Characterizing the Endoribonuclease Activity of APE1

Wan Cheol Kim
BSc, Simon Fraser University, 2007

Thesis Submitted in Partial Fulfillment of
The Requirements for the Degree of
Master of Science
In
Mathematical, Computer, and Physical Sciences
(Chemistry)

The University of Northern British Columbia
June 2009

© Wan Cheol Kim, 2009

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Abstract
Recent evidence shows that mRNA degradation is a major control point in the regulation of
gene expression. APE1, apurinic/apyrimidinic DNA endonuclease 1, has recently been
discovered to possess an endoribonuclease activity against c-myc messenger RNA (mRNA)
in vitro. We showed that APE1 shares catalytic residues to cleave both RNA and AP-DNA.
Our results also suggested that the roles of active site residues in each reaction are not
entirely identical. A consensus RNA secondary structures and sequences that are
preferentially cleaved by APE1 were determined. Our results revealed that APE1 has a
preference for cleaving the single stranded regions or weakly base paired regions of the
RNA. Also, preferred sequences of cleavage were determined to be UA, UG, and CA. When
APE1 was knocked down in HeLa cells, an increased level of c-myc mRNA and half-life was
detected. This demonstrated that APE1 is involved in the regulation of c-myc mRNA
turnover.

TABLE OF CONTENTS
Abstract

i

Table of Contents

iii

List of Tables

vi

List of Figures

viii

Acknowledgements

xi

Candidate's Publications Relevant to this Thesis

xii

Reference List

xiii

CHAPTER 1 - Introduction
1.1

Mechanism of messenger RNA (mRNA) degradation

1

1.2

Role of trans-acting factors in mRNA stability and turnover
1.2.1 Non-coding RNA (miRNA)
1.2.2 RNA-binding proteins (RBPs)
1.2.3 Ribonucleases (RNases)

4
4
5
7

1.3

Ribonucleases (RNases) associated with cancer
1.3.1 RNases from conventional
mRNA decay pathway: CCR4b, PARN, XRN1
1.3.2 RNases from specific contexts: RNase L, IRE1, and PMR1
1.3.3 RNases from miRNA pathway: Drosha, DICER, Argonaute 2
1.3.4 RNases of the nucleus/cytoplasm: Angiogenin, G3BP, and FEN1

7
9
11
18
22

1.4

Apurinic/apyrimidinic endonuclease 1 (APE1) as an endoribonuclease
1.4.1 Possible role of APE1 in cancer
1.4.2 AbasicDNA incision activity of APE1
1.4.3 Variants of APE1 identified in human populations
1.4.4 3'-5'Exonuclease activity of APE 1

27
27
30
34
35

1.5

Research objectives

37

iii

CHAPTER 2 - Identifying the critical amino acid residues for
endoribonuclease activity of APE1 and characterizing its
biochemical properties
2.1

Methodology
2.1.1 Reagents and buffer preparation
2.1.2 PCR generation of linear templates
2.1.3 Standard phenol/chloroform extraction and ethanol precipitation
2.1.4 Generation of unlabeled RNA substrates
2.1.5 Generation of 5'-radiolabeled RNA substrates
2.1.6 Generation of 5'-radiolabeled abasic DNA substrate
2.1.7 SDS-PAGE analysis on recombinant human APE1
2.1.8 Endoribonuclease assays using 5'-radiolabeled RNA
2.1.9 Abasic DNA endonuclease assays using 5'-radiolabeled abasic DNA
2.1.10 Dialysis of APE1
2.1.11 Determination of the nature of 3'-end of the RNA cleavage product
2.1.12 Electrophoretic mobility shit assay (EMSA)

40
40
40
42
42
44
45
47
49
50
50
51
52

2.2

Results and discussion
2.2.1 Generation of unlabeled and 5'-radiolabeled RNA
2.2.2 Identification of essential residues for RNA incision activity of APE1
2.2.3 Assessing the RNA-binding abilities of APE1 structural mutants
2.2.4 RNA incision activity of the population variants of APE1
2.2.5 Assessing the RNA-binding abilities of APE1 population variants
2.2.6 Possible mechanism of RNA incision by APE1
2.2.7 The effect of divalent metal ions in APE1 RNA catalysis
2.2.8 The effect of RNase inhibitors on APE1 activity

55
55
56
65
66
72
74
77
79

CHAPTER 3 - Establishing the RNA structure and sequence
cleaved by APE1
3.1

3.2

Methodology
3.1.1 Plasmid linearization and PCR generation of linear templates
3.1.2 Utilization of enzyme probes for RNA secondary structure
determination
3.1.3 In vitro effects of APE1 RNase activity on DICER product formation

82
82
84
86

Results and discussion
88
3.2.1 Generation of unlabeled and 5'-radiolabeled RNA
88
3.2.2 APE1 cleaves RNA components of SARS-corona virus at specific sites.. .89
3.2.3 APE1 cleaves pri-miR-21 and pri-miR-lOb at specific sites
94
3.2.4 Pre-treatment with APE1 interferes with processing of pre-miR-lOb
and pre-miR-21 by DICER
98

IV

CHAPTER 4 - Assessing the endoribonuclease activity of APE1
in biological systems
4.1

4.2

Methodology
4.1.1 Assessing the steady-state c-myc mRNA level
in APE1 knocked down cells
4.1.2 Assessing c-myc mRNA half-life in APE1 knocked down cells
4.1.3 Preparation of LB-antibiotic plates for Origami cells
4.1.4 Preparation of Origami competent cells
4.1.5 Transformation and induction of protein expression by IPTG
Results and discussion
4.2.1 APE1 knock down in HeLa cells up-regulates
c-myc mRNA expression
4.2.2 Optimization of monitoring c-myc mRNA stability in HeLa cells
4.2.3 Knock down of APE1 in HeLa cells stabilizes c-myc mRNA
4.2.4 Origami cell transformation with pGEX4T3-Thioredoxin A
andRNaseA
4.2.5 Induction of pET15b-WT-APEl in Origami cells is toxic

103
103
110
110
110
Ill
112
112
116
118
119
121

CHAPTER 5 - General discussion
5.1

Role of post-transcriptional control in cancer

125

5.2

Significance of identifying and studying RNases

127

5.3

Benefits of studying RNases in health and therapeutics

128

5.4

Identification of active site residues critical for
RNA-cleaving activity of APE 1

129

5.5

Role of metal ions in APE1 RNA-cleaving activity

135

5.6

Role of N-terminus in APE1

136

5.7

RNA secondary structure and sequences preferentially cleaved by APE 1

139

5.8

Assessment of biological significance of RNA-cleaving activity of APE1

141

5.9

Concluding remarks

144

v

List of Tables
Table 1: Summary of abasic DNA incision activities of
active site mutants (or structural mutants)

32

Table 2: Summary of dissociation constants and
abasic DNA incision activities of
APE1 population variants (Hadi et al. 2000)

35

Table 3: Composition of Reagents used in this chapter

40

Table 4: List of RNA substrates used in assessing the RNA-cleaving activity
of APE1 and its mutants

41

Table 5: PCR primers and their sequences used in the generation of
linear DNA template for c-myc (1705-1792) CRD RNA
by in-vitro RNA transcription

41

Table 6: Composition of reagents for SDS-PAGE

48

Table 7: SDS-PAGE gel staining & destaining reagents

49

Table 8: The composition of reagents used in EMSA experiments

54

Table 9: Summary of fold reduction in WT APE 1 and structural mutants

59

Table 10: Summary of RNA incision activity of
human population variants of APE 1 on
c-myc-1705 -1792 CRD RNA and Oligo IB

68

Table 11: List of RNA substrates generated and their cDNA sequences

83

Table 12: List of primers and their sequences used in the generation of
cDNA templates for in-vitro RNA transcription

84

Table 13: Sequences of Scrambled-Negative
DICER substrate double stranded RNAi (dsRNAi) and
dsRNAi against APE 1 mRNA

104

Table 14: Composition of the reagents used in Western blot

107

Table 15: Primer sequences used in qRT-PCR

109

Table 16: Sample table of CT values generated from qRT-PCR

116

vi

Table 17: Summary of the results of Origami cell transformations
using pET15b based plasmids coding for APE1 mutants
and their RNA cleaving activities in vitro

123

vn

List of Figures
Figure 1: A schematic view of mRNA decay

3

Figure 2: A schematic representation on the modes of action for trans-acting factors... .6
Figure 3: Abasic DNA cleaving active site model of APE1 based on X-ray
crystallography (Mol et al. 2000)

31

Figure 4: Molecular model of the positions of the residues found to be
mutated in human population (Hadi et al. 2000)

34

Figure 5: Chemical structure of abasic site analog represented as F in the cDNA
sequence

45

Figure 6: Agarose and polyacrylamide gel analyses showing
the in vitro transcribed RNA substrate and
the generation of 5'-radiolabeled RNA substrates

56

Figure 7: SDS-PAGE analysis of 1 ugof APE 1 WT and mutants

57

Figure 8: Abasic DNA endonuclease activity of WT APE1 and APE1 structural
mutants

58

Figure 9: Endoribonuclease activity of APE 1 and its structural mutants on
c-myc-1705-1792CRDRNA

59

Figure 10: Endoribonuclease activity of APE 1 and its structural mutants on
OligoIBRNA
Figure 11: Molecular model of APE1 active site (Rothwell et al. 2000)

60
63

Figure 12: RNA-binding abilities of APE 1 and its structural mutants on
c-myc 1705-1886 CRD RNA
Figure 13: Abasic DNA endonuclease activity of population variants of APE 1
Figure 14: Endoribonuclease activity of human population variants APE1 on
c-myc-1705-1792CRDRNA
Figure 15: Endoribonuclease activity of human population variants of APE 1
on OligoIBRNA
Figure 16: Mapping of the cleavage sites generated
by L104R and E126D variants of APE1

66
67

69
70
71

Vlll

Figure 17: RNA-binding abilities of APE 1 and its population variants
on c-myc 1705-1886 CRD RNA

72

Figure 18: RNasin-resistant activity of L104R and E126D
on c-myc 1705-1886 CRD RNA

73

Figure 19: APE1 generates RNA products with 3'phosphates

75

Figure 20: RNA incision activity of APE1 requires 2'-OH on the sugar moiety

76

Figure 21: Effect of divalent metal ions on RNA-cleaving activity of APE1

78

Figure 22: Differing concentrations of magnesium in the reaction mixture
affects the specificity and the incision activity of APE1
Figure 23: RNA incision activity of APE 1 is inhibited by
DEPC and RNase Inhibitor (RNasin)

79
80

Figure 24: Agarose gel analysis showing the digestion of plasmids used for
in vitro transcription and the generation of unlabeled RNA substrates

89

Figure 25: Ribonuclease secondary structure probing of Spike RNA

90

Figure 26: Ribonuclease secondary structure probing Orf lb RNA

92

Figure 27: Ribonuclease secondary structure probing of Orf3 RNA

93

Figure 28: Ribonuclease secondary structure probing pre-miR- 10b RNA

96

Figure 29: Ribonuclease secondary structure probing pre-miR-21 RNA

97

Figure 30: Effect of pre-treatment with APE1 on DICER's ability to process
5'GG-pre-hsa-miR-10b
Figure 31: Effect of pre-treatment with APE1 on DICER's ability to process
5'GG-pre-hsa-miR-21

99
100

Figure 32: The genetic selection system for ribonuclease activity
in Origami cells (Smith and Raines 2006)

102

Figure 33: Western blot illustrating the reduced levels of APE1
after two time points using dsiRNAi against APE 1 in HeLa cells

113

IX

Figure 34: Steady state level of c-myc mRNA is elevated
after transient knock down of APE1
using double stranded siRNA in HeLa cells

114

Figure 35: Level of c-myc mRNA monitored over time after treating cells
with transcriptional inhibitor Actinomycin D, a-amanitin, and DRB

117

Figure 36: Decay of c-myc mRNA over time after treating cells with DRB

118

Figure 37: c-myc mRNA is stabilized after transient knock down of
APE1 in HeLa cells
Figure 38: Transformation of Origami cells with increasing amounts of
pGEX4T3-Thioredoxin A

119
120

Figure 39: Transformation of Origami cells with pGEX4T3 plasmids
carrying cDNAs for Thioredoxin A and Ribonuclease A

120

Figure 40: Transformation of Origami cells with pET15b plasmids
carrying cDNAs for WT-APE 1 and recombinant enzymes

121

Figure 41: IPTG induces expression of APE1 WT, D283A, and H309N
which exerts toxic/non-toxic effects in Origami cells

122

Figure 42: Transformation of Origami cells with pGEX4T3 plasmids
carrying cDNAs for Thioredoxin A, WT-APE1, and E96A-APE1

124

Figure 43: Altered post-transcriptional control in cancer (Kim and Lee 2009)

126

x

Acknowledgements
I would like to thank my MSc. Theis supervisor, Dr. Chow H. Lee for his absolute
support and guidance in completing this thesis. Also, I would like to thank the members of
my MSc. supervisory committee, Drs. Andrea Gorrell and Dezene Huber, for their helpful
suggestions and insights in carrying out this research. I would like to extend my thanks to the
members of the Lee lab, Sang-Eun (Eunice) Kim, Chris Uy, Navkarndeep Bal, Dustin King,
Dana Thomsen, Conan Ma, Mavis Ye, and Maggie Li for their friendship, encouragement,
and technical support. Finally, I would like to say special thanks to my parents and friends in
Prince George who have supported me during my time here at UNBC.

XI

Candidate's Publications Relevant to this Thesis
Articles
Kim, W-C, Li, W-M., Wilson Jr III, D. M., Lee, C. H. (2009) Endoribonuclease activity of
apurinic/apyrimidinic endonuclease 1 (APE1) variants identified in human population.
(Manuscript in preparation)
Kim, W-C, Kim, S-E., Li, W-M., Wilson Jr III, D. M., Lee, C. H. (2009) Endoribonuclease
active site of human apurinic/apyrimidinic endonuclease 1 (APE1). To be submitted to J.
Biol. Chem. (Manuscript in preparation)
Kim, W-C, King, D., Lee, C. H. (2009) RNA-Cleaving Properties of Human
Apurinic/Apyrimidinic Endonuclease 1 (APE1). Submitted to RNA (ID: RNA/2009/017327)
Kim, W-C, and Lee, C. H. (2009) The role of mammalian ribonucleases in cancer. Biochim.
Biophys. Acta. - Reviews on Cancer doi:10.1016/j.bbcan.2009.05.002 In press
Barnes, T., Kim, W-C, Mantha, A., Kim, S-E., Izumi, T., Sankar, M., Lee, C. H. (2009)
Identification of apurinic/apyrimidinic endonuclease (APE1) as the endoribonuclease that
cleaves c-myc mRNA. Nucleic Acids Res. doi: 10.1093/nar/gkp275 In press

Abstracts
Kim, W-C, and Lee, C. H. Understanding endoribonuclease activity of APE1. (2009). Cell
& Molecular Biology Interest Group Seminar, Prince George, BC, Jan. 23,
Speaker
Kim, W-C, and Lee, C. H. Structure and ribonuclease function of apurinic/apyrimidinic
DNA endonuclease 1 (APE1). (2008). RiboClub 2008, Mont Orford, QC,
Sep.22, Poster
Kim, W-C, and Lee, C. H. Biochemical characterization of apurinic/apyrimidinic DNA
endonuclease 1 (APE1) in vitro. (2008). RiboWest 2008, Lethbridge, AB, Jun.
15, Speaker

Patents
U.S. Patent Application No. 83152-9, Title: APE1 AND USES THEREOF, Inventor: LEE,
Chow ; BARNES, Tavish; and KIM, Wan-Cheol: Filed November 29, 2008.

xn

CHAPTER 1
Introduction
The processes conferring mRNA stability, translational inhibition, and transcript
degradation are all intricately related to one another and can be influenced by cz's-acting
elements such as the poly (A) tail and AU-rich elements (AREs) and trans-acting factors
such as non-coding RNAs (ncRNAs), RNA-binding proteins (RBPs), and ribonucleases
(RNases). Alterations of these factors are reported to be implicated in cellular abnormalities
and such link has highlighted the significance of studying these factors. Since RNases play a
significant role in mRNA metabolism and gene expression, further studies are required to
identify

and

characterize

unknown

RNases

in

mammalian

cells.

Recently,

apurinic/apyrimidinic endonuclease 1 (APE1) was identified to possess an endoribonuclease
(endoRNase) activity against c-myc mRNA and this have led to the formulation of the three
major research questions.

1.1

Mechanism of messenger RNA (mRNA) degradation
In eukaryotes, literature emphasizes that mRNA stability and turn-over is an integral

control point in the regulation of gene expression (Parker and Song 2004; Garneau et al.
2007). The stability of various mRNAs within a cell can differ and this results in a magnitude
of difference in mRNA abundance (Ross 1995). Moreover, the stability of individual mRNAs
can be controlled in response to a variety of stimuli, allowing for rapid changes in gene
expression to take place, and this has been viewed to be a critical component in development
and pathogenesis (Wilusz and Wilusz 2004; Audic and Hartley 2004; Steinman 2007).
Therefore, previous studies have focused on discovering major mRNA decay pathways and

1

their players: cis-regulatory elements, protein factors, and enzymes. However, in the last
several years, significant efforts have been put to identifying and studying the enzyme
involved in mRNA degradation.
From studies in yeast and mammals, major pathways of exoribonucleolytic mRNA
decay have been identified. mRNA deadenylases such as Poly A Ribonuclease (PARN) and
CCR4b-NOT can initiate mRNA decay by removing the poly (A) tail from the 3' end of the
mRNA (Figure 1). Also, mRNA degradation can begin by the removal of the 7'methylguanosine cap (m7G) at the 5' end of the mRNA mediated by a complex of decapping
enzymes, Dcplp and Dcp2p (Figure 1). Following these processes, vulnerable transcripts are
subjected to further cleavage and clearance by a 5'-3' exoribonuclease (ExoRNase), Xrnlp,
and a complex of 3'-5' exosome and other 3'-5' exoribonucleases (Garneau et al. 2007;
Schaeffer et al. 2009). In addition, endoribonucleolytic cleavage can also initiate mRNA
degradation pathways. Endoribonucleases (EndoRNases) can cleave in the middle of the
mRNA sequence. This way, the decay and the clearance of the transcript can by-pass the
requirements of deadenylation and decapping step, allowing for a higher efficiency.
Apart from regular mRNA decay, cellular mechanisms can destroy faulty transcripts
and mRNAs undergoing aberrant translation. This mechanism is called an mRNA
surveillance control, which includes three distinct categories of mRNA decay: non-stop
decay, non-sense mediated decay, and no-go decay. The non-stop decay mechanism
recognizes and degrades transcripts without stop codons.

2

m'<

m G

'

m A A AV

?

T>
AAAAAA

PARN
CCR4b-N0T

v

XRN1
AAAAAA
Examples:
PMR1
RNaseL

m'G

1
m?G I
Exosome

AAAAAA

Figure 1: A schematic view of mRNA decay. mRNA decay can be initiated by any of the
three processes shown on the left side of the arrow. Deadenylases such as Poly A
Ribonuclease (PARN) and CCR4b-NOT remove the Poly A tail of the target mRNA, while
decapping enzymes remove the 5'-m7G mRNA cap. EndoRNases such as PMR1 and RNase
L can cleave mRNA in the middle of the sequence. The combinations of these three
processes make mRNA available for ExoRNases to further degrade and clear the transcript in
5'-3' (XRN1) or in 3'-5' direction (Exosome complex) as shown on the right side of the
arrow.
This is mediated by different factors converging on a stalled ribosome at the 3' end of
the mRNA and allows for a recruitment of exosome (Garneau et al. 2007). During non-sense
mediated decay, stalling of the ribosome near the premature termination codon (PTC) allows
for a stable binding of associated factors (UPF1 and SMG1) to the mRNA near the PTC.
These factors interact and further recruit SMG5, SMG6, and SMG7 to promote mRNA decay
via ExoRNases. In addition, EndoRNase activity of SMG6 can directly degrade the aberrant
transcript (Eberle et al. 2009). Finally, in no go mRNA decay, stalling of ribosomes caused
by a stem-loop RNA structure leads to the destruction of the transcript facilitated by Dom34
and Hbsl proteins. Here, in vitro studies have identified dom34 as the responsible
endoRNase (Lee et al. 2007). It was found that dom34 is highly conserved from fruitflies to
mammals, and it plays an important role in the stem cell self-renewal in fruitflies and cell
divisions in mice (Tollervey 2006).

3

1.2

Role of trans-acting factors in mRNA stability and turnover
Having described the pathways of mRNA degradation, it is now important to

understand the roles of trans-acting factors that are notably involved in this process. Hence,
the intricate balance between the actions of mRNA stabilizers which enhance the half-life of
mRNAs and those of destabilizers which exert opposite effects will be further explained.
1.2.1 Non-coding RNA (miRNA)
Micro RNAs (miRNAs) are a class of non coding RNAs (ncRNAs) that have been
revealed to exert an enormous effect on gene expression. This type of RNA (20-25
nucleotides in length) is generated from an endonucleolytic processing of longer transcripts
as in the example of HI9 RNA, a ncRNA, which can act as a miRNA precursor acting on
mRNA degradation and repression (Cai and Cullen 2007). In general, long precursor RNAs,
such as HI9, can form multiple stem-loop structures and an EndoRNase, Drosha, part of
RNase III group, can cleave such RNAs in the nucleus, releasing the pre-miRNA product
into the cytoplasm. Once this RNA has reached cytosol, it is subsequently processed again by
a yet another RNase III type endoRNase called DICER (GroBhans and Filipowicz 2008).
Such processed miRNAs are now deemed mature, and these mature miRNAs can bind to
their target mRNAs by base pairing and recruit proteins of the Argonaute (Ago) family to
form a miRNA ribonucleoprotein (miRNP) complex (Figure 2A). The miRNP complex can
influence the stabilities of their target mRNAs mainly through inhibiting translation while
promoting degradation (Zhang et al. 2007; To et al. 2008). The mRNA decay here can be
mediated by a deadenylation-dependent or -independent exonucleolytic pathway or via other
unidentified decay factors. Reviews on the interplay between miRNA and ExoRNase decay
have been published (Jackson and Standart 2007; Garneau et al. 2007). In some cases, near

4

perfect base-pairing between the miRNA and the target mRNA allows Ago2, one of the four
Ago family members, to endonucleolytically cleave the mRNA and send the fragments for
further degradation as reviewed (Valencia-Sanchez et al. 2006).
1.2.2 RNA-binding proteins (RBPs)
RNA-binding proteins (RBPs) can also dramatically influence the longevity of
mRNAs (Paschoud et al. 2006; Pulcrano et al. 2007). This mechanism not only allows the
half-life of mRNA to vary, but is also capable of regulating the decay rates of individual
mRNA in response to different stimuli, as is the case for transcripts encoding cytokines or
proto-oncogenes bearing AU-rich elements in their 3' untranslated region (UTR) (Chen and
Shyu 1995; Barreau et al. 2006). RBPs can bind to the 3' UTRs as well as to the coding
region of mRNAs and shield them from mRNA degradation machineries (Figure 2B). One
study has reported in breast cancer cell lines that HuR, an RBP, critically mediates the overexpression of oncogenic biomarker by enhancing its mRNA half-life (Guo and Hartley
2006). Similarly, CRD-BP, an RBP that shields mRNA from degradation, has been found to
be elevated not only in the primary samples of melanoma (Elcheva et al. 2008), but also in a
variety of tumor samples including colon (Dimitriadis et al. 2007), colorectal (Ross et al.
2001), mesenchymal (Ioannidis et al. 2001), breast (Ioannidis et al. 2003), Ewing's sarcoma,
brain and non-small cell lung (NSCL) (Ioannidis et al. 2004). Further, it was reported that <30% of human breast cancer cases exhibited amplification of the CRD-BP gene (Doyle et al.
2000), and its ectopic expression in transgenic mice induced the development of mammary
tumors (Tessier et al. 2004). Other RBPs such as KSRP can stimulate the degradation of
several

cellular

mRNAs

that contain

AU-rich

elements

(AREs) by

recruiting

endoribonucleases and deadenylation factors to regulate their expression (Gherzi et al. 2004;

5

Chou et al. 2006; Mazan-Mamczarz et al. 2007). The interactions between mRNAs and
RBPs and their subsequent effects on cells seem complex and vary between studied cell
lines. Recent report shows overexpression of HuR is directly responsible for the increase of
kinase mRNA in fibroblasts (Sobue et al. 2008) while another study reports HuR can
stabilize tumor suppressor mRNA in prostate cancer cell lines and in anti-cancer therapeutics
(Quann et al. 2007; Bandyopadhyay et al. 2008).

Stabilizing
RBP

Figure 2: A schematic representation on the modes of action for trans-acting factors.
(A) Non-coding RNAs can regulate mRNA translation and degradation. Top: miRNAs, a
subclass of non-coding RNA, can inhibit the translational process while recruiting the RNA
decaying machineries such as deadenylase and decapping enzymes. Bottom: Argonaute 2 can
be induced to cleave mRNAs with externally introduced short interfering RNAs (siRNAs).
However, a near to perfect complementary hybridization of miRNAs to mRNAs can also
induce Argonaute 2, complexed with other gene silencing machineries, to cleave the target
mRNA. (B) RNA binding protein (RBP) controls the stability of mRNAs. Top: Destabilizing
RBP can recruit deadenylase machineries or has implications to recruit endoribonucleases to
accelerate the decay of target mRNA. Bottom: Stabilizing RBP can attenuate mRNA decay
by binding either to the coding regions or the 3' untranslated regions of the target mRNA.
Nonetheless, these demonstrate how important these molecules can come into play in
controlling gene expression and tip the balance between decay and survival of diverse range

6

of mRNAs. Articles that provide a complete review on the role of RBPs and how they can
influence gene expression are available (Audic and Hartley 2004; Garneau et al. 2007).
1.2.3 Ribonucleases (RNases)
An efficient interplay between various ribonucleases (RNases) has a major impact on
mammalian mRNA turnover (Wilusz and Wilusz 2004). In mammalian cells, mRNAs are
degraded upon shortening of their poly A tail (deadenylation) or removal of their 5'-m7G cap
(decapping). Exoribonucleases can clear these 'exposed' mRNAs either from its 5' or 3' end
and several of these enzymes are actually involved in the deadenylation step. On the other
hand, endoribonucleases can efficiently by-pass these requirements (decapping and
deadenylation) and internally cleave mRNAs to generate products that are eventually cleared
by exoribonucleases. In the absence of regulatory mechanism, these RNases can have
profound impact on gene expression since a single cleavage on a transcript can result in its
inactivation. Cells have employed mechanisms to prevent potential aberrant degradation of
mRNAs by these endoribonucleases. Some of these (RNase L, IRE1, and PMR1) are tightly
controlled through cellular signaling cascades (Garneau et al. 2007). The role of
ribonucleases (RNases) will be discussed more in depth in the following section.

1.3

Ribonucleases (RNases) associated with cancer
It is noteworthy that some of these aforementioned RNases have been found to

influence or shown potential to affect the turnovers of oncogenic and tumor-suppressive
mRNAs in relation to cancer (Kriiger et al. 2005; Oikawa et a/.2007). Deregulation of
RNases involved in RNAi machinery and miRNA processing also have significant impact on
cell proliferation and tumorigenesis. Some RNases can be turned on by cellular responses

7

that are often observed in cancers such as in misfolded protein response and aberrant tyrosine
kinase signaling.
Suppression of RNases can contribute to the reduction in mRNA decay. mRNA
coding for P-glycoprotein (Pgp) was found to have a prolonged half-life in subcutaneously
grown liver tumors compared to normal tissues (Lee et al. 1998). Pgp is a transport protein
found in plasma membranes that is well associated with multi-drug resistance in many cancer
cell lines and its expression increases during carcinogenesis in rats. In normal liver cells, Pgp
mRNA is thought to be degraded by endonucleases and/or exonucleases because its mRNA
decay intermediates were detected in normal liver cells (Lee et al. 2005). But, significant
decrease in RNase activity in hepatomas (Roth 1963; Sidransky et al. 1978) is considered to
be the reason for the prolonged Pgp mRNA half-life (Lee et al. 1998) and little or absence of
Pgp mRNA decay intermediates in liver tumors (Lee et al. 2005). On the contrary, there are
examples where elevation of RNases can promote tumorigenesis. Angiogenin, an RNase A
family enzyme, can process 28S and 18S rRNAs (Adams and Subramanian 1999) and its
RNase activity is essential for angiogenesis (Crabtree et al. 2007). This activity has been
investigated as the source for inducing angiogenesis and small inhibitors have been tested to
inhibit its RNase activity as an anti-tumor therapeutic (Kao et al. 2002). Studies have found
Angiogenin to be highly over-expressed in numerous tumor samples and its expression
correlated with cancer progression and poor prognosis. Its role in cancer progression is
supported through induction of cell proliferation in melanoma cell lines while having no
effects on normal cells (Crabtree et al. 2007). Finally, in pancreatic cancer cell lines,
increased levels of RNase 1 in patient serums have been reported and the detection of its
elevation is being explored as a distinguishable tumor marker (Fernandez-Salas et al. 2000).

8

As well, different glycosylation states for this RNase have been reported between normal and
tumor tissues though it has not been tested whether different states of glycosylation results in
difference in RNase activity and mRNA levels in cells (Peracaula et al. 2003).

1.3.1 RNases from conventional mRNA decay pathway: CCR4b, PARN,
XRN1
The conventional pathway for mRNA decay plays a critical role in gene expression.
Apart from the actions of endoribonucleases, much of the mRNA decay in mammalian cells
begins with deadenylation. There are diverse sets of enzymes that carry out this important
process (Zheng et al. 2008; Goldstrohm and Wickens 2008) and it is reported that these
enzymes such as, PAN2-PAN3, CCR4-CAF1, and Poly A Ribonuclease (PARN) initiate
mRNA degradation and act on a critical step of determining mRNA turnover rates and gene
expression levels (Parker and Song 2004; Schwede et al. 2008). In mammalian cells, these
complexes carry out exonucleolytic degradation of poly (A) tails on various mRNAs and
making way for 3'-5' exosome complexes to clear off the transcript (Fig. 1C). They can act
ubiquitously on any mRNAs that have poly (A) tails and do not seem to have peculiar
specificity. Genetic and RNAi experiments have demonstrated their broad roles in biological
functions such as fertility and cell growth (Goldstrohm and Wickens 2008). Subsequently,
relatively few reports suggest their implications in cancer.
i)CCR4b
CCR4b, part of CCR4-CAF1 complex, induces growth in fibroblast cell lines by its
activity and is speculated to be a proto-oncogene (Morita et al. 2007). In normal conditions,
this enzyme was reported to regulate the level of p27Kipl mRNA, a tumor suppressor that
inhibits cyclin dependent kinases, and promoted cell growth. Moreover, when the cells were

9

depleted of it, p27Kipl levels were increased and there was growth impairment. CCR4b can
also be suppressed in vitro by a recognized anti-proliferative protein, Tob, supporting
previous findings that its activity promotes cell growth (Miyasaka et al. 2008). Thus far,
there are no reports showing the changes of CCR4b levels from various types of tumors.
Similarly, no association study has been done in primary tumors or its derived cell lines to
assess the levels of CCR4b and its effect on tumor suppressive mRNA turnover.
ii)PARN
RNases have been shown to control the level of mRNAs by interacting with RBPs.
PARN is an example. Its deadenylase activity coupled with destabilizing actions of RBPs can
potentially act as a tumor suppressor, decaying mRNAs coding for growth factors such as IL8 and VEGF. This was seen from a report investigating the role of defective RBP called TTP
that could not recruit PARN in malignant glioma cells (Lai et al. 2003; Suswam et al. 2008).
Also, PARN and exosome complex are shown to be recruited by KSRP and/or DHAU and
account for the destabilization of various mRNAs including c-jun and uPA, of which their
elevation are implicated in cancers (Gherzi et al. 2004; Tran et al. 2004). Another study
looked at the interaction between PARN and CUG-BP in which the RBP recruits the enzyme
to destabilize c-fos and TNF-alpha mRNAs (Moraes et al. 2006).
iii) XRN1
Besides deadenylation, 5'-3' exonucleases can also initiate mRNA decay once their
target transcript is decapped. XRN1 is an example enzyme that is implicated in cancer as a
dysregulated TSG. In primary samples of osteogenic sarcoma (OGS) and its derived cell
lines, there was a reduction and/or depletion of XRN1 mRNA (Zhang et al. 2002). Also, in
one of these cell lines, there was a homozygous loss of function mutation for XRN1

10

suggesting its role as a TSG. From these, we can conclude that such RNases in conventional
mRNA decay pathway have the potential to influence the expression of genes that are
implicated in cancers.

1.3.2 RNases from specific contexts: RNase L, IRE1, and PMR1
This section discusses RNases requiring specific signals to become activated. RNase
L and IRE1 are activated under signals from crisis situations and PMR1 is activated by a
proto-oncogene signal (Garneau et al. 2007; Peng and Schoenberg 2007). RNase L functions
in antiviral defense, but also in tumor suppression as its mutations are revealed to be
implicated in different types of cancer (Li et al. 2007; Bisbal and Silverman 2007). Growing
interests are drawn toward IREl that shows two opposite effects implicated in tumorigenesis
and apoptosis, apart from its role in dealing with ER stress. Finally, evidence indicates an
emerging link between PMR1 and cancer which remains dormant in normal conditions, but
becomes activated to participate in mRNA decay.
i) RNase L
Human RNase L is an endoribonuclease expressed in a number of mammalian cell
types. It has been shown in cell and cell-free systems to cleave both viral mRNA, rRNA, and
several cellular mRNAs in the single stranded regions of UU and UA dinucleotides (Li et al.
2007; Daugherty et al. 2007). RNase L has been shown to control the stability of
mitochondrial DNA-transcribed mRNAs in human and mice cells (Chandrasekaran et al.
2004; Le Roy et al. 2001) and destabilize mRNAs of genes induced by interferon response to
viral infections (Bisbal et al. 2000). It was also reported to control the stability of MyoD
mRNA and the differentiation of muscle cells (Li et al. 2000). Their main role, however, has
been reported to be in innate immunity where its RNase activity can amplify an antiviral

11

response (Malathi et al. 2007; Bisbal and Silverman 2007; Liang et al. 2006). Recently, it has
been shown to regulate interferon-induced apoptosis by controlling the stability of
mitochondrial mRNAs (Le Roy et al. 2007). The activity and expression of RNase L are
under tight regulation (Li et al. 2007; Bisbal and Silverman 2007). In normal conditions, it
resides as a monomeric latent form and its activity is repressed by its inhibitor, RLI (Li et al.
2007). However, upon binding to 2',5'-linked oligoadenylates (2-5A) produced by 2-5A
synthetase enzymes, it can dimerize and act as an activated RNase inside the cytoplasm.
Analysis of its promoter sequence has found several sites for its tissue-specific and general
expression accounting for its wide spread presence among mammalian cells (Zhou et al.
2005). Its expression is reported to be regulated at the mRNA level by ubiquitous RBPs,
including HuR (Li et al. 2007).
Initially, RNase L was mapped to a region linked to susceptibility in hereditary
prostate cancer (HPC) (Bisbal and Silverman 2007) and this is also consistent with the report
that the human variants of RNase L are associated with aggressive metastasis (NoonanWheeler et al. 2006). There are significant differences in RNase activities of these variants
and their effects have been observed in human cancers. A truncating variant, E265X, lacks
2-5A binding domain and this renders it completely inactive (Sun et al. 2007). Subsequently,
higher frequency of heterozygosity in E265X mutation is found in hereditary prostate cancer
(HPC) than in the controls (Rokman et al. 2002). R462Q shows a 3-fold reduction in its
activity (Sun et al. 2007) and two studies have suggested this to be associated with higher
risk of prostate cancer and its early onset (Rennert et al. 2005; Shook et al. 2007). There was
also a correlation between risk of advanced prostate cancer and increased uptake of transfatty acid in carriers (R/Q) and in homozygous groups (Q/Q), showing that this susceptibility

12

gene can also have synergetic effect (Liu et al. 2007). D541E exhibits no loss in its RNase
activity but a homozygous population (E/E) was found to have a significantly higher risk for
developing prostate cancer than the D/D or D/E groups (Shook et al. 2007). The exact cause
of this association still remains to be found and further reviews on population variants of
RNase L to prostate cancer risk and progression are available (Bisbal and Silverman 2007;
Liang et al. 2006; Sun et al. 2007). In addition, these variants have linkages to other types of
cancers. Notably, the R462Q variant has been associated with colorectal (Kriiger et al. 2005),
and pancreatic cancers (Bartsch et al. 2005). Especially in the latter study, R462Q was found
to be associated with increased aggressiveness and metastases of pancreatic cancer, and its
homozygosity (Q/Q) was found to be more common in groups with cancers and possessed a
higher risk.
The tumor suppressive function of RNase L has been well characterized especially
under interferon signaling. RNase L is suggested to degrade mitochondrial mRNA which can
lead to cytochrome-c release and caspase-3 activation during apoptosis induced by
interferon-a or an anti-cancer agent (Le Roy et al. 2007; Naito et al. 2006). RNase L can also
act by up-regulating tumor suppressor genes. Here, it's reported that RNase activity is
required, but may not be sufficient to up-regulate the level of MIC-1/NAG-1. How RNase L
activity raises the level of specific RNAs is still unknown, but there are speculations that it
can destroy inhibitory miRNAs or rRNAs which results in the activation of transcriptional
machineries (Malathi et al. 2005). RNase L was also found to act as a down stream effector
of a tumor suppressor, BRCA1 activated upon interferon-y signaling, to facilitate apoptosis in
cancer cells (Mullan et al. 2005). Similarly, in prostate cells, one pathway that can
antagonize interferon signaling is the dihydrotestosterone (DHT) pathway which promotes

13

cell survival and proliferation. It was shown in cancer cell lines that mutated RNase L can
incur attenuation of interferon signaling and allow DHT pathway to take over (Bettoun et al.
2005). Reduction of RNase L activity may lead to diminished cleavage of DHT induced
mRNAs and allow this pathway to promote cancer development. Accordingly, additive effect
in RNase L activity results in inhibition of tumorigenesis (Kriiger et al. 2005), confirming its
role in suppressing cancer development. Moreover, RNase L shows tumor suppressor
function in a mouse model of fibrosarcoma (Liu et al. 2007). Over-expression of human
RNase L in mouse fibrosarcoma cells significantly delays tumor formation. As well, tumors
over-expressing RNase L have more polygonal cells and shows increased tumor necrosis.
Interestingly, these tumors start to grow once the ectopic expression of RNase L stops. Thus,
it is concluded that RNase L can act as tumor suppressor of which its proper expression and
activity are critical in regulating cell growth and survival.
ii) IRE1
Inositol-requiring enzyme 1 (IRE1) is a transmembrane endoribonuclease found in
endoplasmic reticulum (ER). It has a functional endoribonuclease/kinase domain on the
cytoplasmic side and stress sensor domain on the luminal side. Upon exposure to ER stress,
its dimerization and autophosphorylation results in the activation of its RNase activity (Lee et
al. 2008). To date, identified substrates for IRE1 are mRNAs encoding X-box binding
protein 1 (XBP1), CD59, as well as its own mRNA (Tirasophon et al. 2000; Koong et al.
2006; Oikawa et al. 2007). IRE1 cleavage of XBP1 results in the splicing of this transcript
which encodes a potent transcription factor, XBPls, that activates genes necessary for
unfolded protein response (UPR). UPR is a homeostatic signaling pathway which adjusts ER
protein folding capacity according to the cellular need and, in a variety of contexts, it has

14

been shown to either promote cell survival or apoptosis (Bernales et al. 2006). How this
decision is made is still unknown. Other than IREl-XBPls signaling, two other UPR
activation signals have been found. It is speculated that the overall transcriptional output
from combinations of these three signals ultimately determines the fate of the cell.
Nonetheless, IRE1 acts in upstream to activate the UPR, and is an important RNase that
directs cancer progression as well as cell death.
Its major implication in tumorigenesis comes from its production of XBPls, a potent
transcription factor that activates genes that are necessary for UPR initiation. For example,
oncogene, LMP1, from Epstein Barr Virus, can induce IREl-XBPls pathway to activate
UPR and its own synthesis (Lee and Sugden 2008). As well, its RNase activity has been
suggested to influence the turn over of a significant number of mRNA in mammalian cells,
including whose translation products regulate angiogenic processes (Drogat et al. 2007). For
example, decreased expression of CD59 has been implicated in lung caner and an ectopic
over-expression of IRE1 indeed down regulates its mRNA target (Oikawa et al. 2007).
Indeed, this is consistent with the finding that the maintenance of IRE1 activity can promote
cell survival (Lin et al. 2007). However, mounting evidence has linked XBPls in tumor
growth and progression under hypoxia, which is present in all solid tumors that incur ER
stress. IREl-XBPls signaling has been found to promote cancer survival and progression.
Over-expression of XBPls in ER-positive breast cancer cells leads to estrogen-independent
growth and reduced sensitivity to growth inhibition by the antiestrogens independent of
functional p53 (Gomez et al. 2007). Moreover, signaling through IREl-XBPls is elevated in
breast tumors with enhanced survival (Davies et al. 2008) and enhances the progression of
liver cancer (Shuda et al. 2003).

15

But, UPR activation via IRE1 activation is also shown to mediate tumor suppression
in multiple observations. For example, tumor suppressive mediators BAX and BAK
modulate the UPR by directly interacting with IRE 1 alpha in cells and mice (Hetz et al. 2006).
More importantly, IRE 1-UPR pathway has been found to be used by various agents to induce
cancer cell death and inhibition (Davenport et al. 2007; Guichard et al. 2006; Little et al.
2007). These evidence support IRE1 tumor suppressive activity which is also consistent with
a report that showed its suppression resulting in synovial cell overgrowth (Gao et al. 2008).
Therefore, definitive role of IRE 1-UPR signaling in tumor development still need further
clarifications. Particularly, future works should be focused on discovering a mechanism or a
cellular environment that directs IRE1 activity to tumor progression or inhibition. It was
reviewed that UPR signals are relayed and integrated to re-establish homeostasis in the cell's
protein folding capacity or—if this cannot be achieved—commit cells to apoptosis (Bernales
et al. 2006). Recent study suggests the balance between spliced and un-spliced form of XBP1
plays an important role in restoration of homeostasis, and this underscores the importance of
understanding the regulation of IRE1 activity in deciding the fate of cancer cells (Davies et
al. 2008). Additionally, investigating the regulation of IRE1 expression in cancers and ER
stress can assist in understanding the role of IRE1 in deciding the cell's fate.
iii) Polvsomal ribonuclease 1 (PMR1)
PMR1 is an endoribonuclease originally isolated from Xenopus liver polysomes but
its orthologue was later found to be also present in mammalian cells (Yang et al. 2004).
Uniformly found across the cytoplasm on polysomes, PMR1 has a unique sequence and
structure that differentiate itself from other groups of RNases. Also, PMR1 is activated only
upon being targeted to polysomes, and can initiate mRNA decay during its translation. This

16

critical step in regulation is mediated by c-Src phosphorylation of PMR1 at the tyrosine
residue at C terminus (Peng and Schoenberg 2007). It was also found that this c-Src
phosphorylation can be activated through EGFR signals which are well associated with
tumorigenesis. Like the authors speculated, more external stimuli may be involved in PMR1
activation. Recent report revealed another layer of regulation of PMR1 activity. It was found
that Hsp90 can bind to and stabilize PMR1 from proteasome degradation, sustaining its
mRNA decay capabilities and accumulation inside the cell (Peng et al. 2007).
These findings indicate the potential role of PMR1 in cancer development and
progression. c-Src is a well-known proto-oncogene in humans that is over-expressed or
overtly activated in many cancers (Irby and Yeatman 2000). Its kinase activity can activate
diverse number of transcription factors as well as signaling cascade that lead to cell growth
and proliferation. The mechanistic model from Peng et al. suggests that c-Src may in part use
PMR1 mRNA decay path as a route to destroy transcripts for tumor suppressors. In this,
Hsp90 can prolong the longevity of PMR1 to sustain the process of mRNA decay. Since
Hsp90 serves as an essential mediator for cellular proliferation (Calderwood et al. 2006), the
mechanism mentioned above can be one of its ways to facilitate the destruction of mRNAs
coding for growth regulators. Further studies to identify mRNA substrates of PMR1 should
provide support to this model and the role of PMR1 in cancer. Further, assessing the
relationship between PMR1 and EFGR over-expression and its elevated activity will clarify
the role of PMR1 as a downstream effector in carrying out an oncogenic pathway. In addition,
assessing its expression and activity in tumor samples or cell lines will give further support
for its role in tumorigenesis.

17

1.3.3 RNases from miRNA pathway: Drosha, DICER, Argonaute 2
Growing evidence suggest dysregulation in microRNA (miRNA) expression pattern
is associated with cancers (Hagan and Croce 2007). Subsequently, three major RNases
involved in miRNA maturation and implementation can have critical effects in controlling
miRNA expression and gene expression. This section discusses how alteration of these
RNases can potentially lead to tumorigenesis.
i) Drosha/DGCR8
Maturation of miRNA begins with pri-miRNA processing. This step is carried out by
the endoribonucleolytic function of Drosha complexed with DiGeorge syndrome Critical
Region gene 8 (DGCR8). Drosha is part of the RNase III family responsible for giving the
cleavage activity against double stranded RNAs and evidence suggests its implication as a
proto-oncogene. In one study, elevated expression of Drosha and its mRNAs were found in
esophageal cancers (Sugito et al. 2006). Moreover, its suppression in cancer cell lines
resulted in inhibition of their proliferation. Another study showed a correlation between the
elevation of Drosha mRNAs and gains in genomic copy-numbers in addition to reporting its
increased copy numbers in clinical samples and cell lines of cervical squamous cell
carcinoma (Muralidhar et al. 2007). Recently, in cervical cancer samples, Drosha was one of
the genes that were over-expressed, further supporting its role as an oncoprotein (Scotto et al.
2008).
A few studies outline how Drosha might contribute to tumorigenesis. Its enhanced
activity against pri-miRNAs can result from different signaling molecules, but has been
found to influence the levels of miRNAs implicated in cancers. One study found increased
accessibility of Drosha to pri-miRNA-191 resulted in miR-191 over-expression in leukemic

18

cell lines (Nakamura et al. 2007). Similarly, growth factors such as TGF-P and those from
the family of bone morphogenetic proteins (BMPs) have been found to induce their
downstream transducers which interact with Drosha and enhance its accessibility and/or
activity towards pri-miRNA-21. This resulted in the rapid elevation of miR-21 in vascular
smooth muscle cells (VSMC) and down-regulation of a tumor suppressor PDCD4. (Davis et
al. 2008). The finding is in line with numerous clinical reports of miR-21 over-expression in
different cancers (Iorio et al. 2005; Roldo et al. 2006; Slavy et al. 2007; Dillhoff et al. 2008;
Markou et al. 2008; Connolly et al. 2008). Future studies can identify such signaling
molecules, post-translational modifications, and structural changes in pri-miRNAs that can
affect Drosha accessibility and activity. Further, studies can investigate on what alterations of
these regulations can lead to altered pri-miRNA processing and tumorigenesis.
ii) DICER
DICER processes pre-miRNAs in the cytosol and is part of the RNase III family. A
few studies reported of its role in destabilization of mRNAs and gene expression (Jing et al.
2005; Takahashi et al. 2006). Moreover, mounting evidence reveal its role as critical
regulator of miRNA maturation and its deregulation in tumorigenesis. Whether DICER acts
as a proto-oncogene or a tumor suppressor is unknown, considering its ability to process
nearly all pre-miRNAs. In general, its over-expression has been predominantly associated
with tumorigenesis which points toward its role as a proto-oncogene. In contrast, there is also
evidence in support of DICER's role as a tumor suppressor. Nevertheless, these results
converge on the notion that aberrant DICER function contributes to tumorigenesis. Several
studies have reported its over-expression in a number of cancers including those of the
salivary gland (Chiosea et al. 2008), lung (Chiosea et al. 2007), prostate (Chiosea et al.

19

2006), and ovary (Flavin et al. 2008) as well as in Burkitt's lymphoma (Kaul et al. 2004).
Some of these studies pointed out that DICER over-expression can lead to a global upregulation of oncogenic miRNAs such as miR-21 and others (Chiosea et al. 2006; Volinia et
al. 2006). Future studies can focus on identifying those miRNAs of which their maturations
are affected by DICER over-expression and the reason for their specificity. It has been shown
that DICER can promote angiogenesis in endothelial cells and its expression can be
controlled by miRNA let-7 which is known as a tumor suppressive miRNA (Kuehbacher et
al. 2007; Tokumaru et al. 2008). Thus, these studies support the proto-oncogenic role of
DICER.
In contrast, DICER reduction and subsequent reduction in global miRNA maturation
can enhance tumor development (Blenkiron et al. 2007; Kumar et al. 2007). Reduction in
miRNA species has potential to dysregulate oncogenes. A few clinical studies are consistent
with this finding. Reduction of DICER was found to be a predictive marker for poor
prognosis in lung cancer patients (Karube et al. 2005). Another study showed downregulation of DICER being a strong predictor of relapse among non-small-cell lung cancer
patients (Rosell et al. 2006). Further studies should unravel more on the role of DICER and
its involvement in miRNA processing and in gene expression. For instance, it is warranted to
identify more miRNAs targets affected by aberrant DICER function and elucidate how they
can contribute to tumorigenesis.
iii) Argonaute 2
Pre-miRNAs are processed by DICER to become mature miRNAs that become part
of a miRNA ribonucleoprotein (miRNP) complex. Part of this complex is Argonaute 2
(Ago2) which can destroy mRNAs having complementary sequences to the template miRNA

20

(Blenkiron et al. 2007). If a nearly perfect complementary association can not be reached in
mRNA:miRNA binding, translational inhibition takes place. Ago2 has been reported to
interact with DICER (Song et al. 2004) and suggested to assist its pre-miRNA processing
(O'Carroll et al. 2007). It was shown to localize to the mRNA decay centres, known as
cytoplasmic bodies, to exert miRNA-mediated gene regulation (Sen and Blau 2005). Ago2 is
unique in that it is the only RNase from human Argonaute family comprised of eight
members, four of which are Agol through Ago4 that are closely related and co-expressed in
human cell lines (Meister et al. 2004; Rivas et al. 2005).
Recent reports highlight the altered levels of Ago2 in the deregulation of miRNA
activities in cancer (Kovalchuk et al. 2008). One study showed elevated Ago2 levels in the
more aggressive type of breast tumors (Blenkiron et al. 2007). Also, transfecting MCF7 cell
line with Ago2 resulted in increased cell proliferation as well as transformation of cells into a
phenotype undergoing endothelial-to-mesenchymal transition (Adams et al. 2008). How an
increase in Ago2 levels lead to tumorigenesis is still unknown. Possible explanations include
that the over-expression of Ago2 may relieve the competition implemented between different
miRNA species to regulate gene expression (Vickers et al. 2007) and permit oncogenic
miRNAs to better exert their effects. But, what seems to be more important is the
maintenance of balance of Ago2 levels since its reduction also results in impairment of
DICER's miRNA processing capabilities (O'Carroll et al. 2007). Future studies should
identify unknown facilitating factors that may promote the association of oncogenic miRNAs
to Ago2 and identify regulatory mechanisms exerted on Ago2. Recently, one of the
transducers of the MAPK pathway has been found to phosphorylate Ago2 at a Ser residue
and this enhanced its migration to the mRNA decay sites (Zeng et al. 2008). Identifying more

21

of such pathways will underscore the role of Ago2 as one of the down stream effector of
external signals and may provide reasons behind the correlation between its elevated levels
and tumorigenesis.

1.3.4 RNases of the nucleus/cytoplasm: Angiogenin, G3BP, and FEN1
Efforts are continuing to identify unknown RNases which play key role in the
regulation of gene expression. But, a few proteins with unexpected RNase activity that have
been identified were found to be linked to tumorigenesis and cancer progression. This section
will discuss these evidences. Angiogenin is essential in angiogenic processes whose altered
activity is implicated in cancer development. As well, G3BP is suggested to be the putative
endoribonuclease affecting the turnover of c-myc mRNA in mouse fibroblasts (Tourriere et
al. 2001) and its deregulated expression patterns were reported in cancer. Finally, the two
enzymes, APE1 and FEN1, originally found to act on DNA, are receiving much attention due
to their ability to cleave RNA and altered expressions reported in cancer.
i) Angiogenin
Angiogenin is part of a pancreatic RNase A superfamily. It was originally isolated as
a human tumor-derived angiogenesis factor that has its activity towards UA or CA
dinucleotides. It can specifically hydrolyze cellular tRNA to inhibit protein synthesis and
though its activity is weaker by several orders of magnitude compared to RNase A, it is
essential in angiogenesis (Curran et al. 1993). Unfortunately, its natural mRNA target has not
been identified and its role in mRNA turnover has not been studied. However, it is speculated
that its substrates may reside in the nucleolus of vascular endothelial cell where Angiogenin
normally localizes (Kao et al. 2002). Clinical studies associating Angiogenin and a variety of
cancers have been reviewed previously (Tello-Montoliu et al. 2006). Since it was initially

22

isolated as an angiogenesis factor from tumors, it is not a surprise to learn of its alterations
implicated in different cancers. Several studies concordantly report its higher expression
level correlated with higher proliferation of cancers. A study of a large cohort of
prostatectomy specimens found an increasing expression of Angiogenin in the progression
from benign prostate to high-grade prostatic intraepithelial neoplasia and ultimately to
prostatic adenocarcinoma (Katona et al. 2005). Also, higher Angiogenin mRNA was reported
in gastric carcinoma specimens than in the surrounding non-tumorous tissues (Chen et al.
2006) and recently, serum sample analysis of stage IV malignant melanoma reported marked
elevation of Angiogenin among a sub-group of patients with progressive disease (Vihinen et
al. 2007). Here, higher levels were significantly associated with poor treatment response with
chemo-immunofherapy and the treatment-related survival was shorter in patients with abovemedian levels of Angiogenin than in those with below-median levels. Further, overexpression of Angiogenin was reported to enhance the tumorigenicity and angiogenesis in
prostate cancer cell line while its knock-down exerted an opposite effect, in agreement with
its oncogenic role (Kawada et al. 2007).
In turn, its reduction in activity inhibits cancer proliferation. In melanoma cell line,
basic fibroblast growth factor cannot induce proliferation in the absence of Angiogenin (Song
et al. 2006). Knock down studies in HeLa cells showed notable inhibition of proliferation,
and in prostate adenocarcinoma cells, its knock down caused inhibition of rRNA
transcription, cell proliferation, and xenograft growth in mice (Tsuji et al. 2005; Yoshioka et
al. 2006). Six out of seven known variants are linked to amyotrophic lateral sclerosis and had
marked reduction in RNase activity while three lowest ones showed significant decrease in
angiogenic and proliferative activities (Crabtree et al. 2007). These reports are consistent and

23

demonstrate the importance of its activity in inducing cancer proliferation. It is also in line
with the previous evidence from inhibition studies, which alluded that RNase activity
controls the process of angiogenesis (Wang et al. 2005, Fu et al. 2005). Thus far, however,
the molecular basis of Angiogenin in inducing cell proliferation is unknown and would
require further studies to clearly elucidate its molecular mechanism (Gao et al. 2007).
ii) RasGAP Src homology 3 binding protein (G3BP)
RasGAP Src homology 3 binding protein 1 (G3BP) is an endoribonuclease
ubiquitously expressed in the cytoplasm and in the nucleus. G3BP cleaves the 3' untranslated
region of c-myc mRNA with a preference to cleave in between the CA dinucleotides. Its
influence on mRNA turnover has been suggested when a defective form of G3BP was found
in RasGAP deficient mouse embryonic fibroblasts that also showed delayed c-myc mRNA
decay (Tourriere et al. 2001). However, there has not been a direct proof of G3BP acting as
an endoribonuclease on c-myc mRNA in vivo. The endoribonuclease activity is dependent on
phosphorylation of its serine residues and, in dividing cells, G3BP migrates toward plasma
membrane and can interact with RasGAP, but not in quiescent cells (Tourriere et al. 2001;
Gallouzi et al. 1998). Its essentiality in development is demonstrated by embryonic lethality
and growth retardation in mice after its inactivation (Zekri et al. 2005).
Correlation of G3BP over-expression with a variety of cancers including that of
breast, head and neck, colon and thyroid have been reviewed (Irvine et al. 2004). Recently,
expression of G3BP has been studied in human esophageal squamous carcinoma (ESC)
(Zhang et al. 2007) and in 71% of the cases, there was a positive expression rate for G3BP.
Also, its expression in the lymph node metastasis group was significantly higher than that in
the non-lymph node metastasis group and this expression was associated with shorter

24

survival time. However, like Angiogenin, the molecular mechanism behind this association is
unknown and an opposing observation has also been made. One study reported the reduction
of G3BP was associated with the metastatic lung carcinoma more so than the non-metastatic
cells (Lie et al. 2001). It is possible that G3BP are predominantly found throughout the
cytoplasm, but during arsenite or high temperature stress, it can localize specifically in a
large cytoplasmic structures that resemble stress granules (SGs) (Tourriere et al. 2003). It is
known that SGs are cytoplasmic aggregates of stalled translational complexes that
accumulate during stress. One study proposed that the mRNA released from disassembled
polysomes is sorted and remodeled at SGs, from which selected transcripts are delivered to
processing bodies (PBs) for degradation (Kedersha et al. 2005). Whether G3BP associate
with subset of these transcripts in SGs and can destroy them prematurely is still unknown.
Future studies can clarify this by understanding the regulation of its localization to SGs and
identifying its target mRNA associated with SGs.
iii) Flap endonuclease 1 (FEND
Human flap endonuclease 1 (FEN1) was discovered to have an endoribonuclease
activity against synthetic and native RNAs in vitro (Stevens 1998). Whether FEN1 is capable
of endonucleolytically cleaving RNA in cells is currently unknown. Human FEN1 is
normally found in the nucleus and shows preference to cleave 5' endogenous stem structure
of RNAs in vitro. Multiple functions of FEN1 include RNA primer removal during DNA
replication, dsDNA 5' to 3' exonuclease, gap endonuclease, and RNase H activity (Robins et
al. 1994; Shen et al. 2005). Interestingly, the exonuclease activity of FEN1 seemed to have a
role in apoptotic DNA fragmentation while the gap endonuclease activity showed potential
for repairing stalled DNA replication fork. Similarly to APE1, FEN1 may also be able to

25

regulate the levels of RNAs apart from its well known function in removing RNA primers
during DNA replication. In cancers, it displays two opposing roles. Its over-expression has
been reported to correlate with the aggressiveness of tumors (Sato et al. 2003; Lam et al.
2006). Mainly, its ability to assist in DNA replication via removal of RNA primers may
promote cell proliferation. However, whether its endoribonuclease activity against other
cellular RNAs plays a role in this development still needs to be assessed.
Its reduction in DNA endonuclease activity has also been associated with cancers. Its
activity is reduced by a member of the signal transduction pathway promoting cell growth in
vitro (Henneke et al. 2003). Previous studies have shown its haploinsufficiency contributes to
rapid progression of tumours (Kucherlapati et al. 2002). A recent report from mice model
found FEN-1 deficiency contributes to modest but a significant increase in tumor incidence
and multiplicity (Kucherlapati et al. 2007). The loss in RNA primer removal as well as
repairing stalled DNA replication fork may account for this observation. But, in another
perspective, its ability to cleave RNAs may also play a role in such observations. In order to
fully appreciate the extent of its deficiency, future studies should be carried out to look for its
RNA targets. Also, in certain situations, it is retained in the cytoplasm with an unknown role.
FEN1 has a nuclear localization signal (NLS) located at its C terminus, which directs its
localization during S phase of the cell cycle and in response to DNA damage (Qiu et al.
2001). Truncation or alanine substitutions on the KRK residues of the NLS prevent its
migration from the cytoplasm, but have no effect on its nuclease activity. Further studies can
assess the effects of its retention in the cytoplasm that may involve regulating the levels of
mRNAs.

26

1.4 Apurinic/apyrimidinic endonucleasel (APE1) as an endoribonuclease
c-Myc, a proto-oncogene, has been implicated in the development of virtually all
types of human cancers (Levens 2003). Like other cellular mRNAs, c-myc mRNAs, can be
initiated to decay via exoRNases as described in section 1.1. However, c-myc mRNA can
also be degraded endonucleolytically. Endonucleolytic cleavage of c-myc mRNA was first
shown using a polysome-based in vitro mRNA decay assay (Bernstein et al. 1992). It was
believed that a putative endoRNase targeted the specific exposed region of an endogenous
polysome-associated c-myc mRNA referred to as the c-myc coding region determinant or
CRD. Recent evidence suggested that translational pausing in CRD could result in ribosomedeficient region that is susceptible to endonucleolytic cleavage (20). Other studies have
confirmed that the coding region of c-myc mRNA, including the CRD, is involved in the
regulation of c-myc mRNA stability in cells (Levens 2003). Moreover, endoRNase decay
intermediates for c-myc mRNA have been detected in cells (Hanson and Schoenberg 2001;
Ioannidis et al. 1996; Herrick and Ross 1994) and this provided further support for the
significance of c-myc mRNA decay initiated via an endoRNase. Later, it was discovered that
apurinic/apyrimidinic endonuclease 1 (APE1) was the endoribonuclease that can bind to
CRD region of c-myc mRNA, and specifically cleave in between the single stranded region
of UA, CA, and UG dinucleotides (Barnes et al. 2009).
1.4.1 Possible role of APE1 in cancer
Apurinic/apyrimidinic endonuclease 1 (APE1) is an enzyme that has been found in
the nucleus, cytoplasm, and mitochondria (Evans et al. 2000; Tell et al. 2005; Chattopadhyay
et al. 2006).

Its elevated expression has been observed in some cancers. In multiple

myeloma, positive rate of APE1 expression was found in 65.6% of the patients (Yang et al.

27

2007). Similarly, elevated expression of APE1 was associated in osteosarcoma resistance
and poor prognosis of patients (Wang et al. 2004). Altered subcellular localization of APE1
or its variants have also been reported in a variety of cancers (Tell et al. 2005). The
cytoplasmic expression of APE 1 is significantly higher in hepatocellular carcinoma than the
surrounding tissues (Di Maso et al. 2007). In normal liver, they found no APE1 in the
cytoplasm. A 3-fold increase of cytoplasmic expression in aggressive HCC patients
compared to those with well-differentiated HCC and cytoplasmic localization was correlated
with shorter survival time compared to those with negative cytoplasmic reactivity. APE1 was
considered as a nuclear protein containing a nuclear localization signal near its N-terminus
(Jackson et al. 2005; Chattopadhyay et al. 2006). However, re-distribution of APE 1 between
the nucleus and the cytoplasm in cancerous cells has raised many questions about its role in
the cytoplasm. It is tempting to speculate that the RNase activity of APE1 in the cytoplasm is
correlated with aggressive cancer phenotype. Also, cleavages of cellular RNAs, such as
microRNAs or tumor suppressor mRNAs, can certainly trigger tumorigenesis as envisioned
in the case of RNase L (Malathi et al. 2005). To test the above hypotheses, further studies are
clearly required to identify RNA targets of APE 1.
On the opposite side, APE1 may exert its RNase activity to destroy oncogenic
mRNAs and miRNAs. This may be consistent with the fact that oncogenic Bcl2 suppresses
the expression of APE1 while enhancing that of c-Myc (Jin et al. 2006). The authors were
unclear of the molecular mechanism behind this process, but one model that can be deduced
from this is that APE1 can destroy c-myc mRNA and regulate its turnover. This is supported
by recent evidence that down regulation of APE1 leads to c-Myc over-expression (Barnes et
al. 2009). In addition, there are seven substitution variants of DNA endonuclease domain of

28

APE1 in human population (Hadi et al. 2000). Several of these, notably, L104R, E126D,
R237A, and D283G show 40-90% reductions in apurinic/apyrimidinic DNA endonuclease
(AP-DNase) activity. D148E with an allele frequency of 0.38, surprisingly retains its APDNase activity as well as substrate binding. In cancer, homozygous individuals (E/E) for
D148E variant were more associated with colon cancer (Pardini et al. 2007). Similarly, it was
found in patients that D148E polymorphism was significantly associated with lung cancer
risk (De Ruyck et al. 2007). Therefore, these reports indicate its AP-DNase activity cannot
be responsible for this observation and suggest that alteration in the activity of APE1 may be
associated with these cancers.
The post-translational (PT) modification of APE1 has also been linked to cancers.
One study observed an abundance of post-translationally modified form of APE1 in
leiomyoma extracts, suggesting its association with uterine smooth muscle tumorigenesis
(Orii et al. 2002). It is unclear from this study what modification was done on APE1 but the
authors suggested of multiple processing steps involving phosphorylation and proteolytic
cleavage. Higher level of the modified APE1 (a larger sized APE1 compared to the regular
molecular weight) was also observed in cell lines derived from leiomyosarcomas (Orii et al.
2002). Further, relative abundance of the phosphorylated APE1 was found to correlate with
proliferating cell nuclear antigen levels, suggesting a correlation with increased proliferation.
Here, the authors did not assess the effects of phosphorylation on AP-DNase activity nor on
any of its known functions. At this stage, possibility of alteration in ribonuclease activity
through PT modification can only be speculated. Recent study showed that APE1 nitrosation
of C93 and C310, a consequence of oxidative stress caused by nitric oxide, eliminates its
nuclear export to the cytoplasm (Qu et al. 2007). Preventing the channeling of APE1 to

29

different parts of the cell and interaction with different RNA substrates will most likely affect
mRNA turnover and gene expression. On a similar note, APE1 was found to have 3-fold
increase in its AP-DNase activity when its first 33 amino acid sequence is removed by
proteolysis and exported out of the nucleus to the mitochondria (Chattopadhyay et al. 2006).
1.4.2 Abasic DNA incision activity of APE1
Discovery of APE1 as an endoribonuclease was unexpected and encouraged us to
further characterize this activity in vitro. Its function as an endoRNase in any of the cellular
compartments is completely unknown. Previous studies describe APE1 to possess multiple
activities in cell-free and cellular systems. These activities included the cleavage of
apurinic/apyrimidinic DNA (abasic DNA) (Kane and Linn 1991) and abasic-RNA (Berquist
et al. 2008; Vascotto et al. 2009), redox activation of transcription factors implicated in
apoptosis and cell growth (AP-1, Egr-1, NF-kappaB, p53, c-Jun, and HIF) (Tell et al. 2009),
as well as 3' phosphodiesterase (Chen et al. 1991), 3'-5' exonucleases (Wilson et al. 1995), 3'
phosphatase, and RNase H activities (Barzilay et al. 1995). APE1 is also referred to as redox
effector factor 1 (Ref-1) because it was independently discovered to possess a redox activity.
In regards to its AP-DNA incision activity, it specifically recognizes the abasic site
and facilitates the phosphodiester bond cleavage facilitated by divalent metal ions and water
molecules. It cleaves the phosphodiester bond immediately 5' to the abasic site, generating
products with 3' hydroxyls and 5'-deoxyribose phosphate termini (Demple and Harrison
1994). These products are then processed by enzymes such as DNA polymerases and ligases
that fill the nucleotide gap and seal the nick in the DNA to complete the AP site repair
(Evans et al. 2000). In mammalian cells, APE1 is the major protein involved in initiating the
removal of abasic sites in DNA (Wilson and Barsky 2001).

30

Accordingly, efforts were put into identifying its active site and essential residues
critical in abasic DNA incision. A few biochemical kinetic studies have found APE1 against
abasic sites in ssDNA yielded an apparent Km of 428 nM and a k^, of 4.2 s_1, while against
abasic sites in dsDNA yielded an apparent Km of 24 -33 nM and a kcat of 3-4.1 s_1
(Sokhansanj et al. 2002; Marenstein et al. 2004). Several crystal structures in different
conditions have been reported (Gorman et al. 1997; Mol et al. 2000; Beernink et al. 2001).
Later, it was identified that ten amino acids constitute the active site of APE 1 in carrying out
abasic DNA incision, and was confirmed of their evolutionary conservation across different
species (Tell et al. 2005).

Figure 3: Abasic DNA cleaving active site model of APE1 based on X-ray
crystallography (Mol et al. 2000). The XFIT generated model of APE1 active site. The
orange lines represent phosphate backbone of abasic DNA strand while resides of the active
site of APE1 is indicated by their numbers. The green sphere represents Mn2+ and dotted
lines represent hydrogen bonds.

31

Amino acid residues Asp70, Asp90, Glu96, Tyrl71, Asp210, Asn212, Asp219,
Asp283, Asp308, and His309 have been noted as the critical amino acids in the active site
(Figure 3). Substitution of any of these residues results in varying degrees of reduction in
AP-DNA nuclease activity, and these findings are summarized in Table 1.
Table 1: Summary of abasic DNA incision activities of active site mutants (or structural
mutants).
Mutants
N68A
D70A
E96A
Y171F
D210N
N212A
N212Q
F266A
D283N
D308A
H309N

Fold reduction in abasic DNA
incision activity
600
Reduction
600-100000
5000
25000
Not detectable
300
6
10
5-25
100000

Reference
Nguyen et al. 2000
Nguyen et al. 2000
Erzberger et al. 1999; Chou and Cheng 2003
Erzberger efaZ. 1999
Erzberger et al. 1999
Rothwell and Hickson 1996
Rothwell and Hickson 1996
Erzberger et al. 1998
Hadi et al. 2000
Erzberger et al. 1999; Barzilay et al. 1995
Chou and Cheng 2003

It is noteworthy that substituting Asn212, for example, a polar residue to glutamine,
another polar residue, resulted in a 300 fold reduction whereas mutating it to alanine, a
hydrophobic residue, resulted in a complete abrogation of its nuclease activity (Rothwell and
Hickson 1996). It is predicted that the amino group attached to the side chain of Asn212 can
interact with oxygen on the phosphate backbone on the AP-DNA, helping to orient the
scissile bond for incision to take place. Thus, it is thought that certain charge-charge
interactions between the residues and the substrates are required to give abasic DNA incision
activity.
Previous studies on abasic DNA catalysis and crystal structure of APEl:abasic DNA
complex suggest that APE1 binds to nucleic acid structure that can adopt a uniquely kinked
conformation. It seems that APE1 probes for substrates such as abasic DNA or single

32

nucleotide-gapped DNA that lack base-to-base stacking interaction and those that have
increased backbone flexibility (Wilson and Barsky 2001). It is thought that during the
binding of APE1 to abasic-DNA, the DNA undergoes a large extent of deformation in order
to accommodate for a more stringent structural nature of APE1. APE1 was shown to have a
preformed DNA binding surface that undergoes little conformational change while it uses its
hydrophobic pocket to accommodate abasic site and exclude normal bases (Wilson and
Barsky 2001).
In cleaving abasic DNA, APE1 is dependent on divalent metal ions and displays
maximal activity in the range of 0.1 mM to 2mM Mg2+concentrations under in vitro assay
conditions (6.6mM Tris-HCl, pH 7.5, 0.5mM MgCl2, O.lmM 2-mercaptoethanol) (Barzilay
et al. 1995). It is thought that metal ions have no effect on substrate binding, but their role
becomes critical during catalysis. They are predicted to be involved in substrate orientation,
transition-state stabilization, and the dissociation of the incised product (Rothwell et al.
2000). This is supported by a metal-chelation experiment where EDTA inactivated nuclease
activity of APE1 is rescued to a varying extent by addition of different metal ions. Among
these, magnesium showed the highest rescue rate of nuclease activity followed by
manganese, nickel, zinc, and potassium, in decreasing order. Also, APE1 can be induced to
bind stably to abasic site-containing oligonucleotides (or oligonucleotides with modified
abasic sites such as tetrahydrofuran) by chelation of the active site metal ion (Rothwell et al.
2000). In this way, the activity of APE1 can be 'trapped' in a stable complex with DNA at a
stage subsequent to substrate recognition, but prior to phosphodiester backbone incision.
Hence, the APE1 catalytic cycle can be viewed as being comprised of two steps that are
separable both mechanistically and temporally.

33

1.4.3 Variants of APE1 identified in human populations
A few APE1 variants identified from the human population (Figure 4) were
speculated to be associated with increased disease susceptibility (Hadi et al. 2000).

Figure 4: Molecular model of the positions of the residues found to be mutated in
human population (Hadi et al. 2000). (A) Substitution mutations were found in these
residues indicated by different colours: L104 (orange), E126 (green), D148 (peach), R237
(red), G241 (green), D283 (purple) and G306 (maroon). DNA binding groove is located at
the top, with a metal cofactor represented as a ball. (B) Comparison of local structure in WT
APE1 (left) versus L104R (right). The hydrophobic packaging of the local structure is
predicted to be disrupted by the presence of R104. (C) Comparison of disruption in hydrogen
bonding network of the WT versus R237A. Dashed lines indicate hydrogen bonds.

34

In a total of seven variants tested (L104R, E126D, D148E, R237A, G241R, D283G,
and G306A), reductions in abasic DNA incision activity were observed in L104R, E126D,
and R237A (Table 2). These variants were further discussed to have association with an
increased risk of developing cancer (Hadi et al. 2000). Hadi et al. excluded a few of the
variants that had mutations lying outside of the DNA endonuclease domain (L44C, Q51H,
G57A and I64V). We included in our test Q51H and I64V, notably because these variants
have never been assessed of their DNA endonuclease activities as well as RNA incision
activity. With these, we tested the RNA incision activities of the six human variants except
D283G.
Table 2: Summary of dissociation constants (Kd) and abasic DNA incision activities of
APE1 population variants (Hadi et al. 2000).

WT
L104R
E126D
D148E
R237A
G241R
D283G
G306A

K d (nM)
25.8 + 12.2
54.3 + 19.7
44.0 + 7.2
20.3 ± 3.4
26.5 + 14.1
20.1+0.1
N/A
35.3 ±13.9

Incision
100
56.71 + 26.80
60.43 + 11.16
94.36 + 6.20
35.71+9.68
108.73 + 5.31
N/A
107.22+17.13

1.4.4 3'-5' Exonuclease activity of APE1
It was found that the strength of human APE1 3'-5' exonuclease activity is weaker
than its abasic incision activity by 4-33 fold (Demple et al. 1991; Robson et al. 1991; Wilson
et al. 1995). This is in contrast to the E. coli homolog of APE1, Exo III, which is a far better
3'-5' exonuclease, displaying its abasic endonuclease and exonuclease activities at a similar
efficiency (Wilson et al. 1998). The active site residue difference seemed to account for this
difference between the two enzymes.

35

In terms of substrate specificities, human APE1 showed varying efficiencies (Kcat/Km)
on different DNA substrates (5' overhang dsDNA = 1 nt gapped dsDNA > nicked dsDNA >
2 nt gapped dsDNA » ssDNA = blunt ended dsDNA (Suh et al. 1997; Hadi et al. 2002), but
appeared to exhibit a similar affinity (Km value of -250 nM) for these exonuclease substrates.
It was concluded that human APE1 seemed to be mostly affected by the substrate context at
the catalytic step (Wilson 2003). It was also found that APE1 is a non or poorly processive
exonuclease (Wilson 2003), confirming that this activity of APE1 is weaker than its abasic
DNA incision activity.
Nonetheless, its significance was re-examined after its exonuclease activity showed to
remove various 3' termini blocking groups on DNA. Human APE1 has been found to remove
3'-phosphoglycolates. This excision is highly selective, with marked preference for gapped
DNA substrates in comparison to single-stranded (ss) oligonucleotides or blunt end duplexes
(Suh et al. 1997). It is also noteworthy that human APE1 can remove 3' mismatched
nucleotides at least 50 times more efficiently than those matched correctly (Chou and Cheng
2002) which was recognized as another 3' proofreading function of APE 1.
In linking its activity to biological relevance, human APE1 was also found to be a
strong remover of stereochemically unnatural L-configuration

anticancer nucleoside

analogues (Chou et al. 2000), and beta-L-dioxolane-cytidine (Beta-L-OddC, BCH-4556;
Troxacitabine) (Grove et al. 1995), and other beta-L-configuration nucleoside analogues
from the 3' terminus of DNA in leukaemic cell nuclei. Also, in retroviral therapy against
HIV, certain nucleoside analogues such as 3'-azido-3'-deoxythymidine (AZT) are often used
to block viral DNA replication. APE1, in contrast to other cytosolic exonucleases, can
remove this analogue from the 3' terminus of a nick efficiently (Chou and Cheng 2002). As

36

APE1 is expressed constitutively in human cells, it could therefore be used to limit the
cytotoxicity of AZT and other anti-HIV compounds. Later, APE1 was found to remove 3'tyrosyl residues, which could result from erroneous function of Topoisomerase I, via a
phosphodiesterase or endonucleaselike activity, again depending on the DNA structural
context (APE1 removes 3'-tyrosyl residues from 3'-recessed and nicked DNAs) (Wilson
2003).

1.5

Research objectives
There is increasing evidence that endoribonucleolytic mRNA decay plays a critical

role in mammalian gene expression (Lebreton et al. 2008; Schaeffer et al. 2009; Eberle et al.
2009). Efforts

are being added to identify

and understand novel mammalian

endoribonucleases (endoRNases). APE1 has been recently purified and determined to
possess a novel endoRNase activity in vitro (Barnes et al. 2009). In order to further
understand this activity and its biological significance, the following research objectives were
established.
The first objective was to definitively assess whether the RNA-cleaving activity of
APE1 shares the same active centre as abasic DNA endonuclease activity. To accomplish
this, we assessed the RNA cleaving activities of previously generated active site mutants of
APE1 used in elucidating abasic DNA endonuclease activity. The results obtained from
testing the two RNA substrates (c-myc-1705-1792 CRD RNA, and Oligo IB) were expected
to definitively identify the active site residues of APE1. Also, we intended to assess the RNA
cleaving activities of variants of APE1 found in human population and compare them to the
abasic DNA endonuclease activities. This was expected give us a better understanding in
correlating some of these mutants and their aberrant RNA-cleaving activities in association

37

with tumorigenesis. Finally, to support our findings and to understand the RNA incision
catalysis, the endoribonuclease activity of APE1 was subjected to further biochemical
characterization.
The second objective was to confirm the preferred RNA structure and sequence
specificity for endoRNase activity of APE1 as well as to assess the biological significance of
the endoRNase activity of APE1. The first part of the objective was carried out using five
RNA substrates with different secondary structures and sequence. The experiments were
mainly done using RNA sequencing gels to identify the sites and local RNA structures that
were preferentially cleaved by APE1. The second part of the objective was carried out in
cultured HeLa cells and using siRNA technology to knock down the levels of APE 1 to assess
its influence in c-myc mRNA turnover. Also to support this assessment, we tested the effect
of APE 1 expression in the growth of E. coli Origami cells which has been previously used to
assess the effects of ribonuclease activity of RNase A and Angiogenin (Smith and Raines
2006; Smith and Raines 2008).

38

CHAPTER 2
Identifying the critical amino acid residues for endoribonuclease
activity of APE1 and characterizing its biochemical properties
Preliminary analysis showed APE1 is capable of cleaving at multiple sites of its
substrate RNA (Barnes et al. 2009). However, its active site residues for carrying out its
RNA cleavage were unknown and it is intriguing to assess if these residues were shared
between APE1 endoribonuclease activity and AP-DNA endonuclease activity. By testing
recombinant mutants of APE1 that have amino acid substitutions in critical residues for APDNA endonuclease activity (Table 1), we evaluated their importance during RNA cleavage.
As part of a collaborative effort, we have obtained APE1 structural mutants from Dr. David
Wilson's laboratory at National Institute on Aging, NIH. Also, APE1 variants identified in
the human population were also obtained from Dr. Wilson and assessed for their alterations
in RNA cleaving activity. A few of these variants have been found to possess functional APDNA endonuclease activities (Hadi et al. 2002), however they were reported to be associated
with the development of cancer. Therefore, it was intriguing for us to find out if there were
any disparities in AP-DNA and RNA cleaving activities of these variants.
Also, endoribonuclease activity of APE1 needed to be characterized under various
biochemical settings. This information will provide clues to the catalysis mechanism in RNA
cleavage and may become beneficial in the long run for developing APE1 in applications for
therapeutic purposes. We characterized the effects of metal ions as well as identified the end
of its RNA cleavage product. In addition, robustness of its RNA cleaving activity was
assessed by testing its sensitivity to RNasin, and a known inhibitor of RNase.

39

2.1 Methodology
This section describes the methods implemented in identifying the critical amino acid
residues in endoribonuclease activity of APE1 and understanding its biochemical properties.
2.1.1 Reagents and buffer preparation
These reagents shown in Table 3 were used through out the course of experiments
including those in chapter 3.

Table 3: Composition of Reagents used in this chapter
Composition
Reagent
0.5x TBE
0.45 M Tris-HCl/Boric acid (Sigma) pH
8.3, 5 mM EDTA
lOx T7 Transcription Buffer
400 mM Tris-HCl pH 7.6, 240 mM MgCl2,
20 mM spermidine, 0.1 % Triton X-100
lOx Glycogen
0.2 mg/mL Glycogen, 3M sodium acetate
pH5.2
RNA Elution Buffer
lx TE pH 7.5, 0.01 % SDS, 0.1 M NaCl
lxTE
lOmM Tris-HCl pH 8, ImM EDTA pH 8
lOx Alkali Buffer
500 mM sodium bicarbonate pH 9.2 and 10
mM EDTA
Stopping Dye
9M Urea, 0.01% bromophenol blue, and
0.01% xylene cyanole FF
0.25% bromophenol blue, 0.25% xylene
Agarose Loading Dye
cyanole FF, 15% ficoll

2.1.2 PCR generation of linear templates
The c-myc-1705-1792 CRD RNA was generated in the lab while Oligo IA, dOligo
IA, and Oligo IB were commercially synthesized from IDT, San Diego, CA (Table 4). To
prepare linear DNA templates for in vitro generation of RNA substrates, the pUC19 plasmids
containing cDNAs of c-myc (nucleotides 1705 - 1792) CRD RNA was amplified via PCR in
the region including the T7 promoter and cDNA sequence. The sequences of the primer used
are shown in Table 5.

40

Table 4: List of RNA substrates used in assessing the RNA-cleaving activity of APE 1 and its
mutants. rU and dU indicate ribose and deoxyribose moieties linked to the Uridine bases,
respectively.
RNA Substrate
c-myc
(nts 1705-1792)
CRD RNA
Oligo IA
dOligo IA
Oligo IB

cDNA/Oligonucleotide sequence
5'CCAGATCCCGGAGTTGGAAAACAATGAAAAGGCCC
CCAAGGTAGTTATCCTTAAAAAAGCCACAGCATACA
TCCTGTCCGTCC AAGC-3'
5' -C AAGGTAGTrUATCCTTG-3'
5' -C AAGGTAGTdUATCCTTG-3'
5' -CAAGGrUAGTTATCCTTG-3'

To generate the linear template, a total reaction mixture of 50 jal containing 100 ng of
pUC19 plasmids, 2U of Phusion DNA polymerase (Finnzymes), 0.3 mM dNTPs, lx Phusion
HF Reaction Buffer (Finnzymes), 45 pmoles of forward and reverse primers. The mixture
was put into the PCR MiniCycler PCR machine (MJ Research Inc., Watertown, MA) which
was programmed to incubate the mixture for 18 cycles of 45 sees at 95 °C, 45 sees at 55 °C,
10 mins at 68 °C, 20 min extension at 68 °C, hold at 4 °C to generate linear DNA template.
Once the PCR was finished, the mixture was added with 2 ul of agarose loading dye and run
in 1% agarose gel for lhour at 120V. The PCR fragment was visualized under UV and
excised. The PCR fragment was purified using QIAEX II Gel Extraction Kit (Qiagen Inc.,
Valencia CA) and following the manufacturer's protocol.
Table 5: PCR primers and their sequences used in the generation of linear DNA template for
c-myc (1705-1792) CRD RNA by in-vitro RNA transcription.
Primers
Sequences
c-myc (1705-1792 or 17055' -GGATCCTAATACGACTC ACTATAGGACC A
1886) CRD RNA Forward
GATCCCGGAGTTGG-3'
c-myc (1705-1792) CRD RNA 5' -GCTTGGACGGACAGGATG-3'
Reverse
c-myc (1705-1886) CRD RNA 5' -CGCACAAGAGTTCCGTAG-3'
Reverse

41

2.1.3 Standard phenol/chloroform extraction and ethanol precipitation
To the reaction mixture of 200 jul, Vi volume of phenol (pH = 6.7, Sigma) and Vi
volume of chloroform : isoamyl alcohol (CHCI3 : IAA = 49 : 1, Fluka) were added and
vortexed. Note that for extracting RNA, phenol (pH = 4.3, Sigma) was used. The mixture
was centrifuged at 12,000 rpm for 5 mins at 4 °C and the aqueous layer was aliquoted and
transferred into a fresh tube. Next, 1 volume of CHC13 : IAA = 49 : 1 was added and
vortexed and centrifuged at 12,000 rpm for 5 mins. The aqueous layer was transferred into a
fresh tube and l/10th volume of 3M sodium acetate (NaOAc) pH 5.3 was added followed by
2 volumes of 100% ethanol. The mixture was vortexed and stored at -20 °C for 20 mins. The
mixture was centrifuged at 12,000 rpm for 10 mins to precipitate the plasmid and the
supernatant was poured off. Next, the pellet was rinsed with 200 pi of 70% ethanol and
centrifuged at 12,000 rpm for 5 mins. The supernatant was removed by pouring and the pellet
was dried inside fume hood for 20 mins. The pellet was resuspended with 20 pi of autoclaved
water to give a final concentration of 0.1 - 0.4 ug/ul measured by ND-1000 UVSpectrophotometer (NanoDrop V.3.1.0, Wilmington, Delaware). The concentration was
measured by using the relationship: DNA concentration (ug/ml) = (OD260) x (dilution factor)
x (50 ug DNA/ml) / (1 OD260 unit).
2.1.4 Generation of unlabeled RNA substrates
To generate RNA from linear plasmids, overnight in vitro transcription was
performed. A typical 100 pi reaction contained -4.5 pg of linear plasmid, 10 pi of 100 mM
DTT (Promega, Madison, WI) 1 pi of 40 U/pl RNasin (Promega, Madison, WI), 10 pi of lOx
T7 transcription buffer (Table 3), 5 pi each of 100 mM ATP, CTP, GTP, UTP, 5 pi of 19
U/pl T7 RNA Polymerase (Promega, Madison, WI) and adjusted to the volume with

42

diethylpyrocarbonate (DEPC) (Sigma) treated Milli-Q-ddF^O. The reaction was done at 37
°C for typically 16 hrs. After incubation, the sample was centrifuged at 12,000 rpm for 1 min
and 1 ul of the supernatant was aliquoted into a fresh tube. The rest of the supernatant was
transferred into a fresh tube and 10 ul of 1 U/ul RNase-free RQ1 DNase (Promega, Madison,
WI) was added and incubated at 37 °C for 30 mins. 1 ul of the reaction mixture was aliquoted
into a fresh tube and loaded along with the pre-treated sample (1 ul) on a 2.5% agarose gel to
confirm the digestion of plasmid. The gel was prepared and visualized similarly to the
method described in 2.1.2 except 1.25 g of agarose was used and the gel ran for about 50
mins. Next, 5 ul of phenol (pH = 4.3, Sigma) was added to the DNase treated sample and
vortexed. Then ProbeQuant™ G-50 micro columns (GE Healthcare, Buckinghamshire, UK)
were vortexed and the bottom closure was snapped off followed by centrifuge at 3,000 rpm
for 1 min and flow through discarded. The vortexed DNase treated sample was split into two
50 ul samples and each was applied to the column resin and centrifuged at 3,000 rpm for 2
mins. The flow throughs were collected and combined (-120 ul). Standard ethanol
precipitation was performed. l/10th volume of (12 ul) of 3 M NaOAc pH 5.2 and 2 volumes
(240 ul) of 100% ethanol were added and the sample was stored in -20 °C for 20 mins. After,
the sample was centrifuged at 12,000 rpm for 10 mins and its supernatant removed by
pouring. Then, 200 ul of 70% ethanol was added to rinse the pellets and centrifuged at
12,000 rpm for 5 mins. The supernatant was poured off and the pellet was dried in fume hood
for 20 mins. The pellet was resuspended with 80 ul of DEPC treated water and its RNA
concentration was determined by ND-1000 UV-Spectrophotometer (NanoDrop V.3.1.0,
Wilmington, Delaware). The concentration was measured by using the relationship: RNA
concentration (ug/ml) = (OD26o) x (dilution factor) x (40 ug RNA/ml) / (1 OD26o unit).

43

2.1.5 Generation of 5'-radiolabeled RNA substrates
First, in vitro transcribed RNA (0.2 - 1.0 ug/ul) was dephosphorylated at the 5' end.
Typically two reactions were performed in parallel to produce a higher yield of
dephosphorylated RNA after extraction. A typical 200 ul dephosphorylation reaction
contained 10 ug of RNA (e.g. 333 pmol of c-myc 1705-1792 CRD RNA), 20 ul of lOx NEB
Buffer 3 (New England Biolabs, Beverly, MA), 0.5 ul of 5 U/ul Calf Intestine Phophatase
(New England Biolabs, Beverly, MA) diluted from 10 U/ul in lx NEB Buffer 3, and volume
adjusted with DEPC treated water. The reaction was carried out at 37 °C for 60 mins. Next, 1
ul of 0.5M EDTA pH 8, 4 ul of 5M NaCl, and 10 ul of 20% SDS were added to the mix.
Then, RNA was subjected to a standard phenol/chloroform extraction and ethanol
precipitation as described in 2.1.3. Only exceptions from this protocol were the usage of pH
4.3 phenol (Sigma) and resuspending the RNA from two reactions in 20 ul of DEPC treated
water to give a concentration around 0.25 - 0.8 ug/ul.
The 5'-radiolabeling of the dephosphorylated transcript was performed in a 12.5 ill
final volume consisting of 0.38 ug of the dephosphorylated transcript (e.g. 12.5 pmol of cmyc 1705-1792 CRD RNA) and 38 uCi or 12.5 pmol of y-32P-ATP (PerkinElmer, Boston,
MA), 0.5ul of 1 U/ul of T4 Polynucleotide Kinase (PNK) (New England Biolabs, Beverly,
MA), 1.25 ul of lOx PNK buffer, and DEPC treated water. The reaction mix was incubated
for 1 hour at 37 °C. The reaction mix was stopped by adding 25 ul of Stopping Dye which
contains 9M Urea, 0.01% bromophenol blue, and 0.01% xylene cyanole FF. Subsequently,
the stopped mixture was loaded onto an 8% polyacrylamide

(29:1,

acrylamide:

N,N'=metheylenebisacrylamide) gel containing 7M urea (Invitrogen) and ran at 20 mA with
0.5x TBE buffer (0.45 M Tris-HCl/Boric acid (Sigma), 0.01 M EDTA pH 8.3 for 1 hour or

44

until the bromophenol blue dye was near the bottom of the gel. The gel was exposed onto a
Cyclone Storage Phosphor Screen System (Packard, Meriden, CT) and gel image was
developed using Cyclone Phospholmager and visualized by the OptiQuant software (Hewlett
Packard, Palo Alto, CA). The polyacrylamide gel containing the radiolabeled transcript was
sliced out and put into a tube containing 400 ul of RNA Elution Buffer (lx TE pH 7.5, 0.01
% SDS, 0.1 M NaCl) and 200 ul of phenol (pH = 4.3, Sigma). The elution reaction lasted 5
hrs at 45 °C, followed by standard phenol/chloroform extraction and ethanol precipitation.
The RNA was resuspended in 50 ul of DEPC treated water and diluted appropriately in order
to give a final count of approximately 100,000 c.p.m./ul.
2.1.6 Generation of 5'-radiolabeled abasic DNA substrate
Commercially available 18-mer oligonucleotide, 5'-GTCACCGTGFTACGACTC-3'
(IDT, San Diego, CA) that contained the model analog of an abasic site, tetrahydrofuran (F)
(Figure 5) (Hadi et al. 2000) was used. The sequence of its complementary strand was 5'GAGTCGTAACACGGTGAC-3' (IDT, San Diego, CA). The oligonucleotides were adjusted
to a concentration of 1 ug/ul or 187 pmol/ul for sense strand and 1 (xg/u.1 or 177 pmol/ul for
anti-sense strand by addition of autoclaved water.
5'
\

O—i

0

0=P—O

I
Figure 5: Chemical structure of abasic site analog represented as F in the cDNA
sequence.

45

Dephosphorylation
The oligonucleotide containing the abasic site was dephosphorylated in a typical
reaction of 80 ul that contained 4 ug of oligonucleotide, lx NEB buffer 3 (New England
Biolabs, Beverly, MA), 2 U of Calf Intestine Phophatase (New England Biolabs, Beverly,
MA) diluted from 10 U/ul in lx NEB Buffer 3, and autoclaved water. The reaction was done
at 37 °C for 60 mins
Standard phenol/chloroform extraction
Next, 120 ul of autoclaved water was added to the reaction mix followed by the
addition of 1 ul of 0.5M EDTA pH 8, 4 ul of 5M NaCl, and 10 ul of 20% SDS. Then, the
dephosphorylated oligonucleotide was subjected to a standard phenol/chloroform extraction
and ethanol precipitation as described in 2.1.3. Only exceptions from this protocol were the
replacement of 3M sodium acetate (NaOAc) pH 5.3 with lOx glycogen (0.2 mg/ml), which
was added at a final concentration of 20 ug/mL to the aqueous layer before the addition of
100% ethanol. This was to improve the precipitation of shorter length oligonucleotides. The
precipitated oligonucleotide was resuspended in 20 ul of autoclaved water to give a
concentration of approximately 0.2 ug/ul.
5'-radiolabeling and hybridization
The 5'-radiolabeling of the dephosphorylated oligonucleotide was performed in a 25
ul final volume consisting of 0.13 ug (25 pmol) of the dephosphorylated oligonucleotide and
75 uCi (25 pmol) of y-32P-ATP (Amersham GE Healthcare, Montreal, QC), 10 U of T4
Polynucleotide Kinase (PNK) (New England Biolabs, Beverly, MA), lx PNK buffer (New
England Biolabs, Beverly, MA), and autoclaved water. The reaction mix was incubated for 1
hour at 37 °C. The reaction was stopped by heating at 95 °C for 2 mins and icing the reaction

46

for 1 min. Molar excess (177 pmol) of anti-sense oligonucleotide was added to the stopped
kinase reaction mixture and the mixture was heated to 56 °C for 10 mins. Next, the reaction
was placed at room temperature for 60 mins and at 4 °C overnight for hybridization.
Removal of free y- P-ATP and quantification of radioactivity
ProbeQuant G-50 Micro Column (GE Healthcare) was prepared by vortexing the
resin in the column and spinning the column at 3000 rpm for 1 min. The flow-through was
discarded and 26 ul of hybridization mixture was applied to the resin. The column was
placed into a new eppendorf tube and spun at 3000 rpm for 2 mins. The abasic DNA was
collected in the eppendorf tube which was transferred to another eppendorf tube. 1 ul of
purified abasic DNA was measured of its activity in counts per min (c.p.m.) using Packard
1600 TR liquid scintillation analyzer. The abasic DNA was diluted to give a final count of
100,000 c.p.m./ul.
2.1.7 SDS-PAGE analysis on recombinant human APE1
12 % SDS-PAGE resolving gel mix (12 % acrylamide/0.32 % bisacrylamide, 0.3 M
Tris-HCl pH 8.8, 0.1 % SDS) was made by mixing 2.5 mL of 30% acrylamide/0.8 %
bisacrylamide, 1.5 mL of 4 x Lower Gel Buffer (Table 6), and 2 mL of autoclaved water.
5mL of this mixture was mixed with 10 ul of 20 % ammonium persulfate (APS), and 3 ul of
TEMED (Sigma) and cast into the assembled gel apparatus (Biorad, Hercules, CA). The
surface of the resolving gel was evened out by pipetting in 1 mL isopropanol. The resolving
gel was left to solidify for 30 minutes. After, the isoproanol was removed and the surface of
the resolving gel dried. Similarly, 5 % SDS-PAGE stacking gel mix (5 % acrylamide/0.13 %
bisacrylamide, 0.1 M Tris-HCl pH 6.9, 0.1 % SDS) was made by mixing 0.32 mL of 30%

47

acrylamide/0.8 % bisacrylamide, 0.5 mL of 4 x Upper Gel Buffer, and 1.18 mL of autoclaved
water.
Table 6: Composition of reagents for SDS-PAGE.
Reagents
10 x Running Buffer
4 x Lower Gel Buffer
4 x Upper Gel Buffer
2 x SDS Loading Dye
30 % acrylamide/0.8
% bisacrylamide
20 % ammonium
persulfate

Composition
0.19 M Tris-HCl, 1 % SDS, 1.9 M Glycine
1.15 M Tris-HCl pH 8.8, 0.4 % SDS
0.38 M Tris-HCl pH 6.9, 0.4 % SDS
0.1 M Tris-HCl pH 8.8, 0.4 % SDS, 20 % Glycerol, 0.00004 %
Bromophenol Blue, 10 % p-mercaptoethanol
14.6 g acrylamide, 0.4 g bisacrylamide in 50 mL autoclaved water,
Filter sterilized
2 g ammonium persulfate in 10 mL of autoclaved water

2mL of this mixture was mixed with 5 ul of 20 % ammonium persulfate, and 2 pi of
TEMED (Sigma) and cast on top of the resolving gel already solidified and dried from the
previous step. Immediately, the comb was inserted on to the stacking gel and the gel was left
to solidify for 20 mins.
All protein samples were prepared as follows. 10 ul of protein sample (40 ug) was
mixed with 10 ul of 2 x SDS Loading Dye and boiled in water bath for 4 mins. The 20 pi
mix was spun briefly and subsequently loaded onto the 12 % SDS-PAGE gel. As a control,
10 ul of Full Range Rainbow Molecular Weight Markers (Amersham Biosciences,
Buckinghamshire, UK) was also loaded. The gel was run at 120 V with 1 x Running Buffer
(20 mM Tris-HCl, 0.1 % SDS, 0.2 M Glycine) prepared from its 10 x stock in MilliQ water.
The gel was run for 60 mins or until the Bromophenol Blue dye front had reached the bottom
of the gel.

48

Table 7: SDS-PAGE gel staining & destaining reagents.
Reagents
Composition
Coomassie staining solution
50 % methanol, 10 % acetic acid, 0.1 % (=
O.lg) Coomassie Brilliant Blue R-250
Destaining solution
50 % methanol, 10 % acetic acid
The gel was removed from the apparatus and stained in 50 mL of Coomassie staining
solution (Table 7) overnight with gentle shaking at room temperature. The gel was then
destained using Destaining solution for 1 hr. The gel was visualized and imaged under white
light transillumination using Chemilmager™ System (Alpha Innotech Corporation, San
Leandro, CA).
2.1.8 Endoribonuclease assays using 5'-radiolabeled RNA
The buffer system used in the standard endoribonuclease assay contained Tris-Cl. A
typical 20 ul reaction condition contained the following ingredients: 10 mM Tris-Cl pH 8
and 2 mM DTT, 25 nM of 5'-32P-lableled substrate (100,000 c.p.m.), and varying
concentration of APE1. The reaction is carried out at 37 °C for varying amount of time. Once
the reaction is complete, the reaction mixture is stopped by adding 40 ul of Stopping Dye.
Ten microliters of stopped mixture was loaded onto 8% polyacrylamide (29:1, acrylamide:
N,N'=metheylenebisacrylamide) gel containing 7M urea (Invitrogen) and ran at 20 raA with
0.5x TBE buffer (0.45 M Tris-HCl/Boric acid (Sigma), 0.01 M EDTA pH 8.3) for 60 mins.
The gel was transferred onto a filter paper and dried on a gel dryer (LABCONO, Kansas
City, MO) for 45 mins at 80 °C and exposed overnight to a Cyclone Storage Phosphor Screen
System (Packard, Meriden, CT). On the following day, the gel image was developed using
Cyclone Phospholmager and visualized by the OptiQuant software (Hewlett Packard, Palo
Alto, CA).

49

2.1.9 Abasic DNA endonuclease assays using 5'-radiolabeled abasic DNA
The established protocol for APE1 abasic DNA endonuclease assay was used with
modifications (Mantha et al. 2008). A typical reaction mixture of 15 ul contained 80,000
c.p.m. (0.1 pmoles) of abasic DNA, 25 pg (0.7 fmoles) of APE1, 50 mM Tris-HCl (pH 8), 50
mM KC1, 1 mM DTT, 0.1 m M EDTA, 2 mM MgCl2 and 100 ug/mL bovine serum albumin.
The reaction was carried out at 37 °C for 3 mins. Next, 30 ul of the Stopping Dye was added
to the reaction mixture and 9 ul of the sample was loaded onto an 8% polyacrylamide (29:1,
acrylamide: N,N'=metheylenebisacrylamide) gel containing 7M urea (Invitrogen) and ran at
20 raA with 0.5x TBE buffer (0.45 M Tris-HCl/Boric acid (Sigma), 0.01 M EDTA pH 8.3)
for 40 mins or until the bromophenol blue dye was separated from the xylene cyanole FF by
3 cm. The remainder of the procedure was identical to that described above (section 2.1.8).
2.1.10 Dialysis of APE1
For experiments designed to assess the effects of divalent metal ions, recombinant
APE1 produced from Dr. Mitra's laboratory was dialyzed additionally in a metal-free buffer.
50 ul aliquot of APE1 was treated with 2 ul of 6 M Guanidine-HCl with 20% pmercaptoethanol to allow for denaturation of the protein. A total of 10 ul of the 6M
Guanidine-HCl with 20% p-mercaptoethanol was added in sequential steps by adding 2 ul at
a time. Once the protein was denatured, the aliquots were transferred to the Slide-A-Lyzer
Mini Dialysis Units (Pierce, Rockford, IL) and put into the beaker holding 500 mL of
dialysis buffer (20 mM Tris-HCl pH 8, 300 mM NaCl, 1 mM DTT, 0.1 mM EDTA, and 50%
glycerol) with gentle stirring. The dialysis was carried out in the cold room for a total of 5 hrs
with one change of dialysis buffer after the first 2.5 hrs. The protein was quantified later with
ND-1000 NanoDrop V.3.1.0 Protein A280 method (Wilmington, Delaware).

50

2.1.11 Determination of the nature of 3'-end of the RNA cleavage product
To determine whether APE1 generates RNA products with a 3'phosphate or a
3'hydroxyl group, we adopted the method described by Schoenberg and co-workers
(Chernokalskaya et al. 1997) which uses snake venom exonuclease's (Sigma) ability to
digest only products bearing 3'hydroxyl. The c-myc CRD RNA nts 1705-1792 was used as a
substrate. RNase Tl generates products with 3'phosphates, and these can not be processed by
the exonuclease. In contrast, SI nuclease generates decay products with 3'hydroxyls and
these can be degraded upon treatment with the exonuclease. Hence, APE1 treated RNA
decay products can also be treated the exonuclease to determine the characteristic of their 3'
ends,
i) RNase Tl
A typical 20 jal reaction contained -250,000 c.p.m. of 5'-radiolabeled RNA, 1 U of
RNase Tl in RNase Tl Buffer (25 mM Tris-HCl pH 7.2, 2.5 mM MgCl2, 25 mM KC1). The
reaction was carried out 37 °C for 5 mins.
ii) SI Nuclease
A typical 20 ul reaction contained -250,000 c.p.m. of 5'-radiolabeled RNA, 10 U of
SI Nuclease in lx SI Nuclease Buffer (Invitrogen Life Technologies, Carlsbad, CA). The
reaction was carried out at 37 °C for 5 mins.
iii) APE1
A typical 20 ul reaction contained -250,000 c.p.m. of 5'-radiolabeled RNA, 0.3 ug of
APE1 in APE1 reaction mixture including 10 mM Tris-HCl pH 7.4, 2 mM DTT, and 2 mM
magnesium acetate. The reaction was carried out 37 °C for 5 mins.

51

iv) RNA Product Extraction and Resuspension
After reactions, 130 ul of DEPC treated water was added to raise the volume to 200
(il, followed by a standard phenol/chloroform extraction and ethanol precipitation as
described in section 2.1.2. Exceptions from this section were the usage of pH 4.3 phenol
(Sigma) and the resuspension of the extracted RNA in 12 ul of snake venom phosphatase
reaction buffer (100 mM Tris-HCl pH 8.7, 100 mM NaCl, and 14 mM MgCl2).
v) Snake Venom Phosphatase Reaction
To 4 ul of resuspended RNA, 1 ul of 3.2xl0"5 U/ul of snake venom phosphatase
(Sigma, St. Louis, MO) was added and the reaction was carried out at 25 °C for 10 mins. The
reaction was stopped by adding 5 ul of the Stopping Dye and 3 ul of the stopped mixture was
loaded on to an 8% polyacrylamide (29:1, acrylamide: N,N'=metheylenebisacrylamide) gel
containing 7M urea (Invitrogen) and ran at 20 raA with 0.5x TBE buffer (0.45 M TrisHCl/Boric acid (Sigma), 0.01 M EDTA pH 8.3) for 1 hour or until the bromophenol blue dye
was near the bottom of the gel. The gel was transferred to a filter paper and dried on a gel
dryer (LABCONO, Kansas City, MO) for 45 mins at 80 °C and exposed overnight to a
Cyclone Storage Phosphor Screen System (Packard, Meriden, CT). On the following day, the
gel image was developed using Cyclone Phospholmager and visualized by the OptiQuant
software (Hewlett Packard, Palo Alto, CA).
2.1.12 Electrophoretic mobility shift assay (EMSA)
To assess the RNA binding abilities of APE1 and its mutants, electrophoretic mobility shift
assay (EMSA) was used.

52

i) Generation of internally radiolabeled RNA
For our EMSA, internal radiolabeling of the c-myc 1705-1886 CRD RNA was done through
in vitro transcription reaction. A typical reaction mixture of 20 ul was made up by mixing 4
ul of 5x Transcription Buffer (Promega, Wyoming, MI), 1 ul of 40 U/ul RNasin (Promega,
Madison, WI), 2 ul of 100 mM DTT (Promega, Madison, WI), 1 uls of 10 mM ATP, CTP,
and GTP, and 2.5 ul of 100 uM of UTP, 2 ul of 19 U/ul T7 RNA Polymerase (Promega,
Madison, WI), 0.5 to 1 ug of linear DNA template, and 3 ul of Uridine 5'-triphosphate (a32

P) (PerkinElmer, Boston, MA) and adjusted to the volume with diethylpyrocarbonate

(DEPC) (Sigma) treated Milli-Q-ddH20. The reaction was incubated at 37°C for I hr. Next,
10 ul of 1 U/ul RNase-free RQ1 DNase (Promega, Madison, WI) was added and incubated at
37 °C for 10 mins and the reaction was stopped with 10 ul of Stopping Dye (Table 3).
Subsequently, the sample was loaded onto an 8% polyacrylamide (29:1, acrylamide:
N,N'=metheylenebisacrylamide) gel containing 7M urea (Invitrogen) and ran at 25 mA with
0.5x TBE buffer (Table 3) for 1.5 hour. The gel was exposed onto a Cyclone Storage
Phosphor Screen System (Packard, Meriden, CT) and the image was developed using
Cyclone Phospholmager and visualized by the OptiQuant software (Hewlett Packard, Palo
Alto, CA). The polyacrylamide gel containing the radiolabeled transcript was sliced out and
put into a tube containing 400 ul of RNA Elution Buffer (Table 3) and 200 ul of phenol (pH
= 4.3, Sigma). The elution lasted 5 hrs at 45 °C, followed by standard phenol/chloroform
extraction and ethanol precipitation (refer to section 2.1.3). The RNA was resuspended in 40
ul of DEPC treated water to give a final count of -600,000 c.p.m./ul. The RNA was
quantified by diluting 5 ul of RNA samples in 800 ul of DEPC water in a spectrophotometer
cuvette (VWR International). The sample was loaded onto Ultraspec 1000 (Biochrom)

53

UV/visible spectrophotometer and measured of its absorbance at wavelength 260 nm. The
concentration of the RNA was calculated by the equation, Abs260nm x 6.4 = sample in ug/ul.
A typical concentration was -0.75 ug/ul.
ii) Electrophoretic Mobility Shift Assay (EMSA)
For one EMSA reaction, 6.625 ul of Binding Buffer Cocktail (BBC) was prepared by mixing
4 ul of Binding Buffer (Table 8), 1 ul of 10 mg/ml of yeast tRNA, 1 ul of 10 mg/ml of BSA
(Sigma), 0.125 ul of 40 U/ul RNasin (Promega, Madison, WI), and volume adjusted with
DEPC water. A typical 20 ul EMSA reaction contained 6.625 ul of BBC, 30 nM of internally
radiolabeled RNA and different amounts of APE 1. The mixture was incubated at 35°C for 15
mins. 2 ul of EMSA Loading Dye was added and the mixture was loaded on to pre-running
8% native polyacrylamide (29:1, acrylamide: N,N'=metheylenebisacrylamide) gel. The gel
was run for 2 hrs at 25 mA or until the xylene phenol FF dye reached the bottom of the gel.
The subsequent process was identical to that of the endoribonuclease assay in section 2.1.8.
Table 8: The composition of reagents used in EMSA experiments.
Reagents
Binding Buffer
EMSA Loading Dye

Composition
50 mM Tris-Cl pH 7.4, 12.5 mM EDTA, 10 mM DTT, 25 %
glycerol, 0.01 % Triton-X
250 mM Tris-Cl pH 7.4, 0.07 % Bromophenol blue, 0.07% Xylene
cyanol FF

54

2.2 Results and Discussion
2.2.1 Generation of unlabeled and 5'-radiolabeled RNA
For certain plasmids, it was more convenient to perform PCR amplification to
generate DNA templates for use in in vitro transcription rather than linearizing plasmids. So,
it was decided that the PCR fragment should be used as a DNA template for in vitro
transcription of c-myc 1705-1792 CRD RNA. Primer sets were designed (Table 5) and used
to amplify the DNA templates coding for c-myc-1705-1792 CRD RNA identical in sequence
to the one inserted in pUC19-c-myc-1705-1792. Figure 6A shows the in vitro transcribed
unlabeled RNA substrate generated using the PCR templates. Both lanes 2 and 3 confirmed
the presence of intact RNA substrate before and after DNase I treatment for eliminating DNA
template. Both lanes 2 and 3 confirmed the presence of intact RNA substrate before and after
DNase I treatment for eliminating DNA template. The RNA substrate showed multiple bands
when migrated on a non-denaturing agarose gel. These bands may have been produced due to
the inherent formation of RNA secondary structures under the non-denaturing condition. It
should be noted that the non-denaturing agarose gel analysis is a rapid method to quickly
visualize whether the transcription has occurred, but it does not provide an accurate
estimation on the size of the RNA substrate. To estimate the size of the RNA, the RNA
should be run on a 1% formaldehyde gel. However, later it was confirmed that when c-myc1705-1792 CRD RNA, were phosphorylated with y-32P-ATP, only a single band of high
activity was observed in an 8% denaturing polyacrylamide gel (Figure 6B; left gel),
confirming that the in vitro transcribed RNA is indeed the right product. The size of the
radiolabeled RNA was estimated using the internal standards, xylene cyanol and
bromophenol blue.

55

Commercially obtained RNA oligonucleotides (Table 4) were also radiolabeled in a
similar way and were subjected to purification by excision and elution. Oligo IB is shown in
the right panel of Figure 6B. Other substrates (Oligo IA and dOligo LA.) are not shown.

/ /

A

B
/
— Xylene
cyanol FF
(160 nt)

^
bp
88 n t -

— Bramophenol
Blue(45nt)
200100-

•*•« mm - c-myc CRD RNA (88 nt)

1 2

3

17 nt -

1

2

••
1

2

Figure 6: Agarose and polyacrylamide gel analyses showing the in vitro transcribed
RNA substrate and the generation of 5'-radiolabeled RNA substrates. (A) 2% nondenaturing agarose gel showing in vitro transcribed c-myc 1705-1792 CRD RNA (lane 3)
after DNase I treatment for 30 mins. (B) 8% polyacrylamide denaturing gel showing the
products of 5'-end radiolabeling phosphorylation reactions using y- P-ATP and T4
polynucleotide kinase and unlabeled c-myc-1702-1792 CRD RNA (left). On the right, 8%
polyacrylamide denaturing gel shows the products of Oligo LB RNA substrate 5'radiolabeled in an identical way as c-myc-1702-1792 CRD RNA.

2.2.2 Identification of essential residues for RNA incision activity of APE1
Recombinant APE1, APE1 structural mutants that have mutation on one of the active
site residues, and APE1 population variants obtained from Dr. David Wilson's laboratory
were all run on an SDS-PAGE gel to confirm their purity (Figure 7). All samples showed one
band at an estimated molecular weight of 35.5 kDa with the exception of D308A mutant,
which showed two bands of approximate size of 35.5 kDa and 33.5 kDa. It was known
previously that an N-terminal truncated form of APE1 was observed (Chattopadhyay et al.
2006) and that APE1 can undergo hydrolysis during boiling of the protein samples, losing its

56

N-terminal region of 32-33 amino acids. In fact, previous WT APE1 produced in Dr. Mitra
Sankar's laboratory also showed two bands when ran on an SDS-PAGE gel (Barnes et al.
2009).
A

B
UDaX

#
66
45
36
29
24
20

V5

|1~I':

W

^Sr W SST S"
V ' <y
q? Or Or

illf» *»«««» ,«««i» ^ S * *s^|^#|i ; ||^^fep
s||fc

14.2

6.5

1

2

4

5

6

7

8

9

10

1

2

4

5

6

Figure 7: SDS-PAGE analysis of 1 ug of APE1 WT and mutants. The proteins were
loaded on to each well and the gel was stained with Coomassie Blue. In lane 1 of each gel,
Low Molecular Weight Marker was loaded. (A) APE1 WT (lane 2) and structural (active
site) mutants (lanes 3 to 10). (B) APE1 human population variants (lanes 2 to 7).
The activities of these structural mutants were tested against their typical, 18 nt abasic
DNA, substrate (Figure 8A). When the reduction of their activities were calculated by
measuring the ratio of density of the product over those of the substrate plus product, D70A
and H309S showed 7-fold and 158-fold reduction in their activities, respectively (Table 9).
Compared to the previous reports, our results showed that N68A and D308A exhibited less
reduction in their activities (~5 8-fold and ~ 1.4-fold, respectively compared to the WT)
whereas F266A and D283N showed greater reduction (~30-fold and ~573-fold, respectively
compared to the WT).
This may be due to the different assay conditions that we used in our laboratory as
compared to those used by the other groups. There are disparities in the reported literature in
regards to fold reductions observed in DNA endonuclease activity. For example, E96A
mutant (not tested here) showed variations in reduction of its DNA endonuclease activity up

57

to 100 fold when tested in different enzyme amounts that fell in the range between 100 and
1000 pg (Erzberger et al. 1998; Chou and Cheng 2003). The amount of F266A and D283N
used in our assay were both 25 pg in a reaction volume of 15 ul, which were suggested to
give a linear reaction rate in abasic DNA incision (Mantha et al. 2008). In other studies,
however, the same mutants were used in higher amounts (Erzberger et al. 1998; Masuda et
al. 1998) that the reduction in their activities was reported to be less. Nevertheless,
reductions observed in APE1 structural mutants relative to one another correlated to the
suggested importance of these residues and the roles they play in the DNA endonuclease
activity. These details are discussed further in conjunction with the results on RNA incision
activity.
3'

^

£>•

^ r^^

<?

** <,•*
J? <y^
* <o- ^ ^™

<?x

v

^

<y

<P

^
Substrate (18 nt)
I - Incision Product (9 nt)

5'
C-G
T -A
C-G
A- T
G-C
C-G
A- T
T -A
F-A
G-C
I -A
G-C
C-G
C-G
A- T
C-G
T -A
G-C

S

3'
G
T
G
C
C
A
C
T
G
3'

AP-DNA
l«nt

5'
Product
<tat

Figure 8: Abasic DNA endonuclease activity of WT APE1 and APE1 structural
mutants. (A) 0.14 nM of APE1 (lanes 2 to 10) was tested against 25 nM of 5'- y-32Pradiolabeled abasic DNA in a total reaction volume of 15 ul for 3 mins at 37°C. (B) Structure
and sequence of the abasic DNA (AP-DNA) and its incised product.

58

Table 9: Summary of fold reduction in WT APE1 and structural mutants.
Mutants

N68A
D70A
E96A
Y171F
D210N
N212A
F266A
D283N

600 - 100000

Not tested

Fold reduction
in RNA incision
(Figure 10A)
Abolished
55.6
Not tested

5000
25000
Not detectable

Abolished
Abolished
Not tested

17.7
8312.6
Not tested

6
10

30.4
572.6

Abolished
4.0

Reference

Fold reduction in
Fold reduction in
abasic DNA incision abasic DNA incision
in literatures
(Figure 8A)
Nguyen et al. 2000
600
58.7
Nguyen et al. 2000
Reduction
6.7
Erzberger et al. 1999;
Chou and Cheng 2003
Erzberger et al. 1999
Erzberger et al. 1999
Rothwell and Hickson
1996
Erzberger et al. 1998
Hadi et al. 2000

D308A Erzberger et al. 1999;
Barzilay et al. 1995
H309N Chou and Cheng 2003

5-25

1.4

9.0

100000

Not tested

Nguyen et al. 2000

Not tested

158.3

Reduction
(Barnes et al. 2009)
6.0

H309S

When the structural mutants were challenged with c-myc CRD RNA, most of the
mutants, except D283N, showed reduced or abrogation of RNA incision activity at both
concentrations tested (Figure 9A and B). Compared to the WT, the D283N mutant showed
similar or greater activity, and though reduced, D308A, seemed to retain some RNA-cleaving
activity.
A

^ ^ ^ X < ^ 5 ^ ^ a7lxM

^>#>Vt4<^V" M„M
^1768CA, 1771CA
-J1773UA, 1775CA

1768CA, 1771CA
1773UA, 1775CA

-1757UA
-1751UA
-1747UA
-1742CA

-1730UG
-1727CA

1730UG
1727CA

1

2

3

4

5

6

7

8

9

10

1

2

3

4

5

6

7

8

9

10

Figure 9: Endoribonuclease activity of APE1 and its structural mutants on c-myc-17051792 CRD RNA. (A) 0.7uM of APE1 (lanes 2 to 10) was tested against 25 nM of 5'- y-32Pradiolabeled c-myc-1705-1792 CRD RNA in a total reaction volume of 20 ul for 25 mins at
37°C. (B) Same condition as A with 1.4 uM of APE1 mutants.
59

The test of these mutants showed that the active site for cleaving abasic DNA is also
important in cleaving RNA, despite a subtle difference for D283 which does not seem to be
required for giving RNA incision activity. The same set of enzymes were also challenged
with the 17nt RNA substrate, Oligo IB, which can form a single stem and a hairpin structure
(Figure 10B) and has an identical sequence to the c-myc CRD region of 1741-1757 (Figure
15B). Oligo IB is composed entirely of deoxyribose-phosphate backbone except at base
position 6, where a Uridine base is bonded to a ribose sugar. This was to confirm the
reduction in activity seen with testing c-myc 1705-1792 CRD RNA (88nt). By measuring the
ratio of density of the products (5 nt & 6 nt) over those of the substrate plus product, we
could quantify the degree of reduction observed in these mutants. It was found that the
reduced activities of structural mutants were also observed with Oligo IB (Table 9 and Figure
10A), confirming that these residues are indeed critical for giving RNA incision activity of
APE1.
B

</> f f J* #* <f <f <f ^

"

pM

SubstrateU7nO

T

G

T

A

A
fU

T

rU

G-- C

G

G

G - C

G

G
A

A— I

A

A — T

A

C -

C

G

Product ( 6 n t l
>roduct(Snt)

A

c
5'

Oligo IB

Product

Product

17nt

6nt

Sat

Figure 10: Endoribonuclease activity of APE1 and its structural mutants on Oligo IB
RNA. (A) 1.4 uM of APE1 mutants (lanes 2 to 10) were tested against 25 nM of 5'- y-32Pradiolabeled Oligo IB RNA in a total reaction volume of 20 ul for 25 mins at 37°C. (B)
Secondary structure of Oligo IB predicted through Mfold showing its APE1 RNA incision
product (6nt) and 3' phosphodiesterase product (5nt).

60

One noteworthy observation was that there were two cleavage products of similar
intensities were generated by D70A, Y171F, D210N, and D309S mutants (lanes 4, 5, 6, and
10, Figure 10A). The identity of the 6nt product seemed to have resulted from the RNA
incision activity of APE1 cutting in between rUpA dinucleotides where as the 5nt product
may have resulted from DNA 3' exonuclease activity of APE1 (Chen et al. 1991). The strong
5nt band produced by the WT (lane 2) compared to the weaker bands produced by the
structural mutants showed that the WT can remove rU that hangs on to the recessed strand of
the DNA structure. This suggests that WT APE1 possesses 3' exoribonuclease activity that is
reduced in structural mutants. However, further investigation would be required to confirm
this proposal. The finding is also in line with the suggestion that APE1 shares a common
active site for its DNA endonuclease and 3'-5' exonuclease activities (Chou and Cheng
2003).
In general, human APE1 has been found to be a non- or poorly-processive 3'-5'
exonuclease (Wilson 2003). 3'-5' exonuclease activity of APE1 is about 0.03% of the abasic
DNA incision activity (Wilson et al. 1995) and shows similar affinities (Km) towards 3'
termini, but varying degrees of turnover (kcat) in decreasing order for 3' recessed DNA, 1 nt
gapped DNA, nicked DNA, and 2nt gapped DNA (Wilson 2003). The WT 3'-exonuclease
activity is not detectable on blunt end of duplex DNA (Suh et al. 1997; Hadi et al. 2002) or
on ssDNA (Wilson 2003). Also, human APE1 has been shown to remove 3' mismatched
nucleotides at least 50 times more efficiently than those that are matched correctly. The
mismatched nucleotide was removed more efficiently from nicked DNA than from recessed
DNA (Chou and Cheng 2002). Thus, we speculate that after rUpA is cleaved by the WT
APE1 RNA incision activity, the remaining rU must have been further removed by its 3'-5'

61

exoribonuclease activity of APE1, but this linear deletion could not have proceeded beyond
the removal of rU.
The degree of reduction for each structural mutant in abasic DNA and RNA incision
activity (Table 9) corresponds to the residue's importance in both of the reactions. For
example, reductions seen in D70 and D308 do not seem to be severe as these two residues are
predicted to coordinate metal ions (Erzberger and Wilson 1999) during the abasic DNA
incision that are thought to orient the substrate, stabilize transition state, and facilitate the
dissociation of product. In abasic DNA incision, their mutations are thought to have
relatively minor effects compared to other mutations in other residues with only ~6.7-fold
and ~ 1.4-fold reduction in their DNA endonuclease activities and ~56-fold and ~9-fold
reduction in RNA incision activities (as in Figure 10) for D70A and D308A, respectively. It
can be speculated that the absence of one of these residues may be relieved by the other and
the mutant APE1 would be able to retain its DNA endonuclease activity.
In molecular mapping through crystal structure, N68, E96, Y171, and D283 are
hydrogen bonded to their neighboring residues and create an environment for abasic DNA
incision to occur (Figure 11).

62

ASf 70

Asnljtt

h
\ lm

174

#•--"•

Figure 11: Molecular model of APE1 active site (Rothwell et al. 2000). Blue spheres
represent water molecules and orange sphere represents the bound metal ion. The thick
dotted line represents a hydrogen bond between Asp 210 and Asn 68. The thin dotted lines
represent various hydrogen bonds between side chains of active site residues.

N68 is thought to hydrogen bond to D210 and H309, which supposedly creates a
favorable environment for catalysis (Nguyen et al. 2000) and its mutation to alanine would
logically be detrimental for RNA incision activity (Table 9). E96, originally considered to
serve in the coordination of a metal ion (Beernink et al. 2001; Chou and Cheng 2003), is also
thought to be critical in establishing the appropriate active site environment via a hydrogenbonding network involving Y171 (Erzberger and Wilson 1999). The RNA cleaving activity
of E96A has been found to be non detectable (Barnes et al. 2009) and a mutation of Y171F
also gives reduction in RNA incision activity, confirming that the hydroxyl group on the
phenyl ring is an important component in formation of hydrogen bonds to E96. One
noteworthy observation is that D283N mutant exhibited RNA incision activity and this was a

63

significant contrast to what has been known about D283. Also, D283N showed to some
degree of 3' exonuclease activity and indicated that this residue is not as critical as in RNA
incision as in 3' exonuclease activity of APE1. Its role is to orient H309 through hydrogen
bonding and act to stabilize its positive charge (Hadi et al. 2000), and the presence Oligo LB
incision activity indicated that the significance of D283-H309 dyad in RNA cleavage is not
so critical compared to cleaving abasic DNA. However, H309S mutant showed less RNA
cleaving activity than D283, and this was more evident against c-myc 1705-1792 CRD RNA,
which indicated that the role of H309 is more important than D283 in RNA catalysis.
Similarly, a previous report from Dr. Lee's laboratory also showed a significant reduction in
endoribonuclease activity with H309N, suggesting a critical role for H309 in RNA incision
(Barnes et al. 2009). Its role in abasic DNA cleavage is also thought to be critical in aligning
and polarizing the scissile bond linking the normal base and the abasic sugar (Rothwell et al.
2000), and our observation thus far also pointed to similar role in RNA incision activity.
Initially, the role of D210 was described as the proton donor of the incised leaving
group (Erzberger and Wilson 1999). However, later it was proposed that the reaction
mechanism for abasic DNA incision relied on an elevated pKa for D210 in that the
carboxylate on side chain can be protonated to generate the nucleophilic hydroxyl group
from a water molecule (Mol et al. 2000). Indeed, substitution of this residue strongly
diminishes abasic DNA incision activity, while stabilizing the binding of APE1 to its
substrates (Rothwell et al. 2000). In RNA incision, this trend was also observed (Figures 8
and 9), suggesting that an additional role of D210 may also be in the generation of the
nucleophile. Residue F266 has been found to be a part of a hydrophobic pocket comprised of
F266, W280 and L282 that accommodates abasic sugar moiety during the abasic DNA

64

incision (Mol et al. 2000, Hadi et al. 2002). In RNA incision, we observed significant
reductions both on Oligo IB (Figure 10) and c-myc 1705-1792 CRD RNA (Figure 9), which
suggested that F266 may serve a similar role in recognizing RNA bases. F266A showed
greater reduction in RNA incision with its activity being not detectable compared to cleaving
abasic DNA (30-fold reduction). This suggested that the alteration in this residue or the
hydrophobic pocket were less tolerant for RNA catalysis than for abasic DNA. In addition,
F266A mutant was shown to possess an enhanced DNA 3'-exonuclease activity compared to
the WT on an array of DNA substrates (including single-stranded DNA, and duplex DNAs
containing either a nick, single nucleotide gap, blunt end or 3' flap) (Hadi et al. 2002) and
this fact may account for the appearance of DNA 3'-exonucleolytic product in lane 7 (Figure
10). The difficulty for detecting 3' exonucleolytic products of F266A on abasic duplex DNA
(lane 7; Figure 8A) may be due to its preference for 3' termini in the nicked structure or
within a partial duplex DNA over fully complementary or blunt-ended DNA substrates with
matched 3' ends (Hadi et al. 2002). Hence, F266A showed 3' exonuclease activity on a
single stranded DNA-RNA hybrid (Oligo IB) that can form a dynamic hairpin structure, but
this activity was not observable in an abasic DNA duplex.
2.2.3 Assessing the RNA-binding abilities of APE1 structural mutants
We tested the RNA-binding abilities of a few selected structural mutants using
electrophoretic mobility shift assay (EMSA). Preliminary analysis showed that the WT (lanes
2 to 4) and E96A (lanes 6 to 8) both exhibited similar binding abilities to c-myc 1705-1886
CRD RNA, while H309N (lanes 10 to 12) showed lesser binding (Figure 12A). It also was
found that N68A (lanes 6 to 8), Y171F (lanes 10 to 12), and D210N (lanes 14 to 16)
displayed comparable binding abilities to that of the WT (lanes 2 to 4) to the same substrate

65

(Figure 12B). Interestingly, D283N at lower amounts (250 ng and 500 ng), showed lesser
binding, but at higher amount (1000 ng), showed stronger binding compared to the WT
(lanes 2 to 4) (Figure 12B).

A

B
WT

1 2 3 4 5 6 7 8 9 10 11 12

12

3 4 5

N68A

6 7 8

Y171F

D210N

D283N

9 10 11 12 13 1415 16 1718 19 20

Figure 12: RNA-binding abilities of APE1 and its structural mutants on c-myc 17051886 CRD RNA. Increasing amount of APE1 or structural mutants were tested against 50
nM of 32P-internally radiolabeled c-myc 1705-1886 CRD RNA in a total binding volume of
20 ul for 15 mins at 35°C in a standard electrophoretic mobility shift assay (EMSA). (A)
EMS A gel showing APE1 WT (lanes 2 to 4), E96A (lanes 6 to 8), and H309N (lanes 10 to
12). (B) EMSA gel showing APE1 WT (lanes 2 to 4), N68A (lanes 6 to 8), Y171F (lanes 10
to 12), D210N (lanes 14 to 16), and D283N (lanes 18 to 20).

The role of some of these residues in RNA-binding may not be as critical as our results have
shown. The discussion on specific residues and how they may or may not be involved in the
RNA-binding as well as abasic DNA-binding are further discussed in section 5.4.
2.2.4 RNA incision activity of the population variants of APE1
When these selected population variants were tested on an abasic DNA, they
displayed a similar reduction for all variants previously tested (Hadi et al. 2000), except for
E126D (lane 7) which showed an activity close to that of the WT (Figure 13). The
quantification of the appearance of product over a total density of substrate and product

66

showed that these variants possessed comparable DNA endonuclease activities relative to the
WT (100%) as the following: Q51H (-101%), I64V (-103%), L104R (-74%), E126D
(-102%), D148E (-101%), R237A (-0.1%), G241R (-101%), and G306A (-97%).

/ * fsSSSfSf

^

<+>

•
•
1

2

— Substrate (18 nt)

MB

m *» ** m m-m*

** **

3

10

4

5

6

7

8

9

— Incision Product (9 nt)

12

Figure 13: Abasic DNA endonuclease activity of population variants of APE1. (A) 0.14
nN of 5'- y-32P-radiolabeled abasic DNA
nM of APE
APE11 (lanes 2 to 10) was tested against 25 nM
in a total reaction volume of 15 |jl for 3 mins at 37°C.
It is possible that the increased cleavage activity observed in E126D may have been
due to our increased amount of enzymes in the reaction condition (75 pg vs 2.5 pg) compared
to the previous report (Hadi et al. 2000). Apart from this, we have discovered that Q51H and
I64V showed similar activities as the WT, agreeing with the fact that these residues lie
outside of the DNA endonuclease domain as viewed by X-ray crystallography (Mol et al.
2000). Overall, this assay verified that these population variants possessed abasic DNA
endonuclease activities and could be tested against RNA substrates.

67

Table 10: Summary of abasic DNA and RNA incision activities of human population
variants of APE1. ND indicates non-detectable level of activity. *The ratios of density of the
products (5 nt & 6 nt) over those of the substrate plus product were measured using OptiQuant
software (Hewlett Packard, Palo Alto, CA) to quantify activities
APE1
Proteins
WT
Q51H
I64V
L104R
E126D
D148E
R237A
G241R
G306A

Activity on
Abasic DNA (%)
(Hadi et al. 2000)
100
Not Tested
Not Tested
56.71+26.80
60.43 ±11.16
94.36 + 6.20
35.71+9.68
108.73 + 5.31
107.22 + 17.13

Activity on
Abasic DNA (%)
(Figure
100
101
103
74
102
101
0.1
101
97

Activity on
CRDRNA(%)
(Figure
100
23.1
15.6
60.0
41.4
23.3
6.0
24.3
20.4

Activity on
Oligo IB (%)
(Figure
100
12.5
Abolished
0.9
8.0
6.8
Abolished
12.2
5.5

In endoribonuclease assay against c-myc-1705-1792 CRD RNA as the substrate, we
found that L104R (lane 5 in Figure 14) and E126D (lane 6 in Figure 14) retained RNA
incision activity (-60% and -40%, respectively), but their cleavage sites were altered. In
contrast to the report on abasic DNA incision (Hadi et al. 2000), we found D148E (lane 7),
R237A (lane 8), G241R (lane 9), and G306A (lane 10) to have reduced RNA cleaving
activity (Table 10). This was also confirmed with Oligo IB RNA in lanes 6 to 9 after
quantification of the activities that were reduced down to ~5.5%-12% compared to the WT
(Figure 15 and Table 10). These results showed that these variants have reduced RNA
incision activity and warrants further investigation on their role in human diseases like cancer
and ALS (Hadi et al. 2002).

68

• I Mm j ^ , j i t JS^ j ^ ^fc_ j u t ^
^

^» P r I H P r

pi

TB£PP Mr

^^

HP

]

1766CA, 1768CA,
1771CA, 1773UA,
1775CA

17S7UA
- 1751UA
1747UA
1742CA

1730UG
I - 1727CA
1

2

3

4

5

6

7

8

9

10

Figure 14: Endoribonuclease activity of human population variants APE1 on c-myc1705-1792 CRD RNA. (A) 1.4uM of APE1 (lanes 2 to 10) were tested against 25 nM of 5'y-32P-radiolabeled c-myc-1705-1792 CRD RNA in a total reaction volume of 20 ul for 25
mins at 37°C.
For the first time, we discovered that the activities of Q51H and I64V were also
notably reduced (-23% and -15.6%, respectively) against c-myc RNA (lane 3 and 4, Figure
14) especially those sites of 1730 UG, 1747UA, 1751UA, 1757UA, and 1773 UA, but not
1727 CA, 1742 CA, 1766 CA, 1768 CA, 1771 CA, and 1775 CA sites. The reduction of
RNA incision activity at specific region was confirmed through testing the variants on Oligo
IB RNA designed to mimic the c-myc CRD region of 1741-1757 which also provides an
analogous site of cleavage for APE1 at 1747 UA site. Indeed, the RNA incision activities of
Q51H and 164V were reduced down to -0-13% (Table 10). The reason behind such reduction
for specific dinucleotides sequence is unknown and warrants further investigation on
identifying regions of APE1 that give cleavage specificity. Nonetheless, this showed that
Q51H and I64V have reduced RNA incision activity and warranted further characterization
of their properties.

69

F ff /

V

^

^

f^

0"

©=> 4
— Substrate (17 nl)

I

I—Product (6 nt)
••^jjji. I - Product (S tit)
1

2

3

4

5

6

7

8

9

10

Figure 15: Endoribonuclease activity of human population variants of APE1 on Oligo
IB RNA. 1.4 |JM of APE 1 (lanes 2 to 10) were tested against 25 nM of 5'- y-32P-radiolabeled
Oligo IB RNA in a total reaction volume of 20 jal for 25 mins at 37°C.
In addition, we have mapped the altered cleavage sites of L104R and E126D on a
sequencing gel (Figure 16). For reference, an alkaline hydrolysis ladder was generated as
shown (lane 1). Numbering on the left indicates guanosine residue sites cleaved by RNase Tl
under denaturing conditions (lane 3), whereas numbering on the right indicates the sites
cleaved by APE1 WT (lane 4) and its variants (lanes 5 and 6). It was determined that L104R
produced cleavages at 1727 CA, 1731 GA, 1742 CA, 1747 UA, 1749 GU, 1751 UA, 1768
CA, 1770 GC, 1771 CA, 1773 UA, 1775 CA. It is noteworthy that L104R was capable of
cleaving after guanine bases (1731 GA, 1749 GU, and 1770 GC). For E126D, the cleavage
pattern was suggestive of exoribonucleolytic decays occurring at regions of 1715-1734,
1741-1751, and 1758-1775. Notably, E126D variant was capable of prominently cleaving
1731 GA, 1748 AG, and 1749 GU. Such observation suggested that these residues are not
only important in giving endonuclease activity (abasic DNA and RNA incision), but also in
giving the specificity of cleavage of APE1.

70

B

v^.^>

^ " . ^ / ^
^

N<^ <F *

V

V

1768 CA

?20^,.^*X™G ...

c

*S^U
u

g-

s ... c
*

G . . • C
1790
C • • • G
, /
A C C A G - A U O C
y C C A A G C

/

1710

WT

1 IIMK
Ct26b

1 2

3

4

5

Stwjag-4

'«—•
•—*

«««k

<!
'.=--..
,

6

Figure 16: Mapping of the cleavage sites generated by L104R and E126D variants of
APE1. (A) 1.4 uM of APE1 WT, L104R, and E126D variants (lanes 4 to 6) were tested
against 25 nM of 5'- y-32P-radiolabeled c-myc-1705-1792 CRD RNA in a total reaction
volume of 20 j_il for 25 mins at 37°C. The products were resolved on a 12% polyacrylamide
gel as described in section 3.1.2. (B) Secondary structure of c-myc 1705-1792 CRD RNA
and the positions of cleavage sites for WT, L104R, and E126D.

71

2.2.5 Assessing the RNA binding abilities of APE1 population variants
We tested APE1 population variants in an electrophoretic mobility shift assay
(EMSA) on c-myc 1705-1886 CRD RNA. It was found that the WT and the six population
variants all exhibited similar binding abilities (Figure 17). L104R (lanes 6 to 8, Figure 17A)
and E126D (lanes 10 to 12, Figure 17A) were expected to bind to RNA since they showed
RNA-cleaving activities in previous experiments (Figures 14 to 16). Comparable binding
abilities of D148E (lanes 14 to 16, Figure 17A), R237A (lanes 14 to 16, Figure 17B), G241R
(lanes 10 to 12, Figure 17B), and G306A (lanes 6 to 8, Figure 17B) to that of the WT (lanes 2
to 4, Figure 17A-B) indicated that these residues likely affect the catalytic step of RNA
incision and not the RNA-binding process.

A

B
WT

L104R

E126D

D148E

1 2 3 4 5 6 7 8 9 10 1112 13 14 15 16

WT

G306A

G241R

R237A

1 2 3 4 5 6 7 8 9 1011 12 1314 15 16

Figure 17: RNA-binding abilities of APE1 and its population variants on c-myc 17051886 CRD RNA. Increasing amount of APE1 and its population variants were tested against
50 nM of 32P-internally radiolabeled c-myc 1705-1886 CRD RNA in a total binding volume
of 20 ul for 15 mins at 35°C in a standard electrophoretic mobility shift assay (EMSA). (A)
EMSA gel showing APE1 WT (lanes 2 to 4) and L104R (lanes 6 to 8), E126D (lanes 10 to
12), and D148E (lanes 14 to 16). (B) EMSA gel showing APE1 WT (lanes 2 to 4), G306A
(lanes 6 to 8), G241R (lanes 10 to 12), R237A (lanes 14 to 16).

72

Intriguingly, L104R and E126D appeared to retain RNA-cleaving activities despite in
the presence of RNasin in the EMSA binding reaction. This was visualized by lighter
intensities of APE1-RNA complex (lanes 6-8, 10-12) compared to the WT (lanes 2 to 4) and
D148E (lanes 14 to 16) (Figure 17A). Also, the appearance of RNA incision products from
L104R (lanes 6 to 8) and E126D (lanes 10 to 12) were clearly visible below the APE 1-RNA
complex in the same EMSA gel (Figure 18). Hence, in the binding reaction, the digestion of
RNA took place for L104R and E126D, and we could observe the APE 1-RNA complex
shifting, but at the same time, we could also visualize the presence of decay products.
Therefore, our results have revealed not only an alteration in the cleavage pattern of these
variants (Figure 16), but also a potentially novel RNasin-resistant activity (Figure 18). Future
studies should confirm the existence of this activity and characterize the difference in the
strengths of L104R and E126D compared to the WT and other mutants in the presence of
RNasin. If this novel activity is found to be truly originating from L104R and E126D, the
biological impact of their RNasin-resistant activities should also be assessed.
WT

• | l

L104R

| f

E126D

D148E

I f

A t

M it
jst
1

m

2 3 4 5 6 7 8 9 10 11 12 13 14 15 16

Figure 18: RNasin-resistant activity of L104R and E126D on c-myc 1705-1886 CRD
RNA. From the same EMSA gel in Figure 17A, RNA cleavage products were more visible in
samples treated with L104R (lanes 6 to 8) and E126D (lanes 10 to 12) than those treated with
the WT (lanes 2 to 4) and D148E (lanes 14 to 16).
73

2.2.6 Possible mechanism of RNA incision by APE1
To further understand the RNA cleavage mechanism of APE1, we first tested to see
whether APE1 generated RNA products with a 3'phosphate or a 3'hydroxyl group (Figure
19). This was done using the ability of snake venom exonuclease that digests only the
products bearing 3'hydroxyl group (Chernokalskaya et al. 1997). The distinct product
generated by RNase Tl (asterisk in lane 2) remained visible after digestion with exonuclease
(lane 3). This is consistent with fact that RNase Tl generates products with 3'phosphates. In
contrast, SI nuclease generated decay products (marked with asterisks in lane 5) were
completely degraded upon treatment with the exonuclease (lane 6). This is consistent with
the notion that SI nuclease generates products with 3'hydroxyls. Fig. 16 shows that the
distinct decay products generated by APE1 (marked with asterisks in lane 9) were still visible
upon treatment with the exonuclease (lane 8). However, we also observed the disappearance
of products in the range of 1766 to 1775 after treatment with the exonuclease (Lane 8),
raising the possibility that perhaps these bands were produced by the 3'-5' exonuclease
activity of APE1. What mechanism by which these products were generated is up for
speculation and will require further experimentation. At the least, however, we found that
APE1 can generate endoribonucleolytic products with 3'phosphates.
Determining the 3'end of RNA products generated by APE1 provides insights into
the role of APE1 in mRNA metabolism. APE1 produces those RNA products with 3'
phosphates and 5' hydroxyls (Figure 19), and such products are expected to be less efficient
for clearance by 3'-5' exosome (Chernokalskaya et al. 1997). Nevertheless, APE1 activity
may still be a part of mRNA decay machinery. APE1 generated products can be further
degraded by any one or combinations of the following paths. Activity of exosome itself

74

which possesses both endoribonucleolytic and exonucleolytic activities can clear these
transcripts despite at a reduced rate for 3' phosphate bearing single stranded RNAs
(Schaeffer et al. 2009). XRN1, a 5'-3' exoribonuclease, can degrade these RNAs from their
5' ends after decapping takes place (Garneau et al. 2007).
.- + + . , , .
- - - - , -f- ,

1 2

- - - - KNas«T1
+ •+ „ » . s i Nucleate
- - - + + ReAPEI
, + - 4- » 3*-5* Exonuclease

3 4 5 6

7 8

9

Figure 19: APE1 generates RNA products with 3'phosphates. 25 nM 5'-labeled c-myc
CRD RNA was digested with 1 U RNase Tl (lanes 2 and 3) for 1 min at 37°C, or with 10 U
SI nuclease (lanes 5 and 6) or 8.5 uM purified APE1 (lanes 8 and 9) for 5 min at 37°C under
the standard endonuclease assay. The products were then incubated with 3 x 10"5 U snake
venom exonuclease for 10 min at 25°C (lanes 3, 6, and 8), and electrophoresed on a 8%
polyacrylamide/urea gel. Asterisks indicate the decay of products generated by RNase Tl
(lane 2), SI nuclease (lane 5), or ReAPEI (lane 9).
Since APE1 is ubiquitously expressed and its localization reported to differ perhaps
due to its ability to serve multiple functions in different cell types (Duguid et al. 1995), its
RNA cleaving activity has the potential to play an important role in mRNA decay. On the
other hand, RNA-cleaving activity of APE1 may also have biological significance during
mRNA processing. This speculation is in line with the finding that APE1 serves as a nuclear
enzyme. Also, APE1 has been found to interact with a global regulator of mRNA splicing,

75

hetergenous nuclear ribonucleoprotein L (hnRNP L) which associates with transcribed RNAs
in the nucleus (Kuninger et al. 2002).

A

B
dOligoIA

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Figure 20: RNA incision activity of APE1 requires 2'-OH on the sugar moiety. (A) 1.4
uM of APE1 were tested against 25 nM of 5'- y-32P-radiolabeled dOligo IA (lanes 1-6) or
Oligo IA (lanes 7-12) RNA in a total reaction volume of 20 ul for varying time points at
37°C. (B) Secondary structure of dOligo IA and Oligo IA predicted through Mfold.
In understanding APE1 mechanism of catalysis, producing an RNA product with 3'
phosphate end is different from a typical abasic DNA cleavage product with a 3' hydroxyl
group (Doetsch and Cunningham 1990; Demple and Harrison 1994; Friedberg et al. 1995).
The active site residues in APE1 abasic DNA cleavage could be involved differently in RNA
cleaving, perhaps providing an environment that promotes a general acid-base RNA catalysis
that utilizes 2'-OH group on the sugar moiety as exemplified by RNase A (Raines 1998). If
this were not the case, residues involved in abasic DNA cleavage may still be able to
generate a hydroxyl nucleophile from a water molecule. Next, this nucleophile can attack the
phosphodiester bond from the side opposite to where it is proposed to attack during the
abasic DNA cleavage, generating an 0 5 ' leaving group and a final product with 3'
phosphate.
To test whether 2'-OH group is important in RNA catalysis of APE1, APE1 was
tested on a modified form of Oligo IA, which has a deoxyribose sugar at base position 10

76

(Table 4, Figure 20). APE1 can not cleave the dUA bond in dOligo IA (lanes 2-6) whereas
the rUA bond in Oligo IA was cleaved over time (lanes 8-12). This result suggested that
indeed during APE1 RNA cleavage, 2'-OH is essentially involved in the catalysis, and that
APE1 utilizes residues from the active site, and not H2O molecules to carry out the catalysis.
2.2.7 The effect of divalent metal ions in APE1 RNA catalysis
The abasic DNA incision activity of APE1 has been shown to be divalent metal ion
dependent (Masuda et al. 1998; Erzberger and Wilson 1999; Beernink et al. 2001; Oezguen
et al. 2007). We therefore determined the effect of divalent metal ions on RNA cleavage by
APE1. For the purpose of the following experiments, protein samples were denatured and
refolded in metal ion-free buffer, as described in section 2.1.10, to ensure complete removal
of metal ions. Figure 21 shows that APE1 has the ability to cleave c-myc RNA in the absence
of added divalent metal ions (lanes 1-3). In the assumption that the denaturation/refolding
step had successfully removed all of the divalent metal ions from the APE1 buffer, this result
helped us to conclude that APE1 is able to cleave RNA without the presence of divalent
metal ions. APE1 exhibited activity in the presence of Mg2+ (lanes 4-5), Ca2+ (lanes 6-7), and
Mn2+ (lanes 12-13). In contrast, at 2 mM, Zn2+ (lanes 8-9), Ni2+ (lanes 10-11), Cu2+ (lanes
14-15), and Co2+ (lanes 16-17), all had inhibitory effect on RNA-cleaving activity of APE1.
As in DNA cleavage reactions, RNA cleaving activity of APE1 is present and/or enhanced
by the addition of metal ions, particularly Mg2+. Whether the presence of metal ion enhances
the affinity of APE1 to abasic DNA or RNA is currently unkown and this question needs to
be further explored. However, our observation is in line with reports on APE1 requiring Mg2+
concentration of around 0.1mM-0.5mM for its 3'-5' DNA exonuclease activity and 2 mM-10
mM for its abasic DNA incision activity (Chou and Cheng 2003; Wilson 2005). APE1 has

77

also been shown to exert EDTA-resistant activity towards (Erzberger and Wilson 1999)
acyclic DNA substrates that are considered to have better flexibility than abasic DNA and
even shown a weak incision activity for a regular abasic DNA in the presence of EDTA
(Masuda et al. 1998).

4ST

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1 2 3 4 5 0 7 X 9 W 11 H 13 14 15 1ft 17

Figure 21: Effect of divalent metal ions on RNA-cleaving activity of APE1. 12.5 nM of
5'-labeled c-myc-1705-1792 CRD RNA were treated with purified APE1 for 5 min at 37°C
in the presence (lanes 4-17) or absence (lanes 1-3) of 2raMvarious divalent metal ions as
indicated under the standard endoribonuclease assay. Samples were run on 8%
polyarylamide/7 M urea gel. The background reaction buffere contained (10raMTris-Cl pH
8 and 2 mM DTT).
Single stranded RNA structures that form secondary structures (e.g. hairpin loops) are
particularly more flexible than duplex RNAs or abasic duplex DNAs, which may explain the
RNA cleaving activity observed even at 0 mM Mg2+ (lane 1 and 2). The effect of Mg2+ was
further assessed by testing a range of concentrations (0 mM - 200 mM) (Figure 22). Lane 2
shows that APE1 has the ability to cleave RNA at 0 mM Mg2+ and at 5 mM (lane 4), APE1
showed the most activity in cleaving RNA. The increasing amount of Mg2+ gradually
decreased the activity of APE1 where at 100 mM and 200 mM the activity was clearly
inhibited (lanes 8 and 9). Interestingly, we also observed a change in cleavage pattern at 20

78

raM and 50 mM concentrations (lanes 6 and 7), suggesting metal ions may either alter the
RNA secondary structure or the RNA binding affinity of APE1. This was clarified later by
EMSA experiment using WT APE1 against c-myc-1705-1886 CRD RNA under increasing
concentrations of Mg2+ (Figure 22B). This showed that the binding affinity of APE1 was not
dramatically altered by the increasing divalent metal ion. Hence, our results indicated that the
likely reason behind altered cleavage pattern is due to the alterations in RNA secondary
structure.

A

B
1.4 MM APEI

1

2

3

4

5

6

1

8

9

1

OmM

2mM

2

4

3

5mM 10 mM 20 mM 50mM100mM

S 6

7

8

Mg2*

9 10 11 12 13 14

Figure 22: Differing concentrations of magnesium in the reaction mixture affects the
specificity and the incision activity of APE1. (A) 1.4 uM of APE1 (lanes 2 - 9) were tested
against 25 nM of 5'- y-32P-radiolabeled c-myc-1705-1792 CRD RNA in a total reaction
volume of 20 ul for 25 mins at 37°C. Each lane contains differing magnesium concentration
as indicated. (B) APE1 WT was tested against 50 nM of 32P-internally radiolabeled c-myc
1705-1886 CRD RNA in a total binding volume of 20 ul for 15 mins at 35°C in a standard
electrophoretic mobility shift assay (EMSA) under increasing concentrations of Mg2+.
2.2.8 The effect of RNase inhibitors on APE1 activity
A set of experiments showed that APE1 RNA cleaving activity can be inhibited by
the treatments of typical RNase inhibitors: Diethyl pyrocarbonate (DEPC) and Ribonuclease
Inhibitor (RNasin). It is thought that DEPC is an effective inhibitor that reacts with enzymes
containing -NH, -SH, or -OH groups in their active sites. Hence, APE1 that has active site
residues consisting of Asp, Glu, His, and Asn will likely be inhibited when DEPC reacts with

79

any of these groups. Indeed, titration of DEPC showed that at 1.43 mM of DEPC was enough
to clearly inhibit the activity of APE1 (Figure 23A). Electrostatic forces between
ribonuclease and RNasin are known to allow their binding and sterically hinder substrateribonuclease interaction (Kobe and Deisenhofer 1996). This may also be the case with APE1.
It was found that 1 unit of RNasin was able to inhibit 84 nM of APE1 activity (Figure 23B),
which suggested that RNasin may also interact with APE1 and sterically prevent RNA
substrate from interacting with residues of the active site.

A

B
<P »* s ? s> N- v-

[DEPC]

No RNasin
V

1U RNasin

V

* ^<K <>\ >VA*\ ^ * * N < V \ b \ * V V ' |ReAPEl|
in nM

1 2 3 4 5 6

12

3 4 5 6 7 8 9 10 11 12 13 14 1516 17 18 19 20

Figure 23: RNA incision activity of APE1 is inhibited by DEPC and RNase Inhibitor
(RNasin). (A) 0.2 uM of APE1 (lanes 2 - 6) were tested against 12.5 nM of 5'- y-32Pradiolabeled c-myc-1705-1792 CRD RNA in the presence of increasing concentration of
DEPC dissolved in 0.1% ethanol. (B) Specified concentration of APE1 was tested in the
absence or presence of 1 unit of RNasin against 12.5 nM of substrate RNA.
Future studies can focus on revealing whether this inhibition of RNA cleaving
activity is due to the alterations in RNA binding or in incison step. Testing of APE1 binding
to the RNA substrate in the presence of DEPC or RNasin may give clearer distinction
between the inhibitions via alteration of binding or incision activity of APE 1.

80

CHAPTER 3
Establishing the RNA structure and sequence cleaved by APE1
This chapter presents the methods and discusses the results of the experiments used to
determine the RNA structure and sequences that are preferentially cleaved by APE1. The
purified recombinant human APE1 has thus far been tested only against c-myc CRD RNA
(Barnes et al. 2009). To determine if APE1 can also cleave other RNAs, to further confirm
the sequence and structural cleavage specificity of APE1, and to initiate studies into future
possible use of the enzyme as an anti-viral and anti-cancer agent, we challenged APE1 with
three RNA components, Spike, Orflb and Orf3 RNAs, which are derived from proteins
important for the life cycle of the SARS-corona virus (Tan et al. 2005) and two miRNA
substrates, miR-lOb and miR-21 which are known to be associated with cancer (Ma and
Weinberg 2008; Gramantieri et al. 2008). Spike or S protein mediates binding of the virus to
host receptors and in membrane fusion (Tan et al. 2005). Orflb is one of the replicases
essential for replication of the virus, and Orf3 is a protein with endocytotic properties and
which does not have homology to other coronaviruses and is expressed during SARS-CoV
infection (Tan et al. 2005).
Sequences of human-pri-miR-lOb (pri-miR-lOb) andhuman-pri-miR-21 (pri-miR-21)
were also tested. Notably, miR-lOb has been reported to be associated with metastasis of
breast tumors (Ma et al. 2007), of which its expression has been thought to be induced after
the activation of an oncogenic transcription factor, twist. Similarly,

miR-21 has been

reported to reduce the expression of a tumor suppressor and increases the abilities for cellular
invasion (Asangani et al. 2007). In addition, over expression of miR-21 has been reported in

81

various cancer samples including, cholangiocarcinoma (Selaru et al. 2009), esophageal
squamous cell carcinoma (Hiyoshi et al. 2009), breast carcinoma (Qian et al. 2008; Yan et al.
2008), hepatocarcinoma, (Connolly et al. 2008), and cervical cancer (Lui et al. 2007).

3.1 Methodology
3.1.1 Plasmid linearization and PCR generation of linear templates
To prepare linear DNA templates for in vitro generation of RNA substrates, the
pUC19 plasmids containing cDNAs of Spike, Orflb, and Orf3 were all linearized. The
sequences of these cDNAs are shown in Table 11. These plasmids also contained T7
promoter on the 5' side of cDNA sequences. These plasmids were linearized with the
restriction enzyme EcoRI (Invitrogen). A 35 ul digestion reaction contained 2 to 8 ug of
plasmid, 10 ul of EcoRI (10 U/ul), lx reaction buffer (ReACT 3 from Invitrogen), and made
up to volume with autoclaved water. The digestion was done at 37 °C for 3 hrs in a water
bath. After digestion, 1 ul of linearized samples and 1 ul of original plasmids as well as 1 ul
of 1 kb DNA ladder (Invitrogen) were loaded onto 1% agarose (Invitrogen) gel to check for
completion of the reaction. To prepare the gel, 0.5 g of agarose was added to a 50 ml of 0.5x
TBE buffer (Table 3) and heated by microwave to completely dissolve the residues. The gel
mix was cooled and 15 ul of 10 mg/ml ethidium bromide was added to the mix before
casting on the gel apparatus.

82

Table 11: List of RNA substrates generated and their cDNA sequences from plasmid
RNA Substrate
Spike
Orflb
Orf3
hsa-miR-lOb

hsa-miR-21
5'-GG-pre-miR-10b
5'-GG-pre-miR-21

cDNA Sequence
5' -CTAAACGAAC ATGTTTATTTTCTTATTATTTCTTACT
CTCACTAGTGGTAGTGACCTTGACCGGTGCA-3'
5' -AGGATGTAAACTGAC ATAGCTCGCGTCTC AGTTTC A
AGGAACTTTTAGTGTATGCTGCTGATCCAGCTAT-3'
5' -CGAACTTATGGATTTGTTTATGAGATTTTTTACTCT
TAGATCAATTACTGCACAGCCAGTAAAAATTGACAATG
CTTCTC-3'
5' -CC AGAGGTTGTAACGTTGTCTATATATACCCTGT
AGAACCGAATTTGTGTGGTATCCGTATAGTCACAGATT
CGATTCTAGGGGAATATATGGTCGATGCAAAAACTT
CA-3'
5' -TGTCGGGTAGCTTATC AGACTGATGTTGACTGTT
GAATCTCATGGCAACACCAGTCGATGGGCTGTCTGA
CA-3'
5' -TACCCTGTAGAACCGAATTTGTGTGGTATCCGTA
TAGTCACAGATTCGATTCTAGGGGAAT-3'
5' -TAGCTTATC AGACTGATGTTGACTGTTGAATCTC
ATGGCAACACCAGTCGATGGGCTGT-3'

The gel ran for 1.5 hour at 85 V in 0.5x TBE buffer and visualized under UVtransillumination using Chemilmager™ System (Alpha Innotech Corporation, San Leandro,
CA). Next, 2 ul of Proteinase K (20mg/ml, Roche Diagnostics Inc, Mannheim, Germany)
were added into each linearized samples and incubated at 37 °C for 30 mins. The reaction
mixtures were made up to 200 ul with autoclaved water for standard phenol/chloroform
extraction and ethanol precipitation. For miR substrates, hsa-miR-lOb, hsa-miR-21, 5'-GGpre-miR-lOb and 5'-GG-pre-miR-21, pMIF-cGFP-Zeo-hsa-miR-lOb (System Biosciences,
Mountain View, CA) and pSIF-Neo-Ires-GFP-hsa-miR-21 (gift from Dr. Yong Li,
University of Louisville) were used to generate linear cDNA templates by PCR that included
the T7 promoter and the miR cDNA sequence. The sequences of the primer used are shown
in Table 12.

83

Table 12: List of primers and their sequences used in the generation of cDNA templates by
PCR for in-vitro RNA transcription.
cDNA Template
hsa-miR-lOb Forward
hsa-miR-lOb Reverse
hsa-miR-21 Forward
hsa-miR-21 Reverse
5'-GG-pre-miR10b Forward
5'-GG-pre-miR10b Reverse
5'-GG-pre-miR21 Forward
5'-GG-pre-miR21 Reverse

Sequence
5 '-GGATCCTAATACGACTCACTATAG
GCC AGAGGTTGTAACGTTG-3'
5' -TGAAGTTTTTGC ATCGAC-3'
5 '-GGATCCTAATACGACTCACTATAG
GTGTCGGGTAGCTTATC A-3'
5' -TGTCAGAC AGCCC ATCGA-3'
5 '-GGATCCTAATACGACTCACTATAG
GTACCCTGTAGAACCGAAT-3'
5' - ATTCCCCTAG AATCGAAT-3'
5' -GGATCCTAATACGACTCACTATAG
GTAGCTTATCAGACTGATG-3'
5' -AC AGCCC ATCGACTGGTG-3'

3.1.2 Utilization of enzyme probes for secondary structure determination
The 5'-radiolabeled RNA transcript (50,000 c.p.m.) was partially digested with
RNase Tl (Roche Diagnostics Inc, Mannheim, Germany) which cleaves 3' to single-stranded
guanosines, RNase T2 (Invitrogen Life Technologies, Carlsbad, CA) which preferentially
cleaves 3' to single-stranded adenosines, but also 3' to the other three nucleotides, RNase A
(Ambion, Inc., Austin, Texas) which cleaves 3' to single-stranded uridines and cytidines, and
RNase VI (Ambion, Inc., Austin, Texas) which cleaves 3' to paired nucleotides or stacked
bases (Ehresmann et al. 1987).
i) Alkali digestion
A partial alkaline digestion of the 5'-radiolabeled transcript was carried out to
generate an RNA ladder used to identify the cleavage sites. A typical reaction mixture of 10
ul included 5'-radiolabeled transcript (100,000 c.p.m.), 1 ul of lOx Alkali buffer, and DEPC
treated water. The reaction was carried out at 95 °C for 3 mins, iced for 1 min, followed by
the addition of 5 ul Stopping Dye. Subsequently, 3 ul of the stopped mixture was loaded onto

84

a 12% polyacrylamide (29:1, acrylamide : N,N'= metheylenebisacrylamide) gel containing
7M urea (Invitrogen).
For the following enzymatic reactions, the 5'-radiolabeled transcripts were initially
denatured at 55 °C for 5 mins and re-natured at 4 °C for 5 mins in their respective enzymatic
buffers in the absence of their RNases. This procedure had ensured that all RNAs were
correctly folded into their native secondary structures.
ii) RNase Tl
RNase Tl digestion was carried out under RNA native and denaturing conditions in
order to confirm the strandedness of the guanosine residues. Denaturing condition was
applied by setting up a reaction mixture of 10 ul including 50,000 c.p.m. radiolabeled
transcript, 3 U of RNase Tl, lx sequencing buffer (Ambion, Inc., Austin, Texas) consisting
of 20 mM sodium citrate pH 5, 7 M Urea, and ImM EDTA, and DEPC treated water. The
reaction was carried out under room temperature for 5 mins and stopped by adding 20 ul of
the Stopping Dye. Subsequently, 6 p.1 of the stopped mixture was loaded onto the 12%
polyacrylamide gel described above. Under native conditions, a total reaction volume of 10
ul included 50,000 c.p.m. radiolabeled transcript, 3 U of RNase Tl, 2.5 mM Tris-HCl pH
7.2, 2.5 mM MgCi2, 25 mM KC1, and DEPC treated water. The reaction was carried out
under room temperature for 5 mins and stopped by adding 20 ul of the Stopping Dye (Table
3). Subsequently, 6 ul of the stopped mixture was loaded onto the 12% polyacrylamide gel
described above.
ii) RNases T2, VI. A
For RNase T2, RNase VI, and RNase A digestions, reaction mixtures of 10 ul was
set up that included 50,000 c.p.m. radiolabeled transcript, one of the RNases (1U of RNase

85

T2, 1 mU of RNase VI, 0.2 ng of RNase A), lx structure buffer consisting of 10 mM Tris pH
7, 100 mM KC1, 10 mM MgCl2 (Ambion, Inc., Austin, Texas), and DEPC treated water. The
mixtures were incubated under room temperature for 5 mins and stopped by adding 20 ul of
the Stopping Dye (Table 3). Subsequently, 6 ul of the stopped mixture was loaded onto the
12% polyacrylamide gel described above. For checking the RNA integrity, undigested
transcripts were also loaded and ran on the polyacrylamide gel.
iii) Separation by PAGE
The polyacrylamide gel was ran at 20 mA with 0.5x TBE buffer (0.45 M TrisHCl/Boric acid (Sigma), 0.01 M EDTA pH 8.3 for 45 mins initially, followed by the loading
of another batch of the same stopped reaction mixtures to empty wells and ran under the
same condition for another 1 hour to resolve the cleavage sites near the 5' end. The gel was
transferred to a filter paper and dried on a gel dryer (LABCONO, Kansas City, MO) for 45
mins at 80 °C and exposed overnight to a Cyclone Storage Phosphor Screen System (Packard,
Meriden, CT). On the following day, the gel image was developed using Cyclone
Phosphorlmager and visualized by the OptiQuant software (Hewlett Packard, Palo Alto, CA).
3.1.3 In vitro effects of APE1 RNase activity on DICER product formation
APE 1/DICER reaction
The reaction was carried out in two steps. First, a lOul reaction mix containing 1 uM
of recombinant APE1 and 25 nM of 5'-32P-labeled in vitro transcribed pre-miRNA analog
(50,000 c.p.m/ul) in 8.8 mM Tris-HCl pH 7.4 and 2 mM magnesium acetate was incubated at
37°C for 30 mins. Recombinant APE1 purified and prepared in 20 mM Tris-HCl pH 8, 300
mM NaCl, 1 mM DTT, 0.1 mM EDTA, 50% glycerol was used in this reaction. Second, 10
ul mixture containing 0.4 x DICER Reaction Buffer (Ambion), 0.25 U DICER (Ambion),

86

and DEPC treated water was added to the first reaction and incubated at 37°C for 30 mins.
Addition of 0.4x DICER Reaction Buffer (120 mM NaCl, 8 mM HEPES, 2 mM MgCl2, and
20 mM Tris-HCl pH 9.0) gives the final reaction mix a salt and pH condition similar to that
of the lx DICER Reaction Buffer which allows for optimal DICER activity. DICER
(Ambion) was prepared to 0.25U/ul from 1 U/ul stock in storage buffer (100 mM NaCl, 20
mM Tris (pH7.5), ImM EDTA, and 50% Glycerol) before combining with the DICER
Reaction Buffer. 0.4x DICER Reaction Buffer was prepared from 5X DICER Reaction
Buffer (Ambion, 1.5 M NaCl, 100 mM HEPES, 25 mM MgCl2, and 250 mM Tris-HCl pH
9.0). The reaction was stopped by adding 40 ul of the Stopping Dye (Table 3) and 6 ul was
loaded onto the 8% polyacrylamide gel.
Alkaline Ladder reaction
A 10 ul reaction mix containing 75 nM (100,000 c.p.m) of 5'- P-labeled in vitro
transcribed pre-miRNA analog, 500 mM sodium bicarbonate pH 9.2 and 10 mM EDTA was
incubated at 95 °C for 3 mins and on ice for 1 min before adding 5 ul of Stopping Dye (Table
3). 3 ul of the reaction mixture was loaded onto the 8% polyacrylamide gel
RNase Tl reaction
A 10ul reaction mix containing 25 nM (50,000 c.p.m) of 5'- P-labeled in vitro
transcribed pre-miRNA analog, 0.6x Sequence Buffer (9751G, Ambion), and 0.3 U/ul RNase
Tl was incubated at room temperature for 5 mins and stopped by adding 5 ul of Stopping
Dye (Table 3). 3 ul of the reaction mixture was loaded onto the 8% polyacrylamide gel

87

3.2 Results and discussion
3.2.1 Generation of unlabeled and 5'-radiolabeled RNA
The linearization of pUC19 vectors containing cDNA sequences for Spike and Orf3
with a restriction enzyme, EcoRI, generated discrete single bands sized between 2000bp and
3000bp in lane 3 and 5, respectively (Figure 24A). Undigested plasmids in lane 2 and 4
migrated differently compared to their linearized counterparts. EcoRI digestion result for
Orflb is not shown, however, it can be confirmed that a proper digestion had took place
which resulted in generation of proper sized RNA of -79 nt (Figure 24C). Once the digestion
had taken place, the linearized plasmids were subjected to in vitro transcription to generate
unlabeled RNA substrates (Figure 24B-C). Lanes 3 and 5 in Figure 24B and Lane 3 in Figure
24C confirmed the elimination of DNA templates after transcription. Similarly to c-myc
1705-1792 CRD RNA (Figure 6A), the non-denaturing agarose gel showed multiple bands of
substrate RNAs due to their inherent abilities to form secondary structures. For the two
substrates (5'GG-pre-miR10b and 5'GG-pre-miR21) used for assessing APE1 interference in
DICER activity, their DNA template was PCR amplified that included their T7 promoter and
their cDNA sequences (Table 12). Once we obtained linear PCR fragments, RNAs were in
vitro transcribed following the procedure described in 2.1.4 (Figure 24D).

88

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pUC19-Orflb

Orflb (79 nt)

1 2

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200100-

miR-10b(63nt)
miR-21(61nt)
12

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5

Figure 24: Agarose gel analysis showing the digestion of plasmids used for in vitro
transcription and the generation of unlabeled RNA substrates. (A) 1% agarose gel
showing the undigested plasmids pUC19-Spike and pUC19-Orf3 (lanes 2 and 4) and digested
plasmids (lanes 3 and 5) after treatment with 10 U of EcoRI. (B) Non-denaturing 2% agarose
gel showing the in vitro transcribed RNAs of Spike (lane 3) and Orf3 (lane 5) after DNase I
treatment for 30 mins. (C) Orf lb (lane 3). (D) hsa-miR-21 and hsa-miR-lOb (lane 3 and 5).
3.2.2 APE1 cleaves RNA components of SARS-corona virus at specific sites
The secondary structure of Spike nts 21481-21548 was also predicted using M-fold
program (Zuker 2003), and the RNase structure probing data were then used as a constraint
with the structure generated by Mfold to obtain a structure that best fit the probing data
(Figure 25). In using the 68 nt Spike RNA as substrate, following are the key observations.

89

J

•S\

B

m
G21546- *
G21544 G21543 G21539 -

.

*

G21533 G2X531 G21528G21527G2I525-

RNase T] (DM)
RNase Tl (N)
RNase 12
RNase VI
RNase A
ReAPEl

- 2 1 5 1 4 UA

.21490 CA

-.«,::=

• * —

,—
> •

*~~ e

—

< 1 —

^

21504 UA
21505 AU
21507 UA
21508 AU
1
1

- 2 1 5 0 8 AU
- 2 1 5 0 7 UA

*:§t
A .- U

— 21492 UG

,• ,-21496 UA

A -£&*C

"O'/uj'uli /c Ju\u;

- 2 1 4 9 6 UA

C

G21493 •

Weak

-•—

21490

\

— 21505 AU
-21504 V A

Strong

21500

1H

s VI)
^

G,A

', At, U

0$ +1

•§:A&G-K-*
2

3 4 5 6 7 8

"*>•
C

l y i i G , , , tUfkGtA

U

> \

,TTC "' .U ...»
21540'

I VW
IMC- U

G- C

G21487 -

21514 UA
1
!

'
/

y'\ W

G C

- 2 1 4 9 2 UG
- 2 1 4 9 0 CA

„,_n
! °

2 51

' *••'

/ C

. A \Vs

' ' I VC

21530

Figure 25: Ribonuclease secondary structure probing of Spike RNA. (A) In vitro
transcribed RNA corresponding to the indicated nucleotides was 5'-end-labelled with 32P and
refolded prior to probing with the indicated RNases. For reference, an alkaline hydrolysis
ladder was generated as shown. Numbering on the left indicates guanosine residue sites
cleaved by RNase Tl under denaturing conditions, whereas numbering on the right indicates
some of the sites cleaved by APE1. (B) The RNA secondary structure model shows locations
of bases accessible for enzyme probes. The structure was obtained by using the probing data
in the top panel as constraint with the Mfold program (Zuker 2003). Fully-filled symbols
represent strong cleavage while moderately-filled symbols represent moderate to weak
cleavage. D denotes cleavage by RNase Tl under denaturing conditions, and N denotes
cleavage by RNase Tl under native conditions.

90

'"

"21520

(i) APE1 cleaves strongly at 21490CA, 21504UA, and 21507UA, (ii) APE1 can cut weakly
in between UA at stem region as exemplified by weak cleavage at 21496UA, (iii) APE1 can
cut weakly in 21492UG and 21514UA, (iv) APE1 can cut weakly at AU at weak stem region
as exemplified by cleavages at 21505AU and 21508AU.
The secondary structure of Orflb RNA nts 14439-14508 was predicted using M-fold
program (Zuker 2003), and the RNase structure probing data using RNases Tl, T2, A, and
VI, as shown Figure 26 were then used as a constraint with the structure generated by Mfold
to obtain a structure that best fit the probing data (Figure 26B). APE1 cleavage sites on Orflb
RNA were then mapped as shown in Figure 26A (lane 8). Moderate to major cleavage sites
generated by APE1 are 14445UA, 14453CA, 14455UA, 14473CA, 14484UA, and 14489UA,
while weak cleavages are at 14443UG, 14448AC, 14449CU, 14462CG, 144464UC,
14467CA, and 15587UG. Several key observations can be drawn from this result: (i) there is
preference for UA over UG as exemplified by more intense cleavage at 14445UA over
14443UG, (ii) cleavages can occur at CA, UA, and UC at weak stems as exemplified by
14464UC, 14467CA, 14473CA, and 14484UA, (iii) for the first time, APE1 is shown to cut
very weakly at CU, AC and CG at single-stranded region as exemplified by 14448AC,
14449CU, and 14462CG sites. Similarly, the secondary structure of Orf3 RNA nts 2526025339 was predicted using M-fold program (Zuker 2003), and the RNase structure probing
data as shown in the left panel of Figure 27A were then used to generate a structure that best
fit the probing data (Figure 27B). Moderate to strong cleavage by APE1 are at the following
sites: 25266UA, 25278UA, 25290UA, 25312CA, 25319UA, and 25329CA.

91

&

4^.^

€$

RNase 11 (DN)
RNase Tl (N)
RNase T2
RNase VI
RNase A
ReAPEI

B

{BBM^JUU

Strong

Weak

-«_
*—
•

.«*•».

•

*
—

1

.

—
• «
<'

14480
G14498 G14495G14492G14488 G14486-

* balk

m

-14489 U A
-14487 UG
w -14484 UA

G14477 _
G14476-

14473 C A
-14473 CA

G14469-

-14467 C A
G14463-

-14464 UC

G14461 -

-14462 C G

G14457 —
sf

14443 U G
14445 U A

•14455 UA

14448 A C

-14453 CA

G14451 —

-14449 CU
-14448 A C

•
G14444-

% -14445 UA

14449 CU
14453 CA
14440

A G G A U'jS

ASAUA

-14443 U G
4 5 6 7 8

-14450

Figure 26: Ribonuclease secondary structure probing Orflb RNA (A) In vitro
32
transcribed RNA corresponding to the indicated nucleotides was 5'-end-labelled with JZ
P and
refolded prior to probing with the indicated RNases. For reference, an alkaline hydrolysis
ladder was generated as shown. Numbering on the left indicates guanosine residue sites
cleaved by RNase Tl under denaturing conditions, whereas numbering on the right indicates
some of the sites cleaved by APE1. (B) The RNA secondary structure model shows locations
of bases accessible for enzyme probes. The structure was obtained by using the probing data
in the top panel as constraint with the Mfold program (Zuker 2003). Fully-filled symbols
represent strong cleavage while moderately-filled symbols represent moderate to weak
cleavage. D denotes cleavage by RNase Tl under denaturing conditions, and N denotes
cleavage by RNase Tl under native conditions.
T

92

^s*" <*><?> <?><3><3>^
W^

IP w 6 •# ^H'^H j&*

G25333 -

- ;>

G25327 -

m>

-25329 CA
-25326 UG
-25319 UA

G25318G25314 -

-25312 CA
-25310 CA

G25309 -

-25305 UA
-25301 CA
G25298 -

-25296 UA
*m ._

-25291 A C
-25290 UA

•

-25278 UA

G25283 G25281 -

G25275-

G25270 G25269 -

*

-25268 UG
-25267 AU
-25266 UA

12 3 4 5 6 7 8
Figure 27: Ribonuclease secondary structure probing of Orf3 RNA. (A) In vitro
transcribed RNA corresponding to the indicated nucleotides was 5'-end-labelled with 32P and
refolded prior to probing with the indicated RNases. For reference, an alkaline hydrolysis
ladder was generated as shown. Numbering on the left indicates guanosine residue sites
cleaved by RNase Tl under denaturing conditions, whereas numbering on the right indicates
some of the sites cleaved by APE1.

93

B
Strong
^w™,

RNase T l (DN)
RNase Tl (N)
RNase T2
RNase VI
RNase A
ReAPEl

*—

Weak

25312 CA

- 0 =

•

«

<

4

«m

8

<

^

25310 C A 25310""

Ui U

,c

c

/

G C
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25270
25268 UG
25267 A t
25266 UA

*G
_A~H^T
,G U
G U
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A
G

A U

.25319UA
25305 UA -

25320
l-A-

A

• 25278 UA
A
25301 CA # \ U - A y
25326 UG
25290 UA
25280
25291 AC 25296 UA
^ I ^ a s i i i C A

/

u %*y
C G/
C G A A

*i /

l
;;
-ft -II <U *'U ;U'U'UpUi'C IMC < U U V A * G> A I

1
25260

25290

25300

C M A *'A < U

G'ft
c u uc

u c

25330

Figure 27: Ribonuclease secondary structure probing of Orf3 RNA. (B) The RNA
secondary structure model shows locations of bases accessible for enzyme probes. The
structure was obtained by using the probing data in the top panel as constraint with the Mfold
program (Zuker 2003). Fully-filled symbols represent strong cleavage while moderatelyfilled symbols represent moderate to weak cleavage. D denotes cleavage by RNase Tl under
denaturing conditions, and N denotes cleavage by RNase Tl under native conditions.
Weaker but significant cleavage sites are observed at 25291AC, 25296UA, 25301CA,
25305UA, 25310CA, and 25326UG (Figure 27A). The following observations are noted: (i)
cleavage at UA at weak stem as exemplified by 25266UA, 25278UA, and 25319UA, (ii)
APE1 cuts weakly at AC as exemplified by cleavage at 25291AC, (iii) APE1 can cut weakly
at CA at stem region as exemplified with 25301CA.
3.2.3 APE1 cleaves pri-miR-21 and pri-miR-lOb at specific sites
In our effort to further confirm the sequence and structural cleavage specificity of
APE1 and to initiate studies into the possible future use of APE1 in degrading oncogenic
miRNAs, we challenged in vitro transcribed pri-miR-21 and pri-miR-lOb with the purified

94

recombinant human APE1. The secondary structure of pri-miR-lOb was predicted using
Mfold program (Zuker 2003) and as before, RNase probing experiments were conducted
(Figure 28A). The data was used as a constraint with the structure generated by Mfold to
obtain a structure that best fit the probing data (Figure 28B). The major cleavage sites
generated by APE1 are 11UA, 53UA, and 59UA, while weaker sites are observed at 34UA,
50UG, and 61UA. Key observations for this result are: (i) APE1 cleaves UA at weak stems
as shown by cleavages at 34UA and 53UA, (ii) APE1 does not cut in between UA, CA, UG,
and UC at strong stem regions as exemplified by the absence of cleavage at 21UA, 23UA,
25UA, 48UG, 65CA, and 71UC.
Similarly, RNase probing experiments were performed on pri-miR-21 and the
secondary structure generated upon constraining the experimental data (Figure 29A) with
structure predicted by Mfold, is shown on the right panel of Figure 29B. As shown, the major
cleavage sites generated by APE1 are 34UG, 38UC, and 41CA, while weaker cleavage sites
are seen at 13UA, 16CA, 31UG, 33UU, and 40UC. The following are key observations from
this result: (i) APE1 cleaves CA, UG, and UC at weak stems as shown by 34UG, 38UC, and
41CA, (ii) APE1 does not cut in between UG, CA, and UU at strong stems as indicated by
the absence of cleavage at 12UU, 24UG, 26UU, 27UG, 43UG, 46CA, 49CA, 52CA, and
55UC.

95

B

59 UA

C ^ y <* <s> <$• «£ <$> QV
53 UA

*'*

%i

i

**^«* »*

61 UA

50 UG

G99
G92 —
GS4
C83 =
G82 =
C81

G69—
C63—

- 6 1 UA
- 5 9 UA

G58 —

34 UA

- 5 3 UA
- 5 0 UG

G52 —
G51 —
C49 —
C47 —

Strong

^ft

-

G41 —

A
C36 —

-34 UA

G33 —
*^f

RNase T l (DN)
RNase T1(N)
RNase T2
RNase VI
RNase A
RcAPEl

Weak

-^—
•~«

•
«

•

* «

^
^

O —

<53
u

u
u
CIS-

11 UA

:

-* m-

CIS—

*-^ c

c

A

GIO —

G7 —

-11UA
"

^

•

*

'

«

v'

C6 —
C C A

G
A
G

12 3 4 5 6 7 8

Figure 28: Ribonuclease secondary structure probing pre-miR-lOb RNA. (A) In vitro
transcribed RNA corresponding to the indicated nucleotides was 5'-end-labelled with 32P and
refolded prior to probing with the indicated RNases. For reference, an alkaline hydrolysis
ladder was generated as shown. Numbering on the left indicates guanosine residue sites
cleaved by RNase Tl under denaturing conditions, whereas numbering on the right indicates
some of the sites cleaved by APE1. (B) The RNA secondary structure model shows locations
of bases accessible for enzyme probes. The structure was obtained by using the probing data
in the top panel as constraint with the Mfold program (Zuker 2003). Fully-filled symbols
represent strong cleavage while moderately-filled symbols represent moderate to weak
cleavage. D denotes cleavage by RNase Tl under denaturing conditions, and N denotes
cleavage by RNase Tl under native conditions.

96

y^
^

38 UC

B

A T JS> <& «.* c& , 6 -C*'
A.
«" «" d^ «" J»

34 UG

41 CA
40 UC
38 UC
Strong

34 UG
33 UU
- 3 1 UG
16 CA

RNase Tl (DIN)
RNase Tl (N)
RNase T2
RNase VI
RNase A
Re APE 1

Weak

-*—
—

•

-

•

»

• •
—

13 UA

Figure 29: Ribonuclease secondary structure probing pre-miR-21 RNA. (A) In vitro
transcribed RNA corresponding to the indicated nucleotides was 5'-end-labelled with P and
refolded prior to probing with the indicated RNases. For reference, an alkaline hydrolysis
ladder was generated as shown. Numbering on the left indicates guanosine residue sites
cleaved by RNase Tl under denaturing conditions, whereas numbering on the right indicates
some of the sites cleaved by APE1. (B) The RNA secondary structure model shows locations
of bases accessible for enzyme probes. The structure was obtained by using the probing data
in the top panel as constraint with the Mfold program (Zuker 2003). Fully-filled symbols
represent strong cleavage while moderately-filled symbols represent moderate to weak
cleavage. D denotes cleavage by RNase Tl under denaturing conditions, and N denotes
cleavage by RNase Tl under native conditions.

97

- • • ' •

3.2.4 Pre-treatment with APE1 interferes with processing of pre-miR-lOb and pre-miR21 by DICER
To assess if APE1 can potentially interfere with the processing of pre-miRNAs by
DICER, we treated pre-miR-lOb and pre-miR-21 with 1 uM of APE1 prior to incubation
with DICER. Figure 30A shows the cleavage of APE1 on pre-miRlOb at 26UG, 29UA,
35UA, and 37UA. On the other hand, 0.25 Units of DICER alone cleaves pre-miR-lOb to
generate two major products, 25 nt Product 1 and 26 nt Product 2. Pre-treatment of pre-miR10b with 1 uM of APE1 appeared to substantially suppress the ability of DICER to process
pre-miR-lOb as evident from the significant reduction in Products 1 (-6.2 fold) and 2 (-6.7
fold). Figure 31A shows 1 uM of APE1 cleaves pre-miR-21 at 26UG, 28UU, 29UG, 33UC,
and 36CA. 0.25 Units of DICER cleaves pre-miR-21 to generate a single major product of 26
nt size as shown in Figure 31 A. Pre-treatment with APE1 significantly reduce the ability of
DICER to process pre-miR-21 as evident by the significant reduction of the 26 nt major
DICER product by -6.2 fold. Comparison of the two lanes in Figure 31A and B, notably,
APE1 alone and APE1/DICER treated lanes, showed that APE 1-generated products (29 UG,
33 UC, and 36 CA in lane 4 of Figure 31A and in lane 4 of Figure 3IB) and pre-miR-21
substrate did not differ much, but at the same time, significant reduction in major DICER
product (26nt DICER product in lanes 3 and 4 of Figure 3IB) was observed. This indicated
that prior destruction of the miR-21 substrate by APE1 can substantially abrogate the
production of mature miRNA by DICER. The alkaline ladder reaction (lane 1) was also
loaded onto lane 2, RNase Tl reaction, by accident (Figure 31 A).

98

UtAPF.I

Strong
•+

|
|

Weak
<

B

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/

/

,

/

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Mi

W

A —U
A «• U
G — C
C ••• S
A ... U

G3937 UA
35 UA

O G U A C

G34

A A U

S'-GG-hsa-miR-lOb

«m» - 29 UA

G28
G27-

Major
Products
(25nt, 26nt)

26 VG

G25
G23

G17

10 UA
S'oad G

1

2

3

1 2

3

4

G UA C

Dicer Product 1

S'nid GG UA C

Dicer Product 2

Figure 30: Effect of pre-treatment with APE1 on DICER's ability to process 5'GG-prehsa-miR-lOb. (A) 25 nM 32P-5'-labeled 5'-GG-pre-hsa-miR-10b were treated with purified
APE1 (lane 3) for 60 mins at 37°C under the standard endonuclease assay. For reference, an
alkaline hydrolysis ladder was generated (lane 2). Numbering on the left indicates guanosine
residue sites cleaved by RNase Tl under denaturing conditions (lane 1), whereas numbering
on the right indicates sites cleaved by APE1. (B) The major cleavage products 1 and 2
generated by DICER are shown with numbering on the right. 32P-5'-labeled 5'-GG-pre-hsamiR-lOb pre-treated with (lane 4) or without (lane 3) APE1 for 30 mins at 37°C prior to
incubation with 0.25 units of DICER for additional 30 mins. (C) The RNA secondary

99

structure of 5'-GG-pre-hsa-miR-10b shows locations of cleavage sites by APE1. DICER
products 1 and 2 are shown.

6

4#

£&f

Stnmjj

RcAPEl

•*

Weak

<!
-MC

291JG
- 35 UC
- 36 CA

Li
C
36 CA
- 35 UC
- 33 UC

G27-

2<> l i t ;
28111!
- 26 UG

G
U
U
G

-G »«C

U

- Major
Product
(26 nf)

G
U

c
A
G
C

u
A
U
U
C
G
A
U
G G

5'end

G G

5'-GG-hsa-miR-21
Dicer Product
Figure 31: Effect of pre-treatment with APE1 on DICER's ability to process 5'GG-prehsa-miR-21. (A) 25 nM 32P-5'-labeled 5'GG-pre-hsa-miR-21 were treated with purified
APE1 (lane 4) for 60 mins at 37°C under the standard endonuclease assay. For reference, an
alkaline hydrolysis ladder was generated (lane 1). Numbering on the left indicates guanosine
residue sites cleaved by RNase Tl under denaturing conditions (lane 2), whereas numbering
on the right indicates sites cleaved by APE1. (B) Numbering on the right indicates the major
cleavage product generated by DICER. 32P-5' -labeled 5'GG-pre-hsa-miR-21 pre-treated with
(lane 4) or without (lane 3) APE1 for 30 mins at 37°C prior to incubation with 0.25 units of
DICER for additional 30 mins. (C) The RNA secondary structure of 5'GG-pre-hsa-miR-21
shows locations of cleavage sites by APE1. The major DICER product is shown.

100

CHAPTER 4
Assessing the endoribonuclease activity of APE1 in biological
systems
In chapter 2, we demonstrated that APE1 has the ability to cleave c-myc CRD RNA
in vitro. c-Myc is a major oncogene that is known to promote cell survival and proliferation
(Dang et al. 2005). Similarly, over-expression of c-myc mRNA has been found in virtually
all types of cancers and the fact that APE1 can cleave c-myc mRNA in vitro warranted
further study to confirm this observation in cultured cell line. This would be the first study to
assess the RNA cleaving activity of APE1 in a biological system. The first part of this
chapter investigates the involvement of APE1 in the turnover of c-myc mRNA in cultured
HeLa cervical cancer cells. By treating cells with DICER substrate for RNA interference
specific for APE1 mRNA (APEl-dsRNAi), we could reduce the levels of APE1 expression
in these cells. Total RNA from these cells were harvested and subjected to quantitative RealTime PCR (qRT-PCR) to measure the levels of specific mRNAs of interest. By observing the
relevant ratios of c-myc mRNA to that of a housekeeping gene (P-actin), we could compare
the changes in the levels of c-myc mRNA in APEl-dsRNAi and control-dsRNAi treated
samples. In addition, we assessed the stability of c-myc mRNA in APEl-dsRNAi and
control-dsRNAi treated samples to assess the role of APE 1 in c-myc mRNA stability.
The second part of this chapter tested the validity of applying a bacterial-based
genetic selection system to identify amino acid residues important for the RNA cleaving
activity of APE1 in cells. This was carried out by expressing APE1 and its amino acid
substitution mutants in E. coli Origami cells, a genetic selection system which have been
previously used in determining the essential residues for endoribonucleases such as RNase A

101

and Angiogenin (Figure 32). This system takes advantage of the ability of a ribonuclease to
cleave cellular RNA and hence cause cell death (Smith and Raines 2006). In general, RNase
A and its homologs are not toxic when expressed in typical E. coli strains. This is so because
these ribonucleases contain three or four disulfide bonds, and the reducing environment of a
typical E. coli cytosol will not allow appropriate formation of these bonds. Consequently, the
enzymes are not folded properly to their active state and cellular RNAs are not cleaved.
However, the genetically engineered Origami strain has mutations in two genes, thioredoxin
reductase and glutathione reductase, which shift the reducing potential and therefore allow
disulfide-bond formation in the cytosol. Thus, plasmid-encoded RNase A and Angiogenin
fold and are toxic to Origami cells, allowing the identification of inactive mutants of these
ribonucleases (Smith and Raines 2006).

cleavage

Origami™ E. coli

Figure 32: The genetic selection system for ribonuclease activity in Origami cells (Smith
and Raines 2006). Ribonuclease activity is toxic to the cells, unless missing residues
essential for its activity.
The advantage of this system is that it is fast and has successfully been demonstrated to
detect biologically relevant ribonuclease activities of RNase A and Angiogenin. In chapter 3,
APE1 was shown to cleave multiple RNAs in vitro. In turn, APE1 is expected to have an
impact on Origami cells' survival by cleaving multiple RNAs. When expressed in Origami

102

cells, the mutant forms of APE1 missing essential residues through amino acid substitutions
are expected to result in colony formation.

4.1 Methodology
4.1.1 Assessing the steady-state c-myc mRNA level in APE1 knocked down cells
Cell culture and reagents
HeLa cells were cultured in a tissue culture flask T75CN vent cap red (Sarstedt,
Newton, NC) with 20 mL of Minimum Essential Medium (MEM) (Invitrogen, Carlsbad, CA)
supplemented with 10% fetal bovine serum. The cells were incubated under 5% CO2 and at
37°C. Maintenance of cells were done by splitting the cells in 1:10 ratio into a fresh flask
approximately after two days of growth or when the cells have reached a confluency of 90%.
To plate cells on experimental culture plates, cells in the T75CN flask were washed with 3
mL of PBS pH 7.4 (Invitrogen, Carlsbad, CA) and subsequently treated with 2 mL of 0.25%
Trypsin-EDTA (Invitrogen, Carlsbad, CA) and incubated at 37°C and 5% C 0 2 for 4 mins.
Next, 8 mL of MEM was added to the cells and by pipetting, chunks of cells were separated.
From 10 mL, 0.5 mL of cell sample was taken and from that, 40 ul of cells were mixed with
equal volume of Trypan Blue dye (Sigma). Sample of the mixture was loaded onto the each
side of the Bright Line hemacytometer (Hausser Scientific, Horsham, PA) and the number of
cells in the 16-square quadrant was counted. After counting several quadrants, the numbers
were averaged and the cell concentration in the T75CN flask was determined. Subsequently,
cells in the flask were diluted to 2.5 x 104 cells/mL or 5 x 104 cells/mL and was plated at 2
mL (5 x 104 cells or 10 x 105) onto each wells of 6-well flat bottom plates (Sarstedt, Newton,
NC). The plated cells were grown for 15 - 17 hrs at 37°C and 5% CO2 before transfection.

103

Transfection
Before transfection, cells in the 6-well plates were washed with 1 mL/well PBS pH
7.4 and changed of their media with 2 mL of fresh MEM. 2 ul of Lipofectamine 2000
Reagent (Invitrogen, Carlsbad, CA) and 248 ul of OPTI-MEM (Invitrogen, Carlsbad, CA)
were mixed and incubated under room temperature for 5 mins. As well, 2.5 ul of 20 uM
DICER substrate RNAi against APE1 mRNA (Table 13) or DICER substrate ScrambledNegative control (IDT, San Diego, CA) was mixed with 247.5 ul of OPTI-MEM, and
incubated under room temperature of 5 mins. Next, 250 ul of OPTI-MEM and Lipofectamine
2000 Reagent mix was added to the 250 ul of siRNA or scramble OPTI-MEM mixture,
shaken vigorously, and incubated under room temperature for 20 mins. This allows the
association of lipofectant agent and siRNA to ensure high transfection efficiency. 500 ul of
siRNA or scramble mixture in OPTI-MEM was added to the well of the 6-well plates. The
cells were grown for 24 hrs at 37°C and 5% CO2 before another transfection following the
same protocol.
Table 13: Sequences of Scrambled-Negative DICER substrate double stranded RNAi
(dsRNAi) and dsRNAi against APE1 mRNA.
Oligonucleotide
Sequence
Scrambled Negative-dsRNAi sense
r(CUUCCUCUCUUUCUCUCCCUUGU)dGA
Scrambled Negative-dsRNAi antisense r(UCACAAGGGAGAGAAAGAGAGGAAGGA)
APE1-dsRNAi sense
r(GUCUGGUACGACUGGAGUACCGG)dCA
APE 1-dsRNAi antisense
r(UGCCGGUACUCCAGUCGUACCAGACCU)

Lysis and protein extraction
After 24 hrs or 48 hrs since the first transfection, cells were washed with 1 mL of
PBS pH 7.4. For each well, 200 ul of 8M Urea Lysis Buffer (8M Urea, 100 mM NH2P04, 10
mM Tris-HCl) was added. The cell lysates were collected with cell scraper and the lysates
were transferred to an eppendorf tube. The lysates were stored at -80°C for 15 min before

104

quickly thawing them at 42°C for 5 mins. Subsequently, the lysates were vortexed for 30
sees. The freeze-thaw-vortex cycle was repeated for two additional times. After the final
vortex, lysates were spun at 12,000 rpm for 15 mins at 4°C. The supernatant (~ 200 pi) was
transferred to a fresh eppendorf tube and 3 ul of the sample was taken for Bradford assay to
determine the total extracted protein concentration. The rest of the proteins were stored at 80°C.
Bradford assay
Concentration standards were prepared to generate a linear regression equation. 10
mg/mL of Bovine Serum Albumin (BSA, New England Biolabs, Beverly, MA) was diluted
1:10 in autoclaved water. The resulting stock of 1 mg/mL BSA was subsequently aliquoted
in 5, 10, 15, 20, 25, 30 ul portions to be diluted in 795, 790, 785, 780, 775, 770 ul of
autoclave water, respectively. Next, the 800 pi BSA samples were mixed with 200 pi
Bradford assay reagent (Biorad, Hercules, CA) and incubated at room temperature for 2 to 5
mins to have their colors change. From the total mixture of 1 mL, 200 pi aliquots placed in
triplicates into the wells of the 96-well plate (Sarstedt, Newton, NC). The software for
Multiskan Ascent (Thermo Fisher Scientific, Waltham, MA) was used to quantify the
absorption at a wavelength of 595 nm. Subsequently, the measurements were plotted in
absorption versus ug/mL and the linear equation was derived using Excel spreadsheet. 3 pi
of the lysate sample was mixed with 797 pi of autoclaved water. Then, 200 pi Bradford assay
reagent was added and 200 pi aliquots were placed in triplicates into the wells of the 96-well
plate. Using the software MultiSkan, the absorption value of the sample at a wavelength of
595 nm was obtained. Subsequently, the concentration of the protein sample was determined
using the BSA standard equation.

105

Acetone precipitation
To concentrate the proteins for Western Blot analysis purposes, a calculated volume
of protein sample was mixed with five volumes of 100% acetone. The sample was mixed and
stored in -20°C for 15 mins before spinning at 12,000 rpm for 10 mins at 4°C. The protein
pellet was formed and the supernatant was removed by pouring. 200 pi of 80% acetone was
added to the pellet for rinsing and the sample was spun at 10,000 rpm for 5 mins at 4°C. The
supernatant was removed by pouring and the pellet was dried under fumehood for 15 mins.
Subsequently, the pellet was resuspended in 10 ul of autoclaved water and stored at -20°C.
Western Blot
Prior to transfer, a nitrocellulose membrane and four pieces of filter papers were cut
out in order to have the same dimension as the SDS-PAGE resolving gel. The membrane was
first soaked in de-ionized water for 1 min and subsequently equilibrated in Western Transfer
Buffer for 5 mins at room temperature. SDS-PAGE resolving gel was also equilibrated in
Western Transfer Buffer (Table 14) for 5 mins at room temperature. The transfer sandwich
was assembled in the following order starting from the bottom. A sponge mat was soaked in
Western Transfer Buffer and was placed on the very bottom, followed by the two soaked
filter papers, nitrocellulose membrane, SDS-PAGE resolving gel, two filter papers, and a
sponge mat at the top. The transfer sandwich was placed into the transfer apparatus and the
transfer was done at 90 V for 40 mins at 4°C.

106

Table 14: Composition o : the reagents used in Western blot.
Reagents
Composition
10 x TBS
100 mM Tris-HCl pH 7.4, 1.5 M NaCl
Western Transfer Buffer
37 mM Tris-HCl, 39 mM Glycine, 20 %
methanol
Western Blocking Buffer
10 mM Tris-HCl pH 7.4, 0.15 M NaCl, 5 %
Skim Milk
ST
10 mM Tris-HCl pH 7.4, 0.15 M NaCl, 1 %
Skim Milk, 0.1 % Tween-20
Western Stripping Buffer
62.5 mM Tris-HCl pH 6.7, 2 % SDS, 100
mM P-mercaptoethanol
Following the transfer, the membrane was blocked with Western Blocking Buffer
either at 37°C for 30 mins or at 4°C overnight. Next, the membrane was rinsed twice quickly
in 3 mL of ST and washed for 15 mins at room temperature under gentle shaking with 5 mL
ST. Additional two washes of 5 mins in 5 mL ST was done at room temperature. The
membrane was then incubated with 1° antibody diluted in 1:1500 for monoclonal anti-APEl
antibody raised in mouse (Santa Cruz Biotechnology Inc.) or 1:2500 for monoclonal anti-Pactin antibody raised in mouse (Sigma) in ST for 2 hrs at room temperature. The membrane
was then, rinsed and washed as described previously. Next, the membrane was incubated
with 2° antibody (anti-mouse polyclonal IgG (Promega, Wyoming, MI)) diluted in 1:4000 in
ST for 1 hour at room temperature. The membrane was then, rinsed and washed as described
previously. After the final wash, the membrane was developed by incubation of 1 min with
SuperSignal West Pico Chemilluminescent substrate (Pierce, Rockford, IL) as per
manufacturer's protocol and exposed and visualized using Chemlmager (Alpha Innotech,
San Leandro, CA). The membrane was stripped by incubation with Western Stripping Buffer
at 50°C for 30 mins, and blocked by Western Blocking Buffer as described previously for
another round of immunoblot detection when necessary.

107

Total RNA extraction
Twenty-four hrs or 48 hrs after the first transfection, cells were washed with 1 mL of
PBS pH 7.4. Subsequently, 1 mL of Trizol Reagent (Invitrogen, Carlsbad, CA) was added
and pipetted up and down to homogenize the cells. The mixture was soon transferred to a
fresh eppendorf tube, to which 200 ul of chloroform : isoamyl alcohol (CHCI3: IAA = 49 : 1,
Fluka) was added and the mixture was inverted six times before spinning at 3,000 rpm for 10
mins at 4 °C. The top aqueous layer was transferred to a fresh eppendorf tube and 500 ul of
isopropanol was added and inverted six times. The sample was spun at 11,000 rpm for 15
mins at 4 °C to form an RNA pellet. The supernatant was removed by decanting and 200 ul
of cold 75% ethanol was added to rinse the pellet, and then spun at 9,000 rpm for 5 mins at 4
°C. The supernatant was removed by decanting and the pellet was dried in fumehood for 15
mins. RNA was resuspended in 100 ul of DEPC-treated water and its concentration was
determined

by ND-1000 UV-Spectrophotometer

(NanoDrop V.3.1.0, Wilmington,

Delaware). The concentration was determined by using the relationship:
RNA concentration (ug/ml) = (OD26o) x (dilution factor) x (40 ug RNA/ml) / (1 OD2eo unit).
cDNA synthesis
A total reaction mix of 20 ul included 1 ug of total RNA, 4 ul of 5 x iScript Reaction
Mix (Biorad, Hercules, CA), 1 ul of iScript Reverse Transcriptase (Biorad, Hercules, CA),
and varying amount of nuclease-free water (Biorad, Hercules, CA) to adjust the total volume.
At times, half volume of each reagent was used for the total reaction mix of 10 ul. The
MiniCycler PCR machine (MJ Research Inc., Watertown, MA) was programmed for 5 min at
25°C, 30 min at 42°C, 5 min at 85°C, and hold at 4°C to generate the cDNA.

108

Quantitative Real Time PCR (qRT-PCR)
For one reaction per gene, a 24 ul master mix was made using 12.5 ul of iQ SYBR
Green Supermix (Biorad, Hercules, CA), 1 ul of 10 uM forward primer (Table 15), 1 ul of
10 uM reverse primer (Table 15), and 9.5 ul of autoclaved water.
Table 15; Primer sequences used in qRT-PCR.
RT-qPCR Primers
Sequence
Human APE1 Forward
5'-TGGAATGTGGATGGGCTTCGAGCC-3'
Human APE1 Reverse
5' -AAGGAGCTGACC AGTATTGATGA-3'
Human c-Myc Forward
5' - ACGAAACTTTGCCC ATAGCA-3'
Human c-Myc Reverse
5' -GCAAGGAGAGCCTTTC AGAG-3'
5'-TTGCCGACAGGATGCAGAAGGA-3'
P-actin Forward
5' - AGGTGGAC AGCGAGGCC AGGAT-3'
P-actin Reverse
This master mix was aliquotted into the well of the 96-well PCR microplate (Axygen
Scientific, Union City, CA) at room temperature. Next, 1 ul of cDNA sample was added to
the master mix in the PCR plate and the plate was sealed using the MicroSeal B Adhesive
Sealer (Biorad, Hercules, CA). For one gene, there were triplicates of each reaction. iCycler
Thermal Cycler (Biorad, Hercules, CA) was programmed by using the software, iQ5 Optical
System Software Version 2.0 (Biorad, Hercules, CA). The program was set up for cycle 1 at
95°C for 3 mins, cycle 2 repeated 40 times at 95°C for 10 sees and at 52°C for 30 sees, and
cycle 3 repeated 87 times at 52°C for 10 sees. The CT values obtained from each triplicates
were averaged and used to calculate the relative mRNA levels of interest using the Livak
method (Livak and Schmittgen, 2001). Cj value represents the cycle number at which the
fluorescence generated within a reaction crosses the threshold and this value is inversely
correlated to the logarithm of the initial copy number of cDNAs. A sample calculation
determining the fold changes in mRNA levels using CT values is shown on section 4.2.1.

109

4.1.2 Assessing c-myc mRNA half-life in APE1 knocked down cells
A transcriptional inhibitor, 5,6-dichloro-l-P-D-ribofuranosylbenzimidazole (DRB,
Sigma) was used for this method. The DRB stock of 200 uM x 100 was prepared in 100%
ethanol and stored at -20 °C until further use. The cells were plated at 10 x 104 or 5 x 104
cells/well and transfected with either siRNA against APE1 or scramble over 36 hrs as
described previously in section 4.1.1. After 36 hrs since the first transfection, the cells were
washed with 1 mL PBS pH 7.4. Next, DMEM containing 200 uM of DRB was applied to the
cells and the cells were incubated at 37°C and 5% CO2. At each desired time point, the cells
were washed with PBS pH 7.4 and their RNA harvested using 1 mL of Trizol Reagent
(Invitrogen, Carlsbad, CA). The subsequent steps in extracting RNA are identical to the
described method in 4.1.1.
4.1.3 Preparation of LB-antibiotic plates for Origami cells
The solid LB-agar mixture was liquefied in microwave and appropriate antibiotics
were added. One set of LB-agar plates contained the final concentration of 12.5 ug/mL
Tetracycline and 15 ug/mL Kanamycin. Another set contained 100 ug/mL Ampicillin. Next,
per each 100 mm plates (Fisherbrand), 15 mL of LB-agar was added and allowed to solidify
in room temperature.
4.1.4 Preparation of Origami competent cells
50 ul of Origami B(DE3) cells (Novagen) were streaked on LB-kan-tet plate. The
plate was wrapped on its sides with a strip of plastic plate and incubated overnight at 37°C.
The next day, a single colony was picked from the plate and was grown overnight at 37°C
with shaking in a 50 mL falcon tube containing 10 mL of LB-kan-tet. The next day, 500 mL
LB supplemented with 12.5 ug/mL Tetracycline and 15 ug/mL Kanamycin was prepared and

110

5 mL of the overnight culture was added to the 500 mL LB-kan-tet. The culture was shaken
(200 rpm) at 37°C until the optical density measured at wavelength of 600nm (ODeoonm) had
reached ~0.4. The ODeoonm was measured using DU800 UV/visible Spectrophotometer
(Beckman Coulter Inc., Palo Alto, CA). Next, the culture was split in 250 mL portions and
transferred into sterile centrifuge bottles. The culture was centrifuged for 30 mins at 3500
rpm at 4°C using benchtop centrifuge Allegra X-12R Centrifuge (Beckman Coulter Inc., Palo
Alto, CA). Once, most of the cells were pelleted at the bottom, the cells were resuspended in
30 mL of cold 100 mM CaCl2 and transferred to a 50 mL falcon tube to be incubated on ice
for 30 mins. Again, the cells were centrifuged at 3500 rpm at 4°C for 15 mins. Next, the clear
supernatant was discarded and the cells were resuspended in 4 mL of cold 100 mM CaCb
with 15% glycerol. The cells were aliquotted in 100 ul stocks and stored at -80°C.
4.1.5 Transformation and induction of protein expression by IPTG
The competent Origami cells in 100 ul aliquots were thawed on ice. Next, 100 ng of
plasmids containing either APE1 WT or mutant cDNA were aliquotted into 15 mL round
bottom tubes on ice. The thawed cells were added to the plasmids and the cells were
incubated on ice for 20 mins. The cells were transformed by heat shock at 42°C for 75 to 90
sees and subsequently cooled on ice for 2 mins. 500 ul of LB was added to the cells and they
were allowed to recover at 37°C for 40 mins with shaking (200 rpm). In the meantime, LB
agar plates containing lOOug/mL Ampicillin were pre-warmed at 37°C. Ampicilliln plates
were used to initially screen transformed cells. For IPTG induction, 125 ul aliquots of cells
were further incubated with 0 to 750 uM IPTG for additional 5 mins before being plated on
to LB-ampicillin plates. The plates were incubated at 37°C for 24 hrs. It is noteworthy that,
with Ampicillin alone, we can not distinguish whether the surviving colonies were truly

111

Origami cells. For this purpose, Tetracycline and Kanamycin supplemented plates would be
used. However, in our results testing the effects of mutant APEls, except for H309N, all of
the transformed cells did not survive on LB-ampicillin plates, hence there was no need to
additionally screen them under Tetracycline and Kanamycin supplemented plates.

4.2 Results and Discussion
4.2.1 APE1 knock down in HeLa cells up-regulates c-myc mRNA expression
To assess the RNA cleaving activity of APE1 and its influence on c-myc mRNA
level, we knocked down the expression of APE1 in HeLa human cervical cancer cell line
using DICER substrate RNAi against APE1 mRNA (APEl-dsRNAi). The sequences were
based on siRNA that was successfully used to knock down APE1 (Fan et al. 2003; Table 13).
For the control, DICER substrate Scrambled-Negative (Control-dsRNAi) was used (Table
13). The delivery of these substrates was mediated through Lipofectamine 2000, which had
shown a transfection efficiency of -80% (Bassett et al. 2008). The level of APE1 and c-myc
mRNAs were assessed through quantitative Real-Time PCR (qRT-PCR) and normalized
against mRNA for p-actin, which served as a constitutive expression control.
The reduced levels of APE1 protein and transcripts after 24 hrs and 48 hrs of
transfection were assessed through Western analysis and qRT-PCR. The degree of APE1
reduction at protein levels was quantified by calculating the ratios of density of APE1 in the
control versus the knock down lane using OptiQuant software (Hewlett Packard, Palo Alto,
CA). The reduction was found to be -60% after 24 hrs and -70% after 48 hrs (Figure 33).

112

v

#

24 hrs

48 hrs

/ "

/ "

> /

<? ^ <fK /
'»

***»

I APE1

1 2
3
4
Figure 33: Western blot illustrating the reduced levels of APE1 after two time points
using dsiRNAi against APE1 in HeLa cells. A total of 30 |jg of proteins were loaded onto
the wells and ran on SDS-PAGE gel. The gel was then blotted onto a nitrocellulose
membrane. The identification of proteins was made via monoclonal antibodies against APE1
and beta-actin. Top panel shows the amount of APE1 in dsiRNA-scramble (lanes 1 and 3)
versus the dsiRNA-APEl (lanes 2 and 4) treated cells at two time points. Bottom panel
shows the amount of (3-actin in these cells as a control for loading the same amount.
This was correlated to APE1 reduction at mRNA levels after 24 hrs (-80%) and 48
hrs (-85%) of transfection (Figure 34A). After 24 hrs, the knock down was significant (P <
0.001, student t-test). Although the t-test (P > 0.001) for the knock down at 48 hrs did not
give significance, we could speculate a significant reduction of APE1 mRNA at this time
point verified by Western data (Figure 33). APE1 knock down was also associated with
elevated expression of c-myc mRNA. Using the same total RNA extract, we found increased
level of c-myc mRNA in both 24 hrs (-2.2-fold) and 48 hrs (~4.6-fold) upon APEl-dsiRNAi
transfection (Figure 34B). At 24 hrs, the up-regulation was not significant according to the
student t-test (P > 0.001). However, at 48 hrs, the up-regulation was significant (P < 0.001).
This showed that APE1 was involved in the regulation of c-myc mRNA expression and
warranted further studies to verify whether APE1 was involved in the control of c-myc
mRNA stability.

113

B

A
• Control-dsRNAi

6-i

• APEl-dsRNAi
1.2 -

B Control-dsRNAi
DAPEl-dsRNAi
**

A ratio

5-

1
2 4<

Z 0.8Pi

z

S
.5

E 3-

"g 0.6-

u

i
g 0.4W

a) Z -

<

T

**

0.2-

±
24

48
rime (hrs0

i

1-

1

24

48
Time (hrs)

Figure 34: Steady state level of c-myc mRNA is elevated after transient knock down of
APE1 using double stranded siRNA in HeLa cells. qRT-PCR was carried out on cDNA
samples generated from harvested RNAs at two different time points to measure the relative
levels of (A) APE1 mRNA or (B) c-myc mRNA in both Control-dsiRNAi and APE1dsiRNAi treated HeLa cells. Asterisks indicate the student t-test value was less than 0.001.
Table 16 shows a sample calculation using CT values obtained from qRT-PCR to
assess the fold changes in c-myc mRNA after 24 hrs of transfection with APEl-dsRNAi as
compared to the Control-dsRNAi. To begin the calculation, CT value for 6-actin (15.13) was
subtracted from c-myc (23.46) for the Control-dsRNAi 24h-l sample to give a ACT value of
8.33.
ACT: c-myc CT (23.46) - 6-actin CT (15.13) = 8.33
Next, similar calculations were performed for two other samples for Control-dsRNAi
treated samples as well as the triplicates treated with APEl-dsRNAi to obtain the differences

114

in their CT values (ACT) between c-myc and 6-actin mRNAs as shown under column ACT (Cmyc C T - P-actin CT) (Table 16).
From these values, an average was calculated for the Control-dsRNAi (8.11) and
APEl-dsRNAi (6.97) treated samples.
Average of ACT for Control-dsRNAi: (8.33 + 8.02 + 7.98)/3 = 8.11
Average of AC T for APEl-dsRNAi: (6.58 + 7.11 + 1.22)13 = 6.97
Next, each ACT value of the triplicate in the Control-dsRNAi were subtracted from
the average values for Control-dsRNAi (8.11) and APEl-dsRNAi (6.97), respectively. For
example, from an average value for Control-dsRNAi (8.11), Control-dsRNAi 24h-l (8.33),
Control-dsRNAi 24h-2 (8.02), and Control-dsRNAi 24h-3 (7.98) were all subtracted
separately to give -0.22, 0.09, and 0.13, respectively.
Average of ACT for Control-dsRNAi (8.11) - Control-dsRNAi 24h-l (8.33) = -0.22
Average of ACT for Control-dsRNAi (8.11) - Control-dsRNAi 24h-2 (8.02) = 0.09
Average of ACT for Control-dsRNAi (8.11) - Control-dsRNAi 24h-3 (7.98) = 0.13
Similarly, from an average value for APEl-dsRNAi (6.97), values for ControldsRNAi 24h-l (8.33), Control-dsRNAi 24h-2 (8.02), and Control-dsRNAi 24h-3 (7.98) were
also subtracted separately to give -1.36, -1.05, and -1.01, respectively.
Average of ACT for APEl-dsRNAi (6.97) - Control-dsRNAi 24h-l (8.33) = -1.36
Average of ACT for APEl-dsRNAi (6.97) - Control-dsRNAi 24h-2 (8.02) = -1.05
Average of ACT for APEl-dsRNAi (6.97) - Control-dsRNAi 24h-3 (7.98) = -1.01
These differences (AACT) were averaged for the Control-dsRNAi (0.00) and APEldsRNAi (-1.14) treated samples.
Average of AACT of Control-dsRNAi: (-0.22 + 0.09 + 0.13)/3 = 0.00

115

Average of AACT of APEl-dsRNAi: (-1.36 + -1.05 + -1.01)/3 = -1.14
Finally, by calculating 2~AACT, normalized c-myc mRNA fold changes for the ControldsRNAi (2"(0 00) = 1) and APEl-dsRNAi (2~("u4) = 2.22) treated samples were determined.
Table 16: Sample table of CT values generated from qRT-PCR
AACT
(Avg. ACT
B-actin
mRNA
CT

c-myc
mRNA
CT

ACT
(c-myc
CT

- P-actin

ACT Control-

Normalized
c-myc
mRNA
in
Control-dsRNAi
OR
APEl-dsRNAi
relative to
Control-dsRNAi

dsRNAi)

( 2 -AAC T )

Control-dsRNAi
ACT Control-

dsRNAi) O R

(Avg. ACT
APEl-dsRNAi -

ControldsRNAi 24h-l
ControldsRNAi 24h-2
ControldsRNAi 24h-3
Average
DEV
APEl-dsRNAi
24h-l
APEl-dsRNAi
24h-2
APEl-dsRNAi
24h-3
Average
DEV

15.13

23.46

8.33

-0.22

1.17

14.90

22.92

8.02

0.09

0.94

14.97

22.94

7.98

0.13

0.91

15.00
.12

23.11
.31

8.11
0.19

0.00
0.19

1.01
0.14

16.40

22.98

6.58

-1.36

2.58

15.92

23.02

7.11

-1.05

2.07

15.86

23.08

7.22

-1.01

2.01

16.06
.30

23.03
.05

6.97
0.34

-1.14
0.19

2.22
0.31

4.2.2 Optimization of monitoring c-myc mRNA stability in HeLa cells
By using transcriptional inhibitors, c-myc mRNA decay could be monitored, c-myc
mRNA has been shown decay with a half-life of about 20 to 40 mins (Dani et al. 1984;
Herrick and Ross 1994). A few candidate chemicals were initially selected and applied in our
experiments using HeLa cells. Actinomycin D and alpha-amanitin were tested at a
concentration of 5 ug/mL and 15 ug/mL, respectively (Figure 35A). It was found that at
tested concentrations these two drugs were not able to inhibit transcription. Under such
116

conditions, monitoring c-myc mRNA decay was not feasible. Later, it was found that 5,6dichloro-1-P-D-ribofuranosylbenzimidazole (DRB) at concentrations of 100 uM and 200 uM
were able to inhibit transcription (Figure 35B). Apparent decay c-myc mRNA was
observable after 30 mins of DRB incubation. We therefore chose DRB as the transcriptional
inhibitor for our mRNA half-life studies.

A

B
1.25

1

S S 0.75

II

0.5

0.25 ]

0

20

40
Time (minutes)

60

80

- D R B 100 (cM
- D R B 200 |iM

30

60

Time (minutes)

Figure 35: Level of c-myc mRNA monitored over time after treating cells with
transcriptional inhibitor Actinomycin D, a-amanitin, and DRB. Cells were initially
treated with Control-dsiRNAi over 24 hrs and qRT-PCR was performed on cDNA samples
generated from total RNA extracted at different time points. (A) Data points from two
experiments were pooled in treatment of cells with Actinomycin D (5 ug/mL). For Alphaamanitin treatment (15 ug/ml), only one screening was carried out. (B) mRNA decay
occurring at 60 mins after treatment with DRB (100 uM or 200 uM equivalent to 32 ug/ml or
64 ug/ml).
Cells were plated at a density of 5 x 104 cells/well and 200 uM DRB was applied
after -40 hrs of growth. At different time points, these cells were harvested and their total
RNA were subjected to qRT-PCR to determine their relative levels of c-myc mRNA (Figure
36). It was found that at multiple time points, c-myc mRNA exhibited decay after treating
cells with DRB. Over the course of 75 mins, 200 uM of DRB was able to inhibit
transcription and c-myc mRNA decayed to about 20 % (Figure 36). The faster initiation of

117

mRNA decay compared to the previous observation in Figure 35B may have resulted from
the higher ratio of DRB concentration to the cells. The decay plot showed after -45 mins, cmyc mRNA decayed to -50 % and this observation was in line to the previously reported
half life of c-myc mRNA.

1.75-

- - • - - D R B 100 tiM
—!^— DRB 200 |jM

1.5-

_o
'-fc*

A

<
a 1.25- ^
7 =
S =

s'i

c o
'S o
53

44

-ta*

•*-»

<S
"5
«3 <-

s

•

**^<w^

lt !

"^V^
^S^Jl

0.75-

^^V.

A

0.5

%

*»

^V1*'*--."
U^S.

1

0.25
00

15

30
45
Time (minutes)

**-.

^s.

**•

60

75

Figure 36: Decay of c-myc mRNA over time after treating cells with DRB. Cells were
treated with Control-dsiRNAi over 24 hrs and then treated with DRB at two concentrations
(100 uM & 200 uM). qRT-PCR was performed on cDNA samples generated from total RNA
extracted at different time points.
4.2.3 Knock down of APE1 in HeLa cells stabilizes c-myc mRNA
To investigate the reason behind c-myc mRNA elevation, stability of c-myc mRNA
in APEl-dsiRNAi treated samples was assessed (Figure 37). HeLa cells were plated at a
density of 5 x 104 cells/well and transfected with either APEl-dsRNAi or Control-dsRNAi
for 30 hrs before they were treated with 200 uM of DRB. It was found that the stability of cmyc mRNA in APEl-dsRNAi treated samples were enhanced up to 60 mins compared to the
control which exhibited mRNA half-life of -30 mins which is consistent with the previously

118

reported value (Dani et al. 1984; Herrick and Ross 1994). This experiment suggested that the
increased expression of c-myc mRNA in APEl-knock down cells is in part due to increased
transcript stability.

Time (minutes)

Figure 37: c-myc mRNA is stabilized after transient knock down of APE1 in HeLa cells.
qRT-PCR was carried out on cDNA samples generated from harvested RNAs at three
different time points to measure the relative levels of c-myc mRNA in both scramble dsiRNA
and dsiRNA-APEl treated HeLa cells. The difference in the slopes were determined to be
significant by regression analysis (p < 0.05).
4.2.4 Origami cell transformation with pGEX4T3-Thioredoxin A and RNase A
The pGEX4T3 based vector carrying cDNAs for Thioredoxin A and RNase A were
kindly provided as a gift from Dr. Ronald Raines, University of Wisconsin-Madison. When
pGEX4T3-Thioredoxin A was transformed into Origami cells in an increasing amount of
plasmids (lOng - 750ng), an increase in the numbers of colonies was observed (Figure 38).

119

Figure 38: Transformation of Origami cells with increasing amounts of pGEX4T3Thioredoxin A. (A) 10 ng and 50 ng of plasmids were used for transformation. (B) 100 ng
and 250 ng of plasmids were used for transformation (C) 500 ng and 750 ng of plasmids
were used for transformation.
Indeed, Thioredoxin A did not exert any ribonuclease activity in these cells and
colonies were able to form. Also, this data gave us an idea on what amount of plasmids to use
to observe an optimal level of cell growth. In contrast, when pGEX4T3-RNase A was
transformed into Origami cells (Figure 39), no colonies were observed indicating cell death
imposed by expressed RNase A. This is consistent with a previously reported study (Smith
and Raines 2006). This gave us confidence to proceed with our investigation in assessing the
WT APE1 and its mutants on the growth of Origami cells which would serve as a model to
rapidly assess the endoribonuclease activity of APE 1 in cells.

• • - . '

Nj&-'fi?ti!

Figure 39: Transformation of Origami cells with pGEX4T3 plasmids carrying cDNAs
for Thioredoxin A and Ribonuclease A. lOng of pGEX4T3-RNase A (1) and pGEX4T3Thioredoxin A (2) were used to transform 100 ul of Origami cells. The cells were heat
shocked, recovered, and plated on LB-agar-Ampicillin plate as described in 4.1.5.
120

4.2.5 Induction of pET15b-WT-APEl in Origami cells is toxic
The validity of this genetic selection was evaluated by observing the correlation
between the in vitro activities of APE1 mutants (Table 9) and the death/survival of Origami
cells after expressing these mutants. To test the effects of expressing APE1 WT and its
mutants, we used pET15b based plasmids that carried cDNA sequences for His6X tagged to
the N-terminal end of the APE1 WT or mutants (Figure 40A). We also tested the effects of
IPTG induction (Figure 40B). pET 15b-Acetate Kinase, kindly provided by Dr. Andrea
Gorrell, UNBC, was used as an inert control and was expected to allow cell growth.
«*.-

•w

-

^

I

'.is.. CSV

•fe
No IPTG

,t
n-.i

187.5 pM IPTG
i

i&tffts

;•

.: ^i

Figure 40: Transformation of Origami cells with pET15b plasmids carrying cDNAs for
WT-APE1 and recombinant enzymes. lOOng of plasmids were used carrying cDNAs for
WT APE1 (1), D148E (2), H255A (3), Acetate Kinase (AK) (4) in transforming 100 ul of
Origami cells as described previously. IPTG were applied to Origami cells after their heat
shock recovery of 40 mins at 37 °C. Two concentrations of IPTG are shown in the figure. (A)
0 uM (B) 187.5 uM.
Without IPTG, the WT APE1 allowed cell survival (region 1, Figure 40A), but
inducing its expression instigated cell death as shown in region 1 of Figure 40B. Acetate
kinase showed that this enzyme is not able to induce cell death both at 0 and 187.5 uM IPTG
induction (region 4 in Figure 40A and 40B), which was in line with our expected results.
However, we observed a discrepancy between the in vitro RNA cleavage activities of APE1
mutants and their expected effects in Origami cells. D148E was shown to have a reduced
RNA cleaving activity in vitro, and was expected to have a non-toxic effect in Origami cells.

121

However, we found decreased/retarded growth in Origami cells upon transformation with
D148E and induction with IPTG (Region 2, Figure 40A and B). In addition, H255A, later
found to have reduced RNA cleaving activity in vitro (Kim S.E. et ah, unpublished results)
also exerted toxic effects on Origami cells (Region 3, Figure 40A and 40B), raising the
questions on the validity of this system in assessing the endoribonuclease activity of APE1.

•(

•A

:'

:.3apM.JPT<S. .

'1
;

i \

I,4

\

-

.

.aoOtiMJBTG

:

'
'

"

:

i-j

:

375 yM IPTG

1
.

i

75b^RIP.TG
•

••

".

-'•-.

Figure 41: IPTG induces expression of APE1 WT, D283A, and H309N which exerts
toxic/non-toxic effects in Origami cells. Increasing concentration of IPTG (A) 30 uM, (B)
200 uM, (C) 375 uM, (D) 750 uM were applied to Origami cells after their recovery of 40
mins at 37 °C. The regions of colonies transformed with pET15b-His6x based vectors
carrying cDNA genes for D283A (1), WT APE1 (2), and H309N (3) are noted.
To further test the validity of this system, we tested the effects of pET15b based
vectors carrying cDNAs for D283A and H309N (Figure 41). It was also found that as the
amount of IPTG increased, fewer colonies were visible for the WT and D283A (Region 1
and 2, Figure 41A to D). This indicated that their increased expression was toxic to Origami
cells, possibly via their functional ribonuclease activity. In addition, the number of colonies
for H309N did not vary as much through out different IPTG concentrations tested (Region 3,
Figure 41A to D). This indicated the increased expression of H309N is tolerant for cell
growth, suggesting its ribonuclease activity is reduced or abrogated. Hence, the results of
expressing these two mutants (D283A and H309N) in Origami cells agreed with the expected
results from in vitro endonuclease assays showing D283N having similar activity as the WT

122

(Figure 9 and 10, section 2.2.2) and H309N having a severe reduction in RNA cleavage
(Barnes et al. 2009).
Table 17: Summary of the results of Origami cell transformations using pET15b based
plasmids coding for APE1 mutants and their RNA cleaving activities in vitro.
APEl
Mutants
WT
N68A
D70A
E96A
D148E
Y171F
D210N

Results after
expression
in Origami cells
Death
Death
Death
Death
Death
Death
Death

H255A

Death

F266A

Death

Fold reduction
in RNA incision
(Table 9 and Table 10)
1
Abolished
55.6
Not tested
6.8
17.7
8312.6
Reduction
(Kim S.E. et ah,
unpublished results)
Abolished

D283N

Death

4.0

H309N

Growth

Not tested

Therefore, to rigorously investigate the validity of this system, other mutants of APEl
were tested in Origami cells (Table 17). It was found that pET15b based vectors coding for
N68A, E96A, Y171F, D210N, and F266A, all of which have shown severe reduction in RNA
cleaving activity in vitro, were found to have toxic effects on Origami cells (data not shown).
This suggested that the death/growth of Origami cells upon transformation with APEl
mutants to assess their endoribonuclease activities was not a reliable indicator since the
results from the genetic screening did not correspond to what is expected from the in vitro
assessment of enzymatic activities. Also, cell deaths induced by expressing most of these
APEl mutants meant that the distinction among these mutants were impossible.
The unknown effects of His6X tag in the N-terminus of APEl as well as the different
plasmid (pET15b) system used by us may possibly explain the discrepancies seen in our
system using APEl and the previous report using pGEX-RNase A (Smith and Raines 2006).
123

Apart from the His6X tag in our pET-APEl vector systems, another difference between our
pET-APEl and their pGEX vector system (pGEX-RNase A) was that their pGEX-RNase A
was designed to expresss RNase A without a GST tag. From our observations, pGEX system
also gave similar transformation success comparable to our pET system when used at lower
plasmid amount. Hence, pGEX4T3-WT APE1 and pGEX4T3-E96A were transformed into
Origami cells. We found that Origami cells transformed with these plasmids showed no signs
of cell death even at higher concentrations of IPTG used (Figure 42B and 42C). WT
expression in these cells can not induce cell death, but allows visible growth. This indicated
that the method can not be an effective way to compare the growth of cells expressing APE1
mutants. Our results showed that the growth of cells expressing E96A was less than that of
the WT indicating notable a contrast to what is expected from their in vitro RNA-incision
activities. Conclusively, this showed that genetic selection by plating Origami cells after
transformation with pGEX4T3 based vector (Smith and Raines 2006) is not an optimal
biological system to assess the effects of APE1 RNA cleaving activity.

•

V5', :.lltf!P'.'.,

No t»"

.;
/

•»'

Figure 42: Transformation of Origami cells with pGEX4T3 plasmids carrying cDNAs
for Thioredoxin A, WT-APE1, and E96A-APE1. Increasing concentration of IPTG (A) 0
uM, (B) 375 uM, (C) 750 uM were applied to Origami cells after their recovery of 40 mins
at 37 °C. The areas of colonies transformed with pET-His6x based vectors carrying cDNA
genes for WT APE1 (1), E96A (2), and Thioredoxin A (3) are noted.

124

CHAPTER 5
General discussion
5.1 Role of post-transcriptional control in cancer
A variety of trans-acting factors come into effect in the regulation of gene expression
at the mRNA level. These factors include ncRNAs, RBPs, and RNases which can influence
the degradation of transcripts, and thus the production of proteins (Kim and Lee 2009).
Alterations in this process are caused by aberrant levels or activities of these factors which
are also associated with tumorigenesis and cancer progression. Whether it is due to rapid
degradation or prolonged half-life, several studies have provided the evidence which suggest
that disrupted elements involved in mRNA turnover can give rise to abnormal levels of gene
products as well as sustaining their effects (Kim and Lee 2009). Different trans-acting factors
play their part in regulating the stability of oncogenic and tumor suppressive mRNAs (Figure
43), and alterations have been associated with the attenuation or the enhancement of mRNA
degradation. For instance, HuR can stabilize a diverse population of mRNAs including
oncogenic mRNAs like that of Bcl-2, Mcl-2, ERBB2, and VEGF-A that can elicit an antiapoptotic effect as well as several matrix-metalloproteinases that have been associated with
advance stage tumors (Abdelmohsen et al. 2007; Mook et al. 2004; Bjorklund and Koivunen
2005; Yan and Boyd 2007; Scott et al. 2008; Ido et al. 2008). Similar effects are reported
with destabilizing RNA Binding Proteins as well as miRNAs and RNases. Decreased level of
AUF1 was reported in proliferating lymphoma cells with increased levels of DENR/DRP
mRNA that gives enhanced translation of several oncogenes (Reinert et al. 2006; MazanMamczarz et al. 2007). In addition, miRNA let-7 in early stages of cancer was implicated in

125

the overexpression of oncogenes CRD-BP and HMGA2 (Boyerinas et al. 2008) as well as in
integrin-beta-3 (Muller and Bosserhoff 2008).

Oncogenic mRNA

Tumor suppressive mRNA

1 RNases

t RNases

1

| miRNA

Attenuated
Degradation

t RBP (stabilizing)

\

|RBP (destabilizing)

I

Cancer

Enhanced
Degradation
/

t miRNA

jRBP (stabilizing)

| RBP (destabilizing)

Figure 43: Altered post-transcriptional control in cancer (Kim and Lee 2009).
Attenuated degradation of oncogenic mRNAs can lead to the development of cancer. This
can be caused by the following factors on the left side. Decrease in the activity of
ribonuclease (RNase) targeting such mRNAs as well as miRNAs that inhibit their translation
and promoting mRNA decay can attenuate their degradation. Decrease in the activity of
destabilizing RNA binding protein (RBP) while up-regulation in that of the stabilizing RBP
can also have similar effects. Vice versa, enhanced degradation of tumor suppressive mRNAs
can also lead to the development of cancer (Right side).

Overexpression of miR-135a and miR-135b is implicated in the down regulation of
APC mRNA and its reduction implicated in colorectal cancer (Nagel et al. 2008). Similarly,
miR-29c has been found to down-regulate oncogenic mRNAs implicated in tumor cell
invasiveness and metastatic potential (Sengupta et al. 2008). In contrast, miR-106b family
has been found to degrade p21/CDKNlA mRNA and promote cell cycle progression in
immortalized cell lines (Ivanovska et al. 2008). Also, over-expression of miRNA-373 in a
cancer cell line was shown to contribute to the down-regulation of CD44 mRNA which had
implications in raising cells' metastatic potential (Huang et al. 2008).

126

5.2 Significance of identifying and studying RNases
RNases not only play a critical role in the turn over of various transcripts, but they
also process RNAs that are required for translational control of gene expression. In
eukaryotic mRNA degradation, a number of endonucleolytic cleavage intermediates have
been reported. For example, decay intermediates have been detected for the transferrin
receptor (Binder et al. 1989), albumin (Tharun and Sirdeshmukh 1995), urokinase
(Timofeeva et al. 2000), insulin-like growth factor II (Van Dijk et al. 1998), a-globin (Wang
and Kiledjian 2000), p-globin (Stevens et al. 2002), c-myc (Ioannidis et al. 1996), MDR1
(Lee et al. 2005), hepatitis B virus (Heise et al. 2001), and mRNAs with premature
termination codons (PTC) (Gatfield and Izaurralde 2004).
In the past, only a handful of the responsible endoribonucleases have been identified
presumably due to their specificities to limited number of substrates and having no sequence
homology to known nucleases (Dodson and Shapiro 2002). Selected endoribonucleases have
been described in section 1.1.1 of this thesis. Apart from these, there are also ErEN that was
shown to cleave a-globin (Wang and Kiledjian 2000) and ARD-1 which was proposed to
cleave 3' UTR of c-myc mRNA (Claverie-Martin et al. 1997). The limited numbers of
identified mammalian endoribonucleases therefore warrant further studies to identify and
study these groups of enzymes. In addition, in order to assess their influence on the mRNA
decay and abundance, the mechanism of their catalysis and the significance their activities
need to be studied.
Also, recent reports have highlighted the emerging roles of endoribonucleolytic
mRNA decay in eukaryotic gene expression. For instance, Rrp44, previously found to
possess an 3'-5' exonuclease activity in yeast exosome complex have also been found to

127

possess an endoribonuclease activity (Schaeffer et al. 2009). This finding is expected to
remodel the conventional exonucleolytic mRNA decay revealing another layer of mRNA
abundance control mechanism mediated through endoribonuclease. Another report
demonstrated that SMG6, an enzyme associated in metazoan nonsense-mediated decay
(NMD) pathway, can cleave at or near the site of the PTC endonucleolytically (Eberle et al.
2009). This indicated that for metazoan NMD, endonucleolytic decay can efficiently initiate
this process and bypass the requirements for deadenylating/decapping of the transcript. In
addition, discoveries of novel endoribonucleases, for instance, the DYW domain of the
pentatricopeptide repeat protein (Nakamura and Sugita 2008) and tumor marker human
placental protein 11 (Laneve et al. 2009) are being reported from recent studies. In contrast
to the past where the role of endoribonucleases has been un-recognized, these reports have
highlighted their major potential in mediating the post-transcriptional regulation of gene
expression.
5.3 Benefits of studying RNases in health and therapeutics
The obvious benefit of discovering and studying RNases is in translational research
where these enzymes and/or their properties can be used for the treatment of human diseases
such as cancer. Particularly, exploiting their ability to control gene expression at the RNA
level has been incorporated into similar oncogene inactivation strategies such as antisense
oligonucleotides (ASOs), small interfering RNAs (siRNAs), ribozymes, and DNazymes. For
example, ASOs and DNazyme have been used to specifically target c-myc mRNAs (Tafech
et al. 2007) and inhibit tumorigenesis (Ponzielli et al. 2005; Tack et al. 2008). siRNAs
targeting the mRNA of reverse transcriptase of human telomerase also have demonstrated
their ability to decrease c-Myc levels in nasopharyngeal carcinoma (Wang et al. 2007).

128

Similarly, as a first of its class of anti-cancer agent, an RNase from Rana pipiens oocyte,
Ranpirnase, has been extensively studied as a novel class of cancer chemotherapeutic. It is a
homolog of bovine ribonuclease A that exhibits anti-tumor activity specific for cancer cells
(Lee and Raines 2008). In combination with other agents such as doxorubicin, it is now being
evaluated in treating breast cancers and non-small cell lung cancers, despite its shortcomings
in recent phase Illb trial. Another example is bovine seminal RNase (BS-RNase). The
mechanism of its specificity toward cancer cells is not known yet (Benito et al. 2005), but
against tumours induced in rats and mice, this RNase inhibits the tumours without any
appearance of adverse side effects. Their potential and success thus far have encouraged
development of such RNases to treat cancers. In addition, potencies and specificities of these
RNases are expected to be improved through protein engineering and as the translational
research progresses further, some of the known properties of other RNases may be applied
towards therapeutic purposes. It is with these reasons, we chose to observe and document
some of the important aspects of the novel RNA cleaving activity of APE1.
5.4 Identification of active site residues critical for RNA-cleaving activity of APE1
The first objective of this research was to identify the active site residues of APE1 in
RNA cleavage. Previous studies on APE1 mutants tested against abasic DNA had given us
the rationale to test whether these residues were also involved in RNA catalysis. Results from
DNA incision assays on APE1 mutants revealed varying degrees of reduction in AP-DNA
endonuclease activities amongst different mutants (refer to section 2.2.2). This had provided
us with the basis for assessing these mutants for endoribonuclease activity. Results from
APE1 against c-myc 1705-1792 CRD RNA and Oligo IB revealed moderate to severe
reduction in RNA incision activity exhibited by these mutants. It is noteworthy that moderate

129

reductions observed in D70A and D308A agreed with previous suggestions that these two
residues were involved in the coordination of metals during abasic DNA incision (Erzberger
and Wilson 1999). It can be speculated that the absence of one of these residues may be
relieved by the presence of the other and the mutant APE1 would be able to retain its RNA
incision activity. In addition, the most severe reductions were observed from N68A and
D210N, which were also in agreement with the previous reports. Both of these residues were
previously suggested to have a critical role in the catalysis of abasic DNA incision activity,
with N68 hydrogen bonding to D210 and H309 to allow optimal active site environment,
while D210 participating in the generation of a nucleophile (Nguyen et al. 2000; Erzberger
and Wilson 1999). Hence, the results showed that APE1 shares its active site residues in
cleaving both abasic DNA and RNA. One particular result different from that of abasic DNA
incision assays was the comparable RNA cleaving activity of D283N to that of the WT.
Considering the role of D283 in the maintenance of the D283-H309 dyad (Hadi et al. 2000),
our results indicated the alteration in this dyad may not be critical for RNA incision.
However, it is still a critical component for abasic DNA incision (Hadi et al. 2000). To
further confirm the dispensability of the dyad, we can test the RNA binding and cleavage
ability of D283A, which will have no ability to hydrogen bond to H309, and compare these
results with the data on its effect on abasic DNA.
Previously, H309N was shown to have a 2-fold reduction in its affinity towards
abasic DNA (Masuda et al. 1998). This is in line with our preliminary data that showed its
limited binding ability to RNA compared to the WT enzyme. Also, this indicated that H309
may be involved in the RNA-binding process and its mutation leads to reduced incision
activity. Our results indicated that the residues (N68, E96A, Y171, and D210) from the active

130

site are not directly involved in RNA-binding, but are important in the catalytic step of RNA
incision. D283N mutation may have affected the binding abilities to a certain extent (lanes 18
and 19), but at a higher concentration of the protein (lane 20), it displayed better binding than
the WT (Figure 12B). This was consistent to the previous report testing its binding ability to
abasic DNA where it showed less affinity than the WT at lower concentrations, but at higher
concentrations, it showed better affinity than the WT (Masuda et al. 1998). Hence, the role of
D283 in RNA-binding may not be as critical as H309. The binding abilities of E96A and
D210N to abasic DNA were all comparable to that of the WT (Barzilay et al. 1995;
Marenstein et al. 2004). Although the study on abasic DNA-binding abilities of N68A and
Y171F has not been conducted yet, none of these active site mutants, except H309N, are
expected to show alterations in the binding of DNA. It is because these residues are not part
of the DNA-binding domains that are predicted to interact with abasic DNA (Mol et al.
2000). Similarly, our EMS A and endoribonuclease assays revealed that APE1 uses residues,
N68, E96, Y171, D210, and D283, mainly for RNA incision. Hence, this similarity supports
that APE1 shares these residues to incise abasic DNA as well as RNA.
During the progress of this research, we noticed that APE1 displayed similar RNA
cleavage pattern to that of the RNase A. Hence, we attempted to test whether APE1 also
utilized the two histidine residues to cleave RNA in a manner analogous to RNase A (Kim
S.E. et al., unpublished results). The candidate histidine pair (H255-H289) was chosen based
on the three dimensional structure of APE1 generated by X-ray crystallography (Mol et al.
2000). We generated Ala mutants of these residues as well as generating Ala mutants for
three other histidines (HI 16, H151, and H215) found in the sequence of APE1. It was found
on endoribonuclease assays that the mutant H255A had reduction in its RNA-cleavage

131

activity, however, other mutants (H116A, H151A, H215A, H309N) also displayed moderate
to severe reduction in RNA cleavage. This indicated that the effect of Ala mutations were not
specific to H255-H289 pair, but altering histidines, in general, may have affected the overall
conformation of APE1 that had altered its RNA cleaving activity. This would require further
experimentation to confirm. Hence, results from screening our structural mutants revealed
that APE1 shared its active site residues for abasic DNA incision as well as RNA incision.
In addition, we have screened the RNA cleaving activities of APE1 population
variants. L104R and E126D retained their ability to cleave RNA but reduced activity against
abasic DNA. Interestingly, in RNA cleavage, they also exhibited alterations in their cleavage
sites compared to those of the WT. LI04 is located in the loop region close to the DNA
binding domain (Hadi et al. 2002). It is also speculated to be a part of hydrophobic packing
constituted by L72, L108 and W119. Arg substitution may alter this packing which results in
reduced DNA cleaving activity of APE1. It is also speculated that the DNA binding region
may be altered by this substitution which can result in alternative RNA interaction that may
render its activity to be RNasin-resistant. Alteration in the cleavage site may also result from
this perturbation. This can arise from having Arg that has basicity instead of hydrophobicity.
To further test the role of basicity in giving cleavage site alteration, mutants retaining this
basicity can be generated such as L104N, L104Q. A mutant with a bulkier base, L104K,
could also be tested. Also, the effects of hydrophobic packaging and the requirement of
optimal side chain size similar to Leu could be tested. LI041 which has a similar size but a
different conformation and LI04V which has one less -CH2- group can be tested of their
RNA cleaving activities and binding.

132

El 26 is located on one of the DNA binding domains and is near the AP-DNA
phosphate backbone during AP-DNA binding (Mol et al. 2000). El26 is speculated to
provide an important repulsive interaction, during this binding (Hadi et al. 2002). However,
with E126D, it can be speculated that though it retains its negative charge, its effect may not
be as great since it has been cut short of its side chain by one -CH2-. This may in turn affect
how different secondary structures of RNA may interact with APE1 and lead to alteration in
cleavage specificities. To investigate the role of the negative charge, mutants of El26 may be
generated and tested of their cleavage specificity. Mutants with basicities (E126Q or E126N)
or mutants with no charge (E126L or El261) may be tested. Also, variants of such mutations
bearing smaller groups with no charge (E126G or El26A) or a bulkier group (E126F or
E126W) with no charge may also be tested.
R237A was abrogated of its abasic DNA and RNA incision activities. R237 is located
in a-helix number 9 (Hadi et al. 2002) that is near the DNA binding domain (Figure 4). It is
speculated that R237A substitution imparts structural instability by disrupting the hydrogenbonding network between R237-E216 and R237-E217 (Figure 4 & Hadi et al. 2002). To
confirm the role of this network, mutants E216Q, E216D, E216N, E216A could also be
generated and tested for their RNA and abasic DNA binding and cleavage activities.
Similarly, mutations for E217 that disrupts its electrostatic state or the length of the side
chain arm may also be tested.
The three variants, D148E, G241R, and G306A, all exhibited severe reductions in
their RNA cleaving activities. This was in contrast to their functional abasic DNA incision
activities (Figure 13, section 2.2.4 and Hadi et al. 2002), and to date no study has been done

133

extensively on these residues since their mutations had minimal effects on AP-DNA
endonuclease activity.
D148 is located on the outer surface of APE1 and is likely to contribute in the
interaction with RNA and/or in the maintenance of APE1 structure. The effect of its mutation
to Glu, can further be investigated by generating mutants that can provide useful information.
We can first test the requirement of acidity of D148 with D148N. We know the effect of
gaining -CH2- from D148E showing loss of RNA cleaving activity. But, an additional effect
of losing its acidity can be tested by D148Q. Also, the effect of gain in basicity and side
chain bulk can be tested with D148H D148R, D148K. Further, the steric effects of D148 can
be tested with having a constant hydrophobicity with an increase in side chain bulk (D148G
< D148A < D148L < D148W).
G241 is located on a-helix number 9. Its mutation to Arg is expected to affect the
local helical structure driven by the increase in side chain bulk and basicity. To test the
effects of increase in side chain bulk and RNA catalysis, we can test the mutants with a
gradual increase in side chain size (G241A < G241V < G241L < G241F). Also, the effects of
basicity can be tested by testing mutants with (G241K or G241H) and compared to acidic
residues (G241D or G241E).
G306 is closely located to a critical residue H309 and its Gly to Ala mutation may
perturb the local structure sterically through a gain of -CH3 group. The effect of G306A has
not been described thus far, and by testing the presence charge (acidic and basic) and the
increase in side chain bulk, we could gain more knowledge into its role in maintaining the
local structure. To test the effects of charge, one can generate G306D, G306E, G306H,

134

G306R, and G306K. To test the effects of side chain size, one can generate mutants of
increasing side chain size (G306V < G306L < G306F < G306W).
In the future, assessing the effects of D148E, G241R, or G306A versus R237A in
cultured cells would produce interesting results since the first group showed reductions in
RNA cleavage but functional DNA repair activity and the R237A showed abrogation in both
of these activities. The implications drawn from these biological studies may reveal a novel
link between reductions in APE1 RNA cleavage activity and human diseases.
5.5 Role of metal ions in APE1 RNA-cleaving activity
Many nucleases that cleave DNA or RNA utilize metal ions for their catalysis (Wang
2008). Against a standard abasic DNA substrate, APE1 requires divalent metal ions for its
optimal activity (Erzberger and Wilson 1999; Wilson 2005). It is reported that these metal
ions occupy the active site to stabilize the transition state during DNA incision, and facilitate
the release of the incised product. Previously, it was found that Mg2+, Mn2+, Ni2+, and Zn2+
was able to rescue an EDTA-inactivated APE1 incision activity on an abasic DNA substrate,
but Ca2+ could not (Barzilay et al. 1995). Another study found Ni2+ and Zn2+ ions permitted
the DNA incision activity up to concentrations of 100 uM (Ni2+) and 50 uM (Zn2+) (Wang et
al. 2006). These reports suggest that APE1 RNA-cleaving activity may also require divalent
metal ions for its catalysis.
In comparison with these reports, our results showed a similar trend (refer to section
2.2.7). APE1 was able to cleave RNA without the presence of divalent metal ions and
showed a range of [Mg2+] for its optimal activity in our experimental setting. The results
indicated that APE1 RNA cleaving activity is best enhanced at 5 mM of Mg2+, but other
concentrations also allowed incision to take place. The observable RNA cleaving activity of

135

APE1 at 0 mM Mg + is in agreement with previous reports. APE1 has previously been found
to weakly cleave abasic DNA in the presence of EDTA (Masuda et al. 1998), and
particularly has exhibited EDTA-resistant activity towards acyclic substrates that are
considered to have better flexibility (Erzberger et al. 1999). Single stranded RNA structures
(e.g. hairpin loops) are particularly more flexible than those of RNA stems and abasic DNAs,
which could allow APE1 to cleave such RNA substrates without their divalent metal
cofactors.
APE1 was inhibited by 2 mM of Ni2+, and Zn2+ which was in contrast to the reports
on abasic DNA incision that was active even under 100 uM (Ni2+) and 50 uM (Zn2+) (Wang
et al. 2006). These two metal ions have been shown to interact with amino acid histidine in
other proteins to inhibit their enzymatic activities (Okamura et al. 2003; Kang et al. 2007).
Presence of these metal ions may therefore affect the amino acid residues that are specifically
critical for RNA catalysis of APE1. Alternatively, these metal ions can affect the secondary
structure of RNA substrates and interfere with APE1 binding to these RNAs or interfere with
their release after incision. Further testing by EMSA in the presence of these metal ions may
clarify such questions.
5.6 Role of N-terminus in APE1
Although no experimentation was done in this research on elucidating the role of Nterminus in APE1, it may be still noteworthy to mention some of the previous findings in
better understanding the APE 1-RNA interaction. Recently, co-immunoprecipitation of WT
APE1 had shown APE1 binding to the endogenous rRNAs (47S, 28S, and 18S) in HeLa
cells. Interestingly, the N&33 APE1 deletion mutant had lower ability to bind to rRNAs,
indicating the important role of N-terminus in APE1 RNA binding to rRNAs (Vascotto et al.

136

2009). In vitro EMSA study showed that N&33 APE1 had significantly less ability to bind to
single stranded 34nt RNA compared to the WT at tested enzyme amounts of 0.9 and 1.8 ug.
In addition, endonuclease assays showed the abasic RNA incision activity of Na33 APE1
was weaker than that of the WT. Interestingly, abasic DNA incision activity of N4.33 APE1
was stronger than that of the WT. This indicated that N-terminus is important in RNA
binding ability of APE1. However, it is still unknown, how much is contributed from these
N-terminal residues and the residues from the five APE1 DNA-binding regions (Mol et al.
2000) in RNA-APE1 interaction. Upon assessing their contributions in vitro, the role of these
residues can be further pursued in cells for their effects in RNA decay.
In regards to APEl-protein interaction, it was found that the first 33 N-terminal
amino acids of APE1 are essential for stabilizing the interaction of APE1 with ribosomal and
RNA processing factors such as, NPM1, pre-mRNA-processing factor 19 (PRP19), and 60S
acidic ribosomal protein P0 (Vascotto et al. 2009). In nuclear localization, APE1 was shown
to interact with nuclear importins as demonstrated by co-immunoprecipitation with
karyopherin al and a2. This interaction was found to require the first 20 residues of APE1
(Jackson et al. 2005).
It was found that N.&20 APE1 was re-distributed to the cytoplasm (Jackson et al.
2005). This indicated a presence of nuclear localization signal (NLS) within the 20 residues.
Further, it was found that APE1 possessed two independent NLS with first signal present at
residues 1 to 7 and the second at 8-13. For a significant cytosolic re-distribution, both of
these signals need to be deleted/mutated as demonstrated by visualizing N47-E12A/D13A
APE1 in the cytoplasm. In addition, treating cells with leptomycin B inhibited the nuclear
export of N47-E12A/D13A APE1. Leptomycin B is a common inhibitor for nuclear export

137

protein, CRM1 which was shown to interact with APE1, indicating that APE1 may possess a
nuclear export signal. However, further investigation is required to confirm this hypothesis.
One study reported significant re-distribution of APE1 to the cytoplasm upon RNase
treatment of permeable cells pre-treated with Triton X-100 (Vascotto et al. 2009). This
suggested that the RNA binding was in part a mechanistic factor in APE1 nuclear
localization. This hypothesis may explain NA33 APE1, with presumably less RNA binding
ability, to be predominantly redistributed to the cytoplasm (Chattopadhyay et al. 2006). This
indicated that the RNA binding ability of APE1 is important for its localization in the
cytoplasm and the role of N-terminus may act as an important functional determinant in
different cell compartments. However, it is unclear the mechanistic detail of how this RNA
binding ability of APE1 becomes utilized in nuclear localization of APE1.
In vitro assay showed that APE1 was preferentially ubiquitinated at residues K24 and
K25, and somewhat less at K27. In addition, nuclear export was observed with a fusion
protein, APEl(l-23)-ubiquitin (G76A)-APE 1(24-318). Also, independent of CRM1, a
nuclear exportin, APE1 can be exported out of the nucleus by nitrosylation at residues C93
and C310(Quetal. 2007).
Hence, the role of the first 33 amino acids constituting the N-terminus is still unclear.
APE1 has been shown to be redistributed across nucleus and cytoplasm depending on the
cellular conditions and cell types (Tell et al. 2005). In contrast to the aforementioned results
on NA33 APE1, a similar truncated mutant N&34 APE1 was found to have increased 3'-5'
DNA exonuclease activities that carries out chromatin fragmentation in apoptotic HL-60
cells which also showed predominant nuclear localization (Yoshida et al. 2003). Hence,

138

further characterization on the role of N-terminal residues in RNA binding/cleaving activities
and how it relates to the subcellular localization need to be investigated.
5.7 RNA secondary structure and sequences preferentially cleaved by APE1
The second objective of this research was to establish the RNA structure and
sequences cleaved by APE1. We found that APE1 can cleave multiple RNAs when presented
in vitro (refer to section 3.2.2). APE1 cleavage sites were predominantly located in the single
stranded regions of the RNA substrate with dinucleotide sequences of UA, UG, and CA. In
addition, new cleavage sites were found though very weak compared to the aforementioned
sequences and they were UC, CU, AC, and AU.
To account for the preferred cleavages of single stranded regions or weak stems of
RNA, we have to consider how APE1 recognizes its main substrate, the abasic DNA. In
abasic DNA cleavage, APE1 interacts strongly with an abasic deoxyribose flipped out of the
DNA helix and tightly binds to this moiety facilitated by a hydrophobic pocket composed of
Phe 266, Trp 280, and Leu 282. Such tight packing expels the binding of regular nucleotides
and gives specificity towards the abasic sites (Mol et al. 2000). Similarly in RNA cleavage,
we speculate that APE1 can recognize some of these extra helical bases found in the single
stranded regions or in the weak stems of RNA. It is because the individual RNA nucleotides
in these 'flexible' regions may act similarly in conformational or chemical context to the
abasic nucleotide. It was found that single stranded regions of RNA as in the case of RNA
hairpin loops have their bases flipping outwards from the centre of the loop which allow
RNA binding proteins to easily interact (De Guzman et al. 1998). Such similarity may
account for the preferential activity of APE1 for RNA structures that have weak stems or
single stranded regions. However, RNA nucleotides with normal bases retain polarities, and

139

we expect their interactions with the hydrophobic pocket to be weak and, hence, should
explain the strong preference of APE1 for abasic DNAs over RNAs. It is noteworthy that
F266A was shown to have severe reduction in endoribonuclease activity only (Figure 10A,
section 2.2.2), but not its phosphodiesterase activity (Hadi et al. 2002). This also supported
the proposal that F266 being in the hydrophobic pocket is a critical component in
endoribonuclease activity of APE1 by recognizing RNA bases and accounts for its preference
to cleave in the single stranded regions or weak stems of RNA. Future studies can confirm
the role of this hydrophobic pocket in RNA binding and cleavage by generating Ala mutants
of Trp 280 and Leu 282.
The residues of APE 1 responsible for conferring specificity for pyrimidines still need
to be identified. But, some insights can be drawn from the example of RNase A. In
recognition of its substrate, RNase A uses residue Thr 45 for recognition to cleave poly Cs
and poly Us. Such specificity is facilitated by specific hydrogen bonds between the side
chain -OH with -NH (pyrimidine) and the main chain -NH and =0 (pyrimidine) (Raines
1998). In APE1 RNA cleavage specificity, Thr 268 could serve a similar purpose, which is
located near the hydrophobic pocket of APE1 (Phe 266-Trp 280-Leu 282) for recognizing the
extra helical abasic deoxyribose (Mol et al. 2000) and it also may require further mutagenesis
studies to validate its role. Testing T268G or T268A may show indications of how this
residue plays its role in giving RNA cleavage specificities.
However, we should bear in mind the results from L104R and E126D giving altered
cleavage specificity. The positions of these two residues are also close or on the domain that
is known to interact with DNA (Mol et al. 2000). This indicates that the alteration in the
cleavage site can result from the perturbation of local APE1 structure that can also likely

140

affect the RNA binding ability of APE1 and can not be attributed solely to a specific putative
residue like Thr 268.
Pre-treatment of pre-miRNAs with APE1 significantly reduced the ability of DICER
to process these substrates in vitro (refer to section 3.2.4). This implies that APE1 could
potentially influence the level of mature miRNA in cells. Such speculation could be tested by
looking at the changes in mature miRNA levels after knocking down or over-expressing
APE1. Identification of interaction partners especially with the components of micro RNA
ribonucleoprotein complex or pri- or pre-miRNA processing machineries could reveal a
novel role of APE 1 in miRNA maturation pathway.
5.8 Assessment of biological significance of RNA-cleaving activity of APE1
There were a number of reports from the past that had speculation as to the identity of
the enzyme responsible for the endonucleolytic decay of c-myc mRNA (Swartwout and
Kinniburgh 1989; Wisdom and Lee 1991; Morello et al. 1993; Herrick and Ross 1994;
Ioannidis et al. 1996; Yeilding and Lee 1997; Hanson and Schoenberg 2001). Indeed, APE1
has been found to be present in the nucleus, cytoplasm, and mitochondria of the cell (Tell et
al. 2005; Chattopadhyay et al. 2006). However, for certain cell types such as macrophages,
spermatocytes, hippocampal cells, hepatocytes, hypoglossal motor neurons, and breast cells,
APE1 has been found to be prevalently localized in the cytoplasm (Evans et al. 2000).
Indeed, redistribution of APE1 into the cytoplasm raised further questions in elucidating its
role in that compartment. Previously in our lab, APE1 had been purified from rat liver
polysomal fraction that gave a hint to its role in RNA metabolism (Bergstrom et al. 2006). In
addition, recent report showed that APE1 could cleave abasic RNA (Berquist et al. 2008) in
vitro and this was followed by a report on abasic RNA cleavage in cells (Vascotto et al.

141

2009). These evidence further support the role of APE 1 in RNA metabolism. Hence, the third
objective of this research was to assess the endoribonuclease activity of APE1 in biological
systems.
To investigate this matter, we observed the c-myc mRNA expression after APE1 has
been transiently knocked down by siRNA technology in HeLa cells (refer to section 4.2.1).
Our results revealed that there was an increase in the expression levels of c-myc
mRNA when APE 1 was transiently knocked down. On one note, this result contradicted
with an earlier report that in HeLa cell line, a conditional APE1 expression knockdown by
RNAi leads to a modest reduction in the levels of c-myc mRNA (Vascotto et al. 2009).
However, we speculate that the experimental condition in this particular study had APE1
knocked down for 10 days, and in this setting, unidentified compensatory mechanisms could
reverse the elevation of c-myc mRNA levels.
Hence, it was intriguing to speculate the causes behind the elevation in c-myc mRNA.
First, reduction in APE1 may have resulted in the attenuation of c-myc mRNA decay.
Second, reduction in APE1 may have led to a decrease in the transcriptional repressor of cmyc mRNA. In agreement with the first possibility, our results showed that APE1 knock
down in HeLa cells can enhance the stability of c-myc mRNA, indicating its direct
involvement in c-myc mRNA turnover (refer to section 4.2.3). The mechanistic pathway for
the second possibility is still unknown and requires further investigations. This was the first
report in cells that APE1 can act as a regulator of RNA stability. Microarray studies that can
provide information on the global changes of mRNA abundance for tens of thousands of
genes (Lockhart and Winzeler 2000) can reveal potential APE1 targeted mRNAs. These

142

mRNAs can be further looked at for their turnover in APE1 knocked down cells. Such
studies are expected to provide further insights into the role of APE1 in mRNA decay.
Nonetheless, the role of APE1 should not be limited to RNA decay. Its activity may
have other biological implications that require further studies. There was evidence that
suggested the likeliness of APE1 being involved in RNA processing. One of the initial
findings showed that APE1 can associate with hnRNP-L in cells (Kuninger et al. 2002), a
master regulator in pre-mRNA splicing which was also shown to control the stability of
human VEGF mRNA during hypoxia (Shih and Claffey 1999). Later, APE1 was also shown
to interact with YB-1, an RNA binding protein and a splicing factor (Stickeleref al. 2001;
Chattopadhyay et al. 2008), giving further support to the possibility of APE1 involved in
RNA processing.
Also, APE1 was also found to stably interact with NPM1, an RNA binding protein
involved in the RNA maturation and processing (Vascotto et al. 2009). From this study,
APE1 was found to strongly associate with rRNAs in HeLa cells and had the ability to cleave
abasic RNAs. Presence of NPM1 was inhibitory to this activity and was thought to instigate a
fine tuning of APE1 in the process of rRNA quality control.
Testing the effects of APE1 and its mutants in Origami cells revealed significant
deviations from our in vitro data (refer to section 4.2.4 & 4.2.5). This indicated that
observing the colony formation to assess the RNA-cleaving function of APE1 was not
feasible. The method was adopted from a study that had successfully tested the activities of
the WT versus the active site mutants of RNase A and Angiogenin (Smith and Raines 2006).
RNase A and Angiogenin are both part of a ribonuclease A superfamily that have protein
sizes < 20kDa. Their function is solely dedicated for hydrolyzing RNA and this is in contrast

143

to APE1 which is a larger protein capable of exerting multiple activities in cells including
DNA repair activities, redox activation of transcription factors, 3'-5' exonuclease,
phosphodiesterase, RNase H, and endoribonuclease activities (Tell et al. 2005; Barnes et al.
2009). Hence, APE1 may exert any of these functions in Origami cells making it uncertain
whether the effect on cell growth is solely attributed to its endoribonuclease activity. In
addition, the method of observing colony formation after transforming cells heavily depends
on the transformation efficiency, compromising the evaluation of the effect of protein
expression less clear. A possible way to overcome this problem would be to assess the
growth of these cells in solution supplemented by medium and IPTG. This way we could
monitor the growth of cells not depending on transformation efficiency but on the level of
protein production and its activity.
5.9 Concluding remarks
This research was designed to understand the three main aspects of the
endoribonuclease activity of APE1. First, we have confirmed the importance of catalytic
residues that are required for this activity. APE1 mutants having one of these essential
residues substituted with another amino acid displayed moderate to severe reductions in RNA
catalysis. From these results, it is likely that APE1 shares these residues for cleaving abasic
DNA as well as RNA, however, certain residues may play different roles during each
reaction. We have also documented the RNA cleaving activities of the APE1 variants found
in human populations. Interestingly, a few of these variants displayed severe reductions in
RNA incision in contrast to their functional abasic DNA endonuclease activities.
Hypothetic ally, an APE1 variant with functional DNA repair activity, but dysfunctional RNA
cleaving activity has implications for aberrant RNA processing and turnover. Biological

144

work in understanding the role of these variants in RNA catalysis may generate valuable
insights into understanding a few of their associations with cancers. We have also discovered
important biochemical properties of APE1 in its RNA catalysis. APE1 is active in RNA
cleaving without the presence of divalent metal ions and leaves RNA products with 3'
phosphate as well as suggested to implement an acid-base catalysis mechanism in cleaving
RNA. Second, APE1 is able to cleave multiple RNAs when presented in vitro and has the
preference to cleave in the single stranded regions or weak stems. It prefers to cleave after
pyrimidines and shows its strongest activity towards dinucleotides of UA, UG, and CA.
Further studies are required to identify the residue(s) in giving pyrimidine specificity. Third,
we have assessed the role of APE1 as the regulator of mRNA stability. Our results have
shown that the knock down of APE1 resulted in the elevation of c-myc mRNA and is directly
responsible for its enhanced stability. This finding warrants further studies on the regulation
of its ribonuclease activities in cells. The sub-cellular localization and expression levels of
APE1 were found to be altered by external stimuli (Tell 2005). These include cellular signals
initiated by oxidative stress, hypoxia, and ischemia. The relationship between the changes in
such stimuli and the ribonuclease activity of APE1 has not been investigated. In addition,
more studies are required to reveal how APE1 interacts with RNA binding proteins or
processing enzymes involved in pre-mRNA maturation and processing pathway. The
contribution of ribonuclease activity of APE1 in these interactions and how such activity
affects the mRNA processing or degradation still awaits further experimentation. Future
studies focused on the identification of APE 1-associated cellular RNAs and APE1interacting proteins involved in the post transcriptional controls or translation are expected to
generate further insights.

145

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