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Journal of the American College of Nutrition, Vol. 24, No. 6, 494-502 (2005)
Published by the American College of Nutrition

Chronic Zinc Deficiency in Mice Disrupted T Cell Lymphopoiesis and Erythropoiesis While B Cell Lymphopoiesis and Myelopoiesis Were Maintained

Louis E. King, PhD, Joseph W. Frentzel, James J. Mann and Pamela J. Fraker, PhD

Department of Biochemistry & Molecular Biology (L. E. K., J. W. F., P. J. F.) and *Cell and Molecular Biology Program (J. J. M.), Michigan State University, East Lansing, Michigan

Address reprint requests to: Louis E. King, Ph.D., Department of Biochemistry and Molecular Biology, Room 419, Michigan State University, East Lansing, MI 48824-1319. E-mail: kingL{at}msu.edu


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 ACKNOWLEDGMENTS
 REFERENCES
 
Objective: The purpose of this study was to determine the impact of chronic zinc deficiency (ChrZD) on T and B cell lymphopoiesis, myelopoiesis and erythropoiesis in mice.

Methods: Young adult mice were fed a zinc adequate (ZA) or ChrZD synthetic diet for 34, 45, and 50 days. The cellular composition of the thymus and marrow were determined to assess the impact of ChrZD on lymphopoietic and hematopoietic processes using flow cytometry. Body weights, serum zinc and corticosterone (Cs) were monitored.

Results: For ChrZD mice growth was reduced 10% and serum zinc declined 15% by d 34 compared to ZA mice. By d 50 a 25% decrease in growth and 70% depression in serum zinc was noted though there was never any significant reduction in diet intake. Corticosterone rose 2.5 fold by d 34 and remained elevated in ChrZD mice indicating induction of the stress axis. At d 34 the thymus of ChrZD mice was normal but by d 50 there was a 50% cell loss and a 10% reduction in the proportion of Pre-T cells. Most importantly there was a 60% increase in Pre-T cells undergoing apoptosis in ChrZD mice. Pro-T, T helper, and T cytolytic populations were more resistant to ChrZD. Bone marrow cellularity and granulocyte, monocyte, and lymphocyte compartments remained unchanged in ChrZD mice. However, the erythroid compartment was reduced by 35% at d 50.

Conclusions: The thymus was the most sensitive primary tissue to ChrZD. By d 50 it had atrophied by 36% with significant loss of Pre-T cells via apoptosis such that T-cell lymhopoiesis was disrupted. Significant reductions were also noted in the erythropoietic population by d 50. Conversely the marrow maintained myelopoiesis and B cell lymphopoiesis for the 50 d period indicating greater ability to survive a chronic zinc deficiency and exposure to Cs. The anemia and T cell lymphopenia associated with ChrZD in both rodents and humans may result from a greater sensitivity of their precursor cells to zinc deficiency and elevated Cs.

Key words: chronic zinc deficiency, apoptosis, bone marrow, thymus, lymphopoiesis, myelopoiesis

Abbreviations: ChrZD = chronic zinc deficient • Cs = corticosterone • DAPI = 4', 6-diamidino-2-phenylindole • EDTA = ethylenediaminetetracetic acid disodium salt • FITC = fluorescein isothiocyanate • HEPES = (N-[2-Hydroxyethyl]piperazine-N'-[2-ethanesulfonic acid]) • HIFBS = heat inactivated fetal bovine serum • ICP = inductively coupled plasma-atomic emission spectrophotometer • PE = R-phycoerythrin • PE-Cy5 = R-phycoerythrin-cyanine 5 conjugate • PBS = phosphate buffered saline • SRM = Standard Reference Material • ZA = zinc adequate • ZD = zinc deficient


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 ACKNOWLEDGMENTS
 REFERENCES
 
Mild to moderate zinc deficiency is a common worldwide problem [2]. It has been reported for many diseases including liver disease [3], diarrhea, respiratory illness [4], gastrointestinal disorders, renal disease, sickle cell disease [5,6] and HIV/AIDS [7]. Mild zinc deficiency is also common among women of reproductive age and children who rely heavily on cereal grains for nutrition [8]. It has been estimated by some investigators that 82% of women worldwide may suffer from some degree of zinc deficiency and may benefit from zinc supplementation [9]. Chronic zinc deficiency has also been observed in some elderly, pregnant teenagers, and children from low income homes in the United States [10–12].

Growth retardation [13], reduced serum zinc [10, 14], thymic atrophy, and impaired antibody and cell mediated immunity are common characteristics of an inadequate intake of zinc [15–17]. Although it was known that deficiencies in zinc altered immune function [15] the impact of a chronic deficiency in zinc on T and B cell lymphopoiesis, erythropoiesis, and myelopoiesis is not known. Understanding ChrZD will also permit the identification of any significant differences between ChrZD and acute ZD. Understanding the changes made in these vital hematopoietic processes made by chronic deficiencies in zinc are important and are essential to the designing of nutritional and immunological therapeutic interventions in the clinical setting.

In acute zinc deficiency young adult mice respond quickly within 28–34 days to a zinc deficient diet of 0.5 mg or less Zn/kg diet. Deficient mice were defined as having developed visible alopecia and parakeratosis, being at 72–76% of the control group weight which included a 15% loss in body weight [18]. Serum zinc and thymic weight had fallen 50% and 60% respectively while serum Cs had risen 2.4 fold. During this short period there was a 50% loss in Pre-B cells and 40% loss in Pre-T cells associated with a greater than 50% increase in apoptotic cell death in Pre-T cells [19, 20]. Thus disruption of B and T cell lymphopoiesis occurred quickly. A subgroup of severely zinc deficient mice were also identified that had more extensive parakeratosis, being at 65–68% of control weight which includes a 22% loss from initial weight. Thymic atrophy and Pre-B cell loss were greater and serum Cs levels had risen 4.2 fold. Marrow hematopoietic changes included a 26%–50% reduction in erythropoiesis and a 137%–157% expansion in myeloid cells in acutely zinc deficient mice [18].

The hallmarks of chronic zinc deficiency are a prolonged period of dietary zinc deficiency resulting in growth retardation, reduced serum zinc, and thymic atrophy; all in the absence of alopecia and parakeratosis. This is the definition we use for ChrZD in young adult mice. Young adult mice developed ChrZD when they were fed a zinc deficient diet in multiple experiments of 34, 45, and 50 days. Flow cytometry was used to quantitate changes in thymic T cell lymphopoiesis and apoptotic cell death simultaneously using 4 fluorochromes by first limiting analysis to T cells with CD90.2 and then subdividing this heterogenous population into progenitor Pro-T cells (CD4–CD8–), Pre-T cells (CD4+CD8+) undergoing T cell receptor gene rearrangement, mature T helper cells (CD4+CD8–) and mature T cytotoxic cell (CD4–CD8+). Cell death was followed by examining the cell cycle of each population for the appearance of a subG1 peak of fluorescent DNA resulting from thymic T cell death by apoptosis. The thymus was chosen for analysis of apoptosis because Pre-T cells, which are the most sensitive to apoptosis, comprise approximately 80% of the thymic population making changes in apoptosis easily detectable. Flow cytometry was also used to follow the effects of ChrZD on the composition of bone marrow because it is the site of development for erythroblasts, B cell lymphopoiesis, development of granulocytes and monocytes, and proliferation of progenitor cells to these lineages. Flow cytometry separates and quantitates these 5 populations using CD31 and Ly-6C. They are: erythroblasts (CD31–Ly-6C–), B cells (CD31+Ly-6C–), granulocytes (CD31–Ly-6C+), monocytes (CD31–Ly-6C++), and committed progenitors for all these lineages (CD31+Ly-6C+). Disruption of B cell lymphopoiesis would be shown as a reduction in proportion of this population in the bone marrow. This paper will show ChrZD has a very different impact on primary immune tissues than does acute zinc deficiency.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 ACKNOWLEDGMENTS
 REFERENCES
 
Mice and Diets
Three to four week old female A/J mice were purchased from Jackson Laboratory (Bar Harbor, ME). A pelleted zinc adequate synthetic egg white based diet was provided until mice reached initial experimental weight. Mice were then divided into two groups and housed 3 mice per cage as previously described [20]. Zinc adequate mice (ZA) were fed ad libitum a zinc adequate diet (27–30 mg Zn/kg) and chronic zinc deficient mice (ChrZD) were fed a zinc deficient diet (ZD) (0.5–0.6 mg Zn/kg) (Dyets Inc., Bethleham, PA) for 34, 45, or 50 days performed as separate experiments with measurements taken on individual mice at the completion of the dietary period. The diet compositions are given in Table 1. Diet intake was recorded at 3–4 d intervals. A restricted fed group receiving zinc adequate diet was not included because food consumption was approximately 2.5 g/mouse/d for ZA and ChrZD groups. Procedures used in the course of these experiments were approved by the Michigan State University Laboratory Animal Research Committee.


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  		Table 1. Composition of Synthetic Diets for Mice

 
Serum and Diet Analysis
Each mouse was anesthetized between 0800 and 0900 and blood was taken from the subclavian artery within 90 s of cage contact and collected in zinc free microfuge tubes. Sera was obtained and analyzed for zinc using the method of Rajaram [21] using an inductively coupled plasma-atomic emission spectrometer (ICP) (Thermo Jarrell Ash Corp., Franklin, MA) interfaced to a U-5000 ultrasonic nebulizer (Cetac Technologies Inc,. Omaha, NE). Corticosterone (Cs) was quantified using an 125I Cs radioimmunoassay kit (ICN Biomedicals, Inc., Costa Mesa, CA). Quadruplicate samples of ZA and ZD diet were digested in ULTREX II ultrapure nitric acid (Mallinckrodt Baker, Inc., Phillipsburg, NJ) until digestion was complete. Digested diet was decolorized using hydrogen peroxide and analyzed by ICP [22]. National Institute of Standards and Technology corn starch (SRM 8432) and bovine liver (SRM 1577b) standards were digested in parallel with mouse diet to determine the accuracy of zinc analysis. An itrium internal standard addition was applied to all samples to validate ICP data acquisition.

Preparation and Phenotypic Labeling of Thymic and Bone Marrow Cells
The thymus and femurs were collected from each mouse after anesthetization and were processed individually at room temperature. Each thymus was minced, pressed through a 100 gauge stainless steel screen and thymocytes washed with Hank’s buffered saline containing 10 mmol/L HEPES and 4% heat inactivated fetal bovine serum (HIFBS) (harvest buffer) at pH 7.4 to remove cellular debris. Thymus cellularity and cell viability was determined for each harvested cell suspension and the viability was greater than 95% by trypan blue exclusion. Cells were resuspended in Hank’s buffered saline, 10 mmol/L HEPES, 23 mmol/L sodium azide, with 2% HIFBS (label buffer), pH 7.4 and kept at 4° C throughout phenotypic labeling.

Phenotypic analysis of duplicate thymocyte samples was performed using three color phenotyping with anti-CD90.2-PE (Thy 1.2, 30H-12), anti-CD4-PECy5 (clone RM4-5), and anti-CD8a-FITC (clone 53-6.7) at optimal concentrations. Single color samples were used to set electronic color compensation and three color isotype controls were used to correct for false positive antibody binding during data analysis. The labeled samples were held on ice for immediate flow cytometric analysis. All antibodies and isotype controls were from PharMingen (Los Angeles, CA).

Bone marrow from each mouse was gently aspirated into a single cell suspension as previously described [18]. Marrow cellularity and viability (92%–95%) was determined by trypan blue exclusion. Duplicate samples were processed for three color phenotyping at 4° C in label buffer as previously described [18] except labeling was done prior to fixation in phosphate buffered saline (PBS) (1.9 mM NaH2PO4, 8.1 mM Na2HPO4, 0.15M NaCl, pH 7.2) containing 2% methanol free formaldehyde (v/v). Samples were analyzed within 48h. The antibodies used were: FITC-Ly-6C (ER-MP20), biotinyl-CD31(ER-MP12) (Bachem Bioscience, King of Prussia, PA) detected with Streptavidin R670 (Gibco-BRL, Grand Island, NY.), and one PE conjugated antibody: PE-CD45R (RA36B2) or PE-CD11b (M1/70) or PE-TER119 (PharMingen). Single color and two color (Ly-6C with PE-antibody or PE-antibody with CD31) positive and three color isotype controls were prepared in a parallel process and used to set electronic color compensation and as fluorescent negative controls.

Thymocyte Cell Culture and Phenotypic Identification of Apoptotic Cells
Death signals generated in vivo were allowed to complete the process of apoptosis in vitro in a 6 h cell culture in the absence of phagocytic cells as recently described [20]. Preparation of the thymocytes from mice in 45 and 50 day dietary studies were done as outlined above except that cells were harvested under aseptic conditions using dextran charcoal absorbed fetal bovine serum (Hyclone, Logan, UT) in the harvest buffer and culture medium to limit exposure to exogenous glucocorticoids. A separate set of thymocytes from ZA mice were cultured with 1 µmol/L dexamethasone as a standard for hypodiploid fluorescent DNA associated with apoptotic cells.

Phenotypic labeling and analysis was done as described above except that phenotyped cells were resuspended in PBS containing 50% HIFBS and fixed in 50% ethanol. DAPI DNA staining (2.9 µmol/L 4' 6-diamidino-2-phenylindole with 0.1 mmol/L EDTA in PBS, pH 7.4) and flow cytometric analysis to identify apoptotic cells having subG1 fluorescence has been described [20, 23, 24]. Single color antibody or DAPI stained control thymocytes were prepared for use in color compensation and three color isotype matched controls were prepared for use as fluorescent negative controls.

Flow Cytometric Analysis
A Becton Dickinson Vantage flow cytometer equipped with a G3 Mac PC using CELLQuest software (San Jose, CA) and two I-90 argon lasers were used to gather data from phenotypically labeled samples. Antibodies bearing FITC, PE, or PE-Cy5 were excited with 100 mW of 488 nm light with emission detected at 530 ± 15 nm, 575 ± 13 nm, and 670 ± 14 nm respectively. DAPI was excited with 70 mW of 365 nm light from the second argon laser with DAPI emission detected at 450 ± 27 nm. Fluorescent antibody data was collected in log scale while light scatter and DNA cell cycle fluorescence was collected in linear scale.

A light scatter gate excluding cell debris and aggregates was used in all phenotyping. A second gate using log scaling of DAPI fluorescence was used for live/dead discrimination of three color stained thymocytes used for determination of phenotype distribution of viable thymocytes. A DNA doublet discrimination gate was used to acquire cell cycle profiles for normal and apoptotic fixed cells with percent apoptosis determined from cells defined in a subG, DNA fluorescent peak using the method described by Telford et.al. [23, 25]. The phenotypic identity of the apoptotic cells was accomplished with an additional gate for each T-cell population (Pro-T, Pre-T, CD4, and CD8a) as previously described [20]. Data collection on all populations continued until a minimum of 5000 cells were collected for the least frequent gated population.

Statistics
Data were analyzed using SigmaStat (SPSS, Chicago, IL). A t-test was used to compare physical characteristics between ZA and ChrZD groups. ANOVA was performed to compare ZA and ChrZD groups for all phenotypic data. If a statistically significant difference was found, the Dunnett’s test was executed post hoc to identify data significantly different from the ZA mice. Kruskal-Wallis analysis of variance on ranks was applied to nonparametric data of unequal sample size or variance. Differences between ZA and other groups were considered significant at p < 0.05. Means ± SD are given in the text.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
ACKNOWLEDGMENTS
 REFERENCES
 
Physical Characteristics of Chronic Zinc Deficient Mice
The growth profiles for mice in all experiments is provided in Fig. 1 where ZA and ChrZD mice received diet containing 30 mg Zn/kg and ≤ 0.6 mg Zn/kg respectively. Continual weight gain was seen for ZA mice in all four experiments. Chronic ZD mice essentially failed to grow after 14 days even though their food consumption was equivalent to the ZA group. As a result at d 34, 45 and d 50 ChrZD group weights were at 90%, 80% and 74% respectively of weight of the corresponding ZA group. Chronic ZD mice exhibited no signs of the acrodermatitis or parakeratosis associated with zinc deficiency even at d 50.



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  		Fig. 1. Growth of mice fed synthetic zinc adequate and zinc deficient diets. In two experiments mice weighing 16.0 ± 0.7g were divided into two groups receiving either a zinc adequate (30 mg Zn/kg) or zinc deficient (≤ 0.6 mg Zn/kg) diet in a 50 d dietary regime in which mice receiving zinc deficient diet developed ChrZD. Values are expressed as mean ± SD plotted at each growth point for each experiment. For each experiment ZA n = 9 ChrZD n = 8. *significantly different from ZA mice, p < 0.05.

 
Serum zinc dropped 15% by d 34 to 11.7 µmol/L Zn in ChrZD mice. By d 50 serum zinc fell 70% in ChrZD to 4.1 µmol/L Zn. Serum Cs reached 0.46 µmol/L in ChrZD mice by d 34 and remained elevated at 0.49 µmol/L at d 50 while ZA mice maintained serum Cs at 0.18 µmol/L. This indicated that the hypothalamic-pituitary-adrenal stress axis was active during the prolonged reduction in serum zinc in ChrZD mice as has been noted in more acute ZD [20, 26].

To determine the impact of chronic zinc deficiency and an active stress axis on primary immune tissues, the composition of the thymus was examined at d 34, 45 and 50 for change in T cell lymphopoiesis. Bone marrow was examined at d 50 for changes in B cell lymphopoiesis, myelopoiesis and erythropoiesis.

Effect of chronic Zinc Deficiency on T cell Composition of the Thymus
At d 34 no significant change was seen in thymic weight, cellularity (Table 2) or composition due to chronic zinc deficiency suggesting that zinc homeostasis was being maintained in this otherwise zinc sensitive tissue (Table 3) [20]. By d 45 a statistically significant decrease in thymic weight of 42% and cellularity of 49% had occurred (Table 2). However the cellular composition of the thymus remained unchanged (Table 3). The reduced cell number suggested there was a reduction in thymic lymphopoiesis by d 45 but its impact did not preferentially effect any specific thymic T cell subset.


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  		Table 2. Effects of Chronic Zinc Deficiency on Physical Characteristics of Young Adult Mice1

 

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  		Table 3. Effects of Chronic Zinc Deficiency on Composition of the Thymus at Day 34 and 45 of Chronic Zinc Deficiency1

 
By d 50 a 37% and 50% decrease in thymic weight and cellularity had occurred (Table 2). The ChrZD Pre-T cell population had also decreased by 10% (Fig. 2). Pro-T and mature CD4 and CD8a cells exhibited a small but statistically significant increase in proportion among the remaining T cells of the thymus of ChrZD mice (Fig. 2) When reductions in cellularity are also taken into account, it indicated Pro-T and mature T cell numbers had fallen by less than 20% while Pre-T cell numbers had fallen by 50%. Clearly T cell lymphopoiesis was disrupted at d 50.



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  		Fig. 2. Effect of ChrZD on the phenotypic composition of the thymus in mice at d 50. The phenotypic composition of individual mice are presented by T cell subset. Pro-T (CD90.2+CD4–CD8–), Pre-T (CD90.2+CD4+CD8+), CD4 (CD90.2+CD4+CD8–), CD8 (CD90.2+CD4–CD8+). Each bar is the group mean ± SD. ZA n = 9, ChrZD n = 8. *significantly different from ZA mice, p < 0.05."

 
Apoptotic Response of the Thymus to Chronic Zinc Deficiency
To evaluate the role of apoptotic cell death in the reduction in thymic cell numbers and loss of Pre-T cells, thymocytes were harvested from ZA and ChrZD mice at d 45 and d 50 and placed in culture to allow cells induced into apoptosis to complete the process in vitro in the absence of phagocytic cells that quickly remove dying cells. At d 45 the percentage of apoptotic cells present in ZA thymic subpopulations were: 7.7 ± 0.8% Pro-T, 16.4 ± 4.2% Pre-T, 4.5 ± 1.2% CD4 and 4.8 ± 1.4% CD8 cells. The percentage of apoptotic cells in ChrZD thymic subpopulations were: 5.7 ± 0.5% Pro-T, 18.3 ± 3.2% Pre-T, 3.0 ± 0.8% CD4, and 3.7 ± 0.7% CD8 cells. Not surprisingly the percentage of apoptotic cells in each T cell subset in ChrZD mice at d 45 was not significantly different than ZA mice (p < 0.05). Therefore at d 45 no specific thymic subpopulation was altered, rather all subpopulations continued to decrease proportionately as the ChrZD thymus underwent atrophy. However at d 50 Pre-T cell apoptosis increased 60% in ChrZD mice in comparison to the normal rate of cell death seen for ZA mice (Fig. 3). This indicated that the greater loss of Pre-T cells in ChrZD mice at d50 was due to apoptosis. ChrZD apoptosis among Pro-T cells increased by a smaller amount (18%) with normal cell turnover seen for CD4 and CD8a mature cells. Since apoptotic losses were smaller for Pro-T, CD4, and CD8a cell populations; percentage gains seen in Fig. 2 for these populations were accounted for by the diminution of the larger Pre-T cell population. In summary, by d 50 heighten apoptosis, especially in Pre-T cells, contributed significantly to altered T cell lymphopoiesis.



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  		Fig. 3. Assessment of apoptotic cell death in thymic cell populations at d 50 in ZA and ChrZD mice by flow cytometry. Apoptotic cell death for individual mice are presented by T cell subset: Pro-T (CD90.2+CD4–CD8–), Pre-T (CD90.2+CD4+CD8+), CD4 (CD90.2+CD4+CD8–), CD8 (CD90.2+CD4–CD8+). Each bar is the group mean ± SD. ZA n = 9, ChrZD n = 8. *significantly different from ZA mice, p < 0.05."

 
The Effect of Chronic Zinc Deficiency on Hematopoietic Processes
Chronic zinc deficiency did not statistically alter marrow cellularity in the femur at d 50 in either of two experiments (Table 2). Since the marrow is the site of development for erythrocytes, B cells, monocytes and granulocytes; absence of change in nucleated cell number cannot be taken to indicate lymphopoiesis or hematopoiesis was unchanged. Therefore, the cellular composition of marrow was evaluated at d 50 (Fig. 4). In replicate 50 d dietary experiments, ChrZD mice gave similar phenotypic results. The only statistically significant change in the marrow was a 35% decrease in the nucleated erythroid cell population in ChrZD mice. Small increases noted for other marrow populations in ChrZD mice were insignificant in Fig. 4. Thus B cell lymphopoiesis and myelopoiesis remained intact in d 50 ChrZD mice. The former is in contrast to significant changes observed in lymphopoiesis during acute ZD [19]. The identity of cells within the marrow compartments were confirmed using the markers TER-119, CD45R, and CD11b for the erythroid, B-cell, and myeloid lineages respectively. Thus, zinc homeostasis appears to have been maintained in the marrow.



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  		Fig. 4. Effect of chronic zinc deficiency on bone marrow hematopoietic composition in mice at d 50. Bone marrow of ZA and ChrZD mice were individually phenotyped to identify five marrow populations: erythroblasts (CD31–Ly-6C–), lymphocytes (CD31+Ly-6C–), granulocytes (CD31–Ly-6C+), monocytes (CD31–Ly-6C++), and committed progenitors (CD31+Ly-6C+). Graphics represent the percentage composition for each population in the marrow with mean ± SD values given. ZA n = 7, ChrZD n = 8. *significantly different from ZA mice, p < 0.05."

 

   DISCUSSION
 
In this study chronic zinc deficiency prevented young adult mice from completing the final stages of growth. Chronic zinc deficiency did not lead to loss of appetite as there was never any significant reduction in food intake through 34, 45 and 50 day dietary experiments. Close examination of Fig. 1 shows an initial weight gain occurred during the first two weeks of zinc deficiency followed by cessation of growth. At d 42 body weight began falling slightly potentially indicating the beginning of catabolism which would free tissue zinc for recycling to other tissues. Serum Cs rose 2.5 fold by d 34 and remained elevated at d 50 of chronic zinc deficiency. Absence of reduced appetite, inanition, alopecia, parakeratosis and a slow decrease in serum zinc in ChrZD is clearly divergent from the development of reduced appetite, inanition, and increasing severity of alopecia and parakeratosis seen in acute ZD as demonstrated by up to 30% loss in body weight, a 50% decrease in serum zinc and a 300% increase in serum Cs content within a 30 d dietary period [18, 19, 27, 28]. Never the less zinc deficiency, as shown by serum zinc content, did develop slowly in ChrZD mice as shown by a 15% reduction in serum zinc at d 34 with a final 70% reduction by d 50. It is important to note both ChrZD and acute ZD initiate significant neuroendocrine changes [26]. Thus ChrZD in mice compares well with symptoms of chronic zinc deficiency in humans where serum zinc levels may fall slowly over a prolonged period and result in growth retardation [6, 29]. Though we emphasize these changes related to zinc deficiency, we recognize that zinc deficiency has pleiotropic effects on leptin and neuroendocrine hormones which may be playing a role here [30, 31].

The thymus of ChrZD mice were unchanged at d 34, but by d 45 an alteration in lymphopoiesis led to equivalent reduction in all T cell subpopulations leading to thymic atrophy. By d 50 accelerated apoptosis and preferential loss of 50% of Pre-T cells resulted in complete disruption of T cell lymphopoiesis and altered thymic composition. Under these same conditions Pro-T, T helper, and T cytolytic cells showed much greater survival and less apoptotic cell death. This is in sharp contrast to acute ZD where there was an overall greater response to zinc deficiency and Cs in 30 d which led to a 50% decrease in thymic weight, an 80% reduction in cell number, and a 40% reduction in proportion of Pre-T cells with 30% of the cells dying by apoptosis [20]. The impact of ChrZD on the thymus was less dramatic at d 34 through d 45, but by d 50 the impact of ChrZD on Pre-T cells and the accompanying accelerated apoptosis was similar to acute ZD. Thus both chronic zinc deficiency and acute ZD can lead to thymic atrophy and T cell lymphopenia seen in human zinc deficiency [16, 32]. Both ChrZD and acute ZD cause preferential loss of Pre-T cells via apoptosis.

In sharp contrast, the marrow of ChrZD mice showed an ability to maintain both B cell lymphopoiesis and myelopoiesis for a prolonged period of time. The B lymphocyte, granulocyte, monocyte and progenitor populations showed no alteration in composition or cell number in the presence of chronic zinc deficiency and elevated Cs. Whole body studies using 65Zn have shown that zinc may be sequestered in marrow when dietary zinc is low [33]. This may explain the ability of the marrow to maintain lymphopoiesis and myelopoiesis. Only erythropoiesis showed signs of disruption with a 35% decrease in the nucleated erythroid population in ChrZD mice that would be sufficient to alter production of erythrocytes.

The lack of change in B cell lymphopoiesis and myelopoiesis in ChrZD is in sharp contrast to acute ZD. In about half the time, the nucleated erythroid compartment was reduced by 26% and B lymphoid compartment was reduced by 50%, primarily in the Pre-B cell population in acute ZD. Conversely, the granulocyte and monocyte populations expanded by 36% and 175% respectively [18, 19].

Erythropoiesis may be the most sensitive hematopoietic function to the presence of elevated Cs found in both ChrZD and acute ZD. Corticosteroids and their receptor play a key role in the regulation of erythroid progenitor self renewal and differentiation [34]. This is accomplished by corticosterone binding the glucocorticoid receptor and cooperating with the erythropoietin and stem cell factor receptors to stimulate erythroid progenitor proliferation and delay terminal differentiation [35, 36]. Thus, any sequestering of zinc in the presence of elevated Cs may allow erythroid cell proliferation and delay terminal differentiation. Cells that fail to differentiate may die resulting in a reduction of the erythroid population. The disruption of erythropoiesis seen here and in acute ZD is complimentary to the observation that anemia accompanies more acute forms of ZD in the human population [5].

It is clear from this study that the thymus has greater sensitivity to zinc deficiency and elevated Cs. The thymus is composed of about 80% Pre-T cells which express little Bcl-2 or Bcl-XL, making them vulnerable to cell death [20, 37]. Thus ChrZD may be limiting zinc required for cell proliferation and differentiation. The elevated Cs in combination with inadequate zinc may enhance Pre-T cell death via apoptosis leading to a more rapid failure of T-cell lymhopoiesis even in chronic zinc deficiency.

It is very important to note ChrZD mice did not develop alopecia, parakeratosis or any external evidence of zinc deficiency including inflammation of mucosal membranes. The gastrointestinal tract is the primary organ for making zinc biologically available to the animal through changing zinc absorption and excretion in response to dietary zinc [11, 38]. Zip family genes and their ZIP and ZnT transport proteins are found in the intestine where they are controlled through transcription and translation by the concentration of zinc [39–43]. Low zinc leads to increased zinc absorption and decreased zinc excretion during zinc deficiency. It appears that assimilation of zinc may have been sufficient in ChrZD mice to support maintenance of B cell lymhopoiesis and myelopoiesis for an extended period of time.

Application of zinc tracer techniques and computer modeling have identified rapid and slow zinc turnover pools including bone which contains 29% of the body’s zinc [44–46]. Zinc is extensively involved in bone formation with bone zinc being slightly elevated when zinc is plentiful [47, 48]. It is also possible that some of this bone zinc [49] may have been released during the period of increasing catabolism after d 42 of ZD, and zinc from the body’s pools may have helped maintain the marrow hematopoietic process and delay the appearance of symptoms of zinc deficiency in this tissue [48]. Alteration of gastrointestinal physiology giving improved recovery of zinc via transport mechanisms discussed plus the possibility of some catabolic zinc provided from bone may explain the development of a more chronic zinc deficiency that allowed for protection of hematopoietic processes. That individuals may be able to maintain lymphopoietic and hematopoietic processes reasonably well for a period of time during low zinc intake was confirmed by Pinna, et al. [50] where moderate zinc deficiency in humans over a 10 week period did not alter peripheral blood mononuclear cell phenotype and many other cellular functions which depend on production of mature immune cell types from thymic and marrow tissues. The exception remains erythropoiesis which may have been more impacted by elevated Cs.

A chronic deficiency in zinc in mice demonstrated that immune functions depending on the thymus will be more quickly affected by the deficiency while innate immunity and T independent B cell functions will remain unaffected for a longer period of time.


   ACKNOWLEDGMENTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 ACKNOWLEDGMENTS
REFERENCES
 
Supported by a grant from the National Institute of Health, DK 52289. The authors thank Michigan State University for support of the flow cytometry facility.

Received March 3, 2005. Accepted October 11, 2005.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 ACKNOWLEDGMENTS
 REFERENCES
 

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