Veterinary Parasitology 101 (2001) 143–153

Ecological influences on transmission rates ofAscaris suum to pigs on pastures

A. Roepstorff a,∗, K.D. Murrell a,J. Boes a,b, S. Petkevicius a,c

a Danish Centre for Experimental Parasitology, Royal Veterinary and Agricultural University,Dyrlægevej 100, Frederiksberg C, DK-1870 Copenhagen, Denmark

b Veterinary and Food Advisory Service, Danish Bacon and Meat Council, Axeltorv 3,DK-1609 Copenhagen V, Denmark

c Lithuanian Veterinary Institute, LT-4230 Kaišiadorys, Lithuania

Received 15 January 2001; received in revised form 30 May 2001; accepted 30 May 2001

Abstract

A study was conducted to determine the distribution and transmission rate of Ascaris suum eggsand Oesophagostomum dentatum larvae in a pasture/pig house facility, which during the precedingsummer was contaminated with helminth eggs by infected pigs. In May, four groups of 10 helminthnaıve tracer pigs were exposed to fenced sections of the facility for 7 days and necropsied forparasite recovery 9–10 days later (trial 1). The highest rate of A. suum transmission (201 eggs perday) occurred in the pig house (A). On the pasture, egg transmission decreased with the distancefrom the house: 8 eggs per day in the feeding/dunging area (B); 1 egg per day on the nearest pasture(C); <1 egg per day on the distant pasture (D). Only a few O. dentatum infections were detected,indicating a poor ability of the infective larvae to overwinter. Soil analyses revealed that the highestpercentage (5.8%) of embryonated A. suum eggs were in the house (A). Subsequently, the facilitywas recontaminated with A. suum eggs by infected pigs. A replicate trial 2 was conducted in thefollowing May. A major finding was the complete reversal of egg distribution between the 2 years(trials 1 and 2). In contrast to previous results, the highest rates of transmission (569 and 480 eggsper day) occurred in pasture sections C and D, and the lowest transmission rates (192 and 64 eggsper day) were associated with the feeding/dunging sections and the house (B and A). Soil analysesagain supported the tracer pig results, as the pasture sections had the highest concentrations ofembryonated eggs. Detailed soil analysis also revealed a non-random, aggregated egg distributionpattern. The different results of the two trials may be due to the seasonal timing of egg depositionand tracer pig exposure. Many eggs deposited during the summer prior to trial 1 may have died

∗Corresponding author. Tel.: +45-35-282775; fax: +45-35-282774.E-mail address: aro@kvl.dk (A. Roepstorff).

0304-4017/01/$ – see front matter © 2001 Elsevier Science B.V. All rights reserved.PII: S0304 -4017 (01 )00506 -4

144 A. Roepstorff et al. / Veterinary Parasitology 101 (2001) 143–153

rapidly due to high temperatures and dessication, especially when they were not protected by thehouse, while deposition in the autumn may have favored egg survival through lower temperatures,more moisture, and greater sequestration of eggs in the soil by rain and earthworms. The latter eggsmay, however, not have become embryonated until turnout the next year. The results demonstratethat yearly rotations may not be sufficient in the control of parasites with long-lived eggs, such as A.suum, and that a pasture rotation scheme must include all areas, including housing. © 2001 ElsevierScience B.V. All rights reserved.

Keywords: Ascaris suum; Oesophagostomum dentatum; Transmission; Pig-nematoda; Outdoor rearing

1. Introduction

Ascaris suum and Oesophagostomum dentatum are both known to be transmitted effec-tively amongst outdoor pigs in Denmark, and both helminths may survive the winter andbe an infection risk for parasite naıve pigs turned out in the following spring (Petkeviciuset al., 1996; Roepstorff and Murrell, 1997a,b). Development and survival of the infectivestages of both helminths are dependent upon microclimate (Rose and Small, 1980, 1981;Larsen, 1996; Larsen and Roepstorff, 1999; Kraglund, 1999), and transmission rates maytherefore be associated with biological factors such as vegetation, and physical featuressuch as wallowing facilities, housing, etc. Pigs tend to defecate in particular dung areas(Mejer et al., 1998), however, in practice very little is known about the variation in trans-mission rates within rearing facilities, e.g. pastures, swine housing, feeding and dungingareas. This information is central to the development of pasture rotation schemes to controlpig ascarosis on pasture. The present experiment was designed to study parasite egg distri-bution patterns and transmission rates by exposing helminth naıve tracer pigs to differentareas of a naturally contaminated pasture and housing facility and thereafter determining thenumbers of up taken worms. The results could help creating the background for elaboratingstrategies for parasitic control by means of pasture management in free-range or extensiveswine rearing systems.

2. Materials and methods

2.1. Experimental pasture and housing

The experimental area was located at a University Research Farm, approximately 20 kmwest of Copenhagen. The climate in Denmark is temperate with a monthly rainfall of38–79 mm throughout the year and the lowest and highest monthly mean temperature being0.0◦C (February) and 15.7◦C (August), respectively (Rosenørn and Lindhard, 1994). Arectangular pasture (1820 m2), containing at one end a primitive non-insulated wood pighouse, was contaminated with A. suum (x = 1564 epg, eggs per gram feces) and O. denta-tum (x = 738 epg) eggs by 10 infected pigs from June to October 1995 (Petkevicius et al.,1996). These pigs were nose-ringed to protect the vegetation from uprooting. In the follow-ing spring (1996), the pasture was divided into four sections using electric fencing; (A) the

A. Roepstorff et al. / Veterinary Parasitology 101 (2001) 143–153 145

Fig. 1. An overview of the A. suum contaminated pasture that was divided into sections A (house); B (feedingarea); C (near pasture), and D (distant pasture). The wallowing facility and the defecation area are shown in sectionB. A W-route with marked sub-sampling sites is visualized on section D.

house (15 m2) with straw bedding on a floor of a mixture of dry soil and old straw, includingsome old pig feces; (B) a section surrounding the house (125 m2), which contained feedingtroughs and drinking water nipples as well as a wallowing facility, and no vegetation dueto pig activity the preceding year; the remaining pasture, featuring good grass cover, wasdivided into a proximal (C) and a distant (D) section both measuring 840 m2. The pasturesections are shown in Fig. 1.

2.2. Experimental pigs and tracer protocol

For trial 1 (1996), Danish Landrace/Yorkshire/Duroc cross-bred female pigs and castratedmales were obtained from a specific pathogen-free (SPF) herd. Repeated fecal examinationsindicated that all weaners produced in the herd were helminth-free and no excretion of eggswas observed in the pigs selected for the current study. One group of 10 tracer pigs wasexposed to each of the four sections (A–D). The pigs were randomized in the groupsaccording to sex and body weight and allowed to adapt to feed and outdoor conditions on ahelminth-free pasture for 12 days prior to turnout to the experimental sections on 13 May.At turn out on the experimental pasture, the pigs weighed 29.6 ± 3.7 kg. The feed (groundbarley, soy meal, minerals and vitamins) has previously been shown to be favorable forestablishment and development of these helminths (Petkevicius et al., 1996, 1997). After7 days of exposure to the pasture sections, the pigs were removed to helminth-free indoorfacilities for a subsequent 9–10 days, after which they were slaughtered and worm recoverieswere carried out. By this protocol, all A. suum and O. dentatum originating from the infectiveeggs/larvae from the experimental pasture sections were 9–17 days old at slaughter, andexpected to be located in the lumen of the small and large intestine, respectively (Christensenet al., 1995; Roepstorff et al., 1997).

The tracer pig experiment was repeated in the spring 1997 (trial 2) in the same pasture/house facility using similar cross-bred pigs (weighing 32.3 ± 3.0 kg) that were turned outon the contaminated sections on 5 May. In between the two trials, the experimental pasturewas used for a separate study, in which nose-ringed pigs were allowed to graze the pasture

146 A. Roepstorff et al. / Veterinary Parasitology 101 (2001) 143–153

from late May to early October (see Boes et al., 1998). These pigs became naturally infectedwith A. suum and very few O. dentatum. They were treated with anthelmintics at week 10post-turnout (late July) and allowed to become reinfected during an additional 10-weekperiod before they were removed. Consequently, sections A–D were recontaminated withA. suum eggs for two 3–4 week periods in July (4 pigs had fecal egg counts increasing from0 to almost 6000 epg) and in September and early October (25 pigs had fecal egg countsincreasing from 0 to approximately 1000 epg).

2.3. Sampling and parasitological techniques

In both trials, individual fecal samples were collected at the time of turnout onto theexperimental pasture (sections A–D) and at slaughter. Fecal egg counts were determinedby a concentration McMaster technique with a lower detection limit of 20 eggs per gramfeces (epg) (Roepstorff and Nansen, 1998).

In each trial, four soil samples were collected from each section, including the house:two at the beginning and two at the end of the tracer pig exposure period. This was done bywalking W-routes over each section and collecting soil from the upper 2 cm at every one tosix steps (at least 20 small sub-samples were pooled for each of the four W-routes in eachtrial). A W-route is visualized in Fig. 1. For egg analyses, the soil was thoroughly mixed and5 g sub-samples were soaked in 0.5 M NaOH overnight, followed by repeated flotation andsieving (see Larsen and Roepstorff, 1999). This gave a lower detection limit of 1 A. suumegg per 5 g of soil. In trial 1, 50 g from each of the four soil samples were also baermannizedovernight at 37◦C for isolation of O. dentatum L3-larvae (Roepstorff and Nansen, 1998);this was not repeated in trial 2. Extra soil samples were obtained 11 days after removalof the tracer pigs of trial 2 (2 June 1997); a W-route of soil sampling was again carriedout on sections B–D (the samples collected from section A were lost due to a laboratoryaccident). The 12, 24, and 24 sub-samples (on B, C, and D sections, respectively) were notpooled but analyzed individually to more precisely determine the distribution of eggs onthe pasture.

Blood was collected from all pigs prior to exposure and at slaughter, and the serawere tested for IgG antibodies specific against A. suum excretory/secretory antigens whichwere obtained from in vitro-cultivated L3-larvae; the ELISA technique employed has beendescribed in detail by Lind et al. (1993). The number of superficial white spots on the liverswere counted at slaughter. The number of A. suum in the whole small intestine (both trials)and of O. dentatum in the whole large intestine (trial 1) were enumerated after isolation bymeans of an agar gel technique (Slotved et al., 1996, 1997).

2.4. Calculations and statistics

All calculations were performed using the SAS® Release 6.12 software package. Trans-mission rates (larvae per day) were calculated as the number of intestinal A. suum larvaeobtained per day of exposure. The parameter k of the negative binomial distribution wasestimated as k = mean2/(variance − mean). Since A. suum worm burdens and white spotnumbers did not fit normal distributions, the differences between pasture areas were testedby the Wilcoxon rank-sum test (PROC NPAR1WAY).

A. Roepstorff et al. / Veterinary Parasitology 101 (2001) 143–153 147

3. Results

One pig in trial 2, section B, died due to erysipelas. No clinical symptoms were observed inany other pigs. No helminth eggs were observed in any of the fecal samples taken throughoutthe study.

No O. dentatum larvae were found in the soil samples collected in trial 1 (1996) and atslaughter two pigs (one from sections B and C, respectively) were found to harbor one singleworm each, indicating an extremely low transmission rate in early spring. This indicated verypoor overwintering ability of infective O. dentatum larvae in Denmark. No examinationsfor O. dentatum were carried out in trial 2.

The A. suum intestinal worm counts as well as numbers of liver white spots recordedat necropsy for both trials 1 and 2 are presented in Table 1. All A. suum recovered wereL3–L4-larvae (<1 cm), verifying that the tracer pigs did not have a previous infection. Intrial 1, the group A tracer pigs (house) harbored significantly more A. suum than any ofthe other groups (P < 0.001). In addition, there was a clear relationship between A. suuminfection level and distance from the pig house (A): the group B pigs (feeding area) weremore heavily infected than the pigs of groups C and D (P < 0.001) which in turn weresignificantly different from each other (P < 0.05). These differences were supported by thenumbers of liver white spots, which were reduced by approximately a factor 1000, movingfrom area A to D, indicating a clear gradient in soil egg transmission to tracer pigs movingfrom the house to the most distant pasture. Analyses of the soil (Table 1) revealed that whilesections A–C had comparable soil egg counts (13–38 eggs per 5 g) the percent of larvatedeggs decreased markedly from sections A to C; only few unembryonated eggs were foundin the soil of area D (distant pasture).

Surprisingly, the results of trial 2 differed markedly from those of trial 1. The A. suumworm burdens in pigs exposed to the different sections were in reverse order: the wormrecovery of tracer pigs in sections C and D, while not significantly different were markedlyhigher (P < 0.05) than that in group B, which in turn was significantly greater (P < 0.01)than that in group A. Again, the numbers of white spots reflected the worm burdens. Thehighest percentage of larvated eggs occurred in section C, the section with highest wormburdens and white spot numbers. Overall, the worm burdens were higher in trial 2, reflectingthe higher level of contamination, resulting from the pigs in the summer between the twotrials.

The ELISA results indicated that all pigs were A. suum naıve prior to exposure, withincreases in OD-values following exposure that correlated with the worm burdens andwhite spot numbers at slaughter (data not shown).

The pigs in groups C and D in trial 2 had mean worm burdens of 3985 and 3362 larvae perpig, respectively, indicating that on average 569 and 480 larvae, respectively, had completedmigration and established in the small intestine per day, with an individual maximum trans-mission rate of >1600 larvae per day in both groups (Table 1). With a single exception (trial1, section B), k of the negative binomial distribution exceeded 1 in all tracer pig groups,indicating that the worms were not heavily overdispersed within the host populations.

The results of the analyses of individual soil samples collected from sections B–D in trial2 are shown in Fig. 2. Although 88% of these samples were positive for Ascaris eggs, almost50% of the eggs were found in 3 of the 60 samples only, indicating a pronounced aggregated

148A

.Roepstorffetal./Veterinary

Parasitology101

(2001)143–153

A. Roepstorff et al. / Veterinary Parasitology 101 (2001) 143–153 149

Fig. 2. Trial 2: second soil sampling. Distribution of unembryonated (gray columns) and embryonated (darkcolumns) A. suum eggs in individual samples of soil collected along W-routes across sections B (feeding area); C(near pasture) and D (distant pasture) of the experimental pasture, on 2 June 1997 (11 days after removal of thetracer pigs). The location of the wallowing facility (mud pool) and the defecation area (dung area) of section B isvisualized. The individual samples are shown in original sequence, and for sections C and D, the left sub-sampleswere nearest the house. Note that three columns have been shortened due to extremely high egg counts.

egg distribution in the soil. The k of the negative binomial distribution was calculated tobe 1.07, 0.23, and 0.11 for sections B, C, and D, respectively, showing a high aggregationon all pasture sections. The samples in the defecation area, that was included in section B,generally contained more eggs than the other part of this section (Fig. 2), but did not havesuch high egg concentrations, as would have been expected from the large amounts of fecesdeposited. The wallowing facility only contained moderate numbers of eggs, none of whichhad embryonated (Fig. 2).

4. Discussion

The present study revealed that the distribution and transmission of overwintering infectiveA. suum eggs from different sections of an outdoor pig facility may vary considerably fromyear to year. In the first year, the egg distribution and transmission rate decreased signifi-cantly from the house to the most distant pasture, while the inverse was true in the secondyear. In both trials there was a strong relationship between the presence of embryonatedeggs in the soil of the various sections of the pig facility (pasture, feeding/dunging area,house) and the transmission rate to tracer pigs.

In an attempt to explain the causes underlying these contrasting results, it is important tolook at the history of the pasture. Prior to 1995, the pasture was almost free of A. suum (seeNansen et al., 1996). In 1995, a high Ascaris egg contamination occurred from late June toearly October 1995 (Petkevicius et al., 1996). During the pig grazing season of 1996, the

150 A. Roepstorff et al. / Veterinary Parasitology 101 (2001) 143–153

pasture was contaminated in two 3–4 week periods during July and September/October,respectively (Boes et al., 1998). Of the two egg cohorts from 1996, only eggs depositedin July may have experienced high enough temperatures to achieve embryonation prior toturnout of the trial 2 tracer pigs in the spring 1997. The eggs in the autumn cohort, however,may have only contributed to the large pool of unembryonated eggs found in the spring 1997(see Larsen and Roepstorff, 1999; Kraglund, 1999). The important difference between trials1 and 2 may be due to the state of egg embryonation between the spring of 1996 and the springof 1997, when eggs have been accumulating for 1 or 2 years, respectively. The explanationfor why so few eggs embryonated on the pasture after their deposition in 1995 (prior totrial 1), may be that the majority of eggs deposited during the summer died within fewweeks due to dessication during the relatively hot and dry July–August 1995, as extremelyhigh mortality of A. suum eggs can occur during summer months (Larsen and Roepstorff,1999; Kraglund, 1999). In contrast, eggs deposited in the autumn may have a much highersurvival rate because they are transported quickly down into the soil by rain and earthworms.These eggs will remain unembryonated until the temperature rises the following summer(Larsen and Roepstorff, 1999; Kraglund, 1999). Therefore, the majority of eggs survivingon the pasture from 1995 may not have become infective and available until the spring 1997(trial 2). The house, however, may have protected eggs deposited during the summer 1995against dessication, UV-light, etc. thereby providing more infective eggs in that section inthe spring 1996 (trial 1). Why so relatively few embryonated eggs were present in the housein the spring 1997 remains unclear.

Hitherto, the natural uptake rate of infective A. suum eggs in pigs has been largelyunknown. Under indoor conditions, an extremely wide range of transmission rates mayoccur, as transmission depends on hygiene level and physical/chemical factors (Nansen andRoepstorff, 1999). Conditions may be such that no transmission occurs (Roepstorff, 1997)or just 1 day of exposure to an A. suum contaminated pen may result in extremely large andlife threatening worm burdens (Marquardt, 1965). Under outdoor conditions, A. suum maybe transmitted effectively to piglets within the first few weeks of life (Roepstorff et al., 1992)and continuous transmission may result in extremely high liver white spot scores at 7–10weeks of age (Jolie et al., 1998). The calculated mean transmission rates in trial 2 (64–569larvae per day) and in the house of trial 1 (210 larvae per day) are considerably higher thanthe means of 61–257 larvae per week (9–37 larvae per day) we have obtained using anidentical tracer protocol with groups of tracer pigs exposed on pastures (Mejer et al., 2000;Thomsen et al., 2001). These pastures were, however, contaminated by pigs at lower stock-ing rates than those in the present experiment and the contamination took place from Juneto September, immediately before the tracer pigs were turned out in October. Consequently,many of the eggs deposited in the warm months may have died and those deposited in theautumn may not have become infective. Despite the estimated high transmission rates fromthe present study, they may actually be underestimations, as only 40–60% of the numberof inoculated infective eggs are normally found to establish successfully in the pig smallintestine (Roepstorff et al., 1997).

Normally, exposed pigs acquire resistance against A. suum (see Nansen and Roepstorff,1999), and a negative relationship between adult worm burdens and dose rates has beendescribed (Andersen et al., 1973; Jørgensen et al., 1975; Coates, 2000). Therefore, mea-suring accumulated worm burdens in helminth naıve tracer pigs as an indication of pasture

A. Roepstorff et al. / Veterinary Parasitology 101 (2001) 143–153 151

transmission potential, underestimates transmission rates if the pigs are examined after themajority of worms have been expelled from the small intestine (see Roepstorff et al., 1997).Indirect measures like liver white spot numbers and serum antibody levels may in that casebe useful, although imprecise, for estimating transmission rates (Roepstorff and Murrell,1997a; Mejer et al., 2000; Thomsen et al., 2001). However, by making use of a recentlydeveloped agar gel technique (see Slotved et al., 1997), approximately 50% of the ingestedviable eggs may easily be recovered by days 10–14 post-infection (pi) as minute larvaeaccumulated in the small intestine (Roepstorff et al., 1997). This high recovery was shownto be independent of dose rate within the range of 100–10,000 eggs, while a beginningreduction in numbers of intestinal worms was observed by day 17 pi at a dose rate of 10,000eggs (Roepstorff et al., 1997). With this knowledge, it was possible to design the presenttracer protocol, in which the pigs were exposed for 7 days and subsequently stabled underhelminth-free conditions for further 9–10 days so that eggs picked up during exposure de-veloped to 9–17 days old larvae at the time of necropsy. To avoid any risk of expulsion whenhigh transmission rates are expected, future tracer pig protocols should consider shorteningthe exposure period to about 4 days.

Although outdoor pigs may eat considerable amounts of soil (Mejer et al., 1998), theworm burdens in this study may indicate that soil analyses can underestimate the eggconcentrations on pastures, because at the mean egg densities found, 1.3 kg soil would havebeen eaten per day to account for the observed mean worm burdens in two of the groups. Forthe other groups, similar estimates are 150–400 g soil eaten per pig per day. If transmissionrates have actually been higher than indicated by the worm counts (see above), the soilanalyses may have been even more imprecise.

Soil sample analysis revealed that A. suum eggs were unevenly distributed on the pasturesections (k < 1), and that ‘hot spots’ with extremely high numbers of eggs could be found,but, surprisingly, neither the defecation area nor the wallowing facility were particularlyhighly infective compared to other areas. An aggregated distribution of eggs may theoret-ically lead to considerable heterogeneity in the individual worm burdens, as suggested byAnderson and Gordon (1982). Nevertheless, the worm distributions within the four groupswere only a little overdispersed (k was generally >1) in both trials, indicating that the aggre-gated distribution of eggs on the different pasture sections did not result in overdispersedworm populations in the pigs. This may reflect the intense grazing behavior of pigs onpasture, which results in individuals visiting most of the area.

The O. dentatum results were less spectacular, as only two specimens were observedin totally 40 pigs (trial 1), giving a mean transmission rate of 0.007 L3 per pig per dayof exposure to the pasture. The pasture was contaminated with O. dentatum eggs fromJune to October 1995 (Petkevicius et al., 1996), but the low transmission rate indicatesthat the L3-larvae were almost eliminated during the winter, both from the non-insulatedhouse and from the pasture sections. This was further confirmed by the very sporadic andlow egg excretion of the pigs grazing the area during the summer of 1996 (Boes, personalcommunication). On the same research farm, Petkevicius et al. (1996) showed a significantoverwintering of infective O. dentatum larvae in the winter 1994–1995, whereas Roepstorffand Murrell (1997b) only found a very low survival rate (like in the present study) inthe winter 1993–1994, while Mejer et al. (2000), and Thomsen et al. (2001) found thatO. dentatum became completely eradicated from heavily contaminated pastures during the

152 A. Roepstorff et al. / Veterinary Parasitology 101 (2001) 143–153

winter 1996–1997. Overall, these results show that pasture infectivity with O. dentatumlarvae is markedly reduced during a typical Danish winter, and larval survival dependsupon weather conditions in combination with the relevant physical/biological factors in thepig facility.

If only trial 1 had been carried out, the conclusion could possibly have been that apasture rotation strategy for control would only have to focus on the house and the feed-ing/dunging area as high risk. However, by experimenting over 2 years, it is clear that apasture rotation control scheme must include the whole pig facility. The study results alsoemphasize the importance of long-term studies when dealing with transmission patternsof resistant, long-lived, but slowly developing eggs, like those of A. suum. As the eggssurvive in considerable numbers from year-to-year, multi-year rotation strategies shouldbe adopted. However, no such strategies have been tested in practice, and therefore thereis a great need for further research on practical pasture management in the control of pighelminths in extensive outdoor systems.

Acknowledgements

The authors gratefully acknowledge the skilful technical assistance of Niels Midtgaard,Morten Cleeman Hors, Gudrun Ellendersen, and Lillian Sztuk. This study was supported fi-nancially by The Danish Environmental Research Programme. The Danish NationalResearch Foundation is acknowledged for the financial support of this and concurrentprojects under the auspices of The Danish Centre for Experimental Parasitology.

References

Andersen, S., Jørgensen, R.J., Nansen, P., Nielsen, K., 1973. Experimental Ascaris suum infection in piglets:inverse relationship between the numbers of inoculated eggs and the numbers of worms established in theintestine. Acta Pathol. Microbiol. Scand. B 81, 650–656.

Anderson, R.M., Gordon, D.M., 1982. Processes influencing the distribution of parasite numbers within hostpopulations with special emphasis on parasite-induced host mortalities. Parasitology 85, 373–398.

Boes, J., Medley, G.F., Eriksen, L., Roepstorff, A., Nansen, P., 1998. Distribution of Ascaris suum in experimentallyand naturally-infected pigs and comparison with Ascaris lumbricoides infections in humans. Parasitology 117,589–596.

Christensen, C.M., Barnes, H.E., Nansen, P., Roepstorff, A., Slotved, H-C., 1995. Experimental Oesophagostomumdentatum infection in the pig: worm populations resulting from single infections with three doses of larvae.Int. J. Parasitol. 25, 1491–1498.

Coates, S., 2000. Population Dynamics of Ascaris suum in Pigs: Modelling and Experimental Studies. Ph.D.Thesis, University of Warwick, p. 161.

Jolie, R., Bäckström, L., Pinckney, R., Olson, L., 1998. Ascarid infection and respiratory health in feeder pigsraised on pasture or in confinement. Swine Health Prod. 6, 115–120.

Jørgensen, R.J., Nansen, P., Nielsen, K., Eriksen, L., Andersen, S., 1975. Experimental Ascaris suum infectionin the pig: population kinetics following low and high levels of primary infection in piglets. Vet. Parasitol. 1,151–157.

Kraglund, H.-O., 1999. Survival, development and dispersal of the free-living stages of Ascaris suum,Oesophagostomum dentatum and Trichuris suis at the pasture. Ph.D. Thesis, Royal Veterinary and AgriculturalUniversity, Copenhagen, p. 124.

A. Roepstorff et al. / Veterinary Parasitology 101 (2001) 143–153 153

Larsen, M.N., 1996. Årstidsvariation i overlevelse, udvikling og spredning for 4 porcine parasitter på friland.M.Sc. Thesis, University of Copenhagen, p. 98.

Larsen, M.N., Roepstorff, A., 1999. Seasonal variation in development and survival of Ascaris suum and Trichurissuis eggs on pastures. Parasitology 119, 209–220.

Lind, P., Eriksen, L., Nansen, P., Roepstorff, A., 1993. Response to repeated inoculations with Ascaris suum eggsin pigs during the fattening period. II. Specific IgA, IgG and IgM antibodies determined by enzyme-linkedimmunosorbent assay. Parasitol. Res. 79, 240–244.

Marquardt, W.C., 1965. Ascarid infection in a specific pathogen-free pig. J. Am. Vet. Med. Assoc. 146, 701–702.Mejer, H., Thomsen, L.E., Wendt, S., 1998. Transmission af helminther hos grise på friland. Betydningen af

næsering, belægningsgrad, fodersammensætning og adfærd. M.Sc. Thesis, University of Copenhagen, p. 145.Mejer, H., Wendt, S., Thomsen, L.E., Roepstorff, A., Hindsbo, O., 2000. Nose-rings and transmission of helminths

in outdoor pigs. Acta Vet. Scand. 41, 153–165.Nansen, P., Roepstorff, A., 1999. Parasitic helminths of the pig: factors influencing transmission and infection

levels. Int. J. Parasitol. 29, 877–891.Nansen, P., Larsen, M., Roepstorff, A., Grønvold, J., Wolstrup, J., Henriksen, S.A., 1996. Control of

Oesophagostomum dentatum and Hyostrongylus rubidus in outdoor-reared pigs by daily feeding with themicrofungus Duddingtonia flagrans. Parasitol. Res. 82, 580–584.

Petkevicius, S., Bach Knudsen, K.E., Nansen, P., Roepstorff, A., 1996. The influence of diet on infections withAscaris suum and Oesophagostomum dentatum in pigs on pasture. Helminthologia 33, 173–180.

Petkevicius, S., Bach Knudsen, K.E., Nansen, P., Roepstorff, A., Skjøth, F., Jensen, K., 1997. The impact of dietsvarying in carbohydrates resistant to endogenous enzymes and lignin on populations of Ascaris suum andOesophagostomum dentatum in pigs. Parasitology 114, 555–568.

Roepstorff, A., 1997. Helminth surveillance as a prerequisite for anthelmintic treatment in intensive sow herds.Vet. Parasitol. 73, 139–151.

Roepstorff, A., Murrell, K.D., 1997a. Transmission dynamics of helminth parasites of pigs on continuous pasture:Ascaris suum and Trichuris suis. Int. J. Parasitol. 27, 563–572.

Roepstorff, A., Murrell, K.D., 1997b. Transmission dynamics of helminth parasites of pigs on continuous pasture:Oesophagostomum dentatum and Hyostrongylus rubidus. Int. J. Parasitol. 27, 553–562.

Roepstorff, A., Nansen, P., 1998. The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine: AFAO Animal Health Manual. Food and Agriculture Organization of the United Nations, Rome, Italy, p. 161.

Roepstorff, A., Eriksen, L., Slotved, H.-C., Nansen, P., 1997. Experimental Ascaris suum infection in the pig:worm population kinetics following single inoculations with three doses of infective eggs. Parasitology 115,443–452.

Roepstorff, A., Jørgensen, R.J., Nansen, P., Henriksen, S.A., Skovgaard Pedersen, J., Andreasen, M.,1992. Parasitter hos økologiske svin. Rapport over projekt finansieret af Jordbrugsdirektoratet underLandbrugsministeriet, p. 36.

Rose, J.H., Small, A.J., 1980. Observations on the development and survival of the free-living stages ofOesophagostomum dentatum both in their natural environments out-of-doors and under controlled conditionsin the laboratory. Parasitology 81, 507–517.

Rose, J.H., Small, A.J., 1981. The relationship between pasture herbage and the development and survival of thefree-living stages of Oesophagostomum dentatum. J. Helminthol. 55, 109–113.

Rosenørn, S., Lindhard, K., 1994. Dansk vejr i 100 år. Lademann, Copenhagen, p. 213.Slotved, H.-C., Barnes, E.H., Bjørn, H., Christensen, C.M., Eriksen, L., Roepstorff, A., Nansen, P., 1996. Recovery

of Oesophagostomum dentatum from pigs by isolation of parasites migrating from large intestinal contentsembedded in agar gel. Vet. Parasitol. 63, 237–245.

Slotved, H.-C., Barnes, E.H., Eriksen, L., Roepstorff, A., Nansen, P., Bjørn, H., 1997. Use of an agar gel techniquefor large scale application to recover Ascaris suum larvae from intestinal contents of pigs. Acta Vet. Scand. 38,207–212.

Thomsen, L.E., Mejer, H., Wendt, S., Roepstorff, A., Hindsbo, O., 2001. The influence of stocking rate ontransmission of helminth parasites in pigs on permanent pasture during two consecutive summers. Vet. Parasitol.99, 129–146.