Progress report for ONE22-421
Project Information
The ultimate goal of this project is to develop management practices and technical guidelines for farmers for effective control of swine parasites in organic systems. The specific objectives are:
- Evaluate parasite prevalence and intensity on organic and pasture-raised pig farms by conducting a robust survey study (including 10 organic farms in the Northeast region primarily in PA and NY)
- Analyze the effects of management practices (i.e. bedding and housing used, access to pasture, acreage used for pigs, size of the herd, breeds used, etc.) on parasite infections
- Disseminate information to farmers and educate them on the best management practices for control and management of swine parasites on organic farms
Pork is the third-highest selling meat product in the United States, with the average American consuming 51 pounds of pork annually (Davis and Lin 2005). To compensate for this demand, hogs are raised in specialized, indoor confinement facilities that maximize efficiency. These facilities, however, often have poor indoor environments that restrict innate behaviors and are associated with symptoms of chronic stress in pigs (Stolba and Woodgush 1981). There is growing evidence that animal welfare is improved when pigs are raised outdoors (Edwards 2003; Miao et al. 2004; Velazco et al 2013; Weary et al. 2016). Organically raised pigs are required to have outdoor access as stated by the USDA National Organic Program (USDA organic regulations Title 7, Part 205, §205.239).
Allowing outdoor access, however, can bring an additional set of challenges due to greater exposure to the infective stages of parasites in the environment. Conventional farms deploy regular anthelmintic prophylaxis and house animals on slatted floors to avoid parasite infection (Nansen and Roepstorff 1999), but these practices are not allowed in organic production (USDA organic regulations Title 7, Part 205, §205.239). Organic farmers must consider the control of swine parasites without prophylactic treatments, which is challenging considering that semi-free-ranged pigs can be infected with a higher diversity of parasite species at higher abundances relative to those in non-organic, indoor industrialized production operations (Nansen and Roepstorff 1999; Roepstorff at al. 2011).
Surveys of European organic farms have shown that pigs are infected with more types of parasites at a higher prevalence and intensity of infection (Roepstorff et al. 2011). Bedded floors and access to outdoors or pastures that are required by the NOP are the major sources of parasite contamination on organic pig farms. Compared to pigs housed on slatted floors, organic pigs harbor more species of parasites with heavier concentrations (Li 2018). For certain parasite species, prevalence reduced from 95% in pigs housed on bedded floors to 3% in pigs housed on fully slatted floors (Morris et al. 1984).
Gastrointestinal parasites in swine are a major challenge in organic production systems because they can cause economic losses for farmers. The most common species of parasites in pigs are Ascaris suum (the large intestinal roundworm) and Trichuris suis (the swine whipworm) in growing pigs, and Oesophagostomum spp. (the nodular worm) in breeding sows (Edwards et al. 2014; Lingren et al. 2014). Thousands to millions of eggs are released by these parasites every day and passed in the feces of infected individuals, and infective eggs and larvae can remain viable in the soil for months or years (Roepstorff and Nansen 1998). Infection with gastrointestinal parasites can cause significantly reduced average daily gain and poor feed efficiency, which results in economic losses because more feed is needed to get infected pigs to finishing weight than uninfected pigs (Kipper et al 2011). The growth potential of feeder pigs can be permanently affected by exposure to parasites (Urban et al. 1989). Knecht et al. (2011) found a significant difference between the meatiness of slaughtered pigs that were infected with parasites versus pigs that were not infected. Additional economic loss comes from liver condemnations at slaughter as the result of granulomas from migrating larvae of Ascaris suum (Roepstorff et al. 2011). Overall, the need for additional feed to get the growing stock to finishing weight in a time of rising organic feed costs causes noteworthy economic damage to organic farmers.
Organic pig farmers have a greater need for effective worm control than their conventional counterparts due to the higher incidence of parasite infections, but little scientific data is available on the effectiveness of organically approved anthelmintics. Organic pork producers often employ natural or herbal deworming treatments like garlic, pumpkin seeds, or black walnut husks (Lans et al. 2007). There have been very few scientific studies on the efficacy of these natural methods, especially in pigs, and studies done on other species of livestock often show little to no success. For example, in vivo studies showed that garlic had no significant effect on the fecal egg counts of infected horses or in goats and lambs (Buono et al. 2019; Burke et al. 2009a), two commercially available herbal dewormers were ineffective in goats (Burke et al. 2009b), and diatomaceous earth did not significantly reduce fecal egg counts in grazing ruminants (Fernandez et al. 1998). Effective natural treatments against swine parasites may exist, but more in-depth research is needed to establish a better understanding of organically approved worm management.
With little evidence supporting organically approved deworming methods, farmers need to focus on improving management practices as preventative measures to manage parasites. Pastures are commonly allowed to rest and recover after having pigs, but resting pastures may not be completely effective at controlling parasites because infective eggs can remain viable for over a decade (Roepstorff and Nansen 1998). Proposed natural methods of parasite control via pasture management include planting biofumigants in pastures and the use of predatory or parasitic fungi (Miao 2004). Other farm variables like herd size, stocking density, and whether the operation is a farrowing farm or an all-in/all-out may influence parasite infection.
We plan to address the lack of understanding about parasite prevalence in the northeastern United States by surveying organic and pastured pork production facilities. Prior to 2019, no comprehensive parasite survey of organically managed pig farms existed, and the status of parasite infection on Pennsylvania organic pig farms is widely unknown. In addition to measuring the level of parasite infection on organic farms, we will collect data from each farm on management practices like herd size, stocking density, housing style, and rotation schedule. The parasite infection data will be analyzed with respect to management practices to evaluate the effects of these practices on parasite infection. The results of these analyses will be disseminated to farmers to educate and enable them to make better-informed herd management decisions.
Cooperators
- - Producer
- (Educator and Researcher)
- - Producer
Research
Ten farms will participate in this project. Data on farm demographics and management will be collected via an in-person written attached survey. Data include housing type, herd size, breeds used, type of outdoor access (concrete pad, dirt lot, pasture, etc.), bedding, organic anthelmintics used, and acreage of dirt lot or pasture if applicable.
Participating farms will be chosen based on the following criteria: 1) farms must be certified organic or follow organic management protocols; 2) pigs must have outdoor access and freshly bedded floors. Specifically, farms cannot employ chemical prophylaxis and pigs cannot be housed on slatted floors or in confinement conditions.
Participating farms will be visited seasonally in March, June, September, and December for sampling including:
- Fecal samples (n = 20-40 per farm) will be collected randomly (Katakam et al., 2016) and labeled. Samples will be collected fresh and directly from the rectum.
- Soil samples will be collected by walking a ‘W’ route through pastures that have been or are occupied by pigs (Roepstorff et al., 2001). Three replicates will be collected per area, each consisting of 20 subsamples (approximately 5 g of soil from 0-5 cm depth every 3 m). Subsamples will be combined and thoroughly homogenized by hand.
- Bedding materials and manure bedpack will be collected based on the methods described by Katakam et al. (2016). Bedded areas will be categorized into clean, intermediate, and dirty areas depending on urine and fecal contamination. The clean area will appear dry and minimally contaminated by feces and urine, while the dirty area will be wet and heavily contaminated with feces and urine. The intermediate area will be between the clean and dirty areas. The top 10 cm of bedding material or bedpack manure will be collected along the route of 3 ‘W’ walks. Subsamples will be combined and thoroughly homogenized by hand.
All samples will be analyzed to identify parasite species and quantify egg counts for each species. The number of eggs per gram (EPG) of samples will be used as a relative measure of infection size. Samples will be processed to isolate swine parasite eggs using a concentration McMaster technique (Roepstorff and Nansen, 1998), with a sensitivity of 20 EPG. The flotation fluid used to quantify parasite eggs will be a solution of saturated NaCl and glucose (50 g NaCl, 75 g glucose monohydrate, and 131 g water), which has a specific gravity of 1.27 g/mL. This solution is suitable to recover swine intestinal parasite eggs and to assess the relative abundance of coccidia oocysts as few, medium, or many (Carstensen et al. 2002). Lab activities include:
- Fecal samples: 4 g of feces will be soaked in 56 mL of tap water for 30 minutes then filtered through a single layer of cheesecloth. 10 mL of filtrate will be transferred to a 15 mL Falcon tube and centrifuged at 2000 rpm for 7 minutes. After drawing off and discarding the supernatant, the pellet will be resuspended in ~4 mL of flotation solution. Both chambers of a McMaster slide will be filled and the slide will rest, undisturbed, on the bench top for 5 minutes to allow eggs to float. All eggs under the engraved grids will be counted. This egg count will be multiplied by 20 to get a calculated eggs per gram (epg) of feces.
- Soil samples: 10 g of homogenized soil will be weighed into a 50 mL Falcon tube and the tube will be filled to the 50 mL mark with 0.5 M NaOH. The soil will soak for 24 hours. After 24 hours, the tubes will be centrifuged at 2000 rpm for 7 minutes. The supernatant will be discarded, and the tubes will be filled to the 50 mL mark with flotation solution and the pellets resuspended via vortexing. The tubes will be centrifuged at 2000 rpm for 7 minutes. The supernatants will be collected in pre-labeled glass beakers. The resuspension of pellets in flotation solution and centrifugation will be repeated three more times. The collected supernatant in each beaker will then be washed with tap water on a 20 µm sieve, and the residue remaining on the sieve will be transferred to a 15 mL Falcon tube and centrifuged at 2000 rpm for 7 minutes. All fluid except the bottom 0.5-1 mL will be discarded. ~2 mL flotation solution will be added, and both chambers of a McMaster slide will be filled. The slide will be allowed to rest for five minutes on the bench top, and then all eggs in the slide will be counted (inside and outside the engraved grid).
- Bedding samples: Bedding material will all be cut to pieces 1-5 cm in length and homogenized. 5 g of bedding material will be soaked in 0.5 M NaOH for 16-18 hours. The sample will then be thoroughly washed with tap water on a 212 µm sieve on top of a 20 µm sieve. 10 mL of material retained on the 20 µm sieve will be transferred to a 50 mL Falcon tube, then the tube will be filled to the 50 mL mark with flotation solution and centrifuged at 2000 rpm for 7 minutes. The supernatant will be removed, and the pellet will be washed on a 20 µm sieve. Material retained on the sieve will be transferred to a 15 mL Falcon tube and centrifuged at 2000 rpm for 7 minutes. The supernatant will be discarded and the pellet resuspended in ~2 mL flotation solution. Both chambers of a McMaster slide will be filled and the slide will be allowed to rest for 5 minutes on the benchtop. All eggs in the slide will be counted.
Data will be analyzed using R statistical software (R Core Team, 2017). To examine variability in the prevalence of infection in samples, each individual sample processed will be classified as either uninfected or infected with at least 1 parasite species, or with each of the 3 nematode parasite species separately. Data will be treated as a series of Bernoulli trials with a binary response variable, which will allow us to examine the likelihood of observing infections between different independent variables (e.g. farms, seasons, ages, sex, breed, management, or area of pen sampled), and their interaction. We will use Linear Mixed Effect Models (Pinheiro & Bates, 2000) to examine the variability in the intensity and abundance of EPG for each species recovered from feces, soil, and bedding samples. The number of eggs hatched will be used as the response variable and examined in relation to the following fixed explanatory variables when analyzing fecal EPG: farm ID, season (fall, winter, spring summer), management practice, age, and sex of pigs. The mean intensity and abundance of EPG from soil samples will be used as the response variable, with farm ID, season, and management practice as the fixed explanatory variables. For the mean intensity and abundance of EPG from pig bedding samples, farm ID, area of pen sampled (clean, intermediate, or dirty), age and sex of pigs in each pen will be used as the fixed explanatory variables. A hierarchy of models will be examined based on diverse combinations of independent variables. The variability between the breeds used by different farms will be included as a random effect.
Nine Pennsylvania farms raising pigs with outdoor access and without the use of synthetic anthelmintics were identified to collaborate with us in this project. The first farm visit and sample collection (fecal, soil, and bedding sampling) occurred between September-November 2022. The first of four surveying events were performed during this visit. Farmers from eight of nine farms filled out surveys with farm demographic and hog management information, including location, type of hog housing and outdoor access, farm production information (i.e., number of hogs raised per year, length of time raising pigs), non-synthetic anthelmintic remedies used, and manure and pasture management. Of eight farms surveyed, seven (87.5%) bought in pigs and followed a feeder-to-finish production system, and one (12.5%) housed breeding stock and followed a farrow-to-finish system. Farms had been raising hogs for 3-10 years (average 7.25 +/- 1 year) and raised between 10-60 hogs annually (average 29 +/- 5 hogs). Seven farms had hogs on-site at the time of sampling. Of those seven farms, three had hogs on pasture, three had hogs in the forest, and one had hogs with access to an outdoor concrete pad. Four of eight farms (50%) used a natural anthelmintic remedy; two farms used diatomaceous earth, one used apple cider vinegar, and one used a combination of both.
Table 1. Participating farm demographic and management data. Survey data was not collected for farm # 8 at the time of sampling.
| Farm ID | Farm Location | Time Raising Pigs on Farm (Years) | # of Pigs Raised Annually | Outdoor Access Type at Time of Sampling | Shelter Type | Non-Synthetic Anthelmintics Used? | Biosecurity Measures Implemented? |
| 1 | Fleetwood, PA | 5 | 10-14 | Pasture | Mobile | Y | N |
| 2 | Nescopeck, PA | 3 | 40 | Forest | Mobile | Y | Y |
| 3 | Barto, PA | 9 | 40 | Forest | Forest Cover | N | N |
| 4 | Oley, PA | 10 | 20 | Concrete Pad | Permanent | Y | N |
| 5 | Kingsley, PA | 3 | 15-20 | Forest | Forest Cover | N | N |
| 6 | Newmanstown, PA | 10 | 25 | Pasture | Mobile | Y | N |
| 7 | Kutztown, PA | 10 | 40-60 | Pasture | Permanent | N | Y |
| 8 | Lenhartsville, PA | ||||||
| 9 | Lancaster, PA | 8 | 20-30 | Pasture | Mobile | N | Y |
A total of 90 fecal samples were collected from eight participating farms (11 +/- 2 fecal samples collected per farm. One farm (farm #9) did not have hogs on site at the time of sampling. Fresh fecal samples were collected from the ground immediately after defecation and placed into a Ziplock bag, labeled with the age, sex, and ear tag number (when applicable) of the hog. Samples were immediately placed in a cooler on ice until transferred to the laboratory, where they were stored at approximately 3°C until processing. Samples were processed following a concentration McMaster technique for parasite examination. 4 g of feces was soaked in 56 mL of water for 30 minutes then filtered through a single layer of cheesecloth. 10 mL of filtrate was transferred to a 15 mL Falcon tube and centrifuged at 2000 rpm for 7 minutes. The supernatant was carefully poured off and the pellet was resuspended in 4 mL of flotation solution (50 g NaCl, 75 g glucose monohydrate, 131 mL H2O). Both chambers of a McMaster slide were filled with this solution and the slides were set to rest on the benchtop for 5 minutes to allow for the flotation of parasite eggs. All eggs within the grid of each chamber were counted under a microscope at 40x magnification. The combined number of parasite eggs was multiplied by 20 to calculate the number of eggs per gram of feces for each parasite species.
Twenty-four fecal samples were collected from feeder pigs (weaning-4 months), 60 were collected from finisher pigs (5 months-slaughter), and 4 were collected from breeding sows. 40 samples were collected from female pigs and 48 were collected from male pigs. Sex and age group were not recorded for 2 samples. Seven of eight sampled farms had hogs that were positive for parasite infection (87.5%). 69 hogs were positive for infection with at least one parasite species (76.7%); of the 69 infected hogs, 39 (56.5%) were infected with one species, 22 (31.9%) were infected with two species, and 8 (11.6%) were infected with three species of parasites. The parasite species identified were the large intestinal roundworm Ascaris suum, the swine whipworm Trichuris suis, and strongyles, likely the nodular worm Oesophagostomum spp. Of the 69 infected hogs, 34 (49%) were infected with strongyles, 60 (87%) were infected with A. suum, and 13 (19%) were infected with T. suis.
Soil samples were collected from eight farms; one farm (farm #4) had pigs on a concrete pad and therefore did not have soil to sample. Soil samples were collected from areas occupied by pigs on seven farms, as well as areas recently occupied by pigs on four farms. In each pasture or paddock, twenty 5 cm deep soil cores were collected while walking a ‘W’ route. Three replicates were collected for each area. Soil samples were immediately placed in a cooler on ice for transport to the laboratory, where they were stored at approximately 3°C until processing. To process, the samples were homogenized by hand and 10 g subsamples were transferred to 50 mL Falcon tubes. The tubes were filled to the 50 mL mark with 0.5 M NaOH and soaked for 24 hours. The samples were centrifuged at 2000 rpm for 7 minutes, and the supernatant was poured off. The tubes were then filled to the 50 mL mark with flotation solution and the pellets resuspended. The samples were centrifuged at 2000 rpm for 7 minutes, and the supernatant collected in labeled glass jars. This resuspension, centrifugation, and supernatant collection was repeated three more times. The collected supernatant was then washed on a 20 µm sieve with tap water, and the material collected on the sieve was transferred to a 15 mL Falcon tube and centrifuged at 2000 rpm for 7 minutes. All supernatant except the bottom 0.5-1 mL was drawn off, and approximately 2 mL of flotation solution was added to the sample. Both chambers of a McMaster slide were filled and set to rest on the benchtop for 5 minutes to allow the flotation of parasite eggs. All eggs in both chambers, including those inside and outside the grids, were counted under a microscope at 40x magnification.
Sixty-nine soil samples were collected from eight farms. Parasite eggs were identified in occupied paddocks on 4 farms (farms # 2, 3, 5, and 7), as well as previous paddocks on 3 farms (farms # 1, 7, and 9). Strongyle and Ascaris suum eggs were identified in 10% and 48% of soil samples, respectively.
Bedding samples were collected from 4 farms. Bedding samples were collected by walking a ‘W’ transect through the bedded area and collecting 5 handfuls of bedding material. When applicable, the bedded areas were separated into dirty, intermediate, and clean areas. Dirty areas were heavily contaminated with urine and feces, clean areas consisted of clean, dry bedding with minimal fecal contamination, and intermediate areas were in between the dirty and clean areas with moderate fecal contamination. On farms (#), bedding samples were not separated into these categories because bedded areas were uniformly contaminated with fecal material. Following collection, bedding samples were placed on ice in a cooler for transport to the laboratory, where they were stored at 3°C until processing. Bedding materials were cut into 1-5 cm pieces and homogenized by hand. 5 g subsamples were soaked in enough 0.5 M NaOH to cover the sample for 16-18 hours. Samples were then thoroughly washed on a 212 µm sieve on top of a 20 µm sieve with tap water. 10 mL of material retained on the 20 µm sieve was transferred to a 50 mL Falcon tube, and the tube was filled to the 50 mL mark with flotation solution and centrifuged at 2000 rpm for 7 minutes. The supernatant was poured off and the pellets were washed with tap water on a 20 µm sieve. Material collected on the sieve was transferred to a 15 mL Falcon tube and centrifuged at 2000 rpm for 7 minutes. The supernatant was carefully drawn off and the pellet resuspended in ~ 2 mL of flotation solution. Both chambers of a McMaster slide were filled with solution and the slide was set to rest on the benchtop for 5 minutes. All eggs in the slide were counted under a microscope at 40 x magnification. Bedding samples are currently being processed so we do not have data to report at this time.
There were a few aspects of the project that differed from the original methods. Sample collection began in fall of 2022 instead of spring 2023 as proposed. Fecal sample collection followed the protocol outlined in the proposal methods section, but fewer samples were collected per farm than were proposed. This was due to the size of the swine herds at the time of sampling; the number of hogs on each farm at the time of each sampling ranged from 0-25 (average 15 +/-3 hogs). One farm did not fill out the written survey during sampling. Parasitological data from the fall sampling of this farm will not be used in statistical analyses due to the lack of farm data to compare.
To begin objective 1, a parasitological field survey was conducted on nine farms in Pennsylvania where hogs were being raised with outdoor access and without the use of synthetic anthelmintics. Participating farms represented a variety of management practices. At the time of sampling, four farms had hogs on pasture, three had hogs in the forest, and one had hogs on an outdoor concrete pad. Four of eight surveyed farms reported using non-synthetic anthelmintic remedies like diatomaceous earth and apple cider vinegar, and three implement biosecurity protocols to prevent disease in their herds. Parasite eggs were found in the feces, soil, and/or bedding on all 9 farms sampled. 76.7% of sampled hogs were infected with at least one species of parasite. Parasite eggs were demonstrated in soil and bedding of both occupied and previous housing and pasture areas, indicating on-farm transmission is possible.
The fecal and soil sample results for each individual farm were shared with participating farmers to educate them on the status of parasites in their herd.
Education & Outreach Activities and Participation Summary
Participation Summary:
Outreach and educational activities will be conducted routinely via personal, phone, and email communications. Additionally, extension will be performed by the project team attending grower’s meetings/conferences and organizing and presenting project information and results at the Rodale Institute annual field day. In 2021, Rodale Institute’s on-site field day drew in around 450 farmers, scientists, educators, students, and consumers from 22 states and Washington D.C., and 250 people enrolled for the virtual field day event. Project findings, farmer experience, and technological developments will be shared with stakeholders through web articles and free webinars on the Rodale Institute website. Videos will be recorded and posted online as a tool that farmers can watch and refer to at their convenience. Fact sheets will be produced to highlight the findings of the study which will be available to the public at no charge. Research personnel will also present the research findings to the stakeholders in local and regional meetings like the annual Pasa Sustainable Agriculture conference in Lancaster, PA. The research results will be published and publicly available in the form of fact sheets, bulletins, and research reports. Data showing the effects of management practices on swine parasite infection will be submitted to relevant peer-reviewed journals.
We have not done any outreach or educational activities during this reporting period because data collection is ongoing.
Learning Outcomes
During farm visits, participating farmers were educated on common swine parasite species, as well as current knowledge on the prevalence of swine parasites in United States organic and pastured pig farms. Fecal and soil sample results were shared with participating farmers to educate them on the level of parasitosis on their farms, which is something that many were unaware of prior to this study.
Project Outcomes
This section is not relevant to our project at this time because data collection is ongoing.