characterization of the e3 ubiquitin ligase pirh2 · the p53 gene encodes a transcription factor...
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Characterization of the E3 Ubiquitin Ligase Pirh2
by
Elizabeth Tai
A thesis submitted in conformity with the requirements for the degree of Doctor of Philosophy
Graduate Department of Medical Biophysics University of Toronto
© Copyright by Elizabeth Tai (2010)
II
Characterization of the E3 Ubiquitin Ligase Pirh2
Elizabeth Tai
Doctor of Philosophy
Department of Medical Biophysics University of Toronto
2010
Abstract
The p53 tumour suppressor gene is inactivated by mutation in over 50% of all human
cancers. The p53 protein is activated and stabilized through several post-translational
modifications in response to various stresses and promotes cell cycle arrest and apoptosis.
Thus, regulation of p53 is critical for normal cellular function. Pirh2 is a p53-regulated
gene recently identified in our laboratory which encodes an E3 RING-finger ubiquitin
ligase that binds to p53 and negatively regulates p53 by targeting it for ubiquitin-
mediated proteolysis. Pirh2 is similar to another well-characterized E3 RING finger
ubiquitin ligase, Mdm2, which also participates in a similar negative feedback loop with
p53. At least seven E3 ubiquitin ligases are known to target p53 for degradation and the
reason for this functional redundancy is unclear. The purpose of this study is to
characterize Pirh2 activity.
This study has two aims the first is to identify additional interacting proteins for
Pirh2, and the second is to delineate Pirh2 regulation of p53. Using several tandem
affinity purification strategies and a GST-pull down approach, we have identified PKC
as a candidate interacting protein.
The second aim is to further characterize Pirh2 regulation of p53. Splenocytes and
thymocytes from Pirh2-/- mice demonstrate a subtle increase in total p53 levels after
irradiation when compared to wild-type controls. Phosphoserine 15 p53 levels are
significantly higher in splenocytes and thymocytes from Pirh2 -/- mice relative to wild-
type counterparts. Cells stably transfected with Pirh2 have decreased levels of
phosphoserine 15 p53 and decreased induction of p21 relative to vector control and
Mdm2 expressing cells.
The stability of the p53 protein is primarily regulated through ubiquitin mediated
proteolysis, and there are multiple ubiquitin ligases targeting p53 for degradation. Here
we are able to address the question of functional redundancy by indicating that Pirh2 can
target serine 15 phosphorylated p53 which is reported to not be regulated by Mdm2.
III
Acknowledgements
This thesis arose from many years of work previously completed in the
Benchimol lab. During my time working on the project, I was fortunate to receive
several contributions from many individuals; without their assistance, I would not have
been able to complete this thesis. I would now like to take the time to humbly
acknowledge those individuals.
I would like to thank my supervisor Dr. Sam Benchimol for his advice, guidance
and patience in helping me to develop scientific skills; it has been a fantastic learning
experience. Above all, he has instilled in me a strong sense of integrity, and enthusiasm
in the pursuit of science.
I would also like to acknowledge my committee members Dr. Cheryl Arrowsmith,
Dr. Razqallah Hakem, and Dr. Linda Penn. Their advice and support have been
instrumental in the development of my project and in my development as a researcher.
I would also like to thank the past and present members of the Benchimol lab for
useful and critical discussion. I would like to thank Weili Ma for her unwavering support
and for sharing her extensive technical expertise. I would also like to acknowledge Dr.
Keith Wheaton, Dr. Wissam Assaily, Dr. Christina Berube and Dr. Jenny Ho for
encouragement and scientific discussion. They are my scientific role models, and set a
high standard for which I strived to achieve.
I would also like to thank Dr. Yi Sheng, Shili Duan, and Dr. Anne Hakem for
sharing their technical expertise.
I would like to thank my mother, father, brother and friends for their steadfast
support. Their encouragement has been instrumental to the completion of my thesis.
IV
Table of Contents
CHAPTER 1: Introduction...............................................................................................1 The p53 Network .................................................................................................................2
p53 as a tumour suppressor......................................................................................2 p53-dependent cell cycle arrest................................................................................3 p53-dependent apoptosis..........................................................................................5 Choice of response...................................................................................................7 Mouse models of p53 tumour suppression ..............................................................8 Regulation of p53 function ....................................................................................11 Phosphorylation of p53 at serine 15.......................................................................16
Ubiquitination ....................................................................................................................17 Ubiquitin ................................................................................................................17 The ubiquitination mechanism...............................................................................18 Ubiquitination and signalling.................................................................................24 Ubiquitination and protein degradation .................................................................26 Deubiquitination ....................................................................................................28
Ubiquitination and regulation of p53.................................................................................30 Pirh2.......................................................................................................................35
Thesis Hypothesis ..............................................................................................................37 CHAPTER 2: Identification of Pirh2 Interacting Proteins .........................................38
Abstract ..................................................................................................................39 Introduction............................................................................................................39 Experimental Procedures .......................................................................................41
Cell Culture................................................................................................41 Plasmids .....................................................................................................41 Generation of Clones Expressing pCMV TAP hPirh2 ..............................42 Generation of Clones Expressing pBabe neo Pirh2-SPA ..........................42 Generation of Clones Expressing pBabe puro Pirh2-HF...........................42 Ubiquitination Assay .................................................................................43 Coimmunoprecipitation .............................................................................43 Western blots .............................................................................................44 TAP Purification ........................................................................................44 SPA Purification ........................................................................................45 His-Flag Purification..................................................................................46 GST-pull down...........................................................................................47
Results....................................................................................................................47 Generation and characterization of TAP-hPirh2 clones ............................47 Purification of TAP and TAP-hPirh2 ........................................................51 Generation of clones and purification of Pirh2-SPA .................................51 Generation of clones and purification of Pirh2-HF ...................................54 Identification of Pirh2 interactors using GST-pull down ..........................56
Discussion..............................................................................................................57
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CHAPTER 3: Pirh2 targets phosphoserine 15 p53 for ubiquitination......................61
Abstract ..................................................................................................................62 Introduction............................................................................................................62 Experimental Procedure.........................................................................................64
Cell Culture................................................................................................64 Splenocyte and Thymocyte Culture Preparation .......................................65 Preparation of MCF-7 p53shRNA Cells....................................................65 Western Blotting ........................................................................................66 Immunohistochemistry ..............................................................................66 Coimmunoprecipitation .............................................................................66 Ubiquitination Assay .................................................................................67 Flow Cytometry .........................................................................................67
Results....................................................................................................................68 Pirh2-/- Tissues Show Elevated Levels of p53 After DNA Damage.........68 Pirh2-/- Tissues Show Elevated Levels of Phosphoserine 15 p53
After DNA Damage .......................................................................68 Apoptotic Rate is Increased in Pirh2-/- Mice ............................................70 Pirh2 Can Regulate p53 Mutants for Serine 15 .........................................73 Cells Overexpressing Pirh2 Decrease Levels of Phosphoserine 15 p53
and p21, But Does Not Affect Cell Cycle Arrest...........................73 Pirh2 Coimmunoprecipitates with Phosphoserine 15 p53 and promotes
Phosphoserine 15 p53 Ubiquitination............................................78 Discussion..........................................................................................................................80 CHAPTER 4: Summary and Future Directions ..........................................................84
I. Summary of Findings: Identification of Pirh2 Interactors..................................85 Future Directions: Identification of Pirh2 Interactors............................................85
Identification of Pirh2 Cofactors and Substrates .......................................85 Characterization of the Interactors.............................................................90 Identification of the Pirh2 Degron .............................................................92
II. Summary of Findings: Pirh2 Regulation of Phosphoserine 15 p53 ..................93 Future Directions ...................................................................................................96
Dissecting the Functional Redundancy of p53 Ubiquitin Ligases.............96 Non-Proteolytic Ubiquitination of p53 ......................................................98 Pirh2-Mediated Ubiquitination of p53.......................................................99
III. Pirh2 and Cancer............................................................................................101 REFERENCES...............................................................................................................105
VI
List of Illustrations Figure 1.1 Post-Translational Modifications of p53 .........................................................12 Figure 1.2 Ubiquitin Linkage Forms ................................................................................19 Figure 1.3 Ubiquitination Reaction Mechanism...............................................................21 Figure 1.4 Ubiquitination in the NF-κB Pathway.............................................................25 Figure 2.1 Generation of TAP-hPirh2 clones in H1299 cells ...........................................48 Figure 2.2 Functional Assays for TAP-hPirh2 Activity ...................................................50 Figure 2.3 Purification of TAP and TAP-hPirh2 ..............................................................52 Figure 2.4 Generation and Purification of Pirh2-SPA Clones..........................................53 Figure 2.5 Generation and Purification of Pirh2-HF Clones ............................................55 Figure 2.6 GST-pull Down to Identify Novel Pirh2 Interacting Proteins.........................58 Figure 3.1 Elevated p53 Levels in Pirh2-/- Tissues..........................................................69 Figure 3.2 Elevated Levels of Phosphoserine 15 p53 in Pirh2-/- Mice ............................71 Figure 3.3 Elevated Levels of Apoptosis in Pirh2-/- Mice ...............................................72 Figure 3.4 Mdm2 and Pirh2 Can Regulate p53 Serine 15 Mutant....................................74 Figure 3.5 Decreased Levels of Phosphoserine 15 p53 and p21
in Pirh2 Overexpressing Cells ...................................................................76 Figure 3.6 Overexpression of Pirh2 Does Not Decrease S-phase
After DNA Damage ...................................................................................77 Figure 3.7 Pirh2 Interacts with and Ubiquitinates Phosphoserine 15 p53........................79 Figure 4.1 Model for Direct Pirh2 Regulation of p53 ......................................................95
VII
List of Abbreviations
ADP – Adenosine Diphosphate ATP – Adenosine Triphosphate BrdU – Bromodeoxyuridine BrdU-FITC – Bromodeoxyuridine – Fluorescein BSA – Bovine Serum Albumin CBP – Calmodulin Binding Peptide cIAP – Cellular Inhibitor of Apoptosis cICAT – cleavable Isotope-Coded Affinity Tags DNA – Deoxyribose Nucleic Acid DTT – Dithiothreitol EDTA – Ethylenediaminetetraacetic acid EGFR – Epidermal Growth Factor Receptor EGTA – Ethylene glycol tetraacetic acid EMSA – Electrophoretic Mobility Shift Assay ERAD – Endoplasmic Reticulum Associated Protein Degradation FACS – Fluorescence Activated Cell Sorting FCS – Fetal Calf Serum GST – Glutathione – S Transferase H&E – Hematoxylin & Eosin HA – Hemaglutinnin HECT – Homologous to E6 – Associated Protein Carboxy-Terminus HEPES – 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid HPV – Human Papilloma Virus IgG – Immunoglobulin G IgH – Immunoglobulin Heavy chain IHC – Immunohistochemistry IR – Ionizing Radiation ITRAQ – Isobaric Tags for Relative and Absolute Quantification LC – MS/MS – Liquid Chromatography – Mass Spectrometry/ Mass Spectrometry LFS – Li-Fraumeni Syndrome MALDI - MS/MS – Matrix Laser Desorption Ionization – Mass Spectrometry/Mass
Spectrometry MEFs – Murine Embryonic Fibroblasts MJD – Machado-Joseph Disease mRNA – messenger Ribonucleic Acid NAC – N-Acetyl Cysteine NP-40 – Nonyl phenoxylpolyethoxylethanol OTU – Otubain proteases PAGE – Polyacrylamide Gel Electrophresis PBS – Phosphate Buffered Saline PCR – Polymerase Chain Reaction PKC – Protein Kinase C RING – Really Interesting New Gene RNAi – Ribonucleic Acid interference
VIII
IX
RPMI – Roswell Park Memorial Institute SDS – Sodium Dodecyl Sulfate SNP – Single Nucleotide Polymorphism SPA – Sequential Peptide Affinity TAP – Tandem Affinity Purification TEV – Tobacco Etch Virus Tris-HCl – Tris(hydroxymethyl)aminomethane – Hydrogen Chloride ts – temperature sensitive TUNEL – Terminal deoxynucleotidyl transferase dUTP nick end labeling UCH – Ubiquitin C-terminal hydrolase USP – Ubiquitin Specific Protease UV – Ultraviolet Radiation
CHAPTER 1
Introduction
1
2
The p53 Network
p53 as a Tumour Suppressor
p53 was initially characterized as a protein that interacts with the large T antigen
from the SV40 virus (Lane and Crawford, 1979; Linzer and Levine, 1979). Early
research suggested that p53 was involved in promoting cell division as p53 could
cooperate with oncogenic ras to transform rat embryo fibroblasts (Eliyahu et al., 1984;
Jenkins et al., 1984; Parada et al., 1984). However, it was later determined that wild-type
p53 actually suppresses cell transformation, and the previously characterized oncogenic
properties of p53 were due to mutations in the gene (Finlay et al., 1989; Hinds et al.,
1989). In fact, rearrangements and point mutations in the p53 gene were observed in the
majority of murine erythroleukemic cell lines (Ben David et al., 1988; Mowat et al., 1985;
Munroe et al., 1990). In vivo, the p53 gene is mutated in over 50% of all human cancers,
and is one of the most common mutations in cancer (Vousden and Lu, 2002). Germline
mutations of p53 have been found, and humans born with one mutated copy of the p53
gene develop Li-Fraumeni syndrome (LFS). Patients with LFS have an increased number
and earlier onset of cancers (Malkin et al., 1990). These results suggested that the wild-
type p53 protein functions as a tumour suppressor, and that mutation of the protein
promotes oncogenesis.
The p53 gene encodes a transcription factor that regulates gene expression in
response to various stresses such as DNA damage and oncogene activation. The p53
tumour suppressor protein contains an N-terminal activation domain, a central DNA
binding domain, and a C-terminal regulatory domain. The N-terminal domain of p53 can
bind various transcriptional activators such as histone acetylates p300 and CBP (Riley et
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al., 2008). The C-terminus contains the regulatory domain that includes the
oligomerization domain as p53 binds to DNA as a homotetramer in the form of a dimer
of dimers (Davison et al., 2001). The p53 consensus sequence consists of at least two
repeats of the consensus 5’- PuPuPuC(A/T)(T/A)PyPyPy - 3’ with 0-13 base pairs of
non-specific sequence in between (el-Deiry et al., 1992); the consensus binding sequence
of p53 supports the model of p53 binding to DNA as tetramer. p53 induces the
expression of genes involved in cell cycle arrest and apoptosis in response to stimuli
(Levine, 1997). The central DNA binding domain of p53 is frequently mutated in
cancers, and suggests that the ability of p53 to transactivate gene expression is critical for
tumour suppression (Vousden and Lu, 2002).
p53-dependent cell cycle arrest
p53 promotes cell cycle arrest through the transcriptional activation of numerous
target genes including p21WAF1, 14-3-3σ, REPRIMO, BTG2, GADD45 (el-Deiry et al.,
1993; Hermeking et al., 1997; Ohki et al., 2000; Rouault et al., 1996; Wang et al., 1999).
p53 induces cell cycle arrest in G1 using p21 and BTG2; GADD45, REPRIMO, and 14-
3-3σ control the G2/M transition (Chan et al., 2000; el-Deiry et al., 1993; Guardavaccaro
et al., 2000; Harper et al., 1993; Wang et al., 1999).
A more recent discovery is that p53 can also modulate cell cycle arrest through
induction of microRNAs (miRNA). Activation of p53 causes expression of the miR34
family of miRNAs that are involved in downregulating genes that promote cell cycle
entry such as CDK4 and MET. Ectopic expression of miR34 causes cell cycle arrest in
4
both G1 and G2 phase, and a decrease in S-phase cells in both tumour-derived cell lines
and normal cell lines (He et al., 2007).
Transcriptional repression by p53 is also an important part of the growth arrest
pathway. p53 mediates transcriptional repression of c-myc by binding to the c-myc
promoter and recruiting histone deacetylases. Repression of c-myc by p53 increases G1
arrest (Ho et al., 2005). These results indicate that p53 is able to cause cell cycle arrest
by both upregulating genes that promote arrest, and downregulating genes that promote
entry into the cell cycle.
Loss of p53 in cells causes inappropriate entry into S phase after treatment with
metabolic inhibitors that normally arrest cells in G1; these cells have a higher incidence
of genomic instability and gene amplification (Livingstone et al., 1992; Yin et al., 1992).
Cell cycle arrest is thought to aid in p53 mediated tumour suppression since it allows the
cell time to repair lesions in response to DNA damage.
Cell cycle arrest is also involved in senescence, a permanent state of G1 cell cycle
arrest. In experiments, replicative senescence is studied by taking normal mature cells
lacking telomerase activity, usually human fibroblasts and culturing them in vitro to
undergo multiple cell divisions until the telomeres at the end of the chromosomes are
shortened. These shortened telomeres trigger the activation of p53, primarily through the
ATM/ATR pathway. It is thought that shortened telomeres mimic double strand breaks
(Herbig et al., 2004). The p53 target gene p21 is elevated in replicative senescence, but it
is not the only p53 target that causes replicative senescence. Further studies are being
performed to determine which mediators of p53-dependent cell cycle arrest are important
for replicative senescence (Artandi and Attardi, 2005; Vaziri et al., 1999). Cell cycle
5
arrest is also involved in p53-dependent tumour suppression as rat embryonic fibroblasts
expressing oncogenic ras undergo p53-dependent oncogene induced senescence;
senescence is a physiologically relevant pathway of p53-dependent tumour suppression in
murine models of sarcoma and hepatocellular carcinoma (Serrano et al., 1997; Ventura et
al., 2007; Xue et al., 2007). Similar to replicative senescence, the p53-dependent cell
cycle target genes involved in oncogene induced senescence are unknown, and research
into the field is ongoing.
p53-dependent apoptosis
The apoptotic arm of p53 tumour suppression eliminates cells with irreparable
DNA damage or cells exhibiting unrestrained growth as a result of oncogene activation
(Vousden and Lu, 2002). Various pro-apoptotic genes are induced by p53 including
PUMA, Noxa, Bax, p53AIP1 and PIDD (Lin et al., 2000; Nakano and Vousden, 2001;
Oda et al., 2000a; Oda et al., 2000b; Yin et al., 1997). PUMA, Noxa, and Bax are located
at the mitochondria, and aid in permeabilization of the outer membrane to induce
cytochrome c release causing activation of caspase-9 resulting in apoptosis (Fridman and
Lowe, 2003). Deletion of Apaf-1 or caspase-9 in cells results in protection from p53-
dependent apoptosis; in fact, loss of either Apaf-1 or caspase-9 can substitute for loss of
p53 to prevent c-myc-induced apoptosis, suggesting that this is the main pathway for
p53-dependent apoptosis (Soengas et al., 1999)
More recently, it has been determined that p53-dependent apoptosis can also
occur in a transcription-independent manner. The p53 protein has been found to be
localized to the mitochondria, and can directly activate the pro-apoptotic protein Bax to
6
induce cell death (Chipuk et al., 2004). p53 has also been found to bind to Bcl-XL
through its DNA binding domain, and mitochondrial p53 can result in decreased viability
(Mihara et al., 2003). The model for apoptosis induced by mitochondrial p53 states that
p53 translocates to the mitochondria upon DNA damage where it is sequestered by Bcl-
XL. Concurrently, nuclear p53 induces the expression of PUMA which then binds Bcl-
XL and releases p53 to bind Bax and induce apoptosis (Chipuk et al., 2005). A second
model of how p53 can directly induce apoptosis at the mitochondria claims that
mitochondrial p53 can bind and activate Bak by causing Bak oligomerization which
results in cytochrome c release and apoptosis (Leu et al., 2004). These models of how
p53 can directly induce apoptosis at the mitochondria need to be resolved. More
importantly, how the transcription-dependent arms and the transcription-independent
arms of p53 cooperate to cause cell death is a key question that remains unanswered.
p53-dependent apoptosis is also an important component of p53-dependent
tumour suppression. Mice that overexpress c-myc from the IgH promoter in p53+/- B-
cells develop B-cell lymphoma, and the remaining wild-type allele of p53 is mutated or
lost. However, the wild-type allele of p53 can be retained by overexpressing Bcl-2 or a
dominant negative caspase-9; these are both methods to inhibit mitochondrial apoptosis
downstream of p53 (Schmitt et al., 2002). Furthermore, a knock-in mouse that contains a
p53 construct that has deleted the proline rich domain has reduced capacity for inducing
p53-dependent cell cycle arrest but retains the ability to induce p53-dependent apoptosis
is still protected from spontaneous tumour formation (Toledo et al., 2006). The
previously described results suggest that the apoptotic arm of p53 is critical for mediating
tumour suppression.
7
Choice of Response
The ability of p53 to promote both cell cycle arrest and apoptosis and how p53
decides between these two cellular responses is one of the key questions in the field. This
question has been extensively addressed by many studies that have outlined several key
determinants that influence this choice. These factors include the intensity of stress, the
efficiency of DNA repair, extracellular survival factors, level of p53 expression and
activating, oncogenic composition of the cell, intracellular death/survival pathways and
selective transactivation/repression of p53-target genes in different cell types (Bouvard et
al., 2000; Fei et al., 2002; Johnson et al., 1993; Lin and Benchimol, 1995; Lin et al., 2002;
Weber and Zambetti, 2003; Wu et al., 2005). A classic example uses fibroblasts as a
model; normal fibroblasts undergo p53-dependent cell cycle arrest in response to DNA
damage. However, fibroblasts that have been transformed by an activated oncogene,
such as adenovirus E1A protein, cell cycle proteins E2F-1 or myc, undergo p53-mediated
apoptosis. Cells ectopically expressing pro-survival proteins Bcl-2 or Bcl-XL are
protected from p53-dependent apoptosis, and constitutively active PI3’K and PKB/Akt
are able to delay p53-induced apoptosis (Chiou et al., 1994; Levine, 1997; Lin et al., 2002;
Nahle et al., 2002; Sabbatini and McCormick, 1999; Schott et al., 1995). The p53-target
gene Slug is p53-responsive and is expressed in hematopoetic cells. Slug binds to the
promoter of the pro-apoptotic p53 target gene PUMA to represses p53-dependent
transactivation. This results in a preference for cell cycle arrest over apoptosis (Wu et al.,
2005). The cellular outcome of the p53 response may also reflect differences in the
affinity of various promoters for p53, such that some are responsive only to high levels of
8
p53 or to certain modified forms of p53 (Resnick-Silverman et al., 1998; Sionov and
Haupt, 1999). p53 promoter selectivity leading to cell cycle arrest or apoptosis can also
be regulated by protein-protein interactions between p53 and other proteins including
ASPP, JMY, WT1, BRCA1, Hzf, p63 and p73 (Das et al., 2007; Flores et al., 2002;
MacLachlan et al., 2002; Maheswaran et al., 1995; Samuels-Lev et al., 2001; Shikama et
al., 1999).
In addition to cell cycle arrest and apoptosis as mediators of p53-dependent
tumour suppression, the antioxidant function of p53 may also play a crucial role. The
tumour prone phenotype observed in p53 knock out mice is abrogated by maintaining the
mice on an N-acetylcysteine (NAC; antioxidant) diet; p53 knockout mice treated with
NAC do not develop the characteristic thymic lymphomas observed in p53-/- mice
(Sablina et al., 2005). The p53 tumour suppressor gene also induces the expression of
antioxidant genes glutathione peroxidase 1, mitochondrial superoxide dismutase 2 and
two sestrins (Sablina et al., 2005). These results suggest that tumour prevention by p53
involves the removal of reactive oxygen species; it remains to be determined whether the
induction of antioxidant genes by p53 contributes to tumour suppression in an oncogenic
environment, or whether the involvement of p53 in dealing with reactive oxygen species
is more prophylactic.
Mouse models of p53 tumour suppression
In animal models, mice that have deleted both copies of the p53 gene are
extremely tumour prone, and die within 6 months of birth as a direct result of developing
cancer. Knockout p53 mice can develop a wide spectrum of tumours, but most
9
commonly develop thymic lymphomas and sarcomas (Donehower et al., 1992). The p53
tumour suppressor gene is rarely deleted in cancers and LFS syndrome; more commonly,
a point mutation in the p53 gene is found. As a result, new models for LFS were
developed in mice where the p53 protein with a common point mutation (273H) was
knocked in. These mice are similar to the p53 knockout mice in that they are tumour
prone. However, p53 273H has a gain of function property since tumours in the mice are
also metastatic in contrast to tumours from the p53 knockout mouse (Lang et al., 2004;
Olive et al., 2004). These observations indicate that loss of p53 function contributes to
the development of malignancy, and normal p53 function is critical for tumour
suppression.
Although loss of p53 activity is one of the most common changes in
tumourigenesis, it is unclear whether loss of p53 simply facilitates the genetic changes
such as increased genetic instability, loss of growth-arresting signals and inappropriate
cell survival that contribute to tumour development, or whether tumour growth is
dependent on keeping the p53 pathway turned off permanently (Kastan, 2007). Three
studies addressed these questions independently. The Evans laboratory generated a
knock-in mouse where p53 is fused to the estrogen receptor, and p53 activity can be
turned on by the addition of tamoxifen. In the Eµ-myc model of lymphomagenesis,
restoration of p53 activity through the addition of tamoxifen increases survival of the
mouse. In addition, the tumours eventually developed resistance to p53 suggesting that
the pathway must be inactivated to ensure tumour proliferation (Martins et al., 2006).
The Jacks laboratory addressed this question using a mouse model where a stop cassette
was placed upstream of the p53 gene, but could be removed by a tamoxifen-inducible
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Cre-recombinase. In the absence of Cre, the mice are functionally p53-deficient and
develop spontaneous or irradiation-induced lymphomas or sarcomas. Restoration of p53
resulted in apoptosis in the lymphomas and senescence in the sarcomas (Ventura et al.,
2007). The Lowe laboratory addressed how p53 affects tumorigenesis by using purified
embryonic liver cells transformed by oncogenic Hras and a short hairpin against p53 that
is repressed by the addition of doxycycline. When these oncogenic cells are injected into
immunodeficient mice, hepatocellular carcinomas quickly develop. However, when p53
expression is restored through doxycycline treatment, the tumour shows signs of
regression after four days of doxycycline treatment, and becomes undetectable after
twelve days of treatment. In this model, the hepatocellular carcinomas undergo Hras
oncogene-induced senescence (Xue et al., 2007). These results suggest that restoration of
wild-type p53 in tumours that are deficient for p53 could be therapeutically advantageous.
Gene therapy approaches that directly introduce wild-type p53 genes into tumours are
already in clinical trials (Roth, 2006). Additional in vivo experiments support the
hypothesis that p53 restoration in tumours would be therapeutically effective. In the
Evans mouse model where p53 activity is directly induced by the addition of tamoxifen,
mice were exposed to whole body irradiation to cause radiation induced lymphoma.
Restoration of p53 shortly after irradiation had no effect on tumour suppression, but did
cause an acute radiation response where the spleen, thymus, and small intestine
underwent massive apoptosis. However, restoration of p53 after the acute radiation
response had subsided abrogated tumour formation. These observations suggest that p53
tumour suppression occurs as a result of the activation of oncogenes (Christophorou et al.,
2006). These observations also support the therapeutic efficacy for restoring p53 in
11
tumours; the therapeutic effectiveness may be maximized by further activating p53 with
DNA damaging agents after it is introduced into the oncogenic environment (Kastan,
2007).
Regulation of p53 function
p53 is primarily regulated at the post-translational level and is very complex;
after stress, the protein is stabilized and accumulates in the cell. Regulation of the protein
can occur by a multitude of post-translational modifications such as phosphorylation,
acetylation, ubiquitination, sumoylation, neddylation, methylation, ADP-ribosylation, and
glycosylation (Figure 1.1) (Kruse and Gu, 2009a; Lu, 2005; Toledo and Wahl, 2006). In
addition, p53 function can also be regulated by cofactors that bind to p53. For example,
binding of 14-3-3σ to the C-terminus of p53 can increase the stability and enhance p53
transcriptional activity, and binding of BRCA1 to p53 also increases p53 activity (Yang
et al., 2003; Zhang et al., 1998). The ASPP family of protein can bind to p53 and
preferentially stimulates p53 induced apoptosis by directing p53 transcriptional activation
towards pro-apoptotic promoters (Samuels-Lev et al., 2001). The p53 target gene Hzf
controls p53 transcriptional activity by binding to the p53 DNA binding domain and
redirects p53 to cell cycle arrest promoters (Das et al., 2007; Wu et al., 2005). Inhibitory
proteins can also bind p53, as both MdmX and nucleophosmin (NPM/B23) can bind to
p53 in the N-terminus to inhibit transcriptional activation (Danovi et al., 2004; Maiguel et
al., 2004). MdmX is a critical negative regulator of p53 since MdmX null mice are
embryonic lethal at day 9.5, but embryonic lethality is completely rescued by loss of p53.
The MdmX and p53 double null mice are completely viable, but are cancer prone similar
Figure 1.1 Post-translational Modifications of p53.
p53 is heavily regulated by post-translational modifications including phosphorylation(P), ubiquitination (Ub), acetylation (Ac), methylation (Me), sumoylation (S), neddylation (N8), glycosylation (O-Glc) and ribosylation (ADP). This figure shows the major sites of these modifications. (adapted from Kruse and Gu, 2009)
Me
N8
Ac Ub
P ADP
O-Glc
S
Methylation
Sumoylation
Acetylation
Phosphorylation
Ubiquitination
ADP-ribosylation
Glycosylation
Neddylation
1 92 100 300 307 355 356 393
Transactivation/Proline Rich
DomainDNA binding domain
Tetramerizationdomain
C-Terminaldomain
P P P
Ac Ac
AcADP ADP ADP
S149 S150 S155 E258 D259 E271 K292
Ac Ac Ac
K120 K164
Ub Ub
UbUb
N8 N8
UbP P P P
K305 S315 K319 K320 K321 K351 K357 S376 S378 S392
P P P P P PP P P
S6 S9 S15 T18 S20 S33 S36 S37 S46 T81
P
P
T55
K370
K372
K373
K386
K382
K381
Me
N8Ac
Ub Me
N8Ac
Ub Me
Ac
Ub
Ac
Ub
N8Ac
Ub
Ac
Ub
S
O-Glc
12
13
to the phenotype of the p53 knock out mice (Parant et al., 2001). Another example of an
inhibitor protein is Parc that binds to p53 and relocates nuclear p53 to the cytoplasm.
Ablation of Parc using RNAi relocalizes p53 to the nucleus and induces cells to undergo
apoptosis (Nikolaev et al., 2003). Protein cofactors play an important role in regulating
p53 stability and function; the exact mechanism of when these protein cofactors come
into play remains a mystery.
Regulation of p53 through post-translational modifications is vitally important,
and extremely complex. Modifications on p53 can have both inhibitory and activating
effects. Acetylation of p53 occurs usually on C-terminal lysines and has been shown to
be an activating modification. Acetylation of p53 promotes apoptosis and cell cycle
arrest, and a mutant p53 that has lost all eight acetylation sites has lost the ability to
induce apoptosis and growth arrest (Tang et al., 2008). p53 can be acetylated on multiple
lysines throughout the protein, but most are located in the C-terminal regulatory region.
Lysines 120, 164, 370, 372, 373, 381, 382, 386 can all be acetylated by several proteins
including Tip60/hMOF, p300/CBP and PCAF; these lysines can also be ubiquitinated,
and it is thought that acetylation may act to block ubiquitin mediated proteolysis of p53
(Li et al., 2002b; Nakamura et al., 2000; Tang et al., 2006; Tang et al., 2008).
Acetylation can be reversed by HDAC or hSIRT1, resulting in deactivation and
destabilization of the p53 protein (Ito et al., 2002; Vaziri et al., 2001). Methylation can
occur on multiple residues and can have both an inhibitory and activating effect.
Methylation of lysine 372 by Set9 can activate p53-dependent apoptosis (Chuikov et al.,
2004). p53 activity can also be inhibited by methylation of lysine 370 by SMYD2, which
causes p53 destabilization. This methylation specific inhibitory effect can be reversed
14
through lysine demethylation by LSD1 (Huang et al., 2007; Toledo and Wahl, 2006).
Sumoylation competes with acetylation and ubiquitination at lysine 386 of p53; the
addition of SUMO-1 to lysine 386 of p53 induces transcriptional activation (Rodriguez et
al., 1999). p53 transcriptional activity can also be inhibited through Mdm2-mediated
neddylation (Xirodimas et al., 2004). Ubiquitination of p53 is complex, but can have
both activating and inhibitory functions. Monoubiquitination of cytoplasmic p53 targets
the protein to the mitochondria where it can be deubiquitinated and induce apoptosis
(Marchenko et al., 2007). However, the p53 protein has a very short half-life, and under
unstressed conditions, p53 stability is regulated primarily through ubiquitin-mediated
proteolysis (Brooks and Gu, 2003).
Multiple kinases target p53 for phosphorylation on several serines and threonines
in the N-terminal transactivation domain, central DNA binding domain and C-terminal
regulatory domain. Phosphorylation of p53 is thought to be an activating modification.
Phosphorylation of serine 37 after DNA damage promotes the transcriptional activity
since mutation of this residue to alanine reduces p53-dependent transcription; in addition,
phosphorylation of serine 37 can be reversed by protein phosphatase 2A (Dohoney et al.,
2004). Also, phosphorylation of p53 on serine 46 has been shown to induce expression
of the pro-apoptotic protein p53-AIP (Oda et al., 2000b).
To determine the role of phosphorylation on p53 activity, knock-in mice have
been created. Phosphorylation of serine 20 after DNA damage by Chk2 kinase is thought
to be an important activating modification (Caspari, 2000). A mouse that has mutated
murine serine 23 to alanine (human serine 20) is defective for apoptosis in B-cells and
develops B-cell lymphomas after a long latency period relative to p53 knockout mice
15
(MacPherson et al., 2004). These results suggest that phosphorylation of a single residue
on p53 may not be sufficient to activate the protein; abrogation of a single
phosphorylation site so far has not been able to recapitulate loss of the p53 protein.
Multiple mutations to delete phosphorylation sites or sites of other post-translational
modifications are probably required to completely mimic loss of function of the p53
protein.
The multiple modifications on several residues of p53 has led researchers to
hypothesize a ‘p53 code’ similar to the histone code (Toledo and Wahl, 2006). Perhaps
these multiple modifications are cell type and stress specific. For example, cells treated
with cisplatin, a DNA damaging agent, and taxol, a microtubule inhibitor, show
differences in phosphorylation at serine 15 and serine 20 (Damia et al., 2001). The
HCT116 p53+/+ colon carcinoma cell line shows differences in phosphorylation and
acetylation in response to various stresses such as ultraviolet radiation, ionizing radiation,
adriamycin, taxol and nocodozole (Saito et al., 2003). This set of experiments performed
in a single cell type suggests that p53 becomes modified in a stress-specific manner.
Specific modifications may occur sequentially; threonine 18 phosphorylation by casein
kinase 1 delta appears to be dependent on phopshorylation of serine 15 (Dumaz et al.,
1999). Regulation of post-translational modifications and the functional consequences of
the post-translational modifications remains a complex problem. Several key questions
include which modifications may be sequential, the prior modifications and how they
contribute to sequential modification, and whether all modifications, or just a subset of
modifications occur on p53 at one time. As well, questions about how cell or tissue type
and stress type influences the modification of the p53 protein are also questions that have
16
yet to be determined. Finally, the question about how these modifications affect p53
activity also needs to be addressed.
Phosphorylation of p53 at serine 15
Phosphorylation of serine 15 is one of the earliest and best characterized
modifications on p53 and is thought to be an activating modification. Serine 15 is
phosphorylated by multiple kinases including ATM, DNA-PK, ATR, and AMPK under
stress conditions such as DNA damage and glucose deprivation (Araki et al., 1999; Banin
et al., 1998; Canman et al., 1998; Toledo and Wahl, 2006). Mouse and stem cell models
for serine 18 (human serine 15) have been made where serine has been mutated to alanine
to prevent phosphorylation. In differentiated murine embryonic stem cells expressing a
knocked-in p53 S18A allele, induction of p53 by both ionizing radiation (IR) and
ultraviolet radiation (UV) were partially impaired, and impaired induction of p21 and cell
cycle arrest in response to IR was also observed (Chao et al., 2000a). Thymocytes in the
p53S18A mice show a reduction in the amount of apoptotic cells when compared to their
wild-type counterparts. In addition, target gene activation in response to IR and UV in
murine embryonic fibroblasts was reduced. The p53 S18A mice, however, were not
prone to spontaneous tumours. Whether these mice develop tumours when challenged
with DNA damaging agents remains undetermined (Chao et al., 2003). The observations
in the mouse model are supported by transfection experiments of human p53S15A into
H1299 cells that show reduced levels of apoptosis compared to the wild-type (Unger et
al., 1999b). These results suggest that phosphorylation of p53 on serine 15 contributes to
17
activation of p53 activity, but that this activation step alone cannot sufficiently explain
the full effects of p53 tumour suppression and activity.
In addition, mice where both the murine serine 18 and serine 23 (human serine 15
and serine 20) sites were mutated exhibit little apoptosis in response to IR in the thymus
comparable to the levels observed in thymocytes from p53-/- mice, and significantly
decreased levels of apoptosis compared to the single S18A mutant thymocytes. p53-
induced neuronal apoptosis is also lost in the S18A/S23A double mutant compared with
wild-type cells. The S18A/S23A mice are also prone to spontaneous tumours, but tumour
formation is significantly slower than that observed in p53 -/- mice (Chao et al., 2006).
The combined effect of phosphorylation of serine 18 and serine 23 suggests that multiple
modifications most likely contribute to activation of p53.
Ubiquitination
Ubiquitin
Ubiquitin is a highly conserved 76 amino acid tag that was discovered in 1978
using rabbit reticulocyte lysates as a signal to mediate proteolysis (Hershko, 1996).
Ubiquitin acts as a tag for protein to signal degradation, subcellular localization,
membrane trafficking, DNA repair and chromatin dynamics (Murata et al., 2009).
Ubiquitin has seven lysines at position 6, 11, 27, 29, 33, 48 and 63. Ubiquitin forms
polymers by forming covalent linkages through any of these lysines (Xu et al., 2009).
Polyubiquitin linkages through lysine 48 and lysine 63 are the most well-characterized.
Polyubiquitinated substrates with a lysine 48 linkages tend to be rapidly degraded.
However, proteins that are polyubiquitinated with lysine 63 linkages are not degraded and
18
tend to be involved in signal transduction pathways (Kirkpatrick et al., 2005; Newton et
al., 2008). The functional consequences of the other ubiquitin linkages remains unclear,
and present research is being conducted to determine the how those lysine linkages affect
the protein substrates.
When ubiquitin is attached to the substrate, it can be attached in multiple forms
(Figure 1.2). Monoubiquitination occurs when a single ubiquitin monomer is attached to
a single lysine on the substrate. A multiubiquitinated substrate has multiple ubiquitin
monomers attached through lysines on the substrate. Polybiquitination of a substrate
indicates that a polymer of ubiquitin is attached to the substrate at one or more lysine
residues (Mukhopadhyay and Riezman, 2007).
The ubiquitination mechanism
The ubiquitination reaction forms a bond between the C-terminus of ubiquitin
(G76) and the ε-amino group of a substrate lysine residue either on ubiquitin or the
substrate to be ubiquitinated. The ubiquitination reaction requires at least three enzymes:
an E1 ubiquitin activating enzyme, an E2 ubiquitin conjugating enzyme, an E3 ubiquitin
ligase; an E4 ubiquitin chain assembly factor may be required if the E3 ubiquitin ligase is
unable to polyubiquitinate the protein (Koegl et al., 1999; Pickart, 2001).
During the ubiquitination reaction, the E1 activating enzyme forms a thiol ester
with the carboxyl group of G76 to activate the C-terminus of ubiquitin for nucleophilic
attack. The E2 conjugating enzyme then transiently carries the activated ubiquitin
molecule as a thiol ester, and the E3 ubiquitin ligase mediates transfer of ubiquitin to the
substrate. All ubiquitination reactions follow the same mechanism regardless of the fate
SubstrateUb
Monoubiquitination
Multiubiquitiantion
Polyubiquitination
Figure 1.2 Ubiquitin Linkage Forms.
Ubiquitin can be conjugated to a substrate in many forms. Monoubiquitinationinvolves a single ubiquitin monomer conjugated to the substrate (A). Multiubiquitination involves several ubiquitin monomers conjugated to the substrate (B). Polyubiquitination involves conjugation of a ubiquitin polymer to the substrate (C).
A
B
C
SubstrateUb
UbUbUb
SubstrateUb
Ub
Ub
Ub
Ub
19
20
of the substrate (Pickart, 2001; Pickart, 2004). The E4 chain assembly factor is important
for catalyzing polyubiquitination. In some specific cases, the E3 enzyme can only
catalyze monoubiquitination of the substrate, and association of the E4 enzyme with the
E3 ligase will promote polyubiquitination (Figure 1.3) (Hoppe, 2005; Koegl et al., 1999).
The circumstances that define when an E4 enzyme is required for ubiquitination are
unclear, and further studies will determine when and how the E4 chain assembly factor
functions.
The E1 ubiquitin activating enzyme activates ubiquitin for all the downstream
conjugating enzymes; there are two human E1 enzymes. The activation occurs when
magnesium-ATP binds ubiquitin to form a ubiquitin adenylate intermediate that serves as
the donor of ubiquitin to a cysteine in the E1 active site. Each fully loaded E1 molecule
carries two molecules of activated ubiquitin: one molecule is conjugated to the E1 and the
other ubiquitin is as an adenylate. The thiol-linked ubiquitin is transferred to the next
conjugating cascade, the E2 (Pickart, 2001).
There are approximately fifty ubiquitin conjugating enzymes, E2s, in humans
(Pickart, 2004). E2 enzymes bind tightly to loaded E1 molecules, but weakly to free
ubiquitin and free E1; this ensures that the ubiquitination reaction is efficiently carried
out. The E2 is also responsible for determining the type of ubiquitin linkage (lysine 6,
lysine 11, lysine 27, lysine 29, lysine 33, lysine 48, lysine 63) that is formed (Chen and
Sun, 2009). The E1 enzyme active site is responsible for transferring ubiquitin from the
E1 to the E2. Each ubiquitin conjugating enzyme can bind to a subset of E3 ubiquitin
ligases. Once the E2 is bound to an E3, the ubiquitin ligase catalyzes transfer of the
ubiquitin from the E2 to either the E3 or the substrate (Pickart, 2001).
E1
E3
E4
Poly-ubiquitination
Substrate
Degradation
Ub-activating enzyme
Ub-conjugating enzyme
Ub-ligase
Ub-chain-assembly
factor
E2
Ub
E2
E3
ATP
AMP + PPi
Ub
Ub
Ub
Ub
Ub
UbUb
UbUb
Figure 1.3 Ubiquitination Reaction Mechanism
ATP is required to activate ubiquitin that binds the E1 ubiquitin activating enzyme. The E1 transfers the E2 to the ubiquitin conjugating enzyme. The E2 binds to the E3 ubiquitin ligase that binds to the substrate to mediate polyubiquitinatin of the substrate. Occasionally, an E3 ubiquitin ligase may require an E4 ubiquitin chain assembly factor to help mediate polyubiquitination, the signal for degradation.
21
22
The E3 ubiquitin ligase confers substrate specificity. In the human system there
are about 600 E3 ubiquitin ligases. The E3 binds to the substrate via the degron – the
ubiquitination signal, conferring substrate specificity, and the E3 also catalyzes the
ligation of one or more ubiquitins to the subtrate (Pickart, 2004). There are two classes
of ubiquitin ligases based on structure: homologous to E6-associated protein carboxy
terminus (HECT) domain containing ligases and really interesting new gene (RING)
domain containing ubiquitin ligases (Pickart, 2001).
The first HECT domain E3 ligase was discovered in cells infected with human
papilloma virus (HPV). The E6 protein promotes degradation of p53 through ubiquitin
mediated proteolysis. The E6 protein of HPV binds to a cellular protein called the E6-
associated protein (E6-AP) to form a complex that ubiquitinates and degrades p53. The
C-terminus of the E6-AP contains the HECT domain for which this class of enzyme is
named. HECT domain E3 ligases are modular: the N-terminus of the protein is
responsible for conferring substrate specificity through binding, and the C-terminus
contains the HECT domain that binds the E2 to mediate the ubiquitination reaction.
HECT domains are not interchangeable, and the HECT domain will covalently bind
ubiquitin before transferring it to the substrate (Pickart, 2001).
The RING finger ubiquitin ligases contain the RING domain that consist of a
series of histidine and cysteine residues with specific spacing that coordinates two zinc
ions in a cross-brace structure with the consensus sequence CX2CX(9-39)CX(1-3)HX(2-
3)C/HX2CX(4-48)CX2C. The zinc ions and ligands are catalytically inert, and the
spacing of the zinc ligands is conserved in the RING finger family which allows the
domain to function as a scaffold for protein-protein interaction (Zheng et al., 2000). The
23
RING finger domain is responsible for binding the E2 conjugating enzyme, and the
ubiquitin is transferred directly from the E2 to the substrate; RING finger ubiquitin
ligases serve to bring the E2 and the substrate into proximity. This class of E3 ubiquitin
ligases can further be subdivided into two classes: single subunit and multi-subunit
ligases (Pickart, 2001).
The multi-subunit ligases compartmentalize the RING finger domain and the
substrate recognition domain. For example, the SCF (Skp1-Cul-F-box) E3 ligase
complex contains Cullin1 (Cul1) as a scaffold protein. The Cullin family of proteins
form scaffolds for the multi-subunit ubiquitin ligases. Cul1 interacts with Rbx1 which
contains a RING finger to bind the E2 conjugating enzyme. Cul1 also interacts with
Skp1 to mediate interaction with the F-box protein that confers substrate specificity. The
multi-subunit ubiquitin ligases have all the components of a single unit ubiquitin ligase
broken into several proteins (Pickart, 2001). Another example of a multi-subunit is the
cyclosome/anaphase promoting complex (APC) that promotes degradation of mitotic
cyclins, some anaphase inhibitors and spindle-associated proteins, which are all degraded
at the end of mitosis and also degrades β-catenin (Hershko and Ciechanover, 1998).
The single-subunit RING finger ubiquitin ligases also interact with the E2 through
the RING finger. The same protein mediates substrate specificity with another part of the
protein through other motifs. RING E3 ubiquitin ligases may also mediate
autoubiquitination resulting in self degradation (Pickart, 2001). Examples of single
subunit RING E3 ligases include cIAP1 that ubiquitinates caspase-3 and caspase-7 and
Cbl that targets receptor tyrosine kinases such as EGFR (Jackson et al., 2000).
24
Ubiquitination and signalling
Although ubiquitin was initially discovered as a tag for proteolytic degradation,
current findings indicate that ubiquitin also plays an important role in non-proteolytic
signalling. In NF-κB signalling, non-proteolytic ubiquitination plays a critical role in
activating the pathway. To activate transcription of downstream targets of NF-κB, the
IκB complex must be phosphorylated by IKK and then degraded through ubiquitin-
mediated proteolysis. There are several mechanisms in which non-proteolytic ubiquitin
plays a role in activation of NF-κB. In one example, TRAF6 has ubiquitin ligase activity,
and catalyzes lysine 63-linked polyubiquitination of IRAK1 and itself. The lysine 63
polyubiquitin chains on TRAF6 bind and activate the TAK1 complex. IRAK1 and
TRAF6 that are both polyubiquitinated through lysine 63 linkages, associate with the IL1
receptor complex. The NEMO/IKK complex binds to the lysine 63-linked polyubiquitin
chains on IRAK1 through the NEMO ubiquitin binding domain. NEMO/IKK binding to
polyubiquitinated IRAK1 brings the IKK complex into proximity with the activated
TAK1 complex that phosphorylates IKK targeting it for degradation (Figure 1.4) (Chen
and Sun, 2009). Based on the upstream receptor and cell type, the substrates that are
ubiquitinated and the ligases that catalyze ubiquitination in the NF-κB pathway may vary
(RIP1, NEMO and MALT1 can also be lysine 63 polyubiquitinated based on the
upstream signalling pathway); however, it remains clear that ubiquitination is an integral
part to the regulation of this pathway (Chen and Sun, 2009).
Ubiquitination represents an important component of chromatin biology and gene
expression. Monoubiquitinated histone H2B resides at the promoter of highly expressed
genes, including the p21 promoter after p53 activation (Minsky et al., 2008). In addition,
IKKβ IKK
TRAF6
IRAK1MyD88
Figure 1.4 Ubiquitination in the NF-κB pathway.
Stimulation of the IL-1 receptor (IL-1R) leads to recuitment of adaptor proteins MyD88,IRAK1 and TRAF6 to the receptor. TRAF6 funcions as an E3 ubiquitinligase that catalyzes the synthesis of K63 polyubiquitin chains with the help of Ubc13/Uev1A E2 complex. TAB2 subunit of the TAK1 kinase complex binds the K63 polyubiquitin chains; this results in activation of TAK1 kinase the phosphorylates IKKβ to activate the kinase. IKKβ is recuited to the receptor complex by binding NEMO; NEMO binds the polyubiquitin chains on TRAF6 or IRAK1. IKKβ activation phosphorylates the inhibitor IκB leading to its degradation and NF-κB activation. (adapted from Chen and Sun, 2009)
Ubc13
Uev1A
Ub
Ub
Ub
Ub
Ub
Ub Ub Ub Ub
NEMO
Ub
TA
B2
TA
K1
PP
p65
p50
IκB p65
p50
IκB
P
P
Degradation
p65
p50
NF-κB target genes
IL-1R25
26
at sites of double strand breaks, chromosomes are ubiquitinated by RNF-8. RNF168 aids
in RNF-8 mediated lysine 63 ubiquitination of H2A and H2AX. Lysine 63 polyubiquitin
chains at sites of double strand breaks are important for retaining 53BP1 and BRCA1,
factors that are required for double strand break repair (Doil et al., 2009). In addition,
ubiquitination also plays an important role in DNA damage signalling.
Ubiquitination on p53 is also important for non-proteolytic regulation. E4F1 is an
E3 ubiquitin ligase that ubiquitinates lysine 320 of p53. E4F1 catalyzes lysine 48-linked
polyubiquitination on p53 that leads to increased binding to chromatin and activation of
p53-dependent cell cycle arrest (Le Cam et al., 2006).
Ubiquitination and protein degradation
The 26S proteasome is responsible for degrading ubiquitinated substrates. The
26S proteasome is a 2.5 MDa complex that is made up of two components: a 20S core
particle and the 19S regulator particle (Hanna and Finley, 2007). The 20S core particle
is a barrel-shaped complex that consists of seven proteins. The peptidolytic activity of
the proteasome is found in the center of the core particle barrel (Groll et al., 1997). There
are three distinct peptidolytic activities in the core particle and each are represented twice;
the enzymes found in the core particle are a tryptic activity, a chymotryptic activity and a
post-acidic activity (Kisselev et al., 2006).
The 19S is the regulatory subunit and is a 1MDa complex. It can be further
subdivided into two subcomplexes the base and the lid. The base aids in substrate
unfolding and is located proximal to the core particle; the unfolding of the protein is
required since the barrel of the 20S proteasome core particle is too small to degrade
27
folded proteins (Hanna and Finley, 2007). The base of the subunit contains ATPases that
have chaperone activity and are thought to be required for unfolding of the protein prior
to degradation (Braun et al., 1999). The lid of the regulatory subunit is involved in
substrate recognition. The regulatory complex contains proteins such as Dsk2 and Rad23
that are responsible for binding to lysine 48 linked ubiquitin and targeting them for
unfolding and subsequently proteolysis by the core particle (Lam et al., 2002; Rao and
Sastry, 2002).
Ubiquitin-mediated proteolysis regulates diverse cellular pathways including cell
cycle progression and NF-B activation. The mammalian G1 regulators cyclin E and
cyclin D1 are targeted for ubiquitination by phosphorylation at specific single sites, and
must be degraded to allow progression into S-phase. In the NF-κB pathway, the IκB
inhibitor is degraded by ubiquitin-mediated proteolysis and this is necessary for NF-B
activation (Hershko and Ciechanover, 1998).
Experiments with a lysine 48-linkage specific antibody indicates that lysine 48-
linked ubiquitin is associated with the 20S proteosome consistent with the hypothesis that
lysine 48-linked ubiquitin usually targets proteins for proteasomal degradation (Newton
et al., 2008). Experiments in yeast, show that lysine 48-linked ubiquitin is stabilized in
the presence of proteasomal inhibition which also supports the hypothesis that lysine 48-
linked polyubiquitination target proteins for degradation (Xu et al., 2009). As well, a
yeast strain that has expresses ubiquitin that has substituted arginine for lysine at position
48 is lethal (K48R), suggesting that lysine 48-linked ubiquitin-mediated proteolysis is
crucial for survival (Spence et al., 1995).
28
Recent experiments have also implicated lysine 11 linked ubiquitin chains in
degradation. In yeast, inhibition of the proteasome results in a four-fold accumulation of
lysine 11 linked polyubiquitin chains; these polyubiquitin chains are implicated in
endoplasmic reticulum-associated degradation (ERAD) as deletion of Doa10, an E3
ubiquitin ligase involved in ERAD, also decreases lysine 11 polybiquitination. In
addition, a yeast strain that expresses ubiquitin with lysine 11 mutated to arginine has
increased sensitivity to endoplasmic reticulum stress-inducing reagents such as
dithiothreiotol and tunicamycin (inhibits the synthesis of N-linked glycoproteins in the
endoplasmic reticulum). These experiments suggest that lysine 11 linked ubiquitin plays
an important role in ERAD (Xu et al., 2009).
The previous observations suggest that although lysine 48 linked ubiquitin chains
are critical for survival and substrate degradation, other ubiquitin linkages are also
important for degradation. In the previously described example, it could suggest that
different linkages may be formed in response to different stresses for targeting to the
proteasome.
Deubiquitination
Deubiquitination is performed by deubiquitinating enzymes, ubiquitin specific
proteases that can reverse target-protein ubiquitination by editing or diassembling
polyubiquitin chains. Deubiquitinasese specifically cleave ubiquitin from ubiquitin-
conjugated protein substrates, ubiquitin precursors, ubiquitin adducts and polyubiquitin.
Deubiquitintases have been implicated in several diseases including cancer and
neurodegenerative disorders (Ventii and Wilkinson, 2008).
29
There are five families of deubiquitinases: four are cysteine proteases and the fifth
is a family of metalloproteases. The four classes of cysteine proteases are subdivided
based on their ubiquitin protease domains: ubiqutin-specific proteases (USP), ubiquitin
C-terminal hydrolases (UCH), otubain proteases (OTU), and Machado-Joseph disease
proteases (MJD) (Ventii and Wilkinson, 2008).
Deubiquitinases are involved in several pathways. USP1 is a deubiquitinase
implicated in Fanconi-anemia related cancer; USP1 deubiquitinates monoubiquitinated
FANCD2 after cells exit S-phase or re-enter the cell cycle after DNA damage (Nijman et
al., 2005). In addition, the deubiquitinase USP11 interacts with BRCA2 and promotes
deubiquitination of BRCA2; downregulation of USP11 through RNAi increases cellular
sensitivity to DNA damaging agent mitomycin C in a BRCA-2 dependent manner
(Schoenfeld et al., 2004).
Deubiquitination plays an important role in regulating the NF-κB pathway.
CYLD mediates negative regulation of the pathway by specifically deubiquitinating
lysine 63 polyubiquitinated proteins TRAF2, TRAF6 and NEMO (Kovalenko et al., 2003;
Trompouki et al., 2003). A special case of a deubiquitinating enzyme is A20, a ubiquitin
editing enzyme that downregulates NF-κB signalling. A20 functions as a ubiquitin
editing enzyme on RIP1, an important mediator of TNFR1 induced NF-κB activation.
A20 removes ubiquitin from RIP1 that is polyubiquitinated through K63 linkages. In
addition, A20 has the ability to catalyze K48 linkages on RIP1 to target it for proteolysis
(Wertz et al., 2004).
At the proteasome, ubiquitin is recycled and only the substrate is degraded. The
26S proteasome is associated with deubiquitinating enzymes to facilitate ubiquitin
30
recycling (Yao and Cohen, 2002). The deubiquitinases Rpn11/S13 and Uch37 are stably
associated with the proteasome and USP14/Ubp6 is transiently associated with the
proteasome. RNAi experiments suggest that Rpn11 and either Uch37 or Ubp6 are
required for proper substrate processing (Koulich et al., 2008). Interestingly, deletion of
Ubp6 in yeast results in increased degradation of ubiquitinated cyclin B relative to the
wild-type yeast strain. Ubp6 is proposed to function through progressive
deubiquitination in which deubiquitination occurs sequentially from the N-terminus to the
C-terminus of the ubiquitin chain one monomer at a time. In the absence of Ubp6, Rpn11
catalyzes a more efficient deubiquitination reaction through en block chain release
(cleavage of the whole ubiquitin chain at the C-terminus in one reaction) (Hanna et al.,
2006).
Deubiquitination enzymes play an important and complex role in many pathways.
The function and regulation of these enzymes remains for the most part to be determined,
but remain vital to the understanding of the function of ubiquitination in the cell.
Ubiquitination and regulation of p53
The p53 protein has a short half-life under unstressed conditions due to regulation
through ubiquitin mediated proteolysis. There are several ubiquitin ligases that regulate
p53 including Mdm2, Pirh2, Cop1, ARF-BP1, TOPORS, Synoviolin, Trim24 CARPs
(Allton et al., 2009; Chen et al., 2005a; Dornan et al., 2004b; Haupt et al., 1997; Honda et
al., 1997; Leng et al., 2003; Rajendra et al., 2004; Yamasaki et al., 2007; Yang et al.,
2007). The physiological role of these ubiquitin ligases in the cell, and in regulating p53
function is unclear at the moment, and further studies need to be performed.
31
ARF-BP1 is HECT E3 ubiquitin ligase. ARF-BP1 is inhibited by the tumour
suppressor ARF. ARF-BP1 ubiquitinates p53 and targets it for proteasomal degradation.
Knockdown of ARF-BP1 by RNAi stabilizes p53, increases p21 levels, and increases
p53-dependent apoptosis (Chen et al., 2005a). ARF-BP1 is also known as Mule, and has
also been found to ubiquitinate and degrade the pro-survival protein Mcl-1 (Zhong et al.,
2005). The physiological function of ARF-BP1/Mule is unclear at the present time since
p53 and Mcl-1 have diametrically opposing functions. Further research needs to be
completed to determine the role of ARF-BP1/Mule in regulating p53 and Mcl-1; perhaps
co-factors help redirect the ubiquitin ligase function of ARF-BP1/Mule resulting in a
specific regulation for each substrate.
Cop1 is a RING finger E3 ubiquitin ligase that targets p53 for degradation. Cop1
siRNA enhances p53-dependent G1 arrest and sensitizes cells to ionizing radiation
(Dornan et al., 2004b). Cop1 is also phosphorylated by ATM on serine 387 after DNA
damage, and this phosphorylation site dissociates the p53-Cop1 interaction (Dornan et al.,
2006). Cop1 has also been found to be overexpressed in breast and ovarian
adenocarcinomas (Dornan et al., 2004a).
Mdm2 is the first ubiquitin ligase found to antagonize p53 (Momand et al., 1992;
Oliner et al., 1992). It is also the best characterized. Mdm2 knock out mice are
embryonic lethal prior to implantation, and these mice die by massive apoptosis;
embryonic lethality is fully rescued by deletion of p53 and partially rescued by deletion
of the pro-apoptotic gene bax (Chavez-Reyes et al., 2003; Jones et al., 1995; Montes de
Oca Luna et al., 1995). Mdm2 gene amplification has also been observed in over one
third of human sarcomas that retained wild-type p53, and patients overexpressing Mdm2
32
in sarcomas have a worse prognosis (Oliner et al., 1992; Onel and Cordon-Cardo, 2004).
Mdm2 regulation of p53 has been targeted for chemotherapy. The nutlin group of
inhibitors abrogates the Mdm2/p53 interaction. Treating tumour xenografts in mice with
nutlins causes cell cycle arrest and apoptosis suggesting that nutlins could eventually be
used to treat tumours that use Mdm2 to inactivate p53 (Vassilev et al., 2004).
Mdm2 regulates p53 stability through ubiquitination and this ability to regulate
p53 function is dependent on the RING finger (Haupt et al., 1997; Honda et al., 1997;
Honda and Yasuda, 2000; Kubbutat et al., 1997). Mdm2 binds to p53 in both of their N-
terminal hydrophobic domains, the interaction is mediated by the hydrophobic residues
phenylalanine 19, tryptophan 23 and leucine 26 on p53 that insert into the hydrophobic
cleft on Mdm2 (Kussie et al., 1996). Binding of p53 to Mdm2 promotes ubiquitinatation
of p53 at six lysine residues in the C-terminus (370, 372, 373, 381, 382, 386) (Kubbutat
et al., 1998; Nakamura et al., 2000; Rodriguez et al., 2000). To catalyze
polyubiquitination of p53, Mdm2 requires an E4 chain elongation factor. To date, there
are two chain elongation factors that have been identified for Mdm2-mediated p53
polyubiquitination the first is histone acetylase p300, and the second is the transcription
factor Yin Yang 1 (Gronroos et al., 2004; Grossman et al., 2003; Sui et al., 2004). Yin
Yang 1 can form a ternary complex with p53 and Mdm2, and ablation of Yin Yang 1 by
siRNA results in increased levels of p53 and p21 and an inhibition of cellular
proliferation (Sui et al., 2004).
Mdm2 dissociates from p53 after DNA damage, and this presumably contributes
to stabilization of the p53 protein (Shieh et al., 1997). There are several mechanisms that
could account for this dissociation including Chk2-mediated phosphorylation of p53 on
33
serine 20 that is proposed to disrupt the binding of Mdm2 to p53 (Chehab et al., 1999;
Unger et al., 1999a).
Regulation of Mdm2 occurs at the transcriptional and post-translational level. At
the transcriptional level, the Mdm2 gene has two promoters: the P1 promoter and the P2
promoter. The P1 promoter is required for maintaining basal levels of Mdm2. The P2
promoter is located in intron 1 and contains a p53 binding site and it is where p53 induces
Mdm2 expression (Iwakuma and Lozano, 2003; Wu et al., 1993). Mdm2 can also be
regulated transcriptionally by a naturally occurring polymorphism in the P1 Mdm2
promoter. SNP 309 contains either thymidine or guanosine bases in the DNA, the
guanosine polymorphism creates a higher affinity binding site for the transcriptional
activator Sp1 and results in increased Mdm2 mRNA and protein at the basal level.
Individuals with the guanosine polymorphism exhibit accelerated sporadic and hereditary
tumourigenesis (Bond et al., 2004).
In addition, Mdm2 also promotes auto-ubiquitination resulting in auto-inhibition
through proteasomal degradation (Honda and Yasuda, 2000). Mdm2 auto-ubiquitination
is antagonized by HAUSP that deubiquitinates Mdm2 and stabilizes it. Since HAUSP
stabilizes Mdm2, p53 becomes destabilized (Cummins et al., 2004; Li et al., 2004).
HAUSP has also been shown to deubiquitinate p53 and MdmX (Li et al., 2002a;
Meulmeester et al., 2005). In addition, p53 and Mdm2 compete for the same binding
pocket on HAUSP, and Mdm2 seems to have a stronger interaction with HAUSP than
p53 (Sheng et al., 2006). How HAUSP functions to regulate p53 levels is unclear at the
moment, perhaps there could be additional cofactors that modulate HAUSP activity either
promoting or inhibiting deubiquitination of a particular substrate.
34
There are also several proteins that can bind to Mdm2 to modulate its activity.
Upon oncogenic stress, ARF binds to Mdm2, and relocates it to the nucleolus so that it is
unable to degrade p53 (de Stanchina et al., 1998; Honda and Yasuda, 1999; Palmero et al.,
1998; Pomerantz et al., 1998). Additional proteins such as PML, and nucleolar protein
NPM, L23, and L11 also bind to Mdm2 and sequester it to prevent Mdm2-mediated
degradation of p53 (Bernardi et al., 2004; Dai et al., 2004; Jin et al., 2004; Kurki et al.,
2004; Zhang et al., 2003).
Regulation of Mdm2 also occurs through post-translational modifications. ATM-
mediated phosphorylation of Mdm2 on serine 395 resulted in a decreased ability of
Mdm2 to degrade p53; Wip1 phosphatase dephosphorylates serine 395 on Mdm2 to
increase the degradation of p53 by Mdm2 (Khosravi et al., 1999; Lu et al., 2007; Maya et
al., 2001). Mdm2 is also phosphorylated by Akt/PKB on serines 166 and 188, and this
modification stabilizes Mdm2. Knockout of Akt/PKB in murine embryonic fibroblasts
increases p53 and p21 levels (Feng et al., 2004). Mdm2 is also phosphorylated by c-Abl
after DNA damage on tyrosines 276 and 294; these phosphorylations inhibit Mdm2
degradation of p53 by promoting association with the inhibitor ARF (Dias et al., 2006).
Regulation of p53 stability is complex, and further experiments are required to
determine not only how p53 is regulated, but also how different stress conditions affect
the proteins that regulate p53. Using Mdm2 as an example, several factors such as
protein cofactors, and post-translational modifications influence Mdm2 activity, and as a
result affect p53 activity. As well, there is increasing complexity for the regulation of
p53 as several ubiquitin ligases also target p53 for degradation, this high degree of
functional redundancy must be addressed through additional experiments and animal
35
models to determine which are the critical ubiquitin ligases, and whether each of these
ligases may regulate p53 under different conditions or in different cell types.
Pirh2
Pirh2 was initially identified as a protein that interacts with the androgen receptor,
and is also known as ARNIP (androgen receptor N-terminal interacting protein) that
functions as a ubiquitin ligase (Beitel et al., 2002). The Benchimol lab identified Pirh2 in
DP16.1 p53ts cells. The DP16 cells are murine erythroleukemia cells transformed by
Friend cell virus, and lack endogenous p53 expression; the DP16.1 p53ts cells are stably
transfected with a mutant p53 allele that encodes valine instead of alanine at codon 135.
The p53 A135V mutant is temperature sensitive (ts) and results in a protein that adopts
the mutant conformation and is inactive at 37ºC and adopts the wild-type conformation
and is active at 32ºC (Johnson et al., 1993; Michalovitz et al., 1990). A differential
display experiment was performed in the DP16.1 p53ts cells to identify novel genes that
are regulated by p53 by comparing DP16.1 p53ts cells grown at 32ºC, with active p53, to
those grown at 37ºC, with mutant p53. One of the targets identified was Pirh2, a gene
that encodes a 261 amino acid, 30 kDa protein with a cysteine-rich RING motif (Leng et
al., 2003).
The Pirh2 gene is expressed in a wide variety of tissues including testis, kidney,
lung, and brain with highest expression observed in both heart and liver. Pirh2 is induced
by DNA damage in a p53-dependent manner in both human and murine cells, though p53
induction appears to be cell-type and stress specific (Feng et al., 2007). Chromatin
immunoprecipiation, luciferase assays and EMSA (electrophoretic mobility shift assay)
36
experiments indicate that p53 binds to intron 3 of Pirh2 to induce its expression.
Antisense experiments also indicate that knockdown of Pirh2 causes an increase in p53
protein, and results in a modest increase in G1 arrest (Leng et al., 2003). Further
experiments have also suggested that Pirh2 can only ubiquitinate the tetrameric form of
p53, and that Pirh2 can also undergo auto-ubiquitination (Sheng et al., 2008). Pirh2
regulation of active p53 suggests that it is a less potent ubiquitin ligase for p53 when
compared to Mdm2.
Pirh2 has also been shown to be overexpressed in both murine and human lung
cancers. Using semi-quantitative analysis of immunoblots, Pirh2 protein expression was
elevated by at least two fold in 84% of tumour samples examined (Duan et al., 2004).
Pirh2 has also been shown to be overexpressed in 89% of prostate cancers, and Pirh2
overexpression was correlated with more aggressive tumours as well (Logan et al., 2006).
Additional targets for Pirh2 have been identified. These targets include HDAC1
(hisone deacetylase 1), the G1 cyclin-dependent kinase inhibitor p27/Kip1, SRβ (signal
recognition particle receptor beta subunit) and ε-COP subunit of the coatomer complex
(Abe et al., 2008; Hattori et al., 2007; Logan et al., 2006; Maruyama et al., 2008). Pirh2
has also been shown to interact with the histone acetyl transferase Tip60, and hNTKL-
BP1 (human N-terminal kinase-like binding protein 1) (Logan et al., 2004; Zhang et al.,
2005). Whether these proteins are substrates, or how they affect Pirh2 activity is unclear
at the present.
Pirh2 stability is enhanced by interaction with the transcription factor PLAG2
(pleomorphic adenoma gene like 2) and MV P (measles virus phosphor protein);
37
interaction with either PLAG2 or MV P stabilizes Pirh2 by inhibiting Pirh2 auto-
ubiquitination (Chen et al., 2005b; Zheng et al., 2007).
Pirh2 can also interact with calmodulin, and is phosphorylated by CaMKII
(calmodulin dependent kinase II) on threonine 154 and serine 155. CaMKII
phosphorylation of Pirh2 decreases Pirh2 stability, and decreases Pirh2 mediated
ubiquitination and degradation of p53 (Duan et al., 2007).
Further studies need to determine the key substrates that are targeted for Pirh2-
mediated degradation, the cofactors that mediate degradation and the physiological role
of Pirh2 in regulating p53 and in the cell.
Thesis Hypothesis
It is known that Pirh2 targets p53 for ubiquitin mediated proteolysis, however,
several other ubiquitin ligases also target p53 for degradation. The reason for such a high
degree of functional redundancy is unclear, and furthermore, the physiological role of
Pirh2 in regulating p53 is also undefined. In addition to regulating p53, it is also well-
known that Pirh2 regulates multiple targets, and it is likely that not all substrates of Pirh2
have been identified, and any protein cofactors that aid in Pirh2-dependent ubiquitin
mediated proteolysis also have yet to be identified.
The purpose of this study is two fold. The first is to identify additional substrates
for Pirh2 and any additional protein cofactors that aid in ubiquitination. The second aim
of this study is to address Pirh2 regulation of p53 function.
38
CHAPTER 2
Identification of Pirh2 Interacting
Proteins
Thank you to Dr. Jack Greenblatt and Dr. Cheryl Arrowsmith who provided the His-Flag
vector used for purification, Stephen Chung for cloning the His-Flag vector, Weili Ma for
generating Pirh2-/- E1A/Ras His-Flag and Pirh2 His-Flag clones and the York Mass
Spectrometry facility performed all Mass Spectrometry analysis.
39
ABSTRACT
Pirh2 was identified as a p53 target gene in a Friend-virus transformed mouse
erythroleukemia cell line (Leng et al., 2003). Pirh2 has also been characterized as having
E3 ubiquitin ligase activity, and can target p53, HDAC1, p27/Kip1, SRβ and ε-COP for
ubiquitin mediated proteolysis. Pirh2 can also bind to the androgen receptor, hNTKL-
BP1, calmodulin, and Tip 60 (Abe et al., 2008; Beitel et al., 2002; Duan et al., 2007;
Hattori et al., 2007; Leng et al., 2003; Logan et al., 2004; Maruyama et al., 2008; Zhang
et al., 2005). Here, I described two approaches to identify additional targets for Pirh2
ubiquitination, and cofactors involved in Pirh2-mediated ubiquitination using tandem
affinity purification and a GST-pull down assay.
INTRODUCTION
Pirh2 is a newly identified gene that is regulated by p53 and can also regulate p53
through ubiquitin mediated proteolysis. Pirh2 has several targets including HDAC1
(hisone deacetylase 1), the G1 cyclin-dependent kinase inhibitor p27/Kip1, SRβ (signal
recognition particle receptor beta subunit) and ε-COP subunit of the coatomer complex
(Abe et al., 2008; Hattori et al., 2007; Logan et al., 2006; Maruyama et al., 2008).
Ubiquitin ligases are known to have several targets for ubiquitination, and require several
cofactors including ubiquitin conjugating enzymes (Deshaies and Joazeiro, 2009). To
identify these interactors, we used two methods: the first is a tandem affinity purification
method and the second a GST-pull down assay.
Tandem affinity purification (TAP) was initially developed in yeast. It involves
fusion of a protein to a protein A and calmodulin binding peptide tag separated by a
40
tobacco etch virus (TEV) protease site. The dual tags allow for sequential purification of
a protein and any associated proteins; the dual tag also provides less background
contaminating proteins and false positives than purification performed with a single tag
(Rigaut et al., 1999). A TAP tagged protein is first purified by binding to IgG beads that
interact with the protein A. The complex is then released by cleavage with TEV protease,
and passed over calmodulin beads that interact with the calmodulin binding peptide. The
complex can then be eluted using EGTA that chelates the calcium ions required to
mediate the calmodulin/calmodulin binding peptide interaction (Puig et al., 2001).
One advantage of the dual tag approach is that it can identify complexes where as
the yeast two hybrid system was designed to identify binary interactions. In comparison,
TAP tagging also purifies proteins under more physiological conditions than the yeast
two hybrid system and can map complexes under different stress conditions that may be
dependent on post-translational modification. In contrast, the yeast two hybrid system
relies on overexpression of proteins and does not account for post-translational
modifications. When TAP tagging was developed in yeast, the tagged gene was knocked
into the genome to replace the normal gene – this strategy is advantageous since it
eliminates competition for interacting proteins from the untagged protein, and the tagged
protein is also regulated by the same promoter as the normal protein so expression levels
would be similar (Puig et al., 2001). In mammalian cells, the gene cannot be easily
knocked into somatic cells, but to express similar levels of the tagged protein to the
endogenous protein, stable transfectants are constructed and clones that express the
protein at similar levels to the endogenous protein are selected (Gingras et al., 2005).
41
Several variations of dual purification tags have emerged. In addition to the
classical TAP-tag, we have used two different variations in an attempt to identify proteins
that interact with Pirh2. The first is the SPA (Sequential Peptide Affinity) tag that
contains a triple flag tag and calmodulin binding peptide separated by the TEV protease
site. The second tag is a triple flag tag and six times histidine separated by the TEV
protease site (Zeghouf et al., 2004). In addition, we also performed a GST-pull down
experiment to identify interacting proteins under normal conditions and conditions of
DNA damage.
EXPERIMENTAL PROCEDURES
Cell Culture. H1299 cells and Pirh2 -/- immortalized kidney fibroblasts were cultured in
-MEM with 10% fetal calf serum (FCS). Pirh2 -/- MEFs were transformed by retroviral
infection with pBABE-E1A/Ras and cultured in Dulbecco’s modified Eagles medium
(DMEM) with 10% FCS. Pirh2 -/- immortalized kidney fibroblasts were prepared by
harvesting the kidney from the mouse. The kidney was separated into cells by passing
through a 70 µm strainer (BD Biosciences). Cells were incubated overnight in -MEM
with 10% FCS. The following morning, cellular debris and media were removed, and
replaced with fresh -MEM with 10% FCS. Adhered cells were washed gently with PBS,
and grown in -MEM with 10% FCS for one week. Cells were then immortalized
through the 3T3 protocol.
Plasmids. Human Pirh2 cDNA was PCR amplified from pcDNA3-hPirh2 (Leng et al.,
2003) and cloned into pCMV-TAP to generate an N-terminal TAP-tagged Pirh2 construct.
42
PCR amplification of hPirh2 was also used to create a 3’ SPA tagged hPirh2 construct
and a 3’ His-Flag tagged hPirh2 construct. PCR amplification of hPirh2-SPA and
hPirh2-His-Flag was used to clone into pBabe neo and pBabe puro respectively.
Generation of Clones Expressing pCMV TAP hPirh2. H1299 cells were plated (4 x
105) in a 10 cm dish the day before transfection. Calcium phosphate was used to transfect
10 µg of pCMV-TAP, 10 µg pCMV-TAP-hPirh2, 10 µg of pcDNA3 SPA or 10 µg
pcDNA3 hPirh2-SPA. H1299 cells were selected with 400 µg/mL neomycin (Invitrogen)
and then transferred to 6-well plates and then screened for clone expression.
Generation of Clones Expressing pBabe neo hPirh2-SPA. 293 cells were plated at a
density of 3 x 105 per 6 cm plate and FuGENE (Roche) was used to transfect 1 µg pCL-
Eco and 1 µg of pBabe neo SPA or pBabe neo hPirh2 SPA. After transfection, media
was removed and replaced with 5 mL of DMEM 10% FCS. After 48 hours, media was
collected and filtered (0.45 µm, Millipore) and used for infection of Pirh2-/- E1A/Ras
MEFs. Pirh2-/- E1A/Ras MEFs were plated at a density of 1 x 105 cells per 10 cm plate.
Cells were selected in 200 µg/mL neomycin for one week. Clones were transferred to 6-
well plates and then screened for clone expression by western blot analysis.
Generation of Clones Expressing pBabe puro hPirh2-HF. 293 cells were plated at a
density of 3 x 105 per 6 cm plate and FuGENE (Roche) was used to transfect 1 µg pCL-
Eco and 1 µg of pBabe puro HF or pBabe puro hPirh2-HF. After transfection, media was
removed and replaced with 5 mL of DMEM 10% FCS. After 48 hours, media was
43
collected and filtered (0.45 µm, Millipore) and used for infection of Pirh2-/- E1A/Ras
MEFs. Pirh2-/- E1A/Ras MEFs were plated at a density of 1 x 105 cells per 10 cm plate.
Puromycin (1 µg/mL, Invivogen) was used to select the cells for two days. Clones were
transferred to 6-well plates and then screened for clone expression by western blot
analysis.
Ubiquitination Assay. H1299 cells, TAP or TAP-hPirh2 clone 11 were plated at a
density of 5 x 105 cells per 10 cm plate the day before transfection. Cells were
transfected using calcium phosphate with 10 µg pcDNA3 or pcDNA3 hPirh2 and 1 µg
HA-ubiquitin vector and harvested 40 hours after transfection for western blot analysis.
Coimmunoprecipitation. H1299 cells, TAP or TAP-hPirh2 clone 5 were plated at a
density of 5 x 105 cells per 10cm plate the day before transfection. Cells were transfected
using calcium phosphate with 1µg pcDNA3 or pcDNA3 p53 and harvested 40 hours after
transfection in IP lysis buffer (50 mM Tris-Cl pH 8.0, 5 mM EDTA, 150 mM sodium
chloride, 0.5% NP-40). 1 mg of protein was loaded on rabbit IgG-agarose beads (GE
Healthcare) and rotated at 4ºC for 2 hours. Beads were washed three times with IP wash
buffer (50 mM Tris-Cl pH 8.0, 5 mM EDTA, 150 mM sodium chloride, 0.1% NP-40).
Beads were boiled in 1x Laemmli SDS buffer and separated by SDS-PAGE for western
analysis.
44
Western blots. Antibodies against hPirh2 (Leng et al., 2003), calmodulin binding
peptide (CBP, Calbiochem), histidine (Qiagen), Flag and β-actin (Sigma) and non-
immune rabbit IgG (Cedarlane) were used for western blots.
TAP Purification. H1299 CMV TAP or H1299 TAP-hPirh2 cells were plated in 20
plates each at a density of 1 x 106 per 15 cm plate. Cells were grown to approximately
90% confluency, and lysed 40 mM HEPES pH 7.5, 120 mM sodium chloride, 1 mM
EDTA, 0.5% NP-40, EDTA-free protease inhibitor tablet (Roche). Lysates were spun at
16,000g. An equal volume and protein concentration of TAP and TAP-hPirh2 lysates
were loaded onto rabbit IgG-agarose beads (200 µL, GE Healthcare) and rotated for 4
hours at 4ºC. IgG beads were washed three times with wash buffer (10 mM Tris pH 8.0,
100 mM sodium chloride, 0.5 mM EDTA, 0.1% NP40) at 4ºC, and once with TEV
cleavage buffer (10 mM Tris pH 8.0, 150 mM sodium chloride, 0.5 mM EDTA, 0.1%
NP40, 1 mM DTT). IgG beads were rotated in 300 µL TEV cleavage buffer with 100 µg
TEV protease at 4ºC overnight. Beads were washed with 200 L TEV cleavage buffer
twice, and the fractions were combined. The eluate was supplemented with 2 L of 2 M
calcium chloride and 400 L calmodulin binding buffer (10 mM Tris pH 8.0, 150 mM
sodium chloride, 1 mM imidazole, 1 mM magnesium acetate, 0.2 mM calcium chloride,
10 mM β-mercaptoethanol, 0.1% NP-40). Calmodulin beads (100 µL, GE Healthcare)
were added to the mixture and rotated at 4ºC for 4 hours. Beads were washed three
times with calmodulin binding buffer and washed once with calmodulin binding buffer
without calcium chloride. To elute, the beads were boiled in 1x Laemmli SDS buffer,
45
and the samples were separated using SDS-PAGE. Silver staining was performed using a
method compatible for mass spectrometry (Shevchenko et al., 1996).
SPA Purification. Pirh2-/- E1A/Ras SPA MEFs or Pirh2-/- E1A/Ras hPirh2-SPA MEFs
were plated into 20 plates each at a density of 3 x 106 per 15 cm plate. Cells were grown
to approximately 90% confluency, and lysed in 40 mM HEPES pH 7.5, 120 mM sodium
chloride, 1 mM EDTA, 0.5% NP-40, EDTA-free protease inhibitor tablet (Roche). The
lysate was cleared by spinning at 16,000 g. An equal concentration and volume of SPA
and hPirh2-SPA lysate was loaded onto 200 µL packed M2-agarose beads (Sigma) and
incubated for 4 hours with rotation at 4ºC. M2 beads were washed three times with wash
buffer (10 mM Tris pH 8.0, 100 mM sodium chloride, 0.5 mM EDTA, 0.1% NP40) and
then washed once with TEV cleavage buffer (10 mM Tris pH 8.0, 150 mM sodium
chloride, 0.5 mM EDTA, 0.1% NP40, 1 mM DTT) at 4ºC. M2 beads were incubated in
300 µL TEV cleavage buffer with 100 µg TEV protease at 4ºC with rotation overnight.
M2 beads were then washed with 200 L TEV cleavage buffer twice and the fractions
were combined. The eluate was supplemented with 2 L of 2 M calcium chloride and
400 L of calmodulin binding buffer (10 mM Tris pH 8.0, 150 mM sodium chloride, 1
mM imidazole, 1 mM magnesium acetate, 0.2 mM calcium chloride, 10 mM β-
mercaptoethanol, 0.1% NP-40). Calmodulin beads (100 µL, GE Healthcare) were added
to the mixture and rotated at 4ºC for 4 hours. Beads were washed three times with
calmodulin binding buffer and washed once with calmodulin binding buffer without
calcium chloride. Beads were then washed once with elution buffer without EGTA (50
mM ammonium bicarbonate). The protein mixture was eluted by rotating at 4ºC with
46
500 µL elution buffer (50 mM ammonium bicarbonate, 25 mM EGTA). A second
elution was performed, and the fractions were combined. Samples were lyophilized, and
proteins were identified using LC-MS/MS.
His-Flag Purification. Pirh2-/- E1A/Ras MEFs expressing either His-Flag or Pirh2-His-
Flag were plated each into 80 plates (15 cm) and grown to 90% confluency. Cells were
harvested and resuspended in 40 mL lysis buffer (10 mM Tris pH 8.0, 100 mM sodium
chloride, 5 mM EDTA pH 8.0, 10 mM sodium diphosphate, 10 mM sodium fluoride, 1
mM sodium orthovanadate 0.35% NP-40, 10% glycerol, EDTA-free protease inhibitor
tablet (Roche)). Cell lysates were cleared by centrifugation at 16,000 rpm. A equal
volume and concentration of His-Flag or Pirh2-His-Flag lysate was added to 1 mL of
M2-agarose beads (Sigma), and rotated for 4 hours at 4ºC. Beads were washed three
times with Flag wash buffer (10 mM Tris pH 8.0, 100 mM Sodium chloride, 5 mM
EDTA, pH 8.0, 0.1% NP-40, EDTA-free Protease inhibitor tablet (Roche)) at 4ºC. The
M2-beads were then resuspended in 3mL of Flag-peptide elution buffer (0.1 mg/mL Flag
peptide (Sigma), 20 mM Tris pH 8.0, 100 mM sodium chloride, 0.1 mM EDTA, 5 mM
Imidazole) and rotated overnight at 4ºC. The M2 beads were washed twice with 2 mL
Flag-peptide elution buffer. Eluates were pooled and incubated with 800 µL nickel-
agarose beads (Sigma), and rotated for 2 hours at 4ºC. Nickel-agarose beads were
washed three times with Ni-NTA wash buffer (20 mM Tris pH 8.0, 100 mM sodium
chloride, 0.1 mM EDTA, 5 mM Imidazole). Three elutions were performed with 2 mL
of Ni-NTA elution buffer (10 mM Tris pH 8.0, 100 mM ammonium bicarbonate, 0.1 mM
EDTA, 0.4 M imidazole). The eluted fractions were pooled, and dialyzed overnight at
47
4ºC against 100 mM ammonium bicarbonate. Eluates were frozen at -80ºC, and
lyophilized for identification of the protein mixture using LC-MS/MS.
GST-pull down. Purification of GST-Pirh2 was performed as described (Sheng et al.,
2008). Pirh2-/- kidney fibroblasts were grown to 85% confluency and harvested or
treated with 200 ng/mL of adriamycin (Sigma) for 24 hours. Cells were resuspended in
GST-pull down lysis buffer (50 mM Tris pH 8.0, 5 mM EDTA, 5 mM EGTA, 100 mM
sodium chloride, 0.4% NP-40, 10% glycerol, 10 mM sodium diphosphate, 10 mM
sodium fluoride, 1 mM sodium orthovanadate, and EDTA-free protease inhibitor
(Roche)). 2 mg of Pirh2-/- immortalized kidney lysate either untreated or treated with
adriamycin was incubated with GST-glutathione beads or GST-Pirh2 beads overnight at
4ºC. Beads were washed three times with wash buffer (50 mM Tris pH 8.0, 5 mM EDTA,
5 mM EGTA, 100 mM sodium chloride, 0.1% NP-40, 10 mM sodium diphosphate, 10
mM sodium fluoride, 1 mM sodium orthovanadate, and EDTA-free protease inhibitor
tablet (Roche)). Protein was eluted by boiling in 1x Laemmli SDS buffer. Samples were
run on 4-12% gradient gel (Invitrogen), and stained with Gel Code Blue Stain Reagent
(Pierce). Bands were excised for identification by mass spectrometry (MALDI-MS/MS).
RESULTS
Generation and characterization of TAP-hPirh2 clones.
To identify proteins that interact with Pirh2, we created an N-terminal TAP tag
Pirh2 construct (Fig 2.1A). Stable transfectants were made in H1299 cells and individual
Pirh2
Protein ACBP
TEV Cleavage
___ 24
___ 35
___ 48
___ 54
Actin
TAP Tag
hPirh2
TAP-hPirh2
H1299
pcDNA3
hPirh
2
TAP hPirh
2 CL2
TAP hPirh
2 CL4
TAP hPirh
2 CL7
TAP hPirh
2 CL9
TAP hPirh
2 CL1
1
pCM
VTAP
A
B
WB: hPirh2
Figure 2.1 Generation of TAP-hPirh2 clones in H1299 cells.
(A) A diagram of TAP-Pirh2 .
(B) Western blot of Pirh2 and TAP-Pirh2 expression in H1299 stable clones (CL) and after transfection in H1299 cells. The top panel shows blotting with Pirh2 antibodies and the bottom panel shows blotting with rabbit IgG.
48
H1299
pcDNA3
hPirh
2
TAP hPirh
2 CL2
TAP hPirh
2 CL4
TAP hPirh
2 CL7
TAP hPirh
2 CL9
TAP hPirh
2 CL1
1
Actin
TAP Tag
TAP-hPirh2
___ 24
___ 35
___ 48
___ 54
WB: non-immunerabbit IgG
pCM
VTAP
49
colonies were examined to identify clones that express TAP-Pirh2 at levels comparable to
endogenous Pirh2. Overexpression of TAP-Pirh2 is not desired since this could lead to
non-physiological interactions with the fusion protein. The results of the Pirh2 western
blot are shown in Fig 2.1B. As a result of the protein A tag, TAP-Pirh2 can be detected
either with Pirh2 antibodies or with rabbit IgG (Fig 2.1B). It is difficult to compare TAP-
Pirh2 levels with endogenous Pirh2 levels because the rabbit Pirh2 antibodies will bind to
the protein A tag as well as to the Pirh2 region so the best one can do is to identify clones
that express TAP-Pirh2 but do not overexpress TAP-Pirh2. Five clones were isolated:
two that express TAP-Pirh2 at relatively low levels (clones 2,4), two at more moderate
levels (clones 7, 9) and a single clone that expresses at high levels (clone 11) (Fig. 2.1B).
The TAP tag comprises about half the molecular mass of TAP-hPirh2. We were
concerned that the tag may interfere with protein function, so we performed two assays to
determine function. The first is a ubiquitination assay; H1299 cells stably transfected
with TAP or TAP-hPirh2 were co-transfected with empty vector and HA-ubiquitin (Fig
2.2A). After performing a western blot for HA, and observing the characteristic smear in
the TAP-hPirh2 indicative of ubiquitination activity, we determined that the TAP-hPirh2
fusion protein construct retains the ability to promote ubiquitination of proteins. We used
HA-ubiquitin cotransfected with either empty vector or Pirh2 as negative and positive
controls, respectively. We also tested to determine if TAP-hPirh2 retained the ability to
bind p53. We transfected either empty vector or p53 into parental H1299 cells, TAP or
TAP-hPirh2 cells, and performed immunoprecipitation assays with rabbit IgG and
blotted for p53 (Fig 2.2B). p53 only coimmunoprecipitated with TAP-hPirh2. This
suggests that TAP-hPirh2 retains the ability to bind p53. Based on the ubiquitination and
Vecto
r + H
A-Ub
TAP: V +
HA-Ub
TAP-h
Pirh2:
V +
HA-Ub
Actin
170 __
130 __
100 __
72 __
55 __
hPirh
2 +
HA-Ub
WB: HA
IP: Rabbit IgGWB: p53
Transfected withempty pcDNA3
Transfected with pcDNA3 p53
H1299
TAP
TAP-hPirh
2
p53
A
B
Figure 2.2 Functional Assays for TAP-hPirh2 Activity.
(A) Parental H1299 cells were transiently transfected with empty vector (pcDNA3) or pcDNA3 hPirh2 and HA-ubiquitin. TAP and TAP hPirh2 CL11 were transiently transfected with empty vector and HA ubiquitin. Western blots were performed for HA ubiquitin and actin as a loading control.
(B) H1299, TAP or TAP hPirh2 CL9 cells were transiently transfected with either empty vector or pcDNA3 p53. Lysates were immunoprecipitated with rabbit IgG and western blotted for p53 (B).
50
51
coimmunoprecipitation assays, we conclude that TAP-hPirh2 retains activity and is a
suitable reagent to identify Pirh2 interacting proteins.
Purification of TAP and TAP-hPirh2.
Purification was optimized using TAP and the high expressing TAP-hPirh2 clone
(CL11). Cells were lysed and fractions were collected at each stage in the purification
process: Cell lysate, lysate unbound to IgG, elution after TEV protease, unbound to
calmodulin beads, and bound to calmodulin beads. Calmodulin binding peptide antibody
was used to track TAP-hPirh2 at each step in the purification (Fig 2.3A). We were able
to successfully purify TAP-hPirh2 from high expressing cells, however, we were unable
to detect purification of TAP-hPirh2 from the lower expressing clones (data not shown).
TAP purification was repeated on a preparative scale using CL11, and the fractions were
separated on SDS-PAGE and silver stained to visualize protein bands. Bands were then
excised and sent for identification by mass spectrometry (Fig 2.3B). Many of the bands
were unidentifiable; those that could be identified (Hsc70, alpha-s-casein, tubulin) appear
to be contaminants or non-specific interacting proteins.
Generation of clones and purification of Pirh2-SPA.
Based on the results of the TAP purification experiment, we decided to create a
Pirh2-SPA (Fig 2.4A). The SPA tag is a triple flag tag, TEV protease site and calmodulin
binding peptide construct. The SPA construct is a smaller tag, and is a C-terminal tag; in
contrast, the protein A section of the TAP tag was about 20 kDa and the construct was an
N-terminal tag. We felt that the TAP-hPirh2 construct may not be folding properly, and
WB: CBP
55 __
40 __
33 __
24 __
72 __
17 __
TAP TAP-hPirh2
CBP–hPirh2
TAP-hPirh2
Lysate
Unbou
nd Ig
G
TEV Elua
te
Unbou
nd
Calm.
Bound
Calm
Lysate
Unbou
nd Ig
G
TEV Elua
te
Unbou
nd
CalmBou
nd C
alm
TAP
CBP
A
Figure 2.3 Purification of TAP and TAP-hPirh2
(A) TAP and TAP-hPirh2 CL11 cells were lysed in NP-40 lysis buffer. After centrifugation, TAP and TAP-hPirh2 were purified by binding to IgG, cleavage with TEV protease, and binding to calmodulin beads. Fractions in each step were separated by SDS-PAGE and transferred to PVDF, and blotted with antibody against CBP.
(B) Eluates from TAP Purification were separated by SDS-PAGE and silver stained. Bands were excised and identified by mass spectrometry.
TAP TAP-hPirh2
55 __
40 __
33 __
24 __
72 __
CBP-Pirh2
Hsc70
Tubulin
Alpha-S-casein
B
52
B
40__
33__
24__
17__
PARENT
SPAhPirh2-SPA
H1299
PARENTSPA
hPirh2-SPAPirh2-/- MEFs
SPA
hPirh2-SPA
WB: Flag
C
D
Actin
55 __
40 __
33 __
SPA Pirh2-SPA
Pirh2-CBP
EWB:hPirh2
Pirh2-SPA55__
40__
33__
24__
17__
11__
Pirh2
WB: hPirh2
H1299
hPirh
2
11 20 24 25 1 2 3 4 9 1914SPA hPirh2-SPA
Pirh2-SPA
SPA
55__
40__
33__
24__
17__
11__
Actin
WB: Flag
11 20 24 25 1 2 3 4 9 1914SPA hPirh2-SPA
H1299
hPirh
2
Figure 2.4 Generation and Purification of hPirh2-SPA Clones
(A) Diagram of hPirh2-SPA fusion protein.
(B) Western blots were performed on lysates prepared from the population stably transfected H1299 cells with SPA and hPirh2 SPA vectors, and Pirh2-/-E1A/ras MEFs infected with pBabe neo SPA or hPirh2-SPA.
(C, D) H1299 cells stably transfected were cloned and western blots were performed as indicated.
(E) Purification of CBP-hPirh2 from Pirh2-/-E1A/ras transformed cells (E).
Pirh2
FlagCBP
TEV Cleavage
A 53
54
therefore it was associated with heat shock proteins; we thought that by reducing the size
and the placement of the tag could help the protein fold properly. Pirh2-SPA clones were
stably transfected into H1299 cells or retrovirally infected into Pirh2-/- E1A/Ras MEFs
(Fig. 2.4B). Expression of Pirh2-SPA was quickly shut down in the Pirh2-/- E1A/Ras
MEFs; as a result, stable Pirh2-SPA expressing clones could not be generated (data not
shown). Pirh2-SPA and SPA clones were generated in H1299 cells, and analyzed by
blotting with Flag (Fig 2.4C) and Pirh2 (Fig 2.4D) antibodies. The Pirh2-SPA fusion
protein was expressed stably, but at significantly lower levels than endogenous Pirh2 (Fig.
2.4C,D). Since there is no endogenous Pirh2 to compete with the tagged construct in the
Pirh2-/- MEFs, we used the Pirh2-SPA-expressing Pirh2-/- MEFs to purify the protein.
We fractionated the eluate using SDS-PAGE, and expression of the tagged construct was
greatly reduced, but not completely lost as we were able to purify Pirh2-CBP (Fig. 2.4E).
The final eluate was lyophilized and sent for analysis by mass spectrometry resulting in
the identification of calpain small subunit 1 and S-100 calcium binding protein beta
subunit. Since purification using the calmodulin binding peptide is a calcium dependent
reaction, calcium associated proteins are often purified as contaminants. We did not
purify heat shock proteins or chaperone proteins suggesting that changing the tag, cell
type, expression level, or a combination of these factors may help to identify potential
interactors.
Generation of clones and purification of Pirh2-HF.
Based on the results of the SPA experiment, and that the construct lost expression
very quickly in the Pirh2 -/- E1A/Ras MEF population, we decided to use the His-Flag
His
CL1 CL2 CL9 CL10 CL11 CL14
Actin
His-Flag Pirh2-His-FlagB
His-Flag Pirh2 – His-Flag
Pirh2 – His - Flag
C
Pirh2
FlagHis
TEV CleavageA
Figure 2.5 Generation and Purification of Pirh2-His-Flag Clones.
(A) Diagram of Pirh2-His-Flag fusion protein.
(B) Pirh2-/- E1A/Ras MEFs were infected with retrovirus made from pBabepuro His-Flag or Pirh2-His-Flag and selected in puromycin. Clones (CL) were generated and lysates from the clones were used for western blot analysis with the indicated antibodies.
(C) A pool of His-Flag and Pirh2-His-Flag clones were used for purification. Successful purification was confirmed using western blot analysis.
His
55
56
expression construct. The new construct uses a six times histidine tag and three time flag
peptide for sequential purification (Figure 2.5A). We suspected that the calmodulin
binding peptide present in Pirh2-SPA may have been toxic to the Pirh2-/- E1A/Ras MEFs
(resulting in rapid loss of expression of Pirh2-SPA), and that replacing this peptide with
the histidine tag might alleviate the toxicity. Using retroviral infection of Pirh2-/-
E1A/Ras MEFs, we generated clones of the empty vector (His-Flag) or Pirh2-His-Flag
and picked clones of variable expression (Figure 2.5B). We then pooled a number of the
Pirh2-His-Flag or His-Flag clones, and performed the dual purification protocol. A pool
of clones 9, 10, 11 and 14 to for Pirh2-His-Flag and clones 1 and 2 for His-Flag was used
for the experiment (Fig 2.5B). We chose to make of pool of the clones for the experiment
to reduce clonal artefacts that may arise from purification from a single clone. Also there
is variability in expression levels in each clone; a pool of the Pirh2-His-Flag clones
equalizes expression levels among the four clones. A Western blot performed on the
eluates from the purification protocol indicates that Pirh2-His-Flag was successfully
purified (Figure 2.5C). Mass spectrometry was performed on the eluates. Only
contaminating proteins (keratin, trypsin) were identified.
Identification of Pirh2 interactors using GST-pull down.
Since we were unable to identify specific Pirh2 interacting proteins using the dual
purification techniques, we additionally used a GST-hPirh2 construct to identify
interacting proteins from immortalized Pirh2-/- kidney fibroblasts. Immortalized kidney
fibroblast cells were used since immortalized cells undergo less morphological and
genetic changes than transformed fibroblasts, and we wanted to identify targets under
57
physiological conditions. Pirh2 is also expressed in the kidney (Leng et al., 2003).
Identification of targets in the kidney is important in determining the role of Pirh2 in this
tissue. GST-hPirh2 and GST were bound to glutathione beads. Lysates from untreated
fibroblasts and fibroblasts treated with adriamycin were bound to GST-hPirh2 or GST to
determine if Pirh2 binds proteins differentially under conditions of stress (Fig 2.6).
Interacting proteins were separated by SDS-PAGE, bands were excised and identified
using mass spectrometry. Based on the results of the mass spectrometry analysis, we
identified PKC as a candidate interacting protein.
DISCUSSION
In an attempt to identify proteins that interact with Pirh2 we used TAP tagging,
and variations on the TAP tag (SPA tag, and His-Flag tag). In all cases, we purified
tagged Pirh2, but had difficulty identifying interacting proteins. We then tried to identify
interacting proteins by using GST-hPirh2 in a GST pull down experiment. We were able
to identify a single candidate interactor PKC that needs to be confirmed.
The difficulties encountered in the dual pull down and GST pull down
experiments may be explained by several reasons. The most likely explanation is that
substrate – ubiquitin ligase interactions and cofactor (i.e. ubiquitin conjugating enzyme) –
ubiquitin ligase interactions are thought to be weak and transient, and, hence, may not
survive the washes involved in the dual purification strategies or the GST-pull down
approach that were used (Deshaies and Joazeiro, 2009). If dual purification strategies are
used to identify substrates and cofactors for Pirh2, crosslinking the proteins would
Figure 2.6 GST-pull Down to Identify Novel Pirh2 Interacting Proteins.
Immortalized Pirh2-/- kidney fibroblasts were either untreated or treated for 24 hours with 200 ng/mL adrimycin. Cells were lysed in non-denaturing conditions, and an equal mass and volume of protein was incubated with either GST or GST-hPirh2 bound to glutathione beads. Bound proteins were eluted by boiling SDS, run on an SDS-page gel and stained with coomassie. Bands appearing only in the GST-hPirh2 lanes were excised and identified by mass spectrometry.
GSTGST-hPirh
2
GSTGST-hPirh
2Untreated Adriamycin
GST***
*
Pirh2
55 __
40 __
33 __
24 __
72 __
17 __
130 __
100 __
170 __
GST-hPirh2
Pirh2
Semaphorin 4D
PKC
* Bands sent for mass spectrometry but not identified
58
59
strengthen the interaction and facilitate the identification of interactors (Gingras et al.,
2005). Based on the results of the GST-pull down and mass spectrometry, PKC is a
candidate interacting protein. This interaction will need to be confirmed by endogenous
coimmunoprecipitation and confocal microscopy. If the interaction is confirmed, the
nature of the interaction between Pirh2 and PKC will also need to be determined.
There are several possibilities the most obvious being that Pirh2 may ubiquitinate PKC
or that PKC may phosphorylate Pirh2. Future experiments will need to address these
issues should the interaction be confirmed by endogenous coimmunoprecipitation.
PKC is a serine/threonine kinase, and activation of PKC is a critical pro-
apoptotic signal (Reyland, 2007). When cells are exposed to apoptotic stress such as
DNA damaging agents, oxidative stress and death receptors, PKC becomes
phosphorylated on multiple tyrosine residues and translocates into the nucleus.
Phosphotyrosine PKC is then cleaved by caspase-3. Caspase-3 cleavage causes the
release of an activated pro-apoptotic C-terminal fragment (Reyland, 2007). PKC and
the active cleavage fragment must be retained in the nucleus to cause apoptosis (DeVries-
Seimon et al., 2007). In a mouse model, PKC -/- mice exhibit a reduction in IR, UV
and etoposide induced apoptosis in the parotid gland (Humphries et al., 2006). Activated
PKC is targeted for ubiquitin-mediated proteolysis (Lu et al., 1998).
The results of the GST-pull down show little if any difference between
adriamycin treated and untreated cells (Fig. 2.6). There could be several possibilities.
We harvested cells from adriamycin treatment after 24 hours, and perhaps this time point
was too late to catch changes in the interaction profile. Additional pull-downs should be
performed with cells treated with adriamycin at various time points; a time course of pull
60
downs would be ideal to track the differences, if any. Also, varying the stress may
produce different interacting proteins; adriamycin causes double strand breaks, UV
radiation, serum starvation, and heat shock could be other options to determine if
interactors vary with the stress response. Again, a time course could also be useful to
determine when these interactions occur.
Clearly, additional approaches to identifying proteins that interact with Pirh2 need
to be performed. The approaches that are to be undertaken must address the transient
nature of the substrate/cofactor and ubiquitin ligase interaction to be able to successfully
purify the interactors.
CHAPTER 3
Pirh2 targets phosphoserine 15 p53
for ubiquitination
Thank you to Dr. Razq Hakem and the Hakem lab that provided the WT and Pirh2-/-
mice, immunohistochemistry for p53 on irradiated spleen (Anne Hakem), and TUNEL on
irradiated spleen and thymus (Benedicte Lemmers).
61
62
ABSTRACT
The p53 protein is activated and stabilized by post-translational modifications in
response to stress and promotes cell cycle arrest and apoptosis. Pirh2 is a p53-regulated
gene which encodes an E3 RING finger ubiquitin ligase that binds to p53 and negatively
regulates p53 by targeting it for ubiquitin-mediated proteolysis. Pirh2 is functionally
similar to Mdm2 a well-characterized E3 RING finger ubiquitin ligase; they both target
p53 for degradation, and are transcriptional targets for p53 activity. The stability of the
p53 protein is primarily regulated through ubiquitin mediated proteolysis, and there are
multiple ubiquitin ligases targeting p53. Here we address the question of functional
redundancy and show that Pirh2 can target serine 15 phosphorylated p53 which is
reported to not be regulated by Mdm2.
Phosphoserine 15 p53 levels are significantly higher in irradiated splenocytes and
thymocytes from Pirh2-/- mice compared with Pirh2+/+ mice and this has a functional
consequence on the kinetics of bax induction and apoptosis. In addition, cells stably
expressing ectopic Pirh2 have decreased levels of phosphoserine 15 p53 and p21 after
DNA damage.
INTRODUCTION
The p53 tumour suppressor is a sequence-specific transcription factor that
promotes cell cycle arrest or apoptosis in response to various forms of cellular stress
(Levine, 1997). Cell cycle arrest normally occurs through transcriptional activation of
target genes such as the p21WAF1 cyclin-dependent kinase inhibitor; induction of p21 is a
key step in causing G1 cell cycle arrest (el-Deiry et al., 1993; Harper et al., 1993). Cells
63
from p21 knockout mice are deficient in undergoing p53-dependent cell cycle arrest in
response to DNA damage (Brugarolas et al., 1995; Deng et al., 1995). p53-dependent
apoptosis is partially regulated by transcriptional activation of target genes including bax
(Yin et al., 1997).
The p53 protein is primarily regulated at the post-translational level. Under
normal conditions, the p53 protein is extremely unstable, and rapidly targeted for
ubiquitin-mediated proteolysis by the E3 ubiquitin ligase Mdm2 (Michael and Oren,
2003). After DNA damage, both Mdm2 and p53 undergo several post-translational
modifications including acetylation and phosphorylation. In the presence of double
strand breaks, activated ATM kinase can phosphorylate both p53 on serine 15 and Mdm2
on serine 395. These modifications are proposed to aid in the dissociation of the Mdm2-
p53 complex and contribute to the stabilization and activation of p53 as part of the stress
response (Banin et al., 1998; Maya et al., 2001; Shieh et al., 1997). Mice that express a
p53 serine 15 to alanine mutant are partially deficient in p53-dependent apoptosis and cell
cycle arrest in response to DNA damage suggesting that phosphorylation of p53 on serine
15 reflects p53 protein activation (Chao et al., 2000a; Chao et al., 2000b).
Several ubiquitin ligases have been identified that target p53 for ubiquitin
mediated proteolysis, and the reason for such a high degree of functional redundancy is
not known (Toledo and Wahl, 2006). Among the E3 ubiquitin ligases that target p53
have been identified is Pirh2 (p53-induced protein with a ring H2 domain) (Leng et al.,
2003). Pirh2 was identified as a p53 target gene in a differential display experiment
performed using the p53-null Friend-virus transformed mouse erythroleukemia cell line
(DP16.1/p53ts) that stably expresses a gene that encodes a temperature sensitive (ts)
64
mutant p53 allele. Pirh2 gene expression is p53-dependent, however there appears to be
tissue or stress specific components to p53-induction of Pirh2. Previous experiments
have shown that Pirh2 can ubiquitinate p53 and target it for degradation; antisense
oligonucleotide-mediated repression of Pirh2 in normal fibroblast cells results in an
increase in p53 protein level, and a modest increase in cell cycle arrest (Leng et al., 2003).
In this study, we used splenocytes and thymocytes from Pirh2 null mice, and
noted that basal levels of p53 in these tissues remain unchanged compared to the wild-
type counterparts. After irradiation, both induction of total p53 levels and phosphoserine
15 p53 levels were higher in the Pirh2 null splenocytes and thymocytes compared to
wild-type splenocytes and thymocytes. This increase in total p53 and phosphoserine 15
p53 affects the kinetics of bax and apoptotic induction – bax and apoptosis are induced
earlier relative to the wild-type controls. In cells that overexpress Pirh2, levels of
phosphoserine 15 p53 and the p53 target gene p21 are reduced after DNA damage
compared to the vector controls. We also show that Pirh2 can coimmunoprecipitate
phosphoserine 15 p53 and Pirh2 null splenocytes show decreased levels of ubiquitinated
phosphoserine 15 p53. These results suggest that Pirh2 may directly regulate
phosphoserine 15 p53, and begins to address the question of functional redundancy
among the ubiquitin ligases that target p53 for degradation.
EXPERIMENTAL PROCEDURES
Cell Culture. MCF-7 cells and H1299 cells were cultured in -MEM supplemented with
10% fetal calf serum (FCS). MCF-7 clones were previously created and published (Leng
et al., 2003). 293 cells were cultured in Dulbecco’s MEM in 10% FCS. Splenocytes and
65
thymocytes were cultured in RPMI supplemented with 10% FCS and 568 µM β-
mercaptomethanol.
Splenocyte and Thymocyte Culture Preparation. To prepare splenocyte and
thymocyte culture, spleen and thymus were harvested from age matched wild-type and
Pirh2-/- mice. The organs were homogenized in 70 µm cell strainers (BD Biosciences).
Cells were resuspended in red blood cell lysis buffer (Sigma) and washed once with
phosphate buffered saline (PBS). Splenocytes and thymocytes were cultured in RPMI,
10% FCS and 568 µM β-mercaptomethanol, and irradiated with 6 Gy of γ-IR from a
cesium 137 source. Wild-type and Pirh2-/- splenocytes and thymocytes were harvested at
time points after irradiation, and lysed in a 1x Laemmli SDS sample buffer for western
blot analysis.
Preparation of MCF-7 p53 shRNA Cells. MCF-7 cells were plated (1 x 105) in a 10 cm
dish one day before transfection and were then transfected with 10 µg of pcDNA3 Eco
plasmid using calcium phosphate. 293 cells (3 x 106) were plated in a 10 cm dish one day
before transfection with calcium phosphate and 5 µg pCl-Eco and 5 µg pSuper.puro
p53shRNA plasmid. The medium was changed one day after transfection, and the viral
supernatant was harvested 24 hours later and used to infect MCF-7 cells expressing the
ecotropic receptor in the presence of 8 µg/mL hexadimethrine bromide (Sigma) for 8
hours during the day at 37ºC. MCF-7 cells were then infected a second time with viral
supernatant and 8 µg/mL hexadimethrine bromide (Sigma) for 16 hours overnight at 37ºC.
66
Infected MCF-7 cells were washed with PBS and incubated for 24 hours in -MEM and
10% FCS. MCF-7 cells were then selected with 2 µg/mL puromycin (Invivogen).
Western Blotting. Antibodies for p53 (DO-1, FL393), Mdm2 (SMP14) and p21 (C-19)
were obtained from Santa Cruz. The phosphoserine 15 p53 antibody was obtained from
Cell Signaling, and β-actin antibody from Sigma. Antibodies for Pirh2 were described
previously (Leng et al., 2003) or were obtained from Bethyl laboratories. Quantitation of
the phosphoserine 15 p53 and p53 signal were performed on the LiCOR Odyssey
fluorescence imager.
Immunohistochemistry. Whole body irradiation (6 Gy) was performed on matched
wild-type and Pirh2-/- mice. Tissues were fixed in buffered formalin, processed for
paraffin-embedded sectioning at 5 µm, and stained with H&E (Fisher). IHC was
performed using anti-p53 (FL393, Santa Cruz). TUNEL staining was performed on
histological sections using an in situ cell death detection kit (Boehringer Mannheim).
Coimmunoprecipitation. Rabbit IgG (Cedarlane), phosphoserine 15 p53 (Cell
Signaling) and human Pirh2 antibodies were irreversibly crosslinked to protein A beads
using DSS (Pierce) as per Seize X immmunoprecipitation kit instructions (Pierce). MCF-
7 cells were treated for 16 hours with 200 ng/mL adriamycin (Sigma), and 4 hours with
50 µM MG132 (Sigma). Cells were lysed in 600 µL IP lysis buffer (50mM HEPES pH
8.0, 5 mM EDTA, 100 mM sodium chloride, 0.35% NP-40, 10 mM sodium diphosphate,
10 mM sodium fluoride, 1 mM sodium orthovanadate, 50 µM MG132, and EDTA-free
67
protease inhibitor tablet (Roche)). Cell lysates were rotated for 30 minutes at 4ºC with
0.625 mM DSP (Pierce), and 150 µL of 1 M Tris pH 8.0 was added and rotated at 4ºC for
30 minutes. Protein lysate (2 mg) was mixed with the crosslinked antibody beads and
rotated at 4ºC for 3 hours. Beads were washed three times with wash buffer (50 mM Tris
pH 8.0, 5 mM EDTA, 100 mM sodium chloride, 0.1% NP-40, 10 mM sodium
diphosphate, 10 mM sodium fluoride, 1 mM sodium orthovanadate). Elution was
performed 1x Laemmli SDS sample buffer with 0.1M DTT and heated at 60ºC for 5
minutes.
Ubiquitination Assay. Splenocyte cultures were prepared as previously described.
Wild-type and Pirh2-/- splenocytes were treated with 6 Gy γ-IR and 50 µM MG132 for 4
hours. Cells were lysed in 1x Laemmli SDS sample buffer with 10 mM sodium
diphosphate, 10 mM sodium fluoride, 1 mM sodium orthovanadate. Lysates were used
for western analysis.
Flow Cytometry. MCF-7 cells and clones were plated in a (1 x 106) 10 cm dish one day
before drug treatment. Cells were treated with 200 ng/mL adriamycin (Sigma) for 14
hours; for the last half hour of adriamycin treatment, cells were also treated with 10 µM
of BrdU (Sigma). Cells were fixed in 70% ethanol overnight at -20ºC. Cells were
counted and 1x106 cells were used per sample. Cells were resuspended in denaturing
solution (2 M hydrochloric acid and 0.5% Triton-X) for half an hour at room temperature.
Cells were then resuspended in 0.1 M sodium borate pH 8.5 for two minutes at room
temperature. Cells were washed in 2% bovine serum albumin (BSA) in PBS. Cells were
68
resuspended in anti-BrdU-FITC antibody (BD Biosciences), RNase A, 0.5% Tween-20
and 1% BSA in PBS and incubated overnight at 4ºC. Cells were washed with 2% BSA in
PBS and resuspended in 10µg/mL propidium iodide (Sigma) and 0.05% Triton-X in PBS
for half an hour at room temperature. Cells were washed with 2% BSA in PBS and
resuspended in PBS for analysis by flow cytometry on a BD FACS Calibur.
RESULTS
Pirh2-/- Tissues Show Elevated Levels of p53 After DNA Damage. Pirh2-/- mice
were provided by Dr. Razq Hakem. Spleen and thymus were harvested from age
matched wild-type and Pirh2-/- mice, and irradiated with 6 Gy IR.
Immunohistochemistry on the spleen indicates that p53 levels are elevated 1 hour post-
irradiation in the Pirh2-/- spleen relative to Pirh2+/+ spleen (Fig. 3.1A). Western blot
analysis of non irradiated spleen (Fig 3.1B) and thymus (Fig 3.1C) revealed no difference
in p53 protein levels between the wild-type and Pirh2-/- strains. After irradiation,
however, p53 levels appeared to increase earlier in the Pirh2-/- spleen and thymus; at
later times after irradiation, the levels of p53 were similar in both strains indicating that
Pirh2 regulation of p53 may affect the kinetics of induction.
Pirh2-/- Tissues Show Elevated Levels of Phosphoserine 15 p53 After DNA Damage.
After DNA damage, p53 is phosphorylated by ATM on serine 15 (Banin et al., 1998).
Pirh2 has also been shown to interact with the DNA binding domain and the
oligomerization domain of p53, which are not affected by serine 15 phosphorylation
(Leng et al., 2003; Sheng et al., 2008). Irradiated splenocytes (Fig 3.1B) and thymocytes
WT Pirh2-/-
Total p53
mPirh2
β-actin
0 0.5 1 2 4 6 0 0.5 1 2 4 6
WT Pirh2-/-Time(hr)
A
B
C
Figure 3.1 Elevated p53 levels in irradiated Pirh2-/- tissues
(A) Age matched WT and Pirh2-/- mice were irradiated with 6 Gy of IR and sacrificed at one hour. Spleens were harvested and stained for p53 levels by immunohistochemistry.
Splenocytes (B) and thymocytes (C) were harvested fro age matched WT and Pirh2-/- mice. Cells were irradiated with 6 Gy IR, harvested and lysed at indicated time points for western blotting
Total p53
mPirh2
β-actin
0 0.5 1 2 4 6 0 0.5 1 2 4 6
WT Pirh2-/-Time(hr)
PhosphoSer 15 p53
69
70
(Fig 3.2A) from Pirh2-/- and wild-type mice were used to determine levels of
phosphoserine 15 p53. Levels of phosphoserine 15 p53 are increased in the Pirh2
knockout mice relative to the wild-type counterparts. Since levels of p53 are also
elevated in the Pirh2-/- mice, quantitation of phosphoserine p53 and total p53 levels were
performed using irradiated splenocytes in order to normalize for the increase in total p53.
The data are summarized in Fig. 3.2B, and are displayed as a ratio of phosphoserine 15
p53 levels to total p53 levels. The ratio of phosphoserine 15 p53 to total p53 is higher in
the Pirh2-/- splenocytes suggesting that the observed increase in total p53 does not
account for the increase in phosphoserine 15 p53. These data suggest that Pirh2 regulates
the level of the transcriptionally active form of p53 that is phosphorylated on serine 15.
Apoptotic Rate is Increased in Pirh2-/- Mice. To determine if the increased levels of
p53 protein, and phosphoserine 15 p53 has functional significance, we measured levels of
the p53 target gene Bax in splenocytes (Fig. 3.3A) and thymocytes (Fig 3.3B) after
irradiation. In accordance with the observed increase in total and phosphoserine 15 p53
levels, Pirh2-/- splenocytes and thymocytes also have increased levels of Bax protein.
Moreover, Bax induction occurs earlier compared with wild-type controls. Levels of
apoptosis were visualized using the TUNEL assay (Fig. 3.3C). Apoptosis in irradiated
spleen and thymus from Pirh2-/- mice occurs earlier than in wild-type cells. At later time
points, the number of apoptotic (TUNEL-positive) cells is similar. Hence the kinetics of
apoptosis induction is increased in Pirh2-/- thymocytes and splenocytes compared with
wild-type cells.
0
0.1
0.2
0.3
0.4
0.5
0.6
0.7
0.8
0.9
1
0 1 2 4 6
Hours
Rat
io o
f S
15p
53/p
53to
t
Pirh2-/-
WT
PhosphoSer 15p53
Total p53
0 1 2 4 6 0 1 2 4 6
WTPirh2-/-Time (hrs)
A
PhosphoSer 15p53
β-actin
0 1 2 4 6 0 1 2 4 6
WT Pirh2-/-Time (hr)
Total p53
B
Figure 3.2 Elevated levels of phosphoserine 15 p53 in Pirh2-/- mice
Thymocytes were harvested from age matched WT and Pirh2-/- mice (A). Cells were treated with 6 Gy IR and harvested at various time points. Westerns were performed using indicaed antibodies and developed using film. Quantitation of splenocytes was performed using the LiCOR Odyssey (B).
71
Thymus Spleenwt -/- wt -/-
0 hr
4 hr
8 hr
12 hr
β-actin
Bax
0 0.5 1 2 4 6 0 0.5 1 2 4 6
WT Pirh2-/-
mPirh2
Time(hr)
β-actin
Bax
0 0.5 1 2 4 6 0 0.5 1 2 4 6
WT Pirh2-/-
mPirh2
Time(hr)
A
B
C
Figure 3.3 Elevated levels of apoptosis in Pirh2-/- irradiated mice
Western blot analysis of Bax levels in irradiated splenocytes (A) and thymocytes (B) from age matched WT and Pirh2-/- mice. Cells were irradiated with 6 Gy IR and harvested at various time points.
(C) TUNEL staining of spleen and thymus harvested from age matched WT and Pirh2-/- mice irradiated with 6 Gy of IR.
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73
Pirh2 Can Regulate p53 Mutants for Serine 15. The observation that the accumulation
of phosphoserine 15 p53 occurs earlier after irradiation in Pirh2-/- cells prompted us to
test whether Pirh2 can regulate p53 serine 15 mutants. Pirh2, Mdm2 or empty vector
(pcDNA3) were cotransfected with wild-type p53 or p53S15A or p53S15D into p53-null
H1299 cells (Fig. 3.4). We used the wild-type p53 as a control, the p53S15A remains
unmodified as the protein cannot be phosphorylated and the p53S15D protein mimics
constitutive phosphorylation at serine 15 due to the acidic aspartic acid residue. Our
hypothesis is that Pirh2 can regulate all three constructs, where as Mdm2 would be
predicted to regulate wild-type p53, and the p53S15A protein, but not the p53S15D
protein. Transfected cells were collected 40 hours after transfection, and cell extracts
were prepared and analyzed by western blotting. As expected, Mdm2 and Pirh2 were
able to degrade wild-type p53 and p53S15A and resulted in decreased levels of p53
protein. Pirh2 is also capable of degrading p53S15D, and surprisingly, Mdm2 can
degrade this form of p53 as well. Results for Mdm2 degradation of p53S15D are
reported in the literature, and suggest that serine 15 phosphorylation p53 is not sufficient
to abrogate the interaction between p53 and Mdm2, and Mdm2 is still able to degrade the
p53 S15D mutant (Ashcroft et al., 1999; Dumaz and Meek, 1999). The ability of Pirh2 to
degrade the p53S15D mutant is consistent with the previous results that suggest that
Pirh2 regulates phosphoserine 15 p53.
Cells Overexpressing Pirh2 Decrease Levels of Phosphoserine 15 p53 and p21, But
Do Not Affect Cell Cycle Arrest. To complement the knock out experiments, MCF-7
cells that overexpress Pirh2 were used (Leng et al., 2003). If loss of Pirh2 results in an
p53
Mdm2
hPirh2
β-actin
pcDNA3(
neo)
Mdm
2
hPirh
2pcD
NA3(ne
o)
Mdm
2
hPirh
2pcD
NA3(ne
o)
Mdm
2
hPirh
2
p53 wt p53 S15A p53 S15D
Figure 3.4 Mdm2 and Pirh2 can regulate p53 serine 15 phosphomutants
H1299 cells were cotransfected using calcium phosphate with either empty vector (pcDNA3), Mdm2 or hPirh2 and a p53 construct (wt, S15A, S15D). Lysates were made from the transfected cells at 40 hours after transfection, and western blots were performed for the indicated antibodies.
74
75
increase in phosphoserine 15 p53 and the p53 target gene bax, in cells that overexpress
Pirh2, we should see an opposing effect: there should be a decrease in phosphoserine 15
p53 and the p53 target gene p21. MCF-7 cells were treated with adriamycin for 16 hours.
Clones overexpressing Pirh2 have decreased levels of phosphoserine 15 p53 and
consequently, p53 target gene, p21 (Fig 3.5) compared to vector controls and a control
that overexpresses Mdm2. However, we do not see a difference in the induced p53 levels
among the two vector controls, the Mdm2 overexpressing clone, and the two clones that
overexpress p53. Also, all clones were able to induce the p53 target gene Mdm2 equally
well suggesting that the effect of reduced phosphoserine 15 p53 protein level in the Pirh2
overexpressing clone may affect specific p53 target genes. To determine if there is a
functional consequence of decreased p21 induction in the Pirh2 overexpressing clones,
cell cycle analysis was performed using bromodeoxyuridine and propidium iodide flow
cytometry analysis. MCF-7 cells that stably express p53shRNA were used as a negative
control for cell cycle arrest. These cells show a reduced levels p53 and p21 in the
absence of stress by western blot and after treatment with adriamycin for 24 hours, the
MCF-7 p53shRNA the cells show no induction of p53 and p21 (Figure 3.6A). Results
are summarized in Fig. 3.6B by plotting the percentage of cells in S-phase. MCF-7 cells
that stably express p53shRNA fail to arrest in G1 in response to adriamycin treatment.
After 14 hours of adriamycin treatment, there is no significant decrease in cell cycle
arrest in response to Pirh2 overexpresssion. We have also examined various time points
using 200 ng/mL adriamycin (12 hours, 16 hours, 20 hours, and 24 hours) and different
concentrations of adriamycin (50 ng/mL, 100 ng/mL, 150 ng/mL adriamycin) and noted
no significant decrease in cell cycle arrest (data not shown). Although, the levels of
p21
Pirh2
β-actin
Total p53
PhosphoSer 15 p53
Mdm2
pcDNA3.1
pcDNA3.
2
Mdm2
Pirh2-1
Pirh2-2
Control 200ng/mL Adriamycin
pcDNA3.1
pcDNA3.
2
Mdm2
Pirh2-1
Pirh2-2
Figure 3.5 Decreased levels of phosphoserine 15 p53 and p21 in Pirh2 overexpressing cells.
MCF-7 clones stably transfected with vector, Mdm2, or Pirh2 were created. Cells were either untreated or treated with 200 ng/mL adriamycin for 16 hours and harvested. Western blots were performed with the indicated antibodies.
76
Figure 3.6 Overexpression of Pirh2 does not decrease S-phase after DNA damage
(A) To generate p53shRNA expressing cells, MCF-7 cells were transiently transfected with the ecotropic receptor, and infected with ecotropic virus that expresses the p53shRNA in the pSuper.retro puro backbone. MCF-7 cells were selected using 2ug/mL puromycin for 24 hours. Validation of the p53shRNA was performed by treating cells with 200µg/mL of adriamycin for 24 hours. Western blots were performed as indicated
(B) The stably transfected MCF-7 cells were either untreated or treated with 200 ng/mL of adriamycin for 14 hours. Cells were pulsed with 10 µM BrdU for half an hour and then fixed and stained with anti-BrdU-FITC antibody and PI. Cell cycle analysis was performed using flow cytometry.
77
0
10
20
30
40
50
60
p53s
hRNA C
ontro
l
pcDNA3.
1 Con
trol
pcDNA3.
2 Con
trol
Mdm
2 Con
trol
Pirh2-
1 Con
trol
Pirh2-
2 Con
trol
p53s
hRNA A
dr
pcDNA3.
1 Adr
pcDNA3.
2 Adr
Mdm
2 Adr
Pirh2-
1 Adr
Pirh2-
2 Adr
Ave
rage
S-p
hase
(%
)
A
B
p21
β-actin
p53
Adr - + - +
MCF-7Parental
MCF-7 p53shRNA
78
phosphoserine 15 p53 and p21 are reduced in MCF-7 cells that overexpress Pirh2, this
does not appear to have an effect on cell cycle progression.
Pirh2 Coimmunoprecipitates with Phosphoserine 15 and Promotes Phosphoserine
15 p53 Ubiquitination. To determine if the regulation of phosphoserine 15 p53 by Pirh2
is direct, we performed coimmunoprecipitation and ubiquitination experiments.
Previous experiments have shown that unmodified p53 can interact with Pirh2, and Pirh2
can target p53 for polyubiquitination in in vitro experiments (Leng et al., 2003). An
endogenous coimmunoprecipitation experiment using MCF-7 cells treated with
adriamycin and the proteasomal inhibitor MG132 was performed. The results presented
in Fig 3.7A show that p53 phosphoserine 15 coimmunoprecipitates with Pirh2. In
addition, Pirh2 coimmunoprecipitates with phosphoserine 15 p53. These results clearly
show that Pirh2 can interact physically with p53 phosphorylated on serine 15.
Ubiquitination of phosphoserine 15 p53 was also performed in vivo using splenocytes
from wild-type and Pirh2-/- mice. Splenocytes were treated with IR and MG132 and
harvested for analysis by western blotting (Fig. 3.7B). Splenocytes from wild-type mice
show an increase in high molecular weight laddering of phosphoserine 15 p53 that
usually suggests polyubiquitination. These experiments indicate that phosphoserine 15
p53 interacts with Pirh2, and suggest that Pirh2 can promote the ubiquitination of
phosphoserine 15 p53.
WB: PhosphoSer 15 p53
IP: R
abbi
t IgG
IP: P
irh2
IP: S
er15
p53
Inpu
t 1%
WB: Pirh2
A
B
Figure 3.7 Pirh2 interacts with and ubiquitinates phosphoserine 15 p53
(A) MCF-7 were treated for 16 hours with 200 ng/mL of adriamycin and for 4 hours with 50 µM MG132. Cells were harvested in 0.35% NP-40 lysis buffer and an equal amount and volume of protein was bound to either rabbit IgG, Pirh2 or phosphoserine 15 p53 antibody. Western blots were performed for the indicated antibodies .
(B) Splenocytes were harvested from age matched WT and Pirh2-/- mice that were irradiated with 6 Gy IR and treated for 4 hours with 50 µM MG132. Lysates were prepared and western blotting was performed with indicated antibodies.
PhosphoSer 15 p53
mPirh2
β-actin
WTPirh2-/-
Phosphoserine
15 p53
PhosphoSer 15 p53 -
Ubiquitin
170 __
100 __
130 __
72 __
55 __
79
80
DISCUSSION
In this study, splenocytes and thymocytes from Pirh2-/- mice were used to
investigate the regulation of p53 by Pirh2. Several key observations arise from these
studies. First, the levels of p53 in unstressed splenocytes and thymocytes remain
unchanged in Pirh2-/- cells compared with wild-type cells. This observation would
suggest that Pirh2 does not have a critical role in regulating basal levels of p53 in
unstressed conditions. The second set of experiments tests whether Pirh2 can regulate
p53 after DNA damage. We determined that levels of p53 are modestly elevated and the
levels of phosphoserine 15 p53 are elevated in Pirh2-/- splenocytes and thymocytes after
DNA damage. The increased levels of total and phosphoserine 15 p53 has a functional
consequence as the rate of apoptosis was increased in Pirh2-/- cells as was the rate of
induction of p53-dependent pro-apoptotic target bax. These results suggest that Pirh2
may play a role in regulating p53 after DNA damage.
In the absence of Pirh2, the rate of p53-dependent apoptosis is increased along
with induction of bax. This observation fits with the model that Pirh2 regulates p53 after
DNA damage. However, in MCF-7 cells overexpressing Pirh2, p53-dependent cell cycle
arrest is not attenuated though levels of the cell cycle inhibitor p21 are reduced. This
discrepancy could be explained by several factors. In the MCF-7 cells, there may be a
low threshold of p21 activity required to produce cell cycle arrest or other
downregulation of other Pirh2 substrates may promote cell cycle progression. Also, we
measured a difference in the kinetics of apoptosis; if Pirh2 affects the kinetics of p53-
induction it may be difficult to measure the kinetics of p53-induced cell cycle arrest. It is
also possible that Pirh2 may regulate cell cycle arrest in specific cell types or under
81
specific cell conditions. In the MCF-7 clones that overexpress Pirh2, phosphoserine 15
p53 protein levels and induction of p21 protein was reduced in response to adriamycin,
but induction Mdm2 protein was unaffected by the decreased phosphoserine 15 p53
levels (Fig 3.5). Further experiments with different cell types and different cell stresses
and additional p53 target genes should be performed to determine whether
overexpression of Pirh2 can affect the kinetics of p53-dependent apoptosis or p53-
dependent cell cycle arrest. We can also harvest fibroblasts from Pirh2+/+ and Pirh2-/-
mice to test various DNA damaging agents for p53-dependent target gene expression, and
to measure cell cycle arrest. In Pirh2-/- fibroblasts we would expect to see an increase in
p53-dependent cell cycle arrest target genes, and an increase in p53-dependent cell cycle
arrest.
Pirh2 was identified as a p53-regulated gene that targets p53 for ubiquitin-
mediated proteolysis. Leng et al. (2003) reported that Pirh2 directly binds to p53 and can
catalyze polyubiquitination of p53; moreover knock down of Pirh2 by antisense modestly
increased p53 dependent G1 cell cycle arrest (Leng et al., 2003). Transfection
experiments also show that Pirh2 only targets the tetrameric form of p53 while Mdm2 is
proposed to target all forms of p53 (Sheng et al., 2008). Our data shows that Pirh2 can
coimmunoprecipitate with phosphoserine 15 p53, and Pirh2 also promotes ubiquitination
of phosphoserine 15 p53, an activating modification on p53 after DNA damage (Fig 3.7)
(Chao et al., 2000a). Based on these observations, we propose that Pirh2 may regulate
the active form of p53. In contrast, the role of Mdm2 is to regulate the basal levels p53.
This is consistent with observations from the literature. Mdm2 knockout mice are
embryonic lethal, and this lethality can be rescued by deletion of p53 (Jones et al., 1995;
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Montes de Oca Luna et al., 1995). Also, a mouse model was made where a hypomorphic
Mdm2 allele was conditionally knocked into the thymus; the thymus in the hypomorphic
Mdm2 mouse was much smaller than the thymus in the Mdm2+/+ mouse under
unstressed conditions. In double mutant p53-/- Mdm2 hypomorphic mice, the thymus is
larger, and is comparable in size to the Mdm2+/+ thymus. These results suggest that
unrestrained p53 activity causes the atrophy of the thymus in the Mdm2 hypomorphic
mice, and that Mdm2 keeps p53 activity in check in unstressed conditions in the thymus
(Mendrysa et al., 2003; O'Leary et al., 2004). The Pirh2 knockout mice are viable and
develop normally, and in unstressed conditions show no difference in basal p53 level
when compared to the wild-type mice in the spleen and thymus. This would suggest that
Mdm2 is the main regulator of p53 under unstressed conditions. Its main role is to keep
p53 levels in check, as unrestrained p53 activity is detrimental to the organism – it would
cause massive apoptosis and cell cycle arrest. There is also significant evidence that
Mdm2 is dissociated from p53 after DNA damage, and as a result could not induce the
degradation of p53 (Maya et al., 2001; Shieh et al., 1997). For example, a time course of
coimmunoprecipitation of p53 and Mdm2 shows that the complex dissociates after DNA
damage (Shieh et al., 1997). In addition, ATM has been shown to phosphorylate Mdm2
on serine 395, and an aspartic acid phosphomimic on serine 395 on Mdm2 shows reduced
ability to degrade p53 in transfection experiments (Maya et al., 2001). The results of Fig
3.4 suggest that phosphorylation of serine 15 p53 is insufficient to prevent degradation of
p53 by Mdm2. Fig 3.5 shows that overexpression of Mdm2 does not prevent activation
of p53 and implies that Mdm2 is dissociated from p53 after DNA damage as levels of
p53, phosphoserine 15 p53 and p21 in Mdm2 overexpressing cells are similar to those
83
from the vector control. We suggest that under conditions of DNA damage where Mdm2
does not degrade p53, Pirh2 degradation of active p53 can be observed. The role of Pirh2
in regulating p53 may be to fine-tune the DNA damage response.
CHAPTER 4
Summary and Future Directions
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85
I. Summary of Findings: Identification of Pirh2 Interactors
The ubiquitin ligase Pirh2 has several known targets for degradation including
p53, HDAC1, p27/Kip1, SRβ and ε-COP (Abe et al., 2008; Hattori et al., 2007; Leng et
al., 2003; Logan et al., 2006; Maruyama et al., 2008). We used several dual purification
strategies, the TAP tag, a flag-calmodulin binding peptide tag, and a histidine-flag tag, to
identify proteins that interact with Pirh2 and planned to characterize the interactors as
cofactors or substrates. We also used a GST-pull down approach to identify interactors
under both normal and stress conditions. We had very little success with the two
approaches, with the exception of potentially identifying PKC, and have determined that
additional approaches to identify substrates and cofactors must be investigated.
Future Directions: Identification of Pirh2 Interactors
Identification of Pirh2 co-factors and substrates.
Additional approaches to identify substrates and cofactors for Pirh2 are required.
These strategies need to account for the weak and transient nature of the interaction
between Pirh2 and either substrate or cofactor. Pull downs, or coimmunoprecipitation
experiments may not identify specific interacting proteins since they could be lost in the
numerous washes. There are several alternative approaches to identify substrates for
Pirh2. The first is to fluorescently label cells from Pirh2+/+ and Pirh2-/- mice with
different fluorochromes. One would mix equal amounts of the protein lysates from both
Pirh2+/+ and Pirh2-/- mice, and separate the proteins using two dimensional gel
electrophoresis. The spots from the protein mixture could be visualized on the Typhoon
or an appropriate fluorescence reader and one would look for proteins that are highly
86
expressed in Pirh2-/- cells. These proteins would be candidate substrates, and one would
need to confirm that they are direct substrates of Pirh2.
A second potential approach relies on advances in the mass spectrometry. This
approach would involve labelling of whole cell lysate with one of two tags cICAT
(cleavable Isotope-Coded Affinity Tags) or ITRAQ (Isobaric Tags for Relative and
Absolute Quantification). In these two labelling techniques, cICAT involves labelling of
cysteine residues with either a heavy or light isotope and a biotin tag. The biotin tag
causes problems in the mass spectrometry analysis and the cICAT tag involves acid
cleavage of the biotin tag. ITRAQ labelling techniques involves labelling of the N-
terminus and amino side chains with a set of amine reactive isobaric tags (Wu et al.,
2006). The two labelled samples are then mixed, trypsinized and analyzed by
multidimensional LC-MS/MS (the lysates would be separated by cation exchange and
C18 reverse-phase column chromatography) (Wu et al., 2006). To identify Pirh2
substrates, one would isolate cells from Pirh2+/+ and Pirh2-/- mice, and label the lysate.
For example, the Pirh2+/+ lysate would be labelled with the heavy isotope, and Pirh2-/-
lysate would be labelled with the light isotope. An equal mass of protein from labelled
Pirh2+/+ and Pirh2-/- splenocyte lysates would be mixed, trypsinized, and analyzed by
multi-dimensional LC-MS/MS. The proteins whose peptides have higher peaks from the
mass spectrometry spectrum from the light isotope sample would be considered candidate
substrates.
A third potential approach to identify substrates would be to perform an in vitro
ubiquitination assay using Pirh2 as the ligase and whole cell lysate as the substrate. One
would perform the ubiquitination assay using histidine tagged 7KR ubiquitin mutant.
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The ubiquitinated substrates can be purified using the histidine tag, and the substrates can
be eluted and then identified by mass spectrometry. The histidine tagged 7KR ubiquitin
is used because the histidine tag allows for purification of substrates under denaturing
conditions that will inactivate proteases and deubiquitinases, and the 7KR ubiquitin
reduces the amount of ubiquitin co-purified with the substrates. In the presence of excess
ubiquitin, substrates may not be identified using mass spectrometry; ubiquitin peptides
will be dominant, and will mask signals from other peptides (Yi Sheng, unpublished data,
personal communication).
A fourth possibility for identifying substrates uses protein arrays. In this approach,
proteins are spotted on an array and these serve as substrates in an in vitro ubiquitination
assay with Pirh2 as the E3 ubiquitin ligase and biotinylated ubiquitin. The protein array
is then washed with SDS to thoroughly remove any non-covalently bound ubiquitin. The
biotinylated ubiquitin is then incubated with a strepavidin conjugated to a fluorochrome.
Potential substrates for the ubiquitin ligase can be identified using a fluorescence imaging
detector (Andrews PS, personal communication, Keystone Conference 2009: The Many
Faces of Ubiquitin).
Each of these approaches has advantages and disadvantages. The main advantage
of the two dimensional gel electrophoresis and cICAT/ITRAQ labelling experiments are
that they rely on physiological conditions; however, the results may be caused by an
indirect mechanism. For example, if Pirh2 were to degrade a transcriptional activator, in
the absence of Pirh2 all the targets of that transcriptional activator would be
overexpressed and appear to be Pirh2 targets; hence their overexpression would be an
indirect result of the loss of Pirh2. Candidate substrates would have to be validated using
88
in vitro studies and coimmunoprecipitation and colocalization studies. The third and
fourth approaches rely on in vitro ubiquitination assays that may give false positives. The
key difference in the third and fourth approach is how the substrates are identified. The
third approach uses mass spectrometry to identify the substrate; the sensitivity of mass
spectrometry is heavily dependent on the number of peptides present in the mixture, and
the presence of ubiquitin peptides may reduce the sensitivity of being able to identify the
protein by mass spectrometry analysis (Steen and Mann, 2004). This limitation of mass
spectrometry analysis is also a disadvantage in the cICAT and ITRAQ approach to
identifying substrates – the labelled peptides that show a difference between Pirh2+/+
cells and Pirh2-/- cells must be in high abundance to be successfully detected using mass
spectrometry. The protein chip approach is limited by the number of proteins on the chip,
and any post-translational modifications that may influence binding to Pirh2 since protein
chips contain unmodified proteins. For example, in the two dimensional gel
electrophoresis, cICAT/ITRAQ and the in vitro 7KR ubiquitination assay approaches to
substrate identification, one can treat the cells with drugs, serum starve or use many other
stresses that could influence the substrate specificity of Pirh2.
Identification of cofactors is more difficult. There are approximately fifty E2
ubiquitin conjugating enzymes in the human genome; the E2s often determines the
pattern of ubiquitin conjugates that are formed, and this may govern the biological
outcome of ubiquitination (Deshaies and Joazeiro, 2009). Each E3 ubiquitin ligase only
uses a subset of these E2s and identifying these E2 enzymes has been a challenge. In pull
down/mass spectrometry experiments, E2s are usually not identified due to the weak
interaction between the ubiquitin conjugating enzyme and the ligase (Deshaies and
89
Joazeiro, 2009). The current method of identifying E2 –E3 partners is through screening
purified E2s to find ones that works best with the particular E3 ubiquitin ligase in an in
vitro experiment. This approach can be problematic for several reasons as there are only
a limited number of E2s that can be expressed, purified and stored as recombinant protein.
In addition, E2 and E3 enzymes that require post-translational modification to become
active would be excluded from these in vitro approaches (Deshaies and Joazeiro, 2009).
A physiological approach would be best to identify the proper E2-E3 pairs. Deshaies and
Joazeiro (2009) suggest an approach using in vivo crosslinking, and potentially
immunoprecipitation of the ubiquitin ligase and identification of the associated proteins
using mass spectrometry. This would identify proteins under physiological conditions,
and also account for post-translational modifications that may be required for activity or
binding of the E2-E3 partners.
The first E4 enzyme, UFD2, was discovered in yeast as a factor that binds to
ubiquitin and promotes chain elongation (Koegl et al., 1999). E4 enzymes were thought
to contain a U-box, such as UFD2 (Hoppe, 2005). However, not all E4 chain assembly
factors contain a U-box, and not all U-box proteins act as E4 ubiquitin chain assembly
factors; U-box proteins have been implicated as E3 ubiquitin ligases (Hatakeyama and
Nakayama, 2003). For example, p300 and Yin Yang 1 are a histone acetyl transferase
and a transcription factor, respectively; these proteins are E4 enzymes for p53 and do not
contain a U-box (Gronroos et al., 2004; Grossman et al., 2003; Sui et al., 2004).
Identification of E4 enzymes then becomes problematic since like substrates and E2
enzymes, they are weakly bound to the ubiquitin ligase. In addition, unlike E2 enzymes
that have already been identified and have a characteristic globular domain, a
90
characteristic domain for E4 enzymes has not yet been determined (Pickart and Eddins,
2004). A similar method used to identify of E2 enzymes could be used to identify E4
enzymes – a yeast two hybrid approach and in vivo crosslinking-immunoprecipitation.
However, characterizing the interaction could be problematic as substrates would also be
identified in these experiments. Candidate E4 enzymes would not be substrates, and
potential ubiquitination activity would have to be tested in a ubiquitination assay.
Characterization of the interactors.
Interacting proteins can broadly be classified into two categories: substrates and
cofactors. To characterize substrates one would compare protein levels in the Pirh2+/+
cells and tissues with those isolated from the Pirh2-/- background using western blot
analysis. One would expect higher protein levels for the candidate substrate in the Pirh2
-/- background. In addition, one would perform in vitro ubiquitination assays using
purified proteins, or transfecting into cell lines. Once the candidate substrate was verified,
one would map the interaction domains on Pirh2 and on the substrate by GST-pull down
experiments. One would also want to determine what type of ubiquitin linkages Pirh2
catalyzes on the substrate. One could take two approaches. The first is an in vitro
approach using ubiquitin constructs where a single lysine residue has been mutated to
arginine to prevent chain elongation at that residue; one would cotransfect Pirh2, the
substrate, and the ubiquitin mutant into cells. One would then immunoprecipitate the
substrate and perform a western blot for ubiquitin. One would compare each of the seven
lysine to arginine ubiquitin mutants to the wild-type ubiquitin to determine if a single
residue is preferred to create polyubiquitin chains. One could also treat Pirh2+/+ and
91
Pirh2-/- cells with proteasomal inhibitor and ubiquitin aldehyde (to inhibit
deubiquitinases), immunoprecipitate the substrate and compare the differences in
linkages found in the Pirh2+/+ cells relative to the Pirh2-/- cells. Pirh2 may not
necessarily target the substrate for ubiquitin-mediated proteolysis, but may change the
subcellular localization, or activity of the protein based on the ubiquitin linkage (Chen
and Sun, 2009). These experiments could determine whether Pirh2-mediated
ubiquitination is solely present to target substrates for degradation, or may have a more
complex role in affecting protein function in the absence of proteolysis.
Cofactors are likely to be identified in crosslinking/immunoprecipitation
experiments or yeast two hybrid experiments. They are also likely not to be found in the
substrate identification experiments. If ubiquitin conjugating enzymes are identified, the
important question then becomes identifying which substrates the E2 helps ubiquitinate.
Knockdown of the E2 in question using RNAi will help determine this as the Pirh2
substrates in question will increase. If one compares Pirh2-/- cells with control RNAi
with Pirh2+/+ cells with E2 RNAi and Pirh2-/- cells with RNAi, one can determine the
E2-Pirh2 substrates. Pirh2 substrates will be increased in all three experiments; since
each E2 will cooperate with several other ubiquitin ligases, targets of those ubiquitin
ligases will also be increased when treated with RNAi against the E2. However, the non-
specific targets will not be increased in the Pirh2-/- cells without RNAi against the E2.
For other cofactors, such as E4 enzymes, one can perform an in vitro ubiquitination assay
using GST-hPirh2 on total cell lysate. One can then titrate levels of GST-hPirh2 so that it
weakly catalyzes ubiquitination; by western blot analysis for ubiquitin, there should be a
faint smear near background levels. If a cofactor promotes Pirh2 activity, addition of this
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cofactor to the GST-hPirh2 ubiquitination reaction should increase the efficiency of the
reaction; i.e. using low levels of GST-hPirh2 with the cofactor should catalyze
polyubiquitination more efficiently. By western blot analysis polyubiquitination of the
cell lysate should be increased. One would also want to map the interaction domains on
the cofactor and Pirh2. Also, one would use RNAi against the cofactor to determine if it
preferentially increases polyubiquitination on specific Pirh2 substrates. RNAi against the
cofactor should increase the protein level of specific Pirh2 substrates, or may generally
increase the protein level of all substrates. Determining the cofactors that regulate Pirh2
expression may help determine which cell types and under which stress conditions Pirh2
activity may be important for the regulation of cellular processes.
Identification of the Pirh2 degron
The degron is the signal for a particular ubiquitin ligase to initiate degradation of
its substrate, and it is also where the ubiquitin ligase binds to the substrate (Deshaies and
Joazeiro, 2009). If the binding site for several Pirh2 substrates were known, one would
compare the substrates to look for sequence similarity in those binding regions. One
would then synthesize ten amino acid long peptides corresponding to the sequence
similarities between the Pirh2 binding sites. For example, if one found a fifteen peptide
long amino acid sequence that is similar among the substrates, one would synthesize five
peptides; the first peptide would correspond to amino acids one to ten, the second would
correspond to two to eleven, etc. One would spot these peptides onto a membrane, and
then perform a far western blot using Pirh2. One would determine which peptides Pirh2
binds to, and this would help us to identify the degron. To test the proposed degron, one
93
would mutate the degron in a substrate, and determine if the mutant degron abolishes
Pirh2-mediated ubiquitination and proteolysis and binding. Once one has confirmed the
degron, one can then use this sequence to potentially predict Pirh2 substrates rather than
experimentally identify them.
II. Summary of Findings: Pirh2 Regulation of Phosphoserine 15 p53
The Benchimol lab has previously published that Pirh2 participates in a negative
feedback loop with p53; under stress conditions, p53 induces Pirh2 expression, and Pirh2
targets p53 for ubiquitin mediated proteolysis. To further characterize Pirh2 regulation of
p53, we used the Pirh2-/- mouse generated by the Hakem lab. Splenocytes and
thymocytes have similar levels of basal p53 in both Pirh2+/+ and Pirh2-/- mice
suggesting that Pirh2 is not a major regulator of p53 in unstressed conditions (Fig. 3.1
B,C). After IR, splenocytes and thymocytes from Pirh2-/- mice show an increase in the
level of p53 and an increase in the level of phosphoserine 15 p53. The relative amount of
phosphoserine 15 p53 is higher in Pirh2-/- cells compared with Pirh2+/+ after
normalization to total p53 suggesting that Pirh2 modulates the level of phosphoserine 15
p53 (Fig. 3.2B). The relatively higher level of phosphoserine 15 p53 in Pirh2-/-
splenocytes and thymocytes has functional consequences since the induction of the pro-
apoptotic p53-target gene bax and induction of apoptosis occurs faster in the Pirh2-/- cells
(Fig. 3.3). To determine if high levels of Pirh2 affect p53 induction, we took cells that
stably express ectopic Pirh2 and measured the levels of p53 and p21 after adriamycin
treatment. We determined that ectopic expression of Pirh2 reduces induction of
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phosphoserine 15 p53 and p21 after DNA damage (Fig. 3.5). Hence, the activation of
p53 is facilitated by Pirh2-depletion and impeded by Pirh2 overexpression.
We also noted that Pirh2 coimmunoprecipitates with phosphoserine 15 p53 and
that in the absence of Pirh2, there appears to be reduced polyubiquitination of
phosphoserine 15 p53 (Fig 3.7). These results suggest that Pirh2 directly regulates
phosphoserine p53. We propose a model where under unstressed conditions Mdm2 and
Pirh2 can both degrade p53, but Mdm2 is the main regulator of p53. After DNA damage,
Mdm2 and p53 both are phosphorylated and this aids in the dissociation of the
Mdm2/p53 complex; Mdm2 no longer regulates p53 after DNA damage and p53 levels
increase. Pirh2 then becomes the main regulator of p53, and modulates the p53 response
(Figure 4.1).
There are many ubiquitin ligases that target p53 for ubiquitin mediated proteolysis:
Mdm2, Pirh2, Cop1, Synoviolin, CARPs, TOPORs, Trim24 and ARF-BP1 (Allton et al.,
2009; Chen et al., 2005a; Dornan et al., 2004b; Honda et al., 1997; Leng et al., 2003;
Rajendra et al., 2004; Yamasaki et al., 2007; Yang et al., 2007). Based on several
findings in the literature, it is generally accepted that Mdm2 is a critical regulator of p53
activity in unstressed cells (Brooks and Gu, 2006; Momand et al., 2000). The role of the
other ubiquitin ligases remains to be determined, but there are some differences among
the ubiquitin ligases. For example, under conditions of stress both Mdm2 and Cop1 can
be phosphorylated by ATM, and are no longer able to target p53 for ubiquitination
(Dornan et al., 2006; Maya et al., 2001). In contrast, both Pirh2 and CARPs can target
p53 for ubiquitination after DNA damage. CARP1/2 targets phosphoserine 20 p53 for
degradation, a modified form of p53 that only appears after DNA damage as a result of
A
B
Figure 4.1 Model for Pirh2 regulation of p53.
Under normal conditions, Mdm2 is the main regulator of p53. Pirh2 can regulate p53, but is a minor contributor (A). Under conditions of DNA damage, both p53 and Mdm2 are phosphorylated; these phosphorylation sites do not affect Pirh2, and it becomes a regulator of p53 (B).
95Mdm2
Mdm2
Pirh2
Pirh2
Ub
p53
p53
Ub
Ub
Ub
Ub
Ub
Ub
Ub
Ub
Ub
Ub
Ub
Ub
Ub
Ub
Ub
P
P
96
phosphorylation by the double strand break activated kinase Chk2 (Kruse and Gu, 2009a;
Yang et al., 2007). Based on the preliminary observations outlined above, it appears that
each ubiquitin ligase may target a subset of p53, or may target p53 in a subset of
conditions or cell types. These factors have yet to be determined, but pose an interesting
challenge for the field.
Future Directions
Dissecting the Functional Redundancy of p53 Ubiquitin Ligases
The issue of functional redundancy among the many p53 ubiquitin ligases is a
complex problem. To being resolving the question, one would look at the tissue and
developmental expression of each ubiquitin ligase to see whether there are differences in
the pattern of expression. This could suggest that there may be tissue or developmental
specific regulation of p53. It may also be possible that each ubiquitin ligase regulates a
different post-translationally modified forms of p53, different oligomeric forms, different
isoforms of p53 or mutant p53. Different post-translational modifications on p53 could
be a response to the exposure to various stresses that would cause differing post-
translational modifications. The p53 protein has been found to have several isoforms, the
Δ133p53 and the Δ40p53 isoforms each contain an N-terminal deletion that eliminates
the Mdm2 binding domain (Bourdon et al., 2005). Presumably these isoforms are not
regulated by Mdm2 and may be targeted by one of the other ubiquitin ligases. Cofactors
could influence the binding of the ubiquitin ligases to p53, the stability of the ubiquitin
ligase or the ability of the ubiquitin ligase to promote ubiquitination; expression of the
cofactors can be influenced by cell type and stress response.
97
Post-translational modifications on the E3 ubiquitin ligases can also influence
binding to p53, stability and may also influence the activity of the ubiquitin ligase. For
example phosphorylation of Mdm2 and Cop1 by ATM reduces their ability to target p53
for ubiquitination as well as phosphorylation of Pirh2 by calmodulin-dependent kinase II
decreases p53 ubiquitination (Dornan et al., 2006; Duan et al., 2007; Maya et al., 2001).
Post-translational modifications can be activating as neddylation of the cullin in the SCF
complex is required for ubiquitination activity (Saha and Deshaies, 2008). Post-
translational modifications may be stress-specific, and may be identified using
bioinformatics to predict phosphorylation sites or potentially by mass spectrometry.
There could also be an organism specific effect to regulating p53. Mdm2-/- mice
are embryonic lethal, and die by massive apoptosis preimplantation; the embryonic
lethality is fully rescued by deletion of p53 (Jones et al., 1995; Montes de Oca Luna et al.,
1995). In addition, amplification of Mdm2 has been found in tumours as a mechanism of
inactivating p53, and anti-cancer therapies are being developed to inhibit the Mdm2-p53
interaction (Onel and Cordon-Cardo, 2004; Vassilev et al., 2004). This finding strongly
supports that Mdm2 is a key regulator of p53 of in mice and humans. However, p53
orthologs are found in Drosophila Melanogaster and Caenorhabditis Elegans, but these
organisms do not have an Mdm2 ortholog (Brooks and Gu, 2006). An ortholog of one of
the other ubiquitin ligases may be important to regulate p53 in these organisms. Trim24
appears to be an important regulator of Drosophila p53 (Allton et al., 2009). Orthologs
of Pirh2 exist in Schizosaccharomyces pombe and Drosophila Melanogaster (Leng et al.,
2003).
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Non-Proteolytic Ubiquitination of p53
Two ubiquitin ligases have been identified that target p53 for non-proteolytic
ubiquitination. E4F1 catalyzes lysine 48 linked polyubiquitination of p53 on lysine 320.
This polyubiquitination occurs after DNA damage and promotes the cell cycle arrest
pathway (Le Cam et al., 2006). The ubiquitin ligase MSL2 catalyzes polyubiquitination
of p53 on lysines 351 and 357. MSL2 mediated polyubiquitination promotes cytoplasmic
translocation of p53 (Kruse and Gu, 2009b).
Lysine 48 linked polyubiquitination traditionally targets the substrate for
proteolysis (Xu et al., 2009). E4F1 mediated ubiquitination of p53 targets cell cycle
arrest promoters instead of promoting polyubiquitination (Le Cam et al., 2006). This
finding raises several questions: how does the E4F1 lysine 48 polyubiquitin chain evade
degradation, how does the polyubiquitin chain redirect p53 to cell cycle arrest promoters,
and whether E4F1 activity is required to induce cell cycle arrest.
MSL2 catalyzes polyubiquitination on lysines 351 and 357. The linkage that is
used to create these polyubiquitin chains is unknown and should to be determined (Kruse
and Gu, 2009b). In addition, how polyubiquitination of these lysines causes
relocalization of p53 to the cytoplasm should be determined. How MSL2 affects p53
biological activity also needs to be determined. MSL2 activity is not affected by DNA
damage. Key questions include whether high levels of MSL2 reduces the level of p53
transcriptional targets, or the amount of p53 bound to promoters. Also, p53 is usually a
nuclear protein and translocation to the mitochondria causes apoptosis (Kruse and Gu,
2009b; Mihara et al., 2003). How p53 translocates from the nucleus to the mitochondria
is still a question that needs to be resolved. Perhaps high levels of MSL2 may promote
99
transcription independent, p53-dependent apoptosis after DNA damage by relocalizing
p53 to the cytoplasm where it can then reach the mitochondria. Confocal microscopy
could determine whether high levels of MSL2 promotes p53 translocation to the
mitochondria. In addition, apoptotic assays could help determine whether the ectopic
expression of MSL2 increases p53-dependent apoptosis.
Pirh2-Mediated Ubiquitination of p53
We have determined that Pirh2 can catalyze ubiquitination of phosphoserine 15
p53. It is possible that Pirh2 can ubiquitinate other post-translationally modified forms of
p53 such as phosphoserine 6, 9, 20, 33, 36, 37, 46 and/or phosphothreonine 18, 55, 81.
Several modifications on p53 occur throughout the protein; one would focus on
modifications that occur in the N-terminus because Pirh2 binds to p53 in the DNA
binding domain and the C-terminal oligomerization domain, and modifications in these
regions may help to dissociate the Pirh2-p53 complex (Leng et al., 2003; Sheng et al.,
2008). In addition, we have shown that phosphorylation of serine 15 on p53 does not
affect Pirh2-mediated degradation. The set of experiments performed to characterize
Pirh2-mediated ubiquitination of phosphoserine 15 p53 can be used to determine whether
Pirh2 can target other p53 phosphoforms. One would expect an increase in
phosphorylated p53 in Pirh2-/- cells, and a decrease in phosphorylated p53 in Pirh2
overexpressing cells. One would also perform two dimensional gel electrophoresis on
irradiated Pirh2+/+ and Pirh2-/- samples, and use western blot for total p53. If Pirh2
regulates phosphorylated p53, one would expect to see an accumulation of the acidic
(phosphorylated) forms of p53 in the Pirh2-/- irradiated cells. The two dimensional gel
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electrophoresis results could also determine whether Pirh2 regulates forms of p53 that
contain multiple phosphorylation sites. Should Pirh2 regulate other forms of activated
p53, it would support the hypothesis that Pirh2 may modulate p53 activation.
In addition, one can look at whether post-translational modifications in the DNA
binding domain and the tetramerization domain can inhibit ubiquitination or binding of
Pirh2 to p53. One would also determine whether post-translational modification of the
C-terminal lysines by acetylation, neddylation or methylation can inhibit ubiquitination or
binding of p53 by Pirh2. Understanding the conditions that determine when Pirh2
regulates p53 will contribute to our understanding of p53 ubiquitination and may begin to
resolve the issue of functional redundancy.
One could also determine whether Pirh2 regulates p53 under different stress
conditions. For example, one could use hypoxic conditions, ER stress, UV irradiation,
and nutritional stress to determine whether Pirh2 affects p53 levels. This can be done by
looking at total p53, post-translationally modified p53, and two dimensional gel
electrophoresis.
There are seven possible lysines on ubiquitin that can form polyubiquitin chains
(Xu et al., 2009). One would determine which lysines on ubiquitin that Pirh2 uses to
catalyzes polyubiquitin chains on p53. One would do this using seven mutant ubiquitin
construct where six lysines are mutated to arginine. One would peform an in vivo
ubiquitination assay using p53 as a substrate, Pirh2 as the ubiquitin ligase, and the mutant
ubiquitin. Laddering of p53 on a Western blot for p53 will only be evident if Pirh2 can
catalyze polyubiquitination using the available lysine on ubiquitin. The polyubiquitin
chain that is catalyzed on p53 could suggest that Pirh2 may target p53 under different
101
conditions. For example, polyubiquitination through lysine 63 could suggest a role in
signalling or subcellular localization, or polyubiquitination through lysine 29 could
suggest a role in ERAD (Xu et al., 2009).
Knock down of several ubiquitin ligases by RNAi increases p53 levels and cell
cycle arrest and apoptosis (Chen et al., 2005a; Dornan et al., 2004b). It is possible that
the ubiquitin ligases may work synergistically to target p53 for ubiquitin-mediated
proteolysis. For example, Mdm2 binds to p53 in the N-terminal domain, and this binding
site does not overlap with the Pirh2 binding site in the oligomerization domain or the
DNA binding domain; it is possible that both ubiquitin ligases can simultaneously bind to
p53. Whether ubiquitin ligases act synergistically can be tested by cotrasfecting siRNAs
to two ubiquitin ligases and determining whether p53-dependent cell cycle arrest or
apoptosis are increased beyond the sum of the single transfections. The dual binding of
these ubiquitin ligases can also be confirmed by coimmunoprecipitation. Possible
synergism between the ubiquitin ligases may suggest that one of the ubiquitin ligases may
function as an E4 or cofactor for the ubiquitination reaction. A cofactor role for one of
the ubiquitin ligases could suggest a cooperative relationship between the ligases in
regulating p53.
III. Pirh2 and Cancer
Pirh2 has been found to be overexpressed in lung cancer, and the overexpression
of Pirh2 does not correlate with p53 mutation status (Duan et al., 2004). In contrast,
Mdm2 has been found to be overexpressed in osteosarcomas, and is a mechanism to
inactivate wild-type p53 by targeting it for degradation (Onel and Cordon-Cardo, 2004).
102
This suggests that p53 may not be the key target for Pirh2-mediate ubiquitination in lung
cancer. In addition, Pirh2 overexpression has also been found in prostate cancer and head
and neck cancers. In the head and neck cancers, overexpression of Pirh2 correlates with
low expression of p27, a G1 cell cycle inhibitor. RNAi against Pirh2 in a head and neck
squamous cell carcinoma cell line increases the expression of p27 and increases the cell
doubling time. High expression of Pirh2 and low expression of p27 in the head and neck
cancers correlate with low survival rates compared with low expression of Pirh2 and high
expression of p27 respectively. This study classified patients into four groups: low Pirh2
and high 27, low Pirh2 and low p27, high Pirh2 and low p27, high Pirh2 and high p27.
The patients with low Pirh2 and high p27 have the best prognosis followed by high Pirh2
and high p27, low Pirh2 and low p27, and high Pirh2 and low p27 have the worst
prognosis (Shimada et al., 2009). Since patients with high Pirh2 and low p27 have a
worse prognosis than patients with low Pirh2 and low p27, these results suggest that
Pirh2 is not only regulating p27 in head and neck squamous cell carcinomas, but has
other targets that contribute to oncogenesis.
In prostate cancer, Pirh2 has been found to be overexpressed and high levels of
Pirh2 expression correlate with the presence of bone metastases. Pirh2 targets HDAC1 in
prostate cancers, HDAC1 is an androgen receptor corepressor. Expression of Pirh2 leads
to reduced levels of HDAC1 resulting in increased transcription of the androgen receptor
target gene prostate specific antigen (PSA) (Logan et al., 2006). In contrast, observations
made when Pirh2 was initially characterized as an ARNIP (androgen receptor N-terminal
interacting protein) suggest that Pirh2 does not affect androgen receptor transactivation.
Cotransfection of Pirh2 and the androgen receptor did not affect androgen receptor
103
transactivation in luciferase assays (Beitel et al., 2002). Beitel et al. (2002) used monkey
cells and mouse cells (Cos-1, CV-1 and NSC-34) while Logan et al (2006) performed
their cotransfection experiments in human 293T cells. While Pirh2 is a highly conserved
protein, perhaps the region of Pirh2 that binds HDAC1 may not be conserved across the
species or the binding site for Pirh2 on HDAC1 is not conserved across the species
leading to the differences in results between the two groups (Beitel et al., 2002).
Overexpression of Pirh2 leads to increased transcription by the androgen receptor and
increased levels of PSA mRNA (Logan et al., 2006). However, when Pirh2 is transfected
into the androgen receptor prostate cancer cell line LNCaP, PSA mRNA levels are
increased while PSA protein levels are decreased since ε-COP is required for PSA
secretion; increased levels of Pirh2 leads to degradation of ε-COP (Maruyama et al.,
2008). PSA is a serine protease that is responsible for liquefying seminal fluid. High
levels of PSA are used as a marker of prostate cancer in the PSA test. High levels of PSA
have not been shown to affect oncogenesis; PSA is an androgen receptor responsive gene,
and high androgen receptor activity is linked to the early stages of prostate cancer
development (Kaarbo et al., 2007; Lilja et al., 2008). If overexpression of Pirh2 is an
early event in prostate cancer, this could hinder the effectiveness of the PSA test for
predicting prostate cancer. As well, these cancers are statistically more aggressive, and
early detection of these cancers could help survival (Logan et al., 2006). Pirh2 also
regulates HDAC1, and derepression of HDAC1 target genes may also contribute to the
progression of cancer. Experiments need to determine whether inhibiting Pirh2
degradation of HDAC1 can potentially be developed as a therapeutic option for prostate
cancer.
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Recent observations have suggested that Pirh2 could act as an oncogene or a
tumour suppressor gene. For example, Pirh2 is overexpressed in head and neck
squamous cell carcinomas and Pirh2 degradation of the p27 G1 cell cycle inhibitor
appears to contribute to tumorigenesis (Hattori et al., 2007; Shimada et al., 2009). Pirh2
degradation of ε-COP decreases PSA protein levels, and may prevent early detection of
prostate cancer (Maruyama et al., 2008). In these examples, Pirh2 would appear to act as
an oncogene. In contrast, Pirh2 degrades HDAC1; in breast cancer, overexpression of
HDAC1 in the breast cancer cell line MCF-7 causes increased cell proliferation and
colony formation (Kawai et al., 2003; Logan et al., 2006). These observations suggest
that Pirh2 has the potential to also mediate tumour suppression. Determining the
differences between cell types and what determines which substrates Pirh2 targets would
be valuable in understanding how Pirh2 functions as a ubiquitin ligase, and how
deregulation of Pirh2 contributes to cancer. Targeting Pirh2 activity in cancer may prove
to be a useful therapeutic target or potentially a prognostic marker.
105
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