DELINEATING HUMAN DENDRITIC CELL DEVELOPMENT

Jaeyop Lee

Submitted in partial fulfillment of the requirements for the degree of Doctor of Philosophy under the Executive Committee of the Graduate School of Arts and Sciences

COLUMBIA UNIVERSITY

2016

© 2016

Jaeyop Lee

All rights reserved ABSTRACT

Delineating Human Dendritic Cell Development

Jaeyop Lee

The origin of human dendritic cells (DCs) has long been debated. DCs are a subset of innate immune cells that are essential for initiating adaptive immune responses. Determining their ontogeny is critical for vaccine development and for unveiling the molecular mechanism of

DC insufficiency, which underlies many primary immune deficiency disorders and leukemia.

Like all blood cells, human DCs develop from hematopoietic stem cells through a sequence of increasingly restricted progenitors. Initially it was assumed that DCs should derive exclusively from myeloid progenitors. However, during the past few decades, a number of myeloid and lymphoid progenitors have been described and shown to have DC potential, instigating the debate of the myeloid vs. lymphoid origin of DCs. Hindering the resolution of this debate, human DC-restricted progenitors had not been identified. Further, the potential of known progenitors could not be interrogated due to the lack of a culture system that supports simultaneous differentiation of all human DCs subsets, in conjunction with other myeloid and lymphoid cells. In this work, we establish a culture system that supports the development of the three major subsets of DCs (plasmacytoid DCs or pDCs, and the two classical DCs, cDCs) as well as granulocytes, monocytes and lymphocytes. This system combines mitomycin C-treated murine stromal cell lines, MS5 and OP9, together with human recombinant cytokines, FLT3L,

SCF and GM-CSF, and supports the differentiation of progenitor cells at a population and single cell level. Using this culture in combination with a phenotypic characterization of CD34+ progenitor cells, we identify four consecutive DC progenitors with increasing degree of commitment to DCs and describe their anatomical location of development. We show that DCs develop from a granulocyte-monocyte-DC progenitor (GMDP), which produces granulocytes and a monocyte-DC progenitor (MDP), which then generates monocytes and a common DC progenitor (CDP), which produces pDCs and a cDC precursor (pre-cDC), which finally produces cDCs only. Lastly, we establish a staining panel that allows the phenotypic identification and separation of newly found DC progenitors as well as all previously described myeloid and lymphoid progenitors. We investigate their inter-developmental relationship and DC potential at the single cell level. We show that each progenitor population with homogeneous phenotype is heterogeneous in developmental potential and prove that cell surface marker expression cannot be directly equated to a specific developmental potential. In order to resolve the unreliability of phenotype to draw developmental pathways, we propose to use the quantitative clonal output in order to delineate the DC developmental pathway. In summary, our studies provide a new tool to determine DC potential in vitro, identify new stages of DC development, and propose a new method for tracing the developmental pathway for DC lineage. This will generate a new model of dendritic cell hematopoiesis that can explain and reconcile the conflicts of data on DC origin and development. TABLE OF CONTENTS

List of Figures ...... iv

List of Tables ...... vi

Chapter I – Introduction ...... 1

1. Classical hematopoiesis model and conflicts ...... 1

2. Dendritic cell: subsets and functions ...... 8

3. Human dendritic cells ...... 14

4. Dendritic cell-restricted progenitors ...... 18

5. Systems for analysis of human DC development ...... 23

6. Summary and Goals ...... 25

Chapter II – Materials and Methods ...... 27

1. Mononuclear cell isolation from human tissue samples ...... 27

2. Stromal cell culture ...... 28

3. Flow cytometry and cell sorting ...... 28

4. RT-PCR ...... 34

5. Cytokine secretion assay ...... 35

6. Colony forming unit ...... 35

7. CFSE staining ...... 35

8. Clonal analysis ...... 35

9. In vivo transfers ...... 36

i

Chapter III – Establishing a culture system for DC development ...... 38

1. Introduction ...... 39

2. Results ...... 41

Stromal cells support human DC development ...... 41

In vitro DCs are phenotypically and functionally equivalent to ex vivo DCs ...... 49

MS5 and OP9 together with FLT3L, SCF and GM-CSF provide optimal condition for DC

development ...... 55

3. Discussion ...... 61

Chapter IV – Identification of human committed DC progenitors and precursors ...... 62

1. Introduction ...... 63

2. Results ...... 66

Single cell culture reveals developmental heterogeneity within GMPs ...... 66

Differential expression of cytokine receptors allows fractionation of GMPs into three

developmentally restricted progenitors ...... 70

Cytokine receptor expressions analysis in CD34- cells allows the identification of

pre-cDCs ...... 75

Physiological localization of early DC progenitors and precursors ...... 82

Interdevelopmental relationship among early DC progenitors and precursors ...... 88

3. Discussion ...... 92

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Chapter V – Revisiting the DC development model ...... 94

1. Introduction ...... 95

2. Results ...... 97

Exhaustive phenotypic characterization of 10 non-overlapping progenitors ...... 97

Determining the developmental potential of CD34+ progenitors ...... 103

Interdevelopmental relationship among CD34+ progenitors ...... 109

Phenotypically uniform progenitors are clonally heterogeneous ...... 113

Correlation between clonal output and lineage potential ...... 118

3. Discussion ...... 121

Chapter VI – Discussion ...... 122

1. Result summary ...... 122

2. Establishing culture method and its importance in elucidating DC ontogeny ...... 124

3. DC development in health and disease ...... 126

4. Clonal heterogeneity in marker-defined progenitors ...... 128

Bibliography ...... 131

Appendix ...... 144

Published paper, 2nd author: Gaëlle Breton, Jaeyop Lee, Kang Liu & Michel C Nussenzweig.

Defining human dendritic cell progenitors by multiparametric flow cytometry. Nature Protocols.

10, 1407–1422 (2015) ...... 145

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LIST OF FIGURES

Chapter 1

Figure 1.1 – Classical model of hematopoiesis ...... 6

Figure 1.2 – First dendritic cells in history ...... 12

Figure 1.3 – Surface marker phenotype comparison between human and mouse DCs ...... 16

Figure 1.4 – Mouse DC development ...... 21

Chapter 3

Figure 3.1 – Isolation of human CD34+ Hematopoietic stem and progenitor cells (HSPCs) ...... 43

Figure 3.2 – Stromal cell culture and human DC development ...... 45

Figure 3.3 – Stromal cell treatment with Mitomycin C enhances cellular output ...... 47

Figure 3.4 – In vitro DCs are phenotypically and transcriptionally identical to ex vivo DCs ...... 51

Figure 3.5 – In vitro generated DCs are functionally active ...... 53

Figure 3.6 – OP9 stromal cells specifically increase CD141 cDC numbers ...... 57

Figure 3.7 – FLT3L, SCF and GM-CSF increase cellular yield ...... 59

CHAPTER 4

Figure 4.1 – GMPs are clonally and phenotypically heterogeneous ...... 68

Figure 4.2 – Identification of progressively restricted DC progenitors in GMPs ...... 73

Figure 4.3 – Identification and developmental potential of cDC precursors ...... 78

Figure 4.4 – Proliferative capacity of pre-cDCs ...... 80

Figure 4.5 – Presence of early DC progenitors in bone marrow and peripheral tissues ...... 84

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Figure 4.6 – Presence of pre-cDCs in bone marrow and peripheral tissues ...... 86

Figure 4.7 – Interdevelopmental relationship among DC progenitors ...... 90

CHAPTER 5

Figure 5.1 – Phenotypic characterization of 10 non-overlapping populations ...... 101

Figure 5.2 – Megaerythrokaryocyte, myeloid and lymphoid potential ...... 106

Figure 5.3 – Interdevelopmental relationship among CD34+ progenitors ...... 111

Figure 5.4 – Phenotypically uniform progenitors are clonally heterogeneous ...... 115

Figure 5.5 – Correlation between clonal output and lineage potential ...... 119

v

LIST OF TABLES

CHAPTER 2

Table 2.1 – List of primers for RT-PCR analysis ...... 34

CHAPTER 5

Table 5.1 – Phenotype, potential and inter-developmental relationship of human CD34+ hematopoietic progenitor populations ...... 99

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ACKNOWLEDGEMENTS

I owe my deepest debt of gratitude to Dr. Kang Liu, under whom I had the privilege of working in a fascinating project for 5 years, which is the most rewarding aspect as a graduate student. Leaving the lab with a still growing interest in human hematopoiesis is a testament to her mentorship. I want to thank her for constant support, encouragement and patience throughout my academic and scientific training. I am deeply grateful for having an advisor who has gone out of her way to actively endorse and cheer as I pursue my career as a physician-scientist. I could not have asked for a better advisor and she will always be the scientist I look up to.

My thesis committee members, Dr. Megan Sykes, Dr. Boris Reizis, Dr. Lei Ding and Dr.

Stephen Emerson have offered not just scientific opinions, but have shaped my thinking through insightful comments, which were crucial in the development of this project. I also owe deep gratitude to Dr. David Fidock, the director of our program, whose mentorship, guidance and relentless support from the beginning to the end of the program has made me reach the present destination. My colleagues Andrew Lee, Sarah Puhr and Naomi Yudanin have been a great source of positive energy. Their friendship has kept me afloat many times during some of my vulnerable days. I keep all our conversations and funny stories as invaluable gifts. Thanks also to the lab members who have worked with me on the ninth floor of Hammer Building: Carolyn Lee,

Ekaterina Zvezdova, Yu Jerry Zhou and Deguang Liang. Special thanks is due to Arafat Aljoufi for unreservedly helping me with the experiments during the last two years, and to whom I wish the best as he continues with his graduate studies.

Often times, fortune plays a major role in science. I believe I am extremely lucky to have my wife Elaine Jungeun Song without whose support I would not have been able to complete

vii any of this work. For unscientific reasons or for reasons I cannot yet explain, I believe it was a miracle that I met such a wonderful woman. Her love and understanding have comforted me after working late into the night and sustained me during the demanding process of graduate studies. To her I owe all my achievements.

Finally, I give thanks to my parents and my sister, without whom none of this would have been possible. I thank my parents for their incessant prayers, and my sister for thinking I am a little brother worth boasting.

viii

DEDICATION

To my wife, Elaine J. Song

To my parents and my sister

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Chapter I. Introduction

1. Classical hematopoiesis model and conflicts.

Human blood cells, including dendritic cells (DCs), develop from hematopoietic stem cells (HSCs) in the bone marrow through a series of progenitors that sequentially lose lineage potential. Blood cells are mainly categorized into two groups according to their function. One category includes all myeloid cells, which are comprised of red blood cells, megakaryocytes, granulocytes (G), monocytes (M) and DCs; G, M and DCs together they create the first barrier of immunity or innate immunity. These are relatively short-lived1, 2, 3, 4 and need constant replenishment from bone marrow progenitor cells5. The other category of blood cells, lymphoid cells, is comprised of B, NK and T cells, which are responsible for forming the adaptive immune response and they are longer lived than innate cells6, 7, 8. Due to the significant difference in their function, it was originally postulated that myeloid and lymphoid cells comprise two separate lineages, originating from either a common myeloid progenitor or a common lymphoid progenitor, respectively, both downstream from HSCs (Fig 1.1).

This model was supported in humans and mice by prospective isolation of committed progenitors that are restricted to lymphoid and myeloid lineages, respectively. Common lymphoid progenitors (CLPs)9 and common myeloid progenitors (CMPs)10 were first identified in mice by the Weissman group. Shortly after finding the murine CMPs, Manz et al. identified the human CMP counterpart in cord blood and bone marrow within the CD34+ hematopoietic progenitor compartment11. Human CMPs were characterized by the expression of CD123, CD38 and lack of CD45RA (CD34+CD38+CD123+CD45RA-). Single CMP cells cultured on methylcellulose together with IL-3, IL-6, IL-11, stem cell factor (SCF), FMS-tyrosine kinase 3

1 ligand (FLT3L), granulocyte/monocyte-colony stimulating factor (GM-CSF), thrombopoietin

(TPO) and erythropoietin (EPO), generated colonies that could produce granulocytes, macrophages and megaerythrokaryocytes simultaneously. However, when more than 2x103 cells were co-cultured with Sys-1 stromal cells supplemented with human SCF, FLT3L, IL-7 and IL-2, supporting the development of B and NK cells, CMPs failed to produce lymphocytes12. This proves that CMPs are the clonogenic progenitor that has the potential to produce strictly to myeloid cells. Furthermore, this study also showed that upon culture, CMPs can give rise to cells having the phenotype of megaerythrokaryocyte progenitors (MEKs) and granulocyte- macrophage progenitors (GMPs) that are restricted to megaerythrokaryocytic and myeloid (G and M) lineages, respectively11, 13. Based on these results, it was concluded that the megaerythrokaryocytic and myeloid (G and M) lineages bifurcate at the CMP. Therefore, the identification and characterization of the CMP showed that the human myeloid cell lineage originated from a common progenitor, distinct from the lymphoid lineage, and supported the classical model of hematopoiesis.

Another piece of evidence that seemed to support the classical model was the identification of the human CLP14. Analysis of human bone marrow revealed the presence of a progenitor population with CD34+CD10+ phenotype. These cells exhibited robust B and NK cell potential, and T cell potential when cultured on Sys-1 and OP9-DL stromal cells, respectively.

Additionally, CLPs failed to produce granulocytes, macrophages or megaerythrokaryocytes in methylcellulose CFU assay, thus are committed to lymphoid lineage and lack myeloid potential.

The identification of human CMPs and CLPs supports that myeloid and lymphoid lineages diverge very early into two mutually independent trajectories.

2

The premise of the classical model is that progenitor populations isolated based on a homogeneous phenotype should also have homogeneous potential. Candidate progenitors were placed on the hematopoietic map based on their progenitor-progeny relationship with other known progenitors and based on their capacity to produce terminally differentiated cells. CMPs, for example, produced megaerythrokaryocytes, granulocytes and macrophages in methylcellulose cultures, as well as DCs in vivo15. Although there was no experimental evidence that these four cell types are generated from the same clone, it was assumed that all CMP cells had the uniform potential to produce all myeloid cells, and hence was considered the common progenitor of the myeloid lineage.

However, the classical tree-like model and this premise have frequently been challenged.

First, irreversibly separated myeloid and lymphoid lineages appear to be able to “converge” and do not seem to be completely mutually exclusive. For example, analysis of Sys-1 stromal cell cultures showed that single CD34+CD10+ CLPs consistently produced CD33+ myeloid cells with

DC morphology that expressed MHC II (HLA-DR) and CD1a14, 16, 17, 18, suggesting that DC and lymphoid lineage potential is shared by the same clone. When CLPs and CMPs were plated onto

Sys-1 stromal cells in parallel, they both produced human DCs19. Furthermore, when human

CLPs and CMPs were transferred into immunodeficient mice to assess their in vivo potential15, they produced DCs that had identical gene expression signature irrespective of the progenitor of origin. Therefore, these results indicated that DCs could arise from mutually exclusive progenitors and questioned the reliability of the classical model in explaining human DC development.

Second, recent studies in humans have provided evidence that progenitor populations are highly heterogeneous in phenotype and clonal potential. Human CLPs are a mix of two

3 populations, which could be further fractionated based on their CD38 expression18, 20. CD38+

CLPs only produced B and NK cells and were termed B/NK progenitors (BNKP), and the CD38-

CLPs produced both B/NK lymphocytes, monocytes and DCs, and termed multilymphoid progenitor (MLPs). Additionally, single GMP cells from cord blood cultured on OP9-DL stromal cells displayed robust T cell potential20. Although whether they also produced myeloid cells simultaneously was not determined, it suggested that the GMPs, as a population, may not be restricted to myeloid lineage alone. Furthermore, work on patients with AML leukemia also revealed the presence of an expanded cell population with CD34+CD38-CD45RA+CD10- phenotype that could produce B, NK, T cells and myeloid cells, such as granulocytes21, which was consequently termed lymphoid-primed multipotent progenitor (LMPP). Interestingly, this population was also able to produce GMPs without going through CMP stage, suggesting that

GMPs could be a mixed population arising from different sources.

Moreover, in contrast to what is predicted by the tree-like bifurcation model, that is multiple myeloid lineages or lymphoid lineages descend from one common progenitor cell, different lineages very rarely descend from the same progenitor in vivo. By labeling mouse

CMPs and LMPPs (the mouse CLP counterpart) with bar-coded viruses to trace the fate of single progenitors in vivo, Perié et al. and Naik et al. found that DCs and other myeloid or lymphoid cells rarely descend from the same cell22, 23. That is, a differentiated DC rarely shared the same barcode with another myeloid or lymphoid cell. This suggested that DC lineage is imprinted at early stages and distinct from lymphoid and myeloid lineages. Therefore, increasing data appears to conflict with, and cannot be explained by, the classic tree-like bifurcation model of hematopoiesis.

4

However, understanding DC development through single cell fate tracing has been hampered in humans by two main factors: (1) human DCs are comprised of three main subsets24 and a system that can faithfully support the development of all three cell types is missing, (2) a culture system that can simultaneously support DC and multilineage cell development is missing.

Therefore a system that can evaluate granulocyte, monocyte, lymphocytes and all three DC subset potential at the single cell level is required in order to conclusively prove whether the DC lineage arises from myeloid progenitors or lymphoid progenitors or stands as a lineage on its own.

5

Figure 1.1 Classical model of hematopoiesis. Schema shows the hierarchical ordering of progenitor cells and lineage bifurcation. Current model of hematopoiesis postulates that the first lineage commitment decision is made at the MPP stage between myeloid vs. lymphoid. LT-HSC, long-term hematopoietic stem cells; ST-HSC, short-term hematopoietic stem cell; MPP, multipotent progenitor; CMP, common myeloid progenitor; MkEP, Megaerythrokaryocyte progenitor; GMP, granulocyte-macrophage progenitor; CLP, common lymphoid progenitor; B and T cells. Figure was adapted from Adolfsson et al.25

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Figure 7. Current and Alternative Models for Hematopoietic Stem Cell and Blood Lineage Commitment

(A) Current model (Reya et al., 2001) for hematopoietic lineage commitment and development, postulating that the first lineage commitment step of HSCs results in a strict separation of myleopoiesis and lymphopoiesis as supported through the identification of CMPs and CLPs, respectively (Akashi et al., 2000; Kondo et al., 1997).

(B) Alternative model, based on the present studies, in which a pluripotent HSC upon loss of Mk and E potential develops into a lymphoid primed multipotent progenitor (LMPP) that upon loss of GM potential generates the CLP (Adolfsson et al., 2001). (C) Composite model, incorporating the experimental evidence for models (A) and (B). Also shown are expression of key genes in ST-HSCs and LMPPs based7 on Q-PCR data. LT-HSC, long-term hematopoietic stem cell; ST-HSC, short-term hematopoietic stem cell; MPP, multipotent progenitor; LMPP, lymphoid- primed multipotent progenitor; CLP, common lymphoid progenitor; CMP, common myeloid progenitor; GMP, granulocyte/macrophage pronenitor; MkEP, megakaryocyte/erythroid progenitor; B, B cell; T, T cell.

Our findings are particularly intriguing in light of re- didate progenitors in question have not been pro- cent gene profiling studies of HSCs (with full myeloid spectively purified and characterized and therefore pre- and lymphoid lineage differentiation potentials), reveal- sent at low frequencies and assayed along with other ing wide expression of myeloid (MkE and GM) but not progenitors showing additional lineage potentials in the lymphoid gene programs (Delassus et al., 1999; Miya- utilized assays. Thus, under such conditions it has not moto et al., 2002). Combined with our finding, this ob- been possible to conclude whether or not progenitors servation strongly supports that HSCs might initially be showing restricted lineage development are in fact lin- primed to undergo myeloid (Mk, E, and subsequently eage restricted or whether it rather reflects the inability GM) commitment (Figure 7B) and that lymphoid com- of the utilized assays to efficiently support the full lin- mitment through an LSK Flt3+ LMPP stage depends on eage potentials of all progenitors investigated. For in- subsequent activation of lymphoid genes. In that re- stance, the granulocyte potential of a multipotent pro- gard, it is noteworthy that LSK Flt3+ cells upregulate genitor might go unnoticed if investigated at the wrong gene expression for FLT3 and IL-7Ra, the two cytokine time or under suboptimal conditions, as granulocytes receptors critically involved in lymphoid and B cell are very short lived in vitro, whereas monocytes accu- commitment (Peschon et al., 1994; Sitnicka et al., mulate with time in such cultures. Thus, to prove the 2002). Furthermore, in agreement with their sustained existence of other lineage-restricted progenitors, pro- G and M potential, multiplex single cell PCR analysis spective purification must be combined with character- demonstrates that IL-7Ra+ LSK Flt3+ cells also sustain ization of their lineage potentials in efficient in vitro and expression of the G-CSFR. in vivo assays for all lineage potentials, as demon- In fetal (but not adult) hematopoiesis, a number of strated for LSK Flt3+ cells in the present studies. Pro- previous observations have implicated the potential spective purification of such populations will also be existence of lympho-myeloid lineage-restricted pro- required to obtain meaningful genetic information re- genitors, primarily with a combined B cell, monocyte garding the genetic determinants of lineage com- (Cumano et al., 1992; Mebius et al., 2001), and in some mitment. cases also T cell (Lacaud et al., 1998; Lu et al., 2002) In conclusion, we have in the adult mouse bone mar- potential. However, as emphasized by others (Ema and row LSK HSC compartment identified a prominent LSK Nakauchi, 2003; Katsura, 2002), definitive conclusions Flt3+ LMPP, which in contrast to true ST- and LT-HSCs have been difficult to reach based on these studies, lacks significant Mk and E potential but sustains other since they have been limited by the fact that the can- blood lineage developmental potentials. This repre- 2. Dendritic cells: subsets and functions

DCs are immune sentinel cells, prevalent throughout the body, essential for initiating the adaptive immune response and for maintaining self-tolerance. DCs were originally identified from mouse splenic preparations26, 27 and distinguished from macrophages by their dendritic morphology, lack of phagocytic vacuoles and superior capacity to prime T cells for antibody production28 (Fig 1.2). Advancing from this historical point, many studies have identified additional DC subsets and expanded our understanding of their surface molecule expression, functional specialization and transcriptional regulation during development. There are currently three main classes of DCs in mice and humans, and together they finely orchestrate the immune system.

Classical DCs (cDCs): There are two major subsets of cDCs in mice, called cDC1 and cDC229, which can be identified in lymphoid30 and non-lymphoid organs31 throughout the body. In lymphoid organs, they are distinguished from other mononuclear phagocytes by the expression of CD11c and MHCII32. Within the CD11c+MHCII+ cells, these two populations can be identified based on their mutually exclusive expression of CD8 and CD11b33, 34: CD8+CD11b- cDCs and CD8-CD11b+ESAMhi cDCs35, which correspond to cDC1s and cDC2, respectively. cDC1s and cDC2s differ from each other not only in phenotype but also in their functional capacities36.

cDC1s are known for their superior capacity to cross-present antigens found in apoptotic cells in the context of MHC class I to cytotoxic CD8 T cells (CTLs)37, 38, 39, which is essential for combating intracellular pathogens. This was first demonstrated when cDC1s isolated from mice that had been immunized with OVA-loaded cells were superior to cDC2s at inducing the

8 proliferation of OT-I Rag-/- CD8+ T cells in vitro. cDC1s also induce the polarization of T cells

40, 41 into Th1 cells in vivo in an IL-12 dependent manner . cDC1s injected into wild-type mice after in vitro priming with Keyhole limpet hemocyanin (KLH) induced T cell division and

42 secretion of IL-2 and IFNγ, characteristic of Th1 response . Furthermore, cDC1s specifically express TLR343, 44, which is a conserved pathogen recognition receptor (PRR) that recognizes endosomal double-stranded RNA45, and induces the secretion of antiviral interferon-λ upon stimulation46. cDC1s can also be identified in non-lymphoid organs as CD103+CD11b- cDCs47,

48, 49.

cDC2s, on the other hand, are professional antigen presenters, primarily in the context of

MHC class II50, 51. Transcriptional profile comparison between cDC1s and cDC2s showed that cDC2s had more transcripts of genes associated with the MHCII antigen processing pathway, such as LAMP1/2, Cathepsin Z and H2-DMb151, 52. Different from cDC1s, cDC2s are also better

53, 54, 55, 56 inducers of Th2 subset polarization , which is specific for combatting extracellular pathogens and inducing antibody immunity. Mice receiving CD8- cDCs, or cDC2s, primed with

KLH demonstrated increased IL-4, IL-5 and IL-10, cytokines necessary for polarizing Th2, in their lymph nodes compared to those receiving cDC1s42. cDC2s are also present in non- lymphoid organs and can be identified as CD103+CD11b+ cells35.

cDCs are indispensable for successful clearance of pathogens and regulating hematopoiesis. Mice lacking cDCs by conditional ablation using diphtheria toxin (DT) cannot mount an effective adaptive immune response57, 58. In particular, these mice fail to activate CTLs against lymphocytic choriomeningitis virus (LCMV) or mouse hepatitis virus (MHV)59, resulting in chronic infections. More importantly, DT-induced, cDC-less mice displayed a myeloproliferative disorder syndrome, whereby peripheral numbers of neutrophils, eosinophils

9 and monocytes significantly increased as well as their respective progenitors in the bone marrow.

Therefore, cDCs are not only important in regulating immune responses against pathogens, but also in regulating leukopoiesis.

It is important to note, and distinguish, that cDCs are different from another type of mononuclear phagocytes that share similar phenotype with cDCs. These are the monocyte- derived DCs. Although monocyte-derived DCs comprise the majority of CD11c+MHCII+ cells in the periphery, such as in the intestine60 and skin29, 61, and can equally activate naïve T cells62, 63, they are ontologically different from cDCs64. Monocyte derived DCs, as their name suggests, develop from Ly6Chi monocytes circulating in the blood61, while cDCs differentiate from specific cDC precursors (more in detail below).

Plasmacytoid DCs (pDCs). pDCs were initially identified in human tonsils65, and later in the blood66, 67, 68. Although pDCs receive their name from their plasma cell-like morphology, they are functionally distinct from B cells. pDCs are glass adherent and have the capacity to produce interferon alpha (IFNα), which distinguishes them from B cells69, 70. Further studies in mouse identified the murine pDC counterpart in the lymph nodes, which is characterized by

CD11c+B220+Gr-1+ expression71, 72, 73. pDCs are round and lack dendritic processes, have extensive ER subcellular structure and their main function is to produce copious amounts of

IFNα in response to different signals74, 75. pDCs express TLR7 and TLR976, 77, both of which sense different forms of nucleic acid material. TLR9 binds to CpG-rich DNA material, which is commonly found in DNA viral genomes. pDCs respond to cytomegalovirus (CMV)78 and herpes simplex virus 2 and 1 (HSV2/1)79. However, when TLR9-/- pDCs were exposed to RNA viruses, they were still able to produce IFNα, suggesting the expression of a separate sensor for RNA

10 molecules80. This lead to the search and identification of TLR7 on pDCs, which recognizes single-stranded RNA from viral sources, such as influenza81, 82 and vesicuolstomatitis virus

(VSV)83. Despite the functional and morphological difference from cDCs, extensive transcriptional analysis and developmental studies (described below) have shown that pDCs are more closely related to cDCs than to other blood cell types, including B cells84.

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Figure 1.2 First dendritic cells in history. Phase-contrast microscopy of a representative glass- adherent DC isolated from (A) mouse spleen26 or (B) human blood85 fixed with glutaraldehyde.

(C) Electron micrograph shows the cellular structure of a plasmacytoid DC86.

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Published April 1, 1982

REVIEW

and 1176 human thymus grafted subcutaneouslyHUMAN44–46. During DENDRITIC adult life, CELLSthat chemokine receptors can heterodimerize, delivering intracellu- pDCs are produced constantly in the bone marrow and migrate from lar signals through pertussis toxin–insensitive pathways that trigger the boneCo., marrowOrangeburg, to lymph NY, nodes, 6 Ci/mM) mucosal-associated at 104-120 lymphoid h (MLR) tis- andcell adhesion56-72 h but(periodate). not chemotaxis Cells 52were. Thus, CXCR3, CXCR4 and suesharvested and spleen inin steady-state a multisample conditions. harvester, Kinetic studiesand the indicate data that displayedother chemokineas mean counts receptors per on minute pDCs may ___ heterodimerize, stabiliz- pDCsstandard in mice have deviation an average of turnoverthe mean. of about 2 weeks21. ing pDC adhesive interactions with high endothelial venules. A pos- Human blood pDCs express L-selectin13, which is downregulated sible requirement for LFA-1 or other integrins expressed on pDCs, in lymphoid organs, where pDCs are particularly abundant in the T such as VLA4 and VLA5, for firm adherence of pDCs to high cell–rich areas around high endothelial venules2,10 (ResultsFig. 1). Thus, endothelial venules remains to be explored. However, involvement pDCs mayA small constitutively percentage emigrate (1-2%) from the ofblood adherent into lymph human nodes PBMCof VLA5 hashad already similar been morphologic found in pDC migration into tumors, as through high endothelial venules, like naive T lymphocytes. tumor microvasculature expresses a counter-ligand for VLA-5, Consistentfeatures with to thisthe hypothesis,DC isolated noninflamed from mouse lymph lymphoid nodes and organsVCAM-1 (1). (ref.These 51). features included: spleensirregular of L-selectin-deficient shape, abundant mice show phase-dense a substantial reductiongranules in (mitochondria), irregular nucleus pDCs18. However, other studies have suggested that circulating pDCs Pathogen sensing preferentiallywith small accumulate nucleoli, in lymphand anodes prominent when they rim are ofexposed heterochromatin. to The ability ofThere pDCs towere secrete few type surface I interferons depends on cellular an inflammatoryruffles, pinocytic stimulus 13,23,47vesicles,. In addition,or lysosomes, pDC attachment organelles to thatsensors were that promptlyabundant detect in theadjacent presence of DNA and RNA viruses. inflamedmonocytes high endothelial (Fig. venules1 a). We is dominated first devised by E-selectin- a multistep rather I nitialprocedure clues regarding for enrichingthe identity ofthese pDC viral sensors came from the than L-selectin-mediated interactions47. Thus, the relative impor- demonstration that pDCs express a subset of Toll-like receptors tance"candidate" of constitutive DC.and inflammation-induced This involved successive pDC migration depletion into ofT(TLRs), cells, including monocytes, TLR7 and and TLR9B cells. (ref. 53). TLR9 expression the lymphWe could nodes isthen still controversial.show that the cytologic, surface, and accountsfunctional for pDC properties responses ofto CpGhuman oligonucleotides, which mimic

Chemokines and their receptors mediate leukocyte trafficking. The bacterial DNA53. TLR7 is responsible for pDC responses to guanosine Downloaded from pDCsDC express were the similar chemokine to receptortheir counterparts CXCR3 (refs. 13,47–50), in rodents. which analogs as well as imidazoquinolines, which are used as antiviral com- promotesEnriched migration Preparations in response of to Human interferon- DC.γ (IFN-Theγ)–inducible purificationpounds of human in the treatment DC was of human monitored papilloma virus infections53,54. All chemokines such as CXCL9 and CXCL10. CXCR3 is required for TLRs expressed in pDCs signal through the adaptor molecule by phase-contrast microscopy of specimens47 fixed in glutaraldehyde. At each stage of

http://www.nature.com/natureimmunology most pDC migration into inflamed lymph nodes . However, pDCs MyD88, which recruits signaling mediators to activate the transcrip- expressthe atprotocol least three (Table additional I), receptors,samples CCR1,were attachedCCR2 and CCR5,to PLL-coatedtion factor cover NF-κ Bslips54. Unlike and DCs,incubated pDCs do not express TLR2, TLR4, for at the 37°C inflammatory to allow chemokines cells to CCL2, spread. CCL3, DC CCL4 exhibited and CCL5. manyTLR5 processes, or TLR3, which mitochondria, explains why pDCsand do not respond to bacterial They also express two receptors, CXCR4 and (after activation) CCR7, products such as peptidoglycans, lipopolysaccharide and flagellin, or for theirregular constitutive nuclei chemokines (Fig. 1 CXCL12b, c). Monocytes and CCL21 spread(refs. 47–49). circumferentiallypoly(I:C), which and mimics had viral clear double-stranded ruffles, RNA. CCR7pinocytic can mediate vesicles, migration lysosomes, of mouse pDCs and in inoval- vitro ch oremotaxis kidney-shapedBecause nuclei DNA viral (Fig. genomes 1 d). contain Small CpG-rich regions, TLR9 was a jem.rupress.org assayslymphocytes47,49, and CXCR4 showed is involved little in pDC spreading migration (Fig. in vivo ,1 at c), least exceptlikely in candidate the case for pDCof B recognition cells, which of viral DNA. The generation of into tumors 51. Moreover, pDCs express the G protein–coupled recep- MyD88- and TLR9-deficient mice54 allowed the demonstration that tor ChemR23, which drives their migration in response to chemerin, pDCs require TLR9 to respond to DNA viruses, including herpes sim- a peptide chemoattractant that is released by inflamed tissues and plex virus 2 and 1 and mouse cytomegalovirus24,55,56. Although tumorsA (S. Sozzani, unpublished data).B Thus, CXCR3 may not be TLR9-deficient pDCs respond to RNA viruses (such as influenza), solely responsible for pDC migration; at least some redundancy MyD88-deficient pDCs do not. This observation, along with the seems likely in vivo in steady-state and/or inflammatory conditions. structural homology between some TLR7 ligands and ribonu- In vitro studies have shown that CCR2, CCR5, CCR7 and CXCR3 cleotides57, indicated that TLR7 is a likely candidate for pDC recogni- ©2004 Nature Publishing Group Group Publishing Nature ©2004 are not functional in human pDCs48. However, CXCR3 and CXCR4 tion of RNA viruses. This has been demonstrated with on March 19, 2016 can mediate pDC migration when simultaneously engaged49,50, sug- single-stranded RNA (ssRNA) viruses such as influenza58,59 and gesting that chemokine receptor function may require cooperation vesiculostomatitis virus60, as well as synthetic ssRNA sequences rich between distinct receptors. Studies in T cells have demonstrated in uridine that mimic ssRNA viruses58,59.

a bc C

Figure 2 Morphology of pDCs. (a) By electron microscopy, pDCs appear as lymphoblasts with a medium-to-large diameter, a slightly eccentric, indented, round or oval nucleus, lightly stained perinuclear areas and well developed rough endoplasmic reticulum. (b,c) By scanning electron microscopy, resting pDCs haveFro. a spherical 1. Phase-contrast shape (b), whereas microscopy CD40L-activated of glutaraldehyde-fixed, pDCs have a dendritic adherent cell–like mononuclear morphology (c ). cells Original (× 1,080).magnifications, ×7,000 (a) and ×3,000 (b,c). Reproduced(a) Fresh from mononuclear ref. 10 by copyrightcells adherent permission to glass. of the A Rockefeller DO is surrounded University Press.by four monocytes, which exhibit typical ruffles, lysosomes, and pinocytic vesicles. (b, c) DO from DC-enriched fractions, after attachment and spreading on PLL-coated glass cover slips. Dendrites extend in several directions, NATUREand IMMUNOLOGY the cytoplasmV OLUMEis full of 5 granules NUMBER that 12 are DECEMBER mitochondria. 2004 At the top of Fig 1 c is a typical small 1221 lymphocyte, which does not have processes or ruffles when attached to PLL. (d) Monocytes adherent to PLL-coated glass. The cells spread more actively than on non-PLL-coated glass or plastic, and theFIG. characteristic1. Phase-contrast micrographs ruffles ofand dendritic organelles cells isolated are from more peripheral readily lymphoid visualized. organs and fixed in glutaraldehyde. Figs. 1 a-d are from spleen, (e) from cervical lymph node, and (f) from Peyer's patch. The nucleus is large, irregular in shape, and has a refractile quality. The cytoplasm is arranged in processes of varying sizes and shapes, many of which contain spherical phase-dense mitochondria. Occasional refractile lipid granules are also present. A medium size lymphocyte in Fig. 1 b can be used as a size comparison. (a) X 4,500; (b) X 3,500; (c) X 3,200; (d) X 4,600;13 (e) X 3,200; (f) X 3,200. 1145 3. Human dendritic cells

Shortly after the identification of mouse DCs in 1973, the quest for identifying and classifying human DCs began. Elegant methods for human blood fractionation by density gradient centrifugation allowed the first identification of human DCs (<1% of total mononuclear cells)85. As in mice, DCs found in the human blood had extended dendritic processes on the cell surface when observed by confocal microscopy. Human blood DCs were drastically different from blood monocytes in their morphology and their capacity to prime T cells.

However, it wasn’t until more than a decade later that all three DC counterparts were thoroughly phenotypically and functionally characterized in human blood24. The main reason

DCs in humans could not be directly isolated from blood using the murine phenotype was due to the poor translation of mouse surface markers to humans. A characteristic of mouse DCs is their expression of CD11c and MHCII (HLA-DR, in humans). CD11c in human blood is not restricted to DCs and is also present on neutrophils87 and monocytes88, 89. HLA-DR, which is the receptor in which foreign antigen is presented to CD4 T cells, and DEC-205 or CD205, a marker that specifically recognizes cDC1s in mouse, are expressed by DCs, monocytes, and even circulating

CD34+ hematopoietic progenitor cells24. This indicated that markers utilized to study mouse DCs could not be directly translated to the study of human DCs, and additional markers were needed.

Further studies with human blood showed that the human DC counterparts could be detected in lineage negative, (non-T, B, NK, CD14, CD16 monocytes) CD11c+HLA-DR+ cells by the differential expression of four blood-DC antigens (BDCAs); BDCA3 (CD141) distinguishes human cDC1s, BDCA1 (CD1c) labels cDC2s, and BDCA2 (CD303) and 4

(CD304) are expressed on pDCs24, 90, 91. Isolation and functional assays have revealed great conservation between human and mouse DCs. Human pDCs express TLR7 and 9 and respond to

14

CpG by producing high levels of IFNα92 while human cDCs are superior to pDCs, monocytes and macrophages in their antigen presentation capacity93, 94, 95. Human cDC1s also detect apoptotic cells by CLEC9a receptor96, 97, 98 and efficiently cross-present antigens after processing99, 100, 101. Furthermore, TLR3 is specifically expressed on human cDC1s, which secrete IFNλ upon polyI:C stimulation46, 102.

Further transcriptional analysis between mouse and human DCs showed that DCs across species were more transcriptionally similar to each other than are other cell populations within the same species103, 104, 105. Subset-specific transcriptional factors were identically expressed in human and in mouse DCs. Human and mouse pDCs express high levels of SPIB and E2-2106, 107, a master regulator of pDC development; cDC1 cells from mice and humans both expressed IRF8 and BATF3, essential transcription factors for cDC1 commitment; both cDCs subsets expressed

ID2 and CIITA, which are a transcription factor that promotes cDC differentiation and a coactivator that regulates the expression of genes involved in antigen presentation96. Additionally, all three DCs express high levels of FMS-tyrosine kinase 3 (FLT3), which is a cytokine receptor essential for DC development. In fact, human subjects injected with FLT3 ligand (FLT3L) showed dramatic increase of DC numbers in the periphery, mimicking the response to Flt3L injection described in mouse.

Therefore, the high degree of conservation of transcriptional profile and function across species provides strong evidence that human DC development may occur through conserved progenitor stages108. However, while the developmental stages for mouse DCs have been relatively well characterized, defining the process of human DC development has met major challenges. The main issue for such a delay is the lack of a system that supports the unbiased interrogation of DC potential.

15

Figure 1.3 Surface marker phenotype comparison between human and mouse DCs. Human and mouse DC are compared side by side. Boxes show the surface phenotype signature for human and mouse DCs as well as their primary biological function. Top left, cDC1s or CD141 cDCs; top right, cDC2s or CD1c cDCs; bottom, plasmacytoid DCs. Figure adapted from Collin et al.109

16

REVIEWS REVIEWS

27 Classical DCs proven by haematopoieticproven stem by haematopoietic cell transplantation stem cell, but transplantation 27, but whether there is indefinite self-renewal of Langerhans whether there is indefinite self-renewal of Langerhans cells cannot be tested in this setting as they are replaced cells cannot be tested in this setting as they are replaced 28 by donor cells within a few months of transplantation . 28 by donor cells within a few months of transplantation . Transplanted limbs, Transplanted however, maintain limbs, however,their own maintain local their own local Langerhans cell populations,Langerhans consistent cell populations, with a process consistent with a process of local self-renewal29. 29 of local self-renewal . Known unknowns inKnown humans. unknowns It is still in unclear humans. whether It is still unclear whether committed DC progenitorscommitted exist DC in progenitors the human exist bone in the human bone marrow, and DC precursorsmarrow, and have DC yet precursors to be identified have yet to be identified in human blood. Inin the human haematopoietic blood. In the compartment haematopoietic compartment of the bone marrow,of monocytoidthe bone marrow, DCs canmonocytoid be gener DCs- can be gener- Plasmacytoid DCs ated from both granulocyte–macrophageated from both granulocyte–macrophage progenitors progenitors (GMPs) and the recently(GMPs) identified and the recently multi-lymphoid identified multi-lymphoid 30 progenitors (MLPs), which retain 30myeloid potential . progenitors (MLPs), which retain myeloid potential . However, distinct precursors with restricted DC poten- However, distinct precursors with restricted DC poten- tial, equivalent to MDPs and CDPs in mice, have not tial, equivalent to MDPs and CDPs in mice, have not been found. Anotherbeen major found. puzzle Another is the major nature puzzle of is the nature of human DC precursors in blood. Logically, a ‘pre-classical human DC precursors in blood. Logically, a ‘pre-classical DC’ or precursor of human myeloid tissue DCs must DC’ or precursor of human myeloid tissue DCs must exist. However, the mononuclear fraction of human exist. However, the mononuclear fraction of human blood has been analysed exhaustively and there is no blood has been analysed exhaustively and there is no room for the discovery of novel compartments31. Rather, room for the discovery of novel compartments31. Rather, human DC precursors must share identity with a known human DC precursors must share identity with a known cell, such as one of the existing human blood DC sub- Figure 1 | DC classification in mice and humans. Human ‘myeloid’cell, dendritic such as one of the existing human blood DC sub- Figure 1 | DC classification in mice and humans. Human ‘myeloid’ dendritic sets or perhaps circulating CD34+ cells that are found cells (DCs) are equivalent to mouse ‘classical’ DCs. Plasmacytoid DCs sets(pDCs) or areperhaps circulating CD34+ cells that are found cells (DCs) are equivalent to mouse ‘classical’ DCs. Plasmacytoid DCs (pDCs) are in human blood in the steady state32. The identification functionally closely equivalent in both species. Human blood containsin at human least three blood in the steady state32. The identification functionally closely equivalentDC subsets: in both CD141 species.+ myeloid Human DCs, blood CD1c contains+ myeloid at DCs least and three pDCs. The non-classical of immunodeficiency conditions in which spontaneous + + DC subsets: CD141 myeloidCD16 +DCs, human CD1c monocyte myeloid is alsoDCs sometimes and pDCs. consideredThe non-classical a blood myeloidof DCimmunodeficiency18 (not deletion conditions of one in or which more ofspontaneous these precursors occurs might + 18 CD16 human monocyteshown). is also Insometimes particular, considered a subset of aCD16 blood+ monocytes myeloid DC that (not express 6-sulphodeletion LacNAc of one or morehelp of to these decipher precursors DC lineages occurs mightin humans and, in par- shown). In particular, a subset of CD16+ monocytes that express 617-sulpho LacNAc (SLAN; a carbohydrate modification of P-selectin glycoprotein ligand 1)help has to been decipher DCticular, lineages might in reveal humans the relationshipand, in par between- monocytes (SLAN; a carbohydrate modification of P-selectin glycoprotein83,84 ligand 1)+ has been characterized as a DC subset . The CD141 myeloid DCs found in humanticular, blood might are reveal andthe relationshipDCs. Current between models monocytes of DC ontogeny in mice and characterized as a DC subset83,84. The CD141+ myeloid DCs found in human blood are thought to have equivalent antigen cross-presenting function to mouseand DCs DCs. that Current can modelshumans of are DC summarized ontogeny in in mice BOX 1and. thought to have equivalentbe identified antigen cross-presentingby CD103 expression function in most to mousetissues andDCs by that CD8 can expression humans in lymphoid are summarized in BOX 1. be identified by CD103 organs;expression in both in most species tissues C-type and lectin 9A by CD8 expression(CLEC9A), XC-chemokine in lymphoid receptor 1 (XCR1) organs; in both species andC-type Toll-like lectin 9A receptor 3 (CLEC9A), (TLR3) XC-chemokine are conserved receptor 1markers of these(XCR1) cells 21–24. Tissue Discovering DC deficiency. Without defined examples Discovering DC deficiency. Without defined examples and Toll-like receptor 3 equivalents(TLR3) are conserved of the blood markers CD141 of+ DCthese have cells not21–24 yet. Tissue been described in humans, but most of DC deficiency it has been difficult to tackle the sim- equivalents of the bloodorgans CD141 contain+ DC have a sizeable not yet proportion been described of migratory in humans, CD1c but+ myeloid most DCsof that DC are deficiency distinct it hasple questionbeen difficult of whether to tackle DCs the havesim- a non-redundant organs contain a sizeablefrom proportion macrophages of migratory85,86 and a CD1c second+ myeloid subset ofDCs ‘monocytoid’ that are distinct DCs that expressple question CD14 of whetherrole in DCsresistance have toa non-redundantinfections in humans in vivo or to from macrophages85,86 andand a CD209 second (also subset known of ‘monocytoid’ as DC-SIGN)87 DCs. Their that origin express from CD14 monocytes remainsrole in unprovenresistance toexplore infections more in complex humans aspectsin vivo ofor their to biology, such as and CD209 (also knownbut as DC-SIGN)they are similar87. Their in manyorigin ways from to monocytes the DCs derived remains from unproven classical monocytesexplore more that complextheir aspects developmental of their biology, origin, relationshipsuch as to monocytes have been described in mice19,20. Human lymphoid tissues also contain CD1a+ but they are similar in many ways to the DCs derived from classical monocytes that their developmentaland origin, homeostatic relationship influence to monocytes on regulatory T (TReg) cells. have been described in lymphoid-residentmice19,20. Human lymphoid myeloid DCstissues and also CD14 contain+ monocytoid CD1a+ DCs88. There is a paucity of and homeostatic influenceIt is perhaps on regulatory surprising Tthat (T RegDC) cells.deficiency has not been lymphoid-resident myeloidinformation DCs and on CD14 mouse+ monocytoid blood DCs. DCsThe only88. There candidates is a paucity appear of to be pDCsIt is perhaps and the surprisingdescribed that DC previously, deficiency given has not that been severe defects in cell- precursor of classical DCs (pre-classical DCs), but the frequency of these cells in information on mouse blood DCs. The only candidates appear to be pDCs and the described previously,mediated given that immunity severe defectswould bein expectedcell- if there is an precursor of classical DCsmouse (pre-classical blood is approximately DCs), but the tenfold frequency lower of than these the cells frequency in of DCs in human blood89. BST2, bone marrow stromal antigen 2 (also known as PDCA1);mediated CX CR1, immunityabsolute would requirementbe expected forif there DCs isin an in vivo T cell prim- mouse blood is approximately tenfold lower than the frequency of DCs in human 3 ing. At least part of the problem is that the antigen- 89 CX C-chemokine receptor 1; FLT3, FMS-related tyrosine kinase 3; iNOS,absolute inducible requirement for DCs in in vivo T cell prim- blood . BST2, bone marrow3 stromal antigen 2 (also known as PDCA1); CX CR1, 3 presenting cell compartment is often overlooked, and a CX C-chemokine receptor 1;nitric oxide FLT3, synthase;FMS-related M-CSFR, tyrosine macrophage kinase 3; iNOS, colony-stimulating inducible factoring. receptor; At least part of the problem is that the antigen- 3 lack of DCs might simply have gone undetected in quite nitric oxide synthase; M-CSFR,TIP, TNF macrophageand iNOS producing; colony-stimulating TNF, tumour factor necrosis receptor; factor. presenting cell compartment is often overlooked, and a severely immunocompromised infants. Alternatively, TIP, TNF and iNOS producing; TNF, tumour necrosis factor. lack of DCs might simply have gone undetected in quite the phenotype of DC deficiency might be less severe severely7 ,8immunocompromised infants. Alternatively, precursors that develop in the bone marrowthe phenotype. In the of DCthan deficiency anticipated might owing be to less overlapping severe and redundant precursors that developmouse, in the a series bone ofmarrow progressively7,8. In the restricted than anticipatedprogenitor owingfunctions to overlapping of antigen-presenting and redundant cells. In this case, the cells has been described, including macrophage and DC deletion of one or more DC subsets passes undetected mouse, a series of progressively restricted progenitor functions of antigen-presenting cells. In this case, the precursors (MDPs), which give rise to both monocytes if it does not dramatically compromise resistance to cells has been described, including macrophage and DC deletion of one or more DC subsets passes undetected and DCs, and common DC precursors (CDPs), which infection and survival. Interpolating the evidence from precursors (MDPs), which give rise to both monocytes if it does not dramatically compromise resistance to have restricted DC potential. As in other haematopoietic inducible DC depletion experiments in mice, specific and DCs, and common DC precursors (CDPs), which infection and survival. Interpolating the evidence from lineages, the differentiation of DCs is controlled by the immune response parameters are attenuated when a have restricted DC potential.sequential As expressionin other haematopoietic of growth factor receptorsinducible coupled DC depletion subset experiments is depleted, inbut mice, this specificdoes not always translate lineages, the differentiationto transcription of DCs is factors controlled that directby the cell immunefate6,26. The response bone parametersinto an increase are attenuatedin the mortality when rate a of infection with sequential expression marrowof growth origin factor of receptors human tissue coupled DCs hassubset been is formally depleted, complexbut this pathogensdoes not 4always. translate to transcription factors that direct cell fate6,26. The bone into an increase in the mortality rate of infection with marrow origin of human tissue DCs has been formally complex pathogens4. 576 | SEPTEMBER 2011 | VOLUME 11 www.nature.com/reviews/immunol © 2011 Macmillan Publishers Limited. All rights reserved 576 | SEPTEMBER 2011 | VOLUME 11 www.nature.com/reviews/immunol © 2011 Macmillan Publishers Limited. All rights reserved 4. Dendritic cell-restricted progenitors

In mice, DCs in the periphery are very short lived and need constant replenishment from bone marrow precursors110. After the identification of the aforementioned mouse CMPs, finding the macrophage-dendritic cell progenitor (MDP) that has lost granulocyte potential was a landmark111. Mouse MDPs can be characterized by their Lin-Sca-KithiFlt3+CD115+ CX3CR1+ phenotype and lack lymphoid, megakaryocyte, and granulocyte activity but can generate monocytes, macrophages and DCs, which are the three main mononuclear phagocyte cell types that connect the innate to the adaptive immune response. Downstream MDPs, monocyte and DC developmental pathways split. MDPs generate a common monocyte progenitor (cMOP), which lacks Flt3 ligand receptor, loses DC potential and gives rise to monocytes only112. Toward the

DC branch, MDPs give rise to a common dendritic cell progenitor (CDP)113, which expresses

Flt3 and M-CSFR (CD115)114, but has lost monocyte potential. Downstream CDPs, two branches diverge to produce pDCs and cDCs115, 116. Along the pDC pathway, CDPs produce pre- pDCs, a progenitor that produces pDCs, retains Flt3 but lacks CD115116. Along the cDC pathway, CDPs differentiate into Siglec-H- pre-cDC1s and pre-cDC2s in the bone marrow through a Siglec-H+ stage52, 117. Both pre-cDCs leave the bone marrow, migrate through the blood, and enter peripheral tissues110, where they finish their differentiation into cDC1s and cDC2s52, 117. Hence, cDC commitment and final differentiation are anatomically separated.

Therefore, mouse DC development goes through a sequence of progenitors with increasing commitment and loss of potential to granulocyte, monocyte and other DC subsets.

Despite the transcriptional and functional conservation between humans and mice, the identification of restricted DC precursors in human is an arduous process with major conceptual and technical hurdles.

18

Human DCs were first proposed to derive from circulating blood monocytes. Sallusto and

Lanzavecchia118 showed that blood monocytes cultured in GM-CSF and IL-4 readily adopted dendritic membrane processes, expressed MHC class II and I molecules (e.g. HLA-DR, CD1a, b, c) on the surface, upregulated T cell costimulatory molecules, CD40 and B7, and lacked the monocyte marker CD14, altogether resembling the cDC2s. These monocyte-derived DCs

(MoDCs) also had high antigen presenting activity and could induce strong stimulation of mixed lymphocyte reactions (MLRs) with either allogeneic or autologous T cells. Hence, monocytes were first considered to be the main precursors for human cDC2s118, 119.

However, it was not until recently that MoDCs have been proven to be a separate cell type not normally found in steady state conditions. Further phenotypic, functional and transcription profile characterization conclusively showed that these cells do not give rise to cDC2s. For instance, MoDCs express the C-type lectin molecule DC-SIGN and MMR, receptors that recognize mannose carbohydrates on bacteria and activate phagocytosis, while cDC2s do

24 not . Furthermore, MoDCs were far less prone to maturation upon CD40L or prostaglandin-E2

94 (PGE2) exposure, and less migratory than blood cDC2s in response to CCL21 . Genome-wide expression comparison among monocytes, MoDCs and DC subsets, showed that MoDCs clustered together with monocytes and macrophages rather than with pDCs and cDCs. More importantly, MoDCs did not express FLT3, a DC-specific marker103, 120. In sum, this strongly suggested that blood monocytes are not the cDC precursors in humans.

Another cDC precursor was proposed in parallel to monocytes: the plasmacytoid DC.

Similar to monocytes, pDCs were considered to be the precursors for cDCs because they gained dendritic morphology and stimulated MLRs after culture with IL-3 and CD40L86. However, neither monocytes nor pDCs in the mouse produce any cDCs upon adoptive transfer in the

19 steady state121. Such confusion was clarified after the identification of murine pre-DC115, the cDC precursor that is distinct from monocytes and pDCs. Developmental studies showed that pDCs and cDCs are two separate DC subsets that bifurcate downstream of the CDPs, with pre- cDC arising from the CDP and being the immediate precursor for cDCs.

When I started my thesis work, there was no knowledge of the human counterparts to these sequential cDC progenitors. A major obstacle was that a system that allowed the simultaneous evaluation of all three authentic DC subsets was lacking. Therefore, to enable identification of human DC progenitors, a method to interrogate steady state DC development needed to be established.

20

Figure 1.4 Mouse DC development. Schema shows the sequence of DC-restricted progenitors and their anatomical compartmentalization in mouse. MP, myeloid progenitor; MDP, monocyte-

DC progenitor; CDP, common DC progenitor; pre-DC, cDC precursor. Adapted from Liu et al.122

21

Liu & Nussenzweig Æ Dendritic cell origin and development

Pre-DC Pre-DC DC Flt3L Pre-DC

Flt3L CDP Flt3L pDC MDP pDC MP M-CSF Mono Mono Mono

Tip-DC BM Blood Periphery Commitment Migration Differentiation

Fig. 1. Dendritic cell and monocyte origin and development. ogy. Later, the progeny of pre-DCs become sessile and progenitor, which seeds the skin directly prior to birth. This develop dendritic morphology and join the lymph node DC hypothesis would place the LC in its own differentiation path- network, where they show exhibit their signature probing way, which branches off from the classical DC in the same motion (58, 64). In conclusion, the DC and monocyte lin- place in development as the monocyte (Fig. 1). eages split in the BM, where MDPs give rise to both mono- The dermal layer of the skin contains two subsets of DCs, cytes and CDP; the latter produces pre-DCs, which migrate CD103+CD11bloLangerin+ DCs and CD103)CD11bhiLanger- from BM through the blood to the periphery to give rise to in) DCs (CD103+ and CD103) dermal DCs, respectively) + DCs (Fig. 1). (10, 72–74) (Table 1). Although both LCs and CD103 der- mal DCs express Langerin, CD103+ dermal DCs are more closely related to spleen CD8+ DCs than to LCs (30, 33, 34). Origin of DCs in non-lymphoid tissues 22 While LC development is dependent on M-CSF and indepen-

Skin dent of Flt3, CD103+ dermal DCs like spleen DCs are M-CSF The skin contains LCs in the epidermis and independent and Flt3 dependent (34). Unlike LCs that can CD103+ CD11bloLangerin+ and CD103)CD11bhiLangerin) self-renew in situ, dermal CD103+ DCs are continually replen- DCs in the dermis. LCs have many of the features of DCs, ished from the same blood-borne pre-DCs that give rise to including morphology, ability to re-distribute large amounts lymphoid organ DCs (74). Like CD8+ splenic DCs, develop- of MHC II from the endocytic system to the cell surface, and ment of CD103+ dermal DCs requires expression of the capacity to stimulate allogeneic T-cell proliferation in vitro after IRF8, Id2, and Batf3 transcription factors (see below). Finally, activation (64–68). Importantly, LCs differ from DCs in that both dermal CD103+ dermal DCs and CD8+ spleen DCs they are resistant to irradiation, they are not replaced by BM specialize in antigen cross-presentation (11, 30). CD103) transfer in the steady state because they are self-renewing in dermal DCs differ from the CD103+ in expressing higher situ, and like monocytes and macrophages, their development levels of M-CSFR and lower levels of Flt3. Consistent with is dependent on macrophage colony-stimulating factor this expression profile, either Flt3L or CD115 (MCSF-R) + + ) (M-CSF) (69). LC precursors, which are CX3CR1 CD45 , deficiency results in a decrease in dermal CD103 DCs (34). colonize the dermis during late embryonic development Finally, pre-DCs give rise to CD103) dermal DCs but less (70, 71). In mice, these precursors complete their differentia- efficiently than to CD103+ DCs. Thus, CD103) DCs may be a tion into LCs by postnatal day 2, whereupon they undergo a heterogeneous collection of cells with distinct origins, or they burst of cell division between postnatal d2-d7, which may come from a single Flt3+ precursor that remains M-CSF accounts for the dramatic increase in LCs during this develop- responsive. mental window. The LC precursor in skin appears to resemble Skin-draining lymph nodes contain a subpopulation of the MDP, which is the bone marrow progenitor for mono- DCs that resemble CD103+ and CD103) dermal DCs (72, 74). + + cytes and cDCs with a CD45 CX3CR1 phenotype (70). We Their migration from the skin to the draining lymph nodes is speculate that the MDP or a closely related cell is the LC dependent on CCR7 but independent of CD62L (74). In the

Ó 2010 John Wiley & Sons A/S • Immunological Reviews 234/2010 49 5. Systems for analysis of human DC development

The history of elucidating human DC development is also the history of an in vitro culture that supports DC development. In order to identify progenitors from which DCs develop, a system that allows the ex vivo detection of DC potential is necessary.

Shortly after the GM-CSF and IL-4 culture to produce DCs from monocytes was established, Flt3L treatment was shown to increase the number of DCs in mice. Flt3L injection dramatically boosted the number of CD11c+MHCII+ DCs, more so than when mice were treated with GM-CSF or with GM-CSF and IL-4123. Following clinical investigations examining the effect of FLT3L injections in human subjects provided additional evidence on the effect of

FLT3L as a strong DC-poietin. Volunteers administered with FLT3L showed significant expansion of CD11c+HLA-DR+ cDCs as well as CD11c-CD45RA+CD123hi cells, which correspond to pDCs94, 95, 124. These results together with advances in mouse genetic manipulation investigations have elucidated the indispensable role of FLT3L in DC development125.

Since then, several liquid cultures using FLT3L alone or in combination with other cytokines have been used in an attempt to produce human DCs. The first culture to differentiate unknown precursors to DCs using FLT3L alone tested the capacity of cord blood derived

CD34+CD45RA+CD123+ cells to produce pDCs126. CD34+CD45RA+CD123+ cells successfully generated high numbers of pDCs after 20 days and produced high levels of IFNα upon HSV infection. Although this culture seemed to support pDC development, it could not reliably facilitate the development of cDCs127.

More recently, a two-step culture was reported to produce massive numbers of cDC1 cells97. CD34+ cells were first expanded in the presence of SCF, FLT3L, IL-3 and IL-6, and then were differentiated in the presence of FLT3L, SCF, GM-CSF and IL-4. This culture supported

23 robust production of CLEC9a+ cDC1s, which were capable of engulfing apoptotic cells and expressing cDC1-specific genes (NECL2, BATF3, IRF8 and TLR3). Although a large number of cells bore the CD1a+HLA-DR+ phenotype in this culture, phenotypic characterization to determine whether they were MoDCs or cDC2 that acquired CD1a was not performed.

Additionally, this culture failed to produce any pDCs. Therefore, a culture that supported the development of all three subsets had not yet been established.

In parallel to liquid cultures, a different type of culture system was using immortalized mouse stromal cell lines. CD34+ progenitor cells on a layer of Sys-1 stromal cells with FLT3L developed into pDCs and cDC2s, but not cDC1s19. Interestingly, Sys-1 cells simultaneously supported the development of B cells together with pDCs and cDC2s. Although this culture was not able to produce all three DC subsets, it was a milestone in being able to interrogate two distinct lineages (the DC lineage and B cell lineage) simultaneously. This provided a tool to study the DC lineage in relation to the B cell lineage.

More importantly, because this culture system supported the development of two distinct lineages, it could be used to assess the developmental potential for these two lineages at the single cell level. Previous assays determined the potential of separate lineages using separate cultures, and it was only assumed that a progenitor had multiple potentials. This approach could not resolve clonal heterogeneity within the progenitor population. In other words, whether a single progenitor cell is capable of producing two lineages, or whether two clones individually produce different lineages can only be resolved by single cell analysis.

One of the latest culture systems used to investigate potential at the single cell level employs MS5 stromal cells together with SCF, TPO, IL-7, IL-2, G-CSF, GM-CSF and M-CSF20.

This culture could detect the potential of CD14+CD1a- macrophages in addition to CD14-CD1a+

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DCs, B cells and NK cells (4 lineages). However, the markers used to detect DCs could not discriminate between MoDCs and cDC2s, and hence cannot be used to convincingly define authentic DC lineage. Thus, a culture that supports all three DCs together with lymphocytes was still needed.

One culture that is crucial for determining whether DC-producing clones share myeloid potential is a culture that nurtures both granulocyte and DC development. The primary culture to support myeloid cell development is a semisolid culture with a cocktail of myeloproliferative cytokines11 (granulocyte-colony stimulating factor G-CSF, GM-CSF, erythropoietin, SCF) in methylcellulose. In this system, CD34+ progenitor cells produce megaerythrokaryocytes, granulocytes and macrophages, as determined by their colony morphology, but never DCs.

Therefore, in order to determine whether a progenitor is able to produce both DCs and other myeloid cells, a culture that could evaluate the potential for granulocytes, macrophages and all three DC subsets needed to be developed.

Another powerful system to studying human DC development is xenotransplantaion into immunodeficient mice. Human HSCs transferred into NOD.Cg-Prkdcscid-IL2rgtmlWjl/Sz

(NOD/SCID/IL2rγnull or NSG) NSG bone marrow or intraveneously can recapitulate the development of all human blood cell types128, 129, 130. These mice are able to produce all three DC subsets, as well as robust B cell and monocyte reconstitution. However, NSG mice show poor granulocyte chimerism even after extended periods of time131. Furthermore, because of the need to transfer a large number of human cells (except for HSCs) in order to obtain readable counts,

DC potential can only be determined at the population level and not at the single cell level.

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Therefore, in order to conclusively determine whether a progenitor is restricted to DCs, or belongs to either the myeloid or lymphoid lineage, a culture supporting granulocytes, monocytes, lymphocytes and all three DC lineages from single cells needs to be established.

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6. Summary and Goals

The classical model of hematopoiesis separates blood cells into two lineages (myeloid and lymphoid) each originating from their respective common progenitors. This model was established based on two premises: (1) the development of the myeloid and lymphoid branches does not overlap or converge, and (2) progenitors isolated on a common phenotype also have homogeneous potential. However, these premises have been heavily contested by studies showing that both myeloid and lymphoid progenitors produce DCs, and by showing that progenitors with homogeneous phenotype have great degree of heterogeneity in their clonal potential both in vitro and in vivo. These studies also proposed that DCs, in mice at least, might be a lineage on their own with intermediate DC restricted progenitors, challenging the standard model of hematopoiesis.

Despite the extensive work on mouse DC development, very little is known regarding human DCs. In humans, DCs have only been thoroughly characterized in the past two decades and a system to interrogate the potential of all three DC subsets is still lacking, delaying the progress in identifying human DC restricted progenitors. Furthermore, many oligopotent progenitors have been reported, yet a coherent cross comparison in phenotype, interdevelopmental relationship and clonal potential has not been performed.

In order to clarify the conflict in DC origin, I first established a culture system that allows the simultaneous interrogation of myeloid and lymphoid lineages: granulocytes, monocytes, cDC1s, cDC2s, pDCs and B cells. Using a combination of human cytokines and mouse stromal cell lines, I tested and optimize an in vitro system that can support the development of single progenitor cells into the different cell types.

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Furthermore, based on the evidence that mouse and human DCs have evolutionarily conserved gene expression signatures, I hypothesize that mouse MDPs, CDPs and pre-cDCs can also be found in humans. I searched for and found these precursors by tracing the expression of

DC-poeitin receptor FLT3 and tested them using in vitro and in vivo assays.

Finally, I established a staining panel that can exhaustively identify all currently known hematopoietic progenitors and determine their progenitor-progeny relationship as well as their developmental potential at the single cell level in order to resolve the conflict of the classical model.

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Chapter II. Materials and Methods

1. Mononuclear cell isolation from human tissue samples

Human umbilical cord blood and leukophoretic peripheral blood (buffy coat) were purchased from New York Blood Center. Human bone marrow was obtained from total hip arthroplasty at Hospital for Special Surgery (New York). Tonsils were obtained from routine tonsillectomies performed at the Babies and Children's Hospital of Columbia-Presbyterian

Medical Center. Informed consent was obtained from the patients and/or exempt from informed consent being residual material after diagnosis and fully de-identified. All samples were collected according to protocols approved by the Institutional Review Board at Columbia

University Medical Center and The Rockefeller University. The specimens were kept on ice immediately after surgical removal. Tonsil samples were minced, treated with 5ml of HBSS

(Invitrogen) with 2,000U of collagenase (Roche) at 37oC for 20 minutes. Digestion was stopped by adding 100ul of 0.5M EDTA (Gibco) and incubated for an additional 5 min. Cells were collected, filtered through a 100nm nylon mesh and washed with complete alpha MEM media

(Invitrogen, see below). Cell suspension were then overlaid on Ficoll-Hypaque (Amersham

Pharmacia Biotech, Piscataway, NJ) and spun for 15 min at room temperature. Buffy coat was then washed and collected for cell analysis. Bone marrow samples were preserved in solution containing 1000 Units/ml heparin (NDC #63323-540-11), and digested in RPMI containing 20 mg/ml collagenase IV (Sigma #C-5138) for 15 min at 37oC. After density centrifugation using

Ficoll-Hypaque (Amersham Pharmacia Biotech, Piscataway, NJ), aliquots of mononuclear bone marrow cells were frozen and stored in liquid nitrogen for future analysis.

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2. Stromal cell culture

S17, OP9 and MS5 murine bone marrow stromal cells were maintained and passed in complete alpha MEM medium supplemented with L-glutamine, ribonucleosides and deoxyribonucleosides (Invitrogen) with 10% FCS and penicillin/streptomycin (Invitrogen). 24 hours before coculture with human cells, stromal cells were seeded at 3.75 x 104 cells per well in a 96-well plate or 1.5 x 105 cells per well in a 24-well plate, in 50ul or 500ul, respectively of complete alpha MEM medium. For MS5 and OP9 mix cultures, a fixed number of MS5 cells

(1.5 x105) was mixed with the increasing numbers of OP9 (Fig 3.6) and then plated on 24-well plates. Human cells and cytokines were added the following day reaching a total final volume of

100ul or 1ml for 96-well plates or 24-well plates. Half media was changed every 7 days of culture.

For mitomycin C treatment, stromal cells were treated with 10ug/ml mytomicin C

(Sigma) for three hours, washed with PBS, dislodged from the plate with Trypsin 0.05% (Gibco) and re-seeded as mentioned above.

Human cell culture was performed with medium supplemented with recombinant cytokines: 100ng of FLT3L/ml (Celldex), 20ng/ml SCF (Peprotech) or 10ng/ml GM-CSF

(Peprotech). Cells were harvested between day1-21 for flow cytometry analysis.

3. Flow cytometry and cell sorting

Samples from cord blood, peripheral blood, bone marrow and tonsil were incubated with fluorescent-labeled antibodies for direct analysis on the BDLSR II flow cytometers (Becton

Dickinson Immunocytometry Systems [BDIS], San Jose, CA) or further purification by fluorescence-activated cell sorting on the Influx, both using HeNe and argon lasers.

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For hematopoietic progenitor analysis, single cell lineage potential, developmental hierarchy relationship experiments, and characterization of progenitor cells from cord blood and bone marrow, CD34+ cells were first enriched using CD34 MicroBead Kit and LS MACS magnetic columns (Miltenyi Biotec, Auburn, CA). Enriched CD34+ cells (70%-95% purity) were incubated with appropriate antibody cocktails to each experiment.

For the isolation of CD34+ cells or the identification of human GMDPs, MDPs and CDPs

(Fig 3.1 through 4.7), enriched CD34+ cells were incubated with CD3 (OKT3, Brilliant Violet

(BV) 650, Biolegend), CD19 (HIB19, BV650, Biolegend), CD56 (HCD56, BV650, Biolegend),

CD14 (TuK4, Qdot-655, Invitrogen), CD66b (G10F5, PerCP-Cy5.5, Biolegend), CD303 (201A,

PerCP-Cy5.5, Biolegend), CD141 (M80, PerCP-Cy5.5, Biolegend), CD1c (L161, APC-Cy7,

Biolegend), CD34 (581, Alexa Fluor (AF) 700, Biolegend), CD38 (HIT2, BV421, Biolegend),

CD45RA (HI100, BV510, Biolegend), CD10 (HI10a, PE-Cy7, Biolegend), CD115 (9-4D2-1E4,

APC, Biolegend), CD123 (9F5, PE, BD), CD116 (4H1, FITC, Biolegend). Alternatively, CD135

(4G8, PE, BD) was used instead of CD123.

For purification of differentiated DCs and monocytes from peripheral blood and culture

(Fig 3.2), single cell cultures or NSG bone marrow, cells were stained with LIVE/DEAD® (Life technologies), CD45 (AF700), CD66b (PerCP-Cy5.5), CD56 (B159, Pacific Blue (PB), BD),

CD19 (HIB19, PB, Biolegend), CD14 (Qdot-655), CLEC9a (8F9, PE, Biolegend), CD1c (L161,

PE-Cy7, Biolegend), CD303 (201A, FITC, Biolegend), CD123 (6H6, Brilliant Violet (BV) 510,

Biolegend), CD141 (AD5-14H12, APC, Miltenyi) for 40 minutes on ice. 4 or 10ul of antibody mix was used to stain cells harvested from 96- or 24-well plates, respectively.

For surface marker analysis (Fig 3.4), CD11c (3.9, Biolegend), HLA-DR (G46-6, BD),

CD80 (2D10, Biotin, Biolegend), CD83 (HB15e, Biolegend), CD86 (IT2.2, Biolegend), DC-

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SIGN/CD209 (DCN46, BD) were used in PerCP-Cy5.5 and CD123 (9F5, BD), CX3CR1 (2A9-

1, Biolegend), SIRP α/CD172 (SE5A5, Biolegend) or CD1a (HI149, BD) were used in PE.

For isolation of rare pre-cDCs from cord blood and peripheral blood (Fig 4.3), an enrichment step was performed prior to sorting. In brief, mononuclear cells were incubated with antibodies against CD135 (4G8, PE, BD) and CD117 (A3C6E2, Biotin, Biolegend) for 40 minutes at 4°C. After washing, antibody against PE (PE001, Biotin, Biolegend) was added and incubated for another 10 minutes at 4°C. Following wash, CD117+CD135+ cells were positively selected using anti-biotin MicroBeads and LS MACS magnetic columns (Miltenyi). For sorting pre-cDCs, enriched CD117+CD135+ cells (from CB or PB) or total mononuclear cells (from BM or Tonsils) were stained for CD14 (TuK4, Qdot-655, Invitrogen), CD3 (OKT3, Brilliant Violet

(BV) 650, Biolegend), CD19 (HIB19, BV650, Biolegend), CD20 (2H7, BV650, Biolegend),

CD56 (HCD56, BV650, Biolegend), CD66b (G10F5, PerCP-Cy5.5, Biolegend), CD303 (201A,

PerCP-Cy5.5, Biolegend), CD1c (L161, PE-Cy7, Biolegend), CD141 (M80, PE-Cy7,

Biolegend), CD34 (581, AlexaFluor700, Biolegend), CD117 (104D2, BV421, Biolegend),

CD135 (4G8, PE, BD), CD45RA (HI100, BV510, Biolegend), CD116 (4H1, FITC, Biolegend) and CD115 (9-4D2-1E4, APC, Biolegend) for 40 minutes on ice. pre-cDCs were isolated as

Lin(CD3/19/20/56/14)-Granulocyte(CD66b)-pDC(CD303)-cDC(CD1c/CD141)-CD34-CD117+

CD135+SSCloCD116+CD115-CD45RA+.

For progenitor-progeny relationship between GMDPs, MDPs and CDPs (Fig 4.7), cultured progenitor cells were harvested at specific time points and stained with LIVE/DEAD®

(Invitrogen), CD45 (HI30, AF700, Biolegend), CD14, CD3, CD19, CD56, CD66b, CD303,

CD1c, CD141, CD34 (581, APC-Cy7, Biolegend), CD117 (104D2, BV421, Biolegend), CD123

(9F5, PE, BD), CD45RA (BV510) and CD116 (FITC) for 40 minutes on ice.

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For progenitor-progeny relationship experiments of CDPs and pre-cDCs (Fig 4.7), cultured CDPs were harvested at specific time points and stained with LIVE/DEAD®, CD45,

CD14, CD3, CD19, CD20, CD56, CD66b, CD303, CD1c, CD141, CD34 (581, APC-Cy7,

Biolegend), CD117, CD123 (9F5, PE, BD), CD45RA and CD116 for 40 minutes on ice.

For hematopoietic progenitor cell analysis, single cell lineage potential, developmental hierarchy relationship experiments, and characterization of progenitor cells from cord blood and adult bone marrow (Fig 5.1 through 5.5), enriched CD34+ cells were stained with CD3 (OKT3,

Brilliant Violet (BV) 650, Biolegend), CD19 (HIB19, BV650, Biolegend), CD56 (HCD56,

BV650, Biolegend), CD14 (TuK4, Qdot-655, Invitrogen), CD66b (G10F5, PerCP-Cy5.5,

Biolegend), CD303 (201A, PerCP-Cy5.5, Biolegend), CD141 (M80, PerCP-Cy5.5, Biolegend),

CD1c (L161, APC-Cy7, Biolegend), CD34 (581, Alexa Fluor (AF) 700, Biolegend), CD38

(HIT2, BV421, Biolegend), CD90 (5E10, PE, Biolegend), CD45RA (HI100, AF488, Biolegend),

CD123 (6H6, BV510, BD), CD10 (HI10a, PE-Cy7, Biolegend), CD115 (9-4D2-1E4, APC,

Biolegend). For culture experiments, progenitors were sorted from Lin(CD3/19/56/14/66b/303/

141/1c)-cells and following Table 5.1 surface phenotypes.

For inter-developmental relationship experiments (Fig 5.3), cells from either culture or

NSG bone marrow were stained for LIVE/DEAD® (Life technologies), CD45 (HI30, AF700,

Biolegend), CD14 (Qdot-655), CD3 (OKT3, BV650, Biolegend), CD19 (HIB19, BV650,

Biolegend), CD56 (HCD56, BV650, Biolegend), CD16 (3G8, BV650, Biolegend), CD11c (3.9,

PerCP-Cy5.5, Biolegend), CD66b (PerCP-Cy5.5), CD303 (PerCP-Cy5.5), CD141 (PerCP-

Cy5.5), CD34 (581, APC-Cy7, Biolegend), CD38 (BV421), CD90 (PE), CD7 (CD7-6B7, PE-

Cy7, Biolegend), CD45RA (AF488), CD123 (BV510) and CD115 (APC). Also, CD45RA

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(HI100, BV510, Biolegend) and CD123 (9F5, PE, BD) were used alternatively for experiments measuring CFSE halving.

4. RT-PCR

mRNA was isolated using RNAqueous®-Micro kit (Life technologies) and cDNA was produced by SuperScript® III First Strand kit (Invitrogen). cDNA concentration was measured by Nanodrop 2000c spectrophotometer (Thermo Fisher Scientific) and quantitative PCR was performed using predesigned primers (Table 2.1) and LightCycler® 480 SYBR Green I Master mix (Roche). CT values for each gene transcript of interest was measured by LightCycler® 480

II (Roche). Briefly, samples were heated to 95°C for 5 min and amplified for 40 cycles (melt at

95°C for 10 sec, anneal at 60°C for 20 sec, elongate at 72°C for 20 sec). Relative gene expression was measured as the difference in CT values between the housekeeping gene and the gene of interest (i.e. ΔCT).

Gene Forward Reverse GAPDH 5'- TGGTCACCAGGGCTGCTTTT-3' 5'- ATCTCGCTCCTGGAAGATGGTG-3' IRF8 5'- CTTCGACACCAGCCAGTTCTTC-3' 5'- ACAGCTCTTCCCAGCCTCTTCT-3' E2-2 5'- CTCTTCCTGTACGCCTCCTG-3' 5'- TAGGAGACGATGAGGCCTGT-3' XCR1 5'- TCAAGACGCATGTAAAGAGGTGTAG-3' 5'- GGCAGGGACGTTTAGAGCAT-3' FLT3 5'- GCGTTCCAGAGCCGATCGTGG-3' 5'- GCACAGCACCTTATGTCCGTCCC-3' TLR2 5'- TACCTGTGGGGCTCATTGTG-3' 5'- TGCAGATACCATTGCGGTCA-3' TLR3 5'- AGTGCCGTCTATTTGCCACA-3' 5'- GCATCCCAAAGGGCAAAAGG-3' TLR7 5'- CTCCTTGGGGCTAGATGGTTTC-3' 5'- TGGTGAGGGTGAGGTTCGT-3' TLR9 5'- GAGTGCTGGACCTGAGTGAG-3' 5'- ATCGAGTGAGCGGAAGAAGATG-3'

Table 2.1 List of primers for RT-PCR gene expression analysis

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5. Cytokine secretion assay

DC subsets were sorted from culture and 40,000 cells/100ul were plated in complete

RPMI medium (Invitrogen) containing 10% FCS and penicillin/streptomycin (Invitrogen). Cells were stimulated with 1uM of CpG ODN2216 (Invivogen), 50ug/ml Poly(I:C) (Invivogen) or

10ug/ml LPS (Sigma). 48hrs hours later, supernantant was collected and IFNα (Mabtech), IL-

12p70 and IL-6 were analyzed using ELISA (eBioscience).

6. Colony forming unit

For colony forming unit cultures, 1 x 103 cells or 5 x102 cells were sorted and plated in non-adherent 35mm dish (Stemcell). Colony forming unit assay was performed using

MethoCultTM (Stemcell, H4434), containing SCF, GM-CSF, IL-3 and EPO. Colony-forming unit–cells (CFU-C) were counted after 14 days of culture.

7. CFSE staining

To determine cellular divisions in culture, input populations were labeled for 15min with

5µM carboxyfluorescein diacetate succinimidyl ester (CFSE; Molecular Probes) at 37°C in PBS.

Cells were washed with complete alpha MEM media and plated on stromal cell cultures.

8. Clonal analysis

Single progenitor cells were directly sorted into pre-plated stromal cells on 96-well plates. Immediately after, media containing cytokine mix was added. Each well was harvested after 7-21 days of culture and stained with LIVE/DEAD®, CD45, CD66b, CLEC9a, CD14,

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CD1c, CD303, CD141, CD19 and CD56. Positive clones were determined by the detection of at least 2 (for CDPs) or 7 (for all other progenitors) events in any of the lineage specific gates.

To determine the setting and efficiency for single cell sorting, human PBMCs were stained for 15min with 5µM of carboxyfluoresceindiacetatesuccinimidyl ester (CFSE; Molecular

Probes) at 37°C and CFSE+ single cells were sorted into each well of a 96-well plate. 90.5% of wells contained 1 cell, 9.5% of wells had 0 cells and none had more than 1 cell as verified by microscopy.

9. In vivo transfers

NOD.Cg-Prkdcscid-IL2rgtmlWjl/Sz (NOD/SCID/IL2rγnull) (NSG) and NOD.Cg-Prkdcscid-

IL2rgtmlWjl Tg(CMV-IL3,CSF2,KITLG)1Eav/MloySzJ (NSG-SGM3) mice were developed at the

Jackson Laboratory (Bar Harbor, ME). All experiments were performed according to the guidelines of IACUC at CUMC. Mice were injected intraperitoneally with busulfan (Sigma,

30ug/g of body weight) to ablate endogenous hematopoietic system 2 days prior to human cell transfer.

Briefly, 45mg of busulfan was diluted into 3ml of N,N-Dimethylacetamide (Sigma). 2ml of such solution was then added to 8ml of solution containing 1:1 ratio of molecular grade water

(Invitrogen) and Polyethylene glycol 400 (Fluka). Final cocktail was filtered through 0.2um PES filter (Corning) and chilled in ice prior to injection.

Human progenitors purified from cord blood were resuspended in 10µl PBS and injected into the tibia bone cavity of anesthetized mice. For fig 4.7A, mice were injected intraperitoneally for 5 consecutive days with 10ug Flt3L, starting at 1 day after transfer. 7 or 14 days after transplantation, bone marrow was harvested from recipient mice and analyzed for human CD45+

36 cells. Briefly, tibia bone marrow cells were flushed with cold PBS into individual tubes and spun down. Red blood cells were lysed with 250ul of ACK lysis buffer (Invitrogen) for 1 min and washed with complete alpha MEM media. Cells were filtered through 100um nylon mesh and stained with corresponding antibodies for analysis.

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Chapter III. Establishing a culture system for DC development

Results in this chapter have been published and can be found in: Lee J, Breton G, Aljoufi A, Zhou YJ, Puhr S, Nussenzweig MC, Liu K. Clonal analysis of human dendritic cell progenitor using a stromal cell culture. J Immunol Methods. 2015 Oct;425:21-6. doi: 10.1016/j.jim.2015.06.004. Epub 2015 Jun 6.

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1. Introduction

In humans, there are three subsets of DCs: pDCs, CD1c cDCs and CD141 cDCs.

Together, they orchestrate the innate and adaptive immune response against foreign pathogens and tolerance to self. DCs in the periphery are short-lived and need constant replenishment from bone marrow progenitors cells through hematopoiesis122.

Different lineages depend on distinct cytokines for lineage development, proliferation and survival. FMS-tyrosine kinase 3 ligand (FLT3L) is essential cytokine for all DC lineages in mouse and humans125. Correspondingly, an Flt3L-based in vitro culture system has been established to differentiate and grow murine DCs from bone marrow stem and progenitor cells132. This culture nurtures the differentiation of all three DC subsets from bone marrow cells and allowed the characterization of DC-restricted progenitors in mouse. Another culture important for tracing mouse DC lineage is the stromal cell coculture. Murine S17 and OP9 stromal cells have been widely utilized to investigate DC potential in the context of other cell lineages. S17 cells were used to promote both myeloid (macrophages) and B lymphoid cell development111, 133, while OP9 cells were used to produce primarily lymphoid cells. It is suggested that this difference is due to the expression of IL-15 and lack of M-CSF in OP9 stromal cells134, 135.

Despite the extensive work on mouse DCs, little is known about human DCs development. This is primarily because of a lack of culture that supports the development of all three DC subsets or together with other myeloid or lymphoid cells. Contrasting the murine example, liquid cultures supplemented with human recombinant FLT3L alone only produced pDCs and failed to nurture cDC subsets from human hematopoietic progenitors126, 127, 136.

Recently, a two-step liquid culture system was proposed which attempted to produce DCs using

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FLT3L, GM-CSF, IL-4 and SCF. However, this culture only generated CD141 cDCs but failed to produce pDCs. Murine stromal cells have also been tried for producing human DCs97. Co- culture of human hematopoietic progenitor cells with Sys-1 stromal cells in the presence of

FLT3L, only produced pDCs and CD1c cDCs, but not CD141 cDCs12. Most studies of human

DC development performed in S17 and MS5 stromal cell culture used only CD11c, HLA-DR and CD1a, as markers for DC detection, could not discriminate different DC subsets or between

DCs and monocyte-derived DCs20, 137.

Another issue from these previous attempts is that because of limited lineage output of these cultures, DC development could not be evaluated simultaneously with and in relation to other cell lineages. Therefore, in order to identify DC-restricted progenitors that lack the potential to other lineages, a culture system that can simultaneously evaluate multiple lineages is required.

In this chapter, we show that a mix of MS5 and OP9 is able to simultaneously support not only three DC subsets, but also monocytes, granulocytes and B lymphocytes from human hematopoietic progenitors in the presence of FLT3L. We also optimized this culture for maximum output of all six lineages by treating stromal cells with mitomycin C and adding GM-

CSF and SCF. Finally, we propose to use this culture in identifying human DC progenitors.

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2. Results

Stromal cells support human DC development

None of the previous in vitro cultures attempting human DC differentiation have successfully shown to support the development of all three DC subsets together. In order to establish a culture that supports differentiation of all DC subsets, we have tested the effect of

FLT3L, an essential DC-poietin, in combination with different murine stromal cell lines that had been previously shown to support the development of separate DC subsets. Therefore, we tested three different murine stromal cells (MS5138, S17 and OP9) in the presence of 100ng of

FLT3L/ml for their capacity to support human DC development from CD34+ hematopoietic progenitors isolated from cord blood (Fig 3.1).

After two weeks of culture, cell output was analyzed for the presence of granulocytes, monocytes, CD1c cDCs, CD141 cDCs, pDCs and B cells among total CD45+ leukocytes (Fig

3.2A). Compared to cultures with FLT3L alone, the presence of stromal cells significantly increased total CD45+ cell yield as well as for all identifiable cell subsets (Fig 3.2B). Among the stromal cells, only MS5 allowed the production of all six cell-types and the highest CD45+ cell output. All three stromal cell cultures generated robust granulocytes, but only MS5 and S17 cultures generated high numbers of monocytes and CD1c cDCs. MS5 cultures also produced the highest numbers of pDCs and B cells. On the other hand, OP9 cultures produced the highest numbers of CD141 cDCs followed by MS5, suggesting that OP9 cultures have a greater bias toward CD141 cDC subset. When DC subset composition was analyzed from all three cultures

(Fig 3.2C), we observed that cultures without stroma were biased toward pDCs, MS5 cultures produced similar ratios to the DC composition in peripheral blood, while S17 and OP9 cultures

41 were drastically biased toward CD1c cDCs and CD141 cDCs, respectively. We conclude that

MS5 are capable to support development of multiple lineages including all three DC subsets.

In order to improve the yield of human leukocytes, we have treated MS5 stromal cells with mitomycin C prior to culturing them with CD34+ cells. We reason this will stop stromal cells from expanding and competing for nutrients against progenitors. After 14 days, we compared cultures with treated MS5 stromal cells to untreated ones and observed an increase of

CD45+ leukocytes by 1.5 fold in post-mitomycin C treatment cultures (Fig 3.3A). Furthermore, all six cell-subsets increased in numbers. In particular, pDCs increased by 1.5-fold, CD1c cDCs by 2-fold, CD141 cDCs by 4-fold, and B cells by 3-fold. There was a less than 10% increase of granulocytes and monocytes (Fig 3.3A). Mitomycin C treatment did not affect the overall cell composition (Fig 3.3B). Therefore, because mitomycin C treatment increases the overall cellular yield without drastically affecting the relative cell composition, stromal cells have been pretreated with mitomycin C for all subsequent experiments.

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Figure 3.1 Isolation of human CD34+ Hematopoietic stem and progenitor cells. Flow cytometry plots show gating strategy for sorting CD34+ HSPCs and post-sort purity.

43

95 45.2 62.8 Post-sort 91.8 purity 96.3 99 SSC SSC SSC SSC SSC

FSC pulse width Trig. FSC CD45 Lin CD34 CD34

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Figure 3.2 Stromal cell culture and human DC development. (A) Plots show representative flow cytometry analysis of human CD45+ composition from each culture at day 14.

Granulocytes, brown; monocytes, orange; pDCs, green; CD1c cDCs, blue; CD141 cDCs, red; B cells and NK cells, purple. (B) Bar plots show the absolute number of specified cell types at day

14 from cultures without stroma (white), with MS5 (light gray), S17 (gray) or OP9 (black) stromal cells. Results are from three independent experiments and error bars are SEM. (C) Pie charts indicate the relative representation of each DC subset in each group.

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A C Relative DC Composition No stroma PBMC

MS5 No Stroma

MS5 S17

S17

OP9 OP9 SSC SSC CD14 CD141 CD123 CD66b CLEC9a CD1c CD303 CD19

B CD45 Granulocytes Monocytes CD1c cDCs CD141 cDCs pDCs B cells 2.5×105 4×103 5×104 6×104 1.2×104 2.0×104 4×104 No stroma 5 4 8.0×103 2.0×10 3×103 4×10 1.5×104 3×104 MS5 4×104 1.5×103 1.5×105 3×104 3 4 4 S17 2×10 3 1.0×10 2×10 5 4 1.0×10

Cell # 1.0×10 2×10 2×104 OP9 1×103 2 5.0×103 1×104 5.0×104 1×104 5.0×10 0.0 0 0 0 0.0 0.0 0

MS5 S17 OP9 MS5 S17 OP9 MS5 S17 OP9 MS5 S17 OP9 MS5 S17 OP9 MS5 S17 OP9 MS5 S17 OP9 No stroma No stroma No stroma No stroma No stroma No stroma No stroma

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Figure 3.3 Stromal cell treatment with Mitomycin C enhances cellular output. (A) Bar plots show absolute counts of human CD45+ cells from 1 × 103 CD34+ HSPC cocultures with mitomycin C untreated (gray) or treated (black) MS5 stromal cells. (B) Bars show the relative frequencies of (from the bottom) granulocytes (brown), monocytes (orange), pDCs (green),

CD1c cDCs (blue), CD141 cDCs (red) and B cells (purple). Results show the mean averages from three independent experiments and error bars are SEM.

47

A CD45 Granulocytes Monocytes pDCs CD1c cDC CD141 cDC B cells B 100%

) 400 6 150 25 150 6 4 3 Untreated 80% 300 20 3 60% 4 100 100 4 Treated 15 200 2 40% 10 2 50 50 2 20% 100 1 Cell#(x10 5

% of total hCD45 0% ------Mitomycin C - +

48

In vitro DCs are phenotypically and functionally equivalent to in vivo DCs

In order to confirm that these cultured-derived DCs are physiologically equivalent to that of the in vivo, we compared the expression levels of DC signature genes. Both in vitro and in vivo cDCs and monocytes expressed CD11c, HLA-DR and CD86, with much lower expression on pDCs (Fig 3.4A). The lack of CD80 and CD83 on all DCs reflects their immature state. pDC’s specific expression of CD123 is also markedly high on culture-derived pDCs as their in vivo counterparts. In vitro CD1c cDCs and monocytes were also high in CX3CR1 and SIRPa expression but not on CD141 cDCs or pDCs either in vitro or in vivo. CLEC9a, a receptor that captures necrotic cells, is specifically expressed on both in vitro and in vivo CD141 cDCs but not on other cells. Of note, culture-derived cDCs did not express CD1a or DC-SIGN (Fig 3.4A), which are markers that identify monocyte-derived DCs in vitro, suggesting that MS5 culture supports steady state DC lineage development.

In addition to their surface markers, we also analyzed the expression of DC subset specific signature genes, which include transcription factors and genes associated with DC functions (Fig 3.4B). Transcription factor IRF8 was expressed on pDCs and CD141 cDCs alone, and E2-2, an essential transcription factor for pDC development, was also specifically detected in in vitro derived pDCs. Chemokine receptor XCR1 was uniquely expressed on both in vivo and in vitro CD141 cDCs and DC specific cytokine receptor FLT3 was also expressed on all DCs but not on monocytes (Fig 3.4B), further confirming that cultured DCs have not trans-differentiated from monocytes. Another distinction between monocytes and different DC subsets is their capacity to initiate discrete immune responses in response to different stimuli. This has been shown to depend on their differential expression of specific toll-like receptors (TLR). The relative expression pattern of TLR2, 3, 7 and 9 in cultured cells were identical to that of their in

49 vivo counterparts. TLR2, a lipoteichoic acid, LPS and peptidoglycan receptor, was highly expressed on monocytes followed by CD1c cDCs. TLR3, a double stranded RNA receptor, was highly expressed by CD141 cDCs followed by CD1c cDCs. TLR7 and 9, an intracellular bacterial DNA receptor, was detected in pDCs alone.

In order to show that their unique TLR expression pattern also reflects their functional specialization, we measured the cytokine secretion levels upon TLR stimulation with their cognate ligands (Fig 3.5). After LPS stimulation (a TLR2 agonist), monocytes produced the highest amount of IL-6. After poly I:C stimulation (a TLR3 agonist), CD141 cDCs produced the highest level of IFNa and IL-12, and none of the cells secreted IL-6. The TLR7/9 agonist CpG induced massive production of IFNa by pDCs alone, and very little to none of IL-12 or IL-6 by any of the other cell types. In conclusion, culture-derived monocytes and DCs are equivalent to their in vivo counterpart in phenotype, transcription and function.

50

Figure 3.4 In vitro DCs are phenotypically and transcriptionally identical to in vivo DCs.

(A) Histograms show cell surface markers of human monocytes and DCs isolated from blood

(top rows) and MS5+FLT3L cultures (bottom rows). (B) Bar plots show the relative mRNA expression levels of indicated genes from three independent experiments and error bars are SEM.

51

A Isotype CD11c HLA-DR CD80 CD83 CD86 DC-SIGN

Monocytes pDCs

In vivoIn CD1c cDCs CD141 cDCs

Monocytes pDCs

In vitro In CD1c cDCs CD141 cDCs Isotype CD123 CX3CR1 CLEC9a SIRPa CD1a

Monocytes pDCs

In vivoIn CD1c cDCs CD141 cDCs

Monocytes pDCs

In vitro In CD1c cDCs CD141 cDCs

B IRF8 E2-2 XCR1 FLT3 TLR2 TLR3 TLR7 TLR9 100000 40000 4000 10000 600 15 15000 15000

80000 30000 3000 8000 400 10 10000 10000 60000 6000 20000 2000 40000 4000 In vivoIn 200 5 5000 5000 20000 10000 1000 2000 0 0 0 0 0 0 0 0

150000 25000 4000 8000 3000 25 15000 20000

20000 3000 6000 20 15000 100000 2000 10000 15000 15 2000 4000 10000

In vitro In 10000 10 RelativeGAPDH to 50000 1000 5000 5000 1000 2000 5 5000 0 0 0 0 0 0 0 0

52

Figure 3.5 In vitro generated DCs are functionally active. Cord blood CD34+ derived DC subsets were cultured for 14 d, purified by FACs, and exposed to the indicated TLR stimuli.

Graphs indicate concentration of IFNa, IL-12p70 and IL-6 in the supernatant measured by

ELISA after 48h. n (number of donors) = 3. Error bars indicate SEM. Orange, Monocytes; green, pDCs; blue, CD1c cDCs; red, CD141 cDC.

53

10µg/ml 50µg/ml 1µM Medium LPS Poly(I:C) CpG 30 30 30 15 X1,000 IFNα 20 20 20 10

(pg/ml) 10 10 10 5

0 0 0 0 60 60 60 60

IL-12p70 40 40 40 40

(pg/ml) 20 20 20 20

0 0 0 0 3000 3000 3000 3000 2400 2400 2400 2400 IL-6 1800 1800 1800 1800 (pg/ml) 1200 1200 1200 1200 600 600 600 600 0 0 0 0

54

MS5 and OP9 together with FLT3L, SCF and GM-CSF provide optimal condition for DC development

Although MS5 stromal cells supported the development of all DC subsets, as well as three other cell types, we observed a slight bias toward CD1c cDCs (Fig 3.2C), with comparatively fewer pDCs and CD141 cDCs. In order to correct this bias, we added titrating numbers of OP9 cells into MS5 culture. We did this because OP9 has shown a preference for supporting CD141 cDCs development (Fig 3.2C). We then cultured CD34+ progenitor cells in these new culture conditions and measured the cell output for multiple lineages as well as for each specific DC subset. After 14 days, we observed that adding OP9 cells specifically increased the number of CD141 cDCs cells by 10-fold without drastically changing the cell output for other cell types (Fig 3.6A). Additionally, we found that a ratio of 1 OP9 -to- 6 MS5 generated relative composition of three DC subsets that most resembles their in vivo counterparts (Fig

3.6B).

Finally, we optimized the output of multiple lineages by adding stem cell factor (SCF) and granulocyte-macrophage colony stimulating factor (GM-CSF) to the MS5+OP9 stromal cell culture (Fig 3.7). Compared to FLT3L alone, adding these two cytokines induced a significant increase in the total output of CD45+ leukocytes by 6-fold. In particular, SCF and GM-CSF increased the yield of granulocytes by 23-fold, monocytes by 7-fold, CD1c cDCs by 6-fold,

CD141 cDCs by 11-fold, pDCs by 1.5-fold and B cells by 5-fold. Considering the robust output of both myeloid and lymphoid cells, including all three DC subsets, we conclude that mitomycin

C treated MS5+OP9 at a 6:1 ratio in combination with FLT3L, SCF and GM-CSF (MP+FSG herein) is optimal for simultaneous multilineage differentiation from CD34+ progenitors.

55

Therefore, we conclude that MS5 stromal cells, together with OP9 (1 OP9 : 6 MS5) and cytokines (100ng FLT3L/ml, 20ng SCF/ml and 10ng GM-CSF/ml), support the differentiation of all three human DC subsets as well as granulocytes, monocytes and B lymphocytes. Clonal cultures described in Chapter V were performed using MP+FSG, however, for Chapter IV we have alternatively used MS5 plus FLT3L, SCF and GM-CSF (MS5+FSG), instead.

56

Figure 3.6 OP9 stromal cells specifically increase CD141 cDC numbers. (A) Dots indicate the cell number output for the specified population (y-axis) from each culture combination (y- axis) from 1x103 CD34+ cells. (B) Bar graphs show the relative composition of each DC subset from each culture combination. Results are mean averages from three independent experiments.

57

106 CD45 A B 100 pDCs Granulocytes 105 80 CD141 cDCs Monocytes 4 60 CD1c cDCs 10 CD1c cDCs

Cell # 40 3 CD141 cDCs 10 20 pDCs 102 % of total DCs 0 B cells 0 0 1:6 1:6 1:30 1:30 1:150 1:150 PBMC OP9:MS5 ratio OP9:MS5 ratio

58

Figure 3.7 FLT3L, SCF and GM-CSF increase cellular yield. Bars show the absolute number of cells from 2 × 103 HSPCs and mitomycin C pretreated MS5 cocultures in FLT3L alone (gray) or FLT3L, SCF and GM-CSF (gray). Bars are mean averages are from three independent experiments and error bars are SEM.

59

CD45 Granulocytes Monocytes CD1c cDCs CD141 cDCs pDCs B cells 6×105 8×104 2.5×105 8×104 2.5×104 1.5×104 2.5×104 2.0×105 4 2.0×104 6×104 6×104 2.0×10 4×105 1.0×104 1.5×105 1.5×104 1.5×104 4×104 4×104 5 4 4 5 1.0×10 1.0×10 3 1.0×10 Cell # 2×10 5.0×10 FLT3L 4 4 2×10 4 2×10 3 3 5.0×10 5.0×10 5.0×10 FSG 0 0 0.0 0 0.0 0.0 0.0 F F F F F F F FSG

60

3. Discussion

This culture system allows us to determine the DC potential of candidate progenitors in the context of both myeloid (i.e. granulocytes and monocytes) and lymphoid (i.e. B cells) lineages. Based on the phenotypic, transcriptional and functional characterization of in vitro

DCs, we have proved that cultured DCs are equivalent to that of their in vivo counterparts and not a result of trans-differentiation from monocytes. The development of steady-state DCs allows us to investigate the DC potential in relation to the monocyte lineage. Finally, optimizing the culture by treating stromal cells with mitomycin C and adding SCF and GM-CSF provides an environment that promotes maximal cell output from early progenitors, a promotion that could be due to increasing proliferation and/or maintaining survival. This culture (MP+FSG) promotes maximal cell output from candidate progenitors with limited proliferation capacity, thereby allowing one to determine developmental potential of human CD34+ hematopoietic progenitors.

61

Chapter IV. Identification of human committed DC progenitors and precursors

Results in this chapter have been published and can be found in: Lee J, Breton G, Oliveira TY, Zhou YJ, Aljoufi A, Puhr S, Cameron MJ, Sékaly RP, Nussenzweig MC, Liu K. Restricted dendritic cell and monocyte progenitors in human cord blood and bone marrow. J Exp Med. 2015 Mar 9;212(3):385-99. doi: 10.1084/jem.20141442.

Breton G, Lee J, Zhou YJ, Schreiber JJ, Keler T, Puhr S, Anandasabapathy N, Schlesinger S, Caskey M, Liu K, Nussenzweig MC.Circulating precursors of human CD1c+ and CD141+ dendritic cells. J Exp Med. 2015 Mar 9;212(3):401-13. doi: 10.1084/jem.20141441.

62

1. Introduction

In mouse, DCs develop from hematopoietic stem cells in the bone marrow through a sequence of progenitors that have increasing restriction to terminal DCs. Identification of these progenitors was made possible by understanding the role of several cytokines and their corresponding receptors in DC development.

Flt3 ligand and its receptor (CD135) are essential for DC development125. In the periphery, CD135 is strictly expressed on DCs and administering Flt3L in vivo dramatically expands DC numbers in lymphoid and non-lymphoid organs123. In mice with genetically ablated

Flt3L, peripheral DCs are significantly decreased and genetic deletion of CD135 also showed similar deficiencies in DC development139. Based on the dependence on Flt3 signaling for DC development, DC progenitors have been searched on the criterion that they should also express

CD135 on their surface.

However, when bone marrow CD135+ cells have been tested for their potential, not only did they produce DCs, but also lymphocytes, monocytes and polymorphonuclear cells140, 141.

Thus, additional cytokine receptors were needed to further fractionate the CD135+ population and identify specific cells with restricted DCs potential.

Two receptors that have been essential in the search for DC progenitors are SCF receptor or CD117, and M-CSF receptor or CD115. SCF receptor is mainly expressed by early bone marrow progenitors and is lost as cells differentiate13, 142. CD117 is essential for maintaining hematopoietic progenitors, and homozygous knockout mice are embryonic lethal due to anemic conditions. This shows that the expression of CD117 and the capacity to respond to SCF is critical for early progenitor cell survival. M-CSF, on the other hand, is a cytokine essential for monocyte development and dispensable for DCs. Mice that lack its receptor, CD115, show

63 symptoms of osteoporosis due to the absence of osteoclasts, as well as a deficiency in monocytes, macrophages and skin Langerhans cells, but not DCs143.

The analysis of CD135, CD115 and CD117 expression in non-terminal (i.e. Lin-), non-

HSC (i.e. sca-1-) bone marrow cells led to the identification of two DC progenitors with different degrees of developmental restriction. One population with CD135+CD115+CD117hi phenotype produced only macrophages and DCs, and was termed macrophage-DC progenitor (MDP). The second cell population with CD135+CD115+CD117lo phenotype only produced all three DC subsets, and was named the common DC progenitor (CDP). In vivo experiments also showed that

CDPs arise downstream from MDPs, therefore identifying for the first time intermediate DC progenitors in mouse.

In a separate investigation, spleen and bone marrow cells with Lin-

CD135+CD11c+MHCII- phenotype have been identified and when tested for their developmental potential in vitro and in vivo, they could only produce cDCs but not pDCs, suggesting an additional step of lineage restriction prior to cDC differentiation110. Furthermore, CDPs in vitro or in vivo showed that they were able to give rise to the newly identified cDC precursors, or pre- cDCs. In sum, the identification of MDPs, CDPs and pre-cDCs by tracing the expression of

CD135 revealed the sequential process of DC differentiation in the mouse.

In humans, DC-restricted progenitors have not yet been identified. However, human DCs share many similarities with mouse DCs. Transcriptional profile analysis of human DC subsets and comparison to their murine counterparts reveals high degree of similarity between the two species. For example, CD135 expression is also specific on terminal DCs but not on other cell types103. FLT3L injection of human subjects dramatically increased the number of DCs in circulation124. Furthermore, functional properties (e.g. T cell activation, TLR expression

64 signature, cytokine secretion) of each DC subset are also highly conserved between humans and mice92. This suggests that human DC development may also follow similar developmental stages.

Hence, we hypothesize that restricted DC progenitors can be identified by analyzing and evaluating the potential of CD135+ cells.

In this chapter, we report four consecutive DC progenitors in human cord blood and bone marrow with increasing degree of commitment toward DCs. Using our culture system and analysis of cytokine receptor expression on CD34+ progenitor and CD34- non-progenitor cells, we have been able to identify the sequential human DC progenitors, from a granulocyte- monocyte-DC progenitor (GMDP), which develops into a more restricted human monocyte-DC progenitor (MDP), which then produces monocytes and human CDPs, which generate pDCs and pre-cDCs, which finally give rise to cDCs. We also determined the anatomical location where each of these progenitors can be found in humans. Lastly, we discussed the possibility to resolve the myeloid or lymphoid origin of DCs by evaluating how myeloid and lymphoid progenitors contribute to the GMDP-MDP-CDP trajectory.

65

2. Results

Single cell culture reveals developmental heterogeneity within GMPs

DCs have been traditionally considered to be myeloid cells and closely related to monocytes. There are currently two identified myeloid progenitors in the human hematopoietic tree: the CD34+CD38+CD10-CD135+CD45RA- common myeloid progenitor (CMP) and the

CD34+CD38+CD10-CD135+CD45RA+ granulocyte-macrophage progenitor (GMP). Both of these have granulocyte, monocyte and DC activity in vitro and in vivo. However, CMP can have megaerythrokaryocyte potential, while GMPs are downstream of CMPs and lack megaerythrokaryocyte potential. Additionally, transcriptional profile of GMP population also shows that it expresses transcription factors essential for DC development144, suggesting that

GMPs may contain cells with DC potential.

Therefore, in order to determine the clonal DC activity in GMPs, we have isolated individual cells from cord blood and cultured them together with MS5+FSG (Fig 4.1A). After two weeks of culture we determined their clonal developmental potential based on the cell type generated. Results show that the clonal efficiency is high (65%), indicating that this culture is appropriate for single cells to proliferate and differentiate. Out of 144 GMP clones, 93 have generated identifiable CD45+ leukocytes.

Surprisingly, there is a remarkable heterogeneity within GMPs as determined by their lineage output (Fig 4.1B). The majority of the 93 clones (84/93) produced myeloid and/or DCs alone and lacked lymphoid potential. Only a small fraction (6/93) produced lymphoid and myeloid cells together, and even fewer clones (3/93) remained undifferentiated. Among the 84 myeloid clones, however, only 6 (6.5%) simultaneously produced granulocytes, monocytes and

DCs. Twenty five clones (26.9%) were bipotent. Of these, 11 clones (11.8%) produced

66 monocytes and DCs, 13 clones (14.0%) generated granulocytes and monocytes, and very rare clones (1.1%) produced granulocytes and DCs. A high percentage of clones (53/93, 57.0%) showed uni-lineage potential, i.e., commitment; these include 25 producing only granulocytes,

11 only monocytes and 16 only DCs. Therefore, our culture system allows testing of the development of granulocytes, monocytes, DCs and lymphocytes from single progenitor cells. In addition, this test reveals that phenotypically homogeneous GMPs are heterogeneous in their clonal potential, especially in DC producing clones.

We found 17.2% of the clones within GMPs were committed to DCs alone (Fig 4.1B), resembling the mouse CDPs. We also found that 11.8% of clones with monocyte and DC potential, resembling the mouse MDPs, as well as multipotent clones (6.5%) with granulocyte, monocyte and DC potential. This indicates that GMPs is comprised of a mix of progenitor cells that differ in their developmental capacity and lineage commitment, and can thus be further identified and isolated. Therefore, further dissection of GMPs using additional surface receptors that can distinguish pure populations with homogeneous potential may allow further purification of lineage-restricted progenitors.

67

Figure 4.1 GMPs are clonally and phenotypically heterogeneous. (A) Flow cytometry plots show gating strategy for sorting cord blood derived human GMPs. (B) Single GMP cells were directly sorted into 96 well plates and culture in MS5+FSG. Bar graphs show the cumulative number of wells giving rise to the specified potential out of the total number of positive clones.

G, granulocyte; M, monocyte; DC, pDCs and/or cDCs; L, lymphoid (B and/or NK cells); Undiff, undifferentiated; +, “and”; /, “or”. (C) low cytometry plots show exhaustive sepa- ration of

CD34+ cord blood cells into six populations, HSCs/MPPs, MLPs, B and NK progenitors

(B/NK), CMPs, megakaryocytic and erythroid progenitors (MEP), and GMPs (a), and expression of CD115, CD116, CD135, and CD123 on each of the cord blood CD34+ populations in a (b).

68

A B % of Total Positive Clones 58.8 17.1 0 0 23.25 25 46.5 50 69.75 75 10093 2.96 GMP 55.2 G 25 M 11 95.4 4.27 CD38 CD10

3.59 CD135 DC 16 CD34 CD45RA CD45RA DC + M 11 C DC + G 1 G + M 13 G + M + DC 6 G/M/DC + L 6 CD116 CD116 CD115 CD115 CD123 Undiff 3 CD45RA

69

Differential expression of cytokine receptors allows fractionation of GMPs into three developmentally restricted progenitors

In order to identify lineage-restricted progenitors we analyzed the differential expression of DC-, monocyte- and granulocyte-associated cytokine receptors within GMPs. Since GM-CSF,

M-CSF and IL-386 have been shown to affect granulocyte, monocyte and DC development, we used their respective receptors (i.e. CD116, CD115 and CD123) to fractionate CD45RA+ GMPs.

Flow cytometry analysis revealed five distinct populations (Fig 4.1C, 4.2A). One population expresses high CD123 (i.e. CD123hi) and the rest CD123int cells can be divided into four additional populations based on the differential expression of CD116 and CD115:

CD116+CD115- (i.e. CD116+), CD116+CD115+ (i.e. DP), CD116-CD115+ (i.e. CD115+) and

CD116-CD115- (i.e. DN). In order to determine their developmental capacity, we have isolated

200 cells from each fraction and put them on our MS5+FSG culture (Fig 4.2B). After 7 days of culture we have analyzed the generation of granulocytes, monocytes, CD1c cDCs, CD141 cDCs, pDCs and lymphocytes (i.e. B and NK cells, data not shown).

In order to identify DC progenitor candidates, one criterion must be met: DC progenitors should be able to produce all three DC subsets. Out of the five populations tested, two failed this criterion and were excluded as potential DC progenitors. CD116+ and DP cells, although they produced CD1c cDCs, did not produce other DC cell types and were disregarded as potential precursors.

On the other hand, CD123hi cells produced all three DC subsets and no other cell-types, resembling mouse CDPs (Fig 4.2B). They constitute 0.20 + 0.13% of CD34+ cells and we named them human CDPs (CDPs herein). CD115+ cells generated all three DC types as well, together with monocytes and very few, if any, granulocytes, resembling murine MDPs. CD115+ cells

70 represent 0.31 + 0.18% of CD34+ cells and can be identified as CD45RA+CD123intCD115+ due to the negligible frequency of DP cells (~0.04% of CD34+ cells). Consequently, we named them

MDPs. Finally, the DN fraction, which represents 4.16 + 0.98% of CD34+, generates granulocytes, monocytes and all three DCs and hence, we named these Granulocyte, Monocyte and DC progenitor (GMDP).

When we analyzed their proliferative capacity, GMDPs and MDPs produced higher yield of cells than CDPs, which reflects their less differentiated status and more proliferative activity

(Fig 4.2C). In addition, upon culture of 4 days, MDP and GMDP expanded and resulted in increased number of CD34+ cells (Fig 4.2D). In contrast, CDPs rapidly lost the expression of

CD34, a marker for early progenitors (Fig 4.2D and data not shown).

We also determined their megaerythrokaryocyte and granulocyte potential by culturing

500 cells of each population in methylcellulose culture assays for 14 days (Fig 4.2E). As expected, CDPs did not produce any detectable colonies since this assay does not support DC development. This indicates that CDPs lack megaerythrokaryocyte, granulocyte and macrophage potential. MDPs, on the other hand, were able to generate monocyte-producing colonies and very few granulocyte colonies. GMDPs generated both monocyte- and granulocyte-producing colonies as well as colonies with mixed cell types, suggesting the presence of oligopotent clones.

MDPs and GMDPs produced comparable colony counts, similar to their comparable clonal efficiency in MS5+FSG cultures (Fig 4.2E). Of note, none of the three populations tested had megaerythrokaryocyte potential when compared to the CMPs.

Therefore, we conclude that human GMPs contain phenotypically and functionally distinct sub-populations that show increasing commitment to DC lineage. Cells with

CD34+CD38+CD10-CD45RA+CD123hi phenotype generated only the three types of DCs, and

71 therefore we called these CDPs. Cells with CD34+CD38+CD10-CD45RA+CD123int CD115+ phenotype generated monocytes and DCs, and thus we called these MDPs. Finally, cells with

CD34+CD38+CD10-CD45RA+CD123intCD115- phenotype produced granulocytes, monocytes and DCs, and thus we called these GMDPs.

72

Figure 4.2 Identification of progressively restricted DC progenitors in GMPs. (A) Flow cytometry plots show gating of cord blood GMP cells and further separation into five separate populations based on CD123, CD115, and CD116 expression: CD123hiCD115 (CD123hi),

CD123intCD115+CD116- (CD115+), CD123intCD115+CD116+ (DP), CD123intCD115-CD116+

(CD116+), and CD123intCD115-CD116- (DN). (B) Differentiation potential of 200 purified cells from each of the five populations indicated in (A) in MP+FSG culture harvested after 7 d. Flow cytometry plots show CD45+CD56-CD19- cells. (C) Graph indicates output/input ratio of total number of CD45+ cells obtained from each of the five populations sorted in (A). Bars and error bars are means and SEM, respectively, from three independent experiments. (D) Graph shows the fold-increase of CD34+ cells after 4 days of culture. Bars are means and error bars are SEM from three independent experiments. (E) Graph indicates the differentiation potential of CDPs,

MDPs, GMDPs, and CMPs in methylcellulose colony formation assays in vitro. Colonies were enumerated at 14 d after culture. BFU-E, burst-forming unit erythroid; GEMM, granulocyte, erythrocyte, macrophage, megakaryocyte; GM, granulocyte and macrophage; G, granulocyte; M, macrophage. Bars are means and error bars are SEM from three independent experiments.

73

A CD123 hi CD116+ DP C GMP 20.1 Output cells 100,000

DN CD115+ CD10

CD123 10,000 CD116 CD116

96.2 CD123 CD45RA CD45RA CD115 CD115 B Granulocyte CD141 cDC Monocyte CD1c cDC pDC 1,000 +

hi DP CDP MDP D GMDPCD116

CD123 6 +

cells 4 +

CD115 2 Foldincrease of CD34

0

DN CDP MDP GMDP

E

+ No. of CFU 150 BFU-E CD116 CFU-GEMM 100 CFU-GM

50 CFU-G DP DP CFU-M SSC CD123 CD141 CD14 0 CD66b CLEC9a CD1c CD303 CDP MDP CMP GMDP

74

Cytokine receptor expressions analysis in CD34- cells allows the identification of pre-cDCs

The identification of GMDPs, MDPs and CDPs in cord blood shows that human DC development resembles that of the mouse. Consequently, we speculate that humans also have a downstream precursor between CDPs and cDCs.

Seeing that CDPs lose CD34 expression to produce cDCs, we reasoned that the human equivalent of pre-cDCs should be present within the CD34- portion. Thus, in order to search for the precursor, we decided to look into the non-terminally differentiated (i.e. Lin(B/T/NK/ granulocytes/monocytes/DCs)-) and non-progenitor (i.e. CD34-) cells (Fig 4.3A). We subdivided the Lin-CD34- fraction using differential expression of cytokine receptors CD117 and CD135, which are expressed on terminally differentiated cDCs. We isolated CD117-, CD117+CD135-

(CD135-) and CD117+CD135+ (CD135+) and tested their DC potential on MS5+FSG cultures

(Fig 4.3B). After 7 days of culture, we compared their output with CD34+ multipotent progenitor cells. As expected, CD34+ progenitors were able to generate mature cells of multiple lineages

(monocytes, DCs and lymphocytes). Among CD34- cells, both CD117- and CD117+CD135- cells failed to produce any identifiable cells, and only CD117+CD135+ cells produced monocytes and cDCs but not pDCs. This suggests that CD117+CD135+ cells contain pre-cDCs and can further be isolated. For this purpose, we subdivided CD117+CD135+ cells according to the differential expression of CD116 and CD115, as well as CD45RA (Fig 4.3C). We isolated CD116-,

CD116+CD45RA- (CD45RA-), CD116+CD45RA+CD115+ (CD115+) and CD116+

CD45RA+CD115- (CD115-) cells and cultured them in MS5+FSG for 7 days. Cell output analysis shows that CD116- and CD45RA- fractions lacked both monocyte and DC activity (Fig

4.3D). Interestingly, CD115+ cells produced a high number of monocytes and a few CD1c cDCs, which resemble the mouse monocyte precursor cMOP. CD115- cells produced only cDCs but

75 failed to produce monocytes or pDCs (Fig 4.3E), and resemble the mouse pre-cDC. These cells were present in cord blood and in the peripheral blood (Fig 4.6A).

Further phenotypic characterization of human pre-cDCs in cord blood shows that they have low expression of CD11c and CD123 but are positive for HLA-DR (Fig 4.3F) at comparable levels with fully differentiated DCs. In conclusion, we have identified the human pre-cDCs and they are characterized by Lin-CD34-CD117+CD135+CD116+CD45RA+CD115-.

We next sought to determine pre-cDC’s proliferative capacity. We isolated pre-cDCs from cord blood and peripheral blood and incubated them in MS5+FSG culture. After 7 days, we compared their cell output with CD34+ hematopoietic stem and progenitor cells (HSPCs) as well as with terminally differentiated CD1c cDCs and CD141 cDCs isolated from peripheral blood

(Fig 4.4A). As expected, CD34+ cells were highly proliferative and expanded approximately 156 fold. In contrast, pre-cDCs, either from cord blood or peripheral blood, had significantly lower proliferative capacity (approximately 8- and 6-fold, respectively). Pre-cDCs were, however, still able to significantly expand more than fully differentiated DCs. We next determined whether the increase in cell number was due to pre-cDCs dividing before differentiating or fully differentiated cDCs dividing after differentiation. For this purpose, we stained cord blood derived pre-cDCs with CFSE and followed their differentiation and division for 2 and 7 days.

Pre-cDCs had generated CD141 cDCs and CD1c cDCs as early as 2 days of culture and increased in numbers by 7 days (Fig 4.4B), while CD34+ HSPCs produced cDCs and monocytes only by day 7. When tracing CFSE dilution at day 2, pre-cDCs produced terminal cDCs expressing equal levels of CFSE, suggesting that division and differentiation happens simultaneously (Fig 4.4C). However, at day 7, precursor-derived cDCs had lower levels of

CFSE than remaining pre-cDCs, which indicates that differentiated cDCs can undergo further

76 divisions. Confirming this notion, CD141 cDCs and CD1c cDCs purified from blood, were able to undergo several divisions in MS5+FSG after 7 days of culture, indicated by the dilution of

CFSE (Fig 4.4C). Therefore, this shows that human pre-cDCs, either in cord blood or in peripheral blood, differentiate into cDCs as they divide and that terminally differentiated cDCs can also continue to proliferate.

77

Figure 4.3 Identification and developmental potential of cDC precursors. (A) Flow cytometry plots show gating of CD45+CD3-CD19-CD56-CD14-CD66b-CD1c-CD141-CD303- cells in human cord blood can be divided into four populations based on CD117, CD34, and

CD135: CD34+CD117+ (CD34+), CD34-CD117- (CD117-), CD34-CD117+CD135- (CD135-), and

CD34-CD117+CD135+ (CD135+). Numbers indicate the frequency of respective gates. (B)

Differentiation potential of 1,000 purified cells from each of the 4 populations indicated in (A) in

MP+FSG cultures for 7 d. Flow cytometry plots show gated live human CD45+ cells. (C) Flow cytometry plots show CD135+ cells indicated as in (A) can be further separated into 4 populations based on CD116, CD115, and CD45RA: CD135+CD116- (CD116-),

CD135+CD116+CD45RA- (CD45RA-), CD135+CD116+CD45RA+CD115+ (CD115+), and

CD135+CD116+CD45RA+CD115- (CD115-). (D and E) Differentiation potential of 100 purified cells from each of 4 populations indicated in (C) in MP+FSG cultures for 7 d. (D) Flow cytometry plots of CD45+ cells gated as in (C). (E) Graphs show mean output of pDC, CD1c cDC, CD141 cDC and monocytes from each population from three independent experiments. (F)

Histograms show expression of CD11c, HLA-DR, CD123, CD135, CD117, CD45RA, CD116, and CD115 on indicated cell-type.

78

A

95.5 0.0251

97 35.3 SSC SSC SSC 81.4 CD45RA 99.4 98.6 0.323 FSC FSC CD45 CD3/19/56/14 CD66b/303 CD141 CD1c Trigger pulse W. pulse W. Trigger

CD135- C CD116- CD34+ 64.3 19.3 96.1 2.96 CD45RA- 0.887 CD135+ 12.8 CD115+ CD117- 14

CD115- SSC SSC CD115 CD115 CD34 87 9.95 64 CD117 CD135 CD116 CD45RA

B CD141 cDC Mono CD1c D CD141 cDC Mono CD1c E 2,100 cDC pDC cDC pDC pDC -

+ CD141 cDC 1,400 B cells B cells CD1c cDC Monocyte 700 CD34 CD116 -

- 2,100 -

1,400

700 CD117

CD45RA - 2,100 + - 1,400

700 CD115 CD135

# of output#of cells /100 cells input - 2,100 - + 1,400

700 CD115 CD14 CD14 CD135 CD141 CD303 CD303 CD141 - CLEC9a CD1c CD19 CLEC9a CD1c CD19

F Unstained CD11c HLA-DR CD123 CD135 CD117 CD45RA CD116 CD115

pDC CD1c cDC CD141 cDC preDC

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Figure 4.4 Proliferative capacity of pre-cDCs. (A) Expansion of purified pre-cDCs or CD34+

HSPCs from peripheral blood (PB) or cord blood (CB) in MP+FSG cultures for 7 d. Graph shows the mean fold change of live human CD45+ cells from 100 input cells from three independent experiments. *, P < 0.005, unpaired two-tailed Student’s t test. (B and C) CD34+

HSPCs and pre-cDCs were purified from CB, labeled with CFSE, and cultured in MP+FSG for 2 or 7 d; proliferation was assessed by flow cytometry. FACs plots show (B) gated CD45+ culture- derived cells, including CD34+ cells (purple), CD34-CD1c-CD141- cells (orange), CD141 cDCs

(red), and CD1c cDCs (blue) and (C) their CFSE dilution. Dotted lines mark last division by pre- cDCs. Plots are representative of three independent experiments.

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A C * Input Cells 1,000.0 155.8 CB CD34+ CB preDC PB CD1c+ DC PB CD141+ DC n.s.

100.0 7.9 CD141 cDC 6.4 * CD1c cDC 10.0 CD34- CD1c- CD141- 2days CD34+ 1.0 Foldexpansion 0.8 0.6

0.1 CD141 cDC + CD1c cDC In vitro output output vitro In CD34- CD1c- CD141- 7days CD34+

CB CD34 CB pre-cDCPB pre-cDC PB CD1c cDC CFSE PB CD141 cDC

B CB CD34+ cell culture output CB preDC culture output

CD141 cDC CD1c cDC CD141 cDC CD1c cDC

+ CD34- + CD34 CD34 Day 2 CD1c- CD141-

Day 7 SSC SSC CD14 CD14 CD141 CD141 CD34 CLEC9a CD1c CD34 CLEC9a CD1c

81

Physiological localization of early DC progenitors and precursors

DC development in mice is anatomically compartmentalized. All early progenitors prior to pre-cDCs are localized in the bone marrow and do not effectively migrate to the periphery110.

DC progenitors restrictively develop in the bone marrow cavity from hematopoietic stem cells until reaching pre-cDC stage. Only when they become pre-cDCs can they migrate through blood to the periphery, where they finish their differentiation into cDCs.

Therefore, we determined whether human progenitors and precursors also followed similar anatomical distribution. We first determined the presence of CD34+ DC progenitors in the adult human bone marrow as well as in the peripheral blood and tonsils. In bone marrow,

GMDPs comprised approximately 5.2% of total CD34+ cells (range of 1.8-7.4%), MDPs approximately 0.6% of total CD34+ cells (range of 0.1 – 1.9%) and CDPs represented approximately 1.9% of total CD34+ cells (range of 0.1 – 7.3%) as determined by flow cytometry analysis (Fig 4.5A). Furthermore, GMDP, MDP and CDP in the bone marrow demonstrated identical developmental potential as those identified in cord blood when cultured on MS5+FSG

(Fig 4.5B). Minor differences between adult bone marrow and cord blood DC progenitors were in GMDPs and CDPs. Bone marrow MDPs were significantly (P <0.05) biased toward producing more monocytes and toward producing less CD1c cDCs (P <0.001) than cord blood GMDP counterparts (Fig 4.5C). Bone marrow CDPs also showed a significant bias toward producing less pDCs than from cord blood derived CDPs.

On the other hand, very little CD34+ cells were detectable in peripheral blood or in tonsils

(Fig 4.5A) (approximately <1% of CD45+ cells), and human CDPs, MDPs and GMDPs were unidentifiable among CD34+ cells in the periphery. Therefore, early DC progenitors are restricted to the bone marrow and do not migrate to the periphery.

82

We next determined the migratory capacity of human pre-cDCs by looking for their presence in the bone marrow and peripheral lymphoid organs (i.e. peripheral blood and tonsils).

Similar to cord blood, human bone marrow contained both pre-cDCs as well as CD115+ monocyte precursors (Fig 4.6A). However, peripheral blood or tonsils lacked monocyte precursors and only contained pre-cDCs, suggesting that human DC precursors develop in the bone marrow and migrate to the periphery, while monocytes finish their development in the bone marrow and do not enter circulation. In order to verify their DC developmental potential, we isolated both pre-cDCs (Fig 4.6B) and CD115+ monocyte precursors (Fig 4.6C) from each tissue, and culture them in MS5+FSG. After 7 days of culture, pre-cDCs were able to strictly produce cDCs but not pDCs, and very few, if any, monocytes. Similarly, CD115+ monocyte precursors from cord blood and bone marrow generated primarily monocytes and a few CD1c cDCs. Pre-cDCs constitute 0.008% (range, 0.001-0.016%) of CD45+ cells in cord blood, 0.117%

(range, 0.056-0.188%) in BM, 0.001% (range, 0.001-0.002%) in peripheral blood and 0.001%

(range, 0.001-0.002%) in tonsil. Thus, this shows that, like in the mouse, human pre-cDCs develop in the bone marrow, enter circulation and reach peripheral lymphoid tissue where they could further differentiate into terminal cDCs.

83

Figure 4.5 Presence of early DC progenitors in bone marrow and peripheral tissues. (A)

Representative flow cytometry plots of gated CD45+Lin(CD3/19/56/14)-CD34+ cells show

GMDPs, MDPs, and CDPs in human BM (n=4), peripheral blood (PBMC; n=4), and tonsils

(n=4). (B) GMDPs, MDPs, and CDPs were purified from human BM, and 2,500 progenitors were cultured in MP+FSG for 7 d. Flow cytometry plots of gated live CD45+ cells show phenotype of output cells, including granulocytes (brown), CD141 cDCs (red), CD1c cDCs

(blue), monocytes (orange), and pDCs (green). Data represent three independent experiments. (c)

Graph shows output/input cell ratio in percentage of the indicated cells derived from BM (n=3) or cord blood (CB; n=3) cultures of GMDPs, MDPs, and CDPs in MP+FSG for 7 d. Statistical significance was determined using unpaired Student’s t test. *, P < 0.05; **, P < 0.001.

84

A CDP 7.2 33.2

BM 74.9 MDP GMDP 86.9 0.0334 3.12

PBMC 78.9 CD38

CD10 66 0.122 CD34 CD45RA 3.57 0

Tonsil

SSC 100 0 CD123 CD123 CD34 CD45RA CD115

B Mono CD66b- Gran CD141 pDC cDC GMDP GMDP CD1c cDC Mono CD66b- Gran Mono CD141 pDC cDC MDP MDP

CD1c cDC Mono CD66b- Gran pDC CD141 cDC CDP CDP CD14

SSC CD1c cDC CD141 CD123 CD66b CLEC9a CD1c CD303

C 100% * 80% ■ Granulocyte ** 60% ■ Monocyte ■ pDC 40% ■ CD1c cDC

Cell % Output ■ CD141 cDC 20% *

0% BM CB BM CB BM CB hGMDP hMDP hCDP

85

Figure 4.6 Presence of pre-cDCs in bone marrow and peripheral tissues. (A) Representative flow cytometry plots of gated Lin(CD3/19/56/14/66b)-DC(CD1c/141/303)-CD34- cells show gating strategy and composition of pre-cDCs (SSCloCD117+CD116+CD135+ CD45RA+CD115-, red) and pre-monocytes (SSCloCD117+ CD116+CD135+CD45RA+ CD115+, green) in human cord blood (CB), BM, peripheral blood (PB), and tonsil. Numbers indicate frequency of cells of parent gate (CB, n=5; BM, n=4, PB, n=5; Tonsil, n=3; n, number of donors). (B) Differentiation potential of purified pre-cDCs indicated in (A) in MP+FSG cultures for 5 d. Flow cytometry plots of gated live human CD45+ cells show culture output of CD141 cDC (red) and CD1c cDC

(blue). Representative results of four (CB), three (BM), three (PB), and two (tonsil) independent experiments are shown. (C) Differentiation potential of purified pre-monocytes from cord blood

(CB) and BM in MP+FSG cultures for 5 d. FACS plots show phenotype of gated live human

CD45+ culture-derived cells including monocytes (orange) and CD1c cDCs (blue).

Representative of three independent experiments are shown.

86

A 18.6 CB

74.4

21 BM

70.3

3.57 PB

96.4

0 Tonsil

SSC SSC SSC 72 CD115 CD115 CD117 CD116 CD135 CD45RA B

2.06 CB

3.58 7.06 22.4 33.3

0.79 BM

93.2 83.8

7.39 PB

10.3 Tonsil CD14 Live/dead CD141 CD123 CD45 CLEC9a CD1c CD303

C CB BM CD14 CD141 CLEC9a CD1c

87

Interdevelopmental relationship among early DC progenitors and precursors

The progressively increasing degree of commitment and loss of lineage potential suggests an inter-developmental relationship between GMDPs, MDPs, CDPs and pre-cDCs. In order to determine their progenitor-progeny relationship we first elucidated the relationship among

CD34+ progenitors. For this purpose, we isolated GMDPs from cord blood and transferred them into the bone cavity of conditioned NSG mice. After 7 days of transfer, we analyzed the phenotype of GMDP-derived progeny cells and found that MDPs and CDPs were generated in vivo (Fig 4.7A). This suggests that GMDPs are upstream of MDPs and CDPs.

We have further refined the inter-developmental relationship using MS5+FSG cultures by incubating each GMDP, MDP and CDP population for 1, 4 and 8 days. Flow cytometry analysis reveals that GMDPs remained as GMDPs in day 1 but soon produced MDPs on day 4 (Fig

4.7B), and later CDPs at day 8 of culture. Initial MDP cells quickly produced CDPs as early as in day 1, increased in numbers by day 4, and failed to generate GMDPs on any day throughout the experiment. By day 8 most MDPs have differentiated, which is marked by the loss of CD34 on the surface. Next, CDPs in culture retained their phenotype only on day 1 but quickly differentiated into CD34- cells by day 4 without prior production of either MDPs or GMDPs. We therefore conclude that GMDPs are upstream of MDPs, and that MDPs are upstream of CDPs.

CDPs are also the last progenitor stage within CD34+ cells able to produce DCs and, therefore, the most likely precursor for pre-cDCs.

In order to determine the developmental relationship of pre-cDCs with CDPs, we have isolated CDPs from cord blood and cultured them in MS5+FSG (Fig 4.7C). Consistent with previous experiments (Fig 4.2D), the majority of CDPs readily lost CD34 expression and very few pDCs and cDCs were detectable in culture on day 1. However, we observed an increase in

88 numbers of CD34- cells expressing CD45RA, CD117 and CD116, which resemble pre-cDCs phenotype. At day 2, all of CDPs have downregulated CD34 expression, pDCs and cDC numbers increased, while pre-cDC numbers remained unchanged. By day 4, CD34+ cells and pre-cDCs were not detectable, and only pDCs and cDCs were produced. In other words, CDPs differentiate first into pre-cDCs prior to becoming cDCs. Thus, we conclude that pre-cDCs are immediately downstream of CDPs and upstream of fully differentiated cDCs.

In sum, we have identified four different progenitors with (1) increasing commitment toward DCs and (2) direct progenitor-progeny relationship.

89

Figure 4.7 Interdevelopmental relationship among DC progenitors. (A) Differentiation of

GMDP in vivo. 10,000 GMDPs were purified from cord blood and adoptively transferred into the bone cavity of the preconditioned NSG mice. Seven days later, bone marrow cells were analyzed. Flow cytometry plots show phenotype of bone marrow cells from NSG mice receiving

PBS or GMDP (n=2, each). (B) Differentiation of GMDP, MDP and CDP in vitro. 200-400

GMDP, MDP and CDP cells were purified and cultured in MP+FSG. Cultures were analyzed on days 1, 4 and 8. Flow cytometry plots gated on live CD45+Lin(CD3/19/20/56/14)-CD66b-

DC(CD1c/141/303)-CD45RA+ cells show cell surface markers and frequency of CDP (red),

MDP (blue) and GMDP (grey). Results are representative of three independent experiments. (C)

Differentiation kinetics of CDPs purified from CB in MP+FSG cultures for 1, 2, or 4 d. FACS plots show culture output of live human CD45+ cells, including CD34+ cells (purple), pDC

(green), CD1c cDC (blue), CD141 cDCs (red), and pre-cDCs (orange). Representative of four independent experiments are shown. Graphs summarize composition of indicated populations among total CD45+ cells. Bars are mean values from four independent experiments, and error bars are SEM.

90

A 87.7 CDP

GMDP MDP SSC SSC

SSC 65.5 CD141 CD123 CD123 Lin CD303 CD1c CD34 CD45RA CD115 B 1d 4d 8d CDP GMDP GMDP GMDP MDP CDP MDP MDP

GMDP MDP CD123 CD123 CDP CD34 CD115 CDP CDP

CD123 CD123 GMDP MDP CD123 CD123 CD34 CD115 CD34 CD115 C % of total CD45+ 75

50 1d 25

0 75

50 2d 25

0 75

50 4d

25 CD141 SSC SSC CD116 CD116 CD117 CD117

CD45RA CD45RA 0 CD34 CD303 CD1c pDC hCDP hCDP hpre-cDC

CD1cDC/CD141

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3. Discussion

In this chapter, we have identified four new sequential myeloid progenitors with increasing commitment toward DCs. By using single cell cultures and cytokine receptor expression analysis, we identified three of these in the GMP fraction. These are the GMDPs

(granulocyte, monocyte and DC progenitor), MDPs (monocyte and DC progenitor) and CDPs

(common DC progenitor), and they can be distinguished by their differential expression of

CD123 and CD115. Similarly, by fractionating non-progenitor and non-terminally differentiated cells based on DC-associated cytokine receptor markers, we identified a committed precursor for cDCs that descend immediately downstream from CDPs. We have also anatomically located each step of DC development in adults. All four progenitors and precursors develop in the bone marrow, but only pre-cDCs migrate through blood to the periphery where they finish differentiation into cDCs.

Furthermore, identification of three progenitors within a myeloid progenitor (i.e. GMP) seems to support that DCs pertain to the myeloid lineage. GMPs have been extensively shown to develop from common myeloid progenitors (CMPs), which are capable of producing megaerythrokaryocytes and myeloid cells (including DCs) but not lymphocytes. Thus, our results suggest that DCs follow the myeloid track of development. This is very similar to what has been observed in mice (reference).

However, despite these data in mouse and humans, some conflict on DC origin and lineage remain unresolved. The main issue regarding DC development is that these cells can also derive from lymphoid progenitors, both in humans and in mice. Previous findings identifying and characterizing lymphoid progenitors in human cord blood and bone marrow have shown that

92

DCs can be generated from progenitors that produce B/NK and T cells but not granulocytes or megaerythrokaryocytes, which led to speculate a lymphoid DC ancestry.

One recent work has shown that single cells with CD34+CD38-CD45RA+CD10+ phenotype were able to generate only B cells and DCs, but not granulocytes, formally proving a common lymphoid ontogeny20. In fact, patients with B/NK/Monocyte/DC deficiency109, 145 who lack all three DC subsets, yet have relatively normal levels of granulocytes, also lack this progenitor population but with reduced GMP numbers in their bone marrow, further suggesting

DC association with the lymphoid lineage instead of the myeloid. Nevertheless, how lymphoid progenitors produce DCs and through which developmental stages it progresses to DCs remains to be elucidated.

In order to resolve the classification of monocyte and DC lineage, all identified progenitors, both myeloid and lymphoid, need to be identified and exhaustively tested for their developmental potential as well as their progenitor-progeny relationship. By analyzing which progenitor is capable of producing GMDPs, MDPs and CDPs, we will be able to trace and determine which progenitor contributes toward the DC pathway the most. Furthermore, single cell cultures also need to be done in order to determine the clonal composition of DC producing clones, as we suspect that lymphoid progenitors are heterogeneous in their clonal potential similar to GMPs.

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Chapter V. Revisiting the DC development model

94

1. Introduction

The origin of human DCs has been contentious. This is primarily because data generated from DC development studies often conflict with the standard model of hematopoiesis. The standard model of hematopoiesis proposes that the myeloid-erythroid lineage and lymphoid lineage split into two separate non-overlapping trajectories downstream the long term hematopoietic stem cells (HSCs) and short term multipotent progenitors (MPPs)146. This map was established after the identification of two progenitors supporting this process: (1) the common lymphoid progenitor (CLP)9 and (2) the common myeloid progenitor (CMP)10, 12.

Following identification of CMPs and CLPs in mouse, CLPs in human were identified as

CD34+CD45RA+CD10+ cells in the bone marrow14. Extensive clonal analysis testing their developmental potential showed that these cells failed to produce myeloerythroid cells but successfully produced B, NK and T cells both in vitro and in vivo. This provided evidence that the lymphoid lineage, indeed arises from a progenitor that lacks myeloerythroid potential.

Interestingly, when single CLPs were tested for DC potential in vitro, they showed significant

DC activity, which suggested a lymphoid origin for human DCs.

Then CMPs11, 13 were identified by the expression of CD34+CD348+CD45RA-CD123+.

CMPs, on the other hand, had megaerythrokaryocytic potential, as well as granulocyte and macrophage potential, but failed to produce any lymphocytes. This same study also showed that

CMPs produce megaerythrokaryocytes and granulocytes-macrophages through two progenitors with mutually exclusive phenotype. CD34+CD348+CD45RA+CD123+ were called megaerythrokaryocyte progenitor (MEK) and the CD34+CD348+CD45RA-CD123- cells were called granulocyte-macrophage progenitor (GMP). These results showed, for the first time, that the myeloid lineage in humans is clearly a separate lineage from the lymphoid.

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CMPs and GMPs were later examined for their DC potential both in vitro and in vivo15, 19.

Not surprisingly, CMP and GMPs both produced DCs. However, when DCs derived from

CMP/GMPs were compared to DCs derived from CLPs, results showed that they were transcriptionally indistinguishable, which suggested that DCs could derive from both myeloid and lymphoid progenitors15. Since then, the central origin of DCs became debated.

With more antibodies available to differentiate surface markers on human cells, additional progenitors have been identified thereafter and placed on the hematopoietic map, adding more layers of complexity. A new progenitor was identified in AML leukemia patients with a CD34+CD38-CD45RA+ phenotype that could produce GMPs, bypassing CMP stage21. We showed in our last chapter that GMPs are, indeed, heterogeneous and identified three consecutive

DC progenitors based on the differential expression of CD123 and CD115. Additionally, CLPs could be further subdivided into CD38+ BNK progenitors and CD38- multilymphoid progenitors

(MLPs), with DC potential residing in MLPs20. However, how these different progenitors developmentally relate with each other has not been thoroughly addressed.

Therefore, to resolve the DC origin conflict, we need to exhaustively separate the 100% of CD34+ human hematopoietic progenitors into phenotypically non-overlapping populations in order to determine their terminal differentiation potential as well as their inter-progenitor relationship at the population and single cell level.

In this chapter, we present first a multi-channel staining panel to characterize ten mutually exclusive populations based on their phenotype. Then, by combining in vitro and in vivo assays we describe the developmental potential of each population as well as their inter- progenitor relationship. Finally, we show single cell culture analysis in order to increase the resolution for tracing the DC developmental pathway.

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2. Results

Exhaustive phenotypic characterization of 10 non-overlapping progenitors

In order to determine the developmental relationship of different myeloid and lymphoid progenitors to DCs, we first established a staining panel able to identify all previously characterized CD34+ hematopoietic precursors in cord blood (Table 5.1). This analysis allows us to exhaustively fractionate and phenotypically compare previously found progenitors, including the recently described GMDPs, MDPs and CDPs (Fig 5.1A and B).

By using only seven surface markers (CD34, CD38, CD45RA, CD90, CD10, CD123,

CD115), we are able to divide CD34+ cells into 10 non-overlapping marker pure progenitors (Fig

5.1A and B). All hematopoietic progenitors and stem cells are uniformly CD34+. Among these, two main groups can be subdivided according to their CD38 expression: CD38- early progenitors, and CD38+ late progenitors. Within the early CD38- cells we can identify the hematopoietic stem cells (HSCs) by their expression of CD90 and lack of CD45RA

(CD34+CD38-CD45RA-CD90+). HSCs have been shown to produce all hematopoietic lineages in culture and in vivo in xenogeneic NSG mice. HSCs can also repopulate secondary recipient mice after serial transplantation, resembling the long-term hematopoietic stem cells in mouse (LT-

HSC).

Within the CD90- cells, we can further identify three populations. The short-term hematopoietic stem cells, also called multipotent progenitor (MPP), and two lymphoid progenitors. The MPPs are characterized by their CD34+CD38-CD45RA-CD90- phenotype and, although they can produce all blood lineages, they can do so only transiently. The two lymphoid precursors are the lymphoid-primed multipotent progenitors (LMPPs, CD34+CD38-CD90-

CD45RA+CD10-) and the multilymphoid progenitors (MLPs, CD34+CD38-CD90-

97

CD45RA+CD10+). These two express CD45RA but differentially express CD10 on their surface.

They both have the potential to produce B, NK cells, monocytes and DCs, but not megaerythrokaryocytes. One difference between these two is that LMPPs have very limited granulocyte potential while MLPs completely lack granulocyte potential.

Among the late progenitors, we can also identify a committed B/NK progenitor (BNKP,

CD34+CD38+CD45RA+CD10+). These, like their parent MLPs, express CD10 and CD45RA on their surface but strictly produce lymphocytes and not myeloid cells of any type, including DCs.

Within the CD10- late progenitors, we can find five myeloid progenitors. The earliest among these five is the common myeloid progenitor (CMP, CD34+CD38+CD45RA-CD123+), which has the potential to produce all myeloid lineages both in vivo and in vitro: megaerythrokaryocytes, granulocytes, monocytes and DCs. CMPs then produce two separate progenitors with distinct developmental potential. One is the megaerythrokaryocyte progenitor (MEK,

CD34+CD38+CD45RA-CD123-), which is restricted to producing red blood cells and megakaryocytes, and the other is the aforementioned GMDPs (CD34+CD38+CD45RA+CD123+), which can produce granulocyte, monocytes and DCs, but not megaerythrokaryocytes or lymphocytes. As previously shown, downstream GMDPs are the MDPs, which produce only monocytes and DCs, and finally the CDPs, which are restricted to DCs alone.

Therefore, this panel allows us to exhaustively fractionate progenitor cells from cord blood and bone marrow into 10 non-overlapping marker-pure populations. This in time will also allow us to determine their separate developmental potential toward DCs and other lineages, as well as their interdevelopmental relationship.

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Table 5.1 Phenotype, potential and inter-developmental relationship of human CD34+ hematopoietic progenitor populations. (A) Name, surface marker phenotype, differentiation potential of 10 non-overlapping CD34+ hematopoietic progenitor populations: HSC, MPP,

LMPP, MLP, CMP, MEP, GMDP, MDP and CDP in human cord blood and bone marrow. CFU, colony-forming units assay; MS5, MP+FSG culture assay.

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Lineage output CB (% of BM (% of Population Phenotype CD34+) CD34+) CFU MS5 Hematopoietic stem cell (HSC) CD34+CD38-CD45RA-CD90+ 3.79 1.70 ME G M DC L Multipotent progenitor (MPP) CD34+CD38-CD45RA-CD90- 18.97 11.30 ME G M DC L Lymphoid-primed multipotent progenitor (LMPP) CD34+CD38-CD45RA+CD10- 1.36 16.50 G M DC L Multilymphoid progenitor (MLP) CD34+CD38-CD45RA+CD10+ 4.88 0.80 M DC L B and NK cell progenitor (BNKP) CD34+CD38+CD45RA+CD123int/-CD115-CD10+ 3.12 2.20 L Common myeloid progenitor (CMP) CD34+CD38+CD45RA-CD10-CD123int 39.84 33.80 ME G M DC Granulocyte/Monocyte/DC progenitor (GMDP) CD34+CD38+CD45RA+CD10-CD123int 11.38 11.20 G M DC Monocyte/DC progenitor (MDP) CD34+CD38+CD45RA+CD123intCD115+ 0.81 5.60 M DC Common DC progenitor (CDP) CD34+CD38+CD45RA+CD123hiCD115- 0.54 3.00 DC Megaerythrokaryocyte progenitor (MEP) CD34+CD38+CD45RA-CD10-CD123- 15.34 11.40 ME ME, erythrocyte and megakaryocyte; G, granulocyte; M, monocyte; DC, dendritic cell; L, B/NK lymphocytes. Colony forming unit assay, CFU; MS5 stromal cell culture assay, MS5.

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Figure 5.1 Phenotypic characterization of 10 non-overlapping populations. The gating scheme of defining populations as in Table 5.1 from a representative human (A) cord blood and

(B) bone marrow sample.

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A BNKP CMP CDP

MDP

CD10 MEP GMDP CD123 CD123 CD45RA CD45RA CD115 late HSC MPP MLP early LMPP CD90 CD10 CD38 CD34 CD45RA CD45RA

B BNKP CMP CDP MDP

CD10 MEP GMDP CD123 CD123 CD45RA CD45RA CD115 late MPP HSC MLP

early LMPP CD90 CD10 CD38 CD34 CD45RA CD45RA

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Determining the developmental potential of CD34+ progenitors

After phenotypic separation of CD34+ progenitors into 10 non-overlapping progenitors, we determined the developmental potential of each population by using three separate assays: methylcellulose culture, MP+FSG culture and cell transfers into NSG-SGM3 immunocompromised mice (Fig 5.2).

In order to determine their myeloerythrocytic potential, we have isolated 500 cells from each progenitor population and cultured them on methylcellulose supplemented with human recombinant cytokines (SCF, GM-CSF, IL-3, EPO) (Fig 5.2A). Colony formation analysis shows that, consistent with previous experiments (Fig 4.2E), CDPs failed to produce any identifiable colonies, MDPs produced only macrophage colonies at high numbers, GMDPs produced both macrophage and granulocyte colonies, and none of the progenitors produced erythrocyte colonies. In contrast, CMPs, HSCs and MPPs had the capacity to produce erythrocyte, macrophage and granulocyte colonies (Fig 5.2A). This confirms that CMP possess megaerythrokaryocyte potential and that GMDPs do not.

As expected, lymphoid progenitors (LMPPs, MLPs and BNKPs) lack megaerythrokaryocytic potential. Furthermore, although LMPPs produced myeloid colonies, the numbers are low compared to GMDPs or MDPs (3-fold) (Fig 5.2A). Of those positive clones, the majority were macrophages (approximately 70%) and the remaining colonies were mixed with both macrophages and granulocytes, which indicates that LMPPs have limited macrophage and granulocyte potential (Fig 5.2A). MLPs produced very few macrophage colonies in methylcellulose (i.e. 2+2 per 500 plated cells) (Fig 5.2A). Finally, BNKPs failed to produce any positive megaerythrokaryocyte and myeloid colonies. We conclude that the lymphoid progenitors do not possess megakaryocyte and erythrocyte potential and have limited, if any,

103 myeloid potential. This follows the traditional model of hematopoiesis whereby lymphoid and myeloid lineages follow mutually exclusive and non-overlapping pathways, with CMPs and

LMPPs bifurcating downstream HSC/MPPs.

We then determined their monocyte, DC and lymphoid potential using the MP+FSG culture (Fig 5.2B). Based on lineage output results, HSCs and MPPs are multipotent, as seen by their robust production of granulocytes, monocytes, lymphocytes and all three DC subsets.

Consistent with previous results (Fig 4.2B), CMPs and GMDPs produced all myeloid cell types except for lymphocytes. MDPs produced robust monocyte and DC output, but very few granulocytes and no lymphocytes. CDPs, on the other hand, were restricted to DCs alone.

LMPPs, on the other hand, had robust lymphoid, monocyte and DC potential, but much fewer granulocyte potential than GMDPs or CMPs (Fig 5.2B). MLPs, produced high number of

B/NK cells, fewer monocytes and no granulocytes. Interestingly, MLPs produced more pDCs than cDCs. Finally, BNKPs only produced lymphocytes and failed to generate any type of myeloid cell.

Therefore, consistent with previous investigations, multipotent progenitors (HSCs,

MPPs) and myeloid progenitors (CMP, GMDP, MDP, CDP) have the potential to produce myeloid cells, including monocytes and DCs. What was surprising was that, lymphoid progenitors, which can produce B/NK cells in culture, also overlap in their potential to produce monocytes and DCs.

In vivo transfers into NSG-SGM3 mice also confirm these results (Fig 5.2C). Multipotent progenitors (HSC/MPPs) were capable of producing all cell types after 14 days of transfer, including B/NK cells. Myeloid progenitors (CMPs and GMDPs) produced granulocytes monocytes and DCs, but very little lymphocytes compared to LMPPs and MLPs. The latter two

104 populations produced robust number of B/NK cells but fewer granulocytes and monocytes.

Especially, MLPs almost lacked any granulocyte and monocyte activity but retained strong DC activity. Although LMPPs produced granulocytes, it was at a lower frequency than those from

CMPs and GMDPs. However, they were still able to produce DCs at higher frequencies than

CMPs and GMDPs. These results further support that lymphoid and myeloid progenitors show persistent developmental overlap on the monocyte and DC lineage.

Thus, we conclude that DC and monocyte potential are present in multipotent progenitors

(HSCs, MPPs), as well as in myeloid (CMPs, GMDPs, MDPs or CDPs) and in lymphoid progenitors (LMPPs, MLPs). Our culture and in vivo experiments show that the developmental potential is not as pure and non-overlapping as the surface phenotype is. The “convergence” of myeloid and lymphoid progenitors on the monocyte and DC lineage suggests the presence of an inter-progenitor relationship between myeloid and lymphoid progenitors with DC restricted progenitors.

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Figure 5.2 Megaerythrokaryocyte, myeloid and lymphoid potential. (A) Stacked columns show colony forming units (CFU) produced by HSC, MPP, CMP, GMDP, MDP, CDP, LMPP,

MLP and BNKP in methylcellulose after 14 days of culture. M, macrophage; G, granulocyte;

GM, granulocyte and macrophage; GEMM, granulocyte, erythrocyte, megakaryocyte and macrophage; E, erythrocyte. Bars are mean averages and error bars are SEM from four independent experiments. (B) Flow cytometry plots show output from 100 cells from each of

HSC, MPP, CMP, GMDP, MDP, CDP, LMPP, MLP and BNKP populations in MP+FSG culture after 14 days. Cells gated initially are live CD45+ and plots are representative of five independent experiments. (C) Plots show in vivo progeny of HSC/MPPs, CMPs, LMPPs and MLPs. 1x104 cells of indicated progenitors were sorted from cord blood and injected into the bone cavity of conditioned NSG-SGM mice. 2 weeks post-transfer, tibias have been harvested and analyzed for the presence of fully differentiated cells. Cells gated initially are live/CD45+ and plots are representative of three independent experiments.

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A 70 CFU-M 60 CFU-G 50 CFU-GM 40 CFU-GEMM 30 CFU-E 20

# of colonies#of 500 / cell 10 0

HSC MPP CMP MDP CDP MLP GMDP LMPP BNKP

B Granulocytes CD141 cDC Monocytes CD1c cDC

pDC B/NK HSC

MPP

CMP

GMDP

MDP

CDP

LMPP

MLP

BNKP SSC SSC CD123 CD14 CD141 CD66b CLEC9a CD1c CD303 CD19/56

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C CD141 CD1c Granulocytes DC Monocytes DC pDC B/NK HSC/ MPP

CMP

GMDP

LMPP

MLP SSC SSC CD14 CD66b CD141 CLEC9a CD1c CD123 CD303 CD19/56

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Interdevelopmental relationship among CD34+ progenitors

Based on the overlapping myeloid and lymphoid potential of different progenitors, we decided to further investigate whether there was a progenitor-progeny relationship among different populations. In order to do this, we first stained CD34+CD38-CD45RA- HSC/MPP cells with CFSE and cultured them on MP+FSG. After seven days, we determined the developmental hierarchy of different populations by analyzing the proliferation cycle at which different downstream progenitors arise (Fig 5.3A): LMPPs appeared at division 1-2, CMPs at division 3,

GMDPs at division 3-4, MLP/BNKPs and MDPs at division 5, and CDPs at division 7, indicating a hierarchy among the marker-pure progenitor populations.

We then investigated whether there was a direct progenitor-progeny relationship between myeloid and lymphoid progenitors. For this, we analyzed by flow cytometry downstream progeny cells derived from HSC/MPPs, CMPs, LMPPs, MLPs and BNKPs separately after

MP+FSG cultures (Fig 5.3B). After 7 days, HSC/MPPs produced CMPs, LMPPs, GMDPs and very few MDPs and MLP/BNKPs. CMPs generated GMDPs, MDPs and very few CDPs, but failed to produce lymphoid progenitors, such as LMPPs or MLP/BNKPs. LMPPs, on the other hand, produced downstream MLP/BNKPs, but more strikingly, GMDPs, MDPs and CDPs.

Similarly, MLPs also produced MLP/BNKPs, as well as GMDPs, MDPs and CDPs. However, neither LMPPs or MLPs produced CMPs. Finally, BNKPs produced negligible CD34+ cells, indicating rapid differentiation. In sum, lymphoid progenitor LMPPs and MLPs can generate

GMDPs, MDPs and CDPs but not CMPs. The latter, which cannot give rise to lymphoid progenitors, also produce GMDPs, MDPs and CDPs. This reveals that lymphoid and myeloid progenitors developmentally “converge” at the GMDP stage without giving rise to each other, which we also corroborated through in vivo transfers into NSG-SGM3 mice (Fig 5.3C).

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Thus, we show by phenotypic analysis three distinct developmental pathways for lymphoid and myeloid cell differentiation: (1) HSC/MPP è LMPP è MLP/BNKP; (2)

HSC/MPP è LMPP è MLP è GMDP è MDP è CDP; (3) HSC/MPP è CMP è GMDP

è MDP è CDP (Fig 5.3D). Although this model supports the classical hematopoiesis model, whereby myeloid (i.e. CMP) and lymphoid progenitors (i.e. LMPPs and MLPs) follow separate pathways immediately downstream MPPs, it presents a new conflict: LMPPs and MLPs, which have limited granulocyte potential, are upstream of GMDPs, a population with robust granulocyte potential. This is incompatible with the dogma of hematopoiesis, i.e., lineage potential is irreversibly lost along hematopoiesis.

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Figure 5.3 Interdevelopmental relationships among CD34+ progenitors. (A) Concatenated

FACS plots show division of indicated populations descending from HSCs. 1 x 103 HSCs defined as Fig. 5.1 were sorted, labeled with CFSE and cultured for 6 days. Populations were gated as in (B), and CFSE dilution was shown in dot plots. Numbers indicate divisions that cells have undergone. Plot is representative of four independent experiments. (B) Representative flow cytometry plots show output populations from 1x103 HSCs/MPPs, CMPs, LMPPs, MLPs and

BNKPs after 7-day culture of 3 independent experiments. Cells were initially gated as live

CD45+Lin(CD3/19/56/14/16/1c/303/141)-CD34+. (C) Plots show the in vivo production of downstream progenitors from HSC/MPPs, CMPs, and LMPPs. 1x104 cells from indicated progenitors were sorted and injected into the tibia of preconditioned NSG-SGM3 mice. Tibias were analyzed after 7 days for the presence of hematopoietic progenitors. Cells were initially gated as live/CD45+Lin(CD3/19/56/14/16/1c/303/141)-CD34+. Plots are representative from three independent experiments. (D) Schematic picture summarizes the inter-progenitor relationship of CD34+ progenitors.

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A C 26.8 26.8 8 7 6 5 4 3 2 1 0 4.57 CDP HSC/ CMP Non-dividing MDP MPP LMPP/ HSC/MPP MLP 11.1 60.3GMDP26.9

LMPP 56 34 2.2

CMP CMP GMDP 0 54 39 MLP/BNKP 7.1 87 20 MDP LMPP CDP 0 53 20 CD38 CD45RA CD123 CD115 CFSE

B Input MLP/ D Cell BNKP CMP LMPP CDP 7.6 41.8 1.0 0.1 MDP HSC/ HSC/ MPP

MPP 85.3 98.4 9.9 12.1 10.6 70.9 GMDP 0.8 0.1 CMP LMPP CMP 52.6 98.5 2.6 43.6

4.9 66.0 4.2 6.5 GMDP MLP LMPP 91.1 9.0 69.4 19.1

10.2 3.4 74.4 3.3 MDP BNKP MLP 86.0 10.4 69.9 23.7

17 0 64 0 CDP BNKP

CD7 78 29 89 0 CD38 CD123 CD45RA CD45RA CD115

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Phenotypically uniform progenitors are clonally heterogeneous

The above results show that there are three populations that can potentially contribute to

GMDPs, MDPs and CDPs. This suggests that phenotypically homogeneous GMDPs are actually heterogeneous, with input from both lymphoid and myeloid progenitor cells. In order to resolve this question, we decided to increase the resolution by performing single cell cultures from each progenitor.

Using our MP+FSG culture and CD34+ staining panel, we isolated a total of >5,000 individual clones from HSCs, MPPs, LMPPs, MLPs, BNKPs, CMPs, GMDPs, MDPs and CDPs from cord blood. Through flow cytometry analysis (Fig 5.4A), we determined their granulocyte

(G), monocyte (M), CD1c cDC (C), CD141 cDC (A), pDC (P) and lymphocyte (B or NK, L) potential at the clonal level.

The high clonal efficiency, as determined by clones that generated identifiable human

CD45+ cells, of HSCs (>40%), MPPs (50%), GMDPs (60%) and MDPs (60%) indicates that

MP+FSG cultures are optimal for survival, proliferation or differentiation of single CD34+ progenitor cells (Fig 5.4B). We obtained a cumulative total of 2,247 positive clones from all nine progenitors.

Analysis of global composition of positive clones from each progenitor does follow the developmental hierarchy observed in Fig 5.3A. HSCs and MPPs have the highest ratio of multipotent clones (6-lineage clones, Fig 5.4C) with the highest average yield (Fig 5.4D) than other progenitor populations. As downstream progenitors differentiate from HSC/MPPs, they contain more committed clones with more limited mean proliferative capacity (Fig 5.4C and D), which is consistent with their developmental distance from HSC/MPPs.

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On the other hand, our analysis also reveals that each population is extremely clonally heterogeneous. The standard model presumes that phenotypically homogeneous populations are also functionally homogeneous. However, our findings demonstrate that each population defined by phenotype is indeed comprised of a mix of clones with distinct differentiation and proliferation capacity (Fig 5.4C and D). For example, LMPP progenitors contain clones with different lineage combinations (Fig 5.4E, horizontal red box), which suggest that different clones with distinct clonal potential may acquire identical phenotype. Furthermore, we observe that the same type of clone can also be present in different progenitors that have different phenotype (Fig

5.4E, vertical red box). This proves that a specific lineage potential does not directly correlate with a specific phenotype.

In fact, we can also observe that lineage commitment, as defined by the presence of unilineage clones, occurs at different stages of hematopoiesis (Fig 5.4C and F) rather than at a single population. Granulocyte-committed clones are concentrated in both CMP and GMDP populations, but can also be found as early as in HSC and MPPs (Fig 5.4F). Monocyte- committed clones are concentrated in MDPs, but can also be found in GMDPs, CMPs and

HSC/MPPs. DC-committed clones are concentrated in CDPs and MDPs, as well as in GMDPs,

MLPs and LMPPs. Finally, B/NK cell-committed clones gradually increase in numbers from

HSC/MPPs, to LMPPs, to MLPs and to BNKPs.

Therefore, we conclude that progenitor populations defined by their homogeneous expression of specific surface markers do not clearly represent their true developmental potential and clonal heterogeneity, and hence, cannot be used as criteria to delineate lineage pathways.

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Figure 5.4 Phenotypically uniform progenitors are clonally heterogeneous. (A)

Representative flow cytometry plot shows phenotype of live, human CD45+ cells produced from a single multipotent positive clone. All clones were classified by their output cells including granulocytes (G, brown), monocytes (M, orange), B/NK cells (L, purple), CD141 cDCs (A, red);

CD1c cDCs, (C, blue) and pDCs (P, green). (B) Single cells with progenitor population markers presented in Table 5.1 were sorted by FACS and cultured for 2-3 weeks. Emerging clones were analyzed by flow cytometry for production of G, M, C, A, P or L. Results are from cumulative clones isolated from 17 cord blood donors. Bar graphs show clonal efficiency. n, number of total seeded clones. (C) Stacked bar graphs summarize global lineage outcome of clones in each population. (D) Scatter plots show CD45+ cell yield of all clones in each population. (E) Scatter plots show lineage outcome (X-axis) and CD45+ cells yield (Y-axis) of all clones for each population (rows). (F) Dot plots show clonal yield of granulocytes, monocytes, CD1c cDC, pDC,

CD141 cDC and lymphocytes from uni-potent (colored) and multi- or oligo-potent (grey) clones in each population.

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A Granulocytes CD141 cDC Monocytes CD1c cDC Culture on pDCs B/NK MP+FSG (7d-14d) SSC SSC CD14 CD141 CD123 CD66b CLEC9a CD1c CD303 CD19/56

B 80 E # of lineages per clone n=360n=408 n=791 n=720 n=542 n=800 n=890 n=357 n=691 60 6 5 4 3 2 1

6 HSC 40 1×10 1×105 1×104 3 20 1×10 1×102

Clonal Efficiency (%) 1×101 1×100 0 1×106 L G C A P M G L G C G A G P P L C L A L C A C P M L A P

5 MPP G M M C M P M A G C L G P L G A L G M L M A L G C A G A P G C P C P L G M P C A L A P L G M A M A P

1×10 C A P M C L M P L G M C M C A M C P G C P L G C A L G A P L G M C L M A P L G M P L G C A P G M A L C A P L G M C P G M A P M C A L M C P L G M C A M C A P G C A P L MLP 4 G M C A L G M C P L G M A P L HSC MPP CMP MDP CDP M C A P L LMPP BNKP 1×10 G M C A P GMDP G M C A P L 3 1×10 Lineage loss C 2 100 # of lineage 1×10 combinations 1×101 0 per clone 1×10 80 1×106 L G C A P M G L G C G A G P P L C L A L C A C P M L A P

5 G M M C M P M A G C L G P L G A L

LMPP G M L M A L G C A G A P G C P C P L G M P C A L A P L G M A M A P

1 1×10 C A P M C L M P L G M C M C A M C P G C P L G C A L G A P L G M C L M A P L G M P L G C A P G M A L C A P L G M C P G M A P M C A L M C P L G M C A M C A P G C A P L

4 G M C A L G M C P L G M A P L M C A P L

1×10 G M C A P

60 2 G M C A P L 3 1×10 Lineage loss 3 1×102 40 4 1×101 1×100 20 5 1×106 L G C A P M G L G C G A G P P L C L A L C A C P M L A P

5 G M M C M P M A G C L G P L G A L

CMP G M L M A L G C A G A P G C P C P L G M P C A L A P L G M A M A P

1×10 C A P M C L M P L G M C M C A M C P % of positive%of clones G C P L G C A L G A P L G M C L M A P L G M P L G C A P G M A L C A P L G M C P G M A P

6 M C A L M C P L G M C A M C A P G C A P L

4 G M C A L G M C P L G M A P L M C A P L

1×10 G M C A P

0 G M C A P L 3 1×10 Lineage loss 1×102 HSC MPP CMP MLP MDP CDP LMPP GMDP BNKP 1×101 D 106 1×100 1×106 L G C A P M G L G C G A G P P L C L A L C A C P M L A P

5 G M M C M P M A 5 G C L G P L G A L G M L M A L G C A G A P G C P

GMDP C P L G M P C A L A P L G M A M A P

1×10 C A P M C L M P L G M C M C A M C P

10 G C P L G C A L G A P L G M C L M A P L G M P L G C A P G M A L C A P L G M C P G M A P M C A L M C P L G M C A M C A P G C A P L

4 G M C A L G M C P L G M A P L M C A P L

1×10 G M C A P cell output G M C A P L

4 + 3 10 1×10 Lineage loss 1×102 3 10 1×101 CD45 cell output 1×100 + 2 10 1×106 L G C A P M G L G C G A G P P L C L A L C A C P M L A P

5 G M M C M P M A G C L G P L G A L G M L M A L G C A G A P G C P C P L G M P C A L A P L G M A M A P

1×10 C A P M C L M P L

MLP G M C M C A M C P G C P L G C A L G A P L G M C L M A P L G M P L G C A P G M A L

1 C A P L G M C P G M A P M C A L M C P L G M C A M C A P G C A P L

10 4 G M C A L G M C P L G M A P L M C A P L G M C A P

CD45 1×10 G M C A P L 1×103 100 Lineage loss 1×102 1×101 HSC MPP CMP MLP MDP CDP LMPP GMDP BNKP 1×100 1×106 L G C A P M G L G C G A G P P L C L A L C A C P M L A P

5 G M M C M P M A G C L G P L G A L G M L M A L G C A G A P G C P C P L G M P C A L A P L G M A M A P

1×10 C A P M C L M P L G M C M C A M C P

BNKP G C P L G C A L G A P L G M C L M A P L G M P L G C A P G M A L C A P L G M C P G M A P M C A L M C P L G M C A M C A P G C A P L

4 G M C A L G M C P L G M A P L M C A P L

1×10 G M C A P G M C A P L 3 1×10 Lineage loss 1×102 1×101 1×100 1×106 L G C A P M G L G C G A G P P L C L A L C A C P M L A P

5 G M M C M P M A G C L G P L G A L G M L M A L G C A G A P G C P C P L G M P C A L A P L G M A M A P

1×10 C A P M C L M P L G M C M C A M C P G C P L G C A L G A P L

MDP G M C L M A P L G M P L G C A P G M A L C A P L G M C P G M A P M C A L M C P L G M C A M C A P G C A P L

4 G M C A L G M C P L G M A P L M C A P L

1×10 G M C A P G M C A P L 3 1×10 Lineage loss 1×102 1×101 1×100 1×106 L G C A P M G L G C G A G P P L C L A L C A C P M L A P

5 G M M C M P M A G C L G P L G A L G M L M A L G C A G A P G C P C P L G M P C A L A P L G M A M A P

1×10 C A P M C L M P L G M C M C A M C P G C P L G C A L G A P L G M C L M A P L G M P L G C A P G M A L

CDP C A P L G M C P G M A P M C A L M C P L G M C A M C A P G C A P L

4 G M C A L G M C P L G M A P L M C A P L

1×10 G M C A P G M C A P L 3 1×10 Lineage loss 1×102 1×101 1×100 L G C A P M G L G C G A G P P L C L A L C A C P M L A P G M M C M P M A G C L G P L G A L G M L M A L G C A G A P G C P C P L G M P C A L A P L G M A M A P C A P M C L M P L G M C M C A M C P G C P L G C A L G A P L G M C L M A P L G M P L G C A P G M A L C A P L G M C P G M A P M C A L M C P L G M C A M C A P G C A P L G M C A L G M C P L G M A P L M C A P L G M C A P G M C A P L Lineage loss

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F Granulocyte (G) Monocyte (M) 1×106 1×105

5 1×10 1×104 1×104 1×103 1×103 1×102 1×102 1 1×101 1×10

1×100 1×100 1×105 CD1c cDC (C) 1×104 CD141 cDC (A) HSC MPP MLP CMP MDP CDP HSC MPP MLP CMP MDP CDP LMPP BNKP GMDP LMPP BNKP GMDP 1×104 Progenitors 1×103 Progenitors 1×103 1×102 1×102 1×101 1×101 Cellnumber output 1×100 1×100 1×104 1×105 HSCpDCMPP (P) MLP CMP MDP CDP HSCLymphocyteMPP MLP (L) CMP MDP CDP LMPP BNKP GMDP LMPP BNKP GMDP Progenitors 1×104 Progenitors 1×103 1×103 1×102 1×102 1×101 1×101

1×100 1×100

HSC MPP MLP CMP MDP CDP HSC MPP MLP CMP MDP CDP LMPP BNKP GMDP LMPP BNKP GMDP Progenitors Progenitors

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Correlation between clonal output and lineage potential

In order to trace each lineage, including that of the DCs and monocytes, with high resolution, we propose to use two clonal parameters: (1) number of lineages a cell produced (G,

M, C, A, P or L) and (2) yield of cells generated per lineage. When we clustered all positive clones according to the number of lineages each cell produced, we see a strong correlation between cell number output and lineage combinations (Fig 5.5A). This shows that the more multipotent a clone is, the more cells a clone can generate. In other words, it suggests that there is a relationship between the proliferative capacity and lineage commitment.

In order to prove this relationship, we have stained HSC/MPPs with CFSE and performed clonal cultures from cells that have undergone 0, 3 or 6 divisions as in Fig 5.3A. After two weeks of culture, we measured their clonal yield as well as the lineage composition. Compared to cells that have not divided (division 0), cells that had divided 3 times showed approximately ten-fold decrease in CD45+ cell number output (Fig 5.5B) as well as a decrease in multipotent clones. Inversely, the relative number of oligopotent (2-5 lineage combinations) and unipotent clones increased (Fig 5.5C). Cells that had divided 6 times showed additional decrease of cellular output and a significant increase of unipotent clones. Thus, this shows that cellular yield and clonal lineage potential can be indicative of the developmental stage and distance from

HSC/MPPs, independent of surface marker phenotype.

Therefore, we propose that by using the cellular output for each lineage generated by a single clone as a metric for lineage direction, we will be able to determine its developmental distance from HSC/MPPs as well as from other lineages.

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Figure 5.5 Correlation between clonal output and lineage potential. (A) Scatter plot shows correlation between lineage potential and CD45+ cell yield of all clones. r, Spearman correlation coefficient. (B and C) 1 x 103 CFSE-labeled HSCs were cultured for 6 days on MP+FSG. Cells were sorted from division 0, 3 and 6 for clonal analysis. (B) Graph shows the yield of clones isolated from dividing HSCs that had undergone 0, 3 or 6 divisions (Div). (C) Stacked bars show global view of lineage outcome for clones in (B). Results are from cumulative clones of 3 independent experiments. Asterisks indicate significance based on Fisher’s exact test on uni- potent cell composition.

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P=0.0014 (**) P<0.0001 (***) A B C P<0.0001 (***) P<0.0001 (***) 107 105 r = 0.7 P=0.02 (*) 106 P<0.0001 104 105

104 103 celloutput cell output + + 103 102 # of lineage CD45 CD45 102 combinations % of positive%of clones per clone 101 101 1 2 3 4 5 6

Lineage Potential Div0 Div3 Div6 Div0 Div3 Div6

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3. Discussion

Using a staining panel that allows the identification of 10 non-overlapping marker pure progenitors in cord blood and bone marrow, we have extensively characterized the developmental potential of each separate population in vitro and in vivo. This revealed consistent with previous findings that both myeloid and lymphoid progenitors retained DC and monocyte potential. We have also elucidated that both myeloid and lymphoid progenitors are capable of doing so by giving rise to DC associated progenitors, namely GMDPs, MDPs and CDPs.

However, these results opened a new question: how can lymphoid progenitors that lack granulocyte activity produce GMDPs, which have granulocyte potential? This suggested a mix contribution to GMDPs by both myeloid and lymphoid potential. Interestingly, clonal analysis of each progenitor revealed with high resolution the heterogeneity in clonal potential within phenotypically homogeneous progenitors, not only within GMDPs, but in all others as well, therefore making phenotypic analysis to trace lineage development inadequate.

In order to accurately trace each lineage irrespective of its phenotype of origin, we proposed to use the cellular output information obtained from clonal cultures. Since the cellular yield and lineage combination produced by each clone reflect the developmental distance and direction from HSC/MPPs, we hypothesize that these values can be used as metrics for determining precise lineage pathways.

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Chapter VI. Discussion

1. Result summary

Our studies have endeavored to overcome three major hurdles deterring our understanding of human DC development: (1) lack of culture that allows the development of the three authentic DC subsets together with multilineage cells, (2) unclear identity of DC restricted progenitors in human bone marrow and in the periphery, and (3) conflict in overlapping DC activity in both myeloid and lymphoid progenitors.

First, we have established a novel culture system (MP+FSG) able to support the development of three subsets of human DCs, together with monocytes, granulocytes and lymphocytes, such as B and NK cells. This allowed us to determine the DC potential of new and previously identified progenitors within the context of other myeloid and lymphoid lineages.

Especially, we have shown that the DCs produced in vitro are phenotypically, transcriptionally and functionally equivalent to those in peripheral blood, and are not derived from monocytes.

Furthermore, this culture also supported robust proliferation from single hematopoietic progenitor cells, allowing us to investigate the developmental potential of individual clones.

Using this culture we have identified the granulocyte, monocyte and DC progenitor

(GMDP), the monocyte and DC progenitor (MDP) and common DC progenitor (CDP), all within the previously described CD34+CD38+CD10-CD45RA+ GMPs. Single cell culture of GMPs revealed clonal heterogeneity within this population, prompting us to fractionate them using DC- associated cytokine receptor expression. Phenotypic characterization and separation of GMPs based on the differential expression of CD116, CD115 and CD123 allowed the identification of three novel DC developmental stages with restricted DC activity. We showed that GMDPs

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(CD123+CD115-) produce granulocytes and MDPs (CD123+CD115+), which produce monocytes and CDPs (CD123hiCD115-), which finally produce all three DC subsets. We also identified one more step of specification between CDPs and cDCs, which we called the pre-cDCs. By fractionating cord blood CD34- cells according to the differential expression of cytokine receptors, we have identified a committed cDC precursor that lacks pDC potential with

CD45+Lin-CD34-CD117+CD135+CD116+ CD45RA+CD115- phenotype downstream CDPs.

Finally, in order to determine the main contribution by either myeloid or lymphoid progenitors to the DC pathway, we have established a staining panel for CD34+ cells that allows the identification of 10 non-overlapping marker-pure progenitors. Using this characterization method for identifying distinct progenitors, we have thoroughly tested the developmental relationship of these 10 progenitor populations for their potential to produce DCs, granulocytes, monocytes and lymphocytes. Our results show that (1) myeloid and lymphoid progenitors have overlapping DC and monocyte potential, (2) myeloid and lymphoid progenitors “converge” at the GMDP stage, and (3) marker pure populations are remarkably heterogeneous in developmental. Such results challenge the reliability of markers as parameters for identifying developmentally pure progenitors, for determining progenitor-progeny relationship, and eventually, for tracing the DC developmental pathway. Toward this end, we propose that quantitative clonal output number for any lineage can be used as a parameter to measure the developmental distance of each lineage from HSCs and from other cell lineages, independent of their cell surface markers. We hypothesize that quantitative clonal output could indicate potential and direction and will allow us to delineate with high resolution the DC developmental pathway.

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2. Establishing culture method and its importance in elucidating DC ontogeny

Despite the importance of human DC hematopoiesis, there has not been a good method to analyze simultaneously the differentiation potential of a single progenitor cell to all three DC subsets as well as other myeloid and lymphoid lineages. This is of high relevance in the field of human DC differentiation for two main reasons. First, DCs have been shown to descend from lymphoid progenitors as well as conventional myeloid progenitors; and although a restricted DC progenitor has been identified in mouse, its equivalent in humans was unknown. Therefore, to identify DC- restricted progenitors, candidate precursors must be tested for their DC differentiation potential in the context of other cell types. This can only be achieved in a culture that simultaneously nurtures myeloid and lymphoid lineages as well as DCs.

Furthermore, the DC lineage has been difficult to distinguish from the monocytic lineage.

Although GM-CSF and IL-4 culture has been widely used to generate human DCs from CD34+ cells or blood monocytes118, these GM-CSF-driven DCs have been shown to descend from monocytes and differ significantly from the authentic DCs in the steady state that depend on

FLT3L103. Therefore, a culture that simultaneously supports the development of both monocytes and authentic DCs was necessary to discern monocyte and authentic DC lineages.

We show that MS5 and OP9 stromal cells together with FLT3L, or with FLT3L, SCF and

GM-CSF, can simultaneously support the development of all three subsets of DCs, monocytes, granulocytes and lymphoid cells from CD34+ hematopoietic progenitors. The MP+FSG culture drives sufficient proliferation and differentiation for clonal assay. This indicates that signals from

MS5 and OP9 stromal cells enable differentiation of human DCs, suggesting that the molecules mediating such effects must be highly conserved between mouse and human.

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Importantly, different stromal cells selectively support the preferential expansion of specific DC populations. OP9 cells show significant bias toward CD141 cDCs and S17 cells toward CD1c cDCs. This suggests that each stromal cell expresses a different molecule or groups of molecules that collectively influence and regulate DC subset development. It is not known whether these are soluble or membrane bound surface proteins, but their identification would shed light in the process of DC subset specification, the developmental stages at which signaling is necessary for subset instruction and potential therapeutic approaches to selectively expand individual DC subsets.

Furthermore, MP+FSG may also support the development of other cell types not analyzed in this work. HSC/MPP cultures consistently showed the presence of a big portion of cells that did not express any of the markers we utilized to categorize fully differentiated cells

(Fig 5.2C). Thorough phenotypic characterization of these “undifferentiated” cells could reveal additional lineages that this culture can support. Additionally, MP+FSG cultures can also be used to elucidate the origin of lymphoid organ resident DCs. Recent studies have identified additional

DC subsets in lymph nodes (CD1c+CD1a+EpCam- DCs, CD1c+CD1a-CD206- DCs and

CD1c+CD1a-CD206+ DCs)88, 147 and in spleen (CD1c+CD14+ DCs)148. By using our stromal cell culture, we can determine the progenitor source and elucidate the molecular mechanism for their differentiation.

Lastly, although the administration of FLT3L, SCF and GM-CSF has provided a highly permissive environment for six lineages to develop, it still has some limitations. MP+FSG cultures do not support megaerythrokaryocytes or T cells, which are the great majority of the blood composition. We hypothesize that adding EPO for erythrocyte development and TPO for thrombocyte development, as well as using OP9-DL instead of OP9 cells135, will rescue this

125 deficit. Therefore, we believe our culture is not exhaustive as is, and there is room for optimization in order to support more lineages than those we described.

3. DC development in health and disease

The number of human diseases involving abnormal numbers of DCs is increasing.

However, our understanding of their developmental aberrations is still very limited. Two main reasons that hindered us from studying anomalous DC development are (1) the lack of knowledge in healthy DC development and (2) the lack of a system to interrogate the DC potential of hematopoietic progenitor cells from patients. Therefore, the identification of restricted monocyte and DC progenitors in human bone marrow, as well as establishing a culture system that supports multilineage cell development, will facilitate our understanding of aberrant

DC hematopoiesis.

For example, patients with genetic mutations in the IRF8 transcription factor gene completely lack DCs in the periphery149. In addition to DCs, these patients have significantly reduced numbers of monocytes while the granulocyte counts are normal. Irf8 has been shown to be necessary for murine DC and monocyte development, and human DCs and monocytes also express IRF8. This suggests that IRF8 is required for the transition from GMDPs to MDPs, but is dispensable for granulocyte development.

Another genetically defined syndrome of DC deficiency is the GATA2 associated

DC/monocyte/B and NK lymphoid (DCML) deficiency145, 150, 151. Patients harboring GATA2 mutations have a combined mononuclear cell deficiency in leukocytes and their interstitial tissue

DCs are lacking. Increased in serum FLT3L concentration and disseminated nontuberculous mycobacterial, viral and fungal infections are characteristic susceptibilities in these patients.

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Analysis of their CD34+ cell compartment revealed a dramatic reduction of CD45RA+ progenitors, which include the GMDPs, MDPs and CDPs, as well as the lymphoid MLP progenitor138. The normal number of granulocytes in these patients suggests that acquiring

CD45RA marks a crucial developmental step in monocyte and DC, but not for granulocytes.

DC deficiency is also common in chronic myelogenous leukemia (CML). CML is caused by a chromosomal translocation between chromosomes 9 and 22, forming the fusion oncogene

BCR-ABL152. cDCs and pDCs are consistently low in all stages of CML progression, while monocytes and granulocytes are present at either normal or higher levels153. CD34+ cell analysis revealed a complete absence of CD34+CD45RA+CD123hi cells154, 155, which correspond to the human CDPs, and a significant decrease of CD34+CD45RA+CD123+ cells, which include

GMDPs and MDPs. We can infer, therefore, that the MDP to CDP transition is blocked in CML patients.

DC deficiency is not restricted to CML, but is also regularly observed in acute myeloid leukemia (AML). Similar to CML, AML is mainly driven by chromosomal translocations leading to the formation of the oncogenes FLT3-ITD and AML1-ETO156. AML patients also showed drastic reduction in pDC and cDC numbers in their periphery, suggesting a decrease in

DC progenitors in the bone marrow157, 158. However, which developmental stage has been terminated still remains unknown. We suggest that by CD34+ cell compartment analysis, we will be able to elucidate the aberrant steps along DC differentiation.

There are also other diseases where DCs are specifically expanded. One such syndrome is the blastic plasmacytoid dendritic cell neoplasm (BPDCN)159. This disease is characteristic of their selective increase of a CD303+CD45RA+CD123hiCD4+CD56+ cell population160, which resembles the human pDCs. These cells have the capacity to produce IFNa and respond to IL-3

127 in vitro by increasing their survival and upregulating costimulatory molecules. Clinical manifestations are cutaneous “bruise-like” lesions and leukemic dissemination161. The genetic cause of BPDCN has been partially attributed to a dysregulated and activated NF-kB pathway162.

Interestingly, BPDCN-derived pDCs express the myeloid markers CD13 and CD33163, 164 on their surface and transcriptional analysis revealed higher resemblance to cDCs than pDCs160, which is indicative of a problem in pDC vs. cDC commitment. Although progenitor analysis in bone marrow has not been done, we believe that phenotypic characterization and isolation of

CD34+ cells will reveal the cellular stages of abnormal development as well as the molecular programs, including NF-kB signaling dysregulation, that induce BPDCN syndrome.

In conclusion, we propose that exhaustive phenotypic characterization of CD34+ cells in

DC-associated diseases together with a culture system that allows the investigation of DC potential, will enable us to investigate the transcriptional programs regulating specific DC developmental stages and will also provide a platform for interrogating potential therapeutic approaches in order to rescue DC-deficiency or expansion.

4. Clonal heterogeneity in marker-defined progenitors

The standard model of hematopoiesis explains that blood cell production begins from hematopoietic stem cells (HSCs), which then proceed through a series of developmental stages hierarchically organized according to their restricted developmental potential, to finally terminally differentiated cells.

Much of our understanding of this progressive process has been built on two main factors: (1) the identification of separate progenitors by the differential expression of surface markers, and (2) in vivo and in vitro culture systems that can measure their developmental

128 potential and their interdevelopmental relationship. For example, HSCs have been phenotypically characterized as CD34+CD38-CD90+ cells and placed at the apex of the hematopoietic tree based on their multipotency measured by in vitro and in vivo experiments146.

Multipotent progenitors (MPPs, CD34+CD38-CD90-), which are also able to produce all blood lineages, are placed downstream HSCs because of their lower self-renewal capacity and because

CD90+ HSCs can produce cells with CD34+CD38-CD90- phenotype, which resemble MPPs, but not in the opposite direction. Using this approach, old and new candidate progenitors have been placed on specific lineage pathways along the hematopoietic map, either upstream or downstream of each other, depending on their developmental potential difference and their progenitor-progeny relationship.

However, this approach fails to explain and place DC-restricted progenitors on the hematopoietic map. We have shown that myeloid progenitor CMP, and lymphoid progenitors

LMPPs and MLPs have the capacity to produce DCs in vitro and in vivo, as well as their intermediate progenitors, without giving rise to each other. This suggested to us that the marker- pure GMDPs, MDPs and CDPs were comprised of a mix of progenitors derived from both myeloid and lymphoid cells. In fact, when we cultured single cells from these progenitors, we were surprised to see the prevalent clonal heterogeneity within each population. Contrary to the traditional model, we also observed that the process of lineage commitment to a single cell type is not restricted to one single progenitor stage, but rather occurs gradually starting as soon as in the HSC/MPP stage. Furthermore, our clonal analysis showed that even clones with identical potential could have drastically different surface phenotype, which could explain why two phenotypically different progenitors had shared or overlapping potential.

129

Previous cultures determining the lineage potential of candidate progenitors have been evaluated primarily by bulk cultures. Although this method provided important information regarding the average power for a progenitor to produce certain lineages, it had failed, until today, to prove whether the resulting lineages were shared by one common cell or whether it was the combined product of two separate cells. Limiting dilution cultures have statistically approximated the shared clonality of separate lineages, but this too could not definitively demonstrate the shared origin.

Therefore, our clonal analyses show that the direct correlation of progenitor phenotype to developmental potential is unreliable, and that the previous developmental assays are only indicative of the overall developmental capacity of a population, and do not accurately trace the lineages for each of blood cell type.

So what parameters can be used to indicate and measure developmental status of progenitors? In order to draw the path for each lineage on the hematopoietic map, we propose to use the quantitative clonal output instead of marker-defined phenotype as an indicator of developmental status. We have shown that cell output, as well as the number of lineages it produces, can be used as a measure of their developmental distance from HSCs. The fewer cells a clone produces, the farther it is in the hematopoietic tree, and the fewer the number of lineages a clone produces the more committed it is. Therefore, we propose that by placing each clone in a multidimensional space using the number of cells each clone generated for each of the six lineages our culture supports, we will be able to delineate the developmental path not only for

DCs, but for all other cell types.

130

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137. Sanz, C. et al. Signaling and biological effects of glucagon-like peptide 1 on the differentiation of mesenchymal stem cells from human bone marrow. Am J Physiol Endocrinol Metab 298, E634-643 (2010).

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PROTOCOL Defining human dendritic cell progenitors by multiparametric flow cytometry Gaëlle Breton1, Jaeyop Lee2, Kang Liu2 & Michel C Nussenzweig1,3

1Laboratory of Molecular Immunology, The Rockefeller University, New York, New York, USA. 2Department of Microbiology and Immunology, Columbia University Medical Center, New York, New York, USA. 3Howard Hughes Medical Institute, Chevy Chase, Maryland, USA. Correspondence should be addressed to M.C.N. ([email protected]).

Published online 20 August 2015; doi:10.1038/nprot.2015.092

Human dendritic cells (DCs) develop from progressively restricted bone marrow (BM) progenitors: these progenitor cells include granulocyte, monocyte and DC progenitor (GMDP) cells; monocyte and DC progenitor (MDP) cells; and common DC progenitor (CDP) and DC precursor (pre-DC) cells. These four DC progenitors can be defined on the basis of the expression of surface markers such as CD34 and hematopoietin receptors. In this protocol, we describe five multiparametric flow cytometry panels that can be used as a tool (i) to simultaneously detect or phenotype the four DC progenitors, (ii) to isolate DC progenitors to enable in vitro differentiation or (iii) to assess the in vitro differentiation and proliferation of DC progenitors. The entire procedure from isolation of cells to flow cytometry can be completed in 3–7 h. This protocol provides optimized antibody panels, as well as gating strategies, for immunostaining of BM and cord blood specimens to study human DC hematopoiesis in health, disease and vaccine settings.

INTRODUCTION DCs are a heterogeneous population of immune cells that hematopoiesis in health, disease and vaccine settings are possi- are essential mediators of immunity and tolerance1–3. Although ble, as well as the exploration of their potential utility in cellular DCs share a common ability to process and present antigen immunotherapies. This can be facilitated by the use of this to naive T cells for the initiation of an immune response, they protocol, in which we describe flow cytometry assays to isolate are not all identical, and their unique abilities and phenotypes and characterize DC progenitors. are current areas of investigation. DCs from humans, which are best defined in the blood, are identified with BDCA markers. Development of the protocol BDCA-2(CD303)+ plasmacytoid DCs (pDCs) have the unique The study of human DC hematopoiesis has been hampered by the ability to rapidly produce abundant type I interferon (IFN) in absence of validated markers to identify and track progenitors. response to viral infection4. BDCA-1(CD1c)+ conventional DCs Human hematopoietic stem and progenitor cells (HSPCs) are Nature America, Inc. All rights reserved. Inc. Nature America, + 5,6

5 (cDCs) have been proposed to excel in CD4 T-cell priming . commonly defined by the expression of the cell surface protein Finally, BDCA-3(CD141)hi cDCs have the ability to capture dead CD34, as well as the non-expression of lineage antigens that cells and to cross-present exogenous antigen, offering a mecha- are present on mature leukocytes. These Lin− (lineage) CD34+ © 201 nism for priming CD8+ T cells specific for pathogens that do not HSPCs comprise only a small and variable fraction of human BM directly infect DCs7–12. cells (2–4%), CB cells (~1%) and peripheral blood (PB; <0.2%). Whereas DC development has been studied extensively in Human HSPCs have been further fractionated using common mice13, the origin of human DCs and their relation to mono- markers such as CD38, CD90, CD45RA, CD117 (stem cell cytes have been long debated. Our group has recently clarified the

pathway for human DC hematopoiesis and shown its sequential Bone marrow Blood Periphery origin from increasingly restricted but well-defined BM progeni- pDC tors14,15 (Fig. 1). Human granulocyte, monocyte and DC lineages pDC originate from a common progenitor, the GMDP. GMDPs develop HSC GMDP MDP BDCA-3hi cDC into a more restricted human MDP. MDPs give rise to monocytes CDP

and a CDP, which loses the potential to produce monocytes and DC progenitors pre-cDC BDCA-1+ cDC is restricted to produce the three major subsets of DCs. These committed DC progenitors reside in BM, as well as in cord blood 14 Granulocytes Monocytes (CB), but not in blood or lymphoid tissues . Finally, CDPs Monocytes give rise to pDCs, as well as to a circulating cDC precursor cell (pre-cDC). Indeed, pre-cDCs develop in the BM, travel through Figure 1 | Schematic view of human dendritic cell (DC) hematopoiesis. the blood and differentiate into the two subsets of cDCs in periph- DC hematopoiesis is initiated in the bone marrow (BM). A granulocyte, eral lymphoid organs. In studies of human volunteers injected monocyte and DC progenitor (GMDP) develops into a monocyte and with Fms-related tyrosine kinase 3 ligand (FLT3L), we showed DC progenitor (MDP). MDPs give rise to monocytes and a common DC progenitor (CDP), which loses the potential to produce monocytes. that human pre-cDCs are mobilized into the blood similarly CDPs give rise to plasmacytoid DCs (pDCs), as well as a circulating cDC 15 to the more differentiated DC subsets . Now that the lineage- precursor (pre-cDC). Pre-cDCs migrate from the BM through the blood committed progenitors and immediate precursors for human to the periphery to produce the two major subsets of conventional DCs have been identified, studies to further define human DC DCs (cDCs)—i.e., BDCA-1+ cDCs and BDCA-3hi cDCs.

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TABLE 1 | List of monoclonal antibodies used in this study.

Marker Fluorochrome Clone Host species and isotype Manufacturer Cat. no.

BDCA-2 (CD303) FITC AC144 Mouse IgG1 Miltenyi 130-090-510 CD116 FITC 4H1 Mouse IgG1 BioLegend 305906 BDCA-3 (CD141) PE AD5-14H12 Mouse IgG1 Miltenyi 130-090-514 CD135 PE 4G8 Mouse IgG1 BD Biosciences 558996 CD45 PE Texas Red HI30 Mouse IgG1 Life Technologies MHCD4517 HLA-DR PE Texas Red TÜ36 Mouse IgG2b Life Technologies MHLDR17 CD45RA PE Texas Red MEM-56 Mouse IgG2b Life Technologies MHCD45RA17 CD3 PE Texas Red S4.1 Mouse IgG2a Life Technologies MHCD0317 CD19 PE Texas Red SJ25-C1 Mouse IgG1 Life Technologies MHCD1917 CD14 PE Texas Red TüK4 Mouse IgG2a Life Technologies MHCD1417 BDCA-2 (CD303) PerCP-Cy5.5 201A Mouse IgG2a BioLegend 354210 CD66b PerCP-Cy5.5 G10F5 Mouse IgM BioLegend 305108 CD335 PerCP-Cy5.5 9E2 Mouse IgG1 BioLegend 331920 CD10 PerCP-Cy5.5 HI10a Mouse IgG1 BioLegend 312216 CD335 PE-Cy7 9E2 Mouse IgG1 BD Biosciences 562101 BDCA-3 PE-Cy7 M80 Mouse IgG1 BioLegend 344110 HLA-DR PE-Cy7 L243 Mouse IgG2a BioLegend 307616 CD11c PE-Cy7 3.9 Mouse IgG1 BioLegend 301608 CD33 PE-Cy7 WM53 Mouse IgG1 eBioscience 25-0338-42 CD38 PE-Cy7 HIT2 Mouse IgG1 BioLegend 303516 Nature America, Inc. All rights reserved. Inc. Nature America,

5 CD117 Brilliant Violet 421 104D2 Mouse IgG1 BioLegend 313216 BDCA-3 (CD141) Brilliant Violet 421 M80 Mouse IgG1 BioLegend 344114 © 201 CD117 Brilliant Violet 510 104D2 Mouse IgG1 BioLegend 313220 CD123 Brilliant Violet 510 6H6 Mouse IgG1 BioLegend 306022 CD45 V500 HI30 Mouse IgG1 BD Biosciences 560777 CD10 Brilliant Violet 605 HI10a Mouse IgG1 BioLegend 312222 CD20 Brilliant Violet 605 2H7 Mouse IgG2b BioLegend 302333 CD123 Brilliant Violet 650 6H6 Mouse IgG1 BioLegend 306019 CD45RA Qdot 655 MEM-56 Mouse IgG2b Invitrogen Q10069 CD14 Qdot 800 TüK4 Mouse IgG2a Invitrogen Q10064 BDCA-2 APC AC144 Mouse IgG1 Miltenyi 130-090-905 CD115 APC 9-4D2-1E4 Rat IgG1 BioLegend 347306 Clec9A Alexa Fluor 700 683409 Mouse IgG1 R&D Systems FAB6049N CD34 Alexa Fluor 700 581 Mouse IgG1 BioLegend 343526 BDCA-1 (CD1c) APC-Cy7 L161 Mouse IgG1 BioLegend 331520 CD3 BUV395 UCHT1 Mouse IgG1 BD Biosciences 563546 CD19 BUV395 SJ25-C1 Mouse IgG1 BD Biosciences 563549

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TABLE 2 | Antibody panels for human dendritic cell progenitors.

Panel #4, 13 Panel #5, Panel #1, 15 color Panel #2, 15 color Panel #3, 11 color color 14 color (culture Panel (identification) (characterization) (isolation) (culture output) proliferation) FITC CD116 CD116 CD116 BDCA-2 CFSE PerCP Cy5.5 CD66b/CD335/BDCA-2 CD66b/CD335/BDCA-2 CD66b/CD335/BDCA-2/CD10 CD66b CD66b PE CD135 CD135 CD135 BDCA-3 BDCA-3 PE Texas Red CD45 CD45RA CD3/CD19/CD14 HLA-DR HLA-DR PE-Cy7 BDCA-3 (antibody of choice) BDCA-3 CD335 CD335 APC CD115 CD115 CD115 — BDCA-2 Alexa Fluor 700 CD34 CD34 CD34 Clec9A Clec9A APC-Cy7 BDCA-1 BDCA-1 BDCA-1 BDCA-1 BDCA-1 BV421 CD117 BDCA-3 CD117 — — V500, BV510 CD123 CD117 CD123 CD45 CD45 BV605 CD10 CD10 — CD20 CD20 Qdot 655, BV650 CD45RA CD123 CD45RA CD123 CD123 Qdot 800 CD14 CD14 — CD14 CD14 Live/Dead Live/Dead blue Live/Dead blue — Live/Dead blue Live/Dead blue BUV395 CD3/CD19 CD3/CD19 — CD3 CD3

factor (SCF) receptor) and CD135 (FLT3 receptor)16–19. However, When working with scarce cells, the simultaneous analysis of all the combination of these markers does not separate DC four DC progenitors seems to be essential in order to provide a lineage–committed progenitors from multipotential progeni- comprehensive view of DC hematopoiesis. Nature America, Inc. All rights reserved. Inc. Nature America,

5 tors. Moreover, it has been reported that a small fraction of cells The proven multicolor panels presented in this protocol are that lack the expression of CD34 have progenitor potential20. based on our original studies14,15 with modifications enabling − −

© 201 These Lin CD34 cells probably represent precursor cells that simultaneous assessment of all DC lineage–committed progeni- have lost CD34 expression but that are still too immature to tors. As the common feature of progenitors that are capable of express lineage markers. Therefore, CD34, as well as the afore- developing into DCs is surface expression of CD117 and CD135, mentioned markers, are not enough to further separate HSPCs, the panel to define them includes the following: (i) antibodies and additional markers are required to identify and isolate DC specific to CD117 and CD135, as well as CD34, CD123, CD115, lineage–committed progenitors. CD116 and CD45RA; (ii) antibodies specific to CD3 (T cell), CD19 As hematopoiesis relies on instructive cues from the BM niche (B cell), CD14 (monocytes), CD335 (natural killer (NK) cell), such as hematopoietins, we hypothesized that the lineage potential CD66b (granulocytes) and CD10 (lymphoid progenitors) to of a progenitor might be determined by the set of growth factor exclude specific lineages (Lin−); (iii) antibodies specific to BDCA-1, receptors that it expresses. We showed that multipotential pro- BDCA-2 and BDCA-3 to exclude differentiated DCs (BDCA−); genitors such as myeloid and lymphoid progenitors were hetero- (iv) a viability dye to exclude dead cells; and (v) an optional geneous, and they could be further fractionated using antibodies antibody specific to CD45 to visualize hematopoietic cells in against receptors for granulocyte–macrophage colony-stimulating nonhematopoietic tissues or in vitro cultures. All antibodies factor (GM-CSFR; CD116), macrophage colony-stimulating (Table 1) and fluorochrome combinations (Table 2) are com- factor (M-CSFR; CD115) and interleukin-3 (IL-3R; CD123) mercially available, and experiments can be run on commonly (refs. 14,15). By using two slightly different antibody panels, we available flow cytometers and flow sorters that are capable of identified the following: (i) the human DC lineage–committed multicolor analysis (see configurations in Tables 3 and 4). GMDP, MDP and CDP in the Lin− CD34+ fraction by combin- Defining human DC progenitors also required that we develop ing antibodies against CD45RA, CD116, CD115 and CD123, as an efficient culture assay to study the ability of the progenitors to well as CD38 (ref. 14), and (ii) the human pre-cDC, which was proliferate and differentiate in response to hematopoietic growth found in the Lin− CD34− fraction and identified on the basis of factors. Through systematic testing of different combinations of the expression or lack of CD45RA, CD116, CD115 and CD123, cytokines, as well as the use of mouse stromal cell lines, we showed as well as CD135 and CD117 (ref. 15). These progenitors are that the in vitro differentiation of CD34+ HSPCs on MS-5 stromal rare, which presents a challenge for the isolation of these cells. cells in addition to FLT3L, SCF and GM-CSF cytokines (referred

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TABLE 3 | Configuration of LSR II. TABLE 4 | Configuration of ARIA III.

Detector Long-pass filter Band-pass filter Fluorochrome Detector Long-pass filter Band-pass filter Fluorochrome Blue laser (488 nm) Blue laser (488 nm) A 685LP 695/40 PerCP-Cy5.5 A 655LP 695/40 PerCP-Cy5.5 B 505LP 530/30 FITC, CFSE B 502LP 530/30 FITC C 488/10 SSC C — 488/10 SSC Yellow/green laser (561 nm) Yellow/green laser (561 nm) A 750LP 780/60 PE-Cy7 A 735LP 780/60 PE-Cy7 D 600LP 610/20 PE Texas Red D 600LP 610/20 PE Texas Red E — 582/15 PE E — 582/15 PE Red laser (633 nm) Red laser (640 nm) A 735LP 780/60 APC-Cy7 A 735LP 780/60 APC-Cy7 B 690LP 710/50 Alexa Fluor B 690LP 730/45 Alexa Fluor 700 700 C — 660/20 APC C — 660/20 APC Violet laser (405 nm) Violet laser (407 nm) A 635LP 655/20 Qdot 655, A 600LP 660/20 Qdot 655 BV650 B 502LP 510/50 BV510 B 595LP 605/20 BV605 C — 450/40 BV421 C 505LP 525/50 V500, BV510 D — 450/50 BV421 affect the composition of the progenitor compartment, CB cells UV laser (355 nm) can also be examined. As CB can be collected quite easily and Nature America, Inc. All rights reserved. Inc. Nature America,

5 A 735LP 780/60 Qdot 800 cryopreserved, it is a good source of human progenitors. When these populations are purified and cultured in MS5+FSG, CB B 410LP 450/50 Live/Dead blue

© 201 progenitors showed a differentiation potential similar to that C — 379/28 BUV395 exhibited by their BM counterparts14,15.

Identification and characterization of DC progenitors. As to as MS5+FSG herein) support the development of multiple described, human DC progenitors can be identified by the lack lineages, including all three major human DC subsets. The resulting of lineage and BDCA markers and by combining CD123, CD34, DC subsets resemble their blood counterparts both in gene CD115, CD116, CD45RA, CD117 and CD135 markers (Table 2; expression and function14. panel #1). As shown in Figure 2b for BM and Figure 2c for CB, the following cells can be distinguished: Experimental design In this protocol, we give a flow cytometry–based workflow for • Cells with the phenotype Lin−BDCA−CD123−/loCD34+CD117+ (i) simultaneous detection of DC progenitors (Table 2, panel #1 CD135+CD116−CD115−CD45RA+ correspond to the GMDPs and Fig. 2), as well as their characterization (Table 2, panel #2 with the ability to generate granulocytes, monocytes and DCs. and Fig. 3); (ii) isolation of DC progenitors to enable in vitro • Cells with the phenotype Lin−BDCA−CD123−/loCD34+CD117+ differentiation (Table 2, panel #3 and Fig. 4); and (iii) evaluation CD135+CD116− CD115+CD45RA+ represent the MDPs, which of DC progenitor in vitro differentiation (Table 2, panel #4 and have the potential to give rise to monocytes and DCs but not to Fig. 5) and proliferation (Table 2, panel #5 and Fig. 6). The pro- granulocytes. cedure also covers various preparatory steps—isolation, freezing • Cells with the phenotype Lin−BDCA−CD123hiCD34+CD117+C and thawing of mononuclear cells (MNCs) and hematopoietic D135+CD116+CD115−CD45RA+ correspond to the CDPs with differentiation on stromal cells. the capacity to give rise to all DCs but not to monocytes and granulocytes. Source tissue. The procedure can be performed on BM or CB • Pre-cDCs have the phenotype Lin−BDCA−CD123−/loCD34− samples. BM samples are typically obtained from elderly human CD117+CD135+CD116+CD115−CD45RA+ and are restricted to donors; however, as age-related developmental changes may the two subsets of cDCs.

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a Stain with identification Analyze on LSR II using Preparation of BM or CB sample 1 2 3 antibody panel (panel #1) identification gating strategy

b 250 K BM 250 K 250 K c 250 K CB 250 K 250 K 200 K 200 K 200 K 200 K 200 K 200 K 150 K 150 K 150 K 150 K 150 K 150 K 100 K 100 K 100 K 100 K 100 K 100 K 50 K 50 K 50 K 50 K 50 K 50 K SSC-A SSC-W SSC-A 0 0 SSC-A 0 SSC-A 0 SSC-W 0 0 0 K K K K K 0 K K K K K 0103 104 105 0 K K K K K 0 K K K K K 0103 104 105 50 50 50 50 100 150 200 250 100 150 200 250 100 150 200 250 100 150 200 250 FSC-A SSC-A Live/Dead FSC-A SSC-A Live/Dead

250 K 250 K 105 105 105 105 200 K 4 200 K 4 10 4 104 150 K 10 150 K 10 3 3 100 K 103 10 100 K 103 10 50 K 0 0 50 K 0 0 CD10 BDCA-3 SSC-A 0 CD10 BDCA-3 SSC-A 0 0 103 104 105 0103 104 105 0103 104 105 0103 104 105 0103 104 105 0103 104 105 CD3/CD19 CD14 BDCA-1 CD3/CD19 CD14 BDCA-1

105 105

104 104

3 103 10 0 0 CD66b/CD335 BDCA-2 CD66b/CD335 BDCA-2 3 4 5 0 10 10 10 0 103 104 105 CD123 CD123

105 105 105 105 4 4 104 104 10 10

3 3 103 103 10 10 0 0 0 0 CD117 CD117 CD117 CD117 0103 104 105 0103 104 105 0103 104 105 0103 104 105 CD34 CD34 CD34 CD34

105 105 105 105 105 105

104 104 104 104 104 104

103 103 103 103 103 103 0 0 0 0 0 0 CD116 CD116 CD116 CD116 CD116 CD116

0103 104 105 0103 104 105 0103 104 105 0103 104 105 0103 104 105 0103 104 105 CD135 CD135 CD135 CD135 CD135 CD135

5 5 5 5 5 5 Nature America, Inc. All rights reserved. Inc. Nature America, 10 10 10 10 10 10

5 4 4 4 104 104 104 10 10 10 MDP CDP 3 3 MDP 3 CDP 3 3 3 10 10 10 10 10 GMDP 10 0 0 0 0 0 GMDP 0 © 201 CD115 CD115 CD115 CD115 CD115 CD115

0103 104 105 0103 104 105 0103 104 105 0103 104 105 0103 104 105 0103 104 105 CD45RA CD45RA CD45RA CD45RA CD45RA CD45RA

250 K 250 K 200 K 200 K 150 K 150 K

A 100 K A 100 K 50 K pre-cDC 50 K pre-cDC SSC- 0 SSC- 0 0103 104 105 0103 104 105 CD135 CD135 d GMDP MDP CDP pre-cDC e – CD115 BM CB 105 105

104 104 5

CD45 103 MDP 103 MDP CD11 0 GMDP 0 GMDP CD45RA 0103 104 105 0103 104 105 0103 104 105 0103 104 105 0103 104 105 0103 104 105

Figure 2 | Gating strategy for the identification of human DC progenitors. (a) Workflow for the identification of DC progenitors. (b,c) Human BM (b) and CB (c) MNCs were stained as indicated in Table 2 (panel #1). Progenitors are identified by first gating on live cells (FSC-A versus SSC-A), and then on singlets (SSC-A versus SSC-W) and live cells (Live/Dead blue versus SSC-A). Lin− BDCA− cells are obtained by successively gating out CD3+, CD19+ cells (CD3, CD19 versus SSC-A), CD14+, CD10+ cells (CD14 versus CD10), BDCA-1+, BDCA-3hi cells (BDCA-1 versus BDCA-3) and CD66b+, CD335+, BDCA-2+ cells (CD66b, CD335, BDCA-2 versus CD123). Lin−BDCA−CD123−/lo and Lin−BDCA−CD123hi populations are further subgated on the basis of the expression of CD34 versus CD117, CD135 versus CD116 and CD45RA versus CD115. In these gated populations, GMDPs are CD123−/loCD34+CD117+CD135+CD116−CD115−CD45RA+; MDPs are CD123−/loCD34+CD117+CD135+CD116−CD115+CD45RA+; CDPs are CD123hiCD34+CD117+CD135+CD116+CD115−CD45RA+; and pre-cDCs are CD123−/lo CD34−CD117+CD135+CD116+CD115−CD45RA+SSC-Alo. FSC, forward scatter; SSC, side scatter. (d) Expression of CD45 by GMDPs, MDPs, CDPs and pre-cDCs from human BM. Histograms show the expression of stained (black line) and unstained (gray line) cells. (e) FMO control for CD115 staining used to discriminate between GMPs and MDPs in CB.

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Figure 3 | Characterization of human DC progenitors. (a) Workflow a Preparation of CB or BM sample for the characterization of DC progenitors. (b) Expression of CD11c, 1 HLA-DR, CD38 and CD33 by GMDPs, MDPs, CDPs and pre-cDCs from human BM. DC progenitors are defined as in Figure 2b. Histograms show Stain with characterization expression (x axis) and number of cells (y axis) of stained (black line) 2 antibody panel (panel #2) and unstained (gray line) cells. Analyze on LSR II using 3 identification gating strategy Of note, we do not include isotype controls in our immunostain- ing, as the use of fluorescent isotype controls is not completely b GMDP MDP CDP pre-cDC accurate, and because we have ascertained that the monoclonal BM

antibodies are specific by carefully titrating them and using CD11c fluorescence minus one (FMO) controls for gating. In nonhematopoietic tissues, we recommend using CD45 to 3 4 5 3 4 5 3 4 5 3 4 5 visualize hematopoietic progenitors, as it may be challenging to 010 10 10 010 10 10 010 10 10 010 10 10 identify these rare cells among other cells such as autofluorescent macrophages. As shown in Figure 2d, most, if not all, DC progeni- HLA-DR tors express CD45. Therefore, one channel in antibody panel #1 remains open to the potential use of CD45. 0103 104 105 0103 104 105 0103 104 105 0103 104 105 As DC progenitors are rare cells in both BM and CB, their iden- tification is a matter of carefully defining the gates separating pos- CD38 itive and negative cells. As shown in Figure 2b,c, the first step in the identification of human DC progenitors consists of excluding 0103 104 105 0103 104 105 0103 104 105 0103 104 105 dead cells and debris by scatter gating. Live cells are then gated for singlets (side scatter(SSC)-W versus SSC-A; W = width, A = area), CD33 and dying cells are excluded on the basis of the uptake of the Live/Dead blue stain. The next step consists in the sequential exclusion of all lineage (CD3, CD19, CD14, CD66b, CD335 and 0103 104 105 0103 104 105 0103 104 105 0103 104 105 CD10)- and BDCA (BDCA-1, BDCA-2 and BDCA-3)-expressing cells. To have a better resolution in our gates, we avoid as much phycoerythrin (PE) channel for this purpose, as PE is bright, and as possible the use of a ‘dump’ channel containing all antibod- most if not all antibodies are available in this color. Because CD135 ies marking all excluded populations. Nonetheless, if the use of staining is crucial in order to identify DC progenitors and because a ‘dump’ channel is considered to accommodate an additional only CD135 PE works well in our hands, we decided to open the antibody of interest (as done in Table 2, panel #3), one should PE-Cy7 channel. PE-Cy7 is a tandem dye that can degrade in the carefully titrate each antibody separately before combining presence of light and heat, and it will then emit in the parent dye Nature America, Inc. All rights reserved. Inc. Nature America,

5 them to avoid any compensation issues and difficulties in discrim- channel (PE). However, by avoiding the exposure of samples to inating between negative cells and cells with low expression. The light and heat, this problem can be largely avoided. The gating

© 201 use of CD123 allows clear separation of cells that express a high procedure is the same as that previously described (Fig. 2b,c), level of CD123 (CD123hi) from cells that express no or a low level and the expression of CD11c, HLA-DR, CD38 and CD33 on of CD123 (CD123lo cells). CDPs are found within the CD123hi DC progenitors is shown (Fig. 3b). HLA-DR, whose expression cells by sequentially gating on CD135+, CD116+, CD45RA+ and is neither specific nor identical within the DC subsets, is also CD115− cells. The CD123lo gate is further analyzed by gating on expressed by DC progenitors. CD11c, which is weakly expressed CD34+ CD117+ (CD34+) cells or CD34− CD117+ (CD34−) cells. on cDCs and completely lacking on pDCs, is upregulated dur- Pre-cDCs are found in the CD34− gate. Indeed, we first gate on ing DC development. CD38, which was initially used to identify cells that express CD135, CD116 and CD45RA but do not express multipotential progenitors16, as well as DC lineage–committed CD115, and then exclude SSC-A-high cells, as these cells contain progenitors14, is also expressed by pre-cDCs. Because CD38 is some monocyte precursors15. In contrast, the CD34+ population expressed by all hematopoietic progenitors, we decided not to contains both GMDPs and MDPs. These progenitors are all use it in our antibody panels and focus more on hematopoietin CD135+, CD116− and CD45RA+; expression of CD115 allows the receptors that allow further separation of these cells. CD33, which separation of GMDPs (CD115− cells) from MDPs (CD115+ cells). is highly specific to the hematopoietic compartment, is expressed Even though the CD115 staining is very faint and it is not possible by all DC progenitors. to get a clear-cut separation of positive and negative cells, the use of FMO control allows for proper gating on CD115+ cells (Fig. 2e). Isolation of progenitors. We developed our flow cytometry Moreover, if we consider that lineage commitment occurs when strategy to identify DC progenitors on a 16-color flow cytometer the capacity to give rise to one cell type, e.g., monocyte and DC, (BD LSR II with blue, yellow/green, red, violet and UV lasers; see is increased and the capacity to produce another cell type, e.g., configuration in Table 3). To isolate DC progenitors, we adapted granulocyte, is lost, dim CD115 expression is sufficient to separate our panel to maximize the capacities of an 11-color cell sorter cells with GMDP potential from cells with MDP potential. (BD Aria III with blue, yellow/green, red and violet lasers; see To characterize DC progenitors, we use a related combination configuration in Table 4). This was done in part by restricting of antibodies (Table 2, panel #2). Panel #2 leaves a channel open the number of channels used for lineage markers and by omitting to accommodate an additional antibody of interest. We prefer the the Live/Dead blue stain to exclude dead cells, relying only on

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| a c Figure 4 Gating strategy for the isolation 250 K 250 K BM 105 of human DC progenitors. (a) Workflow for the 200 K 200 K Preparation of BM or CB sample 104 isolation of DC progenitors. (b) Gate hierarchy 1 150 K 150 K 100 K 100 K 103 for sorting. (c) Human BM MNCs were stained as Stain with isolation 50 K 50 K 0 antibody panel (panel #3) SSC-A

2 BDCA-3 SSC-A 0 0 indicated in Table 2 (panel #3). Progenitors are 3 4 5 0 K K K K K 0103 104105 0 10 10 10 Sort on ARIA III using 50 identified by first gating on live cells 3 isolation gating strategy 100 150 200 250 CD3/CD19/ BDCA-1 (FSC-A versus SSC-A). Lin− BDCA− cells are FSC-A CD14 + Culture or transcriptome obtained by successively gating out CD3 , analysis of sorted populations 4 105 + + CD19 , CD14 cells (CD3, CD19 and CD14 versus 104 + hi SSC-A), BDCA-1 , BDCA-3 cells (BDCA-1 versus 3 b 10 BDCA-3) and CD66b+, CD335+, CD10+, BDCA-2+ 0

Gate hierarchy CD66b/CD335 CD10/BDCA-2 cells (CD66b, CD335, CD10, BDCA-2 3 4 5 All events 010 10 10 − − −/lo versus CD123). Lin BDCA CD123 and 1 Live cells CD123 Lin−BDCA−CD123hi populations are further 2 CD3- CD19- CD14- 3 5 5 subgated on the basis of the expression BDCA-3hi- BDCA-1- 10 10 4 CD123-/int 104 104 of CD34 versus CD117, CD135 versus 5 34+ 117+ 3 3 CD116 and CD45RA versus CD115. In these 6 34+ 117+ 135+ 116– 10 10 0 0 −/lo 7 GMDP gated populations, GMDPs are CD123 CD117 7 MDP 3 4 5 CD117 CD34+CD117+CD135+CD116−CD115−CD45RA+; 0 10 10 10 0 103 104 105 5 34– 117+ CD34 CD34 MDPs are CD123−/loCD34+CD117+CD135+ 6 34– 117+ 135+ 116+ − + + hi + 7 115– 45RA+ CD116 CD115 CD45RA ; CDPs are CD123 CD34 5 5 5 8 pre-cDC 10 10 10 CD117+CD135+CD116+CD115−CD45RA+; and 4 4 4 4 CD123hi 10 10 10 3 3 pre-cDCs are CD123−/loCD34−CD117+ 5 123hi 34+ 117+ 10 103 10 CD135+CD116+CD115−CD45RA+SSC-Alo. 6 123hi 135+ 116+ 0 0 0 7 CDP CD116 CD116 CD116 FSC, forward scatter; SSC, side scatter. 0103 104105 0103 104105 0 103 104105 CD135 CD135 CD135

105 105 105 exclusion by scatter gating (Table 2, panel #3). Moreover, the gating 104 104 104 103 103 MDP 103 hierarchy while sorting is limited to a maximum of an eight-gate 0 0 GMDP 0 CDP CD115 CD115 CD115 sort depth. This limit is intrinsic to the current generation of BD 3 4 5 3 4 5 0 10 10 10 0 103 104 105 0 10 10 10 Aria III sorter hardware and firmware. To limit our gating strategy CD45RA CD45RA CD45RA to eight steps (Fig. 4b), we also removed the step to gate out the sin- 250 K glets (SSC-W versus SSC-A). The gating strategy to identify and sort 200 K 150 K all DC-defined progenitors simultaneously is shown in Figure 4c. 100 K pre-cDC The sorted cells are suitable for various downstream applications 50 K SSC-A 0 such as cell culture or gene expression profiling. 0 103 104 105

Nature America, Inc. All rights reserved. Inc. Nature America, CD135 5 Composition and proliferative capacity of in vitro– differentiated cells. Hematopoietic differentiation can be greater proliferative capacity than DC immediate precursors, i.e., © 201 induced in vitro by coculture of hematopoietic progenitors on pre-cDCs. Moreover, BDCA-1+ and BDCA-3hi cDCs differentiate mouse BM stromal cells plus cytokines. To assess the differentia- as early as 12 h from CDP and pre-cDC progenitors. tion potential of different hematopoietic progenitors and their multilineage or restricted potential, we developed an additional Applications of the protocol antibody panel (Table 2, panel #4). BM CD34+ HSPCs cultured We believe that the protocol described here has multiple applica- in MS5+FSG for 7 d can produce T cells (CD3+), B cells (CD20+), tions and a wide interdisciplinary scope including (i) further defi- NK cells (CD335+), granulocytes (CD66b+), monocytes (CD14+) nition of human DC hematopoiesis in health, disease and vaccine and all three major subsets of DCs, that is, pDCs and both subsets settings, as well as (ii) evaluating DC progenitors’ applications in of cDCs (Fig. 5b). Human CD45 antibody must be included in therapeutic approaches. this panel, as it discriminates between human cells and mouse Our protocol can be used as an initial tool to analyze normal or stromal cells, which are not entirely gated out by scatter analysis. abnormal immunophenotypic changes during DC development in Of note, BDCA markers are often used to classify human DCs. humans. For instance, it can facilitate the study of diseases involv- However, they are not strictly restricted to DCs; we therefore used ing aberrant DC hematopoiesis such as chronic myelogenous them in combination with Clec9A for BDCA-3hi cDCs, CD14 for leukemia (CML)21 or a newly identified form of primary immu- BDCA-1+ cDCs and CD123 for BDCA-2+ pDCs. nodeficiency characterized by recurrent infections and dissemi- To examine the proliferative potential of DC progenitors, nated bacille Calmette-Guérin (BCG) infection after vaccination we purified them from CB and stained them with carboxy- in patients lacking GATA2 (ref. 22) and interferon regulatory factor fluorescein diacetate succinimidyl ester (CFSE) before culturing in 8 (IRF8)23. Because of their capacity to elicit and regulate immune MS5+FSG. By using a 14-color antibody panel (Table 2, panel #5) responses, DCs are essential to any effort for vaccine development. that includes CFSE, we were able to track the cells that were Therefore, our protocol provides important tools to design and expanding in culture by CFSE dilution. As shown in Figure 6b,c, monitor and potentially improve vaccines that rely on DCs. all progenitors are expanding in culture, even though DC lineage– Finally, the methods described above can support the committed progenitors—GMDP, MDP and CDP cells—have a exploration of new cellular therapeutic approaches based on

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a b 250 K BM 250 K 250 K 250 K 200 K 200 K 200 K 200 K 150 K 150 K 150 K 150 K Preparation of BM or CB sample 100 K 100 K 100 K 100 K 1 50 K 50 K 50 K 50 K SSC-W SSC-A SSC-A 0 0 0 SSC-A 0 3 4 5 3 4 5 Stain with isolation 0 K K K K K 0 K K K K K 010 10 10 0110 0 10 50 50 2 antibody mix (panel #3) 100 150 200 250 100 150 200 250 Live/Dead CD45 FSC-A SSC-A Sort on ARIA III using 250 K 250 K 250 K 250 K 3 isolation gating strategy 200 K 200 K 200 K 200 K 150 K 150 K 150 K 150 K Culture the sorted fractions on 100 K 100 K 100 K 100 K 4 MS-5 + FLT3L + SCF + GM-CSF 50 K 50 K 50 K 50 K Granulocyte SSC-A 0 SSC-A 0 SSC-A 0 SSC-A 0 3 4 5 3 4 5 3 4 5 3 4 5 Stain with culture output 0 10 10 10 010 10 10 010 10 10 0 10 10 10 5 antibody panel (panel #4) CD3 CD20 CD335 CD66b + 105 105 Monocyte 105 BDCA-2 pDC

Analyze on LSRII using 4 hi 10 BDCA-3 cDC 4 4 6 culture output gating strategy 10 10 103 103 103 0 0 0 +

Clec9A CD14 BDCA-1 cDC CD123 0 103 104 105 0103 104 105 0103 104 105 BDCA-3 BDCA-1 BDCA-2

Figure 5 | Gating strategy for culture output after hematopoietic differentiation on MS-5 stromal cells of human DC progenitors. (a) Workflow for isolation of DC progenitors. (b) CD34+ HSPCs from BM were isolated as in Figure 3c, and then cultured in MS5+FSG for 7 d. Culture output was assessed by flow cytometry using antibody panel #4, as indicated in Table 2. Cells developing in culture are identified by first gating on live cells (FSC-A versus SSC-A), and then on singlets (SSC-A versus SSC-W) and live cells (Live/Dead blue versus SSC-A). CD45+Lin− cells are obtained by successively gating on CD45+ cells (CD45 versus SSC-A) and gating out CD3+ cells (CD3 versus SSC-A), CD19+ cells (CD19 versus SSC-A) and CD335+ cells (CD335 versus SSC-A). Flow cytometry plots show phenotype of output cells, including granulocytes (brown), BDCA-3hi cDCs (red), BDCA-1+ cDCs (blue), monocytes (orange) and BDCA-2+ pDCs (green).

DC progenitors including stem cell therapies focused on drug candidates. Of note, therapeutic approaches based on embryonic stem cells and induced pluripotent stem cells. The human DC progenitors may require the development of a composition and the properties of the cells that are produced nonxenogeneic culture system; therefore, it would be good after in vitro differentiation of progenitors will differ considerably to explore the feasibility of expanding DC progenitors in an depending on the culture condition, and therefore this system entirely human coculture system using either umbilical vein can be used to evaluate DC progenitors to identify novel endothelial cells for CB progenitors or BM stromal cells for

Nature America, Inc. All rights reserved. Inc. Nature America, regulators of DC hematopoiesis or to test the toxicity of new BM progenitors. 5 © 201 MATERIALS REAGENTS • EDTA, 0.5 M (Life Technologies, cat. no. 15575) Cell samples • ACK lysing buffer (Life Technologies, cat. no. A10492) • Human samples can be obtained from national blood centers for CB • Mitomycin C (Sigma-Aldrich, cat. no. M4287) ! CAUTION Avoid contact and from local hospitals for BM ! CAUTION Universal precautions must and inhalation. Wear gloves, safety glasses and a lab coat. be taken while manipulating human specimens, and all experiments • DMSO (Sigma-Aldrich, cat. no. 154938) must be carried out in at least class II biological safety cabinets and • Trypan blue stain 0.4% (wt/vol) (Life Technologies, cat. no. 15250-061) using appropriate protection equipment. ! CAUTION Parental informed • Carboxy-fluorescein diacetate succinimidyl ester (CFSE; Life Technologies, consent (CB) or patient informed consent (BM) must be obtained before cat. no. C1165) collection, and protocols must conform to your Institutional Review • Recombinant human SCF ligand (SCF; R&D systems, cat. no. 255-SC-050/CF) Board (IRB)’s guidelines. IRB-approved protocols usually require that the • Human FLT3 ligand (FLT3L; Celldex Therapeutics, special order) samples be anonymous and fully deidentified at the site of collection • Leukine (sargramostim) as a human GM-CSF (Genzyme) by hospital personnel. M CRITICAL Leukine is a prescription drug that must to be ordered Cell culture through a pharmacy. • Dulbecco’s PBS 1× (DPBS; Life Technologies, cat. no. 14190) • Human serum albumin (HSA; Sigma-Aldrich, cat. no. A9511) • RPMI-1640 (Life Technologies, cat. no. 21870-076) • MS-5 mouse bone marrow stromal cell line (Creative Bioarray, • A-MEM 1× (Life Technologies, cat. no. 12561-056) cat. no. CSC-C2763) ! CAUTION The cell lines used in your research • FBS, heat inactivated (Life Technologies, cat. no. 16140-071) should be regularly checked to ensure that they are authentic and that M CRITICAL We recommend testing several lots of serum for culture they are not infected with mycoplasma. optimization and then using the same lot for an entire series of experiments. • Benzonase nuclease, 10 kU, 25 U/Ml (Novagen, cat. no. 70664-3) In our case, we always test three or four lots for hematopoietic differentiation • Heparin, cell culture grade (Sigma-Aldrich, cat. no. H3149) on MS-5 stromal cells and select the one that supports the development • Bleach ! CAUTION Bleach is an irritant. Avoid contact with eyes, skin of CD34+ HSPCs with an overall better yield for DCs. and clothing. • Antibiotic-antimycotic solution (Cellgro, cat. no. 30-004-CI) Plasticware • Collagenase type IV (Sigma-Aldrich, cat. no. C-5138) • 50-ml conical tubes (BD Falcon, cat. no. 352098) • Ficoll-Paque plus (GE Healthcare, cat. no. 17-1440-03) • 15-ml conical tubes (BD Falcon, cat. no. 352097) • Trypsin-EDTA, 0.05% (Life Technologies, cat. no. 25300-054) • 5-ml polypropylene round-bottom tube (BD Falcon, cat. no. 352063)

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Figure 6 | Proliferative capacity of human DC a b c progenitors after hematopoietic differentiation d = 0.5 d = 1.5 d = 3 d = 5 1,000 Preparation of BM or CB sample CB 800 on MS-5 stromal cells. (a) Workflow for culture 1 600 GMDP GMDP 400 d5 proliferation of DC progenitors. (b) GMDPs, Stain with isolation 200 MDPs, CDPs and pre-cDCs were purified from CB, 2 antibody mix (panel #3) 0 4,000 as shown in Figure 3c, labeled with CFSE, and 3,000 Sort on ARIA III using MDP isolation gating strategy MDP 2,000 cultured in MS5+FSG for 0.5, 1.5, 3 and 5 d; 3 progenitors input d5 3 proliferation was assessed by flow cytometry 1,000 0 Stain with CFSE using antibody panel #5, as indicated in Table 2. 4 2,000 Flow cytometry plots show gated culture-derived 1,600 Culture the sorted fractions on CDP 1,200 CDP cells, as defined in Figure 5b: CD45+ cells (gray), MS-5 + Flt3l + SCF + GM-CSF hi 800 d3 5 BDCA-3 cDCs 400 BDCA-1+ cDCs (blue) and BDCA-3hi cDCs (red). BDCA-1+ cDCs 0 Stain with culture proliferation 105 + + 4 CD45 cells 25 (c) Graphs showing the output of BDCA-1 cDCs 6 antibody panel (panel #5) 10 20 3 Pre-cDC (blue) and BDCA-3hi cDCs (red) for 103 GMDPs (5 d), 10 Cell number output per 10 15 Pre- Analyze on LSRII using 0 10 cDC d3 5 MDPs (5 d), CDPs (3 d) and pre-cDCs (3 d) BDCA-3 3 4 5 7 culture output gating strategy 0 10 10 10 0 cultured in MS5+FSG, as illustrated in Figure 6b. CFSE

• 5-ml polystyrene round-bottom tube (BD Falcon, cat. no. 352052) • Cryo 1 °C freezing container (Nalgene, cat. no. 5100-0001) • 5-ml polystyrene round-bottom tube with cell-strainer cap (BD Falcon, • LN2 storage systems (Thermo Scientific, cat. no. 7404) cat. no. 352235) • BD LSR II flow cytometer running DIVA 8.0, or equivalent (BD Biosciences, • 25-ml serological pipette (Costar, cat. no. 4489) special order). Our LSR II is equipped with four lasers with a configuration • 10-ml serological pipette (Costar, cat. no. 4488) allowing 16-color, 18-parameter flow cytometry (Table 3) • 5-ml serological pipette (Costar, cat. no. 4487) • BD Aria III cell sorter running DIVA 8.0, or equivalent (BD Biosciences, • Transfer pipette (Sarstedt, cat. no. 86.1171.001) special order). Our Aria III is equipped with three lasers with a configura- • Microcentrifuge tubes (1.5 ml; Eppendorf, cat. no. 022363204) tion allowing 11-color, 13-parameter flow cytometry (Table 4) • Cryogenic screw-cap micro tubes (2-ml; Sarstedt, cat. no. 72.694.006) • BD FACSDiva V8.0.1 software (BD Biosciences) for data acquisition • Micro tubes with screw caps, conical bottom (1.5-ml; RPI Corp, • FlowJo software V9.8.2 (TreeStar) for data analysis cat. no. 144500) REAGENT SETUP • Titertube micro test tubes (Bio-Rad, cat. no. 223-9391) Heparin Dissolve heparin in PBS at a final concentration of 1,000 U/ml. • 96-well micro-well plates V-bottom without lid (Nunc, cat. no. 249570) Filter this solution with a 0.2-Mm syringe filter and divide the solution • 96-well micro-well plates lid (Nunc, cat. no. 263339) into aliquots in a 5-ml polypropylene round-bottom tube. Store the • 96-well tissue culture plates flat-bottom (BD Falcon, cat. no. 357072) heparin at 4 °C for up to 2 years. • 10-cm tissue culture dishes (BD Falcon, cat. no. 353003) Collagenase solution Dissolve 100 mg of collagenase type IV in 50 ml • 15-cm tissue culture dishes (BD Falcon, cat. no. 353025) of RPMI-1640 to obtain a 2 mg/ml collagenase stock solution. Filter this • 150-cm2 tissue culture flasks straight neck with vented cap (BD Falcon, solution with a 0.2-Mm syringe filter. Store the solution in 10-ml aliquots cat. no. 355001) at −20 °C for up to 1 year. • 10-ml syringe (BD Falcon, cat. no. 309604) Freezing medium Prepare freezing medium by adding 10% (vol/vol) • Syringe filter with a 0.2-Mm membrane (Pall, cat. no. 4612) Nature America, Inc. All rights reserved. Inc. Nature America, DMSO in FBS on the day of use. • Cell strainer, 100 Mm (BD Falcon, cat. no. 352360) 5 • Cell strainer, 70 Mm (BD Falcon, cat. no. 352350) Thawing medium Prepare thawing medium by adding 10% (vol/vol) • Precision specimen container, 4 ounce, positive seal indicator (Covidien, FBS in RPMI-1640, and store it at 4 °C for up to 4 weeks.

© 201 cat. no. 2200SA) CFSE stock CFSE is resuspended in DMSO at a concentration of 5 mM. Flow cytometry reagents Store the stocks in aliquots of 50 Ml in Eppendorf tubes at −80 °C for up to • Antibodies (Table 1) M CRITICAL Each antibody must be titrated 1 year. Each batch of CFSE should be titrated before use in the procedure before use (see Reagent Setup) ! CAUTION All antibodies must be kept at (Box 1) on cells to determine the optimal concentration for labeling 4 °C and protected from exposure to direct light. (0.5 MM final was used in the example results shown here). • Live/Dead fixable blue dead cell stain kit for UV excitation (Life MS-5 growth medium Prepare the growth medium by adding 10% Technologies, cat. no. L23105) ! CAUTION Keep it at −20 °C and protect (vol/vol) FBS and 1% (vol/vol) penicillin-streptomycin in A-MEM. it from exposure to direct light. M CRITICAL Prepare the medium in advance and store it at 4 °C for up • Formaldehyde solution 37% (wt/vol) H2O (Sigma-Aldrich, cat. no. 252549) to 2–3 weeks. ! CAUTION Formaldehyde is toxic by inhalation or if swallowed; it is Mitomycin C solution Dissolve mitomycin C in sterile H2O to obtain a irritating to the skin, eyes and respiratory system, and it may be carcinogenic. 0.5 mg/ml stock solution. Store the solution in 500-Ml aliquots at −20 °C for Formaldehyde should be used with appropriate safety measures, such as up to 1 year. protective gloves, glasses, clothing and sufficient ventilation. All waste Cytokine buffer Mix 0.1% (vol/vol) HSA in PBS. Sterilize the solution by should be handled according to hazardous waste regulations. filtration using a 0.2-Mm syringe filter, and store it for up to 6 months at 4 °C. • BD CompBeads set anti-mouse Ig (BD Biosciences, cat. no. 552843) K Reconstitution of cytokines Reconstitute human cytokines in cytokine • BD CompBeads set anti-rat IgK (BD Biosciences, cat. no. 552844) buffer at a stock concentration of 20 Mg/ml for GM-CSF and SCF ligand and • Amine-reactive compensation bead kit (Life Technologies, 100 g/ml for FLT3L. Store the cytokines in 50- l aliquots at −80 °C for up cat. no. A10346) M M to 2 years. M CRITICAL Vials of lyophilized cytokines should be centrifuged EQUIPMENT • Biological safety level 2 (BSL-2) tissue culture facilities in a microcentrifuge at maximum speed for 1 min at room temperature • BSL-2 sorting facilities (20–25 °C) before opening vials. • Class II biological safety cabinet (The Baker Company, cat. no. SG403) Formaldehyde working solution Mix 2% (vol/vol) formaldehyde in PBS. Store the solution at 4 °C for up to 1 month. • Cell incubator (37 °C, 5% CO2; Nuaire, cat. no. NU-5500) • Tabletop centrifuge with rotors and adapters for 50- and 15-ml conical Staining buffer Mix 1% (vol/vol) FBS in PBS. M CRITICAL Staining buffer tubes and 96-well plates (Sorvall, cat. no. 75-004-367) can be prepared in advance and stored at 4 °C for up to 1 month. • Microcentrifuge (Eppendorf, cat. no. 5424) Sorting buffer Mix 1% (vol/vol) FBS and 0.5 M EDTA (2 mM final) • Inverted microscope (Nikon, cat. no. Eclipse TS100) in PBS. M CRITICAL Sorting buffer can be prepared in advance and stored • Hemocytometer (Fisher Scientific, cat. no. 02-671-5) or other cell counter at 4 °C for up to 1 month.

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Box 1 | CFSE staining L TIMING 30 min To assess the ability of cells to proliferate in culture (Steps 36–38), we use a cell proliferation assay. Cells are labeled with the fluorescent dye CFSE. Those cells that proliferate show a reduction in CFSE fluorescence intensity, which is measured directly by flow cytometry. 1. Wash the cells twice in PBS and resuspend them in PBS at a 2 × 107 cells per ml concentration. 2. Prepare a 1 MM working solution of CFSE from the stock (5 mM) by diluting the stock in PBS. Label the cells by adding one volume of working solution of CFSE (1 MM) to one volume of cells. The final concentration of CFSE is then 0.5 MM. 3. Incubate the cells in the dark with gentle mixing at room temperature for 8 min. 4. Quench the reaction by adding an equal volume of FBS for 2 min. 5. Wash the cells twice with PBS and resuspend them in MS-5 growth medium. 6. To assess the proliferation of bulk CD34+ cells or purified progenitors (see Step 36B for isolation of progenitors using panel #3) during hematopoietic differentiation on MS-5 stromal cells, proceed directly to Step 33.

Live/Dead blue viability dye Thaw Live/Dead blue powder by leaving it for for a specific antibody is the amount giving the best fluorochrome signal 10 min at room temperature. Add 50 Ml of DMSO and vortex it thoroughly. and the least fluorochrome spreading. To prepare the antibody staining Store the solution in 1-Ml aliquots in Eppendorf tubes at −20 °C for up to solution, mix the titrated amount of antibody for all antibodies and make it 1 year. Each batch of Live/Dead blue should be titrated before use in the up to 20 Ml with staining buffer (Table 2). The staining volume is 40 Ml total procedure on dying cells, for example, cells kept in culture at a very dense (20 Ml of cell suspension + 20 Ml of antibody staining solution). The staining concentration such as 20 × 106 cells per ml, to determine the optimal dilution volume should be scaled up according to the number of cells. Prepare this for labeling (1/1,000 final in the example results shown here). When you solution on the day of use. M CRITICAL Spin the antibody staining solution in are ready to use Live/Dead blue, thaw and keep on ice a 1-Ml aliquot into a microcentrifuge at maximum speed for 1 min at room temperature which you add 99 Ml of PBS (1/100 Live/Dead blue working solution). to remove antibody aggregates. Use the supernatant only to stain cells. Amine-reactive Live/Dead blue CompBeads Vortex the beads thoroughly M CRITICAL Always prepare the antibody staining solution in excess before use. Resuspend one drop of positive beads and one drop of negative (10% more than needed for the experiment). beads in 500 Ml of PBS. Add 20 Ml of Live/Dead blue working solution to BD CompBeads Vortex the beads thoroughly before use. Prepare a 180 Ml of beads solution in a 5-ml polypropylene round-bottom tube (final positive bead solution and a negative bead solution (typically one drop dilution of Live/Dead blue is 1/1,000). Incubate the mixture for 30 min at of beads is added to 500 Ml of PBS for the positive solution). For the 4 °C in the dark. Wash the beads twice with PBS. Resuspend the beads in negative FACS control, add 100 Ml of negative bead solution in one well 150 Ml of formaldehyde working solution and transfer them to a titer tube. of a 96-well V-bottom plate. For the positive FACS controls, add 100 Ml of Prepare the beads on the day of use. M CRITICAL It is crucial to use PBS alone positive bead solution per well of a 96-well V-bottom plate for each control. and not staining buffer that contains serum, as proteins from the serum Add the titrated amount of antibody. Incubate the 96-well plate for 30 min would prevent the binding of Live/Dead blue to the compensation beads. at 4 °C in the dark. Wash the beads twice with PBS. Resuspend each control

Nature America, Inc. All rights reserved. Inc. Nature America, Antibody staining solution The amounts of antibody needed per staining in 150 Ml of formaldehyde working solution. Transfer the controls to

5 are determined by titration experiments done before performing the procedure. titer tubes. Prepare the beads on the day of use. M CRITICAL Carefully The optimal concentration of an antibody is determined by a titration check that the monoclonal antibody chosen can be recognized by the experiment in which typically 2 × 106 cells in a 20-Ml volume are stained beads (for example, use anti-mouse IgK CompBeads for mouse © 201 with 1×, 2×, 4× and 8× the amount of antibody. The optimal concentration monoclonal antibodies).

PROCEDURE Donor CB and BM collection and initial processing L TIMING 10–45 min ! CAUTION All human blood, body fluids and tissues must be treated as if infectious and universal precautions must be taken by lab workers while manipulating these specimens. ! CAUTION All human biological waste to be discarded should be treated with bleach or it should be autoclaved, and then it can be discarded according to your institution disposal procedures. M CRITICAL All steps should be carried out in a class II biological safety cabinet. 1| Obtain human samples from CB, as described in option A, or obtain and prepare a single-cell suspension of BM, as described in option B. (A) CB sample L TIMING ~10 min (i) Obtain a fresh cord blood unit (CBU; typically 80–140 ml) provided in citrate phosphate dextrose adenine (CPDA-1) preservative solution. M CRITICAL STEP The CBU should be kept at room temperature until processing. For optimum results, processing should take place in <12 h, and the samples that are older than 24 h should not be used. (B) BM sample L TIMING ~45 min (i) Obtain biopsies from surgical procedures such as routine hip arthroplasty. The biopsy should be kept on ice immediately after surgical removal in a container for specimen collection, and it should be preserved in 1–2 ml of heparin (1,000 units per ml).

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(ii) Weigh the specimen for reference. M CRITICAL STEP Samples can be stored at 4 °C for a few hours before starting the isolation procedure. M CRITICAL STEP This protocol has been optimized for solid biopsies and not aspirates—i.e., a sample of the liquid portion of BM. M CRITICAL STEP This protocol is optimized for the processing of specimen weighing 20–40 g. For smaller or larger specimens, adjust the volumes accordingly. (iii) Wash the BM biopsy in PBS and transfer it into a 50-ml tube with 10–20 ml of collagenase solution. Incubate the mixture for 30 min at 37 °C under continuous agitation. ! CAUTION Collagenase digestion of tissues to create single-cell suspensions may result in the destruction of certain cell surface markers. However, we did not observe the destruction of any of the surface markers that we tested. M CRITICAL STEP Add 10 ml of collagenase solution to every 20 g of tissue. Collagenase is thawed and kept on ice. M CRITICAL STEP This short digestion allows the disruption of the bony matrix, as well as the release of the fat constituting the marrow. (iv) Filter the BM cell suspension through a fine mesh nylon screen.

Isolation of CB and BM MNCs via density gradient L TIMING 1.5 h 2| Prepare CB (option A) or BM (option B) density gradients. (A) CB gradient (i) Dilute the entire CBU 1:1 with sterile PBS in an upright appropriately sized sterile tissue culture flask. (ii) Carefully and slowly layer 35 ml of diluted CB onto 15 ml of Ficoll-Paque in a 50-ml conical tube. Repeat the process in additional tubes for the remaining diluted CBU. If the last amount does not total 35 ml, make up the volume to 35 ml with PBS. (B) BM gradient (i) Carefully and slowly layer 35 ml of BM cell suspension onto 15 ml of Ficoll-Paque in a 50-ml conical tube. If the amount does not total 35 ml, make up the volume to 35 ml with PBS. M CRITICAL STEP The more diluted the sample, the better the purity of the MNCs. M CRITICAL STEP To avoid mixing Ficoll-Paque and the cell suspension, tilt the 50-ml conical tube at a 45 °C angle and slowly let the blood run down on the side of the tube onto the Ficoll layer.

3| Centrifuge the mixture at 850g for 30 min at 20 °C in a swinging-bucket rotor with no brake.

Nature America, Inc. All rights reserved. Inc. Nature America, 4| Aspirate the upper layer (yellow layer corresponding to the plasma, in the case of CB) leaving the mononuclear cell 5 layer (white layer at the interphase between the Ficoll and the plasma) undisturbed. ?TROUBLESHOOTING © 201 5| Carefully transfer each mononuclear cell layer using a sterile transfer pipette to a new 50-ml conical tube.

6| Fill the 50-ml conical tubes with RPMI-1640, mix well, and centrifuge the tubes at 480g for 5 min at 20 °C with brake.

7| Carefully remove the supernatant completely. Pool the pellet of all 50-ml conical tubes. Fill the 50-ml conical tube with RPMI-1640, mix it well and centrifuge it at 480g for 5 min at 20 °C with brake. M CRITICAL STEP CB samples sometimes show substantial contamination of the interphase layer with red blood cells. In that case, resuspend the cell pellet in 5 ml of ACK lysing buffer. Allow it to stand for 5 min at 4 °C. Fill the 50-ml conical tube with RPMI-1640, mix it well and centrifuge it at 480g for 5 min at 20 °C with brake. Remove the supernatant; resuspend the cells in 50 ml of RPMI-1640 and centrifuge them once more at 480g for 5 min at 20 °C with brake.

8| Carefully remove the supernatant completely. Resuspend the cells in 50 ml of staining buffer, count the cells using a hemocytometer or other cell counter, and determine the viability by trypan blue stain exclusion. If you are not pausing, proceed directly to Step 35 (staining panel #1, 2 or 3) or Step 33 (staining panel #4 or 5). JPAUSE POINT Keep the cells in staining buffer at 4 °C for 1–2 h until use, or freeze them for long-term storage, as described in Steps 9–12.

Freezing of CB and BM MNCs L TIMING 20 min 9| Centrifuge the cells at 480g for 5 min at 20 °C.

10| Resuspend the cells at a density of 10 × 106 cells per ml in freezing medium.

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11| Transfer 1 ml of cells in freezing medium per sterile cryogenic micro tube. M CRITICAL STEP Label each tube clearly (date, sample name, cell concentration). M CRITICAL STEP The optimal cell concentration for freezing is 10 × 106 cells per ml. However, you can go up to 50 × 106 cells per ml in case of large specimens.

12| Put the cryogenic vial in a freezing container, and store it at −80 °C for 12–24 h before transferring it to the liquid nitrogen tank for long-term storage. JPAUSE POINT Cells can be stored in liquid nitrogen for years.

Thawing CB and BM MNCs L TIMING 30 min 13| Add 10 ml of cold thawing medium in a 15-ml conical tube to thaw one vial or add 40 ml of cold thawing medium in a 50-ml conical tube to thaw up to ten vials. M CRITICAL STEP To obtain the best possible survival, thawing of cells must be performed as quickly as possible, and the thawing medium should be cold to minimize heat shocks.

14| Remove the vial from the liquid nitrogen tank, and hold it at room temperature until the sides are thawed but the center remains frozen. M CRITICAL STEP Do not place the vial into a 37 °C water bath, as it is important not to dramatically heat-shock the cells.

15| Transfer the contents of the vial (~1 ml) into a 15-ml conical tube containing cold thawing medium.

16| Centrifuge the cells at 480g for 5 min at 20 °C.

17| Resuspend the cells in staining buffer for extemporaneous use.

Culture of mouse MS-5 stromal cells L TIMING 8 d M CRITICAL Mouse MS-5 stromal cells are only required for staining panel #4 or 5. 18| Culture MS-5 stromal cells in 10-cm tissue culture dishes with 10 ml of MS-5 growth medium. Place the cells in a CO2 cell incubator. M CRITICAL STEP MS-5 stromal cells grow in monolayers. Examine the tissue culture dish under the microscope; the cells are ready to be split when they are 95% confluent. Nature America, Inc. All rights reserved. Inc. Nature America,

5 19| Aspirate the MS-5 growth medium and wash the cells twice with 10 ml of PBS. © 201 20| Add 2 ml of trypsin-EDTA 0.05% (wt/vol) solution and incubate the cells for 2 min at 37 °C in a CO2 cell incubator. M CRITICAL STEP Inadequate washing of cells with PBS or using an old trypsin solution may result in partial detachment of cells from the dish.

21| Add 10 ml of MS-5 growth medium and collect the cells by pipetting.

22| Transfer the cell suspension in a 15-ml conical tube and centrifuge it at 480g for 5 min at 20 °C.

23| Aspirate the supernatant and resuspend the cells in 10 ml of MS-5 growth medium.

24| Add 0.5 ml of cell suspension to 9.5 ml of MS-5 growth medium, and plate the cells onto a 10-cm tissue culture dish. Place the cells in a CO2 cell incubator.

25| When the culture is 95% confluent, if required, split the dish for maintenance (by repeating Steps 19–24). Confluence should be reached after 3–4 d of culture.

26| To expand the cells for coculture experiment, prepare an extra tissue culture dish by adding 1 ml of cell suspension to 19 ml of MS-5 growth medium into a 15-cm Petri dish. M CRITICAL STEP Do not use cells that have a high passage number or that are growing slowly. The doubling time for healthy MS-5 is ~24 h. JPAUSE POINT It is possible to pause the protocol at this stage by freezing and storing MS-5 stromal cells in liquid nitrogen tank for future use. Cells can be frozen and thawed as described in Steps 9–17 for MNCs.

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Hematopoietic differentiation on MS-5 stromal cells L TIMING 8 d 27| The day before the coculture experiment (day 0), take the extra tissue culture dish of MS-5 stromal cells that was prepared (Step 26), and proceed to mitomycin C treatment. Aspirate the medium without disturbing the MS-5 stromal cell layer. Add 20 ml of fresh MS-5 growth medium containing 10 Mg/ml mitomycin C. Incubate the dish for 3 h in a CO2 cell incubator.

28| Trypsinize the MS-5 stromal cell layer, as described in Steps 19–23.

29| Count the MS-5 cells using a hemocytometer or other cell counters and determine viability by trypan blue stain exclusion.

30| Centrifuge the cells at 480g for 5 min at 20 °C; aspirate the supernatant.

31| Resuspend the pellet at 2.5 × 105 per ml in MS-5 growth medium.

32| Seed 2.5 × 104 cells (100 Ml) per well of a 96-well flat-bottom tissue culture plate. Incubate the cells overnight in a CO2 cell incubator.

33| The following day (day 1), resuspend a maximum of 104 bulk CD34+ cells or purified progenitors (see Step 36B for isolation of progenitors using panel #3) in 100 Ml of MS-5 growth medium containing 40 ng/ml GM-CSF and SCF ligand and 200 ng/ml FLT3L. M CRITICAL STEP If progenitors need to be stained with CFSE (panel #4) before coculture with MS-5 stromal cells, proceed as described in Box 1.

34| Seed the progenitor cells on top of the MS-5 stromal cells prepared in Step 32. Incubate the cells for 7 d in a CO2 cell incubator.

Cell surface staining L TIMING 1–2 h 35| Take all or an aliquot of the cell suspension from Step 8. Use at least 2–4 × 106 cells for simple phenotyping (panel #1) and characterization (panel #2) staining. If you are sorting the cells (panel #3), start with a sufficient number of cells to obtain a reasonable number of progenitors after sorting. We usually sort 100–200 × 106 CB or BM MNCs to achieve sufficient progenitors. Use the whole MS-5 culture from Step 34 for culture output (panel #4) and culture

Nature America, Inc. All rights reserved. Inc. Nature America, proliferation (panel #5) staining. 5 M CRITICAL STEP We recommend working with fresh samples. If you are working with frozen samples, thawing and staining should preferably be performed on the same day. If you are using frozen cells, count the cells (from Step 17) after thawing

© 201 using the hemocytometer or other cell counter and determine the viability by Trypan Blue Stain exclusion. ?TROUBLESHOOTING

36| If you are performing cell surface staining for panel #1 and 2, follow option A; for panel #3, follow option B; and for panel #4 and panel #5, follow option C. ! CAUTION This and all subsequent steps should avoid exposure to direct light to preserve fluorochromes. M CRITICAL STEP Do not forget to prepare the compensation tubes as well (see Reagent Setup). (A) Staining for panel #1 and #2 L TIMING 1 h 30 min (i) Resuspend the cells (from Step 8) in PBS at a density of 100 × 106 cells per ml. (ii) Put 2 × 106 cells (20 Ml) in each well of a 96-well V-bottom plate. (iii) Add 2 Ml of 1/100 Live/Dead blue working solution to each well. (iv) Incubate the cells at 4 °C in the dark for 20 min. (v) Wash the cells by adding 150 Ml of staining buffer per well. (vi) Centrifuge the cells at 480g for 5 min at 20 °C; flick the plate to decant the supernatant. (vii) Add 20 Ml of the antibody staining solution per well. (viii) Incubate the cells at 4 °C in the dark for 30 min. (ix) Wash the cells by adding 150 Ml of staining buffer per well. (x) Centrifuge the cells at 480g for 5 min at 20 °C; flick the plate to decant the supernatant. (xi) Resuspend the cells in 150 Ml of formaldehyde working solution per well. Transfer the samples to titer tubes. (xii) Keep the samples in the dark at 4 °C, and acquire samples on a flow cytometer. J PAUSE POINT Fixed cells can be kept in the dark at 4 °C for up to 48 h before analyzing by flow cytometry.

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(B) Staining for panel #3 L TIMING 1 h (i) Resuspend the cells (from Step 8) in staining buffer at a density of 400 × 106 cells per ml in a 50-ml conical tube. (ii) Add 20 Ml per 8 × 106 cells of the antibody staining solution. (iii) Incubate the cells at 4 °C in the dark for 30 min. (iv) Wash the cells by adding 40 ml of sorting buffer. (v) Centrifuge the cells at 480g for 5 min at 20 °C. Aspirate the supernatant. (vi) Resuspend the cells in sorting buffer at a density of 30 × 106 cells per ml, and transfer them to a 5-ml polypropylene round-bottom tube. M CRITICAL STEP We recommend filtering the cell sample before sorting using a 5-ml polystyrene round-bottom tube with a cell-strainer cap. Transfer the sample in a 5-ml polypropylene round-bottom tube for sorting. (vii) Keep the sample in the dark on ice, and then sort the sample on a cell sorter in a timely manner (within 1 h). (C) Staining for panel #4 and #5 L TIMING 1 h 30 min (i) Collect all cells from MS-5 stromal coculture (from Step 34) by vigorous pipetting, and transfer them to a 96-well V-bottom plate. (ii) Centrifuge the cells at 480g for 5 min at 20 °C; flick the plate to decant the supernatant. (iii) Stain the cells by the standard surface staining method, as in Step 36A(iii–xii). J PAUSE POINT Fixed cells can be kept in the dark at 4 °C for up to 48 h before analyzing by flow cytometry.

Acquisition and cell sorting L TIMING ~1–4 h 37| Run the compensation controls (see Reagent Setup). Create the compensation matrix and apply it to tubes for both acquisition and sorting. M CRITICAL STEP Compensation controls must consist of a negative and a positive population for each single fluorochrome. The negative and positive populations should have the same autofluorescence, which means that each positive bead should be compared with its matching negative bead. M CRITICAL STEP When you first set up a panel, you must include a nonstained tube, as well as FMO control (FMOC) tubes. FMOCs sequentially contain all fluorochrome or tandem dyes minus one; they are used to define proper gates and to control the compensation process. ?TROUBLESHOOTING

38| Run the samples (from Step 36A(xii) and Step 36C(iii)) for acquisition on the flow cytometer. Analyze data using FlowJo, as depicted in Figure 2 (panel #1), Figure 3 (panel #2), Figure 5 (panel #4) and Figure 6 (panel #5).

Nature America, Inc. All rights reserved. Inc. Nature America, M CRITICAL STEP If you do not have FlowJo software, use FACSDiva software or an equivalent. 5 ?TROUBLESHOOTING

© 201 39| Proceed to simultaneous sorting of the four populations of interest—i.e., GMDPs, MDPs, CDPs and pre-cDCs—from MNCs obtained in Step 8 and stained in Step 36B(i–vi) using an 85-Mm nozzle cell sorter. Collect the sorted cells in 1.5-ml conical-bottom tubes with screw caps, in which you add 300 Ml of MS-5 growth medium and vortex the collection tubes to coat the surface. You can collect up to 2.5 × 105 cells per tube. M CRITICAL STEP If you are sorting cells to analyze mRNA expression by gene array, cells should be recovered without interruption and should be kept at 4 °C at all times to avoid changes in gene expression. ?TROUBLESHOOTING

?TROUBLESHOOTING Step 4 If the upper layer is contaminated with RBCs, the samples—that is, upper layer and interphase—should be mixed and layered over a second batch of Ficoll in a new 50-ml conical tube. Repeat this step if necessary.

Step 35 Post-thaw samples may clot. The cells are probably aggregating because of the release of DNA by cells dying during the freezing-thawing process. Include DNase such as benzonase (final concentration 25 U/ml) in thawing medium (Steps 13–17), as well as in staining buffer in Steps 36A(iii,iv), 36A(vii,viii) and 36B(ii,iii).

Step 37 If you have no staining of compensation beads, it may be because not all compensation controls have been prepared in Step 36. You must prepare a compensation control for each fluorochrome or conjugate if you are using tandem dyes. It may also be that you are not using the proper beads; use anti-mouse IgK for mouse monoclonal antibodies and anti-rat IgK for rat monoclonal antibodies.

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Step 38 If you have clogging of the sample line while acquiring samples, the cells are probably aggregating because of poor viability. We recommend resuspending the cells in sorting buffer that contains EDTA and filtering the cells using 5-ml polystyrene round-bottom tubes with cell-strainer caps.

Step 39 If you have clogging of the sample line while sorting samples, the cells are probably aggregating because of poor viability. We recommend diluting the cells and filtering the sample again using a 5-ml polystyrene round-bottom tube with a cell-strainer cap.

L TIMING Staining panel #1: ~3 h–5 h 30 min Steps 1–8, fresh samples: ~2 h or Steps 13–17, frozen samples: 30 min Steps 35 and 36A, staining: 1 h 30 min Steps 37 and 38, acquisition: ~1–2 h Staining panel #2: ~3 h–5 h 30 min Steps 1–8, fresh samples: ~2 h or Steps 13–17, frozen samples: 30 min Steps 35 and 36A, staining: 1 h 30 min Steps 37 and 38, acquisition: ~1–2 h Staining panel #3: 3 h 30 min–7 h Steps 1–8, fresh samples: ~2 h or Steps 13–17, frozen samples: 30 min Steps 35 and 36B, staining: 1 h Steps 37 and 39, sorting: ~2–4 h Staining panel #4: 3 h–5 h 30 min + 8 d coculture Steps 1–8, fresh samples: ~2 h or Steps 13–17, frozen samples: 30 min Steps 18–26, culture of mouse MS-5 stromal cells: 8d Steps 27–34, hematopoietic differentiation on MS-5 stromal cells: 8 d Steps 35 and 36C, staining: 1 h 30 min Steps 37 and 38, acquisition: ~1–2 h Staining panel #5: ~3 h–5 h 30 min + 8 d coculture Steps 1–8, fresh samples: ~2 h or Steps 13–17, frozen samples: 30 min

Nature America, Inc. All rights reserved. Inc. Nature America, Box 1, CFSE staining: 30 min 5 Steps 18–26, culture of mouse MS-5 stromal cells: 8d Steps 27–34, hematopoietic differentiation on MS-5 stromal cells: 8 d

© 201 Steps 35 and 36C, for staining: 1 h 30 min Steps 37 and 38, acquisition: ~1–2 h

ANTICIPATED RESULTS Typically, GMDPs, MDPs and CDPs are found in the Lin− CD34+ cells, which comprise only a small and variable fraction of human BM cells (2–4%) and CB cells (~1%): (i) GMDPs represent 5% (BM) and 10% (CB) of CD34+ cells; (ii) MDPs represent 0.5% (BM) and 0.5% (CB) of CD34+ cells; and (iii) CDPs represent 2% (BM) and 0.4% (CB) of CD34+ cells14. Pre-cDCs, which are found in the Lin− CD34− cells, represent 0.1% (BM) and 0.008% (CB) of CD45+ cells15. Therefore, DC progenitors are rare cells; some variability in the number of progenitors is expected from one donor to another, but progenitors are consistently found in all BM and CB specimens. Variability between subjects is particularly problematic for BM samples that are obtained from elderly donors, in which age-related changes may affect the composition and the phenotype of the cells. As this protocol is based mainly on flow cytometry, it is crucial to verify the proper function and setup of the flow cytometer or cell sorter by running an appropriate cytometer procedure and making sure that laser delays are properly adjusted24. This step is often omitted and an improper setting will give inconsistent results. To perform a full and comprehensive evaluation of BM and CB hematopoietic progenitors using multiparametric flow cytometry (panels #1 and #2), an ideal number of 2 × 106 (BM) or 4 × 106 (CB) cells are required. Therefore, a small fraction of BM or CB is used, allowing the majority of the sample to be used for other studies. This protocol provides an appropriate method for BM and CB analysis that could be applied to samples from healthy or abnormal donors. Moreover, the same procedure could be adapted to other tissues—that is, blood, lymph node or tonsil—in order to identify and sort pre-cDCs for further studies and applications. Of note, the low frequency of pre-cDCs in whole blood (~15 cells per ml of blood) dictates the use of several milliliters of blood for their quantification.

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To ensure successful hematopoietic differentiation of DC progenitors in MS5+FSG culture, cell sorting should be performed under sterile conditions, and sorting parameters such as sheath fluid pressure should be optimized for viability of the cells. Ideally, a maximum of 104 CD34+ HSPCs or DC progenitors are plated on top of MS-5 stromal cells for 7 d. After 2 d of expansion, colonies can be observed under the microscope. The proportion of CD45+ cells substantially increases over 7 d. For early hematopoietic progenitors such as CD34+ HSPCs, the yield of cells can be increased a little bit by extending the culture; however, over 14 d expansion is associated with substantially lower output of CD45+ cells because of competition for cytokines and nutrients, as well as stromal cell death. The efficiency of the hematopoietic differentiation is going to depend on the sample itself, as variability between donors exists, and the number of cells that are initially cocultured on MS-5 stromal cells. After the cells are sufficiently expanded, further characterization can be performed using flow cytometry (panels #4 and #5).

ACKNOWLEDGMENTS This work was supported by US National Institutes of Health 9. Jongbloed, S.L. et al. Human CD141+ (BDCA-3)+ dendritic cells (DCs) (NIH) grant 1U19AI111825-01 (to M.C.N.), NIH grant AI13013, NIH/National represent a unique myeloid DC subset that cross-presents necrotic cell Center for Advancing Translational Sciences (NCATS) grant UL1 TR000043, antigens. J. Exp. Med. 207, 1247–1260 (2010). The Iris and Junming Le Foundation (to G.B.), The Empire State Stem Cell Fund 10. Poulin, L.F. et al. Characterization of human DNGR-1+ BDCA3+ leukocytes through New York State Department of Health Contract no. C029562 (to K.L.) and as putative equivalents of mouse CD8a+ dendritic cells. J. Exp. Med. 207, NIH/National Institute of Allergy and Infectious Disease (NIAID) grant AI101251 1261–1271 (2010). (to K. Liu). M.C.N. is a Howard Hughes Medical Institute investigator. 11. Villadangos, J.A. & Shortman, K. Found in translation: the human equivalent of mouse CD8+ dendritic cells. J. Exp. Med. 207, AUTHOR CONTRIBUTIONS G.B., J.L., K.L. and M.C.N. conceived and developed 1131–1134 (2010). the original strategy to identify the DC progenitors; G.B. modified the protocol 12. Robbins, S.H. et al. Novel insights into the relationships between and performed the experiments; G.B. and M.C.N. wrote the manuscript; and all dendritic cell subsets in human and mouse revealed by genome-wide authors reviewed the manuscript. expression profiling. Genome Biol. 9, R17 (2008). 13. Geissmann, F. et al. Development of monocytes, macrophages, and dendritic cells. Science 327, 656–661 (2010). COMPETING FINANCIAL INTERESTS The authors declare no competing financial 14. Lee, J. et al. Restricted dendritic cell and monocyte progenitors in interests. human cord blood and bone marrow. J. Exp. Med. 212, 385–99 (2015). 15. Breton, G. et al. Circulating precursors of human CD1c+ and CD141+ Reprints and permissions information is available online at http://www.nature. dendritic cells. J. Exp. Med. 212, 401–413 (2015). com/reprints/index.html. 16. Doulatov, S. et al. Revised map of the human progenitor hierarchy shows the origin of macrophages and dendritic cells in early lymphoid 1. Steinman, R.M. & Hemmi, H. Dendritic cells: translating innate to development. Nat. Immunol. 11, 585–593 (2010). adaptive immunity. Curr. Top. Microbiol. Immunol. 311, 17–58 (2006). 17. Hao, Q.L. et al. Identification of a novel, human multilymphoid progenitor 2. Steinman, R.M. & Banchereau, J. Taking dendritic cells into medicine. in cord blood. Blood 97, 3683–3690 (2001). Nature 449, 419–426 (2007). 18. Hoebeke, I. et al. T-, B- and NK-lymphoid, but not myeloid cells arise 3. Manicassamy, S. & Pulendran, B. Dendritic cell control of tolerogenic from human CD34+CD38–CD7+ common lymphoid progenitors expressing responses. Immunol. Rev. 241, 206–227 (2011). lymphoid-specific genes. Leukemia 21, 311–319 (2007).

Nature America, Inc. All rights reserved. Inc. Nature America, 4. Colonna, M., Trinchieri, G. & Liu, Y.J. Plasmacytoid dendritic cells in 19. Majeti, R., Park, C.Y. & Weissman, I.L. Identification of a hierarchy of 5 immunity. Nat. Immunol. 5, 1219–1226 (2004). multipotent hematopoietic progenitors in human cord blood. Cell Stem Cell 5. Jin, J.O., Zhang, W., Du, J.Y. & Yu, Q. BDCA1-positive dendritic cells 1, 635–645 (2007). (DCs) represent a unique human myeloid DC subset that induces innate 20. Wang, J. et al. SCID-repopulating cell activity of human cord © 201 and adaptive immune responses to Staphylococcus aureus infection. blood-derived CD34– cells assured by intra-bone marrow injection. Infect. Immun. 82, 4466–4476 (2014). Blood 101, 2924–2931 (2003). 6. Cohn, L. et al. Antigen delivery to early endosomes eliminates the 21. Boissel, N. et al. Defective blood dendritic cells in chronic myeloid superiority of human blood BDCA3+ dendritic cells at cross presentation. leukemia correlate with high plasmatic VEGF and are not normalized by J. Exp. Med. 210, 1049–1063 (2013). imatinib mesylate. Leukemia 18, 1656–1661 (2004). 7. Bachem, A. et al. Superior antigen cross- presentation and XCR1 22. Bigley, V. et al. The human syndrome of dendritic cell, monocyte, B and expression define human CD11c+CD141+ cells as homologues of murine NK lymphoid deficiency. J. Exp. Med. 14, 227–234 (2011). CD8+ dendritic cells. J. Exp. Med. 207, 1273–1281 (2010). 23. Hambleton, S. et al. IRF8 mutations and human dendritic-cell 8. Crozat, K. et al. The XC chemokine receptor 1 is a conserved selective immunodeficiency. N. Engl. J. Med. 365, 127–138 (2011). marker of mammalian cells homologous to mouse CD8a+ dendritic cells. 24. Perfetto, S.P. et al. Quality assurance for polychromatic flow cytometry J. Exp. Med. 207, 1283–1292 (2010). using a suite of calibration beads. Nat. Protoc. 7, 2067–2079 (2012).

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