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Departments of Poultry Science,
*
Nutrition and
Biochemistry and Molecular Biology, The Pennsylvania State University, University Park, PA 16802-3501
1To whom correspondence should be addressed. E-mail: lnr{at}psu.edu.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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KEY WORDS: • zinc deficiency • chickens • growth plate chondrocyte • proliferation • apoptosis
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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). Chondrocytes within the growth plate continuously proliferate, differentiate and hypertrophy in an orderly process. The hypertrophic chondrocytes are eventually removed and replaced by trabecular bone. These sequential events, called endochondral bone formation, result in elongation of long bones.
Dwarfism is a characteristic symptom of zinc deficiency and has been linked to decreased activity of the epiphyseal growth plate. In addition to a decrease in the width of the growth plate, there are specific pathological lesions associated with zinc deficiency (2
–5
). In the growth plate of affected chickens, the chondrocytes near blood vessels appeared normal, whereas the remote cells had abnormal shapes and were decreased in number (4
,5
). Although the growth plate pathology has been described for decades, the molecular and cellular changes caused by zinc deficiency have not yet been elucidated.
Zinc itself plays a critical role in cell proliferation, differentiation and survival (6
–8
). In a serum-free cell culture system, a zinc supplement stimulates the proliferation of epiphyseal growth plate chondrocytes (9
). In addition, zinc may also indirectly affect cellular activities through hormones and growth factors. It has been shown that zinc deficiency reduces serum insulin-like growth factor-1 (IGF-1)2
levels (3
,10
). The importance of IGF-1 in longitudinal bone growth has been confirmed in gene knockout studies. Mice lacking the IGF-1 gene or the receptor gene demonstrate severe dwarfism (11
–13
). However, recent studies have shown that by abolishing IGF-1 production in the liver and thus reducing circulating IGF-1, no effect is shown in postnatal growth in mice (14
,15
), suggesting that IGF-1 locally produced in tissues plays a more important role in maintaining normal growth. In addition to IGF-1, endochondral bone formation is also influenced by other autocrine/paracrine growth factors such as parathyroid hormone related protein (PTHrP) and fibroblast growth factor-2 (FGF-2) (16
,17
). It is not known whether zinc deficiency changes the production of these local growth factors in the growth plate.
The ability to study specific effects of zinc deficiency is impaired by the fact that zinc deficiency has severe effects on food intake and growth rate. To circumvent this problem, we decided to focus on the earliest detectable events associated with zinc deficiency. Preliminary experiments showed histological changes in growth plates of chickens fed a low zinc diet for 7 d. To elucidate the cellular events responsible for the growth plate pathology, cell proliferation, differentiation and apoptosis were studied in growth plates of zinc-deficient chickens before (d 3) and after (d 7) histological changes. A zinc-deficient state was developed in juvenile chickens with a low zinc soy protein–based diet. Chondrocyte proliferation in the growth plate was evaluated with the bromodeoxyuridine (BrdU; Sigma, St. Louis, MO) labeling technique. Apoptosis was detected using the terminal deoxynucleotidyl transferase-mediated dUTP nick end labeling (TUNEL) method. Chondrocyte differentiation was assessed with immunostaining of osteonectin as a marker of chondrocyte maturation. Because we also hypothesized that the effects of zinc deficiency could be indirectly caused by changes in local growth factors, production of IGF-1, PTHrP and FGF-2 in the growth plate was examined with immunohistochemistry.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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Newly hatched male broiler chickens (Avian x Avian; Longnecker Hatchery, Elizabethtown, PA) raised at the Pennsylvania State University Poultry Education and Research Center were used in this study. Birds were maintained under a 16-h light:8-h dark cycle. All animal care and surgical procedures were in accordance with protocols approved by the Institutional Animal Care and Use Committee of the Pennsylvania State University (IACUC # 94R120D097).
Experimental design.
Newly hatched chickens (n = 96) were randomly divided into three groups with four pens of eight chickens per pen. The chickens in one group (zinc-deficient, -Zn) were fed a soy protein–based diet which contained (by analysis) 10 mg zinc/kg of diet3
(Table 1)
. Chickens in the other two dietary groups (zinc-adequate) received the same soybean-based diet supplemented with zinc carbonate so that it contained 68 mg zinc/kg of diet. Chickens in one of the zinc-adequate groups were allowed free access to diet throughout the experiment (ad libitum consumption, +ALZn). The chickens in the other zinc-adequate group consumed feed ad libitum for the first 4 d. From d 5 to the end of the experiment, they were given the amount of diet consumed in the previous 24 h by their zinc-deficient counterparts (pair-fed, +PFZn). Deionized distilled water was consumed ad libitum. Feed intake was recorded daily. The chickens were weighed at the beginning of the experiment and before killing.
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Four chickens from each pen of the three treatment groups were killed by cervical dislocation after 3 and 7 d of feeding. On each of these days, two chickens per pen were injected intraperitoneally with 50 mg BrdU/kg body weight 1 h before killing. Blood was drawn from two birds per pen by cardiac puncture and collected in 2.7-mL monovette syringes (Sarstedt, Nuembrecht, Germany) with potassium-EDTA (1.6 g EDTA/L blood) as an anticlotting factor. The blood was immediately centrifuged (700 x g for 10 min) to separate the plasma, which was stored at -20°C for zinc analysis. The proximal tibiotarsi were dissected and either fixed in 4% paraformaldehyde in PBS or immediately immersed in isopentane (at melting point -159.9°C), which was prechilled in liquid nitrogen. The tissues fixed in 4% paraformaldehyde were embedded in paraffin and serial 5-µm thick longitudinal sections were prepared and stained with hematoxylin-eosin (H & E). Unstained sections were used for immunohistochemistry. Frozen tissues were mounted in tissue-freezing medium (Triangle Biomedical Sciences, Durham, NC) on dry ice. Serial, 8-µm thick longitudinal and cross sections of frozen tissues were cut on a cryostat (Richard-Allan Scientific, Kalamazoo, MI) and collected on Superfrost/plus microscope slides (Fisher Scientific, Pittsburgh, PA). These frozen sections were stored at -80°C and used for examination of nuclear BrdU incorporation and in situ apoptotic cell labeling.
Mineral analysis.
Diet samples were digested in instra-analyzed nitric acid (J. T. Baker, Phillipsburg, NJ) on a hot plate (Thermolyne 2200; Thermolyne, Dubuque, IA) at 100°C (19
), and plasma samples were diluted four times with deionized distilled water before zinc analysis. Zinc concentration was determined on a flame atomic absorption spectrophotometer (Instrumental Laboratory Video 11; Allied Analytical Systems, Andover, MA) at a wavelength of 213.9 nm. Zinc standard (15.3 mmol/L) (Sigma) was diluted with deionized distilled water to form a series of standards that ranged from 0 to 15.3 µmol zinc/L.
BrdU incorporation.
Nuclei with incorporated BrdU were detected using a procedure described by Farquharson and Loveridge (20
). Frozen sections were thawed and fixed with cold acetone for 5 min and air-dried. Nuclear DNA was denatured with 1.5 mol/L HCl for 30 min at room temperature, followed by three washes with PBS, pH 7.4. Sections were incubated for 2 h at 37°C with a monoclonal antibody (primary antibody) to BrdU diluted 1:25 in PBS. After washing with PBS, the sections were incubated for another hour at room temperature with a secondary antibody, goat anti-mouse immunoglobulin (Ig)G conjugated to fluorescein isothiocyanate (FITC; Molecular Probes, Eugene, OR) diluted 1:25 in PBS. The sections were then washed with PBS and stained for 5 min at room temperature with 4,6-diamindino-2-phenylindole (DAPI; Sigma) at a concentration of 100 µg/L. Finally, sections were counterstained with 0.004% Evan’s blue (Sigma) for 10 min and mounted with Fluoromount-G (Southern Biotechnology Associates, Birmingham, AL). Sections from BrdU-labeled tissues incubated with nonimmune mouse serum or non-BrdU–labeled tissues incubated with the primary antibody served as negative controls. All antibody solutions were clarified by brief centrifugation (1000 x g for 5 sec) before use. Sections were examined under a Eclipse TE200 fluorescence microscope (Nikon, Tokyo, Japan). Excitation and emission wavelengths for fluorescein were 495 and 525 nm, respectively. For DAPI, excitation and emission wavelengths were 360 and 450 nm, respectively.
One tibial section from each BrdU-injected chicken (total of eight sections per treatment on d 3 or 7) was studied. For the determination of BrdU and DAPI labeling, two representative fields of the BrdU-labeled zone in the longitudinal section of each growth plate were photographed with 800 speed color print film (Kodak, Rochester, NY). The distance between the upper and lower limits of the BrdU-labeled zone defined the width of the zone. BrdU- and DAPI-labeled nuclei were counted in the same field of the BrdU-labeled zone (each field of BrdU-labeled zone contained 
400 to 600 DAPI-labeled nuclei in the control growth plates). The BrdU labeling index was calculated as the ratio of the number of BrdU-labeled nuclei to the number of DAPI-labeled nuclei (total nuclei) in the same field.
In situ cell death detection.
In situ cell death was detected with TUNEL labeling described by Darfler and Karaszkiewicz (21
). Briefly, frozen sections were thawed and fixed with 10% buffered formalin (VWR Scientific Products, West Chester, PA) for 15 min at room temperature, washed twice with PBS and 70% ethanol, and air-dried. Sections were then digested for 15 min at 37°C with proteinase K (Sigma) at a concentration of 2 mg/L and washed three times with PBS. Sections were incubated for 15 min at 37°C with 25 µL of a solution containing 250,000 U/L terminal transferase, 50 µmol/L fluorescein-dUTP, 2.5 mmol/L cobalt chloride, 0.2 mol/L potassium cacodylate, 0.25 g/L bovine serum allbumin and 25 mmol/L Tris-HCl, pH 6.6 (Boehringer Mannheim, Indianapolis, IN). After washing with PBS, sections were stained for 5 min at room temperature with DAPI at a concentration of 100 µg/L. Finally, sections were counterstained with 0.004% Evan’s blue for 10 min and mounted with Fluoromount-G. Sections from the tibiae of two chickens in each pen (total of eight sections per treatment on d 3 or 7) were examined and photographs were taken with fluorescence microscopy as described above.
Immunohistochemistry.
Paraffin-embedded sections were deparaffinized and rehydrated before immunostaining with sequential 5-min washes in the following solutions: xylene, xylene, xylene/ethanol (1:1), 100% ethanol, 95% ethanol, 70% ethanol, distilled water and PBS. When not specified, steps in the procedure were carried out at room temperature. Control sections were incubated with nonimmune rabbit serum or nonimmune rabbit IgG instead of the primary antibody. Eight sections (one section/chicken) per treatment were studied in each of the following immunohistostainings.
Immunostaining for osteonectin was performed according to Wu et al. (22
). Briefly, after treatment with hyaluronidase and 10% normal goat serum, the sections were incubated with anti-osteonectin antibody (rabbit polyclonal LF-8; NIH/NIDCR, Bethesda, MD) (23
) in PBS (1:200 dilution) overnight at 4°C. After washing with 0.5% goat serum (diluted in PBS), the sections were incubated with goat anti-rabbit IgG conjugated to FITC (Molecular Probes) in 0.5% goat serum (1:250) for 1 h and washed again with 0.5% goat serum. Finally, the sections were counterstained with 0.004% Evan’s blue for 10 min and mounted with Fluoromount-G. Sections were examined with fluorescence microscopy and photographed.
Immunostaining for PTHrP was performed according to Medill et al. (24
). Deparaffinized sections were pretreated with hyaluronidase, immersed in 0.3% hydrogen peroxide (Sigma) to eliminate endogenous peroxidase activity and then in 10% normal goat serum. The sections were incubated overnight at 4°C with anti-PTHrP antibody (rabbit polyclonal raised against residues 34–53 of human PTHrP; Calbiochem, La Jolla, CA) at a concentration of 4 mg/L in PBS. After washing with PBS, the sections were incubated with biotinylated goat anti-rabbit IgG (Sigma) in PBS (1:100 dilution) for 30 min and washed again with PBS. The sections were then incubated in ABC reagent (Vectastain kit; Vector Laboratories Burlingame, CA) for 20 min. The sections were rinsed with PBS and 1% Triton-X 100 in PBS, then incubated for 8 min with 100 µL of a solution containing 67 µg diaminobenzadine tetrahydrochloride (Sigma) and 0.03% hydrogen peroxide. After a rinse with distilled water, the sections were mounted with 90% glycerol in PBS. The sections were examined under a TMS microscope (Nikon) and photographed with 100 speed color print film (Kodak).
Immunostaining for IGF-1 and FGF-2 was performed with a procedure similar to that described above for immunostaining of PTHrP. Tissue sections were treated with hyaluronidase, hydrogen peroxide and 10% normal goat serum, and then incubated overnight at 4°C with either anti-IGF-1 antibody (rabbit polyclonal UB3–189; NIH, Bethesda, MD) or anti-FGF-2 (rabbit polyclonal; Sigma) in PBS (1:200 dilution). After washing with 0.5% goat serum (diluted in PBS), the sections were incubated with goat anti-rabbit IgG conjugated to horseradish peroxidase (Sigma) in 0.5% goat serum (1:250) for 1 h and washed again with 0.5% goat serum. The sections were subsequently incubated with diaminobenzadine tetrahydrochloride for 8 min and mounted with 90% glycerol in PBS.
Statistical analyses.
The experimental design was a two-factor, split-plot design. Experimental units (pens of chickens) were randomly assigned to three dietary levels and each experimental unit was divided into two subunits, randomly assigned to two levels of a time factor (after 3 d of feeding and after 7 d of feeding). ANOVA was conducted using a general linear model program of Minitab Statistical Software (Minitab, State College, PA). When the main effect was significant, Tukey’s test was used to separate treatment means. Differences were considered to be significant at a level of P < 0.05.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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Neither feed intake nor body weight was different among chickens in zinc-deficient (-Zn), pair-fed (+PFZn) and ad libitum-consumption (+ALZn) groups throughout the 7-d experiment (Fig. 1
). However, plasma zinc concentration in -Zn chickens was 65% lower than that in +PFZn and +ALZn chickens on d 3 and 7 (P < 0.05). No differences were found in plasma zinc levels between +PFZn and +ALZn groups.
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No differences in histology were observed between growth plates from +PFZn and +ALZn chickens on d 3 or d 7 (Fig. 2
). Therefore, +PFZn and +ALZn groups are described as one zinc-adequate group (+Zn). H & E staining did not reveal any histological differences in growth plates of -Zn chickens (-Zn growth plates) on d 3 compared with those of +Zn chickens (+Zn growth plates). However, -Zn growth plates developed characteristic histological changes on d 7 similar to those reported by Westmoreland (5
). First, the columnar structure of chondrocytes was disrupted in the growth plate (Fig. 2D
). Second, chondrocytes in areas away from growth plate–penetrating blood vessels had a variety of shapes, forming distinct histological lesions. Cellular density in those lesions was reduced and, therefore, it seemed that cells were surrounded by larger amounts of extracellular matrix.
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BrdU was used to label the nuclei of chondrocytes undergoing DNA synthesis in the S phase of a cell proliferation cycle. A distinct zone adjacent to articular cartilage was visualized with heavy labeling in the longitudinal sections of +Zn growth plates (Fig. 3A and E
). In contrast, very few chondrocytes in articular cartilage were labeled with BrdU.
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). In the upper proliferative zone, only a few layers of chondrocytes were labeled horizontally across the growth plate. In the lower proliferative zone, only chondrocytes in the vicinity of blood vessels were positive, whereas the vast majority of cells remained unlabeled. A similar labeling pattern was observed on d 7 (Fig. 3D
).
Zinc deficiency dramatically reduced the width of the BrdU-labeled zone by 80% on both d 3 and 7 (Table 2)
. In contrast, the BrdU labeling index was less affected. On d 3, the index in -Zn growth plates (18.9%) was slightly lower than that in +Zn growth plates (22.6%, P < 0.05), whereas no differences were found between +Zn and -Zn treatments on d 7. The width of the BrdU-labeled zone and the labeling index were not different between +ALZn and +PFZn groups on d 3 or 7 (Table 2)
.
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100 µm of blood vessels (Fig. 4C and G
). In +Zn growth plates, BrdU labeling did not show a blood vessel–related pattern (Fig. 4A and E
).
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The TUNEL method was employed to detect DNA cleavage associated with the process of cell apoptosis. In cross sections of the lower proliferative zone of +Zn growth plates, no chondrocytes were found positive, but some cells associated with blood vessels were labeled with the TUNEL method (Fig. 5A and E
). Chondrocytes in the upper proliferative, prehypertrophic and hypertrophic zones of +Zn growth plates were also negative for apoptotic labeling (not shown).
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). Apoptotic chondrocytes were present in areas 100 µm from blood vessels (Fig. 5C and G
), where chondrocytes were not labeled with BrdU (Fig. 4C and G
). Further studies on longitudinal sections revealed that apoptotic chondrocytes were also found in the prehypertrophic zone, but not in the upper proliferative and hypertrophic zones (not shown). Immunohistochemistry of osteonectin, FGF-2, PTHrP and IGF-1.
Chondrocytes positive for osteonectin immunostaining were observed in the prehypertrophic and hypertrophic zones of +Zn growth plates, whereas little staining was present in the proliferative zone (Fig. 6A and C
). Therefore, osteonectin staining marks the event of chondrocyte maturation. In -Zn growth plates, chondrocytes in the vicinity of blood vessels had a staining pattern similar to that in +Zn growth plates. However, in areas remote from blood vessels, osteonectin-positive chondrocytes extended from the prehypertrophic zone into the lower proliferative zone (Fig. 6B and D
), indicating that early differentiation occurred in -Zn growth plates.
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)and PTHrP (Fig. 7B
), whereas only hypertrophic cells were positive for IGF-1 (Fig. 7C
). The staining pattern of these local growth factors in -Zn growth plates did not differ from that of +Zn growth plates on d 3. However, on d 7, staining of FGF-2, PTHrP and IGF-1 in chondrocytes of the growth plate lesion was greatly reduced (Fig. 7
).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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,25
), cell proliferation (26
) and cell apoptosis (27
). In the present study, short-term zinc deficiency did not reduce food intake of chickens. In addition, changes in cell proliferation, differentiation and apoptosis were manifested primarily in areas remote from blood vessels in the growth plate. Thus, it was very likely that cellular changes were directly associated with reduced zinc availability from the blood supply. Furthermore, local growth factors FGF-2, PTHrP and IGF-1 were not affected on d 3, excluding the possible involvement of indirect effects through these autocrine/paracrine factors. Therefore, the results of this study suggest that zinc status directly influences chondrocyte proliferation, differentiation and apoptosis in the growth plate.
Changes in cellular activities in the growth plate eventually resulted in formation of severe pathological lesions, which were located in areas away from blood vessels and characterized by cells with aberrant shapes and reduced cellularity. Comparable lesions have been reported in older chickens (10–28 d of age) with zinc deficiency (4
,5
) and in turkeys with deficiencies of choline or nicotinic acid (28
). It has been postulated that in a nutrient-deficient state, cells near the blood supply take up most of the available nutrients and leave little for remote cells (5
,28
). In addition, it has been demonstrated that the rate of cellular zinc uptake is increased fivefold upon addition of serum to stimulate the proliferation of G1-arrested baby hamster kidney cells (29
). Therefore, proliferating chondrocytes may also have a high requirement for zinc, which would exacerbate the zinc-deficient state in the growth plate.
BrdU labeling clearly demonstrated that areas of cell proliferation (the BrdU-labeled zone) were significantly decreased due to zinc deficiency. In contrast, the labeling index in the BrdU-labeled zone was not severely affected. An index of 21–23% was observed in the control groups, whereas in the zinc-deficient group, the index was 19–20%. Although younger chickens were used in the present study, the results were in line with a BrdU-labeling index of 21–25% in the growth plate of 3-wk-old chickens (30
,31
). The significantly decreased labeling areas and the less affected index indicated that zinc deficiency either completely stopped cell proliferation (in areas away from the blood supply) or did not affect it at all (in areas near the blood supply). This is in agreement with a study of Fujioka and Lieberman (32
), who showed that EDTA-induced zinc deficiency inhibited thymidine incorporation into liver cells of partially hepatectomized rats. The degree of inhibition was directly proportional to the reduction in labeled nuclei, suggesting that for a given liver cell, zinc deficiency blocked the replication of DNA either completely or not at all. As presented in this study, whether or not the proliferation of a chondrocyte was affected depended upon its distance from the blood supply.
This study showed that chondrocytes in the growth plate of control chickens were not labeled with the TUNEL method. This is consistent with the findings of Praul et al. (33
). However, in the growth plate of zinc-deficient chickens, TUNEL-positive chondrocytes were observed as early as d 3 of the study in the lower proliferative and prehypertrophic zones, but not in the hypertrophic zone, indicating that proliferative and prehypertrophic chondrocytes were vulnerable to zinc restriction, whereas the terminally differentiated chondrocytes were resistant.
It is interesting that proliferating and apoptotic chondrocytes were present in mutually exclusive areas in the lower proliferative zone of the growth plate from zinc-deficient chickens. This indicated that at least in some cells, apoptosis was initiated after blockage of proliferation. It has been demonstrated that cells cultured in a zinc-deficient medium lose their proliferation capacity and become apoptotic (34
). In vivo studies have also revealed that zinc deficiency induces apoptosis, particularly in cells undergoing rapid cell division (35
,36
). It is unclear why proliferating cells do not arrest in a quiescent phase instead of "committing suicide." One possibility is that a low level of zinc, which inhibits cell proliferation, is sufficient to trigger apoptosis. It is conceivable that zinc deficiency could affect totally unrelated pathways at the same time. On the one hand, zinc restriction may decrease the activities of thymidine kinase and DNA polymerase (37
) and impair cell commitment into S phase (38
). On the other hand, a lowered zinc level could initiate the process of apoptosis by reducing the inhibitory effect on Ca2+/Mg2+-dependent endonucleases (39
) and caspases (40
–42
), or by changing cellular redox potential (43
). Another explanation could be that cell proliferation and apoptosis are tightly coupled processes (44
). Blockage of proliferation with serum deprivation leads to apoptosis rather than growth arrest in c-myc transfected fibroblasts (45
). However, further studies are required to investigate whether such a mechanism exists in normal proliferating chondrocytes.
In the normal growth plate, immunostaining of osteonectin was observed in the differentiated chondrocytes, whereas little staining was present in the proliferating cells (22
). In the growth plate of zinc-deficient chickens, osteonectin staining extended from the prehypertrophic zone to the lower proliferative zone in areas in which proliferation was blocked, suggesting that cells in these areas started the process of differentiation. This is similar to the finding that zinc restriction with EDTA suppresses proliferation of HL-60 cells (a promyelocytic leukemia cell line) and promotes cell differentiation (46
). It is not known why zinc deficiency forces cell differentiation or whether the process resembles normal maturation. It is tempting to postulate that differentiation may help cells survive in a nutrient-deficient state. However, apoptosis occurred in chondrocytes undergoing maturation, suggesting that differentiation could not prevent cells from dying in a zinc-deficient state.
In summary, the present study demonstrated that dietary zinc deficiency inhibited chondrocyte proliferation, promoted cell differentiation, and induced apoptosis in the chicken growth plate within a few days. Lowered plasma zinc levels primarily affected cells remote from the blood supply. Zinc deficiency has been shown to decrease cell proliferation and cause apoptosis in other tissues such as rat liver, intestine and embryos (26
,27
,35
,37
). However, because the growth plate in animals is an avascular or poorly vascularized tissue, this study suggests that longitudinal bone growth might be especially sensitive to zinc status. Future studies are warranted to investigate the molecular mechanisms responsible for the effects of zinc deficiency on chondrocyte proliferation, differentiation and apoptosis. Because zinc deficiency impairs the transition of proliferating cells from G1 phase to S phase (38
), it would be of interest to study whether zinc deficiency affects chondrocyte levels of cyclin D and cyclin A, which are responsible for the transition. The outcome of these studies will facilitate our understanding of zinc biology and the influence of nutrients on longitudinal bone formation.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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3 The zinc requirement for chickens is estimated to range from 35 to 40 mg zinc/kg diet when soybean is the main source of protein (18
).
Manuscript received 9 October 2001. Initial review completed 17 December 2001. Revision accepted 17 January 2002.
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LITERATURE CITED |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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