complement regulation on vascular endothelial cells ...€¦ · 2.3.8.2 bioflux (fluxion...
TRANSCRIPT
Complement Regulation on Vascular Endothelial
Cells – Insights into the Pathogenesis of Thrombotic
Microangiopathy
by
Damien Noone
A thesis submitted in conformity with the requirements
for the degree of Master of Science
Institute of Medical Science
University of Toronto
© Copyright 2015 by Damien G. Noone
II
Complement Regulation On Vascular Endothelial Cells – Insights
Into The Pathogenesis Of Thrombotic Microangiopathy
Damien Noone
Master of Science
Institute of Medical Science
University of Toronto
2015
Abstract
Atypical hemolytic uremic syndrome (aHUS) is a form of thrombotic
microangiopathy (TMA) that occurs due to defective regulation of the alternative
complement pathway (AP). We developed a novel model system using Blood
Outgrowth Endothelial Cells (BOEC), whereby the response of these cells to
complement challenge could be examined under static and microfluidic conditions, in
order to study aHUS pathogenesis. Complete blockade of the membrane-anchored,
AP regulator CD46/MCP, associated with disease in patients, was insufficient to
cause an ex vivo TMA phenotype. Increasing „complement challenge‟, mimicking
additional genetic „hits‟ in complement regulation achieved the phenotype. In addition,
using BOEC from patients lacking von Willebrand Factor (vWF), a hemostatic
protein released from activated endothelial cells, we showed that contrary to current
opinion, vWF does not amplify complement, rather regulates it. The paucity of vWF
in the glomerular endothelium could explain the vulnerability of the kidney to loss of
complement control in aHUS.
III
Acknowledgements
I would like to express my sincerest appreciation and heartfelt gratitude to Dr
Christoph Licht who has been my supervisor, mentor and career advisor. You have
been considerably greater than the sum of those parts. I am indebted to you for your
unflinching support. It will be an absolute honor, to be your colleague at The Hospital
for Sick Children and I hope that you will continue as my mentor for the rest of my
career.
I wish to thank Dr Lisa Robinson for her support, direction, constructive feedback on
how to communicate scientific information, and most especially, for helping me in
my transition from clinician to academic clinician.
I wish to thank Dr Philip Marsden for taking interest in my research and for asking
the tough, probing questions that undoubtedly have made me, and this work, a great
deal stronger.
I would like to especially thank Dr Magdalena Riedl. You have been a tremendous
help and guidance over the past year. Your work ethic is awe-inspiring.
I would also like to thank Dr Fred G Pluthero. Your scientific acumen and insight,
imaging skills, general knowledge and scrabble skills have always enlivened my
laboratory experience.
To Mackenzie Bowman and Dr Paula James in Kingston, Ontario, for your time and
patience in teaching me the technique of blood outgrowth endothelial cell culture.
To Kathy Liszewski and Dr John Atkinson in St Louis Missouri - for the gift of your
time, hospitality and knowledge I am indebted.
To the students of the Licht Laboratory who helped along the way – Lily Lu, Yi
Emma Quan and Steve Balgobin – thank you and good luck in your respective careers.
To Dr Norman Rosenblum, Dr Rulan Parekh and Dr Binita Kamath, members of my
Clinical Fellowship Research Advisory Committee at The Hospital for Sick Children,
for their advice and mentorship.
I owe my deepest gratitude to my wife. Thank you for your encouragement, your
endless sacrifice and love. Thank you for your acceptance of the uncertainties that
came with a medical career, two fellowships in two new countries and one Masters.
I dedicate this thesis to my three daughters, Jasmine, Molly and Annabelle.
I would like to acknowledge the following funding sources: Restracomp and The
Transplant Centre at The Hospital for Sick Children.
IV
Contributions
Damien Noone (author) solely prepared this thesis. All aspects of this body of work,
including the planning, execution, analysis, and writing of all original research and
publications was performed in whole or in part by the author. The following
contributions by other individuals are formally and inclusively acknowledged:
Dr. Christoph Licht (Primary Supervisor and Thesis Committee Member) –
mentorship; laboratory resources; guidance and assistance in planning, execution, and
analysis of experiments as well as thesis preparation
Dr. Lisa Robinson (Supervisor and Thesis Committee Member) – mentorship;
laboratory resources; guidance and assistance in planning, execution, and analysis of
experiments as well as thesis preparation
Dr. Philip Marsden (Thesis Committee Member) – mentorship; guidance in
interpretation of results as well as thesis preparation
Dr Magdalena Riedl - guidance and assistance in planning, execution, and analysis of
experiments for chapters 2 and 3
Dr Fred G Pluthero - guidance and assistance in planning, execution, and analysis of
experiments for chapters 2 and 3
Dr Walter Kahr - laboratory resources; guidance in interpretation of results
Annie Bang - guidance and assistance in planning, execution, and analysis of
experiments for chapter 2
Lily Lu - assistance in execution, and analysis of experiments for chapter 2
Yi (Emma) Quan - guidance and assistance in planning, execution, and analysis of
experiments for chapter 2
Steve Balgobin - assistance in execution, and analysis of experiments for chapters 2
and 3
V
Table of Contents
ACKNOWLEDGEMENTS………………………………………………………………….……….III
CONTRIBUTIONS………………………………………………………………………….………..IV
TABLE OF CONTENTS………………………………………………………………….……….….V
LIST OF TABLES…………………………………………………………………………………….IX
LIST OF FIGURES…………………………………………………………….…………………..….X
LIST OF ABBREVIATIONS…………………………………………………………………….…XII
Contents
CHAPTER 1. AN INTRODUCTION TO THROMBOTIC
MICROANGIOPATHY .............................................................................................. 1
1.1 Introduction ............................................................................................................ 3
1.2 Thrombotic microangiopathy ............................................................................... 4
1.2.1 Introduction to Thrombotic Microangiopathy ........................................................ 4
1.2.2 The Spectrum of Thrombotic Microangiopathy ..................................................... 4
1.2.3 Thrombotic Thrombocytopenic Purpura, VWF and ADAMTS13 ....................... 6
1.2.3.1 Introduction to TTP .............................................................................................. 6
1.2.3.2 von Willebrand Factor .......................................................................................... 6
1.2.3.3 ADAMTS13 ......................................................................................................... 7
1.2.4 Atypical Hemolytic Uremic Syndrome – a Complement-Mediated TMA ............ 8
1.2.4.1 A Brief Introduction of Hemolytic Uremic Syndrome – Typical and Atypical ... 8
1.2.4.2 The Complement System of Innate Immunity ...................................................... 9
1.2.4.3 Endothelial Cell Protection against Complement-Mediated Injury .................... 12
1.2.4.5 Complement-mediated „atypical‟ HUS............................................................... 13
1.2.4.6 CD46/MCP-associated atypical hemolytic uremic syndrome ............................ 14
1.2.5 From Mutation in CD46 to TMA – An Incomplete Story .................................... 15
1.2.5.1 Functional Studies of CD46/MCP Mutations ..................................................... 15
1.2.5.2 Incomplete Penetrance Suggests a Multiple „Hit‟ Concept ................................ 16
1.2.5.3 An Additional Complement Gene Aberrancy Increases Disease Penetrance ..... 16
1.2.5.4 A Link Between Complement and Coagulation ................................................. 17
1.3 Blood Outgrowth Endothelial Cells (BOEC) .................................................... 20
1.3.1 Introduction to BOEC ............................................................................................. 20
1.3.2 BOEC in the Study of Disease ................................................................................. 20
1.3.1 BOEC as a Model to Study TMA ........................................................................... 22
1.4 Knowledge Gap, Hypothesis and Thesis Aims .................................................. 23
VI
CHAPTER 2 MODELING CD46/MCP-ASSOCIATED ATYPICAL
HEMOLYTIC UREMIC SYNDROME USING BLOOD OUTGROWTH
ENDOTHELIAL CELLS .......................................................................................... 24
2.1 Abstract 25
2.2 Introduction .......................................................................................................... 26
2.3 METHODS ........................................................................................................... 28
2.3.1 BOEC Isolation ........................................................................................................ 28
2.3.2 BOEC Characterization By Flow Cytometry ........................................................ 28
2.3.3 BOEC Characterization By Immunofluorescence ................................................ 29
2.3.4 Surface Expression Of Membrane-Bound Regulators On BOEC ....................... 30
2.3.5 Quantitative Gene Expression Of Membrane-Bound Regulators On BOEC .... 31
2.3.6 Complement Challenge Of BOEC .......................................................................... 32
2.3.6.1 50% normal human serum in alternative pathway buffer ................................... 32
2.3.6.2 Membrane-anchored complement regulator blockade ........................................ 32
2.3.6.3 Complement deposition detected by immunofluorescence ................................ 32
2.3.6.4 Complement deposition detected by FACS ........................................................ 33
2.3.7 Cell death assays ....................................................................................................... 34
2.3.7.1 Trypan blue exclusion ......................................................................................... 34
2.3.7.2 Cell cytotoxicity/LDH assay ............................................................................... 34
2.3.7.3 Apoptosis ............................................................................................................ 34
2.3.8 Platelet Adhesion ...................................................................................................... 35
2.3.8.1 Platelet isolation .................................................................................................. 35
2.3.8.2 BioFlux (Fluxion Biosciences) Microfluidic System ......................................... 35
2.4 RESULTS 37
2.4.1 BOEC Have An Endothelial Cell Phenotype And Retain Their Phenotype Over
Various Passages ............................................................................................................... 37
2.4.2 BOEC Express Key Membrane-Anchored Complement Regulators ................. 39
2.4.3 Cell Surface C3b Deposition Mirrors Incremental Functional Blockade of
CD46/MCP, CD55/DAF and CD59 ................................................................................. 41
2.4.4 Complement-mediated BOEC cytotoxicity increases with incremental
membrane-regulator blockade ......................................................................................... 43
2.4.5 Increasing complement challenge to BOEC results in an increase in platelet
adhesion.............................................................................................................................. 45
2.4.6 Discussion .................................................................................................................. 48
2.4.7 Supplemental Data ................................................................................................... 51
VII
CHAPTER 3. VON WILLEBRAND FACTOR MULTIMERS CONTRIBUTE
TO COMPLEMENT ALTERNATIVE PATHWAY CONTROL ........................ 53
3.1 Abstract 54
3.2 Introduction .......................................................................................................... 56
3.3 Methods 58
3.3.1 Establishing and characterizing blood outgrowth endothelial cells (BOEC) ..... 58
3.3.2 Glomerular Endothelial Cell Culture ..................................................................... 58
3.3.3 Determining BOEC protein expression via Western blot .................................... 59
3.3.4 Quantifying BOEC and GEC Gene Expression via qRT-PCR ............................ 59
3.3.5 Characterization of membrane-bound regulators of BOEC ................................ 60
3.3.5.1 Immunofluorescence ........................................................................................... 60
3.3.5.2 Flow cytometry ................................................................................................... 61
3.3.6 Complement Challenge of BOEC ........................................................................... 62
3.3.6.1 Normal human serum.......................................................................................... 62
3.3.6.2 Membrane-anchored complement regulator blockade ........................................ 62
3.3.7 Assessment of Complement Challenge of BOEC .................................................. 63
3.3.7.1 Complement deposition detected by immunofluorescence ................................ 63
3.3.7.2 Complement deposition detected by FACS ........................................................ 63
3.3.8 Assessing the impact of complement challenge ..................................................... 64
3.3.8.1 LDH cell cytotoxicity assay ................................................................................ 64
3.3.8.1 Platelet adhesion Assay ...................................................................................... 64
3.3.9 Ethics ......................................................................................................................... 65
3.3.10 Statistical analysis .................................................................................................. 65
3.4 Results 66
3.4.1 BOEC Possess Endothelial Cell Characteristics ................................................... 66
3.4.2 Type 3 VWD BOEC Express Similar Amounts of Membrane-Anchored
Complement Regulators as Control BOEC .................................................................... 67
3.4.3 Complement activation products associate with VWF ......................................... 69
3.4.4 Platelet adhesion in response to complement challenge is initially dependent on
VWF release....................................................................................................................... 71
3.4.5 VWD BOEC Show Increased C3b Deposition after Complement Challenge .... 73
3.4.6 VWD BOEC Are More Vulnerable To Complement-Mediated Cytotoxicity .... 74
3.5 Discussion 76
3.6 Supplemental Data -VWF expression in glomerular endothelial cells ........... 79
VIII
CHAPTER 4. UNIFYING DISCUSSION, FUTURE DIRECTIONS AND
CONCLUSIONS ........................................................................................................ 81
4.1 Unifying Discussion .............................................................................................. 82
4.2 Future Directions ................................................................................................. 86
4.2.1 Introduction .............................................................................................................. 86
4.2.2 Do Endothelial Cells Mount a Cytoprotective Response when Challenged by
Complement? ..................................................................................................................... 87
4.2.2.1 Rationale and Hypothesis ................................................................................... 87
4.2.2.2 Methods .............................................................................................................. 88
4.2.2.3 Preliminary Results ............................................................................................. 88
4.2.2.4 Discussion ........................................................................................................... 90
4.2.3 How Does the Fluid Phase Regulator CFH Contribute to Endothelial Cell
Protection? ......................................................................................................................... 91
4.2.3.1 Rationale and Hypothesis ................................................................................... 91
4.2.3.2 Methods .............................................................................................................. 91
4.2.3.3 Preliminary Results ............................................................................................. 92
4.2.3.3 Discussion ........................................................................................................... 94
4.2.4 Will BOEC Isolated From a Patient Further our Understanding of TMA? ...... 95
4.2.4.1 Rationale, Hypothesis and Aims ......................................................................... 95
4.2.4.3 Proposed Methods and Anticipated Results ....................................................... 96
4.2.4.4 Anticipated Results and Implications ................................................................. 96
4.3 Conclusions ........................................................................................................... 98
References 100
IX
List of Tables
Table 1-1. Classification of Thrombotic Microangiopathies……………..……5
Table 1-2. Relevant Complement Regulators…………………………………14
X
List of Figures
Figure 1-1. Overview of complement activation ...................................................... 11
Figure 2-1. BOEC have an endothelial cell phenotype and retain their phenotype
over various passages ................................................................................................. 39
Figure 2-2. BOEC express key membrane-anchored complement regulators ..... 41
Figure 2-3. Cell surface C3b deposition mirrors incremental functional blockade
of CD46/MCP, CD55/DAF and CD59 ...................................................................... 42
Figure 2-4. Complement-mediated BOEC cytotoxicity increases with incremental
membrane-regulator blockade ................................................................................. 44
Figure 2-5. Increasing complement challenge to BOEC results in a stepwise
increase in platelet adhesion ..................................................................................... 46
Figure 2-S1. Functional blocking antibody binding and saturation ...................... 51
Figure 2-S2. Expression of surface-bound regulators on glomerular endothelial
cells compared to BOEC ........................................................................................... 52
Figure 3-1. BOEC possess endothelial cell characteristics ...................................... 67
Figure 3-2. Type 3 VWD BOEC express similar amounts of membrane-anchored
complement regulators as control BOEC ................................................................ 68
Figure 3-3. Complement challenge results in VWF release and the association of
complement activation products and VWF ............................................................. 70
Figure 3-4. Platelet adhesion in response to complement challenge is initially
dependent on VWF release ....................................................................................... 72
Figure 3-5. VWD BOEC show increased C3b deposition after complement
challenge...................................................................................................................... 73
XI
Figure 3-6. VWD BOEC are more vulnerable to complement-mediated
cytotoxicity .................................................................................................................. 75
Figure 3-S1. Glomerular endothelial cells express less VWF ................................. 79
Figure 4-1. VWF and ADAMTS13 as a Complement Regulatory System. ........... 85
Figure 4-2. Response of complement regulators to increasing complement
challenge...................................................................................................................... 90
Figure 4-3. Role of CFH-mediated surface protection against endothelial cell
death ............................................................................................................................ 93
Figure 4-4. Multiple genetic hits in complement and coagulation genes needed to
cause aHUS ................................................................................................................. 96
XII
List of Abbreviations
ADAMTS13 A Disintergrin and Metalloprotease with Thrombospondin-1 Motifs
Type 13
ACD Acid Citrate Dextrose
AHUS Atypical Hemolytic Uremic Syndrome
AP Alternative Pathway
BOEC Blood Outgrowth Endothelial Cell
CC3 Cleaved Caspase 3
CFH Complement Factor H
CFI Complement Factor I
DAF Decay Accelerating Factor/CD55
EC Endothelial Cell
FBS Fetal Bovine Serum
HEPES Hyroxyethyl Piperazineethanesulfonic
MCL Mononuclear Cell Layer
MCP Membrane Cofactor Protein
MFI Median Fluorescence Intensity
NHS Normal Human Serum
PBS Phosphate Buffered Saline
TMA Thrombotic Microangiopathy
TTP Thrombotic Thrombocytopenic Purpura
VWF Von Willebrand Factor
1
CHAPTER 1 AN INTRODUCTION TO THROMBOTIC
MICROANGIOPATHY
2
A hidden connection is stronger than an obvious one.
Heraclitus of Ephesus (535–475 BC)
3
1.1 Introduction
This introductory chapter will discuss the clinic-pathological entity thrombotic
microangiopathy (TMA). TMA is clinically manifest as anemia, thrombocytopenia
and acute kidney injury and pathologically, as swollen, damaged microvascular
endothelial cells coupled with platelet-rich microthrombi. There is an increasing
spectrum of diseases leading to TMA, and to this day, still a lack of clarity in
distinguishing where these diseases overlap and where they differ, in terms of
pathomechanisms. At the core of this thesis is an attempt to better understand a
particular form of TMA, atypical hemolytic uremic syndrome (aHUS), a disease
where a dysregulated alternative pathway (AP) of complement is thought to injure the
microvascular endothelium, thus creating a procoagulant nidus for the aggregation of
platelets. The AP of complement is a constitutively active cascade of intravascular
proteins that can ultimately lead to the formation of a pore-forming protein complex
in the plasma membrane of a cell and lysis of that cell. It can be rapidly amplified if
the cell surface is not recognized as being part of the host, which makes it an efficient
means of killing bacteria, or labeling them for phagocytosis and immune processing.
As such, it can be viewed as a sentinel of immune surveillance. Such a system needs
tight regulation and there are a number of regulators that can arrest the complement
cascade or allow for selective, i.e. site- and time- directed activation. Some of these
are soluble in the circulation, and capable of binding and inactivating complement
proteins that are either in the fluid phase or already attached to the host endothelium,
while others are membrane-anchored. Mutations in both the fluid-phase and
membrane-bound regulators have been linked to the pathogenesis of aHUS in about
two thirds of cases.(Noris and Remuzzi, 2009) Of note, complement regulatory
mutations have in general a low penetrance of about 50%. (Noris and Remuzzi, 2009)
Mutations in one of the endothelial cell-anchored regulators, CD46/Membrane
Cofactor Protein (MCP) have been found in about 10% of aHUS cases but these have
an even lower penetrance than some of the other mutations associated with aHUS,
such as the fluid phase regulator complement Factor H (CFH) for instance. There
remain significant gaps in our knowledge in explaining this variation, and in
understanding the path from a dysregulated complement system to a TMA phenotype.
4
1.2 Thrombotic microangiopathy
1.2.1 Introduction to Thrombotic Microangiopathy
Thrombotic microangiopathy (TMA) is a pathological development arising from the
formation of intravascular thrombi, or clots, in the microvasculature, leading to a
microangiopathic hemolytic anemia, occlusion of the vessels and damage of the
tissues downstream of the occluding thrombi. (Kwaan, 2011) On a pathological level
there is capillary and arteriolar thrombosis accompanied by endothelial cell (EC)
swelling and damage.(George and Nester, 2014) Symmers proposed the term
thrombotic microangiopathy in 1952 to indicate “the location and the most striking
feature of the characteristic histological lesions without mentioning inconsistent and
controversial features of this clinico-pathological entity”. (Symmers, 1952) TMA is a
multisystem disease and can, in principle, affect any organ system including the heart,
lungs, brain, liver, pancreas, skin, bones and kidneys, with effects ranging from
sudden death to progressive organ damage and loss. The latter is a hallmark of kidney
TMA, especially in children, which affects the glomeruli, the blood-filtering vascular
units of the kidney. TMA can be an acute and life-threatening disease with serious
long-term sequelae and in some forms, a risk of recurrence. (Cataland and Wu, 2014;
Clark, 2012; Deford et al., 2013; Radhi and Carpenter, 2012)
1.2.2 The Spectrum of Thrombotic Microangiopathy
TMA is defined by the occurrence of occluding thrombi in the microvasculature here
is an increasing spectrum of diseases associated with a TMA and to date at least nine
separate TMA „syndromes‟ have been defined. (George and Nester, 2014; Riedl et al.,
2014a; Riedl et al., 2014b) Broadly speaking the TMAs can be categorized as being
either hereditary or acquired. (George and Nester, 2014) The principal causes and
associations of TMA are outlined in Table 1-1. Specific causes of TMA include: (i)
thrombotic thrombocytopenic purpura (TTP), (ii) hemolytic uremic syndrome (HUS),
(iii) drugs, such as cyclosporine or quinine, (Kojouri et al., 2001; Miller et al., 1997)
(iv) pregnancy-related TMA, (D'Angelo et al., 2009) (v) the inherited metabolic
disorders of cobalamin such as methyl malonic academia and homocystinuria (Van
Hove et al., 2002) and (vi), transplant associated TMA, particularly in association
with antibody-mediated rejection (Noone et al., 2012; Satoskar et al., 2010) and post
stem cell transplantation.(Jodele et al., 2014; Jodele et al., 2013) Although there is an
5
emerging spectrum of diseases leading to a TMA, TTP and HUS are the two
quintessential diseases associated with TMA. (George and Nester, 2014)
Table 1-1: Classification of Thrombotic Microangiopathies Thrombotic thrombocytopenic purpura (TTP)
TMA associated with ADAMTS13 deficiency
Hereditary
Autoimmune
Infection induced TMA
(Enterohemorrhagic) E.coli- (EHEC)-associated HUS
Shigella dysenteriae HUS
Streptococcus pneumonia HUS
Influenza A/H1N1HUS
HUS due to other pathogens: EBV, CMV, Mycoplasma pneumoniae,
Bordetella pertussis, Parvovirus B19, HIV
Complement-Mediated ‘Atypical’ hemolytic uremic syndrome (aHUS)
Hereditary (mutations in CFH, CFI, MCP, C3 or CFB)
Autoimmune (autoantibodies to CFH)
TMA associated with pregnancy
TMA associated with transplantation
TMA developed de-novo after solid organ transplantation
TMA associated with bone-marrow transplantation or stem-cell
transplantation
TMA associated with metabolic disease
Cobalamin C deficiency
TMA associated with other glomerulopathies/vasculitides
Systemic lupus erythematosus/Antiphospholipid syndrome
C3 glomerulopathy (C3G)
Others: IgA nephropathy, focal segmental glomerulosclerosis (FSGS),
vasculitis
Drug induced TMA
Calcineurin inhibitors
Others: quinine, ticlopidine, chemotherapy
Other forms of TMA
DGKE mutation
Adapted from (George and Nester, 2014; Lemaire et al., 2013; Riedl et al., 2014a; Riedl et al., 2014b))
6
1.2.3 Thrombotic Thrombocytopenic Purpura, VWF and ADAMTS13
1.2.3.1 Introduction to TTP
Eli Moschcowitz first described TTP in a 16-year-old girl who died within two weeks
of presenting with hemolytic anemia, thrombocytopenia, fever and neurological
symptoms. At post-mortem examination hyaline thrombosis of the capillaries was
found. (Moschcowitz, 1924) It was almost 60 years later when unusually large von
Willebrand Factor (VWF) multimers, a hemostatic protein, were identified in TTP
patients‟ plasma. (Moake et al., 1982) Sixteen years later the reason for the presence
of these unusually large VWF multimers in the plasma of TTP patients was
discovered to be a defect in the VWF cleaving protease ADAMTS13 (A Disintegrin
And Metalloproteinase with a ThromboSpondin type 1 motif, member 13). This was
either an acquired phenomenon via an inhibiting antibody or due to a mutation of
ADAMTS13. (Furlan et al., 1998; Tsai and Lian, 1998) TTP occurs either as the
hereditary Upshaw-Schulman syndrome (Rennard and Abe, 1979) where there are
either homozygous or compound heterozygous mutations in ADAMTS13, multimers,
or autoantibodies directed against it (Moschcowitz disease). (Kinoshita et al., 2001;
Levy et al., 2001; Sasahara et al., 2001; Tsai, 2013; Zheng et al., 2001)
Apart from the severe congenital form, Upshaw-Schulman syndrome, TTP is more
common in adults. TTP is manifest clinically with a non-immune microangiopathic
hemolytic anemia, thrombocytopenia, altered neurological status, kidney failure and
fever. This makes up the classically described pentad of TTP. (Knobl, 2014) Activity
levels of ADAMTS13 less than 10% may corroborate the diagnosis. (George and
Nester, 2014)
1.2.3.2 von Willebrand Factor
VWF is a glycoprotein contained as ultra large multimers in Weibel-Palade bodies
(WPB) of the ECs that are released upon cell activation. It is synthesized either in
ECs or megakaryocytes where it is stored in the WPB. VWF is also stored in the
granule of platelets. (Valentijn and Eikenboom, 2013) VWF is initially synthesized as
a monomer that dimerizes via its C-terminus and finally forms multimers through N-
terminal interactions. Multimers may either be released basally into the circulation or
trafficked to the WPB where the multimerization process continues, forming ultra
7
large VWF (ULVWF) greater than 10,000 kDa in size. (De Ceunynck et al., 2013)
ULVWF can be released from a single WPB or from a „secretory pod‟ of coalesced
WPB. (De Ceunynck et al., 2013) Under the influence of blood flow, globular VWF
elongates into strings of between 100 -500 μm in length, (De Ceunynck et al., 2011)
which trap and bind platelets thus initiating clot formation. (De Ceunynck et al., 2013;
Nightingale and Cutler, 2013; Turner et al., 2009; Turner et al., 2012) VWF strings
anchored to ECs are much more efficient at adhering platelets after shear stress
exposes the A1 domain. (De Ceunynck et al., 2013; Schneider et al., 2007) VWF also
serves as a carrier protein for Factor VIII, protecting it from degradation and
clearance, thus further facilitating hemostasis. (Nightingale and Cutler, 2013)
1.2.3.3 ADAMTS13
It was initially recognized that VWF in the circulation was cleaved by a protease,
(Furlan et al., 1996; Tsai, 1996) and that this protease was a zinc protease, termed
ADAMTS13. (Levy et al., 2001; Zheng et al., 2001) ADAMTS13 prevents VWF
multimers from becoming excessively large by cleaving the VWF (Dong, 2005) at a
peptide bond in its A2 domain (Dong et al., 2003) and VWF multimers attached to the
EC undergo structural alterations under shear flow that enable ADAMTS13 to cleave
them. (Lancellotti and De Cristofaro, 2011; Lopez and Dong, 2004) When
ADAMTS13 is congenitally deficient or neutralized by an autoantibody, then
ULVWF and platelet rich thrombi may occlude the microcirculation. Platelets avidly
bind and are firmly fixed to ULVWF multimers that have been unfolded in the
circulation, thus forming a nidus for clot formation. (Levy et al., 2001; Tsai, 2010,
2013; Tsai and Lian, 1998; Tsai et al., 2006)
8
1.2.4 Atypical Hemolytic Uremic Syndrome – a Complement-Mediated
TMA
1.2.4.1 A Brief Introduction of Hemolytic Uremic Syndrome – Typical and Atypical
Hemolytic uremic syndrome (HUS) is a life-threatening disease, typically of
childhood, where the child develops a TMA with anemia, thrombocytopenia, and
subsequent kidney failure after infection by an enterohemorrhagic, Shiga toxin-
producing E coli (STEC) strain. (Gasser et al., 1955; Karmali et al., 1983; Tarr et al.,
2005) About 15% of children infected with STEC will go on to develop HUS 7-10
days after ingestion of the bacteria and this is usually associated with bloody diarrhea.
The triad of a non-immune, microangiopathic hemolytic anemia, thrombocytopenia
and acute kidney injury characterizes HUS. Historically this was classified as
diarrhea-positive HUS and occasionally is associated with epidemics or outbreaks.
(Tarr et al., 2005) Although STEC HUS can present with a severe illness recovery is
usually spontaneous, there are no relapses and overall, the prognosis is good.
(Scheiring et al., 2008)
Following the recognition of HUS as a new disease entity came the realization that
there was a subtype of HUS, that was „atypical‟, in that it was either recurrent (Brain,
1969; Kaplan, 1977) or familial and had a more severe presentation and worse
outcome. (Hagge et al., 1967; Kaplan et al., 1975) These atypical forms were
recognized to occur after a nonspecific prodrome and were diarrhea negative. (Kaplan,
1977) Early in the history of atypical HUS, disturbance and activation of the
complement system was noted. (Barre et al., 1977; Carreras et al., 1981; Drukker et
al., 1975) It had also been noted that plasma therapy could replace some, as yet
unknown and presumed deficient plasma factor. (Remuzzi et al., 1978; Remuzzi et al.,
1979) Furthermore, plasma exchange was effective in cases where infusion seemed
insufficient. (Camba et al., 1985; Remuzzi and Ruggenenti, 1992) Low levels of the
complement regulator Factor H (CFH) had been described in aHUS patients, (Pichette
et al., 1994; Thompson and Winterborn, 1981) but the major breakthrough came when
Warwicker et al mapped the inherited form of HUS to the region on chromosome 1q
that encoded complement Factor H (CFH) and confirmed genetic deficiency in this
complement regulator as being the cause. (Warwicker et al., 1999; Warwicker et al.,
9
1998b) Warwicker and Goodship were the first to propose a “complement-based
theory of microangiopathy”. (Warwicker et al., 1998a)
1.2.4.2 The Complement System of Innate Immunity
Complement proteins form part of the innate immune system where they
„complement‟ and augment antibody-mediated bacterial killing, linking innate and
adaptive immunity, as well as protect the host from various infectious challenges and
aid in the clearance of immune complexes and cellular debris. (Walport, 2001a)
Origins of the complement system can be traced back to 500 million years
ago.(Nonaka, 2014) There are three principal complement activation pathways,
namely the classical (CP), mannose-binding lectin (MBL) and AP.
The classical pathway is activated when C1q, part of the C1 protein complex that
includes the serine proteases C1s and C1r, binds antigen-antibody complexes of either
IgG or IgM, resulting in the cleavage of C4 into C4a and C4b. C4b can then bind C2.
The serine protease C1s cleaves the C4b-bound C2 into C2a and C2b. The remaining
C4bC2a acts as a C3 convertase, splitting it into the anaphylatoxin C3a, and C3b.
C4bC2aC3b acts as a C5 convertase to cleave C5 into the anaphylatoxin C5a, and
C5b, thus initiating the terminal complement cascade (TCC). C5b deposited on cell
surfaces associate with the proteins C6, C7, C8 and C9 to form the heteropolymeric,
membrane attack complex, C5b-9 (MAC). (Degn and Thiel, 2013; Matsushita et al.,
2013; Muller-Eberhard, 1986)
In the mannose-binding lectin pathway, polysaccharides and sugars on the surface of
pathogens are bound by either mannose binding lectin (MBL) or ficolins. The MBL
also have associated serine proteases (MASP 1 and 2) that are phylogenetically
analogous to the C1 complex, and capable of cleaving and activating both C4 and C2.
(Matsushita et al., 2013; Wallis, 2007)
The alternative pathway is constitutively active, amplifiable and targets any
unprotected surface. (Gotze and Muller-Eberhard, 1976) This is in contrast to the CP
and MBL that are activated by binding certain pathogen association molecular
patterns. Spontaneous activation of C3 occurs in the fluid phase via a „tick-over
mechanism‟ related to instability of the thioester bond within the C3 alpha chain. C3
in which the thioester bond has been exposed and hydrolyzed will naturally associate
with H20. Complement Factor B can bind C3(H20) to form C3(H20)B. Factor D
10
subsequently cleaves C3(H20)B, thereby forming the AP C3 convertase C3(H20)Bb
and initiating the AP. (Degn and Thiel, 2013; Ricklin et al., 2010) Properdin stabilizes
the AP convertase thus enhancing amplification of this pathway. (Fearon and Austen,
1975; Muller-Eberhard, 1988; Pillemer et al., 1954) C3b(H20)Bb cleaves more C3
creating an amplification loop and the AP C5 convertase, C3bBbC3b, is formed
allowing progression to the TCC and construction of C5b-9/MAC. (Pangburn et al.,
2008) It has recently been observed that the AP can act as an amplifier of the other
two pathways, increasing the amount of C5 cleaved and C5b-9 formed by up to 80%.
(Harboe and Mollnes, 2008; Harboe et al., 2004)
No matter how the complement cascade is activated, all three pathways converge at
the level of hydrolysis (i.e. activation) of C3, the most abundant complement protein
in plasma, with the formation and deposition of C3b onto any target cell surface. C3
activation can occur both in the fluid phase and on surfaces. The attachment of C3b to
the cell surface is followed by the formation of a C3 convertase (C3bBbP), capable of
cleaving more C3. Within seconds to minutes a cell surface can be coated in C3b.
(Pangburn et al., 2008) When C3b (like C4b) is bound to an antigen or cell it can label
it for processing by antigen-presenting cells or for phagocytosis (opsonization).
Alternatively the terminal cascade of complement can be initiated and propagated by
the C3 and C5 convertases, leading to the formation of the membrane attack complex,
C5b-9. (Pangburn et al., 2008; Walport, 2001a) The deposition of C3b on cells is
indiscriminate, and recognition and protection of the host is achieved principally by a
system of regulators, discussed below. (Pangburn et al., 2008)
11
Figure 1-1. Overview of complement activation
Complement may be activated via the classical, mannose-binding lectin or the APs. All
pathways converge at the level of C3. The classical pathway is activated by the C1 protein
complex of C1q and the serine proteases C1s and C1r. Mannose-associated serine protease
(MASP) 1 & 2 are involved in the initiation of the MBL pathway. The AP is continuously
active and amplifiable. All three converge at the level of C3 and have a common terminal
pathway to formation of the membrane attack complex, C5b-9. (Walport, 2001b)
Recently, a fourth pathway of activation was recognized which, in contrast to the
other pathways, bypasses C3 activation. Thrombin, a serine protease that catalyzes the
conversion of fibrinogen to insoluble fibrin strands to aid coagulation, has been
recognized as this fourth, C3 independent, pathway of complement activation.
(Huber-Lang et al., 2006) Furthermore, thrombin can generate a unique C5 split
(C5bT) product that when acted on by the C5 convertase results in a MAC complex
with increased lytic activity. (Krisinger et al., 2012) Thrombin can also cleave C3 to
form the anaphylatoxin C3a. (Amara et al., 2008)
12
1.2.4.3 Endothelial Cell Protection against Complement-Mediated Injury
Over 60 trillion (1012
) endothelial cells make up the largest interconnected organ in
the human body, the lining of the vasculature, covering as much as 4,000 square
meters. (Aird, 2007a) Host protection of the vascular endothelium is extremely
important, especially against the AP of complement, an innate system of immune
surveillance that is constitutively active, amplifiable and ends with a membrane lysing
protein complex. In fact, all human cells have complement regulators on their surface.
(Zipfel and Skerka, 2009) Regulation of complement occurs with a combination of
fluid phase regulators, including complement Factor H (CFH) and membrane-
anchored regulators including CD35/CR1, CD46/Membrane Cofactor Protein (MCP),
CD55/Decay Accelerating Factor (DAF) and CD59. These are outlined in table 1-2.
(Campbell et al., 2002; Skidgel and Erdos, 2007; Thurman and Renner, 2011;
Tschopp and French, 1994) The overarching functional principle of these regulators is
the interference with the progression of the complement cascade at the level of C3b or
the C3 convertase. CFH is a cofactor for the serine protease complement Factor I that
cleaves and inactivates C3b. It also serves to accelerate the decay of the C3
convertase and as a host recognition molecule. (Ferreira et al., 2010) On the EC
surface CD55/DAF also accelerates the decay of the C3 convertase, while CD59
prevents formation of C5b-9 by binding C8 and C9. (Thurman and Renner, 2011)
Within the kidney, glomerular endothelial cells (GEC) express CD46/MCP,
CD55/DAF and CD59, (Endoh et al., 1993; Ichida et al., 1994; Nakanishi et al., 1994)
but do not express the EC surface regulator CR1. (Thurman and Renner, 2011).
CD46/MCP is a transmembrane protein identified first in 1985 as a C3b binding
protein expressed on human mononuclear cells. (Cole et al., 1985) As stated above,
CD46/MCP binds C3b (and C4b) and has cofactor activity for CFI, allowing for
cleavage and inactivation of the complement activation product C3b deposited on the
host cell surface. (Seya and Atkinson, 1989; Seya et al., 1986) Originally described
and characterized on blood cells, (Seya et al., 1988) and later on EC, (McNearney et
al., 1989) CD46/MCP is coded for by a gene lying within the region of regulators of
complement activation (RCA) gene cluster on chromosome 1q32. (Liszewski et al.,
1991) Within the kidney, CD46/MCP expression is ubiquitous, on glomerular
structural cells, podocytes and tubular epithelial cells. (Endoh et al., 1993; Ichida et
al., 1994; Nakanishi et al., 1994) Its role as a complement regulator was extensively
13
studied (Barilla-LaBarca et al., 2002; Brodbeck et al., 2000; Liszewski and Atkinson,
1996) prior to its discovery to be related to aHUS some 15 years later.(Richards et al.,
2003) Structurally, CD46/MCP consists of an extracellular domain with four
complement control protein modules, and an O-glycosylated region rich in serines,
threonines and prolines termed the “STP” region, a cytoplasmic anchor and tail.
Alternative splicing of the CD46 gene affects both the “STP” region and cytoplasmic
tail resulting in principally four isoforms of CD46/MCP. (Post et al., 1991) The
cytoplasmic tail may be responsible for directing the intracellular protein to the cell
surface. (Maisner et al., 1996)
1.2.4.5 Complement-mediated ‘atypical’ HUS
Atypical HUS (aHUS) can be either sporadic or familial, and has been linked to
defective regulation of the innate immune and coagulation systems. (Delvaeye et al.,
2009; Kavanagh et al., 2006; Warwicker et al., 1998b) Atypical HUS (Noris et al.,
2010) is an early-onset disease where many patients have been identified as having
hereditary (e.g. mutations) and/or acquired (e.g. autoantibodies) predispositions
related to the regulation of the complement AP (AP), which involves both plasma-
borne factors (e.g. factor H; CFH) and proteins on the surface of vascular ECs (e.g.
Membrane Cofactor Protein CD46/MCP). (Malina et al., 2012) Different from STEC
HUS, atypical HUS has a high risk of progression to end stage kidney disease (>50%)
and death (up to 25%), or the need for lifelong dialysis or extremely expensive
treatments because these patients are poor candidates for transplant due to disease
recurrence in the graft. (Noris and Remuzzi, 2009) Children with aHUS have a five
times greater mortality as compared to adults.(Fremeaux-Bacchi et al., 2013) Atypical
HUS has been linked to mutations in several genes, detectable in about 60% of cases.
(Caprioli et al., 2006; Noris et al., 2010) Most of these genes are in the regulator of
complement activation (RCA) region of Chr 1 (1q32). (Rodriguez de Cordoba et al.,
1999) Loss of AP regulation results in the generation of excessive amounts of
anaphylatoxins (C3a, C5a), opsonins (C3b) and membrane attack complexes
(MAC/C5b-9) both in fluid phase and on vascular ECs in glomeruli, (Waters and
Licht, 2011) where damage releases signals promoting inflammation and platelet
14
activation leading to thrombus/clot formation. The resultant TMA in the kidney
microvasculature causes to in acute kidney injury (aHUS). (Zipfel et al., 2006)
Table 1-2: Relevant Complement Regulators
Regulator Function
Fluid Phase Regulators
CFH Accelerates decay of C3 convertase
Cofactor for the serine protease CFI that
cleaves C3b into inactive form iC3b
C4b binding protein Functions as CFH but for the CP
Vitronectin/ S protein Binds to C5b-7 to prevent C5b-9
formation
Clusterin Binds to C5b-7 to prevent C5b-9
formation
Carboxypeptidase N Inactivates the anaphylatoxins C3a &
C5a
Pro-carboxypeptidase R
(proCPR)/thrombin-activatable
fibrinolysis inhibitor (TAFI)
Inactivates the anaphylatoxins C3a &
C5a
Membrane-Anchored Regulators
CD46/Membrane Cofactor Protein Cofactor for the serine protease CFI that
cleaves C3b into inactive form iC3b
CD55/Decay Accelerating Factor Accelerates decay of C3 convertase
CD59 Binds C5b-8 on cell membrane
preventing C5b-9 formation
CD35/CR1 Cofactor for the serine protease CFI that
cleaves C3b into inactive form iC3b &
Accelerates decay of C3 convertase
1.2.4.6 CD46/MCP-associated atypical hemolytic uremic syndrome
The principal EC bound complement regulator first recognized as linked to aHUS is
membrane cofactor protein (MCP/CD46). (Pirson et al., 1987; Richards et al., 2003)
CD46 gene mutations have been found in up to 10% of aHUS patients.(Fremeaux-
Bacchi et al., 2013; Fremeaux-Bacchi et al., 2006) The mutations in MCP related
aHUS are mainly heterozygous, compound heterozygous or homozygous and
translate into a reduced cell surface expression of the CD46/MCP protein in more
15
than 75% of cases. (Fremeaux-Bacchi et al., 2013; Goodship, 2006) This reduced cell
surface expression is due to retention of the immature precursor protein within the cell.
(Goodship, 2006) The remainder of MCP mutations result in a protein expressed at
normal levels but lacking either cofactor activity or C3b binding capacity. (Esparza-
Gordillo et al., 2005; Noris et al., 2003; Richards et al., 2003) Studies in transfected
Chinese hamster ovary cells have found that this reduced expression translates into
reduced regulation of the AP, (Liszewski et al., 2007) ultimately then thought to lead
to enhanced C5b-9/MAC generation.
Although those with CD46/MCP related aHUS occur at a younger age and have a
higher rate of recurrence than those with other forms of aHUS including CFH related
aHUS, they are three to four times more likely to retain normal renal function and
have 75% long-term renal survival. (Fremeaux-Bacchi et al., 2013; Noris et al., 2003;
Noris et al., 2010; Sellier-Leclerc et al., 2007) CD46/MCP related aHUS is somewhat
unique, in that its rate of recurrence post-transplant is minimal, likely owing to the
fact that the kidney allograft will be expressing non-mutant CD46/MCP. (Caprioli et
al., 2006; Fremeaux-Bacchi et al., 2007; Richards et al., 2007)
1.2.5 From Mutation in CD46 to TMA – An Incomplete Story
1.2.5.1 Functional Studies of CD46/MCP Mutations
Functional studies of CD46/MCP mutations associated with aHUS have been
conducted using Chinese Hamster Ovary (CHO) cell transfection, and CHO cells
transfected with MCP mutations were demonstrated to having an increased cell
surface deposition of C3b. However, CHO cells express no other complement
regulators and, therefore, studies performed in CHO cells do not take into account the
contribution of any of the other complement regulators that would be present, and
presumably functional, in vivo, on human EC. Thus, although an excellent way for
assessing the functional impact of individual MCP mutations (Brodbeck et al., 2000;
Liszewski and Atkinson, 1996) (Liszewski et al., 2008; Liszewski et al., 2007) this
system does not allow for interpreting the role of MCP mutations in context of the
physiological setting of multilayered complement regulators e.g. on EC with respect
16
to the manifestation of a TMA phenotype. For the most part it is assumed that this
increase in C3 deposition implies progression to the TCC with more C5b-9
production, EC activation, injury or death and the emergence of a prothrombotic EC
surface upon which platelets aggregate. (Noris and Remuzzi, 2009). However, this
pathogenetic sequence of TMA has never been fully proven and hinges significantly
on assumptions.
1.2.5.2 Incomplete Penetrance Suggests a Multiple ‘Hit’ Concept
Mutations in CD46/MCP associated with aHUS – like other aHUS causing mutations
– are not automatically disease causing but are associated with variable penetrance.
(Dragon-Durey and Fremeaux-Bacchi, 2005) In fact a sibling-pair both harboring a
homozygous deletion in CD46/MCP and lacking CD46/MCP has been reported where
one has recurrent aHUS and the other absolutely no disease.(Couzi et al., 2008) This
incomplete penetrance is not unique to CD46/MCP, and in other forms of
complement-mediated aHUS, a patients‟ family members may also carry the same
mutation and remain unaffected.(Kavanagh et al., 2005; Martinez-Barricarte et al.,
2008; Noris et al., 2010) This variable, or incomplete penetrance, suggests a threshold
for the development of aHUS. It has been suggested that mutations in the complement
system may serve as predisposing factors for the development of aHUS, where some
mutations may be associated with a greater risk. Then, with a certain „trigger‟ event –
either environmental or provided by another genetic defect – the patient may manifest
the disease. Infection seems to be a common trigger event, where children often
present after a nonspecific prodrome suggestive of a viral illness, or have a disease
recurrence seemingly precipitated by an intercurrent illness. (Johnson and Waters,
2012) Other triggers preceding aHUS manifestation include immunization, pregnancy,
surgical operations etc. (ref – if at hand)
1.2.5.3 An Additional Complement Gene Aberrancy Increases Disease Penetrance
Having a combination of mutations in complement genes has been described in aHUS.
(Bienaime et al., 2010; Geerdink et al., 2012; Maga et al., 2010; Noris et al., 2010;
Sellier-Leclerc et al., 2007) In an American cohort, 12% of the patients had mutations
in more than one gene, (Maga et al., 2010) and a relatively small pediatric cohort of
45 children with aHUS found 4 with combined mutations. (Geerdink et al., 2012)
17
Furthermore, certain haplotypes and single nucleotide polymorphisms in complement
genes, genetic variations in DNA or single nucleotides that are very close and
therefore inherited alongside each other, have been identified in aHUS patients and
may act as disease susceptibility factors. (Fremeaux-Bacchi et al., 2005) These
variations can be additive, for instance in one pedigree only individuals possessing
three - a mutation and SNP in CD46/MCP as well as an insertion in the gene for CFI
– developed aHUS. (Esparza-Gordillo et al., 2006) SNPs alone in CD46 are likely
insufficient to cause the disease and a mutation is also required.(Ermini et al., 2012)
Recent evidence has shown that the combination of complement gene „risk haplotypes‟
can increase the disease penetrance. (Bresin et al., 2013) The presence of both a CFH
and CD46 risk haplotype increases disease penetrance to 73% versus 36% among
carriers with zero or one risk haplotype. It also seems that as well as increasing
disease penetrance, the natural course of the disease may also be altered by having a
combination of haplotypes as 50% of patients with combined CD46 mutation
developed ESRD within 3 years from onset as compared to just 19% of patients with
an isolated CD46 mutation in this study.(Bresin et al., 2013)
1.2.5.4 A Link Between Complement and Coagulation
The coagulation and complement systems have been partners in protecting their hosts
for millions of years, although a greater understanding of the extent of the interaction
between the two systems continues to evolve. (Delvaeye and Conway, 2009;
Markiewski et al., 2007; Oikonomopoulou et al., 2012) The convergence of both
systems has recently come to the foreground, especially with the finding of an
additional C3-independent pathway of complement activation by thrombin, a
proteolytic serine protease integral to coagulation. (Huber-Lang et al., 2006) This was
followed by the finding that thrombomodulin (THBD; CD141), a cofactor for
thrombin, also functioned as a cofactor for CFI-mediated C3 cleavage, therefore
acting as a complement regulator and that certain heterozygous missense mutations
could be linked to patients with aHUS. (Delvaeye et al., 2009) Furthermore,
pathogenic variants in coagulation genes, notably PLG, that encodes plasminogen,
were recently found in an aHUS cohort. (Bu et al., 2014)
18
Thrombotic microangiopathy (TMA) is a pathological process where this complex
interplay between the complement system and coagulation has always been prominent
and somewhat controversial, especially in terms of distinguishing TTP and aHUS.
Both clinically and pathologically it can be difficult to distinguish the two. (Fujimura,
2003; Remuzzi, 1987, 2003; Tsai, 2003) As it became evident that the underlying
cause was clearly different, related to defective complement regulation in aHUS, and
failure of ADAMTS13 to cleave VWF multimers in TTP, the two seemed clearly
distinct.(Tsai, 2003) However more recently it has emerged that complement may
play a role in TTP, blurring the lines of separation again. (Noris et al., 2012; Reti et
al., 2012; Ruiz-Torres et al., 2005; Wu et al., 2013; Zipfel et al., 2011) Sera from TTP
patients can activate the AP leading to EC cytotoxicity in vitro. (Ruiz-Torres et al.,
2005) Reti et al demonstrated that the anaphylatoxin C3a and soluble C5b-9 levels
were higher in TTP patients during acute disease. (Reti et al., 2012) Following on
from that, Wu et al analyzed stored blood samples from a cohort of 38 TTP patients,
10 of whom died, for complement activation. Thirty three of 38 (87%) had evidence
of complement activation at presentation and in those that died, there was evidence of
both AP and terminal complement cascade activation with levels of Bb, C3a, C5a and
C5b-9 all significantly higher in this group. (Wu et al., 2013)
Recently, the link between complement and coagulation in aHUS has been expanded
to involve VWF and its cleaving protease, ADAMTS13. In a recent study involving
29 aHUS patients, Feng et al found additional ADAMTS13 polymorphisms
associated with decreased ADAMTS13 activity. (Feng et al., 2013a) 80% of the
cohort also carried at least 1 non-synonymous change in ADAMTS13, and in 38% of
patients, multiple ADAMTS13 variations were found. These ADAMTS13 variants
were likely significant as measured ADAMTS13 activity was less than 60% in half of
the patients studied.(Feng et al., 2013a) Alongside this clinical observation, there have
been a number of experimental studies linking CFH, ADAMTS13 and VWF. Turner
and Moake demonstrated the assembly and activation of AP components, especially
C3 and C5, on ultra large VWF strings secreted from and anchored to ECs.(Turner
and Moake, 2013) CFH has been shown to be a reductase for large soluble VWF
(LsVWF) multimers that are secreted from ECs. (Nolasco et al., 2013) Large soluble
VWF multimers released from complement activated ECs assume a conformation that
makes them inaccessible to ADAMTS13 cleavage, but they can still bind platelets and
19
induce platelet aggregation under conditions of high-fluid shear stress. (Nolasco et al.,
2013) The C-terminus of CFH has been shown to enhance ADAMTS13 VWF
cleavage in vitro. (Feng et al., 2013c) Finally, CFH cofactor activity may be enhanced
by an interaction between VWF and CFH. (Rayes et al., 2014) Putting this all together,
it has recently been postulated that the association between VWF and the AP of
complement may be complement amplifying, which would be very pertinent to TMA
and aHUS. (Turner et al., 2014)
20
1.3 Blood Outgrowth Endothelial Cells (BOEC)
1.3.1 Introduction to BOEC
It has recently become feasible to isolate endothelial progenitor cells (EPCs) from
circulating peripheral blood mononuclear cells (PBMC), and culture them to obtain
“outgrowth” populations known as BOEC (Lin et al., 2000) or endothelial colony-
forming cells (ECFCs). (Reinisch et al., 2009) BOEC or endothelial colony-forming
cells (ECFCs), are considered by some authors to be true EC precursors, having a
distinct transcript signature compared to early endothelial progenitor cells (EPCs) and
a high proliferative potential. Populations of these cells can be expanded in multiple
passages for 3-4 months without them undergoing any transformation and with
retention of their EC traits in terms of genotype and phenotype.(Fuchs et al., 2010;
Fuchs et al., 2006; Timmermans et al., 2009) Their precursor has not yet been
identified and it is possible that they originate from blood vessels in vivo, as they are
phenotypically similar to mature ECs, yet have a much greater proliferative
capacity.(Tura et al., 2013)
1.3.2 BOEC in the Study of Disease
BOEC have proven useful in studies of vascular disease; e.g. transcriptome analysis
detected upregulation of disease-relevant genes in BOEC from patients with
proliferative diabetic retinopathy, whose cells also showed diminished migratory
capacity and integration into retinal endothelium. (Tan et al., 2010) BOEC expression
array analysis detected several genes with roles in endothelial biology that may
influence susceptibility to shear stress and cardiovascular disease. (Ahmann et al.,
2011; Ensley et al., 2012; Mazzolai et al., 2011) BOEC are also well suited to in vitro
modeling of cell interactions and pathological conditions such as high shear stress and
ischemia-reperfusion. These cells can be obtained with high efficiency from small
volumes of peripheral blood using cell separation methods and direct culture, and they
have close affinities to their donor‟s vascular ECs. (Martin-Ramirez et al., 2012;
21
Reinisch et al., 2009) For example BOEC have been used to elucidate the functional
symbiosis between glomerular endothelium and epithelium mediated by podocyte-
secreted VEGF. (Hirschberg et al., 2008) They have also proven useful in studies
relating to thrombotic disease. In a shear stress model BOEC were observed to alter
their cytoskeletal structure and antithrombogenic potential, (Ensley et al., 2012) and
showed changes in expression of receptors for tissue plasminogen activator, urokinase
and adhesion molecules. (Ahmann et al., 2011; Mazzolai et al., 2011) BOEC have
also been used to study the effects mutations in EC-expressed genes (e.g. von
Willebrand Factor [VWF] deficiency(Othman et al., 2010)), and there is evidence that
BOEC and other EPCs may represent populations that play important roles in
recovery from thrombotic disease (e.g. stroke(Yang et al., 2012)). Several groups
have begun to explore the therapeutic potential of EPCs for vascular cell
transplantation(Kaneko et al., 2012) (with or without genetic modification) and
antithrombotic endothelialization. (Kaneko et al., 2012; Yoon et al., 2005) BOEC can
also be used to study the functional effects of a known EC mutation as has been done
by Othman et al, where they characterized the functional significance of a specific
von Willebrand Factor (VWF) mutation. (Othman et al., 2010) BOEC have been used
to study the effects of mutations in EC-expressed genes (e.g. von Willebrand Factor
[VWF] deficiency. (Othman et al., 2010; Wang et al., 2013b)
BOEC represent a potentially valuable source of material for gene expression analysis
and the search for disease-associated gene variants using a transcriptomic approach.
(Fernandez et al., 2005; Medina et al., 2010b; Tan et al., 2010) The application of
global microarray profiling of BOEC from patients with a defined genetic phenotype
can provide insight into the mechanisms underlying that disease at the level of the EC.
(Medina et al., 2010a) Different subjects might exhibit different combinations of
polymorphisms that affect EC gene expression and biologic systems. (Enenstein et al.,
2010) Chang Milbauer et al studied BOEC lysates from sickle cell disease patients at
risk for stroke. (Chang Milbauer et al., 2008) By defining a priori, defined biologic
systems that could be implicated in this risk at the endothelial level (e.g. angiogenesis,
hypoxia response, coagulation, shear stress response and inflammation) (Chang
Milbauer et al., 2008) and then assembling specific gene sets to survey that system,
went on to identify enhanced activity of each biologic system. Of note early
expansion stages of BOEC may reveal a transiently acquired phenotype, especially if
22
taken from patients with active inflammation or disease;(Kahlenberg et al., 2011) for
e.g. in burn patients more BOEC colonies were isolated as compared to normal
controls and these BOEC secreted more VEGF than normal counterparts. (Rignault-
Clerc et al., 2012) However, Chang Milbauer et al have shown that after a 10-fold
expansion in the number of cells (at least one passage), some upregulated genes return
back to normal expression levels. (Chang Milbauer et al., 2008) Thus the „acquired
phenotype‟ may be washed out and expanded BOEC reflect culture conditions and the
inherent genetics of the subject‟s ECs. These cells can also be cryopreserved. A study
has shown that after two years cryopreservation at -80 degrees Celsius, almost 80% of
the cells remained viable.(Wagner and Myrup, 2005)
1.3.1 BOEC as a Model to Study TMA
To date most experimental studies on the pathogenesis of TMA and aHUS have used
EC lines such as human umbilical vein ECs, glomerular EC lines (Frimat et al., 2013;
Louise and Obrig, 1994; Ray et al., 2006) or CHO cells. (Liszewski et al., 2007) The
use of non-endothelial cell-based systems however has limitations; (i) it cannot
entirely recapitulate the physiological endothelial milieu and (ii) there are problems
with overexpression/underexpression of the relevant gene being studied, and (iii) the
effect on the other regulators cannot be easily studied. BOEC are an endothelial
progenitor cell that offer a number of advantages over cell lines such as HUVECs and
offer the unique opportunity to directly study patient specific ECs where they will
reflect the particular genotype of that particular individual.
23
1.4 Knowledge Gap, Hypothesis and Thesis Aims
The principal gap in knowledge revolves around explaining the steps from loss of
complement regulation on the endothelium in association with a mutation in
CD46/MCP, to the manifestation of a TMA phenotype of EC death and platelet
microthrombi. What is known comes from studies in CD46/MCP-transfected CHO
cells, a non-human cell, unrelated to the endothelium, and devoid of other
complement regulators. Published, disease-causing mutations in CD46/MCP are less
efficient at inactivating C3b deposited on complement-challenged CHO cells.
Thereafter, it is assumed that the terminal complement cascade gets activated with
more C5b-9 formation, EC injury leading to a procoagulant surface with platelet
aggregation. The fact that the disease is associated with variable penetrance suggests
additional „hits‟ may be required.
Hence, the specific hypotheses tested are the following:
1. In any individual carrying a functionally relevant CD46/MCP mutation, an
additional complement regulatory defect acts as a second „hit‟ and increases
the likelihood of TMA manifestation.
2. Complement-induced EC activation and VWF release acts as an additional „hit‟
via amplification of the complement cascade.
The primary aim of this thesis is to examine how loss of complement regulation on
the EC surface leads to a thrombotic microangiopathic phenotype in an ex vivo model
of disease that incorporates human blood outgrowth endothelial cells (BOEC), an
endothelial progenitor cell that can ultimately be grown from patients, and an
endothelialized microfluidic system.
The secondary aim is to evaluate the contribution of von Willebrand Factor (VWF) to
the pathogenesis of atypical hemolytic uremic syndrome (aHUS).
24
CHAPTER 2 MODELING CD46/MCP-ASSOCIATED
ATYPICAL HEMOLYTIC UREMIC SYNDROME USING
BLOOD OUTGROWTH ENDOTHELIAL CELLS
25
2.1 Abstract
Atypical hemolytic uremic syndrome (aHUS) is due to defective regulation of the
alternative complement pathway (AP), which is constitutively active and requires a
multi-layered defense system. It remains unclear how this defense system functions as
a whole to protect injured endothelial cells (EC) or why loss/diminished function of
one of these regulators results in disease. Mutations in CD46/membrane cofactor
protein (MCP), a membrane-anchored CAP regulator leads to aHUS but with variable
penetrance. We aimed to use blood outgrowth endothelial cells (BOEC), endothelial
cell precursors that can be isolated from the peripheral blood, to study aHUS
pathogenesis ex vivo. BOECs were cultured from subjects by a standard protocol. EC
phenotype and presence of complement regulators was confirmed by
immunofluorescence (IF), western blot (WB), qPCR, and flow cytometry (FACS).
Cells were challenged with complement, with or without functional blockade of
complement regulators. Cell death and apoptosis were quantified. Calcein-labelled
platelets were perfused across confluent BOEC in a microfluidic system and platelet
adhesion was quantified. BOEC were characterized by FACS as being positive for the
endothelial cell markers CD31/PECAM-1 and CD144/VE-cadherin, while being
negative for the hemangioblast marker CD45 and immature endothelial progenitor
cell marker, CD14. Surface expression of complement regulators was confirmed. A
surrogate phenotype of aHUS modelled ex vivo (endothelial cell death and platelet
adhesion) is not achieved with CD46/MCP blockade alone. Complement deposition
(C3) is increased by blocking complement regulators simultaneously (four-fold
increase when CD46/MCP, CD55/DAF and CD59 are blocked; p <0.001) and results
in cell death, apoptosis and an EC prothrombotic phenotype with platelet adhesion.
We have established an ex vivo method to model aHUS pathogenesis that can now be
expanded to use patient-derived BOEC. Our data supports the hypothesis that an
additional factor is required in order for a patient with a CD46/MCP mutation to
manifest disease.
26
2.2 Introduction
Thrombotic microangiopathy (TMA) is a disease affecting the microvasculature,
especially within the kidney, characterized by endothelial cell injury and occlusion of
capillaries and arterioles with platelet microthrombi. Atypical hemolytic uremic
syndrome (aHUS) is one form of TMA typically with a young age of onset that can be
caused by either inherited, or an acquired, dysregulation of the constitutively active
complement alternative pathway (AP). Genetic causes / mutations giving rise to
aHUS either result in loss of appropriate AP regulation by fluid phase (complement
Factors H and I, CFH and CFI respectively), or membrane-anchored regulators
CD46/membrane cofactor protein (MCP), or amplification of the AP by mutant C3 or
complement Factor B (CFB).(Noris and Remuzzi, 2009)
The principal endothelial cell -anchored complement regulator first recognized as
linked to aHUS is CD46/MCP.(Pirson et al., 1987; Richards et al., 2003) CD46/MCP
binds C3b and has cofactor activity for the serine protease complement Factor I (CFI),
allowing for cleavage and inactivation of the complement activation product C3b
deposited on the host cell surface.(Seya and Atkinson, 1989; Seya et al., 1986)
Mutations in CD46/MCP are found in up to 10% of aHUS patients.(Fremeaux-Bacchi
et al., 2006) Mutations are mainly heterozygous, but homozygous, and null mutations
have also been described and involve reduced expression, impaired C3b binding or
retention of a CD46/MCP precursor within the cell. CD46/MCP related aHUS is
somewhat unique, in that unlike the other forms of aHUS, its rate of recurrence post-
transplant is minimal owing to the fact that the kidney allograft will be expressing
non-mutant CD46/MCP.(Caprioli et al., 2006; Fremeaux-Bacchi et al., 2007;
Richards et al., 2007)
How an individual mutation leads to a TMA phenotype and aHUS is not always clear.
Studies of transfected Chinese hamster ovary (CHO) cells have found that reduced
expression or loss of CD46/MCP translates into reduced regulation of the AP and
more C3b deposition on the cell surface.(Liszewski et al., 2007) This might be
expected to translate in vivo, on the human microvascular endothelium, into more
membrane attack complex/C5b-9 (MAC/C5b-9) formation, endothelial cell activation,
27
injury or death and a prothrombotic endothelial cell phenotype with platelet adhesion
and a TMA. However, as clinically affected probands often have unaffected siblings
or relatives carrying the same mutation, current opinion proposes a
„multiple/cumulative-hit‟ sequence of events leading up to disease precipitation.
Disease may become manifest only with the presence of an additional mutation or risk
haplotype, for instance as combination of CFH and CD46/MCP mutations, which has
been shown to (almost) double disease penetrance over carrying individual mutations
and to be associated with a more severe disease and worse outcome.(Bresin et al.,
2013)
Blood outgrowth endothelial cells (BOEC), are an endothelial progenitor that can be
isolated with high efficiency from small volumes of peripheral blood using cell
separation methods and direct culture, and they have close affinities to their donor‟s
vascular endothelial cells.(Martin-Ramirez et al., 2012; Schiff et al., 2004) Although
phenotypically similar to mature endothelial cells, they have a much greater
proliferative capacity and retain their endothelial cell traits in terms of genotype and
phenotype across many passages.(Fuchs et al., 2006; Timmermans et al., 2009; Tura
et al., 2013) BOEC can be used to study the functional effects of known endothelial
cell-expressed genes, with studies in von Willebrand Factor (VWF) deficient patients
and von Willebrand Disease being most successful to date.(Othman et al., 2010;
Wang et al., 2013a)
In this study, using BOEC, we model the impact of a functional defect in CD46/MCP
and show that loss of function of CD46/MCP alone may be insufficient to result in a
TMA phenotype of increased complement deposition, endothelial cell death and
platelet adhesion.
28
2.3 METHODS
2.3.1 BOEC Isolation
48 ml of whole blood was obtained from healthy adult donors in six cell preparation
tube (CPT) vacutainers (Becton Dickinson, Franklin Lakes, USA). Tubes were spun
at 1600 x g for 30 minutes to obtain the serum and mononuclear cell layer (MCL).
The serum and MCL were added to 8 ml of 10% fetal bovine serum (FBS: Sigma-
Aldrich, St. Louis, USA) in phosphate buffered saline (PBS: Wisent). Cells were
pelleted at 520 x g and resuspended in 10% FBS in PBS twice. Using Trypan Blue
(Sigma-Aldrich, St. Louis, USA) and an hematocytometer (Hausser Scientific,
Horsham, USA) the number of cells were quantified. Cells were pelleted at 520 x g
and resuspended in cEGM-2 media with supplements (Lonza, Walkersville, USA: Cat
no. 362753), 10% FBS and 1% Antibiotic-Antimycotic (Gibco, Invitrogen, Life
Technologies, Carlsbad, USA; containing 10,000 units/mL of penicillin, 10,000
µg/mL of streptomycin, and 25 µg/mL of Fungizone® Antimycotic). 3-5 x 107 cells
were aliquoted to each well in a six-well tissue culture plate (Becton Dickinson,
Franklin Lakes, USA) coated with 0.05 mg/ml of rat tail collagen type I in 0.02 M
acetic acid (Becton Dickinson, Franklin Lakes, USA). Cells were kept at 37°C and in
an environment with 5% CO2. For the first 7 days after seeding the cells, 3.5 mL of
media is aspirated carefully, 2 ml of pre-warmed media is gently added, and then
carefully aspirated before replacing with 4 mL of media. Thereafter media is replaced
once every two days for approximately two weeks to a month until other blood cells
die off and a monolayer of BOEC are identifiable by their endothelial cobblestone
morphology and ready for passaging (Figure 2-1A).
2.3.2 BOEC Characterization By Flow Cytometry
Alexa Fluor 488 conjugated mouse anti-human CD31 (PECAM-1) [5μL/test], PE
conjugated mouse anti-human CD144 (VE-Cadherin) [12μL/test], Alexa Fluor 647
conjugated mouse anti-human CD14 [5μL/test] and APC-H7 conjugated mouse anti-
human CD45 [3μL/test] were used for surface staining of the BOEC, with Alexa
Fluor 488 mouse IgG2 κ, PE mouse IgG1 κ, Alexa Fluor 647 mouse IgG2b κ and
APC H7 mouse IgG1 κ, as isotype controls. Cells were stained with Fixable Viability
Dye eFluor 450 (eBioscience, San Diego, USA), fixed with 2% PFA/PBS and
29
blocked with 200 μg/mL final volume of purified mouse serum IgG (Sigma-Aldrich,
St. Louis, USA) prior to specific or isotype antibody incubations. A minimum of
60000 live cells were collected using a Beckman Coulter Gallios flow cytometer
(Beckman Coulter, Brea, USA), equipped with 4 excitation lasers (405nm, 488nm,
561nm, 633nm). Data was analyzed with Kaluza software (Beckman Coulter).
HUVEC passage 6 were used as a positive control for the endothelial cell markers
CD31 (PECAM-1) and CD144 (VE-Cadherin) and blood as a positive control for
CD45 and CD14. Fluorescence minus one (FMO) controls using BOEC were stained
for each of the four specific antibodies and were used as the gating controls during
data analysis. Antibodies were purchased from BD Pharmingen (San Diego, USA).
2.3.3 BOEC Characterization By Immunofluorescence
BOEC at about 80-90% confluence were washed once with HBSS (Gibco, Life
Technologies, Carlsbad, USA), trypsinized (with 0.05% trypsin/0.53mM EDTA,
WISENT, St Bruno, Canada) then resuspended in cEGM-2 (BOEC) media to a
concentration of 0.5 × 106 cells/ml. 500 μL of the BOEC suspension is added to 2.5
mL of cEGM-2 (BOEC) media already pre-warmed in each well of a 6-well tissue
culture dish containing previously collagen-coated 22x22-mm cover slips (VWR
International, Radnor, USA). BOEC adhere to the cover slips after overnight
incubation (37°C, 5% CO2). Samples were washed with ice-cold PBS then fixed with
4% (w/v) paraformaldehyde (Electron Microscopy Sciences, Fort Washington, USA)
in PBS. Samples were blocked with 2% (w/v) donkey serum (Jackson
ImmunoResearch, West Grove, USA) either alone or with 0.2% (v/v) Triton X-100
(for permeabilization) for 60 minutes. Samples were stained overnight with the
relevant primary antibodies made up in the relevant blocking solution. Samples were
washed in PBS and incubated with respective donkey-anti secondary antibodies with
Alexa Fluor® 488 conjugate (Invitrogen, Life Technologies, Carlsbad, USA). Cell
nuclei were stained by either DAPI (1 μg/ml) or 0.12 μg/ml Hoechst stain (Thermo
Fisher Scientific, Waltham, USA) for ten minutes. Samples were washed in PBS and
cover slips were mounted with Dako Fluorescence Mounting Media (Dako, Glostrup,
Denmark) for analysis with either a Nikon Eclipse Ti microscope (Leika
Microsystems, Wetzlar, Germany) or spinning disk confocal microscopy. The latter is
equipped with an Olympus IX81 inverted fluorescence microscope using a 60./1.35
30
oil immersion objective equipped with a Hamamatsu C9100-13 back-thinned EM-
CCD camera and Yokogawa CSU X1 spinning disk confocal scan head (with upgrade
from Spectral Aurora Borealis, Richmond Hill, Canada). The unit is equipped with 4
separate diode-pumped solid state laser lines (Spectral Applied Research, Richmond
Hill, Canada, 405 nm, 491 nm, 561 nm, and 642 nm) with emission filters: 447 nm ±
60, 525 nm ± 50, 593 nm ± 40, 620 nm ± 60, 676 ± 29 and 700 nm ± 75, and 1.5X
magnification lens (Spectral Applied Research). Confocal images were taken with an
Improvision Piezo Focus Drive. Z-stacks were taken at 0.25 μm. Images taken using
the spinning disk confocal microscope were deconvolved by iterative restoration
using Volocity Software (PerkinElmer, Waltham, USA) with confidence limit set to
95% and iteration limit set to 20.
The following antibodies were used for IF: sheep anti-human VWF (1:1000 dilution,
Abd Serotec, Oxford, UK, AHP062), goat anti-human P-selectin (CD62P, 1:100
dilution; Santa Cruz Biotechnology, Dallas, USA; sc-6943), rabbit polyclonal
antibody to MCP (1:50 dilution; Santa Cruz Biotechnology, Dallas, USA; sc-9098,),
rat polyclonal antibody to CD59 (1:1000 dilution; Abd Serotec, Oxford, UK;
MCA715G), and goat polyclonal antibody to CD55 (1:50 dilution; R&D Systems,
Minneapolis, USA; AF2009). Nuclei of cells were stained with 0.12 μg/ml Hoechst
stain (Thermo Fisher Scientific, Waltham, USA) for ten minutes. All secondary
antibodies utilized were conjugated to either Alexa Fluor 488 or Alexa Fluor 555
(Life Technologies, Carlsbad, USA) dyes.
2.3.4 Surface Expression Of Membrane-Bound Regulators On BOEC
Cells were seeded overnight in a 6 well plate (Falcon, Becton Dickinson, Franklin
Lakes, USA) to confluence, washed 1x with HBSS and then trypsinized (0.05%
Trypsin/0.53mM EDTA, WISENT). The primary antibodies (anti-CD46, Santa Cruz
Technology, Dallas, USA, sc-9098; anti-CD55, R&D, Minneapolis, USA, AF2009;
anti-CD59, AbD Serotec, Oxford, UK, MCA715G) and secondary antibodies (Alexa
fluor 488, Invitrogen, Life Technologies, Carlsbad, USA) were incubated at 4° for 20
minutes. Primary antibodies were used at a dilution of 1:100, secondary antibodies at
a 1:200 dilution. Twenty thousand events were recorded using Attune Acoustic
Focusing Cytometer (Life Technologies, Carlsbad, USA) and analyzed with FlowJo
31
software (FlowJo LLC, Ashland, USA). Results are given as median fluorescence
intensity (MFI). Surface expression was compared with that of glomerular endothelial
cells in passage 6.
2.3.5 Quantitative Gene Expression Of Membrane-Bound Regulators On
BOEC
Cells (BOEC and HUVEC) were seeded overnight in a 6 well plate (Falcon) to
confluence, washed 1x with HBSS before RNA was isolated using TRI Reagent
(Sigma-Aldrich, St. Louis, USA, T9424) according to manufacturer‟s instructions.
RNA concentration and integrity was verified by spectrophotometer (NanoDrop 1000,
Thermo Fisher Scientific, Waltham, USA), and reverse transcribed using
ReadyScript™ cDNA Synthesis Mix (Sigma-Aldrich, St. Louis, USA, RDRT).
Samples (200 ng cDNA in diethyl pyrocarbonate (DEPC, Sigma-Aldrich) treated
water) were amplified by real time polymerase chain reaction (PCR) using StepOne™
System from Life Technologies (Carlsbad, USA). Amplified products were detected
using KiCqStart™ SYBR® Green qPCR ReadyMix™, with ROX™ (Sigma-Aldrich,
Carlsbad, USA, KCQS02) and analyzed as follows: 2–(C
T – C
T GAPDH) – C
T control)
.
The following oligonucleotide primers (Sigma-Aldrich, Carlsbad, USA) were used:
GAPDH: forward, 5‟-ACAGTTGCCATGTAGACC-3‟; reverse, 5‟-
TTTTTGGTTGAGCACAGG-3‟.
VWF: forward, 5‟-TGTATCTAGAAACTGAGGCTG-3‟; reverse, 5‟-
CCTTCTTGGGTCATAAAGTC-3‟.
CD46/MCP: forward, 5‟-AGTGGTCAAATGTCGATTTC-3‟; reverse, 5‟-
ATCCCAAGTACTGTTACTGTC-3‟.
CD55/DAF: forward, 5‟-CAGAGGAAAATCTCTAACTTCC-3‟; reverse, 5‟-
AGTTGGTGAGACTTCTGTAG-3‟.
CD59: forward, 5‟-CATTACCAAAGCTGGGTTAC-3‟; reverse, 5‟-
TTTCTCTGATAAGGATGTCCC-3‟.
32
2.3.6 Complement Challenge Of BOEC
2.3.6.1 50% normal human serum in alternative pathway buffer
Serum was collected from whole blood of adult donors into serum vacutainers
(Becton Dickinson, Franklin Lakes, USA), allowed to clot for 30 minutes and then
centrifuged at 3000 x g at 4°C. Serum was collected and stored at -20°C until needed
as a source for complement. Heat-inactivated serum (HIS), for use as a negative
control, was obtained by incubating the serum for 30 minutes at 56°C.
2.3.6.2 Membrane-anchored complement regulator blockade
In order to simulate a CD46/MCP mutation, a monoclonal mouse anti-human anti-
CD46 antibody (GB24, IgG1, kindly provided by John Atkinson, St. Louis, USA) was
used.(Turner et al., 1996) GB24 efficiently and completely blocks C3b binding and
cofactor activity of CD46/MCP in in vitro assays by binding to the complement
control proteins (CCPs) 3 and 4 of CD46/MCP. (Liszewski et al., 2000; Turner et al.,
1996) Additional blockade of endothelial cell surface complement regulation was
achieved with the use of a monoclonal anti-human CD55/DAF (BRIC216, IgG1) and
monoclonal anti-human CD59 (BRIC229, IgG2b) (International Blood Group
Reference Laboratory, NHS Blood and Transplant, Bristol, UK) functional blocking
antibodies. These antibodies are known to be non-complement activating. Antibodies
were used at a concentration of 5μg/mL (derived from antibody titration experiments
performed by FACS, Supplemental figure 2-S1) and diluted in serum free cEGM-2
(Lonza) media for 30 minutes in all experiments. To ensure antibodies were targeting
the correct receptors, BOECs were grown on 22x22 mm coverslips (VWR) coated
with 0.05 mg/ml of rat tail collagen type I (Becton Dickinson) in six well tissue
culture plates, fixed with 4% paraformaldehyde (Electron Microscopy Sciences) in
PBS and stained with the blocking antibodies and with one of the following
antibodies: goat anti CD55/DAF (R & D Systems), rat anti CD59 (AbD Serotec) or
rabbit anti CD46/MCP (Santa Cruz). (Supplemental figure S1)
2.3.6.3 Complement deposition detected by immunofluorescence
For IF demonstration of complement deposition (C3 and C5b-9), BOEC were grown
on collagen-coated coverslips and then incubated for 60-240 minutes (37°C, 5% CO2)
33
with either cEGM-2 (BOEC) media, 50% normal human serum (NHS) in AP buffer
alone or after complement regulator blockade. After 60-240 minutes the supernatant
was aspirated and the cells were washed 3 times with ice cold PBS, blocked with 1%
(w/v) BSA for 1h and incubated with rabbit polyclonal antibody to C3c, (1:1000
dilution; Abcam, Cambridge, UK, ab15980) or a rabbit polyclonal antibody to C5b-9
(1:1000 dilution; Abcam, Cambridge, UK, ab55811) overnight.
2.3.6.4 Complement deposition detected by FACS
C3b deposition was demonstrated by FACS using a polyclonal rabbit anti-human
antibody to C3c (Abcam, ab15980) that detects C3c as well as the C3c part of native
C3 and C3b. Cells were seeded overnight in a 6 well plate (Falcon) to confluence,
washed 1x with HBSS before blocking antibodies (GB24, BRIC216, BRIC229) were
added for 20 minutes in serum-free media, followed by 50% NHS in AP buffer
(20mM HEPES pH7.4, 144mM NaCl, 7mM MgCl2, 10mM EDTA) for 1 hour. Cells
were trypsinized (0.05% Trypsin/0.53mM EDTA, WISENT) and washed in Flow
buffer (FB, 1% FBS/PBS) twice. The primary antibody in FB and the secondary
antibody (R-Phycoerythrin-conjugated AffiniPure F (ab‟)2 Fragment Donkey Anti-
Rabbit IgG (H + L), Jackson ImmunoResearch Laboratories, West Grove, USA, 709-
116-149, 1:200) together with Fixable Viability Dye eFluor780 (eBioscience, San
Diego, USA, 65-0865, 1:1000) in PBS were incubated at 4° for 20 minutes. At least
ten thousand events of BOEC population were recorded using Attune Acoustic
Focusing Cytometer (Life Technologies) and analyzed using FlowJo software.
Results are given as median fluorescence intensity (MFI). Cells were gated for live
cells (red laser 536nm, emission channel 2), single cells (FSC-A vs. FSC-H) and
finally through forward scatter and side scatter to determine the BOEC population.
C3c was recorded via the blue laser 488nm, emission channel 2. To correct for
spectral overlap during multicolor flow cytometry experiments, color compensation
was performed each time.
34
2.3.7 Cell death assays
2.3.7.1 Trypan blue exclusion
BOECs (between passages 3-14) were grown in 0.05 mg/ml rat-tail collagen type I
coated 96 well tissue culture plates (Sarstedt, Nümbrech, Germany). Depending on
the treatment, cells were incubated for thirty minutes at 37°C with one of anti-
CD46/MCP, anti-CD55/DAF, anti-CD59, or all three regulator functional blocking
antibodies in serum free EGM-2 media with supplements (Lonza) as described in the
BOEC Isolation section. The controls were incubated with media only. Following
incubation, cells were exposed to either media or 50% normal human serum from
adult donors in AP buffer. This treatment lasted thirty minutes at 37°C. Then cells
were washed twice with phosphate buffered saline (PBS: Wisent, St Bruno, Canada).
1:1 mixture of Trypan Blue (Sigma-Aldrich, St. Louis, USA) in PBS was added to
cells for five minutes. 4% paraformaldehyde in PBS was utilized to fix cells for ten
minutes. Two fields of cells at 10x magnification from each of duplicate wells were
counted by using a Leitz DM IL microscope (Leica Microsystems). The values for
dead cells were calculated by finding the percentage of overall cells that were dead.
Standard deviation was calculated to create error bars and the student‟s t-test (two-
tailed) was utilized to determine significance (p<0.05).
2.3.7.2 Cell cytotoxicity/LDH assay
BOEC were seeded overnight in a 96 well ELISA plate (Sarstedt) to confluence,
washed 1x with HBSS (Gibco) before complement-blocking antibodies was added for
30 minutes in serum-free media (cEGM-2). After washing twice, 10% NHS in serum-
free media was added for 4 or 6 hours accordingly. Pierce LDH cytotoxicity assay kit
(Thermo Fisher Scientific, Waltham, USA) was used according to manufactures
instructions. OD was normalized to positive control and displayed in percent.
2.3.7.3 Apoptosis
Cells were seeded in 96-well ELISA plates (Sarstedt) and incubated with functional
blocking antibodies for 30 minutes followed by 50% NHS in serum-free media
(cEGM-2) for four hours. Cells were fixed and blocked as described earlier and
35
stained for cleaved caspase-3 (Cell Signaling, Danvers, USA, 1:400 dilution) and
respective secondary antibody conjugated with Alexa fluor 555.
2.3.8 Platelet Adhesion
2.3.8.1 Platelet isolation
For platelet isolation, whole blood was collected from healthy adult donors and mixed
in a 6:1 ratio with acid citrate dextrose (ACD: 22.9 mM citric acid, 44.9 mM sodium
citrate dehydrate, 74 mM dextrose monohydrate). Platelet rich plasma (PRP) was
collected after centrifugation at 160 x g for ten minutes. PRP was spun at 950 x g for
seven minutes to pellet platelets. Platelets were washed twice with PBS/ACD solution
(20% ACD in PBS; pH 6.1). Platelets were incubated with 2.5 μM calcein (Life
Technologies, Carlsbad, USA) for thirty minutes at 37°C Finally, platelets were
pelleted at 950 x g and resuspended at a concentration of 15x107/ml in Tyrodes buffer
(136 mM NaCl, 2.7 mM KCl, 0.42 mM NaH2PO4, 19 mM NaHCO3, 5.5 mM of
glucose, 1 mM CaCl2, 1 mM MgCl2 and 10 mM of hyroxyethyl
piperazineethanesulfonic acid (HEPES: Invitrogen, Life Technologies, Carlsbad,
USA)). Platelet count was measured using an automatic hematocytometer (Beckman
Coulter, Brea, USA).
2.3.8.2 BioFlux (Fluxion Biosciences) Microfluidic System
For BOEC-Platelet Interaction experiments, BOEC (passage 3-14) were seeded into
the BioFlux System (Fluxion Biosciences, South San Francisco, USA) channels the
day before the actual experiment. BOECs were grown in BioFlux 48 well tissue
culture plates coated with 0.05 mg/ml rat-tail collagen type I (Becton Dickinson). The
collagen was added to the output well and flown backward at a shear rate of
3dyne/cm2 for 1-2 minutes after which the plate was incubated overnight at 37°C.
BOEC were trypsinized and up concentrated to 8 x106 cells/mL. After adding 50 μl of
cEGM-2 to the input well as a balance, 50 μl of the cell solution were transferred to
the output well and then perfused backwards for 15 seconds at 1 dyne/cm2. After
sufficient cell adhesion was observed under the microscope, the plate was put into the
37°C incubator for 1 hour before adding 1 ml of cEGM-2 to each input well. Cells
were left in a 37°C / 5% CO2 incubator overnight. Cells were washed at 1 dyne/cm2
36
with Hank‟s balanced salt solution (HBSS). Complement challenge was achieved by
flowing 50% NHS in alternative pathway (AP) buffer over the cells either, (i) alone,
(ii) after cells were pretreated with an anti-CD46/MCP function blocking antibody, or
(iii) after cells were pretreated with an antiCD46/MCP, anti-CD55/DAF and anti-
CD59 functional blocking antibodies. Blocking antibodies were flown over the cells
in serum free EGM-2 media (Lonza) while incubated at 37°C for thirty minutes
before washing with HBSS. 50% NHS was flowed through the BioFlux chambers at 2
dynes/cm2 for 1-2 hours. For platelet adhesion assays, 100 x 10
6 calcein (Life
Technologies) labeled platelets in Tyrodes buffer (as described in platelet preparation
section) were flowed through the chambers at 2 dyne/cm2 after BOEC exposure to
serum. After 5 to 10 min three pictures per channel were taken with a Leitz DM IL
microscope (Leica) at a magnification of 4x. Adhering platelets were counted using
ImageJ software.
2.3.9 Ethics Approval
Ethics approval was obtained from the Research Ethics Board at The Hospital for
Sick Children, Toronto, ON, Canada.
2.3.10 Statistical analysis
Figures were generated using GraphPad Prism 6 and displayed as mean and standard
deviation. Statistical analysis was performed using either two-way ANOVA with
post-hoc analysis or paired t-test. A p value < 0.05 was considered as statistically
significant. In figures p values are presented as follows: * ≤0.05, ** ≤0.01, ***≤0.001,
****≤0.0001, ns > 0.05.
37
2.4 RESULTS
2.4.1 BOEC Have An Endothelial Cell Phenotype And Retain Their
Phenotype Over Various Passages
BOEC were isolated from the peripheral blood of four healthy adult donors. BOEC
appeared after a mean of 9 (range 7-20) days and displayed a typical cobblestone
appearance when visualized by light microscopy (Figure 2-1 A). An endothelial
phenotype was confirmed by demonstrating the presence of Weibel-Palade bodies
containing fluorescently labelled VWF and P-selectin (CD62P) imaged by confocal
spinning disk immunofluorescence microscopy (Figure 2-1 C, 2-1 D). BOEC can be
confirmed as late outgrowth endothelial colony forming cells as distinct from early
endothelial progenitor cells by the presence of specific endothelial cell markers and
the absence of certain surface markers.(Timmermans et al., 2009; Timmermans et al.,
2007) By flow cytometry cells were positive for the endothelial cell markers
CD31/PECAM-1 (platelet-endothelial cell adhesion molecule-1), and CD144 (VE-
Cadherin) [HUVECs as positive controls] and negative for the hemangioblast marker
CD45 and the immature EPC marker CD14 [blood as negative control]), consistent
with a BOEC phenotype (Figure 2-1 B). In order to confirm the stability of the cell
line across passages we showed that VWF expression by qPCR was the same at an
early (passage 3) and a late passage (passage 10) (Figure 2-1 E).
38
39
Figure 2-1. BOEC have an endothelial cell phenotype and retain their phenotype over
various passages
(A) BOEC are derived from mononuclear cells in peripheral blood that are cultured in cEGM-
2 medium. During the first week early endothelial progenitors (eEPCs) and fibroblasts are
seen. From week 2-3, blood outgrowth endothelial cells appear and form a typical
„cobblestone‟ monolayer as the eEPCs die off. (B) An endothelial phenotype was confirmed
by demonstrating the presence of Weibel-Palade bodies containing fluorescently labeled von
Willebrand factor (VWF, Alexa fluor 555; red) and P-selectin (CD62P, Alexa fluor 488;
green). We used an Olympus IX81 inverted fluorescence microscope using a 60/1.35 oil
immersion objective and a Hamamatsu C9100-13 back-thinned EM-CCD camera with
Yokogawa CSU X1 spinning disk confocal scan head. Confocal images were taken with an
Improvision Piezo Focus Drive. Z-stacks were taken at 0.25 μm. Images taken using the
spinning disk confocal microscope were deconvolved by iterative restoration using Volocity
Software with confidence limit set to 95% and iteration limit set to 20. (C) By flow cytometry
we further confirmed their endothelial phenotype, as BOEC are positive for PECAM/CD31
and VE-Cadherin/CD144 and negative for the hemangioblast marker, CD45 and immature
EPC marker, CD14. (D) By quantitative PCR we confirmed that BOECs are stable at an
early (passage 3) and late passage (passage 10) for one endothelial cell marker (VWF).
mRNA expression was normalized to GAPDH and displayed relative to VWF
expression of passage 3. N=2.
2.4.2 BOEC Express Key Membrane-Anchored Complement Regulators
We next wanted to confirm the presence of the key membrane-anchored complement
regulators on BOEC and compare this to HUVEC, a primary endothelial cell line used
in the study of aHUS pathomechanisms. The presence of these regulators
(CD46/MCP, CD55/DAF, CD59 and CD141/Thrombomodulin) was demonstrated by
using confocal, spinning disk microscopy of immunofluorescently labeled BOEC
grown to confluence on coverslips (Figure 2-2 A-C). The baseline gene expression
levels of CD46/MCP, CD55/DAF and CD59 were similar to that of HUVEC, when
assessed by qPCR (Figure 2-2 D). In both HUVECs and BOECs, there is a
significantly higher level of CD59 as compared to the other regulators CD46/MCP
40
and CD55/DAF. This gene expression pattern was replicated by flow cytometry
assessment of cell surface expression of the proteins, where BOEC demonstrated a
similar amount of CD46/MCP and CD55/DAF (p=0.8) and a 14-fold higher relative
amount of CD59 on their surface (as compared to CD46; p=0.002) (Figure 2-2 E-F).
This was confirmed in three different control BOEC lines. The expression of
CD46/MCP, CD55/DAF and CD59 was also similar to that found on glomerular
endothelial cells. (Supplementary figure 2-S2)
41
Figure 2-2. BOEC express key membrane-anchored complement regulators
(A-C) Complement regulators CD46/MCP (A), CD55/DAF (B) and CD59 (C) were
fluorescently labeled and imaged on BOEC by confocal spinning disk microscopy (see Figure
1 for details). (D) By qPCR we compared the relative expression of complement regulators as
compared to VWF (VWF expression=1). * p<0.05. Results were compared to expression
levels in HUVEC. No significant difference was observed (two-way ANOVA). N=6 for
HUVEC and N=5 for BOEC. (E-F) By flow cytometry we quantified a 14-fold higher
expression of CD59 (green) compared to CD46/MCP (blue) and a 9-fold higher expression
compared to CD55/DAF (orange) in three different control BOEC lines. No significant
difference was observed between surface expression of CD46/MCP and CD55/DAF. ** p<
0.01 compared to CD46 and CD55 (paired t-test). Median fluorescence intensity (MFI) was
calculated as fold increase over unstained control. N=3 In (E) one representative flow
cytometry reading of control BOECs is shown.
2.4.3 Cell Surface C3b Deposition Mirrors Incremental Functional
Blockade of CD46/MCP, CD55/DAF and CD59
The functional relevance of aHUS-associated CD46/MCP mutations has, to date, been
studied in a Chinese hamster ovary cell model system. When CHO cells that have
been transfected with the mutant protein are incubated with normal human serum, a
measurable increase in C3b deposition is demonstrated by flow cytometry, as
compared to wild type CD46/MCP expressing cells. (Goodship et al., 2004;
Liszewski et al., 2007; Richards et al., 2007) By contrast, we employed a model
system of BOEC, as these cells physiologically express membrane-bound
complement regulators, in contrast to CHO cells. 50% NHS (in AP buffer) was added
to confluent BOEC for a one hour period, either alone or after functional blockade of
the membrane regulators. Serum was removed; the cells were washed with HBSS,
then trypsinized and taken into suspension for FACS. Cells were labeled with a
polyclonal rabbit anti-human antibody to C3c and showed no significant increase in
C3 deposition over that seen by incubating the cells with 50% NHS alone when
CD46/MCP complement regulatory function was blocked (Figure 2-3 A-B, p>0.05).
Additional blockade of CD55/DAF and CD59 resulted in a 7-fold increase of C3c
deposition (p<0.05). On immunofluorescence an increase in C5b-9 deposition was
visualized when three surface-bound regulators were blocked and cells were perfused
42
with 50% NHS for one hour in the BioFlux at a rate of 2 dynes/cm2, as compared to
single regulator blockade (Figure 2-3 D-F).
Figure 2-3. Cell surface C3b deposition mirrors incremental functional blockade of
CD46/MCP, CD55/DAF and CD59
(A-B) C3 deposition was evaluated by flow cytometry using an anti-C3c antibody (detects C3,
C3b, C3c but not C3a and C3d). (A) shows a representative figure, with the following
conditions: unstained (grey), 50% NHS (light green), CD46/MCP blocked prior 50% NHS
(dark green) and CD46/55 and 59 blocked prior adding 50% NHS (red) for one hour. In
43
figure B the median fluorescence intensity (MFI) of 7-8 different experiments is shown
(*p<0.05, paired t-test).
(C-E) BOEC, cultured in a microfluidic chamber were perfused with 50% NHS for 1h.
BOEC in channel D were pretreated with an anti-CD46/MCP antibody and cells in channel E
with an anti-CD46/MCP, anti-CD55 and anti-CD59 antibody for 30 minutes. An increasing
C5b-9 deposition (polyclonal rabbit anti human C5b-9, 1:1000, secondary antibody: Alexa
fluor 555, 1:1000) was observed only when CD46/MCP, CD55/DAF and CD59 were blocked.
VWF was stained using a polyclonal sheep anti-human antibody (1:1000, secondary antibody:
Alexa fluor 488) and DNA using Hoechst dye (0.12μg/ml). Images were taken on oil-
immersion 60x magnification with a Nikon Eclipse Ti camera.
2.4.4 Complement-mediated BOEC cytotoxicity increases with
incremental membrane-regulator blockade
Current understanding of aHUS pathogenesis suggests that a dysregulated
complement AP can cause injury to the microvascular endothelium leading to EC
death and retraction. (Noris and Remuzzi, 2009) BOEC death was first assessed by a
Trypan blue exclusion assay. Confluent cells were subjected to an increasing
complement challenge with 30 minutes exposure to 50% NHS either alone, or after
functional blockade of the membrane regulators CD46/MCP, CD55/DAF and CD59.
Single regulator blockade failed to result in an increase in the percentage of dead cells
over that which occurs with 50% NHS alone. Intensifying the complement challenge
by the simultaneous blockade of three regulators, followed by exposure to 50% NHS,
resulted in a statistically significant increase in cell death (p=0.04) (Figure 2-4 A).
The same result was replicated when measuring LDH release after incubation with
10% NHS for 4 and 6 hours. Due to the high amount of LDH in serum this assay
could only be performed supplementing 10% serum to serum-free media (Figure 2-4
B). Furthermore, dead cells at 4 hours post complement challenge stained positive on
immunofluorescence for cleaved caspase-3 implicating apoptosis as the principal
mode of endothelial cell death after complement challenge at this timepoint (Figure 2-
4 C-E).
44
Figure 2-4. Complement-mediated BOEC cytotoxicity increases with incremental
membrane-regulator blockade
(A) BOEC grown to confluence in a 96-well plate were exposed to 50% NHS for 30 minutes
after blocking complement regulators. Trypan-blue was added and cells counted on a Leitz
DM IL microscope (Leica) with a 10x magnification. Per well 2 random fields were counted,
two wells per permutation for a total of 4 assays. A significant increase of dead cells was seen
after blocking CD46/CD55/CD59 when compared to cells treated only with 50% NHS. N=4,
*p=0.04 (paired t-test). (B) This was confirmed by measuring release of LDH after incubating
confluent BOEC with 10% NHS in serum-free media for four and six hours, N=5 (blockade
of CD46/CD55/CD59 versus 50% NHS alone; p=0.03 and blockade of CD46/CD55/CD59
versus CD46 blockade; 0.02 respectively) (C-D) After incubation with 50% NHS for four
hours cells were fixed and stained for cleaved caspase 3 (1:400) and donkey anti rabbit Alexa
fluor A555 (red). DNA was stained using Hoechst dye (0.12μg/ml, blue). Cells were grown in
96-well plates for images taken at 20x magnification. (C) Representative image after
pretreatment with anti-CD46 antibody and (D) was taken with same Nikon Eclipse Ti camera
at 20x after pretreatment with anti-CD46/CD55/CD59 antibodies.
45
2.4.5 Increasing complement challenge to BOEC results in an increase in
platelet adhesion
Atypical HUS is characterized clinically by thrombocytopenia, and pathologically by
the presence of platelet microthrombi.(Noris and Remuzzi, 2009) In order to model an
aHUS/TMA phenotype ex vivo we employed an endothelialized, microfluidic system,
so that calcein-labeled platelets could be perfused across a confluent monolayer of
BOEC and visualized for adhesion, after the BOEC had been exposed to a
complement challenge with 50% NHS alone or after regulator blockade. Minimal
platelet adhesion was demonstrated after the cells were exposed to complement active
serum for one hour or if the function of just CD46/MCP was blocked. However,
significant platelet adhesion occurred within 5-10 minutes of flowing the platelets
through the microfluidic channels when we augmented the complement challenge by
blocking the function of three surface complement regulators concurrently (p=0.028)
(Figure 2-5).
46
Figure 2-5. Increasing complement challenge to BOEC results in a stepwise increase in
platelet adhesion
(A-B) Washed and calcein-labeled platelets in Tyrodes buffer (15 x 107/ml, 100μl/well) were
introduced into a microfluidic flow chamber and perfused at 2 dyne/cm2 from inlet to outlet
well. BOEC were pretreated with respective blocking antibody and 50% NHS in AP buffer
for 1 hour. Images were taken with a Nikon Eclipse Ti camera at 4x magnification after 5
minutes. A white line was added to better visualize upper and lower channel. In figure 5A the
upper chamber represents platelet adhesion after pretreatment with 50% NHS only, the lower
channel after pretreatment with CD46/MCP blocking antibody and 50% NHS. Figure 5B
shows in upper channel platelet adhesion after pretreatment with CD46/MCP blocking
antibody and 50% NHS and in lower channel adhesion after pretreatment with
CD46/CD55/CD59 blocking antibodies and 50% NHS for 1 hour. (C) Per viewing chamber,
three random pictures after 5 minutes of perfusion were taken and adhering platelets were
counted using ImageJ software. Number of platelets was normalized to number counted when
channels were perfused with media. N=6. * p<0.05 (paired t-test) (D) Image was taken in
microfluidic chamber using a Nikon Eclipse Ti microscope at 60x magnification oil
47
immersion after BOEC were pretreated with CD46/55/59 antibodies, followed for 1 hour with
50% NHS and 10 minutes of platelets in Tyrodes buffer. Cells were fixed with 4% PFA/PBS,
blocked with 1% BSA/0.2% Triton X-100/PBS and stained for C5b-9 (rabbit polyclonal
antibody, 1:1000, red) and VWF (sheep polyclonal antibody, 1:1000, green) overnight. Alexa
fluor 488 and 555 (1:1000 for 1 hour) was used as secondary antibodies, Hoechst (0.12μg/ml,
10 minutes, blue) for DNA stain. Arrow indicates adhering platelets.
48
2.4.6 Discussion
CD46/MCP acts as a cofactor for the serine protease CFI to inactivate cell surface
bound C3b. It is one of three principal membrane-anchored complement regulators
present on the surface of endothelial cells but the only one associated with aHUS.
Mutations in CD46/MCP, similar to other mutations that cause aHUS such as CFH
and CFI, are associated with low penetrance, but MCP/CD46-associated aHUS
generally has a more favorable outcome.(Caprioli et al., 2006) A mutation in
CD46/MCP may be the sole mutation demonstrated in patients with aHUS, or
MCP/CD46 variants may also represent a risk polymorphism, increasing the chance
of disease manifestation in a patient with another genetic susceptibility. (Bresin et al.,
2013; Provaznikova et al., 2012) Explaining how a mutation in CD46/MCP actually
leads to a TMA phenotype has not been clearly delineated.
In this study we found that a complete functional blockade of CD46/MCP, using a
monoclonal antibody directed towards its C3b binding site, was not followed by a
significant increase in cell surface C3 deposition, cell death, or platelet adhesion in an
ex vivo endothelial cell-based model system. In order to achieve the purported
sequence of events that lead to a TMA with complement deposition, endothelial cell
activation or death, and platelet adhesion, (Noris and Remuzzi, 2009) we had to
amplify the complement challenge by the synchronous/concomitant blockade of three
surface regulators. Only then, could we demonstrate a measurable increase in cell
surface C3b deposition, platelet adhesion and cell death. This implies that the
endothelial cell is well protected against complement injury and cumulative hits –
either endogenous or exogenous in nature – may be necessary to overcome this
protective barrier.
Experimental evidence of synergy between CD46/MCP and CD55/DAF already
exists, (Liszewski et al., 2007) and clinically, the presence of a risk haplotype in both
CD46/MCP and CFH doubles the penetrance as opposed to having just one, or neither,
and the combination doubles the risk of end stage renal disease. (Bresin et al., 2013)
Of course there may be an inciting „trigger‟, for instance, in one series, 100% of those
with aHUS and an CD46/MCP mutation had so in association with an infectious
49
trigger. (Caprioli et al., 2006)
This study is the first study using blood outgrowth endothelial cells to study the
pathogenesis of a TMA - in this case CD46/MCP associated aHUS. BOEC are
thought to be a true EPC that possess distinctly endothelial cell characteristics and
behaviors. (Medina et al., 2010b; Tura et al., 2013) They are highly proliferative and
as compared to HUVEC they retain their phenotype over passages and are more
resistant to cell death. (Bompais et al., 2004) Already studied in various disease
models, (Medina et al., 2012) BOEC have significantly enhanced the understanding
of von Willebrand Disease by taking advantage of the fact that these cells can be
grown from patients expressing mutant endothelial cell proteins. (Starke et al., 2013;
Wang et al., 2013a) BOEC express the surface complement regulators similarly to
HUVEC, as we have shown, however in contrast to HUVEC, are highly proliferative
and stable over passages. Phenotypically they resemble more closely microvascular
endothelial cells, (Gremmels et al., 2011; Jiang et al., 2007; Toshner et al., 2014)
which make them an ideal cell type to study TMA pathogenesis, a disease of the
microvascular endothelium.
CD46/MCP mutations have been studied functionally, by either quantifying the
expression levels on peripheral blood mononuclear cells, or via transfection of CHO
cells, cells devoid of any complement regulators. (Caprioli et al., 2006) Studies of
transfected Chinese hamster ovary (CHO) cells have found that this reduced
expression translates into reduced regulation of the complement AP with increased C3
deposition and this is then thought to lead to enhanced formation and insertion of the
Membrane Attack Complex/ C5b-9 (MAC/C5b-9) into cells with endothelial cell
activation and/or death. (Liszewski et al., 2007) Furthermore, when CD46/MCP
expression levels were reduced by 50% on CHO cells, thus mimicking the situation of
a heterozygous mutation commonly associated with CD46/MCP-associated aHUS,
less efficient complement regulation and more C3b deposition was also found.
(Liszewski et al., 2007) Why we did not find more C3b deposition in the face of
efficient blockade of MCP and complement activation may speak to a synergistic
defensive effect of the other regulators.
The importance of conducting functional studies to characterize any given mutation as
50
disease causing, even if the mutation is predicted to be pathogenic, has recently been
emphasized for aHUS associated with complement Factor B mutations. (Marinozzi et
al., 2014) Even then, explaining how a mutation leads to disease may not be clearly
evident. We believe that using our system incorporating BOEC and an endothelialized
microfluidic chamber that we are better able to model the in vivo scenario.
Our study has limitations of course, and the precise role of the other complement
regulators such as CFH was not studied. Platelet adhesion was assessed after one hour
of complement challenge and perhaps with more prolonged complement challenge
then a difference in CD46/MCP-blocked cells might have emerged, however time was
limited by having the cells outside of the 5% CO2 incubator.
In conclusion, our study suggests that the effect of a loss of CD46/MCP alone, on an
otherwise normal cell, endowed with the full complement of regulators, is probably
minimal. Our data supports the notion of a „multiple hit‟ – that an additional genetic
abnormality in the complement system (here simulated by blocking three surface
regulators) is needed for an individual with a CD46/MCP mutation to manifest the
disease. BOEC represent a novel tool to study aHUS pathogenesis being more
physiologically relevant than CHO cells, in that they express the complement
regulators, but more importantly, because they can now be isolated from patients with
mutations in CD46/MCP, enabling the study, ex vivo, of the „real‟ protein phenotype.
51
2.4.7 Supplemental Data
Figure 2-S1. Functional blocking antibody binding and saturation
(A-C) Using flow cytometry the antibody saturation of anti-MCP antibody (GB24, A), anti-
CD55/DAF antibody (BRIC216, B) and anti-CD59 antibody (BRIC229, C) was determined.
Antibody concentrations of 2μg/ml (blue), 5μg/ml (orange) and 10μg/ml (green) were
compared to unstained control (red). Subsequently, 5μg/ml was used in all experiments. (D-F)
Co-localization of blocking antibody with epitope was determined using immunofluorescence.
The blocking antibodies were detected with secondary donkey anti mouse labeled with A555
(red) and staining antibody with representative Alexa fluor 488 (green). Hoechst dye (0.12
μg/ml, blue) was used to stain DNA. Pictures were taken with a spinning disk confocal
microscope as described in Methods and Figure legend 2-1.
52
Figure 2-S2. Expression of surface-bound regulators on glomerular endothelial cells
compared to BOEC
(A-C) The expression of surface-bound complement regulators CD46/MCP, CD55/DAF and
CD59 on glomerular endothelial cells (GEC, green) and BOEC (purple) was determined
using flow cytometry. A similar expression pattern was observed for CD46/MCP (A),
CD55/DAF (B) and CD59 (C). A curve in grey represents unstained controls. N=1 (D)
mRNA expression of complement regulators were determined in control BOEC and GEC
using quantitative real-time PCR. Significant increased mRNA levels for GEC were
determined. N=3 in one experiment. * p<0.05, **** p<0.001 (two-tailed ANOVA).
53
CHAPTER 3 VON WILLEBRAND FACTOR
MULTIMERS CONTRIBUTE TO COMPLEMENT
ALTERNATIVE PATHWAY CONTROL
54
3.1 Abstract
Recent in vitro evidence supports a link between the alternative pathway of
complement (AP) and VWF based on the co-localization of complement activation
proteins and VWF multimers. It has been suggested that this link could possibly
amplify EC complement activation and therefore might have significant implications
for diseases where the crossover between complement and coagulation is evident,
especially the thrombotic microangiopathies such as atypical hemolytic uremic
syndrome (aHUS) and thrombotic thrombocytopenic purpura (TTP). Clinical
observations have shown a link between aHUS and TTP, as the phenotypes of both
entities significantly overlap. In addition, aHUS patients show variants in
ADAMTS13 and TTP patients show elevated complement activation parameters in
the acute phase of the disease. We aimed to use blood outgrowth endothelial cells
(BOEC), EC precursors that can be isolated from blood of patients, to further study
the role of VWF in the pathogenesis of aHUS and the link between the AP of
complement and VWF.
BOEC were cultured by a standard protocol from healthy controls and two patients
with type 3 von Willebrand disease patients (VWD3 BOEC), which have no VWF
secretion. EC phenotype and presence of complement regulators (CD46/MCP,
CD55/DAF, CD59) were confirmed by immunofluorescence (IF), western blot (WB),
qPCR and flow cytometry (FACS). Cells were challenged with complement using
50% normal human serum (NHS), with or without functional blockade of
complement regulators. BOEC death was assayed by LDH release.
BOEC isolated from healthy adult controls and two patients with type 3, von
Willebrand disease (VWD3), show similar surface expression of the tested
membrane-bound complement regulators. VWD3 BOEC are distinguishable for
having no visible Weibel-Palade bodies or VWF on immunofluorescence. VWF
secretion was confirmed from control BOEC after complement activation.
Immunofluorescently labelled complement products, including C5b-9, were
visualized in association with/in proximity to VWF released from complement
challenged control BOEC by spinning disk confocal microscopy. However,
complement (C3b) deposition was significantly greater on VWD3 BOEC following
complement challenge and these cells succumb more easily to complement-mediated
cytotoxicity. Taken together we have confirmed an association of VWF and
55
complement AP activation products and the finding of increased complement
activation on BOEC devoid of VWF argues for a protective effect of this association.
56
3.2 Introduction
Atypical hemolytic uremic syndrome (aHUS) and thrombotic thrombocytopenic
purpura (TTP) represent two diseases that present a thrombotic microangiopathy
(TMA) phenotype. Atypical HUS is a TMA that occurs due to dysfunctional
regulation of the complement system with microvascular endothelial cell injury and
the formation of platelet microthrombi. (Noris and Remuzzi, 2009) TTP is caused by
either a deficiency of a disintegrin and metalloproteinase with a thrombospondin type
1 motif member 13 (ADAMTS13) or autoantibodies that functionally inhibit its
function, allowing ultra large VWF multimers to accumulate, bind platelets and
produce occlusive thrombi. (Tsai, 2010)
Although TTP and aHUS have significant clinical overlap, making diagnosis a
challenge, they represent two distinct pathologies, both ultimately leading to a
microangiopathic hemolytic anemia. (Tsai, 2013) However, the line distinguishing
them may not be so clear-cut, and aHUS and TTP may well represent part of a
spectrum of complement-associated diseases. (Noris et al., 2012) The finding of a low
C3/hypocomplementemia in aHUS and TTP patients, (Noris et al., 1999) along with
the experimental observation that serum from TTP patients could result in
complement deposition on endothelial cells, (Ruiz-Torres et al., 2005) and the finding
of increased complement alternative pathway (AP) activity in TTP patients, (Feng et
al., 2013b) seems to confirm an overlap between the two entities. Furthermore, a
recent study found that 80% of an aHUS patient cohort also carried at least 1 non-
synonymous change in ADAMTS13, and in 38% of patients, multiple ADAMTS13
variations were found. These ADAMTS13 variants were thought to be clinically
relevant as measured ADAMTS13 activity was less than 60% in half of the patients
studied. (Feng et al., 2013a) This also argues in favor of an interconnection between
the two diseases. (Feng et al., 2013a)
Over the last year there have been a number of experimental studies further linking
the alternative complement pathway (AP) and its principal regulator, complement
Factor H (CFH), with von Willebrand Factor (VWF) and its cleaving protease,
ADAMTS13. (Feng et al., 2013c; Nolasco et al., 2013; Rayes et al., 2014; Turner and
57
Moake, 2013) CFH has been shown to be a reductase for large soluble VWF
(LsVWF) multimers that are secreted from ECs.(Nolasco et al., 2013) LsVWF
multimers released from complement activated ECs assume a conformation that
makes them inaccessible to ADAMTS13 cleavage, but can still bind platelets and
induce platelet aggregation under conditions of high-fluid shear stress. (Nolasco et al.,
2013) In addition, the C-terminus of CFH has been shown to enhance ADAMTS13
VWF cleavage in vitro. (Feng et al., 2013c) Most interesting perhaps has been the
finding that both the assembly, and activation, of complement AP components on
ULVWF strings secreted from, and anchored to, endothelial cells (ECs). (Turner and
Moake, 2013) We aimed to explore this link further using blood outgrowth
endothelial cells (BOEC) lacking functional VWF, derived from patient with von
Willebrand type 3 disease.
58
3.3 Methods
3.3.1 Establishing and characterizing blood outgrowth endothelial cells
(BOEC)
BOEC were isolated from the blood as described before in chapter 2. Briefly, 48ml
blood was drawn into cell preparation tube (CPT) vacutainers (Becton Dickinson,
Franklin Lakes, USA) and mononuclear cell layer was obtained by centrifugation at
1600xg for 30 minutes. Cells were washed twice in 10% fetal bovine serum (FBS:
Sigma-Aldrich, St. Louis, USA) in phosphate buffered saline (PBS, WISENT, St
Bruno, Canada), resuspended in cEGM-2 media with supplements (Lonza,
Walkersville, USA: Cat no. 362753) and seeded on six-well tissue culture plate
(Becton Dickinson, Franklin Lakes, USA) coated with 0.05 mg/ml of rat tail collagen
type I in 0.02 M acetic acid (Becton Dickinson, Franklin Lakes, USA). BOEC were
identified after 1-2 weeks by their endothelial cobblestone morphology and their
endothelial cell phenotype was confirmed by flow cytometry. BOEC are positive for
the endothelial cell (EC) surface markers CD31 (PECAM-1) and CD144 (VE-
Cadherin) while negative for the hemangioblast marker CD45 and the immature
endothelial progenitor cell marker CD14. BOEC were grown from healthy adult
donors (Control BOEC, CTRL BOEC) and patients with von Willebrand Disease type
3 (VWD BOEC).
3.3.2 Glomerular Endothelial Cell Culture
Glomerular endothelial cells (GEC) were cultured on T75 flasks (Sarstedt, Nümbrech,
Germany) coated with collagen (0.05 mg/ml of rat tail collagen type I in 0.02 M
acetic acid (Becton Dickinson, Franklin Lakes, USA)) in endothelial cell medium
(ScienceCell Research Laboratories, Carlsbad, USA, 1001) supplemented with
Endothelial Cell Growth Supplement (ECGS, ScienceCell Research
Laboratories ,1052), 5% fetal bovine serum (FBS, Sigma-Aldrich, St. Louis, USA)
and 1% Antibiotic-Antimycotic (Gibco, Invitrogen, Life Technologies, Carlsbad,
USA; containing 10,000 units/mL of penicillin, 10,000 µg/mL of streptomycin, and
25 µg/mL of Fungizone® Antimycotic). Cells were kept at 37°C and in an
environment with 5% CO2. Passages 4-7 were used for experiments.
59
3.3.3 Determining BOEC protein expression via Western blot
In order to determine VWF expression levels in control and VWD BOEC, we
performed a western blot analysis of BOEC lysates. BCA protein determination assay
was used prior to preparation of samples for western blot. BOEC lysates were
prepared in 4 x sample buffer (40% (v/v) glycerol, 8% (w/v) sodium dodecyl sulfate
(SDS), 0.26M Tris, pH 6.8, 0.05% (w/v) bromphenol blue) and heated at 95°C for 5
minutes. 50 μg of protein were loaded and separated by an 8% SDS-polyacrylamide
gel electrophoresis running at 200 V for 45 minutes (Power Source 250V, VWR
International, Radnor, USA). Protein gels were transferred onto a 0.45 μm
nitrocellulose membrane (Bio-Rad Laboratories, Hercules, USA) for 70 minutes at
100 V (VWR International, Radnor, USA). Membranes were blocked with 5% (w/v)
skim-milk in Tris-Buffered-Saline, pH 7.6, + 0.05% (v/v) Tween-20 (TBST) for one
hour at room temperature. Membranes were incubated with rabbit polyclonal
antibody to VWF (1:1000 dilution; Dako, Glostrup, Denmark) in 5% (w/v) skim-milk
in TBST overnight at 4°C with shaking. For loading controls, membranes were
incubated with mouse monoclonal antibody to β-actin (β-actin, BA3R, 1:10000
dilution; Pierce Biotechnology, Rockford, USA) in 5% (w/v) skim-milk in TBST for
1 hour at room temperature with shaking. Membranes were washed 3 times, with
TBST for 5 minutes, and incubated with secondary antibody in 5% (w/v) skim-milk
in TBST at room temperature for 1 hour with shaking. Membranes were washed 3
times with TBST for 10 minutes. Proteins were detected using Western Lighting™
Plus-ECL, Enhanced (PerkinElmer, Waltham, USA) and developed on Odyssey FC
Imaging System (chemiluminescence detection, LI-COR Biosciences, Lincoln, USA)
using Image Studio software (LI-COR Biosciences, Lincoln, USA). Quantification of
protein bands was performed using densitometry (ImageJ).
3.3.4 Quantifying BOEC and GEC Gene Expression via qRT-PCR
To quantify VWF mRNA expression we performed quantitative real time polymerase
chain reaction (qRT-PCR). Cells were seeded overnight in a 6 well plate (Falcon) to
confluence, washed once with Hank‟s balanced salt solution (HBSS: Gibco, Life
Technologies, Carlsbad, USA) before RNA was isolated using TRI Reagent (Sigma-
Aldrich, St. Louis, USA) according to manufactures instructions. RNA concentration
and integrity was verified using Nanodrop (NanoDrop 1000, Thermo Fisher Scientific,
60
Waltham, USA), and reverse transcribed using ReadyScript™ cDNA Synthesis Mix
(Sigma-Aldrich, St. Louis, USA, RDRT). Samples (200 ng cDNA in diethyl
pyrocarbonate (DEPC, Sigma-Aldrich) treated water) were amplified by real time
polymerase chain reaction (PCR) using StepOne™ System from Life Technologies
(Carlsbad, USA). Amplified products were detected using KiCqStart™ SYBR®
Green qPCR ReadyMix™, with ROX™ (Sigma-Aldrich, St. Louis, USA) and
analyzed as follows: 2–(C
T – C
T GAPDH) – C
T Control BOEC)
.
The following oligonucleotide primers purchased from Sigma-Aldrich, Carlsbad,
USA were used:
GAPDH: forward, 5‟-ACAGTTGCCATGTAGACC-3‟; reverse, 5‟-
TTTTTGGTTGAGCACAGG-3‟.
VWF: forward, 5‟-TGTATCTAGAAACTGAGGCTG-3‟; reverse, 5‟-
CCTTCTTGGGTCATAAAGTC-3‟.
3.3.5 Characterization of membrane-bound regulators of BOEC
For determination of surface expression of the membrane-bound regulators on VWD
BOEC we performed both immunofluorescence labeled imaging and flow cytometry.
3.3.5.1 Immunofluorescence
BOEC at a concentration of 0.5 × 106 cells/ml were seeded onto collagen-coated
22x22-mm cover slips (VWR International, Radnor, USA) and incubated overnight
(37°C, 5% CO2). Samples were washed with ice-cold PBS followed by fixation with
4% (w/v) paraformaldehyde (Electron Microscopy Sciences, Fort Washington, USA)
in PBS and blocked with 1% (w/v) BSA/PBS for 60 minutes. Samples were stained
overnight with the primary antibodies made up in the 1% (w/v) BSA/PBS solution.
Samples were washed in PBS and incubated for one hour with respective species
specific donkey-anti secondary antibodies conjugated with Alexa Fluor® 488 or
Alexa Fluor 555 (1:1000 dilution, Invitrogen, Life Technologies, Carlsbad, USA).
Cell nuclei were stained using 0.12 μg/ml Hoechst 33342 stain (Thermo Fisher
Scientific, Waltham, USA) for ten minutes. Samples were washed in PBS and cover
slips were mounted with Dako Fluorescence Mounting Media (Dako Canada,
Burlington, Canada) for microscopy. Images were taken with a spinning disk confocal
microscopy equipped with an Olympus IX81 inverted fluorescence microscope using
a 60./1.35 oil immersion objective equipped with a Hamamatsu C9100-13 back-
61
thinned EM-CCD camera and Yokogawa CSU X1 spinning disk confocal scan head
(with upgrade from Spectral Aurora Borealis, Richmond Hill, Canada). The unit is
equipped with 4 separate diode-pumped solid state laser lines (Spectral Applied
Research, Richmond Hill, Canada, 405 nm, 491 nm, 561 nm, and 642 nm) with
emission filters: 447 nm ± 60, 525 nm ± 50, 593 nm ± 40, 620 nm ± 60, 676 ± 29 and
700 nm ± 75, and 1.5X magnification lens (Spectral Applied Research). Confocal
images were taken with an Improvision Piezo Focus Drive. Z-stacks were taken at
0.25 μm. Images were then deconvolved by iterative restoration using Volocity
Software (PerkinElmer, Waltham, USA) with confidence limit set to 95% and
iteration limit set to 20.
The following antibodies were used for IF: sheep anti-human VWF (1:1000 dilution,
AbD Serotec, Oxford, UK), rabbit polyclonal to C5b-9 (1:1000 dilution, Abcam,
1:1000 dilution) to CD46/MCP (1:50 dilution; Santa Cruz Biotechnology, Dallas, TX,
USA), rat polyclonal antibody to CD59 (1:1000 dilution; AbD Serotec, Oxford, UK),
and goat polyclonal antibody to CD55 (5 μg/mL; International Blood Group
Reference Laboratory, NHS Blood and Transplant, Bristol, UK).
3.3.5.2 Flow cytometry
Cells were seeded overnight in a 6 well plate (Falcon, Becton Dickinson, Franklin
Lakes, USA) to confluence, washed 1x with HBSS and then trypsinized (0.05%
Trypsin/0.53 mM EDTA, WISENT, St Bruno, Canada). The primary antibodies
(rabbit anti-CD46/MCP, 1:50 dilution, Santa Cruz Biotechnology, Dallas, USA, sc-
9098; goat anti-CD55/DAF, 1:100 dilution, R&D Systems, Minneapolis, USA,
AF2009; rat anti-CD59, 1:100 dilution, AbD Serotec, Oxford, UK, MCA715G) and
secondary antibodies (Alexa fluor 488, 1:200 dilution, Invitrogen) were incubated at
4°C for 20 minutes. At least 10,000 events were recorded using Attune Acoustic
Focusing Cytometer (Invitrogen) and FlowJo vX.0.7 software (FlowJo LLC, Ashland,
USA). Results are given as median fluorescence intensity (MFI). Cells were gated for
live cells (red laser 536nm, emission channel 2), single cells (FSC-A vs. FSC-H) and
finally through forward scatter and side scatter to determine the BOEC population.
Surface regulators were recorded via the blue laser 488 nm, emission channel 1. To
62
correct for spectral overlap during multicolor flow cytometry experiments, color
compensation was performed each time.
3.3.6 Complement Challenge of BOEC
In order to challenge the BOEC via the alternative pathway of complement, cells were
exposed to 50% normal human serum (NHS) with alternative pathway (AP) buffer (7
mM MgCl2, 10 mM EGTA, 144 mM NaCl, 20 mM HEPES buffer, pH 7.4). To
escalate / enhance the challenge to the cells, the membrane-anchored complement
regulators CD46/MCP, CD55/DAF and CD59 were functionally blocked, prior to the
addition of 50% NHS.
3.3.6.1 Normal human serum
50% normal human serum was used as source of complement. Serum was collected
from whole blood of adult donors into serum vacutainers (BD Biosciences), allowed
to clot for 30 minutes and then centrifuged at 3000 x g at 4°C. Serum was stored at -
20°C until needed as a source for complement. Heat-inactivated serum (HIS), for use
as a negative control, was obtained by incubating the serum for 30 minutes at 56°C.
3.3.6.2 Membrane-anchored complement regulator blockade
In order to simulate a CD46/MCP mutation, a non-complement-activating (Schiff et
al., 2004) monoclonal mouse anti-human anti-CD46/MCP antibody (GB24, IgG1,
kindly provided by John Atkinson, St Louis, MO, USA) was used. Additional
blockade of EC surface complement regulation was achieved with the use of a
monoclonal anti-human CD55/DAF (BRIC216, IgG1) and monoclonal anti-human
CD59 (BRIC229, IgG2b) (International Blood Group Reference Laboratory, NHS
Blood and Transplant, Bristol, UK) functionally blocking antibodies. Antibodies
were used at a concentration of 5 μg/mL and diluted in serum free cEGM-2 (Lonza)
media for 30 minutes in all experiments. The concentration of blocking antibody used
was derived from antibody titration and saturation experiments performed by FACS,
as detailed in chapter 2.
63
3.3.7 Assessment of Complement Challenge of BOEC
3.3.7.1 Complement deposition detected by immunofluorescence
For IF demonstration of complement deposition (C3 and C5b-9), BOEC were grown
on collagen-coated coverslips (VWR) and then incubated for 60-240 minutes (37°C,
5% CO2) with either cEGM-2 (BOEC) media, 50% normal human serum (NHS) in
AP buffer alone or after complement regulator blockade. After 60-240 minutes the
supernatant was aspirated and the cells were washed 3 times with ice cold PBS,
blocked with 1% (w/v) BSA for 1 hour and incubated with rabbit polyclonal antibody
to C3c, (Abcam, Cambridge, UK, ab15980, 1:1000 dilution) or a rabbit polyclonal
antibody to C5b-9 (Abcam, Cambridge, UK ab55811, 1:1000 dilution) overnight.
3.3.7.2 Complement deposition detected by FACS
C3b deposition was demonstrated by FACS using an antibody to C3c (rabbit
polyclonal, Abcam) that detects C3c as well as the C3c part of native C3 and C3b.
Cells were seeded overnight in a 6 well plate (Falcon) to confluence, washed 1x with
HBSS before blocking antibodies (GB24, BRIC216, BRIC229) were added for 30
minutes in serum-free media, followed by 50% NHS in AP buffer for 1 hour. Cells
were trypsinized (0.05% Trypsin/0.53 mM EDTA, WISENT) and washed in Flow
buffer (FB, 1% FBS/PBS) twice. The primary antibody (rabbit anti-C3c, 1:100
dilution, Abcam, ab15980,) in FB and the secondary antibody (R-Phycoerythrin-
conjugated AffiniPure F (ab‟) 2 Fragment Donkey Anti-Rabbit IgG (H + L), 1:200
dilution, Jackson ImmunoResearch, West Grove, USA) together with Fixable
Viability Dye eFluor780 (1:1000 dilution, eBioscience, San Diego, USA, 65-0865) in
PBS were incubated at 4° for 20 minutes. Cells were analyzed using Attune Acoustic
Focusing Cytometer and FlowJo 10.X.0 software. Results are given as median
fluorescence intensity (MFI). Cells were gated for live cells (red laser 536nm,
emission channel 2), single cells (FSC-A vs. FSC-H) and finally through forward
scatter and side scatter to determine the BOEC population. C3 deposition was
recorded via the blue laser 488nm, emission channel 2. To correct for spectral overlap
during multicolor flow cytometry experiments, color compensation was performed
each time.
64
3.3.8 Assessing the impact of complement challenge
3.3.8.1 LDH cell cytotoxicity assay
The LDH cell cytotoxicity assay was used to quantify cell death and compare the
impact of complement challenge on the different cell lines. BOEC from three healthy
controls and from two VWD type 3 patients were seeded overnight in a 96 well
ELISA plate (Sarstedt, Nümbrech, Germany) and grown to confluence. Prior to the
experiment cells were washed once with HBSS before, if indicated, adding the
complement-blocking antibodies diluted in serum-free media (cEGM-2) for 30
minutes. After washing twice, 10% NHS in serum-free media was added for 4 hours.
Pierce LDH cytotoxicity assay kit (Thermo Fisher Scientific, Waltham, USA) was
used according to manufacturer‟s instructions. Optical density (OD) was calculated
using a standard curve and then normalized to positive control and displayed in
percent increase over negative control (10%NHS). The positive control represents
maximal release, which was achieved by solubilizing cells with lysis buffer.
3.3.8.1 Platelet adhesion Assay
Whole blood was collected from healthy adult donors with acid citrate dextrose
(ACD: 22.9 mM citric acid, 44.9 mM sodium citrate dehydrate, 74 mM dextrose
monohydrate) anticoagulation. Platelet rich plasma (PRP) was attained by
centrifugation of the whole blood at 160 x g for ten minutes. The PRP was spun at
950 x g for seven minutes to pellet platelets which were then washed twice with
PBS/ACD solution (20% ACD in PBS; pH 6.1). Platelet count was measured using an
automatic hematocytometer (Beckman Coulter, Brea, USA) and concentration was
adjusted accordingly with PBS/ACD. Platelets were incubated with 2.5 μM calcein
(Life Technologies, Carlsbad, USA) for thirty minutes at 37°C. Finally, platelets were
pelleted at 950 x g and resuspended at a concentration of 15 x 107/ml in Tyrodes
buffer (136 mM NaCl, 2.7 mM KCl, 0.42 mM NaH2PO4, 19 mM NaHCO3, 5.5 mM
of glucose, 1 mM CaCl2, 1 mM MgCl2 and 10 mM of hyroxyethyl
piperazineethanesulfonic acid (HEPES: Invitrogen, Life Technologies, Carlsbad,
USA).
For BOEC-platelet adhesion experiments, control BOEC and VWD type 3 BOEC
(passage 3-14) were grown in collagen-coated microfluidic channels of the BioFlux
65
system (Fluxion Biosciences, South San Francisco, USA) as described before (in
chapter 2). Cells were washed at 1 dyne/cm2 with HBSS (Invitrogen). For
complement challenge experiments the membrane complement regulators were first
blocked with anti-human CD46/MCP, CD55/DAF and CD59 functional blocking
antibodies diluted in serum-free media (cEGM-2) and perfused through the channels
at 1-2 dynes/cm2 for thirty minutes. 50% NHS from healthy adult donors in
alternative pathway (AP) buffer was subsequently perfused through the BioFlux
chambers at a shear rate of 2 dynes/cm2 for 60 minutes. For platelet adhesion assays,
15 x 107/ml calcein labeled platelets in Tyrodes buffer (as described in platelet
preparation section) were flowed through the chamber at 2 dyne/cm2 for 10 minutes
after BOEC exposure to serum. Per time point and experiment three pictures of each
channel ([4x] magnification, NIKON camera) were saved and platelet adhesion was
manually counted using ImageJ software.
3.3.9 Ethics
Ethics approval was obtained from the Research Ethics Board at The Hospital for
Sick Children, Toronto, ON, Canada.
3.3.10 Statistical analysis
Figures were generated using GraphPad Prism 6 and displayed as mean and standard
deviation. Statistical analysis was performed using either two-way ANOVA with
post-hoc analysis or paired t-test. A p value < 0.05 was considered as statistically
significant. In figures p values are presented as follows: * ≤0.05, ** ≤0.01, ***≤0.001,
****≤0.0001, ns > 0.05.
66
3.4 Results
3.4.1 BOEC Possess Endothelial Cell Characteristics
Blood outgrowth endothelial cells are endothelial progenitor cells. BOECs were
isolated from the peripheral blood of healthy adult volunteers (control BOEC) and
from two patients with von Willebrand disease type 3 (VWD BOEC). The latter lack
functional von Willebrand factor (VWF antigen levels were undetectable) due to
compound heterozygous mutations (c. 876delC, c. 1255C>T; c. 3939G>A, c. 5842+1
G>C) in both patients. Firstly, an endothelial phenotype was confirmed by
immunofluorescence and flow cytometry. The presence of Weibel-Palade bodies
(WPB) containing VWF in the control BOEC confirmed an endothelial phenotype.
WPB are present only in endothelial cells and megakaryocytes. An endothelial cell
phenotype of both control and VWD BOEC was corroborated by demonstrating the
presence of key endothelial cell markers by flow cytometry (positive for CD144/ VE-
Cadherin and CD31/Platelet endothelial cell adhesion molecule (PECAM-1), as well
as ensuring that the cells were not of an immature progenitor cell or of a
hemangioblast origin (negative for CD45 and CD14). This has been previously
described in more detail in Chapter 2.
Next we confirmed by western blot and qRT-PCR the minimal expression of VWF in
BOECs isolated from the two patients with type 3 VWD. VWD BOEC show only
5.6% of the mRNA levels of control BOEC (Figure 3-1). Of note, VWF mRNA levels
in glomerular endothelial cells were also decreased compared to BOEC (35% of
mRNA level in control BOEC, p< 0.0007, paired t-test, Supplement 3-1).
67
Figure 3-1. BOEC possess endothelial cell characteristics
(A) Lysates of control BOEC (CTRL) and VWD BOEC (VWD) were probed for von
Willebrand factor (VWF, 1:1000) on an 8% SDS-page. VWF multimers were seen in CTRL
BOEC, whereas a significantly reduced amount of VWF was seen in VWD type 3 BOEC. (B)
This was confirmed by qPCR, where VWD BOEC showed just a minimal mRNA expression
of VWF (5.6% of control, p<0.0001, paired t-test). VWF mRNA levels were normalized to
GAPDH and control BOEC.
3.4.2 Type 3 VWD BOEC Express Similar Amounts of Membrane-
Anchored Complement Regulators as Control BOEC
In order to further characterize the VWD3 BOEC as relevant to study aHUS
pathogenesis we confirmed the presence of the membrane-anchored complement
regulators CD46/MCP, CD55/DAF and CD59 by immunofluorescent labeling and
visualization by spinning disk confocal microscopy (Figure 3-2 A-C). We next
demonstrated using flow cytometry that BOEC isolated from patients with type 3
VWD had quantitatively similar expression levels of the complement regulators
CD46/MCP, CD55/DAF and CD59 on their surface (Figure 3-2 D-G). VWD BOEC
had a 4.5-fold higher level of CD55/DAF and a 35-fold higher level of CD59 as
compared to CD46, exhibiting an equivalent cell surface distribution of the regulators
as compared to control BOEC (Figure 3-2 G).
68
Figure 3-2. Type 3 VWD BOEC express similar amounts of membrane-anchored
complement regulators as control BOEC
By immunofluorescence (A-C) and flow cytometry (D-G) the surface expression of
complement regulators CD46/MCP, CD55/DAF and CD59 was detected. (A-C) Cells were
seeded on cover-slips, stained for CD46/MCP (1:50; A), CD55/DAF (1:200; B) and CD59
(1:1000; C) and the representative secondary antibody (Alexa fluor 488, green) and imaged
using a Olympus IX81 inverted fluorescence microscope with a 60/1.35 oil immersion
objective and a Hamamatsu C9100-13 back-thinned EM-CCD camera with Yokogawa CSU
X1 spinning disk confocal scan head. Confocal images were taken with an Improvision Piezo
Focus Drive. Z-stacks were taken at 0.25 μm. Images were deconvolved by iterative
69
restoration using Volocity Software with confidence limit set to 95% and iteration limit set to
20. Cell nuclei were stained using 0.12 μg/ml Hoechst 33342 stain (blue).
(D-G) For flow cytometry cells were trypsinized off a 6-well plate, and incubated with
primary antibody (CD46, 1:50; CD55, 1:100; CD59; 1:100) and respective secondary Alexa
fluor 488 (1:200). Surface expression of complement regulators was acquired using Attune
Acoustic Focusing Cytometer (Invitrogen) and analyzed using FlowJo software. A similar
surface expression of complement regulators on VWD BOEC (blue) compared to control
BOEC (red) was observed, as shown in representative images D-F. The unstained controls are
displayed in dark grey for VWD BOEC and light grey for control BOEC. (G) Comparison of
the median fluorescence intensity (MFI) of three experiments did not show a significantly
different surface expression of CD46/MCP, CD55/DAF and CD59 (p=0.69, two-way
ANOVA).
3.4.3 Complement activation products associate with VWF
Release of VWF from endothelial cells is known to occur in response to complement
activation. This is dependent on the insertion of the terminal complement complex
C5b-9 in the cell membrane and seemingly independent of the anaphylatoxins C3a or
C5a. (Hattori et al., 1989) Recently the association of complement activation products
with endothelial cell derived VWF, has been shown in both static and fluidic
conditions.(Tati et al., 2013; Turner and Moake, 2013) As a first step we stained
concomitantly for both C5b-9 and VWF by immunofluorescence labeling of fixed
BOEC that had been exposed to 50% NHS in AP buffer. This was performed in static
conditions where BOEC, grown to confluence on coverslips, were incubated for one
hour with 50% NHS. By IF, the presence of positive C5b-9 staining on cells was
associated with VWF released from the cell. Intensification of complement challenge,
achieved by blocking three membrane-anchored regulators (CD46/MCP, CD55/DAF
and CD59) and exposing the BOEC to 50% NHS for either one or two hours was
associated with a reduction in the number of visible VWF-positive WPB. Not all cells
demonstrated C5b-9 deposition and in these cells the WPB were still clearly visible.
This was confirmed also under fluidic conditions where the BOEC were grown to
confluence in the BioFlux microfluidic chamber system prior to exposure to 50%
NHS in AP buffer. Representative pictures of experiments repeated on at least 4-6
separate occasions with 3-4 separate BOEC cell lines are shown in figure 3. Finally,
70
we demonstrated an apparent co-localization of C5b-9 with VWF multimers (Figure
3-3).
Figure 3-3. Complement challenge results in VWF release and the association of
complement activation products and VWF
(A-C) Control BOEC were exposed to 50% heat- inactivated serum (HIS) for 2 hours (A),
50% NHS after functional blockade of complement regulators CD46/MCP, CD55/DAF and
CD59 for 1h (B) and 2h (C). Incremental complement blockade results in VWF exocytosis
and subsequent C5b-9 deposition. (D-F) Exposing control BOEC to shear (2dyne/cm2) in a
71
microfluidic (BioFlux) flow chamber and 50% NHS/AP buffer also resulted in VWF release
and the formation of VWF strings (1:1000, green). (B) C5b-9 deposition was detected using a
rabbit polyclonal antibody (1:1000, red). (C) The merge indicates a co-localization of C5b-9
on VWF strings. Pictures were taken with a Nikon Eclipse Ti camera, 60x magnification/oil
immersion.
3.4.4 Platelet adhesion in response to complement challenge is initially
dependent on VWF release
In the complement-mediated aHUS form of TMA, dysregulation of the complement
alternative pathway (AP) is thought to ultimately lead to endothelial cell activation, a
procoagulant endothelium and platelet aggregation. Whether the platelets adhere to
VWF, injured cells or exposed subendothelial matrix or all of these and at what
timepoint this happens has not been clearly delineated. To investigate the functional
role of VWF in this process we sought to measure platelet adhesion to complement-
challenged BOEC grown in a microfluidic flow chamber while comparing normal
BOEC with those from type 3 VWD patients that lack VWF. In pursuance of this aim,
control and type 3 VWD BOEC were seeded overnight in the flow chamber and
washed, calcein-labeled platelets (15x107/ml) were perfused at 2 dyne/cm
2 for 10-20
minutes. Platelet adhesion on VWF strings (Figure 3-4 B-C) was achieved when
BOEC were treated with only 50% NHS and even more after functional blocking of
surface-bound regulators CD46/MCP, CD55/DAF and CD59 (20 platelets/hpf versus
84 platelets/hpf at 4X magnification). Platelets can be seen to adhere in control
BOEC only and not to VWD BOEC devoid of VWF (p<0.01). (Figure 3-4).
72
Figure 3-4. Platelet adhesion in response to complement challenge is initially dependent
on VWF release
Control and VWD BOEC were seeded overnight in microfluidic (BioFlux) 48-well chamber.
If needed cells were blocked with CD46 or CD46/55/59 blocking antibodies for 30 minutes at
1 dyne/cm2 prior to perfusion with 50% NHS in AP buffer for 1 hour. Washed, calcein-
labeled platelets (2.5μM) were introduced from inlet to outlet well at 2 dyne/cm2. Pictures
were taken with a Nikon Eclipse Ti camera at 4x (A-B) and 20x (C) magnification after 5
minutes. (A) Channels containing VWD BOEC (upper channel) and control BOEC (lower
channel) were exposed to 50% NHS/AP buffer for 1 hour and platelets (15 x107/ml,
100μl/well) in Tyrodes buffer were perfused at 2 dyne/cm2 for 5 minutes before pictures were
taken (4x magnification). Minimal platelet adhesion occurred on control BOEC. (B) With
increasing complement challenge, induced by incremental blockade of complement regulators
CD46/MCP, CD55/DAF and CD59, platelet adhesion only occurred when VWF strings were
secreted by control BOEC (4x magnification). (C) Image of control BOEC (treated as in B)
clearly shows platelet adhesion to VWF strings (20x magnification). (D) Platelet adhesion
was analyzed by counting adherent platelets of four random pictures/channel using ImageJ
software. Significantly more platelets can be seen to adhere in control BOEC as
compared to VWD BOEC devoid of VWF (**
p<0.01, two-tailed ANOVA, Sidak's
multiple comparisons test). N=3
73
3.4.5 VWD BOEC Show Increased C3b Deposition after Complement
Challenge
To investigate whether VWF has an amplifying role for the complement AP on the
endothelium we performed flow cytometry for C3b deposition on control and VWD
BOEC under increasing complement challenge as follows; (i) 50% NHS/AP buffer
alone, (ii) the functional blockade of one complement regulator (CD46/MCP)
followed by 50% NHS/AP buffer and (iii) the functional blockade of three surface
regulators (CD46/MCP, CD55/DAF and CD59). We observed an increase of the
median fluorescence intensity (MFI) in VWD BOEC treated with 50% NHS after
functional blockade of complement regulators compared to control BOECs (MFI
5861±3332 vs. 3450±1599, p<0.05, N=5, Figure 3-5).
Figure 3-5. VWD BOEC show increased C3b deposition after complement challenge
Control and VWD BOEC were seeded overnight in 6-well plates and treated with functional
blocking antibody and 50% NHS in AP buffer. Cells were washed, trypsinized and incubated
with primary antibody polyclonal rabbit anti-C3c (1:100), secondary donkey anti rabbit R-
Phycoerythrin (1:200) and fixable viability dye (eFluor 780, 1:1000) at 4° for 20 minutes.
Surface expression of C3b was acquired using Attune Acoustic Focusing Cytometer
(Invitrogen), after gating for live and single cells. Experiments were performed using two
different control BOEC and two different VWD BOEC. (A) shows one representative figure:
unstained control BOEC (light green), unstained VWD BOEC (dark green), 50% NHS
control BOEC (red), 50% NHS VWD BOEC (orange), CD46/55/59 block + 50% NHS
control BOEC (blue) and CD46/55/59 block + 50% NHS VWD BOEC (purple). (B) The
74
Median Fluorescence intensity (MFI) was calculated using FlowJo software after subtracting
MFI of unstained sample.
3.4.6 VWD BOEC Are More Vulnerable To Complement-Mediated
Cytotoxicity
We next wanted to confirm that this increase in complement deposition seen by flow
cytometry translated into an increase in complement-mediated cell injury and death.
Cell cytotoxicity was quantified by measurement of LDH released from the
complement-challenged BOEC into the supernatant. Cell death of VWD BOEC was
increased as compared to control BOEC with mild (50% NHS alone), moderate (50%
NHS after CD46/MCP blockade) or severe (50% NHS after CD46/CD55/CD59
blockade) complement challenge. The adjuvant block of three surface-bound
complement regulators achieved a cumulative effect. VWD BOEC demonstrated a
46±14% rate of cell death when treated with 10% NHS, a 49±16% when treated with
50% NHS after CD46 block and a 81±20% when incubated with 50% NHS after
block of CD46/MCP, CD55/DAF and CD59. This is compared to a rate of cell death
of 31±15%, 30±12% and 48±28%, respectively in control BOEC (p<0.01, p<0.05,
p<0.05, Figure 3-6).
75
Figure 3-6. VWD BOEC are more vulnerable to complement-mediated cytotoxicity
Cell death was measured by detection of LDH release in supernatant of control and VWD
BOEC after incubation with 10% NHS in serum-free media for 4 hours after pretreatment
with either none or CD46 or CD46/55/59 blocking antibodies. Cell death was calculated using
a standard curve and normalized to positive control (100%), obtained adding lysis buffer 45
minutes prior to incubation end. Background cell death from control BOEC was subtracted
for each value. Data was gathered from three different experiments (mean of 4-8 wells/plate)
using two different control BOEC and two different VWD BOEC. A more profound increase
of cell cytotoxicity was observed in VWD BOEC compared to control BOEC in all conditions.
* p<0.05 **p<0.01, ***p<0.001 (paired t-test).
76
3.5 Discussion
We aimed to study the interaction between complement and VWF in TMA
pathogenesis. Using BOEC isolated from type 3 von Willebrand disease patients we
were able to take advantage of a naturally occurring endothelial cell line with
negligent amounts of VWF, essentially equating to a VWF knockout, to study the role
played by VWF in the pathogenesis of a TMA initiated by complement dysregulation
and the association of VWF and the alternative pathway of complement. It is known
that complement activation leads to EC activation and VWF release. (Hattori et al.,
1989; Nakashima et al., 2002; Ota et al., 2005) The first question we addressed was
how important this exocytosed VWF is for platelet adhesion in the setting of an acute
complement stress to the EC. In a microfluidic flow chamber (BioFlux) we were able
to see that platelet adhesion after complement-induced EC activation was primarily
VWF dependent, as no platelet adhesion was observed in VWD BOEC.
Multimerized VWF dimers are compactly stored in the WPB of endothelial cells.
Exocytosed VWF multimers can remain attached to EC and play a pivotal role in the
initiation of hemostasis by providing a platform on which platelets can
adhere.(Valentijn et al., 2011) When released from EC these multimers can form
(bundles of) strings which unfold in the circulation under the influence of shear flow
thus unveiling the glycoprotein (GP) 1b platelet-binding site.(Nightingale and Cutler,
2013) Normally these strings are subject to proteolysis by ADAMTS13.
Alternatively, released VWF can bind to collagen. (Ruggeri, 2007) Secreted VWF
differs from that in the circulation, which is present in a globular state, thus
maintaining both the platelet GP1b binding site and the ADAMTS13 cleavage site
cryptic/hidden. (Nightingale and Cutler, 2013)
To further investigate the interaction of VWF and the AP of complement, we utilized
BOEC from type 3 von Willebrand disease patients. It was first reported that in static
conditions, complement AP products, including C3 and C5, were present on
endothelial cell secreted VWF strings. (Turner and Moake, 2013) Tati et al showed
that, under shear conditions and in the absence of ADAMTS13, C3 bound to
histamine-induced VWF strings, to VWF adherent platelets, and to the endothelial
cell that secreted the VWF. (Tati et al., 2013) However, the functional downstream
77
effect of this binding remained unclear. (Tati et al., 2013) Expanding on these studies,
we found that when BOEC were subjected to an incrementally increased complement
challenge in a microfluidic chamber that C5b-9 deposition was associated with the
VWF multimers as well as being deposited on the endothelial cell.
This raised the question as to whether the association of the AP and VWF was
actually complement amplifying resulting in enhanced complement deposition on EC
– a theory that has also been recently proposed by others. (Turner et al., 2014)
Considering this hypothesis, we expected to find less complement activation on cells
devoid of VWF. However, after exposing VWD BOEC to a complement challenge
(50% NHS with and without regulator blockade) we demonstrated increased C3b
deposition on their surface as compared with control BOEC. As VWD BOEC
expressed similar amounts of complement regulators CD46/MCP, CD55/DAF and
CD59 on their surface we ruled out other reasons for this result. This was
biologically relevant, as VWF deficient BOEC exhibited decreased survival to
complement-mediated cytotoxicity. Taken together, this is highly suggestive of a
protective role for VWF release or it limits complement deposition on EC surface.
It also raises the question as to whether cells perhaps expressing or containing less
VWF are more vulnerable to complement mediated injury. The heterogeneity of the
vascular endothelium is diverse and known to include VWF expression. (Aird, 2007a,
b) In lung endothelium, for example, VWF expression is strongest in veins and very
weak in pulmonary capillaries. (Kawanami et al., 2000; Muller et al., 2002) The
fenestrated glomerular endothelium had patchy positivity for VWF as detected by
immunohistochemistry on biopsy and post mortem specimens in one study.
(Pusztaszeri et al., 2006) This could partially explain why the glomerular endothelium
is vulnerable to complement attack in aHUS and perhaps why in TTP the kidney is
classically spared. We confirmed that cultured GEC have less VWF by both PCR and
Western blot and on immunofluorescence it was apparent that most GEC completely
lack VWF. (Supplementary Figure 3-6)
Our findings suggest two phases in the pathogenesis of TMA and aHUS. Initially,
complement activation leads to EC activation and VWF release, which is a protective
response, to facilitate platelet adhesion and delivery to the site of microvascular EC
78
injury and to sequester complement activation products. In later stages, with
prolonged complement challenge, EC injury may lead to cell death with tissue factor
release and platelet adhesion to the exposed basement membrane components or
apoptotic/dying cells. Loss of VWF from the EC into the circulation in response to
ongoing complement-mediated WPB exocytosis may leave the cells more vulnerable
to complement injury.
In conclusion, by comparing Type 3 VWD BOEC and control BOEC and their
response to complement challenge we have been able to demonstrate that VWF
multimers released by the vascular EC contributes to EC protection by acting as
negative complement regulator. Similarly, lack of VWF results in increased
complement deposition and cell cytotoxicity. Different from recent claims in the
literature, we here demonstrate for the first time a new principle in EC complement
regulation via VWF. Our results overcome the assumed dichotomy of complement
and VWF (or aHUS and TTP) as separately acting biological pathways and provide
evidence for an intimate functional link between the two systems. This insight not
only contributes to a better understanding of TMA pathogenesis but also points
towards more efficient strategies for the monitoring and treatment of TMA patients.
79
3.6 Supplemental Data -VWF expression in glomerular
endothelial cells
Figure 3-S1. Glomerular endothelial cells express less VWF
(A, B) BOEC and glomerular endothelial cell (GEC) lysates were resolved by 8% SDS-
PAGE and probed for VWF (1:1000). A lower expression of VWF was seen in GEC on
western blot, (B) which was quantified using densitometry, and revealed a VWF expression
of 60% in GEC. N=1. (C) By qPCR the mRNA level of von Willebrand Factor (VWF) in
control BOEC and glomerular endothelial cells (GEC) was measured. GEC show 35% of
VWF mRNA levels than BOEC (*** p<0.001, two-tailed t test, N=3) (D)
Immunofluorescence labeling of GEC with antibodies detecting VWF (1:1000, red) and
CD59 (1:1000, green) revealed partial expression of VWF in glomerular endothelial cells.
80
Image is shown in extended focus and was taken using a Olympus IX81 inverted fluorescence
microscope with a 60/1.35 oil immersion objective and a Hamamatsu C9100-13 back-thinned
EM-CCD camera with Yokogawa CSU X1 spinning disk confocal scan head. Confocal
images were taken with an Improvision Piezo Focus Drive.
81
CHAPTER 4 UNIFYING DISCUSSION, FUTURE
DIRECTIONS AND CONCLUSIONS
82
4.1 Unifying Discussion
TMA is a severe multisystem disease involving endothelial cell damage, thrombosis
of the microvasculature and organ dysfunction. Although significant advances have
occurred in understanding the pathophysiology underlying the expanding spectrum of
diseases associated with TMA, coupled with improvements in its diagnosis and
therapy, there are still significant gaps in knowledge, areas of uncertainty and patients
dying from this disease. (George et al., 2012; George and Nester, 2014) Advancing
the understanding of TMA pathology will inevitably lead to new and improved
treatment strategies and translate into better outcomes for TMA patients. In order to
achieve this goal, we developed a model system to study the pathogenesis of TMA
using an endothelial progenitor cell (BOEC). BOEC were examined under static and
microfluidic conditions and exposing them to a complement challenge, achieved by
incrementally impairing complement control by functionally blocking membrane
regulators. A set of reliable, reproducible and disease-relevant readouts was
established, including demonstration of complement deposition by FACS,
quantification of cell death, and platelet aggregation under microfluidic conditions.
Designed to more closely resemble the patients‟ genotype and phenotype, in the
future, we will utilize patient derived BOEC, serum and other constituents such as
platelets to thus achieve an ex vivo test system that resembles an individual patient‟s
pathology in an unprecedented way.
This system was then used to study CD46/MCP associated aHUS. Loss of just 50%
CD46/MCP function can be associated clinically with recurrent aHUS, while a person
with complete knockout of the gene may or may not manifest disease. (Couzi et al.,
2008) Studies performed in vitro have found these mutations to be disease relevant.
These studies have been performed in transfected CHO cells where these mutated
CD46/MCP forms are associated with failure to inactivate cell surface C3b deposited
on cells after complement challenge. Although the CHO system is a „clean‟ system,
allowing delineation of the specific effect of a given mutation, it does not exemplify
the in vivo situation, as these cells are naturally devoid of other regulators. In our
model system, using BOEC, a cell endowed with the other complement regulators,
completely blocking the function of CD46/MCP failed to lead to a measurable or
significant increase in complement deposition, endothelial cell death or platelet
83
adhesion. Instead, it was necessary to augment the „complement challenge‟ by
blocking three of the membrane-anchored regulators to achieve the phenotype.
This finding firstly provides support for the hypothesis that additional (genetic)
aberrancies in the complement system could increase the penetrance of these
CD46/MCP mutations and even modulate severity of disease. (Bresin et al., 2013)
Additional genetic „hits‟ in complement genes and regulators have been described in
patients with CD46/MCP mutations. (Bienaime et al., 2010; Bresin et al., 2013;
Fremeaux-Bacchi et al., 2013; Geerdink et al., 2012; Provaznikova et al., 2012;
Stevenson et al., 2014) Secondly it highlights the importance of functional studies in
helping to understand how a given mutation may lead to disease, and the utility of
different model systems. This has been recently emphasized and is especially relevant
considering the extent to which predictive software is now used to suggest the
likelihood of disease relevance or pathogenicity of a given mutation. (Marinozzi et al.,
2014)
The two archetypal representations of TMA, namely TTP and HUS, have many
shared elements in terms of clinical symptoms and pathology. More recently
complement has emerged as a common bond between the two. (Noris et al., 2012; Wu
et al., 2013) Expanding on this theme, and considering the recent data linking VWF
and the AP of complement (Turner and Moake, 2013) as well as the postulated
complement activating or amplifying role for that interaction (Turner et al., 2014) we
set about using BOEC naturally deficient of VWF to confirm this. Different from this
prediction, however, complement activation was actually greater on these cells rather
than less and the cells were more vulnerable to complement mediated cytotoxicity.
This suggests a regulatory role for VWF multimers released from EC upon
complement-stimulation.
Considering that glomerular endothelial cells have been shown to have very minimal
VWF, (Pusztaszeri et al., 2006) this finding may explain why glomerular endothelial
cells are more prone to complement-mediated injury, thus explaining the
susceptibility of the kidney to complement-mediated TMA. Furthermore, using the
BOEC lacking VWF and comparing them with normal BOEC, we were able to show
that, after endothelial cell activation by complement, platelet adhesion is at least
initially to VWF. It is known that platelets firmly adhere to endothelial cell-anchored
84
VWF that has been unfurled under shear flow. This ensures platelets are brought in
close proximity to the injured endothelium, in order to initiate thrombosis and repair.
(De Ceunynck et al., 2013) Platelets also contain the complement regulator CFH and
it might be expected to aid in local complement control. (Licht et al., 2009)
There has always been considerable debate as to whether TTP and HUS are the same,
contiguous, or at two ends of a continuum or spectrum. Ultimately, with the discovery
of their respective causes, a dichotomy emerged, with one being considered as
complement-mediated and the other, VWF driven. (Remuzzi, 1987, 2003; Tsai, 2003)
Recently, however, those lines of separation were blurred again (Noris et al., 2012)
with the demonstration of complement activation in TTP patient‟s blood (Reti et al.,
2012; Ruiz-Torres et al., 2005; Wu et al., 2013) and the experimental association of
VWF with the alternative complement pathway. (Turner et al., 2014; Turner and
Moake, 2013)
Taken together this leads us to a new understanding of aHUS pathogenesis, or a two-
phase view of the sequence of events that might lead to TMA. Complement challenge
leads to endothelial cell activation with VWF release. This VWF facilitates platelet
adhesion and the binding of complement activation products. This brings platelets
into contact with the injured endothelium. In the presence of efficient and normal
levels of ADAMTS13 these VWF multimers, laden with complement products, are
cleaved and released into the circulation thus helping to maintain endothelial cell
homeostasis (Phase 1). If there is ongoing or persistent complement challenge,
especially in association with deficient or overpowered complement regulators, or less
VWF clearance (less blood flow or ADAMTS 13) then this initially protective effect
of VWF can be overcome, leading to progression towards a TMA phenotype (phase
2). (Figure 4-1) This might help explain the recent finding of ADAMTS13 mutations
in aHUS patients associated with decreased ADAMTS13 activity. (Feng et al., 2013a)
85
Figure 4-1. VWF and ADAMTS13 as a Complement Regulatory System.
VWF released from EC upon complement activation is unfolded under shear flow allowing
the firm adhesion of platelets. VWF can also bind to complement activation products. In the
presence of normal ADAMTS13, then these multimers can be cleaved. VWF likely has a
complement regulatory role following its release from complement-activated EC.
86
4.2 Future Directions
4.2.1 Introduction
After establishing BOEC as a model to study TMA pathogenesis we found that loss of
CD46/MCP function is not enough to fully explain the occurrence of TMA and that
additional complement aberrancies can significantly increase the likelihood of
progressing to a TMA phenotype. While seeking to better understand how a person
with a mutation in CD46/MCP may manifest a TMA, we next looked at VWF as a
potential amplifier of complement. This hypothesis arose from our own observation
of the association of immunofluorescently labeled VWF and C5b-9 under fluidic
conditions that coincided with a publication that demonstrated the association of
activated AP components on VWF. (Turner and Moake, 2013) Using a BOEC
naturally lacking VWF we actually found the contrary, that VWF more likely
downregulates complement. This exciting finding contradicts current published
opinion that VWF amplifies the complement injury (Turner et al., 2014) and opens up
a completely novel way of looking at the two diseases TTP and aHUS, perhaps
reopening the discussion as to how much they are linked.
Moving on from this, we aim to take a closer look at endothelial cell protection and
explaining how the cell protects itself from loss of CD46/MCP function as well as to
focus on more mechanistic details of complement-mediated EC injury. Specifically,
the model system can be employed to examine whether the cells mount a dynamic
response to complement-induced stress by increasing the expression of their surface
bound complement regulators. We have studied CD46/MCP as one of the two aHUS-
linked membrane-anchored complement regulators but we intentionally did not study
the contribution of the key soluble regulator CFH. To study its role is one of the key
next steps to complete the picture.
While addressing the example of CD46/MCP, we have utilized BOEC and thus
established a model system resembling patient characteristics in an unprecedented
way. Now we are in a position to test our findings in BOEC, not just from
CD46/MCP mutant patients, but to take it beyond that and extend our studies to study
BOEC from various subtypes of aHUS (and other TMA diseases) and to combine
them with patients own serum, plasma and platelets.
87
4.2.2 Do Endothelial Cells Mount a Cytoprotective Response when
Challenged by Complement?
4.2.2.1 Rationale and Hypothesis
There appears to be a variation in the cell surface expression levels of CD46/MCP
and the other membrane-anchored complement regulators in various kidney diseases,
although the literature is inconsistent. In an in vitro study of cultured renal mesangial
cells exposed to either the anaphylatoxins C3a, C5a or the membrane attack complex
(MAC), a slight increase in surface expression of CD46/MCP was demonstrated by
flow cytometry although no change in mRNA could be demonstrated. In contrast, an
increase in CD55/DAF mRNA was observed. (Cosio et al., 1994) Another study of
human kidney tissue specimens found enhanced staining of CD46/MCP in association
with complement C3 split-product deposition in glomerulonephritic kidneys as
detected by immunohistochemistry. (Endoh et al., 1993)
Moreover, there is evidence to suggest that surface expression levels of complement
regulators can vary in vivo on different cell types and that this may play a role in the
etiology or susceptibility to disease including; (i) T-cell mediated rejection post
kidney transplant, (Kakuta et al., 2012) (ii) respiratory disorders,(Grumelli et al.,
2011; Lee et al., 2012) (iii) age-related macular degeneration,(Singh et al., 2012) (iv)
osteoarthritis, (Schulze-Tanzil et al., 2012; Scott et al., 2011) (v) rheumatoid
arthritis,(Pahwa et al., 2012) (vi) pregnancy related thrombophilia, (Wirstlein et al.,
2012) and (vii) systemic lupus erythematosus.(Alegretti et al., 2012; Das et al., 2013;
Ellinghaus et al., 2012) Expression levels of the regulators CD46/MCP and CD59
have been detected on neutrophils of patients after severe trauma. (Amara et al., 2010)
Furthermore in a model of STEC HUS, Shiga toxin 2 was shown to downregulate
expression of CD59 on the surface of glomerular endothelial and tubular epithelial
cells in vitro. (Ehrlenbach, Infect Immun 2013)
This suggests a hypothesis that when an EC is exposed to complement challenge, the
cellular response is to increase protector expression. If this dynamic response is
impaired by an endogenous defect or exogenous factor like toxins, then the cells are
more susceptible to injury.
88
4.2.2.2 Methods
BOEC were seeded overnight in a 6 well plate (Falcon) to confluence, washed 1x
with HBSS before RNA was isolated using TRI Reagent (Sigma-Aldrich, St. Louis,
USA, T9424) according to manufacturer‟s instructions. RNA concentration and
integrity was verified by spectrophotometer (NanoDrop 1000, Thermo Fisher
Scientific, Waltham, USA), and reverse transcribed using ReadyScript™ cDNA
Synthesis Mix (Sigma-Aldrich, St. Louis, USA, RDRT). Samples (200 ng cDNA in
diethyl pyrocarbonate (DEPC, Sigma-Aldrich) treated water) were amplified by real
time polymerase chain reaction (PCR) using StepOne™ System from Life
Technologies (Carlsbad, USA). Amplified products were detected using KiCqStart™
SYBR® Green qPCR ReadyMix™, with ROX™ (Sigma-Aldrich, Carlsbad, USA,
KCQS02) and analyzed as follows: 2–(C
T – C
T GAPDH) – C
T control)
. GAPDH was used as
housekeeping gene.
The following oligonucleotide primers (Sigma-Aldrich, Carlsbad, USA) were used:
GAPDH: forward, 5‟-ACAGTTGCCATGTAGACC-3‟; reverse, 5‟-
TTTTTGGTTGAGCACAGG-3‟.
CD46/MCP: forward, 5‟-AGTGGTCAAATGTCGATTTC-3‟; reverse, 5‟-
ATCCCAAGTACTGTTACTGTC-3‟.
CD55/DAF: forward, 5‟-CAGAGGAAAATCTCTAACTTCC-3‟; reverse, 5‟-
AGTTGGTGAGACTTCTGTAG-3‟.
CD59: forward, 5‟-CATTACCAAAGCTGGGTTAC-3‟; reverse, 5‟-
TTTCTCTGATAAGGATGTCCC-3‟.
Using the following experimental conditions (each n = 3-4) where BOEC were
exposed to; (i) media (baseline), (ii) 50% NHS after CD46/MCP blockade and (iii)
50% NHS after CD46/MCP, CD55/DAF and CD59 blockade, for one, two and four
hours.
4.2.2.3 Preliminary Results
With CD46/MCP blockade alone, there was an initial increase in CD46/MCP
transcription, however, this did not persist, and no significant increase in either
89
CD55/DAF, or CD59 was noted. (Figure 4-2 A) In contrast, by augmenting the
complement challenge with functional blockade of CD46/MCP, CD55/DAF and
CD59, BOEC increased their transcription of CD46/MCP and CD59. (Figure 4-2 B)
This was most prominent in the first 1-2 hours and the effect was lost at 4 hours.
90
Figure 4-2. Response of complement regulators to increasing complement challenge
(A) BOEC were blocked with functional antibody against CD46/MCP for 30 minutes,
followed by incubation with 50% NHS in AP buffer for 1h, 2h and 4 hours. RNA was
isolated using TRI reagent and reverse transcribed to cDNA. CT values were normalized to
GAPDH expression and baseline (cells incubated with media). No significant difference in
gene expression of CD46, CD55 and CD59 up to 4 hours of treatment was observed, except
for an 18-fold increase of CD46 gene expression after one hour. N=3-4. (B) Gene expression
levels of CD46 and CD59 changed significantly within the first hour of complement
challenge when cells were blocked for CD46, CD55 and CD59. N=3-4. A two-way ANOVA
with Tukey‟s multiple comparison test was used for statistical analysis. * ≤0.05, ** ≤0.01,
***≤0.001, ****≤0.0001, ns > 0.05.
4.2.2.4 Discussion
In this preliminary study we see that with the increased complement challenge of
exposing BOEC to normal human serum after blocking three surface regulators as
compared to a single regulator, the mRNA expression of CD46/MCP and CD59
increase in the first one to two hours as evidence for dynamic response to protect the
EC. One study has looked at the expression levels of the other regulators CD55/DAF
and CD59 in eleven patients with CD46/MCP-associated aHUS and found no
difference in the expression levels in the patients as compared to healthy control
participants. The CD55/DAF and CD59 levels were measured on granulocytes,
however, and not endothelial cells. (Fremeaux-Bacchi et al., 2006)
Defining the response of normal EC to increasing complement challenge will be
important for when we study the BOEC of patients with disease, as an inability to
upregulate expression of the complement regulators might confer an increased risk of
manifesting disease.
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4.2.3 How Does the Fluid Phase Regulator CFH Contribute to Endothelial
Cell Protection?
4.2.3.1 Rationale and Hypothesis
All human cells are protected against complement-mediated injury of self by a
combination of fluid phase regulators, including complement Factor H (CFH) and
membrane-anchored regulators including CD46/Membrane Cofactor Protein (MCP),
CD55/Decay Accelerating Factor (DAF) and CD59. (Zipfel and Skerka, 2009) CFH
accelerates the decay of the AP C3 convertase and acts as a cofactor for the serine
protease CFI that cleaves C3b into inactive form iC3b. (Perkins et al., 2014) CFH is
considered the most important complement regulator. (Zipfel and Skerka, 2009) We
have utilized our model so far to demonstrate the effect of the membrane-anchored
complement regulators in cell protection. The effect of the other soluble regulators,
particularly CFH, will be important to analyze. Mutations in both CD46/MCP and
CFH have been described in individual patients presenting with aHUS. (Bresin et al.,
2013; Fremeaux-Bacchi et al., 2013)
The fact that CFH-related aHUS carries a worse prognosis than CD46/MCP with 75%
renal loss in the former as compared to 75% renal survival in the latter suggests that it
is the most important regulator for EC protection. (Fremeaux-Bacchi et al., 2013;
Noris et al., 2003; Noris et al., 2010; Sellier-Leclerc et al., 2007) Furthermore, an
additional CFH mutation in patients already carrying an MCP mutation increased the
likelihood of developing the disease and worsened the decline in renal function. We
postulate that there is a hierarchy of complement regulators and that loss of CFH on
top of CD46/MCP significantly increases the TMA phenotype.
4.2.3.2 Methods
CFH blocking
In order to study the effect of CFH in protecting the endothelium from cell death,
confluent BOEC were incubated with neuraminidase at a concentration of 30 mU/mL
in serum-free media for thirty minutes prior to 50% NHS. This disrupts the
glycocalyx and removes glycosaminoglycans (GAGs) by desialylation, as GAGs are
the physiological endothelial cells‟ binding site for CFH.(Junnikkala et al., 2000)
92
Trypan blue exclusion
BOEC (between passages 3-14) were grown in 0.05 mg/ml rat-tail collagen type I
coated 96 well tissue culture plates (Sarstedt, Nümbrech, Germany). Cells were
incubated for thirty minutes at 37°C with (i) media, (ii) 50% normal human serum
(NHS) in AP buffer and (iii) 50% normal human serum (NHS) in AP buffer after pre-
incubation with functional blocking antibodies of CD46/MCP, CD55/DAF and CD59
in serum free EGM-2 media with supplements (Lonza). This treatment lasted thirty
minutes at 37°C. Then cells were washed twice with phosphate buffered saline (PBS:
Wisent, St Bruno, Canada). 1:1 mixture of Trypan Blue (Sigma-Aldrich, St. Louis,
USA) in PBS was added to cells for five minutes. 4% paraformaldehyde in PBS was
utilized to fix cells for ten minutes. Two fields of cells at 10x magnification from each
of duplicate wells were counted by using a Leitz DM IL microscope (Leica
Microsystems). The percent cell death was calculated by the following formula: dead
cells/total cells * 100.
Apoptosis
Cells were seeded in 96-well ELISA plates (Sarstedt) and incubated with functional
blocking antibodies for 30 minutes followed by 50% NHS in serum-free media
(cEGM-2) for four hours. Cells were fixed and blocked as described in detail earlier
and stained for cleaved caspase-3 (Cell Signaling, Danvers, USA, 1:400 dilution) and
respective secondary antibody conjugated with Alexa fluor 555 (1:1000 dilution).
4.2.3.3 Preliminary Results
We first examined whether neuraminidase (NA) pretreatment of cells followed by
exposure to normal human serum was associated with an increase in cell death over
serum alone. There was a three-fold higher increase in cell death with NA
pretreatment and 30 minutes of 50% NHS. (Figure 4-3 A) This cell death was shown
to be via apoptosis as assessed by fluorescently labeling fixed cells with an anti-
human cleaved caspase-3 antibody. There was an increase in the number of apoptotic
cells after 4 hours in those that had NA and an anti CD46 functional blockade prior to
serum.
93
Figure 4-3. Role of CFH-mediated surface protection against endothelial cell death
(A) Neuraminidase (30mU) was used to mimic functional defect of CFH cell surface
protection. Cells grown to confluence on a 96-well plate were incubated for 30 minutes with
neuraminidase, followed by 30 minutes of 50% NHS/AP buffer. Trypan-blue was added and
positive cells were counted and displayed as percent of total cells. A significant (p=0.03, one-
way ANOVA) increase of dead cells was observed when compared to serum treatment alone.
N=5 (B-C) Apoptosis was determined using a cleaved-caspase 3 (1:400) antibody. Cells were
seeded in a 96-well plate, Neuraminidase (30mU) and a functional anti-CD46 antibody was
added for 30 minutes, followed by 4 hours of 50% NHS. Cells were fixed with 4% PFA and
stained for cleaved caspase 3 (red) and DAPI (blue). Cells were imaged using a Nikon Eclipse
Ti camera at 20x magnification (B) and 40x magnification (C).
94
4.2.3.3 Discussion
These preliminary results confirm the importance of CFH as a regulator, and the
cumulative effect of a combined defect in CFH and CD46/MCP. However, because
thrombomodulin (CD141), also known to increase susceptibility to aHUS (Delvaeye
et al., 2009) is also a constituent of the endothelial cell glycocalyx, where it is bound
to chondroitin sulphate (Boels et al., 2013), neuraminidase might also be expected to
affect its function using this technique.(Boels et al., 2013) As such, this technique
may not be specific enough to clearly discriminate the precise CFH contribution.
Therefore, alternative approaches to more specifically assess the function of CFH
might include the use of CFH depleted serum or blockade of CFH function by either
(i) a CFH 19-20 construct, which binds and competitively inhibits the short consensus
repeats (SCR) 19-20 on the C-terminus of CFH, responsible for C3b binding
(Jokiranta et al., 2000; Jozsi et al., 2007) or (ii) a CFH-function blocking antibody.
95
4.2.4 Will BOEC Isolated From a Patient Further our Understanding of
TMA?
4.2.4.1 Rationale, Hypothesis and Aims
Having established and validated the model system employing BOEC to study aHUS
and TMA, we are now poised to study BOEC isolated from patients with various
mutations and TMA diseases. The first patient that we will undertake this on carries
two heterozygous mutations in CD46/MCP and a heterozygous mutation in
ADAMTS13. (Figure 4-4) The affected proband manifested recurrent aHUS,
responsive to plasma therapy and eculizumab, when he inherited a heterozygous
mutation in CD46/MCP from his father and in CD46/MCP and ADAMTS13 from his
mother, in other words defects in both the complement and coagulation systems.
Family members carrying one or two mutations did not develop the disease so far.
Functional studies, carried out elsewhere, did not show a functional relevance for
either the CD46/MCP or the ADAMTS13 mutations alone and were not able to
explain disease manifestation in this patient. As the combination of inherited
alterations seem to have caused the disease in this individual, studying BOEC isolated
from the patient and relatives, might explain why the patient developed the disease
and his parents/siblings did not.
96
Figure 4-4. Multiple genetic hits in complement and coagulation genes needed to cause
aHUS
The index patient (arrow) inherited two heterozygous mutations in MCP (blue, purple) and
one heterozygous mutation in ADAMTS13 (green). Other family members carry one or two
mutations, but never developed the disease.
4.2.4.3 Proposed Methods and Anticipated Results
BOEC that have been isolated from the patient will first be characterized as
endothelial, then as BOEC (positive by FACS for the endothelial markers CD144 and
CD31 and negative for the hemangioblast markers CD45 and CD14).
Immunofluorescence microscopic, FACS and qPCR examination of the expression of
the complement regulators as compared to normal control BOEC will then be
performed. The amount of complement deposition, cell survival and platelet adhesion
can then be compared to control healthy BOEC.
4.2.4.4 Anticipated Results and Implications
We anticipate that when these cells are exposed to normal human serum that there
may be increased complement deposition. Cell death and platelet adhesion assays will
possibly show an increased propensity to exhibit a TMA phenotype. Ultimately
isolating BOEC from patients will lead to a better understanding of the mechanisms
97
involved in terms of disease susceptibility, manifestation or precipitation. It might
actually be able to explain why the patient developed the disease. Furthermore, in the
future isolating BOEC from patients might contribute to the pathogenetic workup of
aHUS patients. Finally, considering the regenerative capacity of BOEC and their use
as a means of cell therapy, they could become a treatment strategy.
98
4.3 Conclusions
In 1924, Moschcowitz described a girl with microangiopathic hemolytic anemia,
thrombocytopenia, fever, neurological symptoms and hematuria. On post mortem
there were widespread, platelet-rich microthrombi in the microvasculature. This
became known as TTP. In 1955, Gasser, a Swiss pediatric nephrologist, described
patients with a similar presentation but in whom renal failure was prominent and he
termed this disease HUS. It also had a microangiopathic hemolytic anemia,
thrombocytopenia and microvascular thrombosis. Where the two diseases appeared to
differ, was whether the brain or the kidney was more prominently affected. (Gasser et
al., 1955) Over the next half century, as their respective etiologies became more clear,
they have been thought of, at various timepoints, and by various authors, as distinct,
separate and overlapping diseases. (Fujimura, 2003; Remuzzi, 1987, 2003; Tsai,
2003) In 1952, when Symmers coined the term thrombotic microangiopathy, it was
universally fatal. (Symmers, 1952) Although the mortality has significantly improved,
there remain many unmet needs in terms of understanding, specific therapies and
improving renal outcomes for individuals affected by this severe, multisystem disease.
In some ways, distinguishing TTP and aHUS is now more important than ever, as
there is a specific, complement-directed therapy, eculizumab that has revolutionized
the treatment for atypical HUS. (Legendre et al., 2013; Nurnberger et al., 2009)
As we continue to gain insight into aHUS, although a relatively rare disease, the
improved understanding will have significantly more broad-reaching implications for
a much larger group of patients, recognizing the increasing spectrum of diseases with
a TMA phenotype. (Riedl et al., 2014a) In the work presented in this thesis, we have
developed a system to study EC response to complement-mediated injury that can be
expanded upon and used to study the real EC phenotype of TMA patients as well as
model any of the diseases resulting in a TMA. We have confirmed that additional
genetic „hits‟ in complement significantly increase the likelihood of TMA over a
CD46/MCP mutation alone, and in so doing, have shown the importance of
biologically relevant model systems to study disease pathomechanisms. Most exciting
has been the discovery that VWF, released from activated ECs, is not complement
amplifying as proposed in the literature, but rather VWF and ADAMTS13 might well
be considered as down regulators of complement, where VWF laden with
complement can be cleaved by ADAMTS13 and released into the circulation for later
99
clearance. In identifying VWF as complement regulator, this paves the way to a novel
understanding of aHUS pathogenesis. Firstly, complement challenged ECs release
VWF as a cell protective response - in order to bind platelets and help establish
control of complement activation. If this system is overcome, or if there is excessive
stimulation of the EC with loss of VWF, then the endothelium becomes more
vulnerable to complement injury. Thereafter platelets would be expected to adhere to
dead cells and exposed basement membrane.
Considering the heterogeneity of endothelial cell VWF expression and the relative
paucity of VWF in the glomerular endothelium, (Pusztaszeri et al., 2006) as opposed
to the brain microvascular endothelium, (Bernas et al., 2010; Dorovini-Zis et al.,
1991) this might explain why loss of complement control in aHUS particularly affects
the kidney microvasculature.
TMA pathogenesis involves the close interplay of a number of biological systems,
especially complement, coagulation and inflammation. At the center of this
interaction is the microvascular endothelium, and BOEC present a unique opportunity
to learn more about these rare and devastating diseases to ultimately develop safer,
more specific and efficient therapies. Finally, we believe that used as we have done,
BOEC will represent a unique toolset to perform functional studies of genes newly
discovered as linked to TMA.
100
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