K. R. Price,* 1 M. T. Guerin, and J. R. Barta *

Transcription

1 2014 Poultry Science Association, Inc. Success and failure: The role of relative humidity levels and environmental management in live Eimeria vaccination of cage-reared replacement layer pullets K. R. Price,* 1 M. T. Guerin, and J. R. Barta * * Department of Pathobiology, and Deptartment of Population Medicine, University of Guelph, Guelph, ON, Canada N1G 2W1 Primary Audience: Flock Supervisors, Veterinarians, Service Technicians, Researchers SUMMARY Contrary to the traditional view that layer flocks housed in conventional cages are unlikely to suffer coccidiosis caused by Eimeria species, this enteric disease has become an emergent issue. Coccidiosis outbreaks in layers are frequently associated with the failure to develop protective immunity at a young age. Layer hens housed in cages are usually sourced from replacement layer pullets housed in similar cages in the rearing barn. Live coccidiosis vaccines administered to young chicks provide a small dose of vaccine oocysts that infect and replicate within the vaccinated birds, resulting in the release of progeny oocysts into the rearing environment; these oocysts must become infective (i.e., sporulate) and then reinfect the partially immune birds in order for the vaccine to generate complete protective immunity. Three factors are needed for successful oocyst sporulation: temperature (4 to 37 C; optimal 29 C), oxygen access, and adequate RH. The purpose of this study was to assess the degree of protective immunity elicited by birds spray-vaccinated with a live coccidiosis vaccine at the hatchery. During the critical period for oocyst sporulation and cycling, RH levels decreased to 13 to 19%, resulting in inadequate oocyst cycling and minimal protection against homologous challenge at 6 and 12 wk of age. This vaccination failure highlights the need to monitor RH in the barn and modify the barn environment so that conditions promote, rather than impede, the success of live coccidiosis vaccines. Key words: coccidiosis, oocysts per gram, parasitology, poultry management, caged housing, brooding 2014 J. Appl. Poult. Res. 23 : DESCRIPTION OF PROBLEM Coccidiosis, caused by highly host-specific parasites in the genus Eimeria, has plagued the chicken industry since intensification of commercial poultry production in the 1940s [1]. This enteric disease can manifest in subclinical or clinical forms that can affect production (e.g., decreased weight gain, poorer FCR, decreased egg production, and increased mortality) [2 6]. Clinical coccidiosis arises in a flock when immunologically naïve birds ingest large numbers of sporulated oocysts [1]. Coccidiosis outbreaks in a flock are frequently associated with the fail- 1 Corresponding author: pricek@uoguelph.ca

2 524 JAPR: Research Report ure to develop sufficient protective immunity at an appropriately young age [7]. Traditionally, in-feed anticoccidial drugs have been used to prevent the disease. However, live coccidiosis vaccines have been in continuous use since the 1950s [8], and these vaccines have been gaining acceptance, albeit to varying degrees, for different chicken production systems and countries. Changes in production methods and housing systems have altered the ways in which environmental management can influence live vaccine success in poultry rearing. Conventional cages are still used in the commercial egg-laying industry in many parts of the world. Layer hens housed in conventional cages are usually sourced from replacement layer pullets housed in similar cages in the rearing barn. Layer hens housed in cages can be infected by several Eimeria species, including Eimeria acervulina, Eimeria maxima, Eimeria necatrix, Eimeria brunetti, Eimeria praecox, Eimeria mitis, and Eimeria tenella. Contrary to the traditional view that layer hens housed in conventional cages are unlikely to suffer clinical coccidiosis [9], the disease (sometimes complicated by necrotic enteritis caused by Clostridium perfringens) has become an emergent issue [10 12]. Observed coccidiosis in layer hens housed in conventional cages suggests that these birds can ingest sufficient numbers of oocysts to induce clinical disease even when housed on mesh flooring. Coccidiosis outbreaks often occur when the pullets are moved to the layer production barn at 18 to 19 wk of age so called new house syndrome [10, 11]. Live vaccination of replacement layer pullets and proactive management of the rearing environment can be used to prevent clinical coccidiosis in layer hens. Live coccidiosis vaccines exploit the ability of the bird to generate species-specific immunity against these parasites after infection [1, 13]. Live coccidiosis vaccines administered to young chicks provide a small dose of vaccine oocysts that infect and replicate within the vaccinated birds, ultimately resulting in the release of many progeny oocysts into the rearing environment; these oocysts must successfully become infective (sporulate to infectivity) in the environment and then reinfect the partially immune birds (via fecal-oral transmission) at an early age in order for the vaccine to generate complete protective immunity in the vaccinated birds [13 15]. Complete protective immunity is defined functionally as the absence of pathogenic effects with limited parasite reproduction and minimal oocyst output [16]. Less efficient oocyst cycling among pullets reared in cages following live vaccine administration has been shown to affect vaccine performance adversely [11, 14]. A preliminary study found that modification to the cage floor environment at 1 d of age with a biodegradable, durable fiber tray that lasted approximately 5 wk allowed for better protection of vaccinated birds against a homologous challenge infection 6 wk later compared with birds reared with a material that lasted only 2 wk [17]. A subsequent study determined that when this durable fiber tray covered 40% of the cage floor, oral-gavaged, livevaccinated birds had lower mean total oocyst output and lesion scores compared with birds reared with 0, 20, or 60% cage floor coverage (CFC) [18]. The latter study mimicked commercial conditions with standard bird densities; however, commercial vaccine administration techniques were not followed and the replacement layer pullets were only challenged at 6 wk of age [18]. In previous experimental work, it was demonstrated that immunity to multiple Eimeria species can last as long as 18 wk [19]. The purpose of the current study was to assess live coccidiosis vaccination success in layer pullets housed in conventional cages with modifications to the cage environment following standard commercial spray vaccination with a multispecies, nonattenuated live coccidiosis vaccine. Vaccine oocyst cycling within cages with different cage floor treatments (0, 20, 40, or 60% coverage with fiber trays) and resultant protection against homologous challenge infection at 6 or 12 wk of age was assessed. MATERIALS AND METHODS Birds, Standard Commercial Vaccination, and Study Location A total of 2,340 Lohmann LSL Lite 1-dold chicks were obtained from a commercial hatchery [20]; the chicks were vaccinated at the hatchery against Marek s disease and infectious bursal disease using standard protocols. A label dose of a nonattenuated live coccidiosis vac-

3 Price et al.: HUMIDITY AND EIMERIA VACCINES 525 cine was used in this trial. Coccivac-D [21] is a multispecies live coccidiosis vaccine containing a proprietary mix of sporulated oocysts of E. tenella, Eimeria mivati, E. acervulina, E. maxima, E. brunetti, Eimeria hagani, E. necatrix, and E. praecox. At the hatchery, chicks were placed in transport boxes at a standard density of 100 chicks per box and then administered Coccivac-D via a spray cabinet system [21] at the recommended dose before transport to a brooder barn within the Poultry Research Facility at the Arkell Research Station, University of Guelph. Upon arrival at the brooder barn, all birds were individually identified using neck tags [22] before experimentation. Pullets were housed in rearing cages [23] at a density of 20 birds per cage (155 cm 2 /bird) from 0 to 6 wk of age. Pullet housing and density during challenge infection are described herein (see experiment 2). Bird densities were in accordance with Canadian Agri-Food Research Council [24] and institutional Animal Care Committee recommendations. These cages and bird densities were used to simulate typical Canadian commercial replacement layer pullet brooding and growing conditions, respectively [24]. Newspaper was placed on the cage floor from 0 to 13 d of age to prevent the chicks feet from falling through the mesh floor. Light intensity and room temperature were set to standard commercial production conditions as outlined by the Lohmann LSL management guide [25] and the Canadian Agri-Food Research Council [24] and maintained automatically; lighting and temperature was checked manually at least twice daily. In addition to automated controls, portable temperature and RH monitors [26] were fitted in all experimental rooms; these monitors recorded room conditions once every 5 min throughout the trial. All bird housing and handling was approved by the University of Guelph Animal Care Committee and completed in accordance with the Canadian Council on Animal Care guidelines [27]. A standard layer pullet diet and water were provided ad libitum to the birds. Experiment 1: Success of Vaccine Administration One hundred 1-d-old chicks were randomly selected from the 2,340 chicks provided by the hatchery. The chicks were randomly assigned to 1 of 5 cages (20 chicks/cage). To determine the relative uptake of vaccine oocysts, all 100 chicks were placed individually into small paper boxes [28] lined with aluminum foil at 6 d postinoculation (DPI) for approximately 2 h. At the end of 2 h, fecal material, if any, from each bird was collected and weighed, and the chicks were returned to their communal cages. The number of oocysts per gram (OPG) of wet feces was determined from each weighed fecal sample (discussed in more detail herein). Experiment 2: Comparison of the Level of Protective Immunity Between Treatment Groups Experiment 2 consisted of 2 phases (Figure 1). The treatment phase encompassed the period of time from 1 to 42 d of age, during which the pullets were reared with different treatments. The challenge infection phase covered the period of time from challenge with mixed Eimeria species at either 42 or 84 d of age to 54 or 96 d of age, respectively, during which the level of protective immunity was assessed. Experiment 2: Treatment Phase. Durable fiber trays [29] offering different percentages of cage floor covering (CFC; treatment) were compared at (1) 0, (2) 20, (3) 40, and (4) 60% CFC. In total, 2,240 pullets (20 birds per cage 4 treatments 28 cages per treatment) distributed into 2 separate rooms were included in this phase of the experiment. At 1, 31, 72, and 105 d of age, all birds within each cage were weighed individually and the average weight per bird for each cage was determined. On 6, 9, and 12 DPI, brown paper [30] was placed over the newspaper and under the trays to collect feces. From 15 to 27 DPI, aluminum foil was placed beneath the cages to collect feces that dropped through the wire floor. From 6 to 27 DPI, feces were collected for 4 h for each collection; from 30 to 60 DPI, a sample of feces was collected every 3 d to assess cycling of the parasite (discussed further herein). At 42 and 84 d of age, 2 pullets were randomly selected from each cage and weighed individually (prechallenge BW) and assessed for animal welfare using 4 welfare parameters as described by Price et al.

4 526 JAPR: Research Report Figure 1. A general timeline describing the phases, pullet numbers per cage, and timing of measurements in experiment 2 for 1 treatment group replicate. A total of 28 replicates in each of 4 treatments followed this scheme. Color version available in the online PDF.

5 Price et al.: HUMIDITY AND EIMERIA VACCINES 527 [18]: (1) plumage cleanliness, (2) food pad dermatitis, (3) hock burn, and (4) bumble foot [31]. Experiment 2: Challenge Infection Phase. At d 42 (postbrood challenge) and 84 (midgrowth challenge) of age, the 2 randomly selected pullets were challenged. One bird was given a high-dose challenge (220,000 oocysts) and the other was given a low-dose challenge (13,000 oocysts) of mixed Eimeria species as found in Coccivac-D via oral gavage. The number of parasites in the challenge dose was established by 2 separate prechallenge titration experiments on age- and strain-matched, immunologically naïve birds that maximized lesion score values and oocyst excretion without causing mortalities (highdose challenge = lesion scores) or epithelial cell crowding (low-dose challenge = oocyst output; data not shown) [32]. After oral gavage, the pullets were randomly allocated to individual cages. The pullets not selected for challenge were housed in grower cages at a density of 8 birds per cage (387 cm 2 /bird) from d 42 to 84 and then 6 birds per cage (516 cm 2 /bird) from d 84 until the end of rearing, after which they were placed into layer production at the Arkell Research Station as per standard commercial protocol. At 6 d postchallenge infection, the challenged birds were reweighed individually (postchallenge BW). After reweighing, the high-dose challenged birds were killed humanely by cervical dislocation [33]. The low-dose challenged birds were returned to their assigned cages for an additional 6 d (i.e., until 12 d postchallenge infection) and then removed from the experiment. The outcomes of interest for the highdose challenge birds were postchallenge BW (controlling for prechallenge BW) and coccidia-induced lesions of the intestinal tract, which were assigned a value of 0 to 4 using the scoring system of Johnson and Reid [34]. The individual performing the lesion scoring was blinded, as they received the isolated intestinal tract and a tag number that was not indicative of treatment. The outcomes of interest for the low-dose challenge birds were postchallenge BW (controlling for prechallenge BW) and total oocyst output values (discussed further herein). In total, for each challenge dose, 112 pullets (4 treatments 28 cages per treatment) were included in this phase of the experiment at each challenge time (i.e., postbrood and midgrowth). Oocyst Output Fecal samples were stored at 4 C. For experiment 1, and for the treatment phase of experiment 2 from 6 to 27 DPI, OPG counts were determined. To do so, the number of oocysts in a fecal sample was determined by the McMaster counting chamber technique using saturated NaCl as the flotation medium [35]. Each sample was counted twice and the 2 counts were averaged to provide a single mean count; the mean count was then divided by the fecal weight in grams to calculate the OPG for the sample. For the treatment phase of experiment 2, from 30 to 60 DPI, oocyst output was determined by the sedimentation-floatation procedure, with NaCl as the flotation medium [35]. During the postbrood challenge (commencing at 42 d of age), total oocyst output was measured individually from each bird given the low-dose challenge from 6 to 12 days postchallenge infection through sequential 24-h collections. The total number of oocysts in each sample was determined using a McMaster counting chamber technique with saturated NaCl as the flotation medium [35]. For the midgrowth challenge, samples were collected as described previously and then pooled within a treatment group before determining total oocyst counts for each pooled sample. Statistical Analyses The ANOVA program within the statistical software SAS 9.2 [36] was used to compare the mean welfare scores, lesion scores, postchallenge BW (controlling for prechallenge BW), and total fecal oocyst counts between treatment groups as per Price et al. [18]. Random effects were used to control for potential clustering (room, cage, and replication). For total fecal oocyst counts, Duncan s tests were used to make pair-wise comparisons between treatment groups; for postchallenge BW, Tukey-Kramer tests were used. For mean welfare scores as well as lesion scores, t-values were used to make pairwise comparisons between treatment groups. For all tests, P < 0.05 was deemed significant. RESULTS AND DISCUSSION Live Eimeria vaccines are critically dependent on environmental fecal-oral cycling of vac-

6 528 JAPR: Research Report cine organisms for development of protective immunity. When live coccidiosis vaccines are administered to a flock in which insufficient lowlevel fecal-oral cycling occurs in the barn, complete protective immunity against a mixed Eimeria species infection is unlikely to develop [37]. Two main factors are needed for a live vaccine to elicit complete protective immunity against a mixed Eimeria species challenge infection. The first is vaccine administration, which must be a synchronous, uniform exposure to a small controlled dose of vaccine oocysts; second, optimal environmental conditions in the barn must exist, cycling with the minimum numbers of infective vaccine oocysts [18]. Environmental factors that could limit oocyst infectivity, survivability, and transmission dynamics (such as temperature, RH, oxygen access, and bird density) can affect live vaccination success [38 45]. Experiment 1 The use of spray vaccination spraying chicks with a premeasured volume of a coarse spray has been the administration method of choice for live vaccines [15]. Despite the inoculum being calibrated to give a specific number of oocysts per chick, this method relies on direct and indirect (preening) ingestion of the proper dose of parasites [13, 46]. Consequently, an inherent risk exists for natural variation in vaccine oocyst ingestion. Directly measuring the oocysts ingested while accounting for those parasites that will infect the host is a complex process; thus, in experiment 1, vaccine oocyst ingestion was indirectly measured by determining the number of OPG of feces at 6 DPI, when birds would be shedding oocysts of most Eimeria species as a result of the mixed-species vaccine inoculum [47, 48]. For each of the 5 communal cages in the current study, some chicks did not produce enough wet fecal material ( 1 g) during the 2-h collection period for the OPG counting technique to be applied. Fecal samples from 59 chicks were counted; the oocyst output ranged from 0 to 439,013 OPG. The frequency distribution of the number of OPG of feces per chick is shown in Figure 2. The mean number of OPG of feces from 59 chicks was 35,065.4 ± 8,105.6 (mean ± SE). Oocyst shedding was not uniform: 6 chicks shed no detectable oocysts, 17 chicks shed 10,000 OPG, and, at the other extreme, 3 chicks shed >100,000 OPG, with a single chick shedding more than 10 times the mean output of all birds. Despite the range in oocyst output when birds are spray-vaccinated, studies have found that susceptible naïve birds on either litter [49, 50] or in a cage environment [51] can ingest oocysts from an infected cage or penmate and, under controlled management conditions, become protected against challenge infections. Experiment 2 Three factors are needed for successful oocyst sporulation: (1) appropriate temperature (4 37 C; optimal at 29 C), (2) oxygen access, and (3) adequate RH [43, 47]. The temperature inside most poultry barns is usually close to the optimal oocyst sporulation temperature depending on the age of the birds being housed [25]. The standard bird density [15] and natural structure of the cages would provide the oxygen access needed for the parasite to become infective. The average temperatures within the rooms for the current study were in accordance with the Lohmann LSL management guidelines [25]. However, the average RH levels in the rooms (Figure 3) during the first 2 wk of the treatment phase were consistently low (e.g., 19, 16, and 13% RH at 6, 9, and 12 DPI, respectively). The Lohmann LSL management guide recommends an RH in the poultry house of 60 and 70% [25]; however, anecdotally, the RH levels in the barn are dependent on local weather conditions, barn set-up (e.g., cages vs. litter), and barn ventilation. As a result, seasonal and geographical variances can affect oocyst sporulation [41, 42], survivability [38, 40], and parasite transmission [52 54] markedly. During the treatment stage, the mean oocyst output for each treatment group expressed as OPG of feces are recorded in Table 1. For each treatment group, the average OPG of feces decreased from 6 to 15 DPI and the oocyst output was below the limit of detection for the technique from 18 to 60 DPI. The average BW of the birds from each treatment group matched expected average weights for Lohmann LSL pullets at various ages (1, 31, 72, and 105 d

7 Price et al.: HUMIDITY AND EIMERIA VACCINES 529 Figure 2. The frequency distribution of the number of oocysts per gram of feces measured 6 d postinoculation for layer chicks inoculated with Coccivac-D [21] via the spray cabinet technique at 1 d of age for experiment 1. Color version available in the online PDF. of age), as described in the breed management guide [25]. In general, the scores for each welfare parameter were minor at 42 and 84 d of age. However, significant differences between treatment groups were identified (Tables 2 and 3). At 42 d of age, pullets reared with increasing CFC percentage had significantly higher mean plumage cleanliness scores, indicating that feathers were dirtier as the percentage CFC increased from 0 to 60%. Pullets reared with 60% CFC had significantly higher foot pad dermatitis and hock burn scores than pullets reared with 0 or 20% CFC. Conversely, bumble foot scores of pullets reared with any cage floor coverage were significantly lower than pullets reared with no CFC. At 84 d of age, pullets reared with 60% CFC had significantly dirtier feathers (higher plumage cleanliness scores) than pullets reared with 0, 20, or 40% CFC. Pullets reared with 60% CFC had significantly higher foot pad dermatitis scores than pullets reared with 0, 20, or 40% CFC. No significant differences were observed between treatment groups for hock burn and bumble foot scores. Previously, a small-scale trial (a total of 240 pullets) was conducted using a low-dose Coccivac-D inoculum, in which age- and strainmatched pullets were reared on the same CFC configurations [18] as in the present study. Birds reared on 40% CFC, when given a homologous challenge at 6 wk of age, demonstrated a significantly lower total oocyst output per bird compared with the other treatment groups [18]. The results of the present study varied greatly compared with the previous trial, despite the similar experimental design, in that the total mean oocyst output data for the low-dose postbrood challenge demonstrated no significant differences between treatment groups in mean oocyst output at the postbrood or midgrowth periods. One variable that differed considerably between these studies was the atmospheric RH in the study barns, especially immediately after vaccination. In the present study, RH remained below 20% (and reached a low of 12.5% at 12 DPI)

8 530 JAPR: Research Report Table 1. The mean number of oocysts per gram of feces (OPG) from 6 to 60 d postinoculation (DPI) for Lohmann LSL Lite layer pullets spray-vaccinated with Coccivac-D [21] at 1 d of age and housed in conventional brooder cages (20 birds/cage) with different percentages of the cage floor covered with fiber trays until 6 wk of age and then in conventional grower cages (8 birds/cage; n = 28 cages/treatment group) Treatment (% cage floor coverage) 6 DPI 9 DPI 12 DPI 15 DPI DPI DPI 0 218,210 13, TFTC 1 NFF ,468 14, TFTC NFF 40 82,695 19, TFTC NFF ,705 31, TFTC NFF 1 TFTC = too few to count. 2 NFF = negative fecal float. during the first 2 wk, whereas RH was approximately 30% during the same period in the previous study [55]. The present study was conducted in the winter (outside temperatures below 0 C; Figure 3), whereas the previous study was conducted in the summer (outside temperatures between 12 and 25 C; recorded at a local Environment Canada weather station in Toronto). The low RH immediately following vaccination in the present study was likely responsible for the observed poor vaccine oocyst cycling during the treatment phase (Table 1). The low Figure 3. The mean temperature ( C) inside the barn room (long dashed line), mean RH (%) inside the barn room (solid line), mean outside temperature ( C; dotted line), and mean outside dew point ( C; short dashed line) recorded from an Environmental Canada (Toronto) weather station 4 km from the Arkell Research Station (University of Guelph, Guelph, Ontario, Canada) over days postinoculation during the treatment phase.

9 Price et al.: HUMIDITY AND EIMERIA VACCINES 531 Table 2. The least squares means scores (± SE) of welfare parameters at 42 d of age for Lohmann LSL Lite layer pullets spray-vaccinated with Coccivac-D [21] at 1 d of age and housed in conventional brooder cages (20 birds/ cage) with different percentages of the cage floor covered with fiber trays Treatment 1 (% cage floor coverage) Plumage cleanliness (score 0 3) Foot pad dermatitis (score 0 4) Hock burn (score 0 4) Bumble foot (score 0 2) ± 0.0 2A 0 ± 0 A 0 ± 0 A 0.3 ± 0.1 A ± 0.1 B ± 0.0 2A ± 0.0 2A 0.1 ± 0.0 2B ± 0.1 C 0.1 ± 0.0 2AB ± 0.0 2AB ± 0.0 2B ± 0.1 D 0.2 ± 0.1 B 0.1 ± 0.0 2B 0 ± 0 B A D Groups displaying the same letters within a column do not differ significantly (P > 0.05); for the purpose of statistical analyses for ANOVA values (capital letters), the difference in severity between a score of 0 and 1 was assumed to be the same as the difference between a score of 1 and 2 and so on [36]. 1 Twenty-eight birds per treatment 2 replicates indicates a nonzero number that rounds to 0 at 1 decimal place. oocyst output observed during the treatment phase following the initial shedding of vaccine oocysts (i.e., from 12 DPI onward) strongly suggests that in-cage cycling of the oocysts was not sufficient to elicit complete protective immunity in the vaccinated birds. Although no significant differences were seen between treatment groups for mean lesion scores at the postbrood or midgrowth periods among high-dose challenged pullets, some differences in BW between treatments were noted during the postbrood and midgrowth periods. Postchallenge BW for the postbrood challenge are shown in Table 4. For the low-dose challenge, no significant differences were observed between treatment groups in mean postchallenge BW after controlling for the prechallenge BW. For the high-dose challenge, the mean postchallenge BW of pullets reared with 40% CFC was significantly higher than pullets reared with 60% CFC after controlling for the prechallenge BW. Postchallenge BW for the midgrowth challenge is shown in Table 5. For both challenge doses, the mean postchallenge BW was lower than the prechallenge BW for all treatment groups (i.e., the pullets lost weight after challenge infection at 84 d). For the low-dose challenge, the mean postchallenge BW of pullets reared with 60% CFC was significantly higher than pullets reared with 0% CFC after controlling for the prechallenge BW. Similarly, for the high-dose challenge, the mean postchallenge BW of pullets reared with 60% CFC was significantly higher than for pullets reared with 0% CFC after controlling for the prechallenge BW. The differences in BW during the challenge periods showed some evidence of limited protective immunity. For example, depending on the challenge dose and period, the mean postchallenge BW was significantly higher for pullets Table 3. The least squares means scores (± SE) of welfare parameters at 84 d of age for Lohmann LSL Lite layer pullets spray-vaccinated with Coccivac-D [21] at 1 d of age and housed in conventional brooder cages (20 birds per cage) with different percentages of the cage floor covered with fiber trays until 6 wk of age Treatment 1 (% cage floor coverage) Plumage cleanliness (score 0 3) Foot pad dermatitis (score 0 4) Hock burn (score 0 4) Bumble foot (score 0 2) ± 0.0 2A 0 ± 0 A 0 ± 0 A 0 ± 0 A ± 0.0 2A 0 ± 0 A 0 ± 0 A 0 ± 0 A ± 0.0 2A 0 ± 0 A 0 ± 0 A 0 ± 0 A ± 0.1 B 0.2 ± 0.1 B 0 ± 0 A 0 ± 0 A A,B Groups displaying the same letters within a column do not differ significantly (P > 0.05); for the purpose of statistical analyses for ANOVA values (capital letters), the difference in severity between a score of 0 and 1 was assumed to be the same as the difference between a score of 1 and 2 and so on [36]. 1 Twenty-eight birds per treatment 2 replicates indicates a nonzero number that rounds to 0 at 1 decimal place.

10 532 JAPR: Research Report Table 4. The mean pre- and postchallenge BW (± SE) for the postbrood challenge at 42 d of age for the low- and high-dose challenge infected Lohmann LSL Lite layer pullets spray-vaccinated with Coccivac-D [21] at 1 d of age and housed in conventional brooder cages (20 birds/cage) with different percentages of the cage floor covered with fiber trays Treatment (% cage floor coverage) Low-dose challenge 1 (28 birds per treatment) Prechallenge weight (g) Unadjusted postchallenge weight (g) High-dose challenge 2 (28 birds per treatment) Prechallenge weight (g) Unadjusted postchallenge weight (g) ± ± 6.3 A ± ± 6.6 AB ± ± 6.4 A ± ± 6.6 AB ± ± 6.4 A ± ± 6.8 B ± ± 6.2 A ± ± 6.8 AC A C Groups displaying the same letters within a column do not differ significantly (P > 0.05) after controlling for the mean prechallenge BW. 1 Pullets given a low-dose challenge were infected with 13,000 mixed oocysts. 2 Pullets given a high-dose challenge were infected with 220,000 mixed oocysts. reared with 60% CFC than some other treatment groups, suggesting that limited protective immunity had been elicited in this treatment group. The overall mean number of OPG of feces during the treatment phase for pullets reared with 60% CFC was higher, albeit slightly and not significantly, than the other treatment groups. This suggestion of somewhat more successful cycling in the 60% CFC cages might help explain the observed partial protection against homologous challenge in these birds. However, following the postbrood or midgrowth challenges, oocyst output and lesion scores for the 60% CFC treatment group did not differ numerically from the other treatment groups. The strikingly different results obtained in the present study compared with previous observations in a technically similar experiment [18] highlight the essential role of RH in the success of live coccidiosis vaccination, particularly during the crucial first few weeks after vaccination. Litter moisture content and atmospheric RH that maximize oocyst sporulation success have not been established. If RH in the barn is not automatically regulated by the ventilation system, the temperature differential between the inside temperature of the barn and the outside air temperature can affect the RH inside the barn dramatically. If the inside barn temperature is high and the outside temperature is low (as was experienced over d 6 to 12 DPI; Figure 3) maintaining an RH around 35% is difficult. Heating outside air during the freezing Ontario (Canada) winter to supply to a brooder barn, especially one fitted with cages, will almost invariably provide an extremely dry environment. Eimeria species may Table 5. The mean pre- and postchallenge BW (± SE) for the postgrowth challenge at 84 d of age for the low- and high-dose challenge infected layer Lohmann LSL Lite pullets spray-vaccinated with Coccivac-D [21] at 1 d of age and housed in conventional brooder cages (20 birds/cage) with different percentages of the cage floor covered with fiber trays until 6 wk of age Treatment (% cage floor coverage) Low-dose challenge 1 (28 birds per treatment) Prechallenge weight (g) Unadjusted postchallenge weight (g) High-dose challenge 2 (28 birds per treatment) Prechallenge weight (g) Unadjusted postchallenge weight (g) ± ± 10.1 A ± ± 10.1 A ± ± 10.5 AB ± ± 10.5 AB ± ± 10.3 AB ± ± 10.3 AB ± ± 10.3 B ± ± 10.3 B A,B Groups displaying the same letters within a column do not differ significantly (P > 0.05) after controlling for the mean prechallenge BW. 1 Pullets given a low-dose challenge were infected with 13,000 mixed oocysts. 2 Pullets given a high-dose challenge were infected with 220,000 mixed oocysts.

11 Price et al.: HUMIDITY AND EIMERIA VACCINES 533 respond differently to various moisture levels [41, 42]. Graat et al. [42] examined the sporulation of E. acervulina oocysts in different litter conditions (dry, damp, and pure feces) at high or low temperatures (21 or 33 C) and RH (40 or 80%). Litter conditions were found to have the greatest effect on maximum sporulation percentages, whereas temperature and RH affected time and speed of sporulation [42]. Oocysts in dry or damp litter had a higher success rate of completing sporulation compared with oocysts in pure feces, regardless of the temperature or humidity level. However, high RH (80%) and temperature (33 C) were associated with the faster onset of sporulation [42], even though the ultimate success rate of completing sporulation was not affected. Conversely, Waldenstedt et al. [41] found that sporulation of E. maxima occurred most efficiently with low (16%) litter moisture content as opposed to wetter (62%) litter moisture conditions. In the latter study, RH and temperature were constant for all tested samples (75% and 23 C, respectively), suggesting that the litter moisture may have been affecting sporulation through means other than available moisture, such as available oxygen or increased ammonia production in wetter litter [41]. Lack of available oxygen may have decreased the observed success of sporulation of E. acervulina in pure feces in the study by Graat et al. [42]. Neither study [41, 42] examined viability or survivability of the sporulated oocysts (this can only be established through in vivo infection), where the number of surviving oocysts may be higher with higher moisture levels [38, 40]. Marquardt et al. [43] studied the effect of RH in correlation with temperature on the sporulation of Eimeria zuernii, a parasite of cattle. Unsporulated E. zuernii oocysts sporulated well at 75% RH and the percentage of sporulation and survival decreased as RH levels decreased from 25 to 13%; additionally, unsporulated oocysts appeared desiccated at RH levels below 13% [43]. However, maximum sporulation may be of less importance to infection in a commercial flock than the onset of sporulation because the latter may determine how quickly birds are infected [42]. Moreover, Williams [44] found that after 3 d on litter (feces and wood shavings) without cycling, all E. acervulina oocysts that had been shed completed sporulation, yet 30% appeared damaged regardless of the optimal temperature and moisture levels. Consequently, faster sporulation and longer survival in the environment will increase the likelihood that an oocyst will become infective and available for ingestion to contribute to parasite transmission. Conventional poultry barns require a ventilation system that will allow the influx of fresh air from outside and efflux of used air from inside the barn [56]. Consequently, the outside air must be heated or cooled as it enters the barn, which can affect RH inside the barn dramatically. A high barn temperature (e.g., as required during early brooding) combined with below-freezing outside air temperatures (e.g., during winter) can result in low RH inside the barn, particularly for newly placed chicks; for example, heating saturated 10 C air (i.e., 100% RH, 10 C dew point) to 32 C will produce an RH of 16% in the resulting heated air ( if no additional moisture is added. Under such low RH conditions, bird health can be affected negatively [57]. The successful sporulation and subsequent cycling of Eimeria species in live coccidiosis vaccines are likely to be greatly reduced, or eliminated entirely, at such low RH. The resultant minimal postvaccination oocyst cycling may only elicit partial protective immunity, as observed in the present study. Maintaining appropriate RH in the barn environment during live vaccination against coccidiosis is likely necessary for success. CONCLUSIONS AND APPLICATIONS 1. Successful sporulation and subsequent cycling of Eimeria species in live vaccines are partially dependent on RH levels in the barn in addition to temperature, oxygen access, and the pullet s access to infective oocysts. 2. Measuring RH after live vaccination of a flock, regardless of the rearing environment, may be an excellent way of predicting the relative success of Eimeria species oocyst sporulation in the barn and therefore, indirectly, the likely success of the live vaccination process. 3. Practical knowledge of factors influencing the exogenous, in-the-barn portion of the Eimeria life cycle during the

12 534 JAPR: Research Report vaccination process, combined with the ability to monitor and influence housing conditions, such as temperature, RH, or access to cycling parasites, will encourage live Eimeria vaccine success. REFERENCES AND NOTES 1. Reid, W. M History of avian medicine in the United States. X. Control of coccidiosis. Avian Dis. 34: Dalloul, R. A Poultry coccidiosis: Recent advancements in control measures and vaccine development. Expert Rev. Vaccines 5: Kitandu, A., and R. Juranova Progress in control measures for chicken coccidiosis. Acta Vet. (Brno) 75: Lee, J. T., C. Broussard, S. Fitz-Coy, P. Burke, N. H. Eckert, S. M. Stevens, P. N. Anderson, S. M. Anderson, and D. J. Caldwell Evaluation of live oocyst vaccination or salinomycin for control of field-strain Eimeria challenge in broilers on two different feeding programs. J. Appl. Poult. Res. 18: Sharman, P. A., N. C. Smith, M. G. Wallach, and M. Katrib Chasing the golden egg: Vaccination against poultry coccidiosis. Parasite Immunol. 32: Dalloul, R. A., and H. S. Lillehoj Recent advances in immunomodulation and vaccination strategies against coccidiosis. Avian Dis. 49: Chapman, H. D Practical use of vaccines for the control of coccidiosis in the chicken. World s Poult. Sci. J. 56: Williams, R. B Fifty years of anticoccidial vaccines for poultry ( ). Avian Dis. 46: Bell, D. D Cage management for layers. Pages in Commercial Chicken Meat and Egg Production, 5th ed. D. D. Bell and W. D. Weaver, ed. Springer, New York NY. 10. McDougald, L. R., A. L. Fuller, and B. L. McMurray An outbreak of Eimeria necatrix coccidiosis in breeder pullets: Analysis of immediate and possible longterm effects on performance. Avian Dis. 34: Soares, R., T. Cosstick, and E. H. Lee Control of coccidiosis in caged egg layers: A paper plate vaccination method. J. Appl. Poult. Res. 13: Gingerich, E US table egg industry update October 2009 to October Pages in Proc. 114th Ann. Meet. US Anim. Health Assoc., Minneapolis, MN. Richardson Printing, Kansas City, MO. 13. Chapman, H. D., T. E. Cherry, H. D. Danforth, G. Richards, M. W. Shirley, and R. B. Williams Sustainable coccidiosis control in poultry production: The role of live vaccines. Int. J. Parasitol. 32: Price, K. R Use of live vaccines for coccidiosis control in replacement layer pullets. J. Appl. Poult. Res. 21: Williams, R. B Anticoccidial vaccines for broiler chickens: Pathways to success. Avian Pathol. 31: Chapman, H. D., B. Roberts, M. W. Shirley, and R. B. Williams Guidelines for evaluating the efficacy and safety of live anticoccidial vaccines, and obtaining approval for their use in chickens and turkeys. Avian Pathol. 34: Price, K., M. Petrik, J. R. Barta, and L. Newman Live-nonattenuated vaccination in 4-tiered cagereared pullets in Ontario: A comparative study of Eimeria oocyst cycling management. Page 42 in Proc. 10th Meet. Can. Anim. Health Lab. Netw., Ontario Vet. Coll., Univ. Guelph, Guelph, Ontario, Canada. 18. Price, K. R., M. T. Guerin, L. Newman, B. M. Hargis, and J. R. Barta Examination of a novel practical poultry management method to enhance the effect of live Eimeria vaccination for conventionally housed replacement layer pullets. Int. J. Poult. Sci. 12: Long, P. L., and B. J. Millard Eimeria: further studies on the immunization of young chickens kept in litter pens. Avian Pathol. 8: Archer s Poultry Farm Ltd., Brighton, Ontario, Canada. 21. Merck Animal Health, Summit, NJ. 22. Ketchum Manufacturing Inc., Brockville, Ontario, Canada. 23. Ford Dickison Inc., Mitchell, Ontario, Canada. 24. National Farm Animal Care Council Recommended code of practice for the care and handling of pullets, layers and spent fowl: Poultry (layers). National Farm Animal Care Council, Lacombe, Alberta, Canada. Accessed Feb. 2012, pdf. 25. Lohmann Tierzucht GmBh Layer management guide: Lohmann LSL Classic. In Lohmann Tierzucht GmBh. Cuxhaven, Germany. 26. Heaters, Controls & Sensors Ltd., London, Ontario, Canada. 27. Tennessen, T., L. Connor, A. M. de Passille, K. Stanford, M. von Keyerserlingk, and G. Griffin CCAC guidelines on: The care and use of farm animals in research, teaching and testing. Canadian Council on Animal Care, Ottawa, Ontario, Canada. Accessed Feb. 2012, ccac.ca/documents/standards/guidelines/farm_animals. pdf. 28. Velkers, F. C., A. Bouma, J. A. Stegeman, and M. C. de Jong Transmission of a live Eimeria acervulina vaccine strain and response to infection in vaccinated and contact-vaccinated broilers. Vaccine 30: Pactiv LLC, Lake Forest, IL. 30. Fisher Scientific Company, Ottawa, Ontario, Canada. 31. Welfare Quality Consortium Welfare Quality Assessment Protocol for Poultry. Welfare Quality Consortium, Lelystad, the Netherlands. 32. Williams, R. B Quantification of the crowding effect during infections with the seven Eimeria species of the domesticated fowl: Its importance for experimental designs and the production of oocyst stocks. Int. J. Parasitol. 31: Charbonneau, R., L. Niel, E. Olfert, M. von Keyerserlingk, and G. Griffin CCAC guidelines on: Euthanasia of animals used in science. Canadian Council on Animal Care, Ottawa, Ontario, Canada. Accessed Feb. 2012, pdf. 34. Johnson, J., and W. Reid Anticoccidial drugs: Lesion scoring techniques in battery and floor-pen experiments with chickens. Exp. Parasitol. 28:30 36.

13 Price et al.: HUMIDITY AND EIMERIA VACCINES Long, P. L., B. J. Millard, L. P. Joyner, and C. C. Norton A guide to laboratory techniques used in the study and diagnosis of avian coccidiosis. Folia Vet. Lat. 6: SAS SAS System for Windows. Version 9.2 ed. SAS Institute Inc., Cary, NC. 37. Joyner, L. P., and C. C. Norton The immunity arising from low-level infection with Eimeria maxima and Eimeria acervulina. Parasitology 72: Reyna, P. S., L. R. McDougald, and G. F. Mathis Survival of coccidia in poultry litter and reservoirs of infection. Avian Dis. 27: Edgar, S. A Effect of temperature on the sporulation of oocysts of the protozoan, Eimeria tenella. Trans. Am. Microsc. Soc. 73: Edgar, S. A Sporulation of oocysts at specific temerpatures and notes on the prepatent period of several species of avian coccidia. J. Parasitol. 41: Waldenstedt, L., K. Elwinger, A. Lunden, P. Thebo, and A. Uggla Sporulation of Eimeria maxima oocysts in litter with different moisture contents. Poult. Sci. 80: Graat, E. A., A. M. Henken, H. W. Ploeger, J. P. T. M. Noordhuizen, and M. H. Vertommen Rate and course of sporulation of oocysts of Eimeria acervulina under different environmental conditions. Parasitology 108: Marquardt, W. C., C. M. Senger, and L. Seghetti The effect of physical and chemical agents on the oocyst of Eimeria zurnii (protozoa, coccidia). J. Protozool. 7: Williams, R. B Epidemiological studies of coccidiosis in the domesticated fowl (Gallus gallus): II. Physical condition and survival of Eimeria acervulina oocysts in poultry-house litter. Appl. Parasitol. 36: Williams, R. B., J. D. Johnson, and S. J. Andrews Anticoccidial vaccination of broiler chickens in various managment programmes: Relationship between oocysts accumulation in litter and the development of protective immunity. Vet. Res. Commun. 24: Caldwell, D. Y., S. D. Young, D. J. Caldwell, R. W. Moore, and B. M. Hargis Interaction of color and photointensity on preening behavior and ingestion of sprayapplied biologics. J. Appl. Poult. Res. 10: Al-Badri, R., and J. R. Barta The kinetics of oocyst shedding and sporulation in two immunologically distinct strains of Eimeria maxima, GS and M6. Parasitol. Res. 111: Reid, W. M., and P. L. Long A diagnostic chart for nine species of fowl coccidia. Pages 5 24 in Georgia Agric. Exp. Stn. Tech. Bull. N. B. Bowen, ed. College of Agriculture, University of Georgia, Athens. 49. Velkers, F. C., A. Bouma, J. Arjan Stegeman, and M. C. de Jong Oocyst output and transmission rates during successive infections with Eimeria acervulina in experimental broiler flocks. Vet. Parasitol. 187: Velkers, F. C., A. Bouma, E. A. Graat, D. Klinkenberg, J. A. Stegeman, and M. C. de Jong Eimeria acervulina: The influence of inoculation dose on transmission between broiler chickens. Exp. Parasitol. 125: Price, K. R., J. Bulfon, and J. R. Barta Personal communication. University of Guelph, Guelph, Ontario, Canada. 52. Lal, A., M. G. Baker, S. Hales, and N. P. French Potential effects of global environmental changes on cryptosporidiosis and giardiasis transmission. Trends Parasitol. 29: Awais, M. M., M. Akhtar, Z. Iqbal, F. Muhammad, and M. I. Anwar Seasonal prevalence of coccidiosis in industrial broiler chickens in Faisalabad, Punjab, Pakistan. Trop. Anim. Health Prod. 44: Farr, M. M., and E. E. Wehr Survival of Eimeria acervulina, E. tenella, and E. maxima oocysts on soil under various field conditions. Ann. N. Y. Acad. Sci. 52: Price, K. R Personal observation. University of Guelph, Guelph, Ontario, Canada. 56. Scanes, C. G., G. Brant, and M. E. Ensminger Poultry Science, 4th ed. Pearson Education Inc., Upper Saddle River, NJ. 57. Reece, F. N., and B. D. Lott Heat and moisture production of broiler chickens during brooding. Poult. Sci. 61: Acknowledgments The technical assistance of J. Cobean, M. Hafeez, R. Al-Badri, H. Brown, M. DeVisser, C. Pinard, and A. Barta (University of Guelph), as well as the advice of M. Petrik (McKinley Hatchery, St. Marys, Ontario, Canada) is gratefully acknowledged. The advice of W. Sears (University of Guelph) on the statistical analyses is gratefully acknowledged. The donation of Coccivac-D (Merck Animal Health, Summit, NJ) for initial vaccination and subsequent challenge is acknowledged. This research was funded through grants from the Natural Sciences and Engineering Research Council of Canada (NSERC; Ottawa, Ontario, Canada) and the Ontario Ministry of Agriculture and Food (Guelph, Ontario, Canada) to J. R. Barta. K. R. Price obtained funding from a postgraduate scholarship from NSERC and an Ontario Graduate Scholarship, Ministry of Training, Colleges and Universities (Toronto, Ontario, Canada).