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Department of Biochemistry and Molecular Biology, Michigan State University, East Lansing, MI 48824-1319
2To whom correspondence should be addressed. E-mail: kingl{at}msu.edu.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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KEY WORDS: • apoptosis • lymphopenia • pre-T cells • thymocytes • zinc deficiency • mice
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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,2
). Indeed, lymphopenia and atrophy of the thymus are the hallmarks of malnutrition, common to both zinc deficiency (ZD)3
and protein-energy malnutrition (PEM) (1
–4
). The literature contains ample documentation that lymphopenia leads to reductions in cell- and antibody-mediated responses, thereby enhancing the incidence of infection and disease associated with ZD or PEM (1
–4
). Because ZD and PEM are also part of the pathobiology of chronic diseases such as AIDS, cancer, gastrointestinal disorders, renal disease, sickle cell anemia or alcoholism, these patients have greater opportunistic infections caused by reduced host defense, which compromises their recovery (1
–5
).
Taken together, forms of ZD and PEM affect a substantial portion of the population, making them key health issues and making it important to identify the cause of lymphopenia. Using the young adult mouse as a model system for ZD, our laboratory has found several key characteristics concerning lymphopenia. 1) Reduction in the absolute number of splenic lymphocytes correlated closely with the inability of the deficient mice to mount antibody-mediated responses (6
). 2) Zinc deficiency disrupted marrow B cell lymphopoiesis, which led to losses of 30–70% of pre-B cells, whereas mature B-cells [immunoglobulin (Ig)M+, IgD+] and the early pro-B cells exhibited resistance to zinc deficiency (7
,8
). This resulted in a substantial reduction in the size of the lymphoid compartment as the deficiency advanced. 3) Adrenalectomy, which removes the source of corticosterone (CS), provided substantial protection in ZD mice against thymic atrophy and losses among thymic and marrow precursor lymphocytes compared with sham-operated ZD mice in which losses were high (9
,10
). Finally, 4) levels of CS analogous to those generated during ZD and PEM induced apoptosis in precursor and immature B-cells of the marrow both in vitro and in vivo in a pattern of attrition similar to ZD (11
,12
). Thus, in more severe acute forms of ZD, it was evident that CS played a role in the disruption of lymphopoiesis via loss of precursor lymphocytes in marrow and thymus; the hypothesized mechanism was apoptotic cell death.
Direct evidence of elevated apoptotic cell death in vivo in primary lymph tissue in response to zinc deficiency is made difficult by the rapid removal of apoptotic cells via phagocytosis (13
). Study of the thymus was chosen because it contains a high proportion of pre-T cells (CD4+CD8+) (78–85%) that are readily susceptible to apoptosis and a modest number of phagocytic cells (<5%) (13
). Because the phagocytic cells of the thymus are associated with the organ parenchyma and because ZD mice have elevated corticosterone levels, we hypothesized that ZD mice would show greater apoptosis in thymic pre-T cells in culture where apoptotic cells would finish development and accumulate in the relative absence of phagocytic cells. Thus, thymuses were removed from marginally ZD and ZA mice and thymocytes cultured for 6 h; then the cultures were phenotyped and simultaneously assessed for the presence of apoptotic cells.
The data provided herein will show that ZD did indeed accelerate apoptosis especially among pre-T cells by as much as 300% and led to altered thymic population composition with loss of pre-T cells. This is an important finding because it clearly demonstrates that apoptosis is a major cause of the lymphopenia and thymic atrophy associated with acute ZD. It may also explain the lymphopenia associated with PEM and other forms of malnutrition that lead to elevated glucocorticoids.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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A/J (6 wk old) or CAF1/J (5 wk old) mice were purchased from Jackson Laboratory based on availability (Bar Harbor, ME). Both strains perform equivalently under the dietary regimen (12
). Treatment began when mice weighed 17.0 ± 0.5 g. A biotin-fortified egg white–based diet containing AIN-76 vitamin mix and AIN-76 mineral mix, with or without zinc carbonate, was used and prepared as previously described (7
,8
,14
,15
) using components from Bioserv (Frecnchtown, NJ). Zinc-adequate (ZA) diet (30 mg Zn/kg) or ZD diet (0.5 mg Zn/kg) was consumed by mice ad libitum with daily changes of feed and feed jars. The mice were housed in stainless steel hanging cages in a room maintained on a 12-h light:dark cycle at 23.3°C. The mice were given acidified Milli-Q filtered water (Millipore, Bedford, MA) to reduce Pseudomonas infections. All glass and plastic materials were washed in 4 mol/L HCl to remove zinc. Cages and water bottles were changed biweekly and the mice were weighed weekly. Studies were terminated when the majority of ZD mice were only marginally zinc deficient, which is defined as follows: 1) having 71% of the body weight of ZA mice; 2) showing few signs of alopecia or parakeratosis (1
,6
,8
); and 3) having a partially atrophied thymus at 34–38% the thymus weight of ZA mice. In the experiment highlighted herein, ZA mice weighed 22.3 ± 0.7 g, whereas ZD mice weighed 15.9 ± 0.6 (Table 1
). This is typical of the five experiments used to confirm these results, requiring between 27 and 32 d of mice consuming each diet, including experiments cited in Table 2
(7
,8
). A total of 30 ZA and 41 ZD mice were evaluated individually in the five experiments. Body weight, serum zinc, and thymus weight were used to document ZD. All protocols used herein were approved by the University Laboratory Animal Research Committee of MSU.
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Thymuses from each dietary group were processed, labeled and analyzed individually. Each thymus was weighed and aseptically processed through 180-mesh stainless steel screen into Hank’s buffered saline containing 10 mmol/L HEPES and 4% dextran charcoal absorbed fetal bovine serum (Ab-FBS) (Hyclone, Logan, UT) at pH 7.4. The treated Ab-FBS was prepared to reduce the concentration of steroids in the medium, which could increase apoptosis during culture. The cell suspension was washed and cell density and viability (>95%) were determined using trypan blue dye exclusion. Thymocytes were cultured in triplicate wells for 6 h at 2 x 106 cells/(mL · well) in a 24-well culture plate at 37°C with 5% CO2. Few macrophages were observed (<1%) because most were removed during processing. The culture medium was RPMI-1640 containing 10 mmol/L HEPES, 10 mmol/L sodium bicarbonate, 1 mmol/L sodium pyruvate, 2 mmol/L L-glutamine, 0.1 mmol/L nonessential amino acids, 50 µmol/L 2-mercaptoethanol, and 5% Ab-FBS (Sigma, St. Louis, MO). A separate set of thymocytes from ZA mice was treated with 1 µmol/L dexamethasone (Dex) as standard for initiation of apoptosis and to generate a hypodiploid population for DNA analysis on the flow cytometer (16
).
Fluorescence-activated cell sorting (FACS) analysis.
At the end of the culture period, the cells were washed and placed in label buffer consisting of Hank’s buffered saline containing 10 mmol/L HEPES, 23 mmol/L sodium azide and 2% FBS, pH 7.4, at 4°C. The thymocytes were simultaneously labeled for 30 min at 4°C with phycoerythrin-labeled anti-CD4 (CD4-PE) (clone GK1.5, Pharmingen, San Diego, CA) and anti-CD8a labeled with fluorescein isothiocyanate (CD8a-FITC) (clone 53–6.7, Pharmingen) at optimum concentrations for simultaneous epitope labeling. After two washes, the cells were resuspended in PBS containing 50% FBS and fixed by addition of 70% ethanol to a final concentration of 50%. Samples were stored overnight at 4°C and then stained with a 4'6-diamidino-2-phenylindole (DAPI) DNA staining solution (2.9 µmol/L 4'6-diamidino-2-phenylindole with 0.1 mmol/L EDTA in PBS, pH 7.4) for 1 h before flow cytometric analysis. Single antibody or DAPI-stained control thymocytes were prepared to optimize flow cytometric analysis. Conjugated isotype matched rat Ig-stained thymocytes were used as fluorescent negative controls. The aforementioned Dex-treated thymocytes, along with scatter profiles, were used to identify apoptotic cells that appeared in the hypodiploid region of the cell cycle as shown earlier (16
,17
).
A Vantage flow cytometer using Lysis II software (Becton Dickinson, San Jose, CA) and equipped with argon and mixed gas lasers (I-90 and Spectrum, Coherent Laser Group, Santa Clara, CA) was used to analyze samples. FITC and PE fluorochromes were excited at 488 nm and detected at 530 ± 15 and 575 ± 13 nm, respectively. DAPI was excited using UV lines (50 mW) from the Spectrum laser where fluorescence was detected at 450 ± 27 nm. Single-color positive and isotype-matched negative samples were used to define positive and negative fluorescence and set electronic spectral compensation for each fluorochrome. DAPI fluorescence was not present in detectors used for FITC or PE. DAPI-stained normal and Dex-treated thymocytes were used to identify the DNA fluorescent population including apoptotic cells. CD4, CD8, DAPI-stained control samples were used to identify CD4+CD8+ pre-T cells, CD4+CD8- T-helper cells, CD8+CD4- T-cytotoxic/suppressor cells and a heterogeneous CD4-CD8- progenitor T-cell population (16
,17
). A corresponding DNA cell cycle profile was generated for each phenotypic population. Data were collected until a minimum of 1500 cell cycle events had been obtained for the least prevalent population.
Serum zinc and corticosterone assessment.
Sera from the subclavian artery were collected into acid-washed microtubes and diluted 1:10 in 32 mmol/L HCl to be analyzed for zinc content by flame atomic absorption spectrophotometry (Varian AA-20 Plus, Mulgrave, Victoria, Australia) at 213.9 nm with deuterium background correction. Addition of known concentrations of zinc standards to serum samples (standard addition) resulted in > 90% recovery of zinc by this method per past investigations (7
,8
).
Endogenously produced corticosteroids were analyzed by collecting blood between 0800 and 0900 h within 90 s of contact with the mouse cage using ether anesthesia and subclavian artery bleeding as previously described (10
,12
). Blood was collected into heparinized tubes. Corticosterone was extracted from 30 µL of serum using 600 µL of dichloromethane (Aldrich, Milwaukee, WI) and treated with 100 µL of 0.1 mol/L NaOH. The organic phase was separated by centrifugation and fluorescence developed by the addition of 200 µL of a mixture of three parts reagent-grade H2SO4 and one part absolute ethanol. Fluorescence intensity was determined using a fluorometer with an excitation wavelength of 475 nm and emission of 525 nm. Standard additions demonstrated detection of 85–95% of the serum corticosterone per past studies (10
,12
).
Data shown are means ± SD of the mean unless otherwise noted. Nonparametric data were examined using the Kruskal-Wallis ANOVA followed by Tukey’s post-hoc test. Parametric data were analyzed using Student’s t test. Statistical significance was set at P < 0.05.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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When the ZD mice weighed 
16 g, the experiment was terminated to obtain partially atrophied thymuses (31 d) (Table 1)
. This utilized 80% of the mice in the ZD group, which were analogous to the marginally zinc-deficient mice used in many previous studies (1
,6
–8
). Thus, the coats of ZD mice were ruffled, but the deficiency had not induced parakeratosis or alopecia, characteristic of a more severe zinc deficiency (1
,6
–8
). The thymuses of the ZD mice were 38% the weight of those harvested from ZA mice (Table 1)
. The ZA mice weighed 22 g at termination and had thymuses of 33 mg in weight (Table 1)
. Thymic atrophy in the ZD mice was associated with a decrease in thymic cellularity from 69 million for ZA mice to 13 million for ZD mice or 18.8% of the cellularity of ZA mice. At this point, serum CS had risen in the ZD mice to 3.3-fold that of ZA mice. Serum zinc in ZD mice was found to be 44% of the level in ZA mice. This demonstrates that lymphopenia in primary immune tissue is an outcome of ZD (Table 1)
. Clearly ZD had induced the hypothalamus-pituitary-adrenal axis to produce more CS, which can induce apoptosis (9
,13
). Whether apoptosis had been induced by CS in vivo was investigated by allowing induced thymocytes to complete apoptotic differentiation in vitro where removal of apoptotic cells by phagocytosis is reduced.
ZD substantially increased apoptosis in the pre-T cell population (Figure 1
). The CD4+CD8+ or pre-T cells exhibited a threefold elevation in apoptosis when harvested from the thymuses of ZD mice. Although response to elevated CS was highly variable, the degree of apoptosis in all seven ZD mice, which ranged from 18 to 55% apoptosis, exceeded that of the ZA mice, which ranged from 7 to 13% apoptosis. The latter is the expected range of homeostatic apoptosis for thymocytes in normal mice (13
,16
). The increased pre-T cell apoptosis identified here was found in ZD mice demonstrating an 80% decrease in thymic cellularity.
|
Two additional experiments verified that ZD enhanced apoptosis in CD4+CD8+ cells (Table 2)
. In these experiments, the ZA and ZD mice met the definition of marginal zinc deficiency used for the larger dietary study discussed above. In Experiment 1, the rate of apoptosis among pre-T cells from ZD mice was double that of the ZA mice. In Experiment 2, there was a 50% increase in cell death for CD4+CD8+ cells from ZD mice compared with those from ZA mice. Both represent substantial increases in rates of death for pre-T cells of ZD mice.
In Experiments 1 and 2, the basal level of cell death was 15% for ZA mice, which is slightly above the 10% average degree of spontaneous death for CD4+CD8+ cells (13
,16
,17
). This modest elevation in the basal level of apoptosis for ZA thymocytes reduced the apoptotic cell death differential between the rates of death of the two dietary groups. Several changes were subsequently made that led to the more compelling results shown in Figure 1
. Thymuses were harvested and processed more rapidly to reduce the inadvertent release of glucocorticoids (10
). Commercially prepared Ab-FBS, rather than in-house prepared Ab-FBS, was used in culture media to further reduce any glucocorticoids present in the sera (11
). These changes reduced the background apoptosis to 10% for ZA mice (Fig. 1)
. Low release of steroids from mice at the time of killing or the presence of these chemicals in sera could, in principle, enhance apoptosis in thymocytes from both dietary groups. However, it may have affected the ZA cells to a greater degree. Because elevated concentrations of glucocorticoids were already present in ZD mice, their thymocytes may not have responded as extensively to the small amounts of additional exogenous steroid generated during harvesting or present in cultures.
Changes in phenotypic distribution of thymic T-cells.
The changes in phenotypic distribution of cells of the thymus were also dramatic and are shown in Figure 2
. There was a significant decline of 40% in the proportion of CD4+CD8+ pre-T cells within the thymuses of the ZD mice. This is expected given their heightened degree of apoptosis shown in Figure 1
. The greater survival of mature CD4+ and CD8+ cells over time was shown by their increase from 7 and 3% in ZA mice to 26 and 11%, respectively, in ZD mice. There was also an increase in the proportion of pro-T cells of nearly 70% among the surviving cells of the thymuses of the ZD mice. Early pro-T and pro-B cells express some Bcl-2 and mature lymphocytes express greater quantities of this antiapoptotic protein, which would account for their survival (18
). Taken together, the data indicate that marginal zinc deficiency markedly changes the phenotypic distribution of cells within the thymus, causing substantial losses in the pre-T cell population needed to replenish the peripheral immune system.
|
In Figure 3
, flow cytometric phenotypic and cell cycle data for a thymus from a representative ZA or ZD mouse are shown. FITC-labeled anti-CD8a and PE-labeled anti-CD4 were used to identify pro-T, pre-T, T-helper and T-cytolytic cells using the four gated regions (R2–R5) identified in the contour plot. DAPI was used to stain the DNA of these four populations of cells (R1). Because few cells from the thymus are cycling, most resided in the Go/G1 region of the cell cycle. Considerable work from several laboratories has shown that highly fragmented DNA from apoptotic cells appears in the hypodiploid region of the cell cycle designated Ao (see Fig. 3
DNA histogram) (16
,17
). The significant increase in CD4+CD8+ apoptotic cells in the Ao region and the corresponding decline of cells in Go/G1 region of the cell cycle are evident from the DNA histogram of the ZD mouse (Fig. 3
B). Loss of these cells by apoptosis corresponds to the greatest loss of pre-T cells from contour plot region R3 in ZD mice. Conversely, examination of the R2 and R4 regions of the ZD contour plot indicates an increased presence of cells in these regions. These regions, in which CD4+ helper cells and CD8+ mature cytolytic T-cells reside, increased substantially within the thymuses of ZD mice and showed less apoptosis in their corresponding cell cycle histograms. Thus, multiparameter FACS enabled us to simultaneously evaluate changes in phenotypic distribution and apoptosis.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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,6
,8
) and 3) having a partially atrophied thymus at 34–38% the thymus weight of ZA mice. The ZA group displayed 7–13% apoptotic cell death, which is the expected range for homeostatic apoptosis for thymocytes from normal mice (13
,16
). This rate of cell loss maintains the ZA thymic weight and organ cellularity seen in Tables 1
and 2
and the phenotypic distribution for homeostatic T-cell development seen in Figure 2
A.
Zinc deficiency stimulates the hypothalamic-pituitary-adrenal cortex stress axis to produce corticosterone (9
,10
). The ZD mice had serum zinc reduced by 56% and a 3.3 fold increase in CS (Table 1)
. In three experiments reported herein, ZD accelerated the levels of apoptosis 50–300% among CD4+CD8+ thymocytes from ZD mice (Table 2
and Fig. 1
). This led to a highly altered thymic phenotypic composition (Fig. 2
B) strongly indicative of disrupted T cell lymphopoiesis. Even though our data represent a physiologic snapshot at d 27–32 of a zinc-deficient diet regimen, such increased apoptosis in marginally ZD mice occurring over the course of many days would account for the 70–80% loss in cellularity and thymic atrophy noted in the thymus (Table 1)
and the development of lymphopenia. Interestingly, serum, nutrient or cytokine deprivation in vitro heightened apoptosis in thymocytes and other kinds of cells (13
).
Similar events appear to be occurring in the marrow, the primary site for development of B-cells in adult mammals. Indeed, previous studies showed analogous losses in the proportion and number of B220+IgM- early precursor cells in the marrow of ZD mice (7
,8
). This loss is via an apoptotic mechanism (11
,12
). Thus, apoptotic-driven mechanisms appear to be a major factor in B- and T-cell lymphopenia and thymic atrophy, which are the hallmarks of ZD described in rodents, primates, and humans (1
,4
,6
,18
). It is possible that apoptosis plays a similar role in other kinds of nutritional deprivation, including PEM in which serum glucocorticoids become elevated.
The rather substantial change in thymic phenotypic composition seen in Figure 2
for ZD mice corrolated with changes in Bcl-2 expression during lymphocyte development. It is at the CD4+CD8+ pre-T cell stage of development that T-cell receptor gene rearrangement occurs, generating abnormal and anti-self T cells, which must be eliminated. At this developmental stage, Bcl-2 or Bcl-XL is minimally expressed in pre-T cells, leaving the cells prone to apoptotic cell death in the presence of CS (13
,17
,19
). Pro-T, T-helper and T-cytolytic cells express higher intracellular levels of Bcl-2 or Bcl-XL, providing some protection against CS-induced apoptosis (19
). Thus ZD mice with elevated CS show massive reduction in thymic organ cell number and extensive loss of pre-T cells, whereas the primitive and mature T-cell types show relatively greater survival rates by expanding in proportion within the remaining population. This pattern of developmental stage–specific apoptotic cell loss and survival has been duplicated for developing B cells in bone marrow of ZD mice (7
,8
). Because glucocorticoids are elevated in acute ZD in both humans and rodents (1
,10
), we suggest that the disruption of lymphopoiesis by accelerated apoptosis accounts for the altered phenotypes seen in primary lymphoid organs.
The separation of the effect of suboptimal zinc and chronically elevated glucocorticoids is much more difficult. We have demonstrated that zinc deficiency stimulates the hypothalamic-pituitary-adrenal cortex stress axis to produce corticosterone (Table 1)
and that adrenalectomy provides substantial protection against CS-induced apoptosis in ZD mice (9
,10
). We confirmed, both in vivo and in vitro, that CS at concentrations seen in ZD mice will produce very similar losses of pre-T and pre-B cells (9
–12
), thereby making direct involvement of zinc unnecessary. Therefore, these data indicate a role for glucocorticoids in inducing apoptosis and disrupting lymphopoiesis.
The role of zinc deficiency in inducing the secretion of glucocorticoids does not preclude a role for suboptimal zinc alone. Recently we showed that low (nmol/L) levels of intracellular zinc can induce apoptosis directly in thymocytes cultured in vitro (17
). However, zinc can modulate the ability of glucocorticoids to bind to the cytoplasmic glucocorticoid receptor (20
). Thus, extracellular and intracellular zinc may act synergistically with the stress axis to elevate the apoptotic cell loss described here.
It was interesting to note that ZD and PEM both have pronounced effects on the lymphoid branch of the primary tissues of the immune system (1
,3
). As mammals progress from the well-fed to the starved state, there are many characteristic changes in the metabolism of vital tissues that are part of an adaptive response (5
). Indeed, loss of appetite and protein-energy deficits routinely accompany zinc deficiency (1
,4
). Patients with PEM often have suboptimal zinc status (4
,5
). For more than a decade, researchers have recognized that ZD and PEM can induce the hypothalamic-pituitary-adrenal cortex stress axis, causing substantial neuroendocrine changes (3
,4
). This may be part of the reason that the two types of deficiencies have many immunological similarities. This fact prompts us to think that apoptosis plays a role in thymic atrophy in PEM and perhaps other types of malnutrition, especially those in which ZD affects growth, development, and reproduction (1
,5
,18
).
The experiments reported here did not involve PEM because they ended in marginal zinc deficiency in which both ZA and ZD mice were consuming 3–4 g food/d at the end of the experiment. Because PEM was not a factor and did not influence our experimental results, inclusion of pair-fed mice to investigate the effects of PEM was not required. We have noted the development of PEM in experiments ending in severe ZD status.
These results stimulate future inquiry in three areas. First, we will investigate PEM in mice of normal zinc status to study changes in apoptotic cell death, phenotypic composition and lymphopoiesis in primary lymphoid tissues and correlate these changes with glucocorticoid levels. Second, the relationship between zinc deficiency and disrupted lymphopoiesis in both primary immune tissues indicates a need to evaluate hematopoiesis to determine whether zinc deficiency affects the myeloid and erythroid systems as well. These studies will provide insight into how nutritional deprivation may cause a reshaping of the immune system to maximize immune defense at a minimum of energy input. Third, the clear role of glucocorticoids and apoptosis in the disruption of lymphopoiesis in the primary immune organs suggests possible models of intervention in reversing the consequences of glucocorticoids and in the inhibition of apoptosis. Successful intervention at the level of glucocorticoid action or prevention of apoptotic cell loss could dramatically improve the medical outcome of patients facing nutritional deficiencies.
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FOOTNOTES |
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3 Abbreviations used: Ab-FBS, dextran charcoal absorbed fetal bovine serum; CS, corticosterone; DAPI, 4'6-diamidino-2-phenylindole; Dex, dexamethasone; FACS, fluorescence-activated cell sorting; FITC, fluorescein isothiocyanate; Ig, immunoglobulin; PE, phycoerythrin; PEM, protein-energy malnutrition; ZA, zinc adequate; ZD, zinc deficient.
Manuscript received 18 July 2001. Initial review completed 11 September 2001. Revision accepted 31 January 2002.
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LITERATURE CITED |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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1.
Fraker, P., King, L., Laakko, T. & Vollmer, T. (2000) The dynamic link between the integrity of the immune system and zinc status. J. Nutr. 130:1399S-1406S.
2. Fraker, P. (1999) Impact of nutritional status on the immune integrity. Gershwin, M. German, J. B. Keen, C. eds. Nutrition and Immunology 1999:147-156 Humana Press Totowa, NJ. .
3. Kuvibidila, S., Yu, L., Ode, D. & Warrier, R. P. (1993) The immune response in protein-energy malnutrition and single nutrient deficiencies. Klurfeld, D. M. eds. Human Nutrition: A Comprehensive Treatise 1993:121-157 Plenum Press New York, NY. .
4. Fraker, P., King, L., Garvy, B. & Medina, C. (1993) Immunopathology of zinc deficiency: a role for apoptosis. Klurfeld, D. M. eds. Human Nutrition: A Comprehensive Treatise 1993:267-283 Plenum Press New York, NY. .
5. Chesters, J. (1997) Zinc. Odel, B. Sunde, R. eds. Handbook of Nutritionally Essential Mineral Elements 1997:185-230 M. Dekker New York, NY. .
6. Cook-Mills, J. & Fraker, P. (1993) Functional capacity of residual lymphocytes from zinc deficient adult mice. Br. J. Nutr. 69:835-848.[Medline]
7. King, L., Osati-Ashtiani, F. & Fraker, P. (1995) Depletion of cells of the B-lineage in the bone marrow of zinc deficient mouse. Immunology 85:69-73.[Medline]
8. Osati-Ashtiani, F., King, L. & Fraker, P. (1998) Variance in the resistance of murine early B-cells to a deficiency in zinc. Immunology 94:94-100.[Medline]
9. Fraker, P., Osati-Ashtiani, F., Wagner, M. & King, L. (1995) Possible roles for glucocorticoids and apoptosis in the suppression of lymphopoiesis during zinc deficiency. J. Am. Coll. Nutr. 14:11-17.[Abstract]
10. DePasquale-Jardieu, P. & Fraker, P. (1980) Further characterization of the role of corticosterone in the loss of humoral immunity in zinc-deficient A/J mice as determined by adrenalectomy. J. Immunol. 124:2650-2655.[Medline]
11. Garvy, B., Telford, W., King, L. & Fraker, P. (1993) Glucocorticoids and irradiation induced apoptosis in normal murine bone marrow B-lineage lymphocytes as determined by flow cytometry. Immunology 79:270-277.[Medline]
12. Garvy, B., King, L., Telford, W., Morford, L. & Fraker, P. (1993) Chronic levels of corticosterone reduces the number of cycling cells of the B-lineage in murine bone marrow and induces apoptosis. Immunology 80:587-592.[Medline]
13. Cohen, J. & Duke, R. (1992) Apoptosis and programmed cell death in immunity. Annu. Rev. Immunol. 10:267-304.[Medline]
14. Graf, E. & Eaton, J. (1984) Effects of phytate on mineral bioavailability in mice. J. Nutr. 114:1192-1198.
15. King, L. E. & Fraker, P. J. (1991) Flow cytometric analysis of the phenotypic distribution of splenic lymphocytes in zinc-deficient adult mice. J. Nutr. 121:1433-1438.
16. Telford, W., King, L. & Fraker, P. (1994) Rapid quantitation of apoptosis in pure and heterogeneous cell populations using flow cytometry. J. Immunol. Methods 172:1-16.[Medline]
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Telford, W. & Fraker, P. (1995) Preferential induction of apoptosis in mouse CD4+CD8+
ßTCRloCD3
lo thymocytes by zinc. J. Cell. Physiol. 164:259-270.[Medline]
18.
Golub, M., Gershwin, M. E., Hurley, L. S. & Hendrickx, A. (1988) Studies of marginal zinc deprivation in rhesus monkeys VIII. Effects in early adolescence. Am. J. Clin. Nutr. 47:1046-1051.
19. Gratiot-Deans, J., Ding, L., Turka, L. & Nunez, G. (1993) Bcl-2 proto-oncogene expression during human T-cell development. J. Immunol. 151:83-91.[Abstract]
20. Telford, W. & Fraker, P. (1997) Zinc reversibility inhibits steroid binding to the murine glucocorticoid receptor. Biochem. Biophys. Res. Commun. 238:86-91.[Medline]
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H. J. Hosea, E. S. Rector, and C. G. Taylor Zinc-Deficient Rats Have Fewer Recent Thymic Emigrant (CD90+) T Lymphocytes in Spleen and Blood J. Nutr., December 1, 2003; 133(12): 4239 - 4242. [Abstract] [Full Text] [PDF]
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 Z. Islam and J. J. Pestka Role of IL-1{beta} in Endotoxin Potentiation of Deoxynivalenol-Induced Corticosterone Response and Leukocyte Apoptosis in Mice Toxicol. Sci., July 1, 2003; 74(1): 93 - 102. [Abstract] [Full Text] [PDF]
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Z. L. Huang and P. J. Fraker Chronic Consumption of a Moderately Low Protein Diet Does Not Alter Hematopoietic Processes in Young Adult Mice J. Nutr., May 1, 2003; 133(5): 1403 - 1408. [Abstract] [Full Text] [PDF]
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M. L. Failla Trace Elements and Host Defense: Recent Advances and Continuing Challenges J. Nutr., May 1, 2003; 133(5): 1443S - 1447. [Abstract] [Full Text] [PDF]
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B. Woodward Apoptotic Loss of Thymic Lymphocytes in Acute Murine Zinc Deficiency J. Nutr., March 1, 2003; 133(3): 814 - 814. [Full Text] [PDF]
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P. Fraker Response to Dr. Woodward J. Nutr., March 1, 2003; 133(3): 815 - 815. [Full Text] [PDF]
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L. E. King and P. J. Fraker Zinc Deficiency in Mice Alters Myelopoiesis and Hematopoiesis J. Nutr., November 1, 2002; 132(11): 3301 - 3307. [Abstract] [Full Text] [PDF]
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