Fhit Inactivation in Neoplasia: in Vitro Insights Into Cancer Initiation and Progression

Fhit inactivation in neoplasia: in vitro insights into cancer initiation and progression

DISSERTATION

Presented in Partial Fulfillment of the Requirements for the Degree Doctor of Philosophy in the Graduate School of The Ohio State University

Jenna Rose Karras

Graduate Program in Molecular, Cellular and Developmental Biology

The Ohio State University

2016

Dissertation Committee:

Kay Huebner, PhD, Advisor

Joanna Groden, PhD

Amanda Toland, PhD

Debbie Parris, PhD

Mark Parthun, PhD

Copyrighted by

Jenna Rose Karras

2016

Abstract

The fragile FHIT gene, encompassing one of the most active common fragile sites, FRA3B, is frequently altered in preneoplasia and cancer, through gene rearrangements or silencing by hypermethylation of regulatory sequences. Silencing of

Fhit protein expression through activation of fragile gaps and breaks in the FRA3B locus, or locus hypermethylation, causes thymidine kinase down-regulation and dNTP imbalance, resulting in spontaneous replication stress that leads to chromosomal aberrations, allele copy number variations, small insertions/deletions and single-base substitutions. Thus, Fhit, which is reduced in expression in the majority of human cancers, is a genome „caretaker‟ whose loss initiates genome instability in preneoplastic lesions creating the “soil” for cancer development. However, some scientists argue that

Fhit is lost in cancers simply because of its position at a fragile locus that is very susceptible to breakage. This skepticism has hindered consideration of Fhit-associated therapeutic targets for the majority of human cancers exhibiting Fhit loss.

Here we followed this process from loss of Fhit genome caretaker function, through development of genetic alterations to isolation of clones with tumorigenic potential. We established epithelial cell lines from kidney tissues of Fhit-/- and Fhit+/+ mice early after weaning of pups, and subjected some cell lines to nutritional and carcinogen stress; the Fhit+/+ cell lines did not survive either of these additional stresses.

In contrast to +/+ cells, loss of Fhit leads to alterations in apoptotic and EMT signaling ii pathways and oncogene activation. These alterations allow for transformation, selective clonal expansion and development of invasive properties in vitro as well as tumor formation and metastasis in vivo. Thus, Fhit loss-induced genome instability results in alterations in genes and gene expression associated with preneoplastic changes, a downstream consequence of TK1 inactivation, to promote neoplastic progression.

Because deregulation of the Fhit-TK1 pathway results in changes in expression patterns promoting cellular transformation, we also investigated the mechanism of positive regulation of TK1 by Fhit. We demonstrate that loss of Fhit expression leads to decreased TK1 mRNA translation possibly due to accumulation of 5‟-cap dinucleotides that can inhibit cap-dependent translation. Uncovering the mechanism by which Fhit regulates TK1 mRNA has important implications for understanding how Fhit modulates many cancer-associated processes through its function as a tumor suppressor and genome caretaker. Collectively, these findings support a model where deregulation of the Fhit-

TK1 pathway initiates global genome instability in preneoplastic cells to promote cancer development.

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To my husband, Randy Soukup, for all the things you have done to help me, support me,

surprise me and to make me happy.

To my parents for their love and support, for teaching me to never quit and inspiring me

to always do my best.

Acknowledgments

I would especially like to thank my advisor, Kay Huebner, for guiding, supporting and mentoring me throughout my graduate research years. Your constant encouragement and patience while I conquered my fear of public speaking was very much appreciated. I will always remember the confidence-boosting meetings we had prior to many of my presentations - they always put pep in my step!

I thank my thesis committee members, Drs. Joanna Groden, Amanda Toland,

Debbie Parris and Mark Parthun, for all of their guidance through this process: your discussion, ideas, and feedback have been absolutely invaluable.

I also thank current and former members of Dr. Huebner‟s lab for creating a happy and helpful atmosphere to work and learn. I would especially like to thank Morgan

Schrock for all her assistance with mouse projects, Catherine Waters for listening to every practice presentation and Teresa Commisso for her help with troubleshooting experiments and submitting manuscripts.

Vita

October 15, 1987 ...... Born – River Falls, Wisconsin

August 2005 ...... River Falls High School

August 2010 ...... B.S. Biology, University of Wisconsin,

Eau Claire, Wisconsin

September 2010 to present ...... Graduate Research Associate, The Ohio

State University, Columbus, Ohio

Publications

1. Karras JR, Batar B, Schrock M, Ouda IM, Zhang J, La Perle K, Druck T, Huebner K, Fhit loss-associated initiation and progression of neoplasia in vitro. Cancer Science. (submitted).

2. Karras JR, Schrock MS*, Batar B*, Huebner K, Fragile genes that are frequently altered in cancer: players not passengers. Cytogenetics and Genome Research. (In review).

3. Waters CE*, Ouda IM*, Kiss DL*, Saldivar JC, Karras JR, et al. Identification of Fhit as a post-transcriptional effector of thymidine kinase 1 expression. (In revision).

4. Paisie CA, Schrock MS, Karras JR, Zhang J, Miuma S, Ouda IM, Waters CE, Saldivar JC, Druck T, Huebner K, Exome-wide single-base substitutions in tissues and derived cell lines of the constitutive Fhit knockout mouse. Cancer Sci. 2016. doi: 10.1111/cas.12887. [Epub ahead of print]. PMID: 26782170.

5. Karras JR*, Paisie CA*, Huebner K. Replicative stress and the FHIT gene: roles in tumor suppression, genome stability and prevention of carcinogenesis. Cancers (Basel). 2014; 6(2):1208-19. doi: 10.3390/cancers6021208. PMID: 24901304.

6. Miuma S, Saldivar JC, Karras JR, et al. Fhit deficiency-induced global genome instability promotes selective mutation and clonal expansion. PLoS ONE. 2013; 8(11):e80730 doi: 10.1371/journal.pone.0080730. eCollection 2013. PMID: 24244712.

Fields of Study

Major Field: Molecular, Cellular and Developmental Biology

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Table of Contents

Abstract ...... ii

Dedication……………………………………………………………………………… iv

Acknowledgments...... v

Vita ...... vi

List of Tables ...... xi

List of Figures ...... xiiii

Chapter 1: Introduction ...... 1

1.1 Common fragile sites and cloning of the FHIT gene……………………..1

1.2 Cancer and the FRA3B/FHIT locus………………………………….……2

1.3 The FHIT gene and protein……………………………………………….5

1.4 Fhit as a tumor suppressor………………………………………………..6

1.5 Fhit as a genome caretaker………………………………………………..8

1.6 Conclusions………………………………………………………………13

Chapter 2: Fhit loss-associated initiation and progression of neoplasia in vitro……………………………………………………….14

2.1 Introduction………………………………………………………………14

2.2 Materials and methods………………………………………………….16

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2.3 Results……………………………………………………………………23

2.3.1 In vitro model of Fhit loss-associated neoplastic progression...…23

2.3.2 Fhit-/- cells exhibit alterations in apoptotic and EMT signal pathways…………………………………………….….25

2.3.3 Fhit loss-associated cell transformation…………………………29

2.3.4 Classification of genes with altered transcription in a Fhit-/- NS cell line……………….…………………………32

2.3.5 NS3T cells display tumorigenic and metastatic potential……..34

2.3.6 Induced Fhit expression delays tumor onset in vivo…………. 38

2.4 Discussion………………………………………………………………..39

Chapter 3: The Fhit-TK1 signal pathway…………………………………………..43

3.1 Introduction………………………………………………………………43

3.2 Materials and methods………………………………………………….44

3.3 Results……………………………………………………………………50

3.3.1 Thymidine supplementation prevents ongoing DNA damage in Fhit-/- cells…………………………………...... 50

3.3.2 Mutation burden is increased in Fhit-/- kidney cells and tissue…………………………………………………..53

3.3.3 A post-transcriptional role for Fhit………………………………56

3.3.4 Fhit affects ribosome binding and translation of TK1 mRNA…………………………………………………..58

3.4 Discussion……………………………………………………………………….61

Chapter 4: Discussion………………….……………………………………………..69

4.1 Introduction……………………………………………………………..69

4.2 Initiation of genome instability through loss of the FRA3B gene product, Fhit……………………………………………….71

4.3 Expression of Fhit and wt p53 are lost coordinately in many cancers……………………………………………74

4.4 Fhit loss and the mutator hypothesis…………………………………..75

4.5 The Fhit-TK1 pathway in preneoplasia vs cancer……………………..77

4.6 Conclusions……………………………………………………………..80

References……………………………………………………………………………….82

List of Tables

Table 1. Antisera used, dilutions and applications ...... 17

Table 2. Primers used for gene amplification ...... 18

Table 3. Establishment of mouse kidney cell lines………………………………………24

Table 4. Protein expression changes in Fhit-/- and +/+ kidney cell lines……………….28

Table 5. Protein expression in Fhit-/- DS, NS and colony-forming cell lines…………...28

Table 6. Summary of tumorigenicity analyses…………………………………………..38

Table 7. Gene-specific primer pairs for validated SBSs…………………………………47

Table 8. Sequence analysis of Trp53, E2f1, Mcm6 and Herc2………………………..55

List of Figures

Figure 1. The FRA3B/FHIT locus ...... 3

Figure 2. Fhit-mediated apoptotic pathways ...... 7

Figure 3. Model for Fhit loss-associated neoplastic progression ...... 12

Figure 4. Fhit-/- mouse kidney cells exhibit signal pathway alterations ...... 26

Figure 5. Wild-type p53 overexpression restores p21 protein expression ...... 27

Figure 6. Transformation-associated features of Fhit-/- cells ...... 31

Figure 7. Classification of genes with altered transcription in NS3 colony cell line relative to -/- kd3 p48 ...... 33

Figure 8. Mesenchymal spindle cell neoplasm phenotype of NS3 colony tumors ...... 35

Figure 9. NS3T cells display high tumorigenic and metastatic potential……………….36

Figure 10. Thymidine supplementation prevents ongoing DNA damage ...... 52

Figure 11. Increased total single base substitutions (SBSs) and C>T and T>C mutations in Fhit-/- cells and tissues ...... 54

Figure 12. Fhit regulation of TK1 mRNA levels ...... 57

Figure 13. Impact of Fhit on polysome binding of TK1 mRNA ...... 60

Figure 14. Fhit hydrolyzes free m7GpppN cap structures… ...... 64

Figure 15. Model for Fhit as a scavenger decapping enzyme regulating TK1 mRNA translation ...... 66

Figure 16. TK1 overexpression in Fhit-negative lung tumors……………………….…78

Figure 17. TK1 up-regulation in late passage mouse kidney cells……………………..79

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Chapter 1: Introduction

Summarizes the history and background of studies of the FHIT locus and gene product;

this summary contains portions from the following manuscripts:

Karras JR et al., Cytogenet Genome Res. 2016 (in review)

Karras JR et al., Cancers (Basel) 2014; 6(2):1208-19. doi: 10.3390/cancers6021208.

1.1 Common fragile sites and cloning of the FHIT gene

Common fragile sites (CFSs) are specific, conserved chromosomal regions that are highly sensitive to replication stress. They were described in the early 1980‟s as chromosomal loci that show breaks, gaps, and rearrangements that are non-randomly distributed in a variety of normal and cancer cells under conditions that impair DNA replication (Glover et al., 2005). CFSs are expressed in all individuals, as opposed to rare, inherited fragile sites, which affect less than 5% of the human population; FRAXA is an example of an inherited fragile site associated with a form of hereditary mental retardation. CFSs are also mostly late-replicating regions of the genome that lack internal, well-spaced replication origins (Letessier et al., 2011). This requires the flanking replication forks to travel long distances, distances that cannot be covered upon fork slowing, predisposing these regions to breakage. Around the time of fragile site discovery, it was also recognized that human cancers exhibit nonrandom chromosomal 1 alterations. Multiple studies reported that some fragile site breaks mapped to the same cytogenetic location as cancer breakpoints suggesting a biological relationship between fragile sites and cancer development (Hecht and Sutherland, 1984; Yunis and Soreng

1984). As a result, the hunt for tumor suppressor genes located at common fragile sites began. In 1996, the Huebner laboratory identified the first gene to encompass a fragile site, naming the gene FHIT (Fragile Histidine Triad) after its location at the FRA3B fragile locus and its sequence homology to the HIT superfamily of enzymes (Ohta et al.,

1996). Although subsequent investigations characterized FHIT as a cancer suppressor, the inherent instability of CFS loci and the lack of inactivating point mutations within fragile gene products, led some scientists to dismiss the idea that deletions within common fragile regions contribute to clonal expansion of neoplastic clones. 20 years and over 1060 publications later, the debate continues. Are these deletions found in cancers because of their extreme sensitivity to replication stress, or do deletions at some CFSs impart a selective advantage for cancer progression?

1.2 Cancer and the FRA3B/FHIT locus

Aberrant FHIT transcripts are detected in many human cancers and are often due to genomic deletions within a core region of FRA3B. This core region of FRA3B overlaps FHIT exons 4-6 (Durkin et al., 2008). Therefore, cancer deletion endpoints that occur within this core region disrupt the first protein coding exons of Fhit, resulting in reduced or lost expression of the Fhit protein (Figure 1) affecting one or both alleles.

Figure 1. The FRA3B/FHIT locus. (a, left panel) Fluorescent image of chromosome 3 showing the gaps at fragile site FRA3B. (a, right panel) Fluorescent in situ hybridization (FISH) detection of the FHIT locus at FRA3B. Yellow arrows point to the visible gap. (b) Schematic of the FHIT gene at human chromosome 3p14.2. Numbers 1-10 designate the 10 exons, and the black shaded exons 5-9 indicate the protein-coding exons. Horizontal red lines indicate deletion break points commonly seen in cancer cells. The green dotted boxed area marks the FRA3B “core” region where most deletions occur in cancer cells, although the fragility of the locus extends further in both directions.

Silencing of Fhit protein expression can also occur in preneoplastic or neoplastic cells through CpG methylation within the FHIT gene regulatory region. Loss of Fhit protein expression is very frequently observed in epithelia-derived cancers of the lung, esophagus, throat, stomach, breast, skin, pancreas, cervix and kidney.

Many investigations have reported frequent genetic alterations at the

FRA3B/FHIT locus in benign, premalignant and malignant lesions of many types of cancer. Allelic imbalances within the FHIT gene not only occur frequently in early hyperplastic lesions of the lung, but FHIT alterations also precede other allelic imbalances at other sites (Gorgoulis et al., 2005; Bartkova et al., 2005). Loss of heterozygosity (LOH) at the FHIT locus occurs in 76% of esophageal primary tumors, which correlates with lost or reduced Fhit expression. Following Fhit expression during the pathogenesis of esophageal carcinoma, loss of Fhit can be observed in 67% of carcinoma in situ and 43% of dysplastic lesions (Mori et al., 2000). Aberrant FHIT transcripts are also detected in 86% of Barrett‟s metaplasia, a precancerous disorder that precedes the development of esophageal adenocarcinoma (Michael et al., 1997). In ductal carcinoma in situ (DCIS), the most common type of non-invasive breast cancer, reduced Fhit expression is observed in 70% of pure DCIS, with 20% of cases exhibiting decreased expression in the adjacent normal tissue (Guler et al., 2005). LOH accounts for reduced Fhit expression in up to 25% of intraductal hyperplasias (Ahmadian et al.,

1997).

The FHIT locus is also highly sensitive to environmental carcinogens. Loss of

Fhit expression has been reported in up to 93% of precancerous lesions of lung squamous cell carcinoma, with Fhit loss more commonly observed among smokers vs nonsmokers

(Sozzi et al., 1998). LOH at the 3p locus has even been identified in the normal tissue of lung cancer patients and in normal bronchial epithelium of smokers (Mori et al., 2000;

Mao et al., 1997; Wistuba et al., 1997). Loss of Fhit expression has also been documented

4 for preneoplastic lesions of the cervix (Birrer et al., 1999, Butler et al., 2002), oral cavity

(Virgilio et al., 1996, Yuge et al., 2005), and other organs (Luan et al,. 1998; Ozkara et al., 2005; Velickovic 1999) suggesting involvement of loss of Fhit expression in the earliest stages of cancer development in many cancers.

1.3 The FHIT gene and protein

The FHIT gene, located at 3p14.2 encompasses one of the most active common fragile sites, FRA3B. FHIT is a 10-exon gene that spans over 1 megabase of DNA.

Despite being a very large gene, exons 5-9 encode for a small messenger RNA (1.1 kb) that is translated into a 147-amino acid protein (16.8 kDa) (Huebner et al., 1998). Fhit belongs to a family of nucleotide hydrolases and transferases characterized by a conserved histidine triad motif, His-x-His-x-His-xx where x represents a hydrophobic amino acid. Hint, aprataxin, DCPS and GalT are other members of this family of enzymes (Martin et al., 2011). Fhit was initially shown to function as an Ap3A hydrolase, cleaving Ap3A into adenosine 5‟-diphosphate and AMP (Barnes et al., 1996). Recent studies have provided an additional role for Fhit as a scavenger mRNA decapping enzyme (Taverniti and Seraphin, 2014), working in collaboration with DCPS to degrade methylguanosine (m7Gpppn) 5‟ cap dinucleotides that are generated by 3‟ to 5‟ degradation of mRNA bodies (Li and Kiledijan, 2010).

Although Fhit was first identified as a cytosolic protein, it can also be directed to the mitochondrion through interaction with chaperones Hsp60 and Hsp10. Fhit is expressed in most adult tissues, but is most abundantly expressed in the epithelial cells

5 lining or within most organs. During development, murine Fhit is highly expressed in organs derived from the endoderm, such as the bronchi, trachea, esophagus, stomach and intestine (Falvella et al., 2000).

1.4 Fhit as a tumor suppressor

Due to the initial finding that FHIT is homozygously deleted in many cancers and cancer cell lines, which is accompanied by loss or reduction of Fhit protein expression, the gene was characterized as a cancer suppressor soon after it was cloned in 1996.

Subsequent studies provided additional evidence for this suppressor role. Fhit knockout mice are more susceptible to cancer than wild-type (wt) mice, as Fhit knockout mice develop more spontaneous and many more carcinogen-induced tumors (Fong et al., 2000,

Zanesi et al., 2001). Overexpression of exogenous FHIT in Fhit-negative cell lines was reported to suppress xenograft tumor growth in mice (Siprashvili et al., 1997).

Furthermore, viral-mediated FHIT gene therapy prevents and reverts carcinogen-induced tumor development in a mouse gastric cancer model via induction of caspase-dependent apoptosis (Dumon et al., 2001; Ishii et al., 2001). In response to oxidative stress, Fhit is imported into the mitochondrion where it interacts with and stabilizes ferredoxin reductase. In conjunction with Fhit overexpression, this interaction leads to enhanced intracellular ROS production, triggering apoptosis (Trapasso et al., 2008; Pichiorri et al.,

2009; Okumura et al., 2009). Fhit can also influence calcium uptake by mitochondria to promote apoptosis (Rimessi et al., 2009). In response to genotoxic stress, Fhit participates

Figure 2. Fhit-mediated apoptotic pathways. In response to oxidative stress, Fhit protein localizes to the mitochondria via an Hsp complex where it interacts with and stabilizes ferredoxin reductase, leading to enhanced production of reactive oxygen species, stimulation of cytochrome c release and subsequent activation of the caspase cascade. In response to genotoxic stress, Fhit participates in the checkpoint response to DNA damage through Chk1 activation to commit cells to cell cycle arrest or apoptosis. Importantly, Fhit-deficient cells are resistant to checkpoint activation and cell death in response to oxidative and genotoxic agents and thus develop preneoplastic changes. (Image from Karras et al., 2014; copyright license http://creativecommons.org/licenses/by/3.0/).

in the checkpoint response to DNA damage via Chk1 to commit cells to cell cycle arrest and apoptosis (Figure 2).

As an additional mechanism for tumor suppression, many reports have implicated

Fhit in the regulation of tumor invasion, epithelial-mesenchymal transition (EMT) and metastasis (Joannes et al., 2010; Bekar et al., 2007; Zhaoet al., 2002). One study found that silencing Fhit promotes slug-dependent invasion of lung tumor cells by inducing expression of EMT-related genes, MMP-9 and Vimentin, via the EGFR/Src/ERK/Slug

7 signaling pathway (Joannes et al., 2014). Another study demonstrated that Fhit expression minimizes invasiveness and metastatic potential of lung cancer cells through microRNA(miR-30c)-mediated suppression of EMT (Suh et al., 2014). While mechanisms underlying these Fhit-associated functions have not been fully defined, the results of these studies show that Fhit is a negative regulator of EMT minimizing the invasiveness and metastatic potential of cancer cells. Interestingly, the tumor suppression function of Fhit is independent of its in vitro hydrolytic activity but dependent on binding of its substrate, though this finding has not been confirmed for a known in vivo substrate

(Trapasso et al., 2003). Mutation of the central H residue of the histidine triad to N allows tight binding of the FHIT-substrate complex but abrogates cleavage of the substrate. This mutant functions nearly as well as a tumor suppressor as wt protein

(Siprashvili et al., 1997). However, mutations disrupting Fhit-substrate binding do not retain suppressor activity, suggesting that the Fhit-substrate complex is the active signaling molecule for tumor suppression.

1.5 Fhit as a genome caretaker

In addition to its tumor suppressive function, recent evidence has highlighted a new role for Fhit in protecting the genome. Genome instability is a hallmark of human cancer and includes chromosomal instability, microsatellite instability and point mutations. Chromosomal instability refers to anomalies in chromosome number

(aneuploidy) and structure (transversions, deletions, translocations and duplications) and is the main form of instability observed in many epithelial tumors (Bayani et al., 2008;

Gagos et al., 2005). Data has shown that loss of Fhit causes genome instability through activation of other fragile loci and induction of replication stress.

Hosseini et al. (2013) showed that loss of Fhit increases the fragility of other CFS.

Fhit-deficient mouse kidney epithelial cells exhibit a two-fold increase in chromosomal breaks and gaps at fragile loci compared to wild-type cells. Upon silencing of Fhit in human mammary epithelial cells, MCF10A, the cells displayed the same elevated frequency of fragile breaks, at other fragile loci encompassing possible tumor suppressor genes reported to be deleted in cancer cell lines (Hosseini et al., 2013).

Besides increased breaks at CFS loci, Fhit loss also causes increased DNA double strand breaks (DSBs) through replication stress. Saldivar et al. (2012) compared cells with Fhit knockdown to normal, transformed and cancer cells expressing endogenous

Fhit. In all cell types, decreased expression of Fhit resulted in increased levels of spontaneous DNA breaks due to replication fork stalling and collapse. Nucleotide pool analysis revealed that dTTP levels were significantly reduced upon loss of Fhit protein expression and thus unable to support efficient DNA replication. Specifically, Fhit regulates thymidine kinase (TK1) expression level for dTTP pool production at the G1/S border to prepare cells for DNA replication. Importantly, the DNA damage generated from the replication defects in Fhit-deficient cells does not activate cell cycle checkpoints or induce apoptosis, allowing spontaneous DNA damage induced by Fhit loss to be transmitted through mitosis to daughter cells, leading to ongoing genome instability. Fhit loss-induced replication stress eventually leads to the formation of micronuclei and aneuploidy, evidence of this ongoing instability.

Other evidence of ongoing instability includes copy number alterations (CNAs).

Gains or losses spanning >10 kb were identified in three Fhit+/+ and Fhit-/- mouse embryo fibroblast (MEF) cell lines at pre- and post-senescence. CNAs were more frequently observed in Fhit-/- MEFs compared to Fhit+/+ MEFs, where only one CNA was detected.

Two of the three Fhit-/- MEFs exhibited allelic gains within the Mdm2 gene (Miuma et al.,

2013) an oncogene involved in cellular transformation. Mdm2 mRNA expression was also observed to be elevated ~4 fold. CNAs were also detected in Fhit-/- weanling tail tissue, demonstrating the existence of genome instability early in development.

Finally, Waters et al. (2015) identified a mechanism whereby Fhit loss-induced replication stress generates substrates for the APOBEC3B enzyme, resulting in significantly more hypermutations in cancers. The APOBEC family of enzymes defends against viral pathogens by deaminating cytosine bases in ssDNA intermediates of viral replication. Specifically, APOBEC enzymes cause C>T and CT motif hypermutation, suggesting that cooperating factors in addition to APOBEC activation are required for hypermutation (Burns et al., 2013;

Roberts et al., 2013). Following the observation that cancers deficient for Fhit, such as breast and lung, are also frequently associated with A3B overexpression, Waters et al.

(2014) hypothesized that ongoing replication stress-induced ssDNAs caused by Fhit loss could serve as A3B substrate. The rationale was that Fhit loss-induced replication stress would enhance A3B activity by increasing the ssDNA substrate available through helicase and polymerase uncoupling. DNAs of lung adenocarcinomas, available through

The Cancer Genome Atlas (TCGA), were stratified by expression of FHIT and A3B, confirming that Fhit low, A3B high tumors exhibited the highest frequency of mutations.

Fhit-deficient cancers showed an incremental increase in C>T mutations corresponding to increasing A3B expression, while Fhit-normal samples showed no correlation with mutation burden and A3B expression. As confirmation that ssDNAs were generated by replication stress induced by Fhit loss and dTTP pool deficiency, A3B-high/Fhit low cells were supplemented with thymidine to increase dTTP pools. Indeed thymidine supplementation decreased C

Altogether, these findings support a model for Fhit loss-associated neoplastic progression (Figure 3). Early in the transformation process, fragility at FRA3B results in deletions in FHIT alleles in normal or preneoplastic cells. Loss of Fhit protein expression causes TK1 down-regulation and nucleotide imbalance (dTTP pool insufficiency). This causes slowing of replication forks and under-replicated DNA regions as cells enter

G2/M. Chromatin condensation during mitosis leads to DNA deletions in under-

11 replicated regions. Continual cell cycling without checkpoint activation leads to accumulation of genomic changes. Loss of Fhit also causes activation of other fragile loci and provides the ssDNA template for A3B activity to induce a hypermutator phenotype.

FHIT loss-induced genome instability thus increases the likelihood of acquiring activating mutations in oncogenes and inactivating mutations in tumor suppressor genes, as selective pressures allow clonal expansion.

Figure 3. Model for Fhit loss-associated neoplastic progression. Preneoplastic colony expansion initiates with loss of Fhit expression in normal cells, which results in down- regulation of TK1 enzyme and subsequent depletion of TTP pools leading to fork stalling, collapse and DSBs in the preneoplastic stage. Loss of Fhit also causes a 2-fold increase in CFS activity. Subsequent increases in mutations and chromosomal instability (e.g. CNAs, p16 loss, Mdm2 activation) pushes cells into the neoplastic stage. Cooperative A3B activity may occur in the preneoplastic or neoplastic stage to induce a hypermutator phenotype.

1.6 Conclusions

Deletion within the FHIT gene at FRA3B is one of the most common deletion events in human cancers, and loss of Fhit protein is one of the earliest events in cancer initiation. Although there is much evidence validating Fhit as both a tumor suppressor and genome caretaker, the location of FHIT at a CFS, a locus prone to breakage and gap formation under even mild replication stress, has encouraged claims that Fhit loss is a passenger event in cancer development. The central hypothesis of this thesis is that Fhit loss is a driver, not a passenger, in neoplastic initiation and progression. Through an in vitro model that recapitulates the neoplastic process, this research shows that the chromosomal instability caused by loss of Fhit has functional consequences that promote cellular transformation. The results of this research also provide insight into the mechanism by which Fhit regulates TK1, and demonstrate how this Fhit-TK1 connection is related to cancer development.

Chapter 2: Fhit loss–associated initiation and progression of neoplasia in vitro

Submitted manuscript: Karras et al., Cancer Science. 2016.

2.1 Introduction

Chromosome fragile sites, conserved weak links in mammalian genomes, are among the most frequently deleted loci in cancer (Bignell et al., 2010). The fragile gene,

FHIT, was identified by the Huebner laboratory ~20 years ago at a locus that is inactivated in >50% of human cancers (Ohta et al., 1996; Pichiorri et al., 2008).

Deletions in the FHIT locus are among the first genetic changes detected in human preneoplastic lesions (Gorgoulis et al., 2005; Bartkova et al., 2005). Many biological functions have been reported to be altered by Fhit loss in cancers: decreased apoptosis

(Sard et al., 1999), increased epithelial-mesenchymal transition (EMT) (Joannes et al.,

2014; Suh et al., 2014), increased resistance to genotoxic agents (Ottey et al., 2004), altered control of Reactive Oxygen Species (ROS) production (Okumura et al., 2009), ongoing genome instability (Saldivar et al., 2013; Miuma et al., 2015); but the direct mechanisms through which Fhit protein affects these functions has remained elusive.

Lack of a known mechanism of action has slowed general acceptance of a role for Fhit in tumor suppression, despite strong evidence of Fhit association with multiple cancer- associated functions. This skepticism has hindered consideration of Fhit-associated

14 therapeutic targets for the majority of human cancers exhibiting Fhit loss. For example, the accumulation of genome mutations occurring as a direct result of Fhit silencing and associated thymidine kinase 1 (TK1) silencing at post-transcriptional levels, and the ability to stop the accumulation of genome damage in its tracks by thymidine supplementation hint at possible pre-neoplasia prevention strategies (Saldivar et al.,

2012). Also, cancer-associated genome instability is the target of treatment strategies based on genotoxic agents such as radiation and cytotoxic drugs, so stratification of cancers by expression level of Fhit and other „caretaker‟ gene products might improve the outcome of some treatment regimens. In addition, Fhit loss-induced DNA damage creates optimal single-stranded DNA substrates for the APOBEC3B enzyme (a cytidine deaminase that converts cytosines to uracils in ssDNA), illustrating a key role for Fhit loss in the recently described hypermutation genotypes observed in most common cancers, a major source of cancer-associated genetic heterogeneity (Waters et al., 2015;

Burns et al., 2015). The APOBEC3B enzyme, which causes hypermutations selectively in Fhit-deficient cells, is likely a critical diagnostic and therapeutic target. The purpose of the current study was to demonstrate that Fhit-deficiency influences neoplastic progression. We followed expression changes from establishment, through proliferation in the face of selective pressures, to transformation, morphological alterations, and nascent neoplastic changes, in epithelial cells from Fhit knockout and Fhit wild-type mice. We have observed that Fhit loss is followed by genomic and functional changes in response to selective pressures that allow survival of clonally expanded populations,

15 supporting the conclusion that Fhit-loss induced genome instability enables selection for transformation and facilitates neoplastic progression.

2.2 Materials and Methods

Ethics statement

Mice were maintained and animal experiments conducted in accord with institutional guidelines established by the Animal Care and Use Committee at Ohio State University.

Cell lines and reagents

Mouse kidney cell lines were established by culturing minced mouse kidney tissue from

3 Fhit+/+ C57Bl6 (B6 +/+ kd cell lines 1, 2, 3) and 3 Fhit-/- (B6x120SvJ backcross) 5- week-old mice (-/- kd cell lines 2, 3, 4). After emergence of epithelial cells from minced kidney fragments, cells gradually filled the culture vessel and could be split; these epithelial kidney cell lines did not exhibit the growth characteristics of mouse embryo fibroblast lines; e.g., they did not exhibit an obvious crisis phase but rather grew steadily from first subculturing after a long static phase. RNA, DNA and protein were isolated at alternate passages. To establish DMBA survivor (DS) cell lines, late-passage (p40) cells were treated with two sequential 24 hr, 20 µM DMBA doses, followed by plating and culturing of surviving colonies 8 days post treatment; +/+ cells did not survive DMBA treatment. To establish nutritionally stressed (NS) cell lines, early passage cells were maintained without replenishing medium for several months, followed by fresh medium and subculturing of surviving colonies; +/+ cell lines did not survive nutritional stress.

Additionally, some DS and NS cell lines formed colonies in soft agar. Colonies were 16 isolated and re-plated to establish colony-forming cell lines; see Table 1 for description of mouse cell lines. The mouse cell lines were cultured in MEM with 5% FBS and 100

µg/ml gentamicin. H1299, a human non-small cell lung carcinoma, was cultured in MEM with 10% FBS and 100 µg/ml gentamicin.

Immunoblots

Whole cell lysates were prepared in RIPA buffer (Thermo Scientific) supplemented with

Halt Protease cocktail Inhibitors (Thermo Scientific). Proteins were separated by SDS gel electrophoresis, transferred to nitrocellulose membranes and immunoblotted with antisera. The antisera used and the working dilutions are available in Table 1.

Table 1. Antisera used, dilutions and applications.

Sequencing of Trp53 and Ras genes in mouse kidney cell lines

Total RNA was extracted (RNeasy mini kit, Qiagen) and cDNA synthesis performed for

15 min at 42°C using 1 µg of total RNA as template, RT primer and QuantiTect Reverse

Transcriptase (Qiagen). PCR was performed using Trp53 and Nras gene specific

17 primers. Total genomic DNA was isolated using DNeasy Blood and Tissue (Qiagen).

PCR was performed to amplify exons 1, 2 and 3 of Kras and exons 1 and 2 of Hras.

Primers used are available in Table 2. Per reaction, 1 U of Platinum PCR SuperMix High

Fidelity Taq polymerase was used and cycling conditions were: pre-incubation for 5 min at 94°C, 30 cycles of 30 s at 94°C, 30 s at 54°C, 2 min at 68°C and final incubation at

4°C. PCR product quality was assessed on 1% agarose gel and purified for sequencing via NucleoSpin Gel and PCR clean-Up Kit (Macherey Nagel). Products for sequencing contained 6.4 pmol of primer and 40 ng of template DNA in a total of 12 µl with DNase free water; sequencing was done in the OSUCCC Nucleic Acid Shared Resource.

Trp53 plasmid construction

F131L and S151R Trp53 cDNAs were amplified from mouse NS1 and NS2 cell lines, respectively, using the following conditions: 94°C for 1 min, 30 cycles at 94°C for 30 sec, 54°C for 30 sec, 68°C for 2 min, and held at 4°C; Trp53 forward 5‟-GCG

AAGCTTATGACTGCCATGGAGGAGTCA -3‟, and reverse 5‟-

GCGTCTAGATCAGCCCTGAAGTCATAAGAC-3‟ primers were used. Mutant Trp53 18 cDNA was cloned into HindIII and XbaI sites of the pRcCMV vector (Invitrogen) under control of the immediate early human cytomegalovirus (CMV) promoter. Recombinant clones containing the mutant Trp53 gene were sequenced in the OSUCCC Nucleic Acid

Shared Resource.

Plasmid Transfections

The mammalian expression vectors, pRcCMV-empty vector, pRcCMV-wtTP53, pRcCMV-mouseTrp53/F131L, or pRcCMV-mouseTrp53/S151R, were transfected into mouse kidney cells using Lipofectamine 2000 reagent (Invitrogen). Mouse kidney cells were plated at a density of 1.5 x 105 cells per 60-mm dish and cultured in MEM without antibiotics 16-24 h before transfection. Transfections were with 1 µg plasmid DNA and 5

µl Lipofectamine diluted in Opti-MEM (Gibco) and incubated for 30 min at room temperature. Cells were overlaid with the plasmid DNA/Lipofectamine solution and incubated for 24 h at 37° C. Cell lysates were collected 24 h post transfection for western blot analysis.

Microarray expression profiles

Total RNA (+/+ kd2 p14, -/- kd3 p48 and NS3 colony p13) was isolated using RNeasy

Mini Kit (Qiagen). RNA integrity was interrogated using the Agilent 2100 Bioanalyzer

(Agilent Technologies, Palo Alto, CA). A 100 ng aliquot of total RNA was linearly amplified. Then, 5.5 µg of cDNA was labeled and fragmented using the GeneChip® WT

PLUS reagent kit (Affymetrix, Santa Clara, CA) following manufacturer‟s instructions.

Labeled cDNA targets were hybridized to Affymetrix GeneChip® Mouse Transcriptome

Array 1.0 for 16 h at 45°C, with rotation at 60 rpm. The arrays were washed and stained using the Fluidics Station 450 and scanned using the GeneChip Scanner 3000. For gene expression analysis, arrays were normalized using RMA algorithm in Expression Console and comparisons made in Transcriptome Analysis Console (Affymetrix, Santa Clara,

CA). The microarray data have been deposited in the NCBI Gene Expression Omnibus

(accession # not yet known).

Growth in soft agar

The soft agar culture system involved a base layer (MEM, 10% FBS, 100 µg/ml gentamicin, 0.5% low melting point agar) and a top growth layer (MEM, 10% FBS, 100

µg/ml gentamicin, 0.3% low melting agar). The top growth layer included 3 x 104 cells suspended in 12 ml per 100 mm plate. Normal medium was added on top of the growth layer on day 0 and changed every 5 days. Colonies were counted at 25 days.

Invasion assay

A commercially available kit containing polycarbonate membrane inserts coated with a basement membrane matrix solution was used (CytoSelect 24-well Cell Invasion Assay

Kit, Cell Biolabs, San Diego, CA). 1.25 x 105 mouse kidney cells in serum-free medium were transferred into the transwell insert and allowed to invade toward 10% FBS for 72 h at 37° in a humidified incubator. Non-invasive cells were removed from the top of the membrane via cotton-tipped swabs. Invading cells on the bottom surface were stained and quantified at OD 560 nm after extraction.

Tumorigenicity and Metastasis Assays

Male and female Foxn1nu/Foxn1nu nude mice (5 wks old) were purchased from The

Jackson Laboratory and athymic nude mice were obtained from an OSU core facility.

Athymic nude mice were bred within the Target Validation Shared Resource (TVSR) at the OSUCCC and maintained on an outbred background. The original breeders

(Strain#553 and #554) were obtained from the NCI Frederick facility. Mice were maintained under pathogen-free conditions. 1 x 107 (Fhit+/+ and NS3 colony) cells in 100

µl PBS were injected subcutaneously into the right flank of J:nu mice, 4 mice per cell line, in the first round of injections. Subsequent injections used 5 x 106 (Fhit+/+, NS1 colony, NS3 colony, NS3T) cells in a total of 20 athymic nude mice (6 females, 14 males). The animals were monitored twice weekly for tumor formation up to 6 months after inoculation. For assessment of ability to grow in metastatic sites, NS3T cells were injected into 7 athymic nude mice (2 females, 5 males). Briefly, 5 x 106 (or 2x106) cells resuspended in 150 µl PBS were tail-vein injected. The mice were monitored twice weekly up to 2 months after inoculation and sacrificed up to 8 wks post injections.

Inducible clones B28 and B29 were treated with doxycycline (1 µg/ml) 48 h before harvesting 5 x 106 cells in 150 µl PBS for subcutaneous injections into the right flank of

18 athymic male nude mice. Mice were fed either sucrose water (30%) or doxycycline water (1.2 mg/ml) twelve days prior to injections; water was replaced every 6 days.

Tumor volumes were calculated using the formula, volume = ½ (length x width2). Error bars depict standard deviation.

Histopathology and immunohistochemistry

Subcutaneous tumors were measured and fixed in 10% neutral buffered formalin; lungs were insufflated with 10% neutral buffered formalin prior to immersion fixation. Tissues were processed by routine methods and embedded in paraffin wax. Sections (4 m) were stained with hematoxylin and eosin (HE) or deparaffinized and hydrated for immunohistochemistry. Immunohistochemistry was performed as previously described

(Guler et al., 2012). Slides were evaluated with an Olympus BX45 light microscope with attached DP25 digital camera (B & B Microscopes Limited) by a comparative pathologist board certified by the American College of Veterinary Pathologists (ACVP). The antisera used and the working dilutions are available in Table 1.

Lentiviral vector construction

Wild-type human FHIT cDNA was amplified from previously constructed plasmids

(Siprashvili et al., 1997) using the following conditions: 95°C for 3 min, 30 cycles at

98°C for 10 sec, 55°C for 15 sec, 72°C for 5 sec, and held at 4°C; FHIT forward 5‟-

CCCTCGTAAAGAATTCATGTCGTTCAGAT-3‟, and reverse 5‟-

GAGGTGGTCTGGATCCTCACTGAAAGTA-3‟ primers were used. The cDNA was cloned into EcoRI and BamHI sites of the pLVX-TetOne-Puro vector (clontech). This vector allows transgene expression by the doxycycline-inducible TRE3G promoter.

Transgene expression upon doxycycline induction was assessed by immunoblot using

Fhit polyclonal antiserum.

Generation of inducible Fhit transfectants

The recombinant plasmid (pLVX-FHIT) was transfected into mouse kidney cells using

Xfect reaction buffer and polymer reagents (Clontech, Mountain View, CA). Mouse kidney cells were plated at a density of 4 x 105 cells per 60-mm dish and cultured in normal growth medium. Transfections were with 5 µg plasmid DNA diluted with 90 µl of Xfect buffer before addition of 1.5 µl Xfect polymer. The solution was incubated for

10 min at room temperature. Cells were overlaid with the plasmid DNA/polymer solution and incubated for 24 h at 37°C. After 24 h, stable clones were selected in puromycin (2 µg/ml) and tested for inducible Fhit expression after doxycycline treatment.

Statistical Analysis

Nonparametric data was analyzed using the Mann-Whitney rank sum test for single comparisons or using the Kruskal-Wallis test for multiple comparisons. Tumor incidence and latency was analyzed by T-test. Fisher‟s exact test was used to determine significant

IPA pathway associations. P-value <0.05 was considered statistically significant.

2.3 Results

2.3.1 In vitro model of Fhit loss – associated neoplastic progression

To create an in vitro model for Fhit-deficient cellular transformation, we established six mouse kidney epithelial cell lines. Three were derived from Fhit+/+

(+/+kd1, +/+kd2, +/+kd3) and three from Fhit knockout (-/-kd2, -/-kd3, -/-kd4), post- weaning mice. These cell lines were subcultured through tissue culture passage (p50) and

Derived Agar Cell Line From Colonies Fhit +/+ Subcultured +/+ kd1 ♀ ms 3451 No +/+ kd2 ♀ ms 3452 No +/+ kd3 ♀ ms 3453 No

Fhit -/- Subcultured -/- kd2 ♀ ms 3454 No -/- kd3 ♂ ms 3455 No -/- kd4 ♂ ms 3456 No Nutritionally Stressed NS1 -/- kd3 p3 Yes NS2 -/- kd2 p1 No NS3 -/- kd3 p5 Yes NS4 -/- kd3 p8 No

DMBA Survivors DS2 -/- kd2 p38 Yes DS3 -/- kd3 p40 Yes DS4 -/- kd4 p41 No Colony Forming NS1 colony NS1 p24 Yes NS3 colony NS3 p13 Yes DS2 colony DS2 p10 Yes DS3 colony DS3 p15 Yes

Table 3. Mouse kidney cells were established from culture of single kidneys from 3 Fhit+/+ C57Bl6 and 3 Fhit-/- (B6x129SvJ backcross) 5-week-old mice. Nutritionally stressed cell lines were isolated from the early passage -/-kd2 and -/-kd3 lines after maintenance without replenishing medium for 3 months, followed by fresh medium and continued subculture. DMBA survivor cell lines were established from late-passage cell lines after treatment with 20 µM DMBA. All cell lines were tested for anchorage- independent colony forming ability. Colony-forming cell lines were established after excision and plating of colonies that grew in soft agar. +/+ cells did not survive nutritional or carcinogen stress.

24 accumulating alterations examined. The initial cell cultures were also used in the generation of 7,12-Dimethylbenz[a]anthracene (DMBA) survivor (DS) and nutritional stress (NS) survivor cell lines. Fhit+/+ cell lines did not survive these additional stresses.

Thus, there are a total of 3 Fhit+/+ cell lines and 14 Fhit-/- cell lines, in which different selective pressures were applied. See Table 3 summary for description of the Fhit-/- cell lines.

2.3.2 Fhit-/- cells exhibit alterations in apoptotic and EMT signal pathways

To follow the evolution of cells from benign to malignant state in vitro, we assessed changes in proteins in signal pathways that are frequently altered in cancers.

We began by interrogating the Trp53/p21 and EMT pathways, key contributors to neoplastic development. In assessing the unstressed +/+ and -/- kidney cell lines, a reduction in Trp53 protein expression was observed in late-passage -/-kd3 along with a decrease in its downstream target p21 (Figure 4a). Trp53/p21 pathway changes were not observed in the DMBA survivor cell lines (Figure 4b). The most striking changes in

Trp53/p21 pathway proteins occurred in the nutritionally stressed lines; all 4 NS lines displayed Trp53 protein expression but lacked p21 expression, suggesting that these cell lines harbor mutated Trp53 genes, selected for as means of surviving the nutritional stress

(Figure 4c). Indeed, the absence of p21 expression is due to mutation in the DNA binding domain of the Trp53 protein. All NS lines exhibit C to G base substitutions, changing a phenylalanine to a leucine at amino acid position (aa) 131 (F131L) in NS1, NS3 and NS4 lines, and changing a serine to an arginine at aa151 (S151R) in NS2 (Figure 4d).

Figure 4. Fhit-/- mouse kidney cells exhibit signal pathway alterations. Immunoblots of p53, p21, Vimentin, Fhit and Vinculin expression in the mouse kidney cells with (a) progressive in vitro subculture, (b) survival of DMBA, and (c) survival of nutritional stress. (d) Chromatogram of Trp53 sequences in NS cell lines. Heterozygous C>G mutation at amino acid position 151 in NS2 cells, changing a serine to an arginine; homozygous C>G mutation at amino acid position 131 for NS1, NS3 and NS4 cells, changing a phenylalanine to a leucine.

Mutations in the Trp53 DNA binding domain can result in faulty transactivation of the

CDKN1A gene encoding p21 protein. To confirm that p21 protein is down-regulated due to Trp53 mutation, we transfected NS lines with wild-type and mutant Trp53 plasmids to determine if p21 expression could be restored. Re-expression of p21 was observed in both NS1 and NS2 cells that were transfected with wild-type Trp53, but not when transfected with F131L or S151R Trp53 mutants (Figure 5a). Although DS lines did not exhibit changes in the Trp53/p21 pathway, increased expression of the pro-survival protein survivin was observed (Figure 5b). Malignant transformation is often associated with induction of EMT signaling pathways. Therefore, to discern if Fhit-/- cells have acquired pro-tumorigenic activities, we tracked the expression of Vimentin, a marker of the mesenchymal phenotype and a hallmark of EMT. All 4 NS lines exhibited robust expression of Vimentin (Figure 4c), suggesting these cells have undergone EMT and possess migratory abilities. Tables 4 and 5 display expression patterns of other proteins tested.

Figure 5. Wild-type p53 overexpression restores p21 protein expression. (a) NS1 and NS2 mouse kidney cells were transfected with either control vector (EV), wild-type or mutant Tp53 (F131L or S151R) expression vector. p21 protein expression is restored 24 h post wild-type p53 transfection. (b) Immunoblot of Survivin expression in DS cells. 27

Table 4. Protein expression changes in Fhit-/- and +/+ kidney cell lines. Expression analysis of proteins at early (p3-16) and late (p40-50) tissue culture passages. +++, very strongly expressed; ++, strongly expressed; +, moderately expressed; +/-, faint expression; -, absent.

Table 5. Protein expression in Fhit-/- DS, NS and colony-forming cell lines. +++, very strongly expressed; ++, strongly expressed; +, moderately expressed; +/-, faint expression; -, absent.

2.3.3 Fhit loss-associated cell transformation

Because protein expression studies provided evidence of in vitro transformation in

Fhit-/- cells, we compared biological features of +/+ and -/- kd cell lines, by measuring the effect of Fhit deficiency on anchorage-independent growth in soft agar. H1299, a human metastatic lung carcinoma cell line, served as positive control. After 24 days of culture in soft agar, some cells of nutritionally stressed cell lines NS1 and NS3 showed anchorage-independent growth, producing 15 and 13 colonies respectively (Figure 6a).

Additionally, some cells of DMBA survivor cell lines DS2 and DS3 formed large colonies, although fewer than the NS cell lines (Figure 6a). Representative photos of agar colonies at day 24 are shown in Figure 6b. The soft agar assay for H1299 produced >750 colonies, whereas no colony formation was observed for any of the +/+ and unstressed -/- kd cell lines. Moreover, after collection of these agar colonies and propagation, we re- assessed one colony line from each group for colony formation potential. Each new colony cell line showed rapid anchorage-independent growth and formed increased numbers of agar colonies (Figure 6c). Thus, the cell lines subjected to exogenous stress demonstrated anchorage-independent growth and colony formation, properties of transformed cells. Western blot analysis showed that Trp53 overexpression and loss of p21 had also occurred in NS1 and NS3 colony cell lines, demonstrating that the Trp53 missense mutation acquired in NS1 and NS3 was maintained in these colony-forming lines (Figure 6d); the DS2 colony line expressed normal Trp53/p21 pathway expression, while the DS3 colony line had lost Trp53 protein expression, resulting in down- regulation of p21 expression, as observed in the -/- kd3 parent cell line. Furthermore, the

NS3 colony line displayed a dramatic increase in Vimentin expression. The NS1 colony line exhibited a lower level of Vimentin expression; DS2 and DS3 colonies did not express Vimentin (Figure 6d). An invasion assay through a basement matrix coated membrane was performed to determine if colony forming cell lines also showed invasive capacity. Results indicate that the NS3 colony line had significant invasive ability vs +/+ controls (P=0.01) (Figure 6e). The NS1 colony line showed increased invasive potential vs a +/+ control cell line, in accord with the low level increase in Vimentin expression observed (Figure 6d,e). The results suggested that the Fhit loss-associated signal pathway alterations identified are in vitro transformation-associated changes.

Figure 6. Transformation-associated features of Fhit-/- cells. (a) Anchorage- independent colony formation was observed in four exogenously stressed Fhit-/- cell lines at day 24 in soft agar. Fhit+/+ kd1, +/+ kd2, +/+ kd3 and Fhit-/- kd2, -/- kd3 and -/- kd4 cell lines were plated for colony formation at early (p4-7) and late (p46-49; p25 for +/+) tissue culture passage, for three independent experiments, but no colony formation was observed. (b) Images of colonies; some colonies were excised from agar and propagated in liquid culture. (c) Anchorage-independent growth of newly propagated colony-forming cell lines. (d) Western blot analysis of colony-forming cell lines. (e) Invasion assay in triplicate was performed to assess the invasion activity of colony- forming cell lines. 31

2.3.4 Classification of genes with altered transcription in a Fhit-/- NS cell line

To further characterize signal pathway alterations that contribute to Fhit loss- directed cellular transformation, we examined signal pathways identified by mRNA expression profiling. Ingenuity Pathway Analysis (IPA) was used to analyze the differentially expressed genes in the in vitro invasive NS3 colony cell line, relative to its non-invasive progenitor -/- kd3. Using a significance cut off of P value <0.05 and a fold- change cut off of 4, there were 432 differentially expressed genes in NS3 colony cells vs

-/- kd3. An IPA „core analysis‟ was performed to classify this dataset into top biological functions and canonical pathways (Figure 7a,b), several of which revolve around DNA replication, cell cycle control and DNA repair. NS3 colony cells also expressed a subset of genes classified into the “regulation of epithelial-to-mesenchymal transition pathway” demonstrating expression changes in EMT transcription factors and markers. An invasion-associated network was constructed to focus on specific genes influencing the invasive phenotype of this cell line (Figure 7c). A classic attribute of a cell undergoing

EMT and gaining invasive properties is the repression of epithelial markers concomitantly with the activation of mesenchymal gene expression. Relative to the -/- kd3 parent, the NS3 colony cell line displayed a dramatic 35-fold down-regulation of E- cadherin (Cdh1), an epithelial marker, and 55-fold up-regulation of Vimentin (Vim), a mesenchymal marker. Up-regulation of transcription factors known to induce EMT, such as Zeb1, Snai2 and Foxm1 is also observed in NS3 colony cells (Da Craene et al., 2013;

Alves et al., 2009; Spaderna et al., 2008; Bao et al., 2011). Furthermore, genes involved in regulating cell-to-cell contacts and cell junction integrity are differentially expressed in

Figure 7. Classification of genes with altered transcription in NS3 colony cell line relative to Fhit-/-kd3 p48. IPA core analysis was used to classify gene expression data of NS3 colony into (a) biological functions and (b) canonical pathways. IPA gene networks associated with (c) invasion and (d) DNA damage. Up-/down-regulated genes within NS3 colony are denoted by red and green color, respectively. Fold change numbers are represented under the nodes.

favor of facilitating the EMT process, suggesting that NS3 colony cells are gaining invasive properties. Additionally, a network analysis of DNA damage response- associated genes (Figure 7d) identified genes important for replication fork progression such as Top2a, Mcm10, Lig1 and Rrm2 that are up-regulated, possibly participating in maintaining increased proliferative signaling. Chek1, a gene responsible for coordinating 33 the DNA damage response, and DNA double-strand break repair proteins Brca1 and

Rad51 are also up-regulated, suggesting enhanced DNA damage repair in NS3 colony cells. The replication licensing factor and proto-oncogene Cdc6 is also up-regulated, which could lead to hyper-replication and increased chromosomal instability, especially in collaboration with mutant Trp53 (Karakaidos et al., 2004). No expression changes were observed for Myc, Raf, Mek, Erk, Erbb2, Egfr or Ras. Sequence analysis of Kras,

Hras and Nras cDNAs amplified from NS cell RNA detected only wild-type sequence at hotspot regions in all NS lines. In the “cyclins/cell cycle regulation” canonical pathway, cyclins Ccne1, Ccne2 and Ccnb1 are up-regulated 2.78, 5.29 and 10.4 fold, respectively, in NS3 colony cells (Figure 7b), demonstrating that Fhit loss precedes the activation of observed oncogenes in this model system. Collectively, these results suggest that the NS3 colony cells acquired signal pathway alterations to meet the challenges of anchorage- independent growth, changes that are known cancer hallmarks, in accord with the hypothesis that this cell line has undergone malignant transformation.

2.3.5 NS3T cells display tumorigenic and metastatic potential

To assess in vivo behavior, we injected the in vitro–invasive colony cell lines subcutaneously (sc) into 5 to 6-wk old nude mice and observed animals weekly for appearance of tumors. NS1 colony, which exhibited modest invasive ability in vitro, formed tumors in two of two male mice, which grew to 6x7 mm and 5x5.5 mm by days

105 and 125, respectively. NS3 colony was injected into 2 male and 2 female mice. By day 133, one male mouse developed tumors at sites on the shoulder and flank that each

34 received 5 x 106 cells. Both sites developed sizable tumors by day 151 (flank tumor

15x12 mm; shoulder nodules, 5x5 mm and 3x5 mm) that showed a mesenchymal spindle cell neoplasm phenotype (Figure 8).

Figure 8. Mesenchymal spindle cell neoplasm phenotype of NS3 colony tumors. NS3 colony cells were injected sc into the flank and shoulder of one male nude mouse. Both sites developed tumors that display a histological phenotype consistent with a mesenchymal spindle cell neoplasm. All masses are unencapsulated and composed of mesenchymal spindle cells on a fine fibrovascular, with distinct collagen fibrils between individual neoplastic cells. Multinucleated cells are also prominent. Scale bar, 50 µm.

Of 4 mice injected with +/+ kd3 p15, none developed tumors by day 200. NS3 tumors were excised for histopathology, the NS3 flank tumor was cultured in vitro and the tumor outgrowth cell line was designated NS3T. A second round of sc injections was performed to determine whether the cultured NS3T cells showed increased tumorigenicity. 5x106 NS3T cells at p10 and +/+ kd3 p16 cells were injected into the flanks of 4 nude mice each, (2 females, 2 males). Both males injected with NS3T cells formed tumors within 12 days, with mean tumor size ~100.5 mm3 by 19 days (Figure 9a).

The female mice and the +/+ kd3 injected mice did not develop tumors and were sacrificed at day 60. The results suggested that tumor formation was biased towards male

Figure 9. NS3T cells display high tumorigenic and metastatic potential. Tumor growth curves of male athymic nude mice that received a single subcutaneous injection of (a) 5x106 NS3T p10 or +/+ kd3 p16 cells and (b) 5x106 NS3T p7 cells to the right flank. (c) H&E staining and expression of Cytokeratin, E-cadherin and Vimentin by immunohistochemical analysis in subcutaneous tumors and lung metastases. Circle, lung metastases; B, bronchiolar epithelium. Scale bar, 50µm except for H&E lung metastasis where scale bar, 1mm. (d) Tumor growth curve and (e) tumor latency of athymic nude mice that received subcutaneous injection of Fhit inducible NS3T clone B28 or clone B29 cells with and without doxycycline induction. Error bars depict standard deviation. (f) Western blot analysis of Fhit expression levels after 48 h doxycycline (1 µg/ml) treatment in NS3T clone B28 and clone B29 cells. 36 mice, possibly because the androgen-receptor is expressed in the NS3 cells as noted in the expression array profile, and the initial cell line was derived from a male mouse kidney; sc injections using later passage NS3 colony cells resulted in 100% tumor incidence in female mice. A final round of 5x106 NS3T cells was injected sc into 5 male nude mice and 4 of 5 developed tumors by 10 days (Figure 9b). See Table 6 summary of tumor incidence. Two of these subcutaneous tumors were further characterized for an

EMT phenotype by assessment of expression of Vimentin, E-cadherin and Cytokeratin via immunohistochemistry. In accord with our transcriptome and western blot analysis, these tumors were strongly immunoreactive for Vimentin and immunonegative for

Cytokeratin and E-cadherin (Figure 9c). Additionally, the metastatic capacity of the

NS3T cells was evaluated by tail vein injection in male and female nude mice.

Histological examination showed lung micro-metastases in 3 of 5 male and 2 of 2 female mice. Lung tumors were more abundant (up to five nodules/lung) and larger in male mice sacrificed 43 days post-injection, whereas both female mice displayed a small, single neoplastic nodule within one lung lobe at sacrifice 58 days post-injection. Neoplastic cells were distributed within the alveolar parenchyma, around blood vessels and bronchioles, sub-pleurally or intravascularly. Lung tumors in one male were characterized by immunohistochemistry as performed for primary sc tumors. Neoplastic cells in the lung were strongly immunoreactive for Vimentin and immunonegative for

Cytokeratin and E-cadherin, in contrast to normal bronchiolar epithelium which served as internal control (Figure 9c). Thus, the alterations that occurred in vitro contributed to in vivo tumorigenicity and metastasis.

Cell Line Sex Route Days post- Tumor Comments injection frequency NS1 colony 2♂ subcu 125 2/2 NS3 colony 2♂ subcu 200 1/2 2♀ subcu 200 0/2 Early passage (p14) 2♀ subcu 200 2/2 Late passage (p27) NS3T 7♂ subcu 60 6/7 2♀ subcu 60 0/2 NS3T 5♂ iv 43 3/5 2♀ iv 58 2/2 +/+ kd3 3♂ subcu 125 0/3 Control for NS1 colony inject. +/+ kd3 2♂ subcu 200 0/2 Control for NS3 colony inject. 2♀ subcu 200 0/2 Control for NS3 colony inject. +/+ kd3 2♂ subcu 60 0/2 Control for NS3T injections 2♀ subcu 60 0/2 Control for NS3T injections

Table 6. Summary of tumorigenicity analyses. Initial studies showed that NS3 colony cells formed tumors only in male mice, likely related to expression of the androgen receptor (AR). Thus, NS3 colony tumor incidence was 50% in male mice. Late-passage NS3 colony tumor incidence of 100% in female mice suggests loss of AR expression. The NS3T tumor incidence was 85%, and tumor incidence is significant with P-value of p = 0.0109.

2.3.6 Induced Fhit expression delays tumor onset in vivo

To confirm that Fhit loss is responsible for tumor initiation in vitro, we created two stable NS3T clones that are doxycycline inducible for Fhit expression, clones B28 and B29. For both clones, we observed no differences in soft agar colony growth or in vitro invasive potential between Fhit-deficient and Fhit-induced cells. No difference in final tumor volumes was observed between Fhit-deficient and Fhit-induced cells for both clones (Figure 9d). However, clone B28 exhibited a significantly (P=0.0001) delayed onset of tumor formation in mice induced for Fhit expression via doxycycline water. All

B28 control mice developed tumors by day 13, whereas tumors did not start to appear

38 until day 16 in the Fhit-induced mice (Figure 9e). Surprisingly, clone B29 did not display the same effect on tumor latency upon Fhit induction (Figure 9e). Western blot analysis revealed that 48 h doxycycline treatment causes ~10-fold lower level induction of Fhit protein compared to clone B28 (Figure 9f), suggesting that robust Fhit expression is necessary for delaying tumor onset. Since there is no selection for retention of the inducible plasmid in the in vivo environment, loss of Fhit plasmid and thus Fhit expression likely explains eventual tumor development by both clones.

2.4 Discussion

The translational research world places substantial focus on the late stages of cancer and identification of specific cancer „driver‟ genes. But largely because of the extensive genome instability and plasticity of neoplastic cells underlying the extreme clonal heterogeneity of metastatic cancer, treatment frequently fails due to relapse and therapy resistance. Thus, the idea of concentrating on the biology of premalignancy to advance prevention and early diagnosis is gaining interest. Kensler et al. (2016) have proposed a Pre-Cancer Genome Atlas initiative for solid tumors of epithelial origin to investigate the molecular alterations associated with premalignant lesions as they progress toward cancer. Alterations of the FHIT gene, straddling a common fragile site in all human genomes, occur in the preneoplastic lesions preceding development of many human cancers.

While point mutations and small insertions/deletions have long been a focus in tumor sequencing, recent studies have suggested that genome structural variants, such as 39 deletions and translocations, the earliest genetic clues observed in hematopoietic cancers, may play a larger role in cancer progression than previously thought. In 2015, a group examined the contribution of recurrent structural variations in the progression of pancreatic cancer, a cancer with poor prognosis and low survival rates. Analysis of 24 ductal pancreatic adenocarcinomas (PDACs) revealed that the FHIT gene is the second most frequently altered gene, with deletions observed in 50% of PDAC tumors that resulted in reduction of Fhit protein expression (Murphy et al., 2016). Another study performed whole genome sequencing to fully characterize the genomic landscape of gastric and esophageal tumors. Again, structural alterations were detected. Compared to matched normal blood samples, recurrent deletions at the FHIT locus were identified in

46% of tumors (Hu et al., 2016). These and earlier sequencing analyses show that cancer mutation profiles include recurrent chromosomal alterations. Our lab has shown that such alterations at the FHIT locus leads to loss of Fhit protein expression that causes mild replication stress via TK1 down-regulation and subsequent dNTP imbalance (Saldivar et al., 2012).

The current study followed in vitro cellular alterations associated with Fhit absence to illustrate that loss of Fhit genome caretaker function supports in vitro tumorigenic progression. We demonstrate that Fhit loss provides a survival and expansion advantage when selective pressures are applied, which enables selection for preneoplastic properties. As proof that Fhit loss provides a survival advantage, all three

Fhit+/+ cell lines did not survive nutritional stress or carcinogen treatment, while all Fhit

-/- cell lines survived. This demonstrates that the genome instability initiated by Fhit

40 protein absence allows a fraction of Fhit-/- cells to survive these stresses, even at early tissue culture passages. Furthermore, in contrast to Fhit+/+ cells, we demonstrate that loss of Fhit, combined with stressful exposures, leads to alterations in apoptotic and EMT signaling pathways and oncogene activation. These alterations allow for transformation, selective clonal expansion and development of invasive properties in vitro as well as tumor formation and metastasis in vivo. Finally, induction of exogenous wild-type Fhit protein delays the onset of tumor formation in vivo. The documentation of frequent FHIT allele losses in precancerous lesions, in combination with our demonstration that loss of

Fhit expression supports preneoplastic and neoplastic clonal expansion, reveals Fhit loss as a driver of neoplastic progression. Therefore, we propose preclinical research to seek in vivo methods to prevent replication stress at FHIT and other fragile loci, or to prevent the consequences of Fhit loss. We have shown that in vitro thymidine supplementation of

Fhit-/- cells prevents ongoing genome instability, implying that inactivation of the Fhit-

TK1 pathway underlies the generation of genome instability that allows clonal expansion

(Saldivar et al., 2012). Although many studies have shown that dNTP pool imbalances cause replication stress and mutations leading to tumorigenesis, it is not known whether in vivo thymidine supplementation would be a feasible prevention approach (Bester et al.,

2011; Chabosseau et al., 2011; Kumar et al., 2011). Testing of thymidine supplementation to support genome stability may be an important avenue for intensive preclinical research and may inspire additional approaches to inhibit genome instability.

In summary, this study demonstrates that Fhit-deficient cells are more likely to acquire cancer-promoting mutations. Through selective pressures to survive, activating

41 mutations in oncogenes, or inactivating mutations in tumor suppressor genes expedites the cellular transformation process. We conclude that in preneoplastic lesions of human tissues, losing Fhit provides a selective advantage for transformation and cancer progression. The significance of Fhit loss as an alteration that lies at the core of cancer initiation and progression should be exploited as a prevention or therapeutic target due to its relevance in more than 50% of human cancers.

Chapter 3: The Fhit-TK1 signal pathway

Includes portions of manuscripts for which I was coauthor, participating in design and

performance of experiments, writing and editing of manuscripts:

Waters et al., Mol Biol of the Cell. (In revision)

Paisie et al., Cancer Sci. 2016. 107(4):528-35.

3.1 Introduction

Loss of expression of the tumor suppressor, Fhit, occurs early in tumor development of many cancers of many types. This loss leads to initiation and accumulation of DNA damage and alters a number of biological processes associated with cancer development. However, how Fhit loss affects these functions, and whether directly or indirectly, has been difficult to assess, partially because of its cytoplasmic location and very few interacting proteins. The Huebner laboratory recently reported that

Fhit loss is associated with severely reduced TK1 protein expression, a key DNA synthesis enzyme (Saldivar et al., 2012; Karras et al., 2014). We postulate that understanding how loss of Fhit causes this reduction in TK1 expression would provide a strong clue to the mechanism of its tumor suppressor and genome caretaker functions, by leading to identification of other genes regulated by Fhit through a similar mechanism.

TK1 functions in the scavenger pathway of dTTP biosynthesis where it catalyzes the ATP-dependent phosphorylation of thymidine to produce thymidine monophosphate.

Thymidine monophosphate is rapidly catalyzed by additional kinases to produce thymidine triphosphate (TTP) (Hu and Chang, 2007). Expression of TK1 is cell-cycle specific, fluctuating with the growth of the cell. TK1 levels are low in G1, increasing drastically during S phase to ensure sufficient production of dTTP for efficient DNA synthesis, and then decreasing in the G2/M phase (Sherley and Kelly, 1988; Munch-

Petersen et al., 1995). Complex transcriptional, post-transcriptional, and translational regulation of TK1 occurs for precise fine-tuning of dTTP pools in cycling cells (Stewart et al., 1987). Therefore, the positive regulation of TK1 by Fhit may occur at any of these levels. Because Fhit is a cytoplasmic protein, we hypothesize that Fhit regulates TK1 at the post-transcriptional level, downstream of mRNA export from the nucleus. Defining the mechanism of Fhit action may lead to discovery of novel targets for therapeutic intervention.

3.2 Materials and methods

Cell lines and reagents

Mouse kidney cell lines used were described in Chapter 2. FHIT-deficient H1299 lung carcinoma cells were previously transfected with inducible FHIT cDNA and tightly regulated inducible clones were isolated (FHIT wt) and empty vector (EV). H1299 cells were maintained in MEM with 10% FBS, zeocin, gentamicin and geneticin. FHIT

44 expression was induced by addition of ponasterone A (2 μM) (Life Technologies) to the growth medium.

Western blot analysis

Cells were lysed with RIPA buffer (Thermo Scientific) supplemented with Halt Protease

Cocktail Inhibitors (Thermo Scientific). Proteins were separated by SDS gel electrophoresis, transferred to nylon membranes, and immunoblotted with antisera.

Antisera used and working dilutions are listed in Table 1 of Chapter 2.

Comet Assay

Neutral comet assays were performed using the CometAssay kit (Trevigen) and recommended protocol. Images were acquired with a Zeiss Axioscop 40 fluorescent microscope mounted with an AxioCam HRc camera, and using an A-Plan 10x/0.25 objective lens. Images were converted to Bitmap files using Axiovision 3.1 software, and comet tail moments were scored using Comet Score 1.5 (TriTek, autocomet.com).

Immunofluorescence (IF) staining

Cells were grown on 8-chamber slides (Lab-Tek II) and fixed in freshly prepared 4% paraformaldehyde (PFA). Samples were rinsed with PBS, permeabilized in 0.1%

TritonX-100 and blocked in 1% BSA. Cells were incubated with primary antisera, mouse anti-γH2AX, at 1:333 dilution (Millipore) overnight at 4°C. Slides were washed

3x10 min in PBS, and secondary antiserum (AlexaFluor 488 – conjugated donkey anti- mouse IgG, 1:280, Invitrogen Molecular Probes) was added and incubated for 1 h at room temperature. Slides were washed and coverslips mounted using Fluoro-Gel II – 45 with Dapi (Electron Microscope Sciences). Images were acquired at room temperature with an Olympus FV1000 spectral confocal microscope, a UPLFLN 40XO objective lens, NA 1.30, and with Olympus FLOWVIEW acquisition software.

Exome sequencing

Sample preparation and exome sequencing of genomic DNAs from -/- mouse kidney and lung tissue and -/- NS1 and -/- NS4 mouse kidney cell lines were carried out by Genome

Quebec. Samples were prepared using (Agilent, Santa Clara, CA, USA) SureSelect

Mouse All Exon Kit; sequencing was done using (Illumina, San Diego, CA, USA) HiSeq

2000. Reads mapping and variants calling were carried out by Genome Quebec using the current MUGQIC DNA-Seq analysis protocol; BCFtools called the variants using B6 as the reference genome.(15) The average read depth was 89.79 98.71 for kidney and lung tissue DNAs and 78.6–81.37 for kidney cell line DNAs; the number of bases read were

~3.5 times more than the number of bases read for samples sequenced by EdgeBio.

Exome data was filtered to remove single base substitutions (SBSs) with low mapping quality (<10), variants calling score (<10), and reads depth (<10). Exome sequencing of genomic DNAs from mouse liver tissues and quality filtering was as described previously

(Miuma et al., 2013).

Validation of SBSs

Genomic DNA used in PCR amplification reactions to validate SBSs was extracted as described previously (Miuma et al., 2013). Primers were designed to amplify selected loci with predicted individual or multiple SBSs; PCR amplification and sequencing was

46 carried out using gene-specific primers and standard PCR and sequencing conditions. See

Table 7 for a list of primer pairs.

Table 7. Gene-specific primer pairs for validated SBSs.

Reverse Transcription qPCR

Total RNA from the culture was isolated using RNeasy Mini Kit (Qiagen) according to the manufacturer‟s protocol while total RNA from the polysomal fractions were isolated using TRIzol reagent (Invitrogen). cDNA was synthesized from 1µg of total RNA following the instructions of the QuantiTect Reverse Transcription Kit (Qiagen). qPCR was performed using SYBR Green (BioRad) on the IQ5 iCycler (BioRad); each sample was analyzed in triplicate and normalized to GAPDH while the RNA of polysomal fractions normalized to Luciferase. Gene specific primers for TK1 and GAPDH were purchased from Santa Cruz Biotechnology and Luciferase primers were: forward primer,

5'-CTTATGCATGCGGCCGCATCTAGAGG-3'and reverse, 5'-

CAGTTGCTCTCCAGCGGTTCCATCC-3'. The comparative ΔΔCt method was used to calculate the relative mRNA expression levels.

Luciferase reporter construction

Full-length TK1 promoter (NM_003258.4, 437 bp) was amplified using primers that included recognition sequences for SpeI and XhoI restriction endonucleases (Arcot et al.,

1989). TK1 promoter amplicons were then sticky-end cloned into pMCS-Red firefly luciferase plasmid (ThermoFisher Scientific) following restriction digest with SpeI and

XhoI restriction enzymes. Lipofectamine 2000 was used to transiently co-transfect the firefly Luciferase construct (and pTK-Green Renilla luciferase reporter to normalize for transfection efficiency) into H1299 EV cells and induced H1299/FHIT wt cells that had been treated with ponasterone A for 24 hrs prior to transfection.

TK1 promoter luciferase activity in Fhit expressing and non-expressing cells

The TK1 promoter luciferase expression vector was co-transfected with pRL-TK (a

Renilla luciferase reporter to normalize for transfection efficiency) into wt Fhit- expressing H1299 cells and H1299 EV containing cells using Lipofectamine 2000 (Life

Technologies) according to the manufacturer‟s instructions. Three independent transfections were assayed in triplicate for all luciferase experiments. Luciferase output from both Firefly and Renilla luciferase were measured step-wise by luminometry on the

Veritas microplate luminator 48 hrs post-transfection using the Dual-Glo luciferase assay system (Promega). Luminescence values for Firefly Luciferase were normalized to

Renilla values to control for transfection efficiency. Relative firefly luciferase light units

(RLU) in Fhit-deficient or Fhit-expressing H1299 cells were plotted relative to Fhit- deficient cells.

Analysis of TK1 mRNA decay

H1299 EV and FHIT wt stably transfected cell clones were incubated with ponasterone A for 15 hrs followed by 5,6-dichloro-1-β-D-ribofuranosylbenzimidazole (DRB) (Sigma-

Aldrich) addition (50 μM). RNA was extracted 0, 2, 4, 6, 8, 12 and 24 hrs after FHIT expression induction and analyzed by RT-qPCR.

Polysome gradients

H1299 EV and wt cell clones were seeded and treated with ponasterone A overnight.

Cells were treated with cycloheximide (CHX, 100 µg/ml) for 10min at 37°C and then cells were collected. Cells were then washed twice with PBS+CHX (100 µg/ml), scraped from the dish with a cell scraper and pelleted by centrifugation at 1000 xg for 5 minutes.

All subsequent procedures were carried out on ice. Collected cell pellets were lysed in 5 pellet volumes of ice cold lysis buffer (50 mM Tris pH 7.5, 10 mM KCl, 10 mM MgCl2,

150 mM NaCl, 0.2 % NP-40, 2 mM DTT, 0.5 mM PMSF, 1 mM sodium orthovanadate,

200 µg/ml CHX, supplemented with 5 µl/ml RNAseOUT (Life Technologies), 25 µl/ml protease inhibitor cocktail (Sigma), 10 µl/ml each phosphatase inhibitor cocktails 2 and 3

(Sigma)). Lysates were incubated on ice for 10 min with mild agitation every two minutes. Nuclei and other debris were pelleted with a 10 minute centrifugation at 16,000 xg. Supernatants were loaded onto 10-50% linear sucrose gradients and centrifuged for 3 hr at 35,000 rpm in a Sorvall TH641 rotor at 4ºC. Gradients were collected into 0.5 ml fractions with continuous monitoring of absorbance at 254 nm. Alternating fractions were

49 spiked with a luciferase control RNA (Promega) and RNA was recovered using TRIzol according to the manufacturer‟s instructions.

Statistical analysis

Two-sided T-tests were used to determine significance for data with a normal distribution and equal variances. Nonparametric data was analyzed using the Mann-Whitney rank sum test for single comparisons or using the Kruskal-Wallis test for multiple comparisons. Groups with P-values less than a value of 0.05 were considered significantly different.

3.3 Results

3.3.1 Thymidine supplementation prevents ongoing DNA damage in Fhit-/- cells

Fhit-deficient mouse kidney epithelial cells display significantly decreased levels of TK1 protein compared to Fhit+/+ (Figure 10a), and in H1299 cells, a Fhit-/- lung carcinoma cell line, induction of exogenous wt Fhit protein restores TK1 expression levels (Figure 10b). This is in agreement with previous studies in which TK1 expression was reduced upon Fhit silencing in human embryonic kidney 293 and HCT116 colorectal carcinoma cell lines (Saldivar et al., 2013). To confirm the role of the Fhit-TK1 pathway in promoting genome instability, we asked if Fhit-deficient cells exhibit decreased levels of DNA damage upon addition of a continuous supply of thymidine, the substrate for

TK1, despite the reduced TK1 protein expression of -/- cells. We first assessed spontaneous levels of damage by quantifying nuclear γH2AX foci, markers of DNA double-strand breaks (DSBs), by indirect IF in early passage +/+ and -/- kidney cell lines 50

(Figure 10c). The three early -/- kd cell lines exhibited ~2-fold increases in cells with

γH2AX foci vs +/+ cells (Figure 10d). Levels of DNA damage prior to thymidine supplementation were also measured in these cells by neutral comet assay, a method that relies on the principle that fragmented DNA from DSBs migrates faster than intact DNA through a gel when subjected to an electrical current. Subsequent staining with a DNA intercalating agent is used to visualize the nucleoid, seen as a comet head representing undamaged DNA, and the budding “comet tail” that represents the fragmented DNA that migrated out of the nucleoid. DSB levels of individual cells were quantified by measuring tail moment, a measure that combines the amount of DNA within the tail as well as the distance of migration. We observed a significant increase in the mean tail moment of

Fhit-/- vs +/+ cells, indicating that -/- cells exhibit elevated levels of damage (Figure

10e). These results are in accord with previous studies demonstrating that loss of Fhit expression causes spontaneous DSBs (Saldivar et al., 2013). Thymidine supplementation

(10 µM) for 40 days suppressed DSB formation in -/- cells only (Figure 10f), with supplemented cells showing mean tail moments similar to +/+ cells, demonstrating that

-/- cells do not accumulate further damage upon addition of thymidine. Thus, the modest level of TK1 that is present in these cells is capable of converting thymidine into dTTPs at a sufficient level to support normal DNA synthesis.

Figure 10. Thymidine supplementation prevents ongoing DNA damage. Immunoblot of Vinculin, TK1 and Fhit in (a) Fhit-/- and Fhit+/+ mouse kidney epithelial cell lines and (b) H1299 cells containing EV or induced wt FHIT plasmid. Cells were harvested 48 hrs after treatment with ponasterone A. (c) Detection of γH2AX foci in a representative Fhit +/+ and -/- kidney cell line at passage 9 by indirect immunofluorescence, before thymidine supplementation. (d-f) Quantifications of pooled Fhit +/+ (+/+ kidney cell lines 1, 2, 3) and Fhit -/- (-/- kidney cell lines 2, 3, 4) cell lines. (d) Quantification of γH2AX-positive cells before thymidine supplementation; cells containing 3 γH2AX foci were considered positive. Bar graph indicates the means, and error bars represent the standard error. (e) Quantification of neutral comet assay results in mouse kidney cells before thymidine supplementation. Box plots of tail moments include data (+/+, n = 285; -/-, n = 435) from 3 separate experiments. (f) Quantification of neutral comet assay results for mouse kidney cells 40 days post 10 µM thymidine supplementation. Box plots of tail moments include data (+/+ untreated, n = 344; +/+ with thymidine, n = 228; -/- untreated, n = 341; -/- with thymidine supplementation, n = 286) from 3 separate experiments. For all boxplots, bottom and top of the box correspond to the 25th and 75th percentiles, respectively, and whiskers represent data points within 1.5xIQR (interquartile range). The black line extending through the boxplot indicates the mean value, and the black line contained within the boxplot represents the median value.

3.3.2 Mutation burden is increased in Fhit-/- kidney cells and tissue

Whole exome sequencing (WES) was used to examine the effects of the aforementioned instability on mutation frequency. Single base substitutions (SBSs) were evaluated in cell lines and tissues derived from FHIT knockout (ko) vs wild-type (wt) mouse strains. Fhit-deficient liver tissues exhibited a ~4-fold increase in total SBSs, compared to wt liver tissue (Figure 11a) (Paisie et al., 2016). Mouse kidney cell lines that survived nutritional stress (NS1 and NS4; characterized in chapter 2) in tissue culture displayed ~2500 total SBSs, whereas ~1500 SBSs were observed in the FHIT ko lung and kidney tissues (Figure 11b) (Paisie et al., 2016). Further examination of the types of transition and transversion mutations observed among these SBSs demonstrated that Fhit- deficient cells and tissues displayed prominent increases in C>T and T>C mutations, as well as slightly elevated levels of C>A, C>G and T>A mutations (Figure 11c) (Paisie et al., 2016). A spike in C>G mutations was observed in the NS lines, possibly acquired from the stress of subculturing or to allow survival of nutrient deprivation. C>T mutations can be generated by spontaneous deamination or DNA replication errors. T>C mutations may be due to the misincorporation of dUTP in place of TTP, a consequence of

TK1 down-regulation in Fhit-deficient cells. This Fhit loss mutation profile is comparable to previously published mutation signatures reported for bladder cancer, human papillary kidney cancer and an “age at diagnosis” signature, or aging signature, that represents accumulation of mutations in normal tissues over time (Figure 11d, 11e)

(Alexandrov et al., 2013; Paisie et al., 2016). As Fhit-deficient genomes accumulate mutations, it is not surprising then that many of these mutations are found within

Figure 11. Increased total single base substitutions (SBSs) and C>T and T>C mutations in Fhit-/- cells and tissues. Total SBS burden was calculated for (a) liver tissue whole exome sequences of Fhit+/+ and Fhit-/- mice and (b) kidney and lung tissue whole exome sequences of Fhit-/- mice and NS1 and NS4 Fhit-/- kidney cell lines. (c) Levels of transitions (C>T, T>C) and transversions (C>A, C>G, T>A, T>G). (d) Fhit loss mutation signature in -/- kidney tissue. (e) Signature 1b “age at diagnosis” signature from Alexandrov et al., 2013 (Images from Paisie et al., 2016; copyright license http://creativecommons.org/licenses/by-nc/4.0/legalcode).

Table 8. Sequence analysis of Trp53, E2f1, Mcm6 and Herc2 in Fhit+/+ and Fhit-/- NS mouse kidney cell lines. The amino acid changes observed are listed. The single base substitutions are as follows: Trp53 (C>G), E2f1 (C>T), Mcm6 (G>C), Herc2 (G>C), and Ptch1 (G>C). N, no mutation or deletion.

important genes, which when altered, contribute to cellular transformation and preneoplastic changes. For the NS1 and NS4 mouse kidney cell lines, such genes include

Trp53, E2f1, Mcm6, Herc2 and Ptch1, which are essential for cell cycle regulation, cell growth, cell survival and DNA replication and repair. Mutations are listed in Table 8.

These genomic alterations, which are a downstream consequence of TK1 inactivation, confer selective growth advantages to cells and drives cellular transformation changes that have been associated with tumorigenicity. Thus, Fhit functions as a genome caretaker through controlling expression of TK1. But how is Fhit, a cytoplasmic protein, affecting expression of an enzyme essential for efficient DNA synthesis?

3.3.3 A post-transcriptional role for Fhit

In addition to changes in TK1 protein expression, loss of Fhit also causes ~80% reduction in TK1 mRNA levels in mouse kidney cells (Figure 12a), and can be restored upon activation of the Fhit transgene in H1299 cells (Figure 12b). These results corroborate previous reports where silencing of Fhit in 293 and HCT116 cells led to a

60% decrease in TK1 mRNA levels (Waters et al., in revision). Because TK1 mRNA levels are influenced by Fhit absence, Waters et al. hypothesized that Fhit may play a role in either gene transcription or stability of TK1 mRNA. To determine if Fhit controls TK1 transcription, a firefly luciferase assay was performed using a construct containing full- length TK1 promoter regulatory region cloned upstream of the firefly luciferase gene. No significant difference in firefly luciferase activity, and therefore TK1 promoter activity, was observed in Fhit-deficient or Fhit-induced cells (Figure 12c), suggesting that Fhit regulates TK1 at a step downstream from transcription. To determine if Fhit stabilizes

TK1 mRNA, the H1299 system was used to examine the half-life of TK1 mRNA in cells with and without Fhit expression following transcription inhibition. RT-qPCR analysis of

TK1 mRNA revealed that decay was more rapid in the absence of Fhit than in its presence (Figure 12d), exhibiting a half-life of 4 hr. A majority of TK1 mRNA remained after 8 hr in the presence of Fhit. Thus, Fhit expression level has an effect on level of

TK1 mRNA over time.

Figure 12. Fhit regulation of TK1 mRNA levels. RT-qPCR analysis of endogenous TK1 mRNA levels in (a) Fhit-/- and Fhit+/+ mouse kidney cell lines and (b) H1299 cells containing EV or induced wt FHIT plasmid. Cells were harvested 48 hrs after treatment with ponasterone A. TK1 mRNA expression was normalized to GAPDH mRNA levels. (c) Luciferase assay to examine TK1 promoter activity. If Fhit were promoting TK1 transcription, elevated luciferase expression should be detected in the Fhit-expressing cells. The y axis is relative to light units, setting the firefly luciferase/renilla luciferase RLU ratio to 1 to compare the reading in the Fhit positive vs negative cells. (d) RT-qPCR analysis of relative TK1 mRNA expression in EV and induced wt Fhit H1299 cells following treatment with DRB for 0-8 hrs. TK1 mRNA levels were normalized to RPLPOP mRNA levels (ribosomal protein, large, P0 pseudogene; ribosomal protein coding mRNA that is very abundant, for use as internal control transcript to normalize TK1 mRNA levels); sequences of the primers used: RPLP0-For GGAGAAACTGCTGCCTCATATC, RPLP0-Rev CAGCAGCTGGCACCTTATT

3.3.4 Fhit affects ribosome binding and translation of TK1 mRNA

There are many examples of RNA binding proteins that function to protect mRNAs from degradation, however, there are no published reports describing mRNA stabilization as a consequence of Fhit binding. mRNA stabilization can also be achieved through binding of translating ribosomes. Therefore, the impact of Fhit on ribosome binding of TK1 mRNA was assessed in H1299 cells, with and without Fhit expression via polysome fractionation. Briefly, cycloheximide is added to the growth medium of the cells to freeze ribosomes on the mRNA, thereby preventing ribosome runoff. Next, ribosomes are extracted and separated based on their density via sucrose density gradient centrifugation to isolate free ribosomal subunits (40S, 60S), monosomes (80S) and polysomes. RNA can then be extracted from sucrose fractions to analyze the abundance of specific mRNAs in pre-and polysomal fractions. In Fhit-expressing cells, there is a lower 40S peak, a greater 80S peak and a small but reproducible increase in polysomes compared to Fhit-deficient cells. In the context of bulk RNA, these results consistently show a positive correlation between Fhit and increased translation activity (Figure 13).

To determine if ribosome binding affects TK1 mRNA stability and translation efficiency,

RT-qPCR analysis of TK1 mRNA was performed at alternating fractions. In the absence of Fhit, the majority of TK1 mRNA is found in fractions 7 and 9, which correspond to the

80S peak (1 ribosome bound) and the first polysome peak (2 ribosomes bound) respectively; minimal TK1 mRNA is distributed in heavy polysomal fractions containing three or more bound ribosomes. In contrast, Fhit-expressing cells display a shift in TK1 mRNA distribution towards the heavy polysomal fractions with ~25% of total TK1

58 mRNA associated with five to eleven bound active ribosomes. Therefore, Fhit-expressing cells generate more protein per TK1 message. Thus, Fhit affects ribosome binding and translation of TK1 mRNA substantiating the finding that Fhit loss reduces TK1 mRNA stability and decreases TK1 protein levels.

Figure 13. Impact of Fhit on polysome binding of TK1 mRNA. Cytoplasmic extracts from EV and wt Fhit-expressing H1299 cells were separated on a 10-50% sucrose gradient. Gradient fractions were monitored continuously at 254 nm, spiked with equal amounts of luciferase RNA as a control for sample recovery, and odd-numbered fractions were analyzed for TK1 and luciferase by RT-qPCR. The data are plotted as the amount of TK1 mRNA relative to the Fhit negative sample in fraction 5. Free ribosomal subunits (40S and 60S), monosomes (80S) and number of ribosomes in the polysome fractions are indicated. 60

3.4 Discussion

Many laboratories have studied the FHIT gene and the function of Fhit protein since its discovery at a familial translocation breakpoint in 1996 (Ohta et al., 1996).

Although Fhit has been shown to affect many cancer-associated functions, lack of a known mechanism of action for Fhit has encouraged the claim that Fhit loss is a passenger event in cancer development. The first glimpse into the molecular basis of Fhit function was reported in 2012 by the exciting finding that Fhit loss leads to accumulation of DNA damage due to defective nucleotide metabolism and imbalance of dNTP pools

(Saldivar et al., 2012). Specifically, Fhit loss resulted in down-regulation of TK1 protein, an enzyme involved in biosynthesis of thymidine triphosphate through the scavenger pathway. Because Fhit is a small cytoplasmic protein, with very few interacting protein partners (Pichiorri et al., 2009; Trapasso et al., 2008), discovery of mechanisms underlying Fhit functions has been difficult. Therefore, discovery of a direct connection between Fhit and TK1 expression levels was a breakthrough finding, providing a new avenue of investigation for potential mechanisms.

The data presented here show that both TK1 mRNA and protein levels are reduced upon loss of Fhit. Because TK1 mRNA was affected by Fhit loss while stability of the protein was not altered by Fhit status, it was proposed that Fhit either acts at the transcriptional or post-transcriptional level. TK1 gene transcription regulation was ruled out by results showing that Fhit does not control TK1 expression through an effect on the

TK1 promoter. TK1 mRNA turnover results showed that TK1 mRNA is more stable in the presence of Fhit than in its absence. Generally, mRNA stabilization occurs through

61 protein binding to a particular sequence element blocking access of a degradative or destabilizing protein (Schoenberg and Maquat, 2012), but there are no reports identifying

Fhit as an RNA-binding protein (Castello et al., 2012). Instead, polysome fractionation studies demonstrate that Fhit affects mRNA stability through ribosome binding. TK1 mRNA in Fhit-positive cells primarily sediments on polysomal fractions, whereas TK1 mRNA in Fhit-negative cells is most abundant toward the top of the gradient corresponding to non-translating or poor-translating complexes. Thus, reduced ribosome binding to TK1 mRNA leads to TK1 mRNA instability and decreased translation efficiency resulting in reduced TK1 protein levels.

The post-transcriptional repression of TK1 mRNA provides novel evidence for a mechanism connecting loss of Fhit with genome instability and cancer development.

Multiple studies have shown that reduced TK1 protein expression results in depletion of

TTP pools causing replication fork stalling and collapse into DSBs. The checkpoint remains blind to this replication stress-induced damage allowing accumulation of genomic changes. As observed in Chapter 2, this ultimately results in changes in expression patterns promoting tumorigenesis in vitro and in vivo in Fhit-/- cells. The accumulation of damage can be prevented upon addition of thymidine, demonstrating the importance of TK1 expression regulation for genome stability. Another direct consequence of TK1 down-regulation is the increase in T>C and T>G mutations. In Fhit- deficient cells, the decrease in dTTP concentrations allows for increased dUTP misincorporation during DNA synthesis. Excision of dUTP by uracil DNA glycosylase creates an abasic site in the DNA. Guanines and cytosines are frequently inserted by

62 translesion polymerases opposite abasic sites, and following the next round of replication,

T>C or T>G mutations result (Waters et al., 2009; Prakash et al., 2005).

Recently, Taverniti and Seraphin (2015) reported a role for Fhit in metabolizing free mRNA cap (m7GpppN) dinucleotides generated by 3‟ to 5‟ mRNA decay. Initial characterization of Fhit had shown that in vitro it cleaves dinucleoside triphosphates, molecules that resemble mRNA 5' caps (Figure 14a) (Barnes et al., 1996). Interestingly, cancer cell lines that have lost Fhit expression produce excess Ap3A (Murphy et al.,

2000). Eukaryotic mRNAs contain a 7-methyl guanosine cap structure that is co- transcriptionally added to the 5‟ terminus. This cap structure interacts with cap-binding proteins to carry out a variety of important functions such as pre-mRNA splicing, 3‟-end processing, nuclear export and translation initiation. Regulation of gene expression can be tightly regulated through mRNA decay, in which two main pathways exist. Both pathways begin with the exonucleolytic removal of the poly(A) tail. In the 5‟ to 3‟ decay pathway, this deadenylation step is followed by hydrolysis of the cap and subsequent digestion of the mRNA body by a 5‟ exonuclease. In the 3‟ to 5‟ decay pathway, deadenylation is followed by 3‟ exonuclease digestion of the mRNA body, resulting in the release of free cap structures. Free cap (m7GpppN) dinucleotides can be cleared through further hydrolysis by scavenger decapping enzymes. Alterations in free cap concentration has been shown to impact pathological conditions (Gogliotti et al., 2013).

Thus, scavenger decapping enzymes play a pivotal role in clearing free cap dinucleotides from the cell to maintain steady-state levels of translating mRNAs.

Figure 14. Fhit hydrolyzes free m7GpppN cap structures. (a) Structure of diadenosine triphosphate (Ap3A), the first recognized in vitro substrate for Fhit, and the 5‟ 7-methylguanosine cap. (b) Products resulting from the incubation of m7GpppG with recombinant proteins were fractionated by TLC and detected by autoradiagraphy, and western blot of purified proteins (Image from Taverniti and Seraphin, 2015; copyright license http://creativecommons.org/licenses/by-nc/4.0/legalcode ).

Studies have shown that aberrant accumulation of m7GpppN competes with and sequesters translation initiation factor eIF4E from its mRNA substrates to inhibit polysome formation and in vitro translation (Hammond et al., 2016). Through thin layer chromatography, Taverniti and Seraphin (2015) showed that wt FHIT cleaves m7GpppN into m7GDP and m7GMP in vitro, whereas the catalytically dead mutant, H96N, could not cleave m7GpppN (Figure 14b). The yeast homolog, Aph1, also demonstrated cleavage into m7GDP which did not occur following inactivation of its enzymatic

64 function (Taverniti and Seraphin, 2015). Therefore following Fhit loss, a decrease in scavenger decapping activity would result in elevated pools of m7GpppN dinucleotides that might compete with selected capped mRNAs for binding by eIF4E, a cytoplasmic cap-binding protein that is required for cap-dependent translation (Figure 15). This model could explain the decline of TK1 mRNA observed in polysome fractions in Fhit-deficient cells.

Figure 15. Model for Fhit as a scavenger decapping enzyme regulating TK1 mRNA translation. In the 3‟ to 5‟ mRNA decay pathway, the exosome generates free m7GpppN dinucleotides that can be hydrolyzed by a scavenger decapping enzyme. In the presence of Fhit, Fhit binds and hydrolyzes m7GpppN into m7GDP and m7GMP which are cleared from the cell. In the absence of Fhit, free m7GpppN caps accumulate. Preferential binding of translation initiation factors to these free caps instead of capped TK1 mRNAs leads to deregulated translation of TK1 mRNA. Thus, Fhit is a scavenger decapping enzyme that eliminates residual cap structures to promote ribosomal binding and translation of cap-bearing mRNAs.

As mentioned previously, the tumor suppressor function of Fhit is independent of its hydrolytic activity, but dependent on binding of its substrate (Trapasso et al., 2003). Is binding of Fhit to m7GpppN dinucleotides sufficient to prevent further binding by eIF4E? Or is it possible that Fhit and eIF4E could bind these free dinucleotides together, in which case m7GpppN hydrolysis would be necessary to prevent eIF4E sequestration.

Ongoing investigations will determine if both m7GpppN binding and hydrolysis are necessary for Fhit genome caretaker function. TK1 expression and levels of DNA damage will be assessed in the H1299 system where ponasterone A treatment will induce expression of wt Fhit, H96N Fhit or Y114A Fhit. The H96N mutant can bind but not hydrolyze its substrate, whereas the Y114A mutant cannot bind or hydrolyze. If binding alone is sufficient to sequester free caps, then we would expect to see the H96N mutant to restore TK1 expression levels and display decreased levels of DNA damage.

We have proposed for the first time that Fhit affects the „translatome‟ and have identified an mRNA directly affected by loss of Fhit, TK1. We hypothesize that Fhit affects expression of other genes through a similar mechanism which contribute to Fhit tumor suppressor and genome caretaker functions. Ribosome profiling will be used to identify other mRNAs that are affected by Fhit loss. This technique provides a snapshot of mRNAs that are being actively translated in a particular condition or moment in time.

Ribosome-mRNA complexes are isolated and any RNA not protected by ribosomes is digested by ribonucleases. The remaining RNA is then sequenced using library synthesis and data analysis similar to normal RNA-seq. Identifying other genes regulated by Fhit

67 will provide novel targets to selectively eliminate Fhit-deficient cells in the early stages of cancer development.

Chapter 4: Discussion

Conclusions and future perspectives

Contains portions from manuscript: Karras et al., Cytogenet Genome Res 2016 (In

review)

4.1 Introduction

Common fragile sites (CFSs) generally encompass 0.6 to >1 MB of genomic

DNA and are found in regions which replicate late during S-phase (Smith et al., 2006; Le

Beau et al., 1998; Hellman et al., 2000; Le Tallec et al., 2011). For years, specific types of DNA sequences were sought that might explain the differences between fragile and non-fragile genomic regions. Recently the fragility of specific CFSs has been shown to vary based in large part on the frequency and distribution of replication origins, which differ in cells derived from different tissues, such as lymphocytes, epithelia and connective tissue. Thus, if replication origins are only found flanking a very large gene that replicates late, replication might not be completed by the end of S phase, leaving regions of the gene un-replicated, and resulting in loss of portions of the gene in daughter cells (Letessier et al., 2011). Such deletions occur at the two most active CFSs, at FRA3B and FRA16D, within the respective FHIT and WWOX genes, and their encoded proteins are inactivated in many human cancers. Fhit protein loss is reported in at least 50% of

69 preneoplasias and human cancers that develop in epithelial tissues. Likewise, Wwox- deficiency is observed in a wide variety of human cancers, and is often associated with more aggressive cancers at a higher stage and with poorer outcomes (Nunez et al., 2005;

Guler et al., 2009).

Due to the inherent instability of CFS loci and the lack of inactivating point mutations characteristic of “classical tumor suppressors” within these fragile gene products, some investigators dismiss the idea that deletions within CFSs contribute to clonal expansion of neoplastic clones. However, the current explanation for CFS fragility, which relies on a cell or tissue type specific, epigenetically regulated, placement of cellular replication origins, has shed an interesting perspective on this claim. For instance, FRA16D is the most active CFS in the epithelial cell types thus far tested (Le

Tallec et al., 2013; Hosseini et al., 2013), and allele loss within the WWOX locus has been observed most frequently in hormonally-regulated epithelial cancers, such as breast, ovarian and prostate cancers. This suggests some selectivity for Wwox loss in these cancers, in which Wwox loss imparts a selective growth or survival advantage. In contrast, many Fhit-deficient cancers originate from epithelial cells, although the

FRA3B/FHIT locus is not the most active fragile region in the epithelial cells thus far examined for fragility. Since FHIT has been reported as the second most commonly deleted gene among human cancers (Beroukhim et al., 2010; Bignell et al., 2010), but is not the most fragile locus in epithelial cells (LeTallec et al., 2013, Hosseini et al., 2013), this supports the proposal that loss of Fhit expression contributes to clonal expansion

(Saldivar et al., 2012; Miuma et al., 2013). In support of this view, this research work

70 demonstrates that FHIT and its fragile locus FRA3B play unique biological and tumor suppressive roles. This chapter summarizes accumulated evidence that Fhit protein functions as a genome „caretaker‟ required to protect the stability of genomes of normal cells of most tissues from agents causing intrinsic and extrinsic DNA damage. Fhit loss leads to intracellular replication stress and subsequent genome instability, which provides an opportunistic mutational landscape in preneoplasias for selection of a variety of other cancer-driving mutations.

4.2 Initiation of genome instability through loss of the FRA3B gene product, Fhit

Genomic instability is an evolving hallmark of cancer, but when and how genome instability originates during cancer development is a subject of ongoing investigation.

Familial cancers carry germ-line mutations in genes important for the response to and repair of DNA damage (for review, Negrini et al., 2010). These genes help to maintain the integrity of the genome and have therefore been designated „genome caretakers‟

(Kinzler and Vogelstein, 1997). For example, mutations in the DNA mismatch repair pathway can cause familial non-polyposis colon cancer and inherited mutations of

BRCA1 and 2 genes, required for homology directed repair, are associated with familial breast and ovarian cancers. Mutation or silencing of these familial cancer genes do occur but are not common in the context of sporadic cancers. Therefore many models have been proposed for initiation of genome instability, such as telomere erosion, impaired

DNA repair and chromosome segregation errors (Halazonetis et al., 2008). These processes are certainly contributors to genome instability during cancer development but

71 are not necessarily common instability initiating events, as they are observed in more advanced lesions (Halazonetis et al., 2008). Oncogene-induced replication stress has also been proposed as a model for initiating instability. However, while oncogene activation does cause genome instability, it often leads to cellular senescence halting proliferation of the cells unless they receive a second “hit” (Lowe et al., 2004). This suggests that there must be some pre-existing level of instability already present. These hypotheses have overlooked the role of Fhit in initiating genome instability. Two key details that make

Fhit an ideal candidate are 1) loss of Fhit, due to genetic or epigenetic alterations, may be the earliest change observed in hyperplastic lesions (Gorgoulis et al., 2005) and 2) loss of

Fhit promotes replication stress-induced genome alterations without activating the DNA damage response pathway (Saldivar et al., 2012). The lack of checkpoint activation implies that there is a threshold for DNA damage that must be met for the cell to stop progression through the cell cycle. This criterion is not met by Fhit loss-induced replication stress. Therefore Fhit loss causes mild replication stress that persists through each cell division and contributes to development of genome-wide instability (Miuma et al., 2013, Paisie et al., 2016; Karras et al., 2014). Coupled with the fact that loss of Fhit expression is one of the earliest changes in the preneoplasia process, Fhit deletion is a strong candidate for being an initiator of genome instability that fuels the development of many cancers.

Hanahan and Weinberg (2000) described six alterations in cell physiology that most, if not all, cancers acquire for malignant development. These six cancer hallmarks include: sustained proliferative signaling, evasion of growth suppressors, evasion of

72 apoptosis, induced angiogenesis, activation of invasion and metastasis, and replicative immortality (Hanahan and Weinberg, 2000). Following the lifespan of mouse kidney epithelial cells in Chapter 2, each of these traits were observed following loss of Fhit: Up- regulation of replication fork proteins, proto-oncogene Cdc6 and Cyclins E1, E2 and B1 demonstrate that Fhit-deficient cells are capable of maintaining a highly proliferative status; Evasion of growth suppressors and apoptosis can both be achieved through the inactivating point mutation of Trp53 within the DNA binding domain; Since p53 is also an important inhibitor of angiogenesis, this mutation could also support the formation of tumor vasculature. However, other inducers of angiogenesis such as VEGF-A and

VEGF-C are also shown to be up-regulated in the transcriptome profiling data.

Activation of the EMT process with subsequent demonstration of tumor formation and lung colonization in vivo explicitly shows that Fhit-/- cell lines acquire invasive and metastatic capability. Finally, increased telomerase expression and decreased expression of telomerase inhibitor, as observed through transcriptome profiling, provides a means for replicative immortality in Fhit-deficient cells. Although the sequence in which these six cancer hallmarks are acquired may vary, this research shows that they can all follow loss of Fhit. Importantly, the Fhit-expressing mouse kidney cells did not acquire any of these alterations required for malignant growth. Thus, Fhit inactivation can be seen as the first “hit” for tumorigenesis whereby replication stress-induced genome instability facilitates the onset of additional cancer hallmarks.

4.3 Expression of Fhit and wt p53 are lost coordinately in many cancers

Cooperation of p53 mutation and Fhit loss is a frequently observed phenomenon in cancer. In lung cancer, these are the two most commonly altered tumor suppressor genes and have been shown to be candidate prognostic markers (Andriani et al., 2012).

Many non-small cell lung tumors that exhibit FHIT locus LOH also exhibit p53 missense mutations (Burke et al., 1998). The p53 protein has several functional domains that are responsible for sequence-specific DNA binding, transcriptional activation, and oligomerization (Vousden and Prives, 2009; Beckerman and Prives, 2010). In more than half of all sporadic human cancers, p53 is inactivated by a single point mutation, most frequently in the DNA binding domain. These mutations inactivate the p53 protein by altering a residue that directly binds DNA and leads to faulty transactivation of target genes (Joerger and Fersht, 2008). When both p53 and FHIT are inactivated, early stage lung tumors exhibit proliferation irregularities that contribute to the progression of lung cancer, and suggest some form of cross talk between the two tumor suppressors (Andriani et al., 2012). This combined genetic deficit, described as a “double hit”, is also associated with poorer prognosis in head and neck squamous cell carcinomas when compared to the loss of each gene individually. Studies have indicated that this double hit promotes resistance to radiation treatment and increased MMP2/9 activity, proteins that degrade the extra cellular matrix and are essential for metastasis (Raju et al., 2015). Others claim that this double hit provides a selective advantage, as p53 mutations associate with loss of 3p more than any other chromosomal region (Gross et al., 2014). In Chapter 2, an in vitro

74 model of Fhit loss-associated transformation was described. Interestingly, the p53 mutation identified was the first alteration observed following Fhit loss. This mutation resides in the DNA binding domain of p53 impairing transactivation of the CDKN1A gene encoding p21 protein. Again, this double hit provides proliferative advantage and accelerates the acquisition of mutations for preneoplastic changes.

4.4 Fhit loss and the mutator hypothesis

DNA sequencing of tumors has uncovered patterns of mutations and genomic changes across tumor types, providing insight into the evolution of cancer. These analyses have shown that each tumor is unique, often exhibiting thousands of mutations, although a limited number of these alterations are responsible for driving cancer development (Kandoth et al., 2013). How are cancer cells acquiring these exceptionally high numbers of mutations? To explain the discrepancy between numbers of mutations in normal cells vs cancer cells, Lawrence Loeb proposed the mutator hypothesis (Loeb,

2011). This hypothesis states that early in cancer development, mutation of a DNA caretaker gene elevates the mutation rate in all daughter cells. This increase in mutation frequency results in genetic diversity that can accelerate tumorigenesis by increasing the probability of acquiring favorable mutations for cancer progression. This process creates a heterogeneous tumor composed of clonal, subclonal and random mutations that may be selected for when the environment changes. Saldivar et al. (2012) showed that Fhit loss causes genome instability, but does loss of Fhit caretaker function, which occurs in the earliest preneoplastic lesions, constitute a mutator phenotype?

Many studies have implicated replication stress as a source of point mutations and small insertions/deletions in cancer cells. Specifically, imbalance of dNTP pools, which can give rise to replication stress, has been linked to increased mutagenesis. In yeast, base substitutions and insertions/deletion rates were drastically elevated at the CAN1 locus when dNTP pool size or balance was altered (Kumar et al., 2010). Fission yeast also exhibit an increased rate of spontaneous base substitutions and small insertions/deletions with decline in dNTP concentrations (Holmberg et al., 2005). On the flip side, a consequence of replication stress is the collapse of stalled replication forks into DSBs.

Single nucleotide changes and small insertions/deletions can result from the repair of

DSBs via non-homologous end joining (Villarreal et al., 2012) or homologous recombination (Deem and Keszthelyi, 2011). Often, these are observed as non-random, clustered mutations (Nik-Zainal et al., 2012). Loss of the genome stabilizing function of

Fhit causes replication stress through nucleotide imbalance resulting in DNA breaks. This replication stress increases the frequency of point mutations, deletions and chromosome aberrations, classic features of a mutator phenotype (Paisie et al., 2016; Miuma et al.,

2013). Fhit loss-induced replication stress also generates enhanced ssDNA templates which serve as substrates for the A3B deaminase mutator to convert C>T and generate a hypermutator phenotype in Fhit-deficient/A3B-overexpressing cells (Waters et al., 2015).

In addition, Fhit loss is observed in cancers exhibiting hypermutation, such as breast and lung cancer. These results confirm that Fhit loss causes a mutator phenotype through induction of replication stress.

4.5 The Fhit-TK1 pathway in preneoplasia vs cancer

TK1 overexpression in Fhit-negative cancer cells provides a curious twist to the

Fhit-TK1 story. Indeed, expression analysis of 345 lung tumor samples obtained from

The Cancer Genome Atlas revealed a negative correlation between Fhit and TK1 expression (correlation coefficient = -0.251) (Figure 16). Interestingly, the level at which

TK1 is controlled in cancer cells is not at the same post-transcriptional level observed in normal and preneoplastic cells (unpublished data from Huebner lab). Thus, up-regulation of TK1 is not due to an increase in TK1 mRNA stability, but possibly due to protein stabilization (data not shown). This suggests that cancer cells find a way to bypass the translational block imposed by Fhit loss to ramp up dTTP production. Because cancer cells need to sustain a high proliferative status, this would be an advantageous alteration to increase efficiency of DNA synthesis. Therefore, TK1 overexpression is a selected event during Fhit-negative cancer development. In addition to TK1, Fhit-negative cancers also show over-expression of other enzymes involved in dTTP synthesis, such as

TYMS and RRM2 (Figure 16).

Figure 16. TK1 overexpression in Fhit-negative lung tumors. RNA-seq data obtained from The Cancer Genome Atlas shows a negative correlation between Fhit expression and expression of TK1, RRM2 and TYMS, enzymes involved in dTTP synthesis. Red, up-regulated; Green, down-regulated.

Following the lifespan of the mouse kidney cells described in Chapter 2, this shift in TK1 over-expression was observed as the Fhit-/- cells became tumorigenic. Initially, the unstressed Fhit-/- cells display drastically reduced TK1 protein levels compared to

Fhit+/+ cells (Figure 10a). However, transcriptome profiling results reveal that the in vitro-transformed cell line, NS3 colony, exhibits a 3.96 fold elevation in TK1 mRNA expression compared to -/- kd3 p48. RT-qPCR analysis validated this finding (Figure

17a). Furthermore, in these late passage colony cells, thymidine supplementation no longer prevents accumulation of DNA breaks in cells over-expressing TK1 (Figure 17b).

A transient down-regulation of TK1 followed by TK1 over-expression is also observed in an additional mouse kidney cell line described by Miuma et al. (2013). Fhit expression decreases in the MK+/+ cells over several passages and is accompanied by concomitant 78

Figure 17. TK1 up-regulation in late passage mouse kidney cells. (a) RT-qPCR analysis of endogenous TK1 mRNA levels in +/+ kd2 p14, -/- kd3 p48 and -/- NS3 colony p13 mouse kidney cell lines. TK1 mRNA expression was normalized to GAPDH mRNA levels. (b) Assessment of DNA damage in cells over-expressing TK1. Box plots of tail moments: mouse kidney cells -/- kd4 without thymidine, n=73; -/-kd4 + thymidine, n=98; NS3 without thymidine, n=85; NS3 + thymidine, n=94. Thymidine supplementation (10 µM) for 16 days and includes data from two separate experiments. (c) Immunoblot of TK1 expression in MK cells. As Fhit expression decreases in MK+/+, TK1 expression decreases. In MK-/-, loss of Fhit results in TK1 down-regulation. This effect is transient as TK1 expression increases at late passage.

79 loss of TK1 expression. This is due to the increasing instability of TK1 mRNA as Fhit is lost. The MK-/- cells displayed reduced TK1 protein levels at early passages, and TK1 expression was up-regulated at late passage 22 (Figure 17c). These results suggest that selective clonal expansion of a TK1-overexpressing cell dominated the population. Fhit loss-induced genome instability creates an environment that supports mutation and precancerous changes to allow for selective clonal outgrowth of cells carrying advantageous alterations, such as evolution of clones with up-regulated TK1.

4.6 Conclusions

The results of this research, as summarized here in Chapter 4, show that a single genomic or epigenomic alteration leading to loss of Fhit protein expression can cause a cascade of damage and deregulation in cells. Following Fhit loss, replication stress- induced DNA damage is propagated in daughter cells without activating the cell cycle checkpoint and ultimately contributing to global genome instability in the form of mutations, indels, CNAs and aneuploidy. Fhit loss-induced replication stress also generates ssDNA substrates for the A3B enzyme, illustrating the role of Fhit loss in hypermutation genotypes of many cancers. This instability increases the genetic diversity allowing the acquisition of cancer hallmarks required for clonal outgrowth and malignant transformation. Thus, Fhit is a guardian of stability.

The research presented here also provides new insights into the mechanism by which Fhit affects genome stability. For the first time, it has been proposed that Fhit affects the „translatome‟, and an mRNA directly affected by loss of Fhit, TK1, has been

80 identified. Profiling Fhit-deficient associated translatomes may identify additional mRNAs whose altered translation contributes to Fhit tumor suppressor or genome caretaker functions. This will be very important for the future development of novel therapeutic strategies based on protein signals directly altered by loss of Fhit. For example, understanding how the translatome is altered in Fhit negative cells of specific tissue types may identify signal pathways that provide suitable targets for synthetic lethality approaches to selectively eliminate Fhit-deficient cells early during cancer progression.

Collectively, these findings support a model where deregulation of the Fhit-TK1 pathway initiates global genome instability in preneoplastic cells to promote tumorigenesis. The demonstration of neoplastic initiation and progression following Fhit loss, in combination with discovery of a mechanism of action for Fhit, delivers an explicit message that deletion at the FHIT locus is not simply a passenger event. Thus, Fhit is a guardian of stability that when lost, drives cancer development.

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