THE EFFECTS OF PURINE NUCLEOSIDE
PHOSPHORYLASE (PNP) DEFICIENCY ON
THYMOCYTE DEVELOPMENT
by
Taniya Papinazath
A thesis submitted in conformity with the requirements
for the degree of Master of Science
Institute of Medical Science
University of Toronto
© Copyright by Taniya Papinazath 2010
ii
Abstract
THE EFFECTS OF PURINE NUCLEOSIDE PHOSPHORYLASE (PNP)
DEFICIENCY ON THYMOCYTE DEVELOPMENT
M.Sc Thesis, 2010
Institute of Medical Science, University of Toronto
© Copyright by Taniya Papinazath 2010
PNP is a crucial enzyme in purine metabolism, and its inherited defects result in severe
T-lineage immune deficiency in humans. I hypothesized that PNP deficiency disrupts the
development of late CD4-CD8- double negative (DN) thymocytes and induces mitochondrial-
mediated apoptosis of CD4+CD8+ double positive (DP) thymocytes. By using PNP-deficient
(PNP-/-) mice as well as an OP9-DL1 co-culture system simulating PNP-deficient conditions, I
demonstrated that PNP deficiency interferes with the maturation of DN thymocytes at the
transition from DN3 to DN4 stage. Although PNP deficiency does not affect the generation or
proliferation of DP thymocytes, PNP-/- DP thymocytes were observed to undergo apoptosis at a
higher rate. My results suggest that apoptosis is induced through a mitochondrial mediated
pathway. Additionally, re-introduction of PNP into PNP-/- thymocytes protected the cells from
the toxic effects of deoxyguanosine by preventing the formation of deoxyguanosine
triphosphate, indicating that the toxic metabolite in PNP deficiency is deoxyguanosine.
iii
Acknowledgements
First and foremost, I would like to thank my supervisor, Dr Eyal Grunebaum for his
wonderful supervision, guidance and mentoring. In the past two years under his supervision,
he has been encouraging and patient in both my scientific and artistic pursuits. Also, thank
you for allowing me to explore without inhibition and learn from my mistakes, I am truly
grateful for this experience.
I would like to extend my appreciation to the members of my program advisory
committee Dr.Yigal Dror, Dr. Cynthia Guidos and Dr. Chetankumar Tailor. I thank you all
for sitting through the numerous PAC meetings listening to me talk time and time again
about how PNP deficiency causes T-cell immune deficiency, and for giving me valuable
suggestions, without which the successful completion of this masters thesis would not have
been possible.
A special thank goes out to the entire Grunebaum lab. Weixian Min for helping with
all the animal handling, Yongmao Yu for giving timely input on flow data, Alka Arora for
making me feel at home when I first joined the lab, and Alireza Mansouri for bringing an
element of humour and joy into a sometimes frustrating scientific environment (and not to
forget all the entertaining coffee breaks).
I would also like to acknowledge the members of the Dror lab; for being patient in
answering all my western blot queries, for allowing me to borrow the cell separation magnets
God knows how many times, and for hearing me out in times of despair. Thank you all for
making me feel like an “honorary” member of your lab, inviting me for lab luncheons and
other lab celebrations.
My thanks is also extended to members of the PMH flow facility, Casey and Francis
for helping me get started in the world of flow cytometry.
My graduate life would be nothing without all the friends I made along the way. You
are far too many to list but thank you for your camaraderie and friendship. A special shout
out goes to Geneve Awong for clearing my OP9-DL1 co-culture queries.
Last but definitely not the least; I’d like to thank my loving family for all their
support and encouragement, giving me the strength to overcome all obstacles faced during
my graduate studies.
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Table of Contents
Abstract ............................................................................................................................... ii Acknowledgements ............................................................................................................ iii Table of Contents ............................................................................................................... iv List of Figures .................................................................................................................... vi List of Abbreviations ....................................................................................................... viii CHAPTER 1: INTRODUCTION ....................................................................................... 1
1.1. PNP Deficiency .................................................................................................... 2 Clinical Aspects .............................................................................................................. 4 Human PNP structure, chemistry, genome and mutations ............................................. 5 Purine Metabolism .......................................................................................................... 6 Pathophysiology .............................................................................................................. 7 Proposed mechanisms of the cellular immunodeficiency in PNP deficiency ................. 7 PNP as a target enzyme for chemotherapeutic applications ......................................... 10 Treatment Options ........................................................................................................ 11 TAT-PNP ...................................................................................................................... 12 Study Model .................................................................................................................. 13 1.2. Murine Thymocyte Development ...................................................................... 14 Introduction ................................................................................................................... 14 T-Cell Receptor (TCR) ................................................................................................. 15 Thymic Organization .................................................................................................... 16 Murine Thymocyte Development ................................................................................. 18 Notch Signaling ............................................................................................................ 22 OP9-DL1 ....................................................................................................................... 23 IL-7 and Flt3L ............................................................................................................... 24 Thymocyte abnormalities in PNP deficiency ............................................................... 25 Effect of PNP deficiency on T-lymphocytes ................................................................ 25 Selective dGuo toxicity to thymocytes and T-lymphocytes ......................................... 26 Thymocyte apoptosis associated with purine deficiencies ........................................... 27 • Insights gained from ADA deficiency ............................................................... 27 • Insights gained from PNP deficiency ................................................................ 29
CHAPTER 2: RATIONALE, HYPOTHESIS AND EXPERIMENTAL OBJECTIVES 33
2.1. Rationale ............................................................................................................ 34 2.2. Hypotheses ......................................................................................................... 39 2.3. Specific Objectives ............................................................................................ 39
CHAPTER 3: MATERIALS AND METHODS .............................................................. 40
3.1. Animals .............................................................................................................. 41 3.2. PNP enzyme activity .......................................................................................... 41 3.3. Cell culture and media ....................................................................................... 41 3.4. Antibodies and reagents ..................................................................................... 42 3.5 Flow cytometry .................................................................................................. 43
v
3.6. Bone marrow (BM) cell isolation ...................................................................... 44 3.7. OP9-DL1 co-cultures ......................................................................................... 44 3.8. Thymocyte isolation ........................................................................................... 45 3.9. BrdU proliferation assay .................................................................................... 46 3.10. Analysis of DN thymocytes ............................................................................... 46 3.11. Apoptosis studies ............................................................................................... 48 3.12. Mitochondrial membrane potential (MMP) ....................................................... 49 3.13. Re-introduction of PNP into PNP-/- cells ........................................................... 49 3.14. Viable cell counting ........................................................................................... 50 3.15. Statistical analysis .............................................................................................. 50
CHAPTER 4: RESULTS .................................................................................................. 51
4.1. Abnormal lymphocyte subpopulation in thymus of PNP-/- mice. ...................... 52
Aim 1: To determine whether PNP deficiency interferes with the transition of thymocytes from the DN3 to the DN4 stage ................................................................. 54 4.2. PNP deficiency disrupts thymocyte maturation at the DN3 to DN4 stage ........ 54 transition. ........................................................................................................... 54 Conclusion .................................................................................................................... 65
Aim 2: To determine whether the reduced number of DP thymocytes in PNP deficiency is due to increased apoptosis caused by dGuo ............................................ 66 4.3. Increased apoptosis in freshly isolated PNP-/- DP thymocytes. ......................... 66 4.4. PNP Deficiency does not affect proliferation or the maturation of DP thymocytes. ................................................................................................................... 68 4.5. dGuo causes increased thymocyte apoptosis. .................................................... 71 Direct effects of dGuo phosphorylation product dGMP on thymocyte apoptosis. ....... 72 Re-introduction of PNP into the cells increases the survival of PNP-/- thymocytes. .... 74 4.6. dGuo preferentially causes the apoptosis of PNP-/- DP thymocytes. ................. 76 Conclusion .................................................................................................................... 79
Aim 3: To examine whether the increased apoptosis of DP thymocytes in PNP deficiency is initiated in the mitochondria .................................................................... 80 4.7. Fas expression is not upregulated in PNP deficiency. ....................................... 80 4.8. Dissipation of mitochondrial membrane potential (MMP). ............................... 82 4.9. Changes in MMP occur before nuclear DNA fragmentation: ........................... 84 4.10. Caspase inhibition prevents dGuo induced DNA fragmentation but does not .. 87 prevent apoptosis of the cells. ............................................................................ 87 Conclusion .................................................................................................................... 89
CHAPTER 5: DISCUSSION ............................................................................................ 90
Future Directions .......................................................................................................... 99
References ................................................................................................................... 101
vi
List of Figures
Figure 1 Schematic representation of the role of PNP in purine metabolism, salvage of purines from ribo and deoxyribonucleosides.
3
Figure 2 Metabolic effects of PNP deficiency. 9
Figure 3 Structure of BCX-1777. 10
Figure 4 Traffic of thymocytes through the thymus. 17
Figure 5 Schematic representation of murine thymocyte development. 21
Figure 6 A schematic representation of postulated apoptotic mechanisms caused by ADA and PNP deficiencies. 32
Figure 7 Stages in murine thymoctye development. 35
Figure 8 Schematic representation of the PNP pathway. 37
Figure 9 Proposed mechanism of PNP deficiency. 38
Figure 10 Distribution of thymocyte populations in PNP-/- mice. 53
Figure 11 Characterization of maturation of DN thymocytes in PNP- deficient mice. 55
Figure 12 Characterization of maturation of DN thymocytes under PNP- deficient conditions using OP9-DL1 co-culture. 57,58
Figure 13 Characterization of maturation of DN thymocytes under PNP- deficient conditions using excess dGuo. 60,61
Figure 14 Characterization of DN3 and DN4 subsets by upregulation of CD71 expression. 63
vii
Figure 15 Characterization of the DN3 and DN4 subsets by intracellular TCRβ staining. 64
Figure 16 Increased apoptosis in freshly isolated DP thymocytes from PNP-deficient mice. 67
Figure 17 BrdU labeling profiles for PNP-/- mice show normal thymocyte proliferation but indicate a survival defect most strikingly obvious on day 5 after labeling.
69,70
Figure 18 Schematic representation of the role of PNP in purine degradation and salvage. 71
Figure 19 Demonstration of apoptosis of thymocytes after incubation in vitro with different purine metabolites. 73
Figure 20 Introduction of PNP into PNP-deficient cultures increases the rate of thymocyte survival. 75
Figure 21 PNP-/- DP thymocytes undergo increased dGuo-induced apoptosis. 77,78
Figure 22 Fas expression in subpopulations of PNP+/+ and PNP-/- thymocytes. 81
Figure 23 Dissipation of MMP. 83
Figure 24 Effects of dGuo on MMP and DNA fragmentation after 12 hours and 16 hours of treatment. 85,86
Figure 25 Effects of pan-caspase inhibition on dGuo-induced apoptosis and nuclear DNA fragmentation. 88
viii
List of Abbreviations
ADA Adenosine deaminase Ado Adenosine APC Allophycocyanin α Alpha α-MEM Alpha- minimum essential medium β Beta BM Bone marrow BrdU Bromodeoxyuridine CD Cluster of differentiation dAdo Deoxyadenosine dATP Deoxyadenosine triphosphate dCK Deoxycytidine kinase dGuo Deoxyguanosine dGMP Deoxyguanosine monophosphate dGDP Deoxyguanosine diphosphate dGTP Deoxyguanosine triphosphate dGK Deoxyguanosine kinase dIno Deoxyinosine DMSO Dimethyl sulphoxide DN Double negative (CD4-CD8-) DNA Deoxyribonucleic acid dNTP Deoxynucleotide triphosphate Dex Dexamethasone DP Double positive (CD4+CD8+) FACS Fluorescence activated cell sorter FBS Fetal bovine serum FITC Fluorescein isothiocyanate FTOC Fetal thymic organ culture Gua Guanine Guo Guanosine GTP Guanosine triphosphate HGPRT Hypoxathine guanine phosphoribosyl transferase HSC Hematopoietic stem cells Hx Hypoxanthine IL Interleukin Ino Inosine
ix
Lck Lymphocyte-specific protein tyrosine kinase Lin- Lineage negative MACS Magnetic activated cell sorter MHC Major histocompatibility complex μl Microlitre ml Milliliter μM Micromolar MLR Mixed lymphocyte reaction MMP Mitochondrial membrane potential
MNGIE Mitochondrial neurogastrointestinal encephalomyopathy
mtDNA Mitochondrial DNA NaCl Sodium chloride NaOH Sodium hydroxide OP9-DL1 OP9 Delta like- 1 PBS Phosphate buffered saline PE Phycoerythrin PEG Polyethylene glycol PI Propidium iodide PNP Purine nucleoside phosphorylase PNP-/- PNP-deficient pTα Pre-T-alpha PTD Protein transduction domain RAG Recombination activating gene RBC Red blood cell SCID Severe combined immunodeficieny syndrome SP Single positive (either CD4+ or CD8+) T-ALL T cell-acute lymphoblastic leukemia TAT Transactivator of transcription TCR T cell receptor UT Untreated
1
CHAPTER 1: INTRODUCTION
2
1.1. PNP Deficiency
Purine nucleoside phosphorylase (PNP) is a ubiquitous enzyme essential for the degradation
of purine nucleosides; guanosine (Guo), deoxyguanosine (dGuo), inosine (Ino) and
deoxyinosine (dIno) into uric acid, or their salvage into nucleic acids (Figure 1). An inherited
deficiency in PNP causes severe T-cell immune dysfunction in patients, contributing to
increased susceptibility to infections, failure to thrive and death in the first few years of life.
Some patients manifest with autoimmunity and malignancy (Markert, 1991; Cohen et al.,
2000). Few patients also exhibit neurological abnormalities. The fundamental role of PNP
in the functioning of organisms makes it vital to research both the enzyme and the
compounds affecting its activity.
History:
The significance of the purine salvage pathway in the maintenance and function of the
immune system became evident by the discovery that genetic deficiency in the purine
salvage enzyme adenosine deaminase (ADA) causes severe combined immunodeficiency
syndrome (SCID) in infants. Further screening of immunodeficient patients led to the
discovery that deficiency in the next enzyme in the purine salvage pathway, PNP, resulted in
severe T-cell immune deficiency (Giblett et al., 1975), emphasizing the importance of the
purine salvage pathway in lymphoid cells. PNP deficiency is inherited as an autosomal
recessive disorder and is less frequent than ADA deficiency. It has been estimated that 4%
of SCID are due to PNP deficiency (Markert, 1991) and to date more than 50 families have
been reported worldwide with PNP deficiency. However, the wide spectrum of clinical
3
manifestations associated with this disorder suggests that many cases have not been
documented.
dAdoADAADA
dATP
HGPRT
Ado
ATP
dIno, Ino, Guo, dGuo Gua and Hx+ R-1-P+ dR-1-P
dGTP GTP
dGK
PNPPNP
Nucleic Acid
Nucleic Acid
Degradation of purinenucleotides
Guase
GMP
GDP
dGMP
dGDP
dAMP
dADP
Xox
Xan
Xox
Uric acid
ADP
AMP
dAdoADAADA
dATP
HGPRT
Ado
ATP
dIno, Ino, Guo, dGuo Gua and Hx+ R-1-P+ dR-1-P
dGTP GTP
dGK
PNPPNP
Nucleic Acid
Nucleic Acid
Degradation of purinenucleotides
Guase
GMP
GDP
dGMP
dGDP
dAMP
dADP
Xox
Xan
Xox
Uric acid
dAdoADAADA
dATP
HGPRT
Ado
ATP
dIno, Ino, Guo, dGuo Gua and Hx+ R-1-P+ dR-1-P
dGTP GTP
dGK
PNPPNP
Nucleic Acid
Nucleic Acid
Degradation of purinenucleotides
Guase
GMP
GDP
dGMP
dGDP
dAMP
dADP
Xox
Xan
Xox
Uric acid
ADP
AMP
Figure 1: Schematic representation of the role of PNP in purine metabolism, salvage of
purines from ribo- and deoxyribonucleosides (adapted from Bzowska et al., 2000). PNP
catalyzes the phosphorolysis of the products of the ADA reaction, Ino and dIno, to yield Hx
and either R-1-P or dR-1-P. PNP also phosphorolyses Guo and dGuo to Gua and the
respective ribose phosphates. The purine bases, Hx and Gua, may then be recycled by the
action of hypoxanthine guanine phosphoribosyl transferase (HGPRT) to inosine
monophosphate (IMP) and guanosine monophosphate (GMP), which can undergo further
phosphorylation to guanosine triphosphate (GTP) which is salvaged back to the guanine
nucleotide pool. Alternatively, the formed Gua and Hx can be acted upon by guanase
(Guase) and xanthine oxidase (Xox) respectively to get converted to xanthine (Xan) and
finally to uric acid and excreted in the urine.
4
Clinical Aspects:
Immunological abnormalities: The most characteristic immune abnormality in PNP-
deficient patients is a profound T-cell defect which makes them susceptible to various
pathogenic infections within the first two years of their life.
Viral infections including cytomegalovirus, parainfluenza type 3, herpesviruses,
Epstein Barr virus have been documented. Persistent Varicella zoster infections which are
difficult to treat have also been observed (Baguette et al., 2002). Pathogenic bacteria such as
Haemophilus influenzae, Pseudomonus and Streptococcus pneumoniae have been
documented, as well as certain opportunistic infections caused by Candida albicans and P.
jiroveci.
On chest x-rays, the thymus is absent and the tonsils are either absent or reduced in
size (Staal et al., 1980). Immunological evaluation has revealed T-lymphopenia and
markedly decreased CD3+ T-cells (often less than 500 cells/ml or less than 15% of total
lymphocytes), which further decreases with age. T-cell function is similarly reduced as
assessed by response to various mitogens and by the mixed lymphocyte reaction (MLR).
PNP deficiency is associated with an increased risk of autoimmune disorders. The
most commonly reported autoimmune occurrences are autoimmune hemolytic anemia,
idiopathic thrombocytopenic purpura and systemic lupus erythematosis. The abnormal
immune surveillance in PNP-deficient patients also predisposes them to develop
malignancies such as lymphoma and lymphosarcoma.
Non Immunological abnormalities: Patients present with mild hepatomegaly and
splenomegaly. Many PNP-deficient patients also suffer from neurological dysfunction
unrelated to the immunological abnormalities (Market, 1991). These neurological
5
imbalances include ataxia, developmental delay and spasticity, possibly reflecting the
importance of purine homeostasis in brain development, as demonstrated in other diseases
with abnormal purine metabolism such as Lesch-Nyhan syndrome (HGPRT deficiency) and
ADA deficiency (Pelled et al., 1999; Tezcan et al., 1995).
Another example of the role of purine metabolites in other tissues includes the effect
of PNP deficiency on bone marrow. PNP-deficient marrow has been found to be
hypersensitive to irradiation and many patients develop dysplastic morphology (Dror et al.,
2004).
Biochemical findings: Most PNP-deficient patients have undetectable levels of PNP
activity. In addition, the deficiency is strongly indicated by low serum and urinary levels of
uric acid, which is the final breakdown product in the PNP pathway. High serum levels of
PNP substrates; inosine (Ino) and guanosine (Guo) and high urinary levels of Ino,
deoxyinosine (dIno), Guo and deoxyguanosine (dGuo) are also often found (Hershfields and
Mitchell, 2001). The diagnosis of PNP deficiency is established when PNP activity is
undetectable and is confirmed by molecular analysis of the PNP gene.
Human PNP structure, chemistry, genome and mutations:
PNP has been isolated from a wide variety of both mammalian and bacterial species
(Pugmire and Ealick, 2002). PNP is a ubiquitous enzyme and has been purified from many
different tissues (Agarwal et al., 1975; Carson et al., 1977). In humans, the highest activity is
found in the kidney, peripheral lymphocytes, and granulocytes. The enzyme exists in the
trimeric form in which each subunit of the functionally active trimer has a molecular weight
of ~ 32 kDa (Zannis et al., 1978; Osbourne, 1980; Stoeckler et al., 1978; Goddard et al.,
1983). Mutant enzymes can be completely inactive or may exhibit residual activity. The
6
enzyme has been crystallized from erythrocytes, and studies of the active site have shown
specificity for binding of guanine (Gua) and hypoxanthine (Hx) and their analogues but not
adenine (Ealick et al., 1990).
The human PNP gene has been cloned, sequenced and mapped to chromosome 14q13
and spans 9kb of genomic DNA (Ricciuti and Ruddle, 1973; Williams et al., 1984). Over
two dozen mutations have been identified; the bulk are missense mutations. The most
common appeared to be the point mutation Arg234Pro (Markert et al, 1996), but many
mutations have been detected since. Residual PNP activity has been observed to cause a
milder phenotype (Broome et al., 1996; Markert et al., 1997) but the diversity of the
mutations and the small number of patients makes it difficult for genotype-phenotype
correlations.
Purine Metabolism:
Purine nucleotides are synthesized by the de novo or the salvage pathway. The de novo
pathway creates phosphorlayted ring structures from simple precursors such as CO2,
glycine and glutamine. The salvage pathways reutilize purine and pyrimidine bases.
Inosinic acid (IMP, inosine monophosphate) is the central product in both pathways in
purine metabolism and is central to the interconversion of adenine and guanine
nucleotides (Nyhan, 2005). These ubiquitous purine metabolic pathways are responsible
for the proper balance between the production of dephosphorylated purines,
detoxification by further degradation to uric acid, and salvage by metabolism back to the
nucleotide level.
7
Pathophysiology:
PNP (EC 2.4.2.1) catalyses the reversible phosphorolysis of four purine nucleosides
guanosine (Guo), deoxyguanosine (dGuo), inosine (Ino) and deoxyinosine (dIno) to yield
guanine (Gua) and hypoxanthine (Hx) respectively, and ribose-1-phosphate (R-1-P) or 2’-
deoxy ribose-1-phosphate (dR-1-P) (Figure 1). Specifically, PNP catalyzes the cleavage of
the glycosidic bond of ribo and deoxyribonucleosides in the presence of ortho inorganic
phosphate (Pi) in order to generate the purine base and R(dR)-1-P (Bzowska et al., 2000),
schematized in the following reaction:
(2’-deoxy) nucleoside + Pi ↔ base + (2’-deoxy) ribose 1-phosphate
Proposed mechanisms of the cellular immunodeficiency in PNP deficiency:
The central feature of this disorder is profound impairment in cellular immunity
characterized mainly by the ability of T-cells to accumulate dGTP. Several mechanisms
have been proposed to explain the effects of PNP deficiency, but most researchers have
focused on the accumulation of PNP substrates and their respective metabolic products
(Hershfield and Mitchell, 2001). There has been extensive evidence pertaining to T-cells,
showing that dGuo interferes with DNA synthesis via its phosphorylated product dGTP.
dGTP is thought to cause feedback inhibition of ribonucleotide reductase (Carson et al.,
1977; Mitchell et al., 1978; Chan, 1978; Ullmann et al., 1979; Kredich and Hershfield, 1989),
which inturn causes depletion of intracellular dNTP pools resulting in inhibition of DNA
synthesis (Reichard, 1978; Martin and Gelfand, 1981) (Figure 2). Arpaia et al further
suggested a mitochondrial basis for immune deficiency (Arpaia et al., 2000). They postulated
that, similar to the effects of dATP in ADA deficiency, the abnormalities observed in PNP
deficiency are due to the selective accumulation of dGTP in the mitochondria of T-cells,
8
leading to abnormal mitochondrial DNA (mtDNA) synthesis which in turn impedes cell
division resulting in apoptosis. A more detailed explanation of the specific effects of dGuo
on thymocytes/T-cells will be discussed in the thymocyte section of this introduction.
Although T-cells are the most severely affected in PNP deficiency, abnormalities in
other tissues have also been observed but not much attention has been paid on the specific
effects of PNP deficiency on brain and other tissues. However, neurological abnormalities
associated with PNP deficiency indicate a correlation with purine metabolism and normal
brain development. Although increased levels of dGTP has been implicated in the T-cell
dysfunction of this disorder, in contrast, the neurological symptoms associated with PNP
deficiency has been postulated to be caused by a depletion of GTP, similar to the effects seen
in HGPRT deficiency (Simmonds et al., 1987). Reduced GTP maybe because of the inability
to produce Gua, or secondary to over-consumption of phosphoribosyl pyrophosphate,
ultimately interfering with de novo purine synthesis (Barankiewicz et al., 1982). Decreased
intracellular levels of GTP observed in PNP deficiency is unlikely to cause the immune
dysfunction, as a similar decrease in intracellular levels of GTP associated with HGPRT
deficiency has no effect on the immune system (Sidi et al., 1989). Therefore, the two major
clinical consequences of PNP deficiency may reflect differing etiologies; neurological effects
may result from deficiency of the PNP enzyme products (GTP) while the immunodeficiency
might be caused by accumulation of the PNP enzyme substrates (dGuo, dGTP). The former
hypothesis is yet to be elucidated.
9
DNA dNTP dNDP NDP
Ribonucleotidereductase
dGuo
accumulation
dGMP
dGDP
dGTP
Inhibition
Gua
PNP deficiency or
inhibition of PNP
dGKdGK
DNA dNTP dNDP NDP
Ribonucleotidereductase
dGuo
accumulation
dGMP
dGDP
dGTP
Inhibition
Gua
PNP deficiency or
inhibition of PNP
dGKdGK
Figure 2: Metabolic Effects of PNP Deficiency (adapted from Bzowska et al., 2000). PNP
deficiency causes the accumulation of dGTP which subsequently inhibits ribonucleotide
reductase and consequently hinders DNA synthesis. Abbreviations; dGK, deoxyguanosine
kinase, NDP, nucleoside diphosphate.
10
PNP as a target enzyme for chemotherapeutic applications:
Selective T-cell immunodeficiency associated with PNP deficiency suggested that PNP can
be targeted to selectively regulate T-cell immune response. The finding of the enzyme
structure gave way to the discovery of potential PNP inhibitors for chemotherapeutic
applications. The observation that in the absence of PNP there is an accumulation of dGuo
in the plasma and an accumulation of dGTP in T-cells has been the impetus to develop a
number of PNP inhibitors and PNP resistant dGuo analogues. Over the years, studies have
shown the beneficial use of PNP inhibitors as a therapeutic agent for treating
lymphoproliferative malignancies (Furman et al., 2007), certain autoimmune disorders such
as rheumatoid arthritis, psoriasis (Morris and Omura, 2000) and to suppress graft vs host
disease (Bantia et al., 2002). One such effective inhibitor of PNP is BCX-1777
(Forodesine/immucillin-H) (Miles et al., 1998; Bantia et al., 2001) (Figure 3), which was the
inhibitor used over the course of this study.
Figure 3: Structure of BCX-1777 (Bantia et al., 2003). BCX-1777 is a potent inhibitor of
human PNP and designed based on the transition state structure.
Previous studies have shown that the cytotoxicity of BCX-1777 in the presence of dGuo is
specific to T-cells without affecting other non T-cell tumor lines (Kiscka et al., 2001). This
11
lead to a phase 1 clinical trial of BCX-1777 in patients with advanced T-cell malignancies
(Balakrishnan et al., 2006). BCX-1777 in the presence of dGuo inhibited the proliferation of
CEM-SS (a T-acute lymphoblastic leukemia (T-ALL)) cells accompanied by a 154 fold
increase in endogenous dGTP. BCX-1777 also inhibited the proliferation of activated human
T-cell lymphocytes induced with various agents such as IL-2, MLR and phytohemagglutinin
(Bantia et al., 2003).
Individuals with genetic deficiency in PNP maintaining even 5% of the catalytic
activity of the enzyme fail to accumulate dGuo and exhibit T-cell immune deficiency.
Therefore it is essential to completely inhibit PNP (>95%) to achieve cytotoxicity of T-cells
due to increase in dGuo (Morris et al., 1998). The necessity for near complete PNP
inhibition makes BCX-1777 an ideal pharmacological inhibitor to study PNP deficiency as it
binds ≈1000 times more tightly to PNP than previously studied inhibitors (Bantia et al.,
2001).
Treatment Options:
PNP-deficient patients usually succumb to this disorder within the first two decades of life
due to opportunistic infections, autoimmune hematological disturbances or malignancy. As
of the time being, hematopoietic stem cell (HSC) transplant is considered the treatment of
choice. Although it has been able to provide long term immune reconstitution in a few
patients, the rarity of this disorder makes it imperative to have a definitive treatment option.
Hence, research is still underway to develop a treatment modality to better manage this
disorder.
Some of the previous attempts to alleviate symptoms associated with PNP deficiency
include; erythrocyte transfusions as a means of PNP enzyme replacement. This therapeutic
12
approach caused partial reconstitution of T-cell mediated immunity, however, it did not
protect from devastating infections (Rich et al., 1980; Sakiyama et al., 1989). This gave way
to a polyethylene glygol (PEG) bound enzyme replacement therapy.
After the relative success of PEG-ADA in the treatment of ADA deficiency (Markert
et al., 1987; Hershfield et al., 1991), PEG-PNP injections were tested on PNP-/- mice
(Roifman –personal communication). Data suggesting that PEG-ADA provided only partial
and temporary immune reconstitution in ADA-deficiency (Chan et al., 2005; Husain et al.,
2007), probably because the enzyme was not being targeted into the cell, difficulties in
producing PEG-PNP, and moreover the high cost of production, did not make this type of
treatment a favorable one. This prompted us to develop a TAT-PNP treatment, an alternative
enzyme replacement therapy that targeted PNP into the cells.
TAT-PNP: TAT-PNP treatment will be highlighted in more detail as this is the
technique that was used to introduce PNP into the PNP-/- thymocytes in this study. Protein
transduction domains (PTD) are a group of peptides that can cross biological membranes in a
receptor independent manner (Wadia et al., 2003). One such PTD is the HIV– transactivator
of transcription (TAT) protein (Gump and Dowdy, 2007). Since only 11 amino acids of TAT
is required for transduction, and its relatively easy production has made it the PTD of choice
(Schwarze et al., 1999; Nagahara et al., 1998). Various in vitro studies have shown that PTD
can transport cargo molecules into many different cell types (Dietz et al., 2004; Kwon et al.,
2005). Based on this, recently, our lab generated a TAT-PNP fusion protein by replacing the
ATG of the human PNP cDNA with the coding sequence for the 11 amino acids of the TAT-
PTD (Toro et al., 2006). Repeated TAT-PNP injections were able to correct thymocyte
development and T-lymphocyte function in PNP-deficient mice. PNP activity was detected
13
in all the organs tested and also increased survival of these mice (Toro and Grunebaum,
2006). Further, pre-clinical studies are currently being conducted to establish the benefit of
PTD-PNP for PNP-deficient patients.
More recently our lab investigated the possibility of gene therapy using a lentivirus
based vector that expressed human PNP. The preliminary studies carried out in PNP-/- mice
have been promising (Liao et al., 2008). Currently our lab is optimizing conditions to ensure
long term gene expression which can translate into better gene therapy for PNP-deficient
patients.
Study Model:
PNP Knockout mice:
There are no known naturally occurring models of PNP deficiency. This led to the
development of two relevant mouse models recapitulating the human disorder. The first
model by Snyder et al was developed by introducing three point mutations in the PNP gene
locus by male germ cell mutagenesis (Snyder et al., 1997). In their mouse model,
progressive loss of thymic cellularity due to reduction of CD4+CD8+DP thymocytes was
observed, with corresponding reduction in peripheral T-lymphocyte numbers and function.
Although these mice shared some similarities with the human phenotype, the residual levels
of PNP activity, the lack of dGTP accumulation in thymocytes, the delayed onset of the
immune deficiency and the normal life span of the mice were not representative of PNP-
deficient patients (Snyder et al., 1997). This prompted another group led by Dr. Roifman, to
develop a PNP deficient (PNP-/-) mouse model by replacing the catalytic domain of the
murine PNP gene with a neomycin cassette, thereby creating mice with no PNP activity
(Arpaia et al., 2000). These PNP-/- mice exhibited nearly all the metabolic, immune and non-
14
immune abnormalities typical of PNP-deficient patients. Some of the important similarities
include; very early death (8-12 weeks), PNP activity in these mice was found to be <0.2% of
normal, increased concentrations of PNP substrates were detectable in the urine, uric acid
production diminished (Arpaia et al., 2000), and the mice failed to gain weight (Toro and
Grunebaum, 2006). The mice developed a hypoplastic thymus characterized by reduced
number of DP cells, markedly reduced number and function of T-cells and relatively
preserved B-cell numbers (Arpaia et al., 2000). The mice displayed autoimmuity with
splenomegaly and hepatomegaly. Mice also manifest with neurological abnormalities as
tested by their inability to stay on a spinning rotorod, which is indicative of possible ataxia
and abnormal motor coordination (Grunebaum unpublished). The above mentioned
characteristics of the PNP-/- mice generated by Dr. Roifman’s group led us to choose this
model, around which all the experiments for this study were designed.
1.2. Murine Thymocyte Development
Introduction
T-cells form an integral part of the adaptive immune system. T-lymphocytes are derived
from hematopoietic precursors that arise in the fetal liver or adult bone marrow and migrate
through the blood stream to the thymus. In the thymus, hematopoietic precursors go through
well defined developmental steps which are required for formation of mature thymocytes
characterized by a functional T-cell receptor (TCR). These mature T-lymphocytes are
released to the periphery where they can subsequently act on invading antigen and fight
infections when given the proper activation signals.
15
The thymic environment is essential for specifying the T-cell lineage fate for the
hematopoietic precursors. Once the T-cell lineage fate is adopted the thymus further
specifies whether thymocytes become either αβ or γδ TCR bearing cells and whether αβ
cells will express either CD4 or CD8 co-receptors. During each stage of thymocyte
differentiation, there are cellular controls that delegate whether the cells are in a position to
proliferate and move forward in the developmental pathway. Thus, formation of T-cell
repertoire involves stepwise fate determination in anatomically distinct thymic
microenvironments (Nitta et al., 2008).
T-Cell Receptor (TCR):
All mature thymocytes and T-cells express a T-cell recptor (TCR) on their cell
surface. The TCR is a complex of integral membrane proteins responsible for recognizing
antigens bound to major histocompatibility complexes (MHC) when presented by antigen
presenting cells such as dendritic cells, macrophages and B-cells (Mazza et al., 1998). The
TCR is a heterodimer consisting of two polypeptide chains either α and β or γ and δ. The
TCR is also associated with a co-stimulatory CD3 complex which is composed of a
γ, δ ε and ζ subunits (Frank et al., 1990). Once a specific peptide:MHC ligand binding
occurs, the cytoplasmic region of the CD3 complex transduces signals to the nucleus which
initiates a signaling cascade leading to the activation of the T-cell (Kane et al., 2000). The
TCR together with the CD3 subchains forms the TCR complex.
Signalling from the TCR complex is enhanced by the presence of co-receptors CD4
or CD8 (Janeway, 1992). The co-receptor specfies antigen specificity as well as the
functional lineage of the T-cell. The co-receptor CD4 binds to class II MHC molecules and
receptor engagement leads to activation of the adaptive immune system. CD4+ T-cells are
16
termed “helper” T-cells as they have no direct cytotoxic or phagocytic activity and their main
function is in aiding other immune cells. CD8 co-receptor binds to class I MHC and
activation of CD8+ T-cells leads to the direct lysis of the target cell and hence are called
“cytotoxic” T-cells.
Thymic Organization:
The thymus is a bilobed organ found in the upper portion of the chest just behind the sternum
(Miller, 1961). The thymic lobes are separated by mesenchymal septae and are further
divided into the outer cortex and the inner medulla regions each characterized by specific
stromal cell types (Anderson and Jenkinson, 2001). The two regions are separated by the
cortico-medullary junction (CMJ) and the cortex is surrounded by the subcapsular zone
(SCZ) (Penit, 1988; Lind et al., 2001). At each stage of maturation, thymocytes occupy
distinct regions in the thymus. Progenitors enter at the CMJ, migrate through the cortex to the
outer SCZ during the early progenitor stages, and then move back toward the CMJ and into
the medulla (Figure 4) (Lind et al., 2001; Petrie and Zúñiga-Pflücker, 2007). The thymus
builds up a stock of T-cells before birth and early childhood and is largest at puberty, after
which it starts to involute and get replaced by fat (Scollay et al., 1980).
17
HP CD117+CD44+CD25-
CD117+CD44+CD25+
CD117-CD44-CD25+ CD117-
CD44-CD25-
CD4+CD8+
CD4
CD8
Subcapsular zone
Cortico-medullaryjunction
Medulla
+ve selection
-ve selection
Cortex
CD8CD4
Mature T cells
Notch ligand
Notch ligand
DN1
DN2
DN3DN4
DP
SP
HP CD117+CD44+CD25-
CD117+CD44+CD25+
CD117-CD44-CD25+ CD117-
CD44-CD25-
CD4+CD8+
CD4
CD8
Subcapsular zone
Cortico-medullaryjunction
Medulla
+ve selection
-ve selection
Cortex
CD8CD4
Mature T cells
Notch ligand
Notch ligand
DN1
DN2
DN3DN4
DP
SP
Figure 4: Traffic of thymocytes through the thymus. Progenitor cells from the BM travels
through the blood stream to the CMJ of the thymus. The earliest double negative (DN)
population DN1 cells strongly rely on notch signaling from the thymic epithelium to assume
a T-cell fate. DN1 cells differentiate into DN2 and relocate outwards to the subcapsular
region of the thymic cortex where they differentiate into DN3 cells. The process of
β-selection takes place here and these DN4 thymocytes proliferate as a result of pre-TCR
signaling. The cortical epithelium is responsible for the process of positive selection and the
positively selected DPs start relocating towards the medulla. SP cells enter the medulla for
negative selection and ultimately mature T-cells leave the thymus for the periphery. The
irregularly shaped cells represent the thymic epithelial cells. The dark blue cells represent
thymic stromal cells expressing notch ligand. Abbreviations: HP, hematopoietic progenitor.
18
Murine Thymocyte Development:
Lin-Sca1+CD117+ BM derived progenitors seed the thymus, where they undergo
differentiation which is broken down into distinct stages characterized by the presence of co-
receptors CD4 and CD8 (Allman et al.,2003). Cells lacking both CD4 and CD8 have been
identified as double negative (DN) thymocytes (Scollay et al., 1984; Scollay et al., 1988).
The early DN subpopulation can be identified by the expression of CD117 (c-kit) and further
classified based on the expression of adhesion molecules CD44, and the IL-2 receptor α,
CD25 (Godfrey et al., 1993) which are expressed in a stage specific manner. The sequence
of DN transition is as follows: DN1 (CD117+CD44+CD25-), DN2 (CD117+CD44+CD25+),
DN3 (CD117-CD44-CD25+), DN4 (CD117-CD44-CD25-). Thymocyte differentiation is
summarized in Figure 5.
DN1 cells move away from the site of entry into the deeper cortex as they
differentiate into DN2 cells. Progression into the DN2 stage is marked by an upregulation of
CD25 (Porrit et al., 2003). DN2 cells still retain some capacity to become natural killer
(NK), dendritic (DC) cells and macrophages (Wu et al., 1996, Schmitt et al., 2004; Bell and
Bhandoola, 2008). Migration to the outer cortex will lead to further differentiation of DN2
cells to the DN3 stage. The DN3 stage is characterized by a down regulation of CD44.
During the transition from the DN2 to the DN3 stage, recombination of the variable (V),
diversity (D) and joining (J) DNA elements of the antigen receptor genes is initiated at the
β, γ and δ loci. This process is driven by the transient expression of the VDJ recombination
activating gene (RAG)-1 and RAG-2 proteins, to give rise to unique TCRs (Fugmann et al.,
2000). As this occurs the DN3 cells become irreversibly committed to the T-cell lineage
(Ciofani et al., 2006). If the DN thymocytes successfully rearrange the γδ loci, they will
19
follow along the γδ lineage. Alternatively, if they successfully rearrange their β-locus the
cells will adopt the αβ lineage. γδ T-cells reside in the epithelial layers of tissues such as
skin, intestinal epithelium, tongue and lung where they act as the first line of defense.
However, their development and selection processes are less well characterized than the αβ
lineage (Xiong and Raulet, 2007).
Further differentiation of the αβ lineage is triggered once the TCRβ chain assembles
with the invariant pre-Tα (pTα), and CD3 signaling chains to form the pre-TCR complex
(Saint Ruf et al., 1994; von Boehmer et al., 1997). αβ T-cells make up 95% of the T-cells in
the thymus of an adult mouse. Signaling through the pre-TCR complex leads to “β-
selection” (Michie and Zúñiga-Pflücker, 2002), the first checkpoint in thymocyte
differentiation ensuring that only thymocytes that have successfully rearranged their TCRβ
chain continue to differentiate into CD4+CD8+ DP stage. Therefore, activation through the
pre-TCR cements the αβ lineage commitment (Hayday et al., 1999).
Pre-TCR signaling and the resulting process of β-selection is a crucial checkpoint in
thymocyte differentiation. The expression of the pre-TCR complex initiates a number of
events including survival and proliferation (Falk et al., 2001; Vasseur et al., 2001; Kruisbeek
et al., 2000). Many DN3 cells do not productively rearrange their TCR loci and will die by
apoptosis in the thymus. Pre-TCR signaling also results in the down regulation of CD25
expression giving rise to DN4 cells. Upregulation of CD4 and CD8 expression allows the
DN4 cells to transit into the DP stage.
DP cells continue to rearrange the TCRα genes to express an α chain. Pairing of an
α chain with the β chain results in the formation of a mature functional αβ TCR (Petrie et al.,
1993). DP thymocytes expressing TCR αβ on their cell surface are localized in the cortex.
20
At this stage, the DP cells undergo further positive and negative selection that ensures that
only self-MHC restricted and self-tolerant thymocytes will become a mature SP
immunocompetent cell that can leave the thymus for the periphery (Sebzda et al., 1999).
During positive selection, only those DP thymocytes that receive TCR signals with
ligand interaction of weak affinity will survive. The necessity for positive selection is to
ensure that the T-cells reaching the periphery are only reactive to antigen when presented in
context with self-MHC. During the positive selection process most of the cells are induced
to undergo apoptosis and only 3- 5% of DPs within the thymus survive this checkpoint and
are selected for the next selection criteria, negative selection (Ashton-Rickardt et al., 1993;
Surh and Sprent, 1994; Takahama et al., 1994). Positively selected thymocytes migrate from
the cortex to the medulla where majority of the negative selection takes place (Witt et al.,
2005).
Negative selection is a crucial step in eliminating self-reactive T-cells (Ohashi, 2003).
This selection procedure ensures that the T-cells reaching the periphery are self-tolerant,
without which undesirable autoimmunity would ensue. In contrast to positive selection,
TCRs which bind MHC peptide complexes presented by a variety of cells like macrophages
and dendritic cells, with high affinity, will be given an apoptotic signal and induced to die,
thus preventing the exit of a self-reactive T-cell to the periphery (Hoffman et al., 1992;
Sprent and Kishimoto, 2002). The divergence of the CD4+/CD8+ SP lineage is dependent on
the specificity of the TCR to either class I MHC or class II MHC. CD4 is actively repressed
in MHC-I restricted TCRs and CD8 in MHC-II restricted TCRs (Germain, 2002). Figure 5
summaries the development of murine thymocytes.
21
CD117+CD44+CD25+
CD117-CD44-CD25-
CD4+CD8+
CD8+
CD4+
Negative Selection
Pre-TCRpTα
TCRβCD3γCD3εCD3ζ
CD117-CD44-CD25+
CD117+CD44+CD25-
β selection
DN4DN1 DN2 DN3 DP
SP
SP
Positive selection
HSC
Lin-, Sca1+, CD117+, Flt3-
Bone marrowBone marrow
ThymusThymus
γδ TCR
VDJ recombination
Pre-TCRTCR
Blo
od
α gene rearrangement
CD117+CD44+CD25+
CD117-CD44-CD25-
CD4+CD8+
CD8+
CD4+
Negative Selection
Pre-TCRpTα
TCRβCD3γCD3εCD3ζ
CD117-CD44-CD25+
CD117+CD44+CD25-
β selection
DN4DN1 DN2 DN3 DP
SP
SP
Positive selection
HSC
Lin-, Sca1+, CD117+, Flt3-
Bone marrowBone marrow
ThymusThymus
γδ TCR
VDJ recombination
Pre-TCRTCR
Blo
od
α gene rearrangement
Figure 5: Schematic representation of murine thymocyte development (adapted from
Kruisbeek et al., 2000). The CD4-CD8-DN subset is characterized by the expression of cell
surface markers CD117, CD44 and CD25. The DN compartment is subdivided into distinct
stages DN1-DN4. The first developmental checkpoint, β-selection, occurs at the DN3 stage.
Only those cells that have successfully rearranged their β-locus by the process of VDJ
recombination and express the pre-TCR will develop into the DN4 stage and become
CD4+CD8+ DP cells. At the DP stage, TCR signals are activated resulting in either CD4+ or
CD8+ SP lineage fate divergence. DPs expressing mature TCRs undergo two selective steps;
positive selection and negative selection. CD4+ or CD8+ SP cells then move into circulation.
The pre-TCR components are depicted in the box. Abbreviations; HSC, hematopoietic stem
cell.
22
The process of thymocyte development requires a crosstalk between the lymphoid cells of
the thymus and the thymic stroma which constitutes the non lymphoid portion (Petrie et al.,
2001). The interplay of various thymic stromal cells creates microenvironments through
which the cells traverse in an orderly manner. The stroma surrounding thymocytes at each
stage of maturation provides signals which are crucial in directing the cell into distinct stages
of development through direct lympho-stromal interactions, various thymopoeitic cytokines
and other soluble growth factors important for T-cell repertoire selection and maturation
(Petrie and Zúñiga-Pflücker, 2007). Furthermore, recent studies have shown that
chemokines produced by thymic stromal cells are pivotal in guiding the traffic of developing
T-cells in the thymus (Takahama, 2006). Consequently, thymocyte development is not a cell
autonomous process but has several molecular players that enable the thymus to carry out its
function (Di Santo and Rodewald, 1998).
Notch Signaling:
Many pathways have been identified in recent years to be important for determining the fate
of T-lineage cells. Among these, Notch signaling has been extensively studied, as it is an
evolutionarily conserved mechanism that is important in determining different cell fates
during development (Artavanis Tsakonas et al., 1999). In mammals, the Notch
transmembrane receptor family consists of four members (Notch 1–4) which interact with
ligands of the Delta-like (DL) family (DL-1, DL-3, and DL-4) and Serrate-like ligands
(Jagged-1 and Jagged-2) (Guidos, 2002).
The Notch pathway, particularly the Notch-1 plays a vital role in the commitment
of progenitor cells to the T-lineage fate. Mice with a neonatally induced loss of Notch-1
function, or irradiated controls repopulated with Notch-1 deficient BM stem cells,
23
severely inhibited thymocyte development but not other hematopoietic lineages (Radtke
et al., 1999). Accordingly, BM progenitors expressing constitutively active form of
Notch-1 injected into mice favored the formation of DP T-cells and abolished B-cell
development in the BM of these mice (Pui et al., 1999). Mouse thymocytes have been
shown to have modulated expression of Notch-1 throughout thymocyte development
(Hasserjian et al., 1996). Although multiple Notch ligands are expressed either in the
thymus or on thymocytes themselves (Radtke et al., 2004), gain of function approaches
have clearly indicated that DL ligands are capable of promoting T-cell development in
vitro and in vivo (Visan et al., 2006). DL-1 and Jagged-1 in particular have been shown
to promote T-cell development and is expressed in the thymic cortical and medullary
epithelium (Lehar et al., 2005).
OP9-DL1:
In light of these findings, an osteopetrosis (OP) stromal cell line was stably transduced to
express the Notch ligand DL1 to form the OP9-DL1 stromal cell line (Schmitt and Zúñiga-
Pflücker, 2002). The lack of macrophage colony stimulating factor (M-CSF) production by
OP9 cells prevented formation of macrophage cells and favored generation of lymphoid
cells. Embryonic stem cells, fetal liver and bone marrow derived hematopoietic progenitors
cultured on OP9-DL1 stromal cells were induced to commit to the T-cell lineage at the
expense of B-cell lymphopoiesis (de Pooter and Zúñiga-Pflücker, 2007). This culture system
is a valuable tool to study T-cell development in vitro and was used in my studies.
24
IL-7 and Flt3L:
In addition to Notch signaling, lymphoid development is also regulated by a variety of
cytokines. One such factor is IL-7, which is secreted by the thymic stroma. IL-7 is required
mainly during early T-cell development in the thymus to protect DN2 and DN3 stages from
apoptotic cell death (Kim et al., 1998). In IL-7 and IL-7 receptor (IL-7R) knockout mice,
progression of thymocytes beyond DN2 stage is severely diminished (Peschon et al.,1994;
von Freeden et al., 1995; Maki et al.,1996), indicating that IL-7 is essential for the expansion
of the DN2 population to transition to the DN3 compartment. Indeed studies have indicated
a role of IL-7 in thymocyte survival wherein IL-7 signalling was seen to upregulate the
expression of the anti-apoptotic molecule, Bcl-2 (von Freeden et al., 1997; Akashi et al.,
1997). Complimentary to these findings, overexpression of Bcl-2 was partially able to rescue
the defects of IL-7R deficiency (Maraskovsky et al., 1997). More recently, Van De Wiele et
al clarified the role of IL-7Rα in early thymocyte development using a transgenic
background of ectopic IL-7Rα background. Their studies led them to conclude that the role
of IL-7 signaling in early thymocyte development is between the time the thymocytes are
fated to become T-cells and the β-selection stage instigated proliferative burst is initiated
(Van de Wiele et al., 2007).
Another cytokine important for thymocyte development is FMS-like tyrosine kinase 3
ligand (Flt3L). Flt3 deficient mice have also shown a moderate decrease in the number of
DN thymocytes (Mackerehtschian et al., 1995). Furthermore, Wang et al showed using the
OP9-DL1 system; the function of Flt3L in thymopoiesis is primarily to support the survival
and proliferation of progenitors (Wang et al., 2006). Interestingly, mice double deficient for
25
Flt3L and IL7R showed not only reduction in the DN1-DN4 and DP progenitors but also
mature peripheral SP T-cells, demonstrating critical and complementary roles of Flt3 and
IL-7 in early T-cell development (Sitnicka et al., 2007).
Thymocyte abnormalities in PNP deficiency:
Patients present with a small hypoplastic thymus and correspondingly reduced
T- lymphocytes in peripheral blood. T-cell receptor excision circles (TRECs) which
represent new thymic emigrants are also reduced in these patients. Concordant to the human
observations, the number of thymocytes in PNP-/- mice is reduced compared to normal
littermates (58±20 x106 cells/thymus vs 146±19 x106 cells/thymus, n=9 in each, my data).
PNP-/- mice also exhibit increased numbers of DN thymocytes and decreased numbers of DP
and SP thymocytes (Arpaia et al., 2000, my data).
Effect of PNP deficiency on T-lymphocytes:
PNP-deficient patients may exhibit normal T-lymphocyte numbers and function early on in
life but deteriorates over a course of time as there is a buildup of toxic metabolites. The
reduced response of T-cells from PNP-deficient patients to mitogens, and the increased
susceptibility of these cells to ionizing radiation (Dror et al., 2004) are indicative that PNP
deficiency, besides affecting thymocytes also directly affects peripheral T-cells. In vivo
studies have also shown depressed T-cell function.
Similar to human observations, lymphocyte function in PNP-deficient mice is
impaired as indicated by an inability to illicit an immune response in a mixed lymphocyte
reaction (MLR) (Arpaia et al., 2000). In addition, recent findings from our lab showed
increased thymidine incorporation into fresh T-lymphocytes of PNP-/- mice (Yu et al., 2009),
26
suggesting that T-lymphocyte abnormalities in PNP deficiency might result from
exaggerated in vivo activation of peripheral lymphocytes, possibly secondary to
unrecognized infections or uncontrolled systemic inflammation. Alternatively, PNP
deficiency might disrupt lymphocytes intracellular signaling, as studies have shown
abnormal cytokine production by PNP-deficient lymphocytes (Arpaia et al., 2000; Dror et al.,
2004; Toro et al., 2006).
Selective dGuo toxicity to thymocytes and T-lymphocytes:
Although there is accumulation of all 4 substrates (Guo, dGuo, Ino and dIno) in PNP
deficiency, it has been suggested that dGuo plays a central role in mediating the effects of
PNP deficiency. The possibility of dGuo being the potentially lymphotoxic metabolite in
PNP deficiency was postulated back in 1977 by Cohen et al (Cohen et al., 1977). They
found dGTP levels were ten-fold higher in PNP-deficient patients compared to normal
patients (Cohen et al., 1977). It was demonstrated that immature human thymocytes were
more sensitive to dGuo toxicity than peripheral T-lymphocytes (Cohen et al., 1980). The
selective toxicity of dGuo to thymocytes may rest in the ability of these cells to trap dGTP
due to high deoxyguanosine kinase and low nucleotidase activity (Carson et al., 1977;
Gelfand et al., 1979).
Many different in vitro models have been used to investigate the specific effects of
dGuo on T-cell immune function. Some of the earlier studies have shown that continuously
growing T-lymphoblastoid cell lines partially deficient for PNP are more sensitive to the
cytotoxic effect of dGuo than B-lymphoblastoid cell lines (Mitchell et al., 1978). Moreover,
these findings were also applicable to lymphoid cells under conditions simulating PNP
deficiency wherein dGTP formation by thymocytes increased ten-fold (Fairbanks et al.,
27
1990). In fact, it has been postulated that the accumulation of dGTP and the preferential
sensitivity of dGTP makes the T-lineage cells more prone to apoptosis in PNP deficiency.
Evidence for this stems from the observation that dGuo has the ability to inhibit T-cell
growth and in consequence, was associated with increase in intracellular dGTP (Gudas et al.,
1978; Ullman et al., 1979), suggesting that toxicity is in fact due to the formation of dGTP
and not due to the direct effects of dGuo. Furthermore, in vitro and in vivo studies carried
out on lymphocytes from patients with T-cell malignancies in which PNP-deficient
environment was simulated using chemical PNP inhibitors, demonstrated increased dGTP
accumulation and reduced proliferation of malignant cells and normal human T-cells (Bantia
et al., 2003; Conry et al., 1998; Gandhi et al., 2005).
Thymocyte apoptosis associated with purine deficiencies:
Several studies have proposed an apoptotic mechanism for the basis of immunodeficiency in
ADA and PNP deficiency (Thompson et al., 2003; Arpaia et al., 2000). Considering that
apoptosis is the fate of most thymocytes, it is not surprising that this may be the main cause
for immune deficiency associated with these disorders.
• Insights gained from ADA deficiency:
ADA is another enzyme important for purine degradation and salvage. Importantly, similar
to PNP deficiency, ADA-deficient patients suffer from severe T-cell immune deficiency that
has been attributed to the accumulation of toxic purine metabolites. Although ADA
deficiency is also associated with severe perturbations in other immune parameters, the close
similarities between ADA and PNP deficiency suggest similar mechanisms.
28
Murine fetal thymic organ cultures (FTOC) have been used to investigate how ADA
deficiency leads to a failure in thymocyte development (Thompson et al., 2000; Van De
Wiele et al., 2002). FTOC is a technique where murine fetal thymic lobes at day 15 of
gestation are cultured on cellulose ester filters sitting on gelfoam sponges at the air/media
interface for periods of 3-5 days (Jenkinson et al., 1990). FTOCs are usually treated with
dGuo to remove thymocytes and leave intact stroma so as to allow the seeded cells (either
thymocytes or bone marrow progenitors) to repopulate the thymic niche. FTOCs performed
on ADA- deficient fetuses or on normal fetuses treated with a potent ADA inhibitor revealed
profound arrest of development at the DN3 stage (Thompson et al., 2000). This was
associated with an increase in dATP levels suggesting that dATP is the toxic metabolite in
these cultures. Moreover, the thymocytes could be rescued by using an inhibitor of
adenosine kinase, an enzyme that phosphorylates dAdo to dAMP (Jenuth et al., 1996). This
observed inhibition in differentiation was not due to a failure in β-selection as TCRβ and T-
early α locus rearrangement occurred normally (Thompson et al., 2000). Rather, the cells in
culture were found to be dying by apoptosis as the cultures could be rescued by inhibiting
apoptotic pathways with the pan caspase inhibitor z-VADfmk (Thompson et al., 2000). This
rescue was associated with an increase in the survival of cells at the β-selection checkpoint
and normalization of dATP levels (Thompson et al., 2000). Similarly, the cultures were also
rescued by Bcl-2 over-expression (Thompson et al., 2003). The large number of thymocytes
that die coupled with the high ability of the thymus to phosphorylate dAdo and low capacity
to dephosphorylate dATP explains the specificity of this organ to accumulate dATP and its
high sensitivity to ADA deficiency.
29
Initial studies concentrated on the effect of ADA deficiency on development of DN
thymocytes, recently there is evidence that ADA deficiency also affects DP thymocytes (Van
De Wiele et al., 2006). The above results have been extrapolated in PNP deficiency research
(discussed below) and have provided a basis for my study.
• Insights gained from PNP deficiency:
The specific T-lineage abnormalities characteristic of PNP deficiency have been attributed to
the accumulation of dGuo and increased sensitivity to dGTP. Similar to ADA deficiency,
studies using PNP-/- mice demonstrated that thymocyte differentiation is severely affected at
the DP stage, with possible increased apoptosis of DP thymocytes, although the exact stage
of developmental abnormality had still not been elucidated (Arpaia et al., 2000; Snyder et al.,
1997). However, Arpaia et al provided some insight on the mechanism causing this T-cell
phenotype. They indeed pinpointed apoptosis as the cause, by demonstrating that Bcl-2 over
expression protected against dGuo induced apoptosis in normal murine thymocytes treated
with a PNP inhibitor (Arpaia et al., 2000). Further exacerbation of thymocyte damage may
result from locally increased toxic purine metabolites released from dying thymocytes.
These findings led them to investigate the possibility that mitochondrial dGTP accumulation
may predispose PNP-/- T-cells to mtDNA damage.
The reasons for this speculation are as follows; firstly, human thymocytes were found
to have increased deoxyguanosine kinase (dGK) activity and elevated levels of dGTP
compared to peripheral T and B-lymphocytes (Refer Figure 2) (Cohen et al., 1980). dGK is
an enzyme that phosphorylates dGuo to dGMP and finally to dGTP and is localized in the
mitochondrial membrane (Wang et al., 1996; Johansson et al., 1996), as a consequence
designating the site for dGTP formation and its subsequent accumulation. Thus Arpaia et al
30
postulated that the accumulation of dGTP in the mitochondria of PNP-/- T lymphocytes
initiates apoptosis by interfering with the repair of mtDNA damage (Arpaia et al., 2000).
Further support for this hypothesis came from the observation that mitochondria of PNP
expressing thymocytes, incubated with dGuo in the presence of PNP inhibitor accumulated
dGTP (Arpaia et al., 2000). Moreover, dissipation of mitochondrial membrane potential
(MMP) in these cells was resistant to caspase inhibition, suggesting that the apoptotic signals
originated within the mitochondria. Also, increased sensitivity of PNP-/- thymocytes and
splenic T-cells to irradiation offers further evidence of DNA damage (Arpaia et al., 2000).
The importance of the balance in dNTP precursors for DNA replication and repair has been
established for a long time now however, its relevance in human disease has gained
momentum only in the past decade. Similar to the hypothesized mechanism of abnormal
DNA synthesis and repair in PNP deficiency, studies carried out in thymidine phosphorylase
and uridine phosphorylase double knockout mice showed nucleotide pool imbalance in the
tissues of these mutant mice and partial mtDNA deletion in their brains (Lopez et al., 2009).
Correspondingly, in dGK deficiency, the conversion of dGuo to dGTP does not occur and
thus mtDNA reduces proportional to dGTP levels as dGTP is the largest dNTP pool in the
mitochondria (Saada et al., 2008). This deficiency is seen to cause a hepatocerebral variant
of mtDNA depletion syndromes (Ashley et al., 2007; Saada et al., 2008), further
strengthening the importance of mitochondrial dNTP balance in various tisssues.
Mechanisms of apoptosis involved in ADA deficiency may also explain apoptosis in
PNP deficiency. The extracellular pathway via death receptor signaling was examined in
ADA deficiency. T-cells treated with a potent ADA inhibitor and dAdo have demonstrated
accumulation of dATP resulting in the inhibition of ribonucleotide reductase and imbalance
31
in the dNTP pool which leads to abrogation of DNA synthesis and repair (Ullman et al.,
1979; Carson et al., 1977; Cohen A, 1986). Apoptotic signals via Fas was eliminated as a
possible mechanism, nevertheless there is still some speculation regarding an alternative
death receptor pathway (Figure 6, Pathway A) (Thompson et al., 2003). The extrinsic
pathway was not studied in PNP deficiency. Another postulated mechanism is that
accumulation of dATP in ADA deficiency triggers the release of mitochondrial cytochrome c
initiating an apoptotic cascade (Figure 6, Pathway B) (Thompson et al., 2003). Similarly it
has been postulated that mtDNA damage may be caused by the accumulation of dGTP in
PNP deficiency leading to apoptosis via the release of cytochrome c from the mitochondrial
membrane (Figure 6, Pathway B) (Arpaia et al., 2000). It is also possible that accumulated
purine metabolites causes nuclear DNA damage which triggers an apoptotic signaling
cascade (Figure 6, Pathway C). There is also some evidence for a p53-dependent apoptotic
mechanism in ADA deficiency which is preventable by over-expression of Bcl-2 (Benveniste
and Cohen, 1995). Of course it is still possible that pro-apoptotic Bcl-2 family members are
involved, but through a p53- independent mechanism (Figure 6, Pathway D). Taken
together, accumulation of dGTP in PNP deficiency may cause apoptosis by one or more of
the above mentioned pathways.
32
Intrinsic pathway
Cellular stress,
(PNP metabolites?)
Pathway APathway A
Pathway BPathway B
Pathway CPathway C
p53?
Extrinsic pathway
Cellular stress, dATP, (PNP metabolites?)
p53?
Pathway DPathway D
?
Intrinsic pathway
Cellular stress,
(PNP metabolites?)
Pathway APathway A
Pathway BPathway B
Pathway CPathway C
p53?
Extrinsic pathway
Cellular stress, dATP, (PNP metabolites?)
p53?
Pathway DPathway D
?
Figure 6: A schematic representation of postulated apoptotic mechanisms caused by ADA
and PNP deficiencies (adapted from Thompson et al., 2003). Pathway A: abnormal levels
of purine metabolites cause induction of death receptor signaling (e.g. Fas). Pathway B:
accumulated dGTP or dATP causes mitochondrial DNA damage leading to a loss in MMP
causing the release of cytochrome c from the mitochondria. Pathway C: abnormal levels of
purine metabolites lead to nuclear DNA damage finally disrupting the mitochondria, leading
to activation of caspases. Pathway D: abnormal leveles of purine metabolites causes
induction of pro-apoptotic Bcl-2 family members, leading to the release of cytochrome c
from the mitochondria.
33
CHAPTER 2: RATIONALE, HYPOTHESIS AND
EXPERIMENTAL OBJECTIVES
34
Rationale, Hypothesis and Experimental Objectives
PNP deficiency in humans is associated with impaired thymic output; however the effect of
PNP deficiency on thymocytes has been difficult to study due to the inability to obtain thymi
from PNP-deficient patients. Arpaia et al and Snyder et al demonstrated increased numbers
of DN thymocytes and reduced numbers of DP and SP thymocytes in PNP-deficient murine
models, nevertheless, the thymus was not the prime interest of their studies (Arpaia et al.,
2000; Snyder et al., 1997). Therefore little is known on the precise stage(s), cause(s) and
mechanism by which PNP deficiency induces abnormality in thymocyte development, which
was the focus of this study.
2.1. Rationale
The increased number of DN thymocytes and reduced number of DP thymocytes found in
PNP-deficient mice could be due to a defect in one or more of the developmental stages
described in Figure 7. However, two major reasons, outlined below, led us to speculate that
the developmental delay is in the later DN stages, more specifically between the DN3 to
DN4 stage transition. ADA deficiency, which as described earlier shares many metabolic
and immunological features with PNP deficiency, has been shown to disrupt thymocyte
development early within the DN3 stage. Thymocytes taken from 15-day old ADA-deficient
mice and grown ex-vivo on FTOCs, revealed profound reduction in the number of CD44–
CD25+ DN3 thymocytes (Thompson et al., 2000). Secondly, the transition between DN3 to
DN4 is a major “check-point” in thymocyte maturation. Many immunodeficiency models
(targeted deletions in Rag, pre-TCRα, TCRβ genes, lymphocyte specific tyrosine kinase-
35
Lck) associated with reduced thymic cellularity similar to PNP deficiency, affect the
development of thymocytes between the DN3 to DN4 stage, which is the stage of β-selection
(Godfrey et al., 1994; Levin et al., 1993; Fehling et al., 1995; Shinkai et al., 1992;
Mombaerts et al., 1992). Therefore we hypothesized that a developmental delay between the
DN3 to DN4 transition is the most plausible explanation for the reduced numbers of DP
thymocytes associated with PNP deficiency.
CD117+CD44+CD25-
CD17+CD44+CD25+
CD117-CD44-CD25+
CD117-CD44-CD25-
CD4+CD8+
CD4-CD8- CD4-CD8- CD4-CD8- CD4-CD8-
Double Negative Compartment CD8+
CD4+
Double positive
(DP)Single Positive (SP)
β Selectionα gene
rearrangement
DN1 DN2 DN3 DN4
(DNs)
CD117+CD44+CD25-
CD17+CD44+CD25+
CD117-CD44-CD25+
CD117-CD44-CD25-
CD4+CD8+
CD4-CD8- CD4-CD8- CD4-CD8- CD4-CD8-
Double Negative Compartment CD8+
CD4+
Double positive
(DP)Single Positive (SP)
β Selectionα gene
rearrangement
DN1 DN2 DN3 DN4
(DNs)
Figure 7: Stages in murine thymoctye development. The red bar indicates the most likely
stage at which thymocyte development is affected in PNP deficiency.
In addition to the increase in the number of DN thymocytes, there is a striking reduction in
the number of DP cells in the thymus of PNP-/- mice, which could be caused by:
1. Decreased proliferation of early thymocytes, as previous studies have shown reduced
proliferation of T-lymphocytes isolated from PNP-deficient patients and mice (Toro et
al., 2006, Arpaia et al., 2000, Markert et al., 1991) as well as lymphocytes isolated from
36
patients treated in vitro or in vivo with chemical PNP inhibitors (Bantia et al., 2003;
Conry et al., 1998; Gandhi et al., 2005; Kicska et al., 2001).
2. Reduction in the percentage of cycling cells due to the effect of toxic purine metabolites.
Indeed, Thompson et al showed that the percentages of cycling cells are reduced in ADA-
deficient cultures (Thompson et al., 2000). This finding is consistent with the reduced
capacity of ADA deficient cultures to reach the CD44- CD25- DN4 stage that normally
contains the highest percentage of rapidly cycling thymocytes in the DN populations
(Tourigny et al., 1997).
3. Increased apoptosis of DP thymocytes. This is the most likely explanation as majority of
normal DP thymocytes undergo apoptosis due to positive and negative selection (Surh
and Sprent, 1994; Ohashi, 2003). In fact, Arpaia et al showed increased apoptosis in
normal thymocytes that were treated with a PNP inhibitor and dGuo (Arpaia et al.,
2000), and T-cell lines treated with BCX-1777 and dGuo also revealed significant
apoptosis (Bantia et al., 2003). Furthermore, there is extensive apoptosis in the
thymus of ADA-deficient mice (Apasov et al., 2001) and also in ADA-deficient
thymocytes grown in FTOCs (Thompson et al., 2000; Van de Wiele et al., 2006).
Among the metabolic abnormalities found in human and murine PNP deficiency which may
lead to increased apoptosis and reduced number of DP cells, we and others hypothesized that
accumulation of dGuo, or a phosphorylation product of dGuo, such as dGTP is the most
likely cause. Among the four substrates of the PNP enzyme (Figure 8), dGuo is the only
substrate that has an alternate metabolic fate, as it can be phosphorylated by cytosolic dCK or
mitochondrial dGK to form dGTP. Increased dGK activity has been observed in human
thymocytes (Cohen et al., 1980) thus possibly explaining the preferential accumulation of
37
dGTP in thymocytes. Additionally, dGuo is also the only PNP substrate that has specifically
been able to inhibit the growth of T-cells, which was associated with elevated dGTP levels in
the cells (Gudas et al., 1978; Ullman et al., 1979). Furthermore, the inhibitory effect of dGuo
was abolished by either blocking dGuo uptake into the cells or by inhibiting the enzymes
dCK and dGK, thereby preventing the formation of dGTP (Ullman et al., 1979). These
results are indicative that the formation of dGTP, secondary to the accumulation of dGuo,
may be the actual cause for apoptosis.
dAdoHx
dIno
Ado InoADA
PNP
Guo GuadGuo
dATP dGTPGTP
dGKdGK
HGPRT
Xn
Uric acid
dAdoHx
dIno
Ado InoADA
PNP
Guo GuadGuo
dATP dGTPGTP
dGKdGK
HGPRT
Xn
Uric acid
Figure 8: Schematic representation of the PNP pathway. PNP substrates are depicted in
the box.
The central role of dGTP (that is produced by dGK, an exclusive mitochondrial enzyme) in
PNP-induced apoptosis suggests a mitochondrial-dependent apoptotic pathway (Figure 9).
This is further supported by the observations that dGTP accumulated in the mitochondria of
normal thymocytes treated with dGuo and a PNP inhibitor, that the apoptosis of these
thymocytes was initiated in the mitochondria and that it could be inhibited by over-
expression of Bcl-2, an anti-apoptotic factor localized primarily in the mitochondria and
38
important for maintaining mitochondrial membrane integrity (Arpaia et al., 2000). In
addition, similar involvement of the mitochondria was demonstrated also in ADA deficiency,
in which dATP accumulation induced a mitochondria-mediated apoptosis possibly by the
release of cytochrome c (Van de Wiele et al., 2002).
mtDNA
Mitochondria
dGTP
cytC
cytC
dGuo
PNP
dGuanine
dGK
mRR
mtDNA
Mitochondria
dGTP
cytC
cytC
dGuo
PNP
dGuanine
dGK
mRR
Figure 9: Proposed mechanism of PNP deficiency. The accumulated dGuo is
phosphorylated by mitochondrial dGK to dGTP which may interfere with mtDNA synthesis
and repair either directly or by inhibition of the mitochondrial ribonucleotide reductase
(mRR) eventually leading to apoptosis by the release of cytochrome C (cytC).
39
2.2. Hypotheses
Based on the rationale presented above, we hypothesized that PNP deficiency interferes with
thymocyte development during the transition from the DN3 to the DN4 stage. In addition,
we hypothesized that the reduced DP thymocytes was due to increased apoptosis induced by
the accumulation of dGuo. We also hypothesized that the increased apoptosis of DP
thymocytes in PNP deficiency is initiated in the mitochondria.
2.3. Specific Objectives
The overall objective of this work was to understand the effect of PNP deficiency on
thymocyte development by using a PNP-deficient murine model.
The specific aims of this project were;
1) To analyse the effect of PNP deficiency on thymocyte development during the DN3
to DN4 stage transition.
2) To examine whether the reduced number of DP thymocytes is caused by increased
apoptosis that is induced by the accumulation of dGuo.
3) To determine whether apoptosis of PNP-deficient DP thymocytes is initiated in the
mitochondria.
40
CHAPTER 3: MATERIALS AND METHODS
41
Materials and Methods
3.1. Animals:
PNP-/- mice (C57/BL6 background) in which the catalytic domain of PNP was replaced
by a neomycin cassette (Arpaia,et al. 2000) and normal controls were maintained in a
pathogen free environment at the Sick Kids Hospital Research Institute’s animal facility.
Mice used in all the experiments were 6-8 weeks old. PNP deficiency was confirmed by the
absence of PNP activity in the whole blood samples obtained from the tail of 18-20 day old
mice. All procedures and experiments involving animals were approved by the Institutional
Animal Care Committee of the Hospital for Sick Children, Toronto, Canada.
3.2. PNP enzyme activity:
PNP enzyme activity was determined by measuring conversion of inosine to
hypoxanthine and has been previously described (Arpaia et al., 2000). Briefly, 14C-labeled
inosine (Movarek Biochemicals, Brea, CA) was added to whole cell lysate and incubated at
37°C followed by a brief denaturation of the enzyme at 100°C for 1 min. The unconverted
substrate and the product were separated by thin-layer chromatography (Fischer Scientific,
Ottawa, ON) and measured using scintillation counting. One unit was defined as the amount
of enzyme required to convert 1μmol of inosine to hypoxanthine in 1min at 37°C.
3.3. Cell culture and media:
3.3.1. Thymocyte cultures were maintained for 8-24 hours in RPMI 1640 (Invitrogen,
Burlington, ON) containing 5% heat inactivated fetal bovine serum (FBS)
(Hyclone, Logan, Utah).
42
3.3.2. Jurkat cells were maintained in RPMI 1640 supplemented with 10% FBS.
3.3.3. OP9-DL1 cells were kindly provided by J.C. Zúñiga-Pflücker (University of
Toronto, Toronto, Canada). OP9-DL1 and OP9-DL1 co-cultures were
maintained in α-MEM (Invitrogen, Burlington, ON) supplemented with 20%
FBS as previously described (Awong et al., 2008).
All culture media was supplemented with 1% penicillin/streptomycin (Invitrogen,
Burlington, ON).
3.4. Antibodies and reagents:
3.4.1. Antibodies: Antibodies used in this study were biotin conjugated anti-mouse
Gr-1, CD11c, CD3ε, NK1.1, TER-119, CD117, CD25, CD11b, B220,
TCRβ, CD19, Mac-1, CD4, CD8, pycoerythrin (PE) conjugated anti-CD4 and
anti-CD25, anti-CD71 and anti-Fas (eBioscience), fluorescein isothiocyanate
(FITC) conjugated anti-CD8 and anti-CD25, and allophycocyanin (APC)
conjugated anti-TCRβ (eBioscience). All biotin conjugated anti-mouse
monoclonal antibodies were purchased from eBioscience (San Diego, CA) while
APC conjugated anti-biotin antibody was from Miltenyi biotec (Auburn, CA).
All other antibody-fluorochrome conjugates were purchased from BD
Pharmingen (Mississauga, Ontario).
3.4.2. Pan caspase inhibitor: N-benzyloxycarbonyl-Val-Ala-Asp-fluoromethylketone
(z-VADfmk) was purchased from Santa Cruz Biotechology (San Diego, CA).
3.4.3. MitoTracker® Red CMXRos:
Assessment of MMP was performed using MitoTracker® Red CMXRos
(Molecular Probes, Ivitrogen, Carlsbad, CA). This is a cell-permanent probe
43
containing a mildly thiol-reactive chloromethyl moiety for labeling mitochondria.
When it enters an actively respiring cell, it is oxidized and sequestered in the
mitochondria, where it reacts with thiols on proteins and peptides to form an
aldehyde-fixable conjugate.
3.4.4. Apoptosis Inducing agents:
• dGuo, dGMP and dGTP (Sigma Aldrich, Oakville, ON) were dissolved in
10mM NaOH.
• Dexamethasone (Dex) was purchased from Sigma Aldrich and dissolved in
dimethyl sulphoxide (DMSO, Sigma Aldrich).
3.4.5. PNP inhibitor:
The PNP inhibitor BCX-1777 (Forodesine hydrochloride) was kindly provided by
BioCryst (Durham, NC). 10mM solutions were prepared in DMSO.
3.5 Flow cytometry:
1-2x106 cells were washed and resuspended in staining buffer (PBS containing 1%
FBS). Antibodies conjugated to PE, FITC and APC were added after careful titration. Cells
were stained on ice for 30 minutes, and then washed with staining buffer. Live cells were
identified by PI exclusion and by forward and side scatter profiles. All multicolour flow
cytometric analysis was carried out on a dual laser FACScalibur (Becton Dickinson (BD),
Franklin Lakes, NJ), and data acquired was imported into either CellQuest (BD) or FlowJo
(Tree Star, Ashland, OR) software for analysis.
44
3.6. Bone marrow (BM) cell isolation:
BM was isolated from the tibias and femurs of PNP-/- mice and age matched controls.
The bones were crushed and passed through a 40μM nylon mesh filter to get a single cell
suspension in α-MEM containing 1% FBS. Following erythrocyte lysis for 5 minutes with
ammonium chloride (Stemcell Technologies, Vancouver, BC) and subsequent washes with
1x PBS (Invitrogen, Burlington, ON) supplemented with 1% FBS, the bone marrow cells
were counted using a hemacytometer. 40x106 cells were taken for lineage depletion.
Lineage depletion:
Lineage (Lin) depletion was carried out using a mouse lineage cell depletion kit
(Miltenyi Biotec), according to manufacturer’s protocol. Briefly, cells were labeled with
biotin-conjugated monoclonal antibodies to CD5, CD45R (B220), CD11b, Anti-Gr-1 (Ly-
6G/C), 7-4 and Ter-119, followed by anti-biotin antibodies conjugated to magnetic
microbeads. The magnetically labeled cells were then passed through MS MACS column
(Miltenyi Biotec) and separated in a magnetic field. The Lin-negative cells passed through
the column while the positive cells were retained. Lin-negative cells were taken for co-
culture experiments.
3.7. OP9-DL1 co-cultures:
OP9-DL1 co-cultures, maintained in α-MEM medium supplemented with 20% FBS,
were carried out as previously described (Schmitt and Zúñiga-Pflücker, 2002). Briefly, 105
Lin-depleted BM cells or freshly isolated thymocytes were plated onto each well of a 6-well
plate containing 90% confluent OP9-DL1 cells. IL-7 and Flt3L (R&D Systems,
Minneapolis, MN) were added into the culture medium at concentrations of 1ng/ml and
5ng/ml respectively. Every 4-5 days the co-cultures were disrupted by pipette mediated
45
disaggregation and passed through a 40μM cell strainer (BD Falcon, Bedford, MA) to
remove residual OP9-DL1 cells. Cells were pelleted and replated with fresh media and
cytokines, onto fresh OP9-DL1 cultures. The BM co-culture was analyzed by flow
cytometry every 7 days.
OP9-DL1 cells were maintained in α-MEM supplemented with 20% FBS. Culture
plates containing confluent monolayers of OP9-DL1 cells were trypsinized using 0.25%
Trypsin-EDTA (Invitrogen, Burlington, ON), pelleted down and resuspended into fresh
medium. The cells were split into 4 parts into 10cm dishes (BD Falcon) for further
maintenance and split every other day.
3.8. Thymocyte isolation:
PNP-/- mice and age matched controls were sacrificed by cervical dislocation. The skin
surface was sterilized using 70% ethanol and removed to expose the rib cage. The rib cage
was cut bilaterally through the costochondral junction exposing the thymus. The thymus,
which is found to overlay the heart was removed using sterile surgical instruments. To
prepare a single cell suspension of thymocytes, the thymus was crushed through a 40 μM
nylon mesh filter placed in a petri dish containing α-MEM using the tip of a 3ml syringe
(BD, Franklin Lakes, NJ). The single cell suspension was transferred into a 15 ml falcon
tube and centrifuged. The residual erythrocytes in the resulting cell pellet were lysed by
hypotonic lysis in ice-cold ammonium chloride for 5 minutes and subsequently washed with
PBS containing 1% FBS. Live thymocytes were counted by trypan blue exclusion and the
cells were taken for culture or analysis by flow cytomtery.
46
3.9. BrdU proliferation assay:
In this method, 5-bromo-2-deoxyuridine (BrdU), which is an analog of the DNA
precursor thymidine, is incorporated into newly synthesized DNA. The incorporated BrdU
was stained with specific anti-BrdU fluorescent antibodies. The levels of BrdU incorporated
by the cells were then visualized by flow cytometry.
In vivo:
PNP-/- and age matched controls received two intraperitoneal injections of BrdU (1 mg
each, 4 hours apart). Thymocytes were isolated 1, 3 and 5 days later and stained for CD4
(PE), CD8 (APC) and anti-BrdU (FITC) as described in the previous section. BrdU
incorporation into the thymocytes was detected with a BrdU flow kit (BD Biosciences, San
Jose, CA) according to the Manufacturer’s instructions.
In vitro:
BrdU was added to the BM co-culture to obtain a final concentration of 10μM. The
following day the cells were stained for CD4 (PE), CD8 (APC) and anti-BrdU (FITC)
antibodies and analyzed by flow cytometry.
3.10. Analysis of DN thymocytes:
3.10.1. Analysis of DN1-DN4 subsets:
For analysis of DN thymocyte populations, single cell suspensions of thymus (1-2x106
cells/sample) were incubated on ice for 30 minutes with biotinylated antibodies specific
for CD4, CD8ε, TER-119, B220, Mac-1, CD3ε, TCRβ, CD11b, CD11c, CD19 and
Gr-1. Cells were then washed and stained with anti-biotin microbeads (Miltenyi
Biotec) for 15 minutes at 4-8 °C. The prepared cell suspension was added to MS
MACS columns and the unbound cells were collected in a tube. The tubes were
47
centrifuged to obtain a cell pellet which was then appropriately diluted for staining. The
cells were stained with fluorochrome conjugated antibodies specific for CD25 (FITC)
and CD44 (PE) and incubated on ice for 25 minutes. Cells were then washed and
resuspended in staining buffer and analysed by flow cytometry.
3.10.2. Expansion of DN3 thymocytes ex-vivo:
Single cell suspensions of thymus from 6-8 week old mice were prepared and 107
cells/sample were incubated on ice for 30 minutes with biotinylated antibodies specific
for CD4, CD8ε, TER-119, B220, CD19, CD3ε, CD11b, and CD117. Cells were
washed and subsequently incubated with anti-biotin microbeads and mature cells
depleted using MS MACS columns in a magnetic field according to manufacturer’s
instructions. The enriched fraction which is mainly CD25+ was plated onto OP9-DL1
cultures in which PNP was inhibited using 20 μM BCX-1777 and 50μM dGuo
contained in a 24-well plate. The co-culture was supplemented with 5ng/ml of IL-7
and was analyzed three days later by double staining with either anti-CD4 (PE) and
anti-CD8 (FITC) or anti-CD25 (FITC) and anti-CD71 (PE).
3.10.3. Analysis of intracellular TCRb expression on freshly isolated DN
thymocytes:
Another method used to study the DN4 subpopulation involved isolating the desired
DN subsets from fresh thymocytes as described above. Subsequently, the cells were
stained with anti-CD25 (FITC) followed by fixation and permeabilization using BD
cytofix/cytoperm kit (BD Biosciences). The fixed cells were stained for intracellular
48
TCRβ by staining with APC conjugated anti-TCRβ followed by analysis by flow
cytometry.
3.11. Apoptosis studies:
3.11.1. Annexin-V staining:
Thymocytes (5x106 cells/ml) were cultured in RPMI media containing 5% FBS and
treated with increasing concentrations of dGTP, dGMP and dGuo for 8-24 hrs. Apoptosis
was detected by Annexin staining using the Annexin V-FITC Apoptosis Detection Kit from
Biovision (Mountain View, CA), according to manufacturer’s protocol. In some
experiments, thymocytes were stained with a combination of PE and APC conjugated
antibodies before Annexin-V staining.
3.11.2. Analysis of nuclear DNA fragmentation by TUNEL assay:
Apoptotic nuclear DNA fragmentation was measured by the terminal
deoxynucleotidyl transferase (TdT)-mediated dUTP nick end labeling (TUNEL) technique
using a FITC conjugated dUTP kit (Roche Applied Science, Laval, Quebec) according to
manufacturer’s instructions. The frequency of apoptotic cells as detected by fragmented
nuclear DNA was determined by flow cytometry.
3.11.3. Inhibition of apoptosis using a pan-caspase inhibitor:
Apoptosis was inhibited by pre-incubating thymocytes for 30 minutes with 50μM
z-VADfmk followed by induction of apoptosis using either 12.5μM dGuo or 3.75μM Dex.
The cells were cultured for 20-24 hrs, stained for Annexin-V, and analyzed by flow
cytometry.
49
3.12. Mitochondrial membrane potential (MMP):
Thymocytes were incubated with 25nM MitoTracker® Red CMXRos in α-MEM
containing 1% FBS at 37°C for 15-20 mins in the dark. The cells were washed and
resuspended in staining buffer and analyzed by flow cytometry. Cell debris was
electronically gated out based on the forward and side scatter. For experiments involving
staining of cell surface antigens, MitoTracker® staining was carried out initially followed by
staining for the cell surface markers.
3.13. Re-introduction of PNP into PNP-/- cells:
PNP-/- thymocytes were treated with 1μg/ml of TAT-PNP for 1 hour followed by
induction of apoptosis using either 12.5μM dGuo or 125μM dGMP. Thymocytes were
incubated for 20-24 hrs and analyzed by flow cytometry.
TAT-PNP Production: TAT-PNP was produced as previously described (Toro.A et al.,
2006) with slight modifications. Briefly, PTD-PNP inserted into a pRSET vector with an N-
terminal histidine tag was expressed in BL21(DE3) E.coli. PTD-PNP transformed bacteria
were grown in Overnight Express™ Instant TB Medium (Novagen, Madison, WI), pelleted
down and lysed using Bugbuster® Master Mix (Novagen). The lysate was applied to a
TALON resin column (Clontech Laboratories Inc., Mountain View, CA) and washed with
15mM imidazole buffer (50mM sodium phosphate, 300mM NaCl). The proteins bound to
the resin were eluted using 250mM imidazole buffer. The proteins were desalted using PD-
10 columns (GE Healthcare, Piscataway, NJ) and concentrated using Amicon® ultra-15
centrifugal filter units (Millipore, Massachusetts). Protein concentration was determined by
the BioRad assay with bovine serum albumin as a standard. Purity of the TAT-PNP
preparation was determined by SDS-PAGE using a 10-12% tris glycine gel (Invitrogen) and
50
western blotting with rabbit anti-human PNP antibodies generated by our lab (Toro et al., 20
06). The biological activity of the purified proteins was determined by assaying for PNP
activity (as described earlier).
3.14. Viable cell counting:
Viable cells were counted using a hemacytometer (Hausser Scientific, Horsham Rd, PA).
The cells were diluted accordingly and taken for trypan blue staining (Invitrogen). 10μl
of this mixture was added to the wells of the hemacytometer and the number of cells
overlying four 1mm2 areas of the counting chamber was counted using a hand counter.
Cell numbers were calculated using the formula;
Total cells counted in 4mm2 / 4 x (dilution factor) x104 =cells / ml
3.15. Statistical analysis:
Statistical analysis was carried out using a two-tailed Student’s t-test where applicable.
The sample size (n) is indicated for each experiment. Differences were considered
significant if the p value was less than 0.05. Statistical analyses were performed using the
Microsoft Excel software.
51
CHAPTER 4: RESULTS
52
Results
4.1. Abnormal lymphocyte subpopulation in thymus of PNP-/- mice.
We have consistently shown an increase in the percentage of CD4-CD8- DN
thymocytes and decrease in the percentage of CD4+CD8+ DP thymocytes in PNP-/- mice
compared to normal littermates (Figure 10A). Furthermore, there was also an increase in the
absolute number of DN cells with a significant decrease in the total numbers of DP
thymocytes and CD4+ and CD8+ SP thymocytes in PNP-/- mice (Figure 10B- note the
loagarithmic scale).
53
A)
B)
1.0
10.0
100.0
1000.0
DN DP CD4 CD8
Num
ber o
f thy
moc
ytes
(x10
6 )
PNP+/+PNP-/-
Figure 10: Distribution of thymocyte populations in PNP-/- mice. Single cell suspensions
of thymocytes from 6-week old PNP-/- mice and age matched controls were stained with anti-
CD4 (PE) and anti-CD8 (FITC) antibody. (A) Percentage of cells in different subpopulations
is indicated in the respective quadrants. The plot is representative of 9 independent
experiments with one or two mice in each. (B) The graph displays the mean ± SD of the total
number of cells in each thymocyte subpopulation isolated from PNP-/- mice (white bars) and
PNP+/+ controls (black bars) (n= 9 in each, ** p=0.0001, * p<0.001).
CD8
CD
4
PNP+/+ PNP-/-
**
*
*
*
54
Aim 1: To determine whether PNP deficiency interferes with the transition of
thymocytes from the DN3 to the DN4 stage.
4.2. PNP deficiency disrupts thymocyte maturation at the DN3 to DN4 stage
transition.
ADA deficiency is another inherited defect in purine metabolism similar to PNP that
causes abnormal thymocyte development. It was previously shown that ADA deficiency
interferes with maturation of DN3 cells (Thompson et al., 2000). Therefore, we hypothesized
that PNP deficiency also affects development of late DN cells.
To identify the effects of PNP deficiency on differentiation of DN thymocytes, and to
tease out the stage at which development was being inhibited, several strategies were used;
1) To specifically study the DN thymocyte populations, DN cells were isolated by column
purification and analyzed for the expression of CD25 and CD44. A representative example
is provided in Figure 11A, demonstrating higher percentage of DN3 cells in thymi of PNP-/-
mice. Calculation of the absolute numbers of thymocyte subsets confirmed that DN3 cells
were significantly increased in thymi from PNP-/- mice (Figure 11B).
55
A)
65.9026.58
1.805.73
65.9026.58
1.805.73
86.0111.77
1.240.98
86.0111.77
1.240.98
CD25
CD
44
B)
0.00
0.50
1.00
1.50
2.00
2.50
DN1 DN2 DN3 DN4
Num
ber o
f thy
moc
ytes
(x10
6 )
PNP+/+PNP-/-
Figure 11: Characterization of maturation of DN thymocytes in PNP-deficient mice.
CD25 and CD44 expression were compared in freshly isolated thymocytes from PNP-/-
mice and age matched controls. Single cell suspensions were stained with biotin-labeled
antibodies against CD4, CD8, CD3, TER-119, NK1.1, CD19, B220, CD11b and Gr-1
followed by anti-biotin microbeads and separated using MS MACS columns. The resulting
DN fraction was stained with anti-CD44 (PE) and anti-CD25 (FITC). (A) Representative
data show expression of CD25 and CD44 in PNP-/- thymocytes and age matched controls.
(B) Data show DN thymocyte subpopulation numbers in PNP-/- mice (white bars) and
controls (black bars). Cells were characterized as CD44+CD25- (DN1), CD44+CD25+
(DN2), CD44-CD25+ (DN3) and CD44-CD25- (DN4). Data is representative of 5
independent experiments with one animal in each (*p =0.01, **p=0.004).
DN1
DN2
DN4
DN3
*
PNP+/+ PNP-/-
56
2) The strategy described above, in which thymocytes were collected at one point in
time, allowed us to study thymocyte development at a steady state. To study
development of thymocytes over time, I used a different strategy, which relied on
expanding T-cells in vitro from normal BM derived stem cells using the OP9-DL1
co-culture technique. The pharmacological inhibitor of PNP (BCX-1777) was used to
prevent the degradation of toxic purine metabolites by OP9-DL1 cells thereby
simulating PNP-deficient conditions in vitro. dGuo was also added to the cultures to
simulate the excess dGuo found in PNP-/- mice. dGuo or inhibitor added separately
did not have any effect on the development of cells (data not shown). During the first
week of culture, when most of the cells were still in the early DN stages (DN1 and
DN2), there were no significant differences between cultures with normal or
abnormal purine metabolism (Figure 12A). This finding suggested that early DN
differentiation is not affected by PNP deficiency which was in concordance with our
in vivo findings. In contrast, by 2-3 weeks of co-culture, when majority of the
developing cells had matured into the later DN stages, a significant increase in the
percentage of DN3 cells was evident in the PNP-deficient cultures (Figure 12B). Of
note, CD4+ and CD8+ cells were not excluded from analysis, therefore our strategy
can not definitely distinguish between abnormal DN3 to DN4 development or
abnormal DN4 to DP development since the unstained fraction in the DN analysis
constitutes DN4, DP and SP cells. Nevertheless, the significantly increased
percentage of CD25+ DN3 cells and the markedly reduced percentage of DP cells in
cultures with PNP-deficient conditions (Figure 12C), suggests that the confounding
factors described above may have not been significant.
57
A)
0
10
20
30
40
50
60
70
DN1 DN2 DN3 DN4/DPSP
Perc
enta
ge o
f cel
lsUntreateddG+I
B)
0
10
20
30
4050
60
70
80
90
100
DN1 DN2 DN3 DN4/DP/SP
Perc
enta
ge o
f cel
ls
UntreateddG+I
*
**
58
C)
0.0
10.0
20.0
30.0
40.0
50.0
60.0
CD4 CD4/CD8 CD8
Perc
enta
ge o
f cel
ls
UntreateddG+I
Figure 12: Characterization of maturation of DN thymocytes under PNP-deficient
conditions using OP9-DL1 co-culture. BM cells were obtained and lineage depleted using
MS MACS columns. Lin- cells were seeded onto near-confluent OP9-DL1 monolayers and
were allowed to develop over a period of 3 weeks. Cells were harvested after (A) 7 days and
(B) 21 days and analyzed for expression of CD25 (FITC) and CD44 (PE) (*p=0.0004,
**p=0.0001). (C) Cells were analyzed for CD8 (FITC) and CD4 (PE) expression on day 21
of co-culture. Data represents the mean ± SD of 3 independent experiments. White bars
indicate PNP+/+ BM development in PNP-deficient environment, black bars represent PNP+/+
BM development in PNP-proficient environment (dG+I, 50μM dGuo+ 20μM inhibitor).
*
59
3) In order to mimic ex vivo conditions in PNP-/- mice more closely, a third strategy was
used to assess the effects of abnormal purine metabolism in PNP deficiency by following
the development of T-cells in vitro from Lin- PNP-/- BM (in contrast to PNP+/+ BM in
strategy #2) cells using OP9-DL1 stromal cells (in contrast to the strategy 2 which tested
normal BM). This strategy avoided the use of the chemical PNP inhibitor (BCX-1777) and
PNP-deficient conditions were simulated using only excess dGuo (75μM vs 50μM in
strategy 2). Similar to the in vitro findings in strategy 2, in this strategy, by the third week
of co-culture, when most of the cells had transited to the later DN stages and the DP stage,
PNP-deficient BM cultures had a higher percentage of cells in the DN3 stage compared to
normal BM cultures (Figure 13A, 13C). Further, analysis of the DP and SP stages revealed
a lower percentage of cells in the DP stage in the PNP-deficient BM cultures (Figure 13B),
an effect that is markedly exacerbated in the presence of 75μM dGuo both in normal and
PNP-deficient conditions. This suggests that PNP deficiency and the accumulation of
dGuo interferes predominantly after reaching the DN3 stage. The caveats mentioned in
strategy 2 also apply to this method.
60
A)
33.9462.61
0.692.75
33.9462.61
0.692.75
37.2466.45
0.542.24
37.2466.45
0.542.24
37.2460.14
0.691.93
37.2460.14
0.691.9367.9125.66
3.393.04
67.9125.66
3.393.04
49.0147.11
1.961.92
49.0147.11
1.961.92
38.7658.88
0.871.49
38.7658.88
0.871.49
44.3652.75
1.381.51
44.3652.75
1.381.51
84.969.22
4.761.06
84.969.22
4.761.06
B)
6.3040.74
48.334.63
6.3040.74
48.334.63
4.0238.31
50.637.04
4.0238.31
50.637.04
7.1041.11
47.584.21
7.1041.11
47.584.21
4.7578.78
13.253.22
4.7578.78
13.253.22
2.9257.56
36.003.52
2.9257.56
36.003.52
2.1051.27
41.105.53
2.1051.27
41.105.53
5.3946.75
44.733.13
5.3946.75
44.733.13
6.0287.09
5.950.95
6.0287.09
5.950.95
CD8
CD 25
Untreated 25μM dGuo 50μM dGuo 75μM dGuo
PNP+/+
PNP-/-
Untreated 25μM dGuo 50μM dGuo 75μM dGuo
PNP+/+
PNP-/-
CD
4
61
(C)
Analysis of BM co-culture treated with 75μM dGuo
0
10
20
30
40
50
60
70
80
90
DN1 DN2 DN3 DN4/DP/SP
Perc
enta
ge o
f cel
ls
PNP+/+PNP-/-
Figure 13: Characterization of maturation of DN thymocytes under PNP-deficient
conditions using excess dGuo. PNP-/- and PNP+/+ Lin- BM was cultured on OP9-DL1 cells
and analyzed for the expression of CD25/CD44 and CD4/CD8 after 18 days of co-culture.
(A) Data is representative of CD25 (FITC) vs CD44 (PE) profiles. (B) Data is representative
of CD4 (PE) vs CD8 (APC) profiles. Data is representative of two independent experiments.
Percentage of cells in different subpopulations is indicated in the respective quadrants.
(C) Graph is a representative of two independent experiments and depicts the percentage of
DN subsets in PNP+/+ BM (black bar) and PNP-/- BM (white bar) co-cultures after 18 days.
62
4) The fourth strategy employed to specifically delineate the DN3 and DN4 transition
relied on measuring CD71 expression. The upregulation of CD71, the transferrin receptor,
mediates iron uptake and is also critical for cell growth (Schneider et al, 1982; Trowbridge
and Shackelford, 1986). Most DN3 thymocytes express low levels of CD71, whereas surface
expression of this nutrient transporter is high in DN4 cells. Progression through β-selection
is thus accompanied by upregulation of CD71 (Kelly et al., 2007). DN3 thymocytes isolated
from PNP-/- and PNP+/+ mice were co-cultured for 3 days on OP9-DL1 cells in which PNP
deficiency was simulated using BCX-1777 and dGuo as mentioned earlier. The limitation of
this study is that the negative fraction obtained after column purification was enriched for
DN3 and was not a homogeneous DN3 population as the technique employed does not allow
for removal of DN4 cells. This may have been the reason why I was unable to detect
differences in CD71 upregulation among DN4+ cells, and in the development of DP cells
(Figure 14A and 14B). A more appropriate experiment would be to sort for DN3 cells by
flow cytometry before plating onto OP9-DL1 cells, but one can argue that these cells are now
labeled and can skew the experimental results. Further optimization is required to draw any
conclusions from these experiments.
5) The DN4 subset is difficult to delineate due to the absence of a defined cell surface
marker. Therefore, I employed another strategy which examined intracellular TCRβ
expression. Early DN3 cells do not express TCRβ chain intracellularly, late DN3 cells start
expressing TCRβ and DN4 cells will express intracellular TCRβ chain. The results indicate
an increased percentage of early DN3 cells in PNP-/- mice with a significant reduction in the
percentage of DN4 cells expressing TCRβ intracellularly compared to normal controls using
63
lineage depleted freshly isolated thymocytes (Figure 15A, 15B). This is indicative that the
percentage of cells transiting from the DN3 to the DN4 stage is reduced in PNP-/- mice
compared to normal controls.
A)
6.6924.00
40.3328.98
6.6924.00
40.3328.98
6.5115.73
39.4728.29
6.5115.73
39.4728.29
CD25
CD
71
B)
10.9047.25
29.3012.56
10.9047.25
29.3012.56
12.3843.32
28.1916.11
12.3843.32
28.1916.11
CD8
CD
4
Figure 14: Characterization of DN3 and DN4 subsets by upregulaton of CD71
expression. DN3 thymocytes were isolated from PNP-/- mice and normal controls by
depleting for CD4, CD8, CD3, CD11b, B220, CD19, TER-119, Gr-1 and CD117 positive
cells using MS MACS columns, and the enriched DN3 fraction was co-cultured on OP9-DL1
cells for 3 days. The plot is representative of (A) CD71 (PE) vs CD25 (FITC) profiles and
(B) CD4 (PE) vs CD8 (FITC) profiles after 3 days of co-culture for 5 independent
experiments with one animal in each group.
PNP+/+ PNP-/-
PNP+/+ PNP-/-
64
A)
33.654.26
28.8333.25
33.654.26
28.8333.25
44.666.88
30.8817.58
44.666.88
30.8817.58
CD25
Intr
acel
lula
r TC
Rβ
B)
0
10
20
30
40
50
60
Early
DN
3(C
D25
+/TC
Rb-
)
Late
DN
3(C
D25
+/TC
Rb+
)
DN
4 (C
D25
-/T
CR
b+)
Perc
enta
ge o
f cel
ls
PNP+/+PNP-/-
Figure 15: Characterization of the DN3 and DN4 subsets by intracellular TCRβ staining.
DN3 and DN4 cells (characterized as negative for CD4, CD8, CD3, CD11b, B220, CD19,
TER-119, Gr-1 and CD117) were isolated from PNP-/- mice and normal controls and stained
for CD25 (FITC) and intracellular TCRβ (APC). (A) Data is representative of one of four
experiments. Percentage of cells is indicated in each quadrant. (B) The graph depicts the
mean ±SD of the percentage of DN3 and DN4 cells in PNP-/- mice (white) and PNP+/+ mice
(black). Data is representative of 4 independent experiments with one animal in each group
(*p=0.0005).
DN4
(CD25-/TCRβ+)
Late DN3
(CD25+/TCRβ+)
(CD25-/TCRβ-)
Early DN3
(CD25+/TCRβ-)
PNP+/+PNP-/-
*
65
CONCLUSION
The above data is indicative that abnormal PNP function and accumulation of dGuo affects
thymocyte development within the DN3 stage or at the transition from DN3 to the DN4 and
DP compartments. However, thus far I have not been able to identify the cause of abnormal
DN development.
66
Aim 2: To determine whether the reduced number of DP thymocytes in PNP deficiency is
due to increased apoptosis caused by dGuo.
4.3. Increased apoptosis in freshly isolated PNP-/- DP thymocytes.
As described earlier, in addition to the decreased percentage of DN thymocytes in PNP
deficiency, there is also a marked reduction in the number and percentage of DP thymocytes.
We hypothesized that the reduced DP could be caused by increased apoptosis. Therefore I
initially measured apoptosis in freshly isolated thymocytes by flow cytometry. I found higher
apoptosis (as measured by Annexin-V staining) in PNP-/- DP thymocytes than in PNP+/+
thymocytes (Figure 16A), as previously reported by Arpaia et al (Arpaia et al., 2000), which
was statistically significant (p=0.02) although not dramatic (Figure 16B). This moderate
increase in apoptosis can probably be attributed to the ability of resident macrophages to
rapidly clear apoptotic thymocytes. No difference in apoptosis of DN and CD4+ or CD8+ SP
populations was found between controls and PNP-/- mice.
67
A)
CD8
CD
4
PNP+/+
PNP-/-
CD8
CD
4
PNP+/+
PNP-/-
B)
0
0.5
1
1.5
2
2.5
3
3.5
4
4.5
PNP+/+ PNP-/-
Perc
enta
ge o
f DP
thym
ocyt
es u
nder
goin
g ap
opto
sis PNP+/+PNP-/-
Figure 16: Increased apoptosis in freshly isolated DP thymocytes from PNP-deficient
mice. Freshly isolated thymocytes were analyzed by three colour flow cytometry for CD8
(APC), CD4 (PE) and Annexin-V (FITC). (A) Data is representative of Annexin positive
cells among CD4+CD8+ DP thymocytes isolated from PNP+/+ (black) or PNP-/- (white) mice.
Percentage of apoptotic cells are indicated above the bars. (B) Data represents mean ± SD of
apoptotic DP thymocytes (n=5 in each, *p=0.02).
*
68
4.4. PNP Deficiency does not affect proliferation or the maturation of DP thymocytes.
The most obvious consequence of PNP deficiency on thymocyte development is the
reduction in the DP thymocytes. We reasoned that although increased apoptosis is the most
likely cause for the DP phenotype observed in PNP deficieny, the reduction in DP
thymocytes could also reflect a defect in expansion of DN precursors or reduced maturation
of DP thymocytes. Therefore, I monitored the generation and subsequent maturation of PNP-
/- DP thymocytes 1 to 5 days after BrdU injection. One day after BrdU injection, the majority
of PNP+/+ and PNP-/- precursors that were labeled with BrdU were DP cells, in concordance
with previous reports (Lucas et al., 1993; Matei et al., 2006) (Figures 17A, 17B). Similar
proportions of DN and DP thymocytes from PNP-/- and PNP+/+ mice incorporated BrdU one
day after BrdU injection, demonstrating that PNP deficiency did not affect thymocyte
proliferation or the initial generation of DP thymocytes. The percentage of BrdU labeled
progeny becoming DP cells decreased dramatically over the next 5 days as some DP matured
into SP thymocytes, while others died (Figures 17A, 17B, 17C). Importantly, the percentage
of PNP-/- precursors becoming SP thymocytes was similar to control after 5 days, indicating
that maturation of DP into SP was not affected by PNP deficiency. Of note is the marked
reduction in the percentage of BrdU+ DP thymocytes in PNP-/- mice 5 days after BrdU
injection (Figure 17C). This observation, together with the normal percentage of BrdU+ SP
PNP-/- thymocytes, indicates accelerated death of PNP-/- DP cells which further supports our
previous observations.
In conclusion, our data clearly demonstrates that the reduced numbers of DP
thymocytes in PNP deficiency is due to a survival defect and not due to decreased
proliferation of thymocyte precursors or defect in maturation of DP thymocytes.
69
A)
31.51%39.97%
48.81%
6.15%3.36%
61.15%
31.51%39.97%
48.81%
6.15%3.36%
31.51%39.97%
48.81%
6.15%3.36%
61.15%
B)
PNP+/+ PNP-/-
CD8
CD
4
PNP+/+ PNP-/-
CD8
CD
4
BrdU
Day 1
Day 3
Day 5
Cel
l cou
nts
PNP+/+ PNP-/-
Day 1
Day 3
Day 5
70
C)
0
10
20
30
40
50
60
CD4+ DP CD8+
Perc
enta
ge B
rdU
+ ce
lls
PNP+/+PNP-/-
Figure 17: BrdU labeling profiles for PNP-/- mice show normal thymocyte proliferation
but indicate a survival defect most strikingly obvious on day 5 after labeling.
(A) Histograms represent BrdU labeling of total thymocytes from PNP-/- mice and age
matched controls at the indicated time points after BrdU injection. (B) The panels depict
CD4 vs CD8 thymocyte profiles of PNP-/- and PNP+/+ mice gated on BrdU+ cells at days 1, 3
and 5 after injection. Representative results of 3 independent experiments with one or two
mice in each. (C) Graph represents mean ±SD of percentage of BrdU+ SP and DP
thymocytes in PNP+/+ (black) and PNP-/- (white) mice after 5 days of BrdU injection (n=3 in
each,*p=0.001).
*
71
4.5. dGuo causes increased thymocyte apoptosis.
After determining that PNP deficiency induces thymocyte apoptosis, I was interested to
understand the mechanism for this effect. It was thus hypothesized that the increased
apoptosis of DP thymocytes is caused by accumulation of toxic PNP substrates, such as
dGuo and its subsequent conversion to dGTP, and not by depletion of PNP products such as
guanine (Gua) or GTP (Figure 18).
dGuoaccumulation
Guanine+ deoxyribose-1-phosphate
dGMP
dGDP dGTP Alterations in dNTP pools
Thymocyteapoptosis
PNP
dGK
dGuoaccumulation
Guanine+ deoxyribose-1-phosphate
dGMP
dGDP dGTP Alterations in dNTP pools
Thymocyteapoptosis
PNP
dGK
Figure 18: Schematic representation of the role of PNP in purine degradation and
salvage. PNP catalyzes the phosphorolysis of dGuo to Gua. In the absence of PNP, dGuo
accumulates and is phosphorylated by mitochondrial dGK eventually converting it into
dGTP. dGTP inhibits mitochondrial ribonucleotide reductase causing mtDNA damage
ultimately leading to apoptosis. PNP deficiency also prevents the formation of PNP products
such as Gua, resulting in depletion of GTP.
72
I therefore determined the effects of dGuo on PNP-deficient thymocytes. PNP-/- thymocytes
were cultured with increasing concentrations of dGuo for 24 hours and apoptosis was
measured by flow cytometry after staining with Annexin-V. A dose-dependant increase in
apoptosis was observed in PNP-/- thymocytes compared to PNP+/+ thymocytes, most evident
at 6.25μM and 12.5 μM dGuo (Figure 19A), concentrations which are similar to those found
in the serum of PNP-deficient patients and PNP-/- mice. Jurkat T-cells, another representative
of PNP-proficient cells were resistant to dGuo induced apoptosis (Figure 19A), further
confirming that dGuo causes apoptosis specifically in PNP-/- thymocytes.
Direct effects of dGuo phosphorylation product dGMP on thymocyte apoptosis.
We hypothesized that the dGuo phosphorylation to dGMP by mitochondrial dGK and
subsequently to dGTP was the cause for thymocyte apoptosis (Figure 18). Therefore I tested
the effects of increasing concentrations of dGMP on thymocyte apoptosis. dGMP showed a
concentration-dependant loss of cell viability similar to dGuo (Figure 19B). As expected,
dGTP, which is a positively charged molecule can not cross the cell membrane and had no
effect on normal or PNP-/- thymocytes (data not shown).
73
A)
0
5
10
15
20
25
30
35
40
UT 3.125 6.25 12.5
Concentration of dGuo (μM)
Perc
enta
ge o
f Ann
exin
+ ce
lls
PNP+/+PNP-/-Jurkat
B)
0.00
5.00
10.00
15.00
20.00
25.00
30.00
UT 31.25 125Concentration of dGMP (μM)
Perc
enta
ge o
f Ann
exin
+ ce
lls
PNP+/+PNP-/-
Figure 19: Demonstration of apoptosis in thymocytes after incubation in vitro with
different purine metabolites. Thymocytes were isolated from PNP-/- mice (dashed) and age
matched controls (solid) and cultured for 24 hours in the presence of indicated concentrations
of (A) dGuo (*p=0.014, **p=0.0018), (B) dGMP (*p=0.0002). Data represents the mean ±
SD of 3 independent experiments with one or two mice in each experiment (UT, untreated).
*
*
**
74
Re-introduction of PNP into the cells increases the survival of PNP-/- thymocytes.
To further substantiate the hypothesis that dGuo and its phosphorylated products was the
cause of thymocyte apoptosis in PNP deficiency, PNP activity was restored inside PNP-/-
thymocytes using PNP fused to a protein transduction domain (TAT), an enzyme
replacement therapy developed in our lab (Toro et al., 2006). TAT-PNP can restore
phosphorolysis of dGuo to Gua thereby preventing accumulation of dGMP and dGTP (refer
Figure 18). Addition of TAT-PNP protected PNP-/- thymocytes from dGuo induced
apoptosis (Figure 20A). In contrast and as expected, TAT-PNP does not prevent apoptosis of
cells treated with dGMP since the formation of dGMP is past the stage of PNP action (Figure
20B).
75
A)
0
10
20
30
40
50
60
UT UT+ TATPNP
dGuo dGuo+ TATPNP
Perc
enta
ge o
f apo
ptot
ic c
ells
PNP-/-
B)
0
5
10
15
20
25
UT dGMP dGMP+TAT-PNPPerc
enta
ge o
f apo
ptot
ic c
ells
PNP-/-
Figure 20: Introduction of PNP into PNP- deficient cultures increases the rate of
thymocyte survival. Single cell thymocyte suspensions were prepared and cultured in the
presence of (A) dGuo (12.5 μM) or (B) dGMP (125μM) for 24 hours after pre-treating with
TAT-PNP (1μg/ml) for 1 hour. The data (A) represents mean ± SD for 5 independent
experiments with one to two mice in each (*p=0.001). (B) Data is representative of two
experiments (UT, untreated).
*
76
4.6. dGuo preferentially causes the apoptosis of PNP-/- DP thymocytes.
In order to magnify the moderate increase in apoptosis observed in freshly isolated PNP-/-
DP thymocytes, an ex-vivo culture was set-up. Thymocytes were cultured in the presence of
dGuo and apoptosis was measured in the various thymocyte subsets after 24 hours. A
significant difference between PNP-/- and PNP+/+ DP thymocytes was seen. As evident in
Figure 21A and 21B, among PNP-proficient cells maintained 24 hours in culture with dGuo,
only 21.58±4.4% of CD4+CD8+ DP thymocytes were apoptotic, compared to 42.74±8.7% of
apoptosis among PNP-/- thymocytes treated similarly (Figure 21C). There were no differences
observed in the rate of apoptosis of CD4+ SP, CD8+ SP or CD4-CD8- DN subpopulations in
the dGuo treated groups (Figure 21B). These findings were in concordance with our in vivo
results (refer Figure 16A and 16B).
77
A)
B)
Untreated thymocytes cultured for 24 hrs
PNP+/+
PNP-/-
PNP+/+
PNP+-/-
Thymocytes treated with 12.5μM dGuo cultured for 24 hrs
DPs undergoing apoptosis
DPs undergoing apoptosis
78
C)
0
10
20
30
40
50
60
PNP+/+ PNP-/-
Perc
enta
ge o
f DPs
und
ergo
ing
apop
tosi
s
PNP+/+PNP-/-
Figure 21: PNP-/- DP thymocytes undergo increased dGuo induced apoptosis.
Thymocytes were incubated for 24 hours in the presence of 12.5μM dGuo. Cells were
analyzed by three colour flow cytometry after staining with anti-CD4 (PE), anti-CD8 (APC)
and Annexin-V (FITC). (A) Histogram is representative of untreated PNP+/+ and PNP-/- DP
thymocytes undergoing apoptosis after 24 hours. (B) Histogram is representative of PNP+/+
and PNP-/- DP thymocytes undergoing apoptosis after 24 hours of dGuo treatment. Numbers
on the histograms represent the percentage of Annexin+ thymocytes in the DP subpopulation.
The percentage of apoptotic cells in the different subpopulations are indicated beside the
respective quadrants. (C) Graph represents the percentage of PNP+/+ (black) and PNP-/-
(white) DP thymocytes undergoing apoptosis after 24 hrs of dGuo treatment. Data is the
mean ±SD of 4 independent experiments containing one or two mice in each experiment
(*p=0.005).
*
79
CONCLUSION
The above mentioned results are indicative that it is indeed increased apoptosis that causes
the reduced numbers of DP thymocytes in PNP-/- mice and not due to a proliferation defect.
Furthermore, the results also show that dGuo is the PNP substrate that acts as a toxic
metabolite possibly by its conversion to dGTP.
80
Aim 3: To examine whether the increased apoptosis of DP thymocytes in PNP deficiency is
initiated in the mitochondria.
After demonstrating that PNP deficiency increases DP thymocyte apoptosis through
elevated dGuo and likely through its phosphorylated products, I was interested to study
the mechanism leading to apoptosis. As mentioned previously in the introduction chapter
(Figure 6), there are two general apoptosis pathways; the intrinsic pathway which
involves the release of cytochrome c from the mitochondria (mitochondrial pathway) and
the extrinsic pathway or activation of death receptors such as Fas by ligand binding
(death receptor pathway).
4.7. Fas expression is not upregulated in PNP deficiency.
Fas expression was analyzed in different thymocyte subpopulations by three colour
flow cytometry. There was no difference in the expression of Fas in freshly isolated DP or
DN thymocytes from PNP-/- or PNP+/+ mice (Figure 22A). Fas expression in the different
thymocyte subsets was similar to previous reports (Ogasawara et al., 1995). Moreover, even
after 24 hours incubation of thymocytes with dGuo, which I have consistently shown to
significantly increase thymocyte apoptosis in PNP deficiency, there was no difference in Fas
expression (Figure 22B). These results suggest that upregulation of Fas may not be the
mechanism by which PNP deficiency mediates apoptosis.
81
A) B)
Figure 22: Fas expression in subpopulations of PNP+/+ and PNP-/- thymocytes. Single cell
suspension from the thymuses of 6-week old PNP-/- mice and age matched controls were
analyzed by three color flow cytometry for CD4 (APC), CD8 (FITC) and Fas (PE)
expression. Histograms are representative of Fas expression on (A) freshly isolated and (B)
24 hour dGuo treated PNP-/- DN (pink) and PNP+/+ DN (blue) thymocytes as well as PNP-/-
DP (green) and PNP+/+ DP (red) thymocytes. Figures are a representative of 3 independent
experiments with one or two mice in each.
DP PNP+/+- red DN PNP+/+- Blue, DP PNP-/--Green, DN PNP-/-- pink
Fas
Fresh thymocytes
12.5 μM dGuo treated
DP DN
DP DN
DP PNP+/+- red DN PNP+/+- Blue, DP PNP-/--Green, DN PNP-/-- pink
Fas
82
4.8. Dissipation of mitochondrial membrane potential (MMP).
The intrinsic pathway (Figure 6, Pathway B) involves the release of pro-apoptotic proteins
that activate caspase enzymes from the mitochondria ultimately triggering apoptosis (Fulda
et al., 2006; Mayer et al., 2003). If the apoptotic signals are generated intramitochondrially,
the disruption of MMP occurs before nuclear DNA fragmentation. In order to determine
whether PNP deficiency disrupts the mitochondrial membrane, the integrity of the
mitochondrial membrane was analyzed by staining with MitoTracker® Red CMXRos.
Following 24 hours of dGuo treatment, a significantly higher percentage of PNP-/-
thymocytes loses MMP (M2), as indicated by a decrease in fluorescence intensity (Figure
23A). Results demonstrate a significantly higher percentage of dGuo treated PNP-/-
thymocytes losing MMP compared to the controls (Figure 23B).
83
A)
B)
0
5
10
15
20
25
PNP+/+ PNP-/-
Perc
enta
ge o
f cel
ls lo
sing
MM
P
PNP+/+PNP-/-
Figure 23: Disruption of MMP. Thymocytes from PNP-/- mice and age matched controls
were labeled with MitoTracker (a sensitive indicator of MMP) after 24 hours of dGuo
treatment. (A) In the histrograms, M1 indicates the percentage of cells maintaining MMP and
M2 indicates the percentage of cells losing MMP. (B) Graph represents the mean ±SD of the
percentage of PNP-/- (white bar) and PNP+/+ (black bar) thymocytes losing MMP from 3
separate experiments containing one or two mice in each (*p=0.003).
PNP-/-
*
M1= 95.48% M2= 4.34%
M1= 90.27% M2= 9.47%
M1
M1
M2M2
M1= 95.22% M2= 4.55%
M1= 78.95% M2= 20.95%
M1M1
M2M2
PNP+/+
Untreated dGuo treated
84
4.9. Changes in MMP occur before nuclear DNA fragmentation:
When apoptotic signals are initiated within the mitochondria, the disruption of MMP
occurs prior to nuclear DNA fragmentation (Ravagnan et al., 1999). Thus, I was interested to
confirm that indeed change in MMP occurs prior to nuclear DNA fragmentation in PNP
deficiency. Therefore, I measured MMP using a mitochondria specific dye, and DNA
fragmentation by the TUNEL assay in dGuo treated PNP-/- thymocytes at several time points.
The time points that I examined included;
6-8 hours treatment, which I hypothesized is prior to initiation of apoptosis,
12 hours treatment, which is when I hypothesized apoptosis is initiated,
16-24 hours of treatment, when apoptosis is fully developed in both the mitochondria and
nucleus (summarized in Figure 24G).
As expected, 6-8 hours after treating PNP-/- thymocytes with dGuo, there was no
increase in TUNEL+ cells or increase in the percentage of cells with abnormal MMP (Figure
24G). In marked contrast, at 12 hours, a significant higher percentage of cells had disrupted
MMP (Figure 24A is a representative example), without a significant change in DNA
fragmentation (Figure 24B), and the results of 4 separate experiments are summarized in
Figure 24C. At 16-24 hours, the percentage of cells with changes in MMP (Figure 24D) was
similar to the percentage of cells undergoing DNA fragmentation (Figure 24E), with the
results of 4 separate experiments summarized in Figure 24F. In conclusion, these results
suggest that apoptosis in PNP-/- thymocytes exposed to dGuo, is initiated in the mitochondria.
85
A) B)
Mitotracker TUNELMitotracker TUNEL
C)
12 hour treatment
0
2
4
6
8
10
12
14
UT dGuo
Perc
enta
ge o
f cel
ls
DNA fragmentationLoss in MMP (M2)
D) E)
*
86
F)
16 hour treatment
0
5
10
15
20
25
UT dGuo
Perc
enta
ge o
f cel
ls
DNA fragmentationLoss in MMP
G)
0
5
10
15
20
25
6 hours 12 hours 16 hours
Thymocytes treated with 12.5μM dGuo
Perc
enta
ge o
f cel
ls
DNA fragmentationLoss in MMP
Figure 24: Effects of dGuo on MMP and DNA fragmentation after 12 hours and 16 hours of treatment. PNP-/- thymocytes were incubated in RPMI supplemented with 5% FBS in the presence of 12.5μM dGuo for the indicated time points. At the indicated time points the cells were stained with Mitotracker to investigate loss of MMP and nuclear DNA fragmentation was assayed by the TUNEL technique. (A) and (D) demonstrate changes in MMP at 12 h and 16 h respectively. Percentages of cells are indicated above the bars. (B) and (E) demonstrate nuclear DNA fragmentation measured by the TUNEL assay at 12 h and 16 h respectively. Percentages of apoptotic cells are indicated above the bars. (C) and (F) summarize DNA fragmentation (black) and changes in MMP (white) in untreated and 12.5μM dGuo treated samples at 12 h (*p=0.001) and 16 h respectively. The data represents mean ± SD for 4 independent experiments at each time point with one or two mice in each. (G) Graph summarizes the DNA fragmentation (black) and changes in MMP (white) profiles following 6 h, 12 h and 16 h dGuo treatment of PNP-/- thymocytes (*p=0.001) (UT, untreated).
*
87
4.10. Caspase inhibition prevents dGuo induced DNA fragmentation but does not
prevent apoptosis of the cells.
To confirm that changes in MMP indeed occured before DNA fragmentation I used an
alternative strategy which determined the effect of caspase inhibition on MMP and DNA
fragmentation. When the apoptotic signals are initiated within the mitochondria, the
disruption of MMP does not involve the release and activation of caspases. Hence this event
is considered to be a caspase independent process. In contrast, when apoptotic signals
originate outside the mitochondria, then the subsequent disruption of MMP is a caspase
dependant process (Ravagnan et al., 1999). Consequently, if apoptosis originated within the
mitochondria then thymocyte apoptosis would not be affected by caspase inhibition.
However, if apoptosis signals originated due to nuclear DNA damage then the apoptosis
would be haulted by caspase inhibition. Therefore I examined the effect of caspase
inhibition (using the pan caspase inhibitor z-VADfmk) on dGuo induced apoptosis.
I first demonstrated that z-VADfmk prevented apoptosis (measured by the percentage
of Annexin+ cells) induced by dexamethasone (Dex) (Figure 25A), which is a well known
caspase-dependent apoptosis inducing agent. In contrast, z-VADfmk was unable to protect
thymocytes from dGuo induced apoptosis, as indicated by Annexin-V staining (Figure 25B).
Interestingly, z-VADfmk did prevent dGuo induced DNA fragmentation, which is
considered to be a late apoptotic event suggesting that in addition to the early mitochondria
induced apoptosis, dGuo also has a late, DNA fragmentation effect (Figure 25C).
88
A)
42.02
22.77
Annexin
Cel
l cou
nt
Dex Dex+ zVAD
42.02
22.77
Annexin
Cel
l cou
nt
Dex Dex+ zVAD
B)
Annexin
Cel
l cou
nt
dGuo dGuo + zVAD
17.48
25.61
Annexin
Cel
l cou
nt
dGuo dGuo + zVAD
17.48
25.61
C)
14.091.65
TUNEL
Cel
l cou
nt
dGuo dGuo + zVAD
14.091.65
TUNEL
Cel
l cou
nt
dGuo dGuo + zVAD
Figure 25: Effects of pan-caspase inhibition on dGuo induced apoptosis and nuclear DNA
fragmentation. PNP-/- thymocytes maintained in RPMI 1640 medium with 5% FBS, were
pre-treated for 30 minutes with 50μM z-VADfmk followed by the addition of (A) 3.75 μM
Dex or (B) and (C) 12.5μM dGuo. 20-24 hours later, apoptosis was measured by Annexin-V
staining (Figures A and B) and DNA fragmentation was measured by the TUNEL assay
(Figure C). Results are representative of 4 independent experiments with one or two mice in
each. The percentage of apoptotic cells are indicated above the bars.
89
CONCLUSION
In summary, these results demonstrate that a change in MMP occurs prior to nuclear DNA
fragmentation. Also, the loss in MMP induced apoptosis is resistant tocaspase inhibition
suggesting it is a caspase independent mechanism. Taken together, these findings suggest
that the accumulation of dGuo and its phosphorylated products in PNP-/- thymocytes
increased apoptosis in a mitochondrial mediated pathway.
90
CHAPTER 5: DISCUSSION
91
Discussion
PNP deficiency disrupts thymocyte development. However, the lack of availability of
thymus tissue from affected patients has prevented detailed analysis of the precise effects of
abnormal purine metabolism on thymocytes. PNP-/- mice displaying metabolic and immune
phenotype similar to the human disease have helped understand some of the mechanisms
associated with toxic purine metabolites, and hence were extensively used in this study.
Although PNP is a ubiquitous enzyme, its preferential effect on thymocytes and T-cells is not
surprising, as these cells, unlike many other cell types, are unique in that they undergo
constant selection and proliferation during their lifetime. To maintain a constant homeostasis
in these cells requires a balance in the nucleotide pools to compensate for the DNA damage
and repair induced during the process of selection and subsequent proliferation. This may be
be the most likely explaination for the selective toxicity of thymocytes to the effects of PNP
deficiency and thus, has been the area of focus for most researchers. However, preliminary
data from our lab and previous observations by others suggests involvement of cells beyond
the thymus. Neurological dysfunction has been reported in more than half of the patients
(Markert, 1991). Similar to PNP-deficient patients PNP-/- mice manifest with neuroloigal
abnormalities as tested by their inability to sustain on a spinning rotorod and increased
apoptosis indicated by in situ TUNEL staining on purkinje cells from PNP-/- mice
cerebellums (Grunebaum, unpublished). Also, PNP-/- skin fibroblasts from mice have been
seen to proliferate at a slower rate compared to normal controls as tested by the thymidine
uptake assay (Grunebaum, unpublished). Few patients also developed bone marrow
abnormalities possibly due to hypersensitivity to irradiation (Dror et al., 2004). Reduced
92
B- cell numbers and abnormal humoral function have been reported amongst a few patients
(Markert, 1987). These findings are indicative that PNP is essential in the maintainance and
functioning of many cells in the body.
We and others have found significant increases in the number of DN thymocytes in
the thymi of PNP-/- mice, which is particularly impressive when considering that the thymus
of these mice is atrophic with marked reduction in other thymocyte populations. This is
suggestive of abnormal maturation within the DN subset. Therefore our focus was initially
targeted towards studying the development of DN thymocytes. The significant increase in
the percentage and numbers of DN3 and reduction of DN4 and DP cells that I found in
freshly isolated thymocytes and ex-vivo cultures simulating PNP deficiency, clearly
demonstrate that maturation from the DN3 stage to the later stages are affected by PNP
deficiency. The limitations of these experiments as mentioned earlier, is that by the third
week of co-culture when most of the cells had developed into the later DN stages (DN3 and
DN4) and to the DP and SP stage, CD4+/CD8+ were not gated out to analyse the DN subsets.
This makes interpretation of results difficult as the “DN4” subset is really a heterogeneous
population consisiting of DN4, DP and SP cells. However, the significant increase in the
percentage of CD25+ cells at this stage was evidence enough for us to conclude that the delay
in thymocyte development in PNP deficiency begins at the DN3 stage. Another method to
study DN generation and expansion would be to carry out a dynamic analysis by a short term
BrdU pulse chase experiment. This would further show whether the early DNs are being
generated and proliferating at a proper rate. Moreover, investigating the exact stage of
developmental delay, i.e. whether between DN3 and DN4 or between DN4 and DP stage,
proved to be a challenge due to the absence of a distinct cell surface marker for the DN4
93
subset. The transition between DN3 to DN4 is a crucial check-point in thymocyte
maturation, and it has been estimated that 4 out of 9 DN3 cells die at this stage as a result of
failure to rearrange their TCRβ locus (Gärtner et al., 1999) and hence my results are not
unexpected. I therefore also determined the percentage of PNP-/- DN3 and DN4 thymocytes
that express intracellular TCRβ chain. I found a decrease in the percentage of DN3 cells
transiting to the DN4 stage by down regulating CD25 and upregulating intracellular TCRβ
expression in PNP-/- mice compared to controls, further indicating a developmental delay
between the DN3 and DN4 stage. However, the similar percentages of late DN3 cells in
PNP-/- and PNP+/+ mice indicate that TCRβ rearrangement is taking place normally. My
results are also in concordance to those reported in ADA deficiency, wherein ADA
deficiency did not seem to compromise TCRβ chain rearrangement in developing
thymocytes (Thompson et al.,2000). Collectively, these results suggest that rearrangement of
the TCRβ locus and expression of the TCRβ chain is not affected by PNP deficiency. It will
be beneficial to study the TCR-vβ repertoire in the periphery of PNP-/- mice compared to
controls by using a panel of antibodies to different TCR-vβ chains, which is a quantitative
approach. A qualitative and more in-depth analysis of TCR-vβ repertoire is by a PCR
technique such as spectra typing.
Another possibility for the decreased DN4 population could be related to sensitivity
of these cells to factors released by dying thymocytes, such as toxic purine metabolites,
particularly dGuo. Alternatively, the increased death of DN4 thymocytes could be due to
abnormal signalling through the TCR complex, as has been seen in mice deficient in Lck
(Levin et al., 1993), pTα deficient mice (Fehling et al., 1995), absence of ZAP-70 (Sugawara
et al., 1998), RAG-/- mice (Shinkai et al., 1992; Mombaerts et al., 1992) and knock down
94
models of pTα, CD3 ε, γ and ζ chains (Fehling et al., 1995; Haks et al., 1998; Dave et al.,
1997; DeJarnette et al., 1998). This explanation, which needs to be examined by future
experiments, is also supported by studies of peripheral PNP-deficient T-cells that showed
reduced responses and decreased IL-2 secretion, again implicating impaired intracellular
signalling (Arpaia et al., 2000; Dror et al., 2004; Toro et al., 2006).
In addition to the effect that PNP deficiency has on DN thymocytes described above,
striking reductions in the numbers and percentages of DP thymocytes in PNP-/- mice were
found. BrdU incorporation into the DNA of newly formed DP thymocytes in PNP-/- mice, as
well as in DP cells developed ex-vivo, demonstrated that proliferation of PNP-/- DP cells was
not impaired (refer Figures 16B, 16C). Moreover, the DP cells generated ex-vivo in PNP-
deficient conditions was reduced, although there is no selection ex-vivo. Thus I concluded
that proliferation and selection were not the causes for the decreased DP thymocytes. As an
alternative explanation, I studied apoptosis in PNP-/- DP thymocytes. An indication for
decreased cell survival was already provided by the rapid reduction in the percentage of
BrdU labelled DP thymocytes, which was not accounted for by reduced proliferation of DN
thymocytes or enhanced maturation into SP thymocytes. The increased apoptosis of PNP-/-
DP thymocytes was further confirmed by the significantly increased Annexin-V+ PI+ cells
among PNP-/- DP thymocytes. The presence of Annexin-V binding is also an indication that
the cells are truly undergoing apoptosis in contrast to other mechanisms of cell death such as
necrosis. Although the value of apoptosis in DP cells increased (on average) only from 1.5%
in normal control to approximately 3.5% among PNP-/- thymocytes, this increase was
statistically significant, was observed consistently among PNP-/- mice and was similar to that
reported previously (Arpaia et al., 2000). We suspect that the relatively low number of
95
apoptotic cells observed at a steady state among fresh thymocytes reflects both the ongoing
gradual effect that toxic purine metabolites have throughout the 6 week period from birth to
the time thymocytes were collected, and also the quick removal of apoptotic cells by the
abundant scavenger macrophages found in the thymus of PNP-/- mice. This is in
concordance to reports by Thompson et al., wherein a moderate increase in thymocyte
apoptosis was not detectable by PI staining or in situ TUNEL staining in ADA-deficient
cultures (Thompson et al., 2000).
Importantly, the increased apoptosis of PNP-/- DP thymocytes cultured ex-vivo
indicated that the apoptosis was not caused by external events, such as surge of
corticosteroids in PNP-deficient mice, unidentified infections, or enhanced signalling through
the Fas-receptor. The latter hypothesis was also supported by demonstrating that PNP-/- DP
thymocytes did not upregulate the surface expression of Fas receptor, further indicating that
the extrinsic pathway mediated by Fas expression may not be the contributing mechanism for
apoptosis. Moreover, death receptor signaling via Fas was eliminated as the apoptotic
mechanism in ADA deficiency, as lpr mice (mice homozygous for lymphoproliferation
spontaneous mutation (Faslpr)) remained sensitive to ADA deficiency. However, the same
group also showed that the use of a caspase-8 inhibitor prevented the apoptosis of ADA
deficient thymocytes, indicating the possible role of extrinsic apoptotic mechanisms via other
death receptor pathways such as tumor necrosis factor receptor (TNFR) family (Thompson et
al., 2003), which was not tested in this study. It is also possible that p53 may be involved,
since Benveniste and Cohen showed that p53-/- mice were resistant to apoptosis when
induced with ADA deficient conditions using dCF and dAdo (Benveniste and Cohen, 1995).
p53 can cause apoptosis by inducing transcription of pro-apoptotic Bcl-2 family members
96
such as Bax, PUMA and Noxa (Miyashita and Reed, 1995, Yu et al., 2001, Oda et al., 2000).
It was later shown by some preliminary studies that FTOCs from p53 knock-out mice were
susceptible to the consequences of ADA inhibition (Thompson et al., 2003). Of course, it is
still possible that a p53 independent mechanism may involve the pro-apoptotic Bcl-2 family
members. None of the above mentioned apoptotic pathways were addressed in my study
nevertheless, my preliminary experiments in attempt to study the extrinsic pathway, although
not conclusive, together with findings from ADA deficiency studies, lead us to narrow the
cause of apoptosis to an intrinsic mechanism.
We next asked whether the accelerated apoptosis of DP thymocytes in PNP
deficiency were caused by increased dGuo and its phosphorylation products, dGMP and
dGTP, or whether it was the depletion of PNP products (i.e. GTP). Both, increased dGuo
and dGTP concentrations, as well as decreased GTP concentrations were observed in PNP-
deficient thymocytes (Arpaia et al., 2000). Moreover, abnormal peripheral T-cell function
have been associated with increased dGuo and dGTP (Kicska et al., 2001; Bantia et al., 2003)
and reduced GTP. Here I demonstrated that apoptosis of PNP-/- thymocytes was induced by
dGuo at concentrations equivalent to those recorded in the plasma of PNP-deficient patients
(Hershfield and Mitchell, 2001) and mice (Arpaia et al., 2000). Moreover, apoptosis of
thymocytes could be prevented by restoring PNP activity within the cells, which provided
unequivocal evidence to the role of the abnormal purine homeostasis in induction of
apoptosis. Apoptosis was also induced in PNP-/- thymocytes by dGMP, an intermediate
metabolite between dGuo and dGTP, which further implicates the latter as the cause of
apoptosis. Thus our study shows that accumulation of dGuo in PNP deficiency, followed by
the conversion of dGuo to dGMP, and probably to dGTP, causes thymocyte apoptosis in
97
PNP-/- mice. We suspect that dGTP is the cause of apoptosis, as intracellular dGTP pools are
normally kept under tight regulation (Bjursell et al., 1980; Reichard et al., 1985), and PNP
inhibition using BCX-1777 in T-ALL, led to accumulation of dGuo in the plasma and dGTP
in circulating leukemia cells that were induced to die rapidly (Bantia et al., 2003). The
suspected role of dGTP in mediating the apoptosis of PNP-deficient thymocytes, also
suggested that the mitochondria was the cellular compartment most affected by PNP
deficiency, as dGTP formation is dependent on the function of dGK, which is a
mitochondrial enzyme (Johansson et al., 1996). Indeed, I found that accumulation of dGuo
led initially to the dissipation of mitochondrial membrane potential and that inhibition of
caspases, prevented dGuo induced nuclear DNA damage, but not mitochondrial damage. In
the experiments conducted to study changes in MMP, apoptosis induced by dGuo was not
seen immediately but rather took 12 hours to observe the toxic effects, possibly
recapitulating the gradual toxicity of purine metabolites to exert its effects on thymocytes in
vivo. The mitochondrial abnormalities that I found in PNP-/- thymocytes were similar to
previous observations in normal thymocytes treated with a PNP inhibitor (Arpaia et al.,
2000), in ADA-deficient thymocytes (Thompson et al., 2000), and in inherited thymidine
phosphorylase deficiency, which causes mitochondrial neurogastrointestinal
encephalomyopathy (MNGIE) disease (Nishino et al., 1999, Hiarano et al., 2006). Thus, our
findings suggest particular sensitivity of mitochondria to abnormalities in purine
homeostasis, which could be due to interference of purine metabolites with the down
regulation of anti-apoptotic molecules such as Bcl-2, as introduction of a Bcl-2 transgene
into PNP-deficient thymocytes increased their survival (Arpaia et al., 2000). Alternatively,
the depletion of mtDNA due to particular sensitivity of mitochondria to abnormal purine
98
metabolism is also suggested by the mitochondrial abnormalities reported in ADA deficiency
(Van De Wiele et al., 2002) and MNGIE (Lopez et al., 2009). Other observations also
provide additional evidence that thymocyte damage in PNP deficiency may originate in the
mitochondria. PNP-deficient cells were found to be sensitive to gamma irradiation but were
still able to replicate their nuclear DNA in response to mitogen in the presence of IL-2
(Arpaia et al., 2000). Also, over-expression of dGK in the mitochondria was found to
increase sensitivity to anti-cancer deoxyguanosine analogues in cancer cell lines (Zhu et
al.,1998).
In conclusion, by this study, I demonstrated that PNP deficiency affects murine
thymocyte development at two distinct stages, and my data also suggests the possibility of
different mechanisms affecting the two stages. The first is at the maturation of thymocytes
from the DN3 to DN4 stage. The precise mechanism leading to this defect is still not known
and requires further investigation. The second thymocyte population affected in PNP-
deficient mice is the DP population. PNP-/- DP thymocytes are undergoing apoptosis at an
increased rate. The apoptosis is induced by accumulating dGuo, and probably by the
secondary formation of dGTP, which interferes with mitochondrial integrity. Importantly,
restoration of PNP activity within the cells increased survival, suggesting that enzyme or
gene replacement therapy may help in correcting the thymocyte abnormalities in PNP-
deficient patients.
99
Future Directions 1) Identify the cause of decreased DN4 thymocytes in PNP-/- mice. Specifically, determine
intracellular signalling in DN3 and DN4 thymocytes. Alternatively, measure cell cycling
in PNP-deficient DN3 and DN4 thymocytes in vivo and ex-vivo.
2) Determine the contribution of GTP depletion in thymocyte abnormalities by GTP
supplementation.
3) Distinguish between the effects of dGuo and those of dGTP by preventing
phosphorylation of dGuo into dGTP, for example by cross-breeding PNP-/- mice with
dGK-deficient mice. If dGTP is the cause of apoptosis, then the double knockout will
prevent the dGuo-induced thymocyte apoptosis.
4) Assess the importance of anti-apoptosis mechanisms in PNP-/- thymocytes. Specifically,
determine whether over-expression of Bcl-2 protects PNP-/- deficient thymocytes from
dGuo induced apoptosis, by cross breeding of PNP-/- mice with Bcl-2 transgenic mice.
Alternatively, measure pro-apoptotic molecules associated with the mitochondrial
pathway, such as BAX and BAK.
5) Determine the sequence of events triggered by the mitochondrial damage that leads to
cell death. Specifically, release of cytochrome c from the intermembrane space of the
mitochondria to the cytoplasm, and activation of specific caspases, such as caspase 3 and
caspase 9.
6) Improve the ability to follow development of PNP-deficient thymocyte ex-vivo by
100
producing bone marrow stroma cells that do not express PNP, thereby more closely
simulating in vivo conditions in PNP-deficient patients. This can be done by inhibiting
PNP (using siRNA to PNP) in the OP9-DL1 cells.
7) Confirm that dGuo is indeed the metabolite responsible for thymocyte toxicity by
preventing its uptake into thymocytes. This can be achieved by the use of a nucleoside
transport inhibitor.
8) Explore the interactions (cross-talk) between thymocytes and the thymic epithelial cells,
which are also affected by the abnormal purine metabolism and possibly contribute to the
immune dysregulation observed in PNP-deficient patients and mice.
101
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