Ribonucleotide Reductase Inhibitors and Future Drug ... - CiteSeerX

Current Cancer Drug Targets, 2006, 6, 409-431

409

Ribonucleotide Reductase Inhibitors and Future Drug Design J. Shao1,2, B. Zhou1, Bernard Chu and Y. Yen1,* 1Department

of Medical Oncology and Therapeutic Research, City of Hope National Medical Center, Duarte, CA

91010, USA 2Department

of Basic Medical Sciences, Zhejiang University School of Medicine, Hangzhou, Zhejiang 310031,

P.R. China Abstract: Ribonucleotide reductase (RR) is a multisubunit enzyme responsible for the reduction of ribonucleotides to their corresponding deoxyribonucleotides, which are building blocks for DNA replication and repair. The key role of RR in DNA synthesis and cell growth control has made it an important target for anticancer therapy. Increased RR activity has been associated with malignant transformation and tumor cell growth. Efforts for new RR inhibitors have been made in basic and translational research. In recent years, several RR inhibitors, including Triapine, Gemcitabine, and GTI-2040, have entered clinical trial or application. Furthermore, the discovery of p53R2, a p53-inducible form of the small subunit of RR, raises the interest to develop subunit-specific RR inhibitors for cancer treatment. This review compiles recent studies on (1) the structure, function, and regulation of two forms of RR; (2) the role in tumorigenesis of RR and the effect of RR inhibition in cancer treatment; (3) the classification, mechanisms of action, antitumor activity, and clinical trial and application of new RR inhibitors that have been used in clinical cancer chemotherapy or are being evaluated in clinical trials; (4) novel approaches for future RR inhibitor discovery.

Keywords: Ribonucleotide reductase, structure and function, inhibitors; classification, mechanism of action, clinical trial and application, drug discovery. INTRODUCTION Ribonucleotide reductase (RR) catalyzes the reduction of ribonucleotides to their corresponding deoxyribonucleotides, which are the building blocks for DNA replication and repair in all living cells [1-4]. Three main classes of RR have been described based on different metal cofactors for the catalytic activity [1, 5]. Class I enzymes are found in practically all eukaryotic organisms, from yeast and algae to plants and mammals, and some prokaryotes and viruses also express this type. Class I is further divided into three subclasses (Ia, Ib, and Ic) based on polypeptide sequence homologies and their overall allosteric regulation behavior [6, 7]. Human RR belongs to Class Ia. Since the reduction of ribonucleotides is the rate-limiting step of DNA synthesis, inactivation of RR stops DNA synthesis, which inhibits cell proliferation. The important role of RR in DNA synthesis and repair has made it an important target for anticancer and antivirus agents [811]. This review focuses on the recent progress on the structure and function of RR, and the mechanisms of action of RR inhibitors that are in clinical use, undergoing clinical trials, or have recently shown promising results in preclinical studies, and future RR inhibitor discovery. Structure and Function of Ribonucleotide Reductase Structure Most of the structure-function studies on Class I RRs have been performed on E.coli and mouse enzymes. A tetrameric holoenzyme model for Class I RR (α 2 β 2 ) has *Address correspondence to this author at the City of Hope National Medical Center, 1500 East Duarte Road, Duarte, CA 91010, USA; Tel: (626) 359-8111, Ext: 62307; Fax: (626) 301-8233; E-mail: [email protected] 1568-0096/06 $50.00+.00

been proposed (Fig. 1. E.coli RR holoenzyme model) [12, 13]. The large α2-homodimer is called R1 and the small β2homodimer is called R2. The crystal structures have been determined separately for R2 [14, 15] and R1 [13] from E. c o l i , and R2 from mouse [16-18]. However, a crystallographic structure of the R1-R2 holoenzyme complex is not yet available [4, 19]. R1 is the reductase component of the Class I enzyme, which harbors the active site (substrate binding), allosteric sites (overall activity site and substrate specificity site), and redox active disulfides that participate in the reduction of substrates (Fig. 2. E.coli R1) [13, 20, 21]. R2 contains an oxygen-linked diferric iron center and a tyrosyl radical that are essential for enzymatic activity (Fig. 3. E.coli R2) [3, 15]. Formation and stability of the radical depends on the binuclear Fe(III) center. The carboxyl end of R2 is important for the formation of the holoenzyme complex [13]. Catalytic Mechanism A scheme for the catalytic reaction has been proposed using E.coli RR as a model [5, 12, 13, 19, 22-26]. The substrate turnover reaction in R1 is proposed to be initiated by a thiyl radical on a conserved cysteine residue (C439) at the substrate binding site by abstracting a hydrogen atom from the ribose ring of the substrate. In the course of the substrate turnover, a disulfide bridge that has to be reduced before the enzyme can be active again, is formed between C225 and C462 at the active site in R1. This is accomplished by the hydrogen donor proteins, thioredoxin or glutaredoxin, via a second disulfide bridge between C754 and C759 located on the flexible C terminal tail of R1. In R2, the diiron center is capable of oxidizing the tyrosine residue Y122 to a tyrosyl radical following the reactivation with dioxygen. The stable tyrosyl radical in R2 is proposed to be transferred to C439 in R1 to generate the putative thiyl © 2006 Bentham Science Publishers Ltd.

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radical by way of a proton-coupled electron-transfer pathway consisting of a chain of hydrogen bonded amino acid residues (Fig. 4) [12, 27]. Unlike E.coli R2, the iron center

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and the radical in mouse R2 is labile and has to be continuously regenerated in vivo in a reaction, which requires ferrous iron and oxygen [28-31].

Fig. (1). Docking model of the R1 and R2 subunits of E. coli RR. A shows the surface complementarity and B shows the electron/proton transfer pathway (rotated 90 degree). The monomers of R1 are indicated in blue and green. Each monomer has substrate (GDP) and effector (TTP). Also indicated in R1 are the three active-site cysteines (C439, C225, and C462) and two tyrosines (Y730 and Y731) thought to be involved in the electron/proton transfer pathway between R1 and R2. All of these residues are in cyan. The monomers of R2 are indicated in red and gold. The C terminal 15 residues of this peptide, 360-375 of R2, are shown in red. The two irons on each monomer are shown in blue balls. The residues thought to be involved in the electron/proton transfer pathway between R1 and R2 are shown in cyan (Y122, D237, and W48) (From references 12 and 13).

Fig. (2). E. coli R1 protein dimer with the C terminal peptide (20 residues) of R2 (PDB Code: 4R1R). The active site and allosteric sites of R1 and the C terminal peptide of R2 are highlighted (From reference 20).

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Fig. (3). E.coli R2 protein dimer structure (PDB Code: 1RIB). The Tyr122 and di-iron ions are highlighted (From reference 15).

Multiple Regulation of Ribonucleotide Reductase Activity Synthesis of deoxyribonucleotides is an extremely well regulated reaction. A failure in the control of the dNTP levels and/or their relative amounts leads to cell death or genetic abnormalities [32, 33]. There are multiple levels of regulation of RR activity in the mammalian cell [34-36]. Allosteric Regulation RR is regulated by allosteric control of the activity and specificity of the enzyme. The overall enzymatic activity is regulated by ATP (activation) and dATP (feedback inhibition at high concentrations) binding to a general activity site (A-site) in R1 [1]. In order to obtain a balanced supply of dNTPs for DNA synthesis, the specificity for each of the four substrates is determined by the binding of an allosteric effector (dATP or ATP, dTTP, and dGTP) to the specificity site (S-site) [34, 37]. A third site called the hexamerization site (H-site) is also proposed to regulate the activity of Class Ia RR. ATP binding to this H-site promotes the formation of an R16 R2 6 hexamer, which is suggested to be the major active form of RR in mammalian cells [38, 39]. Transcriptional Regulation and Protein Degradation Regulation Large amounts of deoxyribonucleotides are required in proliferating cells during the S-phase of the cell cycle where

genome duplication takes place. The level of RR in cells is therefore closely linked with the cell cycle and growth control mechanisms. In human and mouse cells, the genes encoding the R1 and R2 proteins are located on different chromosomes [40, 41]. The transcription of the R1 and R2 genes is cell cycle-regulated with undetectable mRNA levels in G0/G 1 phase cells. Enzyme activity and the R1 and R2 mRNAs reach maximal levels during S-phase [35, 42-44]. R1 protein has a long half-life of 18-24 h, and its levels are almost constant and in excess throughout the cell cycle in proliferating cells. R2 protein shows an S-phase specific expression and has a shorter half-life of about 3-4 h. R2 protein synthesis starts in early S phase, and it slowly accumulates in the cell up to late mitosis when it is rapidly degraded [45]. Enzyme activity is therefore determined by R2 protein levels. R2 degradation in late mitosis depends on a KEN box sequence recognized by the Cdh1·anaphasepromoting complex active in late mitosis and during G0/G1 [36, 46-48]. Other Regulation Mechanisms In addition, RR activity is controlled by binding of a protein inhibitor, Sml1, to the R1 subunit in budding yeast and by an R2 transcript specific cytoplasmic poly(A) polymerase, Cid13, in fission yeast [49, 50]. RR activity regulation may also be obtained by controlling the amount of R2 tyrosyl free radical. In E.coli, small reduced molecules


(such as flavin), which are usually present inside the cells, can scavenge the tyrosyl free radical and this could be a way to regulate the concentration of the active enzyme [51]. The control of the immediately available pool of iron in cells may also play an important role in regulating RR activity [52, 53].

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cooperation with a variety of oncogenes to determine transformation but also enhances cancer invasive potential [57-63]. Tumor cells are more sensitive to the cytotoxic effect of RR inhibition than normal cells because of the increased need of dNTPs for proliferation and decreased adaptability and low responsiveness to regulatory signals. Thus, the enzyme has long been considered an excellent target for cancer chemotherapy [9, 64]. p53R2 and its Role in Tumorigenesis and Cancer Treatment p53R2, a R2 homolog (~80%), has been identified in human and mouse cells [65, 66]. The p53R2 gene contains a p53-binding site in intron 1. The expression of p53R2, but not R2, is induced by ultraviolet light, γ-irradiation or DNA-damaging agents in a p53-dependent manner. Cells that fail to make p53R2 are more sensitive to killing by DNA-damaging agents. p53R2 forms an active RR in vitro with R1 and the expression of the R1 is induced in resting cells after UV irradiation [67]. A study using p53R2 knockout mice showed that impairment of the p53R2involved DNA repair pathway enhances the frequency of spontaneous mutations and activates p53-dependent apoptotic pathway(s) [68]. These findings lead to the hypothesis that there are two pathways in human cells to supply dNTPs for DNA synthesis: one through the activity of R2, involved in normal maintenance of dNTPs for DNA replication during the S/G2 phases in a cell cycle-dependent manner, and the other through p53R2, supplying dNTPs for DNA repair in G0/G1 cells in a p53-dependent manner [69, 70]. However, it is still unclear how the expression and activity of the two small subunits are coordinately regulated in cells. It has been reported that p53 can interact with p53R2 and R2 at the protein level to regulate RR activity: p53 binds both R2 and p53R2 in resting cells but upon exposure to UV irradiation, R2 and p53R2 dissociated from p53 and bound to R1. p53R2, R2 and R1 transfer to the nucleus, forming an active RR complex to provide dNDPs for DNA repair [71]. In p53 mutant and null cells, R2 complements p53R2 in response to UV-induced DNA repair [72]. On the other hand, cells with mutated p53 might still induce p53R2 in response to treatment with DNA-damaging agents [73]. This implies that there might exist an additional p53-independent induction pathway for p53R2.

Fig. (4). Using the model structure described in Fig. 1, the conserved residues proposed to be involved in the electron/proton transfer pathway between R1 and R2 are shown (From references 12 and 27).

Ribonucleotide Reductase as a Target for Cancer Therapy RR is responsible for maintaining a balanced supply of dNTPs required for DNA synthesis and repair. Therefore, the enzyme plays an important role in cell proliferation [54, 55]. The effect of RR inactivation in cells includes decreases of intracellular concentrations of the dNTPs, inhibition of DNA synthesis, inhibition of DNA repair in quiescent cells, and cell cycle arrest and apoptosis [9, 56]. Increased RR activity has been associated with malignant transformation and cancer metastasis; R2 is not only capable of acting in

Several known functional domains in mouse R2 are also conserved in p53R2 [16, 19, 97, 74, 75], including the iron ligands, the radical site tyrosine, the hydrophobic pocket surrounding the tyrosyl radical site, the radical transfer pathway from small subunit to large subunit, the C terminal sequence for binding to the large subunit, and the hydrophobic channel from the surface to the interior of the protein. The major sequence difference between the two small subunits is that p53R2 lacks 33 amino acid residues in its N-terminus [47, 67]. RR activity depends on the integrity of the dinuclear iron center and the tyrosyl radical. Electron paramagnetic resonance (EPR) studies reveal that the tyrosyl radical signal in p53R2 protein is almost identical with that in R2 protein, indicating that both small subunits may employ the same tyrosyl radical mechanism for enzymatic activity. However, the reaction rate of p53R2 was lower than that of R2, which may be due to its reduced binding affinity to R1 [76].


The discovery of p53R2 raises interest in the role of p53R2 in the development of human cancers [65-70]. Furthermore, because R2 and p53R2 play different roles in cells, inhibitors specific for each subunit may exhibit different clinical values. HUMAN RIBONUCLEOTIDE INHIBITORS

REDUCTASE

Classification Inhibitors of RR have been widely investigated as potential anti-tumoral, anti-viral, and anti-bacterial chemotherapeutic agents [9-11]. Some RR inhibitors have been proved for clinical treatment of cancer; others are being evaluated in clinical trials [9, 19, 77, 78]. The inhibitors of RR can be divided into different categories based on different criteria. (1) According to chemical structures and physic-chemical properties, the inhibitors of RR can be divided into three groups: small molecule compounds, synthetic oligopeptides, and oligonucleotides [79-81]. (2) According to targets and mechanisms of action, RR inhibitors can be grouped in two main categories: gene expression regulation and protein inactivation. Each category can be further divided in several sub-categories. Table 1.


Protein Inactivation These inhibitors inactivate RR by interacting with R1 or R2, or interfering with R1 and R2 assembly. The R1 inhibitors inhibit the active site or induce allosteric malfunction by nucleoside analogs. The R2 inhibitors destroy the essential diiron tyrosyl radical center using radical scavengers or iron chelators. Polymerization inhibitors prevent the formation of RR holoenzyme by oligopeptides corresponding to the carboxyl terminus of R2. R1 Protein Inhibitors Depending on mechanisms of inactivation, R1 protein inhibitors can be divided in two sub-groups: (1) nucleoside analogs (including substrate and allosteric effector analogs); (2) inactivators of sulfhydryl groups (Table 1). Nucleoside Analogs The derivatives of the natural purine and pyrimidine nucleosides are effective in the clinical treatment of human cancer or viral diseases. Arabinosylcytosine (ara-C), gemcitabine, fludarabine, cladribine, and clofarabine are examples of this class of drugs that have therapeutic activity against human malignancies. After intracellular conversion to dNDP or dNTP analogs, nucleoside analogs become active species. The key mechanisms responsible for the cytotoxic

RNR Inhibitors

Mechanism of action

Name

Structure

1. Protein inactivation (1) R1 protein inhibition NH2 N Nucleoside analog-Substrate analog

Gemcitabine HO

N

O

O

F F

HO

NH2 N Tezacitabine

HO

N

O

O

H

HO F NH2

N DMDC

413

HO

O

HO

N

CH2

O


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(Table 1)contd.....

Mechanism of action

Name

Structure

1. Protein inactivation (1) R1 protein inhibition NH2 N Cytarabine

HO

N

O

O

HO

HO

H NH2 N

Nucleoside analog-Allosteric effector analog

Fludarabine

H2 O3 PO

N

N

O

N

F

HO

HO

H NH2 N

Cladribine

HO

N

N

O

N

Cl

H HO

H CH3 CH C C CH HC

Clofarabine

H3C

CH2

CH2

HC

CHCH 3

CH

C CH3

H3C NH2 N

N Nucleoside analog-Bivalent nucleotide inhibitor

S N

ADP-S-HBES-S-dGTP PPO

CH C CH C CH 3

N

O

O

S

S

O

6 O

O N

NH

S N

N

O

OPPP O

HO

OH O H N

O Inactivators of sulfhydryl groups

NH2

N

Caracemide

O HN

O

OH



415

(Table 1)contd.....

Mechanism of action

Name

Structure

1. Protein inactivation (1) R1 protein inhibition Cl

Cl Cisplatin

Pt NH3

NH3 (2) R2, p53R2 protein inhibition O Radical scavengers

Hydroxyurea

OH H2N

N H OH

HO

OH

Trimidox

HON

NH2

OH OH Didox

OH O

N H NO

Nitric Oxide OH

R= methyl, ethyl, allyl, or n-propyl

Alkoxyphenols

OR

N Iron chelators

NH2

Triapine N H2 N

NH S OH

H N

O

N

O O

HO

HN

N

DFO NH2 N OH

O O


Shao et al. (Table 1)contd.....

Mechanism of action

Name

Structure

1. Protein inactivation (2) R2, p53R2 protein inhibition N OH

HO

PIH N

N NH O

HO

311 N

N NH O

action of nucleoside analogs involve one of two major biochemical pathways: 1) inhibition of RR (as substrate or allosteric effector analogs) and 2) incorporation into DNA. DNA polymerase inhibition follows incorporation of the dNTP analog into the synthesized chain. Therefore, the concomitant RR inhibition activity of the agent is a powerful self-potentiation mechanism since the dNTP pools decrease, thus favoring the analog incorporation. The combined effect of the analogs leads to termination of DNA chain elongation, inhibition of DNA synthesis, interference with DNA repair mechanisms, and apoptosis as the most likely final pathway of activity [82, 83]. Substrate Analogs These inhibitors are activated by deoxycytidine kinase (dCK) to become a substrate analog of RR. Cytarabine, gemcitabine, tezacitabine, and DMDC are deoxycytidine analogs. These substrate analogs, modified at the carbon 2’ in the sugar moiety (ribose or arabinose), are known as suicide inhibitors because they are recognized by RR as normal substrates and react in the active site of R1 leading to abnormal products that inactivate the enzyme [84-90]. These compounds were found to be important for the design of mechanism based new inhibitors of RR [88, 91]. Cytarabine (1-β-D-arabinosylcytosine, cytosine arabinoside, ara-C) is one of the first inhibitors that showed anti-tumor properties and is the most active drug in acute myeloid leukemias (AML) [92-94]. Ara-C is a good substrate of dCK and is further metabolized to ara-CDP and ara-CTP in vivo. AraCTP inhibits DNA synthesis after incorporation into DNA by DNA polymerase α. Ara-CDP inhibits RR activity. However, at high concentrations ara-CTP acts as a feedback inhibitor of dCK and has a short half-life in tumor cells. Furthermore, the rapid degradation by deamination causes

ara-C to be ineffective against solid tumors. Novel 2’substituted substrate analogs have been synthesized and their anti-tumor activity has been intensively investigated in the hope to find more effective inhibitors [95-97]. Gemcitabine (2',2'-difluoro-2'-deoxycytidine, dFdC) is metabolized intracellularly by deoxycytidine kinase to its monophosphorylated form, and further by other intracellular kinases (nucleoside monophosphate kinases and nucleoside diphosphate kinases) to the metabolically active forms, gemcitabine diphosphate (dFdCDP) and gemcitabine triphosphate (dFdCTP) [98-101]. The cytotoxic effect is attributed to a combination of two actions performed by those two metabolites. First, dFdCDP inhibits RR. The inhibitor is falsely recognized by the enzyme as a natural substrate, and reacts with the active site, leading to abnormal products and loss of RR catalytic activity [102-105]. The mechanism of inactivation depends on the availability of reductant. In the presence of a reductant [thioredoxin (TR)/thioredoxin reductase (TRR)/NADPH or dithiothreitol], inhibition results from R1 inactivation probably by alkylation. In the absence of reductant with prereduced R1 and R2, inhibition results from loss of the essential tyrosyl radical in R2. Inhibition of RR causes a reduction in the cellular concentration of the four DNA monomers. Second, dFdCTP competes with the natural dCTP for incorporation into the replicating DNA. Once one molecule of dFdCTP is incorporated, an additional deoxyribonucleotide is added to the growing DNA strands, and after that, DNA synthesis can no longer proceed [106]. The decreased intracellular concentration of dCTP, caused by the inhibition of RR, has important consequences: faster phosphorylation of dFdC to the two active forms, decreased metabolic clearance of the gemcitabine nucleotides by deoxycytidine monophosphate deaminase, and enhanced


incorporation of dFdCTP into DNA [107, 108]. This selfpotentiation mechanism should account for the high anticancer efficacy of dFdC as opposed to other nucleoside antimetabolites [109]. Cytotoxicity of gemcitabine in nondividing cells has been attributed to the fact that dFdCTP can disturb RNA metabolism by inhibiting CTP synthetase or by direct incorporation into newly synthesized RNA [110, 111]. Gemcitabine has been approved for treatment of nonsmall cell lung cancer and pancreatic cancer by the FDA [112-115]. In Europe it was approved for the treatment of bladder cancer [116, 117]. Moreover, it continues to be employed in multiple trials that assess the clinical benefit of different therapeutic approaches and combinations in these solid tumors [118-123]. Gemcitabine is also shown to be a potent radiation sensitizer in a variety of solid tumors and tumor cell lines [124-128]. The role of gemcitabine as a single agent or in combination with other anti-tumor drugs in the treatment of breast cancer [129], ovarian cancer [130, 131], squamous cell carcinoma of the head and neck [132], and malignant pleural mesothelioma [133] has also been investigated. Gemcitabine has shown activity in clinical trials in patients with hematologic malignancies [134, 135]. Tezacitabine [(E)-2'-deoxy-2'-(fluoromethylene) cytidine, FMdC], a deoxycytidine analog, is an irreversible RR inhibitor and DNA chain terminator [136-139]. After intracellular phosphorylation to a diphosphate form, the drug irreversibly inhibits the enzyme RR [140, 141]. FMCDP inhibits both R1 and R2 subunits of RR, by destroying the essential tyrosyl radical in R2 and alkylating the active site in R1. Tezacitabine is also phosphorylated intracellularly to a triphosphate form that has significant DNA chain termination activity by inhibiting DNA polymerase α, with consequences for cell cycle arrest and apoptosis [142]. This dual mechanism of action is similar to that of gemcitabine. However, at least two significant features differentiate tezacitabine from gemcitabine. First, unlike that by gemcitabine, the inhibition of RR by tezacitabine is irreversible [143]. Second, tezacitabine is approximately 30fold more resistant than gemcitabine to metabolic inactivation through deamination by cytidine deaminase [144]. The latter could provide an advantage for tezacitabine in the treatment of cancer types in which cytidine deaminase levels are elevated or in individual patients who have tumors with high levels of the enzyme. For example, studies with human tumor xenografts suggest that many gastrointestinal tumors have high cytidine deaminase levels, and thus tezacitabine may be particularly useful in the treatment of this group of malignancies [146]. Tezacitabine shows potent antitumor activity against leukemia (HL-60, MOLT-4, L1210 leukemia, and both MDR-resistant and sensitive P388 cell lines) and solid tumors (breast, lung, stomach, colon, ovary, bladder, pancreas, glioma, and melanoma cell lines) [146, 147]. Tezacitabine has also shown radiosensitizing effects on different cancer cell lines and mice tumor xenograft models [148-150]. Tezacitabine has been investigated in phase I and II trials in patients with advanced solid tumors, in patients with relapsed or refractory hematologic malignancies, and in


417

combination with cisplatin and 5-fluorouracil (5-FU) in patients with advanced cancers [151-152]. DMDC (2'-deoxy-2'-methylidenecytidine) is a novel deoxycytidine analog with structural similarity to ara-C and gemcitabine [153]. DMDC is activated by deoxycytidine kinase to the diphosphate and triphosphate metabolites. DMDC triphosphate inhibits human DNA polymerases with resultant inhibition of additional elongation of DNA; DMDC diphosphate inhibits RR [154, 155]. DMDC is highly resistant to cytidine deaminase, so that it i s minimally converted to an inactive metabolite, 2'-deoxy-2'methylideneuridine [156, 157]. In tumors with high cytidine deaminase activity, the enzyme converts 2'-deoxycytidine to 2'-deoxyuridine and lowers intracellular 2'-deoxycytidine concentrations. Because 2'-deoxycytidine competitively inhibits the activation of DMDC by deoxycytidine kinase, DMDC is phosphorylated more effectively because of the decreased level of intracellular 2'-deoxycytidine. DMDC was highly effective in human cancer xenograft models with high levels of cytidine deaminase activity, whereas gemcitabine was less effective in such tumors [156, 158]. These features distinguish DMDC from other deoxycytidine analogs such as ara-C and gemcitabine. Therefore, non-small cell lung cancer, colon cancer, esophageal cancer, pancreas cancer, and cervical cancer, which show high cytidine deaminase activity, seem to be rational targets for therapy with DMDC [159]. DMDC has a broad spectrum of antitumor activity against murine and human tumor cell lines as well as the xenografts of human tumors in nude mice, including murine L1210 and P388 leukemia, colon 26 carcinoma, and M5076 reticulum cell sarcoma, and in the xenografts of SK-Mel-28 human melanoma and LX-1 human lung cancer [160]. Phase I studies have been conducted in patients with advanced solid tumors [161-165]. Several tumor responses are observed at the highest doses of DMDC, indicating a possible dose-response relationship with this drug. DMDC was found both to inhibit the growth and to induce differentiation of acute promyelocytic leukemia (APL) cell lines. Combination of all-trans retinoic acid (ATRA) and DMDC had more than additive effects in inducing the differentiation of NB4 cells. Similar results were observed in a primary culture of leukemia cells that had been freshly isolated from APL patients. These results suggest that DMDC may play a role in the treatment of APL [166]. Allosteric Effector Analogs dATP is an allosteric inhibitory effector of RR activity. Some synthetic deoxyadenosine analogs have an even higher affinity towards R1 with respect to deoxyadenosine (thus giving a powerful RR allosteric inhibition) and have low affinity for nucleotide-catabolizing enzymes such as adenosine deaminase (thus resulting in resistance to cell inactivation). Cladribine and Fludarabine Cladribine (2-chloro-2'-deoxyadenosine, CdA) and Fludarabine (9-β-D-arabinofuranosyl-2-fluoroadenine 5'monophosphate, Fara-A) are deoxyadenosine analogs and require intracellular phosphorylation for their cytotoxic


actions. In proliferating cells, cladribine triphosphate inhibits RR as well as DNA polymerase [167, 168]. CdATP is also incorporated into DNA, producing strand breaks and inhibition of DNA synthesis [169]. In resting cells, CdATP interferes with the proper repair of DNA strand breaks, which activates poly (ADP-ribose) synthetase, and NAD + is depleted, leading to apoptosis [170, 171]. Fludarabine triphosphate is a potent inhibitor of several important enzymes including DNA polymerase [172-174], DNA ligase, DNA primase [175-176], and RR [177-179]. Although the DNA termination may be a major mechanism for the cytotoxicity of fludarabine triphosphate, the depletion of dNTP pools in tumor cells after fludarabine exposure confirms that RR inhibition has a significant role in fludarabine cytotoxic mechanisms. Fara-ATP is also incorporated into RNA resulting in premature termination of the RNA transcript and impairing its functioning as a template for protein synthesis [180]. Because of structural similarity, purine nucleoside analogs may also function as activators of dATP-dependent caspase activation and apoptosis. Indeed, CdATP as well as Fara-ATP have been shown to cooperate with cytochrome c and induce caspase-3 activation and the caspase proteolytic cascade, leading the cell to apoptosis [181, 182]. These findings suggest a mechanism by which CdATP and Fara-ATP might induce cell death in quiescent cells in the absence of their incorporation into DNA playing a role in the chemotherapy of indolent lymphoproliferative diseases. CdA and Fara-A have many common characteristics with respect to their structures and metabolism, but their differences in the mechanism of action give them unique clinical activity. Cladribine is the current treatment of choice in hairy cell leukemia (HCL) [183-186] It has also shown efficacy in other lymphoproliferative disorders, such as chronic lymphocytic leukemia, low-grade non-Hodgkin’s lymphomas and Waldenström’s macroglubinemia, as well as in childhood acute myelogenous leukemia [187]. CdA also has impressive clinical activity against multiple sclerosis [188]. A phase II trial of CdA confirmed efficacy in patients with chronic myeloid leukemia (CML) in accelerated or blast phase [189]. The combination of CdA and ara-C seems to be effective therapy for pediatric AML [190]. CdA and arabinosylcytosine (araC) combinations exert synergistic cytotoxicity in human H9-lymphoid cell lines sensitive and resistant to araC [191]. Fludarabine (is in the treatment of) chronic lymphocytic leukemia (CLL), acute myelogenous leukemia (AML), low-grade lymphoma and other hematologic malignancies [192-195]. Fara-A has also been shown to have synergistic effects in combination with other chemotherapeutic drugs, e.g. ara-C, cyclophosphamide, and also in combination with radiation [196]. Clofarabine (2-chloro-2'-fluoro-2'-deoxy-9-beta - D - arabinofuranosyladenine, Cl-F-ara-A), a deoxyadenosine analog, has been approved by the US FDA in December 2004 for the treatment of pediatric patients with relapsed or refractory acute lymphoblastic leukemia (ALL) [197-199]. Clofarabine combines the most favorable pharmacokinetic properties of fludarabine and cladribine. It retains the halogenation at the carbon 2’ position of adenine of fludarabine and cladribine, which renders those analogs resistant to deamination by

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adenosine deaminase. A fluorine molecule at the 2'-position of the carbohydrate inhibits cleavage of the glycosidic linkage by bacterial purine nucleoside phosphorylase and stabilizes the compound in acidic environments, which is attributed to increase to its oral bioavailability [200-203]. Like fludarabine and cladribine, clofarabine requires intracellular phosphorylation by deoxycytidine kinase to its active triphosphate form. Clofarabine triphosphate inhibits both DNA polymerases and RR, and has a high affinity to the enzyme deoxycytidine kinase, the rate-limiting step in nucleoside phosphorylation [204-206]. Furthermore, clofarabine triphosphate induces apoptosis through induction of DNA strand breaks and disruption of mitochondrial integrity, resulting in the release of proapoptotic proteins [179, 207, 208]. While these mechanisms are shared with fludarabine or cladribine, significant differences exist: 1) fludarabine triphosphate primarily inhibits DNA polymerases after incorporation into DNA; 2) cladribine triphosphate acts mainly by inhibition of RR; 3) although antiapoptotic effects of all three purine nucleoside analogs occur by induction of DNA strand breaks, only clofarabine and cladribine have a direct impact on mitochondrial integrity and activity. Clofarabine has shown cytotoxicity to a variety of human hematologic and solid tumor cell lines and tumor xenograft models (leukemia, colon, renal, and breast tumor models) [209-212]. In addition to having been approved for treatment of pediatric ALL, clofarabine has also demonstrated antitumor activity against childhood and adult AML (acute myeloid leukemia) in Phase I and II trials [213-215]. The combination of clofarabine with cytarabine was active in myeloid malignancies and was well tolerated in phase I-II trials [216]. Bivalent Nucleotide Inhibitor of R1 A novel nucleotide inhibitor (ADP-S-HBES-S-dGTP) of mouse RR was designed to span the active site and the allosteric specific site of the enzyme. The inhibitor contains ADP and dGTP moieties which are linked by 1,6-hexane(bis-ethylenesulfone) (HBES) [217]. Although biological activity evaluation suggests that this compound binds only to the allosteric site and not to the active site, the results provide grounds for optimism that high affinity bivalent inhibitors of RR are attainable via tether optimization. Inactivators of R1 Sulfhydryl Groups Sulfhydryl groups of the active site cysteines in R1 are directly involved in the reduction of the substrate. Compounds that interfere with these sulfhydryl groups can inhibit RR activity. Caracemide (N-acetyl- N, O - di (methylcarbamoyl) - hydroxylamine, CAR) has been shown to inhibit RR from Novikoff ascites tumor cells [218]. Caracemide and its degradation products, N-acetyl-O-methylcarbamoyl-hydroxylamine, are found to irreversibly inhibit RR of E.coli by specific interactions with R1. No effect on R2 was observed. The R1R2 holoenzyme was approximately 30 times more sensitive to caracemide inactivation than the isolated R1 protein. The modification of R1 occurs at an activated cysteine or serine residue in the active site of the enzyme [219]. The


compound was found to have a stronger inhibitory effect on DNA synthesis in P388 lymphocytic leukemia and Ehrlich ascites carcinoma cells than hydroxyurea [220]. However, caracemide is unstable in either phosphate buffer (pH 7.4) or human plasma and causes neurological and psychiatric effects [221, 222]. Phase I studies [223, 224] and phase II [225-228] clinical trials demonstrate that caracemide has shown little promise in the treatment of human malignancies. Cisplatin It is one of the most potent and widely used anticancer agents in the treatment of various solid tumors [229]. The cytotoxicity of the drug is thought to be determined primarily by forming DNA adducts and interfering with its repair mechanism. It has been reported that cisplatin inhibits E.coli RR activity [230]. The site of inhibition was found to be the R1 subunit and the thiol groups of the conserved active site cysteines seem to be the preferential targets. It has also been shown that cisplatin inhibits mammalian RR. However, the study did not support a hypothesis that inhibition of RR is an important component of cisplatin cytotoxic activity or that it is a major participant in cisplatin resistance mechanisms [231]. R2 and p53R2 Protein Inhibitors This category of RR inhibitors target the small subunits R2 and p53R2 by directly quenching the tyrosyl radical and/or affecting the iron center, such as the radical scavenger hydroxyurea; by destroying the tyrosyl radical via forming a redox-active complex with iron and generating ROS, which results in radical destruction and RR inactivation (i.e., the iron chelator Triapine); or by depleting intracellular iron and preventing the formation of the iron center in R2 (i.e., the iron chelator DFO). The radical scavengers may also react with residues that are transiently radicalized throughout the electron transfer pathway from the tyrosyl radical site of R2 to the active site of R1 [232] (Table 1). Radical Scavengers Hydroxyurea It has been marketed for cancer therapy for decades [233]. Hydroxyurea is a radical scavenger and it inactivates RR by directly reducing the tyrosyl radical of R2 to a normal tyrosine residue via one-electron transfer from the drug [234, 235]. The iron center in mouse R2 is also reduced by the inhibitor and released from the protein in contrast to E.coli R2 [30]. The different sensitivity of mouse and E.coli R2 to hydroxyurea could be due to the different accessibility of the iron-tyrosyl radical center of R2. The -NH-OH segment of hydroxyurea is the minimal structural requirement that triggers the inhibitory properties of this compound [236]. In vitro RR holoenzyme activity assay and EPR measurements on the tyrosyl radical of R2 and p53R2 demonstrate that hydroxyurea inhibits both small subunits [76, 236]. In the absence of iron in the sample solution, hydroxyurea inhibits both small subunits at a similar level. However, the inhibition is reversed significantly in the presence of iron, DTT, and oxygen. The regeneration was much more marked for p53R2, resulting in a selective inhibition of R2 more than p53R2.


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Hydroxyurea is commonly used for the treatment of chronic myelogenous leukemia, polycythemia vera and essential thrombocythemia [237-240]. It is used in nontumoral pathologies such as sickle cell anemia or refractory psoriasis [241, 242]. Hydroxyurea has shown synergism with the nucleoside analogs didanosine (2',3'dideoxyinosine, or ddl) and 3'-azido-3'-deoxythymidine (AZT) for the treatment of HIV-1 infection in vitro and in clinical trials presumably because its alteration of intracellular dNTP pools facilitates incorporation of chain terminating nucleoside analogs into the growing viral DNA chain [243-248]. However, long-term use of hydroxyurea in HIV-1 infection may be limited by its hematopoietic toxicity. Hydroxyurea effectiveness is limited by its relatively low affinity for RR, very high hydrophilicity, small molecular size, short half-life and the development of resistance [249-252]. Resistance to hydroxyurea may be caused by quantitative or qualitative modification of RR subunits [253, 254]. Overexpression of R2 increases resistance to hydroxyurea in cancer cells. R2 subunit gene amplification and alteration in transcriptional regulation may be responsible for the mechanism of the drug resistance. Hydroxyurea resistant cell lines are also shown to have an increased expression of ferritin genes [255]. The increase of iron could favor the reactivation of R2 after the destruction of the protein by the drug. Mutations in the R2 subunit may also confer resistance to hydroxyurea [256]. Efforts have been made to develop hydroxyurea analogs and other small inhibitors to improve the inhibitory efficiency of hydroxyurea [257-260]. Didox and Trimidox Didox (3,4-dihydroxybenzohydroxamic acid) and trimidox (3,4,5-trihydroxybenzamidoxime), two hydroxybenzohydroxamic acid derivatives, are excellent radical scavengers and can form iron complexes [261-267]. They have been shown to inhibit the in vitro activity of partially purified RR enzyme 17- and 100-times more effectively than hydroxyurea respectively and are more effective than hydroxyurea in inhibiting the growth of various tumor cell lines. Hydroxyurea and hydroxybenzo-hydroxamic acids have differential effects upon dNTP pools, and hydroxyurearesistant L1210 cells are not cross-resistant to hydroxybenzohydroxamic acids, indicating that hydroxybenzohydroxamic acids have some properties other than those of hydroxyurea, which relate to their RR inhibitory and cytotoxic effects. Didox has been evaluated in phase I and II clinical trials for toxicity and therapeutic efficacy [268-270]. Phase I studies demonstrated that the compound was well tolerated and was considered safe. However, a phase II study found less positive response in the treatment of advanced breast cancer patients. It was suggested that a higher dose is needed for treatment. Potential use of didox for treating other tumors or combining with other anti-tumor agents cannot be excluded. It has been reported that didox is a potent radiosensitizing agent overcoming Bcl-2 mediated radiation resistance in prostate cancer cell line PC-3 [271]. A combination of BCNU (an alkylating anticancer drug) and didox was proven to act in a synergistic manner in two cell lines, 9L rat gliosarcoma and DAOY human medulloblastoma cells [272, 273].


Trimidox has been shown to exert anti-proliferative activities in human leukemia cell lines. The induction of apoptosis in human ovarian carcinoma cells and HL-60 promyelocytic leukemia cells by trimidox seems to be associated with the induction of c-myc expression and activation of caspases [274, 275]. Enhanced effects of adriamycin by combination with trimidox has been observed in L1210 leukemia bearing mice [276]. Trimidox in combination chemotherapy with cisplatin and cyclophosphamide is highly synergistic in the L1210 and P388D1 leukemia mouse models [277]. Trimidox significantly decreases dCTP pools in HL-60 human promyelocytic leukemia cells and increases the antitumor effect of 1-beta-D-arabinofuranosyl cytosine (Ara-C) on the colony formation of HL-60 cells [278]. Trimidox and didox have shown efficacy in combination with Temozolomide (an alkylating agent) in the treatment of human malignant glioma cell lines [279]. Trimidox and didox suppress retrovirus-induced murine AIDS with more effectiveness and less bone marrow toxicity than hydroxyurea [280-282]. Both of the agents also showed the ability to improve the efficacy of the reverse transcriptase inhibitor didanosine (2,3didoxyinosine; ddI) and abacavir in murine AIDS model. Nitric Oxide (NO) It can inactivate RR by scavenging the tyrosyl radical of R2 [9, 283, 284]. The stimulation of NO synthesis in an adenocarcinoma cell line destroys its R2 tyrosyl radical. Thionitrites are spontaneous NO donors in neutral aqueous solutions. They inhibit RR from E. coli and murine adenocarcinoma TA3 cells [285]. EPR spectroscopy both on whole murine R2-overexpressing L1210 cells and on the pure protein showed that the tyrosyl radical of protein R2 reversibly couples to the NO radical, presumably leading to nitrosotyrosine adducts. NO can also react with R1 resulting in the nitrosation of cysteines. These results identify NO donors as a new class of inhibitors of RR with potential applications as anticancer or antiviral chemotherapy agents. Alkoxyphenols They take part in a radical redox-reaction in which the tyrosyl radical in R2 is quenched [286, 287]. In contrast to E. coli, the iron center in R2 from mouse and herpes simplex virus-2 is reduced by the inhibitors [288]. EPR spectroscopy and growth inhibition assays showed that palkoxyphenols inactivated the enzyme with high efficiency in tumor cells. The enzyme RR, and in particular the catalytically essential tyrosyl radical in the active site, is recognized as an important cellular target for growth inhibition of Novikoff hepatoma cells, human leukemia cells and melanoma cells by p-alkoxyphenols. Thus, palkoxyphenols may be considered as future antiproliferative drugs for the systemic treatment of melanoma as well as leukemia and possibly other malignancies [289]. Iron Chelators Iron is a well-known growth factor and when cells lack iron, cell proliferation stops due to specific inhibition of DNA synthesis. Therefore, iron chelators are of value as therapeutic agents in the treatment of cancer [290, 291]. An early effect of iron lacking in the cell is that RR activity decreases, because reduction of intracellular iron prevents the activation of the newly synthesized apo-R2 and the

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regeneration of the R2 iron center when apo-R2 is formed after losing iron [292]. Some iron chelates are redox active, destroying the tyrosyl radical of R2 [293]. Triapine (3-Aminopyridine-2-carboxaldehyde thiosemicarbazone, 3-AP) It is a new potent inhibitor of RR currently in phase II clinical trials for cancer chemotherapy [294-298]. The compound belongs to a class of powerful iron chelators, heterocyclic carboxaldehyde thiosemicarbazones (HCTs), which coordinate with iron through the N*-N*-S* tridentate ligand system to form a 2:1 ligand to iron molar ratio octahedral complex [299, 300]. Triapine inhibits the enzymatic activity and the tyrosyl radical of both small subunits R2 and p53R2 [294, 295, 301]. Unlike hydroxyurea, the presence of iron is required for effective enzyme inhibition and cytotoxicity, whereas deferoxamine (DFO, an iron chelator) decreases the inhibitory effects [294, 295]. Using spin trapping experiments with DMSO, in vitro RR activity assays, room temperature EPR, and antioxident protection assay, it is demonstrated that Triapine-Fe(II) complex is a much more potent inhibitor of the holoenzyme than Triapine alone and Triapine-Fe(III) complex. The complex activates molecular oxygen to give oxygen reactive species (ROS), which is responsible for quenching of the tyrosyl radical and the enzymatic activity of the small subunits [301]. Triapine-Fe(II) is much more effective at inhibiting p53R2/R1, which may be due to the different sensitivity to ROS between R2 and p53R2 [302]. However, addition of excess iron significantly regenerates p53R2/R1 resulting in a similar inhibition by Triapine for both forms of RR. Triapine has also been shown to cause DNA degradation in the presence of H 2 O 2 and iron [303, 304]. This mechanism may be similar to the antitumor drugs adriamycin and bleomycin, both of which can enter a redox cycle to generate free radicals after binding metal ions and lead to DNA damage [305, 306]. Thus, Triapine is not only a potent inhibitor of DNA synthesis and repair through RR inactivation, but also a DNA damaging agent. The two functions can combine synergistically causing cell cycle arrest and apoptosis. Triapine-related adverse events have been observed including methemoglobinemia and hypoxia, which could cause acute symptoms in patients with limited pulmonary or cardiovascular reserve [296, 301]. The druginduced ROS may oxidize ferrous hemoglobin interfering with its oxygen binding and delivery. Triapine possesses 100~1000-fold higher potency in both enzyme and tumor cell growth inhibition than hydroxyurea. The compound is fully active against wild-type and hydroxyurea-resistant tumors (L1210 leukemia cells and human KB nasopharyngeal carcinoma cells), and has broad spectrum antitumor activity in tumor xenograft models. Combinations of Triapine with various classes of DNAdamaging agents (such as etoposide, cisplatin, doxorubicin, etc.) produce synergistic inhibition of cancer cell growth, possibly resulting from preventing repair of damaged DNA [295]. Phase I clinical trials suggest that Triapine is a promising cancer chemotherapeutic drug in patients with advanced cancers not only as monotherapy but also in association with other anticancer drugs such as gemcitabine [296-298, 307-309]. Phase II trials are still ongoing.


DFO (Desferrioxamine) It is currently used in treatment of diseases related with iron overload and has been shown to have anti-proliferative effects attributed to inhibition of RR activity [310]. DFO is a hexadentate chelator and forms a 1:1 ligand –metal complex with Fe(III) that is redox-inactive [311]. Whether alone or complexed with iron, DFO has no direct effect on the tyrosyl radical. Instead, it inhibits RR through chelating intracellular iron pools and preventing the activation and regeneration of the enzyme activity [29, 304]. The inhibition of RR by DFO was reversible with the addition of iron. RR activity of both small subunits was inhibited by DFO, but p53R2 was much more susceptible to DFO than R2 [76]. This difference may be due to a more accessible iron-radical center in p53R2 to the environment, or the iron center of p53R2 may be less stable than that of R2. DFO increases the inhibition of DNA synthesis and cytotoxicity by hydroxyurea perhaps by binding the exogenous iron and inhibiting the regeneration of RR, while attenuating the inhibitory effects of HCTs because of environmental iron chelation that is necessary for the activity of Triapine. DFO has shown anti-proliferative activity against leukemia and neuroblastoma cells in vitro, in vivo and in clinical trials. However, the efficacy of DFO is severely limited due to its poor ability to permeate cell membranes and a very short serum half-life. These limitations have encouraged the development of other Fe chelators that are far more effective than DFO [312]. Polymerization Inhibition These RR inhibitors are oligopeptides with similar sequences to the C terminus of R2, which is responsible for binding to R1. The peptidomimetic drugs prevent the formation of an active RR by competitively inhibiting the binding of R2 to R1 [313-317]. The difference between the C terminus sequence of R2 of different species is important to develop highly specific inhibitors that can inhibit a parasite RR without interfering with the host RR [313, 314, 318, 319]. A herpes simplex virus (HSV) RR inhibitor, BILD 1633 SE has shown activity against cutaneous acyclovir-resistant HSV-1 infections in an athymic nude mouse model. The first work with mammals was reported that the heptapeptide, FTLDADF, identical in sequence to the last seven amino acid residues of the C terminus of the R2 subunit of mouse RR, and its N-alpha-acetyl derivative (N-Ac-FTLDADF) were able to inhibit calf thymus RR [320]. Several systematic structure-function studies, including residues modification and deletions were performed in order to understand the inhibition mechanism and to achieve better inhibitory activity [321-323]. These studies lead to the conclusion that for inhibition only a small peptide is necessary and that the acetylation of these inhibitors improves, in most of the cases, the ligand affinity and therefore the inhibition efficacy. Fmoc-tripeptides (Fmoc: a-fluorenylmethyoxycarbonyl) were also capable of inhibiting the dimerization of RR subunits [324]. The C terminal sequence of R2 is completely conserved in p53R2. Using a synthetic heptapeptide inhibition assay, it was shown that p53R2 bound to R1 through the same C terminal heptapeptide as R2. However, R2 had a 4.76-fold higher binding affinity for R1 than p53R2, which may


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explain the reduced RR activity of p53R2 relative to R2 [76]. Gene Expression Regulation RR gene expression can be regulated at transcriptional or translational levels with subunit specificity. R1 Antisense Inhibitor-- GTI-2501 This inhibitor is directed against the R1 component of RR for the potential treatment of cancer [325]. In preclinical trials, GTI-2501 was an effective anticancer agent when tested in standard mouse models bearing a variety of different human cancer lines including tumor cells derived from lung, breast, colon, kidney, ovary, pancreas and skin cancers. A phase I clinical trial in patients with lymphomas or solid tumors which have not responded to standard therapy, commenced in the US. It has been reported that herpes simplex virus type 2 growth and latency reactivation by cocultivation are inhibited with antisense oligonucleotides complementary to the translation initiation site of R1 [326]. R1-Based Gene Therapy In addition to being a component of the RR holoenzyme, R1 subunit appears to play an important role in determining the malignant potential of tumor cells. Overexpression of mouse R1 resulted in suppression of tumorigenicity and metastatic potential, whereas expression of antisense RNA, complementary to R1 mRNA, increased anchorageindependent growth of ras-transformed NIH 3T3 cells. These results suggested that elevated expression of mouse Rl leads to suppression of transformation, tumorigenicity, a n d metastatic properties of tumor cells [328]. However, the mechanism underlying the tumor-suppressing activity of R1 remains uncertain. Adenovirus-mediated R1 gene therapy has been investigated in human colon adenocarcinoma using a replication-defective recombinant adenoviral construct carrying the human R1 gene (rAd5-R1) [328]. R1 mRNA and protein were overexpressed in Colo320 HRS cells infected with rAd5-R1. Infection with rAd5-R1 significantly inhibited Colo320 HRS cell proliferation in vitro and Colo320 HRS tumor xenografts growth in CD-1 mice. These results demonstrate gene-specific antitumor effects of R1 and suggest that rAd5-R1 gene therapy has the potential to improve currently available treatments for colon cancer [329]. R2 Antisense Inhibitor-GTI-2040 It is a 20-mer oligonucleotide that is complementary to a coding region in the mRNA of R2. In vitro and in vivo studies have demonstrated that GTI-2040 decreases mRNA and protein levels of R2 and significantly inhibits the growth of a wide range of tumors and inhibits metastasis of human melanoma cells to the lungs in nude mice [330, 331]. GTI-2040 has been evaluated in phase I trials [332], and is undergoing phase II trials for the potential treatment of cancers. Besides, several other strategies have been reported for inhibiting R2 expression. R2 antisense cDNA expression by an inducible vector system can effectively decrease R2 protein expression and enzyme activity, and inhibit human oropharyngeal KB cancer cells growth [333]. An R2-targeted siRNA has been reported to be capable of


stable suppression of the R2 subunit [81]. Retrovirally mediated RNA interference targeting the R2 subunit was shown to be a novel therapeutic strategy in pancreatic cancer [334]. NOVEL DRUG DISCOVERY Strategy and Methodology for Drug Discovery In recent years, many new strategies have emerged to speed up drug discovery research, including genomic and proteomic methodologies (particularly gene expression microarrays); robotic high-throughput screening of diverse compound collections, together with in silico and fragmentbased screening techniques; new structural biology methods for rational drug design (especially high-throughput X-ray crystallography and nuclear magnetic resonance); and advanced chemical technologies, including combinatorial and parallel synthesis [335-337]. These technological advances will provide a new basis for the discovery of more effective RR inhibitors for human cancer therapy in the future. Mechanism-Based Design and Improvement for New RR Inhibitors with High Binding Affinity and Target Specificity For the last decades, intensive efforts have been put on the discovery of novel RR inhibitors. However, all known RR inhibitors have their shortcomings. For examples, hydroxyurea has low affinity for RR. Triapine has high inhibitory potency, but its ROS-based mechanism of action not only causes RR inhibition and DNA damage but also causes side effects such as methemoglobemia and hypoxia. In fact, compounds with several intracellular targets (due to an aspecific mechanism of action) have a lesser probability of discriminating between normal and tumor cells as compared to compounds with one or a few (usually related) targets [8, 9]. Besides, the inhibitory efficiency of RR by small subunit inhibitors is affected by the regeneration of the iron-tyrosyl radical center. Design of novel small subunit inhibitors that target the iron-tyrosyl radical co-factor, but with alternative inhibitory mechanisms is a challenge for future generations of drugs.

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apoptosis [49, 74, 75]. We have developed a holoenzymebased in vitro assay for the determination of the potency and subunit-selectivity of small-molecule inhibitors of RR [236]. The assay was implemented using two forms of recombinant RR (R2/R1 and p53R2/R1). It has been found that the responses of two small subunits to the inhibitors were quite different, perhaps due to differences in protein structure or due to differences in the molecular mechanisms of the inhibitors [76, 236, 301]. The inhibition of the translation of each subunit of RR can be obtained by a single oligonucleotide, complementary to its coding region in the mRNA, and is very interesting since it allows for the development of highly specific RR inhibitors [338]. CONCLUSION The clinical use of RR inhibitors has several decades of history, which has demonstrated that RR inhibitors have anti-tumor activity both as single agents and as enhancers of other anticancer drugs. Recent development of novel RR inhibitors in different categories will further contribute to cancer chemotherapy. ABBREVIATIONS RR

=

Ribonucleotide reductase

R1

=

Large subunit of RR

R2

=

Small subunit of RR

EPR =

ACKNOWLEDGEMENTS We thank Leila Su of City of Hope for help in drawing protein structures. REFERENCES [1] [2] [3] [4]

Development of Subunit-Specific Inhibitors for two Forms of RR R2 and p53R2 play different roles in cells. Development of subunit-specific inhibitors for different clinical applications is an attractive direction. Specific inhibition of R2 is a potential therapeutic strategy to kill cancer cells. Upon DNA damage, cancer cells often cannot induce p53R2 due to the lack of p53, whereas normal cells can repair their DNA with help from induced p53R2. Thus, use of genotoxic chemotherapeutic agents in combination with R2 inhibitors is an attractive treatment plan [18, 80]. On the other hand, inhibitors specific for p53R2 may be useful for targeting tumors that overexpress p53R2, because its inhibition increases the sensitivity of cells to DNA damaging agents [49, 74, 81, 82]. In cells that retain the p53 wild-type gene, it is expected that inactivation of p53R2dependent DNA synthesis would activate p53-dependent

Electron paramagnetic resonance

[5] [6]

[7]

[8] [9] [10]

Reichard, P. From RNA to DNA, why so many ribonucleotide reductases? Science 1993, 260, 1773-1777. Reichard, P.; Ehrenberg, A. Ribonucleotide reductase--a radical enzyme. Science 1983, 221, 514-519. Jordan, A.; Reichard, P. Ribonucleotide reductases. Annu. Rev. Biochem. 1998, 67, 71-98. Kolberg, M.; Strand, K. R.; Graff, P.; Andersson, K. K. Structure, function, and mechanism of ribonucleotide reductases. Biochim. Biophys. Acta 2004, 1699, 1-34. Stubbe, J.; van der Donk, J. W. A. Protein radicals in enzyme catalysis. Chem. Rev. 1998, 98, 705-762. Hogbom, M.; Stenmark, P.; Voevodskaya, N.; McClarty, G.; Graslund, A.; Nordlund, P. The radical site in chlamydial ribonucleotide reductase defines a new R2 subclass. Science 2004, 305, 245-248. 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, 1289212896. Smith, B. D.; Karp, J. E. Ribonucleotide reductase: an old target with new potential. Leuk. Res. 2003, 27, 1075-1076. Nocentini, G. Ribonucleotide reductase inhibitors: new strategies for cancer chemotherapy. Crit. Rev. Oncol. Hematol. 1996, 22, 89-126. Holland, K. P.; Elford, H. L.; Bracchi, V.; Annis, C. G.; Schuster, S. M.; Chakrabarti, D. Antimalarial activities of polyhydroxyphenyl and hydroxamic acid derivatives. Antimicrob. Agents Chemother. 1998, 42, 2456-2458.

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Received: October 15, 2005

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Revised: February 1, 2006

Accepted: March 17, 2006