Introduction

Parasite infection is a common problem in organic pig production (Katakam et al. 2016; Lindgren et al. 2020; Li et al. 2022). In a study investigating parasite infection in organic and alternative swine farms across five states in the USA, Hernandez et al. (2022) reported that 89%, 56%, and 44% of the participating farms were infected with Ascaris suum, Oesophagostomum spp., and Trichuris suis, respectively. Pastures contaminated by infected pigs are sources of parasite transmission because infective eggs of common swine parasites, such as A. suum and T. suis, can survive in soil for 5 to 11 years (Nansen and Roepstorff 1999; Carstensen et al. 2002; Roepstorff et al. 2011). Controlling parasite contamination on organic pastures has been a challenge for farmers because synthetic nematicides are not allowed for organic agriculture production (NOP 2020). Common strategies for controlling parasite contamination in pastures include reducing the number of pigs in the pasture and increasing intervals between grazing rotations (Nansen and Roepstorff 1999; Roepstorff et al. 2001; Lindgren et al. 2014). However, these strategies require more land, which is a limiting factor for organic farmers.

Reducing parasite eggs in soil using natural products could be another method for controlling pasture contamination. Biological fumigation or biofumigation is an alternative to the use of synthetic nematicides in the field to reduce parasitic nematodes (Alabouvette et al. 2006; Ploeg 2008). Biofumigation is the process of incorporating residues of plants in the Brassicaceae family into the soil to suppress soil-borne parasites and plant pathogens because plants in this family contain glucosinolates. When these plants are pulverized, glucosinolates are broken down by an endogenous enzyme (myrosinase or thiogucosidase) to release gaseous isothiocyanates (Cole 1976). Isothiocyanates have biocidal properties, including anthelmintic, and are an active ingredient of some synthetic nematicides (Kirkegaard and Sarwar 1998; Ploeg 2008). While biofumigation has been used to control plant-parasitic nematodes (Alabouvette et al. 2006), it has never been tested as a method to suppress swine parasite eggs in pasture soil. This study aimed to assess if biofumigation with rapeseed can reduce the number of parasite eggs in organic pasture soil.

Materials and methods

The protocol for this study was reviewed and approved by the University of Minnesota Institutional Animal Care and Use Committee (IACUC# 2106-39200A).

This study was conducted at Rodale Institute located in Kutztown, Pennsylvania, USA. The institute operated a hog facility that was surrounded by 2.8 hectares of organic pastures. The hog facility was a hoop structure (12 m × 30 m) with 11 pens bedded with straw on concrete floors. Each pen (4.5 m × 4.5 m) housed eight to nine pigs and was equipped with a single-space dry feeder and four freeze-proof nipple drinkers, each at a different height. All pigs were managed organically according to standards set up by the National Organic Program (NOP 2020) and provided free access to pastures. Parasite contaminations were routinely monitored and had been detected in pigs, bedding, and pasture soil prior to the study (Hernandez et al. 2023). Pastures were managed through crop and grazing rotations. Generally, a perennial legume mix (e.g., alfalfa and clover) was the predominant pasture crop.

Experimental design

Pastures were subjected to four treatments: rapeseed pasture with incorporating plant residues into the soil (biofumigation), rapeseed pasture without incorporating plant residues into the soil, clover pasture with incorporating plant residues into the soil, and clover pasture without incorporating plant residues into the soil. The field study was conducted in two replicates in 2021, from May to August for replicate one and August to November for replicate two. Within each replicate, two side-by-side plots (20 m × 40 m/plot) of pastures that were adjacent to the hoop barn were used. Prior to the study, both plots were managed in similar ways in terms of pasture crop and grazing rotation. One plot was planted to rapeseed, and another was planted to Ladino clover. Rapeseed was chosen as a treatment due to its potential biofumigation properties associated with high concentrations of glucosinolates (Johnson 2009; Visnjevec et al. 2021). Ladino clover was used as control due to its low content of glucosinolates and easy establishment observed at Rodale Institute.

Within each replicate, each plot of clover and rapeseed pasture was divided into two sub-plots (plot 1 and plot 2; 10 m × 40 m/sub-plot), with one for biofumigation treatment by mechanically incorporating plant residues into the soil after pigs were removed, and another for control without mechanically incorporating plant residues after pigs were removed (Fig. 1). Each sub-plot was further split into four paddocks (10 m × 10 m/paddock) that were enclosed using electrified polywire to allow pigs grazing within the perimeters. Eight to nine pigs grazed each paddock for a week (see grazing test pastures below).

Fig. 1
figure 1

Experimental design for four pasture treatments: 1. rapeseed plot (shaded yellow): grazed and mechanically incorporated (biofumigation); 2. rapeseed plot (yellow): grazed only (no biofumigation); 3. clover plot (shaded green): grazed and mechanically incorporated (biofumigation); 4. clover plot (green): grazed only (no biofumigation). Paddocks 1–4 (weeks 1–4): each plot was split into four paddocks, and each paddock was grazed by eight to nine pigs for 1 week sequentially

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Pasture establishment for the study

Before planting, pasture soil was moldboard plowed, disced, and culti-packed. Then pastures were sown with a grain drill at the rate of 2 g/m2 for both rapeseed and Ladino clover. Ladino clover pastures for both replicates one and two were planted in March 2021. Rapeseed plots for replicates one and two were planted in March and July 2021, respectively. All pigs were given access to pastures when rapeseed was at about 50% blooming, 10 to 13 weeks after planting in both replicates.

Grazing test pastures

A total of 66 pigs were used for grazing test pastures in two replicates. Pigs were either purchased from a local farmer (n = 58; Landrace × Duroc or Yorkshire × Duroc) at approximately 4 to 6 weeks of age or born at Rodale Institute (n = 8; Large Black × Tamworth). Pigs were housed in the hoop barn with access to non-study pastures for 16 weeks and 10 weeks in replicates one and two, respectively, until 50% of rapeseed blooming. Then, pigs were given access to the test pastures. In the first replicate, pigs (n = 32, body weight = 95.9 ± 13.7 kg) were 21 weeks of age, and in the second replicate, pigs (n = 34, body weight = 57.5 ± 5.6 kg) were 15 weeks of age when given access to the test pastures.

Eight to nine pigs grazed each paddock for a week, starting from the paddock closest to the hoop barn. Then, pigs were removed from the test pastures. For replicate one, pigs were on pastures between June 30 and July 28, 2021 (Table 1), and between September 21 and October 19, 2021, for replicate two. Throughout grazing periods, all pigs had free access to feed and water in their designated pens in the hoop barn.

Table 1 Soil sampling timeline for determining changes in parasite eggs in pasture soil (replicate one)
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Pasture soil biofumigation

Biofumigation of pasture soil was conducted immediately after pigs were removed from each paddock. The biofumigated paddocks were mowed using a flail mower (John Deere 390, Allentown, PA. USA), then crop residues were immediately incorporated into the soil using a mechanical spader (Celli, Langley, BC. Canada), and then culti-packed. The effectiveness of biofumigation was evaluated by changes in egg counts of A. suum and T. suis in pasture soil for each paddock over the 3 weeks that followed.

Data collection

Fecal sampling

To evaluate parasite infection in pigs before grazing, fecal samples (n = 66) were collected from each pig the day before given access to pastures for analysis of A. suum, T. suis, and O. spp. egg counts. Samples (5 to 10 g/sample) were collected either before the feces was dropped on the ground or from the top of feces recently dropped (still warm) on the ground as described previously (Li et al. 2022). Each sample was placed in a labeled plastic zip-lock bag and stored on ice in a cooler immediately after sampling. Samples were stored at 4 °C in a refrigerator for less than 24 h before fecal egg counts (FEC) for each parasite species were made.

Biomass of pasture crops

To evaluate yield and the availability of pasture crops for biofumigation, above ground plant biomass before and after grazing was evaluated for each paddock of all sub-plots. Within each paddock, biomass was collected at three randomly selected 0.56 m2 (0.75 m × 0.75 m) quadrants before pigs were given access to the paddock, and again immediately after pigs were removed from the paddock. All plants within quadrants, including the pasture crop (rapeseed or clover) and weeds, were collected at 1 cm above the ground and considered one sample. For samples collected from rapeseed sub-plots, rapeseed was separated from weeds to determine the availability of rapeseed for biofumigation. Each sample was weighed as is for wet weight, then dried at 100–120 °C for 1 week and weighed to determine dry matter content.

Soil sampling

To evaluate the effectiveness of biofumigation on reducing parasite contamination in pasture soil, soil samples were collected for evaluation of parasite egg counts. Soil samples (n = 480) were collected from each paddock five times: before grazing, immediately after pigs were removed from pastures and weekly thereafter for 3 weeks. Sampling timeline for replicate one is demonstrated in Table 1. Twenty subsamples were collected from a “W” transect as described by Larsen and Roepstorff (1999) and Katakam et al. (2016). Subsamples consisted of soil cores to a depth of 5 cm that were collected using a tubular soil sampler (Forestry Suppliers, Inc., Jackson, MS, USA). All 20 subsamples were thoroughly mixed to yield a soil sample. Three soil samples were collected in different areas of each paddock. Each soil sample was placed in a plastic zip-lock bag and stored on ice in a cooler after sampling. All samples were stored in a refrigerator at 4 °C before analysis for egg counts of A. suum and T. suis.

Lab analysis for swine parasite egg counts

All parasite egg counts were analyzed morphologically. Fecal samples were analyzed for FEC of three parasite species: A. suum, T. suis, and O. spp. using the concentration McMaster method described by Roepstorff and Nansen (1998). Saturated NaCl-glucose solution (50 g NaCl, 75 g glucose monohydrate, and 131 g water) was used as a flotation fluid, and the detection limit was 20 egg per gram (epg) of fecal sample for all three species of parasites. Samples that were detected with parasite eggs were identified as positive samples. Adjustment for false-positive infection of A. suum was used in this study to exclude pigs that were not infected by only passed A. suum eggs through their digestive tracts. Boes et al. (1997) demonstrated that pigs with FEC < 200 epg were not likely harbor A. suum worms. So, fecal samples with A. suum FEC < 200 epg were considered false-positive in this study. Positive fecal samples were presented as percent of total fecal samples analyzed for each parasite species.

Ascaris suum and T. suis eggs in soil samples were quantified following the concentration McMaster method described by Roepstorff et al. (2001). The detection limit for A. suum and T. suis eggs was 1 egg per 10 g of soil sample. Eggs of O. spp. in soil samples were not counted because the eggs hatch larvae shortly after being shed in feces and are not likely to be recovered from soil (Anderson 2000). Soil samples that were detected with parasite eggs were identified as positive samples that were presented as percent of total soil samples analyzed for each parasite species.

Statistical analysis

All data were analyzed using SAS software (Version 9.4; SAS Institute Inc, Cary, NC). Descriptive data (median, maximum, and minimum) of FEC and soil egg counts were summarized using a univariate procedure. Percent of positive fecal and soil samples was analyzed using a frequency procedure with chi-square tests. The univariate procedure with normality tests (Normaltest) was used to evaluate normal distribution of data of FEC, biomass, and soil eggs. All these data were not normally distributed, so a one-way nonparametric procedure with Kruskal–Wallis rank score tests with replicate serving as a stratifying variable was used for analysis of the data. Consequently, the effect of pasture treatment on FEC, biomass, and soil eggs was tested using one-way nonparametric analysis with Kruskal–Wallis tests. Additionally, positive soil samples and soil egg counts were compared among sampling points (before, immediately after, and 1 week, 2 weeks, and 3 weeks after pigs were removed) using chi-square and Kruskal–Wallis tests, respectively. All tests were two-tailed tests. Differences were considered significant when P for chi-square was smaller than 0.05, and a tendency when 0.05 < P ≤ 0.10.

Results

On average, 35 to 56% of fecal samples were positive for A. suum from pigs before grazing; 31 to 53% positive for T. suis; and 6 and 19% positive for O. spp. (Table 2). There was no difference in percent of positive samples or FEC of any parasite species among pigs before grazing different test pastures (all chi-square ≤ 4.3, df = 3; all P ≥ 0.23).

Table 2 Positive fecal samples and fecal egg counts of pigs before grazing different pastures
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Dry weight of plant biomass differed among pastures with different treatments both pre-grazing (P = 0.05; Table 3) and post-grazing (P = 0.03). Likewise, dry matter content of plant biomass tended to differ pre-grazing (P = 0.06) and differed post-grazing (P < 0.001) among pasture treatments. On rapeseed pastures, 24 to 27% of total dry biomass pre-grazing and 21 to 24% of total dry biomass post-grazing was rapeseed, suggesting the majority of plant biomass was non-rapeseed in rapeseed plots.

Table 3 Above ground biomass of pastures before and after grazed by pigs
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Ascaris suum eggs were detected in 51 to 63% of soil samples and there was no difference in percent of positive soil samples among pasture treatments (Table 4). Compared to soil samples contaminated with A. suum eggs, fewer soil samples were contaminated with T. suis eggs, accounting for 17 to 20% of total soil samples. There were no differences in percent of T. suis positive samples across pasture treatments.

Table 4 Testing the effect of pasture treatment on parasite egg counts in soil samples
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Percent of A. suum positive soil samples was different among sampling timepoints (chi-square = 12.1, df = 4; P = 0.02; Table 5). Compared to soil samples collected at 2 and 3 weeks after pigs were removed, soil samples collected before grazing and immediately after pigs were removed had higher percent of positive samples for A. suum (e.g., 66% positive samples immediately after pigs were removed vs. 43% at 2 weeks after pigs were removed). Additionally, mean soil eggs of A. suum differed across sampling timepoints (Kruskal–Wallis chi-square = 28.2, df = 4; P < 0.001), with more parasite eggs in soil samples collected before grazing, and immediately after and 1 week after pigs were removed from paddocks, compared to samples collected 2 and 3 weeks after pigs were removed. Sampling timepoints also affected T. suis positive soil samples (chi-square = 39.8, df = 4; P < 0.001), with more soil samples being positive for T. suis before grazing (34%) and immediately after pigs were removed (35%), compared to 2 (7%) and 3 weeks (11%) after pigs were removed from pastures. Mean soil T. suis eggs were different across sampling timepoints (Kruskal–Wallis chi-square = P < 0.001), with more eggs in soil samples collected before grazing, and immediately after and 1 week after pigs were removed, compared to 2 and 3 weeks after pigs were removed from pastures.

Table 5 Testing the time effect on parasite egg counts in pasture soil
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Discussion

This study did not confirm the effectiveness of biofumigation on reducing parasite contamination in pasture soil. This could be attributed to several factors, including pasture crop selection, the method and timing of incorporating plant residues into the soil, and soil conditions (Ploeg 2008). In the current study, biomass yield of rapeseed was so low that rapeseed did not dominate the rapeseed pastures, as indicated by the fact that only 24 to 27% of dry biomass on the pastures was rapeseed, and the rest was volunteer plants. Additionally, the effectiveness of biofumigation depends on volatile substances, isothiocyanates, produced during the breakdown of Brassica tissues (Cole 1976; Kirkegaard and Sarwar 1998; Ploeg 2008). Because isothiocyanates are products of enzymatic hydrolysis of glucosinolates, contents of glucosinolates in Brassica plants are important to isothiocyanate production. While the content of gluconisolates varies greatly among Brassica species, it also changes with growth period of the plant. In the current study, we chose to incorporate plant residues when rapeseed flowered and the content of glucosinolates are supposed to be high (Kondra and Downey 1970). Some studies (Angus et al. 1994; Morra and Kirkegaard 2002), however, demonstrated that Brassica species with high content of glucosinolates were not effective biofumigants due to low production of isothiocyanates. It was revealed later that methods to destruct plant tissues are important to the effectiveness of biofumigation (Morra and Kirkegaard 2002). When the plants were cut or chopped, as the method often used under field conditions and in the current study, isothiocyanates produced in the soil was only 5% of isothiocyanates produced when the plants were pulverized (Gardner et al. 1999; Morra and Kirkegaard 2002). Additionally, watering or covering the soil using plastic sheets immediately after incorporating plant residues to prevent isothiocyanate gasses from releasing to the air can enhance the effectiveness of biofumigation (Bello et al. 2004). Furthermore, soil temperature higher than 20 °C can increase the effectiveness of biofumigation (Ploeg and Stapleton 2001; Bello et al. 2004). Based on previous studies and our experiences from the current study, we speculate the major reason for the ineffectivenss of biofumigation in the current study was the method that plant residues were incorporated into the soil. Additionally, while covering the field with plastic sheet is not feasible at the farm level due to large area of patures, the soil could have been watered immediately after incorporating plant residues to seal isothiocyanates produced in the soil and consequently improve the effectiveness of biofumigation. To our knowledge, the current study is the first to explore biofumigation as a method to suppress swine parasite eggs in pasture soil. Future research is needed to identify the optimal conditions under which biofumigation may effectively suppress the number of swine parasite eggs on pasture. Additionally, the effectiveness of biofumigation could be tested by changes in the hatching ability of parasite eggs on pasture which was not examined in the current study. Further research should investigate the effect of biofumigation on hatching ability of swine parasite eggs on pasture.

Percent of positive soil samples and egg counts of each parasite species in soil samples in the current study were comparable to the results reported by Lindgren et al. (2020). To increase the possibility of detecting the differences among pasture treatments, we doubled the stocking density of pigs in the current study (8 to 9 pigs/100 m2) from our normal stocking density (4 to 5 pigs/100 m2) on each paddock. As a result, A. suum eggs were detected in 51 to 63% and T. suis eggs in 17 to 27% of soil samples across pasture treatments. While pasture treatments did not affect percent of positive samples or parasite egg counts in soil, time after pigs were removed from paddocks did affect positive soil samples and soil egg counts. Positive samples (%) and egg counts of both A. suum and T. suis in pasture soil reduced 2 weeks after pigs were removed from paddocks. This is not surprising as Roepstorff et al. (2011) reported that the large majority of A. suum and T. suis eggs died within the first few months after being deposited on pastures. Our results demonstrated that the first few weeks after pigs were removed from pastures can make a difference in A. suum and T. suis egg contamination on pasturess. While the egg counts of both A. suum and T. suis dropped signicantly 2 weeks after pigs were removed from pastures, we still detected that more than 43% and 7% of soil samples were contaminated with A. suum and T. suis, respectively, at the inclusion of the study. This demonstrates the long-term impact of parasite contamination on organic pastures. Future research should focus on identifying effective and feasible strategies for controlling parasite contamination on commercial organic pastures.

Conclusions

Our results suggest that biofumigation was not effective in reducing swine parasite eggs in organic pasture soil under conditions of the current study. Additionally, we observed that eggs of both A. suum and T. suis in pasture soil were reduced 2 weeks after pigs were removed, regardless of biofumigation.