Functional Analysis of the Contribution of HIF-1α Proline 402 to the Interaction with VHL

by

Carlo de Guzman Halnin

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

Department of Laboratory Medicine and Pathobiology University of Toronto

© Copyright by Carlo de Guzman Halnin (2016)

ii

Functional Analysis of the Contribution of HIF-1α Proline 402 to

the Interaction with VHL

Carlo de Guzman Halnin

Master of Science

Department of Laboratory Medicine and Pathobiology

University of Toronto

2016

Abstract

Hypoxia-inducible factor 1 alpha (HIF-1α) is involved in transcription of hypoxia-inducible

genes. In normoxia, it binds von Hippel-Lindau (VHL) resulting in ubiquitin-proteasome

degradation. This requires hydroxylation at prolines 564 (P564) or 402 (P402) within the

oxygen-dependent degradation (ODD) domain. P402 hydroxylation only occurs at higher oxygen

concentrations. Its specific contribution to the HIF-1α-VHL interaction is unknown. It was

hypothesized that loss of P402 in normoxia will dampen the HIF-1α-VHL interaction. HA-

tagged ODD constructs (WT, P402A, P564G, and P402A/P564G) were generated. FLAG-VHL

co-immunoprecipitation was lower for P402A compared to WT. In vitro ubiquitination decreased

with P402 mutation when P564 was also mutated. Cycloheximide chase analysis showed that

P402 loss had no significant effect on ODD half-life. These results indicate a secondary role for

P402 in the HIF-1alpha-VHL interaction. Elucidating the details of P402’s role in the interaction

of HIF-1alpha with VHL will enhance the understanding of mammalian oxygen sensing and

ubiquitin-proteasome pathways.

iii

Acknowledgments

None of this would have been possible without the guidance and support of Professor Michael

Ohh. Professors Jeffrey Lee and Stephane Angers kept these research efforts on the right track

through their work as members of the advisory committee. Members of the Ohh and Lee labs,

past and present, have contributed to this study in many ways. Dr. Severa Bunda, Betty Poon,

and Farshad Azimi were instrumental in terms of training in common techniques and lab

equipment. Drs. Claire Robinson, Pardeep Heir, and Halil Aydin gave pertinent advice and

consultation on experimental procedures and analysis. Much gratitude is extended to Dr. Norma

Masson and the lab of Professor Peter Ratcliffe for the provision of full-length HIF-1α P402 and

P564 hydroxyproline mutants that were the basis for the ODD mutants used in this study. Not to

be forgotten is the material support given to this project by the Canadian federal funding

agencies in the form of an NSERC CGS-M scholarship and CIHR grants to the Ohh lab.

iv

Table of Contents

Acknowledgments ........................................................................................................................ iii

Table of Contents ......................................................................................................................... iv

List of Figures ............................................................................................................................... vi

List of Abbreviations .................................................................................................................. vii

Introduction: HIF-1α proline hydroxylation and the VHL interaction ...............................1 1

1.1 HIF-1α is a major regulator of the response to hypoxia ................................................1

1.1.1 Structure and function of HIF-1α.............................................................................1

1.1.2 HIF-1α in oxygen homeostasis ................................................................................3

1.1.3 HIF-1α in development ............................................................................................4

1.1.4 HIF-1α in cancer ......................................................................................................5

1.1.5 HIF-1α in vascular disease .......................................................................................7

1.1.6 HIF-1α in diabetes ...................................................................................................8

1.1.7 HIF-1α and inflammation ........................................................................................8

1.1.8 HIF-2α, a HIF-1α isoform with distinctive roles .....................................................9

1.1.9 HIF-3α, the least known HIF-1α isoform ................................................................9

1.2 VHL is a negative regulator of HIF-1α ..........................................................................10

1.2.1 VHL degrades HIF-1α as an E3 ubiquitin ligase ...................................................10

1.2.2 VHL-independent HIF-1α degradation ..................................................................11

1.2.3 Perturbations of the VHL- HIF-1α interaction lead to disease ..............................12

1.3 HIF-1α degradation is mainly regulated by two proline hydroxylation sites.............13

1.3.1 PHDs are enzymes that act on HIF-1α proline hydroxylation sites .......................13

1.3.2 Other post-translational modifications of HIF-1α .................................................13

1.3.3 Proline 402 is an alternative but secondary proline hydroxylation site of HIF-1α ...........................................................................................................................14

1.3.4 Hypothesis and aims ..............................................................................................15

v

The contribution of proline 402 to the interaction between HIF-1α and VHL .................17 2

2.1 Materials and Methods ....................................................................................................17

2.1.1 Cell culture, constructs, and antibodies .................................................................17

2.1.2 Immunoprecipitation (IP) and immunoblotting .....................................................18

2.1.3 In vitro ubiquitination ............................................................................................19

2.1.4 Statistical analysis and generation of figures .........................................................19

2.2 Results ...............................................................................................................................19

2.2.1 Loss of P402 reduces HIF-1α binding to VHL ......................................................19

2.2.2 Loss of P402 in the absence of proline 564 decreases HIF-1α polyubiquitination ..................................................................................................21

2.2.3 Loss of P402 does not increase the half-life of HIF-1α .........................................22

2.2.4 Confirmation of ODD proline hydroxylation ........................................................23

2.3 Discussion..........................................................................................................................25

Concluding Remarks ..............................................................................................................27 3

3.1 Summary of Work ...........................................................................................................27

3.2 Future Directions .............................................................................................................27

References .....................................................................................................................................30

vi

List of Figures

Figure 1.1. Comparison between the domains of the three HIF-1 family members. .............. 2

Figure 1.2. Structural layout of SCF and VBC E3 ubiquitin ligase complexes. ................... 11

Figure 1.3. HIF-1α proline hydroxylation pathways and degradation vary according to

ambient oxygen levels. ................................................................................................................ 15

Figure 2.1. Loss of P402 decreases FLAG-VHL pulldown of HA-ODD. ............................... 20

Figure 2.2. Loss of P402 decreases in vitro polyubiquitination of ODD when proline 564 is

absent. .......................................................................................................................................... 21

Figure 2.3. Loss of P402 does not increase the half-life of ODD in a cycloheximide chase

assay. ............................................................................................................................................ 22

Supplementary Figure 2.4. Confirmation of hydroxylation of intact P564 sites on WT and

P402A ODD by anti-HyP564 antibody. .................................................................................... 23

Supplementary Figure 2.5. Immunoblotting for hydroxylation of P402 with an anti-HyP402

antibody. ...................................................................................................................................... 24

Supplementary Figure 2.6. Immunoblotting for proline hydroxylation with an anti-HyP

antibody. ...................................................................................................................................... 24

vii

List of Abbreviations

2-OG 2-OxoGlutarate

ANGPTL4 ANGioPoieTin-Like 4

BTB Broad-complex, Tramtrack, and Bric à brac domain

CBP (Cyclic adenosine monophosphate response element binding protein)-Binding

Protein

CD47 Cluster of Differentiation 47

DMSO DiMethyl SulfOxide

EC Endothelial Cells

EMT Epithelial-Mesenchymal Transition

EPO ErythroPOietin

FH Fumarate Hydratase

HA HemAgglutinin

HEK293 Human Embryonic Kidney 293 cells

L1CAM L1 Cell Adhesion Molecule

LOX Lysyl oxidase

mTOR Mammalian Target Of Rapamycin

ODD Oxygen-Dependent Degradation domain

PHD Prolyl-Hydroxylase Domain

RHOBTB3 RHO-related BTB domain-containing protein 3

ROS Reactive Oxygen Species

viii

SDH Succinate DeHydrogenase

SDS-PAGE Sodium Dodecyl Sulfate PolyAcrylamide Gel Electrophoresis

SHARP1 enhancer-of-Split and HAiry-Related Protein 1

Tat Trans-Activator of Transcription

VEGF Vascular Endothelial Growth Factor

1

Introduction: HIF-1α proline hydroxylation and the 1VHL interaction

1.1 HIF-1α is a major regulator of the response to hypoxia

1.1.1 Structure and function of HIF-1α

Hypoxia-inducible factors (HIFs) are a family of transcription factors regulated by changes in

oxygen tension that mediate the genetic response to hypoxia [1]. Of the three members of the

HIF family, HIF-1 is the most well-known. It was discovered as a nuclear factor which bound a

transcriptional activation site for human erythropoietin (EPO) during hypoxia [2], demonstrating

the canonical upregulation of hypoxia-response genes by the HIF family members. Further

characterization of HIF-1 revealed it to be a heterodimer with α and β subunits [3]. HIF-1α is the

larger subunit which contains DNA-binding and transactivation domains [4]. Its structure (Figure

1.1) has 826 amino acid residues and several regions: a basic helix-loop-helix (bHLH) domain

(residues 17-71), a Per-Arnt-Sim (PAS) domain (residues 85-296), and two transactivation

domains (residues 531-575 and 786-826) [4, 5]. The bHLH domain, common to many

transcription factors, is involved in binding to DNA strands and protein ligands [6]. PAS

domains take their name from the three proteins they were found in: period circadian protein

(Per, a Drosophila clock gene) [7], aryl hydrocarbon nuclear translocator protein (Arnt, involved

in dioxin metabolism and also known as HIF-1β) [8], and single-minded protein (Sim, which

regulates central nervous system development in Drosophila) [9]. Heterodimerization between

these proteins, members of the PAS family, is accomplished via their PAS domains [10]. PAS

domains consist of two internal homology domains called PAS-A and PAS-B. HIF-1α residues

between the N- and C-terminal transactivation domains are inhibitory to the gene induction

function of HIF-1α [5].

2

Figure 1.1. Comparison between the domains of the three HIF-1 family members. HIF-1α

and HIF-2α are similar and contain bHLH, PAS, ODD, NAD, and CAD domains. HIF-3α is

found as a number of alternative splice variants. Domains: bHLH (Basic Helix-Loop-Helix),

PAS (Per-Arnt-Sim), ODD (Oxygen-Dependent Degradation), NAD (N-terminal Activation

Domain), CAD (C-terminal Activation Domain), LZIP (Leucine ZIPper). Adapted from Lee et al

2014 [11].

Dimerization with HIF-1β is essential for activation of HIF-1α targets [12], and this happens

once HIF-1α is translocated to the nucleus [13]. A nuclear localization signal in the C-terminal

transactivation domain [14] of HIF-1α was found to be involved in binding to the alpha subunits

of nuclear transport proteins called importins [15]. HIF-1α also has an N-terminal nuclear export

signal that binds importins 4 and 7 to facilitate nuclear entry [16]. Formation of the heterodimer

serves to maintain both HIF-1 subunits in the nucleus, as the absence of either one leads to

export of the other [13].

HIF-1α transcription is dependent on a number of protein coactivators. Nuclear-localized HIF-

1α recruits the CREB-binding protein (CBP)/p300 coactivator complex to facilitate its

transcriptional function [14], binding to it via the C-terminal transactivation domain [17].

However, abrogation of CBP/p300 binding only reduces HIF-1α transactivation by 30-50%, with

some activity accounted for by inhibition of histone deacetylases [18]. Other factors are required

for the induction of hypoxia-inducible genes by HIF-1α. The Tat interacting protein 60 (TIP60)

histone acetyltransferase needs to be recruited by HIF-1α to chromatin for full activation of

hypoxia-responsive genes [19]. CDK8 (cyclin-dependent kinase 8) recruitment by HIF-1α is

3

needed to initiate the elongation activity of RNA polymerase II at gene targets of HIF-1α [20].

HIF-1β also recruits its own set of associated coactivators, such as transforming acidic coiled-

coil-containing protein 3 (TACC3) which is bound by HIF-1β’s N-terminal region [21]. A

histone methyltransferase called SET9 (Su (var) 3-9, Enhancer of zeste, and Trithorax 9) is

involved in stabilizing HIF-1α at the binding sites of target genes [22].

A number of proteins can modify the transcriptional profile of HIF-1α. Some serve to enhance

HIF-1α activity, such as pyruvate kinase M2 (PKM2) which increases HIF-1α target binding and

p300 recruitment [23]. Pontin increases the recruitment of p300 to a subset of promoters targeted

by HIF-1α [24]. Paired-like homeodomain 1 (PITX1) binds to HIF-1β to enhance the HIF-1α-

dependent expression of some histone demethylases [25]. Many suppress the transcriptional

activity of HIF-1α upon binding. A chromatin remodeling protein called reptin was found to bind

a selection of hypoxia-responsive gene promoters to decrease their transcription in hypoxia [26].

Such downregulation is also the case for transglutaminase 2, which binds to HIF-1β [27]. FHL1

(four-and-a-half LIM domain protein 1) binds to HIF-1α to repress its transcriptional activity,

while FHL2 binds to CBP/p300 instead [28]. In chronic hypoxia, peroxiredoxin 2 (PRDX2) and

PRDX4 binding inhibit HIF-1α binding to target genes [29]. With HIF-1α induction of PRDX2,

this forms a negative feedback loop for HIF-1α activity in chronic hypoxia.

The activated complex created from HIF-1’s α and β subunits binds to genes that contain

hypoxia response elements (HREs) that contain the core sequence RCGTG, where R is either A

or G [30]. Genes activated by HIF-1α are involved in numerous cellular pathways with a focus

on the response to hypoxia, such as glycolysis [31] and angiogenesis [32]. HIF-1α’s involvement

in a wide array of cell functions is further underscored by its ubiquitous expression in all tissues

[33, 34]. However, HIF-1α is rapidly degraded in normoxic conditions [35].

1.1.2 HIF-1α in oxygen homeostasis

HIF-1α is tightly bound to the cellular and physiological context of hypoxia, with mRNA [33]

and protein [2-4, 13] levels both highly upregulated. With the role HIF-1α plays in hypoxia, it is

thus a crucial component of oxygen homeostasis. It is expressed in nearly all multicellular

organisms [36], even in the simplest ones [37]. With the vast majority of multicellular life on

Earth highly dependent on oxygen for metabolism, adaptations to decreases in oxygen levels are

vital. HIF-1α achieves this by mediating a switch to anaerobic metabolism, upregulating genes

4

involved in glycolysis such as phosphoglycerate kinase 1 and PKM2 (an isoform of pyruvate

kinase) [23, 38]. The glucose required for the glycolytic switch is brought into cells through the

membrane by glucose transporter 1 (GLUT1), which is upregulated by HIF-1α [39].

Lactate dehydrogenase A (LDHA), which is involved in the interconversion of pyruvate into

lactate, is upregulated by HIF-1α [40]. This promotes lactic acid fermentation at the expense of

oxidative respiration further downstream. HIF-1α also induces expression of pyruvate

dehydrogenase kinase 1 (PDK1) [41]. PDK1 phosphorylates and inhibits pyruvate

dehydrogenase, preventing pyruvate from entering the citric acid cycle and feeding the electron

transport chain on the mitochondrial membrane. A more drastic means of reducing oxidative

respiration is HIF-1α induction of mitochondrial autophagy (mitophagy) through induction of

BNIP3 (BCL2/adenovirus E1B 19 kDa protein-interacting protein 3) and mediation via Beclin1

and Atg5 (autophagy protein 5) [42]. This was seen to contribute to cell survival and decreased

levels of reactive oxygen species (ROS). Downregulation of mitochondrial ATP synthesis, which

consumes 80% of cellular oxygen [43], plays a dominant role in reducing the demand for

oxygen.

HIF-1α also initiates measures to maintain and increase the supply of oxygen to tissues. HIF-1α

is needed for increased breathing rate in response to hypoxia in mice [44]. The oxygen-carrying

capacity of the blood is boosted by HIF-1α transcriptional activation of EPO, which is involved

in initiating red blood cell proliferation [45]. Vasodilation may assist in increased systemic

distribution of oxygen, since its mediator, atrial natriuretic peptide, is induced by HIF-1α [46].

One downstream target of HIF-1α is NOR-1 (neuron-derived orphan receptor 1), a transcription

factor that may be protective to endothelial cells in hypoxia by upregulating cellular inhibitor of

apoptosis protein 2 (cIAP2) [47]. This ensures the integrity of the endothelium to allow for

efficient distribution of oxygen. As a longer-term measure, vascular endothelial growth factor

(VEGF) induction by HIF-1α [32] induces angiogenesis to improve vessel networks in hypoxic

tissues.

1.1.3 HIF-1α in development

By virtue of its role in mediating angiogenesis, HIF-1α is crucial to development. Knockout is

lethal at embryonic day 11, being marked by gross defects in neural tubes and vascular networks

[48]. HIF-1α recruitment to Notch signaling targets helps to maintain an undifferentiated cell

5

state [49]. HIF-1α is involved in epithelial-mesenchymal transition (EMT), which is involved in

different developmental processes. HIF-1α is important in neural crest migration, with its loss

leading to impairment of this process through the downregulation of Twist, a repressor of E-

cadherin and thus a promoter of EMT [50]. In the coronary endothelium, induction of Snail

(another repressor of E-cadherin) led to EMT, which is involved in the formation of heart valves

[51]. During pregnancy, HIF-1α is essential to the development of placental tissue and its

associated blood vessels [52].

HIF-1α is important to the development of connective tissue and the structural integrity of the

body. HIF-1α in fibroblasts coordinates collagen deposition and extracellular matrix construction

by activating collagen hydroxylase genes [53]. In bone tissue, HIF-1α cooperates with Runx2, an

osteogenic bone factor, to induce VEGF [54]. HIF-1α is essential to the development of

cartilage, with its absence leading to death of chondrocytes in the hypoxic environment [55].

Adipocyte differentiation involves an upregulation of HIF-1α [56]. Wound healing demonstrates

the contribution of HIF-1α to developmental pathways involved in recovery from injury.

Heterozygous-null mice exhibited reduced burn healing due to defective angiogenesis [57].

1.1.4 HIF-1α in cancer

HIF-1α function is heavily implicated in human disease, most prominently in cancer where it is

often overexpressed [58]. Tumors often experience hypoxia [59], and are dependent on

angiogenesis mediated by HIF-1α [60]. HIF-1α in cancer cells mediates the switch to anaerobic

metabolism [61]. HIF-1α also aids cancer by upregulation of cell proliferation factors. Met

(inducer of hepatocyte growth factor, HGF) is induced in hypoxic regions of tumors via hypoxia-

response elements in it promoter [62]. HGF induction by Met promotes invasive growth of tumor

cells. Increased transforming growth factor alpha (TGF-α) expression by HIF-1α is a major

contributing factor to renal cell carcinoma [63]. In prostate cancer, upregulated FoxA2

transcription factor works with HIF-1α to promote a more aggressive neuroendocrine phenotype

through induction of proteins overexpressed in metastatic prostate cancer [64]. This induction

was mediated by recruitment of p300 to target gene promoters by FoxA2.

HIF-1α activity is implicated in the metastatic growth of cancer cells, particularly through the

promotion of EMT. Snail and BM1 (B cell-specific Moloney murine leukemia virus integration

site 1) are examples of a HIF-1α targets involved in promoting EMT [51, 65, 66]. HIF-1α may be

6

involved in helping cancer cells evade immune surveillance, with its direct upregulation of CD-

47 (cluster of differentiation 47) in breast cancer cells leading to reduced phagocytosis by bone-

marrow derived macrophages [67].

Mechanisms that suppress HIF-1α or its activity can be protective in cancer. An example is

SHARP1 (enhancer-of-split and hairy-related protein 1), a factor that degrades HIF-1α and

whose expression is positively correlated with survival and vice versa for metastasis in patients

with triple-negative breast cancer [68].

HIF-1α overexpression in cancer appears to be mediated in a number of ways. Insulin-growth

factor 2 (IGF-2) is upregulated by HIF-1α and vice versa, leading to a potential positive feedback

loop in tumor cells [69]. The tumor suppressor p53 was found to negatively regulate HIF-1α by

acting as a chaperone for MDM2 (Mouse double minute 2 homolog)-mediated degradation [70].

This demonstrates a widely documented mechanism to upregulate HIF-1α in cancer, since p53 is

lost in many cancer types [71]. The PI3K (phosphoinositide 3-kinase)-Akt-mTOR (mammalian

target of rapamycin) pathway, heavily involved in cell proliferation, also induces increased levels

of HIF-1α [72, 73]. The PTEN (phosphatase and tensin homolog) tumor suppressor, antagonist

of the PI3K pathway often lost in cancer [74], acts to downregulate HIF-1α [75]. Dysregulation

of mitochondrial metabolism in cancer can also serve to stabilize HIF-1α by reducing its oxygen-

dependent degradation (discussed later in the section on PHDs). Succinate dehydrogenase (SDH)

mutations occur in some rare cancers [76]. The increase in succinate leads to HIF-1α

stabilization and promotion of oncogenesis [77]. Renal cancer caused by loss of fumarate

hydratase (FH) is promoted by HIF-1α stabilization caused by increased cellular fumarate levels

[78].

Much work has been done on the contributions of HIF-1α to breast cancer. HER2 (human

epidermal growth factor receptor 2, a major breast cancer oncogene) upregulates HIF-1α levels

in a manner dependent on the PI3K pathway [79]. The HIF-1α target genes CD47 and TAZ

(tafazzin) promote a stem cell phenotype [67, 80]. In triple negative breast cancer (TNBC, which

doesn’t express any of the 3 common markers, one of which is HER2), XBP1 (X-box binding

protein 1, involved in ER stress through the unfolded protein response) assembles with HIF-1α in

a transcriptional complex which recruits RNA polymerase II [81]. Patient data reveals gene

expression data that shows correlations between XBP1, HIF-1α, and poor prognosis. Breast

7

cancer metastasis is mediated by HIF-1α gene targets. HIF-1α upregulation of lysyl oxidases

(LOX, LOX-like 2, LOX-like 4), angiopoietin-like 4 (ANGPTL4), and L1 cell adhesion

molecule (L1CAM) promote metastasis to the lung [82, 83]. LOX enzymes catalyze collagen

crosslinking, a preparatory step for lung metastatic niches [82]. These enzymes are correlated

with cancer metastasis to bone, suggesting a role for HIF-1α [84]. Extravasation of metastatic

breast cancer cells through endothelial cells (ECs) occurs via ANGPTL4 inhibition of EC-EC

junctions and L1CAM-mediated adhesion of breast cancer cells to ECs [83]. RhoA and Rho

kinase 1 (ROCK1) upregulation by HIF-1α increases the motility and invasiveness of breast

cancer cells [85]. ROCK1 phosphorylation of RhoA leads to phosphorylation of both myosin

light chain (MLC) and focal adhesion kinase (FAK), which both mediate the motile phenotype.

HIF-1α may even contribute to breast cancer pathology beyond its transactivation activity.

Binding of HIF-1α to the multi-subunit complex γ-secretase promotes the latter’s activity and

leads to breast cancer invasion and metastasis [86]. The mechanisms by which HIF-1α functions

in breast cancer demonstrate the widespread involvement and impact of HIF-1α activity in

physiological and pathological processes.

1.1.5 HIF-1α in vascular disease

HIF-1α’s importance in the system that delivers oxygen to the body translates into a prominence

in vascular disease. The loss of blood flow into tissues, or ischemia, causes transient hypoxia that

upregulates HIF-1α [87]. Mice that are heterozygotes for HIF-1α had decreased angiogenesis and

recovery from ischemia, which can be recovered by adenoviral expression of HIF-1α [88].

Protection may be provided by anti-ROS activities mediated by HIF-1α, since loss in ischemia

was seen to worsen cell injury through creation of a more oxidative environment. It is possible

that this is also mediated in part through induction of heme oxygenase 1 (HO-1) by HIF-1α [89].

However, HIF-1α function is not always protective to vascular tissues. Hypoxic HIF-1α

induction was seen to activate DNA methyltransferases which induced a fibrotic cardiac

phenotype that would hinder cardiac function [90]. Cardiac fibrosis induction by HIF-1α is also

mediated via its promotion of EMT [51]. In atherosclerosis, HIF-1α in hypoxic murine plaque

regions caused lipid accumulation in macrophages [91]. Initial neuronal survival and function in

ischemic stroke was improved in mice that had deficiencies in HIF-1α and HIF-2α [92].

8

1.1.6 HIF-1α in diabetes

Hyperglycemia inhibits the stabilization of HIF-1α in hypoxia [93]. Loss of insulin also leads to

a decrease in HIF-1α, since insulin signaling upregulates HIF-1α [94]. The concomitant decrease

in HIF-1α impacts diabetes pathophysiology through a number of mechanisms. Loss of HIF-1α

negatively affects beta cell insulin secretion and other functions, with rescue in high-fat diet

mouse models possible through inhibition of HIF-1α degradation [95]. Rescue was not possible

in HIF-1α knockouts. These knockouts also displayed greater insulin resistance and glucose

metabolism impairment in response to a high-fat diet [96]. Defective wound healing in diabetes

is caused by decreased proliferation factors due to decreased HIF-1α, with rescue possible

through exogenous HIF-1α expression in mice wound tissue [97].

1.1.7 HIF-1α and inflammation

Hypoxia and inflammation are closely intertwined, with multiple observations indicating that

either one leads to the other [98]. HIF-1α induction in inflammation is caused by the hypoxia in

inflamed tissues and the pro-inflammatory transcription factor NF-κB [99]. The innate

inflammatory response is dependent on HIF-1α in several ways. Knockout was seen to lead to

loss of granulocyte and monocyte function due to impaired ATP production [100]. Macrophages

stimulated by inflammatory stimuli such as lipopolysaccharide change from a proliferative

program to a HIF-1α-dependent glycolytic metabolism [101]. Phagocytosis of pathogens needs

HIF-1α to upregulate pathogen-killing factors such as granule proteases and antimicrobial

peptides [102]. Formation of neutrophil extracellular traps (NETs) was found to depend on a

pathway where inflammatory signaling through mTOR upregulates HIF-1α [103]. This mTOR-

dependent upregulation is thought to depend on inhibition of oxygen-dependent degradation of

HIF-1α [73]. Toll-like receptor induction of HIF-1α in normoxia in dendritic cells activates

certain proinflammatory genes that are not induced with hypoxia, such as nitric oxide synthase 2

(Nos2) and prostaglandin-endoperoxide synthase 2 (Ptgs2) [104]. Interestingly, HIF-1α binds to

the promoter of histone deacetylase 2 (HDAC2) to suppress its transcription, which induces

inflammation through C-X-C motif ligand 8 (CXCL8) [105]. HIF-1α activity may limit tissue

damage in inflammation by upregulating forkhead box P3 (FoxP3), which is a transcriptional

factor for regulatory T-cells which initiate various anti-inflammatory responses [106]. With HIF-

1α closely linked to inflammation and inflammation involved in the previously mentioned and

many other illnesses [107], it can be seen that HIF-1α plays a major role in human disease.

9

1.1.8 HIF-2α, a HIF-1α isoform with distinctive roles

As previously mentioned, HIF-2α is one of three members of the HIF transcription factor family

[1]. HIF-2α was first discovered as a HIF-1α-like transcription factor originally thought to be

expressed exclusively in endothelial cells [108]. HIF-2α is larger but still quite similar to HIF-1α

in terms of structure (Figure 1.1). It was found to have greater expression than HIF-1α in lung,

heart, and liver along with the ability to upregulate VEGF [109]. HIF-2α has a different

transcriptional profile compared to HIF-1α due to differences between their N-terminal

transactivation domains [110]. HIF-2α target specificity may also be due to the specific

coactivators it recruits, such as ETS-1 (erythroblastosis virus E26 oncogene homolog-1) [111]

and ELK-1 [110]. HIF-2α does not seem to be pivotal to upregulating glycolytic genes, with little

effect seen with its loss or induction [110, 112, 113]. Oct4 (octamer-binding transcription factor

4, which maintains stem cell pluripotency) is a HIF-2α-specific target, establishing a unique role

for HIF-2α in development [114]. HIF-2α plays unique roles in the liver not shared with HIF-1α,

such as preferential upregulation of EPO [115] and regulation of lipid metabolism [116].

Like HIF-1α, complete knockout of HIF-2α causes embryonic lethality with gross vascular

defects [117], while selective knockout in endothelial cells led to abnormal vasculature in both

normal and tumor cells [118]. HIF-2α’s greater role in inducing EPO was confirmed in global

knockout restricted to the post-natal period, with said mice developing anemia due to loss of

EPO [119]. Erythrocytosis, a condition marked by an excess of red blood cells, is caused by

mutations in HIF-2α, with no such mutations occurring in the other HIFs [120]. These findings

establish a dominant role for HIF-2α in cardiovascular development. The pathogenesis of renal

cell carcinoma is also impacted by HIF-2α, with its downregulation inhibiting tumorigenesis of

cells lacking a protein called Von Hippel-Lindau (a HIF-α interactor, discussed later) [121].

1.1.9 HIF-3α, the least known HIF-1α isoform

HIF-3α was discovered in the course of searching for HIF-like transcription factors, and is the

smallest HIF family member at only 662 amino acids [122]. While it has most of the typical

domains found in the members of the HIF-α family (Figure 1.1), HIF-3α has a leucine zipper

instead of a C-terminal transactivation domain, and acts to inhibit upregulation of genes by HIF-

1α [122-124]. Up to 8 other splice variants have been found so far, all missing the leucine zipper

and some missing parts of the N-terminal and central regions [124, 125]. The expression patterns

10

of each variant within different human tissues differ widely with low levels in cancer cell lines,

as opposed to HIF-1α’s ubiquity and cancer overexpression [126]. More recent work has shown

that HIF-3α can also activate genes, with some of which, e.g. VEGF, are also targeted by others

HIF-α proteins [127]. However, some of these genes are only upregulated by HIF-3α, and fall

into many categories with no clear pattern apparent. Overall, much is not yet clear about the role

of HIF-3α in physiology and disease [125].

1.2 VHL is a negative regulator of HIF-1α

1.2.1 VHL degrades HIF-1α as an E3 ubiquitin ligase

The gene for Von Hippel-Lindau (VHL) protein was discovered around the same time as HIF-1α

[128]. The protein takes its name from a hereditary disease described decades prior. VHL disease

is named after Eugen von Hippel, a German ophthalmologist who described highly vascularized

eye tumors [129], and Arvid Lindau, a Swedish pathologist who found the same type of tumors

(hemangioblastomas) in the brain and spinal cord [130]. Mutations in VHL disease were found

to coincide with the newly discovered gene [128].

VHL is a tumor suppressor protein, with its expression in VHL-null cells suppressing

tumorigenesis in nude mice [131]. It was found to bind to elongins B and C (eloB and eloC) with

cullin-2 (CUL2) [132], in a complex similar to SCF (Skp1–Cdc53–F-box), a complex that targets

proteins for degradation by binding and ubiquitinating them [133]. EloC and CUL2 resemble

Skp1 (S-phase kinase-associated protein 1) and Cdc53 (cell division cycle 53), respectively

[132]. The F-box protein is the substrate recognition subunit of SCF [133], and thus a similar

role was proposed for VHL in a VHL-eloB-eloC-CUL2 degradation complex (Figure 1.2) [132].

This was confirmed by the crystal structure of VBC (VHL-eloB-eloC) [134]. VHL has two

distinct domains: an N-terminal β-domain enriched in β-sheets and a smaller α-domain

composed of α-helices. The α-domain binds to eloC (which itself binds eloB at a distinct

interface), and is one of the two VHL hotspots which have many mutation sites involved in VHL

disease. The other major area where VHL disease mutations are found is a distinct patch on the

β-domain.

11

Figure 1.2. Structural layout of SCF and VBC E3 ubiquitin ligase complexes. SCF and VBC

bind targets through substrate-recognition subunits F-box and VHL, respectively. The scaffold

proteins Cdc53 and CUL2 both bind Rbx1, which transfers ubiquitin to targets bound by the

complex. This leads to subsequent degradation of targets via the proteasome. VHL binds HIF-1α

through the latter’s ODD domain. Abbreviations: Skp1 (S-phase kinase-associated protein 1),

Cdc53 (cell division cycle 53), SCF (Skp1-Cdc53-F-box), VHL (von Hippel-Lindau), eloC

(elongin C), eloB (elongin B), CUL2 (cullin 2), VBC (VHL-eloC-eloB), Rbx1 (ring-box 1),

ODD (oxygen-dependent degradation domain).

As previously mentioned, HIF-1α is degraded in normoxic conditions. It was found that cells

without VHL express HIF-1α target genes in excess despite the presence of oxygen [135]. VHL-

deficient cells were then found to be unable to degrade HIF-1α, with reintroduction of VHL

restoring normoxic degradation [136]. VHL-mediated degradation of HIF-1α requires binding to

a roughly 200 amino acid stretch in the middle of the HIF-1α protein called the Oxygen-

Dependent Degradation (ODD) domain (Figure 1.1) [137]. VHL’s role as a substrate recognition

subunit in an SCF-like protein degradation complex was demonstrated by its ability to bind and

ubiquitinate HIF-1α [138]. Like SCF, VHL’s complex binds Rbx1 (ring-box 1) to facilitate

transfer of ubiquitin to bound targets [139]. Thus, VHL negatively regulates HIF-1α in normoxia

as an E3 ubiquitin ligase.

1.2.2 VHL-independent HIF-1α degradation

It should be noted that HIF-1α may be degraded in VHL-independent pathways, even in the

absence of oxygen. Hsp70 (heat shock protein 70) along with ubiquitin ligase CHIP (C terminus

of HSC70-Interacting Protein) targets HIF-1α (but not HIF-2α) for ubiquitination and

degradation in prolonged hypoxia [140]. SHARP1 binds HIF-1α and presents it to the

12

proteasome, bypassing the cell’s ubiquitination machinery and causing degradation even in

normoxia [68]. HIF-1α is targeted for lysosomal degradation via autophagy mediated by the

chaperone HSC70 (heat shock cognate 71 kDa protein) and receptor LAMP2A (lysosome-

associated membrane protein 2 a) [141].

1.2.3 Perturbations of the VHL- HIF-1α interaction lead to disease

VHL disease can be divided into two types [142]. Type I VHL disease is marked by a low risk of

pheochromocytoma, a neuroendocrine tumor found in chromaffin cells (mostly located in the

adrenal medulla). Type 2 VHL disease is the opposite, having a high risk of pheochromocytoma,

and has three subtypes: Type 2A (low risk of renal cell carcinoma), Type 2B (high risk of renal

cell carcinoma), and Type 2C (at risk for pheochromocytoma and none of the other

complications of VHL disease such as hemangioblastomas). These VHL disease types have

structural determinants. Type 1 mutations map to the β-domain hydrophobic core of VHL, which

would cause gross misfolding of the VHL protein [134]. Type 2 mutations disrupt binding to

either eloC or HIF-1α.

Loss of the interaction between VHL and HIF-1α has significant impacts on human health. Loss

of the titular protein or its function lends VHL disease its various symptoms. The vascular

central nervous system tumors identified by Lindau are caused by VEGF upregulation are due to

a lack of VHL expression [143]. VHL-heterozygous mice develop similar tumors in their livers

[144]. Deletions of the VHL gene are common in clear cell renal carcinoma [145]. As previously

mentioned, lack of VHL in such cells stabilizes HIF-1α which leads to induction of factors such

as TGF-α that drive growth of renal cell carcinoma [63]. In addition to cancer growth, loss of

VHL’s control of HIF-1α also enhances the spread of such cancers, primarily through loss of E-

cadherin and induction of EMT [65]. Reintroduction of VHL into a renal cell carcinoma cell line

suppressed their tumorigenicity in vivo [131]. A mouse model lacking VHL and TP53 (mouse

homolog of p53) indicated that both HIF-1α and HIF-2α contribute to clear cell renal carcinoma

[146]. Additional deletion of either HIF-1 isoform led to failure of renal tumors to form.

13

1.3 HIF-1α degradation is mainly regulated by two proline hydroxylation sites

1.3.1 PHDs are enzymes that act on HIF-1α proline hydroxylation sites

The degradation of HIF-1α by VHL depends on hydroxylation of proline residues to form

hydroxyproline (HyP). In humans, this hydroxylation is achieved through three enzymes called

prolyl hydroxylase domain-containing (PHD) proteins [147]. These enzymes require Fe2+

and 2-

oxoglutarate (2-OG) cofactors in order to take molecular oxygen and transfer it to a hydroxyl on

proline and to their other product succinate (resulting from the decarboxylation of 2-OG [148].

Increased succinate inhibits the activity of PHDs [77], providing an explanation for SDH

mutations leading to pheochromocytoma, renal cell carcinoma, and other rare cancers [76]. A

similar mechanism also underlies the loss of FH leading to increased fumarate (downstream

metabolite of succinate) which then leads to PHD activity suppression [78]. PHD2 is the main

enzyme responsible for regulating the degradation of HIF-1α through proline hydroxylation

[149]. PHDs also hydroxylate other proteins, such as the HIF-1α coactivator PKM2 [23].

PHD hydroxylation is enhanced by protein co-factors. Rho-related BTB (Broad-complex,

Tramtrack, and Bric à brac) domain-containing protein 3 (RHOBTB3) binds with PHD2 to

promote HIF-1α hydroxylation [150]. Dimerization of RHOBTB3 with LIMD1 (LIM domain-

containing protein 1) forms a RHOBTB3/LIMD1-PHD2-VHL-HIF-1α complex that promotes

maximum degradation of HIF-1α. Formation of this complex is suppressed by hypoxia.

Phospholipase D1 (PLD1) also mediates the formation of a HIF-1α complex containing PHD2

and VHL [151]. PLD1 complex formation promotes HIF-1α proline hydroxylation and results in

dissociation of HIF-1α-VHL from the complex. As with RHOBTB3, these interactions are

abrogated in hypoxia. Regulation of PHD levels by other proteins modulates their activity. PHD2

and PHD3 are directly upregulated by HIF-1α, forming a negative feedback loop for hypoxic

induction upon reoxygenation [152, 153]. Siah2 (seven in absentia homolog 2) targets PHD1 and

PHD3 for degradation via the ubiquitin-proteasome pathway [154]. This degradation is enhanced

in hypoxia, potentially through upregulation of Siah2 transcription.

1.3.2 Other post-translational modifications of HIF-1α

Another hydroxylase acts on asparagine 803 (N803) of HIF-1α, with the normoxic presence of

the hydroxylated asparagine leading to inhibition of transactivation activity [155]. This was

14

found to be done by factor inhibiting HIF-1α (FIH-1), with its production of N803 leading to

impaired binding of p300/CBP to the C-terminal transactivation domain [156]. N803

hydroxylation occurs at oxygen levels even higher than the minimum needed for proline

hydroxylation [157]. It can be seen that hydroxylation regulates not only HIF-1α stability but

also its transcriptional activity.

Non-hydroxyproline post-translational modifications also affect the stability of HIF-1α.

Acetylation via ARD1 (arrest-defective protein 1) enhances the HIF-1α-VHL interaction [158].

In hypoxia, SUMO (small ubiquitin-like modifier) is conjugated to HIF-1α, allowing for

hydroxyproline-independent degradation via VHL [159]. Removal of SUMO by SENP1 (sentrin-

specific protease 1) is essential for the stabilization of HIF-1α in hypoxia. Methylation by

SET7/9 destabilized nuclear HIF-1α, while demethylation by LSD1 (lysine-specific demethylase

1) does the opposite [160]. Stabilization of HIF-1α is accomplished by phosphorylation via

cyclin-dependent kinase 1 (CDK1) [161].

1.3.3 Proline 402 is an alternative but secondary proline hydroxylation site of HIF-1α

Hydroxylation of the proline 564 (P564) residue of HIF-1α was found to be essential for VHL-

mediated degradation [162]. Deficiency (in non-mammalian lysates) or inactivation (temporarily

by hypoxia or permanently by heat) of PHDs abrogated VHL binding of HIF-1α and

ubiquitination, as did mutation of P564 to alanine. The presence of the hydroxylated P564

(HyP564) residue was confirmed using mass spectrometry. HIF-1α residues 555-575, mostly

conserved in vertebrates, constitute a minimal sequence capable of being hydroxylated and

bound by VHL’s E3 ubiquitin ligase complex. The crystal structure of a peptide with HyP564

shows VHL making critical hydrogen bonds with HyP564 and the surrounding residues [163].

N-terminal truncations all the way to the residues L562 and A563 along with the C-terminal

truncation up to I566 led to decreased binding in a far-Western assay. VHL residues contacting

HyP564 were found to be heavily implicated in VHL disease as common mutation sites.

Further work showed that VHL binds, ubiquitinates, and degrades an ODD fragment (344-553)

that does not contain HyP564 [164]. This interaction is lost when proline 402 (P402) is mutated

or when prolyl hydroxylation is inhibited. A peptide with hydroxylated P402 (HyP402) was

found to bind with the same affinity to VHL as a peptide containing HyP564 [165]. HyP402 is

15

present only at higher oxygen tensions (Figure 1.3) and is heavily dependent on the presence of

HyP564 [166]. This was determined by transfection of HIF-1α constructs with mutations in

hydroxyprolines into mouse embryonic fibroblasts (MEFs) with HIF knocked out by a Cre-LoxP

(Causes recombination – locus of X-over P1) system. Mutation of P564 to glycine results in

significantly decreased levels of HyP402, but not vice versa. Upon exposure to hypoxia, HyP402

is lost before HyP564 [157]. These observations show that P402 is an alternative proline

hydroxylation site that is only active at conditions close to normoxia.

Figure 1.3. HIF-1α proline hydroxylation pathways and degradation vary according to

ambient oxygen levels.

1.3.4 Hypothesis and aims

HyP402 is essentially present only in normoxia and in the presence of HyP564. This suggests

that HyP402 may play an auxiliary role in the HIF-1α-VHL interaction in normoxia. The

differential contribution of HyP402 to the interaction of HIF-1α with VHL has not been

investigated. These include changes in binding to VHL, ubiquitination of HIF-1α, and the rate at

which HIF-1α is degraded. Loss of P402 should result in a dampening of the interaction between

ODD and VHL. This will be studied with ODD constructs mutated at the two HyP (P402A and

P564G) sites using in vitro systems and human embryonic kidney 293 (HEK293) cells. HEK293

cells are a standard cell line used for their high efficiency of transfection and ease of use [167].

They are a good model when cell-type specific variation is not an issue, such as in the study of

the biochemistry of a ubiquitously expressed protein. The P402A and P564G mutations have

16

been extensively described and used in many studies [108, 164, 166]. Loss of proline in

exchange for a small aliphatic residue is expected to change the structure of the protein due to

the loss of the proline-induced bend. However, this does not appear to be an issue for the ODD-

VHL interaction as a P394A mutation did not affect ubiquitination of an ODD construct [164].

Two aspects of the ODD-VHL interaction will be studied: binding of ODD to VHL, and the

degradation of ODD by VHL.

17

The contribution of proline 402 to the interaction 2between HIF-1α and VHL

2.1 Materials and Methods

2.1.1 Cell culture, constructs, and antibodies

HEK293 cells were sourced from the American Type Culture Collection (Rockville, MD). Cells

were maintained in Dulbecco’s Modified Eagle’s Medium (DMEM; Wisent, St-Bruno, QC,

Canada) supplemented with 10% fetal bovine serum (FBS; Wisent, St-Bruno, QC, Canada), 100

µg/mL streptomycin (Sigma-Aldrich, Oakville, ON), and 1 U/mL penicillin (Sigma-Aldrich,

Oakville, ON) at 37°C in a humidified 5% CO2 atmosphere.

For cell transfections, HEK293 cells were seeded on 150 mm tissue culture dishes (BD

Transduction Laboratories, Mississauga, ON) and grown to 95% confluency. Plasmid DNA and

polyethylenimine in a 4:1 ratio was added to Opti-Minimum Essential Media (OMEM; Gibco,

Burlington, ON), vortexed, and incubated for 10-15 min at room temperature. Cells were

trypsinized, resuspended in OMEM, and added in equal volumes to the DNA mixtures. Cell

mixtures were incubated for 10 min at room temperature, and then plated in 100 mm dishes (BD

Transduction Laboratories, Mississauga, ON) with 10 ml DMEM. Cells were harvested at 48 h

post-transfection. For chase analysis with cycloheximide (Bioshop, Burlington, ON), a 100

µg/ml concentration was used with treatments at 60, 30, and 5 min before lysis.

HA-tagged ODD constructs spanning HIF-1α residues 354 to 623 were created using a ligation-

independent-cloning-based modification of the pCDNA 3.4 plasmid from the lab of Dr. Jeffrey

Lee. The following primers were used: 5’-

CGCCAAGAGCGGCCCTGCTATGTACCCATACGATGTTCCAGATTACGCTGGCAGCCT

TCAACAAACAGAATGTGTCCTTAAACCG-3’ (forward) and 5’-

GGCACCAGGCGGCCTCATTCATCAGTGGTGGCAGTGGTAGT-3’ (reverse). These

constructs had proline hydroxylation sites mutated: WT, P402A, P564G, and P402A P564G.

Full-length HIF-1α constructs with these mutations that were the templates for ODD constructs

were generous gifts from Dr. Norma Masson and the lab of Dr. Peter Ratcliffe. ODD constructs

were recloned into HA-pCDNA3.1 for in vitro expression using BamHI and XhoI. The following

pair of primers was used: 5’-

18

TATGGATCCATGCTTCAACAAACAGAATGTGTCCTTAAACCG-3’ (forward) and 5’-

TATCTCGAGTCATTCATCAGTGGTGGCAGTGGT (reverse).

Monoclonal rabbit anti-HA (C29F4) and anti-HyP564 (D43B5) antibodies were obtained from

Cell Signaling Technology (Beverly, MA). Monoclonal mouse (F1804) and polyclonal rabbit

(F7425) FLAG antibodies were sourced from Sigma-Aldrich (Oakville, ON). Polyclonal rabbit

anti-PHD2 antibody was sourced from Novus Biologicals (Littleton, CO). Monoclonal mouse

anti-vinculin (ab18058), polyclonal rabbit anti-HyP (ab72775), and polyclonal rabbit anti-

HyP402 (ab37067) antibodies were procured from Abcam (Cambridge, MA). Goat anti-rabbit

(31460) IgG and anti-rabbit (31430) Horseradish-peroxidase-conjugated IgG secondary

antibodies were obtained from Pierce Biotechnology (Rockford, IL).

2.1.2 Immunoprecipitation (IP) and immunoblotting

For treatment with MG-132 (Peptides International, Louisville, KY), cells were incubated with

10 µM for 4 h. HEK293 Cells were then lysed in EBC buffer (50 mM Tris pH 8.0, 120 mM

NaCl, 0.5% NP-40) supplemented with protease inhibitor cocktail tablets (Roche, Laval, QC).

Lysates were sonicated and cleared by centrifugation at 14,000 rpm for 10 minutes at 4°C.

Protein levels were quantified by a Bradford Assay with Bio-Rad Protein Assay Dye Reagent

(Bio-Rad, Mississauga, ON). Equal amounts (1.5 mg) of total protein were subject to IP in a

volume of 1 ml with M2 anti-FLAG antibody immobilized on Protein-A beads (Repligen,

Waltham, MA). IPs were performed with rocking at 4°C for 1.5 h. IPs were washed five times

with NETN (20 mM Tris pH 8.0, 900 mM NaCl, 1 mM ethylenediaminetetraacetic acid (EDTA,

and 0.5% NP-40), and resuspended in sodium dodecyl sulfate polyacrylamide gel electrophoresis

(SDS-PAGE) sample buffer prior to immunoblotting.

Western blotting was performed as previously described [168] with a few modifications. Total

protein (150 µg, 10% of immunoprecipitation input) was boiled in SDS-PAGE sample buffer for

5 min at 95°C prior to SDS-PAGE resolution on bis-acrylamide gels. Transfer to polyvinylidene

fluoride (PVDF) membranes (Bio-Rad, Mississauga, ON) was performed at 450 mA for 4-5 h at

4°C. Membranes were blocked for 1 hour at room temperature in 4% milk diluted with TBST

(Tris-Buffered Saline pH 7.4, with 0.05% Tween-20), washed 5 times in TBST, and then

incubated overnight at 4°C with the indicated primary antibodies prepared in TBST. This was

followed by 5 TBST washes, incubation with secondary antibody for 1 h at room temperature

19

and 5 more TBST washes. Finally, bands were visualized by incubating membranes in enhanced

chemiluminescent substrate and imaged with a VersaDoc imaging system (Bio-Rad,

Mississauga, ON).

2.1.3 In vitro ubiquitination

HA-pCDNA3.1 ODD constructs were expressed using TnT T7 Coupled Reticulocyte Lysate

System (Promega, Madison, WI) with incubation at 30°C for 1.5 h. This was followed by an in

vitro ubiquitination assay as previously described [138]. In brief, 5 µl of in vitro translated ODD

was combined with reagents from Boston Biochem (Cambridge, MA): 10µg/µl human ubiquitin,

140 ng/µl ubiquitin-aldehyde, HeLa S100 cytoplasmic extracts, and ATP regeneration system. A

total volume of 20 µl was incubated at 30°C for 2 h. Assays and inputs were resolved with SDS-

PAGE and blotted with anti-HA antibody.

2.1.4 Statistical analysis and generation of figures

Quantification of band intensities was done using ImageJ with measurements of band size,

average intensity, and background. Statistical analysis and generation of graphs were performed

with GraphPad Prism 5 (GraphPad Software, La Jolla, CA). Two-tailed unpaired t-tests with a

95% confidence interval were used to assess statistical significance. Figures were created with

Inkscape.

2.2 Results

2.2.1 Loss of P402 reduces HIF-1α binding to VHL

To assess the role of P402 in the binding between HIF-1α and VHL, HEK293 cells were

transfected with plasmids encoding HA-tagged ODD hydroxyproline mutants, along with

FLAG-VHL and PHD2. Cells were treated with or without the proteasome inhibitor MG-132,

lysed, and IPed with anti-FLAG antibody. Figure 2.1 shows the resulting immunoblots. HA-

ODD co-IP was decreased in the P402A mutant to 40% of WT, with this difference eliminated in

the presence of MG-132. Binding to the P564G mutant is barely visible only in the presence of

MG-132, with pulldown equal to only 20% of WT. These results suggest that loss of P402

decreases the ability of HIF-1α to bind to VHL.

20

Figure 2.1. Loss of P402 decreases FLAG-VHL pulldown of HA-ODD. HEK293 cells were

transfected with HA-ODD hydroxyproline mutant constructs, FLAG-VHL, and PHD2 plasmids.

At 48 h post-transfection, samples were treated with 10 µM MG-132 or dimethyl sulfoxide

(DMSO) solvent control for 4 h prior to IP with anti-FLAG antibody. IPs and inputs were

immunoblotted with the indicated antibodies. Densitometry analysis was performed on HA IP

blots with values in proportion to WT pulldowns, equal to 1. Results are representative of 3

independent experiments, with error bars representing standard deviations. Differences were

analyzed with an unpaired t-test with a 95% confidence interval. Asterisks (*) indicate p

21

2.2.2 Loss of P402 in the absence of proline 564 decreases HIF-1α polyubiquitination

To measure the effect of P402 loss on the ubiquitination of HIF-1α, HA-pCDNA ODD

hydroxyproline mutant constructs were in vitro translated in rabbit reticulocyte lysate. ODD

constructs were then in vitro ubiquitinated with purified human ubiquitin and HeLa S100

cytoplasmic extracts. Anti-HA blotting of ubiquitination assays and in vitro translation inputs is

presented in Figure 2.2. Quantification of polyubiquitination was achieved by densitometry of

ubiquitin laddering found at and above 100 kDa. While the P402A mutant showed no significant

decrease in polyubiquitination compared to WT, the double mutant (7%) does with regards to

P564G (51%). This analysis of polyubiquitination indicates that a loss of P402 in HIF-1α is

associated with decreased ubiquitination when P564 is not present.

Figure 2.2. Loss of P402 decreases in vitro polyubiquitination of ODD when proline 564 is

absent. HA-pCDNA plasmids encoding ODD hydroxyproline mutants were in vitro translated

using rabbit reticulocyte lysate. In vitro translated HA-ODD was in vitro ubiquitinated using

purified human ubiquitin and HeLa cell S100 cytoplasmic extracts. Ubiquitination assays and in

vitro inputs were immunoblotted with anti-HA antibody. Densitometry of lane areas above 100

kDa was performed, with the value for WT set to 1. Results are representative of 3 independent

experiments, with error bars representing standard deviations. Differences were analyzed with an

unpaired t-test with a 95% confidence interval. Asterisks (*) indicate p

22

2.2.3 Loss of P402 does not increase the half-life of HIF-1α

To quantify the effects of the loss of P402, HEK293 cells were transfected with HA-ODD

hydroxyproline mutants, FLAG-VHL, and PHD2. Cycloheximide at 100 µg/ml concentration

was applied at 60, 30, and 5 minutes prior to lysis of cell plates. Immunoblots are shown in

Figure 2.3. Both WT and P402A ODD have a half-life of less than 5 min, while the P564

mutants are mostly stable for the entire time course. This partitioning of ODD half-lives by the

status of the P564 site argues for the lack of an effect of P402 loss on the half-life of HIF-1α.

Figure 2.3. Loss of P402 does not increase the half-life of ODD in a cycloheximide chase

assay. HEK293 cells were transfected with HA-ODD hydroxyproline mutants, FLAG-VHL, and

PHD2. At 48 h post-transfection, samples were treated with 100 µg/µl cycloheximide or DMSO

solvent control for 60, 30, and 5 min prior to immunoblotting with the indicated antibodies.

Densitometry analysis was performed on HA blots with values in proportion to WT (equal to 1).

Results are representative of 4 independent experiments, with error bars representing standard

deviations.

23

2.2.4 Confirmation of ODD proline hydroxylation

To confirm that the prolines on ODD were fully hydroxylated, three different anti-

hydroxyproline antibodies were used in immunoblotting transfected HEK293 cell lysates. P564

hydroxylation was assayed by transfecting HEK293 cells with HA-ODD hydroxyproline

mutants, FLAG-VHL, and PHD2. Cells were treated with 10 µM MG-132 or DMSO for 4 h

before immunoblotting with the corresponding antibodies (Supplementary Figure 2.4). The anti-

HyP564 blotting shows a signal only for WT and P402A as expected.

Supplementary Figure 2.4. Confirmation of hydroxylation of intact P564 sites on WT and

P402A ODD by anti-HyP564 antibody. HEK293 cells were transfected with HA-ODD

hydroxyproline mutants, FLAG-VHL, and PHD2. At 48 h post-transfection, samples were

treated with 10 µM MG-132 or DMSO solvent control for 4 h prior to immunoblotting with the

indicated antibodies. Results are representative of 3 independent experiments.

Hydroxylation of P402 was assayed by transfecting full-length HIF-1α hydroxyproline mutants

into HEK293 cells. Cells were treated with MG-132 for 4 h at 48 h after transfection, lysed, and

immunoblotted with the corresponding antibodies (Supplementary Figure 2.5). There was no

signal, despite WT and P564G HIF-1α having intact P402 sites.

24

Supplementary Figure 2.5. Immunoblotting for hydroxylation of P402 with an anti-HyP402

antibody. HEK293 cells were transfected with HIF-1α hydroxyproline mutants, treated with 10

µM MG-132 for 4 h, then immunoblotted with the indicated antibodies.

Immunoblotting with a pan-HyP antibody was used as an alternative method of measuring the

levels of HyP402. Full-length HIF-1α hydroxyproline mutants and PHD2 were transfected into

HEK293 cells. At 48 h after transfection, cells were treated with MG-132 for 4 h, lysed, and

immunoblotted with the corresponding antibodies (Supplementary Figure 2.6). There was no

signal at all even from WT HIF-1α. These hydroxyproline-specific antibodies were only able to

verify the hydroxylation of P564, with HyP402-specific and general HyP antibodies both failing

to detect their respective targets.

Supplementary Figure 2.6. Immunoblotting for proline hydroxylation with an anti-HyP

antibody. HEK293 cells were transfected with HIF-1α hydroxyproline mutants and PHD2,

treated with 10 µM MG-132 for 4 h, then immunoblotted with the indicated antibodies.

25

2.3 Discussion

The response of multicellular life to low oxygen conditions involves the activation of HIF-1α,

which upregulates genes that mediate the adaptation to hypoxia [4]. In normoxia, VHL binds

HIF-1α for degradation to abrogate the hypoxic response [135]. This binding requires the

hydroxylation of P564 in the ODD region of HIF-1α with which VHL makes vital contacts

[163]. Further study revealed another site, P402, that is hydroxylated only at higher oxygen

concentrations [166].

The share of HyP402 in HIF-1α binding to VHL and its subsequent degradation is unknown.

This was investigated with HA-tagged ODD constructs mutated at P402 and P564. The current

study shows that the presence of P402 accounts for a small share of VHL binding and

ubiquitination of ODD. FLAG-VHL IP showed less binding for P402A, with this decrease

erased with MG-132 treatment (Figure 2.1). That MG-132 removes the difference in VHL

binding when P402 is lost shows the secondary role played by P402 in VHL binding. P564

binding compensated for the loss of P402 given enough time for VHL binding. P564 loss

essentially reduces VHL binding to a minimal level and P402 does not compensate even with

MG-132 incubation.

The ubiquitination and degradation assays used in this study did not seem to show much of a

difference with loss of P402. This may just be an issue of insufficient sensitivity to detect the

smaller response with P402 loss as opposed to P564 loss. Loss of P402 decreased in vitro

ubiquitination only when P564 was also lost (Figure 2.2). While ubiquitination of WT and P402

mutant was similar, it is possible that a shorter incubation time would reveal a difference in

ubiquitination. Saturation of the ODD ubiquitination sites may be a factor, just as VHL binding

was saturated in the IP when MG-132 was applied. A cycloheximide chase assay showed no

decrease in the rate of ODD degradation when P402 was mutated (Figure 2.3). This may be due

to a lack of sensitivity in using cycloheximide to measure small changes in degradation rate. An

in vitro assay would allow for the use of a defined amount of ODD, along with the option of

adding known amounts of VHL and PHD2 to properly calibrate degradation rates for sensitive

measurement. Such a method would also avoid off-target effects of cycloheximide on the various

components of the ubiquitin-proteasome system that may impede ODD degradation.

26

The question of when P402 is not hydroxylated is important for determining its contribution to

the physiology of oxygen sensing and homeostatic adaptation. RCC4 renal cell carcinoma

cultures lost detectable P402 hydroxylation when incubated at 0.5% O2 but not at 2% [166]. The

dynamics of P402 hydroxylation will differ in other cell lines based on the levels of VHL and

PHDs. With excess VHL, the additional binding affinity provided by P402 is unnecessary and

the presence of HyP564 is sufficient for maximum normoxic degradation. The levels of PHDs

are expected to have a direct relationship with the level of HyP402. The affinity of PHDs for the

HyP sites could also be a factor, as it was found that the N-terminal half of ODD has a higher

dissociation rate with PHD2 [169]. The activity of PHDs depend on the availability of iron and

citric acid cycle metabolites [148], which could be limiting in certain contexts. Oxygen and

glucose deprivation was found to lead to stabilization of HIF-1α even after a period of normoxia,

demonstrating that 2-OG deprivation may have lasting effects on PHD hydroxylation even in the

presence of abundant oxygen [170].

HyP402’s importance is underlined by P402 sequence conservation in higher developed

organisms from amphibians to humans [166]. Hydroxylation of P402 must occur for good

reason, as it is an energy-intensive process to split molecular oxygen, requiring the loss of a

high-energy carbon bond from 2-OG for succinate production [148]. The loss of HyP402 in mild

hypoxia may serve to provide an initial stabilization of HIF-1α due to slightly decreased binding

to VHL. This initial priming of HIF-1α might prepare the cell for further hypoxic insult with

minor activation of hypoxia-inducible genes. Such preconditioning using intermittent hypoxia

has been found to be protective in cardiac and brain ischemic insults [171]. In the same vein, it

may be desirable in a therapeutic context to abrogate P402 hydroxylation artificially with small

molecules that bind to the P402 site. Aside from resistance to ischemia, increased long-term

stabilization of HIF-1α may enhance angiogenesis and oxygen carrying capacity. This may be of

use in treating anemia, building cardiovascular endurance, and acclimatizing to higher altitudes.

HyP402’s ability to induce degradation of HIF-1α without the presence of HyP564 [164, 166] is

a possible means of compensating for an inability of VHL to bind HyP564. This is important if

HyP402 binds to a site that different from that of HyP564.

The mechanistic details of how HyP402 participates in the interface formed by HIF-1α with

VHL are important to understanding HyP402’s contribution to the HIF-1α-VHL pathway. There

are two possibilities for how HyP402 interacts with VHL: either its binds to the same site as

27

HyP564 or it recognizes a different VHL site. Both hypotheses would explain the decreased

P402A pulldown with HA-VHL in the absence of MG-132 (Figure 2.1) differently. With an

identical VHL site, HyP402 may act to increase the association of HIF-1α with VHL by having a

binding site exposed for certain orientations of the substrate vis-à-vis the enzyme. A different

binding site on VHL opens the possibility of a synergistic cooperation between HyP402 and

HyP564. Both sites can promote degradation independently [164, 166], so the order of binding to

VHL should not matter. With both HyP bound, it is possible that the combination of

conformation changes induced by binding enhances the binding of VHL to Cul2, or increases the

rate of ubiquitin ligation by the VHL complex.

Concluding Remarks 3

3.1 Summary of Work

The current study set out to explore what the effects of the loss of P402 would do to the

interaction between HIF-1α and VHL. This was done with HA-tagged hydroxyproline ODD

constructs. IP with FLAG-VHL showed decreased binding when P402 is mutated, with this

decrease erased with MG-132 treatment (Figure 2.1). In vitro ubiquitination of ODD was

decreased by P402 mutation only when P564 was already mutated (Figure 2.2). Together, these

results suggest that the P402 site imparts an enhanced binding of ODD to VHL and subsequent

ubiquitination. However, a cycloheximide chase assay showed no increase in the half-life of

ODD when P402 was mutated (Figure 2.3). This may just be due to the assay not being sensitive

enough to detect the contribution of HyP402 to ODD degradation. If P402A is degraded slower

than WT ODD, this would suggest a model where hydroxylation of P402 in normoxia is essential

for the maximum degradation of HIF-1α. Loss of HyP402 in slight hypoxia would create an

initial HIF-1α stabilization that might prime cells and tissues for more severe hypoxia. The

mechanics and physiology of these implications need to be studied to expand the understanding

of ubiquitin-proteasome degradation and oxygen homeostasis.

3.2 Future Directions

As mentioned before, the sensitivity of some of the techniques used could be improved.

Reducing the incubation time of the in vitro ubiquitination assay should reveal a difference

between WT and P402A. Degradation rate of the ODD constructs may be better studied with an

28

in vitro assay that can be adjusted to detect small differences. Verification of the current results

can be accomplished in a few ways. The IP experiment can be repeated under mild hypoxia to

remove HyP402 from the WT ODD. This should result in pulldown of WT being equal to

P402A. Ubiquitination can be assayed in cells by performing an IP after lysates are boiled in

SDS and diluted. This allows the detection of direct ubiquitination of ODD and not that of VHL

complex or other associated proteins. Verification of proline hydroxylation was only

accomplished for P564 in the current study (Supplementary Figure 2.4), since commercial

antibodies against HyP564 and pan-HyP did not give any signal (Supplementary Figures 2.5 and

2.6). Antibodies may have to be generated [157, 166] against HyP402 to check its levels.

Alternatively, commercially-available ELISAs against hydroxyproline to detect collagen can be

adapted to detect HyP402.

The scope of the current study can be expanded in two directions. The first would be to elucidate

the mechanics of HyP402 binding to VHL. VHL mutants may be used to check if HyP402 binds

to the same site as HyP564 or a different one in IP experiments. Isothermal titration calorimetry

can be used to measure more favorable binding of WT compared to P402A [172]. Quantitative

examination of the heat evolved from a protein-protein interaction in a calorimetric chamber

allows for determination of association constants and reaction stoichiometries. A more

energetically-favored interaction with ODD containing an intact P402 hints at possible

conformational changes that enhance the interaction. Such a conformational change may be

detected by circular dichroism (CD), which relies on the differential absorption of circularly-

polarized light [173]. The far UV CD signature reflects absorption by peptide bonds, revealing

secondary structure such as α-helices and β-sheets. Near UV absorption signatures probe the

chemical context of aromatic residues to indicate whether the protein sample being analyzed is

folded or not. The golden standard for analyzing protein structural details remains X-ray

crystallography with its atomic-level resolution [174]. The previous structure showing the

interaction of HyP564 with VHL only used a 20-amino acid peptide [163]. This is gravely

insufficient to show HyP402 interaction with VHL, much less the full details of ODD binding to

VHL. A crystal structure of ODD bound to VHL is the definitive answer to the question of what

HyP402 does in the interface of HIF-1α and VHL.

The other major avenue for investigating the significance of HyP402 is conducting studies on its

contribution to normal and disease physiology. A priority would be the determination of levels in

29

HyP402 in normal cell lines and tissues in order to establish its dynamics in various contexts.

This includes the dependence of HyP402 on the levels of VHL and PHDs, and other potential

factors that could affect P402 hydroxylation. Potential priming by HyP402 can be studied by

exposing normal cells and tissues to mild hypoxia followed by more severe hypoxia. This will

reveal any initial stabilization of HIF-1α and any protective effects caused by it. In parallel, there

should be examinations of HyP402 levels in disease, especially in cancers involving VHL. Such

studies may demonstrate a compensatory effect of HyP402 when binding to HyP564 is lost by

VHL. All of these physiology experiments would be greatly aided with the creation of a HIF-1α

P402A mouse model. These mice should be investigated for a worse response to hypoxia and

VHL inactivation, along with potential developmental changes.

30

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