Tuning Cell Cycle Regulation With an Iron Key

[Cell Cycle 6:16, 1982-1994, 15 August 2007]; ©2007 Landes Bioscience

Review

Tuning Cell Cycle Regulation With an Iron Key Yu Yu† Zaklina Kovacevic† Des R. Richardson*

Abstract

*Correspondence to: Des R. Richardson; Iron Metabolism and Chelation Program; Department of Pathology; University of Sydney; Sydney, New South Wales Australia; Tel.: +61.2.9036.6548; Fax: +61.2.9036.6549; Email: [email protected]

Key words iron, iron chelators, cell cycle, p53, cyclin D1, p21CIP1/WAF1

.D

Previously published online as a Cell Cycle E-publication: http://www.landesbioscience.com/journals/cc/article/4603

ON

OT D

Original manuscript submitted: 06/19/07 Manuscript accepted: 06/19/07

RIB

†These authors contributed equally to this manuscript.

IST

Iron Metabolism and Chelation Program; Department of Pathology; University of Sydney; Sydney, New South Wales Australia

UT E

.

Iron (Fe) is essential for cellular metabolism e.g., DNA synthesis and its depletion causes G1/S arrest and apoptosis. Considering this, Fe chelators have been shown to be effective anti‑proliferative agents. In order to understand the anti‑tumor activity of Fe chelators, the mechanisms responsible for G1/S arrest and apoptosis after Fe‑depletion have been investigated. These studies reveal a multitude of cell cycle control molecules are regulated by Fe. These include p53, p27Kip1, cyclin D1 and cyclin‑dependent kinase 2 (cdk2). Additionally, Fe‑depletion up‑regulates the mRNA levels of the cdk inhibitor, p21CIP1/WAF1, but paradoxically down‑regulates its protein expression. This effect could contribute to the apoptosis observed after Fe‑depletion. Iron‑depletion also leads to proteasomal degradation of p21CIP1/WAF1 and cyclin D1 via an ubiquitin‑independent pathway. This is in contrast to the mechanism in Fe‑replete cells, where it occurs by ubiqui‑ tin‑dependent proteasomal degradation. Up‑regulation of p38 mitogen‑activated protein kinase (MAPK) after Fe‑depletion suggests another facet of cell cycle regulation respon‑ sible for inhibition of proliferation and apoptosis induction. Elucidation of the complex effects of Fe‑depletion on the expression of cell cycle control molecules remains at its infancy. However, these processes are important to dissect for complete understanding of Fe‑deficiency and the development of chelators for cancer treatment.

Iron (Fe) is a metal that is vital for the sustenance of life.1‑3 It is an essential component of many proteins and enzymes that are involved in cell growth and replication.1,3 For instance, Fe plays a crucial role in the conversion of ribonucleotides into deoxyribonucleotides by participating in the rate‑limiting step of DNA synthesis catalyzed by ribonucleotide reductase (RR).4‑6 Depletion of Fe in cells typically results in a G1/S arrest,7,8 indicating that this metal is essential for cell cycle progression, growth and division.9,10 Under some conditions of Fe‑deprivation, a G2/M arrest has also been identified.11 Iron‑deficiency is a common nutritional problem affecting an estimated 500 million individuals resulting in anemia, lethargy, defective psychomotor development and disturbed cognitive function.12 Despite this and the fact that Fe‑depletion induces a G1/S arrest and apoptosis,8,13 it is surprising that little is known concerning the role of Fe in cell‑cycle regulation. Furthermore, it has become clear that some Fe chelators show promising anti‑cancer activity by inducing cell cycle arrest and apoptosis.14,15 However, the mechanisms involved in these effects remain uncertain and important to investigate in terms of understanding chelator structure-activity relationships. In order to understand the role of Fe in cell cycle control, knowledge of its cellular metabolism is essential and this is briefly described below. For more detailed accounts of Fe metabolism and homeostasis, the reader is referred to specialized review articles.3,16‑18

©

20

07

LA

ND

ES

BIO

SC

IEN

This project was supported by a fellowship and project grants from the National Health and Medical Research Council of Australia, Australian Research Council Discovery Grant, Cancer Institute of New South Wales Foundation grant and grants from Australian Rotary Health Research Fund to D.R.R. Z.K. thanks the Australian Rotary Health Research Fund (Dural Rotary Club) and the Cancer Institute of New South Wales for a Ph.D. Scholarship. Y.Y. appreciates an International Post-Graduate Research Scholarship from the University of Sydney. The authors kindly thank Mr. Yohan Suryo Rahmanto and Miss Danuta Kalinowski for their tremendous assistance with the reference formatting of the article.

CE

Prelude

ACKNOWLEDGEMENTS

1982

Cellular Iron Metabolism: From Uptake to Storage Inorganic Fe in the diet is largely in its ferric state and is absorbed by enterocytes lining the small intestine.19 The uptake of Fe is first performed by a reduction step that may be mediated by the enzyme duodenal cytochrome b (Dcytb; Fig. 1A).19 However, the role of this protein in this process is controversial, as Dcytb knockout mice do not show an Fe‑deficiency phenotype.20 Apart from inorganic Fe uptake from the diet, heme may be transported into enterocytes by the recently identified heme carrier protein 1 (HCP1).21 Again, the function of this molecule in Fe absorption remains unclear, with HCP1 also being suggested to be a folate transporter.22,23 Irrespective of the transport mechanism, Cell Cycle

2007; Vol. 6 Issue 16

Iron and the Cell Cycle

Figure 1. The absorption, transport and metabolism of Fe: (A) Fe is absorbed by enterocytes in the small intestine. In the diet, Fe exists as either inorganic Fe or heme Fe. Inorganic Fe is primarily in the Fe3+ state which may be converted to Fe2+ by the postulated ferrireductase, duodenal cytochrome b (Dcytb), although this remains controversial. The Fe2+ is then transported into enterocytes via divalent metal ion transporter 1 (DMT1). Heme Fe is thought to be trans‑ ported into the cell by heme carrier protein 1 (HCP1), although again this remains uncertain. Once internalized, heme is metabolized by heme oxygenase to release Fe, carbon monoxide (CO) and bilirubin. Irrespective of whether Fe has been derived from dietary inorganic Fe or heme, once inside the enterocyte, Fe2+ is either stored in ferritin or transported out of the enterocyte into the blood via the Fe export protein, ferroportin-1. The intracellular ferroxidase, hepha‑ estin, is thought to be involved in this process, although its exact contribution remains unclear. Once at the surface of the enterocyte, Fe2+ is converted back to Fe3+ by the serum ferroxidase, ceruloplasmin. The Fe3+ then binds to the serum Fe transport molecule, transferrin (Tf). (B) Cells which require Fe express the transferrin receptor‑1 (TfR1) on their surface, which binds two molecules of Tf. The Tf‑TfR1 complex is then internalized by receptor‑mediated endocyto‑ sis. Once in the endosome, the pH decreases, allowing the Fe3+ to dissociate from the Tf-TfR1 complex. The endosomal ferrireductase, six‑transmembrane epithelial antigen of the prostate‑3 (Steap3), is thought to convert Fe3+ to Fe2+ in the endosome, allowing Fe2+ to be transported out of the endosome by DMT1. Once in the cell, Fe2+ can be stored in ferritin or it can enter the poorly characterized intracellular labile Fe pool where it is used in the synthesis of various proteins and enzymes such as ribonucleotide reductase (RR) etc.

heme enters the cell and is then metabolized by heme oxygenase, leading to the liberation of Fe, bilirubin and carbon monoxide (Fig. 1A).24 After transport through the apical membrane of the enterocyte and depending upon the body’s requirement, some Fe is stored in the protein, ferritin.25,26 Iron that is not stored within ferritin is released by enterocytes into the bloodstream via a poorly understood mechanism (Fig. 1A). The intracellular ferroxidase, hephaestin, plays some role in this process,27 while the transporter ferroportin‑1, is known to export cellular Fe into the circulation through the basolateral membrane.28 Hepcidin is a peptide hormone secreted by the liver and is involved in maintaining Fe homeostasis.29 It has been shown to downregulate ferroportin‑1 by causing its internalization and degradation.29 Indeed, this is one step of a homeostatic loop of Fe regulation where an excess of Fe in the diet stimulates hepcidin secretion. Hepcidin then reduces Fe uptake into the bloodstream and consequently promotes Fe storage in ferritin.26 Another recently identified protein involved in Fe homeostasis is hemojuvelin (HJV), that is highly expressed in skeletal muscle and liver.30,31 When it is mutated, HJV has been identified as the gene responsible for juvenile hemochromatosis.30 Mice with mutated HJV fail to express hepcidin, leading to severe Fe overload.31 These observations suggest www.landesbioscience.com

HJV plays an essential role in regulating hepcidin expression in the Fe‑sensing pathway.31 Once the Fe in its ferrous state (Fe+2) has been transported by ferroportin‑1 to the surface of the enterocyte, it is thought to be converted to the ferric form (Fe+3) by the copper‑containing serum ferroxidase, ceruloplasmin (Fig. 1A).26,27 Ceruloplasmin may directly interact with ferroportin‑1 to achieve this reaction, although the mechanism involved remains unclear. Serum transferrin (Tf ) then binds the Fe+3 with high affinity.32 Two Fe+3 atoms become bound to Tf, which then binds to the transferrin receptor‑1 (TfR1) that is found on the surface of most cells and is involved in internalized Fe uptake.33 The uptake of Fe occurs following the binding of two Tf molecules to the TfR1.32 The entire complex is then internalized via receptor‑mediated endocytosis (Fig. 1B).9,26,32,34 Following internalization of the Tf‑TfR1 complex, the pH within the endosome declines due to the presence of a proton pump in the endosomal membrane.35 The so formed acidic environment (pH 5.5) then allows the Fe+3 atoms to dissociate from the complex.35 The Fe+3 is subsequently reduced to the ferrous state (Fe+2) possibly by a ferrireductase known as the six‑transmembrane epithelial antigen of the prostate‑3 (Steap3)36 and is then transported out of the endosome into the cytoplasm via the divalent metal

Cell Cycle

1983


transporter‑1 (DMTI; Fig. 1B).18,35‑37 Apo‑Tf bound to the TfR1 is then recycled back to the cell surface rather than being degraded (Fig. 1B).26 A second Tf receptor (TfR2) gene has been cloned and results in the generation of two transcripts, a and b.38 The TfR2a leads to a membrane‑bound protein that binds Tf, but at a lower affinity than TfR1.38,39 The TfR2a appears to play a role in sensing Fe levels rather than being involved in quantitative Fe uptake.16,40 In fact, mutation of this protein leads to Fe‑loading and hemochromatosis.41 Consistent with the role of TfR1 expression in quantitative Fe uptake, TfR1 is well known to be regulated by intracellular Fe concentration,18,42 while TfR2 is not.39 TfR2 appears to be regulated as a function of the cell cycle, with the highest expression in late G1 phase and no expression in G0/G1.39 Interestingly, CHO cells expressing TfR2a developed into tumors in nude mice, whereas CHO control cells did not.39 While TfR2 is involved in Tf‑binding and Fe uptake,38 it cannot compensate for TfR1‑mediated Fe uptake, since TfR1 knockout mice are not viable.43,44 Once in the cytoplasm, Fe is thought to enter the poorly characterized intracellular labile pool and can either be used in the production of new Fe‑containing proteins and enzymes such as RR and hemoglobin etc, or it can be stored in ferritin.17,26 The labile Fe pool was once thought to be composed of low Mr complexes,45 but more recent studies suggest that active protein‑protein and/or organelle interactions such as contact of the Tf‑containing endosome with the mitochondrion may be involved.46‑48 A significant amount of metabolically active Fe is found in the mitochondrion where it participates in the synthesis of Fe‑S clusters or heme.49,50 In mammalian cells, Fe is thought to be transported into the mitochondrion by the membrane‑bound Fe transporter, mitoferrin.51 Disruption of yeast mitoferrin homologs, MRS3 and MRS4, was found to cause defective mitochondrial Fe uptake, leading to defects in mitochondrial Fe metabolism and Fe‑S cluster formation.51 Hence, mitoferrin appears to play an essential role in mitochondrial Fe uptake.51 The storage of some Fe in the mitochondrion is mediated by mitochondrial ferritin.52 Recently, it was found that overexpression of mitochondrial ferritin alters cellular Fe homeostasis causing a cytosolic Fe‑deficiency, increasing TfR1 expression and leading to enhanced Fe uptake from Tf.53 The role of Fe in a variety of metabolic processes necessitates that its cellular levels are tightly controlled. Two iron regulatory proteins (IRP1 and IRP2) have been identified that play important roles in regulating Fe homeostasis.17,54 Both are mRNA‑binding proteins which play an important role in the post‑transcriptional control of a variety of molecules involved in Fe homeostasis18,34,42 and potentially cell cycle regulation.55 The IRPs bind to stem‑loop structures known as Fe‑responsive elements (IRE) found in the untranslated regions (UTRs) of the mRNA of molecules involved in Fe metabolism e.g., ferritin, TfR1, ferroportin‑1, DMT1 etc.34,42 The RNA‑binding activity of the two IRPs are regulated by Fe via different mechanisms. In the case of IRP1, when cells are Fe‑replete, an Fe‑S cluster forms within the protein that inhibits RNA‑binding activity.17 In contrast, in the case of IRP2, high Fe levels result in its proteasomal degradation.56 During Fe‑deprivation, IRP‑RNA binding activity is high, leading to the binding of IRPs to the IRE on the 5' UTR of ferritin mRNA resulting in inhibition of its translation.55 This leads to less Fe storage and the use of Fe for metabolic requirements. On the other hand, the binding of IRPs to IREs in the 3' UTR of TfR1 mRNA makes it more stable and less susceptible to degradation leading to 1984

greater TfR1 expression.55 Subsequently, this promotes the uptake of more Fe via the binding of Tf to the TfR1. It follows that removal of Fe from the hypothetical intracellular labile Fe pool will result in increased activity of the IRPs, leading to the altered expression of a range of molecules possessing IREs. In contrast, when cells are Fe‑replete, this leads to decreased IRP‑RNA binding activity and an opposite scenario develops, leading to decreased TfR1 expression and increased Fe storage in ferritin.17 The recent identification of an IRE within the 3' UTR of cell division cycle 14A (CDC14A) mRNA implies a potential role for this molecule in the cell cycle arrest observed after Fe‑depletion and this is discussed later in the section on CDC14A. Considering the importance of Fe in vital cellular processes such as DNA synthesis, it is not surprising that the cell cycle is “tuned” to Fe availability. For instance, it is well known that TfR1 expression is increased during the S phase of the cell cycle.57 This is presumably due to the Fe requirement of RR that is critical for DNA synthesis.4,6 In agreement with this, it has been reported that depression of TfR1 expression leads to G1 arrest and an alteration in the expression of genes that regulate the cell cycle.58 The essential requirement of Fe for cellular metabolism has led to investigations on the use of Fe chelators as anti‑cancer agents.2,59 By understanding the effects of these agents on cell cycle control, the rationale design of more active compounds can be envisaged.

Iron as Possible Therapeutic Strategy for Cancer Treatment Compared to normal cells, neoplastic cells require greater amount of Fe because generally they proliferate at a greater rate than their normal counterparts.10,15 This is reflected by the higher expression of TfR132,60 and the higher rate of Fe uptake from Tf in cancer cells.61,62 Furthermore, neoplastic cells express high levels of RR63,64 making them more susceptible to the action of Fe chelators than normal cells.10,65 Early studies showed that the clinically used Fe chelator, desferrioxamine (DFO), had some activity at inhibiting the growth of neuroblastoma and leukemia in cell culture and clinical trials.66‑70 This was despite the fact that this chelator suffers from limited membrane permeability.71,72 Considering that DFO was developed specifically for the treatment of Fe‑loading diseases such as b‑thalassemia major and not cancer treatment, its activity in these later trials was encouraging.15 Indeed, there has been continuing efforts to improve the potency and selectivity of Fe chelators against cancer cells (reviewed in refs. 2, 15, 59 and 73). Among these ligands are 3‑aminopyridine‑2‑carboxaldehyde thiosemicarbazone (Triapine®),74 Tachpyridine,75 O‑Trensox76 and hybrid chelators derived from pyridoxal isonicotinoyl hydrazone (PIH)77,78 and thiosemicarbazones.79‑84 The success of Fe chelators as potential anti‑tumor agents is marked by the entry of Triapine® into phase I and II clinical trials alone or in combination with a range of chemotherapeutics.85,86 In fact, recent results with this agent have shown that it can successfully reduce white blood cell counts by 50% in leukemia patients.87 A structurally different ligand known as Tachpyridine is also currently in pre-clinical development with the National Cancer Institute.88 Another series of chelators are those of the PIH class that led to the development of the highly permeable ligand, 2‑hydroxy‑1‑napthylaldehyde isonicotinoyl hydrazone (311).79,89 This compound shows much greater Fe chelation efficacy and anti‑tumor activity than DFO.72,90,91 Studies with 311 and its derivatives83,92 led to the

Cell Cycle

2007; Vol. 6 Issue 16


Figure 2. Summary of the cell cycle in normal Fe‑replete cells. The cell cycle consists of four main phases: G1, S, G2 and M‑phase. Under normal conditions in Fe‑replete cells, the pro‑ gression of the cell cycle is controlled by a number of molecules including the cyclins A, B, D and E, as well as the cyclin‑dependent kinases (cdks). Cyclin D1 forms a complex with cdk4, while cyclin E binds with cdk2, allowing them to become active enzymes. These complexes are then involved in hyper-phosphorylation of the retinoblastoma susceptibility gene product (pRb), which allows it to release the transcription factor, E2F1. Once free, E2F1 is able to translocate to the nucleus where it mediates the transcription of a range of genes vital for S‑phase progression. One of the most important mediators of this G1/S checkpoint is p53, which is able to cause G1/S arrest under conditions of cell stress or DNA damage. One function of p53 is to transactivate the expression of the cdk inhibitor, p21CIP1/WAF1, which then inhibits the activity of cyclin D1/cdk4‑ and cyclin E/cdk2 complexes, thereby prevent‑ ing entry into S‑phase. However, the activity of p21CIP1/WAF1 can be paradoxical and under some conditions can aid in cell cycle progression (see section on p21CIP1/WAF1). In addition, p53 is also able to inhibit cyclins A and B leading to G2/M arrest.

development of a highly potent series of Fe chelators derived from di‑2‑pyridylketone thiosemicarbazone (DpT).59,84,93,94 A number of these ligands have also demonstrated marked in vitro and in vivo anti‑tumor activity82,84,93,94 and are currently being developed as novel anti‑cancer agents. In an attempt to understand the effects of Fe chelators on cellular proliferation and in order to develop more effective ligands, a number of studies have examined the roles of Fe in regulating the cell cycle.95

The Cell Cycle: A Brief Overview As normal cells grow and divide, they progress through the cell cycle in a regulated manner.96 The main groups of molecules involved in the regulation of the cell cycle include the cyclins, cyclin‑dependent kinases (cdks), cyclin‑dependent kinase inhibitors (e.g., p21CIP1/WAF1) and tumor suppressor genes such as p53 and the www.landesbioscience.com

retinoblastoma susceptibility gene product (pRb).96‑99 The cdks are dependent on cyclins to modulate their phosphorylation activity.97,98 For example, cyclin D1 binds to cdk4/6, while cyclin E binds to cdk2 leading to an activation of kinase activity (Fig. 2).97,98,100 Together, these molecules allow the progression of the cell cycle from G1 to S phase (Fig. 2).97,98,100 The S phase is regulated by cyclin A, while the M phase is controlled in part by cyclin B (Fig. 2).97,98 It is the up‑regulation and degradation of the cyclins and their subsequent interaction with cdks that mediate progression through the cell cycle.97,100 There are a number of major checkpoints in the cell cycle that are present at G1/S, S, G2/M and M‑phase (Fig. 2).97,100 These checkpoints are important to determine whether cells will proceed to the next phase of the cycle.97,98,100 One important regulator of the cell cycle is p53, which is involved in both the G1/S and the G2/M checkpoints (Fig. 2).98 In fact, p53 functions to arrest cells following DNA damage and initiate a repair mechanism, or if the DNA is beyond repair, it will activate an apoptotic pathway.97,100 In response to DNA damage, p53 will transactivate one of its downstream targets, p21CIP1/WAF1, which subsequently inhibits cyclin D/cdk4/6 and the cyclin E/cdk2 complexes resulting in G1/S arrest (Fig. 2).97,100 Other targets of p53 include cell cycle regulatory molecules such as growth arrest and DNA‑damage‑inducible 45 alpha (GADD45a), which is induced upon DNA damage and can arrest the cell cycle (see section on the GADD family).97,100 Another key regulator of the cell cycle is pRb, which controls entry of cells into S‑phase.97,100 When hypo‑phosphorylated, pRb binds to and represses the transcription factor E2F1 (Fig. 2).98 Under normal conditions, accumulating cdk‑cyclin D and ‑E complexes function to hyper‑phosphorylate pRb, resulting in the release of E2F1, which leads to the expression of S‑phase genes and subsequent entry into S‑phase.34 The role of Fe in regulating some of these effector molecules that play roles in cell cycle control are discussed below.

The Molecular Targets of Iron Chelators and their Effects on the Cell Cycle A mechanism by which Fe chelators exert their anti‑proliferative effects on tumors is by targeting molecules that are critical in regulating progression of the cell cycle.101,102 Some of these include: RR,5,91 cyclins,101,103 cdks,101,103 p53,101,104‑106 p21CIP1/WAF1,101,107 p27Kip1,108‑110 GADD45a,72 hypoxia inducible factor‑1a (HIF‑1a), N‑myc downstream regulatory gene‑1 (Ndrg‑1)102 and pRb.98,111 By altering the expression and/or function of the above molecules, Fe‑depletion is able to effectively inhibit the growth of tumor cells. Ribonucleotide reductase. Ribonucleotide reductase is an enzyme that catalyses the de novo biosynthesis of deoxyribonucleotides essential for DNA replication, cell cycle progression and cellular repair.6 This process requires a tyrosyl free radical which acts to reduce the corresponding ribonucleotides to deoxyribonucleotides.112 Hence, structurally, RR consists of a “radical generator” and a reductase.113

Cell Cycle

1985


The enzyme is classified into three classes based on different cofactors, such as Fe or cobalamin for their catalytic activity.6 Class I enzymes are found practically in all eukaryotic organisms and in some prokaryotes and viruses. This class is further divided into three subclasses (Ia, Ib and Ic) based on polypeptide sequence homology and allosteric behaviour.114 Human RR is a tetramer that belongs to class Ia and consists of two non-identical homodimers, R1 and either R2 or p53R2.112 The R1 protein contains the active sites and binding sites for allosteric effectors, while the R2 subunit contains one di‑nuclear Fe centre and one stable tyrosyl radical per monomer that is vital for enzymatic activity.112 The R2 subunit is necessary for “housekeeping DNA synthesis” that is essential for DNA replication during the S/G2 phases. In contrast, the p53R2 subunit supplies dNTPs for DNA repair after DNA damage in G0/G1 phase cells in a p53‑dependent manner.115 It has been reported that there is also an additional p53‑independent induction of p53R2, because cells with mutated p53 still express this molecule in response to DNA‑damaging agents.116 In fact, p53R2 can be a transcriptional target of the p53 family member, p73.117 Both the R2 and p53R2 subunits possess an Fe‑binding site that is important for their enzymatic function.118 Since the reduction of ribonucleotides is the rate‑limiting step of DNA synthesis, inactivation of RR has a number of consequences, such as inhibition of DNA synthesis, cell proliferation and DNA repair, leading to cell cycle arrest and apoptosis.119 On the other hand, increased RR activity has been associated with malignant transformation and tumor cell growth,119 making RR an important but largely ignored target for anti‑cancer agents. As Fe is required for the enzymatic activity of RR, Fe chelation is well known to effectively inhibit the activity of this enzyme.5,91,120 Triapine® is one of the most potent inhibitors of RR that targets the R2 subunit of the enzyme.5,121 However, more recent data also found that Triapine® indiscriminately inhibits the p53R2 subunit as well.118 �� Iron‑depletion�� affects multiple molecular targets involved in cell‑cycle control besides RR and these are described below. Cyclins and cdks. As described above, cyclins and cdks are critical for the normal progression through the cell cycle (Fig. 2).96,99 It is the regulated alterations in the availability and activity of these molecules that governs the transition between the different phases of the cycle.96 Iron‑depletion mediated by chelators was found to affect the expression of several cyclins and cdks.101,103,122 Specifically, in SK‑N‑MC neuroepithelioma cells, Fe‑depletion markedly reduced the expression of cyclins D1, D2 and D3, while having a lesser effect on decreasing cyclin A and B levels.101 This latter reduction in cyclin A protein in tumor cells is in good agreement with the results from normal T lymphocytes, where there was a decrease in cyclin A protein and its kinase activity after incubation with DFO.7 Iron‑depletion has also been shown to decrease the expression of cdk2101,120 or cdk4103 protein depending on the cell type and experimental conditions. Studies examining the effect of DFO in neuroblastoma cells have found that protein levels and kinase activity of p34cdc2 is decreased.8 This is of significance, as p34cdc2 functions in the G2/M and potentially G1/S phase transitions, being the catalytic subunit that complexes with cyclin A, B and E.123,124 The effect of Fe depletion on p34cdc2 may explain, at least in part, the G1/S and G2/M arrest observed after Fe chelation under some experimental conditions. A recent study has identified that the mechanism of the Fe‑depletion mediated reduction in cyclin D1 protein expression is a result of proteasomal degradation, there being no decrease in cyclin D1 1986

mRNA levels (Fig. 3).125 However, in contrast to the ubiquitin‑dependent pathway of cyclin D1 degradation that regulates the expression of this molecule under Fe‑replete conditions, Fe‑depletion induced an ubiquitin‑independent pathway of proteasomal degradation (Fig. 3).125 Importantly, the effects of Fe chelators on cyclin D1 expression were found to be due to Fe‑depletion, as the supplementation of Fe was able to reverse these effects.125 Considering the rate‑limiting role that cyclin D1 plays in G1/S progression,96 its regulation by Fe appears to be important for preventing entrance into the S phase, where Fe is essential for RR activity4 and thus, DNA synthesis. It has been suggested that the Fe‑depletion mediated down‑regulation of cyclin D1 leads to decreased phosphorylation of pRb that may be, in part, responsible for the G1/S arrest observed. The importance of regulating cyclin D1 in terms of cell cycle control is obvious from studies demonstrating that over‑expression of cyclin D1 releases cells from their normal controls and acts as an oncogene.126,127 In fact, pharmacological targeting of cyclin D1 may lead to novel anti‑tumor agents128 and the fact that Fe chelators markedly decrease the expression of this molecule may be important in their anti‑tumor activity. Interestingly, in contrast to other cyclins, cyclin E protein expression was found to be elevated in response to Fe‑depletion in neuroepithelioma cells.101,120 This paradoxical response may reflect an attempt by the cell to maintain cell cycle progression after Fe chelation. However, because cyclin E requires cdk2 for its activity and since the expression of the latter is reduced by Fe‑depletion,101 the increase in cyclin E does not overcome the G1/S arrest after Fe chelation.101,103,111 In fact, several studies have shown that following Fe‑depletion, pRb becomes hypo‑phosphorylated leading to G1/S arrest.111,129 These studies demonstrate that expression of critical effectors of the cell cycle such as some cyclins and cdks are affected by intracellular Fe levels, providing another level of control on cell cycle progression. pRb. pRb is an important molecule that mediates progression of the cells from G1 to the S phase of the cell cycle. As already discussed, the regulation of pRb during the cell cycle is through phosphorylation of the protein by cyclin‑dependent kinases (cdks) (Fig. 2).130 Since the expression of cyclin D1 and cdk2 are reduced during Fe‑depletion,101,120,125 this will prevent the formation of cyclin‑cdk complexes leading to hypo‑phosphorylation of pRb that will contribute to G1/S arrest (Fig. 4).101 Indeed, a number of investigators have reported that Fe chelation resulted in the decrease of hyper‑phosphorylated pRb101,111 in neuroepithelioma cells,101 human breast cancer cells103 and T lymphocytes.111 Hypo‑phosphorylation of pRb during mid to late G1 phase by cdk4‑ or cdk6‑cyclin D complexes prevents the release of transcription factor E2F1 from pRb.130,131 The release of E2F1 is necessary to transcribe genes critical for cell cycle progression, such as cyclin A and cyclin E.131 Hence, Fe availability results in alterations in pRb phosphorylation that may play a role in the G1/S arrest observed after Fe‑depletion. p53. One of the most well‑known and important regulators of the cell cycle is p53.10,129 This transcription factor has a multitude of molecular targets and plays a critical role in the G1/S checkpoint (Fig. 2).129 p53 is activated in response to cellular stress or DNA damage and functions to initiate repair mechanisms or, when the damage is irreparable, apoptotic pathways.101,104,105,132 A number of studies have reported elevated levels of p53 protein expression following Fe‑depletion.105,106,133 This increase appears to be at the post‑transcriptional level as there is no change in p53 mRNA after Fe‑depletion.101,105 In some studies, Fe‑depletion

Cell Cycle

2007; Vol. 6 Issue 16


Figure 3. Iron‑depletion induces Ub‑independent degradation of cyclin D1 and p21CIP1/WAF1. Iron‑depletion results in decreased expression of the key cell cycle regulators, cyclin D1 and p21CIP1/WAF1. When cells are Fe‑replete, the traditional pathway of cyclin D1 and p21CIP1/ WAF1 degradation involves ubiqutination of these proteins, allowing them to be transported to the proteasome for degradation. Upon Fe‑depletion, an Ub‑independent pathway leads to the degra‑ dation of both cyclin D1 and p21CIP1/WAF1 by the proteasome. Iron‑depletion also reduces the transport of p21CIP1/WAF1 from the nucleus to the cytoplasm, further reducing the translation of this protein. Furthermore, cyclin D1 and p21CIP1/ WAF1 compete for the proteasomal binding site, suggesting that decreased cyclin D1 expression may lead to increased p21CIP1/WAF1 degrada‑ tion and vice versa.

resulted in the upregulation of p53 expression only in cells expressing the wild‑type molecule,106 while in other investigations, p53 protein levels increased after Fe chelation irrespective of whether it was mutated or not.105 In whole cell systems, Fe‑depletion was able to induce p53‑transactivational activity and sequence‑specific DNA binding in a dose‑ and time‑dependent manner.106,133 Further studies have revealed that several mechanisms may be involved in the activation of p53 by Fe chelation. These include: (1) an increase in p53 protein expression;106 (2) increased conversion of latent p53 to its active DNA‑binding form;104 (3) phosphorylation of p53 at serine‑15 which increases its stability and prevents proteasomal degradation by mdm‑2.104 The increased p53 phosphorylation at this site may indicate up‑regulation of ataxia telangiectasia mutated (ATM) and/or ATM‑Rad3 related (ATR) after Fe‑depletion;104 and (4) other molecules that are also targets of Fe‑depletion, such as the transcription factor, hypoxia inducible factor‑1a (HIF‑1a), can also increase p53 expression.98 While Fe‑depletion results in an increase in p53 protein expression and transcriptional activity, it is unclear which of its molecular targets are affected. It is known that the expression of both p21CIP1/WAF1 and GADD45 mRNA are increased after Fe chelation, but this occurs not only in cells with native p53, but also in those with mutant p53.72 This indicates that p53‑independent up‑regulation of these genes can occur after Fe‑depletion. p21CIP1/WAF1. The progression of the cell cycle is a strictly regulated process in which a number of crucial molecules are involved.99 One such protein is p21CIP1/WAF1, a cdk inhibitor that is involved in regulating the cell cycle.134‑136 Specifically, when over‑expressed, p21CIP1/WAF1 binds to the cyclin E/cdk2 complex, preventing pRb phosphorylation (Fig. 2)137,138 and preventing the G1/S‑phase transition leading to cell cycle arrest.139 In addition, p21CIP1/WAF1 can also prevent DNA replication and affect other transcription factors involved in cell cycle progression, such as E2F1 and c‑myc.140‑142 Interestingly, and paradoxically, when expressed at very low levels, p21CIP1/WAF1 is required for the assembly of cyclin D/cdk complexes and is therefore an important component of cell cycle progression.143,144 Furthermore, p21CIP1/WAF1 has been shown to directly www.landesbioscience.com

inhibit caspase‑3 activation, thereby preventing apoptosis.143,144 Indeed, down‑regulation of p21CIP1/WAF1 in tumor cells was found to lead to increased apoptosis.145 Although often activated by p53 in response to cellular stress and/ or DNA damage, p21CIP1/WAF1 can also be induced by p53‑independent pathways involving other transcription factors such as AP2, Sp1 or Sp3.146,147 More recently, it was discovered that p21CIP1/WAF1 mRNA can be markedly up‑regulated by Fe‑depletion using chelators such as 311 and DFO by a p53‑independent pathway.72,148 At the same time, it was found that Fe‑depletion actually decreased the expression of p21CIP1/WAF1 protein.107 Subsequent supplementation of these cells with Fe restored p21CIP1/WAF1 protein levels, demonstrating that the effect observed was due to Fe‑depletion.107,148 It has also been shown that the Fe‑depletion mediated decrease in p21CIP1/WAF1 protein was due to two mechanisms: (1) a decrease in nuclear translocation of p21CIP1/WAF1 mRNA to the cytosol and (2) ubiquitin‑independent proteasomal degradation leading to reduced p21CIP1/WAF1 protein levels (Fig. 3).107,148 Interestingly, Fe‑depletion appears to have similar effects on both cyclin D1 and p21CIP1/WAF1 protein expression, where it induces ubiquitin‑independent degradation (Fig. 3).107,125 The precise ubiquitin‑independent pathway of p21CIP1/WAF1 degradation has not been identified. However, it may be mediated by NAD(P)H:quinone‑oxidoreductase‑1149 or antizyme150 that are responsible for degradation of other proteins by this process. Of interest, it has been previously demonstrated that cyclin D1 and p21CIP1/WAF1 compete for the C8a subunit binding site of the proteasome (Fig. 3).151 Therefore, the down‑regulation of cyclin D1 and p21CIP1/WAF1 following Fe‑depletion may facilitate degradation of each of these molecules via this pathway.151 Under very different experimental conditions, Gazitt and colleagues152 identified that HL60 leukemia cells need to be Fe‑replete to transcribe p21CIP1/WAF1 when induced by the phorbol ester, phorbol myristate acetate (PMA). In these studies, Fe‑deprivation induced by DFO blocked PMA‑induced differentiation and induced S‑phase arrest and apoptosis.152 Clearly, the difference on whether there is up‑ or down‑regulation of p21CIP1/WAF1 mRNA after Fe‑depletion can be ascribed to the experimental conditions used

Cell Cycle

1987


in each case. The biological effects of PMA are numerous and do not relate to a physiological or pharmacologically‑induced state in humans. Furthermore, the effect of PMA on p21CIP1/WAF1 has only been found in HL60 cells.152 In contrast, the effects of Fe‑depletion on up‑regulating p21CIP1/WAF1 mRNA have been observed in a variety of cell types72,92,101,107,148,153 and are relevant to the pharmacological effects of chelators as anti‑tumor agents.75,84,93 Examination of the Fe chelator‑mediated downregulation of p21CIP1/WAF1 is important, as apart from being a cdk inhibitor and positive regulator of the cell cycle, this protein also has anti‑apoptotic activity. It is known that high p21CIP1/WAF1 expression in some cancers may provide a growth advantage capable of subverting apoptosis induced by DNA‑damaging chemotherapeutics.143 In fact, by decreasing p21CIP1/WAF1 expression using anti‑sense oligonucleotides, cancer cell apoptosis can be induced.143,154 Hence, p21CIP1/WAF1 has been proposed as a target for developing novel anti‑cancer agents.143 Since Fe chelators effectively inhibit p21CIP1/WAF1 expression, this is important for understanding, at least in part, their marked anti‑tumor activity and ability to induce apoptosis. However, the effect of chelators at inhibiting cancer proliferation and inducing apoptosis is probably due to their influence on multiple molecular targets. This may explain the high anti‑tumor activity of some chelators and their ability to overcome resistance to standard chemotherapeutic agents.84,91 p27Kip1. Another cdk inhibitor that is regulated by Fe‑depletion is p27Kip1. This was first shown by Wang and colleagues108,109 using the chelator mimosine and then confirmed by others.155,156 The upregulation of p27Kip1 by Fe‑deprivation occurred at both the mRNA and protein levels.108 It was suggested that Fe‑depletion also increased the expression of transforming growth factor b1 (TGF‑b1).156 Interestingly, when this factor was neutralized using an TGF‑b1 antibody, it prevented the upregulation of p27Kip1 (Fig. 4).156 In a later more comprehensive study, it was shown that the p170 subunit of the eukaryotic initiation factor 3 (eIF3) was down‑regulated by Fe‑depletion and that this allowed increased p27Kip1 expression (Fig. 4).155 The significance of this observation is that the regulation of translation generally occurs at the initiation step which requires multiple eIF’s and the ribosome. This is of interest, as it suggests that p170 is an early response gene to Fe‑deprivation that regulates the translation of a subset of mRNAs.155 This later finding may have wider implications for understanding the changes in the expression of proteins after Fe‑depletion. The GADD family. The GADD group of genes constitutes a small family of stress response molecules comprised of GADD34, GADD45 and GADD153. The expression of these genes is often increased when cells are subjected to a stress such as nutrient deprivation (e.g., glucose, glutamine, zinc)157‑159 or exposed to DNA‑damaging agents (e.g., peroxynitrite)160 which may cause cell cycle arrest and/or apoptosis. The GADD45 group of genes plays an important role in the G2/M checkpoint and apoptosis.161 This family of genes encodes three structurally‑related proteins, GADD45a, GADD45b and GADD45g.161 However, only GADD45a has been shown to activate p53‑dependent G2/M arrest and inhibit cdc2 kinase.161 Neither GADD45b nor GADD45g have been shown to be downstream targets of p53.161 Interestingly, GADD45a is also known to interact with key cell cycle regulatory molecules, such as p21CIP1/WAF1,162 cdc2/cyclin B1163 and p38 mitogen‑activated protein kinase (MAPK; see section below on p38 MAPK).164 In fact, the cellular function of GADD45a is dependent on its interacting partner.164 For example, 1988

interaction between GADD45a and p38 MAPK has been shown to play a pivotal role in preventing oncogene‑induced growth in part by regulating p53.164 Studies using transfected cells suggest that GADD34 and GADD153 appear to have a direct role in initiating apoptosis rather than inducing cell cycle arrest.165,166 Overexpression of each GADD gene causes growth inhibition and/or apoptosis, while combined overexpression of the three GADD genes leads to synergistic or cooperative effects on anti‑proliferative activity.167 Cellular Fe‑depletion mediated by DFO or 311 has been shown to cause a pronounced concentration‑ and time‑dependent increase in the expression of GADD45 mRNA after 20 h of incubation in three different cell lines, BE‑2 neuroblastoma, SK‑N‑MC neuroepithelioma and K562 erythroleukemia.72 This effect was reversible after the removal of the ligands and also the Fe(III) complexes of DFO and 311 had no effect on GADD45 mRNA levels, suggesting that Fe‑depletion was necessary to increase GADD45 mRNA.72 Recent studies from our laboratory also indicate that GADD153 mRNA is also increased upon Fe‑depletion (Siafakas R, Fu D, Richardson DR, unpublished observations). However, interestingly, there was no appreciable increase in the GADD45 protein level in cells after Fe‑depletion, although only a single incubation time (30 h) was assessed101 and further studies are required. Similar to the effect observed after Fe‑depletion, both GADD45 and GADD153 mRNAs have been shown to be up‑regulated during hypoxia.168 This suggests a potential role for the transcription factor HIF‑1a in the up‑regulation of these genes that may be activated by both hypoxia and Fe‑depletion via prolyl hydroxylases (see section on HIF-1a). Several studies have reported that after Fe‑depletion there are alterations to the level of GADD45 interacting partners, such as p38169 and cdk2/cyclin B.101 It can be hypothesized that GADD45 may mediate growth arrest through inhibiting the activity of cyclin B and cdk2163 and studies have shown that Fe‑depletion decreases the expression of these regulatory molecules.101 p38 MAPK. The signaling molecule p38 MAPK is one of the three members of the MAPK family which also includes extracellular signal‑regulated kinase (ERK) and c‑Jun N‑terminal protein kinase/ stress‑activated protein kinase (JNK/SAPK).170 These proteins are activated by a variety of environmental stresses and inflammatory cytokines and affect processes such as cell differentiation and apoptosis.164 Studying the effect of DFO, Lee et al. showed that Fe‑depletion strongly activated p38 MAPK and ERK, but did not activate JNK.169 In addition, the selective p38 MAPK inhibitor, SB203580, and ERK inhibitor, PD98059, protected cells against Fe chelator‑induced death in oral keratinocytes and cancer cells.169 This suggests that p38 and ERK MAPKs are potential mediators of cell death induced by Fe‑deprivation. It is of interest to note that p38‑mediated growth inhibition has been suggested to involve activation of p53171 and decrease cyclin D1 expression,172 both of which also occur upon Fe‑depleti on.101,103,105,125 However, the effect of p38 MAPK on decreasing cyclin D1 expression was thought to be via decreased cyclin D1 transcription, although a post‑transcriptional mechanism that led to decreased protein levels was not excluded.172 Clearly, as noted previously, Fe‑depletion leads to decreased cyclin D1 protein levels due to proteasomal degradation.125 Later studies by Moon and colleagues173 using vascular smooth muscle cells demonstrated that the p38 MAPK pathway participated in p21CIP1/WAF1 induction after metal chelation, which consequently leads to a decrease of cyclin

Cell Cycle

2007; Vol. 6 Issue 16


However, under conditions of oxygen‑deprivation and/or Fe‑depletion, prolyl hydroxylases fail to function, leading to the accumulation of HIF‑1a in the cell.177,178 HIF‑1a is then able to translocate to the nucleus where it binds to HIF‑1b to form the HIF‑1 complex.176,179 Once assembled, HIF‑1 can regulate a number of genes by binding to their hypoxia‑responsive element (HRE) located in the promoter or enhancer.180 For example, TfR1 is transcriptionally upregulated by HIF‑1180,181 which then functions to increase intracellular Fe levels. Also targeted by this transcription factor is Ndrg‑1 (see section on Ndrg-1),182 which has a role in differentiation183 and may be involved in regulating the cell cycle.184 Under conditions of severe hypoxia, HIF‑1 can also induce apoptotic pathways by stabilizing p53 expression185 and up‑regulating pro‑apoptotic factors such as BNIP3 (Fig. 4).186,187 Interestingly, BNIP3 can also be induced in a HIF‑1‑independent manner in response to Fe‑depletion.188,189 This is thought to be mediated by the transcripFigure 4. The demonstrated and potential effects of iron (Fe) deprivation on cell cycle arrest, apoptosis, metastasis and growth suppression. Iron‑regulatory protein (IRP)‑RNA binding tion factor, pleomorphic adenoma gene like 2 (PLAGL2).188,189 Cells treated with DFO respond activity and the expression of transforming growth factor‑b (TGF‑b1), p53, p38 mitogen activated protein kinase (MAPK) and hypoxia‑inducible factor‑1a (HIF‑1a) are increased by increasing the nuclear expression of PLAGL2188 upon Fe‑depletion. Increased RNA‑binding activity of the IRPs acts to stabilize CDC14A that is then able to induce BNIP3 expression in a mRNA, which may promote its translation, although an elevation in CDC14 protein after HIF‑1‑independent manner, by increasing BNIP3 Fe‑depletion has not been shown. CDC14A can de‑phosphorylate cdk substrates and promoter activity.189 Indeed, transfection of mouse may act as another link to mediate the effect of Fe‑depletion on cell cycle progression. Iron‑depletion also leads to increased expression of TGF‑b1, which may up‑regulate the Balb/c3T3 fibroblasts with PLAGL2 inhibited prolifcdk inhibitor, p27Kip1, under some conditions. Additionally, reduced expression of the eration and led to the induction of apoptosis, perhaps p170 subunit of the eukaryotic initiation factor 3 (eIF3) upon Fe‑depletion also increases as a result of BNIP3 activation.189 It is important to p27Kip1 expression. The reduction in expression of some cyclins such as cyclin D1 and note that Fe‑depletion does not cause apoptosis via cyclin‑dependent kinase (cdk) 2 and/or 4 results in hypo‑phosphorylation of the retinoblas‑ BNIP3 alone, as BNIP3 siRNA transfected cells toma susceptibility product (pRb). This leads to reduced release of the E2F1 transcription incubated with Fe chelators also undergo apopfactor from pRb, preventing transactivation of genes essential for cell cycle progression. tosis.189 This indicates that Fe‑depletion activates a Upregulation of p53 may lead to the increased expression of its down‑ stream targets such as growth arrest and DNA damage‑inducible gene number of different apoptotic pathways and these are further discussed in the section on apoptosis. 45a (GADD45a) or cause apoptosis, although these effects are yet to be established. The depletion of Fe also mimics the hypoxic state, leading to increased expression The depletion of intracellular Fe has crucial reperof HIF‑1a, which acts on its downstream targets, p53, BNIP3 and N‑myc downstream regu‑ cussions, resulting in the activation of HIF‑1a and lated gene‑1 (Ndrg‑1) etc. The increased expression of BNIP3 can lead to apoptosis through its numerous down‑stream targets, ultimately leading the mitochondrial pathway by increasing the expression of pro‑apoptotic molecules. Increased to cell cycle arrest, apoptosis, metastasis suppression Ndrg‑1 expression is known to result in the inhibition of tumor growth and metastasis. and inhibition of growth (Fig. 4).190 Although some CIP1/WAF1 Reduction of p21 protein expression may also be potentially involved in mediating of the effects of HIF‑1a up‑regulation are growth apoptosis observed upon Fe‑depletion. Iron‑depletion also up‑regulates p38 mitogen‑activated and angiogenesis (e.g., through vascular endothelial protein kinase (MAPK) which could be involved in cell cycle regulation and/or apoptosis. factor‑1), potent Fe chelators have been shown to override this and activate apoptotic pathways that are D1/cdk4 and cyclin E/cdk2 complexes. Further studies are obviously induced in part by HIF‑1a.101,190 Ndrg‑1. N‑myc downstream regulated gene 1 (Ndrg‑1) is also essential to determine the role of p38 MAPK in the Fe‑depletion mediated decrease in cyclin D1125 expression that could be impor- known as differentiation related gene‑1 (Drg‑1) and Cap‑43 and is a recently identified metastasis suppressor gene that plays roles in cell tant in mediating the cell cycle arrest observed. HIF‑1a. Hypoxia inducible factor‑1 (HIF‑1) is a transcription differentiation and proliferation.182‑184,191‑194 Iron‑depletion markfactor that is activated under hypoxic conditions and acts to initiate edly up‑regulates Ndrg‑1 mRNA and protein expression in a number a signaling pathway leading to cell survival.174,175 This protein is of neoplastic cell types, and hence, may be significant for undercomposed of two subunits, a constitutively expressed b subunit and standing the mechanisms involved in the inhibition of proliferation an a subunit which is regulated by the hypoxic state.176 Under condi- by Fe chelators.102,110 High expression of Ndrg‑1 is associated with greater survival and tions of normal oxygen tension and Fe levels, HIF‑1a is regulated by prolyl hydroxylase enzymes177,178 which allow its binding to the von less aggressive tumors in prostate cancer patients.192 There is also Hippel‑Lindau (VHL) protein. This protein activates a ubiquitin E3 a significant inverse correlation of Ndrg‑1 expression with depth ligase resulting in the subsequent degradation of HIF‑1a.174,175,178 of invasion in pancreatic adenocarcinoma patients.194 Moreover, www.landesbioscience.com

Cell Cycle

1989


Ndrg‑1 expression was significantly reduced in breast and prostate cancer patients with lymph node or bone metastasis compared to those with localized disease.192,193 Studies examining the function of Ndrg‑1 in a number of different cancers both in vitro and in vivo have found that overexpression of this protein results in smaller tumors that are less aggressive.183,184,192,194 Hence, Ndrg‑1 up‑regulation after Fe chelation may, in part, be an important mediator of the inhibition of proliferation observed after treatment with these agents. The expression of Ndrg‑1 in normal breast epithelial cells was found to be biphasic, being highest during G1 and G2/M phases and lowest during the S‑phase, suggesting a potential role in regulating the cell cycle.184 In contrast, in breast cancer cells, this biphasic expression was abolished, with Ndrg‑1 levels remaining constant throughout the cell cycle.184 Analysis of the nucleotide sequence of the Ndrg‑1 promoter using Genomatix Suite 3.0 software (Genomatix, Munchen, Germany) has revealed a motif for the transcription factor E2F1 (Kovacevic, unpublished results), which plays an important role in the G1 to S‑phase transition. Collectively, this evidence suggests a possible role for Ndrg‑1 in cell cycle regulation. Another important link between Ndrg‑1 and the cell cycle comes from studies examining the relationship between Ndrg‑1 and p53, which suggests that Ndrg‑1 is necessary for p53‑dependent apoptosis.191 In fact, this latter study found that Ndrg‑1 was induced by p53 following DNA damage,191 but this observation has not been repeated in other investigations.102 The precise molecular targets of Ndrg‑1 have yet to be identified, but a recent report suggests that it down‑regulates the expression of activating transcription factor 3 (ATF3)195 which is a stress‑inducible molecule thought to play a role in cell cycle progression and apoptosis.195,196 There is evidence that ATF3 expression is implicated in cancer development,196,197 and thus, its down‑regulation by Ndrg‑1 could be important for the growth and metastasis suppressor function of this latter molecule. Furthermore, there is work demonstrating that Fe‑depletion by DFO can up‑regulate ATF3.198 This latter observation appears somewhat paradoxical, as the observed up‑regulation of Ndrg‑1 after Fe‑depletion102,110 would be expected to downregulate ATF3 levels. Recently, Ndrg‑1 was found to be upregulated by DFO and 311 in a range of neoplastic cell types in an Fe‑dependent but p53‑independent manner (Fig. 4).102 The effect of Fe‑deprivation on up‑regulated Ndrg‑1 levels in HeLa cells was confirmed by others in later studies using the chelator, mimosine.110 Both HIF‑1a‑dependent and ‑independent mechanisms were found to be responsible for the up‑regulation of Ndrg‑1 following Fe‑depletion.102 The addition of Fe salts to cells after Fe chelation was found to reverse Ndrg‑1 upregulation, confirming that Fe‑depletion increased Ndrg‑1 expression.102 Interestingly, the degree of Ndrg‑1 up‑regulation was proportional to the efficacy of the anti‑proliferative activity of the chelator assessed, with the ligand di‑2‑pyridylketone‑4,4,‑dimethyl‑ 3‑thiosemicarbazone (Dp44mT) being particularly effective.102 Again, the effect of Dp44mT was mediated by its ability to bind Fe, as an analogue of this compound (i.e., Dp2mT) that does not chelate Fe had no significant anti‑proliferative activity93 nor the ability to upregulate Ndrg‑1.102 Since Ndrg‑1 potentially plays an important role in cell cycle regulation, its up‑regulation following Fe‑depletion may be one mechanism by which Fe chelators lead to cell cycle arrest and apoptosis. CDC14A. As already discussed, IRPs are important for controlling Fe homeostasis via IREs in the 3'‑ or 5'‑UTRs in mRNAs encoding molecules involved in Fe metabolism.17 More recently, Sanchez 1990

et al.55 have identified a novel IRE in the 3' UTR of CDC14A mRNA that binds IRPs. Iron‑depletion using DFO leads to increased expression of the CDC14A transcript that contains the IRE (Fig. 4).55 Interestingly, CDC14A has been found to de‑phosphorylate cdk substrates such as p27Kip1 and cyclin E, which are critical for the transition from G1 to S‑phase of the cell cycle.199 Hence, it was suggested that CDC14A plays a potential role in the cell cycle arrest observed after Fe‑depletion. However, it should be noted that the effect of Fe‑depletion on CDC14A expression was only reported at the mRNA level. Further investigations are essential to determine whether Fe‑depletion affects the protein expression of CDC14A to confirm the role of this molecule in Fe‑regulated cell cycle control. Apoptosis. Apoptosis involves an orchestrated series of biochemical events leading to cell death.200 Cell cycle regulatory molecules, such as p53, p21CIP1/WAF1 and GADD45 have been shown to play a key role in apoptosis.145,201 Several studies using a range of Fe chelators have demonstrated their ability to induce apoptosis,13 even though the exact signaling pathways that lead to this event upon Fe‑depletion are not completely understood. For instance, DFO has been demonstrated to induce apoptosis in a number of cancer cell lines including ovarian cancer,202 neuroblastoma,203 Kaposi’s sarcoma,122 malignant oral keratinocytes169 and cervical carcinomas.204 Other chelators, including Triapine,205 311,90 Tachpyridine,132,206,207 O‑Trensox208 and Dp44mT93 have also been shown to induce apoptosis in a variety of neoplastic cell types both in vitro and in vivo. The process of apoptosis is controlled by two distinct mechanisms, known as the death receptor (extrinsic) and mitochondrial (intrinsic) pathways.200 Iron chelators have been shown to induce cell death through the activation of apoptosis associated with the mitochondrial pathway.200 The mitochondrial pathway is triggered by a number of stimuli, such as DNA damage, ischemia and oxidative stress.200 This pathway is initialized with the permeabilization of the mitochondrial outer membrane leading to protein release, such as cytochrome c and apoptosis‑inducing factor.200 The release of cytochrome c leads to the induction of Apaf‑1 that activates caspase‑9 by the formation of the apoptosome. Caspase‑9 then proceeds to activate caspases‑3 and ‑7 resulting in the induction of apoptosis.200 Permeability of the mitochondrial membrane is regulated by the Bcl‑2 family of proteins that consist of pro‑ and anti‑apoptotic members.200 The pro‑apoptotic molecules include Bax, Bid, Bad, Puma and Bim, whereas the anti‑apoptotic members include Bcl‑2 and Bcl‑xL. Apoptosis induced by p53 is mediated through the mitochondrial pathway and is linked to pro‑apoptotic signals directed from certain Bcl‑2 members. For example, Bax is a p53‑induced pro‑apoptotic molecule and the loss of p53 (which is common in human tumors) results in a decrease of Bax activity.209,210 A number of studies investigating the ability of Fe chelators to induce apoptosis examined their effects on various caspases. It has been demonstrated that incubation of cells with DFO increased the activity of caspase‑3, ‑8 and ‑9,169,202,211 while Dp44mT markedly increased the activity of caspase‑3.93 More importantly, numerous investigations have revealed that Fe chelators induce apoptosis through the mitochondrial pathway. For example, activation of Bid was necessary for Triapine®‑induced apoptosis.205 Furthermore, apoptosis mediated by Tachpyridine was not prevented by blocking the CD95 death receptor pathway with a Fas‑associated death domain protein dominant‑negative mutant.206 In addition, this chelator‑mediated cell death was blocked in cells pre-treated with Bcl‑XL and a dominant‑negative caspase‑9 expression vector.206

Cell Cycle

2007; Vol. 6 Issue 16


Induction of apoptosis by Dp44mT and DFO was accompanied by a decrease in Bcl‑2 expression, an increase in Bax and also cytochrome c efflux from the mitochondrion.93,169 More recently, and similarly, DFO has been shown to induce apoptosis through down‑regulation of the Bcl‑2 protein and upregulation of Bax in malignant oral keratinocytes.169 It has been reported that incubation of cells with DFO leads to the nuclear accumulation of PLAGL2 which results in the expression of the pro‑apoptotic factor, BNIP3.189 The over‑expression of BNIP3 increases the levels of the downstream targets, Bax and Bak, which leads to cytochrome c release and apoptosis (Fig. 4).212 Despite p53 accumulation and the possible association between p53‑induced apoptosis and Bcl‑2, there has been no direct evidence of a p53‑dependent apoptosis signaling pathway being involved after Fe depletion. In fact, incubation of cells with Tachpyridine led to rapid accumulation of p53 and death but this did not require p53 activation.132 In contrast, other mechanisms have been proposed where Fe‑deprivation activates p38 and ERK MAPK to transduce signals for induction of the apoptotic cascade.169 Hence, it remains unclear whether p53 accumulation upon Fe‑deprivation is necessary for apoptosis.

Conclusions Appropriate intracellular Fe levels are a critical determinant for cell cycle progression. Cellular Fe‑depletion results in G1/S arrest and apoptosis which is probably mediated through the coordinated activity of a number of molecules. These include RR, p53, cyclins (e.g., cyclin D1), cdks (i.e., cdk2 and cdk4), p21CIP1/WAF1, HIF‑1a, Ndrg‑1, CDC14A, p38 MAPK etc. While individual protein “players” have been identified to play some role in regulating the cell cycle after Fe‑depletion, it is still unclear what cellular signaling networks are involved in all the responses observed (Fig. 4). Moreover, it is unknown how the players interact to coordinate the response to induce a G1/S arrest and this remains an important goal for future research. References 1. Hershko C. Control of disease by selective iron depletion: A novel therapeutic strategy utilizing iron chelators. Baillieres Clin Haematol 1994; 7:965‑1000. 2. Buss JL, Greene BT, Turner J, Torti FM, Torti SV. Iron chelators in cancer chemotherapy. Curr Top Med Chem 2004; 4:1623‑35. 3. Andrews NC. Disorders of iron metabolism. N Engl J Med 1999; 341:1986‑95. 4. Thelander L, Graslund A, Thelander M. Continual presence of oxygen and iron required for mammalian ribonucleotide reduction: Possible regulation mechanism. Biochem Biophys Res Commun 1983; 110:859‑65. 5. Nyholm S, Mann GJ, Johansson AG, Bergeron RJ, Graslund A, Thelander L. Role of ribonucleotide reductase in inhibition of mammalian cell growth by potent iron chelators. J Biol Chem 1993; 268:26200‑5. 6. Thelander L, Reichard P. Reduction of ribonucleotides. Annu Rev Biochem 1979; 48:133‑58. 7. Lucas JJ, Szepesi A, Domenico J, Takase K, Tordai A, Terada N, Gelfand EW. Effects of iron‑depletion on cell cycle progression in normal human T lymphocytes: Selective inhibition of the appearance of the cyclin A‑associated component of the p33cdk2 kinase. Blood 1995; 86:2268‑80. 8. Brodie C, Siriwardana G, Lucas J, Schleicher R, Terada N, Szepesi A, Gelfand E, Seligman P. Neuroblastoma sensitivity to growth inhibition by deferrioxamine: Evidence for a block in G1 phase of the cell cycle. Cancer Res 1993; 53:3968‑75. 9. Kwok JC, Richardson DR. The iron metabolism of neoplastic cells: Alterations that facilitate proliferation? Crit Rev Oncol Hematol 2002; 42:65‑78. 10. Le NT, Richardson DR. The role of iron in cell cycle progression and the proliferation of neoplastic cells. Biochim Biophys Acta 2002; 1603:31‑46. 11. Renton FJ, Jeitner TM. Cell cycle‑dependent inhibition of the proliferation of human neural tumor cell lines by iron chelators. Biochem Pharmacol 1996; 51:1553‑61. 12. Cook JD, SB, Baynes RD. Iron deficiency: The global perspective. Adv Exp Med Biol 1994; 356:219‑28. 13. Hileti DPP, Hoffbrand AV. Iron chelators induce apoptosis in proliferating cells. Br J Haematol 1995; 89:181‑7. 14. Buss JL, Torti FM, Torti SV. The role of iron chelation in cancer therapy. Curr Med Chem 2003; 10:1021‑34.

www.landesbioscience.com

15. Kalinowski DS, Richardson DR. The evolution of iron chelators for the treatment of iron overload disease and cancer. Pharmacol Rev 2005; 57:547‑83. 16. Dunn LL, Suryo‑Rahmanto Y, Richardson DR. Iron uptake and metabolism in the new millennium. Trends Cell Biol 2007; 17:93‑100. 17. Hentze MW, Muckenthaler MU, Andrews NC. Balancing acts: Molecular control of mammalian iron metabolism. Cell 2004; 117:285‑97. 18. Richardson DR, Ponka P. The molecular mechanisms of the metabolism and transport of iron in normal and neoplastic cells. Biochim Biophys Acta 1997; 1331:1‑40. 19. Latunde‑Dada GO, Van der Westhuizen J, Vulpe CD, Anderson GJ, Simpson RJ, McKie AT. Molecular and functional roles of duodenal cytochrome B (Dcytb) in iron metabolism. Blood Cells Mol Dis 2002; 29:356‑60. 20. Gunshin H, Starr CN, Direnzo C, Fleming MD, Jin J, Greer EL, Sellers VM, Galica SM, Andrews NC. Cybrd1 (duodenal cytochrome b) is not necessary for dietary iron absorption in mice. Blood 2005; 106:2879‑83. 21. Shayeghi M, Latunde‑Dada GO, Oakhill JS, Laftah AH, Takeuchi K, Halliday N, Khan Y, Warley A, McCann FE, Hider RC, Frazer DM, Anderson GJ, Vulpe CD, Simpson RJ, McKie AT. Identification of an intestinal heme transporter. Cell 2005; 122:789‑801. 22. Qiu A, Jansen M, Sakaris A, Min SH, Chattopadhyay S, Tsai E, Sandoval C, Zhao R, Akabas MH, Goldman ID. Identification of an intestinal folate transporter and the molecular basis for hereditary folate malabsorption. Cell 2006; 127:917‑28. 23. Andrews NC. When is a heme transporter not a heme transporter? When it’s a folate transporter. Cell Metab 2007; 5:5‑6. 24. Watts RN, Ponka P, Richardson DR. Effects of nitrogen monoxide and carbon monoxide on molecular and cellular iron metabolism: Mirror‑image effector molecules that target iron. Biochem J 2003; 369:429‑40. 25. Harrison PM, Arosio P. The ferritins: Molecular properties, iron storage function and cellular regulation. Biochim Biophys Acta 1996; 1275:161‑203. 26. Richardson DR. Molecular mechanisms of iron uptake by cells and the use of iron chelators for the treatment of cancer. Curr Med Chem 2005; 12:2711‑29. 27. Vulpe CD, Kuo YM, Murphy TL, Cowley L, Askwith C, Libina N, Gitschier J, Anderson GJ. Hephaestin, a ceruloplasmin homologue implicated in intestinal iron transport, is defective in the sla mouse. Nat Genet 1999; 21:195‑9. 28. Donovan A, Brownlie A, Zhou Y, Shepard J, Pratt SJ, Moynihan J, Paw BH, Drejer A, Barut B, Zapata A, Law TC, Brugnara C, Lux SE, Pinkus GS, Pinkus JL, Kingsley PD, Palis J, Fleming MD, Andrews NC, Zon LI. Positional cloning of zebrafish ferroportin1 identifies a conserved vertebrate iron exporter. Nature 2000; 403:776‑81. 29. Nemeth E, Tuttle MS, Powelson J, Vaughn MB, Donovan A, Ward DM, Ganz T, Kaplan J. Hepcidin regulates cellular iron efflux by binding to ferroportin and inducing its internalization. Science 2004; 306:2090‑3. 30. Papanikolaou G, Samuels ME, Ludwig EH, MacDonald ML, Franchini PL, Dube MP, Andres L, MacFarlane J, Sakellaropoulos N, Politou M, Nemeth E, Thompson J, Risler JK, Zaborowska C, Babakaiff R, Radomski CC, Pape TD, Davidas O, Christakis J, Brissot P, Lockitch G, Ganz T, Hayden MR, Goldberg YP. Mutations in HFE2 cause iron overload in chromosome 1q‑linked juvenile hemochromatosis. Nat Genet 2004; 36:77‑82. 31. Niederkofler V, Salie R, Arber S. Hemojuvelin is essential for dietary iron sensing, and its mutation leads to severe iron overload. J Clin Invest 2005; 115:2180‑6. 32. Morgan EH. Transferrin biochemistry, physiology and clinical significance. Mol Aspects Med 1981; 4:1‑123. 33. Morgan EH, Oates PS. Mechanisms and regulation of intestinal iron absorption. Blood Cells Mol Dis 2002; 29:384‑99. 34. Richardson DR. Iron chelators as therapeutic agents for the treatment of cancer. Crit Rev Oncol Hematol 2002; 42:267‑81. 35. Klausner RD, Ashwell G, van Renswoude J, Harford JB, Bridges KR. Binding of apotransferrin to K562 cells: Explanation of the transferrin cycle. Proc Natl Acad Sci USA 1983; 80:2263‑6. 36. Ohgami RS, Campagna DR, Greer EL, Antiochos B, McDonald A, Chen J, Sharp JJ, Fujiwara Y, Barker JE, Fleming MD. Identification of a ferrireductase required for efficient transferrin‑dependent iron uptake in erythroid cells. Nat Genet 2005; 37:1264‑9. 37. Fleming MD, Trenor IIIrd CC, Su MA, Foernzler D, Beier DR, Dietrich WF, Andrews NC. Microcytic anaemia mice have a mutation in Nramp2, a candidate iron transporter gene. Nat Genet 1997; 16:383‑6. 38. Kawabata H, Yang R, Hirama T, Vuong PT, Kawano S, Gombart AF, Koeffler HP. Molecular cloning of transferrin receptor 2: A new member of the transferrin receptor‑like family. J Biol Chem 1999; 274:20826‑32. 39. Kawabata H, Germain RS, Vuong PT, Nakamaki T, Said JW, Koeffler HP. Transferrin receptor 2‑alpha supports cell growth both in iron‑chelated cultured cells and in vivo. J Biol Chem 2000; 275:16618‑25. 40. Goswami T, Andrews NC. Hereditary hemochromatosis protein, HFE, interaction with transferrin receptor 2 suggests a molecular mechanism for mammalian iron sensing. J Biol Chem 2006; 281:28494‑8. 41. Mattman A, Huntsman D, Lockitch G, Langlois S, Buskard N, Ralston D, Butterfield Y, Rodrigues P, Jones S, Porto G, Marra M, De Sousa M, Vatcher G. Transferrin receptor 2 (TfR2) and HFE mutational analysis in non-C282Y iron overload: Identification of a novel TfR2 mutation. Blood 2002; 100:1075‑7. 42. Hentze MW, Kuhn LC. Molecular control of vertebrate iron metabolism: mRNA‑based regulatory circuits operated by iron, nitric oxide, and oxidative stress. Proc Natl Acad Sci USA 1996; 93:8175‑82. 43. Levy JE, Jin O, Fujiwara Y, Kuo F, Andrews NC. Transferrin receptor is necessary for development of erythrocytes and the nervous system. Nat Genet 1999; 21:396‑9.

Cell Cycle

1991

Iron and the Cell Cycle 44. Robb AD, Ericsson M, Wessling‑Resnick M. Transferrin receptor 2 mediates a biphasic pattern of transferrin uptake associated with ligand delivery to multivesicular bodies. Am J Physiol Cell Physiol 2004; 287:C1769‑75. 45. Jacobs A. Low molecular weight intracellular iron transport compounds. Blood 1977; 50:433‑9. 46. Richardson DR, Ponka P, Vyoral D. Distribution of iron in reticulocytes after inhibition of heme synthesis with succinylacetone: Examination of the intermediates involved in iron metabolism. Blood 1996; 87:3477‑88. 47. Ponka P, Sheftel AD, Zhang AS. Iron targeting to mitochondria in erythroid cells. Biochem Soc Trans 2002; 30:735‑8. 48. Sheftel AD, Zhang AS, Brown CM, Shirihai OS, Ponka P. Direct interorganellar transfer of iron from endosome to mitochondrion. Blood 2007, [Epub ahead of print]. 49. Ponka P. Tissue‑specific regulation of iron metabolism and heme synthesis: Distinct control mechanisms in erythroid cells. Blood 1997; 89:1‑25. 50. Napier I, Ponka P, Richardson DR. Iron trafficking in the mitochondrion: Novel pathways revealed by disease. Blood 2005; 105:1867‑74. 51. Shaw GC, Cope JJ, Li L, Corson K, Hersey C, Ackermann GE, Gwynn B, Lambert AJ, Wingert RA, Traver D, Trede NS, Barut BA, Zhou Y, Minet E, Donovan A, Brownlie A, Balzan R, Weiss MJ, Peters LL, Kaplan J, Zon LI, Paw BH. Mitoferrin is essential for erythroid iron assimilation. Nature 2006; 440:96‑100. 52. Levi S, Corsi B, Bosisio M, Invernizzi R, Volz A, Sanford D, Arosio P, Drysdale J. A human mitochondrial ferritin encoded by an intronless gene. J Biol Chem 2001; 276:24437‑40. 53. Nie G, Sheftel AD, Kim SF, Ponka P. Overexpression of mitochondrial ferritin causes cytosolic iron depletion and changes cellular iron homeostasis. Blood 2005; 105:2161‑7. 54. Eisenstein RS, Ross KL. Novel roles for iron regulatory proteins in the adaptive response to iron deficiency. J Nutr 2003; 133:1510S‑6S. 55. Sanchez M, Galy B, Dandekar T, Bengert P, Vainshtein Y, Stolte J, Muckenthaler MU, Hentze MW. Iron regulation and the cell cycle: Identification of an iron‑responsive element in the 3’‑untranslated region of human cell division cycle 14A mRNA by a refined microarray‑based screening strategy. J Biol Chem 2006; 281:22865‑74. 56. Guo B, Phillips JD, Yu Y, Leibold EA. Iron regulates the intracellular degradation of iron regulatory protein 2 by the proteasome. J Biol Chem 1995; 270:21645‑51. 57. Neckers LM, Cossman J. Transferrin receptor induction in mitogen‑stimulated human T lymphocytes is required for DNA synthesis and cell division and is regulated by interleukin 2. Proc Natl Acad Sci USA 1983; 80:3494‑8. 58. O’Donnell KAYD, Zeller KI, Kim JW, Racke F, Thomas‑Tikhonenko A, Dang CV. Activation of transferrin receptor 1 by c‑Myc enhances cellular proliferation and tumorigenesis. Mol Cell Biol 2006; 26:2373‑86. 59. Kalinowski DS, Richardson DR. Iron chelators and differing modes of action and toxicity: The changing face of iron chelation therapy. Chem Res Toxicol 2007; 20:715‑20. 60. Larrick JW, Cresswell P. Modulation of cell surface iron transferrin receptors by cellular density and state of activation. J Supramol Struct 1979; 11:579‑86. 61. Richardson DR, Baker E. The uptake of iron and transferrin by the human malignant melanoma cell. Biochim Biophys Acta 1990; 1053:1‑12. 62. Richardson DR, Baker E. The effect of desferrioxamine and ferric ammonium citrate on the uptake of iron by the membrane iron‑binding component of human melanoma cells. Biochim Biophys Acta 1992; 1103:275‑80. 63. Elford HL, Freese M, Passamani E, Morris HP. Ribonucleotide reductase and cell proliferation. I. Variations of ribonucleotide reductase activity with tumor growth rate in a series of rat hepatomas. J Biol Chem 1970; 245:5228‑33. 64. Takeda E, Weber G. Role of ribonucleotide reductase in expression in the neoplastic program. Life Sci 1981; 28:1007‑14. 65. Witt L, Yap T, Blakley RL. Regulation of ribonucleotide reductase activity and its possible exploitation in chemotherapy. Adv Enzyme Regul 1978; 17:157‑71. 66. Donfrancesco A, Deb G, Dominici C, Pileggi D, Castello MA, Helson L. Effects of a single course of deferoxamine in neuroblastoma patients. Cancer Res 1990; 50:4929‑30. 67. Estrov Z, Tawa A, Wang XH, Dube ID, Sulh H, Cohen A, Gelfand EW, Freedman MH. In vitro and in vivo effects of deferoxamine in neonatal acute leukemia. Blood 1987; 69:757‑61. 68. Kaplinsky C, Estrov Z, Freedman MH, Gelfand EW, Cohen A. Effect of deferoxamine on DNA synthesis, DNA repair, cell proliferation, and differentiation of HL‑60 cells. Leukemia 1987; 1:437‑41. 69. Blatt J, Taylor SR, Stitely S. Mechanism of antineuroblastoma activity of deferoxamine in vitro. J Lab Clin Med 1988; 112:433‑6. 70. Blatt J, Stitely S. Antineuroblastoma activity of desferoxamine in human cell lines. Cancer Res 1987; 47:1749‑50. 71. Richardson D, Ponka P, Baker E. The effect of the iron(III) chelator, desferrioxamine, on iron and transferrin uptake by the human malignant melanoma cell. Cancer Res 1994; 54:685‑9. 72. Darnell G, Richardson DR. The potential of iron chelators of the pyridoxal isonicotinoyl hydrazone class as effective antiproliferative agents III: The effect of the ligands on molecular targets involved in proliferation. Blood 1999; 94:781‑92. 73. Yu Y, Wong J, Lovejoy DB, Kalinowski DS, Richardson DR. Chelators at the cancer coalface: Desferrioxamine to Triapine and beyond. Clin Cancer Res 2006; 12:6876‑83. 74. Finch RA, Liu M, Grill SP, Rose WC, Loomis R, Vasquez KM, Cheng Y, Sartorelli AC. Triapine (3‑aminopyridine‑2‑carboxaldehyde‑ thiosemicarbazone): A potent inhibitor of ribonucleotide reductase activity with broad spectrum antitumor activity. Biochem Pharmacol 2000; 59:983‑91.

1992

75. Torti SV, Torti FM, Whitman SP, Brechbiel MW, Park G, Planalp RP. Tumor cell cytotoxicity of a novel metal chelator. Blood 1998; 92:1384‑9. 76. Rakba N, Loyer P, Gilot D, Delcros JG, Glaise D, Baret P, Pierre JL, Brissot P, Lescoat G. Antiproliferative and apoptotic effects of O‑Trensox, a new synthetic iron chelator, on differentiated human hepatoma cell lines. Carcinogenesis 2000; 21:943‑51. 77. Ponka P, Borova J, Neuwirt J, Fuchs O. Mobilization of iron from reticulocytes: Identification of pyridoxal isonicotinoyl hydrazone as a new iron chelating agent. FEBS Lett 1979; 97:317‑21. 78. Ponka P, Borova J, Neuwirt J, Fuchs O, Necas E. A study of intracellular iron metabolism using pyridoxal isonicotinoyl hydrazone and other synthetic chelating agents. Biochim Biophys Acta 1979; 586:278‑97. 79. Richardson DR, Tran EH, Ponka P. The potential of iron chelators of the pyridoxal isonicotinoyl hydrazone class as effective antiproliferative agents. Blood 1995; 86:4295‑306. 80. Chaston TB, Watts RN, Yuan J, Richardson DR. Potent antitumor activity of novel iron chelators derived from di‑2‑pyridylketone isonicotinoyl hydrazone involves fenton‑derived free radical generation. Clin Cancer Res 2004; 10:7365‑74. 81. Becker E, Lovejoy DB, Greer J, Watts R, Richardson DR. Novel aroylhydrazone iron chelators differ in their iron chelation efficacy and anti‑proliferative activity: Identification of a new class of potential anti‑proliferative agents. Br J Pharmacol 2003; 138:819‑30. 82. Kalinowski DS, Yu Y, Sharpe PS, Mohammad Islam M, Liao YT, Lovejoy DB, Kumar N, Bernhardt PV, Richardson DR. Design, synthesis and characterization of novel iron chelators: Structure-activity relationships of the 2‑benzoylpyridine thiosemicarbazone series and their 3‑nitrobenzoyl analogs as potent anti‑tumor agents. J Med Chem 2007, (In Press). 83. Lovejoy DB, Richardson DR. Novel “hybrid” iron chelators derived from aroylhydrazones and thiosemicarbazones demonstrate high anti‑proliferative activity that is selective for tumor cells. Blood 2002; 100:666‑76. 84. Whitnall M, Howard J, Ponka P, Richardson DR. A class of iron chelators with a wide spectrum of potent antitumor activity that overcomes resistance to chemotherapeutics. Proc Natl Acad Sci USA 2006; 103:14901‑6. 85. Yen Y, Margolin K, Doroshow J, Fishman M, Johnson B, Clairmont C, Sullivan D, Sznol M. A phase I trial of 3‑aminopyridine‑2‑carboxaldehyde thiosemicarbazone in combination with gemcitabine for patients with advanced cancer. Cancer Chemother Pharmacol 2004; 54:331‑42. 86. Yee KW, Cortes J, Ferrajoli A, Garcia‑Manero G, Verstovsek S, Wierda W, Thomas D, Faderl S, King I, O’Brien SM, Jeha S, Andreeff M, Cahill A, Sznol M, Giles FJ. Triapine and cytarabine is an active combination in patients with acute leukemia or myelodysplastic syndrome. Leuk Res 2006; 30:813‑22. 87. Gojo I, Tidwell ML, Greer J, Takebe N, Seiter K, Pochron MF, Johnson B, Sznol M, Karp JE. Phase I and pharmacokinetic study of Triapine((R)), a potent ribonucleotide reductase inhibitor, in adults with advanced hematologic malignancies. Leuk Res 2007; 31:1173‑81. 88. Turner J, Koumenis C, Kute TE, Planalp RP, Brechbiel MW, Beardsley D, Cody B, Brown KD, Torti FM, Torti SV. Tachpyridine, a metal chelator, induces G2 cell‑cycle arrest, activates checkpoint kinases, and sensitizes cells to ionizing radiation. Blood 2005; 106:3191‑9. 89. Richardson DR, Bernhardt PV. Crystal and molecular structure of 2‑hydroxy‑1‑naphthylaldehyde isonicotinoyl hydrazone (NIH) and its iron(III) complex: An iron chelator with anti‑proliferative activity. J Biol Inorg Chem 1999; 4:266‑73. 90. Richardson DR, Milnes K. The potential of iron chelators of the pyridoxal isonicotinoyl hydrazone class as effective antiproliferative agents II: The mechanism of action of ligands derived from salicylaldehyde benzoyl hydrazone and 2‑hydroxy‑1‑naphthylaldehyde benzoyl hydrazone. Blood 1997; 89:3025‑38. 91. Green DA, Antholine WE, Wong SJ, Richardson DR, Chitambar CR. Inhibition of malignant cell growth by 311, a novel iron chelator of the pyridoxal isonicotinoyl hydrazone class: Effect on the R2 subunit of ribonucleotide reductase. Clin Cancer Res 2001; 7:3574‑9. 92. Becker EM, Lovejoy DB, Greer JM, Watts R, Richardson DR. Identification of the di‑pyridyl ketone isonicotinoyl hydrazone (PKIH) analogues as potent iron chelators and anti‑tumour agents. Br J Pharmacol 2003; 138:819‑30. 93. Yuan J, Lovejoy DB, Richardson DR. Novel di‑2‑pyridyl‑derived iron chelators with marked and selective antitumor activity: In vitro and in vivo assessment. Blood 2004; 104:1450‑8. 94. Richardson DR, Sharpe PC, Lovejoy DB, Senaratne D, Kalinowski DS, Islam M, Bernhardt PV. Dipyridyl thiosemicarbazone chelators with potent and selective anti‑tumor activity form iron complexes with marked redox activity. J Med Chem 2006; 49:6510‑21. 95. Boldt DH. New perspectives on iron: An introduction. Am J Med Sci 1999; 318:207‑12. 96. Sherr CJ. G1 phase progression: Cycling on cue. Cell 1994; 79:551‑5. 97. Zetterberg A, Larsson O, Wiman KG. What is the restriction point? Curr Opin Cell Biol 1995; 7:835‑42. 98. Golias CH, Charalabopoulos A, Charalabopoulos K. Cell proliferation and cell cycle control: A mini review. Int J Clin Pract 2004; 58:1134‑41. 99. Sherr CJ, Roberts JM. CDK inhibitors: Positive and negative regulators of G1‑phase progression. Genes Dev 1999; 13:1501‑12. 100. Reed SI. Control of the G1/S transition. Cancer Surv 1997; 29:7‑23. 101. Gao J, Richardson DR. The potential of iron chelators of the pyridoxal isonicotinoyl hydrazone class as effective antiproliferative agents, IV: The mechanisms involved in inhibiting cell‑cycle progression. Blood 2001; 98:842‑50. 102. Le NT, Richardson DR. Iron chelators with high antiproliferative activity up‑regulate the expression of a growth inhibitory and metastasis suppressor gene: A link between iron metabolism and proliferation. Blood 2004; 104:2967‑75.

Cell Cycle

2007; Vol. 6 Issue 16

Iron and the Cell Cycle 103. Kulp KS, Green SL, Vulliet PR. Iron deprivation inhibits cyclin‑dependent kinase activity and decreases cyclin D/CDK4 protein levels in asynchronous MDA‑MB‑453 human breast cancer cells. Exp Cell Res 1996; 229:60‑8. 104. Ashcroft M, Taya Y, Vousden KH. Stress signals utilize multiple pathways to stabilize p53. Mol Cell Biol 2000; 20:3224‑33. 105. Fukuchi K, Tomoyasu S, Watanabe H, Kaetsu S, Tsuruoka N, Gomi K. Iron deprivation results in an increase in p53 expression. Biol Chem Hoppe Seyler 1995; 376:627‑30. 106. Liang SX, Richardson DR. The effect of potent iron chelators on the regulation of p53: Examination of the expression, localization and DNA‑binding activity of p53 and the transactivation of WAF1. Carcinogenesis 2003; 24:1601‑14. 107. Fu D, Richardson DR. Iron chelation and regulation of the cell‑cycle: Two mechanisms of post‑transcriptional regulation of the universal cyclin‑dependent kinase inhibitor p21CIP1/ WAF1 by iron depletion. Blood 2007, [Epub ahead of print]. 108. Wang G, Miskimins R, Miskimins WK. Mimosine arrests cells in G1 by enhancing the levels of p27(Kip1). Exp Cell Res 2000; 254:64‑71. 109. Wang G, Miskimins R, Miskimins WK. Regulation of p27(Kip1) by intracellular iron levels. Biometals 2004; 17:15‑24. 110. Dong Z, Arnold RJ, Yang Y, Park MH, Hrncirova P, Mechref Y, Novotny MV, Zhang JT. Modulation of differentiation‑related gene 1 expression by cell cycle blocker mimosine, revealed by proteomic analysis. Mol Cell Proteomics 2005; 4:993‑1001. 111. Terada N, Lucas JJ, Gelfand EW. Differential regulation of the tumor suppressor molecules, retinoblastoma susceptibility gene product (Rb) and p53, during cell cycle progression of normal human T cells. J Immunol 1991; 147:698‑704. 112. Jordan A, Reichard P. Ribonucleotide reductases. Annu Rev Biochem 1998; 67:71‑98. 113. Reichard P. From RNA to DNA, why so many ribonucleotide reductases? Science 1993; 260:1773‑7. 114. Jordan A, Pontis E, Atta M, Krook M, Gibert I, Barbe J, Reichard P. A second class I ribonucleotide reductase in Enterobacteriaceae: Characterization of the Salmonella typhimurium enzyme. Proc Natl Acad Sci USA 1994; 91:12892‑6. 115. Tanaka H, Arakawa H, Yamaguchi T, Shiraishi K, Fukuda S, Matsui K, Takei Y, Nakamura Y. A ribonucleotide reductase gene involved in a p53‑dependent cell‑cycle checkpoint for DNA damage. Nature 2000; 404:42‑9. 116. Byun DS, Chae KS, Ryu BK, Lee MG, Chi SG. Expression and mutation analyses of P53R2, a newly identified p53 target for DNA repair in human gastric carcinoma. Int J Cancer 2002; 98:718‑23. 117. Nakano K, Balint E, Ashcroft M, Vousden KH. A ribonucleotide reductase gene is a transcriptional target of p53 and p73. Oncogene 2000; 19:4283‑9. 118. Shao J, Zhou B, Zhu L, Qiu W, Yuan YC, Xi B, Yen Y. In vitro characterization of enzymatic properties and inhibition of the p53R2 subunit of human ribonucleotide reductase. Cancer Res 2004; 64:1‑6. 119. Tsimberidou AM, Alvarado Y, Giles FJ. Evolving role of ribonucleoside reductase inhibitors in hematologic malignancies. Expert Rev Anticancer Ther 2002; 2:437‑48. 120. Chaston TB, Lovejoy DB, Watts RN, Richardson DR. Examination of the antiproliferative activity of iron chelators: Multiple cellular targets and the different mechanism of action of triapine compared with desferrioxamine and the potent pyridoxal isonicotinoyl hydrazone analogue 311. Clin Cancer Res 2003; 9:402‑14. 121. Sartorelli AC, Agrawal KC, Moore EC. Mechanism of inhibition of ribonucleoside diphosphate reductase by a‑(N)‑heterocyclic aldehyde thiosemicarbazones. Biochem Pharmacol 1971; 20:3119‑23. 122. Simonart T, Degraef C, Andrei G, Mosselmans R, Hermans P, Van Vooren JP, Noel JC, Boelaert JR, Snoeck R, Heenen M. Iron chelators inhibit the growth and induce the apoptosis of Kaposi’s sarcoma cells and of their putative endothelial precursors. J Invest Dermatol 2000; 115:893‑900. 123. Aleem E, Kiyokawa H, Kaldis P. Cdc2‑cyclin E complexes regulate the G1/S phase transition. Nat Cell Biol 2005; 7:831‑6. 124. Kaldis P, Aleem E. Cell cycle sibling rivalry: Cdc2 vs Cdk2. Cell Cycle 2005; 4:1491‑4. 125. Nurtjahja‑Tjendraputra E, Fu D, Phang JM, Richardson DR. Iron chelation regulates cyclin D1 expression via the proteasome: A link to iron deficiency‑mediated growth suppression. Blood 2007; 109:4045‑54. 126. Burnworth B, Popp S, Stark HJ, Steinkraus V, Brocker EB, Hartschuh W, Birek C, Boukamp P. Gain of 11q/cyclin D1 overexpression is an essential early step in skin cancer development and causes abnormal tissue organization and differentiation. Oncogene 2006; 25:4399‑412. 127. Donnellan RC, R. Cyclin D1 and human neoplasia. Mol Pathol 1998; 51:1‑7. 128. Schrump DS, CA, Consoli U. Inhibition of lung cancer proliferation by antisense cyclin D. Cancer Gene Ther 1996; 3:131‑5. 129. Hollstein M, Sidransky D, Vogelstein B, Harris CC. p53 mutations in human cancers. Science 1991; 253:49‑53. 130. Hatakeyama M, Weinberg RA. The role of RB in cell cycle control. Prog Cell Cycle Res 1995; 1:9‑19. 131. Weinberg RA. The retinoblastoma protein and cell cycle control. Cell 1995; 81:323‑30. 132. Abeysinghe RD, Greene BT, Haynes R, Willingham MC, Turner J, Planalp RP, Brechbiel MW, Torti FM, Torti SV. p53‑independent apoptosis mediated by tachpyridine, an anti‑cancer iron chelator. Carcinogenesis 2001; 22:1607‑14. 133. Sun Y, Bian J, Wang Y, Jacobs C. Activation of p53 transcriptional activity by 1,10‑phenanthroline, a metal chelator and redox sensitive compound. Oncogene 1997; 14:385‑93. 134. Harper JW, Adami GR, Wei N, Keyomarsi K, Elledge SJ. The p21 Cdk‑interacting protein Cip1 is a potent inhibitor of G1 cyclin‑dependent kinases. Cell 1993; 75:805‑16.

www.landesbioscience.com

135. el‑Deiry WS, Tokino T, Velculescu VE, Levy DB, Parsons R, Trent JM, Lin D, Mercer WE, Kinzler KW, Vogelstein B. WAF1, a potential mediator of p53 tumor suppression. Cell 1993; 75:817‑25. 136. Xiong Y, Hannon GJ, Zhang H, Casso D, Kobayashi R, Beach D. p21 is a universal inhibitor of cyclin kinases. Nature 1993; 366:701‑4. 137. Luo Y, Hurwitz J, Massague J. Cell‑cycle inhibition by independent CDK and PCNA binding domains in p21Cip1. Nature 1995; 375:159‑61. 138. Waga S, Hannon GJ, Beach D, Stillman B. The p21 inhibitor of cyclin‑dependent kinases controls DNA replication by interaction with PCNA. Nature 1994; 369:574‑8. 139. Vidal A, Koff A. Cell‑cycle inhibitors: Three families united by a common cause. Gene 2000; 247:1‑15. 140. Delavaine L, La Thangue NB. Control of E2F activity by p21Waf1/Cip1. Oncogene 1999; 18:5381‑92. 141. Kitaura H, Shinshi M, Uchikoshi Y, Ono T, Iguchi‑Ariga SM, Ariga H. Reciprocal regulation via protein‑protein interaction between c‑Myc and p21(cip1/waf1/sdi1) in DNA replication and transcription. J Biol Chem 2000; 275:10477‑83. 142. Cheng M, Olivier P, Diehl JA, Fero M, Roussel MF, Roberts JM, Sherr CJ. The p21(Cip1) and p27(Kip1) CDK ‘inhibitors’ are essential activators of cyclin D‑dependent kinases in murine fibroblasts. Embo J 1999; 18:1571‑83. 143. Weiss RH. p21Waf1/Cip1 as a therapeutic target in breast and other cancers. Cancer Cell 2003; 4:425‑9. 144. Weiss RH, Marshall D, Howard L, Corbacho AM, Cheung AT, Sawai ET. Suppression of breast cancer growth and angiogenesis by an antisense oligodeoxynucleotide to p21(Waf1/ Cip1). Cancer Lett 2003; 189:39‑48. 145. Gartel AL, Tyner AL. Transcriptional regulation of the p21((WAF1/CIP1)) gene. Exp Cell Res 1999; 246:280‑9. 146. Wajapeyee N, Somasundaram K. Cell cycle arrest and apoptosis induction by activator protein 2alpha (AP‑2alpha) and the role of p53 and p21WAF1/CIP1 in AP‑2alpha‑mediated growth inhibition. J Biol Chem 2003; 278:52093‑101. 147. Tvrdik D, Dundr P, Povysil C, Pytlik R, Plankova M. Up‑regulation of p21WAF1 expression is mediated by Sp1/Sp3 transcription factors in TGFbeta1‑arrested malignant B cells. Med Sci Monit 2006; 12:BR227‑34. 148. Le NT, Richardson DR. Potent iron chelators increase the mRNA levels of the universal cyclin‑dependent kinase inhibitor p21(CIP1/WAF1), but paradoxically inhibit its translation: A potential mechanism of cell cycle dysregulation. Carcinogenesis 2003; 24:1045‑58. 149. Asher G, Bercovich Z, Tsvetkov P, Shaul Y, Kahana C. 20S proteasomal degradation of ornithine decarboxylase is regulated by NQO1. Mol Cell 2005; 17:645‑55. 150. Newman RMA, Mangold U, Koike C, Diah S, Schmidt M, Finely D, Zetter BR. Antizyme targets cyclin D1 for degradation. J Biol Chem 2004; 279:41504‑11. 151. Coleman ML, Marshall CJ, Olson MF. Ras promotes p21(Waf1/Cip1) protein stability via a cyclin D1‑imposed block in proteasome‑mediated degradation. Embo J 2003; 22:2036‑46. 152. Gazitt Y, Reddy SV, Alcantara O, Yang J, Boldt DH. A new molecular role for iron in regulation of cell cycling and differentiation of HL‑60 human leukemia cells: Iron is required for transcription of p21(WAF1/CIP1) in cells induced by phorbol myristate acetate. J Cell Physiol 2001; 187:124‑35. 153. Fukuchi K, Tomoyasu S, Watanabe H, Tsuruoka N, Gomi K. G1 accumulation caused by iron deprivation with deferoxamine does not accompany change of pRB status in ML‑1 cells. Biochim Biophys Acta 1997; 1357:297‑305. 154. Fan YBA, Weiss RH. An antisense oligodeoxynucleotide to p21(Waf1/Cip1) causes apoptosis in human breast cancer cells. Mol Cancer Ther 2003; 2:773‑82. 155. Dong Z, Zhang JT. EIF3 p170, a mediator of mimosine effect on protein synthesis and cell cycle progression. Mol Biol Cell 2003; 14:3942‑51. 156. Yoon G, Kim HJ, Yoon YS, Cho H, Lim IK, Lee JH. Iron chelation‑induced senescence‑like growth arrest in hepatocyte cell lines: Association of transforming growth factor beta1 (TGF‑beta1)‑mediated p27Kip1 expression. Biochem J 2002; 366:613‑21. 157. Carlson SG, Fawcett TW, Bartlett JD, Bernier M, Holbrook NJ. Regulation of the C/ EBP‑related gene gadd153 by glucose deprivation. Mol Cell Biol 1993; 13:4736‑44. 158. Abcouwer SF, Schwarz C, Meguid RA. Glutamine deprivation induces the expression of GADD45 and GADD153 primarily by mRNA stabilization. J Biol Chem 1999; 274:28645‑51. 159. Fanzo JC, Reaves SK, Cui L, Zhu L, Wu JY, Wang YR, Lei KY. Zinc status affects p53, gadd45, and c‑fos expression and caspase‑3 activity in human bronchial epithelial cells. Am J Physiol Cell Physiol 2001; 281:C751‑7. 160. Oh‑Hashi K, Maruyama W, Isobe K. Peroxynitrite induces GADD34, 45, and 153 VIA p38 MAPK in human neuroblastoma SH‑SY5Y cells. Free Radic Biol Med 2001; 30:213‑21. 161. Zerbini LF, Libermann TA. GADD45 deregulation in cancer: Frequently methylated tumor suppressors and potential therapeutic targets. Clin Cancer Res 2005; 11:6409‑13. 162. Kearsey JM, Coates PJ, Prescott AR, Warbrick E, Hall PA. Gadd45 is a nuclear cell cycle regulated protein which interacts with p21Cip1. Oncogene 1995; 11:1675‑83. 163. Vairapandi M, Balliet AG, Hoffman B, Liebermann DA. GADD45b and GADD45g are cdc2/cyclinB1 kinase inhibitors with a role in S and G2/M cell cycle checkpoints induced by genotoxic stress. J Cell Physiol 2002; 192:327‑38. 164. Bulavin DV, Kovalsky O, Hollander MC, Fornace Jr AJ. Loss of oncogenic H‑ras‑induced cell cycle arrest and p38 mitogen‑activated protein kinase activation by disruption of Gadd45a. Mol Cell Biol 2003; 23:3859‑71. 165. Hollander MC, Sheikh MS, Yu K, Zhan Q, Iglesias M, Woodworth C, Fornace Jr AJ. Activation of Gadd34 by diverse apoptotic signals and suppression of its growth inhibitory effects by apoptotic inhibitors. Int J Cancer 2001; 96:22‑31.

Cell Cycle

1993

Iron and the Cell Cycle 166. Maytin EV, Ubeda M, Lin JC, Habener JF. Stress‑inducible transcription factor CHOP/ gadd153 induces apoptosis in mammalian cells via p38 kinase‑dependent and ‑independent mechanisms. Exp Cell Res 2001; 267:193‑204. 167. Zhan Q, Lord KA, Alamo Jr I, Hollander MC, Carrier F, Ron D, Kohn KW, Hoffman B, Liebermann DA, Fornace Jr AJ. The gadd and MyD genes define a novel set of mammalian genes encoding acidic proteins that synergistically suppress cell growth. Mol Cell Biol 1994; 14:2361‑71. 168. Price BD, Calderwood SK. Gadd45 and Gadd153 messenger RNA levels are increased during hypoxia and after exposure of cells to agents which elevate the levels of the glucose‑regulated proteins. Cancer Res 1992; 52:3814‑7. 169. Lee SK, Jang HJ, Lee HJ, Lee J, Jeon BH, Jun CD, Kim EC. p38 and ERK MAP kinase mediates iron chelator‑induced apoptosis and ‑suppressed differentiation of immortalized and malignant human oral keratinocytes. Life Sci 2006; 79:1419‑27. 170. Pruitt K, Pruitt WM, Bilter GK, Westwick JK, Der CJ. Raf‑independent deregulation of p38 and JNK mitogen‑activated protein kinases are critical for Ras transformation. J Biol Chem 2002; 277:31808‑17. 171. Bulavin DV, Saito S, Hollander MC, Sakaguchi K, Anderson CW, Appella E, Fornace Jr AJ. Phosphorylation of human p53 by p38 kinase coordinates N‑terminal phosphorylation and apoptosis in response to UV radiation. Embo J 1999; 18:6845‑54. 172. Lavoie JN, L’Allemain G, Brunet A, Muller R, Pouyssegur J. Cyclin D1 expression is regulated positively by the p42/p44MAPK and negatively by the p38/HOGMAPK pathway. J Biol Chem 1996; 271:20608‑16. 173. Moon SK, Jung SY, Choi YH, Lee YC, Patterson C, Kim CH. PDTC, metal chelating compound, induces G1 phase cell cycle arrest in vascular smooth muscle cells through inducing p21Cip1 expression: Involvement of p38 mitogen activated protein kinase. J Cell Physiol 2004; 198:310‑23. 174. Semenza GL. Regulation of mammalian O2 homeostasis by hypoxia‑inducible factor 1. Annu Rev Cell Dev Biol 1999; 15:551‑78. 175. Greijer AE, van der Groep P, Kemming D, Shvarts A, Semenza GL, Meijer GA, van de Wiel MA, Belien JA, van Diest PJ, van der Wall E. Up‑regulation of gene expression by hypoxia is mediated predominantly by hypoxia‑inducible factor 1 (HIF‑1). J Pathol 2005; 206:291‑304. 176. Wang GL, Jiang BH, Rue EA, Semenza GL. Hypoxia‑inducible factor 1 is a basic‑helix‑loop‑helix‑PAS heterodimer regulated by cellular O2 tension. Proc Natl Acad Sci USA 1995; 92:5510‑4. 177. Stockmann C, Fandrey J. Hypoxia‑induced erythropoietin production: A paradigm for oxygen‑regulated gene expression. Clin Exp Pharmacol Physiol 2006; 33:968‑79. 178. Ivan M, Kondo K, Yang H, Kim W, Valiando J, Ohh M, Salic A, Asara JM, Lane WS, Kaelin Jr WG. HIFalpha targeted for VHL‑mediated destruction by proline hydroxylation: Implications for O2 sensing. Science 2001; 292:464‑8. 179. Caro J. Hypoxia regulation of gene transcription. High Alt Med Biol 2001; 2:145‑54. 180. Bianchi L, Tacchini L, Cairo G. HIF‑1‑mediated activation of transferrin receptor gene transcription by iron chelation. Nucleic Acids Res 1999; 27:4223‑7. 181. Tacchini L, Bianchi L, Bernelli‑Zazzera A, Cairo G. Transferrin receptor induction by hypoxia: HIF‑1‑mediated transcriptional activation and cell‑specific post‑transcriptional regulation. J Biol Chem 1999; 274:24142‑6. 182. Kovacevic Z, Richardson DR. The metastasis suppressor, Ndrg‑1: A new ally in the fight against cancer. Carcinogenesis 2006; 27:2355‑66. 183. Guan RJ, Ford HL, Fu Y, Li Y, Shaw LM, Pardee AB. Drg‑1 as a differentiation‑related, putative metastatic suppressor gene in human colon cancer. Cancer Res 2000; 60:749‑55. 184. Kurdistani SK, Arizti P, Reimer CL, Sugrue MM, Aaronson SA, Lee SW. Inhibition of tumor cell growth by RTP/rit42 and its responsiveness to p53 and DNA damage. Cancer Res 1998; 58:4439‑44. 185. An WG, Kanekal M, Simon MC, Maltepe E, Blagosklonny MV, Neckers LM. Stabilization of wild‑type p53 by hypoxia‑inducible factor 1alpha. Nature 1998; 392:405‑8. 186. Bruick RK. Expression of the gene encoding the proapoptotic Nip3 protein is induced by hypoxia. Proc Natl Acad Sci USA 2000; 97:9082‑7. 187. Guo K, Searfoss G, Krolikowski D, Pagnoni M, Franks C, Clark K, Yu KT, Jaye M, Ivashchenko Y. Hypoxia induces the expression of the pro‑apoptotic gene BNIP3. Cell Death Differ 2001; 8:367‑76. 188. Furukawa T, Adachi Y, Fujisawa J, Kambe T, Yamaguchi‑Iwai Y, Sasaki R, Kuwahara J, Ikehara S, Tokunaga R, Taketani S. Involvement of PLAGL2 in activation of iron deficient‑ and hypoxia‑induced gene expression in mouse cell lines. Oncogene 2001; 20:4718‑27. 189. Mizutani A, Furukawa T, Adachi Y, Ikehara S, Taketani S. A zinc‑finger protein, PLAGL2, induces the expression of a proapoptotic protein Nip3, leading to cellular apoptosis. J Biol Chem 2002; 277:15851‑8. 190. Le NT, Richardson DR. Competing pathways of iron chelation: Angiogenesis or anti‑tumor activity: Targeting different molecules to induce specific effects. Int J Cancer 2004; 110:468‑9. 191. Stein S, Thomas EK, Herzog B, Westfall MD, Rocheleau JV, Jackson IInd RS, Wang M, Liang P. NDRG1 is necessary for p53‑dependent apoptosis. J Biol Chem 2004; 279:48930‑40. 192. Bandyopadhyay S, Pai SK, Gross SC, Hirota S, Hosobe S, Miura K, Saito K, Commes T, Hayashi S, Watabe M, Watabe K. The Drg‑1 gene suppresses tumor metastasis in prostate cancer. Cancer Res 2003; 63:1731‑6. 193. Bandyopadhyay S, Pai SK, Hirota S, Hosobe S, Takano Y, Saito K, Piquemal D, Commes T, Watabe M, Gross SC, Wang Y, Ran S, Watabe K. Role of the putative tumor metastasis suppressor gene Drg‑1 in breast cancer progression. Oncogene 2004; 23:5675‑81.

1994

194. Maruyama Y, Ono M, Kawahara A, Yokoyama T, Basaki Y, Kage M, Aoyagi S, Kinoshita H, Kuwano M. Tumor growth suppression in pancreatic cancer by a putative metastasis suppressor gene Cap43/NDRG1/Drg‑1 through modulation of angiogenesis. Cancer Res 2006; 66:6233‑42. 195. Bandyopadhyay S, Wang Y, Zhan R, Pai SK, Watabe M, Iiizumi M, Furuta E, Mohinta S, Liu W, Hirota S, Hosobe S, Tsukada T, Miura K, Takano Y, Saito K, Commes T, Piquemal D, Hai T, Watabe K. The tumor metastasis suppressor gene Drg‑1 down‑regulates the expression of activating transcription factor 3 in prostate cancer. Cancer Res 2006; 66:11983‑90. 196. Iyengar P, Combs TP, Shah SJ, Gouon‑Evans V, Pollard JW, Albanese C, Flanagan L, Tenniswood MP, Guha C, Lisanti MP, Pestell RG, Scherer PE. Adipocyte‑secreted factors synergistically promote mammary tumorigenesis through induction of anti‑apoptotic transcriptional programs and proto‑oncogene stabilization. Oncogene 2003; 22:6408‑23. 197. Janz M, Hummel M, Truss M, Wollert‑Wulf B, Mathas S, Johrens K, Hagemeier C, Bommert K, Stein H, Dorken B, Bargou RC. Classical Hodgkin lymphoma is characterized by high constitutive expression of activating transcription factor 3 (ATF3), which promotes viability of Hodgkin/Reed‑Sternberg cells. Blood 2006; 107:2536‑9. 198. Ameri K, Hammond EM, Culmsee C, Raida M, Katschinski DM, Wenger RH, Wagner E, Davis RJ, Hai T, Denko N, Harris AL. Induction of activating transcription factor 3 by anoxia is independent of p53 and the hypoxic HIF signalling pathway. Oncogene 2007; 26:284‑9. 199. Kaiser BK, Zimmerman ZA, Charbonneau H, Jackson PK. Disruption of centrosome structure, chromosome segregation, and cytokinesis by misexpression of human Cdc14A phosphatase. Mol Biol Cell 2002; 13:2289‑300. 200. Lawen A. Apoptosis‑an introduction. Bioessays 2003; 25:888‑96. 201. Fornace Jr AJ, Jackman J, Hollander MC, Hoffman‑Liebermann B, Liebermann DA. Genotoxic‑stress‑response genes and growth‑arrest genes: Gadd, MyD, and other genes induced by treatments eliciting growth arrest. Ann NY Acad Sci 1992; 663:139‑53. 202. Brard L, Granai CO, Swamy N. Iron chelators deferoxamine and diethylenetriamine pentaacetic acid induce apoptosis in ovarian carcinoma. Gynecol Oncol 2006; 100:116‑27. 203. Fan L, Iyer J, Zhu S, Frick KK, Wada RK, Eskenazi AE, Berg PE, Ikegaki N, Kennett RH, Frantz CN. Inhibition of N‑myc expression and induction of apoptosis by iron chelation in human neuroblastoma cells. Cancer Res 2001; 61:1073‑9. 204. Simonart T, Boelaert JR, Mosselmans R, Andrei G, Noel JC, De Clercq E, Snoeck R. Antiproliferative and apoptotic effects of iron chelators on human cervical carcinoma cells. Gynecol Oncol 2002; 85:95‑102. 205. Alvero AB, Chen W, Sartorelli AC, Schwartz P, Rutherford T, Mor G. Triapine (3‑aminopyridine‑2‑carboxaldehyde thiosemicarbazone) induces apoptosis in ovarian cancer cells. J Soc Gynecol Investig 2006; 13:145‑52. 206. Greene BT, Thorburn J, Willingham MC, Thorburn A, Planalp RP, Brechbiel MW, Jennings‑Gee J, Wilkinson JT, Torti FM, Torti SV. Activation of caspase pathways during iron chelator‑mediated apoptosis. J Biol Chem 2002; 277:25568‑75. 207. Zhao R, Planalp RP, Ma R, Greene BT, Jones BT, Brechbiel MW, Torti FM, Torti SV. Role of zinc and iron chelation in apoptosis mediated by tachpyridine, an anti‑cancer iron chelator. Biochem Pharmacol 2004; 67:1677‑88. 208. Rakba N, Aouad F, Henry C, Caris C, Morel I, Baret P, Pierre JL, Brissot P, Ward RJ, Lescoat G, Crichton RR. Iron mobilisation and cellular protection by a new synthetic chelator O‑Trensox. Biochem Pharmacol 1998; 55:1797‑806. 209. Amundson SA, Myers TG, Fornace Jr AJ. Roles for p53 in growth arrest and apoptosis: Putting on the brakes after genotoxic stress. Oncogene 1998; 17:3287‑99. 210. LaCasse EC, Baird S, Korneluk RG, MacKenzie AE. The inhibitors of apoptosis (IAPs) and their emerging role in cancer. Oncogene 1998; 17:3247‑59. 211. Wang D, Liu YF, Wang YC. Deferoxamine induces apoptosis of HL‑60 cells by activating caspase‑3. Zhongguo Shi Yan Xue Ye Xue Za Zhi 2006; 14:485‑7. 212. Kubli DA, Ycaza JE, Gustafsson AB. Bnip3 mediates mitochondrial dysfunction and cell death through Bax and Bak. Biochem J 2007.

Cell Cycle

2007; Vol. 6 Issue 16