The Requirement for Oxygen in the Maturation and

Secretion of Soluble urokinase Plasminogen Activator Receptor (uPAR)

by

Ryan A. Rumantir

Supervised by Dr. Marianne Koritzinsky

A thesis submitted in conformity with the requirements for the degree of Master of Science

Graduate Department of the Institute of Medical Science University of Toronto

© Copyright by Ryan A Rumantir (2013)

 

  ii  

The Requirement for Oxygen in the Maturation and Secretion of Soluble urokinase Plasminogen Activator Receptor (uPAR)

Ryan Allister Rumantir Master of Science

Institute of Medical Science

University of Toronto 2013

ABSTRACT

Tumor hypoxia (poor oxygenation) adversely affects patient prognosis by

promoting therapeutic resistance and an aggressive tumor phenotype. We aimed to

understand how urokinase plasminogen activator receptor (uPAR), a cysteine-rich

protein implicated in the malignant phenotype and poor patient prognosis, matures

in hypoxia.

We hypothesized that secretion of uPAR during hypoxia is conferred by a

superior ability to form disulfide bonds without oxygen.

A model and assay was established to monitor the oxygen-dependency of

suPAR (a soluble secreted isoform of uPAR) folding and secretion. We found that

suPAR maturation involves disulfide formation and N-linked glycosylation in

normoxia. In anoxia, suPAR disulfide formation was impaired, but suPAR was

nevertheless secreted. We propose that suPAR has low dependency on disulfide

formation for efficient secretion in comparison to other disulfide-containing proteins.

Mechanisms supporting protein expression during hypoxia may potentially be

targeted to mitigate the adverse effects of tumor hypoxia and ultimately improve

cancer therapy.

 

  iii  

ACKNOWLEDGEMENTS

Working on this thesis for the past two years has undoubtedly been an

exciting and humbling academic adventure. It has been a journey that would have

remained uncharted if not for my supervisor, Dr. Marianne Koritzinsky. So first and

foremost, I would like to thank Marianne for giving me the opportunity to be a part of

her brilliant project. Marianne, you have been a model supervisor- allowing me to

have the creative freedom to explore the scientific process while offering endless

guidance, support and encouragement. No one has invested more time into this

work, and my personal and professional progress than you have. It will always

amaze me how you provided daily epiphanies and doses of genius. You have been

the major influence in my recent development and for that I am ever thankful.

Secondly, much gratitude is owed to Dr. Brad Wouters. Brad, thank you for

your invaluable ideas and for all of your contributions to this project. You have been

a great role model and a vital supplement to my education and development.

To my program advisory committee members, Dr. David Williams and Dr.

Christine Bear, your dedication has been of utmost importance in my maturation as

a graduate student. Thank you for supporting my progress and for ensuring my

proper bearings.

To our external collaborator, Ineke Braakman, thank you for continuing to

offer your expertise to this project.

To the entire Wouters Koritzinsky Lab, thank you for welcoming me into your

lab family, sharing your practical expertise and being great friends. It has been an

 

  iv  

honour and a true privilege to learn from and work along side such a stellar team of

scientists and I wish you all the best in your future pursuits. In particular, I must

thank Ravi for taking the time to introduce me to countless new techniques,

deliberate experimental design and results, and for sharing his expertise in the field

of protein maturation.

To my parents, Henk and Fransisca, and my brother, Oliver, thank you for

undying support and for always keeping my priorities in check. And to the rest of my

family and friends, I cannot thank you enough for the many roles you have played in

my life. I wish only to make you all proud.

Lastly, I would like to express my appreciation to the Terry Fox Research

Institute for funding and making this project possible.

 

  v  

TABLE OF CONTENTS

1 INTRODUCTION  .........................................................................................................................  1  1.1 Cancer  ..................................................................................................................................................  1  1.1.1  Overview  ..........................................................................................................................................................  1  1.1.2  Carcinogenesis  and  Tumorigenesis  .....................................................................................................  1  1.1.3  Metastasis  ........................................................................................................................................................  2  1.1.4  Cancer  Treatment  ........................................................................................................................................  3  

1.2 Hypoxia  ................................................................................................................................................  4  1.2.1  Overview  ..........................................................................................................................................................  4  1.2.2  Tumor  Hypoxia  .............................................................................................................................................  4  1.2.3  Biological  Resistance  Mechanisms  .......................................................................................................  5  1.2.4  Treatment  Resistance  ................................................................................................................................  6  1.2.5  Targeting  Hypoxia  .......................................................................................................................................  7  1.2.6  Biological  Response  to  Hypoxia  .............................................................................................................  8  

1.3 O2-Sensitive Pathways  ..................................................................................................................  8  1.3.1  Hypoxia  Inducible  Factor  (HIF)  .............................................................................................................  9  1.3.1.1  Urokinase  Plasminogen  Activator  Receptor  (uPAR)  .............................................................  11  1.3.2  mTOR  Signaling  .........................................................................................................................................  15  1.3.3  The  Unfolded  Protein  Response  (UPR)  ...........................................................................................  17  1.3.3.1  IRE1  .............................................................................................................................................................  18  1.3.3.2  Activating  Transcription  Factor  6  (ATF6)  ..................................................................................  20  1.3.3.3  PERK  ...........................................................................................................................................................  21  

1.4 Secretory Protein Maturation  ...................................................................................................  24  1.4.1  N-­‐Linked  Glycosylation  ..........................................................................................................................  25  1.4.2  Disulfide  Bond  Formation  .....................................................................................................................  26  1.4.3  Specificity  of  Folding  Factors  ...............................................................................................................  32  1.4.4  ER-­‐Associated  Degradation  (ERAD)  .................................................................................................  33  1.4.5  Golgi  Apparatus  .........................................................................................................................................  34  

2 RATIONALE, AIMS AND HYPOTHESIS  ..........................................................................  38  3 METHODS  ..................................................................................................................................  42  

3.1 Cell Culture  ......................................................................................................................................  42  3.2 Transfection  ....................................................................................................................................  42  3.3 Hypoxia  .............................................................................................................................................  42  3.4 Pulse Chase Assay  ......................................................................................................................  43  3.5 Immunoisolation  ...........................................................................................................................  44  3.6 Gel Electrophoresis  .....................................................................................................................  45  3.7 EndoH and Brefeldin A Treatment  .........................................................................................  46  3.8 PERK Inhibitor  ...............................................................................................................................  47  3.9 qPCR  ..................................................................................................................................................  47  3.10 Antibodies  .....................................................................................................................................  48  3.11 Protein Quantification  ..............................................................................................................  48  3.12 Immunofluorescence  ................................................................................................................  49  

4 RESULTS  ...................................................................................................................................  50  4.1 Aim 1: To Characterize suPAR Maturation in Normoxic Conditions  .......................  50  4.1.1  Establishing  a  Model  for  uPAR  Maturation  and  Secretion  ......................................................  50  4.1.2  Optimizing  Immunoisolation  Techniques  for  suPAR  ................................................................  51  

 

  vi  

4.1.3  A  Transient  Transfection  suPAR  Overexpression  Model  ........................................................  53  4.1.4  Characterizing  suPAR  Electrophoretic  Mobility  Under  Reducing  and  Non-­‐Reducing  Conditions  ...............................................................................................................................................................  56  4.1.5  Optimizing  Pulse  Time  for  suPAR  ......................................................................................................  58  4.1.6  Exploring  the  Possible  Effect  of  Remnant  DTT  on  Disulfide  Bond  Formation  After  a  Reductive  Challenge  ............................................................................................................................................  58  4.1.7  In  Vivo  Reduction  of  suPAR  Following  Radiolabelling  ..............................................................  61  4.1.8  suPAR  Disulfide  Formation  and  Secretion  in  Normoxia  ..........................................................  64  4.1.9  suPAR  Glycosylation  in  Normoxia  .....................................................................................................  66  

4.2 Aim 2: To Characterize suPAR Maturation in Anoxic Conditions  ............................  67  4.2.1  Investigating  the  Influence  of  Glass  Culture  Dishes  on  Protein  Maturation  ...................  68  4.2.2  Protein  Maturation  with  PERK  Inhibitor  ........................................................................................  68  4.2.3  suPAR  Glycosylation  in  Anoxia  ...........................................................................................................  72  circumstances  where  supplementary  stability  is  required  [452].  Complete  EndoH  digestion  of  suPAR  matured  in  anoxia  did  not  resolve  the  larger  suPAR  species,  suggesting  that  this  differential  species  is  not  a  product  of  additional  N-­‐linked  glycosylation  (Figure  4.9).    The  identity  of  the  species  remains  to  be  found.  Therefore,  it  was  concluded  that  the  glycan  modifications  of  suPAR  in  anoxia  were  largely  analogous  to  that  of  suPAR  matured  in  normoxia.  ............................................................................................................................................................  74  4.2.4  suPAR  Oxidative  Folding  in  Anoxia  ...................................................................................................  74  4.2.5  suPAR  Secretion  in  Normoxic  and  Anoxic  Conditions  ..............................................................  76  

5 DISCUSSION  .............................................................................................................................  79  5.1 Technical Limitations  ..................................................................................................................  79  5.1.1  Transfection  Model  ..................................................................................................................................  79  5.1.2  ER  Cargo  Tags  .............................................................................................................................................  80  5.1.3  The  Influence  of  DTT  on  Protein  Maturation  ................................................................................  81  

5.2 suPAR Maturation  ........................................................................................................................  83  5.2.1  Folding  Co-­‐  and  Post-­‐Translationally  ...............................................................................................  83  5.2.2  Oxygen  Dependency  of  Disulfide  Bond  Formation  ....................................................................  85  5.2.3  Abundance  of  Extracellular  suPAR  in  Anoxia  ...............................................................................  87  5.2.4  Secretion  of  Non-­‐Native  suPAR  ..........................................................................................................  88  5.2.5  Potential  of  Preferential  Disulfide  Formation  of  Other  ER  Cargo  in  Anoxia  ...................  90  5.2.6  Identification  of  a  Higher  Molecular  Weight  suPAR  Species  Matured  in  Anoxia  ..........  90  

6 FUTURE DIRECTIONS  ..........................................................................................................  92  6.1 Alternative Pathways of suPAR Secretion  .........................................................................  92  6.2 Further Characterizing suPAR Maturation  .........................................................................  94  6.3 Comparing suPAR Maturation to Other Proteins in Anoxia  .......................................  95  6.4 Investigating Oxygen Dependency of Co- and Post-Translational Disulfide Formation in suPAR  ...........................................................................................................................  95  6.5 Influence of Specific Folding Factors in Anoxia  ..............................................................  96  

7 CONCLUSIONS  ........................................................................................................................  97  

 

  vii  

LIST OF ABREVIATIONS

4E-BP1 4E binding protein 1

a2HS alpha-2-HS-glycoprotein

AAT alpha-1-anti-trypsin

AMPK AMP-activated protein kinase

ARCON accelerated radiotherapy combined with carbogen

ASK1 apoptosis signal-regulating kinase 1

Asn asparagine

ATF activating transcription factor

ATP adenosine triphosphate

ATR ATM- and Rad3-related protein kinase

BAK Bcl2 homologous antagonist/killer

BAX Bcl2-associated X protein

BCL2 B-cell lymphoma 2

BFA Brefeldin A

BNIP3 BCL2/adenovirus E1B 19kDa interacting protein 3

BSA bovine serum albumin

b-Zip basic leucine zipper

CBP CREB-binding protein

CD cluster of differentiation

CFTR cystic fibrosis transmembrane receptor

CHOP C/EBP homologous protein

CHX cycloheximide

 

  viii  

COP coatomer protein

CREB cAMP response element binding protein

CUPS compartments for unconventional secretion

CXCR4 C-X-C chemokine receptor type 4

DsbA disulfide bond formation protein A

DsbB disulfide bond formation protein B

DTT dithiothreitol

ECM extracellular matrix

EDEM ER degradation-enhancing alpha-mannosidase-like protein

EEF2K eukaryotic elongation factor 2 kinase

EIF2α eukaryotic translation factor 2α

EIF2AK3 eukaryotic translation initiation factor 2-alpha kinase 3

EIF4F eukaryotic translation initiation factor 4 F

EMT epithelial to mesenchymal transition

EndoH endoglucosidase H

ER endoplasmic reticulum

ERManI ER mannosidase I

ERSRE ER stress response element

EGFR epidermal growth factor receptor

Epo erythropoietin

ERAD ER-associated degradation

ERGIC ER-Golgi intermediate compartment

ERK extracellular-signal regulating kinase

 

  ix  

ERN1 endoplasmic reticulum to nucleus signaling 1

ERO ER oxidoreductase

FAD flavin adenine dinucleotide

Flu-HA influenza-hemagglutinin

GADD growth arrest and DNA-damage-inducible gene

GalNAc N-Acetylgalactosamine

GβL G-protein β subunit-like protein

GEF GTP exchange factor

GlcNAc N-Acetylglucosamine

GPI glycosylphospatidyl inositol

GPI-PLD GPI-specific phospholipase D

GPx glutathione peroxidase

GRP glucose-regulated protein/binding protein

GTP guanidine triphosphate

HERPUD1 homocysteine-responsive endoplasmic reticulum-resident ubiquitin-like

domain member 1 protein

HIF hypoxia inducible factor

HRE hypoxia response elements

Hrd1 HMG-CoA reductase degradation protein 1

IgG immunoglobulin G

IRE1 inositol-requiring protein 1

IRES internal ribosome entry sites

JNK c-Jun N-terminal kinase

 

  x  

LAMP3 lysosomal-associated membrane protein 3

LDLR low density lipoprotein receptor

LOX lysyl oxidase

LU Ly6/uPAR

MAPK mitogen activated protein kinase

MAX Myc-associated factor X

MCT4 monocarboxylate transporter 4

MEF mouse embryo fibroblast

mLST8 mammalian lethal with sec13 protein 8

MMP metallomatrix protease

mSIN1 stress activated protein kinase interacting protein 1

mTOR mammalian target of rapamycin

MVBs multivesicular bodies

MXI1 (MAX)-interacting protein 1

NANA N-acetyleneuraminic acid

NEM N-ethylmaleimide

NOS nitric oxide synthase

ORP150 150-kDa oxygen regulated protein

OS-9 Osteosarcoma Amplified 9

OST oligosaccharyl transferase

PAI plasminogen activator-inhibitor 1

PDI protein disulfide isomerase

PERK protein kinase (PKR)-like ER kinase

 

  xi  

PGK phosphoglyerate kinase

PP1 protein phosphatase 1

PPIase peptidyl prolyl cis/trans isomerase

PRAS40 proline-rich AKT1 substrate 40

PRDX4 peroxiredoxin 4

pVHL von Hippel-Lindau protein

QSOX quiescin-sulfhydryl oxidase

RBC red blood cell

REDD1 regulated in development and DNA damage responses 1

Rheb Ras homologue enriched in brain

ROS reactive oxygen species

S1P site-1 protease

S2P site-2 protease

Ser serine

SDF1 stromal cell-derived factor 1

SDS-PAGE sodium dodecyl sulfate polyacrylamide gel electrophoresis

SNARE soluble N-ethylmaleimide sensitive fusion attachment protein receptors

SRP signal recognition peptide

suPAR soluble urokinase activator receptor

SYVN1 synoviolin

TF transferrin

TGN trans-Golgi network

Thr threonine

 

  xii  

TRAF2 TNF-receptor associated factor 2

TSC tuberous sclerosis protein

UGGT UDP-glucose glucosyl-transferase

uORF upstream open reading frame

uPAR urokinase plasminogen activator receptor

UPR unfolded protein response

UTR untranslated regions

VEGF vascular endothelial growth factor

VKOR vitamin K epoxide reductase

XBP1 X-box binding protein 1

 

  xiii  

LIST OF FIGURES

Figure 1.1: uPAR and suPAR Structure

Figure 1.2: Plasminogen Activation System and uPAR Interactions

Figure 1.3: Protein Maturation in The Endoplasmic Reticulum (ER)

Figure 1.4: ER-Localized Disulfide Bond Formation

Figure 4.1: Cell Lines with Stable Overexpression of suPAR

Figure 4.2: Comparing Stably and Transiently Transfected suPAR

Figure 4.3: suPAR Maturation in the Pulse Chase Assay

Figure 4.4: Disulfide Bond Formation in Presence of DTT

Figure 4.5: DTT in vivo Reduction of suPAR

Figure 4.6: suPAR Disulfide Formation, Glycan Processing and Secretion in

Normoxia

Figure 4.7: Glass Petri Dishes Do Not Influence Protein Maturation

Figure 4.8: Protein Translation and Maturation in Anoxia with GSK PERK

Inhibitor

Figure 4.9: suPAR Maturation and Secretion in Anoxia

Figure 4.10: suPAR Secretion in Normoxia and Anoxia

 

  xiv  

LIST OF SUPPLEMENTARY FIGURES

Figure S1: uPAR mRNA Induction in Hypoxia

Figure S2: XBP1 Splicing of suPAR-Myc-DDK Transiently Transfected HCT116

Figure S3: suPAR Cellular Localization

Figure S4: Brefeldin A Toxicity of HCT116

Figure S5: Golgi-Dependence of suPAR Maturation

   

 

 

1  

1 INTRODUCTION

1.1 Cancer

1.1.1 Overview   Cancer is a highly prevalent disease with a broad and profound burden on its

victims, their families and society as a whole. Approximately 13 million people

worldwide are diagnosed with cancer and 8 million die from the disease annually.

Cancer is the leading cause of death in the developed world, accounting for 13% of

all human deaths [1]. Beyond the mortality of cancer is the immeasurable emotional

and physical distress inflicted on patients and their families. Thus, it is of dire

importance to global health care and our society to advance our knowledge of and

therapeutics against cancer to combat the ensuing morbidity and mortality brought

upon by this disease.

1.1.2 Carcinogenesis and Tumorigenesis   Cancer is a collection of related diseases in which a series of genetic

alterations accumulate in a single, normal progenitor cell to instigate profound

growth deregulation. Unless the progenitor cell is of hematopoietic origin, this growth

dysregulation leads to formation of a tumor [2, 3]. Though a single cell can

experience up to 105 natural DNA lesions on a daily basis [4], genetic quality control

systems generally correct these errors proficiently. Nonetheless, this quality control

can be bested, conceding genetic mutations. Genetic alterations that confer growth

and survival advantage accumulate through natural selection. Genetic alterations in

oncogenes and tumor suppressors contribute functionally to driving carcinogenesis,

 

 

2  

altering the hallmarks of cancer: dysregulation of proliferation, senescence, death,

interactions with tumor microenvironment (e.g. angiogenesis) and tissue invasion

and metastasis [5]. The uncontrolled growth and compromised anatomical barriers

result in detrimental effects on tissue, organs and bodily systems. Genetic

alterations may also endow the ability for tumor cells to metastasize to distant sites,

amassing both severe local and systemic effects [2].

1.1.3 Metastasis  

Metastases are the development of secondary implants discontinuous with

the primary tumor. Though cancers do not metastasize with equal capacity,

approximately 30% of patients diagnosed with solid tumors also present with

clinically evident metastases, while an additional 20% have concealed metastases

at diagnosis [6].

The metastatic cascade is a complex multi-step process that is currently of

the poorest understood of the so-called “Hallmarks of Cancer” on a mechanistic

level. Epithelial tumor cells lose polarity and detach from adjacent cells in part

through the loss of E-cadherin function via mutational inactivation [7] or suppression

by transcription factors SNAIL and TWIST [8-11]. This characterizes an epithelial to

mesencyhmal transition (EMT) that favors migration and is necessary for metastasis

[12-16]. These cells then degrade the basement membrane and interstitial

connective tissue by releasing, or stimulating stromal cells to release, proteases

such as metallomatrix proteases (MMPs), cathepsin D and urokinase plasminogen

activator receptor (uPAR) [17-22]. The tumor cells penetrate the basement

 

 

3  

membrane, passing through the extracellular matrix (ECM) and intravasating into

blood vessels or lymphatics [23]. Some tumor cells transit as single cells, others

aggregate to form a tumor embolus, which can also include circulating leukocytes

[24]. Although many tumor cells reach the vascular space, only a few eventually

adhere to the vascular endothelium in capillaries [25, 26]. The basement membrane

is again degraded to enable extravasation out of the vasculature and deposition at a

remote site where angiogenesis is promoted to facilitate metastatic tumor growth

[27] [28].

1.1.4 Cancer Treatment  

Current cancer therapies aim to cure, prolong survival or reduce the

symptoms associated with disease. The current standard of care involves surgical

resection, radiotherapy, chemotherapy, hormonal therapy, or a combination of these

therapeutic modalities, depending on the different type, location and stage of cancer.

Chemotherapy treats cancer with cytotoxic agents that kill rapidly dividing cells while

radiation therapy utilizes ionizing radiation to deliver localized lethal DNA damage.

Hormone therapy is also used to combat prostate and breast cancer. More recently,

molecular targeted agents such as monoclonal antibodies or small molecule kinase

inhibitors have been introduced to inhibit cancer cell proliferation more specifically.

Immunotherapy is another new modality that attempts to treat cancer by stimulating

the body’s immune system to destroy tumor cells [29].

Advances in cancer research have greatly improved prognosis for many

cancer types, and valiant efforts have been made to improve therapeutic dosing and

 

 

4  

targeted delivery to optimally reduce normal tissue toxicity and increase efficacy at

the tumor site. Nonetheless, half of the cancer patients being treated die from the

disease [1], highlighting the need for novel treatment strategies.

1.2 Hypoxia

1.2.1 Overview  

While normal tissue is at approximately 5-7% oxygen, hypoxia is defined as

oxygenation below normal levels (i.e. below ~3% O2) [30]. Physiologically, hypoxia is

manifested in embryonic development, exercise and high altitudes and can exist

pathologically in infarction, stroke and solid tumors [31, 32]. Thus, it is of

fundamental and medical importance to better understand oxygen-dependent

cellular processes that drive critical changes in the cellular phenotype.

1.2.2 Tumor Hypoxia  

The tumor microenvironment is extremely heterogeneous in part as a product

of widely irregular vasculature [33], producing tumor regions that vary in

oxygenation, nutrient supply and pH, which ultimately influence gene expression and

cellular behavior. Most human tumors contain regions of hypoxia [34-43]. Tumor

cells are subject to both chronic, diffusion-limited hypoxia and acute, perfusion-

limited hypoxia [44-47]. In tissue, oxygen diffusion is limited by consumption to 100-

200µm, approximately 10-20 cell layers [33] such that tumor cells situated distally to

supporting vasculature may experience chronic hypoxia. Acute hypoxia occurs when

the immature, disorganized and leaky tumor vasculature experiences dynamic

 

 

5  

changes in red blood cell flux, limiting the oxygen supply to surrounding tumor cells

[48-54].

Currently, no ideal method of measuring tumor hypoxia exists. The Eppendorf

needle electrode represents the current clinical gold standard, but its use is declining

because it is invasive, and not available to most centers [55-57]. Efforts have been

put forth to develop endogenous molecular signatures of hypoxia, with some recent

success [58]. There is great interest in exogenous hypoxia markers, such as the 2-

nitroimidazoles, pimonidazole, fluoroazomycin arabinoside and EF5, that bind to

hypoxic cells after a oxygen-sensitive 1-electron reduction after systemic

administration, and can be detected by immunohistochemistry and non-invasive

positron emission tomography respectively [59-61]. When tumor hypoxia is

measured by any of the above mentioned methods, it is clear that substantial

heterogeneity in tumor oxygenation exists within a tumor and between patients [35,

62-64]. Hypoxia in the tumor microenvironment adversely affects patient prognosis

in numerous types of cancer, including cancers of the head and neck, cervix,

prostate, pancreas, brain and sarcomas [34, 37, 57, 65-73], by causing treatment

resistance [33, 34, 50, 57, 71] and promoting a more aggressive, malignant tumor

phenotype [74, 75], as discussed below.

1.2.3 Biological Resistance Mechanisms  

Tumor cells adapt and survive hypoxic conditions by escaping cell death

pathways. For example, hypoxia promotes the phosphorylation of p53 on Ser15 by

ATR (ATM- and Rad3-related protein kinase) [76], resulting in apoptosis [77].

 

 

6  

Hypoxia thereby provides an environment that is selective for cells with a p53

mutation that allows escape of hypoxia-induced apoptosis and thus, survival [78].

Loss of sensitivity to cell death mechanisms could subsequently affect the

propensity to die from anti-cancer agents. Hypoxia also acts to induce genes

promoting drug resistance directly, such as P-glycoprotein [79] [80], an adenosine

triphosphate (ATP)-binding cassette transporter drug efflux pump with broad

substrate specificity [81].

1.2.4 Treatment Resistance  

Hypoxic tumor microenvironments have a substantial negative effect on

patient prognosis by conferring therapeutic resistance [78]. Gray et al. (1953)

hypothesized that tumor hypoxia could limit the efficacy of radiotherapy [82]. Since

then, tumor oxygenation has been shown to be predictive of poor patient outcome

after radiotherapy [37-39, 83-85]. Radiation produces radicals in DNA, resulting in

temporary DNA damage [54]. Oxygen reacts with these DNA radicals with high

efficiency, producing a ’fixed’ DNA lesion that requires enzymatic DNA repair [54,

71]. In the absence of oxygen, the DNA radicals can receive a hydrogen atom from

non-protein sulfhydryls, effectively restoring an undamaged species [54]. As a result,

DNA lesions produced under anoxia (0.0% O2) are decreased to a third of what is

seen in normoxia [86]. Other anti-cancer agents, such as bleomycin [54, 87], also

require oxygen to efficiently impart DNA damage in target cells [88, 89], and

consequently are less potent in the absence of oxygen.

 

 

7  

1.2.5 Targeting Hypoxia  

The efficacy of systemically delivered therapies, including chemotherapy, are

challenged by restricted delivery to a tumor region that is by nature distal to the

already abnormal and ineffective vasculature [90-95]. This causes decreased drug

toxicity with increasing distance from vasculature [87]. Furthermore, chemotherapy

targets highly proliferative cells, however proliferation decreases as a function of

distance from vasculature [96], making targeting of more distant and likely hypoxic

cells more difficult. Thus, therapeutic agents require further development with

regards to delivering efficient and specific cytotoxicity.

Hypoxia is largely unique to tumors, and provides opportunity for selective

cancer therapy. One approach to overcome hypoxia-induced therapeutic resistance

is the use of ARCON, accelerated radiotherapy combined with carbogen (gas

mixture of carbon dioxide and oxygen) and nicotinamide. Carbogen serves to

decrease chronic hypoxia by delivering hyperoxic gas and nicotinamide lowers acute

hypoxia by increasing perfusion [97, 98] to ultimately increase radiation damage [99,

100]. Hypoxic radiosensitizers, such as nimorazole, act as oxygen substitutes in free

radical reactions with greater diffusion capacity than oxygen as they are not subject

to metabolism [101]. There is also potential in bioreactive prodrugs, such as

tirapazamine [34, 102-104]. These prodrugs require one-electron reduction from a

nontoxic state to an active radical that causes DNA damage [102]. The active

molecule however, is also a substrate for oxidation by oxygen back to the original

prodrug [34, 54, 103, 104], providing selective toxicity against hypoxic cells.

 

 

8  

1.2.6 Biological Response to Hypoxia  

A number of adaptive responses grant a tolerance to the hostile, hypoxic

conditions found in tumors. In an effort to adapt to poor oxygen availability, hypoxic

cells exhibit cell cycle inhibition and decreased proliferation [91, 105], decreased

protein synthesis and increased anaerobic metabolism [106-108]. Hypoxia also

stimulates increased secretion of angiogenic factors [109-112]. In cancer cells,

hypoxia promotes increased metastatic potential [109, 113-115] by regulating

proteins involved in matrix degradation, cell migration, invasiveness and thus the

malignant phenotype [71, 116-119]. In consequence, tumor hypoxia has been

strongly associated with tumor progression, migration, invasion [120-122], and

metastasis [37, 39, 44, 113, 123-127]. Although prolonged hypoxia eventually

results in necrosis, another common feature of solid tumors [128], cancer cells are

generally tolerant.

In an effort to uncover and develop novel anticancer therapeutic approaches,

it is of great interest to better understand the mechanisms by which hypoxia

influences cancer biology.

1.3 O2-Sensitive Pathways Hypoxia tolerance is accomplished via changes in gene expression affecting

cellular responses and behavior through at least 3 major oxygen-sensitive signaling

pathways that act to regulate transcription and mRNA translation in response to low

oxygenation [33]. These pathways regulate angiogenesis, metabolism, autophagy,

 

 

9  

endoplasmic reticulum (ER) homeostasis, metastasis, cell migration and invasion,

ultimately characterizing the hypoxic phenotype and malignant progression.

 

1.3.1 Hypoxia Inducible Factor (HIF)

HIF1 is a heterodimeric protein [129], highly expressed in most tumors [130].

Both subunits of HIF1, HIF1α and HIF1β, are ubiquitously expressed. However, in

normoxic conditions, the alpha subunit of HIF1 is unstable as the E3 ubiquitin ligase

von Hippel-Lindau protein (pVHL), binds to HIF1α, targeting it for rapid destruction

by the 26S proteasome [131-134]. pVHL recognition of HIF1α depends on post-

translational hydroxylation of HIF1α’s proline residues 402 and 564 [133]. HIF1α

prolyl hydroxylases (PHD1-3) utilize oxygen as a co-substrate [135, 136]. In hypoxia,

hydroxylation is therefore limited, allowing for the stabilization of HIF1α and thus its

binding to HIF1β. The HIF1 dimer then translocates to the nucleus to initiate target

gene transcription [35, 137]. HIF1 promotes transcription of its target genes by

binding conserved hypoxia response elements (HRE) in gene promoters together

with its co-activator cAMP response element binding protein (CREB)-binding protein

(CBP)/p300 [138, 139]. The result is the induction of over 60 downstream gene

targets involved in promoting tolerance in hypoxia [140] through glycolysis [141], pH

regulation [142] and angiogenesis [41, 143-147]. HIF targets are also implicated in

tumor growth [148-151] and metastasis [70, 139]. HIF is thus associated with poor

prognosis and therapeutic resistance in numerous cancers [130, 152]. Similar to

HIF1, HIF2 can also activate HRE-dependent gene transcription [153], though these

gene targets only partially overlap those of HIF1 [154].

 

 

10  

HIF1 stimulates glycolysis through glucose transporters GLUT-1 and GLUT-3

[155, 156], glycolytic enzymes, such as phosphoglycerate kinase (PGK) [157], and

lactate exporters, such as monocarboxylate transporter 4 (MCT4) [158] to maintain

cellular ATP levels while oxidative phosphorylation is inhibited. In fact, HIF also

inhibits mitochondrial activity directly by upregulating Myc-associated factor X

(MAX)-interacting protein (MXI1) transcription to repress c-Myc transcriptional

activity and promoting proteasomal degradation of c-Myc [159], decreasing c-myc

dependent mitochondrial biogenesis [160], ultimately conserving oxygen during

hypoxia.

To further combat the hypoxic conditions, HIF1 transcriptionally activates

vascular endothelial growth factor (VEGF) [161] to promote angiogenesis from

existing vasculature, stromal cell-derived factor 1 (SDF1) to stimulate the bone

marrow release of endothelial progenitor cells to restore tumor vasculature [101,

162] and erythropoietin (Epo) [154] to increase red blood cell (RBC) production, all

in an effort to restore oxygen levels [71]. Hypoxia also stimulates nitric oxide

synthase (NOS) [163, 164] via HIF1, resulting in vasodilation and subsequent

increased blood flow [71].

Furthermore, as outlined in more detail below, cellular invasion and migration

is increased in hypoxia by HIF1-dependent transcriptional induction of genes such

as urokinase plasminogen activator receptor (uPAR) [165, 166] and lysyl oxidase

(LOX) [167, 168].

 

 

11  

1.3.1.1 Urokinase Plasminogen Activator Receptor (uPAR)  

Of specific focus in this thesis is the urokinase plasminogen activator

receptor (uPAR), also known as cluster of differentiation 87 (CD87).

uPAR is a glycosylphosphatidyl inositol (GPI) anchored membrane-bound [169, 170]

lymphocyte antigen (Ly-6) family member, containing 3 cysteine rich Ly-6/uPAR

(LU) domains (Figure 1.1) [171-173]. It is involved in embryogenesis, ovulation,

inflammation, wound healing, tissue remodeling [174], angiogenesis, cell adhesion,

migration and tumor growth [174-179]. Hypoxic tumors overexpressing HIF are likely

to also overexpress HIF-target uPAR, [120, 180]. Furthermore, uPAR expression is

also stimulated by epidermal growth factor receptor (EGFR) and Ras-MAPK

(mitogen activated protein kinase) signaling [181, 182]. Consistent with this, uPAR

is expressed in most solid and many hematologic malignancies and is restricted

mainly to cancer tissue [179]. uPAR expression also increases with tumor grade or

cancer stage and in metastases [179]. Furthermore, the upregulation of uPAR has

been observed not only at the mRNA level but also at the cell surface in hypoxia

[165].

Extracellular proteolysis is influenced by the production of the serine protease

plasmin by plasminogen activators such as urokinase (Figure 1.2). When the

inactive, pro-uPA binds the central cavity of its receptor uPAR [183], it is cleaved by

plasmin to yield an active uPA where the C- and N-term are held together by a

disulfide bond [184]. uPA-bound uPAR is a protease that cleaves the amino-terminal

prodomain of zymogen plasminogen to produce the active plasmin, that

subsequently degrades the ECM and basement membranes [185, 186], culminating

 

 

12  

 

 

13  

in a positive feedback loop to activate more pro-uPA (Figure 1.2) [187].

Experiments have shown uPA-induced proliferation is uPAR dependent, and

antagonizing uPA and uPAR binding can prevent growth, invasiveness and

metastasis [186].

Plasmin additionally activates pro-collagenase and MMPs, which also

degrade ECM proteins [188-190]. Pro-uPA bound uPAR, along with its plasmin

product, are pronouncedly expressed at the invading edge of tumors [179, 188, 191,

192], facilitating migration by digestion of ECM [186, 193]. As such, the plasminogen

activation system is hijacked in tumors from its normal function in facilitating the

degradation of old tissue, to promote cancer invasion and metastasis [194-196].

In addition to the uPAR’s role in proteolysis, uPAR also forms complexes with

cell surface β1, β2 and β3 integrins [197-202] that serve as cell-ECM adhesion

molecules. Much of uPAR-integrin interaction occurs between uPAR and integrin of

the same cell, regulating cellular shape, cytoskeletal organization, adhesion,

migration and proliferation and thus potentially promoting malignant progression

[165, 186, 203-206]. uPAR-integrin interactions also stimulate a diversity of

important intracellular signaling events [207]. Integrin-dependent signaling is further

enriched by uPAR binding ECM protein vitronectin, which increases the contact

between ECM and the plasma membrane to facilitate the interaction between

integrins and their ligands [208]. uPAR therby promotes migration through a

complex relationship between ECM degradation and cellular adhesion. uPAR-

integrin binding and signaling also regulates tumor growth involving extracellular-

signal regulating kinase (ERK)/ MAPK signaling with the inhibition p38 MAPK [186].

 

 

14  

 

 

15  

Consistent with its influence on malignant progression, uPAR levels have been

correlated with lower survival and poor prognosis in cancer patients [179, 209, 210].

Alternative splicing at the 7th exon produces a soluble variant of uPAR

(suPAR), consisting of an amino-terminal ligand-binding domain without the carboxyl

GPI anchor (Figure 1.1) [169, 195]. This soluble isoform has the same affinity to uPA

as uPAR [173]. Though its biological function is not yet understood, suPAR is

speculated to inhibit cell surface proteolysis locally by sequestering uPA from uPAR

[174, 195]. It may simultaneously induce plasminogen activation at sites aside from

cell surfaces [195]. Membrane-bound uPAR can also undergo cleavage of its GPI

anchor by GPI-specific phospholipase D (GPI-PLD) [211] and MMPs [212], forming

another soluble uPAR [186] released from tumor cells [213]. Urinary [214] and

serum [215] levels of soluble uPAR are elevated in cancer and have been shown to

correlate with poor patient prognosis [215-218], but it is unclear whether this

primarily reflects secreted suPAR or shed surface uPAR.

1.3.2 mTOR Signaling The kinase mammalian target of rapamycin (mTOR) represents another

central signaling pathway that is sensitive to oxygen availability. mTOR is a critical

integrator of metabolic signaling that regulates cell survival and growth through

metabolism, protein synthesis, autophagy and apoptosis sensitivity [33, 219]. mTOR

has been found in 2 complexes, mTORC1 and mTORC2. mTORC1 contains mTOR,

mammalian lethal with sec13 protein 8 (mLST8), proline-rich AKT1 substrate 40

 

 

16  

(PRAS40) and raptor [33] while mTORC2 consists of mTOR, G-protein β subunit-

like protein (GβL) and stress activated protein kinase interacting protein 1 (mSIN1)

[220]. mTOR kinase activity in nutrient-rich conditions stimulates protein synthesis

and cell growth via phosphorylation of ribosomal protein S6 kinase (p70S6K),

eukaryotic initiation factor 4E binding protein 1 (4E-BP1) and eukaryotic elongation

factor 2 kinase (EEF2K) [221]. Hypoxia inhibits mTORC1 activity via multiple routes,

especially in combination with other stressors or in cases of chronic hypoxia [222].

As cellular energy levels drop, mTORC1 is inhibited through AMP-activated protein

kinase (AMPK)-dependent activation of tuberous sclerosis protein 1 (TSC1) in

complex with TSC2 [223, 224]. The TSC1-TSC2 complex negatively regulates the

small G-protein Rheb (Ras homologue enriched in brain) [225], a GTPase that binds

the active site of mTOR to activate mTOR kinase [226]. TSC1-TSC2 activity is also

promoted via HIF-1-dependent transcriptional regulation of regulated in development

and DNA damange responses 1 (REDD1) [227-229] that suppresses mTORC1 by

releasing TSC2 from its inhibitor, 14-3-3 protein [230]. Furthermore, proteins such as

promyelotic leukaemia tumour suppressor (PML) [231] and proapoptotic B-cell

lymphoma 2 (BCL2)/adenovirus E1B 19kDa interacting protein 3 (BNIP3) [232],

physically associate with mTOR and Rheb, respectively, to block their interation

between mTOR and Rheb, inhibiting mTOR activity.

In hypoxia, loss of mTOR activity causes dephosphorylation of 4E-BP1 and

P70S6K to inhibit cap-dependent mRNA translation and cell growth respectively,

presumably in an effort to conserve cellular energy [233, 234]. Since different

mRNAs rely to varying extents on cap-dependent translation, regulation of mTORC1

 

 

17  

activity during hypoxia also results in differential gene expression [235, 236]. Many

genes involved in regulating cell growth and cell death have internal ribosome entry

sites (IRES) sequences in their 5’ UTR allow ribosomal binding independent of cap-

dependent scanning and thus are able to bypass the requirement for cap/eukaryotic

translation initiation factor 4 F (eIF4F)-mediated translation [237, 238]. It remains

unclear however how this differential gene expression specifically contributes to

phenotypic changes during hypoxia.

1.3.3 The Unfolded Protein Response (UPR)  

A third oxygen-sensitive, hypoxia-activated signaling pathway is the

evolutionary conserved unfolded protein response (UPR) of the endoplasmic

reticulum (ER). The ER is a cellular organelle specialized for the maturation and

folding of secreted and membrane-bound proteins, as discussed in more detail

below. An imbalance between the ER’s folding capacity and its protein load results

in the accumulation of unfolded and misfolded proteins [239]. Consequently, ER

stress ensues, which can be caused by inhibition of protein glycosylation, redox

fluctuations, calcium leakage from the ER and ATP deficiency in addition to

increased ER protein cargo load [240, 241]. Our group and others’ have shown that

hypoxia rapidly and robustly activates the UPR [107, 235, 236, 242-244]. There is

also evidence for UPR activation in tumors, suggesting that the hypoxic, nutrient

deprived and acidic tumor microenvironment can perturb ER function resulting in a

stressed ER [245, 246].

 

 

18  

In mammalian cells, the UPR is mediated by ER transmembrane proteins,

eukaryotic translation initiation factor 2-alpha kinase 3 (EIF2AK3) (commonly known

as Protein Kinase (PKR)-like ER kinase (PERK)), endoplasmic reticulum to nucleus

signaling 1 (ERN1) (known as inositol-requiring protein 1 (IRE1)) and activating

transcription factor 6 (ATF6). These 3 sensors of ER stress are kept inactive via

binding to an abundant ER luminal chaperone, glucose-regulated protein/binding

protein (GRP78), also known as BiP [247]. BiP is composed of a C-terminal binding

domain and an N-terminal peptide-dependent ATPase [248], and thus binds and

releases substrates based on ATP release and binding, respectively [249, 250]. As a

molecular chaperone, BiP binds newly synthesized proteins, shielding hydrophobic

domains from aggregation and thereby facilitating proper cargo folding. Upon the

accumulation of unfolded and misfolded proteins, BiP preferentially binds the

exposed hydrophobic patches of aberrant proteins, consequently dissociating from,

and activating PERK, IRE1 and ATF6 [251].

1.3.3.1 IRE1

IRE1 is a type I ER transmembrane serine/threonine protein kinase and

endoribonuclease encoded by the endoplasmic reticulum to nucleus signaling 1

(ERN1) gene. Upon release from BiP or direct binding to unfolded proteins [252],

IRE1 oligomerization and transautophosphorylation activates its cytosolic

endoribonuclease domain to remove a 26 base intron in the pre-mRNA of X-box

binding protein 1 (XBP1) transcription factor [253, 254]. This allows for the synthesis

of active XBP1, which promotes the transcription of genes containing an ER stress

 

 

19  

response element (ERSRE) within their promoters. This population of genes

includes many targets involved in ER protein maturation, such as BiP and protein

disulfide isomerase (PDI), both of which are induced in hypoxia [255, 256].

In contrast, IRE1 kinase activity has been associated with apoptosis through

multiple mechanisms. IRE1 recruits TNF-receptor associated factor 2 (TRAF2) to

activate apoptosis signal-regulating kinase 1 (ASK1, MAP3K5) [257, 258] and

phosphorylate c-Jun N-terminal kinase (JNK) [241]. This promotes the release of

pro-apoptotic Bcl2-associated X protein (BAX) and inhibition of Bcl2 [259]. Activation

of TRAF2 also promotes cleavage of caspase 12 to caspase 9, ultimately resulting

in cell death [248, 260]. Furthermore, IRE1 has been observed to directly interact

with BAX and Bcl2 homologous antagonist/killer (BAK) [261]. The role of IRE1 in

promoting either cell survival or death is controversial and may be cell- and context

dependent [262]. Although IRE1 can signal to the apoptotic machinery as

mentioned above, prolonged IRE1 activity has been observed to enhance cell

survival in HEK293 human embryonic kidney cells in response to tunicamycin, which

activates the UPR by blocking N-linked glycan conjugation [263]. Another recent

study demonstrated no change in overall survival of mouse embryo fibroblasts

(MEFs) when treated with an IRE1 inhibitor in the presence of tunicamycin [264]. In

line with this, our group has found that the same IRE1 inhibitor does not alter cell

clonogenic survival during hypoxia [265]. However, Koong and colleagues showed

previously that MEFs from XBP1 knockout mice have increased apoptosis and

reduced clonogenic survival in hypoxia, as well as impaired tumor growth [266]. The

 

 

20  

role of IRE1 and XBP1 in mediating cell survival and death during hypoxia therefore

remains elusive.

1.3.3.2 Activating Transcription Factor 6 (ATF6)

ATF6α is a type 2 ER transmembrane protein that is involved in cellular

survival in the face of ER stress [267]. When activated, ATF6α translocates to the

cis-Golgi for proteolytic cleavage by site-1 protease (S1P) and site-2 protease (S2P)

[268, 269]. The resulting N-terminal fragment relocates to the nucleus as an active

transcription factor, upregulating the transcription of ER folding factors. ATF6β is

also an ER transmembrane protein, however with a different N-terminal region, and

is a weaker activator than ATF6α though more slowly degraded [270-273]. ATF6

dimerizes with other transcription factors, including XBP1 [274], to regulate genes

with ERSREs in their promoter regions [33]. Cleaved ATF6α and β contain a basic

leucine zipper (b-Zip) domain that facilitates homo- and heterodimeric binding to

ERSREs [274], however dimerization is not necessary for activity [275]. Even

though the importance of ATF6 for hypoxia tolerance is unknown, our group has

observed direct activation of ATF6 during hypoxia (Koritzinsky et al., in review).

Furthermore, indirectly, ATF6 activity in hypoxia is evidenced by the transcriptional

upregulation of folding factors such as BiP, PDI, 94kDA glucose-related protein

(GRP94), ER degradation-enhancing alpha-mannosidase-like protein 1 (EDEM) and

Derlin [276-278], thought to alleviate ER stress by increasing folding capacity [248].

 

 

21  

1.3.3.3 PERK

PERK kinase is a type 1 ER transmembrane protein kinase with a luminal

stress sensor and cytosolic kinase domain that is involved in cellular adaptation to

ER stress [267]. When released from BiP, PERK dimerizes and undergoes

autophosphorylation [267]. Dimerized active PERK can then phosphorylate

eukaryotic translation factor 2α (EIF2α) at Ser51 [279] [248]. EIF2 is part of a ternary

complex (EIF2-GTP-tRNAMet) that recruits the first tRNA to the mRNA start codon

[33]. When phosphorylated, EIF2α prevents the exchange of GDP for GTP,

mediated by EIF2B [33], inhibiting the delivery of initiator methionyl-tRNAi to

ribosomes, thus negatively regulating overall mRNA translation [107]. This serves to

conserve cellular energy, reduce ER protein load and support differential gene

expression [280].

Because protein synthesis is the highest cellular consumer of ATP [281]

[282], mRNA translation is a highly regulated process that is sensitive to numerous

cellular stresses, including hypoxia [283, 284], during which energy production can

become compromised. Hypoxia rapidly inhibits global mRNA translation, by PERK-

mediated phosphorylation of eIF2α [107, 236, 242], reducing protein synthesis

typically by 60-70% within an hour of hypoxic exposure. This global inhibition of

mRNA translation is rapidly reversible upon reoxygenation [107, 244, 285, 286].

Though mRNA translation is globally inhibited in hypoxia, this influences

individual transcripts to a widely variable degree [235, 243, 283, 287]. This variable

effect is at least in part due to differential sequences in the 5’ and 3’ untranslated

regions (UTRs) of specific transcripts [288]. Due to upstream open reading frames

 

 

22  

(uORFs) in the 5’UTR [289, 290], ER stress induced eIF2α phosphorylation leads to

improved translation efficiency of certain proteins. These uORFs serve as decoys for

the translation machinery under normal conditions, preventing translation of the

bona-fide ORF. However, when eIF2α is phosphorylated, the probability of intitiation

at the uORF decreases, rendering it more likely that the initiation complex scans

through to the correct start codon. Hence, controlling global translation also serves

to rapidly and reversibly alter the cellular proteome in hypoxia, as opposed to slower

transcription effects [287].

Translation of activating transcription factor 4 (ATF4) is governed by such

uORFs [290, 291] and thus ATF4 protein is upregulated by eIF2α phosphorylation.

This pathway influences selective gene translation in response to hypoxia [236].

ATF4 target genes are numerous and broad, including those coding for components

of the ER maturation machinery [33], autophagy [292], amino acid metabolism [280]

and several gene products that function to prevent oxidative stress [280, 293].

C/EBP homologous protein (CHOP), also known as growth arrest and DNA-

damage-inducible gene 153 (GADD153), is a transcription factor also induced by

ATF4, and implicated in promoting apoptosis [294]. In a negative feedback loop,

ATF4 also transcriptionally induces growth arrest and DNA-damage-inducible gene

34 (GADD34), which works with protein phosphatase 1 (PP1) to facilitate the

dephosphorylation of eIF2α and recovery of protein synthesis [267, 295, 296].

Nevertheless, protein synthesis rates remain low during prolonged hypoxia, despite

this negative feedback loop, as hypoxia disrupts EIF4F via regulation by 4E-BP1

 

 

23  

and nuclear import factor 4E-T, preventing mRNA recruitment to polysomes and

ultimately inhibiting translation [236] [233].

Activation of PERK and it’s downstream signaling pathway has been

observed by various genetic and pharmacological approaches to be necessary for

hypoxia tolerance in vitro and to support viable hypoxic tumor areas [107, 236, 244,

265, 283, 292, 293]. The specific mechanisms by which PERK promotes hypoxia

tolerance are not completely understood and likely multifactorial. Loss-of-function

genetic screens for hypoxia tolerance in worms have identified several essential

translation factors, suggesting that the inhibition of protein synthesis per se is

important for hypoxia tolerance [297]. ATF4 transcriptional activity has been shown

to be important for several reasons, including the upregulation of proteins involved in

autophagy [292], redox (glutathione) regulation [293] and the BiP chaperone [298].

However, when hypoxic tolerance is no longer manageable amidst prolonged ER

stress, PERK can promote apoptosis through CHOP. Cancer cells are generally

resistant to apoptosis and hence appear to mainly benefit from PERK’s protective

roles [266].

The hypoxia response pathways and selective induction of certain genes in

hypoxia may also be exploited to improve cancer therapy [54]. One strategy may be

to disrupt hypoxia tolerance by targeting mTOR, UPR [33] and HIF [139] pathways.

It is also of interest to block the activities of specific hypoxia-induced proteins,

including LOX [127, 299-301] and uPAR [207, 302, 303] to combat the adverse

hypoxic phenotype. Furthermore, mechanistic understanding of how these pathways

 

 

24  

are activated and support differential gene expression can yield novel therapeutic

targets.

It is currently unknown why the ER is oxygen-sensitive, however UPR

activation suggests that one or more ER-localized protein maturation processes are

oxygen-dependent.

1.4 Secretory Protein Maturation

Approximately one-third of all proteins produced by eukaryotic cells enter the

ER [304, 305] [306], a reticular structure surrounding the nuclear membrane, and

the first organelle of the secretory pathway. Certain professional secretory cells are

able to secrete twice their own mass in protein on a daily basis, and thus experience

extremely high flux through the ER [307].

The ER lumen is characterized by a high protein and calcium concentration

and a unique oxidizing environment necessary for protein folding [306]. An overly

reducing ER environment is unfavorable for co- or post-translational disulfide

formation [308] while an overly oxidizing environment results in inappropriate

disulfides and misfolded proteins [309]. A high concentration of the tripeptide,

glutathione (glutamic acid, cysteine and glycine), which exists as a reduced

monomer (GSH) or oxidized dimer (GSSG), acts as a cellular redox buffer in the ER

[310].

ER-cargo proteins are targeted to the ER by a hydrophobic N-terminal signal

sequence, which is recognized by signal recognition preptide (SRP) that aligns the

translating ribosome with the Sec61 translocon complex in the ER membrane via the

 

 

25  

SRP receptor [306, 311]. The nascent cargo protein is then imported into the ER

lumen co-translationally, during which the signal peptide is rapidly cleaved. The

ensuing protein maturation involves N-linked glycosylation and the enzymatically

regulated formation of disulfide bonds, processes that are unique to the ER. Though

disulfides can form in other cellular compartments, the activity does not support

conformational isomerization as facilitated by folding factors in the ER [312, 313].

Upon entry in to the ER lumen, nascent protein strands are engaged by chaperones

that transiently shield exposed hydrophobic regions of unfolded proteins to prevent

aggregation and consequently assist protein maturation (Figure 1.3) [249, 267, 314].

1.4.1 N-Linked Glycosylation  

Glycosylation is the conjugation of sugar moieties to cargo proteins and

contributes to protein maturation and secretion. The hydrophilicity of glycans

promotes protein solubility [306, 315], mediates interactions with lectins (sugar-

binding proteins) and also influences other protein interactions due to their size.

N-linked glycosylation is a co-translational modification mediated by

oligosaccharyl transferase (OST) in the ER lumen [316]. N-linked core glycans

consist of 3 glucose, 9 mannose and 2 N-acetylglucosamines, that are conjugated to

a protein’s amino group on asparagine in the consensus sequence Asn-X-Ser/Thr,

where X is any amino acid other than proline [317]. This large, branched glycan is

subject to extensive processing, and is used to promote and monitor protein folding

in the ER. After conjugation, glucosidases I and II each trim 1 glucose from the N-

linked glycan [318, 319]. This trimming results in a single remaining glucose on the

 

 

26  

glycan branch that enables binding of the cargo protein to lectin chaperones, such

as membrane-bound calnexin and soluble calreticulin. These lectin chaperones

facilitate the association of cargo proteins to the protein disulfide isomerase

homologue, PDIA3/GRP58/ERp57, which introduces disulfide bonds into cargo

protein [310, 318, 320, 321]. Following disulfide formation, glucosidase II cleaves the

last glucose on the core glycan, releasing the properly folded substrate from its

supporting chaperones [322].

1.4.2 Disulfide Bond Formation   Disulfides are covalent bonds between cysteine residues introduced co- and

post-translationally into cargo proteins by protein disulfide isomerase (PDI) family

members (Figure 1.3, 1.4) in a redox relay that requires a terminal electron acceptor

[323, 324]. Over 20 PDI homologues exist, and some introduce disulfide bonds into

cargo in a thiol-disulfide exchange that begins with the deprotonation of a free thiol

(sulfhydryl) on the cargo protein, forming a thiolate anion, which then displaces a

sulfur of a cysteine in an existing disulfide bond on the N-terminal Cys-X-X-Cys (C-

X-X-C) motif active site of PDI, reducing the disulfide [325-327]. The resulting

transient mixed disulfide is subsequently resolved by the cargos remaining thiolate

anion [328], resulting in the transfer of a disulfide bond from PDI to the cargo protein

in exchange for 2 electrons (reducing equivalents) [310, 328, 329] (Figure 1.4).

Disulfide bonds confer both intra- and intermolecular stability in protein folding [321,

328-330].

 

 

27  

 

 

28  

 

 

29  

PDI family members are characterized by at least one thioredoxin-like domain

that contains a redox active C-X-X-C dithiol/disulfide site [331]. PDIs generally have

2 redox-active A domains and 2 non-catalytic B domains that lack cysteines but host

hydrophobic pockets to facilitate substrate binding [332]. Though non-native

disulfides can form, PDIs can also form mixed disulfides with their cargo and act as

a place holder in the isomerization and rearrangement of disulfide bonds by

additional rounds of disulfide reduction and oxidation to introduce the correct

cysteine pairing [332-335].

In order to support additional rounds of disulfide formation, reduced PDIs then need

to be reoxidized by ER oxidases [310, 321, 328, 336, 337] (Figure 1.4). ER oxidases

ERO1Lα and ERO1Lβ, primarily located in the ER [338], also contain the same C-X-

X-C motif in their active sites [339-341]. These ER oxidases catalyze the reoxidation

of a reduced PDI and receive 2 electrons in return, which are deposited onto an

oxidized flavin cofactor. The reduced flavin can be oxidized by oxygen, which

thereby can serve as the terminal electron acceptor of this redox relay, yielding a

hydrogen peroxide molecule [339, 342-346]. In bacteria, cytoplasmic thiol-disulfide

oxidoreductase DsbB (disulfide bond formation protein B) functions to oxidize

periplasmic DsbA (disulfide bond formation protein A), the direct donor of disulfide

bonds to cargo protein [347-349], akin to ERO1 and PDI activity. DsbB is similarly

reoxidized by quinone reduction [350], which is then reoxidized by oxygen [342, 349,

351]. Ultimately, these reactions must be driven by a terminal electron acceptor,

potentially represented by oxygen, sulfur or other compounds [342, 352].

 

 

30  

Since a molecule of hydrogen peroxide is produced in disulfide bond

formation, ERO activity is tightly regulated. Sevier et al. describe a homeostatic

feedback system that exploits the competition between oxidized ER cargo and yeast

ERO1p’s regulatory cysteine pair, stimulating disulfide formation within ERO1p’s

regulatory cysteine pair when exposed to an overly oxidizing ER environment. This

results in decreased ERO1p activity. On the other hand, a reducing environment

reduces ERO1p’s regulatory disulfide and increases its activity and subsequent

disulfide bond formation in ER cargo. Thus without cargo, ERO1p oxidizes its

regulatory disulfides to promote its inactivation in cis or in trans [353], protecting

against excessive activity and ensuing reactive oxygen species (ROS) production.

The loss of yeast ERO1p results in a backlog of reduced PDI, thus halting

disulfide bond formation [343]. However, mice lacking both ERO1Lα and ERO1Lβ

are viable with minor phenotypes, suggesting [354] ERO independent disulfide bond

formation [355].

Recently, two ERO1-independent disulfide formation pathways have been

shown to be active in mammalian cells. ER-localized peroxiredoxin 4 (PRDX4) has

been observed to couple the reduction of hydrogen peroxide to oxidation of reduced

PDIs, resulting in disulfide bond formation. In the presence of ERO, this potentially

also serves to detoxify ROS from ERO activity [356-361]. PRDX4 can also oxidize

GSH to GSSG in a PDI-dependent manner [361] to support oxidative folding.

Disulfide formation can also be facilitated by the ER transmembrane protein vitamin

K epoxide reductase (VKOR) [362]. In the reduction of vitamin K epoxides to vitamin

 

 

31  

K hydroquinone, VKOR forms a disulfide in its C-X-X-C motif [363] that can be

relayed to PDI substrates [362, 364-366].

In addition to these pathways with proven functional contributions to disulfide

bond formation in living mammalian cells, other oxidases have been suggested to

contribute on the basis of in vitro studies. The addition of glutathione peroxidase

(GPx) 7 or 8 [367] to an in vitro mixture of reduced denatured protein, PDI and

peroxide resulted in disulfide bond formation, suggesting that the ER localized GPx7

and GPx8 can re-oxidize PDI [368, 369]. Similar to PRDX4, GPx7 and GPx8 are

suggested to process ERO1-produced peroxide to drive further disulfide formation

within the ER, while also eliminating ROS [368]. However, ERO-mediated oxygen

consumption also increases in vitro in the presence of GPx7, suggesting that it may

promote ERO activity [368]. The only study demonstrating a functional role for GPx7

in ER-localized protein folding in living cells suggests a mechanism in which GPx7

forms a mixed disulfide with BiP to promote its chaperone activity, rather than a

direct role in disulfide bond formation [370]. In addition, mammals have 2 orthologs

of transmembrane flavoprotein quiescin-sulfhydryl oxidase (QSOX), QSOX 1 and 2,

which directly facilitate oxidative folding in cargo protein in vitro by reducing oxygen

to hydrogen peroxide [371-374]. However, due to its inability to isomerize non-native

disulfides, QSOX activity benefits from PDI presence [375]. QSOX1 also restores

disulfide formation in ERO1-deficient yeast [372], but overexpression or shRNA

mediated knockdown in mammalian cells have so far yielded no phenotypes [376]

[330]. Interestingly, QSOX1 has also been observed to be secreted from cultured

fibroblasts [377]. The Fass group has recently demonstrated an extracellular role for

 

 

32  

secreted QSOX, being required for the incorporation of basement membrane protein

laminin into the ECM and thus displaying an important role in cell migration [378].

1.4.3 Specificity of Folding Factors  

It has been suggested that PDI homologues vary in substrate specificity

[310], and ERO1Lα and ERO1Lβ differ in tissue distribution and transcriptional

activation [346]. Nonetheless, little is known about the requirements for PDI

homologues and ER oxidases in different conditions or with specific cargo proteins.

Disulfide trapping exploits the mixed disulfide formed between an oxidoreductase

and its substrate by mutating a second cysteine that cannot resolve the mixed

disulfide. These mutants have been utilized to better understand the cargo

specificity of PDI homologues [356, 362, 379, 380]. A few examples of specificity of

folding factors to certain cargo have been described in the literature. Rutkevich et

al., 2010 have shown differential sensitivity of maturation and secretion in albumin,

alpha-fetoprotein, alpha-2-HS-glycoprotien (a2HS), alpha-1-anti-trypsin (AAT) and

transferrin (TF) to the depletion of PDI and PDI homologues ERp57, ERp72 and P5

[381].

Many ER-localized folding factors are upregulated in numerous cancer types

[246]. BiP and GRP94 are upregulated in over 10 cancers [382, 383]. PDI

homologue, ERp29, and ERO1 are increased in skin and breast cancer [384-387],

and calreticulin is overexpressed in colorectal carcinoma [388] and calnexin in

breast cancer [389]. BiP on the cancer cell surface localizes and binds uPAR

resulting in proximal cellular migration and invasion [390]. It is possible that

 

 

33  

upregulation of these UPR targets in solid tumors is a result of hypoxia within the

tumor microenvironment. ER-localized ATPase chaperone 150-kDa oxygen

regulated protein (ORP150) has been shown to be upregulated in hypoxia to support

hypoxic tolerance [391, 392].

1.4.4 ER-Associated Degradation (ERAD)   Proteins folded in the ER fall under the tutelage of a strict quality control

system that mitigates the advancement of unfolded and misfolded cargo proteins in

the secretory pathway. Unfolded and misfolded proteins exhibiting exposed

hydrophobic patches, are recognized by UDP-glucose glucosyl-transferase (UGGT),

which introduces a single glucose onto the glycoprotein and permits re-entry into the

lectin-facilitated protein folding cycle [393]. This recycling can continue until proper

folding is achieved and the protein can be exported to the Golgi apparatus.

Terminally aberrant proteins are slowly trimmed of their mannose residues on N-

linked glycans by ER mannosidase I (ERManI) [394, 395], increasing recognition by

OS-9 (Osteosarcoma amplified 9) and XTP3-B, which target misfolded or unfolded

proteins for ERAD   [396]. After the loss of 3-4 mannose residues, chronically

misfolded or unfolded proteins are retrotranslocated out of the ER to the cytosol,

possibly via the SEC61A1 translocon [397]. Homocysteine-responsive endoplasmic

reticulum-resident ubiquitin-like domain member 1 protein (HERPUD1, also known

as HERP), in complex with E3 ubiquitin-protein ligase HMG-CoA reductase

degradation protein 1 (Hrd1) (also known as synoviolin (SYVN1)) transfers ubiquitin

to malformed cargo tagging them for proteasomal degradation [398] [399, 400].

 

 

34  

ERAD thereby acts as a critical mechanism of detoxification. Otherwise, protein

misfolding results in aggregation that must be cleared by autophagy [306, 401-403].

Many proteins involved in ERAD are also induced in hypoxia, including HERP,

SEC61A1 and Hrd1 [33]. Despite ERAD activity, some cargo, such as low density

lipoprotein receptor (LDLR) and mutated cystic fibrosis transmembrane receptor

(CFTR) [404], mutated AAT [405], V2 vasopressin receptor [406], and cochlin [407]

can escape to the Golgi in misfolded states. ER export signals have been proposed

to contribute to this trafficking of misfolded proteins as it has been suggested that

the ERAD and ER export machineries may compete for misfolded proteins [408].

1.4.5 Golgi Apparatus   The Golgi apparatus is a stack of cisternae, compartmentalized into the ER-

facing cis-Golgi, medial-Golgi, and trans-Golgi networks. Following maturation in the

ER, properly folded proteins are concentrated into coatomer protein COPII-coated

transport vesicles that bud off from the ER membrane and assimilate into the ER-

Golgi intermediate compartment (ERGIC) in anterograde export [409]. After

traversing the ERGIC, COPII-coated vesicles fuse to the cis-Golgi [410] via vesicular

soluble N-ethylmaleimide sensitive fusion attachment protein receptors (V-SNAREs)

on the surface of incoming vesicles and target (T)-SNAREs on the surface of the cis-

Golgi that bind one another, resulting in the fusion of the two membranes [411-413].

Upon entry into the Golgi, all N-linked glycans are modified in a

compartmentalized, stepwise maturation procedure producing a complex glycan

[393, 414]. At the cis cisternae of the Golgi, mannosidase I cleaves multiple

 

 

35  

mannose residues. Upon cleavage, the product is advanced to the medial cisternae,

containing N-acetylglucosamine (GlcNAc) transferase I and mannosidase II, and

finally to the trans cisternae, home to galactose transferase and N-

acetyleneuraminic acid (NANA) transferase. O-linked glycosylation is also a Golgi

localized modification in which a sugar molecule, often N-acetylgalactosamine

(GalNAc), is conjugated to oxygen on serine or threonine, providing protein

solubility, stability and facilitating proper conformation [415]

Finally, mature proteins are transported out of the trans-Golgi via vesicles to

traverse the trans-Golgi network (TGN) to endosomes, lysosomes, cell surface or

back to the ER [416]. Proteins, including those containing the ER retrieval KDEL-

sequence, can be recycled to the ER by retrograde transport from the Golgi to ER

facilitated by COPI- coated vesicles [417].

In contrast to the canonical ER-Golgi secretory pathway, compartments for

unconventional secretion (CUPS) have been suggested to facilitate endosome-

mediated secretion of protein without signaling-sequences [418].

1.4.6 Oxygen Dependency of ER-localized Protein Maturation

The unfolded protein response is initiated within minutes of exposure to

anoxia   [236], suggesting a requirement for oxygen in at least one ER-localized

protein maturation process. It has been speculated that low oxygenation could result

in reduced cellular energy and thereby limit ATP-dependent protein maturation steps

as early in the secretory pathway as translocation by the Sec61 membrane complex

 

 

36  

[419]. ATP is also required for correct disulfide formation [420, 421], folding [421,

422], isomerization [420] and secretion [422-424], likely due to the ATP-dependency

of BiP activity [249, 420, 425]. However, the fact that UPR is activated within

minutes of anoxia while it takes hours if not days to affect ATP levels, renders this

an unlikely explanation for hypoxia-induced UPR.

Another process suggested to be oxygen dependent is disulfide bond

formation. The obligate terminal electron acceptors in this process are not fully

elucidated, but may include oxygen. In solution, molecular oxygen has been shown

to act as a terminal electron acceptor, supplying the oxidative potential for disulfide

bond formation by flavin adenine dinucleotide (FAD)-dependent reoxidation of yeast

Ero1p and human Ero1Lα [339, 426, 427]. Oxygen has been interpreted to be the

preferred terminal electron acceptor of disulfide formation in the yeast ER, as yeast

cells with Ero1 mutants were unable to grow in anaerobic conditions [342]. However,

it was also found that oxygen is not required for Ero1p mediated oxidative folding, as

these Ero1 mutants were capable of anerobic growth with FAD1 overexpression and

thus capable of using alternative terminal electron acceptors [342]. Similarly, an FAD

mutation in wild-type Ero1p yeast cells resulted in decreased growth in anoxia [342].

Overexpression of yeast ER oxidase Erv2p in Ero1p mutant yeast cells also

suppressed the growth defect of Ero1p loss in anoxia, suggesting that Erv2p can

also use terminal electron acceptors other than oxygen [342]. Furthermore, PDI1p

remains oxidized in anaerobic yeast [343].

The discovery of hypoxia-specific protein maturation pathways may provide

promising anticancer therapeutic targets, as many secreted and membrane-bound

 

 

37  

proteins facilitate the adverse hypoxic phenotype. Our group has shown that in the

context of human cells, ER-localized N-linked glycosylation, glycan trimming, Golgi-

localized complex glycosylation and protein transport proceeded independently of

oxygen (Koritzinsky et al., in review). To assess the oxygen dependency of disulfide

bond formation, an assay was used where (partially) folded ER-localized cargo

proteins were reduced with DTT and disulfide bond formation monitored in normoxia

or anoxia after removal of DTT. Influenza-hemagglutinin (Flu-HA) and albumin were

able to form disulfide bonds when matured in normoxia, but oxidative folding was

defective without oxygen (Koritzinsky et al., in review). This result demonstrates that

there is a fundamental difference between disulfide bond formation in normoxia and

anoxia, with post-translational re-folding being completely dependent on oxygen.

As a result, in hypoxia, proteins are trapped in unfolded or non-native

conformations, having a profound impact on global protein secretion (Koritzinsky et

al. in review). The observed requirement for oxygen in disulfide bond formation in

living mammalian cells contrasts reported secretion of hypoxia-induced proteins

during anoxia. It is therefore possible that the requirement for oxygen in disulfide

formation is not absolute for all ER cargo proteins. Preferential maturation of ER

cargo proteins in hypoxia may be a product of a competitive advantage for an

oxygen-independent ER maturation machinery.

Our specific interest in this thesis is on a hypoxia-induced disulfide-containing

glycoprotein, uPAR, important for tumor progression and metastasis as outlined

above. Its extracellular upregulation in conditions of low oxygenation suggests that

its maturation is less dependent on oxygen than other ER cargo investigated to date.

 

 

38  

2 RATIONALE, AIMS AND HYPOTHESIS

As outlined above, tumor hypoxia (poor oxygenation) adversely affects

patient prognosis by promoting a more aggressive, malignant tumor phenotype [33,

34, 71, 428]. Thus, it is of critical importance to uncover the molecular and cellular

responses to low oxygenation with the ultimate goal of targeting hypoxia adaptation

in an effort to improve current anticancer therapeutic strategies.

Hypoxia activates oxygen-sensitive signaling pathways that stimulate cellular

responses, such as angiogenesis, metastasis, cell migration and invasion. These

responses contribute to an aggressive tumor phenotype that associates with poor

patient prognosis [37, 65-69]. Many of the proteins responsible for this aggressive

phenotype are secreted proteins that interact with the ECM, tumor stroma and

surrounding tissues. Before hypoxia-induced proteins exit the cell and can influence

tumor progression, they first need to mature and fold in the ER.

We and others have observed that hypoxia activates the UPR [107, 236, 244,

292, 293, 429]. UPR activation suggests that the ER is unable to support normal

protein maturation capacity in hypoxia, bringing forward the possibility that one or

more ER-localized protein maturation processes are oxygen dependent. Our group

has demonstrated a requirement for oxygen in disulfide bond formation in specific

ER cargo like albumin, low-density lipoprotein receptor and influenza hemagglutinin.

Glycan processing and protein transport were in contrast found to be independent of

oxygen (Koritzinsky et al., in review).

Oxygen-dependent disulfide bond formation in the cargo investigated to date

is contrasted by the reported induction of certain proteins in the extracellular space

 

 

39  

in hypoxia, including uPAR and its soluble isoform (suPAR), disulfide-containing

proteins implicated in the malignant phenotype and poor patient prognosis [210,

218, 430-435].

We hypothesize that secretion of suPAR during hypoxia relies on a superior

ability to introduce disulfide bonds in the absence of oxygen.

This hypothesis will be assessed by the following aims and approaches:

Aim 1: To Characterize suPAR Maturation in Normoxic Conditions

A) To establish a model and optimize the pulse chase assay to facilitate the

assessment of suPAR maturation and secretion

HCT116 human colorectal carcinoma cells will be transfected to overexpress

suPAR tagged with Myc and DDK polypeptides to facilitate protein detection and

isolation. These cells will be used to establish an assay to monitor suPAR

maturation and secretion. We will use a pulse chase assay in which proteins are

radioactively labeled during de novo synthesis (pulse), immunoisolated after various

periods of maturation (chase), resolved on sodium dodecyl sulfate polyacrylamide

gel electrophoresis (SDS-PAGE) gels and detected by autoradiography. We will

optimize the assay for signal strength and specificity, and assess the feasibility of

monitoring suPAR disulfide bond formation by changes in electrophoretic mobility.

 

 

40  

B) To characterize the maturation and secretion of suPAR

Using the tools developed in Aim 1A), the kinetics of disulfide formation,

glycan processing and secretion of suPAR in normoxic conditions will be

characterized.

Aim 2: To Characterize suPAR Maturation in Anoxic Conditions

A) To validate the pulse chase assay for use in Anoxia

Modifications to the pulse chase are necessary for the assessment of suPAR

maturation in anoxia. This includes the use of glass petri dishes that do not release

oxygen and addition of the PERK inhibitor VP2323 to allow for protein synthesis. We

will assess the impact of these modifications on the protein maturation assay

established in Aim 1A.

B) To characterize the oxygen dependency of suPAR maturation

The established pulse chase assay will be utilized to characterize the oxygen

dependency of suPAR maturation, addressing the kinetics of disulfide formation,

glycan processing and secretion in the absence of oxygen.

Understanding the oxygen dependency of ER localized protein maturation in

human cells will provide fundamental knowledge regarding the mechanisms of

protein maturation. Protein folding and disulfide bond formation may serve as a

novel mechanism of regulating extracellular expression during oxygen deprivations

relevant for the tumor microenvironment. This phenomenon may potentially be

 

 

41  

exploited to complement chemo- and radiotherapy by mitigating the adverse effects

of tumor hypoxia and ultimately improve cancer therapy.

 

 

42  

3 METHODS

3.1 Cell Culture

Cell lines used were HepG2 (human hepatocellular carcinoma, ATCC: HB-

8065™), HCT116 (human colon carcinoma, ATCC: CCL-247™), HT29 (human

colon carcinoma, ATCC: HTB-38TM) and ME180 (human cervical carcinoma, ATCC:

HTB33TM). Cells were kept in exponential growth phase as adherent monolayers.

HepG2 and ME180 cells were grown in DMEM, HCT116 in RPMI and HT29 in

McCoy’s 5A, all with 10% fetal bovine serum (Gibco).

3.2 Transfection

Cells were transfected in 6-well plates with 2 µg suPAR-Myc-DDK cDNA

(OriGene) and/or 3 µg albumin cDNA (OriGene) with 4 µl Lipofectamine 2000

(Invitrogen) according to manufacturer’s instructions. For establishment of cell lines

stably expressing the transgene, cells were selected with 2 mg/ml G418 Geneticin®

(Invitrogen).

3.3 Hypoxia

Cells were exposed to hypoxia and anoxia in H35 and H85 HypOxystations

(Don Whitley Scientific), respectively. These sealed glove boxes maintain oxygen at

set concentrations. Oxygen content within the HypOxystations was monitored

internally by oxygen sensors as a part of the real-time feedback systems to ensure

oxygen level accuracy to the set point, and independently by Series 3000 Trace

Oxygen Analyzer/Sensor (Alpha Omega Instruments). On the settings used, this

 

 

43  

instrument measured oxygen concentration with an accuracy of 100 ppm (parts per

million).

All buffers and media were deoxygenated overnight in the anoxic chamber

with volumes readjusted with deoxygenated water to compensate for evaporation. At

introduction to anoxia, cells were washed 3 times with deoxygenated media to

remove oxygen.

3.4 Pulse Chase Assay

The core method used was the pulse chase assay  [436]. Cells were grown in

sterile 30mm tissue culture dishes to 80% confluency in a 37oC, humidified, 5% CO2

incubator. Just prior to experimentation, cells and media were transferred to a 37oC

water bath. Cells were washed with 1ml of PBS (Gibco), and starved for 15 min in 1

ml of starvation media (serum-free DMEM without methionine or cysteine, 10 mM

HEPES pH 7.5). Starvation media was aspirated and 200µl of pulse media

(starvation media + 50 µCi EasyTag™ EXPRESS35S Protein Labeling Mix (Perkin

Elmer)) was added to the dish, for a specified duration. At the end of the labeling,

cells were washed once and incubated 5min with 1ml of reducing chase medium

containing 10% FBS, 10 mM HEPES pH 7.5, 5 mM methionine, 5 mM cysteine, 1

mM cycloheximide (CHX) with 5 mM DTT (all Sigma-Aldrich) to stop the

incorporation of radioactivity and reduce disulfide bonds created during the pulse

period. For anoxic conditions, the cells were then brought into the anoxic chamber

following this step. Cells were then washed 3 times and incubated with 1 ml DTT-

free chase media. The cells were incubated on the water bath, or in the incubator in

 

 

44  

the case of long chase intervals. At the end of the chase duration, chase medium

was collected into a microcentrifuge tube and the dish was placed on ice, washed

once then incubated with 1ml of ice-cold stop buffer (PBS with 20 mM N-

ethylmaleimide (NEM)) to terminate protein maturation and irreversibly alkylate any

free cysteines. The stop buffer was aspirated and 300 µL of RIPA lysis buffer (150

mM NaCl, 1% IPEGAL CA-630 (NP-40), 0.5% Na-deoxycholate, 0.1% SDS, 50mM

Tris pH 7.5) with 20 mM NEM (all Sigma-Aldrich) and Halt Protease Inhibitor

Cocktail (Pierce) was added to the dish.

Cells were scraped off and collected into a microcentrifuge tube. Lysates

were vortexed, left on ice for 5 minutes and vortexed again followed by

centrifugation of lysates and chase media at 14,000g for 15 minutes at 4oC using a

microtube centrifuge 5417R (Eppendorf) to pellet nuclei and debris. The supernatant

was collected. 5µl of lysate was taken as a sample of total lysate and added to 5 µl

of 2x reducing loading buffer (0.5 M Tris-HCl pH 6.9, 10% SDS, 20% glycerol,

0.01% bromphenol blue). The remaining samples were flash frozen and stored at -

80oC.

3.5 Immunoisolation

Dynabeads were washed with 1ml of PBS and resuspended in the original

volume of PBS. 50 µL of Dynabeads Protein G (Invitrogen) and 6 µl of anti-uPAR (R

and D Systems) or 50 µL Dynabeads Protein A (Invitrogen) and 3 µl of anti-Albumin

(Sigma-Aldrich) were added to the lysate or media samples. The mixture was

incubated end over end overnight at 4oC. The suspension was briefly centrifuged to

 

 

45  

remove suspension in the cap, and placed in a magnetic rack (DynaMagTM-2) to

gather the bead-antigen complex. The supernatant was aspirated and the beads

washed 3 times with RIPA wash buffer (10 mM Tris pH 7.4, 150 mM NaCl, 0.02%

NaN3 (Sigma-Aldrich), 0.5% NP40). In experiments of serial immunoisolation, 6 µl

anti-uPAR antibody was incubated with 50 µl Protein G Dynabeads for 2 hours, end

over end, at 4oC. Beads and bound antibody complex were resuspended in an

equivalent volume of PBS and added to lysate or media samples and incubated end

over end at 4oC overnight. Immunoisolated uPAR was collected and supernatant

was immunoisolated for albumin. After the final wash, 30 µl 2x non-reducing sample

buffer was added to the beads and vortexed. Samples were boiled for 5min at 95oC,

vortexed and centrifuged for 1min at 14,000 g at room temperature with the resulting

supernatant representing the non-reduced sample. 15 µl of supernatant was

transferred to a new tube to which 2 µl of 500 mM DTT was added. These samples

were boiled again for 5 min at 95oC, vortexed and centrifuged for 1 min at 14,000 g

producing the reduced sample.

3.6 Gel Electrophoresis

Polyacrylamide gels (0.75 mm thick, 12%) were prepared and 15 µl of the

immunoisolated sample or 5 µl of total lysate was loaded, leaving the outer lanes to

be loaded with 1x loading buffer when possible. When reduced and non-reduced

samples were run on the same gel, 2 lanes between reduced and non-reduced

samples were loaded with 1x non-reducing loading buffer to avoid diffusion of

reducing agents into non-reduced samples. Gels were run at 200 V to a preferred

 

 

46  

resolution and stained with 0.25% Coomassie brilliant blue (in 30% methanol

(Sigma-Aldrich) and 10% acetic acid (Sigma-Aldrich)) on a Belly Dancer shaker

(Stovall) for 5min. The stained gels were washed with 30% methanol and 10%

acetic acid for 30 min, then neutralized (30% methanol in PBS) for 5 min, all on the

shaker. The gels were then dried using Gel Dryer 583 (Bio-Rad) on 2 sheets of

Whatman filter paper. The dried gels were exposed to a phosphor-imaging screen

(GE healthcare) for a minimum of 2 days and signal detected on a Typhoon 9410

Scanner (Amersham Biosciences).

Quantification: Signal detected by the Typhoon Scanner was quantified by the

intensity of a band within defined dimensions of individual lanes, using ImageQuant

(GE Healthcare). Quantification data were analyzed using paired Student’s t-test.

Values of p<0.05 were considered statistically significant.

3.7 EndoH and Brefeldin A Treatment

For cleavage of N-linked glycans, immunoisolated protein samples were split

into two equal fractions that were resuspended in 15 µl 100 mM sodium acetate

(Sigma-Aldrich) + 0.2% SDS, pH 5.4. An additional 15 µl 100mM sodium acetate

was added along with Halt Protease Inhibitor and 0.5 µl Endoglycosidase H (EndoH)

(Sigma-Aldrich) into one fraction. Both the EndoH treated and untreated samples

were incubated at 37oC for 2 hours. For complete digestion, samples were

incubated overnight in 1.5 µl EndoH. Following EndoH digestion, samples were

 

 

47  

resuspended in 15 µl of 2x reducing loading buffer, boiled for 5min at 95oC, vortexed

and centrifuged for 1 min at 14,000 g.

To assess the involvement of the Golgi apparatus in maturation, Brefeldin A

(BFA) (Cell Signaling) at specified concentrations was added to growth, starvation,

pulse and chase media.

3.8 PERK Inhibitor

PERK inhibitor was obtained from GlaxoSmithKlein (G797800) or as a gift

from Dr. Uehling (Medicinal Chemistry, OICR) (VP2323) where indicated. Cells were

pretreated with inhibitor for 1 hour in normoxia and exposed to experimental oxygen

conditions where it was added to starvation, pulse and chase media.

3.9 qPCR

RNA was isolated using TRI Reagent® (Sigma-Aldrich) and reverse

transcribed on Mastercycler® ep Gradient S (Eppendorf) with qScript™ (Quanta

Biosciences) according to the manufacturer’s specifications. qPCR was performed

using Realplex (Eppendorf). Relative expression of genes of interest was calculated

using the standard-curve method and normalized to the expression of HPRT1.

Primers used were the following:

uPAR Forward AATGCATTCGAGGTAACGG

uPAR Reverse AGCCTTACCGGTTGTGTG

uPAR Variant 1 Forward AACCGCCTCAATGTGCCAAC

uPAR Variant 1 Reverse AGGTCTGGGTGGTTACAGC

uPAR Variant 2 (suPAR) Forward ACGCTCACTCTGGGGAAGC

 

 

48  

uPAR Variant 2 (suPAR) Reverse TGGGGCTCTATCTCCACATG

XBP1 Spliced Forward CGCTTGGGGATGGATGCCCTG

XBP1 Spliced Reverse CCTGCACCTGCTGCGGACT

XBP1 Total Forward GGCATCCTGGCTTGCCTCCA

XBP1 Total Reverse GCCCCCTCAGCAGGTGTTCC

HPRT1 Forward CCTGGCGTCGTGATTAGTGAT

HPRT1 Reverse AGACGTTCAGTCCTGTCCATA

3.10 Antibodies

For immunoisolation and immunoblotting, the antibodies used were against

uPAR 1:2000 (R and D systems, BAF807), EIF4E 1:5000 (BD Biosciences),

Albumin, c-Myc 1:2000 and FLAG 1:1000 (Sigma Aldrich; A4033, M4439 and F3165

respectively). Secondary antibodies used were HRP-linked Anti-Goat 1:2000 (R and

D Systems, HAF109) and Anti-Mouse 1:2000 IgG (GE Healthcare, NA9310V).

3.11 Protein Quantification

Protein in lysates for Western blotting was quantified using Pierce BCA

Protein Assay (Thermo Scientific) using the standard test tube protocol and

microplate procedure according to manufacturer’s specifications. Absorbance was

measured by microplate reader (Omega).

 

 

49  

3.12 Immunofluorescence

Glass slips (18mm) were placed in 12 well plates on which 200,000 cells

were seeded for 24 hours. Wells were washed 2 times with PBS for 5 minutes per

wash, then fixed with 4% paraformaldehyde in PBS for 20 min. Fixed cells were

washed 3 times with PBS for 5 min and incubated in 100mM glycine (Sigma-Aldrich)

in PBS for 15min to remove remnant paraformaldehyde. The cells were

permeablized with 0.1% Triton X-100 (Thermo Scientific) in PBS for 15 min and

washed 2 times in PBS for 5 min. Blocking was performed in 2% Bovine Serum

Albumin (BSA) (Sigma-Aldrich) and 2% Blocking Grade Milk (Bio-Rad) in PBS for 1

hour. The blocked cells were then washed twice with PBS for 5 minutes and

incubated in 500µl of 1:500 anti-FLAG (DDK tag) for 1 hour. Following 3 washes

with PBS for 5 min, cells were incubated in 500µl of 1:1000 anti-mouse FITC F9137

(Sigma-Aldrich). The cells were washed an additional 3 times for 5 min with PBS

and mounted with 10µl of DAPI-Fluoromount G (Southern Biotechnology).

Fluorescence was visualized in high resolution using Zeiss LSM700 confocal

inverted microscope (Zeiss) at 60x/1.4 NA oil immersion. Images were captured

using LSM Zen 2009 (Zeiss) acquisition software.

   

 

 

50  

4 RESULTS

4.1 Aim 1: To Characterize suPAR Maturation in Normoxic Conditions  A) To establish a model and optimize the pulse chase assay to facilitate the

assessment of suPAR maturation and secretion

4.1.1 Establishing a Model for uPAR Maturation and Secretion

Our overall goal was to test the hypothesis that suPAR possesses

advantageous maturation and folding in the ER in the absence of oxygen. This

hypothesis was based on reports in the literature of the induction of uPAR mRNA

and increased secretion in hypoxia [165, 210, 437-443]. To first validate that uPAR

is indeed induced under hypoxic conditions in our hands, two cell lines reported to

express uPAR, HCT116 and HT29 human colon carcinoma cells [120, 198] [444,

445], were exposed to 21% O2 (normoxia), 0.2% O2 (hypoxia) or 0.0% O2 (anoxia)

for 24h after which RNA was collected. qPCR analysis confirmed that uPAR (Figure

S1A) mRNA was upregulated by approximately 2- and 5- fold in HCT116 and HT29,

respectively, in hypoxia and 15- 20-fold in anoxic conditions. These qPCR primers

detected all uPAR isoforms. Using isoform-specific primers, we found that mRNA of

uPAR splice variant 2, encoding a GPI-anchor deficient, soluble isoform of uPAR

(suPAR), was upregulated similarly to uPAR variant 1 mRNA (Figure S1B).

We also wanted to confirm that suPAR protein secretion was induced during

hypoxia. However, we were not able to detect endogenous uPAR/suPAR protein by

Western blotting using commercially available antibodies.

 

 

51  

To facilitate other assays requiring Western blotting, including validation of

immunoisolation and monitoring of alterations in gel electrophoretic mobility with and

without protein reduction, we decided to create a cellular model overexpressing

suPAR tagged with both Myc and DDK on its C terminus. Thus, 3 cell lines

(HCT116, HT29 and ME180) were stably transfected to overexpress suPAR tagged

with both a polypeptide from Myc and DDK. ME180 human cervical caricinoma cells

were included since they are used in our lab in an orthotopic cervical xenograft

model of metastasis that is a part of this project’s future direction. We validated

suPAR overexpression at the mRNA (Figure 4.1A-C) and protein levels (Figure

4.1D-F) in pools of cells as well as in isolated clones. Although the pool of stably

transfected HCT116 cells showed approximately 7-fold overexpression of suPAR

(Figure 4.1A), select clones exhibited greater overexpression. In HCT116, 3 clones

were identified to overexpress suPAR mRNA over 10-fold (Figure 4.1A), while in

HT29 and ME180, one clone was identified in each to have 25- and 50-fold mRNA

overexpression, respectively (Figure 4.1B and 4.1C). Expression at the protein

level, as detected by immunoblotting for the myc tag demonstrated good agreement

with mRNA levels (Figure 4.1D-F). On the basis of mRNA and protein

overexpression, HCT116 clone 6, HT29 clone 12 and ME180 clone 12 were used in

the following experiments.

4.1.2 Optimizing Immunoisolation Techniques for suPAR

Since the pulse chase assay utilizes immunoisolation techniques to isolate a

protein of interest, we assessed the ability of antibodies against uPAR, and the Myc

and DDK tags to immunoisolate the overexpressed suPAR-myc-DDK.

 

 

52  

 

 

53  

Immunoisolation using the polyclonal antibody against uPAR provided superior

protein signal when subsequently detected with anti-myc or anti-DDK antibodies. We

therefore decided to use this for future experiments. Using a polyclonal antibody can

also be beneficial when trying to recognize folding intermediates where single

epitopes can be transiently buried. We also assessed the performance of numerous

combinations of lysis and wash buffers in order to optimize the immunoisolation

assay, deeming RIPA lysis buffer and a RIPA-based wash buffer to be best for

suPAR immunoisolation on grounds of most robust specific signal with the least

background signal (Figure 4.2A).

4.1.3 A Transient Transfection suPAR Overexpression Model

To monitor suPAR protein maturation, we needed to immunoisolate

radiolabelled protein to be able to identify the protein of interest in different structural

conformations with high sensitivity. We therefore labeled newly synthesized protein

in the stably transfected HCT116 cells using [35S] cysteine and [35S] methionine and

immunoisolate the overexpressed suPAR. Despite our efforts to improve

immunoisolation efficiency, radioactive signal strength was very low when assessing

stably transfected cells (Figure 4.2B). One alternative solution was to transiently

transfect cell lines to produce greater suPAR signal. We therefore radiolabelled

newly synthesized protein in transiently transfected, stably transfected and parental

HCT116 with a long (1 hour) [35S]-cysteine and -methionine pulse and

immunoisolated protein with antibodies against either uPAR or the Myc tag. The

signal strength when radiolabeling transiently transfected HCT116 cells was

 

 

54  

protein seen in transient versus stably transfected HCT116 as identified by Western

 

 

55  

markedly improved in comparison to the stable transfection model and parental cells

(Figure 4.2B), consistent with the intracellular abundance of suPAR blotting for the

Myc tag (Figure 4.2C). As previously described, the uPAR antibody precipitated

more protein than the myc antibody (Figure 4.2B). Based on the improved protein

signal, we proceeded using the transient transfection model for the assessment of

suPAR maturation by the pulse chase assay.

One concern with overexpressing a protein of interest is the possibility that

the cellular machinery may not be capable of adequately processing the increased

protein load, which may result in ER stress. We assessed ER stress in the

transiently transfected cells by monitoring the splicing of XBP1 mRNA by qPCR.

When the ER is stressed, IRE1 is activated and removes a 26-nucleotide intron of

ubiquitously expressed XBP1, resulting in a more potent transcription factor.

However, relative to untransfected cells, the overexpression of suPAR did not

increase the amount of spliced XBP1 mRNA as a fraction of total XBP1, while

parental HCT116 cells exposed to anoxia for 24 hours, a condition expected to

induced ER stress, exhibited a 5-fold induction of spliced XBP1 (Figure S2). Thus,

overexpressing suPAR tagged with Myc and DDK was inconsequential on splicing of

XBP1, indicating that it did not cause ER stress. Unaltered XBP1 splicing also

suggested that overexpressed and tagged suPAR was not misfolded or left

unfolded.

We also monitored the cellular localization of suPAR in transiently transfected

HCT116 cells by immunofluorescence. The staining pattern of suPAR-myc-DDK was

perinuclear and reticular, consistent with localization in the secretory pathway,

 

 

56  

(Figure S3). Finding no evidence of ER stress or protein mislocalization, we decided

to move forward with this model.

4.1.4 Characterizing suPAR Electrophoretic Mobility Under Reducing and Non-Reducing Conditions

Our general approach was to utilize changes in gel electrophoretic mobility to

monitor protein maturation. The introduction of disulfides into a protein’s structure

results in a more compact configuration and greater electrophoretic mobility in a

denaturing resolving gel. Though suPAR has many disulfides, most are formed

between closely situated cysteines and are expected to result in small changes in

the protein structure. The most distant cysteines forming disulfides and thus those

that are most likely to contribute to observable potential mobility change in suPAR

include Cys25-Cys46, Cys39-Cys67, Cys117-Cys144, Cys137-Cys169 and Cys216-

Cys244 (Figure 1.1). To investigate whether differential electrophoretic mobility can

be detected and hence used to evaluate disulfide formation in suPAR, migration of

radiolabelled immunoisolated suPAR was compared under reducing and non-

reducing conditions. When reduced with DTT, suPAR demonstrated a clearly

discernible decreased electrophoretic mobility in comparison to the non-reduced

protein (Figure 4.3A). Furthermore, the identity of the reduced and non-reduced

suPAR bands were confirmed by comparing to non-transfected samples (Figure

4.3A) and immunoblotting for both DDK and Myc tags (data not shown). Some non-

specific bands were unfortunately also detected in non-transfected cells in close

vicinity to suPAR. Nevertheless, electrophoretic mobility difference could clearly be

used to monitor disulfide bond formation in suPAR maturation.

 

 

57  

 

 

58  

4.1.5 Optimizing Pulse Time for suPAR

Fundamental to the pulse chase assay is radiolabeling of a de novo protein

population to monitor throughout the assay. Shorter pulses have the benefit of

creating a more synchronized protein population   [436], but is associated with lower

signal strength. To determine the duration of radiolabeling necessary to detect

immunoisolated suPAR by phosphor imaging, transiently transfected cells were

exposed to pulse media for 5-60 min and chased for 0 or 6 hours.

As expected, increasing the duration of radiolabelling increased the intensity

of suPAR signal, with no signal visible after 5 minutes of pulse increasing up to a

robust signal after 1 hour (Figure 4.3B). It was determined that a 1 hour pulse

duration was required to produce ample suPAR signal (Figure 4.3B). After 6 hour of

chase, intracellular suPAR signal was noticeably less than the amount found

immediately following 1 hour of radiolabeling, suggesting that some suPAR was

secreted within 6 hours. A minor increase in electrophoretic mobility is also seen in

suPAR in reducing conditions after 6 hours of maturation, the significance of which

will be discussed below.

In conclusion, [35S] cysteine and [35S] methionine labeling of transiently

transfected HCT116 for 1 hour produced a detectable population of suPAR for which

maturation steps could be monitored.

4.1.6 Exploring the Possible Effect of Remnant DTT on Disulfide Bond Formation After a Reductive Challenge

Since a pulse of 1 hour was required to generate sufficient suPAR signal to

assess maturation, the newly synthesized and radiolabelled protein were at different

 

 

59  

stages of maturation at the end of the labeling term. In fact, as shown in Figure 4.3A,

most of the protein in the population already contained disulfide bonds immediately

after the pulse, as suPAR exhibited differential electrophoretic mobility under

reducing and non-reducing conditions. To synchronize the protein population after

radiolabelling, one strategy is to reduce (partially) oxidized proteins by incubating

cells with DTT. Despite washing cell culture dishes with chase media 3 times

following DTT reduction in vivo, remnant DTT may be sufficient to inhibit disulfide

bond formation. To address this possibility, we assessed disulfide formation of

albumin in HepG2 cells, which represents a well-characterized model in which an

endogenously expressed protein gives ample signal intensity [381] (Koritzinsky et

al., in review). As such, HepG2 cells were radiolabeled for 3 minutes in normoxia, in

vivo reduced with 5mM DTT for 5 minutes and chased in media containing

increasing concentrations of DTT up to 3mM. Albumin chased for 15 minutes in

media without DTT was oxidized (Figure 4.4), evidenced by a more rapidly migrating

protein population (confirmed to confer disulfide bonds by in vitro reduction in other

experiments, data not shown). Albumin matured in chase media containing up to

1.25mM DTT could not be distinguished from albumin matured in DTT-free media

(Figure 4.4), suggesting that cells remain competent of oxidative folding even at

these relatively high DTT concentrations. In comparison, if it were estimated that

10% of 5mM DTT reducing chase media remained following wash and aspiration,

after 3 washes the concentration of DTT would be 0.005mM. Hence, it is unlikely

that remnant DTT from in vivo reduction following radiolabeling inhibits the formation

of disulfide bonds in the chase period.

 

 

60  

 

 

61  

4.1.7 In Vivo Reduction of suPAR Following Radiolabelling

DTT, at a concentration of 5mM, was previously shown to reduce (partially)

oxidized albumin [376] (Koritzinsky et al., in review), and we wanted to assess if this

was also sufficient for suPAR. Exposing cells to 5mM-50mM of DTT for 5 minutes

reduced the majority of radiolabelled suPAR, as evidenced by the appearance of a

sharply migrating band in the non-reduced samples (Figure 4.5A, lanes 10-14).

Without DTT reduction, non-reduced suPAR was hard to detect (Figure 4.5A, lane

9), presumably because it forms a smear due to multiple disulfide-linked

conformations. A difference in the migration of reduced suPAR in cells treated with

DTT was also evident (Figure 4.5A, lane 2 versus 3). Likely, this is a result of adding

the alkylating agent NEM to the cells at the end pulse chase. NEM binds free

sulfhydryls and thereby prevents further oxidation. Consequently, more NEM binds a

reduced protein. Although NEM is small (125 Da), this can result in differences in

electrophoretic mobility of sulfhydryl-rich proteins. Increased electrophoretic mobility

in samples from cells where no DTT was added (Figure 4.5A, lane 2) is consistent

with less NEM binding due to the existence of disulfide bonds.

Next, we wanted to assess if introducing a reductive challenge affected the

maturation kinetics of suPAR. To that end, suPAR was reduced in vivo with 10mM

DTT for 5 minutes after radiolabeling (or not) and subsequently chased for 6 hours.

At this time point, suPAR migrated as a smear below the non-specific band under

non-reducing conditions regardless of whether the cells had been subjected to a

reductive challenge (Figure 4.5B, lanes 6 and 7). This smear could be collapsed to a

 

 

62  

 

 

63  

sharply migrating band upon reduction in vitro (Figure 4.5B, lanes 2 and 3),

demonstrating that the smear is a consequence of disulfide bond formation.

Disulfide bond formation had hence occurred over 6 hours regardless of the

reductive challenge, producing similarly migrating protein populations. The

electrophoretic mobility of suPAR also increased after 6 hours of chase when

resolved under reducing conditions (Figure 4.5B, lane 2 and 3 versus 1). The source

of this increase could be two-fold. It could reflect differential NEM binding as

explained before, due to the introduction of disulfide bonds, and/or the trimming of

N-linked glycans. We could not differentiate between these possibilities in this

experiment. However, the increase in electrophoretic mobility was clearly larger if

the cells had not been subjected to a reductive challenge (Figure 4.5B, lane 3

versus 2). This result therefore indicates that glycan trimming or disulfide bond

formation in suPAR occurs more slowly following a DTT challenge. When

immunoisolating suPAR from the growth media, increased extracellular expression

(Figure 4.5B) was evident after 6 hours chase, suggesting suPAR secretion. Though

intracellular suPAR levels seem similar with and without DTT reduction, extracellular

levels of suPAR were higher without DTT reduction (Figure 4.5B). It is possible that

a portion of suPAR could be degraded intra-cellularly if the cells had been subjected

to DTT.

The alterations in maturation and secretion kinetics of suPAR inflicted by a

reductive challenge represented a source of concern. Nonetheless, the requirement

for long labeling times required this approach. We therefore decide to proceed,

 

 

64  

using 5mM DTT as a reductive challenge and keeping this possible experimental

limitation in mind.

In summary, a transient transfection model overexpressing suPAR tagged

with Myc and DDK was established. The pulse chase assay was optimized to

assess suPAR maturation, involving 1 hour radioactive labeling followed by a

reductive challenge with 5mM DTT. This approach could be used to study disulfide

bond formation in suPAR, with the caveat that maturation and secretion was slower

than under conditions where redox balance was unperturbed.

B) To characterize the maturation and secretion of suPAR

4.1.8 suPAR Disulfide Formation and Secretion in Normoxia

To assess the maturation and secretion of suPAR in anoxic conditions, it was

necessary to first be able to characterize its maturation and secretion at regular

oxygenation. To assess the kinetics of disulfide bond formation in suPAR, we used

the transient transfection model and pulse assay developed specifically for suPAR in

HCT116 cells, as in Figures 4.1-4.5. In Figure 4.6 (images representative of multiple

separate experiments), transiently transfected HCT116 cells were radiolabelled for 1

hour, in vivo reduced with 5mM of DTT for 5 minutes and matured for 0, 2 and 6

hours in DTT-free media in normoxia. Radiolabelled suPAR produced in the pulse

phase was found to decrease within the cell through the chase duration (Figure

4.6A), correlating with suPAR observed in chase media after 2 and 6 hours (Figure

4.6B) and indicating that suPAR was being secreted within these timepoints.

Furthermore, intracellular suPAR observed at both 2 and 6 hours exhibited a loss of

 

 

65  

 

 

66  

signal in non-reducing conditions in comparison to 0 hours (Figure 4.6A). Given the

presence of the bands in the corresponding reduced conditions, this indicates that

the protein was indeed present and immunoisolated. Therefore this loss of signal is

due to disulfide bond formation resulting in many conformations and producing a

smear that is too diffuse to be lucidly observed. The data thus demonstrate that

suPAR had formed disulfide bonds by 2hours in normoxia, after which no further

modifications could be observed.

   

4.1.9 suPAR Glycosylation in Normoxia

In addition to disulfide bond formation, enzymes in the ER modify proteins by

N-linked glycosylation and suPAR has been previously described to have 4 N-linked

glycans (Figure 1) [173, 446, 447]. So to further characterize suPAR maturation,

glycan processing in normoxic conditions was assessed utilizing endoglucosidase H

(endoH) to cleave N-linked glycans off of the protein strand, producing a smaller,

more mobile protein. Incomplete endoH digestion of suPAR produced 4 distinct

species with increased electrophoretic mobility in comparison to untreated protein

(Figure 4.6C). This indicated that endoH had cleaved between 1 and 4 N-linked

glycans to yield these distinct bands, consistent with previous reports that suPAR

contains 4 N-linked glycans [173, 446, 447].

Following N-linked glycosylation in the ER and translocation to the Golgi,

glycans undergo trimming by Golgi mannosidase I and II and glycosylation by

GlcNAc transferase [393, 414]. These modifications result in a common core

oligosaccharide, which is then modified to produce a variety of complex glycans that

 

 

67  

confer resistance to endoH cleavage. However, none of the intracellular suPAR was

resistant to endoH digestion at any timepoints (Figure 4.6C), providing no evidence

of complex glycosylation. To investigate the possibility that suPAR was secreted

without first being complex glycosylated, suPAR immunoisolated from chase media

was treated with EndoH. Secreted suPAR, harvested after 6 hours of chase, was

also found to be endoH sensitive (Figure 4.6D), demonstrating that the N-linked

glycans were not complex glycosylated in the Golgi prior to secretion. This

observation contrasts the complex glycans of suPAR observed in Chinese hamster

ovary cells stably expressing suPAR [448, 449]. Although lack of complex

glycosylation is not uncommon and could merely reflect no access for the Golgi-

localized enzymes, we cannot rule out a non-canonical route of secretion for suPAR

that bypasses the Golgi apparatus. In summary, our results suggest that suPAR’s 4

N-linked glycans do not undergo complex glycosylation in these cells, and that

disulfide bonds are (re-) formed within 2 hours of in vivo reduction with DTT,

followed by secretion between 2 and 6 hours.

4.2 Aim 2: To Characterize suPAR Maturation in Anoxic Conditions  A) To validate the pulse chase assay for use in anoxia

The pulse chase assay has long been used for the investigation of protein

folding and maturation in vivo [436]. However, the technical details of the assay’s

use in low oxygenated conditions have yet to be validated. Specifically we wanted to

rule out any unexpected influence of glass petri dishes (necessary due to the

oxygen-content in plastic) and evaluate the use of a PERK inhibitor to allow

 

 

68  

translation in anoxic conditions. Albumin was used as ER cargo in the following

validation because it is highly expressed endogenously by liver cells such as

HepG2, with known folding kinetics (Koritzinsky et al., in review) [381].

4.2.1 Investigating the Influence of Glass Culture Dishes on Protein Maturation

To address if glass petri dishes were comparable to plasticware with regards

to protein maturation and cell attachment during multiple manipulations, HepG2 cells

plated on glass or plastic petri dishes were radiolabeled for 3 minutes in normoxia.

As previously described [381], disulfide bonds formed rapidly in albumin during the

pulse, evidenced by increased gel electrophoretic mobility immediately after the

pulse in non-reduced albumin in comparison to samples run under reducing

conditions (Figure 4.7). No difference could be discerned between plastic and glass

dishes, suggesting that disulfide formation in albumin was unaffected by the

difference in dish material. There were also no problems with these cells remaining

attached on the glass during the manipulations. Thus, glass dishware could

substitute for plastic petri dishes to facilitate the assessment of protein maturation in

anoxia.

4.2.2 Protein Maturation with PERK Inhibitor

A limitation of in vivo reducing protein before maturation in anoxia is the

incapacity to observe co-translational disulfide formation. Also, the ER cargo is

removed from it’s normal route of maturation, and we did observe differences in

maturation kinetics conferred by the reductive challenge. To overcome translational

 

 

69  

 

 

70  

inhibition in anoxia and facilitate the study of both co-translational and post-

translational disulfide formation in anoxia, cells were treated with a PERK inhibitor

[450], which competes for ATP binding in PERK [451]. It was expected that the

PERK inhibitor would prevent phosphorylation of EIF2α and the subsequent

inhibition of global mRNA translation in response to hypoxia, thus enabling

radiolabeling and protein synthesis. To assess translation and maturation in anoxia

with the PERK inhibitor, HepG2 cells were pretreated with PERK inhibitor for 1 hour,

exposed to anoxia for 1 hour and radiolabeled for 3 minutes. One hour of exposure

to anoxia markedly reduced the total amount of radiolabelled protein (Figure 4.8A),

indicating that anoxia severely decreased the overall translational ability of HepG2

cells. In cells treated with PERK inhibitor, the amount of radiolabeled protein in

anoxia was similar to the amount radiolabeled in normoxia (Figure 4.8A). This

showed that translational inhibition was prevented by treating cells with PERK

inhibitor, allowing for translation in anoxia. However, albumin translated and matured

in anoxia in cells treated with PERK inhibitor formed aggregates that were resolved

in a reduced gel (Figure 4.8B), suggesting they were disulfide dependent. In the

region of albumin monomers, there was a faint smear suggesting some oxidation of

albumin under anoxia in the absence of the PERK inhibitor, but less oxidation when

PERK was inhibited. One possible explanation for these observations is that some

disulfides are formed co-translationally if the ER protein load is low, but that the ER

folding machinery cannot tolerate high burden in the absence of oxygen, resulting in

protein aggregation. These results indicated that even though the PERK inhibitor

facilitated translation in anoxia, its use may result in non-physiological protein load

 

 

71  

 

 

72  

that renders experimental results less relevant. For the purpose of this thesis, we

therefore chose to limit ourselves to studying post-translational disulfide bond

formation of suPAR in anoxia after a DTT challenge, using glass dishes to prevent

plastic from supplying oxygen and without the use of PERK inhibitor.

B) To characterize the oxygen dependency of suPAR maturation

4.2.3 suPAR Glycosylation in Anoxia

To characterize suPAR maturation in anoxia, the oxygen dependency of

suPAR glycosylation was first assessed. suPAR radiolabeled for 1 hour and chased

for 0 and 6 hours in anoxia, was treated with endoH and exhibited a shift in

electrophoretic mobility with EndoH digestion, akin to suPAR matured in normoxia

(Figure 4.9). It is suggested that suPAR matured in anoxia undergo N-linked

glycosylation processing comparable to protein matured in normoxic conditions. As

in normoxia, suPAR matured in anoxia remained sensitive to endoH digestion,

providing no evidence of complex glycosylation nor Golgi processing. This is

consistent with our group’s previous observations with Flu-HA and alpha-1-

antitrypsin (Koritzinsky et al., in review), demonstrating that glycosylation and glycan

processing is independent of oxygen.

Interestingly, after 6 hours of maturation in anoxia, a slightly higher molecular

weight suPAR species (denoted with an asterisk) was consistently observed when

resolved in reducing conditions (Figure 4.9 and 4.10A). Glycosylation was

investigated as the potential culprit for the observed electrophoretic mobility

difference as proteins have been described to undergo hyperglycosylation in

 

 

73  

 

 

74  

circumstances where supplementary stability is required [452]. Complete EndoH

digestion of suPAR matured in anoxia did not resolve the larger suPAR species,

suggesting that this differential species is not a product of additional N-linked

glycosylation (Figure 4.9). The identity of the species remains to be found.

Therefore, it was concluded that the glycan modifications of suPAR in anoxia were

largely analogous to that of suPAR matured in normoxia.

4.2.4 suPAR Oxidative Folding in Anoxia

After deciding on an approach for the pulse chase assay in anoxic conditions,

suPAR maturation in anoxia was assessed. HCT116 cells transiently transfected to

overexpress suPAR were radiolabelled for 1 hour in normoxia, reduced with 5mM of

DTT for 5 minutes (as in Aim 1) and brought into the anoxic chamber. The cells were

washed and incubated in deoxygenated DTT-free media, and hence permitted to

mature in anoxia. After 2 hours of maturation in normoxia, suPAR exhibits slightly

increased electrophoretic mobility when resolved under reducing conditions (Figure

4.10A) that is likely attributed to glycan trimming or disulfide bond formation

(conferred by NEM binding), as previously proposed. Interestingly, this increased

electrophoretic mobility is not seen after 2 hours in suPAR matured in anoxia,

though is observable after 6 hours, suggesting delayed processing in anoxia.

Following 2 and 6 hours of exposure to anoxia, suPAR was observed with

unchanged electrophoretic mobility in a non-reducing gel when compared to

reduced samples (Figure 4.10A), suggesting less disulfide bond formation in suPAR

in anoxia (Figure 10A representative of 5 separate experiments). This contrasts the

shift in electrophoretic mobility (or essentially loss of signal) observed in suPAR after

 

 

75  

 

 

76  

2 and 6 hours of maturation in normoxia (Figure 4.3B, 4.5B, 4.6A and 4.10A),

indicating disulfide bond formation in regular oxygenation. Despite being less

capable of forming disulfide bonds in anoxia, intracellular suPAR decreased by 2

and 6 hours in oxygen-limited conditions (Figure 4.10A), suggesting either

degradation or secretion in anoxia. Secretion of suPAR in anoxia was supported by

a corresponding increase in suPAR immunoisolated from growth media (Figure

4.10B) after 6 hours. Interestingly, these preliminary data showed that suPAR

secreted in anoxia exhibited no change in electrophoretic mobility when compared to

reduced protein (Figure 4.10B). The results are therefore consistent with suPAR

being secreted under anoxia with slower kinetics than in normoxia, and with either

less or different disulfide bonds. This suggests that suPAR secretion is not strictly

dependent on disulfide formation.

4.2.5 suPAR Secretion in Normoxic and Anoxic Conditions

High molecular weight aggregates are also observed in the immunoisolated

media resolved under non-reducing conditions (Figure 4.10B). Since these

aggregates are resolved in the reduced gel, they are disulfide-linked. This

aggregation complicates quantification of oxidized secreted suPAR as much of the

protein remains in aggregates, where other proteins may contribute to the signal.

We therefore quantified the secretion of suPAR in normoxic and anoxic conditions,

from multiple experiments where suPAR was resolved under reducing conditions.

Mean intracellular suPAR levels decreased to approximately 55% of the original

protein levels in normoxia and 60% in anoxia after 6 hours of chase (Figure 4.11A),

 

 

77  

 

 

78  

however the differences in decreased intracellular suPAR in normoxic and anoxic

conditions at 2 and 6 hours was found to be non-significant. In conjunction, the

amount of suPAR protein immunoisolated from growth media after 2 hours of chase

in anoxia was slightly less than normoxia (Figure 4.11B). After 6 hours of chase,

suPAR extracellular expression in anoxia was less than 70% than in normoxia,

however this difference was also not statistically significant (p=0.18) (Figure 4.11B).

Thus, the secretion of suPAR in both normoxic and anoxic conditions were

consistently observed over multiple experiments suggesting efficient secretion of

suPAR in anoxic conditions.

In characterizing the oxygen dependency of suPAR maturation, intracellular

disulfide formation in suPAR was impaired in anoxia, while glycosylation proceeded

with no discernable difference to maturation in normal oxygenation. Nonetheless,

suPAR secretion in anoxia was still consistently observed, suggesting that the

efficient secretion of suPAR in anoxia may not be dependent on disulfide formation.

 

 

79  

5 DISCUSSION

This project served to investigate the potential selective maturation capacity

in a key cancer-relevant, hypoxia-regulated protein, uPAR in the absence of oxygen.

To that end we created and characterized a model of suPAR overexpression. Within

this thesis, an assay was established and validated for investigating protein

maturation kinetics in anoxia. We then utilized these tools to study the glycosylation,

disulfide formation and secretion of suPAR in oxic and oxygen-free conditions. The

methods developed here are relevant for further studies in other cargo proteins and

hypoxia-related diseases.

5.1 Technical Limitations

5.1.1 Transfection Model  

To evaluate if suPAR possesses advantageously forms disulfide bonds and

folds in environments devoid of oxygen, transient transfection techniques were

employed to create cellular models overexpressing suPAR tagged with Myc and

DDK. The model serves to afford detection of suPAR, as endogenous protein levels

were insufficient for detection by immunoblotting using commercially available

antibodies. Transient transfection introduces numerous copies of cDNA in the cell

nucleus to produce substantial overexpression. One concern is that transfection also

potentially results in physical and biological stresses on the cell that could affect

viability and function, however no toxicity was observed with transfection in these

experiments. Of particularly high relevance for this thesis is the possibility that cells

have a certain capacity to fold proteins and that overexpression therefore, may

 

 

80  

result in non-physiological protein overload and aggregation [453]. Indeed, high

molecular weight aggregates were observed in non-reducing gels, indicating the

existence of disulfide linked-aggregates. However, suPAR overexpression did not

significantly increase XBP1 splicing, indicating no induction of the UPR, and thus

contesting the presence of ER stress. It is therefore possible that these high-

molecular weight species could represent intermediates in a normal folding pathway

rather than aggregation due to protein overload. Nevertheless, it remains a

consideration that transient overexpression of suPAR may exceed the cell’s folding

capacity and obscure the surveillance of suPAR protein folding.

5.1.2 ER Cargo Tags  

The suPAR protein was tagged with C-terminal polypeptide DDK and Myc

tags that may pose a liability to proper and efficient protein folding. Since these tags

are very small in size, it is unlikely that the tags physically obstruct folding.

Furthermore, both tags are present on the C-terminal end, are translated last, and

thus are less likely to disrupt folding that occurs co-translationally. In addition,

neither the DDK or Myc tags contain cysteines, which could inadvertently form an

intermediate or unresolvable disulfide bond with other cysteines within the suPAR

sequence. In support of this, and as mentioned above, unchanged XBP1 splicing

following transient transfection (Figure S2) indicates a lack of UPR induction and

thus absence of significant misfolding. Furthermore, since the DDK- and Myc-tagged

suPAR does get secreted, this also suggests that it is not misfolded. This evidence

supports the validity of using Myc and DDK tagged suPAR to monitor protein folding.

 

 

81  

Nonetheless, the possibility of these tags affecting folding kinetics cannot be

excluded without direct comparison to untagged protein.

5.1.3 The Influence of DTT on Protein Maturation  

Despite using the lowest effective concentration of DTT in the in vivo

reduction of protein immediately following radiolabelling (Figure 4.5A), treatment of

cells with a potent reducing agent may have widespread consequences for cellular

function and environment. However, in the experiments within this thesis, no cell

death was observed on a gross level. Additionally, proteins were capable of

refolding following a reductive challenge, indicating reversibility of the reduction.

Nonetheless, 5mM of DTT was necessary to reduce the protein population following

radiolabelling as lower concentrations were found to be insufficient to keep reduced

protein from refolding (Figure 4.4). Furthermore, literature has shown that although

DTT reduces disulfides in proteins that have yet to reach their native configuration, it

does not extensively interfere with other cellular functions, including translocation,

signal sequence removal, N-linked glycosylation and protein transport within the

secretory pathway. Evidence for this comes from experiments whereby proteins

without disulfides manage to be synthesized, matured and secreted with similar

kinetics and fidelity as in DTT-free conditions [454-456]. Also, disulfide-containing

proteins, like Flu-HA, refolded rapidly and accurately after DTT challenge despite the

fact that completely reduced Flu-HA is not a normal intermediate in the ER [308]. In

vesicular stomatitis virus G, translation in DTT and subsequent wash out resulted in

non-native interchain crosslinks that delayed normal folding, but proper folding was

 

 

82  

nonetheless eventually achieved [457]. Extended incubations (>30min) in DTT

resulted in only partial refolding efficiency [308], however this duration greatly

exceeds the 5 minute incubation utilized in the protocol developed for suPAR. This

experimental evidence show that many cargo proteins are capable of forming

disulfide bonds and folding following in vivo reduction with DTT, and thus support

the use of such refolding experiments to monitor intracellular folding events.

Interestingly, proteins that have reached their native structures by the time

DTT is added to the cells are resistant to DTT reduction and are secreted

nonetheless [455], as over 70% of disulfides are solvent inaccessible in the native

protein structure [458, 459]. This must generally be considered when employing

longer pulse durations as proteins synthesized early in the timeframe may have

reached a conformation in which its disulfides are not available for DTT reduction,

cannot be reduced and hence complicate the pursuit of a synchronized protein

population following radiolabelling. However, in these experiments, DTT-resistant

suPAR was not observed after a 1 hour pulse, and hence was not a major factor

here.

An additional concern exists regarding whether remnant DTT could affect

folding in the chase, even after multiple washes. Experiments within this thesis show

that remnant DTT following reductive challenge is unlikely to interfere with disulfide

bond formation in DTT-free chase media due to the competence for disulfide bond

formation even at relatively high DTT concentrations (Figure 4.4). The observed

rapid refolding of Flu-HA following washout [308] suggests that the cells metabolize

or otherwise remove any residual reducing agent rapidly upon washout.

 

 

83  

Appenzeller-Herzog and colleagues also showed that disulfide bonds are formed

within seconds following the washout of DTT [355]. These data suggest that

refolding after a DTT challenge can provide a useful model for studying post-

translational maturation events.

5.2 suPAR Maturation

5.2.1 Folding Co- and Post-Translationally  

With regards to the modified pulse chase assay specific for the evaluation of

suPAR maturation, 1 hour of radiolabelling was deemed necessary to produce

sufficient signal for assessment (Figure 4.3B). However, a long pulse duration also

necessitates a DTT challenge to reduce all co-translational disulfide bonds to

synchronize the oxidative state of the protein population. This represents a major

limitation of the experimental approach, because the ER cargo is forced to fold

completely post-translationally. Disulfide bonds form in cargo protein during and

shortly after translocation into the ER lumen [460], facilitating co-translational folding

that may very well be a manifestation in the maturation of all proteins [461]. And as

previously reviewed in the introduction, many factors influence co-translational

protein folding, and inhibiting co-translational disulfide formation may alter the

folding pathway of the protein as suggested by McGinnes and Morrison (1996)

[462], by possibly impeding vectorial, domain-by-domain protein folding [463].

Furthermore, even non-native disulfides that form co-translationally and need to be

isomerized for correct folding can support productive folding by stabilizing

intermediates, as in bovine pancreatic trypsin inhibitor [464] and LDLR [465].

 

 

84  

Another potential issue that may arise from DTT reduction refolding

experiments and potential differences in co- and post-translational folding is the

possibility that N-linked glycans can interfere with post-translational disulfide

formation. Glycan modification and disulfide bond formation participate in a temporal

relationship in which glycans directly influence protein conformation, and disulfide

formation and folding can influence occupancy of N-linked glycan sites (sequons)

[306, 466, 467]. Thus, a nascent protein is modified simultaneously by N-linked

glycosylation and disulfide bond formation, and following reduction of existing

disulfides, the attached bulky N-linked glycans may interfere with the regular

disulfide formation and protein folding kinetics. Though no current evidence

suggests that N-linked glycans may be disrupting post-translational disulfide

formation within the investigated long chase timeframes, we cannot rule out the

possibility of interference given that suPAR has 4 N-linked glycans embedded

between 14 disulfides (Figure 1.1).

In addition to issues potentially arising from preventing the involvement of co-

translational disulfide bond formation on vectorial folding, there are general concerns

with removing cargo from its normal folding pathway. In test tube refolding

experiments, folding may halt at intermediate states or diverge to off pathway

reactions resulting in misfolding and aggregation when not subject to cellular folding

factors [468]. Such evidence demonstrates the importance of the ER folding

machinery. It is a possibility that some folding factors may be exclusively or

preferentially available co-translationally, in which case their influence would be lost

in refolding experiments. In support of this, folding factors have been observed to

 

 

85  

bind nascent clients. For example, BiP plays a role in the translocon complex by

regulating pore gating [469] and protein import into the ER lumen during translation.

Hence, BiP may also be able to act to rapidly inhibit aggregation co-translationally.

Furthermore, calnexin binds hemagglutinin [463] and HSP47 binds procollagen [470]

co-translationally. PDI has also been shown to associate with nascent protein [471,

472] and has been suggested to be essential for co-translational disulfide formation

[473]. Though these observations do not suggest that these folding factors are not

equally involved in post-translational folding, such a possibility cannot be overlooked

in differentiating between protein folding during and following translation.

In spite of these concerns, even ER cargo that folds mainly co-translationally

such as Flu-HA can be successfully refolded after a reductive challenge. The

importance of the co-translational folding phase is likely highly cargo-specific. Since

we, due to experimental limitations, have not been able to study co- versus post-

translational folding in suPAR, we cannot estimate to what degree this issue may be

relevant to suPAR. We must in consequence limit our interpretations of the data to

apply to conditions where suPAR is (re)folded post-translationally, potentially

removed from its normal folding pathway.

5.2.2 Oxygen Dependency of Disulfide Bond Formation

During the work of this thesis, it was in fact discovered that the effect of

oxygen on disulfide bond formation differed as a function of time after protein

synthesis. Using Flu-HA, LDLR and albumin as models, it became clear that

disulfide bonds could be introduced into ER cargo very early in its lifetime in the

 

 

86  

absence of oxygen. This contrasted a later phase of disulfide bond formation, which

was completely oxygen-dependent (Koritzinsky et al., in review). The oxygen-

independent phase of disulfide bond formation coincided with cargo translation, but

was not firmly established to be functionally related to translation or association with

the translocon. Nevertheless, in the absence of oxygen, no post-translational

disulfide bond formation or isomerization was ever observed. This observation

extended to the situation of refolding post-translationally after a reductive challenge,

which was only productive in oxygen replete conditions. The data in this thesis are

consistent with these findings, in that we also did not observe intracellular disulfide

bond formation in suPAR after DTT reduction in anoxia. Furthermore, preliminary

data suggesting that this non-disulfide linked species might pass ER quality control

remains interesting. However, under physiological conditions it is possible that

suPAR is able to fold quickly (co-translationally) under anoxia. In consequence,

superiority for some ER cargo to mature in the absence of oxygen may be

associated with their ability to complete disulfide bond formation quickly, and without

the need for isomerization. These data and considerations highlight the importance

of monitoring the effects of oxygen on protein folding both co- and post-

translationally.

Radioactive labeling could not take place under anoxia in HepG2 cells due to

PERK-dependent inhibition of translation and a PERK inhibitor resulting in protein

aggregation (Figure 4.8A). However, this may not be an equally disrupting issue in

other cell types. Many cell types reduce translation to ~50% of basal levels during

anoxia [236], which would render pulse-chase experiments feasible. However,

 

 

87  

conclusions would be limited to cell types with less severe PERK activation, which

may reflect their overall lower secretory load. Nonetheless, the secretory nature and

PERK activation of different cell types may influence oxygen-independent disulfide

formation and should be considered in experimental development and interpretation.

5.2.3 Abundance of Extracellular suPAR in Anoxia  

It seems that appreciable variability exists between experiments in the

amount of secreted suPAR that is immunoisolated from media. One conceivable

factor is the possibility that following secretion, suPAR may bind to membrane-

bound interactors such as integrins or vitronectin, which may effectively decrease

the amount of soluble suPAR in growth media. Additionally, literature has noted that

suPAR is prone to aggregation without uPA binding [189]. Though it is not

characterized to what extent this aggregation occurs, uPA availability may play a

role in varied suPAR signal as the lack of uPA may detract from protein signal

strength as a result of protein aggregation.

Furthermore, proteostasis describes the fate of a secreted protein in terms of

its folding, secretion, aggregation or degradation [474, 475]. And as described in

more detail below, suPAR may not experience advantageous disulfide formation in

oxygen-deficient conditions but is nonetheless secreted. It is possible that the

secretion of suPAR in anoxia is simply an endpoint of proteostasis in which suPAR

eventually finds a route to extracellular space as it is neither folding, aggregating or

degrading intracellularly. suPAR may present an advantageous ability to bypass

 

 

88  

retention by ER quality control mechanisms, facilitating its secretion without bona

fide disulfide bonds.

5.2.4 Secretion of Non-Native suPAR  

In characterizing the maturation of suPAR in oxygen-deficient conditions,

suPAR was less able to form bona-fide disulfide bonds (Figure 4.9A). Nonetheless,

suPAR was still secreted in anoxia(Figure 4.9B). It is possible that the efficient

secretion of suPAR in anoxia may not depend on completion of disulfide bond

formation, and opposes our hypothesis that suPAR secretion during anoxia relies on

advantageous disulfide formation in the absence of oxygen. Instead, the

extracellular expression of suPAR may be attributed to an ability to bypass the strict

ER quality control system, conferring an advantage to extracellular expression even

without attaining its native conformation. The stringency of ER quality control and

retention seems be cargo-dependent. For example, LDLR [476] and CFTR [404,

477, 478] can escape to the Golgi in non-native conformations, suggesting that

quality control must depend on factors beyond protein folding. Though cargo

subjected to less stringent quality control would have to retain function or fold later in

its lifetime, the superior extracellular expression of select proteins, such as suPAR,

in hypoxia may very well lie in escaping quality control. In conjunction with this idea,

Ilani and colleagues have recently demonstrated an extracellular role for secreted

QSOX [378], which may potentially facilitate post-ER disulfide bond formation and

thus the achievement of native protein following secretion. It is possible that suPAR

 

 

89  

secreted in oxygen-lacking conditions may not have bona fide disulfide bonds but

may introduce disulfides in the extracellular space.

Since non-native suPAR reaches the extracellular space in anoxia, it is

necessary to rule out the possibility that unfolded proteins are expelled out of a cell

by an unknown mechanism as opposed to being actively exported by an intact

system. One concern discussed by Zhang and colleagues is the possibility that

protein reaches the extracellular space not by secretion but due to cellular lysis

[479]. Though minimal cell death was observed during our experimentation, an

important future direction is to assess cellular viability to confirm that suPAR is not

expelled out of the cellular structure into the extracellular space, but that it is indeed

secreted in anoxia.

A key question is whether non-native suPAR reaching the extracellular space

is functional without bona fide disulfides. Some proteins have been described to use

changes in disulfide bond profiles to regulate function [480, 481]. A high priority

future aim is to assess if suPAR secreted in anoxia is, or becomes, functionally

active. Such information could be evaluated by collecting suPAR from growth media

in anoxia and assessing pro-uPA activation to uPA. Furthermore, the proteolytic

activity of uPAR can be assayed by Bodipy-tagged bovine serum albumin (BSA),

which becomes fluorescent upon proteolysis [482]. It is also important to investigate

if this secreted, non-native protein is capable of achieving a native conformation

post-secretion though this may be hindered by the formation of disulfide-linked

aggregates.

 

 

90  

5.2.5 Potential of Preferential Disulfide Formation of Other ER Cargo in Anoxia  

Although we did not observe suPAR to fold in anoxia, other cargo proteins

preferentially secreted in hypoxia may yet exhibit advantageous disulfide formation

without oxygen. The ER maturation machinery includes at least 19 PDI homologues,

a growing number of oxidases and numerous chaperones that co-operate in the

folding of ER cargo protein. Our group has shown that hypoxia results in ER stress,

which activates the UPR to induce folding factors. We have shown that a third of the

ERome is induced after 24 hours of exposure to anoxia in numerous cell lines

(Koritzinsky et al., in review), much of which are known or suggested to be involved

in disulfide bond formation. One possibility is that certain ER maturation machinery

are upregulated to support advantageous maturation in hypoxic conditions,

exhibiting specificity to certain clientele, and thus facilitating the superior expression

of certain cargo in hypoxic environments. It is also possible that the hypoxic-

induction of some folding factors may foster overall protein folding in hypoxia.

Although we did not observe advantageous folding of suPAR in oxygen-deficient

conditions, this does not exclude that disulfides formation is achievable without

oxygen in other cargo proteins.

5.2.6 Identification of a Higher Molecular Weight suPAR Species Matured in Anoxia  

When suPAR was matured in anoxia, a suPAR species of slightly higher

molecular weight appeared after 6 hours of maturation without oxygen, under

reducing conditions (Figure 4.9, 4.10B). Since the slower migrating species was

observed under reducing conditions, the species was not a product of differential

 

 

91  

disulfide bond formation. Furthermore, when treated with EndoH, the differential

species was not resolved (Figure 4.10B), suggesting that the electrophoretic mobility

variance was not due to a difference in N-linked glycosylation either. Another

suspect secretory protein modification is Golgi-localized O-linked glycosylation, in

which a sugar molecule is attached to oxygen of serine or threonine residues,

conferring additional protein stability and facilitating proper conformation [415].

However, O-linked glycans have yet to be described in the suPAR structure. It is

possible that in anoxic conditions, suPAR exists in a different conformation and may

present newly available serine or threonine residues for O-linked glycosylation. It

may thus be valuable to probe the possibility of differential O-linked glycosylation in

anoxia.

Given that the species occurs later in maturation and was not found to be

secreted into media, it would seem reasonable that the differential species may be a

consequence of ERAD. However, processing an ER cargo for degradation involves

the cleavage of a mannose residues from N-linked glycans by ER mannosidase I

(ERManI) [394, 395], and the electrophoretic mobility between the two suPAR

species was found to be independent of N-linked glycosylation. The identity of this

differential suPAR species in anoxia requires further investigation to better

characterize suPAR maturation in oxygen-deficient conditions.

           

 

 

92  

6 FUTURE DIRECTIONS

6.1 Alternative Pathways of suPAR Secretion  

When radiolabelled suPAR was matured for 0, 2 and 6 hours, intracellular

and extracellular suPAR did not exhibit resistance to EndoH treatment. Thus,

complex glycosylation of suPAR nor transit through Golgi could be confirmed. One

possibility may be that suPAR is secreted via a Golgi-independent, non-canonical

pathway.

After proteins reach the ERGIC, they are either forwarded to the Golgi,

returned to the ER or targeted to an endosomal compartment for Golgi-independent

secretion [483, 484]. Non-canonical pathways include direct transit across the

plasma membrane without a cell surface transporter, lysosomal secretion, budding

from the plasma membrane, exosomal secretion by multivesicular bodies (MVBs),

amphisome-mediated secretion, autolysosomal secretion and autophagosome

secretion [479, 485-487]. Interestingly, uPAR has been observed to localize in

lysosomal structures and colocalize with lysosomal enzymes [488], such as

cathespin D [489], though this associate may be a part of protein degradation as

opposed to secretion.

To further characterize suPAR maturation and investigate whether suPAR

transits the Golgi en route to the extracellular space, preliminary work using fungal

metabolite brefeldin A (BFA) are underway. BFA targets GTP exchange factors

(GEFs) to inhibit GTPase ARF1p [490-492] and ensuing retrograde v-SNAREs, thus

limiting recruitment of retrograde COPI vesicles to the ER membrane and retrograde

transport from Golgi to ER [493, 494]. Furthermore, BFA restricts ER cargo protein

from entering the Golgi apparatus by causing the Golgi cisternae to fuse with the ER

 

 

93  

[495, 496] and thus preventing the binding of anterograde COPII vesicles to the

Golgi without directly inhibiting COPII vesicles themselves [497]. We have therefore

started optimizing a protocol for BFA treatment of HCT116 cells that disrupts Golgi

transit without causing substantial toxicity (Figure S4 and S5). HCT116 cells were

treated with 3.5, 5 and 10uM of BFA for 0, 2 and 6 hours. The cells did not appear to

show any gross cellular indications of stress following exposure to 5uM of BFA over

a 6-hour duration (Figure S4). To investigate if suPAR maturation and secretion is

independent of the Golgi, a co-transient transfection model was employed to

compare suPAR and albumin (known to follow canonical secretion through the Golgi

[498]) secretion with BFA treatment. HCT116 were radiolabelled and matured in

5uM of BFA. Preliminary work has shown that though suPAR decreased

intracellularly after 2 hours of maturation with BFA (Figure S5A), suggesting its

secretion, albumin was also found to decrease intracellularly and increase

extracellularly with BFA, though not until the 4-hour chase time point (Figure S5C

and S5D). This may be explained by the metabolism of BFA by 4 hours in culture

[499]. Given suPAR extracellular expression with BFA treatment at the 2 hour time

point at which time albumin had yet to be secreted, this may suggest suPAR

secretion to be independent of Golgi apparatus, though more work is needed to

support this possibility.

To further explore the possibility of canonical secretion, preliminary work has

begun employing sucrose gradient ultracentrifugation techniques to separate

subcellular organelles and identify suPAR’s localization in reference to secretory

structures and machinery in normoxic and hypoxic environments (data not shown).

 

 

94  

Additionally, co-localization experiments can be utilized to better elucidate suPAR

cellular localization relative to secretory structures. Both sucrose ultracentrifugation

and co-localization techniques can be utilized to probe unconventional secretory

pathways including lysosomal, exosomal, amphisomal and autophagosomal

secretion. Characterizing suPAR’s journey along the secretory pathway, whether

canonical or non-canonical will provide fundamental knowledge on the maturation of

a biologically and clinically relevant protein, and may even potentially expose new

prospective targets for anti-suPAR based intervention. Non-canonical secretion may

even play a role in the ability of suPAR to reach the extracellular space in anoxia

without all of its native disulfides.

6.2 Further Characterizing suPAR Maturation  

In the assessment of suPAR maturation, the pulse chase assay employed

was limited to a 6 hour time point after which suPAR was still found in part

intracellularly. To further characterize suPAR maturation, it is of interest to

investigate intracellular and extracellular suPAR levels after longer chase times to

determine how long it takes for all suPAR translated and radiolabelled within an hour

to be secreted. Equally as important is to identify whether all of the radiolabelled

protein is eventually destined for extracellular expression. Furthermore, probing

shorter chase durations may identify more precisely how long it takes for disulfide

bonds to form within suPAR, providing further characterization of suPAR folding.

 

 

95  

6.3 Comparing suPAR Maturation to Other Proteins in Anoxia  

To assess if suPAR possesses a secretory advantage in the absence of

oxygen, suPAR maturation should be directly compared to that of other cargo in the

same cells. Albumin represents a cargo protein whose disulfide bond formation and

secretion are known to be sensitive to low oxygenation in hypoxia in HepG2 cells

(Koritzinsky et al., in review). To ensure protein maturation in identical conditions,

we are working on a co-transfection model overexpressing suPAR and albumin in

HCT116 cells to simultaneously overexpress the two proteins.

6.4 Investigating Oxygen Dependency of Co- and Post-Translational Disulfide Formation in suPAR  

We observed a requirement for oxygen in post-translational disulfide bond

formation in suPAR, but the capacity of suPAR to co-translationally form disulfide

bonds without oxygen has yet to be explored. It would be of great value to

investigate the maturation of suPAR in anoxia without DTT reduction following the

pulse phase to address the potential of suPAR co-translational disulfide bond

formation without oxygen. These experiments need to include controls that verify

stringent anoxia. Furthermore, similar observations in endogenously expressed

proteins would also provide more persuasive evidence to the role of oxygen in co-

translational disulfide bond formation.

 

 

96  

6.5 Influence of Specific Folding Factors in Anoxia  

Though suPAR may not favorably introduce disulfide bonds in anoxic

conditions, this does not eliminate the possibility that other hypoxia-related proteins

may exhibit preferential folding in the absence of oxygen. Hence, it is important to

pursue extracellularly expressed, hypoxia-induced, disulfide-containing candidates,

such as LOX, LAMP3, CXCR4 and VEGF, to assess if they possess an

advantageous ability to form disulfide bonds in the absence of oxygen or if their

extracellular expression in anoxia is also less dependent on disulfide formation.

Furthermore, as previously mentioned, our group has observed the

transcriptional induction of ER maturation machinery in numerous cancer cell lines in

anoxia (Koritzinsky et al., in review). It is possible that these hypoxia-induced folding

factors may be responsible for the postulated preferential disulfide bond formation in

anoxia by folding privileged proteins or benefiting overall protein folding.

Investigating the influence of hypoxia-induced chaperones, PDIs or ER oxidases on

disulfide formation, protein maturation and extracellular expression of cancer-related

proteins in anoxia may identify machinery that may support malignant progression in

hypoxic tumor microenvironment. Successively, it would be intriguing to assess the

role of these folding factors on cell survival, migration, invasion and metastasis in

hypoxia in hopes of identifying novel anti-cancer targets.

         

 

 

97  

7 CONCLUSIONS  

In this thesis, a transient transfection model overexpressing suPAR tagged

with Myc and DDK in HCT116 human colorectal carcinoma cells was established

and the pulse chase assay was optimized to assess suPAR maturation. Using this

model and assay, we have investigated the kinetics of suPAR disulfide formation,

glycan processing and secretion. We described suPAR maturation through the

formation of disulfide bonds and N-linked glycosylation, though no evidence of

complex glycosylation had been observed. Furthermore, the pulse chase assay has

been validated for its utility in hypoxia, and for the characterization of the oxygen

dependency of suPAR maturation.

We hypothesized that secretion of suPAR during hypoxia relies on superior

disulfide introduction in the absence of oxygen. The work of this thesis demonstrates

that in anoxia, disulfide formation in suPAR was impaired after a reductive

challenge. Interestingly, suPAR was still secreted. We propose that efficient

secretion of suPAR, and possibly other hypoxia-induced proteins, is possible due to

reduced dependency on disulfide bond formation and an evolutionary advantage to

bypass ER quality control in oxygen-lacking conditions, enabling its extracellular

expression despite not having achieved its native conformation. In consequence,

differential ability to bypass normal requirements for disulfide bond formation may

serve as a novel level of regulating extracellular expression in hypoxia, facilitating

tumor progression and the malignant phenotype in the poorly oxygenated tumor

microenvironment. This phenomenon may potentially be exploited to complement

 

 

98  

chemo- and radiotherapy by mitigating the adverse effects of tumor hypoxia and

ultimately improve cancer therapy.

 

 

99  

REFERENCES 1.   Jemal,  A.,  et  al.,  Global  cancer  statistics.  CA  Cancer  J  Clin,  2011.  61(2):  p.  69-­‐

90.  2.   Hanahan,  D.  and  R.A.  Weinberg,  The  hallmarks  of  cancer.  Cell,  2000.  100(1):  

p.  57-­‐70.  3.   Knudson,   A.G.,   Two  genetic   hits   (more   or   less)   to   cancer.   Nat   Rev   Cancer,  

2001.  1(2):  p.  157-­‐62.  4.   Hoeijmakers,   J.H.,   DNA   damage,   aging,   and   cancer.   N   Engl   J   Med,   2009.  

361(15):  p.  1475-­‐85.  5.   Croce,  C.M.,  Oncogenes  and  cancer.  N  Engl  J  Med,  2008.  358(5):  p.  502-­‐11.  6.   Kumar,  V.,   et   al.,  Robbins  basic  pathology.   9th   ed.   2013,   Philadelphia,   PA:  

Elsevier/Saunders.  p.  7.   Berx,  G.,  et  al.,  Mutations  of  the  human  E-­‐cadherin  (CDH1)  gene.  Hum  Mutat,  

1998.  12(4):  p.  226-­‐37.  8.   Cano,   A.,   et   al.,   The   transcription   factor   snail   controls   epithelial-­‐

mesenchymal  transitions  by  repressing  E-­‐cadherin  expression.  Nat  Cell  Biol,  2000.  2(2):  p.  76-­‐83.  

9.   Yang,   J.,   et   al.,   Twist,   a   master   regulator   of   morphogenesis,   plays   an  essential  role  in  tumor  metastasis.  Cell,  2004.  117(7):  p.  927-­‐39.  

10.   Yang,   J.   and   R.A.   Weinberg,   Epithelial-­‐mesenchymal   transition:   at   the  crossroads  of  development  and  tumor  metastasis.  Dev  Cell,  2008.  14(6):  p.  818-­‐29.  

11.   Peinado,  H.,  D.  Olmeda,  and  A.  Cano,  Snail,  Zeb  and  bHLH  factors  in  tumour  progression:  an  alliance  against   the  epithelial  phenotype?   Nat   Rev   Cancer,  2007.  7(6):  p.  415-­‐28.  

12.   Karreth,   F.   and   D.A.   Tuveson,   Twist   induces   an   epithelial-­‐mesenchymal  transition  to  facilitate  tumor  metastasis.   Cancer  Biol  Ther,  2004.  3(11):  p.  1058-­‐9.  

13.   Kiemer,  A.K.,  K.  Takeuchi,  and  M.P.  Quinlan,  Identification  of  genes  involved  in   epithelial-­‐mesenchymal   transition   and   tumor   progression.   Oncogene,  2001.  20(46):  p.  6679-­‐88.  

14.   Janda,   E.,   et   al.,   Ras   and   TGF[beta]   cooperatively   regulate   epithelial   cell  plasticity  and  metastasis:  dissection  of  Ras  signaling  pathways.   J   Cell   Biol,  2002.  156(2):  p.  299-­‐313.  

15.   Vincent-­‐Salomon,   A.   and   J.P.   Thiery,   Host   microenvironment   in   breast  cancer   development:   epithelial-­‐mesenchymal   transition   in   breast   cancer  development.  Breast  Cancer  Res,  2003.  5(2):  p.  101-­‐6.  

16.   Xue,   C.,   et   al.,   The   gatekeeper   effect   of   epithelial-­‐mesenchymal   transition  regulates   the   frequency   of   breast   cancer   metastasis.   Cancer   Res,   2003.  63(12):  p.  3386-­‐94.  

17.   Liotta,   L.A.,   et   al.,   Degradation   of   basement   membrane   by   murine   tumor  cells.  J  Natl  Cancer  Inst,  1977.  58(5):  p.  1427-­‐31.  

18.   Liotta,   L.A.,   et   al.,   Metastatic   potential   correlates   with   enzymatic  degradation  of  basement  membrane  collagen.  Nature,  1980.  284(5751):  p.  67-­‐8.  

 

 

100  

19.   Nakajima,   M.,   et   al.,  Degradation   of   basement  membrane   type   IV   collagen  and   lung   subendothelial   matrix   by   rat   mammary   adenocarcinoma   cell  clones  of  differing  metastatic  potentials.  Cancer  Res,  1987.  47(18):  p.  4869-­‐76.  

20.   Salo,  T.,  L.A.  Liotta,  and  K.  Tryggvason,  Purification  and  characterization  of  a   murine   basement   membrane   collagen-­‐degrading   enzyme   secreted   by  metastatic  tumor  cells.  J  Biol  Chem,  1983.  258(5):  p.  3058-­‐63.  

21.   Szpaderska,   A.M.   and   A.   Frankfater,   An   intracellular   form   of   cathepsin   B  contributes  to  invasiveness  in  cancer.  Cancer  Res,  2001.  61(8):  p.  3493-­‐500.  

22.   Lah,   T.T.,   et   al.,  Degradation  of   laminin  by  human  tumor  cathepsin  B.   Clin  Exp  Metastasis,  1989.  7(4):  p.  461-­‐8.  

23.   Wyckoff,   J.B.,   et   al.,   A   critical   step   in   metastasis:   in   vivo   analysis   of  intravasation  at  the  primary  tumor.  Cancer  Res,  2000.  60(9):  p.  2504-­‐11.  

24.   Warren,   B.A.,   Environment   of   the   blood-­‐borne   tumor   embolus   adherent   to  vessel  wall.  J  Med,  1973.  4(3):  p.  150-­‐77.  

25.   Kramer,  R.H.,  R.  Gonzalez,  and  G.L.  Nicolson,  Metastatic  tumor  cells  adhere  preferentially   to   the   extracellular   matrix   underlying   vascular   endothelial  cells.  Int  J  Cancer,  1980.  26(5):  p.  639-­‐45.  

26.   Zetter,   B.R.,   Adhesion   molecules   in   tumor   metastasis.   Semin   Cancer   Biol,  1993.  4(4):  p.  219-­‐29.  

27.   Miles,  F.L.,  et  al.,  Stepping  out  of  the  flow:  capillary  extravasation  in  cancer  metastasis.  Clin  Exp  Metastasis,  2008.  25(4):  p.  305-­‐24.  

28.   Luzzi,   K.J.,   et   al.,  Multistep   nature   of   metastatic   inefficiency:   dormancy   of  solitary   cells   after   successful   extravasation   and   limited   survival   of   early  micrometastases.  Am  J  Pathol,  1998.  153(3):  p.  865-­‐73.  

29.   Rosenberg,   S.A.,  Adoptive   immunotherapy  of   cancer:  accomplishments  and  prospects.  Cancer  Treat  Rep,  1984.  68(1):  p.  233-­‐55.  

30.   Maneet   Bhatia,   T.C.K.   and   G.D.T.a.K.F.   Tonissen,   The   Interaction   Between  Redox  and  Hypoxic  Signalling  Pathways  in  the  Dynamic  Oxygen  Environment  of  Cancer  Cells,  Carcinogenesis,  Dr.  Kathryn  Tonissen  (Ed.).  2013.  

31.   Simon,   M.C.   and   B.   Keith,   The   role   of   oxygen   availability   in   embryonic  development  and  stem  cell  function.  Nature  reviews.  Molecular  cell  biology,  2008.  9(4):  p.  285-­‐96.  

32.   Wouters,  B.G.  and  M.  Koritzinsky,  Hypoxia  signalling  through  mTOR  and  the  unfolded  protein  response  in  cancer.  Nature  reviews.  Cancer,  2008.  8(11):  p.  851-­‐64.  

33.   Wouters,  B.G.  and  M.  Koritzinsky,  Hypoxia  signalling  through  mTOR  and  the  unfolded  protein  response  in  cancer.  Nat  Rev  Cancer,  2008.  8(11):  p.  851-­‐64.  

34.   Brown,   J.M.   and   W.R.   Wilson,   Exploiting   tumour   hypoxia   in   cancer  treatment.  Nat  Rev  Cancer,  2004.  4(6):  p.  437-­‐47.  

35.   Hockel,  M.   and   P.   Vaupel,  Tumor  hypoxia:  definitions  and  current   clinical,  biologic,  and  molecular  aspects.  J  Natl  Cancer  Inst,  2001.  93(4):  p.  266-­‐76.  

36.   Knocke,  T.H.,  et  al.,   Intratumoral  pO2-­‐measurements  as  predictive  assay  in  the   treatment   of   carcinoma   of   the   uterine   cervix.   Radiother   Oncol,   1999.  53(2):  p.  99-­‐104.  

 

 

101  

37.   Hockel,   M.,   et   al.,   Association   between   tumor   hypoxia   and   malignant  progression   in   advanced   cancer   of   the   uterine   cervix.   Cancer   Res,   1996.  56(19):  p.  4509-­‐15.  

38.   Nordsmark,  M.,  M.  Overgaard,  and  J.  Overgaard,  Pretreatment  oxygenation  predicts   radiation   response   in   advanced   squamous   cell   carcinoma   of   the  head  and  neck.  Radiother  Oncol,  1996.  41(1):  p.  31-­‐9.  

39.   Brizel,   D.M.,   et   al.,   Tumor   hypoxia   adversely   affects   the   prognosis   of  carcinoma  of  the  head  and  neck.  Int  J  Radiat  Oncol  Biol  Phys,  1997.  38(2):  p.  285-­‐9.  

40.   Lartigau,   E.,   et   al.,   Intratumoral   oxygen   tension   in   metastatic   melanoma.  Melanoma  Res,  1997.  7(5):  p.  400-­‐6.  

41.   Rofstad,   E.K.   and   K.   Maseide,   Radiobiological   and   immunohistochemical  assessment   of   hypoxia   in   human  melanoma   xenografts:   acute   and   chronic  hypoxia  in  individual  tumours.  Int  J  Radiat  Biol,  1999.  75(11):  p.  1377-­‐93.  

42.   Vaupel,  P.,  et  al.,  Oxygenation  of  human  tumors:  evaluation  of  tissue  oxygen  distribution   in   breast   cancers   by   computerized   O2   tension  measurements.  Cancer  Res,  1991.  51(12):  p.  3316-­‐22.  

43.   Runkel,   S.,   et   al.,   Oxygenation   of   mammary   tumors   as   evaluated   by  ultrasound-­‐guided  computerized-­‐pO2-­‐histography.  Adv  Exp  Med  Biol,  1994.  345:  p.  451-­‐8.  

44.   Le,   Q.T.,   N.C.   Denko,   and   A.J.   Giaccia,   Hypoxic   gene   expression   and  metastasis.  Cancer  Metastasis  Rev,  2004.  23(3-­‐4):  p.  293-­‐310.  

45.   Vaupel,  P.,  F.  Kallinowski,  and  P.  Okunieff,  Blood  flow,  oxygen  and  nutrient  supply,  and  metabolic  microenvironment  of  human  tumors:  a  review.  Cancer  Res,  1989.  49(23):  p.  6449-­‐65.  

46.   Vaupel,  P.,  D.K.  Kelleher,   and  O.  Thews,  Modulation  of  tumor  oxygenation.  Int  J  Radiat  Oncol  Biol  Phys,  1998.  42(4):  p.  843-­‐8.  

47.   Harris,   A.N.   and   P.L.   Mellon,   The   basic   helix-­‐loop-­‐helix,   leucine   zipper  transcription  factor,  USF  (upstream  stimulatory  factor),  is  a  key  regulator  of  SF-­‐1  (steroidogenic   factor-­‐1)  gene  expression   in  pituitary  gonadotrope  and  steroidogenic  cells.  Mol  Endocrinol,  1998.  12(5):  p.  714-­‐26.  

48.   Brown,   J.M.,   Evidence   for   acutely   hypoxic   cells   in   mouse   tumours,   and   a  possible  mechanism  of  reoxygenation.  Br  J  Radiol,  1979.  52(620):  p.  650-­‐6.  

49.   Chaplin,  D.J.,  P.L.  Olive,  and  R.E.  Durand,  Intermittent  blood  flow  in  a  murine  tumor:  radiobiological  effects.  Cancer  Res,  1987.  47(2):  p.  597-­‐601.  

50.   Dewhirst,   M.W.,   Y.   Cao,   and   B.   Moeller,   Cycling   hypoxia   and   free   radicals  regulate   angiogenesis   and   radiotherapy   response.   Nat   Rev   Cancer,   2008.  8(6):  p.  425-­‐37.  

51.   Lanzen,   J.,   et   al.,   Direct   demonstration   of   instabilities   in   oxygen  concentrations   within   the   extravascular   compartment   of   an   experimental  tumor.  Cancer  Res,  2006.  66(4):  p.  2219-­‐23.  

52.   Bennewith,   K.L.   and  R.E.   Durand,  Quantifying   transient  hypoxia   in  human  tumor  xenografts  by  flow  cytometry.  Cancer  Res,  2004.  64(17):  p.  6183-­‐9.  

53.   Cardenas-­‐Navia,  L.I.,  et  al.,  Tumor-­‐dependent  kinetics  of  partial  pressure  of  oxygen   fluctuations   during   air   and   oxygen   breathing.   Cancer   Res,   2004.  64(17):  p.  6010-­‐7.  

 

 

102  

54.   Brown,   J.M.,   Tumor   hypoxia   in   cancer   therapy.   Methods   Enzymol,   2007.  435:  p.  297-­‐321.  

55.   Mortensen,   L.S.,   et   al.,   Accessing   radiation   response   using   hypoxia   PET  imaging   and   oxygen   sensitive   electrodes:   a   preclinical   study.   Radiother  Oncol,  2011.  99(3):  p.  418-­‐23.  

56.   Horsman,   M.R.,   et   al.,   Relationship   between   radiobiological   hypoxia   in  tumors   and   electrode   measurements   of   tumor   oxygenation.   Int   J   Radiat  Oncol  Biol  Phys,  1994.  29(3):  p.  439-­‐42.  

57.   Tatum,   J.L.,   et   al.,   Hypoxia:   importance   in   tumor   biology,   noninvasive  measurement  by  imaging,  and  value  of  its  measurement  in  the  management  of  cancer  therapy.  Int  J  Radiat  Biol,  2006.  82(10):  p.  699-­‐757.  

58.   Nordsmark,  M.,  et  al.,  Differential  risk  assessments  from  five  hypoxia  specific  assays:   The   basis   for   biologically   adapted   individualized   radiotherapy   in  advanced  head  and  neck  cancer  patients.   Radiother  Oncol,   2007.   83(3):   p.  389-­‐97.  

59.   Varia,  M.A.,  et  al.,  Pimonidazole:  a  novel  hypoxia  marker  for  complementary  study  of  tumor  hypoxia  and  cell  proliferation  in  cervical  carcinoma.  Gynecol  Oncol,  1998.  71(2):  p.  270-­‐7.  

60.   Piert,   M.,   et   al.,   Hypoxia-­‐specific   tumor   imaging   with   18F-­‐fluoroazomycin  arabinoside.  J  Nucl  Med,  2005.  46(1):  p.  106-­‐13.  

61.   Koch,   C.J.,   et   al.,   Biodistribution   and   dosimetry   of   (18)F-­‐EF5   in   cancer  patients   with   preliminary   comparison   of   (18)F-­‐EF5   uptake   versus   EF5  binding  in  human  glioblastoma.  Eur  J  Nucl  Med  Mol  Imaging,  2010.  37(11):  p.  2048-­‐59.  

62.   Kaanders,   J.H.,   et   al.,   Pimonidazole   binding   and   tumor   vascularity   predict  for  treatment  outcome  in  head  and  neck  cancer.   Cancer  Res,  2002.  62(23):  p.  7066-­‐74.  

63.   Bollineni,   V.R.,   et   al.,   PET   Imaging   of   Tumor   Hypoxia   Using   18F-­‐Fluoroazomycin   Arabinoside   in   Stage   III-­‐IV   Non-­‐Small   Cell   Lung   Cancer  Patients.  J  Nucl  Med,  2013.  

64.   Nordsmark,   M.,   et   al.,   Measurements   of   hypoxia   using   pimonidazole   and  polarographic   oxygen-­‐sensitive   electrodes   in   human   cervix   carcinomas.  Radiother  Oncol,  2003.  67(1):  p.  35-­‐44.  

65.   Hockel,  M.,  et  al.,  Hypoxia  and  Radiation  Response  in  Human  Tumors.  Semin  Radiat  Oncol,  1996.  6(1):  p.  3-­‐9.  

66.   Nordsmark,   M.,   et   al.,   Prognostic   value   of   tumor   oxygenation   in   397  head  and   neck   tumors   after   primary   radiation   therapy.   An   international  multi-­‐center  study.  Radiother  Oncol,  2005.  77(1):  p.  18-­‐24.  

67.   Fyles,   A.,   et   al.,   Tumor   hypoxia   has   independent   predictor   impact   only   in  patients  with  node-­‐negative  cervix  cancer.  J  Clin  Oncol,  2002.  20(3):  p.  680-­‐7.  

68.   Fyles,   A.,   et   al.,   Long-­‐term   performance   of   interstial   fluid   pressure   and  hypoxia   as   prognostic   factors   in   cervix   cancer.   Radiother   Oncol,   2006.  80(2):  p.  132-­‐7.  

69.   Fyles,   A.W.,   et   al.,   Cervix   cancer   oxygenation  measured   following   external  radiation  therapy.  Int  J  Radiat  Oncol  Biol  Phys,  1998.  42(4):  p.  751-­‐3.  

 

 

103  

70.   Harris,   A.L.,   Hypoxia-­‐-­‐a   key   regulatory   factor   in   tumour   growth.   Nat   Rev  Cancer,  2002.  2(1):  p.  38-­‐47.  

71.   Wouters,  B.G.,  et  al.,  Hypoxia  as  a  target  for  combined  modality  treatments.  Eur  J  Cancer,  2002.  38(2):  p.  240-­‐57.  

72.   Hockel,   M.,   et   al.,   [Combined   surgical   and   radiotherapeutic   treatment   of  pelvic   wall   recurrences:   report   of   experiences   after   3   years].   Geburtshilfe  Frauenheilkd,  1993.  53(3):  p.  169-­‐76.  

73.   Vaupel,   P.,   et   al.,   Oxygenation   of   human   tumors:   the   Mainz   experience.  Strahlenther  Onkol,  1998.  174  Suppl  4:  p.  6-­‐12.  

74.   Hockel,  M.  and  P.  Vaupel,  Biological  consequences  of  tumor  hypoxia.  Semin  Oncol,  2001.  28(2  Suppl  8):  p.  36-­‐41.  

75.   Vaupel,   P.   and   M.   Hoeckel,   Predictive   power   of   the   tumor   oxygenation  status.  Adv  Exp  Med  Biol,  1999.  471:  p.  533-­‐9.  

76.   Hammond,   E.M.,   M.J.   Dorie,   and   A.J.   Giaccia,   ATR/ATM   targets   are  phosphorylated   by   ATR   in   response   to   hypoxia   and   ATM   in   response   to  reoxygenation.  J  Biol  Chem,  2003.  278(14):  p.  12207-­‐13.  

77.   Madan,   E.,   R.   Gogna,   and   U.   Pati,   p53   Ser15   phosphorylation   disrupts   the  p53-­‐RPA70   complex   and   induces   RPA70-­‐mediated   DNA   repair   in   hypoxia.  Biochem  J,  2012.  443(3):  p.  811-­‐20.  

78.   Graeber,   T.G.,   et   al.,   Hypoxia-­‐mediated   selection   of   cells   with   diminished  apoptotic  potential  in  solid  tumours.  Nature,  1996.  379(6560):  p.  88-­‐91.  

79.   Comerford,  K.M.,   et   al.,  Hypoxia-­‐inducible  factor-­‐1-­‐dependent  regulation  of  the  multidrug  resistance  (MDR1)  gene.   Cancer  Res,  2002.  62(12):  p.  3387-­‐94.  

80.   Wartenberg,  M.,  et  al.,  Regulation  of  the  multidrug  resistance  transporter  P-­‐glycoprotein   in   multicellular   tumor   spheroids   by   hypoxia-­‐inducible   factor  (HIF-­‐1)  and  reactive  oxygen  species.  FASEB  J,  2003.  17(3):  p.  503-­‐5.  

81.   Lin,   J.H.   and   M.   Yamazaki,   Role   of   P-­‐glycoprotein   in   pharmacokinetics:  clinical  implications.  Clin  Pharmacokinet,  2003.  42(1):  p.  59-­‐98.  

82.   Gray,  L.H.,  et  al.,  The  concentration  of  oxygen  dissolved  in  tissues  at  the  time  of  irradiation  as  a  factor  in  radiotherapy.  Br  J  Radiol,  1953.  26(312):  p.  638-­‐48.  

83.   Brizel,   D.M.,   et   al.,   Oxygenation   of   head   and   neck   cancer:   changes   during  radiotherapy   and   impact   on   treatment   outcome.   Radiother   Oncol,   1999.  53(2):  p.  113-­‐7.  

84.   Fyles,   A.W.,   et   al.,  Oxygenation  predicts  radiation  response  and  survival   in  patients  with  cervix  cancer.  Radiother  Oncol,  1998.  48(2):  p.  149-­‐56.  

85.   Vordermark,   D.   and   J.M.   Brown,   Endogenous   markers   of   tumor   hypoxia  predictors   of   clinical   radiation   resistance?   Strahlenther   Onkol,   2003.  179(12):  p.  801-­‐11.  

86.   Koch,   C.J.,  Oxygen  effects   in  radiobiology.   Adv  Exp  Med  Biol,   1982.   157:   p.  123-­‐44.  

87.   Kizaka-­‐Kondoh,   S.,   et   al.,   Tumor   hypoxia:   a   target   for   selective   cancer  therapy.  Cancer  Sci,  2003.  94(12):  p.  1021-­‐8.  

 

 

104  

88.   Batchelder,   R.M.,   et   al.,   Oxygen   dependence   of   the   cytotoxicity   of   the  enediyne   anti-­‐tumour   antibiotic   esperamicin  A1.   Br   J   Cancer   Suppl,   1996.  27:  p.  S52-­‐6.  

89.   Teicher,   B.A.,   J.S.   Lazo,   and   A.C.   Sartorelli,   Classification   of   antineoplastic  agents   by   their   selective   toxicities   toward   oxygenated   and   hypoxic   tumor  cells.  Cancer  Res,  1981.  41(1):  p.  73-­‐81.  

90.   Tannock,   I.F.,   Conventional   cancer   therapy:   promise   broken   or   promise  delayed?  Lancet,  1998.  351  Suppl  2:  p.  SII9-­‐16.  

91.   Durand,   R.E.,   The   influence   of   microenvironmental   factors   during   cancer  therapy.  In  Vivo,  1994.  8(5):  p.  691-­‐702.  

92.   Hicks,   K.O.,   et   al.,   Use   of   three-­‐dimensional   tissue   cultures   to   model  extravascular   transport   and   predict   in   vivo   activity   of   hypoxia-­‐targeted  anticancer  drugs.  J  Natl  Cancer  Inst,  2006.  98(16):  p.  1118-­‐28.  

93.   Sutherland,  R.M.,   et   al.,  Tumor  Hypoxia  and  Heterogeneity:  Challenges  and  Opportunities  for  the  Future.  Semin  Radiat  Oncol,  1996.  6(1):  p.  59-­‐70.  

94.   Brown,   J.M.   and   A.J.   Giaccia,   The   unique   physiology   of   solid   tumors:  opportunities  (and  problems)  for  cancer  therapy.   Cancer  Res,  1998.  58(7):  p.  1408-­‐16.  

95.   Chaplin,   D.J.   and   S.A.   Hill,   Temporal   heterogeneity   in   microregional  erythrocyte  flux  in  experimental  solid  tumours.  Br  J  Cancer,  1995.  71(6):  p.  1210-­‐3.  

96.   Tannock,   I.F.,   The   relation   between   cell   proliferation   and   the   vascular  system   in   a   transplanted   mouse   mammary   tumour.   Br   J   Cancer,   1968.  22(2):  p.  258-­‐73.  

97.   Horsman,  M.R.,   D.J.   Chaplin,   and   J.M.   Brown,  Tumor   radiosensitization  by  nicotinamide:  a   result  of   improved  perfusion  and  oxygenation.   Radiat   Res,  1989.  118(1):  p.  139-­‐50.  

98.   Chaplin,  D.J.,  M.R.  Horsman,  and  M.J.  Trotter,  Effect  of  nicotinamide  on  the  microregional   heterogeneity   of   oxygen   delivery   within   a   murine   tumor.   J  Natl  Cancer  Inst,  1990.  82(8):  p.  672-­‐6.  

99.   Kaanders,   J.H.,   J.   Bussink,   and  A.J.   van  der  Kogel,  ARCON:  a  novel  biology-­‐based  approach  in  radiotherapy.  Lancet  Oncol,  2002.  3(12):  p.  728-­‐37.  

100.   Kaanders,   J.H.,   et   al.,   ARCON:   experience   in   215   patients   with   advanced  head-­‐and-­‐neck  cancer.  Int  J  Radiat  Oncol  Biol  Phys,  2002.  52(3):  p.  769-­‐78.  

101.   Brown,  M.,  Henry  S.  Kaplan  Distinguished  Scientist  Award  Lecture  2007.  The  remarkable  yin  and  yang  of  tumour  hypoxia.  Int  J  Radiat  Biol,  2010.  86(11):  p.  907-­‐17.  

102.   Zeman,   E.M.,   et   al.,   SR-­‐4233:   a   new  bioreductive   agent  with   high   selective  toxicity   for   hypoxic  mammalian   cells.   Int   J   Radiat   Oncol   Biol   Phys,   1986.  12(7):  p.  1239-­‐42.  

103.   Zeman,  E.M.,  M.J.  Lemmon,  and  J.M.  Brown,  Aerobic  radiosensitization  by  SR  4233  in  vitro  and  in  vivo.   Int  J  Radiat  Oncol  Biol  Phys,  1990.  18(1):  p.  125-­‐32.  

104.   Brown,  J.M.  and  M.J.  Lemmon,  Potentiation  by  the  hypoxic  cytotoxin  SR  4233  of  cell  killing  produced  by  fractionated  irradiation  of  mouse  tumors.  Cancer  Res,  1990.  50(24):  p.  7745-­‐9.  

 

 

105  

105.   Gardner,  L.B.,   et   al.,  Hypoxia  inhibits  G1/S  transition  through  regulation  of  p27  expression.  J  Biol  Chem,  2001.  276(11):  p.  7919-­‐26.  

106.   Connolly,  E.,  et  al.,  Hypoxia  inhibits  protein  synthesis  through  a  4E-­‐BP1  and  elongation   factor   2   kinase   pathway   controlled   by  mTOR   and   uncoupled   in  breast  cancer  cells.  Mol  Cell  Biol,  2006.  26(10):  p.  3955-­‐65.  

107.   Koumenis,  C.,  et  al.,  Regulation  of  protein  synthesis  by  hypoxia  via  activation  of   the   endoplasmic   reticulum   kinase   PERK   and   phosphorylation   of   the  translation  initiation  factor  eIF2alpha.  Mol  Cell  Biol,  2002.  22(21):  p.  7405-­‐16.  

108.   Casey,   T.M.,   et   al.,   Hypoxia   causes   downregulation   of   protein   and   RNA  synthesis   in   noncontracting   Mammalian   cardiomyocytes.   Circ   Res,   2002.  90(7):  p.  777-­‐83.  

109.   Park,   J.E.,   et   al.,   Hypoxic   tumor   cell   modulates   its   microenvironment   to  enhance   angiogenic   and  metastatic   potential   by   secretion   of   proteins   and  exosomes.  Mol  Cell  Proteomics,  2010.  9(6):  p.  1085-­‐99.  

110.   Liu,   L.,   et   al.,   Activation   of   beta-­‐catenin   by   hypoxia   in   hepatocellular  carcinoma  contributes  to  enhanced  metastatic  potential  and  poor  prognosis.  Clin  Cancer  Res,  2010.  16(10):  p.  2740-­‐50.  

111.   Rofstad,  E.K.  and  T.  Danielsen,  Hypoxia-­‐induced  angiogenesis  and  vascular  endothelial  growth  factor  secretion  in  human  melanoma.  Br  J  Cancer,  1998.  77(6):  p.  897-­‐902.  

112.   Garcia-­‐Roman,   J.,   et   al.,   VEGF   secretion   during   hypoxia   depends   on   free  radicals-­‐induced   Fyn   kinase   activity   in   mast   cells.   Biochem   Biophys   Res  Commun,  2010.  401(2):  p.  262-­‐7.  

113.   Young,   S.D.,   R.S.   Marshall,   and   R.P.   Hill,   Hypoxia   induces   DNA  overreplication   and   enhances   metastatic   potential   of   murine   tumor   cells.  Proc  Natl  Acad  Sci  U  S  A,  1988.  85(24):  p.  9533-­‐7.  

114.   Dai,   Y.,   K.   Bae,   and   D.W.   Siemann,   Impact   of   hypoxia   on   the   metastatic  potential  of  human  prostate  cancer  cells.  Int  J  Radiat  Oncol  Biol  Phys,  2011.  81(2):  p.  521-­‐8.  

115.   Ying,   Q.,   et   al.,   Hypoxia-­‐inducible   microRNA-­‐210   augments   the  metastatic  potential   of   tumor   cells   by   targeting   vacuole   membrane   protein   1   in  hepatocellular  carcinoma.  Hepatology,  2011.  54(6):  p.  2064-­‐75.  

116.   Funasaka,   T.,   et   al.,   Regulation   of   phosphoglucose   isomerase/autocrine  motility  factor  expression  by  hypoxia.  FASEB  J,  2005.  19(11):  p.  1422-­‐30.  

117.   Yanagawa,   T.,   et   al.,   Differential   regulation   of   phosphoglucose  isomerase/autocrine   motility   factor   activities   by   protein   kinase   CK2  phosphorylation.  J  Biol  Chem,  2005.  280(11):  p.  10419-­‐26.  

118.   Karakurum,  M.,  et  al.,  Hypoxic  induction  of  interleukin-­‐8  gene  expression  in  human  endothelial  cells.  J  Clin  Invest,  1994.  93(4):  p.  1564-­‐70.  

119.   Ben-­‐Yosef,  Y.,   et   al.,  Hypoxia  of  endothelial  cells  leads  to  MMP-­‐2-­‐dependent  survival  and  death.  Am  J  Physiol  Cell  Physiol,  2005.  289(5):  p.  C1321-­‐31.  

120.   Krishnamachary,   B.,   et   al.,  Regulation   of   colon   carcinoma   cell   invasion  by  hypoxia-­‐inducible  factor  1.  Cancer  Res,  2003.  63(5):  p.  1138-­‐43.  

 

 

106  

121.   Canning,   M.T.,   et   al.,  Oxygen-­‐mediated   regulation   of   gelatinase   and   tissue  inhibitor  of  metalloproteinases-­‐1  expression  by  invasive  cells.   Exp  Cell  Res,  2001.  267(1):  p.  88-­‐94.  

122.   Sahlgren,   C.,   et   al.,   Notch   signaling   mediates   hypoxia-­‐induced   tumor   cell  migration  and  invasion.  Proc  Natl  Acad  Sci  U  S  A,  2008.  105(17):  p.  6392-­‐7.  

123.   Plasswilm,   L.,   et   al.,   Hypoxia-­‐induced   tumour   cell   migration   in   an   in   vivo  chicken  model.  Pathobiology,  2000.  68(3):  p.  99-­‐105.  

124.   De   Jaeger,   K.,   M.C.   Kavanagh,   and   R.P.   Hill,   Relationship   of   hypoxia   to  metastatic  ability  in  rodent  tumours.  Br  J  Cancer,  2001.  84(9):  p.  1280-­‐5.  

125.   Airley,  R.,  et  al.,  Glucose  transporter  glut-­‐1  expression  correlates  with  tumor  hypoxia  and  predicts  metastasis-­‐free  survival  in  advanced  carcinoma  of  the  cervix.  Clin  Cancer  Res,  2001.  7(4):  p.  928-­‐34.  

126.   Cairns,  R.A.  and  R.P.  Hill,  Acute  hypoxia  enhances  spontaneous  lymph  node  metastasis   in   an   orthotopic   murine   model   of   human   cervical   carcinoma.  Cancer  Res,  2004.  64(6):  p.  2054-­‐61.  

127.   Erler,   J.T.,   et   al.,   Lysyl   oxidase   is   essential   for  hypoxia-­‐induced  metastasis.  Nature,  2006.  440(7088):  p.  1222-­‐6.  

128.   Kunz,  M.   and   S.M.   Ibrahim,  Molecular  responses   to  hypoxia   in   tumor  cells.  Mol  Cancer,  2003.  2:  p.  23.  

129.   Semenza,   G.L.,   Regulation   of   mammalian   O2   homeostasis   by   hypoxia-­‐inducible  factor  1.  Annu  Rev  Cell  Dev  Biol,  1999.  15:  p.  551-­‐78.  

130.   Zhong,   H.,   et   al.,   Overexpression   of   hypoxia-­‐inducible   factor   1alpha   in  common  human  cancers  and  their  metastases.  Cancer  Res,  1999.  59(22):  p.  5830-­‐5.  

131.   Maxwell,   P.H.,   et   al.,  The   tumour   suppressor   protein  VHL   targets   hypoxia-­‐inducible   factors   for   oxygen-­‐dependent   proteolysis.   Nature,   1999.  399(6733):  p.  271-­‐5.  

132.   Ivan,  M.,   et   al.,  HIFalpha  targeted   for  VHL-­‐mediated  destruction  by  proline  hydroxylation:   implications   for   O2   sensing.   Science,   2001.   292(5516):   p.  464-­‐8.  

133.   Jaakkola,   P.,   et   al.,   Targeting   of   HIF-­‐alpha   to   the   von   Hippel-­‐Lindau  ubiquitylation  complex  by  O2-­‐regulated  prolyl  hydroxylation.  Science,  2001.  292(5516):  p.  468-­‐72.  

134.   Schofield,  C.J.  and  P.J.  Ratcliffe,  Oxygen  sensing  by  HIF  hydroxylases.  Nat  Rev  Mol  Cell  Biol,  2004.  5(5):  p.  343-­‐54.  

135.   Semenza,  G.L.,  Oxygen  sensing,  homeostasis,  and  disease.  The  New  England  journal  of  medicine,  2011.  365(6):  p.  537-­‐47.  

136.   Semenza,  G.L.,  Hydroxylation  of  HIF-­‐1:  oxygen  sensing  at  the  molecular  level.  Physiology  (Bethesda),  2004.  19:  p.  176-­‐82.  

137.   Kallio,   P.J.,   et   al.,   Signal   transduction   in   hypoxic   cells:   inducible   nuclear  translocation  and  recruitment  of   the  CBP/p300  coactivator  by  the  hypoxia-­‐inducible  factor-­‐1alpha.  EMBO  J,  1998.  17(22):  p.  6573-­‐86.  

138.   O'Rourke,  J.F.,  et  al.,  Hypoxia  response  elements.  Oncol  Res,  1997.  9(6-­‐7):  p.  327-­‐32.  

139.   Semenza,   G.L.,   Targeting  HIF-­‐1   for   cancer   therapy.   Nat   Rev   Cancer,   2003.  3(10):  p.  721-­‐32.  

 

 

107  

140.   Semenza,   G.L.   and   G.L.   Wang,   A   nuclear   factor   induced   by   hypoxia   via   de  novo  protein  synthesis  binds  to  the  human  erythropoietin  gene  enhancer  at  a  site  required  for  transcriptional  activation.  Mol  Cell  Biol,  1992.  12(12):  p.  5447-­‐54.  

141.   Iyer,   N.V.,   et   al.,   Cellular   and   developmental   control   of   O2   homeostasis   by  hypoxia-­‐inducible  factor  1  alpha.  Genes  Dev,  1998.  12(2):  p.  149-­‐62.  

142.   Wykoff,   C.C.,   et   al.,   Hypoxia-­‐inducible   expression   of   tumor-­‐associated  carbonic  anhydrases.  Cancer  Res,  2000.  60(24):  p.  7075-­‐83.  

143.   Levy,   A.P.,   et   al.,  Transcriptional  regulation  of   the  rat  vascular  endothelial  growth  factor  gene  by  hypoxia.  J  Biol  Chem,  1995.  270(22):  p.  13333-­‐40.  

144.   Semenza,   G.L.,   Regulation   of   hypoxia-­‐induced   angiogenesis:   a   chaperone  escorts  VEGF  to  the  dance.  J  Clin  Invest,  2001.  108(1):  p.  39-­‐40.  

145.   Veikkola,   T.,   et   al.,   Regulation   of   angiogenesis   via   vascular   endothelial  growth  factor  receptors.  Cancer  Res,  2000.  60(2):  p.  203-­‐12.  

146.   Forsythe,  M.,  Purchasing  vascular   services.  Develop   specialised  units.   BMJ,  1994.  309(6958):  p.  881.  

147.   Minchenko,   A.,   et   al.,  Hypoxia   regulatory   elements   of   the   human   vascular  endothelial  growth  factor  gene.  Cell  Mol  Biol  Res,  1994.  40(1):  p.  35-­‐9.  

148.   Tung,   K.H.,   et   al.,   CHC   promotes   tumor   growth   and   angiogenesis   through  regulation  of  HIF-­‐1alpha  and  VEGF  signaling.  Cancer  Lett,  2013.  331(1):  p.  58-­‐67.  

149.   Thomas,  S.,  et  al.,  CD24  is  an  effector  of  HIF-­‐1-­‐driven  primary  tumor  growth  and  metastasis.  Cancer  Res,  2012.  72(21):  p.  5600-­‐12.  

150.   Schwab,   L.P.,   et   al.,   Hypoxia-­‐inducible   factor   1alpha   promotes   primary  tumor   growth   and   tumor-­‐initiating   cell   activity   in   breast   cancer.   Breast  Cancer  Res,  2012.  14(1):  p.  R6.  

151.   Jiang,   H.   and   Y.   Feng,   Hypoxia-­‐inducible   factor   1alpha   (HIF-­‐1alpha)  correlated  with  tumor  growth  and  apoptosis  in  ovarian  cancer.  Int  J  Gynecol  Cancer,  2006.  16  Suppl  1:  p.  405-­‐12.  

152.   Talks,   K.L.,   et   al.,  The  expression  and  distribution  of   the  hypoxia-­‐inducible  factors   HIF-­‐1alpha   and  HIF-­‐2alpha   in   normal   human   tissues,   cancers,   and  tumor-­‐associated  macrophages.  Am  J  Pathol,  2000.  157(2):  p.  411-­‐21.  

153.   Wenger,   R.H.,   Cellular   adaptation   to   hypoxia:   O2-­‐sensing   protein  hydroxylases,   hypoxia-­‐inducible   transcription   factors,   and   O2-­‐regulated  gene  expression.  FASEB  J,  2002.  16(10):  p.  1151-­‐62.  

154.   Rankin,   E.B.,   et   al.,   Hypoxia-­‐inducible   factor-­‐2   (HIF-­‐2)   regulates   hepatic  erythropoietin  in  vivo.  J  Clin  Invest,  2007.  117(4):  p.  1068-­‐77.  

155.   Ebert,  B.L.,  J.D.  Firth,  and  P.J.  Ratcliffe,  Hypoxia  and  mitochondrial  inhibitors  regulate   expression   of   glucose   transporter-­‐1   via   distinct   Cis-­‐acting  sequences.  J  Biol  Chem,  1995.  270(49):  p.  29083-­‐9.  

156.   Maxwell,   P.H.,   et   al.,  Hypoxia-­‐inducible  factor-­‐1  modulates  gene  expression  in   solid   tumors   and   influences   both   angiogenesis   and   tumor   growth.   Proc  Natl  Acad  Sci  U  S  A,  1997.  94(15):  p.  8104-­‐9.  

157.   Semenza,  G.L.,  et  al.,  Transcriptional  regulation  of  genes  encoding  glycolytic  enzymes   by   hypoxia-­‐inducible   factor   1.   J   Biol   Chem,   1994.   269(38):   p.  23757-­‐63.  

 

 

108  

158.   Ullah,  M.S.,   A.J.   Davies,   and   A.P.   Halestrap,  The  plasma  membrane   lactate  transporter  MCT4,  but  not  MCT1,  is  up-­‐regulated  by  hypoxia  through  a  HIF-­‐1alpha-­‐dependent  mechanism.  J  Biol  Chem,  2006.  281(14):  p.  9030-­‐7.  

159.   Zhang,   H.,   et   al.,   HIF-­‐1   inhibits   mitochondrial   biogenesis   and   cellular  respiration   in   VHL-­‐deficient   renal   cell   carcinoma   by   repression   of   C-­‐MYC  activity.  Cancer  Cell,  2007.  11(5):  p.  407-­‐20.  

160.   Li,   F.,   et   al.,   Myc   stimulates   nuclearly   encoded   mitochondrial   genes   and  mitochondrial  biogenesis.  Mol  Cell  Biol,  2005.  25(14):  p.  6225-­‐34.  

161.   Liu,   Y.,   et   al.,   Hypoxia   regulates   vascular   endothelial   growth   factor   gene  expression   in   endothelial   cells.   Identification   of   a   5'   enhancer.   Circ   Res,  1995.  77(3):  p.  638-­‐43.  

162.   Zheng,   H.,   et   al.,   Migration   of   endothelial   progenitor   cells   mediated   by  stromal   cell-­‐derived   factor-­‐1alpha/CXCR4   via   PI3K/Akt/eNOS   signal  transduction  pathway.  J  Cardiovasc  Pharmacol,  2007.  50(3):  p.  274-­‐80.  

163.   Melillo,  G.,  et  al.,  A  hypoxia-­‐responsive  element  mediates  a  novel  pathway  of  activation  of  the  inducible  nitric  oxide  synthase  promoter.   J  Exp  Med,  1995.  182(6):  p.  1683-­‐93.  

164.   Arnet,   U.A.,   et   al.,   Regulation   of   endothelial   nitric-­‐oxide   synthase   during  hypoxia.  J  Biol  Chem,  1996.  271(25):  p.  15069-­‐73.  

165.   Graham,   C.H.,   et   al.,   Hypoxia-­‐mediated   stimulation   of   carcinoma   cell  invasiveness  via  upregulation  of  urokinase  receptor  expression.  Int  J  Cancer,  1999.  80(4):  p.  617-­‐23.  

166.   Rofstad,   E.K.   and   T.   Danielsen,   Hypoxia-­‐induced   metastasis   of   human  melanoma   cells:   involvement   of   vascular   endothelial   growth   factor-­‐mediated  angiogenesis.  Br  J  Cancer,  1999.  80(11):  p.  1697-­‐707.  

167.   Pez,   F.,   et   al.,  The  HIF-­‐1-­‐inducible   lysyl   oxidase  activates  HIF-­‐1   via   the  Akt  pathway   in   a   positive   regulation   loop   and   synergizes   with   HIF-­‐1   in  promoting  tumor  cell  growth.  Cancer  Res,  2011.  71(5):  p.  1647-­‐57.  

168.   Ji,   F.,   et   al.,   Hypoxia   inducible   factor   1alpha-­‐mediated   LOX   expression  correlates   with   migration   and   invasion   in   epithelial   ovarian   cancer.   Int   J  Oncol,  2013.  42(5):  p.  1578-­‐88.  

169.   Ploug,   M.,   et   al.,   Cellular   receptor   for   urokinase   plasminogen   activator.  Carboxyl-­‐terminal   processing   and   membrane   anchoring   by   glycosyl-­‐phosphatidylinositol.  J  Biol  Chem,  1991.  266(3):  p.  1926-­‐33.  

170.   Ploug,  M.,  et  al.,  Protein  structure  and  membrane  anchorage  of  the  cellular  receptor  for  urokinase-­‐type  plasminogen  activator.  Semin  Thromb  Hemost,  1991.  17(3):  p.  183-­‐93.  

171.   Plesner,  T.,  N.  Behrendt,  and  M.  Ploug,  Structure,  function  and  expression  on  blood   and   bone  marrow   cells   of   the   urokinase-­‐type   plasminogen   activator  receptor,  uPAR.  Stem  Cells,  1997.  15(6):  p.  398-­‐408.  

172.   Ploug,   M.   and   V.   Ellis,   Structure-­‐function   relationships   in   the   receptor   for  urokinase-­‐type  plasminogen  activator.  Comparison  to  other  members  of  the  Ly-­‐6  family  and  snake  venom  alpha-­‐neurotoxins.  FEBS  Lett,  1994.  349(2):  p.  163-­‐8.  

173.   Xu,   X.,   et   al.,   Crystal   structure   of   the   urokinase   receptor   in   a   ligand-­‐free  form.  J  Mol  Biol,  2012.  416(5):  p.  629-­‐41.  

 

 

109  

174.   Montuori,   N.,   et   al.,   Soluble   and   cleaved   forms   of   the   urokinase-­‐receptor:  degradation  products  or  active  molecules?   Thromb  Haemost,   2005.   93(2):  p.  192-­‐8.  

175.   Min,   H.Y.,   et   al.,   Urokinase   receptor   antagonists   inhibit   angiogenesis   and  primary   tumor   growth   in   syngeneic   mice.   Cancer   Res,   1996.   56(10):   p.  2428-­‐33.  

176.   Chapman,   H.A.,   Plasminogen   activators,   integrins,   and   the   coordinated  regulation  of  cell  adhesion  and  migration.  Curr  Opin  Cell  Biol,  1997.  9(5):  p.  714-­‐24.  

177.   Ossowski,   L.   and   J.A.   Aguirre-­‐Ghiso,   Urokinase   receptor   and   integrin  partnership:   coordination   of   signaling   for   cell   adhesion,   migration   and  growth.  Curr  Opin  Cell  Biol,  2000.  12(5):  p.  613-­‐20.  

178.   Waltz,   D.A.,   et   al.,   Plasmin   and   plasminogen   activator   inhibitor   type   1  promote   cellular   motility   by   regulating   the   interaction   between   the  urokinase  receptor  and  vitronectin.  J  Clin  Invest,  1997.  100(1):  p.  58-­‐67.  

179.   Mazar,   A.P.,   Urokinase   plasminogen   activator   receptor   choreographs  multiple   ligand   interactions:   implications   for   tumor   progression   and  therapy.  Clin  Cancer  Res,  2008.  14(18):  p.  5649-­‐55.  

180.   Lester,   R.D.,   et   al.,   uPAR   induces   epithelial-­‐mesenchymal   transition   in  hypoxic  breast  cancer  cells.  J  Cell  Biol,  2007.  178(3):  p.  425-­‐36.  

181.   Jo,  M.,  et  al.,  Epidermal  growth  factor  receptor-­‐dependent  and  -­‐independent  cell-­‐signaling   pathways   originating   from   the   urokinase   receptor.   J   Biol  Chem,  2003.  278(3):  p.  1642-­‐6.  

182.   Liu,  D.,  et  al.,  EGFR  is  a  transducer  of  the  urokinase  receptor  initiated  signal  that  is  required  for  in  vivo  growth  of  a  human  carcinoma.  Cancer  Cell,  2002.  1(5):  p.  445-­‐57.  

183.   Huai,  Q.,   et   al.,  Crystal  structures  of   two  human  vitronectin,  urokinase  and  urokinase  receptor  complexes.  Nat  Struct  Mol  Biol,  2008.  15(4):  p.  422-­‐3.  

184.   Skriver,   L.,   et   al.,   Plasminogen   activator   released   as   inactive   proenzyme  from   murine   cells   transformed   by   sarcoma   virus.   Eur   J   Biochem,   1982.  124(2):  p.  409-­‐14.  

185.   Cohen,   R.L.,   et   al.,   Effects   of   urokinase   receptor   occupancy   on   plasmin  generation   and   proteolysis   of   basement  membrane   by   human   tumor   cells.  Blood,  1991.  78(2):  p.  479-­‐87.  

186.   Blasi,  F.  and  P.  Carmeliet,  uPAR:  a  versatile  signalling  orchestrator.  Nat  Rev  Mol  Cell  Biol,  2002.  3(12):  p.  932-­‐43.  

187.   Nielsen,   L.S.,   et   al.,  Purification  of  zymogen   to  plasminogen  activator   from  human   glioblastoma   cells   by   affinity   chromatography   with   monoclonal  antibody.  Biochemistry,  1982.  21(25):  p.  6410-­‐5.  

188.   Roldan,   A.L.,   et   al.,   Cloning   and   expression   of   the   receptor   for   human  urokinase  plasminogen  activator,  a  central  molecule  in  cell  surface,  plasmin  dependent  proteolysis.  EMBO  J,  1990.  9(2):  p.  467-­‐74.  

189.   Behrendt,   N.,   et   al.,   The   human   receptor   for   urokinase   plasminogen  activator.   NH2-­‐terminal   amino   acid   sequence   and   glycosylation   variants.   J  Biol  Chem,  1990.  265(11):  p.  6453-­‐60.  

 

 

110  

190.   Carmeliet,   P.,   et   al.,   Urokinase-­‐generated   plasmin   activates   matrix  metalloproteinases  during  aneurysm  formation.  Nat  Genet,  1997.  17(4):  p.  439-­‐44.  

191.   Estreicher,  A.,  et  al.,  The  receptor  for  urokinase  type  plasminogen  activator  polarizes   expression   of   the   protease   to   the   leading   edge   of   migrating  monocytes  and  promotes  degradation  of  enzyme  inhibitor  complexes.   J   Cell  Biol,  1990.  111(2):  p.  783-­‐92.  

192.   Pyke,   C.,   et   al.,   Receptor   for   urokinase   is   present   in   tumor-­‐associated  macrophages  in  ductal  breast  carcinoma.  Cancer  Res,  1993.  53(8):  p.  1911-­‐5.  

193.   Weaver,   A.M.,   et   al.,  Embryonic   fibroblasts   that  are  genetically  deficient   in  low   density   lipoprotein   receptor-­‐related   protein   demonstrate   increased  activity   of   the   urokinase   receptor   system   and   accelerated   migration   on  vitronectin.  J  Biol  Chem,  1997.  272(22):  p.  14372-­‐9.  

194.   Crowley,  C.W.,  et  al.,  Prevention  of  metastasis  by  inhibition  of  the  urokinase  receptor.  Proc  Natl  Acad  Sci  U  S  A,  1993.  90(11):  p.  5021-­‐5.  

195.   Pyke,   C.,   et   al.,   An   alternatively   spliced   variant   of   mRNA   for   the   human  receptor  for  urokinase  plasminogen  activator.  FEBS  Lett,  1993.  326(1-­‐3):  p.  69-­‐74.  

196.   Fazioli,   F.   and   F.   Blasi,   Urokinase-­‐type   plasminogen   activator   and   its  receptor:   new   targets   for   anti-­‐metastatic   therapy?   Trends   Pharmacol   Sci,  1994.  15(1):  p.  25-­‐9.  

197.   Aguirre  Ghiso,  J.A.,  K.  Kovalski,  and  L.  Ossowski,  Tumor  dormancy  induced  by   downregulation   of   urokinase   receptor   in   human   carcinoma   involves  integrin  and  MAPK  signaling.  J  Cell  Biol,  1999.  147(1):  p.  89-­‐104.  

198.   Ahmed,   N.,   et   al.,   Downregulation   of   urokinase   plasminogen   activator  receptor  expression  inhibits  Erk  signalling  with  concomitant  suppression  of  invasiveness   due   to   loss   of   uPAR-­‐beta1   integrin   complex   in   colon   cancer  cells.  Br  J  Cancer,  2003.  89(2):  p.  374-­‐84.  

199.   Xue,   W.,   et   al.,   Urokinase-­‐type   plasminogen   activator   receptors   associate  with   beta1   and   beta3   integrins   of   fibrosarcoma   cells:   dependence   on  extracellular  matrix  components.  Cancer  Res,  1997.  57(9):  p.  1682-­‐9.  

200.   Degryse,   B.,   et   al.,   Urokinase/urokinase   receptor   and  vitronectin/alpha(v)beta(3)   integrin   induce   chemotaxis   and   cytoskeleton  reorganization   through   different   signaling   pathways.   Oncogene,   2001.  20(16):  p.  2032-­‐43.  

201.   Todd,   R.F.,   3rd   and  H.R.   Petty,  Beta  2   (CD11/CD18)   integrins  can  serve  as  signaling   partners   for   other   leukocyte   receptors.   J   Lab   Clin   Med,   1997.  129(5):  p.  492-­‐8.  

202.   Bohuslav,   J.,   et   al.,   Urokinase   plasminogen   activator   receptor,   beta   2-­‐integrins,   and   Src-­‐kinases   within   a   single   receptor   complex   of   human  monocytes.  J  Exp  Med,  1995.  181(4):  p.  1381-­‐90.  

203.   Wei,   Y.,   et   al.,   Regulation   of   integrin   function   by   the   urokinase   receptor.  Science,  1996.  273(5281):  p.  1551-­‐5.  

 

 

111  

204.   Degryse,  B.,  et  al.,  Domain  2  of  the  urokinase  receptor  contains  an  integrin-­‐interacting   epitope   with   intrinsic   signaling   activity:   generation   of   a   new  integrin  inhibitor.  J  Biol  Chem,  2005.  280(26):  p.  24792-­‐803.  

205.   Gargiulo,   L.,   et   al.,   Cross-­‐talk   between   fMLP   and   vitronectin   receptors  triggered  by  urokinase  receptor-­‐derived  SRSRY  peptide.   J  Biol   Chem,  2005.  280(26):  p.  25225-­‐32.  

206.   Chaurasia,  P.,  et  al.,  A  region  in  urokinase  plasminogen  receptor  domain  III  controlling   a   functional   association   with   alpha5beta1   integrin   and   tumor  growth.  J  Biol  Chem,  2006.  281(21):  p.  14852-­‐63.  

207.   Smith,  H.W.  and  C.J.  Marshall,  Regulation  of  cell  signalling  by  uPAR.  Nat  Rev  Mol  Cell  Biol,  2010.  11(1):  p.  23-­‐36.  

208.   Madsen,   C.D.,   et   al.,  uPAR-­‐induced  cell  adhesion  and  migration:  vitronectin  provides  the  key.  J  Cell  Biol,  2007.  177(5):  p.  927-­‐39.  

209.   Ganesh,  S.,  et  al.,  Urokinase  receptor  and  colorectal  cancer  survival.  Lancet,  1994.  344(8919):  p.  401-­‐2.  

210.   Maity,   A.   and   D.   Solomon,   Both   increased   stability   and   transcription  contribute  to  the  induction  of  the  urokinase  plasminogen  activator  receptor  (uPAR)  message  by  hypoxia.  Exp  Cell  Res,  2000.  255(2):  p.  250-­‐7.  

211.   Wilhelm,   O.G.,   et   al.,   Cellular   glycosylphosphatidylinositol-­‐specific  phospholipase   D   regulates   urokinase   receptor   shedding   and   cell   surface  expression.  J  Cell  Physiol,  1999.  180(2):  p.  225-­‐35.  

212.   Andolfo,  A.,   et   al.,  Metalloproteases  cleave  the  urokinase-­‐type  plasminogen  activator   receptor   in   the   D1-­‐D2   linker   region   and   expose   epitopes   not  present   in   the   intact   soluble   receptor.   Thromb   Haemost,   2002.   88(2):   p.  298-­‐306.  

213.   Holst-­‐Hansen,   C.,   et   al.,   Soluble   urokinase   receptor   released   from   human  carcinoma   cells:   a   plasma   parameter   for   xenograft   tumour   studies.   Br   J  Cancer,  1999.  81(2):  p.  203-­‐11.  

214.   Sier,   C.F.,   et   al.,  Presence  of  urokinase-­‐type  plasminogen  activator  receptor  in   urine   of   cancer   patients   and   its   possible   clinical   relevance.   Lab   Invest,  1999.  79(6):  p.  717-­‐22.  

215.   Brunner,   N.,   et   al.,  The  urokinase  plasminogen  activator   receptor   in  blood  from  healthy  individuals  and  patients  with  cancer.  APMIS,  1999.  107(1):  p.  160-­‐7.  

216.   Bene,   M.C.,   et   al.,   CD87   (urokinase-­‐type   plasminogen   activator   receptor),  function   and   pathology   in   hematological   disorders:   a   review.   Leukemia,  2004.  18(3):  p.  394-­‐400.  

217.   Rasch,  M.G.,  et  al.,  Intact  and  cleaved  uPAR  forms:  diagnostic  and  prognostic  value  in  cancer.  Front  Biosci,  2008.  13:  p.  6752-­‐62.  

218.   Stephens,   R.W.,   et   al.,   Plasma   urokinase   receptor   levels   in   patients   with  colorectal   cancer:   relationship   to   prognosis.   J   Natl   Cancer   Inst,   1999.  91(10):  p.  869-­‐74.  

219.   Guertin,  D.A.  and  D.M.  Sabatini,  Defining  the  role  of  mTOR  in  cancer.  Cancer  Cell,  2007.  12(1):  p.  9-­‐22.  

220.   Frias,  M.A.,   et   al.,  mSin1   is  necessary   for  Akt/PKB  phosphorylation,  and   its  isoforms  define  three  distinct  mTORC2s.  Curr  Biol,  2006.  16(18):  p.  1865-­‐70.  

 

 

112  

221.   Browne,  G.J.   and  C.G.   Proud,  A  novel  mTOR-­‐regulated  phosphorylation  site  in   elongation   factor   2   kinase   modulates   the   activity   of   the   kinase   and   its  binding  to  calmodulin.  Mol  Cell  Biol,  2004.  24(7):  p.  2986-­‐97.  

222.   Schneider,  A.,  R.H.  Younis,  and   J.S.  Gutkind,  Hypoxia-­‐induced  energy  stress  inhibits  the  mTOR  pathway  by  activating  an  AMPK/REDD1  signaling  axis  in  head  and  neck  squamous  cell  carcinoma.  Neoplasia,  2008.  10(11):  p.  1295-­‐302.  

223.   Huang,   J.,  et  al.,  The  TSC1-­‐TSC2  complex  is  required  for  proper  activation  of  mTOR  complex  2.  Mol  Cell  Biol,  2008.  28(12):  p.  4104-­‐15.  

224.   Liu,  L.,  et  al.,  Hypoxia-­‐induced  energy  stress  regulates  mRNA  translation  and  cell  growth.  Mol  Cell,  2006.  21(4):  p.  521-­‐31.  

225.   Yang,   Q.,   et   al.,   TSC1/TSC2   and   Rheb   have   different   effects   on   TORC1   and  TORC2  activity.  Proc  Natl  Acad  Sci  U  S  A,  2006.  103(18):  p.  6811-­‐6.  

226.   Long,  X.,  et  al.,  Rheb  binding  to  mammalian  target  of  rapamycin  (mTOR)  is  regulated  by  amino  acid  sufficiency.  J  Biol  Chem,  2005.  280(25):  p.  23433-­‐6.  

227.   Brugarolas,  J.,  et  al.,  Regulation  of  mTOR  function  in  response  to  hypoxia  by  REDD1   and   the   TSC1/TSC2   tumor   suppressor   complex.   Genes   Dev,   2004.  18(23):  p.  2893-­‐904.  

228.   Reiling,   J.H.   and   E.   Hafen,   The   hypoxia-­‐induced   paralogs   Scylla   and  Charybdis   inhibit  growth  by  down-­‐regulating  S6K  activity  upstream  of  TSC  in  Drosophila.  Genes  Dev,  2004.  18(23):  p.  2879-­‐92.  

229.   Sofer,   A.,   et   al.,  Regulation  of  mTOR  and  cell  growth   in  response  to  energy  stress  by  REDD1.  Mol  Cell  Biol,  2005.  25(14):  p.  5834-­‐45.  

230.   DeYoung,  M.P.,  et  al.,  Hypoxia  regulates  TSC1/2-­‐mTOR  signaling  and  tumor  suppression   through   REDD1-­‐mediated   14-­‐3-­‐3   shuttling.   Genes   Dev,   2008.  22(2):  p.  239-­‐51.  

231.   Bernardi,   R.,   et   al.,   PML   inhibits   HIF-­‐1alpha   translation   and  neoangiogenesis  through  repression  of  mTOR.  Nature,  2006.  442(7104):  p.  779-­‐85.  

232.   Li,   Y.,   et   al.,  Bnip3  mediates  the  hypoxia-­‐induced  inhibition  on  mammalian  target  of  rapamycin  by  interacting  with  Rheb.  J  Biol  Chem,  2007.  282(49):  p.  35803-­‐13.  

233.   Arsham,  A.M.,  J.J.  Howell,  and  M.C.  Simon,  A  novel  hypoxia-­‐inducible  factor-­‐independent   hypoxic   response   regulating  mammalian   target   of   rapamycin  and  its  targets.  J  Biol  Chem,  2003.  278(32):  p.  29655-­‐60.  

234.   Magagnin,   M.G.,   et   al.,  The  mTOR   target  4E-­‐BP1   contributes   to  differential  protein  expression  during  normoxia  and  hypoxia  through  changes  in  mRNA  translation  efficiency.  Proteomics,  2008.  8(5):  p.  1019-­‐28.  

235.   van   den   Beucken,   T.,   M.   Koritzinsky,   and   B.G.   Wouters,   Translational  control  of  gene  expression  during  hypoxia.  Cancer  Biol  Ther,  2006.  5(7):  p.  749-­‐55.  

236.   Koritzinsky,  M.,  et  al.,  Gene  expression  during  acute  and  prolonged  hypoxia  is  regulated  by  distinct  mechanisms  of  translational  control.  EMBO  J,  2006.  25(5):  p.  1114-­‐25.  

237.   Macejak,  D.G.  and  P.  Sarnow,   Internal  initiation  of  translation  mediated  by  the  5'  leader  of  a  cellular  mRNA.  Nature,  1991.  353(6339):  p.  90-­‐4.  

 

 

113  

238.   Stoneley,   M.   and   A.E.   Willis,   Cellular   internal   ribosome   entry   segments:  structures,   trans-­‐acting   factors   and   regulation   of   gene   expression.  Oncogene,  2004.  23(18):  p.  3200-­‐7.  

239.   Ron,  D.,  Translational  control  in  the  endoplasmic  reticulum  stress  response.  J  Clin  Invest,  2002.  110(10):  p.  1383-­‐8.  

240.   Zhang,   K.   and   R.J.   Kaufman,   Protein   folding   in   the   endoplasmic   reticulum  and   the   unfolded   protein   response.   Handb   Exp   Pharmacol,   2006(172):   p.  69-­‐91.  

241.   Urano,   F.,   et   al.,   Coupling   of   stress   in   the   ER   to   activation   of   JNK   protein  kinases  by  transmembrane  protein  kinase  IRE1.   Science,  2000.  287(5453):  p.  664-­‐6.  

242.   Koritzinsky,  M.  and  B.G.  Wouters,  Hypoxia  and  regulation  of  messenger  RNA  translation.  Methods  Enzymol,  2007.  435:  p.  247-­‐73.  

243.   Blais,  J.D.,  et  al.,  Activating  transcription  factor  4  is  translationally  regulated  by  hypoxic  stress.  Mol  Cell  Biol,  2004.  24(17):  p.  7469-­‐82.  

244.   Bi,  M.,  et  al.,  ER  stress-­‐regulated  translation  increases  tolerance  to  extreme  hypoxia  and  promotes  tumor  growth.  EMBO  J,  2005.  24(19):  p.  3470-­‐81.  

245.   Rzymski,  T.   and  A.L.  Harris,  The  unfolded  protein  response  and  integrated  stress  response  to  anoxia.  Clin  Cancer  Res,  2007.  13(9):  p.  2537-­‐40.  

246.   Moenner,   M.,   et   al.,   Integrated   endoplasmic   reticulum   stress   responses   in  cancer.  Cancer  Res,  2007.  67(22):  p.  10631-­‐4.  

247.   Bertolotti,  A.,  et  al.,  Dynamic  interaction  of  BiP  and  ER  stress  transducers  in  the  unfolded-­‐protein  response.  Nat  Cell  Biol,  2000.  2(6):  p.  326-­‐32.  

248.   Wu,   J.   and  R.J.   Kaufman,  From  acute  ER  stress  to  physiological  roles  of   the  Unfolded  Protein  Response.  Cell  Death  Differ,  2006.  13(3):  p.  374-­‐84.  

249.   Flynn,  G.C.,  T.G.  Chappell,  and  J.E.  Rothman,  Peptide  binding  and  release  by  proteins   implicated   as   catalysts   of   protein   assembly.   Science,   1989.  245(4916):  p.  385-­‐90.  

250.   Blond-­‐Elguindi,   S.,   et   al.,   Peptide-­‐dependent   stimulation   of   the   ATPase  activity   of   the   molecular   chaperone   BiP   is   the   result   of   conversion   of  oligomers  to  active  monomers.  J  Biol  Chem,  1993.  268(17):  p.  12730-­‐5.  

251.   Hellman,   R.,   et   al.,   The   in   vivo   association   of   BiP   with   newly   synthesized  proteins  is  dependent  on  the  rate  and  stability  of  folding  and  not  simply  on  the  presence  of  sequences  that  can  bind  to  BiP.   J  Cell  Biol,  1999.  144(1):  p.  21-­‐30.  

252.   Gardner,  B.M.   and  P.  Walter,  Unfolded  proteins  are  Ire1-­‐activating  ligands  that   directly   induce   the   unfolded   protein   response.   Science,   2011.  333(6051):  p.  1891-­‐4.  

253.   Shamu,  C.E.  and  P.  Walter,  Oligomerization  and  phosphorylation  of  the  Ire1p  kinase  during  intracellular  signaling  from  the  endoplasmic  reticulum  to  the  nucleus.  EMBO  J,  1996.  15(12):  p.  3028-­‐39.  

254.   Calfon,   M.,   et   al.,   IRE1   couples   endoplasmic   reticulum   load   to   secretory  capacity  by  processing  the  XBP-­‐1  mRNA.  Nature,  2002.  415(6867):  p.  92-­‐6.  

255.   Koong,   A.C.,   et   al.,   The   regulation   of   GRP78   and  messenger   RNA   levels   by  hypoxia   is  modulated  by  protein  kinase  C  activators  and   inhibitors.   Radiat  Res,  1994.  138(1  Suppl):  p.  S60-­‐3.  

 

 

114  

256.   Graven,   K.K.,   et   al.,   Identification   of   protein   disulfide   isomerase   as   an  endothelial  hypoxic  stress  protein.  Am  J  Physiol  Lung  Cell  Mol  Physiol,  2002.  282(5):  p.  L996-­‐1003.  

257.   Nishitoh,   H.,   et   al.,   ASK1   is   essential   for   endoplasmic   reticulum   stress-­‐induced  neuronal   cell   death   triggered  by   expanded  polyglutamine   repeats.  Genes  Dev,  2002.  16(11):  p.  1345-­‐55.  

258.   Matsuzawa,   A.,   et   al.,   Physiological   roles   of   ASK1-­‐mediated   signal  transduction   in  oxidative  stress-­‐  and  endoplasmic  reticulum  stress-­‐induced  apoptosis:   advanced   findings   from   ASK1   knockout   mice.   Antioxid   Redox  Signal,  2002.  4(3):  p.  415-­‐25.  

259.   Cheng,  H.  and  T.  Force,  Molecular  mechanisms  of  cardiovascular  toxicity  of  targeted  cancer  therapeutics.  Circ  Res,  2010.  106(1):  p.  21-­‐34.  

260.   Yoneda,   T.,   et   al.,   Activation   of   caspase-­‐12,   an   endoplastic   reticulum   (ER)  resident   caspase,   through   tumor  necrosis   factor   receptor-­‐associated   factor  2-­‐dependent   mechanism   in   response   to   the   ER   stress.   J   Biol   Chem,   2001.  276(17):  p.  13935-­‐40.  

261.   Hetz,   C.,   et   al.,   Proapoptotic   BAX   and   BAK  modulate   the   unfolded   protein  response  by  a  direct  interaction  with  IRE1alpha.  Science,  2006.  312(5773):  p.  572-­‐6.  

262.   Agostinis,   P.,   Endoplasmic   reticulum   stress   in   health   and   disease.   1st   ed.  2012,  New  York:  Springer.  

263.   Lin,   J.H.,   et   al.,   IRE1   signaling  affects   cell   fate  during   the  unfolded  protein  response.  Science,  2007.  318(5852):  p.  944-­‐9.  

264.   Cross,   B.C.,   et   al.,   The   molecular   basis   for   selective   inhibition   of  unconventional  mRNA  splicing  by  an  IRE1-­‐binding  small  molecule.  Proc  Natl  Acad  Sci  U  S  A,  2012.  109(15):  p.  E869-­‐78.  

265.   Cojocari,   D.,   et   al.,   New   small   molecule   inhibitors   of   UPR   activation  demonstrate   that   PERK,   but   not   IRE1alpha   signaling   is   essential   for  promoting  adaptation  and  survival  to  hypoxia.  Radiother  Oncol,  2013.  

266.   Romero-­‐Ramirez,   L.,   et   al.,   XBP1   is   essential   for   survival   under   hypoxic  conditions  and   is  required   for  tumor  growth.   Cancer  Res,   2004.   64(17):   p.  5943-­‐7.  

267.   Chakrabarti,   A.,   A.W.   Chen,   and   J.D.   Varner,   A   review   of   the   mammalian  unfolded  protein  response.  Biotechnol  Bioeng,  2011.  108(12):  p.  2777-­‐93.  

268.   Zhu,   C.,   F.E.   Johansen,   and   R.   Prywes,   Interaction   of   ATF6   and   serum  response  factor.  Mol  Cell  Biol,  1997.  17(9):  p.  4957-­‐66.  

269.   Haze,   K.,   et   al.,  Mammalian   transcription   factor   ATF6   is   synthesized   as   a  transmembrane   protein   and   activated   by   proteolysis   in   response   to  endoplasmic  reticulum  stress.  Mol  Biol  Cell,  1999.  10(11):  p.  3787-­‐99.  

270.   Thuerauf,  D.J.,  et  al.,  Coordination  of  ATF6-­‐mediated  transcription  and  ATF6  degradation  by  a  domain  that   is  shared  with  the  viral   transcription  factor,  VP16.  J  Biol  Chem,  2002.  277(23):  p.  20734-­‐9.  

271.   Thuerauf,   D.J.,   et   al.,   Effects   of   the   isoform-­‐specific   characteristics   of  ATF6  alpha   and   ATF6   beta   on   endoplasmic   reticulum   stress   response   gene  expression  and  cell  viability.  J  Biol  Chem,  2007.  282(31):  p.  22865-­‐78.  

 

 

115  

272.   Regier,   J.L.,   F.   Shen,   and   S.J.   Triezenberg,   Pattern   of   aromatic   and  hydrophobic   amino   acids   critical   for   one   of   two   subdomains   of   the   VP16  transcriptional  activator.  Proc  Natl  Acad  Sci  U  S  A,  1993.  90(3):  p.  883-­‐7.  

273.   Yoshida,   H.,   et   al.,   Identification   of   the   cis-­‐acting   endoplasmic   reticulum  stress   response   element   responsible   for   transcriptional   induction   of  mammalian  glucose-­‐regulated  proteins.  Involvement  of  basic  leucine  zipper  transcription  factors.  J  Biol  Chem,  1998.  273(50):  p.  33741-­‐9.  

274.   Yoshida,   H.,   et   al.,   XBP1  mRNA   is   induced   by   ATF6   and   spliced   by   IRE1   in  response   to   ER   stress   to   produce   a   highly   active   transcription   factor.   Cell,  2001.  107(7):  p.  881-­‐91.  

275.   Nadanaka,  S.,  et  al.,  Role  of  disulfide  bridges  formed  in  the  luminal  domain  of  ATF6  in  sensing  endoplasmic  reticulum  stress.  Mol  Cell  Biol,  2007.  27(3):  p.  1027-­‐43.  

276.   Yamamoto,   K.,   et   al.,   Differential   contributions   of   ATF6   and   XBP1   to   the  activation   of   endoplasmic   reticulum   stress-­‐responsive   cis-­‐acting   elements  ERSE,  UPRE  and  ERSE-­‐II.  J  Biochem,  2004.  136(3):  p.  343-­‐50.  

277.   Yoshida,  H.,  et  al.,  ATF6  activated  by  proteolysis  binds  in  the  presence  of  NF-­‐Y   (CBF)   directly   to   the   cis-­‐acting   element   responsible   for   the   mammalian  unfolded  protein  response.  Mol  Cell  Biol,  2000.  20(18):  p.  6755-­‐67.  

278.   Rutkowski,   D.T.   and   R.J.   Kaufman,   A   trip   to   the   ER:   coping   with   stress.  Trends  Cell  Biol,  2004.  14(1):  p.  20-­‐8.  

279.   Ron,   D.   and   P.   Walter,   Signal   integration   in   the   endoplasmic   reticulum  unfolded   protein   response.   Nature   reviews.   Molecular   cell   biology,   2007.  8(7):  p.  519-­‐29.  

280.   Harding,   H.P.,   et   al.,   An   integrated   stress   response   regulates   amino   acid  metabolism  and  resistance  to  oxidative  stress.  Mol  Cell,  2003.  11(3):  p.  619-­‐33.  

281.   Hochachka,   P.W.,   et   al.,   Unifying   theory   of   hypoxia   tolerance:  molecular/metabolic   defense   and   rescue  mechanisms   for   surviving   oxygen  lack.  Proc  Natl  Acad  Sci  U  S  A,  1996.  93(18):  p.  9493-­‐8.  

282.   Sutherland,  R.,  et  al.,  Cellular  growth  and  metabolic  adaptations  to  nutrient  stress   environments   in   tumor  microregions.   Int   J   Radiat   Oncol   Biol   Phys,  1986.  12(4):  p.  611-­‐5.  

283.   Koritzinsky,  M.,   et   al.,  The  hypoxic  proteome   is   influenced  by  gene-­‐specific  changes  in  mRNA  translation.  Radiother  Oncol,  2005.  76(2):  p.  177-­‐86.  

284.   Lu,   X.,   et   al.,   Radiation-­‐induced   changes   in   gene   expression   involve  recruitment   of   existing   messenger   RNAs   to   and   away   from   polysomes.  Cancer  Res,  2006.  66(2):  p.  1052-­‐61.  

285.   Kraggerud,   S.M.,   J.A.   Sandvik,   and   E.O.   Pettersen,   Regulation   of   protein  synthesis  in  human  cells  exposed  to  extreme  hypoxia.  Anticancer  Res,  1995.  15(3):  p.  683-­‐6.  

286.   Erler,   J.T.,   et   al.,   Hypoxia-­‐mediated   down-­‐regulation   of   Bid   and   Bax   in  tumors   occurs   via   hypoxia-­‐inducible   factor   1-­‐dependent   and   -­‐independent  mechanisms  and  contributes  to  drug  resistance.  Mol  Cell  Biol,  2004.  24(7):  p.  2875-­‐89.  

 

 

116  

287.   Wouters,   B.G.,   et   al.,  Control  of  the  hypoxic  response  through  regulation  of  mRNA  translation.  Semin  Cell  Dev  Biol,  2005.  16(4-­‐5):  p.  487-­‐501.  

288.   van   den   Beucken,   T.,   et   al.,   Hypoxia-­‐induced   expression   of   carbonic  anhydrase   9   is   dependent   on   the   unfolded   protein   response.   J   Biol   Chem,  2009.  284(36):  p.  24204-­‐12.  

289.   Vattem,  K.M.  and  R.C.  Wek,  Reinitiation  involving  upstream  ORFs  regulates  ATF4  mRNA  translation  in  mammalian  cells.  Proc  Natl  Acad  Sci  U  S  A,  2004.  101(31):  p.  11269-­‐74.  

290.   Harding,  H.P.,  et  al.,  Regulated  translation  initiation  controls  stress-­‐induced  gene  expression  in  mammalian  cells.  Mol  Cell,  2000.  6(5):  p.  1099-­‐108.  

291.   Schroder,  M.  and  R.J.  Kaufman,  ER  stress  and  the  unfolded  protein  response.  Mutat  Res,  2005.  569(1-­‐2):  p.  29-­‐63.  

292.   Rouschop,  K.M.,  et  al.,  The  unfolded  protein  response  protects  human  tumor  cells  during  hypoxia  through  regulation  of  the  autophagy  genes  MAP1LC3B  and  ATG5.  J  Clin  Invest,  2010.  120(1):  p.  127-­‐41.  

293.   Rouschop,  K.M.,  et  al.,  PERK/eIF2alpha  signaling  protects  therapy  resistant  hypoxic   cells   through   induction   of   glutathione   synthesis   and   protection  against  ROS.  Proc  Natl  Acad  Sci  U  S  A,  2013.  110(12):  p.  4622-­‐7.  

294.   Zinszner,  H.,  et  al.,  CHOP  is  implicated  in  programmed  cell  death  in  response  to  impaired  function  of  the  endoplasmic  reticulum.  Genes  Dev,  1998.  12(7):  p.  982-­‐95.  

295.   Novoa,   I.,   et   al.,   Feedback   inhibition   of   the   unfolded   protein   response   by  GADD34-­‐mediated  dephosphorylation  of  eIF2alpha.  J  Cell  Biol,  2001.  153(5):  p.  1011-­‐22.  

296.   Harding,   H.P.,   et   al.,   Transcriptional   and   translational   control   in   the  Mammalian  unfolded  protein  response.  Annu  Rev  Cell  Dev  Biol,  2002.  18:  p.  575-­‐99.  

297.   Anderson,  L.L.,   et   al.,  Survival  from  hypoxia  in  C.  elegans  by  inactivation  of  aminoacyl-­‐tRNA  synthetases.  Science,  2009.  323(5914):  p.  630-­‐3.  

298.   Koong,   A.C.,   et   al.,   Increased   cytotoxicity   of   chronic   hypoxic   cells   by  molecular  inhibition  of  GRP78  induction.  Int  J  Radiat  Oncol  Biol  Phys,  1994.  28(3):  p.  661-­‐6.  

299.   Bondareva,   A.,   et   al.,   The   lysyl   oxidase   inhibitor,   beta-­‐aminopropionitrile,  diminishes  the  metastatic  colonization  potential  of  circulating  breast  cancer  cells.  PLoS  One,  2009.  4(5):  p.  e5620.  

300.   Nishioka,   T.,   A.   Eustace,   and   C.   West,   Lysyl   oxidase:   from  basic   science   to  future  cancer  treatment.  Cell  Struct  Funct,  2012.  37(1):  p.  75-­‐80.  

301.   Erler,   J.T.   and   A.J.   Giaccia,   Lysyl   oxidase   mediates   hypoxic   control   of  metastasis.  Cancer  Res,  2006.  66(21):  p.  10238-­‐41.  

302.   Chen,   Z.,   et   al.,   Challenges   for   drug   discovery   -­‐   a   case   study   of   urokinase  receptor  inhibition.  Comb  Chem  High  Throughput  Screen,  2009.  12(10):  p.  961-­‐7.  

303.   Bauer,  T.W.,  et  al.,  Targeting  of  urokinase  plasminogen  activator  receptor  in  human   pancreatic   carcinoma   cells   inhibits   c-­‐Met-­‐   and   insulin-­‐like   growth  factor-­‐I   receptor-­‐mediated   migration   and   invasion   and   orthotopic   tumor  growth  in  mice.  Cancer  Res,  2005.  65(17):  p.  7775-­‐81.  

 

 

117  

304.   Kawaguchi,   S.   and   D.T.   Ng,   Cell   biology.   Sensing   ER   stress.   Science,   2011.  333(6051):  p.  1830-­‐1.  

305.   Onn,   A.   and   D.   Ron,  Modeling   the  endoplasmic   reticulum  unfolded  protein  response.  Nat  Struct  Mol  Biol,  2010.  17(8):  p.  924-­‐5.  

306.   Braakman,   I.   and   N.J.   Bulleid,   Protein   folding   and   modification   in   the  mammalian  endoplasmic  reticulum.  Annu  Rev  Biochem,  2011.  80:  p.  71-­‐99.  

307.   Marx,   J.,   Cell   biology.   A   stressful   situation.   Science,   2006.   313(5793):   p.  1564-­‐6.  

308.   Braakman,   I.,   J.   Helenius,   and   A.   Helenius,   Manipulating   disulfide   bond  formation  and  protein  folding  in  the  endoplasmic  reticulum.  EMBO   J,  1992.  11(5):  p.  1717-­‐22.  

309.   Marquardt,   T.,   D.N.   Hebert,   and   A.   Helenius,   Post-­‐translational   folding   of  influenza   hemagglutinin   in   isolated   endoplasmic   reticulum-­‐derived  microsomes.  J  Biol  Chem,  1993.  268(26):  p.  19618-­‐25.  

310.   Appenzeller-­‐Herzog,   C.   and   L.   Ellgaard,  The  human  PDI   family:   versatility  packed  into  a  single  fold.  Biochim  Biophys  Acta,  2008.  1783(4):  p.  535-­‐48.  

311.   Greenfield,   J.J.  and  S.  High,  The  Sec61  complex  is  located  in  both  the  ER  and  the   ER-­‐Golgi   intermediate   compartment.   J   Cell   Sci,   1999.   112   (   Pt   10):   p.  1477-­‐86.  

312.   Riemer,  J.,  N.  Bulleid,  and  J.M.  Herrmann,  Disulfide  formation  in  the  ER  and  mitochondria:   two   solutions   to   a   common   process.   Science,   2009.  324(5932):  p.  1284-­‐7.  

313.   Cumming,   R.C.,   et   al.,   Protein   disulfide   bond   formation   in   the   cytoplasm  during  oxidative  stress.  J  Biol  Chem,  2004.  279(21):  p.  21749-­‐58.  

314.   Hartl,  F.U.,  A.  Bracher,  and  M.  Hayer-­‐Hartl,  Molecular  chaperones  in  protein  folding  and  proteostasis.  Nature,  2011.  475(7356):  p.  324-­‐32.  

315.   Rudd,  P.M.  and  R.A.  Dwek,  Glycosylation:  heterogeneity  and  the  3D  structure  of  proteins.  Crit  Rev  Biochem  Mol  Biol,  1997.  32(1):  p.  1-­‐100.  

316.   Spiro,  R.G.,  Protein  glycosylation:  nature,  distribution,  enzymatic  formation,  and  disease  implications  of  glycopeptide  bonds.   Glycobiology,   2002.   12(4):  p.  43R-­‐56R.  

317.   Helenius,   A.   and   M.   Aebi,   Roles   of   N-­‐linked   glycans   in   the   endoplasmic  reticulum.  Annu  Rev  Biochem,  2004.  73:  p.  1019-­‐49.  

318.   Ellgaard,   L.   and  A.  Helenius,  Quality  control   in   the  endoplasmic  reticulum.  Nat  Rev  Mol  Cell  Biol,  2003.  4(3):  p.  181-­‐91.  

319.   Shailubhai,  K.,  M.A.  Pratta,  and  I.K.  Vijay,  Purification  and  characterization  of   glucosidase   I   involved   in   N-­‐linked   glycoprotein   processing   in   bovine  mammary  gland.  Biochem  J,  1987.  247(3):  p.  555-­‐62.  

320.   Brodsky,   J.L.   and   W.R.   Skach,   Protein   folding   and   quality   control   in   the  endoplasmic   reticulum:   Recent   lessons   from   yeast   and   mammalian   cell  systems.  Curr  Opin  Cell  Biol,  2011.  23(4):  p.  464-­‐75.  

321.   Feige,  M.J.   and   L.M.  Hendershot,  Disulfide  bonds   in  ER  protein   folding  and  homeostasis.  Curr  Opin  Cell  Biol,  2011.  23(2):  p.  167-­‐75.  

322.   Pelletier,   M.F.,   et   al.,   The   heterodimeric   structure   of   glucosidase   II   is  required   for   its   activity,   solubility,   and   localization   in   vivo.   Glycobiology,  2000.  10(8):  p.  815-­‐27.  

 

 

118  

323.   Gilbert,  H.F.,  Protein  disulfide  isomerase  and  assisted  protein  folding.   J  Biol  Chem,  1997.  272(47):  p.  29399-­‐402.  

324.   Tu,   B.P.   and   J.S.   Weissman,   Oxidative   protein   folding   in   eukaryotes:  mechanisms  and  consequences.  J  Cell  Biol,  2004.  164(3):  p.  341-­‐6.  

325.   Hatahet,   F.   and   L.W.   Ruddock,   Protein   disulfide   isomerase:   a   critical  evaluation  of  its  function  in  disulfide  bond  formation.  Antioxid  Redox  Signal,  2009.  11(11):  p.  2807-­‐50.  

326.   Horibe,  T.,   et  al.,  Different  contributions  of  the  three  CXXC  motifs  of  human  protein-­‐disulfide   isomerase-­‐related   protein   to   isomerase   activity   and  oxidative  refolding.  J  Biol  Chem,  2004.  279(6):  p.  4604-­‐11.  

327.   Chivers,  P.T.,  M.C.  Laboissiere,  and  R.T.  Raines,  The  CXXC  motif:  imperatives  for  the  formation  of  native  disulfide  bonds  in  the  cell.  EMBO  J,  1996.  15(11):  p.  2659-­‐67.  

328.   Sevier,   C.S.   and   C.A.   Kaiser,  Formation  and  transfer  of  disulphide  bonds   in  living  cells.  Nat  Rev  Mol  Cell  Biol,  2002.  3(11):  p.  836-­‐47.  

329.   Sevier,  C.S.  and  C.A.  Kaiser,  Ero1  and  redox  homeostasis  in  the  endoplasmic  reticulum.  Biochim  Biophys  Acta,  2008.  1783(4):  p.  549-­‐56.  

330.   Bulleid,   N.J.   and   L.   Ellgaard,   Multiple   ways   to   make   disulfides.   Trends  Biochem  Sci,  2011.  36(9):  p.  485-­‐92.  

331.   Lyles,   M.M.   and   H.F.   Gilbert,  Mutations   in   the   thioredoxin   sites   of   protein  disulfide   isomerase   reveal   functional   nonequivalence   of   the   N-­‐   and   C-­‐terminal  domains.  J  Biol  Chem,  1994.  269(49):  p.  30946-­‐52.  

332.   Kosuri,   P.,   et   al.,   Protein   folding   drives   disulfide   formation.   Cell,   2012.  151(4):  p.  794-­‐806.  

333.   Gleiter,   S.   and   J.C.   Bardwell,   Disulfide   bond   isomerization   in   prokaryotes.  Biochim  Biophys  Acta,  2008.  1783(4):  p.  530-­‐4.  

334.   Kersteen,  E.A.,   S.R.  Barrows,   and  R.T.  Raines,  Catalysis  of  protein  disulfide  bond   isomerization   in   a   homogeneous   substrate.   Biochemistry,   2005.  44(36):  p.  12168-­‐78.  

335.   Hotchkiss,   K.A.,   C.N.   Chesterman,   and   P.J.   Hogg,   Catalysis   of   disulfide  isomerization   in   thrombospondin   1   by   protein   disulfide   isomerase.  Biochemistry,  1996.  35(30):  p.  9761-­‐7.  

336.   Frand,  A.R.  and  C.A.  Kaiser,  The  ERO1  gene  of  yeast  is  required  for  oxidation  of  protein  dithiols  in  the  endoplasmic  reticulum.  Mol  Cell,  1998.  1(2):  p.  161-­‐70.  

337.   Pollard,  M.G.,  K.J.  Travers,  and  J.S.  Weissman,  Ero1p:  a  novel  and  ubiquitous  protein  with  an  essential  role  in  oxidative  protein  folding  in  the  endoplasmic  reticulum.  Mol  Cell,  1998.  1(2):  p.  171-­‐82.  

338.   Pagani,   M.,   et   al.,   Endoplasmic   reticulum   oxidoreductin   1-­‐lbeta   (ERO1-­‐Lbeta),  a  human  gene  induced  in  the  course  of  the  unfolded  protein  response.  J  Biol  Chem,  2000.  275(31):  p.  23685-­‐92.  

339.   Gross,   E.,   et   al.,   Structure   of   Ero1p,   source   of   disulfide  bonds   for   oxidative  protein  folding  in  the  cell.  Cell,  2004.  117(5):  p.  601-­‐10.  

340.   Frand,   A.R.,   J.W.   Cuozzo,   and   C.A.   Kaiser,  Pathways   for  protein  disulphide  bond  formation.  Trends  Cell  Biol,  2000.  10(5):  p.  203-­‐10.  

 

 

119  

341.   Frand,   A.R.   and   C.A.   Kaiser,  Two  pairs  of  conserved  cysteines  are  required  for  the  oxidative  activity  of  Ero1p  in  protein  disulfide  bond  formation  in  the  endoplasmic  reticulum.  Mol  Biol  Cell,  2000.  11(9):  p.  2833-­‐43.  

342.   Tu,  B.P.   and   J.S.  Weissman,  The  FAD-­‐  and  O(2)-­‐dependent  reaction  cycle  of  Ero1-­‐mediated  oxidative  protein   folding   in   the  endoplasmic  reticulum.   Mol  Cell,  2002.  10(5):  p.  983-­‐94.  

343.   Tu,   B.P.,   et   al.,   Biochemical   basis   of   oxidative   protein   folding   in   the  endoplasmic  reticulum.  Science,  2000.  290(5496):  p.  1571-­‐4.  

344.   Gross,   E.,   et   al.,  Generating  disulfides  enzymatically:  reaction  products  and  electron   acceptors   of   the   endoplasmic   reticulum   thiol   oxidase   Ero1p.   Proc  Natl  Acad  Sci  U  S  A,  2006.  103(2):  p.  299-­‐304.  

345.   Baker,   K.M.,   et   al.,   Low   reduction   potential   of   Ero1alpha   regulatory  disulphides   ensures   tight   control   of   substrate   oxidation.   EMBO   J,   2008.  27(22):  p.  2988-­‐97.  

346.   Wang,   L.,   L.   Zhu,   and   C.C.   Wang,   The   endoplasmic   reticulum   sulfhydryl  oxidase   Ero1beta   drives   efficient   oxidative   protein   folding   with   loose  regulation.  Biochem  J,  2011.  434(1):  p.  113-­‐21.  

347.   Bardwell,   J.C.,   et   al.,   A  pathway   for   disulfide   bond   formation   in   vivo.   Proc  Natl  Acad  Sci  U  S  A,  1993.  90(3):  p.  1038-­‐42.  

348.   Kobayashi,   T.,   et   al.,   Respiratory   chain   is   required   to   maintain   oxidized  states   of   the   DsbA-­‐DsbB   disulfide   bond   formation   system   in   aerobically  growing   Escherichia   coli   cells.   Proceedings   of   the   National   Academy   of  Sciences  of  the  United  States  of  America,  1997.  94(22):  p.  11857-­‐62.  

349.   Bader,  M.,  et  al.,  Oxidative  protein  folding  is  driven  by  the  electron  transport  system.  Cell,  1999.  98(2):  p.  217-­‐27.  

350.   Regeimbal,   J.  and  J.C.  Bardwell,  DsbB  catalyzes  disulfide  bond  formation  de  novo.  J  Biol  Chem,  2002.  277(36):  p.  32706-­‐13.  

351.   Kobayashi,   T.,   et   al.,   Respiratory   chain   is   required   to   maintain   oxidized  states   of   the   DsbA-­‐DsbB   disulfide   bond   formation   system   in   aerobically  growing   Escherichia   coli   cells.   Proc   Natl   Acad   Sci   U   S   A,   1997.   94(22):   p.  11857-­‐62.  

352.   Muyzer,   G.   and   A.J.   Stams,   The   ecology   and   biotechnology   of   sulphate-­‐reducing  bacteria.  Nat  Rev  Microbiol,  2008.  6(6):  p.  441-­‐54.  

353.   Sevier,  C.S.,  et  al.,  Modulation  of  cellular  disulfide-­‐bond  formation  and  the  ER  redox   environment   by   feedback   regulation   of   Ero1.   Cell,   2007.   129(2):   p.  333-­‐44.  

354.   Zito,   E.,   et   al.,   ERO1-­‐beta,   a   pancreas-­‐specific   disulfide   oxidase,   promotes  insulin  biogenesis  and  glucose  homeostasis.  J  Cell  Biol,  2010.  188(6):  p.  821-­‐32.  

355.   Appenzeller-­‐Herzog,  C.,  et  al.,  Disulphide  production  by  Ero1alpha-­‐PDI  relay  is  rapid  and  effectively  regulated.  EMBO  J,  2010.  29(19):  p.  3318-­‐29.  

356.   Zito,   E.,   et   al.,   Oxidative   protein   folding   by   an   endoplasmic   reticulum-­‐localized  peroxiredoxin.  Mol  Cell,  2010.  40(5):  p.  787-­‐97.  

357.   Zito,   E.,   et   al.,   Oxidative   protein   folding   by   an   endoplasmic   reticulum-­‐localized  peroxiredoxin.  Molecular  cell,  2010.  40(5):  p.  787-­‐97.  

 

 

120  

358.   Tavender,  T.J.,   J.J.  Springate,  and  N.J.  Bulleid,  Recycling  of  peroxiredoxin  IV  provides   a   novel   pathway   for   disulphide   formation   in   the   endoplasmic  reticulum.  The  EMBO  journal,  2010.  29(24):  p.  4185-­‐97.  

359.   Tavender,  T.J.,   J.J.  Springate,  and  N.J.  Bulleid,  Recycling  of  peroxiredoxin  IV  provides   a   novel   pathway   for   disulphide   formation   in   the   endoplasmic  reticulum.  EMBO  J,  2010.  29(24):  p.  4185-­‐97.  

360.   Zito,   E.,   PRDX4,   an   endoplasmic   reticulum-­‐localized   peroxiredoxin   at   the  crossroads  between  enzymatic   oxidative  protein   folding  and  nonenzymatic  protein  oxidation.  Antioxid  Redox  Signal,  2013.  18(13):  p.  1666-­‐74.  

361.   Tavender,  T.J.  and  N.J.  Bulleid,  Peroxiredoxin  IV  protects  cells  from  oxidative  stress  by  removing  H2O2  produced  during  disulphide   formation.   J   Cell   Sci,  2010.  123(Pt  15):  p.  2672-­‐9.  

362.   Schulman,   S.,   et   al.,   Vitamin   K   epoxide   reductase   prefers   ER   membrane-­‐anchored   thioredoxin-­‐like  redox  partners.   Proc   Natl   Acad   Sci   U   S   A,   2010.  107(34):  p.  15027-­‐32.  

363.   Wajih,  N.,  et  al.,  Engineering  of  a  recombinant  vitamin  K-­‐dependent  gamma-­‐carboxylation  system  with  enhanced  gamma-­‐carboxyglutamic  acid  forming  capacity:   evidence   for   a   functional   CXXC   redox   center   in   the   system.   J   Biol  Chem,  2005.  280(11):  p.  10540-­‐7.  

364.   Dutton,   R.J.,   et   al.,   Inhibition   of   bacterial   disulfide   bond   formation   by   the  anticoagulant  warfarin.  Proc  Natl  Acad  Sci  U  S  A,  2010.  107(1):  p.  297-­‐301.  

365.   Wajih,  N.,  S.M.  Hutson,  and  R.  Wallin,  Disulfide-­‐dependent  protein  folding  is  linked   to   operation   of   the   vitamin  K   cycle   in   the   endoplasmic   reticulum.   A  protein   disulfide   isomerase-­‐VKORC1   redox   enzyme   complex   appears   to   be  responsible  for  vitamin  K1  2,3-­‐epoxide  reduction.  The   Journal  of  biological  chemistry,  2007.  282(4):  p.  2626-­‐35.  

366.   Rutkevich,  L.A.  and  D.B.  Williams,  Vitamin  K  epoxide  reductase  contributes  to   protein   disulfide   formation   and   redox   homeostasis   within   the  endoplasmic  reticulum.  Molecular  biology  of  the  cell,  2012.  

367.   Raykhel,   I.,   et   al.,   A   molecular   specificity   code   for   the   three   mammalian  KDEL  receptors.  J  Cell  Biol,  2007.  179(6):  p.  1193-­‐204.  

368.   Nguyen,  V.D.,  et  al.,  Two  endoplasmic  reticulum  PDI  peroxidases  increase  the  efficiency  of  the  use  of  peroxide  during  disulfide  bond  formation.   J  Mol  Biol,  2011.  406(3):  p.  503-­‐15.  

369.   Bosello-­‐Travain,   V.,   et   al.,  Protein  disulfide   isomerase  and  glutathione  are  alternative   substrates   in   the   one   Cys   catalytic   cycle   of   glutathione  peroxidase  7.  Biochim  Biophys  Acta,  2013.  1830(6):  p.  3846-­‐57.  

370.   Wei,   P.C.,   et   al.,   Loss   of   the   oxidative   stress   sensor   NPGPx   compromises  GRP78   chaperone   activity   and   induces   systemic   disease.   Mol   Cell,   2012.  48(5):  p.  747-­‐59.  

371.   Alon,  A.,  et  al.,  The  dynamic  disulphide  relay  of  quiescin  sulphydryl  oxidase.  Nature,  2012.  488(7411):  p.  414-­‐8.  

372.   Chakravarthi,  S.,  et  al.,  Intracellular  catalysis  of  disulfide  bond  formation  by  the  human  sulfhydryl  oxidase,  QSOX1.  Biochem  J,  2007.  404(3):  p.  403-­‐11.  

 

 

121  

373.   Kodali,   V.K.   and   C.   Thorpe,   Oxidative   protein   folding   and   the   Quiescin-­‐sulfhydryl   oxidase   family   of   flavoproteins.   Antioxid   Redox   Signal,   2010.  13(8):  p.  1217-­‐30.  

374.   Mairet-­‐Coello,   G.,   et   al.,   FAD-­‐linked   sulfhydryl   oxidase   QSOX:   topographic,  cellular,   and   subcellular   immunolocalization   in   adult   rat   central   nervous  system.  J  Comp  Neurol,  2004.  473(3):  p.  334-­‐63.  

375.   Rancy,  P.C.   and  C.  Thorpe,  Oxidative  protein  folding  in  vitro:  a  study  of  the  cooperation   between   quiescin-­‐sulfhydryl   oxidase   and   protein   disulfide  isomerase.  Biochemistry,  2008.  47(46):  p.  12047-­‐56.  

376.   Rutkevich,  L.A.  and  D.B.  Williams,  Vitamin  K  epoxide  reductase  contributes  to   protein   disulfide   formation   and   redox   homeostasis   within   the  endoplasmic  reticulum.  Mol  Biol  Cell,  2012.  23(11):  p.  2017-­‐27.  

377.   Coppock,  D.,  et  al.,  Regulation  of  the  quiescence-­‐induced  genes:  quiescin  Q6,  decorin,  and  ribosomal  protein  S29.  Biochem  Biophys  Res  Commun,  2000.  269(2):  p.  604-­‐10.  

378.   Ilani,   T.,   et   al.,   A   secreted   disulfide   catalyst   controls   extracellular   matrix  composition  and  function.  Science,  2013.  341(6141):  p.  74-­‐6.  

379.   Jessop,   C.E.,   et   al.,   Protein   disulphide   isomerase   family   members   show  distinct  substrate  specificity:  P5  is  targeted  to  BiP  client  proteins.   J  Cell   Sci,  2009.  122(Pt  23):  p.  4287-­‐95.  

380.   Jessop,   C.E.,   et   al.,   ERp57   is   essential   for   efficient   folding   of   glycoproteins  sharing  common  structural  domains.  EMBO  J,  2007.  26(1):  p.  28-­‐40.  

381.   Rutkevich,   L.A.,   et   al.,   Functional   relationship   between   protein   disulfide  isomerase  family  members  during  the  oxidative  folding  of  human  secretory  proteins.  Mol  Biol  Cell,  2010.  21(18):  p.  3093-­‐105.  

382.   Chiu,  C.C.,  et  al.,  Glucose-­‐regulated  protein  78  regulates  multiple  malignant  phenotypes  in  head  and  neck  cancer  and  may  serve  as  a  molecular  target  of  therapeutic  intervention.  Mol  Cancer  Ther,  2008.  7(9):  p.  2788-­‐97.  

383.   Lin,   C.Y.,   et   al.,  Positive  association  of  glucose-­‐regulated  protein  78  during  oral   cancer   progression   and   the   prognostic   value   in   oral   precancerous  lesions.  Head  Neck,  2010.  32(8):  p.  1028-­‐39.  

384.   Tavender,  T.J.  and  N.J.  Bulleid,  Molecular  mechanisms  regulating  oxidative  activity   of   the   Ero1   family   in   the   endoplasmic   reticulum.   Antioxid   Redox  Signal,  2010.  13(8):  p.  1177-­‐87.  

385.   Cheretis,  C.,  et  al.,  Expression  of  ERp29,  an  endoplasmic  reticulum  secretion  factor  in  basal-­‐cell  carcinoma.  Am  J  Dermatopathol,  2006.  28(5):  p.  410-­‐2.  

386.   Gao,  D.,   et   al.,  ERp29  induces  breast  cancer  cell  growth  arrest  and  survival  through   modulation   of   activation   of   p38   and   upregulation   of   ER   stress  protein  p58IPK.  Lab  Invest,  2012.  92(2):  p.  200-­‐13.  

387.   Kutomi,  G.,  et  al.,  Human  endoplasmic  reticulum  oxidoreductin  1-­‐alpha  is  a  novel  predictor  for  poor  prognosis  of  breast  cancer.  Cancer  Sci,  2013.  

388.   Toquet,   C.,   et   al.,   Altered   Calreticulin   expression   in   human   colon   cancer:  maintenance   of   Calreticulin   expression   is   associated   with   mucinous  differentiation.  Oncol  Rep,  2007.  17(5):  p.  1101-­‐7.  

389.   Li,   F.,   et   al.,  Growth  factor  regulation  of   the  molecular  chaperone  calnexin.  Biochem  Biophys  Res  Commun,  2001.  289(3):  p.  725-­‐32.  

 

 

122  

390.   Li,   Z.   and  Z.   Li,  Glucose  regulated  protein  78:  a  critical   link  between  tumor  microenvironment   and   cancer   hallmarks.   Biochim   Biophys   Acta,   2012.  1826(1):  p.  13-­‐22.  

391.   Tamatani,   M.,   et   al.,   ORP150   protects   against   hypoxia/ischemia-­‐induced  neuronal  death.  Nat  Med,  2001.  7(3):  p.  317-­‐23.  

392.   Ozawa,   K.,   et   al.,   Regulation   of   tumor   angiogenesis   by   oxygen-­‐regulated  protein   150,   an   inducible   endoplasmic   reticulum   chaperone.   Cancer   Res,  2001.  61(10):  p.  4206-­‐13.  

393.   Roth,  J.,  Protein  N-­‐glycosylation  along  the  secretory  pathway:  relationship  to  organelle   topography   and   function,   protein   quality   control,   and   cell  interactions.  Chem  Rev,  2002.  102(2):  p.  285-­‐303.  

394.   Anelli,  T.  and  R.  Sitia,  Protein  quality  control  in  the  early  secretory  pathway.  EMBO  J,  2008.  27(2):  p.  315-­‐27.  

395.   Lederkremer,   G.Z.   and   M.H.   Glickman,   A   window   of   opportunity:   timing  protein  degradation  by  trimming  of  sugars  and  ubiquitins.  Trends  Biochem  Sci,  2005.  30(6):  p.  297-­‐303.  

396.   Hosokawa,   N.,   et   al.,   Human   OS-­‐9,   a   lectin   required   for   glycoprotein  endoplasmic   reticulum-­‐associated   degradation,   recognizes   mannose-­‐trimmed  N-­‐glycans.  J  Biol  Chem,  2009.  284(25):  p.  17061-­‐8.  

397.   Tsai,  B.,   Y.   Ye,   and  T.A.  Rapoport,  Retro-­‐translocation  of  proteins  from  the  endoplasmic  reticulum  into  the  cytosol.  Nat  Rev  Mol  Cell  Biol,  2002.  3(4):  p.  246-­‐55.  

398.   Kikkert,   M.,   et   al.,   Human   HRD1   is   an   E3   ubiquitin   ligase   involved   in  degradation  of  proteins  from  the  endoplasmic  reticulum.  J  Biol  Chem,  2004.  279(5):  p.  3525-­‐34.  

399.   Schulze,  A.,  et  al.,  The  ubiquitin-­‐domain  protein  HERP  forms  a  complex  with  components  of  the  endoplasmic  reticulum  associated  degradation  pathway.  J  Mol  Biol,  2005.  354(5):  p.  1021-­‐7.  

400.   Meusser,  B.,   et   al.,  ERAD:  the  long  road  to  destruction.   Nat   Cell  Biol,   2005.  7(8):  p.  766-­‐72.  

401.   Lamark,   T.   and   T.   Johansen,   Aggrephagy:   selective   disposal   of   protein  aggregates  by  macroautophagy.  Int  J  Cell  Biol,  2012.  2012:  p.  736905.  

402.   Bukau,  B.,   J.  Weissman,  and  A.  Horwich,  Molecular  chaperones  and  protein  quality  control.  Cell,  2006.  125(3):  p.  443-­‐51.  

403.   Ravikumar,   B.,   R.   Duden,   and   D.C.   Rubinsztein,  Aggregate-­‐prone  proteins  with  polyglutamine  and  polyalanine  expansions  are  degraded  by  autophagy.  Hum  Mol  Genet,  2002.  11(9):  p.  1107-­‐17.  

404.   Park,   H.J.,   et   al.,  A   soluble   sulfogalactosyl   ceramide  mimic  promotes  Delta  F508   CFTR   escape   from   endoplasmic   reticulum   associated   degradation.  Chem  Biol,  2009.  16(4):  p.  461-­‐70.  

405.   Le,   A.,   K.S.   Graham,   and   R.N.   Sifers,   Intracellular   degradation   of   the  transport-­‐impaired   human   PiZ   alpha   1-­‐antitrypsin   variant.   Biochemical  mapping   of   the   degradative   event   among   compartments   of   the   secretory  pathway.  J  Biol  Chem,  1990.  265(23):  p.  14001-­‐7.  

 

 

123  

406.   Hermosilla,   R.,   et   al.,   Disease-­‐causing   V(2)   vasopressin   receptors   are  retained  in  different  compartments  of   the  early  secretory  pathway.   Traffic,  2004.  5(12):  p.  993-­‐1005.  

407.   Robertson,   N.G.,   et   al.,   Subcellular   localisation,   secretion,   and   post-­‐translational   processing   of   normal   cochlin,   and   of   mutants   causing   the  sensorineural  deafness  and  vestibular  disorder,  DFNA9.   J  Med  Genet,  2003.  40(7):  p.  479-­‐86.  

408.   Kincaid,   M.M.   and   A.A.   Cooper,   Misfolded   proteins   traffic   from   the  endoplasmic   reticulum   (ER)   due   to   ER   export   signals.   Mol   Biol   Cell,   2007.  18(2):  p.  455-­‐63.  

409.   Appenzeller-­‐Herzog,   C.   and   H.P.   Hauri,   The   ER-­‐Golgi   intermediate  compartment  (ERGIC):  in  search  of  its  identity  and  function.   J  Cell  Sci,  2006.  119(Pt  11):  p.  2173-­‐83.  

410.   Barlowe,  C.,  et  al.,  COPII:  a  membrane  coat  formed  by  Sec  proteins  that  drive  vesicle  budding  from  the  endoplasmic  reticulum.   Cell,   1994.   77(6):   p.   895-­‐907.  

411.   Lewis,  M.J.,  J.C.  Rayner,  and  H.R.  Pelham,  A  novel  SNARE  complex  implicated  in  vesicle   fusion  with  the  endoplasmic  reticulum.   EMBO   J,   1997.   16(11):   p.  3017-­‐24.  

412.   Lewis,   M.J.   and   H.R.   Pelham,   SNARE-­‐mediated   retrograde   traffic   from   the  Golgi  complex  to  the  endoplasmic  reticulum.  Cell,  1996.  85(2):  p.  205-­‐15.  

413.   Sollner,  T.,  et  al.,  SNAP  receptors  implicated  in  vesicle  targeting  and  fusion.  Nature,  1993.  362(6418):  p.  318-­‐24.  

414.   Trombetta,   E.S.,   The   contribution   of   N-­‐glycans   and   their   processing   in   the  endoplasmic   reticulum   to   glycoprotein   biosynthesis.   Glycobiology,   2003.  13(9):  p.  77R-­‐91R.  

415.   Van   den   Steen,   P.,   et   al.,  Concepts  and  principles  of  O-­‐linked  glycosylation.  Crit  Rev  Biochem  Mol  Biol,  1998.  33(3):  p.  151-­‐208.  

416.   McNiven,   M.A.   and   H.M.   Thompson,   Vesicle   formation   at   the   plasma  membrane  and  trans-­‐Golgi  network:   the  same  but  different.   Science,   2006.  313(5793):  p.  1591-­‐4.  

417.   Orci,   L.,   et   al.,   Budding   from   Golgi   membranes   requires   the   coatomer  complex  of  non-­‐clathrin  coat  proteins.  Nature,  1993.  362(6421):  p.  648-­‐52.  

418.   Malhotra,   V.,   Unconventional   protein   secretion:   an   evolving   mechanism.  EMBO  J,  2013.  

419.   Sanders,   S.L.,   et   al.,   Sec61p   and   BiP   directly   facilitate   polypeptide  translocation  into  the  ER.  Cell,  1992.  69(2):  p.  353-­‐65.  

420.   Braakman,  I.,  J.  Helenius,  and  A.  Helenius,  Role  of  ATP  and  disulphide  bonds  during   protein   folding   in   the   endoplasmic   reticulum.   Nature,   1992.  356(6366):  p.  260-­‐2.  

421.   Mirazimi,   A.   and   L.   Svensson,   ATP   is   required   for   correct   folding   and  disulfide  bond  formation  of  rotavirus  VP7.  J  Virol,  2000.  74(17):  p.  8048-­‐52.  

422.   Dorner,  A.J.  and  R.J.  Kaufman,  The  levels  of  endoplasmic  reticulum  proteins  and  ATP  affect  folding  and  secretion  of  selective  proteins.  Biologicals,  1994.  22(2):  p.  103-­‐12.  

 

 

124  

423.   Verde,   C.,   et   al.,   Effect   of   ATP   depletion   and   DTT   on   the   transport   of  membrane   proteins   from   the   endoplasmic   reticulum   and   the   intermediate  compartment  to  the  Golgi  complex.  Eur  J  Cell  Biol,  1995.  67(3):  p.  267-­‐74.  

424.   Doms,   R.W.,   et   al.,   Role   for   adenosine   triphosphate   in   regulating   the  assembly  and  transport  of  vesicular  stomatitis  virus  G  protein  trimers.  J  Cell  Biol,  1987.  105(5):  p.  1957-­‐69.  

425.   Stevens,   F.J.   and   Y.   Argon,   Protein   folding   in   the   ER.   Semin   Cell   Dev   Biol,  1999.  10(5):  p.  443-­‐54.  

426.   Tu,  B.P.   and   J.S.  Weissman,  The  FAD-­‐  and  O(2)-­‐dependent  reaction  cycle  of  Ero1-­‐mediated   oxidative   protein   folding   in   the   endoplasmic   reticulum.  Molecular  cell,  2002.  10(5):  p.  983-­‐94.  

427.   Wang,   L.,   et   al.,   Reconstitution   of   human   Ero1-­‐Lalpha/protein-­‐disulfide  isomerase  oxidative  folding  pathway  in  vitro.  Position-­‐dependent  differences  in   role   between   the   a   and   a'   domains   of   protein-­‐disulfide   isomerase.   The  Journal  of  biological  chemistry,  2009.  284(1):  p.  199-­‐206.  

428.   Brown,  J.M.,  Exploiting  the  hypoxic  cancer  cell:  mechanisms  and  therapeutic  strategies.  Mol  Med  Today,  2000.  6(4):  p.  157-­‐62.  

429.   Koritzinsky,   M.,   et   al.,  Phosphorylation  of   eIF2alpha   is   required   for  mRNA  translation   inhibition   and   survival   during   moderate   hypoxia.   Radiother  Oncol,  2007.  83(3):  p.  353-­‐61.  

430.   de   Witte,   J.H.,   et   al.,   Prognostic   impact   of   urokinase-­‐type   plasminogen  activator   receptor   (uPAR)   in   cytosols   and   pellet   extracts   derived   from  primary  breast  tumours.  Br  J  Cancer,  2001.  85(1):  p.  85-­‐92.  

431.   Laerum,   O.D.,   et   al.,   Prognosis   in   adenocarcinomas   of   lower   oesophagus,  gastro-­‐oesophageal   junction   and   cardia   evaluated   by   uPAR-­‐immunohistochemistry.  Int  J  Cancer,  2012.  131(3):  p.  558-­‐69.  

432.   Zheng,  Q.,  Z.  Tang,  and  Z.  Wu,  [Urokinase-­‐type  plasminogen  activator  (uPA),  uPA   receptor   (uPA-­‐R)   and   inhibitors   (PA   I   -­‐1)   expression   in  hepatocellular  carcinoma   in   relation   to   cancer   invasion/metastasis   and   prognosis].  Zhonghua  Zhong  Liu  Za  Zhi,  1998.  20(1):  p.  57-­‐9.  

433.   Rigolin,  G.M.,   et   al.,  Soluble  urokinase-­‐type  plasminogen  activator  receptor  (suPAR)   as   an   independent   factor   predicting   worse   prognosis   and   extra-­‐bone   marrow   involvement   in   multiple   myeloma   patients.   Br   J   Haematol,  2003.  120(6):  p.  953-­‐9.  

434.   Miyake,   H.,   et   al.,  Elevation  of   serum   levels  of  urokinase-­‐type  plasminogen  activator   and   its   receptor   is   associated   with   disease   progression   and  prognosis  in  patients  with  prostate  cancer.  Prostate,  1999.  39(2):  p.  123-­‐9.  

435.   Graf,  M.,  et  al.,  High  expression  of  urokinase  plasminogen  activator  receptor  (UPA-­‐R)   in   acute   myeloid   leukemia   (AML)   is   associated   with   worse  prognosis.  Am  J  Hematol,  2005.  79(1):  p.  26-­‐35.  

436.   Jansens,   A.   and   I.   Braakman,   Pulse-­‐chase   labeling   techniques   for   the  analysis   of   protein  maturation   and   degradation.   Methods   Mol   Biol,   2003.  232:  p.  133-­‐45.  

437.   Rofstad,   E.K.,   et   al.,   Hypoxia   promotes   lymph   node   metastasis   in   human  melanoma   xenografts   by   up-­‐regulating   the   urokinase-­‐type   plasminogen  activator  receptor.  Cancer  Res,  2002.  62(6):  p.  1847-­‐53.  

 

 

125  

438.   Buchler,  P.,  et  al.,  Transcriptional  regulation  of  urokinase-­‐type  plasminogen  activator   receptor   by   hypoxia-­‐inducible   factor   1   is   crucial   for   invasion   of  pancreatic  and  liver  cancer.  Neoplasia,  2009.  11(2):  p.  196-­‐206.  

439.   Rofstad,   E.K.,   B.   Mathiesen,   and   K.   Galappathi,   Increased   metastatic  dissemination   in   human  melanoma   xenografts   after   subcurative   radiation  treatment:   radiation-­‐induced   increase   in   fraction   of   hypoxic   cells   and  hypoxia-­‐induced   up-­‐regulation   of   urokinase-­‐type   plasminogen   activator  receptor.  Cancer  Res,  2004.  64(1):  p.  13-­‐8.  

440.   Graham,   C.H.,   T.E.   Fitzpatrick,   and   K.R.   McCrae,   Hypoxia   stimulates  urokinase  receptor  expression  through  a  heme  protein-­‐dependent  pathway.  Blood,  1998.  91(9):  p.  3300-­‐7.  

441.   Lee,   K.H.,   et   al.,   Involvement   of   MAPK   pathway   in   hypoxia-­‐induced   up-­‐regulation   of   urokinase   plasminogen   activator   receptor   in   a   human  prostatic  cancer  cell  line,  PC3MLN4.  Exp  Mol  Med,  2004.  36(1):  p.  57-­‐64.  

442.   Sprague,  L.D.,  et  al.,  Effects  of  hypoxia  and  reoxygenation  on  the  expression  levels   of   the   urokinase-­‐type   plasminogen   activator,   its   inhibitor  plasminogen  activator  inhibitor  type-­‐1  and  the  urokinase-­‐type  plasminogen  activator  receptor  in  human  head  and  neck  tumour  cells.  Oncol  Rep,  2007.  17(5):  p.  1259-­‐68.  

443.   Kroon,  M.E.,   et   al.,  Urokinase  receptor  expression  on  human  microvascular  endothelial  cells  is  increased  by  hypoxia:  implications  for  capillary-­‐like  tube  formation  in  a  fibrin  matrix.  Blood,  2000.  96(8):  p.  2775-­‐83.  

444.   Buo,   L.,   et   al.,   Expression   and   release   of   plasminogen   activators,   their  inhibitors   and   receptor   by   human   tumor   cell   lines.   Anticancer   Res,   1994.  14(6B):  p.  2445-­‐51.  

445.   Khatib,  A.M.,  et  al.,  Regulation  of  urokinase  plasminogen  activator/plasmin-­‐mediated   invasion   of   melanoma   cells   by   the   integrin   vitronectin   receptor  alphaVbeta3.  Int  J  Cancer,  2001.  91(3):  p.  300-­‐8.  

446.   Llinas,   P.,   et   al.,   Crystal   structure   of   the   human   urokinase   plasminogen  activator  receptor  bound  to  an  antagonist  peptide.  EMBO  J,  2005.  24(9):  p.  1655-­‐63.  

447.   Huai,   Q.,   et   al.,   Structure   of   human   urokinase   plasminogen   activator   in  complex  with  its  receptor.  Science,  2006.  311(5761):  p.  656-­‐9.  

448.   Ploug,   M.,   et   al.,   Glycosylation   profile   of   a   recombinant   urokinase-­‐type  plasminogen  activator  receptor  expressed   in  Chinese  hamster  ovary  cells.   J  Biol  Chem,  1998.  273(22):  p.  13933-­‐43.  

449.   Gardsvoll,   H.,   et   al.,   Characterization   of   low-­‐glycosylated   forms   of   soluble  human   urokinase   receptor   expressed   in   Drosophila   Schneider   2   cells   after  deletion  of  glycosylation-­‐sites.  Protein  Expr  Purif,  2004.  34(2):  p.  284-­‐95.  

450.   Axten,   J.M.,   et   al.,   Discovery   of   7-­‐methyl-­‐5-­‐(1-­‐{[3-­‐(trifluoromethyl)phenyl]acetyl}-­‐2,3-­‐dihydro-­‐1H-­‐indol-­‐5-­‐yl)-­‐7H-­‐p   yrrolo[2,3-­‐d]pyrimidin-­‐4-­‐amine   (GSK2606414),   a   potent   and   selective   first-­‐in-­‐class  inhibitor   of   protein   kinase   R   (PKR)-­‐like   endoplasmic   reticulum   kinase  (PERK).  J  Med  Chem,  2012.  55(16):  p.  7193-­‐207.  

 

 

126  

451.   Atkins,   C.,   et   al.,   Characterization   of   a   novel   PERK   kinase   inhibitor   with  antitumor   and   antiangiogenic   activity.   Cancer   Res,   2013.   73(6):   p.   1993-­‐2002.  

452.   Damme,   M.,   et   al.,   Impaired   lysosomal   trimming   of   N-­‐linked  oligosaccharides  leads  to  hyperglycosylation  of  native  lysosomal  proteins  in  mice  with  alpha-­‐mannosidosis.  Mol  Cell  Biol,  2010.  30(1):  p.  273-­‐83.  

453.   Tartaglia,  G.G.,  et  al.,  Life  on  the  edge:  a  link  between  gene  expression  levels  and  aggregation  rates  of  human  proteins.  Trends  Biochem  Sci,  2007.  32(5):  p.  204-­‐6.  

454.   Chanat,   E.,   et   al.,   Reduction   of   the   disulfide   bond   of   chromogranin   B  (secretogranin   I)   in   the   trans-­‐Golgi   network   causes   its   missorting   to   the  constitutive  secretory  pathways.  EMBO  J,  1993.  12(5):  p.  2159-­‐68.  

455.   Tatu,   U.,   I.   Braakman,   and   A.   Helenius,   Membrane   glycoprotein   folding,  oligomerization  and  intracellular  transport:  effects  of  dithiothreitol  in  living  cells.  EMBO  J,  1993.  12(5):  p.  2151-­‐7.  

456.   Lodish,  H.F.  and  N.  Kong,  The  secretory  pathway  is  normal  in  dithiothreitol-­‐treated   cells,   but   disulfide-­‐bonded   proteins   are   reduced   and   reversibly  retained   in   the   endoplasmic   reticulum.   J   Biol   Chem,   1993.   268(27):   p.  20598-­‐605.  

457.   de   Silva,   A.,   I.   Braakman,   and   A.   Helenius,   Posttranslational   folding   of  vesicular   stomatitis   virus   G   protein   in   the   ER:   involvement   of   noncovalent  and  covalent  complexes.  J  Cell  Biol,  1993.  120(3):  p.  647-­‐55.  

458.   Srinivasan,   N.,   et   al.,   Conformations   of   disulfide   bridges   in   proteins.   Int   J  Pept  Protein  Res,  1990.  36(2):  p.  147-­‐55.  

459.   Thornton,   J.M.,   Disulphide   bridges   in   globular   proteins.   J   Mol   Biol,   1981.  151(2):  p.  261-­‐87.  

460.   Hillson,  D.A.,  N.  Lambert,  and  R.B.  Freedman,  Formation  and  isomerization  of   disulfide   bonds   in   proteins:   protein   disulfide-­‐isomerase.   Methods  Enzymol,  1984.  107:  p.  281-­‐94.  

461.   Komar,  A.A.,  A  pause  for  thought  along  the  co-­‐translational  folding  pathway.  Trends  Biochem  Sci,  2009.  34(1):  p.  16-­‐24.  

462.   McGinnes,   L.W.   and   T.G.   Morrison,   Role   of   cotranslational   disulfide   bond  formation   in   the   folding   of   the   hemagglutinin-­‐neuraminidase   protein   of  Newcastle  disease  virus.  Virology,  1996.  224(2):  p.  465-­‐76.  

463.   Chen,   W.,   et   al.,   Cotranslational   folding   and   calnexin   binding   during  glycoprotein  synthesis.  Proc  Natl  Acad  Sci  U  S  A,  1995.  92(14):  p.  6229-­‐33.  

464.   Weissman,   J.S.  and  P.S.  Kim,  The  pro  region  of  BPTI  facilitates  folding.  Cell,  1992.  71(5):  p.  841-­‐51.  

465.   Jansens,  A.,  E.  van  Duijn,  and  I.  Braakman,  Coordinated  nonvectorial  folding  in  a  newly   synthesized  multidomain  protein.   Science,   2002.   298(5602):   p.  2401-­‐3.  

466.   Allen,   S.,   H.Y.   Naim,   and   N.J.   Bulleid,   Intracellular   folding   of   tissue-­‐type  plasminogen   activator.   Effects   of   disulfide   bond   formation   on   N-­‐linked  glycosylation  and  secretion.  J  Biol  Chem,  1995.  270(9):  p.  4797-­‐804.  

 

 

127  

467.   Shakin-­‐Eshleman,   S.H.,   et   al.,   N-­‐linked   glycosylation   of   rabies   virus  glycoprotein.  Individual  sequons  differ  in  their  glycosylation  efficiencies  and  influence  on  cell  surface  expression.  J  Biol  Chem,  1992.  267(15):  p.  10690-­‐8.  

468.   Jaenicke,   R.,   Protein   folding:   local   structures,   domains,   subunits,   and  assemblies.  Biochemistry,  1991.  30(13):  p.  3147-­‐61.  

469.   Alder,   N.N.,   et   al.,   The   molecular   mechanisms   underlying   BiP-­‐mediated  gating   of   the   Sec61   translocon   of   the   endoplasmic   reticulum.   J   Cell   Biol,  2005.  168(3):  p.  389-­‐99.  

470.   Sauk,   J.J.,   et   al.,   Hsp47   and   the   translation-­‐translocation   machinery  cooperate  in  the  production  of  alpha  1(I)  chains  of  type  I  procollagen.   J  Biol  Chem,  1994.  269(6):  p.  3941-­‐6.  

471.   Roth,  R.A.  and  S.B.  Pierce,  In  vivo  cross-­‐linking  of  protein  disulfide  isomerase  to  immunoglobulins.  Biochemistry,  1987.  26(14):  p.  4179-­‐82.  

472.   Klappa,   P.,   R.B.   Freedman,   and   R.   Zimmermann,   Protein   disulphide  isomerase  and  a  lumenal  cyclophilin-­‐type  peptidyl  prolyl  cis-­‐trans  isomerase  are   in   transient   contact   with   secretory   proteins   during   late   stages   of  translocation.  Eur  J  Biochem,  1995.  232(3):  p.  755-­‐64.  

473.   Bulleid,   N.J.   and   R.B.   Freedman,   Defective   co-­‐translational   formation   of  disulphide   bonds   in   protein   disulphide-­‐isomerase-­‐deficient   microsomes.  Nature,  1988.  335(6191):  p.  649-­‐51.  

474.   Balch,  W.E.,   et   al.,   Adapting  proteostasis   for   disease   intervention.   Science,  2008.  319(5865):  p.  916-­‐9.  

475.   Sifers,  R.N.,  Manipulating  proteostasis.  Nat  Chem  Biol,  2010.  6(6):  p.  400-­‐1.  476.   Pena,   F.,   et   al.,   Calcium   as   a   crucial   cofactor   for   low   density   lipoprotein  

receptor  folding  in  the  endoplasmic  reticulum.   J  Biol  Chem,  2010.  285(12):  p.  8656-­‐64.  

477.   Hegedus,  T.,  et  al.,  F508del  CFTR  with  two  altered  RXR  motifs  escapes  from  ER   quality   control   but   its   channel   activity   is   thermally   sensitive.   Biochim  Biophys  Acta,  2006.  1758(5):  p.  565-­‐72.  

478.   Brown,   C.R.,   L.Q.   Hong-­‐Brown,   and   W.J.   Welch,   Correcting   temperature-­‐sensitive  protein  folding  defects.  J  Clin  Invest,  1997.  99(6):  p.  1432-­‐44.  

479.   Zhang,   M.   and   R.   Schekman,   Cell   biology.   Unconventional   secretion,  unconventional  solutions.  Science,  2013.  340(6132):  p.  559-­‐61.  

480.   Willems,   S.H.,   et   al.,   Thiol   isomerases   negatively   regulate   the   cellular  shedding  activity  of  ADAM17.  Biochem  J,  2010.  428(3):  p.  439-­‐50.  

481.   Jordan,   P.A.,   et   al.,   A   role   for   the   thiol   isomerase   protein   ERP5   in   platelet  function.  Blood,  2005.  105(4):  p.  1500-­‐7.  

482.   Elner,   S.G.,  Human   retinal   pigment   epithelial   lysis   of   extracellular  matrix:  functional   urokinase   plasminogen   activator   receptor,   collagenase,   and  elastase.  Trans  Am  Ophthalmol  Soc,  2002.  100:  p.  273-­‐99.  

483.   Marie,  M.,  et  al.,  The  function  of  the  intermediate  compartment  in  pre-­‐Golgi  trafficking  involves  its  stable  connection  with  the  centrosome.  Mol  Biol  Cell,  2009.  20(20):  p.  4458-­‐70.  

484.   Yoo,   J.S.,   et   al.,   Non-­‐conventional   trafficking   of   the   cystic   fibrosis  transmembrane   conductance   regulator   through   the   early   secretory  pathway.  J  Biol  Chem,  2002.  277(13):  p.  11401-­‐9.  

 

 

128  

485.   Nickel,  W.  and  M.  Seedorf,  Unconventional  mechanisms  of  protein  transport  to  the  cell  surface  of  eukaryotic  cells.   Annu  Rev  Cell  Dev  Biol,   2008.  24:  p.  287-­‐308.  

486.   Nickel,   W.   and   C.   Rabouille,   Mechanisms   of   regulated   unconventional  protein  secretion.  Nat  Rev  Mol  Cell  Biol,  2009.  10(2):  p.  148-­‐55.  

487.   Rabouille,   C.,   V.   Malhotra,   and   W.   Nickel,   Diversity   in   unconventional  protein  secretion.  J  Cell  Sci,  2012.  125(Pt  22):  p.  5251-­‐5.  

488.   Nykjaer,   A.,   et   al.,   Mannose   6-­‐phosphate/insulin-­‐like   growth   factor-­‐II  receptor   targets   the   urokinase   receptor   to   lysosomes   via   a   novel   binding  interaction.  J  Cell  Biol,  1998.  141(3):  p.  815-­‐28.  

489.   Bastholm,   L.,   et   al.,   Confocal   fluorescence   microscopy   of   urokinase  plasminogen   activator   receptor   and   cathepsin   D   in   human   MDA-­‐MB-­‐231  breast  cancer  cells  migrating  in  reconstituted  basement  membrane.  Biotech  Histochem,  1994.  69(2):  p.  61-­‐7.  

490.   Donaldson,   J.G.,   D.   Finazzi,   and   R.D.   Klausner,   Brefeldin   A   inhibits   Golgi  membrane-­‐catalysed   exchange   of   guanine   nucleotide   onto   ARF   protein.  Nature,  1992.  360(6402):  p.  350-­‐2.  

491.   Helms,  J.B.  and  J.E.  Rothman,  Inhibition  by  brefeldin  A  of  a  Golgi  membrane  enzyme  that  catalyses  exchange  of  guanine  nucleotide  bound  to  ARF.  Nature,  1992.  360(6402):  p.  352-­‐4.  

492.   Jackson,  C.L.  and  J.E.  Casanova,  Turning  on  ARF:  the  Sec7  family  of  guanine-­‐nucleotide-­‐exchange  factors.  Trends  Cell  Biol,  2000.  10(2):  p.  60-­‐7.  

493.   Klausner,   R.D.,   J.G.   Donaldson,   and   J.   Lippincott-­‐Schwartz,   Brefeldin   A:  insights  into  the  control  of  membrane  traffic  and  organelle  structure.   J  Cell  Biol,  1992.  116(5):  p.  1071-­‐80.  

494.   Scales,   S.J.,   M.   Gomez,   and   T.E.   Kreis,   Coat  proteins   regulating  membrane  traffic.  Int  Rev  Cytol,  2000.  195:  p.  67-­‐144.  

495.   Ritzenthaler,  C.,  et  al.,  Reevaluation  of  the  effects  of  brefeldin  A  on  plant  cells  using   tobacco   Bright   Yellow   2   cells   expressing   Golgi-­‐targeted   green  fluorescent  protein  and  COPI  antisera.  Plant  Cell,  2002.  14(1):  p.  237-­‐61.  

496.   Hess,  M.W.,   et   al.,  Cryopreparation  provides  new   insight   into   the  effects  of  brefeldin   A   on   the   structure   of   the   HepG2   Golgi   apparatus.   J   Struct   Biol,  2000.  130(1):  p.  63-­‐72.  

497.   Nebenfuhr,  A.,  C.  Ritzenthaler,  and  D.G.  Robinson,  Brefeldin  A:  deciphering  an  enigmatic  inhibitor  of  secretion.  Plant  Physiol,  2002.  130(3):  p.  1102-­‐8.  

498.   Misumi,  Y.,  et  al.,  Novel  blockade  by  brefeldin  A  of  intracellular  transport  of  secretory  proteins  in  cultured  rat  hepatocytes.   J  Biol  Chem,  1986.  261(24):  p.  11398-­‐403.  

499.   Fujiwara,  T.,  et  al.,  Brefeldin  A  causes  disassembly  of  the  Golgi  complex  and  accumulation   of   secretory   proteins   in   the   endoplasmic   reticulum.   J   Biol  Chem,  1988.  263(34):  p.  18545-­‐52.  

 

 

 

129  

 

 

130  

 

 

131  

 

 

132  

 

 

133  

 

 

134