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26 Immunophysiology B. GLICK1 Department of Poultry Science College of Agricultural Sciences Clemson University Clemson, South Carolina 29634

I. Introduction 657 II. Cytoarchitecture and Development of the Immune System 657

continues to the calm (‘‘assimilation of new concepts’’), and is followed by the storm of LeSoir (‘‘challenges leading back to LeMatin’’).

A. Primary Immune Tissue 657 B. Secondary Lymphoid Tissue 659

II. CYTOARCHITECTURE AND DEVELOPMENT OF THE IMMUNE SYSTEM

III. Regulation of Immune Response 662 A. Major Histocompatibility Complex 662 B. Cytokines 664 C. Antibody-Mediated Immunity and B-Cell Repertoire 664 D. Macrophages, Natural Killer Cells, Heterophils, and Thrombocytes 666

The immune system is dependent on specialized microenvironments that (1) offer a primary educational milieu where pluripotent precursors will differentiate into clones of lymphocytes endowed with the ability to respond to self or foreign antigens, and (2) offer a secondary educational milieu in which primary educated lymphocytes gather with various accessory cells to respond to specific cell-associated antigens (or noncell associated antigens) and clonally expand. These microenvironments will be discussed under primary immune tissue and secondary lymphoid tissue.

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I. INTRODUCTION An understanding of avian immunology requires some incite into a history of immunology (Silverstein, 1989) and a comprehension of fundamental immunology (Paul, 1993). The primary purpose of this chapter will be to present a knowledge base for avian immunology by writing a brief narrative accompanied by limited illustrations and citing review articles and selected original research. Immunology, like the musical painting of Haydn’s Sixth, Seventh, and Eighth Symphonies, begins with the sunrise in LeMatin (‘‘new language and ideas’’),

A. Primary Immune Tissue The T- and B-cell concept entered the vocabulary of immunology only after basic research with the chicken model revealed an immunological role for the bursa of Fabricius (Glick et al., 1956; Warner et al., 1962; Cooper et al., 1966) and the avian thymus (Cooper et al., 1966; Figure 1). The B-lymphocyte of the concept was so named to identify its avian bursal origin and mammalian bone marrow origin and the T-lymphocyte of the concept identified its thymic origin (Roitt et al., 1969).

1 Retired, June 1995, as Distinguished Emeritus Professor, Mississippi State University; Emeritus Professor, Clemson University; and Adjunct Professor Biomedical Cooperative Greenville Hospital and Clemson University.

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plied to the vent of the chick, would enter the bursal duct and be pinocytosed by the FAE (Bockman and Cooper, 1973) stimulated a carbon-vent study in a 4week-old chicken that revealed an average of 800 FAE areas/fold or 8,000 to 12,000 bursal follicles per bursa considering the presence of 10 to 15 folds (Olah and Glick, 1978a).

2. Bursal B-Cell Markers

FIGURE 1 Primary immune tissue, thymus, and bursa of Fabricius and secondary lymphoid tissue, pineal, Harderian gland, accessory spleen, spleen, lymph node, and cecal tonsil.

1. Bursa of Fabricius: Morphology The initial descriptive study of the bursa was by Hieronymus Fabricius after whom the gland was named (Adelman, 1942). Bursa growth studies revealed (1) a rapid growth from hatch to 3 or 4 weeks, (2) a plateau period for the next 5 or 6 weeks, and (3) regression occurring before sexual maturity (Glick, 1956). These observations set the stage for the functional studies since they directed that bursectomies be performed prior to 3 weeks of age or as close to hatch as possible. Serendipity then entered the picture when bursectomized birds were injected with Salmonella pullorum to satisfy a student laboratory and not as a part of a designed experiment. A more complete description of these events may be found in selected publications (Glick et al., 1956; Glick, 1977, 1987). The bursa is a dorsal diverticulum of the proctodaeal region of the cloaca ( Jolly, 1915). The ovallike bursa of the chicken contrasts to the elongated bursae of the duck and starling (Sternus vulgaris) and to the ostrich (Struthio camelus australis) and emu (Dromaius novaehollandio) bursae, which are an integral part of the proctodaeal mucosa (Glick, 1986; von Rautenfeld and Budras, 1982). The bursal anlage appears between 3 and 5 days of embryonic development (DE) (Romanoff, 1960; Olah et al., 1986). A major feature of embryonic development is the formation of buds, the forerunner of the bursal follicle. The bud develops into the medulla of the bursal follicle. Scanning electron microscopy revealed two types of surface epithelium: follicleassociated-epithelium (FAE), associated with the medulla, and the interfollicular epithelium (IFE), which is between the follicles (Bockman and Cooper, 1973; Holbrook et al., 1974). The IFE and FAE morphologically may appear at 12 and 15 DE, respectively (Naukkarinen et al., 1978). However, the pinocytotic ability of the FAE (Bockman and Cooper, 1973) was not evident until 19–21 DE. The observation that carbon, ap-

The hallmark of B-cells is the presence of membraneassociated immunoglobulin. Membrane Ig in mammals (rodents/humans) occurs after the appearance of cytoplasmic Ig (pre-B-cells) while in birds there is no such sequence. Gilmour et al. (1976) produced alloantisera that revealed two independent autosomal loci, Bu-1 and Th-1, recognizing an antigen of bursal lymphocyte/ peripheral B-cells and thymic cells/peripheral T-cells, respectively. Allelic forms have been identified, Bu-1a (94 kDa) and Bu-lb (70 kDa), and utilized in bursal follicle colonization studies to suggest entry of no more than three precursors per follicle (Chen et al., 1991). A Bu-2 antigen (66 kDa) was shown to be distinct from Bu-1. The Bu-2 antigen identified both Ig⫹ and Ig- cells and in control bursae identified lymphocytes in the cortex and medulla but not the epithelium. Other B-cell antigens, the CB antigens, have been reviewed by Chen et al. (1987, 1991).

3. Thymus, T-cell Receptors, and Cluster of Differentiation The third and fourth pharyngeal pouches contributed to the formation of the thymus which consisted of seven lobes developed along each side of the jugular veins (Romanoff, 1960; Venzke, 1952). The thymus, like the bursa, possessed cortical and medullary regions. An isolated protein, the putative avian thymic hormone, located in the thymus and blood, stimulated bone marrow cells to express T-cell markers (Murthy et al., 1984) and revealed an amino acid sequence similar to parvalbumin (Brewer et al., 1990). Antigen recognition in T-cells differs from B-cells in that the T-cell receptor (TCR) (1) is not an immunoglobulin molecule, (2) recognizes surface-bound antigen only, and (3) is not secreted but remains an integral part of the cell membrane. A cluster of differentiation (CD) identifies in a cell membrane specific groups of determinates which define stages of cellular differentiation and are detected by monoclonal antibodies (Table 1). The human TCR complex includes an invariant fivepolypeptide complex (웂, ␦, ␧, ␨, ␩,) molecule termed CD3. A similar CD3 molecule was identified in chickens by Chen et al. (1986). The chicken CD3 possessed three

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TABLE 1 T-Cell Antigensa Molecular mass (ⴛ 10ⴚ3)

Antigens CT1 CD3 웂/␦ TCR 움/웁TCR 움/웁 TCR CD2 CD4 CD8 CD5

65 17, 19, and 20 90 (subunits 40 and 50) 90 (subunits 40 and 50) 88 (subunits 40 and 48) 40 64 64 (34 dimer) 56

CD28 CD45 CD25 (IL-2 receptor)

40 200 50

Antibody CT1, Ct1-움 CT3 TCR1 TCR2 TCR3 2–4 CT4 CT8 CTLA-5, CTLA-8, 3–8 AV7 L-17,CL1 INN-CH-16

a All cited in narrative with the exception of CT1, CD5, and CD25 (Chen et al., 1991).

polypeptides of Mr 20,000, 19,000, and 17,000 under nonreducing conditions. Avian homologies to mammalian T-cell receptor 웂/ ␴ (TCR1) and TCR 움/웁 (TCR2) have been identified (Chen et al., 1991). While TCR1- and TCR2-positive thymocytes appeared at 12 DE, only TCR1 was detected by surface staining and these were equivalent to the number of CD3 surface-stained thymocytes (Bucy et al., 1990). TCR1 and TCR2 were heterodimers of 50- and 40-kDa glycoproteins. A third TCR had a lower molecular weight (Chen et al., 1991). Cells positive for TCR1 and TCR2 migrate to the spleen by 15 and 19 DE, respectively, while the TCR3 positive cells do not appear in the spleen until after hatching. The range of TCR1, -2, and -3 in peripheral blood was 15–25, 45–55, and 10–15%, respectively (Chen et al., 1991). Precursor cells entered the thymus in three waves. All three lineages were present in the first and second arrivals of precursor cells (Chen et al., 1991). A CD2 antigen (monomeric, 40 kDa), identified by mAb 2-4, appeared in the avian thymus by 11 DE and may influence T-cell growth and differentiation by way of cell adhesion. Avian thymocytes are 98% CD2 positive. CD2 is a coreceptor cooperating with the TCR and contributing to T-cell binding to antigen-presenting cells and/or T-cell signaling. There may be a question concerning the identification of avian CD2 (Young et al., 1994). Young et al. (1994) identified a 40-kDa molecule with a 50% amino acid sequence identical to mammalian CD28. In mammals, CD28 receptors of primed T-cells bound a B7 epitope present in B-cells, dendritic cells, or macrophages and signaled the induction of interleukin 2 and proliferation of the T-cell (Linsley and Ledbetter, 1993).

Avian T-cells possessed homologs to mammalian CD4 (64-kDa monomeric polypeptide) helper T-cells, and CD8 (a dimer, 31 and 34 kDa) cytotoxic T-cells (Chen et al., 1991). Veillette and Ratcliffe (1991) revealed that like the mammalian systems, the chicken homologies of CD4 and CD8 associated with a 56-kDa tyrosine-specific protein kinase. The thymic hormone, avian thymulin, influenced CD4/CD8 ratios and the expression of CD4 and CD8 based on fluorescent staining (Marsh, 1993). Double-positive CD4 CD8 cells, single-positive CD8⫹ cells, and single-positive CD4⫹ cells appeared in the thymic cortex at 9 or 10 DE, 13 DE,and 15 DE, respectively (Bucy et al., 1990). The CD8⫹ cells appeared earlier in the medulla (15 DE) than did the CD4⫹ cells (17 DE). 4. Origin of Bursal and Thymic Lymphocytes Jaffe and Fechheimer (1966) and Moore and Owen (1965, 1966) utilized sex chromosome techniques and suggested the possibility that immigrant cells were the progenitors of bursal lymphocytes. LeDouarin et al. (1984) took advantage of the one or two large clumps of heterochromatin associated with the cell nucleus of quail and their absence in chick cells to incisively reveal the blood-borne origin of bursal and thymic lymphocytes. Experiments revealed quail basophilic stem cell migration between days 7 and 11 of embryogenesis while the chick basophilic cell migrated into the bursa between 8 and 15 days DE (LeDouarin et al., 1984). Unlike the bursa, the thymic precursor lymphocytes entered the thymus in three waves (Le Douarin et al., 1984, 1990) between 6.5 and 8, 12 and 14, and 18 and 20 DE with a refractory period of 4 days between the first and second and second and third waves (Le Douarin et al. 1984, 1990; Figure 2). With the acceptance that the bursal and thymic lymphocytes originated from a blood-borne stem cell, the origin of the blood-borne stem came into question. Chimeras were developed prior to circulation (⬍14 somite stage) by replacing a chick area pellucida (embryo proper) with that of a quail (Martin, 1990). The developing lymphocyte nuclear characteristics in this embryo chimera resembled those of a quail and were, therefore, of intraembryonic origin. The origin of the intraembryonic stem cells may be intraaortic, paraaortic, or from the coelomic epithelium (Dieterlen-Lievre et al., 1990; Olah et al., 1988).

B. Secondary Lymphoid Tissue 1. Spleen Adjacent to the dorsal surface of the right lobe of the liver and dorsal to the proventriculus is the reddishbrown oval spleen (Nickel et al., 1977). Accessory

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ellipsoid-associated cell (EAC) which bound diverse substances that entered the CS through stomata formed by the endothelial cells of the midregion of penicilliform capillaries (White et al., 1970; Olah and Glick, 1982; Figure 4). The EAC is activated following binding and migrates into the PWP, red pulp, PALS, and GC regions. The EAC appeared to be a messenger cell and may be lineage related to interdigitating dendritic cells of the PALS and follicular dendritic cells of the GC (Olah and Glick, 1982; Gallego et al., 1993). The mammalian spleen possessed a marginal zone (macrophages and lymphocytes) which surrounded the PALS separating this region from the red pulp (Weiss, 1972). An avian marginal zone has been suggested by Jeurissen et al. (1992) and Jeurissen (1993) to include the CS and the surrounding EAC, B-cells, and macrophages. 2. Cecal Tonsil

FIGURE 2 Stem cell (SC) migration to the chick bursa occurs between 7.5 and 14 days of embryogenesis (DE) and in the thymus occurs in three waves at 6.5, 12, and 18 DE. A B-cell repertoire able to recognize 106 different antigens forms in the microenvironment of the bursa while the thymus environment signals the formation of clonally specific T-cell receptors (a) and T-helper (CD4) (b) or Tcytotoxic (CD8) (c) cells. The differentiated B- and T-cells then migrate to the secondary lymphoid tissue.

spleens have been described cranial, adjacent, and caudal to the spleen. Subsequent to splenectomy, the cranial accessory spleen hypertrophied (Glick, 1986). The most rapid rate of splenic growth occurred during the first 6 weeks after hatching with maximum size (spleen-tobody weight ratio) attained by 10 weeks of age (Glick, 1986). The avian spleen like the mammalian spleen possesses red and white pulp. Within the white pulp are located (1) the periarteriolar lymphatic sheath (PALS); (2) germinal centers; and (3) periellipsoid white pulp region (PWP). A central artery arising from the splenic artery was surrounded by the PALS, which contained lymphocytes, macrophages, and dendritic cells and was thymic dependent (Glick, 1986). Germinal centers, bursal-dependent regions, were located at the edge of the PALS. The central artery as it entered the PWP became the penicilliform capillary (PC). The midregion of the penicilliform capillary was surrounded by the capillary sleeve (CS) or ellipsoid (Olah and Glick, 1982; Figure 3). The CS was embroidered by the dendritic

The cecal tonsil is an enlarged patch of tissue (4– 18 mm) in the proximal region of each cecum (Muthmann, 1913; Glick, 1986). The cecal tonsil villi were longer and less broad than those from the remainder of the cecum’s proximal region (Glick, 1986). The polycryptic cecal tissue was similar to the mammalian palatine tonsil (Glick et al., 1981). The location and continuous exposure of the tonsil villi to the fecal content suggested a sentinel role for this peripheral lymphoid tissue. The cecal tonsil possessed approximately 400 spherical units, each with a central crypt, diffuse lymphoid tissue, and germinal centers (Glick et al., 1981). The cecal tonsil possessed T- and B-cells (Albini and Wick, 1974) and IgM, IgG, and IgA plasma cells ( Jeurissen et al., 1989b) and produced antibody to soluble antigens ( Jankovic and Mitrovic, 1967; Orlans and Rose, 1970). 3. Peyer’s Patches Peyer’s patches appeared in 10-day-old chickens along the intestine cranial to the ileocecal junction (Schat and Myers, 1991). They possessed lymphocytes beneath the epithelium but were not polycryptic like the cecal tonsil. There were approximately five or six peyer’s patches, 5 mm in diameter, in the intestine of 12-week-old chickens (Schat and Myers, 1991). The majority of T-cells were TCR-1 (움/웁) and CD4 (T-helper). In general, plasma cells produced each of the three Ig isotypes ( Jeurissen et al., 1989a; Schat and Meyers, 1991). Hormonal bursectomy depopulated the lymphocytes in the subepithelial zone (B-dependent) and central zone. 4. Meckel’s Diverticulum The yolk stalk, Meckel’s Diverticulum (MD), in 2week-old chickens, is 3–6 mm long and 1.7 mm thick (Olah and Glick, 1984a). Its distal end continued as the

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FIGURE 3 Chicken spleen: BC, billroth cord; E, ellipsoid or capillary sleeve; CA, central artery; CV, central vein; GC, germinal center; PALS, periarteriolar lymphatic sheath; PC, penicilliform capillary; PWP, periellipsoid white pulp; S, sinus; TA, trabecular artery; TV, trabecular vein. (From Olah and Glick, 1982, Am. J. Anat. 165, 445–480. Copyright 䉷 1982 Wiley-Liss, Inc. Reprinted by permission of Wiley-Liss, Inc., a subsidiary of John Wiley & Sons, Inc.)

yolk sac (Figure 5). The MD may contribute to the circulating pool of white blood cells and may be a site to isolate colony stimulating factors that lead to monocytic or granulocytic colonies. Olah et al. (1984) identified lymphoid accumulation in MD (yolk stalk) at 2 weeks of age and confirmed Calhoun’s (1933) observation of its absence at 1 day old. Jeurissen et al. (1989a, 1989b) identified leukocytes in MD of late embryos, IgM-positive cells underneath the epithelium at 5 days posthatch, and IgG- and IgApositive cells between 2 and 6 weeks of age. Olah et al. (1984) identified dendritic cells (possibly secretory cells) and suggested that they may initiate germinal center formation and may be follicular dendritic cells. Jeurissen et al. (1989b) also reported the presence of dendriticlike cells, possibly follicular dendritic cells, in germinal centers. FIGURE 4 Capillary sleeve (ellipsoid) embroidered by ellipsoid as-

5. Intestinal Lymphocytes

sociated cell (EAC); PC, penicilliform capillary. (From Olah and Glick, 1982, Am. J. Anat. 165, 445–480. Copyright 䉷 1982 Wiley-Liss, Inc. Reprinted by permission of Wiley-Liss, Inc., a subsidiary of John Wiley & Sons, Inc.)

Intestinal lymphocytes generally resided in the epithelium or lamina propria (Schat and Meyers, 1991). The lamina propria contained IgM- and IgA-positive

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cytes appeared in the pineal and attained maximum concentration by 32 days. Three to 5 days following a carotid injection of bovine serum albumin, pineal plasma cells revealed the presence of antibody to bovine serum albumin (Cogburn and Glick, 1983). Immunohistochemistry revealed an IgA-like substance on the luminal surface of the pineal follicles and in the perifollicular layer (Olah and Glick, 1991). Neonatal bursectomy/ thymectomy or at-hatch administration of cyclophosphamide significantly reduced or eliminated T- and Bcells and germinal centers within the pineal (Cogburn and Glick, 1981). The pineal may contribute immunocompetent cells and their products for immune surveillance of the central nervous system. 8. Harderian Gland FIGURE 5 Schematic of Meckel’s Diverticulum.

B-cells and plasma cells and CD4⫹ T-cells. These intraepithelial cells located between the epithelial cells and basement membrane. They appeared to immigrate from the lamina propria. The intraepithelial cells may be of thymic origin since they declined following thymectomy but not bursectomy (Schat and Myers, 1991). A functional separation of the intestinal lymphocytes occurred between the lamina propria, B- and CD4⫹cells, and epithelium CD8⫹cells (Schat and Myers, 1991). TCR2 predominated in the lamina propria and TCR1 in the epithelium. The presence of an antigen, similar to the expression of 웁7 integrin of human and mice, on avian T-cells after their arrival in the intestine suggested that this antigen retained T-cells in the intestinal epithelium (Haury et al., 1993). 6. Lymph Node The most developed lymphoid accumulations along the posterior tibiopopliteal and lower femoral veins were true lymph nodes possessing afferent and efferent lymphatics, T- and B-cells, germinal centers, and a prominent lymphatic sinus system (Olah and Glick, 1983, 1985). Kampmeier (1969) reported similar nodes, the cervicothoracic node in waterfowl. A footpad injection of sheep red blood cells stimulated enlargement of the nodes and generated more plaque forming cells (20 ⫻ 103 PFC/106 lymphoid cells) than in the spleen (4 x 103 PFC/106 lymphoid cells) (McCorkle et al., 1979). 7. Pineal The pineal gland is a lymphopoietic tissue in the chicken (Romieu and Jullien, 1942; Cogburn and Glick, 1981; Olah and Glick, 1984b). By 9 days of age lympho-

The Harderian gland (HG), located ventral and posteromedial to the eyeball (Wight et al., 1971), performs a sentinel role in immune protection of the chicken (Glick and Olah, 1981). The identification of plasma cells in the HG by Bang and Bang (1968) was followed by the observation of Mueller et al. (1971) that the HG was capable of producing a specific antibody. The HG has been suggested to influence B-cell activation, proliferation, and differentiation (Gallego and Glick, 1988; Mansikka et al., 1989; Olah et al., 1992a; Savage et al., 1992; Scott et al., 1993; Scott and Savage, 1996; Tsuji et al., 1993). Later papers identified the response of the HG to a variety of pathogens (Darbyshire, 1987). Plasma cells within the HG (1) concentrated by 3 to 4 weeks posthatch (Niedford and Wolters, 1978; Gallego and Glick, 1988); (2) experienced a proliferation rate 2 to 3 times higher than in the spleen (Gallego and Glick, 1988); (3) peaked in the rate of S-phase by 6 or 8 weeks of age and declined thereafter (Savage et al., 1992); and (4) markedly declined in numbers following bursectomy (Mueller et al., 1971; Sundick et al., 1973). Immunoglobulin class switch from IgM to IgA or IgG occurred in HG B-cells (Mansikka et al., 1989). Immunohistochemical techniques revealed that IgA produced by plasma cells in the central canal and primary branches was secreted by the epithelium of secondary branches and entered the lumen of the primary branch and central canal (Olah et al., 1992a).

III. REGULATION OF IMMUNE RESPONSE A. Major Histocompatibility Complex The major histocompatibility complex (MHC) is a highly polymorphic cluster of genes that produce membrane-associated products that influence T-cell de-

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velopment (possibly B-cell development), recognition by T-cells of antigen (MHC restriction or allospecificity), graft rejection, disease resistance, and production traits. 1. Membrane-Associated Glycoproteins, Class I and Class II The MHC of the chicken originally described by Briles led to the identification of three interesting MHC loci in birds, B-F (class I), B-L (class II), and B-G (class IV) (Crone and Simonsen, 1987). The B-F antigens, expressed on most chicken cells, and the B-L antigens present on B-cells, macrophage/dendritic cells, and activated T-cells are analogous to class I and class II mammalian antigens, respectively. The B-F antigens possessed a membrane-bound glycosylated polymorphic heavy 움-chain (40 to 45 kDa) noncovalently linked with the invariant 웁2-microglobulin (12 kDa) (Crone and Simonsen, 1987; Chen et al., 1991). The 움-chain possesses three domains, the most distal pair, 움1 and 움2, to form a cleft which presents the processed antigen to T-cytotoxic cells (CD8). The B-L antigen possesses a nonpolymorphic 움-chain (32 kDa) and a polymorphic beta chain (27 kDa) (Pharr et al., 1993). The peptide cleft of MHC class II molecules is formed by 웁1 and 움1, the most distal domains from the membrane. These peptide-associated class II molecules are presented to T-helper (CD4) cells (Figure 6). 2. MHC Restriction, Allospecific Response, and T-Cytotoxic Cells Successful presentation of the peptide to a T-cell requires a similar MHC between the antigen-presenting cell and T-cell (MHC restriction). T-cytotoxic cells (CD8) have been identified in chickens (Glick, 1977). Numerous observations suggested that the cytotoxic response is, in part, dependent on T-cells since B-cells were absent in treated bird. A syngeneic-restricted Tcytotoxic lymphocyte response, MHC restriction, has been demonstrated in vitro with the Schmidt-Ruppin strain of avian sarcoma virus or Reticuloendotheliosis virus-infected cells (Schat and Myers, 1991). Another example of MHC restriction is delayed type hypersensitivity. Several weeks after receiving a specific antigen, a subcutaneous injection of the same antigen will induce a T-cell mediated (CD4) inflammation response within 24 to 72 hr (Glick, 1986). T-cytotoxic cells will react with peptides presented by different (allogeneic) MHC molecules to produce an allograft or graft-vs-host response (Glick, 1986). An allograft is a skin transplant to an animal of the same species but with different MHC and demonstrates the

FIGURE 6 A model depicting events leading to activation of T- and B-cells. The recognition of a complementary epitope by the antigen receptor of the B-cell (g) transmits an initial signal (1) to the B-cell which then internalizes and processes the antigen for presentation (e) to T-cell receptors (a) of a clonally specific T-helper (CD4) cell. Coreceptor signals (e–b and d–c) help to activate the T-cell to release a cytokine (f) (IL-4) that binds to the B-cell supplying a second signal (2) which activates the B-cell to proliferate and differentiate into an antibody-secreting cell.

rejection of the skin by the host’s T-cells. Like the allograft response, the graft-vs-host response is governed by differences at the BF and/or B-L loci. A synergistic effect on the graft-vs-host response occurred when bone marrow and thymic cells were combined (Glick, 1986). 3. Signaling and Coreceptors MHC restriction depends on the TCR complex (see Section II,A,3) recognizing a peptide MHC complex on the surface of an antigen-presenting cell (monocyte/ macrophage B-cell or dendritic cell). The CD8 or CD4 molecules associate with the TCR/CD3 complex and are considered coreceptors (Figure 6). In mammals, optimal T-cell activation depends on direct or indirect association of TCR/CD3 complex with the above coreceptors and a membrane-associated CD45 molecule, leukocyte common antigen. Signals received by the Tcell will aggregate the TCR/CD3 complex together with either coreceptor CD4 or CD8, activating the tyrosine protein kinase associated with coreceptors. CD45 (about 200 kDa) contributes to the activation of tyrosine protein kinase by way of its tyrosine-specific phosphatase ( Janeway and Travers, 1992). The chicken model has revealed similar signaling and coreceptor activation. Avian CD4 and CD8, the only known nonmammalian

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CD4 and CD8 homologs, have been shown to physically associate with a tyrosine phosphatase kinase homologous to mammalian p56lck (Veillette and Ratcliffe, 1991). Taken together, the data of Veillette and Ratcliffe (1991) and Paramithiotis et al. (1991) suggest that signaling by way of the avian TCR/CD3 complex is modulated by the activation of the tyrosine protein kinase-p56lck originating in the coreceptor molecules CD4 and CD8 and the tyrosine-specific phosphatase released by the CD45 surface molecule.

B. Cytokines A variety of protein mediators or cytokines are produced by T-lymphocytes (Howard et al., 1993). Of these cytokines only interferon (IFN), interleukin (Il)-I, Il-2, Il-3, and tumor necrosis factor (TNF-움) have been studied in birds (Lillehoj et al., 1992; Klasing, 1994). In addition, a stem cell factor (SCF) and myelomonocytic growth factor (MGF) (Leutz et al., 1989) have been studied. The MGF, originated from a transformed macrophage cell line, HDII, has been cloned by Leutz and shown to have some homology to mammalian granulocyte-colony stimulating factor (G-CSF). The recent work of Nicholas-Bolnet et al. (1991, correspondence) offers important steps in isolating SCF, Il-3, GCSF, and M-CSF. Interleukin-1 is produced by monocytes and stimulated thymocytes to release Il-2 (Schat and Myers, 1991). Limited or no functional cross-reactivity has been reported with chicken, human, and murine Il-1 or Il-2 (Schat and Myers, 1991). The conditioned medium of a concanavalin-Astimulated culture of adherent splenocytes and thymocytes stimulated proliferation of preactivated Tlymphoblasts and Il-2 activity (Schat and Myers, 1991). The molecular weight of between 15.5 and 30 kDa for avian Il-2 is similar to that of mammals. Like the interleukin studies, IFN studies in chickens are in the very early stages of development (Schat and Myers, 1991; Lillehoj et al., 1992). This is surprising since the first inference of IFN was from the chicken embryo data of Isaacs and Lindemann (1957). Transforming growth factor 웁 which influences cell differentiation, wound repair and bone metabolism and growth and tumor necrotic factor-움 have been identified in the chicken (Klasing, 1994). A lymphocyte inhibitory factor (LyIF) was released from thymic or bursal cells sensitized to purified protein derivative (Glick, 1983). Sensitized thymic cells required macrophages for the release of LyIF and a chemotactic factor while the LyIF response of bursal cells was independent of macrophages ( Joshi and Glick, 1990). These types of experiments illustrate an advanced stage of

maturity of cells within the thymic and bursal microenvironments.

C. Antibody-Mediated Immunity and B-Cell Repertoire Antibody production by B-cells prevents the spread of pathogens by (1) combining with the pathogen and neutralizing it, (2) facilitating uptake and digestion of the pathogen by phagocytic cells, and (3) facilitating cell lysis and death. Most antibody responses in birds are thymic dependent (TD) and require cooperation between T-, B-, and macrophage/dendritic cells (Weinbaum et al., 1973; Thorbecke et al., 1980). Thymicindependent (TI) antigens include pneumococcal polysaccharide and Brucella, examples of TI-type I, and ficoll and dextran, examples of a TI-type II (Golub and Green, 1991). A TI response has been demonstrated in birds by allogeneic bursal cell transfers to cyclophosphamide recipients (Toivanen et al., 1974). Antibody response in these birds to Brucella abortus but not the TD antigen sheep red blood cells was restored. The allogeneic bursal stem cells induced B-cell chimerism and tolerance to donor MHC (Vainio and Toivanen, 1987). These types of allogeneic responses will reconstitute TI but not TD responses. The TD responses of cyclophosphamide birds required transfer of thymic and bursal cells syngeneic to host or possibly one MHC similar to donor B-cell and host T-cell (Vanio and Toivanen, 1987). 1. Immunoglobulins B-cells and plasma cells synthesize and release immunoglobulin. Antibody, a glycoprotein, is structurally identical to Ig and is represented by five distinct classes or isotypes: IgM, IgD, IgG, IgA, and IgE. Each class exhibits at least one monomer of two-differentmolecular-weight polypeptide chains: two heavy (H) and two light (L) chains. Beginning at the N-terminal of the H polypeptide class and extending to its C-terminal there is a variable (VH) domain and three to four constant domains (CH) depending on the isotype. The light chain possesses a VL and a single CL domain. The constant regions differentiate the distinctiveness of each heavy chain which are named 애 (IgM), ␦ (IgD), 웂 (IgG), 움 (IgA), and ␧ (IgE). In the chicken there is a single light or ␬-chain. The VH and VL (N terminal) domains contribute to the antigen binding site or Fab while the CH domains (C-terminal) identify the constant or Fc portion of the antibody molecule (Figure 7). Avian IgM resembled mammalian IgM [(애2L2)5] on the basis of physiochemical, antigenic characteristics; sedimentation coefficients (S, 16.7 to 16.9), and molecu-

Chapter 26. Immunophysiology

FIGURE 7 The immunoglobulin molecules possess two heavy (H) chains with a single variable domain (VH) and three or four constant domains (CH to CH4). Each light (L) chain possess a VL- and CLdomain. The Fab refers to the antigen binding position of the molecule (N-terminus) while Fc identifies the constant portion of the molecule (C-terminus).

lar weight (890 kDa) (Leslie and Clem, 1969). Sequence data for chicken 애 identified a homology with mammalian 애 chain (Dahan et al., 1983); there were five IgM CH allotypes (Benedict and Berestecky, 1987). On the other hand, disagreement exists concerning the name of the major avian 7.1- to 7.8-serum Ig IgG (웂2L2). Leslie and Clem (1969) named this 7SIg IgY, in part, because it possessed 4CH and not 3CH as in human IgG and possessed a greater concentration of carbohydrate (6.0%) than did the human IgG. It has been suggested that avian serum 7SIg resembled mammalian IgA more than IgG (Tenenhouse and Deutch, 1966) and that human IgA cross-reacted with the avian serum 7SIg (Ambrosius and Hadge, 1987). The inference that the avian 7SIg could be a precursor of IgA was weakened with the identification of chicken secretory IgA (Benedict and Berestecky, 1987). The amino acid sequence of the IgY H chain placed it closest to mammalian IgE (Parvari et al., 1988). Functionally, the avian 7SIg (IgG or IgY) was similar to mammalian IgG (Benedict and Berestecky, 1987). Interestingly, the duck possessed two serum Igs, a 5.7-SIgG and a 7.8-SIgG, which may be influenced by a single gene (Magor et al., 1992; Higgins and Warr, 1993). A third avian Ig was termed IgA because of its prevalence in lymphoepithelial tissue and structural similarity

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to mammalian IgA (Benedict and Berestecky, 1987). Cloning of the IgA H chain revealed the greatest structural similarity to mammalian 움H chains and not 애H or ␧H (IgE) chains (Mansikka, 1992). The heavy chain of chicken IgA contained four constant 움 domains or one more than human constant 움 chains. These results suggested the possible occurrence of a deletion during evolution. Avian IgA was polymorphic and highly concentrated in the bile (3–12 mg/ml) (Benedict and Berestecky, 1987). A J-component and secretory component were present in avian IgA. The secretory component (60 kDa) was synthesized by epithelial cells and attached to bile IgA before secretion (Benedict and Berestecky, 1987). While chicken and turkey IgA may be monomeric or polymeric, the secretory IgAs were generally trimeric or tetrameric. Polymeric IgA, which lacked the secretory component, may be transported from blood to bile by combining with membrane secretory components produced by hepatocytes (Schat and Myers, 1991). Immunoglobulin G and IgM antibodies possess complement binding sites. Complement is a complex of 9 major components ( Janeway and Travers, 1994). Avian complement 3 (C3) fragments to C3b after binding to an antibody–antigen complex (classical pathway) or in the presence of a pathogenic surface (alternate pathway, AP) and continues the cascade of complement factors that will remove the immune complexes, lyse the pathogens bound by the antibody, and enhance inflammation (Koch, 1987). It appeared that the B-like protein important in the AP existed in the chicken and was similar to mammalian C2 and component B (Kjalke et al., 1994). 2. V-Gene Repertoire and the Contributions of the Bursal Milieu Antibody diversity in mammals depends on somaticdriven events within the B-cell leading to (1) heavychain rearrangement between D and J segments; (2) rearrangement of several hundred V-genes with the rearranged D-J segments (VDJ); (3) light-chain rearrangement of numerous V and J segments; (4) junctional diversity; (5) nucleotide additions; and (6) combinatorial joining between the rearranged H (VDJ)- and L (VJ)-chains (Golub and Green, 1991). Since the chick H chain possesses a single V and 10 D (similar homologies) and a single J gene and the light chain possesses single V and J genes, rearrangement of the H- and L-chains will not contribute diversity as in mammals. Rather, the chicken depends on gene conversion, a transfer of nucleotide sequences from upstream pseudogenes to the rearranged VH (VDJ) and VL (VJ) genes, to produce antibody diversity (McCormack et al., 1991). Including the values of 104 follicles per bursa, 105 cells per follicle and other assumptions,

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mathematical models have been presented to support a B-cell repertoire (antibody diversity) of about 106 different specificities (Salanti et al., 1989; Langman and Cohn, 1993). The rearrangement of H & L chains may occur in the bursa or at other sites (McCormack et al., 1991). These molecular observations supported the original observation of a bursal-independent site for Ig (Lerner et al., 1971; Glick, 1977; Ratcliffe et al., 1986; McCormack et al., 1991). Schemes to explain the bursal-dependent roles of the expansion of in-frame B-cells (i.e., B-cells with rearranged H- and L-chains) and gene conversion (McCormack et al., 1991) have been discussed and conceptually approached by Langman and Cohn (1993). The microenvironmental milieu necessary for these events appears to include (1) the complex interaction of endodermal and mesodermal germ layers (Houssaint et al., 1976), (2) a singular role of the endodermal epithelium (LeDouarin et al., 1980), and (3) a dark mesenchymal cell that is the precursor of the bursal secretory dendritic cell (BSDC) (Olah et al., 1986; Olah and Glick, 1978b; Olah et al., 1992b). During late embryonic development the only cell to possess IgG on its membrane was the BSDC (Olah et al., 1991). We have proposed a receptor paracrine pathway that involved the interaction of the IgM of the in-frame B-cell with the IgG of the BSDC (Glick and Olah 1993a,b; Glick, 1995). This may lead to activation and proliferation of the B-cells and a BSDC secretion that signals gene conversion of the expanded in-frame B-cells. Alternatively, replication of in-frame B-cells might occur when the B-cell identified a bursal specific antigen (Masteller and Thompson, 1994), possibly expressed by the BSDC. In either case, apoptosis (programmed cell death) (Compton, 1993) of the inframe B-cell would occur if the in-frame B-cell fails to make contact with a specialized cell. The receptor–paracrine hypothesis has an analogy with the activity in mammalian germinal centers where follicular dendritic cells trap antigen–antibody complexes, present them to the B-cell, and then release a secretion that contributes to B-cell differentiation (Gordon et al., 1989). Kaufman and Salomonsen (1993), subsequent to several assumptions, proposed the B–G complex of the MHC as the antigen influencing the selection of germline Ig bursal cells. In their negative-selection model, the germline Ig B-cells bound to the B–G self molecule and were eliminated while the B-cells that experienced gene conversion would not bind and would emigrate. In the positive-selection model, the germline B-cells bind to the B–G self molecule, receive a signal to proliferate, and undergo gene conversion, eventually losing their recognition of B–G self and then emigrating (Kaufman et al., 1991). The influence of cell adhesion molecules on the cells of the bursal microenvironment

has been briefly discussed (Glick, 1995) and experimental evidence offered by Masteller and Thompson (1994). Bu-1-positive cells in the embryonic spleen are committed to the B-cell lineage and express a cell adhesion ligand Sialyl Lewis x (Masteller and Thompson, 1994). By 10 to 12 days of embryonic development, Sialyl Lewis x-positive cells appear in the bursa, proliferate, and then contribute to the developing follicle. These B-cells lose Sialyl Lewis x in parallel with the initiation of gene conversion (15 to 18 days of embryogenesis). The loss of Sialyl Lewis x and the switch to Lewis x high in Igpositive B-cells is followed by emigration to the spleen. This might suggest that migration from the bursa is influenced by a change in expression of Lewis x.

D. Macrophages, Natural Killer Cells, Heterophils, and Thrombocytes The phagocytic avian macrophage, similar to the mammalian macrophage, performs the pivotal role of an antigen-presenting cell (Powell, 1987; Vainio et al., 1988). Several investigators have studied the avian macrophages’ effector functions (Dietert et al., 1991; Qureshi et al., 1994) by employing peritoneal exudate macrophages and a malignantly transformed chicken macrophage cell line, MQ-NCSU. Optimum activation of avian macrophages required two signals, a lymphokine (IFN(?)) and lipopolysaccharide (LPS) (Qureshi et al., 1994). Activated macrophages will release Il-1, tumor necrotic factor (TNF), and colony stimulating factors (CSF). The cytotoxicity of TNF may be species specific (Qureshi et al., 1993). Chicken embryonic bone marrow cells exposed to supernatants from cultured MQ-NCSU macrophage cells produce colonies of granulocytes and macrophage-granulomonocytic cells, suggesting the ability of these macrophages to produce GCSF and GM-CSF, respectively. Macrophage cell lines pulsed with LPS produce reactive nitrogen intermediates (e.g., nitric oxide, ⭈NO) (Qureshi et al., 1993, 1994). Unlike mammalian macrophages that synthesize arginine, avian macrophages require exogenous sources of l-arginine, which are converted to reactive nitrogen intermediates (e.g., ⭈NO) by way of oxidative enzymes (Qureshi et al., 1993). These reactive nitrogen intermediates have an antineoplastic and antimicrobial function. Nitric oxide may have an autocrine effect since it suppresses complexes I and II enzymes in the mitochondria of macrophages and tumor target cells (Sung and Dietert, 1994). Further challenges in understanding avian macrophage will be to identify a homolog to the mammalian differentiation antigen CD14 and the LPS–LBP (lipopolysaccharide binding protein) complex that binds to the membrane-associated CD14 to trigger the release of macrophage cytokines (Martin et al., 1994). Also, will

Chapter 26. Immunophysiology

⭈NO inhibit the expression of avian macrophage Ia (class II MHC), which in mammals results in reduced ⭈NO production (Sicher et al., 1994)? Natural killer (NK) cells are large granular leukocytes exhibiting natural cytotoxicity to a variety of tumor cells. The NK cells found in peripheral blood, intestinal epithelium, and spleen tend to be nonadherent cells. Cells exhibiting spontaneous cytotoxicity, NK cells, were identified by two mAb, K-108 and K-4 (Chung and Lillehoj, 1991). NK cells increase with age and in the presence of viruses. Disease and suppressor cells depress NK cells. Mammalian NK cells respond positively to interferon while avian NK cells may not (Sharma and Schat, 1991). The nonlymphoid heterophil and thrombocyte were active in immunity and infection (Powell, 1987). Heterophils are the early cells of inflammation and are phagocytic. Serotonin inhibited the cytotoxicity of granulocytes while pretreatment with interferon impeded the inhibitory effects of serotonin (Garssadi et al., 1994). The thrombocytes role in hemostasis and phagocytosis was shown initially by Stalsberg and Prydz (1963) and Glick et al.(1964), respectively. Monoclonal antibodies raised against platelets will identify thrombocytes (Kunicki and Newman, 1985) while Kaspers et al. (1993) have described a mAb that appears to identify monocyte/thrombocytes.

References Adelmann, H. B. (1942). ‘‘The Embryological Treatises of Hieronymus Fabricius of Aquapendente,’’ Vol. 1, p. 376, Cornell Univ. Press, Ithaca, NY. Albini, B., and Wick, G. (1974). Delineation of B and T–lymphoid cells in the chicken. J. Immunol. 112, 444–450. Ambrosius, H., and Hadge, D. (1987). Chicken immunoglobulins. Vet. Immunol. Immunopathol. 17, 57–67. Bang, B. G., and Bang, F. B. (1968). Localized lymphoid tissues and plasma cells in paraocular and paranasal organ systems in chickens. Am. J. Pathol. 33, 735–751. Benedict, A. A., and Berestecky, J. M. (1987). Special features of avian immunoglobulins. In ‘‘Avian Immunology: Basis and Practice’’ (A. Toivanen and P. Toivanen, eds.), Vol. 1, pp. 113–126. CRC Press, Boca Raton, FL. Bockman, D. E., and M. D. Cooper (1973). Pinocytosis by epithelium associated with lymphoid follicles in the bursa of Fabricius, appendix, and Peyer’s patches. An electron microscopic study. Am. J. Anat. 136, 455–478. Brewer, J. M., Wunderlich, J. K., and Ragland, W. L. (1990). The amino acid sequence of avian thymic hormone, a parvalbumin. Biochimie 72, 653–660. Bucy, R. P., Chen, Chen-Lo H., and Cooper, M. D. (1990). Ontogeny of T–cell receptors in the chicken thymus. J. Immunol. 144, 1161– 1608. Calhoun, M. L. (1933). The microscopic anatomy of the digestive tract of Gallus domesticus. Iowa St. Coll. J. Sci. 7, 261–381. Chen, Chen-Lo H., Ager, L. L., Gartland, E. L., and Cooper, M. D. (1986). Identification of a T3/T cell receptor complex in chickens. J. Exp. Med. 164, 375–380.

667

Chen, Chen-Lo H., and Cooper, M. D. (1987). Identification of cell surface molecules on chicken lymphocytes with monoclonal antibodies. In ‘‘Avian Immunology: Basis and Practice’’ (A. Toivanen and P. Toivanen, eds.), Vol. 1, pp. 138–154. CRC Press, Boca Raton, FL. Chen, Chen-Lo H., Pickel, J. M., Lahti, J. M., and Cooper, M. D. (1991). Surface markers on avian immune cells: In ‘‘Avian Cellular Immunology’’ ( J. M. Sharma, ed.), pp. 1–22. CRC Press, Boca Raton, FL. Chung, K. S., and Lillehoj, H. S. (1991). Developmental and functional characterization of monoclonal antibodies recognizing chicken lymphocytes with natural killer cell activity. Vet. Immunol. Immunopathol. 28, 351–363. Cogburn, L. A., and Glick, B. (1981). Lymphopoiesis in the chicken pineal gland. Am. J. Anat. 102, 131–142. Cogburn, L. A., and Glick, B. (1983). Functional lymphocytes in the chicken pineal gland. J. Immunol. 130, 2109–2112. Compton, M. M. (1993). Programmed cell death in avian thymocytes: Role of the apoptotic endonuclease. Poult. Sci. 72, 1267–1272. Cooper, M. D., Peterson, R. D. A., South, M. A., and Good, R. A. (1966). The functions of the thymus system and bursa system in the chicken. J. Exp. Med. 123, 75–106. Crone, M., and Simonsen, M. (1987). Avian major histocompatibility complex. In ‘‘Avian Immunology: Basis and Practice’’ (A. Toivanen and P. Toivanen, eds.), Vol. II, pp. 25–42. CRC Press, Boca Raton, FL. Dahan, A., Raynaud, C.-A., and Weill, J.-C. (1983). Nucleotide sequence of the constant region of a chicken 애 heavy chain immunoglobulin mRNA. Nucleic Acids Res. 11, 5381–5389. Darbyshire, J. H. (1987). Immunity and resistance in respiratory tract diseases. In ‘‘Avian Immunology: Basis and Practice’’ (A. Toivanen and P. Toivanen, eds.), pp. 129–141. CRC Press, Boca Raton, FL. Dieterlen-Lievre, F., Pardanaud, L., Bolnet, C., and Cormier, F. (1990). Development of the hemopoietic and vascular systems studied in the avian embryo. In ‘‘The Avian Model in Developmental Biology: From Organism to Genes’’ (N. LeDouarin, F. Dieterlen–Lievre, and J. Smith, eds.), pp. 181–196. Edition Du Centre National De La Recherche Scientifique, Paris, France. Dietert, R. R., Golemboski, K. A., Bloom, S. E., and Qureshi, M. A. (1991). The avian macrophage in cellular immunity. In ‘‘Avian Cellular Immunology’’ (J. A. Sharma, ed.), pp. 71–95. CRC Press, Boca Raton, FL. Gallego, M., and Glick, B. (1988). The proliferative capacity of the avian Harderian gland. Dev. Comp. Immunol. 12, 157–166. Gallego, M., Olah, I., Del Cacho, E., and Glick, B. (1993). Anti S-100 antibody recognizes ellipsoid–associated cells and other dendritic cells in the chicken spleen. Dev. Comp. Immunol. 17, 77–83. Garssadi, S. E., Regely, K., Mandi, Y., and Beladi, I. (1994). Inhibition of cytotoxicity of chicken granulocytes by serotonin and ketamine. Vet. Immunol. Immunopathol. 41, 101–112. Gilmour, D. G., Brand, A., Donnelly, N., and Stone, H. A. (1976). Bu-1 and Th-1, two loci determining surface antigens of B and T lymphocytes in chickens. Immunogenetics 3, 549–563. Glick, B. (1956). Normal growth of the bursa of Fabricius in chickens. Poult. Sci. 35, 843–851. Glick, B. (1977). The bursa of Fabricius and immunoglobulin synthesis. Int. Rev. Cytol. 48, 345–402. Glick, B. (1983). Bursa of Fabricius. In ‘‘Avian Biology’’ (D. S. Farner, R. King, and K. C. Parkes, eds.), Vol. 7, pp. 443–500. Academic Press, New York. Glick, B. (1986). Immunophysiology. In ‘‘Avian Physiology’’ (P. D. Sturkie, ed.), pp. 87–101. Springer-Verlag, New York.

668

B. Glick

Glick, B. (1987). How it all began: A continuing story of the bursa of Fabricius. In ‘‘Avian Immunology: Basis and Practice’’ (A. Toivanen and P. Toivanen, eds.), pp. 1–8. CRC Press, Boca Raton, FL. Glick, B. (1995). Embryogenesis in the bursa of Fabricius: Stem cell, microenvironment and receptor–paracrine pathways. Poult. Sci. 74, 419–426. Glick, B., and Olah, I. (1981). Gut–associated lymphoid tissue of the chicken. In ‘‘Scanning Electron Microscopy III’’ (O. Jahari, ed.), pp. 95–108. SEM, Inc., Chicago, IL. Glick, B., and Olah, I. (1993a). Bursal secretory dendritic-like cell: A microenvironment issue. Poult. Sci. 72, 1262–1266. Glick, B., and Olah, I. (1993b). A bursal secretory dendritic cell and its contributions to the microenvironment of the developing bursal follicle. Res. Immunol. 144, 446–448. Glick, B., Chang, T. S., and Jaap, R. G. (1956). The bursa of Fabricius and antibody production in the domestic fowl. Poult. Sci. 35, 224–225. Glick, B., Sato, K., and Cohenour, F. (1964). Comparison of the phagocytic ability of normal and bursectomized birds. J. Reticuloendothel. Soc. 1, 442–449. Glick, B., Holbrook, K. A., Olah, I., Perkins, W. D., and Stinson, R. A. (1981). An electron and light microscope study of the caecal tonsil: The basic unit of the caecal tonsil. J. Dev. Comp. Immunol. 5, 95–104. Golub, E. S., and Green, D. R. (1991). ‘‘Immunology: A Synthesis,’’ 2nd Edition. Sinauer Associates, Sumderland, MA. Gordon, J., Flores-Romo, L., Cairns, J., Millsum, M., Lane, P., Johnson, G., and MacLennan, I. C. M. (1989). CD23: A multifunctional receptor/lymphokine? Immunol. Today 10, 153–157. Haury, M., Kasahara, Y., Schaal, S., Bucy, R., and Cooper, M. D. (1993). Intestinal T-lymphocytes in the chicken express an integrin–like antigen. Eur. J. Immunol. 23, 313–319. Higgins, D. A., and Warr, G. W. (1993). Duck immunoglobulins: Structure, functions and molecular genetics. Avian Pathol. 22, 211–230. Holbrook, K. A., Perkins, W. D., and Glick, B. (1974). The fine structure of the bursa of Fabricius: ‘‘B’’ cell surface configuration and lymphoepithelial organization revealed by scanning and transmission electron microscopy. J. Reticuloendothel. Soc. 16, 300–311. Houssaint, E., Belo, M., and LeDouarin, N. M. (1976). Investigations on cell lineage and tissue interactions in the developing bursa of Fabricius through interspecific chimeras. Dev. Biol. 53, 200–264. Howard, M. C., Miyajima, A., and Coffman, R. (1993). T-cell derived cytokines and their receptors. In ‘‘Fundamentals of Immunology’’ (W. E. Paul, ed.) pp. 763–800. Raven, New York. Issaacs, A., and Lindenmann, J. (1957). Virus interference. I. The interferon. Proc. Roy Soc. London (Biol.) 147, 258–267. Jaffe, W. P., and Fechheimer, N. S. (1966). Cell transport and the bursa of Fabricius. Nature (London) 212, 92. Janeway, C. A., Jr., and Travers, P. (1994). ‘‘Immunobiology, The Immune System in Health and Disease.’’ Garland Publishing, New York. Jeurissen, S. H. M. (1993). The role of various compartments in the chicken spleen during an antigen–specific humoral response. Immunology 80, 29–33. Jankovic, B. D., and Mitrovic, K. (1967). Antibody producing cells in the chicken, as observed by fluorescent antibody technique. Folia. Biol. 13, 406–410. Jeurissen, S. H. M., Janse, E. M., and Koch, G. (1989a). Meckel’s diverticle: A gut-associated lymphoid organ in chickens. Adv. Exp. Med. Biol. 237, 599–606. Jeurissen, S. H. M., Janse, E. M., Koch, G., and deBoer, G.F. (1989b). Post natal development of mucosa–associated lymphoid tissues in chickens. Cell Tissue Res. 258, 119–124.

Jeurissen, S. H. M., Claassen, E., and Janse, E. M. (1992). Histological and functional differentiation of non-lymphoid cells in the chicken spleen. Immunology 77, 75–80. Jolly, J. (1915). Le bourse de Fabricius et les organes lymphoepitheliaux. Arch. Anat. Microsc. Morphol. Exp. 16, 363–546. Joshi, P., and Glick, B. (1990). The role of avian macrophages in the production of avian lymphokines. Dev. Comp. Immunol. 14, 319–325. Kampmeier, O. F. (1969). ‘‘Evolution and Comparative Morphology of the Lymphatic System.’’ Charles C. Thomas, Springfield, IL. Kaspers, B., Lillehoj, H. S., and Lillehoj, E. P. (1993). Chicken macrophages and thrombocytes share a common cell surface antigen defined by a monoclonal antibody. Vet. Immunol. Immunopathol. 36, 333–346. Kaufman, J., and Salomonsen, J. (1993). What in the dickens is with those chickens? An only slightly silly response to the first draft of Langman and Cohn. Res. Immunol. 144, 495–502. Kaufman, J., Skjodt, K., and Salomonsen, J. (1991). The B-G multigene family of the chicken major histocompatibility complex. Crit. Rev. Immunol. 11, 113–143. Kjalke, M., Welender, K. G., and Koch, C. (1994). Structural analysis of chicken factor B-like protease and comparison with mammalian complement proteins factor B and C2. J. Immunol. 151, 4147–4152. Klasing, K. C. (1994). Avian leukocytic cytokines. Poult. Sci. 73, 1035– 1043. Koch, C. (1987). Complement system in avian species. In ‘‘Avian Immunology Basics and Practice’’ (A. Toivanen and P. Toivanen, eds.), Vol. II, pp. 43–55. CRC Press, Boca Raton, FL. Kunicki, T., and Newman, P., (1985). Synthesis of analog of human platelet membrane glycoprotein 11b-111a complex by chicken peripheral blood monocytes. Proc. Natl. Acad. Sci. USA 82, 7319– 7323. Langman, R. E., and Cohn, M. (1993). A theory of the ontogeny of the chicken immune system: The consequences of diversification by gene conversion hyperconversion and its extension to the rabbit. Res. Immunol. 144, 422–496. Le Douarin, N. M., Michel, G., and Baulieu, E. E. (1980). Studies of testosterone-induced involation of the bursa of Fabricius. Dev. Biol. 75, 288–302. Le Douarin, N. M., Dieterlen-Lievre, F., and Oliver, P. D. (1984). Ontogeny of primary lymphoid organs and lymphoid stem cells. Am. J. Anat. 170, 261–299. Le Douarin, N. M., Martin, C., Ohki-Hamazaki, H., Belo, M., Coltey, M. D., and Corbel, C. (1990). Development of the immune system and self/non-self recognition studied in the avian embryo. In ‘‘The Avian Model in Developmental Biology: From Organism to Genes’’ (N. Le Douarin, F. Dieterlen-Lievre, and J. Smith, eds.), pp. 219–237. Edition Du Centre National De La Recherche Scientifique, Paris, France. Lerner, K. G., Glick, B., and McDuffie, F. C. (1971). Role of the bursa of Fabricius in IgG and IgM production in the chicken: Evidence for the role of a non-bursal site in the development of humoral immunity. J. Immunol. 107, 493–503. Leslie, G. A., and Clem, L. W. (1969). Phylogeny of immunoglobulin structure and function. III. Immunoglobulins of the chicken. J. Exp. Med. 130, 1337–1352. Leutz, A., Damm, K., Sterneck, E., Kowenz, E., Ness, S., Frank, R., Gauseponl, H., Pan, Y. C., Smart, J., Haiman, M., and Graft, T. (1989). Molecular cloning of the chicken myelomonocytic growth factor (cMGF) reveals relationship to interleukin-6 and granulocyte colony stimulating factor. EMBO J. 8, 175–181. Lillehoj, H. S., Kaoporo, B., Jenkins, M. O., and Lillehoj, E. P. (1992). Avian interleukin-2: A review by comparison with mammalian homologues. Poult. Sci. Rev. 4, 67–86.

Chapter 26. Immunophysiology Linsley, P. S., and Ledbetter, J. A. (1993). The role of the CD28 receptor during T-cell responses to antigen. Annu. Rev. Immunol. 11, 191–212. Magor, K. E., Warr, G. W., Middleton, D., Wilson, M. R., and Higgins, J. D. A. (1992). Structural relationship between the two IgY of the duck, Anas platyrhynchos: Molecular genetic evidence. J. Immunol. 149, 2627–2633. Mansikka, A. (1992). Chicken IgA H chains: Implications concerning the evolution of H chain genes. J. Immunol. 149, 855–861. Mansikka, A., Sandberg, M., Veromaa, T., Vainio, O., Granfors, K., and Toivanen, P. (1989). B-cell maturation in the chicken Harderian Gland. J. Immunol. 142, 1826–1833. Marsh, J. (1993). The humoral activity of the avian thymic microenvironment. Poult. Sci. 72, 1294–1300. Martin, C. (1990). Quail chick chimeras, a tool for developmental immunology. In ‘‘The Avian Model in Developmental Biology: From Organisms to Genes’’ (N. Le Douarin, F. Dieterlen-Lievre, and J. Smith, eds), pp. 207–217. Editions Du Centre Nationale De La Recherche Scientifique, Paris, France. Martin, T. R., Mongovin, S. M., Tobias, P. S., Mathison, J. C., Moriarty, A. M., Leturcq, D. J., and Ulevitch, R. J., (1994). The CD14 differentiation antigen mediates the development of endotoxin responsiveness during differentiation of mononuclear phagocytes. J. Leuk. Biol. 56, 1–9. Masteller, E. L., and Thompson, C. B. (1994). B-cell development in the chicken. Poult. Sci. 73, 998–1011. McCorkle, F. M., Stinson, R. S., Olah, I., and Glick, B. (1979). The chicken’s femoral lymph nodules: T&B cells and the immune response. J. Immunol. 123, 667–669. McCormack, W. T., Tjoelker, L. W., and Thompson, C. B. (1991). Avian B-cell development: Generation of an immunoglobulin repertoire by gene conversion. Annu. Rev. Immunol. 9, 219–241. Moore, M. A. S., and Owen, J. J. T. (1965). Chromosome marker studies on the development of the hematopoietic system in the chick embryo. Nature (London) 708, 956,989–990. Moore, M. A. S., and Owen, J. J. T. (1966). Experimental studies in the development of the bursa of Fabricius. Dev. Biol. 14, 40–51. Mueller, A., Sato, K., and Glick, B. (1971). The chicken lacrimal gland, gland of Harder, caecal tonsil, and accessory spleens as sources of antibody producing cells. Cell. Immunol. 2, 140–152. Murthy, K. K., Pace, J. L., Barger, B. O., Dawe, D. L., and Ragland, W. L. (1984). Localization and distribution by age and species of a thymus-specific antigen. Thymus 6, 43–56. Muthmann, E., (1913). Beitrage Zur vergleichenden anatomie des Blindarmes und der lymphoiden organe des darmkanals bei saugetieren und vogeln. Anat. Hefte 48, 67–114. Naukkarinen, A., Arstela, A. U., and Sorvari, T. E. (1978). Morphological and functional differentiation of the surface epithelium of the bursa of Fabricii in chicken. Anat. Rec. 191, 415–432. Nicholas-Bolnet, C., Yassine, F., Cormier, F., and Dieterlen-Lievre, F. (1991). Developmental kinetics of hemopoietic progenitors in the avian embryo spleen. Exp. Cell Res. 196, 294–301. Nickel, R., Schummer, A., Seiferle, E., Siller, W. G., and Wight, P. A. L., (1977). ‘‘Anatomy of the Domestic Birds.’’ SpringerVerlag, New York. Niedorf, H. R., and Wolters, B. (1978). Development of the Harderian gland in the chicken: Light and electron microscopic investigations. Invest. Cell. Pathol. 1, 205–215. Olah, I., and Glick, B. (1978a). The number and size of the follicular epithelium (FE) and follicles in the bursa of Fabricius. Poult. Sci. 57, 1445–1450. Olah, I., and Glick, B. (1978b). Secretory cell in the medulla of the bursa of Fabricius. Experientia 34, 1642–1643.

669

Olah, I., and Glick, B. (1982). Splenic white pulp and associated vascular channels in chicken spleen. Am. J. Anat. 165, 445–480. Olah, I., and Glick, B. (1983). Avian lymph node: Light and electron microscopic study. Anat. Rec. 205, 287–299. Olah, I., and Glick, B. (1984a). Meckel’s diverticulum. I. Extra medullary myelopoiesis in the yolk sac of hatched chickens (Gallus domesticus). Anat. Rec. 208, 243–252. Olah, I., and Glick, B. (1984b). Lymphopineal tissue in the chicken. Dev. Comp. Immunol. 8, 855–862. Olah, I., and Glick, B. (1985). Lymphocyte migration through the lymphatic sinuses of the chicken’s lymph node. Poult. Sci. 64, 159–168. Olah, I., and Glick, B. (1991). An Ig-A-like substance in the chicken’s pineal. Experientia 147, 202–205. Olah, I., Glick, B., and Taylor, R. L., Jr. (1984). Meckel’s diverticulum. II. A novel lymphoepithelial organ in the chicken. Anat. Rec. 208, 253–263. Olah, I., Glick, B., and Toro, I. (1986). Bursal development in normal and testosterone-treated chick embryos. Poult. Sci. 65, 574–588. Olah, I., Medgyes, T., and Glick, B. (1988). Origin of aortic cell clusters in the chicken embryo. Anat. Rec. 222, 60–68. Olah, I., Kendall, C., and Glick, B. (1991). Bursal secretory-dendritic cell express a homologue of IgGFc. Fed. Am. Soc. Exp. Biol. J. 5, A1064. [Abstract] Olah, I., Scott, T. R., Gallego, M. Kendall, C., and Glick, B. (1992a). Plasma cells expressing immunoglobulins M and A but not immunoglobulin B develop an intimate relationship with central canal epithelium in the Harderian gland of the chicken. Poult. Sci. 71, 664–676. Olah, I., Kendall, C., and Glick, B. (1992b). Anti-vimentin monoclonal antibody recognizes a cell with dendritic appearance in the chick’s bursa of Fabricius. Anat. Rec. 232, 121–125. Orlans, E., and Rose, M. E. (1970). Antibody formation by transferred cells in inbred fowls. Immunology 18, 473–482. Paramithiotis, E., Tkalec, L., and Ratcliff, M. J. H. (1991). High levels of CD45 are coordinately expressed with CD4 and CD8 in avian thymocytes. J. Immunol. 147, 3710–3717. Parvari, R., Avivi, A., Lentner, F., Ziv, E., Tel-or, S., Burstein, Y., and Schechter, I. (1988). Chicken immunoglobulin 웂 heavy chains: limited VH gene repertoire, combinatorial diversification by D gene segments and evolution of the heavy chicken locus. EMBO J. 7, 739–744. Paul, W. E. (1993). ‘‘Fundamental Immunology.’’ Raven, New York. Pharr, G. T., Hunt, H. D. , Bacon, L. D., and Dodgson, J. B. (1993). Identification of class II major histocompatibility complex Polymorphisms predicted to be important in peptide antigen presentation. Poult. Sci. 72, 1312–1317. Powell, P. C. (1987). Macrophages and other non-lymphoid cells contributing to immunity. In ‘‘Avian Immunology: Basis and Practice’’ (A. Toivanen and P. Toivanen, eds.), Vol. 1, pp. 195–212. CRC Press, Boca Raton, FL. Qureshi, M. A., Petitte, J. H., Laster, S. M., and Dietert, R. R. (1993). Avian macrophages: Contribution to cellular microenvironment and changes in effector functions following activation. Poult. Sci. 72, 1280–1284. Qureshi, M. A., Marsh, J. A., Dietert, R. R., Sung, Y.-J. NicholasBolnet, C., and Petitte, J. H. (1994). Profiles of chicken macrophage effector functions. Poult. Sci. 73, 1027–1034. Ratcliffe, M. Lassila, O., Pink, J. R. L., and Vainio, O. (1986). Avian B-cell precursors: Surface immunoglobulin expression is an early, possibly bursa-independent event. Eur. J. Immunol. 16, 129–133. Roitt, I. M., Torrigiani, G., Greaves, M. F., Brostaff, J., and Playfair, J. H. (1969). The cellular basis of immunological responses. Lancet 2, 367–371.

670

B. Glick

Romanoff, A. (1960). ‘‘The Avian Embryo: Structural and Functional Development.’’ Macmillan, New York. Romieu, M., and Jullien, G. (1942). Sur l‘existence d‘une formation lymphoid dans l‘epiphyse des Gallinces. C. R. Soc. Biol. 136, 626–628. Salanti, E. P., Pink, J., Richard, L., and Steinberg, C. M. (1989). A model of bursal colonization. Theor. Biol. 139, 1–6. Savage, M. L., Olah, I., and Scott, T. R. (1992). Plasma cell proliferation in the chicken Harderian gland. Cell Prolif. 25, 337–344. Schat, K. A., and Myers, T. J. (1991). Avian intestinal immunity. Crit. Rev. Poult. Biol. 3, 19–34. Scott, T. R., and Savage, M. L. (1996). Immune cell proliferation in the Harderian gland: An avian model. Microscopy Res. Tech. 29, 149–155. Scott, T. R., Savage, M. L., and Olah, I. (1993). Plasma cells of the chicken Harderian gland. Poult. Sci. 72, 1273–1279. Sharma, J. M., and Schat, J. A. (1991). Natural immune functions. In ‘‘Avian Cellular Immunology’’ ( J. M. Sharma, ed.), pp. 51–70. CRC Press, Boca Raton, FL. Sicher, S. C., Vazquez, M. A., and Christopher, Y. Lu (1994). Inhibition of macrophage la expression by nitric oxide. J. Immunol. 153, 1293–1300. Silverstein, A. M. (1989). ‘‘History of Immunology.’’ Academic Press, Orlando, FL. Stalsberg, H., and Prydz, H. (1963). Studies on chick thrombocytes II. Function in primary hemostasis. Thromb. Diath. Haemorr. 9, 291–299. Sundick, Roy S., Albini, B., and Wick, G. (1973). Chicken Harderian’s gland: Evidence for a relatively pure bursa dependent lymphoid cell population. Cell Immunol. 7, 332–335. Sung, Y.-J., and Dietert, R. R. (1994). Nitric oxide (⭈No)-induced mitochondrial injury among chickens ⭈NO-generating and target leukocytes. J. Leuk. Biol. 56, 52–58. Tenenhouse, H. S., and Deutsch, H. F. (1966). Some physical chemical properties of chicken 웂-globulins and their pepsin and papain digestion products. Immunochemistry 3, 11–20. Thorbecke, G. J., Palladino, M. A., and Lerman, S. P. (1980). Lymphoid-cell cooperation in immune responses of the chicken. Contemp. Top. Immunol. 9, 91–107. Toivanen, P., Toivanen, A., and Vanio, D. (1974). Complete restoration of bursa-dependent immune system after transplantation of

semi-allogenic stem cells into immunodeficient chickens. J. Exp. Med. 139, 1344–1349. Tsuji, S., Baba, T., Kawata, T., and Kajikawa, T. (1993). Role of Harderian gland on differentiation and proliferation of immunoglobulin A-bearing lymphocytes in chickens. Vet. Immunol. Immunopathol. 37, 276–283. Vainio, O., and Toivanen, A. (1987). Cellular cooperation in immunity. In ‘‘Avian Immunology: Basis and Practice’’ (A. Toivanen and P. Toivanen, eds.), Vol. II, pp. 1–12. CRC Press, Boca Raton, FL. Vainio, O., Veromaa, T., Eerola, E., Toivanen, P., and Radcliffe, M. J. H. (1988). Antigen presenting cell T-cell interaction in the chicken is MHC Class II antigen restricted. J. Immunol. 140, 2864– 2868. Venzke, W. G. (1952). Morphogenesis of the thymus of chicken embryos. Am. J. Vet. Res. 13, 395–404. Veillette, A., and Ratcliffe, M. J. H. (1991). Avian CD4 and CD8 interact with a cellular tyrosine protein kinase homologous to mammalian p56lck. J. Immunol. 21, 397–401. von Rautenfeld, D. B., and Budras, K. D. (1982). The bursa cloacae (Fabricii) of struthioniformes in comparison with the bursa of other birds. J. Morphol. 172, 123–138. Warner, N. L., Szenberg, A. J., and Burnet, F. M. (1962). The immunological role of different lymphoid organs in the chicken. I. Dissociation of immunological responsiveness. Aust. J. Exp. Biol. Med. Sci. 40, 373–388. Weinbaum, F. I., Glimour, D. G., and Thorbecke, G. J. (1973). Immunocompetent cells of the chicken. III. Cooperation of carrier sensitized T-cells from agammaglobulinemic donors with Hapten immune B-cells. J. Immunol. 110, 1434–1436. Weiss, L. (1972). ‘‘The cells and tissues of the immune system.’’ Prentice-Hall, Englewood Cliffs, NJ. White, R. G., French, V. I., and Stark, J. M. (1970). A study of the localization of a protein integrin in the chicken spleen and its relation to the formation of germinal centres. J. Med. Microbiol. 3, 65–83. Wight, P. A. L., Burns, R. B., Rothwell, B., and MacKenzie, G. M. (1971). The Harderian gland of the domestic fowl. I. Histology, with reference to the genesis of plasma cells and Russell bodies. J. Anat. 120, 307–315. Young, J. R., Davison, T. F., Tregaskes, C. A., Rennie, M. C., and Vainio, O. (1994). Monomeric homologue of mammalian CD28 is expressed on chicken T-cells. J. Immunol. 152, 3848–3851.