MURINE ZINC DEFICIENCY ALTERS T-LYMPHOCYTE SUBPOPULATIONS AND GENE EXPRESSION OF CHEMOK INES AND CYTOKINES IN THE GUTASSOCIATED LYMPHOID TISSUES By KELLI ANN HERRLINGER-GARCIA A THESIS PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORID A IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF MASTER OF SCIENCE UNIVERSITY OF FLORIDA 2006
Copyright 2006 by Kelli Ann Herrlinger-Garcia
iii ACKNOWLEDGMENTS I would like to extend immense appreciation to Dr. Bobbi LangkampHenken, my mentor, boss, and friend. Dr. Bobbi Langkamp-Henken has provided endless support, confidence in my abilities, and encouragement. I could not have pursued a masterÂ’s degree wit hout her. I would also li ke to thank the other members of my committee, Dr. Robert J. C ousins and Dr. Sally A. Litherland, for their time and expertise. In addition, I would like to thank Dr. Harry S. Sitren, my graduate coordinator, for his excellent guidance on numerous occasions. I would also like to thank my fellow students in the laboratory, Shannon DeLucia, Jessica McIntire, Justin Silvestre, and Racquel Ramharrack, for their hours of hard work during the rodent sur geries and for their friendship. I would also like to thank my closest friends, spec ifically Denise Dent on, Connie Brand, and C.J. Nieves, who give constant support and encouragement in this endeavor and all aspects of my life. I would like to thank my parents and sisters who taught me the importance of an education, pride in a job well done, and perseverance. A very special thank you is owed to my son, Austin. He wa s patient and supportive when I spent many hours studying, and he constantly reminds me of what is really important. A final thank you is to God, who makes all things possible.
iv TABLE OF CONTENTS page ACKNOWLEDG MENTS.......................................................................................iii LIST OF TABLES.................................................................................................vi LIST OF FI GURES..............................................................................................vii ABSTRACT ........................................................................................................v iii INTRODUCTION..................................................................................................1 LITERATURE REVIEW........................................................................................3 What is GALT?...............................................................................................3 Cellular Populations in the GALT...................................................................9 T-Cell Signaling and the GA LT.....................................................................13 Cytokines and C hemoki nes.........................................................................17 Cytokines, Chemokines and T heir Importance in GA LT..............................19 Overall Immune Responses Du ring Zinc De ficiency....................................20 Zinc Deficiency and Gene Expres sion.........................................................23 Zinc Deficiency and Cellular Ch anges.........................................................25 Zinc Deficiency and Cy tokine Res ponses ....................................................32 Zinc Deficiency and Gut Associat ed Lymphoid Ti ssue (GAL T)....................34 Zinc and T-Cell Signalin g.............................................................................38 Purpose of th is Work....................................................................................40 MATERIALS AND METHODS............................................................................43 Animal s........................................................................................................43 Determination of Serum Zinc Concent rations ..............................................44 Lymphocyte Extraction from Intestinal Tissue..............................................44 Flow Cyto metry............................................................................................47 Isolation of RNA...........................................................................................48 Microarray....................................................................................................50 Quantitative Real-Tim e PCR (q RT-PCR) ....................................................50 Statistical Analys is.......................................................................................52
v RESULT S...........................................................................................................54 Mouse Wei ghts............................................................................................54 Organ Wei ghts.............................................................................................54 Zinc Status Assessm ent...............................................................................55 Flow Cyto metry............................................................................................55 Microarray A nalysis ......................................................................................61 mRNA Expression of Chemoki ne Ligands and In terleukin s.........................62 DISCUSSI ON.....................................................................................................65 SUMMARY AND CO NCLUSION S.....................................................................88 APPENDIX A DIET COMP OSITION..................................................................................89 B MOUSE INFLAMMATORY CYTOKINE AND RECEPTORS MICROARRAY KEY.....................................................................................91 C WEIGHT DATA OF ZINC ADEQUATE, ZINC DEFICIENT, AND PAIRFED DIET GROUPS ....................................................................................90 LIST OF REFE RENCES..................................................................................107 BIOGRAPHICAL SKETCH ...............................................................................124
vi LIST OF TABLES Table page 1 Normal GALT cellular populations ......................................................................12 2 Q-RT PCR primer sequences ..............................................................................52 3 CD8 + and CD4+ populations. .............................................................................60
vii LIST OF FIGURES Figure page 1 Gut-associated lymphoid tissue and i mmune cells of the intestinal villus ......8 2 Percent baseline weight of BALB/c mice fed a zinc-adequate or zincdeficient diet ...........................................................................................................54 3 Serum zinc concentrations and col onic metallothionein mRNA levels.. .......55 4 Total recovered cell numbers in GALT ...............................................................56 5 Colonic CD3+ intraepithelial lymphocytes in BALB/c mice. .............................58 6 Colonic intraepithelial and small intestinal CD3+ TCR CD8 +...................58 7 Small intestinal Peyer's patch CD3+ TCR + CD8 -.......................................59 8 Mesenteric lymph node CD3+ CD28+ lymphocytes ..........................................61 9 Colonic gene expression. .....................................................................................63 10 Colonic chemokine/cyt okine/receptor mRNA ....................................................64 11 Mechanism for CD4 and/or CD8 cellu lar changes during zinc deficiency ....83 12 Proposed model .....................................................................................................84
viii A Thesis Presented to the Graduate School of the University of Florida in Partial Fulf illment of the Requirements for t he Degree of Master of Science MURINE ZINC DEFICIENCY ALTERS T-LYMPHOCYTE SUBPOPULATIONS AND GENE EXPRESSION OF CHEM OKINES AND CYTOKINES IN GUTASSOCIATED LYMPHOID TISSUES By Kelli Herrlinger-Garcia August 2006 Chair: Bobbi Langkamp-Henken Major Department: Food Science and Human Nutrition Zinc deficiency alters lymphopoeisis and TH1/TH2 cytokine balance. Although these cytokine changes ma y alter lymphocyte subpopulation distribution or homing, no studies have examined this outcome. BALB/c mice (4week old) were fed a zincdeficient (ZD, <1 mg Zn/kg) , zinc-adequate (ZA, 27 mg Zn/kg), or pair-fed (PF) diet for 9 weeks. Percent baseline weight did not differ among the 3 groups in this TH2-dominant mouse strain ; therefore, PF was dropped from further analyses. Serum zi nc (ug/mL) decreased with progressive zinc deficiency and at week 9 was 0.7 Â± 0.1 (ZA) versus . 0.3 Â± 0.2 (ZD, mean Â± SEM, P = 0.001). Gut-associated lymphoi d tissue (GALT) phenotypes were evaluated at 3, 6, and 9 weeks by fl ow cytometry with anti-CD3, CD8ÃŸ, TCR , CD4, and CD28 antibodies. ZD CD3+ colonic intraepithelial lymphocytes (cIEL) normalized to the ZA group increased with progressive zinc deficiency ( P = 0.06), and the percentage of CD3+ T cells was higher at week 9 in ZD (43 Â± 4%) versus
ix ZA (29 Â± 3%, P = 0.04) mice. CD3+ CD8ÃŸ+ TCR cIELs were elevated at all time points in ZD mice ( P = 0.008) and may account for the T-cell increase. This cell population was lower at all time point s in lymphocytes isolated from PeyerÂ’s patches ( P < 0.01). PeyerÂ’s patch CD3+ TCR + CD8 were elevated as a percentage of total cells ( P < 0.05) and as a percentage of CD3+ T cells ( P = 0.002). CD4+ populations and subpopulations were lower as a percentage of CD3+ T cells in ZD versus ZA in cIEL (CD4+, P = 0.02) and PeyerÂ’s patches (CD4+ CD8+, P = 0.003). Populations expressing CD28+ decreased as a percentage of CD3+ T cells in PeyerÂ’s Patches (CD3+ CD28+ CD4CD8 +, P < 0.05), cIEL (CD3+ CD4+ CD28+, P < 0.001), and messenteric lymph nodes (CD3+ CD28+, P = 0.04) in ZD mice. At week 9, colonic inflammatory cytokine and receptor transcript abundance was measur ed using a microarray followed by q RT-PCR for confirmation. Microarray analysi s indicated that CCL25 (TECK) was overexpressed in ZA whereas IL-18 was overexpressed in ZD colon. q RT-PCR confirmed that normalized CCL25 mRNA levels were different between diet groups, whereas IL-18 mRNA levels were unchanged. However, IL-18 transcript levels positively correlated wit h the percentage of ZA and ZD CD3+ CD8ÃŸ+ TCR cells. These data suggest that ZD alters GALT lymphocyte subpopulation distribution, CD28+ populations, and a chemokine li gand important in lymphocyte homing.
1 INTRODUCTION Zinc, a mineral that is important in t he maintenance of health, functions in the body to support cell growth, cell r eplication, gene expr ession, protein metabolism, lipid metabolism, and imm une function, and is also an essential cofactor for more than 200 enzymes. T he body contains between 1.5 and 2.5 g of zinc, and requires approximately 8 to 11 mg/day (1). Early studies in the Middle East identified the relationship bet ween zinc deficiency, short stature and subsequent susceptibility to infectious diseas es (2,3). The initial belief that zinc deficiency was isolated to malnourished in dividuals in underdeveloped countries was abolished when later studies in the Un ited States revealed that pregnant women, low-income children, and elders we re also at risk for zinc deficiency (49). In addition, individuals suffering from chronic ill nesses also were found to have zinc deficiency further exacerbati ng their conditions (4,10). These zincdeficient individuals are ther efore at risk for developing complications resulting in an impairment of their immune responses and a general decline in health status. Many studies link zinc deficiency with increased diarrhea in mice and humans, suggesting disturbance of the intest inal barrier; however , little is known of the impact of zinc deficiency on muco sal immunity (4,11,12). Zinc deficiency is associated with a system ic imbalance between TH1 (IL-2, IFN, TNF, and IL12) and TH2 (IL-3, IL-4, IL-5, IL-6, IL-10, and IL-13) cytokines (13,14). Changes in cytokines, chemokines, and/or their receptor s, if perpetuated into the intestinal
2 tissue, may be one mechanism for pr omoting disturbances of immune populations within the intestinal muco sa. Chemokines, cytokines, and their receptors are essential in gut-associat ed lymphoid tissue (GALT) maintenance and recruitment of cells during an imm une response. Zinc deficiency is also associated with alterations of peripher al, thymocyte, and splenocyte cell populations (4,15-17). Mice exposed to zi nc deficiency have been shown to have fewer pre-T, pre-B, and alterations in T-hel per to T-cytotoxic cell ratios (15,16). Alterations in cytokine responses and lymphocyte populations of ZD mice may inhibit the ability of these mice to main tain and protect their GALT and lead to the development of inflammation. The objective of this study was to exam ine the effects of a progressive zinc deficiency on GALT lymphocyte populati ons and gene expression of cytokines and chemokines in mouse colonic tissue. We propose that zinc status affects the lymphocyte populations and balance of c hemokines, cytokines and/or their receptors, important in maintenance of the GALT. The mucosal immune system is a natural barrier and a major route of entry for invading pat hogens; therefore, maintaining its integrity is critical for prot ection of the body from intestinal flora and potential pathogens. A clear understanding of the changes that occur in the GALT during zinc deficiency, includ ing cytokines, chemokines, and their receptors, and cellular populations ma y suggest mechanisms through which immune function is altered to allow su sceptibility to infectious diseases.
3 LITERATURE REVIEW What is GALT? The gut-associated lymphoid tissue (GALT) is composed of the lymphoid rich regions including the mesenteric lymph nodes, Peyer's patches, epithelium, and the lamina propria. GALT contains 106 lymphocytes/g tissue as reviewed by Jabbar et al. (18) making it the larges t lymphoid organ in the body, with total lymphocytes outnumbering even that found in the spleen. In healthy individuals, the GALT is responsible for ensur ing that an adequate immune response is generated following a pat hogenic exposure and at the sa me time preventing the many non-pathogenic dietary antigens, whic h are ingested daily, from evoking an immune response. The epithelium (compos ed of individual epithelia) covers finger-like projections that protrude into the lumen and enhance the surface area of the intestine and allow for greater absorbative capacity. The epithelium, a single layer of cells, forms a barrier of protection from the ex ternal environment and protects the adult human mucosal su rface, which extends 200 to 300m2 (19). Along the villi, interspersed between the epithelial cells at a 6 to 10:1 (epithelial:intraepithelial cell ratio) are lymphocytes called intraepithelial lymphocytes, usually CD8+ T cells (18). Epithelial cells have absorbative capacity, antigen-presenting capabilities, and transport IgA (the major protective gut antibody) into the lumen. At the base of the villi are crypts, containing stem cells that will develop into either epithelia l, goblet (mucus-secreting), or paneth
4 cells (which secrete antimicrobial agent s). The lamina propria is beneath the epithelial cells, and is a dense network of lymphatic and blood vessels. This area contains dendritic cells, macrophages, pl asma cells and their precursor B cells, and CD4+ and CD8+ lymphocytes. Below the lamina propria are several muscle layers that provide structure and mec hanical function, and are important for peristalsis. PeyerÂ’s patches are present throughout the intestine, although they are present in higher numbers in the sma ll intestine. Peyer's patches are lymph node-like structures and are the sites fo r the initiation of immune responses within the small intestine. Peyer's patc hes contain zones of T cells and B cells, and interactions between these two types of cells occur in areas called germinal centers. Cellular populati ons found within the PeyerÂ’s patches include CD4+ and CD8+ T cells, B cells, macrophages, and dendritic cells. The epithelial side of the Peyer's patch contains specialized epithelia l cells called microfold cells (M cells), which do not secrete mucus and allow for more direct contact with particles in the lumen of the intestine. The intestines dr ain into a series of lymph nodes called the mesenteric lymph nodes. Like the Pe yerÂ’s patches, mesenteric lymph nodes have Tand B-cell zones and germinal cent ers. Mesenteric lymph nodes function to filter antigen out of the lym ph and are composed of T and B cells, macrophages, immature dendritic cells and mature dendritic cells. Each of these GALT tissues has a specific function to pr otect the mucosal barrier from invasion of pathogens. Both the innate and adaptive branc hes of the immune system are represented in the GALT and play import ant roles in the protection of the
5 mucosal barrier. The innate, first line, of defense includes the normal peristalsis or movement of food through the digestive tract, digestive secretions (enzymes, gastric secretions, bile, and mucus), the epithelial barrier itself, phagocytes (macrophages and neutrophils), natural kill er cells, and complement proteins. These defense mechanisms are already pr esent or available within hours after antigenic exposure. The adapt ive components of the i mmune system take longer (days) to provide protection but prov ide a stronger, more specific response following an antigenic event. Component s of the adaptive immune system include the B lymphocytes, which develop into antibody secreting plasma cells, and T lymphocytes, which differentiate into T helper (CD4+), T cytotoxic (CD8+), or suppressor/regulatory T cells. A fracti on of these T, B, and plasma cells will become memory cells, which enable the adaptive immune responses to provide a faster and amplified response upon r epeated exposure to the same antigen. Whereas the innate immune response is e ffective at nonspecific clearance of a large number of challenging agents, the adaptive immune components are responsible for antigen specif ic elimination of pathogens. The innate and adaptive immune system s work together to remove pathogens from the GALT. Pa thogens that are able to penetrate the epithelial surface will encounter cells of the i nnate immune system such as macrophages and neutrophils, which will release chem okines, cytokines, and complement factors in an attempt to control an infection. The chemokines, cytokines, and complement factors will recruit other i nnate system cells into the area or opsonize the pathogen to make it more easily phagocytosed. If this local
6 inflammatory response is not able to clear the pathogen, the specific immune response will take over. The innate response will not be covered in depth here because the cell populations evaluated in this study are a part of the adaptive immune responses. The adaptive immune response begins when an antigen, unable to be cleared by the innate response, is tak en up by an antigen-pres enting cell-usually a resident immature dendritic cell. Wei ner et al. (20) de monstrated that particulate antigens are preferentially tak en up by the M cells (overlying PeyerÂ’s patches); whereas, soluble antigens are ta ken up either by the villus epithelial cells or by passing through the tight junctions between epithelial cells (paracellular movement). Furthermore, Nie ss et al. (21) showed that dendritic cells have the capacity to sample luminal contents by extending tentacle-like projections across the basolateral memb rane between the tight junctions via a chemokine receptor-CX3CR1, dependent process. M cells can endocytose antigens and transport them across the cell to antigen-presenting cells and lymphocytes within the PeyerÂ’s patches . The antigen presenting cell (dendritic cells or macrophages) will then take up ant igen either by pinocytosis or phagocytosis. NaÃ¯ve B and T cells enter the PeyerÂ’s patches (which do not have afferent lymphatics) and la mina propria via interaction of L-selectin and 4 7 on their surface with mucosal adressin cell ular adhesion molecule 1 (MAdCAM-1) on the blood vessel endothelium (22). Wi thin the PeyerÂ’s patches antigen is presented by the antigen-pr esenting cell to B and T lymphocytes which are then activated, proliferate, and travel to the lamina propria and eventually to the
7 mesenteric lymph nodes. In addition to enter ing the lamina propria by way of the PeyerÂ’s patches, antigen can also enter the lamina propria via the villus epithelium directly or paracellularly to interact with antigen presenting cells (macrophages or dendritic cells). The epithelia l cells themselves can present this antigen to antigen presenting cells residi ng in the lamina propria (23). Antigen presenting cells within the lamina propria activate lymphocytes present. Activated lymphocytes and antigen-presenting cells travel from the lamina propria and enter the mesenteric lymph nodes through the afferent lymphatic vessels. The organization of the mesenteric lym ph nodes promotes interactions between antigen presenting cells and T cells, B cells and antigen, and activated T cells and antigen exposed B cells. The lymphocytes differentiate and proliferate within the mesenteric lymph node, leave through t he efferent lymphatics, and eventually dump back into the bloodstream at the thoracic duct (Figure 1). NaÃ¯ve T and B lymphocytes travel to lymphoid organs (such as mesenteric lymph nodes and PeyerÂ’s patches); w hereas, antigen-activated T cells can migrate to non-lymphoid sites. Several studies have shown that GALT derived dendritic cells generate CD4+ and CD8+ cells that preferentially home back to the gut via expression on their surface of chem okine receptors that are specific for ligands expressed withi n the GALT (24-27). CD4+ T cells activated within mesenteric lymph nodes expressing chemok ine receptor 9 (CCR9, whose ligand chemokine ligand 25 is expre ssed by epithelial cells) and 4 7 along with
8 AB AB Figure 1. Gut-associated lymphoid tissue (A) and immune cells of the intestinal villus (B). Samples of the cont ents of the intestinal lumen are transported across the specialized epi thelial cells covering the Peyer's patch and presented to lymphocytes (A). Activated lymphocytes leave the Peyer's patch and travel to the mesenteric lymph nodes where they continue differentiating and dividing . The lymphocytes eventually drain into the systemic circulation via t he thoracic duct and home back to the intestinal mucosa and other muco sal tissues. Lymphocytes populate the lamina propria and in traepithelial spaces withi n the intestinal villi (B). Reprinted from Langkamp-Henken, B., Glezer, J.A ., Kudsk, K.A.. Immunologic structure and function of the gastrointestinal tract. Nutr. Clin. Pract. 1992;7:100-108, with pe rmission from the American Society for Parenteral and Enteral Nu trition (A.S.P.E.N.). A.S.P.E.N. does not endorse the use of this material in any form other than its entirety (28) several other chemokine receptors were isolated within the la mina propria (24). Steinstad et al. (24) showed that CCR9 def icient cells transferred into recipient animals had reduced presence in the la mina propria. Johansson-Lindbom et al. (25) showed that mesenteric lymph node de ndritic cells, but not splenic derived dendritic cells, generated 4 7, CCR9, CD8+ T cells. Injection of this cell population into congenic mice showed that this T-cell population was localized to the small intestinal epithelium (25). A late r study by the same investigators, found that the generation of 4 7+ CCR9+ CD8+ T cells which specifically homed to the GALT was dependent on dendritic cells bearing the CD103+ marker (26).
9 Dendritic cells isolated from PeyerÂ’s patches, and not from spleen or peripheral lymph nodes, also have the ability to stimulate expression of gut homing 4 7 and CCR9 markers on CD8+ T cells (27). The author s termed the ability of dendritic cells to mark the cells which t hey activate for later homing back to the specific tissue in which they were origina lly activated, Â“selective imprintingÂ” (27). These data demonstrate the ab ility of gut-derived dendritic cells to influence the trafficking of lymphocytes back to their areas of activation through the use of chemokine receptors (which will be discusse d in a later section). The return of activated lymphocytes to the GALT tissue in which they were activated allows for a more effective immune response agai nst the pathogen that has invaded a specific area. Cellular Populations in the GALT The T and B lymphoid cells found in the GALT share similarities and differences with those found throughout t he rest of the body. T-cell receptors (TCR) on either CD4 or CD8 positive T cells are typically heterodimers composed of an alpha and beta chain (TCR ) or of a gamma and delta chain (TCR ). In general, the dominant TCR in most of the body is TCR . The TCR T cell has less genetic diversity than TCR T cells and are structurally similar to an immunoglobulin molecule (29).The TCR only accounts for 3.8% of TCR in the peripheral blood of healthy adults (30), but their ti ssue distribution is specifically localized. TCR represent as much as 43% of CD45+ intraepithelial leukocytes (31). The PeyerÂ’s patches in the GALT have T and B cell rich zones, the T-cell zones are compos ed of approximately 65% CD4+ TCR and about 30% CD8+ TCR (32). The B cells in the ge rminal centers of the PeyerÂ’s
10 patches produce predominant ly the antibody isotype IgA and some IgM, and therefore contain a high perc entage of IgA positive B cells (32). Epithelial cells of the gut transport antibody across the cell vi a receptors. Antibody is then released into the gut lumen and a portion of t he receptor remains attached to the antibodies providing stabi lity within the lumen. This secretory component functions to keep the antibody in close pr oximity to the mucus layer and protects against antibody proteolysis by digestive enzymes, the IgA released into the lumen is therefore known as secret ory IgA (sIgA). T-cell populations in mesenteric lymph nodes are mainly composed of TCR + T lymphocytes with a small fraction containing the al terative T-cell receptor, TCR . Like PeyerÂ’s patches and mesenteric lymph nodes, the population of TCRs in lamina propria is also similar in phenotypic distribution to the body, consisting of the previously mentioned CD4+ and CD8+ T-cell populations and B cells. Manzano et al. (33) report that CD45+ leukocytes isolated from t he lamina propria are 67% TCR and 7.5% TCR . Intraepithelial lymphocytes (IEL) residing between the epithelial cells (entero cytes) are mainly CD8+, and comprise four main populations, all of which contain a CD8 homodimer (34). Some cells expressing the CD8 homodimer have been found to have an increased production of cytokines (35). Two populations include the CD4+ and CD8 + TCR T cells, similar to the more convent ional T cells of the PeyerÂ’s patch, mesenteric lymph nodes, and lamina pr opria, except also bearing the CD8 homodimer. TCR T cells are present in high amounts in the mucosal epithelium (34, 2005 #71). This population of TCR is CD4 negative and the
11 conventional CD8 heterodimer negative, but has a CD8 homodimer instead (34). The final main population of T cells composing the intraepithelial lymphocyte population have the TCR but are CD4 negative and the conventional CD8 heterodimer negative, but have a CD8 homodimer (34). Manzano et al. (33) report the TCR composing 45%, whereas the TCR compose 43% of CD45+ intraepithelial leukocytes in 7-week BALB/C mice. A difference between mice and humans is the lack of the TCR that are CD4 and the conventional CD8 heterodimer negative, but have a CD8 (in humans these are present in the human fetal period only) (34). TCR cells, which compose a larger percentage of intraepi thelial lymphocytes, appear to have a role in activation of apoptosis in damaged epithelial cells (35). Cellular populations found in GALT tissues are lis ted in Table 1. All of these cell populations and components of the GALT provide important immune functions that serve to create an effective barrier against potential pathogens.
12Table 1-Normal GALT cellular populations Author (reference) Species/Str ain Gated Population Tissue CD3 (%) CD8 (%) CD8 (%) CD4 (%) TCR (%) TCR (%) Fujihashi (36) Mouse/ BALB/C None (% total cells counted) MLN 81.6 PP 34.7 3.3 7.6 36.1 43.4 0.7 IEL 89.4 15.2 69.1 7.7 45.4 43.3 Manzano (31) Mouse/ BALB/C CD45+ Leukocytes LPL 77.9 17.6 37.1 36.3 66.9 7.5 Szczypka (37) Mouse/ BALB/C None (% total cells counted) MLN 43.4 16.5 26.5 IEL 65.0 26.0 64 3 49 16 Helgeland (38) Rat None (% total cells counted) MLN 55 8 9 47 54 1 Fujihashi (39) Mouse/ C3H/HeN CD3-None: Others presented as % of CD3+ IEL 86.4 74.5 7.6 Laky (40) Mouse/ C57BL/6J X 129 Ola CD3+ cells IEL 18-32 Abbreviations: mesenteric lymph nodes, MLN; PeyerÂ’ s patches, PP; intraepithelial lymphocytes, IEL; lamina propria lymphocytes, LPL
13 T-Cell Signaling and the GALT TCR T cells bearing CD4 or CD8 require at least two signals for activation. The activation of T cells bearing the TCR and TCR have both similar and dissimilar characterist ics. The primary signal for TCR is the interaction between the CD4 or CD8 molecule/TCR complex and the antigen/major histocompatib ility (MHC) complex on t he antigen presenting cell. The antigen-specific TCR + T cells recognize that antigen in complex with the major histocompatibility prot ein and is stabilized by either the CD4 molecule (in the case of the MHC class II) or the CD8 molecule (for the MHC class I). The secondary signal for TCR T cell is often an interaction between CD28 on the T cell and CD28 ligands [B7.1 (CD80) or B7 .2 (CD86)] on the antigen presenting cell. CD28 is a constitutively expre ssed receptor on T cells, but can be upregulated at both mRNA and protein le vels following TCR/CD3 stimulation (41), but decreases in aging (42). B7 -CD28 and MHC-TCR activation of the TCR + T cell promotes IL-2 secretion and in creases the cells responsiveness to IL-2. TCR-MHC complexes occur in lip id rafts formed within the T-cell membranes, which bring CD28, CD3, and CD4 or CD8 receptors in close proximity on the T-cell surface (43). The TCR is associated with a complex of the proteins CD3 and CD4 or CD8, this resulting TCR complex has cytoplasmic regions that are associated with Src-fam ily protein kinases. Upon receiving both TCR complex and CD28 signals and formation of the lipid raft, the Src-family protein kinases p59fyn (Fyn) and p56lck (Lck) are activated by removal of inhibitory phosphate groups. The formation of the lipid raft brings the activated p56lck associated with CD4 or CD8 in close pr oximity to itÂ’s downstream substrate
14 on the CD3 TCR, ZAP-70, beginning a signa ling cascade via phosphorylation of ZAP-70 that culminates in the activation of transcription factors in the nucleus, such as NFB, initiating gene transcription, di fferentiation, and proliferation. The signaling of the T cells bearing TCR is less well characterized. Like TCR T cells, data from Sperling et al. (44) demonstrated that TCR T cells also need 2 signals for activation: the antigen-TCR /CD3 complex and B7CD28. The TCR /CD3 complex does not require major histocompatibility proteins on antigen-present ing cells for recognition of antigen (45). Like their TCR counterpart, CD28 was expressed at low levels until stimulation at which point surface expression of CD28 increased (44). Incubation of TCR T cells with an antigen bearing cell li ne, with and without B7 showed that absence of CD28 costimulation prevented TCR T-cell proliferation and IL-2 production (44). As for TCR T cells, the addition of exogenous IL-2 to TCR T cells and antigen bearing cell lines without B7 in vi tro could replace CD28-B7 signaling as a costimulation (44). Sperling et al . (44) showed that a conventional B7+ APC provided costimulation via CD28 to the TCR T cell; however, nonconventional antigen presenting cells could activate TCR T cells using BB-1, a different CD28 ligand or via antigen presentati on with a non-traditional MHC molecule (46). Sperling et al. (44) proposed that the increased presence of TCR T cells in the epithelial tissues may reflect thei r increased potential to be activated by nonconventional antigen presenting cells , such as epithelial cells. The differences obtained in potential signali ng mechanisms and localization may be due to the presence of di fferent subsets of TCR T cells (47). Two main types of
15 TCR T cells have been reported: V 1 and V 2 (47). TCR T cell V 1 express CD57 and are CD5-/lo, CD28-, while TCR T cell V 2 are CD5hi CD28+ CD57(47). TCR T cell V 2, V 9 subpopulations are present in higher numbers in human peripheral blood (48,49), while TCR T cell V 1 have been shown to be present in higher numbers in intestinal intraepithelial lymphocytes (30). The TCR, like the TCR, is associated with the CD3 protein complex, and forms clusters after signaling to trigger t he downstream signaling cascade (29). Arosa et al. (50) proposed that CD28 is an essential costim ulatory molecule of T cells and stimulation of T cells in t he absence of this costimulatory molecule can lead to T-cell anergy. CD28 costimulatio n is necessary for the formation of the lipid raft around the TCR complex site (51). Jordan et al. (52) proposed that the mechanism for this CD28 function was through cytoskeletal rearrangement. Tavano et al. (53) showed that a CD4+ Jurkat T-cell line incubated with antigen stimulated antigen presenting cells, with and without B7 for cosignal engagement, resulted in Lck accumulati on and localization at the lipid raft (immune synapse) only when the CD 28 molecule was engaged. This phenomena was also seen in the peripheral blood lymphocytes of elders. CD4+ T cells from elders demonstrated 5.5-fold decr eased recruitment of Lck to lipid rafts compared to CD4+ T cells from young individual s following stimulation and only a 34% association of CD28 in the lipid rafts from the cells from the elders versus 71% in the cells from the young (54). Larbi et al. (54) concurred with Tavano et al. (53) and concluded that ligation of the TCR and CD28 induced the recruitment of p56lck to the lipid raft. CD4+ T cells stimulated in the absence of CD28
16 stimulation were unable to avoid clonal anergy as determined by lack of IL-2 production and cellular prolif eration (55). Kundig et al. (56) showed that a transient signal at the TCR in CD28 deficient mice anergized CD8+ T cells leading to a reduction in proliferat ion and cytotoxic activity. The cellular populations evaluated in the studies above did not specify whether the TCR was of or origin; however, the st udies used peripheral CD8+ or CD4+ populations or CD4+ Jurkat cell lines, so most likely these conclusions may be pertinent primarily to TCR and select subsets of TCR populations. Furthermore, two separate studies found that CD28 knockout mice failed to form germinal centers in their PeyerÂ’s patches (57) and CD28-/mice had decreased numbers of PeyerÂ’s patches than heterozygous CD28+/C57BL/6 mice (58). Work by Arosa et al. (50) suggested the CD8+ lymphocytes within the GALT are predominantly CD28-. In this study, adult human peripheral blood CD8+ CD28+ lymphocytes cultured with colonic epithe lial cells resulted in the expansion of the CD8+ CD28population (50). The authors hy pothesized that the production of a CD8+ CD28intraepithelial T-cell population was the result of peripheral blood T cells that migrated to the epithelium and adhered to intestinal epithelial cells via gp180 or CD1d (50). The ability of intestinal epithelial cells to act as antigen presenting cells has been well documented by electron microscopy which showed soluble antigen taken up by endolysosomal pathways, expression of antigen with gp180 and CD1d (a nonclassical MHC molecule), and engagement of the intestinal epithelial cell with the CD8 molecule (23). A complex is formed between gp180 and CD1d on the intestinal epithelial cell and
17 the CD8/TCR on the intraepithelial lym phocyte (50).The epi thelial surface molecule gp180 interacts with T cells via CD8 to activate p56lck (50,59). Campbell et al. (59) proposed that this complex served as a non traditional class I MHC molecule which triggered regulatory rather than cytolytic T-cell functions. This theory was confirmed by studies show ing monoclonal antibodies against gp180 blocked CD8+ T-cell proliferation and the acti vation of the CD8 associated p56lck (23). Campbell et al. (59) suggested that IEC (intestinal epithelial cells) may play a role in the activation of a subset of T cells involved in the suppression of local mucosal, and systemic immune response. The idea of suppressive activity of the CD8+ CD28intraepithelial lymphocytes was al so supported by the lack of the presence of the CD28 molecule. Work by Allez et al. (60) also supported this idea by showing that long-term-cell culture of human intestinal epi thelial cells with human CD8+ T cells caused expansion of the CD8+ CD28subpopulation. Cell and protein analysis by flow cytometry and mRNA expression showed that CD8+ CD28+ T cells stimulated by IL-2 for 68 weeks were almost entirely CD28(61). The identification and characterization of this cell population pr ovides essential insight into the mucosal cellular en vironment available to mount an immune response. Cytokines and Chemokines Cytokines play a crucial role in t he orchestration of immune responses. These small proteins, typically weighing around 25 kDa, are released by immune and some non-immune cells in response to a stimulus and bind to receptor proteins on target cells. Cytokines al ter the microenvironm ent and cell-to-cell communications through their ability to act in an autocrine, paracrine, or
18 endocrine manner, meaning that the cytoki ne released by a cell can influence itself, cells in the immedi ate vicinity, or distant ce lls (depending on the particular cytokine stability), respectively. Cyto kines are divided into groups based on structural characteristics and functional pr operties. Pond et al. (62) showed that the profile of cytokines produced is stim uli-, mouse strain-, and tissue-specific, creating a great diversity in potential immunological responses. Cytokine gene expression in a primary infection diffe rs between lymphocytes isolated from BALB/c mouse PeyerÂ’s patches, mesenter ic lymph nodes, and splenocytes (63). When Svetic et al. (63) enterally chal lenged mice with a paras itic nematode, they saw no differences in cytokine expre ssion in splenocytes, but significant alterations in PeyerÂ’s patch and mesenteric lymph node cytokine expression. Many studies have attempted to def ine the cytokine changes during zinc deficiency, measuring systemic and spleni c cytokine patterns of expression and then using these findings to explain or make interpretations about the parasitic challenge results occurring within the inte stinal tissue. However, studies by Svetic et al. (63) and Pond et al. (62) discredit this methodology, suggesting that due to tissue differences a more direct measurement of tissue cytokines is required to truly support such interpretations. CD4+ T cells are called T-helper (TH) cells. There are two general subclasses of TH cells: TH1 and TH2. Cytokine responses have been generally classified as either a TH1 or a TH2 response, based on the types of cytokines produced by these subsets of T-helper cells after antigen activation. TH1 support
19 more inflammatory cell-mediat ed immune responses and a TH2 response supports T-cell regulation of the humoral ( antibody producing) cellular response. Chemokines (7-12 kDa), a particular fa mily of smaller cytokines which can be further subdivided based on amino acid structure, are also secreted products of T cells and are often the first proteins released by tissues in an immune response. Their primary role is to recruit leukocytes to the site of inflammation. Cytokines and chemokines work together to stimulate the endothelium of vessel walls to express selectins and integrins, slo wing the flow of target cells passing in the bloodstream, aiding in the diapedesis of cells across the endothelium from the blood vessel into the tissue. Chemoki nes then provide a chemical gradient to allow for chemotaxis of the target cells to the area of inflammation. Chemokines and cytokines are an integral part of both the initial innate immune response and the adaptive immune response. Cytokines, Chemokines and Their Importance in GALT The essential role of cytokines and chemokines within the GALT, as in other tissues, is in the recruitment of leukocytes to sites of immune responses and inflammation. The mesenteric lym ph nodes, PeyerÂ’s patches, and lamina propria contain high endothelial venules . Cytokines and chemokines act on endothelial cells lining the venules to pr omote the expression of selectins and integrins. These adhesion molecules al low for the adherence of circulating leukocytes to the walls of the hi gh endothelial venules, followed by the diapedesis and chemotaxis of leukocytes in to the tissues. In this way, cytokines and chemokines can recruit T cells and other populations essential for maintaining integrity of the mucosa l barrier to the lamina propria and
20 intraepithelial spaces within the small in testine and the colon. The chemokines CCL 28 (chemokine ligand 28) and CCL25 ar e produced within the intestine by epithelial cells. These proteins bind to the receptors CCR10 (c hemokine receptor 10) and CCR9, respectively, found on gut homing lymphocytes. Previous studies show that CCR9 deficient mice have a 50% reduction in IgA secreting plasma cells (64), alterations in the developm ent of T and B cells, and a reduction in TCR + IEL resulting in a decreased T ce ll to epithelial cell ratio (65). Furthermore, Hosoe et al. (66) sh owed that blocking of the CCL25-CCR9 pathway with an antibody against CCL25 inhi bited the accumulation of LPL and IEL into the intestinal mucosa. Berin et al. (67) showed that human intestinal epithelial cells produced CCL22, result ing in the recruitment of CCR4+ T cells, and the production of TH2 anti-inflammatory cytokines. Human colon intestinal epithelial cells were also shown to ex press and secrete chemokines that could recruit many different cell types into t he mucosal tissue (68). In this study, the stimulation of epithelial cells by bacteria upregulated mRNA and protein chemokine expression leadi ng the authors to conclude that these chemokine changes may explain the differential appear ance of leukocytes during a mucosal inflammatory response (68). Cytokines and chemokines play an important role as chemoattractants for cell popul ations in the maintenance of the mucosal barrier and in response to an inflammatory challenge. Overall Immune Responses During Zinc Deficiency As reviewed by Oteiza and Mackenzie there are numerous clinical situations that can lead to zinc deficiency, genetic conditions such as acrodermatitis enteropathica (a genetic condition, producing a severe zinc
21 deficiency), early parenteral nutrition, ch ronic diarrhea, malabsorption diseases, CrohnÂ’s disease, aging, and short bowel syndrome (69). Severe deficiency is relatively uncommon unless genetic conditions exist or in older studies evaluating parenteral nutrition; however, mild zinc deficiency can occur due to a number of the conditions listed above. Furthermore , zinc deficiency is associated with a characteristic decline in many parameter s of immune function. Zinc deficiency has been linked to decreases in body weight, thymic weight, lymphocyte population, erythroid populat ions (red blood cell progenitor s), natural killer cell activity, and delayed-type hypersensitiv ity; whereas zinc deficiency can simultaneously increase numbers of gr anulocytes and monocytes, and promote susceptibility to parasitic infections ( 14,16,70-75). Six-week-old A/J strain mice fed a zinc deficient diet (0.8 ppm) for 6 weeks showed a two-fold increase in plasma levels of the glucocorticoid corticosterone when compared to zincadequate mice (>50 ppm) (76). Glucocortico ids have been shown to cause death of immature thymocytes (77), increas e circulating granulo cytes numbers, and extend the life of neut rophils, all of which also occur during zinc deficiency (78,79). Zinc deficiency is also associ ated with increased ox idative stress, DNA fragmentation, altered transcription factor activity, such as NFB, ultimately affecting gene expression, and altered DNA repair mechanisms (71,72,80,81). Due to the importance of zinc structurally and functionally it has the potential to disrupt normal bodily functions at many different levels. Previous studies have shown decreases in thymus weight and body weight as a result of zinc defici ency (70,82,83). Studies involv ing zinc-deficient models
22 are complicated by the fact that intake of a zinc-deficient diet is accompanied by a decrease in food intake; therefore, it is difficult to interpret which conclusions result directly from the zinc defici ency and which are the result of caloric restriction. This analysis problem necessi tates the use of a pair-fed group, i.e. a group fed a zinc adequate diet but at the level of intake of the zinc deficient mice. In young A/J mice, 6 weeks of zinc deficiency (0.8 ppm) led to impaired weight gain in comparison to the zinc adequat e group (>50 ppm), which continued to gain weight as expected according to thei r growth curve (76). King et al. (83) showed that using a 50-day zinc-deficient diet (0.5 to 0. 6 ppm) to model a chronic-zinc deficiency in female A/J mi ce resulted in a 37% decrease in thymic weight, decreased body weight, but no signs of parakeratosis. The final mouse weights of the zinc-deficient mice in th is chronic-zinc deficiency study were 74% of their zinc-adequate study mates that gained weight as would be expected by comparison to growth curves for this strain; however, the weights of the zincdeficient mice were essentially identical to their baseline body weights. This implies that the zinc deficiency resulted in a lack of weight gai n over the course of the 50-day study, but led to no weight loss in comparison to their baseline weights (83). The authors made com parisons between the zinc-adequate and zinc-deficient groups, excl uding the pair-fed group, due to no differences seen in intake between the zinc-adequate and the pair-fed groups. These data are in contrast to earlier work by these investi gators in which the female A/J mice fed a 0.8 ppm zinc-deficient diet experienced an 84% decrease in thymus weight and had body weight being 65% of the zi nc-adequate group and less than their
23 baseline weight by day 24 of this severe zinc deficiency (16). The pair-fed group in this study had 88% of the body wei ght of the zinc-adequate mice (16). Furthermore, the weight loss in these severely zinc deficient mice was accompanied by a high degree of parakerat osis (16). Although the later study showed only impairment of weight gain while the earlier study showed weight loss in zinc deficiency, the authors did not explain the differences in the weight data obtained between the two studies . One possible explanation for the reduction in thymus weights could be attr ibuted to increased apoptosis due to the elevated corticosterone levels that occu r in zinc deficiency, as previously mentioned. Removing the source for the gl ucocorticoid by adrenalectomy of zinc deficient mice (0.8 ppm) re sulted in thymus weights equivalent to zinc-adequate mice (>50 ppm) (76). These adrenalectomiz ed, zinc-deficient mice with normal thymus weights did have alterations in t heir thymus composition after four weeks on the diet (76); zinc-adequate mice had a normal 2:1 (cortical:medullary) ratio, while the zinc-deficient mice had a 1:1 ratio (76). The authors hypothesized that the ability to maintain thymus weight but alter compartment size in zinc deficiency may be due to the redistribution of thymocytes (76). Although there are discrepancies on the exact differences that occur in thymus and body weight due to zinc deficiency, most authors c oncur that there are some negative consequences on growth and the thymus as a result of zinc deficiency. Zinc Deficiency and Gene Expression Zinc deficiency has been shown to alter gene expression in vivo and in vitro. Cousins et al. (84) found that cDNA microarray and quantitative PCR of RNA from human mononuclear cells (THP -1) cultured under zinc-supplemented
24 and zinc-deficient conditions showed appr oximately 5% of the 22,216 genes expression were zinc-responsive. When t he zinc responsive genes were divided into subgroups based on functional characteristics, genes involved in signal transduction, immune/cytokine function (cyt okines & receptors), nucleic acid binding, metabolism, apoptosis, cell grow th/development, and cytoskeleton were identified as being zinc responsive (84). Some of these changes in gene expression, but not all, may be attributed to metal-responsive transcription factor 1 (MTF-1), a transcription fa ctor containing 6 zinc fi ngers (85). MTF-1, in the presence of zinc, binds to metal responsive elements in certain genes and induces gene transcription. Metallothi onein is an example of such a MTF-1 dependent, zinc-responsive gene (86). Zinc deficiency in a non-differentiated Tcell line resulted in decreased binding of the zinc-finger transcription factor NFB to DNA (13). Analysis of the effects of zinc deficiency on gene expression in rat liver via cDNA and oligonucleotide a rrays showed the presence of zincresponsive genes with 66 out of 1550 observable genes altered in zinc deficiency (87). Studies by Moore et al. (71,72) identified zinc-de pendent changes in gene expression in thymocytes from CD-1 mice fed a 3-week zinc-deficient (<1 ppm) versus zinc-adequate diet. Genes important in T-cell development and activation, such as p56lck, heat-shock proteins, MHC class II molecules, and a T-cell cytokine receptor, were altered by zinc deficiency (71,72). One study in rats using differential mRNA display on mRNA from rat intestinal tissue identified 13 zinc-regulated genes important in signaling, growth, and transcription, with a 1.5
25 fold difference (88). Collectively, thes e data suggest a tissue dependent effect of zinc deficiency in the regul ation of gene expression. Zinc Deficiency and Cellular Changes Many changes in the immune function t hat occur during zinc deficiency may be due to tissue specific changes in ly mphocyte populations. Absolute numbers of nucleated cells obtained from the bone marrow of moderately and severely zinc-deficient 6-week-old A/J mice were were not different than mice fed a zincadequate (28 ppm) or a pair-fed diet; how ever, the composition of the subpopulations was altered by zinc deficiency (as discussed below) (70). The moderate and severely zinc deficient mice were distinguished by subdividing the group as follows: after receiving a zincdeficient diet (<1 ppm) for 34 days, a mouse was considered moderately zinc defic ient if the weight was 73-75% of the zinc-adequate group, and severe ly zinc deficient if the mouse weight was 6871% of the zinc-adequate gr oup (70). In this study, t he erythroid ce ll population decreased approximately 25% and 60% in the moderately and severely zincdeficient groups, respectively. The eryt hroid cell population composed 18.5% of the nucleated cells extracted from t he bone marrow in the pair-fed and zincadequate group, but only 13.7% in the moder ate, and 9.1% in the severe zincdeficiency groups (70). The authors sugges t that losses in the erythroid cell compartment may explain the anemia that often occurs in zinc deficiency (70). The lymphoid subpopulation of the bone marrow, which gives rise to both T and B lymphocytes, decreased by 50% and 70% in moderately and severely zincdeficient mice, respectively, in comparison to pair-fed and zinc-adequate mice (70). Evaluation of the pr e-B-cell lineage within the bone marrow found pre-B cell
26 losses were only 15% in moderately zinc-deficient mice, versus 75% for severely zinc-deficient mice (70). The granulo cyte population, which is composed of neutrophils, basophils, and eosinophils, in creased by 36% in the moderately zinc-deficient mice and 57% in the se verely zinc-deficient mice bone marrow (70). Granulocytes, as a per centage of the nucleated ce ll population, represented 40%, 54%, and 62%, in the zinc-adequate, moderately zinc-deficient, and severely zinc-deficient diet groups, respectively (70). Bone marrow monocyte populations were 70% larger than in zi nc-adequate mice in both the moderately and severely zinc-deficient diet groups (70). The decreases in percentages and overall numbers of erythr ocytes and lymphocytes were balanced by increases in the percentages and overall numbers in the granulocyte and monocyte populations, resulting in no change in the ov erall nucleated cell populations in the bone marrow during zinc deficiency. King and Fraker hypothesize that downregulation of the metabolically drai ning lymphopoiesis may allow the body to conserve nutrients in the face of micr onutrient deficiency (70) . A later study by the same investigators evaluated a 50-day feeding cycle to mi mic a chronic-zinc deficiency (83). Evaluation of the nuc leated cell populati ons from the bone marrow identified a 35% decrease in the erythroid cell population and no other changes in other subpopulations (83). If zi nc deficiency leads to increases in neutrophils and monocytes, often the first cells to arrive to the location of in inflammatory challenge, they may enable the innate immune system to compensate for the lymphocyte alterations that have the potentia l to disrupt the adaptive immune response.
27 The effects of zinc deficiency on T-ce ll populations in peripheral blood have also been evaluated in rats (17). Peri pheral blood from 3-week-old Sprague Dawley rats fed a zinc-deficient diet (<1 ppm), zinc-adequate diet (level not stated in paper), or pair-fed diet for 3 weeks was analyzed for the presence and proportions of T-cell subpopulations. No differences in T-cell subpopulations were seen except that fewer CD90+ TCR T cells were found in the blood of zinc-deficient growing rats than in the pair-fed and zinc-adequate groups (17). T cells that have recently undergone maturati on in the thymus express the protein CD90 (17). The peripheral T-cell lymphopenia that occurs in zinc deficiency is linked to pre-T cell losses via apoptosis. Th ymic atrophy also plays a role in the lymphopenia that occurs in zinc deficiency due to inability to replete the dwindling T-cell populations. Furthermore, subpopulati on alterations that occur in zinc deficiency may shift the T-cell subpopula tions selected within the thymus and then released into the periphery. Evaluation of thymocytes following zinc deficiency has resulted in conflicting data. These differences may relate to the interpretation of the data by the investigators and how the data were pr esented. In two separate studies on 3week-old rats, Hosea et al. (17,89) observed decreased thymus weights in zincdeficient (<1 ppm), compared to pai r-fed or zinc-adequate (30 ppm) groups following 3-weeks of diet. However, if thymus weights were expressed as a percentage of body weight, no changes were detectable (17,89). In both of these studies the author failed to identify di fferences in the absolute number of lymphocytes isolated per gram of thymus weight (17,89). Two additional studies
28 using 4-6 week-old A/J mice fed a zincdeficient (0.5 ppm) or a zinc-adequate (28 ppm) evaluated over a time-course of 31 days and longer, found that thymus weights in the zinc deficient mice were 38% (after 31 days of di et), 58% (after 45 days), and 63% (after 50 days) of that of the zinc-adequate mice (15,83). These mice had corresponding decreases in th ymocytes extracted (49-80%) in comparison to zinc-adequate mice (15, 83). Although these data suggest that decreases in thymus weight are asso ciated with decreased cellularity, and that lack of weight change of thymuses resu lts in lack of overall changes in cell numbers, the earlier studies did not repor t overall thymocyte numbers only the normalized (number/gram of organ weight) were reported. Thymocytes from 6week-old A/J mice fed a 3-week zinc-defic ient diet (0.5 ppm) resulted in a threefold increase in apoptosis in CD4+ CD8+ pre-T cells versus zinc-adequate mice, resulting in a 40% decrease in CD4+ CD8+ pre-T cells in the thymus of zincdeficient mice (15). Mature T cells (CD4+ CD8and CD4CD8+) had no increases in apoptosis as a result of the zinc deficiency (15). A 50-day chronic-zinc deficiency (in 4-week-old A/J mice, fed 0.5 ppm) resulted in a 60% increase in apoptosis of pre-T cells (83). Thymocytes fr om zinc-deficient rats (<1 ppm diet for 3 weeks) had increased CD4CD8+ TCR percentages compared to zincadequate diet-fed rats (17). A separate st udy showed that 3 weeks using similar diets with rats resulted in decreased (35-52%) thymocyte pre-T cells (TCR CD4+ CD8+ and TCR + CD4+ CD8+) in both zinc deficient (<1 ppm) and pair-fed rats in comparison to zinc adequate (30 ppm), in addition thymocytes from zincdeficient rats had lower TCR + CD4+ CD8(T helper) populations versus zinc-
29 adequate rats (89). The pre-T-cell ly mphopenia may be caused by increased glucocortocoid levels in zinc defici ency (mentioned above), which may lead to thymic atrophy as a result of apoptosis t hat occurs in zinc deficiency (83,90). The involution of the thymus that occurs during zinc deficiency may also be associated with decreased ability to disti nguish self from nonself and a decrease in the number of mature T cells (although the data are inconclusive). The effect of zinc-deficiency on the sp leen was evaluated. Hosea et al. (89) found that 3-week-old rats fed a zinc-deficient diet (<1 ppm) for 3 weeks had a 20% decrease in spleen weight in com parison to baseline values, while pairfeeding resulted in no change, and spleen weights from zinc-adequate fed mice increased 78% over baseline. However, normalization of the spleen weight to body weight resulted in no differences am ong groups (89). In a separate study by the same authors, rat spleen weights were significantly lower (P<0.05, 336 mg Â± 23, mean Â± SEM) in zinc-deficient mice (<1 ppm, 3-week feeding period) than pair-fed (405 Â± 24) or zinc-adequate mice (743 Â± 37); however, normalization by body weight resulted in no difference am ong groups (0.23% to 0.25%, spleen weight/body weight) (17). Moderately and severely zinc deficient A/J mice, weighing 66% and 72% of the zinc-adequat e group, respectively, both had an approximate 45% decrease in splenic lymp hocytes (91). In this study, the severely zinc-deficient group also show ed parakeratosis (91). Six-week-old A/J mice defined as severely zinc deficien t, weighing only 65% of the zinc-adequate mice and having parakeratosis following 31 days of a 0.8 ppm diet, exhibited a 26% increase in T-helper cells, no c hange in cytotoxic T cells, and a 20%
30 increase in the T-helper/Tcytotoxic ratio in comparison to a zinc-adequate group (16). This same study identified only small decreases (5 and 8%) in the B-cell population (B220+) in moderately, weighing 70-72% of zinc-adequate mice, and severely zinc-deficient mice, respective ly (16). This B-cell population decrease was attributed to the non-IgM and non-IgD, immunoglobulin positive subset (16). Rats fed a zinc-deficient (<1 ppm) di et for 3 weeks showed no changes in splenocyte T-cell subpopulations in comparison to pair-fed and zinc-adequate mice (17). This study identified fewer CD90+ TCR T cells in splenocytes isolated from the zinc-deficient rats versus the pair-fed and zinc-adequate rats (17). These data are in contrast to a la ter study by the same authors which found that splenocytes from 3-w eek-old rats fed a zinc-def icient diet (<1ppm) for 3 weeks had 40% to 63% fewer T-helper cells (TCR + CD4+ CD8-) and fewer Tcytotoxic cells (TCR + CD4CD8+) versus the pair-fed and zinc-adequate diet groups (89). Furthermore, spleen weight and T-cell populations recovered within 7 days during zinc repletion in compar ison to 23 days for thymic weights and subpopulations (89). The spleen is an im portant lymphocyte-rich organ which assists in the ability to fight infections . Hence, alterations in splenic functional capacity during zinc deficiency coul d have negative impact on immune responses. Although lymphopoiesis appears to be affected during zinc deficiency (70,92), the remaining cells have normal immune responses (91). Adult mice with decreased zinc intake show losses of pr e-B and pre-T cells and associations of these losses with decreases in protec tive Bcl-2 levels, suggesting another
31 possible mechanism for altering lymphopoiesis (4). The protein Bcl-2 is present on the outer mitochondrial membrane, and increased BCL-2 levels have been shown to be protective against apoptotic mechanisms (93). The function of the residual lymphocytes remaining after zinc deficiency has been shown to be comparable to pair-fed and zinc adequate mice, therefore, the overall decreased immune response due to zinc deficiency can most likely be attributed to the overall decreases in lymphocyte numbers described above or potentially due to shifts in cell subtypes (91,94). Cook-Mills et al. (91) compared the T-lymphocyte functionalit y after a zinc-deficient diet was fed for 30 days. Splenocytes were isolated and cultured for evaluation of proliferative responses and IL-2 production in response to stim ulation with a mitogen (concanavalin A), and in a mixed lymphocyte culture system (using mitomycin C-treated C57BL/6 target cells). Culturing in vitro allows for normalizing the differences in cell numbers obtained from zinc-deficien t, pair-fed, and zinc-adequate mice by performing each assay with an equal number of cells. Lymphocyte proliferative responses and IL-2 production in response to concanavalin A were similar among the zinc-deficient, pair-fed, and zinc-adequate mice (91). In mixed lymphocyte culture the zinc -deficient splenocytes had a higher proliferative response and IL-2 production than those from zinc-adequate mice (91). This may be related to the exposure of splenocyt es to elongated steroid exposure which has been shown to increase mixed lymphocyt e culture responses (95,96). These data are supported by the work of Dowd et al. (94) in which splenocytes isolated from rats fed a zinc-deficient or a pair-fed diet for 4 weeks had equivalent
32 proliferative responses to concanavalin A and IL-2 production. The in vivo cell culture experiments of Cook-Mills et al. (91) and Dowd et al. (94) created a zincdeficient culture environment by using autologous serum from zinc-deficient animals in the zinc-deficient cell cultur es. Investigators in an earlier study had shown decreases in extracted splenocyte T-ce ll responses, but in vitro cultures of these splenocytes contained fetal-bovi ne serum and not autologous serum from zinc-deficient animals (97). Splenocytes were removed from zi nc-deficient, zincadequate, or pair-fed mice injected 5 da ys earlier with sheep red blood cells (SRBC), and the function of B cells wa s analyzed by their ability to form antiSRBC antibodies, IgM anti-SRBC, and IgG anti-SRBC (91). Although the number of antibody producing cells was lower per spleen, the amount of antibody per antibody-producing cell was equivalent bet ween the zinc-deficient, pair-fed and zinc-adequate mice (91). The lymphopenia that occurs in zinc deficiency may result in immune dysfunction; however , the remaining lymphocytes appear to retain their functional characteristics. No studies have evaluated the changes in cellular populations present in the gastrointestinal tissue as a result of zinc deficiency. The changes in immune function at the level of the gastrointestinal tract during zinc deficiency may be the indirect result of changes to systemic immune system development that are perpetuated into the GALT. Zinc Deficiency and Cytokine Responses Zinc deficiency is associat ed with an imbalance between TH1 and TH2 cytokines (13,14,98). Two studi es evaluating the effect of a mild zinc deficiency on human peripheral blood mo nonuclear cells which were isolated and then
33 stimulated in vitro, resu lted in a decline in the TH1 cytokines IL-2 (at 8,12, and 20 weeks) and IFN(at 20 weeks) (99,100). In one of these studies there was no effect of zinc deficiency on the TH2 cytokines IL-4, IL-6, and IL-10 at 20 weeks (100). Analysis of cytokine production by HuT-78 cells, a human malignant T lymphoblastoid cell line, cultured in zinc -deficient culture conditions showed 40% less IL-2 protein and a 50% decrease in IL-2 mRNA following stimulation (13). BALB/c mice fed a zinc-deficient (<1 ppm ) diet for four weeks were unable to respond to a nematode (parasitic) challenge, which requires a TH2 cytokinemediated immune response, resulting in an increased presence of worms (101). Splenocytes from these mice stimulated in culture with the parasite antigen had decreased IL-4, IL-5, and IFNin the infected group in comparison to the zincadequate mice, and no detectable IL-4 in t he uninfected zinc-deficient mice (101). The same authors, f ound that purified T cells from splenocytes of BALB/c mice exposed to the same zinc-deficient conditions as in the previous study had decreased IL-4 and IL-5 cytokine producti on in comparison to pair-fed and zincadequate mice (102). Furthermore, splenocyt e populations from both the zincdeficient and pair-fed mice had decreased IFNproduction from T cells and decreased IL-4, IL-5, and IFNproduction by antigen presenting cells in comparison to zinc-adequate mice (102). This suggests that food restriction rather than zinc restriction produc ed these changes. These authors also evaluated a primary versus secondary immune response in BALB/c mice exposed to a 4-week zinc-deficient (< 1 ppm), pair-fed, or zinc adequate (60 ppm) diet given a parasitic challenge, which was treated, and then the mice were re-
34 exposed to the same parasitic challe nge (103). Stimulation in culture with parasitic antigen resulted in decreased IL-4 levels in zinc-deficient mice following primary infection but not sec ondary infection, and decreased IFNfollowing secondary challenge but not after primary challenge. Bao et al. (98) evaluated cytokine expression from several different cell lines following 24 days of culture in zinc deficient conditions. They found t hat zinc deficiency altered both cytokine mRNA and protein concentra tions. Decreases were identified in IL-2 and IFN, while increases were found in IL-1 , IL-8, and TNF. The authors concluded that zinc deficiency affected cytokine patte rns at the level of gene expression, and that differences in expression were dependent of the cell li neage. The effect of zinc deficiency on TH1 or TH2 cytokine expression may be dependent on the experimental system evaluated (human versus mouse), type of challenge (primary versus secondary, or parasitic versus bacterial), and the level of zinc deficiency. The impaired ability of zinc -deficient humans or mice to produce TH1 and TH2 cytokines would result in an incr eased susceptibility to immunological challenges, as was seen in the par asitic studies described above. Zinc Deficiency and Gut Associated Lymphoid Tissue (GALT) In addition to the inability of zinc defi cient mice to protect their intestinal tract against parasitic infections as mentioned above, many studies link zinc deficiency with increased diarrhea in mice and humans suggesting disturbance of the gut barrier (104-106). T he gut is an important medi ator in zinc homeostasis (107). The body has no Â“long-term storehouse Â” of zinc (107). However, epithelial cells in the intestinal mucosal system contain transport and trafficking proteins capable of facilitating or limiting zinc translocation from the lumen into the
35 bloodstream. Fraker and Ki ng as recently as Februar y of 2004 noted Â“how little we know of the impact of zinc deficiency on mucosal immunityÂ” (4). The imbalance between TH1 and TH2 cytokines may be one mechanism by which zinc deficiency promot es disturbances of the gut mucosa. Individuals with zinc deficiency are more susceptible to intestinal parasitic infections which require TH2 responses for effective eliminatio n from the gut associated lymphoid tissue (GALT) (4). Â“Many studies have documented the early synthesis, production, and uptake of TH2 cytokines by GALTÂ” as important for the production of effective immune responses (108). As previously described, although Prasad et al. (13) reports only TH1 cytokine differences, Scott et al. (14) showed zinc deficiency decreased TH2 cytokine expression and responses in intestinal tissue following parasitic challeng e. Interestingly, parasitic infections are associated with increased IgE producti on and transport into the lumen of the intestine (63,109). The ability of T cells with TCR origin (which are present in unusually high concentration in the mucosa ) to signal the B cells to switch isotype production to IgE is dependent on the presence of TH2 cytokines (108). This is supported by the work of Shi et al. (103) who showed that BALB/c mice fed a zinc-deficient diet (<1 ppm) for four weeks and exposed to a parasitic challenge had decreased IgE production in comparison to pair-fed or zincadequate mice, but no differences were identified among groups for IgG. The failure of zinc-deficient mice to maintain their TH1 and TH2 cytokine production can inhibit the ability of these mice to protect their gut via IgE antibody production. Furthermore, the inte stinal epithelia is one of the most rapidly turning
36 over tissues in the body and has the potent ial to be affected by zinc deficiency which is associated with increased apopt osis as previously mentioned (110). It is essential that the intestine is able to limit the responsiveness to antigens that are frequently enc ountered in our food to prevent hypersensitivity reactions within the gut. This resulting to lerance is T-cell specific and mediated by the induction of TH2 cytokines and suppression of the TH1 inflammatory cytokines (20). This induction of antigen to lerance also involves regulatory T cells (Treg) and the production of a major suppressive cytokine, TGF. Since TH2 cytokines are known to decrease during zinc deficiency, it has been hypothesized that the ability to induce tole rance in zinc deficient animals would be impaired (111). Finamore et al. (111) ev aluated the sensitivity of the intestine to food antigens during zinc deficiency. Repeated exposure in vivo to ovalbumin failed to induce tolerance in zinc-defici ent mice as measured by stimulation of mesenteric lymphocytes and splenocytes with ovalbumin in vitro compared to pair-fed controls, and a resulting decrease in TH2 cytokines (111). However, the 28 days it took to induce tolerance wa s at a point where many studies have shown adverse intestinal changes as a resu lt of zinc deficiency. The failure to evoke tolerance may have been due to a concurrent inflammatory response, which was found to exist in this study based on neutrophil infiltration and histology (111). The authors concluded that t he inability for zinc-deficient mice to develop tolerance as a result of cytokine changes would lead to a more easily inflamed intestinal mucosa (111). These changes at the cellular level have the capacity to interfere with gut homeostasis.
37 Studies show that zinc-deficient ani mals have increased iNOS expression, and the shift from TH2 to TH1 cytokines may be responsible for this iNOS increase. PCR showed iNOS mRNA levels in the small intestine increased in zinc-deficient mice versus pair-fed mice (106). Immunohistochemistry of the intestinal segments following addition of cytokines known to invoke an acutephase response, showed an increase in antiiNOS staining in the zinc-deficient group (106). This staining was localized to the basal layer and dispersed throughout the villus cells (106). Furthermo re, addition of the iNOS inhibitor NGnitro-L-arginine methyl ester (L-NAME) to the drinking water of mice fed the zincdeficient diet decreased iNOS mRNA ex pression and apoptotic cells in the intestinal villi versus mice fed the zincdeficient diet without L-NAME (105). Nitric oxide (NO), the product of the iNOS en zyme, leads to tissue damage when produced in excess. Canali et al. (112) showed that addition of polyphenols, an antioxidant in red wine, to a zinc-def icient diet decreased the number of macrophages and neutrophils that migrat ed into the intestinal mucosa and decreased the production of inflammatory cytokines. These data again show the association of iNOS through its production of NO with intestinal damage in a zinc-deficient animal model. Another mechan ism for the diarrhea that occurs in zinc deficiency is increased uroguanyli n (113). Levels of preprouroguanylin mRNA were 2.5-fold more abundant in rat intestine during zinc deficiency (113). Uroguanylin, a natriuretic peptide hormone, binds guanylate cyclase C (which is also bound by the Escherichia coli enter otoxin) and alters fluid balance in the intestine (113). The diarrhea that is a ssociated with zinc deficiency in many
38 cases not only exasperates the zinc def iciency itself but leads decreased absorption of other nutrients. Zinc and T-Cell Signaling T-cell signaling in activation involves the interaction of many components, several of which have been shown to be alte red in zinc deficiency. These include IL-2, IL-2 receptor (IL-2R), and p56lck. IL-2 is essential in the activation of T cells, which can then release IL-2 to act in an autocrine and paracrine function to stimulate proliferation. In vi tro stimulation following zinc -deficient culturing of a T lymphoblastic cell line showed decreased IL-2 production (98). Furthermore, mildly zinc-deficient human subjects have also been shown to have lower IL-2 production from TH1 lymphocytes (74). Prasad et al . (13) found that the HuT-78 cell line cultured in zinc-deficient conditions produced 50% less IL-2 mRNA, and had a 70% decrease in the soluble IL-2 receptor (sIL-2R) in comparison with zinc-sufficient cells. These changes in IL-2 and sIL-2R data were confirmed at the protein level via wester n blotting (13). IL-2 and/or IL-2R deficient mice were shown to have an inability to develop self -tolerance, and increased incidence of inflammatory bowel disease (114-116). An increase in p56lck, a zinc-finger protein with an essential role in the perpetuation of a cell surface stimulation signal to the intracellular cascading signaling events, has been shown to incr ease in zinc deficiency (72,117,118). Lepage et al. (117) showed that splenocytes isolated from C57BL/6 mice fed a zinc-deficient diet (<1 pp m) for 4 weeks had increased p56lck protein as visualized via western immunobl otting. Cells isolated from the pancreas of CD-1 mice fed a zinc-deficient diet (<1 ppm) for 3 weeks had increased p56lck as
39 measured by cDNA array (>1.5-fold increase), RT-PCR, and western analysis (>80% increase) in comparison to mice f ed a zinc-adequate diet (72). Lin et al. (119) showed the binding of p56lck to glutathione S-transferase (GST) which is complexed with the cytoso lic portion of CD4 or CD8 is dependent on the zinc. Moore et al. (72) hypothesized that meta llothionein may act as a donor and/or acceptor for zinc in the zinc-dependent p56lck interaction with CD4 or CD8, and that disturbance of the inte raction between CD4/CD8 and p56lck may trigger the cell to upregulate p56lck expression due to a feedback mechanism. Overexpression of p56lck has been shown to alter Tcell maturation (decreasing CD4+ CD8+ generation from CD4CD8precursor cells) (120), increase the incidence of thymic tumors (121), and dec rease the presence of TCRs on the cell surface by 75% due to increased lysoso mal degradation (122). Sohn et al. (123) found that transgenic mice overexpressing p56lck also had decreased double positive CD4+ CD8+ thymocytes, but had an increas ed presence of single positive CD4+ and/or CD8+ thymocytes; however, increased p56lck led to decreased survival of peripheral T cells. Alterations of p56lck may provide one mechanism through which zinc deficiency is able to alter T-cell maturation, signaling, and phenotype charactersitics. Other molecules altered by zinc defic iency that may affect T-cell signaling include TNFand the MHC class II molecule. B ao et al. (98) showed that zinc deficiency in cell culture increased TNFconcentrations. This alteration in TNFcan be attributed to a zinc finger protein, tristetrapolin (TTP), capable of being modulated by zinc concentrations in cell culture (124), which decreases the half-
40 life of TNFmRNA (125). Peritoneal macrophages from TTP deficient mice overproduced TNF(126). Furthemore, in vitro cu lture of a T-ce ll line with TNFfor 8 weeks resulted in the specif ic decreased expression of the T-cell costimulatory molecule CD28 without altering other T-ce ll markers (CD4 or CD3) (127). These data provide a potential me chanism by which zinc deficiency may induce CD28 costimulatory marker c hanges, although no studies have evaluated the direct relationship between zinc deficiency and CD28. The mRNA of the Class II MHC molecule, which presents antigen to the T CR/CD3 complex on CD4+ T cells, was shown to decrease in thymocytes from CD-1 mice fed a zinc deficient (<1 ppm) diet for 3 weeks ( 71). Decreased presence of class II MHC molecule, a necessary antigen presenting molecule throughout the body, in the thymus may alter positive and negative se lection during the maturation process. Moore et al. (71) concluded that r eduction in the MHC Class II receptor contributes Â“to the lymphopenia of zi nc deficiency or the pathogen-specific increased susceptibility to infectious disease seen secondary to a zinc deficiencyÂ”; in addition, alterations in any of the signal or costimulatory molecules that may occur during zinc defici ency could lead to these negative consequences. Purpose of this Work Despite protective mechanisms, in creased susceptibility to parasitic infection and inflammation occur within the gut especially during zinc deficiency. Changes in cellular populations as a result of zinc deficiency are well documented systemically; however, differenc es in their tissue distribution exist based on the differential expression of tiss ue-specific chemokines and cytokines
41 as shown by Pond et al. (62). The purpose of this study is to determine if the systemic cellular lymphopenia and cytokine changes that have been shown to occur are perpetuated into the GALT as mice become progressively deficient in zinc. If these changes occur within the s hort-lived epithelial tissues, this would explain the inability of zinc-deficient animals to develop tolerance (111). Furthermore, zinc-deficiency induced weak ening and thinning of the epithelium could lead to increased uptake and pres entation of molecules as antigenic, resulting in the potential for acti vation and functional changes in T-cell subpopulations in the GALT. Changes in lymphocyte populations were evaluated in mice fed one of three diets, either a zinc-deficient, zinc-adequate, or a pair-fed diet for 9 weeks. Cells were isolated fr om the Peyers patches, mesenteric lymph nodes, and intraepithelial ly mphocytes and lamina propri a lymphocytes of the small intestine, and intraepit helial lymphocytes from the colon, for flow cytometric analysis of T-cell subpopulat ions present. Based on systemic losses of naÃ¯ve cell populations previously observed during zinc deficiency (4), it would be expected that changes in the subpopulations would occur. As a first look at T-cell functional activation, the effects of a 9-week progressive zinc deficiency on expression of the signaling molecule CD 28 on T cells was evaluated in all tissues. Tissue from the colon, the site of the symptomatic effects of zinc deficiency, was assessed for global changes in cytokine and chemokines at the transcript level. Evaluation of thes e changes within the GALT during zinc deficiency could provide insight into the increase in gastrointestinal illnesses that
42 occur during zinc deficiency which are lik ely the result of a breakdown in the mucosal barrier.
43 MATERIALS AND METHODS Animals Specific pathogen free (SPF) BALB/c mice (4-week-old, Harlan) were obtained, acclimated in microisolate r cages for 7 days, and then housed individually in acid-washed, hanging stai nless steel cages on a 12-h light/dark cycle with free access to distilled, deion ized water. Entrance into the mouse facility required gowning and personal protec tive equipment as in SPF facilities, although our housing facilities were not SPF. Mice were fed an AIN-76A-based pelleted diet formulated with egg white protein, containing 30 mg zinc/kg of diet (Research Diets, Appendix A) for the firs t week, 6 mice were killed to obtain baseline values, and the remaining mice were randomly assigned to the same zinc-adequate diet (ZA, n=11/ group) or a zinc-deficient diet (<1 mg zinc/kg diet, ZD, n=11/group). A group was pair-fed (n=11/ group) the ZA diet to the level of the ZD mice, but was later dropped from the analysis because differences in weight between the ZA and ZD diet groups were not observed (Appendix B). At various time points over the 9 weeks mice were anesthetized with halothane and killed by exsanguination via cardiac punc ture and cervical dislocation in accordance with the University of Flori da Institutional Animal Care and Use Committee approved protocol (#D963). The blood was collected, allowed to clot for 30 min at room temper ature, centrifuged at 1850 x g for 15 min, and serum was removed and stored at -20Â°C for meas urement of serum zinc. Mouse thymic
44 and liver tissues were removed and weighed. The small intestines and colons were removed and flushed with 10 and 5 mL, respectively, of 1 X HankÂ’s Buffered Saline Solution (HBSS, without Ca2+ or Mg2+, Fisher Scientific) and placed in HBSS/HEPES [15 mM HEPES (Fisher Scient ific) in HBSS without sodium bicarbonate, pH 7.2] on ice. Sm all intestinal tissue was processed for intraepithelial lymphocytes (IEL), lamina propria lymphocytes (LPL), and PeyerÂ’s patch lymphocytes. Mesenteric lymph nodes (MLN) were removed for isolation of lymphocytes. Colonic tissue was proc essed for the isolation of colon intraepithelial lymphocytes (cIEL only, not LPL). A 5 mm section of the colonic tissue used to isolate RNA was placed in to 10 volumes of RNAlater (Ambion, Inc.) at 4oC for 24 hrs, after which the super natant was removed and the tissue was stored at -80Â°C. Determination of Serum Zinc Concentrations Five-fold dilutions of serum in dH20 were analyzed for zinc content by atomic absorption spectrophotometry us ing an AAnalyst 100 Atomic Absorption Spectrophotometer (Perkin Elmer) and compared to a 0.2-1.0 ppm standard curve. Lymphocyte Extraction fr om Intestinal Tissue The procedure for isolation of lym phocytes from tissues was developed based on modifications and combinations of previously described methods (40,66,128). PeyerÂ’s patches and MLN were processed separately but similarly. MLN and PeyerÂ’s patches were removed from the small intestines, fat was removed, and tissues were cut into sma ll sections and cells were extracted by gentle application of pressure using a 5 -mL syringe plunger. PeyerÂ’s patch tissue
45 homogenate was resuspended in 20 mL of 50 U/mL type I collagenase in RPMI complete [10% fetal bovine serum-heat inactivated (FBS, Mediatech), 15 mM HEPES, 0.1 mg/mL gentamyacin, in RPMI-1640 containing L-glutamine but without sodium bicarbonate, pH 7.2, all reagents were obtained from Fisher Scientific], incubated at 37oC for 1 hr on a rocker. MLN homogenate was resuspended with 20 mL RPMI-complet e. MLN and PeyerÂ’s patches tissue suspensions were then propelled through a 100 um mesh. Extracted lymphocytes from each tissue were washed twice with RPMI complete (the first PeyerÂ’s Patch wash was in 50 mL; all other washes were in 20 mL) at 450 x g for 10 min, resuspended for counting in 1.0 mL of FACS buffe r [1% bovine serum albumin, 0.1% sodium azide, 1 X phos phate buffered saline (PBS), pH7.4], and stained with fluorescent conjugated anti bodies and then analyzed as described below in the Flow Cytometry section. The remaining small intestinal and co lonic (with PeyerÂ’s patches removed) tissues had fat removed, were cut length wise, and then were cut trans-sectionally into 5-mm sections and placed into a petri dish containing 10 mL of HBSS/HEPES (1 X HBSS, 15 mM HEPES, pH 7.2). The contents of the Petri dish were poured over a 250-um nylon-me sh screen held taunt by an embroidery hoop and suspended over a petri dish. These tissue pieces were returned to a petri dish rinsed with 10 mL of HBSS/ HEPES, and again filter ed by pouring over the same 250-um mesh apparatus. This procedure was repeated until the tissue had been washed 6 times with HBSS/HEPE S and filtered through the 250-Âµm nylon mesh each time. The remaining tiss ue pieces were then transferred into a
46 siliconized flask containing 10 mL of 37oC HBSS/FBS/EDTA (10% FBS, 15 mM HEPES, 5 mM EDTA, 0.1 mg/mL gentamycin, in HBSS, pH 7.2). This was stirred at 100 rpm (Fisher Electronic Stirre r Model 2008) for 15 min at room temperature. The contents of the flask were filtered through a fresh 250-Âµm nylon mesh filter (as described above), the tissue pieces were returned to the flask, a fresh 10 mL of room temperature H BSS/FBS/EDTA was added, and the flask was again stirred at 100 rpm for 15 min at room temperat ure. This procedure was repeated until the tissue pieces had been was hed and stirred 4 times. The filtrate from these washings was retained, pool ed, and stored on ice for processing of IEL (the tissue was processed for LPL as described below). The IEL containing filtrate was centrifuged at 400 X g for 5 min and resuspended in 4oC RPMI-1640. A nylon wool column was prepared by t he teasing and packing of 0.3 g of prewashed nylon wool into a 10-mL disposab le column. This column was equipped with a 23-gauge needle attached to a 3-wa y stopcock. The column was prewetted with 20 mL of 4oC RPMI-1640, and the column was not allowed to run dry. The resuspended cells were appli ed to the column, allowed to run completely through the column, and 10 mL of 4oC RPMI-1640 was added for elution. The entire 15 mL of flowthrough was centrifu ged at 400 X g for 10 min, the pellet was resuspended in 0.5 mL of FACS buffer and stained with antibody conjugates for flow cytometric analysis (des cribed in the Flow Cytometry section below). The remaining tissue pieces from the small intestine were processed for LPL (LPL were not isolated from the co lonic tissue). The tissue pieces for LPL processing were washed for 5 min in 20 mL of RPMI complete by stirring at 100
47 rpm at room temperature. This wash was filtered through a 250-Âµm nylon mesh (described above) and the tissue retent ate was digested by incubation at 37oC for 1 hr with 20 mL of 100 U/mL type V III collagenase in RPMI complete. The products of this incubation were filt ered through a 250-Âµm nylon mesh and the filtrate was centrifuged at 850 X g for 10 min, washed twice in 20 mL of CMF/HEPES with centri fugation at 850 x g for 10 min, resuspended in 0.5 mL of FACS buffer, and stained for Flow Cytometric analysis as described in the following section. Flow Cytometry Cell marker expression of T-cell s ubpopulations were evaluated using flow cytometry in LPL, IEL, cIEL, MLN, and small intestinal PeyerÂ’s patch cell populations at various time points over t he 9-week zinc deficiency protocol. Cell suspensions of 1.0 X 106 cells in FACS buffer cont aining 2 ug of unlabeled mouse IgG (Southern Biotech), for Fc re ceptor and non-specific antibody binding block, were prepared from each lympho cyte population. For each of the cell populations isolated and treated with m ouse IgG, a separate tube was prepared for determination of background fluorescenc e for each antibody by addition of an appropriate fluorochrome conjugated to an isotype control (BD Biosciences Pharmingen). The following antib ody conjugates (1 Âµg/1 X 106 cells) were added to stain the cells: fluorescein isothiocyanate (FITC) conjugated anti-TCR , allophycocyanin (APC)-anti-CD3, phycoeryt hrin (PE)-anti-CD2 8, FITC-anti-CD4 and biotinylated-anti-CD8 (BD Biosciences Pharmingen). The anti-CD8 labelled cells were subsequently detected by steptavidin-peridin in chlorophyll-a protein (SAv-PERCP) secondary antibody. In addition to the isotype control
48 tubes, the isolated cell suspensions containing mouse IgG were used as follows: to one tube antibodies were adde d for the evaluation of TCR , CD8 , CD3, and CD28, and a separate tube was prepared with CD4, CD8 , CD3, and CD28 antibodies. After a 15-min room-tem perature incubation, 200 ul of 4% formaldehyde in PBS was added. Samples were incubated for 10 min at room temperature and cells were washed twic e with 1 mL of FACS buffer and centrifuged at 600 x g for 5 min each time . Cells were resuspended in 0.4 mL of FACS buffer for analysis. Isotype controls were used to set the crosshairs to define the negative populati ons, and then analysis of the forward scatter versus the CD3 identified the CD3+ population. Flow cytom etric measurements were performed by counting 10,000 CD3+ cells using a FACSCaliber (BD Biosciences). Data were analyzed as the per cent of total cells and as a percent of the CD3+ population using FCS Express vers ion 3.0 (De Novo Software). Isolation of RNA All experiments were done using RNAse inhibitor spray (Continental Lab Products) and DEPC treated water to mini mize RNAse contamination. RNA was isolated from colonic tissue samples (approximately 16.5 mg, stored as previously described) from mice exposed to 3 and 9 weeks of a ZA or ZD diet, n=1/week/diet group, by homogenizati on and isolation according to the manufacturerÂ’s directions using the RNeasy kit (QIAGEN Inc.) for use in Microarray. RNA was quantitated using a Nanodrop Sp ectrophotometer (Nanodrop Technologies) and 4 ug of RNA was diluted 1:1 v/v with glyoxal sample loading dye (Ambion, Inc.) cont aining ethidium bromide, and treated according to manufacturers directions. Samples were loaded on a 1% (in TAE)
49 agarose gel and electrophoresis was perfo rmed at 100 V. Ethidium bromide intercalation into the RNA was visualized to check quality using a UV transilluminator. RNA from remaining colonic tissue sa mples in mice fed for 9 weeks was extracted for use in quantitative real-tim e PCR (q RT-PCR, 20 mg each, n=4/diet group) using a modification of the C homczynski RNA isolation method (129). Tissue was homogenized on ice in 1.0 mL of RNA-Bee (Tel-Test) using a TissueTearor (Fisher Scientific) on a setti ng of 3 for 30 seconds. The sample was centrifuged for 10 mi n at 10,000 x g at 4oC. The supernatant was removed, 0.2 mL of chloroform was added, and samples were incubated on ice for 5 min. The suspension was centrifuged at 12,000 x g for 15 min at 4oC. The top aqueous phase (0.45 mL) was removed and an equa l volume (0.45 mL) of isopropanol was added. This was mixed well, incubat ed at room temper ature for 10 min, centrifuged for 15 mi n at 12,000 x g at 4oC, and the supernatant was removed. The pellet was washed with 1.5 mL of 75% ethanol and centrifuged for 10 min at 8,000 x g at 4oC. The supernatant was poured off and the pellet was allowed to air dry for several minutes. The pellet was reconstituted with water and RNA was quantitated at 260 nm us ing a Beckman DU640 Spectr ophotometer (Beckman Coulter, Inc.). RNA concentration was determined using the OD260 X 0.04 Âµ g/ Âµ L(because 1 Absorbance Unit at 260 nm=40 ug/mL) X dilution factor = Âµ g RNA/ Âµ L. All RNA to be used for q RT-PCR was DNase treated according to the manufacturerÂ’s directions with TURBO DNase (Ambion, Inc.) before analysis. Absorbance at 260 nm was measured us ing a Nanodrop Spectrophotometer to
50 quantitate nucleic acid content of the extracted and DNase treated RNA for use in q RT-PCR. Microarray RNA prepared as described above from weeks 3 and 9 in ZA and ZD mice for microarray analysis was the template for preparation of cRNA using the TruLabeling-Amp 2.0 kit (Super Array Bioscience Corp.). Briefly, cDNA was prepared from 1.8-6 ug of RNA and used as the template for the subsequent cRNA synthesis reaction. The cRNA synthesis reaction involved the addition of biotinylated UTP. The biotinylated cRNA was purified using the SuperArray ArrayGrade cRNA cleanup kit (Supera rray Bioscience) according to the manufacturerÂ’s recommendations and quantitated on a Nanodrop Spectrophotometer as above. The biotin ylated cRNA was used for a pathwayspecific Oligo GE Array Mouse In flammatory Cytokines and Receptors Microarray according to manufactu rerÂ’s recommendations (Superarray Bioscience Corp., complete gene listi ng in Appendix C). Briefly, after hybridization of 1.5 ug of biotin-labeled cRNA to the microarray, it was washed, blocked to reduce background staining, conjugated with alkaline phosphataseconjugated streptavidin, and the image was captured wi th X-ray film following addition of chemiluminescent substrat e. Data were analyzed with GEArray Expression Analysis Suite (GEASuite) Online Image Data Acquisition and Analysis Software (SuperArray Bioscienc e, version 1.1, Feb. 26, 2005) Quantitative Real-Time PCR (q RT-PCR) DNase treated RNA (500 ng) prepared as described above from the ZA and ZD mice was used as a template for t he preparation of cDNA using the iScript
51 cDNA Synthesis kit (Bio-Rad Laboratories, Inc.). The iScript cDNA Synthesis kit uses random hexamer and oli go dT primers for the synthesis of cDNA from all RNA transcripts by the enzyme reverse trans criptase. Ten-fold se rial dilutions of cDNA simultaneously prepared from samp les known to contain the gene of interest functioned as standards in the q RT-PCR. cDNA samples were diluted 1:50 and used for q RT-PCR performed usi ng cDNAs plus gene-specific primers and iQ SYBR Green Supermix detection system (Bio-Rad). Primers were designed using Primer Express Software (version 2.0, Applied Biosystems) and purchased from MWG Biotech. In constr ucting the primers, the entire sequence for the gene of interest was obtained fr om GenBank database (available through NCBI), the glycine (G) and cyt osine (C) content were lim ited to 30% (with G/CÂ’s at the 5Â’ end), the forward and revers e primers were matched for optimum temperatures, and length was a minimum of 19 and maximum of 22 nucleotides. The forward and reverse primer sequenc es obtained were blasted through the GenBank database to ensure that the primary matches obtained were the genes of interest. Primer sequences des igned for chemokine ligand 25 (CCL25), chemokine ligand 17 (CCL17 also know n as TARC), chemokine ligand 8 (CCL8 also known as MCP-2), interl eukin 18 (IL-18), interleukin 1 (IL-1 ), and 18S rRNA are listed in Table 2. A melt curve was done for each primer set prior to its use for q RT-PCR. Primers for metalloth ionein (MT) are described elsewhere (130). The dilutions of cDNA sample s were compared to threshold cycles obtained from the 10-fold serial diluti ons of standard cDNA curves during the
52 exponential phase of product synthesis. Fluorescence generated from the SYBR Green dye intercalation and primer me lt curves were measured using a Table 2. Q-RT PCR primer sequences Gene Orientation Sequence Chemokine ligand 25 (CCL25) Forward Reverse 5 -ccaccaacgtcccagcatgt-3 5 -ggtgagtgggagggccttta-3 Chemokine ligand 17 (CCL17 or TARC) Forward Reverse 5Â’-gagctggtataagacctcagtggag-3Â’ 5Â’-tggccttcttcacatgtttgtc-3Â’ Chemokine ligand 8 (CCL8 or MCP-2)) Forward Reverse 5Â’-tgctttcatgtactaaagctgaaga-3Â’ 5Â’-ctacacagagagacataccctgctt-3Â’ Interleukin 18 (IL-18) Forward Reverse 5Â’-gaaccccagaccagactgataata-3Â’ 5Â’-cttgttcttacaggagagggtagaca-3Â’ Interleukin 1 (IL-1 ) Forward Reverse 5Â’-tgggcctcaaaggaaagaatc-3Â’ 5Â’-ggtattgcttgggatccacact-3Â’ 18S rRNA Forward Reverse 5Â’-cgaggaattcccagtaagtgc-3Â’ 5Â’-ccatccaatcggtagtagcg-3Â’ Bio-Rad iCycler with the following am plification sequence: 2 min at 50oC, 10 min at 95oC, followed by 40 cycles of 15 seconds at 95oC and 1 min at 60oC. Data were analyzed using iCycler iQ Optical System Software version 3.0a (Bio-Rad Laboratories, Inc.). The threshold for det ection was set 10 times greater than the baseline fluorescence signal. A duplicat e set of samples and standards were similarly and simultaneously plated for enumeration of 18S rRNA levels. Expression levels of all genes were normalized to 18S rRNA transcript levels. Statistical Analysis Differences between ZA and ZD groups were analyzed using an unpaired two-tailed StudentÂ’s t-test analysis when variances were equal. When variances were unequal, data were analyzed usi ng a nonparametric two-tailed t test. Standardized (ZD/ZA) cell phenotype data was analyzed using a one-sample t test with a hypothetical mean=1. Cell subpopulations were correlated with
53 chemokine mRNA transcript levels usi ng the Pearson correlation. Results are expressed as mean Â± SEM.
54 RESULTS Mouse Weights All mice gained weight over the 9week feeding protocol. There was no difference between the percent of baseline weight in the ZD versus the ZA mice at weeks 3, 6, or 9 of the feeding peri od (Fig. 2). Additionally, ZD mice showed no outward signs of zinc deficiency, such as parakeratosis or diarrhea. 50 75 100 125 150123456789Weeks fed diet% baseline weight ZA ZD 50 75 100 125 150123456789Weeks fed diet% baseline weight ZA ZD Figure 2. Percent baseline weight of BALB/c mice f ed a zinc-adequate (ZA) or zinc-deficient (ZD) diet. Val ues are expressed as mean Â± SEM. Baseline values represent 6 mi ce, ZA had n=11/group, and ZD had n=11/group. Numbers decreased over time as mice were killed at various time points over the 9 weeks, ending with an n=4 in each group. There was no difference between the percent of baseline weight in the ZD versus the ZA mice at week 3, 6, or 9 of the feeding period. Organ Weights Livers and thymuses were removed from ZA and ZD mice and weighed. No differences were identified in mean weight at any time point for either the livers or thymuses from the ZA or ZD mice.
55 Zinc Status Assessment Serum zinc concentrations decreased progressively over the 9-week feeding protocol in the ZD versus the ZA mice (Fig. 3A). Serum zinc was significantly lower in ZD mice ( P <0.01) at 6 and 9 weeks than ZA mice. Relative colonic MT mRNA gene expression wa s determined by normalization with 18S rRNA. Relative colonic MT mRNA levels of ZD mice were more than 3-fold lower than ZA mice (Fig. 3B, P <0.05). 0 0.2 0.4 0.6 0.8 1 1.2 1.4 ZA ZD Relative MT values 0.0 0.2 0.4 0.6 0.8 1.0 0246810 Weeks fed DietZinc concentration (ug/ml) ZA ZDA B * ** 0 0.2 0.4 0.6 0.8 1 1.2 1.4 ZA ZD Relative MT values 0.0 0.2 0.4 0.6 0.8 1.0 0246810 Weeks fed DietZinc concentration (ug/ml) ZA ZDA B * ** Figure. 3. Serum zinc concentrations and colonic metallothionein mRNA levels. Serum zinc concentrations over 9 w eeks (A, n=6 at baseline, n=11 ZA, n=11 ZD) and relative colon metallothionein (MT) mRNA gene expression at week 9 (B, n=4/diet group) in BALB/c mice fed a zincadequate (ZA) or zinc-deficient (ZD) diet. Zinc concentrations were measured by atomic absorption spectrophotometry, and each point (Fig. 2A) represents a different mous e. Serum zinc was significantly lower (* P <0.01, ZD vs. ZA) at 6 and 9 w eeks in mice fed the ZD diet. Colon MT expression was evaluated with q RT-PCR and normalized to 18s rRNA. MT values were significantly lower with ZD (** P <0.05, ZD vs. ZA). Values (Fig. 2B) repr esent the relative mean Â± SEM. Flow Cytometry Overall cells numbers obt ained and lymphocyte marker expression were evaluated in cellular populations isol ated from all tissues (MLN, small
56 069 0 5 10 15 20 25 30 35 40 45ZA ZD Weeks fed dietMLN cell numbers (x 106) 069 0 25 50 75 100 125Weeks fed dietPeyer's patches cell numbers (x 105) 069 0 5 10 15 20 25 30 35Weeks fed dietSI IEL cell numbers (x 105) 069 0 10 20 30 40 50 60 70Weeks fed dietLPL cell numbers (X 105) 069 0 10 20 30 40 50 60Weeks fed dietcIEL cell numbers (x 104)A B C D E Figure 4. Total recovered cell numbers in GALT. Total cells recovered from mesenteric lymph nodes (A), Peyer's patches (B), and small intestinal intraepithelial lymphocytes ( SI IEL, C), lamina propria lymphocytes (LPL, D), and colon intraepithelial lymphocytes (cIEL, E) in BALB/c mice fed a zinc-adequate (Z A) or zinc-deficient (ZD) diet. There was no affect of diet on overall number of cells recovered from the various tissues. There was a significant week effect in Peyer's patch and small intestinal intraepithelial lymphocytes ( P <0.05). Data represent the mean Â± SEM, n=2 to 6 mice per diet/week.
57 intestinal PeyerÂ’s patches, IEL, and LPL, and cIEL) during the 9-week zinc deficiency. The total number of cells extracted from each tissue was determined for GALTs from ZA and ZD mice at weeks 0, 6, and 9. There was no effect of diet on total number of cells extracted from the mesenteric lymph nodes (Fig. 4A), PeyerÂ’s patches (Fig. 4B), IEL (Fig. 4C), LPL (Fig. 4D), and cIEL (Fig. 4E). There was a significant week effect, with total number of cells extracted from PeyerÂ’s patches and IEL lymphocytes decreasing with fewer cells in later weeks in comparison to week 0 (baseline), P =0.001 and P =0.03, in PeyerÂ’s patch lymphocytes and IEL, respectively. Relative CD3+ levels (ZD/ZA) as a percent of total cells increased with the progressive zinc deficiency in cIEL (Fig. 5, P =0.059, R2=0.54). cIEL CD3+ cells as a percent of total cells was greater at week 9 in ZD versus ZA mice (42.8 Â± 3.9 versus. 29.1 Â± 2.5, respectively, P =0.04). The percentage of total CD3+ cells when analyzed independently of other cell ma rkers was not significantly different in the other GALT examined, although di fferences were identified for several CD3+ subpopulations in these tissues and are described below. The CD3+ subpopulation TCR CD8 + was altered by zinc deficiency in PeyerÂ’s patches and cIEL. The percentage of these CD3+ TCR CD8 + lymphocytes was greater at all time points (ZD/ZA>1.0, P <0.01) in cIEL, but lower at all time points (ZD/ZA<1.0) in PeyerÂ’s patches lymphocytes in ZD relative to ZA mice as a percentage of total extracted cells (Fig. 6, P <0.01). Alterations in other CD8 + subpopulations at week 9 of the zinc deficiency
58 r = 0.74, P = 0.0590.0 0.3 0.5 0.8 1.0 1.3 1.5 1.8 2.0 0246810Weeks fed dietRelative CD3+cIEL (ZD/ZA) r = 0.74, P = 0.0590.0 0.3 0.5 0.8 1.0 1.3 1.5 1.8 2.0 0246810Weeks fed dietRelative CD3+cIEL (ZD/ZA) Figure 5. Colonic CD3+ intraepithelial lymphocytes (c IEL) in BALB/c mice. cIEL CD3+ mice fed a zinc-deficient diet (Z D) relative to mice fed a zincadequate diet (ZA). The percentage of CD3+ cIEL increased in the colon with progressive zinc defici ency. n=2 at week 3 and 6, n=3 at week 9. Ratios based on percent ages of total cell population. 0 0.2 0.4 0.6 0.8 1 1.2 1.4 1.6 1.8 0246810Weeks fed dietRelative % total cells (ZD/ZA) Colon IEL Peyer'spatches 0 0.2 0.4 0.6 0.8 1 1.2 1.4 1.6 1.8 0246810Weeks fed dietRelative % total cells (ZD/ZA) Colon IEL Peyer'spatches Figure 6. Colonic intraepithe lial and small intestinal CD3+ TCR CD8 + Peyer's patches lymphocytes in BALB/c mice. CD3+ TCR CD8 + colonic intraepithelial and small intestinal Pe yer's patches lymphocytes in mice fed a zinc-deficient diet (ZD) relative to control mice fed a zincadequate diet (ZA). The percentage of CD3+ TCR CD8 + cells was higher in colonic IEL (ZD/ZA >1, n=7, P <0.01) but lower in Peyer's patch lymphocytes (ZD/ZA <1, n=7, P <0.01). Ratios based on percentages of total cell population.
59 protocol were also identified in PeyerÂ’ s patches and MLN as either a percentage of total cells or as a percentage of CD3+ cells and are listed in Table 3. No changes in CD8 + populations were identified at week 9 for the small intestine IEL or LPL populations. CD3+ TCR + CD8 lymphocyte subpopulations in PeyerÂ’s patches were significantly elevated as a percentage of total cells (Fig. 7, P <0.05), and as a percentage of CD3+ cells (9.6 Â± 0.6 versus 4.0 Â± 0.5, ZD and ZA, respectively, P =0.002) in the ZD versus the ZA mice at week 9. Changes were not observed in the CD3+ TCR + T-cell subpopulations, for the phenotypic markers specified above, at week 9 in the other GALT examined. 0.0 0.5 1.0 1.5 2.0 2.5 3.0 3.5 4.0 02369 ZA ZD*% total cellsWeeks fed diet 0.0 0.5 1.0 1.5 2.0 2.5 3.0 3.5 4.0 02369 ZA ZD*% total cellsWeeks fed diet Figure 7. Small intestinal Peyer's patch CD3+ TCR + CD8 -. Small intestinal Peyer's patch CD3+ TCR + CD8 lymphocytes as a percentage of the total recovered cell population in BALB/c mice fed a zinc-adequate (ZA) or zinc-deficient (ZD) diet. * P <0.05, ZD vs. ZA at week 9. Values represent mean Â± SEM, n=6, 1, 2, 2, and 3/group at week 0, 2, 3, 6, and 9, respectively. CD4+ T-cell subpopulations also h ad observed changes. Lymphocytes isolated from PeyerÂ’s patches in ZD mice had lower CD4+ CD8 + populations as
60 a percentage of CD3+ cells versus ZA mice at week 9 (Table 3). cIEL CD4+ lymphocytes as a percentage of CD3+ cells were lower in ZD versus ZA mice at 9 weeks (12.2 Â± 1.8 versus 18.9 Â± 0.5, respectively, P =0.02). Changes were not observed in these CD4+ subpopulations at week 9 in the other GALT examined. Table 3. CD8 + and CD4+ populations in various tissues after 9 weeks of zincadequate and zinc-deficient diet. Phenotype Tissue Gate ZA diet ZD diet P value CD3+ CD8 + MLN NG 20.8 Â± 0.7 18.4 Â± 0.4 0.04 CD3+ CD4CD8 + PP NG 2.8 Â± 0.3 1.5 Â± 0.1 0.01 TCR CD8 + PP CD3+ 11.6 Â± 2.0 5.2 Â± 0.6 0.04 CD4+ CD8 + PP CD3+ 19.6 Â± 0.1 14.3 Â± 0.8 0.003 CD4CD8 + PP CD3+ 10.6 Â± 0.6 5.64 Â± 0.5 0.003 CD4+ cIEL CD3+ 18.9 Â± 0.5 12.2 Â± 1.8 0.02 CD4+ CD8 cIEL CD3+ 8.5 Â± 1.7 3.7 Â± 0.8 0.06 Abbreviations: not gated, NG (% of tota l cells); colon intraepi thelial lymphocytes, cIEL; mesenteric lymph nodes , MLN; PeyerÂ’s patches, PP Changes were observed in T-ce ll subpopulations containing the costimulatory marker CD28+. PeyerÂ’s patches isolated from ZD mice had lower CD28+ CD4CD8 + populations as a percentage of CD3+ cells (14.0 Â± 2.4 versus 22.9 Â± 2.0, ZD and ZA, respectively, P =0.048). CD4+ CD28+ cell populations were lower in ZD mice as a percentage of CD3+ cells (4.4 Â± 0.5 and 10.4 Â± 0.2, ZD and ZA, respectively, P =0.0004) in cIEL. Further confirmation of these data was obtained when the CD3+ CD4+ CD28+ cell population was evaluated as a percentage of total cells, this population was also lower in ZD mice as a percentage of total cells (1.8 Â± 0.04 versus 3.1 Â± 0.3, ZD versus ZA, respectively, P =0.014). Changes were observed for three CD28+ cell populations isolated from the mesenteric lymph nodes. The MLN CD3+ CD28+ cell population was lower at week 9 in ZD versus ZA mice as a percentage of total cells (Fig. 8, P =0.03) or CD3+ cells (39.6 Â± 2.8 versus 62.1 Â± 7. 1, ZD versus ZA, respectively, P =0.04).
61 The MLN TCR + CD8 CD28+ CD3+ population was lower at week 9 in ZD mice as a percentage of total cells (0.1 Â± 0.1 versus 0.9 Â± 0. 2, ZD versus ZA, respectively, P =0.017) and as a percentage of CD3+ cells (13.4 Â± 10.9 versus 57.0 Â± 14.5, ZD versus ZA, P =0.074). The MLN TCR CD8 + CD28+ CD3+ cell population was higher at week 9 in ZD mi ce as a percentage of total cells (1.1 Â± 0.1 versus 0.6 Â± 0.1, ZD versus ZA, respectively, P =0.02) and as a percentage of CD3+ cells (6.1 Â± 0.4 versus 3.3 Â± 0. 6, ZD versus ZA, respectively, P =0.02). 0 10 20 30 40 50 60 70 023469Weeksfed diet% total cells ZA ZD 0 10 20 30 40 50 60 70 023469Weeksfed diet% total cells ZA ZD* Figure 8. Mesenteric lymph node (MLN) CD3+ CD28+ lymphocytes. MLN CD3+ CD28+ lymphocytes as a percentage of the total recovered cell population in BALB/c mice fed a zinc -adequate (ZA) or zinc-deficient (ZD) diet. * P <0.05, ZD vs. ZA at week 9. Data represent the mean + SEM, n=1 to 6 mice per data point. Microarray Analysis Agarose gel electrophoresis of mous e colonic RNA samples isolated for use in microarray showed that RNA prepar ed by this method electrophoresed to show two ribosomal RNA bands, 28S and 18S. Bands in all lanes were sharp and the 28S band was of higher intensity than the 18S band, suggesting that the
62 RNA preparation was intact, had not been degraded, and was of high quality. Genes most changed were determined by vi sual examination of microarrays, evaluation of raw numbers following no rmalization with selected housekeeping genes (GEASuite), and fold changes det ermined by GEASuite scatterplot analysis. Figure 9, A and B, show the microarrays for the w eek 3 ZA and ZD mouse samples. A differential pattern of gene expression was found between a ZA and ZD mouse at week 3 [Fig. 9A (ZA) and 9B (ZD)], indicating that changes in gene expression occur after only 3 weeks of diet consumption. Genes listed are those the most highly expressed in co mparison to the alternative diet at the same time point. At week 3 more genes were overexpressed in the ZA mouse in comparison to a ZD mouse. Microarray re sults of the week 9 colonic samples are shown in Figure 9C (ZA) and 9D (ZD). At week 9 genes that were overexpressed in the ZD mouse outnumbered those genes ov erexpressed in the ZA mouse. The percentage of genes that changed more than 1.5 fold according to the GEASuite scatterplot analysis for the week 3 microa rray comparisons of a ZA versus ZD mouse colonic sample was 36%, in cont rast to a total of 52% of genes differentially expressed for the same week 9 microarray comparisons. mRNA Expression of Chemoki ne Ligands and Interleukins Specificity and integrity of the designed primers was confirmed by the observation of a single product from the me lt curves. Relative levels of colonic mRNA transcript levels were obtained by normalization of mRNA transcript levels for the gene of interest with 18S rRNA transcript levels run simultaneously. Relative CCL25 mRNA levels were lower in ZD mice at week 9 versus ZA mice (Fig. 10, P =0.057). Relative mRNA transcript levels for CCL17, CCL8, IL-18,
63 C D A B A B C D E F G H I J K L M N O P 1 2 3 4 5 6 7 8 A B C D E F G H I J K L M N O P A B C D E F G H I J K L M N O P A B C D E F G H I J K L M N O P 1 2 3 4 5 6 7 8 1 2 3 4 5 6 7 8 1 2 3 4 5 6 7 8 Position Genes Differentially Expressed Panel E: Wk 3 K2Interleukin 6 signal transducer(IL-6st) H3Interleukin 13 receptor, alpha 1(IL-13 Ra1) C2Chemokineligand6 (CCL6) A2Burkittlymphoma receptor 1 (Blr1) I3Interleukin 1 beta (IL-1 ) C7Chemokinereceptor 2(CCR2) C4 Chemokineligand8(CCL8) E6 Chemokineligand14(CXCL14) G7 Interleukin 12A (IL-12A) Panel F: B6Chemokineligand25 (CCL25) Wk 9 H4Interleukin 15(IL-15) G3Interleukin 10(IL-10) A8Chemokineligand19 (CCL19) L2LymphotoxinB (LT-b) I3Interleukin 1 beta (IL-1 ) C4Chemokineligand8(CCL8) E5Chemokineligand13 (CXCL13) B3Chemokineligand21a (CCL21a, Serine) I1Interleukin 18 (IL-18) C D A B A B C D E F G H I J K L M N O P 1 2 3 4 5 6 7 8 A B C D E F G H I J K L M N O P A B C D E F G H I J K L M N O P A B C D E F G H I J K L M N O P 1 2 3 4 5 6 7 8 1 2 3 4 5 6 7 8 1 2 3 4 5 6 7 8 Position Genes Differentially Expressed Panel E: Wk 3 K2Interleukin 6 signal transducer(IL-6st) H3Interleukin 13 receptor, alpha 1(IL-13 Ra1) C2Chemokineligand6 (CCL6) A2Burkittlymphoma receptor 1 (Blr1) I3Interleukin 1 beta (IL-1 ) C7Chemokinereceptor 2(CCR2) C4 Chemokineligand8(CCL8) E6 Chemokineligand14(CXCL14) G7 Interleukin 12A (IL-12A) Panel F: B6Chemokineligand25 (CCL25) Wk 9 H4Interleukin 15(IL-15) G3Interleukin 10(IL-10) A8Chemokineligand19 (CCL19) L2LymphotoxinB (LT-b) I3Interleukin 1 beta (IL-1 ) C4Chemokineligand8(CCL8) E5Chemokineligand13 (CXCL13) B3Chemokineligand21a (CCL21a, Serine) I1Interleukin 18 (IL-18) Figure 9. Colonic gene expression. Coloni c gene expression was evaluated using a pathway specific Oligo GEArray Mouse Inflammatory Cytokines & Receptors Microarray (S uperarray Bioscience) at 3 weeks (9A and 9B) and 9 weeks (9C and 9D) of diet in zinc adequate (ZA, 9A and 9C) and zinc deficient (ZD, 9B and 9D) BALB/c mice. Each microarray represents the RNA isol ated from a single mouse. Genes listed in the tables to the right are those most differentially expressed in ZD vs. ZA mice at week 3 (9E) a nd week 9 (9F) of diet consumption after normalization with selected housekeeping genes. and IL-1 were not different between ZA and ZD mice at week 9 in colonic tissue (Fig. 10). Relationships between cytokines , chemokines, serum zinc, and metallothionein concentrations were ev aluated through the use of correlations. CCL25 and CCL17 positively correlated with MT mRNA levels in ZA and ZD groups combined at 9 weeks [co rrelation coefficient (r)=0.85, P =0.02, n=7 and r=0.80, P =0.02, n=8, for CCL25 and CCL17, re spectively], but neither correlated
64 with serum zinc. Although relative mRNA le vels of IL-18 were not higher in ZD than ZA colonic tissue at week 9, relative IL-18 mRNA transcript levels positively correlated with the CD3+ TCR CD8 + lymphocyte subpopulation as a percentage of total cells in the ZA and ZD groups (r=0.81, P =0.049, n=6). Furthermore, in the ZD mice IL-18 mRNA transcript levels positively with MT mRNA levels (r=0.96, P =0.04, n=4). 0 0.5 1 1 .5 2 2.5*Relative valueCCL25CCL17CCL8IL-18 IL-1 ÃŸZA ZD 0 0.5 1 1 .5 2 2.5*Relative valueCCL25CCL17CCL8IL-18 IL-1 ÃŸZA ZD *Relative valueCCL25CCL17CCL8IL-18 IL-1 ÃŸZA ZD Figure 10 . Colonic chemokine/cytokine/ receptor mRNA. Relative transcript levels in colon of BALB/c mice 9 weeks after a zinc-adequate (ZA) or zincdeficient (ZD) diet. mRNA expr ession was evaluated for chemokine ligand (CCL) 25, CCL17, CCL8, interleukin (IL) 18, and IL-1 with q RT-PCR and normalized to 18s rRNA . Data represent mean Â± SEM, n=4/diet group. * P =0.057, ZD vs. ZA group.
65 DISCUSSION Although previous studies showed al terations of cell populations and cytokines during zinc deficiency (13, 16), the assessment of these changes occurred outside the GALT. In addition, numerous studies have demonstrated the symptomatic effects of zinc deficiency, such as parasitic infections, diarrhea, and intestinal morphological changes ( 14,105,106,131). These sym ptoms of zinc deficiency are specifically disruptive to t he gastrointestinal tract, yet no studies have evaluated the cellular phenotypic changes, or cytokine and chemokine levels in the colonic tissue. The purpose of this study was to determine whether a progressive zinc deficiency altered Tlymphocyte subpopulation distribution or cytokines and chemokines within the GA LT. This study demonstrated that a progressive zinc deficiency changed Tlymphocyte subpopulations distribution within the GALT and altered expression of cytokine genes in mouse colonic tissue, the site where many symptomatic effects of zi nc deficiency are primarily localized. Data from this study showed that BAL B/c mice fed a ZD diet less than 1 ppm for 9 weeks had no decrease in thymic we ight or body weight in comparison to mice fed a ZA diet. These data disagr ee with previously discussed work in A/J mice but are supported by a few studies in BALB/c mice (132, 133). Interpretation of these studies is complicated because most of these incorporate an immune challenge into their protocol. Two studies, Minkus et al. (133) and Boulay et al.
66 (132), found that zinc di ets containing 3-5 ppm zinc supported normal growth, Boulay et al. showed no effect of a ZD diet on thymus weight, and neither study showed that ZD mice had impaired imm une responses to a nematode challenge. In contrast to these studies, several studi es by Shi et al. (73,102) in BALB/c mice showed that a ZD diet containing less than 1 ppm zinc, resulted in impairment of weight gain in BALB/c mice and a subseque nt inability to mount an effective immune response to nematode challenge. Str and et al. (134) found that addition of a stress, a pulmonary pneumococcal c hallenge, to the BALB/c model resulted in weight loss in a low zinc diet (le ss than 2 ppm). Data in BALB/c mice are inconclusive, and may depend on the particular stress and level of zinc deficiency to which the mice are exposed. These data suggest that zinc deficiency in some BALB/c models had no effect on body or thymus weight, but since most of these models contain an immune challenge com ponent the timing and type of challenge may make the in terpretation of the data difficult. Additional factors contributing to the la ck of effect of zinc deficiency on body and thymus weights in the present study could be genetic differences between strains and the housing conditions instituted at the University of Florida. The studies in which zinc deficiency consist ently leads to decreased thymic weight and body weight losses in the absence of a stress were done in female A/J mice. A/J mice have T-cell responses dist inguished by a high level of TH1 cytokines (135). Prasad et al. (13) suggest that zi nc deficiency has a bigger impact on the TH1 cytokine profile, suggesting that a TH1-biased strain may be more impacted by zinc deficiency. BALB/c mice, which appear to have less defined weight loss
67 effects during zinc deficiency, are a TH2-biased strain (136). BALB/c mice may therefore be less susceptible to zinc defic iency. It was also shown that there are genetic differences in retention of 65Zn when exposed to a ZD diet (137). Outbred mice retained more and excreted less 65Zn following exposure to a ZD diet than a TH1-strain (137). One final considerati on is the newer husbandry practices in rodent care at universities. Many init ial studies were done prior to the SPF conditions under which most mice ar e now shipped and maintained through acclimation. In this study, once the mice were given the experimental diet they were housed individually in stainless steel wire hanging cages; however, individuals entering the area still followed SPF gowning procedures. The mice are therefore exposed to fewer immune cha llenges in more recent studies than older studies. This may explain the di fferences obtained in body and thymus weights for the A/J mice in the studies described above. If housing procedures vary at different institutions this may ex plain some of the inconsistencies in data obtained with BALB/c mice. In evaluation of weight differences between ZA and ZD mice in different studies it is import ant to consider the zinc concentration of the diet and weeks mice were fed the die t, strain of mice, and housing conditions (indicating potential pathogen exposure). Liver weight was found to be unaffected by zinc status in the current study. Previous studies evaluating the effect of zinc status on liver weight are contradictory. Again, this may be because many zinc deficient studies incorporate an immunological challenge ma king the data difficult to interpret. Zhou et al. (138) found that rats fed a marginal ZD di et of 7 ppm for 5 weeks
68 showed no change in liver weight. In contra st, Shi et al. (101) found that female BALB/c mice fed a 0.75 ppm ZD diet had dec reased liver weight at all timepoints weeks 6-9. Interestingly, in these mice the ZD diet also led to weight loss, loss of hair, decreases in thymus weight, along with the decreased liver weight at week 7-9 (101). Furthermore, a s ubset of these mice expo sed to a nematode challenge showed the ZD mice had increased worm burdens and were thus more susceptible to parasitic challenge (101). B oulay et al. (132) showed that during a primary infection against a nematode, BALB/c mice f ed a 3 ppm ZD diet had increased liver weight compared to mice fed the 60 ppm ZA diet. The zinc deficient status of the mice was confirmed by serum zinc levels that were 64% lower in ZD mice than t he ZA mice. To further access the zinc status of the mice used in this study , MT mRNA levels were analyzed. mRNA levels for MT, a zinc-regulated zinc storage protein, dec reased during zinc deficiency in agreement with previous studies (71,72,118). These data confirm that the zinc deficient mice were indeed zinc deficient. Data from this study show ed that there was no affect of 9 weeks of diet on the total number of cells extracted from each of the tissues evaluated, i.e. mesenteric lymph nodes, PeyerÂ’s patches, IEL, LPL, and cIEL. Extracted IEL and cIEL were passed through nylon wool co lumns, which are intended to remove contaminating B cell populati ons and epithelial cells; therefore, contrary to the other tissues evaluated (mesenteric lymph nodes, PeyerÂ’s patches, and lamina propria lymphocytes) whose numbers refl ect the entire lym phocyte populations and some additional leukocytes, IEL and cIEL numbers should be representative
69 of a more purified T-lym phocyte population. No studies to date have evaluated the effect of zinc deficiency on GALT numbers; however, systemically zinc deficiency is often associated with decr eased lymphocyte populations. King et al. (15) showed that this system ic lymphopenia in zinc-deficient A/J or CAF1/J mice is associated with a 34-38% thymic involution and a corresponding 50-300% increase in apoptosis of pre-T cells. Si milar apoptotic mechanisms are thought to have the same affect on pre-B cells in the marrow, leading to a loss of pre-B cells in zinc deficiency (139). The lack of changes in the total number of cells extracted from the tissues in this study is consistent with the lack of change in thymus weight observed in these data. The failure to observe changes in cellular numbers is in agreement with King and Fr aker who showed no overall cell number changes, only a redistribution of cell ular populations (15,70). The inability to see observable differences in total ce ll numbers in the GALT extracted from ZA and ZD mice suggests that even if systemic changes did occur, these changes were not reflected in the tissue ce llular concentrations. Additionally, zinc deficiency in the strain of mice used in the current study may not have induced systemic changes in cell numbers in contrast to data obtained in other strains of mice. Flow cytometric analysis indicated a larger percentage of CD3+ cells were found in IEL isolated from the colonic tiss ue of zinc-deficient mice. The higher percentage of CD3+ TCR CD8 + cells isolated from cIEL of ZD mice may account for the observed increase in the CD3+ population. The corresponding decrease in this CD3+ TCR CD8 + cell population in the PeyerÂ’s patches and
70 other CD8 + populations in the PeyerÂ’s patches and MLN in ZD mice may indicate a movement of this cell populat ion from PeyerÂ’s patches and MLNs of the gastrointestinal tissue into the cIEL. The TCR CD8 + cell population, which was elevated in the cIEL and decreas ed in the PeyerÂ’s patches most likely represents conventional cytotoxic TCR + CD8 + lymphocytes, which are typically associated with peripheral immune function and the development of intestinal inflammation. These data are in agreement with Hosea et al. (17) who found a higher percentage of TCR + CD4CD8+ in thymocytes from ZD rats. A decrease in TCR + lymphocytes and increase in CD8 + lymphocytes are associated with increased pr oliferative capacity and IFNproduction, both proinflammatory conditions (140). TCR + CD8 + populations in the IEL often also have CD8 , a marker typically associated wit h IEL or cIEL. In this study we did not determine the CD8 status of the TCR + CD8 + population. CD8 and CD8 have different sensitivities to antigens, attributed to the chains they express on their surf ace and the signaling cascades that are activated following cell stimulation (141). The chain increases the sensitivity of the TCR, potentially contributing to the cytotoxic activity of CD8 + cells (142). IELs expressing CD8 have higher levels of Bcl-2 (a protein that is protective against apoptosis), lower expression of costimulatory marker s (like CD28), and lower production of pro-inflamma tory cytokines (141,143,144). The only changes in the TCR + lymphocyte population observed in this study were in the PeyerÂ’s patches of ZD mice where this population increased. This cell population ma y represent the TCR + CD8 + lymphocytes that are
71 thought to play a role in epithelial main tenance. Komano et al. (145) showed that TCR -/mice failed to regenerate intestinal epithelial cells that are constantly turning over in the gut mucosal epit helium and therefore determined that TCR + lymphocytes are essential for maintaini ng mucosal integrity. This was further supported by the work of Chen et al. ( 146) who showed that the production of keratinocyte growth factor by TCR + cells prevented dext ran sodium sulfate induced colitis. CD4+ lymphocytes were lower in cIEL of ZD mice, and the double positive CD8 + CD4+ (most likely an immature pre-T-cell population) was lower in the PeyerÂ’s patches of ZD mice. These fi ndings are in agreement with the systemic measurements of King et al. (15) which showed an increase in apoptosis of the CD8 + CD4+ cells and subsequent decreases in this pre-T-cell population in ZD mice. The data in the current study s uggest that some of the systemic changes occurring within zinc deficiency are per petuated into the GALT. Overall, these data suggest that within the GALT, zinc deficiency has the greatest impact on conventional T-cell populations, such as TCR CD8 + and CD4+ lymphocytes. Changes were observed in this study in the CD28 T-cell costimulatory receptor. The current data show PeyerÂ’ s patches from ZD mice had reduced CD3+ CD28+ CD4CD8 + populations as a percentage of total T cells (CD3+ cells), this may account for the decreased CD8 + population in this tissue from the ZD mice discussed above. These data also showed the CD3+ CD4+ CD28+ subpopulation decreased in cIEL isolated from ZD versus ZA mice, this population change may be responsible for the decrease identified in this study in
72 the CD3+ CD4+ cIEL population. These data indi cate that the PeyerÂ’s patch CD3+ CD8 + and cIEL CD3+ CD4+ subpopulation losses were specifically also CD28+. The tissue which expressed the larges t number of T-cell costimulatory marker changes was the mesenteric lymph nodes. Although no overall CD3+ changes were identified in the MLN T-cell population in dependent of other markers, decreases were identified in the CD3+ CD28+ cell population from ZD mice. These data suggest that although zi nc deficiency does not decrease the overall T-cell population, the T cells that are present express less CD28+ costimulatory receptor and are therefore less acti vateable suggesting a possibility of lack of stimulati on potential or anergy. The TCR + CD8 CD28+ CD3+ MLN subpopulation was lower in the ZD than the ZA mice, although the TCR + CD8 CD3+ subpopulation remained unchanged during ZD. This decrease identified in a CD28+ subpopulation of the TCR + T-cell population may not represent a large population. The TCR + population composed 7.0 Â± 0.48% of the CD3+ cell isolated from the mesenter ic lymph node in this study and according to Helgeland et al. (38) accounts for only 1% of total cells isolated from MLN of rats whose MLN is mainly composed of the conventional T-cell populations. Of all of the ti ssues evaluated only one CD28+ population, the TCR CD8 + CD3+ population, in the MLN was f ound to increase; however, it may not constitute a large population si nce decreases were obtained for the overall CD8 + CD3+ MLN population (discussed above) independent of the other cell markers. These data suggest that decreases in the CD28+ subpopulation of the conventional T-cell populations, CD4+ (cIEL), CD8 + (PeyerÂ’s patch), and
73 CD3+ (MLN), were found to be the most af fected by zinc deficiency and may be contributing to the losses in these Tcell subpopulations. This study did not identify changes in the CD28 status of the overall CD3+ population in the cIEL or in CD3+ TCR CD8 + cells, both of which were s hown in the current study to increase in the cIEL dur ing zinc deficiency. Previous studies found a lack of CD28 expression in aged patients or individuals with chronic inflamma tory diseases (147,148). The CD28T-cell population increases with increasing age, with newborns expressing only about 1% of CD28T cells, while centenarians express approximately 30% CD28T cells (148). Patients with rheumatoi d arthritis had higher percentages of circulating CD4+ CD28and CD8+ CD28than controls (149). The level of expression of CD28 has also been show n to decrease in ChagasÂ’ disease (a parasitic infection that can cause cardiac dysfunction) and CrohnÂ’s disease (150,151). Stimulation of T cells in the absence of CD28 results in nonresponsiveness in the T cell and Effros et al. (152) suggested that this may lead to a state of anergy. The progressive lack of CD28 expression due to zinc deficient status in the cu rrent study may be indicative of an inflammatory tendency without progression to an observa ble disease state, and may provide insight into a potential mechanism for muco sal barrier dysfunction as a result of zinc deficiency. CD28 is an essential disulfide-link ed homodimer on the su rface of T cells following interaction of the TCR with t he antigen/MHC or MHClike complex. The interaction of CD28 with its ligand is ess ential for the prolifer ation of the T cell,
74 production of IL-2, and formation of cytotox ic T cells (152). Azuma et al. (148) found that in the circulating TCR T-cell population of healthy adults CD4+ CD28T cells were uncommon; however, CD8+ CD28+ and CD8+ CD28were present at a ratio of appr oximately 2:1. There are fe w systemically circulating TCR T cells, but the authors did i dentify the presence of both CD28+ and CD28varieties within this T-cell subset (148). Evaluation of CD3+ CD28T-cell population locations (they were absent in the thymus and decreased in cord blood), large size, and additional marker s (CD54, CD58, and CD11a), led the authors to suggest that these cells may represent a memory T-cell population; however, further evaluation of CD45RO ex pression, the memory T-cell marker, did not identify differences for the memory marker between the CD3+ CD28+ and CD3+ CD28populations (148). Garcia de Tena et al. (150) did identify increases in the CD45RO marker in CD4+ CD28null T cells in patients with active CrohnÂ’s disease, indicating that this populat ion may represent a memory T-cell subpopulation. Functional inconsistencies also make determination of the CD28T-cell activity difficult. CD3+ CD28cells may be considered anergic because they showed lower proliferation, less than 5% expressed cytoplasmic IL-2 in response to anti-CD3 antibodies, and they had less markers of activation (such as CD69 and CD25) (148). This was in contrast to the CD3+ CD28+ cell population which had higher proliferative responses, cytopl asmic IL-2 present in 65% of their population, and expressed activation marker s (148). Frydecka et al. (153) found that CD28+ populations of CD3+ CD4+ and CD3+ CD8+ were lower in patients with B-cell chronic lympho cytic leukemia than healthy control subjects. In
75 addition, this cell population was slower to activate (as measured by CD28 density on the cell) and prolif erate in response to stimulation in culture in comparison to the same populations is olated from healthy control subjects, indicating a hyporesponsiveness (153). T hese data are inconsistent with the finding that CD3+ CD28cells have greater cytotoxic activity than their CD28+ counterparts (148). CD4+ CD28T cells have been shown to be highly autoreactive, and Vallejo et al. (147) s uggest that altering the CD28 subset can lead to Â“disease related immune abnorma litiesÂ”. Although the mechanism by which the CD28T-cell population causes immune dsyregulation is unknown, their increased presence in a number of diseases is well documented. The loss of CD28+ T-cell subpopulations in the PeyerÂ’ s patches, mesenteric lymph nodes, and cIEL of ZD mice in the current study provides a potential mechanism for the disruption of the mucosal ba rrier that occurs in zinc deficiency. These data suggest that zinc deficiency may be asso ciated with impaired T-cell activation within the GALT. Several mechanisms have been proposed for the modulation of CD28 expression in the above referenced studies . One theory is that the loss of CD28 is due to replicative senescence, or ce lls that have reached their Hayflick Limit, referring to the maximum number of cell divisions possible for a particular cell (152,154). This is support ed by cell culture analysis which showed decreases in CD28+ cells with increased population doubli ng (155). Effros et al. (152) proposed that this replicative senescence is due to telomere shortening (loss of DNA) during progresssive rounds of cell di vision. This theory is supported by
76 Monteiro et al. (156) who found that telomeres from CD8+ CD28T cells were 1.4 kb shorter than those from CD8+ CD28+ T cells, and estimated that the CD28T cells had experienced 14 to 28 more population doublings than the CD28+ T cells. Vallejo et al. (147) proposed that in chronic disease states this replicative senescence eventually leads to immune dysfunction. In addition to telomeric shortening, expression of CD28 is downregulated by TNFas previously described (127,153). Bryl et al. (127) us ing annexin and CSFE staining showed that TNFtriggered decreases in CD28 was not due to apoptosis or replicative senescence, respectively. The authors showed that TNFdownregulated CD28 expression by decreased transcription through inhibition of the binding of nuclear proteins to the CD28 gene promoter region (127). Furt hermore, it was mentioned above that patients with rheumat oid arthritis had decreased CD28+ expression on their CD4+ and CD8+ cells (149). Bryl et al. (149) showed that administration of anti-TNFtreatment for 1 week resulted in a 69% increase in CD28 molecules, and 56% of subjects showed improvement in disease. In the case of a repeated antigenic exposure, as might occur due to decreased tolerance or low dose repeated antigenic stimulation, expansion of the CD28population occurs as discussed in the Arosa et al. (50) studies above w here the intraepithelial CD8+ CD28population expanded following incuba tion with epithelial cells functioning as antigen-presenting cells. One additio nal mechanism for the decreased CD28 subpopulations includes the loss due to apoptos is (157). However, this loss is not specific for cell surface markers such as CD28, but would include decreases in
77 CD3, CD4, CD8, and CD56 as well as other cell surface molecules as the whole cell is lost (157). Insight into these mechanisms responsible for loss of CD28 expression and current knowledge about zinc deficiency allows speculation into the mechanism by which zinc deficiency may have caus ed CD28 losses in the current study. It was mentioned earlier that increased TNFis observed in zinc deficiency (98). Taylor et al. (158) showed that TTP-defic ient mice (TTP is highly expressed in the intestine) developed dermatitis and chronic skin inflammation along with other phenotypic changes, and that administration of anti-TNFantibody reversed the TTP-deficient phenotype. This reduction in TTP and resulting increase in TNFthat occurs in zinc deficie ncy may be responsible for the decreased CD28 in this work. Interestingl y, zinc deficiency is associated with parakeratosis and dermatitis and hyperkerato sis with areas of parakeratosis were found in TTP-deficient and STAT/CD28 k nockout mice, respectively (158,159). Additionally, since zinc deficiency has been shown to increase p56lck expression (72,117), this signal may be seen as a r epeated stimulation or a primary signal with no subsequent secondary signal, leadin g to loss of other costimulatory molecules and resulting in the development of an anergic, CD28-, cell population. The documented apoptosis that occurs in zinc deficiency may be an additional mechanism for CD28 loss, however, the CD3, CD4, CD8 populations in this study were not decreased in all tissues which had observed CD28 decreases as you would expect if these changes were due to apoptosis (157).
78 By using the pathway-centric microarray, a number of inflammatory-related genes whose expression in the colon appea red to be altered by 3 and 9 weeks of zinc deficiency were identified. Chem okines, cytokines and interleukins are proteins that act as mediators to orchestrate the homing and inflammatory responses of leukocytes. Their corresponding receptors are necessary for the interpretation of these signals by target ce lls. Relative expression levels of some of these genes were further assessed us ing q RT-PCR with 4 mice/group. q RTPCR confirmed initial microarray data of decreased expression of CCL25 mRNA in ZD mice in this study. CCL25 is a chemokine ligand expressed on intestinal epithelial cells and the endothelial cells of the small venuoles in the intestinal lamina propria in mice (160). CCL25 inte racts with the receptor CCR9, present on almost all T cells in the small inte stine and on a fraction of IgA producing antibody-secreting plasma cells (160). De creases between diet groups in this chemokine, important in homing, did not result in a decrease in overall numbers of IEL and LPL observed in this study (Fig. 4C, D, and E). One possible explanation is that differences in m RNA levels of CCL25 were observed but correlating changes in protein levels of CCL25 may not have been occurring. Future studies should evaluate CCL25 protein levels in colonic tissue during progressive zinc deficiency. The pres ence of IEL and LPL lymphocytes and IgA secreting plasma cells would be important in the response to inflammation or maintenance of the gut barrier. These data suggest a potential inability of ZD mice to produce sufficient transcripts fo r this essential chem okine ligand, leading
79 to possible reductions in IgA secreti ng plasma cells, alterations in the development of T and B cells, and a reduc tion in resident IEL T cells (65). In addition to CCL25, other genes we re selected for comparison of expression using q RT-PCR based on ident ification of differences on the microarray and for a variety of other reasons as well. For example, CCL17 (position A7, Fig. 9) was selected because the CCL17 gene maps to the same locus as MT (161), and therefore it is reasonable to hypothesize that it may be controlled by zinc as is MT. CCL17 mRNA was shown to be more highly expressed in the inflamed mucosa of patients with CrohnÂ’s disease (162). CCL-8 (Position C4, Fig. 9) was chosen becaus e it is secreted by many cell types including epithelial cells, and has an import ant role in influencing the migration of T cells and monocytes during inflammation (1 63). Van Coillie et al. (164) showed that IFN, produced by activated CD8 + cells (the population of cells which increased in the cIEL in ZD mice) and NK cells, cause upregulation of mRNA transcripts for CCL8. IL-1 (Position I3, Fig. 9) was selected because it provides an important cosignal in the activation of T lymphocytes, has a role in the induction of the TH1 response, has anti-tumor affects, and also increases the subpopulation of TCR + CD8+ cells when used for in vivo proliferation (165). IL18 (Position I1, Fig. 9) was chosen because it is a product of activated macrophages, increases the activity of NK and cytotoxic T lymphocytes (CTL), and like IL-1 has a role in the induction of the TH1 response and anti-tumor properties (166,167). The effects of IL18 are most potent when there is also costimulation with IL-12, this st imulates the activation of CD8+ T cells and the
80 subsequent production of IFN(168). Significant differences in CCL17, CCL8, IL-1 , and IL-18 gene expression were not obt ained by addition of more animals per group for q RT-PCR analysis. A lar ger sample population is needed to assess the significance of any of these trends, for example, based on these data the number of mice required to yield a si gnificant difference for the CCL17 based on 80% power and P <0.05 is 8 mice per group. The initial microarray data suggest the possibility that some of the cytokines, chemokines, and interleukins changed during the zinc deficiency, and this may play a role in the observed lymphocyte changes. It is also worth noting that the alternative can occur as well: changes in subpopulation dist ributions and activation can alter the cytokine, chemokine, or interleukin microenvironment. Correlations were identified between severa l of the factors evaluated in this study. A positive correlation was found between CCL25 mRNA levels and MT mRNA levels, this correlation supports t he finding that CCL25 mRNA levels were lower in ZD mice. MT was used in this study as a zinc status marker, because this zinc regulated gene is known to decreas e in zinc deficiency; therefore, it is logical that levels of MT mRNA woul d positively correlate with CCL25 mRNA levels if CCL25 mRNA expression is dependent of zinc st atus. CCL17 mRNA levels did not appear to be affected by dietary zinc status. However, CCL17 mRNA levels did correlate with MT mRNA levels independent of dietary zinc status. This may be attributed to the fact that both genes are found in close proximity to each other (161). Data in th is study showed that IL-18 mRNA levels positively correlated with the CD3+ TCR CD8 + cell population. These data
81 are in agreement with several studies wh ich have shown that increased IL-18 administration is associated with an increased CD8+ T-cell population (169,170). Administration of an antigen along with an adjuvant in BALB/c mice downregulated airway inflammation and led to increased mRNA levels of IL-18 (169). However, administration of ant i-CD8 antibody eliminated the beneficial reduction in the model of reactive ai rway disease, leading the authors to conclude that CD8+ T cells played a role in this reduction (169). An association was made between the increased CD8+ T-cell population and the increased IL-18 mRNA levels (169). Ju et al. (170) showed that genetically modified dendritic cells that overexpressed IL-18 when injected simultaneously with a tumor vaccine in C57BL/6 mice produced a higher percentage of tumor specific CD8+ T cells than when the tumor vaccine was given alone. An IL-18 treated mixed lymphocyte culture stimulated with IL-2 pr oduced a two-fold increase in CD8/CD4 ratio in comparison to an IL-2 stimulated n on IL-18 treated cultur e, with the actual number of CD8+ T cells increasing threefold (171) . Furthermore, the IL-18 treated CD8+ T cells upon stimulation had increased IFNproduction, upregulated IL-2R alpha chain expression, and increased cytotox ic activity (171). Depletion of the CD4+ T-cell population inhi bited expansion of CD8+ cells, indicating that IL-18 induced development of CD8+ T cells is CD4+ T-helper cell dependent (171). The correlations identified in the present st udy are consistent with expected results based on the literature and offer further validity to the data obtained. Combining the information already know n about changes that occur in zinc deficiency with the data pr esented in this study, a mechanism can be proposed
82 for the redistribution of the cellular populations incl uding changes in the cIEL populations (increased overall CD3+ and CD8 +, and decreased CD4+), in PeyerÂ’s patches populations (decreased CD8 + and CD4+), in CD28+ populations, for the changes in cytokines , chemokines, and receptors found via microarray, and for the change in CCL25 confirmed on q RT-PCR (Fig. 11). In the major cell populations for which decreases were identified in this study, i.e. in CD28 populations, CD3+ CD28+ CD4CD8 + (PeyerÂ’s Patches), CD3+ CD4+ CD28+ (cIEL), and CD3+ CD28+ (MLN, CD4 or CD8 status undetermined), zinc deficiency could lead to dysfunctional TTP (a zinc-finger protein which makes TNFmRNA unstable) and could lead to increased TNF, as previously shown to occur in zinc deficiency (98,124,126). TNFthen has the potential to disrupt the transcription of CD28 via alteration of protein binding at the promoter (127). At the same time the p56lck expression, associated with the CD4 or CD8 molecule, is increased in zinc defic iency (72,117); however, the absence of CD28 expression on these cells possibly pr events the formation of a lipid raft (172). Inability to form the lipid raft results in failure of the p56lck kinase to phosphorylate ZAP-70, resulting in no onward perpetuation of the signaling cascade, and a decrease in the transcription within the nucleus. The receipt of the primary signal increased p56lck, and absence of receipt of a secondary signal may render these cells anergic. Alternativel y, the decreased availability of zinc may prevent the p56lck/CD4 or CD8 metal binding complex to form and prevention of secondary signal transduc tion (119). Furthermore, van den Brandt et al. (173) showed that CD4 or CD8 ce lls lacking the costimulatory molecule
83 CD28 were more sensitive to glucocorticoid induced apoptosis. Since glucocortocoids are known to increase in zi nc deficiency (76), it is in this manner that the cell population identified above ma y decrease. The lack of a function in these cell populations, as a result of anergy or apopt osis, may alter chemokine and cytokine microenvironment. Alterations in cytokines and chemokines can influence the distribution of cellu lar populations within the GALT. TCRCD4 or 8 CD28 Transcription factor activity (NF ? B) Antigen presenting cell Costimulatory Molecules Cytokines Chemokines Activated cell Non-activated cell TCR CD4 or 8 p56lckZap70 Transcription Factor activity Antigen presenting cell Activated cell in zinc deficiency1osignal only??? No secondary signal CD28 AC BZnTTP-TNF-a TNF-a TTP TCR CD4/8 CD28 p56lckZap70 p56lckZap70 Lipid raft Zn2+Zn2+Zn2+ TCRCD4 or 8 CD28 Transcription factor activity (NF ? B) Antigen presenting cell Costimulatory Molecules Cytokines Chemokines Activated cell Non-activated cell TCR CD4 or 8 p56lckZap70 Transcription Factor activity Antigen presenting cell Activated cell in zinc deficiency1osignal only??? No secondary signal CD28 AC BZnTTP-TNF-a TNF-a TTP TCR CD4/8 CD28 p56lckZap70 p56lckZap70 Lipid raft p56lckZap70 Lipid raft Zn2+Zn2+Zn2+ Figure 11.Mechanism for CD4 and/or CD8 cellular changes during zinc deficiency. Panel A shows the nonactivated cell, with the T-cell receptor (TCR), CD4 or 8 mole cule, and the CD28 costimulatory molecule. The Zap-70 is not associated with the cytoplasmic tail of the receptor in the non-activated cell, p56lck is not active, and the lipid raft is not formed. The activated cell (Panel B) has engagem ent of the TCR and CD28, allowing proper signaling and formation of the lipid raft. The p56lck that is assocated with the CD4 or CD8, activates Zap-70 which continues this signaling cascade and re sults in increased transcription. In zinc deficiency (Panel C) the cell TTP becomes destabilized, increasing TNF, decreasing the transcription of CD28 (98,124,125). This reduction in the costimulatory molecule CD28, along with increased p56lck results in a primary but no secondary signal, failure to form the lipid raft, and lack of ZAP-70 transmission of signal (72,117,172). This lack of CD28 may result in anergy or increased sensitivity to glucocorticoid-induced apoptosis, altering the cytokine/chemokine mi croenvironment (173).
84 Potential consequences of the changes that occur in zinc deficiency when the data in this study are considered in light of what is already known to occur in zinc deficiency causes concern. In this study increases within the colon in the proinflammatory population CD8 (due to potential infilt ration from the PeyerÂ’s patches and MLNs), decreases in t he conventional CD4 population and CD28 costimulatory molecule, and failure to produce transcripts for CCL25 suggest an increase in inflammatory potential occurri ng simultaneous to decreased ability to respond to immune challenges. This dec reased presence of the conventional CD4 T-helper cell population, costimulator y molecule CD28 necessary for proper signaling, and decreased ability to influence homing via CCL25 mRNA expression may impair the ability of a zinc -deficient animal/human to respond to a mucosal challenge. In this way zinc def iciency tips the scale in priming the system towards potential inflammation, so that when a GALT challenge is encountered the mucosal immune system is more easily overcome (Fig. 12). colonInflammation C D 4 + p r o i n f l a m m a t o r y T c e l l s C D 8+ Zinc deficiency T c e l l a c t i v a t i o n ( C D 2 8 , p 5 6l c k) L y mp h o p e n i a A l t e r a t i o n o f ch e m o ki n e s a n d cy t o ki n e s A colonInflammation p r o i n f l a m m a t o r y T c e l l s C D 8+ Zinc deficiency + Immune challenge T c e l l a c t i v a t i o n ( C D 2 8 , p 5 6l c k) L y m p h o p e n i a A l t e r a t i o n o f c h e m o k i n e s a n d c y t o k i n e sB C D 4 + colonInflammation C D 4 + C D 4 + p r o i n f l a m m a t o r y T c e l l s C D 8+ Zinc deficiency T c e l l a c t i v a t i o n ( C D 2 8 , p 5 6l c k) L y mp h o p e n i a A l t e r a t i o n o f ch e m o ki n e s a n d cy t o ki n e s A colonInflammation p r o i n f l a m m a t o r y T c e l l s C D 8+ Zinc deficiency + Immune challenge T c e l l a c t i v a t i o n ( C D 2 8 , p 5 6l c k) L y m p h o p e n i a A l t e r a t i o n o f c h e m o k i n e s a n d c y t o k i n e sB C D 4 + Figure 12.Proposed model. Pre liminary data suggest that zinc deficiency in the absence of an immune challenge (12A) is associated with changes in T-lymphocyte subtypes, inflammatory chemokine and cytokine and/or their receptor/ligands, and T-cell acti vation markers that increase the susceptibility of the intestinal mu cosa and associated lymphoid tissue to inflammation. Zinc deficien cy with a subsequent immune challenge (12B) could tip the balance toward an overwhelming inflammatory response and tissue injury.
85 These data provide insight into the pathway through which zinc deficiency may increase susceptibility to parasitic challenge and become associated with increased gastrointestinal disorders. Future studies in the area of zinc deficiency and mucosal immunity should follow-up on some of the differences identif ied in this work. This study identified changes in T-cell subpopulat ions within the GALT tissue unique from those occurring systemically, along with GALT changes in the cytokine and/or chemokine expression as a result of zinc deficiency. A mechanism for the potential cytokine and/or chemokine driv en T-cell subpopulations changes should be evaluated for specific cellular populations important for prot ection from the effects of zinc deficiency. Furthermore, p56lck and TNFlevels should be evaluated simultaneously with CD28 expr ession to identify the potential mechanism of CD28 decreased expressi on in ZD mice. An interesting observation in this study was that male BALB/c mice did not experience any weight loss as a result of the zinc deficiency. Due to the TH2-biased responses of the BALB/c strain of mice, if the mechanism by which a zinc deficiency exerts its symptomatic effects is via TH1 pro-inflammatory cytokines, BALB/c mice might be more resistant to the effects of a zinc deficiency and it might be beneficial to repeat this work in a TH1-biased strain. Since many of the side effects of zinc deficiency occur in situati ons where the gut is expos ed to pathogens, the addition of a gut stress, and differential responses by TH1and TH2-biased mouse strains may provide insight into the pathways l eading to breakdown of the gut barrier that occurs during zinc deficiency. T he addition of antibodies against cytokines
86 and/or chemokines that may evoke cellula r changes may provide future therapies to alleviate the negative consequences of zinc deficiency. The BALB/c strain used in this study could be beneficial in that it allows the researcher to create a chronic zinc-deficient mouse model fo r experimentation wit hout induction of negative side effects such as diarrhea and par akeratosis. Further studies in this lab will confirm changes in gene expressi on as documented by the microarray, evaluate the resistance BALB/c mice to th e outward symptoms of zinc deficiency in comparison to other strains, evaluat e differences in the expression of cytokines, chemokines, and receptors betw een ZA and ZD mice with the addition of a stress to this model, and probe for the mechanism leading to CD28 T-cell receptor changes. These data suggest that zinc deficien cy alters cellular subpopulations and genes involved in the regulation of pathw ays important in protection of the mucosal gut barrier. These cellular changes may be the result of changes in the cytokine, chemokine, and/or their recept ors in the microenvironment, which have the potential to selectively attract and st imulate different lymphocyte populations. The mice in this study were exposed to a zinc-deficient diet for 9 weeks, sufficient to cause decreases in serum zinc levels and relative colonic MT levels, alterations to the GALT cytokine and chemokine microenvironment, and changes to conventional T-cell populations along with their costimulat ory marker CD28, but the ZD mice never ex perienced the other expected symptoms of severe zinc deficiency as described in previous studi es with other strains of mice (139). A
87 long-term zinc deficiency with no immune challenge in male BALB/c mice may be more comparable to a moderate zinc def iciency in other strains of mice.
88 Summary and Conclusions The altered microenvironment in the GALT during a 9-week progressive zinc deficiency results in changes in Tcell subpopulations within the MLN, small intestinal PeyerÂ’s patches, and cIEL. Thes e results also indicate that zinc deficiency alters colonic expression of chemokines, cytokines, and their receptors important in inflammatory pathw ays. Furthermore, these results show a decreased presence of the Tcell costimulatory activation marker CD28. Zinc deficiency is associated worldwide with an increase in parasitic infections. During chronic inflammatory diseases, such as viral and parasitic infections, it has been shown that there is an increased percentage of CD8+ CD28T cells (50). Therefore, alterations in chemokines and cytokines, and in T-cell subpopulations along with their decreased activation potential, may lead to increased susceptibility to a mucosal barrier disr uption and subsequent inflammation of the intestinal tissue. Previous evidence for the disturbance of the mucosal barrier in zinc deficiency includes the increased incidence of parasitic infections, morphological changes ident ified by histology, increased mRNA for TNF, and increased iNOS. Future research s hould focus on these altered T-cell subpopulations and identify t he potential mechanism by which zinc deficiency affects these T-ce ll populations.
89 APPENDIX A DIET COMPOSITION Zinc Adequate 0.85 ppm Zinc Zinc Deficient 30 ppm Zinc %gmkcalgmkcal Protein 2020.52020.5 Carbohydrate 66.36866.368 Fat 511.5511.5 Total 100100 kcal/gm 3.93.9 Ingredient gmkcalgmkcal Egg Whites, Spray Dried 200800200800 Corn Starch 150600150600 Sucrose 502.62010.4502.52010.2 Cellulose, BW200 500500 Corn Oil 5045050450 Mineral Mix S19401 350350 Vitamin Mix V10001 10401040 Choline Bitartrate 2020 Biotin, 1% 0.400.40 Zinc Carbonate, 52.1% Zinc000.0560 Total 1000390010003900 Diet contents were obtained from Research Diets.
90 APPENDIX B WEIGHT DATA OF ZINC ADEQUATE, ZINC DEFICIENT, AND PAIR-FED DIET GROUPS 50 75 100 125 150 123456789Week% baseline weightZA ZD PF 50 75 100 125 150 123456789Week% baseline weightZA ZD PF
91 APPENDIX C MOUSE INFLAMMATORY CYTOKINE AND RECEPTORS MICROARRAY KEY Microarray Position Map Gapdh 1 Blr1 2 C3 3 Ccl1 4 Ccl11 5 Ccl12 6 Ccl17 7 Ccl19 8 Ccl2 9 Ccl20 10 Ccl21a 11 Ccl22 12 Ccl24 13 Ccl25 14 Ccl3 15 Ccl4 16 Ccl5 17 Ccl6 18 Ccl7 19 Ccl8 20 Ccl9 21 Ccr1 22 Ccr2 23 Ccr3 24 Ccr4 25 Ccr5 26 Ccr6 27 Ccr7 28 Ccr8 29 Ccr9 30 Cx3cl1 31 Cx3cr1 32 Cxcl1 33 Cxcl10 34 Cxcl11 35 Cxcl12 36 Cxcl13 37 Cxcl14 38 Cxcl15 39 Cxcl2 40 Cxcl4 41 Cxcl5 42 Cxcl9 43 Cxcr3 44 Fcer1g 45 Fcgr1 46 Gpr2 47 Ifna2 48 Ifng 49 Igh-4 50 Il10 51 Il10ra 52 Il10rb 53 Il11 54 Il12a 55 Il12b 56 Il12rb2 57 Il13 58 Il13ra1 59 Il15 60 Il16 61 Il17 62 Il17b 63 Il17e 64 Il18 65 Il1a 66 Il1b 67 Il1r1 68 Il1r2 69 Il1rn 70 Il2 71 Il20 72 Il22 73 Il2rb 74 Il2rg 75 Il3 76 Il4 77 Il5 78 Il5ra 79 Il6 80 Il6ra 81 Il6st 82 Il8ra 83 Il8rb 84 Il9 85 Il9r 86 Itgam 87 Itgb2 88 Lta 89 Ltb 90 Mif 91 Nos2 92 AI875142 93 Rac1 94 Scye1 95 Spp1 96 Tgfb1 97 Tlr1 98 Tlr2 99 Tlr3 100 Tlr4 101 Tlr5 102 Tlr6 103 Tlr7 104 Tlr8 105 Tlr9 106 Tnf 107 Tnfrsf1a 108 Tnfrsf1b 109 Tnfsf5 110 Tollip 111 Xcl1 112 Xcr1 113 Blank 114 PUC18 115 Blank 116 Blank 117 AS1R2 118 AS1R1 119 AS1 120 Rps27a 121 B2m 122 Hspcb 123 Hspcb 124 Ppia 125 Ppia 126 BAS2C 127 BAS2C
92 Microarray Gene Identification Information Position GeneBank Symbol Formal Name Functional Importance 1 NM_008084 Gapdh Glyceraldehyde-3phosphate dehydrogenase Oxidoreductase activity;Mitochondrion;Glyceraldehyde-3phosphate dehydrogenase (phosphorylating) activity;Glucose metabolism;Glycolysis;Glyceraldehyde-3phosphate dehydrogenase activity; 2 NM_007551 Blr1 Burkitt lymphoma receptor 1 Integral to membrane;G-protein coupled receptor protein signaling pathway;Receptor activity;Integral to plasma membrane;Rhodopsin-like receptor activity;Lymph gland development;B-cell activation;C-X-C chemokine receptor activity;Angiotensin type II receptor activity;G-protein coupled receptor activity;Purinergic nucleotide receptor activity, G-protein coupled; 3 NM_009778 C3 Complement component 3 Extracellular;Protein binding;Inflammatory response;Extracellular space;Endopeptidase inhibitor activity;Complement activation, alternative pathway;Complement activation, classical pathway;Complement activation;Positive regulation of phagocytosis;Positive regulation of type IIa hypersensitivity; 4 NM_011329 Ccl1 Expressed sequence BF534335 Signal transduction;Extracellular;Immune response;Protein binding;Extracellular space;Chemokine activity;Chemotaxis;Cytokine activity; 5 NM_011330 Ccl11 Small chemokine (CC motif) ligand 11 Signal transduction;Extracellular;Immune response;Inflammatory response;Extracellular space;Chemokine activity;Chemotaxis;Cytokine activity; 6 NM_011331 Ccl12 Chemokine (C-C motif) ligand 12 Signal transduction;Extracellular;Immune response;Inflammatory response;Extracellular space;Chemokine activity;Chemotaxis;Cytokine activity; 7 NM_011332 Ccl17 Chemokine (C-C motif) ligand 17 Signal transduction;Extracellular;Immune response;Extracellular space;Chemokine activity;Chemotaxis;Cytok ine activity;Cellular defense response; 8 NM_011888 Ccl19 Chemokine (C-C motif) ligand 19 Extracellular;Immune response;Inflammatory response;Extracellular space;Chemokine activity;Chemotaxis;Cytokine activity; 9 NM_011333 Ccl2 Chemokine (C-C motif) ligand 2 Signal transduction;Extracellular;Immune response;Protein binding;Inflammatory response;Extracellular space;Chemokine activity;Chemotaxis;C ytokine activity;Gprotein-coupled receptor binding;
93 10 NM_016960 Ccl20 Chemokine (C-C motif) ligand 20 Signal transduction;Extracellular;Immune response;Protein binding;Inflammatory response;Extracellular space;Chemokine activity;Chemotaxis;Cytokine activity; 11 NM_011335 Ccl21a Chemokine (C-C motif) ligand 21c (leucine) Extracellular;Immune response;Inflammatory response;Chemokine activity;Chemotaxis;Cytokine activity;Immune cell chemotaxis;Negative regulation of myeloid blood cell differentiation; 12 NM_009137 Ccl22 Chemokine (C-C motif) ligand 22 Signal transduction;Extracellular;Immune response;Inflammatory response;Extracellular space;Chemokine activity;Chemotaxis;Cytokine activity; 13 NM_019577 Ccl24 Chemokine (C-C motif) ligand 24 Extracellular;Immune response;Inflammatory response;Extracellular space;Chemokine activity;Chemotaxis;Cytokine activity; 14 NM_009138 Ccl25 Chemokine (C-C motif) ligand 25 Signal transduction;Extracellular;Immune response;Protein binding;Inflammatory response;Extracellular space;Chemokine activity;Chemotaxis;Cytokine activity; 15 NM_011337 Ccl3 Chemokine (C-C motif) ligand 3 Signal transduction;Extracellular;Immune response;Protein binding;Inflammatory response;Extracellular space;Chemokine activity;Chemotaxis;Cytokine activity; 16 NM_013652 Ccl4 Chemokine (C-C motif) ligand 4 Signal transduction;Extracellular;Immune response;Inflammatory response;Extracellular space;Chemokine activity;Chemotaxis;Cytokine activity; 17 NM_013653 Ccl5 Chemokine (C-C motif) ligand 5 Signal transduction;Extracellular;Immune response;Inflammatory response;Extracellular space;Chemokine activity;Chemotaxis;Cytokine activity; 18 NM_009139 Ccl6 Chemokine (C-C motif) ligand 6 Signal transduction;Extracellular;Immune response;Extracellular space;Chemokine activity;Chemotaxis;Cytokine activity; 19 NM_013654 Ccl7 Chemokine (C-C motif) ligand 7 Signal transduction;Extracellular;Immune response;Inflammatory response;Receptor activity;Membrane;Extracellular space;Heparin binding;Chemokine activity;Chemotaxis;Transporter activity;Transport;Cytokine activity; 20 NM_021443 Ccl8 Chemokine (C-C motif) ligand 8 Signal transduction;Extracellular;Immune response;Inflammatory response;Extracellular space;Heparin binding;Chemokine activity;Chemotaxis;Cytokine activity; 21 NM_011338 Ccl9 Chemokine (C-C motif) ligand 9 Signal transduction;Extracellular;Immune response;Extracellular space;Chemokine activity;Chemotaxis;Cytokine activity;
94 22 NM_009912 Ccr1 Chemokine (C-C motif) receptor 1 Integral to membrane;Protein binding;Inflammatory response;G-protein coupled receptor protein signaling pathway;Receptor activity;Rhodopsin-like receptor activity;Immune cell chemotaxis;CC chemokine receptor activity;G-protein coupled receptor activity;Neuropeptide Y receptor activity;Purinergic nucleotide receptor activity, G-protein coupled;Myeloid blood cell differentiation; 23 NM_009915 Ccr2 Chemokine (C-C motif) receptor 2 Integral to membrane;Immune response;Protein binding;Humoral immune response;Inflammatory response;G-protein coupled receptor protein signaling pathway;Chemotaxis;Defense response;Rhodopsin-like receptor activity;Hemopoiesis;C-C chemokine receptor activity;G-protein coupled receptor activity;Cytokine binding;Cellular defense response (sensu Vert ebrata);Perception of pain;Regulation of cell migration; 24 NM_009914 Ccr3 Chemokine (C-C motif) receptor 3 Integral to membrane;Protein binding;Gprotein coupled receptor protein signaling pathway;Receptor activity;Chemotaxis;Defense response;Rhodopsin-like receptor activity;CC chemokine receptor activity;G-protein coupled receptor activity;Neuropeptide Y receptor activity;Purinergic nucleotide receptor activity, G-protein coupled; 25 NM_009916 Ccr4 Chemokine (C-C motif) receptor 4 Integral to membrane;Inflammatory response;G-protein coupled receptor protein signaling pathway;Chemotaxis;Rhodopsinlike receptor activity;C-C chemokine recepto r activity;Angiotensin type II receptor activity;G-protein coupled receptor activity; 26 NM_009917 Ccr5 Chemokine (C-C motif) receptor 5 Integral to membrane;G-protein coupled receptor protein signaling pathway;Chemotaxis;Defense response;Rhodopsin-like receptor activity;CC chemokine receptor activity;Cell surface;G-protein coupled receptor activity; 27 NM_009835 Ccr6 Chemokine (C-C motif) receptor 6 Integral to membrane;Protein binding;Gprotein coupled receptor protein signaling pathway;Receptor activity;Rhodopsin-like receptor activity;C-C chemokine receptor activity;G-protein coupled receptor activity; 28 NM_007719 Ccr7 Chemokine (C-C motif) receptor 7 Integral to membrane;Immune response;Gprotein coupled receptor protein signaling pathway;Receptor activity;Extracellular space;Chemotaxis;Rhodopsin-like receptor activity;C-C chemokine receptor activity;Gprotein coupled receptor activity;
95 29 NM_007720 Ccr8 Chemokine (C-C motif) receptor 8 Integral to membrane;Protein binding;Gprotein coupled receptor protein signaling pathway;Chemotaxis;Defense response;Rhodopsin-like receptor activity;CC chemokine receptor activity;G-protein coupled receptor activity; 30 NM_009913 Ccr9 Chemokine (C-C motif) receptor 9 Integral to membrane;Protein binding;Gprotein coupled receptor protein signaling pathway;Receptor activity;Chemotaxis;Defense response;Rhodopsin-like receptor activity;CC chemokine receptor activity;G-protein coupled receptor activity; 31 NM_009142 Cx3cl1 Chemokine (C-X3-C motif) ligand 1 Integral to membrane;Cell adhesion;Signal transduction;Extracellular;Immune response;Protein binding;Membrane;Chemokine activity;Cytokine activity; 32 NM_009987 Cx3cr1 Chemokine (C-X3C) receptor 1 Integral to membrane;G-protein coupled receptor protein signaling pathway;Receptor activity;Rhodopsin-like receptor activity;C-C chemokine receptor activity;Chemokine receptor activity;G-protein coupled receptor activity; 33 NM_008176 Cxcl1 Chemokine (C-X-C motif) ligand 1 Extracellular;Immune response;Cell growth and/or maintenance;Regulation of cell cycle;Inflammatory response;Intracellular;Extracellular space;Growth factor activity;Chemokine activity;Cytokine activity; 34 NM_021274 Cxcl10 Chemokine (C-X-C motif) ligand 10 Extracellular;Immune response;Inflammatory response;Extracellular space;Chemokine activity;Chemotaxis;Cytokine activity; 35 NM_019494 Cxcl11 Chemokine (C-X-C motif) ligand 11 Extracellular;Immune response;Inflammatory response;Extracellular space;Chemokine activity;Chemotaxis;Cytokine activity; 36 NM_021704 Cxcl12 Chemokine (C-X-C motif) ligand 12 Extracellular;Immune response;Germ cell development;Extracellular space;Growth factor activity;Chemokine activity;Chemotaxis;Cytokine activity;Regulation of cell migration;Brain development;T-cell proliferation;Positive regulation of cell migration;Induction of positive chemotaxis;Germ cell migration; 37 NM_018866 Cxcl13 Chemokine (C-X-C motif) ligand 13 Extracellular;Immune response;Inflammatory response;Chemokine activity;Chemotaxis;Cyt okine activity;Lymph gland development; 38 NM_019568 Cxcl14 Chemokine (C-X-C motif) ligand 14 Extracellular;Immune response;Extracellular space;Chemokine activity;Cytokine activity;
96 39 NM_011339 Cxcl15 Chemokine (C-X-C motif) ligand 15 Signal transduction;Extracellular;Immune response;Inflammatory response;Extracellular space;Chemokine activity;Chemotaxis;Cytokine activity;Hemopoiesis;Neutrophil chemotaxis; 40 NM_009140 Cxcl2 Chemokine (C-X-C motif) ligand 2 Signal transduction;Extracellular;Immune response;Inflammatory response;Chemokine activity;Chemotaxis;Cytokine activity; 41 NM_019932 Cxcl4 Chemokine (C-X-C motif) ligand 4 Extracellular;Immune response;Extracellular space;Heparin binding;Chemokine activity;Chemotaxis;Cytokine activity; 42 NM_009141 Cxcl5 Chemokine (C-X-C motif) ligand 5 Signal transduction;Extracellular;Immune response;Inflammatory response;Extracellular space;Chemokine activity;Chemotaxis;Cytokine activity; 43 NM_008599 Cxcl9 Chemokine (C-X-C motif) ligand 9 Extracellular;Immune response;Inflammatory response;Extracellular space;Chemokine activity;Cytokine activity; 44 NM_009910 Cxcr3 Chemokine (C-X-C motif) receptor 3 Integral to membrane;G-protein coupled receptor protein signaling pathway;Receptor activity;Chemotaxis;Defense response;Rhodopsin-like receptor activity;CC chemokine receptor activity;Chemokine receptor activity;C-X-C chemokine receptor activity;G-protein coupled receptor activity;Purinergic nucleotide receptor activity, G-protein coupled; 45 NM_010185 Fcer1g Fc receptor, IgE, high affinity I, gamma polypeptide Plasma membrane;Integral to membrane;Signal transduction;Cell surface receptor linked signal transduction;Receptor activity;Membrane;Integral to plasma membrane;Transmembrane receptor activity;Humoral defense mechanism (sensu Vertebrata);Neutrophil chemotaxis;Regulation of immune response;Antigen pres entation, exogenous antigen via MHC class II;External side of plasma membrane;Positive regulation of tumor necrosis factor-alpha biosynthesis;Lipid raft;Positive regulation of phagocytosis;Positive regulation of type IIa hypersensitivity;IgE binding;IgG binding;Positive regulation of interleukin-10 biosynthesis;Positive regulation of immune response;Antigen pres entation, exogenous antigen via MHC class I;Defense response to pathogenic bacteria;Phagocytosis, engulfment;Positive regulation of type III hypersensitivity;
97 46 NM_010186 Fcgr1 Fc receptor, IgG, high affinity I Integral to membrane;Cell surface receptor linked signal transduction;Receptor activity;Extracellular space;Receptor mediated endocytosis;Regulation of immune response;External side of plasma membrane;Positive regulation of phagocytosis;Positive regulation of type IIa hypersensitivity;Antigen presentation, exogenous antigen;IgG binding;IgG receptor activity;Antibody-dependent cellular cytotoxicity;Antigen pr esentation, exogenous antigen via MHC class I;Defense response to pathogenic bacteria;Phagocytosis, engulfment;Phagocytosis, recognition;Positive regulation of type III hypersensitivity; 47 NM_007721 Gpr2 G protein-coupled receptor 2 Integral to membrane;Protein binding;Gprotein coupled receptor protein signaling pathway;Receptor activity;Integral to plasma membrane;Chemotaxis;Defense response;Rhodopsin-like receptor activity;CC chemokine receptor activity;Cytosolic calcium ion concentration elevation;Chemokine receptor activity;Gprotein coupled receptor activity; 48 NM_010503 Ifna2 Interferon alpha family, gene 2 Extracellular;Extracellular space;Defense response;Cytokine activity;Hematopoietin/interferon-class (D200-domain) cytokine receptor binding; 49 NM_008337 Ifng Interferon gamma Regulation of cell growth;Extracellular;Immune response;Transcriptional activator activity;Extracellular space;Defense response;Regulation of transcription;Cytokine activity;Interferongamma receptor binding;Neutrophil chemotaxis;Positive regulation of transcription, DNA-dependent;Positive regulation of MHC class II biosynthesis;Positive regulation of isotype switching to IgG isotypes;Regulation of immune response;Programmed cell death, neutrophils;Positive regulation of interleukin12 biosynthesis;Positive regulation of interleukin-6 biosynthesis;Positive regulation of chemokine biosynthesis;Positive regulation of interleukin-1 beta secretion;Programmed cell death, inflammatory cells; 50 XM_111360 Igh-4 Immunoglobulin heavy chain 4 (serum IgG1)
98 51 NM_010548 Il10 Interleukin 10 Extracellular;Immune response;Extracellular space;Cytokine activity;P ositive regulation of MHC class II biosynthesis; 52 NM_008348 Il10ra Interleukin 10 receptor, alpha Integral to membrane;Cell surface receptor linked signal transduction;Receptor activity;Membrane;Integral to plasma membrane;Extracellular space;Hematopoietin/interferon-class (D200domain) cytokine receptor activity;Interleukin receptor activity; 53 NM_008349 Il10rb Interleukin 10 receptor, beta Integral to membrane;Protein binding;Cell surface receptor linked signal transduction;Receptor activity;Membrane;Integral to plasma membrane;Hematopoietin/interferon-class (D200-domain) cytokine receptor activity;Interleukin-10 receptor activity; 54 NM_008350 Il11 Interleukin 11 Extracellular space;Growth factor activity;Cytokine activity; 55 NM_008351 Il12a Interleukin 12A Extracellular;Immune response;Extracellular space;Growth factor activity;Cytokine activity;Interleukin-12 receptor binding; 56 NM_008352 Il12b Interleukin 12B Cell surface receptor linked signal transduction;Membrane;Extracellular space;Cytokine activity;Hematopoietin/interferon-class (D200-domain) cytokine receptor activity;Protein homodimerization activity; 57 NM_008354 Il12rb2 Interleukin 12 receptor, beta 2 Integral to membrane;Cell surface receptor linked signal transduction;Receptor activity;Membrane;Integral to plasma membrane;Extracellular space;Hematopoietin/interferon-class (D200domain) cytokine receptor activity;Interleukin receptor activity; 58 NM_008355 Il13 Interleukin 13 Extracellular;Immune response;Extracellular space;Cytokine activity;Interleukin-13 receptor binding;Hematopoietin/interferonclass (D200-domain) cytokine receptor binding; 59 NM_133990 Il13ra1 Interleukin 13 receptor, alpha 1 Integral to membrane;Cell surface receptor linked signal transduction;Receptor activity;Membrane;Integral to plasma membrane;Extracellular space;Hematopoietin/interferon-class (D200domain) cytokine receptor activity;Interleukin receptor activity;
99 60 NM_008357 Il15 Interleukin 15 Extracellular;Immune response;Cytokine activity;Lymph gland development;Hematopoiet in/interferon-class (D200-domain) cytokine receptor binding;Regulation of T-cell differentiation;Positive regulation of T-cell proliferation;NK T-cell proliferation;Extrathymic T-cell selection;Positive regulation of immune response;Regulation of antiviral response by host; 61 NM_010551 Il16 Interleukin 16 Protein binding;Intracellular;Cytokine activity;Immune cell chemotaxis;Induction of positive chemotaxis; 62 NM_010552 Il17 Interleukin 17 Extracellular space;Cytokine activity; 63 NM_019508 Il17b Interleukin 17B Receptor binding;Protein binding;Extracellular space;Cytokine activity;Neutrophil chemotaxis; 64 NM_080729 Il17e Interleukin 17E Inflammato ry response;Extracellular space;Cytokine activity;Response to pathogenic fungi;Interleukin-17E receptor binding;Eosinophil differentiation;Response to nematodes; 65 NM_008360 Il18 Interleukin 18 Extracellular;Immune response;Interleukin-1 receptor binding;Cytokine activity; 66 NM_010554 Il1a Interleukin 1 alpha Cell proliferation;Extracellular;Immune response;Regulation of cell cycle;Inflammatory response;Signal transducer activity;Growth factor activity;Interleukin-1 receptor binding;Cytokine and chemokine mediated signaling pathway;Cytokine activity; 67 NM_008361 Il1b Interleukin 1 beta Cell prolif eration;Extracellular;Immune response;Regulation of cell cycle;Inflammatory response;Signal transducer activity;Growth factor activity;Interleukin-1 receptor binding;Cytokine and chemokine mediated signaling pathway;Cytokine activity;Neutrophil chemotaxis;Positive regulation of interleukin-6 biosynthesis;Positive regulation of chemokine biosynthesis; 68 NM_008362 Il1r1 Interleukin 1 receptor, type I Integral to membrane;Signal transducer activity;Cell surface receptor linked signal transduction;Receptor activity;Membrane;Integral to plasma membrane;Extracellular space;Transmembrane receptor activity;Interleukin-1, Type I, activating receptor activity;Cytokine and chemokine mediated signaling pathway;Interleukin-1
100 receptor activity;Interleukin receptor activity; 69 NM_010555 Il1r2 Interleukin 1 receptor, type II ATP binding;Integral to membrane;Cell surface receptor linked signal transduction;Receptor activity;Membrane;Integral to plasma membrane;Extracellular space;Interleukin-1, Type II, blocking receptor activity;Transport;ATP-binding cassette (ABC) transporter activity;Interleukin-1 receptor activity;Interleukin receptor activity; 70 NM_031167 Il1rn Interleukin 1 receptor antagonist Extracellular;Immune response;Cell surface receptor linked signal transduction;Receptor activity;Integral to plasma membrane;Interleukin-1 receptor binding;Lipid metabolism;Insulin secretion; 71 NM_008366 Il2 Interleukin 2 Cell proliferation;Extracellular;Immune response;Inflammatory response;Extracellular space;Growth factor activity;Hormone activity;Defense response;Cytokine activity;Cellular defense response;Interleukin-2 receptor binding; 72 NM_021380 Il20 Interleukin 20 Extracellular;Immune response;Extracellular space;Cytokine activity;STAT protein nuclea r translocation; 73 NM_016971 Il22 Interleukin 22 Extracellular;Immune response;Protein binding;Cytokine activity;Oxygen and reactive oxygen species metabolism;Regulation of tyrosine phosphorylation of Stat3 protein; 74 NM_008368 Il2rb Interleukin 2 receptor, beta chain Integral to membrane;Cell surface receptor linked signal transduction;Receptor activity;Membrane;Integral to plasma membrane;Extracellular space;Hematopoietin/interferon-class (D200domain) cytokine receptor activity;Interleukin receptor activity; 75 NM_013563 Il2rg Interleukin 2 receptor, gamma chain Integral to membrane;Cell surface receptor linked signal transduction;Receptor activity;Membrane;Integral to plasma membrane;Extracellular space;Hematopoietin/interferon-class (D200domain) cytokine receptor activity;Interleukin receptor activity; 76 NM_010556 Il3 Interleukin 3 Extracellular;Immune response;Extracellular space;Growth factor activity;Cytokine activity;Interleukin-3 receptor binding; 77 NM_021283 Il4 Interleukin 4 Extracellular;Immune response;Extracellular space;Growth factor activity;Cytokine activity;Interleukin-4 receptor binding;Hematopoietin/interferon-class (D200-domain) cytokine receptor binding;Bcell activation;Negative regulation of
101 osteoclast differentiation;Positive regulation of MHC class II biosynthesis;Positive regulation of isotype switching to IgG isotypes;Regulation of immune response;Regulation of phosphorylation; 78 NM_010558 Il5 Interleukin 5 Extracellular;Immune response;Extracellular space;Growth factor activity;Cytokine activity;Interleukin-5 receptor binding; 79 NM_008370 Il5ra Interleukin 5 receptor, alpha Integral to membrane;Cell surface receptor linked signal transduction;Receptor activity;Membrane;Integral to plasma membrane;Extracellular space;Hematopoietin/interferon-class (D200domain) cytokine receptor activity;Interleukin receptor activity; 80 NM_031168 Il6 Interleukin 6 Extracellular;Immune response;Protein binding;Extracellular space;Acute-phase response;Growth factor activity;Cytokine activity;Interleukin-6 receptor binding;Negative regulation of chemokine biosynthesis;Programmed cell death, neutrophils; 81 NM_010559 Il6ra Interleukin 6 receptor, alpha Integral to membrane;Cell surface receptor linked signal transduction;Receptor activity;Membrane;Integral to plasma membrane;Extracellular space;Hematopoietin/interferon-class (D200domain) cytokine receptor activity;Interleukin receptor activity; 82 NM_010560 Il6st Interleukin 6 signal transducer Integral to membrane;Signal transduction;Receptor activity;Membrane;Extracellular space;Hematopoietin/interferon-class (D200domain) cytokine receptor activity;Regulation of Notch signaling pathway; 83 NM_178241 Il8ra Interleukin 8 receptor, alpha Integral to membrane;G-protein coupled receptor protein signaling pathway;Receptor activity;G-protein coupled receptor activity; 84 NM_009909 Il8rb Interleukin 8 receptor, beta Integral to membrane;G-protein coupled receptor protein signaling pathway;Receptor activity;Integral to plasma membrane;Chemotaxis;Defense response;Rhodopsin-like receptor activity;Interleukin-8 receptor activity;Chemokine receptor activity;Gprotein coupled receptor activity; 85 NM_008373 Il9 Interleukin 9 Extracellular;Immune response;Extracellular space;Growth factor activity;Cytokine activity;Interleukin-9 receptor binding;Hematopoietin/interferon-class (D200-domain) cytokine receptor binding; 86 NM_008374 Il9r Interleukin 9 Integral to membrane;Cell surface receptor
102 receptor linked signal transduction;Receptor activity;Membrane;Integral to plasma membrane;Hematopoietin/interferon-class (D200-domain) cytokine receptor activity;Interleukin receptor activity; 87 NM_008401 Itgam Integrin alpha M Integral to membrane;Cell adhesion;Protein binding;Receptor activity;Cell-matrix adhesion;Integrin-mediated signaling pathway;Integrin complex;Neutrophil chemotaxis;Cellular ex travasation;External side of plasma membrane;Opsonin binding; 88 NM_008404 Itgb2 Integrin beta 2 Integral to membrane;Cell adhesion;Protein binding;Development;Receptor activity;Cellmatrix adhesion;Extracellular space;Integrinmediated signaling pathway;Integrin complex;Neutrophil chemotaxis;Cellular extravasation; 89 NM_010735 Lta Lymphotoxin A Cell proliferation;Cell growth and/or maintenance;Humoral immune response;Inflammatory response;Positive regulation of cell prolif eration;Extracellular space;Tumor necrosis factor receptor binding;Cellular defense response;Lymph gland development;Programmed cell death, transformed cells; 90 NM_008518 Ltb Lymphotoxin B Plasma membrane;Integral to membrane;Lymph gland development; 91 NM_010798 Mif Macrophage migration inhibitory factor Inflammatory response;Cell aging;Regulation of cell proliferation;Cytokine activity;Isomerase activity;DNA damage response, signal transduction by p53 class mediator; 92 NM_010927 Nos2 Nitric oxide synthase 2, inducible, macrophage Electron transporter activity;Oxidoreductase activity;Nitric oxide biosynthesis;Electron transport;Superoxide metabolism;Calmodulin binding;Nitric-oxide synthase activity;NOT ovulation (sensu Mammalia);Defense response to bacteria;FMN binding; 93 NM_011101 AI875142 Expressed sequence AI875142 ATP binding;Transferase activity;Protein amino acid phosphorylation;Calcium ion binding;Nucleus;Protein binding;Cytoplasm;Diacylglycerol binding;Protein serine/threonine kinase activity;Intracellular signaling cascade;Protein-tyrosine kinase activity;Protein kinase activity;Induction of apoptosis by intracellular signals;Negative regulation of protein kinase activity;Neutrophil chemotaxis;Regulation of peptidyl-tyrosine phosphorylation;Positive regulation of inflammatory
103 response;Negative r egulation of glucose import;Protein kinase C activity;Calciumdependent protein kinase C activity;Induction of positive chemotaxis;Inactivation of MAPK;Negative regulation of insulin receptor signaling pathway; 94 NM_009007 Rac1 RAS-related C3 botulinum substrate 1 GTP binding;GTPase activity;Small GTPase mediated signal transduction;Cell adhesion;Protein binding;Cytoplasm;Intracellular signaling cascade;Inflammatory response;Response to wounding;Cell motility;Endocytosis;Dendrite morphogenesis;Cytoplasmic vesicle;Positive regulation of actin filament polymerization;Extrinsic to plasma membrane;Lamellipodium;Hyperosmotic response;Lamellipodium biogenesis; 95 NM_007926 Scye1 Small inducible cytokine subfamily E, member 1 Nucleic acid binding;Cytokine activity;TRNA binding;Protein biosynthesis;RNA binding; 96 NM_009263 Spp1 Secreted phosphoprotein 1 Cell adhesion;Protein binding;Extracellular space;Cytokine activity;Ossification; 97 NM_011577 Tgfb1 Transforming growth factor, beta 1 Protein amino acid phosphorylation;Cell proliferation;Protein binding;Negative regulation of cell prolif eration;Regulation of cell cycle;Inflammatory response;Extracellular matrix;Extracellular space;Organogenesis;Growth factor activity;Skeletal development;Defense response;Regulation of cell proliferation;Necrosis;Transforming growth factor beta receptor signaling pathway;Transforming growth factor beta receptor binding;Cell growth;Lymph gland development;Growth;Myogenesis;Regulation of myogenesis;Regulation of protein-nucleus import; 98 NM_030682 Tlr1 Toll-like receptor 1 Integral to membrane;Immune response;Inflammatory response;Receptor activity;Membrane;Transmembrane receptor activity; 99 NM_011905 Tlr2 Toll-like receptor 2 Plasma membrane;Integral to membrane;Immune response;Inflammatory response;Cell surface receptor linked signal transduction;Receptor activity;Membrane;Transmembrane receptor activity;Positive regulation of tumor necrosis factor-alpha biosynthesis; 100 NM_126166 Tlr3 Toll-like receptor 3 Integral to membrane;Immune response;Inflammatory response;Response
104 to virus;Receptor activity;Membrane;Extracellular space;Transmembrane receptor activity;Defense response; 101 NM_021297 Tlr4 Toll-like receptor 4 Integral to membrane;Immune response;Inflammatory response;Receptor activity;Membrane;Extracellular space;Transmembrane receptor activity;Metabolism;I-kappaB kinase/NFkappaB cascade;Catalytic activity;Toll signaling pathway; 102 NM_016928 Tlr5 Toll-like receptor 5 Nucleic acid binding;Integral to membrane;Immune response;Inflammatory response;Receptor activity;Membrane;Extracellular space;Transmembrane receptor activity;Actin binding; 103 NM_011604 Tlr6 Toll-like receptor 6 Integral to membrane;Immune response;Inflammatory response;Receptor activity;Membrane;Transmembrane receptor activity; 104 NM_133211 Tlr7 Toll-like receptor 7 Integral to membrane;Immune response;Inflammatory response;Receptor activity;Membrane;Extracellular space;Transmembrane receptor activity; 105 NM_133212 Tlr8 Toll-like receptor 8 Integral to membrane;Immune response;Inflammatory response;Receptor activity;Membrane;Extracellular space;Transmembrane receptor activity; 106 NM_031178 Tlr9 Toll-like receptor 9 Plasma membrane;Integral to membrane;Immune response;Inflammatory response;Response to virus;Receptor activity;Membrane;Extracellular space;Transmembrane receptor activity; 107 NM_013693 Tnf Tumor necrosis factor Plasma membrane;Cell proliferation;Immune response;Cell growth and/or maintenance;Humoral immune response;Inflammatory response;Positive regulation of cell prolif eration;Induction of apoptosis via death domain receptors;Signal transducer activity;Development;Positive regulation of I-kappaB kinase/NF-kappaB cascade;Integral to plasma membrane;Organogenesis;Tumor necrosis factor receptor binding;Defense response;Secretory granule;Regulation of cell proliferation;Cytokine and chemokine mediated signaling pathway;Cellular defense response;Lymph gland development;Cellular extravasation;Positive regulation of osteoclast differentiation;Programmed cell death, transformed cells;Regulation of
105 osteoclast differentiation; 108 NM_011609 Tnfrsf1a Tumor necrosis factor receptor superfamily, member 1a Plasma membrane;Integral to membrane;Signal transduction;Protein binding;Intracellular signaling cascade;Apoptosis;Inflammatory response;Cell surface receptor linked signal transduction;Receptor activity;Extracellular space;Cell death;Tumor necrosis factor receptor binding;Defense response;Tumor necrosis factor receptor activity;Lymph gland development;Prostaglandin metabolism; 109 NM_011610 Tnfrsf1b Tumor necrosis factor receptor superfamily, member 1b Integral to membrane;Cell proliferation;Inflammatory response;Cell surface receptor linked signal transduction;Receptor activity;Extracellular space;Necrosis; 110 NM_011616 Tnfsf5 Tumor necrosis factor (ligand) superfamily, member 5 Plasma membrane;Integral to membrane;Immune response;Membrane;Tumor necrosis factor receptor binding;Defense response;Cytokine activity; 111 NM_023764 Tollip Toll interacting protein Immune response;Inflammatory response;Signal transducer activity; 112 NM_008510 Xcl1 Chemokine (C motif) ligand 1 Extracellular;Immune response;Extracellular space;Chemokine activity;Chemotaxis;Cytokine activity; 113 NM_011798 Xcr1 Chemokine (C motif) receptor 1 Integral to membrane;G-protein coupled receptor protein signaling pathway;Receptor activity;Rhodopsin-like receptor activity;Chemokine receptor activity;Gprotein coupled receptor activity; 114 115 L08752 PUC18 PUC18 Plasmid DNA 116 117 118 SA_00005 AS1R2 A rtificial Sequence 1 Related 2 (80% identity)(48/60) 119 SA_00004 AS1R1 A rtificial Sequence 1 Related 1 (90% identity)(54/60) 120 SA_00003 AS1 Artificial Sequence 1 121 NM_024277 Rps27a Ribosomal protein S27a Intracellular;Protein biosynthesis;Ribosome; 122 NM_009735 B2m Beta-2 microglobulinPlasma me mbrane;Integral to plasma membrane;Extracellular space;Defense response;Cellular defense response;Antigen processing, endogenous antigen via MHC class I;MHC class I receptor activity;Antigen presentation, endogenous antigen; 123 NM_008302 Hspcb Heat shock protein ATP binding;Protein
106 1, beta binding;Mitochondrion;Response to heat;Protein folding;Unfolded protein binding;Response to unfolded protein;ATP binding;Protein binding;Mitochondrion;Response to heat;Protein folding;Unfolded protein binding;Response to unfolded protein; 124 NM_008302 Hspcb Heat shock protein 1, beta ATP binding;Protein binding;Mitochondrion;Response to heat;Protein folding;Unfolded protein binding;Response to unfolded protein;ATP binding;Protein binding;Mitochondrion;Response to heat;Protein folding;Unfolded protein binding;Response to unfolded protein; 125 NM_008907 Ppia Peptidylprolyl isomerase A Cytosol;Protein folding;Isomerase activity;Peptidyl-prolyl cis-trans isomerase activity;Cytosol;Protein folding;Isomerase activity;Peptidyl-prolyl cis-trans isomerase activity; 126 NM_008907 Ppia Peptidylprolyl isomerase A Cytosol;Protein folding;Isomerase activity;Peptidyl-prolyl cis-trans isomerase activity;Cytosol;Protein folding;Isomerase activity;Peptidyl-prolyl cis-trans isomerase activity; 127 SA_00007 BAS2C Biotinylated Artificial Sequence 2 Complementary sequence 128 SA_00007 BAS2C Biotinylated Artificial Sequence 2 Complementary sequence
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124 BIOGRAPHICAL SKETCH Kelli Herrlinger-Garcia was born in Bloomington, Indiana, and moved to Florida in 1973. She graduated from Ga inesville High School in 1989. Kelli earned an Associate of Arts degree in the summer of 1991 from Santa Fe Community College, and graduated with hi ghest honors in May of 1993 with a Bachelor of Science in interdiscipl inary studies with a concentration in biochemistry and molecular biology. Kelli worked for two years as a scientist for the ShrinerÂ’s Childrens Hospital-Tampa Un it. Kelli then began work for Dr. Bobbi Langkamp-Henken in the Food Science and Hu man Nutrition Depa rtment at the University of Florida, where she has been employed for more than 10 years. Kelli began the Master of Science program in the fall of 2003, while continuing to work full-time with Dr. Bobbi Langkamp-H enken. KelliÂ’s achievements include publication of 27 original research works and abstracts, and she was a 1997 University of Florida Superior A ccomplishment Award winner. During her graduate education she received the Willia m and Agnes Brown Scholarship; was nominated to Gamma Sigma De lta: The Honor Society of Agriculture; and was an American Society of Nutrition Procter & Gamble Graduate Student Competition Abstract Winner. In addition to her work, Kelli has a son, Austin, whom she coaches in YSI Incorporated and the City of Gainesvi lle soccer teams. Kelli will be awarded a Ma ster of Science degree in food science and human nutrition with a specialization in nutr itional science in August of 2006.