Mechanisms of Angiostatin Formation by Tumour Cells
ANGELINA JAP LAY
A thesis submitted in partial fulfillment of the requirements for the degree of Doctor of Philosophy
University of New South Wales
Australia, 2001 TABLE OF CONTENTS
TABLE OF CONTENTS LIST OF FIGURES viii LIST OF TABLES xi LIST OF ABBREVIATIONS xii LIST OF PUBLICATIONS xiv STATEMENT xv ACKNOWLEDGMENTS xvi DEDICATION xvii SUMMARY xviii
CHAPTER 1 LITERATURE REVIEW 1.1 ANGIOGENESIS 1.1.1 Introduction...... 1 1 . .21 Role of endothelial cells in normal physiology ...... 4 1.1.3 Angiogenesis a cascade of events ...... 5 1.1.3.1 The extracellular matrix remodelling...... 6 1.1.3.1.1 Metalloproteinases in angiogenesis...... 6 1.1.3.1.2 Plasminogen activator (PA)-system in angiogenesis... 7 1.1.3.2 Initiation of angiogenic cascade...... 9 1.1.3.3 Endothelial cell proliferation and migration ...... 10 1.1.3.4 Maturation and stabilisation of the neovasculature...... 11 1. 1.4 Control of angiogenesis the balance hypothesis ...... 12 1.1.4.1 Angiogenic stimulators...... 14 1.1.4.1.1 Direct angiogenic inducers...... 14 1.1.4.1.2 Indirect angiogenic inducers ...... 15 . 1.1.4.2 Angiogenic inhibitors...... 16 . . . 1.1.5 Triggers of angiogenic response ...... 17 . . 1.1.5.1 Hypoxia...... 18 1.1.5.2 Mechanical injury...... 19. . . . 1.1.5.3 Inflammation ...... 19 . . . . 1.1.5.4 Genetic factors ...... 19 . . . . 1.1.6 Tumour angiogenesis ...... 20. . . . 1.1.7 Antiangiogenic therapy...... 22 1.2 PLASMINOGEN/PLASMIN SYSTEM 1.2.1 Introduction...... 24 1.2.2 Structure ...... 24 ...... 1.2.3 Kring le domains of plasminogen ...... 27. . . 1.2.4 Variants of plasminogen...... 28 1.2.4.1 Glu-plasminogen ...... 28 . . . . 1.2.4.2 Lys-plasminogen ...... 28 . . . . 1.2.5 Plasminogen activation...... 30. . . . 1.2.5.1 Plasminogen activators ...... 31 . . . 1.2.5.1.1 Tissue-type plasminogen activator (tPA) ...... 32 1.2.5.1.2 Urokinase plasminogen activator (uPA) ...... 33 1.2.5.1.3 Urokinase plasminogen activator receptor (uPAR) . . . . 35 1.2.5.2 Plasminogen activator inhibitors (PAI) ...... 36 . 1.2.5.2.1 Plasminogen activator inhibitor 1 (PAl1)...... 36 1.2.5.2.2 Plasminogen activator inhibitor 2 (PAl2)...... 37 1.2.5.2.3 Protease-nexin 1 (PN-1) ...... 38 . . 1.2.5.2.4 Alpha 2-antiplasmin ...... 38 . . . 1.2.5.3 Plasminogen receptors...... 40
1.3 ANGIOSTATIN 1.3.1 Introduction...... 42 1.3.2 Structure...... 43 1.3.3 Angiostatin converting activity of various enzymes...... 43 1.3.4 Function...... 45 ...... 1.3.5 Inhibitory activity of kringle domains of angiostatin ...... 46 1.3.6 Role of kringle 5 plasminogen...... 48 1.3.7 Mechanism of action ...... 48. . . . .
1.4 PHOSPHOGL YCERATE KINASE (PGK) 1.4.1 Introduction...... 51 1.4.2 Role of PGK in glycolysis ...... 51. . . . 1.4.3 Structure of PGK...... 53 . . . . . 1.4.3.1 The 3-Dimensional structure of PGK ...... 54 . 1.4.4 Domain movement in PGK...... 56. . . .
ii 1.4.5 PGK isoenzyme ...... 57. . . . 1.4.6 Phosphoglycerate kinase in diseases...... 58.
CHAPTER 2 HYPOTHESIS AND AIMS 2.1 HYPOTHESISANDAIMS ...... 61
CHAPTER 3-CHARACTERISATION OF PLASMIN REDUCTASE 3.1 INTRODUCTION...... 59 . . . .
3.2 MATERIALS AND METHODS...... 62 3.2.1 Cell culture ...... 62 . . . . 3.2.1.1 HT1080 cell culture ...... 62 3.2.1.2 Breast carcinoma cell culture ...... 63. . 3.2.1.3 Preparation of conditioned media (CM) ...... 63 3.2.2 Assay for plasmin reductase activity ...... 63 . 3.2.2.1 Angiostatin generation from HT1080 CM ...... 64 3.2.2.2 MPB blot assay...... 65 3.2.2.3 Microtitre plate assay ...... 65. . . 3.2.3 Endogenous glutathione in HT1080CM ...... 66. 3.2.4 Competitive inhibition of plasmin reductase activity by NADH . . . 67 3.2.4.1 Binding of plasmin reductase to Cibachron Blue-Sepharose 67 3.2.5 Alkylation of thiol groups in plasmin reductase...... 67 3.2.5.1 Binding of plasmin reductase to thiol activated sepharose.. 68
3.3 RESULTS ...... _...... 69 . 3.3.1 Elisa for determining plasmin reductase activity ...... 69 3.3.2 Streptavidin-HRP blot for MPS-labelled angiostatin ...... 71 3.3.3 Plasmin reductase activity secreted by various cultured cells . . . 72 3.3.4 Effect of NADH on plasmin reductase activity ...... 73 3.3.4.1 Plasmin reductase bound to Cibachron Blue-Sepharose... 74 3.3.5 Thiol modifying reagents reduced plasmin reductase activity . . . 75 3.3.6 Plasmin reductase bound to thiol activated sepharose ...... 76
3.4 DISCUSSION...... 77. . . . .
iii CHAPTER 4 PLASMIN REDUCTASE PURIFICATION 4.1 INTRODUCTION...... 80. . . . .
4.2 MATERIALS AND METHODS...... 81 4.2.1 Large scale production of CM...... 81 4.2.1.1 Cell factory...... 81 4.2.2 Quantitative analysis of protein...... 81 4.2.3 ELISA and Streptavidin-HRP blot for plasmin reductase activity. 82 4.2.4 Concentration of HT1080 CM...... 82 4.2.5 8Iue-Sepharose CL-68 column chromatography ...... 82 4.2.6 Thiopropyl-Sepharose 68 chromatography...... 83. 4.2. 7 Gel filtration (Sephacryl S-200 HR) ...... 84. . 4.2.8 NHi-terminal sequence analysis ...... 84. .
4.3 RESULTS ...... 86...... 4.3.1 Cibachron 8Iue-Sepharose 48 chromatography ...... 86 4.3.2 Thiopropyl-Sepharose 68 chromatography...... 88. 4.3.3 Sephacryl S-200 HR gel filtration...... 89 . . 4.3.4 Protein microsequencing ...... 91 . . . 4.3.4.1 Amino acid sequence of human PGK...... 92. 4.3.5 Stages of plasmin reductase purification ...... 93 .
4.4 DISCUSSION...... 94 . . . . .
CHAPTER 5 CLONING AND EXPRESSION OF RECOMBINANT HUMAN PGK (rhPGK) 5.1 INTRODUCTION...... 96 . . . . .
5.2 MATERIALS AND METHODS...... 97 . . . 5.2.1 Enzymes and reagents ...... : ...... 97 5.2.2 Cellular mRNA extraction...... 97 . . . 5.2.3 Construction of human PGK cDNA ...... 98. . 5.2.3.1 PCR reaction...... 98 . . . .
iv 5.2.3.2 cDNA and plasmid vector digestion ...... 99 5.2.3.2.1 cDNA vector digestion ...... 99 5.2.3.2.2 Plasmid vector digestion ...... 99 5.2.3.3 Ligation reaction...... 99 5.2.3.4 Transformation ...... 100 5.2.4 Expression of rhPGK...... 100 5.2.4.1 Analysis of rhPGK expression ...... 100 5.2.4.2 Large scale production of rhPGK...... 101 5.2.5 Purification of rhPGK...... 101 5.2.5.1 Ammonium sulfate precipitation...... 101 5.2.5.2 Cibachron Blue-Sepharose ...... 102 5.2.5.3 S-Sepharose chromatography ...... 102 5.2.6 Mass spectrometry...... 102 5.2.7 ELISA for reductase activity of rhPGK ...... 103
5.3 RESULTS ...... 102 5.3.1 cDNA sequences of rhPGK...... 102 5.3.2 rhPGK purification ...... 103 5.3.3 Mass spectrometry of recombinant human PGK...... 104 5.3.4 Plasmin reductase activity of rhPGK...... 105 5. 3.5 Comparison of plasmin reductase activity of various PGK ...... 106
5.4 DISCUSSION...... 107
CHAPTER 6 IN VIVO STUDY OF rhPGK 6.1 INTRODUCTION...... 109
6.2 MATERIALS AND METHODS ...... 110 6.2.1 Cell culture ...... 110 6.2.2 Coupled kinetic assay to measure PGK secretion ...... 110 6.2.3 Tumour implantation ...... 112 6.2.4 Tumourigenesis model ...... 113 6.2.5 lmmunohistochemical staining for angiogenesis...... 114
V 6.3 RESULTS ...... 115 6.3.1 Assay used to determine PGK secretion ...... 115 . 6.3.2 PGK is secreted by cultured carcinoma cells ...... 116 6.3.3 Plasmin reductase activity of various cultured CM ...... 117 6.3.4 Plasma levels of PGK in tumour-bearing mice ...... 118 6.3.5 The effect of systemic administration of rhPGK on plasma levels of MPB-angiostatin ...... 119 6.3.6 Inhibition of HT1080 tumour growth by rhPGK ...... 120 6.3.7 Inhibition of AsPC-1 tumour growth by rhPGK...... 121 6.3.8 Effect of rhPGK on tumour angiogenesis...... 123 .
6.4 DISCUSSION...... 124. . . . .
CHAPTER 7 MECHANISM OF PLASMIN REDUCTION BY PGK 7.1 INTRODUCTION ...... 127
7.2 MATERIALS AND METHODS ...... 129 7.2.1 Chemicals and proteins ...... 129 7.2.2 Production of mutant rhPGK ...... 129 7 .2.2.1 PCR reaction...... 130 . . . . 7 .2.2.2 Dpn I digestion ...... 130. . . . 7.2.2.3 Transformation of super competent cells (XL-B) ...... 130 7.2.2.4 Transformation of BL21 (DE3) cells ...... 131 7.2.3 Purification of mutant rhPGK ...... 133 7 .2.4 Quantitative analysis of free thiols in PGK...... 133 . 7 .2.5 Alkylation of rhPGK ...... 134. . . . 7 .2.6 Plasmin reduction at various conditions...... 134 . 7.2.6.1 pH ...... 134 7 .2.6.2 Ionic strength...... 134 . . . 7.2.6.3 Temperature ...... 135 7 .2.6.4 3-PG/ATP-MgCl2 ...... 135. . . . 7.2.7 Assay for plasmin reduction ...... 135
vi 7.3 RESULTS ...... 137 7.3.1 Sequence confirmation Cys mutations in PGK ...... 137 7.3.2 Purification of PGK mutants ...... 140 7.3.3 Consequence of mutation of all seven PGK Cys residues to Ala for plasmin reductase activity ...... 141 7.3.4 Consequence of mutation of PGK Cys residues on the susceptibility of PGK to proteolysis ...... 143 7.3.5 Quantitative analysis of Cys379,380 in PGK ...... 145 7.3.6 Effect of alkylation of the fast-reacting Cys of PGK on plasmin reductase activity...... 146 7.3.7 Effect of pH, ionic strength and temperature on the plasmin reductase activity of PGK ...... 149 7.3.8 Effect of 3-PG and ATP-induced conformational change in PGK on the plasmin reductase activity ...... 151
7.4 DISCUSSION...... 153
CHAPTER 8 GENERAL DISCUSSIONS AND CONCLUSIONS...... 159
REFERENCES ...... 169
vii LIST OF FIGURES
Fig. Caption Page
1.1 Schematic representation of the angiogenic process...... 3 1.2 Scheme of the tertiary structure of human plasminogen ...... 26 1.3 Plasminogen activation cascade...... 41 1.4 Mechanism of PGK reaction in glycolysis ...... 52 1.5 Complete amino acid sequence of human PGK...... 53 1.6 Three dimensional structure of human PGK...... 55
3.1 ELISA for MPS-labelled angiostatin fragments...... 70 3.2 MPS-labelled angiostatin fragments blotted with Streptavidin-HRP. . 71 3.3 Plasmin reductase activity secreted by selected primary and transformed cells...... 72 3.4 NAOH competitively inhibited plasmin reductase activity ...... 73 3.5 MPS-labelled angiostatin fragments after Cibachron Blue affinity chromatography ...... 74 3.6 Effect of thiol-modifying reagents on plasmin reductase activity. . . . . 75 3. 7 Plasmin reductase activity after incubation with Thiopropyl-Sepharose ...... 76
4.1 Purification of plasmin reductase on Cibachron Blue-Sepharose Chromatography...... 87 4.2 Plasmin reductase activity eluted from Thiopropyl-Sepharose 4B . . . 88 4.3 Purification of plasmin reductase on gel filtration...... 89 4.4 SOS-PAGE of gel filtrated fractions...... 90 4.5 Sequencing profile for NH2-terminal peptides of plasmin reductase.. 91 4.6 Sequence alignment of human PGK and NH2-terminal peptides of plasmin reductase...... 92 4.7 SOS-PAGE of significant stages of plasmin reductase purification ... 93
5.1 Coding cONA and derived sequences of rhPGK ...... 104 5.2 SOS-PAGE of HT1080 PGK and rhPGK ...... 105 5.3 Purified rhPGK has a molecular mass of m/z 44493 as determined by mass spectrometric analysis ...... 106 5.4 rhPGK has the same plasmin reductase activity as HT1080 PGK ... 107
viii Fig. Caption Page
5.5 Plasmin reductase activity of PGK from various origins ...... 108
6.1 Coupled kinetic assay to measure PGK secretion ...... 113 6.2 Protocol for HT1080 tumour implantation on SCID mice ...... 114 6.3 Model for tumourigenesis ...... 115 6.4 PGK coupled kinetic assay standard curve ...... 117 6.5 Rate of PGK secretion by various transformed cells ...... 118 6.6 Plasmin reductase activity in CM of various cultured transformed cells ...... 119 6. 7 Plasma levels of PGK in mice bearing HT1080 tumours compared to mice without tumours ...... 120 6.8 Plasma levels of angiostatin before and after treatment with rhPGK. 121 6.9 Inhibition of HT1080 tumour growth by rhPGK ...... 122 6.10 Inhibition of AsPC-1 tumour growth by rhPGK ...... 123 6.11 Reduction of AsPC-1 tumour size by rhPGK...... 124 6.12 Microvessels in control and rhPGK-treated AsPC-1 tumours from SCID mice ...... 125
7 .1 Three dimensional structure of human PGK showing the organisation of the N-and C- terminal domains ...... 131 7 .2 Sequence alignment of human PGK DNA against recombinant PGK mutants ...... 140 7.3 SOS-PAGE profile of wt and mutant PGK's ...... 143 7.4 Consequence of all seven PGK Cys mutation on plasmin reductase activity ...... 145 7.5 Consequence of mutation of PGK Cys residues on the susceptibility of PGK to proteolysis ...... 146 7.6 Titration of the fast-reacting thiols of wild-type or C379,380A PGK .. 147 7.7 Alkylation of the fast-reacting thiols in rhPGK ...... 149 7.8 Effect of alkylation of the fast-reacting Cys of PGK on plasmin reductase activity ...... 150 7.9 Effect of pH, ionic strength and temperature on the plasmin reductase activity of PGK ...... 152 7 .1 O Effect of 3-PG and ATP-induced conformational change in PGK on the plasmin reductase activity ...... 154
ix 8.1 Model of the molecular events facilitated by PGK in plasmin kringle 5 ...... 167
X LIST OF TABLES
No. Caption Page
1.1. Angiogenic activators and inhibitors ...... 13 1.2. Angiogenic Molecules acting directly on endothelial cells ...... 15
4.1. Summary of different steps in the purification of plasmin reductase .. 93
7 .1 Primers for PGK mutants (base changes are underlined) ...... 135
xi LIST OF ABBREVIATIONS
ABTS 2,2'-azino-bis(3-ethylbenzthiazoline-6-sulfonic acid) ARNT aryl hydrocarbon receptor nuclear translocator ATP Adenosine triphosphate BAEC Bovine aortic endothelial cell bBBr Dibromobimane BSA Bovine serum albumin cDNA Complementary DNA CM Conditioned media CNBr Cyanogen Bromide DMEM Dulbecco's modified Eagle medium DNA Deoxinucleic acid DTNB 5,5'-dithiobis(2-nitrobenzoic acid) OTT Dithiothreitol EDTA Ethylene-diamine-tetra-acetic acid ELISA Enzyme linked immunosorbent assay EA.CA Epsilon Amino-n-caproic acid FCS Fetal calf serum GAPDH Glyceraldehyde-3-phosphate Dehydrogenase GSH Reduced Glutathione GSSG oxidised glutathione HBSS Hanks Balanced Salt Solution HDMVEC Human dermal microvascular endothelial cell HIF-1 Hypoxia-inducible factor 1 HPLC High performance liquid chromatography HRE Hypoxia-response Element HT1080 Human fibrosarcoma cell line HUVEC Human umbilical vein endothelial cell 1AM lodoacetamide lgG lmmunoglobulin G LOH Lactate dehydrogenase mAb Monoclonal antibody MPB 3-(N-maleimidylpropionyl) biocytin mRNA messenger RNA
xii NEM N-ethylmaleimide PAl1 Plasminogen activator inhibitor type-1 PAl2 Plasminogen activator inhibitor type-2 PBS Phosphate buffered saline PBS/Tween Phosphate buffer saline/Tween-20 PGK ATP:3-phospho-D-glycerate1-phosphotransferase Pig Plasminogen Pm Plasmin PMSF Phenylmethylsulfonyl fluoride PN-1 Protease-nexin 1 PVDF Polyvinylidene difluoride rhPGK Recombinant human phosphoglycerate kinase RNA Deoxyribonucleic acid RT-PCR Reverse transcriptase polymerase chain reaction sctPA single-chain tPA scuPA single-chain uPA SOS-PAGE Sodium dodecyl sulfate-polyacrylamide gel electrophoresis StreptABC/HRP Streptavidin and biotinylated horseradish peroxidase tctPA two-chain tPA tcuPA two-chain uPA TFA Trifluoro acetic acid tPA Tissue type plasminogen activator TT Tetrathionate Tween-20 Polyoxyethylene-sorbitan monolaurate uPA Urokinase plasminogen activator
xiii LIST OF PUBLICATIONS
Lav, A.J., Jiang, X.-M., Kisker, 0., Flynn, E., Underwood, A., Condron, R., and Hogg, P.J. Phosphoglycerate kinase acts in tumour angiogenesis as a disulphide reductase. Nature, 408, 869-873 (2000).
Lav A.J., and Hogg PJ. Measurement of reduction of disulfide bonds in plasmin by phosphoglycerate kinase. Methods in Enzymology, 348, 87-92 (2002).
Lav, A.J., Jiang, X.-M., Daly, E.B., Sun, L., and Hogg P.J. Reduction of plasmin by phosphoglycerate kinase is a thiol independent process Journal of Biological Chemistry, 277, Issue 11, 9062-9068 (2002).
Stathakis, P., Lav, A.J., Fitzgerald, M., Schlieker, C., Matthias, L.J., and Hogg, P.J. Angiostatin formation involves disulphide bond reduction and proteolysis in kringle 5 of plasmin. Journal of Biological Chemistry, 274, Issue 13, 8910-8916 (1999).
xiv STATEMENT
I hereby declare that this submission is my own work and to the best of my knowledge it contains no materials previously published or written by another person, nor material which to a substantial extent has been accepted for the award of any other degree or diploma at UNSW or any other educational institute, except where due acknowledgment is made in the thesis. Any contribution made to the research by colleagues, with whom I have worked at UNSW or elsewhere, during my candidature, is fully acknowledged.
I also declare that the intellectual content of this thesis is the product of my own work, except to the extent that assistance from others in the project's design and conception or in style, presentation and linguistic expression is acknowledged.
Angeli ap Lay December 2001
xv ACKNOWLEDGMENTS
First of all I would like to acknowledge the guidance, support and encouragement of my mentor and supervisor, Professor Philip Hogg. Thank you Phil for giving me the opportunity to be part of the team. It has been a very challenging yet enjoyable journey of science discovery. Thank you for being a great boss. To my co-supervisor, Professor Colin Chesterman. Thank you Prof. for all your help and support throughout the year. I would like to thank Dr. Xing Mai Jiang for his guidance and assistance in all molecular biology work. I would like to also acknowledge Dr. Kathy Quinn and Dr. Mark Raftery for their valuable advice and tips on protein purification. Thank you for your support and friendship.
I am honoured to have been part of the Centre for Thrombosis and Vascular Research, not only because of the scientific achievements but more importantly the many wonderful friends I met here in CTVR. To my past and present lab buddies, Peter Newman, Paul Stathakis, Neil Donoghue, Belinda Creighton, Mary Kavurma, Elise Daly, and everyone in CTVR, thank you guys, for making my years in CTVR enjoyable. To Brian and Helen, thank you for those refreshing morning jogs. To Julie and my bay-can buddy Richie, thank you for your support and friendship.
A very special thank you to my loving parents for their endless support, encouragement, love and patience waiting for me to finish my thesis. To Yolanda, Eugene, Frank, Angela, Gary, Reynald and Nancy, thank you so much for always being there. Finally, to James "Chuggy", thank you for standing by me through highs and lows and thank you for being so wonderful. I wouldn't have made it this far without you all.
xvi To Mum and Dad
xvii Summary
Disulfide bonds in secreted proteins are considered to be inert because of the oxidising nature of the extracellular milieu. The finding that carcinoma cells secrete a reductase that reduces disulfide bonds in the serine proteinase plasmin is an exception to this rule. Reduction of plasmin initiates proteolytic cleavage in the kringle 5 domain and release of the angiogenic inhibitor, angiostatin. Plasmin contains five consecutive kringle domains followed by a serine proteinase module and is processed in the conditioned medium of carcinoma cells producing angiostatin fragments consisting of kringle domains 1 to 4 and parts of kringle 5. Plasmin proteolysis occurs in three stages. Firstly, the Cys461-Cys540 and Cys511- Cys535 disulfide bonds in kringle 5 of plasmin are reduced by a plasmin reductase. Secondly, reduction of the kringle 5 disulfide bonds trigger cleavage at Arg529-Lys530 in kringle 5, and also at two other positions C terminal of Cys461, by a serine proteinase releasing kringle 1-4½. Third, matrix metalloproteinase-dependent trimming of kringle 1-4½ to either kringle 1-4 or 1-3.
The aims of this thesis were to characterise plasmin reductase and investigate the mechanism of angiostatin formation by tumour cells. The plasmin reductase was isolated from fibrosarcoma cell conditioned medium and shown to be the glycolytic enzyme, phosphoglycerate kinase (PGK). Recombinant PGK had the same specific activity as the fibrosarcoma derived protein. Plasma of mice bearing fibrosarcoma tumours contained several fold more PGK than mice without tumours and administration of PGK to tumour-bearing mice caused an increase in plasma levels of angiostatin and decrease in tumour vascularity and rate of tumour growth. These findings indicate that PGK not only functions in glycolysis but also is secreted by
xviii tumour cells and participates in the angiogenic process as a disulfide reductase.
Furthermore, the mechanism of plasmin reduction by PGK was investigated. The chemistry of protein reductants is typically mediated by a pair of closely spaced Cys residues. There are 7 Cys in human PGK and only 2 of the 7 are nearby in the primary or tertiary structure. Cys379 and
Cys380 are close to the hinge region that links the N- and C-terminal domains of PGK. These cysteines have been referred to as 'fast-reacting' as they are amenable to alkylation by several thiol-reactive compounds. The role of all 7 Cys and in particular the two fast reacting Cys, in reduction of plasmin disulfide bonds by PGK has been explored in this study. Mutation of all 7 to Ala did not appreciably affect plasmin reductase activity.
Interestingly, some of the mutations perturbed the tertiary structure of the protein. Alkylation/oxidation of Cys379 and Cys380 by four different thiol reactive compounds reduced plasmin reductase activity to 7-35% of control.
Binding of 3-phosphoglycerate and/or MgATP to the N- and C-terminal domains of PGK, respectively, triggers a hinge-bending conformational change in the enzyme. Incubation of PGK with 3-phosphoglycerate and/or
MgATP ablated plasmin reductase activity, with half-maximal inhibitory effects at -1 mM concentration.
In summary, this study shows that none of the Cys residues in PGK are directly involved in plasmin reduction but alkylation/oxidation of the fast reacting Cys or conformational changes in PGK perturbs plasmin reduction, perhaps by obstructing the interaction of plasmin with PGK or perturbing conformational changes in PGK required for plasmin reduction.
xix CHAPTER 1 LITERATURE REVIEW Chapter 1 - Literature Review
1.1 ANGIOGENESIS
1.1.1 Introduction
There are two main mechanisms by which new blood vessels are formed. The first process known as vasculogenesis involves in situ differentiation of endothelial cells (ECs) from mesodermal precursors and their subsequent organisation into new capillaries. The second process is angiogenesis. This is characterised by the formation of new capillaries from pre-existing vessels. Vasculogenesis occurs primarily in early embryogenesis and does not seem to contribute to repair and disease in postembryonic life. Angiogenesis, on the other hand, occurs during development and throughout postnatal life.
Angiogenesis is fundamental to a large number of biological processes. In adult mammals the vasculature remains quiescent except for transient neovascularisation in the female reproductive system, where it is required for implantation and early embryonic development. In addition to its role in reproduction, angiogenesis is required for the maintenance of functional and structural integrity of the organism during its postnatal life, such as for tissue remodelling, in inflammation and in ischemic conditions.
Angiogenesis in such situations is tightly controlled and limited by the metabolic demands of the tissue concerned [1].
Angiogenesis has been implicated in the development and progression of many pathological processes. Either inadequate or excessive angiogenesis can lead to pathological outcomes. For instance, lack of angiogenesis can lead to impaired healing of wounds, bone and chronic ulcers. In contrast, uncontrolled angiogenesis can promote tissue
1 Chapter 1 - Literature Review
destruction, such as hyperproliferation of blood vessels in the eyes of diabetic patients. In rheumatoid arthritis, for example, excessive angiogenesis contributes to extensive proteolytic activity, resulting in the accumulation and proliferation of inflammatory synovial tissue. This leads to the destruction of the joint surface. In childhood hemangioma, excessive angiogenesis is thought to be the central pathogenetic mechanism underlying this childhood condition. In cancer the growth and development of tumour cells depends on adequate angiogenesis for oxygen and nutrient supply.
Once a tumour has established its own blood vessels, tumour cells can escape and disseminate to other parts of the body. Thus, angiogenesis promotes tumour growth and metastasis. Angiogenesis is now widely recognised as a powerful control point in tumour growth.
The many functions of angiogenesis in different pathological conditions mean understanding the precise mechanism of its regulation is critical. Identifying the key players or factors, which trigger angiogenesis and most importantly understanding how to interrupt this process will not only guide diagnosis but also treatment for angiogenic related diseases.
Research into the significance of angiogenesis to the progression of tumours has been a most focused area of investigation in recent years. The idea of targeting angiogenesis as a regime for cancer therapy has stimulated a tremendous search for potential anti-angiogenic factors. As a result, angiogenic inhibitors have now emerged as a new class of drugs.
The complexity of the angiogenic process, involving both pro- and anti-angiogenic molecules, provides various approaches to target angiogenesis at many different levels. A large spectrum of strategies for
2 Chapter 1 - Literature Review
modulation of angiogenesis has been described. These approaches focus on the functions of ECs during blood vessel formation, such as intervention with EC growth, adhesion and migration. The methods for such intervention involved either the use of inhibitors of angiogenic factors or factors with direct anti-angiogenic effect. The former include inhibitors of vascular endothelial growth factor (VEGF), platelet-derived growth factor (PDGF), receptor tyrosine kinases (Flk-1/KDR, EGFR and VEGFR, PDGFR, FGFR RTK), PD
ECGF/TP, matrix metalloproteinases (MMPs), proteases and the integrin family. The direct anti-angiogenic factors include platelet factor 4 (PF4), thrombospondin, angiostatin and endostatin .
•
1) EC activation • 3) EC proliferation : • 2) BM degradation • • • • • • • 4) ECM proteolysis 5) EC migration 1 I I 6) BM synthesis 7) ECM remodelling I I ' 8) Vascular loop formation J}
Angiogenic stimuli
Figure 1.1. Schematic representation of the angiogenic process. The activation of EC in response to an angiogenic stimulus is followed by BM and ECM degradation. ECs then proliferate and migrate into the surrounding matrix and form into new capillary sprouts. Sprout maturation occurs by the synthesis of new BM followed by formation of a functional capillary loop.
3 Chapter 1 - Literature Review
1.1.2 Role of endothelial cells in normal physiology
Blood vessels have a highly ordered and nonthrombogenic monolayer of ECs which line the intimal surface of the vessel walls. ECs are in direct contact with the blood and subendothelially located pericytes, smooth muscle cells (SMC), fibroblasts, basement membrane (BM), and extracellular matrix
(ECM) [2]. Together, these cellular components maintain the structure and stability of blood vessels.
In healthy vessels, ECs are non-migrating cells and actively involved in a number of regulatory processes. They control blood coagulation by regulating the production of thrombomodulin, tissue factor pathway inhibitor and tissue type plasminogen activator. In addition to being metabolically active, they serve as a permeable barrier for small solutes and peptides.
The vascular system is generally stable and ECs are thought to be quiescent, however they can be rapidly activated in response to tissue damage. Upon stimulation, ECs secrete von Willebrand factor (vWF), which subsequently stimulates platelet adhesion and aggregation and plasminogen activator inhibitor 1 (PAl-1). Studi_es have also demonstrated that ECs are involved in directing cells of the immune system to specific sites in the body.
Furthermore, ECs participate in vascular remodelling during ovulation, wound healing, tumour growth and diabetic retinopathy.
In the normal physiological state, ECs are quiescent with one in every
10,000 cells undergoing cell division at any given time [3]. The turnover time for ECs can exceed 1000 days [4]. By contrast, they can be activated to migrate and replicate in response to appropriate stimuli, such as during angiogenesis, giving rise to new capillaries, with a mean turnover of at least
4 Chapter 1 - Literature Review
five days. This process ceases soon after the tissue demands have been fulfilled by the new vasculature.
1.1.3 Angiogenesis - a cascade of events
Angiogenesis is a complex cellular process that requires the functional activity of many protein components of the ECM, including growth factors/cytokines, adhesion molecules, and proteolytic enzymes. During the process of capillary sprouting the combined actions of these proteins leads to matrix remodelling and EC reorganisation.
Angiogenesis occurs in the capillary bed and post capillary venules.
Upon stimulation, ECs release proteases that degrade the basement membrane and the surrounding structural components [5]. Subsequently, these ECs invade the surrounding matrix (EC migration). They then undergo proliferation, differentiation and further migration to form new capillary sprouts. Thus, angiogenesis is characterised by alterations in at least three
EC functions, first, the modulation of the EC-ECM interaction, which requires alterations of cell-matrix contacts and the production of matrix degrading enzymes; second, change in EC locomotion, which allows them to migrate towards angiogenic stimuli and terminate once their destination is reached, and third, an increase in proliferation to recruit new cells for the growing vessel and subsequent return to the quiescent state once the vessel is established [6]. The formation of BM, which signals the onset of vessel maturation, involves recruitment of pericytes.
5 Chapter 1 - Literature Review
1.1.3.1 Extracellular matrix remodelling
The ECM is composed of proteinaceous fibres and other macromolecules that profoundly influence cellular function and tissue architecture [1]. The ECM influences cell behaviour in a number of ways, such as by mobilising growth factors, modulating cellular responses to these growth factors and signalling via receptors [7].
In the initial step of capillary sprouting, the components of the ECM must first be degraded to clear the path for EC invasion into the stroma. This degradation process is spatially and temporally restricted to ensure adequate proteolysis and to prevent excessive matrix degradation. Proteases responsible for this process are the metalloproteinases and components of the plasminogen activator (PA)-plasmin system. These protease systems can act independently or in cooperation to promote optimal degradation of the ECM. In addition to their proteolytic activity, they can also influence the regulation of cytokine activity in the ECM environment.
1.1.3.1.1 Metalloproteinases in angiogenesis
The role of MMPs and their endogenous inhibitors (TIMPs) in the control of angiogenesis has long been recognised [8, 91). MMPs are members of a multigene family of zinc-dependent enzymes. They can be categorised into four subgroups based on their structural characteristics and substrate specificities. The four subgroups are:
6 Chapter 1 - Literature Review
(i) Interstitial collagenases (MMP-1, MMP-8 and collagenase-3)
(ii) Gelatinases (MMP-2 and MMP-9)
(iii) Stromelysins (MMP-3, MMP-10, MMP-7 and stromelysin-3)
(iv) Membrane type (MT)- MMPs
These enzymes can degrade most protein components of the ECM. These include many different types of collagens, gelatin, laminin, fibronectin, elastin, and the protein core of proteoglycans and entactin [1 O].
MMPs are calcium-dependent endoproteases, which are synthesised as inactive proenzymes and activated via cleavage of a propeptide by proteases such as plasmin and by MMPs themselves. Thus, the activities of
MMPs are controlled at three levels, gene transcription, proenzyme activation and inhibition by specific tissue inhibitors [1 O].
The EC expression of MMPs can be up regulated by a number of factors, such as lnterleukin-1 (IL-1), VEGF, EGF, bFGF, PDGF, TGFa and
TNFa. In addition, oncogenes such as ras and src have been shown to elevate the expression of MMPs at the transcriptional level in various cell types.
1.1.3.1.2 Plasminogen activator (PA)-system in angiogenesis
The PA system is one of the most studied proteolytic enzymes in the
context of angiogenesis. As in the case of MMPs, the activities of PAs are
also controlled at three levels, gene transcription, proenzyme activation and
inhibition by specific tissue inhibitors. Again, similar to the MMPs, a number
of cytokines and growth factors with angiogenic activity influence EC
expression of plasminogen activators and inhibitors both in vitro and in vivo.
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Activated ECs express urokinase plasminogen activator (uPA), uPA
receptor (uPAR) and plasminogen activator inhibitors (PAl-1 and PAl-2).
uPA and tissue type plasminogen activator (tPA) are the principal activators
of plasminogen and are secreted as inactive proenzymes. Binding of the
secreted pro-uPA to uPAR on the cell surface activates uPA. Plasminogen
and plasmin also bind to plasma membranes and co-localisation of uPA and
plasminogen on the cell surface increases the efficiency of plasminogen
activation. This cell surface localisation of plasmin activity functions not only to concentrate proteolysis near the cell surface but also to restrict its activity to the immediate pericellular environment [6]. Binding of plasmin to the cell
surface membrane also protects it from rapid inactivation by inhibitors, in
particular ar antiplasmin.
The concentration of plasminogen in blood is 1.5 µM. Thus, small
amounts of plasminogen activators can lead to high levels of plasmin.
Plasmin is a broad acting protease of tryptic specificity, degrading most
components of the stroma such as fibrin, fibronectin, proteoglycans, gelatins
and laminin.
Besides degrading most components of the ECM, plasmin also
activates several MMPs as well as latent elastase [1 O]. In the context of
angiogenesis, this increase in proteolytic activity results in EC degradation
and invasion of the BM. This increase in proteolytic activity also results in
the generation of ECM products that are chemotactic for ECs and activation
and mobilisation of growth factors from the ECM. These proteolytic events
facilitate EC's migration and invasion during angiogenesis.
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1.1.3.2 Initiation of the angiogenic cascade
Cytokines are released from various cells of the vascular system in response to hypoxia or ischemic conditions. Once released, they cause EC activation. This is thought to be the first event in angiogenesis. Thus, hypoxia or ischemic conditions are thought to be responsible for the initiation of the angiogenic cascade.
VEGF is a potent multifunctional cytokine [11, 12] which is secreted by hypoxic tumour cells and cells of the immune system [13]. It is believed to play a role in the early stage of angiogenic activation. This finding was supported by the study of Forsythe and co-workers, which reported the up regulation of VEGF receptor (VEGFR) expression under hypoxia or ischemic conditions [14]. VEGF induces vessel dilatation and increases vascular permeability allowing the diffusion of plasma proteins into the underlying tissue [15]. In addition, VEGF can stimulate the expression of proteases and receptors [16] important in cellular invasion and tissue remodelling as well as contributing to the maintenance of new vasculature [17].
Following activation, ECs locally degrade the basement membrane and disrupt the ECM. Angiopoietin 2 (Ang2) and MMPs, which are produced by epithelial cells, fibroblasts, inflammatory cells and ECs mediate this process. These proteolytic events clear the passage for migrating ECs to invade and reorganise into the surrounding interstitial matrix to form a capillary tube.
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1.1.3.3 Endothelial cell proliferation and migration
In the newly established environment, ECs begin to proliferate and recruit more ECs for the growing vessels. Migration also occurs allowing
ECs to align with one another to form a new capillary tube. VEGF, angiopoietin 1 (Ang1) and bFGF mediate EC proliferation and migration.
Members of the integrin family of cell adhesion proteins are also involved in cell migration.
The integrins are a family of transmembrane glycoproteins composed of a and B subunits. There are more than 15 a and 8 B subunits that can heterodimerise in over 20 combinations [18]. These heterodimeric complexes serve as cell membrane receptors that form focal adhesion contacts with various ECM ligands. These include fibronectin, laminin, vitronectin, the collagens thrombospondin, entactin, fibrinogen, intercellular adhesion molecule (ICAM) and the vascular cell adhesion molecule (VCAM)
[19, 20]. lntegrin-mediated adhesion leads to intracellular signalling events that regulate cell survival, proliferation and migration [21].
Members of the integrins family are expressed on a wide variety of cells and ·tissues including ECs. Of the various integrins expressed on the cell surface, integrin avB3 is particularly important during angiogenesis. avB3 is only minimally expressed on quiescent ECs but is significantly up-regulated on EC activated by bFGF, VEGF and TGFB 1 [22]. Disruption of avB3 ligation with antibody directed to integrin avB3 disrupts blood vessel formation in quail embryo [23] and rabbit corneal model [24]. Taken together these studies strongly suggest a functional role for integrin avB3 in angiogenesis.
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lntegrin avlh is a receptor for various ECM ligands with an exposed Arg
Gly-Asp (RGD) sequence, such as fibronectin, vitronectin, fibrinogen, thrombospondin, proteolysed collagen, vWF and osteopontin [25]. A recent study showed that it could bind MMP2 in a non-RGD-dependent manner.
Engagement of av~3 and MMP2 serves to localise the proteolytic activity of the enzyme on the surface of angiogenic blood vessels [26]. This enables angiogenic ECs to degrade and remodel the ECM during their invasion.
1.1.3.4 Maturation and stabilisation of the neovasculature
Further migration of ECs near the tip of the sprout increases the length of the new vessel. Subsequently this forms a lumen. These microvascular tubules join with one another and develop into a functioning circulatory network in which blood flow is soon established. Vessel maturation is characterised by the formation of new BM, termination of EC proliferation and maturation of EC [1] as well as investment of new vessels with pericytes and smooth muscle cells. Studies have suggested that TGF~ and Ang1 appear to be the first members of ligands that regulate vessel maturation [27, 28].
The newly formed vessel then undergoes stabilisation. The interaction of ECs with the ECM and mesenchymal cells is critical for the establishment of a structurally stable mature vasculature. The nascent endothelial tube structure consists of precursor cells of pericytes and smooth muscle cells.
Recruitment of these cells and their subsequent differentiation as well as the deposition of ECM proteins is fundamental to vessel stability. ECs can accomplish this via the synthesis and secretion of PDGF-88, a mitogen and chemoattractant for a number of mesenchymal cells [29].
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Angiopoietins and receptor tyrosine kinases, Tie-1 and Tie-2, participate in vessel maturation. Tie-1 is associated with EC differentiation and the establishment of blood vessel integrity. Tie-2 is involved in vascular network formation [30, 31] and with its ligand Ang1 is involved with recruitment of pericytes and smooth muscle cells. This further strengthens the interactions between ECs, surrounding support cells and the matrix [32].
Matrix deposition is also critical at the late stage of angiogenesis. It facilitates vessel maturation and inhibits cell migration and proliferation.
Laminin is thought to regulate this process. In addition, TGFj31 and Ang1 also stimulate ECM production. Together these cellular events contribute to the process of capillary morphogenesis.
1.1.4 Control of angiogenesis - the balance hypothesis
Angiogenesis is driven by numerous mediators produced by a variety
of cells under different conditions. These mediators are either soluble, ECM
or components of the ECM itself.
It is now widely accepted that angiogenesis is regulated by a balance between proangiogenic and anti-angiogenic factors. This balance determines whether the ECs of the vascular wall remain in a state of vascular homeostasis or whether they undergo neovascularisation. Thus, changes in the net balance of these positive and negative signals mediate the angiogenic switch. In activated endothelium, angiogenic activators are up regulated. Conversely, endothelial quiescence is achieved by the dominance of negative regulators [33]. It is possible that different tissues with different
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physiological features may use distinct approaches to activate the angiogenic switch.
At present there are more than a dozen angiogenic and anti angiogenic factors that have been isolated and characterised. Most of the angiogenic factors have direct effects on EC migration, proliferation or both.
Some indirectly stimulate host cells to produce angiogenic factors. Activators and inhibitors of angiogenesis are listed in Table 1.1.
Table 1.1. Angiogenic activators and inhibitors
Angiogenic Activators Angiogenic Inhibitors
Acidic and basic fibroblast growth Soluble VEGFR1 (sVEGFR1) factor (aFGF and bFGF) Ang2 Vascular endothelial growth factors (VEGF-A, VEGF-8) Prothrombin kringle 2, Antithrombin Ill fragment Vascular endothelial growth factors receptor (VEGFR) Thrombospondin 1 and 2
Angiopoietin 1 (Ang1), Tie2 Angiostatin
PDGFBB and receptors Endostatin (fragment of collagen XVIII)
TGFf31 and TGFf3 receptor Vasostatin, Calreticulin
Platelet factor 4 (PF4)
VE-Cadherin, PECAM TIMPs, MMP inhibitors and PEX
Plasminogen activator, Plasminogen IFNa, f3, y, IP10, IL4, 12 and 18 activator inhibitor 1 Meth1 and 2 Metalloproteinases (MMPs) Osteopontin fragment Cyclooxygenase (COX) 16 kDa N-terminal fragment of Nitric oxide synthase (NOS) Prolactin
Chemokines Somatostatin
Fragment of Secreted Protein Acidic and Rich in Cytsteine (SPARC)
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1.1.4.1 Angiogenic stimulators
Angiogenic factors can be classified into two categories, specific and non-specific angiogenic inducers. The former include factors that act directly on ECs while the non-specific angiogenic inducers are those which exhibit their functions via accessory cells, such as inflammatory cells and other non
ECs.
1.1.4.1.1 Direct angiogenic inducers
This group of angiogenic molecules is expressed widely in cells such as macrophages, fibroblasts, smooth muscle cells, ECs and a number of tumour cell lines. The most extensively studied molecules in this category are members of the VEGF family, such as VEGF-A, VEGF-B, VEGF-C and the first relative of VEGF identified, placental growth factor (PIGF). The biological activity of these factors is mainly restricted to ECs via the two structurally related PDGF receptor-like tyrosine kinases, VEGFR-1 and
VEGFR-2.
VEGF regulates permeability and promotes vascular EC proliferation
[34, 35]. These activities are mediated through its tyrosine kinase receptors,
VEGFR-1 (previously known as Flt-1) and VEGFR-2 (previously known as
KOR or Flk-1). VEGFR-1 and VEGFR-2 are expressed mainly in the blood vascular endothelium [36].
VEGF plays an important role in vascular development during embryogenesis and it continues to be critical in early post-natal growth.
Carmeliet and co-workers demonstrated that disruption of a single VEGF allele in mice results in embryonic lethality due to severe vascular
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abnormalities [37). Furthermore, studies have shown that even a subtle alteration in VEGF expression during embryonic development results in severe abnormalities leading to embryonic or early post-natal death [38, 39].
In the adult, perturbation of VEGF expression seems to have a less profound effect. It appears that VEGF affects only those structures that actively engage in vascular remodelling, such as bone [40], ovarian corpus lutei [41] and pregnancy-associated angiogenesis in the mammary gland [42]. Other direct angiogenic inducers are listed in Table 1.2.
Table 1.2. Angiogenic molecules acting directly on endothelial cells
Direct angiogenic inducers bFGF/aFGF Platelet-derived EC growth factor Hepatocytes growth factor Interleukin-a Platelet activating factor Granulocytes-colony stimulating factor Proliferin Cytokine-inducible endothelial gene product (861) Soluble vascular cell adhesion molecule Soluble E-Selectin Tat protein of HIV-1 Angiogenin
1.1.4.1.2 Indirect angiogenic inducers
The angiogenic nature of these factors relates to their ability to stimulate a broad range of target cells other than ECs. For example stimulated monocytes secrete angiogenic factors that directly affect ECs.
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Proteins that belong to this category are erythropoietin, gangliosides, hepatocytes growth factor, insulin-like growth factor, prostaglandin E1/E2,
TGFcx, TGFJ3 and TNFcx. An exception to the above is TGFJ3 and TNFcx, which have been shown to have both stimulatory and inhibitory effects on angiogenesis. In vitro, low concentrations of TGFJ3 potentiates bFGF and
VEGF-induced angiogenesis but inhibits EC tube formation at high concentrations [43]. In vivo, both TGFJ3 and TNFcx stimulate the secretion of direct acting inducers from stromal and chemoattracted inflammatory cells.
1.1.4.2 Angiogenic inhibitors
Maintaining vascular patency and homeostasis is an active and an ongoing process that is dependent on the role of many anti-angiogenic factors. It is now quite clear that angiogenic inhibitors must work in concert with the angiogenic inducers to regulate angiogenesis (33].
The first hint that endogenous anti-angiogenic factors exist came in the early 1980's with the findings that ex Interferon and PF4 could inhibit EC chemotaxis and proliferation, respectively (33]. However, it was not until the discovery of thrombospondin in the early 1990's that the importance of angiogenic inhibitors began to be appreciated. A rapid increase in the number of anti-angiogenic substances is evident in the last decade, many of which have been well characterised. Angiogenic inhibitors have received much attention because of their potential application in the treatment of angiogenic related diseases.
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Thrombospondin was one of the first angiogenic inhibitors to be discovered [44]. Subsequently, a 16 kDa N-terminal fragment of prolactin was found to be a potent angiogenic inhibitor [45].
The more recent discovery of the anti-angiogenic members include angiostatin [46] and endostatin [47]. Other factors with known anti angiogenic effects are listed in Table 1.1. Despite the intense research over the last decade, the precise mechanism of action for these factors remains unknown. Their mechanism(s) of production are also mainly undetermined.
Interestingly, most of the known anti-angiogenic factors derive from larger molecules that have no anti-angiogenic activity, for example the 29 kDa fragment of fibronectin [48], the 16 kDa fragment of prolactin [45], the internal fragment of plasminogen, angiostatin [46], the fragment of collagen XVIII, endostatin [47] and anti-thrombin Ill fragment [49]. This pathway may represent the mechanism by which the release and storage of these factors are regulated. Such a mechanism ensures that these natural inhibitors are available for immediate use by simply breaking down the larger proteins.
1.1.5 Triggers of angiogenic responses
As described earlier, angiogenesis is regulated by many factors and at multiple levels. Many of the signals controlling angiogenesis are mediated by specific interactions between proteins, including ligand-receptor, cell extracellular matrix components and soluble factors such as pro- and anti angiogenic factors. The activation of these factors requires an appropriate stimulus and their activities also depend upon the presence of other factors in the environment of the responding endothelium [50].
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Both environmental and genetic factors are involved in the initiation of the angiogenic response. Examples of environmental factors are hypoxia, inflammation and mechanical stress. Genetic factors like activation of oncogenes and tumour suppressor genes, also play a role in angiogenic activation.
1.1.5.1 Hypoxia
Angiogenesis occurs when there is a sustained and local need for increase blood flow. The two conditions that lead to neovascularisation are increase metabolic need and inadequate blood flow. Under such conditions, the normal physiologic responses will be invoked to increase vascularity.
Low oxygen tension or hypoxia is an important environmental factor that has long been recognised as an angiogenic stimulus. Studies have demonstrated that under hypoxic conditions, there is an up-regulation of genes whose protein products increase 0 2 delivery or facilitate metabolic adaptation to hypoxia [51]. Hypoxia in a tumour, for instance, can specifically induce the expression of various angiogenic activators such as VEGF [52,
53]. Indeed, one of the major drives of malignant progression and metastasis is hypoxia. Tumour cells, like their normal counterparts have the ability to sense oxygen concentration. This activates a signalling pathway, which results in the expression of hypoxia-regulated genes. Semenza et al. demonstrated that hypoxia activates hypoxia inducible transcription factors
(HIFs). The activated HIFs induce the expression of several angiogenic stimulators, such as VEGF, nitric oxide synthase (NOS) and PDGF [54].
Moreover, hypoxia markedly elevates transcriptional activation of the VEGF
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receptor and Flt-1 promoter [55]. The expression of the angiogenic factor,
TGFP 1 is also induced by hypoxia.
1.1.5.2 Mechanical injury
Cellular injury by mechanical force stimulates the release of angiogenic factors. These mechanical influences include shear stress, stretch or deformation of the ECs [56]. Cellular injury or wounding has been shown to stimulate the release of FGF [57]. Furthermore, shear stress elevates the level of VEGF in the heart and in the isolated myocytes [58, 59].
1.1.5.3 Inflammation
Cells of the immune system like monocytes/macrophages, lymphocytes, platelets, mast cells and leucocytes influence angiogenesis by promoting the secretion of angiogenic factors. They produce factors, such as
VEGF, ang1, bFGF, TGFp1, PDGF, TNFa, hepatocytes growth factor (HGF), insulin like growth factor-1 (IGF-1), and monocytes chemoattractant protein-1
(MCP-1) [60]. These factors act directly on ECs or stimulate surrounding cells to release angiogenic factors or modulation of receptor activities [61-63].
In addition, haematopoietic cells contain proteases which facilitate the degradation of the ECM.
1.1.5.4 Genetic influence
Genetic alterations stimulate the onset of angiogenesis. There is increasing evidence that the angiogenic switch is governed in part by oncogenes and tumour suppressor genes [64]. There are at least fifteen
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oncogenes that are known to up-regulate expression of pro-angiogenic proteins and some down regulate expression of angiogenic inhibitors. For example, ras oncogene up regulates the expression of VEGF [65, 66]. Other oncogenes that also regulate VEGF expression include bcl-2 [67, 68], V-src and HER-2. Beside VEGF, mutant ras genes are known to up regulate the expression of a variety of other growth factors that are either direct or indirectly stimulate angiogenesis, such as TGFa. and TGFJ3. Oncogenes c myb and sis also induce the expression of a variety of angiogenic molecules.
Furthermore, activated oncogenes can affect angiogenesis by promoting the secretion of enzymes, which degrade BM and ECM [69, 70].
The p53 tumour suppressor gene plays a role in angiogenesis [71].
Cells lacking p53 function have reduced production of TSP-1, a potent inhibitor of angiogenesis. Dameron and Volpert demonstrated that wild-type p53 regulates TSP-1 expression in fibroblasts and mammary epithelial cells
[72, 73]. In another study, the angiogenic activity of p53 null glioblastoma cells was shown to be inhibited upon the introduction of an inducible form of wild-type p53. These studies consistently demonstrated the correlation between the loss of p53 and enhanced in angiogenic activity. In the context of tumours in vivo, the proliferative pro-angiogenic character of the ECs is most likely the result of a combination of mutations in various tumour suppressors and oncogenes.
1.1.6 Tumour angiogenesis
The first demonstration that tumours are highly vascularised was made as early as the 1900s. In the 1970s, angiogenesis was identified as a
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critical component for tumour growth. A considerable amount of evidence suggests that tumours cannot grow beyond 1-2 mm in diameter, without requiring new blood vessel formation. These newly formed vessels carry essential nutrients, oxygen and growth-regulatory molecules to the cells (74].
In addition to providing cells with their metabolic requirements, these new vessels also allow contact between cells and the circulation. Thus tumour angiogenesis aids the process of metastasis. Neovascularisation is therefore a critical step for rapid expansion of tumour mass.
This observation raised the question of when angiogenesis is activated during the course of tumour progression. Angiogenesis may simply be an inevitable consequence of the tumour becoming size-limited by the lack of vascularisation or perhaps, the ability to switch on angiogenesis is a crucial characteristic that the developing tumour must acquire to survive.
Studies in transgenic mice and human carcinomas support the latter hypothesis (33]. That is, angiogenic activation is a discrete component of the tumour phenotype, and is often activated at the pre-malignant stage and persists in the subsequent stages of tumour development (33].
It is now widely accepted that a switch to an angiogenic phenotype or the initiation of angiogenesis within a tumour depends on the net balance of angiogenic activators and inhibitors that are turned on or off at specific stages of tumour development (75]. The observation that tumour growth is governed by pro- and anti-angiogenic factors has led to the concept of inhibiting tumour growth by blocking tumour angiogenesis (76, 77]. A considerable amount of evidence is now available to support the benefits of manipulating angiogenesis for the control of tumour growth. The use of
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angiogenesis inhibitors is increasingly recognised as being an important part of future cancer therapies. With improved methods of early cancer detection, it might be possible to inhibit angiogenesis at the early stage of tumour development, thus preventing its progression to invasive carcinoma.
1.1. 7 Antiangiogenic therapy
Angiostatin and endostatin are among the most potent of the known anti-angiogenic drugs. Angiostatin targets proliferating capillary ECs, thus stopping new vessels from forming within a tumour while sparing quiescent
ECs lining the healthy vessels. The benefit of such specificity is minimal toxic side effects. In addition, resistance to angiostatin did not appear to develop in animals. Pre-clinical studies of anti-angiogenic agents showed that the efficacy of anti-angiogenic therapy is optimal when it is administered daily or intermittently over a long period of time with no break in therapy.
Thus far, there has been no report of drug resistance associated with long term use of angiogenic inhibitors in animal studies nor in Phase 1/11 clinical trials. Indeed, anti-angiogenic therapy has been proposed as a potential approach to avoid drug resistance in the treatment of cancer.
Furthermore, studies have shown that strategies combining anti angiogenic therapy with conventional chemotherapy in tumour bearing animals can be curative. Thus targeting both the endothelial cell compartment of a tumour as well as the tumour cell compartment is more effective than targeting either cell compartment alone [78]. These findings suggest that angiogenesis inhibitors may eventually be used to broaden conventional therapy. Chronic anti-angiogenic therapy following the
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completion of chemotherapy, radiation or surgery may be used to prolong dormancy of microscopic metastasis or to stabilise residual disease [79]
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1.2 PLASMINOGEN/PLASMIN SYSTEM
1.2.1 Introduction
Plasminogen is the circulating zymogen of the active serine
proteinase, plasmin. The mechanism of plasminogen activation consists of
highly regulated processes involving serine proteases, protease receptors,
and protease inhibitors. The conversion of plasminogen to plasmin is
catalysed by several activators. The two main activators in vivo are
urokinase plasminogen activator (uPA) and tissue-type plasminogen
activator (tPA). Two specific and fast-acting plasminogen activator
inhibitors, Type 1 and Type 2 (PAl-1 and PAl-2) regulate their activities.
The plasminogen system has two main roles. First, it maintains hemostasis and vascular patency by regulating the breakdown of fibrin depositions in the circulation. Second, it degrades ECM protein to assist cellular migration during tissue repair and remodelling. Plasminogen degrades ECM proteins either directly or by activation of matrix-degrading enzymes. In addition, plasminogen is the precursor molecule of the angiogenic inhibitor, angiostatin.
1.2.2 Structure
The amino acid sequence of plasminogen has been reported for human plasminogen [80, 81]. Its nucleotide sequence has also been established [82]. The genetic locus for human plasminogen has been mapped to the long arm of chromosome 6 q26-27 [83].
Plasminogen is a single polypeptide chain of 810 amino acid residues, which is processed to a 790 amino acid mature protein containing
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approximately 2% carbohydrate [84). Plasminogen is calculated to have a molecular mass of 92,000 Da. The native form of plasminogen has NHr terminal glutamic acid, termed Glu-plasminogen. This native plasminogen can further subdivide into Glu-1 and Glu-2 plasminogen, based on the carbohydrate composition and the extent of the glycosylation.
The structure of plasminogen is comprised of an NH2-terminal domain
(the heavy chain) and a protease domain (the light chain) at the C-terminal.
The heavy chain (residues 78-560) is characterised by five homologous triple-disulfide bonded domains referred to as kringles. The light chain
(residue 561-790) contains the catalytic site. Two disulfide bonds link the two chains. One links Cys548 to Cys666 and another links Cys558 to Cys566
(Figure 1.2). In the plasma of a healthy adult, it is present in a concentration of approximately 1.5 µM [85). Plasminogen is synthesised in virtually all tissues, for example in kidney [86], testis [87] and epidermal cells [88]. The major production site is the liver [89].
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LBS 1 LBS 2 LBS 3 LBS4 LBS5
Figure 1.2. Scheme of the tertiary structure of human plasminogen. Sites of cleavage by plasminogen activators as well as by plasmin are shown. Heavy (-) and light (-) chains of plasmin are indicated. Plasmin is formed by cleavage of the Arg560 -Val561 bond by plasminogen activators. Disulfide bonds (t-M), position of the Cys168-Cys296 inter-kringle disulfide bond ( .... ), carbohydrate (-CHO) and the amino acids forming the active side in plasmin (His, Asp, Ser) are shown. Adapted from [90].
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1.2.3 Kringle domains of plasminogen
There are five kringle domains in plasminogen. Each consists of 80
amino acid residues with a molecular weight of approximately 10,000. Each
kringle is held in a loop structure by three disulfide bonds. Kringles share a
high degree of sequence homology with each other. Similar kringle motifs
exist in a number of proteins involved in fibrinolysis, such as prothrombin,
tPA, uPA and factor XII [91].
The kringle domains contain lysine-binding sites (LBS), which interact
specifically with certain amino acids or related compounds such as E-amino
caproic acid (EACA), trans-4-aminomethylcyclohexane-1-carboxylic acid
(tranexamic acid) and other lysine analogues. Each kringle displays
different specificities for these compounds. A study by Markus reported that
Glu-plasminogen contains one strong LBS for EACA of Ko 9x10-6 M and four
weaker sites of average Ko 5x10-3 M [92]. Subsequently, Lerch
demonstrated that K1 plasminogen contains the tightest lysine binding site
for EACA, followed by K4 and KS. K2 interacts weakly, while K3 does not
interact with the ligand to a measurable extent [93].
The LBS contained within the kringle domains are responsible for the binding of plasminogen to various proteins in solution as well as to cells.
They interact with cell-associated actin [94], they are critical for binding of plasminogen to cell-associated a-enolase [95] and they have been implicated in promoting neutrophil adherence to endothelial cells [96]. Generally, they bind proteins through the carboxy-terminal lysines [97]. They are primarily responsible for binding of plasminogen to its substrates, such as fibrinogen and fibrin [98], a2-antiplasmin [99] and thrombospondin [100]. Various
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reagents displaying affinity for kringle domains in plasminogen block the interaction between plasminogen and its ligands [101].
1.2.4 Variants of plasminogen
There are two major forms of plasminogen in the plasma. The native form, Glu-plasminogen (Glu-Pg), contains glutamic acid at the NH2-terminal, and a truncated form, Lysine-plasminogen (Lys-Pg), which has an NH2- terminal starting with either Lys78 or Val79.
1.2.4.1 Glu-plasminogen
Glu-Pg exists in a closed conformation. This form of plasminogen is more stable than its counterpart, Lys-Pg, with a plasma half-life of 2.24 days instead of 0.8 days. The stability of the protein results from a specific interaction between the NHrterminal peptide and the A-chain. This interaction is lost upon removal of the peptide, such as when Glu-Pg is converted to Lys-Pg.
1.2.4.2 Lys-plasminogen
Following plasminogen activation by uPA or tPA, the plasmin generated is responsible for the conversion of Glu-Pg to Lys-Pg. This cleavage leads to the release of a peptide containing 77 amino acids from the NH2-terminal [102]. Subsequently, Hallar and co-workers demonstrated that the conversion of Glu-Pg to Lys-Pg is a cell surface-mediated event, which occurs upon binding of circulating Glu-Pg to the endothelial cell surface [103, 104].
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Glu-Pg becomes extended in association with lysine [105]. This open form of plasminogen has properties similar to Lys-Pg. The conformational change also makes plasminogen more susceptible to cleavage by its activators. Lys-Pg is a preferred substrate for both tPA and uPA and is therefore more efficiently activated by these proteins compared to Glu-Pg
[106]. Lys-Pg also represents a form preferentially associated with the cell surface. Hajjar and co-workers showed that endothelial cells bind Lys-Pg with 2.6 times the affinity for Glu-Pg [104).
The two forms of plasminogen can be further divided into two subgroups, based on the extent of glycosylation. Two variants of Glu plasminogen, Glu1-Pg and Glu2-Pg are separated on lysine sepharose chromatography by gradient elution with EACA. Both forms exhibit microheterogeneity due to varying amounts of sialic acid on both the N- and
O-linked polysaccharides. Glu1-Pg contains a carbohydrate moiety of 10-11 monosaccharide units N-linked to Asn288 and a carbohydrate moiety of 3-4 units O-linked to Thr345. Glu2-Pg has only a single glycosylation site at
Thr345 and contains less carbohydrate per mole of protein than Glu1-Pg
[107, 108]. Further study by Powell and co-workers demonstrated that Glu2-
Pg, despite lacking glycosylation at Asn288, possesses identical amino acid sequence to the corresponding variant 1 peptide at the Asn288 region [109].
This finding provided evidence that the heterogeneity in glycosylation is due to incomplete post-translational modification of plasminogen and not a difference in the primary sequence of the two forms.
A number of analyses have demonstrated that glycosylation affects the functional properties of Glu1 and Glu2 plasminogen. First, glycosylation
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at Asn288 in Glu1-Pg reduces its affinity for lysine sepharose, fibrin and a2- antiplasmin [84, 11 0]. Glycosylation is also thought to be responsible for a reduced affinity of plasminogen for promyeloid leukemic U937 cells [111].
This reduced affinity may be due to shielding of kringle 4, thus disrupting recognition. Second, a study analysing the distribution of plasminogen revealed that a higher concentration of Glu2-Pg was present in the extravascular compartment than the intravascular compartment. This observation led to speculation that the N-linked carbohydrate moiety of Glu1-
Pg may interfere with transport mechanisms of plasminogen. This may be due to the binding of carbohydrate to a receptor in the cell, hence preventing its export from the cell. Alternatively, it may physically hinder the binding of plasminogen to receptors on the cell surface or extracellular matrix macromolecules, thereby disrupting the transport mechanism. Third, the rate of conversion of plasminogen by either uPA or tPA is glycosylation dependent, with Glu1-Pg being more rapidly activated than Glu2-Pg.
1.2.5 Plasminogen activation
Various plasminogen activators convert plasminogen to the two-chain plasmin by cleavage at a single peptide bond, Arg560-Val561, in the serine proteinase module [112]. The heavy-chain of plasmin contains a pre activation peptide and five kringle domains. The light-chain contains an active catalytic triad composed of His602, Asp645 and Ser740 residues.
Plasmin has a molecular mass of approximately 84,000 Da. Two interchain-disulfide bonds at residues Cys548-Cys666 and Cys558-Cys566 of plasmin bridge the two chains of plasmin. Plasmin is one of the few
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proteolytic enzymes which has broad substrate specificity and is capable of degrading a large number of extracellular protein components such as fibronectins, laminin, vitronectin and the protein core of proteoglycans.
Binding of both plasminogen and plasmin to fibrin localises the proteinase activity to the required sites of action. In addition, plasmin binds to a cell surface receptor. This cell surface-bound plasminogen/plasmin is readily activated. Its enzymatic activity is also enhanced. Gonzales and co workers demonstrated that cell surface-bound plasmin exhibits increased catalytic efficiency relative to plasmin in solution, suggesting that receptor binding may induce conformational change favouring plasmin activity [113].
Cell surface-bound plasmin can also activate certain MMPs [114], latent elastases [115] and pro-plasminogen activators [116]. Furthermore, both tPA and uPA also form a complex with cell surface receptors. Together, these interactions bring both plasminogen and its activators into a close, favourable environment for activation.
Specific plasminogen activator inhibitors, PAl-1 and PAl-2, regulate the plasminogen/plasmin system by inhibiting both uPA and tPA. Another level of regulation is achieved by the action of serine protease inhibitors, a2- antiplasmin and ar macroglobulin, which inhibit plasmin once formed.
1.2.5.1 Plasminogen activators
The two principal activators of plasminogen are tPA and uPA. Both are the products of related genes, belonging to the serine protease gene family. They have distinct structural determinants found in the noncatalytic regions of the protein and it is suggested that this difference accounts for
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their different role in the target environment. The activation of plasmin by tPA is primarily involved in fibrinolysis, whereas the activity by uPA is predominantly associated with extracellular matrix proteolysis, such as during wound healing, embryogenesis, tumour metastasis and angiogenesis.
1.2.5.1.1 Tissue-type plasminogen activator
tPA is secreted as a single-chain polypeptide, consisting of 527 amino acids. It is synthesised and secreted by vascular endothelial cells. The single-chain expressed form is cleaved at Arg275-lle276 by plasmin forming a two-chain disulfide-linked molecule. Both forms of tPA have comparable enzymatic activity [117]. The A-chain of tPA consists of four connected structural domains. From the NH2-terminal, these domains include a fibronectin finger, an epidermal growth factor-like domain followed by two kringle motifs (K1 and K2) while the B-chain contains the catalytic active site of the enzyme.
The finger domain in tPA shares a structural similarity to the fibrin binding site in fibronectin. This structure was therefore thought to be partly responsible for the fibrin-binding property of tPA. In subsequent studies, it was reported that kringle 2 of tPA contributes to most of the fibrin affinity of tPA. The first kringle on the other hand, has no known role. tPA binds fibrin with high affinity and it is known that fibrin enhances the efficiency of tPA dependent plasminogen activation on clot surfaces almost 1000-fold, through the formation of ternary complexes [118].
In addition to binding fibrin, tPA also binds thrombospondin, laminin and fibronectin. It has also been shown to bind many cells including human
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endothelial cells, human fibroblasts, rat and human hepatoma cells, bovine alveolar macrophages, human smooth muscle cells, neuronal cells and platelets. Cell surface binding in some cases can also stimulate the enzymatic activity of tPA.
The binding of tPA to cells is mediated by a specific tPA receptor.
Multiple receptors for tPA have been reported by various groups. Hajjar and co-workers identified a specific endothelial cell co-receptor for tPA and plasminogen, a 40,000 Da polypeptide, identical to Annexin II [119). They showed that both tPA and plasminogen interacted with Annexin II in a non competitive manner, implying that they bind to different sites on Annexin II.
Binding of tPA to Annexin II specifically enhances the catalytic efficiency of tPA-dependent plasminogen activation 60-fold [120). In addition, Annexin II appears to be a tPA specific receptor which does not bind uPA.
Furthermore, Dudani and co-workers reported a tPA plasminogen receptor expressed on endothelial cells. This 45,000 Da receptor, unlike
Annexin 11, contains a common binding site for both tPA and plasminogen, since their binding can be mutually inhibited [121).
Another receptor for tPA was identified as a 20,000 Da protein expressed on human umbilical vein endothelial cells (HUVEC) [122]. This receptor specifically interacts with tPA to form a 90,000 Da complex.
1.2.5.1.2 Urokinase type plasminogen activator
uPA is a glycoprotein with a molecular mass of 54,000 Da. Its expressed form is an inactive single chain proenzyme (scu-PA). Cleavage of this precursor by plasmin at Lys158-lle159 results in the formation of a
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catalytically active form. The active uPA consists of two chains, light or A chain and heavy or B chain, bridged by a single disulfide bond. Other than plasmin, a number of proteases can also activate scu-PA, including trypsin proteases, kallikrein, factor Xlla, MMPs (thermolysin), and cysteine proteases
(cathepsin Band L) [123].
The light chain of uPA consists of the NH2-terminal end of the inactive form. It is characterised by two structural determinants, a growth factor domain which is homologous to the receptor-binding region in epidermal growth factor and a kringle motif resembling the kringle structures of plasminogen but lacking LBS. The B-chain contains the catalytic site. uPA is produced and secreted by many cell types, in particular those engaged in angiogenesis and migratory process, such as endothelial and carcinoma cells. Certain angiogenic factors are known to stimulate endothelial cell production of uPA.
In the extravascular compartment, uPA and plasmin play a direct role in the degradation of ECM components. uPA can also directly activate latent growth factors such as the precursor form of hepatocyte growth factor/scatter factor (pro-HGF/SF) and indirectly activate latent transforming growth factor
P (pro-TGFp) [124], which suggests uPA may function in tumour cell migration and modulating matrix degradation.
The actions of uPA are mediated while the protease is bound to a specific membrane receptor on the cell surface, the uPA receptor (uPAR).
This receptor-bound active uPA converts plasminogen to plasmin.
Furthermore, uPA and plasmin have been shown to activate single-chain
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uPA thus generating a self-maintained feedback mechanism of single-chain uPA and plasminogen activation.
1.2.5.1.3 Urokinase plasminogen activator receptor
uPAR is a highly glycosylated 55-60,000 Da protein anchored to the
plasma membrane by a glycosyl-phosphatidylinositol (GPI) moiety [125].
uPAR has been identified on cells of the immune system, vascular
endothelial cells, smooth muscle cells, keratinocytes and placenta
trophoblasts as well as malignant cells. A variety of growth factors and
tumour promoters, including TGF~, EGF, HGF/SF and phorbol esters have
be shown to upregulate the expression of uPAR [126].
uPAR binds catalytically active uPA and its precursor pro-uPA via the
growth factor domain of the A-chain [127, 128]. This binding is specific,
saturable and of high affinity (Kci=0.1-1nM) on most cells studied [129].
Behrendt and co-workers demonstrated that the formation of receptor-ligand
complexes does not result in internalisation or receptor down regulation and
that the proteolytic activity is maintained [125].
uPA dissociates very slowly from its receptor and remains active on
the cell surface for several hours [130]. This, together with the
simultaneous binding of plasminogen and plasmin to membrane associated
receptors potentially stimulates a highly efficient proteolytic cascade. First,
bound uPA enhances the activation of pro-uPA, rapidly converting
plasminogen to plasmin. Subsequently, plasmin causes further activation of
pro-uPA. Second, membrane-bound plasmin is protected from its inhibitor,
a2-antiplasmin, and has higher catalytic efficiency than plasmin in solution.
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Collectively, these events localise protease activity to the cell surface and
restrict the activity to the pericellular zone.
1.2.5.2 Plasminogen activator inhibitors
PAl-1 and PAl-2 belong to the serine proteinase inhibitor (serpin) gene superfamily. PAl-1 inhibits both uPA and tPA, whereas PAl-2 is a more effective inhibitor of uPA than tPA. Neither inhibitor interacts with single chain uPA.
PAl-1 and PAl-2 form complexes with uPA in solution, which inactivate
the catalytic site of the 8-chain of active uPA. When uPA is bound to u
PAR, PAl-1 and PAl-2 are less effective inhibitors. PAl-1 and PAl-2 regulate
the proteolytic cascade at the initial stage of plasminogen activation, while
a2-antiplasmin regulates the level of plasmin once it is formed. Thus, they
contribute to the proteolytic control of the plasminogen system.
1.2.5.2.1 Plasminogen activator inhibitor type 1 (PAl-1)
PAl-1 is a glycoprotein of 379 amino acids with a molecular mass of
45-50,000 Da. PAl-1 exhibits a unique conformational flexibility, in which it can convert between three conformations, an active, a latent and a substrate form [131]. It is synthesised as an active molecule but is rapidly converted to an inactive or 'latent' form with a half-life of approximately 2 hours under physiological conditions. The latent form can be partially reactivated by denaturing reagents such as sodium dodecyl sulfate, guanidinium chloride and urea. A third conformation of PAl-1, reacting as a non-inhibitory substrate for various target proteases has also been suggested [132, 133].
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Blood PAl-1 is mostly found in platelets and a small pool is present in plasma. Endothelial cells are believed to be the main source of plasma PAl-
1, although smooth muscle cells have also been suggested to synthesise
PAl-1. In addition, a variety of cells in culture produce PAl-1. A number of cytokines, growth factors, hormones and endotoxins regulate the expression of PAl-1.
PAl-1 binds both uPA in solution and receptor bound uPA. The uPA/PAl-1 complex binds to the cell surface with the same binding specificity as uPA alone but with a slightly lower affinity [134]. Upon binding of PAl-1 to uPA/uPAR, the complex is rapidly internalised and degraded. This rapid removal of active proteolytic complexes from the surface of cells is one way by which PAl-1 controls the extent of ECM degradation. In addition to binding to uPA in solution and the receptor bound form, active PAl-1 binds vitronectin and heparin. The latent form of PAl-1 also binds vitronectin but with low affinity. Binding of PAl-1 to vitronectin and heparin protects it from oxidative inactivation and stabilises its functional activity in solution [135,
136].
1.2.5.2.2 Plasminogen activator inhibitor type 2 (PAl-2)
PAl-2 is a secreted 60,000 Da protein but accumulates as a 47,000 Da intracellular protein [137]. Most newly synthesised PAl-2 resides intracellularly in an unglycosylated form. It has been suggested that it undergoes glycosylation before secretion [138]. Unlike PAl-1, PAl-2 is stable in soluble form for a prolonged period. PAl-2 is produced by a variety of cell
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types such as monocyte/macrophage, fibroblasts, tumour and endothelial cells.
1.2.5.2.3 Protease-nexin 1 (PN-1)
PN-1 is another member of the serpin family known to inhibit plasmin.
PN-1 is a 45,000 Da glycoprotein containing 392 amino acids. It is an effective inhibitor of two-chain uPA but is a slow inhibitor of both single-chain tPA and two-chain tPA. It does not interact with single-chain uPA. PN-1 forms covalent complexes with many serine proteases, such as trypsin, thrombin, urokinase and plasmin. Proteases bound to PN-1 are readily endocytosed via high-affinity nexin receptors on the plasma membrane of cells and is rapidly degraded in the lysosome.
1.2.5.2.4 Alpha 2-antiplasmin
a.2-antiplasmin is an extremely efficient protease inhibitor of the serpin superfamily. It is a single-chain glycoprotein of 67,000 Da and is the principal physiological inhibitor of plasmin. a.2-antiplasmin is present in plasma at a concentration of approximately 1µM, which translates to a plasmin half-life of
0.01-0.1 seconds. Inactivation occurs by the rapid formation of 1:1 equimolar complex [139]. This rapid inactivation can occur at a much slower rate if plasmin activity is generated by receptor bound uPA [140]. Inactivation of on plasmin by a.2-antiplasmin occurs when plasmin dissociates from the cell surface. The overall effect is that plasmin activity is restricted to the cell surface.
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The inhibition of plasmin by a2-antiplasmin occurs in two steps, a fast reversible second-order binding step followed by a slower irreversible first order reaction [99]. C-terminal lysine residues of a2-antiplasmin bind to LBS of plasmin is followed by the formation of a covalent complex involving the active site of plasmin [99]. Thus, occupation of LBS protects plasmin from inhibition by a2-antiplasmin.
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1.2.5.3 Plasminogen receptors
The plasminogen receptors represent a class of structurally diverse molecules that function to direct plasminogen activity to focal areas on the surface of cells, or are involved in the clearance of plasminogen activators.
Thus, they serve to regulate endogenous fibrinolytic activity. These receptors are characterised by their low affinity, high density and widespread distribution.
Plasminogen receptors are expressed by a wide variety of cells, including inflammatory cells, cultured rat hepatocytes and glioma cells, human tumour cells and endothelial cells from several species. The plasminogen binding capacity of many cells has been reported to range between 106 to 107 molecules bound per cell.
To date, there are approximately ten different plasminogen binding proteins reported on various types of cells. Among these receptors are annexin II, a-enolase, amphoterin, ganglioside and members of the low density lipoprotein receptor family, such as glycoprotein 330 and lipoprotein receptor-related protein (LRP).
Both a-enolase and Annexin II bind plasminogen via C-terminal lysines. Amphoterin represents a class of receptor without carboxy terminal lysines. It is not certain, therefore whether C-terminal lysines play a critical role in plasminogen binding to all identified receptors.
The functional consequences of plasminogen receptor occupancy include enhanced plasminogen activation. Receptor binding of plasminogen
(a-enolase and Annexin II) enhances the rate of uPA- and tPA-dependent plasminogen activation. Once activated, plasmin can further increase
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plasminogen activation. This is accomplished by proteolysis of membrane proteins, generating new carboxy terminal lysine residues, which can then serve as additional plasminogen binding sites. Furthermore, bound plasmin is protected from neutralisation by a 2-antiplasmin. Finally, the enzymatic activity of plasmin is enhanced while binding to cell surface receptors.
-. Inhibitors Processes:Fibrinolysis PAl-1& PAl-2 Ovulation Angiogenesis Wound healing Tumour metastasis * Plasminogen uPA
tPA Fibrin l Degradation of: Fibrinogen Plasmin ------____ • Laminin i Pro-collagenase ---·~ Collagenase --- _...,. Collagens
Figure 1.3. Plasminogen activation cascade. Contribution of plasminogen system to the breakdown of ECM components in diverse physiological processes. Adapted from [141 ].
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1.3 ANGIOSTATIN
1.3.1 Introduction
The importance of angiogenesis in diseases such as rheumatoid arthritis, atherosclerosis and tumour growth is now well recognised.
Recognising angiogenesis as being a useful target for therapy began in the early 1970s, when Folkman and Denekamp noted that tumours are highly vascularised and may be vulnerable to disruption of their blood supply.
However, it was not until the discovery of angiostatin in the 1990s that the therapeutic potential of anti-angiogenic molecules was fully appreciated.
It has been observed that resection of the primary tumour in cancer patients was sometimes followed by accelerated growth of its secondary metastasis. A similar phenomenon was described in a number of metastases models. In an effort to explain this phenomenon, multiple theories were proposed. These include the escape of tumour cells as a result of surgical procedures. Another proposed theory related to the depletion of available nutrients by the primary tumour. None of these hypotheses has led to a universally accepted mechanism. However, with the discovery of angiostatin in 1994, a new theory was put forward. O'Reilly and co-workers hypothesised that the primary tumour regulates the growth of its secondary tumours. They suggested that the primary tumour induced angiogenesis in its own vascular bed by generating excess angiogenic stimulators, thus allowing vascularisation and tumour growth. However, at distant sites, angiogenic inhibitors accumulate in excess of angiogenic stimulators, due to their longer plasma half life, thus angiogenesis is inhibited
(46].
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A review of the structure and functions of angiostatin as an anti angiogenic molecule will be discussed in the following sections.
1.3.2 Structure
Angiostatin was isolated from urine and plasma of mice bearing 3LL
Lewis lung carcinoma (LLC-LM) [46]. The purified angiostatin has a molecular weight of 38,000 and shares more than 98% sequence homology to plasminogen. Microsequence analysis revealed an NH2-terminus beginning at amino acid 98. On the basis of its mass, angiostatin was estimated to have an approximate C-terminus at amino acid 440. Thus the fragment would consist of the first four of the five kringle domains of plasminogen.
Furthermore, the same authors demonstrated that fragments derived from proteolytic cleavage of human plasminogen by elastase in vitro were essentially identical to its murine counterpart. Cleavage of plasminogen released three fragments with apparent molecular weights of 40,000, 42,000 and 45,000. They have similar NH2-termini at amino acids 97 or 99 and have comparable anti-proliferative activity on endothelial cells proliferation.
1.3.3 Angiostatin converting activity of various enzymes
Significant progress has been made in an attempt to understand the mechanism of angiostatin formation in vivo. Although angiostatin was detected in association with 3LL Lewis lung carcinoma and subsequently in other tumours, the source and mechanism of formation of angiostatin is not clear. It appears that tumour cells do not produce angiostatin directly, as
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they lack a detectable amount of mRNA for angiostatin or plasminogen [142].
O'Reilly suggested that the angiostatin molecule is formed by proteolytic cleavage of circulating plasminogen, possibly mediated by enzymes produced by tumour cells [46]. This hypothesis was supported by consistent reports using various human carcinoma cell lines. Gately and co-workers first reported the enzymatic activity expressed by human prostate carcinoma cell lines (PC-3, DU-145 and LN-CaP) [143]. All three tumours produce proteolytic activity that generates angiostatin from plasmin. Studies with protease inhibitors revealed the role of serine proteinase in the cleavage of plasminogen to form angiostatin. PC-3-derived angiostatin was shown to inhibit endothelial cell proliferation, migration and tube formation. A subsequent study by Heidtmann and co-workers reported the angiostatin converting activity of prostate-specific antigen (PSA), a serine proteinase secreted by the human prostate and human prostate cancer cells in vitro
[144].
In addition to serine proteases, matrix metalloelastase (MME) produced by Lewis lung carcinoma was shown to be involved in angiostatin generation from plasminogen. Dong and co-workers proposed that increased expression of cytokine, GM-CSF, in 3LL Lewis lung carcinoma
(LLC-LM) up-regulated the metalloelastinolytic activity in macrophages [145].
They used high GM-CSF producing cells in a metastasis model to support this data. They showed cells producing a high level of GM-CSF significantly inhibit lung metastasis of 3LL Lewis lung carcinoma, UV-2237 fibrosarcoma, metastatic melanoma K-1735 M2 and B16-F10 [146].
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Other members of the MMP family can also mediate angiostatin generation in vitro. These include stromelysin-1 (MMP-3), matrilysin (MMP-
7), gelatinase B/type IV collagenase (MMP-9) and MMP-12 [147, 148].
O'Reilly and co-workers identified gelatinase A (MMP-2) as an enzyme involved in angiostatin formation in LLC-LM carcinoma [149].
1.3.4 Function
O'Reilly showed that although angiostatin is derived from plasminogen, the anti-angiogenic activity was not found in plasminogen.
Angiostatin was shown to inhibit the proliferation of bFGF-stimulated bovine capillary endothelial (BCE), bovine aorta endothelial (BAE) and human umbilical vein endothelial (HUVEC) cells in a dose-dependent manner [46].
In contrast, it does not inhibit the proliferation of normal and neoplastic non endothelial cell lines, such as 3T3 fibroblasts, bovine retinal pigment epithelial cells, human fetal fibroblasts and 3LL murine Lewis lung carcinoma cells [46]. Until recently, the anti-proliferative properties of angiostatin were thought to be endothelial specific. However, Walter and co-workers reported that, in addition to ECs, angiostatin binds to smooth muscle cells of human coronary arteries [150]. Their findings suggest that binding is mediated by a plasminogen receptor via lysine binding sites, since excess plasminogen and
EA.CA inhibits the interaction. Furthermore, they showed that angiostatin inhibits HGF-induced proliferation and migration of rabbit aortic SMC. These effects could contribute to the anti-angiogenic role of angiostatin.
In vivo, angiostatin inhibits bFGF-induced angiogenesis in the chick
chorioallantoic membrane (CAM) assay over a concentration range of 0.1-
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100µg/embryo and Lewis lung metastases after removal of the primary tumour. In the mouse corneal micropocket assay, systemic administration of human angiostatin at 50mg/kg every 12 hours, inhibits bFGF-stimulated corneal neovascularisation by 85% compared to untreated mice. The inhibitory activity of angiostatin in the circulation could be detected five days after the removal of the primary tumour [151 ].
Systemic administration of angiostatin to immunodeficient mice bearing human carcinoma results in almost complete inhibition of tumour growth [151). It resulted in a 95 and 97% inhibition in the rate of growth of human breast and colon carcinoma respectively, while completely regressing the growth of prostate carcinoma. No apparent toxicity or resistance was detected even with treatment of angiostatin at 100mg/kg/day concentration for up to 60 days.
Subsequently, the suppressive effects of angiostatin have been reported in primary hemangioendothelioma [152), murine fibrosarcoma [153), rat C6 glioma and human MDA-MB-231 breast carcinoma in mice [154).
Angiostatin appears to also affect the growth of colorectal liver metastasis
(155) and ovarian cancer in mice [156).
1.3.5 Inhibitory activity of kringle domains of angiostatin
The endogenous murine angiostatin consists of K1-4. Studies using elastase digested fragments (K1-4) and recombinant K1-4 showed comparable potency to the native angiostatin, with a 90% inhibition of Lewis lung carcinoma in mice [46, 157). Thus, the anti-angiogenic activity is believed to reside within these kringle domains. Interestingly, smaller
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fragments of human angiostatin have been reported to display different inhibitory effects on endothelial cell proliferation and migration. Recombinant
K1 and K3 display similar inhibitory activity on endothelial cell proliferation with half-maximal effects (ED50) at 320 and 460 nM, respectively.
Recombinant K2 also displays significant inhibition but less than K1 and K3.
Recombinant K4 did not affect cell proliferation. Cao and co-workers also showed that recombinant K1-3 has greater inhibitory activity (<2-fold) compared with K1-4, and that removal of K4 from angiostatin enhances its inhibitory effect. It was thus suggested that K1, K2 and K3 are critical for the inhibitory activity of angiostatin [158]. A similar finding has also been documented using recombinant angiostatin containing only K1-3 [159, 160].
However, Ji and co-workers found that K4 has a potent inhibitory effect on cell migration despite having a marginal anti-proliferative activity [161]. In contrast, K1-3 was found to have only a modest anti-migratory effect. Thus, it was suggested that the overall anti-angiogenic activity of angiostatin is a combination of activities of different kringle domains. In addition, a plasminogen fragment containing K1-5 has also been identified to be a potent inhibitor of endothelial cell proliferation with a half-maximal effect at 50 pM [162]. This effect is approximately 50-fold greater than that of angiostatin. Furthermore, the co-incubation of angiostatin and KS produce a comparable effect to that produced by K1-5 alone.
Ji and co-workers suggested that conformational integrity of angiostatin is important for its inhibitory activity [161]. They showed that disruption of kringle structures, by reducing agents, markedly attenuates its activity. Their finding suggests that folding of kringle structures, as tandem
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domains held together by disulfide bonds, is critical for angiostatin to maintain its full inhibitory function. Lee and co-workers, however, demonstrated that mutation of Cys169 and Cys297 in recombinant K1-3 induced a conformational change and the loss of K2 lysine binding ability but no effect was observed on its anti-angiogenic activity [163]. Furthermore,
Cao and co-workers noted that a combination of K2 and K3 has more inhibitory activity than a recombinant K2-3 fragment [158]. They suggested that reduction in the interkringle disulfide bond between K2 and K3 may be necessary for the maximal inhibitory effect of K2-3. These reports clearly indicate that further research is necessary.
1.3.6 Kringle 5 of plasminogen
The KS domain of human plasminogen is not part of the angiostatin fragments described by O'Reilly. KS shares some sequence homology with
K1 (57.5%), K2 (46.25%), K3 (48.75%), and K4 (52.5%) and was shown to have potent inhibitory effects on both endothelial cell growth [164] and migration [165]. The KS fragment inhibits bovine capillary endothelial cell growth with half-maximal effect at approximately 50 nM and recombinant mouse KS also displayed comparable potency. These findings suggest that
K5 of plasminogen is also a selective inhibitor for endothelial cells.
1.3.7 Mechanism of action
Recently, Griscelli and co-workers suggested that the anti-proliferative effect of angiostatin may result from inhibition of cell cycle progression [154].
Previous reports indicated that the anti-angiogenic effect of angiostatin is the
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result of increased rate of EC apoptosis, in comparison to an unchanged rate of proliferation [33, 151].
The mechanism by which angiostatin exhibits its anti-angiogenic effect remains undetermined. Attempts to understand this mechanism have long been of interest to researchers. For example, Claesson-Welsh and co workers investigated the effect of angiostatin on growth factor induced signal transduction in EC. Both FGF-2 and VEGF induce angiogenesis by binding to specific cell surface-expressed receptors, leading to activation of a cascade of signalling proteins, resulting in specific cellular responses such as proliferation, migration or differentiation. Based on this, they explored the possibility that angiostatin functions by interfering with receptor binding and hence downstream signal transduction. They observed that angiostatin, while capable of stimulating apoptosis of endothelial cells, has no effect on the growth factor-induced signal transduction in these cells [166]. Thus the mechanism by which angiostatin induces EC apoptosis is independent of growth factor-induced endothelial cell proliferation.
A study by Redlitz and co-workers reported that angiostatin transiently reduces phosphorylation of the mitogen-activated protein kinases, ERK1 and
ERK2, in human microvascular ECs. Activation of ERK1 and ERK2 is required for the regulation of a number of cellular functions in angiogenesis.
Inactivation of ERK1 and ERK2 leads to inhibition of tube formation by sinusoidal ECs. The findings by Redlitz and co-workers indicate that the anti angiogenic effect of angiostatin might, at least in part, be mediated by interference of ERK1 and ERK2 activation [167].
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Furthermore, Moser reported the identification of an angiostatin binding receptor on the cell surface. ECs express cell surface receptors for plasminogen and since angiostatin specifically affects EC function, it raises the question whether angiostatin exerts its function by binding to a plasminogen receptor or other EC surface protein. Moser and co-workers found angiostatin bound to a 55,000 Da protein on the surface of HUVEC.
This angiostatin binding protein was identified as the a/~-subunits of ATP synthase [168]. Antibody to a-subunits of ATP synthase blocked the inhibitory effect of angiostatin on HUVEC by as much as 90%. These findings provide evidence that angiostatin binds to EC surface proteins and this binding may mediate its anti-angiogenic effects, inhibiting EC proliferation and migration [168].
Recently, angiostatin has also been reported to act as a non competitive inhibitor of ECM-enhanced, tPA-catalysed plasminogen activation with a Ki of 0.9+/-0.03 µM. Direct binding demonstrated that angiostatin binds tPA with an apparent Kd of 6.7+/-0.7nM. These observations suggest that tPA, when bound to angiostatin, prevents matrix enhanced plasminogen activation, thus providing indirect evidence that angiostatin affects cellular migration and invasion through the regulation of plasmin [169].
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1.4 PHOSPHOGLYCERA TE Kl NASE (PGK)
1.4.1 Introduction
PGK (adenosine-3-phospho-D-glycerate-1-phosphotransferase) (EC
2.7.2.3) was first recognized in yeast extracts in 1939 and was purified, in crystalline form, eight years later. The human PGK was initially isolated from erythrocytes. A variety of tissues such as liver, skeletal muscle and granulocytes have also been identified as a rich source of human PGK.
PGK is an important enzyme required for ATP generation in the glycolytic pathways of aerobes and anaerobes, and for carbon fixation in plants. It is found in high concentrations in the cytoplasm. Recently, its presence in the cell nucleus has also been revealed, along with other glycolytic enzymes such as lactate dehydrogenase (LOH) and glyceraldehyde-3-phosphate dehydrogenase (GAPDH). In addition to its catalytic role in glycolysis, PGK in the nucleus is thought to play a role in
DNA replication, transcription and DNA repair [170, 171). PGK also stimulates viral mRNA synthesis in the cytosol [172) and was detected on the surface of Candida albicans where it extends through the cell wall [173).
PGK binds Annexin II with high affinity. Annexin II is one of the major cell surface receptors for plasminogen.
1.4.2 Role of PGK in glycolysis
PGK is known as the sixth enzyme in glycolysis, where it catalyses the high-energy transfer of a phosphoryl group from 1,3-bisphosphoglycerate
(1,3-BPG) to ADP, generating the first ATP molecule in the glycolytic
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pathway with 3-phosphoglycerate (3-PG) produced as a biproduct (Figure
1.4).
0~ /0- 0~ P-, ~C-0/ "o- 1 H-C-0H I H-C-0 0 + HI '~P, -o/ "o-
1,3-Blsphosphoglycerate ADP
Phosphoglycerate Kinase
0~ C-0- 1 H-C-0H ATP I H-C-0 0 I '~ H /P'-. -0 0-
3-Phosphoglycerate ATP
Figure 1.4. Mechanism of PGK reaction in glycolysis.
Like other enzymes within the kinase family, PGK has an absolute requirement for divalent metal ions for glycolytic activity. Magnesium ions
(Mg2•) fulfil this role. Other divalent metal ions such as Mn2•, Ca2•, and Co2• can also replace Mg2• to a varying degree. Furthermore, various anions, such as sulfate, have been observed to affect PGK activity. In the low milimolar range, sulfate tends to increase Vmax as well as Km for both substrates, but at higher concentrations it has an inhibitory effect.
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1.4.3 Structure of PGK
The primary amino acid sequence of PGK isolated from various species has been determined. Structural comparison of these sequences indicates that PGK from different origins share a high degree of sequence homology and their catalytic modes are also similar.
1 Ser-Leu-Ser-Asn-Lys-Leu-Thr-Leu-Asp-Lys-Leu-Asp-Val-Lys-Gly-Lys-Arg-Val-Val-Met- 21 Arg-Val-Asp-Phe-Asn-Val-Pro-Met-Lys-Asn-Asn-Gln-lle-Thr-Asn-Asn-Gln-Arg-Lys-lle- 41 Lys-Ala-Ala-Val-Pro-Ser-lle-Lys-Phe-Cys-Leu-Asp-Asn-Gly-Ala-Lys-Ser-Val-Val-Leu- 61 Met-Ser-His-Leu-Gly-Arg-Pro-Asp-Gly-Val-Pro-Met-Pro-Asp-Lys-Tyr-Ser-Leu-Glu-Pro- 81 Val-Ala-Val-Glu-Leu-Lys-Ser-Leu-Leu-Gly-Lys-Asp-Val-Leu-Phe-Leu-Lys-Asp-Cys-Val- 101 Gly-Pro-Glu-Val-Glu-Lys-Ala-Cys-Ala-Asn-Pro-Ala-Ala-Gly-Ser-Val-l le-Leu-Leu-Glu- 121 Asn-Leu-Arg-Phe-His-Val-Glu-Glu-Glu-Gly-Lys-Gly-Lys-Asp-Ala-Ser-Gly-Asn-Lys-Val- 141 Lys-Ala-Glu-Pro-Ala-Lys-l le-Glu-Ala-Phe-Arg-Ala-Ser-Leu-Ser-Lys-Leu-Gly-Asp-Val- 161 Tyr-Val-Asn-Asp-Ala-Phe-Gly-Thr-Ala-His-Arg-Ala-His-Ser-Ser-Met-Val-Gly-Val-Asn- 181 Leu-Pro-Gln-Lys-Ala-Gly-Gly-Phe-Leu-Met-Lys-Lys-Glu-Leu-Asn-Tyr-Phe-Ala-Lys-Ala- 201 Leu-Glu-Ser-Pro-Glu-Arg-Pro-Phe-Leu-Ala-l le-Leu-Gly-Gly-Ala-Lys-Val-Ala-Asp-Lys- 221 lle-Gln-Leu-lle-Asn-Asn-Met-Leu-Asp-Lys-Val-Asn-Glu-Met-lle-lle-Gly-Gly-Gly-Met- 241 Ala-Phe-Thr-Phe-Leu-Lys-Val-Leu-Asn-Asn-Met-Glu-l le-Gly-Thr-Ser-Leu-Phe-Asp-Glu- 261 Glu-Gly-Ala-Lys-lle-Val-Lys-Asp-Leu-Met-Ser-Lys-Ala-Glu-Lys-Asn-Gly-Val-Lys-lle- 281 Thr-Leu-Pro-Val-Asp-Phe-Val-Thr-Ala-Asp-Lys-Phe-Asp-Glu-Asn-Ala-Lys-Thr-Gly-Gln- 301 Ala-Thr-Val-Ala-Ser-Gly-lle-Pro-Ala-Gly-Trp-Met-Gly-Leu-Asp-Cys-Gly-Pro-Glu-Ser- 321 Ser-Lys-Lys-Tyr-Ala-Glu-Ala-Val-Thr-Arg-Ala-Lys-Gln-lle-Val-Trp-Asn-Gly-Pro-Val- 341 Gly-Val-Phe-Glu-Trp-Glu-Ala-Phe-Ala-Arg-Gly-Thr-Lys-Ala-Leu-Met-Asp-Glu-Val-Val- 361 Lys-Ala-Thr-Ser-Arg-Gly-Cys-lle-Thr-lle-lle-Gly-Gly-Gly-Asp-Thr-Ala-Thr-Cys-Cys- 381 Ala-Lys-T rp-Asn-Thr-Glu-Asp-Lys-Val-Ser-H is-Val-Ser-Thr-Gly-Gly-Gly-Ala-Ser-Leu- 401 Glu-Leu-Leu-Glu-Gly-Lys-Val-Leu-Pro-Gly-Val-Asp-Ala-Leu-Ser-Asn-lle
Figure 1.5. Complete amino acid sequence of human PGK.
On the basis of this observation, it was hypothesised that the tertiary structure and active site regions of PGK have been well conserved during evolution.
There are seven cysteine residues in mammalian PGKs. Studies have reported that two of these cysteines contain reactive thiol groups,
53 Chapter 1 - Literature Review
whereas the remaining five react at a much slower rate with 5,5'-dithiobis (2- nitrobenzoic acid) (Nbs2). In horse muscle PGK, the two fast-reacting thiols correspond to the two closely spaced cysteines Cys378 and Cys379 located in helix 13 around the hinge region [174]. The same arrangement of thiols is found in pig, mouse and human PGK [175] (Figure 1.5). None of the seven thiols form disulfide bonds in the fully active enzyme or are essential for its enzyme activity. Yeast PGK, on the other hand, has only a single cysteine residue at position Cys99.
Until recently, PGK was always known as a monomeric enzyme consisting of 417 amino acid residues with a molecular mass of 42-45,000
Da, as determined by SOS-PAGE. However, dimeric and tetrameric PGK have been identified in archae Pyrococcus woesei and Su/pho/obus so/fataricus, respectively [176, 177].
1.4.3.1 The 3-Dimensional structure of PGK
PGK from horse [178], yeast [179], Bacillus stearothermophilus [180], and pig muscle [175], have a very similar three-dimensional structure. The structure of PGK as determined by X-ray crystallography is best described as an open bilobal molecule with N-acetylserine at the NH2-terminus and isoleucine at its COOH-terminus. These two structural units, termed N- and
C- terminal domains, represent the two halves of the polypeptide chain, which are folded in a/~ structures. Each consists of a central ~-sheet of six parallel strands that is shielded from solvents by helices. In total there are 15 helices, 12 internal ~-strands and at least five surface ~-strands in horse
PGK molecule [178].
54 Figure 1.6. Three-dimensional structure of human PGK. This structure represents the open conformation of the enzyme. The polypeptide chain is organised into two continuous structural domains (N- and C-domains). a-helices are represented by cylinders and ~- strands by arrows showing their direction. The C-domain contains the ATP binding C.11 C.11 site. The 3-PG binding site is thought to be situated on the interdomain face of the N-domain. The predicted hinge region is circled. This structure was constructed using SWISS-MODEL PDB file [181, 182). The image was generated using Weblab Viewerlite software (Molecular simulation). Chapter 1 - Literature Review
The two domains in PGK are linked by a highly-conserved region, known as the hinge region shown in Figure 1.6. This region is in the junction between ~-strand F and helix 7. The active site of PGK is located deep in the cleft formed between the two domains. In addition to the hinge region, a minor link of the two domains has also been proposed. Part of this structure is represented by residues 404-406, which connect helices 14 and 15.
Substrate binding studies have shown that Mg-ATP and Mg-ADP are bound on the inner surface of the C-domain in the region above the cleft (the
Rossmann fold), which is predicted to be approximately 10 A from the binding site of phosphoglycerate substrate on the NH2-terminal domain. To date, the precise binding site for 1,3-BPG is unknown. Direct determination of 1,3-BPG binding has been difficult due to the presence of a sulfate ion in
PGK crystals grown from ammonium sulfate solution. It is known that sulfate ions interfere with 1,3-BPG binding.
Despite the lack of consistency in the literature, a number of studies suggest that 1,3-BPG binds to the NH2-terminal domain of the enzyme. This binding is thought to be via a network of hydrogen bonds to a cluster of basic amino acid residues, and by electrostatic interactions between the negatively charged phosphate and positively charged arginines [175]. The distance between these bound substrates was estimated to be between 10-12 A for the open form of PGK.
1.4.4 Domain movement in PGK
Based on studies of the tertiary structure of PGK, it was thought that the binding locations of the two substrates are too far apart to favour direct
56 Chapter 1 - Literature Review
phosphoryl transfer during glycolysis (10-12 A). This observation led to the proposal that following substrate binding to the enzyme, the two domains undergo conformational changes, involving complete hinge bending to generate a closed form of PGK. This synergistic combination of 3-PGA and
Mg-ADP-induced conformational changes serve to bring the two ligands, 3-
PG and ADP, into close proximity in a water free region, creating a favourable environment for catalysis. A similar hinge bending mechanism has also been documented for hexokinase [183], pyruvate kinase and phosphofructokinase [184].
1.4.5 PGK isoenzyme
There are two PGK isoforms in mammalian species, encoded by the
PGK1 and PGK2 genes. Although the two genes produce protein, which are essentially similar in both structure and function, their expression is regulated differently. PGK1 is X-chromosome-linked and expressed in all somatic cells, oogenic cells, and premeiotic spermatogenic cells, and contains 1 O introns [185]. The PGK2 is autosomal and is expressed only in meiotic and postmeiotic spermatogenic cells and contains no introns [186].
Both enzymes contain the same number of amino acid residues, but differ in 53 sites along the polypeptide chain. These locations correspond with non-conserved positions in the total PGK sequence data and do not affect its glycolytic function.
57 Chapter 1 - Literature Review
1.4.6 Phosphoglycerate kinase in diseases
PGK is essential for the regulation of cell energy metabolism. It is also responsible for the maintenance of many physiological functions within cells.
Deficiency in PGK has been shown to be associated with a number of metabolic as well as neurological disorders such as chronic hemolytic anemia and mental disorder in human.
The expression of PGK mRNA in tumours is elevated perhaps due to the hypoxia environment of a tumour. This increase in the production is an adaptive response by tumour cells to sustain ATP production for their growth and development. Recently, an inducible transcriptional complex, hypoxia inducible factor 1 (HIF-1) was identified. It was postulated that HIF-1 mediates transcriptional responses to low cellular oxygen by binding to hypoxia-response elements (HREs) of target genes. HIF-1 is a heterodimer consisting of the constitutively expressed aryl hydrocarbon receptor nuclear translocator (ARNT) and the HIF-1a subunit. Although the precise cellular sensoring mechanism for hypoxia is unknown, it was demonstrated that HIF-
1 activation depends primarily on redox-sensitive stabilisation of its a subunit,
HIF-1a. HIF-1a is required for the basal expression of both PGK and LDH.
58 CHAPTER2 HYPOTHESIS AND AIMS Hypothesis and Aims
2.1 HYPOTHESIS AND AIMS
Angiogenesis is a complex process, which plays a critical role in a variety of normal physiological events such as the female reproductive system and tissue repair. Abnormal angiogenesis has also been implicated in a number of pathological conditions such as in arthritis, diabetic retinopathy and tumour growth.
It is now well established that tumour expansion is angiogenic dependent. The growth of solid tumours beyond 1-2 mm in diameter requires new blood vessel formation. In fact, tumour angiogenesis appears to be one of the crucial steps necessary for tumour progression. The degree of vascularisation of many solid tumours correlates with a poor prognosis and an increased risk of metastatic diseases. Thus the idea of targeting angiogenesis offers a novel and practical approach for long term control of the disease.
Convincing evidence has been generated to suggest that angiogenesis is regulated by a balance between several protein activators and inhibitors. Angiostatin is one such inhibitor produced by tumour cells.
The mechanism of angiostatin formation from plasmin is not well understood.
A number of tumour-associated proteins have been shown to play a role in angiostatin formation. Taken together, these experimental reports suggest that the generation of angiostatin from plasmin is the result of combined action of serine proteinases and metalloproteinases. We suggested that plasmin proteolysis occurs in three stages. First, one or more disulfide bonds in kringle 5 of plasmin were reduced by a plasmin reductase secreted by cultured HT1080 cells. Second, reduction of these disulfide bonds in kringle
59 Hypothesis and Aims
5 triggers cleavage at Arg529-Lys530 in kringle 5, and also at two other positions C-terminal of Cys461, by a serine proteinase [187]. Third, the kringle 1-4½ fragments are cleaved by matrix metalloproteinases to produce either kringle 1-4 or 1-3 [145, 149, 188].
I hypothesised that the reduction of plasmin by plasmin reductase is the rate limiting step in the formation of angiostatin by transformed cells. The aims of this study were:
(1) To characterise the plasmin reductase activity secreted by various
cultured transformed cells,
(2) To isolate and clone plasmin reductase secreted by cultured human
fibrosarcoma cells or the HT1080,
(3) To examine the plasma levels of plasmin reductase in tumour bearing
mice and investigate the effect of systemic administration of recombinant
plasmin reductase on plasma levels of angiostatin, tumour angiogenesis
and tumour growth,
(4) To define the mechanism of action of plasmin reductase.
60 CHAPTER3 PLASMIN REDUCTASE CHARACTERISATION Chapter 3 - Characterisation of Plasmin Reductase
3.1 INTRODUCTION
Tumour expansion and metastasis is dependent on tumour neovascularisation, or angiogenesis [33, 189]. Angiogenesis is balanced by several protein activators and inhibitors [33]. One such inhibitor is angiostatin [46], an internal fragment of the plasma zymogen, plasminogen.
Plasminogen contains five consecutive kringle domains and a serine proteinase module. uPA or tPA converts plasminogen to plasmin by
hydrolysis of the Arg560-Val561 peptide bond in the serine proteinase module. It has been shown that plasmin can be processed in CM of tumour cells to produce angiostatin fragments consisting of kringle domains 1 to 4
and approximately half of kringle 5 [187, 190, 191]. Stathakis and co workers demonstrated that the generation of angiostatin from plasmin
involves reduction of at least two disulfide bonds in kringle 5 of plasmin by
plasmin reductase secreted by HT1080 cells [187]. In addition, they showed that reduction of these disulfide bonds in K5 of plasmin triggers proteolytic
attack at Arg529-Lys530 and at two other positions, C-terminal of Cys461by
a serine proteinase [187].
The aim of this study was to characterise plasmin reductase secreted by HT1080 cells. This chapter describes techniques I developed for measuring reduction of disulfide bonds in plasmin. A modified ELISA has been designed to quantitate plasmin reductase activity by .measuring the formation of free thiols in angiostatin. Preliminary studies on the characterisation of plasmin reductase will also be described
61 Chapter 3 - Characterisation of Plasmin Reductase
3.2 MATERIALS AND METHODS
3.2.1 Cell culture
Human foreskin fibroblasts (HFF) [192], blood monocytes/macrophages [193], human umbilical vein endothelial cells
(HUVEC) [194], human dermal microvascular endothelial cells (HDMVEC)
[195], bovine aortic endothelial cells (BAEC) [196] and rat vascular smooth muscle cells (RVSMC) [197] were harvested and cultured as previously described. Bovine vascular smooth muscle cells (BVSMC) were purchased from Cell Applications (San Diego, CA). Human fibrosarcoma (HT1080),
Chinese hamster ovary (CHO-K1), breast carcinoma cells (MCF-7, BT20,
MDA-231) and THP-1 cells were purchased from American Type Cell Culture
(Rockville, MD). All media components were from Gibco BRL Life
Technologies (Grand Island, NY). Cell culture flasks (25 and 75 cm2) were purchased from Falcon (Franklin Lakes, NJ). Flat-bottomed tissue culture plates (96 well) were obtained from Corning (Corning, NY).
3.2.1.1 HT1080 cell culture
HT1080 cells were maintained in DMEM containing 10% (v/v) fetal calf serum (FCS), 10 units/ml penicillin streptomycin and 2 mM L-glutamine, at a pH of 7.2 in a 5% CO2 humidified atmosphere at 37°C. A total of 1x106 cells was suspended in 15 ml of complete DMEM and plated onto a 75cm2 flask.
Confluent cells were passaged 1:4 and re-seeded twice a week.
62 Chapter 3 - Characterisation of Plasmin Reductase
3.2.1.2 Breast carcinoma cell culture
MCF-7, BT20 and MDA-231 cells were maintained in RPMI 1640 containing 20 mM HEP ES, 10% (v/v) FCS, 10 units/ml penicillin streptomycin and 2 mM L-glutamine, 400 units of insulin, pH 7.2 in a 5% CO2 humidified atmosphere at 37°C. A total of 1x106 cells was suspended in 15 ml of complete RPMI 1640 and plated onto a 75cm2 flask. Confluent cells were passaged 1 :4 and re-seeded twice a week.
3.2.1.3 Preparation of conditioned medium (CM)
Various cultured cells at -80% confluence were washed three times with warm phosphate buffered saline (PBS). CM was collected by incubating cells with HBSS containing 25 mM HEPES at a pH of 7.2 for 24 hours. The ratio of number of cells to volume of CM was 1-3 x 106 cells per ml of media.
The CM was centrifuged at 1200 rpm for five minutes to remove cell debris and subsequently sterilised using a 0.22 µm filter prior to storage at -80°C.
3.2.2 Assay for plasmin reductase activity
Plasmin reduction was measured after formation of free thiols in angiostatin using an avidin-biotin interaction in a microtitre plate or
Streptavidin-HRP blot. Briefly, plasm in was incubated with HT1080 CM and the resulting angiostatin fragments labelled with the biotin-linked maleimide,
3-(N-maleimidylpropionyl) biocytin (MPB) [187, 191 ]. Maleimides reacts specifically and rapidly [198), second order rate constant of -104 M·1s·1 at pH
8, with free thiols. The MPS-labelled angiostatin fragments were collected on lysine-Sepharose and resolved in one of two ways. The labelled fragments
63 Chapter 3 - Characterisation of Plasmin Reductase
were separated on SDS-polyacrylamide gel electrophoresis (SOS-PAGE) and transferred to a polyvinylidene difluoride (PVDF) membrane or captured by an anti-angiostatin monoclonal antibody coated on microtitre plate wells.
The incorporated MPB was measured by incubation with peroxidase conjugated streptavidin and a suitable substrate, such as 2,2'-azino-bis (3- ethylbenzthiazoline-6-sulfonic acid) (ABTS).
3.2.2.1 Angiostatin generation from HT1080 CM
uPA used in this study was a gift from Serono (Australia). MPB were purchased from Molecular Probes Inc (Eugene, OR). Lysine-Sepharose was from Pharmacia (Uppsala, Sweden). Reduced glutathione, N-ethylmaleimide and E-amino-caproic acid were from Sigma Chemical (St Louis, MO).
Plasminogen was purified from frozen human plasma using lysine sepharose according to published procedures [81]. Plasmin was generated by incubating plasminogen (20 µM) and uPA (20 nM) in reaction buffer containing 20 mM HEPES, 0.14 M NaCl, 1 mM EDTA, 0.05% (v/v) Tween, pH 7.4 for 30 minutes at 37°C. 500 µI of HT1080 CM was incubated with plasmin (2-10 µg per ml) for 30 minutes at 37°C. Angiostatin fragments were labelled with MPB (100 µM) for 30 minutes at room temperature, followed by quenching of the unreacted MPB with reduced glutathione (GSH) (200 µM) for 10 minutes at room temperature. Unreacted GSH and other free sulfhydryls were blocked with N-ethylmaleimide (400 µM) for 10 minutes at room temperature. The MPS-labelled plasmin kringle products were collected on 50 µI of lysine-sepharose beads by incubation on a rotating wheel for one hour at room temperature. The beads were washed three
64 Chapter 3 - Characterisation of Plasmin Reductase
times with 1 ml of buffer and the plasmin kringle products eluted with 50 µI of reaction buffer containing 50 mM EACA.
3.2.2.2 MPB blot assay
All pre-cast 8-16% SOS-PAGE gels were purchased from Gradipore
(Sydney, Australia). PVOF membrane was obtained from Millipore (Bedford,
MA). Streptavidin horseradish peroxidase was from Amersham (Sydney,
Australia) and NEM Life Science (Boston, MA) provided chemiluminescence.
MPS-labelled angiostatin fragments were resolved on 8-16% SOS-PAGE under non-reducing conditions [199), transferred to a PVOF membrane, blotted with a 1:2000 dilution of streptavidin horseradish peroxidase, then developed and visualised using enhanced chemiluminescence according to the manufacturer's instructions.
3.2.2.3 Microtitre plate assay
96-well plates were purchased from Nunc PolySorp (Nalgene Nunc
International, Naperville, IL). StreptABComplex/HRP was from Oako
(Carpinteria, CA) and ABTS were from Sigma Chemical (St Louis, MO).
MPS-labelled angiostatin fragments were immobilised on wells coated with a murine anti-angiostatin monoclonal antibody designated 8.19. The antibody was generated against human angiostatin [200), which binds with equal affinity to soluble angiostatin and plasmin but has an approximately 10-fold lower affinity to plasminogen. 100 µI of 8.19 diluted in 0.1 M NaHC03, 0.02%
(w/v) NaN3, pH 8.3 (5 µg per ml) was adsorbed to a 96 well plate overnight at
4°C in a humid environment. The wells were washed twice with reaction
65 Chapter 3 - Characterisation of Plasmin Reductase
buffer. Any non-specific binding sites were blocked by adding 200 µI of blocking buffer containing 20 mM HEPES, 0.14 M NaCl, 1 mM EDTA, 2%
(w/v) bovine serum albumin, 0.02% (w/v) NaN3, pH 7.4 and incubated at
37°C for 90 minutes. The washing step was repeated. MPS-labelled angiostatin fragments were diluted 1: 10 in reaction buffer and 100 µI aliquots were added to antibody coated wells and incubated for 30 minutes at room temperature with orbital shaking. The wells were then washed three times with reaction buffer. Subsequently, 100 µI of 1:100 StreptABComplex/HRP in reaction buffer were added and incubated for 30 minutes at room temperature with orbital shaking. Wells were washed three times with reaction buffer and colour developed with 100 µI of 0.003% H20 2, 1 mg/ml
ABTS in substrate buffer (50 mM citrate, pH 4.5) for 10 minutes at room temperature with orbital shaking. Absorbances were determined at 405 nm.
Results were corrected for control reactions containing no plasmin or no
HT1080 CM.
3.2.3 Endogenous glutathione in HT1080 CM
HT1080 cells secreted the most plasmin reductase activity in 24 hours. This cell line, therefore was used to characterise plasmin reductase.
To confirm that the plasmin reductase activity in HT1080 CM was not due to the presence of low molecular weight thiols, the level of thiol components in the CM was measured using the fluorescent compound, 7-benzo-2-oxa-1,3- diazole-4-sulfonic acid followed by HPLC analysis as described by Dudman and co-workers [201].
66 Chapter 3 - Characterisation of Plasmin Reductase
3.2.4 Competitive inhibition of plasmin reductase activity by NADH
At this point in our investigation, the activity of plasmin reductase appeared to resemble the activity of a NAOH oxidase. On this basis I examined the effect of NAOH on plasmin reductase activity in HT1080 CM.
One ml of CM was incubated with 1 mM NAOH at room temperature for one hour. Angiostatin generated by incubation with plasmin (10 µg/ml) was determined following MPB labelling as previously described.
3.2.4.1 Binding of plasmin reductase to Cibachron Blue-Sepharose
The finding showed that NAOH competitively inhibited plasmin reductase activity in the HT1080 CM. This implied that plasmin reductase interacted with NAOH and I hypothesised that plasmin reductase would bind to Cibachron Blue-Sepharose due to the structural similarity between NAOH and Cibachron Blue. HT1080 CM (5 ml) was applied to a 1O ml Cibachron
Blue-Sepharose column equilibrated with 20 mM HEPES, 0.05 M NaCl, 1 mM EOTA, pH 7.4. Bound proteins were eluted with HEPES buffer containing 2 mM NAO followed by buffer containing 1 M NaCl. Eluates were assayed for plasmin reductase activity using ELISA and Streptavidin-HRP blot methods.
3.2.5 Alkylation of thiol groups in plasm in reductase
Protein disulfide isomerase (POi) and thioredoxin (TR) are secreted by cultured cells and participate in the redox reaction of disulfide bonds. Both enzymes can cleave plasminogen to form angiostatin, however neither was responsible for the plasmin reductase activity observed in HT1080 CM [187).
67 Chapter 3 - Characterisation of Plasmin Reductase
In view of this, plasmin reductase was believed to share similar features to
POi or TR.
The effect of thiol modifications of HT1080 CM plasmin reductase was examined. Thiols in plasmin reductase were alkylated by reaction with N ethylmaleimide or iodoacetamide. Five mM of either N-ethylmaleimide or iodoacetamide was added to HT1080 CM and incubated for 14 hours at room temperature. Treated CM was dialysed twice against one litre of buffer containing 20 mM HEPES, 0.14 M NaCl, 1 mM EDTA, pH 7.4 and plasmin reductase activity was determined as previously described.
3.2.5.1 Binding of plasmin reductase to thiol activated sepharose
I examined the binding of plasmin reductase to Thiopropyl-Sepharose.
HT1080 CM ( 1 ml) was added to 200 µI of 1: 1 slurry of Thiopropyl-Sepharose resin and incubated at room temperature for one hour. The supernatant was collected via centrifugation and assayed for plasmin reductase activity. To ensure that the loss of plasmin reductase activity in the CM was not due to the removal of low molecular weight thiols, the endogenous GSH was reconstituted by adding 2 µM GSH to one of the supernatants before assaying the plasmin reductase activity.
68 Chapter 3 - Characterisation of Plasmin Reductase
3.3 RESULTS
3.3.1 ELISA for determining plasmin reductase activity
Plasmin reductase activity in tumour cell CM was measured by formation of free thiols in angiostatin using an avidin-biotin interaction in a microtitre plate format. The MPS-labelled angiostatin fragments were immobilized on wells coated with a murine anti-angiostatin monoclonal antibody and detected using StreptABComplex/HRP. This assay was specific for plasmin reductase activity, since there was no detectable signal in the control without HT1080 CM (Figure 3.1 A). To determine the specificity of the assay, plasmin (2 µg/ml) was incubated in either buffer containing 20 mM HEPES, 0.14 M NaCl, 1 mM EDTA and pH 7.4 or HT1080 CM for 30 minutes at 37°C and the angiostatin fragments labelled with MPB (Figure 3.1
A solid bars). For controls, HT1080 CM was incubated without plasmin or with plasmin but without MPB labelling (Figure 3.1 A open bars).
There was a linear relationship between the absorbance at 405 nm and the concentration of HT1080 CM after a threshold concentration of plasmin reductase was achieved (Figure 3.1 B). Plasmin (2 µg/ml) was incubated in dilutions of HT1080 CM in 20 mM HEPES, 0.14 M NaCl, 1 mM
EDTA pH 7.4 buffer for 30 minutes at 37°C. The angiostatin fragments were labelled with MPB and detected as described. The data was corrected for background absorbance of plasmin in HEPES buffer alone.
69 Chapter 3 - Characterisation of Plasmin Reductase
A B 1.4 1.2
1.2 E 1.0 E C C an 1.0 II) i i 0.8 iv 0.8 Cl) CJ B o.6 C C _! 0.6 .! .. ~ 0.4 0 _! 0.4 c( ! 0.2 0.2 o.o 0.0 • 0.0 0.2 0.4 0.6 0.8 1.0 Dilution of HT1080cm
Figure 3.1. ELISA for MPB-labelled angiostatin fragments. (A) Plasmin (2 µg/ml) was incubated with either HEPES buffer for negative control or HT1080 CM (solid bars). Controls were CM in the absence of plasmin and CM/plasmin reaction without MPB labelling (open bars). Results are the mean and SEM of triplicate determinations. (B) Dotted line represents linear least squares fit to the three right hand data points (r2 = 0.99).
70 Chapter 3 - Characterisation of Plasmin Red uctase
3.3.2 Streptavidin-HRP blot for MPS-labelled angiostatin
MPS-labelled angiostatin fragments were detected by blotting with
Streptavidin-HRP. HT1080 CM (1 ml) was incubated with plasm in (10 µg) followed by MPS labelling of angiostatin fragments as previously described.
MPS-labelled angiostatin fragments were separated on 8-16% SOS-PAGE under non-reducing conditions, transferred to PVDF membrane and blotted with Streptavidin-HRP then developed and visualized using chemiluminescence. Angiostatin fragments are represented by 3 bands,
(molecular mass - 38, -43 and -48,000 Da) (Figure 3.2).
8
6
43
29
20
1 2
Figure 3.2. MPS-labelled angiostatin fragments blotted with Streptavidin-HRP. Plasmin incubated with HEPES buffer (lane 1), plasmin incubated with 1 ml of HT1080 CM (lane 2). The positions of Mr markers in kDa are shown at left. The arrows indicate the three angiostatin fragments.
71 Chapter 3 - Characterisation of Plasmin Reductase
3.3.3 Plasmin reductase activity secreted by various cultured cells
Primary cells (HFF, macrophages, HUVEC, HDMVEC, and BAEC) did not secrete detectable plasmin reductase except RVSMC and BVSMC, which secreted low levels of plasmin reductase. In contrast, all transformed cells tested (HT1080, CHO, MCF-7, BT20, and MDA-231) secrete plasmin reductase. The level of plasmin reductase activity in CM of transformed cells varied significantly. The reductase activity of CHO and MCF-7 was approximately 60% of HT1080 cells, while BT20 cells and MDA-231 cells were 27% and 20% respectively. Figure 3.3 shows relative plasmin reductase activities.
0.5
E C C 0.4 It) 0..,. Cl) g 0.3 .0"'... 0 Cl) 0.2 .0 c( 0.1 0.0 -----·· II_ u. G) Sil O "':" u. Ii> c5 :c u. :c .c .- 0 0 c. !i: :I
:Ii
Figure 3.3. Plasmin reductase activity secreted by selected primary and transformed cells. All CM were generated by incubating cells at -80% confluence with HBSS for 24 hours. CM was assayed for plasmin reductase activity using ELISA as described previously. Results are mean and SEM of triplicate determinations.
72 Chapter 3 - Characterisation of Plasmin Reductase
3.3.4 Effect of NADH on plasmin reductase activity
NADH competitively inhibited plasmin reductase function. Incubation of HT1080 CM with 1 mM NADH for one hour at room temperature caused a
95% loss of plasmin reductase activity as indicated by a significant reduction in the formation of angiostatin fragments (Figure 3.4).
68- 43- .. 29- 20-
16-
1 2 3
Figure 3.4. NADH competitively inhibited plasmin reductase activity. Streptavidin-HRP blot analysis of plasm in incubated with HEP ES buffer (lane 1) ; HT1080 CM (lane 2) and HT1080 CM treated with 1 mM NADH. The positions of Mr markers in kDa are shown at left. The arrows indicate the three angiostatin fragments.
73 Chapter 3 - Characterisation of Plasmin Reductase
3.3.4.1 Plasmin reductase bound Cibachron Blue-Sepharose
The finding that plasmin reductase activity can be competitively
blocked by NADH suggested that plasmin reductase interacted with
Cibachron Blue-sepharose. The majority of plasmin reductase was displaced with either 2 mM NAO or 1 M NaCl. No plasmin reductase activity was detected in the flow through of the Cibachron Blue chromatography (Figure
3.5 A and B).
A
0.15
0.10
0.05
B o.oo
HT1080 CM Unbound NAD NaCl
68-
43- 29- 20-
16-
1 2 3 4
Figure 3.5. MPS-labelled angiostatin fragments after Cibachron Blue affinity chromatography. (A) ELISA, (8) Streptavidin-HRP blot of starting material-HT1080 CM (lane 1) , fraction containing unbound proteins (lane 2) and bound proteins displaced by 2 mM NAO or 1 M NaCl (lanes 3 and 4, respectively).
74 Chapter 3 - Characterisation of Plasmin Reductase
3.3.5 Thiol modifying reagents reduced plasmin reductase activity
HT1080 CM (500µ1) was treated with 5 mM of NEM or 1AM to block thiols in plasmin reductase. Alkylation of thiol groups in plasmin reductase caused 50-70% reduction in plasmin reductase activity as determined by
ELISA (Figure 3.6).
0.5
-E 0.4 C: II) 0 "'llt' 0.3 -G) CJ C: ea .Cl... 0.2 0 ,n .Cl 0.0 Figure 3.6. Effect of thiol-modifying reagents on plasmin reductase activity. Reduced angiostatin formation was observed after incubation of HT1080 CM with N ethylmaleimide or iodoacetamide compared to untreated HT1080 CM. 75 Chapter 3 - Characterisation of Plasmin Reductase 3.3.6 Plasmin reductase bound to thiol activated sepharose As thiol-modifying reagents were shown to inhibit plasmin reductase activity, I examined the binding of plasmin reductase to Thiopropyl Sepharose. Incubation of HT1080 CM with Thiopropyl-Sepharose caused a marked reduction in plasmin reductase activity. Addition of 2 µM GSH to the CM after incubation with Thiopropyl-Sepharose did not restore plasmin reductase activity (Figure 3.7). 0.5 -E 0.4 C II) 0 ~ 0.3 uCD C C'I -e 0.2 0 rn .0 c( 0.1 0.0 - Figure 3.7. Plasmin reductase activity after incubation with Thiopropyl Sepharose gel. HT1080 CM was batch absorbed to Thiopropyl-Sepharose and the supernatant was assayed for plasmin reductase activity in the presence or absence of 2 µM reduced glutathione (GSH). 76 Chapter 3 - Characterisation of Plasmin Reductase 3.4 DISCUSSION The formation of angiostatin from plasmin in CM of transformed cells is a three step process [191]. First, one or more disulfide bonds in plasmin are reduced by a protein disulfide bond reductase, which we have called plasmin reductase. The second step involves proteolytic cleavage of the reduced plasmin by a serine proteinase to form angiostatin. Three angiostatin fragments are produced with apparent molecular masses of -38, -43, and -48,000 Da on non-reducing SOS-PAGE. Angiostatin fragments contain free thiols as a result of disulfide bond reduction in plasmin. These free thiols may be labelled with the biotin linked thiol specific compound MPB and detected using ELISA or Streptavidin-HRP blot analysis. Traditionally, protein thiols are measured after derivitization with reagents such as 5,5'-dithiobis (2-nitrobenzoic acid) (DTNB). However, this method is limited to reactions containing micromolar concentrations of thiols. This necessitated the development of the novel ELISA used to measure the activity of plasmin reductase secreted by tumour cells in culture, which is in the nanomolar range. This assay did not provide an absolute measure of angiostatin formation. A determining factor is the accessibility of MPB for the protein thiols. The proprionyl spacer arm that links biotin to the maleimide moiety makes the maleimide more accessible to partly buried protein thiols. It is possible, however, that a fraction of free thiols on angiostatin will be inaccessible and will not be efficiently labelled with MPB. It is also possible that dithiols may rapidly oxidise to form inter or intra molecular disulfide bonds. Even so, this assay provides a relative measure of plasmin reductase activity. The latter possibility was tested by excluding oxygen radicals formed 77 Chapter 3 - Characterisation of Plasmin Reductase metal ions in solution by adding a metal chelator and anti-oxidant to the buffer. This will slow or prevent the oxidation process. No significant difference in MPB labelling free thiols was observed in the presence or absence of 1mM ascorbic acid and 1 mM EDTA, suggesting that oxidation of free thiols in angiostatin is not a significant event in our reaction. Quantitation of plasmin reductase secretion by various cultured cells indicated that all transformed cell lines tested secrete plasmin reductase. The level of plasmin reductase secretion varied considerably among cells, with HT1080 secreting the highest level of plasmin reductase activity after 24 hours in serum free media. In contrast, primary foreskin fibroblasts, macrophages and endothelial cells do not secrete detectable levels of plasmin reductase. Interestingly, both cultured rat and bovine vascular smooth muscle cells secrete low levels of plasmin reductase. Cellular transformation appeared to be associated with the secretion of plasmin reductase and angiostatin formation. This suggested that angiostatin formation is driven by tumour cells in vivo. Production of angiostatin by VSMC is an interesting observation and suggested that angiostatin may function in the atherosclerotic vessel wall. O'Reilly and subsequent investigators have shown that angiostatin formation was driven by tumour cells [46, 145, 147, 148, 190, 202]. They speculate that the cleavage of plasminogen to form angiostatin involves serine proteinase activity, presumably secreted by tumour cells. Among those enzymes shown to play a role in the generation of angiostatin are the MMP's [145, 147-149] and serine proteinases [187, 190]. The fact that 78 Chapter 3 - Characterisation of Plasmin Reductase various enzymes can cleave plasminogen to form angiostatin suggests that different cell types may employ distinct pathways for angiostatin formation. Gateley and co-workers have shown that high concentrations (100 µM) of small thiols alone can generate angiostatin from plasmin [190], however these concentrations are not achievable in cell culture. Subsequently, Stathakis and co-workers demonstrated that 10 µM GSH had 6% of the angiostatin generating activity of HT1080 CM [187]. The concentration of GSH in our HT1080 CM was 0.82 µM and was the only small thiol detected in the medium. Taken together, these observations implied that plasmin reduction in HT1080 CM was catalysed by plasmin reductase and not by GSH directly. These results support our proposal that plasmin reductase is responsible for the angiostatin converting activity in HT1080 CM and that the activity was not due to the presence of high concentrations of free sulfhydryls. A further defining characteristic of plasmin reductase activity was its inhibition by both NADH and thiol modifying reagents. This implied that plasmin reductase would have an affinity for both Cibachron Blue-Sepharose and Thiolpropyl-Sepharose. Binding studies confirmed this observation. This formed the basis for the selection of chromatography matrices used in Chapter 4 for plasmin reductase purification. 79 CHAPTER4 PLASMIN REDUCTASE PURIFICATION Chapter 4 - Plasmin Reductase Purification 4.1 INTRODUCTION It has been shown that various transformed cell lines secrete plasmin reductases, initiating the formation of the antiangiogenic molecule, angiostatin [187]. As reviewed in Chapter 1, angiostatin is an internal fragment of plasminogen. Stathakis and co-workers demonstrated that plasmin reductase facilitated the reduction of Cys461-Cys540 and Cys511- Cys535 disulfide bonds in kringle 5 of plasmin. They also showed that cultured HT1080 cells secreted three serine proteinases where one or more of these proteins were responsible for cleaving the reduced plasmin to form angiostatin. This Chapter describes the isolation of plasmin reductase. Plasmin reductase was purified from HT1080 CM. HT1080 cells secreted plasmin reductase significantly more than the other transformed cells investigated, and therefore this cell line was selected for the isolation of plasmin reductase. 80 Chapter 4 - Plasmin Reductase Purification 4.2 MATERIALS AND METHODS 4.2.1 Large scale production of CM Twenty litres of HT1080 CM was prepared by culturing approximately 6x1010 HT1080 cells using a high yield culture system as described below. Cells were maintained and CM prepared, as described earlier in Chapter 3. 4.2.1.1 Cell factory Cells were cultured in cell factories (Nalgen Nunc International, Rochester, NY), according to the manufacturer's instructions. Each factory had a growth surface area of 6320 cm2 and produced 2 litres of CM. 4.2.2 Quantitative analysis of protein Total protein concentrations were determined using the Bicinchoninic acid (BCA) protein assay kit (Pierce Chemicals, Rockford, IL). BSA at concentrations ranging from 5-320 µg/ml was used to construct a standard curve. Unknowns and standards were assayed in 96 well microtitre plates. All procedures were performed according to the manufacturer's instructions. To minimize variation, all assays were incubated at 37°C for 30 minutes before measuring the absorbance at 562 nm on a microplate reader (SpectraMax, Molecular Devices). 81 Chapter 4 - Plasmin Reductase Purification 4.2.3 ELISA and Streptavidin-HRP blot for plasmin reductase activity Plasmin reduction was measured after formation of free thiols in angiostatin using the avidin-biotin interaction in a microtitre plate assay or Streptavidin-HRP blotting as described in Chapter 3. 4.2.4 Concentration of HT1080 CM Twenty litres of HT1080 CM was concentrated to a final volume of 350 ml using a spiral-wound cartridge concentrator (Amicon Division, W.R. Grace & Co., Beverly, MA). All procedures were performed at 4°C in the presence of a cocktail of protease inhibitors: leupeptin (10 µM), PMSF (1 mM), EDTA (2 mM) and soybean trypsin inhibitor (10 µg/ml). The concentrated CM was dialysed twice against 4 litres of 20 mM HEPES, 0.05 M NaCl, 0.02% (w/v) NaN3, 1 mM EDTA, pH 7.4 and stored at -80°C until used. 4.2.5 Blue-Sepharose CL-68 column chromatography The dialysed sample was applied to a Blue-Sepharose chromatography matrix (80 ml, 2.5 x 20 cm column) equilibrated with starting buffer containing 20 mM HEPES, 0.05 M NaCl, 1 mM EDTA, 0.02% (w/v) NaN3, pH 7.4. The sample was loaded at a flow rate of 1 ml/min and the matrix washed with three column volumes of starting buffer, until the baseline was reached. Bound proteins were eluted with a linear NaCl gradient of 0.05 M to 2 M at a flow rate of 0.5 ml/min. Fractions of 1.5 ml were collected and assayed for plasmin reductase activity. Fraction 78-89 contained plasmin reductase activity. These fractions were pooled (17 ml of 15 mg/ml) and 82 Chapter 4 - Plasmin Reductase Purification dialysed three times with 1 litre buffer containing 20 mM HEPES, 0.14 M NaCl, 1 mM EOTA, pH 7.4. 4.2.6 Thiopropyl-Sepharose 6B chromatography The dialysed sample was batch absorbed to 6 ml of activated Thiopropyl-Sepharose gel (Pharmacia, Uppsala, Sweden) in a capped tube and incubated at room temperature on a rotating wheel for one hour. The beads were packed into an 8 ml column (1 x 10 cm) at a flow rate of 0.6 ml/min. The packed gel was washed, to remove unbound and non specifically adsorbed proteins, with three column volumes of buffer containing 20 mM HEPES, 0.14 M NaCl, 1 mM EOTA, pH 7.4. The absorbance of flow through was monitored until the baseline was reached. The covalently bound proteins were sequentially eluted from the gel with buffer containing low-molecular weight thiols. They were used at a flow rate of 0.1 ml/min in the order of increasing reducing potential starting with 5 mM L-Cysteine, 50 mM GSH and finally 20 mM OTT. In each elution step, one void volume of reducing buffer was loaded into the column and left to react for 30 minutes at room temperature. Elution was continued with 10 ml HEPES buffer followed by washing with 20 ml of buffer. The absorbance of flow through was monitored until the baseline was reached before loading the next reducing buffer. Three fractions, each containing proteins displaced by L-Cysteine, GSH and OTT respectively, were dialysed extensively in 20 mM HEPES, 0.14 M NaCl, 1 mM EOTA, 0.02% (w/v) NaN3, pH 7.4 to remove all trace of the reducing agents before assaying for plasmin reductase activity. The 20 mM OTT eluate was concentrated to 2.4 ml in an Amicon 83 Chapter 4 - Plasmin Reductase Purification concentrator with PM-10 cutoff membrane (Amicon Division, W.R. Grace & Co., Beverly, MA). 4.2.7 Gel filtration (Sephacryl S-200 HR) A 120 ml Sephacryl S-200 HR gel was washed with buffer containing 20 mM HEPES, 0.14 M NaCl, 1 mM EDTA, 0.02% (w/v) NaN3, pH 7.4 and then packed into a 15 x 700 mm column at a flow rate of 0.5 ml/min. The dialysed sample from the activated Thiopropyl chromatography (2.4 ml) was run at a flow rate of 0.2 ml/min. The first 40 ml eluate was collected as one fraction, thereafter, 1ml fractions were collected. Three protein peaks were eluted from the column. Plasmin reductase activity was found in the final peak, which corresponded to fractions 37-44. These fractions were pooled and analysed on SOS-PAGE. 4.2.8 NH2-terminal sequence analysis The purified intact protein was found to be NH2-terminally blocked and was digested with cyanogen bromide (CNBr) followed by purification of the peptides. A 300 µI of sample, containing approximately 7 µg of protein, was concentrated to 10 µI using a speed-Vac and diluted to 50 µI with 88% formic acid. A small crystal of CNBr was added to the sample and the tube was flushed with nitrogen, capped, wrapped in aluminium foil and incubated in the dark at room temperature for 22 hours. The sample was then diluted with 500 µI of distilled H20 and concentrated to 50 µI using a speed-Vac. Fifty µI of CNBr digested fragments were diluted with 250 µI of milli-Q H20, filtered, and applied to a reverse-phase HPLC column (VYDAC C8, 84 Chapter 4 - Plasmin Reductase Purification 1x250 mm). The column was previously equilibrated with solvent A containing 0.1 % (v/v) TFA in H20 and solvent B containing 0.09% (v/v) TFA, 70% acetonitrile mixture in H20 and passed at a flow rate of 40 µI /min. A linear gradient of 0-50% acetonitrile was developed for the initial 100 minutes, followed by a linear gradient of 50-70% acetonitrile for a further 20 minutes. Fractions were collected and the peptides indicated were sequenced on a Hewlett-Packard G1005 A protein sequencing system. 85 Chapter 4 - Plasmin Reductase Purification 4.3 RESULTS 4.3.1 Cibachron Blue-Sepharose 48 chromatography In the binding study, it was shown that plasmin reductase had a strong affinity for Cibachron Blue-Sepharose. Thus, this chromatography was chosen for the first stage of purification. Twenty litres of HT1080 CM was concentrated (840 mg of protein) using a spiral-wound cartridge concentrator with a 10,000 molecular weight cut-off membrane. The sample was then applied to Cibachron Blue-Sepharose. Bound proteins were eluted from the column with a linear NaCl gradient between 0.05 and 2 M. Plasmin reductase activity eluted as a broad peak starting at approximately 1 M NaCl (Figure 4.1 ). The leading edge of this peak containing fractions between 1- 1.2 M NaCl was pooled. It contained 6 mg of total protein with a 120-fold increase in specific activity. 86 Chapter 4 - Plasmin Reductase Purification 90 / .5 I Plasmin Reductase / / .,, / -Protein / / Di' 80 / ---NaCl Gradient / tn / / / 3 / .4 70 / ::::, / / / / a 60 / c. C E .3 (') -C) 50 I» :i tn ~ - / CD C / / I» ·-Cl) 40 / (') / / .2 0 / / -... / -<' 30 / a. / / ~ / / / 20 / C / .1 ::::, / / / / i 10 / / / 0 .0 0 40 80 120 160 200 240 Elution volume, ml Figure 4.1. Purification of plasmin reductase on Cibachron Blue-Sepharose chromatography. CM from HT1080m cells was applied to Cibachron Blue Sepharose and the bound proteins were eluted with a linear gradient of 0.05-2 M NaCl, pH 7.4. Horizontal bar indicates pooled fractions. 87 Chapter 4 - Plasmin Reductase Purification 4.3.2 Thiopropyl-Sepharose 6B chromatography In addition to binding to Cibachron Blue-Sepharose, plasmin reductase also displayed affin.ity for thiol activated-Sepharose. The pooled sample was batch absorbed to activated Thiopropyl-Sepharose. The matrix was sequentially developed with buffers of increasing reducing potential to elute covalently bound proteins. The majority of plasmin reductase activity was eluted with 20 mM OTT (Figure 4.2). The protein yield was 2 mg with a 710-fold increase in specific activity. 0.20 -E C 0.15 II) 0... -uCD C 0.10 -eftl 0 en .c c( 0.05 0.00 Figure 4.2. Plasmin reductase activity eluted from Thiopropyl-Sepharose 48. Eluate from the Cibachron Blue-Sepharose was batch absorbed to Thiopropyl Sepharose and bound proteins were eluted with buffers of increasing reducing potential. Eluates were dialysed against HEPES buffer and assayed for plasmin reductase activity using ELISA as previously described. Results are mean +/- SEM of triplicate determinations. 88 Chapter 4 - Plasmin Reductase Purification 4.3.3 Sephacryl S-200 HR gel filtration Fractions containing proteins displaced by 20 mM OTT were pooled, concentrated and extensively dialysed. The pooled fraction was separated using Sephacryl S-200 HR. Three major peaks were separated (Figure 4.3). Plasmin reductase activity was found in the third peak. 25 .5 I Plasmin reductase .,., --Protein 2 ii' tn 20 3 .4 -·::::s ell c. - C 15 3 ..EC, .3 n ::l S' A tn C CD ·-Cl) m e 10 .2 ~ a. 1 <" ~ 5 .1 ;= g 0 ...... -----.....------. 0 20 40 60 ..--.------.-.---~•.O80 100 120 Elution volume, ml Figure 4.3. Purification of plasmin reductase on gel filtration. OTT eluate from the activated Thiopropyl-Sepharose chromatography was gel filtered on Sephacryl S-200 HR. Horizontal bar indicates pooled fractions. 89 Chapter 4 - Plasm in Reductase Purification The purity of protein was determined by SOS-PAGE. Samples of each protein peak were resolved under reducing conditions and silver stained (Figure 4.4). The plasmin reductase was associated with a single polypeptide chain of approximately 42,000 Da. A total of -20 µg of plasmin reductase was purified from 20 litres of HT1080 CM. 120- 80- •--•· J 50- ~ - ,,,_.. 40- ._ Plasmin reductase --.,, 20- ...--· ,_..__, ~ ~· 10- 1 2 3 4 5 Figure 4.4. SOS-PAGE of gel filtrated fractions. Fractions were resolved on reducing 8-16% SOS-PAGE and silver stained. Molecular weight markers (lane 1), OTT eluate from activated Thiopropyl-Sepharose chromatography (lane 2), eluates representing peaks 1, 2 and 3 of Sephacryl S-200 HR gel filtration (lanes 3, 4 and 5 respectively). 90 Chapter 4 - Plasmin Reductase Purification 4.3.4 Protein microsequencing The intact -42-44,000 Da protein was refractive to NH2-terminal sequencing. The protein was digested with CNBr and fragments resolved using reverse-phase HPLC (Figure 4.5). A total of 34 NH2-terminal amino acid residues from four internal peptides were determined. Data base sequencing revealed 100% homology with phosphoglycerate kinase (Figure 4.5). mAU 2 4 3 40 E C 0 3) CX) !::!- 8 C 20 l1l .0... 0 r/1 .0 0 20 eo 80 100 min Elution time (min) Figure 4.5. Sequencing profile for NHrterminal peptides of plasmin reductase. Sequenced peptides are numbered 1-4 on the HPLC chromatogram. 91 Chapter 4 - Plasm in Reductase Purification 4.3.4.1 Amino acid sequence of human PGK 1 MSLSNKLTLD KLDVKGKRW MRVDFNVPMK NNQITNNQRI KAAVPSIKFC RVDFN 51 LDNGAKSWL MSHLGRPDGV PMPDKYSLEP VAVELKSLLG KDVLFLKDCV SHLGRPDG 101 GPEVEKACAN PAAGSVILLE NLRFHVEEEG KGKDASGNKV KAEPAKIEAF 151 RASLSKLGDV YVNDAFGTAH RAHSSMVGVN LPQKAGGFLM KKELNYFAKA VGVN LPQKAGG 201 LESPERPFLA ILGGAKVADK IQLINNMLDK VNEMIIGGGM AFTFLKVLNN AFTFLKVLNN 251 MEIGTSLFDE EGAKIVKDLM SKAEKNGVKI TLPVDFVTAD KFDENAKTGQ 301 ATVASGIPAG WMGLDCGPES SKKYAEAVTR AKQIVWNGPV GVFEWEAFAR 351 GTKALMDEW KATSRGCITI IGGGDTATCC AKWNTEDKVS HVSTGGGASL 401 ELLEGKVLPG VDALSNI Figure 4.6. Sequence alignment of human PGK and NH2-terminal peptides of plasmin reductase. Sequence of NHrterminal peptides of plasmin reductase (yellow) and its corresponding sequence in human PGK (aqua). 92 Chapter 4 - Plasmin Reductase Purification 4.3.5 Stages of plasmin reductase purification 120- 80- ...... 50- 40- - .- Plasmin reductase -... 20- ... 10- -..--. 1 2 3 4 5 6 Figure 4.7. SOS-PAGE of significant stages of plasmin reductase purification. (1) Molecular weight markers; (2) unconcentrated; (3) concentrated HT1080 CM; (4) Cibachron Blue-Sepharose eluate; (5) Thiopropyl-Sepharose eluate and (6) eluate of Sephacryl S-200 HR gel filtration containing purified enzyme. Table 4.1. Summary of different steps in the purification of plasmin reductase Total Plasmin reductase specific Purification stage Protein activity (fold increase) (mg) Starting Materials 840 1 Concentrated CM 101 8 Cibachron Blue-Sepharose eluate 6 120 Thiopropyl-Sepharose eluate 2 710 Sephacryl S-200 HR gel filtration eluate 0.02 5000 93 Chapter 4 - Plasmin Reductase Purification 4.4 DISCUSSION The HT1080 cell line was selected for the isolation of plasmin reductase described herein. It was observed that plasmin reductase activity in HT1080 CM was inhibited by the adenine nucleotides, NADH or ATP, and inactivated by the thiol modifying reagents, 1AM and NEM. These results implied that plasmin reductase would have affinity for Cibachron Blue Sepharose and activated Thiopropyl-Sepharose chromatography matrices. Twenty litres of CM from cultured HT1080 cells (840 mg) was concentrated and applied to Cibachron Blue-Sepharose. The bound proteins were eluted with a linear NaCl gradient. Plasmin reductase activity eluted as a broad peak starting at approximately 1 M NaCl (Figure 4.1). The leading edge of the peak of plasmin reductase activity was pooled (6 mg with a 120-fold increase in specific activity) and batch adsorbed to activated Thiopropyl Sepharose. The matrix was developed with buffers of increasing reducing potential to elute the covalently bound proteins. The majority of the plasmin reductase activity was eluted with 20 mM DTT (2 mg with a 710-fold increase in specific activity) (Figure 4.2). The eluate was concentrated and gel filtered on Sephacryl S-200 HR (Figure 4.3). Three major components of the pooled sample were resolved. Plasmin reductase activity was well resolved and associated with a single polypeptide chain of 42,000 Da by SDS-PAGE (Lane 5, Figure 4.4). Approximately 20 µg of plasmin reductase was purified with a 5,000-fold increase in specific activity. Sequence analysis on the four internal peptides revealed that plasmin reductase has an exact (100%) sequence identity to phosphoglycerate 94 Chapter 4 - Plasmin Reductase Purification kinase [203] (PGK; ATP:3-phospho-D-glycerate 1-phospho-transferase, EC 2.7.2.3). PGK has been traditionally studied as a glycolytic enzyme that equilibrates phosphate transfer between 1,3 BPG and the y-phosphate of MgATP. More recently, PGK was shown to influence DNA replication and repair in mammalian cell nuclei [170, 171], stimulate viral mRNA synthesis in the cytosol [172], and was detected on the surface of Candida albicans where it extends through the cell wall [173]. PGK is present in high concentration in the cytoplasm. The finding that transformed cells secrete PGK is an interesting observation. Most secreted proteins contain a traditional consensus signal peptide, which is thought to be responsible for targeted transport across cell membranes. The fact that PGK lacks this signal sequence implies that PGK may be secreted via an as yet unknown pathway. 95 CHAPTERS CLONING AND EXPRESSION OF HUMAN RECOMBINANT PGK Chapter 5 - Cloning and Expression of Human PGK 5.1 INTRODUCTION The plasmin reductase secreted by HT1080 cells in culture is identical to PGK. To further confirm this finding, recombinant human PGK was cloned and expressed in Escherichia coli followed by purification of the recombinant protein using similar protocols to the HT1080 PGK purification. I compared the functional assays of recombinant and HT1080 PGK in angiostatin formation. Human PGK cDNA was cloned by RT-PCR from HT1080 RNA. The recombinant protein was produced in Escherichia coli, extracted and purified by ammonium sulfate precipitation, Cibachron Blue-Sepharose and Heparin Sepharose chromatographies. The plasmin reductase activity of rhPGK was assayed using ELISA and Streptavidin-HRP blotting techniques as described previously in Chapter 3. 96 Chapter 5 - Cloning and Expression of Human PGK 5.2 MATERIALS AND METHODS 5.2.1 Enzymes and reagents Total RNA was extracted from cultured HT1080 cells using an RNA extraction kit (Rneasy Mini Kit, QIAGEN). DNA polymerase Pfx and primers were obtained from Life Technology. All other enzymes and materials for DNA manipulation were purchased from Roche Molecular Biochemicals. A T7 promoter/RNA polymerase expression system including plasmid vector, pET11a, and bacterial strain BL21 (DE3), were provided by Novagen. B PER reagent for bacterial lysis and protein extraction was purchased from Pierce (Rockford, IL). Automated DNA sequencing was performed by the Department of Biological Sciences, University of New South Wales, Sydney. 5.2.2 Cellular RNA extraction Cultured HT1080 cells at -30% confluence were washed with cold PBS, harvested and centrifuged at 2000 rpm for 2 minutes. Cell pellet was suspended in 200 µI of chilled lysis buffer containing 10 mM Tris-HCI, 150 mM NaCl, 1.5 mM MgCI2, 0.65% (v/v) NP-40, pH 7 .5 and vortexed thoroughly for 15 seconds. The suspension was then centrifuged immediately for a further 15 seconds to remove the nuclear pellets. Cytoplasmic lysate was dissolved in 200 µI of RNA extraction buffer containing 7 M urea, 1% (w/v) SOS, 350 mM NaCl, 10 mM EDTA, 10 mM Tris-HCI, pH 7 .5. PCI (phenol:chloroform:isoamyl) solution at a ratio of 25:24:1 (400 µI) was added to the suspension, vortexed thoroughly and centrifuged at 97 Chapter 5 - Cloning and Expression of Human PGK 12,000 rpm for 2 minutes. The upper layer was transferred to a tube containing 1 ml of chilled 95% ethanol and incubated overnight at -20°C before being centrifuged at 13,000 rpm for 15 minutes and washed with 70% ethanol in DEPC H2O. The RNA was dissolved in 30 µI of RNAse free H2O and the concentration determined at 260 nm. An absorbance of one unit at 260 nm corresponds to 40 µg of RNA per ml (A250 =1 =40 µg/ml). 5.2.3 Construction of human PGK cDNA A 1.33 kb PGK cDNA was cloned by reverse transcriptase polymerase chain reaction (RT-PCR) from HT1080 RNA using two gene specific primers. A forward primer, 5'-A GTA CAT ATG TCG CTT TCT AAC AAG CTG-3' (positions 80 to 100) and a reverse primer, 5'-A GTA GGA TCC CTA ATG CCA AGT GGA GAT GCA-3' (positions 1409 to 1389) were designed based on the published DNA sequence of human PGK [204]. Ndel and BamHI sites were incorporated into the forward and reverse primers respectively (underlined) to facilitate sub-cloning. In addition, four bases were added to the 5' end of each primer to enhance recognition by the restriction enzymes. These primers were used to amplify the full-length cDNA of hPGK by RT PCR using DNA polymerase Pfx. 5.2.3.1 PCR reaction Each 50 µI PCR reaction contained 10 µg of reverse transcriptase product, 0.4 µM of each primer, 1unit of Pfx polymerase, 1 mM MgSO4, and 0.3 mM dNTPs. Samples were overlaid with mineral oil (Sigma) to prevent evaporation. The initial denaturing step was at 95°C for 2 minutes. The 98 Chapter 5 - Cloning and Expression of Human PGK cyclic parameters consisted of 20 seconds denaturing at 95°C, 45 seconds annealing at 52°C, and 100 seconds extension at 68°C. This was repeated for 18 cycles with a final extension period of 10 minutes at 68°C. 5.2.3.2 cDNA and plasmid vector digestion The PCR product was extracted twice by PCI and precipitated with ethanol. The cDNA insert and the plasmid vector, pET11 a, were cleaved by Ndel and BamHI followed by extraction and purification from agarose gel. 5.2.3.2.1 cDNA digestion cDNA digestion was performed in a 200 µI reaction containing 10 µI of cDNA (2-3 µg) and 5µ1 of each restriction enzyme. The reaction was incubated at 37°C for 3 hours in aliquots of 50 µI. 5.2.3.2.2 Plasmid vector digestion pET11a vector was digested with Ndel and BamHI. The digested vector was separated via gel electrophoresis and the vector purified using a gel extraction kit (QIAGEN) followed by ethanol precipitation. 5.2.3.3 Ligation reaction The digested cDNA was cloned into the Ndel and BamHI sites of the plasmid expression vector, pET11 a, by ligation. The reaction contained 2 µI of insert (hPGK//Ndel-BamHI) and 2 µI of vector (pET11a//Ndel-BamHI), 1 µI 99 Chapter 5 - Cloning and Expression of Human PGK of T- 7 ligase, 1 µI of ligase buffer and 4 µI of dH2O. The reaction mixture was incubated overnight at 4°C. 5.2.3.4 Transformation Escherichia coli strain, BL21 (DE3), was transformed with the recombinant plasmid (pET11 a/hPGK) and rhPGK expression, BLpET11 a hPGK was then selected from the derived transformants. BL21 (DE3) cells were gently thawed on ice. The recombinant plasmid (pET11 a/hPGK) was diluted in 10 mM Tris, pH 8.0 and 80 µI of BL21 (DE3) cells added. Cells were suspended gently and incubated on ice for 30 seconds, heated at 42°C for 2 minutes and then incubated on ice for a further 30 seconds. Cells were added to 1 ml of LB containing 75 µg/ml of Ampicillin (LB/Amp75) and incubated at 37°C for 1 hour. Cells (100 µI) were spread onto an agar plate containing 75 µg/ml of Ampicillin and cultured overnight at 37°C. 5.2.4 Expression of human recombinant PGK 5.2.4.1 Analysis of rhPGK expression A single colony from the selecting agar plate was inoculated in 1 ml of LB/Amp75 and grown overnight at 37°C. This culture was diluted to a ratio of 1:20 in 1 ml of fresh LB/Amp75 and incubated for 4 hours at 37°C. Protein expression was induced by adding IPTG to a final concentration of 0.5 mM and incubated for a further 5 hours at 37°C. The culture was centrifuged at 6,000 rpm at 4°C for 10 minutes and the pellet was suspended in 200 µI of 8- PER reagent. The lysate was clarified by centrifugation at 15,000 rpm for 20 100 Chapter 5- Cloning and Expression of Human PGK minutes and the supernatant containing soluble rhPGK protein collected and analysed on SOS-PAGE. 5.2.4.2 Large-scale preparation of rhPGK (2 litre cultures) To prepare large-scale rhPGK expression, 1 ml of culture was added to 40 ml of LB/Amp75 media and cultured overnight at 37°C shaker. Cells were diluted to a 1:50 ratio in LB/Amp75 media and incubated for 4 hours or until 0D600 = 0.6-0.8 before adding 0.5 mM IPTG to induce protein expression. Cells were collected by centrifugation at 6000 rpm for 10 minutes. Sixty millilitres of 8-PER bacterial extraction reagent per litre of culture was used to lyse the pellet. Cells were mixed thoroughly followed by gentle shaking at room temperature for 20 minutes. The lysate was sonicated four times at 20 seconds each and clarified by centrifugation at 15,000 rpm for 20 minutes at 4°C. A cocktail of protease inhibitors (10 µM leupeptin, 1 mM PMSF, 2 mM EDTA and 10 µg/ml soybean trypsin inhibitor) was added to the sample to prevent proteolytic cleavage of rhPGK. 5.2.5 Purification of rhPGK 5.2.5.1 Ammonium sulfate precipitation The supernatant was precipitated with ammonium sulfate starting with 25% up to 100% saturation. In each step, the sample was centrifuged at 15,000 rpm for 20 minutes at 4°C. The 100% ammonium sulfate precipitate pellet was dissolved in 20 ml of HEPES buffer containing 20 mM HEPES, 1 mM EDTA, 0.05 M NaCl, 0.02% (w/v) NaN3, pH 7.4. Ammonium sulfate was 101 Chapter 5 - Cloning and Expression of Human PGK subsequently removed by dialysing three times against 2 litres of HEPES buffer. 5.2.5.2 Cibachron Blue-Sepharose chromatography The dialysed sample was applied to a Cibachron Blue-Sepharose chromatography matrix (80 ml, 2.5 x 20 cm) and equilibrated with starting buffer containing 20 mM HEPES, 0.05 M NaCl, 1 mM EDTA, 0.02% (w/v) NaN 3, and pH 7.4. The sample was loaded at a flow rate of 1 ml/min and matrix washed with three column volumes of starting buffer until a stable baseline was achieved. Bound proteins were eluted by a linear gradient of 0.05-2 M NaCl at a flow rate of 0.5 ml/min. Fractions of 1.5 ml were collected and assayed for reductase activity. Fractions containing rhPGK were pooled and dialysed twice with one litre of buffer containing 20 mM HEPES, 0.14 M NaCl, 1 mM EDTA, pH 7.4. 5.2.5.3 S-Sepharose chromatography rhPGK was further purified on S-Sepharose chromatography. A 2 ml sample was applied to a 1 ml S-Sepharose column equilibrated with 20 mM MES, 1 mM EDTA, 0.05 M NaCl, pH 6.0. Bound proteins were eluted with a 0.05-1 M NaCl linear gradient. 5.2.6 Mass spectrometry Mass Spectrometry analysis was performed in collaboration with Dr. Mark Raftery (Cytokine Research Unit, UNSW). rhPGK (4 mg/ml) was further purified using C4 Reversed-phase HPLC, proteins were then dried 102 Chapter 5 - Cloning and Expression of Human PGK using a Speedvac concentrator and dissolved in CH3CN/H2O/CH3CO2H (1: 1 1%). Mass spectra were acquired on a triple quadrupole mass spectrometer (TSQ 7000, Finnigan, San Jose, CA) equipped with an electrospray ionisation source. Sample was administered into a moving solvent (10µI/min; 50:50 CH3CN/H2O, 0.1 % CH3CO2H) coupled directly to the ionisation source via a fuse silica capillary (50 µm x 25 cm). Sample droplets were ionised at a positive potential of 4.5 kV. The ions were extracted into the analyser through a heated capillary (T=180°C). Full scan mass spectra were acquired over the mass range of m/z 600 to 1800 over 2 seconds and deconvoluted using the instrument software. 5.2.7 ELISA for reductase activity of rhPGK The reductase activity of rhPGK was determined using ELISA and Streptavidin-HRP blotting techniques as described previously in Chapter 3. 103 Chapter 5-Cloning and Expression of Human PGK 5.3 RESULTS 5.3.1 cDNA sequences of rhPGK The complete cDNA sequence for rhPGK was verified by automated sequencing. The hPGK cDNA open reading frame consists of 1251 nucleotides, which codes for 417 amino acids starting with a Methionine residue (ATG) (Figure 5.1 ). AAG CCT CCG GAG CGC ACG TCG GCA GTC GGC TCC CTC GTT GAC CGA ATC ACC GAC CTC TCT CCC CAG CTG TAT TTC CAA A l!Q ATG TCG CTT TCT AAC AAG CTG ACG CTG GAC AAG CTG GAC GTT AAA GGG AAG CGG GTC GTT 1 M s L s N K L T L D K L D V K G K R V V ATG AGA GTC GAC TTC AAT GTT CCT ATG AAG AAC AAC CAG ATA ACA AAC AAC CAG AGG ATT 21 M R V D F N V p M K N N Q I T N N Q R I AAG GCT GCT GTC CCA AGC ATC AAA TTC TGC TTG GAC AAT GGA GCC AAG TCG GTA GTC CTT 41 K A A V p s I K F C L D N G A K s V V L ATG AGC CAC CTA GGC CGG CCT GAT GGT GTG CCC ATG CCT GAC AAG TAC TCC TTA GAG CCA 61 M s H L G R p D G V p M p D K y s L E p GTT GCT GTA GAA CTC AAA TCT CTG CTG GGC AAG GAT GTT CTG TTC TTG AAG GAC TGT GTA 81 V A V E L K s L L G K D V L F L K D C V GGC CCA GAA GTG GAG AAA GCC TGT GCC AAC CCA GCT GCT GGG TCT GTC ATC CTG CTG GAG 101 G p E V E K A C A N p A A G s V I L L E AAC CTC CGC TTT CAT GTG GAG GAA GAA GGG AAG GGA AAA GAT GCT TCT GGG AAC AAG GTT 121 N L R F H V E E E G K G K D A s G N K V AAA GCC GAG CCA GCC AAA ATA GAA GCT TTC CGA GCT TCA CTT TCC AAG CTA GGG GAT GTC 141 K A E p A K I E A F R A s L s K L G D V TAT GTC AAT GAT GCT TTT GGC ACT GCT CAC AGA GCC CAC AGC TCC ATG GTA GGA GTC AAT 161 y V N D A F G T A H R A H s s M V G V N CTG CCA CAG AAG GCT GGT GGG TTT TTG ATG AAG AAG GAG CTG AAC TAC TTT GCA AAG GCC 181 L p Q K A G G F L M K K E L N y F A K A TTG GAG AGC CCA GAG CGA CCC TTC CTG GCC ATC CTG GGC GGA GCT AAA GTT GCA GAC AAG 201 L E s p E R p F L A I L G G A K V A D K ATC CAG CTC ATC AAT AAT ATG CTG GAC AAA GTC AAT GAG ATG ATT ATT GGT GGT GGA ATG 221 I Q L I N N M L D K V N E M I I G G G M GCT TTT ACC TTC CTT AAG GTG CTC AAC AAC ATG GAG ATT GGC ACT TCT CTG TTT GAT GAA 241 A F T F L K V L N N M E I G T s L F D E GAG GGA GCC AAG ATT GTC AAA GAC CTA ATG TCC AAA GCT GAG AAG AAT GGT GTG AAG ATT 261 E G A K I V K D L M s K A E K N G V K I ACC TTG CCT GTT GAC TTT GTC ACT GCT GAC AAG TTT GAT GAG AAT GCC AAG ACT GGC CAA 281 T L p V D F V T A D K F D E N A K T G Q GCC ACT GTG GCT TCT GGC ATA CCT GCT GGC TGG ATG GGC TTG GAC TGT GGT CCT GAA AGC 301 A T V A s G I p A G w M G L D C G p E s AGC AAG AAG TAT GCT GAG GCT GTC ACT CGG GCT AAG CAG ATT GTG TGG AAT GGT CCT GTG 321 s K K y A E A V T R A K Q I V w N G p V GGG GTA TTT GAA TGG GAA GCT TTT GCC CGG GGA ACC AAA GCT CTC ATG GAT GAG GTG GTG 341 G V F E w E A F A R G T K A L M D E V V AAA GCC ACT TCT AGG GGC TGC ATC ACC ATC ATA GGT GGT GGA GAC ACT GCC ACT TGC TGT 361 K A T s R G C I T I I G G G D T A T C C GCC AAA TGG AAC ACG GAG GAT AAA GTC AGC CAT GTG AGC ACT GGG GGT GGT GCC AGT TTG 381 A K w N T E D K V s H V s T G G G A s L 1279 GAG CTC CTG GAA GGT AAA GTC CTT CCT GGG GTG GAT GCT CTC AGC AAT ATT TAG 401 E L L E G K V L p G V D A L s N I 1333 TAC TTT CCT GCC TTT TAG TTC CTG TGC ACA GCC CCT AAG TCA ACT TAG CAT TTT CTG CAT 1393 CTC CAC TTG GCA TTA GCT AAA ACC TTC CAT GTC AAG ATT CAG CTA GTG GCC AAG AGA TGC 1453 AGT GCC AGG AAC CCT TAA ACA GTT GCA CAG CAT CTC AGC TCA TCT TCA CTG CAC CCT GGA 1513 TTT GCA TAC ATT CTT CAA GAT CCC ATT TGA ATT TTT TAG TGA CTA AAC CAT TGT GCA TTC 1573 TAG AGT GCA TAT ATT TAT ATT TTG CCT GTT AAA AAG AAA GTG AGC AGT GTT AGC TTA GTT 1633 CTC TTT TGA TGT AGG TTA TTA TGA TTA GCT TTG TCA CTG TTT CAC TAC TCA GCA TGG AAA 1693 CAA GAT GAA ATT CCA TTT GTA GGT AGT GAG ACA AAA TTG ATG ATC CAT TAA GTA AAC AAT 1753 AAA AGT GTC CAT TG Figure 5.1. Coding cDNA and derived sequences of rhPGK. rhPGK sequence starts at Met1 and ends at lle417. Amino acid numbers are in bold. 104 Chapter 5 - Cloning and Expression of Human PGK 5.3.2 rhPGK purification The rhPGK was purified to homogeneity using ammonium sulfate precipitation, Cibachron Blue and S-sepharose ion exchange chromatography matrices. The protein concentration of the purified PGK was determined spectrophotometrically at 280 nm with an absorption index of EJ~ nm = 6.9 (1-cm light path) [205]. Approximately 30 mg of rhPGK protein resulted from one litre of bacterial culture. Purified rhPGK resolved at the same size as the HT1080 derived PGK on an 8-16% SOS PAGE gel stained with Coomassie Brilliant Blue and has an approximate molecular mass of 44,000 Oa (Figure 5.2). The molecular mass of rhPGK protein was also determined by mass spectrometry (Figure 5.3). 120 80- 50 - 40 --. 1111 ._ Plasmin reductase 20 ._ 10 ~ 1 2 3 Figure 5.2. SDS-PAGE of HT1080 PGK and rhPGK. rhPGK (2 µg) was resolved on an 8-16% SOS-PAGE gel and stained with Coomassie Brilliant Blue. Molecular weight markers (lane 1); HT1080 derived PGK (lane 2) and rhPGK (lane 3) . 105 Chapter 5 - Cloning and Expression of Human PGK 5.3.3 Mass spectrometry of recombinant human PGK The experimental mass determined by electrospray mass spectrometry of rhPGK was 44,493 which compared well with the theoretical mass (44498) derived from its cDNA sequences (Figure 5.3). 8908 10 0: 98 .8 44493.0 C C C 7 5' 825. B 1141.7 C ta "Cl C 810. :::, 203.5 0 ~ 5 C 795. I 1309.7 i 2 5' 13 1.4 788.: l. l1589.8 ,~~ 11 ~~ ~ .. ·-· l. j l178r 800 1000 1200 1400 1600 1800 44000 44200 44400 44600 44800 45000 45200 45400 m/z mass Figure 5.3. Purified rhPGK has a molecular mass of m/z 44493 as determined by mass spectrometric analysis. (A) Multiply charged ions formed by electrospray, (B) Deconvoluted mass spectrum of recombinant wild-type PGK. 106 Chapter 5 - Cloning and Expression of Human PGK 5.3.4 Plasmin reductase activity of rhPGK rhPGK has the same plasm in reductase activity as the HT1080 derived PGK. Both proteins convert plasmin to angiostatin fragments as shown in Figure 5.4 A. All three fragments of angiostatin contained free thiols as they incorporated MPS and detected on Streptavidin-HRP blotting analysis (Figure 5.4 B) . A B ...... 120 120- -I ... - 85 c;4I 51 ,_ 43 36 *= ...... '* 25 20- --- 10 ~ 10-- 1 2 3 4 1 2 3 Figure 5.4. rhPGK has the same plasmin reductase activity as HT1080 PGK (A) Molecular weight markers (lane 1), Plasm in (20 µg) was incubated with HEPES buffered saline (lane 2), 500 µI of HT1080 CM (lane 3), or with 40 µg of rhPGK (lane 4) . The plasmin products were separated on 8-16% SOS-PAGE gel and silver stained. (B) Streptavidin-HRP blot of angiostatin generated by HT1080 PGK and rhPGK. Plasm in (20 µg) was incubated with HEPES-buffered saline (lane 1), with 500 µI of HT1080 CM (lane 2), or with 40 µg of rhPGK (lane 3) and the angiostatin fragments labelled with MPB. The plasmin products were separated on 8-16% SOS-PAGE gel and blotted with Streptavidin-HRP. Arrows indicate three angiostatin fragments. 107 Chapter 5 - Cloning and Expression of Human PGK 5.3.5 Comparison of plasmin reductase activity of various PGK The reductase activity of human, rabbit and yeast PGK were compared using ELISA and Streptavidin-HRP blotting techniques. Human and rabbit PGK had comparable plasmin reductase activity, whereas yeast PGK had negligible plasmin reductase activity (Figure 5.5 A and B). A B 0.6 0.5 ,_.. 0.4 120- E C 80- "' ~ 0.3 51- C1J u C 43- .Q"' 0.2 36- 0 1/) -·- .Q <( 25- 0.1 10- 0.0 Iii 2 3 4 5 Figure 5.5. Plasmin reductase activity of PGK from various origins. (A) Plasm in (2 µg/ml) was incubated with either HT1080 PGK, rhPGK, rabbit PGK or yeast PGK (5 µg/ml) in HEPES-buffered saline for 30 minutes at 37°C and the angiostatin fragments labelled with MPB and detected by ELISA. Results are mean and SEM of triplicate determinations. (B)Angiostatin fragments generated in part A; plasm in alone (lane 1) , plasm in with either HT1080 PGK (lane 2), rhPGK (lane 3), rabbit PGK (lane 4) or yeast PGK (lane 5) were separated on an 8-16% SOS-PAGE gel and transferred to PVDF membrane. The MPB labelled fragments were blotted with Streptavidin-HRP and visualised using chemiluminescence. 108 Chapter 5 - Cloning and Expression of Human PGK 5.4 DISCUSSION In Chapter 4, I described the purification of plasmin reductase and its identity as PGK. To confirm this finding, I cloned and expressed rhPGK and studied its role in angiostatin formation from plasmin. Two primers at the 5' and 3' positions of hPGK DNA were used to amplify the hPGK cDNA by RT PCR from HT1080 RNA. The resulting PCR product was sub-cloned into the plasmid vector, pET11 a, which was then transfected into Escherichia coli strain BL21 (DE3). The complete hPGK cDNA sequence (1767 bp) consists of an open reading frame of 1251 bp, which codes for 417 amino acids (Figure 5.1) and has an approximate molecular mass of 44,000 Da. The expressed protein was extracted and purified using ammonium sulfate precipitation, Cibachron Blue and S-Sepharose chromatography matrices. The expression of rhPGK accounted for more than 70% of the total protein expressed. rhPGK had the same molecular weight as the HT1080- derived protein on an 8-16% SOS-PAGE gel under non reducing conditions (Figure 5.2) and had the same plasmin reductase activity (Figures 5.4). This finding demonstrated that plasmin reductase secreted by HT1080 cells is PGK. The plasmin reductase activity of PGK from various sources was examined. These findings showed that rabbit muscle PGK had comparable specific plasmin reductase activity to human PGK, while yeast PGK had negligible activity (Figures 5.5 A and B). Human and rat PGK share 97% sequence identity while the human and yeast enzyme share only 65% sequence identity. There are seven cysteine residues in human or rabbit PGK molecule while yeast PGK has only a single cysteine residue. This 109 Chapter 5 - Cloning and Expression of Human PGK difference could account for the lack of plasmin reductase activity observed in yeast PGK. This observation is consistent with the finding that plasmin reductase activity in HT1080 CM was inhibited by alkylating reagents such as N-ethylmaleimide or iodoacetamide, which suggests that one or more thiols in PGK may play a role in plasmin disulfide bond reduction. 110 CHAPTERS PGK SECRETION IN VIVO Chapter 6 - PGK Secretion in vivo 6.1 INTRODUCTION In this investigation I measured the rate of PGK secretion by various transformed cells using a coupled kinetic assay. I also attempted to characterise the role of PGK in angiostatin-mediated suppression of tumour angiogenesis and growth in vivo. In Chapter 3, the secretion of plasmin reductase was studied in a number of transformed cells. An additional panel of transformed cells was investigated to determine the levels of PGK secretion. It was found that cultured HT1080 cells secrete significant levels of plasmin reductase. To determine if HT1080 tumours secrete PGK in vivo, plasma levels of PGK in mice with HT1080 tumours and in control mice without tumours were investigated. Furthermore, I examined whether PGK could reduce plasmin to form angiostatin in vivo. I hypothesised that rhPGK would facilitate the reduction of plasmin produced by tumours and result in increased plasma levels of angiostatin containing free thiols. Mice with and without HT1080 tumours were injected with rhPGK and the plasma levels of angiostatin before and after treatment were determined. This experiment led to the investigation of the role of PGK in a tumourigenesis model. Mice bearing HT1080 or AsPC-1 tumour were administered PGK. The effect of PGK treatment on tumour growth and angiogenesis was examined. 111 Chapter 6 - PGK Secretion in vivo 6.2 MATERIALS AND METHODS 6.2.1 Cell Culture The HT1080, BT20, MDA-231 and MCF-7M cells were cultured as described in Chapter 3. Human pancreatic cancer cell lines (BxPC-3, SU.86.86, HS 776T and HS 776Tf) were maintained in RPMI 1640 containing 10% (v/v) FBS, 10 units/ml penicillin streptomycin and 2 mM L-glutamine, pH 7.2. The AsPC-1 cells were maintained in RPMI containing 20% heat inactivated FBS. Confluent cells were passaged 1:4 twice a week. Colon carcinoma cell lines (Lim 1215, Lim 2412 and L1863) were maintained in RPMI 1640 containing 20 mM HEP ES, 10 mM NaHCO3, 5% (v/v) FCS, 40 µg/ml of gentamycin, pH 7.2. Cells in suspension were passaged 1:3 on a weekly basis. All cells were maintained at 37°C in a 5% CO2 humidified atmosphere. 6.2.2 Coupled kinetic assay to measure PGK secretion The rate of PGK secretion in CM of various transformed cells was determined spectrophotometrically at 340 nm based on the reaction illustrated below. The 1,3-bisphosphoglycerate (1,3-BPG) formed from 3- phosphoglycerate (3-PG) and ATP is further converted to glyceraldehyde 3- phosphate (GA-3-P) by coupled enzyme assay with glyceraldehyde -3- phosphate dehydrogenase (GAPDH) in the presence of NADH. The reaction measures the oxidation of NADH by 1,3-BPG as the decrease of absorption at 340 nm. 112 Chapter 6 - PGK Secretion in vivo PGK GAPDH 3-PG ~IJJi 1,3-BPG ...11111-7---,-----==~----. GA-3-P ~Mg~ ATP ADP NADH NAO+ ,l.. OD at 340 nm Figure 6.1 Coupled kinetic assay to measure PGK secretion. ATP, NADH, 3-PG and GAPDH were purchased from Sigma (St Louis, MO). GAPDH was dissolved in buffer containing 20 mM HEPES, 1 mM EDTA, 1mM OTT, 50% (w/v) ammonium sulfate, pH 7.4. Other enzymes were dissolved in buffer containing 50 mM HEPES, 50 mM KCI, 0.2 mM EDTA, 0.07 M NaCl, pH 7.4. The assay was performed in a 96 well microtitre plate in 200 µI of buffer containing 0.15 mM NADH, 10 mM ATP, 10 mM 3-PG, 1.2 mM MgCb, 0.4 mg/ml GAPDH. The reaction was initiated by addition of rhPGK (0-125 ng/ml) and incubated at room temperature for 1 hour with orbital shaking. The rate of oxidation of NADH was then measured for 15 minutes at 340 nm on a microtitre plate reader. A standard curve was constructed. 113 Chapter 6- PGK secretion in vivo 6.2.3 Tumour implantation The following protocol was used to study the effect of systemic administration of PGK on plasma PGK and angiostatin in mice with or without a HT1080 tumour. 40 days• 1 hour.. 2 hours.. HT1080 transplant bleed 10 mg/kg rhPGK IP bleed Figure 6.2. Protocol for HT1080 tumour implantation in SCID mice HT1080 tumours (3x106 cells) were established subcutaneously in the proximal midline dorsum of SCID mice [151]. After 40 days, blood was collected from the retroorbital plexus, then one hour later they were administered via intraperitoneal (i.p.) injection of 10 mg/kg of rhPGK (0.2 ml in PBS). Mice were bled 2 hour-post treatment. Plasma was separated and assayed to determine the level of PGK and angiostatin. For MPB labelling angiostatin assay, blood was collected in PBS containing EDTA (20 mM), aprotinin (5 µM) and MPB (200 µM). Cysteine (400 µM) was added after 60 minutes to quench unreacted MPB. 114 Chapter 6- PGK secretion in vivo 6.2.4 Tumourigenesis model To investigate the effect of PGK administration on tumour development and tumour angiogenesis the following procedures were employed. HT1080 or AsPC-1 tumours were implanted subcutaneously in the proximal midline dorsum of SCID mice. Daily treatment of 5 mg/kg of rhPGK or PBS began 10 days after tumour induction via i.p. injection. The smallest (a) and the largest (b) diameters across the tumour were measured every three days by calipers and tumour volume was calculated using the following formula: Tumour volume (mm 3) = a2 x b x 0.52 HT1080 or AsPC-1 in SCID p~ -10 days ... p~ tumour transplant treatment 3 x 106 cells 5 mg/kg/day IP Figure 6.3. Model for tumourigenesis The experiment was terminated (after 25 days) when the tumours of control mice reached 10% of their respective body weights. Tumours were resected for immunohistochemical analysis. 115 Chapter 6 - PGK Secretion in vivo 6.2.5 lmmunohistochemical staining for angiogenesis Tumours were fixed and embedded in paraffin according to standard histological procedures [151]. Endogenous peroxidase activity was blocked by incubation with 0.3% H202 in PBS followed by a triple rinse with PBS. Endothelial cells were stained using antibodies against PECAM (DAKO, Carpinteria, CA) diluted in PBS containing 5% serum. The bound antibody was detected by sequential incubation of the sections with biotinylated anti CD31 antibody followed by streptavidin-peroxidase complex. Positive staining was detected by substrate reaction with diaminobenzidine. Vessel density was determined by counting the number of capillary blood vessels/microvessels (capillaries and venule) per high power field section. 116 Chapter 6 - PGK Secretion in vivo 6.3 RESULTS 6.3.1 Assay used to determine PGK secretion rhPGK (0-125 ng/ml) was used in the coupled kinetic assay to construct the standard curve. The initial rate of reaction (i.e. change in absorbance per minute at 340 nm) was plotted against known concentrations of PGK. The level of PGK secretion in CM of various transformed cells was then determined from the standard curve (Figure 6.4). 0.03 C E E -C ..,....0 0.02 <<] i C'G 0::: -C'G E 0.01 -C 0.00 0 50 100 150 rhPGK, ng/ ml Figure 6.4. PGK coupled kinetic assay standard curve. Standard curve used to determine the rate of PGK secretion in CM of transformed cultured cells. 117 Chapter 6 - PGK Secretion in vivo 6.3.2 PGK is secreted by cultured carcinoma cells The rate of PGK secretion by various carcinoma cells varied considerably (Figure 6.5). Cultured HT1080 cells secreted 0.43 ± 0.01 µg of PGK per 106 cells in 24 hours. SU.86.86 cells secreted the most PGK, 2.09 ± 0.10 µg per 106 cells in 24 hours, while AsPC-1 cells secreted the least, 0.08 ± 0.01 µg per 106 cells in the 24 hour period. In addition, a cell line HS776Tf cloned from HS776T secreted 40% less PGK in 24 hours than the parent line . ... :, 2.0 0 .c •N C 1.5 .!! a, c., IO ...0 1.0 ...Ee C, 0.5 -::t -~ C) D. 0.0 I Iii 111 11 I 0 .... C") co I- 0 .... ~ N C") CIC) -I I CIC) ~ N -C") :E co ...... co .... CIC) 0 0 0 co I- N I N "It U> .... I .... D. D...... m LL .... N .... I- >< ~ .... <( :::c ~ m ::::, :::cUJ UJ C 0 E E E UJ :::c., :E :E :J :J :J ' V ~ '-y------J pancreatic breast colon Figure 6.5. Rate of PGK secretion by various transformed cells. Serum-free CM was collected after 24 hours and the amount of PGK secreted from 1x106 cells determined [206]. Results are mean of triplicate determinations. 118 Chapter 6 - PGK Secretion in vivo 6.3.3 Plasm in reductase activity of various cultured CM Conditioned media of the HT1080, breast (BT20, MDA-231, MCF-7M) and colon (Lim 1215, Lim 2412, Lim 1863) carcinoma cells were also analysed for plasmin reductase activity by ELISA. The plasmin reductase activity correlated with the rate of PGK secretion by these cell lines (Figure 6.6). 0.3 -E C: an 0 -.::I' 0.2 -Cl) (.) C:ea .c.. 0.1 0 ,n .c 0 0 :I!: 10 N CW) 00 N CW) .... co 0 ~ -N N "'1111' 00 m Figure 6.6. Plasmin reductase activity in CM of various cultured transformed cells. CM collected after 24 hours incubation with serum free media. Plasmin reductase activity was assayed using ELISA. 119 Chapter 6 - PGK Secretion in vivo 6.3.4 Plasma levels of PGK in tumour-bearing mice Plasma levels of PGK in mice bearing HT1080 tumours were determined using the coupled kinetic assay. The level of PGK secretion is 6.6-fold greater in mice bearing HT1080 tumours than mice without tumours (Figure 6.7). 3 E t 2 c. en :::1. -:L ~ 1 0 Figure 6.7. Plasma levels of PGK in mice bearing HT1080 tumours compared to mice without tumours. Blood was collected from the retro-orbital plexus of mice without (n=4) and with (n=4) tumours. Plasma levels of PGK were measured. Results are mean of triplicate values. 120 Chapter 6 - PGK Secretion in vivo 6.3.5 The effect of systemic administration of rhPGK on plasma levels of MPB-angiostatin Plasma levels of MPB-angiostatin were determined using ELISA as described previously. Administration of 10 mg/kg of rhPGK to tumour bearing mice caused a significant increase in plasma levels of MPB angiostatin after two hours (P < 0.05). In contrast, administration of rhPGK to mice without tumours did not affect the plasma level of MPB-angiostatin (Figure 6.8). c::::J -1 h E 0.4 -2h C: II) 0 ...,'111:t' 0.3 ns Cl) (J C ns 0.2 .c... 0 u, .c 0.1 0.0 ~ $- ~o 0 vo<:- ~~~ )C Figure 6.8. Plasma levels of angiostatin before and after treatment with rhPGK. Blood was collected 1 hour before (open bars) and 2 hours after (solid bars) the administration of 1 Omg/kg of rhPGK and assayed to determine the level of MPS-labelled angiostatin. Results are the mean of triplicate values. Double asterisks: P < 0.05. 121 Chapter 6 - PGK Secretion in vivo 6.3.6 Inhibition of HT1080 tumour growth by rhPGK Treatment of mice with 5 mg/kg/day of rhPGK caused 50% reduction in the rate of HT1080 tumour growth (Figure 6.9). 4 --0-PBS M ---- 5 mg/kg/day rhPGK E CJ ft Cl) E 3 ::::s -0 ..> ::::s 0 2 E ...::::s co0 0 .... 1 ...:::c 0 5 10 15 20 25 Number of days after tumour injection Figure 6.9. Inhibition of HT1080 tumour growth by rhPGK. Tumor volumes were determined 5 days post-implantation and every 3 days thereafter. When tumours reached -0.1 cm 3 mice were randomised into two groups (n=4). rhPGK (5 mg/kg/day) or PBS alone was administered via an i.p. injection. The arrow indicates the start of treatment. Each point represents mean +/- SEM. 122 Chapter 6 - PGK Secretion in vivo 6.3.7 Inhibition of AsPC-1 tumour growth by rhPGK Administration of rhPGK (5 mg/kg/day) caused 70% reduction in the rate of AsPC-1 tumour growth compared to the control mice treated with PBS (Figure 6.10). 4 --0-- PBS ---e---- 5 mg/kg/day rhPGK C"I E CJ.. CD 3 E :::I -0 ...> :::I 0 2 E :::I 'I"" -I 0 0. 1 u, <( 0 5 10 15 20 25 30 Number of days after tumour injection Figure 6.10. Inhibition of AsPC-1 tumour growth by rhPGK. When the tumours were -0.1 cm3 in volume the mice were randomised into two groups (n=4). rhPGK (5 mg/kg/day) or PBS alone was administered by an i.p. injection. The arrow indicates the start of treatment. Each point represents mean +/ SEM. 123 Chapter 6 - PGK Secretion in vivo AsPC-1 tumours were resected from both groups of mice. Tumours of rhPGK treated mice were significantly reduced compared to control mice treated with PBS. There were no apparent adverse side effects of rhPGK administration to the mice. There were no macroscopic or microscopic signs of tissue necrosis (Figure 6.11 and 6.12). PBS treated rhPGK treated i1 cm I Figure 6.11. Reduction of AsPC-1 tumour size by rhPGK. Mice were sacrificed after 25 days of treatment with 5 mg/kg/day of rhPGK or PBS alone and tumours were excised. 124 Chapter 6 - PGK Secretion in vivo 6.3.8 Effect of rhPGK on tumour angiogenesis AsPC-1 tumours were stained for endothelial cells using CD-31 antibody as previously described [207). Mice treated with rhPGK show a marked reduction in tumour microvessel density compared to PSS-treated (Figure 6.12 A). Microvessel counts in the immunohistochemical section are shown (Figure 6.12 B). There were 80 ± 13 vessels per field in the rhPGK treated tumours compared to 173 ± 9 in the PBS treated tumours. PBS treated tumour rhPGK treated tumour A \ . ,, ,. . .. : . .. I • I • I . • .. I , , . ~ I I• •, • / \ • I \ .. ' I , •) • . ' ' \ .: . .. , • I , _,. , ' , I J- I OI'"\' ' , It' ~ t • ... ' l ,. - · ,, . . .. ' l " , . , ' '' ' • r, " I, • , , ' ,/"* . \ "" ' ' \ . . . , / . '' ) . . •• :-1 / , , . " t 1" ' '·. '. '' ' ' ' . ~ /.1 I , ..ii, . ' _, ' .. f ,.' ' / ,,, 1" • . B 200 "C -~ 150 L1.... (1) c. u, 100 -a; ** j 50 0 PBS rhPGK Figure 6.12. Microvessels in control and rhPGK-treated AsPC-1 tumours from SCID mice. Microvessels are stained brown (A). Microvessel density (B) ** P < 0.05. 125 Chapter 6 - PGK Secretion in vivo 6.4 DISCUSSION In the coupled kinetic assay I found that the rates of secretion of PGK by different carcinoma cells varied considerably (Figure 6.5). Cultured HT1080 cells secreted PGK at a rate of 0.43 ± 0.01 µg per 106 cells in 24 hours. This rate was compared with rates from four human pancreatic (AsPC-1, BxPC-3, SU.86.86 and HS776T), three human breast (BT20, MDA- 231 and MCF-7M) and three human colon (Lim1215, Lim2412 and Lim1863) carcinoma cell lines. The secretion rate varied by as much as 26-fold among the cell lines. SU.86.86 cells secreted the most PGK, 2.09 ± 0.10 µg per 106 cells in 24 hours, while AsPC-1 cells secreted the least, 0.08 ± 0.01 µg per 106 cells in the 24-hour period. In addition, a cell line doned from HS776T, HS776Tf, secreted 40% less PGK in 24 hours than the parent line. There was no discernible decrease in cell viability or significant release of lactate dehydrogenase from the cells following a 24-hour culture (data not shown). These observations suggested that secretion of PGK was a property of living cells and was not due to leakage from dead cells. In addition, the rate of secretion of PGK by the HT1080, breast and colon carcinoma cells correlated with their plasmin reduciase activity (Figure 6.6). Furthermore, the rate of PGK secretion by the pancreatic carcinoma cells was inversely correlated with the tumourigenicity of the cells grown subcutaneously in SCID mice. The tumourigenicity was found in the following order: AsPC-1 > BxPC-3 >> SU.86.86 (not shown). The plasma of mice bearing HT1080 tumours contained 6.6-fold more PGK than mice without tumours (Figure 6. 7). This finding suggested that relatively small amount of PGK was secreted by normal tissue. This is in 126 Chapter 6 - PGK Secretion in vivo accordance with the observation that transformed cells secrete much more PGK (Figure 3.3) than primary cells. To determine whether PGK could reduce plasmin in vivo, mice with and without HT1080 tumours were injected with rhPGK. I reasoned that the rhPGK would facilitate reduction of plasmin produced by the HT1080 tumours and result in increased plasma levels of angiostatin containing free thiols. The plasma levels of MPS-labelled angiostatin were 48% higher, on average, in tumour-bearing mice compared to mice without tumours (Figure 6.8). More importantly, administration of rhPGK to tumour-bearing mice resulted in a 43% increase in the plasma levels of MPS-labelled angiostatin after 2 hours (p < 0.05). In contrast, administration of rhPGK to mice without tumours had no significant effect on plasma levels of MPS-labelled angiostatin. These findings support the proposed role for PGK in angiostatin formation and imply that levels of angiostatin in the blood can be increased by systemic administration of PGK. This led us to test whether treatment of tumour-bearing mice with PGK would inhibit tumour angiogenesis and growth. The growth of human fibrosarcoma (HT1080) and pancreatic carcinoma (AsPC-1) tumours in immunocompromised mice was suppressed by the systemic administration of rhPGK. lntraperitoneal administration of 5 mg rhPGK per kg per day caused a -50% inhibition in the rate of HT1080 tumour growth (Figure 6.9) and -70% inhibition in the rate of AsPC-1 tumour growth (Figure 6.10). There were no apparent adverse effects of rhPGK administration. Nor were there any macroscopic (Figure 6.11) or microscopic (Figure 6.12 A) signs of necrosis in rhPGK-treated tumours. lmmunohistochemical staining of the AsPC-1 tumours for vascularity (Figure 127 Chapter 6 - PGK Secretion in vivo 6.12 A) showed a marked reduction in microvessel density in the rhPGK treated tumours. There were 80 ± 13 vessels per field in the rhPGK-treated tumours compared to 173 ± 9 in the vehicle-treated tumours (p < 0.05) (Figure 6.12 B). It is interesting that AsPC-1 tumourigenesis was more susceptible to suppression by systemic administration of PGK than was HT1080 tumourigenesis. This may reflect the 5-fold reduction in PGK secretion by AsPC-1 cells compared to HT1080 cells (Figure 6.5). 128 CHAPTER 7 PLASMIN REDUCTION BY PGK IS A THIOL INDEPENDENT PROCESS Chapter 7 - Plasmin Reduction by PGK is a Thiol-lndependent Process 7.1 INTRODUCTION Disulfide bonds of certain cell surface proteins can interchange between the oxidised and reduced state [208-21 0]. These observations suggest that the function of some secreted proteins may be controlled by interchange of one or more disulfide bonds [210]. The reduction of disulfide bonds in plasmin by a tumour cell-derived protein was the first example of disulfide exchange in a secreted soluble protein [187, 191, 200]. A second example is disulfide exchange in von Willebrand Factor, which is facilitated by thrombospondin-1 [211]. Plasmin reduction is the first step in formation of the tumour angiogenesis inhibitor, angiostatin. Plasmin proteolysis occurs in three stages. First, the Cys462-Cys541 and Cys512-Cys536 disulfide bonds in kringle 5 of plasmin are reduced by a plasmin reductase [187]. Second, reduction of the kringle 5 disulfide bonds triggers cleavage at Arg530-Lys531 in kringle 5, and also at two other positions C-terminal of Cys462, by a serine proteinase [187]. Autoproteolysis can account for the cleavage [187, 190], although another serine proteinase is responsible in human fibrosarcoma cell conditioned medium [187]. Third, the kringle 1-4½ fragments are cleaved by matrix metalloproteinases to produce either kringle 1-4 or 1-3 [145, 149, 188]. All three kringle-containing fragments have been shown to inhibit endothelial cell proliferation in vitro and angiogenesis in vivo [162]. The plasmin reductase isolated from human fibrosarcoma cell conditioned medium was shown to be the glycolytic enzyme, phosphoglycerate kinase (PGK; ATP:3-phospho-D-glycerate 1- phosphotransferase, EC 2.7.2.3). The finding that PGK facilitates reduction of 129 Chapter 7 - Plasmin Reduction by PGK is a Thiol-lndependent Process disulfide bond(s) in plasmin was the first demonstration of reduction of disulfide bonds in a secreted protein in vivo. The active site of reductases such as glutathione reductase, protein disulfide isomerase and thioredoxin are characterised by the presence of a pair of closely spaced Cys residues, typically in the consensus sequence of CysGlyXxxCys. Mammalian PGK's contain 7 Cys and only 2 of the 7 are nearby in the primary or tertiary structure [17 4]. Cys379,380 are close to the hinge region that links the N- and C-terminal domains of PGK [203] and have been referred to as 'fast-reacting' as they are amenable to alkylation by several thiol-reactive compounds [212]. This study examined the mechanism of plasmin reduction by PGK. The role of all 7 Cys and in particular the two fast-reacting Cys, in reduction of plasmin disulfide bonds by PGK has been explored in this study. The positions of all seven Cys residues in PGK are highlighted in Figure 7.1. 130 Figure 7.1 Three-dimensional structure of human PGK showing the organisation of the N- and C terminal domains. Cysteine residues are highlighted in yellow and their positions are labelled. The predicted hinge region is _. circled. This structure was constructed using SWISS-MODEL PDB file [181, 182]. The image was generated using <,.)_. Weblab Viewerlite software (Molecular simulation). Chapter 7 - Plasmin Reduction by PGK is a Thiol-lndependent Process 7 .2 MATERIALS AND METHODS 7 .2.1 Chemicals and reagents MPB and dibromobimane (bBBr) were purchased from Molecular Probes (Eugene, OR). Mercury Chloride (HgCl2), sodium tetrathionate (TT), reduced glutathione (GSH), oxidised glutathione (GSSG), DTNB and NEM were obtained from Sigma (St Louis, MO). Plasminogen was purified from human plasma and separated into its two carbohydrate variants according to published procedures [81]. Glu(1)-plasminogen was used in the experiments described herein. uPA was a gift from Serono Australia. Plasmin was generated by incubating plasminogen (20 µM) with uPA (20 nM) for 30 minutes in buffer containing 20 mM HEPES, 0.14 M NaCl, 1 mM EDTA, pH 7.4 at 37°C. 7 .2.2 Production of mutant rhPGK A 1.33 kb hPGK cDNA was isolated by RT-PCR from total RNA extracted from HT1080 cells as described previously. The seven Cys in wt PGK were mutated to Ala in a cumulative fashion. The two fast-reacting Cys, Cys379,380, were mutated first and the resulting C379,380A PGK cDNA was then used to mutate Cys367. The corresponding C379,380,367A PGK cDNA was then used to mutate Cys316 and so forth. The Cys were mutated in the order, Cys379, 380, 367, 316, 108, 99 and 50. The Cys379-380 mutation was generated by replacing thymidine bases at positions 1214 and 1217 with guanine and guanine bases at positions 1215 and 1218 with cytidine using QuikChange Site-Directed Mutagenesis kit from Stratagene (La Jolla, CA). The primers used were 5'-GAC ACT GCC ACT GCC GCT GCC AAA TGG 132 Chapter 7- Plasmin Reduction by PGK is a Thiel-Independent Process AAC AC-3' (forward, positions 1202 to 1233) and 5'-GTG TTC CAT TTG GCA GCG GCA GTG GCA GTG TC-3' (reverse, positions 1233 to 1202). All primers used for the subsequent mutants are listed in Table 7.1. A single Cys to Ala mutation at position 50 was also generated using wt PGK as the template. Integrity of the mutant DNA's was confirmed by automated DNA sequencing. 7 .2.2.1 PCR reaction Each 50 µI PCR reaction contained 200 ng of target plasmid DNA, 400 ng of each primer, 5 µI of 10 x reaction buffer, 0.3 mM dNTPs and 1.25 units of Pfu DNA polymerase. Samples were overlaid with 30 µI of mineral oil to prevent evaporation. The initial denaturing step was at 95°C for 30 seconds. The cycling parameters consisted of 30 seconds denaturing at 95°C, 1 minute annealing at 55°C, and 160 seconds extension at 68°C. This was repeated for 18 cycles with a final extension period of 8 minutes at 68°C. Following temperature cycling, reaction mixture was cooled to -37°C. 7 .2.2.2 Dpn I digestion One microlitre of Dpn I (1 0U/µI) was added to the PCR product. Reaction mixture was then centrifuged for 1 minute and incubated at 37°C for 1 hour to digest the parental DNA template. 133 Chapter 7 - Plasmin Reduction by PGK is a Thiol-lndependent Process 7 .2.2.3 Transformation of super competent cells (XL-B) Epicurian Coli XL-Blue super competent cells (50 µI) were gently thawed on ice and transformed with 1 µI of Dpnl-treated DNA. The transformation reaction was suspended gently and incubated on ice for 30 minutes, heated at 42°C for 45 seconds and then incubated on ice for a further 2 minutes. Cells were added to 0.5 ml of NZY broth containing 75µg/ml of Ampicillin and incubated at 37°C for 1 hour. Cells (100 µI) were spread onto an agar plate containing 75 µg/ml of Ampicillin and cultured overnight at 37°C. 7 .2.2.4 Transformation of BL21 (DE3) cells Three colonies were selected from overnight culture. The mutant plasmid DNA was purified and transformed into Escherichia coli strain, BL21 (DE3) (Novagen, Madison, WI). The expression of BLpET11 a-mutant hPGK was then selected from the derived transformants. Recombinant mutant protein expression was induced by 0.5 mM isopropyl-P-D-thiogalacto-pyranoside (IPTG), for 5 hours at 37°C. Bacteria were collected by centrifugation at 6000 rpm for 10 minutes and re-suspended in lysis buffer containing 50 mM Tris-HCI, 2 mM EDTA, 2 mM PMSF, 10 µM leupeptin, 100 µg/ml lysozyme and 50 µg/ml of DNAse, pH 8.0. Lysis buffer was used at a ratio of 50 ml/I of culture. Cells were lysed on ice for 40 minutes followed by gentle shaking for a further 30 minutes at 4°C. Cells were sonicated in the presence of 0.5% (v/v) Triton X-100 for 20 seconds and then allowed to cool for 1 minute this process was repeated five times. The sample was clarified by centrifugation 134 Chapter 7 - Plasmin Reduction by PGK is a Thiol-lndependent Process at 15,000 rpm for 20 minutes and the supernatant was dialysed twice in 2 litres HEPES buffer containing 20 mM HEPES, 0.14 M NaCl, 1 mM EDTA, pH 7.4. Table 7 .1. Primers for PGK mutants (base changes are underlined) Mutant Forward (F) and reverse (R) primers F 5'-GACACTGCCACTGCCGCTGCCAAATGGAACAC-3' (1202 to 1233) C379,380A R 5'-GTGTTCCATTTGGCAGCGGCAGTGGCAGTGTC-3' (1233 to 1202) F 5'-GCCACTTCTAGGGGCGCCATCACCATCATAGG-3' (1163 to 1194) C379,380,367A R 5' -CCTATGATGGTGATGGCGCCCCTAGAAGTGGC-3' (1194 to 1163) F 5'-GGATGGGCTTGGACGCTGGTCCTGAAAGCAG-3' (1011 to 1041) C379,380,367,316A R 5' -CTGCTTTCAGGACCAGCGTCCAAGCCCATCC-3' (1041 to 1011) F 5'-GAAGTGGAGAAAGCCGCTGCCAACCCAGCTGC-3' (386 to 417) C379,380,367,316, 1OBA R 5'-GCAGCTGGGTTGGCAGCGGCTTTCTCCACTTC-3' (417 to 386) F 5'-CTGTTCTTGAAGGACGCTGTAGGCCCAGAAG-3' (359 to 389) C379,380,367,316, 108,99A R 5'-CTTCTGGGCCTACAGCGTCCTTCAAGAACAG-3' (389 to 359) F 5' -CCAAGCATCAAATTCGCCTTGGACAA TGGAGC-3' (212 to 243) C379,380,367,316, 108,99,50A R 5' -GCTCCATTGTCCAAGGCGAA TTTGATGCTTGG-3' (243 to 212) F 5'-CCAAGCATCAAATTCGCCTTGGACAATGGAGC-3' (212 to 243) C50 PGK R 5' -GCTCCATTGTCCAAGGCGAA TTTGA TGCTTGG-3' (243 to 212) 135 Chapter 7 - Plasmin Reduction by PGK is a Thiol-lndependent Process 7 .2.3 Purification of mutant rhPGK Solid (NH4)2S04 was added to the dialysed sample to give a concentration of 25% and mixture stirred for 20 minutes at 4°C. The pellet was collected by centrifugation at 15,000 rpm for 20 minutes, then additional (NH 4)2S04 added to give a final concentration of 100%. After stirring for 20 minutes at 4°C the pellet was collected by centrifugation at 15,000 rpm for 20 minutes, dissolved in buffer containing 20 mM HEPES, 0.05 M NaCl, 1 mM EDTA, 0.02% (w/v) NaN 3, pH 7.4 and dialysed extensively against HEPES buffer. The solution was applied to Cibachron Blue-Sepharose column (80 ml, 2.5x17 cm) (Amersham Pharmacia Biotech, Uppsala, Sweden) equilibrated with HEPES buffer. The column was washed with three bed volumes of HEPES buffer at a flow rate of 0.5 ml/min until the baseline was reached. Bound proteins were eluted with a linear NaCl gradient of 0.05-2 M. rhPGK eluted at approximately 1 M NaCl. The eluate was dialysed against buffer containing 20 mM Tris-HCI, 150 mM NaCl, 0.5 mM EDTA, 1 mM OTT, pH 7.9 and applied to a Heparin-Sepharose column (5 ml column) (Amersham Pharmacia) equilibrated with the same buffer. The column was washed with 20 ml of Tris buffer at a flow rate of 0.5 ml/min, bound proteins were then eluted with a linear NaCl gradient from 0.15-1 M in Tris buffer. rhPGK eluted at approximately 0.8 M NaCl. The purified rhPGK was dialysed against 20 mM HEPES, 0.14 M NaCl, 1 mM EDTA, pH 7.4 and stored at -80°C until use. 136 Chapter 7 - Plasmin Reduction by PGK is a Thiel-Independent Process 7 .2.4 Quantitative analysis of free thiols in rhPGK rhPGK (10 µM) was incubated with DTNB (200 µM) in 0.1 M HEPES, 0.3 M NaCl, 1 mM EDTA, pH 7.0 buffer for up to 120 minutes at room temperature. The absorbance at 412 nm due to the formation of the TNB dianion was measured using a Molecular Devices Thermomax Plus (Palo Alto, CA) microplate reader. The extinction coefficient for the TNB dianion at pH 7.0 is 14,150 M-1cm-1 at 412 nm [213]. 7.2.5 Alkylation of rhPGK rhPGK (10 µM) was incubated with a 1, 2 or 4-fold molar excess of HgCli, bBBr or TT in buffer containing 0.1 M HEPES, 0.3 M NaCl, 1 mM EDTA, pH 7.0 for 60 minutes at room temperature. wt PGK (90 µM) was also incubated with a 1,000-fold molar excess of GSSG in HEPES buffer for 2 hours at room temperature and the unreacted GSSG was removed by dialysis against HEPES buffer. The remaining reactive thiols in the rhPGK were determined using DTNB as described earlier. 7 .2.6 Plasm in reduction at various conditions 7.2.6.1 pH Plasmin (2 µg/ml) was incubated with rhPGK (1 µg/ml) in buffers of different pH. The incubation buffers were (1) 50 mM MES, 0.125 M NaCl, 2 mM EDTA, pH 6.0, (2) 50 mM HEPES, 0.125 M NaCl, 2 mM EDTA, pH 7.0 or (3) 50 mM HEPES, 0.125 M NaCl, 2 mM EDTA, pH 8.0. 137 Chapter 7 - Plasmin Reduction by PGK is a Thiel-Independent Process 7 .2.6.2 Ionic strength Plasmin (2 µg/ml) was incubated with rhPGK (1 µg/ml) in HEPES/Tween buffer containing various concentrations of NaCl (0.14-2 M) for 30 minutes. The control reactions were plasmin or rhPGK incubated alone in HEPES/Tween of various NaCl concentrations. 7 .2.6.3 Temperature Plasmin (2 µg/ml) was incubated with rhPGK (1 µg/ml) in HEPES/Tween buffer for 30 minutes at temperature between 20 and 50°C. The control reactions at each temperature were plasmin or rhPGK incubated alone in HEPES/Tween. 7 .2.6.4 3-PG/ ATP-MgCl2 rhPGK was incubated with 3-PG (0-13 mM) and/or ATP (0-13 mM) and 1 mM MgCl2 in HEPES buffer containing 20 mM HEPES, 0.14 M NaCl, 0.05% (v/v)Tween-20, pH 7.4 (HEPES/Tween) for 10 minutes prior to addition of plasmin (2 µg/ml). The control reactions were plasmin or rhPGK incubated alone in HEPES/Tween buffer. 7.2.7 Assay for plasmin reduction Free thiols generated in plasmin/angiostatin were labelled with MPB (100 µM) for 30 minutes at 37°C, followed by quenching of the unreacted MPB with GSH (200 µM) for 10 minutes at 37°C. Unreacted GSH, and other free sulfhydryls in the system, were blocked with NEM (400 µM) for 10 minutes at 37°C. The plasmin kringle products were collected on 50 µI of 138 Chapter 7 - Plasmin Reduction by PGK is a Thiel-Independent Process packed lysine-Sepharose beads by incubation on a rotating wheel for one hour at room temperature. Beads were washed three times with HEPES/Tween, and eluted with 50 µI of 50 mM E-amino-caproic acid in HEPES/Tween. The plasmin reductase activity was determined by ELISA or Streptavidin-HRP blotting analysis as described earlier in Chapter 3. 139 Chapter 7 - Plasmin Reduction by PGK is a Thiel-Independent Process 7.3 RESULTS 7.3.1 Sequence confirmation of Cys mutations in PGK The presence of the desired mutations was confirmed by automated DNA sequencing. Mutation of thymidine and guanine bases to guanine and cytidine bases, respectively are shown in bold (Figure 7.2 A-C). (A) hPGKDNA x C379,380,367 A 1022 GACTGTGGTCCTGAAAGCAGCAAGAAGTATGCTGAGGCTGTCACTCGGGC 1071 I :: I I I : I I I : I : I : I I I I I I I I I I I I I I I I I I I I I I I I I I I I 1 GNNAGTGNTCC . .. . NTCCNCNAGAAGTATGCTGAGGCTGTCACTCGGGC 46 1072 TAAGCAGATTGTGTGGAATGGTCCTGTGGGGGTATTTGAATGGGAAGCTT 1121 I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I 47 TAAGCAGATTGTGTGGAATGGTCCTGTGGGGGTATTTGAATGGGAAGCTT 96 1122 TTGCCCGGGGAACCAAAGCTCTCATGGATGAGGTGGTGAAAGCCACTTCT 1 171 I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I 97 TTGCCCGGGGAACCAAAGCTCTCATGGATGAGGTGGTGAAAGCCACTTCT 146 Cys367-Ala 1172 AGGGGC GCATCACCATCATAGGTGGTGGAGACACTGCCACT GCTGTGC 1221 I I I I I I I I I I I I I I I I I I I I I 11 I 11111 111 I I I I I I I I I I I I 147 AGGGGCGCGATCACCATCATAGGTGGTGGAGACACTGCCACTGCCGCTGC 1 96 1222 CAAATGGAACACGGAGGATAAAGTCAGCCATGTGAGCACTGGGGGTGGTG 1271 I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I 197 CAAATGGAACACGGAGGATAAAGTCAGCCATGTGAGCACTGGGGGTGGTG 246 1272 CCAGTTTGGAGCTCCTGGAAGGTAAAGTCCTTCCTGGGGTGGATGCTCTC 1321 I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I 247 CCAGTTTGGAGCTCCTGGAAGGTAAAGTCCTTCCTGGGGTGGATGCTCTC 296 1322 AGCAATATTTAGTACTTTCCTGCCTTTTAGTTCCTGTGCACAGCCCCTAA 1371 I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I 11 I I I I I I I I 297 AGCAATATTTAGTACTTTCCTGCCATTTAGTTCCTGTGCACAGCCCCTAA 346 1372 GTCAACTTAGCATTTTCTGCATCTCCACTTGGCATTAG 1409 I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I 347 GTCAACTTAGCATTTTCTGCATCTCCACTTGGCATTAG 384 Figure 7.2. Sequence alignment of human PGK DNA against recombinant PGK mutants. (A) cDNA sequence of C379,380,367A rhPGK showing mutations at Cys 379,380 and 367, (B) C379,380,367,316A rhPGK showing mutations at Cys367 and 316 and (C) C379,380,367,316,108,99,50A rhPGK showing mutations at Cys 108, 99, 50 (base changes are in bold). 140 Chapter 7 - Plasmin Reduction by PGK is a Thiel-Independent Process (8) hPGKDNA x C367,316A 759 TGCTGGACAAAGTCAATGAGATGATTATTGGTGGTGGAATGGCTTTTACC 808 I I I I I I : I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I 474 TGCTGGNCAAAGTCAATGAGATGATTATTGGTGGTGGAATGGCTTTTACC 425 809 TTCCTTAAGGTGCTCAACAACATGGAGATTGGCACTTCTCTGTTTGATGA 858 I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I 424 TTCCTTAAGGTGCTCAACAACATGGAGATTGGCACTTCTCTGTTTGATGA 375 859 AGAGGGAGCCAAGATTGTCAAAGACCTAATGTCCAAAGCTGAGAAGAATG 908 I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I 374 AGAGGGAGCCAAGATTGTCAAAGACCTAATGTCCAAAGCTGAGAAGAATG 325 909 GTGTGAAGATTACCTTGCCTGTTGACTTTGTCACTGCTGACAAGTTTGAT 958 I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I 11 I I I I I I I I I I I I I 324 GTGTGAAGATTACCTTGCCTGTTGACTTTGTCACTGCTGACAAGTTTGAT 275 959 GAGAATGCCAAGACTGGCCAAGCCACTGTGGCTTCTGGCATACCTGCTGG 1008 I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I 27 4 GAGAATGCCAAGACTGGCCAAGCCACTGTGGCTTCTGGCATACCTGCTGG 225 Cys316-Ala 1009 CTGGATGGGCTTGGACTGTGGTCCTGAAAGCAGCAAGAAGTATGCTGAGG 1058 I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I 11 I 224 CTGGATGGGCTTGGAC TGGTCCTGAAAGCAGCAAGAAGTATGCTGAGG 175 1059 CTGTCACTCGGGCTAAGCAGATTGTGTGGAATGGTCCTGTGGGGGTATTT 1108 I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I 174 CTGTCACTCGGGCTAAGCAGATTGTGTGGAATGGTCCTGTGGGGGTATTT 125 1109 GAATGGGAAGCTTTTGCCCGGGGAACCAAAGCTCTCATGGATGAGGTGGT 1158 I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I 124 GAATGGGAAGCTTTTGCCCGGGGAACCAAAGCTCTCATGGATGAGGTGGT 75 Cys367-Ala 1159 GAAAGCCACTTCTAGGGGC~G ATCACCATCATAGGTGGTGGAGACACTG 1208 1111111111111111111 1111111111 111 11 11111 I: 7 4 GAAAGCCACTTCTAGGGGCGCOATCACCATCATAGGTGGTGAAANGGNNT 25 Figure 7.2. Sequence alignment of human PGK DNA against recombinant PGK mutants (continued). 141 Chapter 7 - Plasmin Reduction by PGK is a Th iel-I ndependent Process (C) hPGKDNA x PGKC379,380,367,316,108,99,50A 78 AAATGTCGCTTTCTAACAAGCTGACGCTGGACAAGCTGGACGTTAAAGGG 127 I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I 40 ATATGTCGCTTTCTAACAAGCTGACGCTGGACAAGCTGGACGTTAAAGGG 89 128 AAGCGGGTCGTTATGAGAGTCGACTTCAATGTTCCTATGAAGAACAACCA 177 I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I 90 AAGCGGGTCGTTATGAGAGTCGACTTCAATGTTCC TATGAAGAACAACCA 139 178 GATAACAAACAACCAGAGGATTAAGGC TGCTGTCCCAAGCATCAAATTC 227 I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I 140 GATAACAAACAACCAGAGGATTAAGGCTGCTGTCCCAAGCATCAAATTCG 189 228 GOTTGGACAATGGAGCCAAGTCGGTAGTCCTTATGAGCCACCTAGGCCGG 277 I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I 190 CCTT GGACAATGGAGCCAAGTCGGTAGTCCT TATGAGCCACCTAGGCCGG 239 278 CCTGATGGTGTGCCCATGCCTGACAAGTACTCCTTAGAGCCAGTTGCTGT 327 I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I 240 CCTGATGGTGTGCCCATGTCTGACAAGTACTCCTTAGAGCCAGTTGCTGT 289 328 AGAACTCAAATCTCTGCTGGGCAAGGATGTTCTGTTCTTGAAGGAC TGTG 377 I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I 290 AGAACTCAAATCTCTGCTGGGCAAGGATGTTCTGTTCTTGAAGGACGCTG 339 378 TAGGCCCAGAAGTGGAGAAAGCCTG GCCAACCCAGCTGCTGGGTCTGTC 427 I I I I I I I I I I I I I I I I I I I I I I I I I I 11111111111I11111111 I I 340 TAGGCCCAGAAGTGGAGAAAGCCGCTGCCAACCCAGCTGCTGGGTCTGTC 389 428 ATCCTGCTGGAGAACC TCCGCTTTCATGTGGAGGAAGAAGGGAAGGGAAA 477 I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I 390 ATCCTGCTGGAGAACCTCCGCTTTCATGTGGAGGAAGAAGGGAAGGGAAA 439 478 AGATGCTTCTGGGAACAAGGTTAAAGCCGAGCCAGCCAAAATAGAAGCTT 527 I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I 440 AGATGCTTCTGGGAACAAGGTTAAAGCCGAGCCAGCCAAAATAGAAGCTT 489 528 TCCGAGCTTCACTTTCCAAGCTAGGGGATGTCTATGTCAATGATGCTTTT 577 I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I 490 TCCGAGCTTCACTTTCCAAGCTAGGGGATGTCTATGTCAATGATGCTTTT 539 578 GGCACTGCTCACAGAGCCCACAGCTCCATGGTAGGAGTCAATCTGCCACA 627 I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I 540 GGCACTGCTCACAGAGCCCACAGCTCCATGGTAGGAG TCAATCTGCCACA 589 628 GAAGGCTGGTGGG .TTTTTGATG.AAGAAGGAGCTGAACTACTTTGCAAA 675 111 : 111111111 111111111 11111111111111111111111111 590 GAANGCTGGTGGGTTTTTTGATGAAAGAAGGAGCTGAACTACTTTGCAAA 639 676 GGCCTTGGAGAGCCCAGAGCGACCCTTCCTGGCCATCCTGGGCGGAGCTA 725 1111111111111111 : I 11111111 1111 Ill 111111111111 : 640 GGCCTTGGAGAGCCCANAACGACCCTT .CTGGGCATTCTGGG CGGAGCTN 688 726 AAGTTGCAGACAAGATCCAGCTCATCAATAATATGCTGGAC .AAAGTCAA 774 I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I : I 689 AAGTTGCAGACAAGATCCAGCTCATCAATAATATGCTGGACAAAAGTCNA 738 Figure 7.2. Sequence alignment of human PGK DNA against recombinant PGK mutants (continued). 142 Chapter 7 - Plasmin Reduction by PGK is a Thiel-Independent Process 7.3.2 Purification of PGK mutants The seven Cys in wt PGK were mutated to Ala in a cumulative fashion. The two fast-reacting Cys, Cys379,380, were mutated first and the resulting C379,380A PGK cDNA was then used to mutate Cys367 and so forth. The wt and mutant PGK's were expressed in E. coli and extracted with lysozyme. The extract was clarified by ammonium sulfate precipitation and the protein purified by affinity chromatography on Cibachron Blue-Sepharose and Heparin-Sepharose. The wt and mutant PGK's were homogeneous by SOS PAGE (Figure 7. 3). 200 120 80 60 50 40 ._PGK 30 10 5 1 2 3 4 5 6 7 8 Figure 7.3. SOS-PAGE profile ofwt and mutant PGK's. Five micrograms of wt and mutant PGK's were resolved on SOS-PAGE under non reducing conditions and stained with Coomassie Brilliant Blue. The positions of Mr markers in kDa are shown at left. Arrow indicates PGK. 143 Chapter 7- Plasmin Reduction by PGK is a Thiel-Independent Process 7 .3.3 Consequence of mutation of all seven PGK Cys residues to Ala for plasmin reductase activity Different concentrations of the wt and mutant PGK's (1-40 µg/ml) were incubated with plasmin and the thiols in the reduced plasmin/angiostatin were labelled with MPB as described previously. The control reactions were incubation of PGK or plasmin alone. These controls tested for any confounding effects of labelling of existing free thiols in plasmin or PGK by MPB. Both reactions resulted in negligible signal. The specific plasmin reductase activity of the wt and mutant PGK's were similar, with the exception of the Cys-less PGK (C379,380,367,316, 108,99,S0A PGK) (Figure 7.4 A). The C50A mutant also had similar specific activity as wt PGK (Figure 7.4 A). The same qualitative results were observed for all the PGK mutants when plasmin reduction was measured by resolving the MPS-labelled proteins on SOS-PAGE and blotting with Streptavidin-HRP to detect the labelled angiostatin fragments (Figure 7.4 8). 144 Chapter 7 - Plasmin Reduction by PGK is a Thiel-Independent Process A O wtPGK O C379,380A PGK D C379,380,367 A PGK . C379,380,367,316A PGK • C379,380,367,316, 108A PGK ~ C379,380,367,316,108,99A PGK O C379,380,367,316,108,99,50A PGK 0_7 O C50A PGK E r:: 0.6 II) ~ 0.5 -"'Cl) 0.4 () 1 ~m 111_~ ::0.0 ~ llt I 1 10 40 rPGK, µg.ml"1 B ~ 273 111 • 80 61 49 (I *angiostatin 36 25 19 13 1 2 3 4 5 6 7 8 9 Figure 7.4. Consequence of all seven PGK Cys mutation on plasmin reductase activity. (A) Plasmin reductase activity of wt or mutant PGK's analysed by ELISA. The results have been corrected for the absorbance of the incubation of plasmin alone, which was negligible. Incubation of PGK alone also resulted in negligible absorbance. The bars are the mean and SO of triplicate determinations. (B) Samples in part A (10 µI) were resolved on SOS-PAGE under non-reducing cond itions and blotted with Streptavidin-HRP. The positions of Mr markers in kOa are shown at left. The arrows indicate the three angiostatin fragments. 145 Chapter 7 - Plasmin Reduction by PGK is a Thiol-lndependent Process 7 .3.4 Consequence of mutation of PGK Cys residues on the susceptibility of PGK to proteolysis Wild-type or mutant PGK's were incubated with plasmin and proteolysis of the PGK's examined by SOS-PAGE. wild-type PGK and the C379,380A, C379,380,367,316A, and C50A PGK mutants were resistant to plasmin proteolysis (Figure 7.5). In contrast, the C379,380,367A, C379,380,367,316, 108A, C379,380,367,316, 108,99A and C379,380,367,316, 108,99,50A PGK mutants were proteolysed by plasm in to different extents. In particular, the 6 and 7 Cys to Ala mutants were completely degraded by plasmin during the incubation. 200 120 80 60 50 40 ._PGK 30 --- - 10 5~------' 1 2 3 4 5 6 7 8 Figure 7.5. Consequence of mutation of PGK Cys residues on the susceptibility of PGK to proteolysis. wt or mutant PGK's (0.5 mg/ml) were incubated with plasmin (0.5 mg/ml) in HEPES/Tween-buffered saline for 60 minutes. Samples of the reactions (20 µI) were resolved on SOS-PAGE under non-reducing conditions and stained with Coomassie Brilliant Blue. The positions of Mr markers in kDa are shown at left. 146 Chapter 7 - Plasmin Reduction by PGK is a Thiel-Independent Process 7 .3.5 Quantitative analysis of Cys379,380 in PGK The two fast-reacting Cys of wild-type PGK were titrated by DTNB. As expected, the C379,380A mutant, which lacks the fast-reacting thiols, did not appreciably react with DTNB (Figure 7.6). 3 --e- Wild-type -0-- C379,380A 2 2' C) ....Ill.. -~ ....,n 1 0 20 40 60 80 100 120 Time (Min) Figure 7.6. Titration of the fast-reacting thiols of wild-type or C379,380A PGK. PGK (10 µM) was incubated with DTNB (200 µM) for up to 120 minutes at room temperature. The absorbance at 412 nm due to the formation of the TNB dianion was measured using a microplate reader. The extinction coefficient for the TNB dianion at pH 7.0 is 14,150 M-1cm-1 at 412 nm. 147 Chapter 7 - Plasmin Reduction by PGK is a Thiel-Independent Process 7 .3.6 Effect of alkylation of the fast-reacting Cys of PGK on plasmin reductase activity The involvement of the PGK fast-reacting thiols in plasmin reduction was tested by modifying them with various alkylating/oxidising reagents. Reaction of PGK with HgCl2, bBBr, TT and GSSG reduced the number of reactive thiols per mol of PGK to -0.2-0.5 (Figure 7.7 A-D). Equimolar concentrations of HgCl2, bBBr and TT were maximally effective, as four-fold molar excess of these agents did not change the number of residual thiols in PGK. Incubation of PGK with a 1,000-fold molar excess of GSSG reduced the number of reactive thiols per mol of PGK to -0.4. The alkylated/oxidised PGK's were tested for plasmin reductase activity using ELISA as described previously. The activity of the modified proteins were reduced to 7-35% of control (Figure 7.8). 148 Chapter 7 - Plasmin Reduction by PGK is a Thiel-Independent Process A B 2.5 3 2.0 52' 52' C) C) 2 0. 1.5 0. I::' I::'.... :c :c e 1.0 e 1 0.5 0.0 0 0 1 2 3 4 0 1 2 3 4 C [HgClzl/[PGK] D [bBBr]/[PGK] 2.5 2.0 --+-- NII 2.0 52' 1.5 C) 52' 0. 1.5 C) ;::: 0. :c !Si: 1.0 e 1.0 :c e 0.5 0.5 0.0 0 2 4 6 8 0.0 [TT]/[PGK] 0 20 40 60 80 100 Time,min Figure 7.7. Alkylation of the fast-reacting thiols in rhPGK. wt PGK (10 µM) was incubated with a 1, 2 or 4-fold molar excess of HgCl2 (A), bBBr (B) or TT (C) for 1 hour and the moles of thiols per mol of PGK calculated using DTNB. PGK (90 µM) was also incubated with a 1,000-fold molar excess of GSSG (D) and the moles of thiols per mol of PGK calculated using DTNB after removal of the unreacted GSSG by dialysis. 149 Chapter 7 - Plasmin Reduction by PGK is a Thiol-lndependent Process The alkylated/oxidised PGK's were incubated with plasmin in HEPES/Tween-buffered saline for 30 minutes at 37°C and the thiols in the reduced plasmin/angiostatin were labelled with MPB. The MPS-labelled plasmin/angiostatin fragments were immobilised on wells coated with monoclonal antibody against angiostatin and the MPB detected using Streptavidin-HRP. The activity of the modified proteins was reduced to 7- 35% of control (Figure 7.8). 1.0 E C 0.8 It) 0 "It CU 0.6 -G) u C .a...CU 0.4 0en .a <( 0.2 0.0 1•• ::II:: N ... (!) (!) - III I= rn c3 III Q, Cl + rn .c (!) ::c + ::II:: + (!) + ::II:: Q, ::II:: (!) ::II:: (!) Q, (!) Q, Q, Figure 7.8. Effect of alkylation of the fast-reacting Cys of PGK on plasmin reductase activity. rhPGK's (1 µg/ml) alkylated by various reagents (20 µM) were incubated with plasmin (2 µg/ml) and assayed for plasmin reductase activity using ELISA. The bars are the mean and SD of triplicate determinations. 150 Chapter 7 - Plasmin Reduction by PGK is a Thiel-Independent Process 7.3.7 Effect of pH, ionic strength and temperature on the plasmin reductase activity of PGK The plasmin reductase activity of rhPGK was optimal at pH 7 in the first 30-minutes of incubation. There was no obvious effect, however, of pH on plasmin reduction after 60 minutes incubation (Figure 7.9 A). Increasing NaCl concentrations reduced plasmin reduction, although the effects were relatively modest. Reductase activity was reduced by 73% when the NaCl concentration was increased from 0 to 2 M (Figure 7.9 B). There was no substantial effect of temperature on plasmin reductase activity of PGK between 20 and 50°C (Figure 7.9 C). 151 Chapter 7 - Plasmin Reduction by PGK is a Thiel-Independent Process A 0.6 -pH& c::::::J pH 7 e o.5 ml!!!lpH8 C :g 0.4 "Ill' -Cl) 0.3 CJ C Cl .a.. 0.2 0 .aU) cC 0.1 1~1 0.0 _o_ J_ l~I 5 10 20 40 60 Time,min B 0.5 -E 0.4 C II) 0 "Ill' 0.3 -Cl) CJ C 0.2 .a..Cl 0 .aU) 0.1 cC 0.0 I 0 0.14 0.5 1 1.5 2 NaCl,M C 0.8 -E c 0.6 II) 0 "Ill' -~ 0.4 C .a..Cl 0 U) 0.2 .a cC 0.0 20 25 30 35 37 40 45 50 Temperature, °C Figure 7.9. Effect of pH, ionic strength and temperature on the plasm in reductase activity of PGK. 152 Chapter 7 - Plasmin Reduction by PGK is a Thiol-lndependent Process 7 .3.8 Effect of 3-PG and ATP-induced conformational change in PGK on the plasmin reductase activity The reactivity of the fast-reacting Cys in pig muscle PGK is reduced upon binding of 3-PG and/or MgATP to PGK [214]. This occurs as a result of substrate induced conformational changes in the enzyme [179]. The consequences of substrate-induced conformational changes in rhPGK for plasmin reductase activity were examined. Incubation of rhPGK with 1 mM 3-PG or MgATP reduced plasmin reductase activity by -60%, while incubation with 1 mM 3-PG and MgATP reduced activity by -90% (Figure 7.10 A). Titration of the inhibitory effects of 3-PG and MgATP on plasmin reductase activity is shown in Figures 7.10 B and C. The half-maximal inhibitory effects of 3-PG and MgATP on plasmin reductase activity were -1 mM. 153 Chapter 7 - Plasmin Reduction by PGK is a Thiel-Independent Process A 0.8 E C in 0.6 0... -CV QIu 0.4 C CV .D... 0 Ill .D 0.2 < 0.0 I C) a. a. z a. I- I- I c( c( M - c:n c:n ~ ~ ~ ....E ~ all ....E C) a.I B M C ~ E 0.8 .... 0.8 i E E C C in 0.6 in 0.6 ...0 ...0 CV CV -QI - u 0.4 ~ 0.4 C C CV .D... !... 0 0 Ill Ill .D 0.2 .D 0.2 < < • • 0.0 0.0 • 0 2 4 6 8 10 12 14 0 2 4 6 8 10 12 14 3-PG,mM MgATP,mM Figure 7.10. Effect of 3-PG and ATP-induced conformational change in PGK on the plasmin reductase activity. PGK (1 µg/ml) was incubated with plasmin (2 µg/ml) in the absence or presence of 3-PG (0-13 mM) and/or ATP (0-13 mM) and 1 mM MgCl2 in HEPES/Tween-buffered saline for 30 minutes. Free thiols in the reduced plasmin/angiostatin were labelled with MPB and assayed using ELISA. (A) the effect on plasmin reduction by 1 mM 3-PG and/or ATP/MgCl2, (B) and (C) the concentration dependence of the 3-PG or ATP/MgCl2 effect, respectively. The bars are the mean and SO of triplicate determinations. 154 Chapter 7 - Plasmin Reduction by PGK is a Thiel-Independent Process 7 .4 DISCUSSION Protein reductant active sites typically contain a redox active dithiol/disulfide with the sequence, CysGlyXCys [215]. The Cys thiols cycle between the reduced dithiol and oxidised disulfide bond in coordination with a dithiol or disulfide of a protein substrate. This can result in reduction, formation or interchange of disulfide bonds in the protein substrate. There are seven Cys in PGK, none of which are involved in disulfide bonds, and only two of the seven are nearby in the primary or tertiary structure [175, 216- 218]. This suggests that the mechanism by which PGK reduces disulfide bonds in plasmin is unconventional. I have reported that the plasmin reductase activity of PGK is inhibited by NEM and iodoacetamide [200], which implies a role for one or more of the PGK Cys residues in plasmin reduction. In this study I have explored the role of all 7 PGK Cys, and in particular the two fast-reacting Cys, in reduction of plasmin disulfide bonds. The seven Cys in PGK were mutated to Ala in a cumulative fashion in the order, Cys379, 380, 367, 316, 108, 99 and 50. The two fast-reacting Cys, Cys379,380, were mutated first and the resulting cDNA was then used to mutate Cys367, and so forth. The specific plasmin reductase activity of the mutant PGK's, with the exception of the Cys-less PGK, was similar to that of the wild-type protein. This result suggested that Cys50, which was the last Cys to be mutated, was required for plasmin reductase activity. To test this hypothesis, the C50A PGK mutant was made and assayed for plasmin reductase activity. This C50A mutant had similar specific activity as wt PGK (Figure 7 .4 A). The same qualitative results were observed for all the PGK mutants when plasmin reduction was measured by resolving the MPB- 155 Chapter 7 - Plasmin Reduction by PGK is a Thiol-lndependent Process labelled proteins on SOS-PAGE and blotting with Streptavidin-HRP to detect the labelled angiostatin fragments (Figure 7.4 B). These results indicated that neither Cys50 nor any of the other PGK Cys were required for plasmin reduction. I hypothesised that the Cys-less PGK had lost plasmin reductase activity due to secondary effects of the mutations on the integrity of the PGK tertiary structure. This theory was tested by examining the susceptibility of the mutant PGK's to proteolysis by plasmin. Some mutations were shown to change the tertiary structure of PGK as measured by susceptibility to proteolysis by plasmin. For instance, the 6 and 7 Cys-less PGK was rapidly degraded by plasmin, which probably accounted for their significant decrease in plasmin reductase activity (Figure 7.5). These results implied that PGK Cys were not directly involved in plasmin reduction. The question remained, therefore, why alkylation of Cys379,380 inhibited plasmin reduction. This question was explored by reacting the Cys residues with different alkylating/o:xidising agents and examining the consequences for plasmin reductase activity. Carboxymethylation, but not methylation, of the fast-reacting Cys of pig PGK inactivates the kinase activity [219]. Reaction of the pig enzyme with DTNB also inactivates the kinase activity [212], while reaction with HgCl 2 or bBBr reduces kinase activity by up to 80% [220]. HgC'2 reacts with free thiols and can facilitate oxidation of a dithiol to a disulfide bond. bBBr is a homobifunctional alkylating agent that can cross-link thiols in close proximity. The fast-reacting thiols of PGK were also reacted with TT and GSSG. Accessible thiols react with these compounds to form mixed disulfides with thiosulfate and glutathione, respectively. Reaction of PGK with all four 156 Chapter 7 - Plasmin Reduction by PGK is a Thiol-lndependent Process alkylating/oxidising agents reduced the number of reactive thiols per mol of PGK to -0.2-0.5. The plasmin reductase activity of the modified proteins was reduced to 7-35% of control (Figure 7.8). These results indicate that alkylation of the fast-reacting thiols perturb plasmin reductase activity, which is consistent with our earlier report of inhibition of plasmin reductase activity by NEM and iodoacetamide [200]. Neither changes in pH, ionic strength nor temperature markedly affected plasmin reductase activity, which implies that the reaction of PGK and plasmin involved predominantly hydrophobic interactions. Kinase substrate binding studies have shown that MgATP and MgADP bind to the inner surface of the C-domain [178-180, 218) while 3- phosphoglycerate binds to the inner surface of the N-domain [175,217,218]. The bound substrates are c: 10 A from each other, too great a distance for direct in-line phosphoryl transfer. The enzyme overcomes this distance by undergoing a 'hinge-bending' conformational change that brings the two substrates closer together. The reactivity of both the fast-reacting thiols is reduced by binding of 3-PG and/or MgADP and MgATP to PGK [214]. Moreover, carboxyamidomethylation of the two fast-reacting Cys in pig PGK blocks the substrate-induced 'hinge-bending' conformational change in the enzyme [221]. These observations indicate that alkylation of the fast-reacting thiols in PGK can perturb the conformational changes required for kinase activity. I tested the effect of 3-PG/MgADP-induced conformational changes in PGK for plasmin reductase activity. Incubation of PGK with 1 mM 3-PG or MgATP reduced plasmin reductase activity by -60%, while incubation with 1 157 Chapter 7 - Plasmin Reduction by PGK is a Thiol-lndependent Process mM 3-PG and MgATP reduced activity by -90% (Figure 7.10 A). The half maximal effects of 3-PG and MgATP on plasmin reductase activity were -1 mM (Figure 7 .10 B and C), which is in the range of the Michaelis constants for these substrates in the kinase reaction [222]. It is noteworthy that the effects of 3-PG and MgATP were additive, which may reflect cooperation between 3-PG and MgATP in the conformational change in PGK. In summary this study demonstrated that none of the Cys residues in PGK were required for plasmin reduction. This finding implies that the mechanism by which PGK reduces disulfide bonds in plasmin is a thiol independent process. However, alkylation/oxidation of the fast-reacting Cys or hinge-bending conformational changes in the same region of PGK negates the plasmin reductase activity perhaps by obstructing the binding of plasmin to PGK or perturbing conformational change in PGK required for plasmin reduction. 158 CHAPTERS GENERAL DISCUSSION AND CONCLUSIONS Chapter 8 - General Discussion and Conclusions 8.1 GENERAL DISCUSSION AND CONCLUSIONS Angiogenesis is a complex biological process consisting of a series of events. These cellular events can be classified into three major phases: (i) initiation phase, involving the release of growth factors/cytokines from tumour and inflammatory cells, such as macrophages and mast cells, which subsequently activate endothelial cells [223, 224] (ii) invasion/proliferation phase characterised by an increased production of cell adhesion molecules and matrix degrading enzymes, which proteolyse BM and the ECM to provide a permissive microenvironment for endothelial cell proliferation and migration; and (iii) maturation/differentiation phase involving cell-cell and cell ECM interaction, which leads to the formation of capillary sprouts. Normally, angiogenesis is tightly regulated. A process that can be switched on and off for a brief period of time in response to a local angiogenic stimulus. In some pathological conditions however, angiogenesis persists for a prolonged period of time leading to tissue destruction and promotion of tumour progression. Angiogenesis is crucial for tumour growth beyond a critical size. An increase in tumour mass is accompanied by an increase in their metabolic requirements. In response to such demands, tumour cells produce angiogenic factors or influence the surrounding cells to induce angiogenesis. Interestingly, these tumour cells also produce angiogenic inhibitors. One such inhibitor is angiostatin, a potent inhibitor isolated from plasma of mice bearing 3LL Lewis lung carcinoma [46]. A local balance between angiogenic activators and inhibitors maintains endothelial cell quiescence. This balance will shift in favour of 159 Chapter 8 - General Discussion and Conclusions angiogenesis activation when reduced production of angiogenic inhibitors or an over expression of angiogenic factors ensues. Angiostatin is a proteolytic product of the zymogen, plasminogen. A number of human tumours have been shown to cleave plasmin forming angiostatin. Various members of the serine proteinase and matrix metalloproteinase family have been implicated in angiostatin formation by tumours. It appears that different types of tumours express different enzymatic activity for angiostatin formation. Previous studies in this laboratory have shown that angiostatin formation from plasmin involved reduction of one or more disulfide bonds in kringle 5 of plasmin by a tumour secreted enzyme, plasmin reductase. The reduced plasmin then undergoes proteolytic cleavage by a serine proteinase, which is also secreted by tumour cells producing thiol-containing angiostatin fragments. This thesis described the isolation and characterisation of plasmin reductase from the conditioned medium of cultured human fibrosarcoma HT1080 cells. Plasmin reductase was found to have 100% sequence identity to the glycolytic enzyme, phosphoglycerate kinase (PGK). To further confirm the finding, recombinant human PGK was cloned and its plasmin reductase activity examined. The purified recombinant protein had the same molecular mass and specific activity as the HT1080 derived PGK, supporting the proposal that plasmin reductase is PGK. A significant increase in plasma level of PGK was observed in mice bearing HT1080 tumours compared to mice without tumours, suggesting that PGK was secreted by both cultured and implanted HT1080 cells. To investigate the function of rhPGK in angiostatin formation in vivo, mice 160 Chapter 8 - General Discussion and Conclusions bearing HT1080 tumours were administered with rhPGK and the effect on plasma levels of angiostatin before and after PGK administration was measured. The results indicated that systemic administration of rhPGK to tumour-bearing mice up-regulated plasma levels of angiostatin 2-hr post PGK administration. In contrast, no difference in plasma levels of angiostatin was observed in mice without tumours. This finding was consistent with the hypothesis that PGK is the rate limiting step in the formation of angiostatin from plasmin by tumour cells. The finding that systemic administration of rhPGK significantly elevated plasma levels of angiostatin led to the investigation of the effect of administration of rhPGK on tumour angiogenesis and tumour growth. Systemic administration of rhPGK caused a significant inhibition in the rate of HT1080 (-50%) and AsPC-1 (-70%) tumour growth and a marked reduction in microvessel density. It is interesting that AsPC-1 tumours was more susceptible to suppression by systemic administration of PGK than HT1080 tumours. This may reflect the 5-fold reduced secretion of PGK by AsPC-1 cells compared to HT1080 cells. These studies showed that PGK has an additional function in biology by triggering the extracellular production of angiostatin from plasmin. The mechanism of plasmin reduction by PGK was also explored. The active sites of most protein reductases are characterized by the presence of two closely spaced cysteine residues typically in the consensus sequence, CysGlyXxxCys. The cysteine thiols cycle between the reduced dithiol and oxidised disulfide bond in coordination with a disulfide or dithiol of a protein substrate. This can result in reduction, formation or interchange of the 161 Chapter 8 - General Discussion and Conclusions disulfide bonds in protein substrates. The fact that PGK lacks this motif suggests that the mechanism of plasmin reduction by PGK is not conventional. To explore the role of Cys residues in plasmin reduction, all seven Cys in PGK were mutated to Ala in a cumulative fashion in the order, Cys379, 380, 367, 316, 108, 99 and 50. The specific plasmin reductase activity of the mutant PGK's, with the exception of the Cys-less PGK, was similar to that of the wild-type protein. Interestingly, some mutations induced conformational changes in PGK and rendered them susceptible to proteolysis by plasmin. Cys-less PGK was rapidly degraded by plasmin which presumably accounted for the loss of plasmin reductase activity. These results implied that the PGK Cys were not directly involved in plasmin reduction indicating plasmin reduction by PGK is a thiol-independent process. However, the question remained, why alkylation of thiols inhibited plasmin reduction. This question was explored by reacting Cys residues with different alkylating/oxidising agents and examining the consequences for plasmin reductase activity. Alkylation/oxidation of the two fast-reacting Cys in PGK significantly attenuated plasmin reductase activity. These results indicate that alkylation of the fast-reacting Cys perturb plasmin reductase activity, perhaps by obstructing the binding of PGK to plasmin or perturbing conformational changes in PGK required for plasmin reduction. Further evidence of conformational !ability of the plasmin reductase activity of PGK was provided by substrate binding studies. The effect of 3- PG/MgADP-induced conformational changes in PGK for plasmin reductase activity was examined. Incubation of PGK with 1 mM 3-PG or MgATP 162 Chapter 8 - General Discussion and Conclusions reduced plasmin reductase activity by -60%, while incubation with 1 mM 3- PG and MgATP reduced activity by -90%. These findings indicate that hinge-bending conformational change in PGK negates the plasmin reductase activity of the protein. Furthermore, plasmin reductase activity was not affected by changes in pH, ionic strength or temperature, which implies that the reaction of PGK and plasmin involved predominantly hydrophobic interactions. A model of the molecular events facilitated by PGK in plasmin kringle 5 is shown in Figure 8.1. We have previously proposed that both the Cys462-Cys541 and Cys512-Cys536 disulfide bonds in plasmin kringle 5 are cleaved which renders kringle 5 susceptible to proteolysis at either the Arg530-Lys531 peptide bond or two other unidentified peptide bonds [187). The other peptide bonds are likely to be Arg474-Val475 [187] and Lys467- Gly468 [225]. Plasmin undergoes autoproteolysis at alkaline pH producing a catalytically active microplasmin fragment with a Lys531 N-terminus. Wu and co-workers [226, 227] noticed that both the Cys462-Cys541 and Cys512- Cys536 disulfide bonds in K5 must have been cleaved to release microplasmin from K1-4 and they proposed that the increased -oH ion concentration at alkaline pH was responsible for cleaving the Cys462-Cys541 disulfide bond. We have suggested that the mechanism of plasmin proteolysis at alkaline pH is the same as that facilitated by PGK at neutral pH [187). A method for calculating the strain of a disulfide bond uses dihedral angles. A dihedral angle is the angle of rotation about a certain bond. The 163 Chapter 8 - General Discussion and Conclusions dihedral strain energy of a disulfide bond can be calculated from the five dihedral angles of a disulfide bond as described by Weiner et al. [229] and Katz and Kossiakoff [228]. The calculated strain energies only consider the dihedral angles of the disulfide bond and do not include other factors such as bond lengths, bond angles, and van der Waals contacts in calculating energy. The findings of Pjura et al. [232], however, indicate that such calculations can give useful semi-quantitative insights into the amount of strain in a disulfide bond. There is a correlation between the redox potentials of disulfide bonds and their calculated dihedral energies. Those bonds with higher dihedral energies are more easily reduced [228, 229]. Calculation of the dihedral strain energies of the kringle 5 disulfide bonds from the crystal structure described by Chang et al. [230] revealed that the left-handed Cys512-Cys536 bond has a high strain energy (2.63 kcal.mor1) compared to the right-handed Cys462-Cys541 (1.54 kcal.mor1) and left-handed Cys483-Cys524 (0.98 kcal.mor1) disulfide bonds. The average dihedral strain energies for left- and right-handed disulfide bonds are 1.68 and 3.19 kcal.mor1, respectively [228, 229]. These calculations led us to propose, that PGK facilitates cleavage of the Cys512-Cys536 disulfide bond by hydroxide ions, which results in formation of a sulfenic acid at position 512 and a free thiol at Cys536. The Cys536 thiol is then available to exchange with the Cys462-Cys541 disulfide bond resulting in formation of a new disulfide at Cys536-Cys541 and a free thiol at Cys462. Kringle 5 is then susceptible to proteolysis at Arg530- Lys531, Arg474-Val475 and/or Lys468-Gly469. This is the simplest 164 Chapter 8 - General Discussion and Conclusions sequence of events that can explain all the available data. For instance, cleavage of only the Cys512-Cys536 would not enable release of the kringle 1-4 angiostatin fragments from plasmin. The free thiol that is labelled by MPB in the three angiostatin fragments would be Cys462. Cleavage of the Cys483-Cys524 disulfide bond is not excluded, although this is not required to explain the experimental observations. PGK presumably binds to plasmin and induces a conformational change in kringle 5 which facilitates hydroxide ion attack on the Cys512- Cys536 disulfide bond. This suggests that other molecules that interact with plasmin might also facilitate cleavage of the kringle 5 disulfides. There is recent evidence to support this hypothesis. Interaction of a truncated porcine plasminogen activator inhibitor-1 (residues 80-265), but not full-length protein, with plasmin has been shown to result in generation of kringle containing angiostatin fragments [231]. It is possible that the truncated protein is facilitating the same sequence of events in plasmin that are achieved by PGK. The results of my studies and those of others support the following mechanism of angiostatin formation by tumours. tumour tumour tumour tumour MMP-12 MMP-9• l l l MMP-3 plaamln ,..P:?)r K1-4 uPA PGK reduced s.,protetnue K1-4½ / plasminogen --• plasmin + • plasmin • 2 ""-.. MMP4"& K1-3 MMP-12 \., l V blood angiostatin 165 Chapter 8 - General Discussion and Conclusions The sequential order of events leading to angiostatin formation begins with the conversion of plasminogen to plasmin then reduction of plasmin by PGK. This is followed by serine proteinase-dependent release of kringle 1-4½ [143, 187, 190, 191] and finally matrix metalloproteinase-dependent trimming of kringle 1-4½ to either kringle 1-4 or 1-3 [145, 149, 188]. An advantage of this mechanism is the opportunity for fine control of angiostatin formation at the level of both plasmin formation and plasmin reduction. Tumour production of angiostatin, therefore, will be controlled by production and secretion of both urokinase plasminogen activator and PGK. 166 Chapter 8 - General Discussion and Conclusions /-OH s s i s 847 s 1~I I.j ! I 5i T I 462 483 512 524 536 541 Kringle 5 462 483 512 524 536 541 so· ! 462 483 512 524 536 541 Figure 8.1. Model of the molecular events facilitated by PGK in plasmin kringle 5. Numbering is based on the sequence of human plasminogen (791 residues) beginning at Glu1, excluding the 19 amino acid signal peptide that ends at Met. 167 Chapter 8 - General Discussion and Conclusions Future Directions The precise mechanism of plasmin reduction facilitated by PGK is still not known. Study in our laboratory is currently underway to detect and map the sequence of cleavage events in kringle 5 by mass spectrometry analysis. The formation of sulfenic acid is being assessed as well as the role of hydroxide ion in the cleavage. One intriguing possibility is that the disulfide-bond cleavage is not mediated by hydroxide ion. We are exploring a mechanism whereby a negative charge (Asp/Glu) close to the kringle 5 Cys512-Cys536 disulfide bond and a positive charge (Lys) from PGK or plasmin polarizes and then breaks the disulfide bond. Brandt et al. [233] used semi-empirical quantum chemical calculations as well as ab initio studies to show that the interaction of the carboxylic acid and a primary amine with a disulfide bond can polarize and cleave the bond. A similar sequence of events leading to disulfide reduction may also occur in plasmin catalysed by PGK. Therefore, studies investigating the role of the Asp or Glu located near the kringle 5 Cys512-Cys536 will provide a better understanding of the mechanism of PGK catalysed-reduction of disulfide bonds in plasmin. 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