analysis of stress erythropoiesis during …

The Pennsylvania State University

The Graduate School

Department of Veterinary and Biomedical Sciences

ANALYSIS OF STRESS ERYTHROPOIESIS DURING INFLAMMATION:

STIMULATION OF TOLL-LIKE RECEPTORS INDUCES ERYTHROPHAGOCYTOSIS

AND ACTIVATES STRESS ERYTHROPOIESIS

A Dissertation in

Genetics

by

Laura F. Bennett

2017 Laura Bennett

Submitted in Partial Fulfillment

of the Requirements

for the Degree of

Doctor of Philosophy

August 2017

The dissertation of Laura Bennett was reviewed and approved* by the following:

Robert F. Paulson

Professor of Veterinary and Biomedical Sciences

Dissertation Advisor

Chair of the Intercollege Graduate Degree Program in Genetics

Pamela Hankey Giblin

Professor of Immunology

Chair of Committee

K. Sandeep Prabhu

Professor of Immunology and Molecular Toxicology

Ross Hardison

T. Ming Chu Professor of Biochemistry and Molecular Biology

Connie Rogers

Associate Professor of Nutrition and Physiology

Zhi-Chun Lai

Professor of Biology, Biochemistry and Molecular Biology

*Signatures are on file in the Graduate School

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ABSTRACT

Steady state erythropoiesis takes place in the bone marrow and, during homeostasis,

produces new erythrocytes at a constant rate sufficient to replace those being removed from

circulation. Chronic inflammation directly inhibits the proliferation and differentiation of

erythroid progenitors in the bone marrow, but inflammation also skews hematopoietic cells

toward myelopoiesis to produce monocytes and macrophages. The result is a severe decline in

erythroid output from the bone marrow. Previously, our lab has shown that during times of acute

anemic stress, there is a specialized stress response pathway, known as BMP4 dependent stress

erythropoiesis, which generates a large number of new erythrocytes rapidly to overcome the

anemic burden. Stress erythropoiesis relies on a progenitors which reside in the spleen and unique

signals from the microenvironment regulate their proliferation and differentiation. Work by others

has demonstrated that this pathway is activated in response to inflammation and that these stress

erythroid progenitors (SEPs) in the spleen are not inhibited by the presence of IFN-γ like steady

state progenitors in the marrow. We hypothesized that stress erythropoiesis may be activated by

inflammatory stimuli in order to produce new erythrocytes and compensate for the decline in

bone marrow erythropoiesis until homeostasis is restored.

My work has focused primarily on understanding the relationship between inflammation

and stress erythropoiesis, identifying the mechanism by which stress erythropoiesis is activated

during inflammation, and exploring the contribution of glucocorticoids to this response. In

Chapter 2, I characterize the initial stress erythropoiesis response after inducing inflammation,

showing that there is a rapid activation of GDF15 and BMP4, both key regulators of stress

erythropoiesis, and a significant increase in stress BFU-Es. All of this happens in the absence of

overt anemia, suggesting that inflammation activates stress erythropoiesis by a previously

unknown mechanism. I demonstrate that inducing inflammation results in increased

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erythrophagocytosis in macrophages in the spleen. These phagocytosed erythrocytes will be

broken down by the macrophages, resulting in an increase in intracellular heme. We show that

this small increase in phagocytosis is sufficient to lead to changes in heme-regulated genes and

identify Spi-C as a critical regulator of inflammation-induced stress erythropoiesis. In the absence

of Spi-C, GDF15 expression is diminished, resulting in decreased stress BFU-Es, and by

adoptively transferring wild type monocytes into Spi-C-/- mice, we are able to partially rescue

stress erythropoiesis. I also show in Chapter 2 that some pro-inflammatory cytokines promote

stress erythropoiesis by increasing the frequency of stress BFU-Es in in vitro culture. In vivo

erythrophagocytosis and pro-inflammatory cytokines may work together to activate stress

erythropoiesis in the absence of key signals, such as tissue hypoxia and anemia, as a preemptive

measure to offset the loss of bone marrow erythropoiesis.

Chapter 3 focuses on the role of glucocorticoids (GCs) in inflammation-induced stress

erythropoiesis. We show that GC levels increase after inducing inflammation and are necessary in

surviving. Flexed-tail (f/f) mice and GDF15-/- exhibit severe mortality in response to zymosan,

and I demonstate that this is due decreased GC production and results in a decrease in the

frequency of M2 macrophages. I will also show that the reduction in GC levels in both f/f and

GDF15 is the result of decrease transcription of Cyp enzymes in the adrenal glands.

v

TABLE OF CONTENTS

List of Figures .......................................................................................................................... vii

List of Abbreviations ............................................................................................................... x

Acknowledgements .................................................................................................................. xiii

Chapter 1 Introduction ............................................................................................................. 1

Erythropoiesis .................................................................................................................. 2 I. Murine Stress Erythropoiesis ................................................................................ 2 II. Human Stress Erythropoiesis ............................................................................... 5

Infection, Inflammation, and Erythropoiesis.................................................................... 8 I. Infection and Steady State Erythropoiesis ............................................................ 8 II. Effects of Infection and Inflammation on Stress Erythropoiesis ......................... 14 III. Immunomodulation by Stress Progenitors ......................................................... 16 IV. Glucocorticoids and Erythropoiesis ................................................................... 17 V. Glucocorticoids as Immunomodulators ............................................................... 20

Conclusions ...................................................................................................................... 21 Figures .............................................................................................................................. 23 References ........................................................................................................................ 25

Chapter 2 Zymosan-induced generalized inflammation activates stress erythropoiesis

through a novel mechanism. ............................................................................................ 31

Introduction ...................................................................................................................... 31 Results .............................................................................................................................. 33

Stress erythropoiesis is rapidly induced upon inflammatory stimulation ................ 33 Inflammation induces the expression of GDF15, a regulator of BMP4

expression, in the spleen ................................................................................... 35 Stress erythropoiesis generates new erythrocytes immediately following

zymosan treatment, delaying the onset of anemia. ........................................... 36 Inflammation induces the expression of Epo in the kidneys. ................................... 37 Zymosan induces GDF15 expression by increasing erythrophagocytosis by

splenic macrophages ......................................................................................... 38 MyD88 mediates the increase in GDF15 expression following zymosan

treatment. .......................................................................................................... 42 TNFα and IL1-β promote erythroid differentiation under stress erythropoiesis. ..... 43

Discussion ........................................................................................................................ 44 Material and Methods ...................................................................................................... 47 Figures .............................................................................................................................. 51 References ........................................................................................................................ 81

Chapter 3 How Stress Progenitors, Glucocortcoids, and M2 Macrophages Prevent Lethal

Immune Activation in Response to Zymosan-Induced Inflammation. ............................ 83

Abstract ............................................................................................................................ 83 Introduction ...................................................................................................................... 83

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Results .............................................................................................................................. 85 Glucocorticoids are increased in response to zymosan-induced inflammation. ....... 85 Decreased expression of Cyp enzymes diminishes GC production in f/f and

GDF15-/- mice. ................................................................................................ 86 Lower levels of GCs leads to an increase in pro-inflammatory cytokines. .............. 87 Glucocorticoid production promotes M2 macrophages. .......................................... 88

Discussion ........................................................................................................................ 89 Materials and Methods ..................................................................................................... 91 Figures .............................................................................................................................. 93 References ........................................................................................................................ 107

Chapter 4 Sf-Ron plays a role in regulating the differentiation of erythroid progenitors

during recovery during acute anemic stress. .................................................................... 108

Abstract ............................................................................................................................ 108 Introduction ...................................................................................................................... 108 Results .............................................................................................................................. 112

Sf-Ron during recovery from acute anemia. ............................................................ 112 Sf-Ron prevents premature differentiation of stress erythroid progenitors. ............. 113

Discussion ........................................................................................................................ 115 Materials and Methods ..................................................................................................... 117 Figures .............................................................................................................................. 119 References ........................................................................................................................ 133

Chapter 5 Concluding Remarks and Future Directions ........................................................... 136

Conclusions ...................................................................................................................... 136 Future Studies. ................................................................................................................. 139

Expanding our understanding of inflammation-induced stress erythropoiesis in

mouse and human systems. .............................................................................. 139 Identify the mechanism by which pro-inflammatory cytokines affect SEPs. .......... 139 Relationship of Epo and GDF15. ............................................................................. 140 Spi-C’s role in stress erythropoiesis during acute anemia vs inflammation. ............ 141

References ........................................................................................................................ 144

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LIST OF FIGURES

Figure 2-1 A. Zymosan leads to expansion of stress erythroid progenitors in the spleen. ...... 51

Figure 2-2 B. Zymosan leads to expansion of stress erythroid progenitors in the spleen. ...... 52

Figure 2-2 C. Zymosan leads to expansion of stress erythroid progenitors in the spleen. ...... 53

Figure 2-3 D. Zymosan leads to expansion of stress erythroid progenitors in the spleen. ...... 54

Figure 2-1 E-F. Zymosan leads to expansion of stress erythroid progenitors in the spleen. .. 55

Figure 2-1 G-I. Zymosan leads to expansion of stress erythroid progenitors in the spleen. .. 56

Figure 2-2 A-B. Zymosan induces production of new erythrocytes. ...................................... 57

Figure 2-2 C-E. Zymosan induces production of new erythrocytes. ...................................... 58

Figure 2-3 A. Zymosan induces Erythropoietin expression in the absence of anemia. .......... 59

Figure 2-3 B-C. Zymosan induces Erythropoietin expression in the absence of anemia. ...... 60

Figure 2-3 D-F. Zymosan induces Erythropoietin expression in the absence of anemia. ...... 61

Figure 2-4 A-B. SIRPα expression decreases and results in increased

erythrophagocytosis. ........................................................................................................ 62

Figure 2-4 C-F. SIRPα expression decreases and results in increased

erythrophagocytosis. ........................................................................................................ 63

Figure 2-5 A-C. Erythrophagocytosis leads in increased intracellular heme and changes

in heme-dependent gene expression. ................................................................................ 64

Figure 2-5 D. Erythrophagocytosis leads in increased intracellular heme and changes in

heme-dependent gene expression. .................................................................................... 65

Figure 2-5 E-F. Erythrophagocytosis leads in increased intracellular heme and changes

in heme-dependent gene expression. ................................................................................ 66

Figure 2-6 A-B. Loss of Spi-C affects stress erythropoiesis response after treatment with

zymosan. .......................................................................................................................... 67

Figure 2-6 C. Loss of Spi-C affects stress erythropoiesis response after treatment with

zymosan. .......................................................................................................................... 68

Figure 2-6 D. Loss of Spi-C affects stress erythropoiesis response after treatment with

zymosan. .......................................................................................................................... 69

Figure 2-6 E-F. Loss of Spi-C affects stress erythropoiesis response after treatment with

zymosan. .......................................................................................................................... 70

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Figure 2-6 G. Loss of Spi-C affects stress erythropoiesis response after treatment with

zymosan. .......................................................................................................................... 71

Figure 2-7 A-B. Adoptive Transfer of WT BMDMs or monocytes restores stress

erythropoiesis in Spi-C -/- mice. ...................................................................................... 72

Figure 2-7 C-D. Adoptive Transfer of WT BMDMs or monocytes restores stress


Figure 2-7 E-F. Adoptive Transfer of WT BMDMs or monocytes restores stress


Figure 2-8 A-B. MyD88 is required for increased erythrophagocytosis. ................................ 75

Figure 2-8 C-D. MyD88 is required for increased erythrophagocytosis. ............................... 76

Figure 2-9 A-C. TNFα and IL1-β promote erythroid differentiation under stress

erythropoiesis. .................................................................................................................. 77

Figure 2-9 D. TNFα and IL1-β promote erythroid differentiation under stress

erythropoiesis. .................................................................................................................. 78

Figure 2-9 E-F. TNFα and IL1-β promote erythroid differentiation under stress

erythropoiesis. .................................................................................................................. 79

Figure 2-10 A-C. TNFα increases stress erythropoiesis in human BM cultures. ................... 80

Figure 3-1 A-C. Flexed-tail (f/f) mice have increased mortality after zymosan-induced

inflammation. ................................................................................................................... 93

Figure 3-2 D-E. Flexed-tail (f/f) mice have increased mortality after zymosan-induced

inflammation. ................................................................................................................... 94

Figure 3-2. Glucocorticoid production is critical to surviving zymosan-induced

inflammation. ................................................................................................................... 95

Figure 3-3 A. GDF15-/- and f/f mice have diminished GC production due to decreased

expression of Cyp11b1. .................................................................................................... 96

Figure 3-3 B. GDF15-/- and f/f mice have diminished GC production due to decreased


Figure 3-3 C. GDF15-/- and f/f mice have diminished GC production due to decreased


Figure 3-4 A. f/f mice have lightly elevated levels of pro-inflammatory cytokines. .............. 99

Figure 3-4 B. f/f mice have lightly elevated levels of pro-inflammatory cytokines. ............... 100

Figure 3-5 A-D. GCs promote M2 macrophages to prevent immune lethality. ...................... 101

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Figure 3-5 E. GCs promote M2 macrophages to prevent shock. ............................................ 102

Figure 3-5 F-G. GCs promote M2 macrophages to prevent immune lethality. ...................... 103

Figure 3-6 A-C. Decreased M2 macrophages in f/f mice is due to decreased GC levels. ...... 104

Figure 3-6 D. Decreased M2 macrophages in f/f mice is due to decreased GC levels. ........... 105

Figure 3-6 E-F. Decreased M2 macrophages in f/f mice is due to decreased GC levels. ....... 106

Figure 4-1. Model for Friend virus-induced erythroleukemia. .......................................... 119

Figure 4-2. Sf-Ron expression is upregulated during recovery from

phenylhydrazine. ............................................................................................................ 120

Figure 4-3. Sf-Ron plays a critical role in recovery from PHZ-induced anemia. ............ 121

Figure 4-4. Sf-Ron-/- mice have decreased production of stress BFU-Es after PHZ-

induced anemia. .............................................................................................................. 122

Figure 4-5. Schematic for populations of stress progenitors. ............................................. 123

Figure 4-6 A-B. Sf-Ron-/- stress progenitors are more mature in the absence of Epo

than wild type progenitors. ........................................................................................... 124

Figure 4-6 C-D. Sf-Ron-/- stress progenitors are more mature in the absence of Epo


Figure 4-6 E-F. Sf-Ron-/- stress progenitors are more mature in the absence of Epo


Figure 4-6 G-H. Sf-Ron-/- stress progenitors are more mature in the absence of Epo


Figure 4-6 I-J. Sf-Ron-/- stress progenitors are more mature in the absence of Epo


Figure 4-7 A-B. Sf-Ron-/- stress progenitors have no difference in response to Epo

but are less able to form BFU-Es. ................................................................................. 129

Figure 4-7 C-D. Sf-Ron-/- stress progenitors have no difference in response to Epo

but are less able to form BFU-Es. ................................................................................. 130

Figure 4-7 E. Sf-Ron-/- stress progenitors have no difference in response to Epo but

are less able to form BFU-Es. ........................................................................................ 131

Figure 4-8. Model of the effect of Sf-Ron on stress erythropoiesis. ................................... 132

Figure 5-1. Model of the activation of inflammation-induced stress erythropoiesis. ....... 143

x

LIST OF ABBREVIATIONS

ACTH Adrenocorticotropic hormone

ADX Adrenalectomized

AI Anemia of inflammation

BFU-E Burst forming unit- erythroid

BM Bone marrow

BMDM Bone marrow derived macrophages

BMP4 Bone morphogenetic protein 4

BMP-RE Bone morphogenetic protein responsive element

BMT Bone marrow transplant

CFSE Carboxyfluorescein succinimidyl ester

CFU-E Colony forming unit- erythroid

CLP Common lymphoid progenitor

CMP Common myeloid progenitor

COX-2 Cyclooxygenase 2

DBA Diamond Blackfan Anemia

Epo Erythropoietin

f/f Flexed-tail

Fv2 Friend virus susceptibility locus 2

GC Glucocorticoid

GDF15 Growth/differentiation faction 15

GM Granulocyte-macrophage

GMP Granulocyte-macrophage progenitor

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GPA Glycophorin A

GR Glucocorticoid receptor

HbF Fetal hemoglobin

Hh Hedgehog

Hif2-α Hypoxia inducible factor 2 alpha

HSC Hematopoietic stem cell

IFN-γ Interferon gamma

Ihh Indian hedgehog

IL Interleukin

IL-7Rα Interleukin 7 receptor alpha

IRF-1 Interferon regulatory factor 1

KS Kit+Sca+ cells

LMPP Lymphoid primed multipotential progenitor

LPS Lipopolysaccharide

LSC Leukemia stem cell

PBS Phosphate buffered saline

PGE2 Prostaglandin E2

PHZ Phenylhydrazine

RBC Red blood cell

SCF Stem cell factor

SEDM Stress erythroid differentiation media

SEEM Stress erythroid expansion media

SEP Stress erythroid progenitor

Sf-Ron Short form stem cell derived tyrosine kinase

SIRPα Signal regulatory protein alpha

xii

STAT3 Signal transducer and activator of transcription 3

TLR Toll-like receptor

TNF-α Tumor necrosis factor alpha

TRAIL TNF-related apoptosis-inducing ligand

Vhl von Hipple Lindau

ZIGI Zymosan-induced generalized inflammation

xiii

ACKNOWLEDGEMENTS

This journey has been longer, more stressful, and infinitely more rewarding than I ever

could have imagined when I came to Penn State seven years ago. Thankfully, I have been

surrounded by wonderful and understanding friends and family as well as a group of coworkers

who are unfailingly generous with their time and their thoughts. Nothing I have accomplished

would have been possible without the incredible people in my life who have helped to carry me

through this experience.

First and foremost, I would like to thank my advisor Bob Paulson for teaching me to be a

diligent scientist, to ask the right questions, and that you will almost never mix up your sample

tubes. He has shown me through example what kind of mentor I want to be and has taught me

that it is possible to be a good scientist as well as a good person. Thank you for all the advice,

support, and encouragement and for always believing that I was capable even when I had doubts.

I’ve been lucky to be surrounded by incredible mentors. My committee members (Drs. Sandeep

Prabhu, Pam Hankey-Giblin, Ross Hardison, Zhi-Chun Lai, and Connie Rogers) have provided

me with honest comments and questions about my project, invaluable advice about my career and

future plans, and most importantly, for kindness and support which have made this process less

daunting. I would also like to thank Mary Kennett and Peggy Lorah for always having time to

talk about life, science, and giving the best advice about how to deal with daily struggles.

I also want to thank all past and present members of the Paulson lab for creating such a

supportive and welcoming environment and always being willing to share your expertise and

giving feedback over the years.

Our lab neighbors have helped immensely with making sure this project got off the

ground and providing time, reagents, troubleshooting, and laughs over the years. Mike Quickel,

Ashley, Shay, Emily Finch, and Laura Goodfield have been the best colleagues that anyone could

xiv

ask for. They have sacrificed hours of their time to help to me harvest and process samples or

even just to provide some company on a long night. I will miss our milkshake Mondays and

walks to get tea at ABP – many an experiment would not have happened without our science

talks en route.

Finally, I would like to thank all of my friends and family. Having a support system

during graduate school is so important and I have had one of the best. Thank you to my friends in

State College for all the laughs and non-science activities. You made sure I didn’t spend every

second in the lab or being a true introvert curled up at home by myself. Thanks for all the Friday

nights, five dollar Tuesdays, and game nights that kept me sane. Thanks to my 997 family, Will

and Arslan, for taking me in at the end of this journey and being my family away from home.

You two are some of the kindest souls I’ve ever met and are so near and dear to me. Thanks for

keeping me smiling and taking such good care of me over the last two years. To my friends at

home, thank you for not giving up on me – even when I’m not the most communicative. Thank

you for always making the long periods of time between phone calls or visits seem like no time

has passed.

To my Uncle David, you don’t know how much your phone calls and visits to State

College have meant to me. I was so lucky to have come to Penn State for graduate school and

have you and Sue only a couple of hours away. I’ll miss being so close to Pittsburgh and to you.

My grandparents Arthur and Eloween Lewis have been some of biggest fans. They’ve always

given me a soft place to land when going home and welcomed me with open arms, and a special

thanks to Granddad for always showing me that learning is a lifelong pursuit. Thank you Granny

B (Bess Bennett) for pushing me to pursue my dreams because women can be more than nurses,

secretaries, or teachers. Last and definitely not least, I can’t thank my parents enough. You have

always supported me, believed in my dreams and capabilities, and loved me unconditionally.

Thank you for teaching me to be strong and independent. Your unwavering encouragement has

xv

made me believe I can accomplish anything I put my mind to and I owe everything I am today to

both of you.

1

Chapter 1

Introduction

Chronic inflammation inhibits erythropoiesis due to increased levels of pro-

inflammatory cytokines, which directly affect the proliferation of erythroid progenitors,

decreasing the half life of circulating erythrocytes, and increasing hepcidin levels, which

decreases iron availability.1–8 These events culminate in the establishment of anemia,

often referred to as anemia of inflammation (AI). AI is a secondary complication that

often complicates the treatment of the underlying disease and has proven difficult to

effectively treat.9 Developing effective therapeutic strategies will require an

understanding of how inflammatory cytokines affect both steady state and stress

erythroid progenitors. In the absence of disease, steady state erythropoiesis maintains

oxygenation of tissues, but in response to inflammation it is insufficient to maintain

homeostasis. Stress erythropoiesis is able to fill this need during anemic stress, but there

is evidence to suggest that in some situations it may do more than maintain homeostasis.

In this review, we will discuss some recent findings demonstrating that stress erythroid

progenitors are not inhibited by pro-inflammatory cytokines and may contribute to

immunomodulatory effects that prevent lethal immune activation during infection. This

effect may be driven by signals that regulate stress erythropoiesis and modulate immune

responses such as glucocorticoid production which both promote stress erythropoiesis to

alleviate inflammation-induced anemia in addition to resolving inflammation.

2

Erythropoiesis

I. Murine Stress Erythropoiesis

Steady state erythropoiesis is a homeostatic process taking place primarily in the

bone marrow. Its role is to replace old or damaged erythrocytes which are removed from

circulation, and in individuals with normal erythropoiesis, the number of newly produced

erythrocytes is equivalent to those which are phagocytosed by splenic macrophages.10

However, steady state erythropoiesis is unable to produce a sufficient number of new

erythrocytes in times of acute anemic stress, such as hemorrhage or hemolysis. In those

instances, a dedicated response pathway is necessary to create enough new erythrocytes

to restore tissue oxygenation and homeostasis.

This dedicated response is known as the BMP4-dependent stress erythropoiesis

pathway. It provides a robust response to acute anemia by utilizing specialized erythroid

progenitors, which respond to signals in the microenvironment to rapidly produce

sufficient numbers of erythrocytes for survival. Stress erythropoiesis is best understood in

the mouse. Murine stress erythropoiesis is extramedullary, occurring in the spleen and

liver of adults and the fetal liver during embryonic development. Stress erythropoiesis

uses resident progenitors in the spleen, and following recovery from anemia CD34+KSL

stem cells migrate from the bone marrow to repopulate the spleen. Hedgehog (Hh) and

bone morphogenetic protein 4 (BMP4) signaling in the spleen specifies CD34+KSL stem

cells as stress erythroid progenitors (SEPs) which are both self-renewing and erythroid

restricted (Figure 1-1).11–13 These specified progenitors are self-renewing and are unable

to differentiate in the absence of key signals. Work from our lab has also shown that Wnt

3

signaling by splenic macrophages is crucial in maintaining self-renewing SEPs, which

have limited BFU-E potential (Jie Xiang, unpublished).

Unlike steady state progenitors, stress progenitors require signals not typically

associated with erythropoiesis in order to produce mature erythrocytes. These signals

include Hh, which is most likely Indian hedgehog (Ihh) in the adult spleen, BMP4, stem

cell factor (SCF), and growth differentiation factor 15 (GDF15), in addition to

erythropoietin (Epo) and hypoxia. These factors work in concert to ensure that the stress

erythropoiesis pathway is only activated in times of need.11–17 We demonstrated in vivo

and in vitro that loss of any one of these factors results in a significant reduction in the

number of stress BFU-Es. Loss of SCF in vitro resulted in the most significant reduction

of stress BFU-Es with murine bone marrow, but BMP4 had the most significant effect on

colony forming ability in human cultures.12–14

Stress erythropoiesis occurs in two distinct stages. In response to anemic stress,

progenitors cells in the spleen expand but are unable to differentiate. During this first

expansion stage SCF, Shh, BMP4 and GDF15 promote self-renewal in stress progenitors

and allow for expansion of the population to ensure sufficient numbers of new

erythrocytes are produced once differentiation begins.11–14 However, it is unclear which

signal initiates the expansion of SEPs. At this point, cells are unable to differentiate into

mature erythrocytes and have limited capability to form BFU-Es.14 They are capable of

being serially transplanted and maintain their erythroid restriction.14,15

Expansion of SEPs continues until tissue hypoxia induces an increase in serum Epo,

which acts as a transition signal that drives the progenitors from a state of self-renewal to

differentiation. Unpublished data from out lab suggests Epo acts directly on splenic

4

macrophages, resulting in a shift from Wnt production to prostaglandin E2 (PGE2),

which promotes differentiation of SEPs (Jie Xiang, unpublished). Analysis of murine

SEPs in vitro demonstrated Epo and hypoxia act together to promote erythroid

differentiation. Epo alone is sufficient to promote the transition to differentiation, but

hypoxia potentiates the response by promoting the expression of BMP4 as well as other

genes involved in stress erythropoiesis through hypoxia-inducible factor-2α, resulting in a

more robust stress response.14,17 Following the transition, SEPs proceed through erythroid

differentiation to produce a wave of new erythrocytes.

Using a two-stage in vitro culture system to recapitulate the development of stress

progenitors, Xiang et al. showed that cells cultured in the presence of BMP4, GDF15,

SCF and Shh (referred to as stress erythroid expansion media or SEEM) are primarily

CD34+CD133+KS cells with limited stress BFU-E potential. However, the addition of

Epo and 2%O2 (referred to as stress erythroid differentiation media or SEDM) to culture

conditions results in nearly all cells transitioning to CD34-CD133-KS cells, which have

significantly greater potential to form BFU-Es. Gene expression analysis showed that

CD34+CD133+KS cells showed higher levels of expression for genes associated with

self-renewal, such as Yap1 and Pu.1, whereas CD34-CD133-KS cells had increased

expression of genes associated with erythroid differentiation, such as GATA1 and

GATA2. These findings were also shown to be true during recovery from bone marrow

transplant in vivo.14

Stress erythropoiesis is a highly regulated process with numerous signals working

together to ensure that the pathway is only activated in times of need. In addition to

limiting the activation of the stress erythropoiesis pathway, these signals create a

5

response system with tightly regulated expansion of a transient progenitor population

followed by synchronous differentiation and eventually migration of new progenitors

from the bone marrow to replenish the depleted SEP population.

II. Human Stress Erythropoiesis

While stress erythropoiesis has been extensively studied and characterized in

murine models, human stress erythropoiesis is less well understood, due largely to the

fact that studies examining erythropoiesis in humans are both limited and complicated by

underlying disease. It is not possible to directly examine the mechanisms of human stress

erythropoiesis. However, recent studies of human unfractioned bone marrow cells in vitro

has shown that these human cells can differentiate in a manner similar to murine bone

marrow progenitors and are capable of generating stress BFU-Es when grown under

stress conditions.14 Also, observational data from peripheral blood of patients under acute

anemic stress and primate studies using phenylhydrazine-induced anemia have defined

general characteristics of human stress erythropoiesis.18–25

Human stress erythropoiesis more closely resembles fetal erythropoiesis than

normal adult erythropoiesis. It has been observed that there is an increase in fetal

hemoglobin (HbF) expression in patients during recovery from erythropoietic stress, such

as sickle cell anemia, beta-thalassemia, recovering from bone marrow transplants (BMT),

and acute anemic syndromes.21–23,26 Mice lack the γ-globin gene found in humans and do

not produce fetal hemoglobin. Thus it is not possible to make a direct comparison of the

murine system with these observations in humans and non-human primates. However,

6

murine stress progenitors do express high levels of the embryonic globin, βh1, mirroring

the observations in humans.

There are studies which provide some evidence linking the murine BMP4-

dependent stress erythropoiesis pathway with human stress erythropoiesis and fetal

hemoglobin production. Human unfractionated bone marrow cells cultured under hypoxic

conditions generate large numbers of stress BFU-Es and produce more HbF+ F-

cells.14,24,25 Cells in these cultures also preferentially expanded the population of

Kit+CD34+CD133+ cells when grown in SEEM conditions and, when transitioned to

SEDM, were primarily c-Kit+CD34-CD133-, mirroring what occurs in murine

progenitors.14 These cells also expressed lower levels of the fetal hemoglobin repressor,

BCL11a, compared with steady state human progenitors, which explains the increased

expression of γ-globin.14 Murine stress progenitors express higher ratios of βh1:βmajor

and εy:βmajor. In these cultures, decreased levels of BCL11a were observed, which is

consistent with previous work showing that BCL11a regulates murine embryonic globin

(βh1 and εy) in a similar fashion to the regulation of γ-globin in humans.14,27 In vivo

human stress erythropoiesis is associated with increased HbF. Sickle cell anemia and β-

thalassemia patients exhibit increased numbers of CD34+ Kit+GPA+ cells that when

cultured generate HbF+ cells.25,28 These cells have markers in common with the murine

SEPs, but further analysis will be needed to confirm the similarities.

Additionally, examination of bone marrow and peripheral blood from patients

with sickle cell anemia revealed they have significantly more cells which express CD34

and glycophorin A (GPA) on their surfaces than normal individuals, and in culture, these

patient cells produced more HbF+ F-cells.24,25 The CD34+Kit+GPA+ cells observed in

7

patient samples may be similar to the stress progenitors observed when culturing CD34+

cells under murine stress conditions.

Studies with non-human primates also support the idea that stress erythropoiesis

is similar to fetal erythropoiesis. When studying recovery from phenylhydrazine-induced

anemia in baboons, there was an increase in the production of fetal hemoglobin and

erythroid progenitors in the bone marrow adopt a fetal globin synthesis pattern.18–20

Baboons were either treated with phenylhydrazine (PHZ) or exposed to low oxygen

similar to high altitude in a hyperbaric chamber. In both cases, there was an increase in

reticulocytes and in circulating HbF+ cells.19,20 Similar to data in patients, baboons also

exhibit variability in the increase of HbF following anemic stress due to increased

heterogeneity in the population, but HbF responses were reproducible in each animal

following repeated anemic stress.18,19 Bone marrow was isolated from baboon fetuses or

baboons following PHZ treatment and cultured in the presence of Epo to analyze the

globin synthesis pattern. Both PHZ-treated bone marrow and fetal bone marrow had

similar patterns of globin synthesis, producing primarily γ-globin.18

The data suggest that human stress progenitors observed in vitro may be the same

cells observed in patients with sickle cell anemia and are comparable to murine stress

progenitors. While the spleen is the primary site of stress erythropoiesis in the mouse, it

is thought that in humans stress erythropoiesis occurs in the bone marrow, although the

exact site of human stress erythropoiesis is unclear. In spite of this potential difference,

murine stress erythropoiesis shares many characteristics with human stress erythropoiesis

and could provide an excellent foundation for future studies into human stress

8

erythropoiesis. However, more work is still needed to fully understand human stress

erythropoiesis before it can be fully utilized as a therapeutic target.

Infection, Inflammation, and Erythropoiesis

I. Infection and Steady State Erythropoiesis

Stress erythropoiesis has primarily been studied as a response to severe anemia

and hemorrhage where it acts as a final effort to produce enough erythrocytes to survive

the anemic crisis. However, there are other situations where an alternative erythropoiesis

pathway would be required. Typically, hematopoietic stem cells (HSCs) in the bone

marrow progress through a series of steps to form all cells derived from the HSC and

work primarily to maintain homeostatic levels of each cell type (Figure 1-2), but this

process can become skewed toward one lineage or another under stress conditions. For

example, during bacterial infections there is a hematopoietic shift to produce increased

numbers of myeloid cells to combat and clear the infection.29–32 Increased levels of pro-

inflammatory cytokines also result from inflammation during infection, which leads to

the suppression of bone marrow erythropoiesis and a shortened half-life for circulating

red cells.1–4 Both of the conditions can result in the development of anemia, but this

inhibition of erythropoietic activity in the marrow cannot be maintained indefinitely. In

addition to direct suppression of bone marrow erythropoiesis, interleukin 6 (IL-6) induces

hepcidin and results in decreased iron availability, which is necessary for erythroid

maturation.5–8 Erythrocytes are critical to ensuring that all tissues are oxygentated

9

sufficiently and the survival of the organism. In the event of infection, it is necessary to

increase production of myeloid cells to resolve and survive the infection, but the need for

erythrocytes is not diminished by the infection. Consequently, there is a need for a

secondary method of producing erythrocytes until bone marrow erythropoiesis can be

reestablished.

There is a dramatic shift in hematopoietic cell production in the bone marrow in

response to infection, favoring the myeloid lineage over lymphoid and erythroid.

Hematopoietic stem cells (HSCs) express TLR2 and TLR4 on their surfaces, and

stimulation of these TLRs results in a push toward monocyte and macrophage

differentiation and a loss of lymphoid potential, demonstrated by acquisition of the

surface markers of F4/80, Mac-1, and Gr-1 and no detectable B220. HSCs exposed to

TLR ligands also had decreased levels of GATA2 and SCL mRNA expression, consistent

with myeloid differentiation.29 Granulocyte-monocyte progenitors (GMPs) preferentially

differentiate into monocytes/macrophages in response to TLR stimulation, whereas

committed lymphoid progenitors are pushed to become dendritic cells and B

lymphopoiesis is suppressed.29 Interferon gamma (IFN-γ) stimulation during infection

results in increased proliferation of long-term quiescent HSCs in the bone marrow,

contributing to the replenishment of the pool of committed progenitors.30

Traditionally, it has been thought that the first lineage restriction of hematopoietic

stem cells is the commitment to either the myeloid or lymphoid lineages, resulting in the

production of common myeloid progenitors (CMPs) or common lymphoid progenitors

(CLPs) (Figure 1-2).33,34 However, there is evidence to suggest that the first step of

lineage commitment is a division to either the megakaryocyte-erythroid lineage or

10

lymphoid-primed multipotent progenitors (LMPPs), which are primed for the lymphoid

lineage but also have substantial granulocyte/macrophage (GM) potential.34,35 Belyaev et

al. demonstrated P. chabaudi infection results in the appearance of an atypical population

of Lin-IL-7Rα+Kithi myelolymphoid progenitors and contraction of more typical

lymphoid and myeloid subsets such as CLPs, GMPs, and CMPs. These Lin-IL-7Rα+Kithi

progenitors express markers typical of CLPs but also a subset of markers which identify

short-term reconstituting HSCs, defining a unique early hematopoietic progenitor that is

neither CLP, CMP or LMPP.31 Lin-IL-7Rα+Kithi progenitors have no erythroid potential

and robust myeloid potential with cells generating either myeloid-only, lymphoid-only, or

mixed myeloid and lymphoid progeny in vitro.31 These progenitors also showed a

considerable bias toward myeloid potential at the transcriptional level, expressing high

levels of the master regulator of myelopoiesis Cebpa as well as other genes typically

expressed in myeloid progenitors, such as Mpo, Tal1, GATA1 and GATA2.31 Lin-IL-

7Rα+Kithi progenitors transferred into uninfected mice primarily adopted a myeloid fate,

and when transferred into mice infected with P. chabaudi resulted in less parasitemia,

suggesting the myeloid cells produced by these progenitors play an important role in

parasite clearance.31 It was also shown that IFN-γ signaling was critical in the expansion

of Lin-IL-7Rα+Kithi myelolymphoid progenitors.31

In addition to pushing cells toward the myeloid lineage, pro-inflammatory

cytokines produced during infection also directly and indirectly inhibit bone marrow

erythropoiesis. TNF-α suppresses bone marrow erythropoiesis by inhibiting proliferation

and differentiation of erythropoietic progenitors.1,3 In vitro studies of CD34+ bone

marrow progenitors have shown that BFU-E colony formation induced by either Epo

11

alone or Epo in combination with IL-3 is inhibited in the presence of TNF-α. Not only is

TNF-α capable of suppressing BFU-E formation, it also promotes the formation of

granulocyte/macrophage colonies.1 CD34+ progenitor cells cultured with TNF-α and

IFN-γ increased the expression of Fas, which profoundly affected cell function and

viability. Stimulation of Fas both reduced the colony forming ability of CD34+ bone

marrow cells and induced apoptosis. Fas activation was shown to have a synergistic

effect with TNF-α and IFN-γ in suppression of erythropoiesis.3

IFN-γ acts both directly and indirectly on erythropoiesis and mature erythrocytes.

One of the primary ways in which IFN-γ directly suppresses bone marrow erythropoiesis

is by inducing production of TNF-related apoptosis-inducing ligand (TRAIL). Studies

have shown that production of TRAIL negatively affects the growth of erythroid cells by

selectively inducing apoptosis in early erythroid cells. This response reduces colony

forming ability without affecting either monocytic, granulocytic, or megakaryocytic

lineages. 2 Other work has shown that the presence of IFN-γ also inhibits the ability of

erythroid cells to differentiate by inducing interferon regulatory factor-1 (IRF-1), which

results in an increase in PU.1 expression and inhibition of erythroid differentiation.4 In

addition to acting directly on erythroid precursors in the bone marrow, IFN-γ is also

capable of affecting circulating erythrocytes. Transgenic mice overexpressing CD70, a

TNF superfamily member which induces production of IFN-γ-producing T-cells in

response to costimulation with CD27, have chronic overproduction of IFN-γ resulting in

higher turnover of erythrocytes due to increased phagocytosis by splenic macrophages.4

Inflammatory cytokine IL-6 affects erythropoiesis indirectly by promoting the

expression of hepcidin. Hepcidin controls iron availability directly by binding to the iron

12

exporter ferroportin and targeting it for degradation.36 Treating hepatocyte cell lines with

IL-6 in the presence or absence of cyclohexamide revealed that IL-6 induces hepcidin

expression directly, and this induction requires activation of signal transducer and

activator of transcription 3 (STAT3).5–7 Treating mice with turpentine to induce

inflammation results in increased hepcidin expression and lower serum iron in wild type

mice, and these results were abrogated in IL-6 -/- mice, which is consistent with the idea

that IL-6 induces hepcidin expression during inflammation and promotes hypoferremia.8

Wild type mice treated repeatedly with turpentine over a two week period develop a

microcytic anemia, exhibiting decreased hematocrit, hemoglobin levels, and mean

corpuscular volume (MCV). Hepatocyte-specific STAT-3 -/- mice, which are unable to

induce hepcidin expression in response to increased levels of IL-6, do not develop anemia

when repeatedly treated with turpentine and have higher hematocrits, hemoglobin and

MCV compared with WT animals.37 Increased hepcidin reduces the availibity of iron for

maturing erythrocytes by limiting iron-recycling by macrophages and also decreasing

dietary absorption of iron, resulting in iron-restricted erythropoiesis and contributing to

the development of anemia during inflammation.38,39 Recent studies have shown an IL-6

dependent induction of hepcidin mRNA in both mice and primary human hepatocytes in

response to pathogens, such as S. pneumoniae and mouse-adapted influenza A virus

(PR8), and pathogen-derived molecules like PAM3 and LPS.40 Using IL-6 -/- mice and

Hamp -/- mice treated with heat-killed Brucella abortus, recent studies have

demonstrated that loss of either hepcidin or IL-6 also results in milder anemia and

quicker recovery than in wild type animals.41,42 Together, these data demonstrate that

13

induction of hepcidin by IL-6 and STAT3 signaling is required for the establishment of

AI.

Additionally, BMP signaling has also been shown to play a role in hepcidin induction

during inflammation and the development of anemia.6,43–45 Treating with BMP2 is

capable of inducing hepcidin through the BMP responsive element (BMP-RE) located in

the hepcidin promoter in hepatoma-derived cells (HepG2, Hep3b, and Huh7).43 Culturing

Hep3b cells with the BMP inhibitor dorsomorphin reduces basal level expression of

hepcidin and blocks IL-6 mediated induction, suggesting a critical role for BMP signaling

in increasing hepcidin expression during inflammation.44 Injecting zebrafish with iron-

dextran results in increased phosphorylation of Smad1/5/8 and increased expression of

hepcidin mRNA.44 Treatment with BMP inhibitors (LDN-193189, Noggin, and ALK3-

Fc) also blocks induction of hepcidin by IL-6 in HepG2 cells. Treating C57BL/6 mice

with LDN-193189 also attenuated hepcidin induction following injection of IL-6.45

Inhibition of BMP signaling also blocked the induction of hepcidin in vivo after injection

of turpentine in mice. Mice pre-treated with LDN-193189 have increased Hb, MCV and

serum iron compared with wild type mice after three weeks of turpentine injections,

demonstrating that inhibition of BMP signaling attenuates AI in vivo.45

Activation of TLR2 and TLR6 has also been shown to reduce iron availability by

decreasing ferroportin mRNA and protein levels in a hepcidin-independent manner.

Stimulation of these TLRs in vivo was sufficient to induce hypoferremia in the absence of

increased levels of hepcidin, suggesting that there are multiple ways in which the host

reduces iron availability during infection.46

14

As in mice, hepcidin is the primary regulator of iron homeostasis in humans. Both

nonsense and frameshift mutations in hepcidin have been associated with juvenile

hemochromatosis in humans.47 Studies of patients with anemia of inflammation (AI)

revealed increased hepcidin levels and decreased serum iron, consistent with the findings

in mice, and treatment of patients with IL-6 also increased levels of circulating

hepcidin.8,48 Limited iron availability is a concern when treating AI, making the IL-

6/hepcidin pathway a potential therapeutic target. While targeting IL-6 or hepcidin for

therapeutic strategies for anemia of inflammation (AI) would be beneficial for improving

erythropoiesis, this could negatively impact disease prognosis by increasing the

availability of iron for pathogens and decreasing inflammatory cytokines which aid in

eliminating the infection.49

II. Effects of Infection and Inflammation on Stress Erythropoiesis

Much of the current research has focused on how inflammation, particularly pro-

inflammatory cytokine production, affects bone marrow erythropoiesis. However, little is

known about the effect of inflammation on the stress erythropoiesis pathway. Zymosan is

derived from the cell wall of Saccharomyces cerevisiae and stimulates TLR2, inducing a

potent inflammatory response. Millot et al. have shown that the numbers of bone marrow

erythroid cells decreased after treatment with zymosan, and they did not change in

response to Epo treatment.50 However, erythroid precursors in the spleen increased

following zymosan treatment and injections of Epo resulted in even higher numbers of

erythroblasts. Injection of Epo created a prolonged stress erythropoietic response which

15

was able to partially correct anemia induced by inflammation. Whereas IFN-γ inhibits the

formation of bone marrow BFU-Es, there was no change in the ability of splenic

erythroid cells to form BFU-Es in the presence of IFN-γ.50 These observations

demonstrate that steady state and stress erythropoiesis respond in different ways to

inflammation and presence of pro-inflammatory cytokines. Inducing inflammation with

heated-killed Brucella abortus also shows a dramatic increase in splenic erythropoiesis

despite inhibition of bone marrow erythropoiesis.41,42

Whereas steady state erythroid progenitors decline in response to inflammatory

stimuli, an expansion of erythroid progenitors in the spleen has been noted during some

infections, such as Salmonella and malaria, as well as in models of chronic inflammatory

conditions, such as ulcerative colitis.51–56 Previously, this increase in spleen size was

attributed to an expansion of lymphocytes within the spleen, but in actuality, the most

significant increase is observed in erythroid progenitors.51–54,57 Malaria infections in mice

showed drastic changes to splenic architecture which occur primarily in the red pulp.51,52

There is a large and transient expansion of hematopoiesis. At days 6 and 8 during malaria

infection, there is an increase in CD34+ precursors found in the spleen, and at day 10, an

increase in islands of erythroid precursors is found in the red pulp.51,52 Splenic BFU-Es

and CFU-Es are significantly increased as early as day 6 after infection with Plasmodium

chabaudi chabaudi.57

In the case of Salmonella infection, erythroid precursors in the spleen undergo the

largest increase, expanding from approximately 20% of the total population to 80%.53 In

addition to an increase in early erythroid progenitors, there is also an expansion of F4/80

red pulp macrophages, which are important in erythrophagocytosis and iron recycling as

16

well as phagocytosis of lymphocytes.54 Furthermore, red pulp macrophages may play a

role in erythroid maturation by forming erythroblastic islands. Splenomegaly and the

expansion of erythroid cells is not seen when MyD88/TRIF-deficient mice infected are

with Salmonella, indicating this is a MyD88-dependent response and TLR signaling may

drive stress erythropoiesis during infection. MyD88/TRIF deficient animals also had

increased bacterial loads in the spleen at day 8 post-infection.53

III. Immunomodulation by Stress Progenitors

Stress erythropoiesis may also provide critical immunomodulatory effects in

addition to compensating for loss of erythropoiesis in the bone marrow. Recent work has

shown that CD71+ erythroid cells are expanded in the spleens of neonatal mice and

exhibit immunosuppressive abilities.58 Neonatal mice are more susceptible to L.

monocytogenes infection than adult mice. They exhibited increased bacterial levels in

both spleen and liver when compared to adult animals and they show little to no increase

in TNF-α levels. CD71+Ter119+ cells comprise nearly 70% of neonatal spleens and

depletion of this population resulted in a loss of immunosuppression in response to L.

monocytogenes.58 Co-culture of adult splenocytes with purified CD71+ neonatal erythroid

cells suppressed the immune response of the adult cells upon exposure to heat-killed L.

monocytogenes. The authors proposed that in neonates immunosuppression by immature

erythroid cells allows for microbial colonization of the gut following birth. However, it

remains to be seen if stress erythroid progenitors could play a similar role in adults during

infection. CD71+ cells from phlebotomized adults did not appear to confer the same

17

immunosuppressive qualities but it is unclear why that is.58 Arginase-2 (Arg-2) activity

was lower in the adult CD71+ cells and inhibition of Arg-2 activity in neonatal cells

abolished the immunosuppressive qualities and increased levels of TNF-α.58

CD71, the transferrin receptor, is considered a definitive marker of erythroid

precursors, meaning that CD71+ cells encompass a large heterogeneous population of

cells at various stages of maturity.15 In phlebotomized adults, it is possible that the

immunosuppressive progenitors were not enriched in the CD71+ population at those

times. Stress erythropoiesis is a rapid and highly synchronous response where SEPs

expand until Epo signaling results in a transition to differentiation, and these two stages

are phenotypically and functionally distinct but both are CD71med/lo.14 Further

characterization of neonatal cells with immunosuppressive capabilities could provide

more phenotypic markers to aid in identifying a corresponding population in the adult

spleen. While the current studies suggest that this is a unique property of neonatal

erythroid progenitors, it is interesting to speculate that the expansion of the erythroid

population in response to infection in adult mice may play a dual role by producing

erythrocytes to counteract anemia as well as dampening the immune response to prevent

lethality.

IV. Glucocorticoids and Erythropoiesis

Similarly, glucocorticoids could also have dual functionality in response to

infection or chronic inflammation. Glucocorticoids have been shown to both promote

erythropoiesis as well as modulate the immune response.59–65 Glucocorticoids promote

18

the expansion of erythroid progenitors in vitro.59–62 Golde et al. demonstrated that

exposure of erythroid progenitors from murine fetal liver as well as murine and human

adult progenitors to glucocorticoids resulted in increased colony forming ability.59 Early

work has shown that proerythroblasts cultured with Epo and glucocorticoids enhances

proliferation when compared to cells cultured with Epo alone.61 In fact, culture of

purified human CD34+ cells with Epo, SCF, and dexamethasone resulted in the sustained

proliferation of cells through several rounds of re-plating. When analyzed by flow

cytometry, the majority of cells were CD71+ erythroid cells with only 5-7% of the

cultures belonging to myeloid or lymphoid lineages, indicating a selective proliferation of

the erythroid lineage from CD34+ cells cultured with Epo, SCF, and dexamethasone.60

Addition of a glucocorticoid receptor (GR) antagonist resulted in a 10-fold reduction in

the proliferative ability of human CD34+ cells as well as a reduction in the number of

colony forming units, demonstrating that glucocorticoids are essential for increased

proliferation of erythroid progenitors. 60 Recent work has shown purified BFU-E cells

from fetal liver cultured in the presence of dexamethasone have enhanced self-renewal

and increased CFU-E output, and the RNA-binding protein ZFP36L2, a transcriptional

target of the glucocorticoid receptor, is critical in enhancing BFU-E self-renewal.66,67 It

has also been shown that PPAR-α agonists GW7647 and fenofibrate synergize with

dexamethasone to further increase erythroid output in vitro by enhancing PPAR-α

occupancy at GR binding sites in BFU-Es.68 Treating mice with GW7647 also enhanced

recovery from phenylhydrazine-induced anemia.68

Glucocorticoids also are capable of promoting stress erythropoiesis in vivo.69

Bauer et al. showed that erythroid proliferation was severely impaired in either fetal liver

19

suspensions taken from GRnull/null mice or in wild type fetal liver cells treated with a GR

antagonist.69 Cells from GRnull/null fetal livers were also smaller, differentiated, and more

hemoglobinized than their wild type counterparts, demonstrating that in vivo

glucocorticoids are required for maintaining the proliferation of erythroid progenitors.69

Adult GRdim/dim (GR dimerization deficient) mice treated with phenylhydrazine to induce

anemia showed no increase the number of CFU-Es in the spleen or increase in the

percentage of Kit+ cells, indicating that glucocorticoid production is also required for

expansion of erythroid progenitors during stress erythropoiesis.69 Additionally, chronic

physiological stress induces glucocorticoid production and also increases production of

Epo.70 There was also a significant increase in both colony-forming ability and the

number of Ter119+ erythroid progenitors in the spleens of chronically stressed animals,

indicating the chronic stress induces stress erythropoiesis.70,71

Further evidence for the role of glucocorticoids in erythropoiesis is the use of

synthetic corticosteroids, such as prednisone, to treat erythroid hypoplasias, particularly

Diamond-Blackfan anemia (DBA).62,72–75 DBA is, in many cases, caused by

haploinsufficiency of ribosomal subunits which results in bone marrow failure.

According to the DBA registry, approximately 80% of DBA patients are initially

responsive to prednisone treatment, which alleviates anemia and increases hemoglobin

levels.73–75 In vitro culture of erythroid progenitors from the marrow of DBA patients has

also shown that prednisone increases the proliferative ability and the number of CFU-Es

and BFU-Es produced by these progenitors in some patients.72

20

V. Glucocorticoids as Immunomodulators

Glucocorticoids, in addition to promoting erythroid development, also are critical

immunomodulators during infection. Glucocorticoids have been shown to modulate the

immune response during infection and in a model of septic shock by exerting

immunosuppressive effects on various immune cells.63–65 T-cell-specific depletion of the

glucocorticoid receptor (GR) results in higher levels of pro-inflammatory cytokines, such

as IFN-γ and TNF-α, during Salmonella infection and a significant increase in

mortality.64 This increase in mortality was the result of increased levels of

cyclooxygenase 2 (COX-2), a mediator in the production of both pro- and anti-

inflammatory prostaglandins. Treatment with COX-2 inhibitors prevented lethal immune

activation and reduced inflammation in mice lacking GR.64 Similarly, depletion of GR

from dendritic cells has also been shown to result in immune dysregulation. Using an

LPS-induced model of septic shock, selective depletion of GR in dendritic cells (GRCD11c-

Cre) resulted in increased production of pro-inflammatory cytokines and increased

mortality.65 The GRCD11c-Cre animals exhibited increased IL-12 expression. Treatment

with neutralizing IL-12 antibody prevented lethality. Activation of the GR in dendritic

cells therefore suppresses IL-12 production to prevent lethal immune activation.65

Glucocorticoids also affect macrophage responses by promoting an M2 activation,

which is generally anti-inflammatory and associated with tissue remodeling and

resolution.76–80 Dimerization of the GR is required for alternative activation of

macrophages by glucocorticoids and survival of LPS-induced septic shock, which

suggest the activation of transcription by GR plays a role. In the presence of

21

glucocorticoids, macrophages in GR dim/dim mice do not exhibit anti-inflammatory effects

and lack surface markers or gene expression profiles indicative of M2 macrophages.80 In

addition to their role in resolving inflammation, data from our lab suggests M2

macrophages may also be important in establishing a niche during stress erythropoiesis

when there are dramatic changes in splenic architechture to accommodate the expansion

of SEPs and erythroid-macrophage interactions necessary for erythroid differentiation

and maturation. Taken together, this suggests that glucocorticoids are required during

infection to suppress the immune response, maintaining a balance between appropriate

immune responses and immunolethality, as well as playing an essential role in the

proliferation of erythroid progenitors.

Conclusions

Evidence in the field suggests that stress erythroid progenitors are activated and

capable of expanding in response to inflammation and infection. One reason for this

could be to preemptively combat the onset of anemia, which results from inhibition of

bone marrow erythropoiesis by pro-inflammatory cytokines. Alternatively, there is data

to suggest that these progenitors are capable of exerting immunomodulatory effects, and

this could be driven by the production of glucocorticoids, which promote erythropoiesis

as well as dampen the immune response to limit cytotoxicity. This observation creates a

number of interesting questions aimed at understanding the role of stress progenitors in

maintaining the microenvironment and balancing the immune response during infection.

22

An understanding to how erythroid cells effect the immune response will be critical in

moving forward with studies of the anemia of inflammation.

Bone marrow erythropoiesis has been shown to be inhibited in the presence of

pro-inflammatory cytokines and a shift toward production of myeloid immune

responders, critical to resolving inflammation.1–3,31,34,81 However, continued production

of erythrocytes is critical in maintaining sufficient oxygenation and homeostasis. The

expansion of the erythroid compartment in the spleen during Malaria and salmonella

infections suggests active stress erythropoiesis51–53. We propose that activation of stress

erythropoiesis acts as a compensatory mechanism to maintain production of erythrocytes

until there is a return to homeostasis in the hematopoietic niche in the bone marrow. The

work presented here will outline a novel mechanism for the activation of stress

erythropoiesis in response to inflammatory stimuli and show that this burst of stress

erythropoiesis delays the onset of anemia during inflammation.

23

Figures

Figure 1-1. A model of stress erythropoiesis.

Schematic of stress erythropoiesis. Signals required at each stage are shown below

progenitors. Progenitor cells are shown migrating from the bone marrow to the spleen

where they become committed stress progenitors. Signals in the spleen maintain cells in a

state of self-renewal until Epo signaling and hypoxia switch SEPs from expansion and

self-renewal to differentiation.

24

Figure 1-2. Hematopoietic Lineage.

Schematic of hematopoietic lineage showing lineage commitments from LT-HSCs to

mature cell types. This is a traditional depiction of cell fate decisions. However, recent

evidence suggests this model is not always an accurate representation of stem cell and

progenitor decisions. Adapted from Passegue et al. (2003).

25

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31

Chapter 2

Zymosan-induced generalized inflammation activates stress erythropoiesis

through a novel mechanism.

Introduction

Anemia of inflammation (AI) is common among patients with persistent inflammatory

conditions, such as chronic infections, cancer, and autoimmune disorders. Chronic inflammation

results in over production of pro-inflammatory cytokines to combat the infection, and many pro-

inflammatory cytokines are known to inhibit steady state bone marrow erythropoiesis. In

particular, tumor necrosis factor alpha (TNF-α) and interferon gamma (IFN-γ) inhibit the growth

and differentiation of erythroid progenitors in the bone marrow as well as affecting the lifespan of

mature circulating erythrocytes.1–4 In addition to direct inhibition of bone marrow erythropoiesis

during inflammation and infection, there is a shift toward the production of myeloid cells.

Stimulation of Toll-like receptors (TLRs) on hematopoietic stem cells (HSCs) results in increased

monocyte and macrophage production and a loss of lymphoid potential.5 Interlukin-1 (IL-1) has

also been shown inhibit lymphopoiesis and erythropoiesis and chronic exposure to IL-1 drives

HSCs toward myelopoiesis.6 Additionally, pro-inflammatory cytokine interleukin 6 (IL-6)

induces expression of hepcidin, resulting in decreased iron availability and iron-restricted

erythropoiesis.7

Stress erythropoiesis is a stress response pathway capable of rapidly producing large numbers

of mature erythrocytes during acute anemia when steady state production is not sufficient to

correct the anemia. It is best understood in mice where it occurs primarily in the murine spleen

and relies on unique signals from the microenvironment which are distinct from the signals

32

utilized in steady state erythropoiesis.8–11 Progenitor cells migrate from the bone marrow to the

spleen where hedgehog (Hh) and bone morphogenetic protein 4 (BMP4) signaling specify them

as stress erythroid progenitors (SEPs) which are self-renewing and erythroid restricted.9–12 In

addition to Hh and BMP4, growth differentiation factor 15 (GDF15) is a critical regulator of the

expansion of SEPs, and absence of any of these signals results in a reduction in stress erythroid

potential.11 Erythropoietin (Epo) and tissue hypoxia potentiate the response and promote the

differentiation of these progenitors.11,13 All these signals work in concert to tightly regulate the

expansion and differentiation of SEPs and ensure that the pathway is only activated in times of

need.

Recent work has shown that erythroid progenitors in the spleen are expanded after induction

of inflammation with heat-killed Brucella abortus, suggesting that stress erythroid progenitors

also differ from steady state progenitors in their responses to inflammatory stimuli.14,15

Observations in both Salmonella and malaria infections also show a large expansion of the

erythroid population in the spleen following infection.16–18 Stimulation of Toll-like receptor 2

(TLR2) results in an increase in stress erythroid burst forming units (BFU-Es), and unlike bone

marrow erythroid progenitors, stress BFU-E production is not inhibited by the presence of IFN-

γ.19 However, the expansion of erythroid progenitors during inflammation and how they are

affected by pro-inflammatory cytokines is still not well understood.

Here, we demonstrate that activation of TLR2 by Zymosan A results in the activation of the

stress erythropoiesis pathway, and this occurs in the absence of anemia, suggesting a novel

mechanism for activation. A key aspect of this mechanism is erythrophagocytosis. TLR-

dependent signals decrease the surface expression of signal-regulatory protein alpha (SIRPα), the

“don’t-eat-me” signal on macrophage surface, which in turn leads to increased

erythrophagocytosis. Breakdown of erythrocyte hemoglobin releases heme, increasing the

expression of the heme-regulated transcription factor Spi-C. Our data show that Spi-C plays a

33

critical role in inducing stress erythropoiesis during inflammation. Furthermore, we show that the

pro-inflammatory cytokine TNF-α is capable of enhancing stress erythropoiesis in vitro in both

murine and human bone marrow cultures.

Results

Stress erythropoiesis is rapidly induced upon inflammatory stimulation

We utilized the zymosan-induced generalized inflammation model (ZIGI) to study how

inflammatory signals affect stress erythroid progenitors. Zymosan A is a Toll-like receptor-2

(TLR2) ligand which induces an acute peritonitis after intraperitoneal injection. The acute

peritonitis persists for approximately 48 hours before resolving into a chronic inflammatory

disease.19,20 Zymosan is such a potent inflammatory mediator that, in order to prevent mortality

in wild type animals, mice are first given a small dose of lipopolysaccharide to stimulate

inflammatory cells, which feeds back to inhibit inflammatory responses thus buffering the

response to zymosan.20

Using this model, Millot et al. showed that mice developed anemia approximately 7-9 days

after zymosan treatment. Injection with erythropoietin was able to partially correct the anemia.

Steady state bone marrow erythropoiesis was suppressed by zymosan treatment, but BMP4

dependent stress erythropoiesis was induced. Although injections of Epo improved red cell

numbers, this treatment had no effect on bone marrow steady state erythropoiesis. Instead it

promoted a sustained stress erythropoiesis response. Millot et al. demonstrated that bone marrow

and spleen erythroid progenitors display markedly different responses to inflammation. Although

it is not surprising that stress erythropoiesis was induced in response to zymosan-induced anemia,

34

the question that stood out to us was why stress erythropoiesis was not activated earlier in an

effort to prevent the development of anemia during inflammation.

Activation of the BMP4-dependent stress erythropoiesis pathway is characterized by an

expansion of stress BFU-Es in the spleen in response to acute anemic stress and hypoxia plays a

role in regulating this response. Zymosan treated mice do not develop anemia until 7-9 days post-

treatment. We examined stress erythropoiesis at this time to determine when stress erythropoiesis

was actually activated in the spleen. Surprisingly, we observed a significant increase in the

number of stress BFU-Es in the spleen between 24 and 72 hours after zymosan treatment (Figure

2-1 A). Analysis of stress erythroid progenitor populations by flow cytometry revealed an

increased frequency of population I cells (Kit+Sca-1+CD71lo/medTer119lo/-) following treatment

with zymosan within 36 hours and subsequent increases in population II and III cells (Figure 2-1

B). We also observed a dramatic increase in Kit+Sca+CD34+CD133+ cells, which our lab has

defined as an immature population of stress erythroid progenitors contained within population I

(Figure 2-1 C). In contrast to the expansion of stress erythropoiesis in the spleen, bone marrow

steady state erythropoiesis decreased. We observed a decrease in colony formation and early

erythroid progenitors in the bone marrow after inducing inflammation with zymosan, which is

consistent with inhibition of bone marrow erythropoiesis by pro-inflammatory cytokines (Figure

2-1 D). These data demonstrate that zymosan-induced inflammation induces an immediate stress

erythropoiesis response.

We next tested how mice with mutations blocking stress erythropoiesis respond to zymosan.

GDF15-/- mice exhibit normal steady state erythropoiesis but are unable to respond to acute

anemic stress, whereas flexed-tail (f/f) mice have a delayed stress erythropoiesis response due to a

splicing mutation in Smad5, resulting in insufficient BMP4 signaling during stress erythropoiesis.

Both GDF15-/- mice and f/f mice have increased lethality following treatment with zymosan with

90% of GDF15-/- mice and 40% of f/f mice dying within 7 days of treatment compared with only

35

7% of wild type (WT) mice (Figure 2-1 E). GDF15-/- had significantly fewer stress BFU-Es after

inducing inflammation compared to wild type (Figure 2-1 F). At 36 hours, f/f have decreased

numbers of stress BFU-Es compared to WT but by 72 hours the number of stress BFU-Es is

comparable to WT (Figure 2-1 F). Mice that are unable to mount a stress erythropoiesis response

exhibit compromised survival in the ZIGI model of inflammatory disease.

Inflammation induces the expression of GDF15, a regulator of BMP4 expression, in the

spleen

The expansion of stress BFU-E in the spleen is driven by BMP4 dependent signaling.

Treatment with zymosan leads to a 12-fold increase in BMP4 mRNA expression 9 hours after

treatment (Figure 2-1 H). Previously we demonstrated that BMP4 expression in the spleen was

regulated by hypoxia. Furthermore, BMP4 expression is maintained by GDF15 dependent

signaling which down regulates the expression of VHL prolonging the hypoxia response (Xiang

and Paulson, unpublished data). Inflammation does not induce immediate tissue hypoxia, but we

do observe that GDF15 expression is rapidly upregulated within 3 hours after treatment with

zymosan (Figure 2-1 G), which precedes the increase in BMP4 expression at 9 hours after

treatment (Figure 2-1 H). These data suggest that unlike the response to acute anemia where

hypoxia initiates the increase in BMP4 expression, inflammation relies on an increase in GDF15

expression which induces a hypoxic state. In both f/f and GDF15-/- mice there is no increase in

BMP4 expression at 9 hours (Figure 2-1 I).

36

Stress erythropoiesis generates new erythrocytes immediately following zymosan treatment,

delaying the onset of anemia.

Reticulocyte numbers drop steadily on days 2 and 4 after zymosan treatment, consistent with

an inhibition of bone marrow erythropoiesis, before increasing on day 6 and continuing to rise

steadily through day 14 following the burst of stress erythropoiesis in the spleen (Figure 2-2 A).

There is no change in the number of reticulocytes in PBS-treated controls (Figure 2-2 A). In f/f

mice, there is a lag in the increase in reticulocytes and they are significantly lower than wild type

on days 6 and 8. However, reticulocytes begin to increase on day 10 and reach wild type levels by

day 14 (Figure 2-2 B). We also tested the replacement of old erythrocytes by biotinylating RBCs

in vivo. We observed a rapid drop in biotinylated RBCs beginning at day 6 in zymosan-treated

wild type mice compared to unstimulated controls, indicating an influx of new erythrocytes into

circulation (Figure 2-2 C). f/f mice exhibited a significantly higher percentage of biotinylated

erythrocytes than their wild type counterparts from days 6 to 14 after zymosan treatment,

suggesting fewer new erythrocytes having entered circulation (Figure 2-2 D). Hematocrits for

wild type and f/f mice decrease steadily over the first four days post treatment. However between

day 6 and day 8, f/f mice exhibit a rapid drop in hematocrit while control mice show a much

smaller drop (Figure 2-2 E). Hematocrit values seem to increase slightly on day 10, consistent

with the corresponding increase in reticulocytes on that day, before dropping again reaching a

lower nadir than wild type mice. Taken together, these data suggest that activation of the stress

erythropoiesis pathway by inflammation increases reticulocyte production and this influx of new

erythrocytes into circulation acts to help maintain hematocrit values temporarily, and in the case

of f/f mice where induction of stress erythropoiesis is delayed, hematocrit values drop rapidly and

remain lower than wild type mice.

GDF15-/- mice showed a slight increase in reticulocytes at day 6 but plateaued and remained

lower than wild type controls (Figure 2-2 B). However, there was almost no difference in the

37

percentage of biotinylated erythrocytes after treatment with zymosan and hematocrits were also

similar to wild type mice (Figure 2-2 D,E). This is possibly due to the fact that almost 90% of

GDF15-/- do not survive past day 4 and any mice surviving beyond day 4 have escaped, acting

more similar to wild type mice.

Inflammation induces the expression of Epo in the kidneys.

Typically, stress erythropoiesis is activated only during acute anemic stress with tissue

hypoxia and high levels of serum Epo acting on stress progenitors, promoting their transition

from a state of self-renewal to differentiation. Transcription of Epo is regulated by HIF-2α, a

protein which is targeted for ubiquitination by Vhl and degraded at normoxia.21 After treatment

with zymosan, we observed no decrease in hematocrit values during the first 48 hours where we

observe activation of stress erythropoiesis (Figure 2-2 E). However, analysis of protein from the

kidneys revealed a rapid and transient increase in the levels of HIF-2α at 6 hours after zymosan

(Figure 2-3 A). Epo mRNA expression was increased approximately 30-fold at 24 hours, and a

corresponding rise in serum Epo was observed at 24 and 36 hours (Figure 2-3 B,C). This increase

in serum Epo is sufficient to activate stress erythropoiesis in the absence of a severe drop in

hematocrit.

Both f/f and GDF15 mutants have decreased Epo expression and lower levels of serum Epo

compared with wild type controls at 24 hours after treatment with zymosan (Figure 2-3 D,E).

Bone marrow transplants were performed with mice receiving either wild type or f/f donor cells.

Transplanted mice were allowed to recover for 10 weeks before being treated with zymosan to

test the effect of donor cells in a wild type microenvironment. Mice receiving f/f donor BM have

decreased levels of serum Epo compared with mice which received wild type donor cells,

38

suggesting induction of Epo may be partially dependent on signaling from stress progenitors

(Figure 2-3 F).

Zymosan induces GDF15 expression by increasing erythrophagocytosis by splenic

macrophages

After treatment with zymosan, mice were not anemic but exhibited all the hallmarks of

activating stress erythropoiesis, suggesting that inflammatory signals are capable of inducing

stress erythropoiesis via a novel, hypoxia-independent mechanism. The initial feature of this

response is the up-regulation of GDF15 expression in the spleen. One mechanism that is known

to regulate GDF15 expression is heme-dependent activation of the transcription factor Spi-C.22

Bach1 represses Spi-C expression, but in the presence of heme Bach1 is degraded, leading to

increased Spi-C expression. This heme-dependent signal is required for the development of red

pulp macrophages. We tested whether zymosan can induce heme-dependent signaling in the

splenic macrophages. Erythrophagocytosis by red pulp macrophages plays a key role in

maintaining erythroid homeostasis. Signal regulatory protein alpha (SIRPα) is expressed on the

surface of macrophages and negatively regulates erythrophagocytosis by binding with the

erythrocyte surface protein CD47. This interaction acts as a “don’t-eat-me” signal.23,24 As red

cells age, they lose expression of CD47, resulting in less interaction with macrophage SIRPα

leading to phagocytosis of the erythrocyte. Previous work has shown that activation of TLRs

decreases surface levels of SIRPα. We proposed that TLR signaling decreased levels of SIRPα on

macrophage surfaces, leading to increased erythrophagocytosis. The subsequent increase in

heme-regulated transcription factor Spi-C leads to the expression of GDF15. Due to the rapid

upregulation of GDF15 in the spleen after treatment with zymosan, we focused our experiments

on events occurring within the first 3 hours following zymosan treatment.

39

We first examined SIRPα levels on the surface of F4/80+ cells immediately following

treatment with zymosan. SIRPα surface levels decreased significantly by 150 minutes following

zymosan treatment, resulting in a relative loss of 10.7% expression compared to unstimulated

controls (Figure 2-4 A). To address whether this small reduction in SIRPα levels on the surface of

macrophages was capable of increasing phagocytosis of circulating erythrocytes, we measured

erythrocyte uptake by splenic macrophages. Erythrocytes were isolated and labeled with CFSE

before being transfused into recipient mice where CFSE labeled erythrocytes constituted

approximately 5-10% of the erythrocytes (Figure 2-4 B,C,E). The transfused mice were then

treated with zymosan. There was no significant difference in the percentage of CFSE+ cells in the

blood of PBS- or zymosan-treated mice at 3 hours after treatment because CFSE+ and CFSE-

erythrocytes were phagocytosed at equal rates (Figure 2-4 C). However, examination of F4/80+

phagocytes in the spleens revealed there was a small but significant increase in the percentage of

CFSE labeled cells in zymosan-treated mice, indicating a small increase in erythrophagocytosis

that was detectable at 3 hours after treatment with zymosan (Figure 2-4 D). CFSE+F4/80+ cells in

the spleens of zymosan-treated mice continued to increase after treatment compared to PBS-

treated mice (Figure 2-4 F). At no time was there a significant change in the percentage of CFSE+

RBCs in the blood (Figure 2-4 C,E).

Hemoglobin from phagocytosed erythrocytes releases free heme, which can be toxic to the

cell. We observed that following zymosan treatment the expression of Hmox1, which breaks

down heme, was induced at 60 minutes post-treatment (Figure 2-5 A). FLVCR, the heme

exporter, was also induced at later time points (Figure 2-5 B). These data show the concentration

of intracellular heme increases in the spleen following zymosan treatment. We next examined

whether there was a corresponding change in the expression of heme-regulated genes indicative

of increases in intracellular heme. We focused on the expression of Spi-C which is known to

regulate the differentiation of monocytes in mature red pulp macrophages. Spi-C expression is

40

normally repressed by the protein BACH-1, which is negatively regulated by heme. After

treatment with zymosan, Spi-C expression was significantly increased at 90 minutes and with a

peak increase at 150 minutes (Figure 2-5 C). We assessed protein levels of Bach1 after treatment

with zymosan and were able to detect approximately a 2-fold decrease in the amount of Bach1 in

the spleen (Figure 2-5 D). Bach1 is degraded in the presence of heme, and this decrease supports

an increase in erythrophagocytosis in splenic macrophages in response to zymosan. We next

sorted F4/80+CFSE- and F4/80+CFSE+ cells by flow cytometry to determine if changes in heme-

regulated genes were occurring primarily in cells which had phagocytosed CFSE+ RBCs. The

increased expression of Spi-C was limited to CFSE+ macrophages demonstrating that heme

derived from erythrophagocytosis led to increased SpiC expression. Similarly the expression of

GDF15 was significantly increased by 3000 fold, but only in F4/80+CFSE+ macrophages

(Figure 2-5 E,F). Taken together, these data demonstrate that increased erythrophagocytosis

induced by zymosan results in increased levels of intracellular free heme which are sufficient to

drive changes in expression of heme-related genes and in particular lead to the Spi-C dependent

upregulation of GDF15.

In order to further demonstrate this point, Spi-C-/- mice were treated with zymosan. As

previously shown, Spi-C-/- mice have fewer F4/80+ cells in the spleen, but within the F4/80+

population, these mice also exhibit a significant decrease in phagocytosis of CFSE+ erythrocytes

when compared with wild-type controls (Figure 2-6 A,B). GDF15 expression was lower in

F4/80+CFSE+ cells in Spi-C-/- mice than in wild type controls (Figure 2-6 C). Furthermore, Spi-

C-/- mice treated with zymosan exhibit a defect in the expansion of stress BFU-E in the spleen.

Spi-C-/- mice make fewer stress BFU-Es at 36 and 72 hours after zymosan than wild type mice

(Figure 2-6 D). In order to further demonstrate that Spi-C regulates GDF15 expression, we

generated bone marrow derived macrophages (BMDMs) from wild type and Spi-C-/- mice. In

41

contrast to what we observed in vivo, Spi-C-/- BMDMs show no difference in the decrease in

surface levels of SIRPα and the increased erythrophagocytosis in response to zymosan compared

to BMDMs from control mice, which is consistent with those events being upstream of Spi-C

activation (Figure 2-6 E,F). However the ability of zymosan to induce GDF15 expression in vitro

was severely compromised when Spi-C expression was absent in knockout BMDMs (Figure 2-6

G).

To examine the importance of Spi-C expression in splenic macrophages during

inflammation-induced stress erythropoiesis, we depleted native phagocytes in Spi-C-/- and WT

mice using clodronate liposomes and adoptively transferred wild type BMDMs into these

macrophage-depleted recipients (Figure 2-7 A). Mice were treated with zymosan one week after

transferring BMDMs. Colony assays at 36 hours after zymosan treatment shown that transfer of

WT BMDMs into Spi-C-/- mice results in a partial rescue with the number of stress BFU-Es

being nearly equivalent to WT animals and increased compared with control Spi-C-/- mice

(Figure 2-7 B). WT clodronate-depleted mice also showed increased numbers of stress BFU-Es at

36 hours and the response was comparable to that in normal WT animals. There was increased

variability in clodronate-depleted mice, possibly due to differences in the homing of BMDMs to

the spleen, but the increase in stress BFU-Es was still significant compared with untreated

controls (Figure 2-7 B). We also tested the ability of wild type monocytes to rescue

inflammation-induced stress erythropoiesis in Spi-C-/- mice. Recent studies have shown that for

monocytes to develop into red pulp macrophages heme/phagocytosis of RBCs is required. We

pre-treated recipient mice with LPS and 6 days later treated with zymosan and adoptively

transferred 1x106 purified CD11b+Ly6G-Ly6C+ monocytes into each mouse (Figure 2-7 C,D).

The purified monocytes were from CD45.1 donor mice and after 72 hours comprised

approximately 2-5% of recipient spleens (Figure 2-7 E). In Spi-C-/- recipients, the monocytes

were found at slightly lower frequencies that in wild type recipients (Figure 2-7 E). Spi-C-/- mice

42

with wild type donor monocytes exhibited a small but significant increase in stress BFU-Es at

36h hours compared with Spi-C-/- mice receiving no donor cells (Figure 2-7 F). At 36 hours, Spi-

C-/- recipients still have a lower frequency of stress BFU-Es compared to either group of wild

type mice (Figure 2-7 F). However, by 72 hours Spi-C-/- with WT donor monocytes have

numbers of stress BFU-Es similar to wild type mice, indicating that Spi-C expression in

monocytes and red pulp macrophages is critical in the induction of stress erythropoiesis by

inflammatory signaling (Figure 2-7 F).

MyD88 mediates the increase in GDF15 expression following zymosan treatment.

MyD88 is a critical adaptor protein for signaling through TLRs. Previous work showed that

salmonella infection increases the number of TER119+CD71+ erythroid progenitors in the spleen

and that increase was abrogated in MyD88 tm1.1Defr/ tm1.1Defr (MyD88-/-) mice. These data suggest

that increases in splenic stress erythropoiesis by inflammatory stimuli is a MyD88-dependent

event.18 To determine if the response to zymosan was MyD88-dependent, we utilized either wild-

type or MyD88 -/- BMDMs to assess the role of MyD88 in regulating erythrophagocytosis and

upregulation of GDF15. MyD88-/- mutant BMDMs failed to downregulate SIRPα expression

after treatment with zymosan (Figure 2-8 A). Furthermore, the mutant macrophages exhibited

significantly less phagocytosis of CFSE-labeled RBCs (Figure 2-8 B). Their inability to

phagocytose erythrocytes lead to a weak increase in SpiC expression , which was significantly

lower than the SpiC expression expression levels induced in WT BMDMs (Figure 2-8 C). The

lack of robust SpiC expression prevented the increase in GDF15 expression. In fact the level of

GDF15 expression was similar to unstimulated controls (Figure 2-8 D). Although the

homozygous MyD88 mutant cells exhibited a significant defect in their response to zymosan,

MyD88+/- cells exhibited a response that was intermediate between wildtype controls and mutant

43

cells in that MyD88+/- BMDMs show a reduction in the levels of SIRPα on their cell surface

after treatment with zymosan (Figure 2-8 A). Heterozygous MyD88+/- BMDMs demonstrate

slightly lower rates of erythrophagocytosis compared to wild type, which results in a mild but

significant reduction in GDF15 expression (Figure 2-8 B-D).

TNFα and IL1-β promote erythroid differentiation under stress erythropoiesis.

Inflammation inhibits bone marrow steady state erythropoiesis. This effect is in part due to

pro-inflammatory cytokines, such as TNF-α and IFN-γ.1,2,4,25 Furthermore, chronic exposure of

hematopoietic stem cells (HSCs) to IL-1β results in activation of PU.1 which both drives

myelopoiesis and represses erythropoiesis and lymphopoiesis.6 Zymosan induces an increase in

pro-inflammatory cytokine expression in the spleen, and it has been shown by Millot et al. that

the presence of IFN-γ does not affect production of stress BFU-Es, which indicates that stress

erythroid progenitors in the spleen must respond differently to inflammatory signals.19

Following treatment with zymosan in vivo, TNF-α and IL-1β mRNA is significantly

upregulated after 3 hours (Figure 2-9 A). Intracellular levels of TNF-α and IL-1β protein in

F4/80+ cells in the spleen were measured by flow cytometry and were also increased 3 hours

after treatment with zymosan (Figure 2-9 B,C). To determine how exposure to pro-inflammatory

cytokines affects stress erythroid progenitors in the spleen, we utilized a two stage in vitro culture

system developed in our lab which includes cytokines essential in promoting the expansion and

differentiation of stress progenitors. Whole bone marrow cells were cultured in stress erythroid

expansion media (SEEM) and pulsed with TNF-α, IFN-γ, or IL-1β for 24 hours. Transient

exposure to TNF-α had no effect on the expansion of SEPs compared to cells grown only in

SEEM after 7 days culture (Figure 2-9 D). However, both IL-1β and IFN-γ seemed to inhibit the

expansion of cells, though neither effect was significant (Figure 2-9 D). When cells were plated

44

for stress BFU-Es after 7 days in SEEM, cells treated with either TNFα or IL-1B had a significant

increase in stress BFU-Es, indicating that the presence of IL-1β or TNFα enhances erythroid

output (Figure 9E). The increase in stress BFU-Es with TNF- α treatment is even more profound

when total numbers of stress BFU-Es per culture are calculated (Figure 2-9 F). IL-1β has a more

modest effect on stress BFU-Es per culture due to the slight decrease in growth compared to

untreated cells (Figure 2-9 F). IFN-γ not only seems to inhibit the growth of SEPs but also has no

significant effect on erythroid output as there was no significant change in stress BFU-Es (Figure

2-9 E), but the number of stress BFU-Es in the total culture was lower since there were far fewer

cells in cultures treated with IFN-γ (Figure 2-9 F).

Since TNF-α had the most profound effect on mouse BM cultures, we examined its effect on

unfractionated human bone marrow cultured in stress erythropoiesis conditions. It has previously

been shown that culturing human BM cells in SEEM results in the generation of stress BFU-Es

similar to murine stress erythropoiesis.11 Human cultures pulsed with TNFα for 24 hours

exhibited a slight increase in proliferation compared with untreated cells (Figure 2-10 A). There

was also in increase in stress BFU-Es in the culture treated with TNF-α, consistent with our

finding in mice (Figure 2-10 B,C). Taken together, these data suggest that stress erythroid

progenitors respond differently to the presence of pro-inflammatory cytokines than steady state

bone marrow progenitors and that different cytokines produce different responses in SEPs.

Discussion

Our results indicate that inflammatory stimuli increase phagocytosis of circulating

erythrocytes and this drives changes in gene expression in splenic macrophages which activate

the BMP4 dependent stress erythropoiesis pathway. Previous studies have shown expansion in

the erythroid compartment of the spleen during other model of infection and inflammation.

45

However, most of these studies occur after the establishment of anemia of inflammation (AI) and

several days to weeks after the initial infection or inflammatory stimulus. Here we have

demonstrated that there is a rapid stress erythroid response following the initial inflammatory

stimulus occurring before the onset of anemia. Stress erythropoiesis is typically considered a

stress response pathway during acute anemic or hypoxic stress. However, our data suggest that

SEPs also respond early during inflammation in advance of any overt anemia or tissue hypoxia.

We speculate that this effect is potentially to preemptively offset the shift in the bone marrow

from erythropoiesis to myelopoiesis and help maintain levels of circulating erythrocytes for a

short period of time. This sheds light on a novel role for stress erythropoiesis. The production of

myeloid cells types is critical to resolving inflammation and in clearance during infections.

However, new erythrocytes still need to be produced to continue to provide sufficient

oxygenation to tissues, and the ability of SEPs to be activated early during inflammation indicates

the need to shift to extramedullary erythropoiesis temporarily until homeostatic conditions in the

bone marrow can be reestablished.

Our data also outline a novel mechanism for the activation of the stress erythropoiesis

pathway under inflammatory conditions. Stimulation of TLRs increases phagocytosis of

circulating erythrocytes, which has also been observed previously, and we have shown that this is

critical in the activation of stress erythropoiesis. Our data show that small changes in the amount

of SIRPα on the surface of macrophages in the spleen is sufficient to increase

erythrophagocytosis and consequently increase intracellular heme, indicated by changes in

expression of several heme-regulated genes. Our data suggest that increased expression of Spi-C

is required to activate stress erythropoiesis by inducing expression of GDF15. However, it is

unclear if Spi-C regulates GDF15 directly or indirectly and future studies are needed to fully

understand this interaction. Phagocytosis of circulating erythrocytes results in the accumulation of

hemoglobin in splenic macrophages which efficiently recycle iron and heme. In the absence of

46

anemia, subtle changes in the levels of intracellular heme could act as a signal to initiate stress

erythropoiesis and recycle heme accumulating in splenic macrophages for the production of new

erythrocytes.

Our data also clearly demonstrate that SEPs are not affected by pro-inflammatory cytokines

in the same way as steady state erythroid progenitors. Many studies have demonstrated the direct

and indirect inhibition of erythroid progenitors by TNF-α, IFN-γ, and IL-1. Both IFN-γ and IL-1

signaling drive cells toward the myeloid lineage.6,26 TNF-α and IFN-γ have been shown to inhibit

the proliferation and differentiation of erythroid progenitors in the marrow.1–3 Recent work

examining stress erythropoiesis during inflammatory conditions showed that IFN-γ did not affect

BFU-Es in the spleen while it negatively affected numbers of BFU-Es in the bone marrow.19 Our

data shows that TNF-α is capable of enhancing the erythroid potential of bone marrow cells

cultured under stress conditions. In contrast to their effect in the bone marrow, acute exposure to

TNF-α and IL-1β may work in concert with other stress erythropoiesis signals to promote the

expansion of erythroid progenitors. This phenomenon holds true for human bone marrow MNCs

cultured under stress conditions, indicating a similar function for human SEPs as we see in mice

where acute exposure to TNF-α may promote expansion of this population and delay the onset of

anemia. It is unclear if TNF-α and IL-1β are acting on SEPs directly or affecting them indirectly

through macrophages, and more work will be necessary to fully understand the relationship

between SEPs and these pro-inflammatory cytokines. These data provide new insights into the

complex interactions of SEPs with the splenic microenvironment and the unique signals that

affect their proliferation and differentiation.

Current therapies for AI are unsuitable for long-term treatment solutions and often ineffective

in correcting the anemia. The most common treatments are blood transfusions, iron therapy, or

repeated injections with Epo to stimulate red cell production. Blood transfusions can temporarily

correct anemia but long-term transfusion therapy carries the risk of iron overload. Oral

47

supplementation of dietary iron is problematic due to poor absorption of iron as a result of

decreased ferroportin expression. Understanding how SEPs are mobilized during inflammation

and how inflammatory stimuli affect their proliferation is essential in finding novel therapeutics

for the treatment of AI that target this specialized population of erythroid progenitors.

Material and Methods

Mice. C57BL/6 mice were purchased from Taconic Biosciences, Inc. All mice were 6-12

weeks old. GDF15-/- mice were provided by Dr. Se-Jun Lee at Johns Hopkins.27 Spi-C-/- mice

were previously described elsewhere and kindly provided by Dr. Ken Murphy at the Washington

University School of Medicine (St. Louis, MO).28 B6.129P2(SJL)-MyD88tm1.1Defr/J have been

previously described and were provided by Dr. Matam Vijay Kumar at the Pennsylvania State

University (State College, PA). All procedures have been approved by the Institutional Animal

Care and Use Committee of the Pennsylvania State University.

Zymosan-Induced Generalized Inflammation. Mice were first treated with 40µg/200µL of

lipopolysaccharide from Escherichia coli 0128:B12 (Sigma-Alrich L2887) and followed six days

later by Zymosan A from Saccharomyces cerevisiae (Sigma-Aldrich Z4250) at a concentration of

0.48mg/g. All treatments were administered by intraperitoneal injection.

Bone Marrow Transplant. Recipient mice were irradiated with a single dose of 950Rads and

donor cells were administered by IV injection. Recipeints received 5x105 donor cells and were

allowed to recover for 10 weeks prior to being treated with zymosan.

Murine and Human Cell Cultures. Murine bone marrow was isolated from the femurs and

cultured in stress erythropoiesis expansion media (SEEM) for 7 days at normoxia as previously

described and plated for colony assays.11 Briefly, SEEM is composed of Iscove’s modified

48

Dulbecco’s media (IMDM) containing 10%FBS, 10µg/mL Insulin, 200µg/mL holo-transferrin,

2mM L-glutamine, 10µg/mL ciprofloxacin, 1% BSA, 7µL/L 2-mercaptoethanol, GDF15

(30ng/mL; Biomatik), BMP4 (15,ng/mL; R&D Systems), Shh (25ng/mL; GoldBio), and SCF

(50ng/mL; GoldBio). TNFα (GoldBio 1330-01) was added to cultures for 24 hours at a

concentration of 50ng/mL. After 24 hours, cells were washed and replated with fresh SEEM.

Human bone marrow mononuclear cells were purchased from ReachBio (0300-200) and cultured

in SEEM containing human insulin, human holo-transferrin, and human cytokines. Human MNCs

culture conditions were the same as those used for murine bone marrow.

Colony Assays. Murine splenocytes or bone marrow were plated in methylcellulose media

(Stem Cell Technologies, M3334) at a concentration of 1x105 cells/well in a 12-well tissue

culture plate. BMP4 (15ng/mL), SCF (50ng/mL), Shh (25ng/mL), GDF15 (15ng/mL), and Epo

(3U/mL) were added to methylcellulose media to assay stress BFU-E formation. Cells were

incubated at 1% O2 for 5 days before counting BFU-Es. BFU-Es were stained with benzidine

solution for counting. Human bone marrow was plated in methylcellulose media (Stem Cell

Technolonies, H4130) at a concentration of 5x104 cells/well with human cytokines.

mRNA Isolation and Gene Expression Analysis. Total RNA was isolated using TriZol reagent

(Invitrogen 15596). cDNA was generated using the high capacity cDNA synthesis kit (Applied

Biosystems). Quantitative reverse transcription PCR (qRT PCR) was carried out using Taqman

probes and an ABI7300 real-time PCR system. Taqman probes: BMP4 (Mm00432087_m1),

GDF15 (Mm00442228_m1), Epo(Mm01202755_m1), Spi-C (Mm00488428_m1),

FLVCR(Mm01320423_m1), HMOX1(Mm00516005_m1), TNFα(), 18S(Hs99999901_s1).

Western Blotting. Primary antibodies anti-HIF-2α (Novus Biologicals NB100-122) and anti-

BACH1 (R&D Systems, ) were used for western blots. Bands were visualized using Amersham

ECL prime wester blotting detection reagent (GE Healthcare RPN2106).

49

ELISAs. Serum levels of Erythropoietin were determined using the commercially available

mouse Erythropoietin Quantikine ELISA kit (R&D Systems MEP00B) according to

manufacturer’s instructions.

Flow Cytometry. Flow cytometry was performed using a BD Accuri C6 flow cytometer (BD

Biosciences) and data was analyzed in FlowJo v10. Cells were sorted on a Beckman Coulter

MoFlo Astrios. Flow antibodies: c-Kit Alexa 647 (Clone 2B8, Biolegend 105818), Sca-1 PE-Cy7

(Clone D7, BD Biosciences BDB558162), Ter119 PE (Clone Ter119, BD Biosciences

BDB553673), CD71 FITC (Clone C2, BD Biosciences BDB553266), F4/80 PE-Cy7 (Clone

BM8, Biolegend 123112), CD172a APC (Clone P84, BD Biosciences BDB560106), FITC TNFα

(BD Biosciences 506304), FITC IL-1β (R&D Systems IC4013F).

In vivo Biotinylation of RBCs. Biotinylation was done as previously described.24 In short,

1mg of Biotin-X-NHS (Cayman Chemicals) was dissolved in 20µL dimethylformamide and

diluted in PBS. Biotin was administered intravenously daily for three consecutive days.

Biotinylated RBCs were quantified by flow cytometry using a Strepavidin-Pe-Cy7-conjugated

antibody.

In vivo Transfusion Assay. Erythrocytes were isolated by terminal cardiac puncture and

labeled with CFSE (Life Technologies C34554). Approximately 200µL of blood was removed

from recipients by retro orbital bleeding and recipients were then transfused with a total volume

of 200µL labeled RBCs. Mice were treated with either zymosan or PBS 24 hours after

transfusion. Splenic macrophages were examined by flow cytometry to determine levels

internalized CFSE-labeled RBCs.

In vitro Erythrophagocytosis Assay. Bone-marrow derived macrophages (BMDMs) were

grown in 20% L929-conditioned media for 7 days, and then were replated at a density of 2x106

cells/well. BMDMs were stimulated overnight with either IFN-γ (100U/mL, R&D Systems) +

LPS (10ng/mL) or Zymosan (10µg/mL) + LPS (10ng/mL). Red blood cells (RBCs) were

50

collected in an EDTA-coated tube, labeled with CFSE and aged in Hepes buffer (10mM Hepes,

140mM NaCl, 0.1%BSA, pH 7.4) containing calcium (2.5mM) and Ca2+ ionophore (0.5µM,

A23187). RBCs were aged for 16 hours at 37°C. BMDMs were washed twice with PBS before

aged RBCs were added at a concentration of 3x107 cells/mL. BMDMs were incubated with RBCs

at 37°C for 1 hour and were then washed twice with PBS. BMDMs were harvested and

phagocytosis was assessed by flow cytometry.

Adoptive Transfer of BMDMs and Monocytes. Native phagocytes were depleted with

clodronate liposomes, which were administered for 3 consecutive days and mice received 200µL

of clodronate liposomes (approximately 50mg/mouse) according to manufacturer’s instructions.

1x106 bone marrow derived marcohages were adoptively transferred 24 hours after completion of

clodronate treatment and LPS was given the next day. Mice were treated with zymosan as

described above. Monocytes were purified using the EasyStep Monocyte Enrichment kit from

Stem Cell Technologies (19761) according to manufacturer’s instructions. 1x106 monocytes were

transferred into recipient mice and zymosan was administered simultaneously.

Statistics. P-values were determined using the Student’s t-test (2-tailed), Mann-Whitney test,

or two-way ANOVA, as deemed appropriate. Significance was determined as * p<0.05, **

p<0.01, *** p<0.001.

51

Figures

Figure 2-1 A. Zymosan leads to expansion of stress erythroid progenitors in the spleen.

(A) Mice were treated with zymosan and early activation of stress BFU-Es in the spleen was

measured by colony assays. Splenocytes were plated at a concentration of 1x105 cells/well in the

presence of GDF15, BMP4, SCF, Shh and Epo at 1% O2. Stress BFU-Es were counted after five

days. Mean±SD. Student t-test, n=4-11.

52

Figure 2-2 B. Zymosan leads to expansion of stress erythroid progenitors in the spleen.

(B) In addition to an increase in stress BFU-Es in the spleen, we also observe an increase in

Population I cells in the spleen after treatment with zymosan. Flow cytometry analysis of

splenocytes was performed after treatment with zymosan. Cells were gated on Kit+ cells and

frequencies of CD71 and Ter119 populations are shown in representative images. Populations I,

II, and III are indicated on plots (frequencies are represented as bar graphs in lower panel).

Mean±SD. Student t-test, n=3-5.

53

Figure 2-3 C. Zymosan leads to expansion of stress erythroid progenitors in the spleen.

(C) We also observe an increase in Population I cells in the spleen after treatment with zymosan.

Flow cytometry analysis of splenocytes was performed after treatment with zymosan. Cells were

gated on Kit+Sca+ cells and frequencies of CD34 and CD133 populations are shown in

representative images. Images are representative of 2 independent experiments. n=3.

54

Figure 2-4 D. Zymosan leads to expansion of stress erythroid progenitors in the spleen.

(D) BFU-Es in the bone marrow were assessed to confirm a decrease in erythroid activity after

treatment with zymosan. Cells from bone marrow were plated at a concentration of 1x105

cells/well in the presence of IL-3 (25ng/mL) and Epo (3U/mL) at 20% O2. BFU-Es were counted

after seven days. (n=3, **p<0.01) Data were analyzed using student T-test (two-tailed) and are

representative of at least two independent experiments.

55

Figure 2-1 E-F. Zymosan leads to expansion of stress erythroid progenitors in the spleen.

(E) Survival of C57BL/6, flexed-tail (f/f), and GDF15-/- mice was assessed between days 0 and 7

following treatment with zymosan. f/f mice exhibit approximately a 45% mortality rate and

GDF15-/- have a mortality rate of 90%. Mantel-Cox test, n=13-29. (F) Stress BFU-Es were

measured as previously described to determine if f/f and GDF15-/- mice also have defects in

inflammation-induced erythropoiesis. Splenocytes were plated at a concentration of 1x105

cells/well in the presence of GDF15, BMP4, SCF, Shh and Epo at 1% O2. Stress BFU-Es were

counted after five days. Mean±SD. Student t-test, n=3-9.

56

Figure 2-1 G-I. Zymosan leads to expansion of stress erythroid progenitors in the spleen.

(G,H,I) Mice were treated with zymosan and RNA was isolated from splenocytes at indicated

time points and relative expression of GDF15 and BMP4 compared to 18S was determined by q-

PCR. Both GDF15 and BMP4 expression was upregulated in response to zymosan. Mean±SD.

Student t-test, n=3-10.

57

Figure 2-2 A-B. Zymosan induces production of new erythrocytes.

(A,B) To assess if the activation of stress erythropoiesis in the spleen results in production of new

erythrocytes, the frequency of reticulocytes in the blood was measured by flow cytometry using

Thiazole Orange. Mean±SD. Two way ANOVA, n=4-22.

58

Figure 2-2 C-E. Zymosan induces production of new erythrocytes.

(C,D) The influx of new erythrocytes was also measured by in vivo biotinylation of circulating

erythrocytes prior to treatment with zymosan or PBS. Three injections of biotin are sufficient to

label all circulating RBCs. Biotinylation was measured by flow cytometry after treatment with

zymosan or PBS. Mean±SD. Two way ANOVA, n=4-19. (E) To determine if this influx in new

erythrocytes resulted in maintenance of hematocrit values, blood was collected in microcapillary

tubes from wild type, f/f and GDF15-/- mice after zymosan treatment and spun to determine

hematocrit values. Mean±SD. n=2-10

59

Figure 2-3 A. Zymosan induces Erythropoietin expression in the absence of anemia.

(A) Hif-2α regulates transcription of Epo in the kidneys and is typically only stable during

hypoxic conditions. Protein was isolated from kidneys after treating with zymosan for western

blot, and blots were probed for Hif-2α and β-actin. Blots for biological replicates are shown side

by side. Hif-2α (top) and β-actin (bottom). n=2

60

Figure 2-3 B-C. Zymosan induces Erythropoietin expression in the absence of anemia.

(B) RNA was isolated from kidneys after treatment with zymosan and Epo expression was

determined relative to 18S. Upregulation of Epo transcript was detected at 12 and 24 hours after

treating with zymosan, consistent with the stabilization in Hif-2α. Mean±SD, Student t-test, n=2-

5. (C) To determine if increased transcription resulted in higher levels of Erythropoietin, serum

was isolated after zymosan treatment and concentration of Epo was determined by ELISA.

Mean±SD, Mann Whitney test, n=2-8.

61

Figure 2-3 D-F. Zymosan induces Erythropoietin expression in the absence of anemia.

(D) Epo mRNA expression was measured in f/f and GDF15 mutants. RNA was isolated from

kidneys after treatment with zymosan and Epo expression was determined relative to 18S.

Mean±SD, n=3-5. (E) Serum was isolated from mutants after zymosan treatment and

concentration of Epo was determined by ELISA Mean±SD, n=2-4. (F) Mice were transplanted

with 5x105 unfractionated bone marrow cells from either wild type (WT-WT) or f/f (f/f-WT)

donors and allowed to recover for 8 weeks before treatment with zymosan. Serum was isolated

after zymosan treatment and concentration of Epo was determined by ELISA. Mean±SD, Student

t-test, n=3.

62

Figure 2-4 A-B. SIRPα expression decreases and results in increased erythrophagocytosis.

(A) To determine surface expression of SIRPα, splenocytes were isolated at indicated time points

after zymosan treatment and SIRPα levels were measured on F4/80+ cells by flow cytometry.

SIRPα percentages expressed are relative to 0 minutes (freq. of x minutes/freq. of 0

minutes*100). Mean±SD, Student t-test, n=3-4. (B) Schematic depicting experimental design of

CFSE+ blood transfusion. Recipient mice were treated with LPS and 5 days later were transfused

with fresh donor RBCs that had been labeled with CFSE. Mice were allowed to rest for 24 hours

before being treated with zymosan and assessing phagocytosis of CFSE-labeled RBCs.

63

Figure 2-4 C-F. SIRPα expression decreases and results in increased erythrophagocytosis.

(C-F) Mice were transfused with CFSE-labeled RBCs and treated with zymosan. After 3 or 24

hours, blood (C,E) or splenocytes (D,F) from PBS- or zymosan-treated mice were examined by

flow cytometry for CFSE+ cells in circulation or phagocytosed. The frequency of CFSE+ cells in

the spleen was first gated on F4/80+ cells. Mean±SD, Student t-test, n=3-8.

64

Figure 2-5 A-C. Erythrophagocytosis leads in increased intracellular heme and changes in heme-

dependent gene expression.

(A-C) To assess if this increase in phagocytosis represented a corresponding rise in intracellular

heme, RNA was isolated from splenocytes and expression of heme-dependent genes was

measured. HMOX1 (n=4-7), FLVCR (n=4-7), Spi-C (n=3-8) were measured relative to 18S.

Mean±SD. Student t-test.

65

Figure 2-5 D. Erythrophagocytosis leads in increased intracellular heme and changes in heme-


(D) Bach1 protein levels are diminished in the presence of free heme. Splenocytes were isolated

after treating mice with zymosan. Red blood cells were lysed to remove mature erythrocytes from

the population. Protein was isolated from this population after RBC lysis. Bach1 and b-actin

proteins were measured by western blot. Densitometry was performed using ImageJ. Data were

analyzed by student t-test. Blots shown are representative of 3 biological replicates.

66

Figure 2-5 E-F. Erythrophagocytosis leads in increased intracellular heme and changes in heme-


(E,F) Previous experiments examined the expression of GDF15 and Spi-C from whole spleen. To

determine if this upregulation in these two genes was primarily the result of increased

phagocytosis of RBCs in splenic macrophages, splenocytes were sorted into F4/80+CFSE- (non-

phagocytosing macrcophages) or F4/80+CFSE+ (phagocytosing macrophages) populations 3

hours after treatment with zymosan. RNA was isolated and expression of Spi-C and GDF15 were

measured relative to 18S. Data were analyzed by Mann-Whitney U test and are representative of

two independent experiments. n=4.

67

Figure 2-6 A-B. Loss of Spi-C affects stress erythropoiesis response after treatment with zymosan.

(A) Spi-C-/- mice have a decreased frequency of F4/80+ red pulp macrophages. To confirm this,

splenocytes were isolated from untreated mice and percentage of F4/80+ cells were analyzed by

flow cytometry. Mean±SD. n=3. (B) Even though Spi-C is downstream of SIRPα, the frequency

of CFSE+ cells in the spleen after transfusion of labeled RBCs and treatment with zymosan was

determined by flow cytometry. CFSE+ cells were first gated on F4/80+ cells. Mean±SD. Student

t-test, n=4-7.

68

Figure 2-6 C. Loss of Spi-C affects stress erythropoiesis response after treatment with zymosan.

(C) Mice were transfused with CFSE-labeled erythrocytes and treated with zymosan 24 hours

later. Splenocytes were sorted into F4/80+CFSE- or F4/80+CFSE+ populations after 6 hours. RNA

was isolated and expression of GDF15 was measured relative to 18S. Mean±SD. Mann Whitney

test. n=3-7.

69

Figure 2-6 D. Loss of Spi-C affects stress erythropoiesis response after treatment with zymosan.

(D) Wild type and Spi-C-/- mice were treated with zymosan and splenocytes were isolated to

determine if Spi-C-/- mice have defective inflammation-induced stress erythropoiesis. As

previously described, splenocytes were plated in methylcellulose media containing BMP4, Shh,

SCF, GDF15, and Epo and cultured under hypoxic conditions for 5 days. Stress BFU-Es were

scored after staining with benzidine. Student t-test, n=3-5.

70

Figure 2-6 E-F. Loss of Spi-C affects stress erythropoiesis response after treatment with zymosan.

(E) Bone marrow derived macrophages were obtained by culturing either wild type or Spi-C-/-

bone marrow in the presence of 20% L929 conditioned media for 6 days. BMDMs were then

replated at a density of 3x106 cells/well in a 6-well plate and stimuated with zymosan for 12

hours before adding aged erythrocytes. Surface expression of SIRPα was measured by flow

cytometry after 3 hours. Mean±SD. Student t-test, n=3-5. (F) Percentage of CFSE+ cells was

measured in BMDMs by flow cytometry 1 hour after the addition of CFSE-labelled RBCs.

Mean±SD. Student t-test, n=3.

71

Figure 2-6 G. Loss of Spi-C affects stress erythropoiesis response after treatment with zymosan.

(G) To determine if GDF15 expression is dependent on the presence of Spi-C in vitro, RNA was

isolated from BMDMs 3 hours after the addition of aged RBCs and expression of GDF15 was

measured relative to 18S. Mean±SD. Student t-test, n=3-5.

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Figure 2-7 A-B. Adoptive Transfer of WT BMDMs or monocytes restores stress erythropoiesis in

Spi-C -/- mice.

(A) Schematic depicting experimental design of adoptive transfer of BMDMs. BMDMs were

cultured as previously described in 20% L929 conditioned media. Recipient mice were treated

with clodronate for 3 consecutive days and 1x106 BMDMs were adoptively transferred into the

recipients 24 hours after the last clodronate treatment. Mice were treated with LPS the day after

the adoptive transfer of BMDMs and six days later were treated with zymosan. (B) To assess if

wild type BMDMs were capable of rescuing stress erythropoiesis in Spi-C-/- mice, splenocytes

from the adoptive transfer recipeints were plated in methocelluse after treatment with zymosan at

a concentration of 1x105 cells/well in the presence of GDF15, BMP4, SCF, Shh and Epo at 1%

O2. Stress BFU-Es were counted after five days. Mean±SD. Student t-test, n=3-8.

73

Figure 2-7 C-D. Adoptive Transfer of WT BMDMs or monocytes restores stress erythropoiesis in

Spi-C -/- mice.

(C) Schematic depicting experimental design of adoptive transfer of purified CD11+Ly6G-Ly6C+

monocytes. Monocytes were isolated from donor mice using a negative selection magnetic bead

kit from StemCell Technologies. Recipient mice were treated with LPS and 6 days later were

simultaneously treated with zymosan and recieved 1x106 monocytes. (D) Representative flow

cytometry plots of CD11+Ly6G-Ly6C+ monocyte population purified by magnetic bead selection.

74

Figure 2-7 E-F. Adoptive Transfer of WT BMDMs or monocytes restores stress erythropoiesis in

Spi-C -/- mice.

(E) Recipients received 1x106 purified CD11+Ly6G-Ly6C+ monocytes by retro-orbital injection

and were immediately treated with zymosan. Splenocytes were isolated and CD45.1+ donor cells

were measured by flow cytometry to determine percent engraftment in the spleen. Mean±SD.

n=7-8. (F) To assess if adoptive transfer of wild type monocytes to Spi-C-/- mice was capable of

rescuing inflammation-induced erythropoiesis, mice were treated with zymosan and splenocytes

were plated at a concentration of 1x105 cells/well in the presence of GDF15, BMP4, SCF, Shh

and Epo at 1% O2. Stress BFU-Es were counted after five days. Mean±SD. Student t-test, n=3-4.

75

Figure 2-8 A-B. MyD88 is required for increased erythrophagocytosis.

(A) BMDMs were cultures for 6 days in 20% L929-conditioned media as previously described

and replated before being stimulated with LPS+Zymosan for 12 hours. CFSE labeled erythrocytes

were then added to the culture for 1 hour. BMDMs were collected and analyzed by flow

cytometry for the presence of CFSE. (B) BMDMs were harvested after 3 hours and surface

SIRPα was measured by flow cytometry. Mean±SD. Student t-test, n=3-8.

76

Figure 2-8 C-D. MyD88 is required for increased erythrophagocytosis.

(C, D) To determine if GDF15 and Spi-C expression are compromised in the absence of MyD88-

dependent TLR signaling, RNA was isolated from BMDMs using TRIZol reagent and expression

of Spi-C (C, n=1-4) and GDF15 (D, n=3-7) was quantified by q-PCR relative to ribosomal

subunit 18S. Mean±SD. Student t-test.

77

Figure 2-9 A-C. TNFα and IL1-β promote erythroid differentiation under stress erythropoiesis.

(A) To measure TNF expression in vivo, RNA was isolated from splenocytes 3 hours after

zymosan treatment and TNF expression was measured relative to 18S. Mean±SD. Student t-test,

n=4-7. (B,C) After treatment with zymosan, splenocytes were isolated and incubated with

monensin for 4 hours to inhibit the secretion of TNF-α before cells underwent extracellular

staining for F4/80 and were fixed and permeablized to stain for TNF-α. Levels of TNF-α were

measured by intracellular flow cytometry analysis and the percentage of TNF-α+ cells are shown

in B. Representative histrograms are shown in C. Cells were first gated on F4/80+ cells to enrich

the TNF-α producing population. Mean±SD. Student t-test, n=5.

78

Figure 2-9 D. TNFα and IL1-β promote erythroid differentiation under stress erythropoiesis.

(D) Unfractionated bone marrow was isolated from mice and cultured in SEEM for 7 days at

normoxia in the presence of Shh, SCF, GDF15, and BMP4. Bone marrow cells were cultured at a

concentration of 1x106 cells/mL of SEEM and were pulsed with either TNF-α, IL-1β, or IFN-γ

for 24 hours. Cells were then washed with PBS and replated.. Live cells were counted after 7 days

and expansion of cells (Cells in culture on day 7/Cells plated on day 0) is shown in D. Mean±SD.

n=3-6.

79

Figure 2-9 E-F. TNFα and IL1-β promote erythroid differentiation under stress erythropoiesis.

(E-F) Unfractionated bone marrow was isolated from mice and cultured in SEEM for 7 days as

previously described. BM cultures were pulsed with either TNF-α, IL-1β, or IFN-γ for 24 hours.

Cells were then washed with PBS and replated. To assess stress BFU-E potential, cells were

plated at a concentration of 1x105 cells/well in the presence of GDF15, BMP4, SCF, Shh and Epo

at 1% O2. Stress BFU-Es were counted after five days. E represents the number of stress BFU-Es

per 1x105 cells and F represents total BFU-Es per culture (BFU-Es per 1x105 cells x (Cells in

culture/1x105)). Mean±SD. Student t-test, n=3-8.

80

Figure 2-10 A-C. TNFα increases stress erythropoiesis in human BM cultures.

(A-C) Unfractionated human bone marrow was cultured for 7 days in SEEM at normoxia. Cells

were plated at a concentration of 1x106 cells/mL of SEEM and pulsed for 24 hours with TNF-α.

Live cells were counted at day 7 and expansion of cells (A), stress BFU-Es/1x105 cells (B), and

total stress BFU-Es per culture (C) are shown. n=1.

81

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response to acute anemia. 105, 2741–2748 (2005).

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regulate the expansion of murine stress erythroid progenitors. 109, 4494–4502 (2007).

10. Harandi, O. F., Hedge, S., Wu, D., McKeone, D. & Paulson, R. F. Murine erythroid short-

term radioprotection requires a BMP4-dependent, self-renewing population of stress

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during the recovery from acute anemia. PLoS One 5, (2010).

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83

Chapter 3

How Stress Progenitors, Glucocortcoids, and M2 Macrophages Prevent

Lethal Immune Activation in Response to Zymosan-Induced Inflammation.

Abstract

Corticosteroids play roles in both erythropoiesis and immune modulation. Here will be show

that the induction of glucocorticoids (GCs) during stress erythropoiesis provides critical immune

modulation and prevents lethality following zymosan-induced inflammation. Mice with defects in

stress erythropoiesis die rapidly after treatment with zymosan. However, lethality is not due to an

overwhelming anemic burden since all hematocrits are within the normal range at the time of

death. We will show that inflammation induces high levels of GCs 24 hours after treatment with

zymosan, but this response is diminished in f/f and GDF15-/- mice. Adrenalectomized mice

treated with zymosan experience severe mortality at 24 hours after zymosan treatment, indicating

glucocorticoid production is necessary to survive zymosan-induced inflammation. Diminished

glucocorticoid production in flexed-tail mice results in fewer M2 macrophages in the spleen,

which most likely prevents adequate anti-inflammatory responses in f/f mice and results in

increased mortality. Several pro-inflammatory cytokines are also increased in f/f mice 24 hours

after treatment, indicating a delay in shifting from a pro-inflammatory state to an anti-

inflammatory state.

Introduction

Glucocorticoids (GCs) are steroid hormones produced by the adrenal glands.

Adrenocorticotropic hormone (ACTH) is released by the pituitary gland in response to

physiological stress and stimulates GC production. GCs are known to play a role in the response

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to a variety of physiological stresses, including both infection and acute anemia.1–6 Loss of the

ability to signal through the GC receptor on either T-cells or dendritic cells has been shown to

increase lethality in response to immune activation.1,2 GC signaling is essential in decreasing

levels of pro-inflammatory cytokines to prevent shock during infection, and is known to promote

the development of M2 macrophages, which are typically associated with anti-inflammatory

responses, to resolve inflammation, and promote tissue regeneration and healing.3,7–10

In addition to playing a role in resolving inflammation, GCs have also been shown to be

required in the activation of stress erythropoiesis.11 Fetal liver cells lacking expression of the GC

receptor exhibit diminished proliferation in culture and were smaller and more differentiated,

suggesting that GCs are essential in promoting the proliferation of erythroid progenitors in vivo.11

Additionally, adult mice with deficient GC signaling do not respond to phenylhydrazine-induced

anemia, demonstrating that GCs are necessary to promote stress erythropoiesis. Mice undergoing

chronic physiological stress, which induces ACTH production and stimulates GC production,

have been shown to have increased numbers of stress BFU-Es in the spleen despite not

experiencing acute anemic stress.5

Recent work has also suggested that stress erythroid progenitors (SEPs) may be capable of

exerting immunomodulatory effects and suppressing lethal immune activation. CD71+ erythroid

progenitors in the spleens of neonatal mice were shown to suppress the production of TNF-α

during infection with L. monocytogenes.12 It is hypothesized that immunosuppression by CD71+

erythroid progenitors in the spleen allows for colonization of gut bacteria without lethal immune

activation. While adult CD71+ erythroid cells did not seem to confer the same

immunosuppressive qualities as cells from neonates, it is possible that there is crosstalk between

SEPs and immune cells to appropriately modulate the immune response. We have already

85

demonstrated that SEPs have a unique response to inflammatory stimuli, but it is unclear if SEPs

play a role in dampening the immune response by regulating production of GCs.

In this chapter, we will show that induction of GCs is essential for surviving zymosan-

induced inflammation. f/f and GDF15-/- mice have increased lethality resulting from diminished

or absent GC production and not severe anemia. GCs promote a shift from M1 to M2

macrophages in the spleen and decrease production of pro-inflammatory cytokines, such as

TNFα, IFN-γ, and TGF-β1. In f/f mice, there is a significant reduction in both the frequency and

total numbers of M2 macrophages and increased levels of pro-inflammatory cytokines. In mice

transplanted with unfractionated f/f bone marrow, both GC production and the frequency of M2

macrophages in the spleen are restored. We will show that diminished expression of the enzyme

Cyp11b1 in both f/f and GDF15-/- mice results in lower levels of GCs in response to ACTH.

Results

Glucocorticoids are increased in response to zymosan-induced inflammation.

We have previously shown that stress erythropoiesis is activated after inducing inflammation

with zymosan, and we observed significant mortality in mutants with defects in the stress

erythropoiesis pathway (Figure 3-1A). GDF15-/- are unable to launch a stress erythropoiesis

response due to complete loss of BMP4 signaling, whereas flexed-tail mice (f/f) exhibit a delayed

stress response rather than a complete loss. Colony assays after treatment with zymosan showed

that GDF15-/- mice have no significant increase in colonies at 36 or 72 hours after zymosan, but

f/f mice have decreased numbers of colonies at 36 hours but are near wild type levels by 72 hours

(Figure 3-1B). However, the lethality does not appear to be the result of increased anemic burden

since hematocrits do not begin to drop until day 6 and most mutants die by day 4 (Figure 3-1C).

86

Temperature and body weight were measured after treatment with zymosan as indicators of

shock. Body weight drops dramatically in the first 48 hours after treating mice with zymosan, but

slowly recovers over the next week as the acute inflammation resolves (Figure 3-1D). f/f mice

experience a significant delay in regaining weight between day 4 and day 7 (Figure 3-1D). All the

mice developed hypothermia after being treated with zymosan with body temperatures dropping

an average of 2-3°C (Figure 3-1E). WT mice exhibited a slightly milder drop in temperature

compared with f/f mice (Figure 3-1E), which was not significant but taken together with their

increased mortality could indicate f/f mice are more susceptible to shock after zymosan treatment.

Previous work has demonstrated the importance of GCs in both stress erythropoiesis and

responding to inflammation.1,2,4,5,11 In wild type mice, there is a significant increase in

glucocorticoid levels in the serum at 24 hours after treating with zymosan (Figure 3-2A).

However, both f/f and GDF15-/- mice have a severely reduction in the amount of GCs at 24 hours

(Figure 3-2B). Wild type mice have 193ng/mL of GCs compared to 32ng/mL in f/f mice and

1.5ng/mL in GDF15-/- mice (Figure 3-2B). The adrenal glands are solely responsible for

producing GCs so adrenalectomized (ADX) mice were used to determine the effect of complete

loss of GCs during zymosan-induced inflammation. ADX mice experience severe mortality after

treatment with both the full dose (0.48mg/g) and a half dose (0.24mg/g) with the majority of mice

dying within 24 hours of treatment (Figure 3-2C). This data is consistent with the idea that f/f and

GDF15-/- mice experience increased mortality due to decreased production of GCs.

Decreased expression of Cyp enzymes diminishes GC production in f/f and GDF15-/- mice.

We next examined if the inability of f/f and GDF15-/- mice to produce GCs in response to

inflammation is the result of an inability to respond to the hormone which stimulates

glucocorticoid production in the adrenal glands. Serum levels of GCs were measured 30 minutes

87

after stimulation with ACTH, and GC levels were significantly lower in both f/f and GDF15-/-

mice (Figure 3-3A). In f/f and GDF15-/- mice, there is either a complete loss or a significant

delay in the upregulation of BMP4 (Figure 2-1G). However, it is unknown if BMP4 signaling

affects steroidogenesis and glucocorticoid production. ACTH is secreted by the pituitary gland

and, in the adrenal glands, induces the uptake of cholesterol into mitochondria where it undergoes

a series of enzymatic conversions to become cortisol (Figure 3-3B). We measured mRNA

expression of Cyp11a1, Cyp17a1, and Cyp11b1 30 minutes after stimulation with ACTH to

determine if f/f or GDF15-/- mice have intrinsic defects in the adrenal glands which would

decrease production of GCs. Cyp11b1 expression was dramatically reduced in GDF15-/- mice,

consistent with the low levels of GCs (Figure 3-3C). However, f/f mice had inconsistent

expression of this enzyme with some animals having high levels similar to wild type mice and

other having very low levels more similar to GDF15-/- mice (Figure 3-3C). There also seemed to

be a decrease in levels of Cyp11a1 in both mutants (Figure 3-3C). Cyp17a1 was not detectable in

unstimulated controls and showed no difference in expression between wild type and either f/f or

GDF15-/- mice (Figure 3-3C). These data suggest that BMP4 signaling may play a role in

glucocorticoid production by inducing expression of Cyp enzymes.

Lower levels of GCs leads to an increase in pro-inflammatory cytokines.

Previous work has shown that selective depletion of the glucocorticoid receptor on T-cells

results in lethal immune activation and increased levels of pro-inflammatory cytokines, such as

TNF-α, IFN-γ, and IL-6. In wild type mice, there is a rapid and robust induction of many pro-

inflammatory cytokines within 3 hours of zymosan treatment and then downreguation (Figure 3-

4A). We compared the mRNA expression of several pro-inflammatory cytokines in f/f and wild

type at 24 hours. In wild type mice, there is a drastic reduction in expression after the peak at 3

88

hours (Figure 3-4A). However, we observed a trend in f/f mice of higher levels of expression of

TNF-α, IFN-γ, TGFβ-1, IL-17a and IL-6 (Figure 3-4B). However, these differences were not

statistically significant. IL-10, an anti-inflammatory cytokine, showed no difference between wild

type and f/f mice (Figure 3-4B).

Glucocorticoid production promotes M2 macrophages.

One way that GCs regulate the inflammatory response is through the activation of

macrophages to an M2 phenotype, which promotes anti-inflammatory responses and tissue

remodeling. After treatment with zymosan, wild type mice showed a 2-fold decrease in the M1-

associated gene IRF5 at 24 hours and a 200-fold induction of Arg-1, an M2-assocaticiated gene

(Figure 3-5 A,B) While IRF5 was decreased at 24 hours in f/f mice, it was expressed at slightly

higher levels than in WT mice (Figure 3-5C). There was no significant difference in the

expression of Arg-1 in wild type and f/f mice at 24 hours. (Figure 3-5 D). Using flow cytometry

to determine the distribution of splenic macrophages after treatment with zymosan revealed a

shift from M1 macrophages (F4/80+CD45+CD68+CD11b+CD11c+CD206-) to M2 macrophages

(F4/80+CD45+CD68+CD11b+CD11c-CD206+), occurring between 24 and 48 hours (Figure 3-5

E,F). This is consistent with the mRNA expression of IRF5 and Arg-1, indicating a shift from

pro-inflammatory macrophages to more anti-inflammatory macrophages. Flexed-tail mice

experience a similar shift from the M1 macrophage to M2. However, f/f mice have a significantly

lower frequency of M2 macrophages compared to wild type mice (Figure 3-5 E,F). In wild type

mice, M2 macrophages at 48 hours make up over 40% of the F4/80+CD45+CD68+CD11b+

population, but in f/f mice, M2 macrophages are only 25% of this population (Figure 3-5 F).

Additionally, spleens of f/f mice are consistently smaller in terms of total numbers of cells (data

not shown), and as a result, the total number of M2 macrophages is also significantly reduced in

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f/f mice compared to wild type (Figure 3-5 G). In contrast, f/f mice have no difference in either

the percentage or total numbers of M1 macrophages compared with wild type animals (Figure 3-5

E-G).

We transplanted either wild type or f/f bone marrow into wild type recipients. There is no

difference in the frequency of stress BFU-Es per 1x105 cells between mice receiving wild type

donor cells (WT-WT) or mice which received f/f donor cells (f/f-WT) (Figure 3-6 A). However

f/f-WT spleens contain fewer cells than WT spleens, and thus f/f-WT mice have fewer BFU-Es

per spleen than WT-WT mice (Figure 3-6 B). Serum levels of glucocorticoid were measured, and

f/f-WT mice have significantly increased levels of GCs at 24 hours compared with WT-WT mice

(Figure 3-6 C). This suggests that the decrease in GC production seen in f/f mice is not a result of

delayed stress erythropoiesis in the spleen but an intrinsic problem with GC production in the

adrenal glands. In addition to rescuing glucocorticoid production, the frequency of M2

macrophages in the spleens of f/f-WT mice is similar to that of WT-WT mice (Figure 3-6 D,E).

f/f-WT mice do have fewer M2 macrophages in terms of total numbers though due to the smaller

spleens of f/f-WT animals (Figure 3-6 F).

Discussion

We have shown here that GC production is essential for survival of zymosan-induced

inflammation and acts to promote a shift from M1 macrophages to M2 macrophages in the

spleen. Mice with diminished production of GCs all experience severe mortality after treatment

with zymosan. The data suggest that a delay in this shift from M1 to M2 macrophages leads to

overproduction of pro-inflammatory cytokines such as TNF-α, IFN-γ, IL-17a and IL-6. However,

more work is needed to confirm this. f/f mice have a milder phenotype is response to zymosan

than GDF15-/- mice, most likely due to the fact that GDF15-/- have a complete loss of BMP4

90

expression whereas BMP4 expression is delayed but present f/f mice. Therefore, analysis of M1

versus M2 macrophages and profiling cytokine expression in GDF15-/- mice, where the

phenotype is stronger, would provide more evidence about this shift in macrophage populations

and the idea that these mice die as a result of a lethal immune activation. Injections of

dexamethasone, a synthetic corticosteroid, were unsuccessful in rescuing the M1 to M2

macrophage shift in f/f mice (data not shown). However, physiological doses of dexamethasone

were not able to be given and could explain why injections of corticorsteroids were unable to

boost production of M2 macrophages. It is also possible that longer exposure to corticosteroids is

required for the transition from M1 to M2 and this is not reproducible with single injections of

dexamethasone.

We have also demonstrated a previously unknown relationship between BMP4 signaling

and GC production. Our data clearly demonstrate that mice with absent or delayed BMP4

expression have decreased production of GCs both in response to zymosan-induced inflammation

as well as ACTH stimulation in vivo. The transcription of Cyp11b1 is greatly decreased in

GDF15-/- mice stimulated with ACTH. f/f mice show greater variability in the expression of

Cyp11b1 with some mice exhibiting decreased expression and others having expression levels

similar to wild type. Cyp11a1 also appeared to be lower in GDF15-/- and f/f mice after

stimulation with ACTH. The reduction in corticosteroid production in response to ACTH clearly

indicates that inability to signal effectively through BMP4 is important in steroidogenesis in the

adrenal glands, and this is supported by data from bone marrow transplants where there is

decreased GC production in f/f mice is not a result of the delay in stress erythropoiesis but most

likely due to an intrinsic problem in the adrenal glands of f/f animals. However, it is unclear what

the relationship is between Cyp11a1/Cyp11b1 transcription and BMP4, and further studies are

needed to clarify how BMP4 signaling is impacting the expression of Cyp enzymes. In addition to

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understanding the impact on transcription of Cyp enzymes, protein levels should be assessed to

confirm that there is impairment in these enzymes affecting corticosteroid production.

Materials and Methods

Mice. C57BL/6 mice were purchased from Taconic Biosciences, Inc. All mice were 6-12

weeks old. GDF15-/- mice were provided by Dr. Se-Jun Lee at Johns Hopkins.13

Adrenalectomized (ADX) mice were purchased from Jackson Laboratories and were maintained

on 0.9% saline water. All procedures have been approved by the Institutional Animal Care and

Use Committee of the Pennsylvania State University.

Zymosan-Induced Generalized Inflammation. Mice were first treated with 40µg/200µL of

lipopolysaccharide from Escherichia coli 0128:B12 (Sigma-Alrich L2887) and followed six days

later by Zymosan A from Saccharomyces cerevisiae (Sigma-Aldrich Z4250) at a concentration of

0.48mg/g. All treatments were administered by intraperitoneal injection. Surface body

temperature was measured with an infrared thermometer (VWR 36934-182).

Colony Assays. Splenocytes were plated in methylcellulose media (Stem Cell Technologies,

M3334) at a concentration of 1x105 cells/well in a 12-well tissue culture plate. BMP4 (15ng/mL),

SCF (50ng/mL), Shh (25ng/mL), GDF15 (15ng/mL), and Epo (3U/mL) were added to

methylcellulose media to assay stress BFU-E formation. Cells were incubated at 1% O2 for 5 days

before counting BFU-Es. BFU-Es were stained with benzidine solution for counting.

ELISAs. Serum levels of corticosterone were determined using a commercially available

ELISA kit (Enzo ADI-900-097) according to manufacturer’s instructions.


Biosciences) and an LSR-II Fortessa flow cytometer (BD Biosciences). Flow antibodies: F4/80

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PE-Cy7 (Clone BM8, Biolegend 123112), CD68 Brilliant Violet 421 (), CD45 FITC (), CD11b

PerCP-Cy5.5 (), CD11b APC(), CD11c PE (), CD2016 APC ().

ACTH Stimulation. Mice were dosed with 5U/kg of adrenocorticotropic hormone (ACTH)

from porcine pituitary (Sigma A6303). Serum was isolated 30 minutes after stimulation with

ACTH to measure production of corticosteroids.

Transplant of stress progenitors. Recipient mice were irradiated with a single dose of

950Rads and donor cells were administered by IV injection. Recipeints received 5x105 donor

cells and were allowed to recover for 10 weeks prior to being treated with zymosan.

Statistics. P-values were determined using the Student’s t-test (2-tailed) or Mann-Whitney

test, as deemed appropriate. Significance was determined as * p<0.05, ** p<0.01, *** p<0.001.

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Figures

Figure 3-1 A-C. Flexed-tail (f/f) mice have increased mortality after zymosan-induced

inflammation.

A) WT, f/f and GDF15-/- mice were treated with zymosan and survival was assessed over the first

week after treatment. Mantel-Cox test, n=13-29. B) Stress BFU-Es formation after treatment with

zymosan was measured in wild type, f/f, and GDF15-/- mice. Splenocytes were plated at a

concentration of 1x105 cells/well in methylcellulose media in the presence of Shh, SCF, GDF15,

and BMP4 at 1% O2. BFU-Es were scored after 5 days. Mean±SD. Student t-test, n=3-8. C) To

determine if stress erythropoiesis buffers hematocrit values in response to inflammation, blood

was collected in EDTA-coated microcapillary tubes after zymosan treatment and spun to

determine hematocrit values. Mean±SD. n=2-10.

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Figure 3-2 D-E. Flexed-tail (f/f) mice have increased mortality after zymosan-induced

inflammation.

(D,E) Body weight and temperature, used as indicators of shock, were measured both before after

treatment with zymosan in wild type and f/f mice. Body weight is expressed as total weight lost

(g). Temperature reflects body surface temperature measured on the mouse’s underbelly by

infrared thermometer. Mean±SD. Student t-test, n=4-14.

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Figure 3-2. Glucocorticoid production is critical to surviving zymosan-induced inflammation.

(A,B) Glucocorticoids increase in response to inflammation and are also regulated temporally.

For all GC experiments, studies were started between 8 and 9am to account for daily incraeses.

Serum was collected at each time point and serum levels of corticosteroids were measured by

ELISA. Mean±SD. Student t-test, n=3-9 (C) WT and ADX mice were treated with either a full

dose (0.48mg/g) or a half-dose (0.24mg/g) of zymosan due to increased mortality of ADX mice.

Survival is shown over the first week in hours. Mantel-Cox test. n=4-8

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Figure 3-3 A. GDF15-/- and f/f mice have diminished GC production due to decreased expression

of Cyp11b1.

(A) Mice were treated with ACTH between 8 and 9am and serum was isolated 30 minutes after

stimulation to assess whether f/f mice have innate defects in corticosteroid production. Serum

corticosteroid levels were measured by ELISA. Mean±SD. Student t-test, n=3-7.

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Figure 3-3 B. GDF15-/- and f/f mice have diminished GC production due to decreased expression

of Cyp11b1.

(B) Schematic of enzymatic conversion of adrenocorticotropic hormone (ACTH) to cortisol in

mitochondria of the adrenal glands. Enzymes required for each step are shown in red.

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Figure 3-3 C. GDF15-/- and f/f mice have diminished GC production due to decreased expression

of Cyp11b1.

(C) To determine if defects in transcription of these enzymes is the cause of lower GC production

in f/f mice, RNA was isolated from adrenal glands 30 minutes after stimulation with ACTH and

expression of Cyp11a1, Cyp11b1, and Cyp17a1 were measured relative to 18S. Mean±SD.

Student t-test. n=2-6

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Figure 3-4 A. f/f mice have lightly elevated levels of pro-inflammatory cytokines.

(A) Wild type mice were treated with zymosan and RNA was isolated from whole spleen after

treatment between 0 and 48 hours. Expression of IL-6, IL-10, IL-12a, IL-17a, IFN-γ, TNF, and

TGFβ1 were measured relative to 18S to assess the inflammatory response. Mean±SD. n=3-9

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Figure 3-4 B. f/f mice have lightly elevated levels of pro-inflammatory cytokines.

(A) To determine if the high mortality in f/f mice is the result of overproduction or prolonged

production of pro-inflammatory cytokines, RNA was isolated from whole spleen of either WT or

f/f mice after treatment with zymosan. Expression of IL-6, IL-10, IL-12a, IL-17a, IFN-γ, TNF,

and TGFβ1 were measured relative to 18S. Mean±SD.Mann-Whitney test, n=4-10.

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Figure 3-5 A-D. GCs promote M2 macrophages to prevent immune lethality.

(A,B) GCs can result in a shift to M2 macrophages, which produce more anti-inflammatory

cytokines and resolve inflammation. RNA was isolated from whole spleen of wild type mice at 0

or 24 hours after zymosan and expression of IRF5 and Arg-1 was measured to determine if there

is more of an M1 or M2 RNA profile. (C,D) RNA from f/f and wild type mice was isolated from

whole spleen at 24 hours and expression of IRF5 and Arg-1 was measured relative to 18S.

Mean±SD. Student t-test, n=3

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Figure 3-5 E. GCs promote M2 macrophages to prevent shock.

(E) Wild type and f/f mice were treated with zymosan to examine M1 and M2 markers on splenic

red pulp macrophages. Splenocytes were isolated between 0 and 72 hours after treatment and

analyzed by flow cytometry for expression of CD68, F4/80, CD11b, CD45, CD11c, and CD206.

Plots shown are representative images for each group. Cells were first gated on the

CD45+F4/80+CD68+C11b+ population. n=4

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Figure 3-5 F-G. GCs promote M2 macrophages to prevent immune lethality.

(F) Based on previously shown flow cytometry data, frequencies of M1 macrophages

(CD45+F4/80+CD68+C11b+CD11c+CD206-) and M2 macrophages (CD45+F4/80+CD68+C11b+

CD11c-CD206+) populations are shown for each group. (G) Total numbers of M1 or M2

macrophages per spleen were calculated from the population frequencies and cell counts for each

mouse. Student t-test. Mean±SD. Student t-test, n=4.

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Figure 3-6 A-C. Decreased M2 macrophages in f/f mice is due to decreased GC production.

(A) To determine if the decreased frequency of M2 macrophages is the result of lower GC

production in f/f mice, WT or f/f donor unfractionated bone marrow was transplanted into WT

recipients where GC production should be normal. Mice were allowed to recover 8 weeks prior to

being treated with LPS and zymosan. Splenocytes were isolated between 0 and 72 hours after

zymosan treatment and plated at a concentration of 1x105 cells/well in the presence of Shh, SCF,

GDF15, and BMP4 at 1%O2. BFU-Es were scored after 5 days. (B) Total numbers of BFU-Es per

spleen were calculated from cell counts for each spleen and average number of BFU-Es/1x105

cells. (C) Serum was isolated between 0 and 72 hours after zymosan treatment and levels of

glucocorticoids were measured by ELISA. Mean±SD. Student t-test, n=3-4.

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Figure 3-6 D. Decreased M2 macrophages in f/f mice is due to decreased GC production.

(D) Transplant recipients were treated with zymosan and splenocytes were isolated between 0 and

72 hours. Splenic macrophages were analyzed by flow cytometry for expression of CD68, F4/80,

CD11b, CD45, CD11c, and CD206. Plots shown are representative images for each group. Cells

were first gated on the CD45+F4/80+CD68+C11b+ population. n=3-4

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Figure 3-6 E-F. Decreased M2 macrophages in f/f mice is due to decreased GC production.

(E) Frequencies of M1 macrophages (CD45+F4/80+CD68+C11b+CD11c+CD206-) and M2

macrophages (CD45+F4/80+CD68+C11b+ CD11c-CD206+) populations are shown for each group

based on previously shown flow cytometry data. (F) Total numbers of M1 or M2 macrophages

per spleen were calculated from the population frequencies and cell counts for each mouse.


107

References

1. Brewer, J. a et al. T-cell glucocorticoid receptor is required to suppress COX-2-mediated

lethal immune activation. Nat. Med. 9, 1318–1322 (2003).

2. Li, C. C., Munitic, I., Mittelstadt, P. R., Castro, E. & Ashwell, J. D. Suppression of

Dendritic Cell-Derived IL-12 by Endogenous Glucocorticoids Is Protective in LPS-

Induced Sepsis. PLOS Biol. 13, e1002269 (2015).

3. Kleiman, a. et al. Glucocorticoid receptor dimerization is required for survival in septic

shock via suppression of interleukin-1 in macrophages. FASEB J. 26, 722–729 (2012).

4. Vignjević, S. et al. Chronic psychological stress activates BMP4-dependent

extramedullary erythropoiesis. J. Cell. Mol. Med. 18, 91–103 (2014).

5. Vignjevic, S. et al. Glucocorticoid receptor mediates the expansion of splenic late

erythroid progenitors during chronic psychological stress. J. Physiol. Pharmacol. 66, 91–

100 (2015).

6. Jamieson, A. M., Yu, S., Annicelli, C. H. & Medzhitov, R. Influenza Virus-Induced

Glucocorticoids Compromise Innate Host Defense against a Secondary Bacterial

Infection. Cell Host Microbe 7, 103–114 (2010).

7. Gratchev, a., Kzhyshkowska, J., Utikal, J. & Goerdt, S. Interleukin-4 and dexamethasone

counterregulate extracellular matrix remodelling and phagocytosis in type-2 macrophages.

Scand. J. Immunol. 61, 10–17 (2005).

8. Gratchev, A. et al. Activation of a TGF-beta-specific multistep gene expression program

in mature macrophages requires glucocorticoid-mediated surface expression of TGF-beta

receptor II. J. Immunol. 180, 6553–65 (2008).

9. Schmieder, A. et al. Synergistic activation by p38MAPK and glucocorticoid signaling

mediates induction of M2-like tumor-associated macrophages expressing the novel CD20

homolog MS4A8A. Int. J. Cancer 129, 122–32 (2011).

10. Ehrchen, J. et al. Glucocorticoids induce differentiation of a specifically activated, anti-

inflammatory subtype of human monocytes. Blood 109, 1265–74 (2007).

11. Bauer, A. et al. The glucocorticoid receptor is required for stress erythropoiesis. Genes

Dev. 13, 2996–3002 (1999).

12. Elahi, S. et al. Immunosuppressive CD71+ erythroid cells compromise neonatal host

defence against infection. Nature 504, 158–62 (2013).

13. Hsiao, E. C. et al. Characterization of growth-differentiation factor 15, a transforming

growth factor beta superfamily member induced following liver injury. Mol. Cell. Biol. 20,

3742–3751 (2000).

108

Chapter 4

Sf-Ron plays a role in regulating the differentiation of erythroid progenitors

during recovery during acute anemic stress.

Abstract

Short form stem cell-derived tyrosine kinase (Sf-Ron) has been shown to play an essential

role in the development of erythroleukemia. Mice lacking Sf-Ron are resistant to Friend virus-

induced erythroleukemia, which requires the expansion of stress erythroid progenitors in the

spleen.1,2 Friend virus induces BMP4 to drive the expansion of stress progenitors, and activation

of Sf-Ron induces PU.1 expression to promote proliferation.2,3 Here we will show that Sf-Ron is

not only important in expansion of stress progenitors during erythroleukemia, but it also plays a

role in proliferation and differentiation of erythroid progenitors during the recovery from acute

anemic stress. Mice lacking Sf-Ron have increased mortality and delayed recovery from

phenylhydrazine (PHZ)-induced anemia associated with insufficient expansion of early erythroid

progenitors, resulting in the production of too few mature erythrocytes.

Introduction

Friend virus infection causes erythroleukemia in adult mice. Early work in the field led to the

development of a model where Friend virus-induced erythroleukemia developed through a two

stage progression. The first stage was characterized by expansion of infected cells in the spleen,

leading to splenomegaly and erythrocytosis in mice infected with the polycythemia-inducing

strain of Friend virus, FvP. (For this thesis, Friend virus will refer only to the Fv-P strain.) The

109

massive production of new erythrocytes was proposed to occur because the Friend virus envelope

protein gp55 interacted with and activated the Erythropoietin receptor (EpoR), leading to Epo-

independent erythroid development. The transition to leukemia was proposed to occur in the

second stage where new mutations, proviral insertional activation of Spi1/PU.1 and mutation of

p53, occurred in a subset of cells leading to their leukemic transformation.2,4,5

A new model for the pathogenesis of Friend virus grew out of work from our lab analyzing

disease modifying gene variants that segregated in inbred strains of mice. These modifying genes

fell into three broad categories – genes that affect the retroviral life cycle such as Fv1 and Fv4,

genes that affect the immune response to Friend virus such as Fv3 and Rfv1, and genes that

affected the proliferation or differentiation of infected cells such as Fv2, flexed-tail, and W. It is

the study of the latter category that has led to the development of a new model for the progression

of Friend virus-induced leukemia. Flexed-tail and W encode Smad5 and Kit receptor respectively.

Both of these genes and the signaling pathways they regulate are essential for stress

erythropoiesis. Work from a previous student, Aparna Subramanian, showed that Friend virus-

induced the stress erythropoiesis pathway during infection. The stress erythroid progenitors were

the targets for the virus.

Stress erythropoiesis is a process where new erythrocytes are produced rapidly in the spleen

in response to acute anemic stress. It proceeds in three stages. Initially, progenitor cells that have

stem cell characteristics rapidly expand in the spleen. During the second stage, the progenitor

cells transition from proliferating “stem cell like” stress progenitors to progenitors committed to

terminal differentiation. The progenitors at each stage express a distinct set of cell surface

markers. Analysis of Friend virus infected cells showed that cells capable of Epo-independent

development corresponded to late stage stress erythroid progenitors that have already committed

to terminal differentiation. In contrast, the Friend virus infected cells that caused leukemia

corresponded to the early stem cell like stress erythroid progenitors. These cells acted as leukemia

110

stem cells (LSCs). Analysis of the LSCs showed that they have proviral insertional activation of

Spi1/PU.1 but were p53 wildtype.

These observations led to a new model for Friend virus disease where, early in infection, late

stage stress progenitors are infected, causing them to proliferate and differentiate and leading to

erythrocytosis. A less likely event in the infection of an early stem cell like stress erythroid

progenitor, and the rare development of an LSC requires that the proviral insertion activate the

expression of Spi1/PU.1 locus. Once an LSC is generated, its progeny rapidly take over the

population of infected cells (Figure 1A).

During Friend virus infection, expansion of erythroid progenitors in the spleen occurs in an

Epo-independent manner, and the gene Friend virus susceptibility 2 (Fv2) affects the ability of

erythroid cells to proliferate during Friend virus infection. Cloning of the Fv2 locus revealed it

encodes the gene Ron (stem cell-derived tyrosine kinase, also known as STK), a member of the

Met subfamily of receptor tyrosine kinases.6,7 STK/Ron is the murine homologue of the human

Ron and the avian Sea receptors. A retroviral oncogene version of Sea, v-Sea, is known to cause

erythroblastosis.6 The naturally occurring truncated form of STK/Ron (referred to as Sf-Ron) is

highly expressed in hematopoietic stem cells and is required for susceptibility to Friend virus.6,7

Sensitive mouse strains, such as FVB/NJ and DBA2/J, express high levels of Sf-Ron, whereas

resistant strains, which include C57BL/6 mice and related strains, have either decreased or absent

expression of the truncated form of Ron but normal expression of full length Ron. The reduction

in expression was linked to a three nucleotide deletion in the promoter region for Sf-Ron that

impairs GATA binding. 7

The Friend virus envelope protein, gp55, interacts with Sf-Ron, resulting in constitutive

phosphorylation of Sf-Ron and activation of downstream signaling molecules.8 Sf-Ron retains

both the transmembrane domain and the cytoplasmic kinase domain but lacks the extracelluar

domain.6 The cytoplasmic domain, in addition to the kinase activity, also acts as a docking site

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for SH-2 domain containing proteins, such as Grb2, Gab1, PI3 kinase, Ship-1, and SHP2. Grb2

binding to Sf-Ron is required for the recruitment of Gab2 and proliferation of Epo-independent

erythroid cells during Friend virus infection.1,9 Gab2 contains a binding site for Stat3, which is

recruited and subsequently phosphorylated by Sf-Ron.10 Stat3 then activates purine rich box-1

(PU.1), and in Friend virus infection, inhibits differentiation of the proliferating erythroid

progenitors.3

PU.1 not only plays an important role in the development of erythroleukemia but is also a

crucial regulator of hematopoietic development.11–15 It is required for commitment to the myeloid

lineage in later stages of hematopoiesis.11 However, it also is important in the maintenance of

hematopoietic stem cells (HSCs).16,17 PU.1 regulates several genes involved in the cell cycle,

repressing cell-cycle activators and inducing cell-cycle inhibitors to ensure that HSCs are not

exhausted prematurely. HSCs from hypomorphic PU.1 mice exhibit diminished long-term

repopulation in serial transplants, confirming the importance of PU.1 in stem cell maintenance.16

Additionally, analysis of cells from mouse fetal liver, the primary site of definitive erythropoiesis

in the mouse embryo, showed that PU.1-/- mice have reduced numbers of BFU-Es in the fetal

liver at days E14.5 and E16.5, which is attributed to an inability of PU.1-/- mice to maintain

HSCs in the fetal liver and results in failure of multiple hematopoietic lineages.18 For erythroid

differentiation, GATA-1 expression increases and is responsible for repression of PU.1.13,14

Expression of PU.1 in hematopoietic progenitors is dynamic with low levels required to maintain

HSCs and their capacity to self-renew but also acting later to determine lineage commitment by

being either upregulated or repressed.

While Sf-Ron has been studied extensively in context of Friend virus-induced

erythroleukemia, its role in normal hematopoiesis is unclear. The requirement of Sf-Ron in the

proliferation of stress erythroid progenitors during Friend virus through activation of PU.1

expression, a well-established regulator of various aspects of hematopoiesis, suggest that Sf-Ron

112

might also play a role in the proliferation of stress erythroid progenitors under acute anemic stress

as well. Here we show that Sf-Ron also plays a role in regulating the proliferation of stress

progenitors during phenylhydrazine-induced anemia, and in the absence of Sf-Ron, progenitors

have diminished proliferation before differentation, resulting in the production of fewer new

erythrocytes.

Results

Sf-Ron during recovery from acute anemia.

Previous data from our lab showed that mice lacking Sf-Ron have delayed recovery from

bone marrow transplant, but how Sf-Ron affects the proliferation or differentiation of stress

progenitors is not understood (unpublished data, Lei Shi). We sought to determine the role of Sf-

Ron during stress erythropoiesis in vivo by utilizing phenylhydrazine (PHZ) to induce acute

hemolytic anemia and in vitro using a culture system of stress progenitors developed in our lab.19

PHZ treatment lyses approximately half of all circulating erythrocytes within 24 hours of

treatment. This rapid onset of anemia results in a robust activation of the stress erythropoiesis

pathway.

After treatment with PHZ, we observed a significant increase in the expression of Sf-Ron

mRNA in the spleen (Figure 4-2). Sf-Ron was upregulated approximately 17-fold at 24 hours

following administration of PHZ. This rapid increase in mRNA levels suggests that Sf-Ron plays

a role in the expansion of early stress erythroid progenitors during stress erythropoiesis.

Progenitors capable of forming stress BFU-Es do not peak until 36 hours post-PHZ. Utilizing Sf-

Ron-/- mice, we were able to characterize the recovery from PHZ in the absence of Sf-Ron

expression. Sf-Ron-/- mice have increased mortality after inducing anemia (Figure 4-3A), with

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50% of mice dying between days 6 and 7. Comparatively, all wild type FVB mice survive PHZ

treatment and have recovered by day 7. Hematocrit is the percentage of blood volume that is

erythrocytes, and Sf-Ron-/- mice have a delay in hematocrit recovery compared to wild type,

indicating a lag in production of new erythrocytes or a reduction in total number of erythrocytes

produced (Figure 4-3B).

Sf-Ron-/- mice have decreased production of stress BFU-Es compared to wild type mice

during recovery from PHZ-induced anemia. There is a modest increase in the number of stress

BFU-Es following PHZ treatment in Sf-Ron-/- mice, and for the first four days, there is no

significant difference from wild type mice. Stress BFU-E production plateaus in Sf-Ron-/- mice

at day 4 and is significantly lower than wild type mice at days 6 and 8 (Figure 4-4A). This is

consistent with the idea that Sf-Ron-/- mice may experience increased mortality due to a

reduction in the total number of new erythrocytes produced in response to PHZ treatment. There

is no significant difference in the numbers of BFU-Es in the bone marrow, and in both wild type

and Sf-Ron-/- mice, there is a transient dip in BFU-Es after treatment with PHZ (Figure 4-4B).

Sf-Ron prevents premature differentiation of stress erythroid progenitors.

Stress erythropoiesis can be divided into two phases. The first phase involves the

expansion of progenitors cells in the spleen to ensure that a sufficient number of new erythrocytes

are produced to survive the anemic stress. It requires signals such as hedgehog (most likely Ihh in

vivo), stem cell factor (SCF), bone morphogenetic protein 4 (BMP4), and growth and

differentiation factor 15 (GDF15).19–22 In this first phase, cells are primarily

Kit+Sca+CD34+CD133+ and will remain that way until exposed to erythropoietin (Epo). The

second phase of stress erythropoiesis relies on increased levels of Epo to act as a transition signal

and switch progenitors from expansion and self-renewal to differentiation. Once cells shift to

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differentiation, they lose expression of CD34 and CD133, respectively (Figure 4-5 A).19 Although

Epo regulates erythroid differentiation by binging EpoR in progenitor cells, the transition from

expanding to differentiating stress erythroid progenitors is regulated by Epo signaling in splenic

macrophages (Jie Xiang, unpublished data).

Our lab has developed an in vitro culture system which is capable of recapitulating the

development and differentiation of stress progenitors we observe in vivo.19 Whole bone marrow is

first cultured for 7 days in stress erythroid expansion media (SEEM, containing BMP4, GDF15,

Shh, SCF) at normoxia (20% O2). Cells are then shifted to stress erythroid differentiation media

(SEDM, containing BMP4, GDF15, Shh, SCF, Epo) and grown under hypoxic conditions (1%

O2) for 3 days. Cells grown in SEEM are Kit+Sca+CD34+CD133+, and once shifted to SEDM,

they become Kit+Sca+CD34-CD133- (Figure 4-5 A,B).19

Anaylsis of wild type and Sf-Ron-/- bone marrow after culture in SEEM revealed that

there was no difference in the frequencies of population I (Kit+Sca+CD71Ter119) and population

II (Kit+Sca+CD71Ter119) cells after culture with SEEM (Figure 4-6 A,B). However, Sf-Ron-/-

cultures have an increase in Kit+Sca+CD34-CD133+ cells compared to wild type cultures (Figure

4-6 C,D). In wild type mice, Epo signaling is required for cells to begin to lose expression of

CD34.19 CD34 and CD133 are used to distinguish different stages of immature erythroid

progenitors, all of which are contained with population I. Previously, it has been shown that

Kit+Sca+CD34+CD133+ cells form fewer stress BFU-Es than Kit+Sca+CD34-CD133- cells.19 Sf-

Ron-/- bone marrow cells produce significantly more stress BFU-Es after culture with SEEM

than wild type cells (Figure 4-6 E). Total numbers of cells in each culture were equivalent,

indicating there is no difference in the proliferative capacity of the cells (Figure 4-6 F).

Unfractionated bone marrow from either FVB or Sf-Ron-/- mice was labeled with PKH26 and

cultured for 7 days in SEEM to determine if Sf-Ron-/- SEPs divide at a faster rate than FVB

SEPs. There was no significant difference in the frequencies of PKH26+ and PKH26- populations

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after 7 days culture (Figure 4-6 G,I). The PKH26+ population was also split into five fractions (I,

II, III, IV, and V) with V being the most PKH26+ cells and I being the least PKH26+ cells, and

there was no difference in the frequencies between FVB and Sf-Ron-/- cells in any of the five

subsets of PKH26+ population (Figure 4-6 H,J). These data indicate that Sf-Ron-/- SEPs are more

mature than FVB SEPs after 7 days in SEEM, but this is not due to increased proliferation.

After cells were shifted to SEDM and hypoxia, Sf-Ron-/- and wild type progenitors are

primarily Kit+Sca+CD34-CD133- with no differences in the population frequencies (Figure 4-7

C,D). Sf-Ron-/- cultures did tend to have a slight increase in population II cells, a more mature

population, than wild type cultures (Figure 4-7 A,B). However, this difference was not

statistically significant. When plated for stress BFU-Es, Sf-Ron-/- cells have significantly

decreased numbers of BFU-Es compared to wild type (Figure 4-7 E). This is potentially due to

the fact that the early stages of the culture contained more mature cells, which after transitioning

to Epo, continue to mature and are no longer capable of forming BFU-Es. This is consistent with

the trend of Sf-Ron-/- cultures having an increased frequency of population II cells after culture

in SEDM.

Discussion

Our in vivo data suggest that Sf-Ron plays a role in the development of stress erythroid

progenitors (SEPs). Sf-Ron-/- mice experience high levels of mortality during recovery from

PHZ-induced anemia as well as from bone marrow transplant (Lei Shi, data not shown). It was

previously known that Sf-Ron is critical in the development of erythroleukemia, but our data also

shows that it is important in the maintenance, proliferation, and differentiation of SEPs. It has

been noted in the in vivo models, particularly in bone marrow transplant, that if Sf-Ron-/- mice

have little to no blood taken over the course of recovery they have increased survival (Lei Shi,

116

data not shown). This indicates that Sf-Ron-/- are essentially on the cusp of making enough new

red cells to survive the anemic crisis and any additional stress (i.e. repeated blood draws) are

enough to compromise their survival.

Culturing Sf-Ron-/- bone marrow under stress erythropoiesis conditions revealed that there

are more mature cells after 7 days in SEEM compared with wild type controls. This is seen both

in the increased number of stress BFU-Es and the increased frequency of Kit+Sca+CD34-CD133+

cells, which represent a more mature SEP as previously shown by Xiang et al.19 Additionally,

once cells are shifted to SEDM, Sf-Ron-/- show decreased numbers of stress BFU-Es, suggesting

premature exhaustion of the progenitor population. These data support the idea that Sf-Ron plays

a role in maintaining SEPs in an immature, self-renewing state in order to allow for sufficient

expansion of the progenitor population. However, there is no indication that the more mature

phenotype of these cells is due to an increase in proliferation. PKH26 labeling shows that Sf-Ron-

/- SEPs have similar frequencies of PKH26 positive and negative cells, suggesting they are

dividing at a rate comparable to FVB cells.

These data suggest a model where Sf-Ron signaling acts early to maintain a population of

immature SEPs, potentially by regulating the expression of PU.1 and some of its downstream

targets that are cell cycle regulators. In the absence of Sf-Ron, cells prematurely begin to

differentiate, leading to a smaller pool of progenitors and decreased output of new erythrocytes

(Figure 4-8A). This data has demonstrated the importance Sf-Ron signaling in the maintenance of

SEPs during stress erythropoiesis, not just during the establishment of erythroleukemia. However,

it is still unclear how Sf-Ron is exerting its effects on SEPs, and further studies will be required to

understand the mechanism of how Sf-Ron acts on SEPs.

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Materials and Methods

Mice. FVB mice were purchased from Taconic Biosciences, Inc. All mice were 6-12 weeks

old. Sf-Ron-/- mice were previously described and provided by Dr. Susan Waltz at University of

Cincinnati (Cincinnati, OH). All procedures have been approved by the Institutional Animal Care

and Use Committee of the Pennsylvania State University.

Phenylhydrazine-Induced Anemia. Phenylhydrazine (PHZ, Sigma 114715) was dissolved in

PBS and given administered to mice by subcutaneous injection. Mice were given 100mg/kg PHZ.

Blood was collected in heparinized microcapillary tubes and spun to measure hematocrit values

during recovery.


Biosciences) and data was analyzed in FlowJo v10. Flow antibodies: c-Kit Alexa 647 (Clone

2B8, Biolegend 105818), Sca-1 PE-Cy7 (Clone D7, BD Biosciences BDB558162), Ter119 PE

(Clone Ter119, BD Biosciences BDB553673), CD71 FITC (Clone C2, BD Biosciences

BDB553266), F4/80 PE-Cy7 (Clone BM8, Biolegend 123112).

Murine and Human Cell Cultures. Murine bone marrow was isolated from the femurs and

cultured in stress erythropoiesis expansion media (SEEM) for 7 days at normoxia as previously

described and plated for colony assays.11 Briefly, SEEM is composed of Iscove’s modified

Dulbecco’s media (IMDM) containing 10%FBS, 10µg/mL Insulin, 200µg/mL holo-transferrin,

2mM L-glutamine, 10µg/mL ciprofloxacin, 1% BSA, 7µL/L 2-mercaptoethanol, GDF15

(30ng/mL; Biomatik), BMP4 (15,ng/mL; R&D Systems), Shh (25ng/mL; GoldBio), and SCF

(50ng/mL; GoldBio).

Colony Assays. Murine splenocytes or bone marrow were plated in methylcellulose media

(Stem Cell Technologies, M3334) at a concentration of 1x105 cells/well in a 12-well tissue

culture plate. BMP4 (15ng/mL), SCF (50ng/mL), Shh (25ng/mL), GDF15 (15ng/mL), and Epo

118

(3U/mL) were added to methylcellulose media to assay stress BFU-E formation. Cells were

incubated at 1% O2 for 5 days before counting BFU-Es. BFU-Es were stained with benzidine

solution for counting. Epo (3U/mL) and IL-3 (25ng/mL) were added to methylcellulose to assay

BFU-Es in bone marrow. For typical BFU-Es from bone marrow, cells were incubated for 7 days

at normoxia before being stained with benzidine and scored.

mRNA Isolation and Gene Expression Analysis. Total RNA was isolated using TriZol reagent

(Invitrogen 15596). cDNA was generated using the high capacity cDNA synthesis kit (Applied

Biosystems). Quantitative reverse transcription PCR (qRT PCR) was carried out using Taqman

probes and an ABI7300 real-time PCR system.

PKH26 Labeling. A commercially available PKH26 labeling kit (Sigma MINI26) was

purchased and used according to the manufacturer’s instructions. PKH26 labeling was analyzed

using a BD Accuri C6 flow cytometer.

Statistics. P-values were determined using the Student’s t-test (2-tailed). Significance was

determined as * p<0.05, ** p<0.01, *** p<0.001.

119

Figures

Figure 4-1. Model for Friend virus-induced erythroleukemia.

(A) Schematic depicting early and late stages of Friend virus-induced erythroleukemia. Infected

cells from the bone marrow migrate to the spleen and establish an infectious center. BMP4 and

Hh signaling promote the expansion of SEPs, which are FV targets and become infected. Early

SEPs which are infected form leukemia cells with activation of Spi1 whereas late stage SEPs

differentiate into Epo-independent BFU-Es.

120

Figure 4-2. Sf-Ron expression is upregulated during recovery from phenylhydrazine.

(A) Wild type mice were treated with phenylhydrazine (PHZ) and splenic mRNA was isolated

following treatment. Expression of Sf-Ron was measured by qPCR relative to 18S. Mean±SD.

Student t-test. n=3-9.

121

Figure 4-3. Sf-Ron plays a critical role in recovery from PHZ-induced anemia.

(A) FVB mice and Sf-Ron-/- mice were treated with PHZ and survival was assessed after

treatment from 0 to 10 days. n=10. (B) To determine if Sf-Ron-/- mice have defects or delays in

recovery from PHZ, blood was collected every other day in microcapillary tubes and spun to

determine hematocrit values after PHZ. Mean±SD. Student t-test. n=3-9.

122

Figure 4-4. Sf-Ron-/- mice have decreased production of stress BFU-Es after PHZ-induced

anemia.

(A) After treatment with PHZ, splenocytes were isolated from FVB and Sf-Ron-/- mice and

plated in methylcellulose media containing BMP4, Shh, SCF, GDF15, and Epo and cultured

under hypoxic conditions for 5 days. Stress BFU-Es were scored after staining with benzidine.

(B) Bone marrow cells were plated in methycellulose media containing Epo and IL3 and cultured

for 7 days at normoxia. BFU-Es were stained with benzidine and scored. Mean±SD. Student t-

test, n=3-5.

123

Figure 4-5. Schematic for populations of stress progenitors.

(A) Schematic shows the progression of surface markers as cells differentiate into more mature

erythroid progenitors. Early cells are all contained in Population I and are

Kit+Sca+CD71loTer119lo/neg. Immature SEPs enter the spleen and are Kit+Sca+CD34+CD133+ and

as they mature they lose expression of first CD34 and then CD133. (B) Schematic depicts the in

vitro culture system of erythroid progenitors with the expected combinations of cell surface

markers at each stage of the culture.

124

Figure 4-6 A-B. Sf-Ron-/- stress progenitors are more mature in the absence of Epo than wild

type progenitors.

(A) Unfractionated bone marrow cells were isolated from FVB and Sf-Ron-/- mice and cultured

in SEEM for 7 days at normoxia. Cells were analyzed by flow cytometry for Population I markers

(Kit, Sca, CD71, Ter119) after 7 days in SEEM. Cells are previously gated on Kit+Sca+ cells and

show representative plots of CD71 and Ter119 markers. (B) Graphical representation of

population frequencies shown in B. Mean±SD. Student t-test, n=3.

125

Figure 4-6 C-D. Sf-Ron-/- stress progenitors are more mature in the absence of Epo than wild

type progenitors.

(C) After 7 days in SEEM, FVB and Sf-Ron-/- bone marrow cultures were analyzed for

expression of immature SEP markers (Kit, Sca, CD34, CD133) by flow cytometry. Cells are

previously gated on Kit+Sca+ cells and show representative plots of CD34 and CD133 markers.

(D) Graphical representation of population frequencies shown in C. Mean±SD. Student t-test,

n=3-7.

126

Figure 4-6 E-F. Sf-Ron-/- stress progenitors are more mature in the absence of Epo than wild

type progenitors.

(E) Stress BFU-Es in FVB or Sf-Ron-/- cultures were measured after only 7 days culture in

SEEM. Cells were counted and plated in at a concentration of per 1x105/well in methocellulose

containing Shh, SCF, GDF15, BMP4, and Epo for 5 days in hypoxia. (F) Relative increases in

cell numbers are shown for FVB and Sf-Ron-/- cultures at day 7 (cell count D7/cell count D0).


127

Figure 4-6 G-H. Sf-Ron-/- stress progenitors are more mature in the absence of Epo than wild

type progenitors.

(G,H) Unfractionated BM from FVB or Sf-Ron-/- mice was labeled with PKH26 and cultured for

7 days in SEEM. PKH26 labeling was measured by flow cytometry. Bar graphs represent average

frequencies for either PKH26+ and PKH- populations (G) or subsets of PKH26+ populations (H).

Mean±SD. n=3-4.

128

Figure 4-6 I-J. Sf-Ron-/- stress progenitors are more mature in the absence of Epo than wild

type progenitors.

(I, J) Unfractionated BM from FVB or Sf-Ron-/- mice was labeled with PKH26 and cultured for

7 days in SEEM. PKH26 labeling was measured by flow cytometry. Representative images are

shown for either PKH26+ and PKH26- populations or subsets I-V of PKH26+ populations.

129

Figure 4-7 A-B. Sf-Ron-/- stress progenitors have no difference in response to Epo but are

less able to form BFU-Es.

(A) Unfractionated bone marrow was isolated from FVB and Sf-Ron-/- mice and cultured for 7

days in SEEM and then transitioned for 3 days to SEDM. Population I markers were assessed by

flow cytometry. Cells are previously gated on Kit+Sca+ cells and show representative plots of

CD71 and Ter119 markers. (B) Graphical representation of population frequencies shown in A.

Mean±SD. Student t-test, n=5.

130

Figure 4-7 C-D. Sf-Ron-/- stress progenitors have no difference in response to Epo but are

less able to form BFU-Es.

(C) After 7 days in SEEM and 3 days in SEDM, early SEP markers (Kit, Sca, CD34, CD133)

were measured by flow cytometry in FVB and Sf-Ron-/- cultures. Cells are previously gated on

Kit+Sca+ cells and show representative plots of CD34 and CD133 markers. (D) Graphical

representation of population frequencies shown in C. Mean±SD. Student t-test, n=4-6.

131

Figure 4-7 E. Sf-Ron-/- stress progenitors have no difference in response to Epo but are less

able to form BFU-Es.

(E) Stress BFU-Es were plated at a concentration of 1x105 cells/well from WT or Sf-Ron-/-

cultures after 7 days culture in SEEM followed by 3 days in SEDM. Mean±SD. Student t-test,

n=3-5.

132

Figure 4-8. Model of the effect of Sf-Ron on stress erythropoiesis.

(A) Schematic depicts model where loss of Sf-Ron diminishes the total erythroid output of stress

erythropoiesis. The proposed model based on this work suggests a model where early SEPs do

not expand as much in the absence of Sf-Ron and move prematurely into a more mature

differentiated state in the absence of Epo.

133

References

1. Finkelstein, L. D., Ney, P. a, Liu, Q.-P., Paulson, R. F. & Correll, P. H. Sf-Ron kinase

activity and the Grb2 binding site are required for Epo-independent growth of primary

erythroblasts infected with Friend virus. Oncogene 21, 3562–3570 (2002).

2. Subramanian, A. et al. Friend virus utilizes the BMP4-dependent stress erythropoiesis

pathway to induce erythroleukemia. J. Virol. 82, 382–393 (2008).

3. Hegde, S. et al. Stat3 promotes the development of erythroleukemia by inducing Pu.1

expression and inhibiting erythroid differentiation. Oncogene 28, 3349–3359 (2009).

4. Ney, P. a & D’Andrea, A. D. Friend erythroleukemia revisited. Blood 96, 3675–80 (2000).

5. Subramanian, A., Teal, H. E., Correll, P. H. & Paulson, R. F. Resistance to friend virus-

induced erythroleukemia in W/W(v) mice is caused by a spleen-specific defect which

results in a severe reduction in target cells and a lack of Sf-Ron expression. J. Virol. 79,

14586–14594 (2005).

6. Iwama, A., Okano, K., Sudo, T., Matsuda, Y. & Suda, T. Molecular cloning of a novel

receptor tyrosine kinase gene, RON, derived from enriched hematopoietic stem cells. 83,

3160–3169 (1994).

7. Persons, D. a et al. Fv2 encodes a truncated form of the Ron receptor tyrosine kinase. Nat.

Genet. 23, 159–165 (1999).

8. Nishigaki, K., Thompson, D., Hanson, C., Yugawa, T. & Ruscetti, S. The Envelope

Glycoprotein of Friend Spleen Focus-Forming Virus Covalently Interacts with and

Constitutively Activates a Truncated Form of the Receptor Tyrosine Kinase Ron. J. Virol.

75, 7893–7903 (2001).

9. Teal, H. E. et al. GRB2-mediated recruitment of GAB2, but not GAB1, to SF-RON

supports the expansion of Friend virus-infected erythroid progenitor cells. Oncogene 25,

134

2433–2443 (2006).

10. Ni, S., Zhao, C., Feng, G.-S., Paulson, R. F. & Correll, P. H. A novel Stat3 binding motif

in Gab2 mediates transformation of primary hematopoietic cells by the Ron/Ron receptor

tyrosine kinase in response to Friend virus infection. Mol. Cell. Biol. 27, 3708–3715

(2007).

11. Nerlov, C. & Graf, T. PU.1 induces myeloid lineage commitment in multipotent

hematopoietic progenitors. Genes Dev. 12, 2403–12 (1998).

12. Dakic, A. et al. PU.1 regulates the commitment of adult hematopoietic progenitors and

restricts granulopoiesis. J. Exp. Med. 201, 1487–1502 (2005).

13. Chou, S. T. et al. Graded repression of PU.1/Sfpi1 gene transcription by GATA factors

regulates hematopoietic cell fate. Blood 114, 983–94 (2009).

14. Kastner, P. & Chan, S. PU.1: A crucial and versatile player in hematopoiesis and

leukemia. Int. J. Biochem. Cell Biol. 40, 22–27 (2008).

15. Nutt, S. L., Metcalf, D., D’Amico, A., Polli, M. & Wu, L. Dynamic regulation of PU.1

expression in multipotent hematopoietic progenitors. J. Exp. Med. 201, 221–231 (2005).

16. Staber, P. B. et al. Sustained PU.1 Levels Balance Cell-Cycle Regulators to Prevent

Exhaustion of Adult Hematopoietic Stem Cells. Mol. Cell 49, 934–946 (2013).

17. Imperato, M. R., Cauchy, P., Obier, N. & Bonifer, C. The RUNX1-PU.1 axis in the

control of hematopoiesis. Int. J. Hematol. 101, 319–29 (2015).

18. Kim, H. et al. The ETS family transcription factor PU.1 is necessary for the maintenance

of fetal liver hematopoietic stem cells. Blood 104, 3894–3900 (2004).


progenitors identifies distinct progenitor populations and analogous human progenitors.

Blood 125, 1803–12 (2015).

20. Lenox, L. E., Perry, J. M. & Paulson, R. F. BMP4 and Madh5 regulate the erythroid

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response to acute anemia. 105, 2741–2748 (2005).

21. Perry, J. M., Harandi, O. F. & Paulson, R. F. BMP4 , SCF , and hypoxia cooperatively

regulate the expansion of murine stress erythroid progenitors. 109, 4494–4502 (2007).

22. Perry, J. M. et al. Maintenance of the BMP4-dependent stress erythropoiesis pathway in

the murine spleen requires hedgehog signaling. Blood 113, 911–8 (2009).

136

Chapter 5

Concluding Remarks and Future Directions

Conclusions

This work has identified a novel mechanism which activates the stress erythropoiesis

pathway in the absence of anemia and tissue hypoxia. We have shown that inflammatory signals

induce stress erythropoiesis through increased phagocytosis and heme-dependent signals (Figure

5-1 A). Work by others focused on the inhibition of bone marrow erythropoiesis by inflammatory

signals and the development of anemia of inflammation.1–5 We now show that this burst of stress

erythropoiesis is capable of delaying the onset of anemia temporarily, which in the case of acute

inflammation could be sufficient to maintain a normal hematocrit until the inflammation is

resolved. We have defined a new role for stress erythropoiesis, and further studies will be

required to fully understand the relationship between inflammation and SEPs.

We also show that SEPs have a unique response to the presence of certain pro-

inflammatory cytokines. Many studies have been done which show the inhibitory effects of TNF-

α on the proliferation and maturation of erythroid progenitors. Our data suggests that TNF-α and,

to some extent, IL-1β are capable of enhancing stress erythropoiesis and increase the number of

BFU-Es in vitro. TNF-α protein increases in F4/80+ cells after treatment with zymosan, and based

on our in vitro data, may work with typical stress erythropoiesis signals, such as GDF15 and

BMP4, to promote stress erythropoiesis during acute inflammation when there is no overt anemia

or tissue hypoxia.

This new understanding of stress erythropoiesis as a response to inflammatory inhibition

of steady state erythropoiesis could potentially provide new therapeutic targets in the treatment of

anemia of inflammation (AI). Current treatments for AI, such as continuous blood transfusions or

137

injections of Epo to stimulate blood cell production, are either ineffective or unsuitable for

treatment of chronic anemia associated with inflammation. Thus, new treatments are required

which are capable of enhancing erythropoiesis under inflammatory conditions. It is well

established that inflammation inhibits the erythroid progenitors of the bone marrow. However,

our work suggests that stress erythropoiesis may be activated in short bursts following the

establishment of inflammation, providing a wave of new erythrocytes to overcome the temporary

inhibition of erythropoiesis until homeostasis can be reestablished in the bone marrow. Further

studies should be carried out to further understand the relationship of stress erythroid progenitors

and inflammatory signals and to potentially identify ways to amplify an already existing response

to inflammation. Following a burst of stress erythropoiesis, SEPs in the spleen will become

exhausted and require several weeks before another stress erythropoiesis response can be

launched. If SEPs are activated in response to inflammation, attempting to amplify the magnitude

of the response could result in increased erythrocyte production and delayed onset of anemia in

patients.

In addition to the implications in treating AI, this mechanism could also provide insight

in hemoglobinopathies, such as sickle cell anemia, and other blood disorders, such as β-

thalassemia. Sickle cell anemia results from genetic mutations which cause abnormal

hemoglobin, leading to increased rigidity of erythrocytes and a characteristic sickle shape.

Patients with sickle cell anemia will develop anemia and, due to aggregates which impede blood

flow, tissue ischemia. It has also been noted that these sickled red cells are phagocytosed more

readily in the spleen. In beta thalassemia, which is a disease resulting from the low or absent

production of β-globin, splenomegaly and microcytotic anemia are observed. Splenomegaly is

attributed to the small misshapen erythrocytes becoming trapped and phagocytosed in the spleen.

Our findings suggest that in both these diseases there is accumulation of free heme in splenic

macrophages which would promote stress erythropoiesis. Thus, therapeutics useful in the

138

treatment of AI may also be suitable for treating other blood disorders where we observe

increased extramedullary erythropoiesis.

Our findings from Chapter 3 also indicate that production of glucocorticoids (GCs) is

essential in promoting the transition from an M1 macrophage to M2 macrophages. This is an

integral part in the shift from a potent inflammatory response to resolution of inflammation and

tissue remodeling. However, macrophages also play a critical role in erythropoiesis. Previous

work from our lab has shown that macrophages produce signals such as Wnts and prostaglandin

E2 (PGE2) which act on early stress erythroid progenitors (SEPs) and regulate their

differentiation as well as promoting the expansion of this population (Jie Xiang, unpublished

data). Additionally, the role of macrophages later is erythropoiesis has been well established.

Macrophages provide signals which are critical in maturation, enucleation and terminal

differentiation. Our findings here raise the intriguing question of what types of macrophages are

required during stress erythropoiesis and is the shift from M1 to M2 also denoting changes in the

needs of SEPs? Stress erythropoiesis involves drastic changes to splenic architecture to

accommodate the expansion of SEPs, but each of these SEPs also needs to be in contact with a

macrophage in order to undergo enucleation and terminal differentiation. This would suggest that

M2 macrophages, in addition to resolving inflammation, are also involved in remodeling the

spleen to create a microenvironment that is more conducive to stress erythropoiesis. It is possible

that SEPs have adapted to this transition from M1 to M2, with signals from M1 macrophages

(TNF-α and IL-1β) promoting stress erythropoiesis and then M2 macrophages remodeling the

spleen and serving as erythroblast islands for the maturing SEPs.

139

Future Studies.

Expanding our understanding of inflammation-induced stress erythropoiesis in mouse and

human systems.

In Chapter 2, we identified a mechanism by which zymosan, an inducer in a sterile model

of inflammation, activates stress erythropoiesis through TLR2 signaling. Activation of TLR2

leads to rapid changes in gene expression and drives the induction of stress erythropoiesis.

However, pathogens are recognized by the innate immune system and in a variety of ways. In this

work, we did not test the response of stress erythropoiesis to infection or agonists for other TLRs.

Moving forward with this work, it will be important to understand if this is a unique response to

zymosan, which is a potent TLR2 agonist, or if this is a commonly utilized mechanism to

stimulate extramedullary erythropoiesis in response to a wide variety of pathogens.

Future experiments should also include more studies with primary human cells. We

showed that TNF-α exerts similar effects on human bone marrow as we observe in mice,

increasing numbers of BFU-Es in in vitro culture. However, we should also focus on

understanding if erythrophagocytosis plays a role in driving changes in gene expression and

inducing stress erythropoiesis in human cells. Unfractionated human bone marrow cells can be

differentiated into bone marrow derived macrophages, allowing the system utilized for our

murine studies to be adapted to measure SIRPα surface expression, Spi-C and GDF15 mRNA

expression, and overall phagocytosis.

Identify the mechanism by which pro-inflammatory cytokines affect SEPs.

Our work in vitro showed that short exposures of TNF-α and IL-1β but not IFN-γ are

capable of increasing numbers of stress BFU-Es. However, it is unclear if these pro-inflammatory

140

cytokines are acting on SEPs directly or on the microenvironment. Previously, we pulsed

unfractionated marrow with TNF-α and both non-adherent SEPs and the adherent cells which

make up the microenvironment were treated. The in vitro culture system allows for

unfractionated marrow to be pulsed as before, but after removing TNF-α, fresh untreated SEPs

can be purified and added to the microenvironment. If TNF-α is acting on SEPs rather than the

microenvironment, we would expect there to be no difference in SEPs grown on untreated or

TNF-α treated microenvironment. TNFR1 and TNFR2 should also be measured in SEPs to

determine if the receptors are present. It is also possible that TNF-α changes the macrophages

and, in turn, exert changes on the SEPs in the culture. Macrophages from these cultures should be

characterized by gene expression and flow cytometry to determine if there are differences in

phenotype and function of those treated with TNF-α which affect the proliferation of SEPs.

Relationship of Epo and GDF15.

Our data demonstrates that in both f/f and GDF15-/- mice there is a decrease in serum

Epo after treatment with zymosan. We also observed that control mice transplanted with f/f

mutant BM have decreased serum Epo in response to zymosan even though the cells in the kidney

which produce Epo are wild type. This suggests that there is some level of crosstalk between

BMP4 signaling in SEPs in the spleen and Epo production occurring in the kidney. Previous

studies showed that oxygen-sensing by iron regulatory proteins (IRPs) regulate the translation of

HIF-2α and, thereby, controls expression of Epo.6 In situations where there is acute anemia, this

may be the most critical regulator of Epo production. However, in inflammation-induced stress

erythropoiesis where there is no anemia, it seems that BMP4 signaling is important in

upregulating Epo production. To further understand the regulation of Epo in response to

zymosan, we can perform BMTs of either GDF15-/- BM into WT mice (GDF15 KO-WT) and

141

allow mice to recover 10 weeks before treating with zymosan and measuring serum Epo. We

would expect GDF15 KO-WT to have decreased Epo production compared with WT animals in

response to zymosan. If GDF15 KO-WT mice also have defects in Epo production despite having

wild type microenvironment and kidneys, how are SEPs communicating with renal Epo-

producing cells? One possibility is that BMP4 is secreted by cells in the spleen and stimulates

production of Epo. We could begin to test this by measuring serum levels of BMP4 by ELISA

after treatment with zymosan in both WT and f/f mice. If there is increased BMP4 detectable in

the serum, we could attempt to induce Epo expression in HepG2 cells by treating with BMP4. A

subset of neural crest cells isolated from murine embryos are capable of producing Epo in vitro,

and we could attempt to inhibit Epo production by culturing cells with the BMP inhibitor

Noggin.7

Spi-C’s role in stress erythropoiesis during acute anemia vs inflammation.

We have shown that expression of Spi-C in red pulp macrophages is necessary for the

activation of stress erythropoiesis in response to inflammation. In the absence of Spi-C, there is a

significant reduction in the expression of GDF15 and this results in decreased numbers of stress

BFU-Es. However, other work has shown that during acute anemic stress Spi-C is dispensable

with Spi-C-/- mice having no measurable defect in their response to PHZ-induced anemia.8 In our

studies, GDF15 expression is not completely abolished in Spi-C-/- mice or BMDMs, suggesting

that there are other ways of inducing GDF15 in response to inflammation. Is Spi-C really

unimportant during acute anemic stress or are these other mechanisms of activating GDF15

enough to compensate for the loss of Spi-C? Is the role of Spi-C unique to responding to

inflammation? We showed that marcophages express high levels of Spi-C due to phagocytosis of

erythrocytes following treatment with zymosan. We would expect to find expression of Spi-C in

142

splenic macrophages during recovery from either bone marrow transplant or phenylhydrazine-

induced anemia. In addition to measuring expression of Spi-C in wild type macrophages, we

could deplete native macrophages in wild type recipients and adotively transfer either wild type

or Spi-C-/- monocytes before treating with PHZ. We could measure survival, hematocrit, and

stress BFU-Es during recovery as well as measuring GDF15 expression to see if monocytes

deficient in Spi-C diminish the expression of GDF15 during recovery.

143

Figures

Figure 5-1. Model of the activation of inflammation-induced stress erythropoiesis.

(A) Schematic depicts model stimulation of TLRs leads to MyD88-dependent reduction in

surface levels of SIRPα. This drives an increase in erythrophagocytosis and increases in

intracellular levels of heme, which then affects downstream heme-dependent targets. Specifically,

this results in decreases in protein levels of Bach1 and increased transcription of Spi-C, which

regulates the induction of GDF15 and subsequent activation of BMP4. Activation of BMP4 and

GDF15 are key components in inducing stress erythropoiesis and leading to expansion and

differentiation of SEPs.

144

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human erythropoiesis in vitro: role of p55 and p75 TNF receptors. Blood 85, 989–96

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3. Zamai, L. et al. TNF-related apoptosis-inducing ligand (TRAIL) as a negative regulator of

normal human erythropoiesis. Blood 95, 3716–3724 (2000).

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7. Suzuki, N., Hirano, I., Pan, X., Minegishi, N. & Yamamoto, M. Erythropoietin production

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145

CIRRICULUM VITA Laura Bennett

115 Henning Building, The Pennsylvania State University, University Park, PA 16802

[email protected]

EDUCATION

August 2010-August 2017, PhD in Genetics, Intercollege Graduate Degree Program in

Genetics, The Pennsylvania State University, University Park, PA [GPA=3.81]

August 2006-May 2010, B.S. in Biology, University of Mississippi, Oxford, MS

[GPA=3.84]

AWARDS AND HONORS

College of Agriculture Travel Award, Pennsylvania State University, 2015

Huck Endowment Travel Award, Pennsylvania State University, 2015

Awarded NIH T32 Fellowship for Animal Models of Inflammation, Penn State University, 2012

University Fellow, Pennsylvania State University, Fall 2010

Manga Cum Laude, University of Mississippi, 2010

RESEARCH EXPERIENCE December 2010-August 2017, PhD in Genetics, The Pennsylvania State University

Principal Investigator: Dr. Robert F. Paulson

Dissertation: Analysis of Stress Erythropoiesis and Inflammation

August 2008-May 2010, Sally McDonnell Barksdale Honors College Scholar, University of

Mississippi

Research Advisor: Dr. Bradley Jones

Thesis: Investigation of mutations affecting axon guidance at the midline in Drosophila

May 2008-August 2008, May 2009-August 2009, May 2010-August 2010, Student

Researcher, USDA-ARS, Stoneville, MS

Research Advisor: Dr. Mark Weaver

Project: Investigating the efficacy of Myrothecium verrucaria as a biocontrol agent for

Kudzu

PUBLICATIONS AND MANUSCRIPTS 1. Robert F. Paulson, Laura F Bennett, Jie Xiang. “Regeneration after injury – Activation of

stem cell stress response pathways to rapidly repair tissues.” Adult Stem Cells: Location,

Identity, and Potential. Ed. Dr. Kursad Turksen. [Published Jan. 2014]

2. Laura F Bennett, Chang Liao, Robert F Paulson. “Stress erythropoiesis model systems.”

Methods in Molecular Biology: Erythropoiesis-Methods and Protocols. Ed. Joyce A. Lloyd.

[In press. 2017]

3. LF Bennett, Robert Paulson. Zymosan-induced generalized inflammation activates stress

erythropoiesis through a novel mechanism. [Manuscript in prep.]

4. LF Bennett, Lei Shi, Pam Hankey-Giblin, Robert Paulson. Short-form Ron is required for

stress erythropoiesis. [Manuscript in prep.]

POSTER PRESENTATIONS 1. 2016, 45th Annual ISEH Meeting [San Diego, CA]. Poster: TLR stimulation increases

erythrophagocytosis and induces stress erythropoiesis.

2. 2015, 18th GRC on Red Cells [Holderness, NH]. Poster: Sf-STK regulates the differentiation

of stress progenitors during acute anemic stress.

3. 2014, 19th Hemoglobin Switching Conference [Oxford, UK]. Poster: TLR ligands affect

splenic macrophages and lead to activation of the stress erythropoiesis pathway

4. 2013, 17th GRC on Red Cells [Andover, NH]. Stress erythropoiesis plays a key role in the

initial response to inflammation.

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