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Oncogene (2015) 34, 2011–2021
© 2015 Macmillan Publishers Limited All rights reserved 0950-9232/15
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REVIEW
Ribonucleotide reductase and cancer: biological mechanisms
and targeted therapies
Y Aye1,2, M Li3, MJC Long4 and RS Weiss3
Accurate DNA replication and repair is essential for proper development, growth and tumor-free survival in all multicellular
organisms. A key requirement for the maintenance of genomic integrity is the availability of adequate and balanced pools of
deoxyribonucleoside triphosphates (dNTPs), the building blocks of DNA. Notably, dNTP pool alterations lead to genomic instability
and have been linked to multiple human diseases, including mitochondrial disorders, susceptibility to viral infection and cancer.
In this review, we discuss how a key regulator of dNTP biosynthesis in mammals, the enzyme ribonucleotide reductase (RNR),
impacts cancer susceptibility and serves as a target for anti-cancer therapies. Because RNR-regulated dNTP production can
influence DNA replication fidelity while also supporting genome-protecting DNA repair, RNR has complex and stage-specific roles in
carcinogenesis. Nevertheless, cancer cells are dependent on RNR for de novo dNTP biosynthesis. Therefore, elevated RNR expression
is a characteristic of many cancers, and an array of mechanistically distinct RNR inhibitors serve as effective agents for cancer
treatment. The dNTP metabolism machinery, including RNR, has been exploited for therapeutic benefit for decades and remains an
important target for cancer drug development.
Oncogene (2015) 34, 2011–2021; doi:10.1038/onc.2014.155; published online 9 June 2014
DEOXYRIBONUCLEOSIDE TRIPHOSPHATE (dNTP) POOLS AND
GENOMIC INTEGRITY
The importance of ribonucleotide reductase (RNR) for genome
maintenance relates to its central role in regulating dNTP levels. In
mammalian cells, total dNTP pool sizes peak during S-phase to
support nuclear DNA (nDNA) replication and are roughly 10-fold
lower in G0/G1, when dNTPs are needed for DNA repair and
mitochondrial DNA (mtDNA) synthesis.1,2 As DNA polymerase
substrates, dNTPs influence several aspects of the replication
program, including origin choice, fork speed, inter-origin distance,
and dormant origin usage.3–5 Failure of cells to maintain
appropriate dNTP concentrations can be highly detrimental,
leading to DNA breaks, mutagenesis and cell death.6 During
cancer development, uncoordinated cell proliferation can lead to
insufficient dNTPs that cause replication stress and further
promote genomic instability.7,8 Conversely, elevated dNTP pools
also contribute to increased mutagenesis.1,9,10
Imbalanced dNTP pools enhance mutagenesis mainly by DNA
misinsertion and impaired proofreading.6,9 DNA misinsertion can
result from competition between dNTPs for pairing with the
template base, and a dNTP present in excess can be readily
misincorporated. DNA polymerase proofreading is reduced in
the presence of elevated dNTP concentrations through a
phenomenon known as the next-nucleotide effect.11 Under such
conditions, DNA chain extension following a base misinsertion
proceeds before the mismatched nucleotide can be removed.12–14
dNTP imbalances can also stimulate frameshift mutations by
facilitating correct base-pairing following template slippage or
misalignment.15
1
RNR AND dNTP BIOSYNTHESIS
dNTPs can be generated through de novo and salvage pathways. In
mammals, RNR catalyzes the rate-limiting step of the de novo
pathway, reducing the 2′ carbon of a ribonucleoside diphosphate
(NDP) to produce the corresponding deoxy (d)NDP. Subsequently,
dNDPs are phosphorylated by nucleoside diphosphate kinase
(NDPK) yielding dNTPs.16 Importantly, the de novo biosynthesis of
deoxythymidine triphosphate (dTTP) is much more complex,
requiring the conversion of deoxyuridine monophosphate to
deoxythymidine monophosphate (dTMP) by thymidylate synthase
followed by phosphorylation by thymidylate phosphate kinase
(TMPK) and then NDPK. Deoxyuridine monophosphate is generated
either from deoxyuridine triphosphate (dUTP) or deoxycytidine
monophosphate, the biosynthesis of both of which is dependent
on RNR-catalyzed reduction of the corresponding nucleoside
diphosphates.6 RNR enzymes are present in all organisms and
feature-conserved radical-mediated nucleotide reduction.17 Mammalian RNRs, the focus of this review, consist of two subunits α and
β, that associate to form the holoenzyme (Figure 1). α contains the
catalytic (C) site and two different allosteric sites. β harbors a di-iron
cofactor and tyrosyl radical (Y•) essential for RNR activity. Evidence
suggests that catalytically active RNR is minimally an α2β2
complex.18–20 RNR oligomerization is influenced by (d)NTP/ATP
binding to α allosteric sites. The negative regulator dATP induces
inactive α6 (or α6β2) states, highlighting a role for higher-order
oligomeric associations as a regulatory mechanism for RNR.18,21–24
However, as discussed further below, the precise nature of RNR
oligomeric states in the presence of allosteric activators remains
unsettled (Supplementary Table S1).
Department of Chemistry and Chemical Biology, Cornell University, Ithaca, NY, USA; 2Department of Biochemistry, Weill Cornell Medical College, New York, NY, USA;
Department of Biomedical Sciences, Cornell University, Ithaca, NY, USA and 4Graduate Program in Biochemistry, Brandeis University, Waltham, MA, USA. Correspondence:
Assistant Professor Y Aye, 224 Baker Laboratory of Chemistry & Chemical Biology, Ithaca, NY 14853, USA or Associate Professor RS Weiss, Department of Biomedical Sciences,
Cornell University, Veterinary Research Tower, Ithaca, NY 14853, USA.
E-mail: [email protected] or [email protected]
Received 6 January 2014; revised 25 April 2014; accepted 26 April 2014; published online 9 June 2014
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repair.30–33 dNTPs for mtDNA synthesis can also come from
salvage pathways localized within mitochondria,34,35 and RNR
activity has been identified within mitochondria as well.36
In humans, p53R2 mutations result in mitochondrial disease,
including mtDNA depletion syndrome, a lethal condition in
which patients have only 1–4% residual mtDNA in muscle;30,37
mitochondrial neurogastrointestinal encephalopathy, a neurodegenerative disorder associated with mtDNA depletion;38
and autosomal-dominant progressive external ophthalmoplegia,
characterized by accumulation of multiple mtDNA deletions in
postmitotic tissues.39 Furthermore, p53R2 knockout mice show
mtDNA depletion and die due to renal failure.30,40 Together, these
findings indicate that the de novo dNTP biosynthesis mediated
by RNR is essential to maintain not only nuclear but also
mitochondrial genome integrity.
Figure 1. RNR and de novo dNTP biosynthesis. Mammalian RNR
enzymes function to reduce the 2' carbon of NDPs to generate
dNDPs that are subsequently phosphorylated by nucleoside diphosphate kinase, yielding dNTPs for nuclear and mitochondrial DNA
replication and repair. RNR consists of an α subunit encoded by Rrm1
and a β subunit encoded by Rrm2 or p53R2. The subunits interact to
form hetero-oligomers, with the active enzyme believed to adopt
α2β2 quaternary state. α contains the catalytic site (C), an activity site
(A) that governs overall RNR activity through interactions with dATP
(inhibitory) or ATP (stimulatory) and a specificity site (S) that
determines substrate choice. Each β subunit contains a μ-oxobridged di-nuclear iron center (Fe-O-Fe) and protein tyrosyl radical
(Y-O•) that is transferred to the α C site for catalysis. The simplified
schematic depictions of RNR holoenzyme structures shown here
were guided by structural studies.18,21
NDP reduction to dNDP takes place in the α C site and requires
the unpaired electron initially localized as Y• in β (Figure 1).19
Although the di-iron center is present in each β monomer, a single
Y• is generated per β2.17 Protein radical formation occurs via
reaction of the diferrous ions with molecular oxygen in a multistep
process.17,19 How diferrous ions are loaded into β2, including
potential involvement of iron chaperones, remains unknown.
In vitro, β2 can exist in three distinct states: apo-β2 lacks both the
di-iron center and Y•; met-β2 contains the di-iron center but with
Y• reduced; and holo-β2 houses a diferric–Y• cofactor (the
‘metallocofactor’) and is the only state that, when complexed
with α2, is active (Supplementary Figure S1). The β Y• is transferred
through a proton-coupled electron transfer chain to generate a
transient thiyl radical (S•) within the α C site.19 S• initiates
reduction of the NDP ribose ring, ultimately generating dNDP
and resulting in disulfide bond formation on α. Thioredoxin or
glutaredoxin re-reduces the oxidized cysteines on α, facilitating
turnover.25
In mammals, α is encoded by the gene Rrm1, whereas two
different genes, Rrm2 and p53R2, encode distinct β isoforms.
p53R2 was identified as a p53-inducible, DNA damage-responsive
gene.26 Mouse p53R2 and RRM2 share 81% identity and domain
conservation except for amino-terminal residues required for cell
cycle stage-specific RRM2 degradation.27–29 RRM1–RRM2 holoenzyme provides dNTPs for S-phase nDNA replication and repair in
proliferating cells, whereas RRM1–p53R2 contributes dNTPs for
nDNA repair in quiescent cells as well as mtDNA replication and
Oncogene (2015) 2011 – 2021
RNR ACTIVITY REGULATION
The critical importance of dNTP levels demands that RNR is
tightly regulated through several distinct mechanisms, including
allosteric and oligomeric regulation, as well as alterations of the
level and localization of RNR subunits. Allosteric modulation
involves effector binding to two separate allosteric sites in α, a
specificity (S) site that regulates substrate choice and an activity
(A) site that regulates enzyme activity (Figure 1).29 Substrate
specificity is determined by the binding of ATP, dATP, dTTP or
dGTP (deoxyguanosine triphosphate) to the S site; the mechanism
has been extensively reviewed.29,41 Overall RNR activity is
controlled by (d)ATP binding to the A site, with elevated dATP:
ATP ratios associated with inhibition. α hexamerization is coupled
to dATP-induced inhibition.21,22,24,41
A second major mechanism for RNR regulation occurs through
the cell cycle stage-specific control of RNR protein levels. In
mammals, RNR activity is induced during S phase.42–44 Rrm1 gene
expression is mainly regulated transcriptionally, being negligible
in G0/G1 and peaking in S phase.45 RRM1 protein, however,
remains constant throughout the cell cycle owing to its long halflife.42,46,47 RRM2 is regulated both transcriptionally and by protein
degradation. Similar to that of Rrm1, Rrm2 transcription is minimal
in G0/G1 and peaks in S phase.48 Unlike RRM1, RRM2 abundance
correlates with its mRNA level.42,44 Upon S phase exit, RRM2
protein is degraded through two ubiquitin ligases, the Skp1/
Cullin/F-box (SCF) complex and the anaphase-promoting complex
(APC).27,28 During G2 phase, RRM2 is recognized by cyclin F, an SCF
ubiquitin ligase F-box protein, and targeted for degradation.28
During mitosis/G1 phase, RRM2 is degraded by the Cdh1-APC
complex that recognizes a KEN box motif at the RRM2
N-terminus.27 By contrast, p53R2, which lacks a KEN box, is
continuously expressed throughout the cell cycle. After DNA
damage, p53R2 is transcriptionally induced in a p53-dependent
manner and with RRM1 forms an active holoenzyme, providing
dNTPs for DNA repair.26,29,33
RNR subcellular localization affords an additional layer of
regulatory control, although this aspect of mammalian RNR
biology remains controversial. The bulk of RRM1 and RRM2
constitutively localize to the cytoplasm and produce dNTPs that
diffuse into the nucleus for DNA replication.49,50 However, there is
an increasing evidence that RRM1 and RRM2 accumulate at DNA
damage sites in the nucleus.28,51–53 This accumulation is
dependent on interaction between RRM1 and the DNA damage
response protein Tip60.51,52
The work forging a link between RNR translocation and local
dNTP production at DNA damage sites is appealing. However, a
cautionary note is required, as dNTP synthesis requires multiple
enzymes. The reports linking DNA damage to translocation of
dNTP production machinery have focused on only one or two key
players, typically RNR subunits. It is unknown whether the whole
dNTP production pathway responds en masse to DNA damage or
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Y Aye et al
whether increased dNTP synthesis actually occurs at DNA repair
sites. Evidence does exist that the de novo thymidylate synthesis
pathway forms a scaffolded multienzyme complex at replication
sites in the nucleus.54 With respect to RNR, it is possible that
translocation of this rate-limiting enzyme to DNA damage sites is
sufficient to affect local dNTP pools provided all other enzymes
required for dNTP synthesis are within the nucleus. An alternative
hypothesis related to DNA damage-induced RNR translocation is
that RNR has non-nucleotide reduction properties that promote
DNA repair. Parsing out the mechanisms at play during DNA
damage responses will require careful analysis and ultimately new
ways to accurately measure local dNTP fluxes in cells.
RNR AND DNA DAMAGE RESPONSES
RNR genes were among the first DNA damage-inducible genes
to be identified.55 In mammalian cells, genotoxins such as
chlorambucil and ultraviolet light induce Rrm1 and Rrm2
expression by approximately 10-fold.56,57 RRM2 protein stability
is also DNA damage responsive,44 at least in part, through
ATR-dependent downregulation of cyclin F-mediated RRM2
degradation.28 The discovery of p53R2 further connected DNA
damage responses and RNR activity and also resolved the
question of how DNA repair in quiescent cells could be supported
when RRM2 is undetectable. p53 binds the first p53R2 intron
and stimulates p53R2 expression.26,58 In addition, DNA damageinduced phosphorylation by ATM stabilizes p53R2 and confers
resistance to DNA damage.59 Inhibition of p53R2 expression in
cells that have an intact p53-dependent DNA damage checkpoint
reduces RNR activity, DNA repair and cell survival after genotoxin
exposure.26,60 p53R2-null mouse fibroblasts also show severely
attenuated dNTP pools under oxidative stress.40
One aspect of the DNA damage response influenced by
RNR activity is repair pathway choice.61,62 RNR-mediated dNTP
production after DNA damage fuels DNA synthesis during
homologous recombinational repair, whereas RNR inhibition
promotes break repair by non-homologous end joining,
which does not require extensive DNA synthesis.62 However,
RNR-mediated dNTP pool increases are accompanied by higher
mutation rates, which may result from reduced fidelity
of replicative polymerases and/or activation of error-prone
translesion DNA synthesis at elevated dNTP levels.10,63–65
Despite the well-documented increase in RNR subunit levels after
DNA damage, there remains uncertainty about the extent to which
there is a corresponding change in dNTP pools after genotoxic
stress. In non-proliferating cells, a slow, fourfold accumulation of
p53R2 after DNA damage results in less than a two-fold increase in
dNTP pools, a modest change relative to that which occurs upon
S-phase entry.47 Moreover, logarithmically growing cells do not
show significant dNTP pool increases upon DNA damage. This may
reflect the action of additional mechanisms influencing dNTP levels
after DNA damage. It also remains possible that dNTP biosynthesis
is compartmentalized at damage sites during the DNA damage
response,51 such that measurements of total cellular dNTP levels fail
to reveal local changes.
RNR DEREGULATION AND GENOMIC INSTABILITY
Because proper dNTP levels are essential for genomic integrity,
RNR deregulation is mutagenic.1,10,63 Initial insights into the
consequences of mammalian RNR deregulation came from
analysis of mouse T-lymphosarcoma cells selected for deoxyguanosine resistance and determined to have a mutation in the A site
(RRM1D57N) that disrupts dATP-mediated enzyme inhibition.
The mutagenized cells from which RRM1D57N was cloned show
increased dNTP levels and a 40-fold increase in mutation rate.66–68
RRM1D57N overexpression in CHO cells was subsequently shown to
cause a 15–25-fold increase in spontaneous mutation frequency,
© 2015 Macmillan Publishers Limited
although no dNTP pool alterations could be identified.67 Elevated
RNR activity has also been identified in hydroxyurea (HU)-resistant,
RRM2-overexpressing mouse cell lines. In one study with
fibroblasts, a 3–15-fold increase in RNR activity was accompanied
by moderate changes in dNTP pools.69 By contrast, the
HU-resistant, RRM2-overexpressing mouse mammary tumor TA3
cell line shows a 40-fold increase in enzymatically active RRM2,
estimated based on increased Y• content, but no detectable dNTP
pool changes relative to parental cells.43,44 Therefore, the elevated
mutagenesis associated with RNR activity alterations has not been
definitively linked to dNTP pool perturbations. Failure to detect
dNTP pool changes in cells with altered RNR activity could be
because even relatively small changes in dNTP pools can be highly
mutagenic.70 Alternatively, a specific interaction between RNR and
the DNA replication or repair machinery may be involved as noted
above, so that biologically important localized pool alterations are
obscured during analysis of total intracellular dNTP levels. It is also
noteworthy that measurement of holocomplex activity in vivo is
challenging, because cell lysis can disrupt complex equilibria
between subunits and alter cellular (d)NTP allosteric modulator
concentrations.
RNR IN CANCER
Uncontrolled proliferation is a defining feature of cancers and
must be supported by a sufficient dNTP supply. Cancer cells
undergo metabolic reprogramming such that glucose is no longer
metabolized to maximize ATP production as in normal cells but
instead is used to drive the production of macromolecules for cell
replication, including dNTPs.71 Studies of rat hepatomas in the
1970s established that RNR activity is highly correlated with cancer
growth rate; 200-fold differences in enzyme activity were
observed between fast- and slow-growing tumor cells.72 RRM2
overexpression has been observed in gastric, ovarian, bladder and
colorectal cancers.73–77 RRM2 expression is correlated with tumor
grade for both breast and epithelial ovarian cancers, suggesting a
role for RNR in supporting rapid cell division of high-grade
tumors.78,79 Similarly, RRM2 levels are low in benign skin lesions
but are significantly higher in malignant melanoma, with high
RRM2 expression additionally correlating with poor overall
survival.80 Increased p53R2 expression has also been reported
in cancers, including melanoma, oral carcinoma, esophageal
squamous cell carcinoma and non-small cell lung cancer
(NSCLC).81–85 To broadly assess how RNR is affected in a large
collection of human cancers, we surveyed RNR gene expression in
human cancers using the ONCOMINE database (Figure 2).
Remarkably, RRM2 was among the top 10% most overexpressed
genes in 73 out of the 168 cancer analyses. These include sarcoma
and cancers of the bladder, brain and central nervous system,
breast, colorectal, liver and lung. Similarly, RRM1 was among the
top 10% most-overexpressed genes in 30 out of the 170 studies,
including brain and central nervous system cancer, lung cancer
and sarcoma. By contrast, p53R2 was among the top 10% most
overexpressed genes in only 5 out of the 90 studies.
Elevated RNR expression in cancers could be secondary to cell
cycle alterations, as many neoplasms show an increased S-phase
fraction, or the direct result of gene amplification or other genetic
or epigenetic alterations. Therefore, we analyzed RNR gene copy
number changes using the TCGA database (Supplementary
Figure S2). Among the RNR genes, RRM2B was the most frequently
affected by copy number changes and typically showed gains, in
accord with a recent report on human breast cancers.86 It should
be noted that the RRM2B locus (8q22.3) is located near the C-MYC
oncogene (8q24). Most RRM2B copy number increases in cancers
are accompanied by C-MYC amplification, raising the possibility
that they are simply passenger events. However, RRM2B amplification does occur independently of C-MYC amplification at low
frequency in breast, colon, ovarian, prostate and uterine cancers,
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2013
Ribonucleotide reductase and cancer
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2014
Figure 2. RNR expression in human cancers. The bar graph illustrates
altered RNR expression in human cancers. Data were retrieved from
the ONCOMINE cancer gene expression database (version 4.4.4.4,
search done on 27 November 2013). The y axis represents the
number of analyses with differences in gene expression for the gene
of interest. Dark red, red and pink: number of analyses with the gene
of interest among the top 1, 5 and 10% most overexpressed genes,
respectively, in a given study. Dark green, green and light green:
number of analyses with the gene of interest among the top
1, 5 and 10% most underexpressed genes, respectively, in a given
study. RRM2 is among the top 10% of the most overexpressed genes
in 73 out of the 168 analyses, RRM1 is among the top 10% in 30 out
of the 170 studies and RRM2B (encoding p53R2) is among the top
10% in 5 out of the 96 cases. In addition, RRM2 is among the top
10% of the most underexpressed genes in 7 out of the 168 studies,
and RRM1 is among the top 10% in 6 out of the 170 studies. The
expression level of the c-MYC proto-oncogene is shown for
comparison. c-MYC is among the top 10% of the most overexpressed
genes in 40 out of the 174 analyses and among the top 10% of the
most underexpressed genes in 17 out of the 174 studies.
as well as glioblastoma. This correlates well with increased p53R2
expression in breast and prostate cancers (Figure 2) and supports
a potential role for p53R2 as a tumor promoter. RRM2 is amplified
at low frequency in breast, ovarian, prostate and uterine cancers,
malignancies in which RRM2 gene expression is also increased.
RRM1 also underwent rare copy number changes in cancers, with
the direction of change depending on tumor type. This may
reflect the complex and stage-specific roles RRM1 can have in
tumorigenesis, as discussed below.
RRM1: A TUMOR SUPPRESSOR THAT CAN CONFER
CHEMORESISTANCE
RNR-mediated dNTP biosynthesis can have varied and potentially
opposing effects on tumorigenesis. Altered dNTP pools can impair
DNA replication fidelity, leading to tumor-promoting mutations.
On the other hand, RNR-supported dNTP production can protect
against mutations by facilitating DNA repair. Following malignant
transformation, the same repair mechanisms can protect cancer
cells against potentially lethal stresses, such as those caused by
genotoxic chemotherapies. These complexities are reflected in the
data concerning RRM1 in cancer. Several studies point to RRM1 as
a suppressor of tumor initiation. RRM1 overexpression in cultured
Oncogene (2015) 2011 – 2021
cells leads to reduced transformation and suppression of
tumorigenicity and lung metastasis in vivo.87 In addition, RRM1
overexpression in human lung cancer cells induces phosphatase
and tensin homolog (PTEN) expression and suppresses migration
and invasion, as well as overall tumorigenicity and metastasis
following xenotransplantation.88 Rrm1-overexpressing transgenic
mice show reduced urethane-induced lung tumorigenesis,89
although in an independent study Rrm1 overexpression was not
found to alter the background incidence of spontaneous lung
neoplasms in mice.90 In human NSCLC patients who underwent
surgical resection but received no other form of treatment, highlevel tumor-associated RRM1 expression correlated with longer
lifespan and later disease recurrence.91 Co-expression of RRM1
and excision repair protein ERCC1 was significantly associated
with disease-free and overall survival, especially in patients who
underwent lung cancer surgery at early stages, although aspects
of this study were later questioned.92 The published mechanisms
behind tumor suppression by α, such as PTEN induction, require
further investigation and may indicate that functions other than
dNTP production are required for α tumor-suppressor activity.
Additional animal studies also are needed to address the distinct
outcomes reported concerning how α overexpression impacts
lung tumorigenesis.
Consistent with a role for RNR in promoting DNA repair, high
level RRM1 expression also correlates with poor responses to
platinum drugs, which induce DNA damage against which high
RRM1 levels afford protection, and gemcitabine, which directly
targets RRM1.93–102 Although not all studies have found RRM1
levels to affect patient survival after gemcitabine treatment, there
is general consensus that low RRM1 levels improve responsiveness
to gemcitabine therapy. Additionally, RRM1 overexpression
has been reported in gemcitabine-resistant cancer cells.103–106
Consequently, RRM1 expression has been proposed as a
biomarker in patients with advanced NSCLC to individualize
chemotherapy.107
TUMOR PROMOTION BY RRM2 AND P53R2
In contrast to the tumor-suppressing roles of RRM1, RRM2
has oncogenic activity. For instance, RRM2 cooperates with
oncoproteins to increase focus formation and anchorageindependent growth in mouse cells.108,109 Elevated RRM2 expression in human carcinoma cells correlates with higher invasive
potential in vitro 110 as well as decreased thrombspondin-1 and
increased VEGF production, suggesting that RRM2 can promote
tumor angiogenesis.111
The role of p53R2 in mutagenesis and tumorigenesis is less
clear-cut. Based on the p53-inducible nature of p53R2 expression
and its role in DNA repair, it was originally suggested that p53R2
would have tumor-suppressor activity.26 Under genotoxic stress,
p53R2 promotes p21 accumulation and G1 arrest, which may
facilitate repair and prevent mutation accumulation.112 p53R2
expression is negatively correlated with colon adenocarcinoma
metastasis.113 Similarly, elevated p53R2 expression suppresses
cancer cell invasiveness and correlates with markedly better
survival in colorectal cancer patients.114 Nevertheless p53R2 is
highly expressed in some human cancers as noted above, and
experimental suppression of p53R2 expression impairs cancer cell
proliferation in vitro.81
Widespread overexpression of either Rrm2 or p53R2, but not
Rrm1, in transgenic mice induces NSCLC but not other tumors.90
RNR-induced lung neoplasms arise with relatively long latency,
histopathologically resemble human papillary adenomas and
adenocarcinomas and are associated with K-ras proto-oncogene
mutations. Rrm2 or p53R2 overexpression causes an elevated
mutation frequency in cultured cells, suggesting that RNR-induced
neoplasms could arise through a mutagenic mechanism.
Consistent with this possibility, combining RNR deregulation
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with DNA mismatch repair defects synergistically increases
mutagenesis and tumorigenesis.90
The available data raise fundamental questions about the
transforming activity associated with the RNR-β subunit. Given
that increased RNR activity can lead to altered dNTP pools, one
possibility is that RNR-β overexpression promotes error-prone
DNA synthesis, including increased frequency of base misinsertions, insertion–deletion events and uracil misincorporation,
as RNR also reduces UDP to dUDP. Recent evidence indicates
that RNR-generated dNTPs are also necessary for neoplastic cells
to avoid oncogene-induced senescence.80,115,116 Senescence
induced by activated Ras expression or Myc depletion was found
to be associated with repression of Rrm2 expression and could be
bypassed by treatment with exogenous nucleosides or ectopic
Rrm2 expression. Thus aberrant RNR expression could enable cells
to overcome this important barrier to transformation. Apoptosis
is another cancer-related pathway that could be impacted by
RNR-mediated dNTP biosynthesis. In particular, (d)ATP binding to
Apaf-1 and cytochrome c regulates the formation of the
apoptosome, which is crucial for caspase activation and downstream apoptotic events.117,118
Reactive oxygen species (ROS) also may contribute to RNRinduced mutagenesis and transformation. Because Y• within RRM2
is short-lived,119 its lability and reactivity could lead to secondary
reactive species when RRM2 is overproduced.120,121 It remains
unclear whether RRM2 can indeed propagate ROS.119 Involvement
of ROS would be compatible with the lung specificity of
tumorigenesis in RRM2-overexpressing mice as well as the
observed synergy between RRM2 deregulation and the mismatch
repair system, which responds to both base mismatches and
oxidative DNA damage.90 Interestingly, the cooperativity between
Rrm2 and activated oncogenes in inducing transformation is
independent of ribonucleotide reduction.108 Moreover, RRM2 and
p53R2-overexpressing cells show an increased frequency of G → T
transversions, a signature of oxidative DNA damage.90
As noted for RRM1, there also is evidence that RRM2 levels in
cancer cells can influence therapeutic responses. Suppression of
RRM2 expression sensitizes cancer cells to both RNR inhibitors and
cisplatin.110,122 Elevated p53R2 expression correlates with resistance to genotoxic therapies such as radiation and 5-flurouracil,
whereas p53R2 knockdown sensitizes cells to DNA-damaging
agents.81,123–125 Interestingly, p53R2 levels in cancer cells do not
correlate with sensitivity to RNR-targeted therapeutics, such as
triapine.104,122,126 RRM2 is part of a 12 gene set that is predictive of
benefit from adjuvant chemotherapy in NSCLC and is associated
with poor prognosis.127 RRM2 overexpression is also a marker of
poor prognosis in ovarian cancer patients receiving gemcitabine
or other cytotoxic therapies,128 and low RRM2 expression in
pancreatic cancers is predictive of greater response to
gemcitabine.129,130 Similarly, the absence of p53R2 expression
is associated with responsiveness to chemoradiotherapy for
esophageal squamous cell carcinoma.131
Another intriguing example of therapeutic response modulation
by RNR is found in the case of small-molecule inhibitors of TMPK,
which converts dTMP to dTDP in dTTP biosynthesis.52,132 Both RNR
and TMPK localize to DNA damage in the nucleus, and their
activities impact the relative levels of dUTP (produced via RNR)
and dTTP (produced via TMPK) at repair sites. When the dUTP:
dTTP ratio is low, there is minimal uracil misincorporation during
DNA repair. However, if the dUTP:dTTP ratio is elevated, high
levels of uracil misincorporation can lead to futile repair cycles,
DNA breakage and cell death.52 These observations are the basis
for a therapeutic strategy that combines TMPK inhibition with lowdose chemotherapy. It follows that tumors with elevated RNR
activity, dictated in many cases by RRM2 expression, would be
especially susceptible to such a strategy. This prediction holds true
in initial cell culture analyses and offers hope for a targeted
approach that could be effective against the large fraction of
© 2015 Macmillan Publishers Limited
cancers with high-level RRM2 expression. Nevertheless, because
dNTP biosynthesis is a complex process requiring multiple
enzymes, further work is necessary to fortify the link between
TMPK inhibition and uracil misincorporation. For instance, the
presence of another enzyme necessary for dNTP production,
NDPK, has not been established in the same complexes. Whether
enzymes other than the rate-limiting factor RNR must be present
directly at damage sites to allow local dUTP accumulation remains
unknown, but one possibility is that without NDPK recruitment the
dNTP precursors dTDP and dUDP would instead accumulate.
RNR AND CANCER THERAPY
Because RNR is the gatekeeper of dNTP homeostasis,29,133 the
enzyme is long recognized as a cancer therapeutic target.
Although small-molecule inhibitors continue to represent the
primary strategy for RNR inhibition since the last comprehensive
review, studies have uncovered new approaches to smallmolecule-based subunit-specific activity modulation and highlight
the potential of gene therapy.134,135 Small-molecule RNR inhibitors
in active clinical use fall into two classes: nucleoside analogs and
redox active metal chelators (Figure 3). The former class targets
RRM1 (α), in line with the ability of α to bind nucleotides, whereas
the latter group targets RRM2 (β), consistent with the dependence
of β on a redox active metallocofactor.19 The existence of these
two distinct drug classes reflects the inherent biochemical
diversities of the two subunits, both of which are required for NDP
reduction.29,136 Consequently, most drugs targeting α show little
cross reactivity with β and vice versa.
RNR-α INHIBITORS
One of the first nucleoside analogs targeting α to be clinically
approved was gemcitabine (Gemzar, F2C; Figure 3a), which
continues to be a frontline therapy against pancreatic, bladder and
lung cancers.137 RNR inactivation by gemcitabine has been
extensively reviewed.138 The active form, the substrate analog
diphosphate (F2CDP), is an irreversible, suicide inactivator of α,
which results in a covalent complex between α and the sugar of F2
CDP.139 Despite being stereotyped as an α-targeting drug, F2CDP
cannot inactivate α without β; holocomplex assembly is obligatory
for transient S• formation within the substrate-binding C site on α
that then reacts with substrate analog F2CDP, affording a reactive
intermediate capable of irreversibly alkylating α (Figure 3a).
The second approved α-targeting drug to have its RNRtargeting mechanism elucidated was clofarabine (Clolar, ClF;
Figure 3b and Supplementary Figure S3).22,23 ClF belongs to a
clinically successful class of nucleoside prodrugs, including
cytarabine (ara-C), nelarabine, azacitidine, decitabine, cladribine
and fludarabine, used to treat hematological malignancies.140 The
majority of these antimetabolites are thought to target RNR
as part of their cytotoxic spectrum; however, the molecular
mechanisms underlying RNR inhibition remain poorly defined
except for ClF. As the triphosphate (ClFTP) is the most abundant
ClF metabolite in cells, it was initially postulated that α inhibition
occurred through α A site binding by ClFTP, similarly to the
feedback inhibitor dATP.140 However, both the diphosphate and
triphosphate (ClFD(T)P) emerged to be α inhibitors in vitro. ClFTP is
an A-site-binding reversible inhibitor. RNR-α can be completely
inhibited by brief exposure to saturating amounts of ClFTP, but
activity is regained upon prolonged drug exposure, resulting in a
steady-state level of 50% activity. Whether this restoration of
activity is important for ClF’s drug efficacy remains unaddressed.
Unexpectedly, the substrate analog diphosphate, ClFDP, is a
reversible α inhibitor that binds the C site with slow release
properties. Most strikingly, although ClFDP occupies the same site
as F2CDP, α does not chemically engage with ClFDP, and the drug
Oncogene (2015) 2011 – 2021
Ribonucleotide reductase and cancer
Y Aye et al
2016
Figure 3. Inhibitors targeting multiple characteristics of RNR.
(a, b) Nucleotide analog inhibitors interacting with α. (a) Gemcitabine (F2C) is approved for the treatment of lung, pancreatic, breast
and ovarian cancers. RNR catalytic turnover is triggered by forward
proton-coupled electron transfer, resulting in transient C-S• formation that initiates NDP reduction. The active diphosphate (F2CDP)
form, as a substrate analog, can interact with the C-S• to form a
substrate radical that subsequently decomposes, resulting in
enzyme inactivation. Inactivation leads to covalent crosslinking
between α and the sugar of decomposed F2CDP. Formation of
activated RNR holocomplex bearing the transient C-S• is a
prerequisite for F2CDP inactivation. (b) Clofarabine (ClF) is used
for the treatment of refractory pediatric leukemias. The active
diphosphate and triphosphate [ClFD(T)P] forms are reversible
inhibitors that exclusively target α. Inhibition in both cases is
coupled with the assembly of hexameric, catalytically non-viable
quaternary states that are induced by ClFD(T)P either in the
presence or absence of β. See also Supplementary Figure S3.
(c, d) Small-molecule inhibitors interacting with β. (c) HU is used in
the treatment of chronic myeloid leukemia, melanoma, head and
neck and refractory ovarian cancers. Treatment of β with HU leads to
the formation of catalytically incapable apo-β2 that lacks both Y-O•
and the di-iron center. (d) Triapine (3AP) is currently being evaluated
in clinical trials. β − Specific targeting is mediated by the active
form Fe(II)-(3AP) that reduces Y-O•, converting holo-β2 into the
catalytically incapable variant, met-β2 (Y-OH). C-SH, reduced Cys;
C-S•, Cys radical; Y-OH, Tyr; Y-O•, Tyr radical.
Oncogene (2015) 2011 – 2021
can be recovered intact from α even following prolonged
incubation.22
ClF is a hybrid of its predecessors cladribine and fludarabine,
two adenine-containing nucleosides used previously for leukemic
reticuloendotheliosis and hairy cell leukemia or leukemia and
lymphoma, respectively.140–142 These drugs were developed as
analogs of deoxyadenosine, which selectively kills lymphocytes.143
The triphosphate of cladribine is thought to inhibit RNR, albeit
through an unknown mechanism.144 Fludarabine primarily inhibits
DNA polymerases, with minimal RNR inhibition. Cladribine and
fludarabine feature undesirable chemotypes that make them
both susceptible to glycolytic cleavage, reducing their efficacy.
Particularly with fludarabine, hydrolytic and enzymatic cleavage
produces 2-fluoroadenine, which is subsequently converted to the
highly toxic 2-fluoro-adenosine triphosphate.145 ClF represents a
triumph of semi-rational design, incorporating the most desirable
properties of cladribine and fludarabine, with minimized
toxicity.140
Similar to natural ligands, antimetabolites also modulate RNR
activity through changes to α quaternary structure. Although RNR
oligomeric regulation remains poorly understood, significant
phenotypic differences exist between the natural and nonnatural ligand-induced states.41 The α hexameric state induced
by dATP only persists when the A site is saturated with dATP, thus
enabling interconversions between the active and inactive states
as a function of cellular dATP concentration. ClFTP also binds the
A site and hexamerizes α in vitro.22 However, unlike dATP-induced
α-hexamers, ClFTP-induced α-hexamers persist subsequent to
ClFTP dissociation.23 Persistent α hexamerization is also initiated
by ClFDP binding to the C site, suggesting that quaternary
regulation is not uniquely associated with α allosteric sites. F2CDPmediated irreversible RNR inactivation in which the suicide
substrate F2CDP interacts with the C site is also proposed to lead
to α6β6.139 The kinetic stability of the ClFD(T)P-induced hexameric
states was recently exploited to demonstrate that α exists as a
dynamic equilibrium of oligomers in cells.23 α from untreated cells
is a mixture of dimer and monomer, whereas ClF-treated cells
yield α that is mainly hexameric.
The allosteric activator ATP has also been proposed to affect
α oligomerization independent of β. However, there are differing
reports, based on distinct technical approaches, on the functional
outcome of this process (Supplementary Table S1). Kashlan and
Cooperman24 described ATP-concentration-dependent formation
of αm (m = 2,4,6), whereas Dealwis and colleagues21 showed that
3 mM ATP causes α hexamerization. Hofer and colleagues146 also
detected the presence of α2 and α6 states at 0.1 mM ATP. One way
to reconcile these data is that ATP, in a similar way to dATP,
induces weakly associated hexamers that readily collapse to
lower-order oligomers at low ATP levels. This model is consistent
with analysis of α in 0.5 mM ATP showing a mixture of lowerorder species.22 These observations imply that persistent
hexamerization is uniquely induced by antileukemic nucleotides
that inhibit RNR.
Despite their success, nucleoside analogs suffer from
complications.147 Most are administered as inactive prodrugs,
necessitating successive phosphorylation by cellular kinases to
gain activity.140 Depending upon the nucleoside, a different
steady-state equilibrium between the monophosphate, diphosphate and triphosphate is established.148 These variable drug
metabolites can lead to low steady-state concentrations of the
active variant, side effects149 and ultimately toxicity.150 Nucleos(t)
ides are also susceptible to numerous catabolic pathways,
which can generate dangerous side products.151 In addition,
resistance to certain dNTP analogs is linked to differential miRNA
expression.152
Both F2C and ClF are used in combination therapies. The
combination of F2C and carboplatin is widely used.153 The theory
behind this and related approaches is that functional RNR is
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needed to provide dNTPs to repair DNA damage caused by
drugs such as carboplatin. In line with this hypothesis, the
F2C/carboplatin combination is successful in treating carboplatinresistant tumors.154 ClF is commonly used with ara-C, a dCTP
analog that, following phosphorylation to ara-CTP, can block DNA
synthesis. This combination highlights that nucleoside analogs
have complex metabolic pathways: ara-CTP inhibits deoxycytidine
kinase (dCK), an enzyme responsible for phosphorylating ara-C to
ara-CMP.155 Steady-state cellular ara-CTP levels are relatively low,
impairing efficacy. RNR inhibition by ClFD(T)P stimulates dCK
activity,156 leading to higher ara-CTP levels.157 A similar effect is
observed in cells treated with F2C and staurosporine, which
increases dCK activity.158
RNR-β INHIBITORS
The di-iron center and Y• within β are logical targets for anticancer drugs. One structurally simple β inhibitor, HU (Figure 3c), is
a known metal chelator and radical quencher. It is used in the
treatment of CML,159 AML 160 and glioblastoma.161 Although β is a
HU target 162 and β overexpression confers HU resistance,69,163,164
this drug is promiscuous, and other metalloenzymes, such as
carbonic anhydrase and matrix metalloproteinases, are also HU
targets.165 HU attacks both the Y• and di-iron center of
mammalian RNR-β on a similar timescale.166 This contrasts with
the mechanism of HU-induced β-inactivation in bacteria, which
involves exclusively Y• quenching, leading to a met enzyme
state.167 These data underscore the dual properties of HU as a
metal-chelator and single-electron donor.
A second β-targeting drug, triapine (3AP; Figure 3d) is currently
in clinical trials for CML and various solid tumors.168 3AP
represents an important case study, because it highlights the
complexities in deconvoluting inhibition mechanisms. 3AP is the
most successful of a group of thiosemicarbazone β inhibitors and
is active against HU-resistant tumors.169 Initially, it was assumed
that 3AP inhibited β through iron chelation, either from the β
active site170 or from the labile iron pool.171 However, this theory
was questioned with observations that metal-bound 3AP,
particularly Fe(II)-(3AP) complex, under aerobic conditions, is
capable of generating ROS172 that could inhibit β.173 Recent
studies indicate that the Fe(II)-(3AP) complex is the active inhibitor
in vitro and can reduce Y• at a rate faster than iron chelation at the
β active site.119 Cultured K562 cells and HU-resistant TA3 cells
treated with 3AP showed no change in iron content within β but
underwent a rapid loss of Y•. No oxidation of β residues or
accumulation of oxidized cellular proteins could be detected,
suggesting that ROS is not important for β inhibition by 3AP.
These data collectively imply that human β is highly susceptible to
radical-targeting drugs, a finding that opens a range of prospects
for inhibitor design and optimization.
GENETIC STRATEGIES FOR RNR INHIBITION
Although the most effective and widely used approaches for RNR
inhibition center on small-molecule inhibitors, targeted knockdown of RNR subunits using small interfering RNA (siRNA) also has
been developed. RRM2 knockdown impairs tumor cell proliferation174 and sensitizes cancer cells to DNA-damaging agents as
well as the RNR inhibitors HU and 3AP.122 RRM2 knockdown also
overcomes cisplatin resistance in cultured cells.54 Similarly, siRNAs
designed to downregulate RRM1 expression in tumors have been
developed to overcome gemcitabine resistance.175 Although the
application of these siRNA-based strategies in patients is
complicated by challenges related to delivery176 and stability in
plasma,177 encouraging results have been observed. A 20-mer
phosphorothioate oligodeoxynucleotide, GTI-2040,178 that significantly reduces RRM2 mRNA levels, is presently in clinical trials for
various solid tumors.174–176 Delivery of siRNA against RRM2 by
© 2015 Macmillan Publishers Limited
nanoparticles or retroviruses has been shown to suppress cancers
of the head, neck and pancreas.179,180
SUMMARY AND FUTURE PERSPECTIVES
Since the discovery of RNR activity by Reichard et al.181 in 1961,
there have been tremendous advances in understanding the
structure, function and biological significance of this essential
family of enzymes.17,19,41 The next stage in understanding RNR
functions in disease-related contexts likely will require a concerted
interdisciplinary effort that merges biochemical and genetic
analyses and couples in vitro enzymology and structural studies
with relevant cell culture and animal models. Many mechanistic
details remain to be resolved for mammalian RNR, including the
dynamics and functional importance of oligomerization and
subcellular localization, as well as the intricacies of cofactor
assembly and metalloenzyme regulation. RNR, encompassing
both conventional reductase and possible moonlighting (nonreductase) functions, clearly has important, subunit-specific roles
in cancer biology, influencing tumor initiation, progression and
therapeutic sensitivity, while also serving as a target for anticancer drugs. The extent to which the oncogenic impact of RNR
relates to RNR-mediated mutagenesis, suppression of senescence,
ROS production or modulation of apoptosis remains a key
question for future studies. Finally, given that RNR is wellestablished as an effective therapeutic target, cell-based highthroughput phenotypic screening assays represent a promising
avenue for future drug development. Beginning with the
identification of a novel radical-based catalytic mechanism, the
study of RNR has revealed many intriguing biochemical mechanisms over the years, and we anticipate that more surprises are in
store with the continued analysis of the role of RNR in cancer.
CONFLICT OF INTEREST
The authors declare no conflict of interest.
ACKNOWLEDGEMENTS
We thank Professors Rick Cerione and Jennifer Surtees for helpful discussion and
comments on the manuscript. YA acknowledges a faculty development grant from
the ACCEL program supported by NSF (SBE-0547373), an Affinito-Stewart grant from
the President's Council of Cornell Women and a Milstein sesquicentennial junior
faculty fellowship. MJCL acknowledges an HHMI international student predoctoral
fellowship.
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