apoptosis by c-myc - helsinki
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Helsinki University Biomedical Dissertations No.192
Apoptosis by c-Myc:
coupling metabolic reprogramming
and mitochondrial apoptosis
Anni Viheriäranta née Nieminen
Biochemistry and Developmental Biology,
Institute of Biomedicine, Faculty of Medicine &
Research Programs Unit, Translational Cancer Biology,
University of Helsinki
Finland
Helsinki Biomedicum Graduate Program
ACADEMIC DISSERTATION
To be publicly discussed, with the permission of Faculty of Medicine,
University of Helsinki, in Biomedicum Helsinki Lecture hall 2, on March 21st at
12 o´clock noon
Helsinki 2014
Supervisor: Juha Klefström, PhD, Research Director Translational Cancer Biology Research Program & Institute of Biomedicine Faculty of Medicine, University of Helsinki Helsinki, Finland Reviewers appointed by the Faculty: John Eriksson, PhD, Professor Ville Hietakangas,
Institute of Biotechnology University of Helsinki
Helsinki, Finland Thesis follow-up committee: Phillippe Juin, PhD, Professor John Eriksson, PhD, Professor Institut de Recherche Thérapeutique Université de Nantes France Opponent appointed by the Faculty: Martin Eilers, Biozentrum der Universität Würzburg Theodor Boveri Institut Würzburg, Germany
ISBN 978-952-10-9784-3 (nid.) ISBN 978-952-10-9785-0 (PDF) ISSN: 1457-8433 http://ethesis.helsinki.fi Yliopistopaino Helsinki 2014
To my family
Table of Contents ORIGINAL PUBLICATIONS .......................................................................................... 6
ABBREVIATIONS ............................................................................................................. 7
ABSTRACT ...................................................................................................................... 10
INTRODUCTION ............................................................................................................ 12
REVIEW OF THE LITERATURE ................................................................................ 13 1. Molecular mechanism of cancer ....................................................................................... 13
1.1 Hallmarks of cancer ........................................................................................................... 13 1.2 Excessive proliferation signals from oncogenes is guarded by tumor suppressors .......... 15
1.2.1 Mechanism of oncogene function ............................................................................................ 15 1.2.2 Tumor suppressors ................................................................................................................... 17
1.2.2.1 Master regulator p53 ....................................................................................................... 18 1.2.2.2 Tumor suppression by Lkb1 ............................................................................................. 21 1.2.2.3 AMPK as tumor suppressor ............................................................................................. 23
1.2.3 Cancer metabolism ................................................................................................................... 26 1.3 Breast cancer ..................................................................................................................... 29
2. Apoptosis ............................................................................................................................. 31 2.1 Apoptosis as a form of cell death ...................................................................................... 31 2.2 Apoptotic signaling molecules .......................................................................................... 32
2.2.1 Caspases as executioners ......................................................................................................... 33 2.2.2 Bcl-2 family ............................................................................................................................. 36
2.2.2.1 BH domains in the classification and function of Bcl-2 family members ........................ 36 2.2.2.4 BH3:groove interactions and conformations ................................................................... 38
2.3.1 Mitochondrial apoptosis pathway ............................................................................................ 40 2.3.1.1 MOMP and beyond .......................................................................................................... 40 2.3.1.2 Models for Bak/Bax activation by the Bcl-2 family proteins ........................................... 41 2.3.1.3 p53-dependent apoptotic pathway ................................................................................... 43
2.3.2 Death receptor apoptosis pathway ........................................................................................... 45 2.3.2.1 Death receptors and the DICS complex ........................................................................... 45
3. Oncogene Myc pathways regulating cell suicide and cancer ........................................ 48 3.1 Molecular functions of Myc .............................................................................................. 48 3.2 Oncogenic potential of Myc .............................................................................................. 50
3.2.1 Proliferation and differentiation by Myc ................................................................................. 50 3.2.2 Transformation mechanisms by Myc ....................................................................................... 51
3.3 Myc apoptosis .................................................................................................................... 52 3.4 Myc-induced metabolic reprogramming ........................................................................... 55
AIMS OF THE STUDY ................................................................................................... 58
MATERIALS AND METHODS ..................................................................................... 59
RESULTS AND DISCUSSION ....................................................................................... 72 4. The mitochondrial apoptosis pathway is engaged by Myc in a Bak-dependent manner
independently of cell context and death stimuli ................................................................... 72 5. The mitochondrial amplification loop as a mechanism for lethal caspase activation in
death receptor signaling ....................................................................................................... 75 6. Myc-induced Bak N-terminal exposure is a conformational change preceding Bax
activation and MOMP .......................................................................................................... 76 7. BH3 mimetic reactivates mitochondrial apoptosis in Myc mammary tumors ..................... 78 8. Cytoplasmic tumor suppressor p53 is activated by Myc to prime mitochondrial
apoptosis ............................................................................................................................... 79 9. The energy sensor AMPK is behind the Myc-mediated apoptotic sensitivity (III) ............. 81 10. Bak activation coupling oncogene-mediated metabolic stress and apoptosis ................... 82
CONCLUSIONS ............................................................................................................... 85
ACKNOWLEDGEMENTS ............................................................................................. 87
REFERENCES ................................................................................................................. 89
ORIGINAL PUBLICATIONS
The thesis is based of the following original publications, which have been assigned the following roman numerals: I
II
III Nieminen A.I.*, Eskelinen V.M.*, Haikala H.M., Tervonen T.A, Yan Y., Partanen, J.I., Klefström J. (2013). Myc-induced AMPK phospho-p53 pathway activates Bak to sensitize mitochondrial apoptosis. 110, 1839-48. IV Eskelinen VM., Haikala HM, Marques E,. Saarikoski S., Laajala D., Aittokallio T., Nieminen AI**, Klefstrom. J**, BH3 mimetic ABT-737 antagonizes Myc by aggravating apoptosis in luminal mammary tumors. Submitted.
The articles are reproduced with the kind permission of copyright holders.
∗=equal contributions
**=corresponding authors, Nieminen AI also performed experiments
ABBREVIATIONS 14-3-3 stratifin 3-PG 3-phosphoglycerate; 4-OHT 4-hydroxytamoxifen 5-FU 5-fluorouracil α-KG alpha-ketoglutarate A1/Bfl-1 Bcl-2 related protein A1 ACC acetyl-CoA Carboxylase AICAR 5-aminoimidazole-4-
carboxamide ribonucleoside AIF apoptosis-inducing factor AJ adherens junction AKT v-Akt murine thymoma viral
oncogene homolog 1 AMPK 5´adenosine monophosphate-
activated protein kinase ANT adenin nucleotide translocator APAF-1 apoptosis-activating factor-1 ARF alternative reading frame,
p14ARF ASCT2 neutral amino acid transporter ATM ataxia telangiectasia mutated ATR ataxia telangiectasia and Rad3
related protein AURKA aurora kinase A Bal1 brain-specific angiogenesis
inhibitor 1 Bak Bcl-2 homologous
antagonis/Killer Bax Bcl-2 associated X protein Bcl-2 B Cell leukemia 2 Bcl-xL Bcl2-like 1, L isoform Bcl-w Bcl-2-like 2 Bfk C1orf178, chromosome 1
open reading frame 178 BH Bcl-2 homology bHLHZ basic helix-loop-helix-zipper Bik Bcl-2 Interacting killer Bid BH3 Interacting domain death
agonist BL Burkitt´s lymphoma Bim Bcl-2 interacting mediator of
cell death Bmf Bcl-2 modifying factor Bok Bcl-2 - related ovarian killer Boo/Diva Bcl2-like 10, apoptosis facilitator BRCA1 breast cancer 1, early onset CAK cyclin-activating kinase CAM cell adhesion molecule
CARD caspase recruitment domain Casp caspase, cysteine-dependent
aspartate-specific proteases CDK cyclin dependent kinase CED-3 cell death abnormality 3 CED-4 cell death protein 4 CED-9 cell death protein 9 Chk checkpoint kinaseCK casein kinase CML chronic myelogenous leukemia c-Myc Human homologue of vMyc CSN-K COP9 signalosome associated
kinase complex C-terminal carboxy-terminal Cyt c cytochrome c DCIS ductal carcinoma in situ DD death domain DED death effector domain DISC death-induced signaling
complex including DMEM dulbecco´s modified Eagles´
medium DNA deoxyribonucleic acid DNAPK DNA protein kinase DR-4 TRAIL-R1 DR-5 TRAIL-R2 EBV Epstein-Barr virus ECM extracellular matrix EDTA ethylenediamininetetraacetic
acid EGFP green fluorescent protein EGFR EGF-receptor EMT epithelial-to-mesenchymal
transition EndoG Endonuclease G ER estrogen receptor ERBB2 v-erb-b2 avian erythroblastic
leukemia viral oncogene homolog 2, HER2/Neu
ERK extracellular signal-regulated kinase
FADH2 flavin afenine dinucleotide Fas/CD95 TNF receptor superfamily,
member 6 FADD Fas-associated protein death
domain FCS fetal calf serum
FDG-PET glucose 2-(18F)-fluoro-s-deoxy-D-glucose by positron emission tomography
FLIP flice inhibitory protein G0 Gap 0 in cell cycle G1 Gap 1 in cell cycle G6P glucose-6-phosphate
GLS glutaminase GLUT4 Glucose transporter type 4 GSK3β glycogen synthase kinase 3β HAUSP herpervirus-associated
ubiquitin-specific protein, USP7
HER2/Neu human epidermal growth factor receptor 2
HIF-1 hypoxia-inducible factor-1 HIPK2 homeiodomain-interacting HK hexokinase 2 HNPCC hereditary nonpolyposis
colorectal cancer IAP inhibitor of apoptosis ICE interleukin-convertin enzyme Ig immunoglobulin IGF-1 insulin growth factor 1 IkB nuclear factor of kappa light
polypeptide gene enhancer in B-cells inhibitor
IKK inhibitor of nuclear factor kappa-B kinase
JNK c-Jun NH2-terminal kinase kDa kilodalton LDHA lactate dehydrogenase A LKB1 liver kinase B1, PAR-4 LOH loss of heterozygosity MAC mitochondrial apoptosis-
induced channel MAPK mitogen-activated protein
kinase MAX myc-associated factor X MB myc box Mcl-1 myeloid Cell leukemia 1 MDM2 mouse double minute 2,
Hdmx human homolog MIZ-1 Myc-interacting zinc finger
protein 1 MEF mouse embryonic fibroblast MOMP mitochondrial outer
membrane permeabilization mtDNA mitochondrial DNA MYC vMyc avian
myelocytomatosis viral oncogene homolog
MycERtm Myc construct, activated conditionally by 4-hydroxytamoxifen
NADH nicotinamide adenine dinucleotide
NADPH nicotinamide adenine dinucleotide phosphate
NF-κB nuclear factor kappa-light-chain-enhancer of activated B cells
N-terminal amino-terminal OMM outer mitochondrial
membrane OXPHOS oxidative phosphorylation p16INK4A cyclin-dependent kinase
inhibitor 2A p53AIP1 p53-regulated apoptosis-
inducing protein 1 PAK Serine/threonine-protein
kinase PARP Poly(ADP)ribose polymerase PBS phosphate buffered saline PET positron emission tomography PFK2 phosphofructokinase PGC-1β PPARGC1B, peroxisome
proliferator-activated receptor gamma coactivator 1-beta
PI3K phosphoinositol 3kinase PJS Peutz-Jeghers syndrome PKR protein kinase R PKM2 pyruvate kinase M2 PMSF phenylmethylsulphonyl-
fluoride PPP pentose phosphate pathway PR progesterone receptor pRb retinoblastoma protein PS phosphatidylserine PT permeability transition PTP permeability transition pore Puma p53 upregulated modulator of
apoptosis R5P ribose-5 phosphate RIP receptor-interacting protein RNAi RNA interference ROCK-1 Rho-associated, coiled-coil
containing protein kinase 1 ROS reactive oxygen species SDS-PAGE Sodium dodecyl sulfate
polyacrylamide gel electrophoresis
shRNA Short hairpin interfering RNA
SLC7A5 large neutral amino acids transporter small subunit 1
Smac/ DIABLO Direct IAP binding protein SN2 SNAT5, Solute neutral amino
acid transporter 5 SP-1 specificity-protein 1 tBid truncated Bid TAF1 facilitates chromatin
transcription complex 1 TAK1 MAP3K7, Mitogen-activated
protein kinase kinase kinase 7 TCA tricarboxylic acid TGF-α transforming growth factor-α TGF-β transforming growth factor-β TM hydrophobic domain at
COOH-terminal TNF tumor necrosis factor TNFR tumor necrosis factor receptor TNM tumor-node-metastasis
TRAIL TNF-related apoptosis inducing ligand
TRADD TNFRSF1A-associated via death domain
TRAIL TNF-related apoptosis inducing ligand
TRAIL-R TRAIL Receptor TRAP tsartrate-resistant acid
phosphatase TRIM22 tripartite motif-containing 22 TSP-1 thrombospondin-1 VDAC voltage dependent anion
channel VEGF vascular endothelial growth factor v-Myc viral homolog of MYC wt wild type XIAP X-linked IAP
ABSTRACT
Escaping apoptosis is an important, acquired capability of tumor cells and is considered a
hallmark of cancer. Furthermore, tumor suppressors and apoptotic proteins function as
natural barriers for excessive proliferation induced by oncogenes and can be often found
as mutated or lost in cancer. Understanding how tumor cell specific apoptotic pathways
are working, may guide development of future therapeutic strategies inducing apoptosis
specifically in tumor cells. One of the highly overexpressed oncogene in cancers is Myc
that is a global transcription factor regulating variety of cellular functions such as
proliferation and apoptosis.
One of the major aims in this study was to elucidate mechanisms for Myc-induced
sensitization of cells to cell death. Herein, the results presented here provide evidence that
Myc-induced apoptosis is strictly mitochondria-dependent. Furthermore, our findings
expose new evidence for Bcl-2 family member mediated control of Myc-induced
apoptosis showing that Myc expression results in activation of Bak. Interestingly, our data
suggest a novel Bak activation step, where Myc induces Bak to undergo conformational
change and expose its N-termini, however, without further homo-oligomerization or
apoptosis. Hence, we open new ideas for the current Bak/Bax activation models suggested
earlier. In addition, our data indicate that Myc expression results in phosphorylation and
accumulations of mitochondrial p53 that is priming the mitochondria for death stimuli.
Herein, these results indicate that cytoplasmic p53 has a crucial role in Myc-induced stress
responses. The data presented here also suggest that BH3 mimetic drugs can sensitize
tumor cells to Myc-apoptosis in vivo, which also highlights the role of Bcl-xL as a key
controller of Myc-dependent apoptosis in epithelial tumors. Thereby, our results point new
therapeutic possibilities that could be used in combination to treat breast cancer.
Finally, in this study we found an interesting connection between the Myc-induced
metabolic transformation and the apoptotic signaling of Myc. We suggest that Myc
expression induces activation of metabolic pathways specific for cancer cell such as
activating the glutaminolytic pathway and promoting a catabolic mode of metabolism
through increasing fatty acid oxidation. Furthermore, our results indicate that Myc-
induced decline in the ATP contents of the cell is leading to an activation of a cellular
energy sensor AMPK. Consequently, AMPK is able to induce Bak activation and
apoptotic sensitivity via phosphorylation and stabilization of p53. Importantly, our results
bring new insight into oncogenic stress and apoptotic sensitivity by Myc highlighting the
role of metabolism. All together, it will be future challenge to understand how that
knowledge of coupled metabolic stress and apoptotic sensitivity can be used for
therapeutic interventions in cancer.
INTRODUCTION
Apoptosis is one of the key mechanisms regulating tissue homeostasis and is an important
hallmark of cancer. Moreover, apoptosis is a failsafe mechanism for damaged and
uncontrolled cell behavior and when inhibited a massive proliferation of cells can begin.
Therefore, to find efficient drugs it is intensively studied how cancer cells have acquired
the capability to inhibit apoptosis. The mitochondrial apoptosis pathway is a Bcl-2 family
regulated route that amplifies death signals and activates effector caspases that finally
destroy the cell. Increasing evidence from tumor analysis, suggest that especially the Bcl-
2 family and the proteins regulating it are crucial targets for mutations and expression
changes in cancers, leading to apoptotic abrogation.
Oncogene activation and deficiency in the tumor suppressor function are the major drivers
of tumorigenic transformation. The Myc oncogene contributes to the genesis of many
human cancers and as a transcription factor it regulated almost all cell phenotypes;
proliferation, growth, metabolic reprogramming, senescence and apoptosis. Most
importantly, the apoptosis seems to be especially abrogated in Myc-expressing tumors
highlighting the importance of understanding the molecular mechanism of Myc mediated
apoptosis.
Metabolic activity is a crucial determinant of a cell’s decision to proliferate or die. The
biosynthetic and bioenergetic needs are excessively increasing in proliferating oncogene
expressing cells. This requires a metabolic transformation capacity of the cell to bypass
such energetic stress. Furthermore, it is not fully understood how metabolic pathways such
as glycolysis communicate to cell cycle and apoptotic effectors. However, it is clear that a
complex network of signaling molecules is required to integrate metabolic inputs.
Elucidating these pathways is of great importance, as metabolic aberrations and their
downstream effects are known to contribute to the formation of cancer and more
importantly, to the treatment of cancer.
REVIEW OF THE LITERATURE
Tissues and organelles are well organized and arise by dividing from pre-existing stem
cells. Cells are differentiated to their final function and stay quiescent if meant to do so.
However, transformed cells have obtains the capability of excessive proliferation and does
not anymore respond to the programmed cell death signal leading to suicide. Most likely
such cells will form a benign or malignant tumor in future. Furthermore, malignant tumor
cells can migrate through blood and lymph vessels to distant sites in the body and invades
other tissues, form a metastasis, a secondary tumor (Weinberg, 2007).
Hanahan and Weinberg defined the “hallmarks of cancer” as a certain set of physiological
alterations that tumor cells acquire during multistep tumor progression, and which are
common for all malignant cells (Hanahan and Weinberg, 2000). Firstly, “sustained
proliferative signaling” is achieved by self-sufficiency in growth signals that enable a
tumor cell to grow in the absence of stimulatory proliferation signals (Figure 1). There are
several ways to do this such as transmembrane receptor overexpression (EGFR, HER2),
ligand-independent signaling through truncated receptor domains, integrin changes in
receptors and changes in downstream signaling molecules (H-Ras) (Hunter, 1997;
Medema et al., 1993; Slamon et al., 1987). In tumor cells, the proliferation is also
maintained by the capability to “evade growth suppressors” and being insensitive to
antigrowth signal. Often, the cell cycle clock is rescheduled and the cell transits through
the G1 phase instead of entering G0 via changes in CDK:cyclin complexes-mediated
phosphorylation of the retinoblastoma protein (pRb) (Weinberg, 1995). Also, excessive
proliferation signal by oncogenes (c-myc and erbA) induce de-differentiation and thereby
promote growth. One way of releasing proliferation is to lose contact inhibition for
example via Merlin or loss of epithelial polarity proteins (Foley and Eisenman, 1999;
Partanen et al., 2009). One feature of a tumor cell is to “evade apoptosis”, a form of
programmed cell death. Apoptotic machinery can be disrupted in multiple ways in cancer
either by inhibition of pro-apoptotic proteins by mutations in tumor suppressors or
overexpression of anti-apoptotic proteins. For instance, the tumor suppressor, p53 is
inactive in approximately 50 % of cancers either by mutations or by loss (Vogelstein et al.,
2000). In addition, constitutive survival signaling by the AKT pathway in cancer leads to
inhibition of apoptosis, and loss of tumor suppressor PTEN is one of the most commonly
mutated genes affecting apoptosis (Junttila and Evan, 2009). Also, other forms of
programmed cell death, like autophagy and necrosis can be affected (Galluzzi and
Kroemer, 2008; Hanahan and Weinberg, 2011). To be able to proliferate repeatedly, a
tumor cell has to evade senescence, in which cells enter quiescence with long-term loss of
proliferative capacity. This hallmark capability is considered as “limitless replicative
potential” (Hanahan and Weinberg, 2000). The overpassing of telomere-shortening-
induced crisis is also termed immortalization and is achieved through upregulating the
expression of the telomerase enzyme (Counter et al., 1998; Wright et al., 1989).
Figure 1. Hallmarks of cancer. Capabilities that enable tumor growth and metastasis. Modified from (Hanahan and Weinberg, 2011)
In addition, growing tumor needs oxygen and nutrient supply by the vasculature.
Therefore, one hallmark of cancer is “sustained angiogeesis”, a process where new blood
vessels grow. This is achieved by molecular changes resulting in “angiogenic switch”
(Veikkola and Alitalo, 1999). Also pericytes, endothelial supporting cells, and bone
marrow-derived cells contribute to tumor angiogenesis (Hanahan and Weinberg, 2011).
Furthermore, cancer cells need to obtain a capability to “reprogram metabolism” towards
aerobic glycolysis and catabolism. Importantly, the reason why cancer kills in 90% of
cases, is that the cancer has spread; it has migrated, invaded adjacent tissue and made new
colonies, metastases (Sporn, 1996). This “tissue invasion and metastasis” is a multistep
process and is mediated by physical coupling of cells to their microenvironment by
adherence interaction via CAMs and integrins and activation of matrix-degrading
extracellular proteases (Christofori and Semb, 1999; Werb, 1997). In addition, “tumor
promoting inflammation” and “avoiding immune destruction” are key phenotypes in
tumorigenic cell. How cancers acquire these hallmarks vary significantly, both
mechanistically and chronologically, however, “genomic instability” is one feature that
enables multistep tumorigenesis. For instance, the DNA damage is accumulating in tumors
by the loss of gatekeeper protein p53 tumor suppressor resulting in DNA repair system
(Christofori and Semb, 1999; Hanahan and Weinberg, 2011).
1.2.1 Mechanism of oncogene function
Cell growth is regulated in cells by multiple ways keeping cells in balance between
proliferation and cell death and tightly dedicated to their own differentiated function.
However, excessively activated proto-oncogenes can result in uncontrolled proliferation
and transformation of cells. These oncogenes are suggested to be activated by genetic
changes such as gene amplification, activating point mutation, chromosomal translocation
and insertional mutagenesis. These modifications affect either protein expression or
structure and can lead to constitutive activation of the oncogene. More often oncogenes
are involved in signal transductions pathways and execution of mitogenic signal.
Common oncogenes include growth factors (c-Sic), Receptor tyrosine kinases (EGFR,
VEGFR, HER), Cytoplasmic tyrosine kinases (Src), cytoplasmic serine/threorine kinases
(Raf), regulatory GTPases (Ras) and transcription factors (Myc) (Weinberg, 2007).
For instance, amplification of Her2 was reported in 30% of breast cancers. This was
shown to be consequence of the amplification of an entire chromosomal segment; an
amplicon in chromosome 17q (Tal et al., 1987). Another highly amplified oncogene is c-
myc, encoding the transcription factor and originally found as homologue to the avian
myelocytomatosis (leukemia and sarcoma) retroviral oncogene (v-myc) (Alitalo et al.,
1983). c-Myc (herein Myc) belongs to the family of myc genes (c-myc, n-myc, l-myc, s-
myc and b-myc), and its expression is deregulated in approximately one-third of human
cancers, especially in adenocarcinomas (Erisman et al., 1985). In Burkitt´s lymphomas
occurs chromosomal translocation of c-myc, where myc on chromosome 8 is placed under
control of the transcription-controlling enhancer sequences of an immunoglobulin (Ig)
gene on chromosomes 2, 14 and 22 (Dalla-Favera, 1981). The most common translocation
seems to be t(8;14) (Popescu and Zimonjic, 2002). Specifically, translocation occurs in
75% of cases in the heavy chain of Ig (IgH), but also in the k- and l-chains. In addition, n-
myc is amplified in 30% of childhood neuroblastomas. In the case of chronic myelogenous
leukemia (CML), translocation is caused by fusion of two distinct reading frames resulting
in a hybrid protein. In some cases, point mutations in the coding sequence of myc have
been found in translocated alleles of BL, however, most of the myc overexpression
mutations are absent (Bhatia et al., 1993). Less studied L-Myc has been observed in small
cell lung carcinomas (Dang, 2012). However, most often Myc expression is activated
through alterations in signaling pathways that induce or repress Myc transcription.
Another similar kind of oncogenic breakpoint cluster region of Brc-Abl gene fusion
(Philadelphia chromosome) occurs in chronic myelolytic leukemia, inducing a novel
fusion protein (de Klein et al., 1982).
In addition, tyrosine kinase signaling pathways can be constitutively active (self-
sufficiency in growth signals) because of mutation or overexpression. Indeed, the Ras
oncogene was shown to be reading frame point-mutated in 20% of human tumors in a
variety of tissues. It belongs to the family of Ras oncoproteins (h-ras, k-ras and n-ras),
which are small Guanosine-5´-triphosphate hydrolase (GTPases), having enzymatic
activity required for efficient signal transduction by growth factors (Malumbres and
Barbacid, 2003). The Ras pathway induces Raf and MAP kinase signaling leading to ERK
and PI3K activation. Later it was understood that Ras requires additional oncogene
cooperation, for example, with Myc or E1A (Land et al., 1983; Newbold and Overell,
1983). Moreover, KRAS mutations are found from colon, lung and pancreatic carcinomas,
whereas HRAS from breast cancer.
1.2.2 Tumor suppressors
To avoid uncontrolled cell proliferation, vertebrates have a well-designed growth control
system and various tumor suppressive mechanisms. To discriminate accurately between
normal and neoplastic growth, sensors called tumor suppressors function in diverse
cellular activities, including cell cycle checkpoint, DNA damage repair and, for example,
in protein degradation. Mutations in tumor suppressor genes behave recessively and a
single copy of the gene is enough for protein function (Weinberg, 2007). Therefore,
neoplastic growth requires two successive somatic mutations that occur sporadically. First,
the “classic” tumor suppressor gene, retinoblastoma (Rb), which is a transcriptional
repressor of cell cycle genes in the G1 phase, was found in retinoblastoma, childhood eye
cancer (Knudson, 1971; Sherr, 2004). In this cancer, the wild-type allele of the Rb gene
can be eliminated by mitotic recombination and results in a loss of heterozygosity (LOH),
where the normal copy of the gene is lost as well (Weinberg, 2007). Another important
epigenetic mechanism for inactivating tumor suppressor genes is promoter methylation,
where CpG sequences are methylated. Examples of methylated genes in tumors are the
tumor suppressor p16INK4A that was found in smokers and the BRCA1 gene, which is lost
in familial breast and ovarian cancers. From the same locus as p16INK4A, an another tumor
suppressor gene is encoded, the alternative reading frame, ARF protein that protects p53
from MDM2-mediated degradation (Sherr, 2001). Loss of ARF co-operates with
oncogenes in tumorigenesis in mice, since the apoptotic pathway is abrogated. Another
group of tumor suppressors are originally genes known to affect development, such as
Smad4 and PTEN, and misregulation of these might induce de-differentiation and enhance
tumorigenicity (Sherr, 2004).
Importantly, the DNA damage response and genomic instability are major causes for
mutagenesis in cancer. Furthermore, DNA mismatch repair genes contribute to cancer
development and are found mutated in HNPCC colorectal cancer with microsatellite
instability. Importantly, disruption of the mismatch repair system has been suggested to
lead a “mutated” phenotype in which resulting genetic instability ultimately targets other
oncogenes and tumor suppressors (Loeb et al., 2003). In addition, several DNA damage
activated kinases, ATM (ataxia telangiectasia), CHK2, and DNA-PK, are found mutated
in cancers. Furthermore, mutations in the BRCA genes are the main cause of familial
breast and ovarian cancers (Sherr, 2004) .
Transcription factor and tumor suppressor protein TP53 (tumor protein, p53) is a critical
coordinator of wide range of stress responses. Originally, p53 was discovered in 1979
from studies on SV40 T antigen by investigators from six groups (Kress et al., 1979; Lane
and Crawford, 1979). Interestingly, p53 was first described as an oncogene, not tumor
suppressor. Recent evidence again suggest that mutated p53 drives tumorigenesis having
gain-of-function properties (Brosh and Rotter, 2009; Lane and Benchimol, 1990; Muller
and Vousden, 2013). Later on, Lane et al. described p53 as the “guardian of the genome”
and Levine et al. as the “cellular gatekeeper” demonstrating the important role of p53 in
antiproliferative responses (Lane, 1992; Levine, 1997). Furthermore, the p53 is shown to
be mutated in a variety of cancers and is currently considered a most mutated gene in
cancer since one half of human cancers express a mutant p53 (Olivier et al., 2010). In
normal unstressed cells, p53 activity is maintained at low levels through a combination of
p53 degradation in ubiquitin-proteosome, principally mediated by Mdm2 (Hdm2, human
homologue) (Wade et al., 2006). In addition, Mdm2 is transcribed by p53 itself via a
negative feedback loop and it recognizes p53 as a target that should be ubiquitylated
shortly after its synthesis and finally destroyed. This also partly explains why mutated
p53 is accumulating since p53 lose its transcription-activating powers and Mdm2 is not
transcribed anymore.
Oncogenic signaling and cellular stress (hypoxia, nutrient deprivation) or DNA damage
signals increase p53 activity, which triggers apoptosis, senescence and repair programs
(Lowe et al., 2004). In addition, variety of covalent modifications of p53 occurs when p53
is stabilized and activated, many affecting the C-terminal domain (Figure 2). These post-
translational modifications include acetylation, glycosylation, methylation,
phosphorylation, ribosylation, sumoylation and ubiquitination (Weinberg, 2007; Xu,
2003). These modifications affect the ability of p53 to interact physically with other
transcription factors that modulate its transcriptional capacity and finally enable the
transcription of target genes. Furthermore, p53 is phosphorylated at multiple sites at N-
and C-terminus by a number of kinases. For instance, phosphorylation of p53 amino acid
residues at the N-terminal transactivation domain by kinases blocks Mdm2 binding and
stabilizes p53 (Figure 2). Additionally, domains near the N-terminus contribute to the
apoptotic function of p53. Whereas, the center of p53 acts as a DNA binding domain and
are phosphorylated. Moreover, the C-terminus possesses an oligomerization domain and
when phosphorylated allows p53 to form tetramers (Xu, 2003). In addition, near the C-
terminus exist also nuclear localization signals.
Figure 2. Post-translational modifications of p53. TAD, transactivation domain; SH3, Src homology 3-like domain; NLS, nuclear localization signal; DNA binding domain; TET, tetramerization domain; NES, export signal; REG, carboxy-terminal regulatory domain. Kinases regulating major phosphorylation sites of human p53 are shown and their effect on p53 functions (pink). DNA “hot-spot” mutation area in DNA binding domain (red) and areas for other post-translational modification (blue) are shown. Modified from (Bode and Dong, 2004).
However, several mechanisms have been suggested to induce the loss of p53 function in
cancer. Indeed, p53 is deleted for example in colorectal cancer or highly mutated in
inherited cancers, such as in Li-Fraumeni syndrome, and in a variety of sporadic cancers
(Srivastava et al., 1990). Furthermore, some cancer-inducing DNA viruses, such as SV40
or HPV, encode proteins like E6 that target p53 for degradation and, thereby inactivate
p53. Alternatively, some upstream pathways of p53 are mutated in cancer (Muller and
Vousden, 2013). Normally p53 exists as a homotetramer of four identical polypeptide
subunits. Importantly, more than 75% of the mutations result in the expression of a p53
protein that has lost wild-type function and may exert dominant-negative regulation over
any remaining wild-type p53 thereby inhibit its function. Interestingly, mutant p53 also
acquires oncogenic functions, a so-called gain-of-function mutant, that is entirely
independent of wild-type p53 for instance metabolic capacities (Muller and Vousden,
2013). The primary alteration in p53 is a single amino acid substitution in the 393-amino
acid protein. Most of the mutations cluster within the central DNA-binding domain and a
number of hotspots (Figure 2) have been identified (Muller and Vousden, 2013).
Additionally, mutations can occur in the N-terminal transcriptional transactivation domain.
It is also common that the half-life of the mutant p53 is increased because of changes in
MDM2 interactions, and thereby mutant p53 becomes stabilized and expressed to a higher
level in tumor cells (Bartek et al., 1991). In breast cancers, mutated p53 can be found in
the cytoplasm of tumor cells, indicating a transcriptional inactivation of p53 or folding of
the p53 caused by R175H, structural mutants (Moll et al., 1992).
Majority of p53 functions are regulated through transcriptional regulation of target genes.
When p53 is induced by DNA damage, it blocks forward progress at the G1 phase and
thereby, p53 acts as “guardian of the genome” and drives a cell cycle “checkpoint”
function by transcribing. An important target gene of p53 is p21, which is an inhibitor of
CDK2 and Cdc2 that are active in the late G1, S, G2, and M phases of the cell cycle.
Furthermore, p53 is shown to transcribe several genes (Figure 3) when inducing cell cycle
arrest and when mediating DNA repair (reviewed in Riley et al., 2008). In addition, mouse
models of p53 knockouts show increased genomic instability and broader tumor spectrum
(Fridman and Lowe, 2003; Lowe et al., 2004). The p53 has been also suggested to regulate
apoptosis both transcription-independent manner and -dependent manner, where it directly
transcribes proteins like Puma and Bax (Speidel, 2010). Several oncogenes, like E1A,
Myc and E1F, induce p53 and therefore inactivation of p53 severely compromises
oncogene-induced apoptosis. Moreover, studies in mice indicate that disruption of p53-
mediated apoptosis co-operates with oncogenes in multiple mouse cancer models. In
addition, p53 can promote senescence in MEFs and senescence is a prerequisite for the
transformation of cells (Lowe and Sherr, 2003). Also, p53 transcribes for example TSP1,
which is a blocker of the development of new blood vessels and is shown to be inhibiting
angiogenesis. Emerging evidence also suggest that p53 regulates metabolic pathways such
as glycolysis and thereby enable stress adaptation (Berkers et al., 2013). Recently, more
transcription-independent functions of p53 has been emerging such as regulation of the
microRNA expression and maturation pathway and, also, direct role in DNA repair
process (Sengupta and Harris, 2005; Suzuki and Miyazono, 2013). Transcriptional
independent functions of p53 in apoptosis will be discussed more in 2.3.1.3.
Figure 3. The molecular target and effects of tumor suppressor p53. Modified from (Brown et al., 2009).
LKB1 (PAR-4) is a tumor suppressor protein and a serine/threonine kinase 11, lost in an
autosomal inherited Peutz-Jeghers syndrome (PJS) cancer patients (Jeghers et al., 1949;
Martin and St Johnston, 2003). Patients typically develop multiple benign gastrointestinal
polyps, hamartomas, and mucocutaneous pigmentation in different area and, they are at
risk for intestinal or extra-intestinal cancer (Martin and St Johnston, 2003; Vaahtomeri and
Makela, 2011). In addition, Lkb1 is one of the most commonly mutated genes in sporadic
human lung cancer (NSCLC) and, carriers of germline Lkb1 mutations have increased
breast cancer incidence (Hearle et al., 2006; Shackelford and Shaw, 2009). In tumors,
Lkb1 mutations are suggested to correlate with aggressive phenotypes of tumors and cause
increased invasiveness in mouse models (Vaahtomeri and Makela, 2011). Mammalian
Lkb1 has been implicated as a regulator of multiple biological processes: including
angiogenesis, cell cycle arrest, transformation (Wnt- and Ras pathways), p53-mediated
apoptosis, polarity, and energy metabolism (reviewed in Baas et al., 2004b). Lkb1 is
regulated via binding partners that anchors Lkb1 in the cytoplasm. Furthermore, the kinase
activity is crucial for the tumor suppressive function of Lkb1 (Ylikorkala et al., 1999).
Several groups identified Lkb1 as an upstream kinase and regulator of 5´AMP-activated
protein kinase (AMPK), which is a metabolic stress regulator. Thereby, Lkb1 has been
suggested to act as a tumor suppressor of metabolic transformation of cancer cells
(Shackelford and Shaw, 2009). In contrast, Lkb1 was shown to activate AMPK by
phosphorylating Thr172 in the regulatory activation loop, T-loop (Hawley et al., 1996).
The Lkb1 kinase activity is regulated via the Lkb1-STRAD-MO25 complex resulting in
activation of the AMPK and its related kinases (ARKs, 14 in total) (Baas et al., 2004b;
Vaahtomeri and Makela, 2011). In addition, Lkb1 inhibits proliferation and growth by
affecting mitogenic pathways such as mTOR (S6K1). Lkb1 promotes also senescence in a
p53-independent manner via Nuak2 and, in contrast, cell cycle arrest and cell death in a
p53–dependent manner (Humbert et al., 2010; Tiainen et al., 2002). Part of the Lkb1-
induced regulation of p53 is suggested to occur by direct Lkb1-mediated phosphorylation
of p53 Ser15, or its substrate kinases, AMPK, in response to glucose starvation or cell
detachment (Jones et al., 2005).
Lkb1 belongs to a family of partitioning-defective (PAR) protein that are required for
asymmetrical cell division of the C. elegans zygote. In addition, loss of Lkb1 was
discovered to disrupt polarization of the oocyte cytoskeleton resulting in a loss of anterior-
posterior cell polarity in Drosophila melanogaster and is required for apical-basal polarity
in epithelial cells (reviewed in Baas et al., 2004a). In epithelia, the cell-ECM interface
defines the basolateral membrane, whereas highly organized tight (TJ) and adherens
junctions (AJ) form a physical border between the apical and basolateral membrane
domains of a cell (reviewed in Baas et al., 2004b; Shin et al., 2006). In general, junctions
form a physical border between the apical (extending to the lumen) and basolateral
membrane domains of a cell provide strength to the epithelial sheet by serving as
anchoring sites for the cytoskeleton. Polarity is formed when apical microvillis are
assembled, TJs and AJs are formed, and finally the apical and basolateral surface markers
are ordered. Furthermore, cell-ECM contacts by integrins and cell-cell contacts by
cadherins regulate the transition of a nonpolarized to polarized epithelial cell transition
(Drubin and Nelson, 1996). Activation of LKB1 in mammalian epithelial cell lines results
in the cell´s intrinsic polarization via actin cytoskeleton reorganization and junctional
protein ZO-1 re-localization and involves STRAD (Baas et al., 2004a). The potential role
of Lkb1 in the regulation of polarity is supported by data that Lkb1 deletion in mouse
mammary glands leads to development of ductal carcinomas (McCarthy et al., 2009).
Several studies have linked the polarity functions of Lkb1 with its kinase activity and
downstream target AMPK, for example, AMPK null mutations lead to a strong polarity
phenotype with defects in Drosophila (Mirouse and Billaud, 2011). In contrast, in PJS
polyposis Lkb1 is thought to operate via TGF-β, and regulate cell polarity by controlling
microtubule and actin dynamic by Smad4, Nuak2 or p53 (Vaahtomeri and Makela, 2011).
AMPK is an evolutionarily conserved fuel-sensing serine/threonine protein kinase enzyme
that is activated under stress such as: hypoxia, ischemia, glucose deprivation, and exercise
(Steinberg and Kemp, 2009). Additionally, AMPK is suggested to be a metabolic tumor
suppressor (Figure 4), whose activity is often decreased in metabolic disorders like insulin
resistance, obesity, and Type 2 diabetes. Several epidemiological studies indicate that
metabolic syndrome increases the risk of cancer and thus, patients with Type 2 diabetes
taking well-accepted AMPK activator, metformin, have a reduced risk of cancer (Hardie,
2007; Luo et al., 2010). However, very little data exists on reduced AMPK activation in
cancer.. For example, AMPK mRNA levels inversely correlate with clinical prognosis in
breast and ovarian tumors and AMPK activation is reduced in both lung cancer specimen
containing loss-of function mutations of LKB1 and in breast cancer specimens (Conde et
al., 2007). In addition, a large number of studies have shown that activation of AMPK (by
pharmacological agents, exercise and dietary restriction) can attenuate cancer cell growth
in vitro and inhibit tumor development in vivo (Jiang et al., 2008). Conversely, as energy
stress sensor, AMPK activates catabolic pathways, which can promote tumorigenesis and
will be discussed later.
Figure 4. AMPK molecular targets and effects. Modified from (Liang and Mills, 2013).
AMPK is a heterotrimeric complex consisting of three subunits: a catalytic subunit (α),
and two regulatory subunits (β and γ) (reviewed in Hardie, 2007). Under metabolic stress
such as adequate carbon source and oxygen, the intracellular AMP levels or the AMP to
ATP ratio (ADP/ATP goes down) is increased and AMP is bound to the γ-subunit.
Subsequently, AMPK undergoes allosteric activation, thereby protecting it against
phosphatases that dephoshorylate threonine 172 in the activation loop (Hardie, 2007).
Several other AMPK activators are known, even though their exact mechanism is unclear
(cytokines, activating drugs: metformin, phenmorfin, AICAR, A-769662, and some
natural plant products) (Hardie, 2007). Thus far, several kinases have been identified to
phosphorylate threonine 172 on the catalytic subunit, leading to its activation. Previous
data suggests that in most scenarios, the LKB1 is kinase responsible for AMPK
phosphorylation (Shaw et al., 2004). However, AMPK is also a downstream substrate of
calmodulin-dependent protein kinase (CAMKK-β), which is activated due to increased
intracellular Ca2+ levels and TAK1 kinase (Hawley et al., 2005).
As a regulator of energy homeostasis (Figure 4) the activated AMPK inhibits lipogenesis.
It simultaneously stimulates fatty acid oxidation (ACC-2) and inhibits fatty acid synthesis
(ACC-1) in order to generate more ATP to cope with acute energy demands, and also
inhibits anabolic processes that consume ATP (Luo et al., 2010). In addition, AMPK
activation enhances insulin sensitivity, inhibits hepatic glucose production, stimulates
glucose uptake (by GLUT4 translocation) in muscle, and esterification alleviates
hyperglycemia and hyperlipidemia (Luo et al., 2010). The mechanisms behind these
functions involve down-regulated expression of biosynthetic genes involved in
gluconeogenesis and lipogenesis (Figure 4). Controversially, AMPK suppresses glycolysis
in tumor cells by inhibiting mTOR and on the other hand, it stimulates glycolysis by
activating PFK2 (Marsin et al., 2002).
Likewise, AMPK activation can regulate various processes (Figure 4), including cell cycle
checkpoint, cell polarity, senescence, autophagy, apoptosis, and pro-inflammatory
responses (Hardie, 2007; Shackelford and Shaw, 2009). Pharmacological activation of
AMPK is shown to inhibit growth of cancer cells by causing phosphorylation of p53 on
Ser15 and accumulation of p53 (Imamura et al., 2001). Also, glucose deprivation induced
arrest in MEFs is mediated by AMPK-mediated phosphorylation of p53 (Jones et al.,
2005). AMPK switches off protein synthesis by inhibiting elongation-factor-2 or
inhibiting the translation initiation step through rapamycin (TOR) pathway (Inoki et al.,
2003). Inhibition of the TOR pathway is also involved when AMPK is causing autophagy
(Inoki et al., 2003). Furthermore, at least a part of LKB1´s role in regulation of apico-basal
polarity in D. melanogaster undergoes through its downstream target AMPK (Lee et al.,
2007). Likewise, activation of AMPK induces repolarization of epithelial kidney and
intestinal epithelial cells by assembly of TJs (Hardie, 2007; Inoki et al., 2003). Also,
additional mechanisms for AMPK-induced polarity have been suggested recently, such as
the activation of Myosin II or, alternatively, through NUAK1 (Mirouse and Billaud, 2011).
Interestingly, recent studies indicate that AMPK plays a role in linking metabolic
syndrome and cancer. AMPK is an essential mediator of the tumor suppressor LKB
1 and could be suppressed in cancer cells containing loss-of-function mutations of LKB1
or in cancers associated with metabolic syndrome. Finally, AMPK may function as a
metabolic tumor suppressor regulating glucose, lipid, and protein metabolism.
1.2.3 Cancer metabolism
During tumor development, a growing tumor is soon running into nutrient and oxygen
lack and to bypass such metabolic stress and continue proliferation, tumor must undergo a
period of metabolic adaptation to survive or alternatively cells undergo apoptosis. The
altered metabolism found in cancer was discovered by Otto Warburg in 1920s, when he
described a phenomenon termed the ”Warburg effect”; an increase in glycolysis that is
maintained in conditions of high oxygen tension (“aerobic glycolysis”) and gives rise to
enhanced lactate production (Warburg et al., 1927). Now it is recognized that several
changes occur in the tumor cell´s metabolism, which result in a “metabolic
reprogramming” involving aerobic glycolysis, de novo lipid biosynthesis and glutamine-
dependent anaplerosis, glutaminolysis (Figure 5). These changes support growth and
allow necessary nutrients, energy (rapid ATP production,) and biosynthetic activity
(macromolecules) that are required for the increased proliferation of tumor cells
(DeBerardinis et al., 2008). Moreover, redox status is crucial for cancer cells, and low
levels of reactive oxygen species (ROS, byproducts of metabolic processes) can benefit
tumor cell´s proliferation for instance by activating Src kinase or inactivating PTEN.
However, at high levels ROS starts to induce damage and death. To counter this ROS-
induced oxidative stress, cancer cells also have a highly active antioxidant defense-system
still allowing, for example, moderate levels of ROS to induce mutagenesis (reviewed in
Cairns et al., 2011). Another important feature of cancer cells is increased nutrient uptake,
such as glucose uptake, that can be taken advantage of when imaging primary tumors and
metastases in clinics using the FDG-PET technique (Mankoff et al., 2007).
Several reasons explain why enhanced glucose uptake for glycolytic ATP generation or
anabolic reactions constitutes an advantage for tumor growth. Cancer cells rely on aerobic
glycolysis instead of oxidative phosphorylation (OXPHOS) for ATP production and
generate bicarbonic and lactic acids as end products of glycolysis (Pouyssegur et al., 2006).
Once glucose is transported inside of the cells it is converted to pyruvate that subsequently,
enters either the mitochondria for further conversions to acetyl-CoA, or is immediately
converted to lactate (reviewed in Cairns et al., 2011). In normal cells, glucose is converted
into acetyl-CoA and its complete oxidation occurs through the mitochondrion-localized
tricarboxylic acid (TCA) cycle. Furthermore, oxidative phosphorylation produces electron
donors NADH and FADH2, which donate electrons to the respiratory chain complexes I
and II, respectively CO2 and H2O as end products (total net of 38 ATP molecules). In
contrast, in tumor cells glucose is converted only to pyruvate (generates 2 ATP molecules),
and subsequently to waste products. These anaerobic waste components, acids, have their
own effect on the microenvironment like favoring tumor invasion, suppressing anticancer
immune effectors, and/or they are used by other cells (Kroemer and Pouyssegur, 2008).
Moreover, tumors can metabolize glucose through the pentose phosphate pathway (PPP)
to generate NADPH, which is required for macromolecule biosynthesis and also
contributes to fatty acid synthesis. This is also a way in which cancer cells obtain
antioxidant defenses against a hostile microenvironment and chemotherapeutic agents
(Gatenby and Gillies, 2004). Furthermore, tumor cells can express the PKM2 isoform of
pyruvate kinase, than can surprisingly inhibit glycolysis or slow it down, thereby
promoting shuttling of pyruvate to the PPP instead (Schulze and Harris, 2012). Moreover,
cancer cells use or deviate intermediates of the glycolytic pathway for anabolic reaction
(cataplerosis), which in part enables increased growth (Figure 5).
In addition, pyruvate may enter a truncated TCA cycle resulting an exportation of citrate
from the mitochondrial matrix (Figure 5). Subsequently, its cleavage products acetyl-CoA
becomes available for the synthesis of fatty acid and cholesterol and furthermore,
oxaloacetate (OAA) is converted to malate and reimported to mitochondria into a citrate
cycle (reviewed in Kroemer and Pouyssegur, 2008). Importantly, it is speculated that the
reason why tumor cells use glycolysis is that it benefits both bioenergetics and
biosynthesis and if the glycolytic rate is high enough, a similar yield of ATP is produced
as in oxidative phosphorylation. Lactate is not used in the Warburg effect, and it starts to
accumulate in the cells. However, even though more pyruvate could be oxidized, it is not
done in cancer cell and instead, the cells need to get rid of excess pyruvate via a high-flux
mechanism, and pyruvate is converted to lactate by lactate dehydrogenase A (LDH-A)
(DeBerardinis et al., 2008). Furthermore, in glutaminolysis glutamine is oxidized in the
TCA cycle, allowing proliferating cells to use TCA cycle intermediates as precursors for
biosynthesis (Yuneva et al., 2007).
Figure 5. Metabolic alterations in cancer. Metabolic reprogramming of cancer cells involves activation catabolic pathways (b-oxidation, glycolysis, glutaminolysis), which release carbon for bioenergetic reactions and promotes biosynthesis of nucleic acids. In addition, biosynthesis requires enhanced nutrient uptake. Part of the biosynthesis required for cell growth comes from cataplerosis, where metabolic intermediates are directed for synthesis, for example, glucose 6-phosphate of glycogen and ribose 5-phosphate synthesis, dihydroxyacetone phosphate for triglyceride and phospholipid synthesis, and pyruvate for alanine and malate synthesis. Caner cell have Truncated TCA cycle, in which citrate is exported into the cytosol and used for lipogenesis. Tumor cells also produce ATP via oxidative phosphorylation (OXPHOS), which imposes an additional metabolic burden on the TCA cycle. Modified from (Galluzzi et al., 2013). In general, the metabolism in cancer cells is regulated autonomously without the
requirement of growth-factor-induced signals. Variety of molecular changes explains this
cancer-specific “metabolic reprogramming”, such as mitochondrial DNA mutations,
oncogene activation, mutation in the Ras and PI3K/Akt/mTOR pathway and aberrant
activation of the AMPK-LKB1 pathway (Cairns et al., 2011). For instance, constitutive
cellular stabilization of (hypoxic-) stress- and oncogene-induced HIF-1 is one major
reason for aerobic glycolysis as decreasing conversion of pyruvate to acetyl-CoA, and this
compromises OXPHOS (Harris, 2002). In contrast, tumor suppressor p53 maintains the
respiration, OXPHOS, by increasing the expression of cytochrome c oxidase 2 and down-
regulates phosphoglycerate mutase. Moreover, p53 transcriptionally activate hexokinase-2
and TIGAR, which inhibits phosphofructokinase activity, lowers the levels of FBP, and
hence inhibits glycolysis while channeling glucose to the PPP (Bensaad et al., 2006). In
addition to the genetic changes that alter tumor cell metabolism, the abnormal tumor
microenvironment such as hypoxia, pH and low glucose concentrations have a major role
in determining the metabolic phenotype of tumor cells (Cairns et al., 2011). Another
structural feature of tumor cells is that mitochondria are often small, lack cristae, and are
deficient in the F1 subunit of ATP-synthetase. However, there is no evidence that
mitochondrial respiration would be functionally impaired (Kim and Dang, 2006; Kroemer
and Pouyssegur, 2008). Interestingly, the resistance of cancer cells mitochondria to
apoptosis-associated permeabilization and subsequent cytochrome c release, and the
altered contribution of these organelles to metabolism, are closely related. For example,
increased glycolysis in tumor cells is accompanied by increased stability of the
mitochondrial membrane partly by membrane-associated hexokinase (HK) I/II
upregulation in tumors. Furthermore, defects in OXPHOS induce apoptosis resistance
(Tomiyama et al., 2006). Other alterations in cancer and disabled apoptosis are
hyperpolarization of the inner mitochondrial transmembrane potential and deficiency in
voltage-gated plasma membrane K+ channels (Kroemer et al., 2007).
Mammary gland is an organ that produces milk in the female breast and consists of
epithelial cells lining the walls of alveoli, where the milk is secreted, and surrounding
myoepithelial cells. The alveoli join to form groups known as lobules ending in the nipple.
Beneath the epithelial cell layers lies a basement membrane, which separates the epithelial
cells from the underlying layer of connective tissue cells, the stroma. Furthermore, the
basement membrane consists of extracellular matrix (ECM), which together with
adipocute, fibroblast, inflammatory cells constitute mammary stroma (Watson and Khaled,
2008). Importantly, breast cancer affects one in eight women during their lifetime. Breast
cancer can be classified by different schemata, which reflect treatment responses and
prognoses. Categories include histopathological type, grade (1; well-differentiated and
best prognosis, 2; moderately-differentiated and medium prognosis, 3; poorly-
differentiated and worst prognosis), stage (tumor-node-metastasis; TNM), receptor status
and the presence or absence of genes as determined by DNA testing. The majority of
breast cancers derive from epithelia and are classified as mammary ductal carcinoma
(Pinder, 2010). Ductal carcinoma in situ (DCIS, 13%) is proliferation of cancer cells
within the epithelial tissue without invasion of the surrounding tissue and is harmless.
However, 60% of the lesions will become invasive over the course of 40 years in follow-
up (Zorbas et al., 2004). In contrast, invasive ductal carcinoma (55%) and invasive lobular
carcinoma (5-10%) invades the surrounding tissues. Treatment options always include
surgery for removal of the main tumor mass, or mastectomy (removal of the breast) and
adjuvant therapy includes chemotherapy, radiotherapy, hormonal therapy, and/or targeted
therapy.
The receptor status of the breast cancer is identified by immunohistochemistry, and it
defines the presence of estrogen receptor (ER), progesterone receptor (PR), and HER2. In
addition, tumors also exhibit different gene expression profiles and based on mRNA
profiling data, breast cancers have been classified into 5 subgroups: luminal A and luminal
B, HER2 amplified, basal-like, and normal-like (Perou et al., 2000). Of these, tumors of
the luminal type are generally ER/PR positive, whereas the basal-like and normal-like
subtypes are triple-negative (negative for ER, PR and HER2 receptors). In addition to lack
of receptors, the characteristics of the basal-like subtype are high expression of basal
markers such as cytokeratin 5/6 and/or increased expression of EGFR. Receptor status is a
critical assessment for all breast cancers as it determines the suitability of using targeted
treatments. ER positive cancer cells depend on estrogen for their growth, so they can be
treated with drugs to reduce either the effect of estrogen (e.g. tamoxifen) or the actual
level of estrogen (e.g. aromatase inhibitors). HER2 positive cells respond to drugs such as
the monoclonal antibody, trastuzumab (Gonzalez-Angulo et al., 2007). Conversely, triple
negative (ER-, PR-, HER2) cancer, lacking targeted treatments has a poorest prognosis
(Oakman et al., 2010). Additionally, DNA microarrays can be used for further
categorizing of breast cancer subtypes, such as basal-like.
Furthermore, Myc deregulation contributes to breast cancer initiation and progression and
is associated with poor outcomes (Robanus-Maandag et al., 2003). Myc is amplified
approximately in 16 % of all breast cancers and overexpressed 20 45% of cases,
depending on the study (Bieche et al., 1999; Chrzan et al., 2001; Naidu et al., 2002). The
basal-like breast cancer constitutes one of the most challenging subtypes of breast cancers
accounting for 15% of breast cancer cases. In addition, the basal-like breast cancer is
responsible for most breast cancer deaths, since they lack adequate hormonal therapy.
Importantly, high Myc pathway activity has been implicated in basal-like breast tumor
subtype based on pathway analyses from molecular profiling (Xu et al., 2010).
Cell death, or “cell suicide”, is essential for proper tissue homeostasis and adaptation to a
changing environment and the importance of this phenomenon is emphasized by the fact
that the components of cell death are well conserved (Adams 2003). There are many
triggers for cell death, such as physical or chemical stress, parasites or developmental
signals (embryogenesis and metamorphosis). Originally, Kerr, Wyllie and Currie were the
first to introduce the term “apoptosis” as a form of programmed cell death, to describe the
morphological process leading to controlled cellular self-destruction (Kerr et al., 1972).
Importantly, dysfunction or deregulation of the apoptotic program is implicated in a
variety of pathological conditions like cancer, autoimmune diseases, and the spreading of
viral infections, while neurodegenerative disorders, AIDS, and ischaemic diseases are
caused or enhanced by excessive apoptosis. Currently, cell death types involve: apoptosis,
autophagic cell death, necrosis, necroptosis and pyroptosis, which all are characterized by
a number of distinguishing features and morphological changes (Degterev and Yuan,
2008). First of all, in apoptosis the contents of the dying cell remain contained in
membranes. Another feature is that the cell shrinks, shows deformation and detaches from
surrounding cells, (loss of adhesion), the plasma membrane shows blebbing, and finally,
the cell fragments into smaller, membrane-bound structures known as “apoptotic bodies”.
In addition, the nucleus undergoes characteristic changes, including chromatin
condensationand internucleosomal cleavage of DNA leading to an oligonucleosomal
“ladder” appearing in agarose gel (Wyllie et al., 1980). One biochemical feature is
proteolytic cleavage of a number of intracellular substrates, however, organelle integrity is
still maintained. Finally, the apoptotic bodies are phagocytosed in a relatively short time
by macrophages and parenchymal cells, which recognize the externalized PS on the outer
membrane (reviewed in Saraste and Pulkki, 2000). Thus, in most of the cases, apoptotic
bodies are removed from the tissue without a subsequent inflammatory response, however,
if not phagocytosed, the cell will undergo degradation resembling necrosis.
Autophagic cell death (macroautophagy) differs from apoptosis and is less well
understood. It involves a cellular mechanism of self-eating, autophagy, which can finally
lead to cell death. However, autophagy can also be involved in the turnover of long-lived
proteins and whole organelles (mitophagy or reticulophagy), or in the cell´s adaptation to
starvation-induced stress and lack of nutrients (Maiuri et al., 2007). During autophagy, the
cytoplasm is engulfed by specific double- or multi-membraned autophagic vacuoles
(autophagosomes) and later digested by lysosomal enzymes. Importantly, apoptosis and
autophagy are sometimes overlapping and can serve as an alternative choice if the other
one is inhibited. The third type of death is necrosis, in which cells are suffering a major
insult, resulting in a loss of membrane integrity, swelling and disruption of the cells
leading to a release of cellular contents. Thus causing damage to surrounding cells, and a
strong inflammatory response in the corresponding tissue. However, there are forms of
necrosis that are programmed, termed “necroptosis” (Golstein and Kroemer, 2007). In
addition, “pyroptosis “ is a programmed cell death form associated with antimicrobial
responses during inflammation (Fink and Cookson, 2005).
Apoptosis is mediated by signaling pathways, the mitochondrial (intrinsic) and death
receptor apoptotic (extrinsic) pathway, finally leading to disruption of the cell (Fulda and
Debatin, 2006). In addition, this process is regulated by two main groups of proteins;
caspases and the Bcl-2 family. The history of finding key regulators of apoptosis and
vertebrate homologs began with studies on C. elegans and its molecular machinery
required for proper development: proapoptotic caspase CED-3 and adaptor protein CED-4
and anti-apoptotic CED-9 (Bcl-2 homolog) (Ellis and Horvitz, 1986). Currently it is
appreciated that the balance between the Bcl-2 family interactions regulates the
mitochondrial apoptosis pathway, which upon activation further results in caspase
activation. Moreover, caspases can be catalytically activated to initiate a caspase cascade,
leading to degradation of the cellular proteins.
2.2.1 Caspases as executioners Caspases are cysteine-dependent aspartate-specific proteases residing in inactive form as
zymogens. Once activated, these endopeptidases recognize cysteine-residue in peptide
motif, QACRG, within a proteolytic substrate and cut that protein internally after the
aspartic acid residues (Stennicke and Salvesen, 1997). Since finding the human homolog
of C. elegans´s CED-3, the ICE (interleukin-1b-converting enzyme), 14 distinct
mammalian caspases has been identified in total, of which there are 12 in humans.
Subsequently, the family of apoptotic caspases has been divided into two major classes:
the initiator caspases including procaspase-2, -8, -9 and -10 (Dronc and Dredd in D.
melanogaster); and the effector (or executioner) caspases, which include procaspase-3, -6,
and -7 (Drice, Decay, Damm, Dcp1 and Stripa in D. melanogaster) (reviewed in Degterev
et al., 2003; Hay and Guo, 2006; Riedl and Shi, 2004). The ICE family contains caspase-
13, -5, -4, and -1, which mostly mediate cytokine maturation during inflammation,
however caspase-1 also mediate pyroptosis (Bergsbaken et al., 2009).
Inactive procaspases exists as a monomer carrying at its N-terminus the pro-domain,
followed by a large and a small subunit, which are sometimes separated by a linker
peptide (Figure 6). Activation of initiator caspases occurs at signaling complexes via pro-
domains that are protein-protein interaction motifs, whereas effector caspases are activated
by initiator caspases. The maturation of initiator procaspases at signaling complexes
involves dimerization, autoactivation, and proteolytical processing between the large and
small subunit, resulting in a separation of these subunits (reviewed in Riedl and Shi, 2004).
The recruitment to signaling complexes can happen either in response to the ligation of
cell surface death receptors or in response to signals originating from inside the cell.
Moreover, the pro-domains are mediating the interaction between caspases and adaptor
proteins, and caspase homodimerization. In the case of procaspase-8 and -10, the pro-
domain is called the death effector domain (DED) and in the case of procaspase-9 and -2
the caspase recruitment domain (CARD) (Figure 6). DEDs and CARDs are not related by
sequence homology, but they are structurally similar, each having a “death fold” structure.
Furthermore, caspase-8 and -10 are recruited to the death-inducing signaling complex
(DISC), caspase-2 to PIDDosome and caspase-9 to apoptosome (Boldin et al., 1995; Tinel
and Tschopp, 2004; Zou et al., 1999).
Figure 6. Structure and classification of mammalian caspases. Caspases can be classified on the basis of their function in apoptosis, inflammation or differentiation. All caspases are characterized by a catalytic domain (p20 and p10 subunit) that can give rise to large and small subunits upon processing, and an N-terminal prodomain of variable length. The long prodomain caspases contain specific protein-protein interaction domains (CARD, DED) that are involved in the recruitment of caspases in specific multiprotein caspase-activation platforms (DISC, inflammasome, apoptosome, PIDDosome). caspase-12 is expressed as a truncated, catalytically inactive protein in most humans. Modified from (Riedl and Shi, 2004)
According to the “proximity-induced dimerization model”, when several procaspase-8
molecules are bound, for example, to DISC, they are in close proximity to each other and
are therefore assumed to activate and cleave each other by autoproteolysis (reviewed in
Denault and Salvesen, 2002). However, the activation of initiator caspases is still partly
unknown. When active, for example in the case of caspase-8, the heterotetramer consists
of two small and two large subunits, whereas caspase-9 exists predominantly as a
monomer (Degterev et al., 2003). More importantly, the cleavage does not directly
activate initiator caspases instead it brings a specificity-binding pocket into the correct
position, and thereby stabilizing the caspases. Also, other proteins with sequence
similarity, like FLIP and MALT, can dimerize with caspase-8 and form an active site.
However, without a proper proteolytic activity this does not results in caspase-8 cleavage
and apoptosis is inhibited. Furthermore, the FLIP-caspase-8 heterocomplex has been
suggested to mediate non-apoptotic signaling pathway that does not require processing of
caspase-8 (Micheau et al., 2002).
In contrast, effector (executioner) caspases constitutively pre-exist as inactive homodimers
and possess only short pro-domains, therefore lacking the ability to auto-activate (Figure
6). Thus, the activation of effector caspases is carried out by initiator caspases via
cleavage between the large and small subunits (reviewed in Riedl and Shi, 2004). Effector
caspases are responsible for the final step in apoptosis, and many of their target proteins of
them participate in the formation and regulation of the cytoskeleton, function at the cell-
to-cell and cell-to-matrix attachment points or are involved in the regulation of chromatin
structure (Riedl and Shi, 2004). For example, an apoptotic cell undergoes extensive
plasma membrane blebbing, where caspases cut actin regulator proteins resulting in actin
polymerization and, finally, blebbing. Also, “eat met” signals, lipid phosphatidylserine
externalization, of apoptotic cells are also produced by caspase activation (Fischer et al.,
2003). An important caspase-3-mediated step is chromatin fragmentation, where caspase-3
releases nucleases (caspase-activated DNase) by cutting their inhibitor protein, (iCAD)
(Enari et al., 1998). Another final step is chromatin condensation, which is caused by
caspase-3 and -7-mediated cleavage of the acinus protein, and the disruption of the nuclear
envelope resulting from caspase-6-mediated cutting of lamins (Nicholson, 1999). In
addition, caspases have an effect on ATP levels and energy homeostasis since they cut the
mitochondrial electron transport protein, NDUFS1, (Ricci et al., 2004a).
Furthermore, the enzymatic activity of caspases is inhibited by the IAP family of proteins
(XIAP, c-IAP1 and c-IAP2) (Deveraux et al., 1998). In Drosophila DIAP1 is one of the
most important inhibitor of apoptosis and binds DRONC. The cIAPs contain baculovirus
IAP repeat BIR domains 1-3, through which they, for example, compete to bind to active
binding pocket (Vaux and Silke, 2005). In addition, many IAP proteins contain another
domain called RING, that functions as an E3-ligase, responsible for putting ubiquitin
chains on target proteins and targeting them for degradation. During apoptosis, the IAP-
mediated inhibition of caspases is antagonized by another family of proteins (such as
SMAC/Diablo), which contain an IAP-binding tetrapeptide motif (reviewed in Vaux and
Silke, 2005).
2.2.2 Bcl-2 family The Bcl-2 family of proteins were named for the first family member to be described, Bcl-
2, which was discovered as an overexpressed oncogene in B-cell follicular lymphoma
(Tsujimoto et al., 1984). Currently, the Bcl-2 family consists of 12 proteins in mammals,
which regulate the mitochondrial apoptosis pathway, a process called mitochondria outer
membrane permeabilization (MOMP), and cytochrome c release from mitochondria
(Figure 7). The Bcl-2 family is defined by the presence of conserved motifs known as Bcl-
2 homolog (BH) domains (BH1 to BH4), which promotes the interaction and dimerization
within the family (reviewed in Chipuk et al., 2010). Cellular stresses cause transcriptional
and posttranscriptional regulation of the Bcl-2 family resulting in MOMP. According to
the BH domains structure and function, the family of Bcl-2 proteins is divided into pro-
and anti-apoptotic members (Figure 7). Additionally, Bcl-2 family protein may have
alternate functions in other homeostatic pathways including glucose metabolism, cell
cycle and regulation of mitochondrial morphology (Danial, 2007).
The pro-apoptotic Bcl-2 effectors Bak and Bax, (and Bok), contain multiple domains and
are responsible for MOMP, pore forming and the actual cytochrome c release. Once
activated, Bak and Bax bury themselves in the mitochondrial outer membrane, where they
form oligomers of multiple sizes and are suggested create holes (not channels or pores),
which allow large molecules to pass through the membrane (Youle and Strasser, 2008).
However, visualization of these oligomers has shown that large clusters of Bax are
localized near, but not on the OMM, so pore formation is controversial. Bax is shown to
be soluble in the cytosol and during apoptosis it translocates to mitochondria and is
inserted into OMM (Bleicken et al., 2010; Hsu et al., 1997). In contrast, Bak is suggested
to localize at OMM all the time in the cells by its carboxy-terminal region. More precisely,
in viable cells Bak and Bax exist as monomers or dimers and are said to be inactive as
measured by conformation with N-termini being hidden (Letai et al., 2002). Furthermore,
the presence of Bak can facilitate opimal Bax translocation (Mikhailov et al., 2003). Mice
lacking either one, Bak or Bax develop normally and apoptosis is efficient, however,
animals lacking both Bax and Bak are lethal to the embryo. This indicates a molecular
redundancy in which either Bax or Bak can effect the MOMP (Wei et al., 2001).
Figure 7. Classification of Bcl-2 family proteins. BH: Bcl-2 homology domain; TM: transmembrane domain. Modified from (Taylor et al., 2008).
Another class is the proapoptotic BH3-only proteins (Bad, Bid, Bik, Bim, Bmf, Hrk, Noxa,
and Puma), which share only a conserved α-helical BH3 domain through which they
regulate other members. The different BH3-only proteins have varied tissue distributions
and are expressed under different conditions. Furthermore, they are targets of signal
transduction pathways and are therefore considered as sensors for environmental stress.
Bid is a sensor for protease and is unavailable for interaction with the Bcl-2 family
proteins when native. It has a large, flexible loop that can be cleaved by proteases like the
caspases and can subsequently insert into mitochondrial membranes; the BH3 domain
becomes exposed and can interact with other Bcl-2 membranes (Luo et al., 1998). In
contrast, Bim and Bad are sensors for growth factor deprivation (Lomonosova and
Chinnadurai, 2008). For instance, Bad is phosphorylated in response to survival factor,
which results in interaction changes with Bcl-xL and apoptotic resistance. (Zha et al.,
1996). Other BH3 only members have shown to interact with non-mitochondrial proteins,
for instance Bim is binding to the dynein light chain 1, and BMF associates with dynein
light chain 2 (Huang and Strasser, 2000). Additionally, Puma and Noxa are under
trrancriptional regulation by p53 and have important role during DNA damage-induced
apoptosis. The function of BH3-only protein will be discussed more at 2.3.1.2.
Anti-apoptotic (pro-survival) Bcl-2 proteins class includes Bcl-2, Bcl-xL, Bcl-w, Mcl-1
and A1 and all except Mcl-1 possess domains BH1-4 forming a hydrophobic pocket
(Figure 7-8) to which the BH3 amphipathic α-helix of the other members can bind (Tait
and Green, 2010). They can be integrated within the OMM but may also be localized to
the cytosol or ER to inhibit MOMP. Anti-apoptotic members interact or homodimerize
with proapoptotic proteins in order to inhibit their function (Chipuk et al., 2010).
Multidomain Bcl-2 family proteins all have binding pocket present tat is critical for their
functions (Youle and Strasser, 2008). This hydrophobic binding pocket consists of BH1
(portions of α4–α5), BH2 (α7–α8), and BH3 (α2) domains, which all have certain
amphipathic a-helices (Chipuk et al., 2010). Furthermore, anti-apoptotic Bcl-2 proteins are
suggested to inhibit oligomerization of Bak and Bax by binding to their BH3 regions.
When proapoptotic BH3-only proteins activate multidomain Bak/Bax their hydrophobic
face of BH3 helix is exposed. This enables BH4-only protein to interact with the
hydrophobic groove of multidomain dimerization partners. Furthermore, this is thought to
induce series of conformational changes of Bak/Bax: (1) α1/N-terminal exposure, (2)
transient BH3 exposure, (3) protection of the membrane-embedded groove, and (4)
increased proximity of embedded monomers (Chipuk et al., 2010). For example, the BH3
domain of Bak binds to a groove in Bcl-xL formed from a-helices containing BH1-3
domains (Figure 8). Once Bak is activated, it exposes its BH3 domain, which opens a
groove into which the BH3 domain on another Bak molecule can bind, forming a dimer
(Dewson et al., 2009). Bcl-xL, then, may prevent this Bak-Bak interactions and
oligomerization by binding to this exposed BH3 domain. Furthermore, Bcl-2 family
members have binding specificities and, for instance, Mcl-1 and Bcl-xL are major binders
to the Bak BH3 domain, whereas Bcl-2 and Bcl-xL bind well to BH3 of Bax and poorly to
Mcl-1. In addition, Bcl-2-Bax heterodimerization requires also the BH1-3 groove and a
novel interface involving the BH4 motif of Bcl-2 and of Bax containing the BH3 motif
(Ding et al., 2010). Furthermore, the BH3-only member binding profiles varies. Bid and
Bim are interacting with all anti-apoptotic members and also with Bak and Bax, whereas
Bad (to Bcl-2, Bcl-xL, Bcl-w) and Puma (to all) are suggested to bind mainly anti-
apoptotic proteins. In addition, Noxa (to Mcl-1) and Hrk (to Bcl-xL) seem to be very
specific in binding (Chipuk et al., 2010).
Figure 8. BH3:groove interactions in the regulation of Bak activation and oligomerization. According to “direct activation model” (discussed in 2.3.1.2) anti-apoptotic Bcl-2 family members (Bcl-xL) inhibit the activator BH3-only members (Bid) and the multidomain pro-apoptotic Bak/Bax by sequesterating and occupying their BH3 domains. Sensitized BH3-only proteins (Puma) can displace Bid from Bcl-xL and allow Bid to activate (N-terminal and BH3 exposure) and oligomerize Bak.
2.3.1 Mitochondrial apoptosis pathway The mitochondrial (intrinsic) pathway of cell death is engaged by various types of
intracellular stresses including growth factor withdrawal, DNA damage by ultraviolet
(UV) and γ-irradiation stress, cytoskeletal disruption, unfolding stresses in the
endoplasmic reticulum or accumulation of unfolded proteins or hypoxia. Furthermore,
mitochondrial pathway is engaged by death receptor apoptosis pathway and thereby death
signal is amplified as discussed in 2.3.2.2. (Fulda and Debatin, 2006). The key players in
mitochondrial pathway are the Bcl-2 family proteins that respond to apoptotic signals.
During apoptosis, the mitochondria release a number of factors from their intermembrane
space, like cytochrome c, Smac/Diablo, and AIF, which promote and amplify the
apoptotic cascade. More recently, the mitochondrial fragmentation has been shown to be
important early stage during apoptosis and regulated by proteins such as Drp-1 and Fis1
(Karbowski et al., 2006).
Once the mitochondrial apoptosis pathway becomes activated by upstream signals, the
mitochondrial outer membrane permeabilization (MOMP) event occurs. This step is
sudden and rapid, and is considered to be the ‘point of no return’ during apoptosis (Tait
and Green, 2010). MOMP results in the diffusion of soluble molecules (cytochrome c)
residing in the intermembrane space to the cytosol. Of note, MOMP does not involve a
loss of integrity of the inner mitochondrial membrane and therefore, mitochondrial
function is not destroyed completely (Ow et al., 2008). Among the released proteins is
Smac/Diablo that binds same region of XIAP that binds to initiator caspase-9 and thereby
prevents XIAP from inhibiting effector caspases (Deveraux et al., 1997; Verhagen et al.,
2000). In addition, another protein released following MOMP is Omi (HtrA2), which,
like Smac, has an amino-terminal sequence that inhibits XIAP (Verhagen et al., 2002). In
addition, other lethal proteins released by MOMP are endoG, which is a mitochondrial
enzyme that cleave DNA between nucleosomes and AIF, which translocates from the
mitochondria to the nucleus and causes chromatin condensation and DNA fragmentation
(Susin et al., 1999). Importantly, the release of cytochrome c during MOMP results in
caspase activation required for cell death. Moreover, APAF1 is a CARD domain-
containing adaptor protein for initiator caspase-9 (Figure 6) and pre-exists in the cell as a
cytosolic, inactive monomer that cannot bind or dimerize caspase (Bratton and Salvesen,
2010). Whereas, the cytochrome c is a nuclear-encoded protein that is synthesized as apo-
cytochrome c and is transported into the mitochondria to the space between the inner and
outer mitochondrial membranes (the intermembrane space), where it becomes a
holocytochrome c (Ow et al., 2008). Upon cytochrome c´s release, it engages APAF1 and
changes its conformation, leading to exposure of the oligomerization and CARD domain.
Subsequently, the center of the APAF1 oligomer (CARDs) recruits caspase-9 to activate it
(Figure 10); this APAF1-caspase-9 complex is called the “apoptosome” (Reubold and
Eschenburg, 2012). This function of cytochrome c is suggested to be independent of its
function in electron transport, and is required for proper caspase activation during the
mitochondrial apoptosis pathway.
Mitochondrial permeability transition (MPT) occurs in the mitochondrial inner membrane
by a channel called the permeability transition pore (PTP). Exposure to high
concentrations of calcium or signals including reactive oxygen species, changes in cellular
pH and certain drugs can result in MPT. During MPT the voltage potential changes and
solutes enter the matrix and water swells the mitochondria until the inner membrane
ruptures the outer membrane. In addition, PTP is mostly composed of adenosine
nucleotide transporter (ANT) together with voltage-dependent anion channel (VDAC)
(Kroemer et al., 2007). Even originally thought, it is unlikely that MPT is a major
mechanism of MOMP in apoptosis.
Bcl-2 mediated controof MOMP is the major decision that determines whether a cell will
die by engaging the mitochondrial pathway of apoptosis. All of these decisions depend on
interactions among members of the Bcl-2 family of proteins. According to the first model
described, a “rheostatit model”, the balance between anti- and proapoptotic Bcl-2 proteins
defines the apoptotic state of the cell. Later on, two competing models have been
suggested for Bak/Bax activation. According to a “neutralization/indirect model” (Figure
9) BH3-only would primarily bind anti-apoptotic proteins and thereby displace or
neutralize them resulting in Bak/Bax activation and apoptosis (Uren et al., 2007; Willis et
al., 2007). Moreover, one class including Bid, Bim and Puma bind all anti-apoptotic
members and are more potent in inducing apoptosis. Whereas another group, including
Noxa and Bad, engages only selected group of anti-apoptotic proteins (Chen et al., 2005).
For instance, Bak has been shown to be sequestered by Mcl-1 and Bcl-xL until displaced
by BH3 only proteins Noxa and Bad, leading to self-association of Bax/Bak molecules
and MOMP (Willis et al., 2005).
Figure 9. Indirect and Direct activation models for Bak/Bax activation. In the indirect activation model, antiapoptotic proteins bind and inhibit proapoptotic Bax and Bak. Whereas, BH3-only molecules bind antiapoptotic protein to displace them from Bax or Bak, which subsequently can undergo conformational change and oligomerize. In the direct activation model, BH3-only proteins are divided into activator and sensitizer groups. Antiapoptotic Bcl-2 proteins bind and sequester both the activator and sensitizer subclasses. Sensitizer BH3-only proteins occupy the binding pocket of antiapoptotic molecules, allowing activator BH3-only proteins to engage Bax/Bak (Danial, 2007).
However, often neutralization is not enough and additional BH3 activator is required.
Thus, according to a more recent “a direct activator/de-repressor model” (Figure 9) these
proteins are called “direct activators”. For instance, Bid and Bim and their BH3 peptide
can trigger Bax and Bak oligomerization, and permeabilization of synthetic membranes of
isolated mitochondria (Kuwana et al., 2002; Letai et al., 2002; Wei et al., 2000).
Furthermore, this BH3 peptide-induced Bak/Bax activation seems to occur in a stepwise
manner involving a subset of BH3-only proteins (Kim et al., 2009). Puma has also been
shown to promote Bak/Bax activation, however, whether this is via indirect or direct
mechanisms remains controversial (Chipuk et al., 2005). Moreover, anti-apoptotic proteins
can sequester also these direct activators and another BH3-only proteins (“sensitizer or de-
repressor”) can free the direct activator BH3-only protein to activate Bak and Bax (Figure
9). Recently, novel model have been suggested termed “priming-capture-displacement”
model (Strasser et al., 2011). Additionally, other proteins like p53 can exert an ability to
activate Bak and Bax (Kuwana et al., 2005). Importantly, it is suggested that direct
activators do not remain associated with the pro-apoptotic effector protein; instead this
interaction, for example between Bax and Bid, would be physical and transient. According
to the “embedded together model”, the critical step in Bax activation is the initial
membrane insertion and the model emphasizes interaction among Bcl-2 family members
and different conformations (Leber et al., 2007). However, how higher order oligomers
and pores are formed is still controversial. Furthermore, studies using BH3 peptides with
covalently stapled α-helices show that the BH3 domain of Bim binds to the “back” of Bax
and not to the same spot as Bcl-2 (Gavathiotis et al., 2008). Subsequently, Bax undergoes
a conformational change and exposes its BH3 and BH-binding groove and dimerizes.
Tumor suppressor p53 has an important role in regulating the mitochondrial apoptosis
pathway. Particularly, p53 is activated in response to a variety of DNA-damaging agents,
including IR and UV (Kastan et al., 1991). DNA damage results in an accumulation of
p53 protein through post-translational mechanisms, and subsequent increase in p53
activity results in cell cycle arrest or apoptosis (Zilfou and Lowe, 2009). In the nucleus,
p53 can mediate apoptosis by transcriptional activation of pro-apoptotic genes like the
BH3-only proteins for instance (Figure 3) and by transcriptional repression of Bcl-2 and
the IAPs (reviewed in Benchimol, 2001). However, induction of these target gene
products show variable kinetics, with some being delayed. Therefore, evidence for
transcription-independent p53-mediated apoptosis has been accumulating (Mihara et al.,
2003). It has been shown that p53 can directly translocate to the mitochondria after DNA
damage without transactivation domain, and in another analysis this precedes target gene
activation (Erster et al., 2004). In interaction analysis, p53 is suggested to bind Bcl-2 and
Bcl-xL. Furthermore, the p53-Bcl-xL interaction occurs either via the DNA binding domain
of p53 interacting with the α1/BH4 and partial α2/BH3 of Bcl-xL or via the N-terminus of
p53 interacting with Bcl-xL (Petros et al., 2004; Xu et al., 2006). More importantly, p53
interacts with amino acid residues distinct from those that mediate the binding of Bcl-xL to
Bak. In addition, naturally occurring mutations in p53 decrease the p53-Bcl-xL binding,
whereas mutated p53 is still able to bind, for example, Bak but further oligomerization is
abrogated (Mihara et al., 2003; Pietsch et al., 2008). First, p53 was shown to regulate
MOMP by direct activation of Bax, however, later p53 was demonstrated to interact via
the tetramerization domain with Bak and induce oliogmerization of Bak and disruption of
Bak-Mcl-1 complex (Chipuk et al., 2004; Pietsch et al., 2007). Altogether two possible
models are proposed for the regulatory effect of Bcl-xL-p53 interaction. According to Moll
et al., this interaction inhibits Bcl-xL and liberates Bak from Bcl-xL to engage MOMP. In
contrast, Chipuk et al. suggested that the cytoplasmic function of p53 is mediated by Puma
mediated loss of p53-Bcl-xL interaction (Chipuk et al., 2005).
The activity of p53 is mainly regulated by protein stability therefore it is not surprising
that the mitochondrial function of p53 is also shown to be regulated by stabilizing
modifications (Figure 2). Following stress, for example, an E3 ligase Mdm2-mediated
mono-ubiquitination has been suggested to promote mitochondrial translocation of p53
(Marchenko et al., 2007). Subsequently, p53 is immediately de-ubiquitinated by HAUSP
at the mitochondria, enabling interaction with Bcl-2 family members (Marchenko and
Moll, 2007). Moreover, Mdm2 antagonist Nutlin-3a liberates p53 by competitively
binding to the hydrophobic p53-binding pocket within the N-terminus of Mdm2 (Vassilev
et al., 2004). Furthermore, it has been suggested that Nutlin-3a could induce apoptosis via
a transcription-independent manner using mitochondrial p53 (Vaseva et al., 2009).
Controversial data has been published whether phosphorylation would be the determining
factor for mitochondrial translocation of p53 (Nemajerova et al., 2005; Park et al., 2005).
However, certain phosphorylation sites of p53 have been speculated to phosphorylated
and involved in the apoptotic function of p53 (Feng et al., 2006).
2.3.2 Death receptor apoptosis pathway On the surface of mammalian cells are receptors that are activated by death ligands such
as Fas/CD95, TNF-α, and TRAIL (Schmitz et al., 2000). These ligand bound death
receptors mediate apoptosis pathway by transmitting apoptotic signals to the caspases,
subsequently resulting in activation of caspase cascade and death (Ashkenazi and Dixit,
1998).
The major death receptors of the tumor necrosis factor receptor (TNFR) gene superfamily
include TNFR-1, Fas/CD95, DR (Death receptor)-4/TRAIL-R1, DR-5/TRAIL-R2 and less
known non-apoptotic DR3 and DR6 (Ashkenazi and Dixit, 1998; Pan et al., 1997). In
addition, osteoprotegerin is suggested to be one of the TRAIL receptors (Emery et al.,
1998). The death ligands of these receptors are members of the TNF family and play an
important role in the modulation of host defense mechanisms including: inflammation, T-
cell co-stimulation, induction of cell proliferation, macrophage activation, as well as
elimination of unwanted immune cells or tumor cells by apoptosis (Nagata, 1997).
All members of the TNFR family exist as mono- or trimeric receptors located on the
membrane and consist of cysteine-rich extracellular subdomains, which allow them to
recognize their ligands with specificity. Upon binding, trimerized death receptors expose
their conserved death domains (DD), which are located in the intracellular part of the
receptor (Chan et al., 2000). Different death receptors interact via these DDs with different
DD-containing adaptor proteins, like FADD, resulting in a DD-DD interaction and
formation of a death-inducing signaling complex (DISC) (Chinnaiyan et al., 1995). In
addition to its DD, the adaptor proteins also contain a N-terminal death effector domain
(DED), which allows further interaction with procaspase-8. CF95/Fas and TRAIL/Apo2L
ligands results in similar DISC formation and procaspase-8 processing and activation
(Figure 10) (Scaffidi et al., 1998). Signaling through a TNF receptor is more complex and
involves formation of two different complexes (I and II) and caspase-8 activation or
signaling via NF-κB or JNK (Micheau and Tschopp, 2003).
Figure 10. Death receptor signaling and apoptosis. When the CD95/Fas receptor is activated by its ligand CD95, it recruits FADD by DD-DD interaction resulting in DISC formation and DED exposure of FADD. Subsequently, FADD binds the pro-domain of caspase-8 via the DED and enables procaspase-8 dimerization, auto-activation and processing to p44/p41 and p12 subunits. TRAIL forms a DISC consisting of the activated receptor, FADD and caspase-8 or -10. However, the role of caspase-10 in induction of apoptosis is controversial. In TNF-a signaling the initial plasma membrane bound complex (complex I) consists of TNFR1, the adaptor TRADD, the kinase RIP1, and TRAF2 and its formation results in signaling via NF-kappa B or JNK. In a second step, TRADD and RIP1 associate with FADD and caspase-8, forming a cytoplasmic complex (complex II) leading to caspase-8 activation.
In addition, all DISC complex-mediated activations of caspase-8 can be inhibited by
DED-containing FLICE-inhibitory protein long form (FLIP), which consequently
interferes with FADD-caspase-8 interaction, inhibiting both caspase-8 activation within
the DISC and apoptosis (Micheau et al., 2002; Siegmund et al., 2001). Finally, the
activated caspase-8 can proceed to directly activate downstream caspase-3 or engage
mitochondrial apoptosis pathway.
In Type I cells, such as B-lymphocytes, activated caspase-8 can proceed to activate by
proteolytic processing the effectors caspase-3 and -7, which orchestrate apoptosis.
However, the majority of cells belong to Type II cells, like fibroblasts, where the ligand-
initiated apoptosis does not induce caspase activity strong enough for executing apoptosis,
thereby requiring amplification of the signals through the mitochondrial apoptosis
pathway (Rudner et al., 2005). The Bcl-2 family member Bid, which is a caspase-8 and -3
substrate, links the death receptor-induced caspase signaling to the mitochondria (Wang et
al., 1996). Activated caspase-8 cleaves Bid and its truncated form, tBID, translocates from
the cytosol to the mitochondria where it interacts with the pro-apoptotic Bcl-2 family
members Bax and Bak to induce the release of cytochrome c; whereby downstream
apoptotic events can occur (Figure 10) (Luo et al., 1998). Furthermore, the role of the
mitochondrial apoptosis pathway is evident in TRAIL-apoptosis, where inhibition of
TRAIL-induced apoptosis requires ablation of both Bak and Bax, as single knockouts are
still able to release cytochrome c (Kandasamy et al., 2003). However there exists
controversial data about whether Bax or Bak is the major effector in TRAIL-induced
apoptosis. In addition, Bcl-2 overexpression is able to inhibit TRAIL-induced apoptosis,
and interestingly, both Bid and caspase-8 cleavage were sown to be decreased due to Bcl-
2, indicating that caspase-8 activation is amplified by mitochondrial pathway (Fulda et al.,
2002). Also, Mcl-1 cleavage is required for sufficient TRAIL-induced cytochrome c
release (Weng et al., 2005). In addition, TRAIL may also induce activation of other
signaling pathways. For example, there exists evidence that JNK, p38 MAPK, and
IKK/NF-kB kinase pathways are activated downstream of DISC assembly and caspase-8
activation by TRAIL (Sheridan et al., 1997).
Interestingly, death receptor ligands were shown to exert toxicity specifically towards
cancer cells and, for example, recombinant TRAIL suppressed tumor growth without
apparent toxicity (Walczak et al., 1999). However, malignant tumor cells can escape from
treatments thus, a wide spectrum of studies has been focusing on defining the molecules
behind this resistance to treatment. Increasing clinical data however, indicates that TRAIL
is not effective against human cancers. Thus, already some studies have explored whether,
for example, DNA-damaging drugs could enhance the apoptotic effect of TRAIL
(Broaddus et el, 2005). Resistance to TRAIL-apoptosis in the cancer cell remains a
challenging issue for the successful application of TRAIL in cancer therapy.
The c-myc proto-oncogene encodes the c-Myc (herein Myc) transcription factor that was
originally identified as the cellular homologue to the viral oncogene (v-myc) of the avian
myelocytomatosis retrovirus (Vennstrom et al., 1982). c-Myc belongs to the family of Myc
genes (v-Myc, N-Myc, L-Myc, S-Myc and B-Myc) (Dang, 2012). In addition, Myc is a
64.5kDa transcription factor that contains a carboxy-terminal basic helix-loop-helix-zipper
(bHLHZ) domain and a transactivation domain (TAD) in its 150-amino acid amino-
terminus (Dang et al., 1989). In addition, the TAD contains highly conserved elements,
Myc Boxes (MB) I-IV, required for the transactivation of target genes. Myc activates a
diverse group of genes as part of a heterodimer complex with its partner protein, Max
(Blackwood and Eisenman, 1991). Myc-Max heterodimers are capable of binding specific
DNA sequences such as the E-box sequence (CACGTG) and subsequently activate
transcription of target genes (Adhikary and Eilers, 2005). Furthermore, Max association
and DNA binding are required for transcriptional activation of target genes by Myc as
well as its ability to drive proliferation, malignant cell transformation, and apoptosis
(Amati et al., 1993; Blackwell et al., 1990). Another protein family, the Mxd/Mad proteins
(Mad1, Mxi, Mad3, and Mad4) can antagonize Myc function by forming complexes with
Max. Recently, also other Mad-related proteins have been described (Mnt/Rox and Mga)
and some novel Max-related proteins (Mlx and Mondo) that interact with Mad-related
proteins (reviewed in Lutz et al., 2002). In addition, Myc has been suggested to regulate
chromatin by recruiting histone acetylation complexes via MBII-mediated binding to the
TRAP protein and via INI1, which interacts with the chromatin-remodeling complex
(Cheng et al., 1999; McMahon et al., 2000). Furthermore, Myc can act as a transcriptional
repressor with Max by binding directly MBII and the bHLHLZ domain, transcriptional
activators bound to DNA (Kleine-Kohlbrecher et al., 2006). Alternatively, to repress target
genes Myc associates with other proteins like Sp-1 and Miz-1 (Seoane et al., 2001)
Targeted gene disruption of both Myc alleles in embryonic stem cells leads to embryonic
lethality at day 9.5-10.5, which highlights the crucial role of Myc in normal growth
control during mammalian development (Davis et al., 1993). In approximately one-third
of human cancers, the expression of the Myc protein is deregulated, especially in
adenocarcinomas. Overexpression of Myc occurs through diverse mechanisms (discussed
at chapter 1.2.1) including translocation, amplifications, insertional mutagenesis, or
enhanced translation or protein stability (reviewed in Meyer and Penn, 2008). In addition,
point mutations in the coding sequence of the myc gene have been found in translocated
alleles of myc in Burkitt´s lymphoma. However, most often Myc expression is activated
indirectly through alterations in signaling pathways that induce or repress Myc
transcription or stability. Normally Myc transcription is regulated by elongation, and this
block is lost in cancer (Bentley and Groudine, 1986). Furthermore in non-transformed
cells the expression level of Myc is kept low by the extremely low half-life of its mRNA
and protein, approximately 30 minutes (Dani et al., 1984). More recently, it has become
clear that MYC can be deregulated by many additional mechanisms, including activation
by hormones or growth factors, their receptors, second messengers or transcriptional
effectors that enhance MYC expression. Additionally, in tumor cells the mRNA stability
of Myc is increased by direct and indirect mechanisms (Meyer and Penn, 2008). For
instance, increased Myc protein expression is caused by enhanced phosphorylation of
Ser62 by kinases that regulate the stability of Myc together with Thr58 which directs Myc
to proteosomal degradation (Hann, 2006). Furthermore, in cancer the stability of Myc is
enhanced by protein like Ras (Sears et al., 1999).
Myc binds to approximately 10-15% of the genome and is considered a global
transcriptional regulator (Figure 11). The discovery of Myc target genes started from using
fusion of Myc to the hormone-binding domain of the estrogen receptor (ER), MYC-ER,
and later on its tamoxifen regulated form (Eilers et al., 1989; Littlewood et al., 1995).
Importantly, large scale mRNA expression analysis of Myc target genes revealed that the
majority of Myc mRNA changes are relatively low, therefore deciphering true targets
requires more definition from assays such as the chromatin immunoprecipitation assay
(ChIP) (Meyer and Penn, 2008). Furthermore, more evidence suggests that Myc also
regulate target genes via inducing microRNAs (miRNA) from non-coding regions (Chang
et al., 2008).
3.2.1 Proliferation and differentiation by Myc Myc has a crucial role in cell proliferation that contributes to the neoplastic growth of
tumor cells. In quiescent cells in vitro, Myc expression is virtually undetectable. Either
after mitogenic stimulation or via ectopic expression of Myc, c-Myc mRNA and protein
are rapidly induced and subsequently, the cell enters the G1 phase of the cell cycle (Eilers
et al., 1989; Rabbitts et al., 1985). In the cell cycle, Myc is shown to be essential for
GO/G1 to S phase progression and G1 seems to be shortened in the Myc-expressing cells.
Furthermore, Myc allelic deficiency results in defects in cell growth, such as cell mass,
and in the G1 phase (reviewed in Obaya et al., 1999). The cell cycle is regulated by
multiple target genes of Myc. For example, Myc abrogates the transcription of cell cycle
checkpoint genes (GADD45, GADD153) and inhibits the function of cyclin-dependent
kinase (CDK) inhibitors. In addition, Myc promotes cell cycle progression by activation of
cyclin D1, -D2, -E1, -A2, CDK4, Cdc25, E2F1, and E2F2 (Meyer and Penn, 2008).
Recent studies also suggest that Myc might influence the cell cycle by repressing genes,
such as the cell cycle arrest-inducing genes p15 and p16 (Staller et al., 2001). Importantly,
Myc allows cells to enter the S phase and undergo mitosis in the absence of external
growth factor (Eilers et al., 1991).
Myc is broadly expressed during embryogenesis and its expression correlates with growth
in the growing tissues of adults (Pfeifer-Ohlsson et al., 1985). Furthermore, differentiation
is regulated by Myc in both ways; Myc can block the differentiation of a variety of cell
types. In contrast, MYC has been suggested to stimulate cellular differentiation in the case
of stem cells (Gandarillas and Watt, 1997; Langdon et al., 1986). Importantly, the role of
Myc-induced de-differentiation seems to have a major effect on tumor cell phenotype.
Recently, Myc has been shown to increase the capacity of stem cell self-renewal and to
induce pluripotent stem cells (iPS) in terminally differentiated cells (Okita et al., 2007;
Wilson et al., 2004).
Figure 11. Myc-induced molecular and cellular changes.
3.2.2 Transformation mechanisms by Myc The mechanism behind the tumorigenic function of Myc considers proliferation and
differentiation, but also the ability of Myc to increase cell growth (cell mass in size). This
occurs by increasing protein synthesis and reprogramming cell metabolism (Iritani and
Eisenman, 1999). In accordance, enforced expression of the Drosophila c-Myc ortholog in
wing imaginal disc results in increased cell size (Johnston et al., 1999). Importantly,
genomic instability and amplification are increased in Myc-expressing cells, for example
by ROS generation or overcoming the p53 checkpoint (Felsher and Bishop, 1999). More
recently, Myc was suggested to regulate angiogenesis by inducing angiogenic switch via
miR17-92 microRNA cluster or interleukin 1β (Dews et al., 2010; Shchors et al., 2006).
Additionally, Myc can induce immortalization of a cell, however, the mechanisms are
very controversial (Lutz et al., 2002). Specific metabolic pathways are also targeted by
Myc, as discussed in chapter 3.4. Furthermore, Myc is downstream from many signal
transduction pathways like MEK-ERK and PI3K that are responsive to growth factors or
the cellular microenvironment (Lee et al., 2008). Conversely, Myc tumors have been
shown to be resistant to anti-mitogenic TGF-β signaling (Meyer and Penn, 2008).
Several transgenic mouse models have been developed to decipher the mechanism how
deregulated Myc induce tumorigenesis. The mouse data has provided the evidence that
deregulated expression of Myc is sufficient to drive tumorigenesis in a number of
transgenic mouse tissues, but not all (Adams et al., 1985; Leder et al., 1986). For instance,
Myc induced tumor in mammary mouse tissue under Wap- or MMTV-promoter and in
mouse prostate tissue, however, with delayed onset of tumors (Ellwood-Yen et al., 2003;
Schoenenberger et al., 1988; Stewart et al., 1984). Importantly, in each of these cases
additional mutagenic events are necessary for Myc to enable its tumorigenic potential.
Importantly, using inducible systems (tTA Tet-O-Myc, pIns-MycERtm) it has become
evident that withdrawal of Myc ectopic expression is required to maintain tumor, or
otherwise the tumor will regress (Arvanitis and Felsher, 2006; Karlsson et al., 2003).
Importantly, these tumors are considered to be addicted to Myc. Furthermore, increasing
data on mouse models suggest that evading Myc-apoptosis is one of the major
mechanisms behind the oncogenic function of Myc (Dang et al., 2005).
Several laboratories made an intriguing discovery in the early 90´s that oncogenes such as
Myc and the adenovirus E1A, which are both potent inducers of cell proliferation, induce
also apoptosis. The early view of oncogene-induced apoptosis was that it was indirect or
distal response of the cell to an enforced cell cycle entry or to an inappropriate growth
signal generated by Myc. Currently it is evident that Myc-induced apoptosis has a close
link to the Bcl-2-mediated mitochondrial apoptosis pathway. Originally Bcl-2 was found
to cooperate with Myc in tumorigenesis by inhibiting Myc-induced apoptosis (Fanidi et al.,
1992). Similarly, acceleration of lymphomagenesis is found in transgenic mice that
express both Myc and Bcl-2 compared with transgenic mice that harbor just the Myc
transgene (Strasser et al., 1990). Also Myc-null cells were resistant to diverse apoptotic
stimuli. Furthermore, ectopic expression of Myc in the absence of specific survival factors
lead to apoptosis (Askew et al., 1991; Evan et al., 1992). Later Myc was shown to induce
the release of cytochrome c from mitochondria during apoptosis (Juin et al., 1999).
Several studies suggest an important role for Bax in Myc-mediated engagement of the
mitochondrial pathway. For instance, Myc was shown to be required for DNA damage-
induced Bax conformational change and activation and also to Bax oligomerization at
membranes. (Annis et al., 2005; Soucie et al., 2001). Interestingly, in the pIns-MycERtm
pancreas model, the Myc overexpression is able to induce apoptosis of β-cells
simultaneously with the proliferation. However, expression of Bcl-xL or loss-of Bax are
shown to enable Myc-tumorigenesis in these β-cells (Dansen et al., 2006; Pelengaris et al.,
2002). Also, in the MMTV-Myc mammary transgenic mouse model, Myc induces
apoptosis in a Bax-dependent manner. Additionally, Myc has been shown to up-regulate
Bax in growth factor-deprived human cancer cell lines (Mitchell et al., 2000). In non-
epithelial cell tumor models, Myc can co-operate with Bcl-2 and Bcl-xL overexpression in
Εμ-Myc lymphoma and with loss of Bim in Εμ-Myc B-cell leukemia (Egle et al., 2004).
Some studies have shown evidence that Myc suppresses Bcl-2 or Bcl-xL expression both
in MEFs and primary hematopoietic cells (Eischen et al., 2001b; Kelly et al., 2011).
Furthermore, Myc has been proposed to affect the Bcl-2 family network controlling the
mitochondrial pathway, yet the mechanisms by which Myc actually increase the activation
of Bax and Bak remain unknown.
Importantly, tumor suppressor p53 seems to have an essential for Myc-induced apoptosis
(Hermeking and Eick, 1994). For instance, elevated expression of p53 appears to be
associated with apoptosis occurring by Myc (Hermeking and Eick, 1994). Myc-induced
phosphorylation of Serine 15 (murine homologue 18) inhibits Myc tumorigenesis in Εμ-
Myc mice (Sluss et al., 2010). Furthermore, p53 phosphorylated by ATM promotes Myc
apoptosis in squamous epithelial cells (Pusapati et al., 2006). Likewise, Myc co-operates
in tumorigenesis with loss of p53 or ARF in variety of models like in lymphomas, but not,
for example, in mammary carcinomas (Eischen et al., 1999; Elson et al., 1995).
Conversely, existing data in several systems suggest that elevated Myc can induce
apoptosis in the absence of p53, for example, by suppressing Bcl-xL. Thus, Myc-induced
apoptosis is not always dependent on p53 (Hsu et al., 1995; Maclean et al., 2003).
Similarly, Myc can also induce p53 through p14ARF-independent mechanisms in human
fibroblasts (Lindstrom and Wiman, 2003). Importantly, the Myc binding partner Max
seems to be necessary for apoptosis following inappropriate Myc expression, indicating a
transcriptional role in apoptosis (Amati et al., 1993).
Furthermore, increased CDK2 kinase activity is found from Myc-expressing apoptotic
cells and it has been shown to be required for Myc-induced apoptosis in MEFs. Cyclin D3
also sensitizes cells to TNF-induced Myc-dependent apoptosis (Janicke et al., 1996). In
contrast, the mRNA of cyclin A is up-regulated in cells overexpressing Myc but the CDK
activity and apoptosis were shown to be independent events that occur in response to
activation of Myc (Rudolph et al., 1996). Yet the exact role of CDKs in Myc-induced
apoptosis is still unknown. Additionally, the apoptotic properties of Myc are blocked by
cytokines and adhesion signals. For instance, survival signals, either from Ras signaling of
IGF1 receptor, have been shown to inhibit Myc-induced apoptosis e.g. by blocking
cytochrome c release (Kauffmann-Zeh et al., 1997). Whereas, for example, TGF-α
appears to suppress apoptosis by Myc during development of mammary adenocarcinoma
(Harrington et al., 1994). In addition, cell adhesion also influences the susceptibility of
cells to apoptosis by Myc (McGill et al., 1997). Further studies are needed to assess
whether loss of integrin-mediated adhesion is a triggering event or a correlate of apoptosis
by Myc.
Expression of Myc sensitizes cells to a wide range of other pro-apoptotic stimuli, such as
hypoxia, DNA damage, and depleted survival factors as discussed earlier, as well as to
signaling through the Fas/CD95, TNF-α, and TRAIL death receptors (Hueber et al., 1997;
Klefstrom et al., 1994; Ricci et al., 2004b). The sensitization to death receptors signaling
is suggested to occur immediately downstream of these receptor complexes, for example,
by involving p53 and blocking activation of NF-κB or by targeting RIP in TNF-α
signaling (Klefstrom et al., 1997; Klefstrom et al., 2002). Furthermore, one cancer cell line
study suggests that Myc suppressed FLIP and thereby increases caspase-8 processing at
the DISC (Ricci et al., 2004b). Altogether, there seem to be a wide variety of different
ways for Myc to engage apoptosis and this can be explained, for example, by tissue
specificity.
After finding ways to enable transcription and growth, Myc has to organize energy for the
multiplying cell. Glucose and glutamine are the main nutrients that cultured mammalian
cells catabolize for energy production and biosynthesis (Vander Heiden et al., 2009;
Yuneva et al., 2007). Myc directly induces up-regulation of proteins responsible for
glucose uptake and glycolysis (Figure 12). Moreover, Myc has also been implicated in
promoting RNA splicing for the expression of pyruvate kinase M2 that slows down
pyruvate formation and direct glucolytic intermediates to be used in biosynthetic pathways
(David et al., 2010). Moreover, Myc stimulates glutamine uptake and metabolism by
elevating expression levels of glutamine transporters and glutaminase (GLS) via miR23a
and miR23b, (Chang et al., 2008; Gao et al., 2009; Wise et al., 2008). Thereby,
deregulated Myc reprograms metabolism toward using glutamine as the main oxidizable
substrate for maintenance of the tricarboxylic acid cycle and ATP production, thereby
stimulating glutamine consumption and metabolism and also generation of reactive
oxygen species (Gao et al., 2009; Wise et al., 2008). In addition to providing carbon for
bioenergetic reactions, glutamine is also the obligate donor of nitrogen in the biosynthesis
of nucleic acids, and the primary source of nitrogen for the synthesis of non-essential
amino acids (Wise and Thompson, 2010). More importantly, Myc-overexpressing cells
depend on a continual supply of nutrients thereby causing them to be addicted, for
example, to glutamine (Wise et al., 2008). Furthermore, glutamine metabolism appears to
be important for cell survival under especially glucose- or oxygen-deprived conditions (Le
et al, 2012). Another feature of cancer cells is inefficient way of producing ATP, since
nutrients are being incorporated into biomass. In accordance, Myc-mediated metabolic
reprogramming has been associated with lower ATP levels (Vander Heiden et al., 2009;
Yuneva et al., 2007). Concomitantly, Myc is shown to activate genes involved in glucose
and glutamine usage for nucleotide biosynthesis (Mannava et al., 2008). For instance, Myc
induces glycolytic flux from 3-phosphoglycerate for the synthesis of serine and glycine,
which are essential for nucleotide biosynthesis (Vazquez et al., 2011). Glucose and
glutamine are also essential for fatty acid synthesis and Myc is suggested to incorporate
glucose carbons into fatty acids (Morrish et al., 2010). Altogether, these Myc-induced
molecular changes in glycolysis and glutaminolysis are reflecting cell´s adaptation to
abnormal proliferation and requirement of new building blocks.
Figure 12. Metabolic reprogramming by Myc and the metabolically tumor suppressive function of p53 Myc increases glycolysis by upregulating the expression of glucose transporters proteins Glut1 and Glut4, hexokinase (HK2) and pyruvate dehydrogenase kinase 1 (PDK1). Myc increases glutaminolysis by enhancing the expression of glutamine transporters SCT2, SN2 and SLC7A5 and GLS enzyme. Serine and Nucleotide biosynthesis is enhanced by Myc through increased glycolytic flux and upegulation of proteins like PKM2. Mitochondrial biogenesis is affected by Myc via PGC-1β. P53 counterbalances metabolic effects of Myc via increasing OXPHOS, upregulating GLS and inhibiting glycolysis by TIGAR. Modified from (Li and Simon, 2013).
Importantly, Myc regulates mitochondrial biogenesis by increasing oxygen consumption
and mitochondrial mass and function (Li et al., 2005). In accordance, Myc can directly
reactivate genes involved in mitochondrial biogenesis and in mitochondrial protein
synthesis (Ahuja et al., 2010). Additionally, Myc has been suggested to regulate
mitochondrial fusion and fission (Graves et al., 2012). Intriguingly, ARK5, an AMPK
family member, is shown to be essential for Myc-induced metabolic reprogramming and
energy homeostasis, since ARK5 deficiency is lethal with Myc expression (Liu et al.,
2012). All together, recent data emphasizes how widely Myc is affecting energy
homeostasis, however, future studies are needed to fully understand how metabolic
alteration are linked to activation of apoptotic pathways by Myc.
AIMS OF THE STUDY
The specific aims were:
I To elucidate the role of mitochondrial apoptosis pathway engagement in Myc-induced
apoptosis and analyze the potential re-activation of apoptosis in mammary Myc-tumors.
II To distinguish and characterize the crucial mechanisms of Myc-induced Bak activation
and conformational change
III To explore connections of the Myc-mediated mitochondrial apoptosis pathway and
metabolic reprogramming.
MATERIALS AND METHODS
The material and methods used in this study are listed below and are described in detail in
the original publications, which are here referred to using Roman numerals.
1. Materials
Mouse lines Description Source of reference used in
FVB Inbred mouse strain Harlan/Meilahti III, IV
animal facility
Wap-Myc (FVB.Cg-Tg Mouse ine expressing c-Myc under Jax Laboratories III, IV
(Wap-Myc)212Bri/J Whey acidic protein (wap) promoter (stock #002677)
Cells HEK-293-fT Human embryonic kidney fibroblast ATCC I-III
immortalized by hTERT
BT-474 human breast cancer cell line ATCC III
CCD 1102 KERTr keratinocyte ATCC III
HeLa cervical cancer ATCC III
HMEC-hT primary human mammary epithelial ATCC I-III
cell immortalized by hTERT
HepG2 hepatocellular carcinoma ATCC III
MCF10A non-transformed human mammary ATCC I-IV
epithelial cell line
MCF12A human mammary epithelial cell line ATCC III
MCF7 human breast cancer cell line ATCC III
MDA-MB-361 human breast cancer cell line ATCC III
MDA-MB-436 human breast cancer cell line ATCC III
MRC5-hT primary human lung fibroblasts ATCC I, III
immortalized by hTERT
RPE retinal pigmented epithelial cell ATCC III
Sk-Br3 human breast cancer cell line ATCC III
Phoenix Ampho retroviral packaging cell line Dr. Garry Nolan I, II
NMuNG mouse mammary epithelial cell line ATCC IV
ZR-75-30 human breast cancer cell line ATCC III
Expression vectors Description Source of reference used in
pcDNA3.1-caspase-8 encodes caspase-8 provided by Dr.Evan I
pcDNA3.1-caspase-8 C-A encodes catalytic site C->A mutant provided by Dr.Evan I
caspase-8
pC4M-Fv2E- caspase-8 protease domain of casp-8 cloned Prepared in Klefström I
into a FKBP variant construct allowing laboratory
dimerization
pC4M-Fv2E- caspase-8 protease domain of catalytic site Prepared in Klefström I
C->A mutant laboratory
pcDNA3.1-Mycwt encodes Myc provided by Dr.Evan I
Viral vectors Description Source of reference used in
pBabe-puro-MycERtm retoriviral expression vector encoding Myc provided by Dr.Evan I-III
fused with hormone binding domain 4-OHT
pBabe-puroMydcER retoriviral expression vector encoding Myc provided by Dr.Evan I-III
with deletion in aminoacids 106-143
fused with hormone binding domain 4-OHT
pBabe-hygro-Bcl-xL retroviral expression vector encodes provided by Dr.Evan I, II
Bcl-xL
pWZL-hTERT retroviral expression vector endocing provided by Dr.McMahon I-III
catalytic subunit of human telomerase I
pENTR-H1 Gateway entry vector Biocentrum Helsinki I-III
Genome Biology unit
pDSL-hpUGIH Lentiviral shRNA vector with GFP marker Alliance for Cellular I-III
and ires hygromycin selection marker Signalling (AfCs)
pLentiLox3.7 (pLL3.7) Lentiviral shRNA vector with GFP marker Dr. Luk Van Parijs I, II
pLenti6/V5-p53WT Lentiviral expression vector with V5 tag Addgene III
encoding p53 wt
pLenti6/V5-p53R175H Lentiviral expression vector with V5 tag Addgene III
encoding transcription deficient mutant
Addgene III
shRNA vectors antisense sequence (5´-3´´) Source of reference used in
pLL3.7 shcontrol TTAGTAGAGATCTATGAGA prepared by author I, II
pLL3.7 shcaspse-8 CAGGGTCATGCTCTATCAGATTT prepared by author I, II
pLL3.7 shLkb1-1 CCCAAGGCTGTTTGTGTGAAT prepared in Klefström lab II
pLL3.7 shLkb1-2s GCAGAAGATGTATATGGTGAT prepared in Klefström lab II
pDSL-UGIH-shbak CATAGCGTCGGTTGATGTCgt prepared by author I, III
pDSL-UGIH-shbid ACAGAATCCGCACTGTGCT prepared by author I, II
pDSL-UGIH-shbim TGTTGATTTGTCACAACTC prepared by author II
pDSL-UGIH-shbcl-xL UUCACUACCUGUUCAAAGCTT prepared by author III
pDSL-UGIH-shmcl-1 UUCGAUGCAGCUUUCUUGGTT prepared by author III
pDSL-UGIH-sha1 UAUCCAAAUUCACAGUCUGTT prepared by author III
Antibody Description Source of reference used in
α-actin mouse monoclonal Sigma I, II
α6-integrin mouse monoclonal Cymbus Biotechnnologies II
β-actin Santa Cruz III
ACC rabbit polyclonal Cell Signaling Technology III
p-ACCSer79 rabbit polyclonal Cell Signaling Technology III
AMPK rabbit polyclonal Cell Signaling Technology III
p-AMPK Thr172 rabbit polyclonal Cell Signaling Technology III Bak (N-terminal, ab-1) Calbiochem I, III
Bax (N-terminal) rabbit polyclonal Cell Signaling Technology 1, III
Bax (N-terminal, 6A7) mouse monoclonal Trevigen I
Bcl-xL mouse monoclonal BD Transduction Lab. I-III
Bcl-xL rabbit polyclonal Cell Signaling IV
Bim rabbit polyclonal Cell Signaling I, II
Bid rabbit polyclonal Cell Signaling I,
β-catenin mouse monoclonal BD Transduction Lab. II
caspase-3 (Asp175) rabbit polyclonal Cell Signaling Technology I-IV
Caspase-8 IC12 mouse monoclonal Cell Signaling Technology I
CK14 rabbit polyclonal Covance IV
c-Myc (9E10) mouse monoclonal Covance I, III
Cytochrome c mouse monoclonal Pharmingen I, II
COXIV mouse monoclonal Molecular Probes III
Collagen IV rabbit polyclonal Abcam IV
E-cadherin mouse monoclonal BD Transduction Lab. II
EpCAM rabbit polyclonal Abcam IV
GAPDH mouse monoclonal Cell Signaling Technology III
Glutaminase rabbit polyclonal Abcam III
Antibody Description Source of reference used in
GM-130 mouse monoclonal Cell Signaling Technology II
Histone 3 mouse monoclonal Santa Cruz III
Hsp70 mouse monoclonal Santa Cruz III
Ki67 (NCL-Ki67p) rabbit polyclonal Novocostra Lab III
Ki67 (M724901 rat monoclonal DAKO IV
Lamin B goat polyclonal Santa Cruz III
Lkb1 mouse monoclonal Abcam II
Mcl-1 S-19 rabbit polyclonal Santa Cruz III
Mcl-1 600-401 rabbit polyclonal Rockland III
P21 Cell Signaling Technology III
PCNA rabbit polyclonal Santa Cruz III
Phospho S6 Ser223/236 rabbit polyclonal Cell Signaling Technology II
Puma (ab9643) rabbit polyclonal Cell Signaling Technology I
P53 Ab-1 Calbiochem III
P53 Bp53-12 rabbit polyclonal Santa Cruz III
P53 FL-393 mouse monoclonal Santa Cruz III
p-p53 Ser15(18) rabbit polyclonal Cell Signaling Technology III
β-tubulin, rabbit polyclonal Abcam III, IV
glutaminase rabbit polyclonal Abcam III
2. Methods
The following section describes in detail the main methods used by the author.
Administration of ABT-737 to WAP-Myc mice and measure of tumor growth (IV)
WAP-MYC females mice underwent two pregnancies after 8 week of age. ABT-737 was
first diluted in 65% D5W carrier solution (pH 3-4). Animals received either ABT-737
100mg/kg/d solution or equal volume of pure carrier solution administrated by
intraperitoneal injections (i.p) for 21 consecutive days. After treatment period mice were
followed for another 21 days. To measure the tumors, experimental animals were
monitored twice a week in three-day intervals with an electronic caliper (Clas Ohlson).
The formula used for calculating tumor volume is V= d2* D/2, where d= shortest diameter
and D= longest diameter
Cell culture (I‐IV)
Mammary epithelial MCF10A and HMEC-hT cells were cultured in epithelial cell basal
media MCDB 170 (US Biological) supplemented with insulin 5 μg/mL, Bovine Pituitary
Extract (BPE) 70 μg/mL, Hydrocortison 0.5 μL/ mL, Epidermal Growth Factor 5 ng/mL,
Human Transferrin 5 μg/ mL, and Isopropretenol 0.01 μM. MCF12A cells (ATCC) were
cultured in DMEM/F-12 (Gibco) supplemented with 10% (vol/ vol) FCS, epidermal
growth factor 20 ng/mL, hydrocortison 1 μg/mL, insulin 10 μg/mL, human transferrin 5
μg/mL, glutamine, and antibiotics. Mammary breast adenocarcinoma Sk-Br3 cells were
grown in McCoy_s 5a media (Lonca), MDA-MB-361 and MDAMB- 436 in Leibovitz_s
media (Sigma), BT-474, HeLa, HepG2, and MRC-5-hT cells in DMEM, ZR-75–30 cells
in RPMImedium1640 (Lonza), RPE cells in DMEM/F-12 (Gibco), and MCF7 in EMEM
(Lonza) all supplemented with 10% FCS, glutamine, and antibiotics. MDA-MB-436 and
MCF-7 media had 10 μg/mL insulin. CCD 1102 KERTr (Keratinocyte) cells where
cultured with Keratinocyte- SFM Medium (Kit) with L-Glutamine, EGF, and BPE
(Gibco). Phoenix Ampho packaging cells were cultured in high‐glucose containing D‐MEM (Gibco) supplemented with 10% FCS, glutamine and antibiotics (penicillin and
streptomycin). Primary WAP-MYC mouse mammary tumor cells were cultured in .
In three-dimentional organotypic cell culture, the cells were grown in Matrigeltm, which is
a Basement membrane from Engelbreth‐Holm‐Swarm mouse sarcoma (Becton
Dickinson). The mat was prepared according to manufacturers instructions and Matrigel
was overlaid on 8‐chamber slides and left to solidify for 20 minutes at +37°C.
Subsequently MCF10A cells were trypsinized from 2D culture and resuspended in assay
media (MCDB 170 growth media with supplements containing 2% Matrigel) for seeding
(1500 cells/well) on top of the Matrigel. To prepare Collagen I gel 3D cultures, 3 mg/ml
of Collagen I gel (Rat tail Collagen I, Cultrex) and 23 mM NaOH were mixed on ice with
10x phosphate buffered saline (PBS) and subsequently, the mixture was overlaid on 8‐chamber slides and let to solidify in +37°C for 1 hour. Cells (1500 cells/well) were plated
onto Collagen in MCDB media with supplements.
Cloning of shRNA (I, II)
To silence caspase-8 60 bp shRNA oligos were designed and ordered to target caspase-8
mRNA and one scramble (non-targeting control) (Qiagen). The oligonucleotides were
annealed in Annealing Buffer as follows: at +95 °C for 4 min, +70 °C for 10 min and then
temperature was decreased to + 4 °C slowly (6 °C/min). Annealed oligos were clones into
lentiviral vector LentiLox 3.7 (pLL3.7). To silence Bcl-2 family members shRNA oligo
sequences of mRNA designed by Biomedicum Bioinformatics Unit and Biocentrum
Helsinki Systems Biology Initiative and ordered from (Proligo). The annealed shRNA
oligos were cloned first into pENTRY Gateway vector and subsequently using gateway
cloning into lentiviral pDSL_UGIH.
Chemical Cross-Linking (III)
Cells were collected from 15-cm plates into ice-cold PBS and pelleted by centrifugation at
1,500 rpm (Heraeus Megafuge 1.0 R) for 10 min. The cell pellet was resuspended in
SCEB buffer (200 mM sucrose, 10 mM Hepes pH7.4, 50 mM KCl, 5 mM EGTA, 5 mM
MgCl2,) containing DTT, PMSF, and cytochalasin B and incubated for 30 min before
homogenization with Dounce homogenizer. The unbroken cells and debris were removed
by centrifugation at 2,000 rpm (Heraeus Megafuge 1.0 R) for 10 min at +4 °C.
Subsequently, the supernatant was subjected to a cross-linking assay. BMH (1,6-
bismaleimidohexane) was added to a final concentration of 1 mM, and lysates were
incubated for 30 min at room temperature on constant mixing. After quenching the
reaction with DTT, the mitochondria-enriched heavy membrane fraction was pelleted at
13,000 rpm (Heraeus Pico17) for 10 min at +4 °C and lysed in radioimmunoprecipitation
assay buffer (RIPA). From each sample, 20 μg of mitochondrial proteins was subjected to
SDS/PAGE and Western blot analysis.
Co-immunoprecipitation (III)
Cells growing on 15-cm dishes were lysed in 300 μL of IP-lysis buffer (20 mM Tris·HCL
pH 7.4, 135 mM NaCl, 1.5 mM MgCl2, 1 mM EGTA, 10% (vol/vol) Glycerol, 1%
(wt/vol) CHAPS with protein inhibitor mixture (Roche). Lysates were kept on ice for 10
min, and unbroken cells and debris were centrifuged at 13,000 rpm (HeraeusMegafuge 1.0
R) for 15 min at +4 °C. Immunoprecipitations were carried out using the Catch and
Release v2.0 Reversible Immunoprecipitation System kit (Millipore). Columns were
washed according to the protocol, and the reaction mix for immunoprecipitation contained
2 μg of IP-antibody, 2,000 μg of protein, and antibody capture affinity ligand. Samples
were incubated on rotation at +4 °C for 2 h and washed five times with 400 μL of Catch
and Release Washing Buffer. Antibody captured proteins were eluted with 45 μL of Catch
and Release Denaturing Buffer containing 200 μL of β-mercaptoethanol and by
centrifuging at 5,000 rpm (Heraeus Pico17) for 20 s.
Caspase-8 and caspase-3 activity assay (I, III, IV)
To measure caspase activities (I), the tells were lysed in Triton-X cell lysis buffer (10mM
Tris, pH 7.5 130mM NaCl, 1% Triton X-100, 10mM NaPi and 10mM NaPPi) and the
amount of caspase-8 activity was estimated by measuring proteolytic activity towards
synthetic caspase-8 substrate Ac-IETDpNA or caspase-3, Ac-DEVD-pNa peptide
(Calbiochem). To measure caspase-8 or caspase-3 activity, 40 mg aliquot of each lysate
was incubated in a 100 ml total volume of 1_Salvesen optimal caspase buffer (Stennicke
and Salvesen, 1997) supplemented with 200 mM colorimetric pNa-peptide. Samples were
incubated at 37 °C and read at 405nm with an automated plate-reading spectrophotometer
at 1-h intervals. In addition 96-well commercial kit was used (III, IV) and ells were seeded
on 96-well plates and treated, and apoptosis was measured using the Caspase-Glo 3/7
luminescence kit. Luminescence (RLU) was measured using a VICTORX3 (PerkinElmer)
plate reader.
Generation of Fv-caspase-8 constructs (I)
The protease domains of caspase-8 and caspase-8 C-A mutant (catalytic site cysteine
mutated into an alanine) were PCR amplified from pcDNA3.1-Caspase 8 and pcDNA3.1-
Caspase 8 C-A vectors. Resulting fragments were digested and subcloned into pC4M-
Fv2E (Ariad Pharmaceuticals). The two Fv (FKBP variant) domains in the chimeric Fv2E-
caspase-8 construct substitute for the natural DED dimerization domains of caspase-8 and
allow ectopic dimerization of the construct with chemical dimerizer AP20187. The
sequences of pC4M-Fv2E-caspase-8 and pC4M-Fv2E-caspase-8 C-A constructs were
verified by sequencing.
Immunofluorescence of two-dimensional cultures (I‐III) All cells were fixed with 4% paraformaldehyde (PFA) for 10 minutes at room temperature
(RT) before staining. To detect localization of cyt c, fixed cells were washed for 6 times
with PBS to permeabilize and block with PBS containing 3% BSA and 0.05% Saponin.
Next, the cells were incubated for 1h at RT with monoclonal anti-cyt c antibodies (BD
Pharmingen) diluted 1:300 in permeabilization/blocking buffer. After incubation with
primary antibody, cells were washed for 6 times and incubated for 1h with appropriate
Alexa fluor dye (Alexa 488 or 546, Molecular Probes) coupled secondary antibodies. Next,
cells were washed with PBS and counterstained with 1 μg/ml Hoechst 33258 (Sigma) to
visualize the nuclei. For immunostainings with conformation-specific antibodies (Bak and
Bax), cells were were blocked and permeabilized with 5% goat serum+0,06% Saponin and,
the primary and secondary antibodies were diluted 1:300 in 5% goat serum+0,04 %
Saponin. All other staining were permeabilized with 0.1% (vol/vol) Triton X-100 BSA-
PBS for 30 min and primary and secondary antibodies were incubated in 1% (wt/vol)
BSA-PBS for 1 h at room temperature and washed with PBS. Hoechst 33258 (Sigma) was
used to detect the nuclei. Slides were mounted using Immu Mount (Thermo Scientific)
mounting reagent. Images were acquired using Zeiss Axioplan 2 microscope equipped
with mercury lamp and Zeiss AxioCam HRc color camera and Axiovision 4.6 software
using objectives Plan Neofluar 20x (NA=0.5) and 40x (NA=0.75), 64x. (NA=1.25).
Immunofluorescence of three-dimensional cultures (II)
The organotypic cultures were fixed with 2% PFA for 20 minutes at RT and thereafter
washed with PBS. Epithelial structures were permeabilized with 0.25% Triton X‐100 in
PBS for 10 minutes at +4°C, and thereafter washed with PBS. The non‐specific binding
sites were blocked in immunofluoresence (IF) buffer (7.7 mM NaN3, 0.1% BSA, 0.2%
Triton X‐100 (Sigma) and 0.05% Tween20 in PBS) supplemented with 10% normal goat
serum (Gibco) for 1‐2 hours. Subsequently, slides were incubated with primary antibody
in the blocking solution overnight at +4° following three 15 minutes washes with IF buffer.
Then slides incubated with Alexa Fluor secondary antibody diluted in blocking solution
for 40‐50 minutes at RT following washes with IF buffer. The nuclei were counterstained
with Hoechst33258. Slides containing immunostained epithelial structures were mounted
with Immu‐Mount mounting reagent. Images of the structures were acquired using Zeiss
Axiovert 200 microscope equipped with Zeiss Apotome and AxioCam MRm camera
(study IV) and/or Zeiss LSM Meta 510 confocal microscope (studies I‐IV) equipped with
argon (488), helium‐neon (543 and 633) and diode (405) lasers. The objectives used were
Plan‐Neofluar 40x DIC objective (NA=1.3, oil) and Plan‐Neofluar 63x DIC (NA=1.25,
oil).
Immunohistochemistry (III, IV)
Mammary gland tissue and tumors were collected and fixed in 4% (vol/vol)
paraformaldehyde (PFA) overnight and processed into paraffin blocks. Sections (3 μm)
were cut from paraffin blocks on glass slides (Superfrost Plus), deparaffinised and
incubated overnight at +70 °C in antigen retrieval citrate buffer solution (Dako).
Subsequently, sections were incubated in 0.3% (vol/vol) H2O2 solution for 15 min and
washed for 5 min with PBS. Sections were blocked in buffer containing 10% (vol/vol)
normal goat serum (Gibco) in mammary gland histological buffer [0.1% (wt/vol) BSA,
0.2% (vol/vol) Triton-X (Sigma), 0.05% (vol/vol) Tween 20] for 30 min followed by 1 h
incubation with primary antibody and secondary antibody using standard avidin–biotin
complex detection with 3,3-diaminobenzidine (Vector Laboratories), and counter- staining
with hematoxyline (Thermo Scientific). Sections were dehydrated and mounted with
Vectastain permanent mounting medium (Vector Laboratories). TUNEL staining was
performed with ApopTag in situ detection kit (Chemicon/Millipore) according to
manufacturers instructions. For all paraffin embedded tissues, hematoxylin and eosin
(H&E) staining were performed in Biomedicum Genomics Histolab. The
immunohistochemistry (IHC) samples were analyzed and photographed with a Leica DM
LB microscope and software Studio-Lite 1.0.
Primary cell isolations and fat pad transplantations (IV)
To isolate primary WAP-Myc mouse mammary epithelial cells were isolated from 8‐14
weeks old virgin female mice. To isolate tumor cells, mice were euthanized with CO2 and
cervical dislocation and thoracic and tumors were removed and chopped.
(Heraeus Multifuge 1.0)
0.05% Trypsin‐EDTA for 5‐10 minutes
Fat pad transplantations were performed essentially as described previously (Deome et al.,
1959; Welm et al., 2008). 3‐week‐old female FVB mice were anesthetized using 2‐2.5%
Isoflurane (Baxter) and the anterior part of #4 mammary gland containing rudimentary
ductal epithelium, lymph node and bridge to #5 gland was surgically removed. Wap-Myc
tumor 105 cells/gland in 10 μl volume were injected to the remaining fat pad. The wound
was sutured using wound clips (Autoclip Physicians kit, Clay Adams), which were
removed 1 week after the operation. The mice received analgesic pre‐ and postoperatively
(Temgesic 0.2 mg/kg and Rimadyl 5 mg/kg, respectively) and there covery of the animals
was followed 48 hours after surgery. After 1 week, the fat pad transplanted mice were
administered with ABT-737.
Production of retroviruses and retroviral transductions (I-III)
To produce amphotropic retrovirus particles, a retroviral construct was introduced into the
Phoenix Ampho cell line by transfecting 8 μg of DNA using JetPei transfection reagent
(Polyplus) according to manufacturers instructions. The supernatants containing
retroviruses were collected 48 hours post‐transfection and used for transduction. Prior to
retroviral infection, cells were plated at low density on dish and following day incubated
with virus supernatant. During transduction procedure the cells were centrifuged for 30
minutes 2500 rpm. Transduced cells were selected in puromycin (2.5 μg/ml), blasticidine
(10 μg/ml) or hygromycin and surviving 24 hours post-transduction.
Production of lentiviruses and lentiviral transductions (I‐IV)
The VSVg pseudotyped lentivirusparticles were produced by transfecting 293fT cells with
a three plasmid mixture consisting of the lentiviral expression or shRNA construct,
packaging construct CMVDelta8.9 and the VSVg encoding envelope construct phCMVg.
For traditional high‐titer lentivirus production, the supernatants containing lentiviruses
were collected 72 hours post‐transfection and used for transduction. Viral titer was
determined using p24 ELISA test measuring viral capsid protein p24 (Perkin Elmer,
performed in Biomedicum Genomics). Lentiviral infections of 2D cultured cells were
performed similarly as retroviral infections. Transduced MCF10A cells were selected in
blasticidine (10 μg/ml), puromycin (2.5 μg/ml) or hygromycin (250 mg/ml) 72 hours post‐transduction.
RNA isolation and quantitative real‐time PCR (qPCR) (III)
RNA was isolated by RNeasy kit (Qiagen) from cells transfected with siRNA and grown
on six-well plates for 72 h after treatment, and mRNA levels were analyzed by using the
Light Cycler 480 II instrument (Roche) using Universal Probe Library (no. 43 tggggcag)
probes (Roche Applied Science) and LightCycler 480 Probes Master mix and PCR grade
H2O (Roche Applied Science). To analyze the gene silencing activity of siRNA constructs,
the expression level of the gene of interest was compared with that of a housekeeping gene
(β-actin) and normalized to control siRNA-transduced control. Control siRNA samples
were tested in technical duplicates and the gene of interest siRNA samples in technical
triplicates. The analysis was performed by the Biomedicum Functional Genomics Unit.
SiRNA Transfections (III)
The 3′ Alexa Fluor546 siRNAs were ordered from Qiagen and transfected into the cells by
using HiPerFect transfection reagent (Qiagen). Cells were plated on 24-well plates as a
semiconfluent culture and next day transfected with siRNA. Validated siRNA sequences
are listed below.
Site-Directed Mutagenesis (III)
Expression vectors for mutagenesis were pLenti6/V5-p53_WT p53 and pLenti6/V5-p53
R175H (Bernard Futscher laboratory plasmids, Addgene: plasmids 22945 and 22936, ref.
3). Mutagenesis was performed with the QuikChange Site- Directed Mutagenesis kit
(Agilent Technologies) according to the manufacturer’s instructions. QS mutation (L22Q
and W23S) was introduced with the following primers: sense 5′-
agtcaggaaacattttcagaccaatcgaaactacttcctgaaaacaac-3′ and antisense 5′-
gttgttttcaggaagtagtttcgattggtctgaaaatgtttcctgact-3′. The presence of the mutations in the
vector was confirmed by sequencing.
Reagents (I-IV)
The following reagents were used in the experiments: 4-Hydroxytamoxifen, ABT-737 (a
kind gift from Dr Steven W Elmore, Abbott Laboratories), AP20187 (Ariad),
aminoimidazole carboxamide ribonucleotide (AICAR) (III, WB: Cell Signaling, IF:
Sigma), Etoposide, Doxorubicin, checkpoint kinase 2 (Chk2) inhibitor II (Sigma),
mitogen-activated protein kinase (MEK) inhibitor U0126 (Promega), Nutlin-3a (Alexis
Biochemicals), phosphatidylinositol- 4,5-bisphosphate 3-kinase (PI3K) inhibitor
Ly294002 (Cell Sig- naling), mitogen-activated protein kinase 1 (ERK1) inhibitor II,
ataxia telangiectasia mutated (ATM) inhibitor CGK-733 (Santa Cruz), AMPK inhibitor
Compound C, mitogen-activated protein kinase (p38) inhibitor SB203580, protein kinase,
DNA-activated (DNAk-PK) inhibitor C401, AMPK activator A769662 (Bio- Vision), and
recombinant human TNF-related apoptosis inducing ligand (TRAIL) (R&D Systems), z-
VAD-fmk (Enzyme Systems Products).
Subcellular fractionations (III)
Cells were collected into ice cold PBS and pelleted and subsequently mitochondria
enriched fraction and light membrane/cytosol fractions were collected into ice cold PBS
and isolated according to the manufactures instruction using Dounce homogenizer and
Mitochondria/Cytosol Fractionation Kit (Biovision).
Western blot analysis (I‐IV)
Cells grown on 2D cell culture were lysed either in SDS lysis buffer (II) (20% SDS in 250
mM Tris‐HCl pH 6.8 – study I) or non‐ionic Triton‐X cell lysis buffer (I) (10 mM Tris,
pH 7.5, 130 mM NaCl, 1% Triton X‐100, 10mM NaPi and 10 mM Nappi) or RIPA lysis
buffer (2 5mM Tris-HCl, pH 7.6, 150mM NaCl, 1% NP-40, 1% sodium deoxycholate,
0.1% SDS) all supplemented with EDTA‐free protease inhibitor cocktail (Roche). Mouse
mammary tumors were lysed in mammary gland (MG)‐lysis buffer supplemented with
EDTA‐free protease inhibitor cocktail (Roche). An approximately 2 mm x 2 mm piece of
snap frozen tissue was placed in tube containing MG‐lysis buffer and ceramic beads and
homogenized using Precellys homogenizer (Bertin Technologies). Subsequently, samples
were let stand on ice for 30 minutes and centrifuged at 13 000 g for 15 minutes. Protein
concentrations and western blotting was performed as described above. 10‐20 μg samples
were denatured with 5x Laemmli sample buffer and separated in 10% SDS‐PAGE gel,
and subsequently transferred to nitrocellulose filters (Bio‐Rad) followed by antigen
detection with specific antibodies and ECL™ Detection Reagents (Amersham).
RESULTS AND DISCUSSION
4. The mitochondrial apoptosis pathway is engaged by Myc in a Bak-dependent manner independently of cell context and death stimuli
Our aim was to elucidate what the molecular mechanisms are behind the Myc-induced
sensitization in epithelial cells, especially in mammary tissue. Normal or primary cells are
often resistant to drugs like TRAIL, which offers the possibility to use such molecules in
cancer therapy (LeBlanc et al., 2002). However, it is evident that not all tumors respond
well or they relapse soon after treatment. Therefore, it is important to understand which
molecular mechanism in cancer cells enables sensitivity and resistance to drugs.
Previous results show that Myc activation for instance in growth factor deprived cells or
drug/DNA damage stressed cells results in apoptosis in a mitochondrial-dependent manner
(Klefstrom et al., 2002). Likewise, Myc seems to sensitize cells to variety of different
stresses, however the exact mechanism behind mitochondrial pathway engagement
remains diverse. In our study, we analyzed how mammary epithelial cells respond to
ectopic expression of Myc by using MYC-ERtm cells (Eilers et al., 1989; Littlewood et al.,
1995). We show that activation of Myc does not induce apoptosis in normal, two-
dimensional cell culture as shown previously in other non-transformed cell types however,
Myc can prime mitochondria for various drugs (TRAIL and Etoposide) by engaging the
mitochondrial apoptosis pathway (I-III). Furthermore, Juin et al, showed that Myc
activation results in cytochrome c release in Rat-1 cells (Juin et al., 1999). In accordance,
we showed that combined Myc and TRAIL signaling results in cytochrome c release into
the cytosol, however, Myc alone is unable to induce MOMP (I-III). Previous results
suggest that Bax is the major mediator of Myc-mediated apoptosis in primary mouse cells,
pancreatic β-cells, and in Εμ−Myc lymphoma (Annis et al., 2005; Dansen et al., 2006;
Eischen et al., 2001a). Concordantly, our results show that Bax undergoes a
conformational change and its N-terminus is exposed in cells undergoing apoptosis by
Myc and TRAIL. In addition, we show also that Myc and TRAIL activation was able to
induce also Bak activation and N-terminus exposure in apoptotic cells (I and III).
Importantly, this was the first time when Bak was suggested to mediate Myc functions.
Furthermore, we were able to show using RNAi-techniques that Bak is required for Myc-
mediated engagement of mitochondrial pathway in mammary epithelial cells (I and III). In
contrast, similar analysis of Bax-deficient cells showed that Bax is not necessarily
required for Myc apoptosis in these cells. Surprisingly, an evident Bak up-regulation was
noticed in Bax-deficient cells indicating compensational mechanisms could be involved
(our unpublished results) (Mondal et al., 2012). Also, in variety of models tested
abrogation of apoptosis often requires Bax, Bak double deficiency (Wei et al., 2001).
Furthermore, we showed that Myc was not able to induce transcriptional changes in
protein expression levels on Bad, Bax, Bcl-xL, Bid, Bim or Puma as suggested earlier (I
and our unpublished results) (Hemann et al., 2005; Maclean et al., 2003; Mitchell et al.,
2000). In contrast, a transient Bak upregulation was seen at 10-hour time point in Myc-
activated cells (I). However, whether this is a direct transcriptional target by Myc remains
to be elucidated. These results suggest that Myc-induced mitochondrial priming occurs via
the Bcl-2 family members and in a Bak-dependent manner in various cell types: fibroblast,
mammary epithelial cells and variety of other epithelial cells, cancer cell lines, and
primary mouse mammary cells (I-IV).
In addition, to explore how different physiological environments and epithelial integrity
can affect the capability of Myc to induce proliferation and apoptosis we used a three-
dimensional organotypic cell culture model of mammary epithelial cells (Debnath et al.,
2002). In this system, epithelial cells start to proliferate, undergo apoptosis to form a
hollow lumen and possibly migrate to form a rounded, organized, polarized and quiescent
structure called acini. Our results show that Myc activation can induce apoptosis if
activated during early acinar morphogenesis when the cells are unorganized and
proliferating (II). In contrast, fully-formed acini were resistant to Myc activation and
apoptosis and also to death induction by TRAIL alone suggesting that either polarity or
quiescence could prevent Myc functions (II). In accordance, Zhan et al. showed that Myc
functions are suppressed by cell polarity and deregulation of Scribble can promote Myc-
driven mammary tumorigenesis (Zhan et al., 2008). Importantly, activation of Myc in
fully formed acini was able to sensitize quiescent cells to TRAIL indicating that Myc is
still active and can augment apoptosis. More importantly, the proliferative and apoptotic
capacity of Myc seemed to be different in acini at different morphogenic stages. For
instance, if the cells were non-polarized and unorganized at the time of Myc activation,
like in early acinar morphogenesis, activation of Myc resulted in bigger acini. Whereas, in
fully-formed acini Myc was unable to re-initiate the cell cycle (II). These results indicate
that organized epithelial architecture or proliferation driven by acinar morphogenesis is
critical for tumorigenic functions of Myc, however apoptotic sensitivity remains
independently of proliferative status. In this study we also showed that loss of LKB1
inhibited polarization and organization of acini, as measured by GM130, β-catenin and
α6-integrin (II). However, these LKB1-lacking acini did finally undergo quiescence, and
interestingly, Myc was then able to induce cell cycle re-entry. Interestingly, our results
indicate that Lkb1 inhibits tumorigenic functions of Myc by maintaining the polarity of
epithelial cells in acini. Furthermore, in addition to its polarity function, the Lkb1 protein
has been shown to regulate proliferation by inducing growth arrest, and also activation of
metabolic pathways via AMPK (Tiainen et al., 1999). Thus the results indicate that
organized epithelial architecture is one of the critical suppressors of proliferative Myc
function. Furthermore, acinar morphogenic program and differences in the proliferation
status of acini may impact on the cell, for instance, in metabolism. Hence, in this acini
model, the exact role of active cell cycle machinery and differential activation of
metabolic pathways on Myc function remains to be elucidated. Moreover, our result
indicated that Myc also engaged mitochondrial apoptosis pathway in the three-
dimensional acini model since Bim- and Bid-deficient cells were resistant to Myc and
TRAIL synergistic death. Interestingly, Bak seem to be required for Myc-apoptosis in
early acinar structures (II and our personal data).
Finally, the role of mitochondrial apoptosis pathway in Myc-apoptosis was emphasized
since it was inhibited by anti-apoptotic Bcl-xL. Likewise, previous results suggest that Bcl-
xL and Bcl-2 are key regulators of Myc tumorigenesis and apoptosis (Eischen et al.,
2001b). Furthermore, the BH3 mimetic ABT-737, which inhibits anti-apoptotic Bcl-2
family members, Bcl-2, Bcl-xL and Bcl-w, phenocopied the effect of Myc activation and
sensitized cells to apoptosis (I and II). Intriguingly, since the effect of ABT-737
significantly resembled apoptotic activation by Myc, it could be hypothesized that Myc
influences the balance of anti-apoptotic Bcl-2 family members. Altogether, these results
indicate that Myc-mediated priming of apoptosis is mitochondria-dependent.
5. The mitochondrial amplification loop as a mechanism for lethal
caspase activation in death receptor signaling
Previous studies have implied that during TRAIL death receptor signaling, procaspase-8 is
processed and activated at the DISC after ligand binding and in type II cells this apoptotic
signal is directed to the mitochondrial apoptosis pathway by Bid protein (Broaddus et al.,
2005; Kischkel et al., 2000). To understand the apoptotic co-operation of TRAIL and Myc
signaling, initiator caspase-8 and Bid were silenced using RNAi. Indeed, our results
indicate that both caspase-8 and Bid are required for TRAIL signaling (I-II). Furthermore,
our results demonstrate that TRAIL induces caspase-8 activation in these TRAIL-resistant
mammary epithelial cells and fibroblasts, however evident Bid cleavage was seen only in
cells where Myc was also active (I).
In apoptotic cells by Myc and TRAIL signaling, an interesting augmentation of caspase-8
cleavage was observed when the mitochondrial apoptosis pathway was activated. This
enhanced caspase-8 processing and activity was inhibited in cells if mitochondrial
pathway was blocked (I). Previously such post-mitochondrial Bid and caspase-8
processing has been suggested to occur downstream of the mitochondria, possibly by
caspase-3 (Slee et al., 2000).
FLIP is a caspase-8 inhibitor recruited to the death-inducing signaling complex (DISC)
instead of caspase-8. It has been suggested that Myc would directly increase FLIP
expression levels and thereby increase sensitivity of cells to TRAIL (Ricci et al., 2004b).
However, since Myc sensitizes cells to a variety of death stimuli UV, genotoxic drugs and,
for example, to growth factor deprivation, it is highly possibly that FLIP alone does not
explain apoptotic sensitivity (Askew et al., 1991; Juin et al., 1999; Prendergast, 1999). In
addition, we were not able to see any significant mRNA changes of FLIP in gene
expression profiling of Myc-activated cells (our personal results). Therefore, to answer
this question we analyzed Myc-induced caspase processing by using dimerizable caspase-
8 constructs that are independent of DISC formation. We showed that combined
expression of Myc and dimerized caspase-8 was able to induce cell death (I). These results
indicate that mechanisms other than enhanced DISC-recruitment explain synergistic cell
death between Myc and TRAIL, and results in a mitochondrial amplification of caspase
signaling. Furthermore, mitochondrial pathway-specific ABT-737 was also able to
augment caspase-8 processing and activation when combined with TRAIL (I), also
suggesting a mechanism downstream of DISC for Myc.
6. Myc-induced Bak N-terminal exposure is a conformational change
preceding Bax activation and MOMP
Pro-apoptotic Bak normally exists embedded in the mitochondrial outer membrane in a
conformation where its N-terminus and BH domain are buried (Dewson et al., 2008).
Instead, Bax is more loosely attached to mitochondrial membranes and can be translocated
to the mitochondria upon death stimuli in some cells (Schellenberg et al., 2013). When the
mitochondrial apoptosis pathway is activated, both of these proteins are thought to
undergo a conformational change resulting in exposure of the N-terminus and transient
exposure of the BH3 domain, finally enabling a homo-oligomerization of these proteins
(Annis et al., 2005; Dewson et al., 2009). How oligomerization really occurs, and whether
Bak and Bax molecules are interacting with each other and how the exact pore is formed
is still under depate. In our experiments, when Myc was activated in mammary epithelial
cells, N-terminal exposure of Bak was detected in immunofluorescence staining using
non-detergent conditions (I and III). This was clearly different from Bax activation that
occurred when cells were in addition treated with TRAIL and underwent apoptosis.
Importantly, this “preactivation” of Bak did not coincide with Bak homo-oligomerization
and proper oligomers were visible only during apoptosis. Likewise, Moldoveanu et al.
suggest that in Bak activation process and conformational changes, it is Bid that launches
the dimerization and MOMP-inducing oligomerization of Bak (Moldoveanu et al., 2013)
Furthermore, in our analysis Bak and Bax were localized to mitochondria both in control
and Myc-expressing mammary epithelial cells, however Bax was also found from cytosol
(III and our personal results). This indicated that Myc does not enhance Bax translocation,
on the other hand it still might affect on Bax activity as suggested earlier (Soucie et al.,
2001)
According to the neutralization model, sequestration of Bak binding inhibitors would be
enough to enable Bak conformational change (Tait and Green, 2010). Hence, to further
understand how Bak activation is regulated, we analyzed the role of Bak interacting anti-
apoptotic proteins Bcl-xL, Mcl-1, and A1 using RNAi (Willis et al., 2005). Surprisingly,
only Bcl-xL deficiency resulted in Bak conformational change and activation. In addition,
sequestration of Bcl-xL, Bcl-2, and Bcl-w with ABT-737 resulted in Bak activation (III).
This indicated that Bak acitivity preceeding MOMP is regulated by BclxL-interactions.
However, our interaction analysis showed no differences between Bak-Bcl-xL and Bak-
Mcl-1L in Myc-expressing cells, whereas inhibition of Bcl-xL by ABT-737 resulted in loss
of interaction and both are able to activate Bak (III). Importantly, Bcl-xL, overexpression
in mammary cells was still able to abrogate Myc-induced Bak activation and apoptosis,
suggesting that Bcl-xL might have still have inhibiting role in Myc engagement of
mitochondria (I and III). Indeed, in Myc and TRAIL-treated apoptotic cells this Bcl-xL
association to Bak was diminished almost completely, indicating that Bcl-xL regulates
conformational changes and oligomerization of Bak (our personal results). According to
our personal results, the role of Mcl-1 seems to be evident in TRAIL-signaling pathway
but not important regulator of Myc functions, since we could detect Mcl-1 cleavage and
loss of Bak-Mcl-1 interaction in apoptotic cells by Myc and TRAIL as suggested
previously (Weng et al., 2005). In accordance, our results suggest that Mcl-1
overexpression is able to slightly inhibit cell death by Myc and TRAIL, but not Bak
activation induced by Myc (our personal results). These results support the “neutralization
model” for Bak activity regulation, where Bcl-xL-free Bak undergoes conformational
change and exposes its N-terminus. Conversely, Myc did not induce such Bak liberation
from Bcl-xL suggesting a different mechanism. Hence, according to another ”direct
activation” model Bak activation and oligomerization always requires a another direct
activator BH3-only protein like TRAIL-tBid (Tait and Green, 2010). Likewise, our results
indicate that Bak oligomerization and MOMP require additional activator. Furthermore,
another transient conformational change, BH3 domain exposure, is suggested to follow N-
terminal exposure and occurs before final oligomerization of Bak and can also be involved
in Myc priming (Dewson et al., 2009). Moreover, mitochondrial fission during apoptosis
has major role in regulating MOMP (Suen et al., 2008). Thus, it is possible that Myc-
induced changes at mitochondrial structure and fission can affect interactions between the
Bcl-2 family members and, more importantly, Bak activation. Altogether, these results
suggest a hypothetical model where these two models are combined, and highlights that
N-terminal exposure of Bak is a separate step from final activation, which also includes
BH3 domain exposure and homo-oligomerization.
7. BH3 mimetic reactivates mitochondrial apoptosis in Myc mammary
tumors Myc is highly over-expressed in breast cancer, for example, via amplification (Naidu et al.,
2002). Furthermore, Myc-induced tumorigenesis often requires secondary hits such as
Bcl-xL overexpression, or mutations in Ras or p53, which enable Myc to fully induce
proliferation since its apoptotic phenotype is inhibited (Dang, 2012; Kelly et al., 2011).
However, it is possible that there are apoptotic pathways in Myc overexpressing tumor
cells that could be easily re-activated by specific inhibitors or activators, making these
secondary mutations defective. ABT-737 is a BH3 mimetic small molecule inhibitor that
was originally designed to act like a de-repressor of the anti-apoptotic Bcl-2 family
proteins, Bcl-xL, Bcl-2 and Bcl-w (Oltersdorf et al., 2005). Our previous results from Bak
activation studies suggested that key regulators for Myc-induced mitochondrial priming
were Bak and proteins regulating Bak (I, III). Hence, it was not surprising that WAP-Myc
mouse mammary tumors were sensitive to ABT-737. Likewise, human mammary
epithelial cells expressing Myc synergistically induced apoptosis with ABT-737 (our
unpublished results). Furthermore, our analysis showed that Wap-Myc tumors expressed
Bcl-xL (IV). Therefore, we wanted to explore whether BH3 mimetic ABT-737 would be
such re-activator of Myc apoptosis in WAP-Myc mammary tumors. In our experiments
Wap-Myc tumors arose spontaneously with a 3.5-month latency and subsequent
administration of ABT-737 was able to inhibit tumor growth (IV). In accordance, Kelly et
al. also showed that ABT-737 augments apoptosis of preneoplastic B-lymphoid cells of
Eμ-myc transgenic mice and prolonged lymphoma-free survival (Kelly et al., 2011). The
amount of individuals with presence of pulmonary metastases was diminished to half in
ABT-737-treated animals. Importantly, the endogenous Wap-Myc tumors were
heterogeneous in growth rate and probably also with genetic backgrounds, all acquiring
different secondary mutations. Previous results show evidence that resistance to ABT-737
is associated with overexpression of Mcl-1 (Konopleva et al., 2006; van Delft et al., 2006).
However, the survival and growth rates of ABT-73-treated Myc-tumors did not show any
correlation with Bcl-2 family protein expression levels when compared to control group.
Further analysis could reveal, whether ABT-737 induces evolution of tumors involving
changes in Bcl-2 family proteins expression levels. Thereby, tumor might acquire
resistance. Subsequently, another approach was taken and Wap-Myc tumors were
syngrafted. Importantly, the syngrafted tumor resembled endogenous Myc tumor and were
positive for basal marker (CK14) and epithelial cell markers and had basement membrane
disorganized. Similarly, ABT-737 administered syngrafted Myc tumors grew slower and
tumors showed markers for increased apoptosis, however no differences were seen in
survival. ABT-737 was apparently toxic for mice that had undergone a surgery and had
wound still healing at the place of intraperitoneal injections given daily. This resulted in
an unexpected death of ABT-737 treated mice. Of note, one of the side-effects of
treatment with ABT-737 is acute thrombocytopenia (low platelet count) (Wilson et al.,
2010). Thus, the results suggest, that ABT-737 might re-activate apoptosis in Myc tumors,
however, to be able to see an effect on survival, a less toxic orally bioavailable ABT-263
should be used.
8. Cytoplasmic tumor suppressor p53 is activated by Myc to prime
mitochondrial apoptosis Emerging evidence suggests that p53 is involved in Myc-induced tumorigenesis and
apoptosis (Hemann et al., 2005). In addition to growth arrest, p53 has major role in
mediating the mitochondrial apoptosis pathway after DNA damage, for example, by
transcribing Bcl-2 family members Puma and Bax (Chipuk et al., 2005). Furthermore
when inducing apoptosis, p53 can act independently of its nuclear function and directly
interact and activate Bak and/or Bax at mitochondria and, can be inhibited by Bcl-xL
(Eischen et al., 2001a). Surprisingly, in our analysis p53 was found from mitochondrial
and cytosolic compartments in mammary epithelial cells if Myc was activated (III). In
similar experiments, drug-induced DNA damage was able to stabilize p53 at the nucleus
and induce upregulation of growth inhibitory genes like p21 indicating a different
mechanism for p53 action. Concordantly, neither could the DNA damage induce Bak
activation what Myc did. Previously, activation of oncogene Myc and Myc-induced DNA
damage have been shown to result in phosphorylation of Serines 15 and 37 of p53.
Importantly, phosphorylation seems to be required for inhibition of Myc-induced
tumorigenesis (Pusapati et al., 2006; Sluss et al., 2010). Also our results show that Myc
stabilizes p53 and phosphorylates p53 at Serine 15 in various cell types and also Wap-
Myc mouse mammary hyperplastic tissue expressed Ser15 phoshorylated p53 (III).
Interestingly, a small molecule Mdm2 inhibitor, Nutlin-3a, stabilized and accumulated p53
both at nucleus and mitochondria, and also activated Bak. In contrast, Nutlin-3a was able
to able induce p53-mediated transcription of p53 target genes, like TIGAR, whereas Myc
was not. Likewise, Mdm2 inhibitor ARF has been shown to be involved in Myc-induced
apoptosis, however whether ARF in involved in mitochondrial function of p53 remains to
be elucidated (Zindy et al., 1998). All together, this data does not exclude that
phosphorylation of p53 in Myc-activated cells would not involve DNA damage, but is
suggests that drug-induced DNA damage and Myc activation direct p53 at different
compartments at cell and possibly leading to different phenotypes. In addition, ectopic
transcription-deficient p53QS-mutant experiments in p53-null MCF10A cells indicated that
p53-mediated Bak activation was a transcription-independent phenomenon (III). Thus it
was evident that p53 seemed to have a cytoplasmic role in Myc-induced mitochondrial
priming and Bak activation.
The Bak and Bax are regulated either by inhibiting anti-apoptotic interaction, or directly
activating BH3-only protein (Tait and Green, 2010). Importantly, conformational
activation of Bak is shown to be regulated by Bcl-xL and Mcl-1-mediated BH3:groove
interactions (Willis et al., 2005). However in our analysis, Myc activation did not change
Bcl-xL or Mcl-1 association to Bak as measured by co-immunoprecipitation (III).
Interestingly, ABT-737 has been shown to also release Bcl-xL from p53 and indeed, we
saw loss of p53 and Bcl-xL interaction in ABT-737-treated epithelial cells. Previously, p53
has been suggested to bind the “underside” of the Bcl-xL and therefore p53 and Bak
should not compete for the same interaction site, the hydrophobic binding groove (Hagn et
al., 2010). Surprisingly, Myc activation also decreased Bcl-xL association with p53,
possibly indicating free apoptotic p53 that could thereafter be able to directly activate Bak
as suggested earlier (Chipuk et al., 2004). Additionally, interaction analysis using a
proximity-ligation assay of p53 showed that Myc activation results in co-localization of
p53 and Bak, and, a simultaneous decrease in interaction of p53 and Bcl-xL (III). However,
to completely understand the role of Bcl-xL-Bak interaction other approaches should be
considered. In addition, post-translational modification of Bcl-xL has been indicated to
affect the function and interactions of Bcl-xL and could be involved in Bak activation by
Myc (Basu and Haldar, 2003). However, we did not see, for example, phosphorylation of
Serine 62 of Bcl-xL in Myc-activated cells (our personal results). Finally, results from
p53-deficient cells demonstrate that Myc-induced Bak activation is p53-dependent (III).
Furthermore, our RNAi screen to decipher Bcl-2 family members involved in Myc and
TRAIL apoptosis suggested that p53 interaction partner and derepressor BH3-only protein
Puma could be involved in Myc and TRAIL synergistic death (our personal results).
However, the exact role of Puma in regulating Bak activation or p53 functions by Myc
need to be further elucidated. Thus these data indicate that phosphorylated p53 in Myc
expressing cells is Bcl-xL free and accumulates at the mitochondria to possibly directly
mediate activation of Bak.
9. The energy sensor AMPK is behind the Myc-mediated apoptotic
sensitivity (III)
P53 is phosphorylated and stabilized by a variety of kinases involved in the DNA damage
response. Altogether p53 has 15 sites for phosphorylation some increasing tetramerization,
some decreasing Mdm2 association (Ser15) and a few improving acetylation (MacLaine
and Hupp, 2011). The p53 Serine 15 has been suggested to be crucial for the apoptotic
function of Myc in mouse models (Jones et al., 2005). In our analysis, pharmacological
screening of kinases mediating Myc-induced phosphorylation of p53 revealed a tumor
suppressor AMPK as one of the possible kinases involved (III). AMPK is known to be
involved in energy stress-mediated growth inhibition and phosphorylates p53 during stress
(Jones et al., 2005). In addition, AMPK is known to inhibit catabolic pathways (glycolysis,
fatty acid oxidation and mitochondrial biogenesis) and inhibits anabolic pathways
(gluconeogenesis, glycogen and fatty acid and protein synthesis) (Luo et al., 2010). In
contrast, less data exists about the role of AMPK in mediating apoptosis. Furthermore, our
results demonstrate that pharmacological activation of AMPK was able to induce
phosphorylation of p53, and subsequent Bak activation and cell death sensitivity (III).
Interestingly, AMPKT172 and its phosphorylation target ACCSer79 were increasingly
phosphorylated in Myc-expressing cells and Wap-Myc mouse mammary hyperplasia,
indicating AMPK activation as well. Previously, Lkb1 kinase, CAMKK, and TAK have
been suggested to phosphorylate and activate AMPK, however it remains to be elucidated
in the future through which kinase AMPK is phosphorylated in Myc-expressing. However,
based on our results Lkb1 has tumor suppressive function against tumorigenic Myc. Taken
together, the role of AMPK activation in mediating apoptosis seems to be evident. These
results suggest that AMPK is a key protein in mediating activation of p53 in Myc
expressing cells.
10. Bak activation coupling oncogene-mediated metabolic stress and
apoptosis
AMPK activation is an energy sensor mechanism that responds to the increase of AMP or
decrease in ADP levels in the cells (Hardie, 2011). Interestingly, our data suggests that
ATP levels are dropping and ADP/ATP ratio increases (III).
Importantly, AMPK has an emerging role as a nutrient and energy sensor that maintains
energy homeostasis by increasing catabolic pathways and simultaneously acting as tumor
suppressor (Hardie et al., 2012; Luo et al., 2010). Therefore, it was speculated that
AMPK was activated as a response to such metabolic stress induced by Myc. Previous
data show that Myc stimulates glutamine uptake and metabolism through up-regulation of
glutamine transporters and glutaminase (GLS) (Wise et al., 2008). Likewise, in our
analysis Myc was able to up-regulate GLS expression and rendered cell addicted to
glutamine metabolism (III). Furthermore, these data raises a possibility that AMPK is
involved in Myc-induced metabolic reprogramming, which can benefit tumorigenesis
(Liang and Mills, 2013). In accordance with previous publications, Myc was also able to
induce glycolysis in mammary epithelial cells (our personal results). Additionally, nutrient
availability seems to be a determining factor for Myc-induced metabolic pathway
activation since, for instance, under glutamine-deprivation Myc was shown to direct
reprogramming towards glycolysis (our personal results). Taken together, all these
changes are indicative for activation of catabolic pathways and oncogene-induced
metabolic transformation suggesting that Myc-expressing cells are simultaneously
undergoing energy stress and metabolic reprogramming.
Furthermore, Myc, AMPK, or p53 activation, were able to induce Bak conformational
change and sensitize mitochondria to apoptosis (I-IV). Since Myc-induced metabolic
reprogramming was co-incidental with AMPK and Bak activation, it could be
hypothesized that Bak activation is a result of a metabolic stress. Interestingly, breast
cancer cells expressing Myc had an increased level of Bak activation in
immunofluorescence analysis (III), indicating a possible oncogene stress indicated by Bak
N-terminal exposure. Previous results suggest that “oncogene stress” is partly results from
the enforced induction of cell cycle entry (mitotic stress or replicative stress) by oncogene
that makes cells more prone to apoptosis (ref). However, data presented here indicate that
oncogene-induced metabolic stress is not necessarily the result of excessive proliferation,
since Myc activation in fully-formed mammary acini was able to induce apoptotic
sensitivity without increase in proliferation status. Furthermore, neither did Myc activation
in two-dimensional cell culture induce any evident increase in cell population, even it did
induce cell cycle re-entry in growth deprived conditions (II and our personal results).
Importantly, Myc expressing cells had increased catabolic pathway activity, such as
glutaminolysis, fatty acid oxidation and glycolysis, indicating metabolic reprogramming
(Ward and Thompson, 2012). Hence, a more plausible explanation for stress in Myc
overexpressing cells could be the activation of multiple signaling pathways at the same
time and energy-stress deriving from the usage of catabolic pathways and overall increase
in biosynthesis (Dang, 2013). Once these energy-stressed cells are declined with nutrients,
they may not be able to cope with crucial tasks and become more vulnerable to apoptosis.
Already, some discussion has emerged about ideas how metabolism could impinge on
apoptotic signaling pathway and those involve p53, caspase-2 or Bcl-2 family members
(Buchakjian and Kornbluth, 2010). For instance, growth arrest and metabolism have
already been previously linked via AMPK and p53 (Buchakjian and Kornbluth, 2010).
Furthermore, Bcl-2 family members have shown to directly regulate metabolic pathways
via Bad acting as glucokinase in glycolysis and via Noxa-Mcl-1 interaction balancing
apoptotic sensitivity during glucose insufficiency (Alves et al., 2006; Danial et al., 2003).
In addition, our results indicate that nutrient availability can affect on metabolic
reprogramming and apoptotic sensitivity (III, unpublished results). Importantly, as
mediators of apoptotic sensitivity AMPK and p53 are acting as tumor suppressors in Myc-
expressing cells, however their function as regulators metabolic reprogramming is
possible. All together, these data suggest that the metabolic and mitochondrial apoptotic
pathways can be coupled in a novel AMPK-p53-Bak-dependent way (Figure 13).
Figure 13. Model for Myc-effects in mammary cells. Myc-induced activation of signaling
pathways involved in cell growth and proliferation requires metabolic adaptation. This metabolic
reprogramming enables growth, however, it also lowers the cellular energy as measured by ATP
levels, which is recognized by the energy sensor AMPK. Consequently, AMPK phosphorylates the
tumor suppressor p53 and, thereby p53 is stabilized and accumulating at the mitochondrial
compartment. Finally, mitochondrial p53 engages and activates Bak that makes mitochondrial
apoptosis pathway primed for apoptotic stimuli. Furthermore, this model indicates that by targeting
metabolic pathways or Bcl-2 family members in Myc-expressing tumor cells, cell can become
vulnerable to death. All together, this model explains how Myc-induced metabolic reprogramming
and cell death sensitivity can be coupled.
CONCLUSIONS The studies presented here provide evidence that oncogene Myc-induced apoptosis is
mitochondrial dependent in mammary epithelial cells and in other cell types. Our findings
expose new evidence for Bcl-2 family member mediated control of Myc-induced
apoptosis and Bak-dependent priming of the mitochondrial pathway by a mechanism
involving mitochondrial p53 (I-III). Furthermore, data here suggest a novel definition for
Bak activation and N-terminal exposure as a conformational change that is separate from
BH3 exposure and oligomerization of Bak. We suggest that the conformational change of
Bak is an apoptotic priming mechanism whereas the oligomerization of Bak represents the
pore forming activity, considered as a point-of-no-return in the apoptotic process. Hence
we open new ideas for the current “direct activator/derepression” model suggested earlier
(Tait and Green, 2010). The data presented herein also suggest that BH3 mimetic drugs
can sensitize tumor cells to Myc-apoptosis in vivo (IV), which also highlights the role of
Bcl-xL as a key controller of Myc-dependent apoptosis in epithelial tumors and points new
therapeutic possibilities to treat breast cancer.
Emerging data indicate that p53 has transcriptionally independent functions and that
cytoplasmic p53 has apoptotic function involving interactions with Bcl-xL and Bak/Bax
(Chipuk et al., 2004; Sot et al., 2007). Correspondingly, our results indicate that p53
cytoplasmic p53 has a crucial role in Myc-induced priming of the mitochondrial apoptosis
pathway (III). According to our results, p53 is required for Myc-induced engagement of
the mitochondrial apoptosis pathway via transcription-independent mechanisms, by
functioning as a direct activator for Bak (III). Therefore we suggest a Bak activation
model, where Bak is kept in check and the N-terminus hidden by Bcl-xL. Myc or the
pharmacological inhibition of Bcl-xL is enough for the Bak to be able to undergo the first
conformational change, exposure of the N-terminus. In addition, our results indicate that
Myc induced activation of the AMPK and phosphorylation of its downstream target p53.
Furthermore, we hypothesized that the phosphorylated p53 accumulates in the
mitochondria and by altering the interaction between Bak and Bcl-xL, the Bak finally
becomes activated. Moreover, it could be speculated that Myc activation could induce a
partial release of Bak from tight inhibition by Bcl-xL where Bcl-xL is adopting a different,
yet unknown, conformation allowing the first Bak conformational change. However, more
studies are needed to clarify how these molecular level alterations lead to conformational
activation of Bak.
Finally, the results reveal that Myc-induced metabolic reprogramming and apoptotic
sensitivity are tightly coupled. Here we suggest that activation of Myc promotes metabolic
transformation by inducing glycolysis, activating the glutaminolytic pathway (GLS), and
by promoting a catabolic mode of metabolism through inhibiting fatty acid synthesis (III).
Furthermore, Myc-induced metabolic transformation decreased cellular ATP levels,
activating the energy sensor AMPK that releases carbon for bioenergetic reactions and
promotes biosynthesis of nucleic acids. Thus, we show that Myc-induced metabolic stress
and direct activation of AMPK both induce apoptotic sensitivity via p53 and Bak-
mediated pathway. However, future studies are still needed to understand how, for
example, Myc-induced glutamine-dependence (addiction) is linked to the
apoptotic mechanisms described here or how p53 mechanistically activates Bak at the
surface of the mitochondria. Interestingly, these results also indicate that nutrients and
their availability and sensing affect on transformation capacities and apoptotic sensitivity
of cancer cells possibly in a very critical manner and should be considered carefully in
cancer therapies. The contextual role of AMPK as part of the cancel cells metabolic
reprogramming and as a tumor suppressor was emphasized by our results and will be
important question in future. Furthermore, it will be interesting to explore possibilities to
use drugs that modulate AMPK activity as therapeutic sensitizers of tumor cells to Myc-
dependent apoptosis. Finally, our results bring new insight into oncogenic stress and
apoptotic sensitivity by Myc highlighting the role of altered cell metabolism (1-IV). All
together, it will be future challenge to understand how that knowledge of crosstalk
between metabolic stress and apoptotic sensitivity can be used for therapeutic
interventions in cancer.
ACKNOWLEDGEMENTS
The work presented in this dissertation has been carried out at the University of Helsinki in the department of Biochemistry and Developmental biology at the Institute of Biomedicine in Biomedicum Helsinki and under the Translational Cancer Biology Research Program. I would like to express my gratitude to professors Esa Korpi, Olli Jänne, Sampsa Hautaniemi, Lauri Aaltonen, Hannu Sariola and Tomi Mäkelä for providing the excellent working facilities. I am most grateful to my supervisor, Juha Klefström, for all his support and giving me the possibility to start my science career and to work in his group. More importantly, Juha is acknowledged for giving me the opportunity to take responsibility of my own research goals and independently design experiments to test hypothesis. Juha is also thanked for being an example of enthusiastic scientist, always creating an interesting story. I sincerely acknowledge Professors John Eriksson and Ville Hietakangas for their grateful review of the thesis on a tight schedule. Minna Ahvenainen is warmly acknowledged for her revision of the English language. John Eriksson and Philippe Juin are also acknowledged for their guidance and valuable comments as members of the thesis follow-up committee. Also Professor John Eriksson and Tapio Heino are thanked for reviewing my master thesis. I wish to thank also Helsinki Biomedical Graduate School for all financial support. I would like to warmly thank all my collaborators especially Tomi Mäkelä and Outi Monni for their valuable comment and support. Also Panu Kovanen is acknowledged for expertise in histopathology and Daniel Laajala and Tero Aittokallio for their knowledge and input. This study would not have been possible without all the wonderful people, friends and colleagues, around me. I want to thank all the present and former Klefalab members that participated in the lab meetings with intelligent comments and made our working days enjoyable. Especially I like to acknowledge all co-authors: Vilja Eskelinen, Johanna Partanen, Annika Hau, Topi Tervonen, Sirkku Saarikoski, Heidi Haikala, Elsa Marques, for excellent input. Special thanks go to Johanna Englund and Annika Hau who shared with me the early days in the lab and Vilja Eskelinen for being nice and reliable co-worker, you were always there for me if needed. Also borrowed postdoc Niklas Ekman is thanked for sharing knowledge and opinions about everything with us. My deepest gratitude goes to very helpful technical stuff from Klefalab and Biomedicum Genomics: Tiina Neejärvi, Tarja Välimäki, Teija Inkinen and also secretary Anu Taulio is thanked for everything. Many things would have gone very difficult without you guys. Tiina, Vilja and Heidi are especially thanked for enormous helping during my leaves and enabling working from home.
Especially, Professores Tomi Mäkelä and Päivi Ojala are acknowledged for constructive comments during lab meetings and retreats. Also their lab members, specifically Katja, Emma, Kari, Suski, Annika, Pekka, Anou, Liina, Eeva, Tea, Thomas and their helpful technicians Kirsi and Birgitta are thanked for crateful discussion and most importantly, for all the fun we had. Special thanks go to Kari for important comments concerning AMPK. I also want to thanks my colleagues Essi Havula, Emma Kuuluvainen, Anne Mäkelä, Katja Helenius and Vilja Eskelinen for sharing science and life with me. Also all people attending Kalakisat and Ahvis are thanked for the fun and unforgettable memories. My colleagues Maria Rajecki, Henna Heinonen, Minna Jääskeläinen, Mikko Turunen and Tatyana Lepikhova are thanked for giving me the opportunity to be involved in their interesting projects, it has been pleasure. I also want to thanks my colleagues and study friends from Viikki times for fruitful discussions about working in the science world and being always so positive. Special thanks goes to my dearest friends Salla, Janica, Anski and Sointu. Your support and company has been invaluable. It is always relaxing to discuss with you about something else than science, for example, teaching children… Most importantly, you have walked with me during my whole life, listening, teaching, traveling and having fun. Finally, I wish to thank my family for love and support, especially my husband Tero for his patience and for keeping our life busy with football. Most of all, my son Onni and daughter Emmi, are sincerely thanked for teaching me a lot in life – you are my sunshines. I also want to express my gratitude to my parents, Jouko and Marjut, for teaching me to question things in life and not to depend on other people`s opinions and for always encouraging me for challenges. I also would not be here without my wonderful sisters, Elina and Minna, and brothers, Köpi and Aaro, who always are there for me and were in huge help with combining my thesis and children. Also Tero`s parents and my grandparents, Aili and Eino, are thanked for endless care and support for thesis. This work was supported by personal grants rom the Academy of Finland, the Sigrid Juselius Foundation, the Finnish Cancer Organizations, the Biomedicum Foundation, the Emil Aaltonen Foundation, the Ida Montini Foundation, the Orion-Farmos Research Foundation, the Paulo Foundation, University of Helsinki and Helsinki Biomedicum Graduate Program. Helsinki, March 2014
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