Dendritic Cell Development and Function

Washington University in St. Louis Washington University Open Scholarship

Arts & Sciences Electronic Theses and Dissertations Arts & Sciences

Spring 5-15-2020

Vivek Durai Washington University in St. Louis

Follow this and additional works at: https://openscholarship.wustl.edu/art_sci_etds

Part of the Allergy and Immunology Commons, Genetics Commons, Immunology and Infectious Disease Commons, Medical Immunology Commons, and the Molecular Biology Commons

Recommended Citation Durai, Vivek, "Dendritic Cell Development and Function" (2020). Arts & Sciences Electronic Theses and Dissertations. 2179. https://openscholarship.wustl.edu/art_sci_etds/2179

This Dissertation is brought to you for free and open access by the Arts & Sciences at Washington University Open Scholarship. It has been accepted for inclusion in Arts & Sciences Electronic Theses and Dissertations by an authorized administrator of Washington University Open Scholarship. For more information, please contact [email protected]. WASHINGTON UNIVERSITY IN ST. LOUIS

Division of Biology and Biomedical Sciences Immunology

Dissertation Examination Committee: Kenneth M. Murphy, Chair Brian Edelson Takeshi Egawa Daved Fremont Chyi-Song Hsieh

Dendritic Cell Development and Function

Vivek Durai

A dissertation presented to The Graduate School of Washington University in partial fulfillment of the requirements for the degree of Doctor of Philosophy

May 2020 St. Louis, Missouri

Table of Contents List of Figures ...... iv List of Tables ...... vii Acknowledgments ...... viii Abstract ...... ix Chapter 1: Introduction to Dendritic Cells ...... 1 1.1 Abstract ...... 2 1.2 Dendritic cells and the immune response ...... 3 1.3 Mouse models for studying dendritic cells ...... 5 1.4 Functions of dendritic cells ...... 17 1.5 Conclusions ...... 31 1.6 Author contributions ...... 32 1.7 References ...... 32 Chapter 2: Altered compensatory cytokine signaling underlies the discrepancy between Flt3–/– and Flt3l–/– mice ...... 59 2.1 Abstract ...... 60 2.2 Introduction ...... 60 2.3 Results ...... 63 2.4 Discussion ...... 72 2.5 Materials and Methods ...... 74 2.6 Acknowledgements ...... 82 2.7 Author Contributions ...... 82 2.8 References ...... 83 Chapter 3: Cryptic activation of an Irf8 enhancer governs cDC1 fate specification ...... 110 3.1 Abstract ...... 111 3.2 Introduction ...... 112 3.3 Results ...... 114 3.4 Discussion ...... 123 3.5 Materials and Methods ...... 126 3.6 Acknowledgements ...... 140

3.7 Author Contributions ...... 141 3.8 References ...... 142 Chapter 4: Discussion and Future Directions ...... 176 4.1 Abstract ...... 177 4.2 The role of supportive cytokines in hematopoiesis ...... 178 4.3 Evolving enhancer landscapes in hematopoiesis ...... 179 4.4 Future Directions ...... 180 4.5 References ...... 181

iii

List of Figures Figure 1.1: Functions of dendritic cell subsets in the immune response……………………...…58

Figure 2.1: Flt3l–/– mice have a more severe DC deficiency than Flt3–/– mice at two weeks of age ……………………………………………………………………………………………………88

Figure 2.2: DC deficiency in skin-draining lymph nodes and lungs is more severe in Flt3l–/– mice than in Flt3–/– mice…………………………………………………………………………….…89

Figure 2.3: Flt3l–/– mice continue to have a more severe DC deficiency than Flt3–/– mice at eight weeks of age……………………………………………………………………………………...91

Figure 2.4: Flt3L fails to generate DCs in Flt3–/– mice………………………………………….93

Figure 2.5: Flt3–/– DCs are transcriptionally and functionally similar to WT DCs…………...…95

Figure 2.6: Flt3–/– bone marrow generates mature DCs in response to M-CSF or SCF and contains committed DC progenitors……………………………………………………………..97

Figure 2.7: cDCs from M-CSF and SCF cultures are phenotypically similar to cDCs from Flt3L cultures…………………………………………………………………………………………...99

Figure 2.8: M-CSF and SCF support pDC development from Flt3–/– bone marrow…………...100

Figure 2.9: Committed DC progenitors can mature in response to M-CSF and SCF………….101

Figure 2.10: cDC development in Flt3–/– mice is dependent upon CSF1R…………………….103

Figure 2.11: Flt3–/– and Flt3l–/– BM progenitors and cDCs express normal levels of CSF1R and c-Kit.… ………………………………………………………………………………………...105

Figure 2.12: Committed DC progenitors arise in Flt3l–/– bone marrow and do not display increased sensitivity to M-CSF relative to WT progenitors…….……………………………...106

Figure 2.13: Flt3–/– DC progenitors display increased sensitivity to M-CSF...... 107

Figure 2.14: Deletion of Flt3 rescues the severe DC defect in Flt3l–/– mice…………………...108

Figure 2.15: Deletion of Flt3 in Flt3l–/– mice reverses the severe DC defect in skin-draining lymph nodes and lungs………………………………………………………………………….109

Figure 3.1: The +32 kb Irf8 enhancer is required for cDC1 development...... 148

Figure 3.2: AICEs in the +32 kb Irf8 enhancer drive cDC1 development……………………..150

Figure 3.3: Loss of the +32 kb Irf8 enhancer does not impact non-cDC1 immune lineages…..152

Figure 3.4: The +32 kb Irf8 enhancer is required for compensatory cDC1 development but not for cDC1 specification………………………………………………………………………….154

Figure 3.5: The +32 kb Irf8 enhancer is required for all compensatory cDC1 development…..156

Figure 3.6: CD226 uniquely identifies pre-cDC1s in the bone marrow………………………..158

Figure 3.7: The -50 kb Irf8 enhancer is not required for dendritic cell development…………..160

Figure 3.8: Loss of the -50 kb Irf8 enhancer does not impact non-monocyte/macrophage immune lineages…………………………………………………………………………………………162

Figure 3.9: The -50 kb Irf8 enhancer controls IRF8 expression in monocytes and macrophages……………………………………………………………………………………164

Figure 3.10: ATAC-seq identifies the +41 kb Irf8 enhancer as transiently active in cDC1 progenitors……………………………………………………………………………………...166

Figure 3.11: The +41 kb Irf8 enhancer is required for cDC1 specification and development…168

Figure 3.12: Loss of the +41 kb Irf8 enhancer does not impact non-DC immune lineages……170

Figure 3.13: E proteins are required for cDC1 development…………………………………...172

Figure 3.14: Tcf3–/– BM progenitors have reduced cDC1 and pDC potential………………….174

List of Tables Table 1.1: Comparison of DTA/DTR based mouse models for dendritic cell depletion………..51 Table 1.2: Comparison of Cre strains for conditional deletion of genes in dendritic cells……...54 Table 1.3: Comparison of transcription facot knockout mice that affect dendritic cell development…………………………………………………………………………56

Acknowledgments

This thesis was only completed with the support of my family, friends, and colleagues. First, I would like to thank Dr. Ken Murphy, who has provided me with excellent mentorship throughout the course of my doctoral work. He allowed me to pursue the projects I was truly interested in despite my inexperience and supported me through all the ups and downs inherent in science. This has allowed me to truly develop my own independent skills in research and to satisfy my intellectual curiosity to its fullest. I would also like to thank Dr. Theresa Murphy who taught me almost every technical skill I now possess. Discussions with her also helped to sharpen my hypotheses and models. The members of my thesis committees, Drs. Takeshi Egawa, Deepta Bhattacharya, Marco Colonna, Daved Fremont, Chyi-Song Hsieh, and Brian Edelson, have also provided invaluable support and guidance for all of my projects.

The many members of the Murphy lab that I have interacted with throughout the course of my doctoral work have also been treasured friends and collaborators. I owe them all my gratitude, but in particular I would like to thank Prachi Bagadia, Carlos Briseno, Jesse Davidson IV, Arifumi Iwata, Renee Wu, Derek Theisen, Wumesh KC, Tiantian Liu, and Marco Gargaro. I would also like to thank many collaborators outside the Murphy lab who have been critical in my work, including Jeffrey Granja, Ansuman Satpathy, Devesha Kulkarni, Swapneel Patel, Saran Raju, Miriam Wohner, Hiromi Tagoh, and Jeffrey Ward. I have tried to acknowledge other collaborators appropriately within each chapter.

Finally, I would like to thank the most important people in my life, my parents, my brother Naren Durai, and my wife Sruti. I would also like to acknowledge my friends Pavan Surti, Mark Zacharjasz, Arthur Zacharjasz, Jerry Liu, Vidit Sharma, Ravi Gottumukkala, Hannan Qureshi, and Ali Habib. Without the support of these many people throughout my life I would have accomplished nothing, and any achievements I have are as much theirs as mine.

vii

This work was supported by the Howard Hughes Medical Institute (to K.M.) and the National Institutes of Health (F30DK108498 to V.D.)

Vivek Durai

Washington University in St. Louis

May 2020

viii

ABSTRACT OF THE DISSERTATION

Dendritic Cell Development and Function

Vivek Durai

Doctor of Philosophy in Biology and Biomedical Sciences

Immunology

Washington University in St. Louis, 2020

Professor Kenneth M. Murphy, Chair

Dendritic cells (DCs) are a group of immune cells that include both classical dendritic cells (cDCs) and plasmacytoid dendritic cells (pDCs). cDCs are further comprised of two distinct subsets, cDC1s and cDC2s, which play critical roles in the initiation of innate and adaptive immune responses. Understanding how these lineages develop and function is therefore paramount. All DCs require the receptor tyrosine kinase Flt3 and its ligand Flt3L for their development, but the loss of Flt3L in mice leads to a more severe DC deficiency than does the loss of Flt3. This has led to speculation that Flt3L can bind to an alternate receptor that also supports DC development. However, we found that Flt3L administration to Flt3-/- mice does not generate DCs, arguing against a second receptor. Instead, Flt3-/- DC progenitors matured in response to macrophage colony-stimulating factor (M-CSF) or stem cell factor, and deletion of

Csf1r in Flt3-/- mice further reduced DC development, indicating that these cytokines could compensate for Flt3. Surprisingly, Flt3-/- DC progenitors displayed enhanced M-CSF signaling, suggesting that loss of Flt3 increased responsiveness to other cytokines. In agreement, deletion of Flt3 in Flt3l-/- mice paradoxically rescued their severe DC deficiency. We therefore conclude

ix that cytokines play a supportive role in DC development, and that the discrepancy between Flt3-/- and Flt3l-/- mice results from the increased sensitivity of Flt3-/- progenitors to these supportive cytokines.

The specification of immune lineages such as DCs during hematopoiesis is dependent upon the precise induction of lineage-determining transcription factors in multipotent progenitors, such as the induction of Irf8 in the common dendritic cell progenitor (CDP) that drives cDC1 development. To uncover the mechanisms regulating this induction, we identified

Irf8 enhancers via chromatin profiling of DCs and used CRISPR/Cas9 genome editing to test their functions in vivo. A +32 kb Irf8 enhancer active in mature cDC1s was required for the development and anti-tumor functions of this lineage, but was not required for its specification.

To identify enhancers controlling cDC1 specification, we performed ATAC-seq on DC progenitors. Unexpectedly, a +41 kb Irf8 enhancer, which binds E2A and is active in pDCs, was transiently accessible in cDC1 progenitors but not in mature cDC1s. Deleting this enhancer reduced Irf8 expression in pDCs, as expected, but also surprisingly prevented Irf8 induction in

CDPs and completely abolished cDC1 specification. We thus found that cryptic activation of the

+41 kb Irf8 enhancer was responsible for specifying cDC1 fate. Collectively, our studies have provided insight into the transcriptional mechanisms regulating the development of DCs.

Manipulation of these mechanisms in humans may prove useful in enhancing cytotoxic immune responses against viruses and cancer.

CHAPTER 1:

Introduction to Dendritic Cells

Contents of this chapter have been previously published in Immunity.

Durai V, Murphy KM. Functions of Murine Dendritic Cells. Immunity. 2016 Oct 18;45(4):719-

736.

1.1 Abstract

Dendritic cells (DCs) play critical roles in mediating innate and adaptive immune responses. The functions of DCs were originally obscured by their overlap with other mononuclear phagocytes, but new mouse models have allowed for the selective ablation of subsets of DCs and have helped to identify their non-redundant roles in the immune system.

These tools have elucidated the functions of DCs in host defense against pathogens, autoimmunity, and cancer. They have also provided insight into how DCs develop from their multipotent hematopoietic precursors. In this dissertation, we first approached the problem of promiscuity between cytokines and their receptors in regulating DC development. We found that the less severe DC deficiency found in Flt3–/– mice as compared to Flt3l–/– mice was due to the enhanced responsiveness of Flt3–/– progenitors to compensatory cytokines that could also support

DC development. Second, we studied the roles of enhancers in the Irf8 locus in the induction of this critical lineage-determining transcription factor in multipotent progenitors in order to specify classical type I dendritic cell (cDC1) fate. A +32 kb Irf8 enhancer was required for the maturation of cDC1s, but not for their specification. Instead, a +41 kb Irf8 enhancer that was only active in mature pDCs and was also cryptically activated in cDC1 progenitors was required for the specification of this lineage. Analysis of this enhancer found that it bound to E proteins, and consistently E2A-deficient progenitors failed to generate cDC1s. Our analysis of cryptic enhancer activity therefore revealed previously unrecognized transcriptional networks involved in specifying cDC1 fate. Collectively, these advances have provided insight into how DCs differentiate and the mechanisms utilized during lineage specification in hematopoietic precursors.

1.2 Dendritic cells and the immune response

The vertebrate immune system has evolved the remarkable capacity to robustly and precisely eliminate the wide variety of potential threats it encounters, from single cell bacteria to multicellular parasites to even transformed oncogenic versions of its own cellular components.

To achieve this goal, many diverse lineages of effector cells must act together in different capacities throughout the course of the immune response. As with any system possessing such complexity, the careful control and coordination of the numerous components of the immune system is critical for its proper functioning. As our understanding of each cell type acting within this system has grown, it has become increasingly apparent that dendritic cells (DCs) act as the central regulators of the entire immune response, responsible both for sensing the nature of the threats faced and for activating the precise combination of effectors required to eradicate them.

First isolated by Ralph Steinman and Zanvil Cohn, DCs were identified by their stellate morphology and capacity to stimulate naïve T cells 1-3. DCs comprise two major branches, the classical DCs (cDCs) identified by Steinman and the lymphocyte-like plasmacytoid DCs (pDCs) that produce Type 1 interferon in response to pathogens 4-7. cDCs can be further divided into two major subsets recently renamed cDC1s and cDC2s 8. All DCs originate from bone marrow (BM) progenitors arising from hematopoietic stem cells, starting with the macrophage/dendritic cell progenitor (MDP) 9,10, which gives rise to the common dendritic cell progenitor (CDP) 11,12, which finally gives rise to committed progenitors for each branch of DC such as the pre-cDC1 and the pre-cDC2 13,14.

cDCs express the integrin CD11c and MHC class II 15,16, and each subset can be distinguished by additional markers. Resident cDC1s in the spleen and lymph nodes (LNs)

3 express CD8α, CD24, and XCR1, while cDC2s express CD4 and Sirpα 17,18. In nonlymphoid tissues, all cDCs express CD24, which distinguishes them from macrophages that instead express

CD64 19-21. Nonlymphoid tissue cDC1s also express XCR1 and CD103, while cDC2s express

CD11b and Sirpα. Migratory cDCs that traffic from nonlymphoid tissues to LNs express these same markers they expressed in the periphery. There are several exceptions to these rules, however, such as CD11b+ cDC2s in the small intestine that comprise both CD103+ and CD103- fractions 22,23. While these varied markers have historically been used to identify cDC subsets, a recent analysis suggests that a more simple and consistent identification of these cells across most tissues is possible by gating cDCs as CD11c+MHCII+CD26+CD64-F4/80-, and within this population cDC1s as XCR1+ and cDC2s as Sirpα+ 24. pDCs also express CD11c and MHCII, but can be segregated by their additional expression of B220, Siglec-H, and Bst2 25,26.

While cDCs were discovered for their ability to serve as potent antigen-presenting cells

(APCs), it is now clear that they also have non-redundant roles in innate immune responses 23,27.

Their early recognition of pathogens and rapid cytokine production activates innate immune cells such as innate lymphoid cells (ILCs) and natural killer (NK) cells to limit pathogen spread until adaptive immunity can be initiated. Indeed, the heterogeneity of cDCs can itself be viewed as an evolutionary adaptation for the coordinated activation of the specific innate and adaptive effector responses best suited to control various forms of pathogens 28. cDC1s, for example, recognize intracellular pathogens and initiate type 1 immune responses that require the early activation of

ILC1s and NK cells as well as eventual Th1 polarization 27. Some cDC2s, on the other hand, govern type 2 immune responses against parasites in which they activate ILC2s and Th2 cells 29.

Other cDC2s sense extracellular bacteria and initiate type 3 immune responses by activating

ILC3s and Th17 cells 23,30. In this way, each cDC subset acts as the gatekeeper for a specific

4 module of the immune response, recognizing a particular type of threat and activating both the innate and adaptive defenses best suited for overcoming it.

Since the time of their discovery, there has been considerable debate over whether DCs possess unique functions or whether they are redundant with macrophages and other mononuclear cells 31. Fortunately, increasingly sophisticated mouse models for the specific ablation of DCs or for the conditional deletion of genes in these cells have helped resolve some of these controversies, revealing how immune responses are compromised in the absence of DCs and thus the roles these cells play in the normal response. These mouse models have been critical for elucidating the unique functions of DCs and resolving previously ambiguous conclusions. This dissertation will first provide an overview of the mouse models that have been developed to interrogate DC function and how these models have progressively clarified the role of DCs in various types of immune responses. As recent reviews have discussed DC development 17,18, this overview will focus primarily on how DCs function in immune responses to pathogens, autoimmunity, and cancer.

1.3 Mouse models for studying dendritic cells

Numerous models of constitutive and inducible DC depletion have been generated and used to identify the specific functions of DC subsets. In this section, we will first describe the different strains developed, their specificity, and their limitations. We will organize this by categories of DTR/DTA based models, Cre strains, and transcription factor knockout mice. In the next section we will discuss the discoveries made with these models regarding the functions of

DCs in various immune responses.

Depletion of dendritic cells by DTR/DTA systems

The first models of genetic ablation of cell lineages were transgenic mice in which cell type-specific promoters drove expression of the diphtheria toxin (DT) A-chain 32,33. This toxin disrupts protein translation by catalyzing the ADP-ribosylation of polypeptide chain elongation factor 2 and eventually leads to cell death 34-36. While DT efficiently ablates the cells in which it is expressed, it can be problematic as even low levels of off-target expression can lead to death of unintended cells or even to effects on embryogenesis or morphogenesis 37. Later mouse models overcame this problem by expressing the human or simian DT receptor (DTR) under the control of a cell type-specific promoter, with subsequent administration of DT to these mice 38.

As the mouse ortholog of the DTR is orders of magnitude less sensitive to DT 39,40, this allows for the efficient depletion of only DTR-expressing cells and has the added benefit of allowing for inducible depletion of these target cells rather than constitutive ablation.

The first DTR based model used for DCs was the Itgax-DTR strain, a transgenic line expressing a DTR-GFP fusion protein under the control of the murine promoter for the gene

Itgax, which codes for the integrin CD11c 41. DT administration completely depleted CD11c+

DCs within 24 hours, with no observable depletion of splenic B cells or F4/80+ macrophages.

DCs began to reappear 3 days after DT treatment. While this strain was vital for early work confirming the functions of DCs in T cell priming and pathogen responses, several limitations have emerged. First, repeated administration of DT led to death in these mice, restricting the duration through which depletion could be studied 41. This lethality was likely due to off-target expression of the Itgax-DTR transgene in radioresistant or non-hematopoietic tissues, since BM chimeras of WT BM into Itgax-DTR recipients also died after repeated DT treatment. This limitation can be overcome by generating chimeras of Itgax-DTR BM into WT recipients, which

6 tolerate repeated DT treatment 42. Another caveat derives from CD11c expression by non-DCs and the depletion by DT of such cells, which include splenic metallophilic and marginal zone macrophages, LN sinusoidal macrophages 43, alveolar macrophages 44, activated CD8 T cells 41, and plasma cells 45. Ablation of these cells complicates analysis with this strain, since phenotypes observed may result from their depletion rather than that of DCs. Some CD11c+ cells, notably pDCs, are not depleted in this model, indicating that the level of CD11c expressed by cells influences whether they are depleted 46.

A similar DTR strain, Itgax-DOG 47, is a BAC transgenic line in which the Itgax promoter drives expression of a fusion protein composed of DTR, a portion of ovalbumin, and

GFP (DOG). Continuous DT treatment in these mice does not cause lethality, perhaps due to more faithful expression of the BAC compared with the promoter-based transgene. Depletion of splenic macrophages was also observed after DT treatment in these mice, but whether CD8 T cells and plasma cells were similarly affected has not been evaluated.

The recently developed Zbtb46-DTR mouse allows for more specific depletion of cDCs

48. This strain has sequences for an internal ribosome entry site (IRES) and a DTR-mCherry fusion protein inserted into the 3’ untranslated region (UTR) of the endogenous Zbtb46 gene, which codes for the zinc finger transcription factor Zbtb46 (also known as zDC) whose expression in hematopoietic lineages is restricted to cDCs. However, a single administration of

DT into Zbtb46-DTR mice or into chimeras of WT BM into Zbtb46-DTR recipients proved lethal, indicating zDC expression in vital radioresistant or non-hematopoietic cells. Further work indicated these vital cells may be endothelial cells 49. DT treatment of Zbtb46-DTR BM into WT recipient chimeras depleted cDCs alone without ablation of B cells, T cells, neutrophils, natural killer cells, pDCs, or monocytes. This strain therefore allows for the specific and complete

7 deletion of cDCs, with the caveat that BM chimeras must be used. To circumvent this need, the

Zbtb46-LSL-DTR mouse was generated 50, in which the IRES-mCherry-DTR is preceded by a loxP flanked transcriptional Stop cassette (LSL) that terminates transcription of the locus before the DTR. Transcription of the DTR proceeds only after Cre-mediated excision of the Stop cassette. After crossing the Zbtb46-LSL-DTR strain to a Csf1r-cre strain 50, DT administration specifically depleted cDCs and did not lead to lethality. This model allows for the continuous depletion of cDCs for at least 4 weeks without the need for BM chimeras.

Several DTR strains allow for depletion of specific DC subsets. The first were two strains that expressed DTR from the endogenous Cd207 gene that codes for langerin, a C-type lectin expressed on epidermal Langerhans cells and dermal cDC1s. One strain was generated by inserting an IRES DTR-egfp into the 3’ UTR of the Cd207 gene 51, while in the second strain the first Cd207 coding exon was replaced with a DTR-egfp cassette 52. In both strains, DT treatment depletes Langerhans cells as well as cDC1s in skin-draining LNs. Additionally, a BAC transgenic CD207-DTA mouse strain was generated using a BAC in which an IRES DTA was inserted into the 3’ UTR of the human CD207 gene that codes for langerin 53. This strain has constitutive ablation of Langerhans cells but not of cDC1s in skin-draining LNs, suggesting that the human CD207 promoter is controlled differently from its murine equivalent. It was later demonstrated that these mice also lack CD103+CD11b+ cDC2s in the intestinal lamina propria 54.

The Ly75-DTR strain was generated by inserting an IRES DTR-eGFP into the 3’ UTR of the Ly75 gene that codes for CD205, an endocytic type I C-type lectin-like receptor also known as DEC-205 55. DT administration to this mouse also resulted in lethality, necessitating the use of

Ly75-DTR into WT recipient BM chimeras. DT treatment of such chimeras depleted CD205+ cDCs, the majority of which were CD8α+ cDC1s. CD205 is also expressed on B cells at 10-50

8 fold lower levels than on BM derived DCs 56 and accordingly a minor decrease in splenic B cells was observed upon DT treatment to Ly75-DTR mice. However, CD205 is also expressed at much higher levels in germinal center (GC) B cells 57 as well as all migratory cDCs and

Langerhans cells 58, but it was not determined whether DT treatment also depleted these cells.

The Clec9a-DTR BAC transgenic mouse, in which the first Clec9a coding exon in the

BAC was replaced with DTR, also allows for depletion of cDC1s 59. DT treatment completely ablates cDC1s, but also causes a ~50% decrease in pDCs, which may be due to the lower expression of Clec9a in mature pDCs 60,61. Clec9a is expressed in the common dendritic cell progenitor (CDP) 62, so continuous DT treatment may deplete this progenitor and therefore all

DCs over prolonged periods. Though this issue was not examined in the original study, at least one report indicated that cDC2s were unaffected after 15 days of DT treatment 63.

Two more recent DTR strains specifically deplete cDC1s. In the Xcr1-DTRvenus strain, the first Xcr1 coding exon was replaced by a cassette encoding a DTR-venus fusion protein 64.

XCR1 is a chemokine receptor uniquely expressed by cDC1s in humans, mice, and sheep 65,66.

DT treatment to this strain completely depleted cDC1s with no decrease in T cells, B cells, NK cells, granulocytes, monocytes, cDC2s, or pDCs. Expression of the Venus fluorescent protein was likewise restricted to cDC1s. This subset was depleted by 24 hours after DT treatment and began to recover on day 4 after treatment. A similar mouse strain, referred to as Karma, has an

IRES tdTomato-2A-DTR inserted into the 3’ UTR of the endogenous a530099j19rik gene 67, which like XCR1 is highly specific for cDC1s. Both tdTomato expression and cell ablation after

DT treatment were specific to cDC1s.

The Clec4a4-DTR strain, a BAC transgenic line in which an IRES DTR was inserted into the 3' UTR of the Clec4a4 gene on the BAC, allows for ablation of cDC2s 63. Clec4a4 (also known as DCIR2) is a C-type lectin expressed by cDC2s and recognized by the 33D1 antibody first used to define this subset 68,69. DT treatment in this strain severely reduces CD103+CD11b+ cDC2s in the colonic lamina propria and CD11b+ cDC2s in the mesenteric LNs, and partially depletes CD103-CD11b+ cDC2s and CD64+CX3CR1+ macrophages in the colonic lamina propria. Whether depletion of other cells occurs and whether cDC2s in other organs are similarly depleted remains to be determined. A second strain thought to deplete a subset of cDC2s is the

Mgl2-DTR strain, in which the first coding exon of the endogenous Mgl2 gene was replaced by a

DTR-gfp cassette 70. DT treatment in this strain depletes the Mgl2+ population of dermal cDC2s, but whether DCs in other tissues are also depleted remains to be analyzed.

Several DTR strains have been generated for pDC depletion. The first was the CLEC4C-

DTR BAC transgenic strain, which replaced the exons of the human gene CLEC4C in the BAC with the sequence for DTR 71. CLEC4C codes for BDCA-2, a C-type lectin uniquely expressed by human pDCs with no equivalent in mouse pDCs, but its human promoter remains active in mouse pDCs as DT treatment in CLEC4C-DTR mice specifically and completely depleted pDCs.

No reduction was observed in B cells, cDCs, T cells, NK cells, macrophages, monocytes, or neutrophils. Second, a Siglech-DTR strain was generated by knocking an IRES DTR-egfp sequence into the 3' UTR of the endogenous murine Siglech gene 72. Homozygous Siglech-DTR mice lacked expression of Siglec-H on pDCs, implying that the IRES DTR cassette somehow altered native Siglec-H expression. In this strain, DT administration completely depletes pDCs in the spleen, mesenteric LNs, and BM, with no apparent effect on cDCs. However, Siglec-H deficiency has been associated with altered cytokine responses 73 and perhaps abnormal kidney

10 and testes function 74, which may influence the phenotypes of this strain. Third, a Siglech-DTR

BAC transgenic strain was generated on the Balb/c background by replacing the coding exons of

Siglech in a BAC with the sequence for DTR 59. DT treatment in these mice completely depleted pDCs with no effect on cDCs. A separate BAC transgenic Siglech-DTR line on the C57BL/6 background was later generated, and analysis of both Siglech-DTR lines found that Siglec-H was also expressed by marginal zone macrophages and DT treatment ablated these cells as well as pDCs 73. This indicates that cell ablation in Siglech-DTR strains may not be limited to pDCs, complicating analysis with these lines.

Finally, several DTR based systems for the depletion of monocytes and macrophages

(reviewed in 75), have been generated that are driven by genes such as CCR2 and CX3CR1.

These have helped segregate the independent functions of monocytes and cDCs. One useful model for this purpose is the Cx3cr1-LSL-DTR strain 76, in which an LSL-DTR was inserted into the endogenous Cx3cr1 locus. This strain was crossed to Itgax-cre so that DT treatment specifically depletes CD11c+CX3CR1+ cells, which include CD103-CD11b+ cDC2s and monocyte-derived CD11b+ macrophages in the intestinal lamina propria 22. Another useful model is the MM-DTR strain, which was a cross between the Csf1r-LSL-DTR and LysM-cre strains, and in which DTR expression is limited to monocytes and monocyte-derived macrophages 77.

Table 1.1 provides a summary of DTR strains useful for the study of DC function.

An important caveat for some DTR strains is the reduced LN cellularity seen even without

DT treatment 78. In a cross between the Clec9a-cre strain and a Rosa26-LSL-DTR strain 79, reduced cellularity was observed in skin-draining and mesenteric LNs even without DT treatment.

There was also reduced cellularity in skin-draining LNs of Itgax-DTR mice, Cd207-DTR mice, and a cross between Itgax-cre and Rosa26-LSL-DTR mice. There was no decrease observed in

Itgax-DOG mice, so this phenomenon must be evaluated in each individual strain. The basis for this effect is unclear.

Cre strains for conditional deletion of genes in dendritic cells

The Cre-loxP system allows for conditional deletion of genes in specific cell lineages 80.

In this system, two 34-bp loxP sites are inserted on either side of a gene or exon, which is then said to be “floxed”. Cell type-specific promoters are then used to express the bacteriophage P1 cre gene, which encodes an integrase that mediates recombination between two adjacent loxP sites, leading to deletion of the intervening DNA and inactivation of the floxed gene in Cre+ cells.

The earliest Cre strains developed for cDCs were based on CD11c expression. First was an Itgax-cre BAC transgenic line in which the first coding exon of Itgax was replaced with cre

81. When crossed to the Rosa26-LSL-yfp strain 82, in which Cre-mediated excision of a STOP cassette allows YFP expression, splenic DCs were >95% labeled and pDCs were ~86% labeled.

However, background deletion of 5-12% was seen in splenic T, B, and NK cells. A later study also found deletion in 20-40% of blood monocytes, nearly ~100% of alveolar macrophages, 70% of splenic red pulp macrophages, 35% of marginal zone macrophages, and 20% of peritoneal macrophages 83. Therefore, as with Itgax-DTR strains, Itgax-cre is active in non-DC populations.

The second Itgax-cre strain is a transgenic line expressing cre-IRES-gfp under the control of the

Itgax promoter 84. GFP expression in this line was found to be uniformly high in cDC1s and cDC2s as well as in pDCs, but no expression was seen in T cells or B cells. Whether deletion of

12 genes occurred in cell types other than DCs, especially other CD11c+ populations such as macrophages, was not evaluated and remains to be resolved.

The Clec9a-cre strain was generated by replacing the first two exons of the endogenous

Clec9a gene with cre 62. Clec9a is first expressed at the CDP stage and is maintained in cDC1s and pDCs, but not in cDC2s 60-62. The Clec9a-cre strain achieved deletion in ~100% of cDC1s,

~50% of cDC2s, and ~20% of pDCs, suggesting that the transient expression of Cre at the CDP stage is insufficient for full deletion in cDC2 precursors and that pDCs do not express high enough levels of Cre for complete deletion. Deletion was not observed in any other cell types.

Recently a Zbtb46-cre line was produced by inserting an IRES-cre cassette into the 3’

UTR of the endogenous Zbtb46 gene, allowing for more specific deletion of genes in cDCs 85.

When crossed to Rosa26-LSL-yfp mice, the Zbtb46-cre achieved deletion in ~65% of cDCs, which was lower than the ~100% deletion with Itgax-cre, but also demonstrated <10% deletion in T cells, B cells, and monocytes, which was also consistently lower than the deletion seen with

Itgax-cre. Also <10% deletion was seen in red pulp macrophages, pDCs, and small intestinal macrophages, in contrast to the Itgax-cre strain where ~70% of red pulp macrophages and pDCs and ~100% of small intestinal lamina propria macrophages showed deletion. Zbtb46-cre also induced ~95% deletion of a floxed MHCII allele in cDCs, indicating that deletion of some alleles may be more complete than others. It was not evaluated whether Zbtb46-cre is as active in endothelial cells as the Zbtb46-DTR strain is.

The first Cre strain for targeting specific DC subsets was the Cd207-cre, in which a cre cassette was inserted into the second exon of the endogenous Cd207 gene 86. This strain achieved nearly complete deletion in Langerhans cells and langerin+ cDC1s in the dermis, skin-draining

LNs, and lung. A second Cre line useful for targeting particular DC subsets is the Xcr1-cre strain, in which the first exon of the endogenous Xcr1 gene was replaced by cre 87. This was crossed to

Rosa26-lacZbpAfloxDTA mice 88, in which Cre excises a loxP flanked lacZ-polyadenylation (bpA) sequence and allows for DTA expression and cell ablation. This cross resulted in nearly complete ablation of cDC1s in the spleen, MLN, and intestinal lamina propria, but no reduction in cDC2s in these organs. There was no reduction in CD4 or CD8 T cells in the spleen or MLN, but T cells and intraepithelial lymphocytes in the intestinal lamina propria were reduced, which was attributed to the absence of cDC1s. Finally, in the BAC transgenic Siglech-cre strain, the first exon of Siglech was replaced with a cre IRES mCherry cassette 89. However, this strain only achieved deletion in

~30% of pDCs and ~2% of all lymphoid cells, and so is better suited for lineage tracing than functional analysis.

Transcription factor based depletion of dendritic cells

Several transcription factors have been identified whose deletion selectively depletes specific subsets of DCs, and these knockout mice provide useful models for studying DC function. One of the earliest identified was the Irf8-/- strain, which lacks cDC1s 90,91, but also has impairments in B cells 92, monocytes 93, eosinophils 94, and basophils 95. Irf8-/- mice were also thought to lack pDCs 90, but more recently it was determined that pDCs do not require IRF8 for their development but instead that loss of this factor affects their expression of cell-surface markers and their ability to produce interferon 96. Irf8-/- mice therefore do not serve as a model of specific cDC1 depletion, but this can be overcome by crossing a floxed Irf8 allele to the Itgax-

14 cre strain 97, which depletes cDC1s and CD64+CD11b+ macrophages in the intestinal lamina propria, or to the Zbtb46-cre strain 98, which specifically depletes cDC1s.

Id2-/- mice lack cDC1s 99, but also have defects in NK cells 100 and ILCs 101. Nfil3-/- mice similarly lack cDC1s 102, NK cells 103, and ILCs 104,105. The similarity of Id2-/- and Nfil3-/- mice may result from an epistatic action of Nfil3 on the expression of Id2 during DC development, as has been suggested for NK cell development 106.

Batf3-/- mice are selectively deficient in cDC1s 107. Batf3 and the AP-1 transcription factor Jun form a heterodimer that interacts with IRF8 to bind AP-1/IRF consensus elements

(AICEs) 108,109. Batf3 acts to maintain the expression of IRF8 by autoactivation during development of cDC1s, without which cDC1s divert to cDC2 fate 13. Batf3-/- mice on the

129SvEv and Balb/C backgrounds have an almost complete lack of cDC1s in lymphoid and nonlymphoid tissues 110, but interestingly in the C57BL/6 background they retain cells resembling cDC1s in skin-draining LNs 108.

Few known transcription factors selectively control cDC2 development. Relb-/- mice have

DCs with reduced immunogenicity 111, and reportedly a partial cell-intrinsic defect in cDC2 development 112. RelB acts in the noncanonical NFκB signaling pathway activated by lymphotoxin-β receptor (LTβR) signaling, and Ltbr-/- mice also have a cell-intrinsic but partial reduction in CD4+ cDC2s 113. It has not been tested whether the impact of RelB deficiency on

DCs is secondary to deficient LTβR signaling. As Relb-/- mice develop severe multiorgan inflammation and hematopoietic abnormalities 114, and Ltbr-/- mice lack lymph nodes 115, neither strain is a convenient model for testing DC function. Conditional deletion of Ltbr with Itgax-cre has been used to overcome this limitation 116.

Other factors that influence cDC2 development include Notch2 and Klf4. Conditional deletion of the Notch cofactor Rbpj 81 or Notch2 23,30 using Itgax-cre demonstrated a specific depletion of CD11b+ESAM+ cDC2s in the spleen and CD103+CD11b+ cDC2s in the intestinal lamina propria and MLNs, without apparent loss of other cell types. Notch2f/f Itgax-cre mice did exhibit gene expression changes in remaining cDC1s and cDC2s, however, indicating possible effects of Notch2 in these lineages 23. Conditional deletion of the transcription factor Klf4 with

Itgax-cre also led to the depletion of subsets of cDC2s 29 similar to migratory CD24- CD11b- cDC2s in skin-draining LNs 117.

Irf4-/- mice do not lack cDC2s, but instead have partial impairments in the number or functioning of these cells. CD11b+ cDC2s are present in the skin of Irf4-/- mice, but fail to migrate to LNs 118,119. Similarly, conditional deletion of Irf4 with Itgax-cre resulted in a partial reduction in CD11b+ cDC2s in the lung but an almost complete absence of CD11b+ cDC2s in the lung draining LNs 19, as well as a partial reduction in CD103+CD11b+ cDC2s in the intestinal lamina propria and a more complete reduction in mesenteric LNs 19,120. Finally, conditional deletion of Irf4 with Itgax-cre depletes a PDL2+Mgl2+ population of cDC2s in skin-draining LNs but not the dermis 121. But although Irf4f/f Itgax-cre mice can serve as a useful model for cDC2 deficiency, IRF4 also plays a role in other CD11c+ cells such as M2 macrophages 122 and GM-

CSF activated monocytes 123, and these may also be affected in this strain.

Several transcription factors have been identified that regulate pDC development. The first model of specific pDC deficiency was conditional deletion of Tcf4, which encodes the factor E2-

2, with Itgax-cre 124. Recently, conditional deletion of the transcription factor Zeb2 using Itgax- cre also demonstrated selective pDC deficiency 125. Table 1.3 provides a summary of the transcription factor knockout mice useful for the study of DCs.

1.4 Functions of dendritic cells.

Since their discovery, cDCs were recognized as possessing a remarkable capacity to stimulate naïve T cells and to initiate adaptive immune responses 2,3. Subsequent work, based in large part on the models described above, has shown that DCs are also critical for - not just participants in - early innate immune responses, activating innate lymphoid cells (ILCs) and other cells involved in the immediate response to pathogens. Additionally, subsets of cDCs play varied roles in autoimmunity and responses to tumors. Below we discuss the studies that identified these functional aspects of DCs, what tools were used to arrive at these conclusions, and confounding factors that may require further resolution.

General adaptive immune responses

The first study to use Itgax-DTR mice showed that the absence of CD11c+ cells led to the loss of CD8 T cell priming to cell-associated antigens and to the intracellular pathogens Listeria monocytogenes and malaria 41. This provided the first direct in vivo evidence for the role of

CD11c+ cells in T cell priming. Subsequent use of the Itgax-DTR strain showed a requirement for CD11c+ cells in priming CD8 T cells against lymphocytic choriomeningitis virus (LCMV)

126, herpes simplex virus (HSV) 127, vesicular stomatitis virus (VSV) 128, and influenza 129,130, as well as in CD4 T cell priming against HSV 127, Mycobacterium tuberculosis (Mtb) 131, and immune complexes 132. Depletion of CD11c+ cells in Itgax-DTR mice also impaired the expansion of antigen-specific memory CD8 T cells during rechallenge with Listeria, VSV, or influenza 133 or with LCMV 134.

Later studies used additional models to validate the specific role of cDCs and not other

CD11c+ cells in T cell priming. Specific ablation of cDCs with the Zbtb46-DTR strain resulted in a complete inability to prime CD8 or CD4 T cells against soluble antigen 48, and also a failure to prime CD4 T cells against Mtb 135. Finally, cDC specific deletion of MHCII using Zbtb46-cre led to a complete reduction in CD4 T cell priming to soluble antigen 85.

Targeting antigen to specific DC subsets has indicated that there may be a specialization of roles between subsets, with cDC1s preferentially priming CD8 T cells and cDC2s priming

CD4 T cells 69. cDC1s have also been recognized as the cells most efficient in cross-presentation of exogenous antigens on MHC class I 136. Studies using mouse models with selective depletion of one subset have provided evidence to support this dichotomy. Batf3-/- mice failed to prime

CD8 T cells against cell-associated antigen or West Nile virus (WNV), but still had intact priming of CD4 T cells 107. Batf3-/- mice also mounted reduced CD8 T cell responses to cytomegalovirus (CMV) 137, influenza 138,139, cowpox virus 140, HSV 141,142, and malaria 143.

Depleting cDC1s using the Xcr1-DTRvenus strain also abrogated CD8 T cell priming against soluble and cell-associated antigen as well as against Listeria infection 64. CD4 T cells could still be primed against soluble antigen in these mice. A study using Karma mice similarly showed that depletion of cDC1s led to deficient CD8 T cell priming to soluble protein as well as failure to reactivate memory CD8 T cells in response to Listeria, vaccinia virus, and VSV 67. Taken together these studies suggest that cDCs alone are responsible for priming T cells, with a specific role for cDC1s in priming naïve CD8 T cells and activating memory CD8 T cells during recall responses. Future work with specific depletion of cDC2s is needed to confirm their unique role in priming CD4 T cells.

Recent work has also indicated that CD4 T cell help in CD8 T cell responses might be mediated through “licensing” of cDC1s. Analysis of immune responses suggests that CD4 T cells are primed by DCs earlier than CD8 T cells, and that later in the response clusters of these three cell types form during which CD4 T cells may "license" cDC1s in order to enhance priming of CD8 T cells 144,145. In agreement with this model, one study found that mixed bone marrow chimeric mice made of Xcr1-DTR:H2-Ab1-/- BM, in which following DT treatment all remaining cDC1s lack MHCII expression, generated fewer antigen-specific CD8 T cells upon vaccinia virus infection 145. MHCII expression in cDC1s may therefore serve as a mechanism by which they receive CD4 T cell help rather than for their direct priming of naïve CD4 T cells.

Several studies have implicated DCs in the induction of Tfh cells and the initiation of the germinal center (GC) response. Depletion of all CD11c+ cells by Itgax-DTR was found to diminish the number of Tfh cells induced in response to Toxoplasma infection 146. Specific cDC depletion with Zbtb46-DTR mice also abolished GC responses and antibody production against allogeneic red blood cells (RBCs) 147. This was attributed to the function of cDC2s, as Batf3-/- mice had intact antibody responses but Irf4f/f Itgax-cre mice did not 147. Another study found that CD25 expression by cDCs was crucial for Tfh induction 148. In this study, mixed BM chimera mice reconstituted with Zbtb46-DTR:Il2ra-/- BM, in which after DT treatment all remaining cDCs lack the IL-2 receptor alpha chain (CD25), failed to generate antibodies or GC B cells upon sheep RBC immunization. cDC2s in the spleen can induce expression of CD25, suggesting these may be the specific cells involved in this process 148. Finally, IgA class switching in Peyer’s patches was also found to be regulated by DCs 149. This study utilized mixed BM chimeras made from Itgax-

DTR:Ltbr-/- BM, in which following DT treatment all remaining DCs lack the LTb receptor and thus certain cDC2s fail to develop. After DT treatment, these chimeras had fewer IgA class-

19 switched GC B cells in Peyer’s patches. This appears to involve the activation of latent TGF-β by the integrin chains Itgβ8 and Itgαv on DCs and presentation of the active cytokine by DCs to B cells, as Itgax-cre mediated deletion of Itgb8 also resulted in reduced IgA+ GC B cells 149.

Type 1 immune responses

Type 1 immune responses are mounted against intracellular pathogens that require IFN-g activated macrophages and cytotoxic CD8 T cells for their clearance. Early nonspecific sources of

IFN-g are NK cells and ILC1s, while antigen-specific Th1 and CD8 T cells are responsible for producing this cytokine later in the immune response. IL-12 is critical for the activation of type 1 responses, as it induces NK cells and ILC1s to secrete IFN-g 150, and is also responsible for the polarization of naïve T cells to Th1 cells 151. Several studies have established that cDC1s are vital for mounting type 1 responses because of their non-redundant production of IL-12 and their ability to prime CD8 T cells.

The first indication that cDC1s are important in these responses was a study demonstrating that Irf8-/- mice had increased susceptibility to Toxoplamsa gondii infection and reduced serum IL-12 levels 152, though given the myriad defects in this mouse this could not be attributed specifically to cDC1s. Later, Itgax-DTR mice were used to show that CD11c+ cells were required for IL-12 and IFN-γ production and protection against Toxoplasma infection 153.

Finally, Batf3-/- mice that specifically lack cDC1s were found to be susceptible to Toxoplasma, demonstrating that these cells were indeed critical for resistance 27. Mixed chimeras made from

Batf3-/-:Il12a-/- BM, in which all cDC1s that develop are deficient in IL-12 production, were susceptible to Toxoplasma infection, indicating that cDC1s are the non-redundant source of IL-

12 necessary for resistance to infection 27. Additionally, depletion of cDC1s using the Karma strain abolished IL-12 production in response to soluble Toxoplasma antigen 67. Batf3-/- mice also have reduced IFN-γ production from NK cells during Toxoplasma infection, suggesting that IL-

12 from cDC1s is critical for NK cell activation in this response 154.

The role of cDCs in type 1 responses to several other intracellular pathogens has also been documented. During infection by Listeria, Itgax-DTR 155, Batf3-/- 156, Ly75-DTR 55, and

Xcr1-DTRvenus 64 mice all had reduced CD8 T cell responses. However, many of these strains also exhibited a reduced splenic Listeria burden and Batf3-/- mice actually demonstrated increased resistance to infection. This is because Listeria must infect cDC1s in the splenic marginal zone in order to spread to and proliferate in the lymphoid areas of the spleen 155, so

Batf3-/- mice lacking these cells show reduced susceptibility to infection. Finally, constitutive deletion of CD11c+ cells, achieved by crossing Itgax-cre mice to Rosa26-LSL-DTA mice 157, diminished IFN-γ production from NK cells, NKT cells, and T cells during Listeria infection 158.

Leishmania major, another intracellular pathogen, similarly requires cDC1s for its clearance, as Batf3-/- mice exhibit greater disease burden from infection 159 and generate fewer

Th1 cells 160. Another study used Batf3-/- and Cd207-DTR mice to demonstrate that cDC1s are required for the priming of CD8 T cells and Th1 cells against skin infection by the fungus

Candida albicans 161.

Several studies have implicated DCs in NK cell homeostasis. Depletion of CD11c+ cells in

Itgax-DTR mice diminished steady-state numbers of NK cells 162 and reduced their cytotoxicity and IFN- γ production in several infection models 127,163,164. NK cell survival and activation requires trans-presentation of IL-15, whereby a non-NK cell that expresses the non-signaling

21 receptor chain IL-15Rα binds IL-15 and presents to the full IL-15αβγ receptor on NK cells 165.

One study implicated a CD11c+ cell as the trans-presenting cell by using mixed BM chimera mice made from Itgax-DTR:Il15-/- BM or from Itgax-DTR:Il15rα-/- BM 166. In these chimeras, DT treatment causes all remaining CD11c+ cells to lack IL-15 or IL-15Rα, respectively, and after DT treatment these chimeras show severely reduced NK cell activation in vivo. Future studies are needed to clarify whether cDCs are involved in this activity, and if so which subsets.

Type 2 immune responses

Type 2 responses are carried out against multicellular parasites at barrier surfaces in order to aid in their expulsion. Many cytokines play key roles in this response. These include IL-25 and IL-33, which activate ILC2s to produce effector cytokines such as IL-4, IL-5, and IL-13 150, and IL-4, which polarizes naïve T cells to Th2 cells 167. Much remains unknown about the initiation of type 2 immune responses, and it has been controversial whether DCs are even involved directly.

An early study found that depletion of CD11c+ cells in Itgax-DTR mice did not impair

Th2 cell polarization in response to immunization with papain 168, although another study using the same strain found that it did 169. A later study also found that depletion of CD11c+ cells in

Itgax-DTR mice abrogated type 2 responses, this time to inhaled house dust mite (HDM) allergen 170. This study implicated an FcεRI expressing CD11c+MHCII+ cell population as being the principal APC responsible. Subsequent studies also found that CD11c+ cell depletion by

Itgax-DTR reduced numbers of Th2 cells after infections with the helminthes Schistosoma mansoni 171, Heligmosomoides polygyrus 172, and Nippostrongylus brasiliensis 173.

Other models have suggested putative Th2 inducing DC subsets. Two groups demonstrated that depletion of an Mgl2+ subset of cDC2s, either with the Mgl2-DTR strain 70 or by conditional deletion of Irf4 with Itgax-cre 121, diminished Th2 priming in response to papain and Nippostrongylus infection. Irf4f/f Itgax-cre mice also had reduced Th2 responses after HDM allergen challenge 174. Another study found that depletion of a subset of cDC2s in the Klf4f/f

Itgax-cre strain increased susceptibility to Schistosoma mansoni infection and diminished allergic inflammation after intransal HDM challenge 29. Also, a subset of CXCR5+ DCs, which was depleted by using Itgax-DTR:Cxcr5-/- BM chimeras, appeared to be required for Th2 responses to Heligmosomoides infection 175. Finally, Batf3-/- mice develop somewhat stronger

Th2 responses to helminth infection, suggesting that the IL-12 produced by cDC1s regulates Th2 polarization carried out by other DCs 176.

Cytokine production by DCs has not been established in any of these models, so whether they control type 2 responses by this mechanism or some other remains to be determined. Recent studies have suggested that ILC2s may be the critical source of cytokines acting to induce type 2 responses 177,178. Conceivably, ILC2s and DCs may cooperate in Th2 priming, but this area awaits further studies.

Type 3 immune responses

Type 3 immune responses at barrier surfaces such as the lungs and intestines control infections by extracellular bacteria and fungi and require several cytokines, including IL-23 and

IL-6. IL-6 and TGF-β initiate Th17 cell polarization 179, and IL-23 increases the survival and expansion of committed Th17 cells 180. IL-23 is also crucial in innate responses for activating

ILC3s to produce IL-22, which in turn promotes production of bactericidal lectins such as

RegIIIγ from small intestinal epithelial cells 150,181.

Citrobacter rodentium, a mouse pathogen used to study type 3 immune responses, requires IL-23 and IL-22 for its clearance 182-184. An early study that implicated cDCs in response to oral Citrobacter infection found that conditional deletion of Ltbr by Itgax-cre, which depleted a subset of cDC2s, reduced colonic IL-22 production and increased pathogen burden 116. Later studies demonstrated that depletion of all cDCs in Zbtb46-DTR mice 23,77 led to severe susceptibility to Citrobacter infection, as did depletion of cDC2s with Notch2f/f Itgax-cre mice 23.

Mixed BM chimeras reconstituted with Notch2f/f Itgax-cre:Il23a-/- BM were also severely susceptible to Citrobacter infection, indicating that Notch2-dependent cDC2s are the critical source of IL-23 in defense against this pathogen 23. NFκB signaling in CD11c+ cells is also critical, as mice with conditional deletion of Myd88 with Itgax-cre also showed susceptibility to

Citrobacter 185. Importantly, mice depleted of all cDCs or Notch2-dependent cDC2s die at ~day

10 after Citrobacter infection, while Rag2-/- mice lacking B and T cells die at ~day 30 after infection 182. This suggests that cDC2s are critical for activating innate defenses during

Citrobacter infection and not just for initiating adaptive immunity.

Several recent studies have suggested contributions from multiple cDC2 subsets in defense against Citrobacter. First, CD207-DTA mice, which lack intestinal CD103+CD11b+ cDC2s, were not found to be susceptible to Citrobacter infection 54. Thus, the lethality seen with

Notch2 deficiency may be due to an impact on the function of CD103-CD11b+ cDC2s, rather than the depletion of CD103+CD11b+ cDC2s. In accordance with this possibility, Cx3cr1-LSL-

DTR mice crossed to Itgax-cre, in which DT treatment depletes CD103-CD11b+CX3CR1+ cDC2s and CD11b+ macrophages, also showed susceptibility to Citrobacter and reduced IL-22

24 production 185. Finally, MM-DTR mice, which lack monocytes and monocyte-derived macrophages, did not display susceptibility to Citrobacter, suggesting that CD103-CD11b+ cDC2s and not CD11b+ macrophages are the critical cells for defense against this pathogen 77.

However, direct comparison of these different models may be required to eliminate confounding effects such as variable microbiota in the strains used in each study.

Studies have also implicated cDC2s in various other type 3 responses, but the reported mechanisms by which they act have varied. Several groups found that depletion of intestinal

CD103+CD11b+ cDC2s, using Notch2f/f Itgax-cre, Irf4f/f Itgax-cre, or CD207-DTA mice, resulted in fewer small intestinal Th17 cells at steady state 19,30,54. Another study using the Irf4f/f Itgax-cre strain similarly found reduced numbers of Th17 cells at steady state in the small intestine lamina propria and mesenteric lymph nodes, and also reduced Th17 polarization after immunization with antigen plus αCD40 and LPS 120. This study observed reduced levels of IL-6 in Irf4-/- DCs, which might explain the reduced Th17 priming. A second group found deficient Th17 priming in

Irf4f/f Itgax-cre mice after lung infection by the fungal pathogen Aspergillus fumigatus, which was attributed to the loss of CD11b+ cDC2s that normally express high levels of IL-23, IL-6, and

TGF-β 19.

In agreement, another study used BM chimeras to show that CD11c+ cells must produce

IL-6 and TGF-β to induce Th17 responses to Streptococcus pyogenes 186. They found that BM chimeras of Itgax-DTR:Il6-/- or Itgax-DTR:Tgfb1f/f BM, in which DT treatment causes all remaining DCs to be deficient in either IL-6 or TGF-β respectively, had reduced Th17 responses to this pathogen. This was confirmed in Tgfb1f/f Itgax-cre mice and specifically found to involve

Mgl2+ cDC2s, as DT treated Mgl2-DTR:Il6-/- mixed BM chimeric mice also had reduced Th17 induction upon Streptococcus pyogenes infection 186. A separate study used Mgl2-DTR:Il23a-/-

25 mixed BM chimeras to determine that Mgl2+ cDC2s must produce IL-23 in order to activate IL-

17 secretion from dermal γδ T cells during cutaneous Candida albicans infection 187. Finally, a study using the Il23af/f Itgax-cre strain concluded that IL-23 from CD11c+ cells was important for Th17 and Th1 induction in response to Helicobacter hepaticus infection 188, although the relative contributions of cDCs and macrophages in this infection remains to be defined with more specific methods. While depletion of cDC2s has revealed defects in type 3 responses to several pathogens, responses to many of these infections, specifically Citrobacter 23,

Streptococcus 186, and Candida 187,189, are intact in mice lacking cDC1s. This again underscores the functional specialization of different cDC subsets and the non-redundant roles they play in host defense.

Control of type 3 responses against segmented filamentous bacteria (SFB) may differ from those described above. SFB are a commensal organism present in some mice microbiota that induce an antigen-specific Th17 response in colonized mice 190. Conditional deletion of MHCII with Itgax-cre reduced Th17 induction after inoculation with SFB, which was interpreted to mean cDCs were responsible for priming against this organism. But Batf3-/-CD207-DTA mice, which lack cDC1s and CD103+CD11b+ cDC2s, and Flt3l-/- mice, which have substantially reduced numbers of all cDCs 191, had no impairment in Th17 priming 192,193. Instead, depletion of monocytes with the Ccr2-DTR strain abrogated Th17 induction against SFB, and this could be restored by transfer of wildtype monocytes, suggesting that monocyte-derived intestinal macrophages prime this response 194. This example illustrates how some conclusions regarding cDC function drawn from CD11c-based systems may need re-interpretation with follow-up studies using more precise genetic models.

Dendritic cells in autoimmunity

DCs are important for the induction of tolerance to self-antigens and for the pathogenicity of several autoimmune syndromes. Early studies found a relationship between DC and Treg numbers, and that depletion of Tregs with a Foxp3-DTR strain greatly expanded cDCs 195. This expansion was due to increased numbers of cDC progenitors and was dependent on Flt3L, a crucial growth factor for cDCs 196. Conversely, depletion of DCs in Itgax-DTR mice or in Flt3l-/- mice reduced splenic Treg numbers, while expansion of DCs by exogenous Flt3L increased Treg numbers 197. This phenomenon held for certain antigen-specific Treg clones as well 198.

DCs primarily control Treg numbers by regulating their proliferation rather than their induction, as Tregs transferred into Itgax-DTR mice treated with DT divided less than when transferred into WT mice 197. This control of proliferation required MHCII expression by DCs, as both Itgax-DTR:H2-Ab1-/- mixed BM chimeric mice treated with DT and H2-Ab1 f/- Itgax-cre mice also showed a decrease in Treg numbers and proliferation 197. Later work also concluded that DCs do not play a primary role in Treg induction in the thymus, as neither constitutive depletion of DCs in mice with Itgax-cre crossed to Rosa26-LSL-DTA nor H2-Ab1 f/f Itgax-cre mice had differences in the absolute number of Foxp3+ cells that develop in the thymus 199,200.

However, several recent studies have demonstrated that at least some Treg clones are induced in the thymus by DCs, especially clones that recognize self-antigens expressed by the transcription factor AIRE. One group identified certain T cell clones that convert to Tregs in the thymus in response to AIRE-dependent antigens in wildtype mice, but not in mice that were depleted of CD11c+ cells by crossing the Itgax-cre to the Rosa26-LSL-DTA strain 201. These same T cells also did not convert in Batf3-/- mice, suggesting that cDC1s controlled their

27 induction. A subsequent study reported that both polyclonal Tregs and a specific AIRE- dependent Treg were reduced in number in mice with conditional deletion of MHCII by Itgax- cre, but in contrast to the previous study found that this AIRE-dependent Treg still developed in

Batf3-/- mice 198. This suggests that the specific Treg clone or the nature of the antigen it recognizes may determine which cDC subset is responsible for its induction.

DCs also play a role in several models of autoimmunity. Studies using Itgax-DTR mice show that CD11c+ cells are required for pathogenesis in ovalbumin-driven asthma 44 and in Th2- mediated peanut food allergy 202. Conditional deletion of Myd88 with Itgax-cre also diminished colitis in Il10-/- mice 203 and decreased lupus-like symptoms in Lyn-/- mice 204, suggesting that

NFκB signaling in CD11c+ cells contributes to autoimmunity. Constitutively depleting CD11c+ cells in a model of lupus, achieved by crossing the Rosa26-LSL-DTA and Itgax-cre alleles onto the lupus-prone MRL.Faslpr background, reduced disease severity, in part by decreasing T and B cell activation 205. Disease incidence partly involved MyD88 signaling in CD11c+ cells, since

Myd88f/f Itgax-cre mice on this background also had reduced dermatitis and T cell activation, but not reduced nephritis 206. In the nonobese diabetic (NOD) model of murine diabetes, cDC1s are required for disease initiation, as Batf3-/- NOD mice are completely free of lymphocyte infiltration into and destruction of pancreatic islets 207. Conversely, increased activity of DCs can lead to autoimmunity, often from aberrant cytokine production or inappropriate activation of autoreactive T cells. This was seen when Itgax-cre was used to delete inhibitory signaling molecules such as Fas 84 β-catenin 208, FADD 209, A20 210, TRAF6 211, Lyn 212, and Caspase-8 213.

Recent work has also implicated cDCs in establishing oral tolerance to ingested antigens.

Using Zbtb46-DTR mice, one study demonstrated a loss of oral tolerance in mice lacking all cDCs

98. These mice failed to induce Tregs against orally fed antigen and generated an immune response

28 to subsequent peripheral antigen challenge. Oral tolerance was intact in Irf8f/f Zbtb46-cre mice that lack cDC1s, although antigen-specific Treg conversion was partially impaired. This suggests cDC1s are important in establishing oral tolerance but that other cDCs can compensate for their loss. A separate study found that for orally fed antigen to be tolerized it is phagocytosed by

CX3CR1+ macrophages in the intestinal lamina propria and subsequently transferred to

CD103+CD11b+ cDC2s through gap junctions made up of connexin-43 214. Conditional deletion of Gja1, which encodes connexin-43, with Itgax-cre reduced the number of Tregs induced to orally fed antigen and also diminished oral tolerance established to this same antigen. Also, conditional deletion of Itgb8, which encodes the integrin Itgβ8 that activates latent TGF-β, with Itgax-cre resulted in autoimmune inflammatory bowel disease and reduced numbers of colonic Tregs, suggesting that CD11c+ cells are critical sources of active TGF-β for the induction of Tregs in response to gut antigens 215.

Dendritic cells in tumor immunity

Recent work has expanded our understanding of the roles of DCs in antitumor immune responses. Studies using Itgax-DTR mice demonstrated that depletion of CD11c+ cells reduced

CD8 T cell responses against transferred tumor cells 216,217 and reduced survival in oncogene- driven tumor models 218. cDC1s in particular were critical for antitumor responses, as Batf3-/- mice failed to reject transplanted immunogenic fibrosarcomas 107 and failed to induce T cell infiltration into autochthonous oncogene-driven melanoma 219. cDC1s are also required for the antitumor effects of many cancer therapies, as Batf3-/- mice have reduced responses to therapeutic PD-L1 blockade, Flt3L injection, or polyI:C injection 220 as well as to anti-CD137 or

29 anti-PD1 therapy 221. cDC1s must migrate to LNs to activate tumor-specific CD8 T cells, as mixed BM chimeras made with Xcr1-DTR:Ccr7-/- BM, in which DT treatment causes all remaining cDC1s to lack CCR7, are not able to reject tumors 222. Finally, cDCs play a role in opposing the establishment of metastatic foci, as Zbtb46-DTR mice depleted of cDCs mice had increased numbers of melanoma metastases to the lung 223.

Two studies have demonstrated that interferon signaling in DCs is necessary for the initiation of antitumor immune responses. In the first, conditional deletion of the type 1 interferon receptor Ifnar1 with Itgax-cre prevented rejection of transplanted fibrosarcomas 224.

IFNAR deficient cDC1s also displayed deficient cross-presentation of antigens to CD8 T cells, suggesting that an inability to cross-present tumor antigens and activate CD8 T cells underlies the inability to reject tumors in this strain 224. In the second, Ifnar1-/-:Batf3-/- mixed BM chimeras failed to prime CD8 T cells to tumor antigens, suggesting that interferon signaling is specifically necessary in cDC1s 225.

Comparison of Zbtb46-DTR and Itgax-DTR mice reveals a partial redundancy between cDCs and other CD11c+ cells in some aspects of the antitumor response. When immunized with tumor antigens and challenged with melanoma, DT treatment in either strain led to a failure to reject tumors 48. But mice depleted of CD11c+ cells succumbed to tumors faster than mice depleted of cDCs, suggesting that CD11c+ non-cDCs can compensate in driving antitumor responses.

Others studies have distinguished various macrophage and DC populations present within the tumor microenvironment 226,227, and future studies may provide a better understanding of the roles of each in tumor immunity. Figure 1.1 summarizes the many known roles of cDCs in immune responses.

1.5 Conclusions.

The increasing precision by which cDC subsets can be ablated in mouse models has greatly expanded our understanding of their functions. It is now clear that cDCs are a unique lineage comprised of distinct functional subsets critical for both innate and adaptive immunity. Further work is needed to resolve the apparent heterogeneity in cDC2s 228, and to test for their potential activities. In particular, how cDC2s control type 2 immune responses is unclear, including whether they provide instructive signals themselves, whether they are the primary APCs, and whether they, or other cells such as epithelial tuft cells, directly sense the pathogen. While these and other questions remain, it is now clear that DCs are the critical regulators of the entire immune response and that to understand the immune system we must fully analyze these remarkable cells.

In this dissertation, we address key questions regarding dendritic cell development and functions. First, in Chapter 2, we resolved a long-standing paradox regarding the tyrosine kinase receptor Flt3 and its ligand Flt3L. We determined that the less severe DC deficiency in Flt3–/– mice as compared to Flt3l–/– mice was due to the enhanced responsiveness of Flt3–/– progenitors to compensatory cytokines that could also support DC development. This established that Flt3L does not have instructive roles in DC development, and instead that committed DC precursors can mature in response to multiple cytokines. In Chapter 3, we analyzed enhancer usage within the

Irf8 locus to determine how this critical transcription factor was induced in order to specify cDC1 fate. Irf8 is the lineage-determining transcription factor for cDC1s, but it remains unclear how this factor is induced in the multipotent CDP in order to specify cDC1 fate. We therefore identified active enhancers in this locus by chromatin profiling of DCs and used CRISPR/Cas9 to delete

31 these enhancers and identify the effects in vivo. We found that a +32 kb Irf8 enhancer is responsible for the maturation of cDC1s, but not for their specification. Deletion of a previously identified -50 kb Irf8 enhancer thought to be active in early DC precursors also did not affect DC development, but did reduce Irf8 expression in monocytes and macrophages and lead to increased susceptibility to Salmonella typhimurium infection. To identify enhancers involved in specification, we performed ATAC-seq on all DC precursors. This surprisingly revealed that a +41 kb Irf8 enhancer, which we had previously found to be active in mature pDCs, was transiently active in cDC1 progenitors but not in mature cDC1s. Deletion of this +41 kb Irf8 enhancer eliminated Irf8 induction in the CDP and subsequent cDC1 specification. This enhancer was bound by E proteins, and consistently E2A-deficient DC progenitors failed to generate cDC1s. We therefore conclude that transient activation of the +41 kb Irf8 enhancer by E proteins is responsible for cDC1 fate specification.

1.6 Author Contributions.

V.D. and K.M.M. wrote the review article (Immunity).

1.7 References.

1. Steinman,R.M. & Cohn,Z.A. Identification of a novel cell type in peripheral lymphoid organs of mice. I. Morphology, quantitation, tissue distribution. J Exp. Med. 137, 1142- 1162 (1973).

2. Steinman,R.M. & Witmer,M.D. Lymphoid dendritic cells are potent stimulators of the primary mixed leukocyte reaction in mice. Proc. Natl. Acad. Sci U. S A 75, 5132-5136 (1978).

3. Nussenzweig,M.C., Steinman,R.M., Gutchinov,B. & Cohn,Z.A. Dendritic cells are accessory cells for the development of anti-trinitrophenyl cytotoxic T lymphocytes. J. Exp. Med. 152, 1070-1084 (1980).

4. Perussia,B., Fanning,V. & Trinchieri,G. A leukocyte subset bearing HLA-DR antigens is responsible for in vitro alpha interferon production in response to viruses. Nat. Immun. Cell Growth Regul. 4, 120-137 (1985).

5. Heath,W.R. & Carbone,F.R. Dendritic cell subsets in primary and secondary T cell responses at body surfaces. Nat. Immunol. 10, 1237-1244 (2009).

6. Cella,M. et al. Plasmacytoid monocytes migrate to inflamed lymph nodes and produce large amounts of type I interferon [see comments]. Nature Medicine 5, 919-923 (1999).

7. Siegal,F.P. et al. The nature of the principal type 1 interferon-producing cells in human blood. Science 284, 1835-1837 (1999).

8. Guilliams,M. et al. Dendritic cells, monocytes and macrophages: a unified nomenclature based on ontogeny. Nat. Rev. Immunol. 14, 571-578 (2014).

9. Fogg,D.K. et al. A clonogenic bone marrow progenitor specific for macrophages and dendritic cells. Science 311, 83-87 (2006).

10. Auffray,C. et al. CX3CR1+ CD115+ CD135+ common macrophage/DC precursors and the role of CX3CR1 in their response to inflammation. J Exp. Med. 206, 595-606 (2009).

11. Naik,S.H. et al. Development of plasmacytoid and conventional dendritic cell subtypes from single precursor cells derived in vitro and in vivo. Nat Immunol 8, 1217-1226 (2007).

12. Onai,N. et al. Identification of clonogenic common Flt3(+) M-CSFR+ plasmacytoid and conventional dendritic cell progenitors in mouse bone marrow. Nat Immunol 8, 1207- 1216 (2007).

13. Grajales-Reyes,G.E. et al. Batf3 maintains autoactivation of Irf8 for commitment of a CD8alpha(+) conventional DC clonogenic progenitor. Nat Immunol 16, 708-717 (2015).

14. Schlitzer,A. et al. Identification of cDC1- and cDC2-committed DC progenitors reveals early lineage priming at the common DC progenitor stage in the bone marrow. Nat Immunol (2015).

15. Steinman,R.M., Kaplan,G., Witmer,M.D. & Cohn,Z.A. Identification of a novel cell type in peripheral lymphoid organs of mice. V. Purification of spleen dendritic cells, new surface markers, and maintenance in vitro. J Exp. Med 149, 1-16 (1979).

16. Metlay,J.P. et al. The distinct leukocyte integrins of mouse spleen dendritic cells as identified with new hamster monoclonal antibodies. J Exp. Med 171, 1753-1771 (1990).

17. Mildner,A. & Jung,S. Development and function of dendritic cell subsets. Immunity 40, 642-656 (2014).

18. Murphy,T.L. et al. Transcriptional Control of Dendritic Cell Development. Annu. Rev Immunol 34, 93-119 (2016).

19. Schlitzer,A. et al. IRF4 Transcription Factor-Dependent CD11b(+) Dendritic Cells in Human and Mouse Control Mucosal IL-17 Cytokine Responses. Immunity 38, 970-983 (2013).

20. Plantinga,M. et al. Conventional and Monocyte-Derived CD11b(+) Dendritic Cells Initiate and Maintain T Helper 2 Cell-Mediated Immunity to House Dust Mite Allergen. Immunity (2013).

21. Langlet,C. et al. CD64 expression distinguishes monocyte-derived and conventional dendritic cells and reveals their distinct role during intramuscular immunization. J Immunol 188, 1751-1760 (2012).

22. Bogunovic,M. et al. Origin of the lamina propria dendritic cell network. Immunity 31, 513-525 (2009).

23. Satpathy,A.T. et al. Notch2-dependent classical dendritic cells orchestrate intestinal immunity to attaching-and-effacing bacterial pathogens. Nat. Immunol. 14, 937-948 (2013).

24. Guilliams,M. et al. Unsupervised High-Dimensional Analysis Aligns Dendritic Cells across Tissues and Species. Immunity 45, 669-684 (2016).

25. Blasius,A.L. et al. Bone marrow stromal cell antigen 2 is a specific marker of type I IFN- producing cells in the naive mouse, but a promiscuous cell surface antigen following IFN stimulation. J Immunol 177, 3260-3265 (2006).

26. Zhang,J. et al. Characterization of Siglec-H as a novel endocytic receptor expressed on murine plasmacytoid dendritic cell precursors. Blood 107, 3600-3608 (2006).

27. Mashayekhi,M. et al. CD8a+ Dendritic Cells Are the Critical Source of Interleukin-12 that Controls Acute Infection by Toxoplasma gondii Tachyzoites. Immunity 35, 249-259 (2011).

28. Briseno,C.G., Murphy,T.L. & Murphy,K.M. Complementary diversification of dendritic cells and innate lymphoid cells. Curr. Opin. Immunol. 29C, 69-78 (2014).

29. Tussiwand,R. et al. Klf4 expression in conventional dendritic cells is required for T helper 2 cell responses. Immunity 42, 916-928 (2015).

30. Lewis,K.L. et al. Notch2 receptor signaling controls functional differentiation of dendritic cells in the spleen and intestine. Immunity 35, 780-791 (2011).

31. Hume,D.A. Macrophages as APC and the dendritic cell myth. J Immunol 181, 5829-5835 (2008).

32. Palmiter,R.D. et al. Cell lineage ablation in transgenic mice by cell-specific expression of a toxin gene. Cell 50, 435-443 (1987).

33. Breitman,M.L. et al. Genetic ablation: targeted expression of a toxin gene causes microphthalmia in transgenic mice. Science 238, 1563-1565 (1987).

34. Honjo,T., Nishizuka,Y. & Hayaishi,O. Diphtheria toxin-dependent adenosine diphosphate ribosylation of aminoacyl transferase II and inhibition of protein synthesis. J Biol Chem. 243, 3553-3555 (1968).

35. Robinson,E.A., Henriksen,O. & Maxwell,E.S. Elongation factor 2. Amino acid sequence at the site of adenosine diphosphate ribosylation. J Biol Chem. 249, 5088-5093 (1974).

36. Van Ness,B.G., Howard,J.B. & Bodley,J.W. ADP-ribosylation of elongation factor 2 by diphtheria toxin. Isolation and properties of the novel ribosyl-amino acid and its hydrolysis products. J Biol Chem. 255, 10717-10720 (1980).

37. Breitman,M.L., Rombola,H., Maxwell,I.H., Klintworth,G.K. & Bernstein,A. Genetic ablation in transgenic mice with an attenuated diphtheria toxin A gene. Mol Cell Biol 10, 474-479 (1990).

38. Saito,M. et al. Diphtheria toxin receptor-mediated conditional and targeted cell ablation in transgenic mice. Nat Biotechnol. 19, 746-750 (2001).

39. Mitamura,T., Higashiyama,S., Taniguchi,N., Klagsbrun,M. & Mekada,E. Diphtheria toxin binds to the epidermal growth factor (EGF)-like domain of human heparin-binding EGF-like growth factor/diphtheria toxin receptor and inhibits specifically its mitogenic activity. J Biol Chem. 270, 1015-1019 (1995).

40. Pappenheimer,A.M., Jr., Harper,A.A., Moynihan,M. & Brockes,J.P. Diphtheria toxin and related proteins: effect of route of injection on toxicity and the determination of cytotoxicity for various cultured cells. J Infect. Dis 145, 94-102 (1982).

41. Jung,S. et al. In vivo depletion of CD11c(+) dendritic cells abrogates priming of CD8(+) T cells by exogenous cell-associated antigens. Immunity 17, 211-220 (2002).

42. Zaft,T., Sapoznikov,A., Krauthgamer,R., Littman,D.R. & Jung,S. CD11chigh dendritic cell ablation impairs lymphopenia-driven proliferation of naive and memory CD8+ T cells. J Immunol 175, 6428-6435 (2005).

43. Probst,H.C. et al. Histological analysis of CD11c-DTR/GFP mice after in vivo depletion of dendritic cells. Clin. Exp. Immunol 141, 398-404 (2005).

44. van Rijt,L.S. et al. In vivo depletion of lung CD11c+ dendritic cells during allergen challenge abrogates the characteristic features of asthma. J. Exp. Med. 201, 981-991 (2005).

45. Hebel,K. et al. Plasma cell differentiation in T-independent type 2 immune responses is independent of CD11c(high) dendritic cells. Eur. J Immunol 36, 2912-2919 (2006).

46. Sapoznikov,A. et al. Organ-dependent in vivo priming of naive CD4+, but not CD8+, T cells by plasmacytoid dendritic cells. J Exp. Med. 204, 1923-1933 (2007).

47. Hochweller,K., Striegler,J., Hammerling,G.J. & Garbi,N. A novel CD11c.DTR transgenic mouse for depletion of dendritic cells reveals their requirement for homeostatic proliferation of natural killer cells. Eur. J Immunol 38, 2776-2783 (2008).

48. Meredith,M.M. et al. Expression of the zinc finger transcription factor zDC (Zbtb46, Btbd4) defines the classical dendritic cell lineage. J. Exp. Med. 209, 1153-1165 (2012).

49. Satpathy,A.T. et al. Zbtb46 expression distinguishes classical dendritic cells and their committed progenitors from other immune lineages. J. Exp. Med. 209, 1135-1152 (2012).

50. Loschko,J. et al. Inducible targeting of cDCs and their subsets in vivo. J Immunol Methods (2016).

51. Kissenpfennig,A. et al. Dynamics and function of Langerhans cells in vivo: dermal dendritic cells colonize lymph node areas distinct from slower migrating Langerhans cells. Immunity 22, 643-654 (2005).

52. Bennett,C.L. et al. Inducible ablation of mouse Langerhans cells diminishes but fails to abrogate contact hypersensitivity. J Cell Biol 169, 569-576 (2005).

53. Kaplan,D.H., Jenison,M.C., Saeland,S., Shlomchik,W.D. & Shlomchik,M.J. Epidermal langerhans cell-deficient mice develop enhanced contact hypersensitivity. Immunity 23, 611-620 (2005).

54. Welty,N.E. et al. Intestinal lamina propria dendritic cells maintain T cell homeostasis but do not affect commensalism. J. Exp. Med. (2013).

55. Fukaya,T. et al. Conditional ablation of CD205+ conventional dendritic cells impacts the regulation of T-cell immunity and homeostasis in vivo. Proc. Natl. Acad. Sci. U S A 109, 11288-11293 (2012).

56. Inaba,K. et al. Tissue distribution of the DEC-205 protein that is detected by the monoclonal antibody NLDC-145. I. Expression on dendritic cells and other subsets of mouse leukocytes. Cell Immunol 163, 148-156 (1995).

57. Victora,G.D. et al. Germinal center dynamics revealed by multiphoton microscopy with a photoactivatable fluorescent reporter. Cell 143, 592-605 (2010).

58. Idoyaga,J. et al. Specialized role of migratory dendritic cells in peripheral tolerance induction. J Clin. Invest 123, 844-854 (2013).

59. Piva,L. et al. Cutting edge: Clec9A+ dendritic cells mediate the development of experimental cerebral malaria. J Immunol 189, 1128-1132 (2012).

60. Sancho,D. et al. Tumor therapy in mice via antigen targeting to a novel, DC-restricted C- type lectin. J. Clin. Invest 118, 2098-2110 (2008).

61. Caminschi,I. et al. The dendritic cell subtype-restricted C-type lectin Clec9A is a target for vaccine enhancement. Blood 112, 3264-3273 (2008).

62. Schraml,B.U. et al. Genetic tracing via DNGR-1 expression history defines dendritic cells as a hematopoietic lineage. Cell 154, 843-858 (2013).

63. Muzaki,A.R. et al. Intestinal CD103(+)CD11b(-) dendritic cells restrain colitis via IFN- gamma-induced anti-inflammatory response in epithelial cells. Mucosal Immunol 9, 336- 351 (2016).

64. Yamazaki,C. et al. Critical roles of a dendritic cell subset expressing a chemokine receptor, XCR1. J Immunol 190, 6071-6082 (2013).

65. Crozat,K. et al. The XC chemokine receptor 1 is a conserved selective marker of mammalian cells homologous to mouse CD8alpha+ dendritic cells. J Exp. Med. 207, 1283-1292 (2010).

66. Crozat,K. et al. Cutting edge: expression of XCR1 defines mouse lymphoid-tissue resident and migratory dendritic cells of the CD8alpha+ type. J. Immunol. 187, 4411- 4415 (2011).

67. Alexandre,Y.O. et al. XCR1+ dendritic cells promote memory CD8+ T cell recall upon secondary infections with Listeria monocytogenes or certain viruses. J Exp. Med 213, 75- 92 (2016).

68. Nussenzweig,M.C., Steinman,R.M., Witmer,M.D. & Gutchinov,B. A monoclonal antibody specific for mouse dendritic cells. Proc. Natl. Acad. Sci. U S A 79, 161-165 (1982).

69. Dudziak,D. et al. Differential antigen processing by dendritic cell subsets in vivo. Science 315, 107-111 (2007).

70. Kumamoto,Y. et al. CD301b(+) dermal dendritic cells drive T helper 2 cell-mediated immunity. Immunity 39, 733-743 (2013).

71. Swiecki,M., Gilfillan,S., Vermi,W., Wang,Y. & Colonna,M. Plasmacytoid dendritic cell ablation impacts early interferon responses and antiviral NK and CD8(+) T cell accrual. Immunity 33, 955-966 (2010).

72. Takagi,H. et al. Plasmacytoid dendritic cells are crucial for the initiation of inflammation and T cell immunity in vivo. Immunity. 35, 958-971 (2011).

73. Swiecki,M. et al. Cell depletion in mice that express diphtheria toxin receptor under the control of SiglecH encompasses more than plasmacytoid dendritic cells. J Immunol 192, 4409-4416 (2014).

74. Orr,S.L. et al. A phenotype survey of 36 mutant mouse strains with gene-targeted defects in glycosyltransferases or glycan-binding proteins. Glycobiology 23, 363-380 (2013).

75. Lauvau,G., Loke,P. & Hohl,T.M. Monocyte-mediated defense against bacteria, fungi, and parasites. Semin Immunol 27, 397-409 (2015).

76. Diehl,G.E. et al. Microbiota restricts trafficking of bacteria to mesenteric lymph nodes by CX(3)CR1(hi) cells. Nature 494, 116-120 (2013).

77. Schreiber,H.A. et al. Intestinal monocytes and macrophages are required for T cell polarization in response to Citrobacter rodentium. J. Exp. Med. 210, 2025-2039 (2013).

78. van Blijswijk,J. et al. Altered lymph node composition in diphtheria toxin receptor-based mouse models to ablate dendritic cells. J Immunol 194, 307-315 (2015).

79. Buch,T. et al. A Cre-inducible diphtheria toxin receptor mediates cell lineage ablation after toxin administration. Nat Methods 2, 419-426 (2005).

80. Sharma,S. & Zhu,J. Immunologic applications of conditional gene modification technology in the mouse. Curr. Protoc. Immunol 105, 10-13 (2014).

81. Caton,M.L., Smith-Raska,M.R. & Reizis,B. Notch-RBP-J signaling controls the homeostasis of CD8- dendritic cells in the spleen. J Exp. Med. 204, 1653-1664 (2007).

82. Srinivas,S. et al. Cre reporter strains produced by targeted insertion of EYFP and ECFP into the ROSA26 locus. BMC Dev. Biol 1, 4 (2001).

83. Abram,C.L., Roberge,G.L., Hu,Y. & Lowell,C.A. Comparative analysis of the efficiency and specificity of myeloid-Cre deleting strains using ROSA-EYFP reporter mice. J. Immunol. Methods 408, 89-100 (2014).

84. Stranges,P.B. et al. Elimination of antigen-presenting cells and autoreactive T cells by Fas contributes to prevention of autoimmunity. Immunity 26, 629-641 (2007).

85. Loschko,J. et al. Absence of MHC class II on cDCs results in microbial-dependent intestinal inflammation. J Exp. Med 213, 517-534 (2016).

86. Zahner,S.P. et al. Conditional deletion of TGF-betaR1 using Langerin-Cre mice results in Langerhans cell deficiency and reduced contact hypersensitivity. J Immunol 187, 5069- 5076 (2011).

87. Ohta,T. et al. Crucial roles of XCR1-expressing dendritic cells and the XCR1-XCL1 chemokine axis in intestinal immune homeostasis. Sci. Rep. 6, 23505 (2016).

88. Brockschnieder,D., Pechmann,Y., Sonnenberg-Riethmacher,E. & Riethmacher,D. An improved mouse line for Cre-induced cell ablation due to diphtheria toxin A, expressed from the Rosa26 locus. Genesis. 44, 322-327 (2006).

89. Puttur,F. et al. Absence of Siglec-H in MCMV infection elevates interferon alpha production but does not enhance viral clearance. PLoS Pathog 9, e1003648 (2013). 39

90. Schiavoni,G. et al. ICSBP is essential for the development of mouse type I interferon- producing cells and for the generation and activation of CD8alpha(+) dendritic cells. J Exp. Med. 196, 1415-1425 (2002).

91. Aliberti,J. et al. Essential role for ICSBP in the in vivo development of murine CD8alpha + dendritic cells. Blood 101, 305-310 (2003).

92. Wang,H. et al. IRF8 regulates B-cell lineage specification, commitment, and differentiation. Blood 112, 4028-4038 (2008).

93. Kurotaki,D. et al. Essential role of the IRF8-KLF4 transcription factor cascade in murine monocyte differentiation. Blood 121, 1839-1849 (2013).

94. Milanovic,M., Terszowski,G., Struck,D., Liesenfeld,O. & Carstanjen,D. IFN consensus sequence binding protein (Icsbp) is critical for eosinophil development. J Immunol 181, 5045-5053 (2008).

95. Sasaki,H. et al. Transcription factor IRF8 plays a critical role in the development of murine basophils and mast cells. Blood 125, 358-369 (2015).

96. Sichien,D. et al. IRF8 Transcription Factor Controls Survival and Function of Terminally Differentiated Conventional and Plasmacytoid Dendritic Cells, Respectively. Immunity 45, 626-640 (2016).

97. Luda,K.M. et al. IRF8 Transcription-Factor-Dependent Classical Dendritic Cells Are Essential for Intestinal T Cell Homeostasis. Immunity 44, 860-874 (2016).

98. Esterhazy,D. et al. Classical dendritic cells are required for dietary antigen-mediated induction of peripheral T(reg) cells and tolerance. Nat Immunol 17, 545-555 (2016).

99. Hacker,C. et al. Transcriptional profiling identifies Id2 function in dendritic cell development. Nat Immunol 4, 380-386 (2003).

100. Yokota,Y. et al. Development of peripheral lymphoid organs and natural killer cells depends on the helix-loop-helix inhibitor Id2. Nature 397, 702-706 (1999).

101. Serafini,N., Vosshenrich,C.A. & Di Santo,J.P. Transcriptional regulation of innate lymphoid cell fate. Nat Rev Immunol 15, 415-428 (2015).

102. Kashiwada,M., Pham,N.L., Pewe,L.L., Harty,J.T. & Rothman,P.B. NFIL3/E4BP4 is a key transcription factor for CD8{alpha}+ dendritic cell development. Blood 117, 6193- 6197 (2011).

103. Gascoyne,D.M. et al. The basic leucine zipper transcription factor E4BP4 is essential for natural killer cell development. Nat. Immunol. 10, 1118-1124 (2009).

104. Geiger,T.L. et al. Nfil3 is crucial for development of innate lymphoid cells and host protection against intestinal pathogens. J Exp. Med 211, 1723-1731 (2014).

105. Seillet,C. et al. Nfil3 is required for the development of all innate lymphoid cell subsets. J Exp. Med 211, 1733-1740 (2014).

106. Male,V. et al. The transcription factor E4bp4/Nfil3 controls commitment to the NK lineage and directly regulates Eomes and Id2 expression. J Exp. Med 211, 635-642 (2014).

107. Hildner,K. et al. Batf3 deficiency reveals a critical role for CD8alpha+ dendritic cells in cytotoxic T cell immunity. Science 322, 1097-1100 (2008).

108. Tussiwand,R. et al. Compensatory dendritic cell development mediated by BATF-IRF interactions. Nature 490, 502-507 (2012).

109. Glasmacher,E. et al. A Genomic Regulatory Element That Directs Assembly and Function of Immune-Specific AP-1-IRF Complexes. Science 338, 975-980 (2012).

110. Edelson,B.T. et al. Peripheral CD103+ dendritic cells form a unified subset developmentally related to CD8alpha+ conventional dendritic cells. J. Exp. Med. 207, 823-836 (2010).

111. Burkly,L. et al. Expression of relB is required for the development of thymic medulla and dendritic cells. Nature 373, 531-536 (1995).

112. Wu,L. et al. RelB is essential for the development of myeloid-related CD8alpha- dendritic cells but not of lymphoid-related CD8alpha+ dendritic cells. Immunity. 9, 839- 847 (1998).

113. Kabashima,K. et al. Intrinsic lymphotoxin-beta receptor requirement for homeostasis of lymphoid tissue dendritic cells. Immunity 22, 439-450 (2005).

114. Weih,F. et al. Multiorgan inflammation and hematopoietic abnormalities in mice with a targeted disruption of RelB, a member of the NF-kappa B/Rel family. Cell 80, 331-340 (1995).

115. Futterer,A., Mink,K., Luz,A., Kosco-Vilbois,M.H. & Pfeffer,K. The lymphotoxin beta receptor controls organogenesis and affinity maturation in peripheral lymphoid tissues. Immunity 9, 59-70 (1998).

116. Tumanov,A.V. et al. Lymphotoxin controls the IL-22 protection pathway in gut innate lymphoid cells during mucosal pathogen challenge. Cell Host Microbe 10, 44-53 (2011).

117. Ochiai,S. et al. CD326loCD103loCD11blo Dermal Dendritic Cells Are Activated by Thymic Stromal Lymphopoietin during Contact Sensitization in Mice. J. Immunol. 193, 2504-2511 (2014).

118. Bajana,S., Roach,K., Turner,S., Paul,J. & Kovats,S. IRF4 promotes cutaneous dendritic cell migration to lymph nodes during homeostasis and inflammation. J. Immunol. 189, 3368-3377 (2012).

119. Bajana,S., Turner,S., Paul,J., Ainsua-Enrich,E. & Kovats,S. IRF4 and IRF8 Act in CD11c+ Cells To Regulate Terminal Differentiation of Lung Tissue Dendritic Cells. J Immunol 196, 1666-1677 (2016).

120. Persson,E.K. et al. IRF4 Transcription-Factor-Dependent CD103(+)CD11b(+) Dendritic Cells Drive Mucosal T Helper 17 Cell Differentiation. Immunity 38, 958-969 (2013).

121. Gao,Y. et al. Control of T helper 2 responses by transcription factor IRF4-dependent dendritic cells. Immunity 39, 722-732 (2013).

122. Satoh,T. et al. The Jmjd3-Irf4 axis regulates M2 macrophage polarization and host responses against helminth infection. Nat. Immunol. 11, 936-944 (2010).

123. Briseno,C.G. et al. Distinct Transcriptional Programs Control Cross-Priming in Classical and Monocyte-Derived Dendritic Cells. Cell Rep. 15, 2462-2474 (2016).

124. Cisse,B. et al. Transcription factor E2-2 is an essential and specific regulator of plasmacytoid dendritic cell development. Cell 135, 37-48 (2008).

125. Scott,C.L. et al. The transcription factor Zeb2 regulates development of conventional and plasmacytoid DCs by repressing Id2. J Exp. Med 213, 897-911 (2016).

126. Probst,H.C. & Van Den,B.M. Priming of CTLs by lymphocytic choriomeningitis virus depends on dendritic cells. J Immunol 174, 3920-3924 (2005).

127. Kassim,S.H., Rajasagi,N.K., Zhao,X., Chervenak,R. & Jennings,S.R. In vivo ablation of CD11c-positive dendritic cells increases susceptibility to herpes simplex virus type 1 infection and diminishes NK and T-cell responses. J Virol. 80, 3985-3993 (2006).

128. Ciavarra,R.P., Stephens,A., Nagy,S., Sekellick,M. & Steel,C. Evaluation of immunological paradigms in a virus model: are dendritic cells critical for antiviral immunity and viral clearance? J Immunol 177, 492-500 (2006).

129. GeurtsvanKessel,C.H. et al. Clearance of influenza virus from the lung depends on migratory langerin(+)CD11b(-) but not plasmacytoid dendritic cells. J. Exp. Med. 205, 1621-1634 (2008).

130. Kim,T.S., Hufford,M.M., Sun,J., Fu,Y.X. & Braciale,T.J. Antigen persistence and the control of local T cell memory by migrant respiratory dendritic cells after acute virus infection. J Exp. Med 207, 1161-1172 (2010).

131. Tian,T., Woodworth,J., Skold,M. & Behar,S.M. In vivo depletion of CD11c+ cells delays the CD4+ T cell response to Mycobacterium tuberculosis and exacerbates the outcome of infection. J Immunol 175, 3268-3272 (2005).

132. de Jong,J.M. et al. Dendritic cells, but not macrophages or B cells, activate major histocompatibility complex class II-restricted CD4+ T cells upon immune-complex uptake in vivo. Immunol. 119, 499-506 (2006).

133. Zammit,D.J., Cauley,L.S., Pham,Q.M. & Lefrancois,L. Dendritic cells maximize the memory CD8 T cell response to infection. Immunity 22, 561-570 (2005).

134. Lauterbach,H., Zuniga,E.I., Truong,P., Oldstone,M.B. & McGavern,D.B. Adoptive immunotherapy induces CNS dendritic cell recruitment and antigen presentation during clearance of a persistent viral infection. J Exp. Med 203, 1963-1975 (2006).

135. Samstein,M. et al. Essential yet limited role for CCR2(+) inflammatory monocytes during Mycobacterium tuberculosis-specific T cell priming. Elife. 2, e01086 (2013).

136. den Haan,J.M., Lehar,S.M. & Bevan,M.J. CD8(+) but not CD8(-) dendritic cells cross- prime cytotoxic T cells in vivo. J Exp. Med. 192, 1685-1696 (2000).

137. Torti,N., Walton,S.M., Murphy,K.M. & Oxenius,A. Batf3 transcription factor-dependent DC subsets in murine CMV infection: differential impact on T-cell priming and memory inflation. Eur. J. Immunol. 41, 2612-2618 (2011).

138. Helft,J. et al. Cross-presenting CD103+ dendritic cells are protected from influenza virus infection. J. Clin. Invest 122, 4037-4047 (2012).

139. Waithman,J. et al. Resident CD8(+) and migratory CD103(+) dendritic cells control CD8 T cell immunity during acute influenza infection. PLoS One 8, e66136 (2013).

140. Gainey,M.D., Rivenbark,J.G., Cho,H., Yang,L. & Yokoyama,W.M. Viral MHC class I inhibition evades CD8+ T-cell effector responses in vivo but not CD8+ T-cell priming. Proc. Natl. Acad. Sci. U. S A (2012).

141. Nopora,K. et al. MHC class I cross-presentation by dendritic cells counteracts viral immune evasion. Front Immunol. 3, 348 (2012).

142. Zelenay,S. et al. The dendritic cell receptor DNGR-1 controls endocytic handling of necrotic cell antigens to favor cross-priming of CTLs in virus-infected mice. J. Clin. Invest 122, 1615-1627 (2012).

143. Lau,L.S. et al. CD8+ T cells from a novel T cell receptor transgenic mouse induce liver- stage immunity that can be boosted by blood-stage infection in rodent malaria. PLoS Pathog. 10, e1004135 (2014).

144. Hor,J.L. et al. Spatiotemporally Distinct Interactions with Dendritic Cell Subsets Facilitates CD4+ and CD8+ T Cell Activation to Localized Viral Infection. Immunity 43, 554-565 (2015).

145. Eickhoff,S. et al. Robust Anti-viral Immunity Requires Multiple Distinct T Cell- Dendritic Cell Interactions. Cell 162, 1322-1337 (2015).

146. Goenka,R. et al. Cutting edge: dendritic cell-restricted antigen presentation initiates the follicular helper T cell program but cannot complete ultimate effector differentiation. J Immunol 187, 1091-1095 (2011).

147. Calabro,S. et al. Bridging channel dendritic cells induce immunity to transfused red blood cells. J Exp. Med 213, 887-896 (2016).

148. Li,J., Lu,E., Yi,T. & Cyster,J.G. EBI2 augments Tfh cell fate by promoting interaction with IL-2-quenching dendritic cells. Nature 533, 110-114 (2016).

149. Reboldi,A. et al. IgA production requires B cell interaction with subepithelial dendritic cells in Peyer's patches. Science 352, aaf4822 (2016).

150. Sonnenberg,G.F. & Artis,D. Innate lymphoid cells in the initiation, regulation and resolution of inflammation. Nat Med 21, 698-708 (2015).

151. Hsieh,C.S. et al. Development of TH1 CD4+ T cells through IL-12 produced by Listeria- induced macrophages. Science 260, 547-549 (1993).

152. Scharton-Kersten,T., Contursi,C., Masumi,A., Sher,A. & Ozato,K. Interferon consensus sequence binding protein-deficient mice display impaired resistance to intracellular infection due to a primary defect in interleukin 12 p40 induction. J Exp. Med. 186, 1523- 1534 (1997).

153. Liu,C.H. et al. Cutting edge: dendritic cells are essential for in vivo IL-12 production and development of resistance against Toxoplasma gondii infection in mice. J Immunol 177, 31-35 (2006).

154. Askenase,M.H. et al. Bone-Marrow-Resident NK Cells Prime Monocytes for Regulatory Function during Infection. Immunity 42, 1130-1142 (2015).

155. Neuenhahn,M. et al. CD8alpha+ dendritic cells are required for efficient entry of Listeria monocytogenes into the spleen. Immunity 25, 619-630 (2006).

156. Edelson,B.T. et al. CD8a+ Dendritic Cells Are an Obligate Cellular Entry Point for Productive Infection by Listeria monocytogenes. Immunity 35, 236-248 (2011).

157. Voehringer,D., Liang,H.E. & Locksley,R.M. Homeostasis and effector function of lymphopenia-induced "memory-like" T cells in constitutively T cell-depleted mice. J Immunol 180, 4742-4753 (2008).

158. Kang,S.J., Liang,H.E., Reizis,B. & Locksley,R.M. Regulation of Hierarchical Clustering and Activation of Innate Immune Cells by Dendritic Cells. Immunity 29, 819-833 (2008).

159. Ashok,D. et al. Cross-presenting dendritic cells are required for control of Leishmania major infection. Eur. J. Immunol. 44, 1422-1432 (2014).

160. Martinez-Lopez,M., Iborra,S., Conde-Garrosa,R. & Sancho,D. Batf3-dependent CD103+ dendritic cells are major producers of IL-12 that drive local Th1 immunity against Leishmania major infection in mice. Eur. J Immunol 45, 119-129 (2015).

161. Igyarto,B.Z. et al. Skin-resident murine dendritic cell subsets promote distinct and opposing antigen-specific T helper cell responses. Immunity 35, 260-272 (2011).

162. Guimond,M. et al. In vivo role of Flt3 ligand and dendritic cells in NK cell homeostasis. J Immunol 184, 2769-2775 (2010).

163. Lucas,M., Schachterle,W., Oberle,K., Aichele,P. & Diefenbach,A. Dendritic cells prime natural killer cells by trans-presenting interleukin 15. Immunity 26, 503-517 (2007).

164. Schleicher,U. et al. NK cell activation in visceral leishmaniasis requires TLR9, myeloid DCs, and IL-12, but is independent of plasmacytoid DCs. J Exp. Med 204, 893-906 (2007).

165. Burkett,P.R. et al. Coordinate expression and trans presentation of interleukin (IL)- 15Ralpha and IL-15 supports natural killer cell and memory CD8+ T cell homeostasis. J Exp. Med 200, 825-834 (2004).

166. Mortier,E., Woo,T., Advincula,R., Gozalo,S. & Ma,A. IL-15Ralpha chaperones IL-15 to stable dendritic cell membrane complexes that activate NK cells via trans presentation. J Exp. Med 205, 1213-1225 (2008).

167. Le Gros,G., Ben-Sasson,S.Z., Seder,R., Finkelman,F.D. & Paul,W.E. Generation of interleukin 4 (IL-4)-producing cells in vivo and in vitro: IL-2 and IL-4 are required for in vitro generation of IL-4-producing cells. J. Exp. Med. 172, 921-929 (1990).

168. Sokol,C.L. et al. Basophils function as antigen-presenting cells for an allergen-induced T helper type 2 response. Nat Immunol 10, 713-720 (2009).

169. Tang,H. et al. The T helper type 2 response to cysteine proteases requires dendritic cell- basophil cooperation via ROS-mediated signaling. Nat. Immunol. 11, 608-617 (2010).

170. Hammad,H. et al. Inflammatory dendritic cells--not basophils--are necessary and sufficient for induction of Th2 immunity to inhaled house dust mite allergen. J Exp. Med. 207, 2097-2111 (2010).

171. Phythian-Adams,A.T. et al. CD11c depletion severely disrupts Th2 induction and development in vivo. J. Exp. Med. 207, 2089-2096 (2010).

172. Smith,K.A. et al. Chronic helminth infection promotes immune regulation in vivo through dominance of CD11cloC. J Immunol 186, 7098-7109 (2011).

173. Smith,K.A. et al. Type 2 innate immunity in helminth infection is induced redundantly and acts autonomously following CD11c(+) cell depletion. Infect. Immun. 80, 3481-3489 (2012).

174. Williams,J.W. et al. Transcription factor IRF4 drives dendritic cells to promote Th2 differentiation. Nat. Commun. 4, 2990 (2013).

175. Leon,B. et al. Regulation of T(H)2 development by CXCR5+ dendritic cells and lymphotoxin-expressing B cells. Nat. Immunol. 13, 681-690 (2012).

176. Everts,B. et al. Migratory CD103+ dendritic cells suppress helminth-driven type 2 immunity through constitutive expression of IL-12. J Exp. Med 213, 35-51 (2016).

177. Halim,T.Y. et al. Group 2 innate lymphoid cells are critical for the initiation of adaptive T helper 2 cell-mediated allergic lung inflammation. Immunity 40, 425-435 (2014).

178. Halim,T.Y. et al. Group 2 innate lymphoid cells license dendritic cells to potentiate memory TH2 cell responses. Nat Immunol 17, 57-64 (2016).

179. Bettelli,E. et al. Reciprocal developmental pathways for the generation of pathogenic effector TH17 and regulatory T cells. Nature 441, 235-238 (2006).

180. Veldhoen,M., Hocking,R.J., Atkins,C.J., Locksley,R.M. & Stockinger,B. TGFbeta in the context of an inflammatory cytokine milieu supports de novo differentiation of IL-17- producing T cells. Immunity 24, 179-189 (2006).

181. Kinnebrew,M.A. et al. Interleukin 23 production by intestinal CD103(+)CD11b(+) dendritic cells in response to bacterial flagellin enhances mucosal innate immune defense. Immunity 36, 276-287 (2012).

182. Zheng,Y. et al. Interleukin-22 mediates early host defense against attaching and effacing bacterial pathogens. Nat. Med. 14, 282-289 (2008).

183. Basu,R. et al. Th22 cells are an important source of IL-22 for host protection against enteropathogenic bacteria. Immunity 37, 1061-1075 (2012).

184. Mundy,R., Macdonald,T.T., Dougan,G., Frankel,G. & Wiles,S. Citrobacter rodentium of mice and man. Cell Microbiol. 7, 1697-1706 (2005).

185. Longman,R.S. et al. CX(3)CR1(+) mononuclear phagocytes support colitis-associated innate lymphoid cell production of IL-22. J Exp. Med 211, 1571-1583 (2014).

186. Linehan,J.L. et al. Generation of Th17 cells in response to intranasal infection requires TGF-beta1 from dendritic cells and IL-6 from CD301b+ dendritic cells. Proc. Natl. Acad. Sci. U S A 112, 12782-12787 (2015).

187. Kashem,S.W. et al. Nociceptive Sensory Fibers Drive Interleukin-23 Production from CD301b+ Dermal Dendritic Cells and Drive Protective Cutaneous Immunity. Immunity 43, 515-526 (2015).

188. Arnold,I.C. et al. CD11c(+) monocyte/macrophages promote chronic Helicobacter hepaticus-induced intestinal inflammation through the production of IL-23. Mucosal Immunol 9, 352-363 (2016).

189. Trautwein-Weidner,K. et al. Antigen-Specific Th17 Cells Are Primed by Distinct and Complementary Dendritic Cell Subsets in Oropharyngeal Candidiasis. PLoS Pathog 11, e1005164 (2015).

190. Ivanov,I.I. et al. Induction of intestinal Th17 cells by segmented filamentous bacteria. Cell 139, 485-498 (2009).

191. McKenna,H.J. et al. Mice lacking flt3 ligand have deficient hematopoiesis affecting hematopoietic progenitor cells, dendritic cells, and natural killer cells. Blood 95, 3489- 3497 (2000).

192. Goto,Y. et al. Segmented filamentous bacteria antigens presented by intestinal dendritic cells drive mucosal Th17 cell differentiation. Immunity 40, 594-607 (2014).

193. Geem,D. et al. Specific microbiota-induced intestinal Th17 differentiation requires MHC class II but not GALT and mesenteric lymph nodes. J Immunol 193, 431-438 (2014).

194. Panea,C. et al. Intestinal Monocyte-Derived Macrophages Control Commensal-Specific Th17 Responses. Cell Rep. 12, 1314-1324 (2015).

195. Kim,J.M., Rasmussen,J.P. & Rudensky,A.Y. Regulatory T cells prevent catastrophic autoimmunity throughout the lifespan of mice. Nat Immunol 8, 191-197 (2007).

196. Liu,K. et al. In vivo analysis of dendritic cell development and homeostasis. Science 324, 392-397 (2009).

197. Darrasse-Jeze,G. et al. Feedback control of regulatory T cell homeostasis by dendritic cells in vivo. J Exp. Med 206, 1853-1862 (2009).

198. Leventhal,D.S. et al. Dendritic Cells Coordinate the Development and Homeostasis of Organ-Specific Regulatory T Cells. Immunity 44, 847-859 (2016).

199. Birnberg,T. et al. Lack of conventional dendritic cells is compatible with normal development and T cell homeostasis, but causes myeloid proliferative syndrome. Immunity 29, 986-997 (2008).

200. Liston,A. et al. Differentiation of regulatory Foxp3+ T cells in the thymic cortex. Proc. Natl. Acad. Sci. U S A 105, 11903-11908 (2008).

201. Perry,J.S. et al. Distinct contributions of Aire and antigen-presenting-cell subsets to the generation of self-tolerance in the thymus. Immunity 41, 414-426 (2014).

202. Chu,D.K. et al. Indigenous enteric eosinophils control DCs to initiate a primary Th2 immune response in vivo. J. Exp. Med. 211, 1657-1672 (2014).

203. Hoshi,N. et al. MyD88 signalling in colonic mononuclear phagocytes drives colitis in IL- 10-deficient mice. Nat Commun. 3, 1120 (2012).

204. Hua,Z. et al. Requirement for MyD88 signaling in B cells and dendritic cells for germinal center anti-nuclear antibody production in Lyn-deficient mice. J Immunol 192, 875-885 (2014).

205. Teichmann,L.L. et al. Dendritic cells in lupus are not required for activation of T and B cells but promote their expansion, resulting in tissue damage. Immunity 33, 967-978 (2010).

206. Teichmann,L.L., Schenten,D., Medzhitov,R., Kashgarian,M. & Shlomchik,M.J. Signals via the adaptor MyD88 in B cells and DCs make distinct and synergistic contributions to immune activation and tissue damage in lupus. Immunity 38, 528-540 (2013).

207. Ferris,S.T. et al. A minor subset of Batf3-dependent antigen-presenting cells in islets of Langerhans is essential for the development of autoimmune diabetes. Immunity 41, 657- 669 (2014).

208. Manicassamy,S. et al. Activation of beta-catenin in dendritic cells regulates immunity versus tolerance in the intestine. Science 329, 849-853 (2010).

209. Young,J.A., He,T.H., Reizis,B. & Winoto,A. Commensal microbiota are required for systemic inflammation triggered by necrotic dendritic cells. Cell Rep. 3, 1932-1944 (2013).

210. Hammer,G.E. et al. Expression of A20 by dendritic cells preserves immune homeostasis and prevents colitis and spondyloarthritis. Nat Immunol 12, 1184-1193 (2011).

211. Han,D. et al. Dendritic cell expression of the signaling molecule TRAF6 is critical for gut microbiota-dependent immune tolerance. Immunity 38, 1211-1222 (2013).

212. Lamagna,C., Scapini,P., van Ziffle,J.A., DeFranco,A.L. & Lowell,C.A. Hyperactivated MyD88 signaling in dendritic cells, through specific deletion of Lyn kinase, causes severe autoimmunity and inflammation. Proc. Natl. Acad. Sci. U S A 110, E3311-E3320 (2013).

213. Cuda,C.M. et al. Caspase-8 acts as a molecular rheostat to limit RIPK1- and MyD88- mediated dendritic cell activation. J Immunol 192, 5548-5560 (2014).

214. Mazzini,E., Massimiliano,L., Penna,G. & Rescigno,M. Oral tolerance can be established via gap junction transfer of fed antigens from CX3CR1(+) macrophages to CD103(+) dendritic cells. Immunity 40, 248-261 (2014).

215. Travis,M.A. et al. Loss of integrin alpha(v)beta8 on dendritic cells causes autoimmunity and colitis in mice. Nature 449, 361-365 (2007).

216. Casares,N. et al. Caspase-dependent immunogenicity of doxorubicin-induced tumor cell death. J Exp. Med 202, 1691-1701 (2005).

217. Shimizu,K., Kurosawa,Y., Taniguchi,M., Steinman,R.M. & Fujii,S. Cross-presentation of glycolipid from tumor cells loaded with alpha-galactosylceramide leads to potent and long-lived T cell mediated immunity via dendritic cells. J Exp. Med 204, 2641-2653 (2007).

218. Scarlett,U.K. et al. Ovarian cancer progression is controlled by phenotypic changes in dendritic cells. J Exp. Med 209, 495-506 (2012).

219. Spranger,S., Bao,R. & Gajewski,T.F. Melanoma-intrinsic beta-catenin signalling prevents anti-tumour immunity. Nature 523, 231-235 (2015).

220. Salmon,H. et al. Expansion and Activation of CD103(+) Dendritic Cell Progenitors at the Tumor Site Enhances Tumor Responses to Therapeutic PD-L1 and BRAF Inhibition. Immunity 44, 924-938 (2016).

221. Sanchez-Paulete,A.R. et al. Cancer Immunotherapy with Immunomodulatory Anti- CD137 and Anti-PD-1 Monoclonal Antibodies Requires BATF3-Dependent Dendritic Cells. Cancer Discov. 6, 71-79 (2016).

222. Roberts,E.W. et al. Critical Role for CD103+/CD141+ Dendritic Cells Bearing CCR7 for Tumor Antigen Trafficking and Priming of T Cell Immunity in Melanoma. Cancer Cell (2016).

223. Headley,M.B. et al. Visualization of immediate immune responses to pioneer metastatic cells in the lung. Nature 531, 513-517 (2016).

224. Diamond,M.S. et al. Type I interferon is selectively required by dendritic cells for immune rejection of tumors. J. Exp. Med. 208, 1989-2003 (2011).

225. Fuertes,M.B. et al. Host type I IFN signals are required for antitumor CD8+ T cell responses through CD8{alpha}+ dendritic cells. J. Exp. Med. 208, 2005-2016 (2011).

226. Franklin,R.A. et al. The cellular and molecular origin of tumor-associated macrophages. Science 344, 921-925 (2014).

227. Broz,M.L. et al. Dissecting the tumor myeloid compartment reveals rare activating antigen-presenting cells critical for T cell immunity. Cancer Cell 26, 638-652 (2014).

228. Jaitin,D.A. et al. Massively parallel single-cell RNA-seq for marker-free decomposition of tissues into cell types. Science 343, 776-779 (2014).

Table 1.1. Comparison of DTA/DTR based mouse models for dendritic cell depletion

Strain Cells Depleted Caveats Itgax-DTR cDCs, metallophilic, Repeated DT treatment marginal zone, sinusoidal, leads to death, requires and alveolar CD11c-DTR into WT BM macrophages, activated chimeras CD8 T cells, plasma cells

No depletion of pDCs

Itgax-DOG cDCs, splenic Depletion of other cell types macrophages not evaluated

Zbtb46-DTR cDCs DT treatment leads to death, requires zDC-DTR into WT BM chimeras

Zbtb46-LSL-DTR cDCs Requires crossing to Cre strain active in cDC lineage

Cd207-DTR (Knockin) Langerhans cells, cDC1s

Cd207-DTR (BAC) Langerhans cells, cDC1s

CD207-DTA Langerhans cells, CD103+CD11b+ cDC2s in small intestine

Ly75-DTR cDC1s, ~15% splenic B DT treatment leads to death, cells requires CD205-DTR into WT BM chimeras

Depletion of germinal center B cells, migratory cDCs, and Langerhans cells untested but possible

Clec9a-DTR cDC1s, ~50% pDCs

Xcr1-DTRvenus cDC1s

Karma (a530099j19rik-DTR) cDC1s

Clec4a4-DTR cDC2s and DC depletion in other + + CD64 CX3CR1 organs not analyzed macrophages in intestinal lamina propria and MLNs

Mgl2-DTR Mgl2+ dermal cDC2s

CLEC4C-DTR pDCs

Siglech-DTR (Knockin) pDCs Loss of SiglecH expression in homozygous knockin mice

Depletion of marginal zone macrophages untested but possible

Siglech-DTR (BAC) pDCs, marginal zone macrophages

Cx3cr1-LSL-DTR Itgax-cre CD103-CD11b+CX3CR1+ Requires crossing to Cre cDC2s, + + strain active in cDCs CD11b CX3CR1 macrophages

MM-DTR Monocytes and monocyte-derived macrophages

Table 1.2. Comparison of Cre strains for conditional deletion of genes in dendritic cells

Strain Cells Affected Caveats

Itgax-cre (BAC) ~100% of cDCs, pDCs, Germline deletion possible alveolar macrophages, ~70% red pulp macrophages, 20-40% monocytes, marginal zone macrophages, peritoneal macrophages, ~5-12% T, B, NK cells

Itgax-cre (Transgene) cDCs, pDCs Deletion in other cell types was not analyzed, most likely similar to other Itgax- cre strain

Clec9a-cre ~100% of cDC1s, ~50% of Begins deletion at CDP cDC2s, ~20% of pDCs stage, low or transient expression may affect deletion efficiency in cDC2s and pDCs

Zbtb46-cre 65-95% cDCs, <10% in all Deletion efficiency variable other cell types depending on floxed allele

Cd207-cre Langerhans cells, Langerin+ cDC1s in dermis, skin-draining LNs, and lung

Xcr1-cre cDC1s

Siglech-cre ~30% of pDCs, ~2% of Incomplete deletion in other lymphoid cells pDCs

Deletion in marginal zone macrophages untested but possible

Table 1.3. Comparison of transcription factor knockout mice that affect dendritic cell development

Strain Cells Depleted Caveats

Irf8-/- cDC1s, B cells, monocytes, Myeloid neoplasia eosinophils, basophils eventually results with age

Conditional deletion with Itgax-cre or Zbtb46-cre can prevent depletion of other cell types seen in Irf8-/- mice

Id2-/- cDC1s, NK cells, ILCs

Nfil3-/- cDC1s, NK cells, ILCs

Batf3-/- cDC1s In certain genetic backgrounds residual cDC1-like cells remain in skin-draining LNs

Relb-/- CD11b+ESAM+ cDC2s in Fatal multiorgan spleen inflammation

Ltbr-/- CD11b+ESAM+ cDC2s in Loss of lymph nodes, can spleen overcome with conditional deletion with Itgax-cre

Notch2f/f Itgax-cre CD11b+ESAM+ cDC2s in Transcriptional changes spleen, CD103+CD11b+ also evident in cDC1s and cDC2s in small intestine CD11b+ cDC2s lamina propria

Klf4f/f Itgax-cre Subset of cDC2s similar to CD24-CD11b- cDC2s in

skin-draining LNs

Irf4f/f Itgax-cre ~50% CD11b+ cDC2s in cDC2s cannot migrate, lung, ~50% failure to migrate may CD103+CD11b+ cDC2s in explain many observed small intestine lamina phenotypes propria, complete absence of cDC2s in lymph nodes May also affect M2 macrophages or GM-CSF activated monocytes

Tcf4f/f Itgax-cre pDCs

Zeb2f/f Itgax-cre pDCs

Figure 1.1. Functions of dendritic cell subsets in the immune response. Classical dendritic cells (cDCs) play critical roles in both innate and adaptive immunity. Furthermore, each subset of cDC appears to possess unique functions and to control the immune response against specific forms of pathogens. cDC1s control type 1 immune responses against viruses and intracellular pathogens (left panel). In these responses they prime naïve CD8 T cells, reactivate memory CD8 T cells, activate ILC1s, and induce Th1 cells. Their production of the cytokine IL-12 is vital for many of these functions. cDC1s also play a role in inducing Tregs against orally fed antigens and AIRE- dependent self-antigens expressed in the thymus, though it appears that in some cases cDC2s can also mediate Treg conversion to these antigens. Some cDC2s, on the other hand, regulate type 2 immune responses against parasites in which they induce Th2 cells (right panel). The exact mechanism by which they do this is unclear. Other cDC2s control type 3 immune responses against extracellular bacteria and fungi (right panel). In these responses, cDC2s produce IL-23 in order to activate ILC3s and to induce Th17 cells. Their production of IL-6 and TGF-β also contributes to the polarization of Th17 cells. Finally, cDC2s are also responsible for the induction of T follicular helper (Tfh) cells that regulate the germinal center response. DTR, diphtheria toxin receptor; VSV, vesicular stomatitis virus; LCMV, lymphocytic choriomeningitis virus; WNV, west nile virus; HSV, herpes simplex virus; CMV, cytomegalovirus; AIRE, autoimmune regulator; Ag, antigen.

CHAPTER 2:

Altered compensatory cytokine signaling underlies the discrepancy between Flt3–/– and

Flt3l–/– mice

The contents of this chapter have been previously published in the Journal of Experimental

Medicine.

Durai V, Bagadia P, Briseño CG, Theisen DJ, Iwata A, Davidson JT 4th, Gargaro M, Fremont

DH, Murphy TL, Murphy KM. Altered compensatory cytokine signaling underlies the discrepancy between Flt3-/- and Flt3l-/- mice. J Exp Med. 2018 May 7;215(5):1417-1435.

2.1 Abstract

The receptor Flt3 and its ligand Flt3L are both critical for dendritic cell (DC) development, but DC deficiency is more severe in Flt3l–/– mice than in Flt3–/– mice. This has led to speculation that Flt3L binds to another receptor that also supports DC development. However, we found that Flt3L administration does not generate DCs in Flt3–/– mice, arguing against a second receptor. Instead, Flt3–/– DC progenitors matured in response to M-CSF or SCF, and deletion of Csf1r in Flt3–/– mice further reduced DC development, indicating that these cytokines could compensate for Flt3. Surprisingly, Flt3–/– DC progenitors displayed enhanced M-CSF signaling, suggesting that loss of Flt3 increased responsiveness to other cytokines. In agreement, deletion of Flt3 in Flt3l–/– mice paradoxically rescued their severe DC deficiency. Thus, multiple cytokines can support DC development and the discrepancy between Flt3–/– and Flt3l–/– mice results from the increased sensitivity of Flt3–/– progenitors to these cytokines.

2.2 Introduction

Dendritic cells (DCs) are immune cells with critical functions in both the innate and adaptive immune response that develop from hematopoietic progenitor cells 1,2. The earliest committed progenitor with DC fate potential is the macrophage/DC progenitor (MDP) 3,4, which develops into a common DC progenitor (CDP) that can give rise to plasmacytoid DCs (pDCs) as well as the classical DC (cDC) subsets, cDC1s and cDC2s 5,6. Committed cDC progenitors restricted to only the cDC1 or the cDC2 lineage have recently been identified in mice 7,8 and in humans 9-11.

The development of DCs is dependent upon the class III receptor tyrosine kinase (RTK)

Fms-like tyrosine kinase 3 (Flt3) and its ligand Flt3L 12,13. Flt3 was first identified as a gene enriched in hematopoietic stem cells that encoded a protein homologous to the receptor c-Kit 14.

It was later recognized to be expressed on mature DCs and their progenitors as well 15. Flt3 shares structural properties and downstream signaling pathways with c-Kit and CSF1R, other members of the class III RTK family that are also expressed by committed DC progenitors 5,7,16.

The ligand for Flt3, Flt3L, was subsequently cloned and found to induce proliferation in early bone marrow (BM) progenitors 17. Later, a role for Flt3L in DC homeostasis was uncovered from the expansion of DCs in mice and humans who were administered this cytokine 18,19. Further, treatment of BM progenitors in vitro with Flt3L also supports the development of mature DCs

20,21, and Flt3+ progenitors preferentially gave rise to DCs in vivo 22. Finally, genetic inactivation of the Flt3 23 or Flt3l 12 genes in mice was observed to decrease the numbers of DCs 12,13, confirming their importance in DC homeostasis.

These original studies of Flt3–/– and Flt3l–/– mice surprisingly appeared to find DC deficiencies of varying severity in these two strains. Flt3l–/– mice analyzed between 5 and 14 weeks of age have a 4 to 10-fold reduction in splenic CD8α– DCs and a 6 to 14-fold reduction in splenic CD8α+ DCs 12. Meanwhile, an analysis of Flt3–/– mice found that while all DCs are reduced by 85% at two weeks of age, they are reduced by only 43% (cDCs) or 65% (pDCs) at 9 weeks of age 13. Another study that examined both strains between 8 and 12 weeks of age similarly found more severe reductions in CD8α+ DCs and CD11b+ DCs in Flt3l–/– mice compared to Flt3–/– mice 24. This discrepancy has also been noted in the development of pre-pro

B cells, with Flt3l–/– mice demonstrating a severe reduction in this population while Flt3–/– mice have only a two-fold reduction 23,25-27. However, no study has directly compared Flt3–/– mice

61 with Flt3l–/– mice over time, so it is unknown if the defects caused by Flt3L deficiency improve with age, as with Flt3 deficiency, or whether Flt3–/– mice and Flt3l–/– mice simply have distinct phenotypes. Nevertheless, these apparent discrepancies in phenotypes have led to suggestions that Flt3L could act on a second receptor other than Flt3 to support DC or B cell development, explaining the more severe defects in Flt3l–/– mice 27-31.

Cytokines such as Flt3L can play supportive and instructive roles in the development of hematopoietic lineages 32,33, and the actions of a given cytokine may depend upon the progenitor stage and cell type it acts upon 34,35. For DCs, an instructive role for Flt3 was suggested by the redirection of megakaryocyte-erythrocyte progenitors (MEPs) toward DC fate upon forced Flt3 overexpression 36 and by transgenic mice in which Flt3L overexpression leads to a robust expansion of DCs, lymphocytes, and myeloid cells at the expense of erythrocytes and platelets 37.

Nonetheless, Flt3L administration does not increase in vivo DC output from early Flt3+ progenitors such as CLPs, CMPs, or GMPs 38, and the continued presence of DCs in Flt3–/– and

Flt3l–/– mice, albeit at reduced numbers, suggests that DC development can occur through some unidentified mechanism in the absence of this cytokine pathway. Thus, it remains unclear what specific role is played by Flt3L in DC progenitor specification and development, as well as whether any other cytokines can contribute to this process.

To address the apparent discrepancy in phenotypes between Flt3–/– and Flt3l–/– mice, we directly compared DC development in these strains over time. We confirmed that Flt3l–/– mice show a severe and persistent DC defect while Flt3–/– mice have a less severe defect at all ages analyzed. However, we were unable to demonstrate activity for Flt3L on a second receptor as has been proposed. Instead, we found that DC progenitors developed normally without instructional cues from Flt3 signaling, and surprisingly that these progenitors could mature in Flt3–/– mice in

62 response to M-CSF and SCF, ligands for other class III RTKs expressed by these cells. Deletion of Csf1r in Flt3–/– mice also led to a further reduction in DC development in a cell-intrinsic manner, indicating that this cytokine pathway was partially compensating for the loss of Flt3 in vivo.

Although the expression of CSF1R and c-Kit was not altered in Flt3–/– progenitors, we found that these progenitors were more responsive to stimulation by M-CSF, suggesting that the absence of

Flt3 potentiates signaling by other class III RTKs. To test whether this effect was responsible for the discrepancy between Flt3–/– and Flt3l–/– mice, we generated Flt3–/–Flt3l–/– mice and found that the additional deletion of Flt3 paradoxically restored DC numbers in Flt3l–/– mice. We conclude that multiple cytokine pathways can support the development of DCs, and that the discrepancy between Flt3–/– and Flt3l–/– mice is due to the enhanced sensitivity of Flt3–/– BM progenitors to signaling by these alternative cytokines.

2.3 Results

Flt3l–/– mice have a more severe DC deficiency than Flt3–/– mice

The original studies that analyzed Flt3–/– and Flt3l–/– mice did not directly compare these strains at similar ages, so the previously noted phenotypic discrepancy may simply have been due to mismatched comparisons 12,13. We therefore examined matched sets of Flt3–/– and Flt3l–/– mice at both 2 and 8 weeks of age (Figs. 2.1 and 2.3). We confirmed the previous report of severe reductions in the percent (Fig. 2.1, C and E) and numbers (Fig. 2.1, D and F) of splenic cDCs and pDCs at 2 weeks in both Flt3–/– and Flt3l–/– mice. We also found severe reductions in both populations in the skin draining lymph nodes (SLNs) (Fig. 2.2, B and C), and lungs (Fig.

2.2G) of these strains. Even at this age, however, the defect in cDCs and pDCs was more severe

63 in Flt3l–/– mice than in Flt3–/– mice in all organs analyzed. We also observed previously unrecognized heterozygous phenotypes in both Flt3+/– and Flt3l+/– mice compared with wild type

(WT) mice. Flt3l+/– mice showed a 2-fold reduction in DC numbers relative to WT mice, while surprisingly, Flt3+/– mice had significantly more DCs than WT mice (Fig. 2.1, C-F).

By contrast, at 8 weeks, Flt3–/– mice showed only a mild deficiency in the percent and number of splenic DCs, while Flt3l–/– mice continued having severely reduced DC populations compared to both WT and Flt3–/– mice (Fig. 2.3, A-F). Findings were similar for cDCs and pDCs in the SLNs (Fig. 2.3, D and E) and for cDCs in the lungs (Fig. 3H). Heterozygous phenotypes were again observed at 8 weeks, with Flt3l+/– mice showing a 2-fold reduction in DCs compared with WT mice and Flt3+/– mice having consistently greater numbers of DCs compared with WT mice (Fig. 2.3, A-F). In summary, the more severe DC defect in Flt3l–/– mice compared to Flt3–/– mice appears to be an inherent difference between these strains.

Flt3L does not support DC development from Flt3–/– BM or in Flt3–/– mice

To explain the discrepancy between Flt3–/– and Flt3l–/– mice, some have suggested that

Flt3L could support DC development in Flt3–/– mice by acting on a second receptor, thus explaining the less severe phenotype of Flt3–/– mice 27-31. To test for evidence of a second Flt3L receptor, we cultured WT and Flt3–/– BM in vitro with Flt3L and assessed DC development, hypothesizing that Flt3L could bind to its second receptor in Flt3–/– BM and generate DCs.

However, while Flt3L supported robust DC development from WT BM, it did not generate DCs from Flt3–/– BM (Fig. 2.4, A-E). Thus, in vitro, we were unable to provide evidence for a second receptor responding to Flt3L. We next administered Flt3L in vivo (Fig. 2.4, F-J). Flt3L

64 administration to WT mice caused a significant increase in the percent and numbers of splenic

DCs, as expected 18, but failed to increase DCs in Flt3–/– mice (Fig. 2.4, F-J). In summary, we find no evidence for actions of Flt3L on a second receptor.

Since DCs in Flt3–/– mice appeared to arise independently of Flt3L, we wondered whether they were developmentally or functionally impaired. We therefore examined the transcriptional profile of splenic cDC1s and cDC2s from WT and Flt3–/– mice using gene expression microarrays. We found no substantial changes in gene expression in either cDC subset, with the only annotated transcript showing a greater than 4-fold change being Flt3 itself

(Fig. 2.5, A and B). In addition, we found that Flt3–/– cDC1s were able to cross-present cell- associated and soluble antigen to CD8 T cells as efficiently as WT cDC1s (Fig. 2.5, C-E), suggesting they were functionally similar. We next examined the transcriptional profile of splenic pDCs from WT and Flt3–/– mice (Fig. 2.5F). We again found only minor differences between these populations, with Flt3 being the only annotated transcript showing a greater than

4-fold increase in expression in WT pDCs and with several genes encoding Igκ segments being more highly expressed in Flt3–/– pDCs. pDCs have been found to develop from both myeloid and lymphoid progenitors 39, with pDCs derived from lymphoid progenitors showcasing rearrangements of D-J segments in the immunoglobulin genes. The increased expression of these genes encoding Igκ segments in Flt3–/– pDCs could therefore indicate that the pDCs in Flt3–/– mice preferentially arise from lymphoid rather than myeloid progenitors. Other than these genes, we found that the expression of the vast majority of transcripts was similar between WT and

Flt3–/– pDCs (Fig. 2.5F). Finally, we stimulated WT and Flt3–/– pDCs with CpG oligodeoxynucleotides and found that both populations produced a comparable amount of IFN-α

(Fig. 2.5G). Together, these results suggest that the DCs in Flt3–/– mice are transcriptionally and

65 functionally similar to WT DCs, implying that Flt3 signaling is not essential for generating functional DCs.

Committed DC progenitors develop in Flt3–/– BM and can mature in response to M-CSF and SCF

We next considered whether Flt3–/– DCs could be maturing in response to alternative cytokines. The receptors for M-CSF (CD115/CSF1R) and for SCF (CD117/c-Kit) are both expressed by the MDP and CDP 40, suggesting that these cytokines could act on these progenitors, and M-CSF has also been reported to support the development of some DC subsets in vitro 41. To help identify cDCs in the Flt3–/– background, we crossed Flt3–/– mice to

Zbtb46GFP/+ mice to generate Flt3–/–Zbtb46GFP/+ mice, in which we could use Zbtb46GFP expression to distinguish cDCs from other lineages 42-44. We then cultured bulk BM from WT

(Flt3+/+Zbtb46GFP/+), Flt3+/–Zbtb46GFP/+, and Flt3–/–Zbtb46GFP/+ mice with Flt3L, M-CSF, or

SCF and assessed DC development. Surprisingly, we found that all three cytokines supported the development of cDC1s and cDC2s from WT and Flt3+/– BM, while M-CSF and SCF also supported cDC development from Flt3–/– BM (Fig. 2.6, A and B). Notably, M-CSF and SCF generated a greater percent (Fig. 2.6B) and number (Fig. 2.7A) of cDCs from Flt3–/– BM than from WT BM, suggesting that Flt3–/– BM may be more responsive to these cytokines than WT

BM. Importantly, the cDCs generated in all cultures were uniformly positive for Zbtb46GFP expression (Fig. 2.6A), confirming their identity as bona fide cDCs 42-44.

To further characterize these cells and to identify any heterogeneity in the populations, we examined the expression of a variety of functional markers in cDCs from each culture

66 condition. We found that the cDCs generated in response to each cytokine generally expressed similar levels of costimulatory molecules such as CD86, PD-L2, CD40, and CD205, surface markers such as CD24, CD172a, and CD11b, DC specific markers such as Flt3/CD135 (the cDCs from Flt3L cultures expressed low levels of this receptor, potentially due to downregulation after exposure to Flt3L), CCR7, and CD117, and that they all expressed similarly low levels of macrophage markers such as CD115 and CD64 (Fig. 2.7B). Thus, we concluded that the cDCs that mature in response to M-CSF and SCF are generally similar in phenotype to those that are generated by Flt3L. Finally, we found that Flt3L, M-CSF, and SCF could all support pDC development from WT BM and that M-CSF and SCF could also support pDC development from Flt3–/– BM (Fig. 2.8A).

Since mature DCs could be generated from the BM of Flt3–/– mice, we wondered if DC progenitor specification was occurring normally in these mice. Identifying the CDP relies on Flt3 expression 5 and is therefore not possible in Flt3–/– mice. However, two restricted cDC progenitors downstream of the CDP, the pre-cDC1 and pre-cDC2, can be identified with

Zbtb46GFP expression and additional markers without the need for Flt3 7. We therefore analyzed

BM from Flt3–/–Zbtb46GFP/+ mice to identify these progenitors, and found normal frequencies of pre-cDC1s and pre-cDC2s compared with Flt3+/+Zbtb46GFP/+ mice (Fig. 2.6C). As a control, BM from Irf8–/–Zbtb46GFP/+ mice lacked pre-cDC1s and had reduced pre-cDC2s, as expected 7. Thus, cDC progenitor specification appears to be independent of Flt3 signaling.

We next asked whether M-CSF and SCF could act directly on committed DC progenitors to support their maturation. To address this question, we sort purified WT CDPs, which express

Flt3, CSF1R, and c-Kit, and cultured them with Flt3L, M-CSF or SCF. These cytokines each supported development of the CDP into mature Zbtb46GFP-positive cDC1s and cDC2s (Fig.

2.9A) as well as pDCs (Fig. 8B). In addition, pre-cDC2s, which normally express CSF1R and

Flt3, were isolated from Flt3+/+ mice and Flt3–/– mice and cultured with Flt3L and M-CSF. As expected, Flt3L supported development of the pre-cDC2 into mature Zbtb46GFP-positive cDC2s from Flt3+/+ progenitors but not from Flt3–/– progenitors (Fig. 2.9B). In contrast, M-CSF supported development of the pre-cDC2 into Zbtb46GFP-positive cDC2s from both WT and Flt3–

/– progenitors (Fig. 2.9B). When we analyzed cell divisions using cell proliferation dye tracing, we found that in response to M-CSF Flt3–/– pre-cDC2s proliferated more than WT pre-cDC2s

(Fig. 2.9C), again suggesting that Flt3–/– progenitors were more responsive to M-CSF than WT progenitors. Thus, our data suggests that specified DC progenitors that develop in the presence or absence of Flt3 signaling can mature in response to cytokines other than Flt3L.

DC development in Flt3–/– mice is substantially dependent upon CSF1R

To determine whether M-CSF supported the in vivo DC development observed in Flt3–/– mice, we deleted Csf1r 45 in WT and Flt3–/– mice using a tamoxifen-inducible Cre recombinase

46. Deletion of Csf1r in WT mice caused a small decrease in splenic cDC percentage and numbers, consistent with a previous report describing a minor role for CSF1R in cDC homeostasis (Fig. 2.10, A, C, and D) 24. In contrast, deletion of Csf1r in Flt3–/– mice led to a substantial reduction in cDCs (Fig. 2.10, B-D), confirming that cDCs developing in Flt3–/– mice arise in part through the actions of CSF1R. pDC development, on the other hand, was unaffected by the additional deletion of Csf1r (Fig. 2.10, E and F), consistent with prior studies showing that pDC progenitors are enriched in the CSF1R– fraction of BM progenitors 47 and that almost all pDCs are derived from an IL-7R+ progenitor 48. Our gene expression data had also indicated that

68 the pDCs in Flt3–/– mice may be preferentially developing from lymphoid progenitors (Fig.

2.5F), which, combined with the lack of a role for CSF1R, suggests that the pDCs in Flt3–/– mice may be developing in response to compensatory signaling from a receptor expressed on lymphoid progenitors such as IL-7R. In summary, a substantial portion of cDC but not pDC development in Flt3–/– mice is dependent upon CSF1R.

To test whether this action of CSF1R was cell-intrinsic to DC progenitors, we generated mixed bone marrow chimeras with a 50:50 mix of Flt3–/– BM and either Flt3–/–Csf1rf/fR26+/+ BM

(CreERT2–) or Flt3–/–Csf1rf/fR26CreERT2/+ BM (CreERT2+). While all BM progenitors in these chimeras were deficient in Flt3, tamoxifen treatment led to the additional deletion of CSF1R in half of the BM cells in CreERT2+ chimeras but not in CreERT2– chimeras (Fig. 2.10G). Thus, we could directly compare the DC potential of progenitors lacking both Flt3 and CSF1R with progenitors lacking only Flt3. In CreERT2+ chimeras, cDCs preferentially developed from Flt3–

/– BM cells, rather than from Flt3–/–Csf1r–/– BM, while in CreERT2– chimeras there was no skew in DC development between Flt3–/– and Flt3–/–Csf1rf/fR26+/+ BM (Fig. 2.10, G and H). This result suggests that the contribution of CSF1R to DC development in Flt3–/– progenitors is cell- intrinsic.

Flt3–/– progenitors express normal levels of CSF1R and c-Kit but display enhanced signaling from these receptors

Since M-CSF and SCF could support DC development independently of Flt3, we wondered if the discrepancy between Flt3–/– and Flt3l–/– mice might be due to differences in the expression or signaling of CSF1R or c-Kit in these different strains. First, we found no

69 differences in surface expression of CSF1R or c-Kit in total BM isolated from WT, Flt3–/–, and

Flt3l–/– mice (Fig. 2.11A). We next sought to quantify the expression of these receptors in early

DC progenitors, but as Flt3 is required to identify the MDP and CDP 5, we could not analyze these progenitors in Flt3–/– mice. We therefore generated Flt3l–/–Zbtb46GFP/+ mice to identify the pre-cDC1 and pre-cDC2 in Flt3l–/– mice, which would allow us to compare these progenitors in

WT, Flt3–/–, and Flt3l–/– mice. We found that both progenitors still developed in Flt3l–/– mice, but were substantially fewer in number relative to WT and Flt3–/– mice (Fig. 2.12A). This was similar to the reported deficiency in MDPs and CDPs in Flt3l–/– mice 49, suggesting that all DC progenitors are affected by the loss of Flt3L. We found, however, that the expression of CSF1R and c-Kit was not different in pre-cDC1s or pre-cDC2s from Flt3–/– or Flt3l–/– mice relative to

WT mice (Fig. 2.11B), again suggesting that changes in receptor expression did not explain the differences between these strains. Finally, we sought to determine the expression of these receptors in mature splenic cDCs from these strains, but as collagenase digestion of the spleen impairs detection of these receptors by FACS 50, we instead used gene expression microarrays.

We found no differences in Csf1r or Kit gene expression in splenic cDC1 or cDC2s from WT,

Flt3–/–, and Flt3l–/– mice (Fig. 2.11C). Thus, we excluded differences in receptor expression as a cause of the discrepancy between Flt3–/– and Flt3l–/– mice.

The in vitro DC development supported by M-CSF and SCF was greater in Flt3–/– BM relative to WT BM (Fig. 2.6B) and Flt3–/– pre-cDC2s proliferated more than WT pre-cDC2s in response to M-CSF (Fig. 2.9C), suggesting there was some alteration in receptor sensitivity between WT and Flt3–/– mice. Flt3, CSF1R, and c-Kit are all class III RTKs that share signaling adaptor molecules, such as the Src-homology and collagen (SHC) proteins and Grb2 16,31,51,52.

We wondered whether the loss of Flt3 might increase the availability of these adaptors and

70 thereby enhance signaling from CSF1R or c-Kit. To test this, we quantified the sensitivity of

CSF1R signaling in BM progenitors. We treated serum-starved BM cells from Flt3+/+, Flt3+/–, or Flt3–/– mice with varying concentrations of M-CSF and quantified phosphorylation of the

MAPK pathway protein Erk1/2 in pre-cDC2s (Fig. 2.13A). We examined this progenitor because it expresses CSF1R and can be identified independently of Flt3 by using Zbtb46-GFP expression

7. Over a range of concentrations, both the percentage of pre-cDC2s responding to M-CSF and the intensity of signaling in each progenitor was greater in Flt3–/– pre-cDC2s compared to Flt3+/+ pre-cDC2 cells, with Flt3+/– pre-cDC2s displaying an intermediate phenotype (Fig. 2.13, A and

B). Thus, the loss of Flt3 directly enhances the sensitivity and strength of CSF1R signaling in response to M-CSF. Importantly, no differences were observed in M-CSF signaling sensitivity in

Flt3l–/– pre-cDC2s relative to WT for most concentrations of M-CSF (Fig. 2.12B), indicating that the increased sensitivity to M-CSF was specific to Flt3–/– mice and could possibly explain the difference between these mice and Flt3l–/– mice.

Deletion of Flt3 in Flt3l–/– mice paradoxically restores DC development

The increased sensitivity of CSF1R signaling in Flt3–/– DC progenitors could explain the discrepancy in DC numbers between Flt3–/– and Flt3l–/– mice. Flt3–/– DC progenitors could respond more robustly to alternative cytokines such as M-CSF or SCF and thus develop greater numbers of DCs. This hypothesis predicts that the increased CSF1R sensitivity in Flt3–/– mice should occur regardless of the presence of Flt3L, and thus that loss of Flt3 in the Flt3l–/– background should enhance CSF1R signaling and potentially increase DC numbers. To test this hypothesis, we crossed Flt3–/– and Flt3l–/– mice to generate Flt3–/–Flt3l–/– mice and characterized DC development

71 in the resulting littermates. We first found that, unlike Flt3+/– mice, Flt3+/–Flt3l+/– mice had mildly reduced DC numbers (Fig. 2.14, A-E). This suggests that the decrease in DCs we found in Flt3l+/– mice (Fig. 2.1 and 2.3) was more impactful than the increase in DCs observed in Flt3+/– mice (Fig.

2.1 and 2.2), and that together these alleles resulted in a net decrease in DC numbers in Flt3+/–

Flt3l+/– mice. But most surprisingly, we found that Flt3–/–Flt3l–/– mice had an increase in splenic cDC and pDC percentage and numbers relative to Flt3l–/– mice and were indistinguishable from

Flt3–/– mice (Fig. 2.14, A-E). This increase was also observed in cDCs and pDCs in the SLNs (Fig.

2.15, A and B), as well as in cDCs in the lung (Fig. 2.15C). Thus, paradoxically, the severe DC deficiency observed in Flt3l–/– mice was rescued by the additional loss of Flt3. These results suggest that the discrepancy observed between Flt3–/– and Flt3l–/– mice is, at least in part, due to an inherent alteration in the response of DC progenitors to alternative cytokines secondary to the loss of Flt3.

2.4 Discussion

In this study, we have identified the previously unknown cytokine pathways that support the residual DC development seen in Flt3–/– and Flt3l–/– mice, and we provide an explanation for the puzzling discrepancy in phenotypes between these two strains. While both Flt3 and Flt3L have long been recognized to play major roles in DC homeostasis 12,13, it had remained unclear through what mechanism DCs continue to arise in mice deficient in either of these factors. We have demonstrated here that M-CSF and SCF, two cytokines whose receptors are expressed in

DC progenitors, are capable of generating DCs from both WT and Flt3–/– bone marrow progenitors in vitro. We also found that a substantial fraction of the DCs that arise in vivo in

Flt3–/– mice are dependent upon CSF1R for their development, with c-Kit possibly responsible for generating the remainder of the DCs.

Loss of a receptor and its unique ligand would generally be expected to produce the same phenotype. However, our direct comparison of Flt3–/– and Flt3l–/– mice clearly document that loss of the ligand causes a more severe and durable DC deficiency than loss of the receptor. We were unable to find evidence to support previous suggestions that Flt3L might be acting at a second receptor besides Flt3 27-31. Instead, we found that Flt3–/– DC progenitors exhibit an intrinsic alteration in the sensitivity of other cytokine receptors compared with WT progenitors.

This may allow compensatory cytokines such as M-CSF and SCF to generate a greater number of DCs in Flt3–/– mice relative to WT or Flt3l–/– mice. We therefore predicted, and then validated, the paradoxical result that the severe DC defect in Flt3l–/– mice is actually restored upon deletion of Flt3. Thus, it appears that the discrepancy in DC development between Flt3–/– and Flt3l–/– mice arises secondary to a phenotype of increased sensitivity of Flt3–/– DC progenitors to alternative cytokines. It is possible that a similar increase in sensitivity to IL-7 in early B cell progenitors could explain the previously identified discrepancy in B cell development between these strains 27.

The increased sensitivity of Flt3+/– progenitors to M-CSF (Fig. 9) might also explain the increased DC numbers we observed in Flt3+/– mice (Figs. 1 and 2). Flt3+/– BM produced DCs in response to Flt3L as well as to M-CSF (Fig. 5B), so the increased DC numbers seen in Flt3+/– mice conceivably results from the combination of both normal Flt3L signaling and enhanced M-

CSF signaling contributing to DC development in these mice. By contrast, in Flt3–/– mice only enhanced M-CSF signaling but not Flt3L signaling contributes to DC development, and these mice therefore have fewer DCs despite their even greater sensitivity to M-CSF.

We have not explored the biochemical basis for the altered sensitivity observed in Flt3–/–

DC progenitors. Conceivably, loss of Flt3 increases the availability of shared signaling adaptor molecules such as SHC and Grb2 used by other class III RTKs 16,31,51,52. The increased availability of these adaptors could then allow other receptors acting through them to activate downstream pathways more quickly and potently. While the exact mechanism remains to be uncovered, our results highlight the potential for unintended consequences that may result when manipulating the levels of signaling molecules in complex biological systems. The deletion of Flt3 appears to have ramifications beyond simply the loss of signaling by this receptor, and the study of other receptor knockout mice may have similar caveats that should be considered.

2.5 Materials and Methods

Mice.

All mice were bred and maintained on the C57BL/6 background in a specific pathogen- free animal facility following institutional guidelines and with protocols approved by the Animal

Studies Committee at Washington University in St. Louis. Unless otherwise specified, experiments were performed with mice 6–10 weeks of age. No differences were observed between male and female mice in any assays and so mice of both genders were used throughout this study. Within individual experiments mice used were age- and sex-matched littermates whenever possible.

Flt3–/– mice 23 were a gift from Dr. E. Camilla Forsberg. Zbtb46GFP/+ mice have been previously described 7,42. The following mice were purchased from The Jackson Laboratory:

Flt3l–/– (C57BL/6-Flt3ltm1Imx/TacMmjax), CD45.1+ (B6.SJL-Ptprca Pepcb/BoyJ), OT-I

(C57BL/6-Tg(TcraTcrb)1100Mjb/J), Csf1rf/f (B6.Cg-Csf1rtm1.2Jwp/J), and R26CreERT2/+ (B6.129-

Gt(ROSA)26Sortm1(cre/ERT2)Tyj/J).

Dendritic cell preparation.

Lymphoid and nonlymphoid organ DCs were harvested and prepared as described previously 42. Briefly, spleens and SLNs (inguinal, axillary, and brachial) were minced and digested in 5 mL of Iscove’s modified Dulbecco’s media (IMDM) + 10% FCS (cIMDM) with

250 μg/mL collagenase B (Roche) and 30 U/mL DNaseI (Sigma-Aldrich) for 45 min at 37°C with stirring. Lungs were minced and digested in 5 mL of cIMDM with 4 mg/mL collagenase D

(Roche) and 30 U/mL DNaseI (Sigma-Aldrich) for 1.5 hours at 37°C with stirring. After digestion, cell suspensions from all organs were passed through 70-μm strainers and red blood cells were then lysed with ammonium chloride-potassium bicarbonate (ACK) lysis buffer. Cells were counted with a Vi-CELL analyzer (Beckman Coulter) and 3-5x106 cells were used per antibody staining reaction.

For experiments requiring sorting of splenic cDC subsets, cDCs were first enriched from total spleen cells with CD11c microbeads (Miltenyi Biotec). cDCs were then sorted on a

FACSAria Fusion. cDC1s were sorted as CD11c+MHC-II+CD24+CD172a–F4/80– and cDC2s were sorted as CD11c+MHCII+CD24–CD172a+F4/80–. Sort purity of >95% was confirmed by post-sort analysis before cells were used for further experiments.

For experiments requiring sorting of splenic pDC subsets, spleen cells were first depleted of CD3-, B220-, and Ly6G-expressing cells by staining with the corresponding biotinylated antibodies followed by depletion with MagniSort Streptavidin Negative Selection Beads

(Thermo Fisher). pDCs were then sorted on a FACSAria Fusion as B220+CD317+. Sort purity of

>95% was confirmed by post-sort analysis before cells were used for further experiments.

Antibodies and flow cytometry.

Samples were stained at 4°C in MACS buffer (DPBS + 0.5% BSA + 2 mM EDTA) in the presence of Fc block (2.4G2; BD Biosciences). The following antibodies were purchased from

BD Biosciences: CD8α V450 (53-6.7); CD11b PE (M1/70); CD19 Biotin (1D3); CD44 APC-

Cy7 (IM7); CD45R AlexaFluor 488, V450 (RA3-6B2); CD45.2 PerCP-Cy5.5 (104); CD64

Alexa Fluor 647 (X54-5/7.1); CD103 BV421 (M290), CD117 BUV395 (2B8); CD127 BV421

(SB/199); CD135 PE-CF594 (A2F10.1); Ly-6C Alexa Fluor 700 (AL-21); I-A/I-E V500

(M5/114.15.2). From eBioscience: CD11c PE (N418); CD44 APC (IM7); CD105 Biotin

(MJ7/18); CD117 PE/Cy7 (2B8); CD172a PerCP-eFluor710 (P84); CD317 APC (eBio927);

F4/80 APC-eFluor 780 (BM8). From Invitrogen: TCR Vα2 PE (B20.1); CD172a PerCP- eFluor710 (P84). From Tonbo Biosciences: CD3e Biotin (145-2C11); CD11c APC-Cy7 (N418);

CD45.1 PE-Cy7 (A20); I-A/I-E V450 (M5/114.15.2). From BioLegend: CD4 APC (RM4-5);

CD24 PE/Cy7 (M1/69); CD45.1 BV605 (A20); CD45.2 Alexa Fluor700 (104); CD115 BV711

(ASF98); Ly-6G Biotin (1A8); GFP Alexa Fluor488 (FM264G); Siglec H PE (551); Ter-119

Biotin (TER-119); TCR Vα2 PE, PerCP-Cy5.5 (B20.1). From Cell Signaling: P-p44/42 MAPK

(Erk1/2) (T202/Y204) PE (197G2). In general, all antibodies were used at a dilution of 1:200 unless specified otherwise by the manufacturer. Cells were analyzed on a FACSCanto II or

FACSAria Fusion. Data was analyzed with FlowJo software (TreeStar).

Bone marrow isolation and cell culture.

Bone marrow (BM) was harvested from the femur, tibia, and pelvis. Bones were fragmented by mortar and pestle in MACS buffer, and debris was removed by passing cells through a 70-μm strainer. Red blood cells were subsequently lysed with ACK lysis buffer. Cells were counted on a Vi-CELL analyzer (Beckman Coulter), and 3-5x106 cells were stained for analysis. For Flt3L culture experiments, whole BM (8x106 cells in 4 mL of cIMDM) was cultured at 37°C with 100 ng/mL of Flt3L (Peprotech) or equivalent volume of PBS (vehicle) for

9 days. For comparative cytokine culture experiments, whole BM (6x106 cells in 4 mL of cIMDM) was cultured at 37°C with 100 ng/mL of Flt3L (Peprotech), 20 ng/mL M-CSF

(Peprotech), 50 ng/mL of SCF (Peprotech), or equivalent volume of PBS (vehicle) for 7 days.

For all culture experiments, loosely adherent and suspension cells were harvested by gentle pipetting at the indicated time point and stained with fluorescent antibodies for analysis by flow cytometry.

In vivo Flt3L administration.

Flt3+/+ or Flt3–/– mice were injected subcutaneously with sterile saline (vehicle) or 10 ug of purified Flt3L-Fc fusion protein (generated in-house) daily for 8 days and analyzed on the following day.

Expression microarray analysis.

Total RNA was extracted from sorted splenic DCs with an RNAqueous-Micro kit

(Ambion), amplified with an Ovation Pico WTA System (NuGEN), and hybridized to Mouse

Gene 1.0 ST arrays (Affymetrix). Expression values were analyzed after robust multiarray average (RMA) summarization and quantile normalization using ArrayStar 4 software

(DNASTAR). All gene expression microarray data has been deposited in the NCBI Gene

Expression Omnibus database with accession numbers GSE110789 and GSE110790.

Antigen presentation assays.

In vitro cell-associated cross-presentation assays have been described previously 57.

Briefly, Listeria monocytogenes (LM)-OVA (a gift from H. Shen) was grown in Brain-Heart

Infusion broth at 37°C for 6 hours and then frozen overnight after dilution plating for titer enumeration. Bacteria was subsequently thawed and washed three times with PBS before heat killing at 80°C for 1 hour and then freezing at -80°C. OT-I T cells were meanwhile sorted from spleens of OT-I mice as B220–CD4–CD11c–CD45.1+CD8+Vα2+ and labeled with CFSE

(eBioscience). Sort purity of >95% was confirmed by post-sort analysis before cells were used for assays. 10,000 sorted splenic cDCs of the indicated subset from Flt3+/+ or Flt3–/– mice and

25,000 sorted CFSE-labeled OT-I T cells were incubated with various doses of heat-killed LM-

OVA for 3 days at 37°C. After incubation was complete, OT-I cells were assayed for CFSE dilution and CD44 expression.

For soluble OVA presentation assays, 10,000 sorted splenic cDCs of the indicated subset from Flt3+/+ or Flt3–/– mice and 25,000 sorted CFSE-labeled OT-I T cells were incubated with the indicated concentration of OVA (Worthington Biochemical Corporation) for 3 days at 37°C.

After incubation was complete, OT-I T cells were assayed for CFSE dilution and CD44 expression.

Alternatively, sorted splenic cDCs of the indicated subset from Flt3+/+ or Flt3–/– mice were incubated with 0.1 mg/mL of soluble OVA (Worthington Biochemical Corporation) for 45 minutes at 37°C, washed three times with PBS, and then different numbers of cDCs were incubated with 25,000 sorted CFSE-labeled OT-I T cells for 3 days at 37°C. After incubation was complete, OT-I T cells were assayed for CFSE dilution and CD44 expression.

Type I interferon production from pDCs.

25,000 sorted pDCs from Flt3+/+ or Flt3–/– mice were incubated overnight at 37°C in cIMDM with 100 ng/mL of Flt3L plus either 6 ug/mL of CpG-A oligodeoxynucleotides or the equivalent volume of PBS (vehicle). The next day cells were spun down and the supernatant was collected. IFN-α in the supernatant was measured using the PBL Verikine Mouse IFN Alpha All

Subtype ELISA Kit, High Sensitivity (PBL Assay Science).

Progenitor sorting and culture.

For sorting experiments, BM was isolated as described above and depleted of CD3-,

CD19-, CD105-, Ter119-, and Ly6G-expressing cells by staining with the corresponding biotinylated antibodies followed by depletion with MagniSort Streptavidin Negative Selection

Beads (Thermo Fisher). The entirety of the remaining BM was then stained with fluorescent antibodies prior to sorting. Gates used to define CDPs, pre-cDC1s, and pre-cDC2s were based on

79 previous studies 5,7. CDPs were identified as Lin–CD117intCD135+CD115+CD11c–MHCII–; pre- cDC1s as Lin–CD117intCD11c+MHCIIintZbtb46-GFP+; and pre-cDC2s as Lin–

CD117loCD115+CD11c+MHCII–Zbtb46-GFP+. Lineage markers included CD3, CD19, CD105,

Ter119, and Ly6G. A FACSAria Fusion was used for sorting, and cells were sorted into cIMDM.

Sort purity of >95% was confirmed by post-sort analysis before cells were used for further experiments. Sorted cells were cultured at 37°C in 200 μL total volume of cIMDM with 100 ng/mL Flt3L (Peprotech), 20 ng/mL M-CSF (Peprotech), 50 ng/mL SCF (Peprotech), or equivalent volume of PBS (vehicle) for 5 days.

For cell proliferation studies, sorted pre-cDC2s were washed three times with PBS to remove serum and then incubated with 10 μM of eBioscience Cell Proliferation Dye eFluor 450 (Thermo

Fisher Scientific) at 37°C for 10 minutes. Reaction was quenched by the direct addition of FCS and cells were washed three times with cIMDM. Cells were subsequently cultured at 37°C in

200 μL total volume of cIMDM with 20 ng/mL M-CSF (Peprotech) for 5 days.

Tamoxifen administration.

Mice were orally gavaged with 4 mg of tamoxifen (Sigma Aldrich) dissolved in corn oil

(Sigma Aldrich) for an initial loading dose and then placed on tamoxifen citrate chow (Envigo) for 4-5 weeks. Mice were given up to two days of regular chow per week if significant weight loss was observed. After treatment, mice were rested on regular chow for one week before analysis. Deletion of CSF1R in the BM was confirmed in all mice.

Bone marrow chimeras.

Bone marrow cells from donor mice were collected as described above. To generate mixed bone marrow chimeras, bone marrow cells were isolated from mice of the appropriate genotypes, counted with a Vi-CELL analyzer (Beckman Coulter), and mixed at a ratio of 1:1 before transplantation. Recipient mice received a single dose of 1050 rads of whole-body irradiation and then received a transplant of 10x106 total BM cells the following day. Mice were allowed to reconstitute for 8-10 weeks before further experiments or analyses were conducted.

Bone marrow progenitor cytokine stimulation.

BM was isolated and lineage-depleted as described above. Lin– BM cells were then aliquoted at 1x106 cells per condition and serum-starved in plain IMDM for 4 hours at 37°C.

Following starvation, M-CSF was added at the indicated concentrations and cells were incubated at 37°C for 5 minutes for stimulation. Cells were then fixed for 15 minutes at room temperature by adding paraformaldehyde (Electron Microscopy Sciences) directly to cells to a final concentration of 2%. Cells were washed twice with MACS buffer and then permeabilized by adding ice-cold methanol directly to cells with gentle vortexing followed by incubation at 4°C for 30 minutes. Cells were again washed twice with MACS buffer and then stained with fluorescent antibodies against cell-surface and intracellular antigens for 30 minutes at room temperature. Antibodies were previously tested individually to ensure their functionality in fixed and permeabilized cells. Flow cytometric analyses were then performed on stained cells. For analysis, we used integrated MFI, which combines frequency and MFI into a single measure of total functional response 58,59.

Statistical analysis

All statistical analyses were performed using Prism 7 (GraphPad Software). Statistical significance was evaluated using unpaired two-tailed Student's t-test unless otherwise noted.

Differences with P≤0.05 were considered significant. n represents the number of biological replicates.

2.6 Acknowledgments

We thank E. Camilla Forsberg for Flt3–/– mice and the Alvin J. Siteman Cancer Center at

Washington University School of Medicine for use of the Center for Biomedical Informatics and

Multiplex Gene Analysis Genechip Core Facility. We thank Ansuman T. Satpathy and Xiaodi

Wu for their helpful suggestions on this manuscript. We thank Hannah L. Miller for technical help with pDC assays.

This work was supported by the Howard Hughes Medical Institute (K.M. Murphy) and the

US National Institutes of Health (F30DK108498 to V. Durai). The authors declare no competing financial interests.

2.7 Author Contributions

V. Durai, T.L. Murphy, and K.M. Murphy designed the study with advice from D.H.

Fremont; V. Durai and P. Bagadia performed experiments related to cell sorting, cell culture and

82 flow cytometry with advice from C. Briseño and J.T. Davidson; V. Durai, C.G. Briseño, and M.

Gargaro performed in vivo manipulations of mice with advice from A. Iwata; V. Durai and D.J.

Thiesen performed cross-presentation assays; V. Durai, T.L. Murphy, and K.M. Murphy wrote the manuscript with contributions from all authors.

2.8 References

1. Mildner,A. & Jung,S. Development and function of dendritic cell subsets. Immunity 40, 642-656 (2014).

2. Liu,K. et al. Origin of dendritic cells in peripheral lymphoid organs of mice. Nat Immunol 8, 578-583 (2007).

3. Fogg,D.K. et al. A clonogenic bone marrow progenitor specific for macrophages and dendritic cells. Science 311, 83-87 (2006).

4. Auffray,C. et al. CX3CR1+ CD115+ CD135+ common macrophage/DC precursors and the role of CX3CR1 in their response to inflammation. J Exp. Med. 206, 595-606 (2009).

5. Onai,N. et al. Identification of clonogenic common Flt3(+) M-CSFR+ plasmacytoid and conventional dendritic cell progenitors in mouse bone marrow. Nat Immunol 8, 1207- 1216 (2007).

6. Naik,S.H. et al. Development of plasmacytoid and conventional dendritic cell subtypes from single precursor cells derived in vitro and in vivo. Nat Immunol 8, 1217-1226 (2007).

7. Grajales-Reyes,G.E. et al. Batf3 maintains autoactivation of Irf8 for commitment of a CD8alpha(+) conventional DC clonogenic progenitor. Nat Immunol 16, 708-717 (2015).

8. Schlitzer,A. et al. Identification of cDC1- and cDC2-committed DC progenitors reveals early lineage priming at the common DC progenitor stage in the bone marrow. Nat Immunol (2015).

9. Breton,G. et al. Circulating precursors of human CD1c+ and CD141+ dendritic cells. J Exp. Med 212, 401-413 (2015).

10. Lee,J. et al. Restricted dendritic cell and monocyte progenitors in human cord blood and bone marrow. J Exp. Med 212, 385-399 (2015).

11. See,P. et al. Mapping the human DC lineage through the integration of high-dimensional techniques. Science 356, (2017).

12. McKenna,H.J. et al. Mice lacking flt3 ligand have deficient hematopoiesis affecting hematopoietic progenitor cells, dendritic cells, and natural killer cells. Blood 95, 3489- 3497 (2000).

13. Waskow,C. et al. The receptor tyrosine kinase Flt3 is required for dendritic cell development in peripheral lymphoid tissues. Nat Immunol 9, 676-683 (2008).

14. Matthews,W. et al. A receptor tyrosine kinase cDNA isolated from a population of enriched primitive hematopoietic cells and exhibiting close genetic linkage to c-kit. Proc. Natl. Acad. Sci. U. S A 88, 9026-9030 (1991).

15. Miller,J.C. et al. Deciphering the transcriptional network of the dendritic cell lineage. Nat. Immunol. 13, 888-899 (2012).

16. Verstraete,K. & Savvides,S.N. Extracellular assembly and activation principles of oncogenic class III receptor tyrosine kinases. Nat Rev Cancer 12, 753-766 (2012).

17. Lyman,S.D. et al. Molecular cloning of a ligand for the flt3/flk-2 tyrosine kinase receptor: a proliferative factor for primitive hematopoietic cells. Cell 75, 1157-1167 (1993).

18. Maraskovsky,E. et al. Dramatic increase in the numbers of functionally mature dendritic cells in Flt3 ligand-treated mice: multiple dendritic cell subpopulations identified. J. Exp. Med. 184, 1953-1962 (1996).

19. Maraskovsky,E. et al. In vivo generation of human dendritic cell subsets by Flt3 ligand. Blood 96, 878-884 (2000).

20. Brasel,K., De Smedt,T., Smith,J.L. & Maliszewski,C.R. Generation of murine dendritic cells from flt3-ligand-supplemented bone marrow cultures. Blood 96, 3029-3039 (2000).

21. Naik,S.H. et al. Cutting edge: generation of splenic CD8+ and CD8- dendritic cell equivalents in Fms-like tyrosine kinase 3 ligand bone marrow cultures. J Immunol 174, 6592-6597 (2005).

22. D'Amico,A. & Wu,L. The early progenitors of mouse dendritic cells and plasmacytoid predendritic cells are within the bone marrow hemopoietic precursors expressing Flt3. J Exp. Med. 198, 293-303 (2003).

23. Mackarehtschian,K. et al. Targeted disruption of the flk2/flt3 gene leads to deficiencies in primitive hematopoietic progenitors. Immunity 3, 147-161 (1995).

24. Ginhoux,F. et al. The origin and development of nonlymphoid tissue CD103+ DCs. J Exp. Med. 206, 3115-3130 (2009).

25. Sitnicka,E. et al. Key role of flt3 ligand in regulation of the common lymphoid progenitor but not in maintenance of the hematopoietic stem cell pool. Immunity 17, 463-472 (2002).

26. Sitnicka,E. et al. Complementary signaling through flt3 and interleukin-7 receptor alpha is indispensable for fetal and adult B cell genesis. J Exp. Med 198, 1495-1506 (2003).

27. Nagasawa,T. Microenvironmental niches in the bone marrow required for B-cell development. Nat Rev Immunol 6, 107-116 (2006).

28. Helft,J., Ginhoux,F., Bogunovic,M. & Merad,M. Origin and functional heterogeneity of non-lymphoid tissue dendritic cells in mice. Immunol Rev 234, 55-75 (2010).

29. Satpathy,A.T., Murphy,K.M. & KC,W. Transcription factor networks in dendritic cell development. Semin. Immunol. 23, 388-397 (2011).

30. Moore,A.J. & Anderson,M.K. Dendritic cell development: a choose-your-own-adventure story. Adv. Hematol. 2013, 949513 (2013).

31. Tsapogas,P., Mooney,C.J., Brown,G. & Rolink,A. The Cytokine Flt3-Ligand in Normal and Malignant Hematopoiesis. Int J Mol Sci. 18, (2017).

32. Metcalf,D. Lineage commitment and maturation in hematopoietic cells: the case for extrinsic regulation. Blood 92, 345-347 (1998).

33. Enver,T., Heyworth,C.M. & Dexter,T.M. Do stem cells play dice? Blood 92, 348-351 (1998).

34. Mossadegh-Keller,N. et al. M-CSF instructs myeloid lineage fate in single haematopoietic stem cells. Nature 497, 239-243 (2013).

35. Rieger,M.A., Hoppe,P.S., Smejkal,B.M., Eitelhuber,A.C. & Schroeder,T. Hematopoietic cytokines can instruct lineage choice. Science 325, 217-218 (2009).

36. Onai,N., Obata-Onai,A., Tussiwand,R., Lanzavecchia,A. & Manz,M.G. Activation of the Flt3 signal transduction cascade rescues and enhances type I interferon-producing and dendritic cell development. J Exp. Med. 203, 227-238 (2006).

37. Tsapogas,P. et al. In vivo evidence for an instructive role of fms-like tyrosine kinase-3 (FLT3) ligand in hematopoietic development. Haematologica 99, 638-646 (2014).

38. Karsunky,H., Merad,M., Cozzio,A., Weissman,I.L. & Manz,M.G. Flt3 ligand regulates dendritic cell development from Flt3+ lymphoid and myeloid-committed progenitors to Flt3+ dendritic cells in vivo. J. Exp. Med. 198, 305-313 (2003).

39. Sathe,P., Vremec,D., Wu,L., Corcoran,L. & Shortman,K. Convergent differentiation: myeloid and lymphoid pathways to murine plasmacytoid dendritic cells. Blood 121, 11- 19 (2013).

40. Liu,K. et al. In vivo analysis of dendritic cell development and homeostasis. Science 324, 392-397 (2009).

41. Fancke,B., Suter,M., Hochrein,H. & O'Keeffe,M. M-CSF: a novel plasmacytoid and conventional dendritic cell poietin. Blood 111, 150-159 (2008).

42. Satpathy,A.T. et al. Zbtb46 expression distinguishes classical dendritic cells and their committed progenitors from other immune lineages. J. Exp. Med. 209, 1135-1152 (2012).

43. Meredith,M.M. et al. Expression of the zinc finger transcription factor zDC (Zbtb46, Btbd4) defines the classical dendritic cell lineage. J. Exp. Med. 209, 1153-1165 (2012).

44. Wu,X. et al. Mafb lineage tracing to distinguish macrophages from other immune lineages reveals dual identity of Langerhans cells. J Exp. Med 213, 2553-2565 (2016).

45. Li,J., Chen,K., Zhu,L. & Pollard,J.W. Conditional deletion of the colony stimulating factor-1 receptor (c-fms proto-oncogene) in mice. Genesis. 44, 328-335 (2006).

46. Ventura,A. et al. Restoration of p53 function leads to tumour regression in vivo. Nature 445, 661-665 (2007).

47. Onai,N. et al. A clonogenic progenitor with prominent plasmacytoid dendritic cell developmental potential. Immunity 38, 943-957 (2013).

48. Schlenner,S.M. et al. Fate mapping reveals separate origins of T cells and myeloid lineages in the thymus. Immunity 32, 426-436 (2010).

49. Kingston,D. et al. The concerted action of GM-CSF and Flt3-ligand on in vivo dendritic cell homeostasis. Blood 114, 835-843 (2009).

50. Merad,M., Sathe,P., Helft,J., Miller,J. & Mortha,A. The dendritic cell lineage: ontogeny and function of dendritic cells and their subsets in the steady state and the inflamed setting. Annu. Rev. Immunol. 31, 563-604 (2013).

51. Dosil,M., Wang,S. & Lemischka,I.R. Mitogenic signalling and substrate specificity of the Flk2/Flt3 receptor tyrosine kinase in fibroblasts and interleukin 3-dependent hematopoietic cells. Mol Cell Biol 13, 6572-6585 (1993).

52. Rottapel,R. et al. Substrate specificities and identification of a putative binding site for PI3K in the carboxy tail of the murine Flt3 receptor tyrosine kinase. Oncogene 9, 1755- 1765 (1994).

53. Lieschke,G.J. et al. Mice lacking granulocyte colony-stimulating factor have chronic neutropenia, granulocyte and macrophage progenitor cell deficiency, and impaired neutrophil mobilization. Blood 84, 1737-1746 (1994).

54. Wu,H., Liu,X., Jaenisch,R. & Lodish,H.F. Generation of committed erythroid BFU-E and CFU-E progenitors does not require erythropoietin or the erythropoietin receptor. Cell 83, 59-67 (1995).

55. Freeden-Jeffry,U. et al. Lymphopenia in interleukin (IL)-7 gene-deleted mice identifies IL-7 as a nonredundant cytokine. J Exp. Med 181, 1519-1526 (1995).

56. Peschon,J.J. et al. Early lymphocyte expansion is severely impaired in interleukin 7 receptor-deficient mice. J Exp. Med 180, 1955-1960 (1994).

57. Kretzer,N.M. et al. RAB43 facilitates cross-presentation of cell-associated antigens by CD8alpha+ dendritic cells. J Exp. Med 213, 2871-2883 (2016).

58. Shooshtari,P. et al. Correlation analysis of intracellular and secreted cytokines via the generalized integrated mean fluorescence intensity. Cytometry A 77, 873-880 (2010).

59. Darrah,P.A. et al. Multifunctional TH1 cells define a correlate of vaccine-mediated protection against Leishmania major. Nat. Med. 13, 843-850 (2007).

Figure 2.1. Flt3l–/– mice have a more severe DC deficiency than Flt3–/– mice at two weeks of age. (A and B) Splenocytes from mice of the indicated genotypes at two weeks of age were analyzed by flow cytometry for DC populations. Shown are representative two-color histograms of live cells. Numbers specify the percent of cells within the indicated gates. (C-E) Summary data for the percent (C) and number (D) of splenic cDCs and the percent (E) and number (F) of splenic pDCs in mice of the indicated genotypes. Dots represent biological replicates; small horizontal lines indicate the mean. Data are pooled from seven independent experiments (n = 5 mice per genotype). *P<0.05; **P<0.01; ****P<0.0001 using unpaired, two-tailed Student's t- test. 88

Figure 2.2. DC deficiency in skin-draining lymph nodes and lungs is more severe in Flt3l–/– mice than in Flt3–/– mice. (A) Gating strategy used for identifying resident cDCs (CD11c+MHCIIint) and pDCs (B220+CD317+) in the skin-draining lymph nodes by flow cytometry. Numbers specify the percent of cells within the indicated gates. (B-E) Summary data for the percent of cDCs (B and D) and pDCs (C and E) in SLNs of mice of the indicated genotypes at 2 weeks (B and C) or 8 weeks (D and E) of age. Dots represent biological replicates; small horizontal lines indicate the mean. Data are pooled from sixteen independent

89 experiments (n = 5-9 mice per genotype). (F) Gating strategy used for identifying cDCs (CD45.2+CD24+B220–CD11c+MHCII+) in the lung by flow cytometry. (G and H) Summary data for the percent of cDCs amongst CD45+ cells in lungs of mice of the indicated genotypes at 2 weeks (G) or 8 weeks (H) of age. Dots represent biological replicates; small horizontal lines indicate the mean. Data are pooled from sixteen independent experiments (n = 5-9 mice per genotype). *P<0.05; **P<0.01; ***P<0.001; ****P<0.0001 using unpaired, two-tailed Student's t-test.

Figure 2.3. Flt3l–/– mice continue to have a more severe DC deficiency than Flt3–/– mice at eight weeks of age. (A and B) Splenocytes from mice of the indicated genotypes at eight weeks of age were analyzed by flow cytometry for DC populations. Shown are representative two-color histograms of live cells. Numbers specify the percent of cells within the indicated gates. (C-E) Summary data for the percent (C) and number (D) of splenic cDCs and the percent (E) and number (F) of splenic pDCs in mice of the indicated genotypes. Dots represent biological replicates; small horizontal lines indicate the mean. Data are pooled from nine independent

91 experiments (n = 5-9 mice per genotype). *P<0.05; **P<0.01; ****P<0.0001 using unpaired, two-tailed Student's t-test.

Figure 2.4. Flt3L fails to generate DCs in Flt3–/– mice. (A-E) BM cells from WT (Flt3+/+) or Flt3–/– mice were treated with vehicle or Flt3L and cultured for 9 days. Live cells were subsequently analyzed by flow cytometry for DC populations. (A) Shown are representative two- color histograms of live cells pre-gated as indicated above the plots. Numbers specify the percentage of cells within the indicated gates. (B-E) Summary data for the percent (B) and number (C) of cDCs in each culture, and for the percent (D) and number (E) of pDCs in each culture. Dots represent biological replicates; small horizontal lines indicate the mean. Data are

93 pooled from three independent experiments (n = 4 mice per genotype). (F-J) WT (Flt3+/+) and Flt3–/– mice were administered vehicle or Flt3L subcutaneously for 8 days and splenocytes were subsequently analyzed by flow cytometry on day 9 for DC populations. (F) Shown are representative two-color histograms of live cells. Numbers specify the percent of cells within the indicated gates. (G-J) Summary data for the percent (G) and number (H) of splenic cDCs and the percent (I) and number (J) of splenic pDCs. Dots represent biological replicates; small horizontal lines indicate the mean. Data are pooled from two independent experiments (n = 5 mice per group). ns, not significant (P>0.05); *P<0.05; **P<0.01; ***P<0.001; ****P<0.0001 using unpaired, two-tailed Student's t-test.

Figure 2.5. Flt3–/– DCs are transcriptionally and functionally similar to WT DCs. (A and B) Microarray analysis of gene expression in cDC1s (A) or cDC2s (B) from Flt3+/+ and Flt3–/– mice, presented as M-plots. Colors indicate higher (red) or lower (blue) expression in Flt3+/+ cells than in Flt3–/– cells. Annotated genes with a greater than four-fold change in expression are specified (n = 2 biological replicates per subset per genotype). (C) Sorted cDC subsets from Flt3+/+ or Flt3–/– mice were cultured for 3 days with CFSE-labeled OT-I cells and different doses of heat-killed Listeria monocytogenes-OVA (HKLM-OVA) and then assayed for OT-I

95 proliferation (CFSE–CD44+). Summary data for the percent of proliferated OT-I cells in cultures with cDC subsets from the indicated genotypes are shown. Dots represent the mean; error bars represent the s.e.m. Data are pooled from two independent experiments (n = 4 mice per genotype). (D) Sorted cDC subsets from Flt3+/+ or Flt3–/– mice were cultured for 3 days with CFSE-labeled OT-I cells and different doses of soluble OVA as antigen and then assayed for OT-I proliferation (CFSE–CD44+). Summary data for the percent of proliferated OT-I cells in cultures with cDC subsets from the indicated genotypes are shown. Dots represent the mean; error bars represent the s.e.m. Data are pooled from two independent experiments (n = 4 mice per genotype). (E) Sorted cDC subsets from Flt3+/+ or Flt3–/– mice were incubated with soluble OVA, washed, and then different numbers of cDCs were cultured with CFSE-labeled OT-I cells for 3 days after which OT-I proliferation was assayed. Summary data for the percent of proliferated OT-I cells in cultures with the cDC subsets from the indicated genotypes are shown. Dots represent the mean; error bars represent the s.e.m. Data are pooled from two independent experiments (n = 4 mice per genotype). (F) Microarray analysis of gene expression in pDCs from Flt3+/+ and Flt3–/– mice, presented as M-plots. Colors indicate higher (red) or lower (blue) expression in Flt3+/+ cells than in Flt3–/– cells. Annotated genes with a greater than four-fold change in expression are specified (n = 2 biological replicates per subset per genotype). (G) Sorted pDCs from Flt3+/+ or Flt3–/– mice were stimulated with vehicle or CpG-A oligodeoxynucleotides overnight and then the cell supernatant was analyzed for IFN-α production. Summary data for the quantity of IFN-α produced by pDCs of the indicated genotypes are shown. Dots represent biological replicates; small horizontal lines indicate the mean. Data are pooled from two independent experiments (n = 4 mice per genotype).

Figure 2.6. Flt3–/– bone marrow generates mature DCs in response to M-CSF or SCF and contains committed DC progenitors. (A and B) BM cells from mice of the indicated genotypes were treated with vehicle, Flt3L, M-CSF, or SCF and cultured for 7 days. Live cells were subsequently analyzed by flow cytometry for development of DCs. (A) Shown are representative two-color histograms of live cells pre-gated as indicated above the plots and one-color histograms of Zbtb46GFP expression in cells of the indicated subtype. Numbers represent the percent of cells within the indicated gates. (B) Summary data for the percent of cDCs in each culture. Dots represent biological replicates; small horizontal lines indicate the mean. Data are

97 pooled from five independent experiments (n = 5-7 mice per genotype). (C) BM cells from mice of the indicated genotypes were analyzed by flow cytometry for DC progenitor populations; lineage markers include CD3, CD19, CD105, Ter119, and Ly-6G. Representative two-color histograms are shown of live cells pre-gated as indicated above the plots. Numbers specify the percent of cells within the indicated gates. Data are representative of four independent experiments (n = 7 mice per genotype). ns, not significant (P>0.05); **P<0.01; ***P<0.001; ****P<0.0001 using unpaired, two-tailed Student's t-test.

Figure 2.7. cDCs from M-CSF and SCF cultures are phenotypically similar to cDCs from Flt3L cultures. (A and B) BM cells from mice of the indicated genotypes were treated with vehicle, Flt3L, MCSF, or SCF and cultured for 7 days. Live cells were subsequently analyzed by flow cytometry for the development of cDCs and for their surface marker expression. (A) Summary data for the number of cDCs in each culture. Dots represent biological replicates; small horizontal lines indicate the mean. Data are pooled from five independent experiments (n = 5-7 mice per genotype). (B) cDCs (B220–F4/80–CD11c+MHCII+) in cultures were analyzed by flow cytometry for surface marker expression. Shown are representative one-color histograms of the indicated marker in cDCs from the indicated cytokine cultures. Data are representative of two independent experiments (n = 4 mice). ns, not significant (P>0.05); **P<0.01; ***P<0.001 using unpaired, two-tailed Student's t-test.

Figure 2.8. M-CSF and SCF support pDC development from Flt3–/– bone marrow. (A) BM cells from mice of the indicated genotypes were treated with vehicle, Flt3L, MCSF, or SCF and cultured for 7 days. Live cells were subsequently analyzed by flow cytometry to quantify pDC development. Shown are representative two-color histograms of live cells pre-gated as indicated above the plots. Numbers specify the percent of cells within the indicated gates. Data are representative of three independent experiments (n = 5 mice per genotype). (B) CDPs (Lin– cKitintCD135+CD115+CD11c–MHCII–) were sorted from Zbtb46GFP/+ mice and cultured with vehicle, Flt3L, MCSF, or SCF for 5 days. Live cells were subsequently analyzed by flow cytometry to quantify pDC development. Shown are representative two-color histograms of live cells pre-gated as indicated above the plots. Numbers specify the percent of cells within the indicated gates. Data are representative of three independent experiments (sorted cells from 3 to 5 mice pooled in each individual experiment).

100

Figure 2.9. Committed DC progenitors can mature in response to M-CSF and SCF. (A) CDPs (Lin–CD117intCD135+CD115+CD11c–MHCII–) were sorted from Zbtb46GFP/+ mice and cultured with vehicle, Flt3L, M-CSF, or SCF for 5 days. Cells were subsequently analyzed by flow cytometry for development of DCs. Shown are representative two-color histograms of live cells pre-gated as indicated above the plots and one-color histograms of Zbtb46GFP expression. Numbers specify the percent of cells within the indicated gates. Data are representative of three independent experiments (sorted cells from 3-5 mice were pooled in each individual experiment). (B) Pre-cDC2s (Lin–CD115+CD11c+MHCII–Zbtb46+) were sorted from Flt3+/+Zbtb46GFP/+ or Flt3–/–Zbtb46GFP/+ mice as indicated and cultured with vehicle, Flt3L, or M-CSF for 5 days. Cells were subsequently analyzed by flow cytometry for development of DCs. Shown are representative two-color histograms of live cells pre-gated as indicated above

101 the plots and one-color histograms of Zbtb46GFP expression in cells of the indicated subtype. Numbers specify the percent of cells within the indicated gates. Data are representative of three independent experiments (sorted cells from 3-5 mice per genotype were pooled in each individual experiment). (C) Pre-cDC2s were sorted from Flt3+/+Zbtb46GFP/+ or Flt3–/– Zbtb46GFP/+ mice, labeled with Cell Proliferation Dye (CPD), and cultured with M-CSF for 5 days. Cells were subsequently analyzed by flow cytometry for dilution of CPD to determine the number of divisions they underwent. Shown is a representative one-color histogram of CPD levels and the percent of cells at each division. Data are representative of two independent experiments (sorted cells from 3 mice per genotype were pooled in each individual experiment).

102

Figure 2.10. cDC development in Flt3–/– mice is dependent upon CSF1R. (A-F) Mice of the indicated genotypes were treated with tamoxifen to delete Csf1r. After treatment, splenocytes were analyzed by flow cytometry for DC populations. (A and B) Shown are representative two- color histograms of live cells. Numbers specify the percent of cells within the indicated gates. (C-E) Summary data of the percent (C) and number (D) of splenic cDCs and for the percent (E) and number (F) of splenic pDCs in mice of the indicated genotypes. Dots represent biological replicates; small horizontal lines indicate the mean. Data are pooled from seven independent experiments. (n = 3-6 mice per genotype). (G and H) WT CD45.1+ mice were lethally irradiated and reconstituted with Flt3–/– CD45.1+:Flt3–/–Csf1rf/fR26+/+CD45.2+ bone marrow at a 1:1 ratio

103

(CreERT2–) or Flt3–/–CD45.1+:Flt3–/–Csf1rf/fR26CreERT2/+CD45.2+ bone marrow at a 1:1 ratio (CreERT2+). After reconstitution and tamoxifen treatment to delete Csf1r, spleens and BM were analyzed by flow cytometry. (G) Shown are representative two-color histograms (pregating, above plots; splenic B cells, B220+CD24+; splenic cDCs, CD11c+MHCII+; BM CSF1R+ cells, CD115+). Numbers specify the percent of cells within the indicated gates. (H) Summary data of the chimerism ratio of splenic cDCs presented as the ratio of CD45.1+ to CD45.2+ cDCs normalized to the ratio of CD45.1+ to CD45.2+ B cells within the same mouse. Dots represent individual mice; small horizontal lines indicate the mean. Data are pooled from four independent experiments (n = 6-9 chimeras per genotype). ns, not significant (P>0.05); *P<0.05; **P<0.01; ***P<0.001 using unpaired, two-tailed Student's t-test.

104

Figure 2.11. Flt3–/– and Flt3l–/– BM progenitors and cDCs express normal levels of CSF1R and c-Kit. (A) BM cells from mice of the indicated genotypes were analyzed by flow cytometry for cytokine receptor expression. Representative two-color histograms of live cells are shown. Data are representative of at least three independent experiments (n = 4-7 mice per genotype). (B) Pre-cDC1s and pre-cDC2s from mice of the indicated genotypes were analyzed by flow cytometry for cytokine receptor expression levels. Representative one-color histograms of each receptor are shown. Data are representative of three independent experiments. (C) Expression of the indicated genes in splenic cDC1s and cDC2s from mice of the indicated genotypes was determined by gene expression microarray analysis. Dots represent biological replicates (n = 2 mice per subset per genotype).

105

Figure 2.12. Committed DC progenitors arise in Flt3l–/– bone marrow and do not display increased sensitivity to M-CSF relative to WT progenitors. (A) BM cells from mice of the indicated genotypes were analyzed by flow cytometry for DC progenitor populations; lineage markers include CD3, CD19, CD105, Ter119, and Ly-6G. Representative two-color histograms are shown of live cells pre-gated as indicated above the plots. Numbers specify the percent of cells within the indicated gates. Data are representative of three independent experiments (n = 3 mice per genotype). (B) BM from mice of the indicated genotypes was treated with M-CSF and assayed for phosphorylated Erk1/2 (pErk1/2) by intracellular flow cytometry. Shown are summary data presented as the Integrated MFI of pErk1/2 in pre-cDC2s from mice of the indicated genotypes stimulated with the indicated concentration of M-CSF. Dots indicate the mean from three independent experiments; error bars indicate the s.e.m (n = 3 mice per genotype). ns, not significant (P>0.05); *P<0.05 using Student's t-test.

106

Figure 2.13. Flt3–/– DC progenitors display increased sensitivity to M-CSF. (A and B) Serum-starved BM from mice of the indicated genotypes was treated with M-CSF and assayed for phosphorylated Erk1/2 (pErk1/2) by intracellular flow cytometry. (A) Shown are representative one-color histograms of pErk1/2 in pre-cDC2s stimulated with the indicated concentrations of M-CSF. Numbers in black indicate the percent of cells within the gate and numbers in red indicate the geometric mean fluorescence intensity (MFI) of pErk1/2 in the gated population. Data are representative of three independent experiments. (B) Summary data presented as the integrated MFI of pErk1/2 in pre-cDC2s from mice of the indicated genotypes stimulated with the indicated concentration of M-CSF. Dots indicate the mean from three independent experiments; error bars indicate the s.e.m (n = 3 mice per genotype). *P<0.05 using Student's t-test.

107

Figure 2.14. Deletion of Flt3 rescues the severe DC defect in Flt3l–/– mice. (A-E) Splenocytes from mice of the indicated genotypes at eight weeks of age were analyzed by flow cytometry for DC populations. (A) Shown are representative two-color histograms of live cells. Numbers specify the percent of cells within the indicated gates. (B-E) Summary data for the percent (B) and number (C) of splenic cDCs and the percent (D) and number (E) of splenic pDCs in mice of the indicated genotypes. Dots represent biological replicates; small horizontal lines indicate the mean. Data are pooled from six independent experiments (n = 4-5 mice per genotype). *P<0.05; **P<0.01; ***P<0.001; ****P<0.0001 using unpaired, two-tailed Student's t-test.

108

Figure 2.15. Deletion of Flt3 in Flt3l–/– mice reverses the severe DC defect in skin-draining lymph nodes and lungs. (A and B) Summary data for the percent of cDCs (A) and pDCs (B) determined as in Figure 2A in SLNs of mice of the indicated genotypes. Dots represent biological replicates; small horizontal lines indicate the mean. Data are pooled from at least three independent experiments (n = 2-5 mice per genotype). (C) Summary data for the percent of cDCs amongst CD45+ cells determined as in Figure 2F from mice of the indicated genotypes. Dots represent biological replicates pooled from groups of between 2 and 5 mice combined from at least 3 independent experiments; small horizontal lines indicate the mean. Data are pooled from six independent experiments (n = 4-5 mice per genotype). *P<0.05; ***P<0.001; ****P<0.0001 using unpaired, two-tailed Student's t-test.

109

CHAPTER 3:

Cryptic activation of an Irf8 enhancer governs cDC1 fate specification.

The contents of this chapter have been previously published in Nature Immunology.

Durai V, Bagadia P, Granja JM, Satpathy AT, Kulkarni DH, Davidson JT 4th, Wu R, Patel SJ,

Iwata A, Liu TT, Huang X, Briseño CG, Grajales-Reyes GE, Wöhner M, Tagoh H, Kee BL,

Newberry RD, Busslinger M, Chang HY, Murphy TL, Murphy KM. Cryptic activation of an Irf8 enhancer governs cDC1 fate specification. Nat Immunol. 2019 Sep;20(9):1161-1173.

110

3.1 Abstract

Induction of the transcription factor Irf8 in the common dendritic cell progenitor (CDP) is required for classical type 1 dendritic cell (cDC1) fate specification, but the mechanisms controlling this induction are unclear. Here we identified Irf8 enhancers via chromatin profiling of DCs and used CRISPR/Cas9 genome editing to assess their roles in Irf8 regulation. An enhancer 32 kilobases downstream of the Irf8 transcriptional start site (+32 kb Irf8) that was active in mature cDC1s was required for the development of this lineage, but not for its specification. Instead, a +41 kb Irf8 enhancer previously thought to be active only in plasmacytoid DCs was found to also be transiently accessible in cDC1 progenitors, and deleting this enhancer prevented the induction of Irf8 in CDPs and abolished cDC1 specification. Thus, cryptic activation of the +41 kb Irf8 enhancer in DC progenitors is responsible for cDC1 fate specification.

111

3.2 Introduction

The diversification of immune cells relies upon lineage-determining transcription factors that commit multipotent progenitors to a single fate 1,2. While early studies proposed that stochastic variations in the levels of these factors determined the eventual fate of progenitors 3, more recent work has instead suggested that the expression of individual factors is actively induced in order to specify a particular lineage 4. However, the precise mechanisms responsible for such induction remain unclear.

Gene expression is primarily controlled by cis-acting enhancers bound by transcription factors 5. Certain genes are regulated entirely by a single enhancer 6,7, while others, including many genes important for development, contain multiple potentially redundant enhancers as a safeguard for continued expression 8,9. Further, enhancer usage in individual genes dynamically changes as progenitors mature, possibly indicating the actions of distinct transcriptional networks throughout the developmental progression of a cell type 10. Analyzing the enhancers that regulate expression of lineage-determining transcription factors at developmental branch points could therefore identify the transcriptional mechanisms controlling fate choice.

Denritic cells (DCs) are a group of immune cells critical for innate and adaptive immune responses that include ‘classical’ DCs (cDCs) 11 and plasmacytoid DCs (pDCs) 12. cDCs are comprised of two functionally distinct lineages called cDC1 and cDC2 13. cDC1s are critical for priming CD8 T cells during antiviral and antitumor immune responses 14, as well as for effective responses to checkpoint blockade therapy 15,16. cDC1s are also the most promising substrates for cell-based cancer vaccines 17, so understanding their development is paramount.

DCs are derived from hematopoietic precursors in the bone marrow (BM), the earliest of which is the monocyte/DC progenitor (MDP) 18. The MDP gives rise to a common DC

112 progenitor (CDP) 19,20, which produces distinct clonogenic progenitors, the pre-cDC1 and pre- cDC2 21,22. Several transcription factors regulate development of the cDC1 lineage, including

Irf8, Batf3, Nfil3, and Id2 14,23-25. Although cDC1s can be generated in mice deficient in Nfil3,

Batf3, or Id2 under inflammatory conditions 26,27, Irf8–/– mice have an absolute defect in both pre-cDC1 specification and cDC1 development that cannot be rescued by such conditions.

Further, the Irf8 gene contains a super-enhancer in cDC1s, and Irf8 overexpression biases bone marrow progenitors towards cDC1 output21. These properties together suggest that Irf8 is the lineage-determining transcription factor for cDC1 fate. Understanding the enhancers that regulate Irf8 could therefore provide insight into how cDC1 fate specification from its multipotent progenitor, the CDP, is achieved.

Our previous work identified two distinct enhancers within the Irf8 super-enhancer located at +32 kb and +41 kb relative to the Irf8 transcriptional start site (TSS) 21. Using an integrating retroviral reporter, we demonstrated that the +32 kb Irf8 enhancer was selectively active in cDC1s, and that the +41 kb Irf8 enhancer was selectively active in pDCs. The +32 kb

Irf8 enhancer contained several AP1-IRF composite elements (AICEs) that bound IRF8 and

BATF3 in cDC1s by ChIP-seq, suggesting this enhancer might support Irf8 expression through autoactivation. The +41 kb Irf8 enhancer contained several E box motifs, suggesting that E proteins such as E2-2, the lineage-determining transcription factor of pDCs 28, might utilize this enhancer to drive Irf8 expression in pDCs. Finally, an Irf8 enhancer located at -50 kb was identified and analyzed using bacterial artificial chromosome (BAC) reporter transgenic mice, and was predicted to be required for Irf8 expression in MDPs 29. However, until now the functional requirement of these Irf8 enhancers for in vivo DC development has remained untested.

113

In this study, we used chromatin profiling of DCs and CRISPR/Cas9 genome editing to identify and delete enhancers regulating Irf8 in mice. We found that the +32 kb Irf8 enhancer was required for normal and compensatory cDC1 development but not for pre-cDC1 specification. We also found that the -50 kb Irf8 enhancer was not required for Irf8 expression in

MDPs, as was previously predicted, but rather regulated Irf8 levels selectively in monocyte/macrophage lineages. To find other enhancers regulating the cDC1 lineage, we performed ATAC-seq on DC progenitors. We surprisingly found that the +41 kb Irf8 enhancer became transiently accessible during the transition from the MDP to the pre-cDC1 before again closing in the mature cDC1. Deletion of the +41 kb Irf8 enhancer led to decreased Irf8 expression in pDCs, as was predicted, but also surprisingly prevented Irf8 induction in CDPs and resulted in the loss of both pre-cDC1 and mature cDC1 development. Consistently, Tcf3–/– DC progenitors, which lack E2A, had reduced cDC1 potential. Thus cryptic activation of the Irf8

+41 kb enhancer within CDPs is required for the induction of Irf8 and the subsequent specification of cDC1 fate.

3.3 Results

The +32 kb Irf8 enhancer is required for cDC1 development in vivo

The +32 kb Irf8 enhancer was identified by ChIP-seq as a 547 bp region that bound the transcription factors BATF3, IRF8, and p300 in cDC1s and that contained four AICEs in the 5' portion of the region 21 (Figure 3.1A). Reporter analysis indicated that the first three AICEs within this region were sufficient to confer cDC1 specific reporter activity, and that this activity could be abrogated by mutating these three AICEs (Figures 3.2A and 3.2B). The four AICES

114 contained in the 5' portion of the +32 kb Irf8 enhancer also accounted for the entire activity of the full length enhancer (Figure 3.2C). Further, using Flt3L-treated BM cultures of R26Cas9/+ transgenic mice, we found that expression of an sgRNA directed to the central AICE in the 5’ half of the +32 kb Irf8 enhancer caused a reduction in cDC1 development that was as large as the reduction caused by an sgRNA directed at the Irf8 coding sequence itself (Figure 3.2D). We therefore generated mice with deletions in this region by injecting zygotes with Cas9 mRNA and three sgRNAs, two flanking the first three AICEs in the 5’ half of the enhancer and a third downstream of the fourth AICE (Figures 3.1A and 3.2E). This generated two lines of mice with deletions within the +32 kb Irf8 enhancer, one deleting 149 bp and eliminating 3 AICEs (Irf8

+32 5’–/–- mice), and a second deleting 421 bp and eliminating all 4 AICEs (Irf8 +32–/– mice)

(Figure 3.2F). Both lines show severe reductions in cDC1 development in vivo (Figures 3.2G and 3.2H). Deletion of 3 AICEs reduced cDC1 development by 10-fold compared with wildtype mice, and deletion of all 4 AICEs eliminated all residual cDC1s (Figure 3.2H). All subsequent analyses used the more complete Irf8 +32–/– strain.

The defect in splenic cDC1 development in Irf8 +32–/– mice is more complete than in

Batf3–/– mice and as severe as in Irf8–/– mice (Figures 3.1B and 3.1D). Irf8 +32–/– mice showed no defects in other lineages, confirming the selective activity of this enhancer in cDC1s as predicted by our reporter assays. pDCs were normal in frequency and intracellular IRF8 protein levels (Figures 3.1C and 3.1D), as were monocytes, neutrophils, red pulp macrophages, B cells, and T cells (Figures 3.3A-3.3C). cDC1s in Irf8 +32–/– mice were absent in all tissues, such as in the lung (Figures 3.5A and 3.5B). To test for a functional defect in Irf8 +32–/– mice, we analyzed a tumor rejection system that requires cDC1s (Figure 3.1E). The regressor fibrosarcoma 1969, whose rejection relies on cross-presentation by cDC1s 31, was rejected by all

115 wildtype mice, but not by Irf8 +32–/– mice (Figure 3.1E). These data indicated that the +32 kb

Irf8 enhancer was absolutely required for cDC1 development and that its deletion generates a specific and functional defect restricted to this lineage.

Compensatory cDC1 development requires the +32 kb Irf8 enhancer

cDC1 development can occur in Batf3–/– mice due to compensation from BATF and

BATF2, which can replace BATF3 for interactions with IRF8 30. Compensatory cDC1 development in Batf3–/– mice occurs in skin draining lymph nodes (SLNs) and under inflammatory settings such as after bone marrow transplantation or after administration of IL-12

26,27. We examined these settings to test if compensatory cDC1development can occur in Irf8

+32–/– mice (Figure 3.4). We found that cDC1s were clearly present in SLNs from wildtype and

Batf3–/– mice, but were completely absent in Irf8 +32–/– mice (Figures 3.4A and 3.4B). We further found that administration of IL-12 led to a complete restoration of splenic cDC1 in Batf3–

/– mice, but not in Irf8 +32–/– mice (Figures 3.4C and 3.4D). Finally, we tested for cDC1 compensation after BM transplantation 27. We generated BM chimeras using wildtype, Irf8 +32–

/–, or Batf3–/– donor BM transplanted into irradiated wildtype mice (Figures 3.5C and 3.5D). cDC1s developed in chimeras produced from wildtype BM, as expected, and from Batf3–/– BM, as reported27. However, cDC1s failed to develop in chimeras produced from Irf8 +32–/– BM. In summary, all compensatory cDC1 development failed to occur in mice lacking the +32 kb Irf8 enhancer.

While IL-12 injection did not restore cDC1s in Irf8 +32–/– mice, it did increase the numbers of cDC2s (Figures 3.4C and 3.4D). This potentially resulted from specified pre-cDC1s

116 failing to maintain high Irf8 levels in response to IL-12 and their subsequent diversion to the cDC2 fate, as occurs with pre-cDC1s from Batf3–/– mice 21. We therefore sought to determine whether pre-cDC1 specification occurs in Irf8 +32–/– mice. Previous identification of the pre- cDC1 in BM relied upon Zbtb46-GFP expression. However, we developed a method to identify the pre-cDC1 using CD226 expression in place of Zbtb46-GFP (Figure 3.6). CD226 was not expressed in CDPs, but became expressed in pre-cDC1s (Figures 3.6A and 3.6B). Like Zbtb46-

GFP expression, CD226 expression indicates pre-cDC1 specification, as Flt3L culture of Lin–

CD135+CD117intCD226+ BM cells, like Lin–CD135+CD117int Zbtb46-GFP+ BM cells, generated exclusively cDC1s (Figures 3.6C and 3.6D). Also, Lin–CD135+CD117intCD226+ cells develop in wildtype mice, but not in Irf8–/– mice known to lack pre-cDC1s (Figure 3.4E)21. Using CD226 expression, we found that pre-cDC1s develop in Irf8 +32–/– mice in normal numbers but had reduced intracellular IRF8 protein levels (Figures 3.4E, 3.4F, and 3.5E), similar to the findings in

Batf3–/– mice. And in contrast to wildtyppe pre-cDC1s, pre-cDC1s from Irf8 +32–/– mice cultured in Flt3L become cDC2s (Figure 3.4G). These cDC2s derived from Irf8 +32–/– pre- cDC1s were transcriptionally similar to cDC2s arising from wildtype pre-cDC2s and transcriptionally distinct from cDC1s arising from wildtype pre-cDC1s, including in the expression of key cDC2 transcription factors such as Irf4 (Figure 3.6E-G). Though the cDC2s derived from Irf8 +32–/– pre-cDC1s retained CD24 expression, we found that there were no transcriptional differences between CD24+ cDC2s and CD24– cDC2s (Figure 3.6F), indicating that the diverted cDC2s that developed from Irf8 +32–/– pre-cDC1s were bona fide cDC2s. This result also recapitulated the diversion of Batf3–/– pre-cDC1s to the cDC2 fate (Fig. 3.6H). In summary, Irf8 +32–/– mice showed continued development of a specified pre-cDC1 progenitor in

BM that fails to sustain Irf8 expression and diverts to the cDC2 lineage.

117

The -50 kb Irf8 enhancer regulates Irf8 expression in monocytes and macrophages

To identify enhancers regulating pre-cDC1 specification, we performed ChIP-seq of active histone marks in MDPs, CDPs, and pre-cDC1s. This analysis identified a region of active

H3K27 acetylation at -50 kb relative to the Irf8 TSS that overlapped with a previously identified

Irf8 enhancer 29 (Figure 3.7A). Analysis of BAC transgenic Irf8 reporter mice had suggested that this -50 kb region contains two PU.1 binding sites and is required for Irf8 expression in the

MDP29. To test whether this enhancer regulated Irf8 induction and pre-cDC1 specification, we used CRISPR/Cas9 genome editing to generate mice with a 364 bp deletion that eliminates both of the PU.1 sites in this region (Irf8 -50–/– mice) (Figures 3.8A and 3.8B). Irf8 -50–/– mice have normal frequencies of cDCs, pDCs, neutrophils, B cells, T cells, and red pulp macrophages

(Figures 3.7B and 3.8C), all of which express normal levels of intracellular IRF8 protein

(Figures 3.7C and 3.8D). Frequencies and intracellular IRF8 protein levels of MDPs, CDPs, and pre-cDC1s were also normal in Irf8 -50–/– mice (Figures 3.7D and 3.7E), in contrast to the predictions based on the BAC transgenic Irf8 reporter mice29.

However, ATAC-seq analysis suggested that the -50 kb Irf8 enhancer was active in

F4/80+ peritoneal macrophages and Ly6C+ monocytes (Figure 3.9A). In Irf8 -50–/– mice, monocytes were present at normal numbers, but showed reduced intracellular IRF8 protein levels compared to wildtype mice (Figures 3.9B-3.9D). Likewise, F4/80+ peritoneal macrophages in

Irf8 -50–/– mice had greatly reduced intracellular IRF8 protein levels (Figures 3.9E and 3.9F).

Irf8–/– mice have been reported to have decreased survival in Salmonella enterica Typhimurium infection, possibly due to IRF8 regulation of inflammasome activity in macrophages 32.

118

However, it was not determined whether the in vivo defect in germline Irf8–/– mice was cell- intrinsic to macrophages. We found that Irf8 -50–/– mice were more susceptible to Salmonella infection compared to wildtype mice (Figure 3.9G), consistent with a role for the -50 kb Irf8 enhancer in controlling IRF8 protein levels in monocytes/macrophages during infections. In conclusion, the -50 kb Irf8 enhancer was not required for DC development but did regulate Irf8 specifically in monocytes and macrophages.

ATAC-seq reveals transient use of the +41 kb Irf8 enhancer during cDC1 development

To obtain base pair level resolution of accessible chromatin in progenitors, we next performed ATAC-seq on MDP, CDP, and pre-cDC1 populations 33,34 (Fig. 3.10). Pearson correlation analysis of all distal ATAC-seq peaks indicated that MDPs were most closely related to CDPs and least related to pre-cDC1s (Figure 3.10A). During transition from MDPs to CDPs,

687 unique peaks were gained, and 1,522 lost (Figure 3.10B). By contrast, during transition from CDPs to pre-cDC1s, 14,630 peaks were gained, and 13,664 lost. Thus a large shift in chromatin accessibility occurs as CDPs transition to pre-cDC1s, which could potentially be related to the loss of alternative fate potentials in the pre-cDC1, to the differential proliferative capacities of these two cell types, or to a natural consequence of the differentiation process.

Five main k-means clusters of peaks were identified (Figure 3.10C). Cluster 1 included peaks found only in MDPs. Clusters 2 and 3 included peaks shared between MDPs and CDPs.

Cluster 4 contained peaks found in both CDPs and pre-cDC1s, and cluster 5 included peaks present only in pre-cDC1s. We identified the genes closest in linear distance to the peaks within each cluster, and inferred the transcription factor motifs enriched within these peaks (Figures

119

3.10C and 3.10D). Of note, the enrichment of transcription factors reflects the activity of any factor with a similar DNA binding motif rather than the activity of the specific factor that was enriched. For example, enrichment of Tcf3 could reflect the activity of other Tcf factors such as

Tcf4 or Tcf12 within the clustered peaks. Peaks unique to MDPs (cluster 1) were in proximity to genes related to monocyte ontogeny, such as Mpo and Maf. These peaks were enriched in motifs for transcription factors such as Cebpb and Atf that regulate the macrophage/monocyte lineages

35,36 (Figures 3.10C and 3.10D).

The peaks identified in MDPs and CDPs (cluster 2 and 3) were enriched for motifs binding E proteins and Runx factors, both of which are required for DC development 28,37,38

(Figures 3.10C and 3.10D). Runx1 was also among the genes most proximal to peaks within these clusters, and the Runx1 locus contained open chromatin peaks in the MDP and CDP that were lost in the pre-cDC1 (Figure 3.10E).

The peaks found in CDPs and pre-cDC1s (cluster 4) were also enriched in motifs binding

E proteins (Figure 3.10C). Further, cluster 4 included the +41 kb Irf8 enhancer (Fig. 3.10C), which contains six predicted E box motifs and was previously found to be active in pDCs but not cDC1s21 (Figure 3.1A). This unexpected ATAC-seq accessibility of the +41 kb Irf8 enhancer in the pre-cDC1 (Figure 3.10E) suggested that the activity of this enhancer could be important for cDC1 specification.

Finally, in the cluster of peaks found only in the pre-cDC1 (cluster 5) we found that Irf8 and Batf3 were among the genes at closest proximity to open regions and that AP1-IRF motifs and PU.1 motifs were highly enriched within these regions (Figures 3.10C and 3.10D).

Accessible chromatin peaks were also proximally located to genes known to be critically

120 involved in cDC1 function, such as Wdfy4 39 (Figure 3.10E). Finally, we found that the +32 kb

Irf8 enhancer only became accessible at the pre-cDC1 stage (Figures 3.10C and 3.10E).

Together, our data suggests that the +41 kb Irf8 enhancer was transiently active in cDC1 progenitors before being replaced by activity at the +32 kb Irf8 enhancer in mature cDC1s.

The +41 kb Irf8 enhancer is required for cDC1 specification

By ATAC-seq analysis, the +41 kb Irf8 enhancer was inactive in MDPs, became active in

CDPs and pre-cDC1s, and was inactive again in mature cDC1s (Figure 3.11A). This enhancer also bound p300 in pDCs, suggesting it regulated Irf8 expression in these cells21. We targeted this enhancer using CRISPR/Cas9 genome editing with two flanking sgRNAs (Figure 3.11A), producing mice with a 361 bp deletion (Irf8 +41–/–) that eliminated all six predicted E boxes

(Figures 3.12A and 3.12B). As expected, Irf8 +41–/– mice had a pDC phenotype similar to the phenotype reported for Irf8–/– mice40. pDCs were present in Irf8 +41–/– mice, but had low protein levels of IRF8 and CD317 and increased protein levels of IRF4 (Figures 3.11B-3.11D). By contrast, monocytes, neutrophils, red pulp macrophages, B cells, and T cells from these mice were normal in frequency and intracellular IRF8 protein levels (Figures 3.12C and 3.12D).

However, Irf8 +41–/– mice completely lacked mature cDC1s (Figures 3.11B and 3.11D).

We found normal frequencies of MDPs and CDPs in the bone marrow of Irf8 +41–/– mice, indicating that the +41 kb Irf8 enhancer was not required for the development of these progenitors (Figure 3.11E). However, while the transition from the MDP to CDP is usually accompanied by a ~2-fold increase in intracellular IRF8 protein levels in wildtype CDPs, there was no increase in IRF8 levels in Irf8 +41–/– CDPs (Figure 3.11F). Also, pre-cDC1s failed to

121 develop in Irf8 +41–/– mice, similarly to Irf8–/– mice (Figure 3.11G). In summary, the +41 kb

Irf8 enhancer was only transiently active during cDC1 progenitor development, but was absolutely required for Irf8 induction in the CDP and for cDC1 specification.

E proteins are involved in cDC1 and pDC development

Since deletion of the +41 kb Irf8 enhancer abolished cDC1 specification, the factors that bound to this enhancer could be responsible for cDC1 fate commitment. The 454 bp region defining this enhancer contains predicted six E box motifs 21 (Figure 3.11A). ChIP-seq of LPS- activated B cells, pre-plasmablasts, and plasmablasts, in which E proteins and IRF8 are active, showed E2A binding to the +41 kb Irf8 enhancer. ATAC-Seq of these populations also found that this site was accessible in wildtype B cells, but was not in B cells doubly deficient for the E proteins E2A (Tcf3) and E2-2 (Tcf4) (Figure 3.13A).

The E proteins E2A and E2-2 are known to regulate B cell development and pDC development respectively 28,41, but they have not been implicated in cDC development or function.

Using the Tcf3fl/+VavCretg mice, which express an E2A-GFP fusion protein after Cre activity 41, we found that E2A was expressed in MDPs and CDPs, was reduced in pre-cDC1s, and was completely absent in mature cDC1s and pDCs (Figure 3.13B). This indicated that there was a transient period of E2A expression in DC progenitors. E2A was also expressed in pre-B cells, as has been previously described41, and also in CMPs and CLPs (Fig. 3.14A). To test the relevance of this expression, we analyzed DC development from wildtype and Tcf3–/– BM using Flt3L- treated cultures. While wildtype BM showed normal cDC1 and pDC development in vitro, Tcf3–

/– BM showed severely impaired cDC1 and pDC development but normal cDC2 development

122

(Figures 3.13C and 3.13D). However, cDC1s and pDCs continued to develop in Tcf3–/– mice in vivo, suggesting possible compensation for the actions of Tcf3 by another member of the Tcf family as has been previously described in B cells 42 (Figures 3.14B and 3.14C). To further characterize the role of Tcf3 in vivo, we generated bone marrow chimeras with mixed wildtype and Tcf3–/– bone marrow and assessed DC development after reconstitution (Figures 3.14D and 3.14E). We found that wildtype progenitors preferentially gave rise to cDC1s and exclusively gave rise to pDCs when in competition with Tcf3–/– progenitors. These results suggest a potential role for E proteins in cDC1 and pDC development.

3.4 Discussion

Our study analyzed the in vivo function of three different Irf8 enhancers, discovering unexpected and distinct roles in DC development and monocyte/macrophage function for each.

Our initial aim was to understand the development of the cDC1 subset critical for antiviral and antitumor immunity. Irf8 is the lineage-determining transcription factor responsible for cDC1 development23, but the molecular regulation of its transcription has not been well characterized.

Previous studies have proposed that PU.1 regulated the induction of Irf8 in the MDP through a -

50 kb Irf8 enhancer 29, that BATF3 supports Irf8 autoactivation at the +32 kb Irf8 enhancer 21, and that there was a pDC-specific +41 kb Irf8 enhancer as well 21. However, none had tested the roles of any Irf8 enhancer elements in vivo.

In this study, we generated several targeted deletions of Irf8 enhancers using

CRISPR/Cas9 genome editing in mice. Our results have confirmed the +32 kb Irf8 enhancer's role in cDC1 development 21 and excluded a role for the -50 kb Irf8 enhancer in early Irf8

123 expression 29. However, most importantly, our analysis has established a wholly unexpected requirement for the +41 kb Irf8 enhancer in Irf8 induction within the CDP and subsequent cDC1 specification. This further suggested a previously unrecognized role for E proteins in this process as well.

We have confirmed that the +32 kb Irf8 enhancer is required for normal and compensatory cDC1 development. We had originally predicted this result because BATF3, which is required for cDC1 development, bound along with IRF8 to the +32 kb Irf8 enhancer.

However, it was possible that BATF3 could also interact with other unidentified regions that serve as redundant enhancers to support cDC1 development. Our study found that not only was the +32 kb Irf8 enhancer required for normal cDC1 development, it was also required for the compensatory cDC1 development that can occur in Batf3–/– mice under various inflammatory settings. Because of this compensatory cDC1 development in Batf3–/– mice, this strain has not been useful for examining the function of cDC1s in some settings. For example, during infection by Mycobacterium tuberculosis, cDC1s reappear in Batf3–/– mice, so that the role of these cells in protection cannot be evaluated 26. Also, Batf3 has functions in other lineages, such as T cells and perhaps macrophages 43, so Batf3–/– mice might manifest phenotypes in other cell types that could make interpretations from this strain difficult. These limitations have been resolved by the generation of the Irf8 +32–/– mice. In these mice, Batf3 remains expressed normally in all immune lineages, but cDC1 development, both natural and compensatory, is ablated. These mice should now allow for the analysis of cDC1 function in settings where it could not previously be studied.

Our results also demonstrate how the cryptic activation of enhancers by transiently active transcriptional networks can control lineage diversification. Early models of hematopoiesis

124 proposed that the fate choices of progenitors are resolved by stochastic fluctuations in the expression of cross-antagonistic lineage-determining transcription factors. These fluctuations cause one factor to become dominant and impose fate commitment, which is reinforced by autoactivation of this factor and cross-inhibition of opposing factors. One example is the

PU.1/GATA1 circuit that regulates the choice between myeloid and megakaryocyte/erythrocyte fates. PU.1 and GATA1 are able to inhibit one another 44,45 and also to activate their own transcription 46,47. Further, these factors were thought to be co-expressed in progenitor cells that remained uncommitted until fluctuations in the levels of these two factors specified them to a single fate3. But this model was challenged by a recent study that used high-resolution analysis of single progenitor cells 4. Using reporters for PU.1 and GATA1 expression, that study demonstrated that no progenitor cells actually co-express these factors. Rather, PU.1 levels decay before GATA1 is even expressed in progenitors destined for megakaryocyte/erythrocyte fate. This suggested that some unknown mechanism actually drives this particular cell fate by inducing GATA1, rather than stochastic fluctuations randomly generating this outcome. Our data also supports such a deterministic rather than stochastic model of fate divergence, in which transiently active transcriptional networks initiate the higher expression of lineage-determining transcription factors. Specifically, we found that activation of the +41 kb Irf8 enhancer in CDPs, potentially by E proteins, induced Irf8 expression and subsequent cDC1 fate specification.

Similarly, other transiently acting networks may act to induce factors such as PU.1 or GATA1 in progenitors of other lineages. These networks may not be apparent in mature progeny, but might be revealed by the analysis of chromatin states of progenitors during development.

Our study also highlights how different enhancers of the same gene can be required at different stages of development of a single lineage. cDC1 development requires both the +41 kb

125 and the +32 kb Irf8 enhancers, but the requirements are manifested at different developmental stages. The +41 kb Irf8 enhancer is required for the specification of pre-cDC1s from CDPs, but its activity is subsequently extinguished and is not apparent in the fully developed cDC1. On the other hand, the +32 kb Irf8 enhancer is active only after cDC1 specification, and remains active in the fully developed cDC1. The switch between these two enhancers reveals that different transcriptional networks are responsible for Irf8 expression during distinct periods of cDC1 development. Such switches in enhancer activity could occur throughout hematopoiesis, and analyzing them could help elucidate the mechanisms controlling other fate divergences that are still not fully understood.

Finally, our finding that E proteins were potentially responsible for cDC1 specification was wholly unexpected. However, since pDC development also relies on E proteins, it is unclear how cDC1 and pDC fates are separately specified. Other factors important in cDC1 development, such as Id2, could act to exclude pDC potential by blocking E protein activity, but why this would not also block cDC1 specification is unclear. Future work will need to determine the precise sequence of transcriptional events that govern cDC1 development, including our newfound role for E proteins.

3.5 Materials and Methods

Mice.

Batf3–/–, Irf8–/–, and Zbtb46GFP/+ mice have been described previously. Irf8 +32–/–, Irf8 -

50–/–, and Irf8 +41–/– mice were newly generated as described below. The following mice were purchased from Jackson Laboratories: R26Cas9/+ mice (B6N.129(Cg)-Gt(ROSA)26Sortm1.1(CAG-

126 cas9*,-EGFP)Fezh/J), CD45.1+ mice (B6.SJL-Ptprca Pepcb/BoyJ), Tcf3gfp/+ mice (B6.129-

Tcf3tm1Mbu/J), and VavCretg mice (B6.Cg-Commd10Tg(Vav1-iCre)A2Kio/J). Tcf3–/– mice were a gift from B. Kee. All mice were on the C57BL6/J background except for Tcf3–/– mice, which were on the FVB/NJ background. All mice were generated, bred, and maintained in the Washington

University in St. Louis School of Medicine specific pathogen-free animal facility. Animals were housed in individually ventilated cages covered with autoclaved bedding and provided with nesting material for environmental enrichment. Up to five mice were housed per cage. Cages were changed once a week, and irradiated food and water in autoclaved bottles were provided ad libitum. Animal manipulation was performed using standard protective procedures, including filtered air exchange systems, chlorine-based disinfection, and personnel protective equipment including gloves, gowns, shoe covers, face masks, and head caps. All animal studies followed institutional guidelines with protocols approved by the Animal Studies Committee at

Washington University in St. Louis.

Unless otherwise specified, experiments were performed with mice between 6 and 10 weeks of age. No differences were observed between male and female mice in any assays performed and so mice of both genders were used interchangeably throughout this study. Within individual experiments, mice used were age- and sex-matched whenever possible. When mice were tracked for tumor growth or survival after Salmonella enterica Typhimurium infection, the monitoring scientist was blinded as to the genotypes of the mice in the experiment.

Generation of Irf8 enhancer deletion mice

sgRNAs that flanked the enhancers of interest were identified using GT-Scan (http://gt- scan.braembl.org.au/gt-scan/). For deletion of the Irf8 +32 kb enhancer the following sgRNA

127 sequences were used: Irf8 +32 5’: gttgtgatctttgaggtaga, Irf8 +32 Mid: gtctccttctgaaatttcagtt, and

Irf8 +32 3’: gaactggcctggggcaggtc. For the Irf8 -50 kb enhancer the following sgRNA sequences were used: Irf8 -50 5’: ggtgacatctgtctacggag and Irf8 -50 3’: atgcacccaaggcctggctc. For the Irf8

+41 kb enhancer the following sgRNA sequences were used: Irf8 +41 5’: ggcccttgtagtttagctta and Irf8 +41 3’: aaagaagatctggggtatgt. Oligonucleotides that included these desired sgRNA sequences preceded by a T7 polymerase initiation site (ttaatacgactcactataggg) and followed by a portion of the tracrRNA sequence that annealed to the pX330 vector (gttttagagctagaaatagcaag) were then purchased (Sigma Aldrich). For example, the full Irf8 +32 5’ oligonucleotide purchased was

TTAATACGACTCACTATAGGGgttgtgatctttgaggtagaGTTTTAGAGCTAGAAATAGCAAG.

Each oligonucleotide was used in a PCR reaction with the PX330 Common Reverse Primer, aaaagcaccgactcggtgcc, and with the PX330 vector as a template to generate a complete DNA product containing the T7 polymerase initiation site, the sgRNA sequence, and the full tracrRNA sequence in order. RNA was then synthesized from these products using the MEGAscript T7

Transcription Kit (Thermo Fisher Scientific). RNA was purified using the MEGAclear

Transcription Clean-Up Kit (Thermo Fisher Scientific) and eluted into nuclease-free injection buffer. RNA was diluted and stored at -80°C until it was used for microinjection. Purified Cas9 mRNA was a gift from W. Yokoyama.

Day 0.5 single cell embryos from C57Bl/6 mice were isolated and underwent pronuclear micro-injection at the Department of Pathology Micro-Injection Core. Each embryo was injected with 50 ng of each sgRNA and 100 ng of Cas9 mRNA. Injected embryos were then transferred into the oviducts of pseudopregnant recipient mice.

128

The resulting pups were screened by PCR to identify those that had successful deletion of the enhancers of interest. Mice with the desired deletion were then outcrossed to wildtype

C57Bl/6 mice, and the resulting heterozygous mice were intercrossed to generate homozygous enhancer deletion mice.

Dendritic cell preparation.

Lymphoid and nonlymphoid organ DCs were harvested and prepared as described previously48. Briefly, spleens and inguinal skin-draining LNs were minced and digested in 5 mL of Iscove’s modified Dulbecco’s media (IMDM) + 10% FCS (cIMDM) with 250 µg/mL collagenase B (Roche) and 30 U/mL DNaseI (Sigma-Aldrich) for 45 min at 37⁰C with stirring.

Lungs were minced and digested in 5 mL of cIMDM with 4 mg/mL collagenase D (Roche) and

30 U/mL DNaseI (Sigma-Aldrich) for 1.5 hours at 37⁰C with stirring. After digestion was complete, single cell suspensions from all organs were passed through 70-µm strainers and red blood cells were lysed with ammonium chloride-potassium bicarbonate (ACK) lysis buffer. Cells were subsequently counted with a Vi-CELL analyzer (Beckman Coulter) and 3-5x106 cells were used per antibody staining reaction.

For peritoneal cell analysis, 5 mL of MACS buffer (DPBS + 0.5% BSA + 2mM EDTA) was injected into the peritoneum of mice using a 27 g needle. After injection the mice were shaken gently to dislodge peritoneal cells. A 25 g needle was then used to collect the peritoneal fluid. Cells were ACK lysed and counted as described above.

Antibodies and flow cytometry

129

Cells were stained at 4ºC in MACS buffer in the presence of Fc block (2.4G2; BD

Biosciences).

The following antibodies were from BD Biosciences: CD11b PE (M1/70), CD19 Biotin

(1D3), CD45R AlexaFluor488 (RA3-6B2), CD45.2 PerCP-Cy5.5 (104), CD64 AlexaFluor647

(X54-5/7.1), CD103 BV421 (M290), CD117 BUV395 (2B8), CD127 BV421 (SB/199), CD135

PE-CF594 (A2F10.1), I-A/I-E V500 (M5/114.15.2), Ly6C AlexaFluor 700 (AL-21). The following antibodies were from eBioscience: CD11b eFluor 450 (M1/70), CD11c PE (N418),

CD24 PE-Cy7 (M1/69), CD172a PerCP–eFluor 710 (P84), CD317 APC (eBio927), F4/80 APC- eFluor 780 (BM8), IRF4 PE (3E4), IRF8 PerCP-eFluor 710 (V3GYWCH), Siglec H PerCP-

Cy5.5 (eBio-440c). The following antibodies were from BioLegend: CD45R Biotin (RA3-6B2),

CD45.2 PE (104), CD105 Biotin (MJ/718), CD115 BV711 (AFS98), CD127 biotin (A7R34),

CD226 PE (10E5), F4/80 APC-Cy7 (BM8), F4/80 AlexaFluor700 (BM8), Ly6G Biotin (1A8),

Ter119 Biotin (TER-119), XCR1 BV421 (ZET). The following antibodies were from Tonbo

Bioscience: CD45.1 FITC (A20), CD3e Biotin (145-2c11). The following antibodies were from

Invitrogen: CD11c APC-eFluor780 (N418). Cells were analyzed on a FACSCanto II or

FACSAria Fusion flow cytometer (BD), and data were analyzed with FlowJo v10 software

(TreeStar).

For intracellular IRF8/IRF4 staining, cells were stained for surface markers and then fixed/permeabilized with the intracellular fixation and permeabilization buffer kit (eBioscience) for 1 hour-overnight at 4⁰C. Cells were then stained for 1 hr at room temperature with intracellular antibodies. The cells were then washed and analyzed by flow cytometry.

Bone marrow isolation.

130

Bone marrow (BM) was harvested from the femur, tibia, and pelvis of mice. Bones were collected and fragmented by mortar and pestle in MACS buffer, and debris was removed by passing cells through a 70-µm strainer. Red blood cells were lysed with ACK lysis buffer and cells were subsequently counted on a Vi-CELL analyzer (Beckman Coulter). 3-5x106 were used per antibody staining reaction. For BM culture experiments, bulk BM cells were cultured at 37⁰C in 4 mL total volume of cIMDM supplemented with 100 ng/mL Flt3L (Peprotech) for nine days before further analysis.

Progenitor sorting and culture.

For sorting experiments, BM was isolated as described above and depleted of CD3-,

CD19-, CD105-, Ter119-, and Ly6G-expressing cells by staining with the corresponding biotinylated antibodies followed by depletion with MagniSort Streptavidin Negative Selection

Beads (Thermo Fisher). All remaining BM cells were then stained with fluorescent antibodies prior to sorting. Gates used to define MDPs, CDPs and pre-cDC1s were a combination of previously established markers21 and those identified in this study. MDPs were identified as Lin–

CD117hiCD135+CD115+CD11c–MHCII–. CDPs were identified as Lin–

CD117intCD135+CD115+CD11c–MHCII–. Pre-cDC1s were identified as either Lin–

CD117intCD135+Zbtb46-GFP+ or as Lin–CD117intCD135+CD226+. Pre-cDC2s were identified as

Lin—CD117loCD135+CD115+. Lineage markers included CD3, CD19, CD105, CD127, NK1.1,

Ter119, and Ly6G. For retroviral reporter assays and in vitro CRISPR/Cas9 deletion CD117hi cells were sorted. A FACSAria Fusion was used for sorting and cells were sorted into cIMDM.

Sort purity of >95% was confirmed by post-sort analysis before cells were used for further

131 experiments. For culture experiments, DC progenitors were cultured at 37⁰C in 200 µL total volume of cIMDM supplemented with 100 ng/mL Flt3L (Peprotech) for five days before further analysis.

Retroviral infection and culture

Retroviruses were produced by transfecting retroviral vectors into Plat-E cells essentially as described 26 and collecting viral supernatants 2 days later. For reporter assays, Lin—CD117hi

BM cells were sorted as described above and transduced with viral supernatants by 'spin infection' at 1800 RPM for 1 hour in the presence of 2 ug/mL polybrene. Infected cells were then cultured in Flt3L for 8 days before DCs were analyzed by flow cytometry. For in vitro

CRISPR/Cas9 deletion, Lin— CD117hi BM cells from R26Cas9/+ mice were sorted and the same transduction protocol was used.

The retroviral reporter vector (Thy1.1 pA GFP CMVp_min PmeI MCS RV) was generated by replacing the XhoI-EcoRI fragment containing the IRES-GFP from the MSCV

IRES GFP vector49 with a XhoI-EcoRI fragment containing IRES Thy1.1 from the MSCV-

IRES-Thy1.1 vector50 to produce the Thy1.1 only RV. The SalI-BamHI fragment from hCD4 pA

51 GFP RV containing GFP-Kb pA was then blunted using Pfu polymerase and inserted into the blunted EcoRI site of Thy1.1 only RV to produce a Thy1.1 pA GFP RV vector. Annealed oligos containing PmeI sites and NcoI and HindIII overhangs

(CATGGTGGCATCCACTAGTTCTAGGATCCGTTTAAACA and

AGCTTGTTTAAACGGATCCTAGAACTAGTGGATGCCAC) were then ligated into the

NcoI-HindIII digested Thy1.1 pA GFP RV vector to produce the Thy1.1 pA GFP PmeI-MCS

132

RV vector. A PCR product containing the minimal CMV promoter and BamHI/BglII sites was then amplified from the GFP min CMVp vector 43 and ligated into the BamHI site of the Thy1.1 pA GFP PmeI-MCS RV vector to produce the final Thy1.1 pA GFP CMVp_min PmeI MCS RV vector. Enhancer regions were cloned into this vector using HindIII and BamHI digests.

The +32 kb enhancer regions were amplified from genomic DNA using Pfu polymerase. Purified

PCR products were digested with HindIII and BglII and cloned into the HindIII and BamH1 digested retroviral reporter vector (Thy1.1 pA GFP CMVp RV). The following primers were used for amplification. For the +32 kb WT region: GCAAGCTTTGAGGTAGAGGGCCCA and

ACGAGATCTGAGGAACACCAGGTCCCA. For the +32 kb 5' region:

GCAAGCTTTGAGGTAGAGGGCCCA and

GAGCTAAGATCTCCTCAATGTCCAAGTTCACC. For the +32 kb 3' region:

TATCGATAAGCTTACGCCAGCAACTTCCTGAATC and

ACGAGATCTGAGGAACACCAGGTCCCA.

A fragment of the Irf8 +32 kb enhancer containing the first three AICEs, either WT or mutated versions, were cloned into this same vector using annealed oligos with HindIII and BglII overhangs. The following primers were used. For the 3X AICE1 WT:

AGCTTtctcttcttgtttctatttcaggttctccttctgaatttcagtttggctcaagttcctgcA and

GATCTgcaggaacttgagccaaactgaaattcagaaggagaacctgaaatagaaacaagaagagaA. For 3X AICE1

Mut: AGCTTtctcttcttgtttctatcaaaggttcaacttctgaatcaaagtttggctcaagttcctgcA and

GATCTgcaggaacttgagccaaactttgattcagaagttgaacctttgatagaaacaagaagagaA.

For in vitro CRISR/Cas9 deletion a Thy1.1-hU6-gRNA-BbsI stuffer RV vector was used

(Thiesen et al, 2018). The following primers containing the sgRNA sequence and BbsI overhangs were annealed and cloned into the BbsI digested vector. For the αIrf8 sgRNA:

133

CACCGagtttaccgaattgtccccg and AAACcggggacaattcggtaaactC. For the αIrf8 +32 kb sgRNA:

CACCGgagccaaactgaaattcaga and AAACtctgaatttcagtttggctcC. For the scramble sgRNA:

CACCggcactaccagagctaactca and AAACtgagttagctctggtagtgcC.

Tumor implantation

The 1969 regressor fibrosarcoma has been previously described31. Tumor cells were thawed and propagated in R10 medium (RPMI + 10% FBS + 0.1% 2-ME). On the day of injection, cells were harvested by incubation in 0.05% trypsin-EDTA, washed three times with endotoxin-free PBS, and then 1x106 cells were injected subcutaneously in a total volume of 0.15 mL of PBS into the shaved flanks of mice. Tumor size was measured every three days beginning on day 4 and is presented as the surface area of the tumor (length X width).

IL-12 administration.

Mice were injected intraperitoneally with either saline (vehicle) or 500 µg of recombinant

IL-12 (Pfizer) on two consecutive days and then analyzed three days after the second injection.

Bone marrow chimeras.

Bone marrow cells from donor mice were collected as described above. Recipient

CD45.1+ mice received a single dose of 950 rads of whole-body irradiation and then received a transplant of 10x106 total BM cells the next day. For mixed BM chimeras, donor marrow from

134 mice of each genotype was mixed at a ratio of 1:1 before transplantation. Mice were analyzed three to four weeks after transplantation for dendritic cell reconstitution.

Salmonella typhimurium infection.

Salmonella enterica Typhimurium wildtype strain SB300A1 52 was used for infection.

Bacteria were grown with shaking overnight at 37°C in Luria-Bertain (LB) broth, subcultured for

4 hours, and washed with with cold PBS prior to use. Intra-peritoneal infection was performed with an inoculum of 5x102 CFU bacteria.

Expression microarray analysis

Progenitor cells or their progeny from in vitro culture were sorted as described above.

RNA from sorted populations was extracted with a NucleoSpin RNA XS Kit (Machery-Nagel), then was amplified with WT Pico System (Affymetrix) and hybridized to GeneChip Mouse Gene

1.0 ST microarrays (Affymetrix) for 18 h at 45 °C in a GeneChip Hybridization Oven 640. The data was analyzed with the Affymetrix GeneChip Command Console. Microarray expression data was processed using Command Console (Affymetrix, Inc) and the raw (.CEL) files generated were analyzed using Expression Console software with Affymetrix default RMA Gene analysis settings.(Affymetrix, Inc). Probe summarization (Robust Multichip Analysis, RMA), quality control analysis, and probe annotation were performed according to recommended guidelines (Expression Console Software, Affymetrix, Inc.). Data were normalized by robust

135 multiarray average summarization and underwent quartile normalization with ArrayStar software

(DNASTAR).

ChIP

ChIP-seq of mature DCs was described previously21. ChIP-seq of DC progenitors was performed similarly, with minor modifications. MDPs, CDPs, and pre-cDC1s were isolated from the BM of WT mice as described above and crosslinked prior to sorting. For crosslinking, cells were incubated for 8 min at room temperature with 1% formaldehyde. Reactions were then quenched with 1.25 M glycine, cells were washed twice with PBS, and pellets were “flash frozen” for storage at -80°C. Chromatin was sonicated at 4°C in sonication buffer (10 mM Tris-

HCl, pH 8.0; 100 nM NaCl; 1 mM EDTA, 0.5 mM EGTA; 0.1% sodium deoxycholate; and

0.5% N-lauroylsarcosine) for 24 cycles of 20 s on and 50 s off per cycle with a Vivra-Cell

VCX130PB and CV188 (Sonics & Material) to obtain DNA fragments from 140 bp to 500 bp.

Chromatin was then immunoprecipitated overnight at 4°C with Dynabeads Protein A that had been pre-incubated with 2.5 µg of the appropriate antibody: anti-H3K27ac (Ab 4729; Abcam) or

H3K4me1 (Ab 8895; Abcam). Beads containing protein-DNA complexes were washed seven times with RIPA buffer (50 mM HEPES, pH 7.5; 500 mM LiCl; 1 mM EDTA; 1% NP-40; and

0.7% sodium deoxycholage). DNA fragments were eluted, and crosslinking was reversed by incubation in a solution containing 50 mM Tris-HCl, pH 8.0; 10 mM EDTA; 1% SDS; and 1 mg/mL of proteinase K (New England Biolabs) for 5 hours at 65°C. DNA was purified by phenol-chloroform extraction followed by ethanol precipitation. Libraries for ChIP-Seq were

136 prepared with a ThruPLEX-FD kit (Rubicon Genomics) and were sequenced with an Illumina

HiSeq 2500.

ChIP-seq of B cells was described previously42. Briefly, in vitro LPS stimulated B cells were subjected to crosslinking at room temperature for either 10 min with 1% formaldehyde

(single crosslinking) or for 45 min with 2 mM disuccinimidyl glutarate (Sigma) followed by 10 min with 1% formaldehyde (double crosslinking). The chromatin was prepared as previously described53. The pelleted genomic DNA, crosslinked with proteins, were sheared with a

Bioruptor® sonicator (Diagenode) followed by immunoprecipitation using an anti-E2A antibody. The precipitated DNA (1-2 ng) was used for library preparation and sequencing.

ATAC-Seq

ATAC-seq of DC progenitors was performed using the Omni-ATAC protocol as previously described with minor modifications 34. 10,000 MDPs, CDPs, and pre-cDC1s were sorted from bone marrow as described above and lysed in ice-cold ATAC-RSB buffer containing

0.1% NP40, 0.1% Tween-20, and 0.01% digitonin. Cells were incubated at 4° C for 3 min, then washed with ATAC-RSB buffer containing only 0.1% Tween-20. Nuclei were spun down by centrifugation and then incubated in 50 µL of transposition buffer (25 µL 2X TD buffer, 22.5 µL dH2O, 2.5 µL Tn5 transposase (Nextera DNA Library Prep Kit, Illumina)) and incubated at 37°

C for 30 min. If 10,000 cells could not be obtained for a certain population then the quantity of

Tn5 transposase was titrated down proportionately to the number of cells obtained but cells were still incubated in 50 µL total. Transposed DNA was purified with a DNA Clean & Concentrator kit (Zymo Research), eluted in 21 µL of elution buffer, and stored at -20° C until amplification.

Three biological replicates for each cell population were obtained and sequenced. ATAC-seq

137 libraries were prepared as previously described, barcoded and sequenced on an Illumina Nextseq

500.

ATAC-seq of cDC1s, pDCs, monocytes, peritoneal macrophages, neutrophils, and follicular B cells was obtained from the Immunological Genome Project Open Chromatin

Regions54.

ATAC-seq of LPS activated B cells was described previously 42.

Computational analysis

For computational analysis of ATAC-seq of DC progenitors, adapter sequences were trimmed using SeqPurge and aligned to mm10 genome using bowtie2. These reads were then filtered for mitochondrial reads, low mapping quality (samtools flag “-F 1804 -f 2 -q 20”), and

PCR duplicates using Picard tools MarkDuplicates. The bam was then converted to a bed and the

Tn5 corrected insertion sites were obtained (“+” stranded + 4 bp, “-” stranded -5 bp)33. To identify peaks, we called peaks for each sample using MACS2 “--shift -75 --extsize 150 -- nomodel --call-summits --nolambda --keep-dup all -q 0.01” using the insertion beds. To get a union peak set, the peak summits were then extended by 250 bp on either side to a final width of

501 bp, filtered by the ENCODE mm10 blacklist

(https://www.encodeproject.org/annotations/ENCSR636HFF/), and filtered to remove peaks that extend beyond the ends of chromosomes. Overlapping peaks were handled using an iterative removal procedure as previously described55. First, the most significant peak (defined by

MACS2 score) is kept and any peak that directly overlaps with that significant peak is removed.

Then, this process iterates to the next most significant peak and so on until all peaks have either been kept or removed due to direct overlap with a more significant peak. This resulted in a union

138 peak set of 188,509 equal width peaks. These peaks were then annotated using ChIPseeker and computed the occurrence of a TF motif using motifmatchr in R with chromVARMotifs mouse_pwms_v1 set 56. All insertions that fell within each peak were then counted using

“countOverlaps” in R to get a counts matrix (peak x samples). To determined differential peaks, the raw counts matrix was used as input into DESeq2 using the modelMatrixType = "expanded" and were tested for whether or not a peak was greater than a Log2FoldChange of 0.5

(lfcThreshold = 0.5, altHypothesis="greaterAbs") 57. A cutoff of an FDR < 0.1 was used to denote a differential peak. For clustering analyses, the counts matrix was then normalized by using edgeR’s “cpm(matrix , log = TRUE, prior.count = 5)” followed by a quantile normalization using preprocessCore’s “normalize.quantiles” in R. TF motif enrichment were calculated using a hypergeometric test in R testing the representation of a motif (from motifmatchr above) in a subset of peaks vs all peaks. To compute TF motif deviations, chromVAR was used in R with raw counts in distal peaks (defined as greater than 1kb from a

TSS in TxDb.Mmusculus.UCSC.mm10.knownGene) and then the top 100 variable TF motifs were determined using variability scores 56. To create sequencing tracks, the Tn5 corrected insertion sites were read into R and a coverage pileup was created that was binned every 100bp using rtracklayer and normalized by reads in peak such that they were all scaled to 30M total reads in peaks (from counts matrix).

Computational analysis of ATAC-seq of LPS activated B cells was described previously42. ChIP-seq data sets were aligned to the mouse genome (GRCm38/mm10 assembly) by Bowtie software (version 1.1.1) with the following parameters: --sam -p 4 -t --verbose --trim5

3 -m 1 mm10 --chunkmbs 1000. Data were visualized with the ‘makeUCSCfile’ program of the

139

Homer software package with default parameters. Peaks were identified with MACS software, version 1.4.2 (‘model based analysis for ChIP-Seq’) with a P value of 1x10-9.

Motifs in the Irf8 +32 kb and +41 kb peaks were identified using the FIMO motif- identification program at a P-value threshold of 1x10-3.

Statistical analysis.

Statistical analyses were performed with Prism (GraphPad Software). Results from independent experiments were pooled as indicated in figure legends. When comparing only two groups, unpaired two-tailed Student's t-test was used. When comparing more than two groups,

Ordinary one-way ANOVA was used. When comparing survival, Log-rank (Mantel-Cox) test was used.

Data availability.

The sequencing and microarray data generated during the course of this study have been deposited and are available on the GEO database. The ChIP-seq data of DC progenitors utilized in Fig. 3.7 can be accessed with the following accession number: GSE132239. The ATAC-seq data of DC progenitors utilized in Figs. 3.10 and 3.11 can be accessed with the following accession number: GSE132240. The microarrays utilized in Fig. 3.6 can be accessed with the following accession numbers: GSE123747, GSE132767, and GSE132768.

All other primary data and materials that support the findings of this study are available from the corresponding author upon request.

3.6 Acknowledgments

140

This work was supported by the Howard Hughes Medical Institute (K. Murphy and H.

Chang), the US National Institutes of Health (F30 DK108498 to V. Durai; K08 CA23188-01 to A.

Satpathy; P50 HG007735 to H. Chang; R01 AI106352 to B. Kee), the National Science Foundation

(DGE-1745038 to P. Bagadia), the Parker Institute for Cancer Immunotherapy (A. Satpathy and

H. Chang), and Boehringer Ingelheim (M. Wöhner, H. Tagoh and M. Busslinger). A. Satpathy was supported by a Career Award for Medical Scientists from the Burroughs Wellcome Fund. This work benefitted from data assembled by the ImmGen consortium54. We thank the Genome

Technology Access Center in the Department of Genetics at Washington University in St. Louis

School of Medicine for help with genomic analysis. The Center is partially supported by NCI

Cancer Center Support Grant #P30 CA91842 to the Siteman Cancer Center and by ICTS/CTSA

Grant# UL1TR000448 from the National Center for Research Resources (NCRR), a component of the National Institutes of Health (NIH), and NIH Roadmap for Medical Research. This publication is solely the responsibility of the authors and does not necessarily represent the official view of NCRR or NIH.

3.7 Author Contributions

V.D., T.L.M., and K.M.M. designed the study; V.D. generated the enhancer knockout mice with advice from S.J.P.; V.D. and P.B. performed experiments related to analysis of immune populations, cell sorting, and culture with advice from J.T.D., R.W., T.L., X.H., C.G.B., and

G.E.G.-R.; V.D., J.M.G., and A.T.S. performed ATAC-seq of DC progenitors; J.M.G. and A.T.S. performed computational analysis of ATAC-seq data; D.H.K. and R.D.N. performed Salmonella infections; A.I. performed ChIP-seq and computational analysis on DC progenitors; M.W., H.T.,

141 and M.B. provided ChIP-seq and ATAC-seq data from B cells and advice; B.L.K. provided Tcf3–

/– mice and advice; V.D. and K.M.M wrote the manuscript with advice from all authors.

3.8 References

1. Graf, T. and Enver, T. Forcing cells to change lineages. Nature 462, 587-594 (2009).

2. Orkin, S. H. and Zon, L. I. Hematopoiesis: an evolving paradigm for stem cell biology. Cell 132, 631-644 (2008).

3. Miyamoto, T. et al., Myeloid or lymphoid promiscuity as a critical step in hematopoietic lineage commitment. Dev.Cell 3, 137-147 (2002).

4. Hoppe, P. S. et al., Early myeloid lineage choice is not initiated by random PU.1 to GATA1 protein ratios. Nature 535, 299-302 (2016).

5. Long, H. K., Prescott, S. L., and Wysocka, J. Ever-Changing Landscapes: Transcriptional Enhancers in Development and Evolution. Cell 167, 1170-1187 (2016).

6. Sagai, T. et al., Elimination of a long-range cis-regulatory module causes complete loss of limb-specific Shh expression and truncation of the mouse limb. Development 132, 797-803 (2005).

7. Shim, S. et al., Cis-regulatory control of corticospinal system development and evolution. Nature 486, 74-79 (2012).

8. Frankel, N. et al., Phenotypic robustness conferred by apparently redundant transcriptional enhancers. Nature 466, 490-493 (2010).

9. Osterwalder, M. et al., Enhancer redundancy provides phenotypic robustness in mammalian development. Nature 554, 239-243 (2018).

10. Lara-Astiaso, D. et al., Immunogenetics. Chromatin state dynamics during blood formation. Science 345, 943-949 (2014). 142

11. Steinman, R. M. and Cohn, Z. A. Identification of a novel cell type in peripheral lymphoid organs of mice. I. Morphology, quantitation, tissue distribution. J Exp.Med. 137, 1142-1162 (1973).

12. Cella, M. et al., Plasmacytoid monocytes migrate to inflamed lymph nodes and produce large amounts of type I interferon [see comments]. Nature Medicine 5, 919-923 (1999).

13. Durai, V. and Murphy, K. M. Functions of Murine Dendritic Cells. Immunity 45, 719- 736 (2016).

14. Hildner, K. et al., Batf3 deficiency reveals a critical role for CD8alpha+ dendritic cells in cytotoxic T cell immunity. Science 322, 1097-1100 (2008).

15. Gubin, M. M. et al., Checkpoint blockade cancer immunotherapy targets tumour- specific mutant antigens. Nature 515, 577-581 (2014).

16. Salmon, H. et al., Expansion and Activation of CD103(+) Dendritic Cell Progenitors at the Tumor Site Enhances Tumor Responses to Therapeutic PD-L1 and BRAF Inhibition. Immunity 44, 924-938 (2016).

17. Saxena, M. and Bhardwaj, N. Re-Emergence of Dendritic Cell Vaccines for Cancer Treatment. Trends Cancer 4, 119-137 (2018).

18. Fogg, D. K. et al., A clonogenic bone marrow progenitor specific for macrophages and dendritic cells. Science 311, 83-87 (2006).

19. Naik, S. H. et al., Development of plasmacytoid and conventional dendritic cell subtypes from single precursor cells derived in vitro and in vivo. Nat Immunol 8, 1217-1226 (2007).

20. Onai, N. et al., Identification of clonogenic common Flt3(+) M-CSFR+ plasmacytoid and conventional dendritic cell progenitors in mouse bone marrow. Nature Immunology 8, 1207-1216 (2007).

21. Grajales-Reyes, G. E. et al., Batf3 maintains autoactivation of Irf8 for commitment of a CD8alpha(+) conventional DC clonogenic progenitor. Nat Immunol 16, 708-717 (2015).

143

22. Schlitzer, A. et al., Identification of cDC1- and cDC2-committed DC progenitors reveals early lineage priming at the common DC progenitor stage in the bone marrow. Nat Immunol 16, 718-728 (2015).

23. Schiavoni, G. et al., ICSBP is essential for the development of mouse type I interferon- producing cells and for the generation and activation of CD8alpha(+) dendritic cells. J Exp.Med. 196, 1415-1425 (2002).

24. Kashiwada, M. et al., NFIL3/E4BP4 is a key transcription factor for CD8{alpha}+ dendritic cell development. Blood 117, 6193-6197 (2011).

25. Hacker, C. et al., Transcriptional profiling identifies Id2 function in dendritic cell development. Nat Immunol 4, 380-386 (2003).

26. Tussiwand, R. et al., Compensatory dendritic cell development mediated by BATF-IRF interactions. Nature 490, 502-507 (2012).

27. Seillet, C. et al., CD8alpha+ DCs can be induced in the absence of transcription factors Id2, Nfil3, and Batf3. Blood 121, 1574-1583 (2013).

28. Cisse, B. et al., Transcription factor E2-2 is an essential and specific regulator of plasmacytoid dendritic cell development. Cell 135, 37-48 (2008).

29. Schonheit, J. et al., PU.1 level-directed chromatin structure remodeling at the Irf8 gene drives dendritic cell commitment. Cell Rep. 3, 1617-1628 (2013).

30. Platt, R. J. et al., CRISPR-Cas9 knockin mice for genome editing and cancer modeling. Cell 159, 440-455 (2014).

31. Diamond, M. S. et al., Type I interferon is selectively required by dendritic cells for immune rejection of tumors. Journal of Experimental Medicine 208, 1989-2003 (2011).

32. Karki, R. et al., IRF8 Regulates Transcription of Naips for NLRC4 Inflammasome Activation. Cell 173, 920-933 (2018).

33. Buenrostro, J. D. et al., Transposition of native chromatin for fast and sensitive epigenomic profiling of open chromatin, DNA-binding proteins and nucleosome position. Nat Methods 10, 1213-1218 (2013).

144

34. Corces, M. R. et al., An improved ATAC-seq protocol reduces background and enables interrogation of frozen tissues. Nat Methods 14, 959-962 (2017).

35. Heath, V. et al., C/EBPalpha deficiency results in hyperproliferation of hematopoietic progenitor cells and disrupts macrophage development in vitro and in vivo. Blood 104, 1639-1647 (2004).

36. Labzin, L. I. et al., ATF3 Is a Key Regulator of Macrophage IFN Responses. The Journal of Immunology 195, 4446-4455 (2015).

37. Sawai, C. M. et al., Transcription factor Runx2 controls the development and migration of plasmacytoid dendritic cells. J Exp.Med 210, 2151-2159 (2013).

38. Satpathy, A. T. et al., Runx1 and Cbfbeta regulate the development of Flt3+ dendritic cell progenitors and restrict myeloproliferative disorder. Blood 123, 2968-2977 (2014).

39. Theisen, D. J. et al., WDFY4 is required for cross-presentation in response to viral and tumor antigens. Science 362, 694-699 (2018).

40. Sichien, D. et al., IRF8 Transcription Factor Controls Survival and Function of Terminally Differentiated Conventional and Plasmacytoid Dendritic Cells, Respectively. Immunity 45, 626-640 (2016).

41. Kwon, K. et al., Instructive role of the transcription factor E2A in early B lymphopoiesis and germinal center B cell development. Immunity 28, 751-762 (2008).

42. Wohner, M. et al., Molecular functions of the transcription factors E2A and E2-2 in controlling germinal center B cell and plasma cell development. J Exp.Med 213, 1201- 1221 (2016).

43. Iwata, A. et al., Quality of TCR signaling determined by differential affinities of enhancers for the composite BATF-IRF4 transcription factor complex. Nat Immunol 18, 563-572 (2017).

44. Zhang, P. et al., PU.1 inhibits GATA-1 function and erythroid differentiation by blocking GATA-1 DNA binding. Blood 96, 2641-2648 (2000).

145

45. Nerlov, C. et al., GATA-1 interacts with the myeloid PU.1 transcription factor and represses PU.1-dependent transcription. Blood 95, 2543-2551 (2000).

46. Tsai, S. F., Strauss, E., and Orkin, S. H. Functional analysis and in vivo footprinting implicate the erythroid transcription factor GATA-1 as a positive regulator of its own promoter. Genes & Development 5, 919-931 (1991).

47. Chen, H. et al., PU.1 (Spi-1) autoregulates its expression in myeloid cells. Oncogene 11, 1549-1560 (1995). 48. Durai, V. et al., Altered compensatory cytokine signaling underlies the discrepancy between Flt3(-/-) and Flt3l(-/-) mice. J Exp.Med 215, 1417-1435 (2018).

49. Ranganath, S. et al., GATA-3-dependent enhancer activity in IL-4 gene regulation. Journal of Immunology 161, 3822-3826 (1998).

50. Sedy, J. R. et al., B and T lymphocyte attenuator regulates T cell activation through interaction with herpesvirus entry mediator. Nat.Immunol. 6, 90-98 (2005).

51. Zhu, H. et al., Unexpected characteristics of the IFN-gamma reporters in nontransformed T cells. Journal of Immunology 167, 855-865 (2001).

52. McKinney, J. et al., Tightly regulated gene expression system in Salmonella enterica serovar Typhimurium. J Bacteriol. 184, 6056-6059 (2002).

53. Kohwi-Shigematsu, T. et al., SATB1-mediated functional packaging of chromatin into loops. Methods 58, 243-254 (2012).

54. Heng, T. S., Painter, M. W., and Immunological Genome Project Consortium The Immunological Genome Project: networks of gene expression in immune cells. Nat Immunol 9, 1091-1094 (2008).

55. Corces, M. R. et al., The chromatin accessibility landscape of primary human cancers. Science 362, (2018).

56. Schep, A. N. et al., chromVAR: inferring transcription-factor-associated accessibility from single-cell epigenomic data. Nat Methods 14, 975-978 (2017).

146

57. Love, M. I., Huber, W., and Anders, S. Moderated estimation of fold change and dispersion for RNA-seq data with DESeq2. Genome Biol 15, 550-(2014).

147

Figure 3.1. The +32 kb Irf8 enhancer is required for cDC1 development.

148

(A) Normalized sequencing tracks of ChIP-seq with anti-p300, anti-BATF3, or anti-IRF8 antibodies or of ATAC-seq in the indicated populations. Boxed is the +32 kb Irf8 enhancer. Shown below is the +32 kb Irf8 enhancer with the AICE motifs and sgRNA target sequences depicted. Data are pooled from two independent experiments and the Immunological Genome Project Open Chromatin Regions (n = 1 biological replicate per population).

(B) Flow cytometry of live splenocytes from mice of the indicated genotypes was used to identify dendritic cell (DC) subsets. Numbers indicate the percent of cells in the indicated gates. Data are representative of four independent experiments with similar results (n = 5 mice for WT, n = 5 mice for Irf8 +32–/–, n = 3 mice for Batf3–/–, and n = 3 mice for Irf8–/–).

(C) Intracellular staining for IRF8 in splenic pDCs of mice of the indicated genotypes. Numbers indicate the mean fluorescence intensity (MFI) of IRF8 protein levels in pDCs. Data are representative of four independent experiments with similar results (n = 5 mice for WT, n = 5 mice for Irf8 +32–/–, n = 3 mice for Batf3–/–, and n = 3 mice for Irf8–/–).

(D) Statistical analysis of the frequency of splenic cDC1s and pDCs in mice of the indicated genotypes. Small horizontal lines indicate the mean. Data are pooled from four independent experiments (n = 5 mice for WT, n = 5 mice for Irf8 +32–/–, n = 3 mice for Batf3–/–, and n = 3 mice for Irf8–/–).

(E) Growth of 1969 regressor fibrosarcomas in WT and Irf8 +32 kb–/– mice. Data are pooled from two independent experiments (n = 8 mice for WT, n = 8 mice for Irf8 +32–/–). ns, not significant (P>0.05); *P<0.05 and ****P<0.0001 (Ordinary one-way ANOVA (D)).

149

Figure 3.2. AICEs in the +32 kb Irf8 enhancer drive cDC1 development.

150

(A) Shown are the sequences, either wild type (3x AICE1 WT) or mutated (3x AICE1 Mut), of the first 3 AICEs from the +32 kb Irf8 enhancer used in retroviral reporter assays in (B).

(B) The regions from (A) were placed in a reverse strand retroviral reporter and GFP activity was measured in either cDC1s or cDC2s as previously described21. Gray histograms represent the activity of the empty reporter. Data are representative of three independent experiments with similar results (n = 3 biological replicates per group).

(C) The full 574 bp +32 kb Irf8 enhancer (+32kb WT), or the 5’ half containing 4 AICE sites (+32 kb 5’) or the 3’ half containing no AICEs (+32 kb 3’), were assayed for activity in cDC1s using retroviral reporters as in (B). Numbers indicate the mean fluorescence intensity (MFI) of GFP in cDC1s. Data are representative of three independent experiments with similar results (n = 3 biological replicates per group).

(D) CD117hi BM cells from R26Cas9/+ mice were infected with a retroviral vector expressing a control scramble sgRNA (scramble), an sgRNA targeting the Irf8 coding sequence (aIrf8), or an sgRNA targeting the AICE site 4 shown in (A) (aIrf8 +32kb), and then cultured in Flt3L for 8 days. Flow cytometry was then used to assess cDC1 development. Cells are pre-gated as Thy1.1+, to identify infected cells, and as B220—MHC-II+CD11c+, to identify DCs. Numbers indicate the percent of cells in the indicated gates. Data are representative of three independent experiments (n = 3 biological replicates per group).

(E) Shown is the nucleotide sequence of the +32 kb Irf8 enhancer region. The extent of deletion in the two mouse strains are indicated by colored fonts. Pink nucleotides were deleted only in Irf8 +32 5’—/— mice, blue nucleotides were deleted in both Irf8 +32 5’—/— mice and Irf8 +32—/— mice, and red nucleotides were deleted only in Irf8 +32—/— mice. Red lines indicate the sequences of the sgRNAs used for CRISPR/Cas9 induced deletion in mice. Green boxes indicate the AICE motifs.

(F) PCR of the +32 kb locus in wildtype (WT), Irf8 +32 5’—/—, and Irf8 +32—/— mice demonstrating the deletions produced by CRISPR/Cas9 genome editing. Numbers indicate the size of the PCR product from each strain. Data are representative of two independent experiments with similar results (n = 2 biological replicates per population).

(G,H) Flow cytometry of live splenocytes from mice of the indicated genotypes was used to identify dendritic cell (DC) subsets. Numbers indicate the percent of cells in the indicated gates (G). Statistical analysis of the frequency and absolute number of splenic cDC1s in mice of the indicated genotypes (H). Small horizontal lines indicate the mean. Data are pooled from three independent experiments (n = 6 mice for WT, n = 5 mice for Irf8 +32 5'–/–, and n = 5 mice for Irf8 +32–/–).

***P<0.001 and ****P<0.0001 (unpaired two-tailed Student's t-test (H)).

151

Figure 3.3. Loss of the +32 kb Irf8 enhancer does not impact non-cDC1 immune lineages.

152

(A,B) Flow cytometry of live splenocytes from mice of the indicated genotypes was used to examine myeloid and lymphoid cells. Numbers indicate the percent of cells in the indicated gates (A). Statistical analysis of the frequency of the indicated populations within spleens of mice of the indicated genotypes (B). Small horizontal lines indicate the mean. Data are pooled from four independent experiments (n = 5 mice for WT, n = 5 mice for Irf8 +32–/–, and n = 3 mice for Irf8– /–).

(C) Intracellular staining for IRF8 in the indicated populations in mice of the indicated genotypes. Data are representative of four independent experiments with similar results (n = 5 mice for WT, n = 5 mice for Irf8 +32–/–, and n = 3 mice for Irf8–/–). ns, not significant (P>0.05); **P<0.01, ***P<0.001 and ****P<0.0001 (Ordinary one-way ANOVA (B).

153

Figure 3.4. The +32 kb Irf8 enhancer is required for compensatory cDC1 development but not for cDC1 specification.

(A,B) Flow cytometry of SLNs from mice of the indicated genotypes was used to identify DC subsets. Numbers indicate the percent of cells in the indicated gates (A). Statistical analysis of the frequency of cDC1s in SLNs (B). Small horizontal lines indicate the mean. Data are pooled

154 from four independent experiments (n = 5 mice for WT, n = 5 mice for Irf8 +32–/–, n = 3 mice for Batf3–/–, and n = 3 mice for Irf8–/–).

(C,D) Flow cytometry of live splenocytes from mice of the indicated genotypes after administration of vehicle or IL-12 was used to identify DC subsets. Numbers indicate the percent of cells in the indicated gates (C). Statistical analysis of the absolute number of cDC1s and cDC2s in mice of the indicated genotypes treated with either vehicle or IL-12 (D). Small horizontal lines indicate the mean. Data are pooled from two independent experiments (n = 4 mice per group for WT, n = 4 mice per group for Irf8 +32–/–, n = 4 mice per group for Batf3–/–).

(E) Flow cytometry of Lin—CD135+ bone marrow cells from mice of the indicated genotypes was used to identify pre-cDC1s. Numbers indicate the percent of cells in the indicated gates. Data are representative of three independent experiments with similar results (n = 3 mice for WT, n = 3 mice for Irf8 +32–/–, and n = 3 mice for Irf8–/–).

(F) Intracellular staining for IRF8 within pre-cDC1s from mice of the indicated genotypes. Numbers indicate the MFI of IRF8 protein levels in the pre-cDC1. Black shaded histograms depict IRF8 protein levels from Irf8—/— mice. Data are representative of three independent experiments with similar results (n = 3 mice for WT, n = 3 mice for Irf8 +32–/–, and n = 3 mice for Irf8–/–).

(G) CDPs (Lin—CD117intCD135+CD115+), pre-cDC1s (Lin—CD117intCD135+CD226+), or pre- cDC2s (Lin—CD117loCD135+CD115+) from mice of the indicated genotypes were sorted and cultured for 5 days with Flt3L. Cells were then stained to identify DC subsets. Cells are pregated as CD11c+MHCII+ cells to identify DCs. Data are representative of three independent experiments with similar results (n = 3 mice for WT and n = 3 mice for Irf8 +32–/–). ns, not significant (P>0.05); *P<0.05, **P<0.01 and ****P<0.0001 (Ordinary one-way ANOVA (B) or unpaired two-tailed Student's t-test (D)).

155

Figure 3.5. The +32 kb Irf8 enhancer is required for all compensatory cDC1 development. 156

(A,B) Flow cytometry of lung cells from mice of the indicated genotypes was used to identify DC subsets. Numbers indicate the percent of cells in the indicated gates (A). Statistical analysis of the frequency of cDC1s among CD45+ cells from the lungs of mice of the indicated genotypes (B). Small horizontal lines indicate the mean. Data are pooled from four independent experiments (n = 5 mice for WT, n = 5 mice for Irf8 +32–/–, n = 3 mice for Batf3–/–, and n = 3 mice for Irf8–/–).

(C,D) BM cells from the mice of the indicated genotypes was transferred into irradiated WT mice and allowed to reconstitute for three weeks. Flow cytometry of live splenocytes from these chimeras was then used to identify DC subsets. Numbers indicate the percent of cells in the indicated gates (C). Statistical analysis of the frequency of splenic cDC1s in the indicated chimeras (D). Small horizontal lines indicate the mean. Data are pooled from two independent experiments (n = 3 for WT into WT chimeras, n = 5 for Irf8 +32–/– into WT chimeras, n = 5 for Batf3–/– into WT chimeras).

(E) Statistical analysis of the frequency of pre-cDC1s in mice of the indicated genotypes. Small horizontal lines indicate the mean. Data are pooled from three independent experiments (n = 3 mice for WT and n = 3 mice for Irf8 +32–/–). ns, not significant (P>0.05); **P<0.01 (Ordinary one-way ANOVA (B,D)) or unpaired two- tailed Student's t-test (E).

157

Figure 3.6. CD226 uniquely identifies pre-cDC1s in the bone marrow.

158

(A) Cd226 gene expression levels in CDPs and pre-cDC1s quantified by gene expression microarrays. Small horizontal lines indicate the mean. Data are pooled from three independent experiments (n = 3 biological replicates per population).

(B) Flow cytometry of Lin— BM cells from WT mice was used to examine marker expression in pre-cDC1s. Data are representative of four independent experiments with similar results (n = 4 mice).

(C,D) CDPs (Lin—CD117intCD135+CD115+) and Zbtb46-GFP+ pre-cDC1s (Lin— CD117intCD135+Zbtb46-GFP+) (C), or CDPs and CD226+ pre-cDC1s (Lin— CD117intCD135+CD226+) (D) were sorted from mice and cultured for 5 days in Flt3L. Cells were then analyzed by flow cytometry to identify dendritic cell subsets. Data are representative of three independent experiments with similar results (n = 3 biological replicates per population).

(E) Microarray analysis of cDC2s derived from in vitro culture of pre-cDC1s from Irf8 +32–/– mice compared to cDC1s derived from in vitro culture of WT pre-cDC1s (left) or compared to cDC2s derived from in vitro culture of WT pre-cDC2s (right), presented as M-plots. Green lines indicate the threshold for a two-fold difference in expression between samples. Data was pooled from two independent experiments (n = 2 biological replicates per population).

(F) Microarray analysis of WT CD24+CD172a+ cDC2s compared to WT CD24–CD172+ cDC2s (left) or compared to WT CD24+CD172a– cDC1s (right), presented as M-plots. Green lines indicate the threshold for a two-fold difference in expression between samples. Data was pooled from two independent experiments (n = 2 biological replicates per population).

(G) Irf4 expression levels in the indicated populations quantified by gene expression microarrays. Small horizontal lines indicate the mean. Data was pooled from two independent experiments (n = 2 biological replicates per population).

(H) Microarray analysis of cDC2s derived from in vitro culture of Batf3–/– pre-cDC1s compared to cDC1s derived from in vitro culture of WT pre-cDC1s (left) or compared to cDC2s derived from in vitro culture of WT pre-cDC2s (right), presented as M-plots. Green lines indicate the threshold for a two-fold difference in expression between samples. Data was pooled from two independent experiments (n = 2 biological replicates per population).

159

Figure 3.7. The -50 kb Irf8 enhancer is not required for dendritic cell development.

(A) Normalized sequencing tracks of ChIP-seq with anti-p300 or anti-H3K27Ac antibodies or of ATAC-seq in the indicated populations. Boxed is the -50 kb Irf8 enhancer. Shown below is the - 50 kb Irf8 enhancer with the PU.1 motifs and sgRNA target sequences indicated. Data are pooled from three independent experiments and the Immunological Genome Project Open Chromatin Regions (n = 1 biological replicate per population).

(B) Flow cytometry of live splenocytes from mice of the indicated genotypes was used to identify DC subsets. Numbers indicate the percent of cells in the indicated gates. Data are representative of four independent experiments with similar results (n = 5 mice for WT, n = 5 mice for Irf8 -50–/–, and n = 4 mice for Irf8–/–).

160

(C) Intracellular staining for IRF8 in splenic cDC1s and pDCs from mice of the indicated genotypes. Numbers indicate the MFI of IRF8 protein levels in cDC1s or pDCs as indicated. Data are representative of four independent experiments with similar results (n = 5 mice for WT, n = 5 mice for Irf8 -50–/–, and n = 4 mice for Irf8–/–).

(D) Flow cytometry of Lin—CD135+ bone marrow cells from mice of the indicated genotypes was used to identify DC progenitors. Numbers indicate the percent of cells in the indicated gates. Data are representative of three independent experiments with similar results (n = 3 mice for WT, n = 3 mice for Irf8 -50–/–, and n = 3 mice for Irf8–/–).

(E) Intracellular staining for IRF8 within MDPs, CDPs, and pre-cDC1s from mice of the indicated genotypes. Numbers indicate the MFI of IRF8 protein levels in the indicated populations. Black shaded histograms depict IRF8 levels from Irf8—/— mice. Data are representative of three independent experiments with similar results (n = 3 mice for WT, n = 3 mice for Irf8 -50–/–, and n = 3 mice for Irf8–/–).

161

Figure 3.8. Loss of the -50 kb Irf8 enhancer does not impact non-monocyte/macrophage immune lineages.

162

(A) PCR of the -50 kb locus in WT and Irf8 -50—/— mice demonstrating the deletion produced by CRISPR/Cas9 genome editing. Numbers indicate the size of the PCR product from each strain. Data are representative of two independent experiments with similar results (n = 2 biological replicates per population).

(B) Shown is the nucleotide sequence of the -50 kb Irf8 enhancer region. The extent of deletion in Irf8 -50—/— mice is indicated by colored font. Red nucleotides were deleted in Irf8 -50—/— mice. Red lines indicate the sequences of the sgRNAs used for CRISPR/Cas9 induced deletion in mice. Purple boxes indicate the PU.1 motifs.

(C) Statistical analysis of the frequency of the indicated populations gated as in Fig. 3.3A in spleens of mice of the indicated genotypes. Small horizontal lines indicate the mean. Data are pooled from four independent experiments. (n = 5 mice for WT, n = 5 mice for Irf8 -50–/–, and n = 4 mice for Irf8–/–).

(D) Intracellular staining for IRF8 in the indicated populations in mice of the indicated genotypes. Data are representative of four independent experiments with similar results (n = 5 mice for WT, n = 5 mice for Irf8 -50–/–, and n = 4 mice for Irf8–/–). ns, not significant (P>0.05); *P<0.01, **P<0.01, and ****P<0.0001 (Ordinary one-way ANOVA (c)).

163

Figure 3.9. The -50 kb Irf8 enhancer controls IRF8 expression in monocytes and macrophages.

(A) Normalized sequencing tracks of ATAC-seq in the indicated populations. Boxed is the -50 kb Irf8 enhancer. Data are from the Immunological Genome Project Open Chromatin Regions (n = 1 biological replicate per population).

(B) Flow cytometry of live splenocytes from mice of the indicated genotypes was used to identify monocytes. Numbers indicate the percent of cells in the indicated gates. Data are

164 representative of four independent experiments with similar results (n = 5 mice for WT, n = 5 mice for Irf8 -50–/–, and n = 4 mice for Irf8–/–).

(C) Intracellular staining for IRF8 in splenic monocytes of mice of the indicated genotypes. Numbers indicate the MFI of IRF8 protein levels in monocytes. Data are representative of four independent experiments with similar results (n = 5 mice for WT, n = 5 mice for Irf8 -50–/–, and n = 4 mice for Irf8–/–).

(D) Statistical analysis of the percent of splenic monocytes in mice of the indicated genotypes. Data are pooled from four independent experiments (n = 5 mice for WT, n = 5 mice for Irf8 -50– /–, and n = 4 mice for Irf8–/–).

(E) Flow cytometry of peritoneal lavage cells from mice of the indicated genotypes was used to identify peritoneal macrophages. Numbers indicate the percent of cells in the indicated gates. Data are representative of four independent experiments with similar results (n = 5 mice for WT, n = 5 mice for Irf8 -50–/–, and n = 3 mice for Irf8–/–).

(F) Intracellular staining for IRF8 in F4/80+ peritoneal macrophages of mice of the indicated genotypes. Numbers indicate the MFI of IRF8 protein levels in peritoneal macrophages. Data are representative of four independent experiments with similar results (n = 5 mice for WT, n = 5 mice for Irf8 -50–/–, and n = 3 mice for Irf8–/–).

(G) WT and Irf8 -50—/— mice were infected with Salmonella typhi and survival was monitored. Data are pooled from two independent experiments (n = 10 mice for WT and n = 10 mice for Irf8 -50–/–). ns, not significant (P>0.05); ****P<0.0001 (Ordinary one-way ANOVA (D) or Log-rank (Mantel-Cox) test (G)).

165

Figure 3.10. ATAC-seq identifies the +41 kb Irf8 enhancer as transiently active in cDC1 progenitors

166

(A) Pearson correlation of all distal ATAC-seq peaks for the indicated populations. Data are pooled from three independent experiments (n = 3 biological replicates per population).

(B) Differential ATAC-seq peaks in the indicated populations. Colors indicate orders of significance derived from DESeq2 analysis, with grey (fdr>0.1), yellow (0.1>fdr>0.01) , orange (0.01>fdr>0.001), and red (fdr<0.001) . Data are pooled from three independent experiments (n = 3 biological replicates per population).

(C) Heat map of k-means clusters for the top 25,000 varying distal ATAC-seq peaks. Colors indicate z-scores of reads in each peak compared with mean reads across all populations. Genes nearest to the clustered peaks and transcription factor motifs (TFs) enriched within peaks are indicated. Data are pooled from three independent experiments (n = 3 biological replicates per population).

(D) Heat map of k-means clusters of TF deviation z-scores for ATAC-seq profiles of the indicated populations. Data are pooled from three independent experiments (n = 3 biological replicates per population).

(E) Normalized sequencing tracks of ATAC-seq in the indicated populations are shown for genes in each k-means cluster. Mpo was present in k-cluster 1, Runx1 was present in k-cluster 3, Wdfy4 was present in k-cluster 4, and Irf8 was present in k-clusters 4 and 5. Data are representative of three independent experiments with similar results (n = 3 biological replicates per population).

167

Figure 3.11. The +41 kb Irf8 enhancer is required for cDC1 specification and development.

(A) Normalized sequencing tracks of ChIP-seq with anti-p300 antibody or of ATAC-seq in the indicated populations. Boxed is the +41 kb Irf8 enhancer. Shown below is the +41 kb Irf8

168 enhancer with the E box motifs and sgRNA target sequences indicated. ATAC-seq data are representative of three independent experiments with similar results (n = 3 biological replicates for MDP ATAC, CDP ATAC, and pre-cDC1 ATAC) and ChIP-seq data are pooled from two independent experiments and the Immunological Genome Project Open Chromatin Regions (n = 1 biological replicate for cDC1 p300 IP, pDC p300 IP, cDC1 ATAC, and pDC ATAC).

(B) Flow cytometry of live splenocytes from mice of the indicated genotypes was used to identify DC subsets. Numbers indicate the percent of cells in the indicated gates. Data are representative of five independent experiments with similar results (n = 6 mice for WT, n = 7 mice for Irf8 +41–/–, and n = 4 mice for Irf8–/–).

(C) Intracellular staining for IRF8 and IRF4 in splenic pDCs of mice of the indicated genotypes. Numbers indicate the MFI of IRF8 or IRF4 protein levels in pDCs. Data are representative of four independent experiments with similar results (n = 5 mice for WT, n = 5 mice for Irf8 +41–/–, and n = 3 mice for Irf8–/–).

(D) Statistical analysis of the frequency of cDC1s and pDCs in spleens of mice of the indicated genotypes. Small horizontal lines indicate the mean. Data are pooled from five independent experiments (n = 6 mice for WT, n = 7 mice for Irf8 +41–/–, and n = 4 mice for Irf8–/–).

(E) Flow cytometry of Lin—CD135+ BM from mice of the indicated genotypes was used to identify MDPs and CDPs. Numbers indicate the percent of cells in the indicated gates. Data are representative of three independent experiments with similar results (n = 3 mice for WT, n = 3 mice for Irf8 +41–/–, and n = 3 mice for Irf8–/–).

(F) Intracellular staining for IRF8 within MDPs and CDPs from mice of the indicated genotypes. Red lines indicate the IRF8 protein level in MDPs and blue lines indicate the IRF8 protein level in CDPs. Numbers in red indicate the MFI of IRF8 protein levels in the MDP and numbers in blue indicate the MFI of IRF8 protein levels in the CDP. Data are representative of three independent experiments with similar results (n = 3 mice for WT, n = 3 mice for Irf8 +41–/–, and n = 3 mice for Irf8–/–).

(G) Flow cytometry of Lin—CD135+ BM from mice of the indicated genotypes was used to identify pre-cDC1s. Numbers indicate the percent of cells in the indicated gates. Data are representative of three independent experiments with similar results (n = 3 mice for WT, n = 3 mice for Irf8 +41–/–, and n = 3 mice for Irf8–/–). ns, not significant (P>0.05); *P<0.05 and ****P<0.0001 (Ordinary one-way ANOVA (D)).

169

Figure 3.12. Loss of the +41 kb Irf8 enhancer does not impact non-DC immune lineages.

170

(A) PCR of the +41 kb locus in WT and Irf8 +41—/— mice demonstrating the deletion produced by CRISPR/Cas9 genome editing. Numbers indicate the size of the PCR product from each strain. Data are representative of two independent experiments with similar results (n = 2 biological replicates per population).

(B) Shown is the nucleotide sequence of the +41 kb Irf8 enhancer region. The extent of deletion in Irf8 +41—/— mice is indicated by colored font. Red nucleotides were deleted in Irf8 +41—/— mice. Red lines indicate the sequences of the sgRNAs used for CRISPR/Cas9 induced deletion in mice. Blue boxes indicate the E box motifs.

(C) Statistical analysis of the frequency of the indicated populations gated as in Fig. 3.3A in spleens of mice of the indicated genotypes. Small horizontal lines indicate the mean. Data are pooled from five independent experiments (n = 7 mice for WT, n = 7 mice for Irf8 +41–/–, and n = 4 mice for Irf8–/–).

(D) Intracellular staining for IRF8 in the indicated populations in mice of the indicated genotypes. Data are representative of four independent experiments with similar results (n = 5 mice for WT, n = 5 mice for Irf8 +41–/–, and n = 3 mice for Irf8–/–). ns, not significant (P>0.05); ***P<0.001 and ****P<0.0001 (Ordinary one-way ANOVA (C)).

171

Figure 3.13. E proteins are required for cDC1 development

(A) Normalized sequencing tracks of ChIP-seq with anti-E2A antibody or of ATAC-seq in lymph node B cells activated by LPS for the indicated number of days. Boxed are the +32 kb and

172

+41 kb Irf8 enhancers. Populations analyzed were activated B cells (CD22+CD138—; Act B), pre-plasmablasts (CD22—CD138—; pre-PB), and plasmablasts (CD22—CD138+; PB) from either WT or Tcf3—/—Tcf4—/— (E2A E2-2 DKO) mice. B cells were either single (s)- or double (d)- crosslinked followed by ChIP with an E2A antibody as indicated. Data are pooled from two independent experiments (n = 1 biological replicate per population and per condition).

(B) Flow cytometry of Lin—CD135+ BM cells or live splenocytes from mice of the indicated genotypes was used to determine intracellular E2A-GFP levels in the indicated populations. Numbers indicate the MFI of E2A-GFP levels. Data are representative of four independent experiments with similar results (n = 3 mice for Tcf3fl/+ and n = 6 mice for Tcf3fl/+VavCretg).

(C,D) BM cells from WT or Tcf3—/— mice were cultured for 9 days in Flt3L. Cells were then analyzed by flow cytometry to identify DC subsets. Numbers indicate the percent of cells in the indicated gates (C). Statistical analysis of the frequency of cDC1s and pDCs from cultures is shown (D). Small horizontal lines indicate the mean. Data are pooled from three independent experiments (n = 3 mice for WT and n = 3 mice for Tcf3–/–).

**P<0.01 (unpaired two-tailed Student's t-test (D)).

173

Figure 3.14. Tcf3–/– BM progenitors have reduced cDC1 and pDC potential

(A) Flow cytometry of BM cells from mice of the indicated genotypes was used to determine intracellular E2A-GFP levels in the indicated populations. CMPs were gated as Lin–CD117hiSca-

174

1–CD16/32+CD34+, CLPs were gated as Lin–CD127+CD135+CD117loB220–CD11c–, and pre-B cells were gated as Lin–CD19+CD25+. Numbers indicate the MFI of E2A-GFP levels. Data are representative of two independent experiments with similar results (n = 3 mice for Tcf3fl/+ and n = 3 mice for Tcf3fl/+VavCretg).

(B,C) Flow cytometry of live splenocytes from mice of the indicated genotypes was used to identify dendritic cell subsets. Numbers indicate the percent of cells in the indicated gates (B). Statistical analysis of the frequency of splenic cDC1s and pDCs in mice of the indicated genotypes (C). Small horizontal lines indicate the mean. Data are pooled from three independent experiments. (n = 4 mice for WT and n = 3 mice for Tcf3–/–).

(D,E) WT CD45.1 mice were lethally irradiated and reconstituted with Tcf3+/+ CD45.2: Tcf3+/+ CD45.1 BM at a 1:1 ratio or Tcf3+/+ CD45.2: Tcf3–/– CD45.1 BM at a 1:1 ratio. Mice were analyzed after four weeks of reconstitution. Flow cytometry of live splenocytes from the indicated chimeras was then used to identify DC subsets gated as the following: cDC2s, CD11c+MHCII+CD172a+; cDC1s, CD11c+MHCII+CD24+CD172a–; pDCs, B220+CD317+. Numbers indicate the percent of cells in the indicated gates (d). Summary data of the chimerism ratio of splenic cDC1s (left) or pDCs (right) presented as the ratio of CD45.1+ to CD45.2+ cells of each type normalized to the ratio of CD45.1+ to CD45.2+ cDC2s within the same mouse (e). Small horizontal lines indicate the mean. Data are pooled from two independent experiments (n = 6 mice for Tcf3+/+ chimeras and n = 4 mice for Tcf3–/– chimeras). ns, not significant (P>0.05); *P<0.01 and ****P<0.0001 (unpaired two-tailed Student's t-test (C,E)).

175

CHAPTER 4:

Discussion and Future Directions

Some of the contents of this chapter have been previously published in the Journal of Experimental

Medicine and Nature Immunology.

Durai V, Bagadia P, Briseño CG, Theisen DJ, Iwata A, Davidson JT 4th, Gargaro M, Fremont DH, Murphy TL, Murphy KM. Altered compensatory cytokine signaling underlies the discrepancy between Flt3-/- and Flt3l-/- mice. J Exp Med. 2018 May 7;215(5):1417-1435.

Durai V, Bagadia P, Granja JM, Satpathy AT, Kulkarni DH, Davidson JT 4th, Wu R, Patel SJ, Iwata A, Liu TT, Huang X, Briseño CG, Grajales-Reyes GE, Wöhner M, Tagoh H, Kee BL, Newberry RD, Busslinger M, Chang HY, Murphy TL, Murphy KM. Cryptic activation of an Irf8 enhancer governs cDC1 fate specification. Nat Immunol. 2019 Sep;20(9):1161-1173.

176

4.1 Abstract

In this dissertation, we addressed several mechanisms controlling the development and function of dendritic cells. First, we found that promiscuous cytokines and their receptors can support dendritic cell development and differential sensitivity to these cytokines provides the basis for the discrepancy between Flt3–/– and Flt3l–/– mice. Flt3L, long thought to be a cytokine that instructs DC development, is in fact not necessary for the DC lineage and instead supports the maturation of already committed DC progenitors. This supportive action can be replicated by other cytokines whose receptors are expressed by these progenitors, including M-CSF and SCF.

The less severe DC deficiency in Flt3–/– mice is due to the increased sensitivity of progenitors in these mice to these compensatory cytokines. We next examined the mechanisms by which enhancers in the lineage-determining transcription factor for cDC1s, Irf8, control its induction within the multipotent CDP. We found that two separate enhancers are utilized during the maturation of cDC1s. First, a +41 kb Irf8 enhancer is activated during the CDP stage to induce

Irf8 and commit these cells towards cDC1 fate. Next, a +32 kb Irf8 enhancer is activated in cDC1-specified cells to autoactivate Irf8 and drive maturation of these cells. The differential utilization of enhancers allowed us to determine that E proteins are critical regulators of Irf8 induction and are necessary for cDC1 lineage commitment. Collectively our studies have elucidated the pathways and mechanisms controlling DC lineage commitment.

177

4.2 The role of supportive cytokines in hematopoiesis

We found that M-CSF and SCF can support DC development and that there was continued development of committed DC progenitors in Flt3–/– mice, which together suggest that

DC lineage commitment does not require the Flt3 pathway and that multiple cytokines can provide the supportive signals necessary for the maturation of these progenitors. Further, the

DCs that develop in Flt3–/– mice were functional and had transcriptional profiles indistinguishable from WT DCs, suggesting that the actions of Flt3L in DC development can be compensated entirely by other cytokines. Though we had originally hypothesized that WT DCs would possess a gene signature indicative of their maturation in response to Flt3 signaling, that we did not see such a signature or a corresponding gene signature suggestive of M-CSF signaling in the Flt3–/– DCs suggests that these cytokines may not impose any long-lasting genetic instruction upon mature DCs, but instead simply support their development. DCs are therefore another example of a lineage that continues to develop and function in the absence of its principal growth factor, similar to granulocytes 1, erythrocytes 2 and lymphocytes 3,4. Findings such as these help elucidate the precise role of cytokines in hematopoietic lineage commitment and suggest at least some level of promiscuity between cytokine pathways and cell types.

If cytokines are not instructive but rather support the maturation of already committed progenitors, then determining the instrinsic signals that lead to this commitment is paramount to understanding hematopoiesis. Cascades of transcription factors together lead to the exclusion of certain fates in multipotent progenitors and the specification of others. But isolating the exact circuitry in control of these lineage choices has been a challenge, as knocking out factors one by one and determining the effect on development or tracking the expression of single factors using reporter mice is labor intensive and costly. With the rapid acceleration in our ability to globally

178 survey the chromatin landscapes and mRNA expression in cell populations, we may soon have more robust tools for dissecting these problems. With future advances we may soon be able to understand the full complexity of hematopoietic lineage diversification.

4.3 Evolving enhancer landscapes in hematopoiesis

Our findings of differential Irf8 enhancer usage highlight how activation of enhancers by transiently active transcriptional networks can control lineage diversification. Early models of hematopoiesis proposed that progenitor fate choice is resolved through stochastic fluctuations in the expression of lineage-determining transcription factors, such as the PU.1/GATA1 circuit detailed previously that regulates the choice between myeloid and megakaryocyte/erythrocyte fates. PU.1 and GATA1 can inhibit one another 5,6 and can also activate their own transcription

7,8. Further, these factors were thought to be co-expressed in progenitor cells that remained uncommitted until fluctuations in the levels of these two factors specified them to a single fate 9.

But recent studies using high-resolution analysis of single progenitor cells demonstrated that no progenitor cells actually co-express these factors 10. Rather, PU.1 levels decay before GATA1 is even expressed in progenitors destined for megakaryocyte/erythrocyte fate. This suggested that an unknown mechanism “chooses” this particular cell fate by inducing GATA1, rather than stochastic fluctuations randomly generating the outcome. Our findings support such a deterministic rather than stochastic model of fate divergence, in which transiently active transcriptional networks initiate the higher expression of lineage-determining transcription factors. Specifically, we found a novel requirement for an enhancer regulated by E proteins in the induction of Irf8 at the CDP stage that is a wholly unexpected but obligate requirement for

179 cDC1 specification. Other transiently acting networks may regulate the induction of factors such as PU.1 or GATA1 in progenitors of other lineages.

Whether enhancers are redundant in critical developmental genes appears to vary on a gene-by-gene basis 11-14. Our results demonstrate that non-redundant enhancers entirely control

Irf8 expression in different lineages, including cDC1s (+32 kb Irf8 enhancer), pDCs (+41 kb Irf8 enhancer), and monocytes/macrophages (-50 kb Irf8 enhancer). Possibly, these different enhancers allow Irf8 to be expressed in various lineages despite the distinct set of transcription factors regulating each cell type. For example, E proteins expressed by B cells and pDCs can drive Irf8 expression from the +41 kb enhancer, while BATF3/IRF8 can drive its expression in cDC1s via the +32 kb enhancer. Differential enhancer activity thereby allows various lineages to express shared genes while still retaining the unique transcriptional circuitry that defines their cellular identity.

4.4 Future Directions

Our finding that multiple cytokines support development of dendritic cells opens up an avenue to explore how receptor expression on DC progenitors is transcriptionally regulated.

Since multiple receptors are expressed on MDPs and CDPs but only Flt3 is expressed on mature

DCs, identifying the transcription factors that control receptor expression in progenitors and in mature cells could identify additional mechanisms that contribute to lineage specification. For example, as alternative fate potentials are lost during the transition from the MDP to the CDP, the factors that control expression of c-Kit or CSF1R could be silenced while the factors that control Flt3 expression are maintained. Isolating these factors could help to explain how DC fate is separately specified from monocyte or granulocyte fate.

180

The supportive actions of cytokines further raise questions regarding hematopoietic niches in the bone marrow. It is possible that different cyotkines are expressed in different locations within the bone marrow and that this contributes to the development of different lineages only in certain locations. Alternatively, cytokines in different locations could contribute at different stages of progenitor maturation depending upon the movement of progenitors through the stroma during the course of their development. These questions could be addressed using fluorescent reporters for cyotkines that map their precise location within the bone marrow.

Our analysis of Irf8 enhancer usages reveals the mehanisms by which this transcription factor is induced during the transition from the MDP to the CDP, how it is initially expressed remains unclear. Early work had suggested it was initially expressed through activity of the -50 kb Irf8 enhancer, but we had demonstrated that deletion of this enhancer did not affect Irf8 expression in DC progenitors. ATAC-seq analysis of hematopoietic stem cells and CMPs may identify additional enhancers not found by our analysis that could regulate early Irf8 expression.

We identified a switch in enhancer usage from the +41 kb Irf8 enhancer to the +32 kb

Irf8 enhancer during the progression from the pre-cDC1 to the cDC1. The mechanisms by which these enhancers switch could be in trans, as the factors that act at each are induced separately, or they could be in cis, as the activity of one enhancer activates the other intrachromosomally.

Generating mice heterozygous in each enhancer could resolve this question, and could more generally identify the mehchanisms utilized by enhancers to act in succession during the development of a cell type.

4.5 References

181

1. Lieschke,G.J. et al. Mice lacking granulocyte colony-stimulating factor have chronic neutropenia, granulocyte and macrophage progenitor cell deficiency, and impaired neutrophil mobilization. Blood 84, 1737-1746 (1994).

2. Wu,H., Liu,X., Jaenisch,R. & Lodish,H.F. Generation of committed erythroid BFU-E and CFU-E progenitors does not require erythropoietin or the erythropoietin receptor. Cell 83, 59-67 (1995).

3. Freeden-Jeffry,U. et al. Lymphopenia in interleukin (IL)-7 gene-deleted mice identifies IL- 7 as a nonredundant cytokine. J Exp. Med 181, 1519-1526 (1995).

4. Peschon,J.J. et al. Early lymphocyte expansion is severely impaired in interleukin 7 receptor-deficient mice. J Exp. Med 180, 1955-1960 (1994).

5. Zhang,P. et al. PU.1 inhibits GATA-1 function and erythroid differentiation by blocking GATA-1 DNA binding. Blood 96, 2641-2648 (2000).

6. Nerlov,C., Querfurth,E., Kulessa,H. & Graf,T. GATA-1 interacts with the myeloid PU.1 transcription factor and represses PU.1-dependent transcription. Blood 95, 2543-2551 (2000).

7. Tsai,S.F., Strauss,E. & Orkin,S.H. Functional analysis and in vivo footprinting implicate the erythroid transcription factor GATA-1 as a positive regulator of its own promoter. Genes Dev. 5, 919-931 (1991).

8. Chen,H. et al. PU.1 (Spi-1) autoregulates its expression in myeloid cells. Oncogene 11, 1549-1560 (1995).

9. Miyamoto,T. et al. Myeloid or lymphoid promiscuity as a critical step in hematopoietic lineage commitment. Dev. Cell 3, 137-147 (2002).

10. Hoppe,P.S. et al. Early myeloid lineage choice is not initiated by random PU.1 to GATA1 protein ratios. Nature 535, 299-302 (2016).

11. Sagai,T., Hosoya,M., Mizushina,Y., Tamura,M. & Shiroishi,T. Elimination of a long-range cis-regulatory module causes complete loss of limb-specific Shh expression and truncation of the mouse limb. Development 132, 797-803 (2005).

12. Shim,S., Kwan,K.Y., Li,M., Lefebvre,V. & Sestan,N. Cis-regulatory control of corticospinal system development and evolution. Nature 486, 74-79 (2012).

13. Frankel,N. et al. Phenotypic robustness conferred by apparently redundant transcriptional enhancers. Nature 466, 490-493 (2010).

14. Osterwalder,M. et al. Enhancer redundancy provides phenotypic robustness in mammalian development. Nature 554, 239-243 (2018).

182