Effect of Source and Level of Vitamin D on Immune Function in Growing Broilers 1

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1 2004 Poultry Science Association, Inc. Effect of Source and Level of Vitamin D on Immune Function in Growing Broilers 1 C. A. Fritts, G. F. Erf, T. K. Bersi, and P. W. Waldroup 2 Department of Poultry Science, University of Arkansas, Fayetteville, Arkansas Primary Audience: Nutritionists, Veterinarians, Production Managers, Researchers SUMMARY Cholecalciferol and especially 1,25-dihydroxycholecalciferol have been reported to have immunomodulatory effects in various mammals. A study was conducted to evaluate source and level of vitamin D on various aspects of innate and adaptive immunity in broiler chicks. Nutritionally adequate starter (0 to 21 d) and grower (21 to 42 d) diets were fortified with either cholecalciferol (VIT-D3) or 25-hydroxycholecalciferol (25-OH-D3) to provide 125, 250, 500, 1,000, 2,000, or 4,000 IU/kg, based on the conversion of µg to 1 IU. Male birds of a commercial broiler strain were grown in litter floor pens in a house of commercial design with curtain sidewalls. Four pens of 60 birds were assigned to each dietary treatment. Various measures of innate and acquired immunity were conducted. No significant differences were observed related to source or level of vitamin D on macrophage function at 21 d and cutaneous basophil hypersensitivity at 35 d. Significant differences in concentration and proportion among white blood cells were observed but followed no consistent pattern. Feeding levels of 2,000 or 4,000 IU of vitamin D, as commonly fed in the poultry industry, did not positively or negatively affect the immune system within the parameters measured. Key words: immune function, cholecalciferol, 25-hydroxycholecalciferol, broiler 2004 J. Appl. Poult. Res. 13: INTRODUCTION Previous work has demonstrated that, besides its effects on calcium and bone metabolism, cholecalciferol and especially 1,25-dihydroxycholecalciferol, the active form of vitamin D, possesses pronounced immunomodulatory effects in various mammals [1, 2, 3, 4, 5, 6, 7, 8, 9, 10, 11, 12]. Research comparing vitamin D 3 and 1,25-(OH) 2 -D 3 in humans and mice has shown that the use of 1,25-(OH) 2 -D 3 may help treat cancer, skin and immune disorders, and help prevent graft rejection [13]. The 1,25- (OH) 2 -D 3 metabolite is also instrumental in inhibiting interleukin (IL)-2 and interferon (IFN)- γ synthesis in humans and mice, thus suppressing cell-mediated immunoactivity [14]. Diets with adequate 1,25-(OH) 2 -D 3 supplementation were shown to prevent the progression of arthritis in mice compared with untreated controls [15]. Several studies have reported that the immune system of the chick, specifically monocyte and macrophage function, is negatively altered 1 Published with approval of the Director, Arkansas Agricultural Experiment Station, Fayetteville, AR Mention of a trade name, proprietary product, or specific equipment does not constitute a guarantee or warranty by the University of Arkansas and does not imply its approval to the exclusion of other products that may be suitable. 2 To whom correspondence should be addressed: waldroup@uark.edu.

2 264 when diets are deficient in vitamin D [16, 17, 18]. Huff et al. [19] reported that supplementing turkey diets with vitamin D 3 improved body weight and increased disease resistance, when turkeys were repeatedly injected with dexamethasone and challenged with Escherichia coli. Moreover, additional dietary vitamin D 3 supplementation completely prevented the dexamethasone-induced rise in the heterophil to lymphocyte ratio. However, no changes were observed in control nonchallenged birds by vitamin D 3 supplementation. Vitamin D has long been provided to poultry by supplementation of the diet with crystalline forms of cholecalciferol (VIT-D3). This metabolite must undergo additional changes in the liver to become 25-hydroxycholecalciferol (25-OH- D3) and in the kidney to become 1,25-dihydroxycholecalciferol, considered as the active metabolite [20]. In 1995, 25-OH-D3 was given generally recognized as safe status for use as a vitamin D source in poultry diets [21]. With comparable levels of vitamin D 3, this isomer has been shown to improve body weight gain, feed efficiency, bone ash, and breast meat yield and to reduce the incidence of tibial dyschondroplasia and rickets in broilers [22, 23, 24, 25, 26, 27]. However, little research has been conducted comparing the effects of these 2 sources of vitamin D on development and function of the immune system in broilers. The objective of the present study was to compare the effects of cholecalciferol and 25-OH-D3 at levels ranging from slightly deficient to those commonly used in commercial broiler diets on aspects of innate and adaptive immunity. The effect of these dietary treatments on live performance and bone characteristics has been previously reported [28]. MATERIALS AND METHODS Experimental Diets and Birds The University of Arkansas Animal Care Committee approved all procedures used during the study. Corn-soybean meal based-diets were formulated for starter (0 to 21 d) and grower (21 to 42 d) periods, containing the minimum crude protein content suggested by NRC (1994) with a minimum of 110% of the suggested amino acid levels. Calcium and nonphytate phosphorus JAPR: Research Report were provided at NRC [29] suggested levels. All diets were fortified with a vitamin premix that provided adequate amounts of all vitamins except for vitamin D. A complete trace mineral mix provided all minerals in sulfate form. Composition of starter and grower diets is shown in Table 1. Using aliquots of a common mix of starter or grower diets, 12 experimental diets were prepared using a 2 6 factorial arrangement of treatments. A commercially available [30] VIT- D3 product was used to provide 125, 250, 500, 1,000, 2,000, or 4,000 IU/kg vitamin D. A commercial source [31] of 25-hydroxycholecalciferol (25-OH-D3) was used to provide 3.125, 6.25, 12.5, 25, 50, or 100 µg/kg, calculated to be equal to the levels provided by the VIT-D3, based on the conversion of µg to 1 ICU [29]. Respective sources of vitamin D activity were blended with a portion of the basal diet prior to adding to the mixer to enhance distribution in the diet. All diets were pelleted with steam; starter diets were crumbled. Each of the 12 experimental treatments was assigned to 4 replicate pens of 60 male broilers. Samples of mixed feeds were retained for analysis of crude protein, calcium, phosphorus, and vitamin D activity. One-day-old male chicks of a commercial broiler strain [32], originating from a breeder flock that was fed VIT-D3 as a source of vitamin D, were obtained from a local hatchery where they had been vaccinated in ovo for Marek s virus and had received vaccinations for Newcastle disease and infectious bronchitis posthatch via a coarse spray. Birds were randomly assigned to pens in a steel truss poultry house of commercial design. The house had a 1-m sidewall curtain with 2 outside rows of 12 pens each and 2 inside rows of 12 pens each. The inside and outside rows served as blocks in the experimental design. Sixty chicks were randomly allocated to each of 48 pens (5.2 m 2 ). Previously used litter, top-dressed with new softwood shavings, served as bedding over concrete floors. Each pen was equipped with 1 automatic water fount and 2 tube-type feeders. Birds were provided ad libitum access to feed and water during the study with 23 h of light and 1 h of darkness. One 9-W fluorescent light, suspended 198 cm

3 FRITTS ET AL.: VITAMIN D SOURCE AND IMMUNE FUNCTION 265 over the litter, provided supplemental light in each pen. Immune System Measurements Macrophage function (nitric oxide production and cytotoxicity) was examined to determine the effects of dietary vitamin D on aspects of innate immunity. At 3 wk of age, 3 randomly selected birds per pen were injected in the abdominal cavity with Sephadex G-50 [33] to elicit abdominal exudate cells (AEC) as described by Qureshi and Miller [34] and adapted as outlined in References and Notes [47]. To determine AEC nitric oxide production in macrophage culture supernatants, samples were assayed as described by Green et al. [36] and outlined in References and Notes [48]. The average of 4 measurements per sample was used in the final analysis. A standard curve describing the relationship between nitrite concentration and absorbance units (a.u.) of the samples was generated using various concentrations of sodium nitrite dissolved in LM Hahn medium. Nitrite concentration of the culture supernatants, and hence nitric oxide (NO) concentration, were determined using the samples a.u. and the equation of the standard curve. To measure cytotoxic killing activity in AEC culture supernatants, the RP9 tumor cell assay was used [34] as described in References and Notes [49]. The average of 4 measurements per sample was used in the final analysis. Percent cytoxicity was calculated with the following equation: (a.u. negative control) (a.u. test sample) (a.u. negative control) (a.u. positive control) 100 = % cytotoxicity where a.u. negative control = a.u. of RP9 cells cultured in LM Hahn medium (no killing); a.u. positive control = a.u. of RP9 cells cultured in LM Hahn medium with Triton-X (100% killing); a.u. sample = a.u. of RP9 cells cultured in LM Hahn medium plus supernatant from 24-h AEC cultures (with or without lipopolysaccharide LPS). Cutaneous basophil hypersensitivity (CBH) response to phytohemagglutinin-p (PHA-P) was used to assess in vivo cell-mediated immune activity [38, 39, 40, 41, 42]. At 35 d, 8 birds per treatment received 100 µg PHA-P in 0.1 ml saline intradermally in the left wattle and an equal volume of PBS in the right wattle. Wattle thickness was measured before injection and 4, 8, 12, 20 and 24 h postinjection using a digital caliper (in 0.1 mm). The swelling response was determined by subtracting the preinjection thickness from the postinjection response for each wattle. To assess the concentrations of, and proportions among blood leukocytes at 6 wk of age, heparinized blood was collected from 12 birds per treatment. The concentrations of red blood cells, total white blood cells, and thrombocytes were determined using a Cell-Dyne 3500 System [43] automated hematology analyzer and blood smears as described in References and Notes [50]. For each blood smear, 300 leukocytes (lymphocyte, heterophil, monocyte, eosinophil, basophil) were identified in each slide at 1,000 magnification [44]. The proportions among differential leukocytes were expressed as the percentage of each blood leukocytes in a total of 300 cells. The concentration of differential leukocytes was calculated based on the estimated concentration of white blood cells using the automated hematology analyzer and the proportion of a leukocyte estimated by the manual method. The combination of manual and automated estimates to determine differential leukocyte concentrations has been found in our laboratory to be a more reliable estimate than differential leukocyte concentrations determined by the automated system. Statistical Analysis Individual birds were the experimental unit for immunological assays. Data were subjected to ANOVA as a factorial arrangement of treatments with vitamin D source and level and the interaction between them as the source of variation using the GLM procedure [45]. Significant differences among or between means were separated by repeated t-tests using the least squares means option of SAS software. Statements of significant probability were based on P 0.05 unless otherwise noted. RESULTS AND DISCUSSION The analysis of the diets for vitamin D activity indicated that the diets were in reasonable

4 266 JAPR: Research Report TABLE 1. Composition (g/kg) and nutrient analysis of diets Ingredient 0 to 21 d 21 to 42 d Yellow corn Soybean meal (48% CP) Poultry oil Dicalcium phosphate Ground limestone DL-Methionine (98%) Iodized salt Coban-60 A Vitamin premix B Trace mineral mix C Choline chloride (60%) D L-Threonine L-Lysine HCl (98%) Nutrient analysis E ME, kcal/kg 3, , Crude protein, % Crude protein, % (analyzed) Calcium, % Calcium, % (analyzed) Total P, % Total P, % (analyzed) Nonphytate P, % Methionine, % Lysine, % TSAA, % A Elanco Animal Health Division of Eli Lilly & Co., Indianapolis, IN. B Provided per kilogram of diet: 8,800 IU of vitamin A; 20 IU of vitamin E; mg of vitamin B 12 ; 8 mg of riboflavin; 50 mg of niacin; 15 mg of pantothenic acid; 465 mg of choline; 2 mg of vitamin K; 1 mg of folic acid; 2 mg of thiamin; 2.5 mg of pyridoxine; 0.1 mg of D-biotin; 135 mg of ethoxyquin; 0.1 mg of Se. C Provides per kilogram of diet: Mn (from MnSO 4 H 2 0) 100 mg; Zn (from ZnSO 4 7H 2 O) 100 mg; Fe (from FeSO 4 7H 2 O) 50 mg; Cu (from CuSO 4 5H 2 0) 10 mg; I from Ca(IO 3 ) 2 H 2 0), 1 mg. D Provided 236 mg/kg supplemental choline. E Calculated from NRC (1994) adjusted to crude protein and moisture content of ingredients unless noted otherwise. TABLE 2. Calculated and analyzed levels of vitamin D in test diets A 0 to 21 d 21 to 42 d Source of Expected Found Expected Found vitamin D B (µg/kg) (µg/kg) (µg/kg) (µg/kg) VIT-D < <5 VIT-D VIT-D VIT-D VIT-D VIT-D OH-D < <2 25-OH-D OH-D OH-D OH-D OH-D A Assays were conducted by Monsanto Animal Nutrition, Naperville, IL. B VIT-D3 = cholecalciferol; 25-OH-D3 = 25-hydroxycholecalciferol.

5 FRITTS ET AL.: VITAMIN D SOURCE AND IMMUNE FUNCTION 267 TABLE 3. Effects of source and level of vitamin D on in vitro production of nitric oxide and in vitro cytotoxic activity by broiler microphages with and without in vitro lipopolysaccharide (LPS) stimulation in 21-d broilers Nitric oxide C (µm) Cytotoxicity D (%) Vitamin D level B Comparison A (IU/kg) Unactivated LPS-activated Unactivated LPS-activated Vitamin D source Vit-D ± ± ± ± OH-D ± ± ± ± 3.6 Vitamin D level ± ± ± ± ± ± ± ± ± ± ± ± 6.2 1, ± ± ± ± 5.9 2, ± ± ± ± 6.2 4, ± ± ± ± 6.4 Vitamin D source level Vit-D ± ± ± ± 9.0 Vit-D ± ± ± ± 8.5 Vit-D ± ± ± ± 9.0 Vit-D3 1, ± ± ± ± 8.5 Vit-D3 2, ± ± ± ± 8.2 Vit-D3 4, ± ± ± ± OH-D ± ± ± ± OH-D ± ± ± ± OH-D ± ± ± ± OH-D3 1, ± ± ± ± OH-D3 2, ± ± ± ± OH-D3 4, ± ± ± ± 8.5 Source of variance Probability > F Source of D Level of D Source level A Vit-D3 = cholecalciferol; 25-OH-D3 = 25-hydroxycholecalciferol. B Levels of 25-hydroxycholecalciferol were calculated to be equal to similar levels of cholecalciferol based on conversion of µg of cholecalciferol to 1 IU. C Nitric oxide production (mean ± SEM) determined in the presence or absence of 10 µg of LPS per well using a nitrite assay. D Percentage cytotoxicity (mean ± SEM); culture supernatant fluid from abdominal macrophages cultured 24 h in the presence or absence of 10 µg of LPS per well was collected and cytotoxic activity assessed using RP9 tumor cells and MTT (2 parts 2-propanol, 1 part PBS, and parts hydrochloride) assay.

6 268 JAPR: Research Report TABLE 4. Effects of source and level of vitamin D on the phytohemagglutinin-p- (PHA-P) induced wattle swelling response in 35-d broilers A [increase (mm) of PHA-Pinjected wattle increase of saline-injected wattle (mean ± SEM)] Hours postinjection with 100 µg of phytohemaglutinin per wattle Vitamin D level B Comparison A (IU/kg) 4 h 8 h 12 h 20 h 24 h Vitamin D source C Vit-D ± ± ± ± ± OH-D ± ± ± ± ± 0.06 Vitamin D level ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± , ± ± ± ± ± , ± ± ± ± ± , ± ± ± ± ± 0.11 Vitamin D source level Vit-D ± ± ± ± ± 0.16 Vit-D ± ± ± ± ± 0.16 Vit-D ± ± ± ± ± 0.16 Vit-D3 1, ± ± ± ± ± 0.16 Vit-D3 2, ± ± ± ± ± 0.16 Vit-D3 4, ± ± ± ± ± OH-D ± ± ± ± ± OH-D ± ± ± ± ± OH-D ± ± ± ± ± OH-D3 1, ± ± ± ± ± OH-D3 2, ± ± ± ± ± OH-D3 4, ± ± ± ± ± 0.16 Probability > F Source of variance Source of D Level of D Source level A Increase (mm) of PHA-P-injected wattle increase of saline-injected wattle (mean ± SEM). B Vit-D3 = cholecalciferol; 25-OH-D3 = 25-hydroxycholecalciferol. C Levels of 25-hydroxycholecalciferol were calculated to be equal to similar levels of cholecalciferol based on conversion of µg of cholecalciferol to 1 IU.

7 FRITTS ET AL.: VITAMIN D SOURCE AND IMMUNE FUNCTION 269 TABLE 5. Effects of source and level of vitamin D on blood cell counts and heterophyl:lymphocyte ratio in 42-d broilers (mean ± SEM) Cell concentrations (10 3 /µl) Dietary treatments Heterophil: Source of Level of Red blood White blood lymphocyte vitamin D vitamin D cells cells Lymphocytes Heterophils Monocytes Eosinophils Basophils Thrombocytes ratio Vit-D ± ± 0.9 b 8.49 ± ± ± ± ± ± ± OH-D ± ± 0.9 a 9.78 ± ± ± ± ± ± ± ± ± ± ± 0.6 b 2.60 ± ± ± ± ± ± ± ± ± 0.6 b 2.36 ± ± ± ± ± ± ± ± ± 0.6 ab 2.92 ± ± ± ± ± , ± ± ± ± 0.6 ab 2.55 ± ± ± ± ± , ± ± ± ± 0.6 a 3.02 ± ± ± ± ± , ± ± ± ± 0.6 b 2.36 ± ± ± ± ± 0.02 Source level Vit-D ± ± ± ± ± ± ± ± ± 0.03 Vit-D ± ± ± ± ± ± ± ± ± 0.03 Vit-D ± ± ± ± ± ± ± ± ± 0.03 Vit-D3 1, ± ± ± ± ± ± ± ± ± 0.03 Vit-D3 2, ± ± ± ± ± ± ± ± ± 0.03 Vit-D3 4, ± ± ± ± ± ± ± ± ± OH-D ± ± ± ± ± ± ± ± ± OH-D ± ± ± ± ± ± ± ± ± OH-D ± ± ± ± ± ± ± ± ± 0.03 V25-OH-D3 1, ± ± ± ± ± ± ± ± ± 0.03 V25-OH-D3 2, ± ± ± ± ± ± ± ± ± 0.03 V25-OH-D3 4, ± ± ± ± ± ± ± ± ± 0.03 Probability > F Source of variation Source of D Level of D Source level a,b Means with common superscripts do not differ significantly (P 0.05).

8 270 JAPR: Research Report TABLE 6. Effects of source and level of vitamin D on the proportions among white blood cells in 42-d broilers (mean ± SEM) Cell proportions (%) Dietary treatments Level of vitamin D Lymphocytes Heterophils Monocytes Eosinophils Basophils Source of vitamin D Vit-D ± 0.4 b ± ± ± ± 0.07 b 25-OH-D ± 0.4 a ± ± ± ± 0.07 a ± 0.7 a ± 0.8 c 4.46 ± 0.2 b 1.34 ± 0.1 b 0.71 ± ± 0.8 b ± 0.8 ab 4.52 ± 0.2 b 0.95 ± 0.1 c 1.10 ± ± 0.7 c ± 0.8 a 4.65 ± 0.2 b 1.42 ± 0.1 b 0.89 ± 0.1 1, ± 0.8 a ± 0.8 c 4.69 ± 0.2 ab 1.35 ± 0.1 b 0.89 ± 0.1 2, ± 0.8 bc ± 0.8 b 5.30 ± 0.2 a 1.82 ± 0.1 a 0.92 ± 0.1 4, ± 0.7 ab ± 0.8 bc 4.13 ± 0.2 b 1.59 ± 0.1 ab 0.92 ± 0.1 Source level Vit-D ± 1.0 abc ± 1.1 cde 4.59 ± 0.3 cd 1.53 ± ± 0.2 d Vit-D ± 1.1 bc ± 1.1 bcd 4.55 ± 0.3 cd 0.93 ± ± 0.2 a Vit-D ± 1.0 cd ± 1.1 bc 4.80 ± 0.3 bc 1.28 ± ± 0.2 cd Vit-D3 1, ± 1.0 bc ± 1.1 cde 5.55 ± 0.3 b 1.38 ± ± 0.2 cd Vit-D3 2, ± 1.1 de ± 1.1 abc 6.56 ± 0.4 a 1.82 ± ± 0.2 bcd Vit-D3 4, ± 1.0 cd ± 1.1 bc 4.19 ± 0.3 cd 1.55 ± ± 0.2 bcd 25-OH-D ± 1.1 a ± 1.1 e 4.34 ± 0.3 cd 1.16 ± ± 0.2 bcd 25-OH-D ± 1.1 cd ± 1.1 ab 4.49 ± 0.3 cd 0.97 ± ± 0.2 bcd 25-OH-D ± 1.0 e ± 1.1 a 4.50 ± 0.3 cd 1.56 ± ± 0.2 ab 25-OH-D3 1, ± 1.1 a ± 1.2 e 3.84 ± 0.3 d 1.32 ± ± 0.2 ab 25-OH-D3 2, ± 1.1 abc ± 1.2 cde 4.05 ± 0.3 cd 1.81 ± ± 0.2 abc 25-OH-D3 4, ± 1.1 ab ± 1.1 de 4.06 ± 0.3 cd 1.63 ± ± 0.2 abc Probability > F Source of variation Source of D Level of D Source level a e Means with common superscripts do not differ significantly (P 0.05).

9 FRITTS ET AL.: VITAMIN D SOURCE AND IMMUNE FUNCTION 271 agreement with calculated values (Table 2). Therefore, the results of the present study can be used with confidence in evaluating possible effects of the 2 sources of vitamin D on immune status of the broiler. Nitric oxide production at 3 wk of age, assessed as the concentration of nitrite (µm) produced in vitro by AEC, was not significantly affected by dietary source or level of vitamin D, with no interaction between source and level of vitamin D (Table 3). Nitric oxide production of AEC following in vitro stimulation of LPS was higher than that by AEC without LPS stimulation. The macrophage cytotoxic killing of RP9 tumor cells expressed as percent cytotoxicity in AEC culture supernatant fluid was not significantly affected by dietary source or level of vitamin D (Table 3). Cytotoxic activity of AEC following in vitro stimulation with LPS was higher than that by AEC without LPS stimulation. The maximum CBH swelling response in 35-d-old male broilers occurred 4 to 8 h after PHA-P injection and decreased by 12 h. Swelling response to PHA-P injection in the wattle was not significantly affected by dietary vitamin D source or level (Table 4). Red blood cell concentration and thrombocyte concentration (Table 5) were not significantly affected by source or level of vitamin D, with no interaction by source or level of vitamin D. The concentration of white blood cells (10 3 / µl of blood) was significantly higher for birds fed diets supplemented with 25-OH-D3 than for birds fed the VIT-D3 at 42 d (Table 5). Although the various types of white blood cell concentrations were within a physiological normal range for chicks fed both sources of vitamin D, statistical differences were seen in concentrations of heterophils, monocytes, and eosinophils in broilers fed various levels of vitamin D. However, no consistent pattern was seen regarding the effects of vitamin D level on white blood cell concentration. The same was true for the interaction of source and level of vitamin D on the concentration of the individual white blood cells. Similarly, the proportions of heterophils (Table 6) differed significantly when individual treatment effects were examined for each source; however, no consistent trend regarding effects of source or level on the proportions of these cells were observed. Due to significant source by level interactions, the main effects of source and level could not be assessed. Overall, the concentrations of white blood cells were within a physiological normal range (Table 5). The heterophil to lymphocyte ratio was also not significantly affected by vitamin D source or level (Table 5). The present study was conducted to evaluate both VIT-D3 and 25-OH-D3 fed at levels ranging from below NRC recommendations to those commonly fed in commercial formulations to male broilers on immune response. Feeding 2 sources and various levels of vitamin D did not demonstrate immunomodulatory effects in aspects of cell-mediated immunity of broilers. Macrophage NO production and cytotoxic activity, PHA-P induced swelling response, or the concentrations and proportions among white blood cells were not affected by vitamin D source. Although significant differences were seen in both concentration and proportions among white blood cells, nothing consistent was observed and all concentrations and proportions were within a normal physiological range [44]. Also, no differences were seen in the heterophil to lymphocyte ratio. The ratio was within a normal range [46]. Aslam et al. [16] found broilers deficient in vitamin D had a lower CBH response to PHA-P than broilers fed a VIT-D3 diet of 800 ICU/kg. In the present study the diets were not severely deficient in vitamin D, which may explain why no differences were seen. Moreover, amounts of vitamin D in the levels often used in commercial broiler production (2,000 to 4,000 ICU/kg) did not alter the immune system in broilers within the parameters measured. The findings in this study indicate that immunocompetence of broiler chickens was not affected by source or level of vitamin D. However, feeding high levels of vitamin D, which is often done in commercial situations, does not appear to alter immunocompetence in broilers.

10 272 JAPR: Research Report CONCLUSIONS AND APPLICATIONS 1. There were no significant differences observed when feeding low to high levels (125 to 4,000 ICU) of vitamin D to male broilers on macrophage function (nitric oxide production and cytotoxicity) at 21 d and cutaneous basophil hypersensitivity at 35 d. 2. Significant differences were observed for the concentration and proportions among white blood cells, but these differences did not follow a consistent pattern. 3. Feeding levels of 2,000 or 4,000 ICU of vitamin D, which are commonly fed in commercial poultry production, did not positively or negatively affect the immune system within the parameters measured. REFERENCES AND NOTES 1. van Etten, E., B. Decallonne, and C. Mathieu ,25- dihydroxycholecalciferol: Endocrinology meets the immune system. Proc. Nutr. Soc. 61: Kolb, E., S. Kaskous, and J. 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11 FRITTS ET AL.: VITAMIN D SOURCE AND IMMUNE FUNCTION Cobb 500, Cobb-Vantress, Inc., Siloam Springs, AR. 33. Sigma Chemical, St. Louis, MO. 34. Qureshi, M. A., and L. Miller Comparison of macrophage function in several genetic broiler lines. Poult. Sci. 70: Difco Laboratories, Detroit, MI. 36. Green, L. C., D. A. Wagner, J. Glogowski, P. L. Skipper, J. S. Wishnok, and S. R. Tannebaum Analysis of nitrate, nitrite, and ( 15 N) nitrite in biological fluids. Anal. Biochem. 126: Bio-Tek Instruments, Winooski, VT. 38. Stadecker, M. J., M. Lukic, A. Dvorak, and S. Leskowitz The cutaneous basophil response to phytohemagglutinin in chickens. J. Immunol. 118: McCorkle, F., Jr, I. Olah, and B. Glick The morphology of the phytohemagglutinin-induced cell response in the chicken s wattle. Poult. Sci. 59: Edelman, A. S., P. L. Sanchez, M. E. Robinson, G. M. Hochwald, and G. J. Thorbecke Primary and secondary wattle swelling response phytohemagglutinin as a measure of immunocompetence in chickens. Avian Dis. 30: Corrier, D. E., and J. R. DeLoach Evaluation of cellmediated, cutaneous basophil hypersensitivity in young chickens by an interdigital skin test. Poult. Sci. 69: Kean, R. P., and S. J. Lamont Effect of injection site on cutaneous basophil hypersensitivity response to phytohemagglutinin. Poult. Sci. 73: Abbott Diagnostics, Abbott Park, IL. 44. Lucas, A. M., and C. Jamroz Atlas of Avian Hematology. Agriculture Monograph. USDA, Washington, DC. 45. SAS Institute SAS User s Guide: Statistics. Version 6.03 ed. SAS Institute Inc., Cary, NC. 46. Gross, W. B., and H. S. Siegel Evaluation of the heterophil/lymphocyte ratio as a measure of stress in chickens. Avian Dis. 27: A 3% suspension of Sephadex G-50 was prepared in sterile Dulbecco s PBS [33] and injected at a dose of 1 ml/100-g BW. Approximately 42 h after injection of Sephadex G-50, birds were euthanized by pentobarbital injection. The abdominal cavity was injected with heparinized PBS and fluid containing AEC was drawn from the abdominal cavity. Approximately 30 ml AEC was collected in siliconized glass tubes and centrifuged at 10 C for 20 min. The AEC pellets were re-suspended in LM Hahn medium. The concentration of AEC was determined by counting live cells on a hemacytometer using Trypan blue exclusion. Cells ( macrophages/ml) from each bird were cultured in 24-well culture plates (4 wells per bird), with or without 10 µg/ml LPS [35] in 1 ml LM Hahn mediumto determine the effects of diet on the production of NO, (measured as nitrite) and macrophage cytotoxicity (measured as percentage killing of RP9 tumor cells). After 24 h, the culture supernatant from each well was collected. 48. Briefly, 100 µl AEC culture supernatant fluid from each sample were added to 100 µl of Griess reagent (1 part 0.1% naphthylenediamine dihydrochloride to 1 part of 1% sulfanilamide in 5% phosphoric acid) in 96-well plates. After 10 min of incubation at room temperature, color intensity indicative of nitrite was quantified by reading the plates at 540 nm absorbance in a microplate reader [37]. 49. Live cell concentration of RP9 tumor cells was determined using Trypan blue and a hemacytometer. The cell concentration was adjusted to cell/ml using LM Hahn medium. Fifty microliters of RP9 cells were added to each of 48 wells in a 96-well plate. Each 24-h AEC culture supernatant was then added at 50 µl in quadruplicates to wells with RP9 cells. To serve as negative (no killing) and positive (100% killing) controls, 50 µl of LM Hahn medium and 50 µl Triton-X were added to empty wells, respectively. The plates were covered and incubated for 18 h at 41 C, 5% CO 2. After 18 h, 50 µl MTT acid (2 parts 2-propanol, 1 part PBS, and parts hydrochloride) were added to each of the wells to identify live RP9 cells and incubated for another 4 h. After the 4 h incubation, plates were centrifuged at 450 g at 4 C for 4 min. All liquid was then removed from the wells using a multichannel pipettor, being careful not to disturb the crystals. The MTT acid was added again at 150 µl to each well and mixed until all crystals were dissolved. Plates were read at 540 nm in a microplate reader. 50. Blood smears were prepared by placing a small drop (3 to 5 µl) of blood on a clean microscope slide and using another clean slide to push the blood across the slide to spread the blood evenly. Wright stain (adapted for avian blood as described by Lucas and Jamroz [44]) was used to stain leukocytes. Acknowledgments This work was supported in part by a grant from Monsanto Animal Nutrition, Naperville, IL. The support of Alpharma and Monsanto Animal Nutrition in providing the cholecalciferol and the 25- hydroxycholecalciferol products and conducting the cholecalciferol assays is greatly appreciated. The technical suggestions and excellent manuscript review by William E. Huff are acknowledged.