) and 3201 eggs (65% reduction), respectively (Fig. 1). In the sterilised soil, the fungal treatment resulted in a significant reduction in egg recovery when compared to the control (P < 0.0001). In contrast, there was no such difference in the non-sterilised soil (P = 0.5480). This meant that there was a strong significant (χ = 70.72, df =4, P < 0.0001) interaction between soil sterility, fungal treatment and incubation time on egg recov- ery. Regardless of treatment, ~94% of the recovered eggs at day 35 p.i. contained a slender larva that re- sembled the infective stage. The mean ± S.E. moisture loss in the sterilised soil (28 ± 1.8%) was slightly but signifi- cantly higher than in the non-sterilised soil (21 ± 1.4%) (F =8.53, P = 0.0048). (1, 64) Clinical observations and performance On day 0 p.f.e., the overall mean live weight of hens in the four groups was 1.53–1.60 kg with no significant effect of soil sterility (F =2.29, P = 0.1330), fungal (1, 106) treatment (F =1.91, P = 0.1701) and their inter- (1, 106) action (F =0.82, P = 0.3684). By day 36 p.f.e., the (1, 106) overall mean weight gain of hens in the four groups Fig. 2 Mean (+ SE) total worm burdens of Ascaridia galli (a) and Heterakis spp. (b) recovered from four groups of hens 42 days after was -76 to 90 g, but there was no significant effect of the first (of the total four) in-feed exposures to ascarid eggs soil sterility (F = 3.39, P = 0.0686), fungal treat- (1, 106) embryonated in sterilised control soil (SC), sterilised soil with the ment (F = 0.04, P = 0.8367) and their inter- (1, 106) fungus Pochonia chlamydosporia Biotype 10 (SF), non-sterilised action (F = 2.72, P = 0.1022). Most hens started (1, 106) control soil (NC) or non-sterilised soil with P. chlamydosporia (NF). laying eggs from days 3–7 p.f.e. During the study, the Each bar represents the mean of 28–30 hens allocated to three replicate subgroups of 8 (one NC subgroup) to 10 hens. Different hens showed no overt signs of illness but two hens letters above the bars indicate significant differences (P < 0.05, from one of the three NC subgroups died, possibly Tukey-Kramer’s adjustment for multiple comparisons) due to cannibalism. Thapa et al. Parasites & Vectors (2018) 11:319 Page 6 of 11 for SF hens and 2–3% for the three other groups. With reference to the estimated cumulative egg dose of 2214 (SC), 208 (SF), 1670 (NC) and 1281 eggs (NF) that each hen was theoretically exposed to on four exposures, the overall establishments of total ascarid were 20% (SC), 36% (SF), 21% (NC) and 33% (NF). Parasite population composition The overall mean proportion of A. galli recovered from the intestinal lumen and intestinal wall is shown in Fig. 3. In general, A. galli were more prevalent in the intestinal lumen (61–78%) than in the intestinal tissue (22–39%). Fungal treatment had a significant effect on the rela- tive distribution of tissue phase and luminal phase A. galli (F =9.85, P = 0.0022). This resulted in a (1, 106) significantly higher proportion (37 ± 2%, mean ± S.E.) of tissue phase A. galli in hens not exposed to fungal treatments compared to the hens exposed to fungal treatments (27 ± 2%). There were no significant ef- fects of soil sterility (F = 3.05, P = 0.0660) as (1, 106) well as the interaction between fungal treatment and Fig. 4 Mean proportion (%) of Ascaridia galli of different sizes soil sterility (F =0.50, P = 0.4815) on the A. (1, 106) recovered from four groups of hens 42 days after the first (of the galli distribution between intestinal lumen and tissue. total four) in-feed exposures to ascarid eggs embryonated in The proportion of A. galli (of the total A. galli worm sterilised control soil (SC), sterilised soil with the fungus Pochonia chlamydosporia Biotype 10 (SF), non-sterilised control soil (NC) or burden) within each length category was significantly re- non-sterilised soil with P. chlamydosporia (NF). Each bar represents lated to the interaction between soil sterility and fungal the mean of 28–30 hens allocated to three replicate subgroups of 8 treatment (< 0.5 cm: F =29.48, P < 0.0001; 0.5–1.5 (1, 106) (one NC subgroup) to 10 hens. Different letters above the bars cm: F =9.89, P = 0.0022; 1.5–3.0 cm: F = (1, 106) (1, 106) indicate significant differences (P < 0.05, Tukey-Kramer’s adjustment 29.98, P < 0.0001; 3.0–5.0 cm: F =16.58, P < 0.0001; for multiple comparisons) within each length category (1, 106) 5.0–8.0 cm: F =7.24, P = 0.0083) (Fig. 4). Compared (1, 106) to the groups SC, NC and NF hens, the group SF hens hosted a significantly lower proportion of A. galli <0.5 cm (P < 0.0001 in all cases) and significantly higher propor- tions of the three largest length categories (P < 0.0225 in all cases, except P =0.0508for SF vs NF in the category 5.0–8.0 cm). The SC, NC and NF hens hosted nearly equal proportions of all five A. galli length categories (P > 0.0625 in all cases, except P =0.0076 for SC vs NC in the category 1.5–3.0 cm). Irrespective of group, all tissue phase A. galli larvae were < 0.5 cm (~0.5 mm). In groups SC, NC and NF hens, the luminal A. galli worms within the catergory < 0.5 cm (i.e. 5 mm) were approximately 0.5–1.0 mm whereas those in group SF hens ranged ~0.5–4.9 mm. For H. gallinarum, the highest mean ± S.E. proportion of worms > 0.5 cm was hosted by the group SF hens (52 ± Fig. 3 Mean proportion (%) of luminal phase and tissue phase 9%) followed by NC (38 ± 6%), NF (31 ± 5%) and SC hens Ascaridia galli recovered from four groups of hens 42 days after (25 ± 6%). However, the effect of soil sterility, fungal the first (of the total four) in-feed exposures to ascarid eggs treatment and their inferactions on H. gallinarum embryonated in sterilised control soil (SC), sterilised soil with the fungus Pochonia chlamydosporia Biotype 10 (SF), non-sterilised population composition was not possible to analyse control soil (NC) or non-sterilised soil with P. chlamydosporia (NF). using the same statistical model that was used for A. Each bar represents the mean of 28–30 hens allocated to three galli because many hens had only < 0.5 or ≥ 0.5 cm replicate subgroups of 8 (one NC subgroup) to 10 hens H. gallinarum. Thapa et al. Parasites & Vectors (2018) 11:319 Page 7 of 11 Faecal egg counts On day -1 p.f.e., all hens were negative for ascarid eggs. On day 40 p.f.e., 3, 57, 7 and 17% hens of groups SC, SF, NC and NF, respectively, had positive EPG. There was a signifi- cant interaction between fungal treatment and soil sterility regarding day 40 p.f.e. EPG (F =5.17, P < 0.0250) as (1, 106) the overall mean EPG in group SF hens was signifi- cantly higher than in groups SC (P < 0.0001) and NC hens (P = 0.0241) but comparable to group NF hens (P = 0.4040) (Fig. 5). Ascaridia galli IgY titres The overall group mean (+ SE) A. galli IgY titres in hens in the four groups on days -1 and 36 p.f.e. are shown in Fig. 6 Mean (+ SE) Ascaridia galli IgY titre at one day before and 36 Fig. 6. All hens were seropositive at both time-points. days after the first (of the total four) in-feed exposures to ascarid The antibody titer was significantly affected by the inter- eggs embryonated in sterilised control soil (SC), sterilised soil with action between sterility of soil, fungal treatment and the fungus Pochonia chlamydosporia Biotype 10 (SF), non-sterilised control soil (NC) or non-sterilised soil with P. chlamydosporia (NF). sampling time (F = 14.02, P < 0.0001). On day -1 (4, 212) Each bar represents the mean of 28–30 hens allocated into three p.f.e., the group mean ± SE titres ranged between 734 ± replicate subgroups of 8 (one NC subgroup), 9 (two SC and two NF 62 and 1071 ± 115, with no significant differences be- subgroups) or 10 hens. Different italicised letters above the bars tween the groups (P > 0.7380 in all cases). By day 36 indicate significant differences (P < 0.05, Tukey-Kramer’s adjustment p.f.e., the IgY titre had increased significantly in all for multiple comparisons) between the log-transformed titres groups with an overall 8–11 fold increase in groups SC (P < 0.0001), NC (P < 0.0001) and NF (P < 0.0001), but only three fold increase in group SF hens (P < 0.0001). Discussion There were no significant correlations (P > 0.05) Thepresent studyhas forthe firsttimeshown that between IgY titre and individual worm burden (total ascarid transmission to hens exposed to egg contami- ascarid, total A. galli) in all groups except SF where nated soil can be reduced after the soil has been there were significant but weak correlations for the total treated with the fungus P. chlamydosporia, but only ascarid (r = 0.38, P = 0.0362) and A. galli worm bur- in sterilised soil. The reduced exposure resulted in a (30) den (r = 0.40, P = 0.0275). higher rate of development into adult worms and (30) thus more patent infections compared to the more heavily infected control hens. The fungus P. chlamydosporia Biotype 10 substantially reduced the egg recovery in the sterilised soil whereas in non-sterilised soil there was no additional effect when compared to the corresponding controls. This limited effect of P. chlamydosporia in the non-sterilised soil is in line with previous findings for egg-degrading fungi in general [38, 42, 52, 53]. The currently available literature indicates that native soil biota can reduce the establish- ment of a newly added fungus [54–58]. This is probably because the new fungus must compete for the soil re- sources or overcome antagonism by native established soil biota such as other fungi [59–61], bacteria [62–65], protozoa [66], free-living nematodes [67, 68], mites and Fig. 5 Mean (+ SE) number of ascarid eggs per gram of faeces (EPG) dipteran larvae [69]. In future studies, application of of four groups of hens 40 days after the first (of the total four) in- fungi in nutrient-rich substrates (e.g. rice or barley kernels, feed exposures to ascarid eggs embryonated in sterilised control soil decomposed resources etc.) could be explored as this may (SC), sterilised soil with the fungus Pochonia chlamydosporia Biotype help increase fungal establishment in soil [55, 70, 71]. 10 (SF), non-sterilised control soil (NC) or non-sterilised soil with P. chlamydosporia (NF). Each bar represents the mean of 28–30 hens Ascarid eggs are sensitive to dessication [72] and after allocated to three replicate subgroups of 8 (one NC subgroup) to 10 incubation, we found a slightly higher moisture loss in the hens. Different letters above the bars indicate significant differences sterilised soil compared to the non-sterilised soil. How- (P < 0.05, Tukey-Kramer’s adjustment for multiple comparisons) ever, results indicate that this had no major impact as the Thapa et al. Parasites & Vectors (2018) 11:319 Page 8 of 11 moisture loss in the control and the fungus-treated and the effect of host immune responses on parasite sterilised soil was not significantly different and both the population seem important [79]. highest moisture loss and highest egg recovery from a The above findings highlight the basic complication of single Petri dish was found in the sterilised soil. any control strategy that cannot inactivate all parasite The differences in soil egg numbers (i.e. exposure eggs in the environment. Initially there may be a lowered levels) after fungal treatment was reflected in vivo by impact on the hosts present at the time due to lowered parasite burden, establishment rate and population com- exposure, but if the result is associated with altered in- postion within the host. The least exposed group had fection dynamics, and thus an earlier onset of patency, the lowest ascarid worm burdens but a higher parasite environmental recontamination might be higher than if establishment rate compared to the three other groups there had been no intervention. This is further compli- that were more heavily exposed. Similar findings have cated as freshly deposited eggs take weeks to months to been reported for an A. galli trickle infection in chickens develop to infectivity depending on weather and season [32], H. gallinarum single infection in red-necked pheas- [30]. This goes to show that designing and implementing ants [73]and Oesophagostomum dentatum single and control strategies on a farm must take parasite biology trickle infections in pigs [74–76]. In contrast, Permin et and ecology into account to not only offer temporary al. [77] found no differences in A. galli burdens follow- relief, but also be effective long-term. ing a single dose of 100, 500 or 2500 eggs. This may be There is a close phylogenetic relationship between A. because they only quantified the luminal worms and galli and Heterakis spp. [80] with corresponding produc- many larvae in the two higher dose groups may poten- tion of cross-reacting antibodies [81]. However, the tially have been, at least temporarily, arrested in the current contribution in the IgY titre due to H. gallinarum intestinal mucosa [43, 44]. In the current study, we thus is expected to be neglible due to the much lower worm found an increased proportion of tissue phase A. galli burdens compared to A. galli. The individual antibody presumably at the third larval stage (L3) [43, 44] in the levels appeared to increase with increasing exposure level three high exposure groups, and larger worms and pa- and worm burden. However, individual antibody levels tent infections primarily in the lowest exposure group. seemed uninvolved in any immune-related short-term The absence of patent infections in most heavily regulation of A. galli populations. This is in agreement infected hens supports that faecal egg counts can with previous findings of a very weak or a complete lack severely underestimate immature worm burdens and of correlation between systemic/egg-yolk IgY level and A. exposure levels [44, 74, 76]. galli/H. gallinarum worm burden [3, 81]. A similar lack of The current results showed that low exposure may at association between porcine blood IgG level and worm least, in the short-term, lead to mature A. galli popula- burdens has been reported for Ascaris suum [82, 83]and tions in contrast to predominantly immature infections Trichuris suis [84]. Furthermore, we also found that previ- at higher exposure levels. Reduced exposure and lower ous exposure did not protect against subsequent A. galli worm burdens are both desireable to lower the overall reinfection, which is in line with other studies [32, 85]. impact of ascarids on chicken health and productivity Others have reported increased mRNA expression of Th2 but seem to favour the establishment of patent infec- cytokines IL-4 and IL-13 in the intestinal tissues and tions. Density-dependent worm maturation was previ- spleen of A. galli infected hosts [3, 86, 87]. Both cytokines ously documented for H. gallinarum in chickens and play a role in mediating protective immunity against sev- ring-necked pheasants where heavily infected birds eral helminth parasites [88, 89] but it appears that A. galli hosted significantly smaller female worms compared to may evade host immune responses to avoid expulsion as lightly infected birds [73, 78]. It is unknown if, given suggested for O. dentatum in pigs [90, 91]. This could be time, at least some of our arrested larvae, presumably a reason why A. galli prevalence in laying hens kept in L3 in the intestinal tissue and L4 in the intestinal non-cage systems (barn, free-range and organic) seems to lumen [1, 43], would have reached maturity as our hens increase over time during an egg laying period of approxi- were only followed for 30 days after the last exposure. mately one year [34]. However, Ikeme [44] found the development of nearly all To the best of our knowledge, there are no opti- L3 to be arrested for up to 13 weeks post-last-exposure in mal/standardized protocols to establish patent A. galli birds that received a high infection dose. It is therefore infections in chickens. Experimental infection proce- very important to use sensitive recovery techniques to dures vary greatly in relation to infection material minimize the risk of overlooking high immature worm (source, embryonation medium, temperature and dur- burdens. The precise mechanisms responsible for ation of embryonation) and host factors (age, breed, density-dependent effects are not fully understood. The etc.) [10, 32, 92–96]. This makes it extremely difficult combination of intraspecific competition among worms to compare results between different studies. Many for limited resources (e.g. space, nutrients) in the host gut experiments performed earlier by our group could Thapa et al. Parasites & Vectors (2018) 11:319 Page 9 of 11 not establish patent A. galli infections when chickens Ethics approval The animal experiment was approved by the Animal Experiments were either infected with a single dose of 500 eggs Inspectorate, The Danish Ministry of Food, Agriculture and Fisheries [97, 98] or trickle infected twice weekly with 25–100 (Permit Number: 2015-15-0201-00760). The animals were treated according eggs per infection over a period of six weeks [32, 85]. to the Danish ethical guidelines. The lowest infection dose used in the latter studies is Competing interests very similar to the lowest exposure level of the The authors declare that they have no competing interests. current study. We have therefore made some modifi- cations in the current protocol in relation to the pre- Publisher’sNote vious failures. Hens were exposed to ascarid eggs at Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. 20–22 weeks of age. This corresponded to the period when most hens started to lay eggs and it has been sug- Author details gested that hens during this period are more susceptible Section for Parasitology and Aquatic Pathobiology, Department of Veterinary and Animal Sciences, Faculty of Health and Medical Sciences, and may have an increased establishment of A. galli due University of Copenhagen, Dyrlægevej 100, 1870 Frederiksberg C, Denmark. to hormonal changes in the birds [92]. Furthermore, we 2 College of Veterinary Medicine, Inner Mongolia Agricultural University, embryonated (i.e. incubated) ascarid eggs at 22 °C for only Hohhot 010018, People’s Republic of China. Section for Organismal Biology, Department of Plant and Environmental Sciences, Faculty of Science, five weeks compared to the six week incubation protocol University of Copenhagen, Thorvaldsensvej 40, 1871 Frederiksberg C, in the earlier studies. This was because chicken ascarid 4 Denmark. Section for Immunology and Microbiology, Department of Animal eggs develop fully within four weeks of incubation at Science, Aarhus University, Blichers Allé 20, Building P25, 3334, 8830 Tjele, Denmark. Section for Diagnostics and Scientific Advice, National Veterinary 22 °C [29] and we thus provided only one additional Institute, Technical University of Denmark, Kemitorvet, 2800 Kgs. Lyngby, week for the developed eggs to mature to infectivity. Denmark. Received: 15 February 2018 Accepted: 14 May 2018 Conclusions The Biotype 10 strain of P. chlamydosporia was only effective in inactivating ascarid eggs and thereby reducing References 1. Ikeme MM. Observations on the pathogenecity and pathology of Ascaridia the infection level of sterilised soil and thus total worm galli. Parasitology. 1971;63:169–79. burdens of the exposed hens. The consequence was that 2. Ramadan HH, Abou Znada NY. Some pathological and biochemical studies the proportion of early mature/mature A. galli increased on experimental ascaridiasis in chickens. Nahrung. 1991;35:71–84. 3. Schwarz A, Gauly M, Abel H, Daş G, Humburg J, Rohn K, et al. and faecal eggs counts were higher than in all the Immunopathogenesis of Ascaridia galli infection in layer chicken. Dev Comp other groups where hens were exposed to a higher Immunol. 2011;35:774–84. number of infective eggs in the soil. This underlines an 4. Reid WM, Carmon JL. Effects of numbers of Ascaridia galli in depressing weight gains in chicks. J Parasitol. 1958;44:183–6. inherent dilemma and complexity of ascarid control in that 5. Kilpinen O, Roepstorff A, Permin A, Nørgaard-Nielsen G, Lawson LG, hosts may suffer less short-term, but current reduced Simonsen HB. Influence of Dermanyssus gallinae and Ascaridia galli exposure may lead to long-term higher environmental infections on behaviour and health of laying hens (Gallus gallus domesticus). Br Poult Sci. 2005;46:26–34. re-contamination if not all eggs can be eliminated. This 6. Phiri IK, Phiri AM, Ziela M, Chota A, Masuku M, Monrad J. Prevalence needs to be considered in future biological and other and distribution of gastrointestinal helminths and their effects on control strategies in poultry. weight gain in free-range chickens in Central Zambia. Trop Anim Health Prod. 2007;39:309–15. Abbreviations 7. Skallerup P, Luna LA, Johansen MV, Kyvsgaard NC. The impact of natural EPG: eggs per gram of faeces; FEC: faecal egg counts; SE: standard error helminth infections and supplementary protein on growth performance of free-range chickens on smallholder farms in El Sauce, Nicaragua. Prev Vet Acknowledgements Med. 2005;69:229–44. The authors gratefully acknowledge Rothamsted Research Ltd. (United 8. Gauly M, Duss C, Erhardt G. Influence of Ascaridia galli infections and Kingdom) for kindly providing Pochonia chlamydosporia Biotype 10. We are anthelmintic treatments on the behaviour and social ranks of laying hens very thankful to Lise-Lotte Christensen, Suraj Dhakal, Lene R. Dal, Sara (Gallus gallus domesticus). Vet Parasitol. 2007;146:271–80. Almeida, Sophie Stolzenbach, Tina V. A. Hansen and Eline P. Hansen for the 9. Hørning G, Rasmussen S, Permin A, Bisgaard M. Investigations on the technical assistance. influence of helminth parasites on vaccination of chickens against Newcastle disease virus under village conditions. Trop Anim Health Prod. Funding 2003;35:415–24. This research and the PhD fellowship of ST were financed by the Faculty of 10. Pleidrup J, Dalgaard TS, Norup LR, Permin A, Schou TW, Skovgaard K, et al. Health and Medical Sciences, University of Copenhagen, Denmark. Ascaridia galli infection influences the development of both humoral and cell-mediated immunity after Newcastle disease vaccination in chickens. Availability of data and materials Vaccine. 2014;32:383–92. The datasets generated and analysed in the current study can be available 11. Dahl C, Permin A, Christensen JP, Bisgaard M, Muhairwa AP, Petersen from the corresponding author upon reasonable request. KMD, et al. The effect of concurrent infections with Pasteurella multocida and Ascaridia galli on free range chickens. Vet Microbiol. Authors’ contributions 2002;86:313–24. ST, SMT, HM and NVM designed the study. ST, RW, HM and TSD conducted 12. Reid WM, Mabon JL, Harshbarger WC. Detection of worm parasites in the study. ST analysed the data and drafted the manuscript. All authors chicken eggs by candling. Poult Sci. 1973;52:2316–24. discussed the results, contributed in the manuscript and read and approved 13. Fioretti DP, Veronesi F, Diaferia M, Franciosini MP, Proietti PC. Ascaridia galli: the final manuscript. a report of erratic migration. Ital J Anim Sci. 2005;4:310–2. Thapa et al. Parasites & Vectors (2018) 11:319 Page 10 of 11 14. Lee DL. The structure and development of Histomonas meleagridis 39. Manzanilla-López HR, Esteves I, Finetti-Sialer MM, Hirsch PR, Ward E, (Mastigamoebidae: Protozoa) in the female reproductive tract of its Devonshire J, et al. Pochonia chlamydosporia: advances and challenges to intermediate host, Heterakis gallinarum (Nematoda). Parasitology. improve its performance as a biological control agent of sedentary 1969;59:877–84. endo-parasitic nematodes. J Nematol. 2013;45:1–7. 15. Windisch M, Hess M. Experimental infection of chickens with Histomonas 40. Thapa S, Meyling NV, Katakam KK, Thamsborg SM, Mejer H. A method to meleagridis confirms the presence of antibodies in different parts of the evaluate relative ovicidal effects of soil microfungi on thick-shelled eggs of intestine. Parasite Immunol. 2010;32:29–35. animal-parasitic nematodes. Biocontrol Sci Technol. 2015;25:756–67. 16. McDougald LR. Blackhead disease (histomoniasis) in poultry: a critical 41. Braga FR, Araújo JV, Araujo JM, Frassy LN, Tavela AO, Soares FEF, et al. review. Avian Dis. 2005;49:462–76. Pochonia chlamydosporia fungal activity in a solid medium and its crude extract against eggs of Ascaridia galli. J Helminthol. 2012;86:348–52. 17. Dolka B, Żbikowski A, Dolka I, Szeleszczuk P. Histomonosis - an existing problem in chicken flocks in Poland. Vet Res Commun. 2015;39:189–95. 42. Thapa S, Mejer H, Thamsborg SM, Lekfeldt JDS, Wang R, Jensen B, et al. 18. Grafl B, Liebhart D, Windisch M, Ibesich C, Hess M. Seroprevalence of Survival of chicken ascarid eggs exposed to different soil types and fungi. Histomonas meleagridis in pullets and laying hens determined by ELISA. Appl Soil Ecol. 2017;121:143–51. Vet Rec. 2011;168:160. 43. Herd RP, McNaught DJ. Arrested development and the histotropic phase of 19. Stokholm NM, Permin A, Bisgaard M, Christensen JP. Causes of mortality in Ascaridia galli in the chicken. Int J Parasitol. 1975;5:401–6. commercial organic layers in Denmark. Avian Dis. 2010;54:1241–50. 44. Ikeme MM. Effects of different levels of nutrition and continuing dosing of 20. Esquent C, De Herdt P, De Bosschere H, Ronsmans S, Ducatelle R, Van Erum poultry with Ascaridia galli eggs on the subsequent development of J. An outbreak of histomoniasis in free-range layer hens. Avian Pathol. parasite populations. Parasitology. 1971;63:233–50. 2003;32:305–8. 45. Poulsen PHB, Al-Soud WA, Bergmark L, Magid J, Hansen LH, Sørensen SJ. Effects of fertilization with urban and agricultural organic wastes in a field 21. Thapa S, Hinrichsen LK, Brenninkmeyer C, Gunnarsson S, Heerkens JLT, trial - prokaryotic diversity investigated by pyrosequencing. Soil Biol Verwer C, et al. Prevalence and magnitude of helminth infections in organic Biochem. 2013;57:784–93. laying hens (Gallus gallus domesticus) across Europe. Vet Parasitol. 2015;214:118–24. 46. López-Rayo S, Laursen KH, Lekfeldt JDS, Delle Grazie F, Magid J. Long-term 22. Jansson DS, Nyman A, Vågsholm I, Christensson D, Göransson M, Fossum O, amendment of urban and animal wastes equivalent to more than 100 years et al. Ascarid infections in laying hens kept in different housing systems. of application had minimal effect on plant uptake of potentially toxic Avian Pathol. 2010;39:525–32. elements. Agric Ecosyst Environ. 2016;231:44–53. 23. Kaufmann F, Daş G, Sohnrey B, Gauly M. Helminth infections in laying hens 47. Ferdushy T, Nejsum P, Roepstorff A, Thamsborg SM, Kyvsgaard NC. Ascaridia kept in organic free range systems in Germany. Livest Sci. 2011;141:182–7. galli in chickens: intestinal localization and comparison of methods to 24. Ackert JE. The morphology and life history of the fowl nematode Ascaridia isolate the larvae within the first week of infection. Parasitol Res. lineata (Schneider). Parasitology. 1931;23:360–79. 2012;111:2273–9. 48. Lapage G. Mönnig’s veterinary helminthology and entomology. London: 25. Clapham PA. On the life-history of Heterakis gallinae. J Helminthol. Bailliere, Tindall and Cox; 1956. 1933;11:67–86. 49. Soulsby EJL. Textbook of veterinary clinical parasitology, Volume 1. 26. Wharton D. Nematode egg-shells. Parasitology. 1980;81:447–63. Helminths. Oxford: Blackwell Scientific Publications; 1965. 27. Christenson RO, Earle HH, Butler RL, Creel HH. Studies on the eggs of Ascaridia galli and Heterakis gallinae. Trans Am Microsc Soc. 1942;61:191. 50. Roepstorff A, Nansen P. Epidemiology, diagnosis and control of helminth 28. Farr MM. Further observations on the survival of the protozoan parasite, parasites of swine. FAO Animal Health Manual. Rome: Food and Agriculture Histomonas meleagridis and eggs of poultry nematdoes in feces of infected Organization of the United Nations; 1998. birds. Cornell Vet. 1961;51:3–13. 51. Norup LR, Dalgaard TS, Pleidrup J, Permin A, Schou TW, Jungersen G, et al. Comparison of parasite-specific immunoglobulin levels in two chicken lines 29. Thapa S, Thamsborg SM, Meyling NV, Dhakal S, Mejer H. Survival and during sustained infection with Ascaridia galli. Vet Parasitol. 2013;191:187–90. development of chicken ascarid eggs in temperate pastures. Parasitology. 52. Bourne JM, Kerry BR, De Leij FAAM. Methods for the study of Verticillium 2017;144:1243–52. chlamydosporium in the rhizosphere. J Nematol. 1994;26:587–91. 30. Committee for Medicinal Products for Veterinary Use. Flubendazole 53. Monfort E, Lopez-Llorca LV, Jansson HB, Salinas J. In vitro soil receptivity (extrapolation to poultry), summary report (4). In: EMEA/CVMP/33128/2006- assays to egg-parasitic nematophagous fungi. Mycol Prog. 2006;5:18–23. Final; 2006. http://www.ema.europa.eu/docs/en_GB/document_library/ Maximum_Residue_Limits_-_Report/2009/11/WC500014292.pdf. 54. Lockwood JL, Lingappa BT. Fungitoxicity of sterilised soil inoculated with Accessed 21 Mar 2017. soil microflora. Phytopathology. 1963;58:917–20. 31. Committee for Medicinal Products for Veterinary Use. CVMP assessment report 55. Lockwood JL. Fungistasis in soils. Biol Rev. 1977;52:1–43. for Panacur AquaSol (EMEA/V/C/002008/X/00003). 2014. http://www.ema. 56. Toyota K, Ritz K, Young IM. Microbiological factors affecting the colonisation europa.eu/docs/en_GB/document_library/EPAR_-_Assessment_Report_-_ of soil aggregates by Fusarium oxysporum f. sp. raphani. Soil Biol Biochem. Variation/veterinary/002008/WC500165605.pdf. Assessed 21 Mar 2017. 1996;28:1513–21. 32. Ferdushy T, Luna-Olivares LA, Nejsum P, Thamsborg SM, Kyvsgaard NC. The 57. Mauchline TH, Kerry BR, Hirsch PR. Quantification in soil and the rhizosphere use of genetically marked infection cohorts to study changes in of the nematophagous fungus Verticillium chlamydosporium by competitive establishment rates during the time course of a repeated Ascaridia galli PCR and comparison with selective plating. Appl Environ Microbiol. infection in chickens. Int J Parasitol. 2015;45:393–8. 2002;68:1846–53. 33. Tarbiat B, Jansson DS, Tydén E, Höglund J. Comparison between 58. de Boer W, Verheggen P, Klein Gunnewiek PJ, Kowalchuk G, van Veen JA. anthelmintic treatment strategies against Ascaridia galli in commercial Microbial community composition affects soil fungistasis. Appl Environ laying hens. Vet Parasitol. 2016;226:109–15. Microbiol. 2003;69:835–44. 34. Zloch A, Kuchling S, Hess M, Hess C. Influence of alternative husbandry 59. Chen SY, Chen FJ. Fungal parasitism of Heterodera glycines eggs as systems on postmortem findings and prevalence of important bacteria and influenced by egg age and pre-colonization of cysts by other fungi. parasites in layers monitored from end of rearing until slaughter. Vet Rec. J Nematol. 2003;35:271–7. 2018;182:350. 60. Howell CR. Mechanisms employed by Trichoderma species in the biological 35. Braga FR, De Araújo JV. Nematophagous fungi for biological control of control of plant diseases: The history and evolution of current concepts. gastrointestinal nematodes in domestic animals. Appl Microbiol Biotechnol. Plant Dis. 2003;87:4–10. 2014;98:71–82. 61. Harman GE, Howell CR, Viterbo A, Chet I, Lorito M. Trichoderma species- 36. Domsch KH, Gams W, Anderson T-H. Compendium of soil fungi, Volume 1. opportunistic, avirulent plant symbionts. Nat Rev Microbiol. 2004;2:43–56. 1st ed. London: Academic Press; 1980. 62. Lopez Llorca LV, Boag B. Inhibition of Verticillium suchlasporium and other 37. Lýsek H, Fassatiová O, Cuervo Pineda N, Lorenzo Hernández N. Ovicidal nematophagous fungi by bacteria colonising Heterodera avenae females. fungi in soils of Cuba. Folia Parasitol. 1982;29:265–70. Nematol Mediterr. 1990;18:233–7. 38. Siddiqui IA, Atkins SD, Kerry BR. Relationship between saprotrophic growth 63. Ballhausen MB, van Veen JA, Hundscheid MPJ, de Boer W. Methods for in soil of different biotypes of Pochonia chlamydosporia and the infection of baiting and enriching fungus-feeding (Mycophagous) rhizosphere bacteria. nematode eggs. Ann Appl Biol. 2009;155:131–41. Front Microbiol. 2015;6:1416. Thapa et al. Parasites & Vectors (2018) 11:319 Page 11 of 11 64. Rudnick MB, van Veen JA, de Boer W. Baiting of rhizosphere bacteria with 87. Dalgaard TS, Skovgaard K, Norup LR, Pleidrup J, Permin A, Schou TW, et al. hyphae of common soil fungi reveals a diverse group of potentially Immune gene expression in the spleen of chickens experimentally infected mycophagous secondary consumers. Soil Biol Biochem. 2015;88:73–82. with Ascaridia galli. Vet Immunol Immunopathol. 2015;164:79–86. 65. Li L, Mo M, Qu Q, Luo H, Zhang K. Compounds inhibitory to 88. Anthony RM, Rutitzky LI, Urban JF, Stadecker MJ, Gause WC. Protective nematophagous fungi produced by Bacillus sp. strain H6 isolated from immune mechanisms in helminth infection. Nat Rev Immunol. fungistatic soil. Eur J Plant Pathol. 2007;117:329–40. 2007;7:975–87. 89. Finkelman FD, Wynn TA, Donaldson DD, Urban Jr JF. The role of IL-13 in 66. Ekelund F, Rønn R. Notes on protozoa in agricultural soil with emphasis on helminth-induced inflammation and protective immunity against nematode heterotrophic flagellates and naked amoebae and their ecology. FEMS infections. Curr Opin Immunol. 1999;11:420–6. Microbiol Rev. 1994;15:321–53. 90. Andreasen A, Petersen HH, Kringel H, Iburg TM, Skovgaard K, Dawson H, 67. Ruess L, Dighton J. Cultural studies on soil nematodes and their fungal et al. Immune and inflammatory responses in pigs infected with Trichuris hosts. Nematologica. 1996;42:330–46. suis and Oesophagostomum dentatum. Vet Parasitol. 2015;207:249–58. 68. Jaffee BA, Muldoon AE, Didden WAM. Enchytraeids and nematophagous 91. Andreasen A, Skovgaard K, Klaver EJ, van Die I, Mejer H, Thamsborg SM, et fungi in soil microcosms. Biol Fertil Soils. 1997;25:382–8. al. Comparison of innate and Th1-type host immune responses in 69. Ruess L, Lussenhop J. Trophic interactions of fungi and animals. In: Dighton Oesophagostomum dentatum and Trichuris suis infections in pigs. Parasite J, White FJ, Oudemans P, editors. The fungal community - its organization Immunol. 2016;38:53–63. and role in the ecosystem. 3rd ed. Boca Raton: CRC Press; 2005. p. 581–98. 92. Gauly M, Homann T, Erhardt G. Age-related differences of Ascaridia galli egg 70. Kerry B. The use of microbial agents for the biological control of plant output and worm burden in chickens following a single dose infection. parasitic nematodes. In: Jones DG, editor. Exploitation of microorganisms. Vet Parasitol. 2005;128:141–8. London: Chapman & Hall; 1993. p. 81–104. 93. Permin A, Nansen P, Bisgaard M, Frandsen F. Ascaridia galli infections in 71. Luambano ND, Manzanilla-López RH, Kimenju JW, Powers SJ, Narla RD, free-range layers fed on diets with different protein contents. Br Poult Sci. Wanjohi WJ, et al. Effect of temperature, pH, carbon and nitrogen ratios on 1998;39:441–5. the parasitic activity of Pochonia chlamydosporia on Meloidogyne incognita. 94. Rahimian S, Daş G, Gauly M. Maternal protection against Ascaridia galli? Biol Control. 2015;80:23–9. Vet Parasitol. 2017;233:43–7. 72. Tarbiat B, Jansson DS, Höglund J. Environmental tolerance of free-living 95. Ruhnke I, Andronicos NM, Swick RA, Hine B, Sharma N, Kheravii SK, et al. stages of the poultry roundworm Ascaridia galli. Vet Parasitol. 2015;209:101–7. Immune responses following experimental infection with Ascaridia galli and 73. Tompkins DM, Hudson PJ. Regulation of nematode fecundity in the ring- necrotic enteritis in broiler chickens. Avian Pathol. 2017;46:602–9. necked pheasant (Phasianus colchicus): not just density dependence. 96. Marcos-Atxutegi C, Gandolfi B, Arangüena T, Sepúlveda R, Arévalo M, Simón Parasitology. 1999;118:417–23. F. Antibody and inflammatory responses in laying hens with experimental 74. Christensen CM, Barnes EH, Nansen P, Roepstorff A, Slotved HC. primary infections of Ascaridia galli. Vet Parasitol. 2009;161:69–75. Experimental Oesophagostomum dentatum infections in the pig: Worm 97. Ferdushy T, Luna-Olivares LA, Nejsum P, Roepstorff AK, Thamsborg SM, populations at regular intervals during trickle infections with three dose Kyvsgaard NC. Population dynamics of Ascaridia galli following single levels of larvae. Int J Parasitol. 1995;25:1491–8. infection in young chickens. Parasitology. 2013;140:1078–84. 75. Roepstorff A, Nilsson O, Oksanen A, Gjerde B, Richter SH, Örtenberg E, et al. 98. Luna-Olivares LA, Kyvsgaard NC, Ferdushy T, Nejsum P, Thamsborg SM, Intestinal parasites in swine in the Nordic countries: prevalence and Roepstorff A, et al. The jejunal cellular responses in chickens infected with a geographical distribution. Vet Parasitol. 1998;76:305–19. single dose of Ascaridia galli eggs. Parasitol Res. 2015;114:2507–15. 76. Roepstorff A, Bjørn H, Nansen P, Barnes EH, Christensen CM. Experimental Oesophagostomum dentatum infections in the pig: worm populations resulting from trickle infections with three dose levels of larvae. Int J Parasitol. 1996;26:399–408. 77. Permin A, Bojesen M, Nansen P, Bisgaard M, Frandsen F, Pearman M. Ascaridia galli populations in chickens following single infections with different dose levels. Parasitol Res. 1997;83:614–7. 78. Daş G, Gauly M. Density related effects on lifetime fecundity of Heterakis gallinarum in chickens. Parasit Vectors. 2014;7:334. 79. Keymer A. Density-dependent mechanisms in the regulation of intestinal helminth populations. Parasitology. 1982;84:573–87. 80. Wang B-J, Gu X-B, Yang G-Y, Wang T, Lai W-M, Zhong Z-J, et al. Mitochondrial genomes of Heterakis gallinae and Heterakis beramporia support that they belong to the infraorder Ascaridomorpha. Infect Genet Evol. 2016;40:228–35. 81. Daş G, Hennies M, Sohnrey B, Rahimian S, Wongrak K, Stehr M, et al. A comprehensive evaluation of an ELISA for the diagnosis of the two most common ascarids in chickens using plasma or egg yolks. Parasit Vectors. 2017;10:187. 82. Lind P, Eriksen L, Nansen P, Nilsson O, Roepstorff A. Response to repeated inoculations with Ascaris suum eggs in pigs during the fattening period. II. Specific IgA, IgG, and IgM antibodies determined by enzyme-linked immunosorbent assay. Parasitol Res. 1993;79:240–4. 83. Roepstorff A, Eriksen L, Slotved HC, Nansen P. Experimental Ascaris suum infection in the pig: worm population kinetics following single inoculations with three doses of infective eggs. Parasitology. 1997;115:443–52. 84. Nejsum P, Thamsborg SM, Petersen HH, Kringel H, Fredholm M, Roepstorff A. Population dynamics of Trichuris suis in trickle-infected pigs. Parasitology. 2009;136:691–7. 85. Ferdushy T, Schou TW, Norup LR, Dalgaard TS, Thamsborf SM, Nejsum P, et al. Acquisition of resistance after continuous infection with Ascaridia galli in chickens. Parasitology. 2014;141:1603–10. 86. Degen WGJ, Van Daal N, Rothwell L, Kaiser P, Schijns VEJC. Th1/Th2 polarization by viral and helminth infection in birds. Vet Microbiol. 2005;105:163–7. http://www.deepdyve.com/assets/images/DeepDyve-Logo-lg.png Parasites & Vectors Springer Journals

Effect of the nematophagous fungus Pochonia chlamydosporia on soil content of ascarid eggs and infection levels in exposed hens

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BioMed Central
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Copyright © 2018 by The Author(s).
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Biomedicine; Parasitology; Entomology; Tropical Medicine; Infectious Diseases; Veterinary Medicine/Veterinary Science; Virology
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10.1186/s13071-018-2898-1
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Abstract

Background: The nematophagous fungus Pochonia chlamydosporia can degrade ascarid (e.g. Ascaridia galli) eggs in agar and soil in vitro. However, it has not been investigated how this translates to reduced infection levels in naturally exposed chickens. We thus tested the infectivity of soil artificially contaminated with A. galli (and a few Heterakis gallinarum) eggs and treated with P. chlamydosporia. Sterilised and non-sterilised soils were used to examine any influence of natural soil biota. Methods: Unembryonated eggs were mixed with sterilised (S)/non-sterilised (N) soil, either treated with the fungus (F) or left as untreated controls (C) and incubated (22 °C, 35 days) to allow eggs to embryonate and fungus to grow. Egg number in soil was estimated on days 0 and 35 post-incubation. Hens were exposed to the soil (SC/SF/ NC/NF) four times over 12 days by mixing soil into the feed. On day 42 post-first-exposure (p.f.e.), the hens were euthanized and parasites were recovered. Serum A. galli IgY level and ascarid eggs per gram of faeces (EPG) were examined on days -1 and 36 (IgY) or 40 p.f.e. (EPG). Results: Egg recovery in SF soil was substantially lower than in SC soil, but recovery was not significantly different between NF and NC soils. SF hens had a mean worm count of 76 whereas the other groups had means of 355– 453. Early mature/mature A. galli were recovered from SF hens whereas hens in the other groups harboured mainly immature A. galli. Heterakis gallinarum counts were low overall, especially in SF. The SF post-exposure IgY response was significantly lower while EPG was significantly higher compared to the other groups. Conclusions: Pochonia chlamydosporia was very effective in reducing ascarid egg numbers in sterilised soil and thus worm burdens in the exposed hens. However, reduced exposure of hens shifted A. galli populations toward a higher proportion of mature worms and resulted in a higher faecal egg excretion within the study period. This highlights a fundamental problem in ascarid control: if not all eggs in the farm environment are inactivated, the resulting low level infections may result in higher contamination levels with associated negative long-term consequences. Keywords: Ascaridia galli, Heterakis gallinarum, Fungus, Biological control * Correspondence: sundar@sund.ku.dk Section for Parasitology and Aquatic Pathobiology, Department of Veterinary and Animal Sciences, Faculty of Health and Medical Sciences, University of Copenhagen, Dyrlægevej 100, 1870 Frederiksberg C, Denmark Full list of author information is available at the end of the article © The Author(s). 2018 Open Access This article is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated. Thapa et al. Parasites & Vectors (2018) 11:319 Page 2 of 11 Background respectively, of A. galli eggs in laboratory agar assays. Ascaridia galli and Heterakis spp., collectively known as However, the effect in a soil based assay was relatively ascarids, are economically important intestinal nema- lower (~45%) [42]. In the latter study, the fungal effective- todes of chickens worldwide. Ascaridia galli can impair ness was evaluated based on eggs recovered from soil the health [1–3], productivity [4–7] and welfare of before and after the fungal treatment. However, it is un- chickens [8]. Moreover, A. galli can reduce the vaccine known if the recovered eggs judged visually as viable are efficacy against Newcastle disease [9, 10] and increase indeed infective and whether the reduced contamination the susceptiblilty of chickens to other infectious diseases in the soil assay translates to lower worm burdens in such as fowl cholera [11]. On rare occasions, A. galli can chickens exposed to the fungus-treated soil. In addition, leave the host’s intestine, migrate up the oviduct and literature indicates that population composition of A. galli become enclosed inside one of the hen’s eggs, which is in chickens can be dose-dependent as shown by a reduced of aesthetic concern to the consumers [12, 13]. Com- number of inhibited larvae in the host’s intestine and a pared to A. galli, Heterakis spp. are less pathogenic, but shorter prepatent period in lightly infected compared to they can act as a vector for in ovo transmission of the heavily infected chickens [43, 44]. It is thus important to protozoan Histomonas meleagridis to turkeys and chick- use an in vivo infection model to assess how changes in ens [14]. Histomonas meleagridis is pathogenic [15, 16] exposure level as a result of fungal treatment of soil may and re-emerging in layer flocks in many European coun- modulate worm population dynamics within the host as tries, mainly after the ban of the prophylactic use of che- this may in turn alter on-farm transmission dynamics. motherapeutics in the European Union (EU) member The overall aim of this study was to evaluate the in countries [17–20]. vivo infectivity of soil experimentally contaminated with Recent studies have shown that A. galli and Heterakis ascarid eggs and treated with P. chlamydosporia. Both spp. are highly prevalent in European organic laying hen sterilised and non-sterilised soils were used to include flocks [21–23]. Both nematodes have a simple life-cycle any effect inherent to the natural soil biota and thus to that involves a pre-parasitic development phase evaluate the potential of P. chlamydosporia as an (i.e. free-living nematode eggs) in the environment such on-farm biocontrol agent. as litter and soil and a parasitic phase in the chicken’s intestine following ingestion of infective eggs [24, 25]. Methods Ascarid eggs have thick shells [26, 27] and they can Experimental design survive in the outdoor environment for up to 2–4 Unembryonated ascarid eggs were added to Petri dishes years [28, 29], but no effective means of inactivating with sterilised (S) or non-sterilised (N) soil, half of which eggs during or after embryonation in the farm yards were treated with spores of the fungus P. chlamydos- and pastures are currently available. At present, con- poria (F) while the other half were untreated (C). The trol of ascarid infections by farmers therefore solely Petri dishes were incubated at 22 °C for 35 days to allow relies on flock treatment with commercial anthelmin- the fungus to grow and the eggs to reach infectivity. tics. Of the anthelmintics, only flubendazole and After incubation, three subgroups of 10 hens per soil fenbendazole are available for use in layers in the EU treatment were exposed in-feed four times to the soil [30, 31]. As hens are rapidly reinfected due to con- from one of the four treatments (SC, SF, NF and NC). tinuous exposure to eggs present in the surroundings The hens were euthanized on day 42 post-first-exposure and do not appear to acquire protective immunity (p.f.e.) and examined for A. galli and Heterakis spp. To [32–34], repeated treatments are thus necessary. estimate the number of eggs in the Petri dishes, recovery However, overuse of these drugs may over time en- of eggs was tested before [i.e. day 0 post-incubation hance the risk of selecting for anthelmintic resistance. (p.i.)] and after incubation (day 35 p.i.) for each of the To be able to also combat the parasites in the environ- four soil treatments. ment, there is an increasing interest in using naturally occurring soil microfungi as is done to control agricul- Origin and isolation of ascarid eggs tural nematode pests [35]. An isolate of Pochonia Before collecting faeces, the infection status of A. galli chlamydosporia (syn. Verticillium chlamydosporium) and Heterakis spp. in a Danish organic layer farm with a (Ascomycota: Hypocreales), a microfungus of global oc- flock size of 3000 hens was examined through necropsy currence [36–38], that can mechanically and enzymati- of 18 randomly selected hens. The prevalence was 89% cally degrade the egg shell components (protein and for A. galli and 100% for Heterakis spp., and mean ± S.E. chitin) has already been developed as a biocontrol agent worm burdens were 40 ± 9 and 80 ± 21 worms, respect- against plant-parasitic nematode eggs [39]. The same iso- ively. Ascarid eggs (A. galli and Heterakis spp.) were iso- late [40] and two other isolates [41]of P. chlamydosporia lated from fresh hen faeces collected from the ground have subsequently been shown to kill ~70% and > 80%, using only the top part of the faeces as described by Thapa et al. Parasites & Vectors (2018) 11:319 Page 3 of 11 Thapa et al. [29]. The eggs were stored in sterile demi- (i.e. pre-incubation) soil moisture level per dish (23– neralised water at 5 °C for 6 days. Before use, a subsample 24% of the total soil weight) was estimated (105 °C, 24 of eggs was embryonated in 0.1 N H SO at 25 °C for 15 h). Five random dishes per treatment were used to esti- 2 4 days to assess percentage embryonation (i.e. ability to mate day 0 p.i. egg recovery, while the remaining dishes develop larva) of the egg batch [42] which was 95 ± 1% were incubated at 22 °C for 35 days in darkness. The (mean ± SE). weight of each incubated dish was recorded at days 0 and 35 p.i. to determine soil moisture loss (%). On day Preparation of fungal inoculum 35 p.i., the dishes were opened and 4 ml sterile water Parboiled rice initially soaked for 1 h in demineralised was added to each dish, re-sealed with parafilm and water and autoclaved (121 °C, 15 min) inside a polypropyl- stored at 10 °C for up to 18 days. On day 7 post-storage ene bag (Labsolute®, 300 g rice per bag) was inoculated at 10 °C (i.e. after incubation was terminated), egg with 10 ml P. chlamydosporia Biotype 10 spore suspen- recovery (i.e. exposure level) was estimated in five 7 4 sion [5.5 × 10 conidia and 1.6 × 10 chlamydospores in random dishes per treatment (see section on recovery 0.05% Triton® X-100 (Merck KGaA, Darmstadt, Germany) of eggs from soil). Soil from one random dish was harvested from 5-week-old culture in Sabouraud’sdex- selected and administered in the feed to a correspond- trose agar]. After incubating the rice at 25 °C for 25 days ing subgroup of hens (see section on animal exposure in darkness, 30 ml 0.05% Triton® X-100 was added to each to parasites). This exposure was repeated on days 11, 10 g of rice granules in centrifuge tubes and shaken gently 15 and 19 after incubation was terminated. to separate spores from the rice. The mixture was filtered through a 900 μm sieve to remove rice particles and cen- Recovery of eggs from soil trifuged (1831× g, 3 min) three times after re-suspension Fifty millilitres of 0.5 M NaOH was added to each of the in 0.05% Triton® X-100. Spore concentration was adjusted 40 dishes (n = 5 per treatment on day 0 and 35 p.i.) that 8 5 to 4 × 10 conidia and 1.5 × 10 chlamydospores per ml was then stored at 5 °C for 16 h. The soil was washed suspension. Spore germination was determined [42]tobe through 212 and 20 μm sieves and the material on the 96% and 91% for the conidia and chlamydsopores, latter was divided into four 50 ml tubes, centrifuged at respectively. 253× g for 7 min and the eggs were recovered as described by Thapa et al. [29]. For each dish, the egg Preparation of soil quantity and development stage (unembryonated, In April 2016, 15 kg sandy loam soil (pH 6.8) was pre-larvated, larvated or degenerated) was examined in a collected from a Danish experimental plot that was 20% subsample at 100× magnification [29]. established in 2002 and treated anually (2003–2015) with source separated organic household waste compost (CH) and sown with spring cereals [45, 46]. After Experimental animals and housing removing plant material and stones, the soil was sieved One hundred thirty pullets (ISA Warren, 18-weeks-old), (3 mm) and thoroughly homogenised. Three kilograms of raised indoors without previous anthelmintic treatment, soil was sterilised by autoclaving (121 °C, 30 min) inside a were obtained from a commercial breeder. On arrival polypropylene bag (Labsolute®, 200 g soil per bag, (day -15 p.f.e.), 10 randomly selected pullets were eutha- treatment S) while another 3 kg soil was kept without nized and examined for ascarid infections of the breeder autoclaving (treatment N). farm-origin (see section on recovery of worms). All necropsied pullets were found positive for tissue phase Fungal treatment of soil and eggs A. galli larvae (~0.5 mm long) with an overall mean ± For both soils (S, N), 44 replicate Petri dishes (14.5 × 2 cm) SE worm burden of 194 ± 97 A. galli, while only two each containing 46 g soil and 2 ml ascarid egg suspension birds harboured luminal Heterakis spp. giving an with approximately 8000 ± 260 eggs (mean ± SE) in sterile overall burden of 1 ± 1 worm per hen. The remaining water were prepared. Both soil types were randomised into pullets (n = 120), after random allocation into 12 control (C) and fungus treatment (F). To all SF and NF indoor pens (c.2.8 m ,10pullets perpen), were dishes, 2 ml fungal suspension containing approxi- therefore treated with flubendazole (Verminator®, 1.43 8 5 mately 8 × 10 conidia and 3 × 10 chlamydospores of mg flubendazole per kg live weight daily) in the feed P. chlamydosporia in 0.05% Triton® X-100 was added from days -13 to -6 p.f.e. The individual body weight whereas all SC and NC dishes received 2 ml 0.05% of all birds was measured on days -1 and 36 p.f.e. Triton® X-100 without fungus. The SC and SF dishes The birds were given pelleted feed (17.5% crude received an additional 635 μl of sterile water to balance protein, 4.5% crude fat) in two meals (110 g feed per the total moisture level between the S and N soils. The bird per day) and water ad libitum. Crushed oyster dishes were sealed with Parafilm ‘M’® and the initial shells were offered daily. Wood-chips and straw were Thapa et al. Parasites & Vectors (2018) 11:319 Page 4 of 11 used as bedding material. Pens were enriched with a antigens and one replicate serum sample per animal per perch and nests, and cleaned thoroughly once weekly. sampling day. A dilution series of a highly positive serum was used as standard and the highest concentration was Animal exposure to parasites set at the relative value 2. The 12 pens were allocated to the four treatment groups (SC, SF, NC and NF) in triplets (i.e. three subgroups per Statistical analyses treatment group). The hens were exposed to ascarid All statistical analyses were performed using SAS 9.4 contaminated soil on days 0, 4, 8 and 12 p.f.e. to mimic a (Cary, NC, USA). The main and interaction effects of moderate trickle infection. On each exposure, entire soil soil sterility (S, N), fungal treatment (C, F) and incuba- from one Petri dish was transferred to a 500 ml container tion time (days 0, 35 p.i.) on egg recovery from soil were with 150 g feed of the morning meal and 50 ml tapwater, analysed using a generalised linear model fitted with mixed thoroughly and spread in a tray (58 × 21 × 3 cm) in negative binomial distribution of errors (NBD) (proced- each pen. The feed was eaten within 10–15 min and the ure GENMOD). Soil moisture loss during incubation remainder of the meal was then given in the same tray. was analysed with a linear model (procedure GLM) with percent moisture loss as the outcome and soil sterility Recovery of worms (S, N), fungal treatment (C, F) and their interaction as The hens were euthanized by stunning and cervical predictors. Body weight at day 0 p.f.e. and weight gain dislocation on day 42 p.f.e. The A. galli worms in the (days 0 to 36 p.f.e.) in relation to soil sterility (S, N) and small intestinal lumen were isolated using an agar-gel fungal treatment (C, F) were analysed separately with a method [47] and collected using a 20 μm sieve. The linear-mixed model (procedure MIXED) with subgroup tissue phase larvae of A. galli (day -15 and 42 p.f.e.) and (i.e. pen) as a random effect. Worm burden (total, A. Heterakis spp. (day -15 p.f.e.) were isolated from the galli, H. gallinarum), proportion (%) of A. galli in the in- intestinal/caecal tissue by pepsin (1:3000 IU)-HCl (30%) testinal tissue, proportion (%) of A. galli in each length digestion [47] and collected on a 20 μm sieve. To category (< 0.5, 0.5–1.5, 1.5–3.0, 3.0–5.0, 5.0–8.0 cm) recover luminal Heterakis spp. (day -15 and day 42 and at day 40 p.f.e. EPG was analysed with a generalised p.f.e.), the caeca were opened and stored in tap water at linear mixed model (procedure GLIMMIX, NBD) that 5 °C. After 48 h, the caeca and contents were washed on included soil sterility (S, N), fungal treatment (C, F) and a20 μm sieve. All worm samples were stored in 70% their interaction as fixed effects and subgroup as a ran- ethanol and examined using a dissection microscope dom effect. The log-transformed IgY titre was analysed (30–40× magnification). All A. galli worms were cate- with a linear-mixed model (procedure MIXED) with soil gorised as < 0.5, 0.5–1.5, 1.5–3.0, 3.0–5.0 or 5.0–8.0 cm, sterility (S, N), fungal treatment (C, F), sampling time whereas Heterakis spp. were categorised as < 0.5 or ≥ 0.5 (days -1, 36 p.f.e.) and their interaction as fixed effects, cm. Moreover, Heterakis species were determined based subgroup as a random effect and individual bird as a on the length of the spicules [48, 49] of 50 randomly repeated measurement. At group level, the linear selected male worms (1 worm per hen and representing relationships between worm burden (total ascarid or A. all experimental groups) after exposing each worm to a galli) and the IgY titre difference between pre- and drop of 10% lactic acid in water (weight/weight). post-exposures were examined using a Spearman method (procedure CORR). The goodness of fit of each Faecal egg counts GENMOD and GLIMMIX model was assessed with the Individual faecal samples from all birds were collected ratio of Pearson’s χ and corresponding degress of free- on days -1 and 40 p.f.e. Ascarid eggs per gram faeces dom. The normality of residuals of each GLM and (EPG) was determined by a concentration McMaster MIXED model was examined by a q-q plot and a histo- technique (minimum detection limit: 20 EPG) using a gram, and homogeneity of residual variance assessed by flotation fluid of 500 g glucose monohydrate per litre of residual plots. For each model, the post-hoc significant saturated NaCl solution (specific gravity: 1.27) [50]. differences were determined with the differences of least squares means (Tukey-Kramer’s adjustment for multiple Ascaridia galli antibody (IgY) levels comparisons, P < 0.05). To determine the systemic antibody response as an indirect assessment of parasite exposure, individual Results blood samples from all birds were collected on days -1 Recovery of eggs from soil and 36 p.f.e. from a wing vein. Serum was separated by On day 0 p.i., the mean number of eggs recovered from centrifugation at 1000× g for 15 min and stored at -20 °C. the SC, SF, NC and NF soils were 8702–9673 with no sig- The A. galli IgY level was determined by ELISA according nificant differences between the treatments (P >0.9950 in to Norup et al. [51] using crude adult A. galli somatic all cases) (Fig. 1). Irrespective of treatment, > 97% of the Thapa et al. Parasites & Vectors (2018) 11:319 Page 5 of 11 Worm burdens The overall mean worm burdens of A. galli and H. galli- narum in hens in the four groups are shown in Fig. 2a and b, respectively. All 118 hens were A. galli positive, while 115 birds were H. gallinarum positive. The left and right spicules of H. gallinarum males had a mean ± SE length of 2086 ± 26 μm (range: 1554–2417 μm) and 723 ± 8 μm (range: 402–850 μm), respectively. The interaction between the soil sterility and fungal treatment strongly in- fluenced the total ascarid worm burden (F = 100.38, (1, 106 ) P < 0.0001) and the individual worm burdens of both A. galli (F = 96.85, P < 0.0001) and H. galli- (1, 106) narum (F = 10.07, P = 0.0020). Group SF hens (1, 106) Fig. 1 Mean (+ SE) number of ascarid eggs recovered from soil on thus had significantly lower worm burdens of both A. galli days 0 and 35 post-incubation at 22 °C. Approximately 8000 (P < 0.0001 in all cases) and H. gallinarum (P ≤ 0.0001 in unembryonated eggs were added to soil given four different all cases) compared to the three other groups that all had treatments (n =5) (Abbreviations: SC, sterilised control; SF, sterilised comparable A. galli (P > 0.3120 in all cases) and H. galli- with the fungus Pochonia chlamydosporia Biotype 10; NC, non- sterilised control; NF, non-sterilised with fungus). Different letters narum worm burdens (P > 0.9989 in all cases) (Fig. 2). above the bars indicate significant differences (P < 0.05, Heterakis gallinarum represented 6% of the total ascarids Tukey-Kramer’s adjustment for multiple comparisons) recovered eggs were unembryonated. On day 35 p.i., the mean egg number in the SC, SF, NC and NF soils was 5535 (36% reduction), 521 (94% reduction), 4176 (57% reduction) and 3201 eggs (65% reduction), respectively (Fig. 1). In the sterilised soil, the fungal treatment resulted in a significant reduction in egg recovery when compared to the control (P < 0.0001). In contrast, there was no such difference in the non-sterilised soil (P = 0.5480). This meant that there was a strong significant (χ = 70.72, df =4, P < 0.0001) interaction between soil sterility, fungal treatment and incubation time on egg recov- ery. Regardless of treatment, ~94% of the recovered eggs at day 35 p.i. contained a slender larva that re- sembled the infective stage. The mean ± S.E. moisture loss in the sterilised soil (28 ± 1.8%) was slightly but signifi- cantly higher than in the non-sterilised soil (21 ± 1.4%) (F =8.53, P = 0.0048). (1, 64) Clinical observations and performance On day 0 p.f.e., the overall mean live weight of hens in the four groups was 1.53–1.60 kg with no significant effect of soil sterility (F =2.29, P = 0.1330), fungal (1, 106) treatment (F =1.91, P = 0.1701) and their inter- (1, 106) action (F =0.82, P = 0.3684). By day 36 p.f.e., the (1, 106) overall mean weight gain of hens in the four groups Fig. 2 Mean (+ SE) total worm burdens of Ascaridia galli (a) and Heterakis spp. (b) recovered from four groups of hens 42 days after was -76 to 90 g, but there was no significant effect of the first (of the total four) in-feed exposures to ascarid eggs soil sterility (F = 3.39, P = 0.0686), fungal treat- (1, 106) embryonated in sterilised control soil (SC), sterilised soil with the ment (F = 0.04, P = 0.8367) and their inter- (1, 106) fungus Pochonia chlamydosporia Biotype 10 (SF), non-sterilised action (F = 2.72, P = 0.1022). Most hens started (1, 106) control soil (NC) or non-sterilised soil with P. chlamydosporia (NF). laying eggs from days 3–7 p.f.e. During the study, the Each bar represents the mean of 28–30 hens allocated to three replicate subgroups of 8 (one NC subgroup) to 10 hens. Different hens showed no overt signs of illness but two hens letters above the bars indicate significant differences (P < 0.05, from one of the three NC subgroups died, possibly Tukey-Kramer’s adjustment for multiple comparisons) due to cannibalism. Thapa et al. Parasites & Vectors (2018) 11:319 Page 6 of 11 for SF hens and 2–3% for the three other groups. With reference to the estimated cumulative egg dose of 2214 (SC), 208 (SF), 1670 (NC) and 1281 eggs (NF) that each hen was theoretically exposed to on four exposures, the overall establishments of total ascarid were 20% (SC), 36% (SF), 21% (NC) and 33% (NF). Parasite population composition The overall mean proportion of A. galli recovered from the intestinal lumen and intestinal wall is shown in Fig. 3. In general, A. galli were more prevalent in the intestinal lumen (61–78%) than in the intestinal tissue (22–39%). Fungal treatment had a significant effect on the rela- tive distribution of tissue phase and luminal phase A. galli (F =9.85, P = 0.0022). This resulted in a (1, 106) significantly higher proportion (37 ± 2%, mean ± S.E.) of tissue phase A. galli in hens not exposed to fungal treatments compared to the hens exposed to fungal treatments (27 ± 2%). There were no significant ef- fects of soil sterility (F = 3.05, P = 0.0660) as (1, 106) well as the interaction between fungal treatment and Fig. 4 Mean proportion (%) of Ascaridia galli of different sizes soil sterility (F =0.50, P = 0.4815) on the A. (1, 106) recovered from four groups of hens 42 days after the first (of the galli distribution between intestinal lumen and tissue. total four) in-feed exposures to ascarid eggs embryonated in The proportion of A. galli (of the total A. galli worm sterilised control soil (SC), sterilised soil with the fungus Pochonia chlamydosporia Biotype 10 (SF), non-sterilised control soil (NC) or burden) within each length category was significantly re- non-sterilised soil with P. chlamydosporia (NF). Each bar represents lated to the interaction between soil sterility and fungal the mean of 28–30 hens allocated to three replicate subgroups of 8 treatment (< 0.5 cm: F =29.48, P < 0.0001; 0.5–1.5 (1, 106) (one NC subgroup) to 10 hens. Different letters above the bars cm: F =9.89, P = 0.0022; 1.5–3.0 cm: F = (1, 106) (1, 106) indicate significant differences (P < 0.05, Tukey-Kramer’s adjustment 29.98, P < 0.0001; 3.0–5.0 cm: F =16.58, P < 0.0001; for multiple comparisons) within each length category (1, 106) 5.0–8.0 cm: F =7.24, P = 0.0083) (Fig. 4). Compared (1, 106) to the groups SC, NC and NF hens, the group SF hens hosted a significantly lower proportion of A. galli <0.5 cm (P < 0.0001 in all cases) and significantly higher propor- tions of the three largest length categories (P < 0.0225 in all cases, except P =0.0508for SF vs NF in the category 5.0–8.0 cm). The SC, NC and NF hens hosted nearly equal proportions of all five A. galli length categories (P > 0.0625 in all cases, except P =0.0076 for SC vs NC in the category 1.5–3.0 cm). Irrespective of group, all tissue phase A. galli larvae were < 0.5 cm (~0.5 mm). In groups SC, NC and NF hens, the luminal A. galli worms within the catergory < 0.5 cm (i.e. 5 mm) were approximately 0.5–1.0 mm whereas those in group SF hens ranged ~0.5–4.9 mm. For H. gallinarum, the highest mean ± S.E. proportion of worms > 0.5 cm was hosted by the group SF hens (52 ± Fig. 3 Mean proportion (%) of luminal phase and tissue phase 9%) followed by NC (38 ± 6%), NF (31 ± 5%) and SC hens Ascaridia galli recovered from four groups of hens 42 days after (25 ± 6%). However, the effect of soil sterility, fungal the first (of the total four) in-feed exposures to ascarid eggs treatment and their inferactions on H. gallinarum embryonated in sterilised control soil (SC), sterilised soil with the fungus Pochonia chlamydosporia Biotype 10 (SF), non-sterilised population composition was not possible to analyse control soil (NC) or non-sterilised soil with P. chlamydosporia (NF). using the same statistical model that was used for A. Each bar represents the mean of 28–30 hens allocated to three galli because many hens had only < 0.5 or ≥ 0.5 cm replicate subgroups of 8 (one NC subgroup) to 10 hens H. gallinarum. Thapa et al. Parasites & Vectors (2018) 11:319 Page 7 of 11 Faecal egg counts On day -1 p.f.e., all hens were negative for ascarid eggs. On day 40 p.f.e., 3, 57, 7 and 17% hens of groups SC, SF, NC and NF, respectively, had positive EPG. There was a signifi- cant interaction between fungal treatment and soil sterility regarding day 40 p.f.e. EPG (F =5.17, P < 0.0250) as (1, 106) the overall mean EPG in group SF hens was signifi- cantly higher than in groups SC (P < 0.0001) and NC hens (P = 0.0241) but comparable to group NF hens (P = 0.4040) (Fig. 5). Ascaridia galli IgY titres The overall group mean (+ SE) A. galli IgY titres in hens in the four groups on days -1 and 36 p.f.e. are shown in Fig. 6 Mean (+ SE) Ascaridia galli IgY titre at one day before and 36 Fig. 6. All hens were seropositive at both time-points. days after the first (of the total four) in-feed exposures to ascarid The antibody titer was significantly affected by the inter- eggs embryonated in sterilised control soil (SC), sterilised soil with action between sterility of soil, fungal treatment and the fungus Pochonia chlamydosporia Biotype 10 (SF), non-sterilised control soil (NC) or non-sterilised soil with P. chlamydosporia (NF). sampling time (F = 14.02, P < 0.0001). On day -1 (4, 212) Each bar represents the mean of 28–30 hens allocated into three p.f.e., the group mean ± SE titres ranged between 734 ± replicate subgroups of 8 (one NC subgroup), 9 (two SC and two NF 62 and 1071 ± 115, with no significant differences be- subgroups) or 10 hens. Different italicised letters above the bars tween the groups (P > 0.7380 in all cases). By day 36 indicate significant differences (P < 0.05, Tukey-Kramer’s adjustment p.f.e., the IgY titre had increased significantly in all for multiple comparisons) between the log-transformed titres groups with an overall 8–11 fold increase in groups SC (P < 0.0001), NC (P < 0.0001) and NF (P < 0.0001), but only three fold increase in group SF hens (P < 0.0001). Discussion There were no significant correlations (P > 0.05) Thepresent studyhas forthe firsttimeshown that between IgY titre and individual worm burden (total ascarid transmission to hens exposed to egg contami- ascarid, total A. galli) in all groups except SF where nated soil can be reduced after the soil has been there were significant but weak correlations for the total treated with the fungus P. chlamydosporia, but only ascarid (r = 0.38, P = 0.0362) and A. galli worm bur- in sterilised soil. The reduced exposure resulted in a (30) den (r = 0.40, P = 0.0275). higher rate of development into adult worms and (30) thus more patent infections compared to the more heavily infected control hens. The fungus P. chlamydosporia Biotype 10 substantially reduced the egg recovery in the sterilised soil whereas in non-sterilised soil there was no additional effect when compared to the corresponding controls. This limited effect of P. chlamydosporia in the non-sterilised soil is in line with previous findings for egg-degrading fungi in general [38, 42, 52, 53]. The currently available literature indicates that native soil biota can reduce the establish- ment of a newly added fungus [54–58]. This is probably because the new fungus must compete for the soil re- sources or overcome antagonism by native established soil biota such as other fungi [59–61], bacteria [62–65], protozoa [66], free-living nematodes [67, 68], mites and Fig. 5 Mean (+ SE) number of ascarid eggs per gram of faeces (EPG) dipteran larvae [69]. In future studies, application of of four groups of hens 40 days after the first (of the total four) in- fungi in nutrient-rich substrates (e.g. rice or barley kernels, feed exposures to ascarid eggs embryonated in sterilised control soil decomposed resources etc.) could be explored as this may (SC), sterilised soil with the fungus Pochonia chlamydosporia Biotype help increase fungal establishment in soil [55, 70, 71]. 10 (SF), non-sterilised control soil (NC) or non-sterilised soil with P. chlamydosporia (NF). Each bar represents the mean of 28–30 hens Ascarid eggs are sensitive to dessication [72] and after allocated to three replicate subgroups of 8 (one NC subgroup) to 10 incubation, we found a slightly higher moisture loss in the hens. Different letters above the bars indicate significant differences sterilised soil compared to the non-sterilised soil. How- (P < 0.05, Tukey-Kramer’s adjustment for multiple comparisons) ever, results indicate that this had no major impact as the Thapa et al. Parasites & Vectors (2018) 11:319 Page 8 of 11 moisture loss in the control and the fungus-treated and the effect of host immune responses on parasite sterilised soil was not significantly different and both the population seem important [79]. highest moisture loss and highest egg recovery from a The above findings highlight the basic complication of single Petri dish was found in the sterilised soil. any control strategy that cannot inactivate all parasite The differences in soil egg numbers (i.e. exposure eggs in the environment. Initially there may be a lowered levels) after fungal treatment was reflected in vivo by impact on the hosts present at the time due to lowered parasite burden, establishment rate and population com- exposure, but if the result is associated with altered in- postion within the host. The least exposed group had fection dynamics, and thus an earlier onset of patency, the lowest ascarid worm burdens but a higher parasite environmental recontamination might be higher than if establishment rate compared to the three other groups there had been no intervention. This is further compli- that were more heavily exposed. Similar findings have cated as freshly deposited eggs take weeks to months to been reported for an A. galli trickle infection in chickens develop to infectivity depending on weather and season [32], H. gallinarum single infection in red-necked pheas- [30]. This goes to show that designing and implementing ants [73]and Oesophagostomum dentatum single and control strategies on a farm must take parasite biology trickle infections in pigs [74–76]. In contrast, Permin et and ecology into account to not only offer temporary al. [77] found no differences in A. galli burdens follow- relief, but also be effective long-term. ing a single dose of 100, 500 or 2500 eggs. This may be There is a close phylogenetic relationship between A. because they only quantified the luminal worms and galli and Heterakis spp. [80] with corresponding produc- many larvae in the two higher dose groups may poten- tion of cross-reacting antibodies [81]. However, the tially have been, at least temporarily, arrested in the current contribution in the IgY titre due to H. gallinarum intestinal mucosa [43, 44]. In the current study, we thus is expected to be neglible due to the much lower worm found an increased proportion of tissue phase A. galli burdens compared to A. galli. The individual antibody presumably at the third larval stage (L3) [43, 44] in the levels appeared to increase with increasing exposure level three high exposure groups, and larger worms and pa- and worm burden. However, individual antibody levels tent infections primarily in the lowest exposure group. seemed uninvolved in any immune-related short-term The absence of patent infections in most heavily regulation of A. galli populations. This is in agreement infected hens supports that faecal egg counts can with previous findings of a very weak or a complete lack severely underestimate immature worm burdens and of correlation between systemic/egg-yolk IgY level and A. exposure levels [44, 74, 76]. galli/H. gallinarum worm burden [3, 81]. A similar lack of The current results showed that low exposure may at association between porcine blood IgG level and worm least, in the short-term, lead to mature A. galli popula- burdens has been reported for Ascaris suum [82, 83]and tions in contrast to predominantly immature infections Trichuris suis [84]. Furthermore, we also found that previ- at higher exposure levels. Reduced exposure and lower ous exposure did not protect against subsequent A. galli worm burdens are both desireable to lower the overall reinfection, which is in line with other studies [32, 85]. impact of ascarids on chicken health and productivity Others have reported increased mRNA expression of Th2 but seem to favour the establishment of patent infec- cytokines IL-4 and IL-13 in the intestinal tissues and tions. Density-dependent worm maturation was previ- spleen of A. galli infected hosts [3, 86, 87]. Both cytokines ously documented for H. gallinarum in chickens and play a role in mediating protective immunity against sev- ring-necked pheasants where heavily infected birds eral helminth parasites [88, 89] but it appears that A. galli hosted significantly smaller female worms compared to may evade host immune responses to avoid expulsion as lightly infected birds [73, 78]. It is unknown if, given suggested for O. dentatum in pigs [90, 91]. This could be time, at least some of our arrested larvae, presumably a reason why A. galli prevalence in laying hens kept in L3 in the intestinal tissue and L4 in the intestinal non-cage systems (barn, free-range and organic) seems to lumen [1, 43], would have reached maturity as our hens increase over time during an egg laying period of approxi- were only followed for 30 days after the last exposure. mately one year [34]. However, Ikeme [44] found the development of nearly all To the best of our knowledge, there are no opti- L3 to be arrested for up to 13 weeks post-last-exposure in mal/standardized protocols to establish patent A. galli birds that received a high infection dose. It is therefore infections in chickens. Experimental infection proce- very important to use sensitive recovery techniques to dures vary greatly in relation to infection material minimize the risk of overlooking high immature worm (source, embryonation medium, temperature and dur- burdens. The precise mechanisms responsible for ation of embryonation) and host factors (age, breed, density-dependent effects are not fully understood. The etc.) [10, 32, 92–96]. This makes it extremely difficult combination of intraspecific competition among worms to compare results between different studies. Many for limited resources (e.g. space, nutrients) in the host gut experiments performed earlier by our group could Thapa et al. Parasites & Vectors (2018) 11:319 Page 9 of 11 not establish patent A. galli infections when chickens Ethics approval The animal experiment was approved by the Animal Experiments were either infected with a single dose of 500 eggs Inspectorate, The Danish Ministry of Food, Agriculture and Fisheries [97, 98] or trickle infected twice weekly with 25–100 (Permit Number: 2015-15-0201-00760). The animals were treated according eggs per infection over a period of six weeks [32, 85]. to the Danish ethical guidelines. The lowest infection dose used in the latter studies is Competing interests very similar to the lowest exposure level of the The authors declare that they have no competing interests. current study. We have therefore made some modifi- cations in the current protocol in relation to the pre- Publisher’sNote vious failures. Hens were exposed to ascarid eggs at Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. 20–22 weeks of age. This corresponded to the period when most hens started to lay eggs and it has been sug- Author details gested that hens during this period are more susceptible Section for Parasitology and Aquatic Pathobiology, Department of Veterinary and Animal Sciences, Faculty of Health and Medical Sciences, and may have an increased establishment of A. galli due University of Copenhagen, Dyrlægevej 100, 1870 Frederiksberg C, Denmark. to hormonal changes in the birds [92]. Furthermore, we 2 College of Veterinary Medicine, Inner Mongolia Agricultural University, embryonated (i.e. incubated) ascarid eggs at 22 °C for only Hohhot 010018, People’s Republic of China. Section for Organismal Biology, Department of Plant and Environmental Sciences, Faculty of Science, five weeks compared to the six week incubation protocol University of Copenhagen, Thorvaldsensvej 40, 1871 Frederiksberg C, in the earlier studies. This was because chicken ascarid 4 Denmark. Section for Immunology and Microbiology, Department of Animal eggs develop fully within four weeks of incubation at Science, Aarhus University, Blichers Allé 20, Building P25, 3334, 8830 Tjele, Denmark. Section for Diagnostics and Scientific Advice, National Veterinary 22 °C [29] and we thus provided only one additional Institute, Technical University of Denmark, Kemitorvet, 2800 Kgs. Lyngby, week for the developed eggs to mature to infectivity. Denmark. Received: 15 February 2018 Accepted: 14 May 2018 Conclusions The Biotype 10 strain of P. chlamydosporia was only effective in inactivating ascarid eggs and thereby reducing References 1. Ikeme MM. Observations on the pathogenecity and pathology of Ascaridia the infection level of sterilised soil and thus total worm galli. Parasitology. 1971;63:169–79. burdens of the exposed hens. The consequence was that 2. Ramadan HH, Abou Znada NY. Some pathological and biochemical studies the proportion of early mature/mature A. galli increased on experimental ascaridiasis in chickens. Nahrung. 1991;35:71–84. 3. Schwarz A, Gauly M, Abel H, Daş G, Humburg J, Rohn K, et al. and faecal eggs counts were higher than in all the Immunopathogenesis of Ascaridia galli infection in layer chicken. Dev Comp other groups where hens were exposed to a higher Immunol. 2011;35:774–84. number of infective eggs in the soil. This underlines an 4. Reid WM, Carmon JL. Effects of numbers of Ascaridia galli in depressing weight gains in chicks. J Parasitol. 1958;44:183–6. inherent dilemma and complexity of ascarid control in that 5. Kilpinen O, Roepstorff A, Permin A, Nørgaard-Nielsen G, Lawson LG, hosts may suffer less short-term, but current reduced Simonsen HB. Influence of Dermanyssus gallinae and Ascaridia galli exposure may lead to long-term higher environmental infections on behaviour and health of laying hens (Gallus gallus domesticus). Br Poult Sci. 2005;46:26–34. re-contamination if not all eggs can be eliminated. This 6. Phiri IK, Phiri AM, Ziela M, Chota A, Masuku M, Monrad J. Prevalence needs to be considered in future biological and other and distribution of gastrointestinal helminths and their effects on control strategies in poultry. weight gain in free-range chickens in Central Zambia. Trop Anim Health Prod. 2007;39:309–15. Abbreviations 7. Skallerup P, Luna LA, Johansen MV, Kyvsgaard NC. The impact of natural EPG: eggs per gram of faeces; FEC: faecal egg counts; SE: standard error helminth infections and supplementary protein on growth performance of free-range chickens on smallholder farms in El Sauce, Nicaragua. Prev Vet Acknowledgements Med. 2005;69:229–44. The authors gratefully acknowledge Rothamsted Research Ltd. (United 8. Gauly M, Duss C, Erhardt G. Influence of Ascaridia galli infections and Kingdom) for kindly providing Pochonia chlamydosporia Biotype 10. We are anthelmintic treatments on the behaviour and social ranks of laying hens very thankful to Lise-Lotte Christensen, Suraj Dhakal, Lene R. Dal, Sara (Gallus gallus domesticus). Vet Parasitol. 2007;146:271–80. Almeida, Sophie Stolzenbach, Tina V. A. Hansen and Eline P. Hansen for the 9. Hørning G, Rasmussen S, Permin A, Bisgaard M. Investigations on the technical assistance. influence of helminth parasites on vaccination of chickens against Newcastle disease virus under village conditions. Trop Anim Health Prod. Funding 2003;35:415–24. This research and the PhD fellowship of ST were financed by the Faculty of 10. Pleidrup J, Dalgaard TS, Norup LR, Permin A, Schou TW, Skovgaard K, et al. Health and Medical Sciences, University of Copenhagen, Denmark. Ascaridia galli infection influences the development of both humoral and cell-mediated immunity after Newcastle disease vaccination in chickens. Availability of data and materials Vaccine. 2014;32:383–92. The datasets generated and analysed in the current study can be available 11. Dahl C, Permin A, Christensen JP, Bisgaard M, Muhairwa AP, Petersen from the corresponding author upon reasonable request. KMD, et al. The effect of concurrent infections with Pasteurella multocida and Ascaridia galli on free range chickens. Vet Microbiol. Authors’ contributions 2002;86:313–24. ST, SMT, HM and NVM designed the study. ST, RW, HM and TSD conducted 12. Reid WM, Mabon JL, Harshbarger WC. Detection of worm parasites in the study. ST analysed the data and drafted the manuscript. All authors chicken eggs by candling. Poult Sci. 1973;52:2316–24. discussed the results, contributed in the manuscript and read and approved 13. Fioretti DP, Veronesi F, Diaferia M, Franciosini MP, Proietti PC. Ascaridia galli: the final manuscript. a report of erratic migration. Ital J Anim Sci. 2005;4:310–2. Thapa et al. Parasites & Vectors (2018) 11:319 Page 10 of 11 14. Lee DL. The structure and development of Histomonas meleagridis 39. Manzanilla-López HR, Esteves I, Finetti-Sialer MM, Hirsch PR, Ward E, (Mastigamoebidae: Protozoa) in the female reproductive tract of its Devonshire J, et al. Pochonia chlamydosporia: advances and challenges to intermediate host, Heterakis gallinarum (Nematoda). Parasitology. improve its performance as a biological control agent of sedentary 1969;59:877–84. endo-parasitic nematodes. J Nematol. 2013;45:1–7. 15. Windisch M, Hess M. Experimental infection of chickens with Histomonas 40. Thapa S, Meyling NV, Katakam KK, Thamsborg SM, Mejer H. A method to meleagridis confirms the presence of antibodies in different parts of the evaluate relative ovicidal effects of soil microfungi on thick-shelled eggs of intestine. Parasite Immunol. 2010;32:29–35. animal-parasitic nematodes. Biocontrol Sci Technol. 2015;25:756–67. 16. McDougald LR. Blackhead disease (histomoniasis) in poultry: a critical 41. Braga FR, Araújo JV, Araujo JM, Frassy LN, Tavela AO, Soares FEF, et al. review. Avian Dis. 2005;49:462–76. Pochonia chlamydosporia fungal activity in a solid medium and its crude extract against eggs of Ascaridia galli. J Helminthol. 2012;86:348–52. 17. Dolka B, Żbikowski A, Dolka I, Szeleszczuk P. Histomonosis - an existing problem in chicken flocks in Poland. Vet Res Commun. 2015;39:189–95. 42. Thapa S, Mejer H, Thamsborg SM, Lekfeldt JDS, Wang R, Jensen B, et al. 18. Grafl B, Liebhart D, Windisch M, Ibesich C, Hess M. Seroprevalence of Survival of chicken ascarid eggs exposed to different soil types and fungi. Histomonas meleagridis in pullets and laying hens determined by ELISA. Appl Soil Ecol. 2017;121:143–51. Vet Rec. 2011;168:160. 43. Herd RP, McNaught DJ. Arrested development and the histotropic phase of 19. Stokholm NM, Permin A, Bisgaard M, Christensen JP. Causes of mortality in Ascaridia galli in the chicken. Int J Parasitol. 1975;5:401–6. commercial organic layers in Denmark. Avian Dis. 2010;54:1241–50. 44. Ikeme MM. Effects of different levels of nutrition and continuing dosing of 20. Esquent C, De Herdt P, De Bosschere H, Ronsmans S, Ducatelle R, Van Erum poultry with Ascaridia galli eggs on the subsequent development of J. An outbreak of histomoniasis in free-range layer hens. Avian Pathol. parasite populations. Parasitology. 1971;63:233–50. 2003;32:305–8. 45. Poulsen PHB, Al-Soud WA, Bergmark L, Magid J, Hansen LH, Sørensen SJ. Effects of fertilization with urban and agricultural organic wastes in a field 21. Thapa S, Hinrichsen LK, Brenninkmeyer C, Gunnarsson S, Heerkens JLT, trial - prokaryotic diversity investigated by pyrosequencing. Soil Biol Verwer C, et al. Prevalence and magnitude of helminth infections in organic Biochem. 2013;57:784–93. laying hens (Gallus gallus domesticus) across Europe. Vet Parasitol. 2015;214:118–24. 46. López-Rayo S, Laursen KH, Lekfeldt JDS, Delle Grazie F, Magid J. Long-term 22. Jansson DS, Nyman A, Vågsholm I, Christensson D, Göransson M, Fossum O, amendment of urban and animal wastes equivalent to more than 100 years et al. Ascarid infections in laying hens kept in different housing systems. of application had minimal effect on plant uptake of potentially toxic Avian Pathol. 2010;39:525–32. elements. Agric Ecosyst Environ. 2016;231:44–53. 23. Kaufmann F, Daş G, Sohnrey B, Gauly M. Helminth infections in laying hens 47. Ferdushy T, Nejsum P, Roepstorff A, Thamsborg SM, Kyvsgaard NC. Ascaridia kept in organic free range systems in Germany. Livest Sci. 2011;141:182–7. galli in chickens: intestinal localization and comparison of methods to 24. Ackert JE. The morphology and life history of the fowl nematode Ascaridia isolate the larvae within the first week of infection. Parasitol Res. lineata (Schneider). Parasitology. 1931;23:360–79. 2012;111:2273–9. 48. Lapage G. Mönnig’s veterinary helminthology and entomology. London: 25. Clapham PA. On the life-history of Heterakis gallinae. J Helminthol. Bailliere, Tindall and Cox; 1956. 1933;11:67–86. 49. Soulsby EJL. Textbook of veterinary clinical parasitology, Volume 1. 26. Wharton D. Nematode egg-shells. Parasitology. 1980;81:447–63. Helminths. Oxford: Blackwell Scientific Publications; 1965. 27. Christenson RO, Earle HH, Butler RL, Creel HH. Studies on the eggs of Ascaridia galli and Heterakis gallinae. Trans Am Microsc Soc. 1942;61:191. 50. Roepstorff A, Nansen P. Epidemiology, diagnosis and control of helminth 28. Farr MM. Further observations on the survival of the protozoan parasite, parasites of swine. FAO Animal Health Manual. Rome: Food and Agriculture Histomonas meleagridis and eggs of poultry nematdoes in feces of infected Organization of the United Nations; 1998. birds. Cornell Vet. 1961;51:3–13. 51. Norup LR, Dalgaard TS, Pleidrup J, Permin A, Schou TW, Jungersen G, et al. Comparison of parasite-specific immunoglobulin levels in two chicken lines 29. Thapa S, Thamsborg SM, Meyling NV, Dhakal S, Mejer H. Survival and during sustained infection with Ascaridia galli. Vet Parasitol. 2013;191:187–90. development of chicken ascarid eggs in temperate pastures. Parasitology. 52. Bourne JM, Kerry BR, De Leij FAAM. Methods for the study of Verticillium 2017;144:1243–52. chlamydosporium in the rhizosphere. J Nematol. 1994;26:587–91. 30. Committee for Medicinal Products for Veterinary Use. Flubendazole 53. Monfort E, Lopez-Llorca LV, Jansson HB, Salinas J. In vitro soil receptivity (extrapolation to poultry), summary report (4). In: EMEA/CVMP/33128/2006- assays to egg-parasitic nematophagous fungi. Mycol Prog. 2006;5:18–23. Final; 2006. http://www.ema.europa.eu/docs/en_GB/document_library/ Maximum_Residue_Limits_-_Report/2009/11/WC500014292.pdf. 54. Lockwood JL, Lingappa BT. Fungitoxicity of sterilised soil inoculated with Accessed 21 Mar 2017. soil microflora. Phytopathology. 1963;58:917–20. 31. Committee for Medicinal Products for Veterinary Use. CVMP assessment report 55. Lockwood JL. Fungistasis in soils. Biol Rev. 1977;52:1–43. for Panacur AquaSol (EMEA/V/C/002008/X/00003). 2014. http://www.ema. 56. Toyota K, Ritz K, Young IM. Microbiological factors affecting the colonisation europa.eu/docs/en_GB/document_library/EPAR_-_Assessment_Report_-_ of soil aggregates by Fusarium oxysporum f. sp. raphani. Soil Biol Biochem. Variation/veterinary/002008/WC500165605.pdf. Assessed 21 Mar 2017. 1996;28:1513–21. 32. Ferdushy T, Luna-Olivares LA, Nejsum P, Thamsborg SM, Kyvsgaard NC. The 57. Mauchline TH, Kerry BR, Hirsch PR. Quantification in soil and the rhizosphere use of genetically marked infection cohorts to study changes in of the nematophagous fungus Verticillium chlamydosporium by competitive establishment rates during the time course of a repeated Ascaridia galli PCR and comparison with selective plating. Appl Environ Microbiol. infection in chickens. Int J Parasitol. 2015;45:393–8. 2002;68:1846–53. 33. Tarbiat B, Jansson DS, Tydén E, Höglund J. Comparison between 58. de Boer W, Verheggen P, Klein Gunnewiek PJ, Kowalchuk G, van Veen JA. anthelmintic treatment strategies against Ascaridia galli in commercial Microbial community composition affects soil fungistasis. Appl Environ laying hens. Vet Parasitol. 2016;226:109–15. Microbiol. 2003;69:835–44. 34. Zloch A, Kuchling S, Hess M, Hess C. Influence of alternative husbandry 59. Chen SY, Chen FJ. Fungal parasitism of Heterodera glycines eggs as systems on postmortem findings and prevalence of important bacteria and influenced by egg age and pre-colonization of cysts by other fungi. parasites in layers monitored from end of rearing until slaughter. Vet Rec. J Nematol. 2003;35:271–7. 2018;182:350. 60. Howell CR. Mechanisms employed by Trichoderma species in the biological 35. Braga FR, De Araújo JV. Nematophagous fungi for biological control of control of plant diseases: The history and evolution of current concepts. gastrointestinal nematodes in domestic animals. Appl Microbiol Biotechnol. Plant Dis. 2003;87:4–10. 2014;98:71–82. 61. Harman GE, Howell CR, Viterbo A, Chet I, Lorito M. Trichoderma species- 36. Domsch KH, Gams W, Anderson T-H. Compendium of soil fungi, Volume 1. opportunistic, avirulent plant symbionts. Nat Rev Microbiol. 2004;2:43–56. 1st ed. London: Academic Press; 1980. 62. Lopez Llorca LV, Boag B. Inhibition of Verticillium suchlasporium and other 37. Lýsek H, Fassatiová O, Cuervo Pineda N, Lorenzo Hernández N. Ovicidal nematophagous fungi by bacteria colonising Heterodera avenae females. fungi in soils of Cuba. Folia Parasitol. 1982;29:265–70. Nematol Mediterr. 1990;18:233–7. 38. Siddiqui IA, Atkins SD, Kerry BR. Relationship between saprotrophic growth 63. Ballhausen MB, van Veen JA, Hundscheid MPJ, de Boer W. Methods for in soil of different biotypes of Pochonia chlamydosporia and the infection of baiting and enriching fungus-feeding (Mycophagous) rhizosphere bacteria. nematode eggs. Ann Appl Biol. 2009;155:131–41. Front Microbiol. 2015;6:1416. Thapa et al. Parasites & Vectors (2018) 11:319 Page 11 of 11 64. Rudnick MB, van Veen JA, de Boer W. Baiting of rhizosphere bacteria with 87. Dalgaard TS, Skovgaard K, Norup LR, Pleidrup J, Permin A, Schou TW, et al. hyphae of common soil fungi reveals a diverse group of potentially Immune gene expression in the spleen of chickens experimentally infected mycophagous secondary consumers. Soil Biol Biochem. 2015;88:73–82. with Ascaridia galli. Vet Immunol Immunopathol. 2015;164:79–86. 65. Li L, Mo M, Qu Q, Luo H, Zhang K. Compounds inhibitory to 88. Anthony RM, Rutitzky LI, Urban JF, Stadecker MJ, Gause WC. Protective nematophagous fungi produced by Bacillus sp. strain H6 isolated from immune mechanisms in helminth infection. Nat Rev Immunol. fungistatic soil. Eur J Plant Pathol. 2007;117:329–40. 2007;7:975–87. 89. Finkelman FD, Wynn TA, Donaldson DD, Urban Jr JF. The role of IL-13 in 66. Ekelund F, Rønn R. Notes on protozoa in agricultural soil with emphasis on helminth-induced inflammation and protective immunity against nematode heterotrophic flagellates and naked amoebae and their ecology. FEMS infections. Curr Opin Immunol. 1999;11:420–6. Microbiol Rev. 1994;15:321–53. 90. Andreasen A, Petersen HH, Kringel H, Iburg TM, Skovgaard K, Dawson H, 67. Ruess L, Dighton J. Cultural studies on soil nematodes and their fungal et al. Immune and inflammatory responses in pigs infected with Trichuris hosts. Nematologica. 1996;42:330–46. suis and Oesophagostomum dentatum. Vet Parasitol. 2015;207:249–58. 68. Jaffee BA, Muldoon AE, Didden WAM. Enchytraeids and nematophagous 91. Andreasen A, Skovgaard K, Klaver EJ, van Die I, Mejer H, Thamsborg SM, et fungi in soil microcosms. Biol Fertil Soils. 1997;25:382–8. al. Comparison of innate and Th1-type host immune responses in 69. Ruess L, Lussenhop J. Trophic interactions of fungi and animals. In: Dighton Oesophagostomum dentatum and Trichuris suis infections in pigs. Parasite J, White FJ, Oudemans P, editors. The fungal community - its organization Immunol. 2016;38:53–63. and role in the ecosystem. 3rd ed. Boca Raton: CRC Press; 2005. p. 581–98. 92. Gauly M, Homann T, Erhardt G. Age-related differences of Ascaridia galli egg 70. Kerry B. The use of microbial agents for the biological control of plant output and worm burden in chickens following a single dose infection. parasitic nematodes. In: Jones DG, editor. Exploitation of microorganisms. Vet Parasitol. 2005;128:141–8. London: Chapman & Hall; 1993. p. 81–104. 93. Permin A, Nansen P, Bisgaard M, Frandsen F. Ascaridia galli infections in 71. Luambano ND, Manzanilla-López RH, Kimenju JW, Powers SJ, Narla RD, free-range layers fed on diets with different protein contents. Br Poult Sci. Wanjohi WJ, et al. Effect of temperature, pH, carbon and nitrogen ratios on 1998;39:441–5. the parasitic activity of Pochonia chlamydosporia on Meloidogyne incognita. 94. Rahimian S, Daş G, Gauly M. Maternal protection against Ascaridia galli? Biol Control. 2015;80:23–9. Vet Parasitol. 2017;233:43–7. 72. Tarbiat B, Jansson DS, Höglund J. Environmental tolerance of free-living 95. Ruhnke I, Andronicos NM, Swick RA, Hine B, Sharma N, Kheravii SK, et al. stages of the poultry roundworm Ascaridia galli. Vet Parasitol. 2015;209:101–7. Immune responses following experimental infection with Ascaridia galli and 73. Tompkins DM, Hudson PJ. Regulation of nematode fecundity in the ring- necrotic enteritis in broiler chickens. Avian Pathol. 2017;46:602–9. necked pheasant (Phasianus colchicus): not just density dependence. 96. Marcos-Atxutegi C, Gandolfi B, Arangüena T, Sepúlveda R, Arévalo M, Simón Parasitology. 1999;118:417–23. F. Antibody and inflammatory responses in laying hens with experimental 74. Christensen CM, Barnes EH, Nansen P, Roepstorff A, Slotved HC. primary infections of Ascaridia galli. Vet Parasitol. 2009;161:69–75. Experimental Oesophagostomum dentatum infections in the pig: Worm 97. Ferdushy T, Luna-Olivares LA, Nejsum P, Roepstorff AK, Thamsborg SM, populations at regular intervals during trickle infections with three dose Kyvsgaard NC. Population dynamics of Ascaridia galli following single levels of larvae. Int J Parasitol. 1995;25:1491–8. infection in young chickens. Parasitology. 2013;140:1078–84. 75. Roepstorff A, Nilsson O, Oksanen A, Gjerde B, Richter SH, Örtenberg E, et al. 98. Luna-Olivares LA, Kyvsgaard NC, Ferdushy T, Nejsum P, Thamsborg SM, Intestinal parasites in swine in the Nordic countries: prevalence and Roepstorff A, et al. The jejunal cellular responses in chickens infected with a geographical distribution. Vet Parasitol. 1998;76:305–19. single dose of Ascaridia galli eggs. Parasitol Res. 2015;114:2507–15. 76. Roepstorff A, Bjørn H, Nansen P, Barnes EH, Christensen CM. Experimental Oesophagostomum dentatum infections in the pig: worm populations resulting from trickle infections with three dose levels of larvae. Int J Parasitol. 1996;26:399–408. 77. Permin A, Bojesen M, Nansen P, Bisgaard M, Frandsen F, Pearman M. Ascaridia galli populations in chickens following single infections with different dose levels. Parasitol Res. 1997;83:614–7. 78. Daş G, Gauly M. Density related effects on lifetime fecundity of Heterakis gallinarum in chickens. Parasit Vectors. 2014;7:334. 79. Keymer A. Density-dependent mechanisms in the regulation of intestinal helminth populations. Parasitology. 1982;84:573–87. 80. Wang B-J, Gu X-B, Yang G-Y, Wang T, Lai W-M, Zhong Z-J, et al. Mitochondrial genomes of Heterakis gallinae and Heterakis beramporia support that they belong to the infraorder Ascaridomorpha. Infect Genet Evol. 2016;40:228–35. 81. Daş G, Hennies M, Sohnrey B, Rahimian S, Wongrak K, Stehr M, et al. A comprehensive evaluation of an ELISA for the diagnosis of the two most common ascarids in chickens using plasma or egg yolks. Parasit Vectors. 2017;10:187. 82. Lind P, Eriksen L, Nansen P, Nilsson O, Roepstorff A. Response to repeated inoculations with Ascaris suum eggs in pigs during the fattening period. II. Specific IgA, IgG, and IgM antibodies determined by enzyme-linked immunosorbent assay. Parasitol Res. 1993;79:240–4. 83. Roepstorff A, Eriksen L, Slotved HC, Nansen P. Experimental Ascaris suum infection in the pig: worm population kinetics following single inoculations with three doses of infective eggs. Parasitology. 1997;115:443–52. 84. Nejsum P, Thamsborg SM, Petersen HH, Kringel H, Fredholm M, Roepstorff A. Population dynamics of Trichuris suis in trickle-infected pigs. Parasitology. 2009;136:691–7. 85. Ferdushy T, Schou TW, Norup LR, Dalgaard TS, Thamsborf SM, Nejsum P, et al. Acquisition of resistance after continuous infection with Ascaridia galli in chickens. Parasitology. 2014;141:1603–10. 86. Degen WGJ, Van Daal N, Rothwell L, Kaiser P, Schijns VEJC. Th1/Th2 polarization by viral and helminth infection in birds. Vet Microbiol. 2005;105:163–7.

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Parasites & VectorsSpringer Journals

Published: May 29, 2018

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