Classes Of Ribonucleotide Reductase

Published Date: 02 Nov 2017

Deoxynucleoside triphosphates (dNTPs) are the "building blocks" of DNA and are essential for DNA replication and repair. Levels of dNTPs must be strictly regulated. When dNTP levels are too low, DNA replication is slowed down, generating single-stranded DNA (ssDNA), which increases susceptibility to chromosome breakage (Cha & Kleckner, 2002). On the other hand, elevated dNTP levels lead to mutagenesis, accelerated DNA replication, and inhibition of cell cycle arrest (Chabes & Stillman, 2007; Chabes et al., 2003; Poli et al., 2012). These cellular phenomena are frequently associated with carcinogenesis and tumorigenesis. The levels of dNTPs in a cell are dependent upon the activity of the enzyme ribonucleotide reductase (RNR), which catalyzes the rate-limiting step of dNTP synthesis. Saccharomyces cerevisiae (baker’s yeast) containing the mutation rnr1D57N exhibit increased levels of dNTPs (Chabes et al., 2003). Dr. Jennifer Surtees at the University of Buffalo conducted a screen to identify individual genes in the yeast genome that, when deleted and then combined with the rnr1D57N mutation, cause increased or decreased growth relative to the respective single mutant deletion strain. One of the genes that yielded increased growth was XRS2. The protein Xrs2 is a component of the MRX complex, which repairs double-strand breaks (DSBs) in DNA. Based on the results in the screen, I can hypothesize that XRS2, when mutated the presence of altered dNTPs, leads to enhanced growth rates in mammalian cells as well and therefore have implications for cancer research. The overall goal of this project is to confirm the results from the screen regarding XRS2 and to characterize the interaction of Xrs2 with elevated dNTP pools.

Literature Survey

In all cells, new deoxyribonucleic acid (DNA) is formed when individual deoxynucleoside triphosphate (dNTP) molecules lose two of their three phosphate groups and then form phosphodiester bonds with each other via a dehydration reaction. These dNTP "building blocks" of DNA are essential for DNA replication and repair. Levels of dNTPs must be strictly regulated. When dNTP levels are too low, DNA replication is slowed down, generating single-stranded DNA (ssDNA), which increases susceptibility to chromosome breakage (Cha & Kleckner, 2002). On the other hand, elevated dNTP levels lead to mutagenesis, accelerated DNA replication, and inhibition of cell cycle arrest (Chabes et al., 2003; Poli et al., 2012; Chabes & Stillman, 2007). These cellular phenomena are frequently associated with carcinogenesis and tumorigenesis.

In order to acquire dNTPs, a cell must synthesize them by reducing nucleoside diphosphate (NDP) into deoxynucleoside diphosphate (dNDP) and then phosphorylating the dNDP. The reduction of NDP is the rate-limiting step of this process, and it is catalyzed by the enzyme ribonucleotide reductase (RNR). This specific process and the role of RNR is strictly conserved among all known organisms (Torrents et al., 2002). Furthermore, RNR regulates DNA synthesis and influences DNA/cell mass ratios in organisms (Herrick & Sclavi, 2007). All reactions facilitated by RNR proceed via a free radical mechanism(Wijerathna et al., 2011). The substrates for the RNR enzyme are adenosine diphosphate (ADP), guanosine diphosphate (GDP), cytidine diphosphate (CDP), and uridine diphosphate (UDP). The products deoxyadenosine diphosphate (dADP), deoxyguanosine diphosphate (dGDP), and deoxycytidine diphosphate (dCDP) are converted into their dNTP counterparts while deoxyuridine diphosphate (dUDP) is converted into thymidine monophosphate (dTMP) and then thymidine diphosphate (dTDP) in a series of reactions before becoming thymidine triphosphate (dTTP) (Schaaper & Mathews, 2013).

Classes of ribonucleotide reductase

RNR enzymes are divided into three classes based on the metal cofactor they use to generate an initial free radical, which is delivered to a conserved cysteine residue in the enzyme’s active site, leading to the formation of a thiyl radical. Class I enzymes contain two subunits, R1 and R2. R1 has a catalytic site and one or two allosteric sites. R2 has iron ions that are responsible for the oxygen-dependent generation of a tyrosyl free radical, which is necessary for initiating catalysis. Class I enzymes are further divided into Ia and Ib based on differences in their regulation. Class Ia reductases are found in all eukaryotic organisms, in prokaryotes, and in viruses that infect eukaryotes (Wijerathna et al., 2011). Class Ib reductases are present only in bacteria and utilize manganese ions as well as iron ions to generate free radicals. There are two allosteric sites present on Ia enzymes but only one on Ib enzymes. Another difference is that Ia reductases are inhibited by dATP via a feedback inhibition mechanism while Ib reductases are not (Jordan & Reichard, 1998). Since the vast majority of eukaryotes carry class Ia enzymes, mutations in the genes encoding these particular enzymes are useful for studying the effects of RNR on genome stability its connections with mammalian carcinogenesis.

Class II RNR enzymes have the simplest structure among the three classes and are found in prokaryotes and bacteriophages (Jordan & Reichard, 1998). These enzymes use cobalt-containing adenosylcobalamin (also known as cobamamide or dibencozide), an active form of vitamin B12, as a cofactor to generate a deoxyadenosyl free radical in a reaction that can occur with or without oxygen present (Wijerathna et al., 2011). Class II enzymes reduce both NDP’s and NTP’s, contain one allosteric site, and are not inhibited by dATP (Jordan & Reichard, 1998).

Class III RNR enzymes can be found in archae, anaerobic bacteria, and bacteriophages. These enzymes use iron-sulfur clusters coupled with S-adenosylmethionine to produce a glycyl free radical under anaerobic conditions. Class III enzymes reduce NTP’s, have two allosteric sites, and are inhibited by dATP (Jordan & Reichard, 1998). This class is thought to be evolutionarily the most ancient RNR enzymes (Wijerathna et al., 2011). Although the amino acid sequences differ greatly among the three classes, they are believed to have evolved from one common ancestor enzyme of which class III reductases are the closest relative (Reichard, 1993).

Organisms are not limited to having just one of the three classes of RNR enzymes. For example, Escherichia coli houses both class Ia and class III reductases (Jordan & Reichard, 1998). The vast majority of eukaryotic organisms only carry a class Ia enzyme. In the model organism Saccharomyces cerevisiae (yeast), the enzyme is a product of four polypeptides forming a complex. These polypeptides are encoded by the genes RNR1, RNR2, RNR3, and RNR4.

The S. cerevisiae ribonucleotide reductase

Because S. cerevisiae is such a useful and convenient model organism for studying intracellular mechanisms and processes of eukaryotes, much of the work done on RNR has been conducted using this yeast species. The R1 (alpha) subunit of the enzyme is a homodimer produced by the highly homologous genes RNR1, which is essential for mitotic viability, and RNR3, which is non-essential and induced upon DNA damage (Huang & Elledge, 1997). This dimeric subunit contains the catalytic and allosteric sites of the enzyme (Nordlund & Reichard, 2006). It can exist as a Rnr1/Rnr1 dimer, a Rnr1/Rnr3 dimer, or an Rnr3/Rnr3 dimer, however, the latter does not form stable RNR complexes (Domkin et al., 2002). The R2 (beta) subunit is a homodimer in most eukaryotes but is a heterodimer in yeast encoded by the essential genes RNR2 and RNR4 (Huang & Elledge, 1997). This subunit houses the oxygen-bridged dinuclear iron center, which generates the tyrosyl free radical (Nordlund & Reichard, 2006). Yeast cells lacking Rnr3, rnr3Δ strains, have identical dNTP dynamics and dATP pools compared with wild-type counterparts. Studies have also found that rnr3Δ yeast have no obvious phenotype and do not have heightened sensitivity to DNA damage (Chabes et al., 2003). Rnr4 mutants have been shown to be supersensitive to the RNR inhibitor hydroxyurea (HU) and fail to grow due to S-phase arrest at restrictive temperature (Wang et al., 1997). The mutant strain rnr2-314 was also shown to have increased sensitivity to HU as well as to methyl methanesulfonate (MMS), a DNA-damaging agent (Elledge & Davis, 1989).

The rnr1D57N mutation

The yeast mutant strain rnr1D57N is useful for evaluating the effects of elevated dNTP levels on cellular pathways and processes that can lead to carcinogenesis. As a result of this mutation, the 57th amino acid residue of the Rnr1 polypeptide is changed from aspartic acid to asparagine. The Rnr1 polypeptide contains an allosteric site for dATP inhibition. This particular mutation directly affects the dATP binding site and causes the RNR enzymes in the mutant strain to be resistant to feedback inhibition (Chabes et al., 2003).

When the wild-type RNR1 gene is replaced with the rnr1D57N allele in the haploid yeast strain W1588-4C, dNTP pools are 1.6- to 2.0-fold greater compared with the wild-type control. The mutagen 4-nitroquinoline-N-oxide (4-NQO) is known to increase dNTP levels in yeast cells (Chabes et al., 2003). When rnr1D57N cells are treated with 4-NQO, dNTP levels become 20- to 30-fold greater than in untreated wild-type cells and 4-fold greater than in mutagen-treated wild-type cells. This synergistic increase in treated mutant cells suggests that dATP feedback inhibition is the principle mechanism for limiting the increase in dNTP pools following DNA damage. Furthermore, the rnr1D57N strain exhibits an increased rate of G→C and G→T transversions and frameshift insertions upon exposure to 4-NQO. Lastly, cells with the rnr1D57N mutation exhibit a dramatic increase in survival of DNA damage induced by 4-NQO and a modest increase in survival following UV irradiation (Chabes et al., 2003).

In yeast cells that continuously express rnr1D57N via induction of a GAL1 promoter, the DNA-damage checkpoint mediated by Rad53 is inhibited, and the cell cycle is not arrested after exposure to DNA-damaging agents. In the presence of MMS, only a small proportion of these cells slowed down S-phase progression; the majority continue to progress through the cell cycle, in contrast to the wild-type (Chabes & Stillman, 2007). A recent study revealed that increased RNR activity promotes DNA replication and that decreased RNR activity leads to slower DNA replication (Poli et al., 2012). Taking the increased DNA replication, the increased sensitivity to DNA-damaging agents, and the inhibition of the DNA damage checkpoint all into account, it is not surprising that elevated dNTP pools produced by RNR hyperactivity can lead to carcinogenesis in some mammalian tissues when combined with other mutations (Xu et al., 2008).

Effects of RNR activity on carcinogenesis in mice

In mice, the R1 subunit of the RNR complex is coded by the gene RRM1 while the R2 subunit can be produced by either RRM2 or RRM2b (p53R2) independently. The RNR complexes consisting of Rrm1 and Rrm2 polypeptides account for most of the RNR activity during S phase. The p53R2 gene was originally identified as a target of the tumor suppressor protein p53 and is transcribed at increased levels following DNA damage (Tanaka et al., 2000). In addition, p53R2 is transcribed at low levels throughout the cell cycle and serves to produce dNTP molecules for mitochondrial DNA synthesis when complexed with Rrm1 (Thelander, 2007). Transgenic mice carrying mutant alleles for either Rrm2Tg or p53R2Tg overexpress the respective R2 polypeptide.

Rrm2Tg mice and p53R2Tg mice both exhibit greater incidence of lung neoplasms (tumors), and expression of either mutant gene induces mutagenesis in cultured 3TC mouse fibroblasts. This suggests that overexpression of the small RNR subunit, R2, promotes carcinogenesis in mammals. In contrast, overexpression of Rrm1 did not lead to significantly increased lung carcinogenesis, suggesting that lung tumor induction might be specific to the R2 subunit and independent of R1-dependent RNR activity in single-mutant mice. It was also revealed that the deletion of either of two DNA mismatch repair (MMR) genes, MSH2 and MSH6, work synergistically with the rnr1D57N mutation to increase mutagenesis and carcinogenesis. In addition, mutations affecting the proto-oncogene K-ras were frequently found in RNR-induced neoplasms, suggesting another synergistic interaction (Xu et al., 2008).

Identification of genes that interact with increased dNTP levels in S. cerevisiae

Elevated dNTP pools can interact with multiple genetic pathways in cells, including ones involving genome stability and DNA metabolism, and contribute to carcinogenesis in multi-cellular organisms (Xu et al., 2008). Since many genetic pathways have been conserved in eukaryotic evolution, the study of yeast cells serves as an adequate starting point for testing this hypothesis. Interactions between cellular pathways can be evaluated by creating double mutant strains of yeast and comparing the phenotypes of these cells with the corresponding single mutant strains.

In a recent study aimed at identifying other genetic pathways that interact with elevated dNTP pools to contribute to carcinogenesis, the rnr1D57N allele was combined with the S. cerevisiae non-essential open-reading frame (ORF) deletion library. The growth of each double mutant strain was evaluated and compared to the growth of the corresponding single mutant strain from the deletion library. There were 68 unique gene deletions that led to increased growth and 112 deletions that led to decreased growth when combined with the rnr1D57N mutant allele, however, none of the interactions were synthetically lethal (Surtees, 2012, Personal communication).

Xrs2 and the MRX Complex

One of the gene deletions that leads to increased growth in combination with rnr1D57 is that of XRS2. A long with the proteins Rad50 and Mre11, Xrs2 forms the MRX complex in yeast. This complex is responsible for repairing double-strand breaks (DSB’s) in DNA via homologous recombination (HR) and non-homologous end joining (NHEJ). The genes making up the homologous complex in humans, the MRN complex, are Rad50, Mre11, and Nbs1, respectively (Ghodke & Muniyappa, 2013). These genes were originally identified in S. cerevisiae during a genetic screen for mutations that yield hypersensitivity to DNA-damaging agents (Game & Mortimer, 1974). In humans, hypomorphic mutations of any of the MRN complex also lead to hypersensitivity to ionizing radiation, as well as genome instability and immunodeficiency (Rupnik et al., 2010). Furthermore, Mre11, Rad50, and Xrs2 are all required for activation of Rad53 (CHEK2 in humans), a kinase that is involved in checkpoint signaling in response to DNA damage (Grenon et al., 2001). In order to understand how these three components work together, it is important first to examine the biochemical properties of each component.

The Mre11 polypeptide tends to form a u-shaped homodimer with itself, allowing it to bind to DNA when the complex is repairing a DSB. In the C-terminal half of the peptide, there is a Rad50 binding site flanked by two DNA binding sites. Near the N-terminus, there is a domain to which Nbs1/Xrs2 binds (Hopfner et al., 2001). Five conserved phosphoesterase motifs also located near the N-terminus give the peptide its nuclease capabilities (Williams et al., 2008). Specifically, the protein can act as a ssDNA endonuclease, a 3′->5′ ssDNA exonuclease, and a dsDNA exonuclease (Trujillo & Sung, 2001). The nuclease activity of Mre11 is dependent upon ATP, manganese ions (Mn2+), and Nbs1/Xrs2 (Paull & Gellert, 1999). Based on data from x-ray crystallography, the Mre11 dimer can form two types of complexes with DNA: a "synaptic DNA complex" involving the ends of oligonucleotides comprised mostly of dsDNA or a "branched DNA complex", which involves Y-shaped oligonucleotides comprised of dsDNA and ssDNA, resembling a stalled replication fork (Fig. 1). When binding to dsDNA, Mre11 interacts with the minor groove and bends the molecule to enable its nuclease function (Rupnik et al., 2010; Williams et al., 2008).

Figure 1. Interaction of Mre11 homodimers with DNA. (Rupnik et al., 2010).

The Rad50 component of the MRX complex is a member of the Structural Maintenance of Chromosomes (SMC) family of proteins. It contains a long internal coiled coil that folds back on itself via a "hinge" region and allows the N- and C- termini to link together and form an ATP-binding cassette (ABC) ATPase domain, which can be found in many other proteins (Holland & Blight, 1999; Hopfner et al., 2002). Within this domain are two motifs: Walker A at the N-terminus, and Walker B at the C-terminus. These motifs are required for ATPase activity and DNA binding (Chen et al., 2005). Near the ATPase domain is a region to which Mre11 binds. On the other end of the folded peptide lies the hinge region, which contains a zinc hook composed of a Cysteine-X-X-Cysteine (CXXC) motif. This motif allows two Rad50 peptides to dimerize (Fig. 2) (Hopfner et al., 2002; Rupnik et al., 2010). Additionally, Rad50 has the ability to transfer phosphate groups from ATP to AMP, forming ADP nucleotides (Bhaskara et al., 2007). More notably, when two Rad50 peptides are bound with Mre11 at two different ends of DNA, they can hook together and essentially serve as a tether to mediate the repair of a DSB (Rupnik et al., 2010).

Figure 2. Schematic showing dimerization of Rad50. The red circles represent the Walker motifs. (Rupnik et al., 2010).

The remaining member of the MRX complex is Xrs2/Nbs1, known as nibrin in humans. Xrs2 is a DNA-binding peptide that does not have any known enzymatic activity but is important for assembling Mre11 and Rad50 and recruiting Tel1 at broken DNA ends. Mre11 and Tel1 bind at two different motifs near the C-terminus of Xrs2 (Rupnik et al., 2010). Tel1 initiates DNA damage responses and is the S. cerevisiae equivalent of the mammalian ataxia telangiectasia mutated (ATM) kinase (Nakada et al., 2003). The role that Xrs2 plays in DSB repair differs between HR and NHEJ. A region of Xrs2 near the N-terminus called the forkhead-associated (FHA) domain is involved in NHEJ but not HR. Through the FHA domain, Xrs2 is able to associate with Lif1, a part of the ligase IV complex that serves a role in the rejoining step of DSB repair. Lif1 contains two Xrs2-binding domains, and phosphorylation of serine 383 of Lif1 is recognized by the Xrs2 FHA domain, specifically. This interaction recruits the ligase IV complex and promotes NHEJ. In humans, the protein Xrcc4, a homolog of Lif1, associates with the FHA domain of Nbs1, suggesting that this general mechanism for NHEJ is conserved among eukaryotes (Matsuzaki et al., 2008).

In yeast, the MRX complex is not essential for survival and is not required for cell viability. In contrast, lack of any of the three components of the MRN complex in mammals causes embryonic lethality, indicating that the complex is essential for the survival of many organisms (Rupnik et al., 2010). The complex is implicated in multiple functions in eukaryotic cells besides its main role in DSB repair. These other functions include telomere maintenance, checkpoint signaling, meiotic recombination, and response to stalled replication forks and DNA hairpins during DNA replication (Borde, 2007; Slijepcevic, 2006; Williams et al., 2007). The role of the complex in meiotic recombination was first identified in S. cerevisiae and is known to be conserved in the model organism species Schizosaccharomyces pombe, Coprinopsis cinerea (Coprinus cinereus), Caenorhabditis elegans and Arabidopsis thaliana. It is very likely that this role of MRN is also present in mammals (Borde, 2007). Humans with hypomorphic mutations in MRE11 or NBS1 are able to survive past the embryonic stage but suffer from ataxia-telangiectasia-like disorder (ATLD) or Nijmegen breakage syndrome, respectively, and display genome instability (Concannon & Gatti, 1993; Taylor et al., 2004). The latter of the two diseases also has an established link to increased cancer risk (Concannon & Gatti, 1993). Results from multiple studies have demonstrated that the nuclease activity of the complex is required only for DNA repair and not for checkpoint signaling nor for telomere metabolism (Buis et al., 2008; Williams et al., 2008). The most crucial purpose of the complex is likely for repairing spontaneous DSB’s that occur during DNA replication in rapid-cycling embryonic cells (Rupnik et al., 2010). This would also explain why the complex is less important in yeast, which are single-celled and do not rely on the proper development of multiple cells.

Effects of XRS2 deletion (xrs2Δ)

In yeast cells, cell cycle arrest in response to DNA damage relies on the protein kinase Rad53, which is itself activated by phosphorylation. When yeast are exposed to the DNA-damaging agent phleomycin, cells with the xrs2Δ deletion exhibit a partial decrease in Rad53 phosphorylation when compared to the wild-type strain (Nakada et al., 2004). Cellular responses to DSB’s involve many other key proteins. The kinase Mec1 (ATR in humans) forms a complex with Ddc2 (Lcd1) and localizes to sites of DNA damage, specifically sites of persistent ssDNA (Fig. 3). Rad24 forms a separate complex and recruits the sliding clamp complex Ddc1-Mec3-Rad17 to DNA lesions. The Mec1 and Ddc1 complexes are independently targeted to sites of DNA damage, however, activation of the Mec1 signaling pathway is dependent upon the Rad24 and Ddc1 complexes. Furthermore, phosphorylation of Rad53 is dependent on the Mec1, Ddc1, and Rad24 complexes. The Mec1 association with DSB’s is partially decreased in xrs2Δ yeast cells. In contrast, the association of Ddc1 with DSB’s is not affected by the deletion (Nakada et al., 2004).

The pathways previously discussed ultimately influence Rad53 activity. When Rad53 is phosphorylated, it is able to facilitate the phosphorylation of Exonuclease 1 (Exo1), an enzyme that possesses 5′ to 3′ double-stranded DNA (dsDNA) exonuclease activity and plays an important role in DNA repair and HR. This nuclease is inhibited upon phosphorylation. This inhibition is important because it limits the accumulation of ssDNA (Mimitou & Symington, 2009). In addition, Exo1 and the MRX complex are both required for Rad53 phosphorylation, meaning that Rad53 and Exo1 each influence the activity of the other. Deletion of the EXO1 gene does not decrease the association of Mec1 with DSB sites. However, Mec1 association is undetectable in xrs2Δ exo1Δ double mutants. When exposed to HU, exo1Δ single mutant cells display similar checkpoint responses to wild-type cells. Although xrs2Δ single mutant cells show defects in blocking mitotic entry following HU exposure, xrs2Δ exo1Δ double mutants were more defective (Nakada et al., 2004). In summary, a cell’s ability to detect DNA damage and arrest the cell cycle via the Mec1 pathways is dependent upon the activity of Exo1 and Xrs2 as well as the MRX complex as a whole.

http://ars.els-cdn.com/content/image/1-s2.0-S0968000409000681-gr3.jpg

Figure 3. Molecular response to a broken end of dsDNA. The MRX complex recruits Tel1 to the damaged DNA via its association with Xrs2. Cdc28 phosphorylates Sae2, which initiates resection and causes MRX and Tel1 to dissociate with the DNA. Replication protein A (RPA) peptides bind to the single-stranded DNA. Next, Exonuclease 1 (Exo1) degrades a strand from 5’ to 3’, leading to activation of the damage checkpoint pathway. Mec1, Ddc2, and the Rad24 complex associate with the site followed by the 9-1-1 complex, which is loaded by the Rad24 complex. Rad9 mediates the autophosphorylation of Rad53, activating Rad53. The activated Rad53 then facilitates the phosphorylation and inactivation of Exo1. The addition of a small, red circle to a factor indicates phosphorylation. (Mimitou & Symington, 2009).

On another note, the xrs2Δ deletion slows 5’-3’ degradation at DNA ends that is an important step in DSB repair. When combined with exo1Δ, this degradation defect is further enhanced, although some residual degradation still occurs. Another notable effect of the xrs2Δ mutation is that it leads to a defect in the cells’ ability to block mitotic entry following exposure to HU, which is used as a drug therapy to slow and stop the growth of cancer cells, while mitotic entry is partially delayed in the absence of HU (Nakada et al., 2004). Since HU inhibits cell growth mainly by inhibiting RNR, one would expect that addition of an RNR mutation that counteracts this inhibition to xrs2Δ would lead to an even greater incidence of mitotic entry compared with the xrs2Δ mutation alone. In contrast, the xrs2 null mutation leads to a shortening of telomere lengths which would likely correspond with cellular senescence and decreased cell cycle progression (Shima et al., 2005).

Xrs2 and dNTP levels

The decreased vegetative growth of yeast with xrs2 null alleles is likely due to increased cell cycle arrest in response to increased DSB events. When combined with mutations that can inhibit cell cycle arrest, such as rnr1D57N, this decrease in growth could potentially be reversed. Research conducted by Surtees has suggested that rnr1D47N xrs2Δ double mutant yeast do exhibit increased growth relative to xrs2Δ single mutant yeast (Surtees, 2012). If this result is confirmed, then one can hypothesize that XRS2, when mutated the presence of altered dNTPs, may lead to enhanced growth rates in mammalian cells as well and therefore have implications for cancer research.

Research Plan

Specific Aims

Deoxynucleoside triphosphates (dNTPs) are the "building blocks" of DNA and are essential for DNA replication and repair. Levels of dNTPs must be strictly regulated. When dNTP levels are too low, DNA replication is slowed down, generating single-stranded DNA (ssDNA), which increases susceptibility to chromosome breakage (Cha & Kleckner, 2002). On the other hand, elevated dNTP levels lead to mutagenesis, accelerated DNA replication, and inhibition of cell cycle arrest (Chabes & Stillman, 2007; Chabes et al., 2003; Poli et al., 2012). These cellular phenomena are frequently associated with carcinogenesis and tumorigenesis. The levels of dNTPs in a cell are dependent upon the activity of the enzyme ribonucleotide reductase (RNR), which catalyzes the rate-limiting step of dNTP synthesis. Saccharomyces cerevisiae (baker’s yeast) containing the mutation rnr1D57N exhibit increased levels of dNTPs (Chabes et al., 2003). Dr. Jennifer Surtees at the University of Buffalo conducted a screen to identify individual genes in the yeast genome that, when deleted and then combined with the rnr1D57N mutation, cause increased or decreased growth relative to the respective single mutant deletion strain. One of the genes that yielded increased growth was XRS2. The protein Xrs2 is a component of the MRX complex, which repairs double-strand breaks (DSBs) in DNA. Based on the results in the screen, I can hypothesize that XRS2, when mutated the presence of altered dNTPs, leads to enhanced growth rates in mammalian cells as well and therefore have implications for cancer research. The overall goal of this project is to confirm the results from the screen regarding XRS2 and to characterize the interaction of Xrs2 with elevated dNTP pools.

Aim 1: Construct xrs2Δ and rnr1D57N xrs2Δ mutant yeast strains and confirm their growth phenotypes.

Starting with the wild-type yeast strain W4069-4C MATα RNR1 CAN1, I will construct a strain of Saccharomyces cerevisiae in which a KanMX module is inserted within the XRS2 gene, preventing transcription mRNA coding for Xrs2. These xrs2Δ single mutant yeast will be selected via plating on media containing Geneticin, to which the KanMX module confers resistance. A yeast strain that contains the rnr1D57N mutation (W4069-8C MATα rnr1-D57N CAN1) will undergo the same deletion process. The new rnr1D57N xrs2Δ double mutant yeast will be selected and plated. Dr. Surtees has both xrs2Δ single mutant and rnr1D57N xrs2Δ double mutant yeast, however these are not of the W4069-4C strain with which I will be working. Therefore, it is necessary to obtain a KanMX deletion cassette for XRS2 and perform the deletion myself in our laboratory. Once all of the specific strains have been produced, their phenotypes can then be studied.

The growth of the xrs2Δ single mutant cells and the rnr1D57N xrs2Δ double mutant cells will then be evaluated. This will be accomplished by growing each strain in SC-LEU+2% glucose broth and then measuring growth using a spectrophotometer. The absorbance data produced from the machine will be used to construct growth curves.

Aim 2: Confirm increased dNTP levels in rnr1D57N single mutant and double mutant strains relative to the wild type strain.

The yeast mutant strain rnr1D57N is useful for evaluating the effects of elevated dNTP levels on cellular pathways and processes that can lead to carcinogenesis. As a result of this mutation, the 57th amino acid residue of the Rnr1 polypeptide is changed from aspartic acid to asparagine. The Rnr1 polypeptide contains an allosteric site for dATP inhibition. This particular mutation directly affects the dATP binding site and causes the RNR enzymes in the mutant strain to be resistant to feedback inhibition. Assays have confirmed that W1588-4C yeast with this mutation does in fact have greater dNTP pools when compared with wild-type yeast (Chabes et al., 2003).

I expect that my W4069-4C rnr1D57N single mutant and rnr1D57N xrs2Δ double mutant yeast will exhibit increased dNTP levels when compared with the JSY13 wild-type yeast and the xrs2Δ single mutant yeast. If there are significant differences in dNTP levels between the rnr1D57N single mutant and the rnr1D57N xrs2Δ double mutant strains or between the wild-type and xrs2Δ single mutant strains, then I would investigate the cellular mechanism by which the Xrs2 protein affects dNTP pools. This is definitely a possibility since Xrs2 plays a role in DSB repair, and the presence of damaged DNA leads to increased dNTP pools (Chabes et al., 2003).

Aim 3: To determine how elevated dNTP pools rescue growth of yeast with the xrs2Δ deletion and then determine whether rnr1D57N xrs2Δ yeast exhibit different mutation rates compared with both single mutant strains

If I confirm Dr. Surtees’ result that increased dNTP levels lead to increased growth in rnr1D57N xrs2Δ double mutant yeast relative to xrs2Δ single mutant yeast, then I will investigate this interaction in order to determine how growth becomes rescued. Previous research has revealed that elevated dNTP pools inhibit cell cycle arrest at the DNA-damage checkpoint and allow cells that have been exposed to certain DNA-damaging agents to progress through the cell cycle instead (Chabes & Stillman, 2007). I hypothesize that elevated dNTP pools have a similar effect on cells that contain unrepaired DSBs due to the xrs2Δ mutation and are the reason for the enhanced growth observed in rnr1D57N xrs2Δ yeast compared with xrs2Δ single mutant yeast. I aim to investigate this hypothesis and also determine if the xrs2Δ deletion enhances growth in the presence of elevated dNTP pools.

I plan on characterizing the rnr1D57N xrs2Δ yeast in specific detail. In order to do this, I will evaluate the cell cycle progression for each mutant strain via microscopy. In addition, I will measure mutation rates for each mutant strain using a canavanine assay. Lastly, a single cell gel electrophoresis assay ("comet assay") will be used to measure DNA strand breaks in the cells. Since it is known that elevated dNTP pools are mutagenic, it would be interesting to see how mutation rates are affected when the xrs2Δ deletion is added in and then make connections to potential pathways that may lead to carcinogenesis. It is logical to look specifically at DNA breaks as well since the Xrs2 is involved with DSB repair.

Research Design and Methods

Evaluating and comparing growth phenotypes

I will measure the growth phenotypes of each strain in order to confirm or refute Dr. Surtees’ result that increased dNTP levels lead to increased growth in rnr1D57N xrs2Δ double mutant yeast relative to xrs2Δ single mutant yeast. Using a sterile loop, I will pick an individual colony of yeast from a plate and inoculate it in 3 mL of SC-LEU+2% glucose broth in a test tube. The tube will be shaken at 30°C until the broth is saturated. A second tube, also containing SC-LEU+2% glucose broth, will be inoculated with 0.1 mL of broth from the first tube and incubated overnight. One mL of the broth will then be taken and used to record the optical density (OD) at 595 nm using a spectrophotometer as necessary. When the OD595 of the second tube is at 0.35, 0.1 mL of it will be inoculated into 10 mL of SC-LEU+2% glucose broth in a third tube. Approximately every 6 hours after inoculation, 1.0 mL will be removed and used to measure the OD595. Recordings will continue to be taken until the OD exceeds 1.0. I will then plot growth curves for visual comparison and also calculate doubling times for statistical comparison.

Performing the targeted gene deletion of XRS2

Using xrs2Δ yeast and the appropriate primers from Dr. Surtees, I will amplify the XRS2:KanMX deletion cassette. Copies of this cassette will then be transformed into our W4069-4C MATα RNR1 CAN1 yeast.

A standard lithium acetate transformation will be conducted according to Gietz & Woods. First, this involves inoculating the yeast (to be transformed) into liquid medium and then incubating the cells overnight. Once the cell titer is at an adequate level, the cells will be harvested via centrifugation and then resuspended in water. The cells will then be centrifuged again, and the supernatant will be discarded. A "Transformation Mix" will also be prepared according to Table 1.

Table 1. Components and volumes of the Transformation Mix.

Number of Transformations

 Reagents

  1

5 (6X)

10 (11X)

PEG 3500 50% w/v

 240 µl

 1440 µl

2640 µl

 LiAc 1.0 M

 36 µl

216 µl

396 µl

 Boiled SS-carrier DNA

 50 µl

300 µl

 550 µl

 Plasmid DNA plus Water

 34 µl

204 µl

374 µl

 Total

 360 µl

2160 µl

3960 µl

Transformation Mix will be added to each tube, and the cells will be resuspended in the liquid via vortexing. After a subsequent 40-min heat-shock incubation at 42°C, the cells will be separated from the Transformation Mix and resuspended in water. Lastly, the cells will be plated onto synthetic complete (SC) selection medium containing Geneticin and incubated for 3-4 days.

Colonies of cells that are not killed by the Geneticin are assumed to contain the KanMX4 module. In order to verify that the deletion cassette was recombined at the locus of the YDR369C ORF, DNA will be extracted from some of the cells, and PCR will be conducted using four YDR369C-specific "confirmation primers" (Fig. 4). The sequences for these primers are as follows: A primer: 5’GTATTGAAGCAATTTGTAAGCTGGT3’, B primer: 5’CGAACTATTATTGATTTCCCATTTG3’, C primer: 5’ATTTCAAGACTTTTGTCAAGGTACG3’, and D primer: 5’TCCAATTTTAAGATTTTCACTCTGC3’. If this PCR reaction produces a product, meaning that the YDR369C ORF is still present, then the deletion was not successful. If two different PCR reactions using the A primer and the KanB primer (5'-CTGCAGCGAGGAGCCGTAAT-3') and using the KanC primer (5'-TGATTTTGATGACGAGCGTAAT-3') and the D primer both produce a products, then the deletion was successful, and the KanMX4 module replaced the YDR369C ORF.

Figure 4. PCR-based confirmation process. If the PCR reaction in (a) produces a product, meaning that the YDR369C ORF is still present, then the deletion was not successful. If both PCR reactions in (b) produce products, meaning that the KanMX4 module replaced the YDR369C ORF, then the deletion was successful. ("Yeast Deletion Project," 2012)

Assay for testing dNTP levels adapted from Wilson et al. (2011)

I will determine whether or not my W4069-4C rnr1D57N single mutant and rnr1D57N xrs2Δ double mutant yeast will exhibit increased dNTP levels when compared with the JSY13 wild-type yeast and the xrs2Δ single mutant yeast. To begin, a set number of yeast cells will be added to ice-cold 60% methanol, vortexed vigorously, and then incubated at 95°C for 3 min. These cells will then be sonicated for 30 seconds and then centrifuged at 16,000g for 5 min at 4°C to remove cell debris, precipitated protein, and DNA. The supernatant will then be passed through a pre-equilibrated Amicon Ultra-0.5-mL centrifugal filter at 4°C to remove macromolecules (using the manufacturer’s directions). The filtrate will be evaporated under centrifugal vacuum at 70°C, and the pellet will be resuspended in nuclease-free water and either used for the assay immediately or stored at -80°C until needed.

In preparation for the real-time PCR portion of the essay, the volumes of the following reagents will be acquired and thawed on ice:

1 NDP1 primer (10 µmol/L)

1 µL dTTP-DT6 template (10 µmol/L)

1 µL FAM-dTTP probe (10 µmol/L)

2 µL MgCl2 (25 mmol/L)

1 µL dNTPs WITHOUT dTTP (2.5 mmol/L)

2.5 µL of GeneAmp 10X PCR Buffer II

0.175 µL of AmpliTaq Gold Polymerase (5U/µl)

14.825 µL nuclease-free H2O

22.5 µL

Unlike the other reagents, the AmpliTaq polymerase will be kept at -20°C until immediately prior to use. This set of reagents tests the levels of dTTP specifically. If I wish to evaluate the levels of a different dNTP, then a different template, probe, and dNTP mix will need to be used. These are listed in Table 1 of the Wilson et al. (2011) article.

I will aliquot the 22.5 µL of reagent mix into a PCR tube and then add 2.5 µL cell extract. An additional tube will be prepared with dNTP standard added in place of cell extract. Each tube will then be pulse-spinned at approximately 3000g. The tubes will then be transferred to a real-time PCR thermocycler. The machine will be programmed to detect the 6-FAM fluorophore and to perform the following three steps: 10 min at 95°C, 5 min at 60°C, and a fluorescence reading. A minimum of 8 cycles of these steps will be conducted. Lastly, the output from the machine will be analyzed to determine dTTP levels from the cells. By subtracting the fluorescence units for the negative control or "zero pmole" standard from all of the experimental values, I will produce normalized fluorescence units (NFUs). The NFUs from the standard will be plotted to create a calibration curve, which will be used to calculate concentrations for the cell extract-containing tube.

Assay for analyzing cell cycle progression

In order to characterize the rnr1D57N xrs2Δ yeast in specific detail, I will evaluate the cell cycle progression for each mutant strain via microscopy and compare the results with those of the single mutant strains. Approximately 1 mL of log phase (OD = 0.5-1.0) yeast cells cultured in YPD will be removed and transferred into an Eppendorf tube. The tube will be briefly vortexed to ensure suspension of cells. Approximately 30 microliters of the culture will be placed upon a microscope slide and covered with a glass cover slip. Using a light microscope, I will check to see if the cells are clumped up and need to be sonicated. Once a slide containing non-clumped cells has been acquired, I will set up an oil immersion with an appropriate light microscope and find a field of view in which at least 30 cells can be seen. Using Figure 2, the numbers of cells in G1 phase, S phase, and G2/M phase will be counted and converted to percentage values.

Canavanine assay for analyzing mutation rates

I will also evaluate mutation rates in order to characterize the rnr1D57N xrs2Δ yeast relative to the single mutant strains. For each unique strain, 7 colonies will be separately inoculated in liquid SC medium and incubated for approximately 20 hrs in a 30°C shaker. Using sterile dH2O, three dilutions of each culture will be prepared in separate Eppendorf tubes: 10^-2, 10^-3, and 10^-4. Next, 100 microliters of each 10^-4 diluted culture will be inoculated in liquid SC medium and another 100 microliters of each will be plated onto solid SC medium. The total cell numbers will be estimated by measuring the optical density of each liquid culture. A designated volume around 500 microliters will be taken from each undiluted culture, washed once with dH2O, resuspended in 100 microliters of dH2O, and then plated on ArgDO+Canavanine solid medium. For each of the other dilutions, 100 microliters should be removed and immediately plated onto ArgDO+Canavanine solid medium. All of the plates will then be incubated for approximately 3 days at 30°C. After this period, colonies will be counted and the median frequency of CanR among the 7 colonies will be calculated for each dilution. Each CanR value can be calculated by dividing the number of colonies grown on the canavanine plate for a given colony by the number of colonies grown on the regular SC plates for the same colony.

Comet assay adapted from Azevedo et al. (2011)

The final technique will allow me to measure DNA strand breaks in the cells. This is a logical assay to conduct since the Xrs2 is involved with DSB repair. To begin, I will create a culture of yeast in YPD medium and incubate it at 30°C using a 500 or 50 mL Erlenmeyer flask with an air:liquid ratio of 10:1 on a mechanical shaker set to 200 rpm. A liquid preculture of 5-10 mL YPD will be inoculated with a small amount of yeast cells and incubated overnight. Cells will be suspended in fresh medium to a density of 1.2*10^7 cell/mL. The cell will be harvested after two generations by centrifugation (2 min at 4500g, 4°C), washed twice with the same volume of ice-cold dH2O, and diluted back to the same concentration in ice-cold S-buffer (1 M sorbitol, 25 mM KH2PO4, pH 6.5).

Aliquots of the suspension with approximately 10^6 cells will be harvested via centrifugation (2 min, 18000g, 4°C) and mixed with 1.5% w/v low-melting agarose in S buffer) containing approximately 2 mg/ml zymolyase (20T; 20000 U/g). Eighty microliters of this mixture will be spread over an agarose-coated slide (i.e. a slide coated with a water solution of 0.5% w/v normal-melting agarose), covered with a cover slip, and incubated for 20 min at 30°C, after which the cover slips will be removed. All further procedures will be performed in a 4°C refrigerated chamber. Slide(s) will be incubated in lysis solution (30 mM NaOH, 1 M NaCl, 0.05% w/v laurylsarcosine, 50 mM EDTA acid, 10 mM Tris-HCl, pH 10) for 20 min. The slide(s) will be rinsed three times for 20 min each in electrophoresis buffer (30 mM NaOH, 10 mM EDTA, 10 mM Tris-HCl, pH 10) to remove lysis solution. The samples will then be submitted to electrophoresis in the same buffer for 10 min at 0.7 V/cm. After electrophoresis, the slide(s) will be incubated in neutralization buffer (10 mM Tris-HCl, pH 7.4) for 10 min, followed by consecutive 10 min incubation in 76% and 96% ethanol. The slide(s) will then be air-dried and visualized immediately or stored at 4°C. For visualization, the slide(s) will be stained with ethidium bromide (10 micrograms/mL) and images will be acquired using a fluorescent microscope at a magnification of 400x. The images will be analyzed using the free edition of CometScore software, and the analytic parameter tail length (in microns) will serve as the measurement of DNA damage.

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