Identification and Functional Analysis of Novel Genes Associated with Inherited Bone Marrow Failure Syndromes

Anna Matveev

A thesis submitted in conformity with the requirements for the degree of Master of Science Institute of Medical Science University of Toronto

Abstract

Identification and Functional Analysis of Novel Genes Associated with Inherited Bone Marrow Failure Syndromes

Anna Matveev

Master of Science

Institute of Medical Science University of Toronto

2020

Inherited bone marrow failure syndromes are multisystem-disorders that affect development of hematopoietic system. One of IBMFSs is Shwachman-Diamond-syndrome and about 80-90% of patients have mutations in the Shwachman-Bodian-Diamond-Syndrome gene. To unravel the genetic cause of the disease in the remaining 10-20% of patients, we performed WES as well as

SNP-genotyping in families with SDS-phenotype and no mutations in SBDS. The results showed a region of homozygosity in chromosome 5p-arm DNAJC21 is in this region. Western blotting revealed reduced/null protein in patient. DNAJC21-homolog in yeast has been shown facilitating the release of the Arx1/Alb1 heterodimer from pre-60S.To investigate the cellular functions of

DNAJC21 we knocked-down it in HEK293T-cells. We observed a high-level of ROS, which led to reduced cell proliferation. Our data indicate that mutations in DNAJC21 contribute to SDS.

We hypothesize that DNAJC21 related ribosomal defects lead to increased levels of ROS therefore altering development and maturation of hematopoietic cells.

Acknowledgments

I would like to take this opportunity to extend my deepest gratitude to everyone who has helped me throughout my degree. First, I would like to thank my supervisor Dr. Yigal Dror for giving me the opportunity to study under his supervision, guiding me through the experimental design, supporting me through my degree, and without whom this project would not have been possible.

I am grateful for all the time and attention that he always put into ensuring that I was successful every step of the way. I am also appreciative of my committee members, Dr. Herman Yeger and

Dr. Sevan Hopyan for their precious discussion and constructive criticism.

I would also like to thank and to give credit to the members of Dror lab, especially to our lab manager Hongbing Li who always has had time for me. He has a huge knowledge and always knows the answers to my questions, providing constructive criticism. I would like to thank our clinical research project manager Bozana Zlateska for acquiring patient samples with informed consent and for always providing us with all the knowledge she has. In addition, I want to express my gratitude to our former research associate Chetankumar Tailor for producing lentivirus, optimizing lentiviral transduction of hiPSCs, and assisting with the establishment of transduced hiPSCs. Also, I want to thank former M.Sc student Steph NG for teaching me and helping with hiPSCs and our former research fellow Stephanie Heidemann for assisting with

PCR work. Moreover, I want to extend my deepest gratitude to PhD candidate Santhosh Dhanraj for his assistance in my experiments, for teaching me the bioinformatics tools required for my work, and especially for his constructive criticism. I would like to express my appreciation to Dr.

Supanun Lauhasurayotin for her helpful contribution to my experiments and for providing her valuable expertise in medicine related aspects of the project. I would also like to thank M.Ss candidate Alejandra Lagos Monzon for always being there for me in hard times. Finally, I would

iii

like to thank our summer student Brian Bursic for helping me to finish experiments for a manuscript during my last summer in the lab.

I would also like to express my appreciation to the facilities that helped us with our project: the

Centre for Commercialization of Regenerative Medicine (CCRM) for creating reprogrammed hiPSC and their characterization. I would like to express my appreciation to the former Sick Kids

Embryonic Stem Cell Facility for providing mouse embryonic fibroblasts (MEFs) for our experiments. I would like to thank The Centre for Applied Genomics (TCAG) for performing

Sanger sequencing and for generating PCR primers. I would like to thank the Sick Kids-

University Health Network Flow Cytometry Facility for fluorescence-activated cell sorting

(FACS) and purity assessment.

From a personal side, I would like to thank my husband for always being there and supporting me, for my parents that helped me a lot and gave me an opportunity to continue my studies. I express a deep and special gratitude to my daughter who is always there for me with a smile.

Lastly and most importantly, I would like to thank all the families that participated in this study, and the funding agencies that supported my work, the Canadian Institute for Health Research and the Nicola’s Kids Triathlon.

Statement of Contributions

My supervisor Dr. Yigal Dror and my committee members Dr. Herman Yeger and Dr. Sevan

Hopyan contributed to the experimental design, data interpretation, and thesis revision and approval. Hongbing Li, Santosh Dhanraj, Steph NG, and Dr. Chetankumar Tailor contributed to the experimental design and data interpretation.

Bozana Zlateska and the Canadian Inherited Marrow Failure Registry acquired patient samples with informed consent. CCRM reprogrammed and characterized hiPSCs. TCAG performed the

Sanger sequencing, whole genome sequencing, and PCR primer generation. The Sick Kids

Embryonic Stem Cell Facility provided mouse embryonic fibroblasts. The Sick Kids-University

Health Network Flow Cytometry Facility performed FACS and provided flow cytometers.

I contributed to the experimental design; performed cell culture for different cell lines, hematopoietic differentiation, colony assays, RT-PCR, western blot, DNA/RNA/protein preparation, whole exome sequencing analysis and gene prioritization, lentiviral transduction, and FACS; acquired flow cytometry data; prepared DNA for Sanger sequencing; performed statistical analysis; interpreted data; and wrote the thesis. I designed and performed the experiments described in the thesis, unless indicated otherwise.

This work was funded by Canadian Institute of Health Research.

Table of Contents

Abstract ...... ii

Acknowledgments...... ii

Statement of Contributions ...... v

Table of Contents ...... vi

List of Abbreviations ...... x

List of Tables ...... xiv

List of Figures ...... xv

Chapter 1 Introduction ...... 1

1.1. Clinical characteristics of the inherited bone marrow failure syndromes...... 1

1.2. Genetic background of the inherited bone marrow failure syndromes ...... 2

1.3. Shwachman-Diamond syndrome ...... 3

1.3.1. Hematological abnormalities ...... 5

1.3.2. Non-hematological abnormalities ...... 10

1.3.3. Treatment for SDS ...... 17

1.3.4. Characterization of Shwachman-Bodian Diamond Syndrome gene SBDS...... 18

1.3.5. Ribosome Biogenesis and SBDS protein function ...... 20

1.3.6. Extra-ribosomal Consequences of SBDS Deficiency ...... 22

1.3.7. SDS Patients without Biallelic SBDS Mutations ...... 27

1.3.8. Characterization of DnaJ Heat Shock Protein Family (Hsp40) Member C21 gene ...... 28

Chapter 2 Research Aims and Hypothesis ...... 31

2.1 Rationale...... 31

2.2 Hypothesis...... 31

2.3 AIM 1: To identify novel SDS-related genetic alterations and novel SDS genes through comprehensive Genome-wide genetic screen ...... 32

2.4 AIM 2: Studying the mechanism of how novel IBMFS-related gene DNAJC21 can cause Shwahman Diamond Syndrome ...... 32

Chapter 3 Methods ...... 33

3.1 Patients and Controls – The Canadian Inherited Marrow Failure Registry (CIMFR) ...... 33

3.2 DNA extraction ...... 34

3.3 Exome sequencing and analysis...... 34

3.4 Web Resources...... 34

3.5 PCR Conditions ...... 35

3.6 RNA extraction ...... 39

3.7 cDNA preparation: ...... 39

3.8 Sanger sequencing ...... 39

3.9 Cell culture ...... 40

3.10 shRNA expression cassettes and generation of DNAJC21-knockdown cells ...... 40

3.11 Viral vector production and transduction ...... 41

3.12 Western Blotting ...... 41

3.13 Antibody list...... 42

3.14 Evaluating cell growth using Alamar Blue Assay ...... 43

3.15 Intracellular Reactive Oxygen Species (ROS) Levels and evaluating ability of cells to balance excess ROS levels generated by addition of H2O2 ...... 43

3.16 Estimation of Cell death by Propidium iodide ...... 44

3.17 Flow cytometry and analysis ...... 45

3.18 Statistical Analysis ...... 47

Chapter 4 Results ...... 48

4.1 IBMFSs and SDS families analyzed by WES analysis ...... 48

4.1.1 Sample preparation ...... 48

4.1.2 Analysis of genetic alterations ...... 49

4.2 The study of DNHD1 as a candidate SDS gene ...... 51 vii

4.2.1 Clinical phenotype and genomic characterization ...... 51

4.2.2 Compound heterozygosity mutations in DNHD1 may cause to reduced protein level ...... 55

4.3 Biallelic mutations in DNAJC21 cause Shwachman-Diamond syndrome ...... 58

4.3.1 Clinical phenotype and genomic characterization ...... 58

4.3.2 Biallelic mutations in DNAJC21 cause to reduced protein level ...... 69

4.3.3 Establishing DNAJC21-knockdown cells ...... 71

4.3.4 The role of DNAJC21 in promoting cell growth ...... 72

4.3.5 DNAJC21-KD Increases Baseline Intracellular Reactive Oxygen Species ...... 73

4.3.6 DNAJC21 is critical for balancing excess of Reactive Oxygen Species ...... 74

4.3.7 DNAJC21 knockdown leads to accelerated cell death ...... 75

4.3.8 The Anti-oxidants rescue downregulation of cell growth in DNAJC21 KD Cells ...76

Chapter 5 Discussion and Conclusion ...... 81

Chapter 6 Future directions ...... 84

6.1 WES analysis ...... 84

6.3 Stroma model ...... 85

6.4 CD34+ cells ...... 85

6.5 iPSc module ...... 86

Chapter 7 Additional work - Abstract ...... 87

Chapter 7.1 Introduction ...... 88

Chapter 7.2 Methods ...... 95

7.2.1 Cell Culture ...... 95

7.2.2 Characterization of hiPSCs ...... 97

7.2.3 Hematopoietic Differentiation ...... 98

7.2.4 Statistical Analyses ...... 103

Chapter 7.3 Results ...... 104

7.3.1 Characterization of hiPSCs ...... 104 viii

7.3.2 Colony Forming Unit Assay ...... 105

7.3.3 CD34+/CD43- Population ...... 109

7.3.4 Lentiviral Transduction of PARN hiPSCs ...... 109

Chapter 7.4 Discussion and future directions ...... 111

Bibliography ...... 113

List of Abbreviations

AA Ascorbic acid AGM Aorta-Gonad-Mesonephros AKT Serine-Threonine Protein Kinase AML Acute myelogenous leukemia AMP Adenosine Monophosphate AMPK AMP-activated Protein Kinase ATP Adenosine Triphosphate BD Becton Dickinson bFGF Recombinant Human Fibroblast Growth Factor Basic Protein BFU-E Blast Forming Unit-Erythroid BM Bone Marrow BMP-4 Recombinant Human Bone Morphogenetic Protein 4 C Cytosine Cat Catalase CCRM Centre for Commercialization of Regenerative Medicine CD Cluster of Differentiation cDNA Complementary DNA CFSSPS Chou & Fasman Secondary Structure Prediction Server CFU Colony Forming Unit CFU-C Colony Forming Unit, In Culture CFU-E Colony Forming Unit, Erythroid CFU-G Colony Forming Unit-Granulocyte Colony Forming Unit-Granulocyte, Erythroid, Monocyte, and CFU-GEMM Megakaryocyte CFU-GM Colony Forming Unit-Granulocyte and Monocyte CHST11 Carbohydrate (Chondroitin 4) Sulfotransferase 11 CIMFR Canadian Inherited Marrow Failure Registry CLIBS Chaperones linked to protein synthesis CMP Common Myeloid Progenitor C-MYC V-Myc Avian Myelocytomatosis Viral Oncogene Homolog Comp-Het Compound Het CSF Colony stimulating factor DAPI 4',6-Diamidino-2-Phenylindole DBA Diamond-Blackfan Anemia DKC Dyskeratosis Congenita DE Differentially Expressed, Differential Expression del(20)(q) Deletion of the Long Arm of Chromosome 20 DL1 Delta Like 1 DMEM Dulbecco's Modified Eagle Medium DMEM/F-12 Dulbecco's Modified Eagle Medium/Nutrient Mixture F-12 DMSO Dimethyl Sulfoxide DNA Deoxyribonucleic Acid E Exon

EB Embryoid Body EFL1 Elongation Factor Like GTPase 1 EHP Early Hematopoietic Progenitors eIF6 Eukaryotic Translation Initiation Factor 6, EIF6 EMP Erythro-Myeloid Progenitor EPO Recombinant Human Erythropoietin ESC Embryonic Stem Cell FA Fanconi Anemia ESP Exome Variant Server EXAC The Exome Aggregation Consortium FACS Fluorescence-Activated Cell Sorting FBS Fetal Bovine Serum FCF Flow Cytometry Facility FITC Fluorescein Isothiocyanate FITC-CD43 FITC Mouse Anti-Human CD43 Flt-3L Fms-related Tyrosine Kinase 3 Ligand, mouse FMO Fluorescence-Minus-One FYSH Fungal, Yhr087wp, Shwachman FZD2 Frizzled Class Receptor 2 G Guanine GC Guanine and Cytosine GCDR Gentle Cell Dissociation Reagent G-CSF Granulocyte-colony stimulating factor gDNA Genomic DNA GFR Growth Factor Reduced GMP Granulocytic-Monocytic Progenitor H2-DCFDA 2',7'-dichlorodihydrofluorescein diacetate HE Hemogenic Endothelium hESC Human embryonic stem cells HgF Fetal hemoglobin Het Heterozygous hiPSC Human induced pluripotent stem cell Hom Homozygous Hr Hour HSC/Ps Hematopoietic Stem Cell/progenitors HSCT Hematopoietic Stem Cell Transplantation i(7)(q10) Isochromosome of the Long Arm of Chromosome 7 IBMFS Inherited bone marrow failure syndrome ID Identification IGF-I Recombinant Human Insulin-like Growth Factor I IGV Integrative Genome Viewer IL-11 Recombinant Human Interleukin 11 IL-3 Recombinant Human Interleukin 3 IL-6 Recombinant Human Interleukin 6 IMDM Iscove's Modified Dulbecco's Medium

iPSC Induced Pluripotent Stem Cell IQ Intelligence Quotient IRES Internal ribosome entry site Kb Kilobases KD Knock-Down kDa Kilodalton KLF4 Kruppel-like Factor 4 KOSR KnockOutTM Serum Replacement Leu Leucine MDS Myelodysplastic syndrome MEF Mouse Embryonic Fibroblast MOCK Transduction without DNA MOI Multiplicity of Infection MP Myeloid Progenitors MPP Multipotent Progenitor mRNA Messenger RNA mTOR Mechanistic Target of Rapamycin Mut Mutation N Normal NA Not Applicable NAC N-acetylcysteine NADH Nicotinamide Adenine Dinucleotide ncRNAs A non-coding RNA NEAA Non-essential amino acids NK Natural Killer Oct4 Octamer-binding Transcription Factor 4 OP9-DL1 OP9-Delta-like 1 p p-value P Patient p53 Tumor protein p53 PARN Poly(A)-Specific Ribonuclease PBS Phosphate Buffered Saline PCR Polymerase Chain Reaction PE Phycoerythrin PE-CD43 PE Mouse Anti-Human CD43 PE-Cy7 Phycoerythrin-Cyanine 7 PE-Cy7-CD34 PE-Cy7 Anti-Human CD34 PI Propidium Iodide PI3K Phosphoinositide 3-Kinase Pro Proline PSC Pluripotent Stem Cell QD Quality by Depth qRT-PCR Quantitative Reverse Transcription-PCR RBC Red blood cell RIPA Radioimmunoprecipitation assay buffer xii

RNA Ribonucleic Acid RNAi RNA Interference RNA-seq RNA Sequencing ROS Reactive oxygen species rRNA Ribosomal RNA RT-qPCR Real Time Polymerase Chain Reaction SBDS Shwachman-Bodian-Diamond Syndrome SBDSP1 Shwachman-Bodian-Diamond Syndrome Pseudogene 1 scaRNAs Small Cajal body-specific RNAs SCF Recombinant Human Stem Cell Factor SD Standard Deviation SD Standard Bias SDS Shwachman-Diamond Syndrome SEM Standard Error Of Mean SFM Serum Free Medium SIFT Sorts Intolerant From Tolerant amino acid substitutions snoRNAs Small nucleolar RNAs SNP Single Nucleotide Polymorphism SOX2 SRY (sex determining region Y)-box 2 SP34 StemPro®-34 Serum Free Medium SSEA-4 Stage-Specific Embryonic Antigen 4 T# # of day in differentiation TBST Tris Buffered Saline with Tween 20 TCAG The Centre for Applied Genomics TCF T Cell Factor TERC Telomerase RNA Component Tif6 Translation Initiation Factor 6, yeast (human gene EIF6) Tm Melting Temperature TOE1 Target of EGR1 protein 1 TPO Recombinant Human Thrombopoietin Genome browser hosted by the University of California, Santa UCSC Cruz uORF Upstream Open Reading Frame UTR Untranslated Region VDAC1 Voltage Dependent Anion Channel 1 VEGF Recombinant Human Vascular Endothelial Growth Factor WES Whole Exome Sequence WNT Wingless-Type MMTV Integration Site WNT10B WNT Family, Member 10B WT Wild-Type YD# Number of shRNA αMEM Minimum Essential Medium α

xiii

List of Tables

Table 1: Mutation Effect Prediction for family of patient 1...... 52

Table 2: Mutation Effect Prediction for Family 2...... 54

Table 3: Patients with mutations in DNAJC21 meet the clinical criteria for SDS ...... 62

Table 4: Information about DNAJC21 and C2orf71 variants found in the patients in the present study ...... 64

Table 5: Reagents used in the maintenance of OP9-DL1 culture...... 96

Table 6: Reagents used in the hematopoietic differentiation of hiPSCs cultured on MEFs...... 98

Table 7: Fluorophores and cell density of PI only control, FMO controls, and sorting sample of T9 staining reactions...... 102

Table 8: hIPSC lines used in the present study ...... 104

xiv

List of Figures

Figure 1: Flow for gene periodization from WES data...... 50

Figure 2: Exon 7 deletion in DNHD1 in patient 1 was inherited from father...... 52

Figure 3: DNHD1-associated mutations in Patient 1...... 53

Figure 4: DNHD1-associated mutations in Patient 2...... 55

Figure 5: Cellular localization of DNHD1...... 57

Figure 6: Clinical characteristics of patient 1...... 63

Figure 7: Clinical characteristics of patient 3...... 63

Figure 8: DNAJC21-associated mutations in Patient 1...... 65

Figure 9: Region on homozygosity on chromosome 5 overlap the DNAJC21 locus in patients from family 2 ...... 65

Figure 10: DNAJC21-associated mutations in Patient 3...... 66

Figure 11: DNAJC21 protein domains...... 66

Figure 12: The effect of the p.K34E mutation in DNAJC21 on the protein secondary structure. 67

Figure 13: Pedigree of family 3...... 67

Figure 14: Genetic investigation of Family 3...... 69

Figure 15: Expression of DNAJC21 protein ...... 71

Figure 16: DNAJC21 Protein expression in DNAJC21- knockdown HEK 293T cells...... 72

Figure 17: Cell growth rate is impaired in DNAJC21 knockdown cells...... 73

Figure 18: Baseline ROS levels are increased in DNAJC21 knockdown cells...... 74

Figure 19: The ability to balance ROS levels after addition of exogenous H2O2 is impaired in DNAJC21-knockdown cells...... 75 xv

Figure 20: Cell death is increased in DNAJC21-knockdown cells...... 76

Figure 21: Antioxidants decrease ROS levels and improve cell growth rate in DNAJC21- knockdown cells...... 80

Figure 22: PARN protein expression in hiPSCs...... 104

Figure 23: Analysis of hemogenic endothelium containing cells within the CD34+/CD43- population in T9 embryoid bodies derived from normal iPSC by flow cytometry...... 107

Figure 24: Clonogenic potential of PARN-deficient hiPSCs...... 108

Figure 25: Hemogenic endothelium cell population at day 9 of culturing was decreased compared to controls...... 109

Figure 26: hiPSC colonies transduced with a lentivurs carrying wild type PARN...... 110

xvi

Chapter 1 Introduction

1.1. Clinical characteristics of the inherited bone marrow failure syndromes Inherited bone marrow failure syndromes (IBMFSs) are rare genetic diseases characterized by

varying degrees of defective production of erythrocytes, granulocytes, and platelets in the bone

marrow, leading to anemia, thrombocytopenia, neutropenia or aplastic anemia(Burke, Colebatch

et al. 1967)(Bodian, Sheldon et al. 1964, Shwachman, Diamond et al. 1964, Dror 2005, Dror,

Donadieu et al. 2011). The most common IBMFSs include Diamond-Blackfan anemia, Fanconi

anemia and Shwachman-Diamond syndrome(Aggett, Cavanagh et al. 1980) Arbiv et al, Clin

Genet 2018). These disorders may involve multiple lineages (eg, Fanconi anemia) or

predominantly single cytopenias (eg, Kostmann neutropenia, Diamond-Blackfan anemia and

thrombocytopenia with absent radii syndrome) and, hence, have been commonly classified

according to the lineages that were affected. Our lab has shown that about 20% of the cases of

IBMFS cannot be categorized into the known subclasses and were named unclassifiable. The

latter may contain either novel syndromes or atypical presentations of the known IBMFSs. Many

IBMFSs are cancer-predisposition disorders. Various types of malignancies can develop in the

affected patients(Dror 2005). The most common malignancies are myelodysplastic syndrome

and acute myeloid leukemia (Huijgens, van der Veen et al. 1977, Alter, Baerlocher et al. 2007).

In addition to hematological and oncological manifestations, a broad range of physical

abnormalities are associated with IBMFSs, with significant overlap among the disorders. These

physical anomalies include skeletal, cardiovascular, craniofacial, gastrointestinal,

immunological, dermatological, renal, neurological and endocrine anomalies(Dror 2005, Dror,

Donadieu et al. 2011).

1.2. Genetic background of the inherited bone marrow failure syndromes

As of June 30th 2018, 126 genes associated with IBMFS were identified. In some of the disorders

multiple genes are affected. For example, there are currently 21 Fanconi anemia genes(Dong,

Nebert et al. 2015, Bagby 2018), 10 Diamond Blackfan anemia genes and 3 dyskeratosis congenita

genes (Tsangaris, Klaassen et al. 2011, Ghemlas, Li et al. 2015). The IBMFS genes are involved

in numerous cellular or biochemical pathways. For example, The Fanconi anemia genes are

involved in DNA repair; the dyskeratosis congenita genes are associated with telomere

maintenance; both the Diamond-Blackfan anemia and the Shwachman-Diamond anemia genes are

involved in ribosome biogenesis. Many of the genes are multifunctional and play roles in diverse

cellular pathways. Identifying and studying genes that are mutated in IBMFSs is of utmost clinical

importance. First, in simplex families when only one case is identified genetic diagnosis helps to

quickly rule out acquired etiologies for the bone marrow failure and determine the genetic cause.

Second, the inheritance mode among families with similar presentation or similar disease can be

different. For example, dyskeratosis congenita can be inherited in an autosomal dominant,

autosomal recessive or X-linked mode. Therefore, to provide accurate genetic counseling and to

properly predict the risk of recurrence, the specific gene that is mutated has to be identified. Third,

some genotype-phenotype correlations are known to exist in IBMFSs, thus identifying the mutated

genes may help with outcome prediction (for example, whether severe aplastic anemia or cancer

are likely to occur). Fourth, in the novel therapies will likely depend on the genetic background of

the patients. These may include gene therapy, pre-implantation genetic diagnosis, as well as

selection of unaffected and HLA-matched donors for hematopoietic stem cell transplantation

during in vitro fertilization. Hence, a lot of gaps exist in our understanding of the genetic basis of

IBMFSs. Data from our lab indicates that although more than 120 genes mutated in IBMFS have

been identified, genetic diagnosis of IBMFSs can only be made in about 60% of the patients from

our database. Identification of IBMFS related genes in well characterized IBMFSs and in

unclassifiable IBMFSs cases is very challenging and additional diagnostic tools are required.

Metaphase cytogenetics (MC) is a widely used method to identify genomic alterations in several

IBMFSs and coinciding disrupted genes(Draptchinskaia, Gustavsson et al. 1999, Farrar, Nater et

al. 2008). Unfortunately, it has low resolution and cannot determine alterations which are smaller

than 3Mb. In addition, many IBMFS genes have been identified by linkage analysis(Heiss, Knight

et al. 1998, Boocock, Morrison et al. 2003) that requires multiplex families or a large number of

patients from simplex families. Therefore, the rarity of the IBMFSs renders this approach very

challenging (Ginzberg, Shin et al. 1999, Willig, Niemeyer et al. 1999, Teo, Klaassen et al. 2008).

The approach using targeted specific genes has been successful in identifying several IBMFSs

(CMPL, RPL5, RPL11). However, this method has a low probability of producing positive results

in IBMFSs, since multiple genes can possibly be mutated. These limitations can be overcome by

a genome-wide high-resolution approach using whole exome and genome sequencing methods

that were done in this project.

1.3. Shwachman-Diamond syndrome

Shwachman Diamond Syndrome (SDS) is an inherited bone marrow failure syndrome (IBMFS)

first described in 1964 by Shwachman and Bodian(Bodian, Sheldon et al. 1964, Shwachman,

Diamond et al. 1964). It is an autosomal recessive, multisystem disorder, characterized most

commonly by three major clinical symptoms of bone marrow failure, exocrine pancreatic insufficiency, and skeletal abnormalities. Patients may further present with hepatic complications, immunological abnormalities, renal and cardiac disease(Dror and Freedman 1999,

Dror and Freedman 2001, Dror, Ginzberg et al. 2001, Dror 2005, Dror, Donadieu et al. 2011).

Also, patients with SDS can develop insulin-dependent diabetes, growth-hormone deficiency, hypogonadotropic hypogonadism, cognitive impairment and oral disease. Similar to other

IBMFSs, SDS is characterized by a high tendency to develop myelodysplastic syndrome (MDS) and leukemia, specifically acute myelogenous leukemia (AML) – with an estimated risk of 36% at 30 years old (Woods, Roloff et al. 1981, Dror, Squire et al. 1998). The frequency of SDS was estimated as 1 in 77,000 child births, with no gender or racial bias (Ginzberg, Shin et al. 1999,

Goobie, Popovic et al. 2001, Alter, Giri et al. 2010). Data from the Canadian Inherited Marrow

Failure Registry (CIMFR) indicates that SDS is the third most common disorder among IBMFS; with the first one being Diamond-Blackfan Anemia (DBA) and the second being Fanconi

Anemia (FA)(Arbiv et al, Clin Genet 2018). SDS has an estimated median survival rate of about

35 years(Tsangaris, Klaassen et al. 2011). The molecular basis of the pleiotropic phenotype characteristic of SDS remains unclear, however in 2003 significant advances made by Boocock

& colleagues have helped shed some light on the genetic basis of SDS syndrome. In 2003,

Boocock & colleagues identified hypomorphic disease-associated mutations in the Shwachman

Bodian Diamond syndrome gene (SBDS ) in approximately 90% of SDS patients(Boocock,

Morrison et al. 2003). Several groups have related SBDS to ribosome biogenesis, mitotic spindle assembly, cell survival, chemotaxis, and the formation and activity of hematopoietic stem cells and their progenitors in the bone marrow. Also, it was shown that SBDS is important in the regulation of reactive oxygen species (ROS) and the maintenance of genomic stability (Dror and

Freedman 1999, Dror and Freedman 2001, Stepanovic, Wessels et al. 2004, Wessels, Srikantha

et al. 2006, Ganapathi, Austin et al. 2007, Austin, Gupta et al. 2008, Rujkijyanont, Watanabe et

al. 2008, Rujkijyanont, Adams et al. 2009, Watanabe, Ambekar et al. 2009, Sen, Wang et al.

2011, Burwick, Coats et al. 2012). While identification of the alterations in SBDS gene has

become an important tool in the diagnosis of SDS and recent studies have helped elucidate some

of its functions, further studies are required to define the pathogenic link between SBDS

genotype and the SDS phenotype as well as identify additional genes associated with SDS in

about 10% of patients that lack SBDS mutations.

1.3.1. Hematological abnormalities

SDS is the 3rd most commonly recognized inherited bone marrow failure after Diamond-

Blackfan anemia and Fanconi anemia(Dror 2005). The major concerns of mortality in SDS are

Hematological complications. The hematological features associated with SDS include cytopenia

(at least one line affected: neutropenia, anemia, or thrombocytopenia), elevated fetal

hemoglobin, macrocytic red blood cells (RBC), and hypo-cellular bone marrow(Aggett,

Cavanagh et al. 1980). Additional hematological complications include aplastic anemia,

myelodysplastic syndrome, clonal marrow cytogenetic abnormalities, and acute myeloid

leukemia.

Neutropenia. Neutropenia (neutrophil count is below 1.5×109cells/L) is the most commonly

observed cytopenia in SDS(Mack, Forstner et al. 1996). This cytopenia is seen in 74-98% of

SDS cases(Ginzberg, Shin et al. 1999, Hashmi, Allen et al. 2011, Myers, Bolyard et al.

2014).The severity of neutropenia in SDS can be mild (neutrophil count between 1.0-1.5×109

cells /L) , moderate (neutrophil count between 0.5-1.0×109 cells /L), or severe (neutrophil count

below 0.5×109 cells /L)(Hashmi, Allen et al. 2011). Neutropenia in SDS is often periodic but can

also be persistent (Ginzberg, Shin et al. 1999). Neutropenia in SDS patients can be detected later

in life and not necessarily be seen in newborns (Dror, Donadieu et al. 2011).

Anemia. Anemia (hemoglobin levels below 70g/L) is asymptomatic in SDS. It is the second

most commonly observed cytopenia in SDS, manifesting in 42-58% of SDS patients (Ginzberg,

Shin et al. 1999, Hashmi, Allen et al. 2011). The severity of anemia in patients is often mild to

moderate (hemoglobin levels above 70g/L but below the age- and gender-adjusted lower limit)

rather than severe (hemoglobin levels below 70g/L) as it can be seen in neutropenia.

Thrombocytopenia. Thrombocytopenia (platelet count below 150×109cells/L) is the least

commonly observed cytopenia in SDS. This cytopenia is seen in 34-38% of SDS patients

(Ginzberg, Shin et al. 1999, Hashmi, Allen et al. 2011). The severity of thrombocytopenia in

SDS can be mild (platelet count between 100-150×109 cells /L) but is frequently moderate

(platelet count between 10-100×109 cells /L) and is rarely severe (platelet count below 20×109

cells /L).

Lymphopenia. Lymphopenia (lymphocyte count below 1.5×109 cells /L) is usually not observed

in SDS (Dror and Freedman 2001Dror, Ginzberg et al. 2001). Although most SDS patients do

not present with overt lymphopenia, a study of the immune function of SDS patients uncovered

various lymphoid defects, such as reduced numbers of T-cells, B-cells, and natural killer (NK)

cells. In addition, defects in humoral immunity included reduced serum IgG and its subclasses,

reduced B-lymphocyte proliferation in vitro, reduced circulating B-lymphocytes, and reduced

antibody production. Defects in cellular immunity included reduced circulating T-lymphocytes

and reduced T-lymphocyte proliferation in vitro.

Macrocytic red blood cells. Red blood cells can be normocytic or normochromic but are mainly

macrocytic in SDS occurring in 60% of SDS patients after 1 year of age (Dror, Donadieu et al.

2011, Hashmi, Allen et al. 2011).

Elevated fetal hemoglobin levels. Elevated levels of fetal hemoglobin levels are observed in

72% of SDS patients after 1 year of age(Dror, Donadieu et al. 2011, Hashmi, Allen et al. 2011).

Hypocellular bone marrow. Bone marrow cellularity is usually hypo cellular (cellularity below

25% or cellularity below 50% combined with hematopoietic cells below 30%) in SDS with fat

replacement. It is very rare that patients will have normal or hyper cellular bone marrow

(Ginzberg, Shin et al. 1999, Dror, Donadieu et al. 2011, Hashmi, Allen et al. 2011). Patient`s

bone marrow commonly shows reduced granulopoiesis, arrest in the myeloid lineage

differentiation, and reduced megakaryopoiesis or erythropoiesis (Dror, Donadieu et al. 2011,

Hashmi, Allen et al. 2011).

Aplastic anemia. Aplastic anemia is defined as bilineage or trilineage cytopenia combined with

hypocellular bone marrow in the absence of fibrosis, dysplastic morphology, or clonal evolution.

SDS patients can present with more than one hematopoetic line affected (pancytopenia). In 24%

of SDS patients with bi-lineage cytopenia and in 19-24% with pancytopenia, more than two lines

are affected (Ginzberg, Shin et al. 1999, Hashmi, Allen et al. 2011). SDS can progress to aplastic

anemia in 15% of SDS patients at the average age of 3.9 years (Dror, Donadieu et al. 2011,

Hashmi, Allen et al. 2011). The severity of aplastic anemia in SDS can be moderate, when at

least two of reticulocyte counts are below 40×109cells/L, platelet count is below 40×109 cells /L,

or neutrophil count is below l.5×109 cells /L, or severe, when at least two of reticulocyte counts

are below 40×109 cells /L, platelet count is below 20×109 cells /L, or neutrophil count is below

0.5×109 cells /L. Aplastic anemia in SDS can be transient or persist, and has been diagnosed in 7

newborns (Kuijpers, Nannenberg et al. 2004, Dror, Donadieu et al. 2011). SDS patients with

poor prognosis are indicated by age of diagnosis (before 3 months) and low hematological

parameters in the first three complete blood counts (either neutrophil count below 0.5×109 cells

/L, hemoglobin levels below 9g/dL, or platelet count below 100×109 cells /L). These patients

have 59% risk of developing aplastic anemia by the age of 10 to 20 years (Donadieu, Fenneteau

et al. 2012).

Clonal marrow cytogenetic abnormalities. SDS patients may acquire chromosomal

abnormalities in bone marrow cells; i.e., clonal marrow cytogenetic abnormalities (Dror,

Donadieu et al. 2011). The cause for acquisition of cytogenetic abnormalities and genomic

instability in SDS is unknown, but is possibly an outcome of microtubule instability (Austin,

Gupta et al. 2008). Clonal marrow cytogenetic abnormalities in SDS can be stable, trade in

numbers, occur occasionally, regress, or even progress to myelodysplastic syndrome and acute

myeloid leukemia (Dror, Durie et al. 2002, Smith, Shaw et al. 2002). Around 44% of cytogenetic

alterations in SDS are i(7)(q10) (isochromosome of the long arm of chromosome 7), 33% are

other abnormalities in chromosome 7 (deletion, monosomy, and translocations), and the

remaining 16% are del(20)(q) (interstitial deletions of the long arm of chromosome 20) (Dror

2005). Variant i(7)(q10) is the most common cytogenetic alteration in SDS (Dror 2005).

i(7)(q10) leads to duplication of the long arm of this chromosome and overall three copies of

SBDS, which resides on 7q11. If i(7)(q10) coincides with the splice site mutation (c.258+2T>C)

of SBDS, the duplication of c.258+2T>C can result in increased expression of the SBDS splice

variant (Minelli, Maserati et al. 2009). Thus, i(7)(q10) has been proposed to be a compensatory

mechanism to restore SBDS expression. Variant del(20)(q) or int del(20)(q11.21q13.32) is the

second most frequent cytogenetic alteration in SDS and results in the deletion of EIF6 on

20q11.22 (Pressato, Valli et al. 2012). Therefore, del(20)(q) has been proposed to be a

compensatory pathway to restore ribosome biogenesis by bypassing the requirement for eIF6 to

be released from 60S ribosome subunit. The above hypotheses may explain a relatively good

prognoses associated with i(7)(q10) and del(20)(q), although other cytogenetic alterations with

unknown prognoses are often present at the same time (Pressato, Valli et al. 2015). Opposite to

the aforementioned relatively benign course of i(7)(q10) and del(20q) cytogenetic abnormalities,

malignant prognosis is most commonly observed in monosomy 7 cytogenetic abnormality. It is

observed in 50% of SDS patients with acute myeloid leukemia or myelodysplastic syndrome

(Donadieu, Fenneteau et al. 2012).

Myelodysplastic syndrome. Clonal marrow cytogenetic abnormalities in SDS can progress to

malignant transformation to myelodysplastic syndrome (cytopenia and either presence of MDS-

type clonal marrow cytogenetic abnormality or marrow myeloblast count between 5–29%) (Dror,

Donadieu et al. 2011). Around 18% of SDS patients developed malignant and clonal myeloid

transformation by the age of 20 years(Hashmi, Allen et al. 2011). Myelodysplastic syndrome

may feature hypoplastic bone marrow, and may be difficult to distinguish from aplastic anemia.

Mild and transient dysplastic changes can be seen in myeloid, erythroid, and megakaryocytic

lineages, but they frequently cannot help differentiate between plastic anemia and

myelodyspastic syndrome.

Acute myeloid leukemia. Myelodysplastic syndrome in SDS can progress to acute myeloid

leukemia, which is defined by the presence at least 30% marrow myoblast count (Dror, Donadieu

et al. 2011). About 50% of SDS patients with myelodysplastic syndrome undergo advanced

progress to leukemia with high predominance in male (Dror, Donadieu et al. 2011). Multiple

subtypes of acute myeloid leukemia have been reported: M0 - undifferentiated acute

myeloblastic leukemia, M1 - acute myeloblastic leukemia with minimal maturation, M2 - acute

myeloblastic leukemia with maturation, M4 - acute myelomonocytic leukemia, M5 - acute

monocytic leukemia, and M6 - acute erythroid leukemia, M7-acute megakaryocytic leukemia.

M6 is a rare type of acute myeloid leukemia, but has been reported more commonly in patients

with SDS who develop leukemia(Smith, Hann et al. 1996, Dokal, Rule et al. 1997, Lesesve,

Dugue et al. 2003, Mitsui, Kawakami et al. 2004). Acute lymphoblastic leukemia (Strevens,

Lilleyman et al. 1978) and juvenile myelomonocytic leukemia (Caselitz, Kloppel et al. 1979)

have also been reported in SDS patients.

Recurrent infections. As a result of neutropenia, bacterial infections are common in SDS,

especially during early childhood (Ginzberg, Shin et al. 1999, Dror and Freedman 2001, Dror,

Ginzberg et al. 2001, Hashmi, Allen et al. 2011). Recurrent bacterial infections can manifest as

sinusitis, otitis media, or pneumonia. Recurrent viral infections can present as gastroenteritis,

respiratory tract infection, and unknown source of fever. Severe systemic or deep tissue

infections can cause an oral cellulitis, dental abscess, septicemia, abscess, staphylococcal skin

infection, and osteomyelitis.

1.3.2. Non-hematological abnormalities

1.3.2.1. Pancreatic Dysfunction

SDS is the second most commonly observed pancreatic dysfunction after cystic fibrosis and

malabsorption. It is one of the major concerns for mortality and morbidity early in life in SDS

(Dror 2005). About 74 to 85% of SDS patients have pancreatic dysfunction and it is often

diagnosed before 6 months of age (Mack, Forstner et al. 1996, Ginzberg, Shin et al. 1999,

Hashmi, Allen et al. 2011). 10

Pathological changes in the pancreas. About 83% of SDS patients present with various levels

of pancreatic lipomatosis (Ginzberg, Shin et al. 1999, Myers, Bolyard et al. 2014). The pancreas

is fatty with intact islets of Langerhans and ductal structure, small or enlarged, and often not

assessed. Lipomatosis can also develop or disappear with age.

Exocrine pancreatic enzymes. Acinar cells are exocrine cells of the pancreas that produce and

transport enzymes that are passed into the duodenum where they assist in the digestion of food.

In SDS, these cells are reduced in number and size and the exocrine pancreas is replaced by fat.

The secretion of exocrine pancreatic enzymes is insufficient in SDS.

The diagnosis of exocrine pancreatic insufficiency in SDS is based on 1) the levels of exocrine pancreatic enzymes; e.g. serum trypsinogen levels below 16U/L, fecal elastase levels below

200μg/g (Ip, Dupuis et al. 2002, Dror, Donadieu et al. 2011). As a result of exocrine pancreatic insufficiency fecal fat loss using the 72-hr fat balance study is increased (Ginzberg, Shin et al.

1999, Dror, Donadieu et al. 2011). After the age of 3 years, serum isoamylase levels below

6μg/L is also very useful. Ultrasound and computed tomography of the pancreas commonly show the signal characteristic of lipomatosis. About 75-80% of SDS patients showed evidence pancreatic insufficient based on these criteria [Hashmi et al, Clin Genet 2011](Ginzberg, Shin et al. 1999).

Malabsorption. As the result of insufficient secretion of exocrine pancreatic enzymes, about

86% of SDS patients presented with steatorrhea and 58% of SDS patients presented with

diarrhea (Ginzberg, Shin et al. 1999, Hashmi, Allen et al. 2011, Myers, Bolyard et al. 2014).

Digestion and absorption of fats and fat-soluble vitamins (A, D, E, and K) are also reduced.

Pancreatic sufficiency. The severity of exocrine pancreatic insufficiency can change with age

and SDS patients can eventually become pancreatic sufficient as a result of spontaneous

embellishment of nutrient absorption (Ginzberg, Shin et al. 1999, Dror, Donadieu et al. 2011,

Hashmi, Allen et al. 2011). Frequently after 4 years of age, around 50% of SDS patients become

pancreatic sufficient and reach proper trypsinogen levels but fail to reach proper amylase levels

(Mack, Forstner et al. 1996, Ip, Dupuis et al. 2002). It is noteworthy that males (41%) are more

likely to become pancreatic sufficient than females (16%)(Ginzberg, Shin et al. 1999).

Growth delay. Growth delay is commonly observed before one year of age in SDS patients

(Dror, Donadieu et al. 2011). About 73% present with failure to thrive and about 61% have with

growth delay. Growth failure can present in utero, and about 26% of SDS patients suffer from

intrauterine growth delay (Hashmi, Allen et al. 2011, Donadieu, Fenneteau et al. 2012, Myers,

Bolyard et al. 2014). The mean birth weight is at the 25th percentile. Height and weight are

commonly below the 3rd percentile at the first year of age despite enzyme treatment (Mack,

Forstner et al. 1996, Ginzberg, Shin et al. 1999).

Endocrine pancreas. Although the islets of Langerhans (the regions of the pancreas that contain its endocrine, hormone-producing, cells) appear normal in pancreatic biopsies, rare cases of endocrine pancreatic dysfunction causing impaired insulin production have been reported in SDS(Hashmi, Allen et al. 2011, Myers, Bolyard et al. 2014). The frequency of type 1 diabetes mellitus in SDS was predicted to be 30 times higher than in control population(Gana, Sainati et al. 2011).

1.3.2.2. Skeletal Abnormalities

Another common abnormality that can be observed in SDS is skeletal abnormality. Burke et al.

in 1967 and Pringle et al. in 1968 were the first to report occurrences of metaphyseal dysplasia in

SDS. About 76% of SDS patients present with short stature and overall around 49-73% present

with skeletal abnormalities. Metaphyseal dysostosis is a typical finding in SDS (46%) and can

be detected before or at birth (Beser, Cokugras et al. 2014) or be noticed later on. Other

anomalies include metaphyseal osteopenia, thorax narrowing and dystrophy (32%), scoliosis.

Rare cases with polydactyly have been reported. Severe bone complications that required surgery

occur in 9% of the patients (Ginzberg, Shin et al. 1999, Hashmi, Allen et al. 2011, Donadieu,

Fenneteau et al. 2012).

Skeletal development. Skeletal abnormalities in SDS patients are diverse with severity and

location changing with age(Makitie, Ellis et al. 2004). Based on longitudinal radiographic

analyses of patients with SDS a delayed presence of secondary ossification centers can be

observed and that usually normalizes with age. In addition, the growth plates showed advanced

irregularity and thickening that is related to asymmetrical growth. Metaphyseal and growth plate

abnormalities aggravate with age, and epiphyseal maturation usually normalize with age.

Impaired skeletal development in children with SDS causes short ribs, short limbs, narrowed

thorax, coxa valga, bowing legs, and short stature. Already before age of 5 years the thinning of

cortical bone, early signs of osteoporosis and osteopenia have been observed.

Osteoporosis. Patients with SDS have higher risk of early onset osteopenia and osteoporosis that

are typified by reduced bone mineral density and content, fragility fractures, and vertebral

deformities (Toiviainen-Salo, Mayranpaa et al. 2007). Reduced number of osteoblasts and

osteoclasts lead to a low bone turnover that eventually can cause osteoporosis in SDS patients.

This leads to a reduced numbers of osteoids and a reduced trabecular bone volume.

1.3.2.3. Cognitive and Neurological Abnormalities

Cognitive abnormalities. About 59% of patients with SDS present with developmental delay.

Cognitive function in SDS patients is commonly impaired as assessed by lower intelligence

quotient (IQ), behavioral issues, learning difficulties, and social challenges(Kent, Murphy et al.

1990, Kerr, Ellis et al. 2010). Cognitive defect can affect both children (65%) and adults (76%)

with SDS (Perobelli, Nicolis et al. 2012). Children with SDS can also develop behavioral

disorders including attention deficit hyperactivity disorder, oppositional defiant disorder, and

behavioral issues such as somatic complaints, withdrawal, depression, anxiety, acting out, and

impulsive, uncooperative, aggressive behaviors and autism spectrum disorder. Neur(Kerr, Ellis et

al. 2010). Neuro-psychological tests showed lower intellectual reasoning in children with SDS.

About 20% of children with SDS presented with intellectual disability, 20% presented with

severely impaired perceptual intelligence, and 12% presented with severely impaired verbal

intelligence. In addition, perceptual skills, higher-order language skills, attention, memory, and

academic achievement were weaker in children with SDS. Due to combination of cognitive,

behavioral, and medical difficulties functional independence was also lower in children with

SDS.

Neurological abnormalities. It was shown by Toiviainen-Salo et al. in 2008, that the volumes

of the brain, white matter, and grey matter were reduced in SDS patients. Also, brain regions

responsible for learning, verbal memory, cognition, and IQ were reduced in SDS patients

(Toiviainen-Salo, Makitie et al. 2008, Booij, Reneman et al. 2013). However, in 2013, Booij et

al. demonstrated that increased central striatal dopamine transporters in SDS patients are

associated with impaired memory and attention. Furthermore, in 2015, Perobelli et al. showed

that increased cortical thickness and altered connectivity in the white and grey matter were

connected to cognitive impairment. Clusters of altered fibers were shown to interfere with inter-

and intra-hemispheric connections essential for verbal skills, perceptual skills, visual-motor

integration, memory, and executive functions(Perobelli, Alessandrini et al. 2015). In addition to

aforementioned impairments, other abnormalities such as Chiari malformation type I, cerebellar

tonsillar ectopia, and myopathy and hypotonia can be seen in 5% of SDS patients (Myers,

Bolyard et al. 2014).

1.3.2.4. Abnormalities in other biological systems

Cardiac Abnormalities. Congenital heart disease and cardiomyopathy such as, atrial septal

defect and patent foramen ovale, patent ductus arteriosus, atrioventricular septal defect,

ventricular septal defect, and dilated cardiomyopathy have been reported in 11-19% of SDS

patients (Kopel, Gutierrez et al. 2011, Hauet, Beaupain et al. 2013, Le Gloan, Blin et al. 2014,

Myers, Bolyard et al. 2014). Heart failure in children with SDS has been proposed to be

connected with myocardial necrosis and fibrosis (Guerrero, Lopez Barea et al. 1979, Nezelof and

LeSec 1979). Myocardial fibrosis and progression to heart failure have been suggested to be a

result of increased susceptibility to carditis during concomitant viral infections and myocardial

damage in SDS (Savilahti and Rapola 1984, Hauet, Beaupain et al. 2013). Studies in cardiac

function of SDS patients showed differences in the right ventricular diastolic function, left

ventricular contractile reserve, left ventricular systolic function, and circumferential strain, which

were proposed to lead to heart failure (Toiviainen-Salo, Makitie et al. 2008, Toiviainen-Salo,

Pitkanen et al. 2008, Toiviainen-Salo, Raade et al. 2008, Ryan, Jefferies et al. 2015).

Liver abnormalities. Liver abnormalities in SDS are usually asymptomatic and can manifest in

early childhood, however, they often improved with age (Brueton, Mavromichalis et al. 1977,

Ginzberg, Shin et al. 1999). Around 60% of SDS patients have elevated serum aminotransferase

and around 15% of them are presented with hepatomegaly. Microvesicular and macrovesicular

steatosis, periportal and portal inflammation, periportal, portal, and bridging fibrosis and

glycogenosis were seen in liver biopsies of SDS patients.

Endocrine abnormalities. Endocrine abnormalities occur in about 65% of SDS patients (Myers,

Bolyard et al. 2014). Patients can present with low stimulated growth hormone levels,

hypopituitarism, abnormal thyrotropin levels, abnormal glucose levels, hypothyroism, elevated

follicle-stimulating hormone level, adrenal insufficiency, and postprandial hyperglycemia

(Myers, Rose et al. 2013).

Non-pancreatic gastrointestinal abnormalities. Gastrointestinal abnormalities are diverse but

relatively uncommon in SDS. About 8% of SDS patients have gastrointestinal abnormalities

such as bilateral inguinal hemia, malrotation, and imperforate anus (Myers, Bolyard et al. 2014).

Around 19% of SDS patients showed evanescent but severe gastrointestinal symptoms requiring

enteral or parenteral feeding or gastrostomy at a median age of only 7 months (Donadieu,

Fenneteau et al. 2012).

Dental diseases. About 44% of SDS patients showed oral diseases, including carries in primary

and permanent dentitions, enamel hypoplasia, periodontal disease, and delayed dental

development (Ginzberg, Shin et al. 1999, Ho, Cheretakis et al. 2007, Hashmi, Allen et al. 2011).

Others. Eczema is common in SDS (65%), other abnormalities such as pulmonary hypertension,

urological abnormalities (Testicular atrophy and hypospadias), eye anomaly, subglottic stenosis,

labial cleft, and ear anomalies (hearing loss) are rare (Hashmi, Allen et al. 2011, Donadieu,

Fenneteau et al. 2012, Myers, Bolyard et al. 2014).

1.3.3. Treatment for SDS

There is no curative treatment for SDS patients; however, several things can be done to reduce side symptoms and to treat the most severe conditions.

1. Primary and secondary complications of bone marrow failure. Bacterial or fungal skin

infections due to mild to moderate neutropenia may be treated with oral antibiotics; however,

severe infections or infections concomitant with severe neutropenia are treated with

intravenous broad-spectrum antibiotics and granulocyte colony stimulating factor (Dror,

Donadieu et al. 2011). Coagulation defects and bleeding episodes due to thrombocytopenia

or vitamin K deficiency are treated with tranexamic acid, aminocaproic acid, xylometazoline

nose spray or vitamin K supplement, while severe bleeding due to thrombocytopenia are

treated with platelet transfusion. Anemia is treated with chronic transfusion and iron-

chelation.

2. Hematopoietic stem cell transplantation. Severe cytopenia, myelodysplastic syndrome and

acute myeloid leukemia all demand hematopoietic stem cell transplantation (HSCT).

Hematopoietic stem cell transplantation can be done from a matched related or unrelated

donor (Okcu, Roberts et al. 1998), and umbilical cord blood (Vibhakar, Radhi et al. 2005).

The event free 5 year survival rate of post-HSCT is 60% (Cesaro, Oneto et al. 2005,

Donadieu, Michel et al. 2005). It was shown that survival rate of HSCT for non-malignant

severe cytopenia was higher than those for myelodysplastic syndrome or acute myeloid

leukemia (Donadieu, Fenneteau et al. 2012). Furthermore, the survival rate of post HSCT

using identically matched related donor was higher than that using matched unrelated donor

(Dror, Donadieu et al. 2011). Multiple complications including graft-versus-host disease,

graft failure, prolonged severe aplasia, veno-occlusive disease, infections as well as

neurological and renal complications can arise following HSCT (Dror, Donadieu et al. 2011).

The incidence of regimen-related toxicity in SDS patients who undergo HSCT was highest in

heart and lungs (Tsai, Sahdev et al. 1990), To reduce the toxicity, the intensity of

conditioning regimen can be reduced by avoiding the use of busulfan, cyclophosphamide,

and total body radiation. Such modifications can help achieve myeloid engraftment and full

donor chimerism without grade III-IV graft-versus-host disease (Sauer, Zeidler et al. 2007,

Bhatla, Davies et al. 2008).

3. Pancreatic enzyme supplementation. Pancreatic insufficiency in SDS frequently requires

treatment with pancreatic enzyme replacement with all meals, consisting of lipase (2,000-

10000 units/kg body weight/day), protease, amylase, or additional H2-receptor antagonist if

enzyme supplement fails to improve fat absorption (Dror, Donadieu et al. 2011).

1.3.4. Characterization of Shwachman-Bodian Diamond Syndrome gene (SBDS).

In 2003, Boocock et al. group identified the first gene associated with SDS, which they named

Shwachman-Bodian-Diamond syndrome (SBDS) (Boocock, Morrison et al. 2003). SBDS is

located at chromosome 7q11 with five exons spanning 7.9kb. The five exons encode a 1.6kb

mRNA and final protein product is 28.8kDa molecular weight with a pI of 8.9 (Boocock,

Morrison et al. 2003). SBDS mRNA is ubiquitously expressed in diverse fetal and adult tissues.

SBDS has an additional isoform that skips exon 2, which has lower mRNA expression compare

to the full length isoform.

SBDS mutations. The most common SDS-associated mutations are located in exon 2

(c.183_184TA>CT and c.258+2T>C) (Boocock, Morrison et al. 2003). The c.183_184TA>CT

mutation introduces a premature stop codon, resulting in a truncated protein, p.Lys62*. The

second mutation (c.258+2T>C) disrupts the donor splice site of the second intron and an

upstream cryptic donor splice site at c.251_252 is used. This mutation is resulting in either an

mRNA with 8bp deletion (r.251_258del) causing a frameshift and production of a truncated

protein (p.Cys84Tyrfs*4), or an mRNA that lacks exon two (r.129_258del). With both mutations

there is some level of protein expression as experimentally validated by Orelio et al (Orelio and

Kuijpers 2009, Orelio, Verkuijlen et al. 2009). Interestingly, no genotype-phenotype relationship

has been found between SBDS mutations and SDS manifestations (Makitie, Ellis et al. 2004,

Kuijpers, Alders et al. 2005, Hashmi, Allen et al. 2011, Donadieu, Fenneteau et al. 2012).

SBDS protein structure. SBDS gene is highly conserved across 159 species, from archaea to

eukaryotes (Boocock, Marit et al. 2006). Human SBDS protein consists of 3 different domains

(de Oliveira, Sforca et al. 2010, Finch, Hilcenko et al. 2011). The first one is a highly conserved

N-terminal domain. It spans residues S2-S96 and contains a mixture of α-helices and β-sheets in

the sequence ββααββαα. The N-terminal domain was shown to be a SDS-associated mutational

hotspot. The second domain is the central domain spanning residues D97-A170 and containing

three α-helices. The C-terminal domain spanning residues H171-E250; it contains a ferredoxin

fold in the sequence βαββαβ, which is a putative nucleic acid binding motif. The SBDS protein is

localized in both the cytoplasm and the nucleus, with greater concentration in the nucleolus. The

function of SBDS in the nucleus is still unclear (Austin, Leary et al. 2005).

1.3.5. Ribosome Biogenesis and SBDS protein function

In 2001 Koonin et al. analyzed the archaeal genome and found that the archaeal orthologue of

SBDS is clustered with the RNA processing genes (Koonin, Wolf et al. 2001). In 2002 and 2003,

Wu et al. and Peng et al. profiled the transcriptome of Saccharomyces cerevisiae and confirmed

that the yeast orthologue of SBDS, YRL022C or SDO1 is similarly clustered with genes that are

responsible for RNA processing functions(Wu, Hughes et al. 2002, Peng, Robinson et al. 2003).

In 2006, Krogan et al. utilized the high-throughput affinity-capture mass spectrometry to identify

protein interactions and found that Sdo1p is associated with elongation factor like 1 (Efl1) as

well as other proteins involved in ribosome biogenesis (Krogan, Cagney et al. 2006). In 2007,

Menne et al. reported functional role of the SBDS homologue in Saccharomyces cerevisiae in

ribosome biogenesis (Menne, Goyenechea et al. 2007). It was shown that sdo1Δ yeast mutant

strains were either slow in growth or nonviable (Winzeler, Shoemaker et al. 1999, Menne,

Goyenechea et al. 2007). Translation initiation factor 6 (Tif6, the yeast orthologue of human

eIF6) and Efl1 were shown to be involved in ribosome biogenesis being integral in determining

the function of Sdo1p protein. Tif6 is a shuttling factor that is required for the cytoplasmic export

of premature big ribosome subunit 60S. Moreover, TIF6 mutations rescued the defects in sdo1Δ

mutant. Efl1 is a GTPase required for the release of Tif6 from the premature 60S subunit and

efl1Δ is a pheno-copy of sdo1Δ as both strains showed reduced 60S subunits. sdo1Δ has impaired

Tif6 nuclear recycling, pre-60S cytoplasmic export, and pre-rRNA processing. Overall it became

clear that Sdo1p is an essential factor in Tif6 release during ribosome biogenesis. In September

of the same year, Ganapathi et al. confirmed the role of SBDS in 60S maturation using HeLa

cells. (Ganapathi, Austin et al. 2007). SBDS was localized to the nucleolus during active rRNA

transcription and was co-sedimented with 60S and co-precipitated with 28S rRNA (component

of 60S), nucleophosmin (ribosomal chaperon), and other ribosomal proteins (Ganapathi, Austin

et al. 2007). In 2011, Finch et al. showed that the release of the anti-association factor eIF6 from

the pre-60S ribosome subunit, catalyzed by SBDS and the GTPase EFL1, required GTP

hydrolysis(Finch, Hilcenko et al. 2011). In 2013, Gijsbers et al. analyzed enzyme kinetics of

SBDS and classified it as a GTP stabilizing factor in the class of GDP/GTP exchange factors

(Gijsbers, Garcia-Marquez et al. 2013). This class stabilized the binding of GTP to EFL1 on the

pre-60S large ribosome subunit. In 2014, Asano et al. showed through isothermal titration

calorimetry that domains II-III of SBDS interacted with the intrinsically disordered insertion

domain of EFL1(Asano, Atsuumi et al. 2014). In 2015, Weis et al. analyzed cryo-EM structures

of human SBDS and EFL1 bound to Dictyostelium discoideum 60S with or without eIF6 to

uncover conformation changes in SBDS during the release of EIF6. (Weis, Giudice et al. 2015).

SBDS protein domains were shown to be bound to active sites on pre-60S. This suggests that

SBDS regulates maturation of pre-60S by either having a role in proofreading or by physically

protecting the active sites (Weis et al., 2015). Anchored that are located on P site of the pre-60S,

domain I protected and potentially proofread the peptidyl transferase center and polypeptide exit

tunnel to prevent premature translation initiation. In addition, Domain I mimics tRNA to check

the function of P-site through pseudo-translocation as the Sdo1p-bound P-site can operate tRNA

binding to the A-site (Ng, Waterman et al. 2009, Sulima, Gulay et al. 2014). Binding to the

sarcin-ricin loop at the P-stalk base on the pre-60S, domain III protected and potentially

proofread active sites of translational GTPases to prevent premature pre-60S maturation (Weis,

Giudice et al. 2015).

Ribosomopathy SDS can be considered as a ribosomopathy, a disorder caused by a dysfunction

of ribosomal components or factors involved in ribosome biogenesis and maturation (Armistead

and Triggs-Raine 2014). Although their clinical manifestations are heterogeneous,

ribosomopathies share common characteristics of poor growth, bone marrow failure,

developmental impairment, predisposition to malignancy, and extra-ribosomal consequences

such as cell cycle arrest. Phenotypic heterogeneity of ribosomopathies was proposed to be a

consequence of differential expression of ribosome-associated genes (Kondrashov, Pusic et al.

2011), resulting in a cell type-specific ribosomal composition and a translational control of

specific mRNAs. For instance, translation of Hox genes involved in embryonic patterning

requires RPL38 to recruit ribosomes to RNA regulons in the 5’ UTR, similar to the internal

ribosome entry site (IRES) (Xue, Tian et al. 2015). Moreover, IRES-dependent translation is

affected by DKC1 (gene associated with X-linked dyskeratosis congenita) and by ribosomal

proteins that are associated with Diamond-Blackfan Anemia (Yoon, Peng et al. 2006, Horos,

Ijspeert et al. 2012). Similarly, a translational control through RNA elements may explain the

appearance of various defective cell types in SDS.

1.3.6. Extra-ribosomal Consequences of SBDS Deficiency

Affinity capture and mass spectrometry have identified SBDS interacting proteins with functions

other than ribosome biogenesis, suggesting that SBDS may have multiple functions (Ball, Zhang

et al. 2009). SBDS deficiency has been shown to affect multiple biological processes, including

rRNA processing (Ganapathi, Austin et al. 2007, Luz, Georg et al. 2009, Rujkijyanont, Adams et

al. 2009, Moore, Farrar et al. 2010), translation of upstream open reading frame (uORF)-

dependent protein isoforms (In, Zaini et al. 2016), Fas-mediated apoptosis (Dror and Freedman

2001, Dror, Ginzberg et al. 2001, Rujkijyanont, Watanabe et al. 2008, Watanabe, Ambekar et al.

2009, Ambekar, Das et al. 2010), production of reactive oxygen species (Ambekar, Das et al.

2010, Sen, Wang et al. 2011), endoplasmic reticulum stress (Ball, Zhang et al. 2009),

mitochondrial respiration (Henson, Moore et al. 2013, Ravera, Dufour et al. 2016), p53-mediated

apoptosis and senescence (Elghetany and Alter 2002, Kuijpers, Alders et al. 2005, Austin, Gupta

et al. 2008, Tourlakis, Zhong et al. 2012, Tourlakis, Zhang et al. 2015), actin remodeling and

signaling (Wessels, Srikantha et al. 2006, Orelio and Kuijpers 2009, Leung, Cuddy et al. 2011),

microtubule stability (Austin, Gupta et al. 2008, Orelio and Kuijpers 2009, Orelio, Verkuijlen et

al. 2009) and hyperactivation of mTOR and STAT3 (Bezzerri, Vella et al. 2016, Ravera, Dufour

et al. 2016). Whether some of these processes directly involve SBDS or are consequences of

reduced ribosome biogenesis remains to be elucidated.

Fas-mediated apoptosis and reactive oxygen species. Increased Fas-mediated apoptosis in

SDS was first reported in human SDS bone marrow mononuclear cells (Dror and Freedman

2001). In human SDS bone marrow mononuclear cells and SBDS shRNA knockdown HeLa

cells, FAS was constitutively overexpressed, and FAS stimulation resulted in apoptosis as well

as reduced cell growth and reduced CFU-GM colonies formation (Dror and Freedman 2001,

Rujkijyanont, Watanabe et al. 2008). Increased Fas-mediated apoptosis has been shown to result

from increased Fas receptors in the plasma membrane and increased production of reactive

oxygen species (Watanabe, Ambekar et al. 2009, Ambekar, Das et al. 2010). In SBDS shRNA

knockdown HeLa cells and human myeloid cells, Fas stimulation increased production of

reactive oxygen species and induced apoptosis and necrosis, while antioxidant treatment rescued

Fas-mediated cell death (Ambekar, Das et al. 2010). Similarly, in SBDS knockdown K562 cells,

oxidative stress impaired erythroid differentiation, while antioxidant treatment improved cell

growth (Sen, Wang et al. 2011). Endoplasmic reticulum is an oxidizing environment that

promotes protein folding and disulfide bond formation (Malhotra and Kaufman

2007).Endoplasmic reticulum stress has been reported in several models of SDS (in human

embryonic kidney cell line with siRNA knockdown of SBDS (Ball, Zhang et al. 2009) and in

lymphoblastoid cells derived from SDS patients) can contribute to oxidative stress observed in

SDS. Oxidative stress can also be caused by impaired mitochondrial respiration (Henson,

Moore et al. 2013, Ravera, Dufour et al. 2016).

Mitochondrial Respiration. In sdo1Δ yeast mutant, while a global protein production was

reduced, expression of Por1 (anion channel of the mitochondrial outer membrane, yeast

orthologue of human VDAC1) was increased. Increased Por1 protein expression resulted in

reduced growth of sdo1Δ yeast cultured on glycerol and an absence of red pigmentation in sdo1Δ

yeast cultured on glucose. As the red pigment was an intermediate in the adenine synthesis

pathway that required respiration for production, this indicated that an impairment of

mitochondrial respiration. Moreover, an increased protein expression of VDAC1 was confirmed

in SBDS shRNA knockdown of a human erythroleukemia cell line (Henson, Moore et al. 2013).

SBDS knockdown resulted in reduced mitochondrial membrane potential, reduced oxygen

consumption, and increased reactive oxygen species. Impaired respiration in mitochondria has

been shown to result from reduced activity of complex IV in the electron transport chain

(Ravera, Dufour et al. 2016). In human SDS lymphocytes and lymphoblasts, reduced complex

IV activity resulted in reduced ATP production and increased AMP accumulation. This energy

stress led to an activation of response pathways such as AMP-activated protein kinase and

PI3K/AKT/mTOR, which activated glycolysis as an alternative method of energy production.

Hyperactivation of mTOR and STAT-3 pathways were also found in B cells, granulocytes,

monocytes, T cells, NK cells (Bezzerri, Vella et al. 2016).

Actin remodeling. In 1978, Thong first reported impaired neutrophil chemotaxis in SDS patient

(Aggett, Harries et al. 1979, Ruutu, Savilahti et al. 1984, Szuts, Katona et al. 1984, Repo,

Savilahti et al. 1987). Later on, an impaired chemotactic orientation of human SDS

polymorphonuclear leukocytes was shown using spatial gradients of chemoattractant fMLP.

(Dror et al, 2001, Stepanovic, Wessels et al. 2004). In 2006, Wessels et al. showed that SBDS

was localized to an anterior pseudopod and newly formed lateral pseudopods of Dictyostelium

discoideum during chemotaxis(Wessels, Srikantha et al. 2006). In 2009, Orelio and Kuijpers

showed co-localization of SBDS with F-actin and Rac2, signaling components of actin

remodeling in normal human neutrophils, and a delayed directional F-actin polarization in

human SDS neutrophils during chemotaxis(Orelio and Kuijpers 2009). In 2011, Leung et al.

showed that reduced RANKL-TRAF6-NFkB signaling and reduced Rac2 expression lead to

impaired monocyte migration and fusion required for osteoclast differentiation in myeloid cell-

targeted Sbds conditional knockout mice (Leung, Cuddy et al. 2011). As the mechanism of

SBDS/Rac2 signaling remains to be elucidated, it has been proposed that SBDS may mediate

actin remodeling. In addition, it was hypothesized that SBDS may be localized to regions of

increased translation such as actin reorganization.

Microtubule stabilization. In addition to its role in actin remodeling, SBDS affects the stability

of microtubules. Full-length and truncated SBDS were found to be co-localized with the mitotic

spindle and centrosomes in human bone marrow stromal cells, HeLa cells, and cord blood-

derived neutrophils (Austin, Gupta et al. 2008, Orelio and Kuijpers 2009, Orelio, Verkuijlen et

al. 2009). Mitotic abnormalities (multiple centrosomes, multipolar mitotic spindles, and

aneuploidy accumulation), cell cycle arrest at G1, and apoptosis were reported (Austin, Gupta et

al. 2008) in bone marrow stromal cells and lymphoblasts derived from SDS patients as well as in

skin fibroblasts with siRNA knockdown of SBDS,. In particular, SBDS increased microtubule

stability upon dilution-induced microtubule depolymerization in vitro. In a follow-up study,

shortened mitotic spindles and reduced spindle acetylation were observed during SBDS

deficiency, indicating reduced microtubule stability. Wild-type SBDS increased polymerization

rate and promoted microtubule bundling of polymerized microtubule in vitro, while mutant

SBDS reduced microtubule bundling. Treating SBDS knockdown human CD34+ cells with taxol,

a microtubule stabilizer, rescued proliferation, differentiation, and hematopoietic progenitor

colony formation. Both mitotic abnormalities and microtubule instability have been proposed to

be the mechanisms driving genomic instability, cytogenetic abnormalities, and malignant

transformation associated with SBDS deficiency. It has also been proposed that SBDS may

mediate the interconnecting organization of both actin and microtubule.

P53-mediated apoptosis in hematopoietic cells. In 2002, Elghetany and Tarek demonstrated

that p53 protein was overexpressed in 78% of bone marrow biopsies from SDS patients, in

immature cells but was absent in mature cells (band and segmented neutrophils,

megakaryocytes, and nucleated red cells) (Elghetany and Alter 2002). In 2005, Kuijpers et al.

confirmed an absence of apoptotic neutrophils in SDS patients (Kuijpers, Alders et al. 2005). In

2008, Austin et al. subjected bone marrow stromal cells derived from SDS patients to ionizing

radiation and demonstrated that the cells went into a cell cycle arrest at G1, as indicated by

increased p53 protein expression and p21 (p53 downstream mediator) mRNA expression in the

apoptotic G1 population (Austin, Gupta et al. 2008). Austin et al. proposed that the cell cycle

arrest may be a protective mechanism that eliminated abnormal cells with DNA damage (Austin,

Gupta et al. 2008). In 2015, Zambetti et al. created a SDS neutropenia murine model by

transplanting the fetal liver of hematopoietic progenitor-targeted Sbds conditional knockout mice

(Sbdsf/f) into recipient mice. Using this model, he demonstrated that terminal differentiation was

impaired at the myelocyte-metamyelocyte stage due to p53-mediated apoptosis as indicated by

increased mRNA expression of p53 and p53 targets, increased p53 protein expression, and

increased apoptotic cells in myelocytes and metamyelocytes (Zambetti, Bindels et al. 2015).

P53-mediated senescence in pancreatic cells. In 2012, Tourlakis et al. demonstrated that

exocrine pancreatic defects are caused by senescent cell cycle arrest, as indicated by increased

senescence-associated β-galactosidase activity, increased expression of senescence-associated

mRNA, and increased p53 (senescence activator) protein expression in acinar cells in a pancreas-

targeted Sbds conditional knockout mice (SbdsP-/R126T) (Tourlakis, Zhong et al. 2012). Knocking

out p53 expression in pancreas-targeted conditional knockout mice (SbdsP-/R126T;Trp53-/- double

knockout mice) only temporarily rescued SDS pancreatic defects, as apoptotic cells and

dedifferentiated acinar cells increased as double knockout mice matured. However, knocking out

p53 expression in Sbds knockout mice (SbdsR126T/R126T;Trp53-/- double knockout mice) rescued

hematopoietic progenitors in fetal liver and reduced neural apoptosis in fetal brain.

Telomere shortening. Most SDS patients have normal telomere length in lymphocytes.

However, in one study the mean age-adjusted telomere length in hematopoietic cells was found to be reduced by 1.4kb/year compared to the normal maximum of 60bp/year (Thornley, Dror et al. 2002).

1.3.7. SDS Patients without SBDS Mutations

Cases of clinically diagnosed SDS lacking biallelic SBDS mutations have been reported, with

some cases of confirmed normal SBDS protein expression (Nakashima, Mabuchi et al. 2004,

Woloszynek, Rothbaum et al. 2004, Hashmi, Allen et al. 2011, Myers, Bolyard et al. 2014). Data

from the Canadian Inherited Marrow Failure registry (CIMFR) suggested that these patients

presented with more severe hematological dysfunction, but milder pancreatic dysfunction

compared to SDS patients with biallelic SBDS mutations (Hashmi, Allen et al. 2011). These SDS 27

patients had a higher incidence of severe aplastic anemia, severe bone marrow failure, treatment

requirement for cytopenia, reduced hemoglobin levels, and higher mean fetal hemoglobin levels

after the age of 1 year compared to patients with SDS and SBDS mutations. At the time of this

publication, none of the patients with SBDS-negative SDS required pancreatic enzyme

treatment. In contrast, the North American registry suggested that these SDS patients presented

with milder hematological dysfunction but more severe pancreatic dysfunction (Myers, Bolyard

et al. 2014). These patients had a lower incidence of hypocellular bone marrow and clonal

marrow cytogenetic abnormalities and no occurrence of bone marrow dysplasia or malignant

transformation, although there was a higher incidence of enzyme treatment and failure to thrive.

The discrepancy between the two registries may be due to difference in the underlying genetic

groups between both registries. Other findings of the SBDS-negative SDS patients include

increased incidence of skeletal gait irregularities and no occurrence of elevated liver

transaminases (Hashmi, Allen et al. 2011, Myers, Bolyard et al. 2014).

1.3.8. Characteristics of DNAJC21, a member of DnaJ Heat Shock Protein Family (Hsp40)

DNAJC21 is a human gene of unknown function. It belongs to the family of DnaJ heat shock

protein 40 chaperone proteins characterized by a C-terminal DNAJ domain and two N-terminal

tetra-tricopeptide repeat domains. It is ubiquitously expressed in human tissues and was found in

the cytoplasm and in the nucleus of HeLa and 293T cells (Chen, Yin et al. 2004, Craig, Huang et

al. 2006). It was shown that a reduction in protein levels of DNAJC21 causes various effects on

rRNA processing in HeLa cells. In addition, an yeast orthologue of DNAJC21 was discovered

jjj1, (Meyer, Hung et al. 2007)) and better characterized than the human protein (Helser, Baan et

al. 1981, Lebreton, Saveanu et al. 2006, Demoinet, Jacquier et al. 2007).

1.3.8.1. Molecular chaperones

Molecular chaperones facilitate protein folding and oligomeric complex assembly and

disassembly, as well as most other aspects of protein homeostasis such as quality control of

misfolded or stress-denatured proteins. In eukaryotes cells, cytosolic chaperones are organized

into two distinct but overlapping groups: 1. Stress-inducible heat shock proteins that protect the

cellular proteome from misfolding and stress. 2. Chaperones that are linked to protein synthesis

(CLIPS) and cooperate with the translational apparatus. The observed transcriptional co-

regulation of CLIPS with translational apparatus suggested that this chaperone network functions

in protein biogenesis, most likely in de novo folding of newly made polypeptides. Indeed, many

CLIPS chaperones, such as the Hsp70s Ssb1 and Ssb2 (herein referred to as SSB), the prefoldin

GIMc (genes involved in microtubules complex), and the chaperonin TRiC/CCT associate with

newly translated polypeptides. However, not all ribosome-associated CLIPS bind nascent chains,

as neither Zuo1 nor its Hsp70 partner Ssz1 interact directly with nascent chains. The Zuo1–Ssz1

complex, termed ribosome-associated complex, stimulates the ATPase activity of highly

homologous CLIPS Hsp70s SSB via the N-terminal J domain of Zuo1. Another RAC-like

protein, Jjj1, was characterized as a strictly cytosolic protein that binds to Rei1 and helps recycle

the 60S ribosomal export factor Arx1.

1.3.8.2. Characteristics of Jjj1

Jjj1 associates with the large ribosomal subunit in a late step of subunit maturation, removing

biogenesis factors. Numbers of cytosolic proteins have been shown to be important for the

recycling or release to the nucleus of particular pre-mature 60S ribosome sub-unit. For example,

in the case of the Arx1/Alb1 heterodimer, Jjj1 is required. The majority of Arx1/Alb1 complexes are associated with pre-mature 60S ribosome sub-unit particles in the nucleus, consistent with an efficient cytosolic dissociation and transport back to cytosol through the nuclear membrane.

However, in the absence of Jjj1, Arx1 and Alb1 accumulate in the cytosol and fail to translocate to the nucleus. Moreover, in order to regulate recycle or release of the Arx1/Alb1 complex Jjj needs to interact with Rei1. Both Jjj1 and Rei1 are very important factors in 60S subunit biogenesis, specifically as in affecting the dynamics of the cytosolic interactions of Arx1.

However, the exact mechanism of cooperation between the two proteins is not well understood. Jjj1 role in ribosome biogenesis is an example of involvement of Hsp70/J-protein chaperone machinery in remodeling protein complexes (Demoinet, Jacquier et al. 2007, Meyer,

Hung et al. 2007, Sahi and Craig 2007, Albanese, Reissmann et al. 2010, Karbstein 2010, Meyer,

Hoover et al. 2010, Weeks, Shield et al. 2010, Gillies, Taylor et al. 2012, Greber, Boehringer et al. 2012, Schmidt, Dethloff et al. 2014, Kaschner, Sharma et al. 2015, Wu, Zhang et al. 2016,

Whaley, Caudle et al. 2018).

Chapter 2 Research Aims and Hypothesis 2.1 Rationale.

SDS is a rare multi-system genetic disorder characterized by combined exocrine, pancreatic and hematological dysfunction (Bodian, Sheldon et al. 1964, Shwachman, Diamond et al. 1964).

SDS is often diagnosed in early childhood (Hashmi, Allen et al. 2011) and can manifest in neonates (Todorovic-Guid, Krajnc et al. 2006). About 10-20% of SDS patients do not have a mutation in SBDS gene(Boocock, Morrison et al. 2003, Hashmi, Allen et al. 2011). About 18% of patients will develop clonal malignant myeloid transformation at the median age of 20 years

(Hashmi, Allen et al. 2011). Progression to myelodysplastic syndrome and acute myeloid leukemia is the main concern for mortality in SDS. Today the only treatment option for SDS patients with severe hematopoietic dysfunction is hematopoietic stem cell transplantation, for which the survival rate is around 60-65% (Cesaro, Oneto et al. 2005, Donadieu, Michel et al.

2005). Hence, there is a strong need to develop new treatments for this disease. Bone marrow failure disease including SDS, in 95% are inherited disease. Therefore, uncovering mutations in novel genes present in SDS affected families that lack known SBDS mutations and studying the mechanisms underlying the hematopoietic defects in the disorder will help identify new drug targets and develop effective treatments.

2.2 Hypothesis

We hypothesized that SBDS-negative SDS patients have mutations in genes that have not yet been identified. Some of these mutations might affect hematopoietic genes; thereby affecting bone marrow function. All the exons in a genome are known as the exome, and the method of sequencing these exons is known as whole exome sequencing (WES). In addition, as most 31

known mutations that cause disease occur in exons, WES is thought to be an efficient method to identify possible new disease-causing mutations. Genome-wide analysis studies using high resolution WES would help unravel novel genetic alterations in SDS patients that are not present in healthy individuals.

2.3 Research AIMS

AIM 1: To identify novel SDS-related genetic alterations and novel SDS genes through comprehensive genome-wide genetic screen

To identify novel genetic alterations involved in SDS, I analyzed 9 banked samples from the

Canadian Inherited Marrow Failure Registry (CIMFR). These samples were from patients with classified IBMFSs in whom a genetic cause could not be identified after metaphase cytogenetics, chromosomal fragility testing and other specific genetic tests.

Candidate genes will be selected and further validated using quantitative real time polymerase chain reaction (qPCR) for gene expression.

AIM 2: Studying the mechanism of how novel IBMFS-related gene DNAJC21 can cause Shwahman Diamond Syndrome

(i) To characterize hematopoietic colony forming potential of marrow cells from patients with

SDS and DNAJC21 mutations

(ii) To determine whether DNAJC21 is critical for normal hematopoiesis

(iii) To determine whether DNAJC21 maintains hematopoiesis by protecting the cells from cell death (apoptosis and oxidative stress).

Chapter 3 Methods 3.1 Patients and Controls – The Canadian Inherited Marrow Failure Registry (CIMFR)

The patients studied in this work were enrolled in the Canadian Inherited Marrow Failure

Registry (CIMFR) after informed written consent was obtained from patients and controls, or their legal guardians, prior to sample collection. The registry and the studies performed herein were approved by the Hospital for Sick Children Research Ethics Board. Peripheral blood or bone marrow samples were obtained from 26 unrelated patients with the unknown genotype.

Among those patients 8 had clinical diagnosis of SDS but they did not have mutations in the then known SDS gene, SBDS. The diagnosis was based on published clinical criteria (Dror and

Freedman 2001, Dror, Durie et al. 2002, Dror, Donadieu et al. 2011), which include evidence for both hematological and exocrine pancreatic dysfunction.

This study included patients who had been enrolled in the CIMFR from January 2001 until

December 2017. Data available for these patients until January 2017 were analyzed for the project. Patient information at study entry and at yearly checkup was collected and analyzed.

These data included demographics, family history, clinical symptoms, genetic tests, imaging studies, diagnoses, physical malformations, laboratory tests, treatment, and outcomes.

3.2 DNA extraction

DNA was extracted using the Promega Wizard DNA extraction kit (Promega Corporation,

Madison, WI) according to the manufacturer`s protocol. Genomic DNA was then re-suspended in 30-100 ul of TE buffer and aliquots stored at 80C until used.

3.3 Exome sequencing and analysis

Genomic DNA samples of high quality were sent to The Centre for Applied Genomics, The

Hospital for Sick Children, Toronto, for exome library preparation using Agilent SureSelect

Human Exome Library Preparation V5 kit and for sequencing using the Agilent Bravo

Automation System and paired end sequencing on a HiSeq 2500 platform. Sequence reads were trimmed, and subsequently aligned to the reference human genome (hg19/GRCh37) using

Burrows-Wheelchair aligner. PCR duplicate reads were removed (MarkDuplicates, Picard tools v.1.112) and local realignment was performed (GATK 3.2.2). GATK haplotype caller3.2.2 was used to annotate single nucleotide variants (SNVs) and insertions/deletions (indels). Only variants with minor allele frequency ≤ 0.01 reported in external and internal databases were retained. Pathogenic prediction was performed using six different tools that included Polyphen-2,

SIFT, Provean, mutation taster, SNAP2 and conservation values using GERP.

3.4 Web Resources

The URLs for data presented herein are as follows:

ANNOVAR, http://annovar.openbioinformatics.org/en/latest/

CFSSPS, http://www.biogem.org/tool/chou-fasman/

dbSNP, http://www.ncbi.nlm.nih.gov/projects/SNP/

ExAC Browser, http://exac.broadinstitute.org/

GATK v.3.2.2, https://www.broadinstitute.org/gatk/

GenBank, http://www.ncbi.nlm.nih.gov/genbank/

MutationTaster, http://www.mutationtaster.org/

NCBI HomoloGene, http://www.ncbi.nlm.nih.gov/homologene

NextGene 2.4.2: http://www.softgenetics.com/NextGENe.php

NHLBI Exome Sequencing Project (ESP) Exome Variant Server, http://evs.gs.washington.edu/EVS/

OMIM, http://www.omim.org/ Picard tools v.1.112/

MarkDuplicates, http://broadinstitute.github.io/picard/

PolyPhen-2, http://genetics.bwh.harvard.edu/pph2/

RCSB Protein Data Bank, http://www.rcsb.org/pdb/home/home.do

SIFT, http://sift.bii.a-star.edu.sg/

SNAP2, https://www.rostlab.org/services/snap/

UCSC Genome Browser, http://genome.ucsc.edu

3.5 PCR Conditions

Amplification of DNA template was performed using 50 ng template genomic DNA and 100uM of forward and reverse primers specific for desirable DNA part.

The PCR mix also contained 1 PCR buffer, 0.25–1 mM MgCl, and Taq DNA Polymerase

(Invitrogen, Ont.,Canada). Reactions were carried out for 25 cycles using optimized denaturing, annealing, and extension conditions on the Eppendorf Mastercycler Gradient Thermo-cycler.

Primers sequences and conditions for each exon are available. PCR products were purified using a commercial PCR purification kit from Qiagen (Toronto, ON,Canada) and eluted in 30 ul TE buffer.

PCR Primers:

DNAJC21_cDNA_exon 4-7 –F: 5`-TGG AGA TGA AAA GGG ATT T-3`

DNAJC21_cDNA_exon 4-7 –R: 5`-CGT TCT TCA TGG CCT TT -3`

DNAJC21exon1-F: 5`-CGG AGA GGA CTG CCA GC -3`

DNAJC21exon1-R: 5`-CCT CTC TGC TGG CTC CG -3`

DNAJC21exon2-F: 5`-GTG GAG TCT TTC TGC TTG GC -3`

DNAJC21exon2-R: 5`-TTG –ATA GCT CCC AAV ATC -3`

DNAJC21exon3-F: 5`-AAC AGT TCT GTG AAA GAT TAT GAA CTC -3`

DNAJC21exon3-R: 5`-TGA AGG CAA TGC ATC TTC AG -3`

DNAJC21exon4-F: 5`-TTG-CAG GGA CTG AAC TTT G -3`

DNAJC21exon4-R: 5`- GAG GTC ATG GGA TGA GTT CG -3`

DNAJC21exon6-F: 5`-AAG TGA AAA CCA TGC AGA GG -3`

DNAJC21exon6-R: 5`-GCA AGG AGA TTT CCA GCT TC -3`

DNAJC21exon7-F: 5`-TGC TTT ATT TGG AGC CTT TCC` -3`

DNAJC21exon7-R: 5`-ATG ACT GTG CCQA CTG CAC TC -3`

DNAJC21exon8-F: 5`-TCT TGG TTG CAG TTA TCC AGC -3`

DNAJC21exon8-R: 5`-GGG CAA CAG AGT GAT TCT CC -3`

DNAJC21exon9-F: 5`-TCC ATT GAA CTA CAG CCT TGT -3`

DNAJC21exon9-R: 5`-AAG AGC ACT GCA TTA TCG -3`

DNAJC21exon10-F: 5`-TGG TAA ATT TGA ATG TGG GTT G -3`

DNAJC21exon10-R: 5`-TTG-CAG GGA CTG AAC TTT G -3`

DNAJC21exon11-F: 5`-CCG AAG GTA TAT AAC TAT TTG GCA AC -3`

DNAJC21exon11-R: 5`-CCT GGT CAG TCA TGG GAA AG -3`

DNAJC21exon12-F: 5`-GCA CTC AAA TAA TGA TGG TTA AAA G -3`

DNAJC21exon12-R: 5`-CGG AAT GGC TCA CCA AAT AC -3`

DNAJC21exon13-F: 5`-AAC CTC CAA GAA CAT CTG CC -3`

DNAJC21exon13-R: 5`-CAT TCA AGT CAA TTC AGA TGG C -3`

DNHD1-LongISO-Pair1 –F: 5`-CTG GCC AGT ACA CAG AAG CA -3`

DNHD1-LongISO-Pair1 –R: 5`-CAG GGT TCA CGG TCA AT -3`

DNHD1-LongISO-Pair2 –F: 5`-GCA GCA ACA GAC AAT CCT GA -3`

DNHD1-LongISO-Pair2 –R: 5`-ATC AGT TAC CCG GAC CAC AG -3`

DNHD1-LongISO-Pair3 –F: 5`-AGC AGA TGA GCA AGG CAT TT -3`

DNHD1-LongISO-Pair3 –R: 5`-TAG TGC AGA ACA GCC CAG AG -3`

DNHD1-exon25_p559_mut1-F: 5`-GGT GCC CGA ATG AGA ATC TG -3`

DNHD1-exon25_p559_mut1-R: 5`-GAC CAG GTC TGA GGG CTT TT -3`

DNHD1-exon36_p559_mut2-F: 5`-TGG CAG GCT TAC CTG TCA CT -3`

DNHD1-exon36_p559_mut2-R: 5`-ATC AGC ATG GGC TGA GTA GC -3`

DNHD1-exon23_p559_mut3-F: 5`-GGT CCT TGG AAC AGT TGA GC -3`

DNHD1-exon23_p559_mut3-R: 5`-TCC CGA TTG ATT CCT CTG TC-3`

DNHD1-cDNA_L_ISO-F: 5`-CTG ATG GCA GCA GCA TGG -3`

DNHD1-cDNA_L_ISO-R: 5`-CAC TTC CAC TCG CTG ATG TT -3`

PuroR-C-F: 5`- CGA GGT GCC CGA AGG AC -3`

PuroR-C-R: 5`- GCA TTA AAG CAG CGT ATC -3`

DNHD1_ex24-F: 5`-GTG CTG GTA GAG CCA CAT CA -3`

DNHD1_ex24-R: 5`-GGT TGG GTC TGA GGT ACC AA -3`

DNHD1_ex37-F: 5`-GGA CTT TGG GGA TCA TGG TA -3`

DNHD1_ex37-R: 5`-ATT CCA GGT CTG CAA CCA CT -3`

DNHD1_cDNA_exon1-F: 5`-ACC TCC ATC TGG ACC TGC TA -3`

DNHD1_cDNA_exon1-R: 5`-TGC CAA GGG AAG GAG ACT AA -3`

DNHD1_ exon1+2-F: 5`-GCC CTA GAA GAG GCT GTG TG -3`

DNHD1_ exon1+2-R: 5`-AGA GCA GTC ATC ACC TGG CTA -3`

DNHD1_ex25-F: 5`-TTG CAT GAG GCA CAG AGA AC -3`

DNHD1_ex25-R: 5`-CCT CCT CAC TCT CCT GAT GG -3`

3.6 RNA extraction

Total RNA from cells was extracted using RNease Mini Kit (50) and QIAshredder (50)

(QIAGEN Inc., Mississauga, Canada, 74104, 79654) according to the manufacturer’s instructions. In brief, cells were harvested (max 1x107 cells per reaction),buffer RLT and added and centrifuged at maximum speed. Ethanol was added to lysate and mixed. RNAse mini spin column was used to bind RNA. Finally, RNase-free water was used to elute RNA. RNA was store at -80C.

3.7 cDNA preparation:

Omniscript Reverse Transcription Kit was used for preparing cDNA from RNA according to the manufacturer’s instructions.

The principle of Omniscript Reverse Transcriptase (ORT) is that it has a high affinity for RNA, which enables efficient and sensitive reverse transcription of any template, leading to high yields of cDNA. In brief per 20ul reaction: RNA template was varied (50ng- 2ug), 10x Buffer RT 2ul, dNTP Mix (5uM) 2ul, Oligo Ft primer (10uM) 2ul, RNase Inhibitor (10units/ul) 1ul, ORT 1ul,

RNase free water added accordingly. Preperation was mixed and incubated for 1 hr at 37C. cDNA was stored at -80C.

3.8 Sanger sequencing

Genomic DNA from either expanded peripheral blood T-cells, bone marrow fibroblasts, or whole blood cells were extracted and prepared using the Wizard Genomic DNA Purification Kit

(Promega). Sequencing primers were designed for desirable regions by Primer3 tool. PCR amplifications for genomic DNA were carried out using Maxima HotStart Taq DNA polymerase

(Fermentas) (Tsangaris, Klaassen et al. 2011). PCR products were purified by gel extraction or

PCR cleanup using QIAquick PCR purification Kit (Qiagen) and submitted for Sanger sequencing to The Center for Applied Genomics, The Hospital for Sick Children, Toronto

(TCAG). Targeted genes were analyzed by bidirectional sequencing of individual exons and flanking intronic regions after PCR amplification. Sequences were analyzed using Finch TV and aligned with the NCBI gene database accession.

3.9 Cell culture

HEK 293T cells were cultured in RPMI 1640 with 25 mM HEPES (WISENT BIOCENTER,

350-05-CL) supplemented with 10% FBS, and 1% antibiotics (penicillin).

BM stromal cells were cultured in Medium - Nonhematopoietic stem cell (NH Expansion Media,

Miltenyi Biotec,680-091-130 ) or DMEM high glucose with L-Glutamine and Sodium Pyruvate

(Wisent Biobar, 319-005-CL) supplemented with 10% FBS, and 1% antibiotics (penicillin).

T- cells were cultured in Advanced RPMI Medium 1640 or OpTmizer™ T Cell Expansion SFM

(Invitrogen Gibco Cat. no. 0080022SA) with 100 U/ml penicillin/streptomycin and Dynabeads®

Human T-Activator CD3/CD28 and IL2.

3.10 shRNA expression cassettes and generation of DNAJC21- knockdown cells

DNAJC21 short hairpin RNA (shRNA) and a scrambled (SCR) RNA control sequence were cloned into a pGIPZ vector by SPARC facility.

Sense sequence of 2 shRNAs :

YD84: TGGAAGAACATGAACTCAA

YD83: AGCTTAGATTmTGCTACGCT 40

Picture 1: Plasmid Map. HEK293T cells were transfected with the plasmids and cultured for 1 week with puromycin antibiotic. Individual colonies were isolated after 7-8 days. Four knockdown lines (YD83-6,YD83-7, YD84-3 and YD84-4) and 1 control line with scrambled RNA (Mock) were established.

3.11 Viral vector production and transduction

SPARC facility produced lentivirus containing shRNA for DNJAC21 gene. SPARC facility uses the Open Biosystems TransLenti Viral GPIZ Packaging System (which is a part of Dharmacon, began as an entrepreneurial start-up in the Hudson Alpha Institute of Biotechnology offering access to cDNA, ORF and shRNA products through academic partnerships). Hek293T cells were transduced in the presence of polybrene.

3.12 Western Blotting

Cultures were washed once, and cells were scraped into phosphate-buffered saline. After centrifugation, pellets were incubated with RIPA Lysis Buffer System (Santa Cruz, SC-24948) on ice for 15 min and centrifuged at 12,000 rpm for 10 min at 4C. The supernatant was collected and protein concentration was measured using Bradford protein assay solution (BioRad). Equal amounts of cell lysate were loaded onto a 10–15% sodium dodecyl sulfate-polyacrylamide gel

electrophoresis (SDS-PAGE) performed and transferred to nitrocellulose 0.2um membrane.

Membranes were rinsed in Tris-buffered saline (TBS) and incubated in blocking solution (5% non-fat dry milk in TBS-Tween 20) at room temperature for 1 h. Blocked membranes were incubated with a primary antibody at 4C overnight and washed, followed by incubation with a secondary antibody at room temperature for 1.5 h. After washing, the proteins were visualized using ECL.

3.13 Antibody list

DNHD1(s-20)- Santa Cruz, SC-323794,rabbit polyclonal HRP linked, recognize both long and short isoform of gene

DNHD1- NovusBio, NBP1-900741, rabbit polyclonal HRP linked, recognize long isoform only

DNAJC21 – Cederlane, 23411-1-AP, rabbit polyclonal HRP linked

Vinculin – Sigma Aldrih,v9131, monoclonal mouse HRP linked

β-actin - Santa Cruz,SC-1303300, monoclonal mouse HRP linked

β-Tubulin –Cell Signaling, 2146, Rabbit HRP linked

Caspase 3 - Cell Signaling, 9662, Rabbit polyclonal HRP linked, recognize pro-caspase and cleaved caspase.

Cleaved PARP – Cell Signaling, 9541, Rabbit polyclonal HRP linked, ASP214 clone

Annexin V for flow – BD Biosciences, V450 Annexin V 560506

3.14 Evaluating cell growth using Alamar Blue Assay

Alamar Blue is a cell viability indicator that uses the natural reducing power of living cells to convert resazurin to the fluorescent molecule, resorufin. The active ingredient of alamar Blue

(resazurin) is a nontoxic, cell permeable compound that is blue in color and non- fluorescent. Upon entering cells, resazurin is reduced to resorufin, which produces a bright red fluorescence. Viable cells continuously convert resazurin to resorufin, thereby generating a quantitative measure of viability.

HEK293T cells were cultured in RPMI medium with 10% FBS and 1% antibiotics (growth medium) to confluence level of 70-80%. Then, cells were collected and were prepared for plating by dilution to 5x104 cells/ml using growth medium. 5x103 Hek293T cells (i.e. 100ul of cell plating mixture) were plated per 1 well of 96-well plate. Cells were incubated for 2 hr at

370C in 5% CO2 to allow attachment to the plate. After 2hr, alamar blue (10 ul) was added per well. Fluorescence was read in a fluorescent plate reader (ex. 570nm em 585nm) at 4 hr

(considered as baseline (T0) and at 24 hr (T1). No FBS was used during cell plate reading.

3.15 Intracellular Reactive Oxygen Species (ROS) Levels and evaluating ability of cells to balance excess ROS levels generated by addition of H2O2

Intracellular and induced ROS level was measured by H2-DCFDA (Sigma, D6883). H2-DCFDA is a cell-permeable indicator for ROS that is non-fluorescent until the acetate groups are removed by intracellular esterases during intracellular oxidation reactions. When oxidized by ROS H2-

DCFDA is irreversibly converted to the fluorescent form, DCF.

Picture 2: H2DCFDA reaction For ROS measurements, HEK293T cells were cultured in RPMI medium with 10% FBS and 1% antibiotics (growth medium). Then, cells were collected and diluted to 5x104 cells/ml using growth medium to prepare and plated in per 1 well of pre-coated with 1% gelatin of 96-well plate for 24hr. Cells were then washed and incubated in 370C/5% CO2 for 30 min with 50 uM

H2-DCFDA in growth medium without FBS. After 30 min, 100uM H2O2 (30% Hydrogen

Peroxide, 7722841, BioShop) was added to the culture without changing medium, and incubated for 30 more minutes. After incubation, fluorescence was measured in Spectra Max Gemini microplate reader with an excitation at wavelength of 485 nm and emission at 535 nm. Since

DNAJ21 mutant cells grew slower than control cells, at the end of experiment (24 hours after) we normalized ROS fluorescent levels to the total amount of DNA as a surrogate marker of cell number. To do so, after ROS fluorescence reading, cells were treated with 1% Saponin for 30 minutes (to induce total cell death) and then 50ug/ml PI (Propidium iodide) was added for 1 hr to measure the total amount of DNA. Reading was done with 540nm excitation and 635 nm emission.

3.16 Estimation of Cell death by Propidium iodide

Propidium iodide (PI) is a commonly used red-fluorescent nuclear and chromosome counterstain.

Since propidium iodide is not permeant to live cells, it is frequently used to detect dead cells

(Dumitriu, Mohr et al. 2001). PI binds to DNA by intercalating between the bases with little or no sequence preference(Medin, Migita et al. 1996). In aqueous solution, the dye has

excitation/emission maxima of 493/636 nm. Once the dye is bound, its fluorescence is enhanced

20- to 30-fold, the fluorescence excitation peak is shifted ~30–40 nm to the red and the fluorescence emission peak is shifted ~15 nm to the blue, resulting in an excitation maximum at

535 nm and fluorescence emission maximum at 617 nm(Lackey, Press et al. 2002).

HEK293T cells were cultured in RPMI medium with 10% FBS and 1% antibiotics (growth medium). Then, cell plating mixture was made by collecting and diluting 5x104 cells per 1 ml of growth medium. 5x103 Hek293T cells (i.e. 100ul of cell plating mixture) were plated per 1 well of pre-coated with 1% gelatin 96-well plate for 24hr. Next day, the medium was changed to medium with 2% FBS with or without 100uM H2O2, and the cells were incubated further at

370C in 5% CO2 for 24 hr. After incubation, 50ug/ml PI was added to the culture and the cells were incubated for one more hour in 370C/5% CO2. After incubation, fluorescence was measured in a Spectra Max Gemini microplate reader with an excitation at wavelength of 540 nm and emission at 635 nm. Then 1% of Saponin was added for 30 min and incubated at room temperature without changing medium 50ug/ml of another PI aliquot was added to the wells.

Plates were incubated at RT for another 1hr and fluorescent read was done (ex. 540nm em.

635nm).

3.17 Flow cytometry and analysis

Flow cytometry is a technology that is used to analyze the chemical and physical characteristics of cells, particles that were labelled to fluorescent antibody in a fluid as it passes through at least one laser. Cell components are fluorescently labelled and then excited by the laser to emit light at varying wavelengths.

HEK 293T cells were plated ins 6 well plates in confluency of 30%-50%. After 24 hours the medium was changed to normal growth medium or medium with 100uM H2O2 for 16 hours. On

the day of flow, the cells were collected using 0.05% trypsin, washed and stained for Annexin V and PI using the company standard protocol.

1. Wash cells twice with cold PBS and then resuspend cells in 1× Binding Buffer at a concentration of 1 × 10^6 cells/ml.

2. Transfer 100 µl of the solution (1 × 10^5 cells) to a 5 ml culture tube.

3. Add 5 µl of V450 Annexin V.

4. Add 10 µl PI. The optimal concentration of PI may vary among cell lines where 10 µl of a

50 µg/ml stock is most likely the maximum to be required. Less may yield optimal results in some experimental systems.

5. Gently vortex the cells and incubate for 15 min at RT (25 °C) in the dark.

6. Add 400 µl of 1× Binding Buffer to each tube. Analyze by flow cytometry within 1 hr.

The following controls are used to set up compensation and quadrants:

1. Unstained cells.

2. Cells stained with V450 Annexin V (no PI).

3. Cells stained with PI (no V450 Annexin V).

4. Unstained Mock that was used as single color for FITC

The samples were analyzed by using flow cytometry LSRII-CFI, LSRII-GC, or LSRII-Fortessa cell analyzer (BD Biosciences). Cell analyses were analyzed using Flowjo Version 10. Gates were determined using single stain with less than 1% outliers.

3.18 Statistical Analysis

All the experiments were repeated at least three times with 3 or 4 replicates and the results were expressed as mean ± SD. Statistical significance was determined by Student’s t-test. Values with

P < 0.05 were considered significant.

Chapter 4 Results 4.1 IBMFSs families analyzed by WES analysis 4.1.1 Sample preparation

Looking at CIMF R database we have identified 125 patients with unclassifiable syndromes, who could not be assigned a specific syndromic diagnosis and did not have a known genotype.

These patients fulfilled our previously reported criteria for having an IBMFS (Tsangaris,

Klaassen et al. 2011), which mandated having at least 2 features from the first list and at least 1 feature from the second list as described below:

1. Fulfill at least 2 of the following criteria (related to the chronic bone marrow failure) a. Chronic cytopenia(s) detected on at least 2 occasions over at least 3 months b. Reduced marrow progenitors or reduced clonogenic potential of hematopoietic progenitor cells or evidence of ineffective hematopoiesis c. High fetal hemoglobin for age d. Red blood cell macrocytosis (not caused by hemolysis or a nutritional deficiency)

2. Fulfill at least 1 of the following criteria (related to inheritability of the disorder) a. Family history of bone marrow failure b. Presentation at age <1 y c. Anomalies involving multiple systems to suggest an inherited syndrome

d. Positive genetic testing

In addition, there was a large number of patients enrolled in the CIMFR who could be assigned a specific genetic diagnosis but no specific mutations were identified. Among the latter group, 8 patients had Shwachman-Diamond syndrome (SDS) without mutation in SBDS.

We have extracted DNA from peripheral blood, bone marrow fibroblasts or skin fibroblasts from 12 samples from patients or SDS without SBDS mutations. Extracted DNA underwent whole exome sequencing (WES) Illumina Hiseq 2500 platform. WES allows sequencing of all the protein-coding genes in the genome. There are 180,000 exons, which constitute about 1% of the human genome, or approximately 30 million base pairs, but mutations in these sequences are much more likely to have severe consequences than in the remaining 99%. The goal of this approach is to identify genetic variants that are responsible for both mendelian and common multigenic/multifactorial diseases.

4.1.2 Analysis of genetic alterations

Sequencing data was analyzed by workflow summarized in Figure 1. First, we filtered out all variants that did not pass the quality filters, such as standard bias (SB) less than 0, quality by depth (QD) >=5. Next, we chose variants with frequency lower than 0.01 or equal to blank from 1000 genome and ESP database. Then, we removed all homologues variants from parents (or non-affected samples). We divided the remaining variants to 5 groups from high to low prioritized: homologues (variants in this group were highly prioritized), compound hetro variants, denovo hom, and denovo hetero. Under each group we applied additional filters, such as conserved regions (phyloMam: values >=1 is moderate conservation, values higher or equal 2.5 strong conserved, phyloVert100 Values >=1.5 moderate, values >=4 strong). In addition, utilizing 4 different algorithms we tested whether the mutation/variation negatively 49

affected protein function (sift: variants<=0.05 considered damaging, polyphen: values >=0.95 considered damaging, MutantGene: values >=2 damaging). Lastly, we considered whether each gene, if damaged, can cause the symptoms observed in patients. After the described filtering we chose highly prioritized genes for each family (Table 1).DNHD1 and DNAJC21 were the 2 top hits selected for future investigation. Both genes were associated with SDS disease.

Figure 1: Flow for gene periodization from WES data. The data from TCAG was received in excel and bam file. First, all reads that did not pass the quality filters were taking out. Next, we filtered out all frequent variants and synonymous. Lastly, we choose high conserved regions and prioritized genes by their important potential function to our disease. The programs that we using is: 1000 HumanGenome, sift, provean, exac, kaviar, UCSC, dbsnp, genomecard, ncbi,gwas central, protein atlas and more.

4.2 DNHD1 as a candidate SDS gene

4.2.1 Clinical phenotype and genomic characterization

We identified two patients with compound heterozygous variants in DNHD1 gene. The first patient (p1) fulfilled the criteria for SDS had severe aplastic anemia, low pancreatic enzyme

(serum trypsinogen 6mcg/L, fecal elastase 76ug/g), pancreatic lipomatosis, atrophy on CT, echogenic pancreas on US, low vitamin A and E, osteopenia, metaphyseal dysplasia in femur and tibia, development delay, and normal SBDS sequencing. This patient possessed a mono-allelic rare large deletion of 32 Kb confined to the DNHD1 gene that was inherited from father. (Figure

2). To confirm that the deletion was inherited from the father we performed qPCR analysis comparing expression of the exon 7 from two independent stroma controls, patient and parent’s stroma (all samples were base lined to an expression of FOXP2 housekeeping gene). Ratio of >

0.7 indicates 2 copies and ratio < 0.7 indicates 1 copy. Figure 2 demonstrates that both the father and the patient have one copy deletion. Thus, the patient inherited from father a heterozygous deletion in the DNHD1gene. DNHD1 has 41 exons and two isoforms, short 8 exons and long all

41 exons(Anazi, Maddirevula et al. 2017, Wu, Hivert et al. 2017, Xu, Liu et al. 2017). The deletion crosses out 8 first exons (the whole short isoform) and 6772 bases upstream of the methionine.

Patient 1 (p1) was a single child of non-consanguineous parents. The father was reported healthy without any blood disorders; therefore, we checked for a point mutation that could be inherited from a mother. WES of mother`s DNA showed 17 variants in total, and only one heterozygous variant c.8019T>C (cDNA variant), p. Leu2673Pro (protein variant) was considered as potentially damaging (Table 2) since the gene was altered in more than one patient, the amino acid was conserved and several algorithms predicted the variant to be damaging to protein function; albit not all. This variant was validated by Sanger sequencing. In Figure3 we can see a hetero point mutation in the patient and mother’s sample but not in the sample from the father. The nucleotide 51

thymine, “T”, was mutated in one allele to cytosine “C” in the patient and in the mother and gave us a new variant of amino acid substitution (Leu to Pro).

1.1

1 0.9 0.8 0.7

Exon FOXP2 to7 0.6

0.5

0.4

0.3

0.2

RatioDNHD1 of 0.1

0 Cntrl-1 Cntrl-2 PatientP318 1 Mother Father

Figure 2: Exon 7 deletion in DNHD1 in patient 1 was inherited from father. Stromal fibroblast cells from two independent controls, Patient 1, and parents were prepared and analyzed by q-pcr for exon 7 signal. The results were compared to FOXP2 housekeeping gene (gene that was not affected). Patient 1 and father had reduction in exon 7. Ratio of > 0.7 indicates 2 copies and ratio < 0.7 indicates 1 copy.

Table 1: Mutation Effect Prediction for family of patient 1. Prediction of affected variant inherited from mother. SIFT: values>= 0.05 predicted damaging variant. Polyphen: values>= 0.95 predicted damaging variant. MutationTaster >=50 disease causing.

Prediction software Score/prediction Comments

PolyPhen 0.998 Damaging

MutationTaster 98 (0-215) Disease causing

SIFT (SNP, run along with Provean) 0.010 Damaging

SIFT_human protein 0.27 Tolerated

MutationAssessor 0 Neutral

Provean -2.34 (longer isoform) Neutral

phastCons 0.903 (0-1) conserved

GERP 5.28 conserved

PhyloP conservation 0.649 (-14 to +6) Moderate conservation

Proband Mother Father

Figure 3: DNHD1-associated mutations in Patient 1. The Sanger Sequence was done to validate the mutation and its result was analyzed by using IGV program version 2.3. In this program red color shows line/letter as nucleotide “T”, green “A”, Blue “C”, and black “G”. For het variant the program use letter “Y”. Proband had inherited mutation from mother.

The second patient that was identified with two different heterozygous variants in DNHD1 gene.

Each variant was inherited from a different parent. Clinical manifestations of the patient were similar to the ones observed in the first patient. The patient had neutropenia, absence of granulocytes in bone marrow with normal cellularity, echogenic pancreas, low pancreatic enzyme lipase (<3), developmental delay, moderately short telomeres and normal chromosome fragility.

Mother was healthy; no data about the father was available. This patient was compound heterozygous for variants that were predicted to be damaging (Table 2). The first variant is c.

12144G>T that causes early stop codon p. R4048*. The second variant is c.6066G>C that replaces glycine with arginine (p.G2022R). Both variants were validated by Sanger sequencing in the

mother and the proband (Figure 4). Similar to the previous case, different algorithms had contradicting predictions in regards to the degree of damaging the variant has to the function of the protein.

Table 2: Mutation Effect Prediction for Family 2.

Prediction software c.12144G>T c.6066G>C Comments R4048* (mut1) G2022R (mut2) MutationTaster (0-215) 83 98 Disease causing / (Values >= 50 predicts Disease causing damaging effect) SIFT (SNP, run along 0.033 0.187 Damaging / with Provean) Tolerated (Values >= 0.05 predicts damaging effect) SIFT_human protein 0.01 1 Damaging / (Values >= 0.05 predicts Tolerated damaging effect) PolyPhen-2 Probably (Values >= 0.95 predicts damaging / damaging effect) Probably

1 1 damaging Provean -3.63 -1.05 Deleterious / Neutral MutationAssessor 0.345 1.725 Neutral / Low phastCons (0-1) 1 0.999 Conserved PhyloP conservation (- 1.691 2.179 Mod conservation 14 to +6)

Proban Mother

d Mut1 R4048*

Stop codon mutation

Mut2

G2022R

Missense mutation

Figure 4: DNHD1-associated mutations in Patient 2. The Sanger Sequence was done to validate the mutation and its result was analyzed by using IGV program version 2.3. In this program red color shows line/letter as nucleotide “T”, green “A”, Blue “C”, and black “G”. For het variant the program use letter “Y”.

4.2.2 Compound heterozygosity mutations in DNHD1 may reduce the protein level

Most protein damage prediction software programs we used predicted that p.Leu2673Pro variant

is highly damaging; SIFT, MutationTaster, and Polyphen (0.01, 98, and 0.998, respectively).

Mutation Assessor and Provean showed a neutral score for this mutation. The p. R4048* variant

that causes early stop codon appeared in compound heterozygous state with the missense variant

p.G2022R, which had conflicting damage predictions by the various softwares. Based on the

above information we decided to evaluate the effect of the biallelic variants on protein levels. We

performed Western blot analysis using two commercial antibodies: 1) against only long isoform

(BIONOVUS ab) 2) against long and short isoform (Santa Cruz ab). Unfortunately, we failed to

detect any protein using Western blot analysis in either control samples or cells derived from patient material. Therefore, we performed fluorescent immunoblotting using bone marrow stroma cells and the described antibodies. Interestingly, our results showed that DNHD1 protein was increased in patient cells and not decreased as we hypothesized (Figure 5a and b). DNHD1 protein was localized to Golgi as confirmed by co-staining with Golgi specific antibody (GM-130, figure

5a and b). It is possible that the modified protein is produced and is detected by the antibodies, though it may not be properly folded and consequently is accumulated in the Golgi/ER and is not readily transported to the cell membrane. This hypothesis remains to be further investigated. Since at this stage we found an alternative gene that was mutated in SDS patients (including in those with DNHD1 variants) with higher likelihood of pathogenicity we excluded DNHD1 from our prioritized list of SDS related genes in our patient population.

DNHD1 Merge

Control

GM-130 Merge DNHD1

5B Patient

DNHD1 Merge

Control

GM-130

Patient

Figure 5: Cellular localization of DNHD1. Stroma cells from patient and control were used in Fluorescent immunoblotting. (A) Staining with antibody from Santa Cruz, which recognizes both isoforms (the sort and long isoform). (B) Staining with antibody from bio Novus that recognize only long isoform. DAPI (blue signal) was used for nucleus staining. DHND1 localizes to the Golgi region.

4.3 Biallelic mutations in DNAJC21 cause Shwachman-Diamond syndrome

The experiments in 4.3.1 to 4.3.2 were done with Santhosh Dhanraj. Both, Santhosh Dhanraj and Anna Mateev were first co-authors on the publication that summarized these experiments.

4.3.1 Clinical phenotype and genomic characterization

Four patients from three unrelated families were identified with clinical diagnosis of SDS and were confirmed to be negative for mutations in SBDS gene. All four patients carried biallelic mutations in DNAJC21. Patient 1 of Family 1 was of Afghani ancestry from consanguineous parents. She presented with failure to thrive at the age of 2.5 years and was found to have pancytopenia, high mean corpuscular volume, and increased hemoglobin F (Table 3). The bone marrow was hypo cellular (Figure 6A). In addition, patient 1 had exocrine pancreatic dysfunction with a low serum pancreatic enzyme levels (Table 3), markedly increased echogenicity of the pancreas by ultrasound examination, and hypodense pancreas by computed tomography scan, consistent with lipomatosis (Figure 6B/C). The patient was treated with fat-soluble vitamin supplements. Her height was below the third percentile. Skeletal survey showed evidence of metaphyseal dysplasia, as described in SDS. The patient also had gross and fine motor developmental delay and bilateral retinal dystrophy; the latter is an unusual manifestation of SDS.

The second family included two affected children of consanguineous parents of First Nations

(Canada) ancestry. The first sibling (Patient 2) was diagnosed after birth with progressive bone marrow failure (Table 3) and died of Staphylococcus aureus sepsis at the age of 18 months. He was diagnosed with SDS and pancreatic insufficiency (low serum trypsinogen levels). Autopsy showed atrophic exocrine pancreas with fatty infiltration and observance of the endocrine tissue.

The second sibling (Patient 3) had pancytopenia with severe anemia after birth and later had stable moderate pancytopenia. At 8 months of age bone marrow testing showed hypo cellularity (Figure

7A), and a cytogenetic abnormality 46, XY, der(15)t(1;15)(q12;p11). However, the peripheral blood karyotype was normal. Patient 3 developed severe pancytopenia at 7 years of age: bone marrow transplantation was done. He died 2.5 months post-transplant due to Epstein-Barr virus– associated lymphoproliferative disorder. In addition, the patient had feeding problems, low serum pancreatic enzyme levels (Table 3), and a small hyper echogenic pancreas (Figure 7B-C). He was treated with fat-soluble vitamin supplements. Metaphyseal dysplasia in multiple joints (Table 3) and mild flaring of anterior rib ends as described in SDS were found at presentation. The third family included one affected patient (Patient 4) born to parents of Indian descent. The patient developed severe aplastic anemia at 2 years of age (Table 3) and underwent successful bone marrow transplantation. He had exocrine pancreatic dysfunction, short stature, metaphyseal dysplasia, and developmental delay. He was diagnosed with SDS and no SBDS mutation as previous patients. Of note, after transplantation, this patient developed retinal dysplasia, similar to Patient 1.

Whole-exome sequencing on peripheral blood from the patient and the parents of Family 1 was performed. DNAJC21 was considered to be the highest candidate gene causing the disease due to a homozygous stop codon variant (c.520C.T, p.Gln174*) that was not described in studied control populations (Table 4). We validated a mutation in this gene using Sanger Sequencing. We confirmed that patient 1 had homozygosity mutation and heterozygosity in the parents (Figure 8).

In patient 3 from family 2 DNAJC21 was also considered to be a high candidate for causing SDS.

This decision was made based on an identification of an extended run of homozygosity by SNP6.0 array, for both patient 1 and patient 3, and the parents’ consanguinity (Figure 9A-B). We performed Sanger sequencing to all the exons, and discovered a homozygous mutation c.100A.G

(Figure 10) that results in a substitution of a highly conserved amino acid p.Lys34Glu in the J-

domain (Figure 11). In addition, using multiple protein prediction software programs we predicted that this variant is damaging and disease-causing (Table 4). Moreover, CFSSPS protein structure prediction software, predicted that the mutation results in lengthening of a-helical segment and abolishes a turn within the DnaJ domain (Figure 12). Based on the available NMR structure of the J domain of murine polyoma T antigen (1FAF) (http://www.rcsb.org/pdb), residue K34 is exposed to the solvent side of the protein; therefore, the mutation is likely to change the surface charge and may interrupt with protein folding. Interestingly, initial analysis of WES results from

Family 3 did not reveal any nucleotide level mutations; however, careful examination of BAM file and specifically DNAJC21 sequence from Patient 4 revealed only few reads from both exons

5 and 6 in contrast to surrounding exons and parents` reads (Figure 14A); the parents had borderline read numbers in these regions. Furthermore, NextGene 2.4.2 software indicated a deletion of exons 5 and 6 with a high degree of confidence (Figure 14B-C). To validate this finding, we performed polymerase chain reaction (PCR) on genomic DNA from the described samples. It showed no amplification of exons 5 and 6, consistent with a deletion (Figure 14D).

Deletion of exons 5 and 6 is predicted to cause splicing that merges exon 4 with exon 7 resulting in a frameshift and early protein truncation p.(Val148Lysfs*30). Quantitative RT PCR using primers that amplify a cDNA fragment located between exons 4-7 was performed on RNA sample that was extracted from T-cells of members of family 4 revealed two heterozygosity variants in the parents (long (normal) and a short (mutant) alleles) and a homozygosity for the short allele in the patients (Figure 14E). Sequencing confirmed the absence of exon 5-6 in the short fragment (Figure 14F).

Patient 3 (Sibling of patient Patient 1 Patient 2 (Sibling of Patient 3) Patient 4 2) Age at presentation 2.5y After birth After birth After birth Age at diagnosis with SDS 2y, 11m <1y 7m 2y, 2m Age at last follow-up or death 14y 1.5 y 7y 4m 11 y Evidence of bone marrow failure Moderately low blood counts (from Severe bone marrow failure on Severe pancytopenia (7 y), Severe pancytopenia (2y), underwent bone (Age at testing) 2.5 y). chronic transfusion since birth, was underwent bone marrow marrow transplantation planned for transplantation transplantation 0.4 (2y) - Neutrophils (>1.5 × 109/L) 1.08 (3y), 3.54 (14y) Severely low 1.3 (8m), 0.5 (7y) 74 (2y) - Hemoglobin (>120 g/L) 67 (3y), 118 (14y) Severely low 97 (8m), 55 (7y) 17 (2y 2m) - Platelets (>150 × 109/L) 22 (3y), 118 (14y) Severely low 185 (8m), 17 (7y) 3.4 (2y) - Lymphocytes (>1.5 × 109/L) 2.56 (3y), 2.28 (14y) UK 5.83 (8m), 1.82 (7y) 32.5(2y 2m) - Reticulocytes (>40 × 100/L) 96 (14 y) UK 102 (8m), 33 (7y) 87(2y) - MCV (0.5-3y: 70-86 fL, 7-14y: 75- 89 (3y), 94 (14y) UK 77 (8m), 83 (7y) 96) 11.7 (2y 2 m) - HgF (%)(After 1y <1.2%) 29 (2.5 y) UK 33 10 (2y 5m) -Bone marrow cellularity (%) 40-50 (2.5y) Markedly hypocellular Mildly reduced (8m), 10-50% No - Prominent marrow dysplasia No No (7y) N - Marrow cytogenetics N N No 46,XY,der(15)t(1;15)(q12;p11 ) (8m-7y) Evidence of pancreatic dysfunction - Chronic diarrhea No No No No, fecal fat in microscopy - Lipase (23-300 u/L) 22 (2y 11m) UK 15 (8m) <25 (2y 2m) - Amylase (20-110 u/L) 36 (2y 11m) UK 37 (8m) <30 (2y 3m) - Pancreatic isoamylase (>17 u/L)** 10 (2y 11m) UK 2 (8m) NA - Trypsinogen (>16.6 µg/L) 12.7 (2y 11m) 11.9 (12 m), 9.2 (13 m) 20 (8m) 3.4 (2y 2m) - Vitamin A levels (0.7-2.1 mmol/L) 1.0 (2y 11m) UK 1.4 (8m) 1.4 (2y 2m) - Vitamin D levels 70-250 nmol/L) 60 (2y 11m) UK 39 (8m) 69 (5y 10 m) - Vitamin E levels (12-46 µmol/L) 9.3 (2y 11m) UK 5.2 (3y) 8.4 (2y 2m) - INR (0.9-1.1) 0.9 (2y 11m) UK 0.97 (8m) NA - Hyperechogenic pancreas on US Markedly echogenic UK Markedly echogenic Diffusely echogenic - Hypodense pancreas on CT Hypodense pancreas UK Hypodense pancreas NA Patient was diagnosed with Autopsy (7y 4m): pancreatic pancreatic insufficiency before acinar atrophy death. On autopsy (18m) he had atrophy of exocrine pancreas with fatty infiltration (no changes in the endocrine pancreas). Metaphyseal dysplasia Yes Anomaly of the wrist bone (no Yes (distal radius and ulna, No, but have osteopenia and focal metaphyseal further details) distal femur, proximal tibia. irregularity in right proximal femur Cognitive impairment Yes UK Yes Delayed gross motor development

Stature <3% 30% (11m) <5% (7.5m) <3%

Liver No hepatomegaly, no elevated liver Mild fatty changes of the liver (on No hepatomegaly / mild No hepatomegaly, mild elevation of the liver enzymes autopsy) elevation of liver enzymes in enzymes infancy. Other medical problems Retinitis pigmentosa, generalized Died at the age of 18 months due to Retinal dystrophy without pigmentation, low seizures (6m, 10m), severe eczema, staphylococcus aureus sepsis. visual acuity, decrease visual field, persistent asthma, recurrent otitis media, eczema, failure to thrive, kyphosis, mild high- subtle minor physical foot inversion, abnormal dentition, malformation (low anterior hypopigmentation macule at right thigh hairline, deep-set eyes, mildly (developed after transplant) cupped ears, down-turned corners of the mouth, a horizontal crease on the chin, broad neck and redundant skin, increased creases in the palms and deep folds bilaterally, No hearing deficits). Other tests with normal results Chromosomal breakage studies, Chromosomal breakage Chromosomal breakage studies telomere length, next generation studies, Targeted sequencing of SBDS, RMRP, sequencing of a panel of known 72 Targeted sequencing of SBDS, Affymetrix 6.0 Array. IBMFS genes, targeted sequencing RMRP, Affymetrix 6.0 Array. Mitochondrial DNA deletion analysis (for of SBDS, urine organic acid and Pearson syndrome) plasma amino acids, 4X180K oligonucleotide array (Agilent Tech), Affymetrix 6.0 Array.

Table 3: Patients with mutations in DNAJC21 meet the clinical criteria for SDS HgF, hemoglobin F; m, month; MCV, red blood cell mean corpuscular volume; N, Normal; UK, unknown; y, year * Quantification of cellularity in percentages was not available and commonly used clinical terms to describe bone marrow cellularity were reported to the registry. We refer to “markedly reduced” cellularity in children as <25% and “mildly reduced” cellularity as 50-75%.” **Reference levels are from about the age of 3 years onwards

A B C

Figure 6: Clinical characteristics of patient 1. (A) Reduced cellularity is visible in the marrow biopsy. Hematopoietic activities take around 40% of the space between bones, also surrounding fat tissue is seen (magnification ×100) (B) Imaging of the pancreas showed hyper echogenicity (arrow) on ultrasonography. (C) Normal pancreas from 4 years old boy with aplastic anemia and no Shwachman-Diamond syndrome.

A B C

Figure 7: Clinical characteristics of patient 3. (A) Reduced cellularity is visible in the marrow biopsy. Hematopoietic activities take around 40% of the space between bones, also surrounding fat tissue is seen (magnification ×100) (B) Imaging of the pancreas showed hyper echogenicity (arrow) on ultrasonography. (C) Normal pancreas from 4 years old boy with aplastic anemia and no Shwachman-Diamond syndrome (Figure 7C is also shown in 6C).

Table 4: Information about DNAJC21 and C2orf71 variants found in the patients in the present study

MAF Conservation c.DNA Variation Genomic DNA Protein Change type/Zygosity location(hg19) (ExAC) (GERP) location

DNAJC21

Patient 1 SNP/Hom 5:34937512,C>T 0 5.39 c.520C>T p.Gln174*

p.Lys34Glu

Prediction: damaging/disease causing (SIFT, Patient 2,3 SNP/Hom 5:34933922,A>G 0 5.1 c.100A>G Provean, MutationTaster); affect the protein (SNAP2);

probably damaging (Polyphen-2)

Deletion of exon 5 and 6, and merging of Deletion of exon 5-6 exon 4 with exon 7 Patient 4 Biallelic Mutation (specific breakpoints 0 Not applicable c. 438-?_894+?del were not determined) (prediction: p.(Val148Lysfs*30))

C2orf71

-3.66 (conservation of the Patient 1 INDEL/Hom 2: 29293978,T>TG 0.00018 c.3149_3150 insG p.Pro1051Thrfs*56 T nucleotide)

A Family 1 I-1 I-2 II-1 B

c g a c/t a g g c g a c/t a g g c g a c/t a g g I-1 I-2 ◐ ◨ p.Gln174*/WT p.Gln174*/WT

II-1

5 c.520C>T/WT c.520C>T c.520C>T/WT p.Gln174*/● ⋄NT p.Gln174* Figure 8: DNAJC21-associated mutations in Patient 1. (A)Sanger sequencing was done to validate mutations in DNAJC21 (NM_001012339.2) that was found by WES. The figure indicates the position of the mutation (c.520) on exon 5. The nucleotide substitution of C by T is seen in homozygous (patient, II-1) and heterozygous (parents, I-1 and I-2) states, consistent with recessive inheritance. The results of alteration is premature protein truncation (p.Gln174*). (B) Pedigree of Family 1.

Family 2 I-1 I-2 A B ◐ ◨ P1 p.Lys34Glu/WT p.Lys34Glu/WT P3

II-1 II-2 II-3 II-4 ◻ ◻ ◼ ◼ p.Lys34Glu p.Lys34Glu p.Lys34Glu/ p.Lys34Glu/ WT WT p.Lys34Glu p.Lys34Glu

Figure 9: Region on homozygosity on chromosome 5 overlap the DNAJC21 locus in patients from family 2 (A) Multi-Mb regions of homozygosity in DNA from Patients 1 and 3 were detected by Affymetrix SNP 6.0 and overlap the DNAJC21 locus. (B) Pedigree of Family 2.

I- 1 I- 2 g a t a/g a g a t a/g a

c.100A>G c.100A>G II 1 II 2 II 3 II -4 g a t a/g a g a t a/g a g a t g/g a g a t g/g a

Figure 10: DNAJC21-associated mutations in Patient 3. Sanger sequencing was done to validate the portion of exon 2 of DNAJC21 in family members showing segregation of the pathogenic missense variant.

DnaJ Coiled Coil ZnFC2H2 ZnFC2H2

1 100 200 300 400 500 531

p.Lys34Glu p.Gln174* (Patient 2, 3) p.(Val148Lysfs*30) (Patient 1) (Patient 4)

Figure 11: DNAJC21 protein domains. The DNACJ21 protein with its domains and predicted patients’ mutations are shown.

DNAJC21

DNAJC21 Mutant *

Figure 12: The effect of the p.K34E mutation in DNAJC21 on the protein secondary structure. The effect of mutation p.K34E on the secondary structure of the protein was analyzed using a protein structure prediction software CFSSPS (Chou & Fasman Secondary Structure Prediction Server, http://www.biogem.org/tool/chou-fasman/). The mutation was predicted to lengthen an alpha-helical segment and abolish a turn within the DnaJ domain. The asterisk shows the amino acid which was changed.

Family 3

I-1 I-2

del exon◐ 5,6/WT del◨ exon 5,6/WT p.(Val148Lysfs*30)/WT p.(Val148Lysfs*30)/WT

II-1 ◻ ◼ NA del exon 5,6/del exon 5,6

p.(Val148Lysfs*30)/ p.(Val148Lysfs*30)

Figure 13: Pedigree of family 3.

BAM file

Number of reads by WES

B C

D E

Control

Father

Mothe r

Blank

Control

Control Father

Patient

Father Mothe r

Marker

Mothe r Blank

Blan

Patient Patient 698

bp 241bp DNAJC21 Exon 5 DNAJC21 Exon DNAJC21 Exon 6 9

DNAJC21 Exon DNAJC21 Exon GAPDH 68 4 7

Figure 14: Genetic investigation of Family 3. (A) A BAM file showing reduced reads in exon 5 and 6 of the DNAJC21 gene. (B) CMV analysis of exome sequencing data by the NextGene 2.4.2 program (SoftGenetics LLC,) revealed a deletion at chromosome 5p13.2. (C) Analyzing by the NextGene software program revealed that a high level of confidence that the deletion is real. (D) PCR of genomic DNA from patient 4 showing absence of amplification of exons 5 and 6, and normal amplification of surrounding exons and glyceraldehyde-3- phosphate dehydrogenase control. (E) RT-PCR of 4 exons (from exon 4-7) is showing 2 products (small product of 241bp without exon 5 and 6, and a big product of 698 bp with exons 4-7) for parents and one small product for patient4. (F) Amplification of cDNA using fragments flanking the gene sequencing from exon 4 to 7, revealed inclusion of exon 5-6 in the long fragment (upper panels) and exclusion of exon 5-6 in the short mutation fragment (Lower panel).

4.3.2 Biallelic mutations in DNAJC21 reduces protein levels

DNAJC21 encodes two transcripts with a common isoform of 531 amino acids protein(Chen, Yin et al. 2004). This protein contains a highly conserved amino-terminal DnaJ domain (Craig, Huang et al. 2006), with a centrally positioned coiled coil region, and a carboxyl segment with two C2H2- type zinc fingers that outflank 168 amino acids enriched in charged residues (Meyer, Hung et al.

2007). The predicted structure of the protein and the location of the mutation are shown in Figure

11. Immunoblotting uncovered a markedly reduced protein levels in cells from Patient 1 that carried a homozygous nonsense mutation, and in Patient 4 with a biallelic exon 5-6 deletion compared to healthy control subjects. In addition, a 40% reduction in protein levels was observed in a sample from Patient 3, who had a homozygous missense mutation (Figure 15A). Specificity of the antibody was tested by shRNA-mediated DNAJC21 knockdown in HEK-293T cells (Figure

15B). We quantified the levels of SBDS protein in the patients, and they did not have a prominent reduction in the levels of SBDS (about 10% residual protein), as usually seem in SDS patients with mutations in this gene. (Figure 15C).

l 2 l

Mother

Father

Patient1

Patient3

Patient4

Control4

Control3

Control1

Contro

Control2 Control1 DNAJC21 75 kDa

Vinculin 124 kDa

3 (YD85)

1 (YD83)

2 (YD 84)

Mock

HEK

KD KD DNAJC21 75 kDa

Vincullin 124 kDa

1.2

C 0.8 Actin

- 0.6

0.4 SBDS 0.2

SBDS/B 0 β-actin Patient1Patient2Patient3 Control Figure 15: Expression of DNAJC21 protein (A) Immunoblots of protein extracts of cells from controls and members of Patient 1 (left, peripheral blood T cells), Patient 2 (center, peripheral blood T cells), and Patient 4 (right, marrow fibroblasts) with DNAJC21 and loading control antibodies. NT, not tested. (B) Immunoblotting of DNAJC21 in HEK- 293T cells and stable Knockdown cells. Three shRNA sequences targeting DNAJC21 were used for gene knockdown. After transient transfection, cells were stably selected using Puromycin (1ug/ml). Protein was extracted and assayed by an antibody that recognizes DNAJC21 (400-450 aa region) and an antibody that recognizes vinculin as a loading control. (C) Immunoblotting of SBDS in marrow fibroblast cells of patients with SDS and negative SBDS gene mutations shows only moderate reduction in SBDS levels, but not as low as seen in patients with biallelic SBDS mutations (typically <15%). The right figure shows densitometry of blots, was performed with ImageJ software.

4.3.3 Establishing DNAJC21-knockdown cells

To study the effect of DNAJC21-deficiency in affected cells, we generated HEK293-T cells with the DNAJC21 knockdown. For this purpose, HEK293-T cells were stably transduced with three different shRNA against DNAJC21. We chose 2 lines (YD83 and YD84) that showed null protein expression, as confirmed by Western blotting using an anti-DNAJC21 antibody (Figure 15B).

After two weeks in culture, we picked up 12 clones from each line and tested them separately by using the same immunoblotting method, to choose the best clones that maintained no protein expression of DNAJC21. Four best clones were selected for this study, YD83-6, YD83-7, YD84-

3, YD84-4 (Figure 16).

WT Mock YD83-6 YD83-7 YD84-3 YD84-4 DNAJC21 Vinculin

DNJAC21 protein level 1.6 1.4 1.2 1 0.8 0.6

0.4 DNAJC21/Vinculin 0.2 0

WT Mock YD83-6YD83-4YD84-3YD84-4

Figure 16: DNAJC21 Protein expression in DNAJC21- knockdown HEK 293T cells. After transient transfection, cells were stably selected using Puromycin (1ug/ml). Protein was extracted and assayed by an antibody that recognizes DNAJC21 and an antibody that recognizes vinculin as a loading control. Immunoblots of protein extracts of cells from WT, Mock, and 4 clones. The right figure shows band densities, was performed with ImageJ software. All 4 lines showed very low protein levels compared to WT or MOCK: YD83-6, undetected; YD83-4, 5% of controls, YD84-3, undetected; YD84- 4, 18% of controls).

4.3.4 The role of DNAJC21 in promoting cell growth

DNAJC21 was shown to have a role in ribosome biogenesis and could disrupt protein translation, which can lead to impaired cellular growth (Tummala, Walne et al. 2016). To check this hypothesis, we produced DNAJC21 KD HEK293-T cells. To evaluate the effect of the knockdown on cell proliferation, we utilized an Alamar Blue viability dye. We plated 1x104 cells in each well of 96 black well plates in the presence of Alamar blue and incubated in 37C for 4 and 24 hours.

Baseline evaluation 4 hours after plating showed no differences between WT-Mock, Mock- shRNAs, or WT-shRNAs transduced cells. However, after 24 hours we saw a significant reduction in the growth rate of the knockdown cells (YD83-6, YD83-7, YD84-3, YD84-4) to

around 60% of the levels seen in MOCK or WT cells (Figure 17).

Alamar Blue assay WT and Mock underling 30000 * 25000 WT 20000 units Mock 15000 YD83-6

10000 YD83-7

Flouroscent YD84-3 5000 YD84-4 0 Linear (Mock) 0 24 48 Hours

Figure 17: Cell growth rate is impaired in DNAJC21 knockdown cells. Six samples were cultured separately for a couple of days until 70% confluency and plated in 1x104 cells density with Alamar Blue for up to 24hr. Four technical repeats were done in 5 separate biological repeats. All 4 knockdown lines showed reduction in cell growth rate of around 60% compared to the mock line (P-values for all comparisons of knockdown lines and controls were lower than 0.01).

4.3.5 DNAJC21-KD Increases Baseline Intracellular Reactive Oxygen Species

It is known that a variety of cellular activities, such as metabolism, growth, and death, are modulated and regulated by the oxidative status of the environment that is influenced by the

−· presence of Reactive Oxygen Species (ROS), including superoxide (O2 ), hydrogen peroxide

(H2O2) and hydroxyl radical (HO·). Endogenous generation of ROS has been shown to directly activate certain signaling pathways. These signaling pathways activate cellular responses to big changes in the redox state of cellular environment and are required to maintain the oxidative balance to avoid DNA, protein, and lipid damage, which could potentially lead to necrosis, apoptosis, or senescence (Watanabe, Ambekar et al. 2009, Ambekar, Das et al. 2010). Also, studies have shown that oxidative stress can lead to a wide spectrum of responses at the cellular 73

level starting from proliferation and growth arrest to cell death (Austin, Gupta et al. 2008, Sen,

Wang et al. 2011). In addition, our previous study shows that reduction in SBDS protein leads to increased levels of ROS. (Dror and Freedman 2001, Rujkijyanont, Watanabe et al. 2008,

Watanabe, Ambekar et al. 2009, Ambekar, Das et al. 2010). Thus, we hypothesized that the decrease in proliferation observed in DNAJC21 knockdown cells is due to impaired regulation of

ROS. To determine whether DNAJC21-deficiency affects oxidative stress, intracellular ROS levels were measured in the knockdown lines using the H2-DCFDA reagent as described in methods. Figure 18 shows significantly higher levels of ROS in the DNAJC21-knockdown cells

(four shRNAs clones) compared to the WT and MOCK control cells. These data provide evidence that DNAJC21-deficiency results in increased ROS levels.

3.5 * * 3 * * 2.5 2 1.5 1

arbitraryunits 0.5 0 WT Mock YD83-6 YD83-7 YD84-3 YD84-4 Lines

Figure 18: Baseline ROS levels are increased in DNAJC21 knockdown cells. The lines were plated on 96-well plate for 24hr. After 24hr H2-DCFDA with 50uM concentration was added and incubated for 30 min in 37C. Plates were read by using Spectra Max Gemini microplate reader with an excitation at wavelength of 485 nm and emission at 535 nm. Normalized values to 1. Intracellular ROS level were significantly increased in all 4 lines compared to the mock line. Four technical repeats were done in 6 separate biological repeats. All p values were <0.05

4.3.6 DNAJC21 is critical for maintaining balanced levels of Reactive Oxygen Species

Next, we asked whether DNAJC21 plays a role in eliminating excess ROS. For this purpose, we treated the DNAJC21-deficient HEK293 cells with 100uM H2O2 for 1 hr after incubating them

with H2-DCFDAt. We have found that DNAJC21-deficiency led to a significant increase in ROS levels. Figure 19 illustrates that in all four Knock Down (KD) lines the levels of ROS are produced

4-5-fold higher compared to those observed in the MOCK or WT lines. Importantly, stimulation with H2O2 increases ROS levels to a significantly higher degree compared to the base line (without stimulation). Therefore, DNAJC21 does not only play a role in balancing intracellular ROS levels, but is also required for balancing excess ROS levels derived from extracellular sources.

12 * * 10 * * 8 6 4 arbitraryunits 2 0 WT Mock YD83-6 YD83-7 YD84-3 YD84-4 Lines

Figure 19: The ability to balance ROS levels after addition of exogenous H2O2 is impaired in DNAJC21-knockdown cells. The lines were plated on 96-well plate for 24hr. After 24hr H2-DCFDA with 50uM concentration were added and incubated for 30 min in 37C. Then 100uM of H2O2 were added for 1hr. Plates were read by using Spectra Max Gemini microplate reader with an excitation at wavelength of 485 nm and emission at 535 nm. Normalized values to 1. ROS level were significantly increased in all 4 lines compared to treated mock line. Four technical repeats were done in 6 separate biological repeats. All p values were <0.05.

4.3.7 DNAJC21 knockdown leads to accelerated cell death As aforementioned, oxidative stress can cause cellular damage by inducing apoptosis, necrosis or senescence in cells. We hypothesized that the knockdown DNAJC21 inhibits cell growth and inducing death by increasing ROS levels. To assess cell viability of knockdown cells, propidium iodide (PI) assay was performed. Figure 20 shows that all four shRNAs clones have significantly higher PI staining compared to MOCK or WT lines (1 fold to 1.7 folds) indicating that they have

decreased viability as PI can penetrate membranes of damaged/dying cells but not viable cells.

However, we did not observe a significant effect of H2O2 on PI staining/cell viability.

2.5 * * 2 * * 1.5 1

0.5 arbitraryunits 0

Lines

Figure 20: Cell death is increased in DNAJC21-knockdown cells. The lines were plated on 96-well plate for 24hr. After 24hr PI in 50ug/ml concentration were added and incubated for 1hr min in 37C. Plates were read by using Spectra Max Gemini microplate reader with an excitation at wavelength of 540 nm and emission at 635 nm. After first reading the cells were treated with 1% saponin and second read was done. Normalized values to 1. PI level were significantly increased in all 4 lines compared to the mock line. Four technical repeats were done in 8 separate biological repeats. All p values were p<0.05.

4.3.8 Anti-oxidants improve growth of DNAJC21-Knockdown Cells

Antioxidants are molecules that can reduce oxidative stress by inhibiting production of free radicals from their precursors or neutralize them. The above data indicate that DNAJC21 deficiency leads to increased levels of intracellular ROS and to a reduction of cellular proliferation. To test whether DNAJC21 gene is important for normal growth rate in HEK293T cells the following experiment was performed: Cells were exposed to two different antioxidants

(Ascorbic Acid (AA) and Catalase (Cat), 500uM concentration) for 24hr. After 24 hr the cells were washed, and Alamar Blue assay was performed as described earlier. The results indicate a significant levels of (p<0.05) rescued cellular growth (Figure 21A-B) in cells exposed to either

antioxidant. However, even in the presence of the antioxidants, KD cells have a lower growth rate comparing to the WT or MOCK. This result suggests the existence of yet an additional unknown mechanism by which DNAJC21 promotes cellular proliferation. To verify that the observed rescue in cellular proliferation in cells treated with the antioxidants resulted from the reduction in

ROS levels , we used H2-DCFDA reagent to check ROS levels in these cells. Figure 21 (C1,2-

D1,2) shows a significant reduction in levels of ROS in all the cells lines exposed to the antioxidants either with or without H2O2 stimulation. This confirms our hypothesis that DNAJC21 deficient cells undergo growth arrest and die due to oxidative stress.

A 40000 Alamar Bue Cat- * * * * 35000 Alamar Bue Cat+ 30000 25000 20000 15000

Flouriscents units 10000 5000 0 WT Mock YD83-6 YD83-7 YD84-3 YD84-4 Lines

45000 Alamar blue AA- 40000 * * * Alamar blue AA+ 35000 *

30000

25000

20000

15000

10000 Flouriscents units 5000

0 WT Mock YD83-6 YD83-7 YD84-3 YD84-4 Lines

C 1

3 * * * * H2O2 -/AA- 2.5 H2O2 -/A+ * * 2 * *

1.5

1 Normalizationto 1

0.5

0 WT Mock YD83-6 YD83-7 YD84-3 YD84-4

6 C 2 H2O2 +/AA- * * * * * * H2O2 +/A+ 5 *

4 * * * 3 *

2 Normalizationto 1

0 WT Mock YD83-6 YD83-7 YD84-3 YD84-4

3 * * H2O2 -/Cat- * 2.5 * H2O2 -/Cat+

1.5 * *

1 Normalizationto 1

0.5

0 WT Mock YD83-6 YD83-7 YD84-3 YD84-4

6 * * * H2O2 +/Cat- * H2O2 +/Cat+ 5

4 * *

2 Normalizationto 1

0 WT Mock YD83-6 YD83-7 YD84-3 YD84-4

Figure 21: Antioxidants decrease ROS levels and improve cell growth rate in DNAJC21- knockdown cells. Six types of samples were cultured for 2 days until 70% confluent and plated at 1x104 cell density. Four different experiments were performed (A) cell growth analysis by alamar blue after treatment with catalase for 24hr. The cell growth rate significantly increased after using anti-oxidant (B) Cell growth analysis by alamar blue after treatment with ascorbic acid for 24hr, (C1,2) ROS level assay after treatment with the antioxidant ascorbic acid. (D1,2) ROS level assay after treatment with catalase for 24hr. Six technical repeat were done in 3 separate biological repeats (Asterisk* indicates p<0.01)

Chapter 5 Discussion and Conclusion

Frequently, patients who present with bone marrow failure cannot be directly assigned with genetic diagnosis of a specific syndrome. Our group and others showed that using whole exome sequencing results in a high identification rate of causal mutations with sometimes unexpected genetic findings and amendment of clinical diagnoses (Dhanraj, Manji et al. 2013, Dhanraj,

Gunja et al. 2015, Marshall, Farrell et al. 2015).

The present study shows that genome-wide analysis such as whole exome sequencing can reveal causal mutations in several patients with IBMFSs without known gene mutations.

It is well known that patients with nucleotide-level mutations in IBMFS genes can express both hematological and non-hematological symptoms as a result of a single nucleotide mutation in a gene (Teo, Klaassen et al. 2008, Tsangaris, Klaassen et al. 2011). CIMFR database offers the opportunity to assess a large cohort of patients with IBMFSs in Canada. Current analysis included 13 patients tested with a genome-wide approach to identify nucleotide-level mutations which revealed a causal genetic germline mutation in about half of the patients (Hashmi, Allen et al. 2011).

Shwachman-Diamond syndrome is an inherited multisystem disorder (Dror and Freedman 1999,

Dror, Ginzberg et al. 2001, Dror 2005). The clinical diagnostic criteria include bone marrow failure and exocrine pancreatic dysfunction. In addition, the diagnosis of SDS is supported by patients having short stature, metaphyseal dysplasia, and social behavior difficulties (Bodian,

Sheldon et al. 1964, Dror and Freedman 1999, Ginzberg, Shin et al. 1999). Most patients with this diagnosis have mutations in the SBDS gene(Aggett, Cavanagh et al. 1980). This gene has

been associated with multiple cellular pathways. Most prominent role of SBDS is in the releasing of eIF6 from the 60S ribosome subunit enabling a final stage of its maturation and a formation of monosome 80S ribosome. Approximately 10% to 20% of SDS patients do not have mutations in the SBDS gene (Bodian, Sheldon et al. 1964, Aggett, Cavanagh et al. 1980). Herein, we identify four patients with a clinical diagnosis of SDS from three unrelated families, who did not have mutations in SBDS but carried biallelic mutations in DNAJC21. Based on the clinical cases, comparable ribosomal function of DNAJC21 and SBDS, and recently identified association between DNAJC21 mutations and bone marrow failure, we propose that mutations in DNAJC21 cause SDS.

The second part of my work aimed to investigate a pathway through which an impaired

DNAJC21 gene causes SDS. Ours data shows that DNAJC21-knockdown led to significant decrease in cellular proliferation. Various studies have shown that ROS play an important role in mediating or amplifying death signals. In our study DNAJC21-knockdown led to increased ROS levels in HEK293T cells. In addition, by using H2O2 as a stressor to stimulate oxidative stress, we found that DNAJC21 KD cells were more sensitive to the stimulation and had significant increase in the levels of ROS compared to the WT or MOCK lines. These data lead us to conclude that DNAJC21 is not only required for maintaining baseline oxidation but plays a crucial role in reducing an excess of ROS in cells during oxidative stress. Loss of DNAJC21 protein level or function might result in increased ROS levels by various mechanisms. Indirect effect might be due to a reduction in translation of specific proteins by the DNAJC21-mutant ribosomes that balance oxidative stress in the cells, resulting in damage to mitochondrial respiratory increased oxidative stress. Loss of the chaperone activity of DNAJC21 may also affect critical location of proteins that are required for balancing oxidative stress.

Our third objective was to investigate the effects of elevated ROS in cell survival and proliferation in DNAJC21 mutant cells as done for SBDS (Dror and Freedman 1999, Dror and

Freedman 2001, Austin, Gupta et al. 2008, Rujkijyanont, Watanabe et al. 2008, Ambekar, Das et al. 2010, Sen, Wang et al. 2011). We demonstrated that DNAJC21-knockdown led to accelerated cell death with or without oxidative stress. However, we failed to identify the mechanism of death in these cells and are currently in the process of its further investigation. Moreover, we demonstrated that antioxidants can rescue the phenotype of growth inhibition and death in the knockdown cells. Herein, we conclude that DNAJC21-knockdown resulted in elevated levels of

ROS under normal conditions and a significantly increased ROS levels during an oxidative stress.

In summary, we showed for the first time that DNAJC21 is associated with SDS and that antioxidants improve cell growth by decreasing excess ROS levels. High levels of ROS in

DNAJC21-knockdown cells promote accelerated death and enhance sensitivity to oxidative

DNAJC21-knockdown cells in HEK293T cells. To test the application of our study to specific manifestations of SDS such as bone marrow failure, further studies need to be done using hematopoietic cells. Since ROS have been implicated in malignant transformation, a major complication in SDS, it is possible that increased ROS levels are involved in reduced bone marrow cell growth as well as for the progression of malignant myeloid clones observed in patients with SDS.

Chapter 6 Future directions

6.1 WES analysis

It is very important to continue to look for more families with SDS symptoms that lack known

SBDS or DNAJC21 mutations. After Cystic Fibrosis (CF), SDS in the second most common disease that cause pancreatic insufficiency, which results in a lot of problems in digestion and absorption of food. Also, children with SDS are under higher risk for MDS or leukemia.

Moreover, children with SDS have a multisystem disorder that can lead to a shortened lifespan.

By knowing which genes cause SDS and their mechanism we can target them and improve child`s life. Both genes discussed here are associated with the ribosome; therefore, it is important to look for possible alterations in genes involved in ribosomopathy.

Also, learning from this project we need to take in account and to check more carefully not only data from one type of files that we are provided from sequencing facility. We need to consider all data that we can get together in order to find and choose the right gene that is associated with the disease.

6.2 DNAJC21 gene

Our data demonstrates that cells expressing shRNAs against DNAJC21 have a decreased survival. Therefore, we are planning to investigate whether the observed cell death is mediated by apoptosis, necrosis or senescence. In addition, we observed a decreased survival of DNJC21- knockdown cells compared to the control upon stimulation with H2O2. To validate these results

we are planning to stain the cells with PI/annexin V and perform a Flow cytometry analysis. This assay is known to detect apoptotic/necrotic cells.

Next, we plan to further investigate the DNAJC21 functions in the cell, as it is a newly discovered gene for which not much data is yet available. Transcriptome analysis will shed light on the mechanism by which DNAJC21 can cause SDS. Our results demonstrate that the phenotype of DNAJC21 knockdown cells can be rescued by antioxidants, such as AA or

Catalase; therefore, it is important to further investigate whether oxidative pathways in the cell can be affected by the function of DNAJC21 protein. Ultimately, our goal is to discover novel potential therapeutic targets. The effectiveness of such novel therapies will be assessed by looking at their ability to rescue the hematopoietic phenotype or to delay its progression.

6.3 Primary SDS cell models

Until now we checked KD of DNAJC21 in the HEK293T cells model. However, it is very important to validate these results in patients’ cells. Our goal is to investigate whether the patient cells (e.g. lymphoblasts or fibroblasts cells) exhibit similar phenotype upon DNAJC21 knockdown as observed in HEK293T cells. We will replicate all the experiments that were done with HEK293T cells including assessment of cell proliferation, death and sensitivity to oxidative stress using cells derived from SDS patients. Moreover, a sample of cells will be included in the transcriptome analysis.

6.4 Role in Hematopoiesis

CD34 positive cells represent primitive bone marrow-derived hematopoietic stem and progenitor cells, as well as endothelial stem and progenitor cells. We hypothesize that CD34 positive cells derived from SDS patients have decrease hematopoietic colony formation efficiency. We will

investigate this hypothesis by performing colony forming assay with these cells. In addition, we will investigate whether rescuing DNAJC21 protein in patients will restore (fully or partially) the normal phenotype. This analysis will shed light on whether DNAJC21 is the only gene altered in these patients or whether additional genes are affected and work together with DNAJC21 to cause SDS.

A complementary strategy to study the role of DNAJC21 in hematopoiesis it to use the iPSc model.

In this model we will study the gene expression profiles of various genes during different stages of hematopoietic development. We aim to identify gene or genes expression alterations that are responsible for the onset of the hematopoietic defect during HE development. Follow-up studies will include gain-of-function or loss-of-function experiments of the genes identified, to determine the link between gene expression patterns and phenotype, and to help better understand the underlying mechanism responsible for the hematopoietic phenotype in SDS with deficiency in DNAJC21.Also, we will re-introduce DNAJC21 into the iPSCs, or knock-down

DNAJC21 in control iPSCs to assess impact of these genetic manipulations on generation of cells with HE potential as determined by flow cytometry and clonogenic assays. By re-introducing

DNAJC21, we expect to rescue the hematopoietic phenotype, and therefore see an increase in the

HE, EHP and mature blood cell populations in SDS iPSCs during definitive hematopoiesis. By knocking-down DNAJC21 in control iPSCs we expect to recapitulate the SDS hematopoietic phenotype, that we hypothesized would be observed with our iPSCs derived from patients with deficiency in DNAJC21.

Chapter 7 Additional work - Abstract

Inherited bone marrow failure syndromes (IBMFSs) are rare genetic diseases characterized by varying degrees of defective production of erythrocytes, granulocytes and platelets in the bone marrow, leading to anemia, thrombocytopenia, neutropenia or aplastic anemia. Due to limited efficacy or toxicity of currently available treatments and due to reduced life expectancy of patients with IBMFSs, novel therapeutic strategies are needed. Since the main morbidity and mortality are related to blood dyscrasia, studying hematopoiesis will help characterize the hematological phenotype. We hypothesized that the definitive wave of hematopoiesis is markedly impaired. In order to utilize iPSCs to study human IBMFSs disease we generated iPSCs from a patient with PARN deficiency. The PARN-deficient iPSCs showed a defect in definitive hematopoiesis with a marked reduction in the hemogenic endothelium population. Our study sheds a light on a mechanism of the onset and progression of the PARN deficiency hematopoietic phenotype and provides a platform for the development of novel therapeutic targets to improve patient care.

Chapter 7.1 Introduction

Development of tissues during embryogenesis and tissue homeostasis after their formation are tightly controlled and are delicately orchestrated by the expression both coding and non-coding

RNAs at specific time points and at certain levels (Pauli, Rinn et al. 2011). Deadenylation is a major mechanism that regulates RNA function and fate by controlling the turnover and abundance of mRNAs and the maturation of non-coding RNAs (e.g. small nucleolar RNAs, snoRNAs) (Berndt, Harnisch et al. 2012, Virtanen, Henriksson et al. 2013). Several deadenylases have been identified, such as Caf1(Astrom, Astrom et al. 1991, Brown, Tarun et al. 1996,

Korner, Wormington et al. 1998, Tucker, Valencia-Sanchez et al. 2001). Poly(A)-specific ribonuclease (PARN) is one of the major mammalian deadenylases (Virtanen, Henriksson et al.

2013) that trims single-stranded poly(A) tails of mRNAs (Astrom, Astrom et al. 1991, Astrom,

Astrom et al. 1992, Korner, Wormington et al. 1998, Martinez, Ren et al. 2000, Martinez, Ren et al. 2001), oligoadenylated tails of H/ACA box snoRNAs (Berndt et al., 2012) and microRNAs

(Yoda, Cifuentes et al. 2013). It plays a role in a variety of cellular processes (Rigby and

Rehwinkel 2015) including cell migration, stress response, and adhesion (Lee, Lee et al. 2012).

In addition, a recent paper shows an association between PARN and the target of EGR1 protein 1

(TOE1). TOE1 acts redundantly on some ncRNAs, most predominantly on small scaRNAs,

Cajal body-specific RNAs. scaRNAs is downregulated when PARN and TOE1 are both impaired leading to the defects in snRNA and pseudouridylation of small nuclear RNAs. Moreover, they function regularly in the biogenesis of TERC, telomerase RNA component, which has sequence motifs found in H/ACA box scaRNAs. Thus far, the animal model of PARN loss is a

Morpholino model which recapitulates hematopoietic defects seen in patients with biallelic in

PARN (Dhanraj et al, JMG 2015).

RNA biogenesis has revealed as a mechanism underlying several inherited diseases, such as

Diamond Blackfan anemia, Shwachman Diamond syndrome and dyskeratosis congenita

(Mochizuki, He et al. 2004, Flygare, Aspesi et al. 2007, Welting, Mattijssen et al. 2008, Finch,

Hilcenko et al. 2011) that include defects in rRNA maturation, snRNA maturation (Pellizzoni,

Yong et al. 2002), snoRNA transcription (Sahoo, del Gaudio et al. 2008) and mRNA maturation

(Crosby, Patel et al. 2010). Moreover, dysregulation of mRNA adenylation in mice (Stumpo,

Broxmeyer et al. 2009) and in zebrafish (Bolli, Payne et al. 2011) leads to an impaired hematopoiesis. Therefore, it is reasonable to hypothesize, that gene expression regulation by polyadenylation or deadenylation is very important for promoting proper hematopoietic cell expansion and differentiation therefore preventing bone marrow failure.

We are currently continuing our previous work where we identified four patients with developmental delay or mental illness from 3 different families, who carried large monoallelic deletions in PARN. One of the patients had severe bone marrow failure and neurological manifestations and had a biallelic mutation in PARN. We reported the results of genetic interrogation of these patients, the consequences of biallelic PARN mutations on RNA metabolic pathways and demonstrated conserved function of PARN in hematopoiesis using zebrafish model

(Dhanraj, Gunja et al. 2015). These investigations establish a new paradigm for IBMFS disease.

To show that PARN is important for hematopoiesis we utilized an induced pluripotent stem cells

(iPSC) model. iPSCs were first reprogrammed from mouse and human somatic cells to pluripotency by overexpressing a combination of pluripotency transcription factors OCT4 (O),

SOX2 (S), KLF4 (K), and C-MYC (M) (Takahashi and Yamanaka 2006, Takahashi, Tanabe et al. 2007). Other combinations of reprogramming factors are also sufficient, such as OCT4,

SOX2, NANOG, and LIN28 (Yu, Vodyanik et al. 2007), although activation of OCT4 or SOX2

expression is considered essential in reprogramming (Huangfu, Osafune et al. 2008, Marson,

Foreman et al. 2008, Chen, Liu et al. 2011, Buganim, Faddah et al. 2012). “OSK” form a part of the core transcriptional network of pluripotency, which auto-regulate and co-target regulatory regions of pluripotency genes (Nichols, Zevnik et al. 1998, Boyer, Lee et al. 2005, Loh, Wu et al.

2006, Masui, Nakatake et al. 2007, Kim, Chu et al. 2008). Unlike OSK, C-MYC is not a part of the core transcriptional network. It enhances reprogramming efficiency potentially by promoting chromatin remodeling and cell proliferation (Kim, Chu et al. 2008, Wernig, Meissner et al.

2008). During reprogramming of cells, OCT4, KLF4, and C-MYC repress the transcription of somatic genes, while KLF4 is also essential in initiating mesenchymal-to epithelial transition

(Sridharan, Tchieu et al. 2009, Chen, Liu et al. 2011, Polo, Anderssen et al. 2012, Tiemann,

Marthaler et al. 2014). The first generation of iPSCs was reprogrammed using delivery of reprogramming factors by retroviral infection (Takahashi and Yamanaka 2006, Takahashi,

Tanabe et al. 2007). Integrative delivery methods such as retroviruses are dependent on genomic integration of reprogramming factors, which has a risk of insertional mutagenesis and a partial silencing, leading to reactivation of reprogramming factors and to a repression of differentiation

(Toivonen, Ojala et al. 2013, Brouwer, Zhou et al. 2016). Moreover, the transcriptional profile of hESCs is more similar to non-integrated hiPSCs than to the integrated hiPSCs (Soldner,

Hockemeyer et al. 2009). Hence, non-integrative delivery methods are favored, such as Sendai

RNA virus that introduces reprogramming factors in the form of RNA without integrating into the host genome (Fusaki, Ban et al. 2009). Introduction of the reprogramming factors into somatic cells induces a series of transcriptional and epigenetic events that reverse the transcriptome and epigenome to a pluripotent stage (Maherali, Sridharan et al. 2007, Mikkelsen,

Hanna et al. 2008, Smith, Nachman et al. 2010, Koche, Smith et al. 2011, Lister, Pelizzola et al.

2011, Nishino, Toyoda et al. 2011, Buganim, Faddah et al. 2012, Polo, Anderssen et al. 2012,

Zunder, Lujan et al. 2015). During early reprogramming state, quick histone methylation modifies the regulatory regions of pluripotency and developmental genes to an euchromatic state

(lightly packed form of chromatin) and converts somatic genes to a heterochromatic state (tightly packed form of DNA). All these modifications result in transcriptional changes, reduced cell size, increased proliferation, and loss of somatic function. During the intermediate stage, partly reprogrammed population either returns to the original somatic cell type or continues to differentiate to iPSC stage, progressing through mesenchymal-to- epithelial transition. During late reprogramming, DNA demethylation of regulatory regions of pluripotency genes establishes stable iPSCs. The result of complete reprogramming are iPSCs similar to embryonic stem cells

(ESCs). hiPSC characteristics

Before the discovery of iPSCs, the main source of pluripotent stem cells were the ESCs isolated from the inner cell mass of the blastocysts (Evans and Kaufman 1981, Martin 1981, Thomson,

Itskovitz-Eldor et al. 1998). Both iPSCs and ESCs are defined by their ability to differentiate into almost any types of cells as well as an ability to self-renew (Zhao, Li et al. 2009, Tiscornia,

Vivas et al. 2011, Brouwer, Zhou et al. 2016). The morphology of iPSCs is a dense colony of cells with well-defined borders. Also, iPSCs are characterized by high alkaline phosphatase activity, an expression of pluripotency markers and an ability to differentiate into cells of all three germ layers. Epigenetic profiles of iPSCs reveal evaluated DNA demethylation in pluripotency genes as well as methylation of somatic genes that are necessary for complete reprogramming.

Epigenetic and transcriptional characterizations of iPSCs are similar to ESCs, including mRNA and miRNA profiles, chromosome organization and DNA and histone methylation patterns. 91

(Chin, Mason et al. 2009, Doi, Park et al. 2009, Marchetto, Yeo et al. 2009, Chin, Pellegrini et al.

2010, Guenther, Frampton et al. 2010, Stadtfeld, Apostolou et al. 2010, Bock, Kiskinis et al.

2011, Lister, Pelizzola et al. 2011, Ohi, Qin et al. 2011, Mills, Wang et al. 2013, Shao, Koch et al. 2013, Choi, Lee et al. 2015, Krijger, Di Stefano et al. 2016). However, iPSC-specific transcriptional signatures and differentially methylated regions capable of transmission through differentiation were reported. Moreover, by comparison of genetically matched iPSCs with

ESCs, it was demonstrated that previously observed transcriptional differences were mainly a result of a genetic background. However, iPSC inherent variability can be found among iPSC lines derived from the same donor and is likely as result of varying epigenetic reprogramming events (Osafune, Caron et al. 2008, Chen, Egli et al. 2009, Hu, Weick et al. 2010, Bock, Kiskinis et al. 2011, Lister, Pelizzola et al. 2011, Mills, Wang et al. 2013, Wu, Liu et al. 2014). To minimize differentiation variability, standardized differentiation protocols should be used and stringent quality screening for complete reprogramming is required. The criteria used for the quality screening include histone deposition, differentiation potential, and CD30 expression

(Bock, Kiskinis et al. 2011, Boulting, Kiskinis et al. 2011, Abujarour, Valamehr et al. 2013, Wu,

Liu et al. 2014). CD30 marker was found to be a pluripotency marker specifically expressed by hiPSCs with high expression of common pluripotency markers and distinguished undifferentiated or partially differentiated hiPSCs from differentiated cells (Abujarour, Valamehr et al. 2013, Friedel, Jung-Klawitter et al. 2016).

Moreover, due to incomplete silencing and incomplete DNA methylation of somatic genes, iPSCs were shown to preserve residual somatic cell transcriptional and epigenetic memory, which twist the differentiation potential in partiality the original somatic cell type (Ghosh,

Wilson et al. 2010, Bar-Nur, Russ et al. 2011, Lister, Pelizzola et al. 2011, Ohi, Qin et al. 2011,

Hiler, Chen et al. 2015). Upon continuous passage, somatic transcriptional memory and epigenetic variations were shown to either persist or become lost and barely contribute to transcriptional variations (Polo, Liu et al. 2010, Kim, Zhao et al. 2011, Rouhani, Kumasaka et al.

2014). Nevertheless, when iPSCs that were reprogrammed from different genetically-matched somatic cell types were compared, differentiation and transcriptional variation in iPSCs was found to be mainly contributed by genetic and epigenetic differences (Kajiwara, Aoi et al. 2012,

Shao, Koch et al. 2013, Rouhani, Kumasaka et al. 2014). Thus, reprogramming from the somatic cell type closest to the differentiated cell of interest should be preferred. The last criterion to consider when working with the iPSC model is genomic instability. It was shown that frequency of copy number variations, as well as recurrent and non-recurrent karyotypic aberrations were higher in both iPSCs and ESCs compared to cultured cell lines (Hussein, Batada et al. 2011,

Laurent, Ulitsky et al. 2011, Martins-Taylor, Nisler et al. 2011, Taapken, Nisler et al. 2011,

Martins-Taylor and Xu 2012). Genomic deflections can be acquired not only during integrative reprograming but also during culture adaptation, differentiation, and prolonged culture. All this can cause altered gene expression, differentiation potential and proliferation rate (Mayshar, Ben-

David et al. 2010, Hussein, Batada et al. 2011, Laurent, Ulitsky et al. 2011, Martins-Taylor,

Nisler et al. 2011, Taapken, Nisler et al. 2011, Cheng, Hansen et al. 2012, Mills, Wang et al.

2013). Thus, to minimize the effects of genomic instability we need to use non-integrative delivery of reprogramming factors and perform a regular karyotypic monitoring.

Despite the described challenges in using the iPSCs model, it was successfully utilized to recapitulate various Inherited Bone Marrow Failure syndromes as listed below (Jung, Dunbar et al. 2015).

1) Fanconi anemia - hiPSCs required functional DNA repair and an absence of oxidative stress during reprogramming. hiPSCs were able to recapitulate the erythroid and myeloid defects identifying earlier defects in the hemangioblasts as well as in hematopoietic progenitors.

2) Dyskeratosis congenital - hiPSCs were able to recapitulate the impairment in hematopoietic differentiation and telomere maintenance in dyskeratosis congenita.

3)Familial platelet disorder - hiPSCs were able to recapitulate the megakaryocytic defect and a genomic instability.

4) Diamond-Blackfan anemia - hiPSCs were able to recapitulate the impairment in erythroid differentiation and 40S biogenesis, while identifying earlier defects in multipotent hematopoietic progenitors.

5) Congenital amegakaryocytic thrombocytopenia - hiPSCs were able to recapitulate the impairment in megakaryocytic differentiation and in TPO/MPL signaling.

To summarize, a hematopoietic differentiation of hiPSCs is a good platform for studying all the stages of formation of hematopoietic cells, especially the stage of multipotent progenitors as this stage contains scarce number of cells.

Chapter 7.2 Methods

7.2.1 Cell Culture hiPSC reprogramming. Human primary skin fibroblast cells from normal subjects (CCRM25 and CCRM 5R) and PARN patient (P319) were collected with written and informed consent by the Canadian Inherited Marrow Failure Registry. Skin fibroblasts were generated and reprogrammed to hiPSC using Sendai viral expression of OCT4, SOX2, KLF4, and C-MYC genes by CCRM. hiPSC culture. The following method of maintaining hiPSCs cultured on MEFs was adopted from the laboratory of Dr. Gordon Keller. The materials used in the thawing, maintenance, passaging, and freezing of hiPSC cultured on MEFs are listed in methods. hiPSCs were co- cultured with MEFs in growth medium at 37°C in a humidified 5% CO2 incubator with daily medium change. hiPSCs were passaged in aggregates using 0.25% trypsin at room temperature for up to 1 minute. The digestion was stopped with stop medium and aggregates of hiPSCs were lifted gently using a cell scraper. Aggregates were collected with wash medium and pelleted by centrifugation at 800rpm for 3 minutes at room temperature. Next aggregates were plated onto new 6-well MEF plates in a 1:2 to 1:12 ratio. MEFs were extracted at embryonic day 12.5 and treated with mitomycin C by the SickKids Embryonic Stem Cell Facility and plated in our lab.

To improve viability, ROCK inhibitor and additional 10ng/mL bFGF were supplemented in the growth medium after passaging or thawing. hiPSCs were also cultured on matrigel according to manufacturer’s instructions (Stemcell Technologies 05850) for extracting protein, DNA or RNA.

OP9-DL1 Culture. OP9-DL1 cells were gifted by the laboratory of Dr. Eyal Grunebaum. The following method of expanding OP9-DL1 cells was adopted from the laboratory of Dr. Gordon

Keller. The materials used in the thawing, maintenance, passaging, and freezing of OP9-DL1 are listed in methods. OP9-DL1 cells were cultured on gelatin-coated T-75 flasks in OP9-DL1 medium with medium changed every 3-4 days. Cells were passaged in single cells using 0.05% trypsin at 37°C for 3-5 minutes. The digestion was stopped using FBS stop medium. Cells were collected with αMEM and pelleted by centrifugation at 1200rpm for 5 minutes at room temperature. Cells were resuspended in OP9-DL1 medium and irradiated at 3000rads using the

Gammacell® 40 Exactor (Best Theratronics) at Sickkids Laboratory Animal Services. Cells were frozen in OP9-DL1 freezing medium at a density of 106 cells/cryovial. In preparation for hematopoietic differentiation at T9, irradiated OP9-DL1 cells were plated at a density of 106 cells per gelatin-coated 24-well plate 48 hours before using.

Table 5: Reagents used in the maintenance of OP9-DL1 culture. Catalogue Final Reagents Components Company Number Concentration Gelatin from Porcine Skin Type A Sigma-Aldrich G1890-100G 0.1g/100mL Gelatin Sterile Water Wisent 809-115-CL 100% OP9-DL1 FBS Gibco 12483-020 20% Medium Minimum Essential Medium α (αMEM) Gibco 12561-056 80% PBS Phosphate Buffered Saline Wisent 311-010-CL OP9-DL1 OP9-DL1 Medium 50% Freezing FBS Gibco 12483-020 40% Medium DMSO Sigma-Aldrich D8418 10% hiPSC lentiviral transduction. To transduce hiPSCs, hiPSCs were digested with collagenase B at 37°C for 20 minutes and 0.25% trypsin at 37°C for 3 minutes. The digestion was stopped with

FBS stop medium and hiPSCs were resuspended into single cells. Single cells were collected with wash medium and pelleted by centrifugation at 1200RPM for 5 minutes at room temperature. Single cells were filtered through 35µm cell strainer and pelleted again. Single cells were plated onto new MEF plates at a density of 104cells/well and infected with lentiviral

particles at a multiplicity of infection of 100 and with 8µg/mL protamine sulfate for 24 hours.

Protamine sulfate and lentivirus packaged with the rescue plasmid pLVX.SIN.EF1a.PARN-

HA.IRES.ZsGreen or mock plasmid pLVX.SIN.EF1a.IRES.ZsGreen (generated by Dr.

Chetankumar Tailor in our lab). hiPSCs were cultured in growth medium supplemented with additional 10ng/mL bFGF and 10µM ROCK inhibitor for 10 days with daily medium change.

After 10 days, hiPSCs were cultured in growth medium supplemented with additional 10ng/mL bFGF with daily medium change. The clones were picked up and cultured separately.

7.2.2 Characterization of hiPSCs hiPSC characterization. CCRM assessed fidelity to parental identity by polymerase chain reaction (PCR) of 9 regions of short tandem repeats (STR; CSF1PO, D3S1358 or D21S11,

D13S317, D5S818, D7S820, D16S539, THOI, TPOX, and vWA) and gel electrophoresis, assessed gender by PCR of Amelogenin and gel electrophoresis, assessed RNA expression of pluripotency markers by quantitative reverse transcription-PCR (qRT-PCR), assessed protein expression of pluripotent surface markers by flow cytometry or immunocytochemistry (ICC), assessed germ layer differentiation by EB formation and qRT-PCR, assessed Mycoplasma contamination, and assessed chromosomal abnormality by G-band karyotyping. Further

Mycoplasma testing of hiPSCs cultured on matrigel was performed by SickKids Molecular

Biology Laboratory, and further G-band karyotyping of hiPSCs cultured on MEFs was performed by TCAG.

Western blotting. Cells were harvested and lysed by RIPA lysis buffer (Santa Cruz

Biotechnology sc-24948) according to manufacturer’s instructions. Protein concentration was measured using protein assay reagent concentrate (Bio-Rad 5000006) and BioPhotometer

Spectrophotometer (Eppendorf) according to manufacturer’s instructions. Proteins were resolved

using 7.5% polyacrylamide gel electrophoresis (Mini-PROTEAN® 3 Cell 165-3301) according to manufacturer’s instructions. Western blot transfer (Mini Trans-Blot® Electrophoretic Transfer

Cell, Bio-Rad 170-3930) was performed according to manufacturer’s instructions. The membrane was incubated with blocking solution (TBST/5% skim milk) at room temperature for

1 hour on a rocking platform and washed with TBST. Antibodies were stained according to manufacturer’s instructions. Proteins were detected using ECL Western blotting detection reagent (Amersham Biosciences RPN2106) and Kodak X-OMAT film or Hyperfilm ECL

(8”x10”, Amersham Biosciences RPN2114K) according to manufacturer’s instructions.

7.2.3 Hematopoietic Differentiation

The following method of definitive hematopoietic differentiation of hiPSCs cultured on MEFs was adopted from the laboratory of Dr. Gordon Keller. The materials used in feeder depletion,

EB formation, EB digestion, sorting, OP9-DL1 co-culture, and colony forming unit assay are listed.

Table 6: Reagents used in the hematopoietic differentiation of hiPSCs cultured on MEFs. Catalogue Final Reagents Components Company Number Concentration Corning® Matrigel® Matrix Growth VWR CACB354230 50% Factor Reduced (GFR) Matrigel- Iscove's Modified Dulbecco's Medium GFR Gibco 21056-023 49% (IMDM) Penicillin-Streptomycin, 5000U/mL Gibco 15070-063 1% 0.0625% 0.25% Trypsin Wisent 325-043-EL 25% Trypsin PBS Wisent 311-010-CL 75% Low CorningTM Ultra-Low Attachment Thermo Fisher Attachment 07-200-601 Plates 6-well Scientific Plates StemPro®-34 Serum Free Medium Gibco 10639-011 98% (SFM) (1X) (SP34) Penicillin-Streptomycin, 5000U/mL Gibco 15070-063 1% Basal L-Glutamine, 100× Cellgro 25-005-CI 1% Medium L-Ascorbic Acid Sigma-Aldrich 1-4544 50µg/mL Transferrin Roche 10652202001 150µg/mL 1-Thioglycerol Sigma-Aldrich M-6145 0.4mM Basal Medium T0 Medium Recombinant Human Bone R&D Systems 314-BP-050 10ng/mL Morphogenetic Protein 4 (BMP-4) 98

T0 Medium T1 Medium bFGF R&D Systems 234-FSE-025 5ng/mL T1.75 T1 Medium Medium SB 431542 Hydrate Sigma-Aldrich S4317-5MG 6µM Basal Medium bFGF R&D Systems 234-FSE-025 5ng/mL Recombinant Human Vascular Endothelial Growth Factor (VEGF) R&D Systems 293-Ve-050 10ng/mL T4 Medium 165 Recombinant Human Interleukin 6 R&D Systems 206-IL-050 10ng/mL (IL-6) Recombinant Human Interleukin 11 R&D Systems 218-IL-025 5ng/mL (IL-11) T4 Medium Recombinant Human Stem Cell Factor R&D Systems 255-SC-200 50ng/mL T6 Medium (SCF) Recombinant Human Erythropoietin R&D Systems 287-TC-500 2U/mL (EPO) Basal Medium IL-6 R&D Systems 206-IL-050 10ng/mL IL-11 R&D Systems 218-IL-025 5ng/mL SCF R&D Systems 255-SC-200 100ng/mL EPO R&D Systems 287-TC-500 2U/mL Recombinant Human Thrombopoietin R&D Systems 288-TP-200 50ng/mL T8 Medium (TPO) Recombinant Human Fms-related R&D Systems 308-FKN-025 10ng/mL tyrosine kinase 3 ligand (Flt-3L) Recombinant Human Interleukin 3 R&D Systems 203-IL-050 50ng/mL (IL-3) Recombinant Human Insulin-like R&D Systems 291-G1-200 25ng/mL Growth Factor I (IGF-I) Collagenase from Clostridium Sigma-Aldrich C0130-1G 2mg/mL histolyticum Type I Collagenase FBS Gibco 12483-020 20% I PBS Wisent 311-010-CL 80% EMD DNaseI 260913-10MU 10µg/mL Millipore FBS Gibco 12483-020 10% Staining IMDM Gibco 21056-023 90% Medium EMD DNaseI 260913-10MU 10µg/mL Millipore Phycoerythrin (PE) Mouse Anti- BD 560199 5% Human CD43, Clone 1G10 (PE-CD43) Pharmingen PE-Cyanine 7 (PE-Cy7) Anti-Human Affymetrix 25-0349-42 1% T9 Staining CD34, Clone 4H11 (PE-Cy7-CD34) eBioscience Reaction Thermo Fisher Propidium Iodide (PI) P1304MP 1µg/mL Scientific FBS Gibco 12483-020 10% Staining Medium FBS Gibco 12483-020 1.5% Sorting IMDM Gibco 21056-023 98.5% Medium EMD DNaseI 260913-10MU 10µg/mL Millipore HyCloneTM FBS (Canada), GE Healthcare SH30396.03 20% T9+0 to 1 Characterized (HyClone FBS) Medium αMEM Gibco 12561-056 80% BMP-4 R&D Systems 314-BP-050 10ng/mL 99

IL-11 R&D Systems 218-IL-025 5ng/mL SCF R&D Systems 255-SC-200 100ng/mL TPO R&D Systems 288-TP-200 30ng/mL Flt-3L R&D Systems 308-FKN-025 20ng/mL T9+3 to 5 T9+0 to 1 Medium Medium VEGF R&D Systems 293-Ve-050 5ng/mL Methylcellul HyClone FBS GE Healthcare SH30396.03 2% ose IMDM Gibco 21056-023 98% Medium Stemcell MethocultTM H4034 Optimum 04034 Technologies Methocult IL-6 R&D Systems 206-IL-050 10ng/mL Flt-3L R&D Systems 308-FKN-025 10ng/mL

Feeder depletion. Each 6-well plate was coated with a thin layer of cold matrigel-GFR for 30 minutes on ice. After excess matrigel-GFR was removed, the matrigel-GFR-coated plates were incubated at 37°C for at least 4 hours. hiPSCs cultured on MEFs were passaged and plated onto matrigel-GFR-coated plates in a 1:2 ratio. hiPSCs were cultured in growth medium supplemented with 0.0625% matrigel-GFR at 37°C in a humidified 5% CO2 incubator for 2 days with daily medium change.

Hematopoietic differentiation. Feeder depleted hiPSCs were passaged in aggregates using collagenase B at 37°C for 5 minutes and 0.625% trypsin at room temperature for up to 1 minute.

Aggregates were plated onto low attachment plates at a density of 106 cells/well in T0 medium.

The time when the aggregates were placed in a humidified 37°C 5% O2/5% CO2/90% N2 hypoxic incubator was designated as time-point zero (T0). After 24 hours, T1 medium was added and EBs were returned to hypoxia at T1. After 17 hours, EBs were settled for 30 minutes at 37°C and cultured in T1.75 medium. EBs were returned to hypoxia at T1.75, 6 hours ahead of T2.

After 53 hours, EBs were pelleted by centrifugation at 60×g for 3 minutes at room temperature and settled for 30 minutes at 37°C. EBs were cultured in T4 medium and returned to hypoxia at

T4. After 47 hours, EBs were pelleted, settled, cultured in T6 medium, and returned to hypoxia at T6. After 47 hours, EBs were pelleted, settled, cultured in T8 medium, placed in a humidified

37°C 5% CO2 incubator at T8, and cultured in normoxia. 100

Picture 3: Hematopoetic differentiation schema The schema is based on a protocol provided to us from Dr. Gordon Keller, University of Toronto and can be found in the following article: Kennedy et al. 2012)

T9 digestion. Following 20 hours in normoxia, EBs were collected with IMDM and pelleted by centrifugation at 150×g for 3 minutes at room temperature. EBs were digested with 0.25% trypsin at 37°C for 5 minutes and the digestion was stopped with FBS stop medium. EBs were erupted by syringing with a 20G needle 6 times and pelleted by centrifugation at 500×g for 5 minutes at room temperature. EBs were digested with collagenase I at 37°C for 1 hour and further erupted by syringing with a 20G needle 6 times into single cells. Staining medium was added and single cells were pelleted by centrifugation at 500×g for 5 minutes at 4°C. Single cells were filtered through a 35µm cell strainer and pelleted again. Single cells were resuspended in

101

cold staining medium in preparation for staining reactions. The figure and differentiation schema were adapted from Dr. Gordon Keller Lab (Ditadi, Sturgeon et al. 2017).

Preparation of cells for T9 flow cytometry and sorting. T9 single cells were added to T9 staining reactions, including PI only control, fluorescence-minus-one (FMO) controls, and sorting samples and stained on ice away from light for 30 minutes. Staining medium was added, and stained cells were pelleted by centrifugation at 500×g for 5 minutes at 4°C. The controls were resuspended in cold staining medium and sorting samples were resuspended in cold sorting medium at densities.. The controls were analyzed, and the sorting samples were sorted using the

Moflo® AstriosTM or MofloTM XDP cell sorter (Beckman Coulter) at the Sickkids Flow

Cytometry Facility. Each CD34hi/CD43- population was collected in FBS.

Table 7: Fluorophores and cell density of PI only control, FMO controls, and sorting sample of T9 staining reactions. FMOs PI only –PE-CD43 –PE-Cy7-CD34 Sorting Sample PE-CD43 - - + + PE-Cy7-CD34 - + - + PI + + + + Cells (per 100µL) 5×104 5×104 5×104 106

OP9-DL1 co-culture. The CD34hi/CD43- sorted cells were pelleted by centrifugation at 500×g for 5 minutes at 4°C. Sorted cells were cultured in T9+0 to 1 medium at a density of 2×104 cells per well of irradiated OP9-DL1-coated 24-well plate in a humidified 37°C 5% CO2 incubator in normoxia at T9+0. After 24 hours, T9+0 to 1 medium was added and the culture was returned to normoxia at T9+1. After 48 hours, the cells were cultured in T9+3 to 5 medium and returned to normoxia at T9+3. After 48 hours, half of the medium was changed with T9+3 to 5 medium and the culture was returned to normoxia at T9+5.

102

CFU assay. After 46 hours in normoxia, the medium of the culture was filtered through a 35µm cell strainer. Cells were digested with 0.25% trypsin at room temperature for 5 minutes and the digestion was stopped with FBS stop medium. Cells were resuspended to single cells and filtered through 35µm cell strainer. Single cells were pelleted by centrifugation at 500×g for 5 minutes at room temperature. Single cells were plated onto Methocult H4034 according to manufacturer’s instructions. Duplicates of 4×104 cells per 35mm gridded dish were cultured in a humidified

37°C 5% CO2 incubator at T9+7+0. After 14 days, CFU-GEMM, CFU-GM, and BFU-E were scored at T9+7+14. A colony larger than ¼ of a grid (0.5mm × 0.5mm) was considered large, and a colony equal to or smaller than ¼ of a grid was considered small.

7.2.4 Statistical Analyses

Unpaired data was analyzed using unpaired two-tailed t-test. Paired data with equal variance was analyzed using unpaired two-tailed t-test and data with unequal variance was analyzed by unpaired two-tailed t-test with Welch’s correction. Significance was defined as p-value (p) less than 0.05.

103

Chapter 7.3 Results

7.3.1 Characterization of hiPSCs

Primary skin fibroblast cells were obtained from two non-related normal subjects and one PARN deficiency patient and reprogrammed to hiPSCs using Sendai viral expression of OCT4, SOX2,

KLF4, and c-MYC by CCRM. Two independent hiPSC lines from one individual were used in this study) and one line from each normal control. Western blotting showed reduced PARN protein expression in patient lines compared to normal hiPSCs. These results suggest that hiPSC lines were successfully established from the patient.

Table 8: hIPSC lines used in the present study PARN hiPSC Lines Diagnosis Mutations Healthy CCRM5 - control 1 healthy CCRM25 - control 2 R349W + p319A, IBMF deletion of P319D exon 16-17

PARN – 78KDa

Vinculin – 128KDa

Figure 22: PARN protein expression in hiPSCs. Western blot with anti-PARN and anti-Vinculin antibodies as control confirmed reduced PARN expression in PAGE-separated proteins of whole cell extracts of patient compared to control hiPSCs.

104

7.3.2 Colony Forming Assay

CFU assays of PARN hPSCs have shown reduced total colony count but no differences in CFU-

GEMM as reduction could not be seen in both control and patient lines. To determine if our

PARN hiPSCs were able to recapitulate the PARN hematopoietic defects of reduced myeloid colony count, hiPSCs were differentiated towards myeloid hematopoietic lineage by CFU assay. hiPSCs were feeder depleted for 2 days, subjected to definitive hematopoietic differentiation in the form of EBs for 9 days, sorted for CD34+/CD43- populations as it can be seen in figure 23A-

B, co-cultured on OP9-DL1 for 7 days, cultured in methylcellulose for 14 days, and then scored for colonies of CFU-GEMM, CFU-GM, and BFU-E at T9+7+14 (Figure 24A-B). In addition, we distinguished between small and large colonies to see if there any differences in the maturation rate of the cells. A colony larger than ¼ of a grid (0.5mm × 0.5mm) was considered to be large colony, while a colony equal to or smaller than ¼ of a grid was considered to be small. The size of the colony reflected the proliferative potential of each plated cell, while the number of colonies reflected the hematopoietic multipotency of plated cells. To note, CFU-GM in this study also included CFU-G and CFU-M, as the differences among the three were difficult to distinguish. As illustrated in figure 24A-B, while there was no significant reduction in CFU-

GEMM, there was a significant reduction in the number of total colonies, CFU-GM, and BFU-E.

Moreover, there was a significant reduction in the number of small (p = 0.003) and large (p =

0.002) total colonies, small (p = 0.03) and large (p = 0.002) CFU-GM, and small BFU-E (p =

0.02), although there were no large BFU-E colonies produced in all lines and all repeats that were done.

105

106

Figure 23: Analysis of hemogenic endothelium containing cells within the CD34+/CD43- population in T9 embryoid bodies derived from normal iPSC by flow cytometry. At day T9 of differentiation, embryoid bodies derived from normal iPSc CCRM5R (A) and patient line p319D (B) were analyzed for both markers CD34 and CD43.

107

A Colony assay CFU-GEMM 100 CFU-GM * 90 * BFU-E 80 70 60 50 40 30 20 avrg # # avrg ofcolonies counted 10 0 p319A p319D CCRM5R CCRM25 Lines

B Analysis of small and large colonies p319A

70 p319D

60 * CCRM5R

50 N551C

40 * 30

avrof # colonies 20

0 CFU-GEMM CFU-GEMM CFU-GM CFU-GM BFU-E large BFU-E small large small large small type of cells

Figure 24: Clonogenic potential of PARN-deficient hiPSCs. Normal (CCRM25 and CCRM5R) and PARN (P319A and P319D) hiPSCs were feeder depleted for 2 days, subjected to hematopoietic differentiation for 9 days, sorted for CD34+/CD43- population, co- cultured on OP9-DL1 for 7 days, plated into methylcellulose at a density of 40000 cells/dish, cultured in methylcellulose for 14 days, and scored for large (greater than 0.5mm × 0.5mm) and small (equal to or less than 0.5mm × 0.5mm) colonies of CFU-GEMM, CFU-GM, and BFU-E at T9+7+14. Number of colonies from patient were significantly reduced. Bars represent mean count of large and small colonies, error bars represent standard deviation, and p-values (*) comparing total numbers of colonies derived from normal and PARN hiPSCs are P<0.05.

108

7.3.3 CD34+/CD43- Population

To determine if CD34+/CD43- homorganic and vascular endothelial populations were reduced in patient lines, populations from stage T9 (EBs stage) were analyzed. For this purpose, we performed feeder depletion of hiPSCs for 2 days and then subjected cells to definitive hematopoietic differentiation in the form of EBs for 9 days as it was described in methods. The last stage was to sort the cells for PE-Cy7-CD34+/PE-CD43- population using FACS at T9 to be further cultured on OP9-DL1. The CD34+/CD43- cell population, which contains homorganic and vascular endothelial cells, was significantly lower between PARN deficient lines and normal controls in T9 (figure 25), EBs stage. These results suggest an important role of PARN in early hematopoietic differentiation.

CD34+/CD43- population 20000 15000 10000

5000 * * avrof # cells 0 CCRM5R CCRM25 P319A P319D Lines

Figure 25: Hemogenic endothelium cell population at day 9 of culturing was decreased compared to controls. Normal CCRM25/CCRM5R and PARN P319A/D hiPSCs were feeder depleted for 2 days and, subjected to hematopoietic differentiation for 9 days, sorted for CD34+/CD43- population by FACS and analyzed for CD34+/CD43- population by Flowjo V10. CD34+/CD43- population in patient lines were dramatically decreased comparing to normal line *P<0.05, the experiment was done 6 time.

7.3.4 Lentiviral Transduction of PARN hiPSCs

Next, we tried to establish rescued PARN hiPSC lines by lentiviral transduction of P319A and

P319D hiPSCs using the pLVX.SIN.EF1a.PARN-HA.IRES.ZsGreen plasmid for PARN rescue

109

and pLVX.SIN.EF1a.IRES.ZsGreen plasmid for mock control. However, PARN deficient hiPSC lines lost GFP expression during culture compared to mock that had a stable expression of GFP.

We hypothesized that the loss of GFP expression is due to PARN nuclease activity that can affect expression of the plasmid by cutting the plasmid. We have previously encountered a similar problem while trying to establish a rescued line of skin fibroblasts.

Figure 26: hiPSC colonies transduced with a lentivurs carrying wild type PARN. hiPSCs were transduced with pLVX.SIN.EF1a.IRES.ZSGREEN / pLVX.SIN.EF1a.PARN- HA.IRES.ZSGREEN with 8ug/ml protamine sulfate. Left picture represent transduced cells under fluorescent light. Right picture taken under normal light.

110

Chapter 7.4 Discussion and future directions

Our previous study demonstrated that severe deficiency of PARN in patient cells was associated with loss of specific H/ACA box RNAs, snoRNA/scaRNAs and TERC. This loss was accompanied by two snoRNA-dependent and TERC-dependent pathways with essential roles in normal hematopoiesis, ribosome profile and telomere length. The important role of PARN in normal hematopoiesis was confirmed by an observation, that knocking down PARN resulted in deficiency in blood cells from hematopoietic stem cells/progenitors and in zebrafish. Using in vivo PARN-deficient zebrafish model we demonstrated that re-introduction of PARN in PARN- knockdown zebrafish embryos rescued the phenotype. Unfortunately, we were unable to perform

PARN rescue experiments in human PARN-deficient fibroblasts and hiPSCs due to technical difficulties describe above (impaired growth of cells and loss of GFP expression).

Our lab has previously demonstrated that hematopoietic defects are present in earlier hematopoietic progenitors in the IBMFS, Shwachman-Diamond syndrome. Moreover, patient bone marrow had reduced CD34+ hematopoietic stem and progenitor cells (Dror and Freedman

1999) and colony assays using cells derived from patient bone marrow as well as hPSCs have shown reduced colony forming potential (Saunders, Gall et al. 1979, Suda, Mizoguchi et al.

1982, Dror and Freedman 1999, Kuijpers, Alders et al. 2005, Tulpule, Kelley et al. 2013).

Experiments that aimed to induce differentiation towards definitive hematopoiesis in PARN hiPSC revealed significantly reduced potential to form hematopoietic colonies. This reduction was observed in cells derived from patients that had PARN deficiency as well as developmental delay or mental illness and from patients that carried large monoallelic PARN deletions. These data suggest that biallelic PARN mutations in patients result in a hematopoietic phenotype. In

111

addition, we identified a reduction in homorganic and vascular endothelial populations in cells that are characterized by biallelic PARN mutations.

More studies are required to decipher the mechanism underlying reduced colony formation.

Strategies such as RNA-seq at different differentiation time points may reveal potentially disrupted pathways. Such analysis could shed light on the role of PARN in hematopoiesis. In addition, alternative strategies should be implemented to establish rescue line and define the role of PARN in the observed phenotype.

112

Bibliography

Abujarour, R., B. Valamehr, M. Robinson, B. Rezner, F. Vranceanu and P. Flynn (2013). "Optimized surface markers for the prospective isolation of high-quality hiPSCs using flow cytometry selection." Sci Rep 3: 1179.

Aggett, P. J., N. P. Cavanagh, D. J. Matthew, J. R. Pincott, J. Sutcliffe and J. T. Harries (1980). "Shwachman's syndrome. A review of 21 cases." Arch Dis Child 55(5): 331-347.

Aggett, P. J., J. T. Harries, B. A. Harvey and J. F. Soothill (1979). "An inherited defect of neutrophil mobility in Shwachman syndrome." J Pediatr 94(3): 391-394.

Albanese, V., S. Reissmann and J. Frydman (2010). "A ribosome-anchored chaperone network that facilitates eukaryotic ribosome biogenesis." J Cell Biol 189(1): 69-81.

Alter, B. P., G. M. Baerlocher, S. A. Savage, S. J. Chanock, B. B. Weksler, J. P. Willner, J. A. Peters, N. Giri and P. M. Lansdorp (2007). "Very short telomere length by flow fluorescence in situ hybridization identifies patients with dyskeratosis congenita." Blood 110(5): 1439-1447.

Alter, B. P., N. Giri, S. A. Savage, J. A. Peters, J. T. Loud, L. Leathwood, A. G. Carr, M. H. Greene and P. S. Rosenberg (2010). "Malignancies and survival patterns in the National Cancer Institute inherited bone marrow failure syndromes cohort study." Br J Haematol 150(2): 179-188.

Ambekar, C., B. Das, H. Yeger and Y. Dror (2010). "SBDS-deficiency results in deregulation of reactive oxygen species leading to increased cell death and decreased cell growth." Pediatr Blood Cancer 55(6): 1138-1144.

Anazi, S., S. Maddirevula, E. Faqeih, H. Alsedairy, F. Alzahrani, H. E. Shamseldin, N. Patel, M. Hashem, N. Ibrahim, F. Abdulwahab, N. Ewida, H. S. Alsaif, H. Al Sharif, W. Alamoudi, A. Kentab, F. A. Bashiri, M. Alnaser, A. H. AlWadei, M. Alfadhel, W. Eyaid, A. Hashem, A. Al Asmari, M. M. Saleh, A. AlSaman, K. A. Alhasan, M. Alsughayir, M. Al Shammari, A. Mahmoud, Z. N. Al-Hassnan, M. Al-Husain, R. Osama Khalil, N. Abd El Meguid, A. Masri, R. Ali, T. Ben-Omran, P. El Fishway, A. Hashish, A. Ercan Sencicek, M. State, A. M. Alazami, M. A. Salih, N. Altassan, S. T. Arold, M. Abouelhoda, S. M. Wakil, D. Monies, R. Shaheen and F. S. Alkuraya (2017). "Clinical genomics expands the morbid genome of intellectual disability and offers a high diagnostic yield." Mol Psychiatry 22(4): 615-624.

Armistead, J. and B. Triggs-Raine (2014). "Diverse diseases from a ubiquitous process: the ribosomopathy paradox." FEBS Lett 588(9): 1491-1500.

Asano, N., H. Atsuumi, A. Nakamura, Y. Tanaka, I. Tanaka and M. Yao (2014). "Direct interaction between EFL1 and SBDS is mediated by an intrinsically disordered insertion domain." Biochem Biophys Res Commun 443(4): 1251-1256.

Astrom, J., A. Astrom and A. Virtanen (1991). "In vitro deadenylation of mammalian mRNA by a HeLa cell 3' exonuclease." Embo J 10(10): 3067-3071.

Astrom, J., A. Astrom and A. Virtanen (1992). "Properties of a HeLa cell 3' exonuclease specific for degrading poly(A) tails of mammalian mRNA." J Biol Chem 267(25): 18154-18159.

Austin, K. M., M. L. Gupta, Jr., S. A. Coats, A. Tulpule, G. Mostoslavsky, A. B. Balazs, R. C. Mulligan, G. Daley, D. Pellman and A. Shimamura (2008). "Mitotic spindle destabilization and genomic instability in Shwachman-Diamond syndrome." J Clin Invest 118(4): 1511-1518.

Austin, K. M., R. J. Leary and A. Shimamura (2005). "The Shwachman-Diamond SBDS protein localizes to the nucleolus." Blood 106(4): 1253-1258.

Bagby, G. (2018). "Recent advances in understanding hematopoiesis in Fanconi Anemia." F1000Res 7: 105.

Ball, H. L., B. Zhang, J. J. Riches, R. Gandhi, J. Li, J. M. Rommens and J. S. Myers (2009). "Shwachman-Bodian Diamond syndrome is a multi-functional protein implicated in cellular stress responses." Hum Mol Genet 18(19): 3684-3695.

Bar-Nur, O., H. A. Russ, S. Efrat and N. Benvenisty (2011). "Epigenetic memory and preferential lineage-specific differentiation in induced pluripotent stem cells derived from human pancreatic islet beta cells." Cell Stem Cell 9(1): 17-23.

Berndt, H., C. Harnisch, C. Rammelt, N. Stohr, A. Zirkel, J. C. Dohm, H. Himmelbauer, J. P. Tavanez, S. Huttelmaier and E. Wahle (2012). "Maturation of mammalian H/ACA box snoRNAs: PAPD5-dependent adenylation and PARN-dependent trimming." Rna 18(5): 958-972.

Beser, O. F., F. C. Cokugras, T. Erkan, T. Kutlu, I. Adaletli and S. Kurugoglu (2014). "Shwachman-Diamond syndrome with development of bone formation defects during prenatal life." J Pediatr Gastroenterol Nutr 58(4): e38-40.

Bezzerri, V., A. Vella, E. Calcaterra, A. Finotti, J. Gasparello, R. Gambari, B. M. Assael, M. Cipolli and C. Sorio (2016). "New insights into the Shwachman-Diamond Syndrome-related haematological disorder: hyper-activation of mTOR and STAT3 in leukocytes." Sci Rep 6: 33165.

Bhatla, D., S. M. Davies, S. Shenoy, R. E. Harris, M. Crockett, L. Shoultz, T. Smolarek, J. Bleesing, M. Hansen, S. Jodele, M. Jordan, A. H. Filipovich and P. A. Mehta (2008). "Reduced- intensity conditioning is effective and safe for transplantation of patients with Shwachman- Diamond syndrome." Bone Marrow Transplant 42(3): 159-165.

Bock, C., E. Kiskinis, G. Verstappen, H. Gu, G. Boulting, Z. D. Smith, M. Ziller, G. F. Croft, M. W. Amoroso, D. H. Oakley, A. Gnirke, K. Eggan and A. Meissner (2011). "Reference Maps of human ES and iPS cell variation enable high-throughput characterization of pluripotent cell lines." Cell 144(3): 439-452.

Bodian, M., W. Sheldon and R. Lightwood (1964). "Congenital Hypoplasia of the Exocrine Pancreas." Acta Paediatr 53: 282-293.

Bolli, N., E. M. Payne, J. Rhodes, E. Gjini, A. B. Johnston, F. Guo, J. S. Lee, R. A. Stewart, J. P. Kanki, A. T. Chen, Y. Zhou, L. I. Zon and A. T. Look (2011). "cpsf1 is required for definitive HSC survival in zebrafish." Blood 117(15): 3996-4007.

Boocock, G. R., M. R. Marit and J. M. Rommens (2006). "Phylogeny, sequence conservation, and functional complementation of the SBDS protein family." Genomics 87(6): 758-771.

Boocock, G. R., J. A. Morrison, M. Popovic, N. Richards, L. Ellis, P. R. Durie and J. M. Rommens (2003). "Mutations in SBDS are associated with Shwachman-Diamond syndrome." Nat Genet 33(1): 97-101.

Booij, J., L. Reneman, M. Alders and T. W. Kuijpers (2013). "Increase in central striatal dopamine transporters in patients with Shwachman-Diamond syndrome: additional evidence of a brain phenotype." Am J Med Genet A 161A(1): 102-107.

Boulting, G. L., E. Kiskinis, G. F. Croft, M. W. Amoroso, D. H. Oakley, B. J. Wainger, D. J. Williams, D. J. Kahler, M. Yamaki, L. Davidow, C. T. Rodolfa, J. T. Dimos, S. Mikkilineni, A. B. MacDermott, C. J. Woolf, C. E. Henderson, H. Wichterle and K. Eggan (2011). "A functionally characterized test set of human induced pluripotent stem cells." Nat Biotechnol 29(3): 279-286.

Boyer, L. A., T. I. Lee, M. F. Cole, S. E. Johnstone, S. S. Levine, J. P. Zucker, M. G. Guenther, R. M. Kumar, H. L. Murray, R. G. Jenner, D. K. Gifford, D. A. Melton, R. Jaenisch and R. A. Young (2005). "Core transcriptional regulatory circuitry in human embryonic stem cells." Cell 122(6): 947-956.

Brouwer, M., H. Zhou and N. Nadif Kasri (2016). "Choices for Induction of Pluripotency: Recent Developments in Human Induced Pluripotent Stem Cell Reprogramming Strategies." Stem Cell Rev 12(1): 54-72.

Brown, C. E., S. Z. Tarun, Jr., R. Boeck and A. B. Sachs (1996). "PAN3 encodes a subunit of the Pab1p-dependent poly(A) nuclease in Saccharomyces cerevisiae." Mol Cell Biol 16(10): 5744- 5753.

Brueton, M. J., J. Mavromichalis, M. C. Goodchild and C. M. Anderson (1977). "Hepatic dysfunction in association with pancreatic insufficiency and cyclical neutropenia. Shwachman- Diamond syndrome." Arch Dis Child 52(1): 76-78.

Buganim, Y., D. A. Faddah, A. W. Cheng, E. Itskovich, S. Markoulaki, K. Ganz, S. L. Klemm, A. van Oudenaarden and R. Jaenisch (2012). "Single-cell expression analyses during cellular reprogramming reveal an early stochastic and a late hierarchic phase." Cell 150(6): 1209-1222.

Burke, V., J. H. Colebatch, C. M. Anderson and M. J. Simons (1967). "Association of pancreatic insufficiency and chronic neutropenia in childhood." Arch Dis Child 42(222): 147-157.

Burwick, N., S. A. Coats, T. Nakamura and A. Shimamura (2012). "Impaired ribosomal subunit association in Shwachman-Diamond syndrome." Blood 120(26): 5143-5152.

Caselitz, J., G. Kloppel, G. Delling, R. Gruttner, U. Holdhoff and M. Stern (1979). "Shwachman's syndrome and leukaemia." Virchows Arch A Pathol Anat Histol 385(1): 109-116.

Cesaro, S., R. Oneto, C. Messina, B. E. Gibson, A. Buzyn, C. Steward, E. Gluckman, R. Bredius, M. Boogaerts, C. Vermylen, P. Veys, J. Marsh, I. Badell, G. Michel, T. Gungor, D. Niethammer, P. Bordigoni, C. Oswald, C. Favre, J. Passweg, G. Dini, E. S. A. Anaemia and P. Paediatric Diseases Working (2005). "Haematopoietic stem cell transplantation for Shwachman-Diamond disease: a study from the European Group for blood and marrow transplantation." Br J Haematol 131(2): 231-236.

Chen, A. E., D. Egli, K. Niakan, J. Deng, H. Akutsu, M. Yamaki, C. Cowan, C. Fitz-Gerald, K. Zhang, D. A. Melton and K. Eggan (2009). "Optimal timing of inner cell mass isolation increases the efficiency of human embryonic stem cell derivation and allows generation of sibling cell lines." Cell Stem Cell 4(2): 103-106.

Chen, J., J. Liu, J. Yang, Y. Chen, J. Chen, S. Ni, H. Song, L. Zeng, K. Ding and D. Pei (2011). "BMPs functionally replace Klf4 and support efficient reprogramming of mouse fibroblasts by Oct4 alone." Cell Res 21(1): 205-212.

Chen, J., G. Yin, Y. Lu, M. Lou, H. Cheng, X. Ni, G. Hu, C. Luo, K. Ying, Y. Xie and Y. Mao (2004). "Cloning and characterization of a novel human cDNA encoding a J-domain protein (DNAJA5) from the fetal brain." Int J Mol Med 13(5): 735-740.

Cheng, L., N. F. Hansen, L. Zhao, Y. Du, C. Zou, F. X. Donovan, B. K. Chou, G. Zhou, S. Li, S. N. Dowey, Z. Ye, N. C. S. Program, S. C. Chandrasekharappa, H. Yang, J. C. Mullikin and P. P. Liu (2012). "Low incidence of DNA sequence variation in human induced pluripotent stem cells generated by nonintegrating plasmid expression." Cell Stem Cell 10(3): 337-344.

Chin, M. H., M. J. Mason, W. Xie, S. Volinia, M. Singer, C. Peterson, G. Ambartsumyan, O. Aimiuwu, L. Richter, J. Zhang, I. Khvorostov, V. Ott, M. Grunstein, N. Lavon, N. Benvenisty, C. M. Croce, A. T. Clark, T. Baxter, A. D. Pyle, M. A. Teitell, M. Pelegrini, K. Plath and W. E. Lowry (2009). "Induced pluripotent stem cells and embryonic stem cells are distinguished by gene expression signatures." Cell Stem Cell 5(1): 111-123.

Chin, M. H., M. Pellegrini, K. Plath and W. E. Lowry (2010). "Molecular analyses of human induced pluripotent stem cells and embryonic stem cells." Cell Stem Cell 7(2): 263-269.

Choi, J., S. Lee, W. Mallard, K. Clement, G. M. Tagliazucchi, H. Lim, I. Y. Choi, F. Ferrari, A. M. Tsankov, R. Pop, G. Lee, J. L. Rinn, A. Meissner, P. J. Park and K. Hochedlinger (2015). "A comparison of genetically matched cell lines reveals the equivalence of human iPSCs and ESCs." Nat Biotechnol 33(11): 1173-1181.

Craig, E. A., P. Huang, R. Aron and A. Andrew (2006). "The diverse roles of J-proteins, the obligate Hsp70 co-chaperone." Rev Physiol Biochem Pharmacol 156: 1-21.

Crosby, A. H., H. Patel, B. A. Chioza, C. Proukakis, K. Gurtz, M. A. Patton, R. Sharifi, G. Harlalka, M. A. Simpson, K. Dick, J. A. Reed, A. Al-Memar, Z. M. Chrzanowska-Lightowlers, H. E. Cross and R. N. Lightowlers (2010). "Defective mitochondrial mRNA maturation is associated with spastic ataxia." Am J Hum Genet 87(5): 655-660.

de Oliveira, J. F., M. L. Sforca, T. M. Blumenschein, M. B. Goldfeder, B. G. Guimaraes, C. C. Oliveira, N. I. Zanchin and A. C. Zeri (2010). "Structure, dynamics, and RNA interaction analysis of the human SBDS protein." J Mol Biol 396(4): 1053-1069.

Demoinet, E., A. Jacquier, G. Lutfalla and M. Fromont-Racine (2007). "The Hsp40 chaperone Jjj1 is required for the nucleo-cytoplasmic recycling of preribosomal factors in Saccharomyces cerevisiae." RNA 13(9): 1570-1581.

Dhanraj, S., S. M. Gunja, A. P. Deveau, M. Nissbeck, B. Boonyawat, A. J. Coombs, A. Renieri, M. Mucciolo, A. Marozza, S. Buoni, L. Turner, H. Li, A. Jarrar, M. Sabanayagam, M. Kirby, M. Shago, D. Pinto, J. N. Berman, S. W. Scherer, A. Virtanen and Y. Dror (2015). "Bone marrow failure and developmental delay caused by mutations in poly(A)-specific ribonuclease (PARN)." J Med Genet 52(11): 738-748.

Dhanraj, S., A. Manji, D. Pinto, S. W. Scherer, H. Favre, M. L. Loh, R. Chetty, A. C. Wei and Y. Dror (2013). "Molecular characteristics of a pancreatic adenocarcinoma associated with Shwachman-Diamond syndrome." Pediatr Blood Cancer 60(5): 754-760.

Dhanraj, S., A. Matveev, H. Li, S. Lauhasurayotin, L. Jardine, M. Cada, B. Zlateska, C. S. Tailor, J. Zhou, R. Mendoza-Londono, A. Vincent, P. R. Durie, S. W. Scherer, J. M. Rommens, E. Heon and Y. Dror (2017). "Biallelic mutations in DNAJC21 cause Shwachman-Diamond syndrome." Blood 129(11): 1557-1562.

Ditadi, A., C. M. Sturgeon and G. Keller (2017). "A view of human haematopoietic development from the Petri dish." Nat Rev Mol Cell Biol 18(1): 56-67.

Doi, A., I. H. Park, B. Wen, P. Murakami, M. J. Aryee, R. Irizarry, B. Herb, C. Ladd-Acosta, J. Rho, S. Loewer, J. Miller, T. Schlaeger, G. Q. Daley and A. P. Feinberg (2009). "Differential methylation of tissue- and cancer-specific CpG island shores distinguishes human induced pluripotent stem cells, embryonic stem cells and fibroblasts." Nat Genet 41(12): 1350-1353.

Dokal, I., S. Rule, F. Chen, M. Potter and J. Goldman (1997). "Adult onset of acute myeloid leukaemia (M6) in patients with Shwachman-Diamond syndrome." Br J Haematol 99(1): 171- 173.

Donadieu, J., O. Fenneteau, B. Beaupain, S. Beaufils, F. Bellanger, N. Mahlaoui, A. Lambilliotte, N. Aladjidi, Y. Bertrand, V. Mialou, C. Perot, G. Michel, F. Fouyssac, C. Paillard, V. Gandemer, P. Boutard, J. Schmitz, A. Morali, T. Leblanc, C. Bellanne-Chantelot and R. Associated investigators of the French Severe Chronic Neutropenia (2012). "Classification of and risk factors for hematologic complications in a French national cohort of 102 patients with Shwachman-Diamond syndrome." Haematologica 97(9): 1312-1319.

Donadieu, J., G. Michel, E. Merlin, P. Bordigoni, B. Monteux, B. Beaupain, G. Leverger, J. P. Laporte, O. Hermine, A. Buzyn, Y. Bertrand, J. L. Casanova, T. Leblanc, E. Gluckman, A. Fischer and J. L. Stephan (2005). "Hematopoietic stem cell transplantation for Shwachman- Diamond syndrome: experience of the French neutropenia registry." Bone Marrow Transplant 36(9): 787-792.

Dong, H., D. W. Nebert, E. A. Bruford, D. C. Thompson, H. Joenje and V. Vasiliou (2015). "Update of the human and mouse Fanconi anemia genes." Hum Genomics 9: 32.

Draptchinskaia, N., P. Gustavsson, B. Andersson, M. Pettersson, T. N. Willig, I. Dianzani, S. Ball, G. Tchernia, J. Klar, H. Matsson, D. Tentler, N. Mohandas, B. Carlsson and N. Dahl (1999). "The gene encoding ribosomal protein S19 is mutated in Diamond-Blackfan anaemia." Nat Genet 21(2): 169-175.

Dror, Y. (2005). "Shwachman-Diamond syndrome." Pediatr Blood Cancer 45(7): 892-901.

Dror, Y., J. Donadieu, J. Koglmeier, J. Dodge, S. Toiviainen-Salo, O. Makitie, E. Kerr, C. Zeidler, A. Shimamura, N. Shah, M. Cipolli, T. Kuijpers, P. Durie, J. Rommens, L. Siderius and J. M. Liu (2011). "Draft consensus guidelines for diagnosis and treatment of Shwachman- Diamond syndrome." Ann N Y Acad Sci 1242: 40-55.

Dror, Y., P. Durie, H. Ginzberg, R. Herman, A. Banerjee, M. Champagne, K. Shannon, D. Malkin and M. H. Freedman (2002). "Clonal evolution in marrows of patients with Shwachman- Diamond syndrome: a prospective 5-year follow-up study." Exp Hematol 30(7): 659-669.

Dror, Y. and M. H. Freedman (1999). "Shwachman-Diamond syndrome: An inherited preleukemic bone marrow failure disorder with aberrant hematopoietic progenitors and faulty marrow microenvironment." Blood 94(9): 3048-3054.

Dror, Y. and M. H. Freedman (2001). "Shwachman-Diamond syndrome marrow cells show abnormally increased apoptosis mediated through the Fas pathway." Blood 97(10): 3011-3016.

Dror, Y., H. Ginzberg, I. Dalal, V. Cherepanov, G. Downey, P. Durie, C. M. Roifman and M. H. Freedman (2001). "Immune function in patients with Shwachman-Diamond syndrome." Br J Haematol 114(3): 712-717.

Dror, Y., J. Squire, P. Durie and M. H. Freedman (1998). "Malignant myeloid transformation with isochromosome 7q in Shwachman-Diamond syndrome." Leukemia 12(10): 1591-1595.

Dumitriu, I. E., W. Mohr, W. Kolowos, P. Kern, J. R. Kalden and M. Herrmann (2001). "5,6- carboxyfluorescein diacetate succinimidyl ester-labeled apoptotic and necrotic as well as detergent-treated cells can be traced in composite cell samples." Anal Biochem 299(2): 247-252.

Elghetany, M. T. and B. P. Alter (2002). "p53 protein overexpression in bone marrow biopsies of patients with Shwachman-Diamond syndrome has a prevalence similar to that of patients with refractory anemia." Arch Pathol Lab Med 126(4): 452-455.

Evans, M. J. and M. H. Kaufman (1981). "Establishment in culture of pluripotential cells from mouse embryos." Nature 292(5819): 154-156.

Farrar, J. E., M. Nater, E. Caywood, M. A. McDevitt, J. Kowalski, C. M. Takemoto, C. C. Talbot, Jr., P. Meltzer, D. Esposito, A. H. Beggs, H. E. Schneider, A. Grabowska, S. E. Ball, E. Niewiadomska, C. A. Sieff, A. Vlachos, E. Atsidaftos, S. R. Ellis, J. M. Lipton, H. T. Gazda and R. J. Arceci (2008). "Abnormalities of the large ribosomal subunit protein, Rpl35a, in Diamond- Blackfan anemia." Blood 112(5): 1582-1592.

Finch, A. J., C. Hilcenko, N. Basse, L. F. Drynan, B. Goyenechea, T. F. Menne, A. Gonzalez Fernandez, P. Simpson, C. S. D'Santos, M. J. Arends, J. Donadieu, C. Bellanne-Chantelot, M. Costanzo, C. Boone, A. N. McKenzie, S. M. Freund and A. J. Warren (2011). "Uncoupling of GTP hydrolysis from eIF6 release on the ribosome causes Shwachman-Diamond syndrome." Genes Dev 25(9): 917-929.

Flygare, J., A. Aspesi, J. C. Bailey, K. Miyake, J. M. Caffrey, S. Karlsson and S. R. Ellis (2007). "Human RPS19, the gene mutated in Diamond-Blackfan anemia, encodes a ribosomal protein required for the maturation of 40S ribosomal subunits." Blood 109(3): 980-986.

Friedel, T., S. Jung-Klawitter, A. Sebe, F. Schenk, U. Modlich, Z. Ivics, G. G. Schumann, C. J. Buchholz and I. C. Schneider (2016). "CD30 Receptor-Targeted Lentiviral Vectors for Human Induced Pluripotent Stem Cell-Specific Gene Modification." Stem Cells Dev 25(9): 729-739.

Fusaki, N., H. Ban, A. Nishiyama, K. Saeki and M. Hasegawa (2009). "Efficient induction of transgene-free human pluripotent stem cells using a vector based on Sendai virus, an RNA virus that does not integrate into the host genome." Proc Jpn Acad Ser B Phys Biol Sci 85(8): 348-362.

Gana, S., L. Sainati, M. R. Frau, C. Monciotti, F. Poli, Z. Cannioto, M. Comelli, C. Danesino and A. Minelli (2011). "Shwachman-Diamond syndrome and type 1 diabetes mellitus: more than a chance association?" Exp Clin Endocrinol Diabetes 119(10): 610-612.

Ganapathi, K. A., K. M. Austin, C. S. Lee, A. Dias, M. M. Malsch, R. Reed and A. Shimamura (2007). "The human Shwachman-Diamond syndrome protein, SBDS, associates with ribosomal RNA." Blood 110(5): 1458-1465.

Ghemlas, I., H. Li, B. Zlateska, R. Klaassen, C. V. Fernandez, R. A. Yanofsky, J. Wu, Y. Pastore, M. Silva, J. H. Lipton, J. Brossard, B. Michon, S. Abish, M. Steele, R. Sinha, M. Belletrutti, V. R. Breakey, L. Jardine, L. Goodyear, L. Sung, S. Dhanraj, E. Reble, A. Wagner, J. Beyene, P. Ray, S. Meyn, M. Cada and Y. Dror (2015). "Improving diagnostic precision, care and syndrome definitions using comprehensive next-generation sequencing for the inherited bone marrow failure syndromes." J Med Genet 52(9): 575-584.

Ghosh, Z., K. D. Wilson, Y. Wu, S. Hu, T. Quertermous and J. C. Wu (2010). "Persistent donor cell gene expression among human induced pluripotent stem cells contributes to differences with human embryonic stem cells." PLoS One 5(2): e8975.

Gijsbers, A., A. Garcia-Marquez, A. Luviano and N. Sanchez-Puig (2013). "Guanine nucleotide exchange in the ribosomal GTPase EFL1 is modulated by the protein mutated in the Shwachman-Diamond syndrome." Biochem Biophys Res Commun 437(3): 349-354.

Gillies, A. T., R. Taylor and J. E. Gestwicki (2012). "Synthetic lethal interactions in yeast reveal functional roles of J protein co-chaperones." Mol Biosyst 8(11): 2901-2908.

Ginzberg, H., J. Shin, L. Ellis, S. Goobie, J. Morrison, M. Corey, P. R. Durie and J. M. Rommens (2000). "Segregation analysis in Shwachman-Diamond syndrome: evidence for recessive inheritance." Am J Hum Genet 66(4): 1413-1416.

Ginzberg, H., J. Shin, L. Ellis, J. Morrison, W. Ip, Y. Dror, M. Freedman, L. A. Heitlinger, M. A. Belt, M. Corey, J. M. Rommens and P. R. Durie (1999). "Shwachman syndrome: phenotypic manifestations of sibling sets and isolated cases in a large patient cohort are similar." J Pediatr 135(1): 81-88.

Goobie, S., M. Popovic, J. Morrison, L. Ellis, H. Ginzberg, G. R. Boocock, N. Ehtesham, C. Betard, C. G. Brewer, N. M. Roslin, T. J. Hudson, K. Morgan, T. M. Fujiwara, P. R. Durie and J. M. Rommens (2001). "Shwachman-Diamond syndrome with exocrine pancreatic dysfunction and bone marrow failure maps to the centromeric region of chromosome 7." Am J Hum Genet 68(4): 1048-1054.

Greber, B. J., D. Boehringer, C. Montellese and N. Ban (2012). "Cryo-EM structures of Arx1 and maturation factors Rei1 and Jjj1 bound to the 60S ribosomal subunit." Nat Struct Mol Biol 19(12): 1228-1233.

Guenther, M. G., G. M. Frampton, F. Soldner, D. Hockemeyer, M. Mitalipova, R. Jaenisch and R. A. Young (2010). "Chromatin structure and gene expression programs of human embryonic and induced pluripotent stem cells." Cell Stem Cell 7(2): 249-257.

Guerrero, J., F. Lopez Barea, C. Calvo, F. Moreno and I. Solas (1979). "[Myocardial fibrosis in Shwachman's syndrome (author's transl)]." An Esp Pediatr 12(6-7): 542-548.

Hashmi, S. K., C. Allen, R. Klaassen, C. V. Fernandez, R. Yanofsky, E. Shereck, J. Champagne, M. Silva, J. H. Lipton, J. Brossard, Y. Samson, S. Abish, M. Steele, K. Ali, N. Dower, U. Athale, L. Jardine, J. P. Hand, J. Beyene and Y. Dror (2011). "Comparative analysis of Shwachman- Diamond syndrome to other inherited bone marrow failure syndromes and genotype-phenotype correlation." Clin Genet 79(5): 448-458.

Hauet, Q., B. Beaupain, M. Micheau, M. Blayo, V. Gandemer, F. Gottrand, N. Blin, F. Fouyssac, J. P. Lethor, C. Bellanne-Chantelot, D. Bonnet and J. Donadieu (2013). "Cardiomyopathies and congenital heart diseases in Shwachman-Diamond syndrome: a national survey." Int J Cardiol 167(3): 1048-1050.

Heiss, N. S., S. W. Knight, T. J. Vulliamy, S. M. Klauck, S. Wiemann, P. J. Mason, A. Poustka and I. Dokal (1998). "X-linked dyskeratosis congenita is caused by mutations in a highly conserved gene with putative nucleolar functions." Nat Genet 19(1): 32-38.

Helser, T. L., R. A. Baan and A. E. Dahlberg (1981). "Characterization of a 40S ribosomal subunit complex in polyribosomes of Saccharomyces cerevisiae treated with cycloheximide." Mol Cell Biol 1(1): 51-57.

Henson, A. L., J. B. t. Moore, P. Alard, M. M. Wattenberg, J. M. Liu and S. R. Ellis (2013). "Mitochondrial function is impaired in yeast and human cellular models of Shwachman Diamond syndrome." Biochem Biophys Res Commun 437(1): 29-34.

Hiler, D., X. Chen, J. Hazen, S. Kupriyanov, P. A. Carroll, C. Qu, B. Xu, D. Johnson, L. Griffiths, S. Frase, A. R. Rodriguez, G. Martin, J. Zhang, J. Jeon, Y. Fan, D. Finkelstein, R. N. Eisenman, K. Baldwin and M. A. Dyer (2015). "Quantification of Retinogenesis in 3D Cultures

Reveals Epigenetic Memory and Higher Efficiency in iPSCs Derived from Rod Photoreceptors." Cell Stem Cell 17(1): 101-115.

Ho, W., C. Cheretakis, P. Durie, G. Kulkarni and M. Glogauer (2007). "Prevalence of oral diseases in Shwachman-Diamond syndrome." Spec Care Dentist 27(2): 52-58.

Horos, R., H. Ijspeert, D. Pospisilova, R. Sendtner, C. Andrieu-Soler, E. Taskesen, A. Nieradka, R. Cmejla, M. Sendtner, I. P. Touw and M. von Lindern (2012). "Ribosomal deficiencies in Diamond-Blackfan anemia impair translation of transcripts essential for differentiation of murine and human erythroblasts." Blood 119(1): 262-272.

Hu, B. Y., J. P. Weick, J. Yu, L. X. Ma, X. Q. Zhang, J. A. Thomson and S. C. Zhang (2010). "Neural differentiation of human induced pluripotent stem cells follows developmental principles but with variable potency." Proc Natl Acad Sci U S A 107(9): 4335-4340.

Huangfu, D., K. Osafune, R. Maehr, W. Guo, A. Eijkelenboom, S. Chen, W. Muhlestein and D. A. Melton (2008). "Induction of pluripotent stem cells from primary human fibroblasts with only Oct4 and Sox2." Nat Biotechnol 26(11): 1269-1275.

Huijgens, P. C., E. A. van der Veen, S. Meijer and O. G. Muntinghe (1977). "Syndrome of Shwachman and leukaemia." Scand J Haematol 18(1): 20-24.

Hussein, S. M., N. N. Batada, S. Vuoristo, R. W. Ching, R. Autio, E. Narva, S. Ng, M. Sourour, R. Hamalainen, C. Olsson, K. Lundin, M. Mikkola, R. Trokovic, M. Peitz, O. Brustle, D. P. Bazett-Jones, K. Alitalo, R. Lahesmaa, A. Nagy and T. Otonkoski (2011). "Copy number variation and selection during reprogramming to pluripotency." Nature 471(7336): 58-62.

In, K., M. A. Zaini, C. Muller, A. J. Warren, M. von Lindern and C. F. Calkhoven (2016). "Shwachman-Bodian-Diamond syndrome (SBDS) protein deficiency impairs translation re- initiation from C/EBPalpha and C/EBPbeta mRNAs." Nucleic Acids Res 44(9): 4134-4146.

Ip, W. F., A. Dupuis, L. Ellis, S. Beharry, J. Morrison, M. O. Stormon, M. Corey, J. M. Rommens and P. R. Durie (2002). "Serum pancreatic enzymes define the pancreatic phenotype in patients with Shwachman-Diamond syndrome." J Pediatr 141(2): 259-265.

Jung, M., C. E. Dunbar and T. Winkler (2015). "Modeling Human Bone Marrow Failure Syndromes Using Pluripotent Stem Cells and Genome Engineering." Mol Ther 23(12): 1832- 1842.

Kajiwara, M., T. Aoi, K. Okita, R. Takahashi, H. Inoue, N. Takayama, H. Endo, K. Eto, J. Toguchida, S. Uemoto and S. Yamanaka (2012). "Donor-dependent variations in hepatic differentiation from human-induced pluripotent stem cells." Proc Natl Acad Sci U S A 109(31): 12538-12543.

Karbstein, K. (2010). "Chaperoning ribosome assembly." J Cell Biol 189(1): 11-12.

Kaschner, L. A., R. Sharma, O. K. Shrestha, A. E. Meyer and E. A. Craig (2015). "A conserved domain important for association of eukaryotic J-protein co-chaperones Jjj1 and Zuo1 with the ribosome." Biochim Biophys Acta 1853(5): 1035-1045.

Kent, A., G. H. Murphy and P. Milla (1990). "Psychological characteristics of children with Shwachman syndrome." Arch Dis Child 65(12): 1349-1352.

Kerr, E. N., L. Ellis, A. Dupuis, J. M. Rommens and P. R. Durie (2010). "The behavioral phenotype of school-age children with shwachman diamond syndrome indicates neurocognitive dysfunction with loss of Shwachman-Bodian-Diamond syndrome gene function." J Pediatr 156(3): 433-438.

Kim, J., J. Chu, X. Shen, J. Wang and S. H. Orkin (2008). "An extended transcriptional network for pluripotency of embryonic stem cells." Cell 132(6): 1049-1061.

Kim, K., R. Zhao, A. Doi, K. Ng, J. Unternaehrer, P. Cahan, H. Huo, Y. H. Loh, M. J. Aryee, M. W. Lensch, H. Li, J. J. Collins, A. P. Feinberg and G. Q. Daley (2011). "Donor cell type can influence the epigenome and differentiation potential of human induced pluripotent stem cells." Nat Biotechnol 29(12): 1117-1119.

Koche, R. P., Z. D. Smith, M. Adli, H. Gu, M. Ku, A. Gnirke, B. E. Bernstein and A. Meissner (2011). "Reprogramming factor expression initiates widespread targeted chromatin remodeling." Cell Stem Cell 8(1): 96-105.

Kondrashov, N., A. Pusic, C. R. Stumpf, K. Shimizu, A. C. Hsieh, J. Ishijima, T. Shiroishi and M. Barna (2011). "Ribosome-mediated specificity in Hox mRNA translation and vertebrate tissue patterning." Cell 145(3): 383-397.

Koonin, E. V., Y. I. Wolf and L. Aravind (2001). "Prediction of the archaeal exosome and its connections with the proteasome and the translation and transcription machineries by a comparative-genomic approach." Genome Res 11(2): 240-252.

Kopel, L., P. S. Gutierrez and S. G. Lage (2011). "Dilated cardiomyopathy in a case of Shwachman-Diamond syndrome." Cardiol Young 21(5): 588-590.

Korner, C. G., M. Wormington, M. Muckenthaler, S. Schneider, E. Dehlin and E. Wahle (1998). "The deadenylating nuclease (DAN) is involved in poly(A) tail removal during the meiotic maturation of Xenopus oocytes." EMBO J 17(18): 5427-5437.

Krijger, P. H., B. Di Stefano, E. de Wit, F. Limone, C. van Oevelen, W. de Laat and T. Graf (2016). "Cell-of-Origin-Specific 3D Genome Structure Acquired during Somatic Cell Reprogramming." Cell Stem Cell 18(5): 597-610.

Krogan, N. J., G. Cagney, H. Yu, G. Zhong, X. Guo, A. Ignatchenko, J. Li, S. Pu, N. Datta, A. P. Tikuisis, T. Punna, J. M. Peregrin-Alvarez, M. Shales, X. Zhang, M. Davey, M. D. Robinson, A. Paccanaro, J. E. Bray, A. Sheung, B. Beattie, D. P. Richards, V. Canadien, A. Lalev, F. Mena, P. Wong, A. Starostine, M. M. Canete, J. Vlasblom, S. Wu, C. Orsi, S. R. Collins, S. Chandran, R. Haw, J. J. Rilstone, K. Gandi, N. J. Thompson, G. Musso, P. St Onge, S. Ghanny, M. H. Lam, G. Butland, A. M. Altaf-Ul, S. Kanaya, A. Shilatifard, E. O'Shea, J. S. Weissman, C. J. Ingles, T. R. Hughes, J. Parkinson, M. Gerstein, S. J. Wodak, A. Emili and J. F. Greenblatt (2006). "Global landscape of protein complexes in the yeast Saccharomyces cerevisiae." Nature 440(7084): 637- 643.

Kuijpers, T. W., M. Alders, A. T. Tool, C. Mellink, D. Roos and R. C. Hennekam (2005). "Hematologic abnormalities in Shwachman Diamond syndrome: lack of genotype-phenotype relationship." Blood 106(1): 356-361.

Kuijpers, T. W., E. Nannenberg, M. Alders, R. Bredius and R. C. Hennekam (2004). "Congenital aplastic anemia caused by mutations in the SBDS gene: a rare presentation of Shwachman- Diamond syndrome." Pediatrics 114(3): e387-391.

Lackey, C. A., O. W. Press, A. S. Hoffman and P. S. Stayton (2002). "A biomimetic pH- responsive polymer directs endosomal release and intracellular delivery of an endocytosed antibody complex." Bioconjug Chem 13(5): 996-1001.

Laurent, L. C., I. Ulitsky, I. Slavin, H. Tran, A. Schork, R. Morey, C. Lynch, J. V. Harness, S. Lee, M. J. Barrero, S. Ku, M. Martynova, R. Semechkin, V. Galat, J. Gottesfeld, J. C. Izpisua Belmonte, C. Murry, H. S. Keirstead, H. S. Park, U. Schmidt, A. L. Laslett, F. J. Muller, C. M. Nievergelt, R. Shamir and J. F. Loring (2011). "Dynamic changes in the copy number of pluripotency and cell proliferation genes in human ESCs and iPSCs during reprogramming and time in culture." Cell Stem Cell 8(1): 106-118.

Le Gloan, L., N. Blin and J. M. Langlard (2014). "Atrioventricular septal defect in a case of Shwachman-Diamond syndrome." Cardiol Young 24(3): 549-551.

Lebreton, A., C. Saveanu, L. Decourty, J. C. Rain, A. Jacquier and M. Fromont-Racine (2006). "A functional network involved in the recycling of nucleocytoplasmic pre-60S factors." J Cell Biol 173(3): 349-360.

Lee, J. E., J. Y. Lee, J. Trembly, J. Wilusz, B. Tian and C. J. Wilusz (2012). "The PARN deadenylase targets a discrete set of mRNAs for decay and regulates cell motility in mouse myoblasts." PLoS Genet 8(8): e1002901.

Lesesve, J. F., F. Dugue, M. J. Gregoire, F. Witz and Y. Dror (2003). "Shwachman-Diamond syndrome with late-onset neutropenia and fatal acute myeloid leukaemia without maturation: a case report." Eur J Haematol 71(5): 393-395.

Leung, R., K. Cuddy, Y. Wang, J. Rommens and M. Glogauer (2011). "Sbds is required for Rac2-mediated monocyte migration and signaling downstream of RANK during osteoclastogenesis." Blood 117(6): 2044-2053.

Lister, R., M. Pelizzola, Y. S. Kida, R. D. Hawkins, J. R. Nery, G. Hon, J. Antosiewicz-Bourget, R. O'Malley, R. Castanon, S. Klugman, M. Downes, R. Yu, R. Stewart, B. Ren, J. A. Thomson, R. M. Evans and J. R. Ecker (2011). "Hotspots of aberrant epigenomic reprogramming in human induced pluripotent stem cells." Nature 471(7336): 68-73.

Loh, Y. H., Q. Wu, J. L. Chew, V. B. Vega, W. Zhang, X. Chen, G. Bourque, J. George, B. Leong, J. Liu, K. Y. Wong, K. W. Sung, C. W. Lee, X. D. Zhao, K. P. Chiu, L. Lipovich, V. A. Kuznetsov, P. Robson, L. W. Stanton, C. L. Wei, Y. Ruan, B. Lim and H. H. Ng (2006). "The Oct4 and Nanog transcription network regulates pluripotency in mouse embryonic stem cells." Nat Genet 38(4): 431-440.

Luz, J. S., R. C. Georg, C. H. Gomes, G. M. Machado-Santelli and C. C. Oliveira (2009). "Sdo1p, the yeast orthologue of Shwachman-Bodian-Diamond syndrome protein, binds RNA and interacts with nuclear rRNA-processing factors." Yeast 26(5): 287-298.

Mack, D. R., G. G. Forstner, M. Wilschanski, M. H. Freedman and P. R. Durie (1996). "Shwachman syndrome: exocrine pancreatic dysfunction and variable phenotypic expression." Gastroenterology 111(6): 1593-1602.

Maherali, N., R. Sridharan, W. Xie, J. Utikal, S. Eminli, K. Arnold, M. Stadtfeld, R. Yachechko, J. Tchieu, R. Jaenisch, K. Plath and K. Hochedlinger (2007). "Directly reprogrammed fibroblasts show global epigenetic remodeling and widespread tissue contribution." Cell Stem Cell 1(1): 55- 70.

Makitie, O., L. Ellis, P. R. Durie, J. A. Morrison, E. B. Sochett, J. M. Rommens and W. G. Cole (2004). "Skeletal phenotype in patients with Shwachman-Diamond syndrome and mutations in SBDS." Clin Genet 65(2): 101-112.

Malhotra, J. D. and R. J. Kaufman (2007). "Endoplasmic reticulum stress and oxidative stress: a vicious cycle or a double-edged sword?" Antioxid Redox Signal 9(12): 2277-2293.

Marchetto, M. C., G. W. Yeo, O. Kainohana, M. Marsala, F. H. Gage and A. R. Muotri (2009). "Transcriptional signature and memory retention of human-induced pluripotent stem cells." PLoS One 4(9): e7076.

Marshall, C. R., S. A. Farrell, D. Cushing, T. Paton, T. L. Stockley, D. J. Stavropoulos, P. N. Ray, M. Szego, L. Lau, S. L. Pereira, R. D. Cohn, R. F. Wintle, A. M. Abuzenadah, M. Abu- Elmagd and S. W. Scherer (2015). "Whole-exome analysis of foetal autopsy tissue reveals a frameshift mutation in OBSL1, consistent with a diagnosis of 3-M Syndrome." BMC Genomics 16 Suppl 1: S12.

Marson, A., R. Foreman, B. Chevalier, S. Bilodeau, M. Kahn, R. A. Young and R. Jaenisch (2008). "Wnt signaling promotes reprogramming of somatic cells to pluripotency." Cell Stem Cell 3(2): 132-135.

Martin, G. R. (1981). "Isolation of a pluripotent cell line from early mouse embryos cultured in medium conditioned by teratocarcinoma stem cells." Proc Natl Acad Sci U S A 78(12): 7634- 7638.

Martinez, J., Y. G. Ren, P. Nilsson, M. Ehrenberg and A. Virtanen (2001). "The mRNA cap structure stimulates rate of poly(A) removal and amplifies processivity of degradation." J Biol Chem 276(30): 27923-27929.

Martinez, J., Y. G. Ren, A. C. Thuresson, U. Hellman, J. Astrom and A. Virtanen (2000). "A 54- kDa fragment of the Poly(A)-specific ribonuclease is an oligomeric, processive, and cap- interacting Poly(A)-specific 3' exonuclease." J Biol Chem 275(31): 24222-24230.

Martins-Taylor, K., B. S. Nisler, S. M. Taapken, T. Compton, L. Crandall, K. D. Montgomery, M. Lalande and R. H. Xu (2011). "Recurrent copy number variations in human induced pluripotent stem cells." Nat Biotechnol 29(6): 488-491.

Martins-Taylor, K. and R. H. Xu (2012). "Concise review: Genomic stability of human induced pluripotent stem cells." Stem Cells 30(1): 22-27.

Masui, S., Y. Nakatake, Y. Toyooka, D. Shimosato, R. Yagi, K. Takahashi, H. Okochi, A. Okuda, R. Matoba, A. A. Sharov, M. S. Ko and H. Niwa (2007). "Pluripotency governed by Sox2 via regulation of Oct3/4 expression in mouse embryonic stem cells." Nat Cell Biol 9(6): 625-635.

Mayshar, Y., U. Ben-David, N. Lavon, J. C. Biancotti, B. Yakir, A. T. Clark, K. Plath, W. E. Lowry and N. Benvenisty (2010). "Identification and classification of chromosomal aberrations in human induced pluripotent stem cells." Cell Stem Cell 7(4): 521-531.

Medin, J. A., M. Migita, R. Pawliuk, S. Jacobson, M. Amiri, S. Kluepfel-Stahl, R. O. Brady, R. K. Humphries and S. Karlsson (1996). "A bicistronic therapeutic retroviral vector enables sorting of transduced CD34+ cells and corrects the enzyme deficiency in cells from Gaucher patients." Blood 87(5): 1754-1762.

Menne, T. F., B. Goyenechea, N. Sanchez-Puig, C. C. Wong, L. M. Tonkin, P. J. Ancliff, R. L. Brost, M. Costanzo, C. Boone and A. J. Warren (2007). "The Shwachman-Bodian-Diamond syndrome protein mediates translational activation of ribosomes in yeast." Nat Genet 39(4): 486- 495.

Meyer, A. E., L. A. Hoover and E. A. Craig (2010). "The cytosolic J-protein, Jjj1, and Rei1 function in the removal of the pre-60 S subunit factor Arx1." J Biol Chem 285(2): 961-968.

Meyer, A. E., N. J. Hung, P. Yang, A. W. Johnson and E. A. Craig (2007). "The specialized cytosolic J-protein, Jjj1, functions in 60S ribosomal subunit biogenesis." Proc Natl Acad Sci U S A 104(5): 1558-1563.

Mikkelsen, T. S., J. Hanna, X. Zhang, M. Ku, M. Wernig, P. Schorderet, B. E. Bernstein, R. Jaenisch, E. S. Lander and A. Meissner (2008). "Dissecting direct reprogramming through integrative genomic analysis." Nature 454(7200): 49-55.

Mills, J. A., K. Wang, P. Paluru, L. Ying, L. Lu, A. M. Galvao, D. Xu, Y. Yao, S. K. Sullivan, L. M. Sullivan, H. Mac, A. Omari, J. C. Jean, S. Shen, A. Gower, A. Spira, G. Mostoslavsky, D. N. Kotton, D. L. French, M. J. Weiss and P. Gadue (2013). "Clonal genetic and hematopoietic heterogeneity among human-induced pluripotent stem cell lines." Blood 122(12): 2047-2051.

Minelli, A., E. Maserati, E. Nicolis, M. Zecca, L. Sainati, D. Longoni, F. Lo Curto, G. Menna, F. Poli, E. De Paoli, M. Cipolli, F. Locatelli, F. Pasquali and C. Danesino (2009). "The isochromosome i(7)(q10) carrying c.258+2t>c mutation of the SBDS gene does not promote development of myeloid malignancies in patients with Shwachman syndrome." Leukemia 23(4): 708-711.

Mitsui, T., T. Kawakami, D. Sendo, M. Katsuura, Y. Shimizu and K. Hayasaka (2004). "Successful unrelated donor bone marrow transplantation for Shwachman-Diamond syndrome with leukemia." Int J Hematol 79(2): 189-192.

Mochizuki, Y., J. He, S. Kulkarni, M. Bessler and P. J. Mason (2004). "Mouse dyskerin mutations affect accumulation of telomerase RNA and small nucleolar RNA, telomerase activity, and ribosomal RNA processing." Proc Natl Acad Sci U S A 101(29): 10756-10761.

Moore, J. B. t., J. E. Farrar, R. J. Arceci, J. M. Liu and S. R. Ellis (2010). "Distinct ribosome maturation defects in yeast models of Diamond-Blackfan anemia and Shwachman-Diamond syndrome." Haematologica 95(1): 57-64.

Myers, K. C., A. A. Bolyard, B. Otto, T. E. Wong, A. T. Jones, R. E. Harris, S. M. Davies, D. C. Dale and A. Shimamura (2014). "Variable clinical presentation of Shwachman-Diamond syndrome: update from the North American Shwachman-Diamond Syndrome Registry." J Pediatr 164(4): 866-870.

Myers, K. C., S. R. Rose, M. M. Rutter, P. A. Mehta, J. C. Khoury, T. Cole and R. E. Harris (2013). "Endocrine evaluation of children with and without Shwachman-Bodian-Diamond syndrome gene mutations and Shwachman-Diamond syndrome." J Pediatr 162(6): 1235-1240, 1240 e1231.

Nakashima, E., A. Mabuchi, Y. Makita, M. Masuno, H. Ohashi, G. Nishimura and S. Ikegawa (2004). "Novel SBDS mutations caused by gene conversion in Japanese patients with Shwachman-Diamond syndrome." Hum Genet 114(4): 345-348.

Nezelof, C. and G. LeSec (1979). "Multifocal myocardial necrosis and fibrosis in pancreatic diseases of children." Pediatrics 63(3): 361-368.

Ng, C. L., D. G. Waterman, E. V. Koonin, A. D. Walters, J. P. Chong, M. N. Isupov, A. A. Lebedev, D. H. Bunka, P. G. Stockley, M. Ortiz-Lombardia and A. A. Antson (2009). "Conformational flexibility and molecular interactions of an archaeal homologue of the Shwachman-Bodian-Diamond syndrome protein." BMC Struct Biol 9: 32.

Nichols, J., B. Zevnik, K. Anastassiadis, H. Niwa, D. Klewe-Nebenius, I. Chambers, H. Scholer and A. Smith (1998). "Formation of pluripotent stem cells in the mammalian embryo depends on the POU transcription factor Oct4." Cell 95(3): 379-391.

Nishino, K., M. Toyoda, M. Yamazaki-Inoue, Y. Fukawatase, E. Chikazawa, H. Sakaguchi, H. Akutsu and A. Umezawa (2011). "DNA methylation dynamics in human induced pluripotent stem cells over time." PLoS Genet 7(5): e1002085.

Ohi, Y., H. Qin, C. Hong, L. Blouin, J. M. Polo, T. Guo, Z. Qi, S. L. Downey, P. D. Manos, D. J. Rossi, J. Yu, M. Hebrok, K. Hochedlinger, J. F. Costello, J. S. Song and M. Ramalho-Santos (2011). "Incomplete DNA methylation underlies a transcriptional memory of somatic cells in human iPS cells." Nat Cell Biol 13(5): 541-549.

Okcu, F., W. M. Roberts and K. W. Chan (1998). "Bone marrow transplantation in Shwachman- Diamond syndrome: report of two cases and review of the literature." Bone Marrow Transplant 21(8): 849-851.

Orelio, C. and T. W. Kuijpers (2009). "Shwachman-Diamond syndrome neutrophils have altered chemoattractant-induced F-actin polymerization and polarization characteristics." Haematologica 94(3): 409-413.

Orelio, C., P. Verkuijlen, J. Geissler, T. K. van den Berg and T. W. Kuijpers (2009). "SBDS expression and localization at the mitotic spindle in human myeloid progenitors." PLoS One 4(9): e7084.

Osafune, K., L. Caron, M. Borowiak, R. J. Martinez, C. S. Fitz-Gerald, Y. Sato, C. A. Cowan, K. R. Chien and D. A. Melton (2008). "Marked differences in differentiation propensity among human embryonic stem cell lines." Nat Biotechnol 26(3): 313-315.

Pauli, A., J. L. Rinn and A. F. Schier (2011). "Non-coding RNAs as regulators of embryogenesis." Nat Rev Genet 12(2): 136-149.

Pellizzoni, L., J. Yong and G. Dreyfuss (2002). "Essential role for the SMN complex in the specificity of snRNP assembly." Science 298(5599): 1775-1779.

Peng, W. T., M. D. Robinson, S. Mnaimneh, N. J. Krogan, G. Cagney, Q. Morris, A. P. Davierwala, J. Grigull, X. Yang, W. Zhang, N. Mitsakakis, O. W. Ryan, N. Datta, V. Jojic, C. Pal, V. Canadien, D. Richards, B. Beattie, L. F. Wu, S. J. Altschuler, S. Roweis, B. J. Frey, A. Emili, J. F. Greenblatt and T. R. Hughes (2003). "A panoramic view of yeast noncoding RNA processing." Cell 113(7): 919-933.

Perobelli, S., F. Alessandrini, G. Zoccatelli, E. Nicolis, A. Beltramello, B. M. Assael and M. Cipolli (2015). "Diffuse alterations in grey and white matter associated with cognitive impairment in Shwachman-Diamond syndrome: evidence from a multimodal approach." Neuroimage Clin 7: 721-731.

Perobelli, S., E. Nicolis, B. M. Assael and M. Cipolli (2012). "Further characterization of Shwachman-Diamond syndrome: psychological functioning and quality of life in adult and young patients." Am J Med Genet A 158A(3): 567-573.

Polo, J. M., E. Anderssen, R. M. Walsh, B. A. Schwarz, C. M. Nefzger, S. M. Lim, M. Borkent, E. Apostolou, S. Alaei, J. Cloutier, O. Bar-Nur, S. Cheloufi, M. Stadtfeld, M. E. Figueroa, D. Robinton, S. Natesan, A. Melnick, J. Zhu, S. Ramaswamy and K. Hochedlinger (2012). "A molecular roadmap of reprogramming somatic cells into iPS cells." Cell 151(7): 1617-1632.

Polo, J. M., S. Liu, M. E. Figueroa, W. Kulalert, S. Eminli, K. Y. Tan, E. Apostolou, M. Stadtfeld, Y. Li, T. Shioda, S. Natesan, A. J. Wagers, A. Melnick, T. Evans and K. Hochedlinger (2010). "Cell type of origin influences the molecular and functional properties of mouse induced pluripotent stem cells." Nat Biotechnol 28(8): 848-855.

Popovic, M., S. Goobie, J. Morrison, L. Ellis, N. Ehtesham, N. Richards, G. Boocock, P. R. Durie and J. M. Rommens (2002). "Fine mapping of the locus for Shwachman-Diamond syndrome at 7q11, identification of shared disease haplotypes, and exclusion of TPST1 as a candidate gene." Eur J Hum Genet 10(4): 250-258.

Pressato, B., R. Valli, C. Marletta, L. Mare, G. Montalbano, F. L. Curto, F. Pasquali and E. Maserati (2015). "Cytogenetic monitoring in Shwachman-Diamond syndrome: a note on clonal progression and a practical warning." J Pediatr Hematol Oncol 37(4): 307-310.

Pressato, B., R. Valli, C. Marletta, L. Mare, G. Montalbano, F. Lo Curto, F. Pasquali and E. Maserati (2012). "Deletion of chromosome 20 in bone marrow of patients with Shwachman- Diamond syndrome, loss of the EIF6 gene and benign prognosis." Br J Haematol 157(4): 503- 505.

Ravera, S., C. Dufour, S. Cesaro, R. Bottega, M. Faleschini, P. Cuccarolo, F. Corsolini, C. Usai, M. Columbaro, M. Cipolli, A. Savoia, P. Degan and E. Cappelli (2016). "Evaluation of energy metabolism and calcium homeostasis in cells affected by Shwachman-Diamond syndrome." Sci Rep 6: 25441.

Repo, H., E. Savilahti and M. Leirisalo-Repo (1987). "Aberrant phagocyte function in Shwachman syndrome." Clin Exp Immunol 69(1): 204-212.

Rigby, R. E. and J. Rehwinkel (2015). "RNA degradation in antiviral immunity and autoimmunity." Trends Immunol 36(3): 179-188.

Rouhani, F., N. Kumasaka, M. C. de Brito, A. Bradley, L. Vallier and D. Gaffney (2014). "Genetic background drives transcriptional variation in human induced pluripotent stem cells." PLoS Genet 10(6): e1004432.

Rujkijyanont, P., S. L. Adams, J. Beyene and Y. Dror (2009). "Bone marrow cells from patients with Shwachman-Diamond syndrome abnormally express genes involved in ribosome biogenesis and RNA processing." Br J Haematol 145(6): 806-815.

Rujkijyanont, P., K. Watanabe, C. Ambekar, H. Wang, A. Schimmer, J. Beyene and Y. Dror (2008). "SBDS-deficient cells undergo accelerated apoptosis through the Fas-pathway." Haematologica 93(3): 363-371.

Ruutu, P., E. Savilahti, H. Repo and T. U. Kosunen (1984). "Constant defect in neutrophil locomotion but with age decreasing susceptibility to infection in Shwachman syndrome." Clin Exp Immunol 57(1): 249-255.

Ryan, T. D., J. L. Jefferies, C. Chin, J. J. Sticka, M. D. Taylor, R. Harris, J. Moore, E. Goodridge, L. Mount, A. A. Bolyard, B. Otto, A. Jones, A. Shimamura, S. Davies and K. Myers (2015). "Abnormal circumferential strain measured by echocardiography is present in patients with Shwachman-Diamond syndrome despite normal shortening fraction." Pediatr Blood Cancer 62(7): 1228-1231.

Sahi, C. and E. A. Craig (2007). "Network of general and specialty J protein chaperones of the yeast cytosol." Proc Natl Acad Sci U S A 104(17): 7163-7168.

Sahoo, T., D. del Gaudio, J. R. German, M. Shinawi, S. U. Peters, R. E. Person, A. Garnica, S. W. Cheung and A. L. Beaudet (2008). "Prader-Willi phenotype caused by paternal deficiency for the HBII-85 C/D box small nucleolar RNA cluster." Nat Genet 40(6): 719-721.

Sauer, M., C. Zeidler, B. Meissner, K. Rehe, A. Hanke, K. Welte, P. Lohse and K. W. Sykora (2007). "Substitution of cyclophosphamide and busulfan by fludarabine, treosulfan and melphalan in a preparative regimen for children and adolescents with Shwachman-Diamond syndrome." Bone Marrow Transplant 39(3): 143-147.

Saunders, E. F., G. Gall and M. H. Freedman (1979). "Granulopoiesis in Shwachman's syndrome (pancreatic insufficiency and bone marrow dysfunction)." Pediatrics 64(4): 515-519.

Savchenko, A., N. Krogan, J. R. Cort, E. Evdokimova, J. M. Lew, A. A. Yee, L. Sanchez-Pulido, M. A. Andrade, A. Bochkarev, J. D. Watson, M. A. Kennedy, J. Greenblatt, T. Hughes, C. H. Arrowsmith, J. M. Rommens and A. M. Edwards (2005). "The Shwachman-Bodian-Diamond syndrome protein family is involved in RNA metabolism." J Biol Chem 280(19): 19213-19220.

Savilahti, E. and J. Rapola (1984). "Frequent myocardial lesions in Shwachman's syndrome. Eight fatal cases among 16 Finnish patients." Acta Paediatr Scand 73(5): 642-651.

Schmidt, S., F. Dethloff, O. Beine-Golovchuk and J. Kopka (2014). "REIL proteins of Arabidopsis thaliana interact in yeast-2-hybrid assays with homologs of the yeast Rlp24, Rpl24A, Rlp24B, Arx1, and Jjj1 proteins." Plant Signal Behav 9(3): e28224.

Sen, S., H. Wang, C. L. Nghiem, K. Zhou, J. Yau, C. Tailor, M. S. Irwin and Y. Dror (2011). "The ribosome-related protein, SBDS, is critical for normal erythropoiesis." Blood 118(24): 6407-6417.

Shammas, C., T. F. Menne, C. Hilcenko, S. R. Michell, B. Goyenechea, G. R. Boocock, P. R. Durie, J. M. Rommens and A. J. Warren (2005). "Structural and mutational analysis of the SBDS protein family. Insight into the leukemia-associated Shwachman-Diamond Syndrome." J Biol Chem 280(19): 19221-19229.

Shao, K., C. Koch, M. K. Gupta, Q. Lin, M. Lenz, S. Laufs, B. Denecke, M. Schmidt, M. Linke, H. C. Hennies, J. Hescheler, M. Zenke, U. Zechner, T. Saric and W. Wagner (2013). "Induced pluripotent mesenchymal stromal cell clones retain donor-derived differences in DNA methylation profiles." Mol Ther 21(1): 240-250.

Shi, D., L. Liang, H. Zheng, G. Cai, X. Li, Y. Xu and S. Cai (2017). "Silencing of long non- coding RNA SBDSP1 suppresses tumor growth and invasion in colorectal cancer." Biomed Pharmacother 85: 355-361.

Shwachman, H., L. K. Diamond, F. A. Oski and K. T. Khaw (1964). "The Syndrome of Pancreatic Insufficiency and Bone Marrow Dysfunction." J Pediatr 65: 645-663.

Smith, A., P. J. Shaw, B. Webster, A. Lammi, K. Gaskin, S. Diaz and P. Sharma (2002). "Intermittent 20q- and consistent i(7q) in a patient with Shwachman-Diamond syndrome." Pediatr Hematol Oncol 19(7): 525-528.

Smith, O. P., I. M. Hann, J. M. Chessells, B. R. Reeves and P. Milla (1996). "Haematological abnormalities in Shwachman-Diamond syndrome." Br J Haematol 94(2): 279-284.

Smith, Z. D., I. Nachman, A. Regev and A. Meissner (2010). "Dynamic single-cell imaging of direct reprogramming reveals an early specifying event." Nat Biotechnol 28(5): 521-526.

Soldner, F., D. Hockemeyer, C. Beard, Q. Gao, G. W. Bell, E. G. Cook, G. Hargus, A. Blak, O. Cooper, M. Mitalipova, O. Isacson and R. Jaenisch (2009). "Parkinson's disease patient-derived induced pluripotent stem cells free of viral reprogramming factors." Cell 136(5): 964-977.

Sridharan, R., J. Tchieu, M. J. Mason, R. Yachechko, E. Kuoy, S. Horvath, Q. Zhou and K. Plath (2009). "Role of the murine reprogramming factors in the induction of pluripotency." Cell 136(2): 364-377.

Stadtfeld, M., E. Apostolou, H. Akutsu, A. Fukuda, P. Follett, S. Natesan, T. Kono, T. Shioda and K. Hochedlinger (2010). "Aberrant silencing of imprinted genes on chromosome 12qF1 in mouse induced pluripotent stem cells." Nature 465(7295): 175-181.

Stepanovic, V., D. Wessels, F. D. Goldman, J. Geiger and D. R. Soll (2004). "The chemotaxis defect of Shwachman-Diamond Syndrome leukocytes." Cell Motil Cytoskeleton 57(3): 158-174.

Strevens, M. J., J. S. Lilleyman and R. B. Williams (1978). "Shwachman's syndrome and acute lymphoblastic leukaemia." Br Med J 2(6129): 18.

Stumpo, D. J., H. E. Broxmeyer, T. Ward, S. Cooper, G. Hangoc, Y. J. Chung, W. C. Shelley, E. K. Richfield, M. K. Ray, M. C. Yoder, P. D. Aplan and P. J. Blackshear (2009). "Targeted disruption of Zfp36l2, encoding a CCCH tandem zinc finger RNA-binding protein, results in defective hematopoiesis." Blood 114(12): 2401-2410.

Suda, T., H. Mizoguchi, Y. Miura, K. Kubota, K. Ikuta, H. Sasaki and T. Nagao (1982). "Hemopoietic colony-forming cells in Shwachman's syndrome." Am J Pediatr Hematol Oncol 4(2): 129-133.

Sulima, S. O., S. P. Gulay, M. Anjos, S. Patchett, A. Meskauskas, A. W. Johnson and J. D. Dinman (2014). "Eukaryotic rpL10 drives ribosomal rotation." Nucleic Acids Res 42(3): 2049- 2063.

Szuts, P., Z. Katona, M. Ilyes, I. Szabo and M. Csato (1984). "Correction of defective chemotaxis with thiamine in Shwachman-Diamond syndrome." Lancet 1(8385): 1072-1073.

Taapken, S. M., B. S. Nisler, M. A. Newton, T. L. Sampsell-Barron, K. A. Leonhard, E. M. McIntire and K. D. Montgomery (2011). "Karotypic abnormalities in human induced pluripotent stem cells and embryonic stem cells." Nat Biotechnol 29(4): 313-314.

Takahashi, K., K. Tanabe, M. Ohnuki, M. Narita, T. Ichisaka, K. Tomoda and S. Yamanaka (2007). "Induction of pluripotent stem cells from adult human fibroblasts by defined factors." Cell 131(5): 861-872.

Takahashi, K. and S. Yamanaka (2006). "Induction of pluripotent stem cells from mouse embryonic and adult fibroblast cultures by defined factors." Cell 126(4): 663-676.

Teo, J. T., R. Klaassen, C. V. Fernandez, R. Yanofsky, J. Wu, J. Champagne, M. Silva, J. H. Lipton, J. Brossard, Y. Samson, S. Abish, M. Steele, K. Ali, U. Athale, L. Jardine, J. P. Hand, E. Tsangaris, I. Odame, J. Beyene and Y. Dror (2008). "Clinical and genetic analysis of unclassifiable inherited bone marrow failure syndromes." Pediatrics 122(1): e139-148.

Thomson, J. A., J. Itskovitz-Eldor, S. S. Shapiro, M. A. Waknitz, J. J. Swiergiel, V. S. Marshall and J. M. Jones (1998). "Embryonic stem cell lines derived from human blastocysts." Science 282(5391): 1145-1147.

Thornley, I., Y. Dror, L. Sung, R. F. Wynn and M. H. Freedman (2002). "Abnormal telomere shortening in leucocytes of children with Shwachman-Diamond syndrome." Br J Haematol 117(1): 189-192.

Tiemann, U., A. G. Marthaler, K. Adachi, G. Wu, G. U. Fischedick, M. J. Arauzo-Bravo, H. R. Scholer and N. Tapia (2014). "Counteracting activities of OCT4 and KLF4 during reprogramming to pluripotency." Stem Cell Reports 2(3): 351-365.

Tiscornia, G., E. L. Vivas and J. C. Izpisua Belmonte (2011). "Diseases in a dish: modeling human genetic disorders using induced pluripotent cells." Nat Med 17(12): 1570-1576.

Todorovic-Guid, M., O. Krajnc, N. Marcun Varda, B. Zagradisnik, N. Kokalj Vokac and A. Gregoric (2006). "A case of Shwachman-Diamond syndrome in a male neonate." Acta Paediatr 95(7): 892-893.

Toiviainen-Salo, S., O. Makitie, M. Mannerkoski, J. Hamalainen, L. Valanne and T. Autti (2008). "Shwachman-Diamond syndrome is associated with structural brain alterations on MRI." Am J Med Genet A 146A(12): 1558-1564.

Toiviainen-Salo, S., M. K. Mayranpaa, P. R. Durie, N. Richards, M. Grynpas, L. Ellis, S. Ikegawa, W. G. Cole, J. Rommens, E. Marttinen, E. Savilahti and O. Makitie (2007). "Shwachman-Diamond syndrome is associated with low-turnover osteoporosis." Bone 41(6): 965-972.

Toiviainen-Salo, S., O. Pitkanen, M. Holmstrom, J. Koikkalainen, J. Lotjonen, K. Lauerma, M. Taskinen, E. Savilahti, J. Smallhorn, O. Makitie and S. Kivisto (2008). "Myocardial function in patients with Shwachman-Diamond syndrome: aspects to consider before stem cell transplantation." Pediatr Blood Cancer 51(4): 461-467.

Toiviainen-Salo, S., M. Raade, P. R. Durie, W. Ip, E. Marttinen, E. Savilahti and O. Makitie (2008). "Magnetic resonance imaging findings of the pancreas in patients with Shwachman- Diamond syndrome and mutations in the SBDS gene." J Pediatr 152(3): 434-436.

Toivonen, S., M. Ojala, A. Hyysalo, T. Ilmarinen, K. Rajala, M. Pekkanen-Mattila, R. Aanismaa, K. Lundin, J. Palgi, J. Weltner, R. Trokovic, O. Silvennoinen, H. Skottman, S. Narkilahti, K. Aalto-Setala and T. Otonkoski (2013). "Comparative analysis of targeted differentiation of human induced pluripotent stem cells (hiPSCs) and human embryonic stem cells reveals

variability associated with incomplete transgene silencing in retrovirally derived hiPSC lines." Stem Cells Transl Med 2(2): 83-93.

Tourlakis, M. E., S. Zhang, H. L. Ball, R. Gandhi, H. Liu, J. Zhong, J. S. Yuan, C. J. Guidos, P. R. Durie and J. M. Rommens (2015). "In Vivo Senescence in the Sbds-Deficient Murine Pancreas: Cell-Type Specific Consequences of Translation Insufficiency." PLoS Genet 11(6): e1005288.

Tourlakis, M. E., J. Zhong, R. Gandhi, S. Zhang, L. Chen, P. R. Durie and J. M. Rommens (2012). "Deficiency of Sbds in the mouse pancreas leads to features of Shwachman-Diamond syndrome, with loss of zymogen granules." Gastroenterology 143(2): 481-492.

Tsai, P. H., I. Sahdev, A. Herry and J. M. Lipton (1990). "Fatal cyclophosphamide-induced congestive heart failure in a 10-year-old boy with Shwachman-Diamond syndrome and severe bone marrow failure treated with allogeneic bone marrow transplantation." Am J Pediatr Hematol Oncol 12(4): 472-476.

Tsangaris, E., R. Klaassen, C. V. Fernandez, R. Yanofsky, E. Shereck, J. Champagne, M. Silva, J. H. Lipton, J. Brossard, B. Michon, S. Abish, M. Steele, K. Ali, N. Dower, U. Athale, L. Jardine, J. P. Hand, I. Odame, P. Canning, C. Allen, M. Carcao, J. Beyene, C. M. Roifman and Y. Dror (2011). "Genetic analysis of inherited bone marrow failure syndromes from one prospective, comprehensive and population-based cohort and identification of novel mutations." J Med Genet 48(9): 618-628.

Tucker, M., M. A. Valencia-Sanchez, R. R. Staples, J. Chen, C. L. Denis and R. Parker (2001). "The transcription factor associated Ccr4 and Caf1 proteins are components of the major cytoplasmic mRNA deadenylase in Saccharomyces cerevisiae." Cell 104(3): 377-386.

Tulpule, A., J. M. Kelley, M. W. Lensch, J. McPherson, I. H. Park, O. Hartung, T. Nakamura, T. M. Schlaeger, A. Shimamura and G. Q. Daley (2013). "Pluripotent stem cell models of Shwachman-Diamond syndrome reveal a common mechanism for pancreatic and hematopoietic dysfunction." Cell Stem Cell 12(6): 727-736.

Tummala, H., A. J. Walne, M. Williams, N. Bockett, L. Collopy, S. Cardoso, A. Ellison, R. Wynn, T. Leblanc, J. Fitzgibbon, D. P. Kelsell, D. A. van Heel, E. Payne, V. Plagnol, I. Dokal and T. Vulliamy (2016). "DNAJC21 Mutations Link a Cancer-Prone Bone Marrow Failure Syndrome to Corruption in 60S Ribosome Subunit Maturation." Am J Hum Genet 99(1): 115- 124.

Vibhakar, R., M. Radhi, S. Rumelhart, D. Tatman and F. Goldman (2005). "Successful unrelated umbilical cord blood transplantation in children with Shwachman-Diamond syndrome." Bone Marrow Transplant 36(10): 855-861.

Virtanen, A., N. Henriksson, P. Nilsson and M. Nissbeck (2013). "Poly(A)-specific ribonuclease (PARN): An allosterically regulated, processive and mRNA cap-interacting deadenylase." Crit Rev Biochem Mol Biol 48(2): 192-209.

Watanabe, K., C. Ambekar, H. Wang, A. Ciccolini, A. D. Schimmer and Y. Dror (2009). "SBDS-deficiency results in specific hypersensitivity to Fas stimulation and accumulation of Fas at the plasma membrane." Apoptosis 14(1): 77-89.

Weeks, S. A., W. P. Shield, C. Sahi, E. A. Craig, S. Rospert and D. J. Miller (2010). "A targeted analysis of cellular chaperones reveals contrasting roles for heat shock protein 70 in flock house virus RNA replication." J Virol 84(1): 330-339.

Weis, F., E. Giudice, M. Churcher, L. Jin, C. Hilcenko, C. C. Wong, D. Traynor, R. R. Kay and A. J. Warren (2015). "Mechanism of eIF6 release from the nascent 60S ribosomal subunit." Nat Struct Mol Biol 22(11): 914-919.

Welting, T. J., S. Mattijssen, F. M. Peters, N. L. van Doorn, L. Dekkers, W. J. van Venrooij, H. A. Heus, L. Bonafe and G. J. Pruijn (2008). "Cartilage-hair hypoplasia-associated mutations in the RNase MRP P3 domain affect RNA folding and ribonucleoprotein assembly." Biochim Biophys Acta 1783(3): 455-466.

Wernig, M., A. Meissner, J. P. Cassady and R. Jaenisch (2008). "c-Myc is dispensable for direct reprogramming of mouse fibroblasts." Cell Stem Cell 2(1): 10-12.

Wessels, D., T. Srikantha, S. Yi, S. Kuhl, L. Aravind and D. R. Soll (2006). "The Shwachman- Bodian-Diamond syndrome gene encodes an RNA-binding protein that localizes to the pseudopod of Dictyostelium amoebae during chemotaxis." J Cell Sci 119(Pt 2): 370-379.

Whaley, S. G., K. E. Caudle, L. Simonicova, Q. Zhang, W. S. Moye-Rowley and P. D. Rogers (2018). "Jjj1 Is a Negative Regulator of Pdr1-Mediated Fluconazole Resistance in Candida glabrata." mSphere 3(1).

Willig, T. N., C. M. Niemeyer, T. Leblanc, C. Tiemann, A. Robert, J. Budde, A. Lambiliotte, E. Kohne, G. Souillet, S. Eber, J. L. Stephan, R. Girot, P. Bordigoni, G. Cornu, S. Blanche, J. M. Guillard, N. Mohandas and G. Tchernia (1999). "Identification of new prognosis factors from the clinical and epidemiologic analysis of a registry of 229 Diamond-Blackfan anemia patients. DBA group of Societe d'Hematologie et d'Immunologie Pediatrique (SHIP), Gesellshaft fur Padiatrische Onkologie und Hamatologie (GPOH), and the European Society for Pediatric Hematology and Immunology (ESPHI)." Pediatr Res 46(5): 553-561.

Winzeler, E. A., D. D. Shoemaker, A. Astromoff, H. Liang, K. Anderson, B. Andre, R. Bangham, R. Benito, J. D. Boeke, H. Bussey, A. M. Chu, C. Connelly, K. Davis, F. Dietrich, S. W. Dow, M. El Bakkoury, F. Foury, S. H. Friend, E. Gentalen, G. Giaever, J. H. Hegemann, T. Jones, M. Laub, H. Liao, N. Liebundguth, D. J. Lockhart, A. Lucau-Danila, M. Lussier, N. M'Rabet, P. Menard, M. Mittmann, C. Pai, C. Rebischung, J. L. Revuelta, L. Riles, C. J. Roberts, P. Ross-MacDonald, B. Scherens, M. Snyder, S. Sookhai-Mahadeo, R. K. Storms, S. Veronneau, M. Voet, G. Volckaert, T. R. Ward, R. Wysocki, G. S. Yen, K. Yu, K. Zimmermann, P. Philippsen, M. Johnston and R. W. Davis (1999). "Functional characterization of the S. cerevisiae genome by gene deletion and parallel analysis." Science 285(5429): 901-906.

Woloszynek, J. R., R. J. Rothbaum, A. S. Rawls, P. J. Minx, R. K. Wilson, P. J. Mason, M. Bessler and D. C. Link (2004). "Mutations of the SBDS gene are present in most patients with Shwachman-Diamond syndrome." Blood 104(12): 3588-3590.

Woods, W. G., J. S. Roloff, J. N. Lukens and W. Krivit (1981). "The occurrence of leukemia in patients with the Shwachman syndrome." J Pediatr 99(3): 425-428.

Wu, L. F., T. R. Hughes, A. P. Davierwala, M. D. Robinson, R. Stoughton and S. J. Altschuler (2002). "Large-scale prediction of Saccharomyces cerevisiae gene function using overlapping transcriptional clusters." Nat Genet 31(3): 255-265.

Wu, S., M. F. Hivert, A. Cardenas, J. Zhong, S. L. Rifas-Shiman, G. Agha, E. Colicino, A. C. Just, C. Amarasiriwardena, X. Lin, A. A. Litonjua, D. L. DeMeo, M. W. Gillman, R. O. Wright, E. Oken and A. A. Baccarelli (2017). "Exposure to Low Levels of Lead in Utero and Umbilical Cord Blood DNA Methylation in Project Viva: An Epigenome-Wide Association Study." Environ Health Perspect 125(8): 087019.

Wu, T., Y. Liu, D. Wen, Z. Tseng, M. Tahmasian, M. Zhong, S. Rafii, M. Stadtfeld, K. Hochedlinger and A. Xiao (2014). "Histone variant H2A.X deposition pattern serves as a functional epigenetic mark for distinguishing the developmental potentials of iPSCs." Cell Stem Cell 15(3): 281-294.

Wu, X., L. Zhang, X. Jin, Y. Fang, K. Zhang, L. Qi and D. Zheng (2016). "Deletion of JJJ1 improves acetic acid tolerance and bioethanol fermentation performance of Saccharomyces cerevisiae strains." Biotechnol Lett 38(7): 1097-1106.

Xu, M. D., S. L. Liu, Y. Z. Feng, Q. Liu, M. Shen, Q. Zhi, Z. Liu, D. M. Gu, J. Yu, L. M. Shou, F. R. Gong, Q. Zhu, W. Duan, K. Chen, J. Zhang, M. Y. Wu, M. Tao and W. Li (2017). "Genomic characteristics of pancreatic squamous cell carcinoma, an investigation by using high throughput sequencing after in-solution hybrid capture." Oncotarget 8(9): 14620-14635.

Xue, S., S. Tian, K. Fujii, W. Kladwang, R. Das and M. Barna (2015). "RNA regulons in Hox 5' UTRs confer ribosome specificity to gene regulation." Nature 517(7532): 33-38.

Yoda, M., D. Cifuentes, N. Izumi, Y. Sakaguchi, T. Suzuki, A. J. Giraldez and Y. Tomari (2013). "Poly(A)-specific ribonuclease mediates 3'-end trimming of Argonaute2-cleaved precursor microRNAs." Cell Rep 5(3): 715-726.

Yoon, A., G. Peng, Y. Brandenburger, O. Zollo, W. Xu, E. Rego and D. Ruggero (2006). "Impaired control of IRES-mediated translation in X-linked dyskeratosis congenita." Science 312(5775): 902-906.

Yu, J., M. A. Vodyanik, K. Smuga-Otto, J. Antosiewicz-Bourget, J. L. Frane, S. Tian, J. Nie, G. A. Jonsdottir, V. Ruotti, R. Stewart, Slukvin, II and J. A. Thomson (2007). "Induced pluripotent stem cell lines derived from human somatic cells." Science 318(5858): 1917-1920.

Zambetti, N. A., E. M. Bindels, P. M. Van Strien, M. G. Valkhof, M. N. Adisty, R. M. Hoogenboezem, M. A. Sanders, J. M. Rommens, I. P. Touw and M. H. Raaijmakers (2015). "Deficiency of the ribosome biogenesis gene Sbds in hematopoietic stem and progenitor cells causes neutropenia in mice by attenuating lineage progression in myelocytes." Haematologica 100(10): 1285-1293.

Zhao, X. Y., W. Li, Z. Lv, L. Liu, M. Tong, T. Hai, J. Hao, C. L. Guo, Q. W. Ma, L. Wang, F. Zeng and Q. Zhou (2009). "iPS cells produce viable mice through tetraploid complementation." Nature 461(7260): 86-90.

Zunder, E. R., E. Lujan, Y. Goltsev, M. Wernig and G. P. Nolan (2015). "A continuous molecular roadmap to iPSC reprogramming through progression analysis of single-cell mass cytometry." Cell Stem Cell 16(3): 323-337.