A Dissertation

entitled

Discovery of a New Subset Derived from Immature

By

Shuo Geng

Submitted to the Graduate Fculty as partial fulfillment of the requirement for the Doctor

of Philosophy in Biomedical Sciences

______Dr. Akira Takashima, Committee Chair

______Dr. Kevin Pan, Committee Member

______Dr. Stanislaw Stepkowski, Committee Member

______Dr. Anthony Quinn, Committee Member

______Dr. Randall Worth, Committee Member

______Dr. Patricia R. Komuniecki, Dean College of Graduate Studies

The University of Toledo

June 2011

Copyright 2011, Shuo Geng

This document is copyrighted material. Under copyright law, no parts of this document may be reproduced without the expressed permission of the author. An Abstract of

Discovery of a New Dendritic Cell Subset Derived from Immature Granulocytes

By

Shuo Geng

Submitted to the Graduate Fculty as partial fulfillment of the requirement for the Doctor of Philosophy in Biomedical Sciences

Compared to other leukocytes, dendritic cells (DCs) are extremely heterogeneous in terms of developmental pathways and immunological properties. More than 10 DC subsets have been identified in mouse, which are distinguishable from each other by surface phenotypes, functions, tissue distributions, and developmental origins. Here we describe a novel subset of DCs that are derived from immature granulocytes, known as band cells. Large numbers of band cells are rapidly recruited to inflammatory sites, where some of them differentiate into this DC subset termed “grDCs.” In addition to showing a typical DC-like morphology, grDCs express many DC markers (MHC class II, CD11c, and CD205) and efficiently present peptide antigens to both CD8 and CD4 T cells.

Importantly, grDCs retain the surface expression of Ly6G, which is not detectable on any of the currently known DC subsets, as well as several unique features of granulocytes.

GeneChip analyses have revealed a cluster of genes that are selectively expressed by

Ly6G+ grDCs, but not Ly6G- conventional DCs – this cluster includes cathelicidin

(CRAMP), MMP9 and CD62L. CRAMP-deficient grDC exhibited a diminished bacterial killing activity, indicating a functional contribution of grDC-associated CRAMP. In a peritoneal E. coli infection model, grDC were found to internalize bacteria efficiently and present bacterial antigens to CD4 T cells. Not only have our data unveiled a previously

iii unrecognized DC subset and its functions, they also suggest a new concept that granulocytes can directly participate in the induction of adaptive immunity via differentiation into grDCs.

iv DEDICATION

This work is dedicated to my parents, Wu-qun Geng and You-jie Zhu, as well as my grandparents, Ling-guang Zhu and Jie li, for their endless love. They always have faith in me and encourage me to reach my dreams.

This work is also dedicated to my beloved wife, Ran Lu, for her support, understanding and patience. She has been my wonderful cheerleader and makes my life delightful.

v

ACKNOWLEDGEMENT

I would like to thank my major advisor, Dr Akira Takashima, for giving me the opportunity to work on this exciting project. I appreciate his expert training, sagacious guidance, and sustained encouragement during my doctoral education. His enthusiasm in scientific research has always inspired me. He leads me into the field of and

I have benefited a lot from him.

I would like to thank me committee members: Dr. Kevin Pan, Dr. Stanislaw Stepkowski,

Dr. Anthony Quinn and Dr. Randall Worth for their valuable time and constructive suggestions and criticisms during my PhD study.

I am grateful to Dr. Robert Blumenthal and Dr. Kristen Williams for providing the E. coli strains used in this study and teaching me how to perform bacterial assays. I thank Dr.

William Gunning III for helping us take the EM images. I thank Dr. Richard Gallo for providing anti-CRAMP antibody and BM cells from CRAMP KO mice. I thank Sean

Linkes for his service of FACS analysis and sorting.

I thank all the current and former members in Takashima’s lab: Dr. Hironori Matsushima,

Dr. Ram Veerapaneni, Dr. Claudio Cortes, Dr. Masaaki Miyazawa, Dr. Nobuyasu

Mayuzumi, Dr. Taehyung Lee, Ran Lu, Yi Yao, David Leggat, Benjamin Chojnacki,

Rachel Mohr, Alan Rupp and Colleen Krout. It is a great pleasure to work with them.

vi I would like to thank all the administrative personnel of the Department of Medical

Microbiology and Immunology: Diane Ammons, Tamra Chamberlin, Traci McDaniel and Suzanne Payne, who are always ready to help and solve problems.

I acknowledge all the faculty, staff and students in the Department of Medical

Microbiology and Immunology and the Track of Infection, Immunity and transplantation.

They create a very professional and friendly environment.

vii Contents

ABSTRACT……………………………………………………………...... iii

DEDICATION……………………………………………………...……….v

ACKNOWLEDGEMENT…………………………….…….……..…....….vi

CONTENTS……………………...... ……………….…………....…viii

LIST OF ABBREVIATIONS…………………………………………...….ix

INTRODUCTION…………………………………………………………...1

LITERATURE REVIEW……………………………………………………5

MATERIALS AND METHODS…………………………………………..34

RESULTS…………………………………………………………………..46

DISCUSSION…..………………………………………………………….79

SUMMARY………………………………………………………………..90

REFERENCES……………………………………………………………..91

viii

LIST OF ABBREVIATIONS

APC antigen presenting cell BM BMDC bone marrow-derived dendritic cell cDC conventional dendritic cell CDP common Dendritic cell precursor CFU colony forming unit CHS contact hypersensitivity CLP common lymphoid precursor CMP common myeloid precursor CRAMP cathelicidin CTL cytotoxic T lymphocyte DC dendritic cell DT diphtheria toxin EM electron microscopy Ep-CAM epithelial cell adhesion molecule ER endoplasmic reticulum FDC follicular cendritic cell Flt3 fms-related tyrosin kinase 3 GC germinal center GCDC germinal center dendritic cell G-CSF colony-stimulating factor GM-CSF granulocyte colony-stimulating factor grDC granulocyte-derived dendritic cell IFN interferon IKDC interferon-producing killer dendritic cell i.p. intraperitoneal i.v. intravenous KO knock out LC LN lymph node M-CSF macrophage colony-stimulating factor MFI mean fluorescence intensity MHC major histocompatibility complex MMR macrophage mannose receptor NK natural killer moDC -derived dendritic cell OVA ovalbumin OX oxazolone pDC plasmacytoid dendritic cell

ix PEC peritoneal exudate cell PMNp precursor of polymorphonuclear granulocyte RA rheumatoid arthritis ROS reactive oxygen species SP spleen SR scavenger receptor TG transgenic Tip-DC TNF-α/iNOS producing dendritic cell TLR toll like receptor UV ultraviolet WT wild type

x INTRODUCTION

Dendritic cells (DCs) are specialized antigen presenting cells (APCs), which initiate and modulate the immune response as well as maintain the immune tolerance to self-antigens.

Although Langerhans cells (LCs), a DC subset, were observed as early as 1868, the characterization of DC only began about 25 years ago (Banchereau & Steinman, 1998;

Paul, 2007). Immature DCs, abundant in the body surface, have a potent capacity to uptake antigens and subsequently migrate to lymphoid tissues, where they mature. Then the processed antigen peptides will be loaded onto major histocompatibility complex

(MHC) molecules and presented to CD4 or CD8 T cells, resulting in a dramatic expansion of T cells. Activated CD8 cytotoxic T cells migrate back to the inflamed sites to eliminate the infected cells, while CD4 helper T cells secret cytokines to facilitate the activation of , natural killer (NK) cells and or the antibody production of B cells. B cells also can be activated by direct contact with DCs, and produce neutralizing antibodies to the initial pathogen (Banchereau et al., 2000).

Compared to other bone marrow (BM) derived leukocytes, DC represent a highly heterogeneous population, since various DC subsets have been reported in the literature.

Generally speaking, DC can be divided into two families. One is conventional DC (cDC), which has a dendritic morphology and potent antigen processing and presentation capacity. The other one is plasmacytoid DC (pDC), which is a round, non-dendritic shape, and relatively long-lived cell (Facchetti & Vergoni, 2000; Liu 2005). There are at least

1 seven cDC subsets identified in mouse. Recently, a novel DC subset, interferon- producing killer DC (IKDC), was identified from mouse spleen (Bonmort et al., 2008).

Although they share many common features, different DC subsets have been demonstrated to have distinct phenotype, tissue distribution and biological functions. The diversity of DCs permits the immune system to mount various types of responses to different pathogens and stimulus (Shortman & Liu, 2002; Merad & Manz, 2009).

Most of these DC subsets have a short lifespan, based on the evidence that only less than

5% of DCs are dividing and the half-life is 1.5-7 days in the steady state. DCs must be renewed continually, perhaps by the cells originating from hematopoietic stem cells or

BM or some tissue resident precursor cells. The differentiation of DC is crucial not only to the establishment of DC network, but also to the dynamic response against pathogens, self-antigens or tumor cells. The research on DC differentiation and identification of DC precursors is also important to DC-based immunotherapy, which can improve methods to generate immuno-competent DCs more efficiently (Kamath, et al., 2000; Kamath et al.,

2002; Glibos, 2007; Liu et al., 2007). However, the complex of DC populations makes it challenging to clarify the differentiation and origin of DCs because each DC subset may have some specific precursors or developmental pathways. DC development is relatively flexible, and it has been reported that both myeloid and lymphoid progenitors can give rise to both cDCs and pDCs. In addition, DC development is subject to change in inflammatory conditions and is distinguished from that in steady state condition (Manz et al., 2001; Shortman & Nail, 2007). Due to the high heterogeneity and complicated

2 differentiation, it is possible that some unknown DC subset deriving from a novel pathway plays an important role in immune response.

Recently, we constructed a new transgenic (TG) mouse line expressing DsRed fluorescence marker gene under the control of IL-1β promoter (Matsushima et al., 2010).

From the BM culture of TG mice in the presence of GM-CSF, we identified a

DsRed+/CD11b+/CD11c- population, which was able to give rise to CD11c+ DCs. Based on the morphology, surface phenotype and high endocytotic potential, this population resembled the immature granulocytes, known as band cells. We also identified the equivalent band cell population in crude BM cells from wild type (WT) mice. Not only did the band cells differentiate into mature granulocyte in the presence of G-CSF, they also exhibited a potential to give rise to DCs in when cultured with GM-CSF.

Interestingly, the DCs derived from band cells retained the surface expression of Ly6G, a marker for granulocytes. Since no Ly6G+ DC subset has been reported in the literature, this DC population may represent a novel DC subset, which we have termed “grDC.”

Similar to their precursor band cells, grDCs showed significantly higher potential than cDCs in uptaking extracellular particles as well as antigens. Microarray analysis revealed about 300 genes expressed differentially between grDCs and cDCs, and more than 100 genes were expressed predominantly by grDCs, including cathelicidin (cramp), l-selectin

(cd62l) and mmp9. Compared to cDCs, grDCs were able to kill bacteria more efficiently, and this special capacity was possibly due to production of CRAMP, since the grDCs from CRAMP KO mice exhibited significantly diminished efficiency to kill bacteria. We

3 also identified grDCs from different tissues of WT mice, and the number increases dramatically in the inflamed peritoneal cavity. Adoptive transfer experiments demonstrated that band cells differentiated into DCs in vivo only under inflammatory conditions but not under steady state. Importantly, grDCs showed a more potent capacity to present antigens to T cells compared to cDCs after they internalized bacteria in vivo.

This may be due to their ability to uptake a larger amount of bacteria and to be able to loaded more bacterial antigen peptides on MHC molecules.

In conclusion, the present study unveils an alternative pathway for DC development in which immature granulocytes give rise to DCs (grDCs) under inflammatory conditions.

Moreover, the grDCs represent a novel DC subset with distinct phenotype, gene expression profiles and specialized functions.

4 LITERATURE REVIEW

OVERVIEW OF DC

Since the first identification in the mouse spleen by Dr. Ralph Steinmain in 1970’s

(Steinman & Cohn 1973; 1974), DCs have drawn more and more attention, and many efforts have been devoted into the characterization of DCs. It has been well known that

DCs are the most potent APCs, which initiate, regulate and maintain the immune responses. DCs are able to uptake various antigens and process them into peptides, which can be loaded onto the MHC I or MHC II molecules, and then present the antigen to T cells to induce the expansion of antigen specific T cells. Moreover, DCs make contributions to humoral immunity, since they can directly activate B cells to produce antibodies. Also, DCs possess the capacity to activate innate leukocytes, such as NK cells and NKT cells (Banchereau et al., 2000; Guermonprez et al., 2002)

Antigen uptake and DC maturation

In the steady state, DCs are found in most tissues, both peripheral and lymphoid tissues.

Peripheral DCs reside especially at the surface of the body, such as skin, vagina, ectocervix, anus, as well as the internal and mucosal surfaces (Steinman & Banchereau,

2007; Niess et al., 2005). The majority of these DCs are in an immature state, and they express low levels of surface MHC II and co-stimulatory molecules (CD40, CD80 and

CD86), which are required for the activation of T cells. For example, freshly isolated LCs from the epidermis showed a limited antigen presentation capacity due to the low

5 expression of surface MHC and co-stimulatory molecules (Banchereau & Steinman, 1998;

Bell et al., 1999). However, immature DCs can capture extracellular antigens and molecules, mainly by three mechanisms. First, the insoluble particles and microbes, such as latex beads, cell debris, bacteria, virus and some parasites, can be internalized via , an actin dependent procedure (Moll 1993; Reis e Sousa et al., 1993;

Matsuno et al., 1996; Svensson et al., 1997; Albert et al., 1998a; Albert et al., 1998b).

Second, the extracellular fluid and the small molecules within it can be internalized through a process named macropinocytosis (Sallusto & Lanzavecchia, 1995). Third, different kinds of surface receptors on DCs can mediate rapid endocytosis. C-type lectin receptors, including DEC205 (CD205), macrophage mannose receptor (MMR/CD206),

DC-SIGN (CD209) and dectin-2, recognize the glycoproteins expressed on the surface of infectious pathogens (Jiang et al., 1995; Sallusto & Lanzavecchia, 1995; Engering et al.,

1997; Tan et al., 1997; Geijtenbeek et al., 2000; Figdor et al., 2002; Neumann et al.,

2008). CD91, a receptor for heat shock proteins derived from tumor cells or infected cells, has been identified on mouse APCs to mediate the internalization of Hsc70 and gp90

(Basu et al., 2001). Fcγ and Fcε receptors recognize the immune complexes as well as opsonized particles (Esposito-Farese et al., 1995; Jurgens et al., 1995; Fanger et al., 1996).

In addition, scavenger receptors (SR) on DC surface recognize and mediate the endocytosis of particles with modified low-density lipoprotein (Platt et al., 1998;

Nakamura et al., 2001). In other , for example macrophages, macropinocytosis is transiently induced by some growth factors or phorbol easter. In contrast, macropinocytosis is constitutive in immature DCs, and it allows DCs to rapidly and nonspecifically sample large amount of fluids. Because of their potent macropinocytosis

6 and receptor mediated endocytosis, DCs have much higer antigen uptake and presentation efficiency compared to other APCs, even picomolar concentration of antigens is adequate for DCs to activate T cells (Sallusto & Lanzavecchia, 1995).

After encountering pathogen-associated molecular patterns or danger-associated molecular patterns, DCs undergo a process called maturation or activation, in which their antigen uptake ability is dramatically diminished, but the antigen presentation capacity is greatly boosted. A broad range of environmental and endogenous factors have the capacity to induce the maturation of DC. Microbial related molecules, such as glycolipids, lipoproteins, LPS, flagellin, bacterial DNA, single-stranded and double-stranded RNA, activate DCs via toll like receptors (TLRs) (Cella et al., 1999; Hartmann et al., 1999;

Rescigno et al., 1999; Aderem & Ulevitch, 2000; Muzio et al., 2000; Hertz et al., 2001;

Chua et al., 2003). Viral RNA also can be recognized by cytoplasmic RIG-I and Mda5 receptors and induce DC maturation by a TLR independent mechanism (Kawai et al.,

2005; Lo´pez et al., 2006). Innate lymphocytes, NK cells, NKT cells and γδ T cells, recognize tumors or pathogen-derived antigens of infected cells, and then they can interact with DC to induce DC maturation through cell-cell contact (Münz et al., 2005).

Furthermore, many endogenous ligands have the potential to activate DCs. For example, histamine, heat shock proteins, defensins, extracellular ATP and uric acid released from dying cells have been shown to induce the DC maturation (Rovere et al., 1998; Shi et al.,

2000; Singh-Jasuja et al., 2000). In addition, several proinflammatory cytokines, including TNF-α, IL-1β, IL-4 and IL-6, promote DC maturation both in vitro and in vivo

(Aiba, 1998). After maturation, DCs acquire high levels of surface MHC II as well as the

7 co-stimulatory molecules, but lose some endocytic receptors. Furthermore, DC maturation is accompanied by morphological changes via cytoskeleton remodeling, loss of adhesion molecule and up-regulation of a chemokine receptor, CCR7 (Rescigno et al.,

1997; Winzler et al., 1997; Yanagihara et al., 1998; Granucci et al., 1999). These changes facilitate the migration of activated DCs to draining lymph nodes (LNs) and spleen, where DCs complete their maturation.

Migration of DCs from peripheral inflamed sites to lymphoid tissues is mediated by gradients of CCL19 (MIP-3β) and CCL21 (6Ckine), both of which are CCR7 ligands and can be secreted by the cells around T cell area in the draining LNs. The DCs entering the

LNs can subsequently become the new sources of CCL19 and CCL21, thereby amplifying the chemotaxis signal (Dieu et al., 1998; Chan et al., 1999). These two chemokines can also attract not only DCs but also naïve T cells (Gunn et al., 1998; Ngo et al., 1998; Luther, et al., 2002). Also, matured DCs in the T cell area release CX3CL1

(Fractalkine), CXCL8 (IL-8) and CCL22 (macrophage-derived chemokine) to attract T cells as well as other lymphocytes (Kanazawa et al., 1999; Kikuchi et al., 2005; Oz-

Arslan et al., 2006; Piqueras et at., 2006). In other words, this chemokine-mediated mechanism contributes to an environment where antigen-bearing DCs can easily identify antigen-reactive T cells. Upon contact with T cells, DCs receive further maturation signals by CD40L, RANK/TRANCE and 4-1BB, allowing them transform from antigen- capturing cells to antigen-presenting cells (Caux et al., 1994; DeBenedette et al., 1997;

Wong et al., 1997; Alvarez et al., 2008).

8 Antigen processing and presentation

To activate antigen specific T cells, DC need to process antigens into short peptides and load them into the groove of surface expressing MHC II or MHC I molecules. The antigens loaded on MHC I are presented to CD8+ T cells, while the antigens loaded on

MHC II are presented to CD4+ T cells.

Extracellular microorganisms and apoptotic bodies are efficiently captured by immature

DCs and transferred into membrane-enclosed compartments, phagosomes. After a series of modifications, the phagosomes fuse with lysosome to form phagolysosome, in which the antigens can be digested by the acid hydrolase enzymes (Inaba et al., 1998; Vyas et al., 2008). MHC II molecules are accumulated in lysosome-related compartments in immature DCs, known as MHC class II-rich compartments (MIICs) (Kleijmeer et al.,

1995; Nijman, et al., 1995; Cresswell, 1996). The fusion of MIICs with antigen containing phagosome promotes the catalytic removal of MHC II-associated invariant chain (Ii) peptide and allows the antigen peptide to bind to MHC II molecules. The degradation of Ii is regulated by the ratio between cathepsin S, a lysosomal cysteine protease, and its inhibitor cystatin C. Upon DC maturation, the level of cystatin C is downregulated, so the activity of cathepsin S increases, which degradates the Ii and leads to the loading of antigen peptide (Pierre & Mellman, 1998). Then the antigen peptide loaded MHC II molecules are transported to the immunological synpase on the cell surface. This procedure has been proven to involve the transformation of the MHC II- antigen peptide vesicle into a tubular structure, which is directed towards the DC-T cell interaction site at the plasma membrane (Vyas, et al., 2007). Tubulation of the vesicles

9 requires loading of antigen peptide onto MHC II, maturation of DC and contact with the antigen specific T cells; and selected proteins including RAB7A, RILP and spinophilin have been identifed to mediate the polarized movement of the compartments to DC surface (Jordens et al., 2001; Boes et al., 2003; Bloom et al., 2008). Surface MHC II molecules are internalized rapidly in immature DC, because the cytoplasmic tail of MHC

II β-chain, which is essential for recycling MHC II molecules from cell surface, is ubiquitinated in mouse immature DCs. DC maturation is associated with a decreased number of ubiquitylated MHC II molecules, which accumulate on the cell surface for several days, allowing optional recognition by CD4+ T cells (Cella et al., 1997; Pierre et al., 1997; Shin et al., 2006). DC-LAMP, a lysosome-associated membrane glycoprotein, which is restricted to the lysosomes of mature DC, may facilitate the translocation of antigen peptide loaded MHC II molecules to the cell surface (de Saint-Vis et al., 1998;

Barois et al., 2002). TLR signalling is involed in the efficient MHC II antigen presentation since it can induce DC maturation (Blander, 2008; Ramachandra, et al.,

2009). Indeed, DCs pulsed with antigens together with LPS, a TLR4 ligand, induce a much more potent T cell proliferation than DCs pulsed with antigen alone (Blander &

Medzhitov, 2006).

To generate specific CD8+ cytotoxic lymphocytes, DCs present antigen peptides on MHC

I molecules, which can be loaded via both endogenous and exogenous pathways.

Cytosolic proteins are degradated through an ATP-dependent proteolytic system initiated by ubiquitin conjugation. DCs constitutively express ubiquitin, and can process endogenous antigens more efficiently. Ubiquitinated proteins are directed to the

10 proteasome, where they are cleaved into peptides. The peptides are translocated from cytoplasm into ER lumen via TAP transmembrane transportors, and further cut into 8-10 mers by the amino peptidase in ER. The final peptides are subsequently associted with

MHC I molecules and β2-microglobulin, and this complex allows the optimal folding and delivery to the cell surface through Golgi apparatus (Cresswell et al., 1999; Purcell &

Elliott, 2008). It has been shown that maturation enhances the synthesis of MHC I molecules and prolongs their half-life (Rescigno et al., 1998). DC maturation also induces the change of proteasome composition and may upregulate the efficiency of presentation of specific epitopes (Morel et al., 2000; Dannull et al., 2007).

In 1976, Bevan discovered that even the antigens derived from an exogenous source could elicit strong cytotoxic T-lymphocyte responses (Bevan, 1976). The phenomenon that APCs can uptake, process and present extracellular antigens on MHC I molecules to

CD8+ T cells has been termed as “cross-presentation” or “cross-priming” (Kurts et al.,

2010). The different pathways of antigen uptake determine the fate of antigen presentation. The antigens internalized by pinocytosis or scavenger receptor mediated endocytosis are presented to CD4+ T cells, while the antigens internalized by mannose receptor mediated endocytosis are committed to the cross-presentation pathway

(Burgdorf et al., 2007). Two models of cross-presentation have been described, cytosolic pathway and vacuolar pathway (Lin et al., 2008). The cytosolic model proposes that the internalized antigens escape from phagosomes into cytosol, and then they are degraded by proteasome and transported into ER lumen where the peptides are loaded onto MHC I molecules. The exogenous antigens leaked from phagosome utilize the conventional

11 MHC I presentation pathway. This process is dependent on proteasome complex and

TAP transporter (Kovacsovics-Bankowski & Rock, 1995; Rodriguez et al., 1999;

Cresswell et al., 2005). This model is supported by the data demonstrating that cross- presentation of DCs is sensitive to brefeldin A, which disrupts the ER transportation

(Fonteneau et al., 2003; Morón et al., 2003). Furthermore, inhibition of lysosomal acidification by a drug, chloroquine, does not down-regulate but enhances cross- presentation, also suggesting that exogenous peptides loaded on MHC I molecules are not generated in lysosomal compartments (Accapezzato et al., 2005; Belizaire & Unanue,

2009). The vacuolar model of cross-presentation is based on the observation that exogenous antigens also can be presented to CD8+ T cells in the TAP-deficient mice

(Schoenberger et al., 1998; Sigal & Rock, 2000; Ruedl et al., 2002). In contrast, S- deficient mice exhibit diminished cross-presentation, suggesting that endosomal proteases, such as cathepsin S are required. Therefore, the extracellular antigens might be degraded to peptides in phagosomes or phago-lysosomes, loaded on MHC I molecules and transferred to the cell surface (Shen et al., 2004; Kurotaki et al., 2007); however, this pathway is still debatable. Several studies have demonstrated that DCs are the most efficient APCs to cross-present antigens in vivo, and thus DCs employ this MHC I presentation machinery for exogenous antigens to induce CTL responses against tumors and virus infection, and also to induce peripheral tolerance to self-antigens. Manipulation of DC cross-presentation may be a strategy to develop vaccines (Heath & Carbone, 2001;

Jung et al., 2002; Smith et al., 2003).

12 In addition to MHC class II and class I molecules, DCs use CD1 molecules to present lipids and glycolipids-containing antigens to T cells. There are five CD1 members

(CD1a-e) expressed by human DCs, but CD1d is the only CD1 molecule that has been found in mouse DCs. Newly synthesized CD1 molecules are located in the ER lumen, and their association with β2-microglobulin is crucial for their trafficking to cell surface.

The surface CD1d molecules can be recycled into lysosomes, where the exogenous lipids can be loaded. The CD1 presentation is important to anti-tumor responses as well as anti- microbial immunity, although much needs to be studied (Porcelli & Modlin, 1999;

Jayawardena-Wolf & Bendelac, 2001; Matsuda & Kronenberg, 2001; Barral & Brenner,

2007).

DCs develop the machineries that are specialized for antigen presentation. Compared to other APCs, like macrophages, DCs have relatively low levels of lysosomal proteases and express some protease inhibitors. These properties enhance the antigen retention and allow DCs to maintain the peptides in MHC I or MHC II complexes for up to 3 days

(Halfon et al., 1998; Pierre & Mellman, 1998; Delamarre et al., 2005; Savina et al., 2006).

This fits perfectly with the migration period of DCs from peripheral tissues to lymphoid tissues (about several days), giving DCs better opportunities to present antigens to T cells

(Itano & Jenkins, 2003; Trombetta et al., 2003). Furthermore, DCs produce 10- to 100- fold higher levels of MHC molecules and MHC-peptide complexes than other APCs

(Inaba et al., 1997). Dynamic imaging of DC-T cell interaction in LNs has revealed a long-lived (>15 h) association with antigen bearing DC and naïve T cells, which

13 facilitates the completion of the “conversation” between these two cell types (Stoll et al.,

2002).

MHC-peptide complexes on DC recognized by the antigen specific T cell receptors on T cells provide the “signal one” for T cell activation, and several adhesion molecules, including CD2, CD18, CD50 and CD54, mediate the contact between DCs and T cells

(Banchereau et al., 2002; Balkow et al., 2010). The binding of co-stimulatory molecules on DCs and their ligands expressed on T cells provides the “signal two.” The cytokines secreted by DCs, such as IL-6, IL-10, IL-12 and IL-15, provide the “signal three” and also induce the different pathways of T cell differentiation (Akbari etal., 2001; Dodge et al., 2003; Martien, 2003; Feili-Hariri et al., 2005; Mortier et al., 2009).

Regulation of B cells and innate leukocytes by DC

Besides naïve T cells, DCs are also able to activate naïve or memory B cells. Through secretion of IL-12 together with IL-6/IL6Rα complex, DCs help the differentiation of naïve B cells to plasma cells. CD40-activated DCs promote the differentiation of CD40- activated memory B cells towards IgG-secreting cells, and DCs induce the surface expression of IgA by B cells in the presence of TGF-β (Fayette et al., 1997; Dubois et al.,

1998; Wykes & Macpherson, 2000). DCs have been found to express BAFF, a member of TNF family, and B cells are known to express several receptors for BAFF. BAFF signaling initiated by DCs can induce the B cell activation, proliferation and immunoglobulin secretion (Craxton et al., 2003; Massacand et al., 2008). Germinal center

(GC) is an environment where B cells undergo proliferation, somatic hypermutation and

14 Ig isotype switching, and GC contain T cells, follicular DCs (FDCs) and germinal center

DCs (GCDCs) other than B cells. It has been shown that GCDCs are different from FDCs in phenotype and function (Liu et al., 1996; van Nierop & de Groot, 2002). GCDCs stimulate the proliferation of CD40-activated B cells and lead their differentiation to plasma cells in an IL-12 dependent manner. In vivo studies have demonstrated that un- processed antigens can be captured by GCDCs and then transferred to naïve B cells to prompt a specific antibody response, thus, GCDCs are described as the “antigen transporting cells” (Grouard et al., 1996; Wykes et al., 1998).

DCs can regulate innate leukocytes via direct cell-cell contact and indirect cytokine mediated interaction. NKT cells can secret abundant IFN-γ after recognizing the α- galactosyloceramide (α-GalGer)-CD1d complex on DCs, and α-GalGer is expressed by some bacteria and tumor cells (Tangri et al., 1996; Fujii et al., 2003). DCs can also regulate the activity of NKT cells through the release of IL-12, IL-15 and IL-18

(Kitamura, et al., 1999; Banchereau, et al., 2000). DCs enhance the antiviral and antitumor activity of NK cells by production of IL-12, IL-18 and IFN-α, and activated

NK cells can, in turn, promote the DC maturation in spleen marginal zones by positive regulatory signals (Pan et al., 2004; Ferlazzo & Münz, 2009; Persson & Chambers, 2010).

DC SUBSETS IN MICE

DCs are a highly heterogeneous leukocyte population, and large numbers of DC subsets have been identified in mice based on their phenotype, function and tissue distribution.

Generally, DCs are divided into two populations: 1) conventional DC (cDC) and 2)

15 plasmacytoid DC (pDC). cDCs contain lymphoid tissue resident DCs and non-lymphoid tissue migratory DCs. They express high levels of DC marker CD11c, and have a dendritic morphology, potent antigen processing and presentation capacities (Lin, et al.,

2008; Merad & Manz, 2009). Mouse pDCs are round, non-denritic, relatively long-lived cells, expressing low levels of CD11c, but high levels of B220, Ly6C and PDCA-1 on cell surface (Ferrero et al., 2002; Zucchini et al., 2008). Importantly, pDCs recognize viral and bacterial nucleic acids via TLR7 and TLR9, and secret large amounts of type I interferon (IFN-α) in response to viral or other microbial infections (Liu, 2005; Szabo &

Dolganiuc, 2008; Cao et al., 2009).

DC subsets in spleen

The spleen is the primary source of lymphoid tissue-resident DCs, and the DC subtypes in the mouse spleen have been well characterized. There are three cDC populations identified based on surface expression of CD4 and CD8 (Table I, Liu, 2001). They are

CD4-/CD8+, CD4-/CD8- and CD4+/CD8-. The majority of DCs in spleen are CD4+/CD8- population mostly located in the marginal area, and CD4-/CD8+ DCs mostly located in the T cell area. The CD4-/CD8+ cDCs express high levels of CD205 but not CD11b or

Sirp-α. On the other hand, the CD4-/CD8- and CD4+/CD8- cDCs lack the surface expression of CD205, but they are CD11b and Sirp-α positive (Wu & Dakic, 2004;

Maldonado-López et al., 1999). CD8+ DCs constitutively cross-present soluble and cell- associated antigens to T cells, while CD8- DC subsets are specialized for MHC II presentation (den Hann & Bevan, 2002). Interestingly, the potent cross-presentation capacity of CD8+ DCs is not due to the antigen uptake, since they show comparable

16 endocytic potential as CD8- DCs. It indicates that CD8+ DCs must employ a unique cross-presentation machinery, which is absent in CD8- DCs (Schnorrer et al., 2006). It has been reported that CD8+ DCs preferentially activate T cells towards Th1 differentiation by producing IFN-γ, whereas CD8- DCs preferentially activate T cells towards Th2 differentiation (Maldonado-López et al., 1999). In addition to cDCs, pDCs are also found in mouse spleen, defined as CD11clow, CD45RA+ and B220+ −they are the major cell type producing IFN-α against viral infection (Asselin-Paturel et al., 2001).

Recently, a novel DC subtype, IKDC, was identified in mouse spleen. These IKDCs display a “hybrid” surface phenotype of NK cells, cDCs and pDCs, including CD11c+,

NK1.1+ and B220+. IKDCs are potent producers of IFN-γ and are cytotoxic to NK target cells. Like pDCs, IKDCs are able to secret IFN-α when activated by CpG, a TLR9 agonist. They are comparable to cDCs in their antigen presenting abilities (Chan et al.,

2006; Taieb, et al., 2006). A TNF-α/iNOS producing DC (Tip-DC) subset has been identified in the spleen of mice infected with Listeria monocytogenes. Tip-DCs are not crucial for T cell priming, but are the predominant source of TNF-α and iNOS during

Listeria monocytogenes infection, and they are required for the clearance of primary infection (Serbina et al., 2003).

17 Table I. cDC subsets in mouse spleen.

Liu, 2001

DC subsets in

Mouse blood DCs have not been well characterized, and only two DC subtypes are identified so far. Cells with the surface phenotype of CD11c+/CD11b+/CD45RA- are immature DCs and they rapidly transform to CD8+ cDCs after TNF-α stimulation. The other subtype with the surface phenotype of CD11clow CD11b– CD45RA+ resembles pDC in their morphology and high ability to produce IFN-α after CpG stimulation (O’Keeffe et al., 2003).

18

DC subsets in skin

Langerhans cells (LCs) are the first described DC subset, uniquely distributed to the epidermis. Besides the expression of CD11c, CD11b, MHC II and CD205, LCs are characterized by high expression of epithelial cell adhesion molecule (Ep-CAM) and langerin (CD207), a member of C-type lectins, which contributes to the formation of cytoplasmic Birbeck’s granules. LCs construct a tight network that covers the entire body surface and serve as the primary sentinels of the skin. Compared to the other DC subsets,

LCs have much longer life span in the steady condition (Kamath et al., 2002; Valladeau et al., 2003; Kaplan, 2010). It has been reported by our lab that the half-life of LCs ranges from 53 to 78 days (Vishwanath et al., 2006). In the quiescent skin, LCs are maintained by the local radio-resistant precursors that self-renew in situ, while after substantial injury,

LCs are lost and subsequently generated by circulating precursors (Merad et al., 2002).

Upon inflammation induced by hapten application, ultraviolet (UV) irradiation or microbial infection, LCs become mature and migrate through lymphatic vessels into the draining LNs where they present antigens to the antigen specific T cells. Therefore, this type of DC is categorized as a “migratory DC” (Johnston et al., 2000; Xu et al., 2001;

McGee et al., 2006). The rate of LCs migration is relatively low, as it takes about 3 days for LCs to enter LNs, possibly because the crossing basal layer of the epidermis delays the migration of LCs (Kissenpfennig et al., 2005). LCs migrate to skin-draining LNs even in the absence of inflammation, and this spontaneous LC migration is considered to play a crucial role in the maintenance of peripheral tolerance (Villadangos & Schnorrer, 2007;

Merad et al. 2008).

19

Previously, it was considered that only langerin-/MHC II+ DCs reside in the dermis.

Although langerin+ DCs were indentified in the dermis, they were regarded as the LCs en route to the skin draining LNs (Henri et al., 2001). Recently, several studies have revealed that a population of LC-independent langerin+ dermal DCs, which are recruited from blood in the steady state, migrate to the draining LNs after antigen uptake. This DC population expresses high level of CD103, but it is negative for CD11b and Ep-CAM, which are highly expressed by epidermal LCs. Therefore, langerin expression is not restricted to LCs. After one-time administration of diphtheria toxin (DT) to langerin-DT receptor-EGFP mice, 50% dermal langerin+ DCs recovered after 7 days, when epidermal

LCs were still absent. Using this model, it has been discovered that contact hypersensitivity (CHS) responses were diminished when both LCs and langerin+ dermal

DCs were depleted. However, CHS responses were not affected in the absence of only

LCs. It indicates that langerin+ dermal DCs are the main mediators for CHS (Bursch et al.,

2007; Ginhoux et al., 2007; Poulin et al., 2007). The langerin+/CD103+ DCs in the dermis resemble CD8+ cDC in the spleen, because both of these subsets show preferential capacity to cross-present antigens to CD8+ T cells. Depletion of the transcription factor

Batf3 knocks out these two DC subsets but not other DC subsets (Hildner et al., 2008;

Bedoui et al., 2009).

DC subsets in other non-lymphoid tissues

In addition to skin, DCs have been identified in various non-lymphoid tissues, such as heart, pancreas, liver, kidney, islet, lung and gut. Two cDC populations are identified in

20 most of the tissues. One is CD103+/CD11blow cDC subset, which resembles the CD8+ DC subset in lymphoid tissues, and the other is CD103-/CD11bhi cDC subset, which resembles the CD8- DC subset in lymphoid tissues. The physiological functions of cDCs in non-lymphoid tissues have been well studied. For example, CD103+ DCs in the lung are required for optimal influenza virus responses. Furthermore, pDCs, which are the major source of INF-α after microbial infection, can also be identified in non-lymphoid tissues (Ginhoux et al., 2009; Merad & Manz, 2009).

DC subsets in LNs

The DC subsets found in the skin-draining LNs are more complicated. In addition to the three phenotypically and functionally equivalent resident cDC populations found in spleen, three additional migratory populations have been identified: CD8lo/CD205hi population corresponding to the LCs that migrate from epidermis; CD8-

/CD205int/CD103- population corresponding to the cDCs that migrate from dermis; and

CD11blow/CD103+ population corresponding to the migratory langerin+ dermal DCs

(Shortman & Liu, 2002; Guilliams et al., 2010). Upon infection of Gram- bacteria, a monocyte derived DC subset, which expresses DC-SIGN and MMR, arises in the skin- drain LNs. This DC subset exhibits a potent capacity to present bacterial antigen to both

CD4+ and CD8+ T cells (Cheong et al., 2010).

The three resident cDC populations also can be identified in mesenteric LNs, and the

CD11b+ DCs exhibited higher capacity to produce IL-10 and induce the Th2 differentiation of T cells. Besides, CD103+ DCs migrate to mesenteric LNs from lamina

21 propria in a CCR7 dependent manner (Shortman & Liu, 2002; Mendlovic & Flisser,

2010).

DC DEVELOPMENT

Myeloid origin of DC

DCs were generally believed to derive only from myeloid origin, based on their similarities to and macrophages. In fact, BM myeloid progenitors can generate DCs, macrophages and granulocytes in the presence of granulocyte-macrophage colony-stimulating factor (GM-CSF). Because these three cell types could develop from a single BM progenitor-derived colony, it was confirmed that they shared a common progenitor (Inaba et al., 1993). Further study revealed that mouse blood monocytes gave rise to DCs in vitro in the presence of GM-CSF and IL-4 (Schreurs et al., 1999). This differentiation process also was observed in vivo, since subcutaneous monocytes give rise to DCs (Randolph et al., 1999). After mouse BM common myeloid progenitors (CMPs), with the phenotype of Lin-/CD34+/c-kit+/CD16/32low/Sca-1-/IL-7Rα-, were transplanted into irradiated recipients, they were able to generate both cDC and pDC. This provided the direct evidence for myeloid origin of DCs. CMPs are heterogeneous in fms-related tyrosin kinase 3 (Flt3) expression, and it has been proven that the Flt3+ fraction is the major progenitor for both DC populations in mouse spleen and thymus (D’Amico & Wu,

2003).

22 Lymphoid origin of DC

The early implication that DCs might be related to a lymphoid lineage came from the observations that typical lymphoid markers, including CD8, CD2, and CD25, were found on some DC subsets in mouse spleen and thymus (Vremec et al., 1992). The first evidence of the lymphoid origin of DC is the research showing that the lymphoid- restricted CD4low/c-kit+/Sca-1+/Sca-2+ precursors, which have no myeloid development potential, give rise to CD8+ DCs together with T, B and NK cells in mouse spleen (Wu et al., 1996). These lymphoid-restricted precursors can also generate DCs in culture without

GM-CSF, the essential cytokine for DC development from myeloid pathway in vitro

(Saunders et al., 1996). Subsequent studies using mouse BM common lymphoid progenitors (CLPs), which are Lin-/IL-7Rα+/c-kitint/Sca-1int/Thy1.1-, have demonstrated that CLPs possess the potential to generate all cDC subsets identified in the spleen as well as pDCs, despite a bias to the development of CD8+ DCs. The Flt3+ fraction in CLPs is the primary source for DC development from the lymphoid pathway. CLPs are more potent than CMPs in DC generation on a single cell basis, because almost 70% of CLPs are Flt3+, while only 30% of CMPs are Flt3+. However, CLPs are less numerous than

CMPs in mouse BM, and, thus the overall contribution of CMP and CLP-derived DCs may be similar (Manz et al., 2001; Wu et al., 2001; D’Amico & Wu, 2003).

DC precursors

Monocytes have been known as DC-committed precursors for a long time. After internalize antigens, about 25% of phagocytic monocytes that reside subcutaneously migrate to the T cell area of draining LNs where they differentiate into DCs, while the

23 other 75% monocytes differentiate into macrophages and remain in the subcutaneous tissues (Randolph et al., 1999). The monocytes circulating in mouse peripheral blood are separated into two subpopulations. One is CX3CR1low/CCR2+/Gr-1+, which is short-lived and actively homing to the inflamed tissues; the other one is CX3CR1hi/CCR2-/Gr-1-, which is relatively long-lived and recruited to non-inflamed tissues (Table II, Geissmann et al., 2003). Accordingly, they were termed as “inflammatory monocytes” and “resident monocytes,” and both of them have the potential to differentiate into DCs under inflammatory or steady condition, respectively (Geissmann et al., 2003). The Gr-1+ monocytes in blood also serve as direct precursors for LCs in vivo because they are recruited to the inflamed dermis and epidermis, where they proliferate and then differentiate into dermal macrophages or epidermal LCs (Ginhoux et al., 2006). A recent study has revealed that monocytes give rise to a DC-SIGN+/MMR+ DC subset in vitro and in vivo after Gram negative bacterial infection. These monocyte-derived DCs co- express TLR4 and CD14, and have a potent antigen presentation capacity (Cheong et al.,

2010).

24 Table II. Phenotype of mouse monocyte subsets

Geissmann et al., 2003

Besides monocytes, other DC precursor populations have been identified in mice. These populations have different phenotypes, are distributed to different tissues and generate different DC subsets as well as some other leukocytes. A homogeneous cell population in

BM and blood termed pre-immunocytes, which is CD11c+/CD32+/Ly6C+, can give rise to both macrophage and cDCs in vitro (Bruno et al., 2001). Lin–/CD8+/CD11c- cells isolated from spleen are capable of differentiation specifically into CD8+ DCs but not CD8- DCs,

T cell, NK cells or the myeloid lineage cells (Wang et al., 2002). Lin–/c-Kit+/CX3CR1+ cells in BM exclusively differentiate into macrophages and cDCs both in vitro and in vivo

25 and thus, these cells were termed “MDP” for macrophage and DC progenitor (Fogg et al.,

2006). A cell population called common DC precursor (CDP), which is Lin-/c-

KitLow/CD115+ (M-CSFR+)/Flt3+, can generate both cDCs and pDCs (Onai et al., 2007).

Ken Shortman and his colleagues proposed a model of DC development based on two

DC-committed precursor cells isolated from BM culture with Flt3 ligand (Flt3L). In this model, a dividing DC progenitor population, “pro-DC,” gives rise to a transitional non- dividing DC precursor population, “pre-DC,” which is en route to differentiating into the three distinct DC subtypes, pDCs, CD8+ cDCs and CD8- cDCs (Naik et al., 2007). A DC- committed precursor population, which is CD11c+/MHC II-/ B220+/CD43+/Gr-1-/F4/80-, has been found in peripheral blood. They have the capacity to generate cDCs and pDCs but not the other cells in vivo (del Hoyo et al., 2002). The CD11cint/MHC II-

/CD45RAlow/CD43int/SIRP-αint/CD4-/CD8- spleen cells, which comprise about 0.05% of , only serve as precursors for cDCs in vivo in the steady-state. This population has been termed pre-cDC (Naik et al., 2006; Liu et al., 2009). The pre-cDCs migrate from blood not only to lymphoid tissues, but also to non-lymphoid tissues, where they differentiate into CD103+/CD11blow as well as CD103-/CD11b+ cDCs (Ginhoux et al.,

2009).

Some DC subsets can turn into other DC subsets under certain conditions. For example, pDCs in the BM lose expression of B220 and Ly6C, but acquire the surface expression of

CD11b in the mice infected with lymphocytic choriomeningitis virus. Moreover, pDCs show a traditional dendritic morphology and enhance the antigen presenting function under viral infection, suggesting that pDCs have differentiated into cDCs (Zuniga et al.,

26 2004). Some experimental data indicate that CD8- DCs develop into CD8+ DCs in the spleen and those CD8- DCs up-regulate the CD8 expression in vitro when cultured with

Flt3L and activated by LPS (Ardavín, 2003).

Interestingly, granulocyte-committed cells can be reprogrammed to acquire DC characters in vitro. The immediate precursors of polymorphonuclear granulocytes

(PMNp), including band cells, and , were purified from peripheral blood of patients with chronic myeloid leukemia, bacterial infections or treated with granulocyte colony-stimulating factor (G-CSF), and then were cultured in the presence of GM-CSF, IL-4 and TNF-α for 9 days. The PMNp-derived cells showed a typical DC morphology, acquired surface expression of MHC II, CD1 molecules and co- stimulatory molecules, and exhibited a potent antigen presentation capacity, suggesting that PMNp cells were able to develop into DCs or DC-like cells (Oehler et al., 1998).

Cytokines in DC development

GM-CSF was the first cytokine reported to efficiently support DC differentiation in vitro, and GM-CSF can induce abundant CD11b+ DCs when added into BM culture or monocyte culture. Although commonly used for research, the DCs generated in the presence of GM-CSF are different from the DCs isolated in steady state in vivo. In the steady state, almost no GM-CSF can be detected in the peripheral blood (Inaba et al.,

1992; Xu et al., 2007; Schmid et al., 2010). In the GM-CSF deficient mice or GM-CSFR

β chain knockout mice, there is only a slight decrease in cDC numbers in the spleen and

LNs compared to the numbers found in WT mice (Vremec et al., 1997). Only a small

27 increase in cDC number is observed in the transgenic mice over-expressing GM-CSF.

Therefore, it is believed that GM-CSF-generated DCs are more like inflammatory DCs rather than DCs under steady state (Shortman & Naik, 2007).

Flt3L is a critical cytokine for DC development in the steady state. Mice with the deleted

Flt3L or Flt3 exhibit a dramatic reduction in cDCs and pDCs, and similar DC loss has been observed in the mice that are deficient in STAT3, an important transducer in Flt3L signaling pathway. Administration of Flt3L to mice or over-expression of Flt3L results in significantly increased numbers of DCs and myeloid cells, but not T or B cells.

Furthermore, both cDCs and pDCs can be generated in BM cultures supplemented with

Flt3L alone (Maraskovsky et al., 1996; Laouar et al., 2003; McKenna et al., 2000;

Waskow et al., 2008).

Macrophage colony-stimulating factor (M-CSF) has also been shown to promote DC development. The mice lacking M-CSF show a two- to three-fold reduction in the DC number in spleen compared to WT mice. The M-CSF receptor deficient mice lack both monocytes and LCs (Ginhoux et al., 2006; Schmid et al., 2010). In addition, both cDCs and pDCs can be generated in the BM culture in the presence of M-CSF alone.

Administration of M-CSF to Flt3L deficient mice increases the number of cDCs and pDCs, suggesting that DC differentiation induced by M-CSF is Flt3-independent (Fancke et al., 2008).

28 Our lab has reported recently that IL-33, a member of IL-1 superfamily, promotes the development of CD11b+ DCs in the BM cultures. Based on the phenotype, the DCs generated with IL-33 are similar to the DCs generated with GM-CSF except for their moderate expression of MHC II. The data demonstrate that IL-33 may have the indirect effects on DC development by triggering GM-CSF produced by the and their precursors in the BM cultures (Mayuzumi et al., 2009).

pIL1-DsRed TRANSGENIC MICE

IL-1β is a prototype cytokine with multiple functions, and it plays a major role in acute and chronic inflammation. IL-1β has been shown to mediate the production of a broad range of inflammatory molecules, including leukotrienes, prostaglandin, - activating factor and pro-inflammatory cytokines. IL-1β also affects cell proliferation, leukocyte recruitment, tissue damage, blood pressure, and many other functions. Many diseases are related to IL-1β, such as Alzheimer’s disease, diabetes, rheumatoid arthritis

(RA), periodontitis and acute myeloid leukemia. Macrophages, monocytes and are the primary source of IL-1β, and DCs, epithelia cells, fibroblasts and keratinocytes also have the potential to produce this cytokine (Seymour & Gemmell,

2001; Delaleu & Bickel, 2004; O’Neill, 2008).

It has been reported by our lab that IL-1β promoter in DCs is rapidly activated by a variety of chemical and biological agents, including TLR agonists, purinergic type 2 receptor ligands and necrotic cells (Mizumoto et al., 2005). A 4.1 kb 5’-flanking fragment isolated from the murine IL-1β gene was used to drive the expression of the

29 yellow fluorescence protein gene in a stably transduced DC clone. By analyzing the yellow fluorescence, this DC biosensor system has allowed us to test large numbers of natural and synthetic compounds for their potential to induce DC activation (Tanaka et al.,

2009).

Recently, we constructed a transgenic mouse line with the DsRed fluorescent protein gene expression under the control of the same IL-1β promoter (Figure 1). This mouse line enables us to visualize the IL-1β promoter activation in living animals by using advanced intravital optical imaging technology (Matsushima et al., 2010).

Figure 1. The construction of the pIL1-DsRed mouse line. To generate a red fluorescent protein- expressing vector, a PCR fragment was amplified from pDsRed-Express-DR plasmid, ligated into a TA- cloning vector, and then subcloned into pBK-CMV-SG. pBK-CMV-SG-Red, carried a CMV immediate early promoter upstream of the β-globin intron/exon-RFP fusion gene. The CMV promoter region was removed by digestion with VspI and NheI, followed by blunting of both ends with Klenow fragment and self-ligation. The 4,138 bp BamHI fragment of the murine IL-1β promoter was inserted into the BamHI site to generate the plasmid pBK-SG-IL-1b-Red. The plasmid pBK-SG-IL-1-Red was digested with SalI and NotI. The transgene fragment was purified and microinjected into fertilized eggs of C57BL/6 mice.

30

As shown in Figure 2a, very few cells expressing DsRed were detected in the ear skin of the transgenic mice under the microscope in the steady state. After application of oxazolone (OX), a skin sensitizer used to induce inflammation, the ear skin samples were harvested and examined under the fluorescence microscope. The number of DsRed+ cells in the skin increased dramatically, reaching the maximum level at 48 h, and the DsRed signal then declined sharply at 72 h. The DsRed signals also can be visualized in the OX- painted ears of living animals by confocal microscope and the distribution of DsRed+ cells can be analyzed by the z-axis scanning (Matsushima et al., 2010).

a

b

31 Figure 2. Emergence of DsRed+ cells in the skin under inflammatory condition. (a) Ear skin samples of pIL1-DsRed transgenic mice were harvested and examined under the macro-zoom fluorescence microscope at the indicated time points after OX application. (b) At 24 h after OX treatment, pIL1-DsRed transgenic mice were anesthetized and examined under a confocal microscope. Data shown are compiled x–y plane images of DsRed+ cells in the indicated 5 mm z-axis depth range from the skin surface (Matsushima et al., 2010).

To determine the surface phenotype of the cells expressing DsRed fluorescent signals, ear skin samples were harvested at different time points after OX painting, and epidermal layer was separated from dermal layer. The single cell suspensions from the epidermis were prepared and analyzed by flow cytometry. Consistent with the imaging data, the number of DsRed+ cells increased in a time-dependent manner, with a sharp decline at 72 h (Figure 3a). Majority of the DsRed+ cells expressed CD45, a common leukocyte marker.

The number of CD45+ cells increased after OX application, indicating the immigration of inflammatory leukocytes into epidermis (Figure 3b). Almost all the DsRed+ cells expressed CD11b, suggesting they are of myeloid lineage. MHC II and F4/80 molecules were only expressed by a small fraction of DsRed+ cells, whereas more than 75% of the

DsRed+/CD45+ cells expressed high level of Gr-1 (Figure 3c). Those leukocytes expressing DsRed signals are likely to include granulocytes, inflammatory monocytes and myeloid suppressor cells (Matsushima et al., 2010).

32

Figure 3. Surface phenotype of DsRed+ cells emerging in the epidermis after inflammation. (a) Single cell suspensions from epidermal layers were prepared from the ear skin of pIL1-DsRed transgenic mice at the indicated time points after topical application of OX and examined for DsRed expression. (b) Epidermal cell suspensions from WT mice or pIL1-DsRed transgenic mice were also stained with anti- CD45 mAb or isotype-matched control IgG and then examined for expression of CD45 (y axis) and DsRed (x axis). (c) The CD45+ populations in the above experiments were examined for the expression of the indicated surface markers (y axis) and DsRed (x axis) (Matsushima et al., 2010).

33 MATERIALS AND METHODS

Mice

C57 BL/6 mice (CD45.2+), B6 SJL (CD45.1+) mice, OT II and OT I transgenic mice were purchased from Jackson Laboratory. The pIL1-DsRed transgenic mice were constructed in our lab as described earlier. The transgene expression was confirmed by genomic PCR for DNA isolated from tail tissues. Transgene-positive mice were bred with WT C57BL/6 mice and heterozygous offspring were used in the studies. The mice were maintained and handled according to institutional guidelines.

Cell culture and purification

Cells in the femurs and tibiae were flushed from WT and pIL1-DsRed mice, and the red blood cells were depleted by the lysis buffer (Sigma-Aldrich, St Louis,

MO). The BM cells were cultured in 6-well plates (2×106 cells/ml; 3 ml/well) with complete RPMI supplemented with 10 ng/ml GM-CSF (PeproTech, Rocky Hill, NJ). The complete RPMI medium consisted of RMPI 1640 medium (Hyclone, Logan, UT) supplemented with 10% fetal bovine serum (Sigma-Aldrich, St. Louis, MO), 2 mM L- glutamine (Sigma-Aldrich, St. Louis, MO), 1 mM sodium pyruvate (Sigma-Aldrich, St.

Louis, MO), 50 μM 2-β-mercaptoethanol (Invitrogen, Carlsbad, CA) and antibiotics mixture (100 U/ml penicillin, 100 μg/ml streptomycin and 0.25 μg/ml amphotericin B,

Sigma-Aldrich, St Louis, MO). For both cDCs and grDCs, the medium was changed on day 2 and day 4, when floating cells were returned back to the wells with fresh media.

34 For BM-derived DC (BMDC), the floating cells of day 2 and day 4 were discarded and the fresh media was added back only to adherent cells according to Inaba’s protocol

(Inaba et al., 1992). On day 6, the phenotype of CD11c+ DC populations from the BM cultures was determined. The fraction of grDC, which is CD11c+/MHC II+/Ly6G+, was purified by FACS sorting, and the cDC fraction which is CD11c+/MHC II+/Ly6G-, was also purified simultaneously. Band cells from BM cultures of pIL1-DsRed mice were purified on day 2 by FACS sorting based on the phenotype of DsRed+/CD11b+/CD11c-, and then were kept in culture in the presence of 10 ng/ml GM-CSF for another 3 days, allowing differentiation into DCs. The CD48-/Gr-1hi band cell population of WT mice was isolated from the crude WT BM cells by FACS sorting and cultured with complete

RPMI supplemented with 20ng/ml G-CSF (PeproTech, Rocky Hill, NJ) for 4 days to confirm their differentiation into granulocytes. The CD11b+/Ly6G-/CD11c- monocyte population was also isolated from crude BM cells of WT mice and cultured in the presence of 10 ng/ml GM-CSF to obtain monocyte-derived DCs (moDCs). For establishing the co-culture system, band cells purified from C57BL/6 mice (CD45.2+) were cultured together with the crude BM cells from B6 SJL mice (CD45.1+), which served as “feeder” cells, in the presence of GM-CSF. After 6 days, CD45.2+/CD45.1- cells derived from band cells were purified for subsequent analyses.

Flow cytometry and cell sorting

Multiparameter analyses and purification of stained cell suspensions were performed on

FACSCalibur or FACSAria (BD, Franklin Lakes, NJ). The results were analyzed by

35 CellQuest or FlowJo. The fluorochrome- or biotin-conjugated monoclonal antibodies used for FACS are listed in the table below.

Table III. Antibodies used for FACS

Antigen Clone Isotype Vendor

CD11c HL3 Hamster IgG1 BD

Ly6G 1A8 Rat IgG2a BD, BioLegend

MHC II AF6-120.1 Mouse IgG2a BD

CD205 NLDC-145, 205yekta Rat IgG2a BMA, eBioscience

CD11b M1/70 Rat IgG2b BD

CD40 3/23 Rat IgG2a BD

CD80 16-10A1 Rat IgG2 BD

CD86 GL1 Rat IgG2a BD

CD1d 1B1 Rat IgG2a BD

CD3 145-2C11 Hamster IgG1 BD

CD4 GK1.5 Rat IgG2a BD

CD8 53-6.7 Rat IgG2a BD

CD11a 2D7 Rat IgG2a BD

CD14 rmC5-3 Rat IgG1 BD

CD18 C71/16 Rat IgG2a BD

CD19 1D3 Rat IgG2a BD

CD23 B3B4 Rat IgG2a BD

CD24 M1/69 Rat IgG2b BD

CD25 7D4 Rat IgM BD

CD31 390 Rat IgG2a BD

CD34 RAM34 Rat IgG2a BD

36 CD38 90 Rat IgG2a BioLegend

CD43 S7 Rat IgG2a BD

CD45 33-F11 Rat IgG2b BD

CD45.1 A20 Mouse IgG2a BD

CD45.2 104 Mouse IgG2a BD

CD45RA 14.8 Rat IgG2b BD

CD48 HM48-1 Hamster IgG1 BD

CD49a H129.19 Rat IgG2a BD

CD49b DX5 Rat IgM BD

CD49d R1-2 Rat IgG2b BioLegend

CD54 3E2 Hamster IgG1 BD

CD62L MEL-14 Rat IgG2a BD

CD69 H1.2F3 Hamster IgG1 BD

CD74 In-1 Rat IgG2b BD

CD93 AA4.1 Rat IgG2b eBioscience

CD103 M290 Rat IgG2 BD

CD106 429 Rat IgG2a BD

CD115 AFS98 Rat IgG2a eBioscience

CD117 2B8 Rat IgG2b BD

CD135 A2F10.1 Rat IgG2a BD

CD150 TC15-12F12.2 Rat IgG2a BioLegend

CD153 RM153 Rat IgG2b BD

CD157 BP-3 Rat IgG2b BD

CD172a P84 Rat IgG1 BD

CD184 2B11 Rat IgG2b BD

CD192 E68 Rabbit IgG Abcam

CD193 83103 Rat IgG2a BD

37 CD195 C34-3448 Rat IgG2c BD

CD196 29-2L17 Hamster IgG BD

CD197 4B12 Rat IgG2a BioLegend

CD207 eBioRMUL.2 Rat IgG2a eBioscience

CD209 LWC06 Rat IgG2a eBioscience

CD275 HK5.3 Rat IgG2a BD

CD281 TR23 Rat IgG2a eBioscience

CD282 6C2 Rat IgG2b eBioscience

CD284 MTS510 Rat IgG2a eBioscience

CD317 eBio927 Rat IgG2b eBioscience

CD326 G8.8 Rat IgG2a BioLegend

33D1 33D1 Rat IgG2b BioLegend

B220 RA3-6B2 Rat IgG2a BD

F4/80 BM8, CI:A3-1 Rat IgG2a, Rat IgG2b BioLegend

Gr-1 RB6-8C5 Rat IgG2b BD

Ly6C AL-21 Rat IgM BD

Mac-2 M3/38 Rat IgG2a BioLegend

Mac-3 M3/84 Rat IgG1 BioLegend

MD-1 MD14 Rat IgG2a BD

MDL-1 226402 Rat IgG2a R&D Systems

MHC I AF6-88.5 Mouse IgG2a BD

Sca-1 D7 Rat IgG2a BD

Ter119 TER-119 Rat IgG2b BD

38 Morphological analysis

Band cells from BM cultures of pIL1-DsRed mice were purified on day 2 as described earlier. These samples were embedded and processed to ultrathin section for electron microscopy (EM) analysis. The morphology of the band cells isolated from BM cultures was also examined under the confocal microscope after fixation, permeabilization and staining with Alexa Fluo 488-phalloidin and DAPI. Band cells purified from crude BM cells or peritoneal exudate cells (PECs) were cytospun and stained by using HEMA 3 kit

(Fisher, Kalamazoo, MI), a modified Wright-Giemsa staining, and examined under a inverted light microscope. Mature granulocytes and grDCs were also stained with HEMA

3 kit for morphological analysis.

Genechip analysis

The cDC and grDC subsets were purified from BM cultures as earlier described. Total

RNA was extracted by RNeasy Plus kit (Qiagen, Valencia, CA) and genechip analysis was performed with Affymetrix Mouse Genome 2.0 Array. The data was analyzed by

GeneSifter and TreeView. The results were based on three independent genechip analyses comparing grDCs versus cDCs. The genes that are expressed predominantly in one population in terms of the magnitude of expression (>2-fold) and the statistical significance (n=3, P<0.05) are shown in the heat map.

Real-time RT-PCR

Total RNA from cDC and grDC subsets from BM cultured was extracted, and 25 ng

RNA was applied to generate the first-strand cDNA by reverse transcription using

39 Superscript III (Invitrogen, Carlsbad, CA). Real-time PCR was performed in LightCycler system (Roche, Indianapolis, IN) using SYBR Green (Molecular Probes, Eugene, OR), master mixture (Idaho Technology, Salt Lake City, UT) and Platinum Taq DNA

Polymerase (Invitrogen, Carlsbad, CA). The expression of β-actin was used as house keeping gene control. The primer pairs were as follows: CRAMP forward,

AAGGAGACTGTATGTGGC-AAGGCA; CRAMP reverse, TTTCTTGAACCGAA-

AGGGCTGTGC; MMP9 forward, TGAACAAGGTGGACCATGAGGTGA; MMP9 reverse, TAGAGACTTGCACTGCACGGTTGA; β-actin forward, TGTGATGGT-

GGGAATGGGTCAGAA; β-actin reverse, TGTGGTGCCAGATCTTCTCCATGT.

Western blotting

The protein was extracted from cDCs and grDCs from BM culture by RIPA buffer

(Pierce, Rockford, IL), and protein concentration was measured by BCA Protein Assay kit (Thermo, Rockford, IL). Equal amounts of protein from the two DC subsets were loaded for SDS-polyacrylamide gel electrophoresis (Bio-Rad, Hercules, CA). After transferring to a nitrocellulose membrane, blots were probed with rabbit anti-mouse

CRAMP primary antibody (provided by Dr. Richard Gallo, University of California, San

Diego) followed by horseradish peroxidase-conjugated goat anti-rabbit IgG secondary antibody (Bio-Rad, Hercules, CA). The membranes were visualized after being developed with SuperSignal West Pico kit (Thermo, Rockford, IL).

40 Immunofluorescence analysis

The cDCs and grDCs purified from BM cultures were cytospun, fixed with 4% PFA and permeabilized with 0.1% Triton X-100. The cells were then incubated with rabbit anti-

CRAMP primary antibody (Abcam, Cambridge, MA) or rabbit IgG as control. After washing to remove the unbound antibody, the cells were incubated with Alexa Fluor 488- conjugated goat anti–rabbit IgG antibody (Invitrogen, Carlsbad, CA). Finally, the cells were washed and stained with DAPI. The samples were examined under FSX100 microscope (Olympus, Japan).

ELISA

The cDC and grDC populations purified from BM cultures were incubated in 24-well plates at a density of 1×106 cells/ml in complete RPMI for 24 h. The supernatants were measured for MMP-9 by ELISA (R&D Systems) according to the instructions provided by manufacturer.

Bacterial killing assays

For testing the intracellular bacterial killing capacity, cDCs and grDCs were purified from BM cultures of WT mice as well as CRAMP KO mice (provided by Dr. Richard

Gallo, University of California, San Diego) (Nizet et al., 2001). 5x104 DCs were incubated with living E.coli K-12 (MOI=10) at 37ºC for 1 h in the complete RMPI media without antibiotics supplemented with 1% human plasma. The extracellular bacteria were killed by 30 min incubation with kanamycine (50 μg/ml, Invitrogen, Carlsbad, CA). After extensive washing, DCs were kept in the RPMI media, and lysed by saponin (1%) and

41 sonication at indicated time points. The cell extracts containing intracellular E.coli were diluted and cultured on LB agar overnight. The number of bacterial colonies was counted and colony-forming units (CFU) were calculated. The extracts of cDCs and grDCs were also used to test for their bacterial killing capacity. Cell extracts (50 μg/ml) or synthetic

CRAMP peptide (8 μM) were added in the E.coli cultures supplemented with 10% LB.

Blocking antibody against CRAMP (200 μg/ml) (Abcam, Cambridge, MA) as well as control IgG (Jackson ImmunoResearch Laboratories, West Grove, PA) were added to the bacterial cultures. After incubation at 37ºC for 4 h, the bacterial numbers were counted as above.

Identification of grDCs in spleen

Total splenocytes were harvested by mechanically dissociation of the spleen. After lysing red blood cells, the samples were labled with anti-CD11b magnetic particles, and BD

IMag system (BD, Franklin Lakes, NJ) was used to purify the CD11b+ fraction, which was subsequently stained with anti-CD11c, anti-MHC II and anti-Ly6G antibodies for

FACS analysis.

Experimental peritonitis models

As reported earlier (Segal et al., 2002), 3% thioglycollate was i.p. injected in WT or pIL1-DsRed mice to induce acute inflammation in peritoneal cavity. At different time points mice were sacrificed and PECs were harvested by washing the peritoneal cavity with 7 ml PBS. The PECs were stained with antibodies to identify or purify different leukocyte populations by FACS. The morphologies of band cells and grDCs were

42 examined under an inverted light microscope after HEMA 3 staining. In some experiments, living E.coli K-12 (1×106-5×106/mouse) were i.p. injected to WT mice to induce peritonitis.

Endocytosis assays

Band cells, cDCs and grDCs were purified from BM cultures as described before and the cell densities were adjusted to 4×105-1×106 cells/ml. Then cells were incubated with

FITC-dextran (MW 70,000; 10mg/ml) or fluorescent latex beads (diameter = 1 μm;

10×106 beads/ml) (Ploysciences, Warrington, PA) for 30 min at 4ºC or 37ºC. After removing unbound dextran or beads by washing, the cells were analyzed by FACS. The endocytotic potential was expressed as the mean fluorescence intensity (MFI) of the internalized probes. For in vivo bacterial uptake assay, thioglycollate was injected to induce peritonitis 48 h before i.p. injection of 1x106 E.coli K-12 TOP 10 constitutively expressing GFP (provided by Dr. Robert Blumenthal, University of Toledo). After 1 h,

PECs were harvested and washed to remove extracellular bacteria. The MFI of green signal was analyzed between cDC (CD11c+/Ly6G-) and grDC (CD11c+/Ly6G+) population by FACS to compare the internalization of E.coli between these two DC subsets.

APC function assays

Band cells and DsRed+ DCs (DsRed+/CD11b+/CD11c+) were purified from BM cultures of pIL1-DsRed mice on day 2, and grDCs were purified from BM cultures of WT mice on day 6. Cell density was adjusted to 1×106 cells/ml in complete RPMI and pulsed for 1

43 h with ovalbumin (OVA)-derived peptides, OVA257–264 or OVA323–339 (10 μg/ml). The

OVA specific CD8 or CD4 T cells were isolated from the spleen of OT I (Hogquist et al.,

1994) or OT II (Barnden et al., 1998) transgenic mice, respectively, using Dynal isolation kit (Invitrogen, Carlsbad, CA), and were plated in 96-well plates (5×104 T cells/well).

The peptide-pulsed band cells or DCs were added to the well at different numbers and co- cultured with T cells for 3 days. After labeling with 3H-thymidine in the last 18 h, cells were harvested with Micro96 Harvester (Molecular Devises, Sunnyvale, CA) and counted for radioactivity with a scintillation counter. For in vivo experiments, thioglycollate was i.p. injected to induce peritonitis and 48 h later, 1×106 E.coli expressing full length OVA cDNA (provided by Dr. Robert Blumenthal, University of

Toledo) or WT E.coli were i.p. injected. PECs were collected 1 h after E.coli injection and cDCs (CD11c+/Ly6G-) as well as grDCs (CD11c+/Ly6G+) were purified by FCAS sorting. The DCs were added to OT II CD4 T cell cultures and the APC function was tested in the same way described before.

Adoptive transfer experiments

B6 SJL recipient mice (CD45.1+) were injected with thioglycollate to induce peritoneal inflammation, and 6 h later, band cells (5×105-2×106 cells) purified from crude BM of

C57BL/6 mice (CD45.2+) were transferred into the recipients by i.v. injection. After 42 h,

PECs were harvested and the CD45.2+/CD45.1- population derived from the transferred band cells was analyzed for their CD11c and MHC II expression by FACS.

44 Statistical analysis

All the experiments have been independently performed at least three times and representative results are shown. The data were assessed by Student’s t-test, and two- tailed P-values were used to determine statistical significance.

45 RESULTS

DsRed+ cells emerge in BM cultures of pIL1-DsRed mice

The newly constructed TG mouse line, which expresses DsRed protein gene under the control of IL-1β promoter, has provided us with a powerful tool to directly observe the

IL-1β-producing cells in vivo. Indeed, a large number of DsRed+ cells emerged in the inflamed skin, and majority of the DsRed+ cells expressed the myeloid marker CD11b and Gr-1 (Matsushima et al., 2010).

Based on these observations, we sought to test whether DsRed signal can be detected in the BM cells from the TG mice. Therefore, we isolated the BM cells from pIL1-DsRed

TG mice and cultured them in the presence of GM-CSF, a prototypic myeloid cell growth factor. Very few DsRed+ cells were detected in the BM cells before culture, and the

DsRed+ cells uniformly expressed CD11b, suggesting these cells were of myeloid lineage

(Figure 4a). Many DsRed+ cells became detectable after 2 day, and more than 90% of the

DsRed+ cells were CD11b positive. Although most DsRed+ cells were CD11c negative, we observed a small number of DsRed+/CD11c+ cells (Figure 4b). The DsRed signal cannot be detected in the BM culture of the WT mice, which served as control.

46 a b

c

Figure 4. Emergence of DsRed+ cells in BM cultures. BM cells isolated from pIL1-DsRed TG mice or WT C57BL/6 mice were cultured for 2 days in the presence of GM-CSF (10 ng/ml) and then examined for the expression of DsRed as well as CD11b (top panels) and CD11c (bottom panels). The phenotype was compared between day 0 (a) and day 2 (b). (c) The numbers of DsRed+/CD11b+/CD11c- population (●) and CD11b+/CD11c+ DC population (○) were counted at different time points in the BM culture. The numbers were normalized to 1×106 total cells. Data shown are the means ± SD from three independent cultures.

47 The time kinetics analysis revealed that very few BM cells expressed DsRed at time 0.

When cultured in the presence of GM-CSF, a large number of DsRed+/CD11b+/CD11c- cells emerged express DsRed within 24 h. The number of DsRed+/CD11b+/CD11c- cells reached to a peak level on day 2 and declined thereafter. The CD11b+/CD11c+ DCs in the culture showed a delayed kinetics with a peak on day 5 or day 6 (Figure 4c). We interpreted these observations to suggest that the DsRed+/CD11b+/CD11c- population might give rise to DCs.

DsRed+/CD11b+/CD11c- cells differentiate into DCs

To test this possibility, we purified the DsRed+/CD11b+/CD11c- population from the BM cultures on day 2, and cultured them in the continuous presence of GM-CSF for 3 additional days. This population uniformly acquired the surface expression of CD11c. In addition, the DsRed expression was maintained during the differentiation of

DsRed+/CD11b+/CD11c- cells (Figure 5).

48 Figure 5. The differentiation of DsRed+/CD11b+/CD11c- into DCs. BM cells isolated from pIL1-DsRed TG mice were cultured for 2 days in the presence of GM-CSF (10 ng/ml) and the DsRed+/CD11b+/CD11c- population was purified and examined by FACS (blue dots). After an additional culture of 3 days in the presence of GM-CSF, the expression of DsRed and CD11c was examined (red dots).

Morphology and surface phenotype of DsRed+/CD11b+/CD11c- cells

Indeed, the DsRed+/CD11b+/CD11c- cells can serve as precursors of DCs, and spontaneously we wondered what was identity of these cells. To answer this question, we first examined the morphology of this population. We observed the cells, which were purified from BM cultures on day 2, under both transmission electron microscope and confocal microscope after staining the cells with phalloidin for actin filaments and DAPI for nucleus (Figure 6a and b). Both EM and fluorescence images demonstrated that the

DsRed+/CD11b+/CD11c- cells are highly homogeneous population with a small cell size and not fully segmented nucleus and contain a low to moderate number of granules. The morphological features suggested its granulocyte lineage.

a b

49 c

MHC II Gr-1 Ly6C

Ly6G CD48 CD62L

Figure 6. The morphology and surface phenotype of DsRed+/CD11b+/CD11c- population. The DsRed+/CD11b+/CD11c- population was sorted by FACS from BM cultures of pIL1-DsRed TG mice. The purified population was either processed for EM imaging (Bar = 10μm) (a) or stained with Alexa Fluo 488- phalloidin and DAPI for confocal imaging (Bar = 20μm) (b). (c) Data shown are the expression profile of indicated surface markers (red histogram) or the isotype-matched control antibody (blue histogram) within the DsRed+/CD11b+/CD11c- population.

We also examined the surface phenotype of this population by testing the expression of various surface markers (Table IV). This DsRed+/CD11b+/CD11c- population was MHC

II-/Gr-1+/Ly6C+/Ly6G+/CD48-/CD62L+ (Figure 6c). The negative expression of CD11c and MHC II confirmed that this population differed from DCs. The high expression of

CD11b, Gr-1, and especially Ly6G was consistent with the phenotype of granulocytes.

50 Table IV. The surface phenotype of DsRed+/CD11b+/CD11c- population.

CD1d (–) CD45RA (–) CD196 (–) CD4 (–) CD48 (–) CD197 (–) CD8 (–) CD49a (–) CD205 (–) CD11a (+) CD49b (–) CD209a (–) CD11b (+) CD62L (+) CD275 (–) CD11c (–) CD80 (–) CD281 (–) CD14 (–) CD86 (–) Gr-1 (+) CD19 (–) CD103 (–) Ly6C (+) CD24 (+) CD117 (–) Ly6G (+) CD25 (–) CD135 (–) F4/80 (–) CD31 (–) CD172a (+) Ly86 (–) CD34 (–) CD184 (–) PDCA-1 (–) CD40 (–) CD185 (–) Sca-1 (–/+) CD43 (+) CD192 (–/+) MHC I (–) CD45 (+) CD193 (–) MHC II (–) CD45R (–) CD195 (–)

−: negative +: positive −/+: low to negative

Endocytic potential and APC function of DsRed+/CD11b+/CD11c- cells

Immature and mature granulocytes have been well known as potent phagocytes in that they are able to internalize large amount of extracellular molecules rapidly (Segal 2005).

The endocytic potential of the DsRed+/CD11b+/CD11c- population was tested by FTIC- dextran uptake assay. The results demonstrated that the population had robust potential to uptake FITC-dextran at 37ºC as determined by strong FITC fluorescence (Figure 7a).

51 a

b

Figure 7. The endocytic potential and APC function of DsRed+/CD11b+/CD11c- population. (a) The DsRed+/CD11b+/CD11c- population was sorted by FACS from BM cultures of pIL1-DsRed TG mice. The purified cells were incubated with FITC-dextran at 4ºC (blue histogram) or 37ºC (red histogram) for 30 min and analyzed by FACS for the MFI of internalized dextran. (b) DsRed+ band cells and DsRed+ DCs were FACS-purified from BM culture on day 3, pulsed with OVA257–264 or OVA323–339 peptide (10 μg/ml), and

52 washed extensively before addition to T cell cultures. CD4 T cells purified from OT-II TG mice or CD8 T cells purified from OT-II TG mice were plated in 96-well plates (5×104 T cells/well). Peptide-pulsed band cells or DCs were added to T cell cultures at the indicated cell numbers and incubated for 3 days. Data shown were the mean ± SD (n=3) of 3H-thymidine uptake by T cells stimulated with OVA peptide-pulsed cells (●) or the base line levels of 3H-thymidine uptake by T cells in the absence of OVA peptide pulsing (○).

Based on the characteristic morphology, surface phenotype as well as potent endocytic capacity, we concluded that the DsRed+/CD11b+/CD11c- population indentified from the

BM cultures of pIL1-DsRed TG mice represents the band cell population.

We also compared the APC function between these DsRed+ band cells versus DsRed+

DCs. Band cells and DCs were purified from the BM cultures on day 3, subsequently pulsed with OVA-derived peptides and co-cultured with OVA specific CD4 T cells or

CD8 T cells collected from OT II or OT I TG mice, respectively (Figure 7 b). As expected, peptide-pulsed DCs efficiently induced the proliferation of both CD4 and CD8

T cells. In contrast, the peptide-pulse band cells failed to activate either CD4 or CD8 T cells, probably due to their low or negative expression of surface MHC I and MHC II molecules.

Band cells isolated from WT mice

Based on the data from experiments with pIL1-DsRed TG mice, we hypothesized that the

DsRed+/CD11b+/CD11c- population representing the immature granulocytes, band cells, was able to differentiate into DCs. To test our hypothesis, we sought to purify band cells from WT mice without using DsRed as a marker. According to the phenotype of the band cells we had known (Table IV, Figure 6c), we isolated the Gr-1high/CD48- population

53 from the fresh BM cells of C57BL/6 mice, and analyzed their morphology and surface phenotype. This population was highly homogeneous and showed a unique morphology

(Figure 8a). Especially, they had a characterized ring-shaped nucleus, which was reported to be the typical morphology of the mouse band cells (Santiago et al., 2001, 2010). This population was also characterized with small cell size, low granularity, and a uniform surface phenotype of MHC II-/CD11b+/CD11c-/Ly6C+/Ly6G+ (Figure 8b). The morphology as well as the phenotype of this Gr-1high/CD48- population was consistent with the mouse band cells.

54

Figure 8. The Gr-1high/CD48- band cells isolated from BM cells of WT mice. (a) The Gr-1high/CD48- population was purified by FACS sorting from fresh BM cells of C57BL/6 mice. The purified cells were stained with HEMA 3 kit after cytospin, and detected under an inverted light microscope. Bar = 20 μm. (b) Data shown are the expression profiles of indicated surface markers (red histogram) or background signals with isotype-matched control antibody (blue histogram) of the Gr-1high/CD48- population. Relative cell size and granularity are also shown with FSC and SSC. The Gr-1high/CD48- band cells were cultured for 4 days in the presence of G-CSF (20 ng/ml). The morphology of resulting population was examined after cytospin and HEMA 3 staining. Bar = 20 μm (c), and the surface phenotype was examined with FACS (d).

Band cells have long been known as the precursors of mature granulocytes or neutrophils

(Bainton et al., 1971). We purified the Gr-1high/CD48- band cell population from crude

BM cells, and cultured them in the presence of G-CSF, a cytokine that can effectively induce the differentiation of granulocytes. After 4 days culture with G-CSF, band cells

55 exhibited significant morphological changes, as they increased cell size and acquired highly segmented nuclei (Figure 8c). In addition, they maintained the high level surface expression of CD11b and Ly6G, but still lacked the expression of CD11c (Figure 8d).

Therefore, band cells were functionally competent in the granulocyte lineage differentiation when treated with G-CSF, although they can also differentiate into DCs when treated with GM-CSF (Figure 5). It suggested that cytokines might play a determinative role to induce the different development pathways of band cells, at least in vitro.

Construction of the co-culture system

Although band cells were capable of differentiating into DCs as shown in Figure 5, they started to die upon purification even in the presence of GM-CSF (data not shown).

Therefore, we reasoned that other cell types in BM cultures may serve as feeder cells for band cells. To test this possibility, we constructed a co-culture system (Figure 9). Band cells purified from C57BL/6 mice (CD45.2+) were co-cultured with crude BM feeder cells from B6 SJL mice (CD45.1+) in the presence of GM-CSF. After 6 days, we were able to identify or purify the band cell-derived cells from the feeder cells by differential staining with anti-CD45.1 and anti-CD45.2 antibodies. The viability of the band cells or band cell-derived cells was sustained in this co-culture system (data not shown). This co- culture system enabled us to study the differentiation of DCs from band cells.

56 Band Cells (CD45.2) BM Feeder Cells (CD45.1)

6 Days (+) GM-CSF

Purification CD45.2

CD45.1

Figure 9. Construction of the co-culture system. The Gr-1high/CD48- band cells were purified from the BM of the C57BL/6 mice and then co-cultured with the crude BM cells harvested from B6 SJL mice in the presence of GM-CSF (10 ng/ml). After 6 days, the whole cells in the co-culture were stained with anti- CD45.1 and CD45.2 antibodies for FACS. The band cell-derived cells, which were CD45.2+/CD45.1-, were identified and purified by FACS sorting for the following analyses.

Band cells give rise to a novel DC subset, “grDC”

After 6 days, we FACS-purified the band cell-derived population (CD45.2+/CD45.1-) and analyzed their morphology (Figure 10a). Compared to the starting population, these band cell-derived cells showed a larger cell size and round-shaped nucleus, and importantly, they showed characteristic morphology of DCs. The phenotype analysis revealed that these cells expressed the conventional markers for DCs, such as CD11c, MHC II and

CD205. Interestingly, they retained surface expression of Ly6G, which is a typical marker for granulocytes (Figure 10b). To our knowledge, there is no report of Ly6G+ DC subset in the literature. Therefore, we believe that this Ly6G+ DC population, derived from band cells, is a novel DC subset, which we have termed granulocyte-derived DC or

57 grDC. We also acquired the BM-derived DCs (BMDCs) by a traditional BM culture protocol and monocyte-derived DCs (moDCs) by culturing BM monocytes with GM-

CSF and examined their Ly6G expression. The Ly6G expression was undetectable on

BMDCs or moDCs, compared to the high expression of Ly6G on grDCs (Figure 10c).

Thus, in the mouse DC populations, we can use Ly6G as a marker to distinguish grDCs from the conventional DCs (cDCs).

a b

MHC II CD11c

CD205 Ly6G

c d

58 Figure 10. Band cells give rise to a novel DC subset. (a) The Gr-1high/CD48- band cells were purified from the BM of the C57BL/6 mice and then co-cultured with the crude BM cells harvested from B6 SJL mice in the presence of GM-CSF (10 ng/ml). After 6 days, the band cell-derived cells (CD45.2+/CD45.1-) were purified by FACS sorting and stained by HEMA 3 after cytospin. The morphology of the band cell- derived cells was detected under an inverted light microscope. Bar = 20 μm. (b) Data shown are the expression profile of indicated surface markers (red histogram) or the isotype-matched control antibody (blue histogram) of the cell population derived from band cells. (c) BMDCs were acquired by culturing BM cells with GM-CSF (10 ng/ml) according to Inaba’s protocol (Inaba et al., 1992); the floating cells in the BM culture were removed on day 2 and day 4, and fresh media was added to the adherent cells. On day 6, the CD11c+ BMDC population was collected and analyzed for Ly6G expression. Monocytes (CD11b+/CD11C-/Ly6G-) were purified from crude BM cells by FACS sorting, and then cultured in the presence GM-CSF (10 ng/ml). On day 6, the CD11c+ moDC population was collected and analyzed for Ly6G expression. The samples were stained with anti-Ly6G (red histogram) or the isotype-matched control antibody (blue histogram). (d) The grDC population was FACS-purified from the co-culture system on day 6, pulsed with OVA323–339 or OVA257–264 peptide (10 μg/ml), washed extensively, and then co-cultured with CD4 T cells purified from OT-II TG mice or CD8 T cells purified from OT-II TG mice, respectively. Peptide-pulsed band cells or DCs were added to T cell cultures at the indicated cell numbers and incubated for 3 days. Data shown were the means ± SD (n=3) of 3H-thymidine uptake by T cells stimulated with OVA peptide-pulsed cells (●) or the base line levels of 3H-thymidine uptake by T cells in the absence of OVA peptide pulsing (○).

To test whether grDCs are competent antigen-presenting cells, we purified grDC population on day 6 from the co-culture system, pulsed them with OVA-derived peptides and then added them into the cultures of CD4 or CD8 cells, which were isolated from OT

II or OT I TG mice, respectively. As shown in Figure 10d, grDCs efficiently stimulated the proliferation of antigen specific CD4 and CD8 T cells, indicating grDCs can effectively present antigen peptides to T cells. Furthermore, after pulsed with OVA protein, grDCs can also trigger the proliferation of OT II CD4 T cells and OT I CD8 T cells, although not as strongly as pulsing with OVA peptides (data not shown). Thus, we concluded that the band cells can give rise to fully functional DCs that are characterized by Ly6G expression.

Gene expression profiles differ markedly between grDCs and cDCs

To further test the difference between Ly6G+ grDCs and Ly6G- cDCs, we preformed the genechip analysis. According to Inaba’s BMDC culture protocol (Inaba et al., 1992), the

59 floating cells, including immature and mature granulocytes, were discarded on day 2 and day 4, so we cannot obtain grDCs with this protocol. In our protocol, we kept both floating and adherent cells in the BM cultures with GM-CSF for 6 days. Therefore, we were able to obtain both cDCs and band cell-derived grDCs. The total RNA was extracted from FACS-purified grDCs (CD11c+/ MHC II+/ Ly6G+) and cDCs (CD11c+/

MHC II+/ Ly6G-) in the BM cultures, and was used for the analyses by Affymetrix

Mouse Genome 2.0 Array. The results demonstrated that grDCs differed from cDCs in their gene expression profiles, in that more than 300 genes expressed differentially between these two DC subsets. We identified a cluster of 170 genes expressed predominantly by grDCs, and this cluster included CRAMP, CD62L and MMP9 (Figure

11).

60 grDC cDC

cathelicidin (cramp)

cd62l mmp9

Figure 11. The gene expression profiles are different between grDCs and cDCs. The BM cells were cultured in the presence of GM-CSF (10 ng/ml), and the floating cell together with fresh media was put back to the cultures every 2 days. On day 6, the grDCs (CD11c+/ MHC II+/ Ly6G+) and cDCs (CD11c+/ MHC II+/ Ly6G-) were purified by FACS sorting and the total RNA was extracted for Affymetrix Mouse Genome 2.0 Array. The relative expression levels of genes were presented by a color scale. The results were based on three independent genechip analyses. The genes that are expressed predominantly in one population in terms of the magnitude of expression (>2-fold) and the statistical significance (n=3, P<0.05) are shown in the heat map.

61 grDCs produce large amount of CRAMP

CRAMP is a potent antimicrobial peptide with a strong bacterial killing potential

(Pestonjamasp, et al., 2001). Several cell types have been shown to produce CRAMP under different conditions, but there is no report about DCs producing CRAMP. Our genechip data demonstrated that mRNA level of CRAMP was immense in grDCs. This observation was confirmed by real-time RT-PCR; we observed that the expression of

CRAMP RNA by grDCs was 10-fold higher than cDCs (Figure 12a). Western blotting further revealed abundant CRAMP proteins in the extracts of grDC but not cDC (Figure

12b). Interestingly, grDCs exhibited 16.4 and 15.0 kD CRAMP bands, which correspond to the previously reported CRAMP polypeptides produce in mouse (Bergman et al.,

2006). In the immunofluorescence staining, we observed that grDCs showing dendritic morphology indeed produced CRAMP, and once again, the cDC population showed little immuno-reactivity to anti-CRAMP antibody (Figure 12c).

a c

b

62 Figure 12. CRAMP produced by grDCs but not cDCs. The grDCs (CD11c+/MHC II+/Ly6G+) and cDCs (CD11c+/MHC II+/Ly6G-) were purified from BM cultures by FACS sorting as described before. (a) The total RNA was extracted for Real-Time RT-PCR, and the results were normalized by the RNA expression of β-actin. Data shown are the means ± SD from three independent experiments. ***P<0.001. (b) Protein was extracted from FACS-purified grDCs and cDCs. The concentration was measured and the equal amount of the protein from grDCs and cDCs were used for western blotting to examine the expression of CRAMP. The molecular weight was determined by the protein standard in the same gel. (c) FACS-purified grDCs and cDCs were stained with rabbit anti-mouse CRAMP antibody (left two panels) or control rabbit serum (right two panels), followed by staining with Alexa Fluor 488 goat anti-rabbit IgG antibody. The cells were then observed under a fluorescence microscope to examine the production of CRAMP. Bar = 20 μm.

grDCs have higher bacterial killing capacity than cDCs due to the production of CRAMP

To study the functional contributions of grDC-associated CRAMP molecules, we compared the bacterial killing capacity between grDCs versus cDCs. We first examined the intracellular bacterial killing capacity of these two DC subsets by infecting the same number of grDCs or cDCs with living E.coli K-12 (MOI=10). After 1 h incubation, the extracellular E.coli were killed, and DCs were lysed to determine the CFU of the intracellular bacteria. At time 0, the CFU of E.coli inside grDCs was about two times higher than in cDCs, indicating that more bacteria were internalized by grDCs. The CFU dramatically decreased in the grDC lysate after 30 min, suggesting a rapid killing of intracellular bacteria by grDCs. The number of E.coli inside grDCs decreased at the later time points, indicating a constant bacterial killing. In contrast, cDCs were also able to eliminate the intracellular bacteria in a time-dependent manner, but the bacterial killing was not as efficient as grDCs, since the reduction of CFUs in cDCs was not as fast as in grDCs (Figure 13a). To test whether this rapid bacterial killing by grDCs is due to their production of CRAMP, we purified grDC and cDC populations from the BM cultures of

CRAMP KO mice (Nizet et al., 2001) as well as WT mice. The intracellular bacterial

63 killing assays were performed in the same way. As shown in Figure 13b, the intracellular bacterial elimination was retarded in grDCs from CRAMP KO mice compared to grDCs from WT mice. It suggested that the CRAMP played an important role in the bacterial killing by grDCs. The intracellular E.coli killing by cDCs was almost comparable between CRAMP KO mice and WT mice.

a b

c

Figure 13. Efficient bacterial killing capacity of grDCs is due to the production of CRAMP. (a) The grDCs (CD11c+/MHC II+/Ly6G+) and cDCs (CD11c+/MHC II+/Ly6G-) were purified from BM cultures of WT mice by FACS sorting. 5x104 DCs were incubated with living E.coli (MOI=10) at 37ºC for 1 h in the antibiotics-free complete RMPI media supplemented with 1% human plasma. The extracellular bacteria were killed by a 30 min exposure to kanamycine (50 μg/ml). After removing kanamycine (Time 0), DCs were incubated in the antibiotics-free RPMI media and lysed at different time points. The cell lysates were

64 diluted and cultured on LB agar overnight. The numbers of bacterial colonies was counted and colony- forming units (CFU) were calculated. The CFUs in the grDC lysates (●) were compared to the CFU in the cDC lysate (○) at each time point. (b) The grDCs and cDCs were purified from BM cultures of CRAMP KO mice as well as WT mice by FACS sorting. The intracellular E.coli killing was tested in the same way as described in (a), and CFU at different time points were normalized to the CFU at time 0. The CFU in the lysates of DCs from the WT mice (●) were compared to the CFU in the lysates of DCs from the CRAMP KO mice (○) at each time point. (c) The grDCs and cDCs were purified from BM cultures of WT mice by FACS sorting. As indicated, equal amounts of the cell extracts or synthetic CRAMP peptide (8μM) were added in the E.coli cultures supplemented with 10% LB, and blocking antibody against CRAMP as well as control IgG were added in some cultures. After incubation at 37ºC for 4 h, the media was harvested and the CFUs were calculated. Data shown are the means ± SD from three independent experiments. *P<0.05, **P<0.01, ***P<0.001.

We also tested the bacterial killing by the cell extracts of grDCs and cDCs (Figure 13c).

The same amount of cDC or grDC extracts were added into the E.coli cultures, and the anti-CRAMP blocking antibodies or control antibodies were added to some cultures together with cell extracts. The synthetic CRAMP peptide was used as a positive control.

After 4 h incubation, the CFUs of E.coli with different treatment were calculated. As expected, the synthetic CRAMP peptide effectively killed bacteria. The grDC extracts, which contained 5-10 μM CRAMP as estimated in western blotting, also showed a potent bacterial killing activity. The observed activity was much stronger than that in cDC extracts. Importantly, anti-CRAMP blocking antibodies but not control antibodies significantly inhibited the bacterial killing activity of grDC extracts.

On basis of these observations, we concluded that grDCs had potent ability to kill bacteria compared to cDCs, and that the CRAMP production by grDCs contributed to their bacterial killing potential.

65 Expression of CD62L and MMP9 by grDCs

According to our GeneChip data, grDCs expressed higher levels of CD62L and MMP9 mRNA than did cDCs. To confirm these findings, we tested CD62L protein expression by grDCs and cDCs. FACS analysis revealed that grDCs expressed abundant CD62L, whereas the CD62L expression on the surface of cDCs was marginal (Figure 14a).

CD62L (also known as L-selectin) is a cell adhesion molecule and acts as the “homing receptor” for leukocytes to migrate to lymphoid tissues (Khan et al., 2003). Predominant expression of CD62L on grDCs might facilitate their migration through endothelial venules.

a

b

Figure 14. The grDC population expresses high level of CD62L and MMP9 (a) Data shown are the expression profile of CD62L (red histogram) or the isotype-matched control antibody (blue histogram) within the Ly6G+/CD11c+ grDC population (left) or Ly6G+/CD11c- cDC population (right) from BM

66 cultures. (b) The grDCs (CD11c+/MHC II+/Ly6G+) and cDCs (CD11c+/MHC II+/Ly6G-) were purified from BM cultures by FACS sorting. For Real-Time RT-PCR, total RNA was extracted and the results were normalized by β-actin mRNA expression of (left). For ELISA, grDCs and cDCs were incubated in complete RPMI for 24 h (1×106 cells/ml). The supernatants were collected and used to measure secreted MMP-9 (right). Data shown are the means ± SD from three independent experiments. ***P<0.001.

We also compared grDCs versus cDCs for their mRNA level of MMP9 by real-time RT-

PCR, and their MMP9 protein secretion by using ELISA. The results demonstrated about

8-fold higher MMP9 mRNA expression in grDCs than cDCs, and that grDCs secreted significanly higher amounts of MMP9 (Figure 14b). MMP9 is type IV collagenase involed in breaking down the extracellular matix and it has multiple fuctions in the normal and disease conditions (Nagase & Woessner, 1999). The role of MMP9 produced by grDCs needs to be determined in further studies.

Comparison of surface phenotype between grDCs and cDCs

To distinguish grDCs versus cDCs, we tested the expression of various surface molecules by these two DC subsets (Figure 15). The surface phenotypes of grDCs and cDCs are generally comparable to each other, except that Ly6G and CD62L were displayed only by grDCs.

67

68 Figure 15. The surface phenotype between grDCs and cDCs. Data shown are the FACS profiles with antibodies against the indicated surface markers (red histogram) or with isotype-matched control antibodies (blue histogram) within the Ly6G+/CD11c+ grDC population or Ly6G+/CD11c- cDC population from BM cultures.

grDCs possess potent endocytic potential

Since grDCs are derived from band cells, which exhibit a potent endocytic ability, we hypothesized that grDCs might maintain this fuctional property. To test this, we purified grDCs and cDCs from BM cultures, and incubated them with FITC-dextran, fluorescent latex beads or living E.coli, and then compared the uptake activity between grDCs and cDCs. The uptake of FITC-dextran and fluorescent latex beads was determined by the

MFI generated by the internalized molecules or particles and the uptake of E.coli was determined by the CFU in the DCs. The results of the three experiments demonstrated that grDCs were able to uptake more extracellular soluble molecules, particles and bacteria compared to cDCs (Figure 16).

a b c

Figure 16. Endocytic potential between grDCs versus cDCs. The grDCs (CD11c+/MHC II+/Ly6G+) and cDCs (CD11c+/MHC II+/Ly6G-) were purified from BM cultures by FACS sorting. The cells were incubated with FITC-dextran (MW 70000) (a) or fluorescent latex beads (b) for 30 min at 37ºC, and then analyzed by FACS. The endocytotic potential was determined based on the mean fluorescence intensity (MFI) generated by the internalized molecules or particles. (c) The uptake of E.coli was examined as described in Figure 13a. Data shown are the means ± SD from three independent experiments. ***P<0.001.

69 Identification of grDCs in vivo

Having identified grDCs in vitro, we next sought to determine whether a Ly6G+ grDC equivalent population exists in the mouse tissues. We examined the Ly6G expression in the CD11b+/CD11c+/MHC II+ DC population in the spleen. As shown in Figure 17, a small but significant fraction (~2.5%) in the splenic DCs was found to express Ly6G, indicating the existence of grDC. We also detected small numbers of Ly6G+ DCs in mesenteric LNs and Ly6G+/MHC II+ DCs in peripheral blood (data not shown). Thus, it appears that grDCs do exist at least in selected tissues in vivo.

CD11b+ Population CD11b+/MHC II+/CD11c+ MHC II MHC

CD11c Ly6G

Figure 17. Identification of grDCs in mouse spleen. Total splenocytes were harvested by dissociating the spleen, followed by red blood cells lysis. The cells were labeled with anti-CD11b magnetic particles and the CD11b+ fraction was purified by magnetic selection. The enriched population was stained with anti- CD11c, anti-MHC II and anti-Ly6G antibodies for FACS analysis. The gated population in the left panel is the CD11b+/CD11c+/MHC II+ splenic DC. The right panel shows the FACS profile with antibody against Ly6G (red histogram) or the isotype-matched control antibody (blue histogram) within this DC population.

DsRed+ band cells and DCs emerge in PECs under inflammation

We have observed a robust increase in DsRed+ cells in the skin of pIL1-DsRed TG mice under inflammatory conditions (Matsushima et al., 2010), and we also observed a dramatic increase of DsRed+ band cells when BM cells from the same TG mice were

70 cultured GM-CSF. We speculated that DsRed+ cells may emerge in vivo in other inflammation models. To test this, we employed a well established peritonitis model achieved by i.p. injecting thioglycollate (Segal et al., 2002). Twenty-four hours after thioglycollate injection, large numbers of DsRed+ cells became detectable in the peritoneal cavity. Similar to the DsRed+ cells found in the BM cultures on day 2, almost all the DsRed+ cells in PECs expressed CD11b and most of them were CD11c- (Figure

18a). We analyzed the surface phenotype of the DsRed+/CD11b+/CD11c- population in

PECs, and the results revealed that this population was MHC II-/Ly6G+/Ly6C+/CD48-

(Figure 18b). This phenotype observed in vivo was identical to that of the

DsRed+/CD11b+/CD11c- band cells that we identified in the BM cultures (Figure 6c).

Furthermore, the morphology of this population highly resembled the band cells, based on small cell size, low granularity and characteristic nucleus (Figure 18c). Thus, the

DsRed+/CD11b+/CD11c- population emerging in PECs one day after induction of inflammation represented band cells, which may serve as the precursors of grDCs.

71 a b

c

d

Figure 18. DsRed+ band cells and DCs emerge in PECs under inflammation. Thioglycollate (3%) was i.p. injected in pIL1-DsRed TG mice to induce acute inflammation in peritoneal cavity. Twenty-four hours later, mice were sacrificed and the PECs were harvested by washing peritoneal cavity with 7 ml PBS. (a) The PECs were washed and stained with anti-CD11b and anti-CD11c antibodies for FACS analysis. (b) Data shown are the expression profiles of indicated surface markers (red histogram) or the isotype-matched control antibodies (blue histogram) within the DsRed+/CD11b+/CD11c- population. (c) The DsRed+/CD11b+/CD11c- population was purified from PECs by FACS sorting. The cells were subsequently stained by HEMA 3 kit after cytospin, and the morphology was detected under an inverted light microscope. Bar = 20 μm. (d) Data shown are the expression profile of Ly6G (red histogram) or the isotype-matched control antibody (blue histogram) within the DsRed+/CD11c+DC population (left) or DsRed-/CD11c+DC population (right).

As shown in Figure 18a, there were two DC populations in the PECs with different

DsRed expression, DsRed+/CD11c+ DCs and DsRed-/CD11c+ DCs. We examined the

Ly6G expression in both DC populations in order to indentify grDCs (Figure 18d). Only

72 DsRed+/CD11c+ DCs contained a significant fraction of Ly6G+ cells, indicating the existence of grDCs in PECs under inflammation. The observation that grDCs maintained the DsRed expression was consistent with the results that the DsRed+ band cells gave rise to DsRed+ DCs in vitro (Figure 5). It suggested that IL-1β promoter was activated in grDCs after differentiation from band cells.

Emergence of grDCs in vivo under inflammatory conditions

Inflammation induced by thioglycollate i.p. injection caused the rapid recruitment of a large number of band cells to the peritoneal cavity. The same model was employed to determine whether grDCs may emerge in the inflamed sites. We harvested the PECs at different time points after i.p. injection of thioglycollate, and counted the numbers of

Ly6G+/CD11c+/ MHC II+ grDC population (Figure 19a). Very few grDCs were detected in the peritoneal cavity in the steady state on day 0. We observed striking increases in the number of grDCs with a peak on day 2. In contrast, no increase was observed after injection of sterilized water as a vehicle control. Two days after thioglycollate injection, the Ly6G+/CD11c+/ MHC II+ population in PECs was FACS-purified for morphological analysis. The results demonstrated that the grDCs isolated in vivo showed a typical DC morphology (Figure 19b). We also used living E.coli to induce the peritonitis. One day after i.p. injection of E.coli, we collected the PECs and detected the expression of Ly6G within the CD11c+/ MHC II+ DC population. Only E.coli injection but not PBS injection resulted in a significant Ly6G+ fraction in the DC population, indicating the emergence of grDCs (Figure 19c). Thus, inflammation is associated with rapid and remarkable emergence of grDCs.

73 a

b

74 c

Figure 19. Inflammation induces grDC emergence in vivo. (a) Thioglycollate (3%) (●) or sterilized water (○) was i.p. injected to WT mice. The PECs were harvested and analyzed by FACS at the indicated time points. The number of Ly6G+/CD11c+/MHC II+ grDC population was counted and compared to time 0. Data shown are the means ± SD from three independent experiments. ***P<0.001. (b) Ly6G+/CD11c+/MHC II+ grDCs in PECs were purified by FACS sorting 2 days after thioglycollate injection. The cells were subsequently stained by HEMA 3 kit after cytospin, and the morphology was detected under a phase contrast microscope. Bar = 20 μm. (c) Living E.coli (1×106-5×106/mouse) or PBS was i.p. injected in WT mice. After one day, PECs were harvested and the expression of Ly6G (red histogram) or the isotype-matched control antibody (blue histogram) within CD11c+/MHC II+ DC population was examined.

Band cells give rise to DCs in vivo under inflammatory conditions

We have shown that band cells can give rise to grDCs in vitro. The key question concerned whether band cells are able to differentiate into DCs in vivo. Because inflammation promotes grDCs emergence, we employed the peritoneal inflammation model to study band cells differentiation in vivo. We first i.p. injected thioglycollate to

B6 SJL mice (CD45.1+) to induce peritoneal inflammation. After 6 h, we transferred the band cells (which were CD11c-/MHC II-) isolated from the BM of C57BL/6 mice

(CD45.2) by i.v. injection, and 42 h later, we analyzed the surface phenotype of

CD45.2+/CD45.1- population in the PECs, which were derived from the band cells. As demonstrated in Figure 20, the transferred population acquired the expression of CD11c as well as MHC II, indicating the band cells differentiated into DCs in the inflammatory

75 sites. Band cells were also adoptively transferred to the recipient mice without inducing inflammation, and they failed to differentiate into DCs (data not shown). It suggested that only under inflammatory conditions, can band cells give rise to DCs in vivo.

Figure 20. Band cells give rise to DCs in vivo under inflammatory conditions. B6 SJL recipient mice were injected with thioglycollate to induce peritoneal inflammation, and 6 h later, band cells (5×105-2×106 cells) purified from crude BM of C57BL/6 mice were transfer to the recipients by i.v. injection. After 42 h, PECs were harvested and the CD45.2+/CD45.1- population was analyzed for their CD11c (left) and MHC II (right) expression by FACS. The expression profiles of indicated markers and isotype-matched control antibodies are shown in red dot plots and blue dot plots, respectively.

Comparison of in vivo bacterial uptake and antigen presentation capacity by grDCs versus cDCs

Compared to cDCs, grDCs exhibited more efficient bacterial uptake in vitro. To study in vivo bacterial uptake, we employed the above peritonitis model. Two days after thioglycollate i.p. injection, we i.p. injected E.coli engineered to express GFP. After 1 h, we harvested PECs and analyzed the GFP signals within Ly6G+ grDCs and Ly6G- cDCs.

The MFI of the GFP signal in grDCs was significantly stronger than that in cDCs, suggesting that grDCs uptake more bacteria in vivo (Figure 21a). Our findings implied that grDCs may play an important role in the clearance of bacteria upon infection.

76

Since grDCs captured more bacteria in vivo, they should be loaded with more bacteria- associated antigens. We speculated further that grDCs may be able to present the antigens to T cells efficiently. The peritonitis model was employed to test this possibility.

Inflammation was induced by thioglycollate injection, and after 2 days, WT E.coli or

E.coli expressing full length OVA cDNA were injected to the inflamed peritoneal cavity.

One hour later, Ly6G+ grDCs and Ly6G- cDCs were purified and subsequently co- cultured with CD4 T cell isolated from OT II TG mice (Figure 21b). Both grDCs and cDCs purified from the mice infected by OVA-E.coli were able to present the antigen to

OVA-reactive CD4 T cells; however, the grDCs were more effective than cDCs in this capacity. As expected, neither grDCs nor cDCs purified from the mice infected by WT

E.coli induced the proliferation of OT II CD4 T cells. We also employed the same strategy to compare the cross-presentation capacity by co-culturing grDCs and cDCs with

CD8 OT I T cells; however, neither of them showed a potent capacity to induce CD8 T cell proliferation in this model (data not shown).

77 a

b

Figure 21. Comparison of in vivo bacterial uptake and antigen presentation capacity by grDCs versus cDCs. (a) Thioglycollate (3%) was i.p. injected to induce peritonitis 48 h before the i.p. injection of 1x106 E.coli constitutively expressing GFP. After 1 h, PECs were harvested and washed to remove extracellular bacteria. The MFI of GFP signal was analyzed within grDC (CD11c+/Ly6G+) and cDC (CD11c+/Ly6G-) population by FACS to compare the amount of internalized E.coli. (b) Thioglycollate (3%) was i.p. injected to induce peritonitis and after 48 h, 1×106 E.coli expressing full length of OVA or 1×106 WT E.coli were i.p. injected. PECs were collected 1 h later and grDCs (CD11c+/Ly6G+) as well as cDCs (CD11c+/Ly6G-) were purified by FCAS sorting. CD4 T cells purified from OT-II TG mice were plated in 96-well plates (5×104 T cells/well). After extensive washing, DCs (3×103 DCs/well) were added to OT II CD4 T cell cultures as indicated and incubated for 3 days. Data shown were the mean ± SD (n=3) of 3H- thymidine uptake. Data shown are the means ± SD from three independent experiments. *P<0.05, ***P<0.001.

78 DISCUSSION

DCs exhibit high complexity in terms of phenotypically and functionally distinct subsets, and plasticity in their differentiation, compared to the other leukocytes (Wu & Liu, 2007;

Liu & Nussenzweig, 2010). A variety of DC precursor populations have been identified in mice, and they have different phenotypes, tissue distributions as well as differentiation commitment (Table V).

Table V. DC precursors in mice

Tissue Differentiation Phenotype Reference distribution commitment

Subcutaneous Randolph et al., CD11b+/F4/80−/CD205− cDCs, Macrophages tissue 1999

CD11b+/F4/80+/CX3CR1lo/CCR2 cDCs Geissmann et al., Blood +/Gr-1+ (inflamed tissues) 2003

CD11b+/F4/80+/CX3CR1hi cDCs Geissmann et al., Blood /CCR2-/Gr-1- (non-inflamed tissues) 2003

CD11b+/F4/80+ Ginhoux et al., Blood LCs, Macrophages /Gr-1hi/CD115+ 2006

CD11b+/Ly6G-/CD115+ BM, Blood DC-SIGN+/MMR+ DCs Cheong et al., 2010

Bruno, et al., CD11c+/CD31+/Ly6C+ BM, Blood cDCs, Macrophages 2001

CD11c+/MHC II-/ Blood cDCs, pDCs del Hoyo et al., 2002 B220+/CD43+/Gr-1-/F4/80-

79 Fogg et al., Lin–/c-Kit+/CX3CR1+ BM cDCs, Macrophages 2006

CD11cint/MHC II- Naik et al., Spleen cDCs /CD45RAlo/CD43int 2006

CD11cint/MHC II- Naik et al., Spleen CD8+ cDCs /CD45RAlo/CD43int/CD24hi 2006

Lin−/CD34+/CD16/32− Mende et al., BM cDCs, LCs /c-Kithi /Sca-1− Flt3 + 2006

Onai et al., Lin−/c-Kitint/Flt3+/CD115+ BM cDCs, pDCs 2007

In 1998, Oehler and his colleagues reported that the precursors of human PMNs (PMNp), which were isolated from peripheral blood of patients with chronic myeloid leukemia, bacterial infections or treated with G-CSF, acquired some characteristics of DCs after they were cultured with a cocktail of three cytokines (GM-CSF/IL-4/TNF-α) for 9 days.

The PMNp-derived DCs exhibited conventional DC markers, such as HLA-DR, HLA-

DQ, CD1 molecules and co-stimulatory molecules (CD40, CD80 and CD86), but lost the expression of CD15 and CD54, which are marker for granulocytes. In addition, these

DCs showed APC functions as they were effective to induce proliferation of allogeneic T cells. The authors concluded that these PMNp-derived DCs closely resembled “classical”

DC populations, because they were indistinguishable from conventional DCs in terms of morphology, phenotype or function (Oehler et al., 1998). Several studies have also revealed that human PMNs acquire some features of DCs either in vitro culture or under certain pathological conditions in vivo. PMNs purified from blood of healthy donors escaped from rapid apoptosis and expressed MHC II, CD80, CD83 and CD86, when they

80 were cultured with GM-CSF, IL-4 and IFN-γ (Iking-Konert et al., 2001). In another study,

PMNs acquired the expression of MHC II, CD40, CD83 and CCR6 when cultured with

GM-CSF, TNF-α and IFN-γ (Yamashiro et al., 2000). Stimulation of IL-15 was demonstrated to induce the surface expression of MHC II, CD83 and CD14 on human

PMNs as well (Abdel-Salam & Ebaid, 2008). Up to 15% of the peripheral PMNs of the patients with Wegener's granulomatosis were found to express MHC II, and this expression was closely related to the progress of disease, because it decreased immediately after therapy. Therefore, the MHC II expression on PMNs surface can be used as a diagnostic marker for Wegener's granulomatosis (Hänsch et al., 1999). During acute bacterial infections, the expression of CD83 was dramatically increased in the human PMNs, while that of MHC II, CD80 and CD86 remained negative (Iking-Konert et al., 2002). A paper published recently showed that in the synovial fluid collected from inflamed joints of patients with RA, the infiltrated neutrophils acquired the expression of

MHC II and CD83. After exposure to the synovial fluid from RA patients, PMNs collected from the healthy donors up-regulated CD83 expression. IFN-γ and GM-CSF were involved in the transdifferentiation of the neutrophils. Nevertheless, the authors still considered these MHC II+/CD83+ cells to be “dendritic-like” PMNs, a portion of neutrophils that escaped from apoptosis and acquired DC markers under stimulation

(Iking-Konert et al., 2005).

The results in our present study have demonstrated that immature granulocytes, band cells, acquire characteristics of DCs when cultured with GM-CSF in vitro and under inflammatory conditions in vivo. Importantly, the DCs derived from band cells, which we

81 have termed grDCs, show distinct phenotype, gene expression profiles and specialized functions. Thus, we have concluded that the band cell-derived grDC population represents a previously unknown DC subset.

Granulocytes, especially neutrophils, are generally regarded as highly competent phagocytes that provide the first line of defense of the (Segal,

2005). In the steady state, mature granulocytes are circulating in the blood and a marginating pool called “physiological regional granulocyte pool” (Peters, 1998). In inflammatory conditions, the precursor of granulocytes (band cells, metamyelocytes and myelocytes) are released from BM and they become readily detectable in the blood as well as the inflamed tissues. Meanwhile, the level of key cytokines for , G-

CSF and GM-CSF are up-regulated at sites of inflammation (Marsh et al., 1967;

Hamilton, 2008). As a result, the number of mature granulocytes is dramatically boosted and they rapidly migrate to the inflamed sites. The migration of granulocytes is dependent on selectins and integrins, in which L-selectin and CD11b/CD18 play a most crucial role in their trafficking through blood vessels (Doerschuk, 1992; Hogg &

Doerschuk, 1995; Tedder et al., 1995).

Neutrophils are the first phagocytes arriving at the inflamed sites, and they efficiently engulf and kill the invading microorganisms. The killing and degradation of the internalized microbes by neutrophils is mediated by reactive oxygen species (ROS) dependent and independent mechanisms. The activation of neutrophils triggers the membrane-associted NADPH oxidase system to generate a large amount of ROS and

82 release the ROS into phagolysosomes. Subsequently, ROS can react with organic molecules and kill the internalized microbes that reside in the phagolysosomes (Segal,

2005; Dale et al., 2008). Uptake of microbes also causes the degranulation of neutrophils to release the antimicrobial peptides and proteolytic proteins stored in the granules. The granules either fuse with phagolysosomes containing microbes or are secreted to the extracellular traps (Faurschou & Borregaard, 2003: Borregaard et al., 2007). This process is very fast because granule peptides and proteins are not synthesized de novo at the inflamed sites, but are synthesized and stored during granulocyte differentiation

(Theilgaard-Mönch et al., 2006). Three types of granules are formed during granulocyte differentiation. The primary granules, which contain azuzocidin and cathepsin, are formed at the stage of . The secondary granules, which contain cathelicidin and lactoferrin, are formed at the stage of myelocytes and metamyelocytes. The tertiary granules, which contain peptidoglycan recognition preoteins and urokinase plasminogen activator, are formed in band cells (Soehnlein, 2009). Because grDCs are derived from band cells, and cathelicidin is synthesized and stored within secondary granules that have been formed in band cells (Sorensen et al., 1997; Soehnlein, 2009). The grDCs may retain some secondary granules which are the source of their cathelicidin production.

Recent studies have shown that granulocytes directly and indirectly participate in the innate immunity. Not only do granulocytes produce chemokines to recruit monocytes,

DCs and T cells to the inflamed sites, they also produce a variety of cytokines to regulate the function of APCs (Nathan 2006). By secreting TNF-α or glycosylation-dependent cell interaction with immature DCs, activated neutrophils are able to induce DC maturation,

83 and subsequently initiate T cell responses (Bennouna & Denkers, 2005; van Gisbergen et al., 2005). Parasite triggered neutrophils produce a number of chemokines, such as CCL3,

CCL4, CCL5 and CCL20, which have strong chemotaxis effects to recruit immature DCs to the sites of infection (Bennouna et al., 2003). Bacteria infected neutrophils can serve as substrates for cross-presentation of bacterial antigens by DCs (Tvinnereim et al., 2004).

After pulsed with OVA protein, neutrophils even directly cross-prime OT I CD8 T cells in vivo (Beauvllain et al., 2007). Thus, granulocytes can function as bi-functional leukocytes linking innate immunity to adaptive immunity. Our data demonstrate that immature granulocytes can differentiate into grDCs, further expanding this view.

We believe that grDCs represent a novel subset of DCs but not a mal-differentiated granulocyte subset. They expresse well accepted markers for mouse DCs, such as CD11c,

MHC II and CD205. The further express co-stimulatory molecules (CD40, CD80 and

CD86) and CD1d at the levels comparable to those on conventional DCs.

Morphologically, grDCs purified in vitro and in vivo are indistinguishable from conventional DCs and quite different from band cells or mature granulocytes. Moreover, grDCs are found to be fully functional APCs in presenting peptides to both CD4 and CD8

T cells.

We are able to acquire considerable number of grDCs by culturing BM cells with GM-

CSF, but we keep adherent and floating cells together in our culture system. In the traditional BMDC cultures, the floating cells, mainly immature and mature granulocytes, are removed every 2 day by washing (Inaba et al., 1992). Therefore, the precursor

84 population of grDCs is missing, and it may be one of the reasons why grDCs have been neglected.

Importantly, grDCs retain some features of granulocytes. The expression of Ly6G, a granulocyte marker, distinguishes grDCs from any of the currently known DC subsets.

Compared to cDCs, grDCs express high levels of CD62L, also known as L-selectin, which is highly expressed by granulocytes (Tedder et al., 1995). As a member of matrix metalloproteinase family, MMP9 plays key roles in extracellular matrix remodeling

(Opdenakker et al., 2001). Granulocytes are the most major source of MMP9 and release

MMP9 upon stimulation with G-CSF, Vitamin E, IL-8, TLR2 agonist and kinin peptides

(Heissig et al., 2002; Renò et al., 2004; Chakrabarti & Patel, 2005; Page et al., 2009;

Ehrenfeld et al., 2009). Our data have demonstrated grDCs constitutively secret larger amounts of MMP9 even without treatment. Whether the secretion of MMP9 by grDCs is enhanced after stimulation, and what are the functions of grDC-associated MMP9 needs further examination.

CRAMP (mouse cathelicidin) production by grDCs is of great interest in the present study. Cathelicidin is the major class of antimicrobial peptides, which can interact with anionic components of microorganisms. Cathelicidin molecules are inserted into the membrane of microbes, leading to pores formation and bacteria death (Pestonjamasp, et al., 2001). The production of cathelicidins has been found in granulocytes, keratinocytes, epithelia cells and mast cells (Gudmundsson et al., 1996; Turner et al., 1998; Frohm et al.,

1997; Di Nardo et al., 2008; Ménard et al., 2008); however, there is no report about DCs

85 producing cathelicidins in the literature. As mentioned above, cathelicidins are synthesized and stored within secondary granules, which already exist in band cells

(Soehnlein, 2009), and grDCs may retain some secondary granules after developing from band cells. The CRAMP produced by grDCs makes contribution to the bacterial killing capacity. Intracellular E.coli are killed much more rapidly by grDCs than cDCs, which do not express CRAMP. The cell extracts of grDCs kill E.coli more effectively as well. The grDCs from CRAMP KO mice show diminished intracellular bacterial killing and anti-

CRAMP block antibody also decreases the bacterial killing activity of grDC extracts.

Band cells exhibit a potent endocytic potential, and grDCs preserve this property. We show that grDCs internalize all tested probes (dextran, latex beads and live bacteria) more efficiently than do cDCs, indicating they are optimally equipped with the endocytosis machinery. Since grDCs engulf more bacteria than do cDCs, it is reasonable that grDCs can be loaded with more bacteria-associated antigens. Indeed, compared to cDCs, grDCs induce more robust antigen specific T cell activation after uptake bacteria in the inflamed sites.

The CRAMP production, efficient endocytic potential and potent capacity to present bacterial antigen to T cells suggest that grDCs may play an important role in clearance of invaded bacteria and initiate the adaptive immune responses against bacterial infection.

The present study suggests an alternative pathway for DC development under the inflammatory conditions, in which band cells give rise to DCs, and also unveils a novel

86 member of the DC subset family, grDC. Inflammation recruits abundant band cells to the inflamed tissues and provides an environment for band cells to differentiate into grDCs.

The discovery of the present study reveals that the innate immunity and adaptive immunity synchronize closely during inflammation, and grDCs may be a direct bridge connecting them.

Because of their efficient antigen uptake and potent APC capacity, grDCs can be a good tool for vaccination. They may be easily loaded with target antigens and induced stronger

CD4 and CD8 T cell responses. Especially, compared to other DC precursors which are always in a rare number and difficult to isolate, band cells are a relatively large number and are refreshed frequently in human body (Summers et al., 2010). Therefore, it may be more effortless to acquire enough band cells to generate grDCs in vitro for clinical usage.

By monitoring the frequency of grDCs, we may use this population as a diagnostic marker for bacterial infections or inflammatory diseases. Therefore, we are able to estimate the progress of diseases and therapies. Furthermore, we may develop methodologies to regulate the number of grDC population, and thus we can expand or inhibit their function as needed with different conditions.

We hope this study opens up a new field of immunology research. Many questions still need to be answered, which may be the topics of the following works.

We have wondered whether some cytokines other than GM-CSF are more effective to promote the differentiation of grDCs from band cells. Since the library of a variety of

87 cytokines is available, we can add them individually into the BM cultures and analyze the frequency of CD11c+/Ly6G+ grDCs by FACS and compare to the effect of the BM cultured with GM-CSF.

To further study the in vivo functions of grDCs, we need to construct a system to selectively eliminate CD11c+/Ly6G+ population. To achieve this, we will breed the mouse expressing Cre recombinase under the control of CD11c promoter, with the Ly6G- loxP-flanked-STOP cassette-DTR-EGFP mouse. In F1 generation, only CD11c+/Ly6G+ grDCs express DTR and generate green fluorescent signal. Thus, we can conditionally deplete grDC population by injecting DT. This mouse model may also provide a tool to visualize the migration and behavior of grDCs in vivo by using confocal microscopy.

Antigen specific CD4 T cells can be efficiently activated by grDCs, but it is not clear whether Th1 or Th2 differentiation is polarized. To answer this question, we will pulse grDCs with OVA protein or bacteria expressing OVA, and then co-culture them with

CD4 T cells purified from OT II mice. By measuring the secretion of cytokines with

ELISA, we may tell the polarization of T cells mediated by grDCs. In addition, we may employ the peritonitis model, and inject bacteria expressing OVA to the inflamed peritoneal cavity, then purify grDCs in PECs and perform similar experiments.

The experiments performed in our lab have demonstrated that band cells purified from human BM can give rise to DCs when cultured in vitro (data not shown). Different from

Oehler’s work, the band cell-derived DCs generated in our lab retained some features of

88 granulocytes, like the expression of CD15. This may be because the blood samples used in Oehler’s study were collected from patients with chronic myeloid leukemia, pneumonia, osteomyelitis, non-Hodgkin lymphoma or from individuals treated with G-

CSF (Oehler et al., 1998), while we used BM samples from healthy donors. In future studies, we need to indentify the grDC equivalent population in human tissues and find their functions in steady state as well as in diseases.

89 SUMMARY

1. DsRed+/CD11b+/CD11c– band cells emerge in BM cultures from pIL1-DsRed TG

mice in the presence of GM-CSF.

2. Band cells differentiate into a novel Ly6G+ DC subset, termed “grDC”.

3. The gene expression profiles are different between grDCs and cDCs.

4. Compared to cDCs, grDCs kills bacteria more efficiently due to the production of

CRAMP.

5. The grDC population is found in normal tissues and their numbers increase under

inflammatory conditions.

6. Compared to cDCs, grDCs exhibit a strong capacity to uptake and present

bacterial antigens to T cells.

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