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저 시-비 리- 경 지 2.0 한민

는 아래 조건 르는 경 에 한하여 게

l 저 물 복제, 포, 전송, 전시, 공연 송할 수 습니다.

다 과 같 조건 라야 합니다:

l 하는, 저 물 나 포 경 , 저 물에 적 된 허락조건 명확하게 나타내어야 합니다.

l 저 터 허가를 면 러한 조건들 적 되지 않습니다.

저 에 른 리는 내 에 하여 향 지 않습니다.

것 허락규약(Legal Code) 해하 쉽게 약한 것 니다.

Disclaimer

저 시. 하는 원저 를 시하여야 합니다.

비 리. 하는 저 물 리 목적 할 수 없습니다.

경 지. 하는 저 물 개 , 형 또는 가공할 수 없습니다.

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CCCTC-binding factor

controls the homeostasis of

epidermal Langerhans cells and

adult hematopoietic stem cells

Taegyun Kim

Department of Medical Science

The Graduate School, Yonsei University

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CCCTC-binding factor

controls the homeostasis of

epidermal Langerhans cells and

adult hematopoietic stem cells

Directed by Professor Hyoung-Pyo Kim

The Doctoral Dissertation

submitted to the Department of Medical Science,

the Graduate School of Yonsei University

in partial fulfillment of the requirements for the degree of

Doctor of Philosophy

Taegyun Kim

December 2016

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This certifies that the Doctoral

Dissertation of Taegyun Kim is approved.

_________________________________________

Thesis Supervisor: Hyoung-Pyo Kim

_________________________________________

Thesis Committee Chair: Kwang Hoon Lee

_________________________________________

Thesis Committee Member: Jongsun Kim

_________________________________________

Thesis Committee Member: Chae Gyu Park

_________________________________________

Thesis Committee Member: Sun-Young Chang

The Graduate School

Yonsei University

December 2016

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“Never Lose A Holy Curiosity”

Albert Einstein

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ACKNOWLEDGEMENT

First of all, I thank God for his love and guidance on my life.

I am deeply indebted to my supervisor, professor Hyoung-Pyo Kim,

whose stimulating suggestion and encouragement helped me

throughout my work. His invaluable scientific inputs and our exciting

discussions were the highlight during my PhD life. Thank you for

guiding me how to ask questions and do the science.

I wish to express my sincere thanks to my committee members,

professors Kwang Hoon Lee, Jongsun Kim, Chae Gyu Park, and Sun-

Young Chang for their constructive advices and guidance.

I would like to deeply thank my dermatology mentor, professor Min-

Geol Lee, who firstly showed me the importance of basic science in

medicine and encouraged me to accomplish this research career.

I could not forget the warm supports from the lab members, Sueun

Kim, Mikyoung Kim, Soyeon Jung, Min-Ji Song, Yeeun Choi, Jong-

Joo Lee, Youn Wook Chung, Keunhee Park, and Joo-Hong Park. I

sincerely wish their great success in science.

I would like to thank my parents and parents-in-law for their never-

ending support. Their generous sacrifices have greatly allowed my

scientific pursuits and led me to devoting myself to the nature of

science.

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Last but not least, I wish to deeply thank my beloved wife, Sue and

adored daughter, Saeon for their generous understanding and endless

belief on my choices in life. Without their personal sacrifices and

supports, I could not finish this long journey. I love you.

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TABLE OF CONTENTS

Title ····················································································i

Signature page ······································································ ii

Proverb ·············································································· iii

Acknowledgement ································································· iv

Table of contents ·································································· vi

List of figures ······································································ xi

List of tables ······································································xiv

ABSTRACT ········································································· 1

I. INTRODUCTION ····························································· 3

1. Langerhans cells····························································· 3

2. Hematopoietic stem cells··················································· 5

3. CCCTC-binding factor ···················································· 8

II. MATERIALS AND METHODS···········································10

1. Mice···········································································10

2. Generation of human monocyte-derived Langerhans cells·········11

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3. Viruses and Vpx-containing virus-like particle production ········12

4. Tissue preparation ·························································13

5. Antibodies and flow cytometry ··········································13

6. Whole-mount epidermal sheet staining ································15

7. Immunofluorescence ······················································15

8. Electron microscopy ·······················································15

9. Whole-skin explant culture···············································16

10. In vivo bromodeoxyuridine labeling and analysis ····················17

11. Fluorescein isothiocyanate-induced cutaneous DC migration ·····17

12. Contact hypersensitivity and hapten-specific T-cell responses ····17

13. Croton oil-induced irritant dermatitis ·································18

14. Epicutaneous sensitization and T cell adoptive-transfer············18

15. Clinical and histological evaluation of epicutaneously sensitized

skin············································································19

16. Enzyme-linked immunosorbent assay for OVA-specific antibody

detection ·····································································19

17. Chemotaxis migration assay ·············································20

18. Quantitative real-time polymerase chain reaction ···················20

19. Western blot·································································21

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20. Blood cell count·····························································22

21. Bone marrow transplantation ···········································22

22. Cell cycle assays ····························································23

23. Measurement of reactive oxygen species·······························23

24. Colony-forming unit assay ···············································23

25. Microarray ··································································24

26. Statistics ·····································································24

III. RESULTS ·····································································26

PART A. CCCTC-binding factor controls the homeostatic maintenance

and migration of Langerhans cells ············································26

1. Decreased pool of multiple DC lineages in DC-specific CTCF-

deficient mice································································26

2. Reduced cell number with distinct morphological and surface

phenotypes of CTCF-deficient LCs ·····································33

3. CTCF-deficient LCs exhibit reduced homeostatic self-turnover ··36

4. CTCF deficiency impairs the emigration of LCs from the epidermis

·················································································41

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5. CTCF inactivation in LCs augments cell adhesion molecule gene

expression····································································49

6. CTCF-dependent LC homeostasis is required for optimal immunity

of the skin····································································53

PART B. CCCTC-binding factor regulates adult hematopoietic stem cell

quiescence in mouse ·····························································60

1. Loss of CTCF leads to rapid BM failure·······························60

2. Acute depletion of CTCF results in rapid shrinkage of c-Kithi HSC

and progenitor pool························································64

3. Loss of CTCF in hematopoietic compartment results in rapid

exhaustion of HSCs ························································68

4. Defective HSC-driven myeloid lineage development by deletion of

CTCF in vivo································································73

5. CLPs and lymphoid lineage cells are less sensitive to CTCF

ablation in vivo ·····························································77

6. CTCF-deficient HSCs are unable to maintain quiescence··········79

IV. DISCUSSION·································································83

V. CONCLUSION·······························································89

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REFERENCES·····································································90

ABSTRACT (IN KOREAN)··················································· 105

PUBLICATION LIST ·························································· 107

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LIST OF FIGURES

Figure 1. Schematic diagram showing Ctcffl/fl mice generation

and CTCF deletion efficiency········································· 27

Figure 2. Decrease in the general pool of DCs in mice with DC-

specific CTCF depletion in vivo ······································ 30

Figure 3. DC-selective CTCF deletion results in a decreased

general DC lineage ······················································ 32

Figure 4. LC-selective loss of CTCF leads to decreases in

epidermal LC numbers and distinct phenotypic characteristics

of LCs ······································································ 34

Figure 5. CTCF-deficient LCs exhibit self-proliferation

impairment································································ 37

Figure 6. Early epidermal LC network formation is comparable

between WT and huLang-cKO mice································ 40

Figure 7. CTCF-deficient LCs are significantly less distributed

in their sequential migratory compartments······················ 42

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Figure 8. CTCF-deficient LCs fail to egress from the epidermis

··············································································· 44

Figure 9. CTCF-silenced MDLCs showed a migration defect

toward CCL19 ··························································· 47

Figure 10. CTCF-deficient LCs express higher levels of cell

adhesion molecules ······················································ 51

Figure 11. Prolonged and exaggerated CHS responses with

higher priming of IFN-γ-producing CD8+ T cells in huLang-

cKO mice ·································································· 55

Figure 12. Attenuated epicutaneous sensitization to OVA in

huLang-cKO mice······················································· 58

Figure 13. Acute CTCF ablation leads to hematopoietic failure

··············································································· 62

Figure 14. Acute depletion of CTCF results in severe decrease

in c-Kithi stem/progenitors············································· 66

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Figure 15. CTCF loss in the hematopoietic system leads to

severe BM failure and consequent lethality ······················· 69

Figure 16. CTCF deficiency-mediated HSC depletion is a cell-

autonomous effect ······················································· 72

Figure 17. Loss of CTCF results in exhaustion of myeloid

progenitors and their differentiation ······························· 75

Figure 18. CLP-derived lymphoid differentiation is less

sensitive to CTCF depletion··········································· 78

Figure 19. CTCF-deficiency leads to an augmented cell cycle

progression in HSCs ···················································· 81

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LIST OF TABLES

Table 1. Primer sequences used for real-time qPCR············ 21

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ABSTRACT

CCCTC-binding factor controls the homeostasis of

epidermal Langerhans cells and adult hematopoietic stem cells

Taegyun Kim

Department of Medical Science

The Graduate School, Yonsei University

(Directed by Professor Hyoung-Pyo Kim)

CCCTC-binding factor (CTCF) is a highly conserved DNA-binding zinc-finger protein

that regulates higher order chromatin organization and is involved in various gene

regulation processes. Although its DNA occupancy is widely distributed throughout the

mammalian genome, there exist cell type-specific CTCF binding patterns which partly

explain its heterogeneous biological functions. Therefore, a functional analysis of CTCF in

each cell type would further expand our knowledge of CTCF. In this dissertation, we focus

on the homeostatic role of CTCF in epidermal Langerhans cells (LCs) and adult

hematopoietic stem cells (HSCs).

LCs are skin-resident dendritic cells (DCs) that orchestrate skin immunity mainly

through presenting cutaneous antigens to T lymphocytes. By using DC- and LC-specific

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CTCF conditional knockout (cKO) mouse, we demonstrate that CTCF regulates the

homeostatic maintenance of LCs via supporting LC self-turnover in situ. Functionally,

CTCF is required for the efficient cellular emigration of LCs both in the steady and

inflammatory state by suppressing the transcriptional expression of cell adhesion

molecules. This functional feature is closely associated with the enhanced mouse contact

hypersensitivity, but also with the reduced epicutaneous protein sensitization.

Hematopoiesis is a series of lineage differentiation programs initiated from HSCs in the

bone marrow (BM). To maintain lifelong hematopoiesis, HSCs should undergo tightly

regulated self-proliferation and cellular differentiation. By using a tamoxifen-induced

CTCF cKO mouse system and BM transplantation approaches, we demonstrate that CTCF

is a critical regulator for the homeostatic maintenance of adult HSCs by retaining HSC cell

cycle quiescence.

Our study uncovers that CTCF is a pivotal molecular determinant in the homeostatic

maintenance of epidermal LCs and adult HSCs in vivo.

Key words: CCCTC-binding factor, conditional knockout mouse, dendritic cells,

hematopoietic stem cells, homeostasis, Langerhans cells

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CCCTC-binding factor controls the homeostasis of

epidermal Langerhans cells and adult hematopoietic stem cells

Taegyun Kim

Department of Medical Science

The Graduate School, Yonsei University

(Directed by Professor Hyoung-Pyo Kim)

I. INTRODUCTION

1. Langerhans cells

Langerhans cells (LCs) are migratory dendritic cells (DCs) that reside in the epidermis.

Although LCs were first described as neuronal cells, it is now clear that they are a member

of the antigen-presenting cell family.1 The outermost residence of LCs among the

cutaneous DC subsets may be evolutionally associated with their primary immune-

surveillance of the skin surface where they directly contact with the environmental phase.2

Accordingly, LCs extend their dendrites toward the breaches of epidermal barrier and

actively uptake exterior antigens.2 By doing this, LCs can induce a strong humoral

immunity which confers our body to be protected against certain pathogenic microbial

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challenges.3 However, this immunological process is also involved in particular allergic

inflammations of the skin including atopic eczema.4

LCs had been the most extensively studied cutaneous DC subset until dermal Langerin+

DCs were characterized. LCs are strong antigen-presenting and T cell-stimulating DCs in

the in vitro system, and these cardinal properties led us to consider that LCs were solely

immunogenic.5 However, the resultant contact hypersensitivity (CHS) studies using

genetically engineered LC-ablating mouse strains have suggested that LCs are not only

immunogenic but also tolerogenic.6,7 In addition, other key studies indicate that there is a

functional compensation and/or redundancy between LCs and Langerin+ dermal DCs in

CHS settings, and both LCs and Langerin+ dermal DCs are somehow involved in CHS

induction. Thus, the role for LCs in diverse skin immune responses is quite context

dependent and further elaboration is still needed to elucidate a categorized function of LCs.

Unlike other cutaneous DC populations, which are continuously replaced by bone

marrow-derived precursors entering the skin, LCs have distinct developmental and

homeostatic properties. LCs are derived from skin-infiltrating myeloid precursors that seed

fetal epidermis at the late stage of embryogenesis.8,9 After birth, LC precursors actively

proliferate and start to gain mature LC markers such as MHC-II and Langerin within the

first week of postnatal period. Subsequent elegant lineage-tracing study showed that those

LC precursors derive from at least two cellular origins, embryonic fetal liver monocytes

and yolk-sac macrophages. After completion of epidermal LC network formation, LCs

maintain their cellular pool by slow self-turnover without any blood precursor input.8,10

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LCs also demonstrate a unique cytokine dependency for their development and

homeostasis different from those of other conventional DCs. Although most of

conventional DC homeostasis largely depends on Fms-like tyrosine kinase 3 (Flt3) ligand

cytokine signaling, LCs are independent of Flt3 ligand for their homeostasis and

development. Rather, similar to tissue-resident macrophages, LC development strictly

depends on colony-stimulating factor 1 (CSF-1) signaling.11 Recent studies have revealed

that IL-34 was specifically expressed in the stromal components of skin and brain, and IL-

34 was an actual ligand for CSF-1 receptor that supports LC and brain microglia

development in situ.12,13 Another major cytokine that supports LC development and

homeostasis is transforming growth factor beta 1 (TGF-β1). TGF-β1 null mice showed a

lack of epidermal LC, and both LC autocrine and paracrine sources of TGF-β1 were

essential for LC homeostasis.14,15 Interestingly, TGF-β1 signaling on LCs is not only

involved in LC development but also in actively suppressing homeostatic LC migration out

of the epidermis.16 TGF-β1 dependency in LCs has further been supported as a result of

defective LC development in Id2, Runx3, and PU.1 knockout mouse strains, the genes that

are TGF-β1-associated signaling transcription factors.17-19

2. Hematopoietic stem cells

Hematopoiesis in our body is primarily maintained by a complex differentiation

program beginning in hematopoietic stem cells (HSCs).20 The unique and defining

characteristic of HSCs is the ability for self-renewal and multi-lineage differentiation.

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HSCs give rise to multiple differentiated progenies following a series of hierarchical

differentiation steps through the multipotent progenitors and lineage-restricted precursors.

The most primitive HSCs with the highest multipotent activity are long-term repopulating

HSCs (LT-HSCs) which show self-renewal capacity. In contrast to LT-HSCs, short-term

repopulating HSCs (ST-HSCs) reveal the capacity to contribute transiently to the

production of lymphoid and myeloid cells. The group of heterogeneous multipotent

progenitors (MPPs) is featured by a more limited proliferative potential, but retained

ability to differentiate into various lineage of cells.

HSCs and MPPs can be distinguished by the surface marker expression within the

lineage-negative and Sca-1/c-Kit-positive population which is called LSK fraction

(Lin−Sca-1+c-Kit+) in bone marrow (BM). The SLAM (signaling lymphocyte activation

molecule) family receptors CD150 and CD48 are useful surface markers allowing us to

characterize LT-HSCs (CD150+CD48−), ST-HSCs (CD150−CD48−), and the more lineage-

restricted MPPs (CD48+). Under homeostatic conditions, about 60- 70% of LT-HSCs

remain in a quiescent G0 phase that contributes to their long-term maintenance. The small

fraction of LT-HSCs is in an active dividing state and differentiates into all hematopoietic

cells. This cycling population not only maintains HSC pool by self-renewal but also

produces all blood cells.

The highly regulated differentiation process of the quiescent HSCs is associated with a

variety of cell fate-determining factors. Lifelong HSC perpetuation and function is

preserved by a specific set of transcription factors such as RUNX1, GATA2, GFI1, and

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TAL1,20-22 and several signaling pathways including Wnt/β-catenin and Notch

pathway.23,24

HSCs should undergo a tightly coordinated molecular regulation to support both HSC

self-renewal and differentiation, and in this regards, there have been several reports

emphasizing the critical role for epigenetic and chromatin modifications.25-27 DNA-

methyltransferases such as Dnmt1 and Dnmt3a are required for HSC homeostasis and

differentiation via down-regulation of myeloid

progenitor-related factors comprising Gata1, Id2, and Cepba.28-30 Dnmt1 was crucial for

HSC self-renewal and lineage-commitment. The ablation of reduced expression of Dnmt1

in murine HSCs led to diminished repopulating capacity of HSCs and decreased

production of lymphoid progenitors accompanied with retained myeloid progenitor

development.28,29 In contrast to Dnmt1, Dnmt3a/3b deficiency affected only the long-term

reconstituting ability of HSCs, but not their differentiation.31 Polycomb group proteins

represent a major class of epigenetic regulators in diverse developmental contexts and

participate in transcriptional repression process.32 The components of polycomb-repressive

complexes including Bmi1,33 Rae28,34 and Ring1B35 have been identified to modulate HSC

fate and function. In addition, histone H2A deubiquitinase Mysm1 is also unveiled to be

necessary to maintain the function of HSCs.36 In line with an emerging importance of

epigenetic changes in the pathogenesis of myelodysplastic syndrome and acute myeloid

leukemia,37 further molecular understanding how HSC homeostasis is maintained by other

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epigenetic processes would be important for developing new therapeutic strategies in

hematologic diseases.

3. CCCTC-binding factor

CCCTC-binding factor (CTCF) is a highly conserved, 11-zinc finger protein that

contains a DNA-binding domain which shows about 100% homology among the different

mammals.38 Its DNA occupancy is widely distributed throughout the mammalian genome,

and 30–60% of binding is cell type specific, partly due to differences in DNA

methylation.39 It is thought that the extent of CTCF occupancy relies on intrinsic zinc

finger clusters that recognize specific DNA sequences and extrinsic factors that either

support or destabilize its DNA-binding affinity which likely underlies the diverse functions

of CTCF in different cell types.40 CTCF was originally identified as a negative regulator of

Myc gene expression41 and was subsequently proven to function as an insulator42 and

recently it has been shown as a coordinator of higher-order chromatin structure at beta-

globin locus.43 A growing body of evidence highly supports the idea that CTCF function is

largely attributed to its specific ability to mediate long-range DNA interactions that affect

the three-dimensional chromatin structure.38,44 Topological genetic remodeling can

potentially regulate the expression of cell differentiation- and function-associated genes.

Functional implications of CTCF in the mammalian biology have been discovered by

using conditional CTCF gene knockout mouse line as CTCF-null embryos displays

prenatal lethality.45 CTCF plays multiple roles in hematopoietic cell lineages; it regulates

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early thymocyte development,46 T cell differentiation,47 and cytokine production in helper

T cells,47,48 as well as T-cell receptor alpha gene recombination.49 A CTCF-mediated

chromatin organization essentially modulates immunoglobulin gene rearrangement in B

lymphocytes.50,51 In myeloid lineage, CTCF is involved in common myeloid progenitor

differentiation52 and macrophage development and function.53 It also has been shown that

overexpression of CTCF alters DC survival, proliferation, and differentiation.54 However,

currently the possible role for CTCF in regulating LC and HSC homeostasis is largely

unknown. Therefore, we focus on the functional role of CTCF in epidermal LCs and BM

HSCs in vivo by generating conditional knockout mouse system.

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II. MATERIALS AND METHODS

1. Mice

Mice carrying the targeted CCCTC-binding factor (Ctcf) allele, Ctcftm1a(EUCOMM)Wtsi, were

purchased from European Conditional Mouse Mutagenesis Consortium. For deletion of the

lacZ-neomycin-resistance cassette and the generation of mice with a loxP-flanked Ctcf

allele (Ctcffl/+), Ctcftm1a(EUCOMM)Wtsi mice were bred to ACT-FLPe C57BL/6 mice (Jackson

Laboratory, Bar Harbor, ME, USA), which express a transgene of an enhanced form of the

recombinase FLP that is driven by a chicken β-actin-rabbit β-globin hybrid promoter and

the human cytomegalovirus-IE enhancer.55 Ctcffl/+ mice were intercrossed to generate

Ctcffl/fl mice. Mice were genotyped by polymerase chain reaction (PCR) with the following

oligonucleotides: forward primer 5’- AACTCCCTGGTTCCACTCCT -3’ and reverse

primer 5’- GACCCTAGTGCTGTTTCGGC -3’; wild-type product, 420 bp; Ctcffl product,

539 bp. To generate mice with DC-specific Ctcf deficiency (CD11c-cKO), Ctcffl/fl mice

were crossed with CD11c-Cre BAC transgenic mice.56 To generate mice with LC-specific

Ctcf deficiency (huLang-cKO), we crossed Ctcffl/fl mice with human Langerin-Cre BAC

transgenic mice (obtained from NCI Mouse Repository).15 After crossing Ctcffl/fl mice with

Cre mice, the deletion efficiency of the conditional Ctcf allele was confirmed by genomic

DNA PCR with the following oligonucleotides: forward primer 5’-

AACTCCCTGGTTCCACTCCT-3’ and reverse primer 5’-

TGGAGACCCAGATTCAAATG-3’; Ctcffl product, 1226 bp; CtcfΔ product, 435 bp.

CD45.1 congenic mice and OT-II transgenic mice expressing T-cell receptor specific for

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OVA323-339 were purchased from the Jackson Laboratory. CD45.1 and OT-II mice were

interbred to obtain CD45.1+OT-II mice. Ctcffl/fl mice were crossed to Rosa26-CreER mice

(CreER, kindly provided by Weonsang Ro, Institute of Gastroenterology, Yonsei

University College of Medicine) to generate a tamoxifen-induced systemic Ctcf

conditional knockout strain (CTCF-cKO).57 For inducing Cre-mediated recombination in

CreER mice, tamoxifen (Sigma, St. Louis, MO, USA) was dissolved in corn oil (Sigma)

and intraperitoneally injected for four to five consecutive days (2mg/day). Wild-type

C57BL/6 (CD45.2) mice were purchased from The Jackson Laboratory. Six- to 12-week-

old mice that were bred in specific pathogen-free facilities at Yonsei University College of

Medicine were used for all experiments. Age- and sex-matched Ctcffl/fl or CreER littermate

mice were used as wild-type (WT) controls throughout the study. All animal studies were

approved by the Department of Laboratory Animal Resources Committee of Yonsei

University College of Medicine.

2. Generation of human monocyte-derived Langerhans cells (MDLCs)

Peripheral blood mononuclear cells (PBMCs) were obtained from normal volunteers

(n=15; age range 25-40) under the Yonsei University IRB-approved protocol. Written

informed consent was obtained and study was performed following the Declaration of

Helsinki Principles. PBMCs were isolated by gradient centrifugation with Ficoll-Paque

PLUS (GE Healthcare, Piscataway, NJ, USA) and CD14+ monocytes were purified by

anti-human CD14 microbeads (Miltenyi Biotec, Bergisch Gladbach, Germany). Cell purity

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was confirmed by flow cytometry (>92%). Total 1 x 106 monocytes were placed in 24-well

tissue culture plates (SPL, Seoul, Korea) and cultured in complete RPMI medium, which

consisted of RPMI 1640 (HyClone, Logan, UT, USA) that was supplemented with 10%

heat-inactivated fetal bovine serum (FBS) (HyClone), 50 μM 2-mercaptoethanol, 2 mM L-

glutamine, 1 mM nonessential amino acids, 1 mM sodium pyruvate, 100 units/mL

penicillin, and 100 μg/mL streptomycin. For MDLC differentiation, 100 ng/ml rhGM-CSF,

20 ng/ml rhIL-4 (Creagene, Seoul, Korea), and 10 ng/ml TGF-β1 (Peprotech, Rocky Hill,

NJ, USA) were used. MDLCs were harvested at day 6 and stimulated with 1 μg/ml of

lipopolysaccharide (LPS) (Invivogen, San Diego, CA, USA) for additional 2 days.

3. Viruses and Vpx-containing virus-like particle (Vpx-VLP) production

Due to the strikingly low transduction efficiency of monocytes and DCs with HIV-1-

derived lentiviruses, we took advantage of co-transduction of Vpx-VLP which drastically

assisted MDLCs with lentiviral infection.58 The pGIPz and pGIPz-shCTCF lentiviral

vectors were obtained from Dr. Jianrong Lu (University of Florida College of Medicine).59

pSIV3+ plasmid was kindly provided by Dr. Kwangseog Ahn (Seoul National

University).60 Lentiviruses were generated from HEK-293 cells co-transfected with

psPAX2 and pMD2.G (Addgene, Cambridge, MA, USA) using polyethylenimine

(Polysciences, Warrington, PA, USA). Vpx-VLP was generated by co-transfection of

pSIV3+ with pMD2.G.61 Lentiviral infections were carried out during the first 2 days of

monocyte culture in the presence of 8 μg/mL polybrene (Sigma). Green fluorescent protein

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(GFP)-positive transduced MDLCs were underwent FACS sorting and used for further

analyses.

4. Tissue preparation

For epidermal and dermal single-cell suspensions, excised ears were placed in 2.5

mg/mL dispase II (Roche Diagnostics, GmbH, Mannheim, Germany) for 30 min at 37°C.

Isolated epidermal and dermal sheets were incubated in 1 mg/mL collagenase IV

(Worthington, USA) for 1 hr at 37°C. In some experiments, mouse ears were treated with

20 μL of 1% oxazolone (Sigma) dissolved in acetone/olive oil (4:1), and after 24 hr, the

skins were prepared and subsequently examined. Total cells from the skin-draining lymph

nodes (SDLNs) and spleen were incubated in 1 mg/mL collagenase IV for 1 hr at 37°C.

5. Antibodies

Fluorochrome-conjugated anti-mouse CD11b, CD11c, CD40, CD44, CD45, CD45.1,

CD69, CD80, CD86, CD103, CD115 (colony-stimulating factor-1 receptor [CSF-1R]),

CD135, CD207 (Langerin), Annexin V, C-C chemokine receptor 7 (CCR7), E-cadherin,

epithelial cell adhesion molecule (EpCAM), F4/80, Foxp3, I-Ab (MHC II), interferon

(IFN)-γ, Ki-67, Siglec-H, and signal-regulatory protein alpha (Sirpα) monoclonal

antibodies (mAbs) were obtained from eBioscience (San Diego, CA, USA), and anti-

mouse CD4 and CD8α antibodies were from BD Biosciences (San Jose, CA, USA).

Biotin-conjugated anti-mouse CD3, CD19, NK1.1, Ter119, and B220 mAbs were used for

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discriminating lineage-positive populations in the bone marrow to analyze pre-cDCs

(eBioscience). Anti-mouse CTCF (Abcam, Cambridge, MA, USA) and transforming

growth factor-beta receptor II (TGF-βRII; R&D Systems, Minneapolis, MN, USA)

antibodies were also used in this study. For analyzing HSCs in BM, total BM cells were

flushed and harvested from two hind legs. Fluorochrome-conjugated anti-mouse B220,

CD3, CD4, CD8, CD11b, CD11c, CD19, CD16/32, CD34, CD45.1, CD45.2, CD48,

CD117/c-Kit, CD135/Flt3, CD150, F4/80, IL-7Rα, Ki-67, Ly-6G, Sca-1, Siglec-H were

used for the surface immunephenotyping (eBioscience). Lineage-positive cells in the BM

were excluded using the following biotinylated antibodies followed by APC-Cy-

conjugated streptavidin: B220, CD3, CD4, CD8, CD11b, CD11c, CD19, Gr-1, NK1.1, and

Ter119 (all from eBioscience). Tissue suspensions containing cells were incubated in

violet fluorescent live/dead dye (Molecular Probes, Eugene, OR, USA) for 30 min on ice

and stained with the antibodies described above at the appropriate dilutions using FACS

buffer (phosphate-buffered saline supplemented with 1% bovine serum albumin). For the

intracellular detection of IFN-γ and Langerin, cells were fixed and permeabilized with BD

Cytofix/Cytoperm (BD Biosciences) and incubated for 30 min on ice with antibodies

diluted in Perm/Wash buffer (BD Biosciences). Intracellular detection of Foxp3 and Ki-67

was performed using Foxp3/Transcription Factor Staining Kit (eBioscience). Cell death

and apoptosis were analyzed by Annexin V/PI staining kit (eBioscience). For human

MDLC staining, we used fluorochrome-conjugated anti-human CCR7, CD11c, CD14,

CD80 (BD Biosciences), CD86, and E-cadherin (eBioscience) mAbs. Stained cells were

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acquired by the FACSVerse or LSRFortessa flow cytometer (BD Biosciences). All flow

cytometry data were analyzed with the FlowJo software (Treestar, Ashland, OR, USA).

6. Whole-mount epidermal sheet staining

Separated epidermal sheets were fixed with cold acetone for 15 min at room temperature.

Epidermal sheets were incubated with a purified anti-mouse I-Ab mAb (clone M5/114,

Biolegend, San Diego, CA, USA), and signals were amplified by Alexa 488-conjugated

goat anti-rat IgG (Invitrogen, Carlsbad, CA, USA). After staining, sheets were mounted on

glass slides with mounting medium (Vector Laboratories, Burlingame, CA, USA). Images

were acquired with the LSM 700 confocal microscope (Zeiss, Oberkochen, Germany) and

analyzed with the ZEN software (Zeiss).

7. Immunofluorescence

Tissue sections (6 μm thick) from frozen ear samples that were preserved in optimal

cutting temperature (Sakura, Tokyo, Japan) were prepared by cryostat. Skin sections were

fixed in ice-cold acetone and blocked in 10% goat serum. After antibody staining, images

were generated with the LSM 700 confocal microscope (Zeiss) and analyzed with the ZEN

software (Zeiss). The TUNEL assay was performed using the In Situ Cell Death Detection

Kit, according to the manufacturer’s instructions (Roche Diagnostics).

8. Electron microscopy

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For immersion-fixation, mouse ear tissues were fixed with a mixture of 3%

formaldehyde (freshly prepared from paraformaldehyde) and 1% glutaraldehyde (vacuum

distilled) in 0.1 M cacodylate buffer (pH 7.2) for 15 min at 37°C. Afterwards, the tissues

were sliced into thin pieces, followed by immersion fixation in the same fixative at

ambient temperature for 4 hr. To stop fixation, free aldehyde groups were blocked by

soaking the tissue slices in 50 mM NH4Cl in buffer for 1 hr. The tissue slices were then

post-fixed with 1% osmium tetroxide in distilled water for 1 hr, followed by en bloc

staining with 1% aqueous uranyl acetate for 1 hr. After dehydrations in a series of graded

ethanol, embedding in Epon-Araldite (Fluka) was performed, according to the standard

protocol. Ultrathin 80-nm sections were prepared on copper slot grids, stained with uranyl

acetate and lead citrate, and observed with a Hitachi H-7650 electron microscope (Hitachi,

Tokyo, Japan) at 80 kV. Electron micrographs were taken with an 11-megapixel CCD

XR611-M digital camera (Advanced Microscopy Techniques, Woburn, MA, USA).

9. Whole-skin explant culture

Ears of mice were cut off the base and split into dorsal and ventral halves. Both skin

halves were cultured in 12-well plates containing complete RPMI culture medium. After 3

days, supernatants were pooled from the four skin explants and analyzed by flow

cytometry. Epidermal sheets were prepared from the cultured explants and subsequently

assessed by whole-mount epidermal sheet staining.

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10. In vivo bromodeoxyuridine (BrdU) labeling and analysis

Each mouse received daily intraperitoneal injections of 1 mg BrdU (Sigma) for 7 days.

One day after the final injection, isolated LCs were stained using the BrdU Flow Kit,

according to the manufacturer’s instructions (BD Pharmingen).

11. Fluorescein isothiocyanate (FITC)-induced cutaneous DC migration

Mice were shaved on the abdomen, and 200 μL of the 0.5% FITC (Sigma) solution in

acetone/dibutylphthalate (1:1) was applied to the shaved area. The draining axillary and

inguinal LNs were harvested, and cells were analyzed by flow cytometry at the indicated

time points.

12. Contact hypersensitivity and hapten-specific T-cell responses

2,4-dinitrofluorobenzene (DNFB; Sigma) was used to induce CHS. Mice were

sensitized by the application of 25 μL of 0.5% (wt/vol) DNFB in acetone/olive oil (4:1) to

their shaved abdomens on day 0 and challenged on the right ear on day 5 with 20 μL of 0.3%

(wt/vol) DNFB. Ear thickness was measured before and after challenge using a thickness

gauge. For hapten-specific T-cell responses, mice were sensitized with 25 μL of 0.5%

DNFB, and inguinal and axillary LNs were harvested 5 days later. Total LN cells were

stained with 2.5 μM carboxyfluorescein succinimidyl ester (CFSE; Molecular Probes) for

15 min at 37°C and washed, and 1×106 LN cells/well in a 96-well plate were cultured in

the complete RPMI medium with or without 50 μg/mL 2,4-dinitrobenzene sulfonic acid

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(DNBS; Alfa Aesar, Ward Hill, MA, USA), which is a water-soluble compound with the

same antigenicity as DNFB, for 3 days. The cells were then harvested and re-stimulated

with 50 ng/mL phorbol myristate acetate (PMA; Sigma) and 500 ng/mL ionomycin (Sigma)

in the presence of GolgiStop (BD Biosciences) for 5 hr at 37°C, and subsequent

intracellular staining for IFN-γ was performed. The secretion of IFN-γ was examined by

using the Cytometric Bead Array (BD Biosciences) following the manufacturer’s

instructions.

13. Croton oil-induced irritant dermatitis

Twenty μL of 1% (vol/vol) croton oil (Sigma) in acetone was applied to ear skin and ear

thickness was measured before and after treatment at the indicated time points.

14. Epicutaneous sensitization and T cell adoptive-transfer

Mice were shaved on the back with an electric razor and a 1 x 1 cm size of skin site was

tape-stripped at least 4 times using cellophane tape (Nichiban, Tokyo, Japan). One hundred

μg of ovalbumin (OVA) (Sigma) in 50 μL of normal saline was applied using Finn

Chamber (SmartPractice). A total of three 2 days exposures separated by 1 day intervals

were performed. For T cell adoptive-transfer, CD4+ T cells were isolated from

CD45.1+OT-II mice using beads-conjugated anti-CD4 mAb and magnetic bead separation

(Miltenyi Biotec), and labeled with 10 μM of CFSE. Totally, 4 x 106 labeled cells were

adoptively transferred into recipient mice via retro-orbital routes and mice were

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epicutaneously sensitized with OVA 24 hr later. Brachial SDLNs were harvested 72 hr

after OVA sensitization, and cell suspensions were analyzed for diminution of CFSE by

flow cytometry.

15. Clinical and histological evaluation of epicutaneously sensitized skin

The sum of severity of each clinical category for skin lesions was calculated, graded as 0

(none), 1 (mild), 2 (moderate), and 3 (severe), for the symptoms of erythema, edema,

erosion, scaling, and pruritus. Skin tissues were fixed in 10% formalin and embedded in

paraffin for the histological assessment. Skin sections (6 μm thick) were prepared and

stained with hematoxylin and eosin. As epicutaneously sensitized skins revealed the

features of eczematous dermatitis, we added a score for epidermal spongiosis for the

histological grading system. A grading scale we used is as follows: 0, no response; 1,

minimal response; 2, mild response; 3, moderate response; and 4, marked response.

16. Enzyme-linked immunosorbent assay (ELISA) for OVA-specific antibody

detection

OVA-specific IgE/IgG1/IgG2a levels were measured by an indirect ELISA method

using mouse IgE/IgG1/IgG2a ELISA kit (eBioscience) with some modifications. ELISA

plates were coated with OVA (500 ng/well) at 4˚C overnight. After blocking, 50 μL of 20x

diluted serum was added to each well and incubated for 2 hr. IgE levels were determined

using biotin-conjugated anti-mouse IgE Abs and streptavidin-conjugated horseradish

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peroxidase (eBioscience). Alkaline phosphatase-conjugated anti-mouse IgG1/IgG2a Abs

(Santa Cruz Biotechnology, Santa Cruz, CA, USA) were also used to detect OVA-antibody

complex. Optical density was measured at 450nm.

17. Chemotaxis migration assay

Cell migration was evaluated using the 24-well, 5 μm pore size polycarbonate Transwell

system (Costar, Cambridge, MA, USA). Total of 1 x 105 LPS-stimulated GFP+ MDLCs

were placed on the top of the Transwell in complete RPMI medium and incubated with or

without the indicated concentrations of recombinant human C-C chemokine ligand 19

(CCL19) (Peprotech) in the bottom chamber for 3 hr at 37˚C. The number of migrating

cells was counted for a fixed time period of 1 min using flow cytometry. Chemotaxis index

was calculated as the ratio of the number of cells migrating toward CCL19 divided by the

number of migrating cells in the negative control medium.

18. Quantitative real-time polymerase chain reaction

Total RNA from purified cells was isolated with the Hybrid-R Total RNA kit (GeneAll

Biotechnology, Seoul, Korea). cDNA was synthesized using PrimeScript™ RT Master

Mix (Takara Bio, Shiga, Japan). Quantitative real-time PCR reaction was performed with

the ABI StepOnePlus real-time PCR system (Applied Biosystems, Foster City, CA, USA)

by monitoring the synthesis of double-stranded DNA during the various PCR cycles using

SYBR Green (Takara Bio). For each sample, duplicate test reactions were analyzed for the

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expression of the gene of interest, and results were normalized to Gapdh mRNA. Primer

sequences are listed in Table 1.

Table 1. Primer sequences used for real-time qPCR

Genes Forward Reverse

Bub1 GAGATGCTCAGTAACAAGCCA GGTTTCCAGACTCCTCCTTCA

Cadm3 AGGGATTGTGGCTTTCATTG TCGCTTCGTGTGTCAGGTAG

Ccl3 GTACCATGACACTCTGCAACC GTCAGGAAAATGACACCTGGCTG

Ccnd1 CCCCAACAACTTCCTCTCCT GGCTTCAATCTGTTCCTGGC

Ccr7 CCAGCAAGCAGCTCAACATT GCCGATGAAGGCATACAAGA

Cd69 GGATTGGGCTGAAAAATGAA CTCACAGTCCACAGCGGTAA

Cdh17 CACGACCAATGCAGGATATG CACCAGTGGCTCAGGATTTT

Cdk1 CATCCCACGTCAAGAACCTG GGTGCTTCAGGGCCATTTTG

Ctcf (mouse) AGCGCTATCATGATCCCAAC CGGCTCAGCATTTTCTTCAC

CTCF (human) TGACACAGTCATAGCCCGAAAA TGCCTTGCTCAATATAGGAATGC

Dmc1 GTGGACACATTCTGGCTCAC CATTTTCAGGCATCTCGGGG

Emb CCCCGGTACAGAAAAACTGA ACGTTGAGGGCATCTTTGTC

Gapdh (mouse) TGGCCTTCCGTGTTCCTAC GAGTTGCTGTTGAAGTCGCA

GAPDH (human) AGCCAAAAGGGTCATCATCTC GGACTGTGGTCATGAGTCCTTC

Klrc1 TGCAAAGGTTTTCCATCTCC TGCTTCGGTATATGGTGTGG

Mnd1 GGAGGAAAAGAGAACCCGGA CTTCCTTCACTGACATGGCG

Nid1 TGGTTCCTCCTACACCTGCT TGTTGTAGCAGAAGGCATCG

Sema7a GGCGGAAGCTCTATGTGACC GACTGCAGCACTGATCGTTGG

19. Western blot

Total splenocytes were labeled with magnetic bead-conjugated anti-CD11c mAb

(Miltenyi Biotec), and DCs were purified by two rounds of positive selection on

paramagnetic columns (Miltenyi Biotec). Cells were lysed in 100 μL cell lysis buffer

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containing 50 mM Tris-HCl (pH 7.5), 150 mM NaCl, 1% nonyl

phenoxypolyethoxylethanol, 1 mM ethylenediaminetetraacetic acid, 5% glycerol, and

protease inhibitor cocktail (Sigma). Whole cell lysates were resolved on sodium dodecyl

sulfate-polyacrylamide gels and transferred onto a polyvinylidene fluoride membrane.

After blocking with 5% skim milk, the membrane was incubated with the antibodies,

followed by incubation with the horseradish peroxidase-conjugated secondary antibody.

Target proteins were visualized using SuperSignal West Pico Chemiluminescent Substrate

(Pierce, Rockford, IL, USA).

20. Blood cell count

Whole blood was collected from the superficial facial vein into EDTA-treated tube at

the indicated time points. Total blood cell count was evaluated by HEMAVET 950FS

(Drew Scientific).

21. Bone marrow transplantation

Total 5 x 106 BM cells from CreER and CTCF-cKO mice (CD45.2) were transplanted

into lethally irradiated (10Gy) B6.SJL mice (CD45.1). For generating mixed BM chimeras,

2.5 x 106 BM cells from CreER or CTCF-cKO mice (CD45.2) were mixed with equal

numbers of BM cells from B6.SJL (CD45.1) mice and transferred into the irradiated

B6.SJL (CD45.1) mice. Six to eight weeks after transplantation, Cre-mediated

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recombination was induced by daily tamoxifen injection, and BM and spleen cells were

harvested from reconstituted mice and analyzed.

22. Cell cycle assays

Cells were incubated in the complete RPMI medium containing 10 μg/ml of Hoechst

33342 (Molecular Probes) for 45 min at 37°C. After surface antigen staining, cells were

fixed and permeabilized using the Foxp3 staining kit (eBioscience) and subsequent nuclear

staining of Ki-67 was performed.

23. Measurement of reactive oxygen species

Total BM cells were incubated with 1 μM of 5-(and-6)-carboxy-2’,7’-

Dichlorofluorescein diacetate (carboxy-DCFDA; Molecular Probes) for 30 min at 37°C in

the dark. Then, cells were stained for lineage and HSCs markers at 4°C and assayed by

flow cytometer.

24. Colony-forming unit assay

Colony-forming unit assay was performed using Methocult (M03434, Stem Cell

Technologies, Vancouver, Canada). The number of rising colonies was counted for CFU-

GEMM, CFU-GM, and BFU-E at the indicated time points according to the

manufacturer’s instructions. In brief, 2 x 104 total BM cells were plated in methylcellulose

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medium containing recombinant mouse SCF, IL-3, IL-6, and erythropoietin and cultured in

the 37℃ and 5% CO2 incubator for 10 to 14 days.

25. Microarray

Epidermal LCs and BM LSKs were sorted by the FACSAria II cell sorter (BD

Biosciences) at the flow cytometry core in the Avison Biomedical Research Center

(Yonsei University College of Medicine). Sorted cells were immediately collected in 500

μL TRIzol (Invitrogen), and total RNA was extracted using the isopropanol precipitation

method. Isolated RNA was linearly amplified by the WT Expression Kit (Ambion, Austin,

TX, USA) and labeled using WT Terminal Labeling Kit (Ambion). Labeled cRNA was

hybridized on the Affymetrix Mouse GeneChip 2.0 ST Arrays (Affymetrix, Santa Clara,

CA, USA). Acquired expression data were analyzed by the R software (www.R-

project.org). Quality control was done using a different R package from the Bioconductor

project (www.bioconductor.org). The Significance Analysis of Microarray (SAM) was

used to determine genes that were significantly and differentially expressed between WT

and CTCF-deficient LCs (FDR; q<5%). We performed functional annotation with the

Gene Ontology biological process using the DAVID Functional Annotation tool

(http://david.abcc.ncifcrf.gov) from an expanded subset of gene whose expression differed

by at least 1.5-fold.

26. Statistics

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Data were analyzed with the unpaired Student’s two-tailed t-test, unless otherwise stated,

using the Prism software (GraphPad Software Inc., San Diego, CA, USA). P < 0.05 was

considered to be significant.

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III. RESULTS

PART A. CCCTC-binding factor controls the homeostatic maintenance and

migration of Langerhans cells

1. Decreased pool of multiple DC lineages in DC-specific CTCF-deficient mice

To examine the role of CTCF in the DC lineage in vivo, we generated DC-specific

CTCF-deficient mice (CD11c-cKO) (Figure 1A). Depletion of endogenous CTCF in the

DC lineage was confirmed in purified splenic CD11c+ DCs (Figure 1B).

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Figure 1. Schematic diagram showing Ctcffl/fl mice generation and CTCF deletion

efficiency. (A) Mice carrying the targeted Ctcftm1a(EUCOMM)Wtsi allele were bred to ACT-

FLPe transgenic mice to remove the LacZ/NeoR cassette and generate mice carrying a

conditional Ctcffl allele in which exon 8 was flanked by loxP sites. Mice carrying a

conditional Ctcffl allele were bred to Cre-expressing transgenic mice to generate

conditional Ctcf-deleted (CtcfΔ) knockout mice. Exons are represented by black boxes. (B)

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The efficiency of CTCF deletion was determined by western blotting of sorted CD11c+

cells of the spleen from WT and CD11c-cKO mice. Beta-actin served as the protein

loading control. Data represent two independent experiments with three mice per group.

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To assess whether CTCF deletion influenced overall DC homeostasis, we compared the

proportions of individual DC subsets in the lymphoid and non-lymphoid organs. Lymphoid

conventional DCs (cDCs) in the spleen and skin-draining lymph nodes (SDLNs) were

decreased by ~50% in CD11c-cKO mice as compared with wild-type (WT) mice, and the

numbers of plasmacytoid DCs (pDCs) were slightly reduced (Figure 2A, C, Figure 3).

Despite a decrease in lymphoid cDCs, their subset division into CD8α+ and CD11b+ cells

was comparable. We then analyzed heterogeneous DC populations in the skin and their

corresponding migratory counterparts as a representative of non-lymphoid tissue DCs,

which are comprised of epidermal LCs and dermal DCs (dDCs) (Figure 2B, C, Figure 3).

LCs were diminished by up to 40% in the epidermis of CD11c-cKO mice. Interestingly,

they exhibited far greater declines in the dermis and SDLNs, indicating impaired

distribution of CTCF-deficient LCs in their migrating compartments. CD103+ and CD103−

dDCs were also reduced in the dermis of cKO mice and were further decreased in SDLNs.

Lymphoid cDCs commonly develop from pre-cDCs, which are late-stage committed

progenitors. However, the percentage of pre-cDCs was unchanged in WT and CD11c-cKO

bone marrow (BM), suggesting that CTCF-deficient cDC quantities are not affected by

pre-cDC level (Figure 2D). These findings demonstrate a critical role for CTCF in

maintaining the overall population size of terminally differentiated DCs in vivo.

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Figure 2. Decrease in the general pool of DCs in mice with DC-specific CTCF

depletion in vivo. (A) Lymphoid organ DCs including pDCs (Siglec-H+CD11cint), cDCs

(Siglec-H−CD11c+MHC II+), CD8α+ cDCs (Siglec-H−CD11c+MHC II+CD8α+CD11b−),

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and CD11b+ cDCs (Siglec-H−CD11c+MHC II+CD8α−CD11b+) from the spleen and SDLNs

were analyzed by flow cytometry. (B) Population frequencies of cutaneous DCs in single-

cell suspensions derived from the epidermis, dermis, and SDLNs were analyzed by flow

cytometry. Epidermal LCs (CD45+MHC II+), DC populations among CD45+ leukocytes in

the dermis (including LCs [CD11c+MHC II+Langerin+CD103−], CD103+ dDCs

[CD11c+MHC II+Langerin+CD103+], and CD103− dDCs [CD11c+MHC

II+Langerin−CD103−]), and migratory skin DCs (CD11c+MHC IIhi) in SDLNs were

analyzed. (C) Normalized ratios (relative frequencies) of the percentages of DCs in

CD11c-cKO mice against the percentages of the corresponding WT DC subsets. (D) The

proportion of pre-cDCs (Lin−CD11c+MHC II–CD135+Sirpαlo) in the BM. Lin, lineage

(CD3, CD19, NK1.1, Ter119, and B220). Data were pooled from two independent

experiments with four to five mice per group. Error bars indicate standard error of the

mean (SEM).

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Figure 3. DC-selective CTCF deletion results in a decreased general DC lineage. The

proportion of lymphoid DCs in the spleen and SDLNs and non-lymphoid skin DCs were

analyzed by flow cytometry. Error bars represent the mean±SEM of pooled data from two

experiments with four to five mice per group. *, P<0.05; **, P<0.01; ***, P<0.001; ns, not

significant.

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2. Reduced cell number with distinct morphological and surface phenotypes of

CTCF-deficient LCs

Because the homeostasis of LCs is distinct from that of other DCs, we next established

LC-specific CTCF-deficient mice (huLang-cKO). The efficient excision of the Ctcf locus

was confirmed by genomic DNA polymerase chain reaction (PCR) using LCs sorted by

flow cytometry (Figure 4A). Whole epidermal sheet staining revealed fewer epidermal

MHC II+ LCs in huLang-cKO mice, which was similar to results obtained in the CD11c-

cKO epidermis (Figure 4B). The reduced number of LCs did not affect the frequency of

dendritic epidermal T cells (DETCs), in accordance with a previous report (Figure 4C).

CTCF-deficient LCs increased in cell size and projected more elongated dendrites, and the

MHC II staining intensity was higher in cKO LCs than WT cells. In the steady state,

CTCF-deficient LCs displayed slightly lower basal levels of CD86 but profoundly higher

CD80 and CD40 expressions (Figure 4D). Moreover, in vivo oxazolone (Oxa) treatment

also led to an altered pattern of CD86 and CD80 expression in cKO LCs, indicating that

CTCF differentially regulates each costimulatory molecule in LCs (Figure 4E). Langerin,

which is a marker of LCs, was decreased in CTCF-deficient LCs, and a sparse number of

Birbeck granules (BGs) in CTCF-ablated LCs was observed, thus confirming a positive

relationship between langerin and BG formation (Figure 4F).

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Figure 4. LC-selective loss of CTCF leads to decreases in epidermal LC numbers and

distinct phenotypic characteristics of LCs. (A) FACS gating strategies for isolating LCs

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and dendritic epidermal T cells (DETCs). Genomic DNA PCR for isolated DNA from LCs

and DETCs from WT and huLang-cKO mice as separated by agarose gel electroporation.

(B) Epidermal whole mounts from WT and huLang-cKO mice were stained for MHC II

(green), and the cellular densities for MHC II+ LCs were calculated (bar graph). (C) The

epidermal CD103+ DETC population was assessed by flow cytometry. (D) Expression of

MHC II, CD86, CD80, CD40, and Langerin in WT and CTCF-deficient LCs

(CD45+CD11c+). The shaded line represents the fluorescence minus one (FMO) control. (E)

Surface CD86 and CD80 expressions of LCs were assessed by flow cytometry with or

without 1% oxazolone (Oxa) treatment for 24 hr. (F) Tennis-racket-like Birbeck granules

(black arrows) from WT and CTCF-deficient LCs in the epidermis were examined by

electron microscopy. Bar=100 nm. Data represent at least two independent experiments

with three to six mice per group. Error bars indicate SEM. *, P<0.05; **, P<0.01; ***,

P<0.001; ns, not significant.

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3. CTCF-deficient LCs exhibit reduced homeostatic self-turnover

Since CTCF deletion led to fewer epidermal LCs, we determined whether CTCF was

responsible for maintaining the epidermal LC pool. CTCF variably contributes to apoptosis

in a cell type-specific manner. Enhanced CTCF expression leads to an increase in the

apoptosis of BM-derived DCs in vitro. However, no increases in apoptosis were observed

in CTCF-ablated LCs in situ (Figure 5A). Moreover, we did not observe differences

between annexin V+ population of WT and cKO LCs regardless of LC activation,

indicating that apoptosis does not contribute to the decrease of CTCF-deficient LCs

(Figure 5B). LCs sustain their pool via slow self-renewal in the steady-state epidermis.

Notably, significantly lower numbers of bromodeoxyuridine (BrdU)-incorporated LCs

were quantified in huLang-cKO mice (Figure 5C). Intracellular Ki-67 staining revealed a

lower proportion of cells within the active phases of the cell cycle in CTCF-deficient LCs

(Figure 5D). However, surface TGF-βRII and CSF-1R were similarly expressed in WT and

cKO LCs. This suggests that the CTCF deficiency-induced reduction in LC number is due

to an intrinsic defect of self-perpetuation, rather than insensitivity to external cytokine

signals.

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Figure 5. CTCF-deficient LCs exhibit self-proliferation impairment. (A) In situ

TUNEL assays were performed using MHC II and DAPI co-staining in frozen ear skin

sections from WT and huLang-cKO mice. Original magnification, ×200. (B) LC activation

was induced by 1% Oxa treatment, and apoptosis of LCs (CD45+MHC II+) in epidermal

single-cell suspensions from mouse ears was analyzed by flow cytometry. (C) Intracellular

BrdU levels in LCs (CD45+MHC II+Langerin+) were assessed by flow cytometry. (D) The

actively cycling LC subpopulation was analyzed by detecting Ki-67 positivity. (E) TGF-

βRII and CSF-1R expression levels in LCs were analyzed. Data represent at least two

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independent experiments with three to seven mice per group. Error bars indicate SEM. **,

P<0.01.

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We next examined the initial phase of LC development. Similar infiltrations of

CD11b+F4/80+ myeloid precursors were observed in the WT and huLang-cKO postnatal

day 1 epidermis (Figure 6A, B). Equivalent expansion of MHC II+ cells was seen in day 7

epidermis, and almost all epidermal myeloid leukocytes strongly expressed MHC II and

Langerin (Figure 6B, C). Thus, the decreased number of CTCF-deficient LCs was largely

attributed to reduced LC proliferation capacity after completion of the LC network.

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Figure 6. Early epidermal LC network formation is comparable between WT and

huLang-cKO mice. (A) The frequency of myeloid LC precursors (CD45+CD11b+F4/80+)

in the epidermis on postnatal day 1 was assessed by flow cytometry. (B and C) The

numbers of MHC II+ cells (green) in the epidermis from postnatal days 1 and 7 were

quantified by immunofluorescence (B), and the expressions of MHC II and intracellular

langerin were examined by flow cytometry (C). Data were pooled from three experiments

with four to six mice per group. Error bars indicate SEM. ns, not significant.

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4. CTCF deficiency impairs the emigration of LCs from the epidermis

As the relative LC frequencies in the dermis and SDLNs were lower than those in the

epidermis of CD11c-cKO mice, we next examined the efficacy of LC migration. In

huLang-cKO mice, discordant relative ratios of LCs in the epidermis (~30%), dermis

(<10%), and SDLNs (<10%) were found, suggesting hampered migration of CTCF-

deficient LCs out of the epidermis (Figure 7).

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Figure 7. CTCF-deficient LCs are significantly less distributed in their sequential

migratory compartments. (A-C) DC populations in the epidermis (A), dermis (B), and

SDLNs (C) from WT and huLang-cKO mice were analyzed by flow cytometry, using the

same gating strategies described in Fig 1. (D) Normalized ratios (relative frequencies) of

the percentages of LCs in huLang-cKO mice against the percentages of corresponding WT

LCs in the epidermis, dermis, and SDLNs. Data were pooled from five experiments (n=11).

Error bars indicate SEM. ***, P<0.001; ns, not significant.

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To assess the full migratory capacity of CTCF-deficient LCs in vivo, we evaluated

antigen-bearing LCs in regional LNs by painting the skin with fluorescein isothiocyanate

(FITC). Consistent with a previous report, the number of FITC+ migrating LCs gradually

increased in WT mice (peak at 72 to 96 hr) (Figure 8A, B). Importantly, the draining LNs

of huLang-cKO mice had a remarkably smaller number of FITC+ LCs, whereas similar

proportions of FITC+ dDC migrants were observed. C-C chemokine receptor 7 (CCR7) is

critical for peripheral DC migration toward draining LNs. Notably, CTCF-deficient LCs

expressed a significantly lower level of surface CCR7 (Figure 8C).

Next, we sought to examine whether CTCF-deficient LCs could leave the epidermis. We

cultured mouse ear skin explants for 72 hr to induce LC migration and performed flow

cytometry for crawl-out cells and epidermal sheet staining for remnant ear tissues. The

number of crawl-out LCs from cultured huLang-cKO ear skins was strikingly reduced

compared with WT skin cultures (Figure 8D). Moreover, CTCF-deficient LCs were

retained in the cultured epidermis, while WT epidermal LCs considerably disappeared after

72 hr of skin culture (Figure 8E). Therefore, our in vivo and in vitro findings demonstrate a

critical role for CTCF in efficient LC egress from the epidermis.

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Figure 8. CTCF-deficient LCs fail to egress from the epidermis. (A, B) SDLNs were

analyzed by flow cytometry to quantify migrating DCs after FITC application.

Representative FACS plots for the migrating cutaneous DCs at the indicated time points

are presented (three to four mice per group). (C) Surface CCR7 expression of

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CD45+CD11c+MHC II+ LCs. The number indicates the mean fluorescence intensity ± SEM

for CCR7 staining. (D) Mouse ear skin explants were cultured for 72 hr, and crawl-out

cells in the supernatants were analyzed by flow cytometry to quantify migrating LCs

(MHC II+CD103–CD11c+Langerin+). (E) Epidermal sheets were prepared from resting and

cultured skin, and the number of LCs in the epidermis was assessed by

immunofluorescence after staining for MHC II (green). Data are representative of two to

three independent experiments with three to six mice per group. Error bars indicate SEM.

**, P<0.01; ***, P<0.001.

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Human LCs can be derived from peripheral monocytes in vitro (Figure 9A). We further

assessed the role of CTCF in human LCs using a lentiviral vector-mediated RNA

interference of CTCF. The use of Vpx-containing virus like particles (Vpx-VLP)

significantly enhanced lentiviral transduction efficiency, and human monocyte-derived

LCs (MDLCs) that were green fluorescent protein (GFP)-positive (indicative of successful

viral transduction) were subjected to fluorescence-activated cell sorting (FACS) (Figure

9B). The expressions of CTCF mRNA and protein were significantly decreased in MDLCs

infected with lentiviruses containing shRNA against CTCF (Figure 9C, D). Although the

transduction efficiencies between control and shCTCF-expressing lentiviruses were

comparable, the recovered MDLC number was reduced in the CTCF knockdown group

(Figure 9E). CTCF-diminished human MDLCs showed greater increases in CD80

expression after treatment with lipopolysaccharide (LPS), but CD86 levels were modestly

affected by CTCF silencing (Figure 9F). Notably, CTCF knockdown in LPS-treated

MDLCs resulted in a lower level of surface CCR7 compared to the control cells, similar to

results obtained with murine CTCF-deficient LCs (Figure 9G). Moreover, in vitro

chemotaxis assay revealed that MDLC migration toward C-C chemokine ligand 19

(CCL19) was significantly abrogated by CTCF knockdown (Figure 9H). These data

suggest that CTCF plays an important role in the CCR7-dependent migration of LCs in

both humans and mice.

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Figure 9. CTCF-silenced MDLCs showed a migration defect toward CCL19. (A)

Differentiation of MDDCs and MDLCs assessed by flow cytometry. (B) MDLCs were

infected with pGIPz control lentivirus in the absence or presence of Vpx-VLP, and

transduction efficiency was determined by frequency of GFP+ populations at day 8

(multiplicity of infection=2.0). (C and D) Transduced MDLCs were sorted, and the

knockdown efficiency of CTCF was determined by real-time qPCR (C) and

immunoblotting (D). (E) Representative FACS plot for the transduction efficiency of

MDLCs with control and shCTCF-containing lentiviruses (left), and the number of

recovered MDLCs after gene transduction (right). (F) Surface CD80 and CD86 levels of

FACS-sorted GFP+ MDLCs were analyzed after 2 days of LPS treatment. Data are

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representative of five individuals. (G) CCR7 levels of the transduced human MDLCs

stimulated with lipopolysaccharide (LPS) (representative data of three individuals). (H)

Chemotaxis indices of the GFP+ MDLCs migrating toward the indicated concentrations of

human CCL19 (pooled data from four individuals). Error bars indicate SEM. *, P<0.05.

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5. CTCF inactivation in LCs augments cell adhesion molecule gene expression

To identify CTCF-regulated genes in LCs, we purified WT and CTCF-deficient LCs by

FACS and performed microarray-based gene expression profiling (Figure 10A, B). We

found lower Ccr7 expression in cKO LCs, which was confirmed by flow cytometry. In

particular, systematic Gene Ontology analysis showed that cell adhesion-associated genes

were significantly different (Figure 10C). Among them, genes that promote cellular

adhesion such as cell adhesion molecule 3 (Cadm3), cadherin 17 (Cdh17), and nidogen-1

(Nid1) were highly expressed in CTCF-deficient LCs. To explore whether these adhesion

molecules were physiologically relevant to LC migration, we treated mouse ears with Oxa

to initiate LC migration and analyzed gene expression changes from the sorted LCs.

Notably, Cadm3, Cdh17, and Nid1 levels were significantly decreased in WT LCs upon

Oxa treatment. However, both resting and stimulated CTCF-deficient LCs expressed

strikingly higher levels of those adhesion molecules compared to WT LCs (Figure 10D).

Interestingly, the level of Ctcf in LCs was not significantly influenced by LC activation in

vivo (Figure 10E). This suggests that those adhesion molecules are not regulated by direct

changes in CTCF expression level, but possibly through other mechanisms such as a

differential protein interaction of CTCF which modulates its target gene’s chromatin

structures.

It has been known that LC motility in the epidermis is partly mediated by some adhesion

molecules including E-cadherin and epithelial cell adhesion molecule (EpCAM). Although

no differences in E-cadherin were observed, CTCF-deficient LCs expressed significantly

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lower levels of EpCAM, which led to enhanced binding of LCs to keratinocytes (Figure

10F). Collectively, these results indicate that CTCF is indispensable for efficient LC

emigration by regulating the balanced expression of CCR7 and cell adhesion molecules.

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Figure 10. CTCF-deficient LCs express higher levels of cell adhesion molecules. (A)

LCs (CD45+CD11c+MHC II+) were sorted from epidermal single-cell suspensions. Scatter

plots show raw expression levels (log2), and the selected genes are indicated in red

(upregulated in cKO LCs) or blue (upregulated in WT LCs). Green lines indicate the

twofold boundary. Black indicates the number of genes that exhibited greater than twofold

increases in WT (lower) or cKO (upper) LCs with a false discovery rate <0.05. (B)

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Expressions of the indicated genes in FACS-sorted WT and cKO LCs (same gating

strategies from Fig E3, A) were analyzed by real-time qPCR (top). CD69 protein

expression in WT and cKO LCs was assessed by flow cytometry (bottom). (C) The heat

map shows genes that belong to the cell adhesion biological process, as revealed by the

Gene Ontology analysis. Genes that were confirmed by real-time quantitative polymerase

chain reactions (qPCR) are indicated in bold. (D and E) Epidermal LCs were sorted with or

without the treatment with 1% Oxa, and the expression levels of Cadm3, Cdh17, Nid1 (C),

and Ctcf genes (D) were examined by real-time qPCR. (F) Surface E-cadherin and EpCAM

levels in LCs. Data were pooled from two experiments with three to six mice. Error bars

indicate SEM. *, P<0.05; **, P<0.01; ***, P<0.001; ns, not significant.

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6. CTCF-dependent LC homeostasis is required for optimal immunity of the

skin

Recent studies have proposed an immune-suppressive role for murine LCs by showing

enhanced delayed-type hypersensitivity (DTH) to contact allergens in mice with selective

LC deficiency and impaired LC migration, although the exact role for LCs remains

controversial. As our huLang-cKO mice showed decreases in LC number and defective LC

migration, we firstly assessed 2,4-dinitrofluorobenzene (DNFB)-induced contact

hypersensitivity (CHS) responses. DNFB treatment induced similar extents of ear swelling

in WT and huLang-cKO mice at 24 hr after challenge. However, cutaneous inflammation

was significantly sustained in huLang-cKO mice from 48 hr post-treatment (Figure 11A).

Ear swellings without DNFB sensitization and croton oil-induced irritant dermatitis were

equivalent between WT and cKO mice, indicating that the residual CTCF-deficient LCs

were sufficient to reduce irritant reaction as opposed to the results from mice with

complete LC depletion (Figure 11A, B). Because interferon (IFN)-γ-producing CD4+ and

CD8+ T cells mediate CHS in response to DNFB, we examined the effect of LC-specific

CTCF deficiency on the proliferation and priming of IFN-γ-producing T cells in draining

LNs. Antigen-specific T-cell proliferation was not significantly different between

sensitized LN cells from WT and cKO mice (Figure 11C). However, total sensitized LN

cells from huLang-cKO mice produced greater amounts of IFN-γ than those from WT

mice (Figure 11D). This was associated with significantly enhanced induction of IFN-γ-

producing CD8+ T cells in huLang-cKO LNs (Figure 11E). In contrast, the proportion of

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the IFN-γ+ population within CD4+ T cells was marginally affected by LC-selective CTCF

deficiency. In addition, the frequency of Foxp3+ regulatory T cells in SDLNs was

comparable between WT and cKO mice irrespective of sensitization (Figure 11F). These

findings suggest that the migratory failure of LCs due to CTCF deficiency facilitates the

cross-priming of IFN-γ-producing effector CD8+ T cells and, in turn, enhances CHS

reactions, which indicate a suppressive role for LCs against the haptenized antigen.

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Figure 11. Prolonged and exaggerated CHS responses with higher priming of IFN-γ-

producing CD8+ T cells in huLang-cKO mice. (A) Changes in ear thickness after DNFB

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challenge were measured at the indicated time points. (B) Changes in ear thickness after

application of croton oil or vehicle were measured at the indicated time points. (C) DNFB-

sensitized total LN cells were labeled with CFSE and cultured with or without DNBS for

72 hr. CD44+CFSElow antigen-specific T-cell proliferation was examined by flow

cytometry. (D) DNFB-sensitized total LN cells were cultured for 72 hr with or without

DNBS. IFN-γ levels in the supernatants were evaluated. (E) Cultured total LN cells were

re-stimulated with PMA/ionomycin, and intracellular IFN-γ production among CD8+ and

CD4+ T cells was assessed by flow cytometry. (F) Foxp3+ regulatory T cells among the

CD4+ T cell population in SDLNs were analyzed by flow cytometry 5 days after treatment

with vehicle or DNFB. Data represent three independent experiments with five to seven

mice per group. Error bars indicate SEM. *, P<0.05; ***, P<0.001; ns, not significant.

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Aside from a CHS response, LCs are known to elicit Th2 and humoral immunity to

protein antigens that encounter the epidermal barrier. To further understand the

consequence of CTCF deficiency in LCs, we epicutaneously sensitized mice with

ovalbumin (OVA) patches. Repeated OVA application led to local dermatitis in WT mice;

however, huLang-cKO mice showed a less severe inflammation, which correlated with the

reduced Th2-associated OVA-specific serum IgE and IgG1 levels in cKO mice (Figure

12A-C). Furthermore, huLang-cKO mice exhibited decreased OVA-specific T cell

proliferation after a single epicutaneous exposure to OVA (Figure 12D). Altogether, our

data demonstrate that CTCF-dependent LC homeostasis is functionally required for full

induction of cutaneous adaptive immunity to a protein antigen.

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Figure 12. Attenuated epicutaneous sensitization to OVA in huLang-cKO mice. (A)

Total clinical severity scores. (B) Hematoxylin and eosin staining of the patch-applied

back skins. The sum of each histological score is presented. Scale bar=100 μm. (C) Serum

OVA-specific antibodies were assessed by ELISA. OD values for IgE, IgG1, and IgG2a

were measured at a wavelength of 450 nm. (D) CFSE-labeled CD45.1+OT-II T cells were

adoptively transferred to WT and huLang-cKO mice, and mice were treated with an OVA

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patch. SDLNs were analyzed to measure OVA-specific T cell proliferation. Numbers in the

histogram indicate the number of CD45.1+OT-II T cell divisions. Data represent at least

two independent experiments with five to seven mice per group. Error bars indicate SEM.

*, P<0.05; ***, P<0.001.

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PART B. CCCTC-binding factor regulates adult hematopoietic stem cell

quiescence in mouse

1. Loss of CTCF leads to rapid BM failure

To examine the relative expression levels of Ctcf mRNA in primitive hematopoietic

stem/progenitor cells, we analyzed the Immunological Genome microarray database,

which encompasses a large group of immune cell and their progenitor populations62. Ctcf

mRNA was expressed in all subsets of HSCs and progenitor cells examined, although

GMP expressed a slightly lower Ctcf expression level (Figure 13A). To determine whether

CTCF is required for general hematopoiesis in vivo, we used the Cre-loxP system to

deplete Ctcf gene by injecting tamoxifen (TMX) in adult mice. We crossed ROSA26-

CreER mice with conditional Ctcf-floxed mice63 (hence, CTCF-cKO) and induced an

efficient Ctcf ablation by intraperitoneal injection of TMX for 5 consecutive days (Figure

13B, C). Within 10 to 14 days after TMX treatment, all CTCF-cKO mice showed a rapid

lethality while the survival of WT mice was not affected by TMX (Figure 13D). To

evaluate the overall hematopoiesis in the absence of CTCF, we analyzed peripheral blood

cell counts. Although both genotypes of mice showed normal white blood cell and platelet

counts before CTCF deletion, TMX treatment led to a significant reduction of those

hematological parameters in CTCF-cKO mice (Figure 13E). Red blood cell counts were

slightly declined both in the TMX-treated WT and CTCF-cKO groups without a

significant difference between the groups, indicating that TMX alone would have a

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possible adverse effect on erythropoiesis. Importantly, CTCF-ablated mice had a markedly

lower total BM cellularity than did the WT control mice (Figure 13F). These data indicate

that CTCF is critically involved in hematopoiesis and acute loss of CTCF results in rapid

hematopoietic failure.

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Figure 13. Acute CTCF ablation leads to hematopoietic failure. (A) Relative

expression level of Ctcf gene among the various stem/progenitors (Data from ImmGen). (B,

C) WT and CTCF-cKO mice were treated with TMX for 5 consecutive days and genomic

DNA and total RNA were isolated at day 8. Efficient Ctcf gene deletion by TMX treatment

in CTCF-cKO mice at DNA (B) and RNA (C) level is shown. (D) Kaplan-Meier curve

plotting survival of WT and CTCF-cKO mice (n = 10) is shown. (E) Peripheral blood

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counts of controls and CTCF-cKO mice 8 and 10 days after first TMX injection (n = 4−5)

are shown. (F) Bone marrow counts (femurs and tibia) of WT and CTCF-cKO mice 8 days

after TMX treatment (n = 5) are shown. Data were from at least three independent

experiments with four to five mice per group. Error bars indicate standard error of the

mean (SEM). **, P<0.01; ***, P<0.001; ns, not significant.

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2. Acute depletion of CTCF results in rapid shrinkage of c-Kithi HSC and

progenitor pool

Since TMX-induced systemic depletion of CTCF led to rapid BM failure, we next

examined heterogeneous HSC population by flow cytometry. CD117/c-Kit is a stem cell

factor (SCF) receptor which is highly expressed on the most primitive stem/progenitor

cells in BM64,65. Surprisingly, within 8 days after CTCF ablation in vivo, we found a

marked shrinkage of Lineage−c-Kithi population in BM (Figure 14A). CTCF-ablated

lineage-negative BM cells also revealed a decreased expression of CD135/Flt3 and CD34,

but the level of surface Sca-1 and CD16/32 were rather increased than did WT cells

(Figure 14D). Lineage−c-Kithi population can be divided into LSK (Lineage− Sca-1+c-Kithi)

and LK (Lineage− Sca-1−c-Kithi) population which is HSC- and early myeloid progenitor

(CMP, GMP, and MEP)-enriched compartment, respectively. In line with the acute

hematopoietic failure, we found a marked decrease in both LSK and LK populations after

CTCF ablation (Figure 14B). Since HSCs are comprised of a hierarchical stage of stem

cells20, we further assessed HSCs in the LSK compartment using different surface markers.

The most primitive long-term reconstituting HSCs (LT-HSCs), which are able to be

determined by CD150+ CD48− surface characteristics within LSKs were profoundly

diminished in TMX-treated BM of CTCF-cKO mice (Figure 14C). In addition, the number

of short-term HSCs and other primitive progenitors in LSKs were also generally decreased

by CTCF deletion compared to those of WT controls. Thus CTCF is required for the

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homeostatic maintenance of c-Kithi primitive stem/progenitor cells and regulates variable

surface receptor expressions of the undifferentiated hematopoietic compartment.

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Figure 14. Acute depletion of CTCF results in severe decrease in c-Kithi

stem/progenitors. (A) Surface c-Kit level of lineage-negative population is shown (B) The

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frequencies and absolute numbers of LSK (lineage− Sca-1+ c-Kithi) and LK (lineage− Sca-

1− c-Kithi) population of WT and CTCF-cKO mice 8 days after TMX injection are shown.

(C) LT-HSCs (CD150+ CD48−), ST-HSCs (CD150− CD48−), MPP1 (CD150+ CD48+), and

MPP2 (CD150− CD48+) populations within LSKs are shown. (D) Surface expression level

of Flt3, CD34, Sca-1, CD16/32 are evaluated by flow cytometry from lineage-negative

populations of TMX-treated WT and CTCF-cKO mice. Data were from at least three

independent experiments with three to four mice per group. Error bars indicate SEM. *,

P<0.05; **, P<0.01; ***, P<0.001; ns, not significant.

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3. Loss of CTCF in hematopoietic compartment results in rapid exhaustion of

HSCs

Since TMX-responsible CreER is expressed in ubiquitous tissues in ROSA26-CreER

mice, the observed hematopoietic failure and rapid lethality after inducing CTCF depletion

in CTCF-cKO mice might result from dysfunction of non-hematopoietic compartment. To

determine this issue, we generated BM chimeras using lethally irradiated CD45.1+

recipients reconstituted by WT or CTCF-cKO BM, respectively. BM chimerism was

achieved constantly ~98% (Figure 15A). We found that TMX-treated chimeras

transplanted with CTCF-cKO BM revealed a markedly decreased survival and total BM

cellularity (Figure 15B, C) compared to those of control chimeras. Peripheral blood

analysis demonstrated a similar hematopoietic failure in TMX-treated CTCF-cKO BM

chimeras as seen in the systemic CTCF-deficient mice (Figure 15D). In line with the

decreased number of red blood cells and platelets in blood, CTCF-deficient BM-harboring

chimeras revealed severe anemic and hemorrhagic clinical features (Figure 15E).

Furthermore, TMX treatment led to a rapid loss of c-Kithi HSC/myeloid progenitor

populations and CD150+CD48− LT-HSCs in the BM-specific CTCF-null chimeras (Figure

15F). These results indicate that ablation of CTCF in hematopoietic compartment is

sufficient to induce rapid exhaustion of HSCs followed by resultant hematopoietic failure

and mortality.

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Figure 15. CTCF loss in the hematopoietic system leads to severe BM failure and

consequent lethality. (A) Chimerism efficiency analyzed from lineage-negative BM cells

of CD45.1+ recipient transplanted with CD45.2+ donor BM. (B) Kaplan-Meier curve

plotting survival of WT → CD45.1 and CTCF-cKO → CD45.1 chimeric mice (n = 9) is

shown. (C) Total BM cellularity from each chimera is shown. (D) Peripheral blood counts

of controls and CTCF-cKO chimeric mice 9 days after first TMX injection (n = 5) are

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shown. (E) Clinical pictures of WT and CTCF-cKO chimeras showing anemic (upper and

middle) and spontaneous hemorrhagic (lower) features. (F) The frequencies and absolute

numbers of LSK and LT-HSCs of WT and CTCF-cKO chimeras 8 days after TMX

injection are shown. Data were from at least two independent experiments with three to

four mice per group. Error bars indicate SEM. *, P<0.05; **, P<0.01; ***, P<0.001; ns, not

significant.

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HSC homeostasis is finely regulated by heterogeneous cross-talk between HSCs and

surrounding stromal components within the BM niche.66 As TMX-induced ablation of

CTCF in ROSA26-CreER mice also occurred in BM stromal cells, CTCF deficiency-

mediated HSC exhaustion and accompanied BM failure might result from the HSC-

extrinsic effects. To exclude this possibility, we transplanted CD45.2+ WT or CTCF-cKO

BM cells together with CD45.1+ supporting BM cells at 1:1 ratio into the lethally irradiated

CD45.1+ recipient mice. Six weeks after reconstitution, the mixed BM chimeras were

injected with TMX for inducing CTCF depletion in vivo and BM cells were analyzed at

least after 4 wks (Figure 16A). We found a scarce number of LSKs and CD150+ CD48−

LT-HSCs derived from CTCF-null donor, indicating that exhaustion of CTCF-deficient

HSCs was not rescued by normal BM niche (Figure 16B, C). Thus, genetic loss of CTCF

results in HSC exhaustion in a cell-autonomous manner.

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Figure 16. CTCF deficiency-mediated HSC depletion is a cell-autonomous effect. (A)

Experimental scheme for the generation and use of mixed BM chimeras. (B, C) The

frequencies of CD45.2+ BM-derived cells from the TMX-treated mixed chimeric mice are

shown. Data were from three independent experiments with four to five mice per group.

Error bars indicate SEM. ***, P<0.001.

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4. Defective HSC-driven myeloid lineage development by deletion of CTCF in

vivo

Because CTCF depletion led to a marked reduction of c-Kithi stem/progenitor population

in BM, we next determined the development of HSC-driven early myeloid progenitors and

subsequent peripheral myeloid cells in the absence of CTCF. In the spleen, all subsets of

dendritic cells (DCs), including plasmacytoid (Siglec-H+ CD11c+), conventional CD8α+

(Siglec-H− CD11chi CD8α+ CD11b−), and CD11b+ (Siglec-H− CD11chi CD8α− CD11b+) DCs

derived from CTCF-deficient BM were profoundly decreased than did WT BM (Figure

17A, C). Splenic monocytes (CD11bhi F4/80− SSClo Ly-6G−) and neutrophils (CD11bhi

F4/80− SSChi Ly-6G+), which develop from HSC-CMP-GMP axis also showed a marked

reduction arising from CTCF-ablated BM cells (Figure 17B, C). However, F4/80+ red pulp

macrophages in the spleen (CD11bint F4/80+), which are maintained by self-renewal

without substantial monocyte input, demonstrated rather partial decline after inducing

CTCF depletion, confirming their distinct homeostasis in the steady-state.67,68 In line with

the diminution of HSC-driven myeloid compartment in the spleen after CTCF deletion,

there was a negligible population of CTCF-deficient LK cells in the BM (Figure 17D). To

determine the role of CTCF in colony-forming capacity of stem and myeloid progenitor

cells, we performed in vitro colony-forming unit assays. To adjust the starting number of

stem/progenitor cells, total untouched BM cells either from WT and CTCF-cKO mice were

plated in methylcellulose medium supplemented with SCF, IL-3, IL-6, and erythropoietin

followed by in vitro TMX treatment. TMX-treated CTCF-cKO BM cells showed markedly

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decreased colony-forming units than did WT ones and there were no discernible sorts of

colonies derived from the CTCF-null BM cells (Figure 17E). These data indicate that

CTCF is critical for HSC-driven myeloid lineage development both in vivo and in vitro.

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Figure 17. Loss of CTCF results in exhaustion of myeloid progenitors and their

differentiation. (A, B) Gating strategies for the dendritic cells (A), red pulp macrophages,

monocytes, and neutrophils (B) in spleen are shown. (C) The frequencies of CD45.2+ BM-

derived myeloid lineage cells from the spleen of TMX-injected mixed chimeric mice are

shown. (D) The proportion of CD45.2+ BM-derived LK cells is shown. (E) The number of

total colonies from WT and CTCF-cKO mice cultured in methylcellulose medium

supplemented with myeloid-driving factors at day 9 is shown. Data were from at least three

independent experiments with four to five mice per group. Error bars indicate SEM. **,

P<0.01; ***, P<0.001.

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5. CLPs and lymphoid lineage cells are less sensitive to CTCF ablation in vivo

We next examined the role of CTCF in lymphoid lineage development in vivo.

Surprisingly, CTCF-ablated BM cells gave rise to both CD4+ and CD8+ T cells in the

spleen with a significant amount in contrast to the myeloid lineage (Figure 18A, C). B

lymphocyte development was also relatively preserved as the frequency of B cells from

CTCF-deficient BM was only reduced by 50% than that of WT CD45.2+ BM-derived B

cells (Figure 18B, C). Furthermore, we observed a discrete population of CTCF-deficient

CLPs from CTCF-ablated BM in the mixed chimeras similar to T and B lymphocytes

(Figure 18D). Therefore, our results implicate that the lymphoid lineage development is

only partially affected by CTCF depletion in vivo.

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Figure 18. CLP-derived lymphoid differentiation is less sensitive to CTCF depletion.

(A, B) Gating strategies for the T cells (A) and B cells (B) in spleen are shown. (C) The

frequencies of CD45.2+ BM-derived lymphoid lineage cells from the spleen of TMX-

treated mixed chimeric mice are shown. (D) The proportion of CD45.2+ BM-derived CLPs

is shown. Data were from at least three independent experiments with four to five mice per

group. Error bars indicate SEM. *, P<0.05; **, P<0.01; ***, P<0.001.

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6. CTCF-deficient HSCs are unable to maintain quiescence

To identify the genes regulated by CTCF in hematopoietic stem/progenitor cells, we

performed microarray-based gene expression analyses. LSK population was purely sorted

by flow cytometry after three times of TMX treatment and subjected to the microarray

experiment. Acute loss of CTCF led to changes in gene expression profile between WT

and CTCF-deficient LSKs (Figure 19A). Subsequent systematic gene ontology analysis

revealed that CTCF-depleted LSKs expressed high level of cell cycle-promoting gene

expression program (Figure 19B, C). In the steady state, maintenance of HSC quiescence

is critical for regulating the stem cell pool. To determine whether the HSC’s quiescence is

affected by CTCF deletion, we performed Ki-67/Hoechst staining. Importantly, in line

with the microarray results, CTCF-deficient LSKs showed a significantly increased

proportion of cells in the G1 and S-G2-M phases. In addition, CTCF deficiency led to a

marked reduction of G0 phase and an increased G1 and S-G2-M phases of LT-HSCs which

represented a quiescence defect (Figure 19D). Maintaining low ROS levels is essential to

HSC quiescence and apoptosis. Intracellular ROS production was examined using the CM-

H2DCFDA combined with flow cytometry. Importantly, CTCF-depleted CD150+ HSCs

displayed markedly increased levels of intracellular ROS both in acute systemic or BM-

confined CTCF ablation settings (Figure 19E, F). Furthermore, CTCF-deficient LSKs

demonstrated a significantly increased dead/apoptotic population than did WT cells

(Figure 19G). Therefore, our data indicate that CTCF is critically involved in maintaining

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HSC pool through sustaining HSC quiescence possibly downregulating intracellular ROS

level in vivo.

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Figure 19. CTCF-deficiency leads to an augmented cell cycle progression in HSCs. (A)

LSKs were sorted from BM 24 hr after the last TMX treatment. Scatter plots show raw

expression levels (log2), and the selected genes are indicated in red (upregulated in cKO

LSKs) or blue (upregulated in WT LSKs). Green lines indicate the twofold boundary. (B)

The heat map shows genes that belong to the cell cycle biological process, as revealed by

the Gene Ontology analysis. (C) Expression levels of the selected genes from (B) were

confirmed by real-time quantitative polymerase chain reactions (qPCR). (D) Ki-

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67/Hoechst-based cell cycle analysis of LT-HSCs from WT and CTCF-cKO mice 8 days

after TMX treatment is shown. (E, F) Intracellular level or ROS was measured by CM-

H2DCFDA from TMX-treated straight cKO mice (e) and chimeric mice (f) at day 8. Mean

fluorescence intensity (MFI) of ROS levels were calculated. (G) Cellular death/apoptosis

was analyzed by Annexin V/propidium iodide staining. Data were from at least three

independent experiments with four to five mice per group. Error bars indicate SEM. *,

P<0.05; **, P<0.01; ***, P<0.001.

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IV. DISCUSSION

In the steady state, most DCs, except for epidermal LCs and brain microglia, are

continuously replenished by precursors from the blood.51,52 Moreover, DCs undergo a

small number of divisions in the peripheries that are important for maintaining

homeostasis.69-71 In this study, we used DC-specific CTCF-deficient mice to demonstrate a

common but differential extent of reduction in each DC subset. As gene recombination in

CD11c-Cre mice fully occurs in differentiated DCs,56 an overall decrease in DC quantity

may result from CTCF deficiency in DCs. Similar proportions of pre-cDCs in the WT and

CD11c-cKO BM and decreases in pDCs and epidermal LCs, which develop independent

from pre-cDCs,72 highlight a critical role for CTCF in sustaining terminally differentiated

DC pools in vivo.

We then utilized a LC-specific CTCF-deficient mouse line to show that CTCF is

essentially involved in epidermal LC homeostasis. The reduction in epidermal LC number

in huLang-cKO mice was not attributed to a failure of the initial LC network formation or

a subsequent increase in cell apoptosis. Instead, CTCF-deficient LCs demonstrated a

significant deterioration in self-renewal capacity. Previous studies have shown that

conditional CTCF deletion in T cells leads to cell cycle blockage,46 and impaired

proliferation upon T-cell receptor stimulation.47 In line with those results from lymphoid

lineage cells, our findings suggest that CTCF played an important supporting role in

myeloid LC proliferation. However, CTCF overexpression during granulocyte macrophage

colony-stimulating factor (GM-CSF)-supplemented BMDC culture was reported to lead to

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the diminished proliferation.54 These discrepant observations may be due to the fact that

LC development strictly depends on TGF-β and IL-34, but not GM-CSF.

CTCF-deficient LCs demonstrated unexpected impairments in their egress from the

epidermis. LC migration is initiated by the downregulation of cell adhesion molecules in

response to pro-inflammatory cues, which facilitates the release of LCs from the

epidermis.73 A previous report suggested a role for CTCF in the migration of transformed

corneal cell lines in vitro, but the exact regulatory mechanisms were unclear.74 In our

microarray experiments, the expression levels of cell adhesion molecules including Cadm3,

Cdh17, and Nid1 were significantly greater in CTCF-deficient LCs, which may augment

LC attachment within the epidermis. Cadm3 is associated with the cell-cell adhesion of

neuronal cells.75 Cdh17 is a non-canonical cadherin that is frequently up-regulated in

cancer cells and mediates their adhesion.76,77 Notably, CTCF-deficient LCs highly

expressed Nid1, which can bind to laminin and collagen IV in the basement membrane.78

Because CTCF-deficient LCs were unable to efficiently transmigrate the dermo-epidermal

junction, we speculate that cKO LCs may be entrapped within the epidermis by an

increased affinity to the juxtaposed keratinocytes and basement membrane structure.

Importantly, those adhesion molecules were consistently diminished in WT LCs following

in vivo activation. Collectively, these results strongly suggest that CTCF intrinsically

supports the efficient egress of LCs from the epidermis by negatively regulating LC

adhesion intensity. In addition to the adhesion molecules, both mouse and human CTCF-

depleted LCs showed lower levels of CCR7 expression and impaired migration toward

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CCR7 ligand-rich areas. Thus, our findings provide relevant evidence supporting the

hypothesis that CTCF is involved in LC migration. At molecular level, the constitutive

expression of CTCF even after LC activation proposed that CTCF in LCs may work

through the differential DNA occupancy patterns16 and/or by collaboration with other

interacting proteins, such as transcription factors, cofactors, and cohesin complexes.79,80

Further studies will be necessary to dissect global CTCF binding profiles and its protein

interactomes in LCs to illuminate CTCF-dependent molecular mechanisms for regulating

its target genes.

Although there are conflicting results regarding the role of LCs, several recent studies

have shown that LCs can suppress CHS and other types of inflammation. In huLang-cKO

mice, few antigen-laden LCs could reach the draining LNs, whereas dDC migration was

comparable to that in WT mice. A recent study using a langerin-targeting antigen delivery

strategy demonstrated that priming of CD8+ T cells was mediated by langerin+ dDCs, but

surprisingly, LCs actively suppressed immunity.81 As our data revealed greater priming of

DNFB-specific IFN-γ+CD8+ T cells in huLang-cKO mice, LCs are likely to restrict the

cross-priming of cytotoxic T cells when antigens are captured and presented by both LCs

and dDCs. Another murine model that employed epicutaneous sensitization allowed us to

demonstrate a distinct role for migratory LCs in promoting allergic immunity to protein

antigens. In the skin, LCs are major players that uptake external protein antigens and

activate antigen-specific immune responses,2,82 but their function is somewhat redundant,

suggesting the potential contribution of other types of APCs.83 Nevertheless, our huLang-

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cKO mice showed a lower degree of epicutaneous sensitization to OVA, indicating that

CTCF-dependent LC homeostasis is pivotal for cutaneous allergic responses to proteins.

Interestingly, an association between an increased risk of asthma and a disease-related

allele linked to changes in CTCF DNA-binding was reported, opening a possible causality

between allergic diseases and CTCF.84 Moreover, DCs in different barrier tissues possess

an important role in allergy pathogenesis.85 Therefore, future studies are needed to explore

the link between genome-wide binding patterns of CTCF and immunologic functions of

DCs from allergic patients to clarify the functional implications of the CTCF-DC axis in

allergic inflammation.86

Hematopoietic system differentiation is a multistep process tightly regulated and

controlled by complex molecular and cellular networks. Although CTCF has been shown

to regulate cellular development and differentiation of human erythroid cells,87 murine

thymocytes,46 mature T cells,46,48 B cells,50 and myeloid lineage cells including dendritic

cells,52,54,63 whether CTCF controls a homeostasis of adult HSCs has been unknown. As

CTCF-null embryo exhibited prenatal lethality,45 we generated a TMX-induced CTCF

cKO mouse system to conditionally ablate CTCF in the adult hematopoietic system.

Our combined genetic and BM transplantation approaches reveal that acute systemic and

hematopoietic-specific ablation of CTCF leads to a rapid hematopoietic failure. c-Kit is the

surface receptor of SCF, a cytokine essential for HSC self-renewal, growth, and survival.88

Importantly, CTCF ablation results in a severe decrease in heterogeneous c-Kithi

stem/progenitor populations including the most primitive LT-HSCs. A resultant exhaustion

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of HSCs by CTCF depletion is a cell-autonomous effect as co-transplantation with

supporting BM was not able to revert a decreased frequency of c-Kithi cells after CTCF

deletion. To the best of our knowledge, these results are the first description which shows

the pivotal requirement of CTCF in maintaining HSC pool in vivo. Recent study has

revealed the requirement of MED12, which is a component of mediator, for the

transcriptional regulation of c-Kit in HSCs.89 As our CTCF-ablated mice show a similar

reduction of c-Kithi HSCs, it would be intriguing to assess whether CTCF regulates c-Kit

transcript expression in our model in association with mediator and/or chromatin modifiers.

In the absence of specific mitogenic stimuli, HSCs remain in a quiescent or dormant

state.90 Our microarray analyses demonstrated that CTCF-deficient hematopoietic

stem/progenitor cells showed a defective quiescence and a strikingly elevated level of cell

cycle-promoting gene expression, including Mnd1, Dmc1, Ccnd1, Cdk1, and Bub1. CTCF

has been associated with either cell cycle progression or cell cycle arrest in the cell type-

dependent manner. CTCF promotes cell cycle and proliferation of αβ T cells in the

thymus,46 epidermal Langerhans cells,63 and human embryonic stem cells,91 while it

negatively regulates the cell cycle in several types of tumor cells.92-94 As our transcriptome

analysis has been done using the LSK population prior to the loss of surface c-Kit

expression, an increased cell cycle program is in part directly regulated by CTCF-

deficiency at the transcription level. HSC quiescence is critically maintained together with

the low level of intracellular ROS under the hypoxic conditions in vivo.95-97 An increased

ROS level eventually results in HSC proliferation and differentiation.97,98 CTCF-ablated

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HSCs demonstrated a highly elevated ROS level which is related to the considerably

decreased quiescence phase. In line with this, recent study has revealed a role for CTCF in

collaboration with Cockayne syndrome group B protein to protect cells from the oxidative

stress condition.99 Thus, CTCF exhibits an antioxidative activity in certain cell type

including HSCs, although the biological effects would be different according to its target

genes.

HSCs continuously give rise to both myeloid and lymphoid progenitor populations

through the intermediate multipotent progenitor cells.20,21 Interestingly, our mixed chimera

experiments demonstrate that CTCF-deficient HSCs fail to develop into multiple types of

myeloid populations, while lymphoid development is significantly preserved. Previous

study using CTCF-silenced CMPs shows that CTCF knockdown increases that rate of

CMP differentiation in vitro.52 Since CMPs directly differentiate from HSCs and there is a

lack of further input of HSC-derived CMPs from in vitro system, our data suggest that

CTCF is actually required for the myeloid lineage development through sustaining HSC

pool in vivo. It is currently unclear how lymphoid lineage development is maintained by

CTCF-deficient HSCs. It is possible that CTCF directs the balanced diversification

between myeloid and lymphoid lineage development. Therefore, future work will be

definitely needed to clarify an unexpected role for CTCF in the lineage commitment.

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V. CONCLUSION

In Part A, we elucidate a role for CTCF in the DC lineage using genetic approaches.

CTCF supports quantitative maintenance of the systemic DC pool and promots LC self-

turnover. We also demonstrate that CTCF facilitates LC egress from the epidermis and

confers them to differentially regulate cutaneous immunity. In Part B, we elucidate a role

for CTCF in the HSC homeostasis using an inducible gene depletion system. CTCF

supports the homeostatic maintenance of HSC pool through sustaining HSC quiescence

likely in the ROS-dependent manner. This study expands our knowledge of CTCF, which

is actively involved in multiple facets of LC and HSC homeostasis. Further study is

required to fully understand the underlying mechanisms of CTCF in the control of the

development and function of each DC subset and their associations with inflammatory

disorders. Also we anticipate that our knowledge would be translated to the clinical

settings to improve the result of HSC transplantation or anti-leukemic therapies through

modulating CTCF activity in vivo.

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90

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ABSTRACT (in Korean)

피랑게 한스 포 골수 포항상 지에 어

CCCTC-binding factor 역할

< 지도 수 >

연 학 학원 과학과

태균

CCCTC-binding factor (CTCF)는 진핵생물에 간에 보 어 는 DNA 결합

zinc-finger 단 질로 고차원 크로마틴 결 에 핵심 로 여하여 다양한

생물학 통해 한다고 알려 다. CTCF DNA

결합 체에 걸쳐 재하지만 포 특 CTCF 결합 가 재함

알려 는 CTCF 다양한 생물학 능 해하는 도움 다.

본 피 랑게 한스 포 골수 포 항상 지에 어 CTCF

단 질 생물학 능 연 하 다.

랑게 한스 포는 피에 재하는 수지상 포로 피 항원 T 포에

달하여 피 역 한다고 알려 다. 본 수지상 포

랑게 한스 포 특 CTCF 결핍 마우스 하여 CTCF가 랑게 한스 포

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106

가 열 진하여 포수 지에 핵심 로 여함 하 다.

랑게 한스 포 특 CTCF 결핍 마우스 능 통해 CTCF 가

포 착 감 시켜 랑게 한스 포 동 진시킴 알 수

었다. CTCF 결핍 로 한 랑게 한스 포 수 포 동 감 는 증가

과민 감 경피 단 감 과연 하 다.

과 골수 포에 시 는 련 과 로

포가 생한다. 생 동안 과 해 골수 포 가증식

과 엄격하게 어야 한다. 본 타 시 에 한 CTCF 넉아웃

마우스 하고 골수 식 실험 통해 골수 포 항상 지에 어

CTCF가핵심 로 여함 하 다.

연 통해 CTCF 랑게 한스 포 골수 포 항상

지과 립할수 었 , 향후 CTCF 도 통해랑게 한스 포

골수 포 능 어하여 다양한 역피 질 골수 식치료에 할

수 시하 다.

핵심 는말: CCCTC-binding factor, CTCF 특 넉아웃 마우스, 수지상 포,

골수 포, 항상 , 랑게 한스 포

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PUBLICATION LIST

1. Kim TG, Jee H, Fuentes-Duculan J, Wu WH, Byamba D, Kim DS, et al. Dermal

clusters of mature dendritic cells and T cells are associated with the CCL20/CCR6

chemokine system in chronic psoriasis. J Invest Dermatol 2014;134:1462-5.

2. Kim TG, Kim DS, Kim HP, Lee MG. The pathophysiological role of dendritic cell

subsets in psoriasis. BMB Rep 2014;47:60-8.

3. Song MJ, Lee JJ, Nam YH, Kim TG, Chung YW, Kim M, et al. Modulation of

dendritic cell function by Trichomonas vaginalis-derived secretory products. BMB

Rep 2015;48:103-8.

4. Kim TG, Kim M, Lee JJ, Kim SH, Je JH, Lee Y, et al. CCCTC-binding factor

controls the homeostatic maintenance and migration of Langerhans cells. J Allergy

Clin Immunol 2015;136:713-24.

5. Kim TG, Kim S, Jung S, Kim M, Lee MG, Kim HP. CCCTC-binding factor is

essential to the maintenance and quiescence of hematopoietic stem cells in mice.

(Submitted)