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All birds were maintained at the Experimental Unit PEAT of INRA (Nouzilly, France). The Experimental Unit is registered by the ministry of Agriculture with the license number B-37-175-1 for animal experimentation. All experiments were approved by the Ethic Committee in Animal Experimentation of Val de Loire CEEA Vdl (permit number 2011-02-8). The CEEA vdl is registered by the National Committe ‘Comité National de Réflexion Ethique sur l'Expérimentation Animale’ under the number 19. All experiments were performed in accordance with the European Communities Council Directive 2010/63/UE.
Forty 20-week old White leghorn hens (Gallus gallus domesticus) from the experimental unit PEAT (INRA, Nouzilly) were equally stratified into 2 groups. The groups were balanced for the hen mass and egg mass. The hens of the 2 groups were housed in 2 similar thermo-regulated rooms (40 m2). Each bird was placed in an individual wire home-pen (100×100×50 cm) (length × width × height) containing wood shavings on the floor, a nest, perch, drinker, and trough. In each room, birds had tactile, visual, and vocal contacts with the others. Water and food (calculated content: ME = 2800 kcal/kg; crude protein = 16%; lysine = 0.7%; calcium = 0.4%) were available ad libitum during a 14:10 h light:dark cycle. All the birds were maintained at a temperature of 21.0±0.5°C for a period of 2 weeks. After this period, the room temperature of the experimental group was gradually elevated to 30.0±0.5°C over 3 days. This temperature was maintained for the following 5 weeks. This temperature is considered as a moderate heat challenge for White leghorn hens. These birds had a physiological potential to maintain their laying rate under 30°C. Detrimental effects are commonly known to appear with temperatures above 32°C [32]. The control group was maintained at a temperature of 21±0.5°C (standard breeding temperature) throughout the experiment.
Each hen was weighted 5 times: once a week before the beginning of the treatment and once every week during the first 4 weeks of treatment. Rectal temperature of each bird was recorded twice: once before the treatment and once after 3 weeks of treatment; the temperature was measured using a digital thermometer (Testo, Forbach, France). The daily feed intake was measured once before the treatment and once during the first 3 weeks of treatment. The weight of each trough was measured after every 24 h to determine the daily intake of each hen. Plasma corticosterone levels were measured to evaluate the impact of the heat treatment on hens. Blood samples were collected twice for each hen: once the day before the beginning of the treatment and once after 3 weeks of treatment. Hen's wing was held motionless for 30 seconds in order to sample blood from the wing vein. At each sampling time, 2 mL of blood was obtained, collected in tubes containing EDTA (2 mg/mL) and kept on ice. For both sampling days, all animals were sampled between 9:30 and 11:00 am. The blood samples were then centrifuged at 2,000× g for 15 min at 4°C. The plasma was collected and stored at −20°C before the measurement at the Biological Center of Chizé (France). Corticosterone was extracted by adding 3 mL diethyl-ether to 100 µL of each plasma sample, followed by vortexing and centrifugation. The diethyl-ether phase containing the steroid was decanted and discarded after snap freezing the tube in an alcohol bath at 38°C. The organic solvent was then evaporated. The dried extracts were redissolved in 300 µL of phosphate buffer, and corticosterone concentration was assayed in duplicate. Next, 100 µL of the extract was incubated overnight with 5,000 cpm of the appropriate 3H-steroid (Perkin Elmer, US) and polyclonal rabbit corticosterone-21-thyroglobulin antiserum (Sigma, US). The bound fraction was then separated from the free fraction by adding dextran-coated charcoal and radioactivity of the antibody bound fraction was measured using a tri-carb 2810 TR scintillation counter (Perkin Elmer, US). Tests were performed to validate the corticosterone assay on plasma. Inter- and intra-assay variations were respectively 9.99% and 7.07%. The lowest detectable concentration of corticosterone was 0.14 ng/mL. Two samples were serially diluted in the assay buffer, and their displacement curves were parallel to the standard curve. The mean recovery of the standard spiked in a sample was 92%.
Since heat stress was shown to increase the tonic immobility duration in laying hens [33], which could indicate a state of stress in animals, all the hens were tested once before the heat period and after 3 weeks of treatment. This test followed a procedure similar to that described by Jones [34]. Animals were caught individually and carried to another room. Each hen was then placed on its back in a U-shaped wooden cradle and held by the experimenter with one hand over the sternum and one gently covering the bird's head. Each hen was restrained for 10 s and then released. When more than 10 s passed between release and the bird's escape, duration of tonic immobility was measured. If not, another induction attempt was conducted. A score of zero second was given when tonic immobility could not be attained after 5 induction attempts. The test was stopped, and a maximum duration of 300 s was allocated when the hen did not stand up within 300 s. The observer remained out of the hen's sight during the test. Duration of tonic immobility and number of induction attempts were recorded. The duration of the tonic immobility reaction is considered to be a standard and robust measure of fearfulness [34]. This manipulation induces a reversible catatonic state, the duration of which is positively correlated with the general underlying fearfulness [35], [36].
Eggs from all females were collected daily for 6 consecutive weeks; i.e., 1 week before the heat period and during the 5-week treatment. The laying rate was calculated as the number of laid eggs/female/day.
Incubation and yolk hormonal assays: Since the formation of individual yolks requires 21 days and the hens were progressively acclimated to heat, egg collection for incubation and hormonal assays was started on the 21st day after the beginning of the heat period [37]. Eggs were collected during 15 days in each group and stored at 17°C for incubation. The hens were fertilized by artificial insemination on the fifth and sixth days before egg collection and then once a week during two weeks. To that end, each hen was gently maintained approximately 30 s and 50 µL of a pure mixture of sperms, originating from 10 males, was deposited at the entrance of the cloacal vent with a pipette. Each female contributed on an average 4.27±0.36 eggs, and a total of 270 eggs were collected. Of the 270 eggs collected, 171 were fertile and maintained in the incubator (n = 90 control eggs; n = 81 experimental eggs). All eggs were placed in alternative rows on each shelf of the incubator. They were maintained at 37.8°C and 56% relative humidity and turned automatically and continuously. At day 14 of incubation, the eggs containing dead embryos were eliminated. Three days before hatching, the rotation was stopped, and the temperature was decreased to 37.6°C. Eggs were then placed in a grid constructed of a wire mesh and cardboard dividers so that chicks from both the sets could be identified. During the collection period, 2 eggs per female were used for hormonal assays and stored at −20°C. Eggs sampled for hormonal assay were weighted, and the frozen yolk was separated from the albumen. Eggshells were separated, dried for 24 h, and weighed. The weight of the albumen was determined by subtracting the weight of the eggshell and the yolk from that of the whole egg. The 2 eggs per female were used to determine the mean egg weight and mean relative proportion of each component (yolk, albumen, shell) for each female. Although most research on yolk hormones has focused on the effects of testosterone levels [38], concentrations of yolk progesterone and androstenedione might also play important roles in precocial birds [22], [39]. In addition, the light/dark cycle or housing conditions were found to modulate yolk estradiol levels in laying hens [28], [30]. Therefore, whether yolk testosterone, androstenedione, progesterone, and estradiol levels were modulated by maternal environment was investigated. The presence of corticosterone in egg yolk remains controversial [40] and was not found to be modulated by environmental conditions in laying hens [29], [30] nonetheless its potential implications were also investigated to provide a more complete picture of yolk content. One yolk sample per female was analyzed for corticosterone and estradiol concentrations (Biological Center of Chizé, France). Another yolk per female was analyzed for measuring immunoreactive progesterone, androstenedione, and testosterone concentrations (Veterinary University of Vienna, Austria).
Yolk corticosterone and estradiol radio-immunoassays: Yolk concentrations of corticosterone and estradiol were assayed using the same procedure. Briefly, 100 mg of each sample was homogenised in 1 mL of distilled water. Steroids were extracted by adding 3 mL of diethyl-ether to 300 µL of the mixture, followed by vortexing and centrifugation. The diethyl-ether phase containing steroids was decanted and discarded after snap freezing the tube in an alcohol bath at 38°C. This procedure was repeated twice for each yolk, and the organic solvents were then evaporated. The extracts were redissolved in 600 µL of phosphate buffer, and each hormone was assayed in duplicate. Next, 100 µL of the extract was incubated overnight with 5,000 cpm of the appropriate 3H-steroid (Perkin Elmer, US) and polyclonal rabbit antiserum. Anti-11-HS-corticosterone antiserum was supplied by P.A.R.I.S. (France) and anti-estradiol by Sigma (US). The bound fraction was then separated from the free fraction by adding dextran-coated charcoal, and the activity was counted on a tri-carb 2810 TR scintillation counter (Perkin Elmer, US). Tests were performed to validate both the hormone assays on egg yolk samples. Inter-assay and intra-assay variations for corticosterone and estradiol were 20.98% and 16.70 and 17.18% and 13.13% respectively. The lowest detectable concentrations of corticosterone and estradiol were 1.55 pg/mg, and 0.69 pg/mg respectively. Two yolk samples were serially diluted in the assay buffer, and their displacement curves were found to be parallel to the standard curve. The mean recoveries of the standard spiked sample for corticosterone and estradiol were 120% and 90% respectively.
Yolk progesterone, testosterone, and androstenedione assays: The concentrations of progesterone (P4) and androgens (androstenedione = A4 and testosterone = T) were assayed using a method similar to that described by Lipar et al. [41], Möstl et al. [42] and Hackl et al. [43]. Each yolk was cut in half. Because the distribution of hormones vary between egg layers [41], [42], the mixed half yolk was assayed. After thawing the yolk was mixed and 0.5 g of each yolk was diluted with 1.5 ml of water and vortexed for 30 sec. The emulsion was diluted with 8 ml methanol, vortexed for 30 min and stored overnight at minus 20 °C. After centrifugation (minus 10 °C 1500 g, 15 min) the supernatant was diluted with assay buffer (1+10) and used for enzyme immunoassays (EIAs) for measuring the concentrations of immunoreactive progesterone, testosterone and androstenedione [44]. Inter-assay coefficients of variation were 12.2%, 10.7% and 10.1% respectively. The intra-assay variation was 8.5%, 4.2% and 9.2%.
We kept 96 chicks (48 controls and 48 experimental), all hatched on the 21st day of incubation for the experiment. Each chick was identified with a numbered ring on its leg. The chicks were weighted, and their body temperature under the wing was recorded using an infrared thermometer (TES1326S). The temperature was also recorded at 6 days of age. Approximately 2 h after the chicks hatched, their quality was assessed by 2 experimenters. Of the 8 parameters completely described by Tona et al. [44], 6 were used in this study. Each chick received a quality score that varie from 0 (bad quality) to 40 (good quality) (Table 1). The chicks were weighed at hatching, and at 6, 13 and 20 days.
Table data removed from full text. Table identifier and caption: 10.1371/journal.pone.0057670.t001 Allocation of scores to morphological parameter observations (completely described by Tona et al. [41]). Between 17 and 25 weeks of age, female offspring were weighted and placed in individual cages to record the onset of laying and laying rate.
The 96 chicks were placed in pairs in plastic tubs measuring 50×40×40 cm with a wire top and a floor covered by wood shavings; the chicks were divided in 2 groups. Each group consisted of 24 pairs of chicks: 24 pairs of control chicks (C21) and 24 pairs of experimental chicks (C30). The pairs of chicks were randomly allocated to 2 rooms of equal size and maintained on a 10-h:14-h light:dark cycle; water was provided ad libitum. Chicks were fed ad libitum by using a conventional starter mash (Experimental Unit PEAT, INRA centre de Tours, France). The food was dispensed in 50-cm-long feeding troughs. The troughs were covered with a metallic roof having 12 circular holes (diameter, 5 cm); these holes allowed sufficient access for the chicks to the feed while avoiding food spillage. Two opaque drinking bottles (1 L) with pipettes were placed in each cage. Ambient temperature was maintained at 33.0±1.5 °C from hatching until the chicks were 8-days-old; subsequently, it was decreased progressively to 24±1 °C, until the chicks were 25-days-old. The sex of the animals was determined by observation of the comb and legs at 3 weeks of age. The control and experimental groups consisted of 26 females and 22 males, and 21 females and 27 males, respectively.
The daily intake was measured for each pair of chicks at 3, 11, and 17 days. The weight of each trough was recorded every 24 h to determine the food consumption by each pair. Food preference: the preference of chicks for their standard food (fat content: 6.6%) or an energetic food (mash insectivore; La Ferme de Manon, France, fat content: 13%) was tested when they were 10 days old. For four consecutive days before the test, all the pairs of chicks were exposed to 50 g of the energetic mash placed in a cup in addition to their regular food in order to acclimate them to the food to allow them to assimilate the nutritional value of the mash [45]. Because chicks are extremely distressed when isolated, the study was conducted in the pairs of chicks (n = 24 pairs per treatment). The test box, identical to the home box, was located in a different room. Each pair of chicks was placed in the test box after 1 h of food deprivation. They were transported in a 15×15×15 cm container, deposited in the center of the test box, and observed for 5 min. The test box contained 2 troughs identical to their familiar trough; 1 containing 100 g of the familiar food and another containing 100 g of the energetic food. The location of the troughs was balanced across the test trials. An observer (blind to the treatment), hidden behind a curtain with small observation windows, recorded the behavior of 1 focal bird of each pair. Focal birds were chosen randomly beforehand and identified by a colored mark on the head. The experimenter recorded the latencies to explore each type of food (the bird touch the food with its beack), the latencies to ingest each type of food and the time spent eating on each trough (the bird was considered be eating when the movements of the mandible, neck, and throat due to swallowing were observed). Because the food particles weigh very less, the quantity of food eaten could not be weighed with precision over such a short period. The behavior of chicks toward energetic item was also tested using a choice test between a control solution (600 mL water) and a solution (600 mL water) containing 5% sucrose placed in the home cage. A concentration of 5% sucrose is detectable by chicks and do not engender a preference or an avoidance compared to tap water [46]. The localization of the drinking bottle containing the sweet solution was counterbalanced between the cages. The liquid intake was recorded for each pair by weighing the bottles on a 24-h scale when the chicks were 21–22 days of age. Two pairs of C30 chicks and 3 pairs of C21 chicks drank exclusively from 1 out the 2 drinkers and were excluded from the analysis. Food neophobia: food neophobia (i.e. fear of novel foods) is particularly well described in birds [47] and can be influenced by maternal effects [48]. Food neophobia in offspring and habituation to a novel food were investigated using a protocol described by Bertin et al., [49]. Chicks were exposed twice to a novel food; once at 12 days of age and once at 19 days of age. After 1 h of food deprivation, pairs of chicks were exposed for 3 min to a familiar trough containing 100 g of millet seeds following the same protocol as the food preference test. An experimenter recorded the latency to ingest the novel food and the number of distress calls.
Analysis of the emotional reactivity: A tonic immobility test, as described for adult hens, was performed on all chicks when they were 7 days of age. The number of inductions and the tonic immobility duration were recorded. Open field test: the open-field test involves transferring the birds from a familiar home cage to an unfamiliar and open environment. Chicks were individually placed in the middle of an open cylindrical arena (diameter, 120 cm; height, 35 cm) on a linoleum floor for 5 min. To assess their locomotor activity, 2 perpendicular lines were drawn in the arena, dividing it into 4 equal parts. The latency of the first step, number of distress calls, and activity of the birds (number of times when a bird crossed a line) were recorded by an experimenter hidden behind a curtain with observation windows. Animal activity in an open-field test is considered inversely correlated with emotional reactivity. Quiet and inactive animals are commonly considered to have a higher level of emotional reactivity than active animals [50]. This test was performed when the chicks were 23–24 days of age.
Kolmogorov-Smirnov tests determined whether the data were normally distributed. In the hens, all the morpho-physiological and behavioral measures were analyzed using one-way analysis of variance (ANOVA) for repeated measures (treatment × time). When required, PLSD Fisher post-hoc tests were performed. The relative proportions of egg components (yolk and eggshell) were determined and analyzed using multivariate analysis of variance (MANOVA). If Wilks' Lambda tests showed a significant multivariate effect, individual one-way ANOVAs were performed for each dependent variable. Yolk hormone concentrations were analyzed using a MANOVA and individual one-way ANOVAs. Hatching success and the number of hatchlings obtaining the maximum morphological quality score were tested using a Chi-square test. The body temperature of chicks, the laying rate of the female offspring, and feed intake were tested using ANOVA for repeated measures with the treatment and time as factors. The mass of chicks was analyzed using ANOVA for repeated measures with the treatment and sex as factors. In the food choice test, a chi-square test was conducted on the number of animals choosing to first ingest the energetic food. T-tests were conducted within each group on the other variables (the same for the choice test with water). Inter-group comparisons were analyzed using ANOVA on latency scores (latency to eat the energetic food–latency to eat the standard food), and the total time spent eating during testing. The number of distress calls and the latencies to ingest in the novel food test were analyzed using ANOVA for repeated measures with treatment and sex as factors. Variables recorded in the tonic immobility and open-field tests were analyzed using ANOVAs. Data are presented as mean ± SEM. All analyses were performed using Statview software (SAS, Cary, NC), with significance accepted at P≤0.05.
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