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Hematopoietic remodeling during immune activation: connecting hematopoiesis and immunity

de Bruin, A.M.

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Citation for published version (APA):de Bruin, A. M. (2012). Hematopoietic remodeling during immune activation: connecting hematopoiesis andimmunity.

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Alexander M. de Bruin

Hem

atopoietic remodeling during im

mune activation: connecting hem

atopoiesis and imm

unity A

lexander M. de B

ruin


Alexander M. de Bruin

Hematopoietic remodeling during immune activation: connecting hematopoiesis and immunity Thesis, University of Amsterdam, The Netherlands The printing of this thesis was financially supported by: University of Amsterdam Cover: Vincent van Gogh’s The Langlois Bridge at Arles, 1888

Hematopoietic remodeling during immune activation:

connecting hematopoiesis and immunity

ACADEMISCH PROEFSCHRIFT

ter verkrijging van de graad van doctor

aan de Universiteit van Amsterdam

op gezag van de Rector Magnificus

prof.dr. D.C. van den Boom

ten overstaan van een door het college voor promoties

ingestelde commissie,

in het openbaar te verdedigen in de Agnietenkapel

op vrijdag 27 april 2012, te 14:00 uur

door

Alexander Mathieu de Bruin

geboren te Rotterdam

Promotiecommissie:

Promotor: Prof.dr. R.A.W. van Lier Co-promotor: dr. M.A. Nolte Overige leden: Prof.dr. J.G. Borst Prof.dr. P.J. Coffer Prof.dr. G. de Haan Prof.dr. M.H.J. van Oers Prof.dr. S. Repping Prof.dr. I.P. Touw

Faculteit der Geneeskunde

Contents

Chapter 1 Introduction 8

Chapter 2 CD27-triggering enhances HSC self renewal and accelerates

ageing of the HSC compartment 27

Chapter 3 Interferon-gamma impairs the self renewal of hematopoietic

stem cells 48

Chapter 4 Eosinophil differentiation in the bone marrow is inhibited by

T cell-derived IFN 64

Chapter 5 Interferon-gamma induces monopoiesis and inhibits neutrophil

development during inflammation 88

Chapter 6 CD70-driven chronic immune activation is protective against

atherosclerosis 113

Chapter 7 Discussion 128

Appendix 1 Nederlandse samenvatting 146

Appendix 2 List of publications 150

Appendix 3 Dankwoord 152

Chapter 1 Introduction

Introduction

9

The hematopoietic system

Hematopoiesis is the formation of blood cells by the hematopoietic system. The hematopoietic

system generates white blood cells, which are the immune cells that fight invading pathogens upon

infection, red blood cells, which transport oxygen throughout the body and platelets, which are

crucial during hemostasis. Most of these mature cells have a limited life span and to maintain

sufficient numbers of cells to perform these tasks, the hematopoietic system continuously

replenishes these cells. This system is hierarchally organized, with the most potent cells, the

hematopoietic stem cells (HSCs) on top of the pyramid. HSCs reside in the bone marrow and they

have the capacity to give rise to new HSCs, a process called self renewal, and to generate all blood

cell lineages via step-wise differentiation to downstream progenitors and fully mature cells1. A

large fraction of HSCs remains quiescent, non-proliferating, during homeostasis and thus do not

contribute to hematopoiesis2. Due to the ability to self renew and differentiate, a single HSC can

generate the entire hematopoietic system and maintain hematopoiesis for the life of an organism3.

HSCs are located in a specific place in the bone marrow, termed the HSC niche. The HSC niche is

a specialized microenvironment composed of supporting cells that produce soluble and membrane-

bound factors that regulate HSC activity. Multiple niches, located at different anatomical sites in

the bone marrow and providing different signals to HSCs are suggested to be involved in the

regulation of hematopoiesis, although the exact composition and effects of these niches on HSC

biology are poorly described and still under debate4. A combination of cell-intrinsic (transcription

factors) and external (cytokines, membrane-bound molecules) regulatory mechanisms control the

balance between quiescence, self renewal and differentiation, which is critical in preserving HSCs

and generating sufficient numbers of differentiated mature blood cells5.

Although the balance between HSC quiescence, self renewal and differentiation is tightly

regulated, hematopoietic stress conditions demanding an increased output of the hematopoietic

system can alter this balance2. Infection results in depletion of mature immune cells due to

consumption of the cells fighting an invading pathogen and/or by suppression of the hematopoietic

system6. In either case a rapid compensatory response in hematopoiesis is required to quickly

replenish the blood system. How these events are regulated are not yet fully understood, but

evidence is emerging that crosstalk exists between the activated immune system and the

hematopoietic compartment in the bone marrow7-9. In this chapter we will describe the related

aspects of this crosstalk, namely the immune system itself, regulation of HSC self renewal and

differentiation during the steady state situation and upon hematopoietic stress and how immune

activation regulates these processes.

The immune system

The immune system is composed of different specialized cell types that collectively protect the

body from bacterial, fungal, parasitic and viral infections and from growth of tumor cells. Multiple

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Chapter 1

10

cells of the immune system are involved in the immune response to a specific pathogen and two

distinct types of immunity can be recognized: innate and adaptive.

The innate immune system is the first line of host defence against microbial infections and gives

protection to a variety of pathogens based on the recognition of pathogen-associated molecular

patterns constituted of molecules commonly present on the surface of microorganisms. These

relative invariant molecular patterns are detected by germ line encoded pattern recognition

receptors (PRRs), like Toll like receptors (TLRs), which serve as the microbial sensors of the cells

of the innate immune system10. The cellular arm of the innate immune system is composed of

neutrophils, eosinophils, basophils, mast cells, monocytes/macrophages, dendritic cells (DCs), and

natural killer (NK) cells. Defence against pathogens is mediated by engulfing the microorganisms

by a process called phagocytosis and by secreting a multitude of mediators with pro-inflammatory

and/or anti-microbial activity. Besides having a role in innate immunity, macrophages and DCs can

process and present antigens to cells of the adaptive immune system.

The cellular arm of the adaptive immune system is composed of various types of B and T cells and

every cell employs a unique antigen receptor that is not germ line encoded but custom generated in

each B and T cell in every individual11. Whereas the innate immune response has relatively low

specificity and is able to recognize only a limited range of pathogenic molecular patterns via PRRs,

the adaptive immune response is very specific: it can recognize an infinite range of specific

pathogens and mounts a stronger immune response upon a second encounter of a pathogen via the

generation of immunological memory. Upon generation of B cells and T cells in their primary

lymphoid organs, i.e. bone marrow and thymus, respectively, they circulate through the body and

to secondary lymphoid organs like spleen and lymph nodes, which are sites where captured

antigens from blood and lymph are presented by antigen presenting cells (APCs), like DCs, to the

antigen specific B or T cell. Upon recognition of a specific antigen by B or T cells via their B or T

cell receptor (BCR and TCR), respectively, the cell is activated which results in expansion and

differentiation of cells into effector cells and memory cells.

Following activation by crosslinking of their BCR, B cells undergo an intricate differentiation

process to become plasma cells and secrete soluble BCRs, which are called antibodies and that can

opsonize and neutralize pathogens by binding antigens on their surface. For this differentiation

process to occur, B cells undergo a germinal center reaction, in which they proliferate and

randomly mutate their BCR in order to increase the affinity for the antigen12. CD4 T cells play an

important role in this process, as they provide key survival and differentiation signals to high-

affinity B cells, which allow them to undergo immunoglobulin class switching and further

differentiate into either antibody-producing plasma cells or memory B cells. Next to their

supportive role in the germinal center reaction, CD4 T cells also provide essential signals for

activated CD8 T cells to generate a pool of memory cells. Effector CD8 T cells can kill infected or

transformed cells through secretion of cytotoxic molecules, but they are also potent producers of

1

Introduction

11

the pro-inflammatory cytokine interferon-gamma (IFN), which enhances immune functions of a

variety of immune cells. Essential for the activation, survival, expansion and differentiation of both

CD4 and CD8 T cells are activated DCs, as they present antigenic peptides in the context of MHC

molecules and thereby allow activation of T cells specific for the particular antigen. Activated DCs

further drive the activation and differentiation process of antigen-specific T cells by secreting

particular cytokines, such as IL-12, and by expressing co-stimulatory molecules, like CD80, CD86

and CD4013. Another co-stimulatory molecule expressed on activated DCs and other activated

immune cells is CD70, which interacts with its unique receptor CD27. CD27 is a member of the

tumor necrosis factor receptor superfamily and expressed on T cells, NK cells, subsets of B cells

and hematopoietic progenitors. CD27-CD70 interactions on both CD4 and CD8 T cells contribute

to effector cell formation during primary and secondary infections by inducing survival and

proliferation of activated T cells14;15. CD8 effector T cell survival has been shown to be supported

by CD27-dependent induction of autocrine IL-2 production16. In addition, survival is further

supported by upregulation of the antiapoptotic protein Bcl-xL and the serine/threonine kinase Pim-

1, which has both antiapoptotic and prometabolic effects17. Recently, it has been shown that CD27-

driven costimulation supports the survival of low-affinity CD8 effector and memory T cells during

viral infection, expanding the degree of variety to viral antigens, which is beneficial upon

encountering a similar but mutated pathogen18.

HSC Self renewal

Self renewal of HSCs is the process where at least one daughter cell of a dividing HSC retains the

HSC fate. This process is critical in sustaining the pool of HSCs and is a prerequisite for lifelong

hematopoiesis. HSCs can divide symmetrically or asymmetrically. Symmetric cell division of

HSCs results in two identical fated cells, whereas asymmetric division yields two distinct daughter

cells; one cell retaining HSC fate, and one cell contributing to the hematopoietic process via

multilineage differentiation. Although these two distinct types of divisions have been shown in in

vitro studies19;20, their occurrence has not been shown in vivo. The continuous presence of HSCs

and differentiating cells suggests that asymmetric HSC divisions occur in vivo, however, the fate of

two identical HSC daughter cells might also be differentially directed by environmental factors in

the HSC niche.

Estimations about the frequency of HSC divisions range from once every 28-193 days21, indicating

that HSCs are mostly in a quiescent state and need to be awakened in order to self renew. Niche

factors play an important role in keeping HSC quiescent, thus preventing self renewal and

differentiation, which is essential to prevent HSC exhaustion from excessive proliferation22. In

addition, limiting the number of divisions prevents the accumulation of DNA damage and thereby

reduces the risk of malignant transformation of HSCs23. Given the close relationship between

quiescence and self renewal, one might reason that self renewal is merely a consequence of the loss

1

Chapter 1

12

of HSC quiescence, in stead of an actively induced process. In this respect, signaling molecules

maintaining or inducing a quiescent state can be regarded as inhibitors of self renewal. However,

maintaining a quiescent state is a crucial hallmark of HSCs and essential to preserve HSC fate,

suggesting that quiescence of HSCs is a prerequisite for HSC self renewal.

Although the cellular components of the HSC niches are not fully identified yet, the osteoblastic

niche, found near the endosteal lining of the bones, contains osteoblasts, stromal fibroblasts,

osteoclasts perivascular structures and sympathetic neurons, whereas the vascular niche, located

more centrally near blood vessels in the bone marrow, is build up of CXCL12-abundant reticular

cells, vascular endothelial-cadherin+ sinusoidal endothelial cells, reticular cells and

megakaryocytes. In addition, mesenchymal stem cells and macrophages are thought to be part of

the HSC niches4;22. Receptors that bind soluble or membrane-bound ligands expressed in the niche

and that are crucial for the maintenance of HSCs include Tie2, CXCR4, c-Kit and MPL (the

abbreviation of this receptor is based on the disease it is frequently associated with:

myeloproliferative leukaemia). Interaction between these receptors and their ligands, namely

angiopoietin-1 (Ang1), CXCL12, stem cell factor (SCF) and thrombopoietin (TPO), respectively,

are associated with HSC localization in the niche, thus preserving the quiescent state of HSCs24.

Ang1 is produced by osteoblasts, and signaling through Tie2 on HSCs activates 1-integrin and N-

cadherin, promoting the interaction of HSCs with the components of the niche25. CXCL12 is

produced by stromal cells in the niche and signaling through CXCR4 on HSCs is required to keep

HSCs in their niche and thereby quiescent26. In addition, both Ang1 and CXCL12 were shown to

inhibit HSC proliferation in vitro, suggesting that these factors also inhibit HSC proliferation

independently of other signaling components in the HSC niche25;27.

Osteoblasts express membrane-bound SCF (mbSCF) and produce TPO. mbSCF-c-Kit and TPO-

MPL interactions between HSCs and osteoblasts are essential for HSC maintenance, as mice with a

partial loss of c-Kit28 or loss of TPO or MPL29;30 progressively lose HSCs due to a loss of

quiescence. However, treatment of HSCs in vitro with soluble SCF and TPO supports their survival

and self renewal31, indicating that the effect of these factors on HSC biology depends on the

microenvironment of the HSCs. Moreover, whereas injection of TPO in mice initially decreases the

self renewal of HSCs, proliferation of HSCs increases later after injection. In addition, dose

dependent effects of TPO on HSCs quiescence and self renewal in vivo are described29. This

suggests that mbSCF and TPO maintain HSC quiescence in vivo by adhering HSCs to the niche

and supporting their survival, whereas soluble SCF and niche-independent exposure to TPO

supports HSC self renewal.

The signaling pathways mediated via Notch, Wnt and Sonic hedgehog (Shh) have also been

implicated in HSC self renewal. Although Notch ligands Jagged and Delta1, as well as Wnt

proteins, support the self renewal of HSC in vitro32-34, studies with knock out mice suggest that

none of these pathways strongly affect HSC biology in vivo35-38. Hedgehog signaling is activated

1

Introduction

13

via BMP-4 and mediated via downstream mediator Smad5. Shh signaling has been reported to

induce HSC proliferation in vitro39, but Smad5 deficiency in mice does not affect hematopoiesis40.

However, it can not be excluded that integration of combined signaling through these pathways

does regulate HSC function in vivo.

Transforming growth factor (TGF) is a negative regulator of HSC proliferation in vitro. TGF

inhibits cytokine-mediated clustering of lipid rafts, which is essential for amplification of cytokine

signaling, alters cytokine receptor expression and increases the expression of cell cycle inhibitors41.

Interestingly, studies with mice deficient for the receptor for TGF did not reveal critical roles for

TGF on the regulation of HSC self renewal in vivo42.

The HSC niche is characterized by its hypoxic nature, suggesting that oxidative stress is an

important factor in HSC quiescence. Indeed, increase in reactive oxygen species (ROS) increases

the cycling of HSCs via a p38/MAPK pathway, eventually resulting in exhaustion of HSCs43. ROS

increases mammalian target of rapamycin (mTOR) signaling, and genetic deletion of various genes,

including FoxO1, FoxO3, FoxO4, TSC1, PML and Fbw7, result in increased mTOR signaling and

HSC proliferation44-48. Therefore, increase in the metabolic activity of HSCs and concomitant

increase in ROS levels are sufficient to increase the proliferation of HSCs, and the hypoxic state of

the HSC niche contributes to keeping HSCs in a quiescent state.

Gene knockout studies in mice have implicated a number of other genes and signaling pathways

involved in the regulation of HSC quiescence and self renewal, including Pbx1, members of the

Retinoblastoma family, Mi-2, Egr1, Bmi-1, c-Cbl, Evi-1, Menin, Mll5, Myc24. Although these

genes are associated with regulation of expression of relevant transcription factors, ROS

production, integrin expression, gene silencing and activation, the molecular mechanisms behind

the regulation of these genes are largely unknown.

Lineage specific differentiation of HSCs

HSCs are multipotent, capable of generating all the different cell types of the hematopoietic

system. Differentiation of HSCs occurs in a stepwise process via various intermediate progenitors

and is regulated by cytokines and lineage-specific transcription factors. Various models of the

hematopoietic hierarchy exist and are still changing. Especially the branching point between

myeloid and lymphoid fate is under debate49. The most immature HSCs, long term self renewing

HSCs (LT-HSCs, identified as Lineage-c-Kit+Sca-1+(LKS) CD34-Flt3-, or LKS (CD34-

)CD150+CD48-) give rise to short term self renewing HSCs (ST-HSCs, LKS CD34+Flt3-), which

are still self renewing and multipotent cells, but have lost the capacity to self renewal for the life of

an organism. According to the classical model, ST-HSCs give rise to multipotent progenitors

(MPPs, LKS CD34+Flt3+), which have lost self renewal capacity, but are still multipotent3;50-52.

MPPs give rise to common myeloid progenitors (CMPs, Lineage-c-Kit+Sca-1-(LKS-)

1

Chapter 1

14

CD34lowCD16/32low)53 and common lymphoid progenitors (CLPs, Lineage-IL-7Rα+c-KitlowSca-

1low)54. CMPs subsequently differentiate into granulocyte macrophage progenitors (GMPs, LKS-

CD34+CD16/32+) and megakaryocyte erythrocyte progenitors (MEPs, LKS- CD34-CD16/32)54.

Moreover, it has been suggested that ST-HSCs give rise to two populations of MPPs based on

differential expression of Flt3. Whereas MPPs expressing low levels of Flt3 can give rise to CMPs

and CLPs, Flt3hi MPPs, denoted lymphoid-primed MPPs (LMPPs), have lost erythroid potential but

can still generate lymphoid and myeloid progeny via GMPs and CLPs55. Platelets and erythrocytes

are derived from MEPs, while B, T and NK cells and DCs originate from CLPs. GMPs

differentiate into basophilic, eosinophilic and neutrophilic granulocytes and into mast cells and

monocytes (Fig. 1). Additionally, intermediate progenitors for basophils, mast cells and eosinophils

downstream of CMPs and GMPs have been described56-58.

Lineage commitment is dependent on lineage-specific transcription factors and cytokines. CLPs are

supported by the cytokines IL-7 and Flt3-ligand (Flt3L) and transcription factors required for

lymphoid specification include Ikaros, E2A, EBF1, Bcl11a and Pax559. Myeloid differentiation is

mainly regulated by PU.1, C/EBPα, C/EBP, C/EBP, ICSBP and the GATA factors. PU.1 is

essential for the earliest myeloid specification, although PU.1 expression is also found in CLPs.

Whereas high levels of PU.1 block lymphoid differentiation, PU.1 is required to generate CMPs

from HSCs. C/EBPα and PU.1 induces the production of GMPs from CMPs, whereas GATA

factors instruct differentiation to MEPs. Production of platelets and erythrocytes is supported by

TPO and EPO respectively. Monocyte differentiation from GMPs is controlled by high levels of

PU.1 and ICSBP, whereas granulocytic and mast cell differentiation is regulated by the timed

expression of GATA factors, C/EPBα and C/EBP. Eosinophil, basophil and mast cell

development require expression of GATA2, whereas neutrophil differentiation is supported by

GFI1 and C/EBP. Monocyte, neutrophil and eosinophil development are supported by the

cytokines M-CSF, G-CSF and IL-5 respectively, whereas GM-CSF and Il-3 more broadly support

the expansion and maturation of progenitor cells56;60-63.

Lineage-instructing transcription factors induce the expression of cytokine receptors and often

actively suppress genes expressed in or required for other lineages. Forced expression of

transcription factors is therefore often sufficient to direct the differentiation pathway of progenitors.

Since transcription factors control the expression of cytokine receptors, this suggests that cytokines

support the survival and proliferation of committed progenitor cells. Correspondingly, forced

expression of cytokine receptors can instruct the differentiation of progenitors towards a certain

lineage, suggesting that both intrinsic and extrinsic factors are capable of directing lineage

commitment. Moreover, depending on the progenitor, cytokines can be permissive (support

committed progenitors), instructive (induce commitment of progenitors) and/or restrictive

(inhibiting differentiation to a particular lineage)60;64-67. Recently, it was demonstrated that GMPs,

which express receptors for both M-CSF and G-CSF, can be directed towards monocytes or

1

Introduction

15

Figure. 1. Schematic overview of hematopoiesis. Self renewing LT-HSCs give rise to ST-HSCs which produce MPPs with the ability to generate all hematopoietic lineages via intermediate progenitors like CMPs, GMPs and CLPs.

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Chapter 1

16

neutrophils, respectively, depending on the cytokine provided68. This demonstrates that cytokines

alone can direct lineage commitment in vitro. However, lineage commitment in vivo is likely

dependent on the cell-autonomous transcriptional profile, the level of

supportive/instructive/restrictive cytokines and other external factors affecting the transcriptional

network of uncommitted progenitors.

Stress regulation of HSC proliferation

Hematopoietic stress conditions that induce a decrease in the number of circulating blood cells alter

the homeostatic control of hematopoiesis. Loss of blood cells can be caused by chemotherapeutic

agents, bleeding and infections. Infections cause activation of immune cells and many cells fighting

an invading pathogen eventually go into apoptosis. In addition, inflammatory mediators produced

upon infection can directly cause death of immune cells or suppress the hematopoietic system in

the bone marrow. As a result, an increase in the production of blood cells is required to maintain or

restore homeostatic levels of immune cells in times of hematopoietic stress conditions. Although its

is broadly recognized that depletion of hematopoietic cells by the toxic agent 5-fluorouracil (5-FU),

irradiation, bleeding and various types of infection elicit a compensatory response in hematopoietic

activity, the cellular and molecular mechanisms underlying these changes are poorly understood.

Replenishment of the blood system requires an increased differentiation of HSCs to increase the

output of mature cells from the bone marrow. However, increasing numbers of differentiating

HSCs require an increase in self renewing HSCs, and thus a decrease in quiescent HSCs, to

maintain sufficient numbers of HSCs. Indeed, increased proliferation of HSCs is observed after 5-

FU injection, bleeding and infection of mice2;69;70. This suggests a positive feedback loop to induce

HSC cell cycle entry upon blood cell loss. Which factors are involved in this feedback loop is

largely unknown and few associations with the pathways involved in the homeostatic control of

HSC biology are described.

5-FU injection induces cell death of cycling cells, ablating a large portion of bone marrow cells.

Expression of c-Kit on HSC decreases after 5-FU injection and the loss of c-Kit expression and

increased proliferation of HSCs might result from the increase in availability of cytokines in the

bone marrow71. In addition, since c-Kit facilitates adherence of HSCs in the osteoblastic niche, a

decrease in c-Kit expression can free HSCs from their niche, resulting in loss of quiescence and

increased proliferation28.

Although a feedback mechanism between loss of blood cells upon infection and HSC proliferation

has been suggested, very little is known about the effects of infection on HSC biology, which is

mainly caused by an infection-induced phenotypic change in HSCs. Although multiple reports have

suggested that the number of HSCs increase after infection72;73, the number of HSCs is frequently

overestimated in these papers due to an incorrect definition of HSCs in infected mice. The reason

for this inaccuracy is that the stem cell marker Sca-1, used to identify the LKS subset of

1

Introduction

17

stem/progenitor cells, is strongly upregulated on many bone marrow cells upon production of type I

(IFN and IFN) or type II interferons (IFN) during infection and thereby ‘contaminates’ the LKS

compartment. Flow cytometric analysis of LT-HSCs using the LKSCD34-Flt3- definition is

therefore incorrect and should exclude Sca-1 and instead include other discriminating HSC markers

that are not regulated by interferons, such as CD48 and CD150. Loss of hematopoietic progenitor

activity has been reported in vitro and in vivo following infection, and a critical role for IFNα has

been shown in the decrease in hematopoietic activity upon infection6;74. However, because of the

absence of accurate phenotypical characterization of HSC, it can not be concluded from these

studies if the number of HSCs or their functional capacity diminishes after infection. Recently it

was proposed that both IFN and IFNα, described to have negative effects on HSC/progenitor

function in vitro, directly induce proliferation of HSCs in vivo69;74. Nevertheless, a direct effect of

these interferons on HSC quiescence or self renewal was not demonstrated, due to technical

limitations in the performed experiments.

G-CSF can effectively mobilize HSCs and also induces proliferation of quiescent HSCs75. G-CSF

induces release of proteolytic enzymes from neutrophils that degrade and cleave factors like

VCAM1, VLA4, CXCL12 and CXCR476. Several of these proteins are involved in anchoring

HSCs in their niche and loss of these anchoring factors releases HSCs, inducing their proliferation

and enabling their release into circulation. Infections can result in increasing levels of G-CSF,

stimulating the production of neutrophils, in a process called emergency granulopoiesis77. If and to

what extend this excessive infection-induced production of G-CSF affects HSC proliferation has

not been studied so far.

IL-1 and TNFα are typically produced in response to inflammation. IL-1 induces proliferation of

HSC in indirect manner, since IL-1R expression on radioresistant host cells is sufficient to induce

HSC proliferation. Increased proliferation of HSCs likely results from IL-1R-dependent induction

of emergency granulopoiesis or from IL-1R dependent induction of hematopoietic factors

stimulating proliferation of HSCs78. In contrast, TNFα decreases HSC activity both in vitro and in

vivo, as shown by transplantation and colony assays using TNF receptor deficient mice. Although

the mechanism of suppression is not clear, TNFα mainly targeted cycling HSCs in vivo, rather than

quiescent HSCs, and the in vitro suppression could not be attributed to increased cell death79.

HSC express TLRs, which sense microbial antigens, suggesting that pathogens might directly

affect HSC function. Indeed, TLR ligands could directly induce proliferation of HSCs in vitro80.

However, it has not been demonstrated that TLRs on HSCs are directly involved in the regulation

of the hematopoietic stress response during infection.

All together, hematological stress situations trigger a proliferative response of HSCs. However,

inflammatory mediators stimulating and inhibiting proliferation of HSCs have been demonstrated,

although the mechanisms of these effects are largely unclear. Stimulation of HSC proliferation

facilitates the rapid replenishment of blood cells, while inhibition of proliferation might protect

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HSCs from exhaustion, resulting from excessive compensatory proliferation during immunological

challenges.

Stress regulation of HSC differentiation

In contrast to the regulation of HSC quiescence and self renewal, multiple studies have

demonstrated lineage-specific effects on differentiation of progenitors during pathogenic

challenges. Different pathogenic challenges request a response from particular immune cells, and

the hematopoietic system is directed to increase the output of the appropriate immune cells, best

suited to combat the invading pathogen. Infection with extracellular pathogens, like Candida

Albicans, increases the production of neutrophils81, while infection with intracellular pathogens,

like Listeria Monocytogenes, increases the production of monocytes.77 Emergency granulopoiesis

is stimulated by increased levels of G-CSF, while increased levels of M-CSF increase the

production of monocytes upon infection. Responses to parasitic worms and allergic responses are

accompanied by an increase in IL-5, supporting the production of eosinophils82.

Thus, increased production of a specific immune cell is regulated at least by increases in the

lineage-specific cytokine levels. However, other factors in the bone marrow are also involved in the

modulation of hematopoiesis during inflammation. B cell lymphopoiesis is frequently diminished

during infection, which is mediated by factors like TNFα, IL-1 and IFN83-85, whereas

myelopoiesis is enhanced. Although IL-7 is the main lymphopoiesis-driving cytokine,

inflammation does not decrease IL-7 levels in the bone marrow. Surprisingly, levels of SCF in the

bone marrow decreased after inflammation, and it was demonstrated that lymphopoiesis strongly

depends on SCF, while myelopoiesis is not affected by decreasing SCF levels83. In addition,

CXCL12 has been shown to support lymphopoiesis in the bone marrow, but the production of this

chemokine also decreases upon inflammation84. These findings suggest that lymphoid and myeloid

cells compete for common developmental resources. A decrease in shared growth factors may

decrease the support for cells heavily depending on high levels of these factors, resulting in loss of

these particular populations in the bone marrow, thereby enabling the expansion of other lineages

that are directly required for combating the infection.

Next to increased lineage-specific cytokine levels and increased support in developmental niches in

the bone marrow, differentiation of granulocytes during stress conditions has also been shown to be

regulated by other transcription factors than those directing homeostatic granulopoiesis. While

C/EBPα is required and STAT3 is dispensable for homeostatic control of neutrophil development,

C/EBP and STAT3 are both essential to increase neutrophil production during emergency

granulopoiesis86;87.

Recent evidence suggests that pro-inflammatory cytokine IFN is involved in the modulation of

hematopoietic differentiation during immunologic stress situations. It has been previously shown

1

Introduction

19

that IFN has a detrimental effect on B lymphopoiesis85;88. Studies on infections in IFN or IFNR

deficient mice have demonstrated that IFN is also involved in shaping infection-induced

myelopoiesis, since infections of mice lacking IFN signaling results in an uncontrolled expansion

of the neutrophil compartment. In concert with this, it was recently suggested that IFN is involved

in the regulation of monopoiesis and granulopoiesis during intracellular bacterial infection,

although the underlying molecular mechanism is poorly understood89-91.

Besides the effect of soluble factors on hematopoiesis during stress responses, inflammation-

dependent expression of surface molecules has been demonstrated to modulate hematopoiesis.

CD27 and 4-1BB ligand (4-1BBL) are both constitutively expressed on hematopoietic progenitors,

whereas immune activation induces expression of CD70 on APCs, B cells and T cells, 4-1BBL on

APCs and 4-1BB on myeloid progenitors14;15. Interaction of these ligand-receptor pairs has specific

effects on differentiation of progenitors: 4-1BB engagement has been shown to reduce

myelopoiesis8 and CD27 triggering reduces the colony formation of hematopoietic progenitors in

vitro9, suggesting an overall inhibiting effect on differentiation of progenitors. Moreover, CD27

triggering reduces the differentiation of HSCs in vivo9. However, it remains unclear if this is a

direct effect of CD27 triggering on progenitors or a bystander effect of CD27-mediated T cell

activation.

Linking immune activation and hematopoietic stress regulation

Strikingly, although the described hematopoietic and inflammatory cytokines can be produced by

multiple cell types, most of these factors are also produced by T cells. The different subsets of T

cells can produce IL-3, IL-4, IL-5, IL-13, M-CSF, GM-CSF, TNFα, TGF and IFN and can

stimulate the production of IL-6 and G-CSF by stromal cells via secretion of oncostatin M and IL-

177. Although most of these cytokines are involved in the regulation of the immune response, they

also play an important role in stress hematopoiesis. It has been demonstrated that T cell deficient

mice have impaired myelopoiesis, suggesting that T cells are also involved in homeostatic

regulation of hematopoiesis92;93. During infection, activated T cells migrate to the bone marrow and

can thus locally affect hematopoiesis by production of cytokines. In addition, memory T cells

reside in the bone marrow where they can be primed with their cognate antigen94. Besides T cells,

DCs present in the bone marrow can be activated by and locally present blood-born antigens95,

whereas activated B cells/plasmablasts migrate to the bone marrow, where they further differentiate

to and reside as antibody-producing plasma cells12. As such, activated leukocytes in the bone

marrow do not only produce cytokines, but can also express membrane-bound molecules, like

CD70 or 4-1BBL, and can thereby also directly affect hematopoiesis. The presence of activated

immune cells in the bone marrow and the infection-dependent production of cytokines involved in

both directing the immune response and the hematopoietic system suggests a strong association

1

Chapter 1

20

between immune activation and hematopoiesis. As different pathogens elicit specific immune

responses, the effects on the hematopoietic system will likely depend on the type of immune

response. So far, the presence of such a feedback mechanism is poorly studied and the molecular

mechanisms linking the immune and hematopoietic systems are still largely unknown.

Scope of this thesis

Thorough comprehension of the processes that regulate hematopoiesis during inflammation will

allow us to better understand the occurrence of anemia associated with a variety of chronic

inflammatory diseases and develop new intervention methods to prevent or cure these kind of

pathologies. Therefore we decided to study the factors involved in the feedback mechanism linking

hematopoiesis and immune activation (Fig. 2). In this thesis, we have investigated how CD27-

CD70 interactions and the inflammatory cytokine IFN modulate stress hematopoiesis by affecting

Migration of activated lymphocytes to BM

HSC

Myeloid Lymphoid

Self renewal

Differentiation

Osteoblastic niche

Vascular niche

Progenitors

A B

C

Activated lymphocytes

Bone

HSC

Migration of activated lymphocytes to BM

HSC

Myeloid Lymphoid

Self renewal

Differentiation

Osteoblastic niche

Vascular niche

Progenitors

A B

C

Activated lymphocytes

Bone

HSC

Figure 2. Schematic overview of the possible feedback mechanisms of activated cells of the immune system on hematopoietic processes like HSC self renewal (A), HSC differentiation (B) and lineage fate of HSCs/progenitors (C).

1

Introduction

21

HSC self renewal and/or (lineage specific) differentiation. Chapter 2 focuses on the in vivo role of

CD27 and CD70 in the regulation of HSC self renewal and differentiation and the effect of chronic

immune activation on the ageing of the hematopoietic compartment. Using infection models and in

vitro experiments the effect of IFN on HSC self renewal is studied in chapter 3. How IFN affects

eosinophil development is investigated in chapter 4 and chapter 5 focuses on the mechanistic

effects of IFN in directing monocyte and neutrophil development during viral infection. In chapter

6 we show how chronic immune activation affects monocyte formation and the development of

atherosclerosis.

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1

Chapter 2 CD27-triggering enhances HSC self-renewal and

accelerates ageing of the HSC compartment

Alexander M. de Bruin1, Cláudia I. Brandão Silva1,2, Peter A.C. ’t Hoen3, Martijn A. Nolte1,2

1Department of Experimental Immunology, Academic Medical Center, Amsterdam, 2Department

of Hematopoiesis, Sanquin Research and Landsteiner Laboratory, Amsterdam, The Netherlands, 3Center for Human and Clinical Genetics, Leiden University Medical Center, Leiden, The

Netherlands.

Chapter 2

28

Abstract

Ageing of the hematopoietic stem cell (HSC) compartment is characterized by an accumulation of

less productive HSCs with impaired lymphoid differentiation capacity, which contributes to age-

dependent hematological abnormalities including anemia, myeloproliferative disorders and

impaired adaptive immunity. HSC ageing has been associated with inflammation, but the

underlying mechanism has not been fully elucidated. Since HSCs express the costimulatory

receptor CD27, we investigated the impact of CD27-triggering by its inflammatory ligand CD70 on

HSC maintenance. We found that stimulation of CD27 during CD70-driven immune activation in

young mice increased HSC self-renewal, leading to an accumulation of HSC numbers comparable

with aged control mice. Moreover, CD27-triggering negatively affected HSC differentiation to the

lymphoid lineage and increased myeloid differentiation, which is characteristic for aged HSCs.

This functional change was mirrored by a corresponding difference in gene expression, as CD27-

triggered HSCs have a strongly myeloid-biased gene signature. CD27 signaling also increased

expression of genes involved in cellular responses to DNA damage/repair and reactive oxygen

species (ROS), which are associated with HSC ageing and related to increased proliferation. These

findings demonstrate that CD27-stimulation contributes directly to ageing of the hematopoietic

compartment, which identifies this receptor as a novel connection between inflammation and

ageing.

Introduction

Differentiation and self-renewal of HSCs is critical in maintaining homeostatic levels of circulating

blood cells and sufficient HSC numbers for the lifetime of an organism. Murine HSCs can

repopulate the blood system for multiple lifetimes by transplantation to successive recipients1,

which indicates that HSCs are intrinsically protected from exhaustion. However, it is now evident

that age-associated changes do accumulate in the hematopoietic compartment over time. The

number of HSCs increases steadily during ageing, but their repopulation capacity declines,

suggesting that the expansion of HSCs later in life is a compensatory mechanism for a decline in

HSC activity2-6. Furthermore, the HSC pool produces a balanced output of myeloid and lymphoid

cells in young mice, but an increased myeloid lineage commitment at the expense of lymphoid

lineage differentiation potential has been observed in aged HSCs5-7. In accordance, an age-

progressive decline in the number of lymphoid progenitors has been reported5;8. These functional

changes in lineage differentiation are reflected by corresponding changes in gene expression, since

old HSCs express increased levels of genes associated with myeloid differentiation, whereas genes

related to lymphoid specification are typically downregulated5;9. Furthermore, cell intrinsic ageing

of HSCs has been linked to genes involved in DNA damage10, oxidative stress11 and inflammation9,

suggesting that accumulating DNA damage and stress responses contribute to HSC ageing and that

an inflammatory environment may accelerate the ageing process. The functional consequences of

2

CD27-triggering enhances HSC self-renewal and accelerates ageing of the HSC compartment

29

these processes are also clinically important, since ageing of the hematopoietic compartment in

humans is characterized by a decline in adaptive immunity12 and an increase in myeloproliferative

diseases13 and anemia14.

Whereas the majority of the HSC compartment is normally quiescent, this non-proliferative state is

rapidly lost during hematopoietic stress conditions, such as severe blood loss or bone marrow

ablation, due to the increased demand for new hematopoietic cells. Infection-induced cytopenia

also stresses the hematopoietic system and requests robust HSC differentiation and

proliferation15;16. How the function of HSCs and progenitor cells is regulated during inflammatory

stress responses is largely unknown, but both membrane bound molecules17;18 and inflammatory

cytokines19-22 can influence lineage-specific differentiation and HSC self-renewal. In striking

similarity with ageing, infection induces an increase of myelopoiesis over lymphopoiesis in the

bone marrow23. However, whereas acute infections induce a transient increase of inflammatory

mediators, high levels of inflammatory cytokines are known to be maintained during chronic

diseases in the elderly24.

A key player at the interphase between immune activation and hematopoiesis may be the TNF-

receptor superfamily member CD27. CD27 is expressed on T cells and can be triggered by its

costimulatory ligand CD70, which is only expressed during immune activation (reviewed in25).

Enhanced CD27-triggering through overexpression of CD70 increases activation and expansion of

effector T cells19;26;27, ultimately leading to exhaustion of the T cell pool and causing lethal

immunodeficiency in mice28. This is an important observation, since elevated expression of CD70

has been demonstrated in a number of chronic clinical conditions, including rheumatoid arthritis

(RA)29, systemic lupus erythematosus (SLE)30, HIV31 and various malignancies32. Interestingly,

CD27 is not only expressed on T cells, but also on hematopoietic progenitors cells17;33. We have

previously shown that CD27-stimulation impairs the differentiating ability of these progenitors, in

particular to the B cell lineage17. Because of the association between inflammation and HSC ageing

in general and the immunological consequences of CD27-triggering in particular, we investigated

whether CD27-signaling contributes to the ageing process of HSCs. Since CD70 is not expressed

under non-inflammatory conditions, we made use of CD70-transgenic (CD70TG) mice in which

CD70 is constitutively expressed on B cells19, which allowed us to test the consequence of both

chronic CD27-mediated immune activation and de novo CD27 triggering on HSC function in vivo.

Given the large number of IFN- producing effector T cells in CD70TG mice19;20 and the

detrimental effects of IFN- on the hematopoietic compartment19-22;34, we performed these

experiments on an IFN--deficient background (see also17).

We demonstrate here that CD27 is expressed on all HSC subsets and that CD27 triggering results

in increased HSC self-renewal, reduced but myeloid-biased differentiation and increased

expression of genes associated with HSC ageing. These CD27-mediated changes in gene

expression and HSC function strongly resemble the major characteristics of ageing HSCs and

2

Chapter 2

30

provide evidence that CD27 triggering during infection and chronic inflammation accelerates the

natural ageing of HSCs.

Results

CD27-mediated immune activation changes the composition of the hematopoietic

compartment

To examine which hematopoietic progenitors can be directly affected by CD27 signaling, CD27

expression on hematopoietic progenitors from control and CD70TG mice (both on an IFN--

deficient background) was analyzed by flow cytometry. We found that CD27 is expressed on long-

term self-renewing HSCs (LT-HSCs), short-term self-renewing HSCs (ST-HSCs), multipotent

progenitors (MPPs), common lymphoid progenitors (CLPs), granulocyte-monocyte progenitors

(GMPs) and common myeloid progenitors (CMPs), but not on megakaryocyte-erythrocyte

progenitors (MEPs) (Fig. 1A&B). Expression of CD27 was decreased on all progenitor subsets in

CD70TG mice, indicative of CD27-CD70 ligation35. To determine if CD27 triggering resulted in

changes in the hematopoietic progenitor compartment, we calculated absolute numbers of these

progenitor cell subsets. Strikingly, although the total number of Lineage-c-Kit+Sca-1+ (LKS) cells

was not changed (Fig. 1C), CD70TG mice had an aberrant composition of this HSC compartment

(Fig. 1D). The fraction of self-renewing LT-HSCs and ST-HSCs was increased, whereas the

percentage of both MPPs and lymphoid-primed MPPs (LMPPs; fraction of Flt3hi MPPs) was

dramatically reduced (Fig. 1E), which was mirrored by corresponding changes in absolute numbers

of these cells (Fig. 1F). Moreover, the increase in LT-HSCs and decrease in MPPs and LMPPs in

young (12 week old) CD70TG mice reflected the changes observed in 1 year old control mice,

though this was not the case for ST-HSCs (Fig. 1F). The increase in LT-HSC in CD70TG mice was

confirmed by using an alternative identification of LT-HSCs with the SLAM markers CD48 and

CD15036 (Fig. 1G&H). No changes were found in the number of myeloid progenitors (Fig. 1I),

whereas CLP numbers were severely diminished (Fig. 1J). This strong decrease in CLPs

corresponds with the almost complete absence of LMPPs in CD70TG mice, since LMPPs are a

direct upstream progenitor of CLPs37. These analyses show that CD27-signaling has a particular

impact on the hematopoietic progenitor compartment, as it reduces the formation of (L)MPPs and

lymphoid progenitors, while it increases the number of self-renewing HSCs, which correlates to the

composition of the ageing progenitor compartment in aged mice.

LT-HSCs from CD70TG mice show a high proliferative and a myeloid-biased expression

profile

To address the effect of CD27-mediated immune activation and CD27 signaling on the gene

expression signature of self-renewing HSCs, microarray analysis was performed on LT-HSCs from

control and CD70TG mice. In total we found 2123 genes differentially expressed (p<0.01) in LT-

2


31

CD34

Flt

3

Lineage

% o

f M

ax

Sca-1

c-K

it

0.000

0.005

0.010

0.015Control

CD70TG

Nu

mb

er o

f S

LA

M

HS

Cs

(x10

3 )

Control CD70TG

CD150

CD

48

IL-7

Rα

Lineage Sca-1

c-K

it

CD34

CD

16/3

2

CD27-/-

Control

CD70TG

LT-HSC

% o

f M

ax

CD27

ST-HSC MPP CLPGMP CMP MEP

GMP

CMP

MEP

CLP

GMP CMP MEP0.0

0.1

0.2

0.3Control

CD70TG

Nu

mb

er o

f ce

lls (

x106

)

0.000

0.005

0.010

0.015

0.020Control

CD70TG

Nu

mb

er o

f C

LP

s (x

106)

0.00

0.05

0.10

0.15

0.20Control

CD70TG

Nu

mb

er o

f L

KS

cel

ls (

x106

)

LKSca-1+ LKSca-1- Lin-IL-7Rα+A

B

C D E

F G H

I J

CD34

Flt

3

Control CD70TG

Control CD70TG0

20

40

60

80

100LT-HSCST-HSCMPPLMPP

% o

f L

KS

**

*

*

***

***

***

Lin-Total BM Total BM

11.7 46.5

17.5

39.4

74.2

7.59

0.88

17.5

8.2 15.3

Lin-

MPP

ST-HSCLT-

*

*

* ***

***

*

**

LT-HSC ST-HSC MPP LMPP0.00

0.02

0.04

0.06

0.08

0.10

0.12Control

CD70TG

Aged control

Nu

mb

er o

f ce

lls

(x

10

6 )

CD34

Flt

3

Lineage

% o

f M

ax

Sca-1

c-K

it

0.000

0.005

0.010

0.015Control

CD70TG

Nu

mb

er o

f S

LA

M

HS

Cs

(x10

3 )

Control CD70TG

CD150

CD

48

Control CD70TG

CD150

CD

48

IL-7

Rα

Lineage Sca-1

c-K

it

CD34

CD

16/3

2

CD27-/-

Control

CD70TG

LT-HSC

% o

f M

ax

CD27


CD27-/-

Control

CD70TG

CD27-/-

Control

CD70TG

LT-HSC

% o

f M

ax

CD27


GMP

CMP

MEP

CLP

GMP CMP MEP0.0

0.1

0.2

0.3Control

CD70TG

Nu

mb

er o

f ce

lls (

x106

)

0.000

0.005

0.010

0.015

0.020Control

CD70TG

Nu

mb

er o

f C

LP

s (x

106)

0.00

0.05

0.10

0.15

0.20Control

CD70TG

Nu

mb

er o

f L

KS

cel

ls (

x106

)

LKSca-1+ LKSca-1- Lin-IL-7Rα+A

B

C D E

F G H

I J

CD34

Flt

3

Control CD70TG

Control CD70TG0

20

40

60

80


% o

f L

KS

Control CD70TG0

20

40

60

80


% o

f L

KS

**

*

*

***

***

***

Lin-Total BM Total BM

11.7 46.5

17.5

39.4

11.7 46.5

17.5

39.4

74.2

7.59

0.88

17.5 74.2

7.59

0.88

17.5

8.2 15.3

Lin-

MPP

ST-HSCLT-

*

*

* ***

***

*

**


0.02

0.04

0.06

0.08

0.10

0.12Control

CD70TG

Aged control

Nu

mb

er o

f ce

lls

(x

10

6 )

*

*

* ***

***

*

**


0.02

0.04

0.06

0.08

0.10

0.12Control

CD70TG

Aged control

Nu

mb

er o

f ce

lls

(x

10

6 )

Figure 1. CD27-mediated immune activation changes the composition of the hematopoietic stem and progenitor cell compartment. (A) Gating strategy for identification of the various hematopoietic progenitors. (B) Representative histograms showing expression of CD27 on hematopoietic progenitors of CD27-/- (shaded histogram), control (bold line) and CD70TG mice (thin line). (C) Number of LKS cells in the bone marrow, (D) representative plot and (E) bar graph showing composition of the LKS compartment in control and CD70TG mice. (F) Absolute numbers of LT-HSCs, ST-HSCs and MPPs in young control, young CD70TG and 1-year old control mice. (G) Representative plot of staining for SLAM HSCs and (H) absolute numbers of these cells. Absolute numbers of (I) myeloid and (J) lymphoid progenitors in control and CD70TG mice. Data represent mean ± s.d. from 3-5 mice per group. Experiments were performed at least twice with similar results. *, p < 0.05, **, p < 0.01, ***, p < 0.001.

2

Chapter 2

32

HSCs from CD70TG compared to control mice. Identification of biological processes in the

signature of upregulated genes demonstrated enriched categories linked to cell cycle progression,

DNA replication and DNA damage/repair responses, myeloid differentiation and responses to

inflammation, stress and ROS (Table 1). Categories representing biological processes of

downregulated genes included cell death and cell cycle arrest, regulation of adaptive immune

responses and lymphocyte proliferation (Table 1). Detailed analysis showed increased expression

of a large number of genes positively regulating cell cycle progression (such as c-Myb, PCNA,

polo-like kinases, E2F1 and various cyclins and cell division cycle (associated) genes) and a

corresponding decrease of several cell cycle inhibitors (like Cdkn1c/p57 and E2F5) (Table 2).

Furthermore, in accordance with the decreased differentiation of HSCs to LMPPs and CLPs,

several lymphoid-related genes were downregulated (such as Bcl11b/CTIP2, IL-7,

Tnfrsf13c/BAFF-R and Pbx1), whereas an abundance of genes associated with myeloid

differentiation (like Cathepsin g, myeloperoxidase, C/EBP, CD163, Csf1r/CD115, CD68, CD14

and several FcR-genes) was upregulated in CD70TG LT-HSC (Table 3). These data suggest that

LT-HSCs in CD70TG mice have an increased proliferative rate, which would correlate with the

observed increase in cell numbers. Furthermore, these HSCs display a myeloid-biased

differentiation profile and they have induced expression of genes associated with inflammatory and

metabolic stress responses, which resembles the expression profile of aged HSCs.

Upregulated Downregulated

Biological process Fold change Biological process Fold change

Cell cycle progression 16.6 Regulation of T-helper 2 type immune

Response -9.0

Negative regulation of DNA replication initiation

12.4 Positive regulation of adaptive immune

Response -4.0

DNA damage/DNA repair responses 7.6 Positive regulation of caspase activity -3.8

Neutrophil mediated immunity 7.1 Cell cycle arrest -3.3

Myeloid leukocyte mediated immunity 4.9 Negative regulation of cell proliferation -2.5

Cellular response to reactive oxygen species

4.6

Regulation of lymphocyte proliferation -2.4

Phagocytosis, engulfment 4.4 Positive regulation of cell death -2.0

Oxygen and reactive oxygen species metabolic process

3.2

Regulation of lymphocyte activation -1.9

Protein folding 3.1

Response to oxidative stress 2.5

Cellular response to stress 2.4

Inflammatory response 1.5

Oxidation reduction 1.3

Table 1 Enrichment of selected biological process associated with up- or downregulated genes in LT-HSCs from CD70TG compared to control mice.

2


33

Positive regulators Negative regulators

Gene Fold

change

P value Gene Fold

change

P value Gene Fold

change

P value

Myb 10.03 5.5E-03 Cdc7 2.34 1.8E-05 Cdkn1c -4.71 2,5E-07

Pcna 8.83 8.2E-04 Cdc25c 2.20 1.0E-06 Bcl11b -4.58 2,3E-04

Wee1 8.04 4.5E-03 Cdc25b 2.14 1.6E-05 Ndn -4.35 5,9E-07

Melk 6.59 3.0E-07 Cops5 2.14 8.2E-03 E2F5 -1.57 5,0E-03

Plk4 6.56 9.2E-07 Dnmt1 2.14 2.7E-04 Cdkn2c 1.75 2.0E-04

Plk1 5.23 2.1E-06 Brca1 2.14 2.5E-05

Cdc20 4.95 9.1E-04 Fzr1 2.00 1.0E-03

Ttk 4.87 6.9E-07 Caprin1 1.99 3.0E-04

Ywhae 4.82 2.3E-03 Orc4l 1.99 7.1E-03

Ccnb1 4.68 2.2E-06 Cdca2 1.98 4.2E-03

Gtse1 4.37 1.1E-06 Tcp1 1.97 3.7E-04

Aurka 4.08 8.8E-07 Cdc45l 1.95 1.5E-03

Gmnn 3.98 6.4E-04 Mcm6 1.95 3.6E-04

Skp2 3.89 1.8E-05 Mcm5 1.94 5.2E-04

Cdc2a 3.87 2.1E-04 Ywhaq 1.84 1.5E-03

Cdca5 3.39 3.2E-08 Cdca4 1.79 1.4E-04

Mad2l1 3.35 2.4E-06 Bub3 1.79 9.1E-04

Bub1b 3.26 2.9E-07 Pkmyt1 1.78 1.8E-04

Chek1 3.21 8.1E-03 Orc1l 1.73 1.9E-03

Dbf4 3.20 8.7E-07 E2f1 1.68 2.4E-03

Espl1 3.02 8.7E-06 Mybl2 1.61 4.0E-03

Clspn 2.96 1.8E-06 Ccnd3 1.56 2.4E-03

Cdca3 2.92 1.0E-03 Orc6l 1.54 9.0E-03

Cdc6 2.60 6.6E-06 G0S2 1.47 4.4E-03

Foxm1 2.55 3.8E-06 Gspt1 1.47 2.3E-03

Rbl1 2.51 1.9E-04 Ywhab 1.41 5.5E-03

Ccna2 2.40 1.1E-03 Bin1 1.35 5.5E-03

Cdca8 2.40 3.4E-04

Table 2 Selected cell cycle genes differentially expressed in LT-HSCs from CD70TG compared to control mice.

2

Chapter 2

34

Increased proliferation of LT-HSCs in CD70TG mice

To determine whether the observed changes in gene expression correlated with corresponding

functional changes in the HSC compartment, we first tested whether enhanced CD27-stimulation

influenced HSC proliferation. Cell-cycle analysis of purified LKS cells revealed that an increased

fraction of LKS cells from CD70TG mice was in S/G2/M phase compared to control mice (Fig.

2A-B). Furthermore, since the majority of LT-HSCs is normally in the quiescent G0 phase of the

cell cycle, it lacks expression of the proliferation marker Ki6716. We found that an increased

percentage of CD70TG LT-HSCs expressed Ki67 compared to control mice and had thus lost its

quiescent state (Fig. 2C-D). ST-HSCs and MPPs are typically not quiescent and already 80% of

these cells express Ki67 in control mice, though we found that this percentage was even

significantly increased in CD70TG mice (Fig. 2D). Furthermore, to investigate HSC proliferation

in vivo, mice were injected with 5-bromo-2'-deoxyuridine (BrdU) and incorporation of BrdU was

measured 24 hours later. We observed a significant increase in BrdU+ LT-HSCs (and MPPs) in

CD70TG mice, which demonstrates that CD70TG mice have more proliferating LT-HSCs than

control mice (Fig. 2E-F). Finally, to confirm the increased proliferation of HSCs in vivo, control

and CD70TG mice were injected with 5-fluorouracil (5-FU), as this treatment ablates proliferating

cells. As expected, a strong reduction in HSC numbers was found in CD70TG mice after

administration of 5-FU (Fig. 2G). All together, these experiments suggest that increased CD27-

triggering through its ligand CD70 enhances self-renewal of LT-HSCs.

Myeloid genes Lymphoid genes

Gene Fold

change

P value Gene Fold

change

P value Gene Fold

change

P value

Ctsg 46.85 1.00E-07 S100a10 2.69 2.13E-03 Bcl11b -4.58 2.29E-04

Lyzs 30.77 8.56E-06 Cebpe 2.50 1.90E-03 Ebi3 -2.68 6.36E-04

Mcpt8 13.85 3.21E-08 Fcgr2b 2.47 1.54E-06 Igbp1 -1.98 2.73E-03

Ela2 11.44 1.17E-05 S100a8 2.47 3.36E-03 Tnfrsf13c -1.95 8.02E-04

Lpl 10.78 2.76E-06 Camp 2.46 9.62E-03 Il7 -1.70 2.42E-04

Emr1 7.91 2.03E-07 Tyrobp 2.26 3.12E-04 Zpbp -1.51 4.75E-03

Mgl1 6.79 9.27E-07 Prtn3 2.20 2.14E-03 Elmo1 -1.38 9.84E-03

Mpo 6.39 8.06E-07 Csf1r 2.19 4.31E-04 Pbx1 -1.34 8.14E-03

Clec7a 6.30 1.03E-06 Fcgr3 2.12 3.37E-05

Fcna 5.98 3.04E-09 Cd68 2.07 5.87E-04

Cd163 5.67 6.58E-06 Emb 1.95 8.77E-05

Tgfbr1 4.18 2.04E-03 Ncf1 1.93 1.86E-03

Cd53 3.39 1.40E-04 Cd14 1.77 1.72E-03

Chi3l3 3.25 2.20E-03 Ptpn12 1.75 7.02E-03

Fcgr4 3.07 3.85E-03

Table 3 Selected myeloid and lymphoid marker genes differentially expressed in LT-HSCs from CD70TG compared to control mice.

2


35

LT-HSC ST-HSC MPP0

10

20

30

40

% o

f B

rdU

+ c

ells

G0/G1 S/G2/M0

10

20

3060

80

100

% o

f L

KS

cel

lsLT-HSC ST-HSC MPP

0

20

40

60

80

100 Control

CD70TG

% o

f K

i-67

+ c

ells

B

D

F

Control

24.8

75.5

CD70TG

61.5

38.2

PI (DNA)

% o

f M

ax

Control

4.62

95.4

CD70TG

12.3

87.7

Control

11.9

CD70TG

26.2

A

C

E

G

FSC

Ki6

7

FSC

Brd

U**

**

**

***

*

*

***

0

20

40

60Control

CD70TG

Nu

mb

er o

f H

SC

s (x

103 )

ControlCD70TG

ControlCD70TG

***

LT-HSC ST-HSC MPP0

10

20

30

40

% o

f B

rdU

+ c

ells

G0/G1 S/G2/M0

10

20

3060

80

100

% o

f L

KS

cel

lsLT-HSC ST-HSC MPP

0

20

40

60

80

100 Control

CD70TG

% o

f K

i-67

+ c

ells

B

D

F

Control

24.8

75.5

Control

24.8

75.5

CD70TG

61.5

38.2

CD70TG

61.5

38.2

61.5

38.2

PI (DNA)

% o

f M

ax

Control

4.62

95.4

Control

4.62

95.4

4.62

95.4

CD70TG

12.3

87.7

CD70TG

12.3

87.7

12.3

87.7

Control

11.9

Control

11.9

CD70TG

26.2

CD70TG

26.226.2

A

C

E

G

FSC

Ki6

7

FSC

Brd

U**

**

**

***

*

*

***

0

20

40

60Control

CD70TG

Nu

mb

er o

f H

SC

s (x

103 )

***

0

20

40

60Control

CD70TG

Nu

mb

er o

f H

SC

s (x

103 )

ControlCD70TG

ControlCD70TG

***

Figure 2. Increased proliferation of LT-HSCs in CD70TG mice. (A) Representative histogram of DNA staining in LKS cells and (B) quantification of cell cycle phases of these cells from control and CD70TG mice. (C) Representative plot showing Ki67 expression in LT-HSCs and (D) quantification of Ki67 expression in LT-HSCs, ST-HSCs and MPPs from control and CD70TG mice. (E) Representative plot showing BrdU incorporation in LT-HSCs and (F) quantification of BrdU incorporation in LT-HSCs, ST-HSCs and MPPs from control and CD70TG mice 24 hours after BrdU injection. (G) Number of LKS cells in control and CD70TG mice 5 days after 5-FU injection. Data in B represent mean ± s.d. from 3 independent experiments. Data in D, F and G represent mean ± s.d. from 4-5 mice per group. *, p < 0.05, **, p < 0.01, ***, p < 0.001.

2

Chapter 2

36

Changes in the HSC compartment are a direct consequence of CD27 signaling in these cells

To test whether the observed effects on LT-HSCs in CD70TG mice are indeed a direct result from

CD27 signaling on LT-HSCs and not due to CD27-mediated T cell activation19, we co-transferred

LT-HSCs from CD27+/+ (CD45.2) mice and CD27-/- (CD45.1) into lethally irradiated CD27-/-

(control chimeras) or CD70TG x CD27-/- (CD70TG chimeras) recipient mice (both CD45.1.2; Fig.

3A). These two recipient mouse strains are identical regarding HSC phenotype and hematopoietic

status, since the overexpression of CD70 is without any effect in the absence of its receptor

CD2717;19;20. Therefore, the competitive transplantation of HSCs in either of these two mouse

strains allowed us to analyze the direct consequence of CD27 triggering on HSC maintenance and

function. HSC donor chimerism was assessed 16 weeks after transplantation and analysis of

HSC/progenitor subsets within the LKS fraction revealed that CD27+/+ and CD27-/- LKS cells had a

similar composition in control chimeras (Fig. 3B). In contrast, CD27 triggering of CD27+/+ HSCs in

CD70TG chimeras strongly increased the percentage of LT-HSCs and ST-HSCs and reduced the

percentage of MPPs within the pool of LKS cells (Fig. 3C). These data corroborate our findings in

full CD70TG mice (Fig. 1). This change in the HSC-compartment was a direct consequence of

CD27 triggering on HSCs, since the LKS composition of CD27-/- derived cells was comparable

between control and CD70TG chimeras (Fig. 3B-C).

Furthermore, to evaluate the competitive effect of CD27-triggering on HSC reconstitution and

maintenance, we calculated the ratio of CD27+/+: CD27-/- HSCs in control and CD70TG chimeras.

If CD27+/+ and CD27-/- HSCs would equally contribute to engraftment, a donor HSC chimerism

ratio of 1 is expected. However, CD27+/+: CD27-/- ratios for all HSC/progenitor subsets in the

control chimeras were all higher than 1 (Fig. 3D), demonstrating that CD27-expression on HSCs

contributes to a higher competitive reconstitution capacity. We found that the CD27+/+: CD27-/-

ratio of LT-HSCs in CD70TG recipients were comparable to the ratio in control recipients, though

the CD27+/+: CD27-/- ratios of ST-HSCs and MPPs were strongly decreased in CD70TG chimeras,

suggesting that CD27 signaling impairs the differentiation of LT-HSCs to the more mature ST-

HSCs and MPPs.

To test if the observed increased proliferation of LT-HSCs in full CD70TG mice was also a direct

consequence of CD27 triggering on HSCs, we measured Ki67 expression in LT-HSCs of these

mice. We found that more CD27+/+ LT-HSCs expressed Ki67 and had thus lost their quiescent state

compared to CD27-/- HSCs in both control and CD70TG chimeras (Fig. 3E), which provides a

likely explanation for the enhanced reconstitution capacity of CD27+/+ HSCs in control chimeras.

Whereas the fraction of proliferating CD27-/- HSCs was not different between both groups of

chimeras, an increase of Ki67+-expressing CD27+/+ HSC was observed in CD70TG chimeras

compared to control chimeras (Fig. 3E), which further supports the hypothesis that enhanced CD27

signaling increases proliferation and self-renewal of HSCs. These data demonstrate that the

observed changes in the hematopoietic compartment of CD70TG mice are a direct result from

2


37

CD27 triggering on HSCs and demonstrate that CD27 signaling increases the self-renewal of LT-

HSCs.

CD27 triggering results in an overall reduced and myeloid-skewed differentiation

To examine the extent by which CD27 triggering modulates HSC differentiation and whether this

process is indeed functionally biased towards the myeloid lineage, we analyzed the myeloid and

250 CD27-/- HSCs (CD45.1)

CD27-/- (CD45.1.2)

250 CD27+/+ HSCs (CD45.2)

1 x 106 CD27-/- total bone marrow cells (CD45.1.2)

250 CD27-/- HSCs (CD45.1)

CD70TG x CD27-/- (CD45.1.2)

250 CD27+/+ HSCs (CD45.2)

1 x 106 CD70TG x CD27-/- total bone marrow cells (CD45.1.2)

Control chimeras CD70TG chimeras

A

E

*

***

CD27-/- CD27+/+0

20

40

60

80

100LT-HSCST-HSCMPP

% o

f L

KS

**

***

B C D

CD27-/- CD27+/+0

20

40

60

80

100LT-HSCST-HSCMPP

% o

f L

KS

Control CD70TG

***

LT-HSC ST-HSC MPP

*****

0

1

2

3

4

5

6

7

8Control

CD70TG

Rat

io C

D2

7+/+

:CD

27-/

-

Control CD70TG0

20

40

60

80

100CD27-/-

CD27+/+

% o

f K

i-6

7+L

T-H

SC

250 CD27-/- HSCs (CD45.1)

CD27-/- (CD45.1.2)

250 CD27+/+ HSCs (CD45.2)

1 x 106 CD27-/- total bone marrow cells (CD45.1.2)

250 CD27-/- HSCs (CD45.1)

CD70TG x CD27-/- (CD45.1.2)

250 CD27+/+ HSCs (CD45.2)

1 x 106 CD70TG x CD27-/- total bone marrow cells (CD45.1.2)

Control chimeras CD70TG chimeras

A

E

*

***

CD27-/- CD27+/+0

20

40

60

80

100LT-HSCST-HSCMPP

% o

f L

KS

**

***

B C D

CD27-/- CD27+/+0

20

40

60

80

100LT-HSCST-HSCMPP

% o

f L

KS

Control CD70TG

***

LT-HSC ST-HSC MPP

*****

0

1

2

3

4

5

6

7

8Control

CD70TG

Rat

io C

D2

7+/+

:CD

27-/

-

LT-HSC ST-HSC MPP

*****

0

1

2

3

4

5

6

7

8Control

CD70TG

Rat

io C

D2

7+/+

:CD

27-/

-

Control CD70TG0

20

40

60

80

100CD27-/-

CD27+/+

% o

f K

i-6

7+L

T-H

SC

Control CD70TGControl CD70TG0

20

40

60

80

100CD27-/-

CD27+/+

% o

f K

i-6

7+L

T-H

SC

Figure 3. Changes in the HSC/progenitor compartment are a direct result from CD27 signaling in these cells. (A) Schematic overview of the generation of control and CD70TG chimeric mice. Composition of the CD27-/- and CD27+/+ LKS compartment in (B) control and (C) CD70TG chimeras 16 weeks after transplantation. (D) CD27+/+:CD27-/- ratios of HSCs and progenitors in control and CD70TG chimeric mice. (E) Ki67 expression in CD27-/- and CD27+/+ LT-HSCs in control and CD70TG chimeras. Data represent mean ± s.e.m. from 8 mice per group. A comparable experiment was performed with similar results. *, p < 0.05, **, p < 0.01, ***, p < 0.001.

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lymphoid progenitor compartments 16 weeks after transplantation and peripheral blood of control

and CD70TG chimeras at different time points after HSC transplantation. This revealed that the

CD27+/+: CD27-/- ratio of myeloid progenitors was almost 5-fold lower in CD70TG chimeras,

whereas the ratio for CLPs decreased almost 13-fold (Fig. 4A). This demonstrates that CD27

signaling impairs LT-HSC differentiation in general, but most profoundly to the lymphoid lineage

and thus inducing a myeloid-biased differentiation of HSCs.

Analysis of peripheral blood demonstrated a two- to three-fold increase of WBCs derived from

CD27+/+ compared to CD27-/- donor cells in control chimeras (Fig 4B-E), which followed the ratios

found in the HSC compartment (Fig 3D). The myeloid/lymphoid composition of WBCs was

comparable between the two donor genotypes, although the percentage CD27+/+ B cells was

somewhat increased at the expense of T cells (Fig. 4F). These findings were in striking contrast

with CD70 chimeric mice, in which differentiation of CD27+/+ HSCs to WBCs was four- to five-

fold impaired compared to its CD27-/- counterparts (Fig. 4B-E) and strongly skewed towards

myeloid cells (Fig. 4G). Although myeloid output of CD27+/+ HSCs in CD70TG chimeras was also

hampered early after transplantation, a steady increase in CD27+/+: CD27-/- ratios was observed

over time reaching a ratio of one at 16 weeks (Fig. 4E), which corresponds with the ratio of

myeloid progenitors in these mice (Fig. 4A). Differential analysis of CD27-/- donor HSC-derived

peripheral blood cells demonstrated similar distribution of B, T and myeloid cell lineages in control

and CD70TG chimeras, thus excluding indirect effects of CD27-mediated immune activation on

HSC differentiation (Fig. 4F-G). These experiments demonstrate that also in this adoptive transfer

model CD27 triggering reduces the differentiation capacity of HSCs and induces a myeloid-skewed

differentiation profile, which indicates that CD27-triggering enhances HSC ageing.

Discussion

The causal relationship between inflammation and HSC ageing is not yet clearly understood. The

functional impairment of aged HSCs to generate new immune cells might well increase the

susceptibility for infections and thereby cause more inflammation12. On the other hand it could also

be that the increase in inflammatory mediators during ageing (due to e.g. increased amount of fat

tissue, smoking, subclinical infections, cardiovascular diseases etc.)24 negatively influences the

HSC compartment and causes the induction of inflammation-related genes in aged HSCs9. Here we

describe a new parallel between inflammation and HSC ageing, as we demonstrate that prolonged

triggering of CD27 on HSCs, which can occur during chronic inflammatory conditions, rapidly

ages the HSC compartment. We demonstrate that CD27 is expressed on all hematopoietic

progenitors, except for MEPs, and that CD27 triggering by CD70 results in changes in the

composition of the hematopoietic compartment similar to those observed in aged mice, i.e. an

accumulation of HSCs that have a decreased differentiation capacity, in particular towards the

lymphoid lineage. Although CD27-triggering on progenitor cells other than HSCs could also have

2


39

Myeloid cells

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CD27-/-

CD27+/+

CD27

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CD27+/+

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3 7 10 13 160.01

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-

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Figure 4. CD27 triggering results in an overall reduced and myeloid-skewed differentiation of HSCs. (A) CD27+/+:CD27-/- ratios of myeloid progenitors (MPs; Lin-c-Kit+Sca-1-) and CLPs in control and CD70TG chimeras. CD27+/+:CD27-/- ratios of (B) total WBCs, (C) B cells, (D) T cells and (E) myeloid cells in peripheral blood of control and CD70TG chimeras at the indicated time points. Composition of CD27-/- and CD27+/+ donor-derived peripheral blood cells in (F) control and (G) CD70TG chimeras at the indicated time points. Data represent mean ± s.e.m. from 8 mice per group. A comparable experiment was performed with similar results. In A-E: **, p < 0.01, ***, p < 0.001. In F-G: *, p < 0.05, **, p < 0.01, #, p < 0.001

2

Chapter 2

40

influenced lineage differentiation, the strongly myeloid-biased gene-signature in LT-HSCs from

CD70TG mice indicates that CD27-triggering on HSCs already primes these cells for myeloid

rather than lymphoid differentiation. Next to influencing differentiation, we found that CD27

stimulation also affects HSC maintenance, since LT-HSCs in CD70TG mice are less quiescent and

have an increased turn-over. This functional change correlates with the gene expression profile in

these cells, since many genes that positively regulate mitosis are upregulated, whereas expression

of several cell cycle inhibitors is decreased. Of particular interest are the strong induction of the

proto-oncogene c-Myb and the downregulation of the cyclin-dependent kinase inhibitor p57

(Cdkn1c; Table 2), since these genes have recently been shown to be crucial for HSC self-

renewal38 and quiescence39, respectively. We also found strong downregulation of Necdin, a gene

that restricts excessive HSC proliferation during hematopoietic regeneration40. Importantly, the

functional changes observed in LT-HSCs from CD70TG mice resulted from direct CD27 triggering

on HSCs rather than from a general CD27-mediated inflammatory environment, as CD27+/+ HSCs

did show the ageing phenotype when transferred to CD70 chimeric mice, but CD27-/- HSCs in the

same mouse did not.

While HSCs are multipotent, capable of generating both lymphoid and myeloid progeny, genes

involved in myeloid and lymphoid commitment are already expressed in LT-HSCs5;7 and the

presence of myeloid- and lymphoid-biased HSCs has been acknowledged41;42. With age, myeloid-

biased HSCs accumulate and considering the heterogeneous population at young age it has been

suggested that this accumulation results from an age-related clonal expansion of myeloid-biased

HSCs, rather than a common progressive change in the entire young pool of HSCs7;43. The

decreased lymphoid potential of HSCs in aged mice is cell intrinsic and irreversible, since it can

not be restored by transplanting these cells in young recipients5. We have previously shown that

CD70TG HSCs are impaired in their capacity to reconstitute B cells upon transplantation in WT

mice17, which correlates with the altered gene expression profile found in these HSCs (Tables 1-3).

Importantly, our current study using CD70 chimeric mice demonstrates that de novo CD27-

triggering also rapidly inhibits the lymphoid potential of HSCs, suggesting that the impaired

lymphoid differentiation by HSCs in CD70TG mice is a direct consequence of CD27-triggering on

these cells, rather than a reduction in the number of lymphoid-biased HSCs.

Interestingly, the changes in HSC function during inflammation and ageing have striking

similarities. Previous studies have demonstrated that inflammation transiently modulates the steady

state hematopoiesis in the bone marrow and alters the production of myeloid cells and

lymphocytes. Inflammatory cytokines produced during infection decreases the supportive

conditions for B lymphopoiesis and promotes the production of myeloid cells, in order to increase

the number of cells required to fight the invading pathogen, while normal B lymphopoiesis is

restored after infection23. It was recently shown that mammalian target of rapamycin (mTOR)

activity is elevated in aged HSCs and that mTOR activation increases ROS levels in HSCs from

2


41

young mice, inducing an HSC phenotype comparable to aged HSCs44. mTOR can be activated by

the inflammatory cytokines IL-6 and TNFα45 and elevated levels of these cytokines are found in the

elderly, suggesting that these cytokines contribute to the ageing of HSCs24. Our experiments show

that next to these inflammatory cytokines, the interaction between CD70 and CD27 can also

contribute to ageing of the hematopoietic compartment.

Infection induces hematopoietic stress15 and the resulting increased demand for peripheral immune

cells requires an increased differentiation of HSCs and progenitors. However, uncontrolled

differentiation of HSCs will deplete the HSC pool, and therefore the rate of HSC differentiation

and compensatory proliferation requires strong regulation. Our data demonstrate that CD27-

signaling both diminishes HSC differentiation and induces proliferation to maintain sufficient

numbers of HSCs during inflammation. Although overall differentiation of CD27+/+ HSCs in CD70

chimeric mice is decreased, myeloid differentiation steadily increases over time. Therefore, while

short term CD27 triggering might transiently dampen HSC differentiation and increase

proliferation, the observed myeloid skewing in CD70TG chimeras over time suggests that

continuous CD27 triggering and the resulting increased proliferation accelerates the natural process

of HSC ageing, resulting in an accumulation of myeloid-biased HSCs. Stress responses and DNA

damage have been associated with HSC ageing and genes overexpressed in CD70TG LT-HSCs

were associated with proliferation and DNA replication, responses to ROS and DNA damage/repair

responses. Both DNA replication and ROS contribute to DNA damage and likely play an important

role in the accelerated HSC ageing observed in CD70TG and CD70 chimeric mice, since CD27-

triggered HSCs have a high proliferative rate and metabolic activity. Other studies have also

demonstrated that inflammatory conditions increase the proliferation of HSCs46;47. Increased HSC

proliferation upon hematopoietic stress is required to maintain sufficient HSCs and an increased

output of the bone marrow, however, chronic immune activation might ultimately accelerate HSC

ageing resulting from accumulating DNA damage, due to a heightened proliferative and metabolic

state.

Acknowledgements

We thank Sten Libregts for technical assistance, Berend Hooibrink for cell sorting and the staff of

the animal facility of the AMC for animal care. We thank Prof. Dr. René van Lier for stimulating

discussions and Drs. Paula B. van Hennik and Gerald de Haan for critical reading of the

manuscript. This work was supported by a VIDI grant (MAN; 917.76.310) from The Netherlands

Organization of Scientific Research and an FCT grant (CIBS; SFRH/BD/46946/2008). The authors

have no conflicting financial interests.

2

Chapter 2

42

Materials and methods

Mice

CD27-/-, CD70TGxCD27-/-, IFN-/- (denoted as control) and CD70TGxIFN-/- (denoted as

CD70TG) mice were all on a C57BL/6 background and housed at the animal research institute of

the AMC under specific-pathogen-free conditions. Animal experiments were approved by the

Animal Ethics Committee and performed in accordance with institutional and national guidelines.

Flow cytometry and cell sorting

Single cell suspensions of bone marrow were obtained by crushing tibias and femurs in a mortar

and pestle and filtering through 40 μm cell strainers. Single cell suspensions of spleens were

obtained by mincing the organ through 40 μm cell strainers. Erythrocytes in spleen and peripheral

blood were lysed with an ammonium chloride solution. For purification of LT-HSCs, bone marrow

cells were stained with biotin-conjugated antibodies against the lineage markers CD4 (GK1.5),

CD8α (53-6.7), B220 (RA3-6B2), CD11b (M1/70), Gr1 (RB6-8C5), Ter119 (Ly-76) and lineage

cells were depleted using streptavidin microbeads (Miltenyi Biotec) and MACS LS-columns

(Miltenyi Biotec). Lineage depleted cells were incubated with antibodies against CD34 (RAM34),

Flt3, Sca-1 (D7), c-Kit (2B8) and fluorochrome conjugated streptavidin and LT-HSCs were sorted

on a FacsAria (BD Biosciences). For analysis of hematopoietic progenitors by flow cytometry,

bone marrow cells were stained with antibodies against lineage markers, CD34, Flt3, Sca-1, c-Kit,

Cd16/32 (93), IL-7Rα (A7R34), CD48 (HM-48-1), CD150 (TCF15-12F12.2, Biolegend) and CD27

(LG.7F9) and identified as: LT-HSC (Lin-c-Kit+Sca-1+CD34-Flt3-), ST-HSC (Lin-c-Kit+Sca-

1+CD34+Flt3-), MPP (Lin-c-Kit+Sca-1+CD34+Flt3+), SLAM HSC (Lin-c-Kit+Sca-1+CD48-CD150+),

CMP (Lin-c-Kit+Sca-1-CD34lowCD16/32low), GMP (Lin-c-Kit+Sca-1-CD34+CD16/32+), MEP (Lin-c-

Kit+Sca-1-CD34-CD16/32-), CLP (Lin-c-KitlowSca-1lowIL-7Rα+). For identification of mature cells,

cells were stained with CD11b (M1/70), CD4 (GK1.5), CD8 (53-6-7), B220 (RA3-6B2) and

identified as: myeloid cells (CD11b+), T cells (CD4+/CD8+), B cells (B220+).For identification of

donor cells in chimeric mice, cells were stained with additional antibodies against CD45.1 (A20)

and CD45.2 (104). Where possible, cells were stained in the presence of anti-CD16/CD32 block

(2.4G2; a kind gift from Dr. Louis Boon, Bioceros BV) and dead cells were excluded by stringent

gating on single cells in combination with the use of propidium iodide. All antibodies and

secondary reagents were obtained from eBioscience, unless otherwise specified. Flow cytometry

analyses were performed on FACSCanto II (BD Biosciences) and data were analyzed using FlowJo

software (Tree Star).

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43

Transplantations

For transplantations, LT-HSCs from CD45.1 CD27-/- and CD45.2 CD27+/+ IFN-/- mice were

purified and mixed with whole bone marrow cells from CD27-/- (control chimeras) or CD70TG x

CD27-/- (CD70TG chimeras) recipient mice, washed and 250 donor CD27-/- and 250 CD27+/+ HSCs

together with 1 x 106 whole bone marrow cells from CD45.1.2 recipient mice were injected

intravenously into CD45.1.2 CD27-/- or CD70TG x CD27-/- mice lethally irradiated with a split

dose of 10 Gy. Recipient mice received 1.75 gram/liter neomycin in sterile water for the first 3

weeks after transplantation. For analysis of peripheral blood chimerism, blood was collected by

puncturing the vena saphena at indicated time points.

BRDU

BRDU (5 mg/mouse, Sigma) was injected intraperitoneally in 200 μl PBS. For BrdU detection,

cells were collected 24 hours after injection, stained and subsequently fixed and permeabilized

overnight with the Foxp3 staining buffer set (eBioscience), treated with DNAse (Sigma) as

described elsewhere48 and stained with anti-BrdU (B44, BD Biosciences) for 30 minutes at room

temperature.

5-Fluorouracil (5-FU)

5-FU (5 mg/mouse, Sigma) was injected in 200 l PBS intraperitoneally and bone marrow cells

were collected and analyzed 5 days later. For identification of HSCs after 5-FU injection, HSCs

were characterized as Lin-c-Kit+Sca-1+ without using CD11b in the lineage definition.

Micro-array

RNA was isolated from purified LT-HSCs from 8 control and 8 CD70TG mice (2 mice pooled per

microarray) with the RNeasy Plus Mini Kit (Qiagen). A two-round RNA amplification and labeling

was performed with Ambion’s MessageAmp kit for Illumina arrays. The labeled cRNA was

applied to Illumina’s MouseRef-8 v2.0 arrays, and hybridized, washed and scanned according to

the manufacturer’s protocol. Probe signals were summarized with Illumina’s BeadArray v1.3

software and subsequently normalized with the R package vsn49. Differential expression between

the two groups of mice was analyzed with the limma package in R50. Benjamini-Hochberg’s

method was used to correct the p-values for multiple testing. Raw and normalized microarray data

are available in NCBI’s Gene Expression Omnibus using entry nr. GSE31574.

Gene Ontology

To detect significantly over-represented and under-represented biological processes, gene ontology

analysis was performed using the functional annotation clustering tool DAVID (Database for

Annotation, Visualization, and Integrated Discovery)51. Significant enrichment of biological

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44

processes associated with up- and downregulated genes in CD70TG LT-HSCs was demonstrated

with a modified Fisher’s Exact p-value < 0.05.

Statistics

Mean values ± s.d. or s.e.m. are shown. Statistical analysis was performed using a two-tailed

Student’s t-test with GraphPad Prism software and significance is indicated by *, p < 0.05, **, p <

0.01, ***, p < 0.001.

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20. de Bruin AM, Buitenhuis M, van der Sluijs KF et al. Eosinophil differentiation in the bone marrow is inhibited by T cell-derived IFN-gamma. Blood. 2010;116(14):2559-2569.

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2

Chapter 3 Interferon-gamma impairs the self-renewal of

hematopoietic stem cells

Alexander M. de Bruin1, Berend Hooibrink2, Martijn A. Nolte1,3

1Department of Experimental Immunology, 2Department of Cell Biology and Histology, Academic

Medical Center, University of Amsterdam, Amsterdam, The Netherlands, 3Department of

Hematopoiesis, Sanquin Research and Landsteiner Laboratory, Amsterdam, The Netherlands.

Interferon-gamma impairs the self-renewal of hematopoietic stem cells

49

Abstract

Balancing the processes of hematopoietic stem cell (HSC) differentiation and self-renewal is

critical in maintaining life-long supply of blood cells. The bone marrow (BM) produces a stable

output of newly generated cells, but immunological stress conditions inducing leukopenia increase

the demand for peripheral blood cell supply1;2. How HSC self-renewal and commitment are

modulated during immune activation to meet the increased demand for hematopoietic

differentiation is largely unknown. It has previously been proposed that the pro-inflammatory

cytokine interferon- (IFN-) plays an important role in this process by promoting HSC

proliferation, thereby acting as a positive regulator of HSCs during chronic infection3. However,

we demonstrate here that IFN- rather impairs the maintenance of HSCs by directly reducing their

self-renewal capacity both in vitro and in vivo, and that IFN- impairs restoration of HSC numbers

upon viral infection. We show that IFN- reduces thrombopoietin (TPO)-mediated phosphorylation

of signal transducer and activator of transcription (STAT)5, which is an important positive

regulator of HSC self-renewal4. Furthermore, IFN- deregulates the expression of the STAT5-

mediated cell cycle genes CyclinD1 and p57. These findings demonstrate that IFN- is a negative

modulator of HSC self-renewal by modifying cytokine responses and expression of genes involved

in HSC proliferation. We postulate that the occurrence of BM failure in chronic inflammatory

conditions, such as aplastic anemia, HIV and graft-versus-host disease is related to a sustained

impairment of HSC self-renewal caused by chronic IFN--signaling in these disorders.

Introduction

Hematopoietic stress conditions, like immune activation, change the hematopoietic output from the

BM. Activated lymphocytes have been postulated to influence differentiation of hematopoietic

progenitor cells through direct cell-cell interactions mediated by costimulatory molecules5;6.

Additionally, activated T cells can suppress the formation of several hematopoietic lineages, such

as B cells7, erythrocytes8 and eosinophilic granulocytes9 through the production of the pro-

inflammatory cytokine IFN-. BM failure in multiple chronic inflammatory diseases has been

associated with elevated IFN- levels10-13 and the beneficial effect of immune suppressive drugs on

hematopoietic function in BM-suppressed patients might result from a reduction in IFN--

producing lymphocytes14. Which hematopoietic precursors are affected by IFN- in these diseases

is not known, though the collapse of multiple hematopoietic lineages suggests the failure of a

multipotent progenitor. It was recently implied that IFN- produced during chronic infection

promotes proliferation of long-term repopulating HSCs in vivo and that IFN- thereby positively

contributes to the maintenance and restoration of blood cell homeostasis upon immunological

stress3. However, this concept does not explain the negative impact that IFN- has on

3

Chapter 3

50

hematopoiesis in chronic inflammatory diseases. Besides, the mechanism by which IFN- can

influence HSC self-renewal has not yet been resolved.

Results

To address the effect of IFN- on HSC function, we cultured highly purified HSCs (Lin-c-Kit+Sca-

1+CD48-CD150+) with cytokines supporting both self-renewal and differentiation of HSCs with or

without IFN-. IFN- increased the percentage but not the absolute number of differentiated cells

(Lin+), whereas it decreased both the percentage and absolute number of progenitor cells (Lin-c-

Kit+) and HSCs (Lin-c-Kit+CD48-CD150+; Fig. 1A-B). To assess the functional capacity of these

remaining phenotypical HSCs, control and IFN-cultured cells were mixed and transferred to

lethally irradiated recipient mice, using the congenic marker CD45 to distinguish IFN--treated

(CD45.1.2) and control donor cells (CD45.1) from host cells (CD45.2; Fig. 1C). These experiments

showed long-term, multilineage reconstitution from control cells, but a complete failure from IFN-

-treated cells (Fig. 1D), thus demonstrating that in vitro treatment with IFN- reduces HSC

numbers both phenotypically and functionally.

To analyze at what level IFN- affects HSC maintenance in these cultures, purified HSCs were

labeled with Carboxyfluorescein succinimidyl ester (CFSE), cultured for 4 days with IFN- and

analyzed daily. IFN- impaired the expansion of total cells in culture (Fig. 2A) as well as HSC

numbers (Fig. 2B), which could not be attributed to increased cell death (data not shown). Based on

CFSE-dilution we conclude that the reduction in HSC numbers was due to an inhibitory effect of

IFN- on HSC proliferation (Fig. 2C-D). Reduction in HSC self-renewal was not due to an

increased commitment to differentiate, since IFN- did not change the percentage of self-renewing

CD48-CD150+ HSCs and more committed CD48+CD150+ progenitors, only their absolute number

(Fig. 2E&F). As HSCs are predominantly quiescent in naïve mice, we tested whether IFN-

influences this non-proliferative state. However, overnight culture with IFN did not change

expression of the proliferation marker Ki67 (Fig. S1A&B), which is absent in quiescent HSCs2, nor

the incorporation of 5-bromo-29-deoxyuridine (BrdU) in HSCs and downstream progenitors (Fig.

S1C&D). This demonstrates that IFN- does not directly affect HSC quiescence or cell cycle entry,

which corresponds with the observation that significant differences in HSC proliferation could only

be observed after three days of culture (Fig. 2A-D). We therefore conclude that IFN- does not

affect recruitment of quiescent HSCs into proliferation, but rather inhibits their subsequent self-

renewal divisions.

To examine whether IFN- has the same effect on HSC function in vivo, we infected WT and IFN-

-/- mice with the Armstrong strain of lymphocytic choriomeningitis virus (LCMV). LCMV

infection reduces BM-output and induces leukopenia, which is completely dependent on production

of type I interferons early after infection1. LCMV thereby poses significant pressure on the

3


51

Lineage

C-K

itControl

IFN

CD150

CD

48

33.7 65

73.4 25

2764.7

0.54

2278

C-K

it

Lineage CD150

CD

48

CD45.2

17.50.1

81.9

25.40.2

73.9

11.10.03

88.7

16.70.08

83.1

B cells T cells

CD

45.1

CD45.2

Myeloid cells

0.50

0.49

Lin-c-Kit+

Lin-c-Kit+ HSC

WBC

Control

Host

IFN

A B

C

E

% o

f M

ax

N.D. N.D. N.D.

Transfer cultured cells + 200.000 CD45.2 bonemarrow cells into lethally irradiated CD45.2 host

Control: CD45.1

IFN: CD45.1.2

*** ** *

D

Lin-c-Kit+ Lin+0

5000

10000

15000

20000

Control

IFN-

Nu

mb

ero

f ce

lls

0

50

100

150

Nu

mb

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Control

IFN-

Week 4 Week 8 Week 160

5

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25Control

IFN-

% o

f d

on

or-

der

ived

WB

C

******

Lineage

C-K

itControl

IFN

CD150

CD

48

33.7 6533.7 65

73.4 2573.4 25

2764.7 2764.7

0.540.54

2278 2278

C-K

it

Lineage CD150

CD

48

CD45.2

17.50.1

81.9

17.50.1

81.9

25.40.2

73.9

25.40.2

73.9

11.10.03

88.7

11.10.03

88.7

16.70.08

83.1

16.70.08

83.1

B cells T cells

CD

45.1

CD45.2

Myeloid cells

0.50

0.49

0.50

0.49

Lin-c-Kit+

Lin-c-Kit+ HSC

WBC

Control

Host

IFNControl

Host

IFN

A B

C

E

% o

f M

ax

N.D. N.D. N.D.

Transfer cultured cells + 200.000 CD45.2 bonemarrow cells into lethally irradiated CD45.2 host

Control: CD45.1

IFN: CD45.1.2

*** ** *

D

Lin-c-Kit+ Lin+0

5000

10000

15000

20000

Control

IFN-

Nu

mb

ero

f ce

lls

Lin-c-Kit+ Lin+0

5000

10000

15000

20000

Control

IFN-

Nu

mb

ero

f ce

lls

0

50

100

150

Nu

mb

ero

f H

SC

s

Control

IFN-

0

50

100

150

Nu

mb

ero

f H

SC

s

Control

IFN-

Week 4 Week 8 Week 160

5

10

15

20

25Control

IFN-

% o

f d

on

or-

der

ived

WB

C

******

Figure 1. IFN- reduces HSC maintenance in vitro. HSCs (Lin-c-Kit+Sca-1+CD48-CD150+) were purified and cultured for 7 days with SCF/TPO/IL-3/IL-6/Flt3-L with or without IFN-. (A) Representative plots showing the analysis for progenitors (Lin-c-Kit+), differentiated cells (Lin+), and HSCs (Lin-c-Kit+CD48-CD150+) and (B) absolute numbers of these cells. Data represent three independent experiments with 4-5 wells per condition. (C) HSCs from CD45.1 and CD45.1.2 mice were cultured, (CD45.1 HSCs without IFN-, CD45.1.2 HSCs with IFN-), pooled and analysed and injected into CD45.2 recipient mice. (D) Representative plots showing donor contribution to total white blood cells (WBC) and B, T and myeloid cell lineages in peripheral blood of lethally irradiated CD45.2 recipient mice at 4 weeks after transplantation. (E) Donor contribution to total white blood cells at indicated weeks (n = 8). Experiment was repeated with CD45.1.2 HSCs cultured without IFN- and CD45.1 HSCs with IFN- with similar results. Mean values ± s.e.m. are shown. *, p < 0.05, **, p < 0.01, ***, p < 0.001. N.D.: not detectable.

3

Chapter 3

52

96.6

CD

48

CFSE

Day 1 Day 2 Day 3 Day 4Day 0

55 20 13 10

Control

61 26 14 11

IFN-

1 2 3 40

200

400

600

800

1000 Control

IFN-

Day

Exp

ansi

on

of

cells

in %

0 1 2 3 40

50

100 Control

IFN-

Day

% H

SC

1 2 3 40

1

2

3

4Control

IFN-

Day

Div

isio

n In

dex

1 2 3 40

50

100

150Control

IFN-

Day

Ex

pa

ns

ion

of

HS

C i

n %

Day 1 Day 2 Day 3 Day 4

E

F

A B

C D

*

**

*

**

***

IFN-Control

CD150

*

96.696.6

CD

48

CFSE

Day 1 Day 2 Day 3 Day 4Day 0

5555 2020 1313 1010

Control

6161 2626 1414 1111

IFN-

1 2 3 40

200

400

600

800

1000 Control

IFN-

Day

Exp

ansi

on

of

cells

in %

0 1 2 3 40

50

100 Control

IFN-

Day

% H

SC

1 2 3 40

1

2

3

4Control

IFN-

Day

Div

isio

n In

dex

1 2 3 40

50

100

150Control

IFN-

Day

Ex

pa

ns

ion

of

HS

C i

n %


E

F

A B

C D

*

**

*

**

***

IFN-Control

CD150

*

Figure 2. IFN- impairs self-renewal of HSCs. Purified HSCs (Lin-c-Kit+CD48-CD150+) were labelled with CFSE and cultured for 4 days with SCF/TPO/IL-3/IL-6/Flt3-L with or without IFN-. (A) Expansion of total cells in culture relative to day 1 and (B) expansion of HSCs (Lin-c-Kit+CD48-CD150+) relative to day 1. (C) Histograms and (D) division index (defined as the average number of divisions that a cell (that was present in the starting population) has undergone) of CFSE-labelled HSCs cultured with or without IFN-. (E) Representative plots showing percentage of HSC (Lin-c-Kit+CD48-CD150+) in cultures and (F) quantification of these data. Histograms and plots are representative of three independent experiments. Graphs represent data pooled from three independent experiments with duplicate cultures. Mean values ± s.e.m. are shown. *, p < 0.05, **, p < 0.01, ***, p < 0.001.

3


53

hematopoietic system to restore blood cell homeostasis. As expected, HSC (Lin-c-Kit+CD48-

CD150+) numbers were equally reduced in WT and IFN--/- mice at day 4 of infection. However,

HSC numbers were restored to normal levels in IFN--/- mice at day 8, while only a slight recovery

of WT HSCs was observed at day 12 (Fig. 3A). Impaired recovery of WT HSCs around day 8

paralleled the occurrence of IFN--producing LCMV-specific CD8 T cells (data not shown and 15).

To exclude indirect effects of IFN- on HSCs, we generated mixed chimeric mice with BM from

both WT and IFN-R1-/- mice. Two months after transplantation, donor chimerism of HSCs from

IFN-R1-/- mice was higher than from WT mice (Fig. 3B, day 0), suggesting that homeostatic levels

of IFN- already influence HSC self-renewal. Upon LCMV infection, HSC numbers of both donors

dropped equally early after infection, but IFN-R1-/- HSC recovered much better than their WT

counterparts within the same host (Fig. 3C). Importantly, there was no difference in HSC

quiescence (Fig. 3D), indicating that the impaired recovery of WT HSCs numbers was due to an

IFN--dependent reduction in HSC self-renewal capacity, rather than a decrease in the fraction of

cycling HSCs. We therefore conclude that IFN- directly reduces HSC self-renewal, both in vitro

and in vivo.

0 4 8 120.000

0.002

0.004

0.006

0.008WT

IFN--/-

Day

Nu

mb

er o

f H

SC

s (x

106)

0 4 8 120

20

40

60

80

100WT

IFN-R1-/-

Day

% o

f d

on

or-

de

rive

d H

SC

s

0 4 8 120.000

0.002

0.004

0.006WT

IFN-R1-/-

Day

Nu

mb

er o

f

do

no

r-d

eriv

ed

HS

Cs

(x1

06)

0 4 8 120

20

40

60

80

100WT

IFN-R1-/-

Day

% K

i-6

7+ H

SC

of

do

no

r

A B

C D

** **

**

******

***

**

0 4 8 120.000

0.002

0.004

0.006

0.008WT

IFN--/-

Day

Nu

mb

er o

f H

SC

s (x

106)

0 4 8 120

20

40

60

80

100WT

IFN-R1-/-

Day

% o

f d

on

or-

de

rive

d H

SC

s

0 4 8 120.000

0.002

0.004

0.006WT

IFN-R1-/-

Day

Nu

mb

er o

f

do

no

r-d

eriv

ed

HS

Cs

(x1

06)

0 4 8 120

20

40

60

80

100WT

IFN-R1-/-

Day

% K

i-6

7+ H

SC

of

do

no

r

A B

C D

** **

**

******

***

**

Figure 3. IFN- inhibits HSC recovery after LCMV infection. (A) WT and IFN--/- mice were infected with LCMV and the number of HSCs (Lin-c-Kit+CD48-CD150+) was measured by flow cytometry at indicated days (n = 3-5). Data represent three independent experiments. Mixed BM-chimeric mice were generated with WT (CD45.1) and IFN-R1-/- (CD45.2) BM (1:1 ratio), infected with LCMV and (B) donor HSC chimerism, (C) donor HSC numbers and (D) proliferation of donor HSCs were measured at indicated days (n = 3-5). Data are representative for two independent experiments. Mean values ± s.e.m. are shown. *, p < 0.05, **, p < 0.01, ***, p < 0.001.

3

Chapter 3

54

To investigate the molecular mechanism by which IFN- regulates HSC self-renewal, we examined

both extrinsic and intrinsic pathways that mediate this process. The cytokine thrombopoietin (TPO)

is important for HSC self-renewal, as its receptor, c-mpl, can induce STAT5 phosphorylation and

thereby regulate transcription of self-renewal genes4;16;17. Competitive repopulation capacity of

STAT5-/-or c-mpl-/- HSCs18;19 is severely impaired, while constitutively active STAT5 enhances the

self-renewal and repopulation activity of HSCs4. We found that IFN- reduced TPO-mediated

phosphorylation of STAT5 in purified HSCs (Fig. 4A,B,C) and fully annihilated TPO-driven HSC

expansion (Fig. 4D). IFN- induced IRF-1 expression in HSCs, thus confirming direct activation of

IFN-R signaling, but also induced expression of suppressor of cytokine singling 1 (SOCS1; Fig.

4E), a negative regulator of IFN-R signaling through its ability to inhibit STAT1

phosphorylation20. However, SOCS1 can also inhibit STAT5 phosphorylation20, which can thereby

explain the perturbed TPO-signaling in HSCs by IFN-.

Next, we investigated the impact of IFN- on molecular mediators of HSC proliferation and found

that IFN- significantly reduced the expression of Cyclin D1 and inhibited TPO-mediated

downregulation of cell cycle inhibitor p57 (Fig. 4F). CyclinD1 and p57 are both important

mediators of HSC self-renewal and their expression is associated with TPO-signaling and STAT5

expression16;17. These findings are also relevant for the in vivo situation, as Cyclin D1 and p57 were

significantly down- and upregulated, respectively, in WT HSCs, but not IFN-R1-/- HSC from

mixed BM chimeric mice upon LCMV infection (Fig. 4G). The combined decrease in STAT5

phosphorylation and changes in expression of these key cell cycle genes provide a molecular

explanation on how IFN- negatively affects the self-renewal capacity of HSCs both in vitro and in

vivo.

Discussion

Our findings challenge a recent report in Nature, which concluded that IFN- induces HSC

proliferation in vitro and upon infection with Mycobacterium Avium, and caused a concomitant

decrease in myeloid progenitors3. We postulate that part of these conflicting findings can be

explained by indirect effects of IFN- on other cell types, which we have carefully excluded by

treating HSCs with IFN- after rather than before cell-sorting and by using WT:IFN-R1-/- mixed

chimeric mice for in vivo studies. Another possible explanation is related to the use of Sca-1 for

identification of HSCs and progenitor cells. Sca-1 is an interferon-responsive molecule that is

highly upregulated by IFN- on all hematopoietic progenitors upon treatment in vitro (Fig. S2A) or

infection in vivo (Fig. S2B). As a consequence, myeloid progenitor cells (normally Sca-1-), of

which a fraction expresses CD150, become Sca-1+ and thereby substantially contaminate the HSC

pool and reduce the myeloid progenitor fraction (Fig. S2C). We ruled out this contamination by not

using Sca-1 when identifying progenitor cells that had been exposed to interferons and by

3


55

excluding CD48+ myeloid progenitors. As shown previously21, Sca-1 is not required when HSCs

are identified as Lin-c-Kit+CD48-CD150+ cells, both in vivo (Fig. S2D) and in vitro (Fig. S2E).

Importantly, Sca-1 upregulation possibly also explains other studies that have reported infection-

and interferon-induced increases in HSC numbers22-24.

Cyclin D1

0.0

0.5

1.0

1.5

2.0

SCF SCF/TPO

Rel

ativ

e ex

pre

ssio

n

P57

0

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SCF SCF/TPO

Rel

ativ

e ex

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D

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IFN-

% o

f ce

lls p

ST

AT

5+

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1000

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3000Control

IFN-

Geo

MF

I pS

TA

T5

pSTAT5

% o

f M

ax

TPO induced pSTAT5

TPOTPO + IFN-

medium

*

**

Cyclin D1

0.0

0.5

1.0

1.5

LCMVControl

WT IFN-R1-/-

Rela

tive e

xp

ressio

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p57

0.0

0.5

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LCMVControl

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Rela

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B C

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E

***

*IFN-Control

SOCS1

0

5

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SCF SCF/TPO

Rel

ativ

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***IFN-

Control

IRF1

0

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SCF SCF/TPO

Rel

ativ

e ex

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ssio

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*

IFN-:TPO:

0

50

100

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Nu

mb

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f H

SC

s

- + +- - +

**

Cyclin D1

0.0

0.5

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SCF SCF/TPO

Rel

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Rel

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Geo

MF

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T5

pSTAT5

% o

f M

ax

TPO induced pSTAT5

TPOTPO + IFN-

medium

*

**

Cyclin D1

0.0

0.5

1.0

1.5

LCMVControl

WT IFN-R1-/-

Rela

tive e

xp

ressio

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LCMVControl

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Rela

tive e

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*

*IFN-Control

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IFN-Control

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*** **

E

***

*IFN-Control

SOCS1

0

5

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15

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SCF SCF/TPO

Rel

ativ

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pre

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n**

***IFN-

Control

IRF1

0

5

10

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SCF SCF/TPO

Rel

ativ

e ex

pre

ssio

n

*

IFN-:TPO:

0

50

100

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Nu

mb

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f H

SC

s

- + +- - +

** *

IFN-:TPO:

0

50

100

150

Nu

mb

er o

f H

SC

s

- + +- - +

**

Figure 4. IFN- reduces TPO-mediated STAT5 phosphorylation and affects expression of genes involved in cell cycle regulation. (A) Histogram showing pSTAT5 staining of purified HSCs (Lin-c-Kit+CD48-CD150+) cultured for 24 hours with SCF (shaded histogram), SCF and TPO (bold histogram) or SCF, TPO and IFN- (thin histogram). (B) Percentage of cells stained positive for pSTAT5 and (C) geometric mean fluorescence intensity of pSTAT5. Data represent triplicate cultures and are representative for two independent experiments. (D) Purified HSCs were cultured as in Fig. 1 with or without IFN- and/or TPO and HSC numbers were measured. Data are representative for two experiments with four wells per condition. (E,F) mRNA expression levels of indicated genes in HSCs cultured for 24 hours with SCF or SCF and TPO with or without IFN-. Expression levels are relative to the expression in SCF/TPO cultured HSCs. qPCRs were performed in duplicate and data is pooled from two independent experiments with 1-3 cultures per condition. (G) mRNA expression levels of indicated genes in WT and IFN-R1-/- HSCs from chimeric mice 8 days after LCMV infection. Expression levels are relative to the expression in WT and IFN-R1-/- HSCs from naïve WT: IFN-R1-/- chimeric mice. qPCRs were performed in duplicate of 2-3 pooled samples of purified HSCs. Mean values ± s.d. are shown. *, p < 0.05, **, p < 0.01, ***, p < 0.001.

3

Chapter 3

56

The observation that IFN- directly inhibits HSC self-renewal, but neither HSC quiescence nor

differentiation supports previous studies demonstrating that IFN- impairs HSC function, without

affecting cell cycle entry or viability25;26. Although we did not find evidence that IFN- induced

differentiation of HSCs in vitro, we do not exclude that IFN- acts as an inducer of HSC

differentiation in vivo. Different BM niches regulate the diverse fates of HSCs, like quiescence,

self-renewal and differentiation27. IFN- signaling might be required to impair HSC self-renewal in

order to relocalize them to a microenvironment supporting differentiation. Alternatively, sustained

production of IFN- might reduce HSC self-renewal in order to prevent exhaustion and loss of HSC

activity resulting from excessive HSC proliferation during hematopoietic stress. In both cases,

chronic exposure to IFN- results in declining HSC numbers and impaired hematopoietic output. It

has been postulated that IFN-α induces proliferation of HSCs24, but the use of Sca-1 and frequent

omission of CD48 upon HSC identification in that study could also have resulted in an

overestimation of HSC numbers. Moreover, part of the observed HSC proliferation upon IFN-

injection was found to be an indirect effect, most likely resulting from feedback mechanisms

triggered by the IFN-α-induced leukopenia24. Therefore, it remains unclear if and how IFN-α

affects HSC proliferation directly. However, IFN-α might have similar inhibitory effects on HSC

self-renewal as IFN-, since IFN-α also upregulates SOCS1 and represses megakaryopoiesis by

reducing TPO-mediated STAT5 phosphorylation28.

Although temporary inhibition of HSC self-renewal during acute infection will not immediately

influence peripheral blood cell numbers, prolongation of this process will threaten the maintenance

and/or restoration of blood cell homeostasis. Better understanding of the underlying mechanism is

also clinically important, because increased IFN- production has been associated with bone

marrow-failure in patients with chronic inflammation, such as aplastic anemia Fanconi anemia,

GvhD and HIV10-13. In fact, IFN- neutralization improves the in vitro capacity of hematopoietic

progenitors from patients with aplastic anemia29. Our study suggests that the negative impact of

IFN- on HSC self-renewal contributes to the impaired hematopoietic function in multiple human

diseases and we speculate that repression of IFN- signaling could alleviate the hematopoietic

suppression in patients with chronic inflammation.

Acknowledgements

We thank Claudia Brandão, Klaas van Gisbergen and Sten Libregts for technical assistance and

Prof. Dr. Rene van Lier, Dr. Marieke von Lindern, Dr. Esther Nolte-‘t Hoen and Dr. Monika

Wolkers for critical reading of the manuscript. AMdB and MAN are supported by The Netherlands

Organization of Scientific Research (VIDI grant 91776310).

3


57

Materials and Methods

Mice

Wild-type C57BL/6 (CD45.2), C57BL/6.SJL (CD45.1), litters of wild-type C57BL/6 and

C57BL/6.SJL (CD45.1.2), and IFN--/- C57BL/6 (CD45.2) were used. IFN-R1-/- C57BL/6

(CD45.2) (Stock no. 3288) mice were obtained from The Jackson Laboratory. All animals were

housed at the animal research institute of the AMC under specific-pathogen-free conditions.

Animal experiments were approved by the Animal Ethics Committee and performed in accordance

with institutional and national guidelines. For LCMV infection, mice were injected

intraperitoneally with 1 x 105 PFU of LCMV clone Armstrong in 200 l PBS.

Transplants

For transplantation of cultured HSCs, CD45.1 and CD45.1.2 cultured cells were pooled, thoroughly

washed in PBS, mixed with 2 x 105 CD45.2 whole BM cells and injected intravenously into

CD45.2 mice lethally irradiated with a split dose of 10 Gy. For WT:IFN-R1-/- chimeras, 2 x 106

WT CD45.1.2 whole BM cells were mixed with 2 x 106 IFN-R1-/- CD45.2 and injected into

irradiated WT CD45.1 recipients. Recipient mice received 1.75 gram/liter neomycin in sterile water

for the first 3 weeks after transplantation. Chimeric mice were infected with LCMV 2 months after

transplantation.

Quantitative real-time PCR

RNA was extracted using Trizol (Invitrogen) and cDNA was made with random hexamers and

Superscript II reverse transcriptase (Roche). Quantitative real-time PCR was performed in

duplicate with Express SYBR GreenER reagents (Invitrogen) on the StepOnePlus RT-PCR system

(Applied Biosystems) and data was normalized using 18S as a reference gene. Primer sequences

available on request.

Cultures

For 7-day cultures, 150 HSCs (Lin-c-Kit+Sca-1+CD48-CD150+) were sorted directly in 96-well

plates cultured in X-VIVO 15 medium (Lonza) supplemented with 5 ng/ml TPO, SCF, IL-3, IL-6

and Flt3-L. IFN- (20 ng/ml) was added when indicated. Cells were cultured at 37°C in a

humidified incubator at 5% CO2 For STAT5 analysis and RNA collection, HSCs (Lin-c-Kit+Sca-

1+CD48-CD150+) were cultured for 24 hours with SCF or SCF and TPO in the presence or absence

of IFN-. All cytokines were obtained from Peprotech. For BrdU incorporation, HSCs (Lin-c-

Kit+Sca-1+CD34-), ST-HSCs and MPPs (Lin-c-Kit+Sca-1+CD34+) and CMPs (Lin-c-Kit+Sca-1-

CD34+CD16/32low) were purified and cultured for 18 hours with SCF or SCF and IFN- for 18

3

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58

hours. 10 M BrdU (Sigma) was added for 18 hours (HSCs) or for the last 3 hours of culture

(CD34+LKS and CMP). For CFSE-labeling, purified HSCs (Lin-c-Kit+Sca-1+CD48-CD150+) were

washed in PBS, labeled with 0.5 M CFSE (Invitrogen) in PBS for 12 minutes at 37C, washed

and subsequently cultured.

Flowcytometry

FACSCanto II and FACSAria were used for flow cytometric analysis and cell sorting. For cell

sorting, whole BM was stained with biotinylated lineage markers CD4 (GK1.5), CD8α (53-6.7),

B220 (RA3-6B2), CD11b (M1/70), Gr1 (RB6-8C5), Ter119 (Ly-76) and IL-7Rα (B12-1) and

enriched for progenitors by negative depletion of labeled lineage cells using streptavidin (SA)

microbeads and MACS LS-Columns (Miltenyi Biotec). Cells were stained with CD34 (RAM34),

Sca-1 (D7), fluorochrome conjugated SA, CD48 (HM-48-1), CD150 (TCF15-12F12.2, Biolegend),

CD16/32 (93) and c-Kit. HSCs in all experiments were purified as Lin-c-Kit+Sca-1+CD48-CD150+.

For purification of HSCs from LCMV infected chimeric mice cells were stained with additional

antibodies against CD45.1 (A20) and CD45.2 (104) and purified as CD45.1+ or CD45.2+ Lin-c-

Kit+CD48-CD150+. For the BrdU incorporation assay, cells were purified as Lin-c-Kit+Sca-1+CD34-

(HSCs), Lin-c-Kit+Sca-1+CD34+ (ST-HSCs and MPPs) and Lin-c-Kit+Sca-1-CD34+CD16/32low

(CMPs). For immunophenotypic characterization of HSCs in culture or in mice, cells were stained

as above, with additional antibodies against CD45.1 (A20) and CD45.2 (104) when required, and

identified as Lin-c-Kit+CD48-CD150+ in all experiments. Dead cells were excluded by stringent

gating on single cells and by using propidium iodide. For staining of phosphorylated STAT5,

cultured cells were directly fixed for 10 minutes at 37°C by adding an equal volume of BD

Cytofix/Cytoperm buffer (BD Biosciences), chilled on ice for 1 minute, washed, and fixed with -

20°C 90% methanol for a least 1 hour. Cells were washed and incubated with pSTAT5 antibody

(47, BD Biosciences) at room temperature for 45 minutes. For detection of incorporated BrdU,

cells were treated as described elsewhere30 and BrdU and DNA content was visualized using anti-

BrdU (PRB-1) and propidium iodide. All antibodies were obtained from eBioscience, unless

indicated otherwise.

Statistics

Mean values ± s.d. or s.e.m. are shown. Statistical analysis was performed using a two-tailed

Student’s t-test with GraphPad Prism software and significance is indicated by *, p < 0.05, **, p <

0.01, ***, p < 0.001.

References 1. Binder D, Fehr J, Hengartner H, Zinkernagel RM. Virus-induced transient bone marrow

aplasia: major role of interferon-alpha/beta during acute infection with the noncytopathic lymphocytic choriomeningitis virus. J.Exp.Med. 1997;185(3):517-530.

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2. Wilson A, Laurenti E, Oser G et al. Hematopoietic stem cells reversibly switch from dormancy to self-renewal during homeostasis and repair. Cell. 2008;135(6):1118-1129.

3. Baldridge MT, King KY, Boles NC, Weksberg DC, Goodell MA. Quiescent haematopoietic stem cells are activated by IFN-gamma in response to chronic infection. Nature. 2010;465(7299):793-797.

4. Kato Y, Iwama A, Tadokoro Y et al. Selective activation of STAT5 unveils its role in stem cell self-renewal in normal and leukemic hematopoiesis. J.Exp.Med. 2005;202(1):169-179.


6. Nolte MA, Arens R, van OR et al. Immune activation modulates hematopoiesis through interactions between CD27 and CD70. Nat.Immunol. 2005;6(4):412-418.




10. Dufour C, Corcione A, Svahn J et al. TNF-alpha and IFN-gamma are overexpressed in the bone marrow of Fanconi anemia patients and TNF-alpha suppresses erythropoiesis in vitro. Blood. 2003;102(6):2053-2059.

11. Fuchs D, Zangerle R, Artner-Dworzak E et al. Association between immune activation, changes of iron metabolism and anaemia in patients with HIV infection. Eur.J.Haematol. 1993;50(2):90-94.

12. Iwasaki T, Hamano T, Saheki K et al. Graft-versus-host-disease-associated donor cell engraftment in an F1 hybrid model is dependent upon the Fas pathway. Immunology. 2000;99(1):94-100.

13. Zoumbos NC, Gascon P, Djeu JY, Young NS. Interferon is a mediator of hematopoietic suppression in aplastic anemia in vitro and possibly in vivo. Proc.Natl.Acad.Sci.U.S.A. 1985;82(1):188-192.

14. Selleri C, Maciejewski JP, Catalano L et al. Effects of cyclosporine on hematopoietic and immune functions in patients with hypoplastic myelodysplasia: in vitro and in vivo studies. Cancer. 2002;95(9):1911-1922.

15. Binder D, van den Broek MF, Kagi D et al. Aplastic anemia rescued by exhaustion of cytokine-secreting CD8+ T cells in persistent infection with lymphocytic choriomeningitis virus. J.Exp.Med. 1998;187(11):1903-1920.

16. Matsumura I, Kitamura T, Wakao H et al. Transcriptional regulation of the cyclin D1 promoter by STAT5: its involvement in cytokine-dependent growth of hematopoietic cells. EMBO J. 1999;18(5):1367-1377.

17. Yoshihara H, Arai F, Hosokawa K et al. Thrombopoietin/MPL signaling regulates hematopoietic stem cell quiescence and interaction with the osteoblastic niche. Cell Stem Cell. 2007;1(6):685-697.

18. Bunting KD, Bradley HL, Hawley TS et al. Reduced lymphomyeloid repopulating activity from adult bone marrow and fetal liver of mice lacking expression of STAT5. Blood. 2002;99(2):479-487.

19. Kimura S, Roberts AW, Metcalf D, Alexander WS. Hematopoietic stem cell deficiencies in mice lacking c-Mpl, the receptor for thrombopoietin. Proc.Natl.Acad.Sci.U.S.A. 1998;95(3):1195-1200.

20. Krebs DL, Hilton DJ. SOCS proteins: negative regulators of cytokine signaling. Stem Cells. 2001;19(5):378-387.


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23. Singh P, Yao Y, Weliver A et al. Vaccinia virus infection modulates the hematopoietic cell compartments in the bone marrow. Stem Cells. 2008;26(4):1009-1016.




27. Ehninger A, Trumpp A. The bone marrow stem cell niche grows up: mesenchymal stem cells and macrophages move in. J.Exp.Med. 2011;208(3):421-428.

28. Wang Q, Miyakawa Y, Fox N, Kaushansky K. Interferon-alpha directly represses megakaryopoiesis by inhibiting thrombopoietin-induced signaling through induction of SOCS-1. Blood. 2000;96(6):2093-2099.

29. Laver J, Castro-Malaspina H, Kernan NA et al. In vitro interferon-gamma production by cultured T-cells in severe aplastic anaemia: correlation with granulomonopoietic inhibition in patients who respond to anti-thymocyte globulin. Br.J.Haematol. 1988;69(4):545-550.


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Supplemental figures

96 0.09

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Supplemental figure S1. IFN does not affect quiescence of HSCs. (A) Representative plots showing gating of HSCs (Lin-c-Kit+CD48-CD150+) and expression of Ki-67 in WT HSCs after overnight culture of total bone marrow with SCF in the presence or absence of IFN and (B) quantification of these experiments. Data represent two independent experiments with triplicate cultures. (C) Representative plots showing DNA content and BRDU incorporation in purified HSCs (CD34-LKS), short term-HSCs (ST-HSCs) and multipotent progenitors (MPPs) (CD34+LKS) and common myeloid progenitors (CMPs) (Lin-c-Kit+Sca-1-CD34+CD16/32low) cultured for 18 hours with SCF or SCF and IFN- in the presence of BRDU and (D) quantification of these experiments. Plots are representative of three independent experiments. Bar graphs represent data pooled from three independent experiments with 1-3 cultures per condition. *, p < 0.05, **, p < 0.01, ***, p < 0.001.

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Supplemental figure S2. IFN- induces Sca-1 expression resulting in aberrant HSC identification when CD48 expressing cells are not excluded. (A) Representative plots showing Sca-1 expression on purified LKS cells and myeloid progenitor cells after overnight culture with or without IFN- and (B) expression of Sca-1 on Lin-c-Kit+ cells in WT and IFN--/- naïve or LCMV-infected mice. (C) Representative plots showing identification of HSCs as Lin-c-Kit+(Sca-1+/-)CD150+ or Lin-c-Kit+(Sca-1+/-)CD150+CD34- in WT naïve mice and after infection. HSC gating is shown for Lin-c-Kit+ cells (which equals Lin-c-Kit+Sca-1- cells in infected mice), Lin-c-Kit+Sca-1+ cells and Lin-c-Kit+Sca-1- cells and shows the contamination of HSCs by progenitors

Sca-1

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0.127 0.127 0

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0.005020.00611 0.337 0.301

Lineage-c-Kit+ Lineage-c-Kit+Sca-1+ Lineage-c-Kit+Sca-1+Lineage-c-Kit+

Lineage-c-Kit+ Lineage-c-Kit+Sca-1+ Lineage-c-Kit+Sca-1- Lineage-c-Kit+ Lineage-c-Kit+Sca-1+ Lineage-c-Kit+Sca-1-

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0.005020.005020.006110.00611 0.3370.337 0.3010.301

Lineage-c-Kit+ Lineage-c-Kit+Sca-1+ Lineage-c-Kit+Sca-1+Lineage-c-Kit+

Lineage-c-Kit+ Lineage-c-Kit+Sca-1+ Lineage-c-Kit+Sca-1- Lineage-c-Kit+ Lineage-c-Kit+Sca-1+ Lineage-c-Kit+Sca-1-

CD150

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B

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D E

myeloid progenitors when HSCs are identified as Lin-c-Kit+Sca-1+CD150+ or Lin-c-Kit+Sca-1+CD150+CD34- cells in mice undergoing an infection. Representative plots showing near identical numbers of HSCs when identified as Lin-c-Kit+Sca-1+CD150+CD48- or Lin-c-Kit+CD150+CD48- (D) in mice and (E) after culture. Numbers in all plots indicate percentages of total viable cells.

3

Chapter 4 Eosinophil differentiation in the bone marrow is

inhibited by T cell-derived IFN

Alexander M. de Bruin1, Miranda Buitenhuis2, Koenraad F. van der Sluijs1,3, Klaas P.J.M. van

Gisbergen1, Louis Boon4 and Martijn A. Nolte1

1 Department of Experimental Immunology and 3 Department of Pulmonology, Academic Medical

Center, University of Amsterdam, Amsterdam, The Netherlands. 2 Department of Hematology,

Erasmus Medical Center, Rotterdam, The Netherlands. 4Bioceros BV, Yalelaan 46, 3584 CM

Utrecht, The Netherlands.

Blood 2010 Oct 7;116(14):2559-69

Eosinophil differentiation in the bone marrow is inhibited by T cell-derived IFN

65

Abstract

To explore if and how T cells can affect myelopoiesis, we investigated myeloid differentiation in a

model for T cell-mediated immune activation. We found that CD70-transgenic (CD70TG) mice,

which have elevated numbers of IFN-producing effector T cells in the periphery and bone

marrow, are almost devoid of eosinophilic granulocytes. Induction of allergic airway inflammation

in these mice failed to induce eosinophilia as well as airway hyperresponsiveness. CD70TG mice

also have strongly reduced numbers of eosinophil lineage-committed progenitors, whereas

granulocyte/macrophage progenitors (GMPs) from these mice are unable to generate eosinophils in

vitro. We found that GMPs express IFNR1 and that IFN is sufficient to inhibit eosinophil

differentiation of both murine and human progenitor cells in vitro. We demonstrate that inhibition

of eosinophil development in CD70TG mice is IFN-dependent and that T cell-derived IFN is

sufficient to inhibit eosinophil formation in vivo. Finally, we found that IFN produced upon anti-

CD40 treatment and during viral infection can also suppress eosinophil formation in wildtype mice.

These data demonstrate that IFN inhibits the differentiation of myeloid progenitors to eosinophils,

indicating that the adaptive immune system plays an important role in orchestrating the formation

of the appropriate type of myeloid cells during immune activation.

Introduction

Differentiation of hematopoietic progenitor cells to all separate lineages of mature red and white

blood cells is tightly controlled by cytokines locally produced by stromal cells in the bone marrow.

However, this process can also be influenced by bone marrow external factors. In particular, T cells

have been implicated in steady state hematopoiesis, as T cell-deficient mice have impaired

maturation and accumulation of myeloid progenitors in the bone marrow, which can be restored by

reconstitution with CD4+ T cells1. The bone marrow harbors a relatively large number of memory

T cells and it also recruits antigen-specific effector T cells during the course of an immune

response (reviewed by Di Rosa and Pabst2). We have previously postulated that activated

lymphocytes could influence the function of hematopoietic progenitor cells through direct cell-cell

contact3, but it is also clear from a variety of disease models that T cells can modulate myelopoiesis

during immune activation via production of cytokines like IL-3, IL-5, IL-17, Oncostatin M or GM-

CSF (reviewed by Dent and Kaplan4). These data imply that the adaptive immune system is able to

directly modulate the formation of new blood cells during immune activation by the migration of

activated T cells to the bone marrow.

The ability of activated T cells to modulate the formation of myeloid cells is particularly relevant in

asthma, as the characteristic occurrence of eosinophilia in this disease is accompanied by an

increase of IL-5 producing T helper 2 (Th2) cells in the bone marrow5;6. Moreover, induction of

Th1 responses prior to the induction of asthma decreases the levels of IL-5 in the lung and reduces

4

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local eosinophilia7;8. It has therefore been proposed that IL-5 production by Th2 cells induces

eosinophil accumulation and that Th1 cells merely inhibit lung eosinophilia by antagonizing the

formation of Th2 cells. However, it is not clear from these studies whether Th1 cells can also

directly affect the formation of eosinophils in the bone marrow. This is important, because the

prototypical Th1 cytokine IFN can directly affect proliferation of hematopoietic progenitor cells

and their differentiation towards myeloid cells9-11. Since eosinophils play an important role both in

allergic diseases and during helminth infections, it is important to understand how eosinophil

formation is regulated. Therefore, we investigated if and how IFN-producing T cells affect

eosinophil formation in vivo.

Myeloid cells in the bone marrow originate from a well-defined population of precursor cells,

known as the common myeloid progenitor (CMP), which can give rise to monocytes, mast cells

and granulocytes via the intermediate granulocyte/macrophage progenitor (GMP) and to platelets

and erythrocytes via the megakaryocyte/erythrocyte progenitor12. GMPs give rise to eosinophilic

granulocytes through an intermediate eosinophil lineage-committed progenitor (EoP) in mice13,

whereas the human EoP is derived from the CMP or its upstream multipotent progenitor14.

Expression of the IL-5 receptor on EoPs is a result of commitment of GMPs to the eosinophil

lineage, which makes that IL-5 can support eosinophil development from EoPs rather than instruct

GMPs to commit to the eosinophil lineage13.

To study how eosinophil development in the bone marrow is regulated by activated T cells, we

made use of a mouse model in which the formation of effector T cells is enhanced due to

overexpression of the costimulatory ligand CD70. Under normal circumstances, CD70 is only

transiently expressed on activated DCs, T and B cells, but constitutive expression of CD70 on

either of these cell types induces strong type 1 effector T cell formation due to enhanced

costimulation through its receptor CD2715-18. CD70-transgenic (CD70TG) mice develop high

numbers of IFNγ-producing effector CD4 and CD8 T cells, which is dependent on CD27-

expression and T cell receptor stimulation15;19. Although the antigens driving this T cell activation

are not known, CD70TG mice mount a protective T cell response against a challenge with

influenza virus or tumor cells20. NK cell numbers are strongly reduced in these mice and the

remaining NK cells produce less IFN than WT mice21. Here we describe that CD27-mediated T

cell activation has a strong and specific impact on myelopoiesis, as CD70TG mice are almost

devoid of eosinophils, while the formation of neutrophils and monocytes is actually increased.

Detailed analysis revealed that this defect in eosinophil formation can be fully attributed to the

increased production of IFNγ by effector T cells in these mice. We show that IFN is sufficient to

block eosinophil development in vitro and in vivo by affecting the formation of EoPs from GMPs,

which demonstrates that IFN is inhibitive for eosinophil development. These data reveal that type

4


67

1 effector T cells can directly regulate hematopoiesis by influencing the differentiation capacity of

specific myeloid progenitor cells.

Results

CD70TG mice lack eosinophils and have increased numbers of neutrophils and monocytes

To examine the influence of CD27-driven T cell activation on myelopoiesis, we analyzed the

presence of myeloid cells in bone marrow, blood and spleen of WT and B cell-specific CD70TG

mice. In all these compartments, we found that CD70TG mice were almost completely devoid of

eosinophils, based on their characteristic phenotype of Siglec-F+ and Sideward Scatter (SSC)hi 22

(Fig. 1A&B). On the other hand, the numbers of neutrophils and monocytes were in most cases

increased (Fig. 1A). In the periphery, neutrophil and monocyte numbers were increased in the

spleen and a strong increase of monocytes was observed in blood (Fig. 1A). Eosinophils were also

absent from the peritoneal cavity, which normally serves as a reservoir for eosinophils23, while no

differences were found for macrophages, mast cells and neutrophils at this site (Fig. 1A). As

eosinophils lose their typical high SSC profile when they degranulate, we also stained for other

eosinophil-related markers24;25, such as CCR3 (Fig. 1B), FIRE, CD125 (IL-5Rα), F4/80, Gr1, or

CD16/32 (data not shown). This analysis confirmed that eosinophils are virtually absent in

CD70TG mice, which was also found in a second, independent founder line (data not shown) and

corroborated by May-Grünwald-staining of bone marrow smears (Fig. 1C). This demonstrates that

CD27-mediated immune activation has a particular impact on the myeloid compartment, as it

inhibits the formation of eosinophils, while it increases the numbers of monocytes and neutrophils.

CD70TG mice fail to produce eosinophils during experimental asthma

To test whether eosinophil production could be induced in CD70TG mice, we made use of an

ovalbumin (OVA)-induced asthma model, as this treatment typically enhances eosinophil

production in the bone marrow and induces eosinophilia in blood, lung and spleen6;26-28. Because

this model works most effectively in BALB/c mice, we used CD70TG mice that were backcrossed

on a BALB/c background; importantly, the decrease in eosinophil numbers in CD70TG mice is not

dependent on their genetic background (Fig. 2A&B). WT and CD70TG BALB/c mice were

sensitized (day 0 and 14) with OVA/Alum and intranasally challenged with OVA at day 28, 29 and

30. This treatment provoked a robust influx of eosinophils in the lungs of WT mice, but not of

CD70TG mice, as judged by flow cytometric analysis (Fig. 2C&D). Histology on the lungs of

CD70TG mice confirmed the almost complete absence of eosinophils and showed a strong

reduction in perivascular or peribronchial infiltrates and mucus production compared to WT mice

(Fig. 2E). Analysis of bone marrow, spleen and blood also showed that eosinophil numbers were

systemically increased in OVA challenged WT mice, but not in CD70TG mice (Fig. 2F). Finally,

and in line with the absence of eosinophilia in the lung, we found that CD70TG mice lacked the

4

Chapter 4

68

3.65 3.45

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12.517.522.527.5 WT

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um

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Siglec-F

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BM Spleen BloodBM Spleen Blood

CC

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A.

B.

**

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Blood


1

2

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mb

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es

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0.00

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12.517.522.527.5 WT

CD70TGN

um

ber

of

cell

s (

x1

06 )

C. WT CD70TGWT CD70TG

Figure 1. CD70TG mice lack eosinophils and have increased numbers of neutrophils and monocytes. (A) Absolute numbers of neutrophils (CD11b+, Gr1+, CD115-, F4/80-), monocytes (CD11b+, CD115+, F4/80+) and eosinophils (SSChi, Siglec-F+) in bone marrow (two femurs and two tibiae), spleen and blood and of macrophages (CD11b+, F4/80+, Gr1-), mast cells (c-Kit+, FcεR1+), neutrophils and eosinophils in the peritoneum. (B) The presence of eosinophils was determined by analyzing expression of Siglec-F on SSChi and on CCR3+ cells (percentages are shown). (C) May-Grünwald-Giemsa-staining of bone marrow smears from WT and CD70TG mice. Mean values ± SD for three individual mice per group are shown. Results are representative of three independent experiments. *, p < 0.05, **, p < 0.01.

4


69

C.

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se

OV

A/P

BS

WTCD70TG

*

PBS OVA0

5

10

15

20

Nu

mb

er o

f E

osi

no

ph

ils(x

105 )

WTCD70TG

***

**

Bone marrow Spleen0.0

0.2

0.4

0.6

0.8

1.0

Nu

mb

er

of

Eo

sin

op

hil

s(x

106 )

WTCD70TG

D.

B.

C.

E.

G.

WT CD70TG

HE

PAS

WT CD70TG

HE

PAS

CD70TG

Siglec-F

0.54 8.42

0.48 0.48

PBS OVA

WT

SS

C

CD70TG

Siglec-F

0.54 8.42

0.48 0.48

PBS OVA

WT

SS

C

**

0 3.1 6.3 12.5 250

200

400

600

800

1000

WT PBS

CD70TG PBS

WT OVA

CD70TG OVA


Pen

H(%

of

bas

elin

e)

**

0 3.1 6.3 12.5 250

200

400

600

800

1000

WT PBS

CD70TG PBS

WT OVA

CD70TG OVA


Pen

H(%

of

bas

elin

e)

Siglec-F

2.09

0.25

0.45

0.09

Bone marrow Spleen

WT (BALB/c)

CD70TG (BALB/c)

SS

C

Siglec-F

2.09

0.25

0.45

0.09

Bone marrow Spleen

WT (BALB/c)

CD70TG (BALB/c)

SS

C

A.

F.

Bone marrow

Spleen Blood

*** **

0

5

10

15

Fo

ld i

ncr

ea

se

OV

A/P

BS

WTCD70TG

Bone marrow

Spleen Blood

*** **

0

5

10

15

Fo

ld i

ncr

ea

se

OV

A/P

BS

Bone marrow

Spleen Blood

*** **

0

5

10

15

Fo

ld i

ncr

ea

se

OV

A/P

BS

WTCD70TG

*

PBS OVA0

5

10

15

20

Nu

mb

er o

f E

osi

no

ph

ils(x

105 )

WTCD70TG

*

PBS OVA0

5

10

15

20

Nu

mb

er o

f E

osi

no

ph

ils(x

105 )

*

PBS OVA0

5

10

15

20

Nu

mb

er o

f E

osi

no

ph

ils(x

105 )

WTCD70TGWTCD70TG

***

**


0.2

0.4

0.6

0.8

1.0

Nu

mb

er

of

Eo

sin

op

hil

s(x

106 )

WTCD70TG***

**


0.2

0.4

0.6

0.8

1.0

Nu

mb

er

of

Eo

sin

op

hil

s(x

106 )

WTCD70TGWTCD70TG

D.

B.

Figure 2. CD70 mice fail to produce eosinophils during experimental asthma and lack airway hyperresponsiveness. (A) Representative plots of Siglec-F staining on bone marrow and spleens of WT BALB/c and CD70TG BALB/c mice showing eosinophils as SSChiSiglec-F+ cells (percentages eosinophils of all cells are shown) and (B) absolute numbers of eosinophils in the bone marrow and spleens of these mice. vcvc (C) WT BALB/c and CD70TG BALC/c mice were sensitized and challenged with OVA (OVA) or treated with PBS as control (PBS). Contour plots display Siglec-F expression on eosinophils in the lung gated on CD4- cells (percentages are shown) from PBS and OVA treated WT and CD70TG mice and (D) absolute numbers of eosinophils in the lung were calculated. (continued)

4

Chapter 4

70

induction of airway hyperresponsiveness, as assessed by a challenge with increasing doses of

metacholine (Fig. 2G). These data demonstrate that CD70TG mice not only have strongly reduced

numbers of eosinophils, but are also unable to generate them under strongly stimulating

pathophysiological conditions in vivo.

CD70TG mice lack eosinophil-specific progenitors and are unable to generate eosinophils

from GMPs

To determine if the defect in eosinophil development could be explained at the level of progenitor

cells, the presence of various precursor subsets was examined in the bone marrow of CD70TG and

WT mice. Absolute numbers of CMPs (Lineage-c-Kit+CD16/32intCD34+) and GMPs (Lineage-c-

Kit+CD16/32hiCD34+) was not different between CD70TG and WT mice (Fig. 3A&B; see M&M

for more information on the phenotypic definition of these cells). However, we did observe a

dramatic decrease in the numbers of EoPs (Lineage-c-KitlowCD16/32hiCD34+IL-5Rα+) in the bone

marrow of CD70TG mice (Fig. 3C&D), which indicates a block in the commitment of GMPs to the

eosinophil lineage. This was supported by QPCR analysis of key transcription factors in GMPs and

EoPs, as we found that GATA-1 was not induced in the few EoPs of CD70TG mice compared to

GMPs (Fig. 3E), which is a key event in eosinophil development13. C/EBP expression was also

reduced in EoPs of CD70TG compared to WT mice, while the reverse was true for C/EBP

expression (Fig. 3E). GATA-2 and PU.1 were not differentially expressed in EoPs from CD70TG

mice (data not shown). These data indicate CD70TG mice not only have a strong reduction of

EoPs, but that the remaining cells lack the required transcription factors for eosinophil

development.

To test the capacity of myeloid progenitors from CD70TG mice to generate eosinophils in vitro,

CMPs or GMPs were sorted and cultured for 9 days with stem cell factor (SCF) and IL-5. This

induced the development of eosinophils that contain eosinophilic granules and a characteristic

donut-shaped nucleus, as observed by cytology (Fig. 3F). Flow cytometric analysis confirmed the

development of eosinophils, as these cells were F4/80+, Siglec-F+ and Gr1+ (Fig. 3F). We found

that the formation of eosinophils from CD70TG GMPs was dramatically reduced compared to WT

controls, whereas CMPs from CD70TG mice did not show this defect (Fig. 3G). CD70TG GMPs

(E) Representative HE and PAS staining on tissue slides of lungs from OVA treated WT and CD70TG mice. (F) Eosinophil numbers in PBS and OVA treated mice from both groups were determined in bone marrow, spleen and blood. Fold increase in eosinophil numbers compared to PBS treated mice is shown. (G) Airway hyperresponsiveness to increasing concentrations of metacholine was measured in PBS and OVA treated WT and CD70TG mice. Data presented are from five mice per group for OVA treated animals and three mice per group for PBS treated animals. An identical experiment with comparable results was performed in WT and CD70TG mice on C57BL/6 background. Means ± SD are shown. *, p < 0.05, **, p < 0.01, ***, p < 0.001. * in figure D represents difference between OVA treated WT and CD70TG mice.

4


71

9.04 1.71

IL-5R

c-K

it

WT CD70TG

WT CD70TG

CD34

CD

16/3

212.6

27.3

11.3

28.9

A.

C.

F.

H.Gr1

93

Sig

lec-

F

F4/80

G.

WTCD70TG

**Eosinophil Progenitor

0.0

0.5

1.0

1.5

2.0

2.5

Nu

mb

er o

f c

ell

s (

x1

05)

After 9 days culture with SCF& IL-5:

CD34

CD

16/3

2 GMP

CMP

B.

D.CMP GMP

0

2

4

6

8

Nu

mb

er o

f c

ell

s (

x1

05)

WTCD70TG

*

CMP GMP0

10

20

30

40

50

Nu

mb

er o

f E

os

ino

ph

ils

(x

103

)

WT

CD70TG

CMP GMP0

10

20

30

40

Nu

mb

er o

f c

ell

s (

x1

03) WT

CD70TG

% o

f M

ax

E.

Rat

io G

AT

A-1

/18S

(x1

0-6 )

Rat

io C

/EB

P

/18S

(10

-6)

Rat

io C

/EB

P/

18S

(x1

0-6 )

C/EBPWTCD70TG

GMP EoP0

2

4

6

8GATA-1

*

GMP EoP0

5

10

15C/EBP

*

GMP EoP0

5

10

15

20

25

*

9.049.04 1.711.71

IL-5R

c-K

it

WT CD70TG

WT CD70TG

CD34

CD

16/3

212.6

27.3

12.6

27.3

11.3

28.9

11.3

28.9

A.

C.

F.

H.Gr1

9393

Sig

lec-

F

F4/80

G.

WTCD70TG


0.0

0.5

1.0

1.5

2.0

2.5

Nu

mb

er o

f c

ell

s (

x1

05)

WTCD70TGWTCD70TG


0.0

0.5

1.0

1.5

2.0

2.5

Nu

mb

er o

f c

ell

s (

x1

05)


0.0

0.5

1.0

1.5

2.0

2.5

Nu

mb

er o

f c

ell

s (

x1

05)

After 9 days culture with SCF& IL-5:

CD34

CD

16/3

2 GMP

CMP

B.

D.CMP GMP

0

2

4

6

8

Nu

mb

er o

f c

ell

s (

x1

05)

CMP GMP0

2

4

6

8

Nu

mb

er o

f c

ell

s (

x1

05)

WTCD70TG

*

CMP GMP0

10

20

30

40

50

Nu

mb

er o

f E

os

ino

ph

ils

(x

103

)

WT

CD70TG *

CMP GMP0

10

20

30

40

50

Nu

mb

er o

f E

os

ino

ph

ils

(x

103

)

*

CMP GMP0

10

20

30

40

50

Nu

mb

er o

f E

os

ino

ph

ils

(x

103

)

WT

CD70TG

WT

CD70TG

CMP GMP0

10

20

30

40

Nu

mb

er o

f c

ell

s (

x1

03)

CMP GMP0

10

20

30

40

Nu

mb

er o

f c

ell

s (

x1

03) WT

CD70TGWTCD70TG

% o

f M

ax

E.

Rat

io G

AT

A-1

/18S

(x1

0-6 )

Rat

io C

/EB

P

/18S

(10

-6)

Rat

io C

/EB

P/

18S

(x1

0-6 )

C/EBPWTCD70TG

GMP EoP0

2

4

6

8GATA-1

*WTCD70TGWTCD70TG

GMP EoP0

2

4

6

8GATA-1

*

GMP EoP0

5

10

15C/EBP

*

GMP EoP0

5

10

15C/EBP

*

GMP EoP0

5

10

15

20

25

*

GMP EoP0

5

10

15

20

25

*

Figure 3. CD70TG mice lack EoPs and CD70TG GMPs are unable to generate eosinophils. (A) Representative contour plot of staining for CMPs (CD16/32intCD34+) and GMPs (CD16/32hiCD34+) within the Lin-c-Kit+ compartment in bone marrow of WT and CD70TG mice (percentages are shown) and (B) absolute numbers of CMPs and GMPs present in two femurs and two tibiae per mouse. (C) Representative contour plot of staining for EoPs (c-KitlowIL-5Rα+) within the Lineage-CD34+CD16/32hi compartment of WT and CD70TG mice (percentages are shown) and (D) absolute numbers of EoPs. (E) Amount of mRNA of GATA1, C/EBP and C/EBP relative to 18S in GMPs and EoPs from WT and CD70TG mice. (continued)

4

Chapter 4

72

were not deficient in generating other types of myeloid cells in vitro, as these cells were fully

capable of generating neutrophils and monocytes when GM-CSF was added to the cultures (Fig.

3H). These experiments indicate that differentiation towards the eosinophilic, but not other myeloid

lineages, is specifically inhibited in CD70TG GMPs.

Development of eosinophils from myeloid progenitors is inhibited by IFN

CD70TG mice have an accumulation of T cells in the bone marrow, which are mainly effector

memory-like (CD44+CD62L-) T cells (Fig. 4A) and which are potent producers of the typical Th1

cytokine IFNγ (Fig. 4B). To address the possibility that IFNγ can directly inhibit the development

of eosinophils from myeloid progenitors, we analyzed the expression of the ligand-binding chain of

the IFNγ receptor (IFNγR1) on CMPs and GMPs by flow cytometry. Expression of IFNγR1 could

be detected on both CMPs and GMPs, with the highest expression on GMPs. Expression of

IFNγR1 was downregulated on CMPs and GMPs of CD70TG mice (Fig. 4C&D), indicative of

recent signaling through this receptor29.

To determine if IFNγ can directly block development of eosinophils, we analyzed the effect of

IFNγ on the outgrowth of eosinophils from purified WT progenitors. Indeed, the addition of IFNγ

to WT CMPs (Fig. 4E) as well as GMPs (data not shown) severely reduced their ability to generate

eosinophils in a dose-dependent manner. This was not due to a generally inhibitory effect of IFNγ

on the differentiation capacity of these precursors, as neutrophils and monocytes developed

normally when GM-CSF was added to these cultures (Fig. 4F). Together with the observation that

CD70TG GMPs, but not CMPs, are impaired in generating eosinophils in vitro, we postulate that at

least GMPs are sensitive to the inhibitive effect of IFN on eosinophil differentiation; the observed

impact of IFN on WT CMPs is probably due to the fact that CMPs first give rise to GMPs before

differentiating to eosinophils in these cultures.

The strongly reduced numbers of EoPs, but not GMPs, in CD70TG mice indicates that IFNγ

specifically blocks the differentiation of GMPs to this downstream eosinophil-specific progenitor.

To test this hypothesis, WT GMPs were cultured with SCF, IL-3, IL-5 and GM-CSF with or

without IFNγ. After 3 days in the absence of IFN, we found expression of IL-5R on a small

fraction of the undifferentiated cells, which were phenotypically similar to the EoP (Fig. 4G).

(F) When CMPs or GMPs (left) are sorted and cultured for 9 days with SCF and IL-5, they develop into a near homogeneous population of eosinophils, as determined by May-Grunwald Giemsa-staining on cytospins (middle) and flow cytometric analysis for expression of Siglec-F, F4/80 and Gr1 (right). Filled histogram shows fluorescence minus one control staining. (G) Absolute number of eosinophils derived from 1.0x104 CMPs or GMPs from WT and CD70TG mice cultured for 9 days with SCF and IL-5. (H) Absolute number of cells derived from 2.5x103

CMPs or GMPs from WT and CD70TG mice cultured for 9 days with SCF, IL-5 and GM-CSF. Mean values ± SD for three individual mice per group are shown in B and D and for 2-3 mice in E. Mean values ± SD for two cultures per condition are shown in G and H. Data in B, D and G are representative of three and data in H of two independent experiments. *, p < 0.05, **, p < 0.01.

4


73

CMP GMP0

200

400

600

800

1000WT

CD70TG

IFNR

1 (G

eoM

FI)

CD

34

0 20 1000

10

20

30

40

IFN concentration (ng/ml)

Nu

mb

er o

f ce

lls (

x103 )

IL-5Rα

- IFN + IFN

29.7

70.3

14.1

85.9

74.1

25.8

74.7

25.1

IFN

CD4+ CD8+

WT

CD70TG

WT

CD70TG

CMP GMP

IFNR1

WT

CD70TG

% o

f max

A.

E.

B.

C.

G.

0 800 40000

50

100

150

IFN concentration (U/ml)

Exp

ansi

on

(%

of

con

tro

l)

0 800 40000

50

100

150


Exp

ansi

on

(%

of

con

tro

l)***

***

K.J.I.

0.64±0.05 0.02±0.01

0 20 1000

5

10

15


Nu

mb

er

of

Eo

sin

op

hils

(x1

03 )

*

* **

*** ** * *

**

**

Total EM CM Naive0.0

0.5

1.0

WT

CD70TG

Nu

mb

er

of

T c

ells

(x

10

6 )

Day 5 Day 18 Day 18

D.

H.

0 800 40000

10

20

30

40

IFN concentration (U/ml)Dif

fere

nia

ted

eosi

no

ph

ils

(%)

IL-5Rα

% o

f m

ax

IL-5

IL-5+IFNγ

F.CMP GMP

0

200

400

600

800

1000WT

CD70TG

IFNR

1 (G

eoM

FI)

CD

34C

D34

0 20 1000

10

20

30

40


Nu

mb

er o

f ce

lls (

x103 )

IL-5Rα

- IFN + IFN

29.7

70.3

14.1

85.9

74.1

25.8

74.7

25.1

IFN

CD4+ CD8+

WT

CD70TG

29.7

70.3

14.1

85.9

74.1

25.8

74.7

25.1

IFN

CD4+ CD8+

WT

CD70TG

WT

CD70TG

WT

CD70TG

CMP GMP

IFNR1

WT

CD70TG

% o

f max

A.

E.

B.

C.

G.

0 800 40000

50

100

150


Exp

ansi

on

(%

of

con

tro

l)

0 800 40000

50

100

150


Exp

ansi

on

(%

of

con

tro

l)***

***

K.J.I.

0.64±0.05 0.02±0.01

0 20 1000

5

10

15


Nu

mb

er

of

Eo

sin

op

hils

(x1

03 )

*

* **

0 20 1000

5

10

15


Nu

mb

er

of

Eo

sin

op

hils

(x1

03 )

*

* **

*** ** * ** *

**

**


0.5

1.0

WT

CD70TG

Nu

mb

er

of

T c

ells

(x

10

6 )

**

**


0.5

1.0

WT

CD70TG

Nu

mb

er

of

T c

ells

(x

10

6 )

Day 5 Day 18 Day 18

D.

H.

0 800 40000

10

20

30

40


fere

nia

ted

eosi

no

ph

ils

(%)

0 800 40000

10

20

30

40


fere

nia

ted

eosi

no

ph

ils

(%)

IL-5Rα

% o

f m

ax

IL-5

IL-5+IFNγ

F.

Figure 4. Development of eosinophils from myeloid progenitors is inhibited by IFNγ. (A) Absolute numbers of total T cells and T cell subsets in the bone marrow (one femur and one tibia) of WT and CD70TG mice (EM: effector memory, CM: central memory). (B) Intracellular expression of IFNγ in CD3+CD4+ gated and CD3+CD8+ gated T cells from bone marrow of WT and CD70TG mice upon PMA and ionomycin stimulation (percentages are shown). (C) Expression of IFNγR1 on CMPs and GMPs of WT (thin line) and CD70TG mice (bold line). Filled graph shows fluorescence-minus-one control staining. (D) Expression of IFNγR1 on CMPs and GMPs as the average geometric mean fluorescence intensity (GeoMFI). (continued)

4

Chapter 4

74

Strikingly, the addition of IFN inhibited the appearance of IL-5R+ cells in these cultures and

reduced the fraction of EoP-like cells by 95% (Fig. 4G). These observations indicate that IFNγ can

inhibit the outgrowth of eosinophils from myeloid progenitor cells by preventing the development

of GMPs to EoPs. To test whether IFN can directly affect the expression of the IL-5R, we cultured

the IL-5 responsive erythroleukemic cell line TF-1 with IL-5 in the presence or absence of IFN.

We found that IFN indeed inhibited the surface expression of the IL-5R by 30% (Fig. 4H), but

not the mRNA expression (data no shown). In accordance, this treatment inhibited the IL-5

dependent proliferation of these cells, but not the GM-CSF dependent proliferation (data not

shown).

Finally, to determine whether IFN could also inhibit eosinophil development from human

hematopoietic progenitors, cord blood derived CD34+ cells were differentiated to eosinophils in the

absence or presence of IFN. Cells were expanded for 3 days in the presence of SCF, FLT-3, GM-

CSF, IL-3 and IL-5 and subsequently cultured for another 15 days in the presence of IL-3 and IL-5

alone to induce eosinophil differentiation30. Expansion of CD34+ progenitor cells during the first

period of culture was not affected by the presence of IFNγ (Fig 4I). However, after 18 days of

culture, both the cellular expansion (Fig. 4J) and the eosinophil differentiation (Fig. 4K) were

strongly reduced by IFNγ. This demonstrates that IFNγ does not inhibit the expansion of early

hematopoietic stem and progenitor cells, but rather the differentiation and expansion of more

restricted eosinophil precursors.

Reduced eosinophil development during CD70-mediated immune activation is IFN-

dependent

Thus far, we have shown that CD70TG mice are almost devoid of eosinophils and that IFN is

sufficient to impair eosinophil development in vitro. To establish whether the block in eosinophil

(E) Number of eosinophils generated from 5.0x103 WT CMPs after 9 days of culture with SCF and IL-5 with or without addition of various concentrations of IFNγ. (F) Number of cells generated from 2.5x103 WT CMPs after 9 days of culture with SCF, IL-5 and GM-CSF with or without addition of various concentrations of IFNγ. (G) Representative plot of staining for CD34+ IL-5Rα+ EoPs gated on the Lineage-CD16/32+c-Kitlow compartment after a 3 day culture of GMPs in the presence of SCF, IL-3, IL-5 and GM-CSF with or without the addition of IFNγ (data indicate percentage of EoPs from total cells) (H) Flow cytometric analysis of IL-5Rα expression on TF1 cells cultured with IL-5 in the presence (thick line) or absence (thin line) of IFNγ. Isotype control is shown in shaded grey. (I-K) Human CD34+ progenitor cells were cultured for 3 days in the presence of SCF, FLT-3, GM-CSF, IL-3 and IL-5 followed by culture for 15 days in IL-3 and IL-5. Expansion of progenitor cells at day 5 (I) and day 18 (J) in cultures in the presence of IFNγ, expressed as percentage of expansion in the cultures without IFNγ. (K) Percentage of differentiated eosinophils after 18 days of culture analyzed after May-Grünwald Giemsa staining of cytospins. Data presented in A and D represents the mean of three mice per group. Mean values ± SD for two cultures per condition are shown in E and F and for 4 cultures per condition in H. Data in A-H are representative of at least two independent experiments. Data presented in I, J and K represent the mean of three independent experiments from different donors. Means ± SD are shown. *, p < 0.05, **, p < 0.01, ***, p < 0.001.

4


75

differentiation in CD70TG mice is a direct consequence of the increased IFN production, we

backcrossed CD70TG mice on an IFNγ-deficient background. Indeed, eosinophil differentiation

was largely restored in CD70TGxIFNγ-/- mice, as absolute eosinophil numbers in bone marrow and

spleen were comparable to WT and IFNγ-/- mice (Fig. 5A&B). Although less pronounced than in

CD70TG mice, CD70TGxIFN-/- mice still had an accumulation of effector memory-like T cells in

the bone marrow compared to control mice (Fig. 5C). Yet, the restoration of eosinophil

differentiation in these mice indicates that the inhibition in eosinophil formation in CD70TG mice

can be attributed to the enhanced production of IFNγ.

IFNγ producing T cells in the bone marrow can block eosinophil development

To determine if IFNγ derived from T cells is sufficient to induce the observed block in eosinophil

development, we used a previously described adoptive transfer model of T cells to CD70TG mice

that are deficient for CD27. CD27-/- mice have normal myeloid differentiation and since CD27 and

CD70 are a unique receptor-ligand pair, the strong phenotype of CD70TG mice is completely

nullified when backcrossed on a CD27-/- background15. Transfer of T cells to these

CD70TGxIFN-/-

3.58

0.49

4.52

4.3

0.68

0.1

1.04

1.45

Siglec-F

SS

C

BM Spleen

WT

CD70TG

IFN-/-

A. B.

C.

**

Bone marrow Spleen0

1

2

3

4WTCD70TG

IFN-/-

Nu

mb

er o

f e

osi

no

ph

ils

(x10

6 )

CD70TGxIFN-/-

***

* ***

* ****

*

*

*WTCD70TG

IFN-/-CD70TGxIFN-/-


0.2

0.4

0.6

0.8

1.0

Nu

mb

er o

f T

cel

ls (

x106

)

CD70TGxIFN-/-

3.58

0.49

4.52

4.3

0.68

0.1

1.04

1.45

Siglec-F

SS

C

BM Spleen

WT

CD70TG

IFN-/-

3.58

0.49

4.52

4.3

0.68

0.1

1.04

1.45

Siglec-F

SS

C

BM Spleen

WT

CD70TG

IFN-/-

A. B.

C.

**

Bone marrow Spleen0

1

2

3

4WTCD70TG

IFN-/-

Nu

mb

er o

f e

osi

no

ph

ils

(x10

6 )

CD70TGxIFN-/-

**

Bone marrow Spleen0

1

2

3

4WTCD70TG

IFN-/-

Nu

mb

er o

f e

osi

no

ph

ils

(x10

6 )

CD70TGxIFN-/-

***

* ***

* ****

*

*

*WTCD70TG

IFN-/-CD70TGxIFN-/-


0.2

0.4

0.6

0.8

1.0

Nu

mb

er o

f T

cel

ls (

x106

) ***

* ***

* ****

*

*

*WTCD70TG

IFN-/-CD70TGxIFN-/-

WTCD70TG

IFN-/-CD70TGxIFN-/-


0.2

0.4

0.6

0.8

1.0

Nu

mb

er o

f T

cel

ls (

x106

)

Figure 5. Reduced eosinophil development during CD70-mediated immune activation is IFNγ-dependent. (A) Representative plots of Siglec-F staining on bone marrow and spleens of WT, CD70TG, IFNγ-/- and CD70TGx IFNγ-/- mice showing eosinophils as SSChiSiglec-F+ cells (percentages are shown) and (B) absolute numbers of eosinophils in the bone marrow (two femurs and two tibiae) and spleens of these mice. (C) Absolute numbers of total T cells and T cell subsets (percentages are shown) in the bone marrow of WT, CD70TG, IFNγ-/- and CD70TGx IFNγ-/- mice (EM: effector memory, CM: central memory). Data presented in B and C represents the mean of three individual mice per group. Means ± SD are shown. Experiments were performed at least three times with similar results. *, p < 0.05, **, p < 0.01, ***, p < 0.001.

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CD70TGxCD27-/- mice is sufficient to induce activation of the donor T cells and results in CD27-

dependent, IFN-mediated depletion of B cells in the bone marrow. To assess whether this

treatment also affects eosinophil formation, we transferred T cells to CD70TGxCD27-/- mice (Fig.

6A), which differentiated to effector cells and accumulated in the bone marrow (Fig. 6B). Although

CD70TGxCD27-/- mice have normal numbers of eosinophils, the transfer of WT T cells is

sufficient to significantly inhibit eosinophil differentiation in the bone marrow within a period of

three weeks (Fig. 6C&D). These effects could be attributed to CD70-mediated activation of the

donor T cells, as transfer of WT T cells to CD27-/- mice did not lead to effector T cell formation

(Fig. 6B), nor did it inhibit eosinophil development (Fig. 6C&D). Importantly, the observed block

in eosinophil differentiation in CD70TGxCD27-/- recipient mice was fully dependent on the

production of IFNγ by the donor T cells, since eosinophil formation was not affected when IFNγ-/-

T cells were transferred (Fig. 6C&D). Neutrophil numbers did not change in the bone marrow of

CD70TGxCD27-/- mice receiving WT T cells, while the number of monocytes was slightly

increased (data not shown). These results demonstrate that T cell-derived IFNγ is sufficient to

specifically block the development of eosinophils in vivo.

IFN produced during immune activation in WT mice also suppresses eosinophil

development

To test whether IFN can also suppress development of eosinophils in vivo in a non-transgenic

setting, we treated WT and IFNγ-/- mice with an agonistic anti-CD40 antibody, which is frequently

used to boost immunizations and increases the number of IFNγ-producing T cells31;32. Indeed, we

found that this treatment upregulated the expression of IFN-responsive molecules such as MHC-II

and Sca-1 on a large proportion of bone marrow cells in WT mice, but not IFN-/- mice (data not

shown and 3;15). Importantly, anti-CD40-treated WT mice displayed a strong decrease in eosinophil

numbers in both bone marrow and blood, whereas IFNγ-/- mice were not affected (Fig. 7A-C). In

addition, EoP numbers were also decreased in an IFN-dependent manner (Fig. 7D), indicating that

IFN also impaired development from bone marrow progenitors in this model.

Finally, to test whether IFNγ produced in response to a natural infection also inhibits eosinophil

development, we infected WT and IFNγ-/- mice with lymphocytic choriomeningitis virus (LCMV)

Armstrong. Analysis 8 days after infection revealed that eosinophil numbers were strongly reduced

in peripheral blood and to a lesser extent also in bone marrow of WT mice (Fig. 7E-G). In contrast,

eosinophils were not changed in peripheral blood and even increased in the bone marrow of IFNγ-/-

mice. These data demonstrate that IFNγ also negatively affects the development of eosinophils

during the course of an anti-viral immune response.

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77

Discussion

In this report we show that T cell activation and in particular the ensuing IFN production, can

inhibit eosinophil differentiation in vivo and thereby modulate myelopoiesis in a very specific

manner. Impaired eosinophil production was found in two independent B cell-specific as well as a

T cell-specific CD70TG founder line, and in either case dependent on CD27 and IFN (data not

A.

C.

0.26

3.11

3.38

2.81

2.96

CD27-/- CD70TGxCD27-/-

-

WT

IFN-/-

Donor T cells:

Siglec-F

SS

C

Recipient mice:

- WT - WT IFN-/-

* ****

0.0

0.5

1.0

1.5

2.0

Donor T cells:

Recipient mice:

Nu

mb

er o

f E

osi

no

ph

ils

(x10

6)


CD

44

IFN-/- WT

CD62L

12 10.2

77.6

14.6 11.4

73.7

WT donor T cells 3 wks after transfer to recipient:

13.7 16.2

69.7

CD62L

CD27-/-79.7 10.7

7.0

CD70TGxCD27-/-

CD

44

B.

Analyse recipients 3 weeks after T cell transfer

IFN-/- T cells

CD27-/-

CD70TGxCD27-/-

WT T cells

D.

A.

C.

0.26

3.11

3.38

2.81

2.96


-

WT

IFN-/-

Donor T cells:

Siglec-F

SS

C

Recipient mice:

- WT - WT IFN-/-

* ****

0.0

0.5

1.0

1.5

2.0

Donor T cells:

Recipient mice:

Nu

mb

er o

f E

osi

no

ph

ils

(x10

6)


- WT - WT IFN-/-

* ****

0.0

0.5

1.0

1.5

2.0

Donor T cells:

Recipient mice:

Nu

mb

er o

f E

osi

no

ph

ils

(x10

6)


CD

44

IFN-/- WT

CD62L

12 10.2

77.6

14.6 11.4

73.7


13.7 16.2

69.7

CD62L

CD27-/-79.7 10.7

7.0

CD70TGxCD27-/-

CD

44

CD

44

IFN-/- WT

CD62L

12 10.2

77.6

14.6 11.4

73.7CD

44

IFN-/- WT

CD62L

12 10.2

77.6

14.6 11.4

73.7


13.7 16.2

69.7

CD62L

CD27-/-79.7 10.7

7.0

CD70TGxCD27-/-

CD

44

B.


IFN-/- T cells

CD27-/-

CD70TGxCD27-/-

WT T cells


IFN-/- T cells

CD27-/-

CD70TGxCD27-/-

WT T cells

D.

Figure 6. T cell-derived IFNγ is sufficient to block differentiation of eosinophils. (A) Schematic overview of the experiment. (B) Distribution of subsets of donor T cells derived from WT (CD45.1+) and IFNγ-/- mice (upper panel) and phenotype of transferred CD45.1+ WT T cells in the bone marrow of CD27-/- and CD70TGxCD27-/- host mice (lower panel; gated on CD45.1+ cells). (CD44+CD62L-: effector memory, CD44+CD62L+: central memory, CD44-

CD62L+: naïve) (C) Representative plots of Siglec-F staining of bone marrow of CD27-/- and CD70TGxCD27-/- host mice injected with PBS, WT or IFNγ-/- T cells (9x106 cells/mouse) showing eosinophils as SSChiSiglec-F+ cells (percentages are shown). (D) Absolute numbers of eosinophils in bone marrow (one femur and one tibia) of host mice after injection with PBS or WT or IFNγ-/- T cells. B, C and D represents data of three and five individual mice per group, respectively and are representative of two independent experiments. Means ± SD are shown. *, p < 0.05, **, p < 0.01.

4

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shown). These findings demonstrate that the block in eosinophil differentiation in CD70TG mice is

a direct result of enhanced IFN production in these mice and not due to an intrinsic impairment in

eosinophils or their precursors, nor to enhanced CD27 triggering on early hematopoietic progenitor

F.E. G.

4.3 0.4

4.2 4.6

Siglec-F

SS

C

WT

IFN-/-

Control LCMV

C. D.

A. B.

IFNγ-/- 3.84.6

4.2 0.6

Siglec-F

SS

C

WT

Control IgG αCD40

**

***Control LCMV Control LCMV

0.0

0.2

0.4

0.6

0.8

WT IFN-/-

Nu

mb

ero

f eo

sin

op

hil

s(x

106

)/m

l Blood

**

**

0

1

2

3

Nu

mb

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f eo

sin

op

hil

s(x

106

) BM

Control LCMV Control LCMV

WT IFN-/-

***IgG CD40 IgG CD40

0.0

0.5

1.0

1.5

WT

Nu

mb

er o

f eo

sin

op

hil

s(x

106

)/m

l

IFN-/-

Blood

IgG CD40 IgG CD400.0

0.5

1.0

1.5

2.0

2.5

WT IFN-/-

Nu

mb

er o

f eo

sin

op

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s(x

106

)

BM

*


0.005

0.010

0.015

WT IFN-/-

Nu

mb

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f E

oP

s(x

106

) BM EoPs

F.E. G.

4.3 0.4

4.2 4.6

Siglec-F

SS

C

WT

IFN-/-

Control LCMV

4.34.3 0.40.4

4.24.2 4.64.6

Siglec-F

SS

C

WT

IFN-/-

Control LCMV

C. D.

A. B.

IFNγ-/- 3.84.6

4.2 0.6

Siglec-F

SS

C

WT

Control IgG αCD40

IFNγ-/- 3.84.6

4.2 0.6

Siglec-F

SS

C

WT

Control IgG αCD40

**

***Control LCMV Control LCMV

0.0

0.2

0.4

0.6

0.8

WT IFN-/-

Nu

mb

ero

f eo

sin

op

hil

s(x

106

)/m

l Blood

**

**

0

1

2

3

Nu

mb

ero

f eo

sin

op

hil

s(x

106

) BM


WT IFN-/-

**

**

0

1

2

3

Nu

mb

ero

f eo

sin

op

hil

s(x

106

) BM


WT IFN-/-


0.0

0.5

1.0

1.5

WT

Nu

mb

er o

f eo

sin

op

hil

s(x

106

)/m

l

IFN-/-

Blood


0.0

0.5

1.0

1.5

WT

Nu

mb

er o

f eo

sin

op

hil

s(x

106

)/m

l

IFN-/-

Blood


0.5

1.0

1.5

2.0

2.5

WT IFN-/-

Nu

mb

er o

f eo

sin

op

hil

s(x

106

)

BM

*


0.005

0.010

0.015

WT IFN-/-

Nu

mb

er o

f E

oP

s(x

106

) BM EoPs

*


0.005

0.010

0.015

WT IFN-/-

Nu

mb

er o

f E

oP

s(x

106

) BM EoPs

Figure 7. IFNγ produced upon immune activation in WT mice suppresses eosinophil development. (A) Representative plots of Siglec-F staining on blood of WT and IFNγ-/- mice, showing eosinophils as SSChiSiglec-F+ cells (percentages are shown) 4 days after the first injection of αCD40 or control IgG. Numbers of eosinophils in (B) blood and (C) bone marrow (total from 2 femurs and 2 tibiae). (D) Numbers of EoPs in the bone marrow of these mice. (E) Representative plots of Siglec-F staining on blood of WT and IFNγ-/- mice showing eosinophils as SSChiSiglec-F+ cells (percentages are shown) 8 days after infection with LCMV-Armstrong. Numbers of eosinophils in (F) blood and (G) bone marrow (total from 1 femur and 1 tibia). Data in A-D are from five mice per group and in E-G 5-6 mice per infected group and 3-4 mice per control group. Both experiments have been performed twice with comparable outcome. Means ± SEM are shown. *, p < 0.05, **, p < 0.01, ***, p < 0.001.

4


79

cells in these mice3. Serum levels of IFN are below detection limit in CD70TG mice, though these

mice express high levels of MHC-II and Sca-1 and gradually lose their B cells, which are known

consequences of IFN triggering and indeed do not occur in CD70TG*IFN-/- mice15. Moreover,

our in vitro experiments in which IFN is added to purified progenitor cells demonstrate that IFN

can directly inhibit the development of both murine and human eosinophils (Fig. 4). This is

corroborated by the fact that IFN induced by anti-CD40 treatment or LCMV infection strongly

decreased eosinophil numbers in WT mice (Fig. 7), but also in CD27-/- mice (data not shown),

which demonstrates that CD27-triggering is sufficient, but not required for IFN-mediated

inhibition of eosinophil development.

The eosinophil defect in CD70TG mice could be traced back to a strong decrease in the number of

IL-5Rα+ eosinophil progenitors in the bone marrow. Moreover, IFN could inhibit formation of

these EoPs from GMPs and downregulate IL-5R expression on TF-1 cells, which could indicate

that the eosinophil defect is due to a direct effect of IFNon the expression of the IL-5R. However,

IL-5Rα expression is not a cause, but rather a consequence of commitment of myeloid progenitors

to the eosinophil lineage through the expression of GATA-1 and GATA-213. We observed that the

remaining EoPs from CD70TG mice lacked GATA-1 expression, indicating that IFN actually

blocks the commitment of GMPs to the eosinophilic lineage by inhibiting the induction of GATA-

1. This will make these cells unable to upregulate essential genes, such as the IL-5R, to respond to

environmental cues and differentiate to eosinophils. We did not observe induction of apoptosis of

GMPs by IFNγ, but it is most likely that IFN will reduce the viability of such precursors when

they do not receive the appropriate signals to differentiate further. The finding that addition of GM-

CSF to IFN-treated GMPs enabled normal differentiation to monocytes and neutrophils supports

this view (Fig. 4F). Furthermore, the paradoxical finding that CD70TG CMPs were able to

generate eosinophils, but WT CMPs incubated with IFN were not, is most likely due to the fact

that GMPs derived from CD70TG CMPs were not exposed to IFN in these cultures, while CMP-

derived WT GMPs were. GMPs are probably also more sensitive to IFN, as they have higher

expression of the IFNR (Fig.4C). Whether IFN does indeed inhibit the upregulation and/or

function of GATA-1 in GMPs and by what molecular mechanism is currently under investigation.

An important question is why the production of IFN inhibits eosinophil formation. Considering

the fact that CD70TG mice not only have strongly reduced eosinophil numbers, but also increased

monocyte numbers, we speculate that IFN inhibits eosinophil development at the benefit of other

myeloid cells. This would be highly relevant during viral infections, where monocytes rather than

eosinophils are required to combat the infection. Indeed, we found that IFN produced during

LCMV-infection inhibits the numbers of eosinophils in bone marrow and blood (Fig. 7). The

increase in eosinophils in bone marrow of LCMV-infected IFN-/- mice is probably the result of a

4

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more general increase in myelopoiesis induced by the infection, in which eosinophil formation is

not inhibited due to the absence of IFN. This is supported by the finding that large numbers of

eosinophils accumulate in the lungs of IFNR-deficient mice upon infection with respiratory

syncytial virus33 and in the brains of IFN-deficient mice upon infection with Borna disease virus34,

but not in WT controls. Since virus-specific CD8 T cells migrate to the bone marrow during the

course of a viral infection with LCMV in mice35, SIV in monkeys36 and HIV in humans37, we

postulate that these cells actively inhibit the local development of eosinophils through the

production of IFN and thereby balance myeloid output from the bone marrow. Furthermore,

studies in which mice were treated with IL-128 or challenged with MCMV7 prior to induction of an

allergic airway response, showed that Th1-induction reduced the ensuing airway eosinophilia. IL-

12 treatment also prevented systemic eosinophilia induced either by challenging sensitized mice

with an allergen38 or by infecting mice intraperitoneally with a parasite39, which is in both cases

IFN-dependent. All together, these data support the hypothesis that IFN-producing T cells that

develop during anti-viral responses can actively prevent the development of those myeloid cells

that are not required and possibly even counterproductive to eradicate the virus.

IL-5 plays a crucial role in the development of eosinophils, as IL-5 overexpression is sufficient to

increase eosinophil formation in the bone marrow and induce eosinophilia in blood40;41.

Conversely, IL-5 deficiency decreases eosinophil production in the bone marrow and prevents

induction of eosinophilia upon allergen challenge or parasite infection42;43. Importantly, CD70TG

mice have no defect in IL-5 production and even have more serum IL-5 (62.8 ± 14.3 pg/ml; n=3)

than WT controls (<10 pg/ml; n=3). Thus, our data demonstrate for the first time that IFNγ can,

even in the presence of sufficient levels of IL-5, directly inhibit eosinophil development, both in

vitro and in vivo. A regulating role for IFN rather than IL-5 in the development of eosinophilia

and asthma is supported by studies in humans. Smart et al. showed that both patients with ongoing,

severe asthma and patients with resolved asthma had increased house dust mite-induced IL-5

production. Whereas IFNγ responses were decreased in patients with persistent asthma, IFNγ

production was back to normal in subjects with resolved disease, demonstrating that normalization

of IFN responses was associated with resolution of asthma44. This finding is supported by other

studies demonstrating decreased IFN production in response to rhinovirus or house dust mite and

an inverse correlation between IFN levels and asthma severity45-47. In addition, a genetic

functional polymorphism in the promoter of suppressor of cytokine signaling 1 (SOCS1), a

negative regulator of IFNγ signaling, resulting in increased levels of SOCS1 protein, was

associated with adult asthma48. Mice deficient in SOCS1 tend to have reduced numbers of

eosinophils in the blood, which is restored to normal in SOCS1-deficient mice on an IFNγ-/-

background49. Collectively, these studies corroborate our findings that increased IFN signaling is

negatively associated with asthma development and severity by affecting eosinophil formation.

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81

Taken together, we have demonstrated that IFN is a potent inhibitive cytokine in eosinophil

differentiation, even in the presence of the eosinophil-permissive cytokine IL-5. As such, IFNγ-

producing T cells are important modulators of myeloid output of the bone marrow, as they

specifically limit the generation of eosinophils by inhibiting the commitment of myeloid precursor

cells to the eosinophil lineage.

Acknowledgements

We thank Sten Libregts, Cláudia Brandão Silva and Ronald van Olffen for technical assistance,

Berend Hooibrink for cell sorting and the staff of the animal facility of the AMC for excellent

animal care. We thank Prof. Dr. Rene van Lier for stimulating discussions and Drs Esther Nolte-‘t

Hoen, Kris Reedquist, Christian Geest and Prof. Dr. Rene van Lier for critical reading of the

manuscript. This work was funded by a VENI (MB; 91676137) and a VIDI grant (MAN;

91776310) from The Netherlands Organization of Scientific Research.


Mice

WT, IFN-/- and CD70TG15 C57BL/6 and BALB/c mice were housed at the animal research

institute of the AMC under specific-pathogen-free conditions. C57BL/6 mice were used for all

experiments, except for the OVA-induced allergic asthma model, in which BALB/c mice were

used. Therefore, CD70TG C57BL/6 mice were backcrossed 10 times to BALB/c. Animal

experiments were approved by the Animal Ethics Committee and performed in accordance with

institutional and national guidelines.


Single cell suspensions were obtained by mincing the organ through 40 μm cell strainers.

Erythrocytes were lysed with an ammonium chloride solution. Purification of CMPs and GMPs

was based on described methods12. Briefly, c-Kit+ cells were enriched using anti-c-Kit microbeads

(Miltenyi Biotec) and MACS MS-columns (Miltenyi Biotec). Enriched cells were incubated with

antibodies for CD34, CD16/32 and c-Kit and progenitors were sorted on a FacsAria (BD

Biosciences). Sca-1 could not be included in our identification of myeloid progenitors, due to a

systemic IFNγ-mediated upregulation of Sca-1 in CD70TG mice3;15. While this implies that the

CMP and GMP fraction in our experiments also contains some Sca-1+ stem/progenitor cells

(~10%), this did not affect our functional analyses, since c-Kit+Sca-1+ progenitors do not contribute

significantly to the outgrowth of eosinophils in our culture system (data not shown).

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For identification of progenitor cells by flow cytometry, cells were stained with a lineage cocktail

of unlabeled or biotin-conjugated antibodies directed against CD4 (GK1.5), CD8α (53-6.7), B220

(RA3-6B2), CD11b (M1/70), Gr1 (RB6-8C5), Ter119 (Ly-76) and IL-7Rα (B12-1). For

identification of eosinophil progenitors anti-CD11b was excluded from the lineage definition. The

antibodies used were CD4-FITC (GK1.5), CD45.1-PE (A-20), CD44-PE (IM7), CD44-Alexa Fluor

700 (IM7), CD3ε-APC (145-2C11), CD4-PE-Cy5.5 (RM4-5), CD62L-APC, CD62L-PE-Cy7

(MEL-14), CD8-APC-Cy7 (53-6.7, Biolegend), IFNγ-APC (XMG1.2), CD34-FITC (RAM34),

F4/80-FITC (BM8), Gr1-FITC, Gr1-PE (RA3-6B2, both from BD Pharmingen), CD115-biotin

(AFS98), Siglec-F-PE (E50-2440, BD Pharmingen), CCR3-Alexa Fluor 647 (BD Pharmingen),

CD16/32-PE-Cy7 (93), CD11b-APC (M1/70), CD11b-Alexa Fluor 750 (M1/70), c-Kit-Alexa Fluor

750 (2B8), unlabeled IL-5Rα (H7, Wako Pure Chemicals) and CD119-biotin (Biolegend). Biotin-

conjugated and unlabeled antibodies were visualized by streptavidin-PE-Cy7, streptavidin-PE,

streptavidin-APC or anti-Rat Alexa Fluor 633 (Invitrogen). Where possible, cells were stained in

the presence of anti-CD16/CD32 block (2.4G2; purified from hybridoma supernatant). Intracellular

cytokine staining was performed as described previously18. All antibodies and secondary reagents

were obtained from eBioscience, unless otherwise specified. Flow cytometry analyses were

performed on FACSCalibur or FACSCanto (BD Biosciences) and data were analyzed using FlowJo

software (Tree Star).

Culture of murine progenitors

Sorted CMPs and GMPs were cultured for 9 days at 37°C in a humidified incubator at 5% CO2 in

96-well plates in IMDM (Lonza) containing 10% FCS at a density of 5.000 or 10.000 cells per

well. For eosinophil differentiation, medium contained IL-5 and stem cell factor (SCF). For

differentiation towards monocytes/macrophages, neutrophils and eosinophils, medium contained

IL-5, GM-CSF and SCF. Differentiation of GMPs towards eosinophil progenitors was assessed

after 3 days of culture in IL-3, IL-5, GM-CSF and SCF. IFNγ was added to cultures when

indicated. Concentrations used were: IL-5 (40 ng/ml), SCF (5 ng/ml), GM-CSF (2 ng/ml), IL-3 (5

ng/ml), IFNγ (20 or 100 ng/ml). All cytokines were obtained from Peprotech.

Culture of human CD34+ and TF-1 cells

Human eosinophils were differentiated from CD34+ cells purified from umbilical cord blood and

analyzed as previously described30. TF-1 cells were cultured with IL-5 (100 pmol/L; Peprotech) in

the presence or absence of IFN (50 ng/ml; Peprotech). After 2 days cells were stained with a PE-

labeled antibody against IL-5R or an isotype control (BD Pharmingen).

4


83


RNA was extracted using Trizol (Invitrogen) and reverse transcribed to cDNA using random

hexamers and Superscript II reverse transcriptase (Roche). Quantitative real-time PCR was

performed in duplicate using Express SYBR GreenER (Invitrogen) on the StepOnePlus RT-PCR

system (Applied Biosystems). Data was normalized using 18S rRNA as a reference gene. Primer

sequences available on request.

Allergic asthma model

WT and CD70TG BALB/c mice were sensitized with i.p. injections of 20 μg OVA with alum (200

μg) at days 0 and 14. Mice were intranasally (i.n.) challenged with 50 μl OVA (2 mg/ml) on days

28, 29 and 30. Mice receiving intraperitoneal (i.p.) injections with alum and i.n. challenge with

PBS were used as controls. On day 31 airway responsiveness to inhaled metacholine was measured

using whole body plethysmography. Responsiveness to aerosolized PBS was used to set a baseline

value, followed by increasing concentrations of aerosolized metacholine. PenH values were

measured after each metacholine challenge.

Adoptive transfer, LCMV infection and anti-CD40 injection.

T cells were obtained from lymph nodes and spleens of WT (CD45.1+) and IFNγ-/- mice by

incubating single cell suspensions with CD4 and CD8 microbeads (Miltenyi Biotec) and sorting by

MACS positive bead selection with LS columns (Miltenyi Biotec). Cells were washed and

resuspended in PBS and 9x106 cells in 200 ul PBS were injected intravenously into CD27-/- and

CD70TGxCD27-/- recipient mice. Purity of isolated T cells was over 95% as determined by flow

cytometric analysis. Mice were sacrificed and analyzed 3 weeks after transfer of T cells. For

LCMV infection, mice were infected i.p. with 1.5 x 105 PFU of LCMV-Armstrong and analyzed 8

days after infection. An agonistic antibody to CD40 (100 μg, clone FGK-45) or a rat control

antibody (100 g, GL113) was injected i.p. at day 1 and 3 and mice were analyzed at day 5.

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maintained by activated bone marrow CD4+ T cells. Blood. 2005;105(4):1484-1491. 2. Di Rosa F, Pabst R. The bone marrow: a nest for migratory memory T cells. Trends

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interactions between CD27 and CD70. Nat.Immunol. 2005;6(4):412-418. 4. Dent AL, Kaplan MH. T cell regulation of hematopoiesis. Front Biosci. 2008;13(6229-6236. 5. Minshall EM, Schleimer R, Cameron L et al. Interleukin-5 expression in the bone marrow of

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7. Wu CA, Puddington L, Whiteley HE et al. Murine cytomegalovirus infection alters Th1/Th2 cytokine expression, decreases airway eosinophilia, and enhances mucus production in allergic airway disease. J.Immunol. 2001;167(5):2798-2807.

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9. Caux C, Moreau I, Saeland S, Banchereau J. Interferon-gamma enhances factor-dependent myeloid proliferation of human CD34+ hematopoietic progenitor cells. Blood. 1992;79(10):2628-2635.




13. Iwasaki H, Mizuno S, Mayfield R et al. Identification of eosinophil lineage-committed progenitors in the murine bone marrow. J.Exp.Med. 2005;201(12):1891-1897.

14. Mori Y, Iwasaki H, Kohno K et al. Identification of the human eosinophil lineage-committed progenitor: revision of phenotypic definition of the human common myeloid progenitor. J.Exp.Med. 2009;206(1):183-193.


16. Keller AM, Schildknecht A, Xiao Y, van den Broek M, Borst J. Expression of costimulatory ligand CD70 on steady-state dendritic cells breaks CD8+ T cell tolerance and permits effective immunity. Immunity. 2008;29(6):934-946.


18. van Gisbergen KP, van Olffen RW, van BJ et al. Protective CD8 T cell memory is impaired during chronic CD70-driven costimulation. J.Immunol. 2009;182(9):5352-5362.


20. Arens R, Schepers K, Nolte MA et al. Tumor rejection induced by CD70-mediated quantitative and qualitative effects on effector CD8+ T cell formation. J.Exp.Med. 2004;199(11):1595-1605.

21. De Colvenaer V, Taveirne S, Hamann J et al. Continuous CD27 triggering in vivo strongly reduces NK cell numbers. Eur.J.Immunol. 2010;40(4):1107-1117.

22. Zhang JQ, Biedermann B, Nitschke L, Crocker PR. The murine inhibitory receptor mSiglec-E is expressed broadly on cells of the innate immune system whereas mSiglec-F is restricted to eosinophils. Eur.J.Immunol. 2004;34(4):1175-1184.


24. Voehringer D, Shinkai K, Locksley RM. Type 2 immunity reflects orchestrated recruitment of cells committed to IL-4 production. Immunity. 2004;20(3):267-277.

25. Voehringer D, van RN, Locksley RM. Eosinophils develop in distinct stages and are recruited to peripheral sites by alternatively activated macrophages. J.Leukoc.Biol. 2007;81(6):1434-1444.

26. Kung TT, Jones H, Adams GK, III et al. Characterization of a murine model of allergic pulmonary inflammation. Int.Arch.Allergy Immunol. 1994;105(1):83-90.

27. Ohkawara Y, Lei XF, Stampfli MR et al. Cytokine and eosinophil responses in the lung, peripheral blood, and bone marrow compartments in a murine model of allergen-induced airways inflammation. Am.J.Respir.Cell Mol.Biol. 1997;16(5):510-520.

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28. Hogan MB, Weissman DN, Hubbs AF et al. Regulation of eosinophilopoiesis in a murine model of asthma. J.Immunol. 2003;171(5):2644-2651.

29. Wietzerbin J, Gaudelet C, Aguet M, Falcoff E. Binding and cross-linking of recombinant mouse interferon-gamma to receptors in mouse leukemic L1210 cells; interferon-gamma internalization and receptor down-regulation. J.Immunol. 1986;136(7):2451-2455.

30. Buitenhuis M, Baltus B, Lammers JW, Coffer PJ, Koenderman L. Signal transducer and activator of transcription 5a (STAT5a) is required for eosinophil differentiation of human cord blood-derived CD34+ cells. Blood. 2003;101(1):134-142.

31. Martin DL, King CL, Pearlman E, Strine E, Heinzel FP. IFN-gamma is necessary but not sufficient for anti-CD40 antibody-mediated inhibition of the Th2 response to Schistosoma mansoni eggs. J.Immunol. 2000;164(2):779-785.

32. Gorbachev AV, Fairchild RL. CD40 Engagement Enhances Antigen-Presenting Langerhans Cell Priming of IFN-{gamma}-Producing CD4+ and CD8+ T Cells Independently of IL-12. J Immunol. 2004;173(4):2443-2452.

33. Boelen A, Kwakkel J, Barends M et al. Effect of lack of Interleukin-4, Interleukin-12, Interleukin-18, or the Interferon-gamma receptor on virus replication, cytokine response, and lung pathology during respiratory syncytial virus infection in mice. J.Med.Virol. 2002;66(4):552-560.

34. Hausmann J, Pagenstecher A, Baur K et al. CD8 T cells require gamma interferon to clear borna disease virus from the brain and prevent immune system-mediated neuronal damage. J.Virol. 2005;79(21):13509-13518.

35. Slifka MK, Whitmire JK, Ahmed R. Bone marrow contains virus-specific cytotoxic T lymphocytes. Blood. 1997;90(5):2103-2108.

36. Kuroda MJ, Schmitz JE, Seth A et al. Simian immunodeficiency virus-specific cytotoxic T lymphocytes and cell-associated viral RNA levels in distinct lymphoid compartments of SIVmac-infected rhesus monkeys. Blood. 2000;96(4):1474-1479.

37. Kulkosky J, Bouhamdan M, Geist A et al. Pathogenesis of HIV-1 infection within bone marrow cells. Leuk.Lymphoma. 2000;37(5-6):497-515.

38. Rais M, Wild JS, Choudhury BK et al. Interleukin-12 inhibits eosinophil differentiation from bone marrow stem cells in an interferon-gamma-dependent manner in a mouse model of asthma. Clin.Exp.Allergy. 2002;32(4):627-632.

39. Finkelman FD, Madden KB, Cheever AW et al. Effects of interleukin 12 on immune responses and host protection in mice infected with intestinal nematode parasites. J Exp.Med. 1994;179(5):1563-1572.

40. Dent LA, Strath M, Mellor AL, Sanderson CJ. Eosinophilia in transgenic mice expressing interleukin 5. J Exp.Med. 1990;172(5):1425-1431.

41. Lee NA, McGarry MP, Larson KA et al. Expression of IL-5 in thymocytes/T cells leads to the development of a massive eosinophilia, extramedullary eosinophilopoiesis, and unique histopathologies. J Immunol. 1997;158(3):1332-1344.

42. Foster PS, Hogan SP, Ramsay AJ, Matthaei KI, Young IG. Interleukin 5 deficiency abolishes eosinophilia, airways hyperreactivity, and lung damage in a mouse asthma model. J Exp.Med. 1996;183(1):195-201.

43. Kopf M, Brombacher F, Hodgkin PD et al. IL-5-deficient mice have a developmental defect in CD5+ B-1 cells and lack eosinophilia but have normal antibody and cytotoxic T cell responses. Immunity. 1996;4(1):15-24.

44. Smart JM, Horak E, Kemp AS, Robertson CF, Tang ML. Polyclonal and allergen-induced cytokine responses in adults with asthma: resolution of asthma is associated with normalization of IFN-gamma responses. J.Allergy Clin.Immunol. 2002;110(3):450-456.

45. Brooks GD, Buchta KA, Swenson CA, Gern JE, Busse WW. Rhinovirus-induced interferon-gamma and airway responsiveness in asthma. Am.J.Respir.Crit Care Med. 2003;168(9):1091-1094.

46. Leonard C, Tormey V, Burke C, Poulter LW. Allergen-induced cytokine production in atopic disease and its relationship to disease severity. Am.J.Respir.Cell Mol.Biol. 1997;17(3):368-375.

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47. Papadopoulos NG, Stanciu LA, Papi A, Holgate ST, Johnston SL. A defective type 1 response to rhinovirus in atopic asthma. Thorax. 2002;57(4):328-332.

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49. Alexander WS, Starr R, Fenner JE et al. SOCS1 is a critical inhibitor of interferon gamma signaling and prevents the potentially fatal neonatal actions of this cytokine. Cell. 1999;98(5):597-608.

4

Chapter 5 Interferon-gamma induces monopoiesis

and inhibits neutrophil development during

inflammation.

Alexander M. de Bruin1, Sten F. Libregts1,2, Marijke Valkhof3, Louis Boon4, Ivo P. Touw3 and

Martijn A. Nolte1,2

1Department of Experimental Immunology, Academical Medical Center, 1105 AZ Amsterdam, The

Netherlands, 2Department of Hematopoiesis, Sanquin Research and Landsteiner Laboratory, 1066

CX Amsterdam, The Netherlands, 3Department of Hematology, Erasmus Medical Center, 3000 CA

Rotterdam, The Netherlands, 4Bioceros BV, 3584 CM Utrecht, The Netherlands.

Blood 2012 Feb 9;119(6):1543-54

Interferon-gamma induces monopoiesis and inhibits neutrophil development during inflammation

89

Abstract

Steady-state hematopoiesis is altered upon infection, but the cellular and molecular mechanisms

driving these changes are largely unknown. Modulation of hematopoiesis is essential to increase

the output of the appropriate type of effector cell required to combat the invading pathogen. Here

we demonstrate that the pro-inflammatory cytokine interferon-gamma (IFN) is involved in

orchestrating inflammation-induced myelopoiesis. Using both mouse models and in vitro assays we

show that IFN induces differentiation of monocytes over neutrophils at the level of myeloid

progenitors. Infection with Lymphocytic Choriomeningitis Virus (LCMV) induces monopoiesis in

WT mice, but this infection rather causes increased neutrophil production in IFN-/- mice. We

demonstrate that IFN enhances expression of monopoiesis-inducing transcription factors IRF8 and

PU.1 in myeloid progenitor cells, whereas it reduces G-CSF-driven neutrophil differentiation via a

SOCS3-dependent inhibition of STAT3 phosphorylation. These results establish a critical role for

IFN in directing monocyte versus neutrophil development during immune activation.

Introduction

Tightly regulated proliferation and differentiation of hematopoietic progenitors in the bone marrow

(BM) is critical for maintaining homeostatic levels of peripheral blood cells and is controlled by

intrinsic and extrinsic factors. Immunological stress conditions, like infections, induce changes in

the magnitude and composition of hematopoietic output, which are essential to meet the increased

demand for immune cells and to induce proper immune defense against invading pathogens1. As

such, it has been demonstrated that infection with the extracellular fungal pathogen Candida

albicans induces peripheral neutrophilia and an increase in BM neutrophils2. Conversely, increased

production of monocytes has shown to be essential in controlling Listeria monocytogenes and

Toxoplasma gondii infections3;4. Yet, the molecular and cellular mechanisms that underlie these

hematopoietic changes in myeloid differentiation remain to be elucidated.

Both neutrophils and monocytes are derived from the granulocyte-macrophage progenitor (GMP)

and differentiation to either of these myeloid cell types is driven by a particular cytokine5;6. In

steady state conditions, G-CSF is crucial for maintaining appropriate neutrophil numbers, as loss of

G-CSF or its receptor decreases the number of circulating neutrophils, whereas injection of G-CSF

increases neutrophil numbers7;8. G-CSFR signaling induces phosphorylation of signal transducer

and activator of transcription (STAT) 3, a member of the STAT family, which are signaling

proteins that regulate gene expression in response to cytokines9. The occurrence of emergency

granulopoiesis has been linked to increased levels of granulopoiesis-supporting cytokines, like G-

CSF, and is dependent on STAT3 phosphorylation10;11. Conversely, production of monocytes is

supported by M-CSF and the transcription factors PU.1 and IRF8, which are essential for monocyte

differentiation (reviewed by Friedman12). As such, the development of monocytosis after Listeria

5

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infection could be driven by the corresponding increase in M-CSF levels in serum10, possibly in

conjunction with TLR-mediated signals13. On the other hand, the increase of monocytes in a burn

sepsis model has been associated with the upregulation of the M-CSF receptor on myeloid

progenitor cells14;15, whereas the occurrence of neutropenia in this model is due to recognition of

LPS by early precursors16.

Evidence is emerging that activated T cells also play an important factor in modulating

hematopoiesis during an immune response. Large numbers of effector T cells are known to enter

the BM parenchyma during viral infections17 and T cells are thought to modulate hematopoietic

progenitors either through direct cell-cell contact18;19 or by the secretion of pro-inflammatory

cytokines such as TNF or IFN20;21. IFN is typically produced in response to infection with

intracellular pathogens and has important stimulatory functions in innate and adaptive immunity22.

Using a chronic inflammatory mouse model in which transgenic expression of CD70 on B cells

(CD70TG mice) induces CD27-driven formation of IFN-producing effector T cells, we have

previously demonstrated that T cell-derived IFN can also suppress the development of B cells23,

erythrocytes24 and eosinophilic granulocytes20. Other reports have also shown lineage-specific

effects of IFN on infection-induced changes in myelopoiesis25-27.

The cellular and molecular mechanism by which IFN differentially affects formation of various

types of myeloid cells is largely unknown. Here we investigated if and how IFN affects the

mechanisms regulating neutrophil and monocyte differentiation at the level of myeloid progenitors.

We report that IFN promotes monopoiesis and suppresses neutrophil production in the BM.

Correspondingly, IFN elevates expression of the monocyte-inducing transcription factors PU.1

and IRF8 in GMPs and reduces G-CSF-mediated phosphorylation of STAT3 in a SOCS3-

dependent manner. These data demonstrate that IFN can directly regulate the balance between

monocyte and neutrophil production by affecting cytokine responses and expression of lineage-

specific transcription factors in GMPs.

Results

CD70TG mice have more monocytes and increased monopoiesis over granulopoiesis

To examine the impact of T cell-driven immune activation on the development of monocytes and

neutrophils, we analyzed the myeloid compartment of CD70TG mice, as these mice have large

numbers of effector T cells due to enhanced costimulation through CD2723. We found that

CD70TG mice contained relatively more monocytes (CD115+Gr1dim) than neutrophils (CD115-

Gr1high) within the fraction of CD11b+ cells in peripheral blood (Fig. 1A). Quantification of these

data revealed that CD70TG mice had a strong increase in the absolute numbers of circulating

monocytes, whereas neutrophil numbers were not different from WT mice (Fig. 1B). This increase

in monocytes was also found in BM, suggesting that monocyte production was increased in

5


91

Figure 1. CD70TG mice have increased numbers of monocytes and increased monopoiesis over granulopoiesis. (A) Representative plots of staining for monocytes (Gr1lowCD115+) and neutrophils (Gr1+CD115-) within the CD11b+ compartment in peripheral blood of WT and CD70TG mice and absolute numbers of monocytes and neutrophils in peripheral blood (B) and BM (C) of WT and CD70TG mice. Data in B and C represent mean ± s.d. from at least 3 mice per group. (continued)

CD115

31.3

31.9

71.1

14.1G

r1WT CD70TG

Neutrophils Monocytes0

2

4

6WTCD70TG

Nu

mb

er o

f ce

lls (

x106

)/m

l

0 12 18 240

5

10

15

20WT

CD70TG

Hours

% B

rdU

+ m

on

oc

yte

s

0 1 2 3 4 50

20

40

60

80WT

CD70TG

Days

% B

rdU

+ n

eu

tro

ph

ils

A

B


10

20

30

40WTCD70TG

Nu

mb

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f ce

lls (

x106 )

WT CD70TG

0.0

0.2

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0.6

0.8

Rat

io M

:N

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0

2

4

6

Rat

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:N

C

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F G H

***

***

**

*****

*** ***

0

50

100

**

*

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0

20

40

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# C

olo

nie

s/ 2

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BM

cel

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GMGM

CD115

31.3

31.9

31.3

31.9

71.1

14.1

71.1

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2

4

6WTCD70TG

Nu

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)/m

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0 12 18 240

5

10

15

20WT

CD70TG

Hours

% B

rdU

+ m

on

oc

yte

s

0 1 2 3 4 50

20

40

60

80WT

CD70TG

Days

% B

rdU

+ n

eu

tro

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ils

A

B


10

20

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40WTCD70TG

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0.0

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0.8

Rat

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:N

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0

2

4

6

Rat

io M

:N

C

D E

F G H

***

***

**

*****

*** ***

0

50

100

**

*

% o

f m

yelo

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lon

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WT CD70TG

0

20

40

60

# C

olo

nie

s/ 2

5000

BM

cel

ls

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GMGM

0

20

40

60

# C

olo

nie

s/ 2

5000

BM

cel

ls

WT CD70TG

GMGM

5

Chapter 5

92

CD70TG mice (Fig. 1C). To test this, WT and CD70TG mice were injected with BrdU and the

appearance of circulating monocytes that had incorporated BrdU was followed over time. We

found more BrdU+ monocytes in blood from CD70TG than WT mice at 12, 18 and 24 hours after

BrdU injection (Fig. 1D). In contrast, the release of BrdU+ neutrophils was not altered (Fig. 1E),

which confirmed that the production of particularly monocytes was increased in CD70TG mice.

Myeloid differentiation was further analyzed by performing colony formation of total BM. These

experiments reveal an increase of monocyte/macrophage colonies in CD70TG mice compared to

WT mice and a small decrease in the contribution of neutrophil colonies (Fig. 1F). Although higher

in number, we found that the monocyte/macrophage colonies in CD70TG mice were smaller in size

and thus contained fewer cells than the corresponding colonies in WT mice (data not shown).

Finally, to better visualize the balance between monocyte and neutrophil output in vivo, we

calculated the ratio between monocytes and neutrophils (M:N ratio), which showed an increase in

M:N ratio in both BM (Fig. 1G) and blood (Fig. 1H) of CD70TG mice. These data demonstrate that

CD27-driven immune activation differentially impacts myelopoiesis, shifting the formation of

myeloid cells towards the monocyte lineage.

IFN induces monopoiesis over granulopoiesis in vivo and in vitro

Since CD70TG mice have increased numbers of IFN-producing T cells in BM20;23, we tested

whether the increased monopoiesis over neutrophil development in CD70TG mice was dependent

on IFN. We found that the increased M:N ratio in CD70TG mice was restored to WT levels when

these mice were backcrossed on an IFN-deficient background, both in BM (Fig. 2A) and blood

(Fig. 2B), demonstrating that IFNγ is indeed responsible for the observed skewing of monocytes

over neutrophils in this model. To test whether T cell-derived IFN is also sufficient for altering the

balance between monocyte and neutrophil formation, we used a previously described adoptive

transfer model of T cells to CD27-deficient CD70TG mice23. CD27-/- and CD70TGxCD27-/- mice

have normal myeloid differentiation, and transfer of T cells into these mice results in CD70-

(D) Percentage of BrdU+ monocytes within the CD11b+ compartment in peripheral blood of WT and CD70TG mice measured 12, 18 and 24 hours after injection of BrdU and (E) percentage of BrdU+ neutrophils in peripheral blood of WT and CD70TG mice measured at indicated days after BrdU injection. Data in D and E represent mean ± s.e.m. from 5 mice per group. (F) Type of colonies derived from total BM from WT and CD70TG mice cultured for 8 days in semi-solid medium. Data are displayed as the absolute number of CFU per 25.000 BM cells (left) or as percentage from total colonies (right). M: monocyte/macrophage, G: granulocyte, GM: mixed monocyte/macrophage and granulocyte. Data represent mean ± s.d. from 6 mice per group. M:N ratios in (G) BM and (H) peripheral blood of WT and CD70TG mice were measured by dividing the number of monocytes by the number of neutrophils. Data in G and H represent mean ± s.d. from at least 3 mice per group. BM represents cell numbers per 2 femurs and 2 tibiae. All experiments were performed at least 2 times with similar results. *, p < 0.05, **, p < 0.01, ***, p < 0.001.

5


93

dependent activation and accumulation of IFN-producing effector T cells in BM20;23. Transfer of

WT T cells to CD70TGxCD27-/- mice increased the M:N ratio, which was completely dependent

on IFN production by these transferred T cells (Fig. 2C). To test the effect of IFNγ on the M:N

0.0

0.2

0.4

0.6

0.8

CD70TG

IFN -/-

CD70TGxIFN -/-

WTR

atio

M:N

0

2

4

6

CD70TG

IFN -/-

CD70TGxIFN -/-

WT

Rat

io M

:N

- WT - WT IFN-/-

0.0

0.2

0.4

0.6

Donor T cells

Host CD27-/- CD70TGxCD27-/-

Rat

io M

:N

Isotype CD40 Isotype CD400.0

0.2

0.4

0.6

WT IFN -/-

Rat

io M

:N

A B

C D

E

******

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20

40

60

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*****

# C

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0

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Total BM

CFU-GMCFU-GCFU-M

0.0

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0.6

0.8

CD70TG

IFN -/-

CD70TGxIFN -/-

WTR

atio

M:N

0

2

4

6

CD70TG

IFN -/-

CD70TGxIFN -/-

WT

Rat

io M

:N

- WT - WT IFN-/-

0.0

0.2

0.4

0.6

Donor T cells

Host CD27-/- CD70TGxCD27-/-

Rat

io M

:N

Isotype CD40 Isotype CD400.0

0.2

0.4

0.6

WT IFN -/-

Rat

io M

:N

A B

C D

E

******

** **

GMPs

Medium IFN0

20

40

60

80**

*****

# C

olo

nie

s/ 2

50 G

MP

s GMPs

Medium IFN0

20

40

60

80**

*****

# C

olo

nie

s/ 2

50 G

MP

s

0

50

100

*

***

% o

f m

yelo

idc

olo

nie

s

Medium IFN0

50

100

*

***

% o

f m

yelo

idc

olo

nie

s

Medium IFN

Total BM

CFU-GMCFU-GCFU-M

CFU-GMCFU-GCFU-M

Figure 2. IFN induces monopoiesis over granulopoiesis in vivo and in vitro. M:N ratios in (A) BM and (B) peripheral blood of WT, CD70TG, IFN-/- and CD70TGxIFN-/- mice were measured by dividing the number of monocytes by the number of neutrophils. Data in A and B represent mean ± s.d. from at least 3 mice per group. (C) M:N ratios in BM of CD27-/- and CD70TGxCD27-

/- control mice and 3 weeks after transfer of WT or IFN-/- T cells. (D) M:N ratios in BM of WT and IFN-/- mice 4 days after the first injection of αCD40 or control IgG. Data in C and D represent mean ± s.d. from 5 mice per group. (E) Type of colonies derived from total WT BM (left; mean ± s.d. from 6 mice per group) or from 250 purified GMPs (right; mean ± s.d. from 4 mice per group) cultured for 8 days in semi-solid medium with or without IFN. M: monocyte/macrophage, G: granulocyte, GM: mixed monocyte/macrophage and granulocyte. All experiments were performed at least 2 times with similar results. *, p < 0.05, **, p < 0.01, ***, p < 0.001.

5

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ratio in a T cell activation model that is not driven by transgenic CD70-expression, we injected WT

and IFN-/- mice with an agonistic anti-CD40 antibody, which increases the number of IFN-

producing T cells20;28. We found that the M:N ratio increased in BM of WT mice injected with anti-

CD40, but not in IFN-/- mice (Fig. 2D). In both experimental settings we observed the same

pattern of M:N ratio in peripheral blood (data not shown). Finally, since both monocytes and

neutrophils are derived from GMPs and because we have previously shown that GMPs express the

IFNreceptor20, we tested if IFN directly affects monocyte vs. neutrophil differentiation from

GMPs. We found that addition of IFN to WT GMPs did indeed increase the formation of

monocyte/macrophage colonies at the cost of neutrophil colonies (Fig. 2E). All together, these

experiments demonstrate that IFN is sufficient to enhance monocyte over neutrophil development

both in vitro and in vivo and that this effect can be mediated at the level of the GMP.

IFN is required to increase monocyte numbers and suppress neutrophilia during LCMV

infection

To test if IFN also modulates the development of monocytes and neutrophils during a viral

infection, WT and IFN-/- mice were infected with LCMV Armstrong. Infection with this virus

results in a type I interferon-dependent leukopenia early after infection, while IFN-producing T

cells can only be found 7-8 days after the onset of infection29. Importantly, it was previously shown

that IFN is not required for a proper adaptive immune response against LCMV or for viral

clearance29. Upon LCMV infection, virus-specific T cells can be found in BM, which produce IFN

upon stimulation with LCMV peptide (Fig. 3A) or PMA/ionomycin (Fig. 3B), indicating that

locally produced IFN could affect myelopoiesis in BM. WT and IFN-/- mice responded similarly

to the leukopenia during the first days after infection, showing a reduction in the numbers of

circulating monocytes, but not neutrophils (Fig. 3C-E), thereby reducing the M:N ratio during the

first days after infection (Fig. 3F). However, at day 8 of the infection, monocyte numbers strongly

increased in peripheral blood of WT mice, but not in IFN-/- mice (Fig. 3D). In contrast, neutrophil

numbers had returned to baseline levels in WT mice at day 8 (Fig. 3D), but were significantly

increased in IFN-/- mice (Fig. 3E). In accordance with these findings in peripheral blood,

monocyte numbers were increased in BM of WT mice 8 days after infection (Fig. 3G), whereas

neutrophil numbers were increased in IFN-/- mice (Fig. 3H), resulting in increased M:N ratio in

BM of WT mice and a decreased M:N ratio in IFN-/- mice (Fig. 3I). Remarkably, the observed

changes in circulating monocyte and neutrophil numbers between WT and IFN-/- mice were found

at day 8, which corresponds with the peak of the anti-viral T cell response (Fig. 3J). IFN

deficiency did not affect the kinetics and magnitude (Fig. 3J), nor the specificity (Fig. 3K) of the T

cell response. In addition, equal numbers of LCMV-specific T cells were found in BM of WT and

IFN-/- mice (Fig. 3L), suggesting that the distinct IFN-dependent effects on myelopoiesis do not

5


95

0 4 6 80

1

2

3

4WT

IFN -/-

Day

Nu

mb

er o

f n

eutr

op

hils

(x1

06)/

ml

0 4 6 80

1

2

3

4WT

IFN -/-

Day

Nu

mb

er o

f m

on

ocy

tes

(x10

6)/

ml

0 4 6 80

1

2

3

4WT

IFN -/-

Day

Rat

io M

:N

Control LCMV Control LCMV0

2

4

6

8

WT IFN-/-

Nu

mb

er o

f m

on

ocy

tes

(x10

6 )


10

20

30

WT IFN-/-

Nu

mb

er o

f n

eutr

op

hils

(x1

06)

Control LCMV Control LCMV0.0

0.1

0.2

0.3

0.4

0.5

WT IFN-/-

Rat

io M

:N

46.6 78.1 63.3 14.5

29.9

42

18.8

70.3

11.8

78

17.3

72

0 4 6 80

5

10

15

20WT

IFN -/-

Day

Nu

mb

er o

f C

D8

T c

ells

(x1

06)/

ml

WT IFN-/-0.00

0.01

0.02

0.03

0.04

Nu

mb

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f T

et+

CD

8 T

cel

ls (

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)

5.49

0.17

3.6 9.7

20 50

WT IFN-/-0

2

4

6

8

% T

et+ o

f C

D8

T c

ells

A

CD115

Gr1


D E F

G H I

J K L

IFN

CD4 CD8

IFN

B

C

WT

IFN-/-

FSC FSC

Control

LCMV

WT

IFN-/-

34.4 16 24.1 63.4

*** *** ***

*** *** ***

***

0 4 6 80

1

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*** *** ***

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Figure 3. IFN is required to increase monocyte numbers and suppress neutrophilia during LCMV infection. Representative plot of intracellular staining for IFN in T cells from BM of WT and IFN-/- mice stimulated in vitro 8 days after LCMV infection with (A) LCMV peptide or with (B) PMA/ionomycin. (C) Representative plot of staining for monocytes (Gr1lowCD115+) and neutrophils (Gr1+CD115-) in peripheral blood before LCMV infection and at the indicated days after infection. Number of (D) monocytes and (E) neutrophils in peripheral blood of WT and IFN-/-

mice at indicated days after LCMV infection and (F) M:N ratios in peripheral blood of these mice. Data in D, E and F represent mean ± s.e.m. from 5 mice per group. Number of (G) monocytes and (H) neutrophils in BM of WT and IFN-/- naïve mice and LCMV infected mice 8 days after infection and (I) M:N ratio in BM of these mice. Data in G, H and I represent mean ± s.d. from 5 mice per group; BM represents cell numbers per 2 femurs and 2 tibiae. (J) Number of CD8 T cells in peripheral blood of WT and IFN-/- mice at indicated days after LCMV infection. (K) Percentage of tetramer binding cells of total CD8 T cells in peripheral blood and (L) number of tetramer binding CD8 T cells in BM of WT and IFN-/- mice 8 days after LCMV infection. Data in J represent mean ± s.e.m. from 5 mice per group and data in K and L represent mean ± s.d. from 5 mice per group. All data shown are representative of at least 2 independent experiments. *, p < 0.05, **, p < 0.01, ***, p < 0.001.

5

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result from an altered T cell response. These experiments demonstrate that IFN induces an

expansion of the monocyte compartment and is required to suppress an increase in neutrophils

during LCMV infection.

IFN directs myelopoiesis during LCMV infection

To investigate whether the IFN-induced changes in monocyte and neutrophil numbers upon

LCMV infection were indeed due to corresponding changes in myelopoiesis, we examined myeloid

progenitors and the production of new progeny. Analysis of GMPs by flow cytometry revealed that

the absolute number of these myeloid-committed progenitors was increased in LCMV-infected

mice, independently of IFN (Fig. 4A). However, BrdU incorporation experiments demonstrated a

strong increase of BrdU+ monocytes in blood (Fig. 4B&C) and BM (Fig. 4D) of WT mice at day 8

of infection, but not in IFN-/- mice. Although very low numbers of circulating BrdU+ neutrophils

could be detected in naïve mice or WT LCMV-infected mice, a strong increase in BrdU+

neutrophils was observed in blood of IFN-/- infected mice (Fig. 4E). Correspondingly, we found a

much stronger increase of BrdU+ neutrophils in BM of IFN-/- than WT mice upon infection (Fig.

4F&G). These data demonstrate that the increase of monocytes in LCMV-infected WT mice and

neutrophils in IFN-/- mice is indeed the direct result of increased production of these cells in BM

and that IFN is able to redirect myelopoiesis towards the monocytic lineage during viral infection.

IFN impairs the proliferation and differentiation of myeloid progenitors in response to G-

CSF

Thus far, we have shown that IFN induces the production of monocytes, while it negatively affects

neutrophil development. Monocyte and neutrophil production from hematopoietic progenitors is

induced by M-CSF and G-CSF, respectively6. To test if IFN directs monocyte and neutrophil

differentiation by affecting cytokine responses, purified common myeloid progenitors (CMPs) and

GMPs were cultured for 3 days in the presence of M-CSF or G-CSF with or without IFN. We

found that IFN slightly increased the outgrowth of CMPs (Fig. 5A) and GMPs (Fig. 5B) in

response to M-CSF, whereas IFN strongly reduced the outgrowth of these cells in response to G-

CSF; these effects could not be explained by corresponding changes in cell death (Fig. S1A&B).

We tested whether IFN impairs the proliferative response to G-CSF by labeling purified

progenitors with CFSE and subsequently culturing these cells as described above. Indeed,

proliferation of CMPs and GMPs in response to G-CSF was significantly reduced by IFN (Fig.

5C-D). In contrast, IFN did not negatively affect, but even slightly enhanced, the proliferative

response to M-CSF (Fig. 5E). Upon differentiation of progenitor cells, expression of c-Kit is

gradually lost and expression of mature lineage markers is increased. Flow cytometric analysis of

cultured CFSE-labeled cells indeed revealed that IFN impaired G-CSF-induced differentiation of

5


97


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18.118.1

44.444.430.630.6

19.619.6

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WT IFNy-/-

Control

LCMV

15.615.6

37.537.5

18.818.8

20.220.2

FSC

Brd

U

WT IFNy-/-

Control

LCMV

* *

***

*

***

***

***

Figure 4. IFN directs myelopoiesis during LCMV infection. (A) Number of GMPs in BM of WT and IFN-/- naïve mice and 8 days after LCMV infection. (B) Representative plot of BrdU staining in monocytes in peripheral blood of naïve and LCMV infected mice 8 days after infection and absolute numbers of BrdU+ monocytes in (C) peripheral blood and (D) BM of these mice. (E) Number of BrdU+ neutrophils in peripheral blood of naïve and LCMV infected mice 8 days after infection. (F) Representative plot of BrdU staining in neutrophils in BM and (G) number of BrdU+ neutrophils in BM of these sasa mice. Data in A, C, D, E represent mean ± s.d. from 5 mice per group; BM represents cell numbers per 2 femurs and 2 tibiae. Experiments were performed at least 2 times with similar results. *, p < 0.05, **, p < 0.01, ***, p < 0.001.

5

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progenitors, as the cells in culture were mainly c-Kit positive and did not express lineage markers

to the same extend as the cells cultured without IFN (Fig. 5F). These experiments demonstrate that

IFN directly inhibits the proliferation and differentiation of myeloid progenitors in response to G-

CSF, while it promotes M-CSF-induced proliferation/differentiation.

IFN induces expression of IRF8 and PU.1

Induction of monopoiesis is mainly controlled by the transcription factors PU.1 and IRF812. IRF8 is

directly inducible by IFN30 and we have previously reported that IFN increases PU.1 expression

in primary progenitor cells in an IRF1-dependent manner24. To test if the observed monopoiesis-

promoting effect of IFN on hematopoietic progenitors was related to increased expression of PU.1

and IRF8, mRNA levels were measured in GMPs from naïve and LCMV-infected mice. Both PU.1

(Fig. 6A) and IRF8 (Fig. 6B) mRNA levels were elevated in WT infected mice, but unaffected in

IFN-/- infected mice. To examine if IFN directly affects the expression of these factors, purified

GMPs were stimulated with IFN and mRNA levels of PU.1 and IRF8 were measured. We found

that IFN increased the expression of both PU.1 (Fig. 6C) and IRF8 (Fig. 6D). These observations

demonstrate that IFN is sufficient to elevate the expression of these monopoiesis-inducing

transcription factors in GMPs both in vitro and in vivo.

IFN reduces G-CSF-mediated STAT3 phosphorylation by a SOCS3-dependent mechanism

The observed IFN-mediated reduction in the responsiveness of progenitors to G-CSF could have

resulted from reduced expression of the G-CSF receptor (G-CSFR). However, IFNγ did not affect

the expression of G-CSFR on progenitors in CD70TG mice, LCMV infected mice or cultured

progenitors (data not shown). Therefore, we examined whether signaling through the G-CSFR was

altered by IFNγ. G-CSFR signaling induces phosphorylation of STAT3, which induces expression

of transcription factors required for expansion and differentiation of progenitor cells towards the

neutrophil lineage. To test if IFN reduced the phosphorylation of STAT3 in response to G-CSF,

total BM cells were cultured for 5, 15 or 30 minutes with G-CSF, with or without a 45 minute pre-

incubation with IFN, and the levels of phosphorylated STAT3 (pSTAT3) were measured by flow

cytometry. Indeed, we found that G-CSF induced a time-dependent increase in phosphorylation of

STAT3, which was strongly inhibited by IFN (Fig. 7A). Importantly, IFN also reduced the G-

CSF-induced phosphorylation of STAT3 in purified GMPs (Fig. 7B).

G-CSF induces STAT3-dependent expression of suppressor of cytokine signaling (SOCS) 331,

which serves as a negative regulator of G-CSFR signaling by reducing phosphorylation of

STAT332. IFN is known to induce expression of multiple SOCS family members in a variety of

cell types33. To test if IFN increases SOCS3 expression in myeloid progenitors, GMPs were

purified and stimulated with recombinant IFN for 30 minutes. We found that SOCS3 mRNA

5


99

Figure 5. IFN impairs the proliferation and differentiation of myeloid progenitors in response to G-CSF. Absolute cell numbers derived from (A) 1.000 CMPs and (B) 1.500 GMPs after 3 days of culture with M-CSF or G-CSF with or without IFN. (C) Histograms showing CFSE dilution of CMPs and GMPs cultured for 3 days with M-CSF or G-CSF with or without IFN and absolute number of cells in each CFSE dilution peak of (D) G-CSF and (E) M-CSF cultured CMPs and GMPs. (continued)

A B

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5

Chapter 5

100

levels in GMPs increased almost 20-fold after stimulation with IFN (Fig. 7C). To determine

whether the impaired phosphorylation of STAT3 is indeed dependent on IFN-induced expression

of SOCS3, the effect of IFN on G-CSF-mediated STAT3 phosphorylation was measured in the

murine myeloid progenitor cell line 32D, expressing WT or mutant human G-CSFR34. The

cytoplasmic tail of the human G-CSFR contains four conserved tyrosine-based motifs (with

tyrosine residues at positions 704, 729, 744 and 764), which are involved in recruiting factors

regulating G-CSFR signaling. Y729 has been shown to be the recruitment site for SOCS3 and a

mutant G-CSFR lacking Y729 is insensitive to SOCS3-mediated inhibition of G-CSF-induced

signaling35;36. We compared pSTAT3 levels in cells expressing the WT G-CSFR with a mutant in

which all four tyrosine motifs were substituted with phenylalanine (mNull), a truncated mutant

lacking that part of the cytoplasmic tail of the G-CSFR that contains Y729, Y744 and Y764 (D715)

and four different mutants in which only one tyrosine motif was intact and the other three were

replaced with phenylalanine (Y704, Y729, Y744 and Y764). Although less pronounced compared

to primary BM cells, IFN significantly reduced pSTAT3 levels in G-CSF-stimulated 32D cells

(F) Representative plots showing CFSE dilution vs. c-Kit or lineage expression of CFSE-labelled CMPs cultured for 3 days with M-CSF or G-CSF with or without IFN. Data in A, B and D represent mean ± s.d. from triplicate cultures and all data is representative of 3 independent experiments. *, p < 0.05, **, p < 0.01, ***, p < 0.001.

A C

B


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2.5Control

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Rel

ativ

e P

U.1

exp

ress

ion

D

*

**

*

***

Figure 6. IFN induces expression of IRF8 and PU.1. Quantitative PCR analysis of expression levels of (A) PU.1 and (B) IRF8 in purified GMPs from naïve mice and from mice 8 days after LCMV infection. Expression is relative to the levels in naïve WT GMPs. Relative expression of (C) PU.1 and (D) IRF8 in WT GMPs after in vitro stimulation with IFN. Expression is relative to the levels in unstimulated GMPs. Data in A and B represent mean ± s.d. from 3 independent experiments with 2 mice per group. Data in C and D represent mean ± s.d. from triplicate stimulations measured in duplicate. *, p < 0.05, **, p < 0.01, ***, p < 0.001. 5


101

expressing WT G-CSFR (Fig. 7D; GeoMFI, mean ± SD, G-CSF: 1146 ± 0.7 vs. IFN + G-CSF:

963 ± 21.2, p < 0.01). IFN did not affect STAT3 phosphorylation when all four G-CSFR tyrosine

motifs were substituted (mNull), the truncated mutant (D715) nor when only Y704, Y744 or Y764

was present (Fig. 7D). However, IFN did significantly reduce pSTAT3 levels in cells that still

expressed the SOCS3-recruiting motif Y729 (Fig. 7D; GeoMFI, mean ± SD, G-CSF: 484 ± 1.7 vs.

IFN + G-CSF: 373 ± 13.2, p < 0.001). All together, these experiments demonstrate that IFN

impairs G-CSF-induced STAT3 phosphorylation, which is dependent on recruitment of IFN-

induced SOCS3 to tyrosine motif Y729 of the G-CSFR.

Discussion

Multiple factors modulate hematopoiesis during stress responses like infection and inflammation in

order to mount a proper immune response to the particular type of invading pathogen. Previously,

effects of IFN on HSC self renewal37, B cell lymphopoiesis23, eosinophil development20 and

erythropoiesis24 have been reported, suggesting that IFN modulates hematopoiesis at multiple

levels. Here we demonstrate that IFN is also an important factor in modulating differentiation of

myeloid progenitors to neutrophils and monocytes. Using various mouse models and in vitro

systems we show that IFN directly induces monopoiesis over neutrophil development. Moreover,

IFN is required to induce monopoiesis and suppress neutrophil development during acute viral

infection with LCMV. Previous studies have shown severe neutrophilia after infection of IFN-/-

mice with various pathogens25-27 and an inhibiting effect of IFN on granulocyte colony formation

from human progenitor cells in vitro38, supporting our finding that IFN suppresses neutrophil

production. Similar to viral infection, infection with intracellular bacteria induces profound IFN-

production, which might explain the observed increase in monocyte numbers after infection with

such pathogens13;25. A recent study demonstrated that IFN-signaling is also required to increase

the frequency of monocytes in circulation after infection with the intracellular bacterium Ehrlichia

muris, confirming the important role for IFN in inducing monopoiesis25. Whereas IFN is required

for control of persistent LCMV infection, it is not required for clearance of LCMV clone

Armstrong during acute infection39, excluding confounding effects of viral infection in IFN-/- mice

in our experiments. During LCMV infection, T cells have been shown to enter the BM17 and inhibit

both granulopoiesis and erythropoiesis through IFN-production21. Together with our current and

earlier findings20;23;24, we postulate that IFN-producing T cells adjust the hematopoietic process

during viral infection to improve the anti-viral response. These findings have important clinical

relevance, since IFN-producing T cells have been associated with BM failure in patients with

aplastic anemia40, suggesting that transient production of IFN is required to modulate

5

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Figure 7. IFN reduces G-CSF-mediated STAT3 phosphorylation in a SOCS3-dependent way. (A) Representative histograms showing intracellular staining for pSTAT3 in total BM cells stimulated for indicated times with G-CSF with or without 45 minutes pre-incubation with IFN. (B) Representative histograms showing overlays of intracellular staining for pSTAT3 in total BM cells and GMPs after 30 minutes stimulation with G-CSF with or without pre-incubation with IFN. (C) Relative expression of SOCS3 in WT GMPs after 30 minutes in vitro stimulation with IFN measured by qPCR analysis. Expression is relative to the levels in unstimulated GMPs. Data represent mean ± s.d. from triplicate cultures. (D) Representative histograms showing overlays of intracellular staining for pSTAT3 in 32D cells expressing WT or a mutant human G-CSFR after 30 minutes stimulation with G-CSF with or without pre-incubation with IFN. The SOCS3-recruiting tyrosine motif Y729 is only present in cells expressing WT G-CSFR or the mutant receptor denoted as Y729. All data are representative of at least two independent experiments. ***, p < 0.001.

A

B C

D

G-CSF

IFN + G-CSF

Unstimulated

pSTAT3

% o

f m

ax

WT D715

Y764

mNull

Y729Y704 Y744

G-CSF

IFN + G-CSFUnstimulated

Unstimulated

G-CSF

0

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OC

S3

exp

ress

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Control

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5 min. 15 min. 30 min.

pSTAT3

% o

f m

ax

3.97

19.9

55.5

6.51 8.29

28

Total BM GMPs

pSTAT3

% o

f m

ax

A

B C

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G-CSF

IFN + G-CSF

Unstimulated

pSTAT3

% o

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ax

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Y764

mNull

Y729Y704 Y744

G-CSF

IFN + G-CSFUnstimulated

Unstimulated

G-CSF

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OC

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0

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IFN

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OC

S3

exp

ress

ion

***

Control

IFN

5 min. 15 min. 30 min.

pSTAT3

% o

f m

ax

3.973.973.97

19.919.919.9

55.555.555.5

6.516.516.51 8.298.298.29

282828

Total BM GMPs

pSTAT3

% o

f m

ax

Total BM GMPs

pSTAT3

% o

f m

ax

5


103

hematopoiesis during infection, whereas prolonged production of IFN results in suppression of

multiple hematopoietic lineages and anemia41.

The observed effects of IFN on differentiation of GMPs in vitro and in LCMV-infected WT mice

demonstrate that IFN promotes monopoiesis and suppresses neutrophilia. We found that IFN

elevates the mRNA expression levels of IRF8 and PU.1 in GMPs. IRF8 is an important regulator in

neutrophil versus monocyte development. IRF8-deficient mice display an increase in neutrophils

and IRF8-deficient progenitors have an impaired macrophage differentiation42;43. Moreover, forced

expression of IRF8 in IRF8-deficient myeloid progenitor cells induces differentiation of these cells

into macrophages and represses granulocyte-specific genes as well as G-CSF-mediated

differentiation into granulocytes44. PU.1 is well known as the master regulator in myeloid cell

development12. Low PU.1 levels induce granulopoiesis while higher PU.1 levels induce

monopoiesis45. Phosphorylation of PU.1 allows interaction with IRF846, and increased macrophage

development in zebrafish resulting from overexpression of IRF8 is dependent on the presence of

PU.147. Finally, we have recently shown that IFN induces an IRF-1 mediated upregulation of PU.1

in CMPs, which caused a reduction in erythroid and an increase in myeloid output24. Thus, the

stimulating effect of IFN on monocyte development can be explained by an increase of PU.1 and

IRF-8 in myeloid progenitor cells. The IFN-induced increase in expression of these transcription

factors likely predisposes myeloid progenitors for monocyte differentiation, as the increased CFU-

M capacity of total BM from CD70TG mice is maintained in vitro in the absence of exogenous

IFN.

Besides the positive impact of IFN on monocyte-inducing factors such as PU.1 and IRF-8, IFN

inhibits G-CSF-mediated neutrophil differentiation. G-CSF induces proliferation, differentiation

and survival of progenitor cells and neutrophils and elevated levels of G-CSF are involved in

emergency granulopoiesis10;48. G-CSF-mediated phosphorylation of STAT3 induces expression of

CCAAT/enhancer-binding protein and accelerates cell cycle progression and maturation of

neutrophils during emergency responses11;48. STAT3-induced expression of SOCS3 serves as a

negative feedback loop by interfering with phosphorylation of STAT331;32. In accordance, SOCS3-

deficient cells display prolonged STAT3 activation and hyperresponsive to G-CSF. Injection of G-

CSF in SOCS3-deficient mice results in peripheral neutrophilia and inflammatory infiltration of

neutrophils in various tissues, demonstrating that SOCS3 is also required to suppress G-CSF-

induced emergency granulopoiesis32. Recruitment of SOCS3 to tyrosine 729 of the cytoplasmic tail

of the G-CSFR is essential for the inhibiting effect of SOCS3 on STAT signaling35. Our

experiments demonstrate that IFN induces SOCS3 expression in GMPs and that the observed

IFN-mediated reduction in G-CSF-dependent STAT3 phosphorylation is dependent on SOCS3

recruitment to Y729. All together, these observations support the hypothesis that IFN acts both as

a promoter of monopoiesis and a suppressor of neutrophil development.

5

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Infection with intracellular viral and bacterial pathogens induces production of IFN and both type

of infections result in increased monocyte production. It is tempting to speculate that the IFN-

dependent mechanisms underlying the induction of monopoiesis and suppression of neutrophil

production are similar in both types of intracellular infection. Macrophages, derived from BM

monocytes, are crucial in controlling infection with intracellular pathogens and antigen presenting

monocyte-derived dendritic cells contribute to shaping adaptive immunity against these

pathogens49. Thus, there is strong rationale to induce monocyte production over neutrophil

development during intracellular pathogenic challenges. Previously it was suggested that

neutrophils found at sites of viral infection contribute to the suppression of viral replication50;51.

However, in these studies, characterization of neutrophils ex-vivo and depletion of neutrophils in

vivo was performed using an antibody against the neutrophil marker Gr1 (clone RB6-8C5).

Although the neutrophil-specific surface molecule Ly6G is the major antigen detected by this

antibody, Gr1-antibody also binds to the Ly6C antigen, which is moderately expressed on

monocyte during homeostasis and expression increases significantly upon infection (also seen in

Fig. 3C). Therefore, injection of Gr1-antibodies in mice depletes neutrophils as well as monocytes

and other Ly6C-expressing leukocytes particularly during infections, which questions the

conclusions of previous findings on the contribution of Gr1-characterized or depleted neutrophils in

controlling viral infection50;51. Correspondingly, injection of the neutrophil-specific Ly6G antibody

(1A8) in mice depletes neutrophils, but not Gr1+ monocytes, making these antibodies a useful tool

in studies on the effect of neutrophil depletion during viral infection52. Indeed, depletion of

neutrophils with the Ly6G-specific antibody had no effect on replication of HSV-1, while depletion

with the Gr1-antibody exacerbated virus replication, which suggests an important role for

monocytes in controlling this viral infection53. However, another recent study reported increased

replication of influenza virus after depletion of neutrophils with the Ly6G-antibody, suggesting that

the contribution of neutrophils in controlling viral infection may also depend on the type and/or site

of viral infection54.

Taken together, we have demonstrated that IFN is an important cytokine in directing myelopoiesis

during acute viral infection. IFN directly stimulates monopoiesis by inducing expression of

lineage-specific regulators of myelopoiesis and it suppresses neutrophil production by perturbing

G-CSF-mediated signaling.

Acknowledgements

We thank Cláudia Brandão Silva for technical assistance, B. Hooibrink for cell sorting, the staff of

the animal facility of the AMC for excellent animal care, Prof Dr René van Lier for stimulating

discussion and critical reading of the manuscript. The authors declare no competing financial

interests. This work was supported by a VIDI grant from The Netherlands Organization of

5


105

Scientific Research (MAN; 917.76.310) and a project grant from the Landsteiner Foundation for

Bloodtransfusion Research (MAN; 0607).

Materials and methods

Mice

WT, IFN-/-, CD70TGxIFN-/- and CD70TG C57BL/6 mice were housed under specific-pathogen-

free conditions. Animal experiments were approved by the Animal Ethics Committee and

performed in accordance with institutional and national guidelines.


Single cell suspensions of BM were obtained by crushing both tibias and femurs using a mortar and

pestle and filtering through 40 μm cell strainers. Erythrocytes in peripheral blood were lysed with

an ammonium chloride solution. Purification of CMPs and GMPs was performed as described

previously20. Briefly, BM cells were stained with biotin-conjugated antibodies against the lineage

markers CD4 (GK1.5), CD8α (53-6.7), B220 (RA3-6B2), CD11b (M1/70), Gr1 (RB6-8C5),

Ter119 (Ly-76) and lineage cells were depleted using streptavidin microbeads (Miltenyi Biotec)

and MACS LS-columns (Miltenyi Biotec). Lineage depleted cells were incubated with antibodies

for CD34, CD16/32, Sca-1 and c-Kit and progenitors were sorted on a FacsAria (BD Biosciences).

Sca-1 could not be included in the identification of myeloid progenitors in CD70TG mice and

LCMV-infected mice, due to a systemic IFNγ-mediated upregulation of Sca-119. Within

experiments, similar definitions were used for the progenitors of all mice included.

The antibodies used for identification of cells by flow cytometry were CD62L-PE-Cy7 (MEL-14),

IFNγ-APC (XMG1.2), CD34-FITC (RAM34), Gr1-FITC (RB6-8C5), Ly6G-PE (1A8, both from

BD Pharmingen), CD115-biotin (AFS98), CD16/32-PE-Cy7 (93), CD11b-APC and CD11b-APC

eFluor-780 (M1/70), c-Kit-PE (BD Biosciences) and c-Kit-APC eFluor-780 (2B8), Sca-1-PE and

Sca-1-PE-Cy7 (D7), BrdU-FITC (PRB-1), CD127-FITC (A7R34), PD-1-PE (J43), CD8-PE and

CD8-PerCP-Cy5.5 (53-6-7), LCMV GP33-41 tetramer-APC, CD4-Alexa Fluor 700 (L3T4), CD4-

PE-Cy7 (GK1.5), CD3-APC eFluor-780, BrdU-FITC (PRB-1), STAT3-Alexa Fluor 647 (4/P-

STAT3, against the phosphorylated Y705 of STAT3). Expression of the G-CSF-R was measured

with biotinylated G-CSF36. Biotin-conjugates were visualized by streptavidin-PerCP-Cy5.5 (BD

Pharmingen), streptavidin-PE-Cy7 or streptavidin-PE. Where possible, cells were stained in the

presence of anti-CD16/CD32 block (2.4G2) and dead cells were excluded by propidium iodide.

Intracellular cytokine staining of stimulated T cells was performed as described previously55. All

antibodies and secondary reagents were obtained from eBioscience, unless otherwise specified.

Flow cytometry analyses were performed on FACSCanto (BD Biosciences) and data were analyzed

using FlowJo software (Tree Star).

5

Chapter 5

106

Cell culture and stimulation

For liquid cultures, CMPs and GMPs were cultured for 3 days at 37°C in a humidified incubator at

5% CO2 in 96-well plates in X-VIVO-15 (Lonza) at a density of 1.000 (CMPs) or 1.500 (GMPs)

cells per well in the presence of SCF and M-CSF or G-CSF. IFNγ was added to cultures when

indicated. Concentrations used were: SCF (5 ng/ml), G-CSF (5 ng/ml), M-CSF (5 ng/ml), IFNγ (20

ng/ml). CFSE staining of (100.000-150.000) progenitors was performed with 0.5 μM CFSE

(Invitrogen) in PBS for 12 minutes at 37°C. The number of cells in culture was quantified by flow

cytometry and for CSFE histograms between 5.000 and 20.000 cells measured. For colony-forming

assays, 250 purified GMPs were plated in duplo in methylcellulose media (Methocult M3434,

StemCell Technologies) in 6-well plates, supplemented with IFNγ when indicated, and colonies

were scored at day 8. When cells were stimulated with IFNγ and/or G-CSF for pSTAT3 staining or

RNA isolation, cells were starved for at least 1 hour in serum free X-VIVO 15 medium prior to

stimulation. Cytokines were obtained from Peprotech. For in vitro stimulation of BM T cells, total

BM was stimulated for 5 hours with LCMV peptide GP33-41 (1 μg/ml, KAVYNFATC, Genscript) in

the presence of Brefeldin A (10 μg/ml, Sigma) in IMDM containing 10% FCS or with

PMA/ionomycin as described previously55.


RNA extraction was done with Trizol (Invitrogen) and reverse transcribed to cDNA using random

hexamers and Superscript II reverse transcriptase (Roche). Quantitative real-time PCR was

performed in duplicate using Express SYBR GreenER (Invitrogen) on the StepOnePlus RT-PCR

system (Applied Biosystems). Data was normalized using 18S rRNA as a reference gene. Primer

sequences are available on request.

Adoptive transfer, LCMV infection and anti-CD40 injection

T cells were isolated from lymph nodes and spleens using CD4 and CD8 microbeads (Miltenyi

Biotec) and MACS positive bead selection with LS columns (Miltenyi Biotec). Cells were

resuspended in PBS and 10x106 cells were injected intravenously into CD27-/- and

CD70TGxCD27-/- recipient mice, which were analyzed 3 weeks later. For LCMV infection, mice

were infected i.p. with 1x105 PFU of LCMV clone Armstrong and analyzed at indicated days. For

sequential analysis, a drop of blood was collected by puncturing the vena saphena and analyzed by

flow cytometry. An agonistic antibody to CD40 (100 g, clone FGK-45) or a rat control antibody

(100 g, clone GL113) was injected intraperitoneally in 200 l PBS at day 1 and 3 and mice were

analyzed at day 5.

5


107

BrdU

BrdU (5 mg) was injected intraperitoneally in 200 μl PBS. Collected cells were stained with the

appropriate antibodies and BrdU was visualized as described elsewhere56.

pSTAT3 phosphoflow Staining

32D cells expressing WT or mutant human G-CSFR34 were maintained in RPMI 1640 medium

(Lonza) supplemented with 10% FCS and IL-3. When indicated, cells were pre-incubated with

IFNγ for 45 minutes and subsequently stimulated with 5 ng/ml murine G-CSF (for primary murine

cells) or 10 ng/ml human G-CSF 36 (for 32D cells) for 30 minutes. Cells were fixed for 10 minutes

at 37°C by adding an equal volume of BD Cytofix/Cytoperm buffer (BD Biosciences), chilled on

ice for 1 minute, washed, and fixed with -20°C 90% methanol for a least 1 hour. Cells were washed

and incubated with pSTAT3-Alexa Fluor 647 antibody at room temperature for 45 minutes.

Statistics

Mean values ± s.d. or s.e.m. are shown. Statistical analysis was performed using either a paired or

unpaired two-tailed Student’s t-test with GraphPad Prism software and significance is indicated by

*, p < 0.05, **, p < 0.01, ***, p < 0.001.

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35. Hortner M, Nielsch U, Mayr LM et al. Suppressor of cytokine signaling-3 is recruited to the activated granulocyte-colony stimulating factor receptor and modulates its signal transduction. J.Immunol. 2002;169(3):1219-1227.

36. Hermans MH, van de Geijn GJ, Antonissen C et al. Signaling mechanisms coupled to tyrosines in the granulocyte colony-stimulating factor receptor orchestrate G-CSF-induced expansion of myeloid progenitor cells. Blood. 2003;101(7):2584-2590.

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40. Sloand E, Kim S, Maciejewski JP et al. Intracellular interferon-gamma in circulating and marrow T cells detected by flow cytometry and the response to immunosuppressive therapy in patients with aplastic anemia. Blood. 2002;100(4):1185-1191.

41. Young NS, Scheinberg P, Calado RT. Aplastic anemia. Curr.Opin.Hematol. 2008;15(3):162-168.

42. Tsujimura H, Nagamura-Inoue T, Tamura T, Ozato K. IFN consensus sequence binding protein/IFN regulatory factor-8 guides bone marrow progenitor cells toward the macrophage lineage. J.Immunol. 2002;169(3):1261-1269.

43. Holtschke T, Lohler J, Kanno Y et al. Immunodeficiency and chronic myelogenous leukemia-like syndrome in mice with a targeted mutation of the ICSBP gene. Cell. 1996;87(2):307-317.

44. Tamura T, Nagamura-Inoue T, Shmeltzer Z, Kuwata T, Ozato K. ICSBP directs bipotential myeloid progenitor cells to differentiate into mature macrophages. Immunity. 2000;13(2):155-165.

45. Dahl R, Walsh JC, Lancki D et al. Regulation of macrophage and neutrophil cell fates by the PU.1:C/EBPalpha ratio and granulocyte colony-stimulating factor. Nat.Immunol. 2003;4(10):1029-1036.

46. Meraro D, Hashmueli S, Koren B et al. Protein-protein and DNA-protein interactions affect the activity of lymphoid-specific IFN regulatory factors. J.Immunol. 1999;163(12):6468-6478.

47. Li L, Jin H, Xu J, Shi Y, Wen Z. Irf8 regulates macrophage versus neutrophil fate during zebrafish primitive myelopoiesis. Blood. 2011;117(4):1359-1369.

48. Hirai H, Zhang P, Dayaram T et al. C/EBPbeta is required for 'emergency' granulopoiesis. Nat.Immunol. 2006;7(7):732-739.

49. Dominguez PM, Ardavin C. Differentiation and function of mouse monocyte-derived dendritic cells in steady state and inflammation. Immunol.Rev. 2010;234(1):90-104.

50. Tumpey TM, Chen SH, Oakes JE, Lausch RN. Neutrophil-mediated suppression of virus replication after herpes simplex virus type 1 infection of the murine cornea. J.Virol. 1996;70(2):898-904.

51. Tate MD, Brooks AG, Reading PC. The role of neutrophils in the upper and lower respiratory tract during influenza virus infection of mice. Respir.Res. 2008;9(1):57.

52. Daley JM, Thomay AA, Connolly MD, Reichner JS, Albina JE. Use of Ly6G-specific monoclonal antibody to deplete neutrophils in mice. J.Leukoc.Biol. 2008;83(1):64-70.

53. Wojtasiak M, Pickett DL, Tate MD et al. Depletion of Gr-1+, but not Ly6G+, immune cells exacerbates virus replication and disease in an intranasal model of herpes simplex virus type 1 infection. J.Gen.Virol. 2010;91(Pt 9):2158-2166.

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54. Tate MD, Deng YM, Jones JE et al. Neutrophils ameliorate lung injury and the development of severe disease during influenza infection. J.Immunol. 2009;183(11):7441-7450.

55. van Gisbergen KP, van Olffen RW, van Beek J et al. Protective CD8 T cell memory is impaired during chronic CD70-driven costimulation. J.Immunol. 2009;182(9):5352-5362.

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Supplemental figure

B

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Supplemental figure 1. Percentage of viable cells (i.e. propidium iodide-negative) after culture of (A) CMPs and (B) GMPs for 3 days with M-CSF or G-CSF in the absence or presence of IFN. Data represent mean ± s.d. from 3 independent experiments with triplicate cultures.

5

Chapter 6 CD70-driven chronic immune activation is protective

against atherosclerosis.

Ronald W. van Olffen1, Alex M. de Bruin1, Mariska Vos2, Anna D. Staniszewska1, Jörg Hamann1,

René A. W. van Lier1, Carlie J. M. de Vries2, and Martijn A. Nolte1

1 Department of Experimental Immunology and 2 Department of Medical Biochemistry, Academic

Medical Center, University of Amsterdam, Meibergdreef 9, 1105 AZ, Amsterdam, The

Netherlands.

Journal of Innate Immunology 2010;2(4):344-52

Chapter 6

114

Abstract

Chronic infection and inflammation are strongly associated with the development of

atherosclerosis. To investigate whether chronic inflammation in the absence of an infectious cause

also predisposes to the development of atherosclerosis, we used a mouse model in which sterile

inflammation is driven by enhanced costimulation. Constitutive triggering of CD27 on T cells

through overexpression of CD70 on B cells increases the numbers of IFNγ-producing effector T

cells, which reduces the numbers of B cells. However, despite these pro-atherogenic features, we

found that CD70-transgenic (CD70TG) mice on an ApoE*3-Leiden background were strongly

protected against the induction of atherosclerotic lesions, with a normal increase in serum

cholesterol level and the absence of atheroprotective antibodies. We found that circulating

monocytes in CD70TG mice were activated and increased in numbers, in particular the pool of

inflammatory (Ly6C+) monocytes. Importantly, monocytes from CD70TG mice had no defects in

phagocytosis nor in TNF production, but they were more prone to apoptosis, which was IFN-

dependent. These data indicate that sterile pro-inflammatory conditions can be protective against

atherosclerosis development, possibly due to a reduced viability of circulating monocytes. This

unexpected outcome provides a new insight in the consequences of costimulatory signals and their

impact on innate immunity.

Introduction

Infection and inflammation are strongly implicated in the progression of atherosclerotic disease. A

role for inflammation in the pathogenesis of this process is supported by the presence of

monocytes/macrophages and T cells in atherosclerotic plaques. Furthermore, a large number of

studies in humans and animal models has demonstrated that infection with persistent pathogens is

associated with an increased risk of vascular disease (reviewed in1-3). Infectious pathogens may

affect atherosclerotic disease in several, not mutually exclusive, ways. First, pathogens may infect

cells residing in lesions, e.g. endothelial cells, thereby causing alterations of cellular function or

even cell destruction. Second, antigens delivered by microbes will activate specific T and B cells

that may cross-react with (altered) self-antigens, such as modified lipo-proteins, present in the

lesions. In this scenario, part of the progression of vascular disease takes place via specific (auto-

)immune reactions. Third, pathogens will, via the activation of the innate and adaptive immune

system, induce systemic inflammation resulting in increased plasma levels of inflammatory

cytokines, together with enhanced numbers of effector T cells. Both soluble and cellular mediators

of immunity localize in the atherosclerotic plaques and amplify the inflammatory process in an

antigen non-specific fashion4.

To test the influence of immune activation on the development of atherosclerosis without the

confounding effects of infectious agents, we examined atherosclerosis development in a murine

model of sterile T cell-mediated immune activation, driven by overexpression of the costimulatory

6

CD70-driven chronic immune activation is protective against atherosclerosis

115

molecule CD70. This member of the TNF-superfamily is normally only transiently expressed on

activated T cells, B cells and dendritic cells (reviewed in5) and under these conditions it provides

important pro-survival signals to T cells through its receptor CD276;7. We have previously shown

that transgenic overexpression of CD70 on B cells is sufficient to induce strong T cell activation

and a concomitant increase of IFN-producing effector T cells, which provides protection against a

challenge with influenza virus as well as tumor cells8;9. However, there is also a severe downside of

this enhanced immune activation, as these CD70-transgenic (CD70TG) mice gradually loose their

B cells due to the chronic exposure to IFN8 and in the long term even exhaust their naïve T cell

pool10. We used this non-infectious model of chronic immune activation to determine whether

persistent sterile inflammation is sufficient to enhance the development of atherosclerosis.

Results

CD70TG mice are protected against atherosclerosis

To examine the extent by which chronic immune activation could affect atherosclerotic plaque

development, we crossed CD70TG mice with atherosclerosis-prone ApoE*3-Leiden mice.

ApoE*3-Leiden mice build up severe hypercholesterolemia following a high cholesterol/fat-diet,

leading to atherosclerotic lesions in the aorta and larger arteries11. CD70TGxApoE*3-Leiden and

ApoE*3-Leiden control mice were analyzed for vascular lesion formation in the aortic root after a

12, 16 or 20 week period of high cholesterol/fat-diet. In contrast to what we expected,

CD70TGxApoE*3-Leiden mice were fully protected against atherosclerosis development, while

control ApoE*3-Leiden mice did develop large atherosclerotic lesions (Figure 1A-B). Both groups

of mice did develop diet-induced hypercholesterolemia, although serum cholesterol levels were

slightly lower in CD70TGxApoE*3-Leiden compared to ApoE*3-Leiden control mice (Figure 1C;

p=0,026 in a 2-way ANOVA between the two groups of mice, but no statistical differences per

time point); however, this small difference in cholesterol level is not sufficient to explain the large

difference in atherosclerosis development between these mice. As the production of antibodies

against oxidized LDL (ox-LDL) plays an important protective role in initiation of

atherosclerosis12;13, we tested whether the observed protection against atherosclerosis could be

explained by an increase of these athero-protective antibodies. However, we found a strongly

reduced production of anti-ox-LDL antibodies in CD70TGxApoE*3-Leiden mice compared to

ApoE*3-Leiden mice (area under the curve (average ± SD): 3808 ± 312 vs. 20920 ± 6347; p<0.01)

(Figure 1E), which correlated with a profound B cell depletion in these mice (Figure 1D). Thus,

despite the development of strong pro-atherogenic features, CD70-mediated chronic inflammation

induced a sustained protective effect against the development of atherosclerosis.

CD70 overexpression induces monocyte accumulation and activation

6

Chapter 6

116

Since monocytes/macrophages play

an important role in the development

of atherosclerosis, we tested whether

circulating monocytes were affected

in CD70TG mice, as a defect in this

cellular compartment could explain

the observed protection against

atherosclerosis (reviewed in14). Yet,

we found that CD70TG mice had

even more monocytes, defined as

Sideward ScatterlowCD11bhiF4/80+ cells, in peripheral blood than WT controls, not only percentage

wise (Figure 2A), but also in absolute numbers (Figure 2B). Moreover, these monocytes had an

activated phenotype, based on the high expression of MHC class II (Figure 2C), which was a direct

effect of the enhanced production of IFN in these mice, as it could not be seen in CD70TG mice

that were backcrossed on an IFN-deficient background (Figure 2C). To substantiate the activation

state of these cells, we examined more features of cellular activation. Indeed, monocytes in

0

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ApoE*3Lcontrol

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0

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p<0.01

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Figure 1. CD70 overexpression is protective for development of atherosclerotic lesions. (A) Representative lipid stainings (in red) of the aortic roots of ApoE*3-Leiden and CD70TGxApoE*3-Leiden mice after 12, 16 and 20 weeks of cholesterol-rich diet. (B) Average atherosclerotic lesion area in the aortic roots (performed on total plaque area) and (C) average serum cholesterol levels of ApoE*3-Leiden (in grey) and CD70TGxApoE*3-Leiden mice (in black). Black diamond symbol indicates serum cholesterol of control mice fed normal chow for 12 weeks. (D) Average percentage of splenic T and B cells in ApoE*3-Leiden (in grey) and CD70TGxApoE*3-Leiden mice (in black). (E) Serum levels of antibodies against ox-LDL in ApoE*3-Leiden (in grey) and CD70TGxApoE*3-Leiden mice (in black). Error bars indicate the standard deviation of 8 to 10 mice per group. Asterisks denote a significant difference (* p<0.05) as determined by a Student’s t-test.

6


117

F4/

80

C

A

E

MHC-II

geoMFI: 61 + 6 geoMFI: 976 + 90

13.1% 42.6%

WT CD70TG

CD

11b

F4/80

SSClowCD11bhiF4/80+:

7.9 58.6

6.227.4

3.8 80.6

3.012.6

Ly6C

CD62L

WT CD70TG

BSSClow:

SSClowCD11bhi:

WT CD70TG

*

0

10

20

30

40

50

% m

onoc

ytes

of a

ll W

BC WT

CD70TG

0

1

2

3

4

5

6

# of

mon

ocyt

es(*

106 )

/ml

WTCD70TG

*

0

1

2

3

4

# of

mon

ocyt

es(*

106)

/ml

Ly6C+ Ly6C-

WTCD70TG

F*

D

CD95 CCR5 CX3CR1 CD11a

Exp

ress

ion

(Geo

MF

I)

10

100

1000WTCD70TGIFN-/-CD70TGxIFN-/-

* **

* **

* **

* **

IFN-/- CD70TGxIFN-/-

geoMFI: 48 + 6 geoMFI: 59 + 2

F4/

80

C

A

E

MHC-II

geoMFI: 61 + 6 geoMFI: 976 + 90

13.1% 42.6%

WT CD70TG

CD

11b

F4/80

SSClowCD11bhiF4/80+:

7.9 58.6

6.227.4

3.8 80.6

3.012.6

Ly6C

CD62L

WT CD70TG

BSSClow:

SSClowCD11bhi:

WT CD70TG

*

0

10

20

30

40

50

% m

onoc

ytes

of a

ll W

BC WT

CD70TG

*

0

10

20

30

40

50

% m

onoc

ytes

of a

ll W

BC WT

CD70TG

0

1

2

3

4

5

6

# of

mon

ocyt

es(*

106 )

/ml

WTCD70TG

*

0

1

2

3

4

5

6

# of

mon

ocyt

es(*

106 )

/ml

WTCD70TG

*

0

1

2

3

4

# of

mon

ocyt

es(*

106)

/ml

Ly6C+ Ly6C-

WTCD70TG

F*

D

CD95 CCR5 CX3CR1 CD11a

Exp

ress

ion

(Geo

MF

I)

10

100

1000WTCD70TGIFN-/-CD70TGxIFN-/-

* **

* **

* **

* **

* **

* **

* **

* **

IFN-/- CD70TGxIFN-/-

geoMFI: 48 + 6 geoMFI: 59 + 2

Figure 2. CD70TG mice have increased numbers of activated circulating monocytes. (A) Representative staining for monocytes (CD11bhiF4/80+ cells) in peripheral blood of WT and CD70TG mice, gated on SSClow cells to exclude F4/80+ eosinophils. (B) Relative and absolute numbers (per ml) of monocytes in peripheral blood in WT (grey bars) and CD70TG mice (black bars). (C) Representative staining for MHC class II expression on monocytes in peripheral blood, gated on SSClowCD11bhi cells. Average geometric mean fluorescent intensity (GeoMFI) for MHC class II expression on the gated F4/80+ monocytes is indicated. (D) Mean expression of several activation-dependent cell surface molecules on peripheral blood monocytes in WT (white bars), CD70TG (dark grey bars), IFN-/- (light grey bars) and CD70TGxIFN-/- mice (black bars). (E) Representative staining for Ly6C and CD62L on peripheral blood monocytes (gated on SSClowCD11bhiF4/80+ cells). (F) Absolute numbers (per ml of blood) of Ly6C+ and Ly6C- monocytes (as defined above) in WT (grey bars) and CD70TG mice (black bars). Shown are averages ± standard deviation of 3 mice per group. In B and F, asterisks denote a significant difference (* p<0.01) as determined by a Student’s t-test, while in D they denote a significant difference (* p<0.05) as determined by a 1-way ANOVA with Bonferroni posttests.

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CD70TG mice had increased expression of CD95 (Fas), CCR5 (receptor for the inflammatory

chemokines MIP-1α/β, RANTES and MCP-2), CX3CR1 (receptor for the inflammatory chemokine

fractalkine) and CD11a (the chain of the 2-integrin LFA-1) (Figure 2D), which are all

upregulated on monocytes during activation. This increase in monocyte activation was due to an

increase in IFN production, as it was absent in CD70TGxIFN-/- mice (Figure 2D).

Circulating monocytes can be separated in two distinct subsets based on Ly6C expression, which

have been attributed different functions: Ly6ChiCD62L+ monocytes are regarded as inflammatory

monocytes that are actively recruited to inflamed tissues, whereas Ly6CloCD62L- monocytes

migrate to non-inflamed tissues to differentiate into resident macrophages15. In a pro-atherogenic

setting, Ly6Chi monocytes are recruited to atherosclerotic plaques and locally develop into lipid-

laden macrophages and are hence recognized as the key monocyte subset in the development of

atherosclerosis16;17. Intriguingly, we found that the increase in monocytes in CD70TG mice was

due to a rise in Ly6Chi monocytes, whereas the number of Ly6Clo cells was not altered (Figure 2E-

F). Thus, these data demonstrate that CD70TG mice have high numbers of activated, inflammatory

monocytes in circulation.

Monocytes from CD70TG mice have no defect in phagocytosis nor TNF production

Next, we examined if the enhanced activation state of circulating monocytes in CD70TG mice was

accompanied by changes in monocyte function. First, the phagocytic capacity of these cells was

measured in vitro, by culturing full blood from WT and CD70TG mice in the presence of ox-LDL

or zymosan (heat-killed yeast particles). We found that monocytes from CD70TG mice

phagocytosed ox-LDL and zymosan equally well, if not better, than WT monocytes (Figure 3A-B).

Moreover, 5 day cultures of monocytes with ox-LDL also indicated that monocytes from CD70TG

mice could develop normally into foam cells (data not shown). Finally, we determined whether

monocytes in CD70TG mice could still produce TNF, since this pro-inflammatory cytokine is a

potent activator of endothelial cells and a key factor in the development of atherosclerosis18;19. We

found that CD70TG mice were not defective in their production of TNF and even produced more

TNF per cell than WT mice, when stimulated with either LPS (Figure 3C) or PMA/ionomycin

(Figure 3D).

Monocytes from CD70TG mice are more susceptible to apoptosis

Since monocytes in CD70TG mice are more activated and express a higher level of CD95 (Figure

2D), we considered the possibility that chronic immune activation affected the viability of these

cells. Therefore, we cultured full blood of WT and CD70TG mice overnight and determined

cellular viability the next day by measuring the mitochondrial membrane potential, as this potential

is rapidly lost when cells go into apoptosis20. Although no major differences could be observed in

6


119

cellular viability when measured directly ex vivo (data not shown), we found that CD70TG-

derived monocytes died more rapidly than cells from WT mice and that this feature was specific

for monocytes, as we found no difference for granulocytes in these same cultures (Figure 4A-

B). This increased apoptosis susceptibility was seen in monocytes from CD70TG mice, but not

from CD70TGxIFN-/- mice, thus demonstrating that this process was dependent on IFN

(Figure 4A-B). Finally, we tested whether the enhanced propensity to die was intrinsic to the

monocytes and therefore we sorted Ly6Chi and Ly6Clo monocytes from peripheral blood of both

mice. Upon overnight culture, we observed an increase in monocyte apoptosis in both

populations of CD70TG mice, measured either with Mitotracker to determine mitochondrial

membrane potential (Figure 4C) or with propidium iodide to test cell membrane integrity

(Figure 4D). These findings demonstrate that the enhanced susceptibility to apoptosis is indeed

cell-intrinsic and affects both Ly6Chi and Ly6Clo monocyte subsets in CD70TG mice.

Figure 3. Monocytes of CD70TG mice can phagocytose and produce high amounts of TNFUptake of (A) DiI-labeled ox-LDL or (B) FITC-labeled zymosan by CD11bhiF4/80+ monocytes after 3 hours incubation at 37oC of full blood from WT (dotted line) or CD70TG mice (black line) in the absence (filled grey histogram) or presence of the indicate compounds. Data are representative stainings of two independent experiments of 3 mice per group. TNF expression (in GeoMFI) in CD11bhiF4/80+ monocytes from WT (grey bars) and CD70TG mice (black bars) after 3 hours stimulation with (C) LPS or (D) PMA/ionomycin in the presence of Brefeldin A. Bars indicate average ± standard deviation of 3 mice per group. Asterisks denote a significant difference (* p<0.01) as determined by a Student’s t-test.

% o

f Max

ZymosanFITC

ControlWTCD70TG

Zymosan

% o

f Max

DiIOx-LDL

ControlWTCD70TG

Ox-LDL BA

0

10000

20000

30000

40000

50000

TN

F

exp

ress

ion

(Ge

oMF

I) WTCD70TG

LPS stimulation

0

200

400

600

800

1000

TN

F

exp

ress

ion

(Ge

oMF

I) WTCD70TG

PMA/iono stimulationDC

**

% o

f Max

ZymosanFITC

ControlWTCD70TG

ControlWTCD70TG

Zymosan

% o

f Max

DiIOx-LDL

ControlWTCD70TG

ControlWTCD70TG

Ox-LDL BA

0

10000

20000

30000

40000

50000

TN

F

exp

ress

ion

(Ge

oMF

I) WTCD70TG

LPS stimulation

0

200

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600

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1000

TN

F

exp

ress

ion

(Ge

oMF

I) WTCD70TG

PMA/iono stimulationDC

**

6

Chapter 6

120

A

C

Mitotracker

WT CD70TG

73% 30%

97% 86%

% o

f Max

Monocytes

Granulocytes

B

WT CD70TG

Ly6C+

Ly6C-

86 % 61 %

97 % 97 %

CD70TGxIFN-/-IFN-/-

0

50

100

150

% L

ive

cells

Monocytes Granulocytes

WTCD70TGIFN-/-CD70TGxIFN-/-

*

WT CD70TG

Ly6C+

Ly6C-

D

Pro

pidi

umIo

dide

F4/80

48% 27%

61% 23%

Mito

trac

ker

F4/80

40% 21%

47% 14%

A

C

Mitotracker

WT CD70TG

73% 30%

97% 86%

% o

f Max

Monocytes

Granulocytes

B

WT CD70TG

Ly6C+

Ly6C-

86 % 61 %

97 % 97 %

CD70TGxIFN-/-IFN-/-

0

50

100

150

% L

ive

cells



0

50

100

150

% L

ive

cells




*

WT CD70TG

Ly6C+

Ly6C-

D

Pro

pidi

umIo

dide

F4/80F4/80

48%48% 27%27%

61%61% 23%23%

Mito

trac

ker

F4/80F4/80

40%40% 21%21%

47%47% 14%14%

Figure 4. Monocytes from CD70 TG mice are more prone to apoptosis. (A) Representative staining for mitochondrial potential with Mitotracker for monocytes (CD11b+F4/80+) and granulocytes (CD11b+F4/80-) of WT, CD70TG, IFN-/- and CD70TGxIFN-/- mice after overnight culture in full blood; percentages of live (Mitotrackerhi) cells are indicated. (B) Average percentage of live (Mitotrackerhi) cells within the monocyte and granulocytes fraction after overnight culture for WT (white bars), CD70TG (dark grey bars), IFN-/- (light grey bars) and CD70TGxIFN-/- mice (black bars). Error bars indicate the standard deviation of 3 mice per group. Asterisk denotes a significant difference (p<0.01) as determined by Student’s t-test. (C-D) Representative staining for mitochondrial potential using Mitotracker (C) or cell permeability using propidium iodide (D) of sorted Ly6C+ and Ly6C- peripheral blood monocytes from WT and CD70TG mice after overnight culture. Percentages of live monocytes is indicated.

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Discussion

Although chronic immune responses to persistent pathogens are associated with an increased risk

of atherosclerosis, most likely due to the chronic and bi-directional activation of cells from the

innate and acquired immune system, we show here that sterile chronic immune activation driven by

enhanced expression of CD70 is in fact highly protective against atherosclerosis. This would argue

that the anti-pathogenic response itself, rather than the ensuing chronic inflammatory response

plays an important factor in this response. It would therefore be interesting to determine

atherosclerotic susceptibility of CD70TG mice that are chronically infected with a persistent

pathogen, such as murine cytomegalovirus or lymphocytic choriomeningitis virus. Yet, this

approach does require that these mice do not clear the virus, which is not trivial as CD70TG mice

did display increased anti-viral responses against infection with influenza virus9. From a different

perspective, it would also be interesting to test whether overexpression of other TNF-superfamily

members, which generally leads to sterile chronic immune activation as well (summarized in5), also

induces protection against atherosclerosis. This might enable us to unravel the molecular pathways

that underlie the observed effects and establish whether they can be generalized to chronic immune

activation or that they are specific to CD27-CD70 interactions.

Development of atherosclerotic plaques is driven by the recruitment of monocytes to the intima of

inflamed blood vessels, which is mediated by activated endothelial cells and locally produced

chemokines. Many studies have shown that defects in this recruitment can prevent the onset of

atherosclerosis (reviewed in4) and it has also been shown that CCR5, CX3CR1 and CCR2 are all

required for the transendothelial migration of Ly6Chi monocytes in atherosclerosis 17;21. Yet, we

observed no defects in the expression of these receptors on monocytes in CD70TG mice and even

found that expression of CCR5 and CX3CR1 was increased (Figure 2D). CCR2 was not

differentially expressed and in vitro transwell migration assays with its ligand MCP-1 also showed

no differences compared to WT controls (data not shown). Furthermore, CD62L and 2-integrins,

required for rolling and firm adhesion, respectively, were not differentially expressed on monocytes

from CD70TG mice (Figure 2). These data make it highly unlikely that a defect in transendothelial

migration of monocytes is the cause of the observed atheroprotective effect in these mice.

Previous experiments with CD70TG mice have revealed that production of IFN by T cells plays

an important role in the phenotype of these mice, in particular the gradual loss of B cells8. Although

serum levels of IFN are below detection limit in CD70TG mice (data not shown), the dramatic B

cell loss was still observed on the ApoE*3-Leiden background, which indicates that IFN is also

increased in this compound model. We demonstrate that IFN is responsible for the activated

phenotype as well as the increased apoptosis susceptibility of monocytes from CD70TG mice

(Figure 2D and 4A&B); this is important as it also demonstrates that triggering of CD27 on

hematopoietic progenitor cells and subsequent changes in hematopoiesis22 are not the cause of the

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observed phenotype. Importantly, overnight incubation of WT monocytes with IFN was not

sufficient to induce monocyte apoptosis (data not shown), which means that the IFN-activated

environment of the CD70TG mice increases their sensitivity to apoptosis, rather than the exposure

to IFN itself. Expression of the death receptor CD95 (FAS) was upregulated on monocytes from

CD70TG mice in an IFN-dependent manner (Figure 2D), but we were unable to prevent apoptosis

with blocking antibodies against CD95L (FASL), nor with the pan-caspase inhibitor QVD (data not

shown). Although the mode of apoptosis of CD70TG-derived monocytes remains elusive at this

stage, it is conceivable that the underlying mechanism plays an important role in the protection

against atherosclerosis seen in CD70TG mice, especially since selective depletion of

monocytes/macrophages is sufficient to inhibit early atherogenesis23. Premature monocyte death

upon entry into the premature lesion will preclude accumulation of pro-inflammatory macrophages

and dendritic cells and thereby prevent or at least attenuate subsequent development of the

atherosclerotic plaque24;25. Examining atherosclerosis development in CD70TG mice in the absence

of chronic exposure to IFN would therefore be a very attractive idea to examine the contribution

of IFNγ in this process, but these experiments are hampered by the fact that IFNγ is essential for

monocyte maturation and thereby indispensable for the development of atherosclerotic lesions in

this model26. Moreover, it has been shown that TNF-deficient mice have a reduced percentage of

apoptotic cells inside atherosclerotic lesions27, which could indicate that the enhanced TNF

production of monocytes in CD70TG mice (Figure 3C-D) also plays an important role in the

increased level of apoptosis and thereby inhibiting lesion development. However, neutralizing

antibodies against TNF in the in vitro cultures could not prevent the increased monocyte

apoptosis (data not shown), which makes it difficult to substantiate this hypothesis. Further

investigation is required to assess whether the pro-apoptotic profile of CD70TG-derived monocytes

is indeed causally related to the observed protection against atherosclerosis of these mice.

In conclusion, despite the prevailing notion that chronic inflammatory conditions are contributing

to the development of atherosclerotic lesions, we describe here a model of costimulation-driven

chronic immune activation that is highly protective against the development of this disease, despite

the occurrence of several pro-atherogenic features. It will be important to establish the nature of

this strongly protective mechanism and to investigate whether it can be exploited for future

therapeutic use.

Acknowledgements

We thank Sten Libregts and Dr Klaas van Gisbergen for technical assistance and the staff of the

animal facility of the AMC for excellent animal care. CJMdeV was supported by the Netherlands

Heart Foundation (grantnr. 2008B037). MAN and RAWvL were supported by a VIDI and a VICI

Grant from The Netherlands Organization of Scientific Research, respectively.

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Mice

B cell-specific CD70TG mice, IFN-/- and CD70TGxIFN-/- mice 8, as well as ApoE*3-Leiden

mice 11, all on a C57Bl/6 background, were maintained at the animal department of the Academic

Medical Center (Amsterdam, The Netherlands) and used for experiments between 8-12 weeks of

age. Non-transgenic littermates were used as controls. For the induction of atherosclerosis,

CD70TG mice were backcrossed on a ApoE*3-Leiden-background. Identification of mutant mice

was performed by PCR analysis of tail DNA or by FACS analysis of peripheral blood cells. From

these crosses, only female ApoE*3-Leiden+ littermates (WT and CD70 TG) mice were used and

from the age of 8 weeks these mice were fed a high cholesterol/fat diet (1% cholesterol/18% fat,

Purif Diet W, 4021.36, Hope Farms, The Netherlands) for 12, 16 or 20 weeks. All mice were

handled in accordance with institutional and national guidelines and all experimental protocols

were approved by the institutional Ethics Committee for Animal Experiments.

Antibodies & Flow cytometry

Antibodies used in this study were obtained from Pharmingen: allophycocyanin (APC)-conjugated

anti-CD11b (clone M1/70), PE-conjugated anti-Fas (clone Jo2) and PE- or APC- conjugated anti-

CD62L (clone MEL-14), from eBioscience: FITC-conjugated anti-F4/80 (clone BM8) and PerCP-

Cy5.5-conjugated anti-Ly6C (clone HK1.4) or from Sanbio: PE-conjugated anti-CX3CR1 (clone

2A9-1). Antibodies against CCR5 (clone MC-68, a kind gift from Dr. Matthias Mack), CD11a

(clone H154.163; a kind gift from Dr. Yvette van Kooyk) and MHC class II (clone M5-114) were

purified from hybridoma supernatants. Anti-MHC class II was conjugated to biotin according to

standard procedures and detected with streptavidin-PerCP-Cy5.5 (Pharmingen), while PE- or

FITC-conjugated donkey anti-rat (Jackson) was used to detect non-conjugated antibodies. When

possible, Fc-receptor-mediated binding was blocked by co-incubation with anti-FcRII/III receptor

(clone 2.4G2; kind gift from Dr. Louis Boon, Bioceros, Utrecht, The Netherlands).

Blood (± 700 μl) was obtained by heart puncture and mixed with 10 μl heparine (5000 IE/ml, LEO

Pharma). Upon lysis of erythrocytes with an ammonium chloride solution, white blood cells were

resuspended in staining buffer (PBS with 0.5% bovine serum albumin) and stained with antibodies

for 30 min at 4˚C in a 96-well V-bottomed plate. For analysis of apoptosis, cells were incubated for

30 min at 37oC with Mitotracker Orange (250 nM; Invitrogen). For TNF stainings, 100 ul of

heparinized blood was cultured for 4 hours at 37˚C with LPS (1,875 ug/ml) or PMA (1 ng/ml) and

ionomycin (1 μM) in the presence of brefeldin A (1 μg/ml; all reagents obtained from Sigma).

After lysis of erythrocytes, cells were washed and stained with antibodies against F4/80 and

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CD11b, followed by fixation and permeabilization (Cytofix/Cytoperm; BD Biosciences) and

staining for TNFα. Data acquisition was done with a FACS Calibur or FACS Canto (BD) and data

were analyzed with Flowjo software (Tree Star).

Phagocytosis assay and foam cell formation

To determine the phagocytic capacity of blood monocytes, 100 µl heparinized blood was cultured

for 3 hours at 37˚C in a total volume of 1 ml Iscove’s modified Dulbecco’s medium (IMDM) +

10% FCS in the presence or absence of DiI-labeled oxidized LDL (5 µg/ml, Intracel) or FITC-

conjugated zymosan (5 mg/ml sterilized zymosan (Sigma) was incubated with 2.5 g/ml filter-

sterilized FITC (Sigma) for 45 min at RT, subsequently washed with sterile PBS and stored at -

20oC). Next, cells were washed, erythrocytes were lysed with an ammonium chloride solution and

leukocytes were subsequently stained for analysis by flow cytometry as described above.

For foam cell formation, erythrocytes were lysed from fresh blood and the remaining cell fraction

was cultured in IMDM/10% FCS on coverslips in the presence of 100 g/ml ox-LDL (RP-047,

Intracel). After 5 days of culture, coverslips were washed, fixed with 3.5% formaldehyde, stained

with oil red O (Sigma) and counterstained with hematoxylin, both according to standard protocols.

Foam cells, identified as large cells laden with red lipid droplets, were counted and analyzed for

morphology with light microscopy.

Serum cholesterol analysis

Mice underwent a fasting period of 4-12 hours after which blood was isolated. The concentrations

of total cholesterol in the serum were determined according to the manufacturer’s instructions

(bioMerieux). A cholesterol calibrator (standardized serum; bioMerieux) was used as internal

standard.

Measurement of oxidized LDL antibodies

To determine anti-oxidized LDL antibodies in serum, 96 wells plates were coated overnight at 4○C

with oxidized LDL (10 µg/ml, RP-047, Intracel) in coating buffer (0.1 M NaH4CO3). Subsequently,

plates were blocked with 5% BSA for 2 hours at RT. Sera from CD70TGxApoE*3-Leiden and

ApoE*3-Leiden mice were added into duplicate wells in various dilutions and incubated for 2

hours at RT. Plates were subsequently incubated with a biotin-conjugated goat anti-mouse IgG

(1034-08, SBA), followed by an incubation with streptavidin-conjugated horse radish peroxidase

(P0397, DAKO). Between the various steps the plate underwent extensive washing. Finally, plates

were developed with 50 µl substrate (10 ml NaAc, 100 µl tetramethylbenzidine (6 mg/ml in

DMSO, VWR), 10 µl 3% H2O2) and terminated with 2N H2SO4. Optical density was measured at

450 and 540 nm using a microplate reader.

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Histological analysis of hearts and aortas

Atherosclerotic lesion development was analyzed after completion of the high cholesterol/fat diet.

Mice were anesthetized and perfused with PBS for 10 min via a cannula in the left ventricle,

followed by a 10 min post-perfusion fixation with 1% neutral-buffered formalin (Formal-Fix,

Shandon Scientific) and subsequent storage of the hearts and aortas in formalin. Hearts were

bisected at the level of the atria and the base of the heart was taken for analysis. Cryostat 7 µm

cross-sections of the aortic root were made and stained with oil red O (Sigma). Quantification of

the atherosclerotic lesion area in the sections was performed on total plaque area, using Q-win

analysis (Leica). The mean lesion area was calculated (in μm2) from ten sections, starting at the

appearance of the tricuspid valves.

Statistical analysis

For measurement of lesion size and serum cholesterol levels at different time points, we used a 2-

way ANOVA with Bonferroni posttests and Area Under the Curve analysis to compare anti-

oxidized LDL antibodies in serum. For comparison of cell surface marker expression between 4

different groups of mice we used a 1-way ANOVA with Bonferroni posttests. For all other

analyses, we used an unpaired Student’s t-test, in which p<0.05 was considered to be significantly

different. For all statistical tests we made use of GraphPad Prism 5 software.

References 1. Becker AE, de Boer OJ, van der Wal AC. The role of inflammation and infection in coronary

artery disease. Annual Review of Medicine. 2001;52(1):289-297. 2. Epstein SE, Zhu J, Burnett MS et al. Infection and Atherosclerosis : Potential Roles of

Pathogen Burden and Molecular Mimicry. Arterioscler Thromb Vasc Biol. 2000;20(6):1417-1420.

3. Leinonen M, Saikku P. Evidence for infectious agents in cardiovascular disease and atherosclerosis. The Lancet Infectious Diseases. 2002;2(1):11-17.

4. Hansson GK, Libby P. The immune response in atherosclerosis: a double-edged sword. Nat.Rev.Immunol. 2006;6(7):508-519.


6. Hendriks J, Xiao Y, Borst J. CD27 promotes survival of activated T cells and complements CD28 in generation and establishment of the effector T cell pool. J.Exp.Med. 2003;198(9):1369-1380.

7. Peperzak V, Xiao Y, Veraar EA, Borst J. CD27 sustains survival of CTLs in virus-infected nonlymphoid tissue in mice by inducing autocrine IL-2 production. J.Clin.Invest. 2009


9. Arens R, Schepers K, Nolte MA et al. Tumor rejection induced by CD70-mediated quantitative and qualitative effects on effector CD8+ T cell formation. J.Exp.Med. 2004;199(11):1595-1605.


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11. van Vlijmen BJ, van den Maagdenberg AM, Gijbels MJ et al. Diet-induced hyperlipoproteinemia and atherosclerosis in apolipoprotein E3-Leiden transgenic mice. J.Clin.Invest. 1994;93(4):1403-1410.

12. Caligiuri G, Nicoletti A, Poirier B, Hansson GK. Protective immunity against atherosclerosis carried by B cells of hypercholesterolemic mice. J.Clin.Invest. 2002;109(6):745-753.

13. Major AS, Fazio S, Linton MF. B-lymphocyte deficiency increases atherosclerosis in LDL receptor-null mice. Arterioscler.Thromb.Vasc.Biol. 2002;22(11):1892-1898.

14. Galkina E, Ley K. Immune and inflammatory mechanisms of atherosclerosis (*). Annu.Rev.Immunol. 2009;27(165-197.

15. Geissmann F, Jung S, Littman DR. Blood monocytes consist of two principal subsets with distinct migratory properties. Immunity. 2003;19(1):71-82.

16. Swirski FK, Libby P, Aikawa E et al. Ly-6Chi monocytes dominate hypercholesterolemia-associated monocytosis and give rise to macrophages in atheromata. J.Clin.Invest. 2007;117(1):195-205.

17. Tacke F, Alvarez D, Kaplan TJ et al. Monocyte subsets differentially employ CCR2, CCR5, and CX3CR1 to accumulate within atherosclerotic plaques. J.Clin.Invest. 2007;117(1):185-194.

18. Boesten LS, Zadelaar AS, van NA et al. Tumor necrosis factor-alpha promotes atherosclerotic lesion progression in APOE*3-Leiden transgenic mice. Cardiovasc.Res. 2005;66(1):179-185.

19. Ohta H, Wada H, Niwa T et al. Disruption of tumor necrosis factor-alpha gene diminishes the development of atherosclerosis in ApoE-deficient mice. Atherosclerosis. 2005;180(1):11-17.

20. Poot M, Gibson LL, Singer VL. Detection of apoptosis in live cells by MitoTracker red CMXRos and SYTO dye flow cytometry. Cytometry. 1997;27(4):358-364.

21. Gautier EL, Jakubzick C, Randolph GJ. Regulation of the Migration and Survival of Monocyte Subsets by Chemokine Receptors and Its Relevance to Atherosclerosis. Arterioscler Thromb Vasc Biol. 2009;29(10):1412-1418.

22. Nolte MA, Arens R, van OR et al. Immune activation modulates hematopoiesis through interactions between CD27 and CD70. Nat.Immunol. 2005;6(4):412-418.

23. Stoneman V, Braganza D, Figg N et al. Monocyte/Macrophage Suppression in CD11b Diphtheria Toxin Receptor Transgenic Mice Differentially Affects Atherogenesis and Established Plaques. Circ Res. 2007;100(6):884-893.

24. Tabas I. Macrophage death and defective inflammation resolution in atherosclerosis. Nat Rev Immunol. 2010;10(1):36-46.

25. Swirski FK, Weissleder R, Pittet MJ. Heterogeneous in vivo behavior of monocyte subsets in atherosclerosis. Arterioscler.Thromb.Vasc.Biol. 2009;29(10):1424-1432.

26. Leon ML, Zuckerman SH. Gamma interferon: a central mediator in atherosclerosis. Inflamm.Res. 2005;54(10):395-411.

27. Canault M, Peiretti F, Poggi M et al. Progression of atherosclerosis in ApoE-deficient mice that express distinct molecular forms of TNF-alpha. J.Pathol. 2008;214(5):574-583.

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Chapter 7 Discussion

Discussion

129

Introduction

All blood cells are derived from a common multipotent hematopoietic progenitor, the hematopoietic

stem cell (HSC). HSCs constitute a rare population and reside in the bone marrow (BM) in specific

niches which support and regulate the two main functions of HSCs: self renewal, which is essential to

maintain adequate HSC numbers, and differentiation, required to supply sufficient numbers of mature

blood cells1. Although proliferation of HSCs is essential for the maintenance of homeostatic levels of

blood cells and HSCs, the main fraction of HSCs is in a non-proliferating or quiescent state2, to protect

the cells from loss of HSC activity resulting from exhaustion and to reduce the risk of oncogenic

transformation. Hematopoietic stress conditions caused by toxic agents, irradiation, bleeding and

infections require a response of both HSC proliferation and differentiation to increase the production

of blood cells in the BM while preserving sufficient numbers of HSCs2-4. How stress hematopoiesis is

activated and regulated during immunologic challenges is largely unknown, but evidence is emerging

that inflammatory mediators are involved in the regulation of this response. Recent evidence suggests

an important role for the type II interferon interferon- (IFN) in the regulation of HSC proliferation as

well as lineage-specific differentiation3;5-7. IFN is typically produced by T, NK and NKT cells during

an immune response to intracellular pathogens, like mycobacteria and viruses. Although IFN is an

important cytokine in inducing and orchestrating a vast array of immunological responses, its

modulating effect on hematopoiesis are plainly recognized but poorly studied and understood. IFN is

historically known as a suppressor of hematopoiesis in both human and mice and BM failure

syndromes resulting from chronic inflammation are associated with IFN production. Increased levels

of IFN in circulation and BM are detected in patients with aplastic anemia (AA)8 and it was shown

that AA patients display increased production of IFN by circulating T cells9. Moreover, a

polymorphism in the gene encoding IFN that results in overproduction of IFN, is associated with the

risk of AA10. Immunosuppressive drugs that decrease the production of T cell-derived IFN improve

the hematopoietic function in most AA patients9. Furthermore, the in vitro hematopoietic capacity of

BM from AA patients is severely attenuated and addition of anti-IFN to these cultures increases the

colony forming potential of BM cells8. Decreased numbers of IFN-producing lymphocytes have also

been associated with hematological improvement following immunosuppression in patients with

hypoplastic myelodysplasia11. In addition, IFN is associated with the hematopoietic suppression

observed in patients with Faconi anemia, HIV and graft versus host disease12-14. Although IFN is

strongly associated with impaired hematopoietic function, the molecular and cellular mechanisms

behind this suppression are less well understood. Here we discuss the early studies on the impact of

IFN on hematopoiesis and how more recent studies have extended our understanding on IFN as a

modulator of infection-induced hematopoiesis, both on HSC proliferation and differentiation.

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IFN inhibits HSC self renewal in vitro

Whereas studies investigating the effect of IFN on specific complex processes like HSC self renewal

and differentiation are rare, multiple studies have shown general inhibitory effects of IFN on

hematopoiesis in vitro. Selleri et al.15 showed that IFN is an inhibitor of hematopoiesis using a long-

term culture-initiating cell (LTC-IC) assay combined with colony forming unit (CFU) assays. Total

bone marrow (BM) cells or flow sorted CD34+ cells from BM were cultured for 5 weeks on BM

stromal cells producing IFN or with addition of exogenous IFN and subsequently CFU capacity of

the total pool of cells in culture was measured. Both exogenous IFN and IFN produced by the

stromal cell layer had a potent inhibiting effect on the number of LTC-ICs. It was demonstrated that

chronic exposure of IFN was required for the inhibiting effect, which was also shown to be dose-

dependent. However, while it was found that the number of LTC-ICs was decreased, from these

studies it could not be concluded if the clonogenicity of IFN-cultured HSCs was also impaired, as the

cultures containing LTC-ICs were not phenotypically characterized. Although it was suggested that

IFN negatively affected the percentage of cells in G1 and S phase of the cell cycle and increased the

percentage of cells in apoptosis, no quantification of the absolute number of cells was shown,

therefore, it remained unclear if IFN induces apoptosis or differentiation of LTC-ICs or only affects

their proliferation15. Similar observations were done when murine BM was used. Culture of total BM

on IFN-producing stromal cells decreased CFU activity and impaired the in vivo reconstitution ability

of the cultured cells, demonstrating a loss of HSC/progenitor activity16. Others also demonstrated an

inhibiting effect of IFN on the hematopoietic activity of human BM cells and showed that IFN acts

in synergy with IFNα17 and TNFα18 in the suppression of CFU capacity.

Surprisingly, addition of IFN to IL-3-supported cultures enhanced proliferation and CFU activity of

lineage-depleted cells from peripheral blood, while in the same study it was shown that IFN

suppressed G-CSF-supported cultures of BM cells19, suggesting that the source of progenitor cells and

supplemented cytokines affect the response to IFN. Even though the synergistic effect of IFN on IL-

3-supported cultures was also shown using CD34+ cells derived from cord blood (CB) and BM20, it

was previously shown that IL-3 alone was not sufficient to maintain CD34+CD38- LTC-ICs from

BM21, suggesting that IFN might act in synergy with IL-3 on the growth of more mature progenitors

and not immature HSCs. However, in contrast to most other studies using multiple supporting

cytokines, one report demonstrated stimulating effects of IFN on the growth of CD34+ progenitor

cells in cultures supplemented with various cytokines22. While studies reporting a negative effect of

IFN in these cultures used progenitor cells from BM, this particular study used CD34+ cells enriched

from peripheral blood, again suggesting that IFN-signaling differentially affects progenitors from BM

and blood.

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Most studies have addressed the effect of IFN on hematopoiesis by using total BM, lineage depleted

cells or CD34+-enriched progenitors, which does not address the direct effect of IFN on immature

HSCs and does not exclude indirect effects of IFN on progenitor cells. Few reports have focused on

the direct role of IFN on the function of the most immature LT-HSCs. Snoeck et al21. used a 2-step

single cell culture system to study the effect of IFN on purified CD34+CD38- human HSCs. Cells

were cultured for 14 days in liquid cultures with or without IFN and subsequently CFU activity of

these cultured cells was assessed in semi-solid cultures. This revealed that IFN strongly diminished

expansion of cells in the primary liquid cultures, indicating that IFN decreased self renewal of HSCs.

Although secondary colony formation of these IFN-cultured cells was also reduced, the authors used

the complete primary cultures in the secondary colony assays, not correcting for the number of cells in

these cultures, and therefore clonogenicity of the primary cells could not be addressed. Surprisingly,

IFN did not affect primary colony formation of more mature CD34+CD38+ cells, although the number

of cells per colony was not indicated and an IFN-mediated effect on the degree of cellular expansion

can thus not be excluded. In addition, IFN was found to support the survival of HSCs when cultured

in cytokine-free medium, as more secondary colonies were recovered from cells precultured in the

presence of IFN than in the absence of IFN21.

More recently it was shown that IFN inhibits the cellular expansion of CD34+CD38- cord blood (CB)-

derived human HSCs in liquid cultures and impairs the maintenance of LTC-ICs in these cultures, as

shown by CFU assays and reconstitution of NOD/SCID mice. Using cultures with CD34+ cells, it was

shown that neither apoptosis nor cell cycle status was affected by IFN, as Ki67 expression and DNA

content was not changed. While previous studies only used a functional read out of progenitor cells

when cultured with IFN, this study also performed phenotypical analysis of the cultured cells. This

revealed that IFN increased the percentage of cells expressing lineage markers and a decrease in the

percentage of CD34+ expressing cells, suggesting that IFN increased the differentiation of

progenitors, rather than affecting their proliferation7.

Recently, another study suggested that IFN induces in vitro proliferation of murine HSCs. It was

shown that in a 12 hour culture of total BM with BrdU, the percentage of BrdU+ LKSCD150+ cells

was increased3. However, several inflammatory cytokines, including IFN, induce expression of Sca-1

on hematopoietic progenitors. As a result, myeloid progenitors (normally not expressing Sca-1) of

which a considerable fraction lacks CD34 and expresses CD150, contaminate the HSC compartment

when HSCs are identified as LKS in combination with CD34, Flt3 and/or CD150. This contamination

can be easily prevented by excluding CD48+ cells, because all long term repopulating HSC activity is

restricted to CD48- cells. In addition, HSCs were sorted and analyzed after culture of total BM cells

with IFN, allowing indirect effects of IFN mediated by other cells present in the BM.

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Using in vitro and in vivo assays we have shown in Chapter 3 of this thesis that IFN decreases the

maintenance of highly purified murine HSCs (LKS CD150+CD48-), both phenotypically and

functionally. IFN did not affect cell cycle entry, differentiation or apoptosis of HSCs in vitro but

rather decreased the number of self renewing cell divisions. We have shown that IFN perturbs

thrombopoietin (TPO)-induced phosphorylation of STAT5 and expression of STAT5-associated genes

involved in the regulation of HSC proliferation (Chapter 3). The combined decrease in STAT5

phosphorylation and changes in expression of cell cycle genes provide a molecular explanation on

how IFN- negatively affects HSC self renewal.

All together, in vitro studies using highly purified human CD34+CD38- or murine LKS CD150+CD48-

HSCs have shown a suppressing effect on HSC activity, both in in vitro and in vivo assays, which can

be attributed to an inhibition of self renewal divisions of HSCs. Conflicting observations on the effect

of IFN on the hematopoietic capacity of more mature progenitors likely depend on the source of

progenitor cells, the purity and phenotype of progenitors, the type and read out of in vitro assays and

the supplemented cytokines in these cultures.

IFN impairs HSC activity in vivo

Although multiple studies have shown that IFN-cultured cells have an impaired reconstitution

capacity in vivo7;16, the direct effects of IFN on HSC activity in vivo is poorly studied. Several murine

knockout studies have suggested that IFN-signaling has suppressive effect on HSC function in vivo.

Mice deficient for ADAR123 or IRGM124, genes induced by IFN, have an hyperproliferative HSC

compartment and a decreased reconstitution capacity, indicating that IFN-induced genes inhibit HSC

proliferation and are required for normal HSC activity in vivo. Overexpression of IFN in IFN

transgenic mice decreased the number of CFU-GEMM colonies derived from BM, indicating a loss of

functional multipotent HSCs25. It was demonstrated that injection of IFN induces a phenotypical

expansion of LKS cells in mice26, however, as described earlier, HSCs can not be adequately

identified using the LKS definition after exposure to IFN, and these observations can therefore be

explained by an overestimation of HSC numbers due to Sca-1 upregulation on myeloid progenitor

cells. Others have demonstrated positive effects of IFN-injection on proliferation of HSCs,

characterized using vital dye staining (side population), however, since many cells express IFNR,

indirect effects of IFN-signaling on HSCs proliferation can not be excluded3. It has also been

postulated that injection of type I interferon IFN-α induces proliferation of HSCs, but the use of Sca-1

and frequent omission of CD48 upon HSC identification in that study could also have resulted in an

overestimation of HSC numbers. Moreover, part of the observed HSC proliferation upon IFN-

injection was found to be an indirect effect, most likely resulting from feedback mechanisms triggered

7

Discussion

133

by the IFN-α-induced leukopenia27. Therefore, it remained unclear if IFN (or IFNα), injected or in the

context of an immune response, directly modulates HSC function in vivo.

To address the direct effect of infection-induced production of IFN on HSCs, defined as Lineage-c-

Kit+CD150+CD48-, we have analyzed the HSC compartment in WT:IFNR1-/- chimeric mice during

infection with lymphocytic choriomeningitis virus (LCMV). We found that IFN- reduced the HSC

self-renewal capacity and modulates expression of genes (CyclinD1, p57) involved in the regulation of

HSC self renewal in vivo. These observations demonstrated that IFN directly impairs the self renewal

of HSCs during viral infection (Chapter 3).

We can not exclude that IFN induces HSC differentiation in vivo, which was recently suggested by a

study from Baldridge et al.3 Moreover, although we did not find an effect of IFN on the

differentiation of the most immature HSCs in vitro, differentiation of more mature HSCs and

downstream progenitors was accelerated by IFN (Fig. 1A&B). However, an in vivo feedback

mechanism on HSC differentiation activated by accelerated differentiation of progenitor cells can be

excluded, as only WT HSCs were impaired in their recovery after LCMV infection of WT:IFNR1-/-

chimeric mice. Therefore, effects of IFN on HSC differentiation (and self renewal) are expected to be

directly on the level of HSCs. More studies using WT:IFNR deficient chimeric mice are required to

elucidate the effects of IFN on HSC self renewal and differentiation in vivo.

36.17.56

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Figure 1. IFN induces differentiation of hematopoietic progenitors. Progenitor subsets were purified and cultured for 2 days in the presence of stem cell factor with or without IFN. (A) Representative plots of lineage and c-Kit staining and (B) absolute numbers of differentiated (lineage+) cells. *, p < 0.05, **, p < 0.01.

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The effect of IFN on hematopoietic differentiation

B lymphopoiesis

Next to its effect on HSC self renewal, IFN has been reported to have both suppressive and

stimulating effects on differentiation of specific hematopoietic lineages. Using IFN transgenic mice it

was demonstrated that IFN has a suppressing effect on B cell development25. In addition, using

transgenic mouse overexpressing CD70, which results in a CD27-mediated immune activation and

expansion of effector T cells, our lab has shown that inflammation-induced production of IFN by T

cells is sufficient to decrease B lymphopoiesis in the BM28. It has been demonstrated that IFN impairs

the proliferation of IL-7 responsive pre-B cells29. Whereas IFN alone does not directly induce

apoptosis of pre-B cells in vitro, increased IFN-mediated cell death was observed in IL-7-

supplemented cultures of pre-B cells29;30, which could be rescued by using cells from BCL2 transgenic

mice30. While IFN did not affect IL-7 mRNA expression, a decreased binding of IL-7 to its receptor

was reported, which can explain the negative effect of IFN on IL-7-induced survival and

proliferation29. Furthermore, IFN inhibited the IL-7-mediated differentiation of pre-B cells in vitro.

Many cytokines, including IL-7, but also IFN-, induce expression of suppressor of cytokine signaling

(SOCS) molecules, which induces a negative feedback to the receptor that is transmitting the signal in

order to terminate the signaling process31. We have shown that IFN induces SOCS1 and 3 in

hematopoietic progenitor cells, thereby perturbing TPO and G-CSF responses, respectively (Chapter 3

and 5). It was recently demonstrated that IFN also induces SOCS1 expression in immature B cells,

thereby abrogating signaling through the IL-7 receptor and decreasing the responsiveness of these

cells to IL-732, thus providing an explanation for above mentioned observations. In fact, SOCS-

mediated inhibition of cytokine signaling could be a unifying mechanism that explains why IFN

negatively affects the differentiation of many hematopoietic lineages.

Furthermore, although decreased IL-7-signaling might cause the block in differentiation, it can not be

excluded that IFN also changes the transcriptional profile required for B cell differentiation. We have

previously shown that IFN directly increases expression of PU.1 in myeloid and erythroid

progenitors6. Whereas high levels of PU.1 induce myeloid and inhibit B cell differentiation of

progenitors, low levels of PU.1 are required for B cell differentiation33. Therefore, next to an IFN-

mediated decreased responsiveness to IL-7, IFN might also increase PU.1 expression in B cells and

thereby contribute to the block in B cell differentiation.

Erythropoiesis and thrombopoiesis

Chronic infections can result in anemia, and a role for IFN has been recognized in the reduction of

red blood cell (RBC) levels. IFN disturbs iron homeostasis and thereby the erythroid balance by

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Discussion

135

inducing iron retention in macrophages34. Furthermore, IFN-activated macrophages contribute to

RBC loss by increased hemophagocytosis35. However, erythroid colony formation of hematopoietic

progenitors is also directly suppressed by IFN17;18 and it has been shown that IFN primarily inhibits

the earliest stages of erythroid differentiation and proliferation36. Erythroid progenitors are obtained

from megakaryocyte-erythroid progenitors (MEPs), which, in contrast to myeloid progenitors, express

low levels of PU.1 and high levels of GATA-1. Using human erythroid progenitors, we have recently

shown that IFN induces PU.1 in an IRF1-dependent manner and thereby suppresses the CFU

potential of these cells. Inhibition of either IRF-1 or PU.1 expression was sufficient to overcome the

IFN-induced reduction in erythroid colony formation. In addition, chronic exposure to IFN increases

PU.1 protein and mRNA expression in murine MEPs and CD71+ erythroid progenitors in vivo, which

was a direct effect of IFN, since similar effects were observed when these cells were purified and

exposed to IFN in vitro6. In conclusion, since PU.1 and GATA-1 physically interact and inhibit each

others function, an IFN-mediated increase of PU.1 alters the transcriptional profile of hematopoietic

progenitors, thereby blocking their erythroid differentiation.

In addition, IFN might interfere with the cytokine response to erythropoietin (EPO), which is an

essential regulator of erythropoiesis. EPO signaling results in phosphorylation of STAT5 and, like

IFN, induces expression of SOCS1 and SOCS331. We have shown that IFN reduces STAT5

phosphorylation in response to TPO, and therefore, it is not unlikely that, next to PU.1, SOCS1 and 3

are involved in the IFN-mediated suppression of erythropoiesis37.

Strikingly, although IFN diminishes TPO signaling in HSCs and impairs HSC proliferation, IFN

does not interfere with the strongly TPO-dependent process of megakaryopoiesis ((Fig. 2A&B) and38).

A possible explanation for this dissimilarity is the requirement of STAT1 for megakaryopoiesis, which

is a downstream target of GATA1 but also activated by IFN signaling39. In addition, the IFN target

genes IRF1 and IRF2 are involved in the regulation of megakaryopoiesis by inducing expression of

the integrin CD4139;40. Whereas IFN is not required for megakaryopoiesis, IFN-induced factors do

not reduce but in stead contribute to the process of platelet formation. Although platelet counts are

decreased twofold in CD70TG mice, but not CD70TG*IFN-/- mice (Fig. 2C), this is most likely due

to increased phagocytosis of platelets by IFN-activated macrophages in the liver and spleen (data not

shown and6). We found that the percentage of reticulated platelets is increased twofold in CD70TG

mice (Fig. 2D), which indicates that the IFN-mediated decrease in platelet counts is the result of

increased removal of platelets rather than reduced megakaryopoiesis. In addition, culture of WT and

CD70TG BM, enriched for c-Kit expressing cells, revealed that megakaryopoiesis of CD70TG

progenitors is indeed not impaired, but even slightly elevated (Fig. 2E).

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Myelopoiesis

Although IFN suppresses B lymphopoiesis and erythropoiesis in the BM, both promoting and

inhibiting effects of IFN have been described on myelopoiesis, suggesting that IFN also plays an

important role in directing lineage-specific myeloid differentiation during immunological challenges.

A critical role for IFN in the regulation of infection-induced myelopoiesis was demonstrated by

infecting IFN deficient mice with Mycobacterium Bovis, which displayed a severe granulocytosis in

BM, peripheral blood and spleen41. Similar increases in granulocytes have been reported after

infection of IFN deficient mice with Mycobacterium tuberculosis42 and Toxoplasma gondii43.

Recently it was demonstrated that Ehrlichia muris infection induces expansion of peripheral blood

monocytes in WT mice, whereas neutrophils expanded in IFN deficient mice, suggesting that IFN

has a directing role in monocyte and neutrophil differentiation44. Similarly, we have observed that

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E Figure 2. Megakaryopoiesis is not inhibited by IFN. BM progenitors were purified and cultured for 5 days in the presence of TPO with or without IFN. (A) Representative histograms of DNA staining and (B) polyploidy distribution in the cultured cells. (C) Number of platelets and (D) percentage of reticulated platelets in indicated mouse strains. (E) Number of megakaryocytes with indicated polyploidy in cultures of WT and CD70TG progenitors. *, p < 0.05, **, p < 0.01.

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Discussion

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LCMV infection of WT mice induces predominantly monopoiesis, whereas, in contrast, neutrophil

development was strongly increased in IFN deficient infected mice. Using in vitro and in vivo assays,

we have shown that IFN is directly involved in both monocyte and neutrophil production. IFN

suppresses both progenitor proliferation and differentiation towards neutrophils in response to G-CSF

by interfering with the G-CSF signaling pathway, whereas M-CSF-mediated proliferation and

differentiation was not affected. We found that IFN-induced SOCS3 expression impairs

phosphorylation of STAT3, which is an essential transcription factor during emergency

granulopoiesis45. In addition, IFN elevates expression of the monocyte-promoting transcription

factors PU.1 and ICSBP in myeloid progenitors, both in vivo and in vitro, demonstrating that IFN

directs monocyte and neutrophil development by regulating expression of transcription factors and

cytokine responses (Chapter 5). These findings are in agreement with a previous report demonstrating

that IFN stimulates monocytic colony formation from human progenitor cells when cultured with G-

CSF, GM-CSF or IL-3, while G-CSF-induced granulocytic colony formation was impaired in the

presence of IFN46.

Multiple studies have demonstrated suppressive effects of IFN on eosinophil development upon viral

infection or sensitization with an allergen. Mice deficient for INF signaling show an accumulation of

eosinophils in the lungs or brains upon infection with respiratory syncytial virus47 or Borna disease

virus48, respectively. In addition, IL-12-mediated induction of IFN signaling inhibits eosinophilia

upon allergen challenge of sensitized mice49 or after parasite infection50. We have shown that IFN has

a direct inhibiting effect on the differentiation of eosinophils, as IFN reduces expression of IL-5Rα

and GATA-1 in progenitor cells, which are both essential factors in eosinophil development. Whereas

IFN impairs the IL-5-mediated differentiation of myeloid progenitors to eosinophils, IFN did not

alter the response to GM-CSF-supported cultures, again demonstrating that IFN has lineage-specific

effects on myelopoiesis (Chapter 4). A recent report demonstrated that malaria infection induces the

generation of a novel type of progenitor cell, which was dependent on IFN signaling. Although these

cells expressed IL-7Rα, like lymphoid progenitors, transplantation studies revealed that these cells

produce myeloid progeny in vivo, which contribute to the clearance of malaria-infected cells51,

emphasizing the importance of IFN in the regulation of myelopoiesis during immunological

challenges.

These observations demonstrate that while IFN suppresses B cell lymphopoiesis and erythropoiesis, it

has suppressing and promoting effects on the development of different types of myeloid cells. IFN

induces expression of specific transcription factors, while it impairs responses to certain cytokines.

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Conclusions and implications

Although very few studies, both in human and mice, have used highly purified HSCs or WT:IFNR1

deficient chimeric mice to study the direct effects of IFN on HSCs in vitro and in vivo, respectively,

those particular studies have all shown a suppressing effect of IFN on HSC function (Chapter 3

and7;21. Contradicting reports on the effects of IFN on the hematopoietic capacity of more mature

progenitor cells in vitro may be explained by a number of technical reasons, including the use of

different cytokine cocktails supplemented to these cultures, as IFN negatively affects certain cytokine

signaling pathways, while it does not affect or even promotes the response to other cytokines.

Important players in the IFN-dependent modulation of cytokine responses are the SOCS proteins

which are involved in directing cytokine responses regulating HSC function and development of

various hematopoietic lineages (Chapter 3 and 5 and31;32). In contrast, IFN induces expression of

transcription factors like PU.1 and IRF8 (Chapter 5) which directly promotes differentiation of

hematopoietic cells into specific lineages52;53 (Fig. 3&4).

We postulate that the differential affects of IFN on particular hematopoietic lineages in vivo

facilitates the expansion of the appropriate immune cells to combat the invading pathogen. Myeloid

cells are the first line of defense against pathogens, and in contrast to RBCs, have a very limited

lifespan. A transient decrease in erythropoiesis would therefore not affect oxygen transport, while a

HSC

Self renewal

Differentiation

Progenitors

Self renewal

Lymphoid Myeloid

Eosinophils Neutrophils Monocytes

CD27

CD27

IFN

IFN

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IFN

Figure 3. Schematic overview of the studied effects of IFN and CD27 triggering on HSC self renewal and differentiation of HSCs/progenitors. IFN inhibits self renewal of HSCs, promotes monopoiesis and reduces neutrophil and eosinophil development. CD27 signaling induces HSC self renewal and impairs lymphoid differentiation of HSCs/progenitors.

7

Discussion

139

decreased competition for growth factors and space in the BM would facilitate the expansion of

myeloid cells. Likewise, and as shown by Ueda and colleagues, decreased B lymphopoiesis allows an

increased myelopoiesis as both cells compete for common developmental resources54. Moreover,

whereas monocytes are strongly involved in immune responses to intracellular pathogens, which

typically induce IFN production and thereby monopoiesis, neutrophils and eosinophils are more

suited to clear extracellular infection, which do not elicit the production of IFN. Because these

particular myeloid cells are derived from the same myeloid progenitor, IFN plays a critical role in

specifically promoting monocyte expansion and perturbing the differentiation pathways of cells not

required for clearance of the pathogen. Although IFN is not required for maintaining steady state

hematopoiesis, it is essential in the regulation of hematopoiesis during immunological challenges,

indicating that IFN is an important factor in controlling hematopoietic homeostasis during stress

situations.

IFN STAT1 IRF1 PU.1

GATA1 IL-5Ra

IRF8

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HSC self renewal

Monocytes

Figure 4. Overview of the molecular effects of IFN signaling on HSC self renewal and differentiation. IFN induces expression of SOCS molecules which impair self renewal of HSC and development of neutrophils by inhibiting phosphorylation of STAT5 and STAT3 by TPO and G-CSF, respectively. INF induces expression of PU.1 and IRF8, promoting monopoiesis. PU.1 blocks GATA1 function, inhibits expression of IL-5Rα and thereby decreases eosinophil development.

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We have shown that IFN reduces the self renewal of HSCs (Chapter 3), suggesting that IFN protects

HSCs from exhaustion resulting from excessive proliferation during inflammation-induced

hematopoietic stress conditions. However, as mentioned earlier, we can not exclude that IFN

decreases HSC self renewal in order to facilitate differentiation of HSCs. In addition, various reports

have suggested that an IFN-dependent upregulation of Fas, resulting in increased cell death of

hematopoietic progenitors, contributes to a decline in hematopoietic output55-57. Although short term

exposure to IFN might thus preserve sufficient HSCs in times of immunological stress or,

opposingly, increase the hematopoietic differentiation, chronic exposure to IFN in both cases result in

declining HSC numbers and impaired hematopoietic output. This is evident in patients with aplastic

anemia (AA), graft versus host disease (GVHD) and HIV suffering from BM failure8-10;12;14. Whereas

most information regarding the role of IFN on BM failure in human disease comes from studies on

AA, chronic immune activation is generally the underlying cause of IFN-associated BM failure58.

Increased IFN production is found in BM and circulating T cells of AA patients, and the presence and

loss of these IFN producing lymphocytes correlated well with the response to immunosuppressive

therapy and subsequent recovery9. Moreover, anti-IFN- antibodies improve the in vitro capacity of

hematopoietic progenitors derived from patients with AA59, suggesting that next to suppression of the

immune system, blocking IFN can directly alleviate immune mediated hematopoietic suppression.

Although recombinant IFN60 and anti-IFN antibodies61 both are used clinically in diseases associated

with a declined or exacerbated immune response, respectively, anti-IFN antibodies are not used to

reduce hematopoietic suppression in patients with BM failure. However, the use of anti-IFN

antibodies might be a promising therapeutic intervention, as survival of mice with T cell-induced BM

failure, characterized by severe pancytopenia and BM hypercellularity, was rescued by administration

of anti-IFN antibodies62. Altogether, whereas more studies are needed to elucidate the exact effects of

IFN on intricate processes like HSC self renewal and differentiation, we have shown that, while

required for the modulation of hematopoiesis during viral infection, IFN negatively affects HSC

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7

Appendices

Nederlandse Samenvatting

List of publications

Dankwoord

Appendix I

146

Nederlandse samenvatting

De aanmaak van bloedcellen in het beenmerg, een proces dat hematopoiese wordt genoemd, zorgt

ervoor dat er een continue aanvoer is van nieuwe bloedcellen. Omdat bloedcellen een beperkte

levensduur hebben is dit proces cruciaal voor het functioneren en gezond houden van ons lichaam.

Alle bloedcellen ontstaan uit een hele kleine groep bijzondere cellen, de hematopoietische stamcellen.

Deze stamcellen hebben de eigenschap om door celdeling, of proliferatie, cellen te genereren die

identiek zijn aan de moedercel, een proces dat wordt aangeduid met de term ‘zelfvernieuwing’. Dit

proces is nodig om ervoor te zorgen dat we gedurende ons leven over voldoende stamcellen

beschikken. Een andere eigenschap van stamcellen is dat ze door stapsgewijze uitrijping, of

differentiatie, en celdeling alle verschillende soorten bloedcellen van ons lichaam genereren. Deze

processen werken zo efficiënt en de stamcellen zijn zo potent dat een enkele stamcel alle bloedcellen

van een muis kan maken na transplantatie van een stamcel in een bestraalde muis. In de normale

situatie zijn stamcellen het grootste deel van de tijd inactief en delen maar heel sporadisch. De

inactieve staat, zelfvernieuwing en differentiatie van stamcellen is sterk gereguleerd om een gezonde

balans te houden tussen het aantal stamcellen en de mate van bloedcel productie. Factoren buiten

(excentriek) en binnen (intrinsiek) de stamcellen in het beenmerg beïnvloeden de hematopoiese.

Cytokines, eiwitten die uitgescheiden worden door cellen en signalen overbrengen naar een cel

waaraan de cytokine kan binden via een receptor, zijn een belangrijke component van de excentrieke

regulatie van stamcellen. Intrinsieke regulatie vindt vooral plaats door transcriptiefactoren, eiwitten

die de activiteit van een gen bepalen en daarmee specifieke processen in de cel reguleren.

De stamcellen genereren witte bloedcellen, de cellen van het immuunsysteem die ziekteverwekkers, of

pathogenen, opruimen na infectie, rode bloedcellen, die zuurstof transporteren, en bloedplaatjes,

hoofdzakelijk betrokken bij het in stand houden van de hemostase door bloedstolling. De witte

bloedcellen van het immuunsysteem kunnen worden onderverdeeld in twee groepen: de cellen van het

aangeboren immuunsysteem en de cellen van het adaptieve (of verworven) immuunsysteem. Het

aangeboren immuunsysteem bestaat uit verschillende types immuuncellen, zoals granulocyten en

monocyten (ook wel myeloide cellen genoemd), en ruimt pathogenen op door ze op te eten en door het

uitscheiden van stoffen die de pathogenen vernietigen. Pathogenen worden herkend door myeloide

cellen via moleculaire patronen die veelvoudig voorkomen op deze micro-organismen. Het adaptieve

immuunsysteem bestaat uit B en T cellen (lymfocyten) en deze cellen hebben als taak de pathogenen

op te ruimen die niet door het aangeboren immuunsysteem vernietigd worden. B cellen scheiden

antistoffen uit die specifiek de pathogeen herkennen, de humorale afweer, terwijl T cellen de cellulaire

afweer vormen en geïnfecteerde cellen kunnen doden. Daarnaast scheiden T cellen nog cytokines uit

die belangrijk zijn in de regulatie van de immuunrespons.

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Witte bloedcellen, zoals T cellen, scheiden stoffen af en brengen eiwitten to expressie op hun

celmembranen die betrokken zijn bij de zowel de directe afweer tegen pathogenen als de vormgeving

van de immuunrespons in bredere zin. Daarnaast zijn er verscheidene eiwitten beschreven die zowel

een effect hebben op de immuunrespons als op de hematopoiese. Ook is het bekend dat immuuncellen

tijdens en na infectie verblijven in het beenmerg, waar ook de stamcellen zich bevinden, wat

suggereert dat immuuncellen een feedback effect hebben op de hematopoiese tijdens een infectie. Een

van de stoffen die geproduceerd wordt door onder andere T cellen tijdens een infectie en ook een

effect heeft op hematopoiese is het cytokine Interferon-gamma (IFN). IFN is betrokken bij de

activering van verscheidene immuuncellen en meerdere oudere studies hebben aangetoond dat het

tevens een remmend effect heeft op de hematopoiese, hoewel niet bekend is wat het mechanisme van

dit effect is. Een ander molecuul dat geassocieerd is met immuunregulatie en hematopoiese is het

membraaneiwit CD70. CD70 komt alleen tot expressie op enkele types geactiveerde immuuncellen en

is betrokken bij onder andere de activatie van T cellen door binding aan het eiwit CD27, de receptor

voor CD70. Opmerkelijk is dat CD27 ook tot expressie komt op stamcellen en de functie van deze

cellen kan beïnvloeden na binding van CD70 aan de receptor. Tijdens een immuunrespons is er een

verhoogde vraag naar immuuncellen om de pathogenen te bestrijden, wat een stress respons van het

hematopoietisch systeem induceert. Aangezien de verschillende types immuuncellen een specifieke

functie hebben is het waarschijnlijk dat de reactie van het hematopoietisch systeem op een infectie

afhangt van het specifieke pathogeen. In dit proefschrift is het onderzoek beschreven dat zich richt op

het mogelijke feedback mechanisme van geactiveerde immuuncellen op de hematopoiese.

In hoofdstuk 2 hebben we onderzocht wat het effect van CD70 stimulatie via CD27 op stamcellen is.

Eerst hebben we aangetoond dat CD27 tot expressie komt op alle verschillende subsets van stamcellen

en de meeste andere voorlopercellen. Omdat CD70 alleen voorkomt op immuuncellen na activatie

hebben we speciale muizen gebruikt die constant CD70 op B cellen tot expressie brengen,

zogenaamde CD70 transgene (CD70TG) muizen, die dus een chronische ontsteking nabootsen.

Analyse van de stamcellen van deze muizen liet zien dat er een verhoging is van het aantal stamcellen.

Daarnaast vonden we dat het aantal voorlopers voor de lymfocyten enorm was afgenomen, terwijl het

aantal specifieke voorlopers voor de andere bloedcellen niet was veranderd. Analyse van genen die tot

expressie kwamen in stamcellen van CD70TG muizen suggereerde dat deze cellen vaak deelden,

minder goed in staat waren om lymfocyten te maken en dat ze net zo onder stress stonden als

stamcellen tijdens ontstekingen. Met een reeks functionele experimenten met normale en CD70TG

muizen konden we aantonen dat de stamcellen ten gevolge van de CD70 stimulatie inderdaad meer

prolifereerden, minder lymfocyten maakten en meer myeloide cellen produceerden. Een verhoogde

proliferatie is gunstig ten tijde van ontsteking als er een verhoogde vraag is naar immuuncellen. Maar

een verhoogde myeloide productie en het genenprofiel dat geassocieerd is met stress en DNA schade

zijn eigenschappen die normaal geassocieerd zijn met oude stamcellen. Dit suggereert dat chronische

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ontstekingen een versnelde veroudering van stamcellen induceert, mogelijk doordat ze zoveel vaker

moeten delen.

IFN is van oudsher beschreven als een cytokine dat een negatief effect heeft op de hematopoietische

capaciteit van stamcellen. Maar dit effect is nauwelijks onderzocht in vivo, in een levend organisme,

en hoe IFN de hematopoiese beïnvloedt is grotendeels onbekend. In hoofdstuk 3 hebben we laten

zien dat IFN een negatief effect heeft op de functie van stamcellen, zowel in vitro als in vivo. Door

stamcellen te kweken met IFN en daarna in muizen te transplanteren lieten we zien dat IFN de

capaciteit van stamcellen enorm verlaagt. Met in vitro experimenten toonden we aan dat IFN de

zelfvernieuwing van stamcellen remt. Dit wordt mogelijk veroorzaakt doordat IFN een remmend

effect heeft op de activatie van STAT5, een transcriptiefactor die belangrijk is in de regulatie van

zelfvernieuwing van stamcellen. Om te onderzoeken of IFN ook in vivo tijdens infecties een negatief

effect heeft op stamcellen werden normale muizen en muizen die geen IFN kunnen maken (IFN-/-

muizen) geïnfecteerd met LCMV, een virus dat voornamelijk voorkomt bij knaagdieren en dat bij

mensen hersenvliesontsteking veroorzaakt. Infectie met LCMV induceert de activatie van T cellen en

deze cellen produceren vervolgens IFN. Deze experimenten toonden aan dat IFN inderdaad een

remmend effect heeft op de zelfvernieuwing van stamcellen tijdens virale infectie. Daarnaast vonden

we dat IFN de expressie van een aantal genen die betrokken zijn bij stamcel zelfvernieuwing

beïnvloedde.

In hoofdstuk 4 en hoofdstuk 5 hebben we het effect van IFN op de differentiatie van myeloide

cellen onderzocht. IFN wordt met name geproduceerd tijdens virale infecties. Omdat sommige types

immuuncellen meer betrokken zijn bij het opruimen van een virale infectie (monocyten) dan andere

cellen (granulocyten) hebben we onderzocht of en hoe IFN hier bij betrokken is. Dit deden we onder

andere door gebruik te maken van CD70TG muizen, die verhoogde productie van IFN hebben, en

door muizen te infecteren met LCMV. In hoofdstuk 4 hebben we met verschillende modellen laten

zien IFN direct de productie van eosinofielen remde, een type granulocyt dat betrokken is bij

allergische reacties en het bestrijden van infecties met parasieten. IFN zorgde ervoor dat de directe

voorloper van eosinofielen veel minder gemaakt wordt. Wij vonden dat IFN de expressie van een

aantal genen, dat direct betrokken is bij de productie van eosinofielen uit voorlopercellen, zodanig

beïnvloedde, dat deze cellen veel minder goed in staat zijn om uit te rijpen tot volwassen eosinofielen.

In hoofdstuk 5 hebben we onderzocht of IFN een effect heeft op de uitrijping van neutrofielen, een

granulocyt betrokken bij het opruimen van bacteriën, en monocyten, die wel een duidelijke functie

hebben in het bestrijden van virale infecties. Deze twee celtypes ontstaan beide uit dezelfde voorloper.

We vonden dat IFN de uitrijping van monocyten stimuleert ten gunste van de uitrijping van

neutrofielen. Door normale muizen en IFN-/- muizen te infecteren met LCMV vonden we dat IFN de

productie van monocyten verhoogt, maar ook nodig is om de productie van neutrofielen niet te laten A


149

verhogen, en dus eigenlijk een remmend effect heeft op de productie van neutrofielen. We vonden dat

IFN de neutrofiel productie remt door zowel de proliferatie als de uitrijping van de voorlopercellen te

verminderen. Verder toonden we aan dat IFN dit doet door een de productie van een eiwit (SOCS3)

te stimuleren dat de activatie van STAT3 remt, een transcriptiefactor betrokken bij de proliferatie en

uitrijping van neutrofielen. Ook induceerde IFN de expressie van een aantal genen (PU.1, IRF8) die

een stimulerend effect hebben op de productie van monocyten uit voorlopercellen, en daardoor

indirect neutrofiel ontwikkeling remmen. Deze studies tonen aan dat IFN betrokken is bij de regulatie

van de hematopoiese tijdens virale infectie.

Chronische infecties zijn geassocieerd met het optreden van atherosclerose, of aderverkalking. In

hoofdstuk 6 hebben we met CD70TG muizen, ons model voor chronische ontsteking, onderzocht of

deze muizen gevoeliger zijn voor het optreden van aderverkalking. Opvallend was dat deze muizen

juist geen enkele mate van aderverkalking toonden na een cholesterol- en vetrijk dieet. Omdat

monocyten een belangrijke rol spelen in het proces van aderverkalking werden de monocyten in deze

muizen bestudeerd. CD70TG muizen hadden meer monocyten hadden en deze cellen waren extra

geactiveerd. In vitro experimenten toonden echter aan dat deze cellen heel gevoelig waren voor

celdood, wat mogelijk verklaart waarom de muizen tegen verwachting in beschermd waren tegen

atherosclerose.

In dit proefschrift hebben we het effect van immuun activatie op de hematopoiese onderzocht. We

hebben aangetoond dat er meerdere feedback mechanismen zijn waarbij geactiveerd immuuncellen

een regulerend effect hebben op stamcellen of latere voorlopers. Dit mechanisme is betrokken bij

zowel de zelfvernieuwing van stamcellen als de specifieke uitrijping van stamcellen/voorlopercellen.

Op deze manier reguleren de geactiveerde immuuncellen de productie van cellen in het beenmerg die

nodig zijn om een specifieke infectie te bestrijden. Hoewel stamcellen veel bestudeerd worden, zijn

alle factoren en hun effect op stamcellen nog verre van duidelijk. Ons onderzoek draagt onder andere

bij aan het begrijpen van ziekten die gekenmerkt worden door een falen van de hematopoiese in het

beenmerg als gevolg van een chronisch geactiveerd immuunsysteem.

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Appendix III: Dankwoord

152

Dankwoord

Ik zou veel woorden kunnen gebruiken om vele mensen te bedanken, of weinig woorden om iedereen

te bedanken die in enig opzicht heeft bijgedragen aan het tot stand komen van dit proefschrift. Omdat

de waarde van woorden vaak omgekeerd evenredig is met de mate van gebruik kies ik hier voor de

laatste optie: bedankt iedereen. Ook al is je naam hier niet vermeld, weet dat ik aan je dacht en

dankbaar ben voor je bijdrage.

Maar enkele mensen wil ik toch in het bijzonder noemen. Beste Martijn, je gaf me een tweede kans als

AIO. Ik heb geen spijt gehad dat ik 010 heb ingeruild voor 020. In korte tijd heb ik enorm veel van je

geleerd. Ik respecteer de wijze waarop je mij meestal m’n gang liet gaan en toch erg betrokken was en

oog voor de details had. Zonder die vrijheid had ik vast niet zo’n goede tijd gehad. Bedankt voor je

enthousiasme, vooral in tijden wanneer ik het zelf even niet had. Ik heb genoten van onze

wetenschappelijke discussies. Je bent een fijne vent. Veel succes met je groep.

René, ik vind het knap hoe je betrokken blijft bij zoveel projecten tegelijk. De wekelijkse meeting met

jou was altijd nuttig om de grote lijnen te bewaken en het doel niet uit het oog te verliezen.

Stan jonguh, jij kwam mij als eerste vergezellen bij de BM-posse. Bedankt voor de gezellige sfeer die

je mee bracht en je gemoedelijkheid. Je hulp bij experimenten was waardevol, maar samen lol maken,

knuffelen, bakkies doen en natuurlijk de woensdag-hakkendag (Rotterdam Terror) was ook erg

aangenaam. Succes met het afronden van je experimenten en het schrijven.

Cláudia, dear friend, you’ve been a very helpful colleague. Many huge experiments were manageable

because of your help. Thanks a lot. I enjoyed all the discussions about everything and nothing during

the experiments. Good luck in the final period of your PhD, and enjoy.

Klaas en Natasja, ook al waren jullie officieel geen deel van BM-groep, toch voelde het wel zo. Klaas,

jij was altijd een hele nuttige vraagbaak. Bedankt voor je hulp bij experimenten en het binnenhalen

van LCMV. Dat virus is erg waardevol geweest voor mijn studies. Natas, jij stond altijd klaar om te

helpen, of het nou om experimenten ging of om bestellingen. Bedankt.

Dianne, bedankt voor je hulp bij menig papierwerk en voor de talrijke gezamenlijke saffies en bakkies.

En natuurlijk de hoeders op het ARIA. Bedankt voor de goede zorg voor de muizen en de

gezelligheid.

Het ga jullie allen goed/all the best,

Alex

A

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