The Role of Cytosolic 5’-Nucleotidase II (NT5C2) in Drug Resistance
and Relapse of Acute Lymphoblastic Leukemia
Gannie Tzoneva
Submitted in partial fulfillment of the requirements for the degree of Doctor of Philosophy under the Executive Committee of the Graduate School of Arts and Sciences
COLUMBIA UNIVERSITY
2016
© 2016
Gannie Tzoneva
All rights reserved ABSTRACT
The Role of Cytosolic 5’-Nucleotidase II (NT5C2) in Drug Resistance and Relapse
of Acute Lymphoblastic Leukemia
Gannie Tzoneva
Acute lymphoblastic leukemia (ALL) is an aggressive hematological cancer which arises from the malignant transformation of B-cell or T-cell progenitors. Despite recent pioneering improvements in intensified combination chemotherapy, 20% of pediatric and 50% of adult ALL patients present with primary drug-resistant leukemia or develop relapse. Treatment of refractory and relapsed ALL has remained a significant clinical challenge with survival rates following relapse of only 40%, highlighting the need to understand the mechanisms which drive drug resistance and relapse of ALL.
Through extensive sequencing analyses of matched diagnostic, remission and relapsed DNA samples from patients with B-precursor ALL (B-ALL) and T-cell ALL (T-ALL) we have identified recurrent relapse-specific gain-of-function mutations in the cytosolic 5'-nucleotidase II (NT5C2) gene in 25% of relapsed T-ALLs and 6% of relapsed B-ALLs. NT5C2 is a highly conserved, ubiquitously expressed enzyme which regulates intracellular purine nucleotide levels by dephosphorylating purine monophosphates. NT5C2 also dephosphorylates key metabolites in the activation of purine analog prodrugs such as 6-mercaptopurine and 6-thioguanine which are routinely used in the treatment of ALL, allowing purine analog nucleosides to be readily exported out of the cell.
Here we show that mutant NT5C2 proteins have increased 5’-nucleotidase activity and confer resistance to 6-mercaptopurine and 6-thioguanine chemotherapy when expressed in leukemic cells.
Consistently, NT5C2 mutations correlate with early relapse and relapse while under therapy. We present a novel T-ALL conditional inducible knock-in mouse model of the highly recurrent NT5C2 R367Q mutation and show that expression of one Nt5c2 R367Q allele from the endogenous locus in primary T-ALL lymphoblasts induces overt resistance and disease progression under therapy with 6-mercaptopurine in vivo, while surprisingly conferring reduced growth and decreased leukemia initiating activity in the absence of chemotherapy. Metabolically we show that the observed loss of fitness in Nt5c2 R367Q tumors can be explained by a severe depletion of endogenous purine monophosphate metabolites as a result of increased Nt5c2 5’-nucleotidase activity. Consistently, using ultra-sensitive mutation analyses we show that relapse-associated NT5C2 mutations are not detectable at initial disease presentation, indicating that NT5C2-mutant tumor cells are negatively selected by clonal competition in the early stages of disease development and only positively selected under prolonged 6-mercaptopruine chemotherapy which is the backbone treatment for ALL following remission. Our findings present the first known example of chemotherapy resistance and disease progression driven by a tumor clone with decreased leukemia initiating activity, highlighting the intense selective pressure of chemotherapy in the clonal evolution of tumors from diagnosis to relapse.
Through extensive biochemical and structural characterizations of recombinant NT5C2 mutant proteins, we have grouped relapse-specific NT5C2 activating mutations into 3 different classes, each conferring unique enzymatic behavior in basal conditions and in response to allosteric activation, and each with unique structural features which mediate increased 5’-nucleotidase activity. Moreover, we identify a novel auto-regulatory switch-off mechanism of the NT5C2 enzyme involving movement of an unstructured flexible loop, and present the first crystal structure view of the NT5C2 C-terminal acidic tail, implicating it as an auto-inhibitory brake to the allosteric activation of the enzyme. The presence of multiple mutational mechanisms of activating such a highly conserved enzyme, especially in light of the inherent loss of fitness to the tumor cells, indicates a strong convergent evolution towards activating
NT5C2. This is supported by our discovery that patients can harbor multiple leukemic clones with NT5C2 mutations at relapse.
Overall our findings highlight NT5C2 as a major driver of drug resistance and relapse of ALL and pinpoint metabolic susceptibilities which could be exploited therapeutically to target NT5C2-mutant tumors in the future. Our in-depth structural and enzymatic knowledge of mutant NT5C2 proteins will serve as an essential tool in the rational drug development of novel NT5C2 inhibitors with increased specificity and selectivity for mutant NT5C2, while our novel Nt5c2 R367Q knock-in mouse model will serve as a platform for the pre-clinical testing of both NT5C2 inhibitors and alternative compounds selective for Nt5c2-mutant leukemias which can be used for prevention and treatment of relapsed ALL. TABLE OF CONTENTS
List of Figures...... iv List of Tables...... vii
CHAPTER 1: Introduction...... 1 I. Acute Lymphoblastic Leukemia (ALL)...... 1 History...... 1 Epidemiology...... 1 Clinical presentation...... 1 Molecular and genetic basis of ALL...... 2 Inherited genetic determinants of leukemogenesis...... 9 II. Treatment of Acute Lymphoblastic Leukemia...... 9 History of combination chemotherapy...... 10 Contemporary therapy for ALL...... 13 Prognostic factors and risk stratification...... 16 Toxic effects of treatment...... 17 Inherited genomic determinants of therapy response and toxicity...... 19 III. Drug Resistance and Relapse of Acute Lymphoblastic Leukemia...... 20 Risk and prognostic factors...... 21 Biology of drug resistant and relapsed ALL...... 22 Current therapy for relapsed ALL...... 27 IV. Novel Agents in the Treatment of ALL...... 28 Improvements in current chemotherapy...... 28 Precision medicine...... 29 Immunotherapy...... 30 V. The Role of Cytosolic 5’-Nucleotidase II (NT5C2) in Health and Disease...... 32 Physiological role...... 32 Structural basis of NT5C2 activity and allosteric regulation...... 34 Aberrant activity of NT5C2 in human disease...... 38 VIII. Specific Aims...... 40
CHAPTER 2: Materials and Methods...... 41 I. Chapter 3...... 41 Patient samples...... 41 Whole-exome capture and next generation sequence analysis...... 41 Mutation validation and analysis of recurrence...... 42
i Structural depiction and analysis (Z. Carpenter) ...... 43 Site-directed mutagenesis...... 44 Cell lines...... 44 Lentiviral production and infection...... 44 Western blot...... 44 Cell viability and chemotherapy drug response...... 44 Recombinant protein production and purification...... 45 5′-nucleotidase assays...... 45 Single cell PCR analysis...... 45 Statistical analyses...... 46 II. Chapter 4...... 46 Mouse procedures and generation of Nt5c2 R367Q conditional knock-in...... 46 Generation of NOTCH1-induced leukemias...... 47 In vivo mouse experiments...... 47 Drugs and administration...... 48 Cell culture...... 49 Retroviral and lentiviral infections...... 49 Genotyping of conditional Nt5c2 R367Q induction...... 49 In vitro cell viability and chemotherapy responses...... 50 Patient samples...... 50 Targeted ultra-deep sequencing of diagnostic and remission samples...... 50 Quantitative NT5C2 p.R367Q allele specific qPCR assay...... 51 Metabolomic analyses...... 52 Statistical analyses...... 52 III. Chapter 5...... 52 Recombinant protein production and purification...... 52 5′-nucleotidase assays...... 54 Structural modeling analyses (Z. Carpenter) ...... 54 Crystallization and structure determination (F. Forouhar) ...... 55
CHAPTER 3: Activating mutations in the NT5C2 nucleotidase gene drive chemotherapy resistance in relapsed ALL...... 57 Introduction...... 57 Results...... 58 T-ALLs undergo clonal evolution from diagnosis to relapse...... 58 NT5C2 mutations are recurrently found in relapsed ALL and associate with early relapse and relapse while on treatment...... 59
ii Relapse-associated NT5C2 mutations are gain-of-function alleles which increase NT5C2 nucleotidase activity...... 61 Relapse-associated NT5C2 mutations confer resistance to 6-mercaptopurine and 6-thioguanine in leukemic cells...... 64 Relapsed ALL tumors exhibit convergent evolution towards NT5C2 activation...... 65 Discussion...... 68
CHAPTER 4: Chemotherapy resistance-driving NT5C2 mutations impair leukemia initiating cell activity in relapsed acute lymphoblastic leukemia...... 70 Introduction...... 70 Results...... 72 Establishment of a conditional inducible Nt5c2 R367Q model of T-ALL...... 72 Conditional Nt5c2 R367Q primary murine T-ALLs show marked resistance to thiopurine chemotherapy in vitro...... 75 Mice bearing Nt5c2 R367Q T-ALLs show resistance and disease progression under 6-MP chemotherapy in vivo...... 76 Nt5c2 R367Q primary murine leukemias have decreased fitness and leukemia initiating activity...78 NT5C2 mutations are not present in primary leukemic tumors at diagnosis...... 79 Nt5c2 R367Q primary leukemic cells have depleted pools of endogenous purine metabolites...... 80 Discussion...... 82
CHAPTER 5: Unique structural mechanisms of NT5C2 mutational hyperactivation in relapsed ALL and discovery of a novel NT5C2 auto-regulatory switch...... 84 Introduction...... 84 Results...... 86 Activating NT5C2 mutations in relapsed ALL comprise three unique enzymatic classes...... 86 Class I NT5C2 mutations are neomorphic alleles which directly mimic allosteric activation...... 89 Class II mutations conserve the architecture of the allosteric activation center of NT5C2 but block an auto-regulatory switch-off mechanism...... 93 The Class III truncating mutation NT5C2 Q523* removes a C-terminal acidic regulatory patch which serves as a brake to the allosteric activation of NT5C2...... 100 Discussion...... 105
CHAPTER 6: Conclusions...... 108 References...... 111 Appendix: Supplementary Figures and Tables...... 147
iii LIST OF FIGURES
CHAPTER 1 Figure 1.1. Proposed sequential acquisition of genetic alterations contributing to the pathogenesis of B- ALL...... 3 Figure 1.2. The NOTCH1 signaling pathway...... 5 Figure 1.3. Prevalence of NOTCH1 activating mutations in T-ALL...... 6 Figure 1.4. Overall survival among children with ALL who were enrolled in Children’s Cancer Group and Children’s Oncology Group clinical trials, 1968-2009...... 13 Figure 1.5. Model of clonal evolution of relapsed ALL...... 25 Figure 1.6. Purine de novo biosynthesis and salvage pathways, highlighting metabolic conversions catalyzed by NT5C2...... 33 Figure 1.7. NT5C2 is an HAD family enzyme...... 35 Figure 1.8. Mechanism of activation of NT5C2...... 37
CHAPTER 3 Figure 3.1. NT5C2 mutations in relapsed ALL...... 60 Figure 3.2. Structure-function analysis of the NT5C2 K359Q mutant protein...... 62 Figure 3.3. Structure-function analysis of the NT5C2 S360P mutant protein...... 63 Figure 3.4. Increased 5'-IMP nucleotidase activity in NT5C2 mutant proteins...... 63 Figure 3.5. Expression of NT5C2 mutations in CCRF-CEM ALL cells induces resistance to chemotherapy with 6-MP and 6-TG...... 65 Figure 3.6. Expression of NT5C2 mutations in CUTLL1 ALL cells induces resistance to chemotherapy with 6-MP and 6-TG...... 66 Figure 3.7. Expression of NT5C2 mutations in CCRF-CEM ALL cells does not induce resistance to chemotherapy with nelarabine or araG...... 66
CHAPTER 4 Figure 4.1. NT5C2 inactivates the cytotoxic metabolites of 6-mercaptopurine (6-MP) and 6-thioguanine (6-TG) ...... 71 Figure 4.2. Conditional knock-in targeting of Nt5c2 in mouse embryonic stem cells...... 73 Figure 4.3. Experimental strategy for developing conditional inducible Nt5c2 R367Q primary mouse T- ALL tumors and assessment of the role of Nt5c2 R367Q in established T-ALL in the presence and absence of chemotherapy...... 74 Figure 4.4. Expression of a single Nt5c2 R367Q allele from the endogenous mouse locus leads to overt 6-MP and 6-TG resistance in vitro...... 75
iv Figure 4.5. Nt5c2 R367Q induces T-ALL drug resistance and disease progression under 6-MP therapy in vivo...... 77 Figure 4.6. Nt5c2 R367Q tumor cells have reduced proliferation and leukemia initiating cell activity in the absence of chemotherapy...... 78 Figure 4.7. NT5C2 R367Q allele specific PCR...... 80 Figure 4.8. Metabolic profiling analysis shows a significant metabolic disadvantage in Nt5c2wt/R367Q cells, with decreased levels of Nt5c2 purine substrates and increased export of downstream purine metabolites...... 81
CHAPTER 5 Figure 5.1. NT5C2 mutant proteins have altered responses to allosteric activation...... 87 Figure 5.2. NT5C2 mutant proteins can be separated into three classes based on their enzymatic behavior and response to allosteric activation by ATP...... 89 Figure 5.3. Crystal structure of NT5C2 K359Q shows a shift in the hydrogen bonding network at the allosteric region resulting in direct helix A stabilization and formation of the active enzyme conformation in the absence of allosteric activator or substrate...... 90 Figure 5.4. Class I mutants NT5C2 K359Q and NT5C2 L375F have a tighter dimer interface than wildtype NT5C2...... 91 Figure 5.5. In silico model and crystal structure of NT5C2 L375F show additional stabilizing π-π stacking interactions at the monomer-monomer interface...... 92 Figure 5.6. Crystal structure of NT5C2 L375F shows a shift in the hydrogen bonding network at the allosteric region resulting in direct helix A stabilization and formation of the active enzyme conformation in the absence of allosteric activator or substrate...... 92 Figure 5.7. Class I NT5C2 mutations are neomorphic alleles which directly alter the architecture of the NT5C2 allosteric region, resulting in constitutive NT5C2 activation in the absence of allosteric activator...... 93 Figure 5.8. Class II NT5C2 mutations map to two regions on the homodimer functional unit...... 94 Figure 5.9. Flexible unstructured loop of NT5C2 enters into the basic intermonomeric pocket which opens up upon NT5C2 activation...... 95 Figure 5.10. Following NT5C2 activation, D407 in the flexible loop invades into the basic pocket and interacts with K361 on the neighboring subunit, resulting in helix A destabilization and enzyme de- activation...... 96 Figure 5.11. Novel crystal structure of inactive wildtype NT5C2 shows the flexible loop interacting with R238 on the neighboring monomer...... 97 Figure 5.12. NT5C2 R238W structure shows formation of a dense hydrophobic plug near the entrance of the intermonomeric basic cavity...... 98
v Figure 5.13. Structure of active NT5C2 R367Q active structure shows conservation of hydrogen bonding interactions supporting allosteric helix A...... 98 Figure 5.14. Class II NT5C2 mutations show strict conservation of the allosteric architecture surrounding helix A while removing positive charges which interact with the auto-inhibitory flexible loop of the neighboring monomer...... 99 Figure 5.15. Antibodies against the NT5C2 flexible loop increase 5’-nucleotidase activity in the presence of ATP...... 100 Figure 5.16. Antibodies against the NT5C2 C-terminal acidic tail increase 5’-nucleotidase activity in the presence of ATP...... 101 Figure 5.17. Novel crystal structure of inactive full length NT5C2 R367Q shows the C-terminus wrapped around the neighboring monomer with the acidic tail tucked into the basic intermonomeric pocket.....102 Figure 5.18. In the inactive (apo) state, the NT5C2 acidic C-terminal tail interacts with basic residues from both adjacent NT5C2 dimer subunits, including R367 in the allosteric region...... 103 Figure 5.19. Interchanging occupation of a surface groove on NT5C2 by the N-terminus of the same monomer and the C-terminus of the adjacent monomer in active and inactive enzyme states respectively...... 103 Figure 5.20. Cys547 in the NT5C2 C-terminus can form a disulfide bridge with Cys121 on the neighboring monomer under oxidizing conditions...... 104
APPENDIX I. Chapter 3 Supplementary Figure A3.1. TP53, RPL11 and BANP mutations in relapsed pediatric ALL...... 147 Supplementary Figure A3.2. Western blot analysis of NT5C2 expression in CCRF-CEM and CUTLL1 cells transduced with wild type and mutant NT5C2 expressing lentiviruses...... 148 Supplementary Figure A3.3. . 5'-IMP nucleotidase activity in NT5C2 mutant proteins...... 148 Supplementary Figure A3.4. Expression of NT5C2 R238W, L375F in ALL cells does not induce resistance to 6-MP or 6-TG...... 149 II. Chapter 4 No Supplementary Figures III. Chapter 5 Supplementary Figure A5.1. C-terminally truncated recombinant NT5C2 proteins used in crystallographic studies retain activating phenotypes of relapse-associated NT5C2 mutations...... 161
vi LIST OF TABLES
CHAPTER 1 Table 1.1. List of commonly used chemotherapeutic agents in the treatment of ALL...... 14 Table 1.2. Toxic side effects of chemotherapy used in the treatment of ALL...... 18
CHAPTER 2 No Tables.
CHAPTER 3 Table 3.1. Concurrent NT5C2 mutations identified in relapsed ALL...... 67 Table 3.2. Clonal cDNA analysis of relapsed ALL samples harboring concurrent NT5C2 R238W and L375F mutations...... 67 Table 3.3. Single-cell sequencing analysis of a relapsed ALL tumor sample harboring concurrent NT5C2 R238W and L375F mutations...... 68
CHAPTER 4 Table 4.1. Targeted deep sequencing of NT5C2 in matching diagnostic and remission DNA samples from patients with NT5C2-mutated relapsed ALL...... 79 Table 4.2. Allele specific PCR analysis of NT5C2 R367Q presence in matching diagnostic and remission DNA samples from patients with NT5C2-mutated relapsed ALL...... 80
APPENDIX I. Chapter 3 Supplementary table A3.1. Summary of patient samples and genetic analyses...... 149 Supplementary Table A3.2. Mutations identified by exome sequencing in diagnosis and relapse T- ALL...... 150 Supplementary table A3.3. Genomic regions of LOH in diagnostic and relapsed T-ALLs analyzed by whole exome sequencing...... 151 Supplementary Table A3.4. NT5C2 mutations in T-ALL...... 152 Supplementary Table A3.5. NT5C2 mutations in B-ALL...... 153 Supplementary Table A3.6. Correlative clinical data on NT5C2-mutated relapsed T-ALLs treated in AIEOP clinical trials...... 154 Supplementary Table A3.7. Correlative clinical and molecular data on NT5C2-mutated relapsed T-ALLs in BFM based protocols...... 154 Supplementary Table A3.8. Correlative clinical data on rescue treatment for NT5C2-mutated relapsed T- ALLs in BFM based protocols...... 155
vii Supplementary Table A3.9. Association of NT5C2 mutations with clinical characteristics, response,
outcome and genetics in T-ALL patients treated in BFM based protocols...... 156 Supplementary Table A3.10. Association of frontline treatment with NT5C2 mutations...... 158 Supplementary Table A3.11. Multivariate binary logistic regression with time of relapse (on/off treatment) as outcome variable...... 158 Supplementary Table A3.12. Nucleoside analog IC50 values (μM) of T-ALL cell lines expressing relapse- associated NT5C2 mutant alleles...... 159 II. Chapter 4 Supplementary Table A4.1. Nucleoside analog IC50 values (μM) of isogenic Nt5c2 wildtype and Nt5c2 R367Q primary murine T-ALL tumors...... 160 Supplementary Table A4.2. Results of Leukemia Initiating Cell assay performed by limiting dilution transplantations of isogenic Nt5c2 wildtype and Nt5c2 R367Q primary mouse T-ALL tumor cells...... 160 III. Chapter 5 Supplementary Table A5.1. Summary of crystallographic information for NT5C2 537X proteins...... 161 Supplementary Table A5.2. Summary of crystallographic information for full length NT5C2 proteins.....162
viii ACKNOWLEDGMENTS
“If I have seen further than others, it is by standing upon the shoulders of giants.”
Isaac Newton
First and foremost, I would like to thank my adviser Dr. Adolfo Ferrando for his advice, support and mentorship over the years, for his confidence in my abilities and for providing me the opportunity to grow as a scientist while developing this exciting multi-disciplinary project. Through his guidance Dr.
Ferrando has taught me the importance of taking bold risks in research and of asking the less obvious scientific questions. It has been a great privilege to learn from his incredible intellect, his enthusiasm for discovery and his talent for choosing the best direction when exploring the unknown.
I would like to thank my thesis committee, Dr. Ramon Parsons, Dr. Shan Zha and Dr. Laura
Pasqualucci, for their continued guidance, advice and support, as well as Dr. Elli Papaemmanuil for participating as external examiner for my defense.
I would like to thank Dr. Raul Rabadan and his group, especially Dr. Hossein Khiabanian for the exome sequencing analyses and Dr. Zachary Carpenter for the structural modeling analyses. I would also like to thank Dr. Liang Tong for the highly successful NT5C2 crystallography collaboration and especially
Dr. Farhad Forouhar for the crystallographic analyses and in-depth discussions on the complex structural regulation of NT5C2 activity. In addition, I would like to thank Dr. Victor Lin at the Columbia Transgenic
Mouse Shared Resource for his help in the development of the Nt5c2 R367Q knock-in mouse. A big thank you also goes to our collaborators overseas for providing us with patient samples and especially Dr.
Renate Kirschner-Schwabe and Dr. Jana Hof for the clinical correlative analyses.
I would like to thank all the members of the Ferrando lab, past and present, for their help and advice over the years and for making the lab a stimulating and supportive environment in which to train as a scientist. I would especially like to thank Dr. Ana da Silva Almeida for her continued advice on protein purification and characterization, Dr. Arianne Perez Garcia for her guidance and help with our first publication on NT5C2, Kevin Xu for his assistance with mouse experiments, Dr. Lucile Couronné and Dr.
Marta Sanchez-Martin for assistance with deep sequencing studies and Dr. Alberto Ambesi-Impiombato
ix for sequencing analyses. Finally I would like to give special thanks to Chelsea Dieck for taking up the torch for the NT5C2 project and for her continuous willingness to learn and readiness to help. Chelsea’s positive disposition, assistance and encouragement have been invaluable during the last year of my doctoral studies, and it has been a great pleasure passing on my knowledge to her and working alongside her.
I would also like to thank Dr. Ronald Liem of the Pathobiology and Molecular Medicine program for his advice and support and Dr. Maria Luisa Sulis for sharing her time in the clinic as part of the Med into Grad initiative. A special thank you to my program coordinator Zaia Sivo, whose continuous care and guidance from my very first day in the US have brought me great comfort as an international student.
Finally, I would like to thank my family, loved ones and friends near and far for the unending support, encouragement and advice, especially through the most challenging times. Thank you for cheering me on through thick and thin and for always reminding me that I can achieve anything to which I set my mind. I am grateful to have walked this journey with you by my side.
x DEDICATION
"An investment in knowledge pays the best interest."
Benjamin Franklin
This work is dedicated to my parents. You endowed me with a strong set of values, a love of learning and
a determination to use my knowledge to help others. You taught me to think critically, to work hard to
achieve my goals, and to never give up in the face of challenge. Your unending love, support,
encouragement and guidance are the foundation of my strength.
xi
CHAPTER 1
Introduction
I. Acute Lymphoblastic Leukemia (ALL)
History
Acute lymphoblastic leukemia (ALL) is an aggressive hematological cancer which results from the malignant transformation of immature T-cell or B-cell progenitors in the bone marrow (Inaba, Greaves et al. 2013). Leukemia means “white blood” in Greek and reports in the literature exist as early as 1800s, including Cullen’s description in 1811 of a case of “splenitis acutus in which the serum of the blood drawn from the arm had the appearance of milk” (Cullen 1811). The most notable and clear descriptions of leukemia came simultaneously in 1845 in two independent reports by John Bennett in Scotland and
Rudolph Virchow in Germany. Bennett was the first to show illustrations of leukemic cells, describing the suppuration of the blood and terming it “leukocythemia” (Bennett 1845), while Virchow described a similar case linking it to splenic enlargement and referring to the disease as “Weisses Blut” and later “leukemia”
(Virchow 1845).
Epidemiology
ALL affects both adults and children, with approximately 6000 cases in the United States diagnosed annually (1.7 per 100,000 persons) and a peak incidence between 3 to 5 years of age (Smith,
Seibel et al. 2010). It is the most common malignancy in children and adolescents (4-5 per 100,000), accounting for approximately 30% of all childhood cancers and being the greatest cancer-related cause of death before 20 years of age (Linabery and Ross 2008, Smith, Seibel et al. 2010). B-precursor ALL (B-
ALL) is the more prevalent form, with T-cell ALL (T-ALL) accounting for 15% of pediatric and 25% of adult
ALL cases and being more prevalent in males (55%) (Linabery and Ross 2008).
Clinical presentation
Patients with ALL typically present with fatigue, pallor, easy bruising or bleeding and possibly infection. Analysis of their blood shows a high percentage of blasts, accompanied by anemia, thrombocytopenia and neutropenia. There is commonly diffuse infiltration of the bone marrow by
1
Chapter 1 immature B- or T-cell precursor cells, as diagnosed by microscopy and flow cytometric immunophenotype assessment. Frequently this is accompanied by leukemic infiltration of the spleen, liver, lymph nodes, mediastinum (with pleural effusions) and in some cases the central nervous system (CNS) or testicles
(Hunger and Mullighan 2015).
Molecular and genetic basis of ALL
Early cytogenetic studies and recent advances in genomics, transcriptomics and epigenomics are providing us with great insight into the biology of ALL. Genetically ALL is a heterogeneous disease, with a number of subtypes classified recently (Pui, Yang et al. 2015). Most cases harbor large structural genetic alterations such as aneuploidy (gains or losses of chromosomes) and chromosomal translocations resulting in the deregulated expression of hematopoietic regulators, tumor suppressors or oncogenes, or in the expression of novel fusion proteins with leukemogenic potential. Additional cooperating genetic hits in the form of DNA sequence mutations are also frequently observed.
In B-ALL the two most common alterations are hyperdiploidy (>50 chromosomes) in 25% of cases and the t(12;21)(p13;q22) translocation which encodes ETV6-RUNX1 in 25% of cases (Shurtleff,
Buijs et al. 1995). This rearrangement alters the activities of the ETV6 and RUNX1 transcription factors which are important regulators of hematopoiesis, and predisposes B-precursor cells to leukemic transformation (Morrow, Horton et al. 2004). Other B-ALL translocations include the BCR-ABL1
Philadelphia (Ph) chromosome (3% of pediatric and 25% of adult cases) which results in oncogenic constitutive tyrosine kinase signaling (Ribeiro, Abromowitch et al. 1987) , TCF3-PBX1 (5%) which disrupts hematopoietic differentiation (Hunger 1996, Lu and Kamps 1997), and translocations of MLL (KMT2A) with various partners (5%) (Meyer, Schneider et al. 2006). MLL encodes a histone H3K4 methyltransferase that plays an essential role during early development and hematopoiesis by transcriptional co-activation of HOX genes, and its chromosomal rearrangements result in epigenetic dysregulation of lymphoblast self-renewal (Armstrong, Staunton et al. 2002). MLL translocations are very common in ALL patients younger than 1 year (75%) and occur with very few additional genetic alterations
(Andersson, Ma et al. 2015).
2 Chapter 1
The advent of array comparative genomic hybridization, single nucleotide polymorphism (SNP) microarrays and next generation sequencing allowed for the identification of submicroscopic genetic alterations that can co-operate with the large structural lesions in ALL to drive leukemogenesis. In over two thirds of B-ALL these newly discovered targets of mutation/rearrangement/deletion include genes encoding transcriptional regulators of B-cell differentiation such as PAX5 (in over 30% of cases), EBF1 and IKZF1-3, as well as signaling components, cell cycle and apoptosis regulators, and tumor suppressors (Figure 1.1) (Bousquet, Broccardo et al. 2007, Nebral, Denk et al. 2009, Zhang, Mullighan et al. 2011).
Figure 1.1. Proposed sequential acquisition of genetic alterations contributing to the pathogenesis of B- ALL. Some common inherited variants or, rarely, deleterious germline mutations confer a predisposition to ALL. Initiating lesions, commonly translocations, are acquired in a lymphoid progenitor. Secondary sequence mutations and structural genetic alterations contribute to an arrest in lymphoid development and perturbation of multiple cellular pathways, resulting in clinically manifest leukemia. PI3K denotes phosphatidylinositol 3- kinase. Adapted from Hunger and Mullighan, 2015.
Transcriptional profiling studies allowed for the identification of a novel subtype of B-ALL which exhibits a gene expression profile very similar to that of BCR-ABL1 rearranged B-ALL but does not bear the BCR-ABL1 fusion. This subtype, named BCR-ABL1-like (Ph-like), accounts for 10-15% of B-ALL cases and frequently exhibits deletions or mutations of the IKZF1 gene (encoding the lymphoid transcription factor IKAROS) and rearrangements in the CRLF2 cytokine receptor gene which lead to
3 Chapter 1 overexpression of the receptor on the cell surface (Den Boer, van Slegtenhorst et al. 2009, Mullighan,
Collins-Underwood et al. 2009, Russell, Capasso et al. 2009). A recent study identified a diverse set of genetic alterations in Ph-like B-ALL which can activate tyrosine kinase and cytokine receptor signaling pathways, including fusions involving the ABL-class kinases (e.g. NUP214-ABL1, EBF1-PDGFRB) and fusions/deletions/mutations which activate JAK-STAT signaling (e.g. IL7R mutations, BCR-JAK2)
(Roberts, Morin et al. 2012).
Other characteristic subtypes of B-ALL include intrachromosomal amplification of chromosome 21
(iAMP21) in 2% of cases resulting in gain of at least three copies of RUNX1 (Harewood, Robinson et al.
2003), hypodiploid ALL (3%) with less than 44 chromosomes (Harrison, Moorman et al. 2004) and focal deletions in part of the ERG transcription factor gene in 3% of cases (Clappier, Auclerc et al. 2014).
In T-ALL the most prominent alterations leading to T-cell transformation involve constitutive activation of NOTCH1 signaling (Weng, Ferrando et al. 2004). NOTCH1 is a ligand-activated heterodimeric receptor that is a crucial regulator of cellular lineage commitment during hematopoietic development, being absolutely required for the development of T-cells from hematopoietic progenitors
(Greenwald 1998, Pui, Allman et al. 1999, Radtke, Wilson et al. 1999). The extracellular domain of
NOTCH1 interacts with Delta-like and Jagged ligands on neighboring cells, which results in a conformational change that triggers two consecutive proteolytic cleavages of the receptor, first by an
ADAM metalloprotease and then by the γ-secretase complex, which releases the intracellular domains of
NOTCH1 (ICN1) (Figure 1.2). ICN1 translocates to the nucleus where together with RBPJ/CSL it binds to
NOTCH1 recognition motifs on DNA and recruits the transcriptional co-activator MAML1, leading to expression of NOTCH1 target genes (Figure 1.2) (Tzoneva and Ferrando 2012). The C-terminus of ICN1 contains a PEST domain (rich in proline, glutamic acid, serine and threonine), which targets the activated receptor for polyubiquitination and subsequent proteasomal degradation in the nucleus, thereby acting as a switch-off mechanism of the NOTCH1 signal (Rechsteiner 1988, Fryer, White et al. 2004).
The first observation of increased NOTCH1 signaling in the pathogenesis of T-ALL came from the analysis of a rare, recurrent t(7;9)(q34;q34.3) chromosomal translocation in T-ALL, which placed an N- terminally truncated allele of NOTCH1 next to the TCRB (T-cell receptor beta) locus, resulting in aberrant
4 Chapter 1
Figure 1.2. The NOTCH1 signaling pathway. Schematic representation of NOTCH1 activation. Interaction of the NOTCH1 receptor with Delta- like and Jagged ligands expressed on the surface of neighboring cells triggers the proteolytic cleavage of the receptor, first by and ADAM metalloprotease (S2 cleavage) and subsequently by the γ-secretase complex (S3 cleavage), which releases the intracellular domains of NOTCH1 (ICN1) from the membrane. ICN1 translocates to the nucleus and interacts with RBPJ/CSL DNA binding protein and recruits the MAML1 coactivator to activate the expression of NOTCH1 target genes. Reproduced from Tzoneva and Ferrando, 2012.
expression of a constitutively active form of NOTCH1 (Ellisen, Bird et al. 1991). The exact location of the translocation breakpoint and the translation initiation site in each case results in expression of either membrane-bound derivative NOTCH1 receptors that are constitutively processed by γ-secretase, or intracellular proteins without a transmembrane domain (akin to ICN1) that are constitutively active and independent of γ-secretase processing (Ellisen, Bird et al. 1991, Palomero, Barnes et al. 2006).
Subsequent mouse studies demonstrated that expression of N-terminally truncated, constitutively active
Notch1 in hematopoietic progenitors leads to potent induction of murine T-ALL (Pear, Aster et al. 1996).
The full significance of NOTCH1 in T-ALL was revealed after genetic sequencing revealed the presence of NOTCH1 activating mutations in over 60% of human T-ALLs (Weng, Ferrando et al. 2004).
The majority of the mutations disrupt the architecture of the heterodimerization (HD) domain and its association with the LNR repeats, resulting in increased or constitutive cleavage by ADAM metalloproteases (Malecki, Sanchez-Irizarry et al. 2006, Gordon, Roy et al. 2009) (Figure 1.3).
Juxtamembrane expansion mutations similarly increase enzymatic processing of NOTCH1 by displacing the HD-LNR complex away from the cell membrane (Sulis, Williams et al. 2008).
5 Chapter 1
Figure 1.3. Prevalence of NOTCH1 activating mutations in T-ALL. The majority of NOTCH1 activating mutations in T-ALL are NOTCH1 HD and ΔPEST. An additional 1% of patients harbor translocations involving NOTCH1 and TCR loci. About 15% of T-ALL cases harbor deletions or mutations in FBXW7, which impair the degradation of activated NOTCH1 in the nucleus and are functionally related to NOTCH1 ΔPEST alleles. Reproduced from Tzoneva and Ferrando, 2012.
Another important class of NOTCH1 mutations consists of frameshifts or nonsense nucleotide substitutions which lead to partial or complete truncations of the PEST domain (NOTCH1 ΔPEST)
(Figure 1.3)(Weng, Ferrando et al. 2004) . The loss of the PEST domain prevents NOTCH1 targeting for proteasomal degradation by the FBXW7 ubiquitin ligase, resulting in increased levels of activated
NOTCH1 (Thompson, Buonamici et al. 2007). More recent studies found that 15% of T-ALLs harbor loss- of-function mutations or deletions in FBXW7 itself, resulting in impaired substrate recognition (Figure 1.3)
(O'Neil, Grim et al. 2007, Thompson, Buonamici et al. 2007, Asnafi, Buzyn et al. 2009). Hence the
FBXW7 mutations have the same biological effect as the NOTCH1 ΔPEST mutations by promoting increased stability of activated NOTCH1 (O'Neil, Grim et al. 2007, Thompson, Buonamici et al. 2007), but since FBXW7 can also target a number of other oncoproteins for degradation such as c-MYC, JUN and mTOR (Nateri, Riera-Sans et al. 2004, Welcker, Orian et al. 2004, Yada, Hatakeyama et al. 2004, Mao,
Kim et al. 2008), the consequences of FBXW7 inactivation are likely much broader. Notably, 15% of T-
ALLs carry both NOTCH1 HD and ΔPEST mutations in cis (Weng, Ferrando et al. 2004), while 5% have a
NOTCH1 HD and a FBXW7 mutation (O'Neil, Grim et al. 2007, Thompson, Buonamici et al. 2007)(Figure
1.3). Both of these combinations of constitutive activation and an impaired switch-off mechanism of
NOTCH1 result in even higher levels of NOTCH1 signaling, illustrating the evolutionary pressure for
NOTCH1 during T-cell transformation (Weng, Ferrando et al. 2004, Thompson, Buonamici et al. 2007).
Despite the high prevalence of oncogenic NOTCH1 activating alterations, there is a second highly notable genetic alteration found in 70% of T-ALLs, namely deletion of the CDKN2A locus which encodes both the p16/INK4A and the p14/ARF tumor suppressor genes (Hebert, Cayuela et al. 1994, Ferrando,
Neuberg et al. 2002). Thus the majority of T-ALLs likely arise from the cooperation of NOTCH1 activation
6 Chapter 1 and loss of p16/INK4A and p14/ARF.
In addition, similarly to the NOTCH1 t(7;9)(q34;q34.3) translocation, T-ALLs can harbor other chromosomal rearrangements which place transcription factor oncogenes such as TAL1, TAL2, LYL1,
LMO1, LMO2, TLX1, TLX3, NKX2.1, NKX2.2, NKX2.5, MYC and MYB under the control of T-cell specific enhancers in the TCRB or TCRA-TCRD loci, leading to strong overexpression of these oncogenes in T- cell progenitors (Van Vlierberghe and Ferrando 2012). The TLX3 locus is also (more frequently) translocated to T-cell regulatory regions near the BCL11B locus (Bernard, Busson-LeConiat et al. 2001), while TAL1 overexpression and LMO2 activation can also arise as a result of other small intrachromosomal deletions (Aplan, Lombardi et al. 1990, Van Vlierberghe, van Grotel et al. 2008).
As observed with B-ALL, there are also translocations in T-ALL which result in expression of transcription factor oncogene fusions, e.g.MLL-AF4 and MLL-MLLT1 in about 5% of T-ALLs, PICALM-
MLLT10 in 5-10% and the rare SET-NUP214 (Ferrando and Look 2000).
Notably, while MYC is activated by translocation only in 1% of T-ALLs, its activity is prominently upregulated in many T-ALLs as a result of being a direct target of NOTCH1 (Palomero, Lim et al. 2006,
Margolin, Palomero et al. 2009, Herranz, Ambesi-Impiombato et al. 2014). TAL1 is a key regulator of hematopoietic development and, overall, as a result of various rearrangements it is aberrantly expressed in 60% of T-ALL cases and associated with a molecular signature indicating developmental arrest at the late stage of thymocyte development, with CD4 and CD8 double positivity (Bash, Hall et al. 1995,
Ferrando, Neuberg et al. 2002). TLX1 and TLX3 are both HOX-family transcription factors crucial for development (Dear, Sanchez-Garcia et al. 1993). Leukemias with aberrant TLX1 and TLX3 expression have similar mechanisms of transformation and bear specific cooperating mutations that are nearly TLX- exclusive, such as expression of the NUP214-ABL1 fusion oncogene and mutations in the PTPN2, WT1 and PHF6 tumor suppressors (Graux, Cools et al. 2004, Tosello, Mansour et al. 2009, Kleppe, Lahortiga et al. 2010, Renneville, Kaltenbach et al. 2010, Van Vlierberghe, Palomero et al. 2010). Other tumor suppressors are also targets of inactivation in T-ALL, including monoallelic or biallelic deletions in LEF1 in
15% of cases (Gutierrez, Sanda et al. 2010) and frequent loss of function mutations and heterozygous deletions of BCL11B (De Keersmaecker, Real et al. 2010, Gutierrez, Kentsis et al. 2011).
7 Chapter 1
Recent studies have unveiled novel alterations in crucial chromatin remodeling factors in T-ALL, including loss of function mutations and deletions in EZH2 and SUZ12, two critical subunits of the polycomb repressive complex 2 (PRC2), a major writer of the repressive histone H3 lysine 27 trimethylation mark, in up to 25% of T-ALLs (Ntziachristos, Tsirigos et al. 2012, Zhang, Ding et al. 2012).
PHF6, a factor with proposed epigenetic function, is mutated and deleted in 16% of pediatric and 38% of adult T-ALL cases (Van Vlierberghe, Palomero et al. 2010). The PHF6 gene is located on the X- chromosome, and notably, mutations in T-ALL are observed almost exclusively in males (Van
Vlierberghe, Palomero et al. 2010). Recent studies also showed that different subtypes of ALL have distinct DNA methylation signatures which correlate with their gene expression profiles (Figueroa, Chen et al. 2013).
Finally, T-ALLs also bear genetic alterations in important signal transduction pathways, such as deletion mutations in the PTEN tumor suppressor in 5-10% of cases (Palomero, Sulis et al. 2007), ABL1 rearrangements including EML1-ABL1, ETV6-ABL1 and the already mentioned NUP214-ABL1 in 8% of cases (Van Limbergen, Beverloo et al. 2001, Graux, Cools et al. 2004, De Keersmaecker, Graux et al.
2005), prototypical RAS-activating mutations in 5-10% of cases (Bar-Eli, Ahuja et al. 1989, Van
Vlierberghe, Ambesi-Impiombato et al. 2011, Zhang, Ding et al. 2012), and alterations leading to
JAK/STAT pathway activation such as IL7R gain of function mutations in 10% of cases (Shochat, Tal et al. 2011, Zenatti, Ribeiro et al. 2011), activating mutations in JAK1 and JAK3 (Flex, Petrangeli et al. 2008,
Asnafi, Le Noir et al. 2010, Zhang, Ding et al. 2012), and a rare rearrangement resulting in expression of a constitutively active ETV6-JAK2 kinase fusion oncogene (Lacronique, Boureux et al. 1997). Gene expression profiling studies of these heterogeneous T-ALL tumors allowed the classification of T-ALL into distinct molecular subtypes based on the predominant oncogenic pathway which is activated: LYL1,
HOX11 or TAL1 (Ferrando, Neuberg et al. 2002).
Another important classification of T-ALL based on immunophenotype, genetics and pathology yielded a distinct subgroup called early T-cell precursor ALL (ETP-ALL), accounting for approximately
12% of cases (Coustan-Smith, Mullighan et al. 2009, Van Vlierberghe, Ambesi-Impiombato et al. 2011,
Zhang, Ding et al. 2012). ETP-ALLs show an early block at the double negative stage of thymocyte development with activation of transcriptional programs related to stem cells and myeloid progenitors, as
8 Chapter 1 well as subgroup-specific changes including aberrant expression of MEF2C, mutations in acute myeloid leukemia (AML) oncogenes and tumor suppressors, and inactivation of key transcription factors like
RUNX1, GATA3 and ETV6 (Coustan-Smith, Mullighan et al. 2009, Van Vlierberghe, Ambesi-Impiombato et al. 2011, Zhang, Ding et al. 2012, Zuurbier, Gutierrez et al. 2014).
Inherited genetic determinants of leukemogenesis
Most patients who develop ALL have no recognizable inherited predisposing factors. Patients with Down syndrome, however, have a 20-fold increased risk of developing ALL, although the exact role of the chromosome 21 in leukemia development is unknown (Hasle, Clemmensen et al. 2000). Notably,
50% of Down syndrome ALL cases harbor CRLF2 rearrangements which are frequently associated with
JAK mutations activating the JAK-STAT pathway (Mullighan, Collins-Underwood et al. 2009). Recently, rare germline mutations in PAX5, ETV6 and the cytokine signaling regulator SH2B3 have been linked to familial ALL (Perez-Garcia, Ambesi-Impiombato et al. 2013, Shah, Schrader et al. 2013, Zhang, Churpek et al. 2015), and germline TP53 mutations (as found in Li-Fraumeni syndrome) have been linked to low- hypodiploid ALL (Kleihues, Schauble et al. 1997, Holmfeldt, Wei et al. 2013). On the other hand, germline heterozygous RUNX1 mutations resulting in RUNX1 haploinsufficiency have been shown to cause familial thrombocytopenia with an increased risk of developing myeloid malignancies (Song, Sullivan et al.
1999).
Genome-wide association studies have also identified specific polymorphisms associated with a high risk of developing ALL, a high risk of relapse and/or with specific ALL subtypes in a few genes including GATA3, ARID5B, CEBPE, and IKZF1 (Papaemmanuil, Hosking et al. 2009, Trevino, Yang et al.
2009, Perez-Andreu, Roberts et al. 2013). The correlation between certain germline variants and specific
ALL subtypes (e.g. ARID5B variants and hyperdiploidy) indicates that inherited and acquired genetic traits may interact during leukemia development (Papaemmanuil, Hosking et al. 2009, Trevino, Yang et al.
2009).
II. Treatment of Acute Lymphoblastic Leukemia
The treatment of ALL has historic significance as a pioneering step in the use of chemotherapy
9 Chapter 1 for cancer and has served as a paradigm for the progressive improvement of patient outcome with incremental advances in treatment regimens and risk-based therapy.
History of combination chemotherapy
The term “chemotherapy” was coined by Paul Ehrlich in the early 1900s, who was one of the first to classify leukemia as myelogenous or lymphocytic and who pioneered the use of animal models to screen chemical compounds for the treatment of infectious diseases and cancers (Ehrlich 1877, Ehrlich
1880). In the early 1910s George Clowes developed the first transplantable tumor systems in rodents, allowing for standardized screening of multiple compounds for potential cancer treatment at the National
Cancer Institute (NCI) (Zubrod 1966). However, screening efforts were unsuccessful, with surgery and radiotherapy remaining the dominant anti-cancer treatment regimens and acute lymphoblastic leukemia remaining uniformly fatal (Southam, Craver et al. 1951).
The next development in the use of chemotherapy for leukemia came from serendipitous findings during World War I and II. Classified research on the effects of vesicant nitrogen and sulfur mustard gases during World War I showed that soldiers killed by sulfur mustard gas had developed leukopenia and lymphoid hypoplasia (Krumbhaar and Krumbhaar 1919). In World War II an accidental spill of sulfur mustard from a bombed ship in Bari Harbor, Italy, led to the exposure of over 1000 crewmen who developed resulting bone marrow and lymphoid hypoplasias (Alexander 1947). After the war the findings stimulated research on these alkylating agents as potential therapies for leukemias and lymphomas.
Alfred Gilman and Louis Goodman at Yale tested nitrogen mustard in mice with a transplanted lymphoid tumor and observed marked tumor regressions, prompting nitrogen mustard administration to lymphoma patients (Gilman 1946). Marked regressions were observed in these patients, encouraging the synthesis and testing of other alkylating agents, including cyclophosphamide, an anti-cancer drug in use today
(Gilman 1946, Goodman, Wintrobe et al. 1946). Unfortunately remissions induced by nitrogen mustard in lymphoma patients were brief and incomplete, with no evidence of efficacy against leukemias (Gilman
1963).
A crucial development in anti-leukemic chemotherapy was the discovery of folic acid from nutritional research by Lucy Wills, who found that yeast and liver extracts contained a factor that could
10 Chapter 1 alleviate anemia and leukopenia in pregnant women, and that when this factor was omitted from the diet of non-human primates it resulted in decreased bone marrow function (Wills, Clutterbuck et al. 1937).
Early studies had also suggested that this factor can induce breast tumor regression in animal models
(Leuchtenberger, Leuchtenberger et al. 1945). A team of chemists at Lederle Laboratories led by
Yellapragada SubbaRow identified the structure of folic acid and synthesized it (Angier, Boothe et al.
1945, SubbaRow Y 1946). Sidney Farber and Louis Diamond then tested folic acid on 90 patients with different late-stage cancers at the Children’s Medical Center in Boston, and found that folic acid actually leads to an acceleration of the leukemic process (Farber 1949). Thus Farber hypothesized that reduction of folic acid levels in leukemic patients might suppress the growth of malignant hematopoietic cells and collaborated with Lederle Laboratories to develop a series of folic acid antagonists known as “antifolates”
(Seeger, Smith et al. 1947). The antifolates included aminopterin and amethopterin (methotrexate) and were shown to induce folic acid deficiency in mice and chicks (Franklin, Stokstad et al. 1947). These results prompted the first clinical study of aminopterin in children critically ill with acute leukemia by Farber in 1948, which showed unquestionable remissions for the first time (Farber and Diamond 1948). While remissions were temporary and accompanied by serious side-effects, this seminal study demonstrated that a chemotherapeutic agent can efficiently suppress proliferation of malignant cells and allow for a restoration of normal hematopoiesis in the bone marrow. The aminopterin analog methotrexate is now a key antifolate drug in the treatment of ALL (Hunger and Mullighan 2015).
At the start of the 1950s Gertrude Elion and George Hitchings isolated a substance which inhibited adenine metabolism, leading them to develop two crucial nucleoside analog drugs, 6- mercaptopurine and 6-thioguanine, which were shown to be efficacious in treating leukemia by Joe
Burchenal and others, and which became a core part of the treatment of acute leukemias (Burchenal,
Murphy et al. 1953, Elion, Singer et al. 1954, Hitchings and Elion 1954). Increased screening efforts yielded more valuable drugs, including the Vinca alkaloids, anthracyclines and platinum drugs, and trials in acute leukemia occurred with the newly discovered corticosteroids, but these were also found to produce only brief responses when used alone (Pearson, Eliel et al. 1949). By 1960 complete remissions could be achieved with single agents in 25% of children with leukemia, but they only lasted some months
(DeVita and Chu 2008).
11 Chapter 1
Furth and Kahn had previously demonstrated that a single implanted leukemic cell was sufficient to induce leukemia and death in mice (Furth 1937) and Skipper proposed the “Cell Kill” hypothesis that a given dose of chemotherapy kills a constant fraction of tumor cells, such that drug efficacy depends on the number of tumor cells at the start of treatment (Skipper, Schabel et al. 1964). The importance of chemotherapy scheduling and dosage had also been demonstrated (Goldin, Mantel et al. 1954), and together these observations prompted the introduction of more aggressive treatment regimens.
In 1961 Emil Frei and Jay Freireich at the NCI reported the first study of combination chemotherapy with 6-mercaptopurine and methotrexate in 39 pediatric leukemia patients, achieving a complete remission rate of 59% and a 2-year survival rate of 20% (Freireich, Gehan et al. 1961). This was followed by the first cyclically administered treatment regimen called “VAMP” which included the newly discovered Vinca alkaloid vincristine, amethopterin (methotrexate), 6-mercaptopurine and prednisone
(Frei, Karon et al. 1965). Subsequent combination chemotherapy studies progressively increased the remission rate to 60% and the remission duration from months to years by 1970, which for the first time was compatible with cure in some patients (Frei and Freireich 1965, Frei 1967, Holland 1970). Increased intensity regimens were accompanied by the development of improved supportive care, including platelet transfusions and prophylactic administration of antibiotics (Gaydos, Freireich et al. 1962, Hersh, Bodey et al. 1965), while the occurrence of central nervous system (CNS) relapses prompted the development of concurrent CNS prophylaxis/treatment (Evans, Gilbert et al. 1970).
St Jude Children’s Research Hospital developed a curative approach to ALL treatment including multiple components, namely remission induction, central nervous system (CNS)-directed therapy to prevent/treat CNS metastases, intensification (consolidation) therapy, and continuation (maintenance) therapy (Pinkel 1971). This regimen proved highly successful, increasing cure rates to 50% (Aur, Simone et al. 1971), and subsequent studies in the US and Germany found that administration of intensification therapy soon after remission induction and addition of high-dose L-asparaginase could achieve cure rates of almost 70% (Henze, Langermann et al. 1981, Sallan, Hitchcock-Bryan et al. 1983). The regimen developed in Germany which included intense eight-drug, eight-week induction and consolidation phases, known as Berlin-Frankfurt-Münster, became the basis for most current therapies of ALL.
12 Chapter 1
At the same time concerns were raised over the potential developmental side effects and increased risk of secondary brain tumors by the CNS therapy to date, which had included prophylactic cranial irradiation. This led to the development of combination chemotherapy for the CNS and studies showed that triple intrathecal therapy with methotrexate, hydrocortisone and cytarabine, as well as higher doses of intravenous methotrexate were equally effective in preventing CNS relapse (Sullivan, Chen et al.
1982, Freeman, Weinberg et al. 1983, Jones, Freeman et al. 1991). This allowed for the removal of prophylactic cranial irradiation from the treatment of almost all patients with ALL (Pui, Campana et al.
2009).
Overall, these seminal studies in ALL provided a proof-of-concept and a model for achieving progressive increases in survival by step-wise improvements in combination chemotherapy.
Contemporary therapy for ALL
Continued improvements in multi-agent chemotherapy and risk-based treatment stratification have achieved remarkable success particularly in pediatric ALL, with overall survival reaching 80-90% in most protocols (Figure 1.4) (Hunger and Mullighan 2015).
Figure 1.4. Overall survival among children with ALL who were enrolled in Children’s Cancer Group and Children’s Oncology Group clinical trials, 1968-2009. Reproduced from Hunger and Mullighan, 2015.
13 Chapter 1
Current therapy for ALL consists of four discreet phases: remission induction, consolidation or intensification, maintenance therapy and, in parallel with these, CNS-directed therapy (Hunger and
Mullighan 2015).
Induction therapy lasts four to six weeks, with an aim to induce complete remission with a restoration of normal hematopoiesis and less than 5% blasts detectable microscopically. The chemotherapy agents used are vincristine, asparaginase and glucocorticoids (prednisone or dexamethasone), with optional addition of an anthracycline (doxorubicin or daunorubicin) (Table 1.1)
(Cooper and Brown 2015). Clinical trials have shown similar efficacy and toxicity for doxorubicin and daunorubicin (Escherich, Zimmermann et al. 2013), while glucocorticoid studies have shown that dexamethasone has improved CNS penetration and leads to reduced risk of CNS relapse, but at the cost of increased toxicity (Mitchell, Richards et al. 2005). CNS-directed intrathecal therapy/prophylaxis is also
Table 1.1. List of commonly used chemotherapeutic agents in the treatment of ALL.
Drug name Type of drug Mechanism Vincristine Mitotic spindle poison Blocks cell division, leading to apoptosis. Asparaginase Enzyme Removes circulating asparagine which is needed by leukemic cells since they cannot synthesize it. Dexamethasone/ Glucocorticoids Activate glucocorticoid receptor signaling and transcription, Prednisone repress AP-1 or NFκB transcription, resulting in cell cycle arrest and apoptosis. Doxorubicin/ Topoisomerase II Intercalate into DNA, block topoisomerase II and DNA Daunorubicin poisons replication, thereby inducing DNA damage response. Methotrexate Antifolate Inhibits DHFR, thereby blocking folate synthesis which is needed for de novo nucleoside biosynthesis. Cytarabine Antimetabolite Analog of deoxycytidine, which is incorporated into DNA and blocks DNA synthesis. 6-mercaptopurine/ 6- Antimetabolite Metabolized to deoxyguanosine analogs which block DNA thioguanine replication. Methylated 6-mercaptopurine metabolites block de novo purine biosynthesis. Cyclophosphamide Alkylating agent Adds an alkyl group to DNA, resulting in DNA crosslinking and (nitrogen mustard) disruption of DNA replication. Etoposide Topoisomerase II Blocks topoisomerase II and disrupts DNA replication, thereby poison inducing DNA damage response.
Abbreviations: AP-1, activator protein 1; NFκB, nuclear factor kappa-B; DHFR, dhydrofolate reductase.
14 Chapter 1 administered during the induction phase, including methotrexate, cytarabine and hydrocortisone (Cooper and Brown 2015). About 95% of patients successfully achieve post-induction remission and enter into the consolidation phase, which lasts 6 to 9 months and aims to eradicate submicroscopic residual leukemic burden. Consolidation consists of an intensive combination of chemotherapy agents aimed to minimize drug resistance, including some drugs not yet administered, such as 6-mercaptopurine, 6-thioguanine, cytarabine, methotrexate, cyclophosphamide and etoposide (Cooper and Brown 2015). The repeated use of high-dose methotrexate followed by folinic acid (leucovorin) rescue of normal tissues from toxicity is a key component of this therapy (Hunger and Mullighan 2015). Consolidation is followed by an 8-week delayed-intensification (re-induction) regimen with drugs similar to those used during remission-induction, which has proven particularly beneficial for the outcome of patients with high-risk disease (Seibel,
Steinherz et al. 2008).
Patients then enter into the final phase of treatment, maintenance therapy, which can last 2-3 years and includes low-intensity, mostly anti-metabolite-based chemotherapy which has been shown to reduce the risk of relapse (Frei, Karon et al. 1965, Lonsdale, Gehan et al. 1975). The cornerstone of maintenance therapy is daily oral 6-mercaptopurine and weekly methotrexate. While 6-thioguanine has shown some improvements in event-free survival in select groups compared to 6-mercaptopurine, it does not provide improvements in overall survival and leads to greater liver toxicity, making 6-mercaptopurine the thiopurine drug of choice (Escherich, Richards et al. 2011). Some regimens also include monthly 5-7 day pulses of vincristine and glucocorticoids, although there is not much evidence for added benefit
(Eden, Pieters et al. 2010). A crucial factor in the success of maintenance therapy is compliance, with studies showing that 20% of patients are less than 90% adherent, and that non-adherence is correlated with a four-fold higher risk of relapse (Bhatia, Landier et al. 2012, Bhatia, Landier et al. 2014).
Finally, the efficacy of allogeneic hematopoietic stem cell transplantation at different stages of the disease is still being assessed, with its use currently reserved for the most high-risk groups and for relapsed cases (Hochberg, Khaled et al. 2013, Peters, Schrappe et al. 2015).
The recent therapeutic achievements in pediatric ALL have not been reflected in the adult setting, where current therapy is only able to cure 40-50% of patients (Sive, Buck et al. 2012). The treatment for adult ALL is largely the same as that for pediatric ALL, with the exception that adult patients receive lower
15 Chapter 1 cumulative doses of chemotherapy, while also presenting more frequently with high-risk features at diagnosis (Jabbour, O'Brien et al. 2015). Recent studies have indicated that outcomes for adolescents and young adults are better if they are enrolled in pediatric-inspired protocols with intensified exposure to non-myelosuppressive agents like steroids, vincristine and asparaginase (Stock, La et al. 2008, Huguet,
Leguay et al. 2009, Rytting, Thomas et al. 2014). In the older adult population, however, tolerance of high-intensity chemotherapy is greatly decreased, with toxic side effects limiting the possibility for increased drug exposure (O'Brien, Thomas et al. 2008, Huguet, Leguay et al. 2009).
Prognostic factors and risk stratification
A hallmark of ALL treatment is the risk-based stratification of patients according to a set of prognostic factors including clinical features, leukemic cell biology or genetics and initial response to therapy, with patients at a higher risk of treatment failure receiving more intensive treatments, while lower- risk patients being spared unnecessary chemotherapy and associated toxicity (Cooper and Brown 2015,
Hunger and Mullighan 2015).
Clinical features include age and white blood cell count (WBC) at presentation, with age between
1 and 10 years considered standard risk, while infants, patients older than 10 years and patients with a
WBC greater than 50,000/µL are considered to have high-risk disease (Moricke, Zimmermann et al. 2005,
Schultz, Pullen et al. 2007). Leukemic involvement of extramedullary anatomic locations at diagnosis such as the CNS or testes (termed “sanctuary sites” due to being difficult to penetrate with systemic chemotherapy) is also a high-risk clinical feature (Pui and Thiel 2009, Cooper and Brown 2015).
The features of the leukemic cells also play an important role in prognosis. Immunophenotype classification as B-cell or T-cell ALL is performed based on the expression of cell-surface and cytoplasmic lineage markers, with T-ALL historically considered as high-risk and associated with reduced survival compared to B-ALL, although intensified chemotherapy has narrowed this difference in recent years
(Hunger, Lu et al. 2012). Exceptions are ETP-ALL and T-ALL with absence of biallelic TCRG deletion (an early event in T-lineage development), which are still associated with a very poor prognosis (Coustan-
Smith, Mullighan et al. 2009, Gutierrez, Dahlberg et al. 2010).
Genetic alterations in the leukemic cells which have been linked to favorable prognosis include
16 Chapter 1 high hyperdiploidy (51-65 chromosomes/cell) and the ETV6-RUNX1 translocation, which form the two most common subtypes of B-ALL (Schultz, Pullen et al. 2007). On the other hand, hypodiplody (< 44 chromosomes/cell) is associated with poor prognosis (Nachman, Heerema et al. 2007), as are the BCR-
ABL1 (Ph) fusion (Arico, Schrappe et al. 2010), Ph-like ALL (Den Boer, van Slegtenhorst et al. 2009,
Mullighan, Su et al. 2009), MLL rearrangements (Moorman, Ensor et al. 2010), intrachromosomal amplification of chromosome 21 (iAMP21) (Harrison, Moorman et al. 2014) and CRLF2 rearrangement
(Harvey, Mullighan et al. 2010). In T-ALL, aberrant expression of TLX1 is associated with a favorable prognosis (Ferrando, Neuberg et al. 2002, Ferrando, Neuberg et al. 2004), as is the MLL-MLLT1 fusion
(contrary to MLL rearrangements in B-ALL) (Rubnitz, Camitta et al. 1999, Ferrando, Neuberg et al. 2002).
In addition to all of these features observed at diagnosis, the initial patient response to early induction chemotherapy has powerful independent prognostic value (Coustan-Smith, Sancho et al. 2000).
The assessment of post-induction remission by microscopic morphology is relatively insensitive and is now supplemented by the evaluation of minimal residual disease (MDR) in the bone marrow by flow cytometry or polymerase chain reaction (PCR), allowing the assessment of submicroscopic disease at a detection threshold of 1 leukemic blast per 10,000-100,000 cells (Campana 2012, Pui, Pei et al. 2015).
The end-of-induction MRD analysis has become standard in risk-stratification for ALL treatment, with late or incomplete response to induction therapy indicating a high risk of relapse (Borowitz, Devidas et al.
2008, Schrappe, Valsecchi et al. 2011). The risk of treatment failure and death is 3-5 fold higher in cases with post-induction MRD of 0.01% or greater (Borowitz, Devidas et al. 2008, Conter, Bartram et al. 2010,
Schrappe, Valsecchi et al. 2011), but intensification of therapy for these patients can improve their outcome (Vora, Goulden et al. 2014, Pui, Pei et al. 2015). Studies are also investigating the utility of post- consolidation MRD measurement, which may be particularly informative of outcome for T-ALL (Schrappe,
Valsecchi et al. 2011). Finally, new deep-sequencing methods for MRD analysis are being developed with improved precision and sensitivity, which may also prove useful in the monitoring of tumor clonal evolution (Faham, Zheng et al. 2012).
Toxic effects of treatment
Intensified combination chemotherapy has led to a great improvement in overall survival, but this
17 Chapter 1 has come at a cost of short-term and long-term toxic side effects for the patient (Table 1.2). While serious toxicities are relatively uncommon, about 1-2% of patients die from treatment-related toxicity while in remission (Blanco, Beyene et al. 2012). The most serious side effect observed is osteonecrosis in the hips, knees, shoulders and ankles in 5 to 10% of cases, often requiring surgical management and joint replacement (Te Winkel, Pieters et al. 2014). The risk is much higher among older patients and females, but can be decreased by modifying the schedule of glucocorticoid administration (Mattano, Devidas et al.
2012). Other side effects include liver, kidney and cardiovascular toxicity, CNS and peripheral neuropathy, hypertension, obesity and metabolic syndrome (Table 1.2) (Cooper and Brown 2015). In general dose-limiting toxicities are observed more frequently in the adult population (O'Brien, Thomas et al. 2008, Huguet, Leguay et al. 2009).
Table 1.2. Toxic side effects of chemotherapy used in the treatment of ALL.
Drug name Side effects Vincristine Inappropriate diuretic hormone, neuropathy (foot/wrist drop, paresthesias, constipation, ptosis, vocal cord paresis) Asparaginase Hypersensitivity reactions, pancreatitis, thrombosis Glucocorticoids Osteonecrosis, hypertension, hyperglycemia, fluid retention, psychosis Doxorubicin/ Cardiotoxicity Daunorubicin Methotrexate Mucositis, nephrotoxicity, hepatotoxicity, encephalopathy Cytarabine Conjunctivitis, flu-like symptoms 6-mercaptopurine Hepatotoxicity 6-thioguanine Hepatotoxicity including sinusoidal obstruction syndrome and portal hypertension Cyclophosphamide Nephrotoxicity, hemorrhagic cystitis, hyponatremia, fluid retention Etoposide Nephrotoxicity, hepatotoxicity, hypersensitivity reactions
Adapted from Cooper and Brown, 2015.
Unfortunately many long term survivors experience chronic toxic effects (Oeffinger, Mertens et al.
2006) and potential neurocognitive deficits become more apparent with increase in age (Krull, Brinkman et al. 2013). Pharmacokinetic, pharmacodynamic and pharmacogenomic studies have begun to identify patient-specific determinants that influence the efficacy and the risk of toxicity of each chemotherapy drug, with the ultimate goal to tailor the drug exposure for each patient according to both the risk of treatment failure and the risk of toxicity (Pui 2015).
18 Chapter 1
Inherited genomic determinants of therapy response and toxicity
An important early pharmacodynamics study of high-dose methotrexate treatment for ALL found that steady-state serum concentrations (an indication of the patient’s systemic exposure to the drug) directly correlated with treatment outcome (Evans, Crom et al. 1986). A subsequent randomized trial showed that patient outcome can be improved by tailoring high-dose methotrexate, teniposide and cytarabine therapy according to each patient’s rate of clearance of the drugs (Evans, Relling et al. 1998), thereby showing that treatment failure could be the result of suboptimal exposure of the leukemic cells to the chemotherapy and establishing the principle of individualized or “personalized” dosing in cancer treatment.
A crucial finding about inter-patient variability in 6-mercaptopurine efficacy and toxicity came soon after, with the identification of inherited polymorphisms in the gene encoding thiopurine methyltransferase
(TPMT), an enzyme responsible for the S-methylation of thiopurines to inactive methylated metabolites
(Lennard, Lilleyman et al. 1990, Krynetski, Schuetz et al. 1995). About 10% of individuals inherit either one or two variant alleles of TPMT which encode for an unstable or less active enzyme, leading to excessive accumulation of cytotoxic thiopurine metabolites following 6-mercaptopurine treatment (Relling,
Hancock et al. 1999). This results in increased risk of hematopoietic toxicity and development of therapy- related secondary leukemia (Relling, Hancock et al. 1999, Evans, Hon et al. 2001, Schmiegelow, Heyman et al. 2010). Notably, the cumulative incidences of myelosuppression for patients with wildtype, heterozygous and homozygous deficient TPMT alleles are 7%, 35% and 100% respectively (Relling,
Hancock et al. 1999). These findings have led to the implementation of new pharmacogenetic guidelines for tailoring 6-mercaptopurine treatment according to TPMT genotype (with as much as a 90% dose reduction for cases with homozygous deficient TPMT), resulting in equivalent systemic exposure, efficacy and tolerance between patients (Relling, Gardner et al. 2011).
Genome-wide SNP analyses have uncovered a number of other germline polymorphisms which affect response to therapy. One study identified 102 variants in 71 unique genomic loci that associate with chemotherapy disposition and MRD at the end of remission induction therapy (Yang, Cheng et al. 2009).
Notably, a fifth of the SNPs related to high MRD were linked to increased drug clearance of methotrexate
19 Chapter 1 and etoposide or reduced accumulation of cytotoxic methotrexate polyglutamate metabolites inside the leukemic cells (Yang, Cheng et al. 2009). Subsequent studies showed that methotrexate clearance is affected by germline SNPs in the organic anion transporter gene SLCO1B1, while methotrexate polyglutamate accumulation is affected by variants in ARID5B (Trevino, Shimasaki et al. 2009, Ramsey,
Bruun et al. 2012, Xu, Cheng et al. 2012, Ramsey, Panetta et al. 2013). Other studies have found inherited genetic variations associated with increased risk of allergy and hypersensitivity to asparaginase
(Fernandez, Smith et al. 2014), steroid-induced osteopenia and osteonecrosis (French, Hamilton et al.
2008, Jones, Kaste et al. 2008), vincristine-induced neuropathy (Diouf, Crews et al. 2015) and anthracycline-relayed cardiomyopathy (Wang, Liu et al. 2014). New polymorphisms affecting 6- mercaptopurine toxicity have also been discovered, including variants of PACSIN2 associated with mercaptopurine-related gastrointestinal toxicity (Stocco, Yang et al. 2012) and a NUDT15 variant associated with thiopurine-induced leukopenia which is most commonly found in East Asians (Yang,
Landier et al. 2015). Moreover, 10% of Caucasians have been found to carry reduced-function SNPs in the ITPA gene which encodes an enzyme involved in thiopurine metabolism, leading to an increased risk of 6-mercaptopurine-induced toxicity (Stocco, Cheok et al. 2009, Stocco, Crews et al. 2010).
As our knowledge of important host polymorphisms grows, new guidelines for prospective pharmacogenetics testing are being implemented for patients with ALL, which will increasingly individualize leukemia treatment for achieving the best outcome with minimal toxic effects (Hoffman,
Haidar et al. 2014).
III. Drug Resistance and Relapse of Acute Lymphoblastic Leukemia
Drug resistance and relapse of ALL remain the biggest causes of treatment failure, with 15-20% of cases showing primary resistance or relapsing after remission (Bhojwani and Pui 2013). Moreover, treatment regimens for patients with relapsed or refractory ALL are still highly ineffective and can only achieve a cure in 40% of cases (Bhojwani and Pui 2013). Relapsed T-ALL has a particularly dismal prognosis with 3-year event-free survival rates of less than 15% (Einsiedel, von Stackelberg et al. 2005,
Raetz, Borowitz et al. 2008).
20 Chapter 1
Risk and prognostic factors
In addition to germline variants linked to drug efficacy and toxicity, studies have identified polymorphisms that are associated with increased risk of relapse, such as inherited variants in GATA3
(Perez-Andreu, Roberts et al. 2013), PYGL, which is essential for response to 6-mercaptopurine and methotrexate (Yang, Cheng et al. 2012), and PDE4B and ABCB1, which are important for methotrexate pharmacodynamics and sensitivity to steroids (Yang, Cheng et al. 2012). Deletion of IKZF1 in the leukemic cells is also a strong predictor of relapse (Mullighan, Su et al. 2009, Kuiper, Waanders et al.
2010).
Risk factors linked to overall outcome after relapse have also been identified, such as the length of first complete remission (CR1), with early relapses (<18 months from diagnosis) having the worst prognosis while those occurring late (>3 years from diagnosis) having the best prognosis (Chessells
1998). The site of relapse is another prognostic factor, with isolated bone marrow relapses occurring in
50-60% of cases and having the worst prognosis, isolated extramedullary relapses (20% occurring in the
CNS and 5% in the testes) having the best prognosis, and the remainder accounting for combined marrow and extramedullary relapses which have intermediate prognosis (Nguyen, Devidas et al. 2008).
Similarly to newly diagnosed ALL, the early response of relapsed ALL to re-induction (MRD level) carries strong prognostic significance (Coustan-Smith, Gajjar et al. 2004, Eckert, von Stackelberg et al. 2013), and the presence of ETV6-RUNX1 is associated with improved survival following relapse (Seeger, von
Stackelberg et al. 2001, Gandemer, Chevret et al. 2012). On the other hand, the presence of TP53 mutations or IKZF1 deletions in the relapsed leukemia is associated with a poor outcome (Hof, Krentz et al. 2011, Krentz, Hof et al. 2013).
A crucial factor associated with risk of relapse is the systemic exposure to 6-mercaptopurine and methotrexate during maintenance therapy, where treatment adherence and pharmacogenetics play a major role (Relling, Gardner et al. 2011, Bhatia, Landier et al. 2012, Bhatia, Landier et al. 2014). Studies have shown that the degree of myelosuppression from these drugs during maintenance therapy correlates with the risk of relapse (Schmiegelow, Schroder et al. 1995, Schmiegelow, Heyman et al.
2010) and that personalized tailoring of methotrexate and 6-mercaptopurine dosages to the limits of tolerance is associated with improved outcome (Pui, Mullighan et al. 2012). As a result, some treatment
21 Chapter 1 centers now administer weekly maintenance methotrexate intravenously rather than orally in order to circumvent inter-patient differences in bioavailability and poor treatment adherence (Pui and Evans 2013).
Biology of drug resistant and relapsed ALL
As our knowledge of the host/germline genetic factors linked to drug resistance grows, so does our understanding of the biology of the drug resistant leukemic cells themselves, with early genome-wide expression studies showing that the mRNA expression of a relatively small number of genes in the tumor cells is associated with de novo sensitivity to glucocorticoids, L-asparaginase, vincristine and daunorubicin (Holleman, Cheok et al. 2004), response to high-dose methotrexate (Sorich, Pottier et al.
2008), and leukemic cell multi-drug cross-resistance (Lugthart, Cheok et al. 2005). Recently de novo resistance to thiopurine therapy in 11% of newly diagnosed ALL was attributed to somatic deletions of genes required for maintaining protein stability of MSH2, a DNA mismatch repair enzyme that is important for thiopurine activity (Diouf, Cheng et al. 2011).
Resistance to glucocorticoid therapy is of key importance in ALL, with dexamethasone or prednisone resistance frequently observed in relapsed ALL tumor cells and primary glucocorticoid resistance conferring a much higher risk of relapse (Riehm, Reiter et al. 1987, Klumper, Pieters et al.
1995, Kaspers, Wijnands et al. 2005). Recently an extensive study of prednisolone sensitivity in 444 primary tumor samples from newly diagnosed ALL patients revealed that ALL tumors with primary glucocorticoid resistance expressed significantly higher levels of caspase 1 (CASP1) and its activator
NLRP3 due to decreased somatic methylation of their promoters (Paugh, Bonten et al. 2015). Greater caspase 1 expression led to glucocorticoid resistance by increasing cleavage of the NR3C1 glucocorticoid receptor (GR), which resulted in decreased GR-induced transcriptional response to glucocorticoids (Paugh, Bonten et al. 2015).
Earlier research found that patient-derived B-ALL xenografts which exhibit glucocorticoid resistance in vitro had similar expression levels and ligand affinity of the GR as glucocorticoid-sensitive xenografts (Bachmann, Gorman et al. 2005). Similarly, rates of GR translocation to the nucleus in response to dexamethasone were equivalent in glucocorticoid sensitive and resistant tumors, indicating that glucocorticoid resistance may occur downstream of the GR in those cases (Bachmann, Gorman et al.
22 Chapter 1
2005). Glucocorticoid-mediated lymphoblast cytotoxicity has been well-documented to occur through the intrinsic apoptotic pathway (Wang, Malone et al. 2003, Schmidt, Rainer et al. 2006), and analyses of the apoptotic machinery in glucocorticoid-resistant xenografts showed that resistant leukemic cells have significantly attenuated induction of the pro-apoptotic BCL-2 family member BIM in response to dexamethasone (Bachmann, Gorman et al. 2005). Reduced expression levels of BIM in glucocorticoid- resistant patient samples were found to be due to epigenetic silencing of the BIM gene, which is effectively reversed by histone deacetylase inhibition with vorinostat, resulting in resensitization of the leukemic cells to dexamethasone treatment (Bachmann, Piazza et al. 2010).
New evidence is emerging that non-apoptotic cell death such as autophagy-dependent necroptosis may play an important role in resensitizing resistant ALL cells to glucocorticoid therapy
(Bonapace, Bornhauser et al. 2010) and that dexamethasone-induced autophagy may even be required for the apoptotic response (Laane, Tamm et al. 2009).
A few studies have shown that glucocorticoid-resistant ALL cells have increased glucose/carbohydrate metabolism, oxidative phosphorylation and cholesterol biosynthesis (Holleman,
Cheok et al. 2004, Samuels, Heng et al. 2014), and that glycolysis inhibition with 2-deoxy-D-glucose (2-
DG) synergizes with prednisolone or dexamethasone treatment to increase cell death specifically in glucocorticoid-resistant ALL cells (Eberhart, Renner et al. 2009, Hulleman, Kazemier et al. 2009,
Samuels, Heng et al. 2014). Similar synergistic effects with glucocorticoids have been observed with co- administration of the oxidative phosphorylation inhibitor oligomycin or the inhibitor of cholesterol metabolism simvastatin in resistant cells (Samuels, Heng et al. 2014).
Studies on T-ALL cells have uncovered that NOTCH1 pathway activation plays a key role in mediating resistance to glucocorticoids, which can be reversed with GSI treatment (Real, Tosello et al.
2009). These findings were recently confirmed in a preclinical study which showed that a combination of dexamethasone and a clinically-relevant GSI had a synergistic anti-leukemic effect in vivo (Samon,
Castillo-Martin et al. 2012). Glucocorticoid resistance in T-ALL can also arise from loss of the PTEN tumor suppressor, which leads to increased AKT-mediated phosphorylation of the glucocorticoid receptor
NR3C1 that blocks its translocation to the nucleus (Piovan, Yu et al. 2013). This can be effectively restored by pharmacological AKT1 inhibition, which reverses glucocorticoid resistance in vitro and in vivo
23 Chapter 1
(Piovan, Yu et al. 2013). Other groups have similarly shown PI3K/AKT/mTOR pathway inhibition to be efficacious in sensitizing glucocorticoid-resistant B- and T-ALL cells to dexamethasone (Wei, Twomey et al. 2006, Bornhauser, Bonapace et al. 2007).
Other studies have suggested that glucocorticoid resistance may be mediated by decreased expression of SWI/SNF chromatin-remodeling complex subunits (Pottier, Yang et al. 2008), by increased
MAPK pathway activity (Jones, Gearheart et al. 2015), by loss-of-function alterations in the tumor suppressor IKZF1 (Marke, Havinga et al. 2016) or by deletions in TBL1XR1, an F-box like protein responsible for regulating the nuclear hormone repressor complex stability (Jones, Bhatla et al. 2014).
Early studies comparing diagnostic and relapsed ALL cells showed that relapsed leukemic blasts are more resistant to a number of chemotherapy drugs (Hongo and Fujii 1991, Klumper, Pieters et al.
1995). However until recently not much was known about the molecular mechanisms underlying ALL relapse. Gene expression analysis of matched diagnosis and relapse ALL samples has revealed some valuable differences in the expression levels of genes regulating DNA repair, cell cycle progression, apoptosis and drug response (Beesley, Cummings et al. 2005, Bhojwani, Kang et al. 2006). An example is the increased expression of survivin, an inhibitor of apoptosis, at relapse (Bhojwani, Kang et al. 2006).
Overexpression of survivin is sufficient to induce chemoresistance in primary ALL cells, while combination of chemotherapy and a survivin inhibitor is sufficient to eliminate primary patient-derived drug-resistant
ALL cells (Park, Gang et al. 2011).
It was discovered early on that ALL is a clonally heterogeneous disease which undergoes cytogenetic and immunophenotypic drift from diagnosis to relapse (Pui, Raimondi et al. 1986, Raimondi,
Pui et al. 1993, van Wering, Beishuizen et al. 1995). Several mechanisms have been implicated in the emergence of relapse clones, such as the positive selection of more aggressive malignant cells (Clappier,
Gerby et al. 2011), the presence of rare quiescent leukemia stem cells which can escape chemotherapy and reseed tumor populations (Lapidot, Sirard et al. 1994, Konopleva, Zhao et al. 2002, Guan, Gerhard et al. 2003, Chiu, Jiang et al. 2010, Eppert, Takenaka et al. 2011, Meyer, Eckhoff et al. 2011, Shlush, Zandi et al. 2014), or the protection and support of tumor cells by microenvironment niches (Konopleva,
Konoplev et al. 2002, Iwamoto, Mihara et al. 2007, Yang, Mallampati et al. 2013). New genome-wide analyses of matched germline, diagnosis and relapsed ALL samples have shown that the majority of
24 Chapter 1 relapses occur via clonal evolution from a heterogeneous mix of tumor clones found at diagnosis, with outgrowth of a subclone which is related to but distinct from the original major tumor clone (Figure 1.5)
(Mullighan, Phillips et al. 2008, Yang, Bhojwani et al. 2008, Clappier, Gerby et al. 2011, Ma, Edmonson et al. 2015). DNA copy number analyses with lesion-specific backtracking initially suggested that about half of relapsed cases originate from a pre-leukemic clone (possibly a leukemic stem cell) which is ancestral and related to the primary leukemic cells and represents a minor subpopulation at diagnosis, while about a third of relapses harbor the same genetic lesions as those present in the predominant diagnostic clone together with new acquired genetic alterations, indicating linear evolution of the primary leukemic clone.
Lastly, a small percentage of patients relapse with a second malignancy which bears completely different genetic alterations to their original leukemia (Figure 1.5) (Mullighan, Phillips et al. 2008). Similar findings have been reported on different ALL subtypes, namely ETV6-RUNX1-positive ALL, high-hyperdiploid ALL and T-ALL (Davidsson, Paulsson et al. 2010, Kuster, Grausenburger et al. 2011, Szczepanski, van der
Velden et al. 2011, van Delft, Horsley et al. 2011). A more recent study using deep whole exome
Figure 1.5. Model of clonal evolution of relapsed ALL. A pre-leukemic stem cell (light blue) gives rise to ALL (dark blue). Intrinsically drug-resistant clones (red) may be present at diagnosis at low levels. Leukemic stem cells and other subclones may survive initial treatment and acquire additional lesions that result in acquired drug resistance. Rarely, the relapsed tumor is genetically distinct from that at diagnosis (second malignancy). Adapted from Bhatla, 2014.
25 Chapter 1 sequencing analysis to study the clonal evolution in 20 matching diagnostic, remission and relapsed B-
ALL cases showed that 75% of relapsed leukemias are descendants of minor tumor subclones found at diagnosis, indicating that in most cases the predominant diagnostic clones are eradicated by therapy (Ma,
Edmonson et al. 2015). Thus, clonal evolution of the tumor population could arise from direct positive selection of a pre-existing clone found at diagnosis, or from the acquisition of new genetic alterations during induction, consolidation or maintenance therapy.
Indeed, early studies showed that drug-resistant ALL can emerge by the acquisition of secondary genetic alterations which directly promote chemotherapy resistance, such as amplification of the dihydrofolate reductase (DHFR) gene in resistance to methotrexate (Alt, Kellems et al. 1978, Goker,
Waltham et al. 1995). In this context, we recently discovered novel secondary genetic alterations which directly drive resistance to 6-mercaptopurine in relapsed ALL (described in Chapter 3) (Tzoneva, Perez-
Garcia et al. 2013). Through sequencing analyses of matched diagnostic, remission and relapsed ALL samples we identified recurrent relapse-specific activating mutations in the cytosolic 5’-nucleotidase II
(NT5C2) gene in 19% of relapsed T-ALLs and 3% of relapsed B-ALLs (Tzoneva, Perez-Garcia et al.
2013), with another group concurrently identifying NT5C2 mutations in 10% of relapsed B-ALLs (Meyer,
Wang et al. 2013). NT5C2 encodes a ubiquitously expressed and highly conserved enzyme that dephosphorylates purine monophosphates such as inosine monophosphate (IMP) and guanosine monophosphate (GMP) (Spychala, Madrid-Marina et al. 1988, Oka, Matsumoto et al. 1994), but which is also capable of dephosphorylating and inactivating the cytotoxic metabolites of the key anti-leukemic purine analog drugs 6-mercaptopurine and 6-thioguanine (Brouwer, Vogels-Mentink et al. 2005).
Subsequent studies have reported NT5C2 mutations in 45% (9/20) of early-relapsed B-ALLs (Ma,
Edmonson et al. 2015) and 38% (5/13) of relapsed T-ALLs (Kunz, Rausch et al. 2015). In a recent collaborative study we also identified recurrent relapse-associated mutations in another purine metabolic pathway gene, PRPS1 (phosphoribosyl pyrophosphate synthetase 1) in 6.7% (24/358) of relapsed childhood B-ALLs, which also drive resistance to 6-mercaptopurine and 6-thioguanine (Li, Li et al. 2015).
Other relapse-acquired lesions are enriched in tumor suppressor pathways (TP53 mutations occur in 10% of relapsed B-ALLs and T-ALLs) (Hof, Krentz et al. 2011), Ras signaling (NRAS, KRAS,
NF1, PTPN11), and epigenetic regulators like CREBBP, SETD2, KDM6A, MLL2, WHSC1 and SUZ12
26 Chapter 1
(Mullighan, Zhang et al. 2011, Mar, Bullinger et al. 2014, Kunz, Rausch et al. 2015, Ma, Edmonson et al.
2015). Mutations in the Ras pathway are associated with high-risk ALL features and poor outcome, being frequently present at low levels at diagnosis and enriched at relapse (Lubbert, Mirro et al. 1990, Irving,
Matheson et al. 2014). Relapse-specific CREBBP mutations (observed in 18% (17/71) of cases) have been linked to glucocorticoid resistance (Mullighan, Zhang et al. 2011), and have been found at a higher frequency in a small cohort of high-hyperdiploid ALL (10/16 cases) (Inthal, Zeitlhofer et al. 2012). Recent epigenetic studies have also reported an increase in global promoter methylation at relapse which can be reversed with demethylating agents to confer enhanced chemosensitivity (Hogan, Meyer et al. 2011,
Bhatla, Wang et al. 2012).
Relapsed ALLs can also harbor focal deletions in the MSH6 mismatch repair gene which have been implicated in resistance to 6-mercaptopurine and prednisone (Yang, Bhojwani et al. 2008, Hogan,
Meyer et al. 2011), and deletions in glucocorticoid signaling components such as BTG1, TBL1XR1 and the glucocorticoid receptor NR3C1 (Mullighan, Goorha et al. 2007, Mullighan, Phillips et al. 2008, Yang,
Bhojwani et al. 2008, Hogan, Meyer et al. 2011, Kuster, Grausenburger et al. 2011), which have been linked to steroid resistance (Perissi, Aggarwal et al. 2004, Perissi, Scafoglio et al. 2008, van Galen,
Kuiper et al. 2010). In addition to IKZF1 deletions, relapsed B-ALL cases show enrichment in somatic copy number alterations of other regulators of B-cell development and differentiation, including
CDKN2A/B, PAX5 and EBF1 (Mullighan, Phillips et al. 2008, Yang, Bhojwani et al. 2008, Hogan, Meyer et al. 2011).
Current therapy for relapsed ALL
The current treatment of relapsed ALL resembles therapy for primary ALL and employs mostly the same chemotherapy agents; however hematopoietic stem cell transplantation now plays a larger role.
In general, patients with low risk or standard risk relapse are assigned to receive 2 more years of chemotherapy with a re-induction phase that can include three to nine blocks of intensive combination chemotherapy (consisting of steroids, vincristine, asparaginase and an anthracycline) and subsequent blocks which include not-yet-administered drugs, such as etoposide and cytarabine (Raetz, Borowitz et al. 2008, Tallen, Ratei et al. 2010). Patients with high-risk relapse, such as early marrow relapse or T-cell
27 Chapter 1 phenotype, are assigned to transplantation following a re-induction multi-drug chemotherapy phase to reduce disease burden. There is a concerted effort to remove MRD prior to the transplant, since studies have shown that MRD level is negatively associated with survival (Bader, Kreyenberg et al. 2009). Cases of CNS or testicular relapse are treated with intensive local and systemic chemotherapy (including high- dose methotrexate) followed by cranial or testicular irradiation respectively (Wofford, Smith et al. 1992,
Barredo, Devidas et al. 2006, Domenech, Mercier et al. 2008).
A novel nucleoside analog, nelarabine, was shown to be selectively cytotoxic to T-lineage cells without affecting B lymphocytes (Gandhi, Keating et al. 2006). Following promising results in clinical trials, nelarabine is now approved as salvage therapy for relapsed T-ALLs and is being investigated in the treatment of newly diagnosed high-risk T-cell malignancies (Berg, Blaney et al. 2005, Kurtzberg, Ernst et al. 2005, DeAngelo, Yu et al. 2007, Gokbuget, Basara et al. 2011, Dunsmore, Devidas et al. 2012).
Despite numerous clinical trials and some promising results, the last 30 years have seen only marginal improvements in survival rates following relapse of ALL (Fielding, Richards et al. 2007, Bailey,
Lange et al. 2008), making the improvement of current therapy and the discovery of novel efficacious and non-toxic anti-leukemic agents ever more pressing.
IV. Novel Agents in the Treatment of ALL
Improvements in current chemotherapy
A number of developments have been made in improving the formulation and delivery of existing chemotherapy drugs which form the core of ALL treatment. It was found that the originally used asparaginase product derived from E. coli was frequently allergenic, leading to serious hypersensitivity reactions and limited efficacy, which translate into increased risk of bone marrow and CNS relapse
(Kawedia, Liu et al. 2012, Liu, Kawedia et al. 2012 26:2303-2309). This was replaced by a pegylated form of E. coli asparaginase (PEG-asparaginase) for first-line treatments, which reduces the possibility of antibody generation while also having a prolonged half-life and increased efficacy (Avramis and
Panosyan 2005). For patients with hypersensitivity reactions or silent antibody-mediated inactivation of E. coli asparaginase or PEG-asparaginase, a third form has been developed from Erwinia chrysanthemia
28 Chapter 1 which is not cross-reactive with the other 2 preparations and can be used as second-line therapy (Pieters,
Hunger et al. 2011).
Administration of vincristine has also been greatly improved with the development of a liposomal preparation (vincristine sulfate liposome injection) which encapsulates the drug in sphingomyelin- and cholesterol-containing nanoparticles (Webb, Logan et al. 1998). This formulation allows for administration of higher doses of vincristine and increased concentration of the drug in relevant tissues and organs, showing greater anti-leukemic activity with decreased toxicity (O'Brien, Schiller et al. 2013).
A number of clinical studies are investigating new chemotherapeutic agents for relapsed ALL, such as the novel nucleoside analog clofarabine, which has shown some benefit in combination treatments (Hijiya, Thomson et al. 2011). A recent study compared two different anthracyclines, mitoxantrone and idarubicin, with early results indicating improved survival after mitoxantrone treatment during remission induction, but this was coupled with a very high risk of severe adverse events (Parker,
Waters et al. 2010).
Precision medicine
The identification of specific molecular lesions and oncogenic pathways in leukemia has led to the development of remarkable targeted therapies for certain cancers, a great example being imatinib, the
ABL1 tyrosine kinase inhibitor that has revolutionized the treatment of chronic myeloid leukemia (CML) which is driven by the BCR-ABL1 (Ph) fusion oncoprotein (Kantarjian, Sawyers et al. 2002). The great success of imatinib in CML prompted the initiation of clinical trials with imatinib and other tyrosine kinase inhibitors like dasatinib in Ph-positive ALL and Ph-like ALL, which are already showing great improvements in outcome for these high-risk groups and could potentially spare some patients from transplantation (Schultz, Bowman et al. 2009, Biondi, Schrappe et al. 2012, Lengline, Beldjord et al. 2013,
Weston, Hayden et al. 2013, Zwaan, Rizzari et al. 2013, Roberts, Li et al. 2014, Schultz, Carroll et al.
2014). Similarly, tyrosine kinase inhibitors are being investigated for the treatment of T-ALLs bearing
ABL1 translocations such as NUP214-ABL1, EML1-ABL1 and ETV6-ABL1 (Quintas-Cardama, Tong et al.
2008)
29 Chapter 1
The activation of the JAK-STAT signaling pathway in Ph-like ALL, Down Syndrome ALL and T-
ALLs (especially ETP-ALL) has prompted studies with JAK inhibitors (Maude, Tasian et al. 2012, Roberts,
Morin et al. 2012, Zhang, Ding et al. 2012, Treanor, Zhou et al. 2014), while the presence of alterations in epigenetic regulators provides great promise for the use of chromatin modifiers like the histone deacetylase inhibitor vorinostat (Burke and Bhatla 2014, Peirs, Van der Meulen et al. 2015). In addition, hypodiploid ALLs and T-ALLs exhibit activation of PI3K/mTOR signaling, suggesting that PI3K and mTOR inhibitors could be efficacious for those subtypes (Gutierrez, Sanda et al. 2009, Holmfeldt, Wei et al.
2013).
The high prevalence of NOTCH1-activating mutations in T-ALL and the role of NOTCH1 in resistance to glucocorticoids (Real, Tosello et al. 2009) have spurred on the development and clinical testing of a number of GSIs, with recent studies showing positive results with minimal toxicity (Samon,
Castillo-Martin et al. 2012). The long-term efficacy of GSI therapy still remains to be seen, with a recent discovery that T-ALL tumor cells can reprogram their metabolism to build resistance to anti-NOTCH1 therapies (Herranz, Ambesi-Impiombato et al. 2015). Other novel potential therapies for T-ALL include targeting of calcineurin activation (Medyouf, Alcalde et al. 2007, Gachet, Genesca et al. 2013) and of the cyclin D3:CDK4/6 complex (Sawai, Freund et al. 2012).
Another exciting development in the search for novel anti-leukemic drugs stemmed from in vitro studies which showed that leukemia cells display high levels of proteasomes and are easily predisposed to undergo apoptosis upon proteasomal inhibition (Kumatori, Tanaka et al. 1990). Bortezomib is a proteasome inhibitor which has shown great response rates in combination with induction chemotherapy for pediatric relapsed or refractory ALL, and is now being investigated in the adult population (Messinger,
Gaynon et al. 2012).
Immunotherapy
The field of immunotherapy (harnessing the power of the immune system to target and kill cancer cells) has grown very rapidly, with a number of promising antibody-based treatments for leukemia showing success in the clinic (Hoelzer and Gokbuget 2012). The majority of current immunotherapies are designed against B-cell malignancies. The first to be approved by the FDA was rituximab, a genetically
30 Chapter 1 modified antibody that recognizes CD20, a molecule found exclusively on B cells (Thomas, O'Brien et al.
2010), which was improved upon to generate ofatumumab and obinutuzumab, which have a higher binding affinity and cytotoxicity (Lemery, Zhang et al. 2010, Sehn, Assouline et al. 2012). CD22 is another
B cell antigen, which has been targeted by the monoclonal antibody epratuzumab (Raetz, Borowitz et al.
2008). CD22 is internalized by B cells after binding by ligand or antibody, which makes it a convenient target for antibody-drug conjugates that can directly deliver a cytotoxin into the leukemic cells (Raponi, De
Propris et al. 2011). This prompted the development of inotuzumab ozogamicin, an anti-CD22 monoclonal antibody conjugated to the DNA-damaging agent calicheamicin, which is currently in clinical trials (de Vries, Zwaan et al. 2012, Kantarjian, Thomas et al. 2012).
Another novel form of immunotherapy is bispecific T-cell engagement (BiTE), which directs the patient’s cytotoxic T-cells to the tumor cells which present a specific antigen. Blinatumomab is the first bispecific antibody that recognizes both CD19 (an antigen found on B-ALL cells) and CD3 (an antigen found on autologous T cells), thereby bringing the T-cells in direct contact with tumor cells and activating their cytotoxic activity (Hoffmann, Hofmeister et al. 2005). Blinatumomab has already shown great efficacy in B-ALL with increased relapse-free survival in a number of studies (Bargou, Leo et al. 2008,
Topp, Kufer et al. 2011, Topp, Gokbuget et al. 2012).
As for T-ALL, there have only been a few studies with alemtuzumab, a monoclonal antibody against CD52, an antigen present on both B and T malignant cells (Rowan, Tite et al. 1998). So far alemtuzumab has shown limited efficacy coupled with significant side effects, which might limit its future application in the treatment of leukemia (Angiolillo, Yu et al. 2009).
Lastly, while BiTE therapy like blinatumomab can effectively direct T-cells to malignant B cells, it is limited in ensuring the expansion of the tumor-directed T cells. This is where recently developed chimeric antigen receptor (CAR)-modified T cells have achieved great success (Kalos, Levine et al.
2011). In CAR T cell therapy, autologous T cells are harvested from the patient and transduced with a
CAR which contains an anti-CD19 antibody fragment connected to intracellular signaling domains which activate the T-cell. This redirects the T-cell to identify and kill B-ALL cells which express the CD19 antigen. The genetically modified CAR T cells are expanded ex vivo and then re-introduced to the patient.
Initial tests in CLL and B-ALL patients showed on-target toxicity, with CAR T cells able to expand in vivo
31 Chapter 1 and home to the bone marrow, where they continue to express functional CARs for at least 6 months and persist as memory CAR T cells (Kalos, Levine et al. 2011, Porter, Levine et al. 2011). Most patients treated with CAR T cells experience a severe cytokine release syndrome which requires very close monitoring but has been successfully managed in the clinic (Grupp, Kalos et al. 2013). CAR T cells have shown remarkable efficacy in inducing remissions in up to 90% of heavily pre-treated ALL patients who have had multiple relapses, however the durability of the remissions varies and is still under investigation long-term (Grupp, Kalos et al. 2013, Maude, Frey et al. 2014).
As our understanding of ALL biology and the mechanisms of ALL relapse grows, so will our arsenal of chemical and biological therapeutic approaches which can effectively complement current risk- based therapy and bring ALL treatment into the era of personalized medicine. This in turn will provide new opportunities for the prevention and treatment of relapsed ALL.
V. The Role of Cytosolic 5’-Nucleotidase II (NT5C2) in Health and Disease
Physiological role
The maintenance of a balanced nucleotide pool is crucial for cellular homeostasis and dependent on a series of catabolic and anabolic pathways of both purines and pyrimidines. Cellular nucleotides are derived either by de novo synthesis, or by uptake and processing of external nucleosides via the salvage pathway. The degree to which de novo synthesis and the salvage pathway are active differs between cell types and depends on the state of each cell (Murray 1971, Pilz, Willis et al. 1984). While de novo nucleoside biosynthesis occurs only in the S phase of the cell cycle, the salvage pathway remains active throughout the cell cycle. Moreover, the salvage pathway is also responsible for the metabolic activation of nucleoside analog prodrugs which are routinely used in the treatment of cancer and viral diseases
(Hunsucker, Mitchell et al. 2005).
Salvaged nucleosides are phosphorylated for incorporation into anabolic pathways by nucleoside kinases, while excess nucleotides are dephosphorylated by intracellular 5’-nucleotidases allowing them to be excreted out of the cell (Bianchi, Ferraro et al. 1994, Gazziola, Ferraro et al. 2001). Cytosolic 5’- nucleotidase II (NT5C2, also known as cN-II) belongs to a family of eight 5’-nucleotidases which all dephosphorylate 5’-nucleoside monophosphates, but have different purine or pyrimidine substrate
32 Chapter 1 specificities and subcellular localizations (Hunsucker, Mitchell et al. 2005). NT5C2 preferentially dephosphorylates the 6-hydroxypurine monophosphates inosine monophosphate (IMP), guanosine monophosphate (GMP) and xanthosine monophosphate (XMP), as well as the deoxyribose forms of IMP and GMP (dIMP and dGMP) (Itoh, Mitsui et al. 1967, Spychala, Madrid-Marina et al. 1988, Tozzi, Camici et al. 1991, Pesi, Turriani et al. 1994, Banditelli, Baiocchi et al. 1996, Allegrini, Pesi et al. 1997). NT5C2 can also process adenosine monophosphate (AMP) and uridine monophosphate (UMP), although far less efficiently (Itoh, Mitsui et al. 1967, Spychala, Madrid-Marina et al. 1988, Tozzi, Camici et al. 1991,
Allegrini, Pesi et al. 1997). The primary substrate of NT5C2, IMP, is a central metabolite which exists at the cross-roads of de novo purine biosynthesis and the purine salvage pathway. IMP generated from either pathway is required for the synthesis of both GMP (via XMP) and AMP (via adenylosuccinate)
(Figure 1.6). Thus, while NT5C2 directly processes IMP, XMP and GMP as part of the purine salvage pathway, NT5C2 can also affect the result of de novo purine biosynthesis through its action on IMP.
Synthesized GMP and AMP are phosphorylated to GDP and ADP which are central precursors to GTP and ATP (for energy and for incorporation into RNA) and, via ribonucleotide reductase (RR), to the deoxyribose forms dGTP and dATP (for use in DNA replication and repair) (Figure 1.6) (Hunsucker,
Mitchell et al. 2005).
Figure 1.6. Purine de novo biosynthesis and salvage pathways, highlighting metabolic conversions catalyzed by NT5C2.
The human NT5C2 gene is located on chromosome 10q24.32 and the cDNA codes for 561 amino acids with a predicted molecular weight of 65 kDa (Oka, Matsumoto et al. 1994). Early studies
33 Chapter 1 indicated that NT5C2 exists as a homotetramer (Spychala, Madrid-Marina et al. 1988, Itoh 1993,
Marques, Teixeira et al. 1998, Spychala, Chen et al. 1999), with recent functional studies and crystal structures confirming that it forms a tetramer made up of a dimer of dimers (Wallden, Stenmark et al.
2007, Pesi, Allegrini et al. 2010, Wallden and Nordlund 2011). Notably, NT5C2 is ubiquitously expressed and highly conserved, with the primary NT5C2 protein sequence sharing 99% identity with other mammalian NT5C2 proteins (cow, dog, brown rat), 95% identity with the enzyme in birds, 65% identity with the Drosophila melanogaster (common fruit fly) enzyme and 51% with the Caenorhabditis elegans
(nematode) enzyme (Itoh 2013). This indicates that NT5C2 plays a very central role in the maintenance of a balanced intracellular nucleoside pool in response to different physical and metabolic stimuli.
Structural basis of NT5C2 activity and allosteric regulation
All 5’-nucleotidases have very similar catalytic mechanisms, which proceed via formation of a phosphoenzyme intermediate (Collet, Stroobant et al. 1998, Allegrini, Scaloni et al. 2001). The intracellular 5’-nucleotidases, while having low sequence homology, are all Mg2+ dependent and share three conserved motifs which place them in the diverse Haloacid Dehalogenase (HAD) superfamily
(Koonin and Tatusov 1994). These three motifs form the catalytic center of the enzyme (Figure 1.7). Motif
I, DXDX(V/T)(L/V), is most integral to the reaction mechanism, in which the first Asp makes a nucleophilic attack on the phosphate of the substrate and the second Asp donates a proton to the departing nucleoside (Koonin and Tatusov 1994, Rinaldo-Matthis, Rampazzo et al. 2002). The first nucleophilic Asp was identified and confirmed to be Asp52 in NT5C2 by functional and structural studies (Figure 1.7)
(Allegrini, Scaloni et al. 2001, Wallden and Nordlund 2011). Motif II is a (S/T) residue (Thr249 in NT5C2) which stabilizes the phosphoenzyme intermediate, and motif III, K(Xx)D(X0-4)D, contains a highly conserved Lys (Lys292 in NT5C2) which also contributes to the stabilization of the intermediate by countering some of the negative charges. In addition, motif III contains Asp351 and Asp356 in NT5C2 which interact with the essential Mg2+ ion involved in the catalysis. Asp54 in motif I also helps to coordinate the Mg2+ ion, while Met53 and Thr56 (motif I) are important for the correct orientation of all Asp residues (Figure 1.7).
34 Chapter 1
The formation of the phosphoenzyme intermediate allows the nucleoside to leave, after which the phosphoenzyme intermediate is hydrolyzed to release the phosphate group. This hydrolysis can be done by a water molecule, resulting in the release of free phosphate, although studies have shown that under specific conditions NT5C2 can also act as a phosphotransferase by donating the phosphate to another nucleoside (Worku and Newby 1982). Recent crystallographic studies have also shed some light on the substrate selectivity of the active site, highlighting Arg202, Asp206, Phe157 and Phe354 as important determinants of purine/pyrimidine selectivity (Wallden and Nordlund 2011).
Figure 1.7. NT5C2 is an HAD family enzyme. (a) Catalytic motifs in the NT5C2 protein sequence. Blue amino acids are catalytic residues conserved in other HAD enzymes; yellow amino acids are uncharged residues with similarities in other HAD enzymes; orange amino acids are C-terminal residues involved in activity regulation. In parentheses is the number of omitted residues. (b) Catalytic amino acids in a schematic active site. Adapted from Bretonnet et al, 2005.
NT5C2 is regulated by a number of allosteric activators, including ATP, dATP, ADP, 2,3- bisphosphoglycerate (2,3-BPG), and diadenosine tetraphosphate (Ap4A, two adenosine molecules connected by four phosphates). ATP/dATP and Ap4A are the strongest activators at physiological concentrations, resulting in decreased Km and increased Vmax (Pinto, Canales et al. 1986, Spychala,
Madrid-Marina et al. 1988, Marques, Teixeira et al. 1998). ATP has been shown to protect the enzyme from heat inactivation and trypsin digestion (Itoh, Usami et al. 1978, Itoh 1982) and to promote auto- oligomerization (Spychala, Chen et al. 1999). On the other hand, inorganic phosphate acts as an NT5C2 inhibitor, by increasing Km without altering Vmax (Pesi, Turriani et al. 1994).
The activation of NT5C2 by ATP and ADP suggests that NT5C2 responds to changes in the adenylate energy charge of the cell (Itoh 1981, Itoh, Oka et al. 1986). However it is also possible that
Ap4A is the more major physiological effector since it activates the enzyme at two orders of magnitude
35 Chapter 1 lower than ATP (Pinto, Canales et al. 1986). While not much is known about the physiological role of
Ap4A, studies have suggested that it is a key signaling molecule during the induction of apoptosis
(Vartanian, Suzuki et al. 2003). Levels of Ap4A increase during apoptosis, while levels of Ap3A (shorter by one phosphate) decrease (Vartanian, Prudovsky et al. 1997), and interestingly NT5C2 is not activated by
Ap3A (Marques, Teixeira et al. 1998). Thus it is possible that Ap4A specifically activates NT5C2 during apoptosis to allow catabolism of the excess purine nucleotides which are released from DNA and RNA degradation, so that membrane-permeable nucleosides can be released for uptake by neighboring cells.
The recent crystal structures of human NT5C2 carrying a substrate-trapping mutation at the catalytic nucleophile Asp52 (D52N) showed the allosterically activated state of NT5C2 with both the IMP substrate and ATP activator bound, which provided a structural understanding of the allosteric mechanism of activation (Wallden and Nordlund 2011). NT5C2 functions as a dimer of dimers, with the dimer being the smallest functional unit with any enzymatic activity (Spychala, Madrid-Marina et al. 1988,
Wallden and Nordlund 2011). The effector binding site is located close to the subunit interface, so that two ATP molecules bind back-to-back to each subunit of the dimer, with their phosphate groups bridged
2+ by a Mg ion (Figure 1.8). Ap4A binds in the same fashion, linking the two neighboring monomers, but without Mg2+ due to the presence of its four phosphates.
Comparison of the inactive (apo) and active structures of NT5C2 revealed a crucial difference in a stretch of residues, Gly355-Glu364, which connects the active site to the effector site. Notably, this stretch is disordered in the apo form, while it constitutes an ordered α-helix (helix A) in the activated enzyme. Binding of an effector (e.g. ATP) in the allosteric pocket results in a conformational change involving the disorder-to-order transition of helix A, which is stabilized by electrostatic interactions between Lys362 and the phosphate moieties of the effector, and by a π-stacking interaction between
Phe354 and the nucleobase of the effector (Figure 1.8). The ordering of helix A results in a rotation of
Phe354 out of the active site, allowing Asp356 of motif III to move into the catalytic center where it helps to organize the active site and to coordinate the Mg2+ ion by rearranging Lys292 and His352 of motif III.
This creates an electrostatically favorable environment which facilitates binding and catalysis of the substrate IMP (Figure 1.8) (Wallden and Nordlund 2011).
36 Chapter 1
Figure 1.8. Mechanism of activation of NT5C2. Left: active site of the effector-free apo NT5C2 enzyme. Dotted line indicates disordered residues. Right: active site and effector site of activated NT5C2. Binding of an allosteric activator (here ATP), results in a disorder-to-order transition of G355-E364 of NT5C2 forming α-helix A. Helix A stabilization organizes the active site, enabling Mg2+ and subsequently the substrate (here IMP) to bind. Adapted from Wallden and Nordlund, 2011.
A second effector site has been tentatively proposed (Pesi, Baiocchi et al. 1998, Wallden,
Stenmark et al. 2007) where only an adenosine has been visualized with a very disordered ribose moiety, however this effector site could not be confirmed by subsequent crystallography studies (Wallden and
Nordlund 2011). Of note, the protein structure around this proposed site is less ordered, including a disordered loop near the surface of the protein (residues 375-417), which is not visible in the reported crystal structures (Wallden and Nordlund 2011).
Finally, the C-terminal end of NT5C2 has a prominent acidic stretch of 13 Asp/Glu residues which suggests that it has a regulatory role (Figure 1.7a). Studies have indicated that it may be involved in subunit association or dissociation, with three positively charged regions proposed as possible interaction partners for the C-terminal acidic tail, K(25)KYRR, K(359)SKKRQ and Q(420)RRIKK (Spychala, Chen et al. 1999). Another study showed that directly upstream of the C-terminal acidic stretch is an important
Cys547 which forms a disulfide bridge with Cys175 under oxidative conditions, leading to auto-inhibition of NT5C2 enzymatic activity (Allegrini, Scaloni et al. 2004). Therefore the C-terminal region of NT5C2 may also act as an auto-regulatory redox switch, but not much is known about it because it appears to be unstructured and flexible and has not been visualized in a crystal structure to date.
37 Chapter 1
Aberrant activity of NT5C2 in human disease
A balanced nucleoside pool is crucial for the development and function of many tissues, and perturbation of nucleoside homeostasis can lead to disease. Examples in humans include immunodeficiencies which result from decreased function of nucleoside-metabolizing enzymes such as adenosine deaminase (ADA) (Giblett, Anderson et al. 1972) or purine nucleoside phosphorylase (PNP)
(Cohen, Gudas et al. 1978). Purine metabolism also plays a large role in the nervous system, with a number of neurological disorders, such as Lesch-Nyhan syndrome, linked to deficiencies in enzymes of the purine biosynthesis pathway (Jinnah, Sabina et al. 2013). Notably, a recent study identified recurrent loss-of-function mutations in NT5C2 and two other genes encoding purine metabolism enzymes, AMPD2 and ENTPD1, in hereditary spastic paraplegias (Novarino, Fenstermaker et al. 2014).
In addition, several enzymes involved in nucleoside metabolism have been implicated in drug resistance to nucleoside analogs used in cancer treatment, particularly nucleoside analog prodrugs which require 5’-phosphorylation for their activation (Galmarini, Graham et al. 2001). For example, drug resistance to cytarabine (a cytosine analog containing an arabinose sugar) has been linked to decreased activity of deoxycytidine kinase (dCK) due to alternative splicing or mutational inactivation, and also to increased expression of drug-inactivating enzymes such as cytidine deaminase and the family of 5’- nucleotidases (Galmarini, Graham et al. 2001).
Several studies have implicated NT5C2 in resistance to nucleoside analog chemotherapy drugs.
Early studies on drug resistant cell lines generated by prolonged incubation with increasing concentrations of nucleoside analogs showed increased NT5C2 expression in those lines (Carson,
Carrera et al. 1991, Schirmer, Stegmann et al. 1998, Dumontet, Fabianowska-Majewska et al. 1999, Lotfi,
Mansson et al. 2001). Clinical reports have shown that high levels of NT5C2 correlate with poor clinical outcome and cytarabine-resistant relapses in AML (Galmarini, Graham et al. 2001, Galmarini, Thomas et al. 2002, Galmarini, Thomas et al. 2003, Galmarini, Cros et al. 2004, Mitra, Crews et al. 2011), decreased efficacy of cladribine in leukemic patients (Kawasaki, Carrera et al. 1993), and poor prognosis in high-risk myelodysplastic syndrome (Suzuki, Sugawara et al. 2007). Additionally, an early study in B-ALL and T-
38 Chapter 1
ALL patients correlated high levels of NT5C2 enzyme activity with resistance to 6-MP and 6-TG (Pieters,
Huismans et al. 1992).
Studies on recombinant NT5C2 showed that it is capable of dephosphorylating the 5’- monophosphate metabolites of a few nucleoside analogs, albeit only at 1-11% the rate of its natural substrate, IMP (Mazzon, Rampazzo et al. 2003). Surprisingly, NT5C2 does not hydrolyze cytarabine monophosphate, which suggested that higher expression levels of NT5C2 may induce cytarabine resistance indirectly though changes in the nucleoside pool (Mazzon, Rampazzo et al. 2003). On the other hand, NT5C2 is able to dephosphorylate 6-thioinosine monophosphate (6-TIMP), 6-thioxanthosine monophosphate (6-TXMP) and 6-thioguanosine monophosphate (6-TGMP) – the key metabolic intermediates in the activation of 6-thioguanine and 6-mercaptopurine (Brouwer, Vogels-Mentink et al.
2005). This highlights NT5C2 activity as a direct determinant of 6-mercaptopurine and 6-thioguanine efficacy and underscores its importance in the treatment of ALL.
The mounting evidence on the role of NT5C2 in drug metabolism prompted the search for NT5C2 inhibitors, with several attempts reported to date. The first described NT5C2 inhibitors were two 5’- modified analogs of adenosine and inosine which were designed to inhibit the release of inorganic phosphate from the phosphoenzyme intermediate (Skladanowski, Sala et al. 1989). The two molecules,
5’-deoxy-5’-isobutylthioadenosine (IBTA) and 5’-deoxy-5’-isobutylthioinosine (IBTI) were characterized as noncompetitive inhibitors to IMP but inhibited enzyme activity at relatively high millimolar concentrations
(Skladanowski, Sala et al. 1989). More recent attempts have investigated a series of non-hydrolyzable substrate analogs, with the identification of a few ribonucleoside phosphonate compounds which show competitive inhibition of NT5C2 activity in the high micromolar to low millimolar range (Gallier, Lallemand et al. 2011, Meurillon, Marton et al. 2014). However, these inhibitors are likely to have poor specificity and are too hydrophilic to cross through the cell membrane. As a result these phosphonates have not yet been evaluated in cell models and any further studies with them would require their development into prodrugs which can be taken up into the cell. Another series of NT5C2 inhibitors consisting of anthraquinone derivatives was identified from virtual screening using the crystal structure of NT5C2
(Jordheim, Marton et al. 2013). These compounds showed competitive NT5C2 inhibition in vitro and mildly sensitized cancer cells to purine nucleoside analogs, yet they are predicted to have poor specificity
39 Chapter 1 and, due to the very high concentrations required to achieve inhibition, have not been tested in vivo
(Jordheim, Marton et al. 2013). Therefore the development of novel, effective and specific NT5C2 inhibitors which can potentiate nucleoside analog therapies still remains a major challenge.
VIII. Specific Aims
Acute lymphoblastic leukemia is an aggressive hematological malignancy which represents the most common cancer in children and adolescents. While intensified chemotherapy has achieved great improvements in overall survival, the outcome of patients with relapsed ALL remains very poor. My central hypothesis is that chemotherapy treatment induces clonal selection of specific mutations which drive disease progression and chemotherapy resistance in relapsed ALL. Thus, my central goal for this work was to identify and functionally characterize the genetic mechanisms driving chemotherapy resistance and clonal selection during treatment and relapse of ALL.
To achieve this objective I proposed the following specific aims:
Aim 1: To identify recurrent genetic alterations in relapsed ALL.
Aim 2: To functionally characterize the role of relapse-specific genetic alterations in promoting drug resistance and relapse of ALL.
Aim 3: To model the process of clonal selection, drug resistance and leukemia relapse in vivo using mouse models of ALL.
40
CHAPTER 2
Materials and Methods
I. Chapter 3
Patient samples
DNAs from leukemic ALL blasts at diagnosis and relapse and matched remission lymphocytes were provided by the Hemato-Oncology Laboratory at University of Padua, Italy; the Eastern Cooperative
Oncology Tumor Bank Laboratory in New York, New York, USA; the Department of Pediatric
Oncology/Hematology at the Charité-Universitätsmedizin Berlin in Berlin, Germany; Saitama Children’s
Medical Center, Saitama, Japan; the Asturias Central Hospital, Oviedo, Spain and the Pediatric Oncology
Division at Columbia University Medical Center. Informed consent was obtained at study entry. We collected and analyzed samples under the supervision of the local Columbia University Medical Center
Institutional Review Board.
Whole-exome capture and next generation sequence analysis
For whole-exome sequencing we selected samples on the basis of the availability of sufficient
DNA from diagnosis, remission and relapse samples, and we evaluated high tumor content at relapse on the basis of copy number analysis of T cell receptor–associated deletions. We used matched diagnostic, remission and relapsed DNA samples from five patients with T-ALL from the University of Padua treated under Associazione Italiana Ematologia Oncologia Pediatrica (AIEOP) protocols (Supplementary Table
A3.1) for exome capture with the SureSelect 50 Mb All Exon kit (Agilent Technologies) following standard protocols. We performed paired-end sequencing (2 × 100 bp) by using HiSeq2000 sequencing instruments at Centrillion Biosciences. Illumina HiSeq analysis produced between 60 million and 120 million paired-end reads per sample. We mapped reads to the reference genome hg19 using the
Burrows-Wheeler Aligner (BWA) alignment tool version 0.5.9. The mean depth (defined as the mean number of reads covering the captured coding sequence of a haploid reference) was 50×, with 80% of the genome covered more than 10× and 57% covered more than 30×. We identified sites that differed from the reference (called here variants) in each sample independently. We constructed empirical priors for the
41
Chapter 2 distribution of variant frequencies for each sample. We obtained high-credibility intervals (posterior probability ≥1–10−5) for the corresponding change in frequency between tumor and normal samples using the SAVI (Statistical Algorithm for Variant Identification) algorithm developed at Columbia University
(Pasqualucci, Trifonov et al. 2011, Tiacci, Trifonov et al. 2011). The number of germline SNPs in the coding region was 18,000, which is comparable with previous reports (Tiacci, Trifonov et al. 2011). Most of the candidate germline SNPs (16,000, or ~90% of germline variants) were reported in the dbSNP database. We identified candidate somatic variants using the following criteria: variant total depth in tumor and normal >10× and <300×, variant frequency >15% in tumor and <3% in normal and ≥1% change in frequency from the normal with high posterior probability (≥1–10−5). Also, to remove systematic errors, we excluded all variants that were found in unaffected individuals. In addition, to eliminate ambiguous mapping from captured pseudogenes and regions of low complexity, each variant with a flanking 20-base context sequence around its genomic position was mapped to the hg19 reference using the BLAST algorithm. We kept in the list only those with unique mappability; that is, we required the 41-base sequence to uniquely map to the reference genome with only one mismatch.
To discern the regions of loss of heterozygosity (LOH), we used the SAVI-calculated high- credibility intervals for the variants in dbSNP, which correspond to the change in their frequency between tumor and normal samples. In an LOH event, depending on whether the reference or the dbSNP allele was lost, at least a 1% or at most a −1% change in frequency from the normal is expected. Therefore, by segmenting the regions covering more than ten dbSNP variants with significantly changed frequencies, we were able to identify the LOH regions.
Mutation validation and analysis of recurrence
We designed primers flanking exons containing candidate somatic variants using Primer3
(http://frodo.wi.mit.edu/primer3/) and used them for PCR amplification from whole genome–amplified
(WGA) tumor, relapse and matched normal (remission) DNAs. We analyzed the resulting amplicons by direct bidirectional dideoxynucleotide sequencing with a validation rate of 97%. After exome sequence analysis of 5 diagnostic relapse and remission T-ALL AIEOP samples from the University of Padua
(Supplementary Table A3.1), we used 18 additional patient samples from the same institution for the
42 Chapter 2 analysis of recurrence of TP53, BANP, RPL11, NRAS and NT5C2 (Supplementary Table A3.1). We subsequently extended this series to additional relapse T-ALL samples from the University of Padua (n =
13) and the Charité-Universitätsmedizin Berlin (n = 67) (Supplementary Table A3.1) and to relapsed patients with B-precursor ALL from the University of Padua (n = 35) and from Charité-Universitätsmedizin
Berlin (n=466, complete results to be published soon) for extended mutation analysis of NT5C2
(Supplementary Table A3.1). Subsequently we have performed exome sequencing analyses of matching diagnosis, remission and relapsed T-ALL samples from the Children’s Oncology Group (n=20) and University of Padua (n=1), and of matching triplet B-precursor ALL samples from Saitama Children’s
Medical Center (n=7), Hospital Central de Asturias (n=2) and Columbia University Medical Center (n=5), the complete results of which will be reported in an upcoming manuscript (submitted). We used two cohorts of diagnostic patients with T-ALL from ECOG (n = 23) and diagnostic patients with B-precursor
ALL from the University of Padua (n = 27) to verify the absence of NT5C2 mutations in diagnostic ALL specimens (Supplementary Table A3.1).
Structural depiction and analysis (Z. Carpenter)
We identified structural coverage of the NT5C2 protein through use of the PSI-Blast and SKAN algorithms; we subsequently mapped viable structures to all NT5C2 isoforms and analyzed them using
Chimera Suite (Altschul, Madden et al. 1997, Pettersen, Goddard et al. 2004). We aligned structurally the
PDB structures 2XCW, 2XCX, 2XCB, 2XCV, 2XJB, 2XJC, 2XJD, 2XJE, 2XJF, 2J2C and 2JC9 and subsequently analyzed the composite structure to assess conformational flexibilities (Pettersen, Goddard et al. 2004). We structurally modeled NT5C2 mutations using the I-TASSER software suite and subsequently refined and analyzed them by minimization and rotamer library analysis in Chimera
(Pettersen, Goddard et al. 2004, Roy, Kucukural et al. 2010). We predicted protein stability changes resulting from mutation through use of the SDM potential energy statistical algorithm and associated software (Worth, Preissner et al. 2011). We created all structural images using UCSF Chimera
(Pettersen, Goddard et al. 2004).
43 Chapter 2
Site-directed mutagenesis
We generated the NT5C2 mutations by site-directed mutagenesis on the mammalian expression pLOC-NT5C2 vector (Open Biosystems) using the QuikChange II XL Site-Directed Mutagenesis Kit
(Stratagene) according to the manufacturer's instructions.
Cell lines
We cultured CCRF-CEM and CUTLL1 cells in RPMI-1640 medium supplemented with 10% FBS,
−1 −1 100 U ml penicillin G and 100 μg ml streptomycin at 37 °C in a humidified atmosphere under 5% CO2.
We maintained HEK293T cells under similar conditions in DMEM media.
Lentiviral production and infection
We transfected the lentiviral constructs containing pLOC-NT5C2 (NT5C2 wildtype or NT5C2 mutant), or the pLOC-RFP control plasmid with Gag-Pol– and V-SVG–expressing vectors into HEK293T cells using JetPEI transfection reagent (Polyplus). We collected viral supernatants after 48 h and used them for infection of CCRF-CEM and CUTLL1 cells by spinoculation. After infection, we selected cells for
5 d in blasticidin and ficolled them the day before experiments.
Western blot
Western blot analysis was performed using a rabbit polyclonal antibody to NT5C2 (1:1,000,
Abcam, ab96084) and a goat polyclonal antibody to glyceraldehyde 3-phosphate dehydrogenase
(GAPDH) (1:1,000, Santa Cruz Biotechnology, sc-20357) using standard procedures.
Cell viability and chemotherapy drug response
We determined cell viability by measurement of the metabolic reduction of the tetrazolium salt
MTT using the Cell Proliferation Kit I (Roche) following the manufacturer's instructions. We performed experiments in triplicate. We analyzed viability at 48 h or 72 h after initiation of treatment with 6-MP, 6-TG, nelarabine and AraG.
44 Chapter 2
Recombinant protein production and purification
We cloned full-length complementary DNA constructs encoding wild-type or mutant NT5C2 with an N-terminal hexahistidine (His6) tag in the pET28a-LIC expression vector using the In-Fusion HD PCR cloning system (Clontech) as per the manufacturer's instructions. We expressed recombinant proteins from Rosetta 2(DE3) Escherichia coli cells by induction with 0.5 mM isopropyl-β-D-thiogalactopyranoside for 3 h at 37 °C. We harvested cells and lysed them in lysis buffer (50 mM sodium phosphate, pH 7.4, 100 mM NaCl, 10% glycerol, 5 mM β-mercaptoethanol, 1% Triton X-100, 0.5 mg ml−1 lysozyme and 20 mM imidazole) supplemented with Complete EDTA-free protease inhibitor (Roche). We purified His6-tagged
NT5C2 proteins by binding them to nickel-Sepharose beads and eluting them with 50 mM sodium phosphate, pH 7.4, 100 mM NaCl, 10% glycerol, 5 mM β-mercaptoethanol and 300 mM imidazole. We removed imidazole by buffer exchange using PD-10 desalting columns (GE Healthcare). We assessed protein expression and purity by SDS-PAGE and Coomassie staining.
5′-nucleotidase assays
We assessed 5′-NT activity of purified recombinant wild-type and mutant NT5C2 proteins using the 5′-NT Enzymatic Test Kit (Diazyme) according to the manufacturer's instructions. The assay measures the enzymatic hydrolysis of inosine 5′-monophosphate to inosine, which is reacted further to hypoxanthine by purine nucleoside phosphorylase and then to uric acid and hydrogen peroxide by xanthine oxidase. H2O2 is quantified using a Trinder reaction. We calculated 5′-NT activity levels using a calibrator of known 5′-NT activity as standard. We performed assays in triplicate in an Infinite M200 Tecan plate reader.
Single cell PCR analysis
To perform single cell PCR analyses on NT5C2 exon 9 (to test for R238W mutation) and NT5C2 exon 13 (to test for L375F mutation), we collected relapsed tumor cells by scraping from a viably frozen vial into PBS containing 1% BSA. We washed and resuspended cells in 500ul PBS with 1% BSA and sorted single cells by flow cytometry into 96-well PCR plates, containing 4µL PBS sc per well from Repli-g
Single Cell Kit (Qiagen). We performed whole genome amplification on the sorted single cells in the 96-
45 Chapter 2 well plates using Repli-g Single Cell Kit (Qiagen), following the manufacturer’s instructions with a few changes. Specifically, we used 2µL of buffer D2, 2 µL of STOP solution and 20 µL of DNA polymerization master mix per reaction. We performed PCR reactions for NT5C2 exon 9 and 13 using 2 µL of a 1:100 dilution of the amplified DNA as template and analyzed PCR products by Sanger sequencing.
Statistical analyses
We evaluated differences in the percentages of wild-type and mutant NT5C2 in patients with ALL in different relapsed categories using Fisher's exact test. We analyzed the equality of categorical and continuous variables by Fisher's exact test and Mann-Whitney U test, respectively.
II. Chapter 4
Mouse procedures and generation of Nt5c2 R367Q conditional knock-in
We maintained all animals in specific pathogen-free facilities at the Irving Cancer Research
Center at Columbia University Medical Campus. The Columbia University Institutional Animal Care and
Use Committee (IACUC) approved all animal procedures. To generate conditional Nt5c2 R367Q knock-in mice we used homologous recombination in C57BL/6 embryonic stem cells to introduce the R367Q point mutation (AG→CA) in exon 14 (two nucleotide changes required due to alternate codon usage in the mouse) as well as a LoxP-flanked wildtype mini-gene cassette (1958 bp) inserted 233bp upstream of exon 14, composed of the fusion of exons 14-18 and the flanking genomic sequences upstream of exon
14 and downstream of exon 18. Immediately downstream of the mini-gene we introduced a FRT-flanked neomycin selection cassette. We generated chimeras in C57BL/6 albino blastocysts using three independent targeted embryonic stem cell clones identified by PCR analysis and verified by Southern blot
(inGenious Targeting Laboratory). We verified germline transmission in the offspring of highly chimeric male mice crossed with C57BL/6 females. To remove the neomycin selection cassette we crossed mice harboring the targeting construct with a Flp germline deleter line (B6;SJL-Tg(ACTFLPe)9205Dym/J, the
Jackson Laboratory) and crossed the resulting mice with wild type C57BL/6 to breed out the Flp allele. To generate conditional inducible Nt5c2 R367Q knock-in mice we bred animals harboring the conditional
46 Chapter 2
Nt5c2 R367Q allele with Rosa26-CreERT2 mice, which express a tamoxifen-inducible form of the Cre recombinase from the ubiquitous Rosa26 locus (Guo, McMinn et al. 2007).
Generation of NOTCH1-induced leukemias
To generate NOTCH1-induced T-ALL tumors in mice, we performed retroviral transduction of bone marrow cells (from Rosa26-CreERT2 Nt5c2 R367Q wt/flox mice) enriched in lineage negative cells isolated using magnetic beads (Lineage Cell Depletion Kit, Miltenyi Biotec) with an activated form of the
NOTCH1 oncogene (ΔE-NOTCH1) (Schroeter, Kisslinger et al. 1998) and transplanted them via intravenous injection into lethally irradiated isogenic recipients (6-8 week old C57BL/6 females, Taconic
Farms) as previously described (Chiang, Xu et al. 2008). The ΔE-NOTCH1 retroviral construct also expresses green fluorescent protein (GFP) allowing identification of tumor cells by FACS analysis.
We assessed T-ALL tumor development by monitoring for a transient increase in CD4+CD8+ double positive cells in peripheral blood at 3 weeks post-transplant by incubating blood samples with red blood cell lysis buffer (155 mM NH4Cl; 10 mM KHCO3; 0.1 mM EDTA) for 3x 5 min at room temperature before staining with APC-Cy7-conjugated antibodies against mouse CD4 (BD Pharmingen-552051) and
PE-Cy7-conjugated antibodies against mouse CD8a (eBioscience-25-0081-82). Flow cytometry analyses were performed in a FACSCanto flow cytometer (BD Biosciences).
In vivo mouse experiments
For all subsequent in vivo studies, we harvested fresh Rosa26-CreERT2 Nt5c2 R367Q wt/flox T-
ALL tumor cells from the spleens of donor mice and transplanted them into sublethally irradiated (4 Gy) secondary recipients (C57BL/6 females, 6-8 weeks old, Taconic Farms). Animals were randomly assigned to different treatment groups and no blinding was done. For survival and leukemia initiating cells experiments, two days following transplantation we treated the mice with tamoxifen (3mg/mouse via intraperitoneal injection) to induce mini-gene deletion and expression of Nt5c2 R367Q in the leukemic cells, or with corn oil vehicle for control groups. Mice were then observed for incidence and time of onset of leukemia.
47 Chapter 2
For acute treatment studies we used Rosa26-CreERT2 Nt5c2 R367Q wt/flox T-ALL tumor cells which were infected with lentiviral particles expressing the red cherry fluorescent protein and luciferase
(FUW-mCherry-Luc-puro) and selected with puromycin. Following transplantation, C57BL/6 recipients were monitored by in vivo luminescence bioimaging with the In Vivo Imaging System (IVIS, Xenogen) and analysis of GFP positive cells in the peripheral blood. Once mice had 50% GFP positive cells in the peripheral blood and a detectable baseline tumor burden by bioluminescence, we treated them with 3mg tamoxifen or corn oil vehicle as described above and allowed them to recover over two days. We then initiated treatment with a range of doses of 6-mercaptopurine (0, 50, 60, 80, 100 mg/kg/day) via intraperitoneal injection for 5 consecutive days, monitoring disease progression and response to chemotherapy by bioluminescence imaging on days 0, 3 and 6 after start of 6-mercaptopurine treatment.
We sacrificed mice on day 6 and analyzed spleen weight, then processed spleens through a 70 μm mesh to obtain single cell suspensions, incubated the samples with red blood cell lysis buffer and analyzed spleen cellularity and GFP content.
Drugs and Administration
We purchased tamoxifen, 4-hydroxytamoxifen, 6-mercaptopurine (6-MP) and 6-thioguanine (6-
TG) from Sigma-Aldrich. For in vitro assays we dissolved 4-hydroxytamoxifen in 100% ethanol, 6-MP in
DMSO and 6-TG in 1M NaOH. For in vivo studies we resuspended tamoxifen in 100 uL of ethanol/100 mg and added corn oil to reach a final concentration of 3mg/100ul. We then rotated the tamoxifen suspension for 1 hour at 55°C and froze in aliquots at -20°C. We administered tamoxifen as a single
100ul intraperitoneal injection per mouse. For intraperitoneal injections of 6-MP we prepared frozen aliquots of 5 mg/ml 6-MP in 0.1N NaOH and immediately prior to each round of treatment we prepared fresh final solutions of 6-MP by buffering the stock solution down to pH 8 with 0.2M NaH2PO4. This resulted in a 6-MP concentration of 3.48 mg/ml, which we diluted to various final concentrations using a solution made from 0.05N NaOH and 0.2M NaH2PO4 adjusted to pH 8. Due to large injection volumes required to achieve intraperitoneal doses of 6-MP greater than 50mg/kg/day, we treated mice twice daily.
For groups receiving 0 mg/kg/day 6-MP we prepared vehicle by dissolving 0.254g NaCl per 50ml 0.05N
48 Chapter 2
NaOH and adjusting to pH 8 with 0.2M NaH2PO4. We weighed mice and adjusted injections to correct for any differences in mass.
Cell culture
We harvested primary mouse tumor cells from the spleens of leukemic mice by processing spleens through a 70 μm mesh to obtain single cell suspensions and incubating with red blood cell lysis buffer. Tumor cells were then placed in culture in OptiMEM media supplemented with 10% FBS, 1%
Penicillin/Streptomycin, 1X β-mercaptoethanol, 10 ng/ml IL7 and 10 ng/ml IL2. Subsequent passages of tumor cells did not include IL2. In case initial monoculture proved difficult, we also placed some of each tumor sample on a monolayer of OP9 feeder cells which were gradually removed over 2 weeks.
We purchased HEK 293T cells for viral production from ATCC and cultured them in standard conditions in DMEM media supplemented with 10% FBS and 1% Penicillin/Streptomycin.
Retroviral and lentiviral infections
We transfected retroviral or lentiviral plasmids together with gag-pol and V-SVG expressing vectors into HEK293T cells using JetPEI transfection reagent (Polyplus). We collected viral supernatants after 48h and used them to infect mouse bone marrow progenitors or primary tumor cells by spinoculation. We selected infected primary mouse tumor cells with puromycin for 5 days.
Genotyping of conditional Nt5c2 R367Q induction
To detect tamoxifen-inducible mini-gene deletion we purified DNA from primary Rosa26-CreERT2
Nt5c2 R367Q wt/flox mouse tumor cells treated with 1µM 4-hydroxytamoxifen or ethanol vehicle (in vitro) and tamoxifen or corn oil vehicle (in vivo) and PCR-amplified in a three-primer reaction the minigene cassette (primer immediately upstream of proximal LoxP site and reverse primer in exon 17), the deleted mini-gene allele and the wild type allele (primer immediately upstream of proximal LoxP site and reverse primer in intron 14). The deleted and wild type alleles differ by the size of the remaining LoxP site (49 bp).
We visualized PCR products with ethidium bromide on a 1.5% agarose gel.
49 Chapter 2
To detect tamoxifen-inducible expression of Nt5c2 R367Q mRNA we purified total RNA from mouse tumor cells with the RNeasy kit (Qiagen), prepared complementary DNA (cDNA) by reverse transcription using the SuperScript First-Strand Synthesis System for RT-PCR (Invitrogen) and PCR amplified the Nt5c2 exon 14 cDNA region using primers spanning neighboring exons. We sequenced resulting PCR products with the PCR primers and assessed the presence of the engineered Nt5c2
R367Q nucleotide substitutions (AG→CA).
In vitro cell viability and chemotherapy responses
We measured cell growth and chemotherapy responses of primary mouse tumors in vitro by measurement of the metabolic reduction of the tetrazolium salt MTT using the Cell Proliferation Kit I
(Roche) following the manufacturer’s instructions. We analyzed chemotherapy responses of tumor cells following 72-hour incubation with increasing concentrations of 6-mercaptopurine or 6-thioguanine.
Patient samples
DNAs from leukemic T-ALL blasts at diagnosis and relapse and matched remission lymphocytes were provided by the Hemato-Oncology Laboratory at University of Padua, Italy and the Children’s
Oncology Group. All samples were collected in clinical trials with informed consent and under the supervision of local IRB committees. Consent was obtained from all patients at trial entry according to the
Declaration of Helsinki. Genomic DNA was extracted from patient leukemic blasts or from the lymphoid fraction of peripheral blood at remission using the Qiagen DNeasy Blood & Tissue Kit.
Targeted ultra-deep sequencing of diagnostic and remission samples
We designed primers for amplifying genomic regions (200 bp ± 20 bp) of relapse-associated
NT5C2 mutations according to Fluidigm recommendations and included Fluidigm-specific adapter sequences at the 5’ ends. After PCR amplification from 200 ng of diagnostic or remission DNA, we added
Fluidigm barcodes so that all amplicons corresponding to one sample carry the same index. We pooled all indexed amplicons and quantified the resulting library by quantitative PCR using a Kapa Library
Quantification Kit (Kapa Biosystems) in a 7500 Real-time PCR system (Applied Biosystems). To increase
50 Chapter 2 amplicon diversity we spiked the amplicon library with ~25% PhiX genomic library. We sequenced a library containing samples T-ALL 5, 11, 15, 17, 22 and 35 in a MiSeq instrument to generate 2×251 bp paired end reads using a sequencing protocol for custom primers. Given that the amplicon sequences
(200 bp ± 20 bp) were shorter than the read length, we stitched together paired end reads using FLASH version 1.2.6 (Fast Length Adjustment of SHort reads), This step increases the quality of the reads correcting for mismatches in the overlap by selecting the base with higher quality. Then we trimmed 5' and 3' adaptors and PCR primer sequences using cutadapt and aligned trimmed reads to the UCSC hg19 reference genome using BWA-MEM as single-end reads. We used SAVI (Statistical Algorithm for Variant
Identification) to analyze reads for variants, selecting variants based on coverage depth and frequency.
We sequenced a library containing samples T-ALL11, T-ALLs C4, C5, C7, C10, C11, C14, C17 and C20, and B-ALL Ko4 using a NextSeq instrument to generate 2×251 bp paired end reads and we improved the quality of the reads by masking any mismatch within each read-pair. Stitched reads were trimmed to exclude primer, adapter, and over-hang sequences. Trimmed reads were then aligned to reference genome hg19 as single-end reads using STAR aligner and analyzed using the SAVI algorithm.
Quantitative NT5C2 p.R367Q allele specific qPCR assay
We quantitatively assessed the presence of NT5C2 p.R367Q in diagnostic and remission DNA samples from matching relapsed cases with the NT5C2 p.R367Q mutation using a custom Mutation
Detection Competitive Allele-Specific TaqMan® PCR (castPCR™) Assay (Life Technologies). We followed the manufacturer’s instructions using 30ng of DNA, 2x Taqman Universal PCR Master Mix and
20x custom Taqman assay (NT5C2 wild type or NT5C2 p.R367Q) in a reaction volume of 20µl. We performed all analyses in a 7500 real-time PCR system (Applied Biosystems) using the manufacturer’s recommended cycling conditions. We determined a detection ΔCT cutoff value for the assay by running the NT5C2 wild type and NT5C2 p.R367Q assays on three wild type cell line genomic DNA samples and calibrated both assays by running on increasing concentrations of NT5C2 wild type or NT5C2 p.R367Q mutant plasmids respectively. We determined the NT5C2 p.R367Q assay sensitivity by running on wild type cell line genomic DNA spiked with increasing concentrations of NT5C2 p.R367Q plasmid. We analyzed experimental data using Mutation Detector™ Software (Life Technologies), calculating the ΔCT
51 Chapter 2 value between the NT5C2 wild type and NT5C2 p.R367Q assay reads for each sample, and comparing to the predetermined ΔCT cutoff value. Based on these results we determined the assay mutation detection threshold as 1:1000.
Metabolomic analyses
To analyze metabolic differences between Nt5c2wt/wt and Nt5c2wt/R367Q primary mouse tumors we treated tumor cells in triplicate with 1µM 4-hydroxytamoxifen for 48h in vitro to induce expression of the
Nt5c2 R367Q allele or with vehicle for wildtype controls, after which we diluted out the 4- hydroxytamoxifen or vehicle and allowed cells to recover over 72h. We then harvested cells into packed pellets of 50-100µl size and took samples of conditioned media from each. We flash froze pellets and media samples, which were then extracted and analyzed on the GC/MS and LC/MS/MS platforms in
Metabolon facilities. The dataset comprised a total of 459 named biochemicals in cells and 252 named biochemicals in media. We first normalized results to protein concentration, log transformed and imputed any missing values with the minimum observed value for each compound. We then used Welch’s two- sample t-test to identify biochemicals that differed significantly between experimental groups. To account for the multiple comparisons that occur in metabolomics studies we also calculated an estimate of the false discovery rate (q-value), which indicates the fraction of biochemicals that would meet a given p- value cut-off by random chance.
Statistical analyses
We performed statistical analysis by Student’s t-test. Survival in mouse experiments was represented with Kaplan-Meier curves and significance was estimated with the log-rank test (GraphPad
Prism). We analyzed serial limited dilution leukemia-initiating cell data using the ELDA software (Hu and
Smyth 2009).
III. Chapter 5
Recombinant protein production and purification
For 5’-nucleotidase assays in the absence and presence of allosteric activators we cloned,
52 Chapter 2 expressed and purified recombinant wildtype and mutant NT5C2 proteins as previously described
(Section I of this Chapter) (Tzoneva, Perez-Garcia et al. 2013).
For X-ray crystallography analyses we cloned full-length (561 amino acids) or C-terminally truncated (amino acids 1-536) complementary DNA constructs encoding wild-type or mutant NT5C2 with an N-terminal hexahistidine (His6) tag in the pET28a-LIC expression vector. We expressed recombinant proteins from Rosetta 2(DE3) Escherichia coli cells by induction with 0.5 mM isopropyl-β-D- thiogalactopyranoside overnight at 16 °C. We resuspended harvested cells in lysis buffer (50 mM sodium phosphate pH 7.4, 500 mM sodium chloride, 10% glycerol, 0.5 mM TCEP, 20 mM imidazole) supplemented with Complete EDTA-free protease inhibitor (Roche) and lysed cells by sonication. We purified recombinant proteins using an ӒKTA fast protein liquid chromatography system (GE Healthcare) using a 2-step protocol adapted from one previously described (Wallden, Stenmark et al. 2007). We first performed affinity chromatography using a 1 ml Ni2+-charged His-Trap HP column (GE Healthcare) equilibrated in lysis buffer. We eluted NT5C2 proteins from the His-Trap column in a step-wise method with elution buffer (lysis buffer with 500mM imidazole) by first setting the buffer ratio to 25% elution buffer for 8 column volumes and then switching to a linear gradient to 100% elution buffer over 10 column volumes. We pooled NT5C2-containing fractions and purified further by size exclusion chromatography using a HiLoad 16/60 Superdex 200 gel filtration column (GE Healthcare) equilibrated in 50 mM sodium phosphate, pH 7.4, 100 mM NaCl, 10% glycerol and 0.5 mM TCEP. We assessed protein expression and purity by SDS-PAGE and Coomassie staining and concentrated protein samples to 4-9 mg/ml.
For 5’-nucleotidase assays of NT5C2 proteins incubated with Loop or C-terminal polyclonal antibodies and/or specific blocking peptides, we expressed proteins as described for standard 5’- nucleotidase assays (Section I of this Chapter) (Tzoneva, Perez-Garcia et al. 2013). We purified His6- tagged NT5C2 proteins in a 2-step process. First we bound proteins to nickel-Sepharose beads and eluted them with 50 mM sodium phosphate, pH 7.4, 100 mM NaCl, 10% glycerol, 5 mM β- mercaptoethanol and 300 mM imidazole. We then further purified NT5C2 protein eluates by size exclusion using a HiLoad 16/60 Superdex 200 gel filtration column (GE Healthcare) equilibrated in 50 mM sodium phosphate, pH 7.4, 100 mM NaCl, 10% glycerol and 0.5 mM TCEP.
53 Chapter 2
5′-nucleotidase assays
We assessed 5′-NT activity of purified recombinant wild-type and mutant NT5C2 proteins using the 5′-NT Enzymatic Test Kit (Diazyme) according to the manufacturer's instructions as described previously (Section I of this Chapter) (Tzoneva, Perez-Garcia et al. 2013). For assays with allosteric activators ATP, ADP and Ap4A (all from Sigma Aldrich) we dissolved powders directly in Reagent 2 of the test kit (containing the substrate IMP) and made serial dilutions to achieve a range of concentrations. For assays with antibodies and peptides (custom-generated by Covance), both antibodies and peptides were resuspended in phosphate buffered saline (PBS) and all corresponding samples and controls were diluted with PBS accordingly. We performed statistical analysis by Student’s t-test.
Structural modeling analyses (Z. Carpenter)
Computational modeling analyses were performed by Dr Zachary Carpenter as described previously (Section I of this Chapter) (Tzoneva, Perez-Garcia et al. 2013) and images were generated using Chimera suite (Pettersen, Goddard et al. 2004). Additional modeling was conducted using Modeller and I-TASSER webserver (Eswar, Eramian et al. 2008, Roy, Kucukural et al. 2010). Loop models were built, refined and scored using Modeller Software suite (Fiser, Do et al. 2000, Eswar, Eramian et al.
2008). Top models for figures were selected by ranking 5000 iterations by DOPE score as described elsewhere (Fiser, Do et al. 2000).
Protein cavity analysis was performed using CASTp Webserver and Chimera Interface (Dundas,
Ouyang et al. 2006). NT5C2 structure PDB files were re-written to contain quaternary structure coordinates. These files were subsequently used as input into the CASTp algorithm with a probe radius set at the default 1.4 Angstroms. HET groups were included in calculations that were set to proceed on any combination of chains from each structure.
Protein stability changes upon mutation were assessed using the SDM potential energy algorithm and webserver on re-written tetrameric NT5C2 structure PDB files (Worth, Preissner et al. 2011). In-silico mutagenesis was conducted with Modeller Software suite utilizing the SDM output as a starting template structure (Eswar, Webb et al. 2006, Eswar, Eramian et al. 2008). All secondary structure prediction was conducted using the Predict Protein platform and webserver (Rost, Yachdav et al. 2004).
54 Chapter 2
Path prediction and molecular dynamics of NT5C2 models were predicted using UCSF Chimera software (Pettersen, Goddard et al. 2004). Minimization of mutants was conducted using a 6-angstrom sphere to specify fixed and unfixed atoms, 1000 steepest descent steps, steepest descent step size of
0.02 angstroms and 100 conjugant gradient steps of 0.02 angstroms each. Residues with one or more atoms included in a sphere were included in their entirety regardless of full inclusion in 6 angstroms from any point of the mutant residue. Hydrogen bonding networks and Van der Waals clashing were predicted using UCSF Chimera Software (Pettersen, Goddard et al. 2004).
Crystallization and structure determination (F. Forouhar)
We grew all crystals of the human truncated wildtype (WT) and mutant NT5C2 537X proteins
(residues 1-536) using the microbatch method at 18 °C. We grew all crystals of full-length WT and mutant
NT5C2 proteins at 4 °C for the first week, followed by transferring the crystallization plates to 18 °C for the next 3 weeks. We crystallized the apo WT NT5C2 537X protein by mixing 2 µl of protein solution (8.5 mg/ml in 50 mM Na3PO4 pH 7.4, 100 mM NaCl, 10% glycerol, 0.5 mM TCEP) with 2 µl precipitant solution
(2 M ammonium sulfate and 100 mM sodium acetate trihydrate pH 4.6). To cryoprotect the crystals we supplemented the crystallization solution with 20% (v/v) glycerol and flash-froze it in liquid nitrogen for data collection at 100 K. We collected a single-wavelength native data set to resolution 2.8 Å at the X4C beamline of the National Synchrotron Light Source (NSLS) and processed the diffraction images with the
HKL package (Otwinowski 1997). The crystals of WT apo enzyme belong to space group C2 and there are two protomers of the apo enzyme in the asymmetric unit (ASU) of the crystal. We used the structure of human NT5C2 537X in complex ATP, Mg2+, and IMP (PDB id: 2XCW) (Wallden and Nordlund 2011) to determine the structure of apo WT NT5C2 537X using the molecular replacement method with the program COMO (Jogl 2001). We manually built the NT5C2 flexible loop and four additional residues (489-
492) of the C-terminal domain with the program XtalView (McRee 1999). All stages of the structure refinement were performed using CNS 1.3 (Schroder, Levitt et al. 2010). For any dataset diffracting better than 2.5 Å, we used the crystallographic program PHENIX in the last stage of refinement (Adams,
Afonine et al. 2010).
55 Chapter 2
We grew crystals of NT5C2 D52N 537X in complex with ATP, Mg2+, and IMP using the same methodology, except the precipitation reagents were composed of 100 mM ammonium citrate tribasic (pH
7) and 12% (w/v) PEG 3350. The crystals of complexed NT5C2 D52N 537X diffract to 2.9 Å and belong
2+ to space group P6522. There are two protomers in complex with ATP, Mg , and IMP per ASU of the crystals. For crystallization of NT5C2 mutant proteins D52N R238W, D52N R367Q, D52N K359Q and
D52N L375F we used four different crystallization conditions, each having a buffer near the physiological pH (i.e., pH 7-7.5). We grew crystals of NT5C2 D52N R367Q 537X in the presence of ATP, Mg2+ and IMP using the crystallization cocktail consisting of 100 mM ammonium nitrate, 100 mM HEPES (pH 7.5), and
20% (w/v) PEG 1000. Crystals diffract to 2.34 Å and belong to space group C2. There are two protomers in ASU of the crystal. IMP has well-defined electron density, whereas ATP and Mg2+ both have poor electron densities. We grew crystals of NT5C2 D52N K359Q 537X in a crystallization cocktail comprising
0.1 M ammonium bromide, 100 mM HEPES (pH 7.5), and 20% (w/v) PEG 1000. Crystals diffract to 1.8 Å and belong to space group I222 with one protomer in ASU. We grew crystals of NT5C2 D52N L375F
537X apo enzyme using the crystallization cocktail consisting of 1.8 M ammonium citrate tribasic (pH 7).
Crystals diffract to 2.9 Å and belong to space group I222 with one protomer per ASU of the crystal. We grew crystals of NT5C2 D52N R238W 537X in the presence of ATP, Mg2+ and IMP using the crystallization cocktail consisting of 35% (v/v) tascimate (pH 7). Crystals diffract to 1.9 Å and belong to space group I222, in each ASU of which there is one protomer in complex with ATP, Mg2+, and IMP.
We crystallized the full-length WT enzyme in the presence of ATP, Mg2+ and IMP using the seeding method and a crystallization cocktail comprising 100 ammonium nitrate, 100 mM HEPES (pH
7.5) and 20% (w/v) PEG 1000. Crystals diffract to 2.54 Å and belong to space group C2 with one dimer
(two protomers) in ASU. The structure is strikingly similar to 2XCW. We grew crystals of full-length apo
NT5C2 D52N R367Q using the crystallization condition consisting of 200 mM L-proline, 100 mM HEPES
(pH 7.5), and 10% (w/v) PEG 3350. Crystals diffract to 2.32 Å and there are two protomers in ASU of the crystals with mostly ordered C-terminal domains not observed previously. We obtained crystals of full- length apo NT5C2 D52N R39Q by the seeding method with a condition of 100 mM HEPES (pH 7.5) and
12% (w/v) PEG 3350. Crystals diffract to 2.9 Å and belong to space group C2221, and there are two protomers in ASU of the crystals. This mutant structure also reveals mostly ordered C-terminal domain.
56
CHAPTER 3
Activating mutations in the NT5C2 nucleotidase gene drive chemotherapy
resistance in relapsed ALL
A large component of the work described in this chapter was published in “Tzoneva et al. Activating
mutations in the NT5C2 nucleotidase gene drive chemotherapy resistance in relapsed ALL. Nat Med,
2013, 19(3):368-71”.
Introduction
Acute lymphoblastic leukemia (ALL) is an aggressive hematological tumor resulting from the malignant transformation of lymphoid progenitors. Despite intensive chemotherapy, 20% of pediatric and over 50% of adult ALL patients fail to achieve a complete remission or relapse after intensified chemotherapy, making disease relapse and resistance to therapy the most significant challenge in the treatment of this disease (Moricke, Zimmermann et al. 2010, Salzer, Devidas et al. 2010).
Therapy of ALL includes initial treatment with high dose combination chemotherapy, which obtains clinical and hematologic remission in over 90% of cases. This is typically followed by additional rounds of highly intensive therapy aimed to further reduce disease burden; and then by a 2 year long lower intensity maintenance therapy in which treatment with oral 6-mercaptopurine plays a particularly important role (Koren, Ferrazini et al. 1990, Relling, Hancock et al. 1999). Patients with relapsed ALL generally receive a more intense treatment. However, despite these efforts, their outcome remains unsatisfactory with cure rates of less than 40% (Pui and Evans 2006). This is particularly the case in patients with relapsed T-ALL and in cases with primary resistance or early relapse, which is associated with higher risk of failure to achieve a second complete remission, shorter duration of chemotherapy response and poor survival (Gaynon, Qu et al. 1998, Tallen, Ratei et al. 2010).
Much effort has been spent on the study of the molecular basis of relapse and chemotherapy resistance in ALL. However the specific mechanisms mediating escape from therapy, disease progression and leukemia relapse remain largely unknown. In this chapter I describe our use of whole exome and targeted sequencing analysis to understand the emergence of relapsed ALL, with which we identify recurrent relapse-specific gain-of-function mutations in the cytosolic 5'-nucleotidase II (NT5C2)
57
Chapter 3 gene in 25% of relapsed T-ALLs and 6% of relapsed B-precursor ALLs. I show that these mutations increase NT5C2 nucleotidase activity and confer resistance to 6-mercaptopurine and 6-thioguanine chemotherapy when expressed in ALL lymphoblasts. These results support a prominent role for activating mutations in NT5C2 and increased nucleoside analog metabolism in disease progression and chemotherapy resistance in ALL.
Results
T-ALLs undergo clonal evolution from diagnosis to relapse
To identify the genetic mechanisms which may drive relapse of ALL we performed whole exome sequencing of matched diagnosis, remission and relapse DNA samples from 5 pediatric T-ALL patients
(Supplementary Table A3.1). This analysis (by Dr. Hossein Khiabanian) identified a mean mutation load of 13 somatic mutations per sample (range 5 – 17) (Supplementary Table A3.2). Out of 60 somatic mutations identified in total, 17 mutations were present at diagnosis and at relapse, 24 genes were selectively mutated in relapsed T-ALL samples and 19 mutations were present only at diagnosis.
Moreover, 4 of the 5 relapsed cases analyzed showed the presence of at least one somatic mutation present also at diagnosis, together with secondary mutations specifically acquired at the time of relapse.
In addition, 4 out of the 5 cases showed absence of at least one mutation marker present at diagnosis during disease progression leading to relapse. Single nucleotide polymorphism analysis of exome sequencing results ruled out that loss of these markers was due to loss of heterozygosity at relapse
(Supplementary Table A3.3). This result is consistent with previous studies based on copy number alteration analyses (Mullighan, Phillips et al. 2008, Tosello, Mansour et al. 2009, Clappier, Gerby et al.
2011) and supports that relapsed ALLs can originate as derivatives of ancestral subclones related to, but distinct from the main leukemic population present at diagnosis.
Somatically mutated genes at diagnosis included known T-ALL tumor suppressor genes such as
FBXW7 (Thompson, Buonamici et al. 2007), WT1 (Tosello, Mansour et al. 2009) and DNM2 (Zhang, Ding et al. 2012) in addition to numerous new genes not implicated before in the pathogenesis of this disease.
Analysis of mutant alleles found at the time of relapse identified mutations in three genes encoding proteins involved in positive regulation of TP53 signaling, including TP53 itself (TP53 R213Q), BANP
58 Chapter 3
(BANP H391Y) (Jalota, Singh et al. 2005) and RPL11 (RPL11 R18P) (Zhang, Wolf et al. 2003)
(Supplementary Figure A3.1). Notably, mutations in TP53 have been reported in about 10% of relapsed
ALL cases and are associated with a particularly poor prognosis (Hof, Krentz et al. 2011). Given the prominent role of TP53 pathway in DNA damage induced apoptosis (Bunz, Hwang et al. 1999), we performed extended mutation analysis of the TP53, BANP and RPL11 genes in 18 additional diagnostic and relapsed T-ALL samples (Supplementary Table A3.1). This analysis failed to identify additional
TP53 or BANP mutations, but showed the presence of two additional somatic RPL11 mutant alleles; one present both at diagnosis and relapse (RPL11 X178Q); and the other one (RPL11 G30fs) specifically mutated at relapse (Supplementary Figure A3.1). Relapse-associated mutations also included a prototypical activating mutation in the NRAS oncogene (NRAS G13V). Notably, NRAS mutations in ALL have been associated with poor outcome (Lubbert, Mirro et al. 1990) and are particularly prevalent in early T-cell precursor ALLs (Van Vlierberghe, Ambesi-Impiombato et al. 2011, Zhang, Ding et al. 2012), a group of high risk leukemias with poor prognosis (Coustan-Smith, Mullighan et al. 2009). Extended mutation analysis of NRAS in relapsed T-ALL cases demonstrated the presence of 2 diagnostic and relapse sample pairs harboring a prototypical NRAS G12S activating allele and a third patient with a heterozygous activating NRAS G12R mutation, which was present at diagnosis and showed loss of heterozygosity at the time of relapse.
NT5C2 mutations are recurrently found in relapsed ALL and associate with early relapse and relapse while on treatment
The most remarkable finding in our exome sequence analysis was the presence of a relapse- associated heterozygous mutation in the NT5C2 gene (NT5C2 K359Q). NT5C2 is a ubiquitous enzyme responsible for the final dephosphorylation of 6-hydroxypurine nucleotide monophosphates such as IMP, dIMP, GMP, dGMP and XMP before they can be exported out of the cell (Spychala, Madrid-Marina et al.
1988, Oka, Matsumoto et al. 1994). In addition, and most notably, NT5C2 can also dephosphorylate and inactivate 6-thioinositol monophosphate and 6-thioguanosine monophosphate which mediate the cytotoxic effects of 6-mercaptopurine (6-MP) and 6-thioguanine (6-TG) (Brouwer, Vogels-Mentink et al.
2005), two nucleoside analogs commonly used in the treatment of ALL.
59 Chapter 3
Mutation analysis of an extended panel of 119 relapsed T-ALL and 515 relapsed B-precursor ALL samples (Supplementary Table A3.1) identified 34 additional NT5C2 mutations in 30 T-ALL patients and
32 additional NT5C2 mutations in 30 B-precursor ALL patients in first relapse (Figure 3.1 and
Figure 3.1. NT5C2 mutations in relapsed ALL. (a) Schematic representation of the structure of the NT5C2 protein. The haloacid dehalogenase (HAD) domain is indicated. NT5C2 mutations identified in relapsed ALL samples are shown. Filled circles represent heterozygous mutations. Multiple circles in the same amino acid position account for multiple patients with the same variant. Alleles in light blue font are believed to be passenger mutations based on functional analysis. (b) DNA sequencing chromatograms of paired diagnosis and relapse genomic ALL DNA samples showing representative examples of relapse specific heterozygous NT5C2 mutations, with the mutant allele sequence highlighted in red.
60 Chapter 3
Supplementary Tables A3.4 and A3.5), bringing the overall frequency of NT5C2 mutations to 25%
(31/124) of relapsed T-ALLs and 6% (30/515) of relapsed B-precursor ALLs. Strikingly, 33 of these samples harbored the same NT5C2 R367Q mutation, 9 cases showed a recurrent NT5C2 R238W mutation and 3 cases harbored NT5C2 R238L. In addition we observed 5 cases harboring mutations in
NT5C2 D407 to different amino acids, 3 cases with NT5C2 R39Q, and 2 samples with a L375F single amino acid substitution (Figure 3.1 and Supplementary Tables A3.4 and A3.5). A number of these
NT5C2 mutations have been reported in other studies of relapsed B-precursor ALLs (Figure 3.1) (Meyer,
Wang et al. 2013, Ma, Edmonson et al. 2015). In each of the 19 cases for which original diagnostic DNA was available for analysis, NT5C2 mutations showed to be specifically acquired at the time of relapse. No
NT5C2 mutations were identified in 23 T-ALL and 27 B-precursor ALL additional diagnostic samples, further supporting the specific association of NT5C2 mutations with relapsed disease. Analysis performed by Dr. Renate Kirschner-Schwabe and Dr. Jana Hof of clinical and molecular features associated with
NT5C2 mutant relapsed T-ALLs treated in Berlin Frankfurt Münster (BFM) group based clinical trials
(ALL-BFM 95, ALL-BFM 2000, COALL 06-97, NHL-BFM 95 and Euro-LB 02) (Supplementary Table
A3.1) showed an association of NT5C2 mutations with early disease recurrence (very early or early relapse vs. late relapse, P <0.05) and relapse under treatment (P = 0.002) independently of treatment protocol (Supplementary Tables A3.6 – A3.11).
Relapse-associated NT5C2 mutations are gain-of-function alleles which increase NT5C2 nucleotidase activity
Given the described role of NT5C2 in the metabolism and inactivation of nucleoside analog drugs
(Galmarini, Jordheim et al. 2003, Brouwer, Vogels-Mentink et al. 2005, Hunsucker, Mitchell et al. 2005); the recurrent finding of the NT5C2 R367Q, NT5C2 R238W and other NT5C2 mutated alleles; and the reported association of increased levels of nucleotidase activity with thiopurine resistance and worse clinical outcome (Pieters, Huismans et al. 1992), we hypothesized that relapse-associated NT5C2 mutations may represent gain of function alleles with increased enzymatic activity. Detailed structure- function analysis of the NT5C2 K359Q mutation further supported this hypothesis. Thus, comparison of the wild type NT5C2 structure and models of the mutant NT5C2 K359Q protein by Dr. Zachary Carpenter
61 Chapter 3 showed that this mutation could result in increased NT5C2 activity by mimicking the effect of positive allosteric regulators (Figure 3.2). Allosteric activation of NT5C2 is mediated by binding of ATP, dATP,
Ap4A and 2,3-BPG to an allosteric pocket proximal to the NT5C2 active site (Figure 3.2a). Occupancy of this regulatory site results in increased ordering of an alpha helix formed by residues G355-E364 (Helix
A), which in turn displaces F354 from the catalytic center and moves D356 into the active site of the protein (Figure 3.2b, c). Similarly, our model predicts that the NT5C2 K359Q mutation could increase the
Helix A stability and reduce its solvent accessibility, resulting in an active configuration with displacement of F354 out of the NT5C2 active site and positioning D356 into the catalytic center of the enzyme (Figure
3.2d-e). Similar analysis of the S360P mutation observed by Ma et al. (Figure 3.1a) indicated that, while this mutation introduces a helix-breaking proline residue which disrupts the distal end of Helix A, it
Figure 3.2. Structure-function analysis of the NT5C2 K359Q mutant protein. (a) Molecular surface representation of NT5C2 protein structure. The position of the NT5C2 K359Q mutation found is highlighted in red. The substrate inosine monophosphate (IMP) is depicted in purple; the ATP allosteric activator is shown in yellow. (b) Structure representation of the NT5C2 catalytic center and allosteric regulatory site devoid of substrate or ligands (PDB 2XCX). (c) Structure representation of the NT5C2 catalytic center and allosteric regulatory site bound to IMP and ATP, respectively (PDB 2XCW). (d) Structure representation of the NT5C2 K359Q mutant model corresponding to the catalytic center and allosteric regulatory sites. (e) Overlay of the structures shown in b–d. The white arrow indicates the repositioning of Phe354 from the inactive NT5C2 configuration to the active – ATP-bound NT5C2 and NT5C2 K359Q– structures. Mg2+ ions are depicted as green spheres. Courtesy of Z. Carpenter.
62 Chapter 3 promotes organization of the proximal end of Helix A and may activate NT5C2 in a similar fashion to the
K359Q allele (Figure 3.3).
Figure 3.3. Structure-function analysis of the NT5C2 S360P mutant protein. Structural representation of the NT5C2 S360P mutant model corresponding to the catalytic center and allosteric regulatory sites. Courtesy of Z. Carpenter.
Consistent with this prediction, 5'-nucleotidase assays using NT5C2 K359Q and NT5C2 S360P recombinant proteins demonstrated a 48-fold and 16-fold increase in enzymatic activity compared wild type NT5C2, respectively (Figure 3.4). In addition, and despite the lack of clear structural cues on the
Figure 3.4. Increased 5'-IMP nucleotidase activity in NT5C2 mutant proteins. 5'-Nucleotidase activity levels of recombinant mutant proteins relative to wild type NT5C2 control are shown. Data are means ± s.d.
63 Chapter 3 role of other mutations in NT5C2 activation, nucleotidase activity analysis showed 10-25 fold increases in enzymatic activity for six more recombinant NT5C2 mutant proteins (R39Q, R238W, R367Q, D407A,
S408R, S445F, R478S) as well as a second highly-hyperactive mutant like K359Q with more than 55-fold increase in enzymatic activity, NT5C2 L375F (Figure 3.4). An additional structurally interesting allele is the NT5C2 Q523* nonsense mutation, which removes a proposed inhibitory region located in the C- terminal segment of the NT5C2 protein (Allegrini, Scaloni et al. 2004). However, it appears to be a very weakly activating allele in functional assays, which is supported by its lack of recurrence in NT5C2 mutational analyses of relapsed ALL samples. Lastly, in our sequencing analyses we also observed three
NT5C2 alleles encoding NT5C2 R291W, NT5C2 D384N and NT5C2 V454M, (Figure 3.1a) which we believe to be passenger mutations, since they had no effect or a very slightly deleterious effect on NT5C2 activity.
Relapse-associated NT5C2 mutations confer resistance to 6-mercaptopurine and 6-thioguanine in leukemic cells
To formally test the role of NT5C2 mutations in chemotherapy resistance we analyzed the effects of wild type and relapse-associated mutant NT5C2 expression in the response of CCRF-CEM T-ALL and
CUTLL1 T-ALL cells to 6-mercaptopurine (6-MP) and 6-thioguanine (6-TG) (Figures 3.5 and 3.6). Cell viability analysis in the presence of increasing drug concentrations demonstrated increased resistance to
6-MP and 6-TG therapy in cells expressing hyperactive NT5C2 mutants compared with empty vector and wild type NT5C2 controls (Figures 3.5 and 3.6, Supplementary Figure A3.2 and Supplementary Table
A3.12).
Finally, we tested the effects of relapse-associated NT5C2 mutations in the response to nelarabine – an araG precursor highly active in relapsed T-ALL– and araG (Berg, Blaney et al. 2005, DeAngelo, Yu et al.
2007, Sanford and Lyseng-Williamson 2008, Gokbuget, Basara et al. 2011). Strikingly, both nelarabine and AraG showed to be equally active in cells expressing relapse-associated NT5C2 mutations compared to controls (Figure 3.7).
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Figure 3.5. Expression of NT5C2 mutations in CCRF-CEM ALL cells induces resistance to chemotherapy with 6-MP and 6-TG. (a) Viability assays in CCRF-CEM T-ALL cells expressing wild type NT5C2, relapse- associated mutant NT5C2 alleles or a red fluorescent protein control (RFP), treated with increasing concentrations of 6-mercaptopurine (6-MP). (b) 6-Thioguanine (6-TG) dose response cell viability curves. Data is shown as means ± s.d.
Relapsed ALL tumors exhibit convergent evolution towards NT5C2 activation
In our sequencing analyses we discovered a number of cases harboring more than one NT5C2 mutation (Table 3.1). In particular we noted two samples which both showed NT5C2 R238W and NT5C2
L375F mutations. SNP analysis of seven independent loci confirmed that these samples are from unique individuals. This prompted us to investigate whether these two mutated alleles occur in cis (on the same chromosome) or in trans (on different chromosomes of the same cell, or in different cells of the tumor).
We obtained RNA material from both patients and generated cDNA from which we amplified and subcloned the stretch including the coding regions for amino acids 238 and 375. Sequencing analysis of individual cDNA clones from both patient samples showed the presence of either R238W or L375F alone, and the absence of any double-mutant clones (Table 3.2), indicating that the R238W and L375F mutations occur in trans. This was supported by enzymatic and cell-based analyses which showed that
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Figure 3.6. Expression of NT5C2 mutations in CUTLL1 ALL cells induces resistance to chemotherapy with 6-MP and 6-TG. Viability assays in CUTLL1 T-ALL cells were performed with (a) 6-MP and (b) 6-TG as described in Figure 3.4.
a b
Figure 3.7. Expression of NT5C2 mutations in CCRF-CEM ALL cells does not induce resistance to chemotherapy with nelarabine or araG. Representative viability assays in CCRF-CEM T-ALL cells expressing wild type NT5C2 or relapse-associated mutant NT5C2 alleles compared to a red fluorescent protein control (RFP), treated with increasing concentrations of (a) nelarabine or (b) araG. Data is shown as means ± s.d.
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Table 3.1. Concurrent NT5C2 mutations identified in relapsed ALL.
Sample NT5C2 mutation 1 NT5C2 mutation 2 T-ALL 17 R238L R367Q T-ALL B9 R238W L375F T-ALL B64 R238W L375F T-ALL C20 R478S R367Q B-ALL B3499 R238W R367Q B-ALL B3477 D407H D415G
Table 3.2. Clonal cDNA analysis of relapsed ALL samples harboring concurrent NT5C2 R238W and L375F mutations.
Patient 1 (no. of Patient 2 (no. of NT5C2 Mutation cDNA clones) cDNA clones) R238W only 1/18 1/22 L375F only 5/18 2/22 R238W + L375F 0/18 0/22
the double-mutant enzyme does not show hyperactivity and that it does not confer resistance to 6-MP or
6-TG in ALL cell lines (Supplementary Figures A3.3 and A3.4). In addition, a 1:1 mix of recombinant
NT5C2 R238W and NT5C2 L375F did not yield any synergistic increase in enzymatic activity, suggesting that there is no biological benefit for these two mutated alleles to appear in trans within the same tumor cell (Supplementary Figure A3.3).
To assess if the concurrent NT5C2 R238W and L375F mutations were found on different chromosomes of the same cell or if they were in different cells of the relapsed tumor, we performed single-cell DNA sequencing analyses of the corresponding exons 9 and 13 of NT5C2 from sorted viable relapsed tumor cells (material available for only one patient). These results showed that the relapsed tumor population was composed of a mix of clones which had either wildtype NT5C2 R238 and L375
(49%), only NT5C2 R238W (37%), or only NT5C2 L375F (14%) (Table 3.3).
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Table 3.3. Single-cell sequencing analysis of a relapsed ALL tumor sample harboring concurrent NT5C2 R238W and L375F mutations.
NT5C2 R238 NT5C2 L375 No. of cells (%) WT WT 32/65 (49%) R238W WT 24/65 (37%) WT L375F 9/65 (14%)
This analysis indicates that the NT5C2 R238W and L375F mutations in at least one patient had arisen in individual tumor clones which had independently gone through positive selection to emerge in the relapsed tumor population.
Discussion
Drug resistance and relapse of ALL are significant clinical challenges in the treatment of this aggressive malignancy, however the genetic mechanisms underlying chemotherapy resistance, disease progression and leukemia relapse have remained largely unknown. Prolonged maintenance treatment with 6-mercaptopurine is essential to obtain durable remissions in the treatment of ALL (Koren, Ferrazini et al. 1990, Relling, Hancock et al. 1999). Indeed, low-adherence to 6-mercaptopurine treatment, defined as less than 95% compliance, results in increased relapse rates and may account for as much as 59% of all ALL relapses (Bhatia, Landier et al. 2012).
In this context, our results highlight the prominent role of relapse-specific activating mutations in
NT5C2 as a direct mechanism of resistance to 6-mercaptopurine and a recurrent genetic driver of relapse in ALL. Moreover, the presence of multiple tumor clones with NT5C2 gain-of-function mutations at relapse emphasizes the selective pressure from 6-mercaptopurine chemotherapy during maintenance treatment as a major force of convergent evolution of tumor populations from diagnosis to relapse. This convergent mechanism is echoed by the presence of NT5C2 activating mutations in both B-precursor and T-cell ALL relapses, which represent two unique molecular subtypes of ALL.
The observed difference in NT5C2 mutational recurrence between relapsed T-ALL (25%) and relapsed B-ALL (6%) is yet to be understood, although it may be explained by inherent differences in
68 Chapter 3 purine metabolic pathway activity between malignant B-cells and malignant T-cells. Indeed, NT5C2 enzymatic activity is approximately 5-fold lower in T-cells than in B-cells (Rosi, Agostinho et al. 1998,
Hunsucker, Mitchell et al. 2005), suggesting that mutational hyperactivation of NT5C2 may provide a greater evolutionary benefit of chemotherapy resistance to malignant T-cells than malignant B-precursor cells.
Given the importance of achieving effective systemic exposure to 6-mercaptopurine in minimizing the risk of ALL relapse (Koren, Ferrazini et al. 1990, Relling, Hancock et al. 1999), it would be worthwhile to investigate if inadequate dosing or poor patient adherence to 6-mercaptopurine treatment may promote the emergence of NT5C2-mutant tumor clones at relapse, and similarly, whether dose intensification of 6- mercaptopurine during maintenance therapy may be beneficial in reducing the rate of NT5C2-mutant relapses.
Notably, the lack of nelarabine cross-resistance in cells expressing activating NT5C2 alleles analyzed here suggests that these mutations may not impair the effectiveness of nelarabine-based salvage therapies currently in use for relapsed T-ALL (Berg, Blaney et al. 2005, DeAngelo, Yu et al. 2007,
Gokbuget, Basara et al. 2011).
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Chemotherapy resistance-driving NT5C2 mutations impair leukemia initiating cell
activity in relapsed acute lymphoblastic leukemia
Introduction
Relapse and resistance to chemotherapy remain the most difficult challenges in the treatment of pediatric acute lymphoblastic leukemia (ALL) (Bhojwani and Pui 2013). While improved and intensified chemotherapy regimens have pushed overall survival to over 80% for newly diagnosed pediatric ALL
(Schultz, Pullen et al. 2007, Mitchell, Richards et al. 2010, Vrooman, Stevenson et al. 2013, Hunger and
Mullighan 2015), patients with relapsed or refractory ALL still have disappointingly poor outcomes, with a cure rate of only 40% (Bhojwani and Pui 2013, Hunger and Mullighan 2015).
Several drivers of leukemia relapse and drug resistance have been proposed, such as the presence of rare leukemia stem cells capable of self-renewal (Lapidot, Sirard et al. 1994, Konopleva,
Zhao et al. 2002, Guan, Gerhard et al. 2003, Chiu, Jiang et al. 2010, Eppert, Takenaka et al. 2011,
Meyer, Eckhoff et al. 2011, Shlush, Zandi et al. 2014), protective microenvironment safe-haven niches
(Konopleva, Konoplev et al. 2002, Iwamoto, Mihara et al. 2007, Yang, Mallampati et al. 2013) and clonal evolution (Mullighan, Phillips et al. 2008, Ma, Edmonson et al. 2015, Swaminathan, Klemm et al. 2015) with acquisition of secondary genetic alterations which directly promote chemotherapy resistance (Goker,
Waltham et al. 1995, Mullighan, Zhang et al. 2011, Meyer, Wang et al. 2013, Tzoneva, Perez-Garcia et al. 2013, Li, Li et al. 2015). In this context, heterozygous activating mutations in the NT5C2 nucleotidase gene are present in 19-38% of relapsed pediatric T-cell ALL cases (Tzoneva, Perez-Garcia et al. 2013,
Kunz, Rausch et al. 2015) and 3-10% of relapsed B-precursor ALLs (Meyer, Wang et al. 2013, Tzoneva,
Perez-Garcia et al. 2013, Ma, Edmonson et al. 2015).
NT5C2 (cytosolic 5’-nucleotidase II) is a highly conserved, ubiquitously expressed enzyme that maintains a balanced nucleotide pool in the cell by catalyzing the 5’-dephosphorylation of purines such as inosine monophosphate, xanthosine monophosphate and guanosine monophosphate before they are exported out of the cell (Spychala, Madrid-Marina et al. 1988, Oka, Matsumoto et al. 1994). In addition,
NT5C2 is capable of metabolizing the cytotoxic metabolites of purine analogs like 6-mercaptopurine (6-
MP) and 6-thioguanine (6-TG) (Brouwer, Vogels-Mentink et al. 2005) (Figure 4.1), and expression of
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Figure 4.1. NT5C2 inactivates the cytotoxic metabolites of 6-mercaptopurine (6-MP) and 6-thioguanine (6-TG).
relapse-associated hyperactive NT5C2 alleles in leukemic cell lines confers resistance to 6-MP and 6-TG in vitro (Meyer, Wang et al. 2013, Tzoneva, Perez-Garcia et al. 2013).
6-MP and 6-TG are thiopurine prodrugs which are metabolically activated inside leukemic cells via the purine salvage pathway (Lennard 1992). The hypoxanthine-guanine phosphoribosyl transferase enzyme (HPRT) processes 6-MP to thio-IMP, which is then converted to thio-XMP and thio-GMP, while 6-
TG is processed by HPRT directly into thio-GMP (Figure 4.1). Subsequent metabolism of thio-GMP by kinases and reductases yields thio-dGTP which is incorporated into replicating DNA strands and triggers the DNA mismatch-repair machinery, leading to cell cycle arrest and apoptosis (Figure 4.1) (Swann,
Waters et al. 1996). The anti-leukemic effects of 6-MP are also attributed to a second metabolic pathway whereby thiopurine S-methyl transferase (TPMT) methylates thio-IMP to form methylthio-IMP (MeTIMP), which is a potent inhibitor of amidophosphoribosyltransferase (ATase), an enzyme catalyzing the committed step of de novo purine biosynthesis (Figure 4.1). NT5C2 antagonizes the actions of both 6-MP and 6-TG due to its ability to dephosphorylate thio-IMP, thio-XMP and thio-GMP, which serves as the last step before export of these key drug metabolites out of the cell (Figure 4.1) (Lennard 1992). Notably, 6-
71 Chapter 4
MP is a mainstay chemotherapy drug in the treatment of ALL, particularly during a 2-year long maintenance therapy phase which follows onset of remission (Inaba, Greaves et al. 2013) and has been shown to reduce rates of relapse (Koren, Ferrazini et al. 1990, Relling, Hancock et al. 1999, Inaba,
Greaves et al. 2013, Siegel, Naishadham et al. 2013).
Here we show that a highly recurrent, relapse-associated activating mutation in the NT5C2 nucleotidase gene expressed from the endogenous mouse locus induces marked resistance and progression under therapy with 6-MP in primary mouse leukemia in vivo, while surprisingly conferring a growth and metabolic disadvantage and reduced leukemia initiating cell activity in the absence of chemotherapy. Given the importance of long-term daily 6-MP treatment during maintenance therapy for
ALL, our results are consistent with a model whereby NT5C2 mutations are negatively selected in the early stages of disease development and only positively selected during maintenance therapy where chemotherapy pressure plays a crucial role in the clonal evolution of the tumor. This is supported by a lack of detectable NT5C2 mutations at diagnosis and serves as the first known example of a tumor clone with reduced fitness driving drug resistance and relapse of cancer.
Results
Establishment of a conditional inducible Nt5c2 R367Q model of T-ALL
To assess the role of mutant NT5C2 in the evolution of leukemic resistance in vivo we generated a conditional knock-in mouse model of the highly recurrent NT5C2 R367Q allele. We used homologous recombination in mouse embryonic stem cells with a targeting vector that includes Nt5c2 exon 13 followed by a floxed minigene cassette consisting of the remaining downstream Nt5c2 wildtype cDNA
(exons 14-18) (Figure 4.2). The floxed minigene is followed by the remainder of the Nt5c2 gene from exon 14 downstream, including an engineered R367Q mutation in exon 14. In this model, the floxed knock-in allele would express wildtype Nt5c2 via splicing to the minigene, whereas expression of Cre recombinase would lead to excision of the minigene and splicing from the endogenous exon 13 through to the mutated exon 14, resulting in expression of Nt5c2 R367Q from the endogenous mouse locus. We obtained four engineered knock-in embryonic stem cell lines from InGenious Technologies and performed blastocysts injections and subsequent mouse line development at Columbia University (Figure 4.2).
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Figure 4.2. Conditional knock-in targeting of Nt5c2 in mouse embryonic stem cells. (a) Schematic representation of the targeting strategy for generation of Nt5c2 R367Q conditional knock-in mice (b) Southern blot analysis of Nt5c2 targeted embyonic stem cells in DNA digestions with BamHI restriction enzyme hybridized with a DNA probe external to the long arm. (c) Southern blot analysis of Nt5c2 targeted embryonic stem cells in DNA digestions with ApaI restriction enzyme hybridized with a DNA probe in the short arm.
To study the role of Nt5c2 R367Q expression in the chemotherapy resistance and maintenance of established T-ALL, we generated primary murine T-ALL tumors by retroviral expression of an oncogenic constitutively active form of NOTCH1 (ΔE-NOTCH1) (Schroeter, Kisslinger et al. 1998) in hematopoietic progenitors from a tamoxifen-inducible conditional heterozygous knock-in donor line (Rosa26-CreERT2
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Figure 4.3. Experimental strategy for developing conditional inducible Nt5c2 R367Q primary mouse T-ALL tumors and assessment of the role of Nt5c2 R367Q in established T-ALL in the presence and absence of chemotherapy.
Nt5c2 R367Q wt/flox) and subsequent transplantation into sublethally irradiated isogenic C57BL/6 recipients
(Figure 4.3). Consistent with previous studies (Pear, Aster et al. 1996, Herranz, Ambesi-Impiombato et al.
2014), mice transplanted with ΔE-NOTCH1 infected progenitors displayed a transient increase in the frequency of CD4+CD8+ double-positive cells in the peripheral blood 3 weeks post transplantation, which resulted in overt CD4+CD8+ T-ALL development with 100% penetrance at 5-6 weeks post-transplant
(Figure 4.4a).
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Figure 4.4. Expression of a single Nt5c2 R367Q allele from the endogenous mouse locus leads to overt 6- MP and 6-TG resistance in vitro. (a) Representative FACS plots of Rosa26-CreERT2 Nt5c2 R367Q wt/flox primary T-ALL tumors showing CD4, CD8 double positivity indicative of T-ALLs driven by hyperactive ΔE- NOTCH1. (b) Genotyping PCR from genomic DNA of Rosa26-CreERT2 Nt5c2 R367Q wt/flox primary T-ALL tumors treated with 4-hydroxytamoxifen or vehicle in vitro showing Cre-mediated deletion of Nt5c2 wild type mini-gene. (c) Sequencing analysis of Nt5c2 mRNA from 4-hydroxytamoxifen or vehicle-treated tumor cells, showing Cre- mediated expression of the engineered Nt5c2 R367Q mutant allele. (d,e) Viability assays showing drug responses of isogenic Nt5c2wt/wt and Nt5c2wt/R367Q primary mouse T-ALLs to increasing doses of (d) 6-MP and (e) 6-TG in vitro. Data are shown as the means ± s.d.
Conditional Nt5c2 R367Q primary murine T-ALLs show marked resistance to thiopurine chemotherapy in vitro
Treatment of primary Rosa26-CreERT2 Nt5c2 R367Q wt/flox T-ALL tumors with 4- hydroxytamoxifen in vitro resulted in efficient deletion of the minigene cassette (Figure 4.4b) and expression of Nt5c2 R367Q mRNA at a 1:1 ratio with wildtype Nt5c2 mRNA within 48 hours (Figure
75 Chapter 4
4.4c), thereby serving as a highly accurate model of the heterozygous mutations observed in relapsed human T-ALLs. We assessed the sensitivity of 4-hydroxytamoxifen-treated (Nt5c2 wt/R367Q) or vehicle- treated (Nt5c2 wt/flox) isogenic tumors to increasing concentrations of 6-MP and 6-TG in vitro and found that expression of a single copy of Nt5c2 R367Q in these established leukemias led to overt chemoresistance to both purine analogs (Figure 4.4d, e; Supplementary Table A4.1).
Mice bearing Nt5c2 R367Q T-ALLs show resistance and disease progression under 6-MP chemotherapy in vivo
To assess the role of 6-MP dosage on the clonal expansion of Nt5c2 R367Q leukemias in vivo, we transplanted secondary recipients with Rosa26-CreERT2 Nt5c2 R367Q wt/flox T-ALL lymphoblasts expressing lentiviral firefly luciferase, allowing analysis of leukemia progression via live bioluminescence imaging. Once recipients had developed detectable leukemic tumor burden, we treated them with tamoxifen or vehicle to generate isogenic Nt5c2 wildtype or R367Q tumors, and after recovery for 48 hours we treated both groups intraperitoneally with a range of doses of 6-MP daily for 5 days. We found that Nt5c2 R367Q induces resistance and disease progression under therapy in vivo under a broad range of 6-MP doses (50, 60, 80 mg/kg/day), with partial therapeutic response observed only at the highest dose of 100 mg/kg/day (Figure 4.5a-c). In addition, after 5 days of 6-MP treatment, mice bearing wildtype leukemias displayed clear dose-response reductions in spleen size, cellularity and percentage of GFP- positive tumor cells, while these effects were greatly diminished in mice bearing isogenic Nt5c2 R367Q tumors (Figure 4.5d-g). These results are consistent with the strong association between NT5C2 mutations and early relapse or relapse while on treatment shown in a number of studies (Meyer, Wang et al. 2013, Tzoneva, Perez-Garcia et al. 2013, Ma, Edmonson et al. 2015) and highlight the importance of
6-MP dose optimization and compliance to therapy which have been shown to affect risk of relapse
(Relling, Hancock et al. 1999, Bhatia, Landier et al. 2012).
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Figure 4.5. Nt5c2 R367Q induces T-ALL drug resistance and disease progression under 6-MP therapy in vivo. (a) Average changes in tumor load by bioimaging in mice allografted with NOTCH1-induced isogenic Nt5c2wt/wt and Nt5c2wt/R367Q primary mouse T-ALLs treated with a range of 6-MP doses (n = 5 per group). (b) Representative images and (c) quantitation of tumor burden assessed by bioimaging. (d-g) Assessment of tumor burden at the end of 5 days of 6-MP treatment by (d) spleen size, (e) spleen weight, (f) spleen cell density and (g) percentage of GFP+ve tumor cells in the spleen. P values were calculated using two-tailed Student’s t-test. Data are shown as the means ± s.d. Scale bar, 1 cm.
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Nt5c2 R367Q primary murine leukemias have decreased fitness and leukemia initiating activity
Previous studies have linked resistance-driving mutations to more aggressive leukemia growth and proliferation (Clappier, Gerby et al. 2011), clonal expansion at early stages of tumor development and increased leukemia stem cell activity (Shlush, Zandi et al. 2014, Chen, Chen et al. 2015, Wong,
Ramsingh et al. 2015). We assessed the effect of Nt5c2 R367Q on leukemic cell growth and surprisingly discovered that in the absence of chemotherapy Nt5c2 R367Q tumors have decreased proliferation compared to isogenic wildtype counterparts in vitro (Figure 4.6a), and mice transplanted with Nt5c2
R367Q leukemic cells display slower disease progression compared to controls bearing isogenic wildtype tumors in vivo (Figure 4.6b). In addition and most notably, in limiting dilution transplantation assays Nt5c2
R367Q tumors displayed a 17-fold reduction in leukemia initiating cell activity compared to isogenic wildtype controls (Figure 4.6c; Supplementary Table A4.2). These paradoxical results indicate that
NT5C2 R367Q decreases fitness and self-renewal of the leukemic clone in the absence of 6-MP
Figure 4.6. Nt5c2 R367Q tumor cells have reduced proliferation and leukemia initiating cell activity in the absence of chemotherapy. (a) In vitro cell growth of isogenic Nt5c2wt/wt and Nt5c2wt/R367Q primary mouse T- ALLs. (b) Kaplan-Meier survival curve of mice harboring Nt5c2wt/wt and Nt5c2wt/R367Q isogenic leukemias. (c) Serial dilution leukemia initiating cell analysis plot in mice treated with vehicle only (Nt5c2wt/wt) or tamoxifen (Nt5c2wt/R367Q) after transplantation with serial dilutions of Rosa26-CreERT2 Nt5c2 R367Q wt/flox cells (left), and confidence intervals showing 1/(stem cell frequency) (right) (n=6 per group). Data in (a) are shown as the means ± s.d. and P values calculated using two-tailed Student’s t-test. **** P ≤0.001; n.s., not significant.
78 Chapter 4 chemotherapy. A corollary of this is that NT5C2 mutant tumor cells could be negatively selected by clonal competition at the early stages of tumor development and would only have an advantage during the selective pressure of maintenance therapy with 6-MP.
NT5C2 mutations are not present in primary leukemic tumors at diagnosis
We tested for the presence of NT5C2 mutations at disease presentation by ultra-deep sequencing of matching diagnostic material from 15 relapsed cases with NT5C2 mutations (sensitivity of
1:200). In all cases analyzed we did not uncover any of the corresponding relapse-associated NT5C2 mutations at diagnosis (Table 4.1). We then performed an allele specific PCR for the highly recurrent
NT5C2 R367Q with a sensitivity of 1:1000 (Figure 4.7) and again did not detect the mutation in any of the diagnostic samples analyzed (Table 4.2). These results show that NT5C2 mutant clones are not present in the original tumor or they represent very minor, undetectable populations.
Table 4.1. Targeted deep sequencing of NT5C2 in matching diagnostic and remission DNA samples from patients with NT5C2-mutated relapsed ALL
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Figure 4.7. NT5C2 R367Q allele specific PCR. (a) Calibration of NT5C2 WT and R367Q allele-specific PCR assays. NT5C2 WT and R367Q assays were run on increasing amounts of 100% WT or 100% R367Q plasmid template respectively. A calibration ΔCT value was determined as the difference between the average CT values for the WT and R367Q assays at a given amount of DNA template. This value was used to normalize the ΔCT detection cutoff between the WT and R367Q assays. (b) Determination of NT5C2 R367Q assay sensitivity. NT5C2 R367Q allele specific PCR assay was performed on WT NT5C2 genomic DNA from JURKAT cells spiked with increasing amounts of NT5C2 R367Q plasmid DNA. Data are shown as the means ± s.d.
Table 4.2. Allele specific PCR analysis of NT5C2 R367Q presence in matching diagnostic and remission DNA samples from patients with NT5C2-mutated relapsed ALL.
Nt5c2 R367Q primary leukemic cells have depleted pools of endogenous purine metabolites
Since the normal function of NT5C2 is to dephosphorylate key 6-hydroxypurine nucleoside monophosphates before they are exported out of the cell (Spychala, Madrid-Marina et al. 1988, Oka,
Matsumoto et al. 1994), we hypothesized that NT5C2 activating mutations would result in increased processing and depletion of the endogenous intracellular pool of purine metabolites, which could mediate the loss of fitness phenotype observed in Nt5c2 R367Q mutant leukemia cells.
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To test this hypothesis we performed broad-based LC-MS/MS metabolomic analyses on isogenic
Nt5c2 wildtype and Nt5c2 R367Q primary mouse ALL lymphoblasts as well as the corresponding conditioned media from each (Metabolon Inc.). We found that Nt5c2 R367Q cells had significantly
Figure 4.8. Metabolic profiling analysis shows a significant metabolic disadvantage in Nt5c2wt/R367Q cells, with decreased levels of Nt5c2 purine substrates and increased export of downstream purine metabolites. (a) Purine salvage pathway, highlighting NT5C2-catalyzed reactions. (b) GC/MS and LC/MS/MS metabolic profiles (mass spectrometry scaled intensity, arbitrary units) of Nt5c2wt/wt and Nt5c2wt/R367Q isogenic primary mouse T-ALL cells and conditioned media, showing effects on Nt5c2 substrates and downstream metabolites. (c) Metabolic changes in adenosine pathway as a secondary effect of reduced IMP levels in Nt5c2wt/R367Q tumor cells. Box plots represent the upper quartile to lower quartile distribution. + Sign indicates mean value and horizontal line the median value. Whiskers indicate the maximum and minimum values of the distribution. Open circles indicate extreme data points.
81 Chapter 4 decreased levels of endogenous Nt5c2 substrates (IMP, XMP and GMP), coupled with increased downstream products and metabolites in the conditioned media (inosine, hypoxanthine, xanthosine, xanthine, guanine and uric acid) as compared to wildtype controls (Figure 4.8a, b). Moreover, as a result of the decreased levels of IMP which is required for the synthesis of AMP, we observed a reduction in all downstream adenosine metabolites (adenylosuccinate, AMP, adenosine, adenine) in Nt5c2 R367Q cells
(Figure 4.8a, c). These results are consistent with a significant metabolic loss of fitness due to increased
NT5C2 5’-nucleotidase activity in leukemias harboring activating NT5C2 mutations – an evolutionary cost paid for attaining resistance to 6-mercaptopurine chemotherapy.
Discussion
Drug resistance and relapse of ALL remain the most significant challenges in the treatment of this aggressive hematological malignancy. Until recently not much was known about the genetic basis and evolutionary mechanisms of leukemia relapse. The rapid development of next-generation sequencing technology has allowed us to identify activating mutations in the NT5C2 nucleotidase gene as the most prominent relapse-specific genetic alterations in ALL, with highest recurrence in relapsed T-ALLs (Meyer,
Wang et al. 2013, Tzoneva, Perez-Garcia et al. 2013, Kunz, Rausch et al. , Ma, Edmonson et al. 2015).
Our murine model of T-ALL with conditional inducible expression of the highly recurrent Nt5c2
R367Q allele from the endogenous mouse locus has allowed us to accurately study the role of NT5C2 in drug resistance and disease progression of T-ALL in vivo. These studies have highlighted the overt resistance to 6-mercaptopurine chemotherapy induced by expression of a single allele of Nt5c2 R367Q in leukemic cells and the potential of 6-mercaptopurine dose intensification as an approach to minimize
NT5C2-mutant leukemic relapses. Indeed, reduced intensity and low-adherence to 6-MP maintenance therapy have been associated with a greatly increased risk of relapse (Relling, Hancock et al. 1999,
Bhatia, Landier et al. 2012).
Remarkably, we have discovered that the clonal advantage of NT5C2 activating mutations during disease progression is strictly dictated by selection pressure from 6-mercaptupurine chemotherapy and comes at a great biological cost to the tumor cells, with severe dysregulation of their purine nucleotide pools, decreased fitness and delayed growth in the absence of chemotherapy. Thus our studies
82 Chapter 4 demonstrate for the first time that less aggressive tumor clones can drive disease progression under chemotherapy in vivo despite having decreased fitness and impaired leukemia-initiating ability. These results are consistent with the lack of detectable NT5C2 mutations at disease presentation, supporting a model whereby NT5C2 mutations are negatively selected in the early stages of leukemia development and positively selected only under prolonged 6-mercaptopurine pressure. These findings are especially pertinent given the importance of long-term daily 6-mercaptopurine treatment during maintenance therapy for ALL (Koren, Ferrazini et al. 1990, Relling, Hancock et al. 1999). Thus, our results highlight the significance and dominance of chemotherapy as a selective pressure in the Darwinian evolution of
NT5C2 mutant tumors from diagnosis to relapse, and implicate dosage and compliance to chemotherapy as key factors in limiting disease progression and relapse of ALL.
Finally, the decreased metabolic fitness of NT5C2 mutant tumor cells offers potential therapeutic opportunities for targeting parallel metabolic pathways to which NT5C2 mutant cells may be sensitized. In line with this, the Nt5c2 R367Q T-ALL mouse model we have developed will serve as an excellent platform for drug screening approaches for cytotoxic or chemosensitizing agents with selectivity for
NT5C2 mutant leukemias.
83
CHAPTER 5
Unique structural mechanisms of NT5C2 mutational hyperactivation in relapsed
ALL and discovery of a novel NT5C2 auto-regulatory switch
Introduction
Cytosolic 5’-nucleotidase II (NT5C2) is a highly conserved and ubiquitously expressed enzyme which plays a central role in maintaining intracellular nucleotide pool homeostasis. It removes the 5’ phosphate moiety of 6-hydroxypurine monophophates, showing highest preference for inosine monophosphate (IMP), guanosine monophosphate (GMP) and xanthosine monophosphate (XMP) as well as the deoxyribose forms dIMP and dGMP (Itoh, Mitsui et al. 1967, Spychala, Madrid-Marina et al. 1988,
Tozzi, Camici et al. 1991, Pesi, Turriani et al. 1994, Banditelli, Baiocchi et al. 1996, Allegrini, Pesi et al.
1997). The removal of the 5’ phosphate group allows excess purine nucleosides to leave the cell and is a major component of the regulation of cellular nucleotide pool composition in response to changing metabolic demands.
In addition, NT5C2 can dephosphorylate 6-thioinosine monophosphate (6-TIMP), 6- thioxanthosine monophosphate (6-TXMP) and 6-thioguanosine monophosphate (6-TGMP) – key metabolic intermediates in the activation of two anti-leukemic nucleoside analog drugs, 6-thioguanine (6-
TG) and 6-mercaptopurine (6-MP) (Brouwer, Vogels-Mentink et al. 2005). In this respect, increased expression levels of NT5C2 have been correlated with nucleoside analog drug resistance phenotypes
(Carson, Carrera et al. 1991, Schirmer, Stegmann et al. 1998, Dumontet, Fabianowska-Majewska et al.
1999, Lotfi, Mansson et al. 2001) and poor patient outcomes after treatment with cytarabine, cladribine, 6-
MP and 6-TG (Pieters, Huismans et al. 1992, Kawasaki, Carrera et al. 1993, Galmarini, Graham et al.
2001, Galmarini, Thomas et al. 2002, Galmarini, Thomas et al. 2003, Galmarini, Cros et al. 2004, Mitra,
Crews et al. 2011). More recently, the role of NT5C2 in chemotherapy drug resistance was revealed to be of far greater significance, with the discovery by our group and others of recurrent relapse-associated activating mutations in NT5C2 in relapsed T-cell and B-precursor acute lymphoblastic leukemia samples
(Figure 3.1, Chapter 3) (Meyer, Wang et al. 2013, Tzoneva, Perez-Garcia et al. 2013, Kunz, Rausch et al. 2015, Ma, Edmonson et al. 2015).
84
Chapter 5
NT5C2 belongs to a family of Mg2+ dependent intracellular 5’-nucleotidases with varying substrate specificities and subcellular localizations (Hunsucker, Mitchell et al. 2005), which all share a common reaction mechanism that places them in the haloacid dehalogenase (HAD) superfamily (Koonin and
Tatusov 1994). Catalysis involves formation of a phosphoenzyme intermediate via nucleophilic attack by a highly conserved Asp (Asp52 in NT5C2) on the phosphate of the substrate, and donation of a proton by a second conserved Asp (Asp54 in NT5C2) to the departing nucleoside (Koonin and Tatusov 1994,
Collet, Stroobant et al. 1998, Allegrini, Scaloni et al. 2001, Rinaldo-Matthis, Rampazzo et al. 2002,
Wallden and Nordlund 2011).
The human NT5C2 gene codes for a peptide of 561 amino acids with a molecular weight of about
65 kDa (Oka, Matsumoto et al. 1994). Functional and structural studies have shown that NT5C2 exists as a homotetramer made up of a dimer of dimers, with homodimerization strictly required for enzymatic activity (Spychala, Madrid-Marina et al. 1988, Itoh 1993, Marques, Teixeira et al. 1998, Spychala, Chen et al. 1999, Wallden, Stenmark et al. 2007, Pesi, Allegrini et al. 2010, Wallden and Nordlund 2011).
NT5C2 is allosterically activated by ATP, dATP, adenosine diphosphate (ADP), 2,3- bisphosphoglycerate (2,3-BPG), and Ap4A (two adenosine molecules connected by four phosphates), with ATP/dATP and Ap4A being the strongest activators at physiological concentrations, resulting in decreased Km and increased Vmax (Pinto, Canales et al. 1986, Spychala, Madrid-Marina et al. 1988,
Marques, Teixeira et al. 1998). Moreover, ATP protects the enzyme from heat inactivation and trypsin digestion (Itoh, Usami et al. 1978, Itoh 1982) and promotes auto-oligomerization (Spychala, Chen et al.
1999), suggesting that the oligomerization state is an important factor for NT5C2 activation.
Crystallography studies of a substrate-trapping human NT5C2 mutant protein, where the catalytic
Asp52 was mutated to Asn (D52N), provided a structural basis for its allosteric activation (Wallden and
Nordlund 2011). Comparison of the inactive (apo) and active (effector bound and substrate trapped) structures of NT5C2 revealed a crucial difference in a stretch of residues, Gly355-Glu364, which connects the active site to the effector site. Notably, this stretch is disordered in the apo form, while it constitutes an ordered α-helix (helix A) in the activated enzyme. Thus, binding of an effector (e.g. ATP) in the allosteric pocket results in a conformational change involving the disorder-to-order transition of helix A, which is stabilized by electrostatic interactions between Lys362 and the phosphate moieties of the effector, and by
85 Chapter 5 a π-stacking interaction between Phe354 and the nucleobase of the effector. The ordering of helix A results in a rotation of Phe354 out of the active site, allowing Asp356 to move into the catalytic center where it helps to organize the active site and to coordinate the Mg2+ ion. This creates an electrostatically favorable environment which facilitates binding and catalysis of the substrate (Wallden and Nordlund
2011).
The high prevalence of NT5C2 mutations in relapsed T-ALL (25%) and relapsed B-ALL (6%) prompted us to understand the biochemical and structural mechanisms of NT5C2 hyperactivation in these samples. Our prior modeling analysis of NT5C2 K359Q indicated that this mutation increases helix A stability and thereby acts as a mimic of the allosterically activated state (Figure 3.2, Chapter 3). Here I describe protein biochemical studies of 11 different relapse-associated NT5C2 mutant alleles through which we identified three enzymatic classes of NT5C2 activating mutations, as well as crystallographic studies through which we uncovered the unique underlying structural mechanisms of each class. In addition, and most notably, we have discovered a novel auto-regulatory switch-off mechanism of NT5C2, which is targeted by activating mutations in different regions of the NT5C2 functional dimer unit. These results will facilitate the rational targeted drug design of NT5C2 inhibitors with high specificity, low systemic toxicity and increased selectivity for mutant NT5C2.
Results
Activating NT5C2 mutations in relapsed ALL comprise three unique enzymatic classes
To understand the enzymatic effects of different relapse-specific NT5C2 mutations we characterized the 5’-nucleotidase (5’-NT) activity of purified recombinant NT5C2 wildtype and mutant proteins in the absence and in the presence of increasing concentrations of allosteric activators ATP,
ADP and Ap4A (Figure 5.1). We found that wildtype and mutant NT5C2 proteins are most responsive to allosteric activation by Ap4A and ATP (Figure 5.1a-d) with relatively weaker responses to ADP (Figure
5.1e, f), as has been described for the NT5C2 wildtype protein (Hunsucker, Mitchell et al. 2005). Our results are also consistent with previous reports that Ap4A is capable of activating NT5C2 at lower
86 Chapter 5
Figure 5.1. NT5C2 mutant proteins have altered responses to allosteric activation. 5’-nucleotidase activity of wildtype and mutant recombinant NT5C2 proteins is shown in response to increasing concentrations of allosteric activators ATP (a,b), Ap4A (c,d) and ADP (e,f). Specific activity responses are shown on the left (a,c,e) and fold responses relative to each protein’s activity at basal conditions are on the right (b,d,f). Data are shown as means ± s.d.
87 Chapter 5 concentrations than ATP, suggesting that it may be a more relevant physiological regulator (Pinto,
Canales et al. 1986).
NT5C2 responses to ATP and Ap4A had similar reaction profiles, with the majority of NT5C2 mutant proteins, while already hyperactive in basal conditions, also showing reactivity to ATP and Ap4A at low concentration ranges and reaching early saturation at the physiological concentration of 3 mM ATP and 1-2 mM Ap4A as compared to wildtype NT5C2 (Figure 5.1a, c). This indicates that mutant NT5C2 proteins are able to achieve far greater enzymatic activity than wildtype NT5C2 when the intracellular
ATP and Ap4A concentrations are low. The similarities between ATP and Ap4A activity are consistent with the two activators binding in the same allosteric pocket. This is supported by the crystal structure of activated wildtype NT5C2, which shows two ATP molecules bound back-to-back at the interface of neighboring NT5C2 monomers, so that the phosphate groups of the two activator molecules are bridged by a Mg2+ ion (Wallden and Nordlund 2011). In this context, a single molecule of Ap4A would be equivalent to two ATP molecules, with the two adenosine moieties binding in each subunit of a NT5C2 dimer and bridged by the polyphosphate linker.
Compared to the wildtype protein, NT5C2 mutant proteins have decreased relative fold responses to allosteric activation by ATP and Ap4A, since they are already much closer to enzymatic activity saturation in basal conditions (Figure 5.1b, d). A clear exception is the NT5C2 Q523* truncated protein. Notably, in our sequencing analyses we have only observed this single case of a NT5C2 truncating mutation, suggesting that it is a very weakly activating allele. Consistent with this prediction, inducible expression of NT5C2 Q523* in human T-ALL cell lines leads to moderate 6-mercaptopurine resistance, but is less effective than relapse-associated NT5C2 single amino acid substitutions (by
Chelsea Dieck in our lab, unpublished). Enzymatically, NT5C2 Q523* does not show notable hyperactivation in basal conditions, yet it is more responsive to allosteric activation than the remaining
NT5C2 mutant proteins (Figure 5.1b, d). This property allows it to reach a higher 5’-NT enzymatic rate than the wildtype protein when allosteric activators are present (Figure 5.1a, c).
In contrast to ATP and Ap4A, NT5C2 mutant proteins display slow, gradual activation responses to increasing ADP concentrations (Figure 5.1e) and similar relative fold responses to ADP as the wildtype
88 Chapter 5 protein (Figure 5.1f). An exception is NT5C2 S445F which has a slightly sharper specific activity response to increasing ADP concentrations, with earlier saturation than the other NT5C2 mutant proteins.
Our analysis of the effects of NT5C2 mutations on the enzyme’s allosteric activation allowed us to group NT5C2 mutant proteins into three classes based on their responses to ATP: Class I – hyperactive and ATP independent, Class II – hyperactive and ATP hyper-responsive, and Class III – ATP hyper- responsive (Figure 5.2). Class I includes NT5C2 K359Q and NT5C2 L375F, which are both highly hyperactive in the absence of allosteric activation, with limited further increase in 5’-NT activity in response to ATP. Class II is the largest class of NT5C2 mutant enzymes (R39Q, R238W, S360P, R367Q,
D407A, S408R, S445F, R478S) which show a moderate degree of hyperactivity in the absence of allosteric activation, but also a strong response to increasing ATP concentrations. Class III consists of the unusual truncated mutant protein NT5C2 Q523*, which shows very weak hyperactivity in the absence of allosteric activation, but is hyper-responsive to increasing ATP concentrations.
Figure 5.2. NT5C2 mutant proteins can be separated into three classes based on their enzymatic behavior and response to allosteric activation by ATP. Representative NT5C2 mutant proteins are shown from Classes I, II and III. Class I mutant proteins are hyperactive and ATP-independent. Class II mutant proteins are hyperactive and ATP hyper-responsive. Class III contains an unusual truncated mutant protein which is not significantly hyperactive at basal conditions but is hyper-responsive to allosteric activation by ATP.
Class I NT5C2 mutations are neomorphic alleles which directly mimic allosteric activation
To confirm our modeling study of the Class I mutant NT5C2 K359Q we obtained the crystal structure of human NT5C2 K359Q bearing a substrate-trapping D52N mutation (Wallden and Nordlund
2011). For crystallography studies we used NT5C2 constructs of amino acids 1-536 out of 561 due to
89 Chapter 5 improved solubility as used in previous studies of wildtype NT5C2 (Wallden, Stenmark et al. 2007,
Wallden and Nordlund 2011), and which we confirmed retained the activating phenotype of relapse- associated NT5C2 mutations (Supplementary figure A5.1). Notably, our analysis (in collaboration with
Dr. Farhad Forouhar and Dr. Liang Tong at Columbia University) showed NT5C2 K359Q in the active conformation with helix A organized in the absence of IMP or ATP (Figure 5.3, Supplementary Table
A5.1), which is consistent with our previous modeling analysis and with the constitutive, allosterically independent hyperactivity of this mutant which we have observed. Closer examination of the allosteric architecture of NT5C2 K359Q revealed a shift in the hydrogen bonding network around helix A. In wildtype NT5C2 helix A is stabilized by a crucial bidentate interaction between K361 on helix A and D459, which is anchored in place by R367 and R39, both of which are recurrently mutated in relapsed ALL
(Figure 5.3a). The presence of Gln at position 359 results in a twist of D459, which loosens its hydrogen bonding with R367 and alters the neighboring network of interactions allowing for direct stabilization of helix A (Figure 5.3b). When we compared the quaternary tetramer structure of NT5C2 K359Q to that of wildtype NT5C2 we noticed that surprisingly NT5C2 K359Q has a much tighter dimer interface (Figure
a b
Figure 5.3. Crystal structure of NT5C2 K359Q shows a shift in the hydrogen bonding network at the allosteric region resulting in direct helix A stabilization and formation of the active enzyme conformation in the absence of allosteric activator or substrate. (a) Crystal structure of active wildtype NT5C2 with ATP activator and IMP substrate bound. Binding of ATP leads to ordering of allosteric helix A which is stabilized by a crucial interaction between K361 on helix A and D459, which in turn is supported by hydrogen bonding interactions with R39 and R367. (b) Crystal structure of NT5C2 K359Q showing ordering and stabilization of allosteric helix A in the absence of ATP or IMP. Notably, D459 is rotated and has lost one hydrogen bonding interaction with R367, resulting in stronger stabilizing interactions with K361 on helix A. Images courtesy of F. Forouhar.
90 Chapter 5
5.4a, b ,d), indicating that the K359Q mutation causes long-range structural changes in the protein architecture of NT5C2.
Interestingly, the second Class I mutation, NT5C2 L375F, is located at the NT5C2 monomer- monomer interface, in a highly hydrophobic pocket (Figure 5.5a). We modeled the effect of the L375F mutation on the NT5C2 structure, which suggested that the presence of Phe at position 375 would enhance inter-monomeric hydrophobic interactions to H486 on the neighboring subunit through favorable
π-stacking (Figure 5.5b). Thus we predicted that L375F would also lead to increased NT5C2 dimerization as we had observed for NT5C2 K359Q. We obtained the crystal structure of activated NT5C2 L375F
(Supplementary Table A5.1) which confirmed our modeling analysis, showing that the L375F results in four additional π-π stacking interactions: three within the same monomer to F36, F441 and Y461; and the fourth to H486 on the neighboring subunit (Figure 5.5c). Consistent with our prediction, this resulted in a tighter dimer interface but also in a tighter tetramer interface compared to wildtype NT5C2 (Figure 5.4a, c, d).
a b c d
Dimer Tetramer NT5C2 Interface interface (Å2) (Å2) WT 4,059 1,681 K359Q 4,984 1,557 L375F 4,142 1,740
Figure 5.4. Class I mutants NT5C2 K359Q and NT5C2 L375F have a tighter dimer interface than wildtype NT5C2. (a) Wildtype NT5C2 dimer showing contact area residues as shaded surfaces with color coding by monomer. (b) NT5C2 K359Q dimer showing contact surface area. (c) NT5C2 L375F dimer showing contact surface area. (d) Summary of dimer and tetramer contact surface areas for wildtype (WT) NT5C2 and Class I mutant proteins NT5C2 K359Q and NT5C2 L375F. Images courtesy of F. Forouhar.
When we assessed how the L375F change at the dimer interface translates to the allosteric activation center of the enzyme we again found that helix A is stabilized in the absence of ATP or IMP (Figure 5.6).
Remarkably, the distal L375F substitution leads to a significant change in the hydrogen bonding network around the K361 of helix A, with an almost 90° rotation of D459, resulting in complete loss of its hydrogen bonding interaction with R367 and a much tighter bidentate interaction with K361 which firmly stabilizes
91 Chapter 5 helix A (Figure 5.6). This structure is again consistent with our observation of NT5C2 L375F as an allosterically independent and highly hyperactive Class I mutant enzyme.
a b c
Figure 5.5. In silico model and crystal structure of NT5C2 L375F show additional stabilizing π-π stacking interactions at the monomer-monomer interface. (a) NT5C2 wildtype structure indicating location of L375 at the center of a densely hydrophobic pocket facing H486 on the neighboring monomer. (b) In silico model and (c) crystal structure of NT5C2 L375F, showing surrounding residues F36, F441 and Y461 on the same monomer and H486 on the neighboring monomer which take part in four new stabilizing π-π stacking interactions with F375. Images courtesy of Z. Carpenter and F. Forouhar.
a b
Figure 5.6. Crystal structure of NT5C2 L375F shows a shift in the hydrogen bonding network at the allosteric region resulting in direct helix A stabilization and formation of the active enzyme conformation in the absence of allosteric activator or substrate. (a) Crystal structure of active wildtype NT5C2 with ATP activator and IMP substrate bound. Binding of ATP leads to ordering of allosteric helix A which is stabilized by a crucial interaction between K361 on helix A and D459, which in turn is supported by hydrogen bonding interactions with R39 and R367. (b) Crystal structure of NT5C2 L375F showing ordering and stabilization of allosteric helix A in the absence of ATP or IMP. Notably, D459 is rotated at 90° and has lost both hydrogen bonding interactions with R367, resulting in much closer and stronger stabilizing interactions with K361 on helix A. Images courtesy of F. Forouhar.
92 Chapter 5
An overlay of the active NT5C2 wildtype structure with those of the two Class I mutant proteins
NT5C2 K359Q and NT5C2 L375F clearly shows the rotation and slight displacement of D459, R367 and key neighboring residues which partake in a dense hydrogen bonding network to support the crucial
K361-D459 bidentate interaction stabilizing helix A (Figure 5.7). Thus, the Class I (NT5C2 K359Q and
NT5C2 L375F) alleles can be considered as neomorphic structural alterations, in that they directly alter the resting state of the NT5C2 protein to induce enzyme activity. The presence of increased dimerization in these mutants indicates that conformational changes in the quaternary structure of NT5C2 are important for its activation, which is consistent with reported observations of increased oligomerization of wildtype NT5C2 in response to ATP (Spychala, Chen et al. 1999).
Figure 5.7. Class I NT5C2 mutations are neomorphic alleles which directly alter the architecture of the NT5C2 allosteric region, resulting in constitutive NT5C2 activation in the absence of allosteric activator. Overlay of crystal structures of active wildtype NT5C2 and Class I mutant proteins NT5C2 K359Q and NT5C2 L375F highlights subtle shifts in the hydrogen bonding network around allosteric helix A. In the Class I NT5C2 mutant structures the crucial D459 residue is rotated, resulting in a decrease or complete loss of its interaction with R367 and a closer stabilizing interaction with K361 on helix A. These changes lead to constitutive mimicking of the activated state of the enzyme independently of allosteric activation. Image courtesy of F. Forouhar.
Class II mutations conserve the architecture of the allosteric activation center of NT5C2 but block an auto-regulatory switch-off mechanism
By mapping the locations of all discovered and reported Class II mutations onto the structure of wildtype NT5C2 we noticed that they cluster in two distinct regions of the functional dimer unit. Some of
93 Chapter 5 the mutations (R367Q, R39Q, R238W/L/G/Q, S445F, R446Q, R478S) are located in a positively charged pocket between the two adjacent monomers which opens up once the enzyme is activated, while another set of mutations (K404N, 404-405insD, D407A/Y/E/H, S408R, P414S, D415G and delS408-D415) is located in a disordered stretch of amino acids (residues 395-417) which is not visible in the reported structure of wildtype NT5C2 (Wallden and Nordlund 2011) and which forms a flexible loop facing the entrance of the intermonomeric pocket, supported by two α-helices in a crane-like structure (Figure 5.8a).
This unstructured loop is 100% conserved in vertebrates and shows a highly specific ordering of polar, negatively charged, and hydrophobic residues (Figure 5.8b).
a b
Figure 5.8. Class II NT5C2 mutations map to two regions on the homodimer functional unit. (a) NT5C2 Class II mutations (shown in red) are located in a basic intermonomeric pocket which opens up when the enzyme is activated, and in an unstructured flexible loop supported by two α-helices in a crane-like structure facing the opening to the basic pocket. (b) NT5C2 orthologue alignment from humans to nematodes, showing invariant conservation of the flexible unstructured loop (residues 395-417) from humans to zebrafish. Images modified from Z. Carpenter.
Given that mutations in the pocket and the loop have the same enzymatic behaviors, we hypothesized that these two regions form part of a dynamic regulatory mechanism of NT5C2 activity. To test this hypothesis, we first performed an in silico analysis of the preferred conformation and localization of the unstructured loop in both the apo inactive and the IMP, ATP-bound active structures of wildtype
94 Chapter 5
NT5C2 (performed by Zachary Carpenter). We found that in the inactive state of NT5C2 the disordered loop is shorter (L(405)DSSSNERPD) and lies near the surface of the enzyme. However, in the activated
ATP-bound state of NT5C2, the unstructured loop becomes longer (E(399)LYKHLDSSSNERPD) which allows it to move dynamically into the inter-monomeric pocket which opens up upon activation, interacting with the basic residues in the cavity along the way (Figure 5.9).
a b
Figure 5.9. Flexible unstructured loop of NT5C2 enters into the basic intermonomeric pocket which opens up upon NT5C2 activation. Modeling analysis is shown indicating the top 20 preferred conformations of the flexible unstructured loop in (a) the inactive NT5C2 enzyme dimer and (b) the allosterically activated NT5C2 enzyme. Images modified from Z. Carpenter.
Our modeling analysis showed that the key residue in the loop mediating interaction with the positively charged pocket is D407, which is also recurrently mutated to different amino acids in relapsed
ALL. Notably, the most favorable (lowest potential energy) configuration of D407 in the activated state of
NT5C2 is near the bottom of the basic cavity, interacting via hydrogen bonding with the K361 on the neighboring monomer (Figure 5.10). This dynamic engagement of K361 by D407 would destabilize its bidentate interaction with D459 which is critical for the organization of the allosteric helix A (Figure 5.10).
In this manner D407 competes with D459 for the K361, leading to a destabilization of helix A and inactivation of NT5C2 (Figure 5.10). Thus, based on this model, allosteric activation of NT5C2 results in a conformational change which increases accessibility of the positively charged pocket between adjacent monomers (including helix A and surrounding residues) and allows the flexible loop of one monomer to destabilize helix A of the neighboring monomer, resulting in NT5C2 deactivation and thereby acting as an auto-regulatory switch-off mechanism. Hence, the NT5C2 mutations observed in the loop and in the
95 Chapter 5
a b
Figure 5.10. Following NT5C2 activation, D407 in the flexible loop invades into the basic pocket and interacts with K361 on the neighboring subunit, resulting in helix A destabilization and enzyme de- activation. (a) Structure of active wildtype NT5C2 protein showing key hydrogen bonding interactions stabilizing helix A in the active state and modeling analysis of the unstructured loop of the neighboring monomer (green) showing the most favorable configuration of D407 in the active state. (b) Schematic representation of the dynamic interaction between the D407 loop and K361 in helix A. Disruption of the helix A stabilizing interaction between K361 and D459 by D407 would return NT5C2 to its basal inactive configuration. Images modified from Z. Carpenter.
residues lining the basic pocket would interfere with the motion of the loop, blocking its auto-inhibitory action, which is consistent with our enzymatic data. This is further supported by our discovery of an in- frame partial deletion of the flexible loop (delS408-D415) in a relapsed case of ALL (Figure 5.8), which we have shown is activating and induces resistance to treatment with 6-MP in leukemic cells (Chelsea
Dieck, unpublished).
Our modeling analysis indicated that in the inactive state of NT5C2 the flexible loop rests on the surface of the homodimer and is more structured. To confirm this prediction we crystallized the inactive apo wildtype NT5C2 protein using improved crystallization conditions at physiological pH. We obtained a novel crystal structure of wildtype NT5C2 in its resting inactive state (Supplementary Table A5.1), which showed clear density for the flexible loop, highlighting the tight association between adjacent inactive
NT5C2 monomers, with the flexible loop straddling the top of the intermonomeric surface (Figure 5.11a).
Most notably, we observed a strong intermonomeric hydrogen bond network involving D407,
96 Chapter 5
a b
Figure 5.11. Novel crystal structure of inactive wildtype NT5C2 shows the flexible loop interacting with R238 on the neighboring monomer. (a) Structure of inactive wildtype NT5C2 dimer (monomers color-coded cyan and orange) highlighting each R238 residue located opposite the visible flexible loop of the neighboring monomer near the surface of the dimer. (b) Close-up view of the inactive wildtype NT5C2 structure indicating the hydrogen bond interactions between R238 and loop residues from the neighboring monomer. Images modified from F. Forouhar.
S408, S409 and S410 on the loop and R238 on the neighboring subunit (Figure 5.11b). Given the high prevalence of mutations in R238 to uncharged and hydrophobic amino acids (Chapter 3), we hypothesized that R238 is an important guide for the flexible loop which controls entry of the loop into the basic intermonomeric pocket upon NT5C2 activation. To confirm this hypothesis we solved the crystal structure of active NT5C2 R238W (Supplementary Table A5.1), which showed that the presence of a hydrophobic residue at position 238 is a much better fit for the surrounding local environment of the monomer, which includes a number of hydrophobic residues (L234, L235, L469, F473, M19, L24) (Figure
5.12). Thus, R238W results in the formation of a dense hydrophobic plug near the surface of the monomer, which would block entry of the flexible loop from the neighboring subunit into the positively charged intermonomeric pocket.
To study the structural effects of Class II mutations located at the bottom of the positively charged pocket we focused on the most recurrent NT5C2 mutant allele, NT5C2 R367Q. We solved the crystal structure of active NT5C2 R367Q (Supplementary Table A5.1) and found that, remarkably, the presence of a Gln at position 367 maintained the same hydrogen bonding interactions stabilizing helix A as are present in the active wildtype protein (Figure 5.13). Overlay of both Class II mutant structures, NT5C2
R238W and NT5C2 R367Q, with that of active wildtype NT5C2 showed that unlike Class I mutants, Class
II mutant proteins show strict conservation of the allosteric architecture supporting helix A in the active
97 Chapter 5 state of NT5C2 (Figure 5.14). These results indicate that Class II mutations lead to aberrant NT5C2 activation primarily by blocking the auto-regulatory switch-off mechanism of the flexible loop.
Figure 5.12. NT5C2 R238W structure shows formation of a dense hydrophobic plug near the entrance of the intermonomeric basic cavity. Overlay of wildtype (WT) NT5C2 and NT5C2 R238W structures indicates that W238 is a much better fit for the surrounding protein architecture than R238 due to the dense hydrophobic pocket around residue 238 made up of residues L234, L235, L469, F473, M19 and L24. The R238W mutation creates a strong hydrophobic plug near the entrance of the basic pocket which would block entry of the neighboring flexible loop, disrupting the ability of the loop to de-activate NT5C2. Image courtesy of F. Forouhar.
a b
Figure 5.13. Structure of active NT5C2 R367Q active structure shows conservation of hydrogen bonding interactions supporting allosteric helix A. (a) Structure of active wildtype NT5C2 showing hydrogen bonding interactions stabilizing allosteric helix A in the active state. (b) Structure of active NT5C2 R367Q showing that Q367 removes the basic charge of residue 367 while strictly conserving the hydrogen bonding interactions which stabilize helix A in the wildtype NT5C2 protein. Images courtesy of Z. Carpenter and F. Forouhar.
98 Chapter 5
Figure 5.14. Class II NT5C2 mutations show strict conservation of the allosteric architecture surrounding helix A while removing positive charges which interact with the auto-inhibitory flexible loop of the neighboring monomer. Overlay of crystal structures of active wildtype (WT) NT5C2 and Class II mutant proteins NT5C2 R238W and NT5C2 R367Q highlights complete conservation of the allosteric region architecture, indicating that the main mechanism of action of Class II mutations is to remove the positive charges that interact with D407 and S408 on the flexible loop. Image courtesy of F. Forouhar.
To functionally test this model we designed rabbit polyclonal antibodies against the flexible loop
(E(399)LYKHLDSSSNERPDI) and hypothesized that antibody-mediated sequestration of the loop would mimic the enzymatic activation effects of Class II mutations. Peptide-purified antibodies efficiently recognized recombinant purified NT5C2 protein (Figure 5.15a) and, consistent with our prediction, incubation of NT5C2 protein with either of two loop antibodies resulted in significant increase in 5’- nucleotidase activity, which was especially evident at increasing ATP concentrations (Figure 5.15b).
Moreover, addition of the corresponding loop blocking peptide specifically rescued the effect of each loop antibody (Figure 5.15b). These results phenocopy the enzymatic behavior of Class II mutant NT5C2 proteins and support our model of the flexible loop as a key switch-off mechanism required for the de- activation of NT5C2 following allosteric activation.
99 Chapter 5
a b
Figure 5.15. Antibodies against the NT5C2 flexible loop increase 5’-nucleotidase activity in the presence of ATP. (a) Western blot analysis showing recognition of recombinant purified NT5C2 protein by polyclonal antibodies designed against the NT5C2 flexible loop. Detection of an N-terminal hexahistidine tag (6x-His) on recombinant NT5C2 proteins serves as loading control. (b) 5’-nucleotidase assays of NT5C2 proteins pre- incubated with either buffer alone, with loop antibodies, or with loop antibodies plus loop blocking peptide at increasing concentrations of ATP allosteric activator. Antibody-mediated sequestration of the flexible loop phenocopies Class II NT5C2 mutations by increasing 5’-nucleotidase activity especially in the presence of ATP, and is efficiently abrogated by addition of antibody-specific blocking peptide.
The Class III truncating mutation NT5C2 Q523* removes a C-terminal acidic regulatory patch which serves as a brake to the allosteric activation of NT5C2
The NT5C2 C-terminus (including residues 488-561) appears to be poorly structured and has not been visualized to date (Wallden, Stenmark et al. 2007, Wallden and Nordlund 2011). While the non- recurrent NT5C2 Q523* mutation is a weakly activating allele with decreased impact on 6-mercaptopurine resistance in leukemic cells, it indicates that the C-terminus of NT5C2 has an auto-inhibitory role. Notably, the last 13 residues (549-561) constitute a poly-Asp/Glu acidic stretch, suggesting that this is an important regulatory element of the enzyme. To functionally test the role of the C-terminus we designed polyclonal rabbit antibodies against the NT5C2 C-terminal acidic patch
(Q(542)EITHCHDEDDDEEEEEEEE), which efficiently recognized the full-length recombinant NT5C2 protein but did not recognize the C-terminally truncated NT5C2 Q523* protein, indicating specific binding to the C-terminus and no non-specific binding to other polar regions on the protein (Figure 5.16a). Similar to results obtained with antibodies against the flexible loop, incubation of the full length NT5C2 protein
100 Chapter 5
a b c
Figure 5.16. Antibodies against the NT5C2 C-terminal acidic tail increase 5’-nucleotidase activity in the presence of ATP. (a) Western blot analysis showing recognition of recombinant purified NT5C2 full-length protein (WT), and lack of off-target recognition of NT5C2 Q523* truncated protein, by polyclonal antibodies designed against the NT5C2 C-terminal acidic tail. Detection of an N-terminal hexahistidine tag (6x-His) serves as loading control. (b) 5’-nucleotidase assays of NT5C2 proteins pre-incubated with either buffer alone, with C- terminus antibodies, or with C-terminus antibodies plus C-terminus blocking peptide at increasing concentrations of ATP allosteric activator. Antibody-mediated sequestration of the NT5C2 C-terminal acidic tail phenocopies the Class III NT5C2 truncating mutation by increasing 5’-nucleotidase activity in the presence of ATP, and is efficiently abrogated by addition of antibody-specific blocking peptide. (c) NT5C2 C-terminal antibodies show no off-target effects on NT5C2 Q523* truncated protein.
with either of two unique C-terminus antibodies resulted in significant increase in 5’-nucleotidase activity especially evident in the presence of allosteric activation by ATP (Figure 5.16b). This effect was efficiently abrogated by addition of the C-terminal specific blocking peptide and was not observed in the
C-terminal truncated NT5C2 Q523* protein (Figure 5.16b, c). Overall, these results recapitulate the enzymatic behavior of the NT5C2 Q523* mutation and confirm the auto-inhibitory role of the acidic C- terminal region of NT5C2.
To understand the structural basis of NT5C2 C-terminal auto-regulation, we pursued crystallization of full-length NT5C2 proteins despite challenges in protein solubility as encountered by
Wallden et al. (Wallden, Stenmark et al. 2007). We obtained the crystal structure of the activated full- length wildtype NT5C2 protein (with D52N substrate-trapping mutation) complexed with IMP, Mg2+, and
ATP (Supplementary Table A5.2). This structure was identical to the already reported structure of the C- terminally truncated wildtype NT5C2 (Wallden and Nordlund 2011) with the C-terminus not visible, suggesting that the C-terminus is not ordered in the activated state of NT5C2. To compare this structure
101 Chapter 5 to the inactive state of NT5C2 we pursued crystallization of apo, uncomplexed full-length NT5C2 proteins.
We obtained the crystal structure of apo, inactive full-length NT5C2 R367Q (Supplementary Table A5.2) which yielded partial density for the C-terminus. Notably, we found the C-terminal acidic tail to occupy the intermonomeric positively charged cavity, interacting favorably with basic residues from both subunits of the NT5C2 dimer (Figures 5.17a and 5.18a). We obtained a similar structure of inactive full-length
NT5C2 R39Q (Supplementary Table A5.2) which had poorer resolution but also showed the acidic tail bound in the basic pocket, in this case confirming direct interaction of the C-terminal tail with the R367 residue (Figure 5.18b) and indicating that the C-terminus communicates directly with the allosteric activation center of the enzyme.
a b
Figure 5.17. Novel crystal structure of inactive full length NT5C2 R367Q shows the C-terminus wrapped around the neighboring monomer with the acidic tail tucked into the basic intermonomeric pocket. Structure of inactive NT5C2 R367Q dimer (monomers color-coded yellow and cyan) showing(a) the C-terminal acidic tail interacting with the positively charged interface between adjacent subunits and (b) rotated view showing partial structure for the C-terminal region directly upstream of the acidic tail, including a short α-helical segment from E501-R509 and an unstructured sequence (T510-L536) predicted to be a disordered α-helix. Images courtesy of F. Forouhar.
A section of the C-terminus from R509 to F537 (predicted to be a disordered α-helix) is not visible in the NT5C2 R367Q structure, but seems to encircle the neighboring monomer, with E501-R509 forming a visible short α-helix bound in a groove on the adjacent subunit (Figure 5.17b). Interestingly, we noticed that in this full-length inactive structure the N-terminus of NT5C2 is not visible, whereas the crystal structures of active NT5C2 proteins we have obtained show a completely organized N-terminus.
Comparison of these structures showed us that the same groove occupied by the C-terminal E501-R509
102 Chapter 5
a b
Figure 5.18. In the inactive (apo) state, the NT5C2 acidic C-terminal tail interacts with basic residues from both adjacent NT5C2 dimer subunits, including R367 in the allosteric region. (a) Structure of inactive NT5C2 R367Q dimer (monomers color-coded cyan and yellow) highlighting in pink the basic residues from each monomer which interact with the acidic C-terminal tail. (b) Structure of inactive NT5C2 R39Q dimer (monomers color-coded cyan/yellow and pink) highlighting hydrogen bonding interaction between D551 on the C-terminal acidic tail (cyan) and the R367 of the same monomer. Images courtesy of F. Forouhar.
a b
Figure 5.19. Interchanging occupation of a surface groove on NT5C2 by the N-terminus of the same monomer and the C-terminus of the adjacent monomer in active and inactive enzyme states respectively. (a) Overlay of active and inactive NT5C2 structures showing surface groove on subunit A occupied by its own N- terminal α-helix (starting at T3) in the active state (pink), whereas in the inactive state the N-terminal region of subunit A is not visible (cyan) and the groove is occupied by the C-terminal E501-R509 α-helix of subunit B (yellow). (b) Model showing conformational changes in key regulatory regions of the NT5C2 dimer upon activation. In the inactive state each C-terminus is wrapped around the neighboring monomer, holding the dimer tightly shut with the flexible loop resting on the surface. When the enzyme becomes activated the C-terminus loosens and is replaced by the monomer’s own N-terminus. The basic pocket is opened allowing catalysis to occur and the flexible loop to enter and de-activate the enzyme by destabilizing the allosteric region.
103 Chapter 5 residues of the neighboring monomer in the inactive state is occupied by the N-terminus from the same monomer in the active state (Figure 5.19a). Thus there appears to be an interchange between the N- and
C-termini of the two neighboring monomers during activation. In the inactive state each C-terminus is wrapped around the adjacent subunit, holding the enzyme in a tightly shut conformation, with the N- termini unstructured. In the activated state a conformational change is induced which leads to displacement of the C-terminus from the neighboring monomer and binding of each monomer’s own N- terminus in the previously occupied groove. This may leave the C-terminus in a largely unstructured conformation, which could explain our inability to visualize it in the active state of NT5C2. A loosened or absent state of the C-terminus from the basic cavity would facilitate allosteric activation and allow for the subsequent dynamic de-activating motion of the flexible loop (Figure 5.19b). This model is consistent with our enzymatic data, indicating that the C-terminus acts as a brake to the allosteric activation of
NT5C2, whereby its absence does not directly confer increased 5’-nucleotidase activity but lowers the threshold for allosteric activation.
Finally, we noted that Cys547 located immediately upstream of the C-terminal acidic tail lies within very close proximity to Cys121 on the neighboring subunit (Figure 5.20), indicating that a disulfide
Figure 5.20. Cys547 in the NT5C2 C-terminus can form a disulfide bridge with Cys121 on the neighboring monomer under oxidizing conditions. Structure of full length inactive NT5C2 R367Q with one monomer shown in yellow and C-terminal region of neighboring monomer shown in blue. Cys121 and Cys547 which are within bonding distance are shown in red. In green is shown Cys175 which has been proposed to form a disulfide bridge with Cys547 by Allegrini et al, 2004, but which is not within bonding distance to Cys547 in this structure.
104 Chapter 5 bridge may form between these two residues under oxidizing conditions, which would effectively lock the dimer in a tight inactive state, suggesting that the role of the C-terminus may be strongly influenced by the biochemical and redox state of the cell.
Discussion
We have shown that recurrent activating mutations in the NT5C2 nucleotidase gene drive resistance to 6-mercaptopurine chemotherapy in up to 25% of relapsed T-ALLs and 6% of relapsed B-
ALLs. Our extensive sequencing analyses showed that NT5C2 mutations occur in a number of regions of the NT5C2 protein, with a few hotspots including R367, R238, R39Q and D407. Through enzymatic and structural studies we have identified three different classes of NT5C2 activating mutations, each with unique enzymatic behaviors and structural features.
Class I mutations are structurally neomorphic alleles which directly alter the allosteric region architecture of the enzyme to mimic the activated state even in the absence of allosteric activator such as
ATP, resulting in high levels of hyperactivity at baseline which are largely independent of ATP.
Class II mutations represent the biggest group, targeting both the positively charged intermonomeric cavity and the flexible loop which faces the entrance to this basic pocket. Our modeling, enzymatic and structural studies identified these two regions as part of a novel regulatory switch-off mechanism of NT5C2, whereby activation of the enzyme results in a conformational change that lengthens the flexible loop and opens the basic intermonomeric cavity, allowing the loop to dynamically interact with and displace positively charged residues in the pocket such as K361 which maintain the allosterically activated state of helix A. This motion results in destabilization of helix A and de-activation of the enzyme. Class II mutations in the NT5C2 basic cavity and flexible loop both result in moderate NT5C2 hyperactivity in baseline conditions and increased responses to allosteric activation, supporting a joint role for these two regions as an auto-regulatory switch-off mechanism of the enzyme. Moreover, NT5C2 mutations at the entrance and at the bottom of the positively charged pocket between adjacent monomers show strict conservation of the allosteric activation center architecture, which is in stark contrast to Class I mutations and indicates that the primary mechanism of Class II mutations is to block the dynamic motion of the auto-inhibitory flexible loop.
105 Chapter 5
Finally, we identified Class III representing a unique truncating mutation, NT5C2 Q523*, which shows negligible hyperactivity in baseline conditions yet is hyper-responsive to allosteric activation. Our enzymatic studies showed that antibody-mediated specific sequestration of the C-terminal acidic tail of
NT5C2 phenocopies the NT5C2 Q523* mutation, indicating that this acidic patch has an auto-inhibitory role. Our novel structural studies of the C-terminus show that the C-terminal acidic patch is bound in the intermonomeric basic pocket in the inactive state but is disordered in the active state of NT5C2, suggesting that the C-terminus helps to hold each dimer in a tightly closed state when inactive, acting as a brake to the allosteric activation of the enzyme. This model is consistent with our observations that
NT5C2 Q523* does not directly activate the enzyme but does result in increased response to ATP.
Although we are yet to visualize the C-terminus in a structure of wildtype NT5C2, the location of the C- terminus in the R367Q and R39Q structures is likely to be highly representative of that in the wildtype protein. Our findings that the NT5C2 C-terminus can wrap around the adjacent monomer with the acidic tail in the intermonomeric pocket are consistent with the proposed role of the C-terminal acidic tail in subunit association (Spychala, Chen et al. 1999).
Our observation of potential disulfide bridge formation between Cys547 and Cys121 on neighboring subunits supports a role for the C-terminus as a redox-dependent mechanism that can lock the NT5C2 dimer in a tight, inactive state under oxidizing conditions. The involvement of Cys547 in a disulfide bridge has previously been proposed as an auto-inhibitory mechanism, however as partner to
Cys175 rather than Cys121 (Allegrini, Scaloni et al. 2004). Our structural analyses of full length NT5C2
R367Q and NT5C2 R39Q showed that, while Cys547 lies within bonding distance to Cys121 on the neighboring subunit, it is not close enough to Cys175 for disulfide bridge formation. Further structural studies on the NT5C2 C-terminus in both active and inactive states should provide more insight on its function, yet our results thus far indicate that the C-terminus plays a subtle and context-specific role in the auto-regulation of NT5C2 activity, dictated by changing biochemical and redox conditions.
The activity of NT5C2 may indeed be closely linked to the state of the cell. While NT5C2 activation by ATP and ADP indicates that NT5C2 responds to changes in the adenylate energy charge of the cell (Itoh 1981, Itoh, Oka et al. 1986), our enzymatic analyses showed a stronger reactivity of both wildtype and mutant NT5C2 proteins to Ap4A, a molecule which has been proposed as a key signaling
106 Chapter 5 factor during apoptotis (Vartanian, Suzuki et al. 2003). Our results are consistent with previous studies of wildtype NT5C2 (Pinto, Canales et al. 1986) and suggest that Ap4A may specifically activate NT5C2 during apoptosis to increase catabolism of excess purine nucleotides from DNA and RNA degradation, which can be exported for uptake by neighboring cells.
While both ATP and Ap4A induced strong levels of NT5C2 activation, our analyses showed that
ADP was a far weaker allosteric activator of both wildtype and mutant NT5C2 proteins, with NT5C2
S445F being an exception and showing a slightly stronger response. Interestingly, S445 lies near a proposed second effector site on NT5C2, where an adenosine has been visualized with a completely disordered ribose moiety (Pesi, Baiocchi et al. 1998, Wallden, Stenmark et al. 2007), but which has not been confirmed by subsequent studies (Wallden and Nordlund 2011). It is possible that this site may mediate NT5C2 allosteric activation by ADP, although given the very weak response of NT5C2 wildtype protein to ADP, and the much stronger effects of relapse-associated NT5C2 mutations on allosteric activation by ATP and Ap4A, it is unlikely that ADP plays a large role in the activation of NT5C2.
Given the central role of NT5C2 in purine nucleotide homeostasis and its high degree of conservation within and beyond mammals, our results uncover a notable case of convergent structural evolution, where we have identified three different mutational mechanisms of activating this ubiquitous enzyme, each with unique enzymatic and structural characteristics. These findings highlight the intense selective pressure of 6-mercaptopurine chemotherapy as a Darwinian evolutionary force on leukemic cells during maintenance therapy. In addition, our novel structural understanding of mutant NT5C2 hyperactivation will serve as an essential tool in the design of NT5C2 inhibitors with increased specificity and selectivity for leukemia-associated mutant hyperactive NT5C2 enzymes.
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CHAPTER 6
CONCLUSIONS
“...for the shield may be as important for victory, as the sword or spear.”
Charles Darwin, The Origin of Species
Acute lymphoblastic leukemia (ALL) is an aggressive hematological malignancy that affects both children and adults but is especially prevalent in the pediatric population, representing the most common childhood malignancy and the greatest cancer-related cause of death before 20 years of age (Linabery and Ross 2008, Smith, Seibel et al. 2010). Despite pioneering advances in treatment which have transformed ALL from a uniformly fatal malignancy into a curable disease, 20% of pediatric patients and
50% of adult patients either fail to achieve remission or develop relapsed disease. Relapsed ALL remains a great clinical challenge with a survival rate of only 40% and with little progress seen in recent trials
(Raetz, Borowitz et al. 2008, Bhojwani and Pui 2013).
The advent of next-generation sequencing has allowed us to uncover the genetic mechanisms driving disease progression and relapse of ALL and to gain insight into the clonal diversity and evolution of this malignancy from diagnosis to relapse. We have discovered that in most cases ALL exists as a heterogeneous disease comprised of multiple competing leukemic clones which drive the continuous evolution of the tumor population in response to various changing selective pressures. Most notably we have identified activating mutations in the NT5C2 nucleotidase gene as the most recurrent relapse- specific alterations in ALL and as major drivers of chemotherapy resistance to 6-mercaptopurine.
Clinically we have found that NT5C2 mutations associate with disease progression under therapy and with early relapse, indicative of poorer outcome. Given that daily oral 6-mercaptopurine constitutes the core 2-year maintenance therapy for ALL following remission (Inaba, Greaves et al. 2013), our findings reveal a key role for chemotherapy as a selective pressure in the evolution of relapsed ALL. The evolutionary impact of 6-mercaptopurine therapy has recently become even more evident with our co- discovery of activating mutations in the PRPS1 purine biosynthesis enzyme as another mechanism of 6- mercaptopurine resistance in 6.7% of relapsed ALL (Li, Li et al. 2015).
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Chapter 6
Remarkably, we have discovered that while NT5C2 mutations provide leukemic cells with the advantage of resistance to 6-mercaptopurine therapy, they result in a severe metabolic disadvantage which translates into reduced tumor cell growth and decreased leukemia initiating activity in the absence of chemotherapy. Our findings provide the first known example of a genetic alteration which drives chemotherapy resistance yet results in decreased leukemic cell fitness, highlighting the dominance of chemotherapy pressure over clonal competition in the positive selection of leukemic clones harboring
NT5C2 mutations.
The strong impact of chemotherapy as a selective pressure on tumor populations is further illustrated by our recurrent findings of relapsed ALL samples containing multiple clones with NT5C2 activating mutations. These cases represent a remarkable example of chemotherapy-driven convergent evolution towards activating NT5C2, a ubiquitously expressed and highly conserved enzyme from vertebrates to nematodes (Itoh 2013). This convergent evolution is also evidenced by the presence of relapse-specific NT5C2 mutations in both B-precursor and T-cell ALLs, two unique molecular subtypes of
ALL with differing metabolic needs, yet both treated with prolonged 6-mercaptopurine therapy following onset of remission (Inaba, Greaves et al. 2013).
Finally, we have also identified three convergent mutational mechanisms of activating NT5C2 on a protein structural level, with a few mutations directly hyperactivating the enzyme in basal conditions
(Class I), the majority of mutations abrogating an auto-regulatory switch-off mechanism to cause moderate hyperactivation and hyper-responsiveness to allosteric activation (Class II), and a unique truncating mutation which is weakly activating and removes a C-terminal auto-inhibitory brake to the enzyme’s allosteric activation (Class III). These multiple mechanisms of activating a structurally conserved enzyme which is subject to tight and complex regulation further highlight the intense chemotherapy pressure on tumor cells which drive relapse of ALL.
Interestingly, the difference in prevalence of each class of NT5C2 activating mutations could be explained by a fine balance between providing resistance to chemotherapy and conferring a metabolic disadvantage to the tumor cells. Consistently, Class I NT5C2 mutations which cause very high levels of allosterically independent NT5C2 hyperactivation are very rare, most likely because they result in a severe imbalance of the endogenous purine nucleotide pool. The Class II NT5C2 mutations are most
109 Chapter 6 frequently observed and result in moderate NT5C2 hyperactivation which allows for efficient processing of active 6-mercaptopurine metabolites while allowing responsiveness to changing concentrations of allosteric activators in the cytosol. Finally, the Class III truncating mutation would be too weak to provide sufficient resistance to 6-mercaptopurine chemotherapy, which explains its lack of recurrence in our extended series of relapsed ALL cases.
Our in-depth enzymatic and structural characterizations of the NT5C2 activating mutations in relapsed ALL and of the previously unsolved NT5C2 C-terminus will serve as an essential foundation for the rational drug design of NT5C2 inhibitors with increased efficacy and selectivity for mutant NT5C2 proteins. In particular, our expanded knowledge of NT5C2 will uniquely allow us to search for novel allosteric or mixed enzyme inhibitors in addition to NT5C2 substrate analogs.
The conditional inducible Nt5c2 R367Q knock-in mouse model we have developed has allowed us to accurately study the effects of heterozygous NT5C2 mutations in primary isogenic T-ALL tumors, demonstrating not only the associated loss of fitness and leukemia initiating activity in the absence of chemotherapy, but also that expression of a single allele of mutant Nt5c2 is sufficient to induce overt resistance and progression under therapy with 6-mercaptopurine in vivo. Moreover these studies have suggested that dose-intensification of 6-mercaptopurine may overcome mutant NT5C2-induced resistance. In this respect, our Nt5c2 R367Q mouse model can serve as a key tool for future studies on maintenance therapy dosing and compliance. Moreover, this model will be an excellent platform for the preclinical testing of NT5C2 inhibitors and other compounds to which NT5C2 mutant tumor cells may be sensitized.
As the new era of precision medicine is upon us, our studies on NT5C2 will invaluably inform both the future development of diagnostic testing for mutant NT5C2 and the personalized treatment of NT5C2- mutant leukemias. Given the presence of NT5C2 mutations not only in relapsed B-precursor ALLs but also in a quarter of relapsed T-ALLs – a subtype with particularly poor prognosis – our findings stand to make a vital contribution in the prevention and treatment of currently intractable drug-resistant and relapsed leukemias.
110
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APPENDIX
SUPPLEMENTARY FIGURES AND TABLES
I. Chapter 3
Supplementary Figures
Supplementary Figure A3.1. TP53, RPL11 and BANP mutations in relapsed pediatric ALL. Schematic representation of the structure of the TP53, RPL11 and BANP proteins. Missense mutations identified in these genes in relapsed pediatric patients are indicated with filled circles. Open circles represent frameshift mutations.
147
Appendix
Supplementary Figure A3.2. Western blot analysis of NT5C2 expression in CCRF-CEM and CUTLL1 cells transduced with wild type and mutant NT5C2 expressing lentiviruses.
Supplementary Figure A3.3. 5'-IMP nucleotidase activity in NT5C2 mutant proteins. 5'-Nucleotidase activity levels of NT5C2 R238W, NT5C2 L375F and NT5C2 R238W, L375F (cis) recombinant mutant proteins, as well as a 1:1 mixture of NT5C2 R238W and NT5C2 L375F proteins (trans) relative to wild type NT5C2 control are shown. Data shown are means ± s.d.
148 Appendix
Supplementary Figure A3.4. Expression of NT5C2 R238W, L375F in ALL cells does not induce resistance to 6-MP or 6-TG. Viability assays in (a) CCRF-CEM T-ALL cells and (b) CUTLL1 cells expressing wild type NT5C2, NT5C2 R238W, L375F or a red fluorescent protein control (RFP), treated with increasing concentrations of 6- mercaptopurine (6-MP) or 6-Thioguanine (6-TG). Data is shown as means ± s.d.
Supplementary Tables
Supplementary table A3.1. Summary of patient samples and genetic analyses
Sample cohorts N Source Analyses T-ALL diagnosis / 5 AIEOP (University of Padua) Whole exome remission / relapse sequencing T-ALL diagnosis / 18 AIEOP (University of Padua) TP53, BANP, remission / relapse RPL11, NRAS, NT5C2 mutation analysis T-ALL diagnosis / 21 COG (n=20) Whole exome remission / relapse* AIEOP (University of Padua) (n=1) sequencing* T-ALL relapse 80 AIEOP (University of Padua) (n=13) NT5C2 mutation BFM (Charité-Universitätsmedizin Berlin) (n=67) analysis B-precursor ALL 14 Saitama Children's Medical Center, Japan (n=7) Whole exome diagnosis / remission / Hospital Central de Asturias, Spain (n=2) sequencing* relapse* Columbia University Medical Center (n=5) B-precursor ALL 501 AIEOP (University of Padua) (n=35) NT5C2 mutation relapse BFM (Charité-Universitätsmedizin Berlin) (n=466)* analysis T-ALL diagnosis 23 ECOG NT5C2 mutation analysis B-precursor ALL 27 AIEOP (University of Padua) NT5C2 mutation diagnosis analysis
* Complete results of whole exome and targeted sequencing will be published separately (manuscript submitted)
149 Appendix
Supplementary Table A3.2. Mutations identified by exome sequencing in diagnosis and relapse T- ALL
150 Appendix
Supplementary table A3.3. Genomic regions of LOH in diagnostic and relapsed T-ALLs analyzed by whole exome sequencing
Sample Chromosome Start End Cytoband Sample T-ALL 16 12 116408538 133768553 12q24.21 - 12q24.33 Relapse T-ALL 38 6 30954873 31323353 6p21.33 Diagnosis T-ALL 47 1 949608 54644859 1p36.33 - 1p32.3 Diagnosis T-ALL 47 1 949608 120437718 1p36.33 - 1p12 Relapse T-ALL 47 9 117713 32425910 9p24.3 - 9p21.1 Diagnosis T-ALL 11 9 214864 21816758 9p24.3 - 9p21.3 Diagnosis
151 Appendix
Supplementary Table A3.4. NT5C2 mutations in T-ALL
Amino acid Ref. Seq Source Sample ID Exon change NM_012229 AIEOP (University of Padua) Relapse T-ALL 17 9 R238L 713G>T AIEOP (University of Padua) Relapse T-ALL 22 9 R238W 712C>T AIEOP (University of Padua) Relapse T-ALL 29 11 R291W 871C>T AIEOP (University of Padua) Relapse T-ALL 11 13 K359Q 1075A>C AIEOP (University of Padua) Relapse T-ALL 4 13 R367Q 1100G>A AIEOP (University of Padua) Relapse T-ALL 17 13 R367Q 1100G>A AIEOP (University of Padua) Relapse T-ALL 35 13 R367Q 1100G>A AIEOP (University of Padua) Relapse T-ALL 37 15 D407A 1220A>C AIEOP (University of Padua) Relapse T-ALL 30 17 Q523* 1567C>T BFM (Charité-Universitätsmedizin Berlin) Relapse T-ALL B60 2 R39Q 116 G>A BFM (Charité-Universitätsmedizin Berlin) Relapse T-ALL B9 9 R238W 712C>T BFM (Charité-Universitätsmedizin Berlin) Relapse T-ALL B29 9 R238W 712C>T BFM (Charité-Universitätsmedizin Berlin) Relapse T-ALL B64 9 R238W 712C>T BFM (Charité-Universitätsmedizin Berlin) Relapse T-ALL B11 13 R367Q 1100G>A BFM (Charité-Universitätsmedizin Berlin) Relapse T-ALL B15 13 R367Q 1100G>A BFM (Charité-Universitätsmedizin Berlin) Relapse T-ALL B30 13 R367Q 1100G>A BFM (Charité-Universitätsmedizin Berlin) Relapse T-ALL B37 13 R367Q 1100G>A BFM (Charité-Universitätsmedizin Berlin) Relapse T-ALL B39 13 R367Q 1100G>A BFM (Charité-Universitätsmedizin Berlin) Relapse T-ALL B44 13 R367Q 1100G>A BFM (Charité-Universitätsmedizin Berlin) Relapse T-ALL B48 13 R367Q 1100G>A BFM (Charité-Universitätsmedizin Berlin) Relapse T-ALL B52 13 R367Q 1100G>A BFM (Charité-Universitätsmedizin Berlin) Relapse T-ALL B53 13 R367Q 1100G>A BFM (Charité-Universitätsmedizin Berlin) Relapse T-ALL B9 13 L375F 1123C>T BFM (Charité-Universitätsmedizin Berlin) Relapse T-ALL B64 13 L375F 1123C>T BFM (Charité-Universitätsmedizin Berlin) Relapse T-ALL B63 15 D407Y 1219 G>T Children's Oncology Group Relapse T-ALL C5 9 R238W 712C>T Children's Oncology Group Relapse T-ALL C10 9 R238W 712C>T Children's Oncology Group Relapse T-ALL C4 13 R367Q 1100G>A Children's Oncology Group Relapse T-ALL C7 13 R367Q 1100G>A Children's Oncology Group Relapse T-ALL C11 13 R367Q 1100G>A Children's Oncology Group Relapse T-ALL C17 13 R367Q 1100G>A Children's Oncology Group Relapse T-ALL C18 13 R367Q 1100G>A Children's Oncology Group Relapse T-ALL C20 13 R367Q 1100G>A Children's Oncology Group Relapse T-ALL C14 15 D407E 1221 C>A Children's Oncology Group Relapse T-ALL C20 16 R478S 1434 G>C
152 Appendix
Supplementary Table A3.5. NT5C2 mutations in B-ALL
Amino acid Ref. Seq Source Sample ID Exon change NM_012229 AIEOP (University of Padua) Relapse pre-B ALL 16 13 R367Q 1100G>A BFM (Charité-Universitätsmedizin Berlin) Relapse pre-B ALL B2993 2 R39Q 116G>A BFM (Charité-Universitätsmedizin Berlin) Relapse pre-B ALL B3544 2 R39Q 116G>A BFM (Charité-Universitätsmedizin Berlin) Relapse pre-B ALL B3089 9 R238W 712C>T BFM (Charité-Universitätsmedizin Berlin) Relapse pre-B ALL B3181 9 R238W 712C>T BFM (Charité-Universitätsmedizin Berlin) Relapse pre-B ALL B3499 9 R238W 712C>T BFM (Charité-Universitätsmedizin Berlin) Relapse pre-B ALL B2633 9 R238L 713G>T BFM (Charité-Universitätsmedizin Berlin) Relapse pre-B ALL B3374 9 R238L 713G>T BFM (Charité-Universitätsmedizin Berlin) Relapse pre-B ALL B2554 13 R367Q 1100G>A BFM (Charité-Universitätsmedizin Berlin) Relapse pre-B ALL B2738 13 R367Q 1100G>A BFM (Charité-Universitätsmedizin Berlin) Relapse pre-B ALL B2823 13 R367Q 1100G>A BFM (Charité-Universitätsmedizin Berlin) Relapse pre-B ALL B2863 13 R367Q 1100G>A BFM (Charité-Universitätsmedizin Berlin) Relapse pre-B ALL B2897 13 R367Q 1100G>A BFM (Charité-Universitätsmedizin Berlin) Relapse pre-B ALL B2899 13 R367Q 1100G>A BFM (Charité-Universitätsmedizin Berlin) Relapse pre-B ALL B2916 13 R367Q 1100G>A BFM (Charité-Universitätsmedizin Berlin) Relapse pre-B ALL B2925 13 R367Q 1100G>A BFM (Charité-Universitätsmedizin Berlin) Relapse pre-B ALL B3246 13 R367Q 1100G>A BFM (Charité-Universitätsmedizin Berlin) Relapse pre-B ALL B3348 13 R367Q 1100G>A BFM (Charité-Universitätsmedizin Berlin) Relapse pre-B ALL B3388 13 R367Q 1100G>A BFM (Charité-Universitätsmedizin Berlin) Relapse pre-B ALL B3407 13 R367Q 1100G>A BFM (Charité-Universitätsmedizin Berlin) Relapse pre-B ALL B3454 13 R367Q 1100G>A BFM (Charité-Universitätsmedizin Berlin) Relapse pre-B ALL B3499 13 R367Q 1100G>A BFM (Charité-Universitätsmedizin Berlin) Relapse pre-B ALL B3430 13 D384N 1150G>A BFM (Charité-Universitätsmedizin Berlin) Relapse pre-B ALL B3077 15 K404N 1212G>T BFM (Charité-Universitätsmedizin Berlin) Relapse pre-B ALL B2816 15 D407Y 1219G>T BFM (Charité-Universitätsmedizin Berlin) Relapse pre-B ALL B3477 15 D407H 1219G>C BFM (Charité-Universitätsmedizin Berlin) Relapse pre-B ALL B3212 15 del S408-D415 del 1223_1246 BFM (Charité-Universitätsmedizin Berlin) Relapse pre-B ALL B2577 15 P414S 1240C>T BFM (Charité-Universitätsmedizin Berlin) Relapse pre-B ALL B2885 15 P414S 1240C>T BFM (Charité-Universitätsmedizin Berlin) Relapse pre-B ALL B3477 15 D415G 1244A>G BFM (Charité-Universitätsmedizin Berlin) Relapse pre-B ALL B3145 16 V454M 1360G>A Saitama Children's Medical Center, 1334 C>T, Relapse pre-B ALL C4 16 S445F, R446Q Japan 1337 G>A
153 Appendix
Supplementary Table A3.6. Correlative clinical data on NT5C2-mutated relapsed T-ALLs treated in AIEOP clinical trials.
Clinical protocols are described in Conter et al, J Clin Oncol 1997; 15:2786-91 (AIEOP LAL91); Lo Nigro et al, Med and Pediatr Oncol 2000; 35:449-455 (AIEOP LAL95) and Cario et al, Blood 2010; 115:5393-5397 (AIEOP LAL2000).
Supplementary Table A3.7. Correlative clinical and molecular data on NT5C2-mutated relapsed T- ALLs in BFM based protocols.
Clinical protocols are described in Möricke et al, Blood 2008; 111:4477-4489 (NHL/ALL-BFM 95); Escherich et al, Hematologica 2011; 96:854-862 (COALL 06-97); Scherey et al, Pediatr Blood Cancer 2010; 54:952-958 (ALL- BFM2000); Oschlies et al, Am J Surg Pathol 2011; 35:836-44 (Euro-LB 02).
154 Appendix
Supplementary Table A3.8. Correlative clinical data on rescue treatment for NT5C2-mutated relapsed T-ALLs in BFM based protocols.
Clinical protocols are described in Von Stackelberg et al, Med Pediatr Oncol 2002; 39: 236 (ALL-REZ BFM 96) and at clinicaltrials.gov (http://clinicaltrials.gov/show/NCT00114348) (ALL-REZ BFM 2002).
155 Appendix
Supplementary Table A3.9. Association of NT5C2 mutations with clinical characteristics, response, outcome and genetics in T-ALL patients treated in BFM based protocols.
156 Appendix
Supplementary Table A3.9 continued
1) time of relapse: very early, <18 months after initial diagnosis of ALL; early, ≥18 months after initial diagnosis of ALL; late, ≥6 months after completion of primary treatment. 2) site of relapse: isolated BM, no evidence of extramedullary disease; combined BM, more than 5% leukemic cells in the BM combined with CNS, testis, or other extramedullary disease. 3) cytologic response: early, remission after first induction course; normal, remission after second induction course; st st late, remission after 1 R2 block/day 15 protocol II-IDA; non-response, no remission after 1 R1 block/day 29 protocol II-IDA. Cytologic remission was defined as less than 5% leukemic blasts in regenerating bone marrow, no peripheral blasts cells and no extramedullary involvement. 4) ALL-BFM includes patients treated according to protocols ALL-BFM 90, ALL-BFM 95 and ALL-BFM 2000. 5) COALL includes patients treated according to protocols CoALL 05-92, CoALL 06-97 and CoALL 07-03. 6) BFM – lymphoblastic lymphoma includes patients treated according to protocols NHL-BFM 90, NHL-BFM 95 and Euro-LB 02. 7) high-risk treatment includes patients treated according to the high-risk arm of protocols ALL-BFM 90, ALL-BFM 95, ALL-BFM 2000, CoALL 06-97 and CoALL 07-03. 8) non high-risk treatment includes patients treated according to the standard risk or intermediate risk arm of protocols ALL-BFM 90, ALL-BFM 95 and ALL-BFM 2000.
Abbreviations: BM, bone marrow; CR2, second complete remission; CCR, complete continuous remission; MRD, minimal residual disease in bone marrow aspiration after induction phase (week 5); HSCT, hematopoietic stem cell transplantation.
157 Appendix
Supplementary Table A3.10. Association of frontline treatment with NT5C2 mutations.
1) Patients without data regarding frontline treatment were excluded from the statistical analysis. 2) ALL-BFM includes patients treated according to protocols ALL-BFM 90, ALL-BFM 95 and ALL-BFM 2000. 3) COALL includes patients treated according to protocols CoALL 05-92, CoALL 06-97 and CoALL 07-03. 4) BFM - lymphoblastic lymphoma includes patients treated according to protocols NHL-BFM 90, NHL-BFM 95 and Euro-LB 02. 5) high-risk treatment includes patients treated according to the high risk arm of protocols ALL-BFM 90, ALL-BFM 95, ALL-BFM 2000, CoALL 06-97 and CoALL 07-03. 6) non high-risk treatment includes patients treated according to the standard risk or intermediate risk arm of protocols ALL-BFM 90, ALL-BFM 95 and ALL-BFM 2000. 7) Equality of categorical variables was analyzed by Fisher’s exact test (2-sided).
Supplementary Table A3.11. Multivariate binary logistic regression with time of relapse (on/off treatment) as outcome variable.
1) The total number of patients included in the analysis is 64 of 67 ALL-REZ BFM T-ALL relapsepatients. Three patients were excluded because of missing data on frontline treatment.
158 Appendix
Supplementary Table A3.12. Nucleoside analog IC50 values (μM) of T-ALL cell lines expressing relapse-associated NT5C2 mutant alleles.
6-mercaptopurine 6-thioguanine
CCRF-CEM CUTLL1 CCRF-CEM CUTLL1
RFP* 3.07 0.71 RFP* 0.77 0.45
NT5C2 WT# 3.54 0.41 NT5C2 WT# 1.10 0.51
NT5C2 R39Q >10 1.85 NT5C2 R39Q 5.45 6.82
NT5C2 R238W 7.05 >10 NT5C2 R238W 2.41 3.98
NT5C2 K359Q 6.47 >10 NT5C2 K359Q >10 3.37
NT5C2 S360P 5.25 1.44 NT5C2 S360P 1.55 1.09
NT5C2 R367Q 5.57 >10 NT5C2 R367Q >10 2.60
NT5C2 L375F 4.57 >10 NT5C2 L375F 2.04 2.14
NT5C2 D407A 5.67 >10 NT5C2 D407A 10.00 2.56
NT5C2 S408R 8.02 1.88 NT5C2 S408R 3.10 6.22
NT5C2 S445F 6.76 2.13 NT5C2 S445F 2.07 1.83
NT5C2 R478S >10 1.56 NT5C2 R478S 4.51 5.21
NT5C2 Q523X 4.18 1.39 NT5C2 Q523X 1.36 0.57
*RFP, empty vector control #NT5C2 WT, wild type NT5C2
159 Appendix
II. Chapter 4
Supplementary Tables
Supplementary Table A4.1. Nucleoside analog IC50 values (μM) of isogenic Nt5c2 wildtype and Nt5c2 R367Q primary murine T-ALL tumors.
6-mercaptopurine 6-thioguanine Mouse Tumor Nt5c2wt/wt Nt5c2wt/R367Q Nt5c2wt/wt Nt5c2wt/R367Q T-ALL 1 0.85 251.30 0.36 1.11 T-ALL 2 0.19 36.30 0.12 0.89 T-ALL 3 0.51 23.34 0.21 1.08
Supplementary Table A4.2. Results of Leukemia Initiating Cell assay performed by limiting dilution transplantations of isogenic Nt5c2 wildtype and Nt5c2 R367Q primary mouse T-ALL tumor cells.
160 Appendix
III. Chapter 5
Supplementary Figures
Supplementary Figure A5.1. C-terminally truncated recombinant NT5C2 proteins used in crystallographic studies retain activating phenotypes of relapse-associated NT5C2 mutations. Comparison of relative 5’-nucleotidase activity of purified full-length (residues 1-561) mutant NT5C2 proteins (black bars) and C-terminally truncated (residues 1-536) mutant NT5C2 proteins (537X, red bars) indicates that the C-terminus is not required for the activating phenotype of relapse-associated NT5C2 mutations. WT, wildtype.
Supplementary Tables
Supplementary Table A5.1. Summary of crystallographic information for NT5C2 537X proteins.
Protein WT R367Q K359Q L375F R238W
ATP+Mg2+ Ligands P IMP+P P P i i i i +IMP Space group C2 C2 I222 I222 I222 a=144.5, a=182.4, a=91.5, a=92.2, a=92.7, Cell parameters b=124.1, b=91.9, b=126.8, b=127.1, b=127.3, in (Å) and (°) c=90.7, c=127.8, c=130.2 c=130.6 c=131.1 β=116.2 β=134.4 Maximum resolution (Å) 2.8 2.34 1.8 2.9 1.9 Number of observations 99,269 219,723 919,574 119,045 585,022 a Rmerge (%) 12.0 (56.7) 6.0 (40.1) 7.4 (51.0) 9.6 (39.6) 7.7 (53.2) I/σI 8.8 (1.9) 16.8 (2.5) 44.1 (2.7) 23.9 (4.4) 31.0 (2.2) Resolution range used 50.0-2.8 50.0-2.34 50.0-1.8 50.0-2.9 50.0-1.90 for refinement Number of reflectionsb 30,955 59,892 66,693 15,106 Completeness (%) 88.3 (79.5) 93.9 (84.4) 94.9 (85.3) 86.9 (78.2) R factor (%)c 23.8 (30.3) 20.3 (25.3) 19.5 (24.5) 23.6 (33.4) 19.6 (25.4) Free R factor (%) 28.0 (35.8) 24.0 (29.8) 23.1 (28.9) 28.7 (39.7) 23.5 (29.7) A&B:3-405, A:3-401, 416- A:3-401, 416- Model A&B:26-492 A:3-488 414-488 488 488 rms deviation in bond 0.008 0.007 0.006 0.008 0.006 lengths (Å) rms deviation in bond 1.4 1.2 1.2 1.4 1.3 angles (°)
161 Appendix
a . The numbers in parentheses are for the highest resolution shell. Rmerge Ihi Ih Ihi h i h i b The number for the selenomethionyl protein includes both Friedel pairs. c o c o . R Fh Fh Fh h h
Supplementary Table A5.2. Summary of crystallographic information for full length NT5C2 proteins.
Protein WT R367Q R39Q
Ligands Pi Pi Pi Space group P212121 C2221 C2221 a=97.1, a=121.8, a=120.8, Cell parameters b=121.9, b=171.4, b=171.0, in (Å) and (°) c=191.8 c=122.4 c=121.9
Maximum resolution (Å) 3.35 2.32 2.9 Number of observations 470,823 687,855 422,770 a Rmerge (%) 17.3 (96.6) 14.2 (81.0) 17.4 (91.0) I/σI 7.5 (1.9) 19.9 (2.6) 7.5 (0.9) Resolution range used 50.0-3.35 50.0-2.32 50.0-2.9 for refinement Number of reflectionsb 29,069 52,756 25,428 Completeness (%) 88.3 (79.5) 94.8 (85.2) 89.7 (80.8) R factor (%)c 25.4 (33.3) 21.5 (25.8) 21.9 (28.5) Free R factor (%) 31.0 (40.3) 25.9 (31.8) 29.3 (38.9) A&B:24-509, A&B:24-511, Model A-D:26-492 537-557 537-553 rms deviation in bond 0.009 0.006 0.008 lengths (Å) rms deviation in bond 1.4 1.3 1.4 angles (°)
a . The numbers in parentheses are for the highest resolution shell. Rmerge Ihi Ih Ihi h i h i b The number for the selenomethionyl protein includes both Friedel pairs. c o c o . R Fh Fh Fh h h
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