Tumor hypoxia and genetic alterations in sporadic cancers

Abstract

Cancer is a disease of a cell that gains the ability to multiply in an uncontrolled way, to invade from the primary site to surrounding tissues, and to metastasize to distant sites. Throughout the past three decades, the field of cancer genetics has identified critical genes and the pathways1 whose dysfunction leads to major cancer phenotypes: self-sufficiency in growth signals, insensitivity to anti-growth signals, evading apoptosis, limitless replicative potential, sustained angiogenesis, tissue invasion and metastasis.2 Thus, cancer is a multi-genetic disease and exhibits a progressive process where the genetic or epigenetic alterations responsible for these phenotypes accumulate in time and space. To date, about 350 cancer genes have been identified.3 Results of recent systematic DNA sequencing of the cancer genome have shown the following characteristics. There are two types of mutations in cancer cells: ‘driver’ and ‘passenger’. Driver mutations contribute to tumor cell growth and survival under restricted conditions and are positively selected during the course of cancer development. The rest of the mutations are ‘passenger’ mutations, which have not contributed to cancer development or been positively or negatively selected. Most of the mutations found in the cancer genome are passengers. The frequency of mutation in each driver is low, suggesting that the number of ‘drivers’ in common adult sporadic cancers could be greater than the 5–7 which have been estimated by age-incidence statistics. There is heterogeneity in the number of gene mutations among cancers; some genomes have >100 000 point mutations whereas others have fewer than 1000.3 This suggests that more than 99% of mutations in the cancer genome do not contribute to the carcinogenic process. So how do cancer cells gain such excessive numbers of mutations? There are three types of cancer genes: oncogenes, tumor suppressor genes and stability genes.1 Oncogenes encode proteins that promote cell multiplication and survival. Their expression or functions are activated by point gene mutation, fusion to another gene by chromosomal translocation and/or gene amplification. About 90% of cancer genes are dominant-acting oncogenes.3 Tumor suppressor genes encode proteins that inhibit cell multiplication and promote cell death. Inactivation of tumor suppressor genes is achieved by point mutation, gene deletion or insertion, or by epigenetic silencing. Activation of oncogenes or inactivation of tumor suppressor genes confers cell growth and gives the cancer cell a survival advantage. On the other hand, stability genes encode proteins whose loss or over-expression increases genetic alterations all over the genome. Stability genes include DNA repair genes, DNA damage sensor genes and cell cycle checkpoint genes. Malfunction of stability genes could be the driving force of the carcinogenic process.4–6 Alternatively they may not be necessary for carcinogenesis, but may merely promote this process.7 This topic is one of issues that will be discussed in this review. Most solid tumor tissues, even when they are microscopically small, contain acute and chronic hypoxic and/or anoxic areas where oxygen pressure is lower than is physiologically normal.8,9 As an adaptive response to the lack of oxygen, cancer cells may change their genome to increase their survival. In 1996, Glazer's group first presented evidence that the tumor microenvironment, especially hypoxia, induces high levels of gene mutations in cancer cells. This study was based on their hypothesis that ‘the microenvironment may give conditions that either increase DNA damage or compromise the DNA repair process’.10 Since then, this hypothesis has been tested by many research groups.11 The results of these studies generated a new concept that the microenvironment (hypoxia) induces genetic instability.12 This hypothesis accepts the idea of ‘genetic instability as a hallmark of cancer’; however, the extension of the hypothesis does not necessarily require the idea that cancer, especially sporadic cancer, gains gene mutations in putative stability genes that may drive the carcinogenic process.4–6 If hypoxia (the microenvironment) plays a role for the carcinogenic process, it should contribute to the progression stage of tumor development when a part of the tumor tissue starts to receive less oxygen. In fact, many clinical and experimental observations indicate that hypoxia is associated with the aggressiveness of tumor cells, leading to poor prognosis and metastasis in a variety of human cancers. Within tumor tissues, oxygen concentrations fluctuate both spatially and temporally. Hypoxic tumor cells may be re-exposed by a higher concentration of oxygen (re-oxygenation), which can alter the cancer genome and contribute to tumor progression. In this review, mechanisms by which hypoxia and re-oxygenation induce genetic alterations in sporadic cancer will be considered. Toward this goal, literature relating to tumor hypoxia, cellular pathways affected by hypoxia, types of genetic alterations and DNA repair systems affected by hypoxia and re-oxygenation has been compiled. The impact of hypoxia on human cancer in medicine was first recognized by radiologists. In the 1930s, the presence of hypoxia in solid tumor tissues was first hypothesized based on the observation that low levels of oxygen (hypoxia) protect a cell from the lethal effects of ionizing radiation and that some solid tumors are resistant to radiation.13 In 1955, Thomlinson and Gray reported histological observations of tumor cords with and without central necrosis in human lung tumors, suggesting the presence of an oxygen gradient within a tumor cord. They found that: (i) all of the tumor cords surrounded by the stroma and >200 µm in radius contained central necrosis; (ii) none of the tumor cords <160 µm in radius contained central necrosis; and (iii) no intact tumor cells were found at a distant of 180 µm from the stroma. Based on these results and the calculated distance of oxygen diffusion (150 µm), they proposed the presence of radio-resistant hypoxic cells at the edge of the necrotic area.14 Until the late 1980s when polarographic electrodes were used to directly measure levels of oxygen in human cancer tissues, the presence of tumor hypoxia was speculative.15,16 During the 1990s, several key findings were made using various methods for directly detecting tumor hypoxia in human tumor tissues.9,15 These findings are as follows: (i) hypoxic and anoxic areas exist in most solid tumors (areas with <2.5 mmHg of oxygen pressure); (ii) there is no predictable association between tumor hypoxia and other clinical factors, including size, stage, grade and site; (iii) tumor hypoxia may be an adverse prognostic factor;9,17 and (iv) tumor hypoxia not only induces radiation-resistance, but it may also induce resistance to chemotherapeutic agents.9,18 Using DNA-binding chemical Hoechst 33 432, cell sorting and radiation, Chaplin et al. first demonstrated that two types of hypoxia exist in solid tumor tissues.19 Because of the abnormal structure, distribution and function of microvessels formed by unregulated angiogenesis within tumor tissues, an inadequate blood flow occurs in geometrically different parts of tumor tissues. This type of hypoxia, called acute hypoxia, lasts from minutes to hours and is followed by re-oxygenation.16,19 Another type of hypoxia is caused by the reduction of oxygen diffusion due to an increase in the distance of the tumor cells from the tumor or host vasculature. This type of hypoxia is called diffusion-limited hypoxia or chronic hypoxia. It may last days, followed by re-oxygenation or cell death.16 It has been suggested that a different biology may exist between acute and chronic hypoxia and this might influence the interpretation of clinical and experimental data, and the design of treatments for hypoxic tumors.20 While struggling to overcome the radiation-resistance of hypoxic tumors, many aspects of the cellular response to hypoxia have been recognized and studied. These hypoxic responses are related to angiogenesis, glycolysis, metastasis, stress response, erythropoiesis and genomic stability.20,21 Hypoxia-inducible factors (HIFs) play a central role in these responses to hypoxia. In 1995, Wang et al. identified one of the HIFs, HIF1, a complex between HIF1α and HIFβ subunits, which is stabilized in response to hypoxia and regulates transcription of its target down-stream genes.22 HIF1 binds to the hypoxia response elements (HREs), 5′-G/ACGTG-3′, in the promoter region of target genes, such as EPO,23VEGF,24Aldolase, Enolase and LDHA.25 Currently, transcription of at least 70 known genes, and probably more, is regulated by HIFs through recognition of HREs.26 There are three HIFα family subunits, HIF1α, HIF2α and HIF3α, and they form a heteroduplex with a common constitutive HIFβ subunit. Both the HIF1 and HIF2 heteroduplexes function as transcription factors for genes containing HREs under hypoxia. HIF1α and HIF2α, but not the HIFβ subunits, are rapidly degraded by the ubiquitin–protease pathway in normoxic conditions through oxygen-dependent degradation domain.27 A tumor suppressor protein, von Hippel-Lindau (VHL), binds to HIFα subunits and promotes oxygen-dependent degradation of HIF.28 VHL is a part of the E3 ubiquitin ligase complex and binds directly to HIFα subunits and a ubiquitinates the subunits.29 The binding between the VHL and HIFα subunits is regulated through hydroxylation of a proline residue within HIFα subunits by the family of prolyl hydroxylases (PHDs or HPHs).30,31 Because the enzyme activity of PHDs requires oxygen and iron, the lack of oxygen or iron in a cell leads to the accumulation of HIFs. Another oxygen- and iron-sensitive enzyme, FIH1 (factor inhibiting HIF1), which catalyzes hydroxylation of asparagine residue on HIFα subunits, inhibits the interaction of HIFα subunits and their transcription co-activators, such as p300/CREB. Hypoxia impairs FIH1 activity, which results in formation of a HIF1/CBP/p300 complex and leads to enhanced transcription of HIF target genes.32 In sustained hypoxia (chronic hypoxia), HIF activity is attenuated by the following negative feedback mechanisms: (i) HIF up-regulates CITED2 (transcription of a CBP/p300-interacting transactivator with a Glu/Asp-rich carboxy-terminal domain), which binds to CBP/p300 and blocks interaction between HIF and CBP/p300 and the transactivation of HIFs in hypoxic cells;33 (ii) HIF hydroxylase levels are up-regulated by PHD activation, leading to destruction of HIF even if oxygen levels are low;34 and (iii) antisense RNA against HIF1α is transcribed from the HIF1 locus in an HRE-dependent manner.34 A major mechanism for a cell to adapt to hypoxia is by using the HIF pathway that activates target pathways regulating the delivery of oxygen and its utility. However, as can be seen below, HIF1 also directly or indirectly regulates the expression of other genes involved in stability of the cellular genome. There are two other cellular signaling pathways in response to hypoxia. These include the mammalian target of rapamycin (mTOR) pathway and the endoplasmic reticulum stress pathway. Repression of the mTOR pathway and activation of the endoplasmic reticulum stress pathway by hypoxia regulates protein synthesis through inhibition of mRNA translation.35 Although there have been only a few studies reporting the involvement of these pathways in the stability of the cellular genome, it is worthwhile to briefly review these pathways. The mTOR is a Ser/Thr protein kinase and forms mTOR complex 1 (mTORC1) with Raptor and GβL. Raptor is a scaffolding protein that mediates interaction between mTOR kinase and its substrates to promote mTOR signaling. GβL plays a role in stabilizing mTOR and Raptor binding. When cells are under nutrient- and energy-replete conditions, the mTORC1 activates downstream proteins, including ribosomal protein S6 kinase (p70S6K), eukaryotic initiation factor 4E binding protein 1 (4E-BP1) and eukaryotic elongation factor 2 kinase (EEF2K). Phosphorylation of these proteins promotes protein synthesis, cell growth, cell proliferation and cell metabolism.35,36 Chronic hypoxia down-regulates mTORC1 signaling through multiple pathways to maintain cellular protein synthesis levels appropriate for suboptimal conditions. Hypoxia inhibits mTORC1 signaling through the accumulation of the tuberous sclerosis protein 1 and 2 (TSC1-TSC2) complex. TSC1 stabilizes TSC2 by forming a complex with TSC2. TSC2 is a GTPase-activating protein (GAP) and regulates the Ras homolog enriched in brain (RHEB). RHEB activates mTORC1 when it is GTP-bound. Since the TSC1-TSC2 complex promotes conversion of RHEB-GTP to RHEB-GDP, this results in the cessation of mTORC1 activity.36 Accumulation of the TSC1-TSC2 complex is achieved through competitive inhibition of complex formations between 14 and 3-3 and TSC2 by DNA-damage-inducible transcript 4 (DDIT4 or REDD1). REDD1 is up-regulated by HIF1 under hypoxic conditions, binding to 14-3-3, and it dissociates TSC2 from the 14-3-3/TSC2 complex.37–39 Hypoxia also activates the AMP-activated protein kinase (AMPK) pathway. Hypoxic cells switch respiration from the aerobic mitochondrial chain to anaerobic glycolysis to generate adenosine triphosphate (ATP). This results in an increase in the adenosine monophosphate (AMP)/ATP ratio and activates AMPK activity. AMPK phosphorylates and activates GAP in TSC2 leading to inhibition of mTORC1 through a decrease in RHEB-GTP.40 It has been demonstrated that the Bcl2/adenovirus E1B 19-kDa interacting protein 3 (BNIP3), which is up-regulated by HIF1, interacts with RHEB and decreases the level of GTP-bound RHEB. This results in inhibition of mTORC1 activity and subsequent cessation of protein synthesis.41 It has also been reported that the promyelocytic leukemia tumor suppressor (PML) inhibits mTORC1 by binding and transporting it to a nuclear body under hypoxia.42 The endoplasmic reticulum (ER) is a cellular organelle for protein folding and maturing. When a cell faces a number of biochemical, physiologic or pathologic environments, including nutrient depletion, oxidative stress, DNA damage, energy perturbation or hypoxia, the process of protein folding and correct assembly of mature proteins is disrupted in the ER. As a result, unfolded or misfolded proteins accumulate within the ER (termed ‘ER stress’). In response to ER stress, the ER generates signals that alter transcriptional and translational programs that ensure the fidelity of protein folding and maturation, effectively eliminating the unfolded and misfolded proteins, and selectively allowing translation of mRNAs whose products promote the cell's survival under hypoxic conditions. This response is called the unfolded protein response (UPR).36,43 Hypoxia triggers UPR by activating three ER stress sensors, including the inositol-requiring protein 1 (IRE1), activating transcription factor 6 (ATF6) and PKR-like ER kinase (PERK).36,43 The inactive forms of these three proteins are bounded by the chaperone immunoglobulin heavy chain-binding protein (BIP) and embedded in the ER membrane. Unfolded or misfolded proteins activate these sensors by binding to BIP and dissociating BIP from these sensor proteins or by directly binding to the sensors. Activated PERK phosphorylates eukaryotic initiation factor 2 subunit α (EIF2α), resulting in inhibition of global mRNA translation and selective translation of ATF4 and other hypoxia-inducible mRNAs. Activation of IRE1 results in endoribonuclease activity against the X-box-binding protein 1 (XBP1) pre-mRNA and in the selective expression of XBP1. Activation of ATF6 results in its translocation to the Golgi apparatus and its cleavage to gain transcriptional activity. ATF4, XBP1 and ATF6 transactivate genes whose products increase protein folding and maturation in the ER and genes whose products remove unfolded and misfolded proteins from the ER.36,43 Re-oxygenation is a component of hypoxia-induced genetic alterations. In mammalian cells, hypoxia followed by re-oxygenation (H/R) increases the production of reactive oxygen species (ROS) from affected cells,44 which can damage DNA, proteins and lipids, leading to gene mutations, apoptosis and necrosis. Therefore, hypoxic cancer cells have to deal with the toxic effect of ROS; however, if cancer cells have already acquired gene mutations, for instance mutated p53, which overcomes apoptosis signals triggered by H/R,45 these cells have an increased probability of gaining additional mutations. Although ROS can generate various types of modified bases in DNA, 7,8-dihydro-8-oxoguanine (8-oxo-G) is frequently generated.46 For example, the hypoxic human cervical cancer cells, HeLa, placed under 1% oxygen for 24 h, produced excessive amounts of ROS at 30 min after reoxygenation.47 This overproduction of ROS was transient and lasted for 2 h after re-oxygenation. Simultaneously, the same cell population generating ROS also exhibited extensive DNA damage with 8-oxoguanine.47 The 8-oxo-G:C pair, if not repaired, generates G:C > T:A or A:T > C:G transversions. These mutations are frequently found in sporadic human cancers, including lung, breast, ovarian, gastric and colon cancers.48 In in vivo and in vitro hypoxia models, an increase in transversion mutations, such as G:C > T:A and A:T > G:C, has been reported,10 suggesting an important carcinogenic role of ROS generated by H/R in tumor tissues. Reactive oxygen species also induce DNA slippage mutations at microsatellite sequences in human cells. When human lung cancer cells carrying plasmid vector with cytosine-adenine (CA) repeats were treated with ROS generating chemicals, paraquat and H2O2, a significant increase in deletion or insertion mutations was observed within CA repeats.49 Similarly, Gasche et al. showed that the frequency of microsatellite mutations (CA repeats) in transfected plasmids was increased by H2O2 treatment in human colon cancer cells.50 Yamada et al. examined the effect of H2O2 treatment on mutation frequencies of mononucleotide (A or G repeats) and di-nucleotide repeats (CA repeats) in non-cancer human diploid cell lines. They found that H2O2 treatment decreased the mutation frequency of mononucleotide repeats, but increased the mutation frequency of di-nucleotide repeats in non-cancer diploid human cells. They speculated that ROS induces low levels of mutations in di-nucleotide repeats.51 In accordance with the effect of ROS on microsatellite loci in human cells, Chang et al. reported that non-toxic levels of H2O2 impair mismatch repair activity,52 which leads to DNA slippage mutations at microsatellite loci (see below). In order to faithfully transmit genetic information to a progenitor cell, the cell is equipped with mechanisms that sense DNA damage in the genome (sensor), transmit a DNA damage-signal to repair system and cell cycle machinery (signal), and target a cell for apoptosis if damage is not repaired (effector). There is some evidence that H/R activates DNA damage response. Ataxia telangiectasia mutated (ATM) and ataxia telangiectasia and Rad3-related (ATR) are DNA damage signal transducers. A double-strand break is recognized by the sensor protein complex MRN (MRE11-RAD50-NBS1). The sensor recruits ATM, which further activates its targets CHK1/CHK2. A single-strand DNA is sensed by ATRIP (ATR interacting protein) and recruits ATR. ATR also activates CHK1/CHK2. It has been found that acute severe hypoxia (<0.02% O2 for less than 24 h) activates both ATR and ATM without DNA damage.53 It is assumed that the activation of ATR is not transducing DNA damage but directed toward maintaining replication folk stability during severe hypoxia by phosphorylating the replisome components, MCM2 and MCM3.54 However, when cells are re-exposed to oxygen, reactive oxygen species (ROS) are very quickly generated and damage cellular DNA. In response to the damage, ATM is activated and phosphorylates a downstream protein, CHK2.55,56 The activated CHK2 causes G2 cell cycle arrest through phosphorylation of Cdc25C and Cdc2.56 There is a possibility that cancer cells may propagate new genetic alterations caused by reoxygenation-induced ROS if the cells are insensitive to the G2 arrest.54 The concept of ‘genetic instability’ was introduced to define the cancer cells' property of new mutations with each cell division. Using tissue cultured cancer cells, Lengauer and Vogelstein first demonstrated that some, but not all, cancer cells continuously change their chromosome numbers with each cell division.57 They termed this type of genetic instability as chromosome instability (CIN). Later, CIN was extended to characterize persistent changes, not only in the number of whole or part chromosomes (whole chromosome instability, W-CIN), but also changes in the structure of chromosomes (amplification, deletion and translocations: segmental chromosome instability, S-CIN) during the lifetime of cancer cells. Based on CIN observed in tissue cultures, it is assumed that the frequent occurrence of the chromosomal abbreviations observed in human tumor tissues is caused by CIN mechanisms. Great progress in understanding the molecular basis of CIN has been made through the use of experimental in vitro and animal models.58 These studies have shown that W-CIN is caused by failures in the correct transmission of chromosomes into daughter cells or the spindle mitotic checkpoint.57 On the other hand, some inherited conditions, such as ataxia telangiectasis, Bloom syndrome, Fanconi anemia and Nijmegen breakage syndrome, are called chromosome instability syndromes and associated with S-CIN and a predisposition to certain types of cancer. Through identification of the genes responsible for these conditions, it is known that S-CIN is caused by mutations of the genes involved in replication, repair and S-phase checkpoints.59 Before CIN was fully understand, another type of genetic instability, microsatellite instability (MSI or MIN), had been recognized in a small fraction of cancers. MSI is characterized as an expansion or contraction of repeat units within the microsatellite locus. The origin of MSI is thought to be replication mistakes by DNA polymerase at the microsatellite followed by failed mismatch repair.60 Therefore, the main cause of MSI found in human cancers is due to inactivation of the mismatch repair system.61 Recently, an additional form of genetic instability, point mutation instability (PIN), was proposed by Loeb's lab. This is based on their DNA sequencing data that showed that cancer exhibits a 200-fold higher mutation rate than normal at the nucleotide level;62 however, the corresponding mechanism for this type of instability is not known. W-CIN can be induced by the disturbance of the mitotic checkpoint, a mechanism ensuring a faithful segregation of copied chromosomes to a daughter cell, or by abnormalities in spindle and centrosome functions. The experimental evidence using animal models supports this hypothesis. A partial loss of mitotic checkpoint genes, including mad2l1, mad1l1, fzr1, plk4, bub1b, bub3, bub1 and cenpe causes aneuploidy in cells derived from heterozygous mice.58 Over-expression of genes, including mad2 and hec1, also leads to CIN.58 Moreover, these mitotic checkpoint mutant mice are predisposed to various type of cancers.58 The genes responsible for the chromosome instability syndromes mentioned above are AMT, BLM and FANC genes and NBS1; the loss of these gene products in a cell induces S-CIN and a predisposition to cancer.63–66 Germline mutations in BRCA1, BRCA2, PALB2, RAD50 and BRIP1 are found in hereditary forms of breast cancers and linked to S-CIN.67 All these genes are involved in DNA damage checkpoint, cell cycle checkpoint, and homologous and non-homologous recombination repair. However, recent data from cancer genome sequencing has showed that gene mutations in these CIN genes are rare in sporadic human cancers.68 Mutations in other DNA repair genes involved in nucleotide excision repair and mismatch repair (MMR) are also rare in sporadic human cancers.68 Despite the lack of mutations in stability genes, aberrant expression of stability genes has been observed in sporadic human cancers. For example, some mitotic checkpoint gene products, including AURKA, AURKB, MAD2L1, PLK4, BUB1B and BUB3 are over-expressed in various types of human cancers.58 BRCA1 is down-regulated and BRCA2 is up-regulated in sporadic breast cancers.69,70FANC genes are down-regulated in head and neck squamous cell carcinoma.71 If up- or down-regulation of stability gene products is responsible for genetic instability in sporadic tumors, it is necessary to clarify how these genes are regulated in human cancer tissues. A strong candidate for controlling the expression of stability genes in tumor tissues is tumor hypoxia/reoxygenation.11,12 The following is evidence that hypoxia affects the stability of the cellular genome. Several early studies demonstrated that the exposure of cultured mammalian cells to hypoxia followed by re-oxygenation results in DNA over-replication and gene amplification.72–74 For instance, Rice et al. showed that over-replication of cellular DNA is induced by H/R, which is followed by amplification of the dihydrofolate reductase gene under methotrexate selection.73 Hypoxia followed by re-oxygenation also induces fragile sites that trigger DNA breakages and gene amplification.75 Fragile sites are chromosomal sites that show gaps and breaks after inhibition of DNA synthesis.76 They are usually associated with repetitive sequences with tri-, tetra- and dodeca-nucleotide repeats or with adenosine-thymidine (AT)-rich repeats. These repeats form DNA secondary structures. Based on these unique sequences in fragile sites, Durkin and Glover proposed a molecular model for fragile site instability.77 In this model, first, a dissociation of DNA-unwinding by the helicase/topoisomerase complex and DNA synthesis occurs when the action of DNA polymerase is inhibited. This creates a long stretch of single-strand DNA around the fragile site. Second, AT-rich-repeats within a single strand of DNA form a hairpin structure by self annealing. This structure further causes replication fork stalling. Although most of these structures will be detected and repaired by DNA repair machinery, some forks collapse, resulting in formation of single or double stand breaks, and present themselves as gaps or breaks on metaphase chromosomes at fragile sites.77 In support of this model, Pires et al. demonstrated that acute and severe hypoxia (<0.02% O2 for <8 h) blocks DNA synthesis of human cancers through inhibition of replication initiation and elongation. This blockage is due to the reduction of levels of the four dinucleotide triphosphate molecules that are required for DNA synthesis.54 A break at a hypoxia-induced fragile site may initiate gene amplification through the breakage-fusion-bridge mechanism.78 Another example of H/R-induced chromosomal alterations was reported by Rofstad et al.79 They examined the effects of severe hypoxia (<0.01% O2 for 24 h) on chromosome contents of diploid as well as hyperdiploid human melanoma cell lines. They found that a subpopulation of diploid cells was arrested at the G2/M boundary during hypoxia exposure. During the first M phase after re-oxygenation, they observed a cell population which showed tetraploid chromosomes where homologous chromosomes were grouped in pairs (diplochromosomes), suggesting that severe H/R may disturb cell mitosis.79,80 Lee et al. placed phytohemagglutinin-stimulated normal human lymphocytes from 40 healthy donors under mild hypoxia (3% oxygen concentration) for 12 h or 24 h.81 After hypoxia exposure the cells were subjected to chromosomal analysis. They found that the frequency of sister chromatid exchange (SCE) (recombination between homologous sister chromatids) was higher in hypoxia treated cultures than normoxia cultures.80 The mechanism for SCE by H/R is not clear, however, because a perturbation of DNA synthesis results in SCE, these results suggest that even moderate levels of hypoxia followed by reoxygenation affects the DNA synthesis of a normal cell. The effect of hypoxia on gene mutations has been examined by several mutation assay systems. Reynolds et al. transplanted tumorigenic mouse cells into nude mice or placed the cells under hypoxic conditions in vitro.10 These cells were marked with a lambda shuttle vector containing supF as a reporter for mutations. The results showed a significant increase in point mutations and small deletions in DNA rescued from hypoxic cells transplanted into nude mice, as well as in cells exposed to hypoxia in tissue cultures. Sixty-two percent of point mutations showed transversion (G > T, G > C and A > C) and 38% were transitions (G > A) in DNA from hypoxic cells. In contrast, the percentage of transition (62%) mutations dominated over transversion mutations (38%) under normoxic conditions.10 Because the major oxidative DNA damage product, 8-oxo-G, can produce transversion mutations (G > C or G > T),46 the observed increase in mutation frequency may be caused by oxidative damage. This was supported by Keysar et al., who showed that the free radical scavenger dimethyl sulfoxide blocked hypoxia-induced gene mutations.82 Because hypoxia itself does not cause DNA damage,55 oxidative stress must be generated during re-oxygenation. Similarly, Rapp-Szabo et al. reported that hypoxia/re-oxygenation increased the mutation frequency of a reporter gene, lacI, integrated into the cellular DNA of cell lines derived from the BigBlue rat.83 They observed a small bias of transversion mutations against transition mutations in hypoxic cells in tissue cultures. These results suggest that H/R increases mutation frequency through oxida