UNIVERSITY OF CINCINNATI
Date: 10-Nov-2009
I, Keith J Gaddie , hereby submit this original work as part of the requirements for the degree of: Doctor of Philosophy in Molecular, Cellular & Biochemical Pharmacology It is entitled: Structural Elements that Regulate Interactions between the
Extracellular and Transmembrane Domains of Human Nucleoside
Triphosphate Student Signature: Keith J Gaddie
This work and its defense approved by: Committee Chair: Terence Kirley, PhD Terence Kirley, PhD
Scott Belcher, PhD Scott Belcher, PhD
Paul Rosevear, PhD Paul Rosevear, PhD
John Maggio, PhD John Maggio, PhD
Ronald Millard, PhD Ronald Millard, PhD
11/23/2009 294
Structural Elements that Regulate Interactions between the Extracellular and Transmembrane Domains of Human Nucleoside Triphosphate Diphosphohydrolase 3
A dissertation submitted to the
Division of Research and Advanced Studies of the University of Cincinnati
in partial fulfillment of the requirements for the degree of
DOCTOR OF PHILOSOPHY (Ph.D.)
in the Department of Pharmacology and Cell Biophysics of the College of Medicine
2009
By Keith J. Gaddie
B.S. Fisk University, 2003
Committee Chairperson: Terence L. Kirley, Ph.D.
Abstract
The nucleoside triphosphate diphosphohydrolases (NTPDases) are a family of
constitutively expressed, endogenous nucleotidases, some of which regulate purinergic signaling
by divalent cation-dependent hydrolysis of nucleotides acting as agonists at purinergic receptors.
Due to the scarcity of specific inhibitors and genetically modified animals, the functions of most
individual NTPDases are poorly understood and still under investigation. However, the
NTPDases have been implicated in many biological and physiological processes, including
secretion, cell adhesion, pain perception, cancer and malignant transformation, adenosine
recycling, and platelet aggregation.
Membrane-bound NTPDase3 expressed on the cell surface has a large extracellular
domain, a “linker region”, and a transmembrane domain. Evidence exists for interactions
between the transmembrane domain and the active site lobes that govern the function of rat
NTPDase1. However, the specific mechanism(s) regulating this cross-talk for NTPDases, as well as how the N- and C-terminal transmembrane helices in NTPDase3 interact within and
between monomers to mediate oligomerization and modulation of enzymatic activity is still
unknown.
The roles of the conserved proline residues of human NTPDase3, located in the “linker
region” that connects the N- and C-terminal transmembrane helices with the extracellular active
site, were examined by proline to alanine substitutions coupled with single cysteine substitutions
strategically placed in the transmembrane domain to serve as cross-linking “sensors” of helical
interactions. Mutation of several proline residues resulted in decreased nucleotidase activities
and some “uncoupled” the effect of ATP binding on TMD movements. The data suggest a role
for proline residues 53 and 481 in the linker region of human NTPDase3 for “coupling”
iii nucleotide binding and hydrolysis at the enzyme active site to movements and/or rearrangements of the transmembrane helices necessary for optimal nucleotide hydrolysis.
To investigate the structural/functional roles of the conserved polar residues in the transmembrane helices of human NTPDase3, each was singly mutated to alanine. All mutants were properly glycosylated and had specific activities similar to wild-type, except Q44A. The
Q44A mutation decreased specific activities by approximately 50% - 70%, and nearly eliminated
Triton X-100 detergent inhibition. The same conserved polar residues were mutated to cysteine, singly and in pairs, to allow a disulfide cross-linking strategy to map potential inter- and intra- molecular hydrogen bond interactions. The results support the centrality of Q44 for the strong inter-molecular interactions driving the association of the N-terminal domains of two NTPDase3 monomers in a dimer, while S39 and T495 may contribute to helical interactions involved in forming higher order oligomers. These results suggest a model for putative hydrogen bond interactions of the conserved polar residues in the transmembrane domain of native, dimeric
NTPDase3 that are important for protein expression, activity, and susceptibility to membrane perturbations.
This dissertation provides additional insights to the structural elements that regulate the enzymatic activity of the NTPDases. With the recent elucidation of the crystal structure of the extracellular portion of rat NTPDase2, the major remaining structural questions regarding the cell membrane NTPDases are how the TM helices interact and how these interactions are coupled to modulation of enzyme activity and NTPDase function. This dissertation addresses these questions for human NTPDase3, and suggests that similar structure-function relationships are relevant to the family of NTPDases controlling purinergic signaling.
iv Acknowledgements
My graduate experience was a big adjustment, a learning process of my strengths and
weaknesses, and a time where I was always pushed to excel to new heights. Luckily, I was
surrounded by talented and accomplished individuals who cared about my scientific future and
were willing to share their expertise as I traveled this challenging journey. First, I would like to
express my sincere gratitude to my mentor and advisor, Dr. Terry Kirley. Dr. Kirley possesses
so many qualities of a great scientist, disciplined, the ability to pay a great attention to detail, and excellent time management skills. In addition, through our casual conversations and some observation, he is an excellent family man.
Dr. Kirley was very instrumental in developing my skills as a research scientist and he always encouraged me to work and think in an independent manner, but more importantly, to
pay attention to the small details. This has allowed me to grow as a scientist and to think critically about my data. I am truly indebted to Dr. Kirley for all his hard work, support, and guidance. In addition, I owe thanks to my committee members, Drs. Scott Belcher, John
Maggio, Ronald Millard, and Paul Rosevear, for providing constructive criticism and insight over the years. Their input and suggestions were instrumental in shaping the progression of my research projects.
I would also like to thank all the members of the Kirley laboratory for their assistance and guidance throughout the years. When I first joined the laboratory, Dr. Patrick Crawford was very instrumental in teaching me the basic laboratory protocols and worked closely with me on his project regarding the NTPDase3 splice variant, NTPDase3β. Dr. Vasily Ivanenkov instructed and motivated me to interpret my data from several points of view. In addition, he discovered that Tween 20 stimulated NTPDase3 activity and I applied in my dissertation
v research. I would also like to thank Dr. Min Yang and Clifford Cookman for all of their assistance during their time in the laboratory.
I would like to thank everybody in the department of Pharmacology, especially Nancy
Thyberg, Damita Jo Harris, and Carol Ross for always handling the seen and unseen obstacles of graduate school from a paperwork point of view.
Finally, I am so thankful for having a supportive family (Godsister’s Stephanie Ford and
Janelle Henson) and friends (Linda Smith and Dr. Bradley Sheares) who never gave up on me along the way. I would not have successfully completed my studies if it were not for my mom
(Pamela J. Gaddie) and dad’s (Keith Taylor) unconditional love, my aunts’ (Brenda, Dee Dee,
LaVada, and Margo) unwavering support, as well as my best friend’s (Derrick G. Demmons) and roommate’s (Mark A. Edmonson) encouraging words.
vi Table of Contents
Page
Abstract iii
Acknowledgements v
Table of contents vii
List of figures and tables xiii
List of abbreviations xvii
Chapter I: Introduction 1
1. Evolutionary Appearance of Nucleosides 1
1.1. Nucleotide structure 2
1.2. Nucleotide synthesis 2
1.2.1. De novo purine nucleotide synthesis 2
1.2.2. De novo pyrimidine nucleotide synthesis 4
2. The Concept of Purinergic Receptor Signaling 8
3. ATP: The Neurotransmitter 12
4. Purinergic Receptors 15
4.1. P1 (Adenosine) Receptors 18
4.2. P2X Receptors 18
4.3. P2Y Receptors 19
5. Functional Significance of Purinergic Receptor Signaling 20
6. Ecto-nucleotidases 21
6.1. Alkaline phosphatase (AP) 21
6.2. Ecto-5’-nucleotidase/CD73 23
vii 6.3. Ecto-nucleotide phosphodiesterase/pyrophosphatase (E-NPP) 23
6.4. Nucleoside Triphosphate Diphosphohydrolase (NTPDase) 24
7. Molecular Cloning, Sequencing, Expression, and Characterization of the NTPDases 26
7.1. The membrane-bound, cell surface NTPDases (NTPDase1-3 and 8) 26
7.2. The intraluminal NTPDases (NTPDase4-7) 27
8. The Biochemical and Structural Characterization of the NTPDases 30
8.1. Enzymology of the NTPDases 30
8.2. Inhibitors of the NTPDases 31
8.3. Site-directed mutagenesis and computational comparative protein modeling of the
membrane-bound, cell surface NTPDases 36
8.3.1. The structural features of the extracellular domain of the NTPDases 38
8.3.2. The structural features of the transmembrane domain of the NTPDases 41
9. Functional Significance of the NTPDases 44
10. Dissertation Scope and Objectives 46
Chapter II: Methods and Materials 49
1. General Methods and Materials 49
1.1. Purification of oligonucleotides for site-directed mutagenesis 49
1.2. Site-directed mutagenesis of NTPDase3 50
1.3. Bacterial transformation and inoculation of PCR products 50
1.4. Isolation of DNA from XL1-Blue Supercompetent Cells 51
1.5. Restriction enzyme digestion of DNA 52
1.6. Transient transfection of COS-1 cell membranes 52
viii 1.7. COS-1 cell crude membrane preparations 53
1.8. Protein concentration of COS-1 cell crude membrane preparations 54
1.9. Specific nucleotidase activity of NTPDase3 expressed in COS-1 cells 55
1.10. Malachite green procedure for nucleotide assays 55
1.11. SDS-PAGE and Western blotting 56
1.12. Deglycosylation 57
2. Project Specific Methods and Materials 58
2.1. NTPDase3 “linker region” proline residue mutagenesis 58
2.1.1. List of oligonucleotides/mutants made 58
2.1.2. Oxidative cross-linking in the presence and absence of substrate 59
2.2. NTPDase3 transmembrane domain polar residue mutagenesis 59
2.2.1. List of oligonucleotides/mutants made 59
2.2.2. Triton X-100 nucleotidase assays 61
2.2.3. Oxidative cross-linking and alkylation of cysteine 61
2.2.4. NTPDase3 nucleotidase assays in the presence of Tween 20 62
2.3. NTPDase3 alternatively spliced variant (NTPDase3β) 63
2.3.1. IMAGE clone encoding NTPDase3β 63
2.3.2. List of oligonucleotides/mutants made 63
2.4. NTPDase3 single nucleotide polymorphisms (SNPs) 64
2.4.1. Source of SNPs 64
2.4.2. List of oligonucleotides/mutants made to mimic known SNPs 64
2.4.3. Treatment of wild-type and mutant NTPDase3 with Concanavalin A 65
ix Chapter III: Results 66
1. Mutagenesis of NTPDase3 “linker region” proline residues 66
1.1. Rationale for selection and analysis of NTPDase3 “linker region” proline residues 66
1.2. Characterization of the NTPDase3 proline to alanine mutants 69
1.3. Rationale for selection of transmembrane “sensor” mutations and experimental
approach 76
1.4. Characterization of the TM “sensor” mutations (V42C and G489C NTPDase3) 77
1.5. Characterization of proline to alanine mutants in the V42C or G489C NTPDase3
background 81
2. Mutagenesis of NTPDase3 transmembrane domain polar residues 87
2.1. Rationale for selection and analysis of TM polar residue mutants 87
2.2. Characterization of the NTPDase3 polar residue to alanine mutants 90
2.3. Characterization of the polar residues singly mutated to cysteine 96
2.4. Paired cysteine mutagenesis and generation of a model incorporating the inter-
and intra-molecular hydrogen bonding pattern of the conserved polar residues
in the transmembrane helices of NTPDase3 100
2.5. Tween 20 stimulation of NTPDase3 107
3. NTPDase3 alternatively spliced variant 117
3.1. Rationale and experimental approach 117
3.2. NTPDase3β alternatively spliced isoform 118
3.3. Expression and enzymatic activities of NTPDase3 isoforms 122
3.4. Co-transfection of NTPDase3a and NTPDase3β 123
4. NTPDase3 single nucleotide polymorphisms (SNPs) 127
x 4.1. Rationale and experimental approach 129
4.2. Characterization of NTPDase3 SNPs 129
Chapter IV: Discussion 135
1. Regulation of the enzymatic activity of membrane-bound, cell surface NTPDases 135
2. Structural and functional roles of specific structural elements in NTPDase3 137
2.1. The conserved proline residues are important components of the linkage between
the active site and transmembrane domain movements 137
2.2. The conserved polar residues stabilize transmembrane domains and promote
oligomerization 139
2.3. NTPDase3β: A possible modulator of nucleotidase activity and purinergic receptor
signaling 144
2.4. Biochemical characterization of NTPDase3 single nucleotide polymorphisms 145
3. Summary and Future Directions 145
3.1. Determine if a polyproline II helix exists in the linker region of NTPDase3 and
identify the specific amino acids involved in its formation/stability 146
3.2. To further define the structural motifs that mediate oligomerization via the
transmembrane domain of the NTPDases 147
3.3. Further biochemical characterization of the single nucleotide polymorphisms of
NTPDase3 148
Chapter V: References 149
xi Appendix: Publications and Manuscripts under Review 175
xii List of Figures and Tables
Figures: Page
1. The structural elements of the most common nucleotides 3
2. De novo synthesis of purine nucleotides 5
3. Synthesis of AMP and GMP from IMP 6
4. De novo synthesis of pyrimidine nucleotides 7
5. Timeline of highlights of ATP signaling 9
6. General methods of nucleotide release 14
7. Purinergic signaling pathways 16
8. Major extracellular nucleotide hydrolyzing enzymes 22
9. Intracellular nucleotidase control of protein folding and processing 29
10. The three structural regions of the NTPDases 37
11. Multiple sequence alignment for portions of NTPDase sequences between the N-
and C-terminal transmembrane helices and the extracellular domain containing
the enzyme active site 67
12. Alanine substitution at P53, P472, P475, and P485 caused a decrease in
expression level in mutant NTPDase3 enzymes 70
13. P53A and P481A NTPDase3 mutants exhibit a decrease in nucleotidase
activities as compared to the C10S NTPDase3 background mutant 72
14. Alanine substitution of NTPDase3 proline residues are properly processed and
trafficked to the cell membrane 73
15. Double alanine substitution of the proline residues cause a decrease in
xiii expression level in mutant NTPDase3 enzymes 74
16. P53A/P481A and P484A/P485A NTPDase3 mutants exhibit a decrease in
nucleotidase activities as compared to the C10S NTPDase3 background mutant 75
17. Cysteine substitution at V42 and G489 in NTPDase3 exhibits similar nucleotidase
activities as the C10S/C501S/C509S NTPDase3 background mutant 78
18. ATP binding at the extracellular domain decreased the cross-linking efficiency
of V42C and G489C NTPDase3 mutants 79
19. Oxidative cross-linking decreases ATP hydrolysis 80
20. Ca-ATPase activity of the V42C proline to alanine mutants 82
21. Ca-ATPase activity of G489C proline to alanine mutants 83
22. P53A and P481A abolish the ATP-induced decrease in cross-linking efficiency 84
23. P481A abolishes the ATP-induced decrease in cross-linking efficiency 86
24. Rationale for NTPDase3 mutations generated and analyzed in this study 88
25. Alanine substitution of the polar residues cause a decrease in expression level,
but similar nucleotidase activities in most mutant NTPDase3 enzymes 91
26. Alanine substitution of NTPDase3 polar residues are properly processed and
trafficked to the cell membrane 94
27. Alanine substitution at Q44 eliminates the detergent inhibition of Triton X-100
on NTPDase3 enzymatic activities 95
28. Characterization of NTPDase3 polar residues to single cysteine mutants 97
29. Specific activities of NTPDase3 double cysteine mutants 101
30. Oxidative cross-linking of NTPDase3 polar residue hydrogen bonding pair
(double cysteine) mutants 103
xiv 31. Model of the transmembrane domain helical interactions in the NTPDase3 dimer 105
32. NTPDase3 S39C/Q44C, T41C/Q44C, and T495C/Q44C mutant enzymatic activities 108
33. Analysis for higher order oligomeric structure by oxidative cross-linking of
S39C/Q44C, T41C/Q44C, and T495C/Q44C NTPDase3 double mutants 109
34. The stimulatory effect of Tween 20 is specific for NTPDase3 and conserved among
species 110
35. Tween 20 promotes oligomerization mediated by the TMDs of NTPDase3 112
36. Detergents Tween 20 and Triton X-100 do not change the enzymatic activity of
sNTPDase3 115
37. Effect 0.1% Tween 20 or Triton X-100 on CuPhen cross-linking efficiency of the
TMDs of a mutant NTPDase3 116
38. cDNA and translated amino acid sequences of NTPDase3β, the truncated splice
variant of NTPDase3 119
39. A diagram of the exon map of the human NTPDase3 gene representing the
splicing events that occur 121
40. Western blot analysis of co-transfection experiments using the KLH1 or KLH11
antibody 125
41. Percent activity of co-transfections normalized per molecule active protein 128
42. Enzymatic activity of NTPDase3 SNPs 130
43. ConA stimulatory effect on NTPDase3 SNPs 131
44. Triton X-100 detergent effect on NTPDase3 SNPs 132
45. Tween 20 stimulatory effect on NTPDase3 SNPs 133
xv Tables: Page
1. Members of the E-NTPDase family 25
2. Enzymatic characterization of NTPDase1-3 and 8 32
3. Results of co-transfection of different NTPDase3 proteins in COS cells 124
xvi List of Abbreviations
3-(N-morpholino)propanesulfonic acid (MOPS)
4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES)
5-phosphoribosyl α-diphosphate (PRPP)
5’-aminoimidazole ribonucleotide (AIR) acetylcholine (ACh) adenosine (Ado) adenosine 5’-diphosphate adenosine 5’-monophosphate (AMP) adenosine 5’-triphosphate (ATP) alkaline phosphatases (AP) apyrase conserved regions (ACRs) arylazidoaminopropionyl-ATP (ANAPP3)
ATP-binding cassette (ABC) proteins
cyclic adenosine monophosphate (cAMP)
cyclic guanosine monophosphate (cGMP)
concanavalin A (Con A)
copper phenanthroline (CuPhen)
cytidine 5’-monophosphate (CMP; cytidylate)
cytidine 5’-triphosphate (CTP)
deoxyribonucleic acid (DNA)
disuccinimidyl suberate (DSS)
dithiothreitol (DTT)
xvii ecto-nucleoside triphosphate diphosphohydrolases (eNTPDases) ecto-nucleotide phosphodiesterase/pyrophosphatases (E-NPPs) endoglycosidase H (EndoH) endothelium-derived relaxing factor (EDRF) ethylenediaminetetraacetic acid (EDTA) formylglycinamide ribonucleotide (FGAR) formylglycinamidine ribonucleotide (FGAM) gamma-amino butyric acid (GABA) glycinamide ribonucleotide (GAR) guanosine 5’-diphosphate (GDP) guanosine 5’-monophosphate (GMP) guanosine 5’-triphosphate (GTP) hypoxanthine (Hyp) inorganic phosphate (Pi) inosine (Ino) inosine 5’-diphosphate (IDP) inosine 5’-monophosphate (IMP; inosinate) inosine 5’-triphosphate (ITP)
International Union of Pharmacology (IUPHAR) isotonic wash buffer (IWB) maleimide polyethylene glycol-5000 (MalPEG) multi-drug resistance (MDR) glycoprotein
N-ethylmaleimide (NEM)
xviii N-succinylo-5-aminoimidazole-4-carboxamide ribonucleotide (SAICAR) nicotinamide adenine dinucleotide (NAD+; NADH) non-adrenergic, non-cholinergic (NANC) noradrenaline (NA) or norepinephrine (NE) orotidine 5’-monophosphate (orotidylate; OMP) p-chloromercuriphenylsulfonate (pCMPS) phosphatidylinositol (PI) phospholipase C (PLC) plasma cell differentiation antigen-1 (PC-1) polymerase chain reaction (PCR) purine nucleoside phosphohydrolase (PNP) real time-polymerase chain reaction (RT-PCR) ribonucleic acid (RNA) ribose phosphate pyrophosphokinase (PRPP synthetase) single nucleotide polymorphisms (SNPs) sodium azide (NaN3) sodium dodecyl sulfate (SDS) tissue homogenization buffer (THB) transmembrane (TM) transmembrane domain (TMD) uridine 5’-diphosphate (UDP) uridine 5’-monophosphate (UMP; uridylate) uridine 5’-triphosphate (UTP)
xix xanthine 5’-monophosphate (XMP ; xanylate)
xx Chapter I: Introduction
1. Evolutionary Appearance of Nucleotides
Nucleotides are the basic structural units that form ribonucleic acid (RNA) and deoxyribonucleic acid (DNA), the compounds that contain the genetic instructions used in the development and functioning of all known living organisms. Nucleotides also play important roles in cellular metabolism by serving as sources of chemical energy, participate in cell signaling, and are utilized as cofactors of enzymatic reactions. In the early stages of the evolution of life, it is thought that thermal processes mediated phosphorylation of nucleosides and the formation of pyrophosphate bonds (1). These resulting compounds became the starting materials for nucleotide synthesis in terrestrial and aquatic life (2). Since adenosine 5’- triphosphate (ATP) has a propensity to bind the metal ion magnesium, which promotes dephosphorylation and the generation of energy, it is thought to be particularly suitable for early development. Inosine 5’-triphosphate (ITP) and guanosine 5’-triphosphate (GTP) also show a metal ion-promoted dephosphorylation, but their reaction rates are lower. The pyrimidines were also available on the early earth. It is suggested they were incorporated into living systems in a passive way, possibly directed by the purines (3). Therefore, purines such as ATP may have played a crucial and active role in early evolution. The formation of ATP from adenosine 5’- monophosphate (AMP), or other nucleosides, seems to have paved the way for the synthesis of oligonucleotides, then RNA, and eventually DNA, a molecule more reliable for storing genetic information (4).
1 1.1. Nucleotide structure
As shown in Figure 1, a nucleotide is composed of a nitrogenous base, a five carbon
sugar (either ribose (RNA) or deoxyribose (DNA)), and one to three phosphate groups (at either
the 2’, 3’, or 5’-carbon atom of the sugar moiety, the 5’-carbon site occurring most commonly).
Together, the nitrogenous base and the sugar constitute a nucleoside. Nucleosides can contain
two different types of nitrogenous bases, purines (adenine and guanine) or pyrimidines (cytosine,
uracil, and thymine).
1.2. Nucleotide synthesis
Nucleotides can be synthesized in vitro and in vivo. In vivo nucleotides are synthesized
via the de novo pathway or the salvage pathway. De novo synthesis of nucleotides begins with
their metabolic precursors: amino acids, ribose 5’-phosphate, CO2, and NH3. Salvage pathways
recycle free bases and nucleosides released from nucleic acid breakdown. Both pathways are important in cellular metabolism.
1.2.1. De novo purine nucleotide synthesis
The de novo pathway for purine nucleotide synthesis seems to be identical in nearly all living organisms. The two parent purine nucleotides are AMP and guanosine 5’-monophosphate
(GMP), containing the purine bases adenine and guanine. Eleven enzymatic reactions lead to the first intermediate with a complete purine ring,
2
Figure 1. The structural elements of the most common nucleotides. Nucleotides have three characteristic components: 1. a nitrogenous base, 2. a pentose sugar, and 3. a phosphate. The molecule without the phosphate group is called a nucleoside. The nitrogenous bases are derivatives of two parent compound classes, pyrimidines (cytosine, uracil, and thymine) and purines (adenine and guanine). The base of the nucleotide is joined covalently (at N-1 of pyrimidines and N-9 of purines) in an N-β-glycosyl bond to the 1’ carbon of the pentose, and the phosphate is most commonly esterified to the 5’-carbon atom.
3 inosine 5’-monophosphate (IMP; inosinate) (Figure 2). From IMP, AMP and GMP are synthesized in separate two step reactions (Figure 3).
1.2.2. De novo pyrimidine nucleotide synthesis
The common pyrimidine nucleotides are cytidine 5’-monophosphate (CMP; cytidylate) and uridine 5’-monophosphate (UMP; uridylate), which contain the pyrimidines cytosine and uracil. During de novo pyrimidine nucleotide synthesis (Figure 4), the six-member pyrimidine ring is made first and then attached to ribose 5’-phosphate. Pyrimidine nucleotide synthesis begins with the formation of N-carbamoylaspartate from aspartate and carbamoyl phosphate and yields orotate through subsequent reaction steps. Orotate reacts with 5-phosphoribosyl α- diphosphate (PRPP) to yield orotidine 5’-monophosphate (orotidylate; OMP) which is decarboxylated to form UMP. From UMP, other pyrimidine nucleotides are derived through kinase reactions to yield uridine 5’-triphosphate (UTP). Cytidine 5’-triphosphate (CTP) is derived from the conversion of UTP by cytidylate synthetase. Upon the accumulation of excess concentrations of CTP, pyrimidine nucleotide synthesis is also regulated by a feedback inhibition, where CTP inhibits the enzyme aspartate transcarbamoylase, the first reaction in the process.
4
5 5-Phosphoribosyl-1-pyrophosphate (PRPP) N -CAIR Glutamine 1 Glutamate 7
PPi 5-Amino-4-carboxyamino- 5-Phospho-β-D-ribosylamine Glycine Imidazole ribonucleotide 2 ATP Aspartate ATP ADP + Pi 8 ADP + Pi Glycinamide ribonucleotide (GAR) N-Succinylo-5-aminoimidazole-4- 10 N -Formyl H4 carboxamide ribonucleotide (SAICAR) 3 folate
Formylglycinamide ribonucleotide (FGAR) 9 Fumarate Glutamine 5-aminoimidazole-4-carboxamide 4 Glutamate ATP ribonucleotide (AICAR) ADP + Pi 10 10 N -Formyl H4 Formylglycinamidine ribonucleotide (FGAM) folate ATP N-Formylaminoimidazole-4- 5 ADP + Pi carboxamide ribonucleotide (FAICAR) H20 5-Aminoimidazole ribonucleotide (AIR) 11 H2O HCO3 ATP 6 Inosine-5’-monophosphate (IMP); ADP + Pi 5 5 Inosinate N -Carboxyaminoimidazole ribonucleotide (N -CAIR)
Figure 2. De novo synthesis of purine nucleotides. Formation of 5-phosphoribosylamine, the
product of the first reaction, is the first committed step in purine synthesis. The enzymes are: 1.
glutamine-PRPP amidotransferase, 2. GAR synthetase, 3. GAR transformylase, 4. FGAR
amidotransferase, 5. FGAM cyclase (AIR synthetase), 6. N5-CAIR synthetase, 7. N5-CAIR mutase, 8. SAICAR synthetase, 9. SAICAR lyase, 10. AICAR transformylase, and 11. IMP synthase.
5 Fumarate Adenosine-5’- GDP + Pi GTP Adenylosuccinate monophosphate (AMP); 2 Adenylate Aspartate 1
Inosine-5’-monophosphate (IMP); Inosinate
H2O NAD+ NADH + H+ 3 Gln Glu ATP AMP + PPi Xanthine-5’- Guanosine-5’- monophosphate (XMP); monophosphate (GMP); Xanylate 4 Adenylate H2O
Figure 3. Synthesis of AMP and GMP from IMP. The reaction enzymes are: 1. adenylosuccinate synthetase, 2. adenylosuccinate lyase, 3. IMP dehydrogenase, and 4. XMP- glutamine amidotransferase.
6 Aspartate Carbamoyl aspartate Phosphate transcarbamoylase Pi N-Carbamoylaspartate dihydroorotase
H2O L-Dihydroorotate + dihydroorotate NAD
dehydrogenase NADH + H+ Orotate orotate PRPP phosphoribosyl- transferase PPi Orotidylate orotidylate decarboxylase CO2 Uridylate (UMP) kinases 2 ATP 2 ADP Uridine-5’-triphosphate Gln cytidylate Glu synthetase ATP ADP + Pi Cytidine-5’-triphosphate (CTP)
Figure 4. De novo synthesis of pyrimidine nucleotides. The pyrimidine is constructed from carbamoyl phosphate and aspartate. Then the ribose 5’-phosphate is added to the completed pyrimidine ring by orotate phosphoribosyltransferase.
7 2. The Concept of Purinergic Signaling
In 1929 (Figure 5), ATP was identified in muscle extracts independently by Karl
Lohmann at the Kaiser Wilhelm Institute for Biology in Berlin and by Cyrus Hartwell Fiske and
Yellagaprada Subbarow at Harvard University (5, 6). This led individuals to believe ATP was primarily involved in muscle contraction. In the years following the discovery of ATP it became clear that the three chemically and functionally different parts of the molecule were synthesized by independent metabolic routes and that ATP was generated during the breakdown of glucose to lactic acid (anaerobic glycolysis), within the citric acid cycle, and predominantly during aerobic oxidation (oxidative phosphorylation) (7). The recognition that ATP was utilized for energy requiring functions allowed Lippman to introduce his general hypothesis for energy transfer in living cells (8). As a result, ATP was soon recognized as the universal source of readily available chemical energy for all living cells.
It was the essential intracellular roles of ATP that led to considerable resistance to the concept that ATP could exert any extracellular physiological action. The main argument was that cells would not release a molecule so fundamental to life under normal physiological conditions. This idea was reinforced by the size and charge of ATP, indicating it could not cross the plasma membrane by simple diffusion (9, 10). However, the biological effects of extracellular nucleotides were discovered early. In 1929, Drury and Szent-Gyorgyi at the
University of Cambridge published the first paper demonstrating the extracellular actions of
AMP and adenosine on the heart and coronary blood vessels (11), causing negative chronotropic
effects and dilation of coronary vessels. Several studies followed, confirming the role
8
2010 2008 sol NTPDase2 crystal structure 2005 First 3-D NTPDase3 model proposed 2000 1998 NTPDase3 identified 1996 NTPDase1 identified 1993 P2Y receptor, 1994 P2X receptor 1990 1990 5’-nucleotidase identified 1985 P2X and P2Y receptors differentiated 1985 AP and NPP1 identified 1980
1974 ATP in cholinergic synaptic vessicles 1970 1970 ATP recognized as a neurotransmitter
1960 1957 Term “ecto-ATPase” coined
1950 1948 Chemical synthesis of ATP 1945 Extracellular hydrolysis of ATP 1940
1934 Biological activity of ATP 1930 1929 ATP discovered
Figure 5. Timeline of highlights of ATP signaling. ATP was discovered in 1929 and its biological activity recognized in 1934. In 1957, the term “ecto-ATPase” was coined. In 1970,
ATP was recognized as a neurotransmitter and in the mid 1980s and 90s the purinergic receptors and the ecto-nucleotidases were discovered. In 1998, NTPDase3, the enzyme used throughout this dissertation, was cloned, sequenced, expressed, and characterized by our laboratory (12).
9 of purine nucleotides in the cardiovascular system (13). In 1934, J.H. Gillespie from Queen’s
University in Belfast extended these studies, providing evidence that extracellular ATP affects a
variety of tissues including heart, vasculature, and smooth muscle (14). Non-vascular effects of
purine nucleotides were later established in the gut, bladder, vagina, and other visceral organs
and vascular beds (15, 16).
Initially, adenosine (17) or AMP (18) were the purines thought to mediate the
extracellular actions of nucleotides, which are both degradation products of ATP. Studies soon established the length of the phosphate chain conferred differences in activity and ATP often had stronger effects than the other purinergic compounds (19, 20). In 1948, ATP was chemically synthesized by Alexander Todd, and scientists no longer isolated ATP from muscle extracts for use in their experiments (21). More detailed reports followed on the hydrolysis of ATP and ADP by intact yeast cells (22), human erythrocytes (23), ascites tumor cells (24), and nucleated avian erythrocytes (25). This led to the term “ecto-ATPase”, which was coined in 1957 by W.A.
Engelhardt from the Academy of Sciences in Moscow, during his findings of surface-located
ATPase activity on avian blood cells (25) as well as he introduced the terms “ecto-enzyme” and
“ecto-apyrase” (25, 26).
For many years the concept of neurotransmission was dominated by the dogma that each neuron releases only a single transmitter, known as “Dale’s Principle” (27). This arose from a widely adopted misinterpretation of his suggestion that the same neurotransmitter was stored in and released from all terminals of a single neuron. However, Dale’s Principle did not specifically exclude the possibility that more than one transmitter may be associated with the same neuron. In 1953 and 1954, Holton observed that ATP could be released from stimulated sensory neurons, which raised the possibility that ATP may be a neurotransmitter at sensory
10 nerve endings (28, 29). Soon after, it was recognized that adenosine and ATP were co-stored
with catecholamines in adrenal chromaffin cells (30, 31). Subsequently, ATP was shown to be
co-released with adrenaline from chromaffin cells (32, 33). In 1970, Geoffrey Burnstock, then working at the University of Melborne, proposed the existence of a “non-adrenergic, non- cholinergic” (NANC) regulation of smooth muscle contraction in the gut and bladder, in which
ATP was identified as the principal transmitter (34). Soon after, Burnstock and colleagues
provided the first evidence for sympathetic co-transmission involving ATP together with
noradrenaline (NA). In that study, they demonstrated that stimulation of periarterial sympathetic
nerves leads to release of tritium from guinea-pig taenia coli pre-incubated in 3H-adenosine
(which is taken up and largely converted to 3H-ATP), and that the release of both tritium and NA
was blocked by guanethidine (35). This led Burnstock and his colleagues to propose the use of
the term “purinergic” (36), and demonstrate that ATP fully fulfills the criteria for a
neurotransmitter as well as a co-transmitter.
Since the mid 1980’s, scientists have identified and functionally characterized many
purinergic (and pyrimidinergic) receptors (37) and the ecto-nucleotidases that regulate these
receptors by hydrolysis of their extracellular nucleotide agonists. We now know that not only
ATP, but also adenosine 5’-diphosphate (ADP), UTP, uridine 5’-diphosphate (UDP),
dinucleoside polyphosphates, UDP glucose, and even nicotinamide adenine dinucleotide (NAD+) activate nucleotide receptors and act as signaling molecules between cells. These signaling molecules are rapidly degraded to terminate their action on purinergic receptors by ecto- nucleotidases. The first ecto-nucleotidases cloned were an alkaline phosphatase (38), nucleotide phosphodiesterase/pyrophosphatase 1 (NPP1) (39), followed by ecto-5’-nucleotidase (40). In
1996, the first nucleoside triphosphate diphosphohydrolase (NTPDase), human CD39/NTPDase1
11 was identified (41), and in 1998, human NTPDase3, the enzyme studied in this dissertation, was
identified by our laboratory (12). Now it is widely recognized that purinergic signaling is an
evolutionarily ancient system involved in many neuronal and non-neuronal mechanisms, in both
short-term and long-term (trophic) events (42).
3. ATP: The Neurotransmitter
Intracellular ATP is primarily utilized to drive energy-requiring processes such as active
transport, cell motility, and biosynthesis, whereas extracellular ATP functions as a signaling
molecule at purinergic receptors. There is a multi-step cascade of ATP “release-signaling-
inactivation” that includes: 1. transient release of ATP and ADP; 2. triggering of signaling events
through a series of ligand-gated ionotropic (P2X) and metabotropic (P2Y) receptors; 3.
nucleotide breakdown by membrane-bound and soluble nucleotidases; 4. interaction of the
resulting adenosine with its own nucleoside-selective (P1) receptors; and 5. extracellular
adenosine re-uptake and metabolism (43).
ATP is a ubiquitous molecule synthesized and released by almost every mammalian cell.
Various excitatory/secretory tissues such as nerve terminals and chromaffin cells (44, 45), pancreatic acinar cells (46), and circulating platelets (47) store ATP and ADP, together with other neurotransmitters in specialized granules (called synaptic vesicles, chromaffin granules, or platelet dense granules) at concentrations up to 1 M. Specifically, platelet dense granules contain very high concentrations of both ATP (up to 1 M) and ADP (up to 0.5 M) (47). ATP is also co- stored in the synaptic vesicles with acetylcholine (ACh) at concentrations in the millimolar range. Cholinergic synaptic vesicles can also store (at lower concentrations) GTP, ADP, and
12 AMP (chromaffin granules also UTP (48)). There is always more ACh in the isolated vesicle fractions than ATP, with the molecular ratio of ACh to ATP in cholinergic vesicles varying
between 5 and 10, depending on the tissue source (49). It is also well established that
noradrenaline (NA) and ATP exist as co-transmitters in sympathetic nerves in a molar ratio
estimated to be 7:1 to 12:1, NA:ATP (50-54). However, the molar proportions of NA:ATP vary in different tissues, and species, during development and aging, and in different pathophysiological conditions (55).
It was previously assumed the main source of ATP acting on purinergic receptors originated from damaged or dying cells. It is now accepted that ATP release from healthy cells is a physiological or pathophysiological response to hypoxia, inflammation, mechanical stress, and some agonists (56). At basal level, many cells release nanomolar concentrations of ATP, and it is likely different mechanisms underlie constitutive versus stress-stimulated nucleotide release (67-69). ATP is also released from the following non-excitatory tissues: astrocytes and glial cells (44); bone cells, joints, and keratinocytes (44, 61); cardiomyocytes (62); epithelial (57) and endothelial cells (58); erythrocytes (63); fibroblasts (59); hepatocytes (60);
monocytes/macrophages (65); neutrophils (64); and other hematopoietic cells (66).
There are three general mechanisms by which intracellular ATP can be released (Figure
6) from cells. The first is cell lysis, which may occur due to cell death or tissue damage.
Although this is not a physiological release of ATP, it contributes to pathophysiological
mechanisms. Second is vesicular release, resulting from neuronal and non-neuronal cells taking
up and packaging ATP. The vesicular ATP is then released upon stimulation (70) via both Ca-
dependent and Ca-independent mechanisms (71, 72). The third major mechanism for
13
ATP Extracellular ATP ATP ATP ATP Pool ATP (nanomolar) ATP
ATP ATP ATP NE ACh ATP Intracellular Cell ATP ATP Pool Vesicular -3 ATP ATP-binding (~10 M) ATP Damage ATP ATP ATP ATP ATP Release Cassette (ABC) ATP ATP ATP ATP proteins
Figure 6. General methods of nucleotide release. There are three main mechanisms for release of intracellular ATP: 1. cell lysis/damage; 2. vesicular release; and 3. ATP binding cassette (ABC) proteins.
14 intracellular ATP release is by activation of ATP-binding cassette (ABC) proteins. ABC proteins constitute a family of transport ATPases found in virtually all cells including bacteria, plants, and animals (73). One such ABC protein is the P-glycoprotein (multi-drug resistance
(MDR) glycoprotein), which is responsible for microorganisms acquiring resistance to a variety of normally toxic drugs via extrusion of the drugs from the microorganism (74).
Once released from the cell, extracellular adenosine nucleotides are rapidly hydrolyzed to adenosine by ecto-nucleotidases to terminate P2 receptor signaling and to limit desensitization and internalization of the P2 purinergic receptors. When adenosine levels generated by the action of ecto-nucleotidases in the extracellular space are high, adenosine is further metabolized to inosine by adenosine deaminase, or transported into cells by means of transporters and phosphorylated to AMP by adenosine kinase. Additional intracellular phosphorylation events convert AMP to ADP and finally, back to ATP.
4. Purinergic Receptors
The concept of extracellular nucleotides acting as agonists at specific receptors was first
proposed by Dr. Geoffrey Burnstock in 1976 (75). In 1978, Burnstock suggested a subdivision
of purinergic receptors into P1 (adenosine) and P2 (ATP/ADP) receptors based on several
criteria (76) (Figure 7). At the P1 receptors, an agonist potency order of adenosine > AMP >
ADP > ATP was proposed. Methylxanthines such as caffeine and theophylline were proposed to
be selective antagonists and activation of the P1 receptor was suggested to lead to changes in
intracellular cAMP levels. At the P2 receptors, an agonist potency order of ATP > ADP > AMP
15
Ecto- Nucleotidase Nucleotidase 5’-nucleotidase ATP ADP AMP Ado
Extracellular
COOH NH2 P2X P2Y 1-7 1,2,4,6,11,12,13,14 P11,2A,2B,3 Intracellular
Figure 7. Purinergic signaling pathways. The purinergic receptors are divided into P1
(adenosine) and P2 (ATP/ADP) receptors. The P2 receptors, directly modulated by ecto- nucleotidase hydrolysis of nucleotides, are divided into the non-selective P2X ligand-gated ion channels and the P2Y G-protein coupled receptors. ATP is the only agonist at the P2X receptors, while either ATP or ADP can act as agonists at some P2Y receptors. Membrane-bound and soluble nucleotidases quickly terminate agonist response by hydrolyzing NTPs and NDPs to
NMPs and the ecto-5’-nucleotidase specifically hydrolyses AMP to adenosine.
16 > adenosine was proposed. Methylxanthines were not antagonists at P2 receptors and activation
of the P2 receptor was suggested not to lead to changes in intracellular cAMP levels, but in some
cases to promote prostaglandin synthesis. In 1985, Burnstock proposed a pharmacological basis
for distinguishing subtypes of P2 receptors (77), designated P2X and P2Y. The P2X receptors have an agonist potency order of α,β-methyleneATP > β,γ-methyleneATP > ATP = 2-
methylthioATP; antagonism by arylazidoaminopropionyl-ATP (ANAPP3), and selective
desensitization following administration of α,β-methyleneATP. These contraction mediating
P2X receptors were localized in the vas deferens and urinary bladder of guinea-pig, rat, frog, as
well as in rat ventricle, and also in the smooth muscle of the rat femoral artery and rabbit central
ear artery. The P2Y receptors were characterized as having an agonist potency order of 2-
methylthioATP >> ATP > α,β-methyleneATP = β,γ-methyleneATP; weak antagonism by
ANAPP3 and desensitization following administration of α,β-methyleneATP. These P2Y
receptors were present in the guinea-pig taenia coli and the longitudinal muscle layer of the
rabbit portal vein, where they mediate relaxation, and also caused relaxation in the rat vascular endothelial cells, femoral artery, and pig aorta.
Subsequent identification of P2X and P2Y receptor subtypes led Abbracchio and
Burnstock to propose two major P2 receptor families: the non-selective P2X family of ligand- gated ion channel receptors and a P2Y family of G protein-coupled receptors (78). Currently there are seven P2X and eight P2Y receptors recognized in the human genome. Most of these receptors are activated by nucleotide concentrations within the range of 10-6 to 10-4 M (66) and
are regulated by the ecto-nucleotidases via hydrolysis of extracellular nucleotides. The
generation of adenosine and its action on the P1 receptors is an important, indirect affect of
extracellular nucleotide hydrolysis.
17 4.1. P1 (Adenosine) Receptors
P1 receptors are seven transmembrane G-protein coupled receptors. Adenosine is their
preferred, endogenous agonist. To date, four P1 receptor subtypes have been cloned and
characterized, namely: A1, a ubiquitously expressed receptor that couples to Gi/o, decreasing
cAMP, and causing inhibitory actions in most tissues; A2A, an abundant receptor in basal
ganglia, vasculature, and platelets that couples to Gs to increase cAMP levels; A2B, which interacts with Gs to increase cAMP levels; and A3, which interacts with Gi/o to decrease cAMP
levels (79-82).
4.2. P2X Receptors
P2X receptors are non-selective, ligand-gated ion channels that open in response to the
binding of extracellular ATP, but do not respond to any other naturally occurring nucleotides.
There are seven genes in humans (P2X1-7) that encode P2X subunits. Each subunit shows a
topology similar to the NTPDases, in which there are: two transmembrane helices with an
intracellular amino- and carboxyl-terminus; a large extracellular loop with 10 conserved cysteine
residues forming 5 disulfide bonds; and an ATP-binding site, which may involve regions of the
extracellular loop adjacent to the TM helices. All P2X receptors are thought to form functional
trimers (83-86).
The P2X receptor family shows many pharmacological and operational differences (87).
The kinetics of activation, inactivation, and deactivation vary considerably among P2X receptor
subtypes. The pharmacology of the recombinant P2X receptor subtypes expressed in oocytes or
18 other cell types is often different from the pharmacology of P2X receptor-mediated responses in
naturally occurring tissues. This is partly due to the existence of hetero- and homo-trimers of
various P2X receptor subtypes that form the ion pores. Alternatively spliced variants of P2X
receptor subtypes may also add to the diversity of P2X receptor properties. In addition, the
presence of ecto-nucleotidases in tissues that rapidly hydrolyze ATP is usually not a factor when
examining recombinant receptors, but is in vivo.
4.3. P2Y Receptors
Metabotropic P2Y receptors (P2Y1, P2Y2, P2Y4, P2Y6, P2Y11, P2Y12, P2Y13, and P2Y14) are seven transmembrane G-protein coupled receptors. Most P2Y receptors couple to a single G protein (P2Y1,2,4,6, all couple to Gq/11; P2Y11 couples to both Gq/11 and Gs; P2Y12,13 couple to Gi;
and P2Y14 couples to Gi/o) (67). The P2Y receptor subtypes differ in agonist activation, where:
P2Y1,6,12 are activated principally by nucleoside diphosphates; P2Y2,4 are activated mainly by
nucleoside triphosphates; some P2Y receptor subtypes are activated by both purine and
pyrimidine nucleotides (P2Y2,4,6), and others are activated by purine nucleotides alone
(P2Y1,11,12) (67). In response to nucleotide activation, recombinant P2Y receptors either activate
phospholipase C (PLC) and release intracellular calcium, or modulate adenylyl cyclase and alter
cAMP levels (67). P2Y receptor subtypes can also form heteromeric complexes and most
recently, adenosine (P1) receptors were shown to form a heteromeric complex with P2Y1 receptors (67, 88). There is little evidence that P2Y5,9,10 sequences are nucleotide receptors or
that they affect intracellular signaling cascades. Consequently, these proteins were dropped from
19 International Union of Pharmacology (IUPHAR) P2Y receptor nomenclature and termed
“orphan” receptors.
5. Functional Significance of Purinergic Receptor Signaling
The majority of the acute signaling events of purinergic receptors revolve around ATP as
a neurotransmitter, co-released with norepinephrine (NE), acetylcholine (ACh), and other
neurotransmitters and neuropeptides. Acute purinergic signaling also plays a role in exocrine
and endocrine secretion by regulation of ion transport in epithelial cells to stimulate Cl- transport and maintain moist airway surfaces (89); vascular tone, mediated not only by P2X, but also P2Y receptor subtypes that cause the release of endothelium-derived relaxing factor (EDRF) and potent vasodilatation (90-92); and insulin secretion from pancreatic β-cells (93). Other acute signaling roles for the purinergic receptors include: ionotropic, chronotropic, and arrhythmogenic effects in the myocardium (62), immune responses and control of leukocyte traffic between the blood and tissues (66); as well as activation and aggregation of platelets at sites of vascular injury (47).
There is increasing experimental data demonstrating the long-term (trophic) roles purines and pyrimidines have in embryonic development (94), cell proliferation and growth (95), induction of apoptosis and anticancer activity (66, 96), atherosclerotic plaque formation (97), as
well as wound healing and bone formation and resorption (61).
20 6. Ecto-nucleotidases
Extracellular nucleotides are rapidly hydrolyzed to adenosine to terminate nucleotide
signaling (Figure 8) at P2 receptors. There are several enzyme families involved in the metabolism and control of extracellular nucleotide concentrations, including: alkaline
phosphatases (AP); the ecto-5’-nucleotidase; the ecto-nucleotide
phosphodiesterase/pyrophosphatases (E-NPPs); and finally, the ecto-nucleoside triphosphate
diphosphohydrolases (eNTPDases), which are considered the key players in extracellular
nucleotide hydrolysis.
6.1. Alkaline phosphatase (AP)
Alkaline phosphatases are ubiquitous enzymes present in many organisms from bacteria
to humans. As their name implies, APs have a pH optimum for hydrolysis in the alkaline range,
from pH 8 to 11 (98). With few exceptions, these enzymes are homodimeric and each catalytic
site contains three metal ions (two zinc and one magnesium) necessary for enzymatic activity
(98). Three isoenzymes are tissue-specific (intestinal AP, placental AP, and germ cell AP) with
90-98% homology. The fourth isoenzyme, tissue-nonspecific alkaline phosphatase (TNAP) is only ~50% identical in sequence to the other three isoenzymes, and is abundantly expressed in bone, liver, kidney, and at lower levels in other tissues (99). Mammalian APs display broad substrate specificity towards different phosphomonoesters and other phosphorylated compounds, including various fatty acid chains, inorganic polyphosphates, glucose-phosphates, β-
21 1
345 ATP ADP AMP Ado Ino Hyp
2 2
Figure 8. Major extracellular nucleotide hydrolyzing enzymes. There are five major
families of enzymes that hydrolyze extracellular nucleotides and nucleosides: 1. E-NPP: ATP Æ
AMP + PPi; 2. E-NTPDase: ATP Æ ADP + Pi and ADP Æ AMP + Pi; 3. Ecto-5’-nucleotidase:
AMP Æ adenosine (Ado) + Pi; 4. Adenosine deaminase: Adenosine Æ Inosine (Ino); and 5.
Purine nucleoside phosphohydrolase (PNP): Inosine Æ hypoxanthine (Hyp). In addition,
alkaline phosphatase (AP) is able to catalyze the hydrolysis of all three phosphates of ATP, resulting in adenosine production.
22 glycerophosphate, and bis(p-nitrophenyl) phosphate, catalyzing release of inorganic phosphate from these varied substrates (98).
6.2. Ecto-5’-nucleotidase/CD73
Currently, there are seven human 5’-nucleotidases isolated and characterized. Five of the enzymes are located in the cytosol, one in the mitochondrial matrix, and one attached to the outer
plasma membrane. This latter enzyme, known as ecto-5’-nucleotidase, is expressed to a variable extent in different tissues, with abundant expression in the colon, kidney, brain, liver, heart, and lung (100). Cell surface-associated ecto-5’-nucleotidase, also known as CD73, efficiently hydrolyses 5’-AMP with Km of 3-50 µM, and shows no activity towards either nucleoside 2’-
and 3’-monophosphates or nucleoside di- and triphosphates (100). The mammalian ecto-5’-
nucleotidase consists of two glycoprotein subunits that are tethered by non-covalent bonds (101), bind zinc and other divalent metal ions at the N-terminal domain (102), and is anchored to the plasma membrane at the C-terminus by phosphatidylinositol (PI) (100, 103). This ecto-enzyme belongs to a large superfamily of dinuclear metallophosphoesterases acting on diverse substrates such as Ser/Thr phosphoproteins, various nucleotides, and sphingomyelin (102).
6.3. Ecto-nucleotide phosphodiesterase/pyrophosphatase (E-NPP)
The E-NPP family of enzymes consists of seven structurally related ecto-enzymes (NPP1
– NPP7) that possess broad substrate specificity. They are capable of hydrolyzing
pyrophosphate and phosphodiester bonds in di-nucleotides, nucleic acids, nucleotide sugars, as
23 well as choline phosphate esters and lysophospholipids (104). Only the first three members of
this family, NPP1 (previously called plasma cell differentiation antigen-1, PC-1), NPP2
(autotoxin, aka phophodiesterase 1α), and NPP3 (gp130RB13-6, aka B10 or phosphodiesterase 1β),
are capable of hydrolyzing various nucleotides, and therefore relevant in the context of
purinergic signaling (105).
6.4. Nucleoside triphosphate diphosphohydrolases (NTPDases)
In 1999, at the “Second International Workshop on Ecto-ATPases and Related
Ectonucleotidases” held in Belgium, the participants established a unifying nomenclature for the
NTPDases, and other extracellular nucleotide hydrolyzing enzymes. The protein nomenclature
for nucleoside triphosphate diphosphohydrolase is “NTPDaseX”, where X is 1-8 (106) (Table 1).
The gene names for the NTPDases are “ENTPDX”, where X is 1-8, with the gene numbers corresponding to the protein numbers (Table 1).
There are eight members of the human NTPDase family of enzymes that hydrolyze extracellular and intra-luminal nucleotides in a divalent cation (Ca+2 and Mg+2) dependent
manner. There are six, membrane-bound enzymes (NTPDases 1-4, 7 and 8), whose carboxyl-
and amino-terminal ends are each anchored to the membrane (107). Two enzymes lack a
carboxyl-terminal transmembrane (TM) helix (NTPDase5 and NTPDase6), and can be secreted
as soluble enzymes after cleavage of their respective N-terminal signal sequences (108, 109).
NTPDase 1-3 and 8 reside on the cell surface, thereby modulating purinergic receptors and
nucleotide signaling.
24
Protein Name Previously used protein names Comments (Gene Name)
NTPDase1 CD39, ecto-ATP diphosphohydrolase, Cell-surface membrane protein, hydrolyzes ATP and (ENTPD1) ectoapyrase ADP almost equally well, maintains hemostasis
NTPDase2 CD39L1, ecto-ATPase Cell-surface membrane protein, hydrolyzes ATP much (ENTPD2) faster than ADP (10-50 X as fast)
NTPDase3 CD39L3, HB6 Cell-surface membrane protein, hydrolyzes ATP (ENTPD3) slightly faster than ADP (3-5 X as fast)
NTPDase4 Golgi UDPase, LALP70 Found in Golgi, and in lysosomal membranes, may (ENTPD4) function in glycosylation of proteins
NTPDase5 CD39L4, PCPH proto-oncogene Found in internal membranes and can be secreted as (ENTPD5) a soluble enzyme after cleavage of signal sequence
NTPDase6 CD39L2 Found in internal membranes and can be secreted as (ENTPD6) a soluble enzyme after cleavage of signal sequence
NTPDase7 LALP1 Very similar to NTPDase4 (ENTPD7)
NTPDase8 ATPDase Cell-surface membrane protein, hydrolyzes ATP (ENTPD8) (chicken liver, oviduct and stomach, slightly faster than ADP (2-3 X as fast), less sensitive and mouse liver) to detergent inactivation
Table 1. Members of the E-NTPDase Family. There are eight members of the human
NTPDase family of enzymes, NTPDase1-8. The protein nomenclature is "NTPDaseX", for
"ecto-nucleoside triphosphate diphosphohydrolase, where X is 1-8. The gene names for these enzymes are "ENTPDX", where X is 1-8, with the gene numbers corresponding to the protein numbers.
25 7. Molecular Cloning, Sequencing, Expression, and Characterization of the NTPDases
7.1. The membrane-bound, cell surface NTPDases (NTPDase1-3 and 8)
NTPDase1-3 and 8 are all membrane-bound, cell surface enzymes that hydrolyze extracellular nucleotides. NTPDase1/CD39 was originally identified as the major surface marker of EBV-transformed B lymphoblastoid cells, and later was found primarily on activated immune cells (110). In 1996, NTPDase1/CD39 became the first human gene reported to encode an
NTPDase (41). By transfection of COS-7 cells with NTPDase1 cDNA, Wang et al demonstrated the 510-amino acid, 70 kDa protein hydrolyzed ATP and ADP equally well in the presence of divalent cations (Ca+2 and Mg+2). In addition, its enzymatic activity was not inhibited by P- or
V-type ATPase inhibitors (41).
Hicks-Berger and Kirley expressed and characterized NTPDase2/CD39L1 in mammalian
COS-1 cells, demonstrating NTPDase2 is a ~66 kDa glycoprotein that hydrolyzes NTPs at a
ratio of 20:1 as compared to NDPs (111). The activity of NTPDase2 depends on its
glycosylation state (treatment with PNGase-F decreases enzyme activity) and also rapidly
decreases in activity with time during incubation at 37oC. To the contrary, the activity of
NTPDase2 is stimulated by lectin concanavalin A (Con A) and the chemical cross-linking agent disuccinimidyl suberate (DSS). Based on Northern blot hybridization, NTPDase2 is expressed in the heart, brain, placenta, skeletal muscle, kidney, and pancreas (112).
Smith and Kirley isolated the NTPDase3 (aka HB6 and CD39L3) clone from a human brain cDNA library and expressed the protein in mammalian COS-1 cells (12). NTPDase3 encodes a 529-amino acid glycoprotein, with a molecular weight of ~80 kDa. It has seven
26 putative N-glycosylation sites, one cAMP- or cGMP-dependent protein kinase phosphorylation
site, four protein kinase C (PKC) phosphorylation sites, and two casein kinase phosphorylation
sites. The nucleotide hydrolysis ratio of the expressed protein is approximately 3:1
(ATPase:ADPase activity). Based on Northern blot hybridization, NTPDase3 is highly
expressed in the brain, pancreas, spleen, and prostate, with moderate to low expression in other
tissues (112). Similar to other NTPDases, NTPDase3 is sensitive to inhibition by millimolar
concentrations of sodium azide (NaN3).
NTPDase8 is the newest member of the NTPDase family that was cloned from human
liver RNA by real time-polymerase chain reaction (RT-PCR) and expressed in HEK293 cells
(113). It is a 495-amino acid protein that is sensitive to detergent inhibition by Triton X-100 and
NP-40. Consistent with many other membrane-bound NTPDases, NTPDase3 preferentially
hydrolyzes ATP over ADP, and has higher enzyme activity in the presence of Mg+2 as compared
to Ca+2.
7.2. The intraluminal NTPDases (NTPDase4-7)
NTPDase4 and 7 are intracellular, membrane-bound proteins found in the Golgi and
lysosomal membranes. NTPDase4 was cloned from a human brain cDNA library and expressed
in mammalian COS-7 cells (114). It encodes a 610-amino acid protein with two transmembrane
domains (TMDs). The enzyme preferentially hydrolyzes uridine 5’-diphosphate (UDP) as a
substrate. It is stimulated by divalent cations in the following order: Ca+2 >> Mg+2 > Mn+2. Due to its localization, it is suggested NTPDase4 may play a role in glycosylation of proteins and
27 lipids by hydrolyzing UDP, an inhibitor of galactosyltransferase, decreasing its accumulation in
the Golgi apparatus (Figure 9).
NTPDase7/LALP1 was cloned from human brain and expressed in HEK293 cells (115).
It encodes a 604-amino acid protein with a molecular weight of approximately 70 kDa.
NTPDase7 preferentially hydrolyzes NTPs (UTP, GTP, and CTP) and may play a role in the
salvage of nucleotides from the lysosomal/autophagic vacuole lumen.
NTPDase5 and 6 are found in intracellular membrane organelles, but can also be secreted
as soluble enzymes upon cleavage of their respective N-terminal signal sequences (108, 109).
Therefore, these enzymes can hydrolyze both extracellular and intraluminal nucleotides.
NTPDase5 was cloned from a human brain cDNA library as the first reported human, secreted
NTPDase. It was also expressed in bacteria and refolded from inclusion bodies. The soluble form of the enzyme is 410-amino acids long with a molecular weight of 45.5 kDa. It prefers the hydrolysis of NDPs (specifically GDP) over NTPs, and Mg+2 over Ca+2 as the divalent cation
supporting nucleotide hydrolysis. NTPDase6 was expressed in mammalian COS-1 cells where
90% of the enzyme activity was found in the cell media, indicating the presence of a soluble
protein. NTPDase6 is released as a 50 kDa soluble protein, resulting from cleavage of its N-
terminal sequence after Ala-78. It has four cysteine residues that are conserved among the other
NTPDases, which presumably form two disulfide bonds (109). In addition, NTPDase6 has two
potential N-glycosylation sites (glycosylation not necessary for activity) and prefers the
hydrolysis of NDPs (specifically GDP and IDP) over NTPs.
28
Cytoplasm
ER/Golgi Lumen
Folded protein NDPase (NTPDase 4, 5 or 6) UDP UMP UMP ( - Antiporter ) Glucose Transferase + UDP-glucose UDP-glucose Folding protein
Folding protein
Chaperones ATP AMP AMP Misfolded protein ADP Protein Synthesis ATP
Figure 9. Intracellular nucleotidase control of protein folding and processing? Intraluminal
nucleotides are hydrolyzed by NTPDase4, 5, and 6. These nucleotidase, which preferentially
hydrolyze UDP and GDP, are located in the lumen of the ER and Golgi. After protein synthesis
on ribosomes, each protein must be folded correctly. Chaperone proteins, which require ATP, are expressed in the lumen of the ER and Golgi to assist in this process. During the folding
process, many proteins must be glycosylated in the Golgi to be properly processed. NTPDase4,
5, and 6 hydrolyze UDP to UMP. Accumulation of UDP present in the lumen of the ER and
Golgi causes a feedback mechanism that inhibits glucose transferase, the enzyme necessary for
adding glucose molecules to proteins. Thus, intracellular NTPDases regulate the amount of
intraluminal UDP present, likely modulating glycosylation of proteins.
29 8. The Biochemical and Structural Characterization of the NTPDases
8.1. Enzymology of the NTPDases
The NTPDases are the key players in extracellular nucleotide hydrolysis to control the
level of these molecules at the cell surface. ATP, ADP, UTP, and UDP are the biologically most
relevant substrates of the NTPDases, which initiate a number of cellular responses via the
activation of ionotropic P2X and metabotropic P2Y receptors. Therefore, the NTPDases regulate
many physiological processes by altering and controlling the concentration of extracellular
nucleotide agonist at P2 (and P1) receptors.
The NTPDases exhibit E-type ATPase activity, meaning: their nucleotidase activity is
+2 dependent upon millimolar Ca or Mg+2; they are insensitive to specific inhibitors of P-type, F-
type, and V-type ATPases (vanadate, ouabain, and oligomycin, respectively); and sensitive to
millimolar NaN3 inhibition of enzyme activity (116). Typically, the NTPDases have a low
specificity for the hydrolysis of different nucleoside tri- and di-phosphates, but nucleoside
monophosphates and non-nucleoside phosphates are not substrates. The NTPDases also
hydrolyze nucleotides over a broad range of pH, and do not form phosphorylated intermediates during catalysis. In addition, the NTPDases are glycoproteins whose enzyme activity is stimulated by lectins. They are also relatively insensitive to inactivation by proteases, but most are inactivated by many biological detergents, being especially sensitive to Triton X-100 inactivation. Currently, there is a lack of potent and specific inhibitors for both the family of enzymes and for the individual NTPDase isoenzymes.
30 Four out of the eight members of the NTPDase family (NTPDase1-3 and 8) are relevant
to the control of P2 receptor signaling. These enzymes are localized at the surface of the plasma
membrane and hydrolyze nucleotides in the range of concentrations that activate P2 receptors.
Considering each of the membrane-bound, cell surface NTPDases (NTPDase1-3 and 8) were
partially characterized, expressed in different laboratories, and under different conditions,
comparison of the biochemical properties of these enzymes was complicated until Kukulski et al
expressed them all together (117) (Table 2). Even though NTPDase5 (108) and NTPDase6 (109)
can be present at the surface of the plasma membrane and secreted as soluble enzymes following
a proteolytic cleavage, their high Km values and relatively low specific activities make it unlikely
that these enzymes regulate P2 receptor signaling (in comparison to purified, full-length rabbit
NTPDase2 (118), which has a specific activity of approximately 400,000 µmol/mg/hr).
Therefore NTPDase5 and NTPDase6 were not included in the comparative analysis shown in
Table 2.
8.2. Inhibitors of the NTPDases
Modulation of nucleotide signaling has mainly focused on identifying inhibitors
(antagonists) for the purinergic receptors. Over the last decade, the NTPDase family of enzymes
has emerged as a potential new pharmacological target. It is evident that inhibition or
modulation of the NTPDases may prolong the effect of nucleotides at their respective receptors.
Currently, nucleotide analogues (particularly P2 receptor agonists/antagonists) and molecules related to the suramin-Evans blue dye families are being used to inhibit the NTPDases. Gendron
31
max V UDP
m M) (U/mg prot.) (U/mg M) ND ND K µ 67 67 ± 3 ± 0.17 0.01 135 ± 10 ±0.79 0.04 ( 1780 ± 40 ± 1.37 0.08
max V
UTP
m M) (U/mg prot.) (U/mg M) K µ ( 58 ±6 0.57 ±0.03 0.57 ±6 58 47 ± 4 1.05 ± ±47 1.05 0.03 4 393 ± 30 ±3.9 0.2 124 ± 10 ±1.17 0.06
max V ADP
m M) (U/mg prot.) (U/mg M) K ND ND ND µ ( 31 31 ± 1 0.18 ± 0.01 22 22 ± 1 0.75 ± 0.01 265± 20 ±0.61 0.03 max V ATP m M) (U/mg prot.) (U/mg M) µ K ( 75 ± 0.79 10 ± 0.03 70 ± 2 2.3 ± 0.03 17 ±17 ± 0.94 1 0.02 46 ±46 ± 0.74 5 0.04 7 – 10 4.5 –4.5 11 4.5 –4.5 8.5 4.5 –4.5 8.5 pH range Enzyme activity > 50% activity ~2:1 ADP ATP: Ratio ~3-4:1 ~1-2:1 ATPase Apyrase Apyrase Apyrase 10 – 40:1 +2 +2 +2 +2 = Mg = Mg > Mg > Mg +2 +2 +2 +2 Ca Ca Ca Ca Divalent Cation Preference NTPDase hNTPDase1 hNTPDase2 hNTPDase3 rNTPDase8
32 Table 2. Enzymatic characterization of membrane-bound, cell surface human NTPDase1-3 and rat 8 (from reference (119)). Reactions were carried out for 10 minutes in the presence of
5 mM CaCl2 and 80 mM Tris buffer pH 7.4. In all assays, less than 10% of the substrate was hydrolyzed. Km and Vmax values were estimated with nucleotide concentrations ranging from
0.02 to 1.5 mM (119). The results are expressed as the mean ± SEM of three separate experiments, each performed in triplicate.
33 et al described some inhibitory nucleotide analogue compounds (120). Brunschweiger et al
have synthesized a nucleotide analogue derived from uridine-5’-carboxamide that is the first
potent and selective human NTPDase2 inhibitor (121). However, FPL 67156 (6-N,N-diethyl-
β,γ-dibromomethylene-D-ATP) was the first ATP analogue described as a novel, selective
inhibitor of the NTPDases with some 50-fold selectivity over the purinergic receptors (122).
More recently, ARL 67156 (the only commercially available inhibitor of the NTPDases; aka FPL
67156) was shown to be a weak competitive inhibitor of human NTPDase1 (Ki = 11 ± 3 µM) and
NTPDase3 (Ki = 18 ± 4 µM) (123). However, human NTPDase2 and NTPDase8 were less
efficiently inhibited by ARL 67156. At high concentrations of substrate, ARL 67156 was not
able to efficiently inhibit any of the NTPDases, and high concentrations of ARL 67156 were
shown to inhibit the P2Y2, P2Y4, P2Y12, and P2X receptors. Overall, inhibitory nucleotide analogues face the seemingly insurmountable obstacle of the high likelihood of affecting other nucleotide handling enzymes and purinergic receptors as well as their intended NTPDase target.
Non-nucleotide NTPDase inhibitors also exists. Our laboratory was the first to discover that several P2 receptor antagonists such as suramin, Cibacron Blue, and Coomassie Blue G-250 inhibited a purified NTPDase (124). Three structure-activity requirements were found for this class of inhibitors: 1. the position of the sulfonate groups on the naphthalene core is important
(the 2-sulfonate group was found to be required for inhibitory activity); 2. the amino-hydroxy- naphthalene disulfonate structures of these antagonists are involved in the binding to the
NTPDases; and 3. the inhibitory potencies and affinities of these compounds acting as antagonists at P2 receptors increase with size (125).
Polyoxometalates hold some promise as small molecule NTPDase inhibitors (64), but are not yet sufficiently potent or specific to be truly useful. A recent publication (126) gives rise to
34 hope that specific and fairly potent small molecule inhibitors for NTPDases might be possible.
One inhibitor (compound #21) had a sub-micromolar IC50 for rat NTPDase1 and another very
intriguing compound (#20) seems to be specific for NTPDase3 over NTPDase1 and NTPDase2,
at least for the rat enzymes (126). However, this compound does not show the same specificity
for human NTPDase3 (personal communication from the senior author of that published study,
Dr. C.E. Muller, and confirmed by unpublished experiments performed in our laboratory).
Lastly, Munkonda et al are the first to report inhibitory monoclonal antibodies specific
for human NTPDase3 that inhibit the enzyme activity by 60 – 70% (127). In this study,
Munkonda et al demonstrates the importance of the native conformation of NTPDase3 for
epitope recognition as denaturation of the protein with SDS or reducing agents, indication
abolished binding of the monoclonal antibodies. Additional data demonstrated the SDS-resistant
parts of the inhibitory epitope were located in two fragments of the C-terminal lobe (i.e. Leu220-
Cys347 and Cys347-Pro485) and site-directed mutagenesis revealed the importance of Ser297 and the fifth disulfide bond (Cys399-Cys422) for antibody binding. As yet unpublished epitope localization data has further pinpointed the amino acids critical for antibody binding. This could serve as a springboard for development of antibody inhibitors of other NTPDases, if like
NTPDase3, each NTPDase has a specific epitope for antibody binding, and consequent inhibition.
Regardless, the potential therapeutic outcomes of specific NTPDase inhibition are substantial, as delineated by Gendron et al (120, 128). It is evident the ideal NTPDase inhibitor should not be a P2 receptor agonist/antagonist and should not be dephosphorylated by the enzyme. Unfortunately, the use of non-specific purinergic receptor and NTPDase inhibitors has led to the misinterpretation of some purinergic signaling data; specifically, which purinergic
35 receptors are involved in the many and varied physiological responses to exogenously added nucleotides.
8.3. Site-directed mutagenesis and computational comparative protein modeling of the
membrane-bound, cell-surface NTPDases
Site-directed mutagenesis and computational comparative protein modeling are useful tools to characterize and elucidate key structural features of the NTPDases. The general topography of the membrane-bound, cell surface NTPDases resemble a variety of channel forming proteins such as the P2X receptor or members of the epithelial Na+ channel (129). In addition, cross-linking experiments have indicated NTPDase1-3 form homo-oligomeric complexes starting with dimers. Rat and human NTPDase1 were found to form up to tetrameric oligomers (130, 131), whereas chicken NTPDase2 (132) and human NTPDase3 (133) preferentially form dimers. However, no clear picture has emerged regarding how the N- and C- terminal TM helices interact within and between monomers to form the native oligomeric complexes of the NTPDases.
As shown in Figure 10, membrane-bound, cell surface NTPDases can be divided into three major structural regions: a large extracellular domain, a “linker region”, and a transmembrane domain (TMD) (Figure 10). The extracellular region contains the enzyme active site, five apyrase conserved regions (ACRs), and five conserved disulfide bonds (107).
36 472 476 53 481 P52 484 P Extracellular 485
Cell Membrane
Intracellular NH COOH 2
Figure 10. The three structural regions of the NTPDases. Using NTPDase3 as an example, the three structural regions of an NTPDases monomer are shown. The extracellular domain is a group of multi-colored alpha helices and beta sheets. The “linker” region is depicted as red (C- terminal) and blue (N-terminal) lines labeled with the conserved proline residues of NTPDase3.
The TM helices are depicted as yellow cylinders
37 Through extensive site-directed mutagenesis of human NTPDase3, many of the structural and functional elements of the extracellular domain of the NTPDases are defined (134). The “linker region” is a stretch of amino acids in the primary sequence of NTPDase3 that connects the TMD
(the N- and C-terminal TM helices) and extracellular domain (location of the enzyme active site), and contains several conserved proline residues (N-terminal: P52 and P53; C-terminal:
P472, P476, P481, P484, P485). Interestingly, proline residues confer unique structural rigidity needed for some conformational changes in other proteins, and may facilitate the cross-talk between the extracellular and TM domains of NTPDase3. Lastly, the TMD is thought to mediate substrate specificity (135), sensitivity to membrane perturbation (136, 137), and oligomerization of the membrane-bound, cell surface NTPDases.
8.3.1. The structural features of the extracellular domain of the NTPDases
The extracellular domain contains five ACRs that share nearly identical sequence and spatial conservation with related members of the NTPDase family (139). Based on the conservation of the amino acids within the ACRs of the E-type ATPases (specifically two
“DXG” phosphate binding motifs) with the phosphate binding motifs of the actin/heat shock protein/sugar kinase superfamily of proteins, ACR1 and ACR4 were identified as phosphate binding domains in the extracellular domain, suggesting their role in nucleotide binding and subsequent hydrolysis (139). The importance of these residues in ACR1 and ACR4 was established by mutation of the aspartic acids and glycines, which resulted in the inactivation of
NTPDase3 without a change in the protein expression level (140). There are also two highly conserved tryptophan residues (W187 and W459), which are the only two tryptophan residues
38 conserved in the extracellular domain of nearly all the NTPDases (141). Alanine substitution at
W187 (located in ACR3) resulted in inactivation of NTPDase3 and adversely affected the
cellular localization of the enzyme. Data supported the possibility of altered tertiary structure,
causing NTPDase3 to be improperly folded and retained within the cell. To the contrary, alanine
substitution at W459 (located in ACR5) resulted in an increased ATPase:ADPase ratio (an enhanced NTPase activity, but diminished NDPase activity). The W459A mutant was coupled with a previously described mutant, D219E (located in DXG2/ACR55), which together nearly converted NTPDase3 to an NTPase. In addition, the double mutant (D219E/W459A) was substantially less sensitive to inhibition by NaN3, a more potent inhibitor of ecto-apyrases than ecto-ATPases. This data indicated that a major change in nucleotide tri- and di-phosphate specificity of an NTPDase is possible with only a few changes in the amino acid sequence.
In actin, the conserved glycine residues are very important for its structure and function.
Kirley et al mutated the most highly conserved glycine residues (excluding the two DXG
glycines previously mutated in ACR1 and ACR4) to alanine in the extracellular domain of
NTPDase3 to determine their structural and functional significance (142). Alanine substitution
at G98 (in ACR1a) resulted in an approximately 60% decrease in enzyme activity. Alanine
substitution at G263 (in ACR4a) resulted in an inactive protein with severe folding/processing
problems. Alanine substitution at G462 (in ACR5) resulted in an approximately 85% decrease in
enzymatic activity with some enzyme folding and processing problems. This data led to the
identification of two additional ACR regions, ACR1a and ACR4a, in the extracellular domain of
NTPDase3.
The NTPDases are also heavily glycosylated in their extracellular domain. To determine
the role of glycosylation at the most conserved glycosylation site of the NTPDases (located
39 between ACR1 and ACR1a), N81 and T83 were mutated in NTPDase3 (143). Mutation of either
the asparagine residue (N81D or N81E) where the sugars are attached, or the threonine residue
(T83A) that is part of the consensus sequence for glycosylation, resulted in a decreased
nucleotidase activity (more severely affecting Mg-ATPase than Ca-ATPase activity). In addition, the inhibitory affect of treatment with a deglycosylation enzyme (PNGase-F) as well as
stimulation of activity by concanavalin A (ConA) was attenuated. This data suggests that
glycosylation is important for the activity and may maintain the oligomeric structure of
NTPDase3 and other membrane-bound, cell surface NTPDases, since glycosylation is not
important for the activity of the soluble, monomeric NTPDases.
It is known that the membrane-bound, cell surface NTPDases form dimers and higher
order oligomers (144). Lysine 79 is conserved in all cell surface NTPDases (NTPDase1-3 and
8), but not in the soluble (NTPDase5 and 6) and intracellular (NTPDase4 and 7) enzymes. In human NTPDase3, all substitutions at this site resulted in a loss of 70 – 80% of activity (145).
Only K79R, which maintains the positive charge at this residue, remained able to be stimulated/stabilized by Con A. Interestingly, the residual activity of all the mutants at the K79 site became resistant to Triton X-100 detergent inhibition. Since Triton X-100 is known to dissociate NTPDase oligomers into monomers, this data suggests K79 may be involved in the dimer interface of the membrane-bound, cell surface NTPDases, partially mediated by the extracellular domain.
Recently, Ivanenkov et al determined the ten conserved cysteine residues in the
extracellular domain of all membrane-bound, cell surface NTPDases form five disulfide bonds.
When these disulfide bonds are disrupted in NTPDase3, the protein demonstrates five distinct
biochemical phenotypes (107). This data strongly suggests disulfide bonds between C92-C116,
40 C261-C308, C289-C334, C347-C353, and C399-C422. Due to the conservation of the 10 cysteines, it is likely the disulfide bonds exist in all cell surface NTPDases. The third (C289-
C334) and fifth (C399-C422) disulfide bonds are also present in the soluble NTPDases and are
critical for biosynthetic processing and function. In addition, computational comparative protein
modeling of the NTPDase3 sequence revealed a high degree of structural fold similarity with a
bacterial exopolyphosphatase (PDB 1T6C). This established a 3-D model of the extracellular
portion of NTPDase3, which is consistent with the assignment of several of the disulfide bonds.
A recent crystal structure of the extracellular region of rat NTPDase2 (146) confirmed almost all the site-directed mutagenesis conclusions using NTPDase3, and forms the basis for the much more accurate 3-D models for the extracellular domain of NTPDase3 and other NTPDases currently in use.
8.3.2. The structural features of the transmembrane domain of the NTPDases
.
Many ecto-proteins are anchored to the cell membrane by a single transmembrane domain (TMD) or a lipid link. One of the most intriguing features of the membrane-bound, cell surface NTPDases is the presence and function of its two TM helices, one at both the N- and C- terminal of the enzymes. Most studies regarding the role of the TMD have focused on oligomerization. The predominant, active form of these enzymes appear to be a dimer, although higher-order oligomers are also observed, and the soluble NTPDases (which lack a TMD) are monomeric (131, 133, 144). However, no structural data exists and very limited site-directed mutagenesis studies were performed to perturb and analyze helix interactions in the TMD of the
NTPDases.
41 Wang et al deleted the amino terminal, carboxyl terminal, or both TM helices of
NTPDase1/CD39 and compared the activity of each truncated protein with native NTPDase1
(131). Their goal was to determine whether the TM helices are required for enzymatic activity, and whether disruption of the interactions between the helices is the mechanism for detergent inhibition effects. Truncation of both TM helices resulted in a catalytically active, secreted, soluble protein with an enzyme activity ~90% lower than native NTPDase1. Deletion of either
TM helix resulted in a catalytically active enzyme that was properly processed and trafficked to the cell surface, but both mutant proteins had reduced enzymatic activities similar to the double truncation mutant. Wang et al further proposed that not only the presence and membrane insertion of the TM helices, but also the interactions between them are required for optimal activity (131). Glutaraldehyde cross-linking and sucrose density gradient sedimentation in the presence of detergents showed that NTPDase1 formed oligomers. In addition, detergents that reduced the enzyme activity of NTPDase1 dissociated the oligomeric structure (i.e. Triton X-
100), and that the soluble extracellular domain lacking both TM helices existed only as monomers. One detergent, digitonin, had only a small effect on enzyme activity and preserved the oligomers. Taken together, this data suggested that both TM helices are necessary for full enzymatic activity and they may mediate oligomer formation via TM helix interactions.
Murphy et al have conducted one of the few studies to date that use site-directed mutagenesis to determine the function of a specific set of amino acids in the TMD of NTPDase3
(147). Cysteine to serine substitution was used to test the functional and structural significance of the three non-extracellular cysteine residues (C10, located on the short N-terminal cytoplasmic tail; and C501 and C509, located in the C-terminal TM helix of NTPDase3). None of these cysteines were found to be essential for enzymatic activity. However, cysteine 10 was
42 found to be responsible for dimer formation occurring via oxidation during membrane preparation, as well as for dimer cross-linking resulting from exogenously added sulfhydryl- specific cross-linking agents. Cysteine 501 was found to be the site of chemical modification by
a sulfhydryl-specific reagent, p-chloromercuriphenylsulfonic acid (pCMPS), leading to inhibition
of enzyme activity. The latter result was further confirmed by Chiang et al using NTPDase2, in
which pCMPS also decreases its enzymatic activity via cysteine modification in the TMD (148).
In 2004, Grinthal and Guidotti used a cysteine substitution/copper phenanthroline
disulfide bond-induced cross-linking strategy to probe for possible intra- and intermolecular interactions in the TMD of NTPDase1 (149). They demonstrated that the TM helices of rat
NTPDase1 have strong intra- and inter-molecular interactions between the N- and C-terminal
TM helices, and that these interactions are stronger near the extracellular side of the membrane.
In addition, they showed that the TM helices undergo dynamic motions that are necessary for
maximal enzymatic activity, and that these TM interactions and motions are regulated by
nucleotide binding at the extracellular active site in NTPDase1 (149). Lastly, Chiang et al (148)
using human NTPDase2 and our laboratory using human NTPDase3 (138) also noted a similar
reduction in enzymatic activity of these NTPDases upon oxidative cross-linking of introduced
cysteine residues in their respective TM helices. These data suggest the TM helices of the
membrane-bound, cell surface NTPDases must move or re-orient relative to one another during
nucleotide hydrolysis to facilitate optimal enzymatic activity.
43 9. Functional Significance of the NTPDases
The physiological and pathophysiological functions of the NTPDases are currently under
investigation. Many putative functions have been suggested. Data has implicated a role for the
NTPDases in secretion (150), cell adhesion (151), cancer and malignant transformation (152,
153), pain perception, purinergic receptor system modulation, adenosine recycling, and platelet
aggregation (for a review, see (116)).
Strong evidence was presented that NTPDase1, found in vascular endothelial cells, is one of the three important endothelial thromboregulatory systems that inhibit platelet aggregation in response to agonists, along with nitric oxide (NO) and the eicosanoids (154, 155). This proposed function is supported by the generation of NTPDase1 null mice having defects in blood clotting and related processes (156, 157).
NTPDase2 is expressed in the germinal zones of the brain and suggested to be important for the development of certain areas of the brain (158). It is also expressed in the glial cells of the peripheral nervous system (159). In addition, NTPDase2 is expressed in smooth muscle, where contraction is under the control of ATP via purinergic signaling. More recently,
NTPDase2 was shown to play a critical role in the development of the eye, since over-expression of NTPDase2 in the frog caused ectopic eye-like structures to form, and occasionally even complete duplication of the eye (160).
NTPDase3 is expressed at a relatively high level in brain (112). Immunolocalization using three different anti-peptide antibodies demonstrated NTPDase3 localization in rat brain is highly restricted to axons and associated with synapse-like structures, a pattern that strongly suggests NTPDase3 acts as a regulator of synaptic function (161). The high density of
44 NTPDase3 positive terminals on noradrenergic neurons suggests that NTPDase3 may play an
important role in regulation of noradrenergic autonomic function. Moreover, very high densities
of NTPDase3 were detected in the mediobasal hypothalamus, with the overall mesencephalic
pattern of staining associated closely with hormone responsive brain regions. This pattern of expression, in addition to the observed co-localization with hypocretin-1/orexin-A, suggests that
NTPDase3, by regulating the extracellular turnover of ATP, may modulate feeding, sleep-wake, and other behaviors through diverse homeostatic systems. Appelbaum et al (162), using zebra fish, confirmed our laboratory’s localization of NTPDase3 in the brain to the hypothalamus.
They suggested that NTPDase3 in this location functions in conjunction with P2X8 receptors. In
addition, they also found expression of NTPDase3 in primary sensory neurons in the spinal cord,
where it is likely involved in ATP-mediated pain transmission in conjunction with P2X2 and/or
P2X3 receptors.
Most likely, NTPDase3 has other physiological functions outside the nervous system.
Interestingly, "virtual Northern" data (available via
https://escience.invitrogen.com/ipath/search.do) suggests that pancreatic beta cells, responsible
for insulin production, may have the highest expression level of NTPDase3 of any tissue/cell type. Consistent with this, our laboratory has confirmed the presence of NTPDase3 protein in these cells by Western blot and immunoprecipitation (unpublished results). We have also demonstrated that NTPDase3 endogenously expressed in human pancreatic beta cells can be inhibited in situ by a monoclonal antibody specific for human NTPDase3 (127). Since insulin secretion is modulated by purinergic signaling, this suggests that regulation of NTPDase3 may be of clinical significance for the development and treatment of diabetes.
45 Lastly, soluble NTPDase6 was reported to be co-released with ATP in some mammalian sympathetic nerves (163, 164). Therefore, soluble NTPDase6 may be involved in terminating the effects of the co-released ATP neurotransmitter.
10. Dissertation Scope and Objectives
Site-directed mutagenesis and computational comparative protein modeling have served as useful tools to formulate testable hypotheses, generate models for the enzyme active site, and to elucidate the structural elements that modulate the enzymatic properties of the NTPDase family of enzymes. The recently published crystal structure of the extracellular domain of rat
NTPDase2 (146) confirmed and extended much of the site-directed mutagenesis work done by our laboratory using human NTPDase3 (134). Now it seems the major structural and functional features of the extracellular active site of the NTPDases are fairly well defined. However, this crystal structure does not tell the complete story of the structural elements that regulate the enzymatic mechanism of the NTPDases. The expressed and refolded extracellular domain of rat
NTPDase2 that was crystallized is monomeric. However, the membrane-bound, cell surface
NTPDase enzymes are oligomeric, and the oligomeric state effects enzyme activity. Our laboratory has determined that the membrane-bound, cell surface NTPDases can be divided into three main domains: 1. the extracellular domain, consisting of two lobes between which is the location of the enzymatic active site; 2. the “linker region”, a stretch of amino acids in the primary sequence of NTPDase3, between the TM and extracellular domain; and 3. the TMD, containing the N- and C-terminal TM helices and short cytoplasmic tails (Figure 10).
Questions regarding the structural element(s) that facilitate the cross-talk between the
46 extracellular and transmembrane domains during nucleotide hydrolysis as well as the orientation
and/or interactions of the TM helices are still unanswered.
The objectives of this dissertation are: 1. to better understand the structural elements that mediate the intra-protein signal transduction mediated by substrate binding at the extracellular domain, causing movements and/or re-orientations of the TM helices during nucleotide hydrolysis, and 2. to determine how the N- and C-terminal TM helices interact in human
NTPDase3 to form the native, dimeric enzyme. The rationale for these studies and the specific objectives are described below:
A) The “linker region” of NTPDase3 joins the N- and C-terminal lobes of the extracellular domain to the N- and C-terminal helices in the TMD, respectively. Several conserved proline residues (N-terminal: P52 and P53; C-terminal: P472, P476, P481, P484, and P485) are located in the linker region of NTPDase3. Due to the unique structure of proline and its involvement in domain rearrangements in other proteins, we investigated the role of the conserved proline residues in mediating the linkage of substrate binding in the extracellular domain to movements of the TM helices during nucleotide hydrolysis in human NTPDase3.
B) Polar residues are known to drive the association of TM helices and have effects on protein processing and function. NTPDase3 contains several conserved polar residues (N-terminal: S33,
S39, T41, and Q44; C-terminal: T490, T495, and C501) in its TMD. Therefore, we investigated the role(s) of these conserved polar residues in the TMD of NTPDase3, and strove to understand how the N- and C-terminal TM helices interact within and/or between monomers, in order to
47 enhance the knowledge of NTPDase function and its modulation by oligomerization and
membrane perturbation.
C) The existence of alternatively spliced variants is a common phenomenon among enzymes,
including some NTPDases. The cDNA encoding a truncated variant of NTPDase3, lacking the
C-terminal TM helix, was found in a human lung library by polymerase chain reaction (PCR).
We expressed and characterized the enzymatic properties of this alternatively spliced variant to
test the hypothesis that truncation would result in an enzyme with diminished enzymatic
properties as compared to full-length NTPDase3.
D) Single nucleotide polymorphisms (SNPs) can alter the primary structure and cause a distinct
biochemical phenotype, which may be physiologically important. Human NTPDase3 has five
known SNPs, namely: I24V, R264Q, A496V, L505F, and E440D. Three of these SNPs (I24V,
A496V, and L505F) are located in the TMD, and R264Q is located in the enzymatic active site.
We expressed these mutant proteins to test our hypothesis that one or more of these SNPs would
substantially modify the enzymology, and therefore possibly the function, of human NTPDase3.
48 Chapter II: Materials and Methods
1. General Materials and Methods
1.1. Purification of oligonucleotides for site-directed mutagenesis
350 µL TE Buffer was added to the commercial custom synthetic oligonucleotides in an eppendorf tube, followed by vortexing and incubating in a 37°C water bath for 15 minutes. The samples were vortexed again, and then sonicated in a water bath sonicator for 5 minutes. After centrifugation of the samples for 3 minutes at maximum speed, the supernatants were transferred into a clean eppendorf tube. 35 µL of 3 M NaOAc pH 7.0 was added to each, the samples vortexed after adding 900 µL of 100% ethanol and mixing, and the samples were incubated at -
20°C for 75 minutes. Afterwards, the samples were centrifuged for 5 minutes at maximum speed in a microcentrifuge at room temperature (22oC). The supernatants were carefully removed by
pipetting, and the pellets were gently washed with 1 mL 70% ethanol, followed by centrifugation
for 3 minutes. The supernatants were discarded, the pellets were centrifuged again for 3 minutes,
the rest of the ethanol was removed, and the pellets were air dried for 30 minutes. The
oligonucleotide pellets were then dissolved in 200 µL 1X TE buffer. To calculate the
oligonucleotide concentrations, 20 µL of each stock solution was diluted with 730 µL 1X TE buffer and the OD260 was measured using a Beckman DU-800 spectrophotometer (1 OD260 = 35
µg/µL oligonucleotide). The oligonucleotide solutions were stored at -20°C.
49 1.2. Site-directed mutagenesis of NTPDase3
Mutagenesis of NTPDase3 in the mammalian pcDNA3 vector was performed using a
QuikChange Site-Directed Mutagenesis kit (Stratagene). 10 ng of template DNA (encoding
NTPDase3) was added to 5 µL 10X buffer, 125 ng sense oligonucleotide, 125 ng anti-sense
oligonucleotide, and 1 µL dNTP mix and brought to a total volume of 50 µL with MilliQ H2O in a PCR tube. The solution was gently mixed and 1 µL PfuTurbo DNA polymerase added. For a control sample, template DNA was omitted. The samples were placed in a Techne TC-3000
Thermocycler for 16 cycles of PCR amplification: 30 seconds at 95°C, 1 minute at 55°C, and 10 minutes at 68°C. The samples were cooled to room temperature (22oC) and the parental DNA
digested with 1 µL DpnI at 37°C for 1 hour.
1.3. Bacterial transformation and inoculation of PCR products
A 15 mL Falcon polystyrene round-bottom tube (Fisher) was pre-chilled on ice for 5
minutes. 50 µL XL1-Blue Supercompetent Cells (Stratagene) and 1 µL PCR product was added
to the tube, swirled gently, and incubated on ice for 30 minutes. The bacteria were heat shocked
in a 42°C water bath for 30 – 45 seconds, incubated on ice for 2 minutes, and 500 µL of LB
media (preheated to 42°C) was added and the tube incubated in a 37°C air shaker for 1 hour.
100 µL of the sample was spread onto an LB-carbenicillin plate, incubated at room temperature
for 10 minutes, and then incubated overnight at 37°C upside down. The following day, several
bacterial colonies were picked from the plate and added to 5 mL LB containing 5 uL (100 ng/uL)
50 carbenicillin in a 15 mL Falcon polystyrene round-bottom tube. The colonies were grown overnight in a 37°C air shaker for plasmid cDNA isolation the following day.
1.4. Isolation of DNA from XL1-Blue Supercompetent Cells
Isolation of plasmid cDNA from the bacterial XL1-Blue Supercompetent Cells
(Stratagene) was performed using a QIAprep Spin Miniprep kit (Qiagen). From the 5 mL overnight sample of LB and bacteria, 1.5 mL was placed into an eppendorf tube, microcentrifuged in a Marathon micro A (Fisher) at maximum speed for 3 minutes, and the supernatant discarded. This was repeated twice, using the same liquid culture and the same tube.
250 µL of Buffer P1 was added to re-suspend the cells. 250 µL Buffer P2 was added to lyse the cells and the eppendorf tube was gently inverted 4-6 times to mix. 350 µL Buffer N3 was added to neutralize the reaction and the eppendorf tube was immediately, but gently, inverted 4-6 times.
The samples were centrifuged for 10 minutes at maximum speed. A QIAprep spin column was placed in a 2 mL collection tube (both supplied in the kit) and the supernatant from the eppendorf tube was applied to the QIAprep spin column by pipetting. The QIAprep spin column was centrifuged for 1 min and the flow through discarded. The QIAprep spin column was washed with 750 µL Buffer PE and centrifuged for 1 minute. The flow through was discarded and the spin column was centrifuged for 1 minute to remove any residual wash buffer. The
QIAprep spin column was placed in a clean 1.5 mL eppendorf tube and 50 µL Buffer PE was added to the spin column, incubated at room temperature (22°C) for 1 minute, and centrifuged for 1 minute to collect the plasmid cDNA. To calculate the cDNA concentration, 5 µL of the cDNA preparation was added to 195 µL of MilliQ H2O, and the absorbance at 260 nm was
51 measured using a Beckman DU-800 spectrophotometer (1 OD260 = 50 ng/µL). The plasmid
cDNA was stored at -20°C until used.
1.5. Restriction enzyme digestion of DNA
1 µL isolated cDNA, 1 µL KpnI (New England BioLabs), 1 µL XbaI (New England
BioLabs), 1 µL 10X multi-core buffer (Fisher), and 6 µL MilliQ H20 were mixed in an
eppendorf tube. The samples were centrifuged in a microcentrifuge (National Labnet Company) for 1 minute and incubated in a 37°C water bath for 1 hour. 5 µL of DNA loading buffer was added and the restriction enzyme digested samples were loaded onto a 0.8% agarose gel. (Make
a 0.8% agarose gel by adding 0.4 grams of electrophoresis grade agarose (Bethesda Research
Laboratories) to 1 mL 50X TAE buffer (60.5 grams Tris Base, 14.3 mL glacial acetic acid, and
25 mL 0.5 M EDTA, pH 8 in 250 mL MilliQ H2O) and 49 mL MilliQ H2O. The solution was
heated for 1.5 minutes, swirled, and heated for an additional 45 seconds in a microwave. The
solution was cooled for 5 minutes, at which time 10 µL of ethidium bromide (5 mg/mL) was added. After mixing, the solution was poured into the gel cast and allowed to cool and solidify.
250 mL of 1X TAE Buffer was added to the gel chamber. The samples were electrophoresed at room temperature (22°C) for 1 hour at 80V.
1.6. Transient transfection of COS-1 cells
COS-1 cells were grown in Dulbecco’s modified Eagle’s medium (DMEM) with 10% bovine serum and 2% mixture of antibiotics and antimycotics (Invitrogen), known as complete media. 4 µg plasmid cDNA encoding NTPDase3 (or empty pcDNA3 vector as a control) was
52 diluted into 750 µL of 37oC serum-free, antibiotic-free DMEM. The solution was mixed after the addition of 20 µL Plus Reagent (Invitrogen) and incubated at room temperature (22oC) for 15 minutes. 30 µL of Lipofectamine (Invitrogen) was added to another 1.5 mL eppendorf tube containing 750 µL of serum-free, antibiotic-free DMEM. The diluted Lipofectamine Reagent,
Plus Reagent, and plasmid cDNA were combined into one eppendorf tube and incubated at 22oC for 15 minutes. At this point, plates of COS-1 cells for transfection were removed from the incubator, the media removed by aspiration, and 5 mL of 37oC serum-free, antibiotic-free
DMEM added. This washing was repeated a second time. The third time the 37oC serum free, antibiotic free DMEM was added, the DNA complexes (DNA, DMEM, Lipofectamine, and Plus
Reagent mixture) were also added to the cells. The plates were swirled and allowed to incubate at 37oC for five (5) hours. 13 mL of complete media was then added and the plates were placed back in the incubator. The next morning, the media was removed by aspiration and 10 mL of
37oC complete media was added to the plates. The cell culture plates were returned to the incubator and the COS-1 cells were harvested the next day.
1.7. COS-1 cell crude membrane preparation
Media was removed from the culture dishes by aspiration, and each culture dish was rinsed twice with 10 mL isotonic wash buffer (IWB; 20 mM MOPS, pH 7.4, 140 mM NaCl, and
5 mM KCl, prepared the day of use). 1 mL tissue homogenization buffer (THB; 250 mM sucrose, 30 mM MOPS, 2 mM EDTA, adjusted to pH 7.4 with NaOH), prepared fresh the day of use, was added to each cell culture plate. The cells were removed from the culture plate with a scraper and the cell suspension transferred to a glass-Teflon homogenizer on ice. The culture
53 plates were rinsed with 1 mL THB, the remaining cells scraped off and added to the cell
suspension in the homogenizer. Each sample was homogenized with 25 passes of the Teflon
pestle (on ice). The homogenized extract was centrifuged in a 50 Ti rotor for 1 hour at 40,000
RPM at 4oC in a Beckman centrifuge. The supernatant was discarded and the COS cell pellet re- suspended in 150 µL THB. The cell suspension was transferred to a homogenizer on ice. The centrifuge tube was rinsed with an additional 150 µL THB, which was added to the homogenizer. The COS cell membranes were homogenized with 25 passes of the Teflon pestle and transferred to a 1.5 mL eppendorf tube on ice. The COS cell membranes were aliquoted and stored at -80oC until used.
1.8. Protein concentration of COS-1 crude cell membrane preparations
Protein concentrations were determined using the Bio-Rad protein assay reagent with the
modifications of Stoscheck (165). The crude COS-1 cell membrane preparations were diluted
1:5 with THB. 20 µL of COS-1 cell membranes (or water for blanks, or bovine serum albumin
for standards) was added to a 12 x 75 mm glass test tube (Fisher). 20 µL of 5 M NaOH was
added to the test tube and vortexed. 960 µL of diluted (1:5) commercial BioRad protein assay
dye reagent was added to each tube and vortexed. The samples were incubated for at least 5 (but
not more than 30) minutes at room temperature (22oC) and the absorbance at 595 nm versus
water was measured using a SPECTRONIC 20 GENESIS spectrophotometer. Protein
concentrations were determined using the “LOWRY” program included in “A collection of
radioligand binding analysis programs” by G.A. McPherson, using a non-linear curve fit.
54 1.9. Specific nucleotidase activity of NTPDase3 expressed in COS-1 cells
For most experiments, nucleotidase activities were determined by measuring the amount
o of inorganic phosphate (Pi) released from nucleotide substrates (Sigma) at 37 C using modifications of the Amidol technique of Fiske and Subbarow (166) as previously described
(140). Either 5 mM MgCl2 or 5 mM CaCl2 (both in 20 mM MOPS pH 7.1 buffer) was used for cation-dependent nucleotidase activities. The reactions were initiated by the addition of nucleotide to a final concentration of 2.5 mM in the 0.3 mL assay solutions. Hydrolysis was allowed to proceed for 30 minutes or 1 hour, depending on the substrate used. The activities were corrected for pcDNA3/COS-1 cell membrane background (membranes from COS-1 cells transfected with an empty vector), as well as differences in expression levels as determined by quantitative Western blotting of each sample.
1.10. Malachite green phosphate determination procedure for nucleotidase assays
Malachite green solution (167) was made fresh on the day of use by combining, with stirring, 10 mL color reagent (60 mL concentrated sulfuric acid to 300 mL MilliQ H2O, cool to
22oC, add 0.44 grams Malachite green), 2.5 mL ammonium molybdate solution (dissolve 7.5
grams ammonium molybdate in 100 mL MilliQ H2O), and 220 µL 10% Tween 20. The solution was incubated at room temperature (22oC) with stirring for 30 minutes. 0.4 µg crude COS-1 cell
membrane protein was typically assayed. The calculated volume of MilliQ H2O (620 µL minus the volume of sample) and 160 µL of 5X buffer (resulting in 5 mM MgCl2 or 5 mM CaCl2 in 20 mM MOPS, pH 7.1 buffer after dilution) was added to a test tube (Fisher). The calculated
55 volume of sample was added, gently mixed, and pre-incubated at 37oC for 3 minutes. Substrate
was added to a final concentration of 0.25 mM to start the reaction. Hydrolysis was typically
allowed to proceed for 8 minutes (Ca-ATPase), 16 minutes (Ca-ADPase), 16 minutes (Mg-
ATPase), or 32 minutes (Mg-ADPase) at 37oC. The reaction was quenched by adding 200 µL
Malachite green solution and incubated at 22oC for 10 minutes. The absorbance at 630 nm was
measured to determine the amount of inorganic phosphate (Pi) released.
1.11. SDS-PAGE and Western blotting
Pre-cast 10-well or 15-well 4-15% polyacrylamide gradient mini-gels (Bio-Rad) were used to resolve aliquots of crude COS membrane proteins (0.5 – 2 µg, depending on the sample and the experimental purpose), usually after boiling for 5 minutes in SDS sample buffer containing 30 mM dithiothreitol (DTT). Following SDS-PAGE electrophoresis, the proteins were electro-transferred to Immun-Blot PVDF membrane (Bio-Rad) for 3 hours at 33 V in cold
10 mM CAPS/NaOH, pH 11. After transfer, the PVDF membrane was incubated for 1 hour in blocking solution (5% nonfat dry milk in Tris-buffered saline, TBS) at room temperature (22oC), and then incubated overnight at room temperature in blocking solution containing 0.02% sodium azide and a 1:5000 dilution of rabbit polyclonal primary antisera against human NTPDase3
(either KLH1 antibody, generated against the cytoplasmic C-terminal peptide (amino acid residues 515-529) of human NTPDase3 (133) or KLH11 antibody, generated against an amino acid sequence in the extracellular domain (amino acid residues 311-327) of human NTPDase3
(107)). After washing the blot in a TBS containing 0.05% Tween 20 (TTBS), a goat anti-rabbit
HRP-conjugated secondary antibody (Pierce) was applied at a 1:4000 dilution for 1 hour at room
56 temperature followed by washing in TTBS and applying the Pierce SuperSignal West Dura
Extended Duration Substrate to the PVDF membrane for 5 minutes to detect immunoreactivity by chemiluminescence. Chemiluminescence was recorded and quantified using a FluorChem IS-
8800 system (Alpha Innotech).
1.12. Deglycosylation
Partial deglycosylation was performed using Endo H (New England Biolabs) according to the manufacturer’s instructions. Briefly, COS-1 cell crude membrane protein was solubilized with 0.2% Triton X-100 containing 1 mM EDTA for 30 minutes at 22oC. The solubilized samples were then centrifuged in a Beckman air-driven centrifuge for 20 minutes at 100,000g.
The supernatant was combined with 10X Glycoprotein Denaturing Buffer (to yield final concentrations of 0.5% SDS and 1% β-mercaptoethanol) and boiled for 10 minutes. After cooling to room temperature (22oC), each sample was combined with the appropriate reaction buffer (NEBuffer G5), aliquoted, and incubated with, or without (controls), Endo H at 37oC for 1 hour or 3 hours. After incubation, the samples were combined with SDS sample buffer containing 100 mM DTT, run on an SDS-PAGE gel, and analyzed by Western blot as described above.
57 2. Project Specific Materials and Methods
2.1. NTPDase3 linker region proline residue mutagenesis
2.1.1. List of oligonucleotides/mutants made
V42C, 5’-GGTACTTGTGAGTATCACTTGCATCCAGATCCACAAGCAAG-3’;
G489C, 5’-CCACCTGTCTTTGTGTGCACCCTCGCTTTC-3’;
P52A, 5’-CACAAGCAAGAGGTCCTCGCTCCAGGACTGAAGTATG-3’;
P53A, 5’-CAAGAGGTCCTCCCTGCAGGACTGAAGTATGC-3’;
P472A, 5’-CTGACCAACCAGATCGCAGCTGAAAGCCCTC-3’;
P476A, 5’-GATCCCAGCTGAAAGCGCTCTGATCCGTCTGC-3’;
P481A, 5’-CCTCTGATCCGTCTGGCCATAGAACCACCTG-3’;
P484A, 5’-CGTCTGCCCATAGAAGCACCTGTCTTTGTGGG-3’;
P485A, 5’-CTGCCCATAGAACCAGCTGTCTTTGTGGGCAC-3’
P52A/P53A, 5’-GCAAGAGGTCCTCGCTGCAGGACTGAAGTATG-3’
P481A/P484A, 5’-CTCTGATCCGTCTGGCCATAGAAGCACC-3’
P484A/P485A, 5’-CTGCCCATAGAAGCAGCTGTCTTTGTGGGC-3’
The altered codons used to introduce the mutations are bolded and underlined (the
complementary anti-sense oligonucleotides also needed for mutagenesis are not shown). The
mutants were made in the wild-type like “free sulfhydryl-less” C10S/C501S/C509S
NTPDase3 background to eliminate all other free sulfhydryls (147). The resulting cDNA
58 constructs were sequenced by the University of Cincinnati DNA Core Facility to verify the
presence of the desired mutation and the absence of any unwanted changes.
2.1.2. Oxidative cross-linking in the presence and absence of substrate
Copper phenanthroline (CuPhen) was the oxidative cross-linker used and was prepared
fresh each day, just before use, by combining cupric sulfate and 1,10-phenanthroline (Sigma)
at a 1:3 ratio in 20% ethanol (168). Cysteine-substituted protein from COS-1 cell crude
membranes was diluted in 50 mM Tris-HCl, pH 7.5 to a final concentration of 0.1 mg/mL
and oxidized with a final concentration of 5 mM CuPhen in the absence or presence of a final
concentration of 5 mM ATP (Sigma) at 37oC for 5 minutes. The reaction was stopped by
adding an equal volume of non-reducing SDS loading buffer containing 20 mM N-
ethylmaleimide (NEM) and 20 mM EDTA. The samples were then loaded on a gel for SDS-
PAGE and Western blotting as described above.
2.2. NTPDase3 transmembrane domain polar residue mutagenesis
2.2.1. List of oligonucleotides/mutants made
S33A, 5’-GGTGGTCTTGCTTGTGGCTATTGTGGTACTTGTG-3’;
S39A, 5’-GAGTATTGTGGTACTTGTGGCTATCACTGTCATCCAGATCC-3’;
T41A, 5’- CTTGTGAGTATCGCTGTCATCCAGATCCACAAGC-3’;
Q44A, 5’-GTGAGTATCACTGTCATCGCGATCCACAAGCAAGAGG-3’;
T490A, 5’-CCTGTCTTTGTGGGCGCCCTCGCTTTCTTC-3’;
59 T495A, 5’-CCTCGCTTTCTTCACAGCAGCGGCAGCCTTG-3’;
C501A, 5’-CACAGCGGCAGCCTTGCTGCTCTGGCATTTCTTGCATAC-3’;
S33C, 5’-CTTGGTGGTCTTGCTTGTGTGTATTGTGGTACTTGTGAG-3’;
S39C, 5’-GTGAGTATTGTGGTACTTGTGTGTATCACTGTCATCCAGATC-3’;
T41C, 5’-GTGGTACTTGTGAGTATCTGTGTCATCCAGATCCACAAG-3’;
Q44C, 5’-CTTGTGAGTATCACTGTCATCTGTATCCACAAGCAAGAGGTCCTC-3’;
T490C, 5’-CACCTGTCTTTGTGGGCTGCCTCGCTTTCTTCACAGC-3’;
T495C, 5’-CACCCTCGCTTTCTTCTGCGCGGCAGCCTTGCTGTC-3’;
Q44C/S39C, 5’-GTGAGTATTGTGGTACTTGTGTGTATCACTGTCATCTGTATC-3’;
Q44C/T41C, 5’-GTGGTACTTGTGAGTATCTGTGTCATCTGTATCCACAAG-3’
The altered codons used to introduce the mutations are underlined and bolded and the
complementary anti-sense oligonucleotides also required for mutagenesis are not shown.
The polar residue to alanine substitution mutants were made in the “wild-type like” C10S
NTPDase3 background and the single as well as double cysteine mutants were made in the
“free sulfhydryl-less” C10S/C501S/C509S NTPDase3 background (which has enzymatic
properties similar to WT C10S (147)), except for the S33C/C501 mutant which was made in
the C10S/C509S NTPDase3 background (also having enzymatic properties similar to WT
C10S (147)). DNA Analysis, LLC sequenced the resulting cDNA constructs to verify the
presence of the desired mutation and the absence of any unwanted changes.
60 2.2.2. Triton X-100 nucleotidase assays
In some experiments, COS-1 cell crude membrane protein (0.05 mg/mL) was
solubilized in 1% Triton X-100 and 25 mM MOPS buffer, pH 7.1 at 22oC for 10 minutes
with occasional vortexing. In most experiments, the effects of Triton X-100 on nucleotidase
activities were measured by dilution of COS membranes into assay solutions containing
0.1% Triton X-100, using either 5 mM CaCl2 or 5 mM MgCl2 (both in 20 mM MOPS, pH
7.1 buffer) and nucleotide at a final concentration of 0.25 mM. The malachite green
phosphate assay described above (167) was used, due to the interference of Triton X-100
with the modified Amidol (Fiske and Subbarow) assay described above.
2.2.3. Oxidative cross-linking and alkylation of cysteine
Copper phenanthroline (CuPhen) was used as the oxidative cross-linker to form
disulfides from pairs of properly positioned free cysteine sulfhydryls. The reagent was
prepared just before use by combining cupric sulfate and 1,10-phenanthroline (Sigma) at a
1:3 ratio in 20% ethanol as described above (168). Cysteine-substituted COS-1 cell crude
membrane protein (0.1 mg/ml) was cross-linked with a final concentration of 0.5 mM
CuPhen in 50 mM HEPES, pH 7.5 at 37oC for 20 minutes. For samples directly analyzed by
SDS-PAGE, the reactions were stopped by adding an equal volume of non-reducing SDS
loading buffer containing 20 mM EDTA. For samples alkylated with maleimide
polyethylene glycol-5000 (MalPEG), the cross-linking reaction was stopped with 0.5 volume
of a solution to yield final concentrations of 20 mM EDTA and 1% SDS in 20 mM HEPES,
61 pH 7.5. The samples were incubated at room temperature (22oC) for 10 minutes with
occasional vortexing, followed by the addition of MalPEG to a final concentration of 5 mM.
The samples were alkylated at room temperature for 20 minutes with occasional vortexing,
and the reaction was stopped by adding an equal volume of non-reducing SDS-PAGE
loading buffer. The samples were then heated at 60oC for 10 minutes and loaded onto a gel
for SDS-PAGE followed by Western blot analysis as described above.
2.2.4. NTPDase3 nucleotidase assays in the presence of Tween 20
COS-1 cell crude membrane protein (0.25 µg/ml) was pre-incubated for 5 minutes at
o 37 C in 0.1% Tween 20 and 5 mM MgCl2 or 5 mM CaCl2 (both in 20 mM MOPS, pH 7.1
buffer). After addition of nucleotide to a final concentration of 0.25 mM to start the reaction,
hydrolysis was typically allowed to proceed for 5 minutes (Ca-ATPase), 10 minutes (Ca-
ADPase), 10 minutes (Mg-ATPase), or 20 minutes (Mg-ADPase) at 37oC. Nucleotidase
activities were measured using a malachite green inorganic phosphate assay as described
above. The absorbance at 630 nm was measured to determine the amount of inorganic
phosphate (Pi) released.
62 2.3. NTPDase3 alternatively spliced variant (NTPDase3β)
2.3.1. IMAGE clone encoding NTPDase3β
A bacterial stock of the NTPDase3β IMAGE clone (an alternatively spliced human
NTPDase3, IMAGE clone ID 5174918; GenBank Accession BC029869) in pCMV-SPORT6
vector was obtained from Invitrogen. Plasmid DNA was purified (Qiagen Miniprep) and
sequenced (University of Cincinnati DNA Core Facility), and digested with NotI and KpnI
(Fisher Scientific). A 1% low-melting point agarose gel was used to purify the gene-
containing plasmid insert after endonuclease digestion. The resulting fragment was ligated
using T4 DNA ligase (Invitrogen) into mammalian expression vector pcDNA3, which was
also digested with the same endonucleases. The resulting plasmid was transformed into
XL1-Blue Supercompetent bacterial cells (Stratagene) and plated on carbenicillin containing
LB-Agar plates.
2.3.2. List of oligonucleotides/mutants made
A511stop, sense primer: 5’-GCATACCTGTGTTCATTGAACCAGAAGAAAGAGGC-3’
A511stop, antisense primer: 5’-GCCTCTTTCTTCTGGTTCATGAACACAGGTATGC-3’
The A511Stop mutant was made in the wild type-like C10S NTPDase3 background.
The DNA Core facility at the University of Cincinnati produced the synthetic oligonucleotide
primers needed to make the NTPDase3 mutant lacking the 19 amino acids comprising the C-
63 terminal cytoplasmic tail, and sequenced the resulting cDNA construct to verify the presence
of the desired mutation and the absence of any unwanted changes.
2.4. NTPDase3 single nucleotide polymorphisms (SNPs)
2.4.1. Source of SNPs
The SNPs were identified based on data obtained from the HapMap database
(www.hapmap.org). This information is catalogued at the National Center for Biological
Technology Information website (http://www.ncbi.nlm.nih.gov) and describes the different
nucleotide variants, where they occur in the DNA, and how they are distributed among
people within populations and among various human populations in different parts of the
world.
2.4.2. List of oligonucleotides/mutants made to mimic known SNPs
I24V, 5’-CCGAACTCCAACCGTCATTGCCTTGGTG -3’;
A496V, 5’-CGCTTTCT TCACAGTGGCAGCCTTGCTG-3’;
L505F, 5’-GCTGTGTCTGGCATTTTTTGCATACCTGTGTTC-3’;
E440D, 5’-CGGTTACAAATTCACAGATGAGACTTGGCCCCAAATAC-3’;
R264Q, 5’-CTTCCAGTGCTATGGCCAGAATGA GGCTGAGAAG-3’
The altered codons used to introduce the mutations mimicking the SNPs are bolded and
underlined. The complementary anti-sense oligonucleotides necessary for the mutagenesis
64 are not shown. A QuikChange Site Directed Mutagenesis kit (Stratagene) was used to
mutate wild-type NTPDase3 in the mammalian pcDNA3 vector. The mutagenized cDNA
was transformed into Stratagene XL1 Blue Supercompetent Cells and grown overnight in 5
mL of LB broth. The mutagenized plasmid DNA was purified from the overnight cultures
using the Qiagen Miniprep system. The plasmid DNA constructs were sequenced by the
University of Cincinnati DNA Core Facility to verify the presence of the desired mutation
and the absence of any unwanted changes.
2.4.3. Treatment of wild-type and mutant NTPDase3 with concanavalin A
Concanavalin A (Con A) lectin (Sigma) was prepared at a concentration of 5 mg/mL in
20 mM MOPS buffer containing 100 mM NaCl, 1 mM MnCl2, and 1 mM CaCl2, pH 7.4.
Wild-type and mutant crude membrane preparations (2 µg) were incubated in a total volume
of 300 µL containing 5 µL of 5.0 mg/mL Con A at 37oC for 15 minutes. Afterwards,
nucleotidase assays were performed in the presence of 5 mM MgCl2 and 2.5 mM substrate
(ATP) as described above.
65 Chapter III: Results
1. Mutagenesis of NTPDase3 “Linker Region” Proline Residues
1.1. Rationale for selection and analysis of NTPDase3 “linker region” proline residues
The “linker region” is one of three structural domains in NTPDase3. It consists of a
stretch of amino acids between the TMD (location of the N- and C-terminal TM helices) and the
extracellular domain (location of the enzyme active site). It also contains several conserved
proline residues (N-terminal: P52 and P53; C-terminal: P472, P476, P481, P484, P485 – see
Figure 11). Previous studies have demonstrated that interactions between the extracellular and
TM domains occur in NTPDases (135, 169, 170); however, the specific structural features that
couple these interactions are unknown.
Proline is a structurally unique amino acid with a distinctive cyclic structure. As a result
of its side-chain binding to the backbone amide position, proline acts only as a hydrogen bond
acceptor since the amide proton is replaced by a -CH2 group, eliminating its ability to be a
hydrogen bond donor. The bulkiness of the N-CH2 group places restrictions on the conformation
of the residue preceding proline (171), thereby destabilizing α-helix secondary structures (172,
173). The elimination of the hydrogen bond donor, together with the bulkiness of the side-chain,
produces the well-known “helix-breaker” (and β-sheet breaker) secondary structure effect of
proline residues. Lastly, the ring structure of proline locks the available backbone φ dihedral angle at approximately 650, resulting in a conformational rigidity unique to
66 ___
TLAF LMVL 481 489 489 G S S G 481
VFLGLLII VFLGLLII AFLGILAF AFLGILAF VFMGVLAF VFMGVLAF VFMGVLAF VFMGVLAF VFVGTLAF VFVGTLAF VFV VFVGTLAF VFVGTLAF VFV VFVGTLAF VFVGTLAF VFVGTLAF VFVGTLAF VFMGVLAF VFMGVLAF VFMGVLAF VFMGVLAF AFLGILAF AFLGILAF VFLGLLII VFLGLLII SAFRCTVAF SAFRCTVAF QVFTGVMFF SAFRCTVAF SAFRCTVAF QVFTGVMFF QVFTGVMFF P P PP PP PP PP PP PP PP PP PP PP PP PP PP P P PP LK MR MK MD IQ IQ LPHSTYI IE IE LPHSTYI LSHSTYVFLMVL LSHSTYVFLMVL IE IE LSHSTYVFLMVL IE IE IQ IQ MD LK MR MK P P P P P P P P P P P P P P P P P P P P P P LSP LSP LST LST LIRL QIQL LIHV LIRL LIQL LIHL LIRL LIRL GLRKGTDFSSWVVLLLL GLRKGTDFSSWVVLLLL AQWRAESYGVMVAKVVF LIRL LIRL GLRKGTDFSSWVVLLLL AQWRAESYGVMVAKVVF LIRL LIRL LIHL LIQL LIRL LIRL QIQL LIHV P P P P P P P P. P P. P P P P. P P. P P P P P P P P P P AES AEN AES AES AGS AGS AES AEQ ___ C-term.TM______C-term.TM_ AES ADP LRDIQQEAFRASHTHWRGVSF ADA AES AEQ AEQ AES AEQ ADP LRDIQQEAFRASHTHWRGVSF ADA AES AES AGS AGS AES AES AEN AES P P P P P P P P P P P P P P P P P P P P P P P P P P P P 472 476481 484/5 472 476481 484/5
ACR5______ACR5______TNQI TNQI TNMI ___ TNMI ___
Horse NTPDase3 NTPDase3 Horse TNQI Opossum NTPDase3 TNMI TNMI NTPDase3 Platypus Dog NTPDase3 NTPDase3 Dog TNQI Rat NTPDase3 NTPDase3 Rat TNQI Mouse NTPDase3 NTPDase3 Mouse TNQI
Human NTPDase3 Human NTPDase1 Rat Human NTPDase4/7 NTPDase4/7 TRFL Human Human NTPDase2 NTPDase2 Human TNLI NTPDase8 Human TGMI Chimpanzee NTPDase3 TNQI NTPDase3 Chimpanzee NTPDase1 Human TNMI Macaque NTPDase3 TNQI Human NTPDase3 Human NTPDase1 Rat Macaque NTPDase3 NTPDase3 TNQI Macaque Human NTPDase2 NTPDase2 Human TNLI Human NTPDase4/7 TRFL Chimpanzee NTPDase3 TNQI NTPDase3 Chimpanzee NTPDase1 Human TNMI NTPDase8 Human TGMI Mouse NTPDase3 TNQI NTPDase3 Mouse Rat NTPDase3 NTPDase3 Rat TNQI Dog NTPDase3 NTPDase3 Dog TNQI Horse NTPDase3 NTPDase3 Horse TNQI Opossum NTPDase3 NTPDase3 TNMI Opossum Platypus NTPDase3 TNMI NTPDase3 Platypus
C-terminal Conserved Prolines Conserved C-terminal
GLKYGIVFDAG GLKYGIVFDAG GLKYGIVLDAG GLKYGIVLDAG GLKYGVVLDAG GLKYGVVLDAG GLKYGIVLDAG GLKYGIVLDAG GLKYGIV-DAG GLKYGIV-DAG GLKYGIV-DAG GLKYGIV-DAG GLKYGIVLDAG GLKYGIVLDAG ALKYGIVLDAG NVNYGIVVDCG P P P P P TDIKFGIVFDAG ENVKYGIVLDAG ENVKYGIVLDAG PP PP PP PP P PP P P PP ___ACR1______52 53 IQIHKQEVL VGLTQNKAL 34 A V N-term.TM___ SIT ALL N-terminal Conserved Prolines Prolines Conserved N-terminal Opossum NTPDase3 AITLIQINQKEVLA NTPDase3 TITLVQINQRETLS Platypus Rat NTPDase3 TLTLIQIHHPQVLS Dog NTPDase3 AITLIQFHHKEVLL NTPDase3 Horse TITVIQIHQKEVL Mouse NTPDase3 TLTLIQIRHPQVL Macaque NTPDase3 SITVIQIHKQEVL Chimpanzee NTPDase3 SITVIQIHKQEVL NTPDase3 Chimpanzee NTPDase4/7 LARVTDIEATDTNN Human Human NTPDase8 LLLVE..ATSVLL ___ 42 Human NTPDase3 Human NTPDase1 Rat Human NTPDase1 NTPDase1 Human ALLAVGLTQNKAL Human NTPDase2 LLCVP..TRDVRE
67 Figure 11. Multiple sequence alignment for portions of NTPDase sequences between the N- and C-terminal TM helices and the extracellular domain containing the enzyme active site.
Conserved proline residues found in human NTPDase3 are numbered and bolded. The locations of the cysteine substitutions useful in rat NTPDase1/CD39 (A34C and S481C, (149)), are bolded and underlined, and guided the choice of locations of cysteine substitutions used in this work as cross-linking “sensors” (V42C and G489C, which are underlined and bolded in the human
NTPase3 sequence (top line of each alignment)). To give a perspective as to the location of these conserved proline residues in the overall structure of NTPDase3, the locations of the ends of the N-terminal TM helix, ACR1, ACR5, and the C-terminal TM helix are indicated by lines and italic labeling under the alignments.
68 proline (174, 175). Thus, proline residues confer unique structural characteristics needed for some conformational changes in proteins.
Multiple sequence alignments of the N- and C-terminal linker regions that connect the
TM helices with the extracellular active site lobes of several known NTPDase3 sequences (rat
NTPDase1 and human NTPDase 1, 2, 4/7, and 8) revealed complete conservation of P472, a
high degree of conservation of P53, P476 and P481 (completely conserved in the NTPDase3
enzymes), and a lesser degree of conservation of P52, P484, and P485 (Figure 11). This led to
the hypothesis that these proline residues may provide the structural rigidity in the linker region
of NTPDase3 that is necessary to couple the movements of the active site to the transmembrane
domain during nucleotide hydrolysis. To investigate the role of the conserved proline residues in
coupling the two domain movements, proline to alanine substitutions were coupled with specific
TM helix residue to cysteine “sensor” substitutions used to monitor TMD cross-linking
efficiency in the presence and absence of ATP.
1.2. Characterization of the NTPDase3 proline to alanine mutants
Single proline to alanine mutants were made in the “wild-type like” C10S NTPDase3,
background (denoted as C10S) to eliminate the artificial dimerization via oxidation of cysteine
10 (147), and expressed the mutants in COS-1 cells. The mutants were characterized for
enzymatic activity, expression level, and proper glycosylation and delivery to the cell surface in their native oligomeric state. As seen in Figure 12, alanine substitution caused a significant decrease in the expression level of P53, P472, P476, and P485, as compared to
69 1.0
0.8 * * * * 0.6
0.4
0.2 Relative Expression Level
0.0 C10S P52A P53A P472A P476A P481A P484A P485A
Figure 12. Alanine substitution at P53, P472, P475, and P485 causes a decrease in
expression level in mutant NTPDase3 enzymes. One microgram (µg) of COS-1 cell crude membrane protein for each mutant and 0.25 – 1.0 µg of the C10S NTPDase3 as a standard curve was loaded onto an SDS-PAGE gel and detected and quantified by chemiluminescence. The data is presented as the mean relative expression level ± standard deviation from three separate experiments. The asterisks represent statistical significance (p ≤ 0.05), comparing the C10S background control mutant to the proline to alanine mutants.
70 the background C10S NTPDase3. However, once corrected for expression level, all the mutants
exhibit enzymatic activities similar to C10S except: P53A, which has a significant decrease in
both Ca- and Mg-dependent nucleotidase activities; P481A, which has a significant decrease in
both Ca- and Mg-dependent nucleotidase activities; and mutants P484A and P485A, which
demonstrate a small, but significant decrease in Ca-ATPase activity (Figure 13).
During studies which established the location of 5 extracellular disulfide bonds in human
NTPDase3, endoglycosidase H (EndoH) deglycosylation analysis was shown to be a simple and reliable assay for proper protein processing and delivery of human NTPDase3 to the cell membrane (107). All of the proline to alanine mutants were properly processed and trafficked to the cell-surface as assessed by endoglycosidase H (EndoH) (Figure 14).
Often a structural motif in proteins necessary for protein function consists of multiple amino acids. Therefore, double proline to alanine substitutions (P52A/P53A, P53A/P481A,
P481A/P484A, and P484A/P485A) were made in the C10S background to determine if replacing two prolines on the same linker arm or one on each linker arm would cause a greater decrease in enzymatic activity of NTPDase3. As shown in Figure 15, each of the double proline to alanine mutants experienced a decrease in expression level as compared to C10S. However, once corrected for expression level, all the double mutants had specific activities similar to C10S except: P53A/P481A, which demonstrated a significant decrease in Ca-dependent nucleotidase activities and Mg-ATPase; and the P484A/P485A mutant, which demonstrated a significant decrease in Ca-ADPase and Mg-dependent nucleotidase activities (Figure 16).
Taken together, this data demonstrates a clear role for P53 and P481 in facilitating the full enzymatic activity of NTPDase3. The effect of decreased enzymatic activity was also evident in the P484A/P485A double mutant, eliminating two vicinal proline residues near the C-
71
CaATPase 350 CaADPase MgATPase MgADPase 300 * * 250 *
mol/mg/hr) 200 µ
150 *
100 * * 50 * * Specific Activity ( * * 0 C10S P52A P53A P472A P476A P481A P484A P485A NTPDase3
Figure 13. P53A and P481A NTPDase3 mutants exhibit a decrease in nucleotidase activities as compared to the C10S NTPDase3 background mutant. Nucleotidase activities were assayed as described in the methods. Values represent the mean ± standard deviation from three separate experiments. Asterisks represent statistical significance (p ≤ 0.05), comparing the
C10S background control mutant to the proline to alanine mutants.
72 - + - + - + - + - + - + - + - + EndoH
Native glycosylation
C10S P52A P53A P472A P476A P481A P484A P485A
Figure 14. Alanine substitution of NTPDase3 proline residues are properly processed and presumably trafficked to the cell membrane. C10S and NTPDase 3 proline to alanine substituted mutant proteins were treated with (+) and without (-) Endo H for 1 hour at 37oC.
NTPDase3 has seven putative glycosylation sites, one being an immature high-mannose glycan sensitive to Endo H, resulting in a small shift in the electrophoretic mobility of C10S following
Endo H treatment (+ C10S lane). Protein bands with electrophoretic mobility similar to that of
C10S are marked as “native glycosylation” since they exhibit similar shifts after Endo H digestion, and this band represents the enzyme that is delivered to the cell surface. None of the mutants demonstrated any non-glycosylated “core protein” at about 59 kDa.
73 1.0
0.8 * * * 0.6 *
0.4
Relative Expression Level Expression Relative 0.2
0.0 C10S P52A/P53A P53A/P481A P481A/P484A P484A/P485A
Figure 15. Double alanine substitution of the proline residues causes a decrease in
expression level in mutant NTPDase3 enzymes. One microgram (µg) of COS-1 cell crude membrane protein for each mutant and 0.25 – 1.0 µg of the C10S NTPDase3 as a standard curve was loaded onto an SDS-PAGE gel and detected and quantified by chemiluminescence. The data is presented as the mean relative expression level ± standard deviation from three separate experiments. The asterisks represent statistical significance (p ≤ 0.05), comparing the C10S background control mutant to the proline to alanine mutants. The standard deviation for the
P484A/P485A mutant is too small for representation on this graph.
74
CaATPase CaADPase MgATPase 300 MgADPase
250
200 *
mol/mg/hr) µ
150
* 100 *
Specific Activity ( 50 * * *
0 C10S P52A/P53A P53A/P481A P481A/P484A P484A/P485A
NTPDase3
Figure 16. P53A/P481A and P484A/P485A NTPDase3 mutants exhibit a decrease in
nucleotidase activities as compared to the C10S NTPDase3 background mutant.
Nucleotidase activities were assayed as described in the methods. Values represent the mean ±
standard deviation from three separate experiments. Asterisks represent statistical significance
(p ≤ 0.05), comparing the C10S background control mutant to the double alanine substituted mutants.
75 terminal TM helix. More importantly, the results suggest that P53 on the N-terminal linker and
P481 on the C-terminal linker are functionally important, and may play a role in coupling the movements of the extracellular domain (site of nucleotide binding and hydrolysis) to the TM helices that facilitate full enzymatic activity of NTPDase3.
1.3. Rationale for selection of transmembrane “sensor” mutations and experimental
approach
To test the hypothesis that some of the conserved proline residues couple the movements of the extracellular domain to the TM helices, cysteine residues (V42C and G489C) were singly introduced in the C10S/C501S/C509S WT-like NTPDase3 (147), which is devoid of any free sulfhydryls to allow site-specific, oxidative, intermolecular cross-linking of the TM helices of human NTPDase3. These amino acids correspond to those of rat CD39/NTPDase1 that were shown to be excellent “sensors” of ATP binding effects in the extracellular domain on TM helix cross-linking (149). Thus, V42 (corresponding to A34 in rat NTPDase1) and G489
(corresponding to S481 in rat NTPDase1) in human NTPDase3 were singly mutated to cysteine
(Figure 11). These mutants were used as the “background” and “sensors” for subsequent proline to alanine mutations that were characterized for nucleotidase activity and changes in copper phenanthroline (CuPhen) cross-linking efficiency in the presence and absence of ATP.
76 1.4. Characterization of the transmembrane “sensor” mutations (V42C and G489C
NTPDase3)
The nucleotidase activities of the V42C and G489C mutants were similar to the
C10S/C501S/C509S NTPDase3 control, with a slight, but significant increase in the Ca-ATPase of V42C and a small, but significant decrease in the Ca- and Mg-dependent nucleotidase activities for G489C (Figure 17). The cross-linking efficiency of V42C and G489C in the presence and absence of 5 mM ATP was then assessed, to determine if these mutants could be used as TM specific, cross-linking “sensors” for the subsequent proline residue mutations. In the presence of 5 mM CuPhen, both V42C (Figure 18A) and G489C (Figure 18B) mutants were cross-linked to form dimers (it should be noted that the KLH1 antibody reacts much stronger with dimeric than monomeric NTPDase3, and thus only monomer bands were used in the calculation of the cross-linking efficiency). In the presence of 5 mM ATP, CuPhen-induced cross-linking efficiency of both V42C and G489C mutants decreased (the percent monomer remaining increased). Oxidative cross-linking of V42C and G489C also caused a decrease in the enzymatic activity of NTPDase3 (Figure 19), which is consistent with analogous experiments previously published using rat NTPDase1 (149). These data demonstrate the importance of the
TM helices of NTPDase3 to move relative to one another during nucleotide hydrolysis to achieve
full enzymatic activity, since CuPhen-induced disulfide restrictions caused a decrease in
enzymatic activity. Also, the data confirms the use of V42C and G489C NTPDase3 mutants as
appropriate “sensors” for active-site dependent changes of the TM helices of NTPDase3. This
provides the necessary “background” and “sensor” mutants to test the hypothesis that the cross-
talk between the active site and the TM helices may depend on the presence of some of the
77 CaATPase CaADPase MgATPase 100 * MgADPase
80 *
mol/mg/hr)
µ 60
* 40
20 * * Specific Activity (
0 C10S/C501S/ V42C G489C C509S NTPDase3
Figure 17. Cysteine substitution mutants V42C and G489C in NTPDase3 exhibits similar nucleotidase activities as the C10S/C501S/C509S NTPDase3 “background”. Nucleotidase activities were assayed as described in the methods. Values represent the mean ± standard deviation from three separate experiments. Asterisks represent statistical significance (p ≤ 0.05), comparing the C10S/C501S/C509S background control mutant to the V42C or G489C mutant.
78 100
80 * 60 D 40
M 20 Percent Monomer (M) 0 - + + - + + CuPhen
- - + - - + ATP 100
80 *
D 60
40
20 M Percent Monomer (M) Monomer Percent 0 - + + - + + CuPhen - - + - - + ATP
Figure 18. ATP binding at the extracellular domain decreases the cross-linking efficiency of V42C and G489C NTPDase3 mutants. Cross-linking was performed and the amount of monomer quantified as described in the methods. Top Panel. V42C NTPDase3 mutant.
Bottom Panel. G489C NTPDase3 mutant. The monomer (M) and dimer (D) bands are indicated. Values represent the mean percent monomer remaining ± standard deviation from three separate experiments. Asterisks represent statistical significance (p ≤ 0.05), comparing the amount of monomer remaining between the CuPhen alone and the CuPhen + ATP treated samples.
79
Control CuPhen
1.0
0.8
0.6 Activity
0.4 * Abs 660 nm * 0.2
0.0 C10S/ V42C G489C C501S/C509S NTPDase3
Figure 19. Oxidative cross-linking decreases ATP hydrolysis. Oxidative cross-linking was
conducted as described in the methods with CuPhen at a final concentration of 5 mM. The
nucleotidase reaction was carried out in 5 mM CaCl2 and ATP to a final concentration of 2.5 mM
as described in the methods. The values represent the average Abs 660 nm (Ca-ATPase activity)
± standard deviation from three separate experiments. Asterisks denote statistical significance (p
≤ 0.05), comparing the enzyme activity of the sample in the presence and absence of CuPhen.
80 conserved proline residues located in the “linker region” of NTPDase3.
1.5. Characterization of proline to alanine mutants in the V42C or G489C NTPDase3
background
To determine if the proline residues play a role in “coupling” ATP binding at the enzyme active site to movements of the TM helices, site-directed mutagenesis was used to singly substitute the conserved proline residues in the “linker regions” to alanine in both the V42C and
G489C NTPDase3 backgrounds. Proline to alanine substitutions at positions 53 and 481 resulted in significant decreases in Ca-ATPase activity as compared to the V42C background control
(Figure 20). In addition, proline to alanine substitutions at positions 484 and 485 resulted in smaller, but significant decreases in Ca-ATPase activity as compared to the V42C background control (Figure 20). Proline to alanine substitutions at positions 53, 481, and, to a lesser extent,
484, resulted in significant decreases in Ca-ATPase activity as compared to the G489C background control (Figure 21). Alanine substitution at P472 resulted in a significant increase in
Ca-ATPase activity as compared to G489C, while the P485A mutant demonstrated a small, but significant increase in Ca-ATPase activity as compared to G489C (Figure 21).
Each mutant was also oxidized with 5 mM CuPhen in the presence and absence of 5 mM
ATP to assess CuPhen cross-linking efficiency (measured by quantifying the monomer band remaining relative to the untreated control). In the V42C background, the ATP-induced decrease in CuPhen cross-linking efficiency was abolished by proline to alanine substitution at positions
53 and 481 (Figure 22). Proline to alanine substitutions at position 481 abolished the ATP-
81 450
375
* 300 * *
mol/mg/hr) µ
225 *
150
75
Ca-ATPase Activity ( 0 V42C P52A P53A P472A P476A P481A P484A P485A
V42C NTPDase3 Mutants
Figure 20. Ca-ATPase activity of the V42C proline to alanine mutants. Values represent the mean ± standard deviation from three separate experiments. Asterisks represent statistical significance (p ≤ 0.05), comparing the V42C background control mutant to the proline to alanine mutants.
82
375 ) *
300 *
mol/mg/hr µ ( 225 * *
150 *
75
Ca-ATPase Activity 0 G489C P52A P53A P472A P476A P481A P484A P485A
G489C NTPDase3 Mutants
Figure 21. Ca-ATPase activity of G489C proline to alanine mutants. Values represent the
mean ± standard deviation from three separate experiments. Asterisks represent statistical
significance (p ≤ 0.05), comparing the G489C background control mutant to the proline to alanine mutants.
83
CuPhen CuPhen + ATP 70 * * 60 *
50 * * 40 *
30
20
(M) Monomer Percent 10
0 V42C P52A P53A P472A P476A P481A P484A P485A
V42C NTPDase3
Figure 22. P53A and P481A abolish the ATP-induced decrease in cross-linking efficiency.
Cross-linking was performed and the amount of monomer quantified as described in the methods. Values represent the mean percent of monomer remaining ± standard deviation from three separate experiments. Asterisks represent statistical significance (comparing each mutant in the presence and absence of ATP, p ≤ 0.05).
84 induced decrease in CuPhen cross-linking efficiency in the G489C background (Figure 23).
Interestingly, alanine substitution at position 485 in the G489C background resulted in an inability of the mutant to cross-link at G489C by CuPhen, both in the presence or absence of
ATP (Figure 23). Taken together, these data strongly suggest specific proline residues, namely
P53 and P481, facilitate the cross-talk between binding and hydrolysis of substrate at the enzyme active site to movements or reorientations of the TM helices that are necessary to achieve full enzymatic activity of NTPDase3.
85
CuPhen
CuPhen + ATP
140
120
100
80 * * *
60 * * * 40
Percent Monomer (M) Monomer Percent 20
0 G489C P52A P53A P472A P476A P481A P484A P485A
G489C NTPDase3
Figure 23. P481A abolishes the ATP-induced decrease in cross-linking efficiency. Cross- linking was performed and the amount of monomer quantified as described in the methods.
Values represent the mean percent of monomer remaining ± standard deviation from three separate experiments. Asterisks represent statistical significance (comparing each mutant in the presence and absence of ATP, p ≤ 0.05).
86 2. NTPDase3 Transmembrane Domain Polar Residue Mutagenesis
2.1. Rationale for selection and analysis of TM polar residue mutants
Generally, TM α-helices consist of 20-30 hydrophobic amino acids that have a central region rich in aliphatic residues and phenylalanines, and short border regions enriched in tryptophan and tyrosine (176). Strongly polar residues such as arginine, asparagine, aspartic acid, glutamic acid, glutamine, histidine, and lysine are under-represented in TM helices (177,
178). Mutations of these residues often cause adverse effects on enzyme function, suggesting conserved structural and functional roles (179-181). In addition, the non-covalent association of native TM helices is reported to be mediated by several mechanisms, including: side chain packing interactions mediated by a heptad motif of leucine (182); a G-X-X-X-G motif, first described in human glycophorin A (183); and the formation of hydrogen bonds via TMD polar residues (184).
Multiple sequence alignments of the N- and C-terminal TM helices of all known
NTPDase3 sequences to date revealed complete conservation of polar residues serine 33 (S33) and glutamine 44 (Q44), high conservation of threonine 41 (T41) and cysteine 501 (C501), and a much more limited degree of conservation for serine 39 (S39), threonine 490 (T490), and threonine 495 (T495) (Figure 24A). Helical wheel predictions of the TM helices revealed that 3 pairs of polar residues, namely Q44 and T490, S39 and T495, and S33 and C501 are at approximately the same depth in the membrane, and located on the same face of the TM alpha helices (Figure 24B), suggesting that each pair has the potential for inter- or intra-molecular hydrogen bonding (Figure 24C). Therefore, the polar residues were singly mutated to alanine
87
N-terminal TM Helix C-terminal TM Helix A 33 39 41 44 490 495 501 Human 24 IIALVVLLVSIVVLVSITVIQI 45 486 VFVGTLAFFTVAALLCLAFLAYLC 509 Rat IVTLVVLLVSIVVLVTLTLIQI VFMGVLAFFTAIALLCLAFLFYLC Monkey IIALVVLLVSIVVLVSITVIQI VFVGTLAFFTAAALLCLAFLAYLC Mouse IVTLVVLLVSIVVLVTLTLIQI VFMGVLAFFTAIALLCLAFLLYLC Cat IIALVVLFLSVVVLVTITLIQF AFVGTLAFFTAVALLCLAFLVYLC Chicken VVAQAFLLLSVILIIAIAVIQI LFAGLLIFLTVLALLCLFFLVYLC Chimpanzee IIALVVLLVSIVVLVSITVIQI LFAGLLIFLTVLALLCLFFLVYLC Cow IIALVVLLVSIVVLVSITVIQI Dog IIALVVLLLSVVVLVAITLIQF AFLGILAFFTAVALLCLAFLVYLY Platypus IVALVLLLVSVVVLVTITLVQI VFLGLLIISSAVALLCLVFLVYLC Fish FVRFLCTAASIAALVAVVVIQ– VFAGLVFLFSALTIIMVVFVFIIL Frog IFSLLFLIISVSVIFAIAYVQI VFTGLLTLFTVICALCLIVLLILF Horse IVALVFLLLSVVVLVTITVIQI AFRCTVAFFSVVTLLCLAFLVYLC Oppossum VVALVVFLVSVAVLMAITLIQI VFTGVMFFFAGVALLCLLFLVYLC Pufferfish AGVMLLLLASIAALVAVAVIQ– VFAGLVFLFSALTIV------Shrew IITLVVLLVSVVVLVTITIIQM AFWGILIFFMVMAMMCLAFLVYFY Zebrafish VLAAAFMLASIAVIISIAVVQT LFTGLLLLFSVLTIITLTYLVIAL
T490 Q44 B L37 S33 A497 C501 C L30 I40 F504 F494 T41 F493 Potential TM polar V29 L505 residue hydrogen L500 A498 I34 N-TM V36 C-TM bond pairings: L27 Y507 Helix Helix I43 L491 Q44 – T490 V38 V496 V32 L502 S39 – T495 L31 A503 S39 T495 V28 A506 S33 – C501 V42 V35 A492 L499
88 Figure 24. Rationale for NTPDase3 mutations generated and analyzed in this study. Panel
A. Multiple sequence alignment of the N- and C-terminal TM helices in NTPDase3. The polar residues present in human NTPDase3 which were mutated are bolded and underlined. The sequence in GenBank for the cow NTPDase3 terminates prior to the C-terminal TM helix, and is therefore not known and not shown in the figure. Panel B. Helical wheel analysis of human
NTPDase3 TM helices. The polar TM amino acids are represented by filled black circles. The helices are depicted as viewed from the extracellular side of the cell membrane, with the size of the circles decreasing, representing the amino acids increased distance from the cell surface.
Panel C. The potential TM polar residue hydrogen bond pairings. Based on positions on the helical wheel diagram Q44, S39, and S33 on the N-terminal TM helix, are predicted to be on the same face and at the same depth as T490, T495, and C501, respectively (Panel B). Note that intra-molecular, inter-molecular (or both in the case of Q44 which can form 2 hydrogen bonds) interactions are possible for these polar residues present in the NTPDase3 dimer, which includes
2 N-terminal and 2 C-terminal TM helices.
89 and analyzed for the effect of eliminating each polar residue in the TM helices of NTPDase3.
Then the polar residues were mutated to cysteine, both singly and in pairs, in a “free sulfhydryl-
less” NTPDase3 background (C10S/C501S/C509S NTPDase3) to probe possible hydrogen
bonding pairs involving these residues.
2.2. Characterization of the NTPDase3 polar residue to alanine mutants
To explore the structural and/or functional significance of the conserved polar residues in
the N-terminal (S33, S39, T41, Q44) and C-terminal (T490, T495, and C501) TM helices of
NTPDase3, a site-directed mutagenesis strategy was utilized to singly substitute an alanine
residue (a non-polar amino acid) for each of the polar residues, and the mutants were expressed
in COS-1 cells. The “wild-type like” C10S NTPDase3, denoted as WT (C10S), was used as the
background for these mutations to eliminate the artificial dimerization via oxidation of cysteine
10 which is observed when homogenizing COS membranes expressing wt human NTPDase3
(147). The mutants were characterized for enzymatic activity, expression level, and proper
glycosylation and delivery to the cell surface in their native oligomeric state. As seen in Figure
25A and B, alanine substitution caused a significant and substantial decrease in the expression
level of S33, S39, Q44, T495, and C501 as compared to WT (C10S). However, after correcting for expression level, most of the mutants have specific enzymatic activities similar to the WT
(C10S) enzyme (Figure 25C). Exceptions include: S39A, which, although expressed relatively poorly (26% the level of WT (C10S)), exhibits significant increases in Ca- and Mg-dependent nucleotidase activities; T490A, which exhibits small, but significant decreases in Ca-ATPase specific activity and Mg-dependent nucleotidase activities; T495A, which displays relatively
90
1.0 WT (C10S)
A S33A T41A Q44A 0.25 0.5 1.0 B 0.8
* * 0.6
* * 0.4 WT (C10S) * S39A T490A T495A C501A 0.25 0.5 1.0 0.2
Protein Expression Relative
0.0 WT S33A S39A T41A Q44A T490A T495A C501A C10S NTPDase3
CaATPase CaADPase 600 MgATPase MgADPase C 500 * 400 *
300 *
mol/mg/hr) µ * 200 * * 100 * * * * 75 * * * * 50 *
25
Specific( Activity * 0 WT S33A S39A T41A Q44A T490A T495A C501A C10S NTPDase3
91 Figure 25. Alanine substitution of the polar residues causes a decrease in expression level, but similar nucleotidase activities in most mutant NTPDase3 enzymes. Panel A. Western blot analysis of NTPDase3 mutants detected by the KLH1 C-terminal antibody. One microgram
(µg) of COS-1 cell crude membrane protein for each mutant and 0.25 – 1.0 µg of the WT (C10S)
enzyme as a standard curve was loaded onto an SDS-PAGE gel, blotted, and detected and
quantified by chemiluminescence. Panel B. Relative protein expression as compared to WT
(C10S) NTPDase3. The bar graph represents the relative expression level for each mutant calculated from the WT (C10S) standard curve. Values represent the mean ± standard deviation from three separate experiments. Panel C. Activity of NTPDase3 polar residue to alanine substituted mutants, normalized for expression levels as compared to WT (C10S). Nucleotidase activities were assayed using the same amount of COS-1 cell crude membrane protein per assay.
The activities were determined by measuring the amount of inorganic phosphate (Pi) released from nucleotide substrates (ATP or ADP) in the presence of 5 mM CaCl2 or MgCl2 buffer.
Values represent the mean ± standard deviation from three separate experiments. The asterisks represent statistical significance p ≤ 0.05.
92 small, but significant increases in Ca-dependent nucleotidase activities and Mg-ADPase specific
activity, but a significant decrease in Mg-ATPase specific activity; and C501A, which shows
significant increases in Ca-ADPase and Mg-ADPase specific activities. However, the largest
change is seen for the Q44A mutant, which exhibits significant decreases in both Ca- and Mg-
nucleotidase activities.
Previous studies have shown mutation of a polar residue to a non polar residue may result in improper protein processing and/or abnormal biochemical properties of an enzyme (179). Due
to the significant decreases in expression level and changes in nucleotide hydrolysis as compared
to WT (C10S), possible changes in processing and trafficking of the mutants to the cell-surface
were examined by Endo H (107). As seen in Figure 26, all of the mutants migrated at
approximately 70 kDa with a small electrophoretic shift in mobility upon treatment with Endo H.
This data suggests that all the mutants were properly processed and presumably trafficked to the
cell-surface.
The monomeric effects of alanine substitution on the enzyme were monitored by treating
the mutant proteins with Triton X-100. This detergent is known to cause a decrease in enzymatic
activity by dissociation of NTPDase3 oligomers into monomers (147). As seen in Figure 27,
Q44A, unlike the WT (C10S) and other polar residue to alanine mutants, is insensitive to Ca-
dependent Triton X-100 detergent inhibition and the Mg-dependent Triton X-100 inhibition is
greatly attenuated. Thus, alanine substitution at Q44 may result in compromised TM helix
interactions between two different monomers, presumably accounting for the substantial
decrease in Ca- and Mg-dependent nucleotidase activities (as compared to the WT (C10S) in the
absence of detergent), and explaining the attenuation of further decrease in enzymatic activities
after Triton X-100 treatment (Figure 27). Therefore, the highly conserved and strongly polar
93 - - + - - + - - + - - + - - + - - + - - + - - + Endo H 97-
66- Native glycosylation Core protein
45- WT S33A S39A T41A Q44A T490A T495A C501A (C10S)
Figure 26. Alanine substitution of NTPDase3 polar residues are properly processed and trafficked to the cell membrane. WT (C10S) and NTPDase 3 polar reside to alanine substituted mutant proteins were treated with Endo H for 1 hour at 37°C. There are two negative
(-) control samples: the first represents total COS-1 cell membrane protein and the second one is the Triton X-100 extract control (heated at 37°C for 1 hour; neither is treated with Endo H).
NTPDase3 has seven putative glycosylation sites, one being an immature high-mannose glycan sensitive to Endo H, resulting in a shift in the electrophoretic mobility of WT (C10S). Protein bands with electrophoretic mobility similar to that of WT (C10S) are marked as “native glycosylation” since they exhibit similar shifts after Endo H digestion, and this band represents the enzyme that is delivered to the cell surface. A small amount of non-glycosylated “core protein” is seen in the T490A, T495A, and C501A mutants.
94
160 CaATPase CaADPase 140 MgATPase MgADPase 120
100
80
60
40
20
Percent No Triton Control Activity Control Triton No Percent 0 WT S33A S39A T41A Q44A T490A T495A C501A C10S NTPDase3
Figure 27. Alanine substitution at Q44 eliminates or attenuates the detergent inhibition of
Triton X-100 on NTPDase3 enzymatic activities. Triton X-100 is known to cause dissociation of wt NTPDase3 oligomers to monomers, inhibiting enzyme activity. Therefore, effects of the alanine substitutions of the polar residues on the monomeric enzyme were assessed by measuring activities after Triton X-100 solubilization. The nucleotidase activities were performed by
incubating equal amounts of COS-1 cell crude membrane protein in 0.1% Triton X-100 as
described in the methods. The activities are expressed as the percent control activity measured in
the absence of Triton X-100. Values represent the mean ± standard deviation from three separate
experiments. The dashed horizontal line indicates no change of activity in the presence of Triton
X-100.
95 Q44 residue may play a dominant role in driving TM helix associations to form the native, dimeric NTPDase3.
2.3. Characterization of the polar residues singly mutated to cysteine
The introduction of a free sulfhydryl by cysteine substitution may cause potential problems for protein folding and processing (due to inappropriate disulfide bond formation), so each polar residue was first singly substituted to cysteine in the “free sulfhydryl-less”
C10S/C501S/C509S NTPDase3 background (ensuring that the introduced cysteine is the only free sulfhydryl available for chemical reactivity). All these mutants were enzymatically active
(Figure 28A). Once corrected for expression level, only T41C exhibited specific enzymatic activities similar to the sulfhydryl-less background, unlike: S33C, which exhibited a statistically significant increase in Ca- and Mg-dependent specific activities; S39C, which exhibited a statistically significant increase in Ca- and Mg-dependent specific activities; Q44C, which exhibited an ~60% decrease in Ca- and Mg-dependent specific activities; and T490C, T495C, and C501, which all exhibited small, but statistically significant decreases in Ca- and Mg- dependent specific activities (Figure 28A). Interestingly, similar to Q44A, the Q44C mutant exhibited many of the same biochemical properties, including its attenuation of Triton X-100 detergent inhibition, unlike the other mutants (Figure 28B).
96
160 AB600 CaATPase CaATPase
) * CaADPase CaADPase MgATPase 140 MgATPase 500 MgADPase MgADPase 120
400 * * mol/mg/hr 100 µ ( 300 80 * * * * 60 200 * * * * 40 100 * * * * * * 20
Specific Activity * * * * * * * 0 0
C10S/ S33C S39C T41C Q44C T490C T495C C501 No Triton Control Activity Percent C10S/ S33C S39C T41C Q44C T490C T495C C501 C501S/C509S C501S/C509S NTPDase3 NTPDase3 C D + - + - + - + - + - + - + - DTT BME DTT NR
D D
M M
S33C S39C T41C Q44C T490C T495C C501 Q44C
97 Figure 28. Characterization of NTPDase3 polar residues to single cysteine mutants. Panel
A. Activity of NTPDase3 mutants normalized to the expression level of the C10S/C501S/C509S
NTPDase3 background enzyme. Nucleotidase activities were determined as described in the
methods. Values represent the mean ± standard deviation from three separate experiments. The
asterisks represent p ≤ 0.05. Panel B. Effects of cysteine substitution on the monomeric enzyme
activities. Equal amounts of COS-1 cell crude membrane protein were assayed in the presence
of 0.1% Triton X-100 as described in the methods. The activities are expressed as the percent
control activity measured in the absence of Triton X-100. Values represent the mean ± standard
deviation from three separate experiments. The dashed horizontal line indicates no change of
activity in the presence of Triton X-100. Panel C. Western blot analysis for the presence of
spontaneous, intermolecular disulfide bond formation. Equal amounts of COS-1 cell crude
membrane protein were treated with or without 30 mM DTT and boiled for 5 minutes as
described in the methods. The monomer (M) and dimer (D) bands are labeled. Bands migrating
above the dimer are higher order oligomers. Panel D. Analysis of the Q44C inter-molecular
disulfide linked dimer which is resistant to reduction by 30 mM DTT (Panel C). Equal amounts
of Q44C COS-1 cell crude membrane protein was either treated with no reductant (NR), 200 mM
DTT, or 2.86 M BME, and boiled for 5 minutes prior to electrophoresis, as described in the
methods.
98 In order to better understand the contributions the TMD make to the oligomeric structure
of NTPDase3, it is important to elucidate if, and how, the conserved polar residues interact between the N- and C-terminal TM helices by the cysteine substitution/disulfide bond formation strategy. This should allow development of a working model for TMD interactions. To explore the orientation and spatial proximity via the introduced cysteine residues for spontaneous, inter- molecular disulfide bond formation, the mutants were analyzed by SDS-PAGE with and without reduction by 30 mM DTT. As seen in Figure 28C, most of the cysteine mutants form some spontaneously linked disulfide dimers and higher order oligomers, however the extent of this spontaneous cross-linking is small when quantitatively comparing the two monomer bands of the treated versus untreated samples for each mutant (it should be noted that the KLH1 antibody reacts much more strongly with oligomeric forms of NTPDase3 than with monomeric NTPDase3
on Western blots). Furthermore, each of the spontaneously cross-linked cysteine mutants was
sensitive to DTT reduction, except Q44C. To ensure the spontaneous cross-linking observed in
Q44C was indeed disulfide-mediated, the sample was treated with 200 mM DTT or 2.86 M
(20%) β-mercaptoethanol (BME) at 100oC for 5 minutes prior to SDS-PAGE. As seen in Figure
28D, the spontaneously cross-linked Q44C dimer was still somewhat resistant to reduction by
this very high concentration of DTT, but was fully reduced by the very high concentration of
BME. The spontaneous cross-linking of the Q44C mutant demonstrates proper orientation and
close proximity needed for effective hydrogen bonding, and suggests that Q44 in the N-terminal
TM helix of one monomer interacts with Q44 in the N-terminal helix of the other monomer in
dimeric NTPDase3.
99 2.4. Paired cysteine mutagenesis and generation of a model incorporating the inter- and
intra- molecular hydrogen bonding pattern of the conserved polar residues in the
transmembrane helices of NTPDase3
The helical wheel analysis predicted that Q44 and T490, S39 and T495, and S33 and
C501 are on the same helical face and at the same depth in the membrane of their respective TM helices (Figure 24B). Thus, in a dimeric structure, these residues have the potential for inter- or intra-molecular hydrogen bonding with themselves or each other. To test this hypothesis, each of these putative polar residue hydrogen bonding pairs were replaced with cysteine residues and introduced into the “free sulfhydryl-less” C10S/C501S/C509S NTPDase3 background (the
C10S/C509S background for the S33C/C501 mutant). These mutants have only two free cysteines available for sulfhydryl specific cross-linking via copper phenanthroline (CuPhen) oxidation. Each double mutant cysteine pair was enzymatically active (Figure 29). To assess the proximity and orientation of each cysteine residue to one another in the TMD of NTPDase3, and to determine if each pair might preferentially interact via inter- or intra-molecular TM helix hydrogen bonding, each double cysteine mutant was alkylated with maleimide polyethylene glycol-5000 (MalPEG), oxidized with CuPhen, or treated with the combination of CuPhen followed by MalPEG. MalPEG reacts covalently to alkylate free sulfhydryls and will increases the molecular weight of NTPDase3 approximately 5 kDa for each free cysteine alkylated by
MalPEG. Each double cysteine mutant was first alkylated with MalPEG before oxidative cross- linking with CuPhen to allow differentiation of cysteine(s) present as free sulfhydryls as opposed to those being involved in spontaneous, intra-molecular cross-linking.
100
220 CaATPase A 200 CaADPase 180 MgATPase MgADPase 160
mol/mg/hr) 140 µ 120 100 80 60 40
Enzyme Activity( 20 0 C10S/C501S/C509S S39C/T495C Q44C/T490C NTPDase3
250 ) CaATPase B CaADPase MgATPase 200 MgADPase
mol/mg/hr
µ 150 (
100
50
Enzyme Activity 0 C10S/C509S S33C/C501 NTPDase3
Figure 29. Specific activities of NTPDase3 putative hydrogen bonding pair (double
cysteine) mutants. Panel A. Nucleotidase activities of NTPDase3 mutants normalized to the expression level of C10S/501S/C509S NTPDase3 background enzyme. Activity measurements were done as described in the methods and expressed as the mean ± standard deviation from three separate experiments. Panel B. Nucleotidase activities of NTPDase3 S33C/C501 mutant
normalized to the expression level of the C10S/C509S NTPDase3 background enzyme.
Nucleotidase activities were determined as described in the methods and expressed as the mean ±
standard deviation from three separate experiments.
101 As seen in Figure 30A, neither the S33C nor C501 single cysteine mutants exhibit inter- molecular cross-linking (dimer formation) after oxidation with CuPhen. Furthermore, the
S33C/C501 paired cysteine mutant was insensitive to alkylation by MalPEG or oxidation with
CuPhen as the mutant protein migrated as a monomer on the SDS-PAGE gel under all conditions. This suggests that the cysteines present at positions 33 and 501 (in the S33C/C501 mutant) are spontaneously and quantitatively intra-molecularly cross-linked, and therefore unavailable for exogenous sulfhydryl chemical reactions.
In Figure 30B, both the S39C and T495C single cysteine mutants were shown to inter- molecularly cross-link to a relatively small extent, with S39C cross-linking more efficiently than
T495C. When the S39C/T495C double mutant was oxidized with CuPhen, dimers were formed also indicating inter-molecular cross-linking. Oxidation of the S39C/T495C double cysteine mutant followed by alkylation with MalPEG, reiterates the dimer formation and shows an electrophoretic shift for the monomer, indicating the presence of free sulfhydryls even after oxidative cross-linking. This data suggests that S39 and T495 may favor inter-molecular hydrogen bonding either within or between dimeric NTPDase3.
As seen in Figure 30C, the Q44C single cysteine mutant inter-molecularly and quantitatively cross-links with CuPhen (i.e., no monomer remains). The T490C single cysteine mutant was also shown to inter-molecularly cross-link, but with lesser efficiency. When the
Q44C/T490C double cysteine mutant was alkylated with MalPEG alone, 2 bands were seen with shifted mobilities, indicating the presence of free sulfhydryls in both the monomer and the spontaneously cross-linked dimer. After CuPhen cross-linking, at least 4 bands were seen. The lowest band indicates intra-molecular cross-linking, as evidenced by its electrophoretic shift to a
102
AB- + - + + - + CuPhen - + - + + - + CuPhen - - + - + - - MalPEG - - + - + - - MalPEG
-200 kD -200kD
D D
M M S33C S33C/C501 C501 S39C S39C/T495C T495C - + - + + - + CuPhen C - - + - + - - MalPEG
-200 kDa
D
M
Q44C Q44C/T490C T490C
Figure 30. Oxidative cross-linking of NTPDase3 polar residue hydrogen bonding pair
(double cysteine) mutants. Cysteine-substituted protein (0.1 mg/ml) from COS-1 cell crude
membranes was cross-linked by CuPhen and treated with MalPEG as described in the methods.
Panel A. Lack of incorporation of MalPEG and lack of dimer formation indicates quantitative, intra-molecular cross-linking of the S33C/C501 mutant. Panel B. Inter-molecular interactions of the S39C/T495C mutant. Panel C. Inter- and intra-molecular interactions of Q44C/T490C
mutant. The monomer (M) and dimer (D) bands are indicated. Bands migrating above the dimer
are higher order oligomers.
103 position slightly above the monomer, and the lack of any shift seen after alkylation with
MalPEG. The most dense band, second from the bottom, represents the spontaneously and
CuPhen-induced, inter-molecularly cross-linked Q44C dimer, which contains two free sulfhydryls that are alkylated when treated with MalPEG as indicated by electrophoretic mobility shift higher than the CuPhen-induced Q44C dimer. The third band from the bottom represents a dimer which is apparently inter-molecularly cross-linked between both the N-terminal
(Q44C/Q44C) and C-terminal (T490C/T490C) TM helices of two monomers, since alkylation with MalPEG does not change the electrophoretic mobility of the band treated with CuPhen alone. The faint fourth band from the bottom may represent higher order oligomers.
Integrating all the cross-linking data leads to the putative hydrogen bonding pattern of the conserved polar residues as illustrated in Figure 31. The polar residues mutated, the labeling of the monomer 1 helices (denoted as N1 (N-terminal domain) and C1 (C-terminal domain)) and the solid lines representing the inter- and intra-molecular interactions of monomer 1, are bolded.
Monomer 2, containing TM helices denoted N2 and C2, and the dotted lines representing the intra-molecular interactions of monomer 2 are labeled without bolding. The filled black circles represent the polar residues with the circle size decreasing as each amino acid increases in distance through the cell membrane towards the cytosol, away from the extracellular surface (see
Figure 24 legend for a complete description of the representations in Panel A). This model places S39, T495, and, to a lesser extent, T41 on the “outside” of the interacting faces of the TM helices forming dimeric NTPDase3 (i.e., they are not directly involved in making TM polar dimeric contacts). To test this model, which suggests that S39, T495, and possibly T41, are not involved in hydrogen bonding within a dimer, but may instead be involved in higher order oligomer interactions involving multiple dimers, we used site-directed mutagenesis to make
104 A B
N2 S39 C2 Q44 N2 T41 T495 S33 Q44 T490 T490 T490 C2 C501 C501 C1 Q44 T490 Q44 S33 T495 T41 N1 S33C S33C C501 C501
S39 C1 N1
Figure 31. Model of the transmembrane domain helical interactions in the NTPDase3
dimer. Panel A. Helical wheel model depicting the putative inter- and intra-molecular
hydrogen bonding pattern of the conserved polar residues in the TMD of NTPDase3. The polar
residues mutated, the labeling of the helices, and the solid lines representing the inter- and intra-
molecular interactions of monomer 1, denoted as N1 (N-terminal domain) and C1 (C-terminal
domain), are bolded. Monomer 2, containing TM helices denoted N2 and C2, and the dotted lines representing the inter- and intra-molecular interactions are labeled without bolding. The filled black circles represent the polar residues with the circle size decreasing as each amino acid increases in distance into the membrane from the extracellular surface (see Figure 24 legend for a complete description of the representation in Panel A). Data pairing the Q44C mutation with
S39C, T41C, or T495C mutations suggests that S39 and T495 may contribute to interactions involved in forming higher order oligomers (e.g. tetramers formed by interactions between dimers), since they are available for cross-linking dimers into higher order oligomers. T41 may
105 provide specificity of helical association by taking part in the Polar-X-X-Polar (T41-V42-I43-
Q44) NTPDase3 motif, but is not available for sulfhydryl specific cross-linking of dimers into
higher order oligomers, as shown in the model by its relative inaccessibility from the surface of
the dimer helical bundle. Panel B. A 3-D cartoon model depicting the putative inter- and intra-
molecular hydrogen bonding pattern of the conserved polar residues in the TMD of NTPDase3.
Monomer 1, denoted as N1 (N-terminal domain) and C1 (C-terminal domain), is represented as
solid cylinders with polar residues important for dimer interactions underlined and solid lines
representing the helical interactions. Monomer 2, denoted as N2 and C2, is represented as dotted cylinders with dotted lines representing the helical interactions. Q44 is the “lynch-pin” of interactions responsible for dimer formation, forming presumed hydrogen bonds between the
Q44 residues present in each monomer. In addition, Q44 interacts with T490, each strongly polar glutamine being capable of forming two separate hydrogen bonds. S33 and C501 form a strong intra-molecular association within each monomer.
106 Q44C/S39C, Q44C/T41C, and Q44C/T495C double mutants. Our hypothesis was that upon
oxidative cross-linking by CuPhen, these “outside-facing” polar residues, coupled with the
spontaneous, inter-molecularly cross-linked Q44C mutant (revealing the presumed native dimer),
could form higher order oligomers upon oxidation by CuPhen. Each of the double mutants was
enzymatically active (Figure 32). As seen in Figure 33, the Q44C/S39C (and to a lesser extent,
the Q44C/T495C mutant) forms a small amount of higher order oligomers (above 200 kDa), and
these bands do not electrophoretically shift upon the addition of MalPEG, suggesting the absence
of free sulfhydryls. No higher order oligomers were evident after oxidative cross-linking of the
Q44C/T41C mutant. This data suggests that while Q44 is a main player in driving the
association of the TM helices of dimeric NTPDase3, S39, and possibly T495, may contribute to
higher order oligomer formation. T41 does not seem to play a role in promoting higher order
oligomerization, but instead is most likely teamed with Q44 to form a “Polar-X-X-Polar” motif mediating TMD association (185).
2.5. Tween 20 Stimulation of NTPDase3
Generally, the enzymatic activity of the NTPDases is inhibited by most detergents used to solubilize membrane-bound proteins (116). However, during the course of this study, we discovered that incubation of NTPDase3 with 0.1% Tween 20 increases the Mg-dependent nucleotidase activities of the human enzyme by more than 2.5 fold (Figure 34A). In contrast,
Tween 20 was shown to inhibit the enzymatic activity of human NTPDase1 and 2 by about 50 –
70% (Figure 34A), suggesting the stimulation of activity by Tween 20 is selective for
107
140 CaATPase CaADPase 120 MgATPase MgADPase 100
mol/mg/hr) µ 80
60
40
20 Enzyme Activity (
0 C10S/ Q44C S39C/Q44C T41C/Q44CT495C/Q44C C501S/C509S
NTPDase3
Figure 32. NTPDase3 S39C/Q44C, T41C/Q44C, and T495C/Q44C mutant enzymatic activities. Nucleotidase activities were determined as descried in the methods and corrected to the expression level of the C10S/C501S/C509S NTPDase3 background enzyme. Values are expressed as mean ± standard deviation from three separate experiments.
108
--++--++ - - + + - - + + CuPhen - + - + - + - + - + - + - + - + MalPEG
-200 kDa
D
M
Q44C Q44C/S39C Q44C/T41C Q44C/T495C
Figure 33. Analysis for higher order oligomeric structure by oxidative cross-linking of
S39C/Q44C, T41C/Q44C, and T495C/Q44C NTPDase3 double mutants. The mutants were treated with CuPhen and MalPEG as described in the methods. Since, unlike the parent Q44C, a small amount of higher order oligomers are observed in the S39C/Q44C and T495C/Q44C mutants (indicated by asterisks), S39 and T495 may participate in hydrogen bond interactions involved in forming these oligomers. The monomer (M) and dimer (D) bands are labeled and band(s) above 200 kDa represent a tetramer.
109
A 300
250
200
150
100
50
Percent No Tween Control Activity 0 NTPDase1 NTPDase2 NTPDase3
NTPDase Isoenzyme
B 450 400 350
300 250 200
150 100
Percent No Tween Activity No Tween Percent 50
0 Human Mouse Rat NTPDase3
110 Figure 34. The stimulatory effect of Tween 20 is specific for NTPDase3 and conserved
among species. Panel A. Mg+2-ATPase assay of human, cell surface NTPDase isoenzymes in
the presence of 0.1% Tween 20. Equal amounts of COS-1 cell crude protein were pre-incubated
with 0.1% Tween 20 for 5 minutes at 37oC before starting hydrolysis of 0.25 mM ATP for 10
minutes in 5 mM MgCl2 buffer, pH 7.5. The activities are expressed as the percent control
activity measured in the absence of Tween 20. Values represent the mean ± standard deviation
from three separate experiments. The dashed horizontal line indicates no change of activity in
the presence of Tween 20. Panel B. Mg+2-ATPase assay of NTPDase3 from 3 species in the
presence of 0.1% Tween 20. Human, rat, or mouse NTPDase3 was pre-incubated for 5 minutes
o in 0.1% Tween 20 at 37 C before hydrolysis of 0.25 mM ATP for 10 minutes in 5 mM MgCl2 buffer, pH 7.5. The activities are expressed as the percent control activity measured in the absence of Tween 20. Values represent the mean ± standard deviation from three separate experiments. The dashed horizontal line indicates no change of activity in the presence of
Tween 20.
111 + + + + Glutaraldehyde - + - - Digitonin - - + - Tween 20 -- - + Triton X-100
Dimer
Monomer
Figure 35. Tween 20 promotes oligomerization mediated by the TMDs of NTPDase3.
Panel A. Effect of Tween 20 on glutaraldehyde (lysine specific) cross-linking efficiency via the non-transmembrane regions of NTPDase3. WT (C10S) NTPDase3 was diluted to 0.1 mg/ml in
20 mM MOPS, 5 mM MgCl2 buffer, pH = 7.5. WT (C10S) was solubilized with and without
0.1% detergent (digitonin, Tween 20, or Triton X-100) for 10 minutes at 22oC prior to incubation with 5 mM glutaraldehyde at 22oC for 10 minutes. Note less monomer remaining, and more oligomers formed, in the presence of Tween 20, in sharp contrast to the large attenuation of cross-linking observed with Triton X-100.
112 NTPDase3. To confirm this observation, Figure 34B demonstrates that incubation of human,
mouse, and rat NTPDase3 with 0.1% Tween 20 results in a greater than 2.5 fold increase in Mg-
ATPase activity, indicating the stimulatory effect is not species-specific. Therefore, we hypothesized the detergent Tween 20 stimulates the enzyme activity of NTPDase3 by enhancing the oligomeric stability of the enzyme, mediated by the TM helices. Glutaraldehyde (a lysine specific cross-linker) has been frequently used to assess the oligomerization status of NTPDases
(143). NTPDase3 (WT-like C10S) was therefore glutaraldehyde cross-linked in the absence of
any detergent, as well as after treatment with 0.1% of the detergents digitonin, Tween 20, or
Triton X-100, and analyzed on a reducing SDS-PAGE gel. As seen in Figure 35, NTPDase3
treated with digitonin (which maintains activity and native oligomeric structure (116)) resulted in efficient cross-linking into dimers and higher order oligomers. NTPDase3 treated with Tween
20 resulted in more dimer and higher order oligomer formation following glutaraldehyde cross- linking as compared to the NTPDase3 treated with no detergent or treated with digitonin. In contrast, NTPDase3 treated with Triton X-100, a detergent known to disrupt the oligomeric state of NTPDase3, dramatically decreased the cross-linking efficiency of the normally dimeric
NTPDase3. This data suggests Tween 20, unlike Triton X-100 (and most other detergents), enhances NTPDase3 oligomer formation, resulting in increased nucleotidase activity.
Previous studies have shown that detergents elicit their inhibitory effects on the
membrane-bound NTPDases, but do not inhibit the soluble, extracellular domains of these same
enzymes (131). To test the hypothesis that Tween 20 stimulation of NTPDase3 activity may be
mediated by the TM helices, a soluble NTPDase3 (sNTPDase3) construct (lacking the TM
helices and short cytoplasmic N- and C-terminal tails) was treated with Tween 20 (and Triton X-
100) to assess their effect on activity. No change in the absorbance at 630 nm in the Ca-ATPase
113 Malachite green assay was observed for soluble NTPDase3 (Figure 36), indicating the Tween 20
stimulatory effect is indeed mediated by the TM helices of NTPDase3.
The S39C/T495C mutant was used to further test the hypothesis that Tween 20 increases
the enzymatic activity of NTPDase3 by promoting oligomerization of the enzyme at the TMD
level. The S39C/T495C mutant protein (in the free sulfhydryl-less NTPDase3 background) was treated with or without 0.1% Tween 20 or Triton X-100, followed by oxidative cross-linking with 0.5 mM CuPhen for 20 minutes at 37oC. As seen in Figure 37, treatment of the
S39C/T495C mutant with 0.1% Tween 20 followed by CuPhen oxidation resulted in the monomeric enzyme efficiently intra-molecularly cross-linked, and more efficiently inter- molecularly cross-linked to oligomeric forms. To the contrary, 0.1% Triton X-100 treatment of
S39C/T495C followed by CuPhen oxidation resulted in complete attenuation of dimer and higher order oligomer formation, but intra-molecular (within a monomer) cross-linking was still evident as a decrease in the electrophoretic mobility of the monomer. This data suggests the NTPDase3- selective stimulatory effect of Tween 20 might be explained by this detergent promoting oligomerization via the TM helices of NTPDase3, which contain a set of unique, conserved polar residues not present in NTPDase1 or NTPDase2.
114
0.5
0.4
0.3
0.2
Activity
0.1
0.0 NosNTPDase3 Detergent Tween 20 Triton X-100 Control
Figure 36. Detergents Tween 20 and Triton X-100 do not change the enzymatic activity of sNTPDase3. The Ca-ATPase activity of a soluble construct of human NTPDase3 was conducted as described in the methods using the Malachite green assay. The data represented the mean activity ± standard deviation from three separate experiments.
115
Tween 20 Triton X-100 0 0 0.1 0 0 0.1 [Detergent (vol %)] - + + - + + CuPhen
Dimer
Monomer
Figure 37. Effect of 0.1% Tween 20 or Triton X-100 on CuPhen cross-linking efficiency of
the TMDs of a mutant NTPDase3. S39C/T495C NTPDase3 was diluted to 0.1 mg/ml in 50
mM Hepes, pH = 7.5 and solubilized with and without 0.1% Tween 20 or Triton X-100 for 10
minutes at 22oC prior to oxidative cross-linking with or without 0.5 mM CuPhen at 37oC for 20 minutes. Note the stark contrast in results obtained using Tween 20 versus Triton X-100.
116 3. NTPDase3 Alternatively Spliced Variant
Note: This work is the culmination of a project begun and performed to a large extent by
Dr. Patrick Crawford. My role in this work, which resulted in authorship for me on the resulting paper, was to design and perform the necessary experiments requested by the reviewers and to re-write the manuscript accordingly for acceptance for publication.
3.1. Rationale and experimental approach
Alternative splicing occurs as a normal phenomenon in eukaryotes, where it greatly increases the diversity of proteins that are encoded by the genome. In humans, over 80% of gene products are alternatively spliced, including some NTPDases, namely NTPDase1, 2, and 4 (186,
187). The truncated splice variant of NTPDase1 was obtained from a human placental cDNA library, and is not yet fully characterized (188). Three splice variants of NTPDase2 have been reported: NTPDase2α, the full-length, active form of NTPDase2, which has ATPase activity and is properly processed and trafficked to the cell-surface (189); NTPDase2β, the first
NTPDase2 isoform reported (which lacks 23 amino acids in the extracellular domain) (189), and
NTPDase2γ, which is a C-terminally truncated isoform (190). These two variants of NTPDase2
(2β and 2γ) are due to differential splicing of exon 8 that results in expression of inactive proteins lacking a portion of the extracellular domain near ACR5. These splice variants are not fully biosynthetically processed and are retained in the endoplasmic reticulum. Lastly, an active, alternatively spliced form of the intracellular NTPDase4 has been described that differs in only 8 amino acids, but has broader nucleotide substrate specificity (188). Our laboratory discovered
117 and characterized (191) an alternatively spliced form of human NTPDase3, which is described
below.
3.2. NTPDase3β alternatively spliced isoform
NTPDase3β is a C-terminally truncated, alternatively spliced variant of human
NTPDase3, which was discovered by searching the IMAGE (192) clone libraries (IMAGE ID
5174918; GenBank accession BC029869) and subsequently isolated from a human lung cDNA library. The complete 1608 bp cDNA encoding NTPDase3β was sequenced and found to be a novel, isoform of NTPDase3 with a shorter 5`-untranslated region resulting in a truncated protein, maintaining the 10 extracellular cysteines forming 5 disulfide bonds (107), but lacking
apyrase conserved region 5 (ACR5), a potential N-glycosylation site (Asn 454), the C-terminal
TM helix, and the short cytoplasmic C-terminal peptide. The coding region of NTPDase3β is
identical to the full-length NTPDase3α except for two notable exceptions (Figure 38): a single A
to G point mutation at the beginning of the N-terminal TMD (a known single nucleotide
polymorphism in NTPDase3 according to the HapMap database at http://www.hapmap.org/),
which changes isoleucine 24 to valine in the N-terminal TM helix; and a premature truncation of
NTPDase3β immediately after glutamate 452, which is a valine in NTPDase3α. This truncation is generated using an alternative exon in the genomic DNA of NTPDase3 (Figure 39). Thus, both isoforms of NTPDase3 have 11 exons, the first 10 of which are the same. The alternative exons, exons 11 and 12, are the terminal exons of NTPDase3β and NTPDase3α, respectively.
The difference in the 5’-untranslated regions results from NTPDase3β having only nucleotides
59 through 68 of exon 1, presumably due to the use of an alternative promoter.
118 M F T V L T R Q P C E Q A 1 CTCCGCACAGCTAGGAGAAAAGATGTTCACTGTGCTGACCCGCCAACCATGTGAGCAAGC 60
G L K A L Y R T P T V I A L V V L L V S 61 AGGCCTCAAGGCCCTCTACCGAACTCCAACCGTCATTGCCTTGGTGGTCTTGCTTGTGAG 120
I V V L V S I T V I Q I H K Q E V L P P 121 TATTGTGGTACTTGTGAGTATCACTGTCATCCAGATCCACAAGCAAGAGGTCCTCCCTCC 180 ACR1 G L K Y G I V L D A G S S R T T V Y V Y 181 AGGACTGAAGTATGGTATTGTGCTGGATGCCGGGTCTTCAAGAACCACAGTCTACGTGTA 240
Q W P A E K E N N T G V V S Q T F K C S 241 TCAATGGCCAGCAGAAAAAGAGAATAATACCGGAGTGGTCAGTCAAACCTTCAAATGTAG 300 ACR1a V K G S G I S S Y G N N P Q D V P R A F 301 TGTGAAAGGCTCTGGAATCTCCAGCTATGGAAATAACCCCCAAGATGTCCCCAGAGCCTT 360
E E C M Q K V K G Q V P S H L H G S T P 361 TGAGGAGTGTATGCAAAAAGTCAAGGGGCAGGTTCCATCCCACCTCCACGGATCCACCCC 420 ACR2 I H L G A T A G M R L L R L Q N E T A A 421 CATTCACCTGGGAGCCACGGCTGGGATGCGCTTGCTGAGGTTGCAAAATGAAACAGCAGC 480
N E V L E S I Q S Y F K S Q P F D F R G 481 TAATGAAGTCCTTGAAAGCATCCAAAGCTACTTCAAGTCCCAGCCCTTTGACTTTAGGGG 540 ACR3 A Q I I S G Q E E G V Y G W I T A N Y L 541 TGCTCAAATCATTTCTGGGCAAGAAGAAGGGGTATATGGATGGATTACAGCCAACTATTT 600
M G N F L E K N L W H M W V H P H G V E 601 AATGGGAAATTTCCTGGAGAAGAACCTGTGGCACATGTGGGTGCACCCGCATGGAGTGGA 660 ACR4 T T G A L D L G G A S T Q I S F V A G E 661 AACCACGGGTGCCCTGGACTTAGGTGGTGCCTCCACCCAAATATCCTTCGTGGCAGGAGA 720
K M D L N T S D I M Q V S L Y G Y V Y T 721 GAAGATGGATCTGAACACCAGCGACATCATGCAGGTGTCCCTGTATGGCTACGTATACAC 780 ACR4a L Y T H S F Q C Y G R N E A E K K F L A 781 GCTCTACACACACAGCTTCCAGTGCTATGGCCGGAATGAGGCTGAGAAGAAGTTTCTGGC 840
M L L Q N S P T K N H L T N P C Y P R D 841 AATGCTCCTGCAGAATTCTCCTACCAAAAACCATCTCACCAATCCCTGTTACCCTCGGGA 900
Y S I S F T M G H V F D S L C T V D Q R 901 TTATAGCATCAGCTTCACCATGGGCCATGTATTTGATAGCCTGTGCACTGTGGACCAGAG 960
P E S Y N P N D V I T F E G T G D P S L 961 GCCAGAAAGTTATAACCCCAATGATGTCATCACTTTTGAAGGAACTGGGGACCCATCTCT 1020
C K E K V A S I F D F K A C H D Q E T C 1021 GTGTAAGGAGAAGGTGGCTTCCATATTTGACTTCAAAGCTTGCCATGATCAAGAAACCTG 1080
S F D G V Y Q P K I K G P F V A F A G F 1081 TTCTTTTGATGGGGTTTATCAGCCAAAGATTAAAGGGCCATTTGTGGCTTTTGCAGGATT 1140
119 Y Y T A S A L N L S G S F S L D T F N S 1141 CTACTACACAGCCAGTGCTTTAAATCTTTCAGGTAGCTTTTCCCTGGACACCTTCAACTC 1200
S T W N F C S Q N W S Q L P L L L P K F 1201 CAGCACCTGGAATTTCTGCTCACAGAATTGGAGTCAGCTCCCACTGCTGCTCCCCAAATT 1260
D E V Y A R S Y C F S A N Y I Y H L F V 1261 TGATGAGGTATATGCCCGCTCTTACTGCTTCTCAGCCAACTACATCTACCACTTGTTTGT 1320
N G Y K F T E E T W P Q I H F E K E E * 1321 GAACGGTTACAAATTCACAGAGGAGACTTGGCCCCAAATACACTTTGAAAAAGAAGAATA 1380
1381 GCTGATATGAAGCCTGAAGGATGTCTGGAGTCAGCCTGTTCAATACCAGCTTCAGCAATA 1440 1441 ATTAGATTAAACCAATCAATATCCTGCATTAGGCCTCTGACTATCTACTTTTACATAAAT 1500 1501 AAAATTGAGTCATCTACAAAAAAATAAGGATGTCTGGAGACATAAGGACTGTTGGTAGCA 1560 1561 TGTACTCTCTAAGCTTTGCTTCTAATAAATCTAATTACCAGTAAAGTA 1608
Figure 38. cDNA and translated amino acid sequences of NTPDase3β, the truncated splice variant of NTPDase3. The amino acids that differ between NTPDase3β and full-length
NTPDase3α are indicated in bold. The N-terminal transmembrane domain is underlined and the apyrase conserved regions (ACRs) are labeled and shaded.
120 Start 1 68 69 121 122 TM1 250 251ACR1 369 NTPDase3α 12 3 4 NTPDase3β 1 10 11 6364 192 193 311
P-2 P-1 ACR2 ACR3 370 521 522 682 683ACR4 907 908 1181 5 6 7 8
312 463 464 624 625 849 850 KLH111123 ACR1a ACR4a A511Stop Stop ACR5 P-α 1182 1293 1294 1432 1433 2793 91011 12
1124 1179 1180 1374 1375 1608 TM2 KLH1 P-β St op
Figure 39. A diagram of the exon map of the human NTPDase3 gene representing the splicing events that occur. The boxes represent the 12 exons, which are numbered (exon numbers inside ovals), and the introns are depicted as peaks and valleys connecting the exons above and below the boxes. The cDNA sequence is indicated by the dark line with NTPDase3a above and NTPDase3b below the exons. The seven ACRs are marked by crosshatched boxes within the exons. The transmembrane helices (TM1 and TM2) are represented by boxes filled with vertical lines. The two antibodies (KLH1 and KLH11) used are indicated by shaded boxes within the exons.
121 3.3. Expression and enzymatic activities of NTPDase3 isoforms
Ca- and Mg-dependent nucleotidase assays were performed on the crude membrane preparations of COS-1 cells transiently transfected with NTPDase3α or NTPDase3β. ATP and
ADP nucleotide hydrolysis by NTPDase3α was, as expected, detected in the membrane preparations (12, 107, 140). However, there was no measurable ATPase or ADPase activity detected for NTPDase3β in the membrane preparations. COS cell membrane fractions expressing NTPDase3β protein were further tested for activity using alternative nucleotide substrates (AMP, GDP, UDP, UTP, IDP, CDP, and CTP), but no detectable hydrolysis was observed using any of the nucleotide substrates (191). The cell culture media was assayed for nucleotidase activity and NTPDase3 protein, to determine if the truncated form of NTPDase3 was being released as a soluble enzyme after cleavage of the single TM helix, similar to what has been observed for NTPDase5 and 6. Neither nucleotidase activity nor NTPDase3 protein was detected in any of the soluble fractions (191). Western-blot analysis performed utilizing an antibody (KLH11) against an internal peptide sequence (191), demonstrated that the COS cell crude membrane preparations contained NTPDase3β protein. There was a clear difference in the molecular masses of the two isoforms of NTPDase3 as predicted from their respective amino acid sequences (191). The non-glycosylated theoretical molecular mass of the full-length
NTPDase3α isoform is 59,190 Da, compared to 50,748 Da for the non-glycosylated
NTPDase3β isoform.
122 3.4. Co-Transfection of NTPDase3α and NTPDase3β
Membrane-bound NTPDases, NTPDase3 being one, are expressed on the cell-surface
mostly as homodimers. To assess the effect of expressing NTPDase3α and
NTPDase3β together, COS-1 cells were transfected with NTPDase3α, NTPDase3β, the
A511Stop-NTPDase3 mutant, or a combination of the three DNAs, as shown in Table 3. The co-transfected COS cell membranes were characterized based on the expression level of
NTPDase3 protein (Figure 40) and nucleotidase activities (Table 3 and Figure 40). The
A511Stop-NTPDase3 mutant is a fully active, truncated NTPDase3α protein lacking only the cytoplasmic C-terminal sequence (i.e. amino acids 511-529). This NTPDase3 mutant is not recognized by the KLH1 C-terminal antibody, but still reacts with the internal KLH11 antibody.
These properties allowed the A511Stop mutant to be used as a control in the co-transfection experiments.
Co-transfection with NTPDase3β caused a significant decrease in Ca- and Mg-dependent
nucleotidase specific activities that were substantially greater than can be explained by
NTPDase3β simply being an inactive isoform and having no effect on NTPDase3α expression and activity (Table 3). These activities are approximately 40% of the nucleotidase activities
measured for COS cells transfected with NTPDase3α alone (Table 3, compare the “8,0,0” row
with the “4,4,0” row). Smaller, but similar decreases in activity are also seen when NTPDase3α
is co-transfected with NTPDase3β and A511Stop (the “4,2,2” row). More importantly, when the
123 Expression levels relative to NTPDase3α transfected alone Specific activity (µmol/mg/hr, µg cDNA used to transfect COS cells (from Figure 4) uncorrected for expression levels)
Using Using KLH11 KLH1 detected detected active, upper Ca- Mg- NTPDase3α NTPDase3β A511Stop bands bands Ca-ATP ADP Mg-ATP ADP
8 0 0 100 100 203±31 38±3 119±29 32±6
4 0 4 60±14* 82±1** 173±20+ 34±3+ 98±24# 28±4#
4 4 0 30±6* 34±5** 73±7+ 15±2+ 41±9# 14±1#
4 2 2 31±4 49±6 93±4 19±1 56±6 17±1
Table 3. Results of co-transfection of different NTPDase3 proteins in COS cells. This table summarizes the expression level and enzymatic properties of NTPDase3α when co-transfected with either or both the A511Stop-mutant or NTPDase3β. The relative protein expression level and enzyme activity measurements of NTPDase3α in the presence of A511Stop are statistically different than it is in the presence of the same amount of DNA encoding NTPDase3β. The p- values given for this table were calculated by comparing row “4,0,4” with “4,4,0” and each have an n=3. (*p-value <=0.04, **p-value <=0.003, +p-value<=0.004, #p-value<=0.03)
124
KLH1 KLH11
1 2 3 4 5 6 7 8 9 10
NTPDase3α + NTPDase3α A511Stop NTPDase3β
µg NTPDase3α 8 4 4 4 0 8 4 4 4 0 µg NTPDase3β 0 0 4 2 8 0 0 4 2 8 µg A511Stop 0 4 0 2 0 0 4 0 2 0
Figure 40. Western blot analysis of co-transfection experiments using the KLH1 or KLH11 antibody. COS-1 cells were transfected with a total of 8 µg of NTPDase3 cDNA. Primary antibodies used were KLH1 for lanes 1-5 and KLH11 for lanes 6-10. KLH1 is reactive against the C-terminal cytoplasmic amino acid sequence and only detects NPTDase3α. KLH11 is reactive against a portion of the extracellular domain and recognizes NTPDase3α and A511Stop
(active, upper bands) as well as NPTDase3β (inactive, lower bands).
125 A511Stop-NTPDase3 mutant was co-transfected with full-length NTPDase3α, relatively small decreases in specific activity and expression levels were noticed. This was substantially and statistically significantly different in every case than those observed when co-expressing
NTPDase3α with NTPDase3β (compare the “4,0,4” and “4,4,0” rows in Table 3). The comparison of these two rows is the most important aspect of Table 3 and demonstrates a clear difference between a “control”, non-interfering, co-transfected DNA (A511Stop in the “4,0,4” row) from a co-transfected DNA that results in inhibition of expression of a full-length, wild- type enzyme (the NTPDase3β isoform, in the "4,4,0" row). Thus, co-transfection of NTPDase3β causes a specific effect of decreased expression level of NTPDase3α that is not seen in the presence of the active, normally processed, artificially truncated A511Stop-NTPDase3 mutant.
The A511Stop-NTPDase3 mutant only decreases NTPDase3α expression level by about 40%
(60 ± 14 %) (see Table 3 and lanes 1 and 2 and 6 and 7 of Figure 40), while NTPDase3β more dramatically decreases the expression level of NTPDase3α by about 70% (30 ± 6 %), in the presence and absence of the A511Stop-NTPDase3. Therefore, the active and properly processed
A511Stop was used as a control for non-specific effects that might be caused by any other DNA present during co-transfection, affecting the amount of DNA taken up by the COS cells, or the ability of the COS cells to express the transiently transfected NTPDase3α DNA.
The KLH11 antibody is able to detect all NTPDase3 proteins used in this set of experiments (Figure 40). On the basis of size, by Western blot analysis, NTPDase3α and
NTPDase3β, as well as the A511Stop-NTPDase3 and NTPDase3β are distinguished from one another, but NTPDase3α and the A511Stop-NTPDase3 mutant cannot be distinguished from one another (Figure 40). Thus, KLH11 can be used to distinguish between the active forms of
NTPDase3 (NTPDase3α and A511Stop – the upper bands in lanes 6-9 in Figure 40) and the
126 inactive NTPDase3β (the lower bands in lanes 8-10 in Figure 40). Therefore, the data plotted in
Figure 41 is calculated from the nucleotidase activities reported in Table 3, after correcting for the amount of active and properly processed NTPDase3 by dividing the specific activities in
Table 3 by the relative densities of the upper, active band detected by the KLH11 antibody (as depicted in lanes 6-9 of Figure 40 and quantified in Table 3). Thus, as seen in Figure 41, the activity per molecule of fully processed NTPDase3 (NTPDase3α plus A511Stop) is not changed by the presence of either 2 or 4 µg of NTPDase3β DΝΑ (or by the presence of the A511Stop
DNA), suggesting that the co-expression of NTPDase3β does not inactivate NTPDase3α (or
A511Stop NTPDase3α) through a direct protein-protein interaction, as might occur via hetero- dimerization-induced inactivation of NTPDase3α by NTPDase3β contained in the same oligomer. Thus, both NTPDase3α and the active A511Stop mutant exhibit the same nucleotidase activity per molecule expressed, as detected by the upper band reactive with the
KLH11 antibody, resulting in no significant differences between any of the nucleotidase values,
after correction for expression level (see Figure 41, where none of the values are statistically
different from the “8,0,0” row controls). This suggests that the presence of NTPDase3β protein
interferes with the biosynthetic processing of NTPDase3α protein, decreasing the amount of
NTPDase3α that is properly processed and delivered to the cell-surface in an active form, but
does not alter the specific activity of the smaller amount of NTPDase3α that is properly
processed.
127
Figure 41. Percent activity of co-transfections normalized per molecule active protein.
Specific activity was calculated from the amount of inorganic phosphate released during the reaction as µmol Pi released per mg active protein per hour (see Table 3) and normalized to the expression level of active forms of the protein, NTPDase3α and A511Stop, quantified using the upper bands observed in the KLH11 Western blot (see Figure 40, lanes 6-9, and Table 3). This normalized specific activity was used to generate the percentage normalized specific activities, defining the NTPDase3α, KLH11 normalized, specific activity as 100%. The data is expressed as the mean percent NTPDase3a specific activity ± standard deviation for multiple assays performed on three independent experiments.
128 4. NTPDase3 Single Nucleotide Polymorphisms (SNPs)
4.1. Rationale and experimental approach
Single nucleotide polymorphisms (SNPs) are single nucleotide base changes that can
alter the primary structure of a particular protein. Often, these SNPs lead to phenotypic changes
that result from alterations of the structure and/or function of a protein. Human NTPDase3 has
five non-synonymous, coding region SNPs (I24V, R264Q, A496V, L505F, and E440D), which
we constructed and expressed in COS-1 cells to characterize these mutant proteins. The mutant proteins were characterized based on their enzymatic activities, expression level, Triton X-100 detergent sensitivity, and concanavalin A (ConA) and Tween 20 stimulatory effects.
4.2. Characterization of NTPDase3 SNPs
Using site-directed mutagenesis, the five SNPs were made in the wt NTPDase3 background. All the mutants exhibited similar Ca- and Mg-dependent nucleotidase activities, except: I24V, which demonstrated a small, but significant increase in nucleotidase activities;
L505F, which demonstrated a small, but significant increase in nucleotidase activities; and
R264Q, which demonstrated a substantial and significant decrease in nucleotidase activities
(Figure 42).
The Mg-ATPase activity of each mutant was stimulated approximately 2.5 fold by ConA
(Figure 43), inhibited approximately 90% by Triton X-100 (Figure 44) and stimulated more than
2-fold by Tween 20 (Figure 45), similar to the wt NTPDase3 enzyme. Taken together, these
129 CaATPase CaADPase MgATPase 200 MgADPase
150 mol/mg/hr) µ * 100
50 * * Specific Activity ( * 0 WT I24V R264Q E440D A496V L505F NTPDase3
Figure 42. Enzymatic activity of NTPDase3 SNPs. The nucleotidase activities were conducted as described in the methods and corrected for the expression level of WT NTPDase3.
The expression levels for the mutants, relative to expressed wt NTPDase3: I24V, 0.97; R264Q,
1.23; E440D, 1.22; A496V, 1.14; and L505F, 0.86. The data is presented as the mean activity ± standard deviation from three separate experiments. The asterisks represent statistical significance (p ≤ 0.05), comparing the wt NTPDase3 with the SNPs.
130 300
250
200
150
100
50 % no Con A Control
0 wt I24V R264Q E440D A496V L505F NTPDase3
Figure 43. ConA stimulatory effect on NTPDase3 SNPs. The COS cell membrane proteins were treated with ConA and the Mg-ATPase activity determined as described in the methods.
The data is presented as the mean Mg-ATPase activity ± standard deviation from three separate experiments.
131 20 18 16 14 12 10 8 6 4 2
% no Triton X-100 Control 0 wt I24V R264Q E440D A496V L505F NTPDase3
Figure 44. Triton X-100 Detergent effect on NTPDase3 SNPs. The nucleotidase activities were performed by incubating equal amounts of COS-1 cell crude membrane protein in 0.1%
Triton X-100 as described in the methods. The activities are expressed as the percent control activity measured in the absence of Triton X-100. Values represent the mean ± standard deviation from three separate experiments.
132 300
250
200
150
100
50 % no Tween 20 Control 0 wt I24V R264Q E440D A496V L505F NTPDase3
Figure 45. Tween 20 Stimulatory effect of NTPDase3 SNPs. The Mg-ATPase activity was conducted as described in the methods using the Malachite green assay. The data represented the mean ± standard deviation from three separate experiments.
133 data suggest that, with the exception of R264Q that is located near the active site, these SNPs may not be functionally important. The substantial decreases in nucleotidase activities of R264Q could cause alterations in purinergic receptor signaling via less efficient hydrolysis of ATP and
ADP agonists. These human NTPDase3 SNPs were not pursued any further at this time, since those SNPs that are relevant to this thesis (due to their location in the TMD of NTPDase3: I24V,
A496V, and L505F) were not different from the wild-type human NTPDase3, based on this preliminary biochemical characterization.
134 Chapter IV: Discussion
1. Regulation of the Enzymatic Activity of the Membrane-bound, Cell Surface
NTPDases
The most direct and obvious way to regulate the activity of an enzyme is via its active site. However, the recent analysis of the crystal structure of the extracellular domain of rat
NTPDase2 (146), which contains the active site, does not tell the entire story of the enzymatic mechanisms regulating the NTPDases. This is because nucleotide hydrolysis of the membrane- bound, cell surface NTPDases appears to be regulated by a complex system of coordinated movements between (dependent on the oligomeric stability and structure) and within (dependent upon interactions between the enzyme active site and the transmembrane domains) monomers.
To better understand the structural features regulating hydrolysis (and therefore function), it is necessary to elucidate what the major structural elements of the membrane-bound, cell surface
NTPDases are, and how they work together for optimal enzyme function.
Membrane-bound NTPDase3 expressed on the cell surface has three major structural domains: a large extracellular domain containing the active site, a “linker region”, and a transmembrane domain consisting of 2 TM helices per monomer (134). The extracellular domain contains five disulfide bonds that are conserved among all cell surface NTPDases, which when disrupted in NTPDase3 yield five distinct biochemical phenotypes (107). There are also five apyrase conserved regions (ACRs) in the extracellular region, which are common to all members of the NTPDase family (139), and share sequence homology with the actin/heat shock protein/sugar kinase superfamily of proteins. ACR1 and ACR4 were identified as phosphate
135 binding domains in the extracellular active site by homology with similar regions in the actin superfamily of proteins (193). Also by homology with the actin superfamily of proteins, it has been hypothesized that the extracellular lobes of the NTPDases, upon functional state changes
(i.e. nucleotide binding), undergo an interdomain hinge (butterfly-like) motion of the two lobes during nucleotide hydrolysis.
The “linker region” is a stretch of amino acids in the primary sequence of NTPDase3, between the TMD (location of the N- and C-terminal TM helices) and the extracellular domain
(location of the enzyme active site). The linker region contains a set of conserved proline residues (N-terminal: P52 and P53; C-terminal: P472, P476, P481, P484, and P485). Grinthal and Guidotti, using rat NTPDase1, demonstrated an intra-protein signal transduction phenomenon involving ATP binding in the extracellular domain, which resulted in movement of the transmembrane (TM) helices during nucleotide hydrolysis (149). Thus, NTPDase activities and substrate specificities are modulated not only by mutation of amino acids in the active site
(134, 194), post-translational modifications such as glycosylation (143, 190) and N-acetylation
(195) in the extracellular domain, but also by membrane perturbations of the TM helical interactions (137, 169) and modification of TMD cysteine residues by p- chloromercuriphenylsulfonate (pCMPS) (147, 148).
The transmembrane domain (TMD) of NTPDase3 contains two TM helices, one near the
N-terminus and one near the C-terminus, each of which contains a set of conserved polar residues (N-terminal: S33, S39, T41, and Q44; C-terminal: T490, T495, and C501). Most studies regarding the role of the TMD of the membrane-bound, cell-surface NTPDases have focused on oligomerization, since the predominant, active form of these enzymes appears to be a dimer, although higher-order oligomers are also observed, and the soluble NTPDases (which
136 lack TMDs) are monomeric (131, 133, 144). However, no structural data exists, and no site- directed mutagenesis studies have been performed, to perturb and analyze helix interactions in the TMD of NTPDase3.
The objectives of this dissertation were to: 1. elucidate the structural elements that mediate the intra-protein signal transduction between substrate binding at the extracellular domain and movements and/or re-arrangements of the TM helices during nucleotide hydrolysis; and 2. determine how the N- and C-terminal TM helices interact within and between monomers to form native, dimeric NTPDase3. The experimental evidence presented in this dissertation provides novel insights regarding the enzymatic mechanism of NTPDase3 via : 1. the conserved proline residues linking the active site to TMD movements; 2. the conserved polar residues stabilizing the TMD and promoting oligomerization; 3. the selective stimulation of the enzymatic activity of NTPDase3 by the detergent Tween 20; and 4. the possible modulation of purinergic receptor signaling via expression of the alternatively spliced variant of human NTPDase3,
NTPDase3β, or the R264Q NTPDase3 SNP.
2. Structural and Functional Roles of Specific Structural Elements in NTPDase3
2.1. The conserved proline residues are important components of the linkage between
the active site and transmembrane domain movements
Similar to the results obtained using rat CD39/NTPDase1 (149), mutations in both TM helices of human NTPDase3 near the extracellular surface (V42C and G489C in NTPDase3) cross-linked efficiently in the presence of CuPhen. In the presence of ATP, the CuPhen-induced
137 cross-linking efficiency of the V42C and G489C NTPDase3 mutants was attenuated, resulting in
an increase in the amount of monomer remaining after treatment. Disulfide cross-linking of
V42C and G489C via CuPhen also inhibited the nucleotidase activity of the mutants. This data suggests the importance of the TM helix rearrangements and/or movements during nucleotide hydrolysis that facilitate full enzymatic activity and may be generalized to all cell surface
NTPDases.
When proline to alanine mutations were added to the cross-linking “sensor” background
NTPDase3 enzymes (V42C and G489C), alanine substitution at positions P53 and P481 in the
V42C sensor background caused a substantial decrease in activity, while a significant, but smaller decrease in activity was noted for P484 and P485. Alanine substitution at positions P53 and P481 in the G489C sensor background resulted in a substantial decrease in activity, while the
P484A mutant exhibited a smaller decrease in activity. This data suggests P53, P481, and P484, which are completely conserved in all sequenced NTPDase3 enzymes, but not in all other
NTPDases, are needed for optimal enzymatic activity of NTPDase3.
Assessment of the proline to alanine mutations for possible “uncoupling” effects revealed that mutations at positions P53 and P481 in the V42C background were indeed capable of uncoupling ATP binding at the active site to reorientations or movements of the TM helices, as assessed by abolishment of the ATP-dependent decrease in TM cross-linking efficiency. In the
G489C background, proline to alanine substitution at position P481 also caused this effect.
Surprisingly, alanine substitution at position P485 abolished the CuPhen-induced cross-linking of G489C both in the presence and absence of ATP, although this was not true for cross-linking of V42C after alanine substitution at the same P485 residue. This suggests that the placement of the cysteine residue used as the cross-linking sensor is important for the ability to measure the
138 effects of the proline mutations, and that such proline to alanine mutations may affect the TM
helix closer to the mutation more than the TM helix on the other end of the protein. Thus,
alanine substitution at P485, which is only 4 amino acids away from the G489C mutation used as
a sensor in the C-terminal helix, changes the orientation or freedom of movement of the C-
terminal TM helix of NTPDase3 in a way that abolishes intermolecular, CuPhen-induced cross-
linking at G489C, but not cross-linking at V42C, which is located in the N-terminal TM helix.
Thus, consistent with our hypothesis, some proline to alanine mutants exhibited a decrease in enzymatic activities, an “uncoupling” of the interactions between the active site and
TMD, or both. Importantly, other proline to alanine mutants also located in the linker region had little, if any, effect, demonstrating that the results obtained are not simply non-specific artifacts, in the sense that any proline to alanine substitution in this area of the protein would result in decreases in activity and changes in ATP-induced cross-linking efficiencies as a result of adverse effects on protein folding or processing. Only alanine substitution at positions P53 and P481 resulted in both decreases in nucleotidase activities and abolishment of the decreased cross- linking efficiency of the TM helices in the presence of ATP, suggesting that these two proline residues are important components of the linkage between movements of the extracellular and
TM domains necessary for optimal function of NTPDase3.
2.2. The conserved polar residues stabilize transmembrane domains and promote
oligomerization
The significant decreases in expression levels of S33A, S39A, Q44A, T495A, and C501A mutants were not surprising, since similar polar-to-apolar amino acid substitutions in the TMDs
139 of several unrelated proteins are disease-phenotypic mutations, often accompanied by adverse effects on processing and normal expression of the mutant proteins (196). However, each of the
NTPDase3 polar residues to alanine mutants was properly glycosylated and apparently trafficked to the cell surface, as suggested by Endo H deglycosylation analyses. Somewhat surprising, most of the polar residue to alanine mutations, once corrected for their lower expression levels, did not adversely affect enzymatic activity. Taken together, these data suggest that the conserved polar residues in the TMD of NTPDase3 are important for proper protein folding and normal protein expression, presumably via TM hydrogen bonding.
Interestingly, the Q44A and Q44C mutants exhibited similar enzymatic characteristics, specifically, a decrease in nucleotidase activities, expression level, and the relative insensitivity to Triton X-100 detergent inhibition (Triton X-100, as well as many other detergents, has been shown to inhibit activity of NTPDase3 as well as most other cell surface NTPDases (116)). In fact, Ca-dependent activities actually increased for the Q44 mutants, probably due to unmasking of active sites by making a sub-population of sealed COS membrane vesicles leaky to the nucleotide substrates. The Mg-dependent activities are decreased by Triton, but this inhibitory effect is greatly attenuated as compared to the WT (C10S) sample and the other polar residue mutants. Thus, it seems there may be either a second component of the inhibitory effect of
Triton X-100 that is detected when assaying for Mg-nucleotidase activities, or a stabilizing effect on NTPDase3 mediated by calcium. Nevertheless, the insensitivity of the Q44A mutant to
Triton inhibition suggests that alanine substitution at Q44 may result in compromised TM helix interactions between two different monomers, presumably accounting for the approximately
50%-70% decrease in specific nucleotidase activities (as compared to the WT (C10S), both in the absence of detergent), and explaining the attenuation of further decreases in enzymatic
140 activities after Triton X-100 treatment (Triton X-100 is known to dissociate the oligomeric
structure of several NTPDases, including human NTPDase3). In addition, the spontaneous,
intermolecular disulfide bond formation observed in the Q44C mutant (resistant to even very
high concentrations of DTT, but could be reduced by high concentrations of the more
hydrophobic reduction agent, BME) most likely limits the dynamic motions of the TM helices.
This decreases activity, since both appropriate hydrogen bonding and movement of the TM
helices during nucleotide hydrolysis are necessary to achieve full enzymatic activity.
Taken together, this data is consist with results and conclusions of Sal-Man et al (184,
185), suggesting the existence of a Polar-X-X-Polar (T41-X-X-Q44) TM-TM association motif in the N-terminal helix of NTPDase3. In that study, Sal-Man et al demonstrated the importance of a strong polar residue present in this motif to drive the association of the TM helices in the E.
coli Tar-1 receptor. We conclude that Q44 of one monomer is in close proximity and in the
correct orientation to the Q44 of the other monomer in a dimer, and that the N-terminal TM
helices of dimeric NTPDase3 interact efficiently near the extracellular surface of the cell
membrane. The current data demonstrating the importance of the strongly polar Q44 residue,
which is located at the interface of the N-terminal TM helix with the extracellular portion of
NTPDase3, is also consistent with data published by Grinthal and Guidotti (149) using rat
NTPDase1. In that study, a cysteine substitution and oxidative cross-linking strategy was also
utilized, and identified strong inter- and intra-molecular interactions between the TM helices of
rat NTPDase1 involving residues near the extracellular surface of the membrane, and further
demonstrated that disulfide bond formation between two TM helices decreased enzymatic
activity of NTPDase1.
141 Our laboratory previously noted the attenuation of Triton X-100 inhibition following site-
directed mutagenesis of lysine 79 in the extracellular domain of NTPDase3 (145). Lysine 79 is
conserved in all membrane-bound, cell surface NTPDases, but not in the soluble, monomeric or
intracellular NTPDases. It is also located in a region hypothesized to stabilize oligomerization
(between ACR1 and a glycosylation site, N81) via the extracellular domain. Mutation of lysine
79 to any amino acid besides arginine (which maintains the positive charge) rendered NTPDase3
insensitive to Triton inhibition. Therefore, it is likely that, in addition to the highly conserved,
and strongly polar, Q44 in the TMD, amino acid residues in the extracellular portion of
NTPDase3 also contribute to interactions that enable and stabilize dimerization of NTPDase3.
Our laboratory previously determined that the dominant oligomeric structure of
NTPDase3 is a dimer, with higher order oligomers also possible, including a tetramer, suggested
to be a “dimer of dimers” (147). In that study, it was noted that the interactions between two
monomers in each dimer is stronger than the interactions between the two dimers forming the
tetramer (147). The cysteine substitution/CuPhen cross-linking strategy used in this study
revealed: 1. Q44C appears to form strong intermolecular disulfides with Q44C and, being a
strongly polar residue with the ability to form two hydrogen bonds, Q44C also formed
intramolecular disulfides with T490C; 2. S33C and C501 appear to spontaneously and
quantitatively form intramolecular disulfides with each other; and 3. S39C and T495C
inefficiently form some intermolecular disulfides, resulting in higher order oligomers. This
cross-linking data suggests a putative hydrogen bonding pattern of the conserved polar residues
that places S39 and T495 on the “outside” of the TM helices forming dimeric NTPDase3.
Additional cross-linking data suggested S39 and T495 may be involved in higher order oligomer interactions (e.g., to form the “dimer of dimers” (147)).
142 These cross-linking results suggest that Q44 is the “lynch-pin” residue in the association
of the N-terminal TM helixes to form dimeric NTPDase3. S39 (and possibly T495) may form intermolecular hydrogen bonds to promote interactions that facilitate formation of the tetrameric
and higher order oligomeric forms of NTPDase3. T41, due to its inability to form higher order
oligomers when combined with Q44C, is not likely to be directly involved in higher order
oligomer formation, but instead may work in concert with Q44 to provide the specificity of the
N-terminal helix associations via a “Polar-X-X-Polar” (T41-X-X-Q44) TM-TM association
motif (197).
Lastly, during the course of this study, the detergent Tween 20 was shown to exhibit an
isoenzyme specific, species-conserved, stimulatory effect on the nucleotidase activity of
NTPDase3. This stimulation was not observed when a soluble construct, comprising the
extracellular domain of human NTPDase3, was treated with Tween 20 under identical
conditions. This discovery of the stimulatory effect of Tween 20 on full-length NTPDase3 may
facilitate future structural studies and serve as a useful tool for identifying the presence and
importance of NTPDase3 in various tissues and processes. Considering this stimulatory effect is
isoenzyme-specific (not observed in the closely related human NTPDase1 or NTPDase2), it is
not unlikely that the unique TMD hydrogen bond interactions mediated by the conserved polar
residues of NTPDase3 promote the association of the TM helices in the presence of Tween 20. It
should be noted that the conserved NTPDase3 TMD polar residues examined in this dissertation
are not conserved in other membrane-bound NTPDases, suggesting that the results reported are
specific for NTPDase3, and that other NTPDases have different mechanisms and amino acids facilitating TM helix interactions and TMD oligomeric associations. This most likely contributes to the differential susceptibility to membrane perturbation (e.g., detergent activation
143 and inactivation) (136, 137) and substrate specificities (119, 170) seen in different family
members of the NTPDase enzymes.
2.3. NTPDase3β: A possible modulator of nucleotidase activity and purinergic receptor
signaling
No activity was found in the media or membranes of COS cells transfected with truncated
NTPDase3β. Co-transfection of full-length NTPDase3α with NTPDase3β decreased the
expression level and enzymatic activity of NTPDase3α detected in the COS cell membranes.
Since NTPDases are known to form active homo-oligomers (198), it is possible that the inactive,
truncated NTPDase3β could form hetero-oligomeric structures with the active, full-length
NTPDase3α isoforms, causing a decrease in the enzymatic activity of NTPDase3α, as suggested
by Mateo et al (190). However, the activity per molecule of full length NTPDase3α, as detected
by the NTPDase3α specific KLH1 antibody, did not decrease in co-transfection experiments
including NTPDase3β. In addition, Endo H deglycosylation revealed that NTPDase3β is not
delivered to the cell surface where it could form hetero-oligomers with NTPDase3α. This is in
contrast to what was found for the P2X7 receptor, which has identical membrane topology to the
cell surface NTPDases (199), and for the human serotonin receptor 5-HT3A (200), which both form hetero-oligomers containing both full length and truncated proteins. In contrast,
NTPDase3β appears to be interfering with the processing of the full-length isoform (as judged by the large decrease in expression levels of the NTPDase3α when co-transfected with
NTPDase3β). To ensure this was not a non-specific effect of the presence of another cDNA co- expressed by the same COS cells the active, A511Stop NTPDase3 mutant (missing just the non- critical C-terminal cytoplasmic tail) was co-transfected with NTPDase3α, supporting the
144 differential effect. It also seems unlikely that the effect of the co-transfection of NTPDase3β on
NTPDase3α expression is mediated via transcriptional means. Thus, we speculate that the
physiological role for the enzymatically inactive, truncated isoform of NTPDase3, NTPDase3β,
might be to decrease the ATPase and ADPase activity at the cell surface by decreasing the
synthesis and cell surface delivery of full-length, active NTPDase3α. In vivo, this would lead to
extended activation of P2 purinergic receptors by extracellular nucleotides, and increased P2 signaling in cells expressing the truncated isoform of NTPDase3, NTPDase3β.
2.4. Biochemical characterization of NTPDase3 single nucleotide polymorphisms
Other than significant decreases in the nucleotidase activity of R264Q, the NTPDase3
SNPs exhibit similar biochemical characteristics to wt NTPDase3. However, if R264Q
NTPDase3 is expressed instead of wt NTPDase3, this would result in slower hydrolysis of extracellular nucleotide agonists, leading to extended activation of P2 receptors by extracellular nucleotides, which may lead to a distinct phenotype.
3. Summary and Future Directions
This dissertation provides new insights regarding the structural elements that modulate
the enzymatic activity and mechanism of NTPDase3, as well as other membrane-bound, cell
surface NTPDases. Specifically, the findings extend our knowledge of two key structural
domains of NTPDase3: 1. the linker region, which contains a set of conserved proline residues
that mediate the intra-protein signal transduction between substrate binding in the extracellular
145 domain and movements and/or rearrangements of the TM helices; and 2. the TMD, which contains a set of conserved polar residues that provide NTPDase3 with an isoenzyme specific hydrogen bonding pattern, facilitating TM helix association, and mediating differential susceptibility to membrane perturbation. This body of work broadens our understanding of how the enzymatic activity of the membrane-bound, cell surface NTPDases is regulated by associations between (affecting the stability of the oligomeric structure of the NTPDase) and within (modulating interactions between the extracellular and TM domains) monomers to regulate nucleotide hydrolysis. These findings suggest the following experiments:
3.1. Determine if a polyproline II helix exists in the linker region of NTPDase3 and
identify the specific amino acids involved in its formation/stability
The conserved proline residues in the linker region of NTPDase3 constitute a proline-rich region that mediates interactions between the extracellular and TM domains, suggesting the possible existence of a polyproline II (PPII) helix. Typically, PPII helices are shorter than five amino acids, contain multiple proline resides, and often contain positively charged amino acids.
(201-203). These basic characteristics are consistent with the amino acid sequences of the linker regions of NTPDase3 that contain the functionally important P53 and P481 residues (N-terminal:
QEVLPP53GLK; C-terminal: QIPAESPLIRLP481IEPP). One approach to investigate this hypothesis is to make synthetic peptides of varying lengths and amino acid compositions to confirm the existence of this putative secondary structure, as well as to determine by mutagenesis the amino acids critical for formation of this secondary structure (in addition to P53
146 and P481), which may be involved in forming the hypothesized PPII helix in the N- and C-
terminal linker regions of human NTPDase3.
3.2. To further define the structural motifs that mediate oligomerization via the
transmembrane domain of the NTPDases
It is well established that the membrane-bound, cell surface NTPDases form homo-
oligomers from dimers up to tetramers (144). In this dissertation, data was presented suggesting
a Polar-X-X-Polar (Q44-X-X-T41) TM-TM motif existed in the N-terminal TM helix of
NTPDase3. There are likely other, non-polar, TM helix interactions involved in TMD
oligomerization and stability. One possibility is a putative leucine zipper motif consisting of L30
and L37 in the N-terminal TM helix of human NTPDase3. According to the helical wheel
analysis, L30 and L37 are on the same face of the TM helix as the presumed Polar-X-X-Polar
motif, possibly giving rise to the hydrophobic pocket making the Q44C mutant dimers
particularly resistant to DTT reduction. Considering these leucine residues are more conserved among the different membrane-bound NTPDases, the putative leucine zipper motif may mediate
TM helix interactions that are not specific for individual NTPDases (drive hetero-
oligomerization), while the Polar-X-X-Polar motif provides specificity for native, homo-
oligomeric NTPDase3.
In addition, the C-terminal transmembrane helix of NTPDase3 contains a set of
conserved phenylalanine residues (F487, F493, F494, F504), which may also promote non-polar
TM helix interactions, possibly by forming a “phenylalanine zipper” utilizing F487 and F494
(204-206).
147
3.3. Further biochemical characterization of the single nucleotide polymorphisms of
NTPDase3
Some SNPs are known to be disease phenotypic (207), and considering the lack of genetically modified animal models, as well as the lack of small molecule specific inhibitors of
NTPDase3, expression and further biochemical characterization of SNPs found in NTPDase3 may lead to a better understanding of the physiological function(s) of NTPDase3. Especially interesting is the SNP that is very close to the active site, R264Q. Of course, genetic studies showing a linkage between a disease phenotype and a particular SNP would be instrumental in determining the physiological function(s) of NTPDase3. Future generation and characterization of an NTPDase3 knockout mouse may indicate physiological functions of NTPDase3, and suggest phenotypes likely to occur with the lower activity R264Q NTPDase3 SNP.
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174 Appendix: Publications and Manuscripts under Review
1. Gaddie, K. J. and Kirley, T. L. Conserved Polar Residues Stabilize Transmembrane
Domains and Promote Oligomerization in Human Nucleoside Triphosphate
Diphosphohydrolase3. Biochemistry. 48(40): 9437-9447. 2009.
2. Gaddie, K. J., and Kirley, T. L. Proline Residues Link the Active Site to Transmembrane
Domain Movements in Human Nucleoside Triphosphate Diphosphohydrolase 3
(NTPDase3). Submitted to Purinergic Signalling, November, 2009.
2. P. Crawford, K. J. Gaddie, T.M. Smith and T. L. Kirley. Characterization of an alternative
splice variant of human nucleoside triphosphate diphosphohydrolase 3 (NTPDase3): A
possible modulator of nucleotidase activity in purinergic signaling. Archives of
Biochemistry and Biophysics. 475(1): 7-15. 2006.
175