Safer conditioning for -encoding transfer to induce

Md Anayet Hasan BSc (Honors), MSc in Biotechnology

https://orcid.org/0000-0003-3796-7031

A thesis submitted for the degree of Doctor of Philosophy at The University of Queensland in Year 2020 Faculty of Medicine

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Abstract Transplanting genetically-engineered hematopoietic stem cells (HSC) encoding antigen targeted to dendritic cells (DC) or other antigen presenting cells (APC) provides an efficient means to induce antigen-specific T-cell tolerance. Low-level, but stable HSC chimerism is sufficient for T-cell tolerance induction and prevention of autoimmune disease in animal models. A current limitation of this approach is, highly-toxic myeloablative and/or immunoablative pre-transfer recipient conditioning regimens are typically used to achieve high chimerism levels with the transferred engineered HSC. Whilst low doses of total body irradiation can be used in order to reduce toxicity, this is still genotoxic, leading to cell and DNA damage. Several alternatives to total body irradiation and other genotoxic (e.g. chemotherapy) approaches have been suggested. One alternative that has been reported is the use of a CD45-targeted immunotoxin (CD45-SAP) that has been shown to deplete >98% of HSC and facilitates 90% of donor chimerism when used at high doses in mice, but CD45- SAP is highly immunoablative at the explored high doses. To promote clinical applicability that might be achieved by avoiding immunodepletion and, as low-levels of stable donor chimerism are suitable to generate T-cell tolerance, I tested whether use of lower doses of CD45.2-SAP that might preserve would be suitable as a low-toxicity conditioning regimen. Titrating the dose of CD45.2- SAP led to a dose-dependent depletion of phenotypically-defined long-term repopulating HSC (LT- HSC) in bone marrow. Dose-dependent CD45.2-SAP- mediated LT-HSC depletion was verified by competitive bone marrow transplantation. In contrast to high dose CD45.2-SAP, a lower dose (0.5 mg/kg) preferentially depleted approximately 45-50% of LT-HSC whereas peripheral leukocytes were largely preserved (approximately 80% retained). A moderate level of hematopoietic mixed chimerism (10-12%) and tolerance induction was achieved after transplanting antigen-encoding bone marrow using this non-genotoxic conditioning. It is also established that tolerance induction can be improved by transplanting large number of gene-modified bone marrow cells. These results were carried forward and hematopoietic stem and progenitor cells (HSPC) specific immunotoxin (2B8- SAP) was tested whether it induce T-cell tolerance without affecting recipients peripheral leukocytes. It is demonstrated that low dose 2B8-SAP (0.5mg/kg) depleted >90% of LT-HSC without depleting peripheral leukocytes and permits approximately 25-30% of donor chimerism which subsequently induce T-cell tolerance. Thus non-genotoxic conditioning approach through targeted immunotoxins (low dose CD45.2-SAP/2B8-SAP) could provide a useful alternative to toxic conditioning approaches for tolerance induction with gene-engineered bone marrow.

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Declaration by author

This thesis is composed of my original work, and contains no material previously published or written by another person except where due reference has been made in the text. I have clearly stated the contribution by others to jointly-authored works that I have included in my thesis.

I have clearly stated the contribution of others to my thesis as a whole, including statistical assistance, survey design, data analysis, significant technical procedures, professional editorial advice, financial support and any other original research work used or reported in my thesis. The content of my thesis is the result of work I have carried out since the commencement of my higher degree by research candidature and does not include a substantial part of work that has been submitted to qualify for the award of any other degree or diploma in any university or other tertiary institution. I have clearly stated which parts of my thesis, if any, have been submitted to qualify for another award.

I acknowledge that an electronic copy of my thesis must be lodged with the University Library and, subject to the policy and procedures of The University of Queensland, the thesis be made available for research and study in accordance with the Copyright Act 1968 unless a period of embargo has been approved by the Dean of the Graduate School.

I acknowledge that copyright of all material contained in my thesis resides with the copyright holder(s) of that material. Where appropriate I have obtained copyright permission from the copyright holder to reproduce material in this thesis and have sought permission from co-authors for any jointly authored works included in the thesis.

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Publications included in this thesis

No publication included

Submitted manuscripts included in this thesis

No manuscript submitted for publication

Other publications during candidature

Research papers No other publication.

Oral*/Poster presentation ➢ Hasan MA, Pettit AR, Steptoe RJ (2019) “Transplanting antigen-encoding bone marrow under non-genotoxic conditioning facilitates hematopoietic chimerism and antigen-specific immune tolerance”- 48th Annual Scientific Meeting of The Australasian and New Zealand Society for , Adelaide, Australia.

➢ Hasan MA, Pettit AR, Steptoe RJ (2019) “Transplanting antigen-encoding bone marrow under non-genotoxic conditioning facilitates hematopoietic chimerism and antigen-specific immune tolerance”- British Society for Immunology Congress, Liverpool, United Kingdom.

➢ Hasan MA, Pettit AR, Steptoe RJ (2019) “Transplanting antigen-encoding bone marrow under non-genotoxic conditioning facilitates hematopoietic chimerism and antigen-specific immune tolerance”- Institute of Medical Sciences, The University of Tokyo (IMSUT), Tokyo, Japan. *

➢ Hasan MA, Pettit AR, Steptoe RJ (2019) “Non-genotoxic conditioning facilitates tolerance induction through stable mixed hematopoietic chimerism of gene modified bone marrow”- 17th International Congress of Immunology, Beijing, China.

➢ Hasan MA, Pettit AR, Steptoe RJ (2019) “Non-genotoxic conditioning facilitates tolerance induction through stable mixed hematopoietic chimerism of gene modified bone marrow”- Princess Alexandra Hospital Health Symposium, Brisbane, Australia.

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➢ Hasan MA, Pettit AR, Steptoe RJ (2019) “Non-genotoxic conditioning facilitates tolerance induction through stable mixed hematopoietic chimerism of gene modified bone marrow”- Translational Research Institute Research Symposium, Brisbane, Australia.

➢ Hasan MA, Pettit AR, Steptoe RJ (2018) “Non-genotoxic conditioning facilitates stable mixed hematopoietic chimerism of gene modified bone marrow”- 47th Annual Scientific Meeting of The Australasian Society for Immunology, Perth, Australia.

➢ Hasan MA, Pettit AR, Steptoe RJ (2018) “Non-genotoxic conditioning facilitates stable mixed hematopoietic chimerism of gene modified bone marrow”- 18th Annual Brisbane Immunology Group Meeting, Gold Coast, Australia.

➢ Hasan MA, Pettit AR, Steptoe RJ (2018) “Non-genotoxic conditioning facilitates stable mixed hematopoietic chimerism of gene modified bone marrow”- Princess Alexandra Hospital Health Symposium, Brisbane, Australia.

➢ Hasan MA, Pettit AR, Steptoe RJ (2018) “Non-genotoxic conditioning facilitates stable mixed hematopoietic chimerism of gene modified bone marrow”- Translational Research Institute Research Symposium, Brisbane, Australia.

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Contributions by others to the thesis

Associate Professor Raymond Steptoe contributed to the design and analysis of all experiments as well as the written portions of this thesis. Professor Allison Pettit contributed to the design of some experiments and some written portion of this thesis.

In chapter 4 and 5, James Barber performed mice immunization (Figure 4.17 Figure 4.21 and Figure 5.11).

Statement of parts of the thesis submitted to qualify for the award of another degree

No works submitted towards another degree have been included in this thesis.

Research involving human or animal subjects

Experiments included in this thesis were approved in the following projects: TRI/UQDI/296/14/NHMRC TRI/UQDI/371/17/NHMRC

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Acknowledgments

First of all I wish to express my heartiest gratitude to my God for giving me the strength to carry out this work with enthusiasm and for being with me every steps of the way. My regards, gratitude, indebtedness and appreciation goes to my respected supervisor Associate Professor Raymond Steptoe for his supervision, constructive criticism, expert guidance, enthusiastic encouragement to pursue new ideas and a good sense of humour and never ending inspiration throughout the entire period of my research work. Without you, this thesis would have been in no way achievable. I know words are not enough to show how thankful I am. I am so lucky to have you as my supervisor. Thank you Ray. I would like to thank my co-supervisor Professor Allison Pettit for her supervision, support, invaluable advice and constructive criticism during my candidature. My thesis committee members, Associate Professor Graham Leggatt, Associate Professor Emma Hamilton-Williams and Dr Roberta Mazzieri, I thank you for your time and valuable feedback during my PhD milestones. I would like to extend my deepest gratitude to all the present and past members of the and Tolerance Laboratory, and very special thanks to Jeremy Brooks, Dino Potato, Peter Murphy, James Barber and Tommy Liu for constructive suggestions, constant help and encouragement during my candidature.

My sincere gratitude to Irina Buckle and Arifur Rahman for your continual help and encouragement. I am thankful to TRI flow cytometry, histology and microscopy facility. Enormous thanks to TRI BRF staffs specially Karen Knox, Christa Sing and Jason Zahra who made my animal experiment so easier.

I would like to express utmost gratitude to my beloved parents and family members for their constant support, encouragement and unlimited help and also grateful to all of my friends and well- wishers for their encouragement, inspiration, good contribution and good feelings to me during the course of the research.

Finally, to my lovely wife Jui who has been sacrificed a lot during my candidature. Thank you so much for your endless love, continual support and understanding. I am blessed to have you in my life.

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Financial support

This research was supported by an Australian Government Research Training Program (RTP) Scholarship formerly known as International Postgraduate Research Scholarship (IPRS) and Australian Postgraduate Award (APA).

Keywords

Autoimmune disease, tolerance, bone marrow transplantation, hematopoietic stem cell, T-cells, immunotoxin

Australian and New Zealand Standard Research Classifications (ANZSRC)

ANZSRC code: 0110708, Transplant immunology, 50% ANZSRC code: 0110704, Cellular immunology, 30% ANZSRC code: 0110703, Autoimmunity, 20%

Fields of Research (FoR) Classification

FoR code: 1107, Immunology, 100%

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Table of Contents

Chapter 1 Literature Review 01 1.1. Autoimmune disease 02 1.2. Mechanism of immune tolerance and its failure 03 1.2.1. Central T-cell tolerance 03 1.2.2. Peripheral T-cell tolerance 04 1.2.2.1. Clonal ignorance 04 1.2.2.2. Peripheral deletion 04 1.2.2.3. 05 1.2.2.4. Regulation 06 1.2.3. Memory T-cell (Tmem) tolerance 06 1.2.3.1. Memory T-cell tolerance in autoimmune disease 07 1.2.4. Immunological tolerance failure triggers autoimmune diseases 08 1.2.4.1. Genetic influences on tolerance breakdown 09 1.2.4.2. Evidence for environmental influences in tolerance failure 10 1.3. Therapeutic approaches for autoimmune diseases 11 1.3.1. Current therapeutic strategies for tolerance induction in autoimmune disease 12 1.3.2. HSCT and HSC mediated gene therapy for autoimmune disease 12 1.3.3. Tolerance induction by autologous HSCT 14 1.3.4. Tolerance induction by allogenic HSCT 15 1.3.5. Hematopoietic chimerism contributes to tolerance induction in autoimmune disease 16 1.3.6. Tolerance induction through gene-modified HSC 18 1.3.7. DC-induced induction 18 1.4. HSC niche and Conditioning regimes 19 1.4.1. HSC niche: Home for hematopoietic stem cells 19 1.4.2. Current conditioning regimes used clinically 20 1.4.3. Specific conditioning regimes 21 1.5. Modulation of the by immunosuppressive conditioning in auto-HSCT 22 1.6. Hypothesis and aims 24

Chapter 2 Materials and Methods 26 2.1. Mice 27 2.1.1. Congenic mice strains 27 2.1.2. Transgenic mice strains 27 ix | P a g e

2.2. Reagents 28 2.2.1. Proteins 28 2.2.2. Peptides 29 2.2.3. Chemicals 29 2.2.4. Buffers 30 2.2.5. Adjuvants 32 2.2.6. Tissue culture media 32 2.2.7. Electrophoresis gels 32 2.2.8. List of monoclonal (mAb) 33 2.2.9. Staining panel 34 2.2.10. Immunotoxin Preparation 35 2.3. Animal procedures 36 2.3.1. Immunization 36 2.3.2. Blood collection 36 2.3.3. Lymph nodes and spleen 37 2.3.4. Bone marrow transplantation 37 2.4. Flow cytometry 38 2.4.1. Surface staining 38 2.4.2. Pulse-chase assay 38 2.5. ELISPOT 39 2.6. Histology 39 2.6.1. Bone tissue collection and fixation 39 2.6.2. Bone decalcification 40 2.6.3. Paraffin processing and embedding 40 2.6.4. Tissue sectioning and staining 40 2.7. Statistical analysis 40

Chapter 3 A leukocyte-targeting immunotoxin preferentially depletes HSC when used at low doses 41 3.1. Introduction 42 3.2. Results 44 3.2.1. CD45.2-SAP depletes long term repopulating HSC (LT-HSC) in a dose dependent 44 manner 3.2.2. Low-dose (0.5mg/kg) CD45.2-SAP depletes LT-HSC 2 days after administration 55 3.2.3. Mechanisms of CD45.2-SAP targeting LT-HSC 66 x | P a g e

3.2.4. Competitive BM transplantation verifies CD45.2-SAP mediated LT-HSC depletion 73 3.3. Discussion 84 3.4. Summary 86

Chapter 4 Non-genotoxic conditioning using leukocyte-targeting immunotoxin facilitates tolerance induction 87 4.1. Introduction 88 4.2. Results 90 4.2.1. Low-dose CD45.2-SAP enables tolerance induction by facilitating stable mixed hematopoietic chimerism of gene-modified BM 90 4.2.2. Increasing the BM cell dose accelerates donor chimerism and subsequently induce 100 tolerance 4.3. Discussion 112 4.4. Summary and Future direction 115

Chapter 5 Non-genotoxic conditioning using HSPC-specific immunotoxin facilitates tolerance induction 116 5.1. Introduction 117 5.2. Results 118 5.2.1. 2B8-SAP potently depletes endogenous long-term repopulating LT-HSC from mouse bone marrow 118 5.2.2. 2B8-SAP facilitates donor chimerism which subsequently induces antigen-specific T-cell tolerance 129 5.3. Discussion 138 5.4. Summary 140

Chapter 6 Concluding Remarks 141

Chapter 7 References 147

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List of Figures and Tables

Figure 1.1 Autoimmune disease is assumed to result from a combined effects of genetic and environmental factors 11 Figure 1.2 Steps involved in gene therapy 14 Figure 1.3 Mechanism of action of saporin 22 Figure 3.1 CD45.2-SAP depletes haematopoietic stem cells and multipotent progenitor cells 47 Figure 3.2 Low-dose CD45.2-SAP (0.5 mg/kg) has limited depletion effect on common lymphoid progenitors and myeloid progenitors 49 Figure 3.3 Low-dose CD45.2-SAP (0.5 mg/kg) does not deplete lineage-committed hematopoietic progenitors 51 Figure 3.4 Low-dose CD45.2-SAP (0.5 mg/kg) has limited depletion effect on T and B cells but myeloid cells and dendritic cells increase after CD45.2-SAP treatment 53 Figure 3.5 Low-dose (0.5mg/kg) CD45.2-SAP partially depletes HSC and HSPC at day 2 57 Figure 3.6 Low-dose CD45.2-SAP (0.5 mg/kg) depletes a significant proportion of lineage- committed progenitors 2 days after administration 59 Figure 3.7 Low-dose CD45.2-SAP (0.5 mg/kg) depletes a proportion of CD8+ and CD4+ T- cells but not B cells and myeloid cells 61 Figure 3.8 number increases after low-dose CD45.2-SAP (0.5 mg/kg) administration 63 Figure 3.9 Immunohistochemistry confirms CD45.2-SAP (0.5 mg/kg) induce macrophage intensity 64 Figure 3.10 Leukocytes express more CD45 than LT-HSC 68 Figure 3.11 Binding of anti-CD45 clone blocks the binding of the same clone but does not block a different clone 69 Figure 3.12 CD45 internalizes in LT-HSC and B cells 71 Figure 3.13 CD45.2-SAP treated BM engrafts in dose dependent manner under non- myeloablative conditioning 76 Figure 3.14 Donor chimerism depends on transplanted donor LT-HSC and HSPC numbers 78 Figure 3.15 Engraftment of CD45.2-SAP treated BM under non myeloablative condition 80 Figure 3.16 Donor derived cell concentrations increased over the time 82 Figure 3.17 Transplanted donor progenitor cells contribute to early mixed chimerism 83 Figure 4.1 CD45.2-SAP allows rapid immune recovery 93 Figure 4.2 Low dose CD45.2-SAP permits donor chimerism 95 xii | P a g e

Figure 4.3 CD45.2-SAP allows chimerism of donor BM derived lineages 96 Figure 4.4 CD45.2-SAP facilitates donor HSPC chimerism 98 Figure 4.5 CD45.2-SAP facilitates antigen specific T-cell tolerance 99 Figure 4.6 Increasing the BM cell dose permits rapid cell recovery 103 Figure 4.7 Higher BM dose accelerates donor chimerism 105 Figure 4.8 Higher BM dose accelerates donor BM derived lineages chimerism 107 Figure 4.9 Higher BM dose permits higher and faster donor chimerism than standard dose 109 Figure 4.10 Higher BM cell dose facilitates donor HSPC chimerism 110 Figure 4.11 Higher BM cell dose induce T-cell tolerance in 17 weeks 111 Figure 5.1 Single i.v. administration of 2B8-SAP (0.5mg/kg) potently depletes long-term repopulating hematopoietic stem cells (LT-HSC) and hematopoietic stem and progenitor cells (HSPC) 120 Figure 5.2 Low-dose 2B8-SAP (0.5mg/kg) does not change CLP and MP numbers 122 Figure 5.3 Low-dose 2B8-SAP (0.5mg/kg) does not deplete lineage-committed hematopoietic progenitors except GMP 123 Figure 5.4 Low-dose 2B8-SAP (0.5mg/kg) preserves peripheral leukocytes 125 Figure 5.5 and number increases after 2B8-SAP treatment but macrophage number remain unchanged 127 Figure 5.6 2B8-SAP (0.5mg/kg) reduces the total leukocyte concentration in blood 128 Figure 5.7 Low-dose 2B8-SAP allows quick total cell recovery 131 Figure 5.8 2B8-SAP facilitates significant donor chimerism 133 Figure 5.9 2B8-SAP facilitates donor-derived leukocyte lineage chimerism 134 Figure 5.10 2B8-SAP allows donor HSPC chimerism 136 Figure 5.11 2B8-SAP mediates significant donor chimerism and subsequently induce T-cell tolerance 137

Table 2.1 List of chemicals 29 Table 2.2 List of buffers 30 Table 2.3 List of mAbs used in flow cytometry 33 Table 2.4 List of secondaries used in flow cytometry 34 Table 2.5 List of used in ELISPOT 39

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List of Key abbreviations used in the thesis

AD Autoimmune disease APC Antigen presenting cell BM Bone marrow BMT Bone marrow transplantation CLP Common lymphoid progenitor DC FACS Fluorescence-activated cell sorting FOXP3 Forkhead box P3 G-CSF Granulocyte colony stimulating factor GMP Granulocyte myeloid progenitor GVHD Graft versus host disease HLA HSC Hematopoietic stem cell HSCT Hematopoietic stem cell transplantation HSPC Hematopoietic stem and progenitor cell IFN-γ Interferon-γ IL Interleukin ISO LT-HSC Long-term repopulating hematopoietic stem cell mAb MEP Myeloid erythroid progenitor MHC Major Histocompatibility complex MP Myeloid progenitor MPP Multipotent progenitor MS Multiple sclerosis MT-PBS Mouse tonicity phosphate buffered saline NSAID Non-steroidal anti-inflammatory drugs OVA Ovalbumin PCR Polymerase chain reaction RA Rheumatoid arthritis SA Streptavidin SAP Saporin xiv | P a g e

T1D Type 1 diabetes TBI Total body irradiation TCR T-cell receptor Tg Transgenic TNF Tumor necrosis factor TRI-BRF Translational Research Institute-Biological Resource Facility

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Chapter 1 Literature Review

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1.1. Autoimmune diseases Autoimmune disease occurs due to the breakdown of self-tolerance mechanism and subsequent induction of aberrant immune responses against self- either in the form of auto- antibodies or autoreactive T-cell responses [1-3]. To date more than 80 different autoimmune diseases have been reported, ranging from mild to severe, depending on which body system is under attack and the degree of attack. Some autoimmune diseases such as type 1 diabetes (T1D) [4], multiple sclerosis (MS) [5] and autoimmune thyroiditis [6] affect mainly one organ (i.e. organ-specific), whilst others such as rheumatoid arthritis (RA) [7], systemic lupus erythematosus (SLE) [8] and systemic vasculitis (SV) [9] have more widespread effects (i.e. systemic). Several mechanisms behind autoimmune diseases have been proposed but the actual causes are still unknown [10]. Genetic predispositions, viral or bacterial infections and some drugs play a crucial role in promoting autoimmune diseases [11, 12]. In addition, chemical and environmental irritants such as tartrazine, hydrazine, trichloroethylene, mercury, silica. can also trigger autoimmune diseases [13-15].

Typically, autoimmune diseases are diagnosed through blood tests (, and organ function), x-rays and sometimes biopsy of affected tissues [16, 17]. Currently autoimmune disease are not completely curable but can be manageable. There are a wide range of treatment options aimed at limiting self-directed immune responses and inflammation. These include non-steroidal anti-inflammatory drugs (NSAID) [18], corticosteroid anti-inflammatories [19] and immunosuppressive agents [20]. Clinically, their application depends on type and stage of autoimmune disease [21]. Some other currently available treatment options are, anti- therapy, therapeutic monoclonal antibodies such as TNF inhibitors [22], and immunoglobulin replacement therapy [23]. The main aim for these treatments is to alleviate symptoms, minimise tissue damage and retain organ function. These non-specific immunosuppressives have limited efficacy, high toxicity and life-threatening side effects, hence more specific therapeutic approaches are required to lower the risk of autoimmune disease and improve tolerability [24]. Tolerance induction through haematopoietic stem cell transplantation (HSCT) could be an effective approach to treat autoimmune diseases. Currently, HSCT is being tested as a treatment option for severe autoimmune disease, such as therapy-resistant MS [25] and RA [26]. Studies using animal models have shown that transfer of HSC can reverse autoimmunity, and numerous mechanistic pathways may explain this phenomenon [27, 28].

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1.2. Mechanism of immune tolerance and its failure

Immunological tolerance is the process by which an individual’s immune system is controlled in order to avoid immune responses to self-antigen or innocuous environmental antigens. One feature of immunological tolerance is that the capability of mounting an immune response against foreign antigens is retained. The mechanisms of immune tolerance are generally divided into two broad categories, central and peripheral tolerance.

1.2.1. Central T-cell tolerance

Central tolerance is a term used to describe the process that occurs during development where self-reactive are stripped from the repertoire. For B cells this occurs in the bone marrow and for T-cells, the . Recombination of the genes encoding the T-cell receptor can theoretically generate up to 1020 different T-cell receptor (TCR), although the actual number of specificities in human has been estimated to be in the order of 2.5 x 106 [29, 30]. While the potential to recognise so many foreign antigens is useful, the possibility that T-cell recognise or cross react with determinants from self-molecules is raised. To avoid unwanted autoimmunity, self-reactive T-cells undergo through [31], become unresponsive through a process known as clonal anergy [32] or are subjected to a secondary TCR gene rearrangement, well-known as receptor editing [33]. Although clonal anergy and receptor editing are believed to have lesser roles, clonal deletion is the key process of keeping the auto-reactive cells in check during T-cell development and thus preventing autoimmunity [34].

As T-cells develop through the CD4+CD8+ double-positive phase in the thymic inner cortex they begin to express TCR. Ligation of ‘productive’ TCRs that bind MHC class I or II leads to transduction of a survival signal and the developing T-cell is ‘positively selected’. Positively- selected mature either into CD8+ or CD4+ single-positive T-cells through the interaction of a peptide–MHC class I or peptide-MHC class II, respectively. [35].

T-cells emerging from positive selection migrate towards the thymic medulla and at the thymic cortical-medullary junction encounter dendritic cells (DC), and other bone marrow derived antigen presenting cells with high expression of self-peptide-MHC complex. T-cells binding with MHC-self-peptide with a higher affinity leads to apoptosis.

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1.2.2. Peripheral T-cell tolerance

Despite being an efficient system, central tolerance is sometimes leaky and is not a foolproof system to prevent the egress of autoreactive T-cells from the thymus because not all autoreactive T-cells are deleted by central tolerance mechanism nor all autoantigens are expressed in the thymus. Therefore the body has an additional mechanism to control these potentially autoreactive T-cells. This is accomplished by peripheral tolerance mechanisms [36]. Since T-cells first encounter different antigens such as autoantigens, food antigens, and microbiota out of the thymus, it is extremely important to have tolerance against these. Four key peripheral tolerance mechanisms have been widely defined, clonal ignorance, peripheral deletion, anergy, and regulation.

1.2.2.1. Clonal ignorance

In some circumstances autoreactive T-cells may not encounter their cognate antigen because it is located in inaccessible tissues [37] and eventually die out of ignorance. This phenomenon is called clonal ignorance. The importance of clonal ignorance was first demonstrated in a double transgenic mouse where T-cells expressed a specific TCR for a viral peptide and were not tolerant in vivo because they ‘ignored’ their target cells. A key mechanisms is that the target antigen is expressed at a level too low to induce the activation of T-cells [38]. Additional mechanisms include, the potential antigens are separated from the T-cells by an immunological barrier, or the lack of costimulatory molecules to promote T-cells activation [39]. Thus, autoreactive T-cells are ignored or kept aside from potential autoantigens to avoid peripheral damage.

1.2.2.2. Peripheral deletion

Autoreactive T-cell deletion or apoptotic cell death is one of the vital mechanisms to achieve central tolerance in the thymus as well as tolerance in the peripheral tissues. Peripheral deletion of autoreactive T-cells is achieved by apoptosis directed by Fas (CD95) and lymphoma- 2 (Bcl-2) interacting mediator (Bim). Although these Fas and Bim mediated pathways are distinct in their mode of action, a combined effect of the pathways ensure the autoreactive T- cells undergo apoptosis in the periphery. in conjunction with activation of death receptors can also result in T-cell deletion in a process commonly referred to as activation induced cell death (AICD). T-cells express Fas (CD95), a death receptor protein which can be activated by the FasL (CD178). Antigenic stimulation or cognate stimulation of

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T-cells activates a Fas mediated cascade of downstream pathways to trigger an intracellular “death-inducing signalling complex” (DISC); which successively, initiates Caspase-8 directed apoptosis of the cells [40]. Besides the Fas/FasL pathway the cytokine TNF-α has been shown to play a role in controlling AICD. Signalling by TNF-α activates TNFR1 which can then induce apoptosis by either a caspase-dependent or intrinsic pathway [41].

It is well established that Bcl-2 family members regulate the intrinsic apoptotic pathway through either inhibiting (anti-apoptotic) or inducing (pro-apoptotic) apoptosis. Bcl-2 family proteins are classified into 3 groups, anti-apoptotic factors (Bcl-2, Bcl-X, Bcl-W, Mcl-1, Bcl- B), pro-apoptotic activators (BIM, BID) and pro-apoptotic effectors (BAD, NOXA, PUMA). The balance between anti-apoptotic and pro-apoptotic factors control cell survival or death. Bim has a paradoxical role as it can induce T-cell activation via IP3R/calcium/NFAT pathway and can simultaneously cause apoptotic death of autoreactive T-cells in the periphery [42, 43]. In addition, Bim can activate the Bax/Bak pathway to directly permeabilize outer mitochondrial membrane to induce autoreactive T-cells apoptosis [44]. Defective apoptosis can trigger autoimmune disease like T1D and leads to aberrant peripheral tolerance. DC have important role in antigen presentation and must be tightly controlled by intrinsic apoptotic pathway. Mice containing Bim deficient DC are unable to induce effective apoptosis and are susceptible to autoimmune diseases [45].

1.2.2.3. Clonal anergy

IL-2 expression as a result of costimulation is one of the crucial requirements for T-cell activation. Furthermore signalling through the IL-2 receptor complex can eventually activate the PI3K/AKT/mTOR pathway for the effector function of T-cells. However, long term functional unresponsiveness can be observed when T-cells are activated in the absence of appropriate co-stimulation and the phenomenon is termed as ‘anergy’[46]. Anergic T-cells are characterized by lack of proliferation and IL-2 production following activation. Research on mTOR pathway inhibition using rapamycin led the scope to reveal the association of the ATP activation dependent AMPK pathway and the involvement of Tregs in the induction of anergy in T-cells [47]. A second signal involving the interaction of program death-1 (PD-1) with its ligands, PDL-1 and PDL-2 plays a significant role in limiting the effector functions of T-cells through inhibition of autoreactive T-cell activation and expansion in the periphery [48, 49]. Another costimulatory molecule CTLA-4 also has an exquisite role in mediating inhibition through the engagement with B7 and subsequently transducing a negative signal for arresting

5 | P a g e the cell cycle. In several studies, mice lacking CTLA-4 and PD-1 has been shown to develop spontaneous autoimmunity [50, 51].

1.2.2.4. Regulation

A distinct subset of CD4+ T-cells with regulatory function known as regulatory T-cells (Treg), first identified as suppressor cells, are considered as the fundamental mediator in suppressing pathogenic effector function of auto-reactive T-cells in the periphery. In the last few decades, they have been studied extensively in the field of tolerance and autoimmunity. Tregs suppress the effector function of Th cells and keep aberrant T-cells in check, both in homeostasis and disease. However, Treg cells encompasses a heterogeneous family of cells. The first group of Treg cells which exert their suppressive effect through the release of transforming growth factor-β (TGF-β) are also known as Th3 cells [52]. Another group of Tregs which co-express CD25 and CD4 and transcription factor forkhead box p3 (Foxp3) is classically known as Tregs cell [53]. Currently these Tregs are divided into two broad subpopulations, one is known as ‘natural’ Tregs (nTregs), which develop in the thymus and the other is induced Tregs (iTregs), that may acquire regulatory properties under certain conditions in the periphery through differentiation from conventional CD4 T-cells [54]. Tregs can utilize multiple suppressive mechanisms to establish normal homeostasis and inhibit self-reactive T-cells in the peripheral organs [55]. Another type of regulatory CD4 T-cells, named type 1 regulatory T-cells (Tr1 cells), typically developed in the peripheral organs and do not require Foxp3 for their function. Although Tr1 cells can exert effects similar to Tregs, evidence suggests that expression of high levels of the immunoregulatory cytokine, IL-10 plays crucial role in suppression of self- reactive T-cells in the periphery [56].

The presence of peripheral autoreactive T-cells in healthy individuals raises the question of whether there are other existing mechanisms to regulate T-cell tolerance in homeostasis and how these regulatory mechanisms are compromised in the context of autoimmune diseases. While the mechanisms of tolerance described above are largely effective in limiting the development of autoimmune disease, alterations in either central or peripheral tolerance can overwhelm the normally effective Treg-mediated mechanisms [57].

1.2.3. Memory T-cell (Tmem) tolerance

A fundamental characteristic of the immune system, is a faster and stronger secondary response to a pathogen or antigen. After activation, antigen-specific naïve T-cells proliferate and

6 | P a g e differentiate into effector T-cells. Many effector cells migrate to an infection site to exert effector function and clear invading microbes. As the microbial threat is eliminated, the population of effector T-cells contracts and undergoes apoptosis but a small percentage (approximately 5-10%) of cells persist as Tmem. Two theories have been proposed to explain the generation of memory T-cells, “stochastic selection” [58, 59] or survival of “straggler” effector T-cells activated late in the immune response [60]. These Tmem cells have significant role in adaptive immunity and can rapidly produce effector T-cells upon antigen re-encounter with reduced dependence on co-stimulatory signal compared to naïve T-cells [61]. It is not known precisely what determines whether effector T-cells undergo apoptosis or survive to become memory cells.

There are two main subsets of Tmem cells, effector memory T-cells (TEM) and central memory

T-cells (TCM) which differ mainly by their homing capabilities. TCM express CD62L (L- selection), C-C chemokine receptor type 7 (CCR7), CD45RO (in human) and CD44

(intermediate to high). The TCM subpopulation recirculates through lymph nodes and lymphoid tissues. TEM, on the other hand do not express CD62L and CCR7 but express CD45RO and CD44 (intermediate to high) and recirculate through non-lymphoid tissues and the blood [62].

Compared to naïve T-cells, TCM are less dependent on costimulatory signals and sensitive to lower antigen doses which allows stronger and faster reactivation [63, 64]. TCM requires more time than TEM for reactivation and proliferation, TCM have a superior capability for proliferation and IL-2 expression compared to TEM. TCM’s capability to produce large numbers of effector T-cells makes them crucial in autoimmune disease where continuous reactivation of antigen- specific T-cells occurs [62].

1.2.3.1. Memory T-cell tolerance in autoimmune disease

Approaches to treating autoimmune disease must address the role of Tmem, either by regulating or inducing tolerance in the population of pathogenic auto-reactive T-cells responsible for tissue damage. Autoreactive Tmem populations can be rapidly recruited into effector populations upon antigen exposure in autoimmune diseases, including T1D [65]. Therefore, treatment effectiveness for Tmem is extremely important [65, 66]. Tregs, effectively control naive T-cell activation but can be unaffected by Treg suppression [67]. Similarly, immunosuppressants that target T-cells via the IL-2 signalling pathway are less effective on Tmem compared to naïve T-cells [68]. Tmem have previously been categorized as resistant to

7 | P a g e peripheral tolerance induction and consequently a hurdle in treatment of autoimmune disease [69].

In the steady state, immature and semi-mature DC play a crucial role in the induction of immune tolerance. Steady-state immature DC are continually processing antigens, including autoantigens and migrate to lymph nodes to interact with naïve T-cells. Immature DC express less co-stimulatory molecules and therefore T-cells specific for any presented autoantigens will be incompletely activated and undergo deletion or be induced into anergy [70]. It was well established that steady state DC can induce peripheral tolerance in naïve T-cells but it was thought that memory T-cells may be resistant to peripheral tolerance induction by the same mechanisms [71, 72]. However in a study in the host lab it was found that steady-state DC expressing cognate antigen terminated the response of memory CD8+ T-cells. It was demonstrated that Tmem were ‘tolerized’ by means of deletion and inactivation [73-75]. Targeting antigen expression to the most appropriate cell lineage is crucial to ensure tolerance induction and limit possible adverse effects. The host lab has performed a series of studies testing induction of tolerance to ovalbumin (OVA)in antigen specific (OT-I) Tmem by DCs or a range of APCs expressing OVA under the control of CD11c or MHC class II promoter, respectively [74]. When OVA expression was targeted to DCs, an initial expansion of OT-I Tmem was observed before unresponsiveness occurred. Such expansion could result in harmful tissue damage and potentially destroy the tissue intended for protection. In contrast, OVA expression targeted to all MHC class II+ APCs using the MHC class II promoter resulted in rapid inactivation of OT-I Tmem with little expansion or effector function [74, 76].

1.2.4. Immunological tolerance failure triggers autoimmune diseases

As discussed above, induction of immune tolerance is crucial for protection from unwanted, detrimental responses to self-antigens. When this fails, autoimmune diseases develop as a result of the aberrant immune responses against self-antigens that arise in the form of auto-antibodies or T-cell responses. Several mechanisms can contribute to failure of self-tolerance, including greater activation or decreased depletion of autoreactive CD4+ Th cells, imperfect modulation of the immune system by CD8+ suppressor T-cells and CD4+ Treg and, molecular mimicry between foreign particles and self-antigens, dysregulated signalling (upregulated pro- inflammatory ). Although how autoimmune disease is developed is not completely understood, autoimmunity is assumed to result from a combined effects of genetic and

8 | P a g e environmental factors such as pathogen-induced infections, smoking, diet and stochastic events (Figure 1.1) [77].

1.2.4.1. Genetic influences on tolerance breakdown

It has been identified in animal models and human diseases that many different gene polymorphism can confer the susceptibility to or the resistance from autoimmune disorders. This provides key information into important pathways of disease development. Immunogenetic loci encode MHC class I and class II, immunoglobulins (antibody), complement proteins, chemokines, cytokines, TCR genes and TAP genes. In humans, both class I and class II MHC genes are extremely polymorphic and located on chromosome 6 [78, 79]. Class III MHC molecules are involved in inflammatory responses, antigen recognition and macrophage activation (e.g. tumor necrosis factor) [80]. On the other hand MHC does not encode T and B cell stimulating cytokines such as interleukin (IL-2), interleukin (IL-12), interleukin (IL-16) and interferons [81, 82]. MHC gene provide a major genetic contribution to type 1 diabetes (T1D), accounting for almost 50% of total genetic influence. HLA-DQ variation acts as key disease influencing locus for T1D. DRB1*03–DQA1*0501–DQB1*0201 and DRB1*04–DQA1*0301–DQB1*0302 haplotypes play major roles in T1D in European populations whereas T1D in Japanese and Korean populations is associated with HLA-DR9 (DRB1*0901–DQB1*0303) and HLA-DR4 (DRB1*0405–DQB1*0401) [83]. Genetic differences in immunoglobulin assembly (antibody allotypes) and TCR could also influence immune responsiveness to self and foreign antigens [84].

Polymorphisms in non-MHC genes such as cytotoxic T lymphocyte antigen 4 (CTLA4), protein tyrosine phosphatase, non-receptor type 22 (PTPN22) and IL-2 promoter region are also associated with susceptibility to autoimmune diseases [85, 86]. CTLA4, is an inhibitor receptor that binds the costimulatory molecules CD80 and CD86 and limits T-cell responsiveness and promotes anergy [87]. CTLA4 competes with the activating receptor CD28 to block costimulation and also transduces inhibitory signals that act to inhibit ZAP70 and AKT activity [88, 89]. Germline CTLA4 knockout mice develop a lethal syndrome of severe lymphoid organ enlargement and multiple organ lymphocytic infiltrates. These symptoms result from systemic autoimmunity against multiple autoantigens. These studies indicate the key role for CTLA4 in immune regulation. In human disease such as autoimmune Addison's disease (AAD), noticeable demonstration of CTLA4 functions have directed a quest for polymorphisms (single nucleotide or multi nucleotide) that are closely linked with autoimmune

9 | P a g e diseases. Surprisingly, several autoimmune diseases such as T1D and MS display a remarkable correlation of CLTA4 polymorphism that produce reduced number of truncated CTLA4, which shows inhibitory functions [90]. The consequences of producing truncated CTLA4 is undefined as yet, but as expected, the biological influence of the underlying allele is more delicate than what is found in knockout mice or in monogenic disorders.

1.2.4.2. Evidence for environmental influences in tolerance failure

The association between autoimmune disease and environmental factors has not only been demonstrated in mice but also in humans (epigenetic changes induced by environment) [91- 93]. Genomic studies on autoimmune diseases have suggested that barely 20% of phenotypic variation is caused by genetic polymorphism [94]. Moreover, monozygotic twins display variable rates of concordance for autoimmune diseases such as psoriasis or celiac disease [95]. In contrast, T1D, RA and ulcerative colitis have greater and stronger environmental effects (concordance rate is around 40%) [96]. The above data indicates the diverse influences and interconnections between environmental and genetic factors in several autoimmune diseases. Chronological interactions between the onset of autoimmunity and environmental exposures, and associations between seasonality birth patterns and autoimmune disease development have been detected [97, 98]. Several environmental determinants, such as exposure to infectious agents, tobacco, UV light, radiation and chemical compounds are connected with the autoimmune disease development [99]. Most of these environmental determinants can indirectly or directly influence epigenetic changes, which controls the expression of gene and is consequently connected with changes in the immune response. Hence, epigenetics is a crucial molecular mechanisms that could describe the environmental influences on the development of autoimmune disease [100-102].

10 | P a g e

Figure 1.1: Autoimmune disease is assumed to result from a combined effects of genetic and environmental factors such as pathogen induced infections, smoking, diet and stochastic events.

1.3.Therapeutic approaches for autoimmune diseases

The therapeutic approach to autoimmune disease comprises two main approaches. The first is to administer NSAIDs, disease-modifying anti-inflammatory drugs (DMARDs), analgesic, and corticosteroids to limit symptoms. This may incorporate immune modulators. For example, adalimumab and infliximab have the ability to interact with TNF-α receptor and block TNF-α signalling and are effective in RA, ulcerative Colitis and Crohn’s disease. Rituximab (anti- CD20) has been used to treat RA and displays promise in T1D [103]. The second approach,

11 | P a g e using biologicals, encompasses immunotherapies and is intended to modify immune responses rather than merely relieving symptoms. These therapies include anti-cytokine therapy, biological agents that inhibit naïve and memory T-cells function and induce B cell depletion [104]. Targeted deletion or inactivation of autoreactive pathogenic T-cells can be crucial for long-term attempt for autoimmune disease cure. In particular, hematopoietic stem cell transplantation (HSCT) and HSC-mediated gene therapy signifies a promising therapeutic approach for autoimmune diseases resistant to existing treatment strategies [103].

1.3.1. Current therapeutic strategies for tolerance induction in autoimmune disease

Immunological tolerance induction and maintenance is a principle therapeutic aim in autoimmune disease treatment. Current approaches for limiting autoimmune diseases are the use of immunosuppressive agents which can lead to severe infections or result in disease reappearance after immunosuppressive withdrawal. Therefore specific targeted approaches are needed [21]. Administration of monoclonal antibodies (mAb) can contribute better outcomes over classical immunosuppressive drugs and it has already been tested as a more refined approach against specific inflammatory mediators [105]. Some mAb showed significant efficacy in clinical applications. Nonetheless, they induce generalised immunomodulation by targeting immune activation pathways and they also generate impairment of regulatory immune responses [106, 107]. Hence, their constant clinical success upon their lifelong administration is still unclear [108].

Regulatory or immunosuppressive cell therapy is a comparatively new strategy with great prospects. So far the use of autologous and allogenic immunosuppressive cells have been limited in patients with inflammation because immunosuppressive cell frequency is low in these type of patients. Therapeutic approaches targeted to expand and/or improving their regulatory role can also provide new opportunities [109]. This has encouraged the exploration of antigen-specific therapeutic approaches that exert their effects directly on disease-causing antigen-specific immune effector cells.

1.3.2. HSCT and HSC mediated gene therapy for autoimmune disease

E. Donnall Thomas first performed an allogenic bone marrow transplantation (BMT) in 1957 [110]. Currently BMT or hematopoietic stem cell transplantation (HSCT) is an indispensable therapeutic option for not only blood disorders such as severe haemolytic anaemia and aplastic

12 | P a g e anaemia but also malignant haematological disorders, for example, lymphoma and leukemia. Even though mismatched HSCT is also successfully used to treat genetically inherited diseases such as thalassemia or sickle cell anaemia its applications are limited because of treatment- associated risks including conditioning regimen-associated toxicities and complications such as infections and graft versus host disease (GVHD) [111].

Combining gene therapy with HSCT (Figure 1.2) holds the potential to induce life-long tolerance [112, 113]. Memory T-cells present a particularly important barrier to immunotherapy of autoimmune diseases due to their ability to continually generate large pools of auto-antigen specific effector T-cells. Enforced expression of auto-antigens through the combined use of gene therapy and HSCT may allow the silencing of disease-causing memory T-cells in an antigen-specific manner. In such a scenario the balance between tolerance and immunity must be carefully regarded as the latter would only exacerbate disease [72, 114]. To achieve this it is likely that antigen-expression will need to be life-long, requiring the use of retroviral or lentiviral vectors. Engineered bone marrow cells were successfully used in first gene therapy for adenosine deaminase deficiency and encouraged immense hope for gene therapy in the future [115-117]. This was followed by HSC-mediated gene therapy for X-linked severe combined (SCID-X1) [118]. HSCT has already been used to treat autoimmune disease with varying degrees of success. To combine these two (gene therapy and HSCT) for treatment of non-life threatening autoimmune disease will require careful planning and extensive proof-of-concept in animal models.

The recipient conditioning is a crucial step for successful HSCT. Typically the recipient is prepared for HSCT by potent immunosuppressive therapy such as irradiation and/or chemotherapy. Successful conditioning is followed by autologous HSCT (auto-HSCT; HSCs harvested from recipients own body prior to conditioning) or allogenic HSCT (allo-HSCT; cells harvested from different donors) to re-establish recipient immune system. Both auto- HSCT and allo-HSCT were successful in experimental models and is currently in clinical trials. Allo-HSCT has greater toxicity and potential for acute and chronic rejection, which is why auto-HSCT is more preferable than allo-HSCT so far. The risk of GVHD in allo-HSCT, which typically arises from donor allogenic T-cells attacking recipient tissues, is associated with significant mortality and morbidity. In recent times, several researchers successfully transplanted allogenic HSC without myeloablative conditioning regimen to induce transplant tolerance and restoration of self-tolerance in animal models of autoimmune disease [119].

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Though such current technological improvements have made allo-HSCT less toxic but presumably it would not be suitable for less severe treatable autoimmune diseases [120].

Figure 1.2: Steps involved in gene therapy. Cells are collected from the patient’s body, engineered through viral vector and then returned back to the body. Injected altered cells produce desired protein. Image is adapted from https://biol325group6.wordpress.com/about/.

1.3.3. Tolerance induction by autologous HSCT

Ven Bekkum et al. first used autologous HSCT (auto-HSCT) in application to autoimmune disease to treat adjuvant induced arthritis in rats. Treatment of rats with syngeneic HSC and cyclosporine did not result in significant recovery from arthritis [121]. In another study, transfer of syngeneic HSC with total body irradiation resulted in a significant decrease in arthritis severity [122]. Several clinical trials are going on in several autoimmune disease including T1D, systemic and multiple sclerosis, crohn’s disease and juvenile arthritis [123, 124]. These clinical trials have demonstrated substantial abolition of autoimmune pathology and also demonstrated the efficiency of auto-HSCT over existing treatment options for those diseases. Research also showed effectiveness of myeloablative and non-myeloablative auto-HSCT in

14 | P a g e systemic sclerosis and multiple sclerosis [125]. In systemic sclerosis, 10 out of 19 auto-HSCT treated patients improved in pulmonary function and Rodnan skin score. Moreover, no disease progression was found in auto-HSCT treated arm compared to non-auto-HSCT control arm patients [25, 125]. An important advantage of auto-HSCT is there is no risk of graft versus host disease (GVHD) and primary complications of auto-HSCT is treatment related mortality (as high as 15%) and treatment related morbidity includes infections, infertility and risk of malignant diseases [126].

1.3.4. Tolerance induction by allogenic HSCT

Successful allogenic-HSCT (allo-HSCT) requires an increase of immunological tolerance against alloantigens. T-cells are key mediators of acute and chronic graft rejection, graft failure, and GVHD. As discussed earlier, immunological tolerance for T-cells is mediated by two key processes, central and peripheral tolerance. Tolerance induction and maintenance could increase the benefits of transplantation. Immune suppressor cells comprising Tregs, veto cells and NKT-cells are promising treatment options and could modulate immunological tolerance [127, 128].

Typically, auto-HSCT is successful in treating autoimmune disease in animal models including spontaneous autoimmune diabetes in NOD mice and murine spontaneous onset lupus-like disease but failed in other spontaneously arised animal autoimmune disorders [129, 130]. For example, auto-HSCT treatment was unsuccessful in mrl/lpr mice with spontaneous onset lupus- like disease and resulted in transitory disease improvement [131, 132]. So, allo-HSCT from a non-autoimmune prone donor is required for the treatment of spontaneous onset autoimmune- like diseases [133, 134]. Allo-HSCT from normal strain donor/non autoimmune prone donor was successful in murine spontaneous onset lupus-like disease and spontaneous onset diabetes in non-obese diabetic (NOD) mice [135-137]. However, HSCs tolerizing effect is best validated by donor specific organ tolerance when treated with a combination of solid organ and HSC transplant from the same donor [138, 139].

Donor specific organ tolerance requires a myeloablative or nonmyeloablative conditioning regime to deplete recipient followed by allo-HSCT [140, 141]. Although full or mixed hematopoietic chimerism is closely associated with donor specific tolerance, the molecular and cellular mechanisms of donor specific tolerance is not fully understood yet [142]. Fas ligand is a type II surface protein from TNF family and binds with its receptor (FasR)

15 | P a g e to induce apoptosis. Fas knockout mice does not show hematopoietic mixed chimerism induced tolerance. Hence, Fas ligand expression is very important for HSC induced donor specific tolerance [143]. It has been suggested that allo-HSCT can induce tolerance by depletion of self- reactive T and B cell repertoire. This phenomenon has been termed graft versus autoimmunity (GVA) [144, 145]. By using non-myeloablative conditioning regimes, natural animal model of autoimmunity have been treated in the hematopoietic mixed chimerism settings [146, 147].

Though, theoretically GVA may be advantageous, the significant toxicity from transferred donor cells acting on recipient’s cells results in a medical complication termed as graft versus host disease (GVHD). Non-myeloablative conditioning has less risk of GVHD related to myeloablative conditioning regimes. The risk of GVHD might be reduced as a result of reduced inflammatory cytokine release, decreased regime related tissue damage, and veto of donor reactive T and B cells principally CD8+ cells by HSCs of host origin [148, 149].

Anecdotal case reports suggest patient go through allo-HSCT for malignancy may achieve long-lasting autoimmune disease remission [150-152]. The majority of patients with autoimmune disease treated in this way maintain this reduction for their entire life after immunosuppressant withdrawal. Rarely have patients relapsed even with 100% donor hematopoietic chimerism. Occasionally, clinically symptom-free donors can also have some subclinical autoimmune disease such as RA and there are reports this can be adoptively transferred to the recipient [153].

1.3.5. Hematopoietic chimerism contributes to tolerance in autoimmune disease

A robust and consistent tolerance might be induced by mixed allogenic hematopoietic chimerism and potential development of toxicity-free regimens may play a vital role in widespread application of this in the clinic. Two examples of such cases have been discussed below where tolerance was induced by mixed hematopoietic chimerism.

In one case, a Stanford University group has developed a conditioning regime based on experiments on mice, animals, non-human primates, and on clinical studies which involves total lymphoid irradiation (TLI) and anti- globulin (ATG) to induce tolerance with either full or mixed chimerism in HLA-matched kidney transplantation. In this conditioning protocol, G-CSF mobilized CD34+ cells were transplanted followed by post-transplant immunosuppression [154-156] to improve the HSCT outcomes for the treatment of

16 | P a g e hematologic malignancies [157, 158]. Almost all of the patients (15 out of 16) treated with the Stanford regime achieved stable mixed chimerism and half of them developed mixed chimerism lasting more than six months after complete immunosuppressive agents withdrawal. Twenty-five percent of the total patients were continued to treat with immunosuppressant (cyclosporine) because of the reappearance of focal segmental glomerulosclerosis and rejection incidents. They also investigated tolerance induction in renal allograft with HLA-non-identical kidney transplantation, which was unsuccessful to induce tolerance in HLA-non-identical hosts by this procedure [154].

A second clinical strategy by Northwestern University was reported where low dose total body irradiation (TBI), high dose Cy and fludarabine were used for conditioning for both pre and post-HSCT. After conditioning, donor HSC was transferred for the treatment of eight HLA- non-identical kidney transplant recipients. This was an improvement on previous protocols and full donor chimerism was attained in several cases without GVHD [159, 160]. The outcomes of this study portends that either the conditioning regime was myeloablative or the donor T- cells destroyed host haematopoiesis through graft versus host (GVH) reaction. Though the study procedure was not completely disclosed it was claimed that tolerogenic CD8+/TCR- facilitating cells were administered to prevent rejection [161, 162] and the standard criteria for prevention of all immunosuppression maintenance was specified as stable donor chimerism (eight out of fifteen patients) without GVHD for one year [160]. In these patients, the highly myeloablative conditioning and potential GVHR associated with the transplantation is highlighted by the extended thrombocytopenia and neutropenia which required G-CSF and platelet transfusion support [159, 160].

These results differ significantly to patients with hematologic malignancies who received HSCT from non-identical donors with a similar regime. In 34% cases acute GVHD occured, along with considerable chronic GVHD was also registered [163, 164]. This makes the Northwestern analysis unexpected if validated with extended follow-up and a greater volume of patients. Though, considerable morbidity because of potential infections were reported and it is possible that these patients might not recover standard immunological competency [165- 167]. It is to be considered that the patients in the HLA-non-identical HSCT clinical trials at Massachusetts General Hospital have not experienced any considerable opportunistic infections.

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1.3.6. Tolerance induction through gene-modified HSC

HSC has the capability to differentiate into different leukocyte lineages such as T-cell, B-cell and DC and could be an attractive target for gene modification to induce immune tolerance. The idea of HSC modification for tolerance induction arose from observations of tolerance induction to donor MHC proteins due to development of host-donor mixed chimerism after allo-HSCT [168, 169]. But, as discussed above most of the cases of allo-HSCT results in graft rejection and GVHD. Then the theory arose that gene-modified auto-HSCT could be a solution to generate mixed chimerism as well as reduce the GVHD risk following transplantation [170- 172]. Several research groups induced immune tolerance through the expression of targeted transgene to a specific cell-lineage by using tissue-specific promoter or microRNA (miRNA) [173-175]. Tolerance induction through gene-modified HSCT has been successful so far in several disease models, e.g. foreign cytosolic reporter gene expression [176, 177] or allogenic MHC I and II gene expression [178, 179] in BM-derived cells was successful to skin grafts. Gene-modified HSCT express myelin oligodendrocyte glycoprotein to DC under myeloablative or non-myeloablative conditioning delayed EAE onset [180, 181]. Gene- modified HSC transfer was successful in inducing antigen-specific immune tolerance in several mouse models including non-obese diabetes mouse models [4], mouse models [182, 183] and haemophilia disease model [184, 185].

1.3.7. DC-induced peripheral tolerance induction

DC is one of the key contributors to control immune response and T-cell tolerance. It controls immune responses through receptor signalling regulation either by activation or by inhibition [186]. T-cells are activated when TCR encounters DC presented peptide-MHC complex in the presence of co-stimulatory interactions (CD28-CD80/CD86 interactions). Interaction between T-cell expressed CD40L and APC-expressed CD40 upregulates CD80/CD86 expression which subsequently activates T-cells [187, 188]. CD40/CD40L interaction express OX40L on DC in the presence of pro-inflammatory cytokines [189]. Th2 differentiation initiates following OX40 and OX40L interaction on T-cells [190]. DC maintains a functionally immature state and continuously process and present autoantigens in the absence of pathogen-associated molecular pattern molecules or damage-associated molecular pattern molecules generated through infection and trauma [191]. In this functionally immature state, DC express little to no co-stimulatory molecules and antigen-specific interaction with T-cells leads to immune tolerance. Low-doses of intact soluble protein expression to steady state DC enables to induce

18 | P a g e antigen-specific T-cell tolerance through T-cell deletion [192, 193]. Several reports show this DC-induced peripheral tolerance through using either transgenic mice express antigen targeted to DC [72, 192] or gene-modified DC express targeted antigen under the control of CD11c promoter [48, 76]. All these approaches lead to DC-induced peripheral T-cell tolerance when CD8+ and CD4+ T-cells encounter antigens under non-infectious conditions. BM-derived immature DC allows alloantigen specific T-cell anergy in vivo and in vitro [194, 195]. Steptoe et al. demonstrated complete prevention of diabetes in NOD mouse model by transferring gene- modified HSC encode proinsulin targeted to MHC class II+ APC [4]. Coleman et al. also showed targeted ovalbumin (OVA) expression to DC or other antigen presenting cells induce antigen-specific T-cell tolerance in naïve and primed recipient however light total body irradiation (300 cGy) was used [173].

1.4. HSC niche and conditioning regimes 1.4.1. HSC niche: Home for hematopoietic stem cells

Bone marrow is a very complex organ, having various types of haematopoietic and non- haematopoietic components that is surrounded by a shell of innervated and vascularized bone [196, 197]. A stem cell niche is a specific site for hematopoietic stem cell where they reside and go through self-renewal and produce large numbers of progeny cells. Structurally, the niche is formed by ancillary cells that provide a microenvironment for stem cells [136, 198]. The niche is perivascular, generated partly by endothelial and mesenchymal stromal cells and often, positioned near trabecular bone [196]. Typically trabecular bones are found throughout the metaphysis such many cells in this region are close to the bone surface. The edge of bone and BM is well-known as endosteum, and enclosed by bone-lining cells that include bone-resorbing osteoclasts and bone-forming osteoblasts. Oxygen, growth factors and nutrients are carried by arteries into the BM, before feeding into sinusoids, which combine as a central sinus to form the venous circulation. Sinusoids are specific and specialized venules that form a reticular network of fenestrated vessels that permit cells to pass in and out of circulation [199]. The bone marrow is cellularly complex with CD150+CD48−CD117+lineage− HSC residing in close interaction not only with perivascular and vascular cells but also megakaryocytes and other haematopoietic cells [196, 200].

HSC originate in body tissues or embryos and migrate to BM during embryonic development [201]. In mammals, lifespans of most blood cells are relatively short. Hence, HSC continuously differentiate into multiple lineages of different blood cell types, simultaneously duplicating

19 | P a g e themselves by self-renewal to maintain the stem cell pool in the BM [201-203]. Self-renewal and differentiation of stem cells are tightly regulated intrinsically by gene expression in a cell- type-specific manner and is modulated through interactions with extrinsic cues from the environment, such as growth factors [204].

However, to achieve successful engraftment of exogenous HSC, the transplanted cells must have access to HSC niche space within the recipient BM. Because free HSC niche space is critical for transplanted HSC self-renewal and differentiation. So, recipient conditioning is required to create space inside HSC niche and donor engraftment is highly dependent on percent of HSC niche space created after recipient conditioning [204-207].

1.4.2. Current conditioning regimes used clinically

The main aim of conditioning regimens for HSCT is to deplete the autoreactive lymphocytes and create space in the bone marrow for immune rescue by HSC infusion. Cyclophosphamide has a long history as a chemotherapeutic agent for cancer treatment and is also commonly used for HSCT conditioning [208]. It is a nitrogen mustard alkylating agent that once metabolized, forms intra- and inter- strand DNA crosslinks which cause cell cytotoxicity. In early 2000, Brodsky et al. treated several autoimmune diseases with higher cyclophosphamide dose (200mg/kg) and without HSC support, but relapse was very common in all cases except SLE. A strong myeloablative regime with HSC infusion was necessary for robust responses [209].

The ideal conditioning regime should create space inside BM for donor HSC and deplete autoreactive lymphocytes and without creating lymphocytopaenia and neutropaenia. But unfortunately these types of conditioning regimes do not exist currently. The conditioning regimes in use currently were principally developed for malignancies. Conditioning regimes typically used in autoimmune diseases include cyclophosphamide [210-213]; cyclophosphamide and antithymocyte globulin (ATG) or rat anti-human monoclonal anti- CD52 (ATG/Cy or Campath-1H/Cy respectively) [214-217]; Etoposide, BEAM (carmustine, etoposide, cytarabine, and melphalan) [218]; total body irradiation and cyclophosphamide (TBI/Cy) [219]; TBI, cyclophosphamide and antithymocyte (TBI/Cy/ATG) [220]; cyclophosphamide and busulfan (Cy/Bu) [221]; cyclophosphamide, busulfan and ATG (Cy/Bu/ATG) [222]; thiotepa and cyclophosphamide (TT/Cy) [223] and fludarabine based regimes. Among these, the Autoimmune Disease Working Party (ADWP) from the European Society for Blood and Marrow Transplantation (EBMT) suggests 200mg/kg

20 | P a g e cyclophosphamide with monoclonal or polyclonal serotherapy for adults and 120mg/kg Cy with fludarabine (150mg/m2) and antithymocyte globulin (ATG) for children [224]. Subsequently, hematopoietic stem cells (HSC) are infused at a minimum dose of 2 million CD34 positive cells/kg [225]. Typically, patients are discharged from hospital within 1-3 weeks after HSC infusion, when number increases. However, most of the patients remain severely lymphocytopenic for several weeks or even months following HSCT while their immune system rebuilds [208]. It is recommended to warn all patients about regimen related late toxicities such as infertility, malignancies and organ damage.

1.4.3. Specific conditioning regimes

Current research has revealed that monoclonal anti c-kit antibodies (ACK2) can deplete a significant amount of HSC from bone marrow niches and permit a successful HSC engraftment in fully immunodeficient mice [226]. Chhabra et al., [227] showed that host HSC depletion is reliant on Fc-mediated antibody effector functions, and augmenting effector function by the blockade of integrin associated protein (CD47) could offer ACK2 conditioning to completely immunocompetent mice. This cumulative approach (ACK2 therapy and CD47 blockade) can lead to the elimination of over 99% of recipient HSPCs and induce healthy multilineage blood re-formation following HSCT [226, 227]. A recent study also suggested valine restricted diet cleared the mice BMN and provided donor HSPC engraftment without myeloablative conditioning [228].

Recent studies suggest an improvement on antibody depletion approaches and saporin based immunotoxins which specifically or non-specifically target hematopoietic cells could be a promising approach to improve recipient conditioning [229, 230]. Saporin is a catalytic N- glycosidase ribosome-inactivating plant toxin extracted from Saponaria officinalis. After cell entry it binds with ribosomes and blocks protein synthesis which ultimately induces cell death (Figure 1.3) [231]. Saporin does not have a cell entry domain and is not toxic unless conjugated with a monoclonal antibody or ligand that targets a receptor or cell-surface molecule that undergoes internalization [232]. When targeted by a specific mAb or ligand it forms a highly effective immunotoxin [231]. It is reported that CD45-SAP consisting of streptavidin-saporin (SAP) conjugated to a biotinylated anti-CD45 monoclonal antibody, is a potent CD45+ cell- specific immunotoxin [229]. Administration of a high-dose (3mg/kg) CD45-SAP resulted in extensive (>98%) depletion of HSC allowing donor HSC to engraft at very high levels when

21 | P a g e transferred to CD45-SAP treated mice [229]. Palchaudhuri et al. also screened a series of antigen targets such as CD49d, CD84, CD90, CD133, CD135, and CD184 which is expressed by both human and mice HSC. But CD45-SAP was most effective to deplete recipient HSC [229]. This targeted immunotoxin-based conditioning could replace genotoxic conditioning and provide proof-of-principle that irradiation and chemotherapy is not required for recipient conditioning. However, CD45-SAP is immunoablative as peripheral leukocytes express significant amounts of CD45.

Figure 1.3: Mechanism of action of saporin. After entering the cell cytosol it binds with ribosome and induces apoptosis. It also cause apoptosis activation by both caspase independent or dependent pathway. Image adapted from Polito et al. 2013 [232].

1.5. Modulation of the immune system by immunosuppressive conditioning in auto- HSCT

Effective immunotherapy has the ability to delete self-reactive T and B cells, thus ‘resetting’ the immunological system after auto-HSCT. Considering the hypothesized character of microbial agents and unknown environmental factors trigger in stimulating autoimmune

22 | P a g e diseases, it is promising that deletion of existing B and T-cell repertoires permit the patient’s immunological system to “start from scratch” and rebuild [120]. Freshly developing T and B cells from the thymus and bone marrow respectively might encounter self-antigen and experience self-tolerization. The concordance rates of autoimmune diseases in identical twins may be used to estimate the reappearance ratio if entire pre-existing self-reactive T and B cells were deleted prior auto-HSCT. Depending on the disease, reappearance range varies from 10 to 35% and suggests that this procedure might result in a consistent reduction in almost all of the recipients if total depletion of self-reactive T and B cells could be achieved [120, 233, 234]. However, it is unclear whether or not self-reactive memory T and B cells and long-lasting plasma cells could be totally deleted from the target organs and through a conditioning approach. In clinical trials, partial immune depletion can cause sub-optimal reactions and greater ratio of initial reversion after auto-HSCT. In spite of partial depletion, maintenance of self-reactions is dependent on multiple cell populations and could lead to disease improvement. For instance, strong chemotherapeutic agents for conditioning stimulates severe and extended CD4+ T-cells suppression characterized by a reversed CD8+:CD4+ T- cells ratio, a predominance of Tmem and predominant extra thymic pathways of T-cell recovery [235]. T-cells recovered after chemotherapeutic conditioning show enhanced susceptibility to apoptotic death and altered function. Hence, altered immunological reconstruction and immunosuppressive therapy after conditioning can result in extended remission of disease [120]. Auto-HSCT using mobilized stem cells, higher chemotherapeutic dose and immunosuppression permits intensely immune depletive therapy and it has been already introduced in several clinical trials in autoimmune disease patients such as MS, RA, SLE and myasthenia gravis [236, 237]. Such transplantation includes several conditioning regimes, exclusion of T-cells, various HSC sources and several HSC mobilization procedures from the bone marrow niche. Auto-HSCT has been shown in clinical trial phase I and II as a practical transplantation procedures and has the ability to reduce autoimmune diseases [123, 238]. However, the efficiency of auto-HSCT varies from one disease to another but over one in three patients have constant and robust reduction without any further need of immunosuppression [237, 239, 240]. Phase III clinical trial has already started in Europe and United States. Preferably the data from phase III clinical trials will help to explain if key advantages of these therapies relates to the depletion capability of self-reactive T and B cells or re-establish immune regulation that restricts autoimmune diseases. Moreover these data will also explain proof of

23 | P a g e efficacy, which is crucial for broader auto-HSCT application in numerous autoimmune diseases [239, 240].

1.6. Hypothesis and Aims

Dysregulated or pathogenic immune responses underlie many autoimmune and inflammatory diseases as well as immune resistance to the effectiveness of protein replacement therapies. Underlying these conditions is a common state where the body’s immune system has mounted a response to normal body components, normally innocuous environmental agents or therapeutically administered proteins. One important goal for alleviating these conditions is the development of effective therapeutic strategies that can turn-off undesirable immune responses that have become established. Transferring gene-engineered HSC expressing antigen targeted to DC is established as a robust method to induce antigen-specific T-cell tolerance. One challenge for this HSC-mediated gene therapy is that the host immune system can ‘resist’ the growth of transferred cells by mounting an immune response, which kills the transferred donor cells. However successful transferred donor HSPC engraftment depends on empty BM niche spaces. Conditioning regimes must create sufficient empty spaces inside BM niche for the donor HSPC. The current conditioning approaches of total body irradiation and/or high-dose chemotherapy creates empty spaces inside BM niche by eliminating resident HSC but these are non-specific and simultaneously induce lymphocytopenia, neutropenia and anemia. Total body irradiation has a key benefit that it reduces competition for the sites in which transferred bone marrow cells grow and provides a ‘niche’ for transferred cells to grow in. However, these procedures are non-specific to other tissues and can cause lifelong complications. To overcome this toxic conditioning for BMT, HSC depletion from bone marrow niches with targeted immunotoxin could be a crucial step and a great opportunity for advancing clinical applicability of HSC-mediated gene therapy to induce antigen-specific T-cell tolerance.

By titrating the dose of CD45.2-SAP, it might be possible to achieve 30-40% donor HSC chimerism, a level previously established as effective for inducing immune tolerance [4, 241]. A benefit of this might be that the pan-leukocyte depleting activity of CD45.2-SAP may be reduced allowing some preservation of immunity. Or the use of HSPC-specific immunotoxin c-kit-SAP might facilitate sufficient amount of donor chimerism for tolerance induction sparing non-hematopoietic cells. So, it is anticipated that saporin based immunotoxin could efficiently deplete recipient LT-HSC and create engraftment ‘space’ for transferred genetically engineered LT-HSC.

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Hypotheses of this thesis:

1. Non-genotoxic conditioning using HSPC non-specific immunotoxin will facilitate tolerance induction through stable mixed hematopoietic chimerism of gene-modified antigen-encoding bone marrow. 2. Hematopoietic stem and progenitor cell specific immunotoxin will induce antigen- specific T-cell tolerance by allowing donor hematopoietic chimerism.

Major aims of this thesis:

1. To define whether non-genotoxic leukocyte depleting immunotoxin preferentially depletes long-term repopulating hematopoietic stem cells (LT-HSC) and enables stable long term molecular chimerism using gene-engineered BM. 2. To develop sufficient level of donor chimerism to induce antigen-specific T-cell tolerance by using HSPC non-specific immunotoxin as recipient conditioning agent. 3. To develop sufficient level of donor chimerism to induce antigen-specific T-cell tolerance by using HSPC specific immunotoxin as recipient conditioning agent.

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Chapter 2 Materials and Methods

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2.1. Mice Mice were bred or housed in specific pathogen-free housing at the Translational Research Institute Biological Resource Facility (TRI-BRF) (Brisbane, Australia). All mice were euthanized by CO2 asphyxiation prior to tissue collection. Unless stated otherwise, 8-16 weeks aged male mice were used and all experiments were approved by The University of Queensland Animal Ethics Committee (UQAEC).

2.1.1. Congenic mice strains

C57BL/6JArc mice C57BL/6JArc express the CD45.2 allelic variant of the CD45 membrane bound pan-leukocyte molecule. C57BL/6J mice were purchased from the Animal Resource Centre (ARC, Perth, Australia) at 6-8 weeks of age.

B6.SJL-PtprcaPep3b/BoyJArc mice B6.SJL-PtprcaPep3b/BoyJArc (henceforth referred to as B6.SJL) mice are commonly used in immunology and adoptive transfer experiments as they express the CD45.1 allelic variant form of the CD45 membrane-bound pan-leukocyte molecule expression which allows identification of transferred cells in CD45.2 congenic mice. B6.SJL mice were purchased from the ARC (Perth, Australia) at 6-8 weeks of age. These mice were used in BMT studies with C57BL/6J to track host and donor chimerism based on their allelic CD45 expression (CD45.2 and CD45.1).

2.1.2. Transgenic mice strains 11c.OVA mice 11c.OVA mice (CD45.2+) with C57BL/6J background express a membrane bound OVA construct under the control of the CD11c (Itgax) promoter. Only CD11chi expressing conventional DC present OVA [76]. 11c.OVA mice were bred and maintained at the Translational Research Institute Biological Research Facility (TRI-BRF, Brisbane, Australia).

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11c.45.1

11c.45.1 mice with B6.SJL-ptprca background express a membrane bound OVA construct under the control of the CD11c (Itgax) promoter. Only CD11chi expressing conventional DC present OVA [76]. These DC exist in steady-state. 11c.45.1 mice express the CD45.1 allelic variant of the CD45 membrane bound pan-leukocyte molecule. Mice were screened by polymerase chain reaction (PCR) to detect OVA transgene. Mice did not possess OVA transgene considered as non-OVA expressing littermates and were used as wild type controls. 11c.45.1 mice were bred and maintained at the TRI-BRF (Brisbane, Australia).

OT-I mice Developed to enable studies on an enlarged population of antigen-specific T cells during an immune response. OVA-specific, TCR transgenic mice that are H-2Kb class I-restricted carry

CD8 T cells specific for the CD8+ immunodominant OVA257-264, commonly referred to as SIINFEKL [242]. OT-I mice express CD45.2 and were bred and maintained at the TRI-BRF (Brisbane, Australia).

2.2. Reagents

2.2.1. Proteins

Saporin Saporin processed from Saponaria officinalis, is a 29 kDa ribosome binding and inactivating protein. Saporin was dissolved in mouse tonicity phosphate buffer saline (MT-PBS) to a concentration of 0.5mg/ml or 1 mg/ml based on experiments and stored at 4oC. Saporin was purchased from Sigma-Aldrich (St. Louis, MA, USA).

Streptavidin Streptavidin (SA) is a 52.8 kDa protein from Streptomyces avidinii. It is a very high affinity protein for biotin. It is a tetrameric protein and each subunit is able to bind a single biotin molecule. Streptavidin was diluted in MT-PBS and used to prepare streptavidin-saporin conjugates. Streptavidin and Streptavidin conjugation kits were purchased from Promega (WI, USA) and Abcam (ab102921, Cambridge, UK) respectively.

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Ovalbumin Ovalbumin (OVA) is a glycoprotein and the predominant protein of chicken egg white. Ovalbumin was dissolved in MT-PBS (1mg/ml), filter sterilized and used on the day of preparation. Mice were immunized subcutaneously with 50 μg OVA in conjunction with 20 μg QuilA adjuvant at the base of the tail. OVA was also used in FACS, ELISPOT, or made up in RPMI culture media supplemented with 5% FCS for splenocyte re-stimulation. OVA was purchased from Sigma-Aldrich (MA, USA).

Bovine serum albumin (BSA) BSA (Fraction V) purified from bovine blood plasma. Lyophilised BSA was dissolved to 1% w/v in MT-PBS for FACS wash. BSA was purchased from Bovogen Biologicals (Melbourne, Australia)

2.2.2. Peptides

OVA257-264 b OVA257-264 (SIINFEKL) is the immunodominant H-2K -restricted determinant of OVA and together with H-2Kb forms the MHC class I/peptide complex recognized by OT-I T cells.

OVA257-264 was dissolved either in MT-PBS or R5 medium (RPMI-1640 5% FCS) to 10µg/ml 0 concentration, aliquoted and stored at -30 C. OVA257-264 was purchased from Auspep (Melbourne, Australia).

OVA323-339 b OVA323--339 (ISQAVHAAHAEINEAGR) is the immunodominant H-2 -restricted determinant of OVA and together with I-Ab or I-Ad forms an MHC class II/peptide complex recognized by

CD4+ T cells. OVA323-339 was dissolved in R5 medium (RPMI-1640 5% FCS) to 10mg/ml concentration before use. OVA257-264 was purchased from Auspep (Melbourne, Australia).

2.2.3. Chemicals Table 2.1. List of chemicals Chemical Name Company Preparation Use Ethidium bromide Sigma-Aldrich Purchased as 10mg/ml Used in agarose gel (MA, USA) electrophoresis to fluorescently label PCR amplified DNA

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GelRed Biotium (CA, Used in agarose gel USA) electrophoresis to fluorescently label PCR amplified DNA Coomassie Gift from Dr Prepared to 0.1% w/v Used in sodium dodecyl Brilliant Blue Jeremy Brooks, in methanol, H2O and sulphate–polyacrylamide The University acetic acid gel electrophoresis of Queensland (SDS-PAGE) (Brisbane, Australia).

Paraformaldehyde Sigma-Aldrich 10g PFA was weighed Used to fix the tibia and (PFA) (MA, USA) and added into 200ml femur for histology pre-warmed deionized H2O. 25µl of 10N NaOH was added and mixed until powder was completely dissolved. After dissolving, 25ml 1M phosphate buffer (pH 7.4) was added and made up to 250ml with deionized water. 4% PFA was prepared fresh just prior to use unless stated otherwise.

Ethylenediamine Sigma-Aldrich 2L stock was prepared Used to decalcify bones tetraacetate (MA, USA) by combining EDTA for histology sample (EDTA) solution (280g), NaOH (40g) preparation. and 2000ml H20. pH was adjusted to 7.2 by using concentrated hydrochloric acid (HCl).

2.2.4. Buffers Table 2.2. List of buffers Buffer Name Preparation Use Mouse-tonicity 10x stock was prepared by combining Used for all experiments phosphate buffered NaH2PO4.1H20 (5.52g/L), saline (MT-PBS) Na2HPO4.2H20 (28.5g/L) and NaCl (87g/L) dissolved in MilliQ water. 1x working concentration was prepared as a 1 in 10 dilution and sterilized by autoclaving prior to use.

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MT-PBS/2.5% MT-PBS supplemented with 2.5% FCS Routinely used for tissue FCS or FBS or FBS collection and cell washes. FCS was added for sterile procedures such as bone marrow cell transfers, in vitro culture and FBS was used for ex- vivo experiments.

ACK red cell lysis ACK buffer was prepared in laboratory Used routinely to lyse buffer and was composed of Na2EDTA (0.0372 red blood cells from g/L), KHCO3 (1 g/L), NH4Cl (8.29 g/L) peripheral blood, spleen dissolved in MilliQ water. pH was and bone marrow. adjusted to ~7.2-7.4, filter sterilized prior to storage at 4°C. FACS wash buffer Prepared in 10x concentrations of 10% Used to wash cells after BSA 200mM EDTA in MT-PBS and antibody staining stored in -20°C freezer. 1x FACS wash was prepared by diluting 50mL of 10x FACS wash in 500mL of sterile MT-PBS and was stored in 4°C refrigerator. FACS block Prepared in laboratory from the Used block the cells supernatant of 2.4G2 hybridoma cells before antibody staining producing anti-CD16/32. BSA (1%) was to prevent non-specific mixed with the supernatant and filter- mAb binding sterilized, made aliquots and stored in - 20°C freezer. ELISPOT wash Prepared from 0.05% Tween 20 in PBS Used in the ELISPOT buffer technique to wash the plates between steps. Tris-acetate-EDTA TAE buffer (50x) was prepared by Used to cast gels for (TAE) buffer dissolving Tris base (242g/1L) and EDTA agarose gel (18.6g/1L) in 700mL MilliQ water. 57mL electrophoresis as buffer of glacial acetic acid was then added and during DNA runs. total volume was adjusted to 1L with MilliQ water. pH was adjusted to 8.0. 1x concentration was prepared by diluting with MilliQ water prior to use. SDS-PAGE Running buffer was prepared at 10x stock Used in SDS-PAGE as a running buffer by dissolving Tris base (144.0g/L) in 1L running buffer MilliQ water with 10% w/v SDS. 1x concentration was prepared by diluting with MilliQ water prior to use. Sample loading Loading buffer was purchased at 2X stock Used to prepare protein buffer concentration. It was mixed in equal samples for SDS-PAGE volume with cell lysate or purified protein. Sample reducing Sample reducing buffer was purchased at Used to reduce buffer 10X stock concentration. It was directly disulphide bridges for added to protein sample to make 1X SDS-PAGE working concentration.

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2.2.5. Adjuvants Quil-A Quil-A purified from Quillaja saponaria is a saponin adjuvant which induces potent CD8+ responses. QuilA was dissolved in pyrogen free H2O (Baxter, IL, USA) to 0.4mg/ml concentration, aliquoted to 500µl per vial, and stored at -30oC. Aliquoted Quil-A was diluted to 0.2mg/mL in sterile MT- PBS and OVA (0.5mg/mL). 50μg of OVA and 20μg of Quil-A per mouse was used for immunization (i.e. 100μL of 0.2mg/mL). Quil-A was purchased from Soperfos Biosector (DK-Vedback, Denmark).

2.2.6. Tissue culture media Complete Roswell Park Memorial Institute 1640 (RPMI-1640) Medium Complete RPMI-1640 comprises of sterile liquid RPMI-1640 (Gibco-Invitrogen, Carlsbad, CA, USA) supplemented with 5x10-5M 2-mercaptoethanol (Sigma-Aldrich, St. Louis, MA, USA), 1mM sodium pyruvate (Gibco-Invitrogen, Carlsbad, CA, USA), 1x10-4M non-essential amino acids (Gibco-Invitrogen, Carlsbad, CA, USA), 100 units/ml streptomycin and 2mM L- glutamine (Gibco-Invitrogen, Carlsbad, CA, USA) and 100 units/ml penicillin.

X-VIVOTM 15 Serum-free Hematopoietic Cell Medium X-VIVO™ (Lonza, Basel, Switzerland) is a serum-free haematopoietic cell medium offer nutritionally balanced and complete environments for HSPC cultivation and proliferation. This medium composed of human albumin (pharmaceutical grade), pasteurised human transferrin and recombinant human insulin. This medium does not have any artificial stimulator for cellular proliferation, growth factors or undefined nutritional supplements.

2.2.7. Electrophoresis gels Agarose gel Agarose gel (2%) was used for separating DNA product amplified by PCR. Molecular Biology Grade agarose (2g) was dissolved in 100mL TAE buffer by using microwave. GelRed (10μL) or ethidium bromide (2μL) was added just before pouring into the tray. SDS-PAGE gels SDS-PAGE gels, comprising of 8% reducing gel and 4% stacking gel (unless stated otherwise) were prepared in laboratory. To prepare 8% reducing gel, 8.5ml H2O, 3.2ml Polyacrylamide

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(40%), 4.0ml resolving buffer (Tris 1.5M pH 8.8), 160µl SDS (10% w/v), 160µl ammonium persulphate (10%) and 16µl N,N,N',N'-tetramethylethane-1,2-diamine (TEMED) were mixed.

And 6.3ml H2O, 1ml Polyacrylamide (40%), 2.5ml resolving buffer (Tris 1.5M pH 8.8), 100µl SDS (10% w/v), 100µl ammonium persulphate (10%) and 10µl TEMED were mixed to prepare 4% stacking gel. Gels were cast in 0.5-1mm glass plates (BioRad CA, USA).

2.2.8. List of monoclonal antibodies (mAb) All mAbs and streptavidin conjugates used throughout this study are listed in Table 2.3 and 2.4

Table 2.3: List of mAbs used in flow cytometry Antibody Fluorophore Host/Isotype Clone Supplier Catalogue Anti-mouse CD3 FITC Armenian Hamster 145-2C11 Biolegend 100306 IgG Anti-mouse CD4 FITC Rat IgG2a k GK1.5 Biolegend 100406 Anti-mouse CD4 PE Rat IgG2a k GK1.5 Biolegend 100408 Anti-mouse CD8α APC Rat IgG2a k 53-6.7 Biolegend 100712 Anti-mouse CD8α FITC Rat IgG2a k 53-6.7 Biolegend 100706 Anti-mouse CD8α PE Rat IgG2a k 53-6.7 Biolegend 100708 Anti-mouse CD8α PerCPCy5.5 Rat IgG2a k 53-6.7 Biolegend 100734 Anti-mouse CD8α PE.Cy7 Rat IgG2a k 53-6.7 Biolegend 100722 Anti-mouse CD11b FITC Rat IgG2b M1/70 Grown in - house Anti-mouse CD11b PerCPCy5.5 Rat IgG2b k M1/70 Biolegend 101228 Anti-mouse CD11c PE Hamster IgG N418 Biolegend 117308 Anti-mouse PECy7 93 Biolegend 101318 Rat IgG2a k CD16/32 Anti-mouse CD19 APC Rat IgG2a k 6D5 Biolegend 115512 Anti-mouse CD19 BV785 Rat IgG2a k 6D5 Biolegend 115543 Anti-mouse CD34 PE Rat IgG2a k RAM34 BD 551387 Pharminogen Anti-mouse CD44 PE Rat IgG2b K IM7 Biolegend 103008 Anti-mouse FITC Rat IgG2a k RA3-6B2 Biolegend 103206 CD45R Anti-mouse APC Rat IgG2a k RA3-6B2 Biolegend 103212 CD45R Anti-mouse CD45 PE.Cy7 Rat IgG2b k 30-F11 Biolegend 103114 Anti-mouse APC Mouse IgG2a k A20 Biolegend 110714 CD45.1 Anti-mouse PE.Cy7 Mouse IgG2a k A20 Biolegend 110730 CD45.1 Anti-mouse FITC Mouse IgG2a k 104 Biolegend 109806 CD45.2 Anti-mouse APC Mouse IgG2a k 104 Biolegend 109814 CD45.2 Anti-mouse PE/Cy7 Mouse IgG2a k 104 Biolegend 109830 CD45.2

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Anti-mouse PE Mouse IgG2a k 104 Biolegend 109808 CD45.2 Anti-mouse Biotin Mouse IgG2a k 104 Biolegend 109804 CD45.2 Anti-mouse CD48 APC Hamster IgG HM48-1 Biolegend 103412 Anti-mouse CD62L FITC Rat IgG2a K MEL-14 Biolegend 104406 Anti-mouse PECy7 CD115 Anti-mouse CD117 FITC Rat IgG2b 2B8 Biolegend 105812 (c-kit) Anti-mouse CD117 PE Rat IgG2b 2B8 Biolegend 105808 (c-kit) Anti-mouse CD117 APC Rat IgG2b 2B8 Biolegend 105812 (c-kit) Anti-mouse CD117 Biotin Rat IgG2b 2B8 Biolegend 105804 (c-kit) Anti-mouse CD117 Biotin Rat IgG2b ACK-2 Biolegend 135129 (c-kit) Anti-mouse CD127 Biotin Rat IgG2a k B12-1 BD 555288 Pharminogen Anti-mouse CD127 BV510 Rat IgG2a k A7R34 Biolegend 135033 Anti-mouse CD150 PE.Cy7 Mouse IgG2a k TC15- Biolegend 115914 12F12.2 Anti-mouse LY- FITC Rat IgG2a k RB6-8C5 Grown in - 6G/C (Gr-1) house Anti-mouse LY- PerCPCy5.5 Rat IgG2a D7 Biolegend 108124 6A/E (Sca-1) Anti-mouse FITC Rat IgG2b k TER119 Grown in - TER119 house Anti-mouse TCR APC Rat IgG2a K B20.1 Biolegend 127810 Vα2 Anti-mouse Ly6C PE Rat IgG2c K HK1.4 Biolegend 128008 Anti-mouse Ly6G FITC Rat IgG2a K 1A8 Biolegend 127606 Anti-mouse F4/80 Biotin Rat IgG2a K BM8 Biolegend 123106

Table 2.4: List of secondaries used in flow cytometry Antibody Fluorophore Host/Isotype Clone Supplier Catalogue Streptavidin PE CALTAG SA1004-4 Streptavidin PE Biolegend 405203 Streptavidin BV510 BD 563261 Streptavidin BV510 Biolegend 405234 Streptavidin APC Biolegend 405207 Streptavidin FITC Biolegend 405202 Streptavidin Percp.Cy5.5 Biolegend 405214

2.2.9. Staining panels The following flow cytometry staining panels were used in analysis of BM, spleen, blood and lymph node cells.

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Lineage FITC refers to a cocktail of CD4, CD8, CD11b, B220, GR-1, TER-119 antibodies and all were conjugated with FITC.

HSC depletion analysis Lin-FITC/c-kit-PE/Sca1-Percpcy5.5/Isotype-APC/CD150-PECy7 Lin-FITC/c-kit-PE/Sca1-Percpcy5.5/CD48-APC/Isotype-PECy7 Lin-FITC/c-kit-PE/Sca1-Percpcy5.5/CD48-APC/CD150-PECy7 HSPC depletion analysis Lin-FITC/c-Kit-APC/Sca1-Percpcy5.5/ Isotype-PECy7/CD34-PE/CD127 (Il7Rα) Bio-BV510 Lin-FITC/c-Kit-APC/Sca1-Percpcy5.5/ CD16/32-PECy7/Isotype-PE/CD127 (Il7Rα) Bio- BV510 Lin-FITC/c-Kit-APC/Sca1-Percpcy5.5/ CD16/32-PECy7/CD34-PE/Isotype Bio-BV510 Lin-FITC/c-Kit-APC/Sca1-Percpcy5.5/ CD16/32-PECy7/CD34-PE/CD127 (Il7Rα) Bio- BV510 Macrophages depletion analysis CD11b-Percpcy5.5/CD115-PECy7/Ly6C-PE/Ly6G-FITC/F480Bio-SA-BV510/VCAM1- BV421 Engraftment (T cells) CD45.2-Fz/ CD45.1-APC/CD4-PE/CD8-PerCP.Cy5.5 Engraftment (Antigen presenting cells) CD45.2-Fz/ CD45.1-PECy7/CD11c-PE/CD11b-PerCP.Cy5.5/ B220-APC HSPC chimerism CD45.2-APC/CD45.1-PE.Cy7/Lin-Fz/c-Kit-PE/ Sca-1-PerCP.Cy5.5 CD45 expression analysis Lin-FITC/c-kit-PE/Sca1-Percpcy5.5/CD48-APC/CD150-PECy7/SA-BV510/CD19- BV785/CD3-Pacblue Internalization analysis (HSC) Lin-FITC/c-kit-PE/Sca1-Percpcy5.5/CD48-APC/CD150-PECy7/ SA-BV510 Internalization analysis (Leukocytes) B220APC/SA-BV510/CD3-FITC/CD11b-Percpcy5.5/CD45 (30-F11) PEcy7

2.2.10. Immunotoxin preparation Biotinylated anti-CD45.2 (clone 104) was purchased from Biolegend and streptavidin-saporin was purchased from Advanced Targeting Systems (ATS), USA. Saporin is a ribosome binding

35 | P a g e and inactivating protein from Saponaria officinalis and lacks a cell entry domain and is therefore not toxic unless conjugated with a targeting monoclonal antibody or other ligand that has the capacity of receptor-mediated cell internalization. When targeted it forms a highly effective immunotoxin. Streptavidin is a tetrameric protein with molecular weight of 55 kDa and each subunit is able to bind a single biotin molecule. The bond between biotin and streptavidin is rapid and virtually irreversible. Streptavidin-saporin becomes a targeted immunotoxin when conjugated with biotinylated anti-CD45.2 antibody. CD45-SAP was prepared by combining biotinylated anti-CD45.2 (molecular weight ~145 kDa) with streptavidin-SAP (molecular weight ~124 kDa) in a 1:1 molar ratio and then this was diluted in MT-PBS. Prepared immunotoxin was mixed by inversion (not vortexing) and allowed to sit for 15 minutes at room temperature before injecting. CD45.2-SAP created fresh on the day of use at least 15 minutes prior to intravenous injection. For higher and lower dosing toxin, antibody and PBS was adjusted accordingly. Control BIG-SAP was prepared by conjugating saporin with an untargeted IgG molecule.

2.3. Animal procedure

2.3.1. Immunization

Mice were immunized with OVA/Quil-A subcutaneously at the base of the tail. For subcutaneous route, OVA/Quil-A were prepared immediately prior to immunization. 50μg of OVA was delivered per immunization per mouse. Mice were restrained and 50μL of OVA/Quil-A was injected on both left and right sides of the tail (100μL total).

2.3.2. Blood collection

For endpoint of an experiment, mice were euthanised by CO2 narcosis and blood was immediately collected by cardiac puncture into Eppendorf tubes and transferred to appropriately labelled Alsever’s solution (Sigma-Aldrich, St. Louis, MA, USA) containing FACS tube, which were then placed on ice. Alternatively, retro-orbital or submandibular bleeding was used to collect blood over multiple time-points from the same animal during the experimental procedures. Mice were lightly anaesthetised, restrained, and an anti-coagulant capillary tube was inserted laterally to the eye to collect blood from the retro-orbital venous sinus. Blood was collected into Eppendorf tube and immediately transferred to Alsever’s

36 | P a g e solution (Sigma-Aldrich, St. Louis, MA, USA) containing FACS tube. 100μL of blood was routinely collected and no sooner than 5 days after a prior bleed.

2.3.3. Lymph nodes and spleen

Lymph nodes (axillary, brachial, inguinal and mesenteric) were harvested and placed in a cold F2.5 filled 15mL Falcon tube (BD Biosciences, CA, USA). Similarly, spleens were aseptically harvested and placed in a cold F2.5 filled 15mL Falcon tube. To prepare single cell suspensions, spleen or lymph nodes were placed into a nylon mesh cell strainer (70 μm) positioned on a 50ml falcon tube (BD Biosciences, CA, USA) in a laminar airflow hood. The rubber end of a nozzle from a 5mL syringe (NIPRO, Japan) was used to manually the disrupt spleen or lymph node cells, with regular flushes with cold F2.5. Cells were then collected by centrifugation at 36.2 × g for 5 minutes at 4°C. Spleen cells were the mixed with ACK red cell lysis buffer to lyse the red blood cells (RBC), while a further washing step was done to collect spleen cells. Lymph nodes were not subjected to ACK lysis. Both spleen and lymph node cells were resuspended in cold F2.5 or different culture media. Trypan blue was used to count viable cells using a hemocytometer and cells concentrations were adjusted as required.

2.3.4. Bone marrow transplantation

Tibia and femurs were collected from mice using aseptic techniques by severing the tibia above ankle and femur at the hip. Muscles were separated from bone before placed into cold F2.5 filled 15ml/50ml falcon tube. After collecting bones from mice, set up a sterile area inside laminar airflow and bone marrow was flushed from bones using scissors, forceps and 21-25g needles based on BM cavity. Upper extremity of tibia were broke by scissors and forceps. Needles were inserted into upper margin of marrow cavity and flush marrow with 2-3ml cold F2.5. Distal end of femur was broken and marrow was flushed by inserting needles similar as tibia. Sometimes a small proportion of upper femur were needed to be cut off, if marrow was not flushing easily. BM cells were triturated by repeated passage through a needle until no visible clumps remained. Cells were transferred to a fresh sterile 50ml falcon tube and centrifuge at 36.2 × g for 5 minutes at 4°C. RBC were lysed through ACK buffer flowed by washing with F2.5. After lysis Trypan blue was used to count viable cells using a hemocytometer and cells concentrations were adjusted as required for injection. Recipient mice were exposed to 300 cGY irradiation (Gammacell Irradiator, MDS Nordion, Canada) at the Translational Research Institute (Brisbane, Queensland) or treated with different doses of CD45.2-SAP, 2B8-SAP or relevant control treatments. Donor cells were administered through

37 | P a g e intravenous tail vein injection. 1x107 and 2.5107 bulk BM cells were injected in 200μl MT- PBS.

2.4. Flow cytometry

2.4.1. Surface staining

FACS surface staining was routinely performed with cells in a single cell suspension in FACS wash. Cells were blocked for 10 minutes before antibody staining to prevent non-specific mAb binding. Cells were incubated with the appropriate biotinylated or fluorescent monoclonal antibodies at optimal staining concentrations for 30 minutes at 4°C, in the dark condition. After 30 minutes, cells were washed with FACS wash to remove un-hybridize antibody. Secondary antibodies were added when needed and left for 30 minutes at 4°C, in the dark condition. As required, propidium iodine/7AAD was added to samples immediately prior to run FACS to assist with dead cell exclusion. For enumeration of cell populations 20μl (unless stated otherwise) of flow count beads (Cat 7547053; Beckman Coulter, FL, USA) were added to a known portion of the total tissue sample immediately prior to FACS run. Flow cytometry was performed on a BD Fortessa X-20 with cells of interest identified based on forward scatter (FSC) and side scatter (SSC) profiles. FACS Diva (BD Biosciences, CA, USA) software was used to analyse data.

2.4.2. Pulse-chase assay

Bone marrow and spleen were harvested and single cell suspension was prepared through standard procedure. Prepared single cells were resuspended on X-VIVO 15 Serum-free Hematopoietic Cell Medium. After resuspension cells were adjusted to 4x106 cell/ml and biotinylated anti-CD45.2 (clone 104) or biotinylated anti-CD45 (clone 30-F11) was added at a concentration of 2.5ug/ml. Antibody stained cells were plated into 48 wells culture plate and incubated (370C) for 0-240 minutes. After culture, cells were transferred to appropriate FACS tube, washed and stained with surface markers for 30 minutes. Flow cytometry was performed on a BD Fortessa X-20 with cells of interest identified based on FSC and SSC profiles. FACS Diva (BD Biosciences, CA, USA) software was used to analyse data.

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2.5. ELISPOT

Polyvinylidene difluoride ELISPOT plates were purchased from Millipore (Massachusetts, USA) and were pre-washed with 70% ethanol and 1X MT-PBS. IFN-γ capture antibody (5µg/ml) was added to the plates and were incubated overnight wrapped in foil at 4°C refrigerator. The following day plates were removed from the freeze and washed with 1X MT- PBS (three times) and blocked with R5 medium (RPMI-1640 5% FCS) for 1 hours at room temperature. Spleen cells were prepared by using standard protocol and resuspended in R5 medium and adjusted at 7.5 x105 or 1.25x106. Then 200μl R5 was added to each well. Cells were left untreated or stimulated with anti-CD3 (0.1μg/ml) or OVA257-264 (0.5μg/ml). Plates 0 were incubated for 2 days at 37 C 5% CO2 in a damp chamber. 2 days later, plates were removed and washed with PBS/0.05% Tween 20 and 1X MT-PBS and IFN-γ detection antibody (2µg/ml) was added. Plates were again incubated overnight wrapped in foil at 4°C. Plates were washed with PBS/0.05% Tween 20 and 1X MT-PBS and sterptavidin-HRP detection enzyme (DAKO) was then added and plates left to incubate at room temperature for 1 hour. After 1 hour the detection enzyme was washed off and AEC substrate (Calibochem, CA, USA) was added. Development was observed under a dissecting microscope and staining was stopped immediately after spot development. Plates were stored in dark condition at room temperature until dry. Spots were detected by using an immuno-spot plate reader (AID GmbH, Strassburg, Germany).

Table 2.5.: List of antibody used in ELISPOT Antibody Fluorophore Host/Isotype Clone Supplier Catalogue IFN- γ AN18 eBioscience 14-7313-85 IFN- γ R4-6A2 eBioscience 13-7312-85

2.6. Histology

2.6.1. Bone tissue collection and fixation

Bones were dissected mostly left hind-limb by cutting the limb just below the hip joint and above the ankle joint (thereby exposing the marrow cavity of both the femur & the tibia). Skin, the majority of muscle, ligaments and tendons and connective tissues were removed and immediately placed in 4% PFA solution for fixation. Tissue was kept on 4% PFA for 72 hours at 40C. Extended periods of time was avoided as it could have damaged tissue and compromise antigen detection.

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2.6.2. Bone decalcification

4% PFA fixed bones were washed with 1X MT-PBS to remove excess paraformaldehyde and then transferred to 14% EDTA solution for decalcification. EDTA solution was changed every week and decalcification took 2 weeks unless stated otherwise. To confirm decalcification 27G needle was inserted into the bone at strategic points to determine any resistance. Once decalcified bones were processed with paraffin and embedded.

2.6.3. Paraffin processing and embedding

Decalcified tissue was dehydrated through a series of graded ethanol baths to displace the H2O and then infiltrated with wax. The infiltrated tissues were then embedded into wax blocks.

2.6.4. Tissue sectioning and staining

Five micron serial sections were cut from the blocks by using a microtome and placed on labelled slides and left for drying overnight at 370C. Then slides were transferred to fridge and were dewaxed in xylene substitute and rehydrated in graded ethanol washes. Tissues were stained with F4/80 and its isotype and counterstain with hematoxylin. Stained slides were scanned at 20X magnification by an Olympus VS120 (Olympus, Tokyo, Japan). Images were prepared by using OlyVIA version 2.6 (Olympus Soft imaging Solution, USA). All sections were analyzed using Visiopharm (Hoersholm, Denmark).

2.7. Statistical analysis

Statistical analyses were performed and all the graphs were generated using GraphPad prism version 7. Student’s t-test was used to compare means between groups, or one-way ANOVA with Tukey’s multiple comparison post-test for multiple comparisons of means. Details of statistical comparisons is listed in each figure legend. Significance was considered for the following p-values: < 0.0001 (****), < 0.001 (***), < 0.01 (**), < 0.05 (*).

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Chapter 3 A leukocyte-targeting immunotoxin preferentially depletes LT-HSC when used at low doses

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3.1. Introduction

Inducing antigen-specific T cell tolerance by transplanting bone marrow (BM) or haematopoietic stem cells (HSC) genetically engineered to encode antigen targeted to dendritic cells (DC) or other antigen presenting cells (APC) may be an effective procedure for autoimmune disease treatment [4, 103, 243]. To achieve long-term tolerance it is likely that gene-engineered antigen expression needs to be life-long. Consequently, long-term repopulating hematopoietic stem cell (LT-HSC) engraftment is required to ensure a continuous supply of tolerogenic antigen-expressing DC/APC in the periphery [244]. Stable HSC engraftment is attainable through pre-transfer myeloablative or non-myeloablative recipient conditioning. But these types of conditioning deplete recipient immunity and have a range of toxic side effects. In recent years, non-myeloablative conditioning such as low-dose irradiation has been introduced for hematologic malignancies to reduce toxicity and transplant-associated risk [245, 246]. But still these conditioning approaches remain genotoxic, leading to cell and DNA damage [247]. So, developing a non-genotoxic conditioning regimen, which avoids DNA mutagenesis, to promote engraftment of transferred HSC, is clinically important because it would be safer and, possibly, more targeted than currently used genotoxic conditioning.

Using biologics such as antibodies and immunotoxins which can specifically target hematopoietic cells and spare non-hematopoietic cells could be a promising approach to improve recipient conditioning. Recently, some approaches to achieving high levels of donor HSC engraftment without the need for irradiation or chemotherapeutics have emerged. For example, administration of an anti-c-kit antibody (ACK2) depletes a significant number of HSC from BM niches and permits successful donor HSC engraftment in immunodeficient mice [226, 248]. Chhabra et al., [227] showed that host HSC depletion is reliant on Fc-mediated antibody effector functions, and that augmenting effector function by the blockade of integrin associated protein (CD47) enhances the depleting effect of ACK2 in immunocompetent mice. This combined approach of ACK2 with CD47 blockade or ACK2 with low dose irradiation can lead to the elimination of over 99% of recipient HSC and permits a high level of transferred donor HSC engraftment which achieves donor-derived multilineage leukocyte development and tolerance but clinical translation of this approach is challenging [226, 227, 249, 250]. CD45-SAP, an internalizing immunotoxin targets hematopoietic-cell restricted CD45 and depletes >98% of HSC which permits >90% donor HSC engraftment after transfer, when used at high dose [229]. However, at high doses CD45-SAP is immunodepleting. To interrogate

42 | P a g e clinical applicability that might be achieved by avoiding immunodepletion, and, as low-levels of engraftment are suitable to generate T-cell tolerance, it is important to test whether use of lower doses of CD45-SAP that might preserve or partially preserve immunity would be suitable as a low toxicity conditioning regimen.

The hypothesis for this chapter is low-dose CD45.2-SAP depletes LT-HSC from BM without largely depleting peripheral leukocytes and creates space for donor exogenous BM cells. If this is the case, treatment with low-dose CD45.2-SAP might subsequently promote donor engraftment. In this chapter, I describe that titrating the dose of CD45.2-SAP gives a dose- dependent depletion of LT- HSC. Using reduced doses of CD45.2-SAP might also have the additional advantage that depletion of leukocytes may be minimised. To strengthen the conclusion, a competitive BM transplantation was performed to verify the depletion of LT- HSC by CD45.2-SAP.

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3.2. Results

3.2.1. CD45.2-SAP depletes long-term repopulating HSC (LT-HSC) in a dose dependent manner

Use of an anti-CD45.2-immunotoxin (CD45.2-SAP) has been shown to lead to a high level of LT-HSC depletion enabling high levels of engraftment of transferred donor LT-HSC in treated mice. However, a side effect of this was depletion of mature peripheral leukocytes [229]. As modest or may be low levels of gene-engineered donor LT-HSC engraftment may be sufficient to induce T-cell tolerance, so a moderate level of recipient LT-HSC depletion might be enough. Therefore, experiments were designed to determine whether reducing leukocyte depletion through titrating CD45.2-SAP treatment could lead to sufficient LT-HSC depletion. Therefore, the dose-dependent depleting effects of CD45.2-SAP was assessed. For these experiments the aim was to determine the depletion effects on LT-HSC, multipotent progenitor (MPP), common lymphoid progenitor (CLP), common myeloid progenitor (CMP), myeloid progenitor (MP), granulocyte myeloid progenitor (GMP) and myeloid erythroid progenitor (MEP) content of BM as well as on mature leukocytes such as CD8+ T cells, CD4+ T cells, B cells, myeloid cells and DC.

CD45.2-SAP were prepared by conjugating biotinylated anti-CD45.2 with streptavidin-saporin and injected as graded doses. A single dose of a control untargeted immunotoxin made with an irrelevant biotinylated antibody (BIgG-SAP, prepared and provided as a control by Advanced Targeting System, San Diego, CA, USA). CD45.2-SAP, BIgG-SAP and PBS were administered by i.v. injection. Untreated mice were also included as controls for depletion. Seven days after CD45.2-SAP administration, spleen and BM were collected for analysis (Figure 3.1A). Flow cytometry was used to determine LT-HSC (Lin-c-kit+Sca1+CD48- CD150+), MPP (Lin-c-kit+Sca1-CD48-CD150-), CLP (Lin-c-kit+Sca1+CD127+), CMP (Lin-c- kit+Sca1-CD127-CD34+CD16/32-), MP (Lin-c-kit+Sca1-CD127-), GMP (Lin-c-kit+Sca1- CD127-CD34+CD16/32+) and MEP (Lin-c-kit+Sca1-CD127-CD34-CD16/32-) content of BM (Figure 3.1A, 3.2A, 3.3A) and a bead-based approach used to determine their absolute numbers. This assay identified that a single dose of CD45.2-SAP had potent LT-HSC-depleting capability depending on the dose administered. BM from untreated and PBS-treated mice contained approximately 4×103 to 6×103 LT-HSC in the BM whereas CD45.2-SAP and BIgG- SAP treated mice showed a significantly reduced number of LT-HSC in BM (Figure 3.1C).

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Similar to a previous report [229], a high dose of CD45.2-SAP (3 mg/kg) depleted on average 90% of LT-HSC (Figure 3.1D). As the dose of CD45.2-SAP was reduced, the extent of LT- HSC depletion was also reduced. A moderate dose of 2 mg/kg or 1 mg/kg CD45.2-SAP depleted approximately 70% and 50% LT-HSC respectively. Notably, with a low-dose of CD45.2-SAP (0.5 mg/kg) approximately 45% of LT-HSC were depleted (Figure 3.1D). The depletion trend of CD45.2-SAP on MPP and HSPC was similar to that for LT-HSC. At 3 mg/kg CD45.2-SAP, 90% of MPP and 50% of HSPC were depleted whereas 1 mg/kg and 2 mg/kg depleted 60-70% of MPP and 35-40% of HSPC respectively. At the 0.5 mg/kg dose depletion was limited, with only 40-45% of MPP and 20-25% of HSPC (Figure 3.1 E-F) being depleted. These data indicate that in vivo administration of CD45.2–SAP efficiently depletes ‘phenotypic’ LT-HSC from the mouse BM and LT-HSC are most affected among early or primitive hematopoietic stem and progenitor cells. The depletion effect was also dose dependent.

To determine the effect of CD45.2-SAP on more committed hematopoietic progenitors, these cells were also analysed by flow cytometry (Figure 3.2 A and 3.3A). As expected, a higher CD45.2-SAP dose (3 mg/kg) displayed significant depleting activity on hematopoietic progenitors except CLP (Figure 3.2B) and GMP (Figure 3.3D). It depleted approximately 55% of MP (Figure 3.2E), 80% of MEP (Figure 3.3B) and 40% of CMP (Figure 3.3C). But lower doses such as 2 mg/kg, 1 mg/kg or 0.5mg/kg showed no significant depletion effects on lineage-committed hematopoietic progenitors except GMP (Figure 3.3D). These data suggest low and moderate CD45.2-SAP doses do not deplete different committed hematopoietic progenitors, therefore higher doses of immunotoxins are required to do so.

As peripheral leukocytes such as CD8+ T cells, CD4+ T cells and B cells express CD45 so it was expected they would also be depleted by CD45.2-SAP treatment. To determine whether CD45.2-SAP depleted peripheral leukocytes, spleen was harvested 7 days after CD45.2-SAP administration. Assessing peripheral leukocytes in CD45.2-SAP-treated spleen revealed a significant depletion of CD8+ T cells (50-70%), CD4+ T cells (60-70%) and CD45R+ B cells (20-40%) at the different higher doses of CD45.2-SAP such as 1 mg/kg, 2 mg/kg and 3 mg/kg. In contrast, a lower dose (0.5 mg/kg) showed very little depletion (~15%) of either CD8+ or CD4+ T cells (Figure 3.4A-B) and no depletion of B cells (Figure 3.4C). Surprisingly CD45.2-SAP did not show any depletion effect on CD11c+ DC (Figure 3.4D) and myeloid cells (Figure 3.4E), but actually increased their absolute numbers.

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Taken together, these data show that in vivo administration of a low-dose of CD45.2-SAP (0.5 mg/kg) efficiently depletes a significant proportion of phenotypic LT-HSC but with substantial preservation of differentiated hematopoietic progenitors and peripheral leukocytes, compared to higher doses.

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CD45.2-SAP A Single dose, IV C57BL/6

Day 0 Day 7 Assess LT-HSC and HSPC depletion in BM and leukocyte depletion in Spleen

B

Cy5.5

-

PerCP Lin Lin FITC

Sca1 APC

-

c-kit-PE c-kit-PE CD48

CD150-PE-Cy7 C D

**** **** ns **** 8 ns 100 * ns ns 75 ns 6 ns ** *** 50 ns 4 ns 25 2 0

0 -25 l i S P .5 1 2 3 depletion control) of LT-HSC (% S P .5 1 2 3 N B A 0 A 0

LT-HSC Absolute Numbers (X10^-3) Numbers Absolute LT-HSC B P -S P -S G G Ig Ig B CD45.2-SAP dose (mg/kg) B CD45.2-SAP dose (mg/kg)

E F

**** **** ns 100 **** ns ns 100 *** 75 * *** ns 75 ns ns 50 50 ** 25 25 0 0

-25 -25

HSPCcontrol) depletion of (%

MPP depletion (% of control) of (% depletion MPP S P .5 1 2 3 P .5 1 2 3 A 0 S A 0 B S B S P - P - G G g Ig I B CD45.2-SAP dose (mg/kg) B CD45.2-SAP dose (mg/kg)

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Figure 3.1. CD45.2-SAP depletes haematopoietic stem cells and multipotent progenitor cells. (A) Schematic experimental outline for assessing depletion of LT-HSC in C57BL/6 mice. Graded doses of CD45.2-SAP (0.5 mg/kg, 1 mg/kg, 2 mg/kg, and 3 mg/kg) were injected i.v. and depletion was analysed 7 days later. (B) Flow cytometric analysis was performed to analyse LT-HSC number in BM (C), LT-HSC (=Lin-c-kit+Sca1+CD48-CD150+), (D) MPP (=Lin-c- kit+Sca1+CD48-CD150-) (E) and HSPC (=Lin-c-kit+) (F) absolute number and depletion using the gating strategy shown. Depletion percentage was calculated as a proportion (% of control) of the average of the absolute number of cells harvested from untreated mice within each experiment. Data are pooled from 4 individual experiments and show individual mice with mean ± SD, n=4-9 mice per group. Statistical significance determined by one-way ANOVA, ****p<0.0001, ***p<0.001, **p<0.01, *p<0.05 and ns= no significance.

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A CLP

MP

FITC -

Lin

CLP

c-kit-APC PerCPCy5.5 - MP

Sca1

CD127-BV510

B C

100 500 ns ns 400 50

300 0 200 -50 100 -100

0 depletion CLP control) of (% l P 5 1 2 3

CLP Absolute Numbers (X10^-3) Numbers Absolute CLP . i S P .5 1 2 3 S A 0 N B A 0 B S P -S P - G G Ig Ig B CD45.2-SAP dose (mg/kg) B CD45.2-SAP dose (mg/kg)

D E

*** 3000 ns 100 *

ns 2000 50

1000 0

0 -50

-1000 depletion MP control) of (% -100

MP Absolute Numbers (X10^-3) l i S P .5 1 2 3 S P .5 1 2 3 N B A 0 B A 0 P -S P -S G G Ig Ig B CD45.2-SAP dose (mg/kg) B CD45.2-SAP dose (mg/kg)

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Figure 3.2. Low-dose CD45.2-SAP (0.5 mg/kg) has limited depletion effect on common lymphoid progenitors and myeloid progenitors. Different doses of CD45.2-SAP were injected intravenously and depletion effects on common lymphoid (CLP) and myeloid (MP) progenitors were analysed 7 days after CD45.2-SAP administration by flow cytometry and shown gating strategy was used to define CLP and MP populations (A). Dose-dependent effects of CD45.2-SAP on (B, C) CLP (Lin-c-kit+Sca1+CD127+), (D, E) MP (Lin-c-kit+Sca1-CD127) were assessed. Depletion percentage were calculated as a proportion (%) of the average number of cells harvested from untreated mice (% of control mice). Data are pooled from 4 individual experiments and represent the individual mice with mean ± SD, n=4-9 mice per group. Statistical significance determined by one-way ANOVA, ****p<0.0001, ***p<0.001, **p<0.01, *p<0.05 and ns= no significance.

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A GMP

CMP

MEP

FITC

-

PerCPCy5.5 Lin

-

Sca1 GMP

PECy7 CMP c-kit-APC CD127-BV510 -

MEP D16/32

C

CD34-PE

B C

**** * *** * ns **** 100 100 * ns ns * 50 50 * ns 0 0

-50 -50

-100 -100

MEP depletion (% of control) of (% depletion MEP CMP control) depletionCMP of (% 5 1 2 3 S P . S P .5 1 2 3 B A 0 B A 0 P -S P -S G G Ig Ig B CD45.2-SAP dose (mg/kg) B CD45.2-SAP dose (mg/kg)

D

100 * ns ** 50 ns 0

-50

-100 GMP depletion (% of control) of (% depletion GMP S P .5 1 2 3 B A 0 P -S G Ig B CD45.2-SAP dose (mg/kg)

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Figure 3.3. Low-dose CD45.2-SAP (0.5 mg/kg) does not deplete lineage-committed hematopoietic progenitors. Different doses of CD45.2-SAP were injected intravenously and depletion effects on different hematopoietic progenitor cells were analysed 7 days after CD45.2-SAP administration by flow cytometry and the gating strategy shown was used to define haematopoietic progenitor populations (A). MEP (=Lin-c-kit+Sca1-CD127-CD34- CD16/32-)(B), CMP (Lin-c-kit+Sca1-CD127-CD34+CD16/32-) (C), GMP (Lin-c-kit+Sca1- CD127-CD34+CD16/32+) (D) were assessed. Depletion percentage mice (% of control) was calculated as a proportion (%) of the average number of cells harvested from untreated mice. Data are pooled from 4 individual experiments and represent individual mice with mean ± SD, n=4-7 mice per group. Statistical significance determined by one-way ANOVA, ****p<0.0001, ***p<0.001, **p<0.01, *p<0.05 and ns= no significance.

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A B

**** **** 100 *** 100 **** ns

* *** ns 50 50 ** ns ns 0 0

-50 -50 CD4+ depletion control) CD4+ of (% CD8+ depletion control) CD8+ of (% S P .5 1 2 3 S P .5 1 2 3 B A 0 B A 0 P -S P -S G G Ig Ig B CD45.2-SAP dose (mg/kg) B CD45.2-SAP dose (mg/kg)

C D

* ** ** 100 * 100 * ns 80 * 50 ns 60 ns 40 0 ns 20 -50 0

-20 control) of (% depletion DC -100 B cell B depletion control) of (% S P .5 1 2 3 S P .5 1 2 3 B A 0 B A 0 P -S P -S G G Ig I B CD45.2-SAP dose (mg/kg) B CD45.2-SAP dose (mg/kg)

E

200 * 100 ns ns 0

-100

-200

-300

P .5 1 2 3 S A 0 Myeloidcell depletion of (% control) B P -S G Ig B CD45.2-SAP dose (mg/kg)

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Figure 3.4. Low-dose CD45.2-SAP (0.5 mg/kg) has limited depletion effect on T and B cells but myeloid cells and DC increase after CD45.2-SAP treatment. Different doses of CD45.2-SAP were injected intravenously and spleen was harvested 7 days after CD45.2-SAP administration to analyse the depletion effects CD45.2-SAP on different leukocytes. Relative levels of peripheral CD8+ (A), CD4+ (B), B cells (C), DC (D) and myeloid cells (E) post CD45.2-SAP treatment were determined. Depletion percentage mice (% of control) was calculated as a proportion (%) of the average number of cells harvested from untreated mice. Data are pooled from 4 individual experiments and represent the individual mice with mean ± SD, n=4-7 mice per group. Statistical significance determined by one-way ANOVA, ****p<0.0001, ***p<0.001, **p<0.01, *p<0.05 and ns= no significance.

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3.2.2. Low-dose (0.5mg/kg) CD45.2-SAP depletes LT-HSC 2 days after administration

Accumulated data show low-dose CD45.2-SAP (0.5mg/kg) gives a moderate level of LT-HSC depletion with mild depletion of peripheral leukocytes 7 days after administration (Figure 3.1 and 3.4). However, CD45.2-SAP might also deplete target cells quickly after administration and this may not be detected by analysis at day 7. Experiments were therefore designed to define the depletion effects of low-dose CD45.2-SAP (0.5mg/kg) on peripheral leukocytes, LT-HSC and different committed progenitors at day 2 post-CD45.2-SAP administration. CD45.2-SAP (0.5mg/kg), BIgG-SAP and PBS were administered by i.v. injection. Untreated mice were used as no depletion controls. BM and spleens were collected 2 days after CD45.2- SAP administration and used to determine the deletion effect on leukocytes, LT-HSC and progenitors by flow cytometric analysis (Figure 3.5A).

This experiment identified that low-dose CD45.2-SAP (0.5mg/kg) had depleted a significant proportion of LT-HSC within 2 days. Flow cytometric analysis showed approximately 5-6×103 LT-HSC in BM from untreated and PBS treated mice whereas the number was reduced in BIgG-SAP and CD45.2-SAP treated groups (Figure 3.5B). Similar to day 7 post-CD45.2-SAP administration, 0.5mg/kg CD45.2-SAP depleted approximately 40-50% of LT-HSC (Figure 3.5C) at day 2 post CD45.2-SAP administration. Interestingly, approximately 50% of HSPC were depleted (Figure 3.5 D, E) at day 2 post CD45.2-SAP administration which was twice the depletion effect (~25%) observed on day 7 (Figure 3.1F). These data indicate that at day 2, the observed effects of CD45.2-SAP differ to day 7 and show that HSPC whilst depleted 2 days after CD45.2-SAP administration, may have recovered by day 7. The depletion effect on LT-HSC at day 2 was similar as that observed at day 7.

To determine the depletion effect of CD45.2-SAP (0.5mg/kg) on different lineage committed hematopoietic progenitors at day 2, these cells were analysed by flow cytometry. As expected, CD45.2-SAP (0.5mg/kg) showed a substantial depleting effect on hematopoietic progenitors except common lymphoid progenitor cells (Figure 3.6A-F). It depleted approximately 60% of MPP (Figure 3.6A), 45% of MP (Figure 3.6C), 50% of MEP (Figure 3.6D), 65% of CMP (Figure 3.6E) and 45% of GMP (Figure 3.6F). BIgG-SAP also showed moderate depletion activity on MPP approximately 30% (Figure 3.6A), 10% on MP (Figure 3.6C), 15% on MEP (Figure 3.6D), 40% on CMP (Figure 3.6E) and ~20% on GMP (Figure 3.6F). PBS showed no considerable effect on these hematopoietic progenitor cells. These data suggest low-dose

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CD45.2-SAP (0.5mg/kg) depletes a significant proportion of different committed hematopoietic progenitor cells at day 2 post administration.

To determine the depletion effects of CD45.2-SAP on peripheral leukocytes, spleens were collected and analysed by flow cytometry. Assessing peripheral leukocytes revealed approximately 50% depletion of CD8+ T-cells (Figure 3.7A) and 40% depletion of CD4+ T- cells (Figure 3.7B). Myeloid cell number was increased by approximately 90% at day 2 post administration (Figure 3.7C) whereas B cells (Figure 3.7D) and DC (Figure 3.7E) were unaffected. The depletion effect on CD8+ and CD4+ T-cell was much higher at day 2 than day 7 post-CD45.2-SAP administration. Myeloid cell numbers also increased more at day 2 than day 7 post-CD45.2-SAP injection.

To further analyse the effect of CD45.2-SAP (0.5mg/kg) on myeloid cell populations such as macrophage, and , BM was assessed by flow cytometry. Macrophage number was increased approximately 50% (Figure 3.8A) by CD45.2-SAP treatment whereas granulocyte number were increased by 35% (Figure 3.8B) and monocytes by 10% (Figure 3.8C). Small number of mice were analysed to understand the effects and data are indicative only. The increase in the number of macrophages was verified by immunohistochemistry. Bones were harvested, processed and stained with anti-F4/80. Stained samples were scanned with Olympus VS120 and analysed on Visiopharm software. Immunohistochemistry assay confirmed the increase in macrophages by CD45.2-SAP treatment (Figure 3.9A-B). As low- dose CD45.2-SAP (0.5mg/kg) depleted large numbers of LT-HSC and HSPC at day 2 post administration, the BM may be expected to contain a large number of dead cells after CD45.2- SAP treatment.

Taken together, these data suggest that in vivo administration of low-dose CD45.2-SAP (0.5mg/kg) depletes a significant proportion of LT-HSC and HSPC at day 2 post administration. While the extent of LT-HSC depletion was similar at days 2 and 7 after CD45.2-SAP administration, HSPC were more substantially reduced at day 2 than day 7.

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A

B C

**** **** 100 10 ** ns ns 8 75 *** ** 6 50

4 25

2 0

0 -25

il P P depletion control) of LT-HSC (% S P P N S A A LT-HSC absolute numbers (X10^3) numbers absolute LT-HSC B A A B S S P -S -S P - - G .2 G .2 g 5 g 5 I 4 I 4 B D B D C C

D E

*** ** 2500 100 * ns ** * 2000 75 ns 1500 50

1000 25

500 0

0 -25 il HSPCcontrol) depletion of (% P P HSPCabsolute numbers(X10^3) S P P S N B A A B A A P -S -S P -S -S G .2 G .2 g 5 g 5 I 4 I 4 B D B D C C

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Figure 3.5. Low-dose (0.5mg/kg) CD45.2-SAP partially depletes LT-HSC and HSPC at day 2. (A) Schematic experimental outline for assessing depletion of LT-HSC in C57BL/6 mice. Low-dose CD45.2-SAP (0.5 mg/kg) were injected i.v. and depletion was analysed 2 days later. Flow cytometric analysis was done to analyse LT-HSC absolute number in BM (B), LT- HSC depletion (C) HSPC number (D) and HSPC depletion (E). Depletion percentage was calculated as a proportion (%) of the average of absolute number of cells harvested from untreated mice (% of control). Data are pooled from 4 individual experiments and show individual mice with the mean ± SD, n=6-8 mice per group. Statistical significance determined by one-way ANOVA, ****p<0.0001, ***p<0.001, **p<0.01, *p<0.05 and ns= no significance.

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A B

**** **** 100 200 ns ** * 75 *** 0 50 -200 25 -400 0 -600 -25

-800

MPPdepletion control) of (% CLP depletion CLP control) of (%

S P P P A A S P A B S S B A S P - - P -S - G .2 .2 g 5 G 5 I 4 Ig 4 B D B D C C D C

* **** ns 100 100 ***

ns

50 50 ns

0 0

-50

MP depletion MP control) of (% -50 MEPdepletion control) of (%

S P P S P P B A A B A A P -S -S P -S -S G .2 G .2 g 5 g 5 I 4 I 4 B D B D C C

E F

**** ** ** ns 100 100 ns * 50 50

0 0

-50 -50

GMP depletion (% of control) of (% depletion GMP CMP control) depletionCMP of (%

S P P S P P B A A B A A P -S -S P -S -S G .2 G .2 g 5 g 5 I 4 I 4 B D B D C C

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Figure 3.6. Low-dose CD45.2-SAP (0.5 mg/kg) depletes a significant proportion of lineage-committed progenitors 2 days after administration. 0.5mg/kg of CD45.2-SAP were injected intravenously and depletion effects on different committed progenitors were analysed 2 days after CD45.2-SAP administration by flow cytometry and depletion effects of CD45.2- SAP on MPP (A), CLP (B), MP (C), MEP (D), CMP (E) and GMP (F) were assessed. Depletion percentage were calculated as a proportion (%) of the average number of cells harvested from untreated mice (% of control mice). Data are pooled from 4 individual experiments and represent individual mice the mean ± SD, n=6-8 mice per group. Statistical significance determined by one-way ANOVA, ****p<0.0001, ***p<0.001, **p<0.01, *p<0.05 and ns= no significance.

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A B

*** * 100 100 * ns ns 50 50

0 0

-50 -50 CD4+ depletion CD4+ control) of (% CD8+ depletion control) CD8+ of (% S P P S P P B A A B A A P -S -S P -S -S G .2 G .2 Ig 5 g 5 B 4 I 4 D B D C C D C

100 100 ns 50 0 0

-100 -50

-200 -100

cell B depletion control) of (% P P P P S S A A B A A Myeloid celldepletion control) of (% B P -S -S P -S -S 2 G .2 G . g 5 Ig 5 I 4 4 B D B D C C

E

ns 100

* 50 **

0

-50

DC depletion control) DC of (% -100

S P P B A A P -S -S G .2 g 5 I 4 B D C

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Figure 3.7. Low-dose CD45.2-SAP (0.5 mg/kg) depletes a proportion of CD8+ and CD4+ T-cells but not B cells and myeloid cells. 0.5mg/kg of CD45.2-SAP were injected intravenously and spleen was harvested 2 days after CD45.2-SAP administration to analyse the depletion effects of CD45.2-SAP on different leukocytes. Relative levels of peripheral CD8+ (A), CD4+ (B), myeloid cells (C), B cells (D) and dendritic cells (E) post CD45.2-SAP treatment were determined. Data are pooled from 4 individual experiments and represent the individual mice with mean ± SD, n=6-8 mice per group. Statistical significance determined by one-way ANOVA, ****p<0.0001, ***p<0.001, **p<0.01, *p<0.05 and ns= no significance.

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A B

100 100

50 50

0 0

-50 -50

-100 -100

-150 -150 P P P P S A A S A A B B S S Macrophage depletion (% of control) of (% depletion Macrophage P -S S P - -

- control) of Granulocytes(% depletion G .2 G .2 g 5 g 5 I 4 I 4 B D B D C C

C

100

50

0

-50

-100 P P S A A

Monocytesdepletion control) of (% B P -S -S G .2 g 5 I 4 B D C

Figure 3.8. Macrophage number increases after low-dose CD45.2-SAP (0.5 mg/kg) administration. CD45.2-SAP (0.5mg/kg) was injected intravenously and BM was harvested 2 days after CD45.2-SAP administration to analyse the depletion effects of CD45.2-SAP on macrophage (A), granulocytes (B) and monocytes (C). Data are pooled from 2 individual experiments and show values for individual mice with mean ± SD, n=2-4 mice per group. Depletion percentage were calculated as a proportion (%) of the average number of cells harvested from untreated mice (% of control mice).

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A

20µm 20µm 20µm

Isotype Untreated PBS treated

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Figure 3.9. Immunohistochemistry confirms CD45.2-SAP (0.5 mg/kg) induces macrophage populations. CD45.2-SAP (0.5mg/kg) was injected intravenously and BM was harvested 2 days after CD45.2-SAP. Five micron serial section were cut from decalcified bone and stained with anti-F4/80 antibody or appropriate isotype control and counterstained with hematoxylin. Stained samples were scanned by an Olympus VS120 (Olympus, Tokyo, Japan). Images were prepared using OlyVIA and analyzed with Visiopharm software. Representatives of immunohistochemistry staining using anti-F4/80 in serial BM sections from PBS, BIgG- SAP and CD45.2-SAP treated mice are shown (A), arrows shows the cells stained with F4/80 (brown colored). Percent of F4/80 staining was quantified using manually established colour threshold using Visiopharm software (B). Data represent single experiment and the mean ± SD, n=2 mice per group. Scale bar in images represent 20µm.

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3.2.3. Mechanisms of CD45.2-SAP targeting LT-HSC

To understand the mechanism of CD45.2-SAP targeting of LT-HSC, a series of ex vivo CD45 expression, blocking and internalization assays were performed using BM and spleen cells. To determine why low-dose CD45.2-SAP preferentially targets LT-HSC and MPP (Figure 3.10A) over peripheral leukocytes (Figure 3.10B), a CD45 expression assay was performed. BM and spleen cells were harvested and mixed in an equal ratio and then stained with different concentrations of biotinylated anti-CD45.2 antibody. After incubation, the antibody binding was detected using streptavidin-BV510 and flow cytometric analysis. As expected, anti- CD45.2 antibody bound to LT-HSC and MPP from BM (Figure 3.10A) and peripheral leukocytes such as B cell and myeloid cells from spleen (Figure 3.10B) in a dose-dependent manner. The pattern of CD45.2 staining for LT-HSC and B cells was compared and an EC50 value was calculated. Comparison of EC50 values shows that B cells require a higher concentration of antibody for a half-maximal staining than LT-HSC and anti-CD45 staining intensity was higher on B cells than LT-HSC. From this it was concluded that both LT-HSC and peripheral leukocytes express significant amount of CD45.2 and B cells express more CD45.2 than LT-HSC (Figure 3.10C).

To determine whether CD45.2-SAP depletes cells through receptor-mediated internalization, a blocking assay was performed where spleen cells were stained and incubated with different concentrations of biotinylated anti-CD45.2 (clone 104) and half an hour later cells were re- stained with anti-CD45-PECy7 either with clone 104 or clone 30-F11. As the concentration of biotinylated anti-CD45 (clone 104) was increased, the extent of binding of the other anti-CD45- PECy7 (clone 104) was reduced both for B cells (Figure 3.11A) and myeloid cells (Figure 3.11B). In contrast, staining with biotinylated anti-CD45 (clone 104) did not block the staining with anti-CD45-PECy7 (clone 30-F11) both for B cells (Figure 3.11C) and myeloid cells (Figure 3.11D). These data indicate that binding of one antibody to the cell surface receptor blocks the binding of the same clone but does not block the alternative clone tested. In all depletion experiments mice were treated with CD45.2-SAP which was prepared by conjugating SAP with anti-CD45.2 (clone 104) and after its treatment cells were collected and re-stained with clone 104. So the possible explanation of how CD45.2-SAP enters into the cell is receptor mediated internalization because if it is not the case then un-internalized CD45.2-SAP persist on the cell surface and blocks the staining of another same clone anti-CD45.

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To define how CD45.2-SAP depletes LT-HSC and peripheral leukocytes, a CD45 internalization assay was performed where BM and spleen cells were cultured in X-VIVO serum-free medium. Anti-CD45 antibody was added to the cells, washed to remove unbound antibody and cultured at 370C for 0 to 240 minutes. To prevent CD45.2 internalization some cells were incubated at 40C and used as negative control. After incubation for 0-240 minutes, cells were re-stained with SA-BV510-secondary antibody and surface bound anti-CD45.2 antibody by T cells, B cells and LT-HSC was measured using flow cytometric analysis. The staining intensity of surface bound anti-CD45.2 was maximal at 0 minutes and decreased with time. The bound anti-CD45.2 significantly decreased over the time (Figure 3.12A) and was reduced to over 50% after 240 minutes, indicating partial internalization of anti-CD45.2. The loss of surface binding was not detectable at 30 minutes but after 30 minutes bound anti- CD45.2 to membrane decreased slowly until 120 minutes and then reduced faster and dropped to approximately 75% in T cells, 65% in B cells and 50% in LT-HSC (Figure 3.12B-D).

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A B BM Spleen 20000 20000 EC50=1.444 LT-HSC EC50=1.385 B cells 16000 EC50=1.499 MPP 16000 EC50=1.411 Myeloid cell

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BM+Spleen 20000 EC50=0.990 LT-HSC 16000 EC50=1.125 B cells **** **** 12000 ****

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0 0 8 3 6 2 0 0 0 0 0 7 5 0 1 5 0 0 0 .0 .0 .1 .3 .6 .2 .5 .0 .0 0 0 0 0 0 1 2 5 0 1 Antibody concentration (µg/ml)

Figure 3.10. Leukocytes express more CD45 than LT-HSC. Samples were prepared by harvesting BM alone, spleen alone and mixing BM and spleen cells and also stained and incubated with different anti-CD45.2 antibody concentrations. BM and spleen only cells were also used. CD45.2 staining is antibody concentration dependent and LT-HSC and MPP (A) stained less than leukocytes such as B cells and myeloid cells (B) in BM and spleen cells respectively. The staining intensity was higher in B cells than LT-HSC in the mix sample of BM and spleen (C). Data are pooled from three individual experiments with mean ± SD. Statistical significance determined by two-way ANOVA, ****p<0.0001, ***p<0.001, **p<0.01, *p<0.05 and ns= no significance.

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A B

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Figure 3.11. Binding of anti-CD45.2 clone blocks the binding of the same clone but does not block a different clone. Spleen cells were harvested and stained with different concentrations of biotinylated anti-CD45.2 (clone 104) and half an hour later cells were re- stained with streptavidin-PE and anti-CD45.2-PECy7 (clone 104) or anti-CD45-PECy7 (clone 30-F11). The bold circle lines (A, B) indicate the increase of CD45.2-Biotin-Streptavidin-PE staining due to the increase of biotinylated antibody concentration. And the square lines (A, B) indicate the decrease of CD45.2-PECy7 (clone 104) staining as cells already bound with CD45.2-Biotin (clone 104) blocks the binding of CD45.2-PECy7 (clone 104). Sample was CD45R+ B cells (A) CD11b+ myeloid cells (B). Clone 104 blocks the binding of another clone 104 but does not block the binding of clone 30-F11 in B cells (C) and myeloid cells (D). X- axis represents biotinylated antibody concentration and Y-axis represents CD45 staining intensity. Data represent mean ± SD of a single or two (pooled) experiments.

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Figure 3.12: CD45 internalizes in LT-HSC and leukocytes. BM and spleen were harvested and resuspended on X-VIVO hematopoietic cell media. Biotinylated anti-CD45 antibody was added at a concentration of 2.5 µg/ml and cells were incubated at 370C for different times to allow internalization. After incubation for 0-240 minutes, cells were stained with SA-BV510- secondary antibody. Cell incubation without anti-CD45 was used as control. (A) Staining intensity of CD45 and the percentage of cell surface bound CD45 in LT-HSC (B) and B cells and myeloids (C) at the indicated time points. After 240 minutes, LT-HSC lost more surface bound CD45 than B and myeloid cells (D). The bound CD45 percentage were calculated as the percentage of mean fluorescence intensity (MFI) values for each time point with respect of the 0 minutes samples. Data are pooled from three individual experiments with mean ± SD.

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3.2.4 Competitive BM transplantation verifies CD45.2-SAP mediated LT-HSC depletion

To understand whether the depletion of LT-HSC and hematopoietic progenitors defined by flow cytometry reflected a functional depletion, CD45.2-SAP-mediated LT-HSC and HSPC depletion was tested using a competitive BM chimerism assay where transferred donor BM cells competed with recipient BM cells. Here, BM harvested from C57BL/6 mice treated with graded doses of CD45.2-SAP (3 mg/kg, 2 mg/kg, 1 mg/kg and 0.5 mg/kg) or control BIgG- SAP (0.5 mg/kg) was transferred to lightly-irradiated (300 cGy) B6.SJL-ptprca recipient mice. BM from untreated C57BL/6 mice was used as a positive control. Donor chimerism was monitored as accumulation of donor leukocytes within peripheral blood of recipients at the indicated time points. Donor chimerism was also determined in spleen at the final time-point (Figure 3.13A).

In recipients of BM from control untreated and BIgG-SAP treated mice, donor BM-derived total cell concentration (Figure 3.13B) and donor-derived total leukocyte chimerism gradually increased in blood over time, reaching approximately 40-50% of total leukocytes in blood 16 weeks after transplantation (Figure 3.13C-D). In spleen the proportion of donor leukocytes reached approximately 50% 16 weeks after BMT (Figure 3.13E). In contrast, recipient mice that received BM from CD45.2-SAP-treated mice showed less extensive development of donor-derived leukocytes in both blood and spleen (Figure 3.13D-E). The development of donor chimerism was approximately 25% in recipients of 0.5mg/kg CD45.2-SAP treated donor whereas recipients of 1mg/kg and 2mg/kg showed approximately 20% and 10-15% donor chimerism respectively. Donor cells treated with 3mg/kg failed to develop detectable donor chimerism. These data indicate development of donor chimerism is highly dependent on the dose of CD45.2-SAP used.

By analysing the total blood cell concentration in recipient mice over the indicated time points, it was observed that at early time points such as week 2 and 4 after BM transfer, recipients of BM from untreated and BIgG-SAP treated donor mice showed a normal range of peripheral blood leukocyte concentrations (~7×103-8×103 leukocytes/µl) whereas the recipients that received CD45.2-SAP treated BM had a reduced cell concentration (~2×103-5×103/µl) that was related to the CD45.2-SAP dose given to donor mice (Figure 3.14A) and development of donor chimerism (Figure 3.13C). Total cell concentration returned to normal at week 8 post-BMT. It is interesting because all the recipient mice were exposed to 300 cGy irradiation and received 10x106 bulk BM cells but hematopoietic recovery differed.

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To investigate the reason behind lower donor chimerism and lower total cell concentration in CD45.2-SAP treated recipients, the number of donor LT-HSC and HSPC that was transferred to each irradiated recipient was calculated. 10x106 bulk BM cells were transferred to each recipient, so theoretically every recipient mouse should get around 1500-2000 LT-HSC (Lin- c-kit+Sca1+CD48-CD150+) as bulk BM cells contain 0.01-0.02% phenotypically defined LT- HSC. But in this experiment, recipient mice received 10x106 BM cells but different doses of LT-HSC due to dose-dependent LT-HSC depletion by CD45.2-SAP (Figure 3.1B). Here, different recipient groups got different number of LT-HSC (Figure 3.14B) and HSPC (Figure 3.14C) and developed donor chimerism based on the dose of LT-HSC and HSPC transferred. BM from untreated donors engrafted quickly and donor-derived cell concentration increased accordingly in blood and reached a maximum (4.5×103/µl) at week 8 post-BMT. Recipients of BM from untreated donors got approximately 1500 LT-HSC (Figure 3.14B) and 400,000 HSPC (Figure 3.14C) and donor chimerism was approximately 45% at week 16 post-BMT. As the dose of CD45.2-SAP was increased the extent of donor-derived cell development was reduced. Particularly, recipient groups that got donor BM cells from mice treated with a high CD45.2-SAP dose (3 mg/kg) failed to establish detectable amount of donor chimerism as these recipients got only 150 LT-HSC (Figure 3.14B) and 170,000 HSPC (Figure 3.14C). Overall other recipient groups received roughly 400 (2mg/kg), 700 (1mg/kg) and 800 (0.5 mg/kg) LT- HSC and 225,000 (2mg/kg), 335,000 (1mg/kg) and 315,000 (0.5mg/kg) HSPC and engrafted at approximately 10-15%, 20% and 25% respectively (Figure 3.13A). To verify the relationship between depletion percentage and chimerism reduction, percentage of chimerism reduction on different doses of CD45.2-SAP was calculated and data clearly state that development of donor chimerism is dependent on depletion percentage. 0.5 mg/kg CD45.2- SAP depleted approximately 45% of LT-HSC and donor development capability reduced by 40% whereas higher doses such as 2 and 3mg/kg depleted roughly 70% and 85% LT-HSC and donor development capacity declined by 75% and 90% respectively (Figure 3.14D).

To determine the rate of development of distinct leukocyte lineages the concentration of donor derived CD8+ T cells, CD4+ T cells, B cells, myeloid cells and DC was tracked in blood over 16 weeks post-BMT. BM from untreated and BIgG-SAP treated donor mice engrafted efficiently, with myeloid cells (Figure 3.15A) and DC (Figure 3.15B) accumulating quickly after two weeks post transplantation. Donor-derived B cells also appeared rapidly at 4 weeks post-BMT and reached 50% donor-derived within 4 weeks post-BMT (Figure 3.15C) and both

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CD8+ T (Figure 3.15D) and CD4+ T (Figure 3.15E) cells established slowly because donor progenitors take time to migrate into the thymus and for subsequent T cell differentiation and selection before exiting to the periphery. The cell concentration accumulation trend of CD8+ T (Figure 3.16A) and CD4+ T cells (Figure 3.16B) was similar as overall cell concentrations except myeloid cells (Figure 3.16C) because the myeloid population was less affected by CD45.2-SAP treatment (Figure 3.4E) and contributes to accumulation of the donor-derived myeloid population at week 2 after transplantation. B cells (Figure 3.16D) accumulated moderately as transferred HSPC contained significant proportion of premature B cells. So these early-differentiated donor myeloid and B cells could have derived from transplanted donor progenitor cells (Figure 3.2B-E and 3.3B-D). These transferred progenitors cells could lead to development of mature myeloid cells only transiently so myeloid cell number went down at week 4 post-BMT and came back at later time point.

All recipient groups had similar CD8+ T (Figure 3.17A) and CD4+ T cell (Figure 3.17B) number in early time points. But B cells (Figure 3.17C) and myeloid cell (Figure 3.17D) number was different and the trend was same as total cell numbers. As bulk BM cells have a substantial amount of pre-mature B cells and early myeloid progenitor cells which differentiates very quickly and contribute to accumulation of the donor derived B and myeloid population at early stage of transplantation. So it is obvious that early detected B and myeloid cells came from transplanted donor BM and it contributes to early mixed chimerism.

Overall, donor-derived cell accumulation clearly demonstrates a dose-dependent LT-HSC depletion has occurred. But interestingly BIgG-SAP treated donors engrafted steadily and donor cells developed as same as untreated control donors (3-5% variation) though BIgG-SAP appeared to deplete around 40% of phenotypic LT-HSC (Figure 3.1C). So these competitive engraftment data show the ‘apparent’ BIgG-SAP mediated LT-HSC depletion was not an actual functional depletion of LT-HSC. It is possible this observation may have been due to BM niche structure changes induced by BIgG-SAP treatment or this non-specific treatment changed the receptor structures on LT-HSC.

Together these data indicate that CD45.2-SAP effectively depletes LT-HSC from the BM niche and reduces engraftment capacity and this competitive BMT verified this outcome.

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A

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5 80 Untreated Untreated BIgG-SAP BIgG-SAP 4 0.5 mg/kg 60 0.5 mg/kg 1 mg/kg 1 mg/kg 2 mg/kg 2 mg/kg 3 3 mg/kg 3 mg/kg 40 2 20 1

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0 0 Donor chimerism Donor (%) boold in chimerism spleen in Donor (%) d P .5 1 2 3 d P .5 1 2 3 te A 0 te A 0 a S a S e - e - tr G tr G n Ig CD45.2-SAP dose (mg/kg) n Ig U B U B CD45.2-SAP dose (mg/kg)

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Figure 3.13. CD45.2-SAP treated BM engrafts in dose dependent manner under non- myeloablative conditioning. (A) Protocol for BM transplantation assay to verify CD45.2-SAP mediated LT-HSC depletion. BM cells were harvested from CD45.2-SAP treated and control C57BL/6 mice and single cell suspensions prepared. 10x106 bulk BM cells were transplanted into lightly irradiated (300 cGy) recipient B6.SJL-ptprca mice. Donor engraftment was assessed by total percentage of donor CD45+ leukocytes (B) within peripheral blood at the indicated time-points. Donor-derived cell concentration was also tracked (C). Proportion of total donor CD45+ in peripheral blood (D) and spleen (E) at 16 weeks. Data show individual mice with mean ± SD of results from four individual experiments, n=4-7 mice per group. Statistical significance determined by one-way ANOVA, ****p<0.0001, ***p<0.001, **p<0.01, *p<0.05 and ns= no significance.

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Figure 3.14. Donor chimerism depends on transplanted donor LT-HSC and HSPC numbers. Donor C57BL/6 mice were treated with different doses of CD45.2-SAP and 10x106 bulk BM cells were transplanted into lightly irradiated (300cGY) recipient B6.SJL-ptprca mice. Total cell concentration was assessed over the indicated time points (A). Different recipient groups received different number of LT-HSC (B) and HSPC (C) cells and engraft accordingly. Transferred LT-HSC and HSPC number was calculated as a proportion of survived LT-HSC after CD45.2-SAP treatment present in total transferred bulk BM cells (10x106) and reduction of chimerism was also calculated (D). Data show individual mice with mean ± SD of results from four individual experiments, n=4-7 mice per group. Statistical significance determined by one-way ANOVA, ****p<0.0001, ***p<0.001, **p<0.01, *p<0.05 and ns= no significance.

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A B

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Figure 3.15. Engraftment of CD45.2-SAP treated BM under non myeloablative condition. 10x106 bulk BM cells from CD45.2-SAP treated and control C57BL/6 mice were transplanted into lightly irradiated (300cGY) recipient B6.SJL-ptprca mice. Development of donor derived myeloid cells (A), DC (B), B cells (C), CD8+ T cells (D) and CD4+ T cells (E) was tracked in peripheral blood at the indicated time points. Different colour represents different recipient groups treated with CD45.2-SAP and BIgG-SAP. Data show individual mice with mean ± SD of results from four individual experiments, n=4-7 mice per group.

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2 4 8 16 2 4 8 16 Donor B Cell conc B (X10^3cells/ul)Donor

Donor Myeloid Cell conc Cell Myeloid (X10^3cells/ul)Donor Weeks after BMT Weeks after BMT

Figure 3.16. Donor derived cell concentrations increased over the time. Donor C57BL/6 mice were treated with different doses of CD45.2-SAP and 10x106 bulk BM cells were transplanted into lightly irradiated (300cGY) recipient B6.SJL-ptprca mice. Donor derived CD8+ T cells (A), CD4+ T cells (B), myeloid cells (C) and B cells (D) recovery were tracked over the indicated time points. Each line represents individual mice and different colour represent different treated groups. Data show individual mice with mean ± SD of results from four individual experiments, n=4-7 mice per group.

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A B

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B Cell B conc (X10^3cells/ul) 0 0 2 4 8 16 2 4 8 16

Myeloid Cell conc (X10^3cells/ul) Weeks after BMT Weeks after BMT

Figure 3.17. Transplanted donor progenitor cells contribute to early mixed chimerism. Donor C57BL/6 mice were treated with different doses of CD45.2-SAP and 10x106 bulk BM cells were transplanted into lightly irradiated (300cGY) recipient B6.SJL-ptprca mice. Total cell concentration was assessed over the indicated time points. Total absolute cell numbers (A) and leukocytes lineages such as CD 8+ T cells (B), CD4+ T cells (C), B cells (D) and myeloid cells (E) numbers were tracked over the time. Data show individual mice with mean ± SD of results from four individual experiments, different colours represent different treated groups, n=4-7 mice per group.

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3.3. Discussion

The ultimate goal of this PhD project is to induce antigen specific immune tolerance through safer and more specific recipient conditioning. Most studies performed to improve tolerance induction through HSC-mediated antigen targeting approaches have used either myeloablative conditioning or non-myeloablative conditioning, fewer have explored non-genotoxic conditioning. It has been previously shown that non-myeloablative conditioning through low- dose irradiation (300cGy) facilitates BM engraftment with subsequent antigen-specific tolerance induction [173, 251]. However, these low-dose irradiation procedures are non- specific, extensively damage healthy cells and are also genotoxic, leading to permanent DNA damage [252]. Immunotoxin-based recipient conditioning is emerging and is anticipated to have less non-specific toxic effect than typical irradiation and/or chemotherapy based conditioning. Hematopoietic cell-targeted immunotoxins would have selective effect on hematopoietic cells avoiding to damage hematopoietic niches and preserve recipient immunity. Previous research targeting hematopoietic cells was successful to achieve high levels of donor engraftment without the need of irradiation or chemotherapeutics. For example, a recent study using high dose of CD45-SAP (3 mg/kg), an internalizing immunotoxin that targets BM restricted CD45 and depletes >98% of HSC which permits >90% donor chimerism. However, high-dose CD45-SAP induces lymphocytopenia by depleting peripheral leukocytes [229].

Here, in this study titrating the dose of CD45.2-SAP demonstrated a dose-dependent depletion of LT-HSC in immunocompetent mice. As the dose of CD45.2-SAP was reduced the extent of peripheral leukocyte depletion was also reduced. CD45.2-SAP induces cell death by receptor- mediated internalization and targets cells that express CD45 and reduces the risk of non- specific and bystander toxic effect to cells that do not express CD45. The effects of low-dose CD45.2-SAP on LT-HSC was significantly different to HSPC and peripheral leukocytes. However, different expression level of CD45 antigen on the surface of LT-HSC, HSPC and peripheral leukocytes and variation of internalization kinetics among those cells were not correlated with relative sensitivity to CD45.2-SAP. So the result of differential depletion effects might be explained by cell-specific differential processing of CD45.2-SAP or, possibly, LT-HSC are more sensitive to CD45.2-SAP than other hematopoietic or non-hematopoietic cells. Further experimentation is required to test these hypotheses and future experiments will focus on differential CD45.2-SAP processing by different cell types.

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This study also demonstrates that CD45.2-SAP administration increases the number of myeloid and myeloid progenitor-derived cells such as macrophages, granulocytes and monocytes. As CD45.2-SAP depletes an enormous number of LT-HSC, HSPC and other BM cells, so the BM is expected to contain a large number of dead cells after CD45.2-SAP treatment. Macrophages are responsible for clearing up the dead cells so it would be a possible explanation why macrophage number was increased after CD45.2-SAP treatment. Previous studies also support this finding that infection, inflammation and toxic treatment induces macrophage expansion [253-256]. However, further experimentation is required to verify this explanation.

In addition, it is also shown that CD45.2-SAP treatment reduced donor BM engraftment capacity indicating functional LT-HSC depletion. Competitive BM transplantation of CD45.2- SAP treated BM to lightly irradiated (300cGy) recipients engrafted according to CD45.2-SAP dose given to donors. But interestingly untargeted immunotoxin (BIgG-SAP) treated donors engrafted steadily and donor cells developed as same as untreated control donors (3-5% variation) though BIgG-SAP appeared to deplete a significant amount of phenotypic LT-HSC. So the BIgG-SAP mediated depletion effect on LT-HSC may be explained by non-specific untargeted toxic treatment forces to change the receptor structures on LT-HSC or untargeted treatment helps to mobilize LT-HSC from BM to periphery that is why BM staining showed reduced LT-HSC number. It would be interesting to see the effect of SAP bound to other protein molecule. But SAP is the toxic portion of BIgG-SAP/CD45.2-SAP and these monoclonal antibodies act as its transporter and target identifier. These monoclonal antibodies help the immunotoxin to enter the targeted cells which ultimately induce cell death by inhibiting protein synthesis [232]. Further experimentation with unconjugated SAP, unconjugated antibody or SAP conjugated with irrelevant protein which does not have any specific receptor such as ovalbumin is required to understand the non-specific effect of antibody and SAP.

Taken together data presented in this chapter provide substantial evidence that a low CD45.2- SAP dose might be advantageous compared to traditional conditioning procedures such as irradiation and/or chemotherapeutics when low-level but stable mixed hematopoietic chimerism is desired. Some previous studies reported moderate or low level of donor chimerism could be sufficient to induce immune tolerance [241, 257]. So it is anticipated that lower CD45.2-SAP dose would be effective to induce immune tolerance by facilitating significant level of donor chimerism. As CD45.2-SAP is non-genotoxic and does not have any DNA damaging effect so it could be a potential alternative for recipient conditioning.

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3.4. Summary

In summary, it is established that titrating the dose of CD45.2-SAP led to a dose-dependent depletion of phenotypic LT-HSC in BM. Dose-dependent LT-HSC depletion by CD45.2-SAP was verified by competitive bone marrow transplantation. In contrast to higher dose CD45.2- SAP, a lower dose (0.5 mg/kg) preferentially depleted 45-50% of LT-HSC whereas peripheral leukocytes were largely preserved (~85% retained). This low-dose CD45.2-SAP is non- genotoxic and does not cause DNA damage. Thus low-doses of CD45.2-SAP could provide a useful alternative to toxic conditioning approaches for tolerance induction with gene- engineered BM. Further experimentation is required to determine the ability of low-dose CD45.2-SAP in gene-engineered bone marrow transplantation. This is investigated in chapter 4.

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Chapter 4 Non-genotoxic conditioning using leukocyte-targeting immunotoxin facilitates tolerance induction

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4.1. Introduction

Several studies have been performed to induce T-cell tolerance by multilineage mixed hematopoietic chimerism [258-260]. In 1984 Ildstad and Sachs induced donor-specific skin graft tolerance by achieving mixed lymphohematopoietic chimerism where hematopoietic and lymphoid cells were derived from both host and donor HSC [170]. Some kidney transplant studies also support this phenomenon [261-263]. For inducing transplant tolerance complete immune depletion and reconstitution is not necessary, as a small percentage of survived donor HSC-derived cells have been found sufficient to induce tolerance [264]. Paralleling studies of mixed chimerism, transplanting genetically-engineered haematopoietic stem cells (HSC) encoding antigen provides an efficient means to induce antigen-specific immune tolerance [4, 243]. It has been found, here too, low-level, but stable HSC engraftment is sufficient for T-cell tolerance induction and prevention of autoimmune disease in animal models [241, 248]. Therefore, highly-toxic myeloablative and/or immunoablative pre-transfer recipient conditioning regimens that are typically used to achieve high engraftment levels for engineered HSC may not be necessary.

To progress this approach to clinical application one requirement will be to minimize conditioning toxicity to avoid transplant-related complications. Although toxicity can be alleviated by reducing the intensity of pre-transplant conditioning regimen [265, 266]. Reduced intensity conditioning regimens currently use chemotherapeutics as key components and this carries the risks of genotxicity. Additionally, reduced intensity conditioning procedures currently used clinically are designed for treatment of malignancies and in this setting malignant cell ablation, and consequently, immune ablation is an important component of the overall effect of the conditioning regimen. Moreover, induction of tolerance is not a focus of currently-used reduced intensity conditioning and, in fact, strong graft vs leukemia effects are desired.

Targeted immunotoxin-based conditioning for depleting recipient HSC has shown promise in genetically-engineered BMT to treat hematologic disease models of sickle cell anaemia and haemophilia [229, 267]. CD45-SAP facilitates >90% donor HSC chimerism and enabled treatment of sickle cell anemia in an animal model when used at high dose [229]. It could be adopted for autoimmune disease but at high dose CD45-SAP is immunodepleting and shows bystander effects. The hope for a less toxic and specific option by using low-dose CD45-SAP as non-genotoxic conditioning approach to induce antigen-specific T-cell tolerance. It is shown

88 | P a g e in Chapter 3 that low-dose CD45.2-SAP (0.5mg/kg) depletes approximately 45% of LT-HSC while depleting a reduced proportion of HSPC and peripheral leukocytes. So theoretically it could create approximately 45% space inside bone marrow for donor exogenous HSC which would be sufficient to permit a sufficient level of donor chimerism required for tolerance induction.

The hypothesis for this chapter is gene-engineered donor HSC engraftment facilitate by non- genotoxic conditioning through HSPC non-specific immunotoxin CD45.2-SAP permits subsequent induction of immune tolerance. In this chapter, I describe that low-dose CD45.2- SAP facilitates sufficient donor HSC chimerism to induce antigen-specific T-cell tolerance. I also describe transplanting a higher number of BM cells accelerates the development of chimerism in recipients in a way that might reduce the time required for tolerance induction.

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4.2. Results

4.2.1. Low-dose CD45.2-SAP enables tolerance induction by facilitating stable mixed hematopoietic chimerism of gene-modified BM

To determine whether CD45.2-SAP-mediated LT-HSC depletion facilitates donor LT-HSC engraftment, bulk BM cells from OVA expressing transgenic 11c.45.1 or non-transgenic control (both CD45.1+) mice which express a membrane bound truncated form of OVA under the control of CD11c promoter were transferred to CD45.2-SAP-conditioned recipient C57BL/6 (CD45.2+) mice. Donor leukocyte development was tracked in blood at the indicated time points. Then, to determine whether this also induced OVA-specific T-cell tolerance, the recipient mice were tested by immunising (OVA/QuilA) 25 weeks after BM transfer and OVA- responsiveness was tested by ELISPOT. As 0.5 mg/kg CD45.2-SAP depletes approximately 45-50% of LT-HSC (Figure 3.1A) but with a reduced effect on peripheral leukocytes (Figure 3.4 A-E), it was hypothesized that 0.5 mg/kg CD45.2-SAP would facilitate sufficient level of donor HSC chimerism that would be sufficient for tolerance induction. Therefore, recipient mice were conditioned with low-dose CD45.2-SAP (0.5 mg/kg) and BM transfer was performed. Untreated mice were used as negative controls and mice conditioned with low-dose (300 cGy) irradiation were used as positive controls for donor BM chimerism. Cell depletion after CD45.2-SAP treatment was assessed by retro-orbital bleeding one day prior to BM transfer (i.e. 6 days after treatment). Donor BM chimerism was assessed as development of donor leukocytes by tracking total leukocyte frequency and the proportion of lineages within peripheral blood. Donor chimerism was also determined in spleen and BM at the final time- point (Figure 4.1A).

Analysis of the peripheral blood following conditioning but before transplantation confirmed the effectiveness of CD45.2-SAP depletion in each mouse treated. Overall, approximately 40- 45% of blood leukocytes were depleted when CD45.2 SAP was administered (Figure 4.1B). 300 cGy radiation depleted 50-60% of peripheral blood leukocytes. Tracking peripheral blood leukocyte concentration across time showed that recovery of peripheral blood leukocyte was more rapid in CD45.2-SAP-conditioned recipients than in irradiated BM recipients (Figure 4.1C). In CD45.2-SAP-conditioned mice peripheral blood leukocytes levels were restored to 80-90% of that in untreated controls within two weeks of CD45.2-SAP treatment and

90 | P a g e transplantation. In contrast, in irradiated mice peripheral blood leukocytes populations did not return to approximately normal levels until more than eight weeks after irradiation and BM transfer (Figure 4.1C).

Therefore, conditioning with CD45.2-SAP (0.5 mg/kg) partially preserved recipient leukocyte populations (Figure 4.1B) indicating that immunity likely remains, at least, partially intact. In CD45.2-SAP treated recipients, both Tg 11c.45.1 and non-transgenic littermate donor BM- derived leukocytes gradually increased in frequency in blood over time reaching approximately 10-15% of total leukocytes 25 weeks after BM transfer and this level was retained also at 1 week -immunization at week 25 (Figure 4.2A-B). In contrast, 300 cGy radiation enabled (Figure 4.2A) 40-50% chimerism after Tg 11c.45.1 donor BM transfer. BM transferred to untreated recipients failed to generate detectable chimerism higher than the background detected for un-transplanted recipient mice (Figure 4.2A). Donor-derived cells (CD45.1+) from CD45.2-SAP treated recipient group comprised approximately 15% in spleen at this time point (Figure 4.2C). To determine the rate of development of distinct leukocyte lineages, donor-derived CD8+ T cells, CD4+ T cells, B cells, myeloid cells and DC were tracked in blood. Both Tg 11c.45.1 and non-transgenic littermate donor-derived cells accumulated slowly, with myeloid cells apparent 2 weeks after BM transfer (Figure 4.3A) and DC at 4 weeks (Figure 4.3B). On the other hand, both CD8+ (Figure 4.3C) and CD4+ T cells (Figure 4.3D) as well as B cells (Figure 4.3E) established very slowly and took 16 weeks to be detected.

Overall, chimerism in blood and spleen was compared with BM chimerism 26 weeks post BM transfer, to confirm that accumulated donor-derived lineages was representative of engrafted donor BM cells. Donor derived CD45.1+ cells were analysed in BM and around 10-13% donor chimerism was observed (Figure 4.4A) indicating proportional chimerism was similar to that observed for donor derived leukocyte accumulation in peripheral blood and spleen. Both Tg 11c.45.1 and non-transgenic littermate donor BM engrafted at similar levels and as expected donor BM did not engraft in untreated recipient mice. Donor-derived HSPC (Lin-/c-kit+) were also measured and approximately 10-12% of donor chimerism were detected which is similar to overall CD45.1+ donor cell chimerism in BM (Figure 4.4B).

Mice were immunized with OVA/QuilA at week 25 after BM transfer and tolerance to OVA was assessed using an IFN-γ ELISPOT assay. One week after immunization, ELISPOT analysis showed that no OVA257-264-specific responsiveness was present in CD45.2-SAP conditioned recipients of OVA encoding BM (Tg 11c.45.1) whereas recipients of non-OVA

91 | P a g e expressing BM (non-Tg 11c45.1) showed a significant IFN-γ response to OVA257-264 (Figure 4.5A). Tg 11c.45.1 BM transferred to untreated recipients failed to generate detectable donor chimerism and showed significant level of IFN-γ response to OVA257-264 (Figure 4.5A) indicating no tolerance induction in the absence of stable donor chimerism. No BMT control groups displayed OVA257-264-specific IFN-γ response. A similar response was observed in CD45.2-SAP conditioned recipients with no BMT groups indicate CD45.2-SAP has no effect on tolerance induction.

Taken together, our data suggest that low-dose CD45.2-SAP (0.5 mg/kg) efficiently facilitates tolerance induction through stable mixed hematopoietic chimerism of gene-modified BM. Additionally, CD45.2-SAP conditioned mice also recovered depleted immune cells more rapidly than the mice conditioned with irradiation.

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Untreated/CD45-SAP treated A C57BL/6 C57BL/6 CD45.2 -SAP

Day -7 Day 0 Week 16 Week 25 Week 26 Day -1 Week 2 Week 4 Week 8 Blood Blood Blood Blood Blood Mice immunisation Blood Spleen Bone marrow Donors Non-Tg/Tg-11c.45.1

B

15 **** ns

10 ns 5

0

Cellconc. (X10-^3 Cells/ul) T T T T T M M M M M B B B B B o .1 o .1 .1 N 5 N 5 5 .4 .4 .4 c c c 1 1 1 1 1 1 g g g T -T T n o N No Rx CD45.2-SAP

C

15

10 No BMT/No Rx Tg 11c.45.1 BMT/No Rx No BMT/CD45.2-SAP Non-Tg 11c.45.1 BMT/CD45.2-SAP 5 Tg 11c.45.1 BMT/CD45.2-SAP Tg 11c.45.1 BMT/300cGY Cell conc. Cell (X10-^3Cells/ul) 0 -1 0 2 4 8 16 25 26 Weeks post BMT

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Figure 4.1. CD45.2-SAP allows rapid immune recovery. (A) BM from Non-Tg 11c.45.1 and Tg 11c.45.1 (CD45.1+) mice was transferred to untreated, CD45.2-SAP (0.5 mg/kg) treated and 300 cGy irradiated C57BL/6 recipients (CD45.2+). (B) 6 days after CD45.2-SAP administration and 1 day prior to BM transplantation blood was collected to analyse the depletion effect of CD45.2-SAP (B). Peripheral blood leukocyte concentration in recipient C57BL/6 at indicated time point post CD45.2-SAP mediated BMT was assessed (C). Data show individual mice with mean ± SD for results from four individual experiments, n=5-7 mice per group. Statistical significance determined by one-way ANOVA, ****p<0.0001, ***p<0.001, **p<0.01, *p<0.05 and ns= no significance. No Rx=No treatment.

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A

60

50 No BMT/No Rx 40 Tg 11c.45.1 BMT/No Rx No BMT/CD45.2-SAP 30 Non-Tg 11c.45.1 BMT/CD45.2-SAP

20 Tg 11c.45.1 BMT/CD45.2-SAP Tg 11c.45.1 BMT/300cGY

Donorchimerism (%) 10

0 2 4 8 16 25 26 Weeks post BMT

B C

**** **** **** **** 50 50

40 40

30 30 **** 20 ns **** 20 ns 10 ns 10 ns

0 0 T T T T T T

Donor chimerism in boold (%)booldDonor chimerism in M M M M M M T T T T T T B B B B B B M M M M M M o .1 o .1 .1 .1 spleen(%) in chimerism Donor B B B B B B N 5 N 5 5 5 o .1 o .1 .1 .1 .4 .4 .4 .4 N 5 N 5 5 5 c c c c .4 .4 .4 .4 1 1 1 1 c c c c 1 1 1 1 1 1 1 1 g g g g 1 1 1 1 T T T T g - g g g T n T -T T o n N o N No Rx CD45.2-SAP 300cGy No Rx CD45.2-SAP 300cGy

Figure 4.2. Low-dose CD45.2-SAP permits donor chimerism. 10x106 bulk BM from Non- Tg 11c.45.1 and Tg 11c.45.1 (CD45.1+) mice was transferred to untreated, CD45.2-SAP (0.5 mg/kg) treated and 300 cGy irradiated C57BL/6 recipients (CD45.2+). Donor chimerism was tracked within leukocytes (A) in peripheral blood at the indicated time points. The proportion of donor (CD45.1+) cells in peripheral blood (B) and spleen (C) is shown. Data show individual mice with mean ± SD for results pooled from four individual experiments, n=7-10 mice per group. Statistical significance determined by one-way ANOVA, ****p<0.0001, ***p<0.001, **p<0.01, *p<0.05 and ns= no significance.

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A B

No BMT/No Rx No BMT/No Rx Tg 11c.45.1 BMT/No Rx Tg 11c.45.1 BMT/No Rx No BMT/CD45.2-SAP 80 80 No BMT/CD45.2-SAP Non-Tg 11c.45.1 BMT/CD45.2-SAP Non-Tg 11c.45.1 BMT/CD45.2-SAP Tg 11c.45.1 BMT/CD45.2-SAP Tg 11c.45.1 BMT/CD45.2-SAP Tg 11c.45.1 BMT/300cGY 60 60 Tg 11c.45.1 BMT/300cGY

40 40

20 20

0 0 2 4 8 16 26 DonorDC (%)Chimerism 2 4 8 16 26

DonorMyeloid (%)Chimerism Weeks post BMT Weeks post BMT

C D

No BMT/No Rx 80 No BMT/No Rx 80 Tg 11c.45.1 BMT/No Rx Tg 11c.45.1 BMT/No Rx No BMT/CD45.2-SAP No BMT/CD45.2-SAP 60 Non-Tg 11c.45.1 BMT/CD45.2-SAP 60 Non-Tg 11c.45.1 BMT/CD45.2-SAP Tg 11c.45.1 BMT/CD45.2-SAP Tg 11c.45.1 BMT/CD45.2-SAP Tg 11c.45.1 BMT/300cGY Tg 11c.45.1 BMT/300cGY 40 40

20 20

0 0

2 4 8 16 26 2 4 8 16 26

Donor CD8+ Chimerism (%) CD8+ Donor Chimerism (%) CD4+ Donor Chimerism Weeks post BMT Weeks post BMT

E

No BMT/No Rx 80 Tg 11c.45.1 BMT/No Rx No BMT/CD45.2-SAP 60 Non-Tg 11c.45.1 BMT/CD45.2-SAP Tg 11c.45.1 BMT/CD45.2-SAP Tg 11c.45.1 BMT/300cGY 40

20

0 2 4 8 16 26 Donor(%)cell B Chimerism

Weeks post BMT

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Figure 4.3. CD45.2-SAP allows chimerism of donor BM derived lineages. 10x106 bulk BM cells from Non-Tg 11c.45.1 and Tg 11c.45.1 mice were transferred into untreated, low-dose CD45.2-SAP treated and lightly irradiated C57BL/6 mice. Development of donor derived myeloid cells (A), DC (B), CD8+ T cells (C), CD4+ T cells (D) and B cells (E) was assessed in peripheral blood at the indicated time points. Data are pooled from four individual experiments and show individual mice with mean ± SD, n=7-10 mice per group.

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A B

**** 60 **** **** 60 ****

40 40

**** ns **** 20 20 ns

ns ns

0 0

Donor chimerism in BMDonorchimerism in (%) T T T T T T T T T T T T M M M M M M HSPC chimerismDonor (%) BM in M M M M M M B B B B B B B B B B B B o .1 o .1 .1 .1 N 5 N 5 5 5 o .1 o .1 .1 .1 4 4 4 4 N 5 N 5 5 5 . . . . .4 .4 .4 .4 c c c c c c c c 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 g g g g g T T T T g g g T - T -T T n n o o N N No Rx CD45.2-SAP 300cGy No Rx CD45.2-SAP 300cGy

Figure 4.4. CD45.2-SAP facilitates donor HSPC chimerism. Overall chimerism of Non-Tg 11c.45.1 and Tg 11c.45.1 was analysed 26 weeks after BM transplantation. Overall donor chimerism in BM (A) and the percentage of donor derived HSPC (Lin-c-kit+) cells were also analysed (B). Data show individual mice with mean ± SD of results from four experiments, n=5-10 mice per group. Statistical significance determined by one-way ANOVA, ****p<0.0001, ***p<0.001, **p<0.01, *p<0.05 and ns= no significance.

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A * **** *** 1000 *

SPC 800 ** 5

600

400 SFU/2.5×10

 200 ns

IFN- 0 T T T T T T e M M M M M M ic B B B B B B M o .1 o .1 .1 .1 .1 N 5 N 5 5 5 5 .4 .4 .4 .4 .4 c c c c c 1 1 1 1 1 1 1 1 1 1 g g g g g T -T T T T n o N No Rx CD45.2-SAP 300cGY

Figure 4.5. CD45.2-SAP facilitates antigen specific T-cell tolerance. 10x106 bulk BM cells from Non-Tg 11c.45.1 and Tg 11c.45.1 mice were transferred into untreated, low-dose CD45.2-SAP treated and lightly irradiated C57BL/6 mice. Mice were challenged with

OVA/QuilA on week 25 and one week later, immune response to OVA257-264 by recipients was assessed by ELISPOT (A). Data show individual mice with mean ± SD of results from four experiments, n=5-10 mice per group. Statistical significance determined by one-way ANOVA, ****p<0.0001, ***p<0.001, **p<0.01, *p<0.05 and ns= no significance.

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4.2.2. Increasing the BM cell dose accelerates donor chimerism and tolerance induction

After transfer of 10x106 bulk BM cells to low-dose (0.5mg/kg) CD45.2-SAP-conditioned recipient, chimerism developed slowly and 26 weeks was required to get a modest, but effective level of donor chimerism to induce T-cell tolerance. It is time consuming and approximately one-third of a mouse life expectancy. There would be two possible options to get faster chimerism, i) to increase CD45.2-SAP dose to create more space for exogenous BM or ii) to increase the number of donor BM cells transferred. Increasing the dose of CD45.2-SAP would lead to a more pronounced ‘bystander’ effect through depletion of a higher proportion of leukocytes. So increasing the number of donor BM cells could be an option to accelerate the level of donor chimerism without increasing bystander effects. To determine whether increasing the dose of transferred BM facilitates accelerated development of donor chimerism 25x106 bulk BM cells from transgenic, OVA expressing 11c45.1 or non-transgenic littermates were transferred to CD45.2-SAP-conditioned (0.5mg/kg) recipient C57BL/6 mice. Donor leukocyte development was tracked in blood at the indicated time points. To test the OVA specific T-cell tolerance, the recipient mice were immunized with OVA/QuilA 16 weeks after BM transplantation and OVA responsiveness was tested by IFN-γ ELISPOT (Figure 4.6A).

The total donor cell dose used (25x106) was split into two separate injections of 12.5x106 BM cells each, to limit potential harmful effects on mice health. BM cells were transferred on day 2 and day 7after CD45.2-SAP administration. Untreated mice were used as negative controls and mice conditioned with low-dose (300 cGy) irradiation were used as positive controls for donor chimerism and also received 25x106 BM cells in two injections. Donor BM chimerism was assessed as development of donor leukocytes by tracking total leukocyte frequency and the proportion of lineages within peripheral blood. Donor chimerism was also determined in spleen and BM at the final time-point.

In CD45.2-SAP-conditioned mice that received BM, peripheral blood leukocyte level had returned to 70-80% of control values within two weeks on transfer and to approximately100% within 8 weeks after transfer. In contrast in irradiated mice, recovery of peripheral blood leukocytes numbers was slower (Figure 4.6B).

In CD45.2-SAP-conditioned recipients of BM from transgenic 11c.45.1 and non-transgenic littermates, donor-derived total leukocyte chimerism gradually increased in blood and reached approximately 10-15% 16 weeks post-BMT (Figure 4.7A-B). In spleen the proportion of

100 | P a g e donor-derived cells was approximately 15% 16 weeks post-BMT (Figure 4.7C). In contrast, transgenic 11c.45.1 BM transferred to untreated recipient showed less than 1% donor chimerism 16 weeks post-BMT. 300cGy radiation enabled roughly 50-55% of donor chimerism at week 16 post-BMT both in blood and spleen (Figure 4.7 A-C). To determine the rate of development of distinct leukocyte lineages, donor-derived CD8+ T cells, CD4+ T cells, B cells, myeloid cells and DC were tracked in blood. In CD45.2-SAP-conditioned recipients of Tg 11c.45.1 and non-Tg littermates, donor derived myeloid cells (Figure 4.8A) and DC (Figure 4.8B) appeared quickly, within two weeks after BMT. On the other hand CD8+ T- cells (Figure 4.8C), CD4+ T-cells (Figure 4.8D) and B cells (Figure 4.8E) established more slowly and took 8 weeks to be detected.

The trend of overall chimerism development was compared between the higher cell number used (25x106) and the standard cell number transferred (10x106) and, as expected, increasing the donor BM dose increased the extent of donor chimerism established. The established overall chimerism of the higher dose BMT was approximately double than the standard dose BMT (Figure 4.9A). To determine whether the extent of donor-derived lineages accumulating in blood and spleen was representative of engrafted donor BM cells, donor chimerism in BM was analysed. Approximately 15% donor chimerism was observed in BM (Figure 4.10A) and the proportion was on par to that observed in blood and spleen. Both Tg 11c.45.1 and non- transgenic littermate donors’ BM engrafted at similar levels and as expected donor BM did not engraft in untreated recipient mice. Donor derived HSPC (Lin-/c-kit+) were also measured and 10-15% donor chimerism was detected which was similar to overall donor chimerism observed in BM (Figure 4.10B).

To test whether OVA-specific T-cell tolerance was induced, mice were immunized with

OVA/QuilA at week 16 post-BMT and OVA257-264 responsiveness was assessed by an IFN-γ ELISPOT assay one week later. IFN-γ ELISPOT analysis displayed increased IFN-γ spot forming unit (SFU) in response to OVA257-264 in no BMT control recipients and CD45.2-SAP conditioned recipients with no BMT (Figure 4.11A) compared to tolerance control Tg- 11c.45.1 mice. A similar SFU number was observed in untreated control recipients of Tg

11c.45.1 BM groups. However, IFN-γ ELISPOT analysis demonstrated no OVA257-264 specific responsiveness in CD45.2-SAP conditioned recipients of Tg 11c.45.1 BM compared to recipients of non-Tg 11c.45.1 BM which showed an increased IFN-γ response to OVA257-264 relative to Tg-11c.45.1 tolerance control groups (Figure 4.11A). 300cGy treated recipients of

Tg 11c.45.1 BM and Tg 11c.45.1 mice failed to show an IFN-γ response to OVA257-264

101 | P a g e indicating tolerance control group behaved as expected (Figure 4.11A). Comparing these data with 10x106 BMT (Figure 4.5A) indicated approximately 10% stable donor chimerism would be sufficient to induce antigen specific T-cell tolerance.

Overall, increasing the donor BM cell numbers increased the proportion of donor derived cells and facilitates tolerance induction by allowing a significant level of stable donor chimerism in 16 weeks.

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A CD45.2-SAP

C57BL/6 Untreated/CD45-SAP treated C57BL/6

Day -7 Day -5 Day 0 Week 2 Week 4 Week 8 Week 16 Week 17 Blood Blood Blood Blood Blood Mice immunization Spleen Bone marrow

Donors Non-Tg/Tg-11c.45.1

B

15

No BMT/No Rx

Tg 11c.45.1 BMT/No Rx 10 No BMT/CD45.2-SAP Non-Tg 11c.45.1 BMT/CD45.2-SAP Tg 11c.45.1 BMT/CD45.2-SAP 5 Tg 11c.45.1 BMT/300cGY

0

Cellconc.(X10-^3 Cells/ul) 2 4 8 16 17

Weeks post BMT

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Figure 4.6. Increasing the BM cell dose permits rapid cell recovery. (A) A total 25x106 bone BM from Non-Tg 11c.45.1 and Tg 11c.45.1 (CD45.1+) mice was transferred to untreated, CD45.2-SAP (0.5 mg/kg) treated and 300 cGy irradiated C57BL/6 recipients (CD45.2+). Total cell dose was split into two shots 12.5x106 each and transferred on day -5 (two days post CD45.2-SAP administration) and day 0 (7 days post CD45.2-SAP administration). (B) Relative levels of peripheral blood cell concentration in recipient C57BL/6 at indicated time point post CD45.2-SAP mediated BMT was assessed. Data show individual mice with mean ± SD of results from two individual experiments, n=4 mice per group. Statistical significance determined by one-way ANOVA, ****p<0.0001, ***p<0.001, **p<0.01, *p<0.05 and ns= no significance. Rx=No treatment.

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A

60

No BMT/No Rx 40 Tg 11c.45.1 BMT/No Rx No BMT/CD45.2-SAP Non-Tg 11c.45.1 BMT/CD45.2-SAP 20 Tg 11c.45.1 BMT/CD45.2-SAP Tg 11c.45.1 BMT/300cGY

Donorchimerism (%) 0 2 4 8 16 17 Weeks post BMT

B C

**** **** **** 60 60 ****

40 40

**** **** ns ns 20 20 ns ns

0 0

T T T T T T T T T T T T

Donor chimerism in boold (%) boold in chimerism Donor M M M M M M M M M M M M B B B B B B Donorchimerism spleen in (%) B B B B B B o .1 o .1 .1 .1 o .1 o .1 .1 .1 N 5 N 5 5 5 N 5 N 5 5 5 .4 .4 .4 .4 .4 .4 .4 .4 c c c c c c c c 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 g g g g g T g g g T T -T T T -T T n n o o N N

No Rx CD45.2-SAP 300cGy No Rx CD45.2-SAP 300cGy

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Figure 4.7. Higher BM dose accelerates donor chimerism. 25x106 bulk BM from Non-Tg 11c.45.1 and Tg 11c.45.1 (CD45.1+) mice was transferred to untreated, CD45.2-SAP (0.5 mg/kg) treated and 300 cGY irradiated C57BL/6 recipients (CD45.2+). Donor chimerism was tracked by total leukocytes (A) within peripheral blood at the indicated time points. Development of total donor CD45+ in peripheral blood (B) and spleen (C) was also assessed. Data show individual mice with mean ± SD of results from two individual experiments, n=4 mice per group. Statistical significance determined by one-way ANOVA, ****p<0.0001, ***p<0.001, **p<0.01, *p<0.05 and ns= no significance.

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A B No BMT/No Rx No BMT/No Rx Tg 11c.45.1 BMT/No Rx Tg 11c.45.1 BMT/No Rx No BMT/CD45.2-SAP No BMT/CD45.2-SAP 80 Non-Tg 11c.45.1 BMT/CD45.2-SAP Non-Tg 11c.45.1 BMT/CD45.2-SAP 80 Tg 11c.45.1 BMT/CD45.2-SAP Tg 11c.45.1 BMT/CD45.2-SAP Tg 11c.45.1 BMT/300cGY Tg 11c.45.1 BMT/300cGY 60 60

40 40

20 20

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DonorDC (%)Chimerism

2 4 8 16 17 2 4 8 16 17 DonorMyeloid (%)Chimerism Weeks post BMT Weeks post BMT

C D

No BMT/No Rx No BMT/No Rx Tg 11c.45.1 BMT/No Rx Tg 11c.45.1 BMT/No Rx 80 No BMT/CD45.2-SAP 80 No BMT/CD45.2-SAP Non-Tg 11c.45.1 BMT/CD45.2-SAP Non-Tg 11c.45.1 BMT/CD45.2-SAP 60 Tg 11c.45.1 BMT/CD45.2-SAP Tg 11c.45.1 BMT/CD45.2-SAP 60 Tg 11c.45.1 BMT/300cGY Tg 11c.45.1 BMT/300cGY

40 40

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DonorCD8+ (%)Chimerism 2 4 8 16 17 Donor CD4+ Chimerism (%) CD4+ Donor Chimerism

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No BMT/No Rx Tg 11c.45.1 BMT/No Rx No BMT/CD45.2-SAP 80 Non-Tg 11c.45.1 BMT/CD45.2-SAP Tg 11c.45.1 BMT/CD45.2-SAP Tg 11c.45.1 BMT/300cGY 60

40

20

0

2 4 8 16 17 Donor(%)cell B Chimerism Weeks post BMT

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Figure 4.8. Higher BM dose accelerates donor BM-derived lineage chimerism. 25x106 bulk BM cells from Non-Tg 11c.45.1 and Tg 11c.45.1 mice were transferred into untreated, low-dose CD45.2-SAP treated and lightly irradiated C57BL/6 mice. Development of donor derived myeloid cells (A), DC (B), CD8+ T cells (C), CD4+ T cells (D) and B cells (E) was assessed in peripheral blood at the indicated time points. Data are pooled from two individual experiments and show individual mice with mean ± SD, n=4 mice per group.

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A

6 Non-Tg 11c.45.1 BMT (10×10 ) 6 20 Non-Tg 11c.45.1 BMT (25×10 ) Tg 11c.45.1 BMT (10×10 6) 6 Tg 11c.45.1 BMT (25×10 ) 15

10

5

DonorChimerism (%) 0

2 4 8 16 17 26

Weeks post BMT

Figure 4.9. Higher BM dose permits higher and faster donor chimerism than standard dose. 25x106 bulk BM from Non-Tg 11c.45.1 and Tg 11c.45.1 (CD45.1+) mice was transferred to untreated, CD45.2-SAP (0.5 mg/kg) treated and 300 cGY irradiated C57BL/6 recipients (CD45.2+). Donor chimerism was tracked by total leukocytes (A) within peripheral blood at the indicated time points. Data show individual mice with mean ± SD of results from two/four individual experiments, n=4 mice per group.

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A B

80 **** 80 **** **** 60 **** 60 40 **** 40 ns **** 20 ns ns 20 ns 0 0 Donor chimerism in BMDonorchimerism in (%) T T T T T T M M M M M M T T T T T T B B B B B B M M M M M M o 1 o .1 1 1 B B B B B B N . N . . 5 5 5 5 (%) BM in chimerism HSPC Donor o .1 o .1 .1 .1 .4 .4 .4 .4 N 5 N 5 5 5 c c c c .4 .4 .4 .4 1 1 1 1 c c c c 1 1 1 1 1 1 1 1 g g 1 1 1 1 g g T g T -T T g g g T n T -T T o n N o N

No Rx CD45.2-SAP 300cGy No Rx CD45.2-SAP 300cGy

Figure 4.10. Higher BM cell dose facilitates donor HSPC chimerism. Overall chimerism of Non-Tg 11c.45.1 and Tg 11c.45.1 was analysed 17 weeks after BM transplantation. Overall donor chimerism in BM (A) and the percentage of donor derived HSPC (Lin-c-kit+) cells were also analysed (B). Data show individual mice with mean ± SD of results from two experiments, n=4 mice per group. Statistical significance determined by one-way ANOVA, ****p<0.0001, ***p<0.001, **p<0.01, *p<0.05 and ns= no significance.

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A

500 ****

ns SPC

5 400 *** **** 300

200

SFU/1.25×10 100

 ns

IFN- 0 T T T T T T e M M M M M M ic B B B B B B M o .1 o .1 .1 .1 .1 N 5 N 5 5 5 5 .4 .4 .4 .4 .4 c c c c c 1 1 1 1 1 1 1 1 1 1 g g g g g T -T T T T n o N

No Rx CD45.2-SAP 300cGY

Figure 4.11. Higher BM cell dose induces T-cell tolerance within 17 weeks. 25x106 bulk BM cells from Non-Tg 11c.45.1 and Tg 11c.45.1 (CD45.1+) mice were transferred to untreated, CD45.2-SAP (0.5 mg/kg) treated and 300 cGy irradiated C57BL/6 recipients (CD45.2+). Mice were challenged with OVA/QuilA on week 16 and one week later, immune response to OVA257-264 by recipients was assessed by ELISPOT (A). Data show individual mice with mean ± SD of results from two experiments, n=4 mice per group. Statistical significance was determined by one-way ANOVA, ****p<0.0001, ***p<0.001, **p<0.01, *p<0.05 and ns= no significance.

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4.3. Discussion

The majority of animal studies aiming to induce immune tolerance through HSC-based genetic antigen targeting approaches have used ubiquitous promoters to drive expression of specific antigen with myeloablative or non-myeloablative recipient conditioning [173, 257]. These myeloablative or non-myeloablative conditioning regimes either completely or partially eliminate recipient immunity. However some studies have targeted antigen to APC populations. Ko and colleagues targeted myelin oligodendrocyte glycoprotein (MOG) to DC controlled by 960bp CD11c promoter and transferred retrovirally transduced HPC to experimental autoimmune encephalomyelitis (EAE) mice as a model of MS. HPC transplantation where MOG expression was controlled by DC promoter delayed EAE onset. Myeloablative conditioning with 1100cGy total body irradiation was used as recipient conditioning [180]. The same group showed that ubiquitous MOG expression prevents EAE in an animal model when using non-myeloablative recipient conditioning (275 cGy radiation) [268]. In 2013, Coleman et al. showed targeted ovalbumin (OVA) expression to DC or other antigen presenting cells induce antigen-specific T-cell tolerance in naïve and primed recipient however light total body irradiation (300 cGy) was used [173]. All studies provide a strong potential of antigen targeting approaches for improving immune tolerance but the major drawback of this approach is radiation and/or chemotherapy mediated transplant related toxicity.

In this chapter, it is shown that low-dose CD45.2-SAP (0.5mg/kg) effectively conditioned recipient mice for transfer of gene-engineered BM and facilitated 10-12% of donor chimerism which subsequently induced antigen-specific T-cell tolerance. Long-term donor chimerism is required but no studies have been done to quantify the minimum proportion of donor chimerism that may induce immune tolerance. Some studies support this finding that a low-level of donor chimerism might be sufficient to induce tolerance but there is no established threshold value. Cippa et al. induced tolerance with approximately 5% donor chimerism and they used Bcl-2 inhibitor ABT-737 and co-stimulatory blockade (anti-CD154 and CTLA4Ig) as recipient conditioning agents [241]. Transfer of proinsulin encoding BM prevents the development of autoimmune diabetes in NOD mice with 6-9% donor chimerism [4]. These data indicate, stable donor chimerism and continual antigen expression targeted to APC is the key for tolerance induction.

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Recovery of immune cells is very important after BMT as transplantation-related complications are associated with morbidity. CD45.2-SAP promoted rapid immune recovery compared to low-dose irradiation. CD45.2-SAP conditioned mice recovered the majority of their depleted peripheral blood leukocytes within two weeks post transplantation. B cells, myeloid cells and dendritic cells accumulated quickly after transplantation and this could be a consequence of large number of pre-mature B cells and early myeloid progenitor cells contained in bulk BM [269-271] and these transferred premature myeloid progenitors likely contribute to early chimerism. Whereas donor-derived T cells established slowly because donor progenitors take time to migrate into the thymus for subsequent T cell differentiation and selection before exiting to the periphery [272]. Such rapid peripheral blood leukocyte recovery limits the risks of anemia, neutropenia and thymopoietic damage. It is also predicted that rapid recovery might also minimize further transplant related co-morbidities such as nausea, anxiety and infertility [230, 273].

In addition, it is shown that transferring a large number of bulk BM cells accelerate donor chimerism and improves the tolerance induction timeframe. According to recent research, only 36% transferred donor bulk BM cells traffic to bone marrow whereas 8% migrate to spleen and 36% to liver [274]. Therefore, increasing the number of transferred BM cells help to traffic more donor BM cells to host BM and can compete more successfully with host HSC leading ultimately to higher levels of chimerism. The trend of the donor development suggests further increases in the number of donor cells transferred might further accelerate development of donor chimerism, but this would not be clinically feasible due to potential risks and challenges of getting large BM harvest from the donor.

All the experiments done in this chapter are related to autologous hematopoietic stem cell transplantation (auto-HSCT), where OVA expression was targeted to DC. Further investigation is required to assess the efficacy of CD45.2-SAP in OVA expression targeted to other APC or ubiquitous OVA expression. It is conceivable that immunotoxins prepared with specific HSC target agents might efficiently deplete HSC whilst completely preserving recipient immunity during conditioning. To deliver this potential conditioning approach to clinics, it is important to investigate its efficiency on human cells. The length of time for establishing a sufficient chimerism established here might be different to that which would occur in humans. If it took 26 weeks to get sufficient chimerism to induce tolerance this might be a problem for mice as 26 weeks is roughly one-third of their life expectancy but might not be an issue for humans. These data also indicate that chimerism might be life-long. But still it is not clear the minimum

113 | P a g e amount of donor chimerism required for inducing tolerance, so it is hard to predict the exact time when tolerance was induced without further studies. Further time course experiments are required to determine the minimum donor chimerism level for inducing tolerance. To test the efficacy of this approach in humans further experiments need to be done with immunotoxins prepared from human CD45 antibodies.

Taken together data presented in this chapter provide substantial evidence that using low-dose CD45.2-SAP enables improved tolerance induction by facilitating stable mixed chimerism of gene-modified BM whilst removing genotoxic components of recipient conditioning. The data also highlight that a low proportion of stable donor chimerism when generated using this approach is sufficient for immune tolerance. Similar protein-based immunotoxins that are specific for HSC/HSPC could be more beneficial to improve immune tolerance without affecting recipient peripheral leukocytes. Chapter 5 will focus on investigating HSPC-specific immunotoxins as conditioning approach for gene-engineered BMT.

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4.4. Summary and Future direction In summary, it is demonstrated that transplanting gene-modified BM encoding antigen targeted to DC induces immune tolerance under non-genotoxic conditioning. This non-genotoxic conditioning approach with low-dose CD45.2-SAP (0.5 mg/kg) permits tolerance induction though stable long-term donor hematopoietic chimerism. Thus gentler conditioning regime that does not cause DNA damage and have gentler effects on peripheral leukocytes could provide useful alternatives to low-dose irradiation for tolerance induction with gene modified BM.

This study also demonstrated that under this setting, gene-engineered BM takes over six months to engraft to a significant level that is sufficient for tolerance induction. However, it was improved by transplanting a higher number of BM cells. More specific recipient conditioning is the ultimate goal to treat established disease, so in order to improve the concept of DC-induced tolerance by engrafting gene-modified BM, it is important to develop HSPC specific immunotoxin which would not have any bystander effect on any other type of cells. This is investigated in Chapter 5.

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Chapter 5 Non-genotoxic conditioning using HSPC- specific immunotoxin facilitates tolerance induction

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5.1. Introduction

Chapter 3 and 4 demonstrated that low-dose CD45.2-SAP (0.5mg/kg) depleted 45-50% of LT- HSC from mouse bone marrow and permitted sufficient donor chimerism for induction of antigen-specific immune tolerance. However, conditioning with low-dose CD45.2-SAP does deplete leukocytes and therefore result in immune deficiency. To induce antigen-specific tolerance by HSC mediated gene therapy, improved and more-specific conditioning whereby HSC can be depleted specifically while preserving immunity would be optimal. c-kit is a receptor tyrosine protein kinase that binds stem cell factor (SCF), and is encoded by the kit gene [275]. It is only expressed on the surface of HSPC and some other cell types such as mast cells [276, 277]. Initially, anti-c-kit (CD117) antibody was used to achieve host HSC depletion which subsequently facilitated donor chimerism in various immunodeficient mouse models [248, 278, 279] but failed in immunocompetent mice [250]. However, combination of anti-c-kit with anti-CD47 or low-dose total body irradiation was successful and allowed donor chimerism in immunocompetent mice [227, 250]. As peripheral leukocytes do not express c- kit, saporin conjugated with anti-c-kit (c-kit-SAP) could be a promising approach to improve recipient conditioning without diminishing recipient immunity. A recent study showed in vivo administration of anti-c-kit-SAP (anti-2B8-SAP) depleted >99% of LT-HSC from mouse BM and permitted >90% donor chimerism, when used at a high dose (1.5mg/kg) [230]. But at high dose 2B8-SAP clears the stem cell niche in BM and therefore recipients can likely be susceptible to infection. Peripheral leukocytes remain present after 2B8-SAP treatment as it targets only c-kit expressing cells but if HSC transplantation fails then the patient can die. As low-level donor chimerism is sufficient to induce T-cell tolerance, complete host HSC depletion may not be necessary. Using low-dose 2B8-SAP might reduce the depletion effect on LT-HSC. So it is important to test whether use of low-dose 2B8-SAP that might preserve some LT-HSC is able to facilitate sufficient donor chimerism to induce tolerance.

The hypothesis for this chapter is “non-genotoxic conditioning through HSPC-specific immunotoxin c-kit-SAP (2B8-SAP) permits gene-modified donor BM engraftment and induction of antigen-specific T-cell tolerance.” In this chapter, I show that low-dose c-kit-SAP (0.5mg/kg) specifically depletes LT-HSC from mouse BM with minimal depletion effect on more differentiated hematopoietic progenitors. Low-dose c-kit-SAP also preserves peripheral leukocytes and allows donor chimerism sufficient to induce immune tolerance.

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5.2. Results

5.2.1. 2B8-SAP potently depletes endogenous long-term repopulating LT-HSC from mouse bone marrow

Low-dose CD45.2-SAP has been shown to facilitate antigen-specific T-cell tolerance through stable donor chimerism of gene-modified bone marrow (BM). However, 0.5mg/kg CD45.2- SAP showed a significant bystander effect on peripheral leukocytes. To avoid this immune depletion, experiments were designed to evaluate whether anti-c-kit-SAP mediates LT-HSC depletion. Anti-c-kit-SAP was prepared by conjugating biotinylated anti-c-kit (clone 2B8) with streptavidin-saporin. A control untargeted immunotoxin isotype control-SAP (ISO-SAP) was prepared by combining biotinylated anti-Rat IgG2b, κ (clone RTK4530) with streptavidin- saporin. PBS, ISO-SAP or 2B8-SAP was injected i.v. to C57BL/6 mice and the depletion effect was assessed 7 days post-injection (Figure 5.1A). A bead-based flow cytometric approach was used to assess LT-HSC (Lin-c-kit+Sca1+CD48-CD150+), HSPC (Lin-c-kit+), CLP (Lin-c- kit+Sca1+CD127+), CMP (Lin-c-kit+Sca1-CD127-CD34+CD16/32-), MP (Lin-c-kit+Sca1- CD127-), GMP (Lin-c-kit+Sca1-CD127-CD34+CD16/32+) and MEP (Lin-c-kit+Sca1-CD127- CD34-CD16/32-) content in BM and CD8+ T-cells, CD4+ T-cells, CD45R+ B cells, CD11b+ myeloid cells and CD11c+ DC content in spleen. This assay revealed that a single, low-dose of 2B8-SAP (0.5mg/kg) resulted in >90% depletion of phenotypic LT-HSC from BM (Figure 5.1B, C). Mice treated with ISO-SAP showed a significantly reduced LT-HSC number and approximately 20-25% of LT-HSC appeared to be depleted from BM (Figure 5.1 B, C). As HSPC express c-kit so it was expected they would also be depleted by 2B8-SAP treatment. 2B8-SAP reduced HSPC number by around 50-60% whereas ISO-SAP reduced approximately 20-25% of HSPC number (Figure 5.1D, E). PBS did not show any depletion effect both on LT-HSC and HSPC (Figure 5.1 B-E). These data indicate 2B8-SAP potently depletes LT-HSC and HSPC from mouse BM and LT-HSC are more affected than HSPC.

To assess the depletion effect of 2B8-SAP on lineage-committed hematopoietic progenitors, BM was harvested and analysed by a bead-based flow count assay. Surprisingly treatment with low-dose 2B8-SAP did not alter CLP (Figure 5.2A, B), MP (Figure 5.2C, D), MEP (Figure 5.3A, B), CMP (Figure 5.3C, D) numbers and only GMP number was changed. The GMP number increased by approximately 60% (Figure 5.3E, F). These data suggest low-dose 2B8- SAP effectively depletes LT-HSC without largely affecting lineage-committed progenitors.

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The major goal of using 2B8-SAP instead of CD45.2-SAP was to avoid immune depletion. As peripheral leukocytes do not express c-kit, it was proposed that they might not be depleted by 2B8-SAP treatment. To assess the depletion effect of 2B8-SAP on peripheral leukocytes, spleen was harvested 7 days after 2B8-SAP treatment. As expected, no depletion effect was observed for CD8+ T-cells (Figure 5.4A), CD4+ T-cells (Figure 5.4B), CD45R+ B cells (Figure 5.4C), CD11b+ myeloid cells (Figure 5.4 D) and CD11c+ DC (Figure 5.4 E). BM was also analysed to assess the effect on F4/80+Ly6G-VCAM1- macrophage, CD11b+Ly6G+granulocytes and F4/80+CD115+Ly6C+CD11b+ monocytes. Macrophage number remain unchanged (Figure 5.5A) whereas granulocytes and monocyte number increased >50% (Figure 5.5B, C). Hence, these data indicate 2B8-SAP preserves peripheral leukocytes.

To determine the effect of 2B8-SAP on different cells, peripheral blood, BM and spleen were harvested and absolute number was counted using a bead-based flow count assay. The total absolute number of peripheral blood cells dropped to 20% after ISO-SAP treatment and 30% after 2B8-SAP treatment (Figure 5.6A), BM number was unchanged by ISO-SAP treatment and slightly increased by 2B8-SAP treatment (Figure 5.6B). Spleen number remain unchanged by ISO-SAP and 2B8-SAP treatment (Figure 5.6C).

Taken together, these data suggest that in vivo administration of low-dose 2B8-SAP (0.5mg/kg) deplete a large proportion of LT-HSC without largely depleting lineage-committed hematopoietic progenitors.

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2B8-SAP A Single dose, IV C57BL/6

Day 0 Day 7 Assess HSC, HSPC and myeloid derived cell depletion in BM and leukocyte depletion in spleen

B C **** **** * **** 10 ns 100

8 75 **** 6 50 * 25 4 0 2 -25

0 HSC depletion HSC control) of (% l HSC Absolute Numbers (X10^-3) Numbers Absolute HSC i S P P S P P N B A A B A A P -S -S P -S -S O 8 O 8 B B IS 2 IS 2

D E

*** **** 100 2500 ** *** 75 2000 * ns * 50 ns 1500 ns 25 1000 0 500 -25

0 l control) of (% depletion HSPC P i S P P S P (X10^-3) Numbers Absolute HSPC N A A B A A B S S P -S -S P - - O 8 O 8 B S B IS 2 I 2

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Figure 5.1: Single i.v. administration of 2B8-SAP (0.5mg/kg) potently depletes long-term repopulating hematopoietic stem cells (LT-HSC) and hematopoietic stem and progenitor cells (HSPC). Experimental outline for assessing 2B8-SAP mediated LT-HSC and HSPC depletion in C57BL/6 mice (A). Low-dose 2B8-SAP (0.5mg/kg), ISO-SAP and PBS was administered i.v. and depletion was assessed through flow cytometry 7 days later. Flow count assay was performed to calculate absolute LT-HSC (Lin-c-kit+CD48-CD150+) number (B) and depletion percentage was also calculated (C). Depletion effect on HSPC (Lin-c-kit+) was also analysed (D, E). Data are pooled from 2 individual experiments and show individual mice with mean ± SD, n=5 mice per group. Statistical significance determined by one-way ANOVA, ****p<0.0001, ***p<0.001, **p<0.01, *p<0.05 and ns= no significance.

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A B

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400 50 ns 300 ns 0 200

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CLP Numbers CLP (X10^-3)

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MP Numbers MP (X10^-3)

0 depletion MP control) of (% -100 l i S P P S P P N B A A B A A P -S -S P -S -S O 8 O 8 S B S B I 2 I 2

Figure 5.2: Low-dose 2B8-SAP (0.5mg/kg) does not change CLP and MP numbers. Low- dose 2B8-SAP, ISO-SAP and PBS was injected i.v. and absolute numbers and depletion effect of common lymphoid progenitors (A, B) and myeloid progenitors (C, D) were analysed 7 days post 2B8-SAP treatment. Common lymphoid progenitors were defined as (Lin-c- kit+Sca1+CD127+) and myeloid progenitors were defined as (Lin-c-kit+Sca1-CD127-). Data show individual mice with mean ± SD, n=5 mice per group and pooled from 2 individuals experiments. Statistical significance determined by one-way ANOVA, ****p<0.0001, ***p<0.001, **p<0.01, *p<0.05 and ns= no significance.

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A B

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Figure 5.3: Low-dose 2B8-SAP (0.5mg/kg) does not deplete lineage-committed hematopoietic progenitors except GMP. 0.5 mg/kg 2B8-SAP, ISO-SAP and PBS was administered i.v. and absolute numbers and depletion effect was calculated 7 days after 2B8- SAP treatment. Bead based flow count assay was performed to analyse MEP=Lin-c-kit+Sca1- CD127-CD34-CD16/32- (A, B), CMP=Lin-c-kit+Sca1-CD127-CD34+CD16/32- (C, D), GMP=Lin-c-kit+Sca1-CD127-CD34+CD16/32+ (E, F). Data are pooled from 2 individual experiments and represent the individual mice with mean ± SD, n=5 mice per group. Statistical significance determined by one-way ANOVA, ****p<0.0001, ***p<0.001, **p<0.01, *p<0.05 and ns= no significance.

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A B

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-50 -200 B cell B depletion of (% control) S P P P P B A A S A A S S control) cell Myeloid of (% depletion B P - - P -S -S O 8 O 8 S B B I 2 IS 2 E

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DC depletion control) DC of (% -100

S P P B A A P -S -S O 8 B IS 2

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Figure 5.4: Low-dose 2B8-SAP (0.5mg/kg) preserves peripheral leukocytes. 2B8-SAP was injected i.v. and spleen was harvested to analyse the depletion effect on different peripheral leukocytes. CD8+ (A), CD4+ (B), CD45R+ B cells (C), CD11b+ myeloid cells (D) and CD11c+ DC (E) were analysed. Data are pooled from 2 individual experiments and represent the individual mice with mean ± SD, n=5 mice per group. Statistical significance determined by one-way ANOVA, ****p<0.0001, ***p<0.001, **p<0.01, *p<0.05 and ns= no significance.

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A B

100 100 *

50 ns 50 * ns

0 0

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-100 -100 P P S P P S A A A A B S S B S S Granulocytedepletion control) of (% P - - Macrophagedepletion control) of (% P - - 8 O 8 O B IS B IS 2 2

C

100 ****

*** 50 *

0

-50

-100

S P P Monocytedepletion control) of (% B A A P -S -S O 8 B IS 2

Figure 5.5: Granulocyte and monocyte number increases after 2B8-SAP treatment but macrophage number remain unchanged. 2B8-SAP (0.5mg/kg) was administered i.v. and BM was harvested 7 days post 2B8-SAP administration. Depletion effect of 2B8-SAP on F4/80+Ly6G-VCAM1- macrophage (A), CD11b+Ly6G+granulocytes (B) and F4/80+CD115+Ly6C+CD11b+ monocytes (C) were analysed by flow cytometry. Data show individual mice with mean ± SD, n=5 mice per group and pooled from 2 individuals experiments. Statistical significance determined by one-way ANOVA, ****p<0.0001, ***p<0.001, **p<0.01, *p<0.05 and ns= no significance.

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A

10

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0 l i S P P N B A A P -S -S O 8 Total Total cellconc. in (cells/ul blood X10^-3) B IS 2

B C

** 100 *** 100 80 ns 80 ns ns 60 60 * 40 ns 40

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0 0 (X10^-6) number absolute BM l i S P P il P P N Spleenabsolute number (X10^-6) S B A A N B A A P -S -S P -S -S O 8 O 8 B B IS 2 IS 2

Figure 5.6: 2B8-SAP (0.5mg/kg) reduces the total leukocyte concentration in blood. 2B8- SAP was administered i.v. in C57BL/6 mice and absolute number was assessed through flow cytometry 7 days post 2B8-SAP treatment. Total cell number was assessed in blood (A), bone marrow (B) and spleen (C). Data show individual mice with mean ± SD, n=3-5 mice per group and pooled from 2 individual experiments for bone marrow and spleen and a single experiment for blood. Statistical significance determined by one-way ANOVA, ****p<0.0001, ***p<0.001, **p<0.01, *p<0.05 and ns= no significance.

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5.2.2. 2B8-SAP facilitates donor chimerism which subsequently induces antigen-specific T- cell tolerance

It was next determined whether LT-HSC depletion achieved by low-dose 2B8-SAP (0.5mg/kg) conditioning facilitated sufficient donor chimerism of gene-modified BM for tolerance induction. Consistent with previous experiments conducted with low-dose CD45.2-SAP conditioning, 10x106 bulk BM from OVA expressing transgenic 11c.45.1 and non-transgenic littermates was transferred to 2B8-SAP conditioned C57BL/6 mice. Untreated recipients were used as negative controls for chimerism and tolerance whereas 300cGy treated recipients were used as positive controls. Cell depletion after 2B8-SAP treatment was assessed through submandibular bleeding one day prior to BM transfer. Donor-derived leukocyte development was tracked in blood at weeks 2, 4, 6, 8 and 9 after BM transfer. Donor chimerism was also determined in spleen and BM at week 9 after BM transfer. To test immune tolerance mice, were immunized with OVA/QuilA 8 weeks after BM cell transfer and OVA responsiveness was tested by IFN-γ ELISPOT (Figure 5.7A).

Peripheral blood analysis after 2B8-SAP treatment and prior to transferring BM indicated the efficacy of 2B8-SAP. Total blood leukocyte number reduced by approximately 20-30%, 6 days after 2B8-SAP treatment (Figure 5.7B). 300cGy radiation reduced over 60% of the peripheral leukocytes from blood (Figure 5.7C). Peripheral blood concentration was tracked over the experiment at the indicated time points and 2B8-SAP conditioned mice recovered over 90% of their leukocyte count within 2 weeks whereas peripheral blood leukocyte levels in 300cGy treated did not fully return to pre-treatment levels (Figure 5.7C).

Using this recipient conditioning setting of low-dose 2B8-SAP (0.5mg/kg), 20-25% donor chimerism was observed within peripheral blood at week 8 after transfer. This level of donor chimerism was also retained one week after immunization (Figure 5.8A, B). Untreated recipients failed to establish detectable chimerism, whereas 300cGy facilitated over 50% donor chimerism (Figure 5.8 A). 2B8-SAP also facilitated approximately 25-30% donor chimerism in spleen at week 9 after BM transfer (Figure 5.8C). To assess the development rate of donor derived leukocyte lineages, donor derived myeloid cells, DC, CD8+ T-cells, CD4+ T-cells and B cells were tracked in peripheral blood over the indicated time points. Donor derived myeloid cells, DC and B cells appeared in 2B8-SAP-conditioned recipients quickly after BM cell transfer and gradually reached to over 25% at week 6 (Figure 5.9 A-C). In contrast, donor

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CD8+ T-cells and CD4+ T-cells accumulated slowly, were detected at week 4 after BM transfer and were established to around 20-25% at week 9 (Figure 5.9D, E).

To compare the donor chimerism in blood and spleen with BM, BM was harvested at the final time point (week 9) and donor chimerism assessed. BM analysis revealed 20-30% donor chimerism (Figure 5.10A) and the proportion was similar to that observed in blood and spleen indicating donor derived leukocyte lineages found in peripheral blood or spleen was actually derived from engrafted donor BM. Both transgenic (Tg) 11c45.1 and non-transgenic (non-Tg) littermate engrafted equally. And as expected untreated recipients failed to generate detectable donor chimerism and 300cGy established over 45% donor chimerism in BM (Figure 5.10A). 20-30% donor HSPC chimerism (Lin-/c-kit+) was observed in 2B8-SAP conditioned Tg and non-Tg 11c45.1 BM transferred groups whereas 300cGy facilitated approximately 45-50% donor HSPC chimerism (Figure 5.10B).

To test tolerance induction, mice were immunized with OVA/QuilA at week 8 after BM cell transfer and OVA257-264 responsiveness was tested by IFN-γ ELISPOT. In no-BMT control recipients and untreated control recipients of Tg 11c45.1 BM, IFN-γ ELISPOT demonstrated an increase in number of OVA257-264 induced IFN-γ spot forming units (SFU) compared to the tolerance control Tg-11c.45.1 mice. A similar SFU number was observed in OVA immunized 2B8-SAP conditioned recipients with no BMT control groups (Figure 5.11A). However, IFN-

γ ELISPOT analysis displayed no OVA257-264 specific responsiveness in 2B8-SAP conditioned recipients of OVA encoding BM whereas recipients of non-OVA encoding BM showed increased IFN-γ response to OVA257-264 relative to Tg-11c.45.1 tolerance control groups (Figure 5.11A). 300cGy treated recipients got Tg 11c.45.1 BM also failed to generate IFN-γ response to OVA257-264 (Figure 5.11A). Comparing the OVA257-264 induced IFN-γ spots between untreated and 2B8-SAP treated recipients of Tg 11c.45.1 BM indicates stable donor chimerism is the key to induce antigen-specific T-cell tolerance in this setting.

Finally, these data suggest low-dose 2B8-SAP (0.5mg/kg) facilitates antigen-specific T-cell tolerance by enabling stable donor chimerism of gene-modified BM.

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Untreated/2B8-SAP treated A C57BL/6 C57BL/6 2B8-SAP

Day -7 Day 0 Week 8 Week 8 Week 9 Day -1 Week 2 Week 4 Week 6 Blood Blood Blood Blood Blood Mice immunisation Blood Spleen Bone marrow Donors Non-Tg/Tg-11c.45.1

B

15

ns **** 10

ns

5

0

T T T T T M M M M M B B B B B o .1 o .1 .1 N 5 N 5 5 Totalconc. ell blood (X10-^3 Cells/ul) .4 .4 .4 c c c 1 1 1 -1 1 -1 g g g T -T T n o N C No Rx 2B8-SAP

15 No BMT/No Rx Tg-11c.45.1 BMT/No Rx 10 No BMT/2B8-SAP Non-Tg 11c.45.1 BMT/2B8-SAP Tg-11c.45.1 BMT/2B8-SAP 5 Tg-11c.45.1 BMT/300cGY

0

Cellconc. (X10-^3 Cells/ul) 0 2 4 6 8 9 Weeks post BMT

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Figure 5.7: Low-dose 2B8-SAP allows quick total cell recovery. 10x106 bulk BM from Tg 11c.45.1 and non-Tg littermate was transferred to untreated, 2B8-SAP treated and 300 cGy irradiated C57BL/6 recipients (A). Blood was collected through submandibular bleeding 6 days post 2B8-SAP treatment. Peripheral blood leukocyte concentration was measured in peripheral blood (B). Cell recovery was tracked over the indicated time period (C). Data are pooled from 2 individual experiments and represent the individual mice with mean ± SD, n=6 mice per group. Statistical significance determined by one-way ANOVA, ****p<0.0001, ***p<0.001, **p<0.01, *p<0.05 and ns= no significance. No Rx = No treatment.

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A

60 No BMT/No Rx Tg-11c.45.1 BMT/No Rx 40 No BMT/2B8-SAP Non-Tg 11c.45.1 BMT/2B8-SAP

Tg-11c.45.1 BMT/2B8-SAP 20 Tg-11c.45.1 BMT/300cGY

chimerism Donor (%) 0 2 4 6 8 9

Weeks post BMT

B C

**** **** **** 60 60 **** **** ns **** 40 40 ns

20 20 ns ns

0 0 T T T T T T

T T T T T T M M M M M M Donor chimerism in spleenDonorchimerism in (%) Donor chimerism in boold (%)booldDonor chimerism in M M M M M M B B B B B B B B B B B B o .1 o .1 .1 .1 o .1 o .1 .1 .1 N 5 N 5 5 5 N 5 N 5 5 5 .4 .4 .4 .4 .4 .4 .4 .4 c c c c c c c c 1 1 1 1 1 1 1 1 -1 1 -1 -1 -1 1 -1 -1 g g g g g g g g T -T T T T -T T T n n o o N N No Rx 2B8-SAP 300cGy No Rx 2B8-SAP 300cGy

Figure 5.8: 2B8-SAP facilitates significant donor chimerism. BM from non-Tg 11c.45.1 and Tg 11c.45.1 (CD45.1+) mice was transferred to untreated, 2B8-SAP (0.5 mg/kg) treated and 300 cGy irradiated C57BL/6 recipients (CD45.2+). Donor-derived leukocyte development was tracked over the indicated time periods (A). Donor chimerism in blood (B) and spleen (C) was measured in week 9 post transplantation. Data represent individual mice with mean ± SD and pooled from 2 separate experiments, n=6 mice per group. Statistical significance determined by one-way ANOVA, ****p<0.0001, ***p<0.001, **p<0.01, *p<0.05 and ns= no significance.

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A B No BMT/No Rx No BMT/No Rx Tg-11c.45.1 BMT/No Rx Tg-11c.45.1 BMT/No Rx No BMT/2B8-SAP 80 No BMT/2B8-SAP 80 Non-Tg 11c.45.1 BMT/2B8-SAP Non-Tg 11c.45.1 BMT/2B8-SAP Tg-11c.45.1 BMT/2B8-SAP Tg-11c.45.1 BMT/2B8-SAP Tg-11c.45.1 BMT/300cGY 60 Tg-11c.45.1 BMT/300cGY 60

40 40

20 20

0 0

(%) chimerism DC Donor 2 4 6 8 9 2 4 6 8 9 (%) chimerism myeloid Donor Weeks post BMT Weeks post BMT

C D

No BMT/No Rx No BMT/No Rx 80 80 Tg 11c.45.1/No Rx Tg-11c.45.1 BMT/No Rx No BMT/2B8-SAP No BMT/2B8-SAP Non-Tg 11c.45.1 BMT/2B8-SAP Non-Tg 11c.45.1 BMT/2B8-SAP Tg-11c.45.1 BMT/2B8-SAP 60 60 Tg-11c.45.1 BMT/2B8-SAP Tg-11c.45.1 BMT/300cGY Tg-11c.45.1 BMT/300cGY 40 40

20 20

0 0

DonorCD8+ chimerism (%) 2 4 6 8 9 Donor(%)cell B chimerism 2 4 6 8 9 Weeks post BMT Weeks post BMT

E

No BMT/No Rx Tg-11c.45.1 BMT/No Rx 80 No BMT/2B8-SAP Non-Tg 11c.45.1 BMT/2B8-SAP Tg-11c.45.1 BMT/2B8-SAP 60 Tg-11c.45.1 BMT/300cGY

40

20

0

DonorCD4+ chimerism (%) 2 4 6 8 9 Weeks post BMT

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Figure 5.9: 2B8-SAP facilitates donor-derived leukocyte lineage chimerism. 10x106 bulk BM cells from Non-Tg 11c.45.1 and Tg 11c.45.1 mice were transferred into untreated, low- dose 2B8-SAP treated and lightly irradiated C57BL/6 mice. Donor derived leukocytes lineages such as myeloid cell (A), DC (B), B cells (C), CD8+ T-cells (D) and CD4+ T-cells (E) was tracked over the time in peripheral blood. Data are pooled from two individual experiments and show individual mice with mean ± SD, n=6 mice per group.

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A B

**** 60 **** **** 60 **** **** ns 40 **** ns 40

20 20 ns ns 0 0 BMDonorchimerism in (%) T T T T T T M M M M M M T T T T T T B B B B B B M M M M M M o .1 o .1 .1 .1 B B B B B B N 5 N 5 5 5 (%) BM in chimerism HSPC Donor o .1 o .1 .1 .1 .4 .4 .4 .4 N 5 N 5 5 5 c c c c .4 .4 .4 .4 1 1 1 1 c c c c -1 1 -1 -1 1 1 1 1 g g g g -1 1 -1 -1 T -T T T g g g g n T -T T T n o o N N

No Rx 2B8-SAP 300cGy No Rx 2B8-SAP 300cGy

Figure 5.10: 2B8-SAP allows donor HSPC chimerism. 10x106 BM from Non-Tg 11c.45.1 and Tg 11c.45.1 mice was transferred to untreated, 2B8-SAP (0.5 mg/kg) treated and 300 cGy irradiated C57BL/6 recipients. Overall donor chimerism in bone marrow was analysed 9 weeks post transplantation. Flow cytometry was done to analyse donor chimerism in bone marrow (A) and HSPC (Lin-c-kit+) was also analysed. Data are pooled from 2 individual experiments and show individual mice with mean ± SD, n=6 mice per group. Statistical significance determined by one-way ANOVA, ****p<0.0001, ***p<0.001, **p<0.01, *p<0.05 and ns= no significance.

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A

**** **** 800 ****

** **** SPC

5 600

400

200 SFU/1.25×10  ns

IFN- 0 T T T T T T e M M M M M M ic B B B B B B M o .1 o .1 .1 .1 .1 N 5 N 5 5 5 5 .4 .4 .4 .4 .4 c c c c c 1 1 1 1 1 1 1 1 1 1 g g g g g T -T T T T n o N No Rx 2B8-SAP 300cGY

Figure 5.11: 2B8-SAP mediates significant donor chimerism and subsequently induce T- cell tolerance. 8 weeks post cell transplantation, mice were immunized with OVA/QuilA and immune response to OVA257-264 was assessed by IFNγ-ELISPOT (A). Data are pooled from 2 individual experiments and show individual mice with mean ± SD, n=6 mice per group. Statistical significance determined by one-way ANOVA, ****p<0.0001, ***p<0.001, **p<0.01, *p<0.05 and ns= no significance.

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5.3. Discussion

Current recipient conditioning used clinically for HSC-mediated gene therapy involves non- specific chemotherapy, that does not target only HSC, either alone and/or in combination with total body irradiation which is genotoxic and cytotoxic is not cell specific and causes extensive bystander damage to healthy cells [252, 280]. Even at lower doses (non-myeloablative) these conditioning regimes induce neutropenia, thrombocytopenia, lymphocytopenia and non- specific organ toxicities [281]. Red blood cells (RBC) and platelet transfusion can alleviate anemia and thrombocytopenia [252] but not the lymphocytopenia and neutropenia that underlies patient susceptibility to severe infection [282, 283]. Additionally, the genotoxic nature of these conditioning approaches can permanently damage DNA which can lead to cancer [252]. Emerging immunotoxin-based recipient conditioning is expected to reduce non- specific toxicity compared to traditional myeloablative conditioning approaches. It is demonstrated in Chapter 4 and 5 that HSC depletion by HSPC non-specific low-dose CD45.2- SAP facilitates long-term donor chimerism of gene-engineered BM which subsequently induces antigen-specific T-cell tolerance. But still, this HSPC non-specific conditioning depletes leukocytes that might reduce immunity. This is despite the rapid rate at which depleted peripheral blood leukocytes are reinstated after treatment. Replacing genotoxic and non- specific recipient conditioning with more improved HSC-specific conditioning is a major medical need in HSC-mediated gene therapy. Here, this study shows antigen-specific tolerance induction through long-term donor chimerism can be achieved by using a HSPC-specific conditioning approach.

In this chapter, it is shown that low-dose 2B8-SAP (0.5mg/kg) achieved a high level of LT- HSC depletion from mouse BM. 2B8-SAP targets c-kit expressing LT-HSC and HSPC which should reduce the risk of collateral damage to other cell types that do not express c-kit. Though both LT-HSC and HSPC express c-kit, surprisingly, LT-HSC are preferentially targeted and depleted by 2B8-SAP over HSPC. There would be at least 4 possibilities to explain this outcome, i) different expression level of c-kit on the surface of HSC and HSPC, ii) variation of internalization kinetics, iii) differential processing of 2B8-SAP by specific cell types iv) or HSC might be more sensitive to 2B8-SAP than HSPC. However, further studies are required to explore these possibilities. As expected, no peripheral leukocyte depletion was observed in spleen as peripheral leukocytes do not express c-kit on their surface. Unlike low-dose CD45.2- SAP, 2B8-SAP treatment did not lead to an increase in myeloid or macrophage numbers nor did 2B8-SAP change total spleen cellularity. Surprisingly, total blood leukocyte concentration

138 | P a g e goes down after 2B8-SAP treatment but it comes back very quickly after transplantation compared to 300cGy irradiated groups. Interestingly total BM cellularity slightly goes up after 2B8-SAP treatment. This surprising and interesting effect can be explained by peripheral blood cell migration. Possibly, 2B8-SAP treatment induces peripheral blood cells to migrate to peripheral organs or BM. Because the reduction in peripheral blood count was reversed very quickly, and if this reduction was phenotypic depletion then it should have taken a while to restore its depleted cells. Further experimentation is required to explore this possibilities.

This study also demonstrates, low-dose 2B8-SAP enables a level of donor chimerism of gene- engineered BM sufficient to induce immune tolerance. Low-dose 2B8-SAP also facilitates more rapid development of donor chimerism than low-dose CD45.2-SAP, because 2B8-SAP depleted more LT-HSC than CD45.2-SAP. Therefore, the competition between transferred donor LT-HSC and remaining host LT-HSC would be reduced compared to CD45.2-SAP- mediated conditioning. As expected myeloid cells, DC and B cells accumulated quickly after transplantation as transferred donor bulk BM cells contained large number of premature myeloid progenitors and B-cells. Donor derived T-cell development was slower because T-cell differentiation and selection takes longer than other peripheral leukocytes. Similar to low-dose total body irradiation 2B8-SAP enables donor chimerism but without inducing neutropenia and lymphopenia.

These data show that a single injection of 2B8-SAP (0.5mg/kg) can facilitate antigen-specific T-cell tolerance in mouse model. This is also anticipated that conditioning with 2B8-SAP not only facilitates robust long-term donor chimerism but is also non-toxic to other cell types. To translate this promising conditioning approach to hospital, it must be effective on human cells. According to these data, I anticipate 2B8-SAP mediated recipient conditioning will be broadly used in experimental HSC-mediated gene therapy and possibly it could be a potential candidate for human HSC-mediated gene therapy.

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5.4. Summary

In summary, it is established that low-dose 2B8-SAP (0.5mg/kg) effectively depletes a significant proportion of LT-HSC from BM and has preferential effects on HSC over HSPC. It is also demonstrated that transferring gene-modified BM encoding antigen targeted to DC induces immune tolerance under 2B8-SAP-mediated non-genotoxic conditioning. This gentler conditioning regimen, which would not cause DNA damage and showed limited effects on peripheral leukocyte homeostasis could provide a powerful alternative to toxic conditioning approaches for tolerance induction with gene-engineered BM.

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Chapter 6 Concluding Remarks

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Autoimmune diseases result from dysregulated immune response to native tissue components and CD8+ T-cells can be key contributors for this autoimmune process [284]. Existing therapeutic approaches have caveats and limitations and rarely achieve sustained remission [24, 285]. An important strategy for alleviating these conditions is the development of effective immunotherapies that can turn-off the dysregulated immune responses that underlie these conditions. Currently bone marrow transplantation (BMT)/hematopoietic stem cell transplantation (HSCT) is an indispensable therapeutic option not only for irredeemable blood disorders such as severe haemolytic anaemia and aplastic anaemia but also malignant haematological disorders. HSCT has also been adapted to treat autoimmune diseases. Autologous HSCT have alleviated disease in experimental models of autoimmune disease and are currently in clinical trials for type 1 diabetes, rheumatoid arthritis and multiple sclerosis [25, 26, 237]. To date, clinical outcomes of autologous HSCT in rheumatoid arthritis and multiple sclerosis are encouraging but relapse rates are high due to re-emergence of pathogenic immune cells or their incomplete depletion during pre-HSCT conditioning [286, 287]. Active tolerance induction through antigen-specific approaches has the potential to overcome such disease relapse [103]. Enforced expression of antigen in DC/APC ‘turns off’ effector and memory T cell responses [73-75] and could be a major therapeutic approach to treat autoimmune diseases driven by pathogenic T-cells. Several groups have utilized this strategy in mouse models of autoimmune diseases [4, 113, 173, 180, 288]. To achieve long-term tolerance it is likely that gene-engineered HSC-encoded antigen expression will need to be life- long and consequently long term HSC engraftment is required to ensure a continuous supply of antigen expressing DC/APC in the periphery [244]. Tolerance induction after gene- engineered HSC transfer and engraftment is attained after myeloablative or non-myeloablative HSCT conditioning [243], but these are typically genotoxic, non-specifically immune suppressive and can cause lifelong complications [247]. Developing a non-genotoxic conditioning to promote engraftment of transferred HSC is clinically important because it would be safer and more targeted than currently used toxic conditioning. This thesis examined the potential of non-genotoxic targeted immunotoxin based conditioning to induce antigen- specific T-cell tolerance in a mouse model.

The overall findings of this thesis support the potential of the non-genotoxic conditioning approach with existing HSC mediated gene-therapy strategies to induce antigen-specific T-cell tolerance. Data presented in Chapter 3 demonstrated that a leukocyte-depleting non-genotoxic immunotoxin (CD45.2-SAP) preferentially depletes LT-HSC without largely affecting

142 | P a g e peripheral leukocytes when used at low doses. This knowledge was applied in Chapter 4 and low dose CD45.2-SAP (0.5mg/kg) used as recipient conditioning and it was shown that transplanting BM encoding OVA targeted to DC induced OVA-specific T-cell tolerance 26 weeks after BMT. 11c.OVA mouse was used as donor which express CD45.1 allelic variant to avoid residual effect of CD45.2-SAP. However, donor chimerism was improved by transplanting a higher number of BM cells. The ultimate aim is to replace genotoxic conditioning with some form of LT-HSC-specific immunotoxin and while progressing this, this data suggest some caveats. Although LT-HSC have self-renewal and differentiation capability they are typically quiescent or dormant, which protects them from cellular insults and functional exhaustion. This fundamental property of LT-HSC facilitates life-long hematopoietic cell production [289]. It would be interesting to further define whether SAP- based immunotoxins that induce cell death by inhibiting protein synthesis are most effective for LT-HSC depletion because they induce cell cycle independent cell death [290], in contrast to other agents for example anti-mitotic small molecule-antibody conjugates [291]. A previous publication reported that CD45 has poor cell internalization capability and therefore might not be suitable for targeted immunotoxin [292]. But, the current study found the loss of surface bound CD45 after incubation indicated partial internalization and also found significant LT- HSC and peripheral leukocyte depletion by CD45.2-SAP, therefore validated its use. CD45 is expressed both in peripheral leukocytes and LT-HSC. CD45 expressed at 200,000 molecules per leukocytes [293], so it is probable that not only the frequency of internalization but also the absolute number of internalized molecules determine target suitability. It remains unclear why CD45.2-SAP preferentially targets LT-HSC for depletion over peripheral leukocytes. An ex vivo expression and internalization experiment was performed and described in Chapter 3 but the data was not conclusive. Further experimentation might uncover the mechanism which could include cell specific differential processing or depleting sensitivity. Finally, Chapter 5 demonstrated a HSPC-specific immunotoxin (2B8-SAP) depletes LT-HSC and creates space for donor exogenous HSC without affecting peripheral leukocytes. It is also demonstrated that transferring antigen encoding BM under 2B8-SAP conditioning facilitated donor chimerism which subsequently induced immune tolerance.

The data presented in this thesis open several avenues to explore in future. In Chapter 3 it was observed that low-dose CD45.2-SAP (0.5mg/kg) showed preferential effect on LT-HSC. Myeloid cells and myeloid progenitor cell-derived population such as monocytes, granulocytes and macrophage were increased after CD45.2-SAP treatment. So, it is also worth investigating

143 | P a g e why myeloid cell number is increasing after CD45.2-SAP treatment. In Chapter 4, the BM transferred to CD45.2-SAP conditioned recipients encoded OVA targeted to DC only. So, BM encoding OVA targeted to other APC or that encode OVA ubiquitously could be investigated in similar conditioning settings. Moreover, experiments described in Chapter 4 were performed in naïve mice where recipients were not exposed to any antigen prior to conditioning. So, BM transfer to CD45.2-SAP conditioned primed mice where recipient mice have been immunized with antigen prior to conditioning needs to be investigated to progress this tolerance induction protocol toward clinical settings. The observed HSPC-specific effects of 2B8-SAP described in Chapter 5 can also be further optimized. LT-HSC and lineage committed hematopoietic progenitors express c-kit although the expression level and internalization frequency is unknown so it is hard to predict why 2B8-SAP affected LT-HSC more than HSPC. That may be due to differences in expression level, variation in internalization kinetics or due to differential 2B8-SAP processing by specific cell types. However, further experimentation is required to understand this surprising effect. The real challenge of this study is to translate its findings in clinical settings. Before starting clinical trials three things will need to be considered. Firstly, test the effectiveness of CD45.2-SAP/2B8-SAP in memory settings. It is very crucial to investigate tolerance induction though transfer of BM encoding OVA targeted to DC/APC or encoding ubiquitously-expressed OVA to CD45.2-SAP/2B8-SAP conditioned primed recipients. Secondly, to deliver this potential conditioning approach to clinics, it is important to investigate its efficiency on human cells. So, further experiments need to be done with immunotoxins prepared from human CD45/2B8 antibodies and test them on humanized mice. Finally, progressing with a clinical perspective with this strategy, transfer of virally- transduced BM encoding a disease relevant antigen controlled by CD11c promoter need to be investigated in autoimmune mouse models to support the data presented in this thesis. Demonstrating effective prevention and therapy in disease relevant mouse models will provide the necessary evidence for moving forward with antigen encoding auto-HSCT to human clinical trials.

The key strength of this thesis is inducing antigen-specific T-cell tolerance by using less-toxic recipient conditioning which spares the majority of hematopoietic progenitor cells and peripheral leukocytes. It is established that low-level long-term donor chimerism is sufficient to induce tolerance [4, 241] although the minimum required donor chimerism level is still unknown. So it is also unclear how much niche space is required for donor transferred cells inside recipient BM to develop sufficient donor chimerism to induce tolerance. Previous

144 | P a g e research indicated specific host LT-HSC depletion is a critical requirement to enable donor derived cell development and that can attained through LT-HSC targeting approaches [248]. This study highlighted long-term donor chimerism can be attained through specific LT-HSC depletion. Besides, donor-derived cell develops without affecting recipient myeloid cells and hematopoietic progenitor cells, highlighting a role for BM transplantation in a non-genotoxic context [248]. Previous studies also used antibody-based conditioning to create niche space inside recipient BM by specifically depleting LT-HSC. For example, administration of an anti- c-kit antibody (ACK2) depletes a significant number of LT-HSC from BM niches and permits successful donor HSC engraftment in immunocompromised mice [226]. Another report showed the combined approach of ACK2 and CD47 blockade can deplete over 99% of recipient LT-HSC and permits a high level donor chimerism [227]. These approaches showed encouraging results in immunocompromised mice but failed to established significant chimerism in immunocompetent mice. To overcome this challenge saporin based immunotoxin is emerging. Here, this thesis demonstrated that single low-dose CD45.2-SAP and 2B8-SAP facilitated sufficient donor chimerism in immunocompetent mice C57BL/6J while largely preserving peripheral leukocytes. 2B8-SAP also preserve intact immunity as no depletion effect was observed on peripheral leukocytes. Total blood cell concentration was slightly decreased after treatment but also recovered very quickly indicated no neutropenia, anemia or lymphopenia. It is also expected that this leukocyte preservation might reduce additional co- morbidities relevant to traditional genotoxic conditioning. Furthermore, it facilitated sufficient amount of gene-engineered donor chimerism to induce T-cell tolerance in 8 weeks. This non- genotoxic conditioning might be very attractive in auto-HSC mediated gene therapy to induce immune tolerance.

All the experiments done in this thesis are related to autologous hematopoietic stem cell transplantation (auto-HSCT), further investigation is required to assess the efficiency of CD45.2-SAP in allogenic hematopoietic stem cell transplantation (allo-HSCT). The major challenge of allogenic transplantation is graft versus host disease (GVHD) where recipient immune system reject donor graft, hence transient immune suppression is required. However, CD45.2-SAP conditioning can provide advantages over conventional conditioning by depleting only CD45 expressing immune cells. Moreover, to reduce genotoxicity and reduce tissue inflammation, CD45.2-SAP might permit use of T-cell depleted grafts which would result in considerably reduced GVHD [294]. This strategy can also permit rapid immune recovery which is a main concern with manipulated donor grafts and additionally enable

145 | P a g e limited immune suppression with reduced infection complications. Additionally, in complementary studies with combined immunotoxin (CD45.2-SAP/2B8-SAP) and transient immune-suppressive (i.e. rapamycin, cyclosporine, ABT-737) recipient conditioning approach can permit human leukocyte antigen (HLA) mismatched transplantation and simultaneous tolerance induction likely due to long-term donor HSC chimerism permitting BM transplantation with various donors.

The data presented in this thesis provide substantial evidence that single low-dose CD45.2- SAP and 2B8-SAP improved tolerance induction protocol without the help of conventional genotoxic conditioning in mice model and might be an attractive choice for humans. To transfer this potential conditioning strategies to the clinic, its efficacy on human cells need to be investigated. Biotinylated antibody-streptavidin-saporin conjugation provides a proof of concept but development and optimization of anti-human CD45-SAP or anti-human 2B8-SAP is required to test this phenomenon against human cells.

In conclusion, this thesis established a reliable targeted immunotoxin based conditioning strategy to induce antigen-specific T-cell tolerance that might be used to reverse autoimmune disease state. This study advanced the conventional conditioning strategies to induce T-cell tolerance and provide potential prospects to explore in clinical trials.

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Chapter 7 References

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1. Katz, U., Y. Shoenfeld, and G. Zandman-Goddard, Update on intravenous immunoglobulins (IVIg) mechanisms of action and off- label use in autoimmune diseases. Curr Pharm Des, 2011. 17(29): p. 3166-75. 2. Cotsapas, C. and D.A. Hafler, Immune-mediated disease genetics: the shared basis of pathogenesis. Trends Immunol, 2013. 34(1): p. 22-6. 3. Walsh, S.J. and L.M. Rau, Autoimmune diseases: a leading cause of death among young and middle-aged women in the United States. Am J Public Health, 2000. 90(9): p. 1463-6. 4. Steptoe, R.J., J.M. Ritchie, and L.C. Harrison, Transfer of hematopoietic stem cells encoding autoantigen prevents autoimmune diabetes. J Clin Invest, 2003. 111(9): p. 1357-63. 5. Fischer, J.S., et al., The Multiple Sclerosis Functional Composite Measure (MSFC): an integrated approach to MS clinical outcome assessment. National MS Society Clinical Outcomes Assessment Task Force. Mult Scler, 1999. 5(4): p. 244-50. 6. Caturegli, P., et al., Autoimmune thyroid diseases. Curr Opin Rheumatol, 2007. 19(1): p. 44-8. 7. Aletaha, D., et al., 2010 Rheumatoid arthritis classification criteria: an American College of Rheumatology/European League Against Rheumatism collaborative initiative. Arthritis Rheum, 2010. 62(9): p. 2569-81. 8. Hochberg, M.C., Updating the American College of Rheumatology revised criteria for the classification of systemic lupus erythematosus. Arthritis Rheum, 1997. 40(9): p. 1725. 9. Singh, R.P., R.T. Waldron, and B.H. Hahn, Genes, tolerance and systemic autoimmunity. Autoimmun Rev, 2012. 11(9): p. 664-9. 10. Maverakis, E., et al., Glycans in the immune system and The Altered Glycan Theory of Autoimmunity: a critical review. J Autoimmun, 2015. 57: p. 1-13. 11. Ceccarelli, F., N. Agmon-Levin, and C. Perricone, Genetic Factors of Autoimmune Diseases. J Immunol Res, 2016. 2016: p. 3476023. 12. Steed, A.L. and T.S. Stappenbeck, Role of viruses and bacteria-virus interactions in autoimmunity. Curr Opin Immunol, 2014. 31: p. 102-7. 13. Martin, S.F., T lymphocyte-mediated immune responses to chemical and metal ions: implications for allergic and autoimmune disease. Int Arch Allergy Immunol, 2004. 134(3): p. 186-98. 14. Dai, R. and S.A. Ahmed, MicroRNA, a new paradigm for understanding immunoregulation, inflammation, and autoimmune diseases. Transl Res, 2011. 157(4): p. 163-79. 15. Hess, E.V., Environmental chemicals and autoimmune disease: cause and effect. Toxicology, 2002. 181-182: p. 65-70. 16. Okazaki, K., et al., Recent advances in the concept and diagnosis of autoimmune pancreatitis and IgG4-related disease. J Gastroenterol, 2011. 46(3): p. 277-88. 17. Chari, S.T., et al., Diagnosis of autoimmune pancreatitis: the Mayo Clinic experience. Clin Gastroenterol Hepatol, 2006. 4(8): p. 1010-6; quiz 934. 18. Ulrich, C.M., J. Bigler, and J.D. Potter, Non-steroidal anti-inflammatory drugs for cancer prevention: promise, perils and pharmacogenetics. Nat Rev Cancer, 2006. 6(2): p. 130-40. 19. Ichai, P., et al., Usefulness of corticosteroids for the treatment of severe and fulminant forms of autoimmune hepatitis. Liver Transpl, 2007. 13(7): p. 996-1003. 20. Ghoreschi, K., et al., Interleukin-4 therapy of psoriasis induces Th2 responses and improves human autoimmune disease. Nat Med, 2003. 9(1): p. 40-6.

148 | P a g e

21. Chandrashekara, S., The treatment strategies of autoimmune disease may need a different approach from conventional protocol: a review. Indian J Pharmacol, 2012. 44(6): p. 665-71. 22. Gross, J.A., et al., TACI and BCMA are receptors for a TNF homologue implicated in B-cell autoimmune disease. Nature, 2000. 404(6781): p. 995-9. 23. Kaveri, S.V., et al., Intravenous immunoglobulins in : more than mere replacement therapy. Clin Exp Immunol, 2011. 164 Suppl 2: p. 2-5. 24. Lallana, E.C. and C.E. Fadul, Toxicities of immunosuppressive treatment of autoimmune neurologic diseases. Curr Neuropharmacol, 2011. 9(3): p. 468-77. 25. Atkins, H.L., et al., Immunoablation and autologous haemopoietic stem-cell transplantation for aggressive multiple sclerosis: a multicentre single-group phase 2 trial. Lancet, 2016. 388(10044): p. 576-85. 26. Snowden, J.A., S. Kapoor, and A.G. Wilson, Stem cell transplantation in rheumatoid arthritis. Autoimmunity, 2008. 41(8): p. 625-31. 27. Burt, R.K., et al., Clinical applications of blood-derived and marrow-derived stem cells for nonmalignant diseases. JAMA, 2008. 299(8): p. 925-36. 28. de Kleer, I., et al., Autologous stem cell transplantation for autoimmunity induces immunologic self-tolerance by reprogramming autoreactive T cells and restoring the CD4+CD25+ immune regulatory network. Blood, 2006. 107(4): p. 1696-702. 29. Arstila, T.P., et al., A direct estimate of the human alphabeta T cell receptor diversity. Science, 1999. 286(5441): p. 958-61. 30. Laydon, D.J., C.R. Bangham, and B. Asquith, Estimating T-cell repertoire diversity: limitations of classical estimators and a new approach. Philos Trans R Soc Lond B Biol Sci, 2015. 370(1675). 31. Palmer, E., Negative selection--clearing out the bad apples from the T-cell repertoire. Nat Rev Immunol, 2003. 3(5): p. 383-91. 32. Hammerling, G.J., et al., Non-deletional mechanisms of peripheral and central tolerance: studies with transgenic mice with tissue-specific expression of a foreign MHC class I antigen. Immunol Rev, 1991. 122: p. 47-67. 33. McGargill, M.A., J.M. Derbinski, and K.A. Hogquist, Receptor editing in developing T cells. Nat Immunol, 2000. 1(4): p. 336-41. 34. Wang, F., C.Y. Huang, and O. Kanagawa, Rapid deletion of rearranged T cell antigen receptor (TCR) Valpha-Jalpha segment by secondary rearrangement in the thymus: role of continuous rearrangement of TCR alpha chain gene and positive selection in the T cell repertoire formation. Proc Natl Acad Sci U S A, 1998. 95(20): p. 11834-9. 35. Petrie, H.T., Role of thymic organ structure and stromal composition in steady-state postnatal T-cell production. Immunol Rev, 2002. 189: p. 8-19. 36. Kamradt, T. and N.A. Mitchison, Tolerance and autoimmunity. N Engl J Med, 2001. 344(9): p. 655-64. 37. Butcher, E.C. and L.J. Picker, Lymphocyte homing and homeostasis. Science, 1996. 272(5258): p. 60-6. 38. Ferber, I., et al., Levels of peripheral T cell tolerance induced by different doses of tolerogen. Science, 1994. 263(5147): p. 674-6. 39. Janeway, C.A., Jr., The immune system evolved to discriminate infectious nonself from noninfectious self. Immunol Today, 1992. 13(1): p. 11-6. 40. Kaufmann, T., A. Strasser, and P.J. Jost, Fas death receptor signalling: roles of Bid and XIAP. Cell Death Differ, 2012. 19(1): p. 42-50. 41. Zhang, M., et al., Transmembrane TNF-alpha promotes activation-induced cell death by forward and reverse signaling. Oncotarget, 2017. 8(38): p. 63799-63812.

149 | P a g e

42. Ludwinski, M.W., et al., Critical roles of Bim in T cell activation and T cell-mediated autoimmune inflammation in mice. J Clin Invest, 2009. 119(6): p. 1706-13. 43. Snow, A.L., et al., Critical role for BIM in T cell receptor restimulation-induced death. Biol Direct, 2008. 3: p. 34. 44. Gross, A., J.M. McDonnell, and S.J. Korsmeyer, BCL-2 family members and the mitochondria in apoptosis. Genes Dev, 1999. 13(15): p. 1899-911. 45. Chen, M., L. Huang, and J. Wang, Deficiency of Bim in dendritic cells contributes to overactivation of lymphocytes and autoimmunity. Blood, 2007. 109(10): p. 4360-7. 46. Appleman, L.J. and V.A. Boussiotis, T cell anergy and costimulation. Immunol Rev, 2003. 192: p. 161-80. 47. Chappert, P. and R.H. Schwartz, Induction of T cell anergy: integration of environmental cues and . Curr Opin Immunol, 2010. 22(5): p. 552- 9. 48. Probst, H.C., et al., Resting dendritic cells induce peripheral CD8+ T cell tolerance through PD-1 and CTLA-4. Nat Immunol, 2005. 6(3): p. 280-6. 49. Keir, M.E., et al., PD-1 and its ligands in tolerance and immunity. Annu Rev Immunol, 2008. 26: p. 677-704. 50. Nishimura, H., et al., Development of lupus-like autoimmune diseases by disruption of the PD-1 gene encoding an ITIM motif-carrying immunoreceptor. Immunity, 1999. 11(2): p. 141-51. 51. Tivol, E.A., et al., Loss of CTLA-4 leads to massive lymphoproliferation and fatal multiorgan tissue destruction, revealing a critical negative regulatory role of CTLA-4. Immunity, 1995. 3(5): p. 541-7. 52. Weiner, H.L., Induction and mechanism of action of transforming growth factor-beta- secreting Th3 regulatory cells. Immunol Rev, 2001. 182: p. 207-14. 53. Sakaguchi, S., et al., FOXP3+ regulatory T cells in the human immune system. Nat Rev Immunol, 2010. 10(7): p. 490-500. 54. Workman, C.J., et al., The development and function of regulatory T cells. Cell Mol Life Sci, 2009. 66(16): p. 2603-22. 55. Vignali, D.A., L.W. Collison, and C.J. Workman, How regulatory T cells work. Nat Rev Immunol, 2008. 8(7): p. 523-32. 56. Zeng, H., et al., Type 1 regulatory T cells: a new mechanism of peripheral immune tolerance. Cell Mol Immunol, 2015. 12(5): p. 566-71. 57. Sojka, D.K., Y.H. Huang, and D.J. Fowell, Mechanisms of regulatory T-cell suppression - a diverse arsenal for a moving target. Immunology, 2008. 124(1): p. 13- 22. 58. Lefrancois, L. and A.L. Marzo, The descent of memory T-cell subsets. Nat Rev Immunol, 2006. 6(8): p. 618-23. 59. Sourdive, D.J., et al., Conserved T cell receptor repertoire in primary and memory CD8 T cell responses to an acute viral infection. J Exp Med, 1998. 188(1): p. 71-82. 60. Opata, M.M., et al., Early effector cells survive the contraction phase in malaria infection and generate both central and effector memory T cells. J Immunol, 2015. 194(11): p. 5346-54. 61. Kaech, S.M., E.J. Wherry, and R. Ahmed, Effector and memory T-cell differentiation: implications for vaccine development. Nat Rev Immunol, 2002. 2(4): p. 251-62. 62. Sallusto, F., J. Geginat, and A. Lanzavecchia, Central memory and effector memory T cell subsets: function, generation, and maintenance. Annu Rev Immunol, 2004. 22: p. 745-63. 63. Barski, A., et al., Rapid Recall Ability of Memory T cells is Encoded in their Epigenome. Sci Rep, 2017. 7: p. 39785.

150 | P a g e

64. Lauvau, G. and S.M. Soudja, Mechanisms of Memory T Cell Activation and Effective Immunity. Adv Exp Med Biol, 2015. 850: p. 73-80. 65. Viglietta, V., et al., GAD65-reactive T cells are activated in patients with autoimmune type 1a diabetes. J Clin Invest, 2002. 109(7): p. 895-903. 66. Monti, P., et al., Islet transplantation in patients with autoimmune diabetes induces homeostatic cytokines that expand autoreactive memory T cells. J Clin Invest, 2008. 118(5): p. 1806-14. 67. Yang, J., et al., Allograft rejection mediated by memory T cells is resistant to regulation. Proc Natl Acad Sci U S A, 2007. 104(50): p. 19954-9. 68. Boyman, O. and J. Sprent, The role of interleukin-2 during homeostasis and activation of the immune system. Nat Rev Immunol, 2012. 12(3): p. 180-90. 69. Lakkis, F.G. and M.H. Sayegh, Memory T cells: a hurdle to immunologic tolerance. J Am Soc Nephrol, 2003. 14(9): p. 2402-10. 70. Lange, C., et al., Dendritic cell-regulatory T-cell interactions control self-directed immunity. Immunol Cell Biol, 2007. 85(8): p. 575-81. 71. Audiger, C., et al., The Importance of Dendritic Cells in Maintaining Immune Tolerance. J Immunol, 2017. 198(6): p. 2223-2231. 72. Bonifaz, L., et al., Efficient targeting of protein antigen to the dendritic cell receptor DEC-205 in the steady state leads to antigen presentation on major histocompatibility complex class I products and peripheral CD8+ T cell tolerance. J Exp Med, 2002. 196(12): p. 1627-38. 73. Kenna, T.J., R. Thomas, and R.J. Steptoe, Steady-state dendritic cells expressing cognate antigen terminate memory CD8+ T-cell responses. Blood, 2008. 111(4): p. 2091-100. 74. Kenna, T.J., et al., Targeting antigen to diverse APCs inactivates memory CD8+ T cells without eliciting tissue-destructive effector function. J Immunol, 2010. 184(2): p. 598- 606. 75. Nasreen, M., et al., Steady-state antigen-expressing dendritic cells terminate CD4+ memory T-cell responses. Eur J Immunol, 2010. 40(7): p. 2016-25. 76. Steptoe, R.J., et al., Cognate CD4+ help elicited by resting dendritic cells does not impair the induction of peripheral tolerance in CD8+ T cells. J Immunol, 2007. 178(4): p. 2094-103. 77. Rioux, J.D. and A.K. Abbas, Paths to understanding the genetic basis of autoimmune disease. Nature, 2005. 435(7042): p. 584-9. 78. Cooper, G.S., F.W. Miller, and J.P. Pandey, The role of genetic factors in autoimmune disease: implications for environmental research. Environ Health Perspect, 1999. 107 Suppl 5: p. 693-700. 79. Trowsdale, J. and J.C. Knight, Major histocompatibility complex genomics and human disease. Annu Rev Genomics Hum Genet, 2013. 14: p. 301-23. 80. Deakin, J.E., et al., Evolution and comparative analysis of the MHC Class III inflammatory region. BMC Genomics, 2006. 7: p. 281. 81. Santamaria, P., Cytokines and chemokines in autoimmune disease: an overview. Adv Exp Med Biol, 2003. 520: p. 1-7. 82. Sun, L., et al., Interleukin 12 (IL-12) family cytokines: Role in immune pathogenesis and treatment of CNS autoimmune disease. Cytokine, 2015. 75(2): p. 249-55. 83. Erlich, H., et al., HLA DR-DQ haplotypes and genotypes and type 1 diabetes risk: analysis of the type 1 diabetes genetics consortium families. Diabetes, 2008. 57(4): p. 1084-92.

151 | P a g e

84. Watson, C.T. and F. Breden, The immunoglobulin heavy chain locus: genetic variation, missing data, and implications for human disease. Genes Immun, 2012. 13(5): p. 363- 73. 85. Tang, J., et al., Association of polymorphisms in non-classic MHC genes with susceptibility to autoimmune hepatitis. Hepatobiliary Pancreat Dis Int, 2012. 11(2): p. 125-31. 86. Minias, P., et al., Extensive shared polymorphism at non-MHC immune genes in recently diverged North American prairie grouse. Immunogenetics, 2018. 70(3): p. 195-204. 87. Greenwald, R.J., et al., CTLA-4 regulates induction of anergy in vivo. Immunity, 2001. 14(2): p. 145-55. 88. Baroja, M.L., et al., Inhibition of CTLA-4 function by the regulatory subunit of serine/threonine phosphatase 2A. J Immunol, 2002. 168(10): p. 5070-8. 89. Wang, X., et al., B7-H4 Treatment of T Cells Inhibits ERK, JNK, p38, and AKT Activation. PLoS One, 2012. 7(1): p. e28232. 90. Ueda, H., et al., Association of the T-cell regulatory gene CTLA4 with susceptibility to autoimmune disease. Nature, 2003. 423(6939): p. 506-11. 91. Heijmans, B.T., et al., Persistent epigenetic differences associated with prenatal exposure to famine in humans. Proc Natl Acad Sci U S A, 2008. 105(44): p. 17046-9. 92. Katari, S., et al., DNA methylation and gene expression differences in children conceived in vitro or in vivo. Hum Mol Genet, 2009. 18(20): p. 3769-78. 93. Waterland, R.A., et al., Season of conception in rural gambia affects DNA methylation at putative human metastable epialleles. PLoS Genet, 2010. 6(12): p. e1001252. 94. Morris, A.P. and E. Zeggini, An evaluation of statistical approaches to rare variant analysis in genetic association studies. Genet Epidemiol, 2010. 34(2): p. 188-93. 95. Ballestar, E., Epigenetics lessons from twins: prospects for autoimmune disease. Clin Rev Allergy Immunol, 2010. 39(1): p. 30-41. 96. Metcalfe, K.A., et al., Concordance for type 1 diabetes in identical twins is affected by insulin genotype. Diabetes Care, 2001. 24(5): p. 838-42. 97. Samuelsson, U. and J. Carstensen, Space-time clustering at birth and at diagnosis of type 1 diabetes mellitus in relation to early clinical manifestation. J Pediatr Endocrinol Metab, 2003. 16(6): p. 859-67. 98. Sarkar, K., et al., Seasonal influence on the onset of idiopathic inflammatory myopathies in serologically defined groups. Arthritis Rheum, 2005. 52(8): p. 2433-8. 99. Anaya, J.M., et al., The Autoimmune Ecology. Front Immunol, 2016. 7: p. 139. 100. Daxinger, L. and E. Whitelaw, Transgenerational epigenetic inheritance: more questions than answers. Genome Res, 2010. 20(12): p. 1623-8. 101. Sandovici, I., et al., Maternal diet and aging alter the epigenetic control of a promoter- enhancer interaction at the Hnf4a gene in rat pancreatic islets. Proc Natl Acad Sci U S A, 2011. 108(13): p. 5449-54. 102. Waterland, R.A., et al., Maternal methyl supplements increase offspring DNA methylation at Axin Fused. Genesis, 2006. 44(9): p. 401-6. 103. Coleman, M.A. and R.J. Steptoe, Induction of antigen-specific tolerance through hematopoietic stem cell-mediated gene therapy: the future for therapy of autoimmune disease? Autoimmun Rev, 2012. 12(2): p. 195-203. 104. Rosato, E., S. Pisarri, and F. Salsano, Current strategies for the treatment of autoimmune diseases. J Biol Regul Homeost Agents, 2010. 24(3): p. 251-9. 105. Focosi, D., et al., Immunosuppressive monoclonal antibodies: current and next generation. Clin Microbiol Infect, 2011. 17(12): p. 1759-68.

152 | P a g e

106. Weiner, G.J., Building better monoclonal antibody-based therapeutics. Nat Rev Cancer, 2015. 15(6): p. 361-70. 107. Benson, J.M., et al., Therapeutic targeting of the IL-12/23 pathways: generation and characterization of ustekinumab. Nat Biotechnol, 2011. 29(7): p. 615-24. 108. Hansel, T.T., et al., The safety and side effects of monoclonal antibodies. Nat Rev Drug Discov, 2010. 9(4): p. 325-38. 109. Janikashvili, N., et al., Immunotherapeutic Targeting in Autoimmune Diseases. Mediators Inflamm, 2016. 2016: p. 1432702. 110. Henig, I. and T. Zuckerman, Hematopoietic stem cell transplantation-50 years of evolution and future perspectives. Rambam Maimonides Med J, 2014. 5(4): p. e0028. 111. Garcia Morin, M., et al., [Bone marrow transplant in patients with sickle cell anaemia. Experience in one centre]. An Pediatr (Barc), 2017. 86(3): p. 142-150. 112. Eixarch, H., et al., Tolerance induction in experimental autoimmune encephalomyelitis using non-myeloablative hematopoietic gene therapy with autoantigen. Mol Ther, 2009. 17(5): p. 897-905. 113. Chan, J., et al., Transplantation of bone marrow genetically engineered to express proinsulin II protects against autoimmune insulitis in NOD mice. J Gene Med, 2006. 8(11): p. 1281-90. 114. Doan, T., et al., Steady-state dendritic cells continuously inactivate T cells that escape thymic negative selection. Immunol Cell Biol, 2009. 87(8): p. 615-22. 115. Blaese, R.M., et al., T lymphocyte-directed gene therapy for ADA- SCID: initial trial results after 4 years. Science, 1995. 270(5235): p. 475-80. 116. Bordignon, C., et al., Gene therapy in peripheral blood lymphocytes and bone marrow for ADA- immunodeficient patients. Science, 1995. 270(5235): p. 470-5. 117. Hoogerbrugge, P.M., et al., Bone marrow gene transfer in three patients with adenosine deaminase deficiency. Gene Ther, 1996. 3(2): p. 179-83. 118. Cavazzana-Calvo, M., et al., Gene therapy of human severe combined immunodeficiency (SCID)-X1 disease. Science, 2000. 288(5466): p. 669-72. 119. Ruiz, P., et al., Transplant tolerance: new insights and strategies for long-term allograft acceptance. Clin Dev Immunol, 2013. 2013: p. 210506. 120. Sykes, M. and B. Nikolic, Treatment of severe autoimmune disease by stem-cell transplantation. Nature, 2005. 435(7042): p. 620-7. 121. van Bekkum, D.W., et al., Regression of adjuvant-induced arthritis in rats following bone marrow transplantation. Proc Natl Acad Sci U S A, 1989. 86(24): p. 10090-4. 122. Knaan-Shanzer, S., et al., Remission induction of adjuvant arthritis in rats by total body irradiation and autologous bone marrow transplantation. Bone Marrow Transplant, 1991. 8(5): p. 333-8. 123. Snowden, J.A., et al., Haematopoietic SCT in severe autoimmune diseases: updated guidelines of the European Group for Blood and Marrow Transplantation. Bone Marrow Transplant, 2012. 47(6): p. 770-90. 124. Arruda, L.C., et al., Resetting the immune response after autologous hematopoietic stem cell transplantation for autoimmune diseases. Curr Res Transl Med, 2016. 64(2): p. 107-13. 125. Burt, R.K., et al., Autologous non-myeloablative haemopoietic stem-cell transplantation compared with pulse cyclophosphamide once per month for systemic sclerosis (ASSIST): an open-label, randomised phase 2 trial. Lancet, 2011. 378(9790): p. 498-506. 126. Daikeler, T., A. Tichelli, and J. Passweg, Complications of autologous hematopoietic stem cell transplantation for patients with autoimmune diseases. Pediatr Res, 2012. 71(4 Pt 2): p. 439-44.

153 | P a g e

127. Hongo, D., et al., Interactions between NKT cells and Tregs are required for tolerance to combined bone marrow and organ transplants. Blood, 2012. 119(6): p. 1581-9. 128. Yi, H. and Y. Zhao, Chemokines, chemokine receptors and CD4+CD25+ regulatory T cells. Expert Rev Clin Immunol, 2007. 3(3): p. 343-9. 129. Li, H., et al., Mixed allogeneic chimerism induced by a sublethal approach prevents autoimmune diabetes and reverses insulitis in nonobese diabetic (NOD) mice. J Immunol, 1996. 156(1): p. 380-8. 130. LaFace, D.M. and A.B. Peck, Reciprocal allogeneic bone marrow transplantation between NOD mice and diabetes-nonsusceptible mice associated with transfer and prevention of autoimmune diabetes. Diabetes, 1989. 38(7): p. 894-901. 131. Karussis, D.M., et al., Immunomodulation of autoimmunity in MRL/lpr mice with syngeneic bone marrow transplantation (SBMT). Clin Exp Immunol, 1995. 100(1): p. 111-7. 132. Kushida, T., et al., Treatment of intractable autoimmune diseases in MRL/lpr mice using a new strategy for allogeneic bone marrow transplantation. Blood, 2000. 95(5): p. 1862-8. 133. van Gelder, M., A.H. Mulder, and D.W. van Bekkum, Treatment of relapsing experimental autoimmune encephalomyelitis with largely MHC-matched allogeneic bone marrow transplantation. Transplantation, 1996. 62(6): p. 810-8. 134. El-Badri, N.S., et al., Successful prevention of autoimmune disease by transplantation of adequate number of fully allogeneic hematopoietic stem cells. Transplantation, 2000. 70(6): p. 870-7. 135. Yasumizu, R., et al., Treatment of type 1 diabetes mellitus in non-obese diabetic mice by transplantation of allogeneic bone marrow and pancreatic tissue. Proc Natl Acad Sci U S A, 1987. 84(18): p. 6555-7. 136. Lin, H., The stem-cell niche theory: lessons from flies. Nat Rev Genet, 2002. 3(12): p. 931-40. 137. Kirzner, R.P., et al., Prevention of coronary vascular disease by transplantation of T- cell-depleted bone marrow and hematopoietic stem cell preparation in autoimmune- prone w/BF(1) mice. Biol Blood Marrow Transplant, 2000. 6(5): p. 513-22. 138. Weissman, I.L., Transfer of tolerance. Transplantation, 1973. 15(3): p. 265-9. 139. Ikehara, S., et al., Long-term observations of autoimmune-prone mice treated for autoimmune disease by allogeneic bone marrow transplantation. Proc Natl Acad Sci U S A, 1989. 86(9): p. 3306-10. 140. Gandy, K.L. and I.L. Weissman, Tolerance of allogeneic heart grafts in mice simultaneously reconstituted with purified allogeneic hematopoietic stem cells. Transplantation, 1998. 65(3): p. 295-304. 141. Kawaharada, N., et al., Mixed hematopoietic chimerism prevents allograft vasculopathy. J Heart Lung Transplant, 1999. 18(6): p. 532-41. 142. Remuzzi, G., Cellular basis of long-term organ transplant acceptance: pivotal role of intrathymic clonal deletion and thymic dependence of bone marrow microchimerism- associated tolerance. Am J Kidney Dis, 1998. 31(2): p. 197-212. 143. George, J.F., et al., An essential role for Fas ligand in transplantation tolerance induced by donor bone marrow. Nat Med, 1998. 4(3): p. 333-5. 144. Burt, R.K. and A. Traynor, Hematopoietic stem cell therapy of autoimmune diseases. Curr Opin Hematol, 1998. 5(6): p. 472-7. 145. Slavin, S., et al., Graft vs autoimmunity following allogeneic non-myeloablative blood stem cell transplantation in a patient with chronic myelogenous leukemia and severe systemic psoriasis and psoriatic polyarthritis. Exp Hematol, 2000. 28(7): p. 853-7.

154 | P a g e

146. Delaney, C.P., et al., Allogeneic hematolymphoid microchimerism and prevention of autoimmune disease in the rat. A relationship between allo- and autoimmunity. J Clin Invest, 1996. 97(1): p. 217-25. 147. Wang, B., et al., Effective treatment of autoimmune disease and progressive renal disease by mixed bone-marrow transplantation that establishes a stable mixed chimerism in BXSB recipient mice. Proc Natl Acad Sci U S A, 1999. 96(6): p. 3012-6. 148. Prigozhina, T.B., O. Gurevitch, and S. Slavin, Nonmyeloablative conditioning to induce bilateral tolerance after allogeneic bone marrow transplantation in mice. Exp Hematol, 1999. 27(10): p. 1503-10. 149. Weiss, L. and S. Slavin, Prevention and treatment of graft-versus-host disease by down- regulation of anti-host reactivity with veto cells of host origin. Bone Marrow Transplant, 1999. 23(11): p. 1139-43. 150. Lopez-Cubero, S.O., K.M. Sullivan, and G.B. McDonald, Course of Crohn's disease after allogeneic marrow transplantation. Gastroenterology, 1998. 114(3): p. 433-40. 151. Olalla, J.I., et al., Disappearance of lupus anticoagulant after allogeneic bone marrow transplantation. Bone Marrow Transplant, 1999. 23(1): p. 83-5. 152. Yin, J.A. and S.N. Jowitt, Resolution of immune-mediated diseases following allogeneic bone marrow transplantation for leukaemia. Bone Marrow Transplant, 1992. 9(1): p. 31-3. 153. Tracy, A., C.D. Buckley, and K. Raza, Pre-symptomatic autoimmunity in rheumatoid arthritis: when does the disease start? Semin Immunopathol, 2017. 39(4): p. 423-435. 154. Millan, M.T., et al., Mixed chimerism and immunosuppressive drug withdrawal after HLA-mismatched kidney and hematopoietic progenitor transplantation. Transplantation, 2002. 73(9): p. 1386-91. 155. Scandling, J.D., et al., Tolerance and chimerism after renal and hematopoietic-cell transplantation. N Engl J Med, 2008. 358(4): p. 362-8. 156. Scandling, J.D., et al., Tolerance and withdrawal of immunosuppressive drugs in patients given kidney and hematopoietic cell transplants. Am J Transplant, 2012. 12(5): p. 1133-45. 157. Lowsky, R., et al., Protective conditioning for acute graft-versus-host disease. N Engl J Med, 2005. 353(13): p. 1321-31. 158. Kohrt, H.E., et al., TLI and ATG conditioning with low risk of graft-versus-host disease retains antitumor reactions after allogeneic hematopoietic cell transplantation from related and unrelated donors. Blood, 2009. 114(5): p. 1099-109. 159. Leventhal, J., et al., Chimerism and tolerance without GVHD or engraftment syndrome in HLA-mismatched combined kidney and hematopoietic stem cell transplantation. Sci Transl Med, 2012. 4(124): p. 124ra28. 160. Leventhal, J., et al., Tolerance induction in HLA disparate living donor kidney transplantation by donor stem cell infusion: durable chimerism predicts outcome. Transplantation, 2013. 95(1): p. 169-76. 161. Kaufman, C.L., et al., Phenotypic characterization of a novel bone marrow-derived cell that facilitates engraftment of allogeneic bone marrow stem cells. Blood, 1994. 84(8): p. 2436-46. 162. Williams, R.C., Jr., Hypothesis: rheumatoid factors are antiidiotypes related to bacterial or viral Fc receptors. Arthritis Rheum, 1988. 31(9): p. 1204-7. 163. Kasamon, Y.L., et al., Nonmyeloablative HLA-haploidentical bone marrow transplantation with high-dose posttransplantation cyclophosphamide: effect of HLA disparity on outcome. Biol Blood Marrow Transplant, 2010. 16(4): p. 482-9.

155 | P a g e

164. Luznik, L., et al., HLA-haploidentical bone marrow transplantation for hematologic malignancies using nonmyeloablative conditioning and high-dose, posttransplantation cyclophosphamide. Biol Blood Marrow Transplant, 2008. 14(6): p. 641-50. 165. Li, H.W. and M. Sykes, Emerging concepts in haematopoietic cell transplantation. Nat Rev Immunol, 2012. 12(6): p. 403-16. 166. Kean, L.S. and B.R. Blazar, Cooking up tolerance: has a new recipe been created? Am J Transplant, 2012. 12(7): p. 1667-9. 167. Strober, S., et al., Translational studies in hematopoietic cell transplantation: treatment of hematologic malignancies as a stepping stone to tolerance induction. Semin Immunol, 2011. 23(4): p. 273-81. 168. Billingham, R.E., L. Brent, and P.B. Medawar, Actively acquired tolerance of foreign cells. Nature, 1953. 172(4379): p. 603-6. 169. Owen, R.D., Immunogenetic Consequences of Vascular Anastomoses between Bovine Twins. Science, 1945. 102(2651): p. 400-1. 170. Ildstad, S.T. and D.H. Sachs, Reconstitution with syngeneic plus allogeneic or xenogeneic bone marrow leads to specific acceptance of allografts or xenografts. Nature, 1984. 307(5947): p. 168-70. 171. Sundt, T.M., 3rd and D.H. Sachs, Applications of molecular biology to transplantation tolerance. Immunol Today, 1988. 9(11): p. 342-4. 172. Wong, W., P.J. Morris, and K.J. Wood, Syngeneic bone marrow expressing a single donor class I MHC molecule permits acceptance of a fully allogeneic cardiac allograft. Transplantation, 1996. 62(10): p. 1462-8. 173. Coleman, M.A., et al., Tolerance induction with gene-modified stem cells and immune- preserving conditioning in primed mice: restricting antigen to differentiated antigen- presenting cells permits efficacy. Blood, 2013. 121(6): p. 1049-58. 174. Brown, B.D., et al., Endogenous microRNA can be broadly exploited to regulate transgene expression according to tissue, lineage and differentiation state. Nat Biotechnol, 2007. 25(12): p. 1457-67. 175. Shi, Q., et al., Syngeneic transplantation of hematopoietic stem cells that are genetically modified to express factor VIII in platelets restores hemostasis to hemophilia A mice with preexisting FVIII immunity. Blood, 2008. 112(7): p. 2713-21. 176. Andersson, G., et al., Engraftment of retroviral EGFP-transduced bone marrow in mice prevents rejection of EGFP-transgenic skin grafts. Mol Ther, 2003. 8(3): p. 385-91. 177. Tian, C., et al., Induction of T cell tolerance to a protein expressed in the cytoplasm through retroviral-mediated gene transfer. J Gene Med, 2003. 5(5): p. 359-65. 178. Bagley, J., et al., Induction of T-cell tolerance to an MHC class I alloantigen by gene therapy. Blood, 2002. 99(12): p. 4394-9. 179. Jindra, P.T., et al., Tolerance to MHC class II disparate allografts through genetic modification of bone marrow. Gene Ther, 2013. 20(5): p. 478-86. 180. Ko, H.J., et al., Targeting MOG expression to dendritic cells delays onset of experimental autoimmune disease. Autoimmunity, 2011. 44(3): p. 177-87. 181. Nasa, Z., et al., Nonmyeloablative conditioning generates autoantigen-encoding bone marrow that prevents and cures an experimental autoimmune disease. Am J Transplant, 2012. 12(8): p. 2062-71. 182. Baranyi, U., et al., Persistent molecular microchimerism induces long-term tolerance towards a clinically relevant respiratory . Clin Exp Allergy, 2012. 42(8): p. 1282-92. 183. Gattringer, M., et al., Engraftment of retrovirally transduced Bet v 1-GFP expressing bone marrow cells leads to allergen-specific tolerance. Immunobiology, 2013. 218(9): p. 1139-46.

156 | P a g e

184. Chang, A.H., et al., Erythroid-specific human factor IX delivery from in vivo selected hematopoietic stem cells following nonmyeloablative conditioning in hemophilia B mice. Mol Ther, 2008. 16(10): p. 1745-52. 185. Shi, Q., et al., Platelet gene therapy corrects the hemophilic phenotype in immunocompromised hemophilia A mice transplanted with genetically manipulated human cord blood stem cells. Blood, 2014. 123(3): p. 395-403. 186. Mellman, I., Dendritic cells: master regulators of the immune response. Cancer Immunol Res, 2013. 1(3): p. 145-9. 187. Cella, M., et al., Ligation of CD40 on dendritic cells triggers production of high levels of interleukin-12 and enhances T cell stimulatory capacity: T-T help via APC activation. J Exp Med, 1996. 184(2): p. 747-52. 188. Grewal, I.S., et al., Requirement for CD40 ligand in costimulation induction, T cell activation, and experimental allergic encephalomyelitis. Science, 1996. 273(5283): p. 1864-7. 189. Burgess, J.K., et al., Detection and characterization of OX40 ligand expression in human airway smooth muscle cells: a possible role in asthma? J Allergy Clin Immunol, 2004. 113(4): p. 683-9. 190. Ito, T., et al., TSLP-activated dendritic cells induce an inflammatory T helper type 2 cell response through OX40 ligand. J Exp Med, 2005. 202(9): p. 1213-23. 191. Reis e Sousa, C., Dendritic cells in a mature age. Nat Rev Immunol, 2006. 6(6): p. 476- 83. 192. Kurts, C., et al., Class I-restricted cross-presentation of exogenous self-antigens leads to deletion of autoreactive CD8(+) T cells. J Exp Med, 1997. 186(2): p. 239-45. 193. Mukhopadhaya, A., et al., Selective delivery of beta cell antigen to dendritic cells in vivo leads to deletion and tolerance of autoreactive CD8+ T cells in NOD mice. Proc Natl Acad Sci U S A, 2008. 105(17): p. 6374-9. 194. Fu, F., et al., Costimulatory molecule-deficient dendritic cell progenitors (MHC class II+, CD80dim, CD86-) prolong cardiac allograft survival in nonimmunosuppressed recipients. Transplantation, 1996. 62(5): p. 659-65. 195. Lu, L., et al., Bone marrow-derived dendritic cell progenitors (NLDC 145+, MHC class II+, B7-1dim, B7-2-) induce alloantigen-specific hyporesponsiveness in murine T lymphocytes. Transplantation, 1995. 60(12): p. 1539-45. 196. Morrison, S.J. and D.T. Scadden, The bone marrow niche for haematopoietic stem cells. Nature, 2014. 505(7483): p. 327-34. 197. Di Rosa, F., Two Niches in the Bone Marrow: A Hypothesis on Life-long T Cell Memory. Trends Immunol, 2016. 37(8): p. 503-12. 198. Zhao, M. and L. Li, Dissecting the bone marrow HSC niches. Cell Res, 2016. 26(9): p. 975-6. 199. Cordeiro-Spinetti, E., R.S. Taichman, and A. Balduino, The bone marrow endosteal niche: how far from the surface? J Cell Biochem, 2015. 116(1): p. 6-11. 200. Nemeth, K. and E. Mezey, Origin of stem cells in the BM niche: new clues from mastocytosis. Blood, 2016. 127(6): p. 670-2. 201. Huang, X., S. Cho, and G.J. Spangrude, Hematopoietic stem cells: generation and self- renewal. Cell Death Differ, 2007. 14(11): p. 1851-9. 202. Seita, J. and I.L. Weissman, Hematopoietic stem cell: self-renewal versus differentiation. Wiley Interdiscip Rev Syst Biol Med, 2010. 2(6): p. 640-53. 203. Wang, T., et al., The control of hematopoietic stem cell maintenance, self-renewal, and differentiation by Mysm1-mediated epigenetic regulation. Blood, 2013. 122(16): p. 2812-22.

157 | P a g e

204. Zon, L.I., Intrinsic and extrinsic control of haematopoietic stem-cell self-renewal. Nature, 2008. 453(7193): p. 306-13. 205. Bhattacharya, D., L.I. Ehrlich, and I.L. Weissman, Space-time considerations for hematopoietic stem cell transplantation. Eur J Immunol, 2008. 38(8): p. 2060-7. 206. Chan, C.K., et al., Endochondral ossification is required for haematopoietic stem-cell niche formation. Nature, 2009. 457(7228): p. 490-4. 207. Chan, C.K., et al., Identification and specification of the mouse skeletal stem cell. Cell, 2015. 160(1-2): p. 285-98. 208. Burt, R.K., et al., Induction of tolerance in autoimmune diseases by hematopoietic stem cell transplantation: getting closer to a cure? Blood, 2002. 99(3): p. 768-84. 209. Brodsky, R.A., et al., Immunoablative high-dose cyclophosphamide without stem-cell rescue for refractory, severe autoimmune disease. Ann Intern Med, 1998. 129(12): p. 1031-5. 210. Rebeiro, P. and J. Moore, The role of autologous haemopoietic stem cell transplantation in the treatment of autoimmune disorders. Intern Med J, 2016. 46(1): p. 17-28. 211. Joske, D.J., et al., Autologous bone-marrow transplantation for rheumatoid arthritis. Lancet, 1997. 350(9074): p. 337-8. 212. Snowden, J.A., et al., A phase I/II dose escalation study of intensified cyclophosphamide and autologous blood stem cell rescue in severe, active rheumatoid arthritis. Arthritis Rheum, 1999. 42(11): p. 2286-92. 213. McColl, G., et al., High-dose chemotherapy and syngeneic hemopoietic stem-cell transplantation for severe, seronegative rheumatoid arthritis. Ann Intern Med, 1999. 131(7): p. 507-9. 214. Burt, R.K., et al., Treatment of autoimmune disease by intense immunosuppressive conditioning and autologous hematopoietic stem cell transplantation. Blood, 1998. 92(10): p. 3505-14. 215. Traynor, A.E., et al., Treatment of severe systemic lupus erythematosus with high-dose chemotherapy and haemopoietic stem-cell transplantation: a phase I study. Lancet, 2000. 356(9231): p. 701-7. 216. Martini, A., et al., Marked and sustained improvement two years after autologous stem cell transplantation in a girl with systemic sclerosis. Arthritis Rheum, 1999. 42(4): p. 807-11. 217. Musso, M., et al., Successful treatment of resistant thrombotic thrombocytopenic purpura/hemolytic uremic syndrome with autologous peripheral blood stem and progenitor (CD34+) cell transplantation. Bone Marrow Transplant, 1999. 24(2): p. 207-9. 218. Mills, W., et al., BEAM chemotherapy and autologous bone marrow transplantation for patients with relapsed or refractory non-Hodgkin's lymphoma. J Clin Oncol, 1995. 13(3): p. 588-95. 219. Burt, R.K., et al., T cell-depleted autologous hematopoietic stem cell transplantation for multiple sclerosis: report on the first three patients. Bone Marrow Transplant, 1998. 21(6): p. 537-41. 220. Wulffraat, N., et al., Autologous haemopoietic stem-cell transplantation in four patients with refractory juvenile chronic arthritis. Lancet, 1999. 353(9152): p. 550-3. 221. Openshaw, H., et al., Peripheral blood stem cell transplantation in multiple sclerosis with busulfan and cyclophosphamide conditioning: report of toxicity and immunological monitoring. Biol Blood Marrow Transplant, 2000. 6(5A): p. 563-75.

158 | P a g e

222. Baron, F., et al., Effective treatment of Jo-1-associated polymyositis with T-cell- depleted autologous peripheral blood stem cell transplantation. Br J Haematol, 2000. 110(2): p. 339-42. 223. Marmont, A.M., et al., Failure of autologous stem cell transplantation in refractory thrombocytopenic purpura. Bone Marrow Transplant, 1998. 22(8): p. 827-8. 224. Swart, J.F., et al., Changing winds in refractory autoimmune disease in children: clearing the road for tolerance with cellular therapies. Curr Opin Rheumatol, 2012. 24(3): p. 267-73. 225. Sureda, A., et al., Indications for allo- and auto-SCT for haematological diseases, solid tumours and immune disorders: current practice in Europe, 2015. Bone Marrow Transplant, 2015. 50(8): p. 1037-56. 226. Derderian, S.C., et al., In utero depletion of fetal hematopoietic stem cells improves engraftment after neonatal transplantation in mice. Blood, 2014. 124(6): p. 973-80. 227. Chhabra, A., et al., Hematopoietic stem cell transplantation in immunocompetent hosts without radiation or chemotherapy. Sci Transl Med, 2016. 8(351): p. 351ra105. 228. Taya, Y., et al., Depleting dietary valine permits nonmyeloablative mouse hematopoietic stem cell transplantation. Science, 2016. 354(6316): p. 1152-1155. 229. Palchaudhuri, R., et al., Non-genotoxic conditioning for hematopoietic stem cell transplantation using a hematopoietic-cell-specific internalizing immunotoxin. Nat Biotechnol, 2016. 34(7): p. 738-45. 230. Czechowicz, A., et al., Selective hematopoietic stem cell ablation using CD117- antibody-drug-conjugates enables safe and effective transplantation with immunity preservation. Nat Commun, 2019. 10(1): p. 617. 231. Bergamaschi, G., et al., Saporin, a ribosome-inactivating protein used to prepare immunotoxins, induces cell death via apoptosis. Br J Haematol, 1996. 93(4): p. 789-94. 232. Polito, L., et al., Saporin-S6: a useful tool in cancer therapy. Toxins (Basel), 2013. 5(10): p. 1698-722. 233. Pestronk, A., et al., Combined short-term immunotherapy for experimental autoimmune myasthenia gravis. Ann Neurol, 1983. 14(2): p. 235-41. 234. Van Bekkum, D.W., Experimental basis for the treatment of autoimmune diseases with autologous hematopoietic stem cell transplantation. Bone Marrow Transplant, 2003. 32 Suppl 1: p. S37-9. 235. Guillaume, T., D.B. Rubinstein, and M. Symann, Immune reconstitution and immunotherapy after autologous hematopoietic stem cell transplantation. Blood, 1998. 92(5): p. 1471-90. 236. Burt, R.K., et al., The promise of hematopoietic stem cell transplantation for autoimmune diseases. Bone Marrow Transplant, 2003. 31(7): p. 521-4. 237. Tyndall, A. and R. Saccardi, Haematopoietic stem cell transplantation in the treatment of severe autoimmune disease: results from phase I/II studies, prospective randomized trials and future directions. Clin Exp Immunol, 2005. 141(1): p. 1-9. 238. Hugle, T. and T. Daikeler, Stem cell transplantation for autoimmune diseases. Haematologica, 2010. 95(2): p. 185-8. 239. Hough, R.E., J.A. Snowden, and N.M. Wulffraat, Haemopoietic stem cell transplantation in autoimmune diseases: a European perspective. Br J Haematol, 2005. 128(4): p. 432-59. 240. Popat, U. and R. Krance, Haematopoietic stem cell transplantation for autoimmune disorders: the American perspective. Br J Haematol, 2004. 126(5): p. 637-49. 241. Cippa, P.E., et al., Targeting apoptosis to induce stable mixed hematopoietic chimerism and long-term allograft survival without myelosuppressive conditioning in mice. Blood, 2013. 122(9): p. 1669-77.

159 | P a g e

242. Hogquist, K.A., et al., T cell receptor antagonist peptides induce positive selection. Cell, 1994. 76(1): p. 17-27. 243. Chan, J., et al., Transplantation of bone marrow transduced to express self-antigen establishes deletional tolerance and permanently remits autoimmune disease. J Immunol, 2008. 181(11): p. 7571-80. 244. Dufait, I., et al., Retroviral and lentiviral vectors for the induction of immunological tolerance. Scientifica (Cairo), 2012. 2012: 694137. 245. Sorror, M.L., et al., Hematopoietic cell transplantation (HCT)-specific comorbidity index: a new tool for risk assessment before allogeneic HCT. Blood, 2005. 106(8): p. 2912-9. 246. Diaconescu, R., et al., Morbidity and mortality with nonmyeloablative compared with myeloablative conditioning before hematopoietic cell transplantation from HLA- matched related donors. Blood, 2004. 104(5): p. 1550-8. 247. Kornblit, B., et al., Haematopoietic cell transplantation with non-myeloablative conditioning in Denmark: disease-specific outcome, complications and hospitalization requirements of the first 100 transplants. Bone Marrow Transplant, 2008. 41(10): p. 851-9. 248. Czechowicz, A., et al., Efficient transplantation via antibody-based clearance of hematopoietic stem cell niches. Science, 2007. 318(5854): p. 1296-9. 249. George, B.M., et al., Antibody Conditioning Enables MHC-Mismatched Hematopoietic Stem Cell Transplants and Organ Graft Tolerance. Cell Stem Cell, 2019. 25(2): p. 185- 192 e3. 250. Xue, X., et al., Antibody targeting KIT as pretransplantation conditioning in immunocompetent mice. Blood, 2010. 116(24): p. 5419-22. 251. Coleman, M.A., et al., Antigen-encoding bone marrow terminates islet-directed memory CD8+ T-cell responses to alleviate islet transplant rejection. Diabetes, 2016. 252. Aiuti, A., et al., Lentiviral hematopoietic stem cell gene therapy in patients with Wiskott-Aldrich syndrome. Science, 2013. 341(6148): p. 1233151. 253. Ruckerl, D. and J.E. Allen, Macrophage proliferation, provenance, and plasticity in macroparasite infection. Immunol Rev, 2014. 262(1): p. 113-33. 254. Meng, F. and C.A. Lowell, Lipopolysaccharide (LPS)-induced macrophage activation and signal transduction in the absence of Src-family kinases Hck, Fgr, and Lyn. J Exp Med, 1997. 185(9): p. 1661-70. 255. Zheng, X.F., et al., Lipopolysaccharide-induced M2 to M1 macrophage transformation for IL-12p70 production is blocked by Candida albicans mediated up-regulation of EBI3 expression. PLoS One, 2013. 8(5): p. e63967. 256. Arteaga Figueroa, L., et al., Comparison between Peritoneal Macrophage Activation by Bougainvillea xbuttiana Extract and LPS and/or Interleukins. Biomed Res Int, 2017. 2017: p. 4602952. 257. Bhatt, K.H., et al., Short-course rapamycin treatment enables engraftment of immunogenic gene-engineered bone marrow under low-dose irradiation to permit long-term immunological tolerance. Stem Cell Res Ther, 2017. 8(1): p. 57. 258. Cosimi, A.B. and D.H. Sachs, Mixed chimerism and transplantation tolerance. Transplantation, 2004. 77(6): p. 943-6. 259. Sachs, D.H., Tolerance: of mice and men. J Clin Invest, 2003. 111(12): p. 1819-21. 260. Sykes, M. and D.H. Sachs, Mixed chimerism. Philos Trans R Soc Lond B Biol Sci, 2001. 356(1409): p. 707-26. 261. Sayegh, M.H., et al., Immunologic tolerance to renal allografts after bone marrow transplants from the same donors. Ann Intern Med, 1991. 114(11): p. 954-5.

160 | P a g e

262. Helg, C., et al., Renal transplantation without immunosuppression in a host with tolerance induced by allogeneic bone marrow transplantation. Transplantation, 1994. 58(12): p. 1420-2. 263. Jacobsen, N., et al., Tolerance to an HLA-B,DR disparate kidney allograft after bone- marrow transplantation from same donor. Lancet, 1994. 343(8900): p. 800. 264. Fuchimoto, Y., et al., Relationship between chimerism and tolerance in a kidney transplantation model. J Immunol, 1999. 162(10): p. 5704-11. 265. Oudin, C., et al., Reduced-toxicity conditioning prior to allogeneic stem cell transplantation improves outcome in patients with myeloid malignancies. Haematologica, 2014. 99(11): p. 1762-8. 266. Gyurkocza, B. and B.M. Sandmaier, Conditioning regimens for hematopoietic cell transplantation: one size does not fit all. Blood, 2014. 124(3): p. 344-53. 267. Gao, C., et al., Nongenotoxic antibody-drug conjugate conditioning enables safe and effective platelet gene therapy of hemophilia A mice. Blood Adv, 2019. 3(18): p. 2700- 2711. 268. Hosseini, H., et al., Non-myeloablative transplantation of bone marrow expressing self- antigen establishes peripheral tolerance and completely prevents autoimmunity in mice. Gene Ther, 2012. 19(11): p. 1075-84. 269. Schafer, R., et al., Quantitation of progenitor cell populations and growth factors after bone marrow aspirate concentration. J Transl Med, 2019. 17(1): p. 115. 270. Weiskopf, K., et al., Myeloid Cell Origins, Differentiation, and Clinical Implications. Microbiol Spectr, 2016. 4(5). 271. Shahaf, G., et al., B Cell Development in the Bone Marrow Is Regulated by Homeostatic Feedback Exerted by Mature B Cells. Front Immunol, 2016. 7: p. 77. 272. Cull, G., et al., Lymphocyte reconstitution following autologous stem cell transplantation for progressive MS. Mult Scler J Exp Transl Clin, 2017. 3(1): p. 2055217317700167. 273. Pollack, S.M., et al., Assessment of the hematopoietic cell transplantation comorbidity index in non-Hodgkin lymphoma patients receiving reduced-intensity allogeneic hematopoietic stem cell transplantation. Biol Blood Marrow Transplant, 2009. 15(2): p. 223-30. 274. Asiedu, K.O., et al., Bone Marrow Cell Trafficking Analyzed by (89)Zr-oxine Positron Emission Tomography in a Murine Transplantation Model. Clin Cancer Res, 2017. 23(11): p. 2759-2768. 275. Andre, C., et al., Sequence analysis of two genomic regions containing the KIT and the FMS receptor tyrosine kinase genes. Genomics, 1997. 39(2): p. 216-26. 276. Ogawa, M., et al., Expression and function of c-kit in hemopoietic progenitor cells. J Exp Med, 1991. 174(1): p. 63-71. 277. Marech, I., et al., C-Kit receptor and tryptase expressing mast cells correlate with angiogenesis in breast cancer patients. Oncotarget, 2018. 9(8): p. 7918-7927. 278. Pang, W.W., et al., Anti-CD117 antibody depletes normal and myelodysplastic syndrome human hematopoietic stem cells in xenografted mice. Blood, 2019. 133(19): p. 2069-2078. 279. Chandrakasan, S., et al., KIT blockade is sufficient for donor hematopoietic stem cell engraftment in Fanconi anemia mice. Blood, 2017. 129(8): p. 1048-1052. 280. Biffi, A., et al., Lentiviral hematopoietic stem cell gene therapy benefits metachromatic leukodystrophy. Science, 2013. 341(6148): p. 1233158. 281. Gooley, T.A., et al., Reduced mortality after allogeneic hematopoietic-cell transplantation. N Engl J Med, 2010. 363(22): p. 2091-101.

161 | P a g e

282. Mikulska, M., et al., Mortality after bloodstream infections in allogeneic haematopoietic stem cell transplant (HSCT) recipients. Infection, 2012. 40(3): p. 271- 8. 283. Marr, K.A., Delayed opportunistic infections in hematopoietic stem cell transplantation patients: a surmountable challenge. Hematology Am Soc Hematol Educ Program, 2012. 2012: p. 265-70. 284. Wang, B., et al., The role of CD8+ T cells in the initiation of insulin-dependent diabetes mellitus. Eur J Immunol, 1996. 26(8): p. 1762-9. 285. Benson, R.A., J.M. Brewer, and A.M. Platt, Mechanisms of autoimmunity in human diseases: a critical review of current dogma. Curr Opin Rheumatol, 2014. 26(2): p. 197-203. 286. Szodoray, P., et al., Autologous stem cell transplantation in autoimmune and rheumatic diseases: from the molecular background to clinical applications. Scand J Rheumatol, 2010. 39(1): p. 1-11. 287. Bakhuraysah, M.M., C. Siatskas, and S. Petratos, Hematopoietic stem cell transplantation for multiple sclerosis: is it a clinical reality? Stem Cell Res Ther, 2016. 7: p. 12. 288. Ko, H.J., et al., Transplantation of -encoding bone marrow cells delays the onset of experimental autoimmune encephalomyelitis. Eur J Immunol, 2010. 40(12): p. 3499-509. 289. Nakamura-Ishizu, A., H. Takizawa, and T. Suda, The analysis, roles and regulation of quiescence in hematopoietic stem cells. Development, 2014. 141(24): p. 4656-66. 290. Rodriguez, R., et al., Identification of diphtheria toxin via screening as a potent cell cycle and p53-independent cytotoxin for human prostate cancer therapeutics. Prostate, 1998. 34(4): p. 259-69. 291. Bouchard, H., C. Viskov, and C. Garcia-Echeverria, Antibody-drug conjugates-a new wave of cancer drugs. Bioorg Med Chem Lett, 2014. 24(23): p. 5357-63. 292. Press, O.W., et al., Retention of B-cell-specific monoclonal antibodies by human lymphoma cells. Blood, 1994. 83(5): p. 1390-7. 293. Bikoue, A., G. Janossy, and D. Barnett, Stabilised cellular immuno-fluorescence assay: CD45 expression as a calibration standard for human leukocytes. J Immunol Methods, 2002. 266(1-2): p. 19-32. 294. Saad, A. and L.S. Lamb, Ex vivo T-cell depletion in allogeneic hematopoietic stem cell transplant: past, present and future. Bone Marrow Transplant, 2017. 52(9): p. 1241- 1248.

162 | P a g e