identification and characterization of small molecule …...ii identification and characterization...
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Identification and Characterization of Small Molecule Inhibitors of Polynucleotide Kinase 3'-Phosphatase
by
Nathalie Moatti
A thesis submitted in conformity with the requirements for the degree of Master of Science
Department of Molecular Genetics University of Toronto
© Copyright by Nathalie Moatti 2012
ii
Identification and Characterization of Small Molecule Inhibitors of
Polynucleotide Kinase 3’-Phosphatase
Nathalie Moatti
Master of Science
Department of Molecular Genetics
University of Toronto
2012
Abstract
DNA lesions arise constantly in cells and are repaired by a variety of DNA repair pathways.
Polynucleotide kinase 3’-phosphatase (PNKP) aids repair by phosphorylating 5’-hydroxyl DNA
termini and dephosphorylating 3’-phosphate DNA termini for the completion of repair by DNA
ligases. This activity is critical in vivo because DNA breaks do not usually possess ligatable
termini.
PNKP knockdown sensitizes cells to several DNA damaging agents, including the topoisomerase
I (TOP1) inhibitor camptothecin - analogs of which are being developed into chemotherapeutic
drugs - because the resolution of stalled TOP1-DNA complexes requires processing by PNKP.
We hypothesize that small molecule inhibitors of PNKP could bolster the effects of radio- and
chemotherapies on cancer cells.
I have identified eight compounds that effectively inhibit human PNKP and, with reduced
potency, T4 PNK in vitro. These compounds act by reversibly inhibiting the substrate-enzyme
interaction but they do not appear to sensitize U2OS cells to camptothecin.
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Acknowledgments
I would like to thank my supervisor, Dr. Daniel Durocher, for his guidance, encouragement, and
support during my time in the lab – without a doubt it helped me get through the long periods of
troubleshooting. I would also like to thank the members of the Durocher lab, not only for their
constant willingness to help, but also simply for being great people to work with. For many
reasons, I am often reminded of how grateful I am to have joined this lab. I am indebted to
Thomas Sun, Fred Vizeacoumar, and Alessandro Datti of the SMART robotics facility for all of
their help with my screen. I also owe many thanks to my supervisory committee members, Drs.
Frank Sicheri and Peter Roy, as well as our collaborators in Dr. Mark Glover’s lab at the
University of Alberta and Dr. David Uehling at the Ontario Institute for Cancer Research. Lastly,
thanks to my family and friends for their support and great efforts to understand what I’ve been
working on these past two years.
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Table of Contents
Acknowledgments.......................................................................................................................... iii
Table of Contents ........................................................................................................................... iv
List of Tables ................................................................................................................................. vi
List of Figures ............................................................................................................................... vii
List of Appendices ......................................................................................................................... ix
1 Introduction .................................................................................................................................1
1.1 DNA repair is critical for maintaining genomic stability and preventing tumorigenesis ....1
1.2 Targeted cancer therapies, like poly-ADP ribose polymerase inhibitors, kill tumor
cells with fewer side effects than chemotherapy .................................................................2
1.3 Polynucleotide kinase 3’-phosphatase processes certain types of damaged DNA
termini to allow for the completion of DNA repair by ligation ...........................................4
1.4 The importance of PNKP for DNA repair during development ..........................................7
1.5 Inhibitors of PNKP’s phosphatase activity would be useful tools for research and
clinical applications .............................................................................................................8
2 Results .......................................................................................................................................10
2.1 Development of screen conditions, Version 1 ...................................................................10
2.2 Development of screen conditions, Version 2 ...................................................................17
2.3 Homogeneous time-resolved FRET-based screen .............................................................24
2.4 Putative inhibitors of hPNKP.............................................................................................26
2.5 Validation and IC50 measurement of putative hPNKP inhibitors by HTRF ......................27
2.6 Optimization of gel-based phosphatase assay ....................................................................29
2.7 Measurement of IC50 values by gel-based phosphatase assay ...........................................30
2.8 Structure activity relationship analysis ..............................................................................31
2.9 Optimization of fluorescence polarization-based substrate-binding assay ........................37
2.10 Substrate-binding assay confirms that compounds prevent a 3'-phosphorylated
oligonucleotide from binding to hPNKP ...........................................................................38
2.11 Inhibitors of hPNKP display different degrees of reversibility .........................................39
2.12 Inhibitors of hPNKP can act on T4 PNK with reduced potency .......................................41
2.13 Inhibitors of hPNKP do not show a clear effect in U2OS cells co-treated with
camptothecin ......................................................................................................................44
v
3 Discussion and future directions ...............................................................................................51
4 Materials and Methods ..............................................................................................................57
4.1 Transformation of E. coli. ..................................................................................................57
4.2 SDS-PAGE. .......................................................................................................................57
4.3 Quantitation of protein .......................................................................................................57
4.4 Cloning of human PNKP-D171A ......................................................................................58
4.5 Purification of human PNKP and PNKP-D171A ..............................................................58
4.6 Purification of T4 DNA Ligase..........................................................................................60
4.7 Preparation of 3'-phosphorylated double-stranded DNA substrates. .................................63
4.8 Measurement of PNKP-dependent ligation by homogeneous time-resolved
fluorescence HTRF) in a 384-well format, Version 1. ......................................................63
4.9 Measurement of PNKP-dependent ligation by HTRF in a 384-well format, Version 2. ..64
4.10 Measurement of PNKP-independent ligation by HTRF in a 384-well format, Versions
1 and 2. ...............................................................................................................................64
4.11 Screening............................................................................................................................65
4.12 Calculation of z’ factor, signal-to-noise (S/N), signal-to-background (S/B), and
percent error. ......................................................................................................................66
4.13 Preparation of polyacrylamide sequencing gels. ...............................................................67
4.14 Phosphatase gel assay. .......................................................................................................67
4.15 Fluorescence polarization (FP) substrate binding assay. ...................................................68
4.16 Measurement of the Kd of hPNKP-D171A in the FP substrate binding assay. .................68
4.17 Reversibility assay. ............................................................................................................69
4.18 Tissue culture. ....................................................................................................................69
4.19 ATP Lite cell viability assay. .............................................................................................70
Tables .............................................................................................................................................71
References Cited ............................................................................................................................70
Appendices .....................................................................................................................................75
vi
List of Tables
Table 1a Buffers ............................................................................................................................ 71
Table 2a Sequences of oligonucleotide substrates (5' to 3') ........................................................ 73
Table 1 Original hit compounds ................................................................................................... 74
Table 2 Analog compounds .......................................................................................................... 76
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List of Figures
Figure 1.3.1 PNKP is critical for the repair of DNA breaks with 3’-phosphate and/or 5’-hydroxyl
termini ............................................................................................................................................. 7
Figure 2.1.1 Optimization of HTRF screen conditions ................................................................ 13
Figure 2.1.2 Optimization of the order of addition, stability of reagents at room temperature, and
hPNKP concentration for HTRF screen conditions ...................................................................... 15
Figure 2.1.3 Screening the Prestwick Chemical Library confirmed that the optimized conditions
were amenable to high-throughput screening ............................................................................... 17
Figure 2.2.1 Further optimization of the HTRF assay was undertaken to find non-saturating
conditions with high statistical validity ........................................................................................ 20
Figure 2.2.2 The T4 Ligase and hPNKP concentrations were optimized to prevent the ligation
reaction from causing a kinetic lag ............................................................................................... 22
Figure 2.2.3 Optimization of DNA concentration, ligation product length, and acceptor
fluorophore concentration to maximize signal-to-background ..................................................... 24
Figure 2.3.1 Screening of the OICR HTS chemical collection under primary and secondary
screen conditions to identify putative hPNKP inhibitors .............................................................. 26
Figure 2.5.1 Several putative inhibitors were validated as inhibitors of hPNKP with a range of
potencies, while others could not be reconfirmed ........................................................................ 28
Figure 2.6.1 A gel-based assay was developed to directly measure the dephosphorylation
activity of PNKP ........................................................................................................................... 30
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Figure 2.7.1 Dose response experiments were performed for each of the five original hits using
the gel assay, and IC50 values were measured .............................................................................. 31
Figure 2.8.1 Structure activity relationship analyses of the heterocyclic ring and the fragment
common to OICR7 and OICR9..................................................................................................... 33
Figure 2.8.2 Structure activity relationship analyses using analogs of OICR79 ......................... 35
Figure 2.8.3 Structure activity relationship analysis of OICR79 uncovered two modifications
that did not abolish inhibition ....................................................................................................... 37
Figure 2.10.1 Fluorescence polarization-based direct binding assays were carried out to
determine if the compounds could compete with a 3’-phosphorylated oligonucleotide substrate
for binding to hPNKP-D171A ...................................................................................................... 39
Figure 2.11.1 Reversibility assays demonstrated that most of the compounds were at least
slightly reversible .......................................................................................................................... 41
Figure 2.12.1 Gel-based assays conducted with T4 PNK demonstrate that some, but not all, of
the hPNKP inhibitors are capable of inhibiting this distant ortholog ........................................... 43
Figure 2.13.1 The viability of U2OS cells co-treated with 5 nM camptothecin (CPT) was not
significantly impaired relative to their DMSO-treated counterparts ............................................ 45
Figure 4.6.1 hPNKP and T4 Ligase were purified from BL21 (DE3) CodonPlus E. coli cells .. 62
Figure 4.16.1 FP reactions were conducted with hPNKP for proof of concept and to establish
optimal reaction conditions ........................................................................................................... 69
ix
List of Appendices
PNKP inhibitors ............................................................................................................................ 75
Analogs that did not effectively inhibit PNKP ............................................................................. 76
1
1 Introduction
1.1 DNA repair is critical for maintaining genomic stability and preventing tumorigenesis
DNA lesions arise constantly as a consequence of cellular metabolism or in response to
genotoxic agents, requiring immediate attention so that the cell may divide safely (Branzei and
Foiani, 2008). In order to repair these lesions effectively, there are a variety of DNA repair
pathways available depending on the nature of the damage, which allow cells to continue
functioning normally (Branzei and Foiani, 2008). However, if these pathways fail to mend even
one DNA lesion, the cell and its lineage can incur mutations or larger-scale genomic aberrations
with potentially disastrous consequences including cell death, especially if that break is double-
stranded (Jackson, 2002).
Cancer is a disease that arises when a cell loses its ability to control proliferation, causing the cell
to divide incessantly and form one or more tumors (Croce, 2008). This unrestrained growth is
borne from genomic translocations and/or mutations that activate oncogenes and disable tumor
suppressor genes (Croce, 2008). The term “oncogene” signifies a gene whose protein product
originally played a role in the regulation of proliferation, differentiation, or apoptosis, but has
since become mutated in such a way that causes it to promote or allow constitutive growth,
which can lead to tumorigenesis (Croce, 2008). Conversely, tumor suppressor genes act to
stabilize the genome and limit proliferation when the cell is stressed; for example, when faced
with a genotoxic agent (Weinberg, 1991). Proteins involved in the DNA damage response
(DDR) fall into this latter category, as they act to maintain the integrity of the genome and
perform checkpoints for cell division, which arrest the cell cycle until genomic stability is
recovered.
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2
In cases of familial breast, ovarian, and prostate cancer, 5-20% are traced to heterozygotic
germline mutations in BRCA1 or BRCA2 (Venkitaraman, 2002), two tumor suppressor genes that
are critical for homologous recombination (HR) - the least mutagenic form of double-strand
break repair (DSBR). Although BRCA1/2+/-
cells are capable of efficiently and accurately
repairing double-strand breaks (DSBs), it has been proposed that tumorigenesis is initiated by a
loss-of-heterozygosity event, wherein the sole functional BRCA1/2 allele acquires a nonsense
mutation (Miki et al., 1994; Wooster et al., 1995). This complete loss of BRCA1/2 activity
creates genomic instability in the cell and acts as a “driver mutation” by promoting the
development of new mutations in other genes, some of which are likely to exacerbate the
problem (Shah, 2008). In fact, it has been suggested that the inactivation of DDR factors is a
critical step in the formation of all cancers, as the aberrant growth of cancer cells creates
excessive DNA replication stress, which would normally activate the DDR and trigger cell cycle
arrest or cell death (Jackson and Bartek, 2009).
1.2 Targeted cancer therapies, like poly-ADP ribose polymerase inhibitors, kill tumor cells with fewer side effects than chemotherapy
When it comes to treating cancer, there are two main types of drugs available – chemotherapies
and targeted therapies. Chemotherapies include DNA damaging agents like cisplatin, which acts
by creating DNA inter-strand cross-links and invokes a number of pathways – cell cycle arrest
and p53-induced apoptosis, for instance – and results in preferential killing of rapidly diving
cells (Siddik, 2003). Consequently, these drugs effectively eliminate cancer cells but they also
harm the body’s naturally rapidly-diving cells, such as hair and gut cells, resulting in a slew of
unwelcome side effects. On the other hand, targeted therapies work on the basis of the specific
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genetic aberrations present in a given patient’s cancer cells so that they theoretically harm only
the abnormal cells and cause minimal side effects to healthy cells, thus creating a much larger
treatment window (Sawyers, 2004). One class of targeted therapies that has garnered massive
attention over the last few years is that of poly-ADP ribose polymerase (PARP) inhibitors.
PARP-1 is an enzyme with myriad roles in vivo, one of which is in the detection and repair of
single-strand breaks (SSBs), formed directly or as a product of base excision repair (BER;
Helleday, 2011; Satoh and Lindahl, 1992). When a base is damaged, the removal of it and a few
adjacent bases forms an unprotected SSB, which is a substrate for rapid recognition and transient
binding by PARP-1 (Strom et al., 2011). Upon binding to the SSB, PARP-1 is activated and
recruits XRCC1-Ligase III to initiate single-strand break repair (SSBR; El-Khamisy et al., 2003).
Importantly, PARP-1 is also hypothesized to mediate a distinct pathway of replication repair by
enabling the restart of stalled replication forks (Bryant et al., 2009). Inhibitors of PARP1 have
shown promise in the treatment of BRCA1/2 defective cancers because PARP-1 and BRCA1/2
exhibit a synthetic lethal relationship; that is, cells can survive the inactivation of PARP-1 or
BRCA1/2 alone, but they die when the two enzymes are lost concurrently (Bryant et al., 2005;
Farmer et al., 2005). As such, PARP inhibitors are effective in killing BRCA1/2 cancer cells,
while doing minimal harm to normal, BRCA1/2 positive cells (Kling, 2009). Thanks to the
promising results seen in tissue culture (Bryant et al., 2005; Farmer et al., 2005), the effects of
several PARP inhibitors, such as Olaparib (Astra Zeneca, UK), Veliparib (Abbott Laboratories,
USA) and Iniparib (Sanofi Aventis, France), on BRCA1/2 tumors are currently being investigated
in clinical trials. Furthermore, because the efficacy of PARP inhibitors in BRCA1/2 cancer cells
is thought to be due to the dual loss of PARP- and HR-mediated replication fork-restart
(Helleday, 2011), there is reason to believe that PARP inhibitors can be useful in treating other
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HR-deficient “BRCA-like” cancers (Anders et al., 2010) or as potentiators of chemo- and radio-
therapy (Zaremba and Curtin, 2007).
In 2008, a siRNA-based study conducted by Turner et al. identified genes whose silencing would
sensitize CAL51 metastatic mammary adenocarcinoma cells (Gioanni et al., 1990) to the potent
PARP inhibitor KU0058948 (Farmer et al., 2005). The CAL51 cells employed by the authors
were diploid, BRCA1/2-wild type (WT) and TP53-WT, which they reasoned would facilitate the
identification of new determinants of sensitivity to PARP inhibitors with minimal interference
from other somatic mutations (Turner et al., 2008). Their analysis revealed that, out of the six on-
target hits identified, loss of polynucleotide kinase 3’-phosphatase (PNKP) was the most
effective in sensitizing CAL51 and cervical carcinoma HeLa cells to PARP inhibition, with a
striking 63-fold increase in sensitivity to the PARP inhibitor (Turner et al., 2008). These results
strongly suggest that inhibitors of PNKP could also be promising drugs for the treatment of
BRCA1/2 or HR-deficient cancers as monotherapies, or as sensitizers to PARP inhibitors,
chemotherapy, and/or radiation therapy.
1.3 Polynucleotide kinase 3’-phosphatase processes certain types of damaged DNA termini to allow for the completion of DNA repair by ligation
Mammalian PNKP was first identified in the 1980s and subsequently characterized in the late
1990s (Habraken and Verly, 1983; Karimi-Busheri et al., 1999; Pheiffer and Zimmerman, 1982).
Like its well-known T4 counterpart, mammalian PNKP contains a 5’-kinase domain and a 3’-
phosphatase domain - together termed the “catalytic domain” – which enable this enzyme to
phosphorylate the 5’- ends of DNA and de-phosphorylate the 3’-ends of DNA (Bernstein et al.,
2005). However, unlike T4 PNK, mammalian PNKP also has a forkhead associated (FHA)
domain, which is responsible for recruiting PNKP to DNA breaks (Bernstein et al., 2005).
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FHA domains are common among a number of DNA repair proteins and are responsible for a
range of protein-protein interactions owing to their affinity for phosphothreonine, but typically
not phosphoserine, residues (Mohammad and Yaffe, 2009). In the case of PNKP, the FHA
domain mediates interactions with X-ray repair cross complementing group 1 (XRCC1) and X-
ray repair cross complementing group 4 (XRCC4), which function as scaffolding proteins during
SSBR and the non-homologous end-joining (NHEJ) arm of DSBR, respectively, to recruit
several DNA repair factors to the site of the break, including PNKP (Lee et al., 2000;
Whitehouse et al., 2001). PNKP is also recruited by XRCC1 to DSBs in the alternative end-
joining pathway of DSBR (Audebert et al., 2004), but not HR (Karimi-Busheri et al., 2007).
The interaction between the PNKP FHA domain and XRCC1 is rather unique, as the FHA binds
to a vicinal phosphothreonine-phosphoserine motif in XRCC1 (XRCC1; Ali et al., 2009), which
is generated by casein kinase 2 (CK2) upon DNA damage (Loizou et al., 2004). Conversely, the
interaction between the FHA domain and CK2-phosphorylated XRCC4 appears to occur via a
specific phosphothreonine residue in XRCC4, with residues at positions +1 and -2 relative to the
pThr (pT+1 and pT-2, respectively) contributing to binding specificity (Chappell et al., 2002;
Koch et al., 2004). Intriguingly, for both XRCC1 and XRCC4, the FHA domain exhibits a lack
of selectivity for the pT+3 residue, which is unique because pT+3 specificity is a feature of all
other FHA domains studied to date (Bernstein et al., 2005; Pennell et al., 2010).
Additionally, it has been shown that the PNKP FHA binds to XRCC1 with a stoichiometry of 2:1
in a hierarchical and cooperative fashion: binding of the vicinal pSer/pThr motif by one PNKP
FHA facilitates the interaction of a second PNKP FHA with another CK2-phosphorylated
threonine nearby (Ali et al., 2009); presumably, this 2:1 stoichiometry aids in increasing the
processivity of the XRCC1-based multiprotein complex at damaged SSBs (Ali et al., 2009).
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Aside from DSBs mended by HR, the DNA end-processing functions of PNKP are critical for
the repair of most DNA breaks because all known DNA polymerases and ligases require 5’-
phosphate and 3’-hydroxyl termini as substrates (Doherty and Suh, 2000), but most SSBs and
DSBs are “dirty”, and thus require processing to produce conventional termini before repair can
be completed (Caldecott, 2008). Specifically, ionizing radiation, reactive oxygen species (ROS),
the process of BER, and Topoisomerase I (Top1) poisons such as camptothecin (CPT) frequently
create 5’-OH and/or 3’-phosphate groups (Figure 1.3.1; Caldecott, 2008). Accordingly, it has
been shown that the recruitment of PNKP to SSBs by XRCC1 is crucial for timely SSBR in cells
following oxidative damage (Breslin and Caldecott, 2009). Notably, the importance of PNKP in
SSBR was attributed to its 3’-phosphatase activity because overexpression of wild-type but not
phosphatase-dead PNKP was sufficient to restore rapid SSBR when the PNKP-XRCC1
interaction was abolished (Breslin and Caldecott, 2009).
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Figure 1.3.1 PNKP is critical for the repair of DNA breaks with 3’-phosphate and/or 5’-hydroxyl
termini. Upon detection of a damaged base (top), APE1 creates an apurinic/apyrimidinic (AP) site with
both types of damaged ends. Reactive oxygen species (ROS; middle), produced by exogenous sources or
as a natural consequence of ATP production, attack the sugars in DNA, which often leads to 3’-phosphate
ends as well as 5-‘hydroxyl ends to a lesser extent. The stalling of topoisomerase I (Top1; bottom), due to
poisons like camptothecin, leads to detection and processing by tyrosyl-DNA phosphodiesterase 1
(Tdp1), which generates one or both types of damaged termini via removal of Top1. Processing of these
breaks by PNKP creates conventional DNA termini, which allows the appropriate polymerases and
ligases to complete the repair process.
1.4 The importance of PNKP for DNA repair during development
During development, the rapid proliferation of stem and progenitor cells creates a great deal of
oxidative stress for the DNA replication machinery and this can lead to mutations and/or cell
death if replication errors are not dealt with appropriately (O'Driscoll and Jeggo, 2008). It has
been proposed that neuroprogenitor cells are particularly sensitive to DSBs, and an accumulation
of DNA damage can induce large-scale apoptosis in this population, resulting in defective
neuronal development; accurate DNA repair is therefore critical at this early stage (Ciccia and
Elledge, 2010). This hypothesis is thought to explain why a number of neurological disorders
have been attributed to mutations in DNA repair proteins; for example, defects in ataxia
telangiectasia mutated (ATM) are responsible for ataxia telangiectasia (Savitsky et al., 1995),
and ataxia oculomotor apraxia is due to a defect in Aprataxin (APTX; Moreira et al., 2001).
8
In line with the findings of Breslin and Caldecott (2009), which outlined the importance of the
phosphatase activity of PNKP in maintaining rapid SSBR after oxidative stress, mutations in
PNKP have been identified as the origin of the autosomal recessive disorder MCSZ
(microcephaly with seizures; Shen et al., 2010). MCSZ is characterized by microcephaly, early-
onset seizures, developmental delays, and variable behavioural problems (Shen et al., 2010).
Furthermore, lymphocytes from MCSZ patients were found to be significantly impaired in their
abilities to repair hydrogen peroxide- and CPT-induced damage (Shen et al., 2010).
1.5 Inhibitors of PNKP’s phosphatase activity would be useful tools for research and clinical applications
The body of research surrounding PNKP points to its importance in effecting rapid and accurate
DNA repair in the SSBR and NHEJ pathways. Taking into consideration that NHEJ is the main
method of DSBR in vivo (Mao et al., 2008), it follows that inhibiting PNKP would severely
hinder most SSBR and DSBR, which could prove to be even more effective than PARP
inhibitors in eliminating HR-impaired tumor cells. Furthermore, evidence for the sensitization of
A549 lung cancer cells to gamma radiation and camptothecin by stable down-regulation of
PNKP (Rasouli-Nia et al., 2004) makes a strong case for the potential to increase the efficacy of
certain classic cancer therapies – such as radiation therapy and Top1 poisons – by combining
them with PNKP inhibitors (Bernstein et al., 2008). Toward this end, as well as to generate new
tools for the study of PNKP, my work has focused on the identification of PNKP inhibitors
through high-throughput screening, and characterization with follow-up assays.
In order to query a variety of chemical structures, I developed a high-throughput screen based on
homogeneous time-resolved fluorescence resonance energy transfer (HTR-FRET or HTRF)
technology, which measured PNKP-dependent ligation of 3’-phosphorylated DNA substrates.
9
Importantly, I used 3’-phosphorylated DNA to search for inhibitors of PNKP’s 3’-phosphatase
activity as opposed to its 5’-kinase activity because 3’-phosphates are far more common than 5’-
hydroxyls at damaged DNA ends in vivo (Buchko and Weinfeld, 1993; Coquerelle et al., 1973;
Henner et al., 1982). Also, despite the fact that Ape1 and Aptx are capable of mending 3’-
phosphates, it has been demonstrated that their phosphatase activities pale in comparison to that
of PNKP (Reynolds et al., 2009; Wiederhold et al., 2004) – specifically, 3’-phosphates are
removed by PNKP at a rate at least ten-fold greater than that of Aptx and nearly 1000-fold
greater than that of Ape1 (Takahashi et al., 2007; Wiederhold et al., 2004).
Primary and secondary HTRF-based screening of more than 142,000 compounds yielded nine
hits, five of which still exhibited inhibition of PNKP when obtained commercially and tested
with freshly dissolved stocks. I measured the relative potencies of these five compounds by
determining the concentration of compound at which dephosphorylation is inhibited by 50% -
termed an “IC50 value” - with HTRF- and gel-based assays. In addition to these five “hits”, I
acquired several analog compounds from commercial sources and tested their potency in order to
perform structure-activity relationship analyses. I further characterized the compounds and active
analogs by assessing their degree of reversibility and abilities to inhibit the distant ortholog T4
PNK. Finally, I assessed the ability of the inhibitors to sensitize U2OS cells to CPT, but they did
not appear to significantly reduce the viability of CPT-treated cells compared to control cells.
The sum of this work reinforces that the five original hits identified, as well as the three active
analog compounds, are true inhibitors of human PNKP and may prove useful in the development
of biologically active PNKP inhibitors for research or clinical purposes, but further study will be
required to determine their effectiveness in cells.
10
2 Results
2.1 Development of screen conditions, Version 1
In an effort to develop a high-throughput assay for measuring the phosphatase activity of PNKP,
several techniques were investigated in our lab by Marella Canny before she found that
homogenous time-resolved fluorescence resonance energy transfer (HTR-FRET or HTRF)
provided high signal-to-background ratios and low variability. HTRF is a FRET-based
technology that employs a Europium cryptate as a donor fluorophore capable of emitting
fluorescence up to 100 times longer than more common donor fluorophores like Cy3 (Bazin et
al., 2002). Because of this long-lived donor emission, the delay between donor excitation and
measurement of acceptor fluorescence is much longer than it would be in a standard FRET
measurement, and this is beneficial because it allows any short-lived background fluorescence to
dissipate before the measurement is taken, which makes for stronger signal-to-background ratios
(Bazin et al., 2002).
The use of cross-linked allophycocyanin (symbolized “XL-665”) as the acceptor fluorophore is
the second feature that makes HTRF unique because the Europium cryptate/XL-665 FRET pair
has an exceptionally large Forster distance of 9 nm, which means that they exhibit a much higher
efficiency of energy transfer (Bazin et al., 2002). The Europium cryptate is conjugated to an anti-
2,4-dinitrophenol (DNP) antibody (“DNPK”), and XL-665 is conjugated to a streptavidin
molecule (“SAXL”) so that the two can be targeted to molecules of interest that are tagged with
DNP or biotin, respectively. In order to measure the phosphatase activity of PNKP, Marella
designed complimentary DNA substrates wherein one double-stranded DNA (dsDNA) has a 5’-
DNP tag, the other dsDNA has a 3’-biotin tag, and they each have a 3’-phosphate end (Table 2).
11
Marella’s early experiments indicated that the dephosphorylation of these DNA substrates could
be measured by HTRF with reactions that contained 44 nM Homo sapiens PNKP (hPNKP) and
74 nM T4 Ligase in HTRF buffer (Table 1) with 0.1 mg/ml BSA. Under these conditions, the
HTRFphos-1 and HTRFphos-2 substrates (Table 2) were incubated at 20 nM. SAXL and DNPK
were diluted 100-fold in PBS (Table 1) and added separately at the end of the
dephosphorylation-ligation reaction. Once hPNKP dephosphorylated the DNA substrates, T4
Ligase was then able to ligate them due to their complimentary 5’ overhangs. The ligation of the
two DNA strands would bring the donor and acceptor fluorophores sufficiently close together to
produce a HTRF signal. The “background” levels of HTRF were then measured by performing
the assay with samples that lack enzyme and should therefore not contain ligated product, which
was then used to normalize HTRF signal between experiments (i.e. “signal/background”).
Though Marella’s studies showed proof of concept, a number of parameters required testing and
optimization in order to adapt this assay for high-throughput compound screening. Because the
compounds to be tested are dissolved in DMSO, it was necessary to determine if the
concentration of DMSO had an effect on the assay. I accomplished this by carrying out the
aforementioned HTRF reactions with 1%, 2.5%, 5%, or no DMSO. The results of this
experiment indicated that the inclusion of up to 5% DMSO in the reaction did not significantly
affect HTRF signal (Figure 2.1.1.a).
Another parameter that required optimization was the concentration of potassium fluoride (KF)
in the DNPK solution: according to the manufacturer (Cisbio Bioassays, USA), reactions
containing Europium cryptate require 100 – 400 mM KF to protect the cryptate from quenchers
that may be present in the reaction at the time of HTRF measurement. Also, the manufacturer
states that KF is only required at the time of HTRF measurement; therefore, I introduced the KF
12
with the DNPK solution (Table 1) because that was the last component to be added to the
reaction before HTRF was read.
I carried out HTRF reactions in the presence of 100, 200, 300, or 400 mM KF. I observed that,
although variability remained the same across KF concentrations of 200 mM or greater, there
was an apparent decrease in signal with decreasing KF concentration (Figure 2.1.1.b). Notably,
HTRF signal was very poor at 100 mM KF. On this basis, I concluded that the final
concentration of KF in HTRF reactions should be 400 mM.
The next condition that I optimized was the duration of the dephosphorylation-ligation reaction.
To achieve this, I performed HTRF reactions as previously described and reactions were allowed
to proceed for 0.5, 1, 1.5, 2, 2.5, or 3 h at room temperature. To ensure that the incubation time
with fluorophores was consistent between time-points, I started the reactions every 30 min, and
then SAXL and DNPK solutions were added to all wells after 3 h. The results of the time course
with 44 nM hPNKP and 74 nM T4 Ligase showed high amounts of signal at each time point
(Figure 2.1.1.c) and I determined that a 1.5 h incubation time would provide the greatest signal
window within a reasonable timeframe.
Another important optimization step was that of hPNKP and DNA concentrations. Although the
aforementioned optimizations were performed under previously established conditions, it was
necessary to verify that the enzyme and substrate concentrations were optimal in my hands.
hPNKP was diluted in PNKP dilution buffer (Table 1) and reactions were carried out as
described with 88, 44, 22, 8.8 or 0.88 nM hPNKP (Figure 2.1.1.d). Because maximizing signal
was considered paramount, I concluded that 44 nM of hPNKP (50 ng) was still an appropriate
concentration to work with. I then performed HTRF reactions at 44 nM hPNKP with final
13
HTRFphos-1 and HTRFphos-2 (Table 2) DNA concentrations of 0, 5, 10, 20, 35, 50, 200, or 250
nM (Figure 2.1.1.e). From this experiment, I found that maximum signal was achieved at 35-50
nM DNA, so I selected 40 nM of DNA substrates as the optimal DNA concentration for this
assay.
Figure 2.1.1 Optimization of HTRF screen conditions. (a) HTRF assays were carried out with different
final concentrations of DMSO to verify that DMSO concentration would not affect signal. (b) HTRF
assays were carried out at 0 - 400 mM KF to determine which concentration produced maximum signal.
(c) A time course for the HTRF reaction was carried out to select an incubation time that maximized
signal. (d) hPNKP was titrated in HTRF reactions to determine which concentration produced maximum
signal. (e) DNA was titrated with 42 nM SAXL and 2 nM DNPK fluorophores in HTRF reactions to
determine the optimal concentration of DNA for use with the aforementioned concentration of
fluorophores. Error bars for (a) and (d) represent standard error of the mean (SEM) for at least two
independent experiments. Error bars for (b), (c), and (e) represent the SEM for two technical replicates.
Until this point, I had prepared SAXL and DNPK solutions by separately diluting each reagent
100-fold in PBS (Table 1), and these were added sequentially at the end of the reaction; however,
to optimize this assay for a high-throughput screen it was necessary to determine if the order of
fluorophore addition could be modified to minimize the number of addition steps. To this end, I
14
carried out HTRF reactions wherein SAXL and DNPK were diluted and added separately as
before, diluted and added together, or diluted and added together with the DNA substrates
(Figure 2.1.2.a). The results of this experiment suggested that SAXL and DNPK could be
diluted and added together at the end of the reaction, albeit with a small loss of signal. I also
subsequently carried out reactions in which SAXL was either added at the end of the reaction
with DNPK or included in the reaction with the DNA substrates, and this experiment
demonstrated that the latter order of addition led to a marked increase in signal (Figure 2.1.2.b),
so I adopted this order of addition for the screen.
Due to the nature of high-throughput screening, it was also important to verify that the HTRF
reagents were stable for long periods of time at room temperature as each screen run can take
several hours. To test this, I prepared dilutions of SAXL and DNPK in PBS (Table 1) every 2 h,
up to 6 h. HTRF reactions were initiated 4.5 h after the first SAXL and DNPK dilutions were
made and incubated for 1.5 h. I ended the reactions at the same time by adding the SAXL and
DNPK solutions that had been prepared at different times (i.e. 2 h, 4 h, 6 h ago or freshly made)
to the wells. The data suggested that the HTRF fluorophores were indeed stable at room
temperature for up to 6 h after being diluted to their working concentrations (Figure 2.1.2.c).
Lastly, I repeated the titration of hPNKP to ensure that the concentration of enzyme was
optimized for the screening conditions, and I determined that a concentration of 8.8 nM hPNKP
per reaction (10 ng) was the most suitable (Figure 2.1.2.d).
15
Figure 2.1.2 Optimization of the order of addition, stability of reagents at room temperature, and
hPNKP concentration for HTRF screen conditions. (a,b) Different methods of preparing HTRF reagents
were tested to determine which order of addition produced maximum signal. (c) HTRF reagents were
tested after being left at room temperature to verify that the reagents would be stable for the duration of a
screen run. (d) hPNKP was again titrated to check that the concentration was appropriate with the newly
optimized HTRF conditions. Error bars represent the SEM for two technical replicates.
To verify that this assay was amenable to high-throughput screening, I used the optimized assay
to screen the Prestwick Chemical Library (Prestwick Chemical, France), which consists of 1200
FDA-approved drugs (Figure 2.1.3.a). According to the manufacturer, this library contains
compounds with a high degree of chemical and pharmacological diversity, as well as
bioavailability. From this screen, I identified seven inhibitory compounds (Figure 2.1.3.b).
However, further testing under hPNKP-independent HTRF conditions showed that these
compounds were inhibiting both PNKP-dependent and –independent ligation reactions (Figure
2.1.3.c).
16
Importantly, the z’ factor was quantified for this screen as a measure of the its statistical validity,
because it represents a screen’s capacity for identifying hits by taking into account the assay’s
signal window as well as dynamic range (Zhang et al., 1999). Specifically, a ratio of 0.5 is the
minimum value considered acceptable for high-throughput screening because it signifies clear
separation of positive and negative controls plus or minus three standard deviations (Zhang et al.,
1999).
Based on this pilot screen’s z’ factor of 0.73 ± 0.02, this trial demonstrated that the assay was
compatible with high-throughput screening. Therefore, despite the fact that this screen did not
identify any putative hPNKP inhibitors, it did prove that the assay produces statistically valid
data and can reliably identify primary hit compounds.
17
Figure 2.1.3 Screening the Prestwick Chemical Library confirmed that the optimized conditions were
amenable to high-throughput screening. (a) Control-based analyses were performed to identify the
compounds that reduced HTRF signal by greater than three standard deviations from the mean positive
control (light blue line) for each plate. The mean negative controls for each plate are represented by the
dark blue line. FA = fluorescence emitted at 665 nm by SA-XL (acceptor); FD = fluorescence emitted at
620 nm by DNPK (donor); HTRF = (FA/FD) × 10,000 (b) Seven compounds were found to inhibit the
HTRF reactions, representing a 0.58% hit rate. (c) A secondary HTRF assay measuring hPNKP-
independent ligation was used to conclude whether or not the reduction in signal observed in the screen
was specific to hPNKP inhibition. Because all of the compounds caused inhibition in this secondary
screen, they were not considered to be inhibitors of hPNKP.
2.2 Development of screen conditions, Version 2
I developed the first version of the screen in such a way as to maximize the signal window with
little regard for the kinetics of the reaction. While these conditions led to a high degree of
statistical validity, they were not ideal for an inhibitor screen because assaying a reaction at
equilibrium can mask the effects of weak inhibitors, which have the potential to be developed
into more potent inhibitors. In the case of such inhibitors, measuring the assay in the linear
portion is key because a weakly inhibited reaction will exhibit a slower initial velocity, but it will
18
ultimately reach a level of signal similar to that of an uninhibited reaction at saturation
(Copeland, 2005). Consequently, I undertook further optimizations to find conditions that were
not saturating but still maintained a high degree of statistical validity.
First, I noted that, although a time course was performed early in the optimization process, the
chosen incubation time of 1.5 h resulted in a highly saturated reaction; therefore, the incubation
time required adjustment to alleviate the reaction’s saturation. It was also important to ensure
that the reactions were being efficiently stopped at the desired time-point.
I tested two approaches for stopping the reactions: addition of EDTA, which is a chelator of
magnesium – the divalent metal necessary for hPNKP and T4 Ligase activity - or addition of the
classic phosphatase inhibitor sodium orthovanadate (Na3VO4, Gordon, 1991). First, I determined
the concentrations of EDTA and Na3VO4 at which the reaction is inhibited by 50%, i.e. their IC50
values, by pre-incubating the reaction mix with EDTA at 0.1 – 50 mM or Na3VO4 at 100 nM –
20 mM (Figure 2.2.1.a-b). I found the IC50 of EDTA to be 7.6 mM (Figure 2.2.1.a), while that
of Na3VO4 was estimated to be about the same (Figure 2.2.1.b). I then carried out time courses
wherein reactions were stopped with either 50 mM EDTA, 10 mM Na3VO4, or 20 mM Na3VO4
(Figure 2.2.1.c). Notably, it appeared that, when reactions were stopped with Na3VO4, the HTRF
ratios (Fl645×10,000/Fl620) were within their typical ranges but the fluorescence intensity in each
channel was about 4-7 times lower than what was typically observed. This suggested that
Na3VO4 was quenching fluorescence, which was further supported by the observation that
fluorescence intensities were 2-fold lower for the wells containing 20 mM Na3VO4 than the wells
containing 10 mM Na3VO4 (data not shown). Furthermore, when reactions were stopped with
Na3VO4, the HTRF ratios at t0 were 2 – 3 times higher than what was typically observed; this
was not the case for reactions stopped with EDTA, so I concluded that addition of 50 mM EDTA
19
was better suited for stopping the dephosphorylation and ligation reactions. Furthermore, to
minimize the number of additions during the screen, EDTA was included in the DNPK solution
(Table 1) without negatively impacting signal.
The presence of BSA in the screen was another condition that I reconsidered. I had added BSA
to the reactions as a carrier for the enzymes of interest, but its high concentration in the screen
could make it capable of binding compounds and sequestering them from the enzymes of
interest, which could produce false negatives and reduce sensitivity. To test if this could be
occurring, I assessed the effects of three putative hPNKP inhibitors by HTRF in the presence and
absence of BSA at 0.1 mg/ml (Figure 2.2.1.d). I found that the inclusion of BSA reduced the
inhibitory effects by 20-25% for two of the three compounds that were tested, with little effect
on the uninhibited reaction (Figure 2.2.1.d). Consequently, I excluded BSA from future HTRF
reactions.
In attempting to find an incubation time that would fall in the linear portion of the reaction curve,
I observed that the reaction proceeded too quickly at 8.8 nM hPNKP, so the concentration of
hPNKP had to be reduced to generate a longer linear portion (Figure 2.2.1.e). However, the
reaction curve under the current conditions also appeared to be biphasic, which suggested that
the concentration of T4 Ligase may not be saturated, as a reporter enzyme should be (Figure
2.2.1.e). Consequently, I optimized both T4 Ligase and hPNKP concentrations once again.
20
Figure 2.2.1 Further optimization of the HTRF assay was undertaken to find non-saturating conditions
with high statistical validity. (a) HTRF reactions were pre-incubated with different concentrations of
EDTA to determine the IC50 value. (b) Using the same technique, the IC50 value of sodium orthovanadate
(Na3VO4) was estimated. (c) A time course was carried out, with reactions stopped with 10 mM or 20
mM Na3VO4, or 50 mM EDTA, to observe which method of halting the reaction would be most effective.
(d) Different putative hPNKP inhibitors identified from an earlier screen were tested in reactions with and
without BSA to evaluate whether or not BSA could mask the inhibitory effects of compounds. (e) New
time courses performed with more early time-points indicated that there was a lag occurring in the first 15
minutes of the reaction. Error bars for (a) and (b) represent the SEM for at least two independent
experiments. Error bars for (d) and (e) represent the SEM from two technical replicates.
I re-optimized the T4 Ligase concentration by performing HTRF reactions with 40, 75, 150, 300,
400, 600, 800, or 1000 nM T4 Ligase (Figure 2.2.2.a). The experiment indicated that the
previous T4 Ligase concentration of 74 nM was far from saturating, as hypothesized. To
21
guarantee that T4 Ligase was sufficiently saturating, I selected 800 nM T4 Ligase as the best
concentration for the remainder of the screen.
Next, I titrated hPNKP in reactions at 0.1, 0.3, 1, 3, and 10 nM (Figure 2.2.2.b). Although the
linearity of the curve within the first 15 minutes had improved, the excessive drop in activity
between 3 nM and 1 nM was curious and suggestive of the protein being unstable at
concentrations below 3 nM in PNKP dilution buffer (Table 2). This observation led me to repeat
the experiment with the addition of 0.02% Tween-20 to PNKP dilution buffer (Table 1), which
could increase the protein’s stability at lower concentrations and also reduce its potential to stick
to the polypropylene tube. Remarkably, the inclusion of Tween-20 in the dilution buffer led to an
increase in the initial velocities across all hPNKP concentrations and appreciable levels of
activity for reactions with less than 3 nM hPNKP. I subsequently repeated the experiment with
the liquid handling robotics platform to ensure that the best hPNKP concentration and incubation
time would be chosen for the screen (Figure 2.2.2.c). Based on these experiments, I determined
that 1 nM hPNKP and 20 minutes for the dephosphorylation-ligation reaction were suitable
parameters for the remainder of the screen.
22
Figure 2.2.2 The T4 Ligase and hPNKP concentrations were optimized to prevent the ligation reaction
from causing a kinetic lag. (a) The T4 Ligase concentration was titrated to find a concentration at which
the ligation reaction would no longer be limiting. (b) hPNKP was diluted in PNKP dilution buffer (Table
1) and titrated into HTRF reactions with 800 nM T4 Ligase. (c) hPNKP was diluted in PNKP dilution
buffer with 0.02% Tween-20 and titrated into HTRF reactions with 800 nM T4 Ligase to increase the
stability of hPNKP at low concentrations. Error bars in (a) represent the SEM from two independent
experiments.
The change of hPNKP and T4 Ligase concentrations meant that the concentration of HTRFphos-
1 and HTRFphos-2 substrates (Table 2) should once again be adjusted. I also tested different
concentrations of the SAXL acceptor fluorophore to determine if signal could be increased
(Figure 2.2.3.a-b). I carried out the experiment with 1 nM (Figure 2.2.3.a) or 3 nM hPNKP
(Figure 2.2.3.b) and a 20 minute reaction time. The results suggested that 120 nM DNA
substrates and 84 nM SAXL fluorophore produced the best signal without using an excessive
amount of the expensive fluorophore at the hPNKP concentrations tested.
Finally, another approach was taken to improve upon HTRF signal: shortening the DNA
substrates. According to the FRET equation - percent energy transfer = 1 / [1 + (r/R0)6, where r is
the distance between acceptor and donor fluorophores, and R0 is the Förster distance, at which
23
50% of the energy is transferred from donor to acceptor – with a Förster distance of 90 Å, only
28% of the energy is transferred from the Europium cryptate in DNPK to the XL-665
fluorophore in SAXL when the ligated DNA is 31 nucleotides (nt) – or about 105 Å - long.
However, by shortening the ligated DNA product to 25 nt (85 Å), the efficiency of energy
transfer more than doubles to 59%, which in theory would increase the signal over background. I
tested the newly optimized conditions side-by-side with the 31 nt and 25 nt substrates, and, as
hypothesized, the shorter substrates more than doubled the signal over background (Figure
2.2.3.c). I carried out a final time course at 1, 2, and 4 nM hPNKP using the newly optimized
conditions and shorter DNA substrates, and 2 nM hPNKP with a 20 minute incubation time was
found to provide sufficient signal without completely saturating the reaction (Figure 2.2.3.d).
Unfortunately, these conditions were not totally amenable to high-throughput screening because
once applied to a test run of ten plates, there was a substantial increase in variability, which
greatly reduced statistical validity such that the z’-factors were hovering around 0.5. In an effort
to reduce variability, 240 ml polypropylene reservoirs were utilized as the reagent source instead
of 2 ml 96-well plates. However, the use of reservoirs as source plates demanded a greater
consumption of reagents, which was excessive in the case of the SAXL fluorophore. To
minimize the costs associated with this increased consumption, I adjusted the concentration of
SAXL once more. I carried out reactions with 42, 60, or 84 nM SAXL, and 2 nM or no hPNKP
(Figure 2.2.3.e). I selected 60 nM SAXL as the best concentration going forward, not only
because it would consume less reagent, but also because it gave better signal-to-background
ratios (Figure 2.2.3.e). The combination of using reservoirs as source plates and reducing SAXL
from 84 nM to 60 nM greatly benefited the statistical validity of the screen, with z’ factors
increasing from ~ 0.5 to a more acceptable level of greater than 0.65.
24
Figure 2.2.3 Optimization of DNA concentration, ligation product length, and acceptor fluorophore
concentration to maximize signal-to-background. (a, b) The concentration of DNA (31 nt product) was
titrated in HTRF reactions at 1 nM hPNKP (a) or 3 nM hPNKP (b). (c) HTRF assays were conducted
with DNA substrates that would create 31 or 25 nt ligation products to verify that the shorter product
would produce greater HTRF signal. (d) A time course was carried out at different hPNKP concentrations
using the HTRFphos-3 and HTRFphos-4 substrates, which generate 25 nt ligation products; inset shows a
close-up on the first 15 minutes of the reaction. (e) Lower concentrations of SAXL were tested under the
optimized conditions. Error bars in (c) and (e) represent the SEM from two technical replicates.
2.3 Homogeneous time-resolved FRET-based screen
To find putative inhibitors of hPNKP, the HTRF assay was used to probe the small molecule
library assembled by the Ontario Institute for Cancer Research (OICR; Toronto, Canada) with
ten, twenty, or thirty plates screened per day. I included positive and negative controls in each
plate so that each plate could be analyzed independently. In order to maintain statistical validity,
I repeated plates with z’-factors below 0.50. Compounds that caused a reduction in signal by
more than three standard deviations were designated as “primary hits”, and these were selected
for secondary screening. At the end of the “first half” of the screen (plates 1-250, Version 1 of
25
the HTRF assay), 574 primary hits were uncovered from the 80,000 compounds that were
assayed, giving a hit rate of 0.72% (Figure 2.3.1.a). From the “second half” of the screen (plates
251-445, Version 2 of the HTRF assay), 357 hits were identified from the 62,400 compounds
tested, giving a hit rate of 0.57% (Figure 2.3.1.b).
Because the primary HTRF-based screen uses phosphatase-dependent ligation of DNA substrates
as the readout for hPNKP activity, the primary screen is inherently going to identify inhibitors of
both ligation and dephosphorylation. There is also the possibility that some apparent inhibitors
are results of non-specific effects, e.g. fluorescence quenching or DNA intercalation.
Consequently, I employed a secondary HTRF-based screen to eliminate the compounds that
reduced the HTRF signal for reasons other than hPNKP inhibition. To do so, the protocol for the
secondary screen was identical to that of the primary screen, except that hPNKP was eliminated
from the reactions and the DNA substrates contained ligatable 3’-hydroxyl groups (HTRFOH-1
and -2, Version 1; or HTRFOH-3 and -4, Version 2; Table 2). With this method, only
compounds that caused hPNKP-independent inhibition will inhibit reactions in both the primary
and the secondary screens; therefore putative hPNKP inhibitors were hits in the primary screen
but showed little to no inhibition in the secondary screen (Figure 2.3.1.c-d). Using this
approach, I found a total of nine putative inhibitors of hPNKP (Table 3), which equals an overall
hit rate of 0.0063%.
26
Figure 2.3.1 Screening of the OICR HTS chemical collection under primary and secondary screen
conditions to identify putative hPNKP inhibitors. (a) 80,000 compounds were tested under saturating
(Version 1) screen conditions. The mean fold-change in activity is represented by the red line, and the
blue lines represent plus or minus three standard deviations away from the mean. (b) More than 62,000
compounds were tested under non-saturating (Version 2) screen conditions. (c) After testing the primary
hits in the secondary screen (Version 1) for inhibition of hPNKP-independent ligation, four hits were
identified as putative hPNKP inhibitors, circled in red. (d) After testing the primary hits in the secondary
screen (Version 2) for inhibition of hPNKP-independent ligation, five hits were identified as putative
hPNKP inhibitors, circled in red.
2.4 Putative inhibitors of hPNKP
Interestingly, with input from Dr. David Uehling of the OICR, we noticed some common
structural motifs among the putative hPNKP inhibitors. One feature was the presence of pyrrole,
pyrazine and/or thiophene rings, which were found in all nine hits. Another feature, which was
found in the first four hits of the screen (OICR1-4, Table 3), was the presence of a carboxylic
acid group at the end of an aliphatic chain attached to a five- or six-membered ring. The final
apparent feature was the presence of an amide group within the structures of the five compounds
27
identified from the “second half” of the screen (OICR5-9, Table 3). Intriguingly, compounds
OICR7 and OICR9 have a common fragment, which contains two of the three aforementioned
features - a pyrrole ring, and a carboxylic acid or amide group attached to a benzene ring
(OICR79, Table 4). The importance of these structural motifs will be revisited later with
structure-activity relationship (SAR) analysis.
2.5 Validation and IC50 measurement of putative hPNKP inhibitors by HTRF
I acquired each putative inhibitor anew as a powder from commercial sources. I dissolved the
compounds in DMSO and assayed them at a range of concentrations in order to establish IC50
values, provided that the compounds still inhibited the reaction. Unfortunately, OICR4 was no
longer available, so it could not be subjected to any follow-up experiments. Furthermore,
compounds OICR1, OICR5, and OICR8 did not reproduce inhibition when tested from freshly
dissolved powders (Figure 2.5.1.a); we posit that the apparent inhibitory effects of these
compounds in the screen were due to breakdown products or impurities present in the library
source plates.
The remaining five compounds did exhibit inhibition of the HTRF reaction after being freshly
dissolved in DMSO, so I measured the IC50 values of these five compounds in order to determine
their relative potencies (Figure 2.5.1.b). I observed a wide range of potencies with these
compounds, with OICR2 having an IC50 value of ~ 0.5 µM, and OICR3 having an IC50 value of
~ 40 µM. Although OICR2 and OICR3 may appear superficially similar, their Tanimoto
coefficient – which describes the similarity of two molecules, and has a minimum of 0 and a
maximum of 1 (Tanimoto, 1957) – is 0.40, which indicates that they are chemically dissimilar,
and this explains the difference in their IC50 values.
28
Figure 2.5.1 Several putative inhibitors were validated as inhibitors of hPNKP with a range of potencies,
while others could not be reconfirmed. (a) Compounds OICR1, OICR5, and OICR8 did not reconfirm
when tested from freshly dissolved powders. (b) The remaining five hits – OICR2, OICR3, OICR6,
OICR7 and OICR9 – did reconfirm with IC50 values from ~ 0.5 µM (strongest inhibitor, OICR2) to ~ 40
µM (weakest inhibitor, OICR3). Error bars represent the SEM from at least two independent experiments.
29
2.6 Optimization of gel-based phosphatase assay
I developed a gel-based assay based on the methods of Freschauf et al. (2009) so that the
putative inhibitors could be validated by directly measuring phosphatase activity, thus abolishing
the need for T4 Ligase and HTRF fluorophores as reporters. Classically, the 3’-phosphorylation
status of DNA that has been 5’-labeled with 32
P would be followed by electrophoresis on a
polyacrylamide sequencing gel. The presence of the 3’-phosphate imparts a greater negative
charge to the phosphorylated DNA, which causes it to migrate faster on the gel (Figure 2.6.1.a).
However, thanks to the modern availability of fluorescent labels, these experiments were carried
out with Cy5-labeled DNA in lieu of radioactively-labeled DNA (GELphos, Table 2).
To maintain consistency and be able to directly compare the results from the gel-based
experiments with those from the HTRF-based experiments, I carried out all of the phosphatase
gel assays in HTRF buffer (Table 1) at 2 nM hPNKP and 120 nM GELphos DNA substrate
(Table 2). However, to ensure that the reactions were not saturated, I conducted a time course to
select an optimal reaction time. I quantified the percentage of de-phosphorylated DNA at each
time-point using ImageQuant 5.0 (Molecular Dynamics, USA) and plotted a reaction curve
(Figure 2.6.1.b). Based on this data, I selected 15 minutes as a suitable reaction time; although
this time point does not fall exactly in the linear portion of the reaction, it does appear that ~ 50%
of the DNA has been de-phosphorylated and the reaction is still progressing, albeit at a slower
rate.
30
Figure 2.6.1 A gel-based assay was developed to directly measure the dephosphorylation activity of
PNKP. (a) The activity of PNKP on the 3’-phosphorylated DNA substrate leads to a band shift because
the loss of the negative charge imparted by the 3’-phosphate causes the DNA to migrate more slowly on
the gel. (b) A time course was performed with 2 nM hPNKP and 120 nM DNA to select a reaction time
that would allow for the dephosphorylation of about 50% of the DNA.
2.7 Measurement of IC50 values by gel-based phosphatase assay
Using the above reaction conditions, each putative inhibitor was assayed at a range of
concentrations in order to establish IC50 values from this direct assay. Remarkably, the IC50
values generated by the gel assay were significantly lower than those from HTRF experiments,
by a factor of about 10-20 (Figure 2.7.1). Since the hPNKP and DNA substrate concentrations
were maintained between the HTRF and gel experiments, I hypothesized that this disparity may
be due to nonspecific - albeit benign - binding of the compounds to the T4 Ligase. Such
nonspecific binding would titrate the compound away from hPNKP and raise the apparent IC50
value. Additionally, the DNA sequences of the HTRFphos and GELphos substrates (Table 2) are
not identical, so this may contribute to the differences in IC50 values. Alternatively, it is possible
that the gel assay is simply more sensitive than the HTRF assay.
31
Figure 2.7.1 Dose response experiments were performed for each of the five original hits using the gel
assay, and IC50 values were measured. (a-e) The phosphatase activity of hPNKP was measured directly by
gel assay, which eliminated the use of a reporter enzyme and thus confirmed that the compounds are
impeding hPNKP from processing 3' phosphate DNA termini. Error bars represent the SEM from at least
two independent experiments.
2.8 Structure activity relationship analysis
The three structural motifs common to the putative hPNKP inhibitors identified by the screen –
amide groups, carboxylic acid groups, and five-membered heterocyclic rings - were of great
interest to us, so I utilized structure-activity relationship (SAR) analysis to assess their roles in
inhibiting hPNKP. Toward this end, I obtained a number of analog compounds in which these
features were individually modified or moved relative to the original structure (Table 4).
Furthermore, I acquired a number of analogs of OICR79 in order to investigate whether this
minimal fragment could be made more potent (Table 4).
32
I employed HTRF experiments as well as subsequent gel-based experiments to assess how the
modifications affected the compounds’ potency. Based on analogs whose pyrrole rings were
removed (OICR9A) or converted to –N(CH3)2 (OICR9B), I determined that the pyrrole ring was
necessary for PNKP inhibition by OICR9 derivatives (Figure 2.8.1.a). The lack of inhibition by
analog OICR7A further reinforced the importance of the pyrrole ring, since changing its position
relative to the rest of the structure also abolished inhibitory activity (Figure 2.8.1.b).
OICR79 - the common fragment from OICR7 and OICR9 - was also tested by HTRF (Figure
2.8.1.c). We surmised that this structure could be responsible for the inhibition of hPNKP by
OICR7 and OICR9, and if this was the case, OICR79 should prove to be at least as inhibitory as
the original hits. The IC50 value of OICR79 was found to be on the same scale as that of OICR7
(compare Figure 2.8.1.c and Figure 2.5.1.b), which suggests that this portion of OICR7 may be
entirely responsible for hPNKP inhibition. However, it should be noted that the IC50 values from
the gel assay are not as consistent, with OICR79 appearing to be ~ 7 times less potent than the
parent compounds. Owing to this discrepancy, the importance of the rest of the OICR7 and
OICR9 structures should not be discounted entirely and their roles should be further studied by
testing new analogs in which they are modified.
Since the common fragment portion of OICR9 contains an amide group on the benzene ring
rather than a carboxylic acid group, we would have liked to test an amide version of this
fragment as well, but such a compound is not currently available.
33
Figure 2.8.1 Structure activity relationship analyses of the heterocyclic ring and the fragment common to
OICR7 and OICR9. (a) Analogs of OICR9 may appear to inhibit hPNKP at concentrations > 100 µM, but
closer inspection of the fluorescence emitted in the acceptor and donor emission channels (below; “SAXL
em.” and “DNPK em.” respectively) suggest that this is due to fluorescence quenching by high
concentrations of compound. (b) HTRF assays demonstrated that shifting the position of the heterocyclic
ring in OICR7 also abolished activity. (c) OICR79, the fragment common to OICR7 and OICR9,
displayed inhibition of hPNKP of a magnitude similar to the parent compounds (left). This inhibition was
confirmed by the phosphatase gel assay (right), although the IC50 value is about seven times greater than
those of the parent molecules. Error bars represent the SEM from at least two independent experiments.
Functional groups in red represent groups that differ between the parent and analog compound.
34
To learn more about the OICR79 fragment, I examined several analogs (Table 4) for their ability
to inhibit hPNKP. The addition of a methyl group at C6 of the benzene ring in OICR791 and
OICR793 (Table 4) caused a reduction in potency to the point that they only produced
observable inhibition at 500 µM in HTRF reactions (Figure 2.8.2.a-b); perhaps the presence of a
methyl group near the pyrrole ring imposes steric hindrance, which impedes the pyrrole ring
from binding effectively to hPNKP.
Another analog that virtually lost its activity was OICR792, which has a C=C(NH2)2 group
attached to C2 of the pyrrole ring (Figure 2.8.2.c; Table 4). The consistent loss of potency
brought on by the addition of functional groups on or near the pyrrole ring of the parent
compound further strengthens the argument for its importance. Interestingly, analog OICR795,
which had the pyrrole and carboxylic acid groups in ortho rather than meta positions on the
benzene ring (Table 4), was less potent but not completely inactive with an IC50 of ~ 200 µM
(Figure 2.8.2.d). This inhibition, although considerably weaker than that of the parent
compound, was curious because it appeared to be greater than that of OICR791 and OICR793,
which have methyl groups added to the same position. I hypothesized that this discrepancy arose
because the carboxyl group bound better to hPNKP in an ortho rather than meta position, and the
inclusion of an ortho methyl group with the meta carboxyl group threw a proverbial wrench in
the works, making binding very difficult for the carboxyl group. The validation of this
hypothesis would be aided by a structure of hPNKP complexed with inhibitor.
35
Figure 2.8.2 Structure activity relationship analyses using analogs of OICR79. (a, b) HTRF dose
response assays demonstrated that the addition of a methyl group ortho to the pyrrole ring was
detrimental to inhibition, with the first ortho position (a, OICR791) impeding inhibition the most. (c)
HTRF dose response assays revealed that adding a –C=C(NH2)2 group to C2 of the pyrrole ring hindered
inhibition, too. (d) Shifting the two functional groups to ortho instead of meta positions also reduced
inhibition almost 100-fold by HTRF. Error bars represent the SEM from at least two independent
experiments.
One analog that displayed a reduced but still appreciable level of inhibition with an IC50 value of
~ 30 µM was OICR796 (Figure 2.8.3.a; Table 4); not only did this compound have a –CHOCH3
group rather than a -COOH group on C3 of the benzene ring, but it also had a –C=NCH3 group
attached to C2 of the pyrrole ring. We chose to test this analog because it had been identified as a
hit in a primary screen for inhibitors of the protein phosphatase Scp1, which is a member of the
36
HAD superfamily of phosphatases like hPNKP (Kamenski et al., 2004). However, because there
are two modifications in this compound relative to OICR79, it cannot be determined what role
each of them plays in the reduced inhibition.
Finally, in contrast to the aforementioned analogs that eliminated or reduced potency, OICR794
led to a significant increase in activity via the addition of another carboxyl group to the second
meta position (Figure 2.8.3.b). I posit that the symmetrical carboxyl group enhances the binding
of the compound to hPNKP, perhaps making additional contacts with the protein, and this
reinforces the importance of the carboxyl group, which is common among the primary hits.
37
Figure 2.8.3 Structure activity relationship analysis of OICR79 uncovered two modifications that did not
abolish inhibition. (a) With a C=NCH3 group added to C2 of the pyrrole ring and CHOCH3 replacing the
COOH group, OICR796 exhibited relatively weak activity by HTRF (left) but appreciable activity
according to the gel assay (right). (b) With an additional COOH group on the second meta position of the
benzene ring, OICR794 demonstrated about two times stronger inhibition than OICR79 by both HTRF
and gel assays. Error bars represent the SEM from at least two independent experiments.
2.9 Optimization of fluorescence polarization-based substrate-binding assay
Fluorescence polarization (FP) is a widely used technique for measuring the binding of a
fluorescently-labeled ligand to a protein because FP is proportional to the tumbling properties of
the fluorophore (Pagano et al., 2011). When a small fluorescently labeled ligand is free in
38
solution, its fluorophore tumbles quickly and therefore does not have a stable orientation; a
consequence of this rapid movement is that the fluorophore has poor capability for polarizing
fluorescent light upon excitation (Pagano et al., 2011). Conversely, when the ligand is bound to a
protein, it becomes stabilized in space because the much larger protein tumbles very slowly, and
this causes a significant increase in FP upon excitation of the fluorophore (Pagano et al., 2011).
Based on the methods of Coquelle et al. (2011), I utilized this technique to measure the binding
of a FAM-labeled 3’-phosphorylated penta-nucleotide substrate to hPNKP. For this assay, I
employed the phosphatase-dead D171A mutant of hPNKP so that it would bind the substrate but
not act on it.
To determine the optimal concentration of the FPphos substrate (Table 2) for this assay, I
measured the fluorescence intensity of substrate in FP buffer (Table 1) with a Beacon 2000
variable temperature fluorescence polarization system (Invitrogen, USA) at 510 nm. I selected a
concentration of 12.5 nM as a suitable working concentration for the FP assay as it gave a
fluorescence intensity of ~1000, which was low enough to prevent saturation of the detectors
(data not shown). I then titrated hPNKP-D171A in FP reactions, and analysis of this data
determined that the Kd of hPNKP under these conditions was 41.6 nM (see Methods, Figure
4.15.1). Based on these results, I chose 50 nM hPNKP-D171A as the concentration of protein to
use in the assay, since it bound ~50% of the substrate at equilibrium.
2.10 Substrate-binding assay confirms that compounds prevent a 3'-phosphorylated oligonucleotide from binding to hPNKP
Using the conditions outlined above, I measured an IC50 value for each of the original hits as
well as OICR79, OICR794 and OICR796 - the three analogs that exhibited measurable levels of
inhibition by HTRF. Remarkably, all of the compounds tested were capable of preventing the
39
substrate from binding to hPNKP-D171A, with IC50 values either similar to or stronger than
those measured by HTRF (Figure 2.10.1a-h).
Figure 2.10.1 Fluorescence polarization-based direct binding assays were carried out to determine if the
compounds could compete with a 3’-phosphorylated oligonucleotide substrate for binding to hPNKP-
D171A. (a-h) A range of compound concentrations were incubated with hPNKP-D171A and substrate
was added to the reaction and allowed to come to equilibrium prior to measurement of FP. The typical
sigmoidal dose-response curves for each compound indicated that these compounds were displacing DNA
from the phosphatase active site, although the mechanism by which this is achieved cannot be concluded.
Error bars represent the SEM from at least two independent experiments.
2.11 Inhibitors of hPNKP display different degrees of reversibility
The extent of reversibility of an inhibitor can provide insight into the mechanism by which the
inhibitor works; for example, irreversible inhibitors generally act by covalently modifying the
target enzyme so that it is permanently incapable of acting on its substrate (Copeland, 2005).
40
Conversely, inhibitors can be slowly or rapidly reversible; in such cases, the compound interacts
with the enzyme through non-covalent bonds such as hydrophobic interactions and ionic bonds
(Copeland, 2005).
To assess the reversibility of the hPNKP inhibitors, I performed an assay in which hPNKP was
incubated at 100-fold over the reaction concentration (i.e. at 200 nM) with ten times the IC50 of a
given compound (e.g. 76 µM OICR7) – which is enough to inhibit the enzyme by 90% - and
then diluted in reaction buffer 100-fold – leaving a concentration of compound at which hPNKP
would be inhibited by 9% (Copeland, 2005). I followed the reactions with a time course and
established the compound’s reversibility based on how quickly the enzyme recovered its activity
upon dilution (Figure 2.11.1). From the reaction curves, I determined that compounds OICR2
and OICR3 are slowly reversible, while the other compounds are very slowly reversible. In fact,
OICR6, OICR7 and OICR79 appear to hinder the recovery of hPNKP activity so much so that
they could be considered virtually irreversible.
41
Figure 2.11.1 Reversibility assays demonstrated that most of the compounds were at least slightly
reversible. (a) OICR2 and OICR3 were slowly reversible because the recovery of hPNKP activity was
slow and gradual over the time course. (b) OICR9 was somewhat reversible because a small amount of
hPNKP activity was recovered after 22 min. OICR6, OICR7 were virtually irreversible. (c) Analogs
OICR794 and OICR796 were also very slowly reversible, with hPNKP recovering to similar levels as
with OICR9. Analog OICR79 was virtually irreversible, with an effect similar to that of OICR7.
2.12 Inhibitors of hPNKP can act on T4 PNK with reduced potency
Although mammalian and T4 PNK(P) are each capable of performing 3’-phosphatase and 5’-
kinase activities, they exhibit a number of key differences; most notably, the presence of an
FHA domain in mammalian PNKP but not T4 PNK (Galburt et al., 2002). It is this FHA domain
that allows mammalian PNKP to be intimately involved in various forms of DNA repair because
the domain binds to phosphorylated XRCC1 and XRCC4, two integral components of the BER
and NHEJ pathways, respectively (Koch et al., 2004; Whitehouse et al., 2001). Additionally, T4
PNK behaves as a tetramer in vivo, whereas mammalian PNKP acts as a monomer (Galburt et
al., 2002). Furthermore, T4 PNK accepts RNA, DNA, and oligonucleotides as substrates, with a
preference for extended 5’ ends (Galburt et al., 2002); mammalian PNK, on the other hand,
42
accepts only DNA that is at least eight nucleotides long, with a preference for recessed DNA
ends (Bernstein et al., 2005).
In the interest of determining how specific the inhibitors were towards hPNKP, if at all, IC50
measurements were performed using T4 PNK in the gel-based assay (Figure 2.12.1).
Remarkably, after factoring in the two-fold increase in T4 PNK concentration over that of
hPNKP in the same assay, the IC50 values for T4 PNK are up to 50-fold higher than those for
hPNKP, thus revealing a substantial degree of specificity for the human protein over its T4
ortholog. It should be noted that the comparisons had to be made between gel-based rather than
HTRF-based experiments because the IC50 values were quite high, and the high concentrations of
compound had a tendency to interfere with the HTRF measurements.
Overall, despite the considerable reduction in potency relative to hPNKP, most of the
compounds were capable of effectively inhibiting T4 PNK, with only three compounds
displaying poor inhibition.
43
Figure 2.12.1 Gel-based assays conducted with T4 PNK demonstrate that some, but not all, of the
hPNKP inhibitors are capable of inhibiting this distant ortholog. (a-h) Gel-based assays were carried out
with 5 nM T4 PNK and 120 nM 3’-phosphorylated GELphos DNA (Table 2); most of the compounds -
OICR2 (a), OICR9 (e), OICR79 (f), OICR794 (g) and OICR796 (h) - exhibit sigmoidal dose-response
curves, indicating that they are functional inhibitors of T4 PNK. Compounds OICR3 (b), OICR6 (c), and
OICR7 (d) appear to inhibit T4 PNK at very high concentrations but their IC50values could only be
approximated because they inhibit so weakly. Error bars represent the SEM from at least two independent
experiments.
44
2.13 Inhibitors of hPNKP do not show a clear effect in U2OS cells co-treated with camptothecin
It has been demonstrated that knockdown or inhibition of hPNKP can potentiate the effects of
certain DNA damaging agents on cells; in particular, the requirement of PNKP for the repair of
damage induced by Topoisomerase I stalling causes hypersensitivity to CPT in hPNKP-impaired
cells (Freschauf et al., 2010; Rasouli-Nia et al., 2004). In order to explore whether or not the
compounds under investigation could induce the same sensitivity, I assessed cell viability in
U2OS cells treated with DMSO or 0.1 – 100 µM compound of interest, and co-treated with
DMSO or CPT (Figure 2.13.1.a-g). I used 5 nM CPT to sensitize the cells because it was found
to kill about 50% of the cells after a 96 h treatment at the seeding density used.
45
Figure 2.13.1 The viability of U2OS cells co-treated with 5 nM camptothecin (CPT) was not significantly
impaired relative to their DMSO-treated counterparts. (a) The indicated number of cells were treated with
a series of CPT concentrations for 96 h prior to the quantification of cell viability with the ATP Lite assay
(Perkin Elmer, USA) (b-i) Cells were treated for 96 h with a range of concentrations of each hPNKP
inhibitor plus DMSO or 5 nM CPT. Error bars represent the SEM from at least two independent
experiments.
Unfortunately, it does not appear that any of the compounds were capable of significantly
impairing the growth of CPT-treated U2OS cells relative to the control DMSO-treated cells
under these conditions. Some compounds did slightly impair cell survival at high doses, such as
OICR3 and OICR794 (Figure 2.13.1.b, f), but these effects were not vastly different between
DMSO and CPT co-treated cells.
There are a number of potential explanations for why the compounds did not sensitize the cells to
CPT as desired – for example, they cannot cross the cell membrane, they break down or become
46
modified in the cell, they preferentially target other enzymes in the cell – but to elucidate why
the compounds did not act as potentiators of camptothecin will require additional experiments
examining each hypothesis. Furthermore, and perhaps most importantly, the assay may not have
been designed optimally for the detection of PNKP inhibition-dependent CPT sensitization, as it
has been shown that cells in S-phase may not require the activity of PNKP to recover from CPT-
mediated damage (Rasouli-Nia et al., 2004). realize
Another limitation of the experimental set up that we later recognized was that the ATP Lite
assay I used to measure cell viability cannot differentiate between live cells and senescent cells.
By overlooking DNA damage-induced senescence (Di Leonardo et al., 1994), it is possible that
an impact of the inhibitors on viability was missed.
51
3 Discussion and future directions
The search for inhibitors of phosphatases has been challenging due to the nature of the enzymes’
structure. A common issue with inhibitors is a lack of selectivity, which is a consequence of
phosphatases’ highly conserved active sites (Barr, 2010). The second major concern regarding
phosphatase inhibitors is their low level of cell permeability, which is also a consequence of the
enzymes’ active sites: the highly polar active sites favor inhibitors with charged anionic
phosphate mimics that do not readily cross the cell membrane (Barr, 2010). However, in spite of
these common difficulties, the frequent finding that one or more phosphatases are involved in a
given disease has fueled countless searches for inhibitors of this class of enzymes. In the case of
PNKP, our search for an inhibitory compound was driven by the discovery that knockdown of
PNKP sensitizes cells to PARP inhibitors (Turner et al., 2008), Topoisomerase I inhibitors
(Rasouli-Nia et al., 2004), and ionizing radiation (Rasouli-Nia et al., 2004), which has
implications for the treatment of many types of cancer. This work was also encouraged by the
identification of selective inhibitors towards proteins in the same haloacid dehalogenase (HAD)
family of phosphatases as PNKP, such as Scp1 (Zhang et al., 2011).
The work described here has identified five unique compounds and several analog compounds
that based on the direct gel-based phosphatase assay (Figure 1.2.7.1), exhibit a range of
potencies, some of which are in the low nanomolar range. It is worth noting that the discovery of
only five inhibitors from a screen of more than 140,000 compounds amounts to a hit rate of
0.0035%, which is considered relatively low, as other high-throughput screens typically produce
hit rates that are greater by one to two orders of magnitude (Copeland, 2005). This may be due to
an unknown structural bias in the library, or it is possible that the structure of hPNKP supports
52
only a limited range of moieties. The likelihood of the former could be assessed by screening
additional chemical libraries to see what sorts of inhibitors they contain, if any.
Despite the low hit rate, the inhibitors identified by this large screen are very informative as they
belie a penchant for hPNKP to bind to heterocyclic rings, amides and carboxylic acid (COOH)
groups – at least two of which were present in each of the compounds and analogs tested (Tables
3, 4). Furthermore, the examination of analogs of OICR9 that lacked the heterocyclic ring or
retained the nitrogen heteroatom but not the full ring structure - OICR9A and OICR9B,
respectively - demonstrated the importance of this motif in effecting inhibition. Conversely, the
importance of the carboxylic acid moiety could be contested, as OICR796 contains a CHOCH3
group in its stead and still displays a moderate level of potency. However, because there are also
modifications made to the heterocyclic ring in OICR796, it is not possible to say for certain what
effect the change from –COOH to –CHOCH3 had on the potency of the compound; to do so, it
would be necessary to test an analog where only a single change was made. Bearing that in mind
though, it raises the possibility that the presence of a moiety with trigonal planar geometry is
more important than a COOH group specifically; to determine if this is the case, there are many
more motifs that could be explored, with the potential for some to increase potency and/or
selectivity. Moreover, the presence of an uncharged group in lieu of the COOH group would be
beneficial because it could lead to an increase in the compound’s membrane permeability.
With respect to the compounds' mechanisms of action, the FP assay confirmed that they do
prevent the DNA from binding to hPNKP, but whether that occurs via competition for the active
site, binding to an allosteric site, or simply denaturing the protein could not be ascertained from
this approach. However, the latter possibility was largely eliminated as a consequence of the
reversibility assays, which demonstrated that the compounds are generally reversible, with some
53
dissociating and relieving hPNKP inhibition more quickly than others. Other biochemical
experiments – for example, using circular dichroism (CD) spectroscopy to monitor changes in
PNKP secondary structure in the presence of substrate and/or inhibitor (Chen et al., 1974) could
provide additional evidence towards a mechanism of action. For example, if the addition of
inhibitor led to an increase in random structure, it would be suggestive of a noncompetitive
mechanism, whereby the binding of compound outside the active site causes a change in enzyme
structure that allosterically inhibits the enzyme.
Further insight into how the compounds work can be gleaned from the dose response
experiments with T4 PNK (Figure 2.12.1). Interestingly, across all dose response experiments
with hPNKP, regardless of the value of the IC50, the trend in relative potencies was maintained;
for example, OICR2 was always the most potent and OICR3, the least. However, this trend was
not completely upheld with T4 PNK; the most apparent difference being that, while OICR9 is
still relatively potent with an IC50 value of 5.74 µM, OICR7 with the common fragment does not
appear to inhibit T4 PNK. This contrast is curious since the compounds each contain nearly
identical active portions (based on the activity seen with OICR79), but this discrepancy is likely
due to structural differences in the vicinity of the common fragment's binding site on each
protein - that is, the binding site on T4 PNK may accommodate OICR9 but not OICR7, thus
preventing the latter from effecting inhibition of this ortholog. In fact, the finding that OICR79
can inhibit T4 PNK with an approximate ten-fold reduction in potency relative to OICR9
strongly implies that the dissimilar portions of OICR7 and OICR9 play an important role in
dictating the compounds' interactions with T4 PNK. These differences in activity towards
hPNKP and T4 PNK have implications for the inhibitors’ mechanisms of action because they
point to a structural difference between the two enzymes as being a component of the compound
54
binding site. This hypothesis is bolstered by preliminary data in which the compounds exhibit a
similar, albeit slightly reduced degree of potency (~ 1.5-2 fold) towards murine PNKP, which is
considerably more similar in sequence and structure to hPNKP than is T4 PNK. A
straightforward method for deciphering the compounds’ mode of action is to crystallize them
with PNKP, preferably with the human homolog for which they are most effectual.
The work presented here collectively demonstrates the discovery of eight compounds capable of
inhibiting hPNKP in vitro with at least moderate potency (i.e. micromolar IC50 values);
unfortunately however, none of the compounds showed an ability to sensitize U2OS cells to
camptothecin in tissue culture. Generally, uncharged polar compounds have the highest degree of
cell permeability since they are best equipped to cross the amphipathic lipid bilayer that
constitutes a cell’s membrane. Because carboxylic acid groups have a net negative charge, their
presence can make compounds less permeable; therefore, it is possible that the hPNKP inhibitors
are not entering the cell efficiently and this would prevent them from being present at high
enough concentrations to have a significant effect on hPNKP. This hypothesis could be tested by
assaying analogs that contain a methoxy group in place of the carboxyl group, provided that such
modifications do not eliminate the compounds’ activity in vitro. There are also well-established
cell-based assays that could be used to investigate the compounds’ membrane permeability, such
as the Caco-2 cell permeability assay, which could also be used to assess the metabolism of the
compounds in these cells (van Breemen and Li, 2005).
Beyond the question of the compounds' membrane permeability, it is not certain that the viability
experiments were designed appropriately to assess the role of PNKP inhibition in surviving CPT-
induced DNA damage: using clonogenic assays to measure the viability of A549 lung cancer
cells, Rasouli-Nia et al. (2004) have demonstrated that the cells exhibit a biphasic response to
55
CPT, with PNKP knockdown only sensitizing cells at doses greater than 1 µM. They postulated
that the drop in viability at lower doses was a consequence of the increased sensitivity of S phase
cells to CPT and that the inability of PNKP knockdown to sensitize these cells indicated that
PNKP was not critical for recovery from CPT during S phase. These findings have important
implications for my assessment of PNKP inhibitors in cells because they suggest that the choice
of CPT treatment conditions determines whether or not PNKP is needed for recovery and thus if
sensitization can take place. The conditions I employed for the cell viability assays - 5 nM CPT
for 72 h with no drug-free recovery time - are very different from those at which Rasouli-Nia et
al. (2004) demonstrated sensitization by PNKP knockdown - ≥ 1 μM CPT for 2 h and 2-3 weeks
recovery; therefore, I hypothesize that treating the cells for several days with a low dose of CPT
likely facilitated cell death in S-phase and caused PNKP activity to be inconsequential. This
hypothesis is bolstered by recently published work from Ray Chaudhuri et al. (2012), which
revealed that low doses of CPT are not associated with DSB formation, but rather with PARP-
dependent replication fork stalling and reversal. Overall, this suggests that the inability of the
PNKP inhibitors to sensitize cells to CPT may simply be a consequence of experimental design.
In order to accurately determine whether or not the compounds are effective in cells, it will be
key to repeat these experiments under conditions that trigger CPT-induced death in non-S phase
cells – that is, at higher doses of CPT with a shorter treatment time and increased recovery time.
Furthermore, the effects of PNKP knockdown on CPT sensitization published by Rasouli-Nia et
al. were shown by clonogenic assays, which quantify viability by measuring colony formation
following drug treatments. As a consequence, clonogenic assays account for the effects of both
apoptosis and senescence on viability, unlike the ATP Lite assay, which measures only
apoptosis. Because it is quite possible that PNKP inhibition decreases viability by promoting
senescence, it would be preferable to re-assess CPT sensitization by clonogenic assay.
56
Additionally, it would be worthwhile to repeat the experiments using the A549 lung carcinoma
cell line, in which Rasouli-Nia et al. demonstrated the CPT-sensitizing effects of hPNKP
knockdown, as well as using ionizing radiation as the source of DNA damage.
In the event that the compounds continue to fail to produce CPT sensitization, it will be
necessary to further investigate their membrane permeability as well as whether or not they are
affecting their intended target, using the alkaline comet assay, for example, which measures the
accumulation of SSBs in cells (Pfuhler and Wolf, 1996). If the compounds are in fact inhibiting
hPNKP, and thus hindering the repair of SSBs, DNA breaks might accumulate in mono-treated
cells and should persist even longer in CPT-sensitized cells.
Furthermore, the selectivity of the inhibitor-hPNKP interaction, and therefore the likelihood of
off-target effects occurring in cells, could be assessed by testing the ability of the compounds to
inhibit other phosphatases within the aspartate-based HAD family of protein tyrosine
phosphatases (PTPs), as well as phosphatases that operate via other mechanisms; for example, a
cysteine-based PTP or a protein serine/threonine phosphatase (PSP). It would also be prudent to
examine whether or not the compounds have an effect on other enzymes that perform similar
processing functions or have similar domains, such as Aprataxin (APTX) and APTX- and
PNKP-like factor (APLF).
Overall, the work that I have outlined herein has identified several compounds that inhibit
hPNKP, and these effects appear to be due to more than simple denaturation based on dose-
response, reversibility, and species-specificity experiments. Although further studies would be
required to determine the actual mechanism by which inhibition is achieved and the degree of
selectivity these compounds show for PNKP, the search for effective hPNKP inhibitors has
certainly moved forward. Alas, these inhibitors failed to demonstrate an ability to kill CPT-
57
sensitized cells, which is not particularly useful for the development of potential cancer
therapies, but there are many tools in medicinal chemistry that can be utilized to improve cell
permeability or reduce the potential for off-target effects. A collaboration that we have
undertaken with Dr. Mark Glover’s group at the University of Alberta may help answer many of
these questions, as they attempt to obtain and solve the structure of crystals of our compounds
bound to murine, and hopefully human, PNKP.
57
4 Materials and Methods
4.1 Transformation of E. coli.
XL1-Blue E. coli cells (Stratagene, USA) were made ultra-competent according to the Inoue
method (Inoue et al., 1990). BL21 (DE3) CodonPlus E. coli cells were made chemically
competent as described by Sambrook and Russell (2001). Cells were incubated in a microfuge
tube with plasmid DNA for 15 min on ice, heat-shocked for 30 sec at 42 °C, then moved
immediately back to ice where they were incubated for 10 min. One ml of Luria Broth (LB)
media (Table 1) was added to each tube and cells were grown for 1 h at 37 °C with shaking.
Cells were pelleted by spinning for 20 sec at maximum speed in a 5415 D table-top centrifuge
(Eppendorf, Germany). The supernatant was discarded, and the cells were resuspended in 100 µl
of LB media (Table 1). For XL1-Blue cells, 10% of the transformation mix was plated on one
LB plate containing 0.1 mg/ml ampicillin (LB + AMP; Table 1), and the remaining 90% was
plated on a second LB + AMP plate. For BL21 cells, 100% of the transformation mix was plated
on a single LB + AMP plate. Plates were incubated upside down in a 37 °C incubator overnight.
4.2 SDS-PAGE.
6X SDS-PAGE sample buffer (Table 1) was added to samples to a final concentration of 1X.
Samples were boiled for 5 min and then run on 8% or 10% SDS-PAGE gels (Table 1) for 45 min
at 200 V. Proteins in the gel were visualized by staining with Coommassie Brilliant Blue solution
(Table 1) for 5 min and then de-stained overnight in de-stain solution (Table 1).
4.3 Quantitation of protein
Protein samples were diluted 10-fold in 6 M guanidinium-HCl. A Nanodrop 8000
spectrophotometer was blanked with 6 M guanidinium-HCl and the concentration of protein was
determined by measuring absorbance at 280 nm.
58
4.4 Cloning of human PNKP-D171A
The Quickchange PCR method was used to generate the phosphatase-dead hPNKP D171A
construct from the pProExHTA-hPNKP plasmid (M. Canny, A. Koch). Seventy-five ng of
plasmid DNA was amplified with 250 ng each of forward (5’-
GGTGAAACCCCAGGGCAAGGTGGCTGGCTTTGCGCTGGACGGGACGCTCATCACCA
CACG-3’) and reverse (5’-
CGTGTGGTGATGAGCGTCCCGTCCAGCGCAAAGCCAGCCACCTTGCCCTGGGGTTTC
ACC-3’) primers. The PCR reaction also contained 100 μM of each dNTP, 2.5 units of Pfu DNA
polymerase (Agilent Technologies, USA), and 1X Pfu buffer [20 mM Tris-HCl, pH 8.8, 2 mM
MgSO4, 10 mM (NH4)2SO4, 10 mM KCl, 0.1% (v/v) Triton X-100, 0.1 mg/ml BSA] for 19
cycles [95 °C for 30 sec, 55 °C for 1 min, 68 °C for 22 min] and a final extension step of 68 °C
for 30 min. The PCR product was incubated with 20 units of DpnI restriction endonuclease (New
England BioLabs, USA) at 37 °C for 3 h. Six individual colonies from freshly transformed XL1-
blue E. coli were picked and grown overnight at 37 °C in 2-ml LB + AMP cultures (Table 1).
The DNA was purified from these cultures using Miniprep Spin Columns (Qiagen, Inc.)
according to the manufacturer’s instructions. The Miniprep columns gave yields of 115-200 ng
plasmid DNA/µl, and plasmids were sequenced by ACGT DNA Technologies Corporation
(Toronto, Ontario) to verify that the mutation was in place and the reading frame was intact.
4.5 Purification of human PNKP and PNKP-D171A
Competent BL21 (DE3) CodonPlus E. coli were transformed with 200 ng pProExHTA-hPNKP
plasmid (M. Canny, A. Koch) or pProExHTA-hPNKP-D171A, plated on LB + AMP (Table 1),
and grown overnight at 37 °C. Freshly transformed E.coli BL21 Codon Plus cells were scraped
into 10 ml of LB media (Table 1) and used to inoculate 2 × 1L of LB + AMP media (Table 1).
When the cultures reached an OD600 of 0.8, hPNKP expression was induced at 16 °C by adding
59
isopropyl β-D-1-thiogalactopyranoside (IPTG; Sigma, USA) to a final concentration of 100 μM.
Cultures were grown overnight at 16 °C and cells were harvested by spinning in a cold JLA8.1
rotor (Beckman Coulter, USA) for 10 min at 6000 × g. Cells were resuspended in 5 ml of cold
PNKP lysis buffer (Table 1) per gram of pellet. The resuspended cells were frozen at -20 °C for
overnight storage. To continue with the protein purification, cells were thawed in warm water
and lysed with a VibraCell sonicator (Sonics, USA) for seven cycles of 30 sec of bursts at 85%
power/30 sec on ice. Lysates were incubated with chicken egg lysozyme (Sigma, USA) at 0.2
mg/ml for 30 min at 4 °C while nutating and then spun at 18,000 rpm for 45 min in a JA25.5
rotor (Beckman Coulter, USA). The supernatants were filtered through a 60 ml syringe fitted
with a 0.8 µm filter (Thermo Scientific, USA) and then through a second 60 ml syringe fitted
with a 0.45 µm filter (Thermo Scientific, USA) into two 50 ml conical tubes. Fifteen-hundred µl
of agarose-Ni2+
-NTA bead slurry (Qiagen, Inc.) was added to a 15 ml conical tube and washed in
5 ml of lysis buffer (Table 1) by spinning at 500 × g for 2 min per wash for a total of three
washes. The washed beads were split evenly between the two 50 ml conical tubes of supernatant
and the tubes were nutated for 1.5 h at 4°C. During this time, a polypropylene 20 ml column
(Bio-Rad Laboratories, Inc.) was assembled at 4 °C and rinsed once with water and then once
with 70% ethanol. When the incubation was complete, 20 ml of bead-supernatant slurry was
transferred to the cold column and incubated for 5 min; the flow-through was then eluted into a
flask and these steps were repeated until all of the bead-supernatant slurry was loaded onto the
column. The column was then washed with 2 × 20 ml of cold PNKP lysis buffer (Table 1),
catching each flow-through in a new flask. Samples of uninduced and induced cells, supernatant
and pellet fractions, column flow-through and washes, and bead slurry from the column were
analyzed by SDS-PAGE in order to confirm the presence of hPNKP (Figure 4.6.1.a). Once the
presence of hPNKP was confirmed, 1 ml of cold PNKP lysis buffer (Table 1) and 100 μl of TEV
60
protease (5 mg/ml; F. Sicheri) were added to the column. The column was tapped to mix and
incubated overnight at 4 °C to release hPNKP from the beads. Following the incubation, the
column contents were eluted into a 1.5 ml microfuge tube. Seven additional eluate fractions were
obtained by adding 1.5 ml cold PNKP lysis buffer (Table 1) to the column and eluting into
separate 1.5 ml microfuge tubes. Samples of beads before and after TEV incubation, and all
eluate fractions were analyzed by SDS-PAGE on a 10% SDS-PAGE gel (Table 1) in order to
confirm that hPNKP had been eluted (Figure 4.6.1.b). The eluate fractions containing hPNKP
(as determined from the gel) were combined and filtered through a 10 ml syringe fitted with a
0.22 µm filter (Millipore, USA) into a 15 ml conical tube. The concentration of protein in this
eluate was determined as described above and then transferred to 12-14 kDa MW cut-off dialysis
tubing (Spectrum Labs, USA). The protein was dialyzed overnight in PNKP dialysis buffer
(Table 1) at 4 °C. Following dialysis, the protein solution was filtered through a 10 ml syringe
fitted with a 0.22 µm filter (Millipore, USA). The concentration of hPNKP was determined as
described above and then divided into 100 μl aliquots in 1.5 ml microfuge tubes for long-term
storage at -20 °C.
4.6 Purification of T4 DNA Ligase.
Competent BL21 (DE3) CodonPlus cells were transformed with 120 ng of pT4 plasmid DNA
(courtesy of F. Sicheri), plated on LB + AMP (Table 1), and grown overnight at 37 °C. Freshly
transformed E. coli were scraped into 10 ml of LB media (Table 1) and transferred to a 15 ml
conical tube. Each of six beveled flasks containing 1 L LB + AMP (Table 1) media were
inoculated with 0.25 ml of cell suspension and grown at 37 °C until the cultures reached an
OD600 of 0.4, at which point the temperature of the incubator was decreased to 18 °C. The
cultures were grown until they reached an OD600 of 0.8, at which point T4 Ligase expression was
induced by adding IPTG (Sigma, USA) to a final concentration of 100 μM. The cultures were
61
grown overnight at 18 °C and then harvested in cold JLA8.1 bottles by spinning in a JLA8.1
rotor (Beckman Coulter, USA) at 6000 rpm for 6 min at 4 °C. Cells were resuspended in 20 ml
of Ligase lysis buffer (Table 1) and transferred to cold 50 ml conical tubes, and stored overnight
at -20 °C. Cells were thawed the following day in warm water and divided into 4 tubes (~18 ml
per tube), which were then each brought up to 40 ml with cold Ligase lysis buffer (Table 1).
Sixty mg of chicken egg lysozyme (Sigma, USA), 400 µl of 8% sodium deoxycholate (Sigma,
USA), 400 µl of 100 mM PMSF (Sigma, USA) in isopropanol, and one large EDTA-free
complete protease inhibitor cocktail tablet (Roche, Canada) were added to each tube. Cells were
lysed with a VibraCell sonicator (Sonics, USA) for eight cycles of 20 sec of bursts at 85%
power/ 90 sec on ice. Lysates were centrifuged for 45 min at 18000 rpm at 4 °C in a cold JA25.5
rotor (Beckman Coulter, USA). The supernatant was filtered through a syringe fitted with a 0.8
µm filter (Thermo Scientific, USA) and then through a syringe fitted with a 0.45 µm filter
(Thermo Scientific, USA) into four 50 ml conical tubes. Three ml of agarose-Ni2+
-NTA bead
slurry (Qiagen, Inc.) was added to each conical tube and the tubes were nutated for 1 h at 4 °C.
Two polypropylene columns (Bio-Rad Laboratories, Inc.) were set up at 4 °C, washed once with
water and then once with 70% ethanol. The bead-supernatant slurry was loaded onto the columns
as described above for hPNKP, with each column containing ~3 ml of packed beads once fully
loaded. Each column was washed six times with 18 ml of cold Ligase lysis buffer (Table 1). T4
Ligase was retrieved from the column by loading 20 ml of Ligase elution buffer (Table 1) and
eluting; this step was repeated once for a total of two eluate fractions. Samples of uninduced and
induced cells, supernatant and pellet fractions, column flow-through and washes, and all eluates
were analyzed by SDS-PAGE in order to confirm the presence of T4 Ligase (Figure 4.6.1.c).
62
Figure 4.6.1 hPNKP and T4 Ligase were purified from BL21 (DE3) CodonPlus E. coli cells. (a) The
presence of hPNKP protein in cells and lysates was confirmed prior to purification of the protein with
agarose-Ni2+
-NTA beads, TEV protease cleavage, and dialysis. (b) Final fractions were checked at the
completion of purification. Eluate fractions 1-6 were combined for dialysis. (c) Fractions were checked
for T4 Ligase prior to and following the application of lysates to agarose-Ni2+
-NTA beads.
Following elution, both eultion fractions were concentrated using an Amicon Ultra 30 kDa cut-
off centriprep column (Millipore, USA) to a volume of ~ 5 ml. To do so, 10 ml of T4 Ligase
elution buffer (Table 1) was first added to the column and spun through at 3,000 × g for 10 min
at 4 °C. The buffer was discarded and the protein was added to the column and similarly spun at
3,000 × g for 10 min at 4 °C. The flow-through was spun through the column repeatedly until it
reached a volume of ~ 5 ml, and then transferred to a cold 15 ml conical tube. The chamber of
the column was then rinsed with 1 ml of T4 Ligase elution buffer (Table 1) and this was added to
the conical tube as well. The concentration of protein in the conical tube was quantified as
described above. Next, two PD-10 desalting columns (GE Life Sciences, USA) were washed five
times with 4 ml of 2X T4 Ligase storage buffer (Table 1). Two and one-half ml of protein were
poured onto each column and eluted into a single cold 15 ml conical tube. The protein was then
63
re-concentrated on an Amicon Ultra 30 kDa cut-off centriprep column (Millipore, USA) that had
been rinsed with 10 ml of 2X T4 Ligase storage buffer (Table 1). As before, the flow-through
was re-applied to the column, this time until it reached a volume of ~ 2.5 ml. The protein
concentration was quantified as above and an equal volume of sterile glycerol (Fischer Scientific,
USA) was added in order to store the purified protein in 50% glycerol at -20 °C.
4.7 Preparation of 3'-phosphorylated double-stranded DNA substrates.
Oligonucleotides were purchased from Integrated DNA Technologies (USA) or Eurofins MWG
Operon (USA). Complementary single-stranded oligonucleotides were diluted to 10-50 µM in 10
mM Tris-HCl, pH 7.5, 50 mM NaCl, and 1 mM EDTA. The tube was heated at 95 °C for 5 min
and then allowed to cool slowly at room temperature for at least 1 h.
4.8 Measurement of PNKP-dependent ligation by homogeneous time-resolved fluorescence HTRF) in a 384-well format, Version 1.
hPNKP was thawed on ice and diluted to 0.1 mg/ml in cold PNKP dilution buffer (Table 1).
HTRF buffer (Table 1) was prepared with 0.1 mg/ml BSA (Sigma, USA), 8.8 nM hPNKP and 74
nM T4 Ligase (HTRF-BLP buffer) in a 50 ml conical tube or a sterile glass bottle. HTRF-BP
buffer was prepared in an identical manner, except lacking T4 Ligase. Fifteen µl of HTRF-BLP
or HTRF-BP master mix was aliquoted into the requisite number of wells of an opaque black
polystyrene 384-well microplate (Corning, model # 3573). Five hundred nl of compound
dissolved in DMSO (Sigma, USA) was added to each reaction (or 500 nl of 100% DMSO for
controls) and incubated for 1 h at room temperature. To initiate reactions, 5 µl of SAXL-DNA
solution (Table 1) with 160 nM HTRFphos-1 (Table 2) and 160 nM HTRFphos-2 (Table 2) was
added to each well, giving final concentrations of 40 nM for each oligonucleotide. Unless the
experiment required a time course, the reaction was allowed to proceed for 1.5 h at room
temperature. To end reactions, 10 µl of DNPK solution (Table 1) was added to each well and
64
incubated for 1 h at room temperature. SA-XL and DNPK fluorophores for HTRF were obtained
from Cisbio Bioassays, USA. HTRF signal was measured on a PheraStar microplate reader
(BMG Labtech, Germany). The donor fluorophore (DNPK) was excited at 337 nm and
fluorescence emission was measured simultaneously at 620 nm (Channel B, DNPK) and 665 nm
(Channel A, SAXL). HTRF ratios were calculated as (FI665×10,000)/FI620.
4.9 Measurement of PNKP-dependent ligation by HTRF in a 384-well format, Version 2.
hPNKP was thawed on ice and diluted to 4.9 μg/ml in cold PNKP dilution buffer (Table 1).
HTRF buffer (Table 1) was prepared with 2.1 nM hPNKP and 800 nM T4 Ligase (HTRF-LP
buffer) in a 50 ml conical tube or a sterile glass bottle. HTRF-P buffer was prepared in an
identical manner, except lacking T4 Ligase. Fifteen µl of HTRF-LP or HTRF-P buffer was
aliquoted into wells of an opaque black polystyrene 384-well microplate (Corning, model #
3573). Five hundred nl of compound dissolved in DMSO (Sigma, USA) was added to each
reaction (or 500 nl of 100% DMSO for controls) and incubated for 1 h at room temperature. To
initiate reactions, 5 µl of SAXL-DNA solution (Table 1) with 160 nM HTRFphos-3 (Table 2)
and 160 nM HTRFphos-4 (Table 2) was added to each well. Unless the experiment required a
time course, the reaction was allowed to proceed for 20 min at room temperature. To end
reactions, 10 µl of DNPK solution (Table 1) was added to each well and incubated for 1 h at
room temperature. HTRF signal was measured on a PheraStar microplate reader (BMG Labtech,
Germany) using the same settings as described above.
4.10 Measurement of PNKP-independent ligation by HTRF in a 384-well format, Versions 1 and 2.
Assays were carried out as described above except that hPNKP was left out of the reactions and
the oligonucleotides used in the SAXL + DNA solutions were either 160 nM HTRFOH-1 and
65
160 nM HTRFOH-2 (Table 2) for Version 1, or 160 nM HTRFOH-3 and 160 nM HTRFOH-4
(Table 2) for Version 2.
4.11 Screening.
All mixes were prepared at the beginning of the day and aliquoted into source plates or reservoirs
[Polystyrene 96-well plates (#351177, Falcon, USA), polypropylene deep-well plates (P-DW-20-
C, Axygen, USA), or polypropylene reservoirs (RES-SW96-HP, Axygen, USA)]. If more than
one source plate was required for a given mix, or if a reservoir needed to be replenished during a
run, the remainder of the mix would be stored at 4 °C until needed.
For the first half of the screen (HTRF, Version 1), Biomek AP96 P20 and P250 tips (Beckman
Coulter, USA) were used. For the second half of the screen (HTRF, Version 2), FX-250-L-R
Maximum Recovery tips (Axygen, USA) and Biomek AP96 P20 tips (Beckman Coulter, USA)
were used. HTRF-(B)LP was dispensed with the 250 µl-capacity tips, while the SAXL + DNA
and DNPK solutions were dispensed with the 20 µl-capacity tips.
The screen was carried out on a Biomek FX platform (Beckman Coulter, USA) using a Thermo
CRS Dimension 4 system (Thermo Fisher Scientific, USA). Plates were kept in a room
temperature incubator for each incubation period. Plates were staggered in order to ensure that
incubation times were kept constant - for example, for a 30-plate screening run, there was a 7
min delay between plates. For each plate, 15 µl of HTRF-(B)LP or HTRF-(B)P mix was
dispensed from the source plate or reservoir into the wells of an opaque black polystyrene 384-
well microplate (Corning, model # 3573). Then, 0.2 µl of 1 mM compounds were pinned into the
wells of the screen plate (columns 3-22) and 0.2 µl DMSO (Sigma, USA) was subsequently
pinned from a “control” polypropylene 384-well source plate into the screen plate (columns 1-2
and 23-24). The compounds and protein(s) were incubated together for 1 h in the room
66
temperature incubator. Next, 5 µl of SAXL + DNA solution was dispensed into each well of the
screen plate and incubated in the room temperature incubator for the appropriate amount of time,
depending on the version of the HTRF assay being applied. Ten µl of DNPK solution (Table 1)
was then dispensed into each well of the screen plate and incubated for 1 h in the room
temperature incubator prior to the HTRF signal being measured on the Pherastar plate reader
(BMG Labtech, Germany). It should be noted that a 384-pin head was used for pinning
compounds and DMSO, but a 96-tip head was used with the tips for dispensing each reagent; as
such, screen reagents were aspirated and dispensed four times per plate to minimize quadrant
effects.
4.12 Calculation of z’ factor, signal-to-noise (S/N), signal-to-background (S/B), and percent error.
The z’ factor was calculated as a measure of the available signal window according to the
methods of Zhang et al. (1999), wherein µp and σp represent the mean HTRF signal and standard
deviation, respectively, for reactions containing PNKP and Ligase (positive control). µn and σn
represent the mean HTRF signal and standard deviation, respectively, in reactions containing
PNKP only (negative control).
z' factor = 1 - [(3σp + 3σn)/(µp - µn)]
S/N = (µp - µn)/√(σp2 + σn
2)
S/B = µp ÷ µn
% error = (σ × 100) ÷ µ
67
4.13 Calculation of Tanimoto coefficient.
The SMILES codes for the compounds of interest were entered together into an online similarity
matrix calculator with default fingerprint settings of 1024 bit length and a search depth of 6
(Guha, 2012).
4.14 Preparation of polyacrylamide sequencing gels.
One large (41.9 cm × 33.3 cm) and one small (39.4 cm × 33.3 cm) siliconized optically clear gel
plate (Gel Company, USA) were aligned with 0.25 mm spacers (Gel Company, USA) between
them, and then clamped together with bulldog clips. The bottom of the gel was sealed with vinyl
tape (VWR, USA). 12% polyacrylamide gel mix (Table 1) was poured between the plates either
with a syringe fitted with an 18-gauge needle or directly from a glass beaker. A 20- or 42-well
0.25 mm comb (Gel Company, USA) was placed between the plates and the top of the gel was
clamped as well. The gel was allowed to polymerize overnight at room temperature.
4.15 Phosphatase gel assay.
hPNKP was thawed on ice and diluted to 4.9 μg/ml in cold PNKP dilution buffer (Table 1).
HTRF-P buffer was prepared in a microfuge tube with 2.1 nM hPNKP. For reactions with T4
PNK, the HTRF-P buffer was prepared in an identical manner except with 5.7 nM T4 PNK in the
place of hPNKP. HTRF-NEG buffer was prepared in an identical manner, except lacking
enzyme. Seventeen and one-half µl of HTRF-P or HTRF-NEG buffer was aliquoted into
microfuge tubes or a generic PCR plate (Sarstedt, Germany). Five hundred nl of compound
dissolved in DMSO (Sigma, USA) was added to each reaction (or 500 nl of 100% DMSO for
controls) and incubated for 1 h at room temperature. To initiate reactions, 2 µl of 1.2 µM
GELphos DNA (Table 2) was added to each well. Unless the experiment required a time course,
the reaction was allowed to proceed for 20 min at room temperature. Reactions were stopped by
adding 20 µl of 2X formamide loading buffer (90% formamide, 25 mM EDTA, 0.02%
68
bromophenol blue, 0.02% xylene cyanol) and heating for 5 min at 95 °C. Four µl (0.2 pmol of
DNA) from each reaction was loaded into a 12% polyacrylamide sequencing gel, which had
been pre-run for 1 h at 40 W. Samples were run in 1X TBE (Table 1) at 40 W for 3 h or until the
bromophenol blue dye reached the bottom of the gel. DNA was visualized by scanning in the
Cy5 channel with a Typhoon 9400 variable mode imager (GE Healthcare, USA) and the
percentage of dephosphorylation was quantified with ImageQuant 5.2 software (Molecular
Dynamics, USA).
4.16 Fluorescence polarization (FP) substrate binding assay.
hPNKP-D171A was thawed on ice. Two-times FP-PNKP buffer was prepared in a microfuge
tube by diluting hPNKP-D171A to 50 nM in FP buffer (Table 1). FP-NEG buffer (2X) was
prepared in an identical manner, except lacking hPNKP. Twelve and one-half µl of 2X FP-PNKP
buffer was aliquoted into wells of an opaque black polystyrene 384-well microplate (Corning,
model # 3573) and was incubated with 0.5 µl compound dissolved in DMSO (or 0.5 µl 100%
DMSO for controls) for 1 h at room temperature. FPphos DNA (Table 2) was diluted to 25 nM
in FP buffer (Table 1) and 12.5 µl was added to all reactions and incubated for 15 min at room
temperature. Fluorescence polarization was measured with an Analyst HT (Molecular Devices,
USA) using the fluorescein filter (samples excited at 485 nm and emission measured at 530 nm).
4.17 Measurement of the Kd of hPNKP-D171A in the FP substrate binding assay.
The fluorescence polarization assay was carried out as above except with the final concentration
of hPNKP titrated from 5 nM to 10 µM, and the percentage of substrate bound was determined
by FP (Figure 2.9.1). GraphPad 5.0 (Prism, USA) software was used to analyze the data, and the
Kd for this reaction was found to be 41.6 nM (Figure 2.9.1).
69
Figure 4.17.1 FP reactions were conducted with hPNKP for proof of concept and to establish optimal
reaction conditions. FP reactions were conducted with 5 nM – 10 µM hPNKP, and with a Kd of 41.6 nM,
50 nM was selected as the concentration of hPNKP at which to perform dose response experiments with
inhibitors.
4.18 Reversibility assay.
hPNKP was thawed on ice and diluted to 0.15 mg/ml (2.6 µM) in HTRF Buffer (Table 1). The
diluted hPNKP was incubated at 100 nM with 0.5 µl compound in HTRF buffer (Table 1) for 30
min at room temperature. The concentration of compound for this incubation was 10-times its
IC50 for 2 nM hPNKP. Five µl of hPNKP with compound was added to 495 µl of SAXL + DNA
solution (Table 1) containing 40 nM HTRFphos3 (Table 2) and 40 nM HTRFphos4 (Table 2),
with 800 nM T4 Ligase in HTRF buffer (Table 1) and quickly inverted a few times to mix.
Twenty µl of reaction mix was aliquoted into the appropriate number of wells of an opaque black
polystyrene 384-well microplate (Corning, model # 3573) and reactions were stopped at
appropriate time points with the addition of 10 µl of REV-SD solution (Table 1). Reactions were
incubated with REV-SD solution for 1 h at room temperature and HTRF signal was measured
with a PheraStar microplate reader (BMG Labtech, Germany) as before.
4.19 Tissue culture.
U2OS human osteosarcoma cells (Ponten and Saksela, 1967) were maintained in McCoy's
medium (Gibco, USA) supplemented with 5% fetal bovine serum (Gibco, USA), 10 IU penicillin
and 0.1 mg/ml streptomycin (Multicell Technologies, Inc., USA). Cells were maintained at 37
70
°C and 5% CO2. Cells were split when they approached 80-90% confluency by treating with a
0.25 % trypsin-EDTA solution (Gibco, USA) and transferring to new flasks.
4.20 ATP Lite cell viability assay.
Cells were lifted from a non-confluent flask by trypsinizing for no more than five minutes, and
transferred to a sterile 15 ml conical tube. The concentration of cells was determined using a
hemocytometer and the appropriate amount of cells were diluted into 10 ml of warm media to
give 15,000 cells/ml. 100 μl of diluted cells (1500 cells) were seeded in the requisite number of
wells of an opaque, white, tissue culture-treated 96-well polystyrene Costar microplate (Corning,
model #3917) using a multichannel pipette and a sterile single-use polypropylene basin. Cells
were then incubated for 24 h at 37 °C/5% CO2 to allow them to attach to the plate. The following
day, hPNKP inhibitors and CPT were diluted to a 500X concentration in tissue culture-quality
DMSO (Sigma, USA) by serial dilution in a biological safety tissue culture cabinet. To prepare
media for treatments, 1 μl of compound (or DMSO for controls) was added for every 500 μl of
media and vortexed thoroughly. After the 24 h incubation, media was suctioned off and 100 µl of
media containing compounds and/or DMSO was added to the appropriate wells in duplicate.
Cells were returned to 37 °C/5% CO2for the 96 h drug treatment. To measure viability, the ATP
Lite cell viability kit (PerkinElmer, USA) was used as directed. ATP Lite Lysis buffer and ATP
Lite Substrate were thawed slowly until they reached room temperature. Fifty μl of Lysis buffer
was added to each well with a multichannel pipette and the plate was shaken at 700 rpm for 5
min. Then, 50 μl of Substrate was added to each well by multichannel pipette and the plate was
shaken again at 700 rpm for 5 min. Chemiluminescence was measured with an EnSpire 2300
multi-label plate reader (PerkinElmer, USA) after a 10 min delay to dark-adapt the plate.
71
Tables
Table 1 Buffers
Coommassie dye
solution
0.2% Coommassie Brilliant Blue R-250, 40% methanol
De-stain solution 30% methanol, 5% glacial acetic acid
DNPK solution (3X) 5 ng DNPK, 0.02% Tween-20, 0.5X PBS (1.2 M KF, Version
1; and 150 mM EDTA pH 8.0, Version 2)
HTRF Buffer (1X) 50 mM Tris-HCl, pH 7.5, 10 mM MgCl2, 10 mM DTT, 1mM
ATP, 0.02% Tween-20
FP buffer (1X) 50 mM Tris-HCl, pH 7.5, 10 mM MgCl2, 10 mM DTT, 0.2%
Tween-20
Luria broth (LB) 0.5% yeast extract, 1% NaCl, 1% tryptone peptone
PBS (1X) 137 mM NaCl, 7.8 mM Na2HPO4, 2.7 mM KCl, 1.5 mM
KH2PO4
PNKP dialysis
buffer
50 mM Tris-HCl, pH 8.0, 150 mM NaCl, 1 mM DTT, 20%
glycerol
PNKP dilution
buffer (1X)
50 mM Tris-HCl, pH 7.0, 100 mM KCl, 1 mM EDTA, 1mM
DTT
PNKP lysis buffer 50 mM Tris-HCl, pH 8.0, 500 mM NaCl, 200 μM PMSF,
0.04% β-mercaptoethanol, and two EDTA-free complete
protease inhibitor cocktail tablets (Roche) per 200 ml buffer
SAXL solution 50 ng SAXL, 1X PBS
SAXL-DNA
solution (4X)
50 ng SAXL, 4X DNA substrates, TE pH 7.0
REV-SD solution 71 ng SA-XL, 5 ng DNPK, 50 mM EDTA, 0.02% Tween; in
1X PBS
SDS-PAGE running
buffer (1X)
25 mM Tris, 200 μM glycine, 0.1% SDS
SDS-PAGE sample
buffer (1X)
83 mM Tris-HCl, pH 6.8, 1% SDS, 0.003% bromophenol
blue, 10% glycerol, 10 mM DTT
SDS-PAGE
separating gel
375 mM Tris-HCl, pH 8.8, 0.1% SDS, 0.05% ammonium
persulfate, 0.1% TEMED, 40% acrylamide:bisacrylamide
(29:1) to desired final concentration
SDS-PAGE stacking
gel
0.06 M Tris-HCl, pH 6.8, 0.1% SDS, 0.1% ammonium
persulfate, 0.1% TEMED, 40% acrylamide:bisacrylamide
(29:1) to desired final concentration
T4 Ligase buffer
(1X)
50 mM Tris-HCl, pH 7.5, 10 mM MgCl2, 10 mM DTT, 1 mM
ATP
T4 Ligase elution
buffer
50 mM sodium phosphate, pH 8.0, 300 mM NaCl, 300 mM
imidazole, 0.1% β-mercaptoethanol
T4 Ligase lysis
buffer
50 mM sodium phosphate, pH 8.0, 300 mM NaCl, 10 mM
imidazole, 0.1% β-mercaptoethanol
72
Table 3 Buffers (continued)
T4 Ligase storage
buffer (1X)
10 mM Tris-HCl, pH 7.5, 50 mM KCl, 1 mM DTT, 50%
glycerol
T4 Ligase storage
buffer without
glycerol (2X)
20 mM Tris-HCl, pH 7.5, 100 mM KCl, 2 mM DTT
TBE buffer (1X) 89 mM Tris base, 89 mM boric acid, 10 mM EDTA
TE buffer 10 mM Tris-HCl, pH 7.0-8.0 as needed, 1 mM EDTA
73
Table 2 Sequences of oligonucleotide substrates (5' to 3')
Bio14 ATTACCGTAATCAG-biotin 14 nt
Bio2 pATTACCGTAATCATG-biotin 15 nt
Cy5PNK Cy5-TAATCGAGCTCGAATTCACT 20 nt
DNP11OH DNP-GAATTCAGTCG 11 nt
DNP11phos DNP-GAATTCAGTCGp 11 nt
DNPOH DNP-AGTGAATTCGAGCTCG 16 nt
DNPphos DNP-AGTGAATTCGAGCTCGp 16 nt
FPphos 6-FAM-TCCTCp 5 nt
GELphos Cy5PNK + Oligo1phos annealed 20 nt dsDNA
HTRFOH-1 DNPOH + Oligo5 annealed 20 nt dsDNA
HTRFOH-2 Bio2 + Oligo2OH annealed 15 nt dsDNA
HTRFOH-3 DNP11OH + Oligo15 annealed 15 nt dsDNA
HTRFOH-4 Bio14 + Oligo10OH annealed 14 nt dsDNA
HTRFphos-1 DNPphos + Oligo5 annealed 20 ntdsDNA
HTRFphos-2 Bio2 + Oligo2phos annealed 15 nt dsDNA
HTRFphos-3 DNP11phos + Oligo15 annealed 15 nt dsDNA
HTRFphos-4 Bio14 + Oligo10phos annealed 14 nt dsDNA
Oligo1phos AGTGAATTCGAGCTCGp 16 nt
Oligo2OH pCATGATTACGG 11 nt
Oligo2phos CATGATTACGGp 11 nt
Oligo5 pTAATCGAGCTCGAATTCACT 20 nt
Oligo10OH pCTGATTACGG 10 nt
Oligo10phos pCTGATTACGGp 10 nt
Oligo15 pTAATCGACTGAATTC 15 nt
74
Table 3 Original hit compounds
Compound Name Structure
OICR1
OICR2
OICR3
OICR4
OICR5
75
Table 1 Original hit compounds (continued)
OICR6
OICR7
OICR8
OICR9
76
Table 4 Analog compounds
Compound Name Structure
OICR7A
OICR9A
OICR9B
OICR79
OICR791
OICR792
77
Table 2 Analog compounds (continued)
OICR793
OICR794
OICR795
OICR796
Table 4 Analog compounds Functional groups in red represent groups that have been modified relative
to the parent compound, OICR79, for the purpose of structure-activity relationship analysis.
70
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Appendices
I. PNKP inhibitors
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II. Analogs that did not effectively inhibit PNKP