RNA Oxidative Damage and Ribosomal RNA Surveillance under Oxidative Stress
by
Min Liu
A Dissertation Submitted to the Faculty of
The Charles E. Schmidt College of Science
in Partial Fulfillment of the Requirements for the Degree of
Doctor of Philosophy
Florida Atlantic University
Boca Raton, FL
August 2012
Acknowledgements
The author wishes to express her sincere appreciation to her advisor,
Dr.Zhongwei Li, for his guidance during her research and study at Florida Atlantic
University. The author is grateful to her other committees, Dr. Kasirajan Ayyanathan, Dr.
David Binninger, Dr. Keith Brew, and Dr. Christopher M. Burns, for their time and
assistance throughout the graduate study. The author also would like to thank her
collegues, friends, and families for their support and encouragement over the years.
iii
Abstract
Author:
Min Liu
Title:
RNA Oxidative Damage and Ribosomal RNA Surveillance
under Oxidative Stress
Institution:
Florida Atlantic University
Dissertation Advisor:
Dr. Zhongwei Li
Degree:
Doctor of Philosophy
Year:
2012
We have studied oxidative damage of RNA, a major type of cellular
macromolecules. RNA is a primary target of reactive oxygen species (ROS). Under
oxidative stress, most nucleic acid damages in Escherichia coli (E.coli) are present in
RNA as shown by the high levels of 8-oxo-G, an oxidized form of guanine. Increased
RNA oxidation is closely correlated to cell death under oxidative stress. Surprisingly,
neither RNA structure nor association with proteins protects RNA from oxidation. When
E. coli cultures were treated with hydrogen peroxide (H2O2), 8-oxo-G forms at higher
levels in ribosomal RNA than in non-ribosomal RNA species. The preferential formation
of 8-oxo-G in ribosomal RNA is related to the high order structure of the RNA since
oxidation produces more 8-oxo-G in native RNA than in denatured RNA, and in a
iv
RNA:DNA duplex than in single-stranded RNA of the same sequence. H2O2-induced 8oxo-G in ribosomes is removed specifically depending on the activities of polynucleotide
phosphorylase (PNPase) and RNase R, two 3’ exoribonucleases capable of degrading
structured RNA. H2O2–treatment of E. coli cultures also causes rRNA degradation in a
dosage dependent manner. In cells lacking the RNA-degradation exoribonucleases,
RNase R, PNPase, and/or RNase II, rRNA fragments accumulated to a high level upon
H2O2-treatment. The pattern of the rRNA fragments suggested a specific rRNA
degradation pathway that is initiated by endonucleolytic cleavages of 16S and 23S rRNA
in the intact ribosomes or subunits of ribosomes, followed by the degradation of the
fragments
exonucleolytically.
Surprisingly,
none
of
the
known
specific
endoribonucleases, RNase E, G, or P, are involved in the initial cleavages of 16S rRNA.
Our results demonstrate a major role for RNA degradation in controlling oxidized RNA.
We have identified activities that may work in specific pathways for selectively
degrading damaged RNA. These activities may play pivotal role in controlling oxidized
RNA and protecting cells under oxidative stress.
v
RNA Oxidative Damage and Ribosomal RNA Surveillance under Oxidative Stress
List of Tables ...................................................................................................................... x
List of Figures .................................................................................................................... xi
1. Introduction ..................................................................................................................... 1
1.1 RNA turnover........................................................................................................... 3
1.2 RNA quality control ................................................................................................. 9
1.2.1 RNA quality control in eukaryotes. .............................................................. 10
1.2.2 RNA quality control in prokaryotes. ............................................................. 14
1.3 RNA oxidative damage and quality control........................................................... 18
1.3.1 Reactive oxygen species and RNA oxidation ............................................... 18
1.3.2 Deleterious effect of RNA oxidation to its function and the detection......... 22
1.3.3 Potential physiological and pathological implications of RNA oxidation.... 23
1.3.4 Control of oxidized RNA .............................................................................. 25
1.4 Hypothesis and approaches of this study ............................................................... 29
2. Materials and Methods ................................................................................................. 31
2.1 Materials ................................................................................................................ 31
2.2 Strains and growth condition ................................................................................. 32
2.3 Mutant strains construction .................................................................................... 33
2.4 Treatment of E. coli cultures with H2O2 ................................................................ 33
vi
2.5 Isolation of RNA and DNA ................................................................................... 34
2.6 Isolation of ribosomal and non-ribosomal RNA .................................................... 35
2.7 Separation of long and short RNA species ............................................................ 36
2.8 Determination of 8-oxo-G level in RNA and 8-oxo-dG level in DNA by
HPLC ..................................................................................................................... 36
2.9 RNA denaturation and oxidation in vitro............................................................... 37
2.10 Preparation of oligomer single-stranded RNA and RNA:DNA duplex............... 37
2.11 Determination of copper binding capacity........................................................... 37
2.12 Determination of rRNA fragmentation ................................................................ 39
2.13 Determination of the 5’ end of rRNA fragments by primer extension ................ 40
2.14 Determination of the 3’ end of rRNA fragments by 3’ RACE ............................ 41
2.15 Effect of H2O2 on the growth of wild-type and mutant cells ............................... 42
2.16 Determination of cell viability ............................................................................. 42
3. Results. .......................................................................................................................... 44
3.1 Characterization of RNA damage under oxidative stress in Escherichia coli. ...... 44
3.1.1 H2O2 causes a quick and dosage-dependent increase of 8-oxo-G in
cellular RNA ................................................................................................. 46
3.1.2 H2O2 induces higher levels of 8-hydroxyguanine in RNA than in DNA. .... 46
3.1.3 The distribution of 8-oxo-G in various RNA species. .................................. 48
3.1.4 Highly folded structure does not protect RNA from being oxidized
in vitro . ......................................................................................................... 51
3.1.5 RNA fragmentation upon H2O2 treatment . .................................................. 53
vii
3.1.6 Cell death in response to H2O2 challenge .................................................... 54
3.1.7 Discussion. .................................................................................................... 55
3.2 RNA structures promote the formation of 8-hydroxyguanosine ........................... 62
3.2.1 H2O2 induces higher levels of 8-oxo-G in ribosomal RNA than in nonribosomal RNA ............................................................................................. 63
3.2.2 Oxidation of rRNA and tRNA is inversely correlated to the extent of
denaturation.................................................................................................... 65
3.2.3 H2O2–treatment generated less 8-oxo-G in single-stranded RNA than in
RNA:DNA duplex ........................................................................................ 67
3.2.4 Cu2+ bound different forms of nucleic acids with different affinity ............. 70
3.2.5 Discussion ..................................................................................................... 72
3.3 Exoribonucleases play an important role in eliminating oxidized RNA in
ribosome.................................................................................................................. 73
3.3.1 H2O2–induced ribosomal 8-oxo-G level decrease after removal of the
oxidant.......................................................................................................... 73
3.3.2 Specific removal of 8-oxo-G is blocked by deficiency in RNA
degradation ................................................................................................... 74
3.4 Degradation of 16S and 23S ribosomal RNA under oxidative stress in
Escherichia coli. .................................................................................................... 78
3.4.1 rRNA fragments are detected in H2O2-challenged E. coli cells.................... 81
3.4.2 The three processive exoribonucleases play a role in degrading rRNA
fragments under oxidative stress .................................................................. 83
viii
3.4.3 Degradation of 16S rRNA under oxidative stress......................................... 83
3.4.4 Degradation of 23S rRNA under oxidative stress......................................... 87
3.4.5 Pre-existing rRNA is degraded under oxidative stress ................................. 91
3.4.6 RNase PH is not required for the initiation of rRNA degradation under
oxidative stress .............................................................................................. 94
3.4.7 RNase E and RNase G are responsible for the major endonucleolytic
cleavage of 23S rRNA under oxidative stress .............................................. 96
3.4.8 Discussion ..................................................................................................... 99
4. Conclusion .................................................................................................................. 109
Appendices ...................................................................................................................... 111
I. Identification of RNA-related proteins that protect cells under oxidative
stress ........................................................................................................................ 111
I.1 Selection of candidate proteins and construction of E. coli mutants
lacking the proteins ......................................................................................... 112
I.2 H2O2-sensitivity of E.coli mutants lacking RNA-related proteins .................. 113
I.3 The protective roles of RNases, RNA helicases, or poly(A)
polymerase under oxidative stress ................................................................. 116
I.4 Discussion........................................................................................................ 120
II. Supplementary Data .............................................................................................. 121
5. References ................................................................................................................... 125
ix
List of Tables
Table 1: Cell death upon H2O2 insult*.............................................................................. 57
Table 2: Steady state levels of RNA oxidative damage in E. coli in response to
H2O2 treatment .................................................................................................... 58
Table 3: Summary of identified proteins that protect cells against oxidative stress ....... 115
x
List of Figures
Figure 1: RNA oxidative damage and cellular defense .................................................... 28
Figure 2: H2O2 treatment causes quick, dosage-dependent increase of 8-oxo-G
content in cellular RNA ..................................................................................... 47
Figure 3: H2O2 treatment causes a higher elevation of 8-oxo-G in RNA than that
of 8-oxo-dG in DNA .......................................................................................... 49
Figure 4: The levels of 8-oxo-G in various cellular RNA species under normal
conditions and in response to H2O2 treatment ................................................... 51
Figure 5: Native RNA structures do not protect RNA from H2O2 - mediated
oxidation in vitro ................................................................................................ 52
Figure 6: RNA fragmentation induced by oxidative stress ............................................... 53
Figure 7: H2O2 treatment causes a dose dependent growth reduction of E. coli .............. 56
Figure 8: H2O2 induces higher levels of 8-oxo-G in ribosomal RNA than in nonribosomal RNA in vivo and in vitro ................................................................... 64
Figure 9: Oxidation of rRNA and tRNA is inversely correlated to the extent of
denaturation........................................................................................................ 66
Figure 10: H2O2 induces higher levels of 8-oxo-G in RNA:DNA duplex than in
single-stranded RNA of the same sequence.................................................... 69
Figure 11: Cu2+ affinities for various nucleic acids .......................................................... 71
Figure 12: Proposed oxidized RNA quality control model. ............................................. 75
xi
Figure 13: The alteration of 8-oxo-G level in ribosomal and non-ribosomal RNA in
wide-type and mutant cells with continuous or pulse H2O2 treatment ........... 77
Figure 14: Accumulation of rRNA fragments at different time courses with various
concentrations of H2O2 ................................................................................... 82
Figure 15: Northern blot analysis of 16S rRNA fragments .............................................. 85
Figure 16: Northern blot analysis of 23S rRNA fragments .............................................. 90
Figure 17: Northern blot analysis of 16S rRNA in the wild type (wt) and mutant
cells with H2O2 and rifampicin treatments....................................................... 92
Figure 18: Northern blot analysis of 16S rRNA in the mutant rnb rnr rph pnpts ............. 95
Figure 19: Northern blot analysis of 16S rRNA in the control strain and mutants
lacking endoribonucleases RNase E and/or RNase G .................................... 97
Figure 20: Northern blot analysis of 16S rRNA in mutants lacking PNPase and
endoribonucleases ............................................................................................ 98
Figure 21: Northern blot analysis of 23S rRNA fragments in mutants lacking
endoribonucleases RNase E and/or RNase G ............................................... 100
Figure 22: Northern blot analysis of 23S rRNA fragments in mutants lacking
endoribonucleases ......................................................................................... 101
Figure 23: Hypersensitivity of E.coli cells to H2O2 treatment ........................................ 114
Figure 24: More sensitivity of combined mutation of RNase, RNA helicase RhlB,
or poly(A) polymerase under oxidative stress. .............................................. 117
Figure 25: More sensitivity of combined mutation of RNase and RNA helicase
under oxidative stress ..................................................................................... 119
xii
1. Introduction
Messenger RNA (mRNA), ribosomal RNA (rRNA), and transfer RNA (tRNA)
are three major types of cellular RNAs that are responsible for translating genetic
information from DNA into proteins. During gene expression, DNA sequences encoding
specific proteins are first transcribed into mRNA. The coding sequence in mRNA is then
translated into corresponding amino acid sequence utilizing ribosomes and tRNA. The
ribosome is composed of rRNAs and ribosomal proteins. tRNA recognizes codons in
mRNA and brings the appropriate amino acids to the growing peptide train.
In addition, many other RNA species have been discovered and they function in
various cellular processes. For instance, RNA enzymes catalyze reactions that process
RNA substrates. The RNA component of telomerase works as a template for the
synthesis of DNA telomeres. snRNA and snoRNA play important roles in splicing
(Valadkhan, 2005), and site-specific methylation and pseudouridylation of other RNAs
(Topkara and Holley, 2011), respectively. More recently, small interfering RNA (siRNA)
and micro-RNA (miRNA) have been unveiled that have been shown to regulate gene
expression at the post-transcriptional level by base-pairing with target mRNA sequences
(Bass, 2000; Topkara and Holley, 2011). In bacteria, tmRNA mediates a trans-translation
mechanism that rescues ribosomes that are stuck on non-stop mRNA (Keiler, 2008).
Numerous small, non-coding RNAs have recently been found in bacteria that regulate a
1
broad spectrum of biological activities through targeting mRNA species. Therefore,
production and maintenance of functional RNA species is of utmost importance for
normal gene expression and cell function. Essentially, all living organisms invest heavily
in RNA metabolism.
RNAs are synthesized as primary transcripts that undergo extensive processing
reactions before being functional. RNA processing includes removing extra sequences
from both 5’ and 3’ termini, splicing of introns, modification and assembly to multimolecular complexes. In eukaryotes, mRNA undergoes splicing, 5’ capping, and 3’
polyadenylation. In some cases, polyadenylation also occurs on other RNA species (Li et
al., 1998). For primary transcripts of rRNA and tRNA, the extra sequences at both 5’ and
3’ termini are removed by RNA cleavage reactions. Eukaryotic tRNAs are transcribed
with introns that are removed by specific endonucleases. In prokaryotes, the primary
transcripts of mRNA are generally translatable; however, polyadenylation can happen
posttranscriptionally. The primary transcripts of prokaryotic rRNA and tRNA undergo
processing to remove the extra sequences at the 5’ and 3’ termini. In all living organisms,
RNA molecules remain functional for various amounts of time, and they are eventually
broken down by RNA degradation activities in the process of normal RNA turnover.
rRNA and tRNA are relatively stable and they usually last multiple cell cycles. In
contrast, mRNA turnover is much faster. Controlling the decay rate of mRNA is an
important mechanism for regulation of gene expression.
2
RNA can be made with errors and become non-functional or even harmful. Such
RNA molecules can be generated by gene mutation, misincorporation of nucleotides,
misfolding, hypo- or hyper-modification, inappropriate editing and degradation, chemical
damage, etc. Such aberrant RNAs have to be identified and eliminated to maintain
normal cell activity. Various RNA surveillance mechanisms have been described.
However, the complexity of RNA quality control has only started to be revealed.
This work focuses on the RNA oxidative damage, and the related degradation of
RNA under normal and stress conditions.
1.1 RNA turnover
mRNA species have characteristic half-lives. In bacteria, the average half-life of
mRNA, approximately few minutes, depends on the needs of cells for specific proteins
and is in favor of adaption for cells to their quickly changing environment (Jain, 2002;
Melselson et al., 1964). The average half-life of mRNA in yeast is around 20 minutes,
while for most human mRNA, it is about 10 hours. The longer half-lives of mRNA in
higher organisms are consistent with their more stable cellular environment. rRNA
remains stable through the exponential growth phase of bacteria (Li and Deutscher, 2009)
and yeast (Cole and Lariviere, 2008). rRNA turns over slowly in L cells growing in tissue
culture. It was also reported that rRNA remains stable up to days in rat liver cells (Loeb
et al, 1965). Similar to rRNA, tRNA is generally stable. The half-life of endogenous
tRNA was reported to be approximately one day in SV3T3 mouse cells, two days in TC7
cells (an African green monkey kidney line) (Schlegel et al, 1978), 36 h in resting cells,
60 h in growing cells of mouse 3T6 cells (Abelson et al, 1974), and 53 h in Drosophila
3
cells (Lengyel and Penman, 1977). In actively growing bacteria cells, tRNA usually lasts
several cell cycles (Li et al, 2006). However, in Vibrio cholerae, a bacterium causing
cholera, tRNA undergoes rapid turnover with an average half-life of 11.8 min
(Mukhopadhyay, 1994).
mRNA decay in eukaryotes can be either deadenylation-dependent or
deadenylation-independent (Beelman and Parker, 1995). In the deadenylation-dependent
pathway, mRNA decay is initiated by shortening the poly (A) tail using deadenylase
activities, followed by removing the 5’ cap with decapping enzymes. 5’ decapping is
catalyzed by a protein consisting of two subunits, Dcp1p and Dcp2p. The decapped
mRNA is rapidly digested by the exoribonuclease Xrn1p, from the 5’-> 3’ direction,
without accumulation of degradation intermediates. Some deadenylated mRNA can be
degraded in the 3’->5’ direction. 3’ to 5’ degradation requires the exosome which is a
large complex of 3’ -5’ exoribonucleases (Van Hoof and Parker, 1999; Mitchell and
Tollervey, 2000; Parker and Song, 2004). In the deadenylation-independent decay, 5’
decapping occurs first and is followed by 5’ to 3’ degradation. Alternatively,
endonucleolytic activity cleaves the 3’ sequence, usually at the 3’ untranslated region.
The cleavage fragments could go through decapping and 5’ to 3’ degradation or 3’ to 5’
degradation directly.
In bacteria, mRNA decay usually involves multiple ribonucleases. In Escherichia
coli (E. coli), mRNA decay is primarily initiated by endonucleolytic cleavage, generally
performed by RNase E. Other endoribonucleases, such as RNase III, RNase G, and
RNase P, have limited activities on mRNA decay. Some endoribonuclease toxins, such as
4
RelE, MazF, and Kid, could also initiate mRNA degradation to turn off global translation
(Li and Deutscher, 2004). RNase E is a multidomain protein and usually cleaves in
unstructured, AU-rich regions (Deutscher, 2006). It was also reported to degrade RNA
substrates with 3’ poly (A) or poly (U) tail (Huang et al, 1998).
The RNA fragments generated by the initial cleavages are then digested by
exoribonucleases, mainly the polynucleotide phosphorylase (PNPase), RNase II, and
RNase R. While both RNase II and PNPase can be stalled by stable stem-loops in RNA
(Spickler and Mackie, 2000), due to the difficulties in attachment to the 3’ ends of RNA
(Jain, 2002), PNPase can be facilitated by poly (A) polymerase (PAP) which adds the
poly(A) tail to the RNA substrate helping PNPase to bind and continue the degradation.
In contrast to PNPase, RNase II may remove the poly(A) tail but cannot work through
structured regions in mRNA, which conversely protects mRNA from further degradation
(Mohanty and Kushner, 2003). RNase R is a universal degrader of structured RNA. It can
degrade extensively structured regions of mRNA (Cheng and Deutscher, 2005) if there is
an overhang of at least 7 nt present at the 3’ end (Vincent and Deutscher, 2006).
Degradation by these exoribonucleases results in final products in the form of
oligoribonucleotides that are 2-5 nt in length. These oligoribonucleotides are degraded to
single nucleotides by an essential enzyme oligoribonuclease (Ghosh and Deutscher,
1999).
Interestingly, RNase E recruits several proteins at its C-terminal domain to form a
large multienzyme complex, the degradosome. The complex is principally composed of
RNase E, PNPase, a RNA helicase RhlB, and a glycolytic enzyme enolase (Carpousis et
5
al, 1999). Other proteins found in the complex are DnaK, GroEL, and PPK, which are
less stably associated with degradosome (Miczak et al, 1996; Blum et al, 1997). The Cterminal region of RNase E interacts with RhlB, PNPase, and enolase, while the Nterminal region accomplishes the catalytic activity. When it binds with mRNA at the 5’
end, RNase E apparently scans downstream and cleaves at AU rich sequences. Instead of
the cleavage products leaving the complex, they can be degraded by PNPase. RhlB
facilitates the PNPase-mediated degradation by resolving the RNA structures. Any
additional roles of other components of the degradosome in RNA degradation have not
been determined yet. Despite the apparent relationship of degradosome with its proposed
function in RNA degradation, formation of such a complex is dispensable for the RNA
decay under normal conditions. The cleavage activity of RNase E is enhanced by the
monophosphorylated 5’ end of the RNA. The conversion of a triphosphated 5’ end to a
monophosphated one is performed by a protein, RppH, also called NudH/YgdP. RppH
initiates mRNA degradation by removing the pyrophosphate from the 5’ end of
triphosphorylated RNA (Deana et al., 2008). Its activity is suspended when a stem-loop is
present at the 5’ end of RNA.
In Bacillus subtilis, neither RNase E nor its homolog is present. mRNA
degradation is taken over by two other essential ribonucleases, RNase Y and RNase J.
RNase Y is an endoribonuclease associated with the membrane, and RNase J has both
endonuclease and 5’ exonuclease activities (Richards et al., 2011). To initiate mRNA
decay, the triphosphorylated 5’ termini of mRNA is converted into monophosphorylated
ends by the protein BsRppH, and then further degraded by RNase J whose activity is
6
monophosphate-dependent. RNase Y also preferentially degrades RNAs with a single
phosphate at the 5’ end. Moreover, RNase Y is important for S-adenosylmethionine
(SAM)-dependent riboswitch RNA turnover (Shahbabian et al., 2009).
It is interesting to note that polyadenylation of mRNA in eukaryotes is usually
implicated in stability, maturation, nuclear transportation etc. However, it signals
degradation in prokaryotes, archaea, mitochondria and chloroplast (Abernathy et al.,
2009). The polyadenylation of rRNA, reported in humans, yeasts, and protozoa, could
initiate the degradation, but not stability, because truncated polyadenylated rRNA
transcripts were detected (Kuai et al., 2004; Slomovic et al., 2006, Abernathy et al.,
2009).
Although rRNA is usually long-lived, its stability may vary depending on growth
conditions of the cells (Deutscher, 2003). It is well established that bulk rRNA is
degraded under starvation of phosphate (Maruyama and Mizuno, 1970), nitrogen (Ben
Hamida and Schlessinger, 1966), glucose (Jacobson and Gillespie, 1968), or a carbon
source (Kaplan and Apirion, 1975). When Salmonella cells reach the stationary phase,
more than 90% of 23S rRNA and ~50% of 16S rRNA are degraded (Hsu et al., 1994). In
E. coli strains, portions of newly synthesized rRNA were degraded after nutritionally
downshifting for 30 min. At very slow growth rates, 70% of the newly synthesized rRNA
was degraded (Deutscher, 2003).
Mutations on rRNA also alter rRNA stability
(Deutscher, 2003). rRNA operons with mutated leader sequences influence the synthesis
of mature 16S rRNA and 30S subunit formation and eventually 16S rRNA stability,
although the leader region is removed during maturation. RNAs with deleted termini
7
cannot form stable ribonucleoprotein (RNP) particles and are eventually degraded.
Moreover, many agents influence RNA degradation, such as streptomycin, mitomycin C,
polymixin E, toluene, dodecyldiethanolamine, Hg2+ ions and so on. These agents may
affect the cell membrane and alter permeability, which leads to the ionic changes, and
cause ribosomal structure alterations. As a result, rRNA becomes more accessible to the
degradative RNases (Deutscher, 2003). During starvation of E. coli cells, the
exoribonuclease RNase PH initiates rRNA degradation by shortening the 3’ end of 16S
rRNA, rendering the ribosome non-functional in translation. This may be followed by
endonucleolytic cleavages that generate rRNA fragments. The rRNA fragments are then
removed by RNase R and PNPase (Cheng and Deutscher, 2003, Basturea et al., 2011).
Normal tRNA is usually quite stable. Even after UV irradiation or during a
productive bacteriophage lambda infection that leads to cell death, tRNA molecules
remain intact (King et al, 1986). The stability of tRNA may be due to its extensive
secondary and tertiary structure and aminoacylation at the 3’ end. The high order
structure of tRNA may confer resistance to nuclease, since tRNAs with specific
mutations affecting their secondary structure were degraded (Smith, 1974; Li et al.,
2002). Moreover, transient association with aminoacyl-tRNA synthetases, elongation
factor, and ribosomes may also protect tRNA from ribonuclease activities. Nevertheless,
in several organisms, tRNA is cleaved in the anticodon loop under specific growth
conditions (Phizicky and Hopper, 2010). tRNA cleavage in the anticodon region was
reported in soil bacterium Streptomyces coelicolor under starvation through an unknown
mechanism (Haiser et al., 2007). In a protozoan, Tetrahymena thermophila, starvation
8
induces cleavage of mature tRNA in the anticodon loop region, and/or in the variable arm
(Lee and Collins, 2005; Phizicky and Hopper, 2010). Cleavage in the anticodon loop of
tRNA was also found in Aspergillus fumigatus (Jöchl et al., 2008). The accumulated
tRNA halves lack 3’ CCA residues that are required for aminoacylation, indicating that
the tRNA was deacylated before being cleaved, and then the 3’ termini could be removed
since the uncharged tRNA is more susceptible to CCA loss (Lee and Collins, 2005). This
cleavage was inhibited when essential amino acids were added.
Cleaved tRNAs were also identified in human cells, although the mechanism is
unknown (Kawaji et al., 2008). tRNA fragments were detected in the urine and sera of
cancer patients, and the level increased as the tumor burden increased (Borek et al., 1977;
Thompson and Parker, 2009). However, the activity for tRNA turnover has not been
identified. In E. coli, several molecules are responsible for the specific tRNA cleavage
(Haiser et al., 2007; Tomita et al., 2000; Ogawa et al., 1999). Colicin D cleaves tRNAArg
in the anticodon loop, while colicin E5 cuts tRNATyr, tRNAHis, tRNAAsn, and tRNAAsp. In
addition, E3 also cleaves 16S rRNA in the 70S ribosome at the 49th phosphodiester bond
from the 3’ end in E. coli (Masaki and Ogawa, 2002). Polyadenylation of normal tRNA
may occur on hypo- or hyper-modified tRNA, which may be related with the degradation
of the defective tRNA (Kadaba et al., 2004).
1.2 RNA quality control
Although there are many factors in RNA metabolism that ensure the formation of
normal RNA, aberrant RNAs are always generated in the lifetime of cells, which could be
caused by environmental stress, nutrient alterations, misincorporation of nucleotides
9
during transcription, mistakes in processing, modification, gene mutation etc. Therefore,
quality control of RNA in both eukaryotes and prokaryotes is required to eliminate the
defective RNA.
1.2.1 RNA quality control in eukaryotes.
mRNA
In eukaryotes, mRNA molecules that are incorrectly processed may contain stop
codons within the normal coding region (non-sense mRNA), or may lack stop codon
(non-stop mRNA). These aberrant mRNAs must be degraded quickly to avoid the
accumulation of abnormal protein products. Specific mechanisms have evolved in the cell
to recognize and remove these defective mRNAs. Nonsense-mediated mRNA decay
(NMD) degrades mutated or defective pre-processing mRNA, such as mRNA harboring a
premature termination codon (Baker and Parker, 2004).
Generally, terminal codons are present in the last exon. If the stop codons are
followed by a downstream intron longer than 50-55 nt from the exon-exon junction closer
to the 3’ end, they are called premature termination codons (PTC) and trigger NMD.
After splicing, a protein complex, called the exon-junction complex (EJC), stays 20-24 nt
upstream of exon-exon junction (Le Hir et al., 2000 a; 2000b). The ribosome displaces
the EJCs as it passes in the 5’ to 3’ direction on mRNA during translation. However,
when the ribosome reaches a PTC, it stops and is unable to displace the EJC present
downstream of the PTC. As a result, the EJC proteins, Y14 and Upf3, are recruited on the
undisplaced EJC and trigger NMD, indicating that the EJC acts as a signal for NMD
10
during translation (Baker and Parker, 2004). However, in several cases in mammalian
genes, the aberrant mRNA is not defined by an EJC (Maquat, 2004). For instance, the Tcell receptor (TCR)-β transcript is degraded by NMD, although the distance between the
PTC and the downstream exon-exon junction is less than 50-55 nt. A PTC within βglobin exon 1 does not trigger NMD, even when it is followed by an intron that is longer
than 50-55 nt.
In the above mentioned cases, mRNA turnover begins with a shortening of the 3’
poly (A) tail, followed by 5’ decapping/5’-3’ degradation or 3’-5’ degradation. In yeast,
NMD triggers rapid shortening of the 3’ poly(A) tail and accelerates the 5’ decapping,
and then aberrant mRNA is degraded by a deadenylation-independent pathway. However,
if decapping and 5’-3’ degradation are unable to proceed, NMD accelerates the 3’-5’
degradation (Baker and Parker, 2004).
Non-stop mRNAs have no termination codon. During translation, the ribosome
stalls at the 3’ end of the mRNA and a poly (A) tail is added, which triggers non-stop
decay (NSD).
Subsequently the dead-end templates are degraded by NSD and the
associated ribosomes are released for the translation of other mRNAs (Isken and Maquat,
2007). To degrade non-stop mRNA, the empty A site of the 80S ribosome is recognized
and bound by an exosome-associated protein, Ski7p, which brings the exosome to the 3’
end of the mRNA and then the exosome degrades this mRNA from 3’-5’ direction (Isken
and Maquat, 2007; Van Hoof et al., 2002). Nonstop mRNA decay (NSD) is also triggered
when transcription aborts (Isken and Maquat, 2007). However, if normal termination
codons are mutated or translating ribosomes fail to recognize the normal termination
11
codons since 3’ UTRs of mRNA contain many other in-frame termination codons, NSD
will not be initiated.
Another degradation pathway of mRNA is called No-go mRNA decay (NGD).
This occurs when mRNA translation stalls, and the endonucleolytic cleavage is close to
the stalled ribosome. Elongation pause could be due to a strong RNA structure such as
the stem-loop, rare codons within the open reading frame (ORF) when translation rates
are slow, or to premature stop codons (Passos et al., 2009; Doma, 2008). In yeast cells,
translation elongation discontinues when an active mRNA harbors a stem-loop structure.
In NGD, the 5’-cleavage product generated by the initial endonucleolytic cleavage is
degraded by the exosome, and the 3’-cleavage product is degraded by Xrn1 (Isken and
Maquat, 2007).
mRNA can also be degraded by processing bodies (P-bodies), which are protein
complexes composed of mRNA decapping and degradation enzymes (Hillebrand et al.,
2007; Parker and Sheth, 2007). Some proteins of nonsense-mediated decay (NMD) are
also observed in P-bodies under stress or overexpression conditions (Parker and Sheth,
2007). mRNA is believed to be degraded in P-bodies (Eulalio et al., 2007; Parker and
Sheth, 2007). However, mRNA can also return to translation from the P-body, when
growth conditions change (Parker and Sheth, 2007).
rRNA
The exosome degrades improperly processed pre-rRNA intermediates in
eukaryotes (Allmang et al., 2000). In yeast, the elimination of mature rRNA containing
12
point mutations in positions important for translation has been detected (LaReviere et al.,
2006). Mature rRNA in ribosome subunits that have a functional problem after
transcription and pre-rRNA processing are removed by nonfunctional rRNA decay
(NRD) (Cole and LaRiviere, 2008). NRD can be divided into two mechanisms; one is
responsible for rRNAs mutated in a decoding site (18S NRD), and another removes
rRNA with mutations in the peptidyl transferase center (25S NRD) (Cole et al., 2009).
18S NRD is a cytoplasmic process and requires ongoing translation elongation. The
essential enzymes mediating 18S NRD include the major cytoplasmic 5’-3’ exonuclease
Xrn1p, the core exosome, Ski7p, Hbs1p, and Dom34p, which also play roles in NGD
mRNA quality control. In contrast to 18S NRD, 25S NRD is not related to any known
translation-dependent mRNA decay pathways, and has no mRNA decay factors
participating, except the core exosome exonuclease, Rrp44p. It could be a late quality
control mechanism to prevent 60S subunits that are functionally defective in the peptidyl
transferase center from interfering with normal protein synthesis, since 25S NRD
substrates cosediment with 60S ribosomal subunits, but not with monosome or polysome
fractions.
25S and 5.8S rRNA are significantly degraded in yeast which undergos
programmed cell death (PCD) (Mroczek and Kufel, 2008). PCD, also called apoptosis, is
trigerred by hydrogen peroxide, acetic acid, hyperosmotic stress (60% glucose), and
ageing. The cleavage of 28S rRNA during apoptosis was reported in metazoans (Degen et
al., 2000). It was observed that rRNA decay was corresponded to the level of ROS
(Mroczek and Kufel, 2008).
13
tRNA
Aberrantly modified tRNA is degraded rapidly in tumor cells, although the
mechanism for eliminating this tRNA is unclear (Borek et al., 1977). Fully modified
tRNAs are more thermally stable than completely unmodified tRNA transcripts
(Chernyakov et al., 2008). In yeast, mature tRNA has to be modified properly. Otherwise,
specific mechanisms proceed to remove the defective tRNAs. It was reported that a
hypomethylated pre-tRNAiMet is polyadenylated by Trf4p, a DNA polymerase with poly
(A) polymerase activity, and is degraded by Rrp6 and the nuclear exosome (Kadaba et
al., 2004; 2006). This pathway also targets a truncated 5S rRNA, aberrant rRNA
processing intermediate, and pre-rRNAs that accumulate in the nucleoli due to blocked
export of ribosomal subunits (Reinisch and Wolin, 2007).
tRNA lacking m7G and m5C modifications is quickly degraded by a mechanism
that does not require Trf4p or the exosome (Alexandrov et al., 2006). This mechanism is
called rapid tRNA decay (RTD) and involves 5’-3’ exonucleases Rat1, Xrn1, and Met22
(Chernyakov, 2008). It can also degrade multiple mature tRNA species lacking various
combinations of modifications.
1.2.2 RNA quality control in prokaryotes.
mRNA
In prokaryotes, the trans-translation surveillance mechanism for degradation of
non-stop mRNA has been well studied. The key factor in this pathway is tmRNA, also
called ssrA RNA or 10Sa RNA, which has the properties of both tRNA and mRNA.
14
During translation of non-stop mRNA and other aberrant mRNA molecules, ribosomes
stuck on the mRNA due to the absence of proper mechanism to terminate the translation.
tmRNA rescues the unproductive stalled ribosome and releases the aberrant mRNAs
which are then degraded by RNases. The polypeptides produced from the aberrant
mRNAs are also released from ribosomes and subsequently degraded (Yamamoto et al.,
2003; Dulebohn et al., 2007).
tmRNA has mRNA-like and tRNA-like domains. The 5’ and 3’ ends of tmRNA
are tRNA-like domains having an acceptor stem, a T arm, and a D-loop with no stem.
This domain is linked with the rest of the tmRNA molecule by a long disrupted stem, and
charged with alanine by alanyl-tRNA synthetase (Komine et al., 1994; Dulebohn et al.,
2007). The mRNA-like-domain of tmRNA is an open reading frame (ORF), encoding a
peptide of 10 amino acids in length and ending with a stop codon. The alanine-charged
tmRNA rescues the stalled ribosome by performing like a tRNA in the first step. It
recognizes stalled ribosomes, binds to the empty A-site, and transfers its charged alanine
to the nascent polypeptide chain. Then it acts as an mRNA that replaces the defective
mRNA with its encoded open reading frame (Keiler et al., 1996; Dulebohn, et al., 2007).
Therefore, the ribosome continues the translation until it reaches the stop codon in the
mRNA-like-domain. Translation is terminated normally at the stop codon on tmRNA,
releasing peptides with this additional 11 amino acid tag. The tagged peptides are
recognized by C-terminal specific cellular proteases, like ClpX and ClpA, (Dulebohn, et
al., 2007). In E. coli, the tag sequence was identified as AANDENYALAA in several
expressed foreign proteins that produced truncated peptides (Tu et al., 1995). These
15
tagged peptides were not produced in an E. coli mutant that had a disrupted ssrA gene.
tmRNA is widely present in most bacteria. In most cases, a small protein, SmpB, forms
complex with tmRNA.
SmpB stabilizes tmRNA and facilitate binding of stalled
ribosomes.
Degradation of non-stop mRNA released by trans-translation involves the
exoribonuclease RNase R (Richards et al., 2006; Ge et al., 2010), although other
activities for mRNA turnover may also be involved. This tmRNA-facilitated mRNA
decay prefers to degrade aberrant mRNAs which promotes ribosome stalling. The Cterminal lysine-rich domain of RNase R interacts with the stalled ribosome associated
with SmpB and tmRNA.
rRNA
rRNA incorporated into ribosomes are stable under normal conditions. rRNAs
that are overexpressed or synthesized more than ribosomal proteins are
degraded,
suggesting that rRNA in ribosomes are protected by ribosomal proteins (Siehnel and
Morgan, 1985; Lewicki et al., 1993; Deutscher, 2003). Therefore, degradation of rRNA
molecules that are not completely or correctly assembled in ribosomes is an important
mechanism for rRNA quality control.
Emerging evidence suggests that in E. coli, a quality control mechanism initiates
rRNA degradation by yet unidentified endonucleolytic activity(ies) at defined positions.
The resulting rRNA fragments are degraded by RNase R and PNPase. In the absence of
PNPase and RNase R, 16S and 23S rRNA fragments accumulated, suggesting that these
16
two enzymes are responsible for the degradation of defective rRNAs (Cheng and
Deutscher, 2003). Consistent with the idea that incompletely assembled rRNA are
degraded by quality control mechanisms, it was reported that RNase R and PNPase are
responsible for degradation of only newly synthesized rRNA, but not rRNA in mature
ribosomes under conditions requiring rRNA quality control (Basturea et al., 2011). As
stated above, RNase R and PNPase may also be the key enzymes that degrade rRNA
fragments during rRNA turnover under conditions such as starvation.
In addition, polyadenylation may also play a role in the degradation of aberrant
rRNA. In the absence of sufficient RNA 3’ processing exoribonucleases, precursors of
stable RNAs including 5S and 23S rRNA are polyadenylated (Li et al., 1998). rRNA
fragments associated with the degradosome were degraded by the degradosome in vitro
(Bessarab et al., 1998), suggesting a possible role of this mRNA decay complex in rRNA
quality control.
tRNA
Aberrant tRNA may be eliminated by degradation. However, information about
prokaryotic tRNA quality control is relatively scarce. The precursor of a mutated tRNATrp
is rapidly degraded in E. coli (Li et al, 2002). It could be due to, but not limited to,
PNPase and PAP, since defective tRNAs are still degraded in the absence of PNPase,
indicating that other ribonucleases substitute for the activity of PNPase.
A quality control pathway of tRNA is reported for uncharged or unaminoacylated
tRNA. In this process, the A residue of the CCA sequence at the 3’ end of the tRNA was
17
removed by RNase T, and tRNA dissociates from RNase T, and finally repaired by
nucleotidyltransferase (Deutscher, 2003).
1.3 RNA oxidative damage and quality control
Although there has been considerable progress toward understanding the quality
control of aberrant RNAs that contain wrong sequences and structure, almost nothing is
known about RNAs that are chemically damaged. This is partly due to the ignorance of
the potential importance of RNA damage by chemicals. Furthermore, since it is believed
that damaged RNA can be eliminated by RNA turnover mechanisms, cells would never
need to make an additional investment in controlling chemically damaged RNA. We
argue that this is a misconception. As discussed below, RNA can be significantly
damaged under oxidative stress conditions. Oxidized RNA may be a threat to normal cell
function and cell viability, and specific mechanisms may be employed to eliminate such
RNA.
1.3.1 Reactive oxygen species and RNA oxidation
Oxidative stress is a common stress condition for the cell in various organisms. It
is related with many diseases, such as cancer, cardiovascular disease, Down’s syndrome,
Friedreich’s ataxia, rheumatoid arthritis, autoimmune disease, and AIDS (Temple et al.,
2005).
Reactive oxygen species, ROS, are byproducts of normal oxygen (O2)
metabolism, in which O2 receives electrons and becomes superoxide (O2-). It can also be
generated by ionizing and ultraviolet radiation, as well as exposure to carcinogens and
18
chemotherapeutic agents (Cooke et al., 2003, Bellacosa and Moss, 2003). ROS includes
oxygen free radicals, such as superoxide radical anion (O2●-), hydroxyl radical (●OH), and
non-radical oxidants, such as hydrogen peroxide (H2O2) and singlet oxygen (1O2). ROS is
a major source of damage to cellular components, such as DNA, RNA, proteins and
lipids. Essentially all living organisms have developed some antioxidant defenses to
remove ROS, such as superoxide dismutase (SOD), catalase, and glutathione peroxidase
(GP). SOD catalyses the conversion of superoxide radical O2-• into H2O2 (Oberley and
Buettner, 1979; Chaudière and Ferrari-Iliou, 1999). Catalase decomposes H2O2 into O2
and water, while GP reduces H2O2 into water and organic hydroperoxides into alcohols
by converting reduced glutathione (GSH) to oxidized glutathione (GSSG) (Maier and
Chan, 2002; Chaudière and Ferrari-Iliou, 1999). Such mechanisms greatly reduce the
level of ROS. On the other hand, genetic defects and environmental hazards may cause
oxidative stress characterized by increased production of ROS. Therefore, the level of
ROS that actually cause damage to macromolecules is determined by the both ROS
production and antioxidant activities. It has been widely accepted that increased damage
to macromolecules under oxidative stress is the major cause of cell cycle arrest and
finally cell death, which eventually may lead to cancer, accelerated ageing, and agerelated degenerative diseases (Mroczek and Kufel, 2008).
Although RNA has a greater chemical oxidative stability than DNA (Thorp,
2000), many reagents causing DNA damage also damage RNA. In DNA, more than 20
different types of oxidatively altered bases have been detected including 8hydroxyguanine,
8-hydroxyadenine,
2,6-diamino-4-hydroxy-5-formamidopyrimidine
19
(FapyGua) from guanine, 4,6-diamino-5-formamidopyrimidine from adenine, and
cytosine glycol (Cook et al., 2003; Li et al., 2006). Counterparts of these compounds
should also be produced in RNA by oxidative stress (Bellacosa and Moss, 2003). Among
these lesions, 8-hydroxydeoxyguanosine (8-oxo-dG) in DNA or 8-hydroxyguanosine (8oxo-G) in RNA is most deleterious (Ames and Gold, 1991), allowing them to be used as
markers of oxidation level in DNA or RNA. 8-oxo-G in RNA can mispair with adenine
(A) or thymine (T) at similar or higher efficiency than with cytosine (C), causing
nucleotide mis-incorporation during DNA and RNA synthesis (Taddei et al., 1997).
The level of 8-oxo-G can be determined by separating RNA nucleosides on
HPLC. This method is adopted from determination of 8-oxo-dG in DNA by HPLC-ECD,
in which deoxynucleosides are separated and 8-oxo-dG was detected by an
electrochemical detector (ECD) with in-line detection of dG by a UV detector. 8-oxo-dG
per 105 dG was calculated (Floyd et al., 1989). Similarly, 8-oxo-G and G in RNA can be
detected by ECD and UV detectors, respectively (Fiala et al., 1989; Shen et al., 2000).
The level of RNA oxidation can be presented as 8-oxo-G/105 G. 8-oxo-G can also be
detected by monoclonal antibodies (Nunomura et al., 1999a and 1999b; Shan et al.,
2003). This antibody-based analysis is especially useful for detecting 8-oxo-G in specific
regions of tissue samples or in a subpopulation of total RNA.
Other damage occurring in DNA may also happen in RNA, including
modification to other bases and ribose, base excision, and strand break. Therefore, under
both normal and oxidative stress conditions, the total damage in RNA must be much
higher than the detected 8-oxo-G level. Reverse transcription can be blocked by many
20
types of oxidative damage to an RNA template (Rhee et al., 1995). Increased blockage of
reverse transcription by oxidative damage of rRNA was detected (Gong et al., 2006).
Abasic site, a sugar moiety without its base, in DNA is induced chemically by DNA
damage or oxidizing agent such as alkylating agent or ionizing radiation. They are also
intermediates in the repair pathway initiated to eliminate oxidized bases by DNA Nglycosidases (Tanaka et al., 2011a and b). Abasic sites in DNA can be determined by an
aldehyde reactive probe (ARP). Interestingly, abasic sites in RNA were recently detected
also by ARP (Tanaka et al., 2011b). ARP reactivity was increased in RNA after in vitro
oxidation by Fenton reaction, γ-radiation or reactive nitrogen species. It also increased in
RNA under oxidative stress by H2O2 or peroxynitrite, suggesting that abasic site could be
used as RNA oxidation marker (Tanaka et al., 2011a).
Greater oxidation of RNA than DNA has been shown in cell lines and tissues,
such as human leukocytes (Shen et al., 2000), human skin fibroblasts (Wamer and Wei,
1997), human lung epithelial cells (Hofer et al., 2005), rat skeletal muscle (Hofer et al.,
2008), and rat liver (Fiala et al., 1989; Hofer et al., 2006). However, in contrast to DNA,
RNA oxidative damage has not attracted adequate attention yet.
Large amounts of oxidized mRNAs were detected in cells from Alzheimer disease
(AD) frontal cortex, and amyotrophic lateral sclerosis (ALS) post-mortem tissues (Shan
et al., 2003; Shan and Lin, 2006; Chang et al., 2008). The ROS-induced oxidation to
mRNA is not random but highly selective, and some mRNA species are more susceptible
to oxidative damage. More recently, it has been shown that various mRNA species in
21
yeast also contain different levels of 8-oxo-G using a global RNA analysis (McKinlay et
al., 2012).
The majority of 8-oxo-G detected in total RNA could be present in rRNA, since
rRNA is highly abundant, ~ 80% of cellular RNA, and turns over slowly. Indeed, rRNA
was shown to be predominately oxidized among various RNA species, presumably due to
higher redox iron binding to rRNA than to other RNA species (Ding et al., 2005; Honda
et al., 2005). When ROS level increases, 25S and 5.8S rRNAs in yeast were significantly
degraded, and specific intermediates accumulated (Mroczek and Kufel, 2008; Thompson
et al., 2008).
tRNA fragments resulting from endonucleolytic cleavage were detected in yeast,
plants, and mammalian cells under oxidative stress, especially during entry into
stationary phase. The fragmentation does not significantly reduce the pool of full-length
tRNAs in yeast or human cells (Thompson et al., 2008). However, compared to rRNA,
tRNA generated much less 8-oxo-G when exposed to oxidant (Honda et al., 2005).
1.3.2 Deleterious effect of RNA oxidation to its function and the detection
Oxidation has been shown to be highly disruptive of RNA function. Up to 50% of
mRNAs purified from AD frontal cortices are oxidized. Oxidized mRNA is not translated
efficiently, since the level of corresponding proteins is significantly decreased (Shan et
al., 2003 and 2007; Chang et al., 2008). The reduced protein expression is possibly
because the ribosomes slowed down or stalled on the oxidized bases in mRNA. Oxidized
mRNA also induces aggregated proteins (Shan et al., 2003), full-length proteins with no
22
or reduced activities, short peptides because of premature termination or translation errorinduced degradation (Tanaka et al., 2007), and cell death (Shan et al., 2007). It is
speculated that oxidized bases on mRNA alter pairing capacity with tRNA and produce
mutated proteins (Kong et al., 2008).
The oxidation of rRNA, the majority of total RNA, would be a serious problem
that may cause cell dysfunction (Nunomura et al., 2006). Oxidized rRNA becomes
nonfunctional in protein synthesis in Alzheimer Disease (Ding et al., 2005).
Accompanied with the ribosomal dysfunction, decreased rRNA and tRNA levels and
increased 8-oxo-G in the total RNA pool, especially in rRNA, are detected. The oxidation
damage of RNA related with decreased capacity of protein synthesis may be a contributor
of the onset and development of Alzheimer’s disease (Ding et al., 2005).
1.3.3 Potential physiological and pathological implications of RNA oxidation
Oxidative damage of RNA has been described in several neurological diseases,
suggesting many detrimental effects of RNA oxidation (Nunomura et al., 2006). Based
on HPLC-ECD analysis, the levels of 8-oxo-G in RNA from mammalian cell cultures or
tissues increased five to ten fold after oxidative stress, which is 10-25 times higher than
those of 8-oxo-dG in DNA under the same conditions (Hofer et al., 2005 and 2006; Shen
et al., 2000). Remarkable increases of 8-oxo-G levels in RNA were detected in patients
over a range of diseases, such as Parkinson’s disease (Zhang et al., 1999; Kikuchi et al.,
2002), Alzheimer’s disease (Nunomura et al., 1999; Abe et al., 2002; Ding et al., 2006),
dementia with Lewy bodies (Nunomura et al., 2002), myopathies (Tateyama et al., 2003),
atherosclerosis (Martinet et al., 2004), multiple system atrophy (Kikuchi et al., 2002),
23
Down’s syndrome (Nunomura et al., 1999), ALS (Chang et al., 2008), hemochromatosis
(Broedbaek et al., 2009), hepatic encephalopathy (Görg et al., 2010), and schizophrenia
(Che et al., 2010). Recently, RNA oxidation adducts were shown to be differentially
correlated with insoluble amyloid-β42, a causative factor of Alzheimer disease (Weidner
et al., 2011), suggesting that different forms of oxidized RNA may be involved in
different stages of this disease (Abe et al., 2002; Weidner et al., 2011). Furthermore,
increased RNA oxidation was reported under physiological conditions associated with
aging (Liu et al., 2002; Seo et al., 2008; Hofer et al., 2008a, 2008b). These findings
suggest that oxidative damage of RNA is a common problem that can affect many
systems. RNA oxidative damage may contribute to the process of aging and disease
development. Whether RNA oxidation damage directly causes diseases or aging and how
much it contributes to these processes have yet to be revealed.
It has been shown that the amount of 8-oxo-G is increased in the brains and
cerebrospinal fluid of Alzheimer Disease (AD) patients (Gabbita et al., 1998; Lovell and
Markesbery, 2001), and is restricted to vulnerable neurons in AD (Nunomura et al.,
1999). The oxidized mRNA and reduced protein expression indicate that RNA oxidation
may be directly associated with neuronal deterioration. mRNA oxidation may play an
important role in the pathogenesis of AD (Shan and Lin, 2006). Many identified oxidized
mRNA species are related to AD, either the transcripts have been characterized in AD or
their protein functions have been implicated in the pathogenesis of AD.These proteins
include p21ras, mitogen-activated protein kinase 1, carbonyl reductase, SOD1,
apolipoprotein D, glutamate dehydrogenase, etc (Kong et al., 2008).
24
rRNA in AD is oxidized by bound redox-active iron, which impaired protein
synthesis by altering ribosomal nucleic acids and the polyribosomal complex itself, and
occurred in the earlier stage of AD (Ding et al, 2005; Hoda et al., 2005).
1.3.4 Control of oxidized RNA
As described previously, RNA can be a major target of ROS among cellular
macromolecules. Compared to DNA, RNA is several times more abundant. Moreover,
RNA is present mostly in cytosol, and is in close proximity to mitochondria where the
majority of ROS produces. It is reported that in various organisms, levels of oxidative
damage in RNA are higher than those found in DNA. For instance, irradiation of human
skin fibroblasts with 765-kJ/m2 UV A induced more oxidation in RNA than in DNA
(Wamer and Wei, 1997). Based on HPLC-ECD analysis, the normalized levels of 8-oxoG in RNA from mammalian cell cultures or tissues increased five to ten fold after
oxidative stress, which is 10 - 25 times higher than those of 8-oxo-dG in DNA under the
same condition (Fiala et al., 1989, Hofer et al., 2005 and 2006; Shen et al., 2000).
Therefore, we would conclude that there are many more damaged lesions in RNA than in
DNA within the same cell at any time.
Such a high number of oxidized RNA lesions may cause major problems to cells
if the lesions are not efficiently removed. However, little is known how oxidized RNA is
eliminated in any living organism. One may argue that oxidized RNA is naturally
destroyed during RNA turnover, and therefore is not a real threat to cell. This is a
misconception. Oxidative damage caused by ROS occurs in only minutes. As described
previously, mRNA turnover generally takes longer length of time. The majority of
25
cellular RNA is in the form of more stable rRNA and tRNA. There must be more
efficient and specific mechanisms than normal RNA turnover for elimination of oxidized
RNA. It is well documented that oxidatively damaged DNA is actively repaired by a
number of mechanisms. In contrast, repair activity of oxidized RNA has not been
reported.
The existence of RNA surveillance mechanisms to control oxidized RNA and to
protect cells against oxidative stress is proposed in Figure 1. This idea is supported by the
observation that 8-oxo-G level of RNA drops dramatically after removal of oxidative
stress insult. Pulse exposure of cultured human lung epithelia cells to H2O2 increased the
8-oxo-G level immediately and sharply, and the 8-oxo-G level decreased after 24 h
(Hofer et al., 2005). 8-oxo-G level of RNA in HeLa cells was increased first after a pulse
treatment of H2O2, followed by 50% drop within 1 h and then continuously decreased to
baseline (Wu and Li, 2008). In E. coli, after a sharp increase, 8-oxo-G level quickly drops
close to the normal level when the cells were subsequently grown in fresh medium (Liu
et al., 2012). Since the H2O2 is degraded rapidly and reduced to basal level in minutes
after adding (Wu et al., 2009), the sustained high level of 8-oxo-G in cultures of
continuous H2O2 treatment is likely caused by oxidized components of the medium.
These results suggest that high levels of oxidized RNA are not tolerated by cells and the
cell tends to remove the oxidized RNA assembly by specific mechanisms.
As described in the previous section, RNA quality control mechanisms usually
degrade aberrant RNA using multiple RNases and other facilitating enzymes. It is natural
to propose that similar RNA control activities may also be responsible for removing
26
oxidized RNA. In E. coli, oxidatively damaged RNA could be degraded by ribonucleases,
poly (A) polymerase, and RNA helicases. Recently, we have shown that PNPase binds
oxidized RNA with high affinity to help the cell reduce 8-oxo-G in RNA and to survive
oxidative stress (Wu et al., 2009). Interestingly, this function is not dependent on PNPase
association with the degradasome or with RhlB. It has also been shown that human
PNPase specifically binds an 8-oxo-G-containing RNA with a high affinity (Hayakawa et
al., 2001, 2006). Similar to its E. coli homolog, human PNPase reduces 8-oxo-G and
protects HeLa cells under oxidative stress (Wu et al., 2008). Studies in our laboratory
have also demonstrated that RNase II, R, and PAP play protective roles under oxidative
stress by controlling the level of oxidized RNA (unpublished observations). However,
much remains unknown about how oxidized RNA is recognized, and how specific
degradation of oxidized RNA is initiated.
Oxidized ribonucleotides can be generated by direct oxidation of ribonucleotides
or by degradation of oxidized RNA. In fact, leukocytes produced more 8-oxo-G in the
nucleotide pool under oxidative stress (Shen et al., 2000). If the oxidized riboucleotides
were reused in transcription, damaged RNA could be generated during RNA synthesis. In
fact, cells have developed mechanisms to reduce the incorporation of damaged
ribonucleotides into RNA (Hayakawa et al., 1999). In E. coli, RNA polymerase can
utilize 8-oxo-GTP as a substrate, which could be prevented by the MutT protein. MutT
was originally found to degrade 8-oxo-dGTP, and prevent the misincorporation of 8-oxodGTP during replication (Maki and Sekiguchi, 1992). Later, it was found that MutT also
degrades 8-oxo-GTP to 8-oxo-GDP and 8-oxo-GMP (Taddei et al., 1997). Moreover, 827
oxo-GTP was degraded more efficiently than 8-oxo-dGTP by MutT. Guanylate kinase,
which converts GMP to GDP, is inactive on 8-oxo-GMP, blocking reuse of 8-oxo-GMP
in RNA synthesis. In addition, the E. coli RNA polymerase incorporates 8-oxo-GMP
from 8-oxo-GTP into RNA at a much lower efficiency than incorporating GMP from
GTP (Taddei et al., 1997).
Normal ROS level
Antioxidant
mechanisms
Environmental
stress
Increased ROS
RNA
Increased RNA
Normal level of
oxidative damage
RNA damage
surveillance?
Normal
Related cell
dysfunction
Related diseases
(Adapted from Li et al., 2006)
Figure 1.
RNA oxidative damage and cellular defense. Reactive oxygen species
(ROS) are produced in normal oxygen metabolism. The imbalance of antioxidant
capacity and the generation of ROS will cause increased RNA oxidative damage. RNA
surveillance mechanisms should exist to decrease the RNA damage to normal level;
otherwise the oxidative damage of RNA may cause cell dysfunction and related disease.
28
1.4 Hypothesis and approaches of this study
We propose that ribosomal RNA (rRNA), although being highly structured and
associated with ribosomal proteins, can be oxidized under oxidative stress; and damaged
rRNA under oxidative stress is selectively degraded.
To examine these hypotheses, I first characterized oxidation levels of various
RNA species. We anticipated that different RNA species may undergo different levels of
oxidation and quality control processes. Therefore, the steady-state levels of RNA
oxidation may vary among rRNA, tRNA and other RNAs. Factors that may cause the
variation may include RNA structure, association with proteins, binding to redox metals
(Honda et al., 2005), etc. I approached this goal by isolating various RNA species from
cultures that were treated with or without H2O2, and analyzed 8-oxo-G levels in these
RNA preparations. This experimental set up also enabled us to examine the effect of
H2O2 dosage and length of treatment time on the level of 8-oxo-G in various RNA
species. The results of this study helped us understand if RNA structure and association
with protein or redox metals play a role in RNA oxidation under oxidative stress.
Second, I investigated whether 8-oxo-G-containing RNA is selectively eliminated
over time. To understand if different RNA species undergo different mechanisms of 8oxo-G elimination, ribosomal and non-ribosomal 8-oxo-G levels were studied separately
in a time course following a pulse H2O2 treatment. Furthermore, the rate of removal was
studied in wild type and mutant strains lacking RNA decay enzymes. The results
provided insight on the rate of selective removal of 8-oxo-G by specific RNA
degradation activities.
29
Third, I studied the role of several RNases in the degradation of the 16S and 23S
rRNA under oxidative stress conditions. This will further help us understand how
damaged rRNA can be removed within the cell.
Finally, I have tried to identify additional activities that may control RNA quality
and protect cells under oxidative stress. I have worked on candidate genes whose
potential roles have been suggested by proteomics and bioinformatics searches. The
results of this work revealed a broader spectrum of proteins that play roles in RNA
quality control and cell survival under oxidative stress conditions.
The specific aims of my work are: i. Characterize the RNA oxidative damage in
cells under oxidative stress. ii. Analyze the pattern of elimination of oxidized rRNA. iii.
Identify proteins that play roles in rRNA degradation and quality control under oxidative
stress.
30
2. Materials and Methods
2.1 Materials
Chloramphenol, lysozyme, diethyl pyrocarbonate (DEPC), deferoxaminemesylate
(DFOM), guanosine (G) and Chelex 100 were purchased from Sigma-Aldrich (St. Louis,
MO). Tetracycline was obtained from MP Biomedical Inc (Solon, OH). 8hydroguanosine (8-oxo-G) was from Calbiochem (La Jolla, CA). TRI Reagent was from
Molecular Research Center (Cincinnati, OH). Sodium deoxycholate was purchased from
Avocado Research Chemicals Ltd (Lancashire, LA). α -32P-dATP was purchased from
GE Healthcare Inc (Piscataway, NJ). γ -32P-ATP was purchased from PerkinElmer
(Waltham, MA). ThermoscriptTM RNase H- reverse transcriptase, dNTPs, Taq DNA
polymerase, and DTT used in PCR reactions were from Invitrogen (Carlsbad, CA).
QIAquick Gel Extraction kits were purchased from QIAGEN Science (Germantown,
MD). T4 Polynucleotide Kinase was purchased from New England BioLabs (Ipswich,
MA). M-MLV Reverse Transcriptase and 5 X M-MLV buffer were from Promega
(Madison, WI). Kanamycin and RNain were obtained from Fisher Scientific (Pittsburgh,
PA). Rifampicin was purchased from Duchefa Biochemie (Saint Louis, MO). All other
chemicals are reagent grade. Double distilled water (ddH2O) was treated with DEPC and
Chelex 100. Buffers, phenol (EMD Chemicals, Inc. Gibbstown, NJ), ethanol (Sigma-
31
Aldrich, Saint Louis, MO) and isopropanol (ACROS Organics, Fair Lawn, NJ) were also
treated with Chelex 100 as appropriate.
The following RNA and DNA oligonucleotides were synthesized by Integrated
DNA Technologies (Coralville, IA) and were used as primers for rRNA specific PCR
products and to produce RNA:DNA duplex: 23S-1, 5 ′ -AGC GAC TAA GCG TAC
ACG GT-3 ′ ; 23S -2, 5 ′ -AAG ACC AAG GGT TCC TGT CC-3 ′ ; 23 S-3, 5 ′ -TTA
GAG GCT TTT CCT GGA AGC-3 ′ ; 23S-4, 5 ′ -AGC CTC ACG GTT CAT TAG TAC
C-3 ′ ; 16S-R, 5 ′ -TAT TCA CCG TCC CAT TCT GA-3 ′ ; 16S-F, 5 ′ -TGC AAG TCG
AAC GGT AAC AG-3 ′ ; 5’-rCrGrG rArGrA rGrUrA rArArA rArUrG rArArA rGrUrA
rCrGrU rGrCrU rUrCrC rGrUrG rArArG rUrArA rUrUrU rUrUrU rCrGrC rArU-3’, 5’ATG CGA AAA AAT TAC TTC ACG GAA GCA CGT ACT TTC ATT TTT ACT
CTC CG-3’. More 16S and 23S rRNA oligo probes and primers, and RNA linker
synthesized by Integrated DNA Technologies (Coralville, IA) are in supplementary data.
2.2 Strains and growth condition
Escherichia coli K-12 strain BW25113 mutant strain (lacIq rrnBT14 ΔlacZWJ16
hsdR514 ΔaraBADAH33 ΔrhaBADLD78) was derived from the F- E coli K-12 strain
BD792, a two-step descendent of ancestral E.coli K-12, and has no other known
mutations (Baba et al., 2006, and Datsenko et al., 2000 ). The mutants of BW25113
having one gene missing used in this study were constructed by Wanner’s and Mori’s
Group (Beba et al., 2006). Single or double gene deletion mutants of E.coli K-12
derivative CA244 rna (lacZ trp relA spoT rna, tetR) (Li and Deutscher, 1994) used as
wild type were constructed by P1 transduction. Cultures from individual colonies were
32
usually grown in a Yeast extract-Trypton (YT) medium (BD Diagnostic Systems, Sparks,
Maryland, USA) with respective antibiotic, and incubated at 37°C with shaking.
Antibiotics kanamycin (kan) was added to the medium at 25 μg/ml, chloramphenicol
(cam) 25 μg/ml, and tetracycline (tet) 10 μg/ml when needed.
2.3 Mutant strains construction
The open reading frames of the corresponding gene were replaced with a
kanamycin cassette (kan) when originally constructed in E.coli K-12 strain BW25113
(Beba et al., 2006). In order to compare with other mutations that we have been studying,
these mutant alleles were transferred by P1 transduction into another E.coli K12
derivative CA244. A single colony from each BW25113 mutant was inoculated into YT
medium and grown overnight at 37 °C. P1 phage was prepared by incubating the culture
with CaCl2 and starter P1 lysate at 37 °C. Chloroform was used to help lyse the cells. P1
phage containing the antibiotic cassette was stored at 4 °C. The recipient strain of CA244
Δrna was grown overnight at 37°C and infected with the P1 phage preparation. Genetic
recombination, catalyzed by enzymes of the recipient strain, will incorporate the bacterial
fragments into the recipient chromosome (Thomason et al, 2007). The cell was resuspended in YT with 20mM NaLitrate and spread onto the antibiotic plates. For each
strain, two colonies were picked and re-streaked onto antibiotics plates.
2.4 Treatment of E. coli cultures with H2O2
Overnight cultures grown in YT medium with shaking at 37 °C were diluted with
fresh, pre-warmed YT medium and incubated until OD550 reached 0.5. H2O2 was added to
33
the culture at the desired final concentration. An equal volume of H2O was added to the
control cultures. The cultures were continually incubated with shaking and samples were
collected at various time points.
2.5 Isolation of RNA and DNA
Cells were collected and resuspended in lysis buffer (10 mm Tris · Cl, 10 mm
EDTA pH 8.0, 1 % SDS, 10 % glycerol and supplemented with freshly added DFOM to a
final concentration of 10 mm. Total RNA was routinely prepared for 8-oxo-G analysis.
The lysates were diluted 10 fold with ddH2O and an equal volume of a mixture of
liquefied phenol (pH ~ 4) and chloroform (9:1) was added. The mixture was applied to a
vortex intermittently for 10 min at room temperature. After centrifugation, the upper
layer of aqueous phase was transferred to a new tube and was extracted one more time
with phenol-chloroform. To the recovered aqueous phase, 1/10 volume of 3 M potassium
acetate (KAc, pH 5.2) and an equal volume of isopropanol were added. The tubes were
filled with nitrogen, mixed well and kept at -80 ° C for 1 h. RNA was collected by
centrifuging at 20,000 g for 10 min at 4 °C. The RNA pellets were washed twice with 75
% cold ethanol and were vacuum-dried for 10 min. The dried RNA was directly
dissolved in a digestion mixture in which the RNA was converted to nucleosides (Wu et
al., 2009). When RNA and DNA were isolated simultaneously, cell lysates were
extracted with phenol and chloroform and then the nucleic acids were precipitated by
ethanol, as previously described (Wu et al., 2009).
34
2.6 Isolation of ribosomal and non-ribosomal RNA
Exponentially growing cells were harvested by centrifugation at 10,000 g for 3
min. The cell pellet was resuspended in 1 ml saline solution, transferred into a
microcentrifuge tube and collected by centrifugation at 10,000 g for 1 min. The pellet
was resuspended in 800 μl cell extract buffer (10 mm Tris · HCl, pH 7.75, 100 mm
NH4Cl and 1 mm DFOM; Vaidyanathan et al., 2007). The cell suspension was
supplemented with 12.5 μl of lysozyme (40 mg/ml) and the mixture was incubated for 1
min at room temperature (Ron et al., 1966). The mixture was frozen in a dry ice/ethanol
bath and then thawed completely in a water bath at 37 °C. The freeze-thaw process was
repeated three times to completely lyse the cells (Ron et al., 1966). To the cell lysate, 30
μl of 10 % sodium deoxycholate was added and the solution was incubated in an ice
water bath for 3 min (Ron et al., 1966). The lysate was then centrifuged at 25,000 g for
40 min at 4 ° C. The supernatant was transferred to an ultracentrifuge tube
(Microcentrifuge Polyallomer, Part No. 357448, Beckman Coulter, Indianapolis, IN,
USA) and centrifuged at 200, 000 g using the Beckman TLA-100.4 rotor and
ultracentrifuge at 4 °C for 2 h to pellet the ribosomes (Vaidyanathan et al., 2007, Székely
et al., 1973). The supernatant was transferred to a microcentrifuge tube. Subsequently,
the ribosome pellet was briefly rinsed with cell extract buffer, and re-suspended in the
same buffer supplemented with 0.5 % SDS (Li and Deutscher, 2009). RNA was isolated
from the ribosome suspension and the non-ribosomal supernatant fraction as previously
described (Li and Deutscher, 2009).
35
2.7 Separation of long and short RNA species
Long and short RNAs were separated by differential precipitation using
isopropanol (Deutscher and Hilderman, 1974; Li and Deutscher, 2009). RNA was added
together with sodium acetate (NaAc, pH 7.0) to 0.3 M and DFOM to 1 mM. To the RNA
solution, 0.54 volume of isopropanol was added and the solution was mixed immediately.
The mixture was centrifuged at 21,100 g for 15 min. The RNA pellet contained long
RNA, mainly rRNA and mRNA species. The aqueous phase was transferred into a new
microcentrifuge tube, and additional isopropanol was added to a final 0.98 volume. The
mixture was kept at -80°C for at least 1 h. RNA was collected by centrifugation at 21,100
g for 10 min. The resulting small RNA was almost pure tRNA (Deutscher and
Hilderman, 1974). The pellets of long and short RNAs were washed twice with precooled 75 % ethanol at 21, 100 g for 3 min before further analysis.
2.8 Determination of 8-oxo-G level in RNA and 8-oxo-dG level in DNA by HPLC
After digestion, the 8-oxo-G level in RNA was determined by HPLC as
previously described (Gong et al., 2006; Wu et al., 2009). For simultaneous detection of
8-oxo-G in RNA and 8-oxo-dG in DNA, digestion was carried out under the same
conditions for RNA. The resolving time for HPLC is 70 min to allow the separation of
both DNA and RNA nucleosides. Chemical standards of G, dG, 8-oxo-G and 8-oxo-dG
were used. The normalized level 8-oxo-G/105 G in RNA or 8-oxo-dG/105 dG in DNA
was calculated.
36
2.9 RNA denaturation and oxidation in vitro
The highly structured rRNA and tRNA were isolated by the method described
above. RNA dissolved in double-distilled water (ddH2O) was denatured by incubating at
95°C for 2 min, followed by chilling immediately in an ice water bath. In vitro oxidation
was performed as described previously with modifications (Gong et al., 2006). Briefly,
40 μg of native or denatured RNA were incubated in a buffer containing 10 mM H2PO4
/HPO4 , pH 7.4, 1 μM CuSO4 , 10 μM ascorbic acid and various concentrations of H2O2
at 37°C for 1 h. RNA was then precipitated and washed before analysis of 8-oxo-G
levels.
2.10 Preparation of oligomer single-stranded RNA and RNA:DNA duplex
The single-stranded 50-mer RNA and complementary DNA were synthesized
chemically. To generate the RNA:DNA duplex, equal molar amounts of RNA and
complementary DNA oligonucleotides were mixed in 50 µl annealing buffer containing
10 mM Tris chloride, pH 7.5, and 25 mM NaCl (Schein, 2001). The mixture was heated
to 94°C and was then gradually cooled to room temperature.
2.11 Determination of copper binding capacity
20 µg rRNA or tRNA dissolved in oxidative buffer containing 10 µM ascorbic
acid, and 10 mM H2PO4 /HPO4, pH 7.4, was denatured by incubating at 95°C for 8 min
and then kept in ice water for 4 min. 40 µM CuSO4 was added to the mixture. At the
same time, an equal amount of rRNA or tRNA dissolved in the same buffer was kept in
ice water as the native RNA. Single stranded RNA and RNA:DNA duplex were mixed
37
with the oxidative buffer too. When all samples were ready, 40 µM CuSO4 was added. A
G25 column was prepared as instructed by spinning at 2,700 rpm for 1 min and then 50
µl oxidation buffer was used to calibrate the column by spinning another 1 min at 2,700
rpm. The nucleic acid in oxidative buffer with 40 µM CuSO4 was applied to the
calibrated G25 column and the elute was collected in a 1.5 ml microcentrifuge tube after
centrifuging at 2,700 rpm for 2 min, which included the nucleic acid and the bound
copper. The free Cu2+ ion was trapped in the resin of G25 columns. The saturation of G25
column was determined by applying 50 µl oxidation buffer with 40 µM CuSO4 but
without nucleic acid. The oxidation buffer without the nucleic acid or application of G25
column, but only with 40 µM CuSO4, was used as the control. A copper assay kit was
used to determine the copper concentration in the elute. As the manufacture instructed,
100 µl DEPC- and Chelex 100- treated ddH2O was used as blank. 20 µl provided 1.5
mg/dL copper concentrate was mixed with 80 µl DEPC- and Chelex 100- treated ddH2O
to make 46.5 µM Cu2+, and lower concentrations of 20 µM, 10 µM, 5 µM, and 1 µM
Cu2+ were prepared for a standard curve. 35 µl Reagent A was mixed with every tube
including the blank, Cu2+ in various concentrations for the standard curve, and RNAs
eluted from the G25 column. When Reagent A was mixed with the eluted RNA, the
mixture was first centrifuged at 20,000 g for 10 min at 4 °C and then 100 µl clear
supernatant was transferred to separated wells of a clear flat-bottom 96-well plate. When
Reagent A was mixed with the blank or various concentrations of Cu2+, 100 µl mixture
was directly transferred to the 96-wells plate without centrifugation. 5 µl Reagent B and
150 µl Reagent C were mixed as Working Reagent and 150 µl of the Working Reagent
was transferred to each well of the 96-well plate. The plate was tapped to mix all the
38
solutions and incubated for 5 min at room temperature. The optical density was read by
SpectraMax M5e plate reader at 359 nm, and each copper concentration was evaluated by
using the standard curve.
2.12 Determination of rRNA fragmentation
Total RNA was extracted with the TRI Reagent and was dissolved in ddH2O. The
RNA solution was added to 1/5 volume of 6 × RNA loading buffer containing 0.25 %
Bromophenol blue, 0.25 % xylene cyanol, 30 % glycerol, 1.2 % SDS, 60 mm sodium
phosphate (Kevil et al. , 1997). RNA in the loading mixture was denatured by incubating
at 75 °C for 5 min and immediately chilled in an ice water bath (Cheng and Deutscher,
2003). RNA was separated by electrophoresis on a 1.5 % agarose gel in 0.5 × TBE
buffer, first at 60 V for 10 min and then at 100 V for 1 h. The gel was stained with SYBR
Gold and photographed under UV light. Northern analysis was carried out according to
the procedure described previously (Cheng and Deutscher, 2003) with some
modifications. RNA in the gel was transferred to GeneScreen Plus ® membrane (Pall
Life Science, Pensacola, FL, USA) by electroblotting at 15 V overnight in 1 × TBE
buffer. RNA was fixed to the membrane by UV irradiation for 2 min, prehybridized for
2.5 h at 68 °C in 1 × Denhardt’s solution and then hybridized overnight at 55 ° C with
probes specific for 23S or 16S rRNA. The 23S and 16S specific DNA products were
generated by PCR using specific primers and E. coli genomic DNA. The PCR products
coverd the entire rRNA sequences. Because 23S rRNA is nearly 3 kilobases in length, we
first made two halves using the two pairs of primers. 23S-1 and 23S-3 oligos in Materials
were one pair of primers to synthesize the 5′ end part products. 23S-2 and 23S-4 oligos in
39
Materials were another pair of primers to synthesize the 3′ end part products. Then all
labeled [32P]-labeled probes were generated by PCR reactions using the PCR products of
16S or 23S rRNA as template, the primers complementary to the rRNA (23S-3, 23S-4,
and 16S-R in Materials), and dNTPs plus α -[32P]-dATP. RNAs in the membrane were
hybridized with one 16S rRNA probe or two 23S probes, 5’ half and 3’ half, to label the
related rRNA products. After hybridization, the membrane was washed twice for 10 min
at 55 °C, each with 1 X SSC and 0.1% SDS, prior to autoradiography.
2.13 Determination of the 5’ end of rRNA fragments by primer extension
The protocol is based on the method described (Li and Deutscher, 1995) with
some modifications. 16S and 23S primers, 10 bp, and 25 bp DNA ladders were labeled
by γ-32P-ATP with T4 Polynucleotide Kinase at 37 °C for 1 h and then 65 °C for 20 min.
Four micrograms of rRNA was mixed with 2 µM of 32P-labeled primers in 10 µl of buffer
containing 10 mM Tris HCl (pH 7.5), 300 mM NaCl, 2 mM EDTA (pH 8.0). The mixture
was heated to 80°C for 4 min and then at 55°C for 1 h. Two microliters of reverse
transcriptase (M-MLV, 200 u/µl) in 8 µl 5 X M-MLV buffer, 0.38 µl of RNasin (15
units), 10 µM DTT, and 30 µl DEPC-H2O were added to the hybridization mixture. The
primer was extended at 37 °C for 30 min. The reaction was stopped by addition of 1 ul of
0.5 M EDTA (pH 8.0). DNA was precipitated with 1/10th vol of 3 M sodium acetate (pH
4.8) and 3 vol of cold absolute ethanol by incubating at -80 °C overnight. The pellet was
recovered after centrifugation, washed with cold 75% ethanol, and vacuum dried.
Samples were redissolved in 6 µl 96% formamide containing 1 mM EDTA, xylene
40
cyanol, and bromphenol blue. The products were separated on 8% polyacrylamide/8 M
urea gels and detected by autoradiography.
2.14 Determination of the 3’ end of rRNA fragments by 3’ RACE
Several 16S and 23S oligonucleotides were designed to be used as the primers of
RT-PCR and listed in the supplementary data. Total RNA from mutant rnb rnr pnp was
extracted by TRI Reagent. 1 µg RNA was linked with a 10 pmol linker by T4 RNA
lygase, incubating at 37 °C for 1 h. First-strand complementary cDNA was synthesized
using ThermoscriptTM RNase H- reverse transcriptase to transcribe RNA with the primer
of the linker. The primer pair of 16S- or 23S- oligo/primer of the linker was used to
amplify the first 16S or 23S rRNA cDNA fragment. The PCR buffer contains 1 X Taq
buffer, 0.2 mM dNTPs, 0.04 U Taq polymerase, and 1 µl 10 mM of each primer. The
PCR conditions included an initial step at 90 °C for 3 min, followed by 20 cycles at 90 °C
for 3 s, 55 °C for 30 s, and 72 °C for 1 min. A final extension step was performed at 72
°C for 10 min after which it was cooled to 4 °C. A nest PCR was then performed to avoid
non-specific products, whose conditions are the same with 1st PCR described above,
excepting the 16S- or 23S oligos is around 100 nt downstream of 1st PCR corresponding
one, and the amplification was proceeded 15 cycles. The nest PCR products were
subjected to electrophoresis on a 2% agarose gel to confirm the purity, and purified using
QIAquick Gel Extraction kit following the manufacturer’s instructions. The sequences of
extracted fragments were determined by MCLAB, CA.
41
2.15 Effect of H2O2 on the growth of wild-type and mutant cells
One colony from the wild-type or mutant strain was inoculated into fresh YT
medium and incubated at 37 °C for overnight. The overnight culture was diluted with
pre-warmed YT and incubated again until OD550 ≈ 0.5. This culture in exponential phage
would be diluted ten times into OD550 = 0.05, and then continually serially diluted five
times with fresh YT. Two microliters of each dilution was spotted onto YT agar plate
supplemented with H2O2. The H2O2 concentration was 0 mM, 0.4 mM, 0.5 mM, and 0.6
mM. The cell culture on the plates was incubated for 14 h at 37 °C. Pictures of cell
growth on the plate were taken by UV spectrometry.
2.16 Determination of cell viability
When the OD550 of the culture reached 0.5, H2O2 or H2O was added. 100 μl of the
cultures were dispensed into each well of a 96-well plate. The plate was incubated at 37
°C by shaking at 150 rpm. The optical density value at 550 nm was detected by a plate
reader in a time course. CFU was determined in a time course after addition of H2O2 to
exponentially grown cultures at different densities. 10 % (v/v) AlamarBlue (Life
Technologies, Grand Island, NY, USA) stock solution (final concentration of resazurin:
17.5 mm) was made in YT medium. The serially 10-times diluted AlamarBlue was also
prepared and kept on ice. 100 μl of each dilution was placed in all wells of a 96-well plate
and one well contained 100 μl stock AlamarBlue as the blank. When a desired
concentration of H2O2 was added to the cell in the exponential phase, at each time point, a
10 μl culture was transferred into one well containing the supplemented AlamarBlue
dilutions. The plate was put in a plate reader (Molecular Devices SpectraMax M5e,
42
Sunnyvale, CA, USA) and the fluorescence was detected (excitation, 544 nm; emission,
590 nm, automix, 5 s) in arbitrary units (FSU). Finally, the cell culture would reach the
maximum fluorescence. FSU of each well was normalized by subtracting the
fluorescence data of the blank. The linear curve of FSU and time was plotted. On the
curve, the time to F50 (half of maximal fluorescence), T50, was pinpointed for each
dilution. Then T50 was plotted versus log CFU per ml.
43
3. Results
3.1 Characterization of RNA damage under oxidative stress in Escherichia coli
This chapter is taken from a recently published paper (Liu et al., 2012). Reactive
oxygen species (ROS) are constantly produced in normal metabolic processes and
become more abundant under oxidative stress (OS). It has been shown that RNA is a
major target of ROS. Under oxidative stress, RNA oxidation increases markedly,
resulting in elevated levels of 8-hydroxyguanine (8-oxo-G) (Hofer et al., 2005, 2006; Liu
et al., 2012), increased termination of reverse transcription (Gong et al., 2006), and
increased abasic sites (Tanaka et al, 2011a, 2011b). 8-oxo-G level in RNA was much
greater than 8-oxo-dG in DNA in mammalian cells, tissues, and E.coli with H2O2
challenge (Fiala et al. , 1989 ; Shen et al. , 2000 ; Hofer et al. , 2005, 2006; Liu et al.,
2012). These findings suggest that RNA oxidation is a predominant feature under
conditions of ROS attack.
Mounting evidence suggests that RNA oxidation affects translation. Oxidized
mRNA causes a reduction in protein synthesis and the formation of aggregated protein
products in cells (Shan et al., 2003, 2007). It was further shown that ribosomes stall
during translation elongation of oxidized mRNA and produce truncated proteins (Tanaka
et al., 2007; Shan et al., 2007). Oxidation also causes ribosome dysfunction (Ding et al.,
2005; Honda et al., 2005). Oxidized RNA may be controlled by cellular surveillance
44
activities that prevent damaged nucleotides from being incorporated into RNA (Taddei et
al., 1997; Hayakawa et al, 1999; Ishibashi et al., 2005), degrade, or repair damaged RNA
(Wu and Li, 2008; Wu et al., 2009; Kong and Lin, 2010). Consistently, PNPase was
shown to reduce RNA oxidation and protect E. coli and HeLa cells under oxidative stress
(Wu and Li, 2008; Wu et al., 2009). Deficiency in RNA surveillance, or environmental
conditions that cause OS, may result in a surge of RNA oxidation which leads to a loss or
alteration of RNA function in protein synthesis and other processes (Li et al., 2006).
RNA oxidation has been recently recognized as an important biological process
that is strongly implicated in deficient cellular functions and in development of human
diseases (reviewed in Nunomura et al., 2009; Wurtmann and Wolin, 2009, Paulsen et al.,
2012). Despite the increasing interest in studying RNA oxidative damage and quality
control, little is known about the level of RNA damage under oxidative stress and
whether different RNA species and structures are differentially oxidized by ROS. In a
growing E. coli cell, highly structured rRNA and tRNA account for nearly 80 % and 15
% of the total RNA, respectively. In addition, most of the rRNA molecules are present in
ribosomes where rRNAs are tightly bound with ribosomal proteins. One would expect
that RNA would be protected from oxidation by the presence of highly folded structures
or by association with proteins. In this work, we examined the level and distribution of
oxidized RNA in E. coli under normal and oxidative stress conditions to answer these
questions.
45
3.1.1 H2O2 causes a quick and dosage-dependent increase of 8-oxo-G in cellular
RNA
In order to determine the level of RNA oxidation in response to H2O2 treatment,
we have measured 8-oxo-G content in cellular RNA in a time course after addition of
H2O2 in E. coli cultures. Under conditions described in Materials and methods, the basal
level of 8-oxo-G in E. coli RNA is slightly lower than one 8-oxo-G per 105 G. After
adding H2O2 to the cultures, the 8-oxo-G level increases in only minutes and remains
high for hours (Fig. 2A). It should be noted that the concentration of H2O2 reduces
rapidly in the culture media, becoming close to the basal level 5 min after addition of the
oxidant (Wu et al., 2009). Interestingly, after a pulse treatment with H2O2, the level of 8oxo-G initially rises and then quickly drops to almost normal level when the cells were
subsequently grown in fresh medium (Fig. 2A). This observation suggests that the
sustained high level of 8-oxo-G in cultures of continuous H2O2 treatment is likely to be
caused by oxidized components of the medium.
The increase of 8-oxo-G in RNA depends on the dosage of H2O2, from three 8oxo-G/105 G when treated with 1 mm H2O2 to nearly ten 8-oxo-G/105 G in the presence
of 5 mm H2O2 (Fig. 2B). Cells grown in rich and minimal media contain similar levels of
basal and H2O2 induced 8-oxo-G in RNA (data not shown).
3.1.2 H2O2 induces higher levels of 8-hydroxyguanine in RNA than in DNA
To compare the level of oxidation in DNA and RNA, we measured 8-oxo-G
content in RNA and 8-hydroxydeoxyguanosine (8-oxo-dG) in DNA simultaneously in the
46
A.
B.
(A By Xin Gong and B by Zhongwei Li)
Figure 2. H2O2 treatment causes quick, dosage-dependent increase of 8-oxo-G
content in cellular RNA. A. 8-oxo-G levels in RNA in a time course after addition of
H2O2 or H2O. Exponentially grown cultures of E. coli CA244 rna strain (OD550 = 0.5)
were treated with 1 mm H2O2 or an equal volume of H2O at 0 min. Total RNA was
extracted and 8-oxo-G levels were analyzed as described in Materials and methods. Pulse
treatment was carried out by exposing the cultures to H2O2 for 10 min. Cells were
pelleted and grown in fresh H2O2 -free medium for the rest of the time course. B. Increase
of 8-oxo-G levels in RNA depends on the dose of H2O2. Exponentially grown cultures
were treated with indicated concentrations of H2O2 for 15 min. Total RNA was isolated
and 8-oxo-G level was measured.
same culture treated with or without H2O2. This was done by a modified procedure that
enabled us to extract the DNA and RNA together. The nucleic acids were digested and
the resulting nucleosides were analyzed by HPLC under conditions that guanosine (G),
deoxyguanosine (dG), 8-oxo-G and 8-oxo-dG were separately detected in a single
47
sample. As shown (Fig. 3), this modified procedure caused an elevation in the basal level
of 8-oxo-G in RNA to above three 8-oxo-G/105 G, probably due to spurious oxidation
during preparation of the nucleic acids (de Souza -Pinto and Bohr, 2002). Nonetheless,
treatment with 5 mm H2O2 for 15 min increases 8-oxo-G to approximately ten per 105 G
in RNA when both RNA and DNA were prepared simultaneously (Materials and
methods). This result is consistent with the level of 8-oxo-G induction by the same H2O2
concentration following the procedure for RNA isolation only (Fig. 2B).
The basal level of 8-oxo-dG in DNA is lower than 8-oxo-G in RNA (Fig. 3).
However, it must still be higher than the actual level in DNA due to the same spurious
oxidation observed in RNA. Importantly, the level of 8-oxo-dG in DNA only increases
slightly after treatment with 5 mM H2O2, contrasting the large increase of 8-oxo-G in
RNA. Similarly, treatment with 1 mM H2O2 causes moderate increase of 8-oxo-G in
RNA but no change of 8-oxo-dG in DNA (data not shown). These data suggest that at a
steady state, OS induced oxidative damage of RNA is higher than that of DNA,
consistent with the results from similar studies using mammalian samples (Fiala et al. ,
1989 ; Shen et al. , 2000 ; Hofer et al. , 2005, 2006).
3.1.3 The distribution of 8-oxo-G in various RNA species
We first tried to understand the oxidation level of RNA of different sizes. Long
and short RNAs were isolated from total RNA. The long RNA is predominantly
composed of rRNA, and the short RNA is almost pure tRNA. Under normal conditions,
8-oxo-G level in the long RNA fraction is slightly lower than that in the short RNA. After
cells were treated with 3 mM H2O2, both RNA fractions contain elevated levels of 8-oxo48
G (Fig. 4 A). These results suggest that RNA size does not significantly affect oxidation
level and that structured RNA species can be oxidized efficiently. The latter point was
examined further below.
(By Xin Gong)
Figure 3. H2O2 treatment causes a higher elevation of 8-oxo-G in RNA than that of
8-oxo-dG in DNA. Exponential phase (OD550 = 0.5) cell culture was treated with and
without 5 mm H2O2 for 15 min. RNA and DNA were isolated together, then 8-oxo-G in
RNA and 8-oxo-dG in DNA were measured as described (Wu et al., 2009). Treatment
with lower concentrations of H2O2 does not cause a detectable increase of 8-oxo-dG in
DNA under these conditions (data not shown).
The rRNA species present in ribosomes constitute the majority of cellular RNA.
In order to understand if the highly folded structure of rRNA and tight association with
ribosomal proteins protect RNA from oxidation, we have determined 8-oxo-G levels in
RNA from ribosome and nonribosome fractions. To isolate the RNAs, cells were lysed
by freeze and thaw, after which, the ribosomes were prepared as described in Materials
and methods. RNA samples were prepared from both ribosomal and non-ribosomal
49
(supernatant) fractions and were examined by gel electrophoresis. The RNA from
ribosome fractions contains essentially pure rRNA. RNA from the non-ribosomal fraction
contains all RNA species including rRNAs that are not incorporated into ribosomes in
addition to RNA degradation intermediates. Non-ribosomal RNAs can also be further
fractionated into long and short RNAs; the latter is almost pure tRNA (Deutscher and
Hilderman, 1974; Li and Deutscher, 2009).
Under normal conditions, RNA isolated from ribosomes contains approximately
0.4 8-oxo-G per 105 G (Fig. 4B). This is significantly lower than the level of 8-oxo-G in
total RNA (Fig. 2). In contrast, the level of 8-oxo-G in RNA from the non-ribosomal
fraction is almost three times higher than the level in ribosomal RNA and is also higher
than the level in total RNA (Fig. 4B). There is no difference between the long and short
RNAs in the non-ribosomal RNA fraction. In various experiments, RNA isolated from
ribosomes is approximately 60% of total RNA and RNA from non-ribosomal fractions
constitutes the remaining 40%.
After cells are exposed to H2O2 for 15 min, 8-oxo-G levels increase in all RNA
fractions. Interestingly, RNA from ribosomes contains 8-oxo-G at the same and in some
cases higher levels than non-ribosomal RNA depending on the H2O2 dosage (Fig. 4B).
The results suggest that the exceeding complex structures do not protect rRNA from
oxidation, nor do the surrounding ribosomal proteins. 8-oxo-Gs were also generated in
non-ribosomal RNAs, without showing differences in the long and short RNA fractions.
Note that in the experiments shown in Figure 4B, the levels of 8-oxo-G are generally
50
higher than those shown in Figure 2, presumably due to differences in cell growth and
treatment conditions.
A.
B.
(A by Ravi Kumar Alluri)
Figure 4. The levels of 8-oxo-G in various cellular RNA species under normal
conditions and in response to H2O2 treatment. A. 8-oxo-G levels in long and short
RNA fractions of total RNA isolated from cultures that were treated with and without 3
mm H2O 2. B. 8-oxo-G levels in RNA from ribosome, non-ribosome and long and short
non-ribosome fractions. Exponentially grown cells were treated with 0.5 or 1 mm H2O2
for 15 min. Various RNA fractions were prepared and 8-oxo-G level was measured as
described in Materials and methods. The mean and standard error of triplicates were
plotted.
3.1.4 Highly folded structure does not protect RNA from being oxidized in vitro
In order to examine if RNA structure has any protective role against oxidation, we
have determined the levels of 8-oxo-G generated by H2O2 treatment in vitro of tRNA and
rRNA in either native or denatured forms. Such treatment introduces hundreds of 8-oxo51
G per 105 G using millimolar levels of H2O2 (Fig. 5). Surprisingly, native tRNA contains
slightly more 8-oxo-G than denatured tRNA at every concentration of H2O2 used in this
experiment (Fig. 5A). As for rRNA, 8-oxo-G level in the native form is not significantly
different from that in the denatured form, although the latter appears slightly lower at the
1 mM and 10 mM H2O2 dosages (Fig. 5B). The results are consistent with the
observation of RNA oxidation in ribosomes in vivo (Fig. 4) and contradict the proposition
that RNA structure can protect RNA from oxidative damage.
A.
B.
(A by Ravi Kumar Alluri)
Figure 5. Native RNA structures do not protect RNA from H2O2 - mediated
oxidation in vitro. A. tRNA was isolated from total RNA by isopropanol differential
precipitation (see Figure 4). Native and denatured tRNA samples were incubated with
indicated concentrations of H2O2 in vitro and 8-oxo-G levels were determined as
described in the Materials and methods section. B. rRNA was prepared from ribosomes
as shown in Figure 4. Native and denatured rRNAs were treated with indicated
concentrations of H2O2 in vitro and 8-oxo-G level was determined. The mean and
standard error of at least three replicates were plotted.
52
3.1.5 RNA fragmentation upon H2O2 treatment
Oxidation is also able to cause RNA strand breaks (Li et al., 2006). Alternatively,
oxidized RNA might undergo degradation. These processes could cause elevated levels
of RNA fragments. Here, we have examined if such fragments are produced and
accumulated upon H2O2 treatment. After cells were treated with H2O2, RNA products
increased below the full length 16S rRNA band (Fig. 6, left panel). These can be
fragments of 23S or 16S rRNAs. We analyzed 23S rRNA, 16S rRNA and their fragments
by Northern Blotting as described in Materials and methods. As shown (Fig. 6), middle
and right panels, short products of 23S and 16S rRNAs are increased by H2O2 treatment,
especially in areas below the full-length 16S rRNA band. Note that the increase of
fragmentation by H2O2 treatment is not dramatic. Considering that most cells are viable
under this condition (see below) and that some fragments of the rRNAs might be
degraded, the detectable increase in rRNA fragmentation does present another problem
that cells have to handle under OS.
Figure 6. RNA fragmentation induced by oxidative stress. Left panel. H2O2 treatment
causes a slight increase in the amount of RNA products that are shorter than 23S or 16S
rRNAs. Total RNA was prepared using TRI Reagent from exponential cultures treated
53
with and without 1 mm H2O2 for 15 min and was separated by electrophoresis on a 1.5 %
agarose gel. The major RNA species are marked on the left. Middle panel. Northern blot
of 23S rRNA showed a slightly darker smear under the full-length 23S upon H2O2
treatment, representing a slight increase in 23S rRNA fragments. Right panel. Northern
blot of 16S rRNA showing increased levels of 16S fragments in response to H2O2
treatment.
3.1.6 Cell death in response to H2O2 challenge
Escherichia coli cell viability under H2O2 treatment was studied by measuring
optical density of cultures and by determining colony forming unit (CFU). H2O2 causes
reductions of cell density at concentrations of 2 mM or higher (Fig. 7A). The slight
reduction of density in response to 2 and 4 mM H2O2 mainly occurs in the first 30 min.
Very little change in OD550 was observed when H2O2 was added to 7 or 10 mM,
suggesting substantial cell death (Fig. 7A).
Viable cell counts confirm that the reduction of culture density is actually due to
cell death, which happens mainly in the first 60 min after H2O2 addition (Fig. 7B).
Similar to 8-oxo-G levels in RNA, cell death depends on the dosage of H2O2 (Table 1).
When H2O2 was added at 1 to 5 mM, cells die at rates that increase depending on H2O2
concentration. A higher rate of cell death was observed at 60 min than at 30 min after
H2O2 addition at every concentration. H2O2 at 10 mM kills cells completely, consistent
with the results of cell density reduction (Fig. 7A). We noted that 1 mM H2O2 causes
detectable cell death based on CFU but it does not affect culture density. The difference
between CFU and culture density reduction responding to 1 mM H2O2 is presumably due
54
to the fact that dead cells contribute to density reading of the cultures. As expected, H2O2
at 2 mM or higher concentrations caused a reduction to both CFU and density of culture.
CFU analysis also demonstrated that the effect of H2O2 depends on the density of
the culture. As shown (Fig. 7C), where OD550 = 0.01 cultures were treated with 2 mM
H2O2, only a small percentage of cells (4-5 %) survived after 60 min. CFU was also
reduced by 60 % in the presence of only 0.5 mM H2O2. It is likely that low density
cultures hydrolyze H2O2 much slower than high density cultures, resulting in sustained
oxidation and more cell death in low density cultures.
3.1.7 Discussion
In this work we have demonstrated that cellular RNAs are quickly and highly
oxidized under oxidative stress, as indicated by the rise of 8-oxo-G levels in RNA, which
is dependent on H2O2 dosage. Oxidation occurs to all RNA species. RNA structures and
association with proteins do not appear to be able to protect RNA from being oxidized by
H2O2. In addition, a small amount of ribosomal RNA fragments can be identified upon
H2O2 treatment.
Fenton reaction during preparation, storage and processing causes spurious
oxidation of nucleic acids, resulting in variations of reported basal levels of oxidized
nucleobases in the order of magnitude (ESCODD et al., 2005). This variation could also
greatly interfere with the results in response to oxidants (de Souza -Pinto and Bohr,
2002). In order to reduce spurious oxidation in our experiments, we have adopted
55
procedures to minimize exposure to oxygen and to reduce the level of contaminating
metal ions (Shen et al., 2000; ESCODD et al., 2005; Hofer et al., 2006; Wu et al., 2009).
However, it is likely that the true basal level of 8-oxo-G is lower than reported in this
work because spurious oxidation might not be completely avoided during HPLC analysis
(ESCODD et al., 2005). In fact, elevated basal levels were occasionally observed when
an alternative preparation method was used (Fig. 3). Nonetheless, our results
demonstrated 8-oxo-G levels that respond well to all dosages of H2O2.
A.
B.
C.
(A by Jinhua Wu, B and C by Xin Gong)
Figure 7. H2O2 treatment causes a dose dependent growth reduction of E. coli. A.
H2O2 was added into exponential-phase cultures (OD550 = 0.5) at 0 min to final
concentrations indicated in the Figure. The cultures were then immediately dispensed into
the wells of a 96-well plate. The plate was incubated with shaking and the OD550 was
recorded by a plate reader in a time course. Four replicates were read and the mean and
standard error were plotted. Note the actual OD550 shown is lower than 0.5 initially due to
the thinner cultures in the wells than in a regular cuvette. B. Colony forming unit in
OD550 = 0.5 cultures in a time course after addition of H2O2 to the indicated final
56
concentrations at 0 min. Colony forming units were determined as described in Materials
and methods. The mean and standard error of triplicates were plotted. C. Colony forming
unit in OD550 = 0.01 cultures in a time course after treatment with indicated
concentrations of H2O2. The mean and standard error of triplicates were plotted.
Table 1. Cell death upon H2O2 insult*
H2O2 (mM)
% of CFU ± standard error
30 min
60 min
0
100 ± 1.2
100 ± 1.0
1
91.8 ± 0.2
83.5 ± 2.5
2
82.0 ± 0.9
70.6 ± 1.1
5
57.8 ± 0.2
42.4 ± 0.4
10
0.8 ± 0.6
0.04± 0.04
Cultures of OD550 = 0.5 were treated with the indicated concentration of H2O2. The CFU
of control cultures (0 mm H2O2) was set at 100 %.
(Data was taken from Fig. 7B)
57
Table 2. Steady state levels of RNA oxidative damage in E. coli in response to H2O2
treatment
8-oxo-G/105
Ga
8-oxo-G RNAb
Damaged RNAc
Cell viability
(CFU)a
Normal aeration
1.0 ± 0.1
0.25%
2.4%
(set to 100%)
Plus 1 mM H2O2
3.9 ± 0.1
0.98%
9.3%
83%
Plus 5 mM H2O2
10.9 ± 0.5
2.7%
24%
42%
Growth
condition
a
8-oxo-G levels were determined 15 min after addition of H2O2 to exponentially growing
cultures. CFU was determined after treatment with H2O2 for 60 min. bBased on the
assumption that the length of RNA is 1 kb in average, the GC content is 50%, and 8-oxoG is randomly distributed so that the percentage of 8-oxo-G containing RNA can be
described by Poisson distribution.
c
Based on the assumption that total damage
occurrences can be 10 times of 8-oxo-G content.
(8-oxo-G/105G data was taken from Fig. 1B, all other calculations were done by
Zhongwei Li)
Oxidation of cellular RNA by exogenous H2O2 takes a very short time to happen,
suggesting that RNA is a direct target of ROS (Fig. 2). After H2O2 addition, 8-oxo-G
levels remain increased throughout the entire time course examined (Fig. 2A), although
H2O2 levels in the culture quickly became undetectable in minutes (Wu et al., 2009; data
not shown). A small decrease in the level of 8-oxo-G 60 min after the addition of H2O2
has been consistently observed. The H2O2 –induced 8-oxo-G levels reduced rapidly after
the cells were shifted to fresh H2O2 -free medium. It is likely that degradation or repair
58
reduces the level of 8-oxo-G containing RNA, whereas residual ROS derived from H2O2
would cause an increase in the level of 8-oxo-G. The observed steady state levels of 8oxo-G in RNA induced by continuous H2O2 treatment must reflect the equilibrium of the
two processes.
An unexpected observation was that rRNA and tRNA are not protected by their
structures or by proteins associated with rRNA (Fig. 4). The inability of RNA structure to
protect RNA from oxidation was further shown in vitro by exposing purified RNA to
H2O2 (Fig. 5). These findings are surprising because one would expect that RNA
structure and protein binding would limit the accessibility of RNA to ROS. Apparently,
ROS can reach the bases in these RNAs efficiently. One possible explanation for the lack
of protection is the association of the highly structured RNAs with Fe2+, the ion known to
generate oxidative chemicals and free radicals from H2O2 by Fenton chemistry
(Wardman and Candeias, 1996). Indeed, such association of rRNA with Fe2+ has been
reported to play a role in promoting rRNA oxidation and inactivation in translation
(Honda et al., 2005).
Importantly, under normal conditions rRNAs isolated from the ribosomes contain
much lower levels of 8-oxo-G than the rest of the cellular RNA (Fig. 4), suggesting that
rRNA in ribosomes are normally kept with low oxidative damage. This phenomenon
could be important for optimal ribosome function to support cell growth because
oxidative damage might hinder protein synthesis or generate errors in the protein
products.
59
A number of the observations in this work also suggest the existence of RNA
quality control activities that remove oxidized RNA and support cells surviving OS
insults. First, the high levels of 8-oxo-G generated by H2O2 treatment must eventually be
reduced to a normal level, at least in the viable cells. Second, given the fact that
ribosomal RNA normally contains low 8-oxo-G but is efficiently oxidized by H2O2,
oxidized rRNA species might be identified and removed from ribosomes. Such
mechanism(s) might not only be responsible for the removal of 8-oxo-G in ribosomes
(Min Liu and Zhongwei Li, unpublished observations) but could also be used for
reducing 8-oxo-G in rRNA under normal conditions.
Degradation of oxidized RNA can be a major quality control pathway because
until now a repairing mechanism is only found for alkylated RNA. As reported
previously, E. coli polynucleotide phosphorylase (PNPase) plays a pivotal role in
controlling 8-oxo-G levels and supporting cell viability under OS (Wu et al., 2009). In
addition, other RNA degradation activities also are important for reducing 8-oxo-G and
protecting cells under OS (unpublished results from our laboratory). In addition to
elevated levels of 8-oxo-G, we have also observed increased fragmentation of rRNA in
response to H2O2 treatment (Fig. 6). These rRNA fragments might be removed from
ribosomes and eventually degraded. The detailed molecular mechanisms for the
elimination of oxidized RNA, possibly involving specific recognition and targeted
degradation of such RNA molecules have yet to be elucidated.
Under the same conditions that H2O2 causes ~ 0.7 8-oxo-dG/105 dG in DNA, 8oxo-G rises by 6-7 per 105 G in RNA at steady state (Fig. 3). The relatively lower level of
60
H2O2 induced 8-oxo-dG in DNA might be due to strong DNA repair activities that
quickly remove 8-oxo-dG after its formation. Such repair activities have not yet been
reported for RNA. In contrast, oxidized RNA might undergo rapid degradation resulting
in the observed steady state level of 8-oxo-G in RNA. Considering that the amount of
RNA is approximately four times that of DNA, the oxidized guanine in RNA can be
much higher than that in DNA under OS conditions used in this work. The presence of
such large amount of oxidized nucleotides in RNA and the possible turnover of oxidized
RNA might be an important antioxidant mechanism which consumes the majority of
nucleotide oxidizing agents and reduces DNA damage (Radak and Boldogh, 2010).
These potential mechanisms should be defined in further studies.
Moreover, increased RNA damage might attenuate cell growth. A correlation
between the levels of RNA oxidation and cell growth reduction is proposed (Table 2).
Assuming that RNA is oxidized randomly and that total damage in RNA is
approximately ten times higher than the levels of 8-oxo-G (Gajewski et al., 1990; Rhee et
al., 1995; Cooke et al., 2003; Li et al., 2006), a significant fraction of RNA molecules
might be damaged by H2O2 at millimolar concentrations at steady state. It is likely that
RNA oxidative damage contributes, at least partly, to the observed cell death under the
same OS conditions (Table 2). Currently, there is no clear explanation about how
oxidized RNA would reduce cell viability under oxidative stress conditions. It was
previously reported that oxidized rRNA and mRNA are defective in protein synthesis in
vitro or in cells transfected by oxidized RNA (Ding et al., 2005; Honda et al., 2005,
Tanaka et al., 2007; Shan et al., 2007). Lack of protein synthesis and production of
61
aberrant proteins due to RNA oxidation are likely a main cause of cell death. We have
reported that PNPase, an exoribonuclease that binds to 8-oxo-G RNA with specificity
(Hayakawa et al., 2001) and degrades defective RNA (Li et al., 2002), is important for
controlling the level of 8-oxo-G in RNA and for protecting E. coli cells under OS (Wu et
al., 2009). In the absence of this enzyme, E. coli cells become hypersensitive to H2O2 and
other oxidants, and contain elevated levels of 8-oxo-G in RNA (Wu et al., 2009). These
findings strongly support the notion that oxidized RNA is deleterious to cells.
3.2 RNA structures promote the formation of 8-hydroxyguanosine
Despite the apparent importance of RNA oxidation to living organisms, little is
known about factors that may affect the extent of RNA damage caused by ROS. It has
been suggested that different RNA species may undergo different levels of oxidation
(Shan et al., 2003; Honda et al., 2005). If this is true, RNA function may be affected by
oxidation differently depending on its sequence or structure. It has been postulated that
RNA structure and its association with proteins may reduce ROS accessibility and
prevent RNA oxidation (Li et al., 2006; Wurthmann and Wolin, 2009). In a growing cell,
highly structured rRNA and tRNA account for nearly 80% and 15% of the total RNA
respectively. Most of the rRNA molecules are tightly bound with ribosomal proteins to
form ribosomes. In contrast to the abovementioned postulation, we have recently shown
that RNA structure and its association with proteins do not seem to protect RNA from
oxidative damage. After Escherichia coli cells were exposed to hydrogen peroxide
(H2O2), RNA isolated from ribosomes often contained higher levels of 8-oxo-G than
RNA from the non-ribosomal fraction (Liu et al., 2012). In addition, highly structured
62
RNA species are not protected from oxidation in vitro (Liu et al., 2012). These
observations prompted us to investigate the potential relationship between RNA structure
and levels of RNA oxidation.
3.2.1 H2O2 induces higher levels of 8-oxo-G in ribosomal RNA than in nonribosomal RNA
Recently, we have reported that exposure of E. coli cells to H2O2 for 15 min
generated somewhat higher levels of 8-oxo-G in RNA isolated from ribosomes than in
RNA from non-ribosomal fraction (Liu et al., 2012). While RNA prepared from
ribosomes is essentially pure rRNA, RNA from non-ribosomal fraction contains mRNA,
tRNA, non-coding RNA, intermediates of RNA processing and degradation, and rRNA
that has not been assembled into mature ribosomes. In order to better understand this
phenomenon, we have examined 8-oxo-G levels in these two RNA fractions in a time
course after E. coli cultures were exposed to H2O2. Figure 8A shows 8-oxo-G levels in
control and H2O2–treated E. coli cultures. In control cultures (-H2O2), 8-oxo-G is lower in
RNA from ribosomes than in RNA from non-ribosomal fraction during the entire time
course, suggesting that cells normally maintain low damage levels in ribosomal RNA.
Upon treatment with H2O2, the RNA 8-oxo-G reached its highest levels in only 5 min.
The levels of 8-oxo-G in ribosomes were higher than those in non-ribosomal fractions
throughout the entire time course. Importantly, the greatest difference in 8-oxo-G levels
between these two RNA preparations was found at the earliest time point. Such quicklygenerated difference must have been caused directly by more efficient oxidation of
ribosomal RNA. It should also be noted that the level of H2O2-induced 8-oxo-G in
63
ribosomes decreased ~20% at 15 min from the level observed at 5 min, while those in
non-ribosomal fraction remained the same during this period of time. Whether the
reduction of 8-oxo-G-containing RNA at the later time points is due to its degradation
remains to be determined. It is likely that 8-oxo-G-containing RNA in ribosomes is first
released into non-ribosomal fraction before it is completely eliminated.
A.
B.
6
4
200
100
0
H2O2
2
0
H2O2
300
-
Time (min)
+
5
+
-
+
15
+
-
+
60
+
-
+
-
+
Non-ribosomal
RNA
Ribosomal RNA
Non-ribosomal RNA
400
Ribosomal
RNA
8-oxo-G/105 G
8
8-oxo-G/105 G
500
Figure 8. H2O2 induces higher levels of 8-oxo-G in ribosomal RNA than in nonribosomal RNA in vivo and in vitro. A. Exponentially growing cultures of E. coli
CA244 rna (OD550= 0.5) were supplemented with H2O2 to 1mM (+H2O2) or an equal
volume of YT (-H2O2) at 0 min. Ribosomal and non-ribosomal RNA were extracted and
8-oxo-G levels were analyzed as described in Materials and Methods. B. Purified
Ribosomal and non-ribosomal RNA preparations were treated in vitro with (+H2O2) or
without (-H2O2) 1 mM H2O2 in a 100 µl mixture containing 20 µg RNA. 8-oxo-G levels
were analyzed as described in Materials and Methods. The mean and standard error of at
least three replicates were plotted.
64
Based on the above observations, we predicted that H2O2 may cause greater
oxidation to ribosomal RNA than to non-ribosomal RNA in their purified forms. Sure
enough, after incubating these RNA preparations with H2O2, ribosomal RNA produced
two-fold higher 8-oxo-G than non-ribosomal RNA (Fig. 8B). The results provided direct
evidence for more efficient formation of 8-oxo-G in ribosomal RNA than in nonribosomal RNA, contrary to what has been previously speculated (Li et al., 2006).
The data suggest that the observed differences in H2O2-induced 8-oxo-G levels in
these RNAs is most likely caused by the difference in RNA composition, although an
effect of the RNAs’ respective cellular environments cannot be eliminated under in vivo
conditions. Ribosomal RNA is rich in higher-order structures, raising the possibility that
certain structural features may enhance the formation of 8-oxo-G under OS. In addition,
tRNA tends to form less 8-oxo-G compared to rRNA (see below), contributing to the
lower 8-oxo-G levels observed in non-ribosomal RNA.
3.2.2 Oxidation of rRNA and tRNA is inversely correlated to the extent of
denaturation
In order to test if the preferential formation of 8-oxo-G in ribosomal RNA is
related to its highly structured feature, native and heat-denatured rRNA were oxidized in
vitro and 8-oxo-G levels were determined. As shown in Fig. 9A, H2O2-induced 8-oxo-G
level is higher in native rRNA than in denatured rRNA. Longer heat treatments result in
lower 8-oxo-G levels, presumably due to more complete denaturation of rRNA. In the
control reactions without H2O2, heat treatment per se did not have much effect on 8-oxoG levels. These results clearly demonstrated a role of RNA structures for promoting 865
oxo-G formation. The effect of RNA structures may account for the production of at least
~40% of 8-oxo-G comparing the levels of 0 and 8 min denaturation (Fig. 9A). Because
the RNA was denatured by heating prior to incubating with H2O2, it is likely that some of
the denatured RNA molecules may re-nature during oxidation, making it likely that the
actual level of structural effect is even higher than shown here.
A.
B.
200
rRNA + H2O2
150
100
50
8-oxo-G/105G
8-oxo-G/105 G
100
80
tRNA +H2O2
60
40
20
rRNA - H2O2
tRNA -H2O2
0
0
0
2
4
6
0
8
Heat denaturation (min)
2
4
6
8
Heat denaturation (min)
(tRNA by Ravi Kumar Alluri)
Figure 9. Oxidation of rRNA and tRNA is inversely correlated to the extent of
denaturation. rRNA and tRNA were prepared from purified ribosomes and total RNA
respectively as described in Materials and Methods. The RNA samples were dissolved in
the buffer for in vitro oxidation without H2O2. RNA denaturation was carried out by
incubating at 95 °C for the indicated amount of time and then immediately chilled in an
ice water bath. The heat-denatured RNA was then immediately oxidized by the addition
of H2O2 to a final concentration of 1 mM in a 100 µl mixture containing 20 µg rRNA or
tRNA. 8-oxo-G levels were determined as described in Materials and Methods. The mean
66
and standard error of at least three replicates were plotted. A. 8-oxo-G levels generated in
rRNA. B. 8-oxo-G levels generated in tRNA.
Interestingly, similar patterns of H2O2-induced 8-oxo-G production were observed
for native and denatured tRNA samples. In this case, denaturation by 8 min heating
caused a reduction of ~60% of H2O2-induced 8-oxo-G (Fig. 9B).We noted that tRNA is
oxidized to a lower level than rRNA in their native forms, even though higher-order
structures in tRNA are still important for 8-oxo-G formation.
3.2.3 H2O2–treatment generated less 8-oxo-G in single-stranded RNA than in
RNA:DNA duplex
Although the data shown in Figure 8 and 9 strongly suggest that certain RNA
structures promote 8-oxo-G formation on H2O2 treatment, a few things need to be
clarified. First, cellular RNA changes constantly due to the synthesis of new RNA and the
degradation of existing RNA, making it difficult to evaluate steady state levels of 8-oxoG. Second, purified RNA samples from cells may contain various components that affect
differently the efficiency of H2O2-induced 8-oxo-G production. Finally, both rRNA and
tRNA contain various types of higher-order structures. It is difficult to evaluate how
different structural features in these RNA species affect their ability to be oxidized.
To better understand the problem, we have analyzed H2O2-induced 8-oxo-G
production in synthesized RNA in vitro. The materials used in this experiment are a
synthetic 50-mer RNA oligonucleotide that does not show any predictable structure, and
an RNA:DNA duplex formed by this RNA and a complementary DNA. The synthetic
67
oligonucleotides are free of contamination of any cellular components that may
complicate the outcome of in vitro oxidation. In addition, the effect of double-stranded
structure on oxidation can be directly assessed without interference of other structures.
The reason for using an RNA:DNA duplex rather than an RNA:RNA duplex is to avoid
complications regarding the origin of 8-oxo-G.
Under the conditions used for this experiment, DNA only produces negligible
amount of contaminating guanosine and 8-oxo-G. Therefore, the 8-oxo-G detected in
RNA:DNA duplex must be from RNA. Interestingly, treatment with H2O2 generated
more 8-oxo-G in the RNA:DNA duplex than in the single-stranded RNA (Fig. 10). In
oxidation reactions containing the same amount of total nucleic acids, the RNA:DNA
duplex produced a much higher level of 8-oxo-G than the single stranded RNA. We also
noted a reproducible increase of 8-oxo-G in the RNA:DNA duplex in the control reaction
without H2O2.When the same amount of RNA was used for both the single-stranded
RNA and the RNA:DNA duplex, the duplex containing double amount of total nucleic
acids still produced a moderately higher level of 8-oxo-G than the single-stranded RNA.
In the same reaction, a fraction of H2O2 must have been consumed by the oxidation of the
DNA strand although the level of DNA oxidation cannot be evaluated at this time. It is
therefore safe to conclude that compared to single stranded RNA, the same amount of
RNA in the duplex was oxidized to higher level by less H2O2.The data demonstrate that
the double-stranded structure is able to promote RNA oxidation, and suggest that doublestranded structure is probably a major cause of the more efficient 8-oxo-G formation
observed in highly structured cellular RNA.
68
8-oxo-G/105 G
800
600
400
200
0
H2O2
-
+
-
+
-
+
RNA RNA:DNA RNA:DNA
0.5X RNA 1X RNA
Figure 10. H2O2 induces higher levels of 8-oxo-G in RNA:DNA duplex than in
single-stranded RNA of the same sequence. The 50-mer single-stranded RNA and
RNA:DNA duplex were treated with 1 mM H2O2 or with buffer alone in a 100 µl mixture
as described in Materials and Methods. After incubation with H2O2, the oligonucleotides
were precipitated through the addition of 20 µl 1 M NaAc and 720 µl absolute ethanol,
and was kept at -80 °C for 4 h. Nucleic acids were pelleted at 20,000g for 15 min at 4 °C,
washed with 80% cold ethanol and dissolved in DEPC- and Chelex 100-treated H2O
before the analysis of 8-oxo-G levels. 8-oxo-G levels in all samples were determined and
the mean and standard error of at least three replicates were plotted. Single-stranded RNA
(21 µg or 655 pmol, labeled as RNA), an equal amount of RNA:DNA (327.5 pmol RNA
plus 327.5 pmol DNA, labeled as RNA:DNA 0.5 X RNA) and a double amount of
RNA:DNA (655 pmol RNA plus 655 pmol DNA, labeled as RNA:DNA 1 X RNA) were
shown.
69
3.2.4 Cu2+ bound different forms of nucleic acids with different affinity
Previous studies indicated that the difference between the 8-oxo-G levels in native
and denatured RNA is related with the varied iron binding capacity. It was suggested that
higher iron binding may cause more efficient oxidation of RNA by the ROS generated
through iron-mediated Fenton reaction (Honda et al., 2005). It is reported that iron
binding capacity was significantly higher in rRNA than that in tRNA and mRNA.
Moreover, iron-binding was dramatically decreased in the denatured rRNA and tRNA to
the same level, suggesting a role for the secondary or a higher structure of RNA (Honda
et al., 2005). In our experiment, Fe2+/Fe3+ was replaced by copper (Cu2+) which is known
to catalyze Fenton reaction similarly by the Cu+/Cu2+ conversions. In order to understand
whether Cu2+ binding capacity is related to 8-oxo-G levels observed in various RNA
preparations in this work, we have determined the affinity of these RNAs and copper.
As shown in Figure 11, native rRNA bound three-fold more copper than
denatured rRNA. This result is consistent with the notion that higher affinity to Cu2+ is
correlated with the resulting higher levels of 8-oxo-G in rRNA. In addition, native tRNA
bound three-fold less copper than native rRNA, which may be partially responsible for
the lower 8-oxo-G level of native tRNA compared to native rRNA. In contrast, native
tRNA bound similar level of copper with denatured tRNA. Furthermore, RNA:DNA
duplex bound less copper than the single stranded RNA. In the latter cases, 8-oxo-G
levels in the more structured forms of RNA, i. e., native tRNA and RNA:DNA duplex,
were higher than those in the corresponding less structured forms, i. e., denatured tRNA
or single-stranded RNA. Therefore, affinity with copper is not related to 8-oxo-G levels
70
produced in tRNA and RNA:DNA duplex. Our results are essentially different with the
previously reported results of a strong correlation between iron binding capacity and
metal ion binding (Honda et al, 2005). Copper binding capacity may have affected the
level of 8-oxo-G only in the case of rRNA, but not in the cases of other RNAs tested
here. Alternatively, copper binding may not be related to 8-oxo-G formation at all. For
instance, the variation in 8-oxo-G levels may have been due to other reasons such as
RNA structures per se. The effect of RNA structure on 8-oxo-G formation remains to be
studied further.
0.2
1.5
(µmol/10 µg
RNA:DNA)
0.8
4
0.6
4
2
(µmol/10 µg RNA)
3
C.
2+
B.
2+
Cu
Cu
0.4
Cu
1
2+
4
(µmol/10 µg RNA)
A.
0
0.0
N
D
rRNA
N
D
tRNA
1.0
0.5
0.0
RNA
RNA:DNA
Figure 11. Cu2+ affinities for various nucleic acids. 40 µM CuSO4 was incubated with
different forms of nucleic acid as described in Materials and Methods. Copper
concentrations were determined by the plate reader. The copper concentrations per unit
amount of rRNA, tRNA, single stranded RNA, and RNA:DNA (0.5X RNA) were shown
as mean and standard error for three replicates. A. copper binding to the native and
denatured rRNA. B. copper binding to the native and denatured tRNA. C. copper binding
to the single stranded RNA and RNA:DNA duplex. N: native; D: denatured.
71
3.2.5 Discussion
The results of this study suggest a surprising role for higher-order structure of
RNA in promoting oxidative damage. Such structural features may include, but are not
limited to, double-stranded structures. Compared to single strand RNA, double strand
may recruit ROS more efficiently or react with ROS at higher rate. Other structural
feature and sequence context may also affect oxidation of RNA, making it possible for
“hot spots” of ROS target to exist. For the same reason, some RNA molecules may
contain higher levels of oxidative lesions than others, causing differential effect on RNA
function under OS. The exact nature of how each of the sequence and structural features
affect the formation of various oxidative damages in RNA deserve to be studied in the
future.
The observation that ribosomal RNA is highly oxidized shortly after the cells’
exposure to H2O2 suggests that preferential oxidation of highly structured RNA may also
happen in vivo. Because highly structured rRNA and tRNA constitute the majority of
cellular RNA, high levels of oxidation of these RNAs would present a major challenge to
any living organism. Efficient RNA surveillance mechanisms may play pivotal roles on
cell survival under OS, but such important mechanisms remain to be elucidated.
Some metal ions in the cell may copurify with rRNA and tRNA, which affected
the copper binding and also the 8-oxo-G level in RNA. 8-oxo-G levels in native rRNA
and tRNA were higher than that in the corresponding denatured RNA even without
adding copper, although copper dramatically increased the 8-oxo-G level in the native
RNA (data not shown). However, metal ion contamination did not occur in single
72
stranded RNA and RNA:DNA duplex. Single stranded RNA bound more copper than the
RNA:DNA duplex, but gained less 8-oxo-G than the duplex. Therefore, it is difficult to
evaluate the effect of copper binding capacity to the oxidation of nucleic acid.
3.3 Exoribonucleases play an important role in eliminating oxidized RNA in
ribosome
One important fact is that under normal growth conditions, RNA in ribosome
fractions contains lower 8-oxo-G than RNA from the non-ribosome fraction, indicating
that cell maintains a high quality of RNA in ribosomes. When cells are exposed to H2O2,
RNA in the ribosome is quickly oxidized to higher levels than non-ribosomal RNA. This
result suggests that under normal conditions, the low 8-oxo-G content in ribosomal RNA
is not due to protection of rRNA against oxidation. It also suggests that once oxidized
RNA forms in ribosomes, it can be removed efficiently from ribosomes. As shown in
Figure 6 and 15, H2O2 treatment causes rRNA degradation, suggesting a plausible
mechanism by which selective degradation of oxidized RNA in the ribosomes helps
maintain rRNA quality under oxidative stress.
3.3.1 H2O2–induced ribosomal 8-oxo-G level decrease after removal of the oxidant
To test the idea of selective elimination of 8-oxo-G in ribosomal RNA, we have
carried out analysis of 8-oxo-G levels in ribosomal RNA in a time course after cells were
exposed to a continual or a pulse H2O2 treatment. As shown in Figure 13A, continuous
H2O2 treatment produced increased levels of 8-oxo-G in both ribosome and non-ribosome
fractions which lasted the entire time course, with a slight reduction at the end of 60 min
73
after an addition of H2O2. Ribosomal 8-oxo-G level is 60% to 100% higher than nonribosomal 8-oxo-G at every time point. This is similar to the pattern observed in Figure
8A.
Importantly, a pulse H2O2 treatment in the first 15 min caused an initial increase
of 8-oxo-G, which was followed by a sharp reduction of 8-oxo-G after removal of the
oxidant (Fig. 13A). The level of ribosomal 8-oxo-G was increased in the first 15 min in
the presence of H2O2. After removal of H2O2, it was reduced by ~45% at 30 min and
~80% at 60 min time points, respectively. The level of non-ribosomal 8-oxo-G was also
initially increased after 15 min and then reduced by ~70% at 30 min and 60 min. At the
end of 60 min, 8-oxo-G levels in ribosome and non-ribosome fractions were at almost the
same low levels, though they were still higher than the levels before H2O2 treatment. This
behavior strongly suggested that H2O2-induced 8-oxo-G is quickly removed in cellular
RNA. It should be noted that the experimental condition allowed cells to grow after the
pulse H2O2 treatment. Considering that the generation time of the E. coli strain is ~30
min under this condition, dilution of 8-oxo-G from newly synthesized RNA may explain
part of the observed decrease in 8-oxo-G. However, the sharp reduction of 8-oxo-G in
both ribosome and non-ribosome fractions must also be caused by selective elimination
of 8-oxo-G-containing RNA in vivo.
3.3.2 Specific removal of 8-oxo-G is blocked by deficiency in RNA degradation
In order to understand if degradation is responsible for the selective elimination of
oxidized RNA, we have studied the rate of 8-oxo-G reduction in mutants lacking RNA
degradation activities. Two RNA degradation pathways are speculated to play critical
74
roles in the removal of damaged RNAs (Fig. 12). One is composed of RNase R and PAP,
and another is PNPase by RhlB. PAP adds a poly(A) tail that helps RNase R bind and
degrade structured RNA. RhlB opens up RNA double-strand to facilitate degradation by
PNPase.
These two pathways may both play important roles in the exonucleolytic
degradation of RNA fragments. In the absence of one pathway, the other can take over
the task. Because a cell lacking both PNPase and RNase R is non-viable, we have chosen
to study the behavior of pnp pap and rnr rhlB mutants which lack one of the RNases and
an enzyme facilitating the activity of the other RNase. Therefore, we anticipated that at
least some structured RNAs accumulate in these mutant cells.
Figure 12. Proposed oxidized RNA quality control model. Oxidized RNA could be
firstly accessed by endo-RNase and cleaved into fragments. These fragments are
continually degraded by two pathways, RNase R and PAP or PNPase and RhlB, and
eventually converted to mononucleotides by oligoribonucleases. Dark closed rectangle:
the oxidation damage.
75
We have studied 8-oxo-G levels in ribosomal and non-ribosomal RNA in the
mutants after a continuous or pulse treatment with H2O2 (Fig. 13B and 13C). The results
from the mutants were compared to those from the wild type (Fig. 13).
In the mutant lacking both RNase R and RhlB, continuous and pulse H2O2
treatments caused a sharp increase of ribosomal 8-oxo-G levels at first and then a slow
decrease later (Fig. 13B). Importantly, after a pulse treatment with H2O2, the 8-oxo-G
levels in the ribosomal RNA remained much higher in the mutant than in the wild type.
This indicates that the deficiency of the RNA degradation pathways affects the removal
of oxidized rRNA.
In the case of non-ribosomal RNA, 8-oxo-G levels remained high after continuous
treatment with H2O2 in both rnr rhlB and wild type cells. In contrast, 8-oxo-G levels after
pulse H2O2 treatment decreased much slower in the mutant than in the wild type. This
strongly suggests that RNase R and RhlB also play a role in the removal of nonribosomal 8-oxo-G.
Deficiency in PNPase and poly(A) polymerase also affected 8-oxo-G reduction
even more dramatically (Fig. 13C). After continuous H2O2 treatment, both ribosomal and
non-ribosomal 8-oxo-G levels were higher in the mutant than in the wild type. After the
pulse treatment, ribosomal 8-oxo-G decreased very slowly whereas non-ribosomal 8-oxoG increased slightly over time in the pnp pap cells. These results suggest that PNPase and
poly(A) polymerase plays a somewhat more important role than RNase R and RhlB in the
degradation of 8-oxo-G containing RNA.
76
A. wild-type (wt)
wild-type
6
5
8-oxoG/10 G
5
4
continuous H2O2 ribosomal RNA
3
continuous H2O2 non-ribosomal RNA
2
pulse H2O2 ribosomal RNA
1
pulse H2O2 non-ribosomal RNA
0
0
15
30
45
60
time (min)
B. rnr rhlB
C. pnp pap
rnr rhlB
6
pnp pap
6
5
8-oxoG/10 G
4
5
5
8-oxoG/10 G
5
3
2
1
4
3
2
1
0
0
15
30
45
0
60
0
time (min)
15
30
45
60
time (min)
Figure 13. The alteration of 8-oxo-G level in ribosomal and non-ribosomal RNA in
wild-type and mutant cells with continuous or pulse H2O2 treatment. Exponentially
grown cultures of E. coli strains, wt (CA244 rna) and mutants (rnr rhlB and pnp pap),
were treated with 1 mM H2O2 for 15 min when OD550 reached 0.5. Cells were grown
continually and collected in the indicated time course (continuous H2O2); or cells were
centrifuged, resuspended in fresh H2O2 -free YT medium and collected in the indicated
time course (pulse H2O2). Ribosomal and non-ribosomal RNAs were extracted, and 8oxo-G levels were measured as described in Materials and methods. A. 8-oxo-G levels in
77
ribosomal RNA and non-ribosomal RNA in wild-type (wt) cells. B. 8-oxo-G levels in
ribosomal RNA and non-ribosomal RNA in rnr rhlB. C. 8-oxo-G levels in ribosomal
RNA and non-ribosomal RNA in pnp pap.
It should be noted that the exonucleolytic activities supposedly degrade RNA
fragments that are produced by initial endonucleolytic cleavages. As shown in 3.4,
deficiency in PNPase and RNase R caused the accumulation of rRNA fragments. The
observation that inactivation of these RNases affects ribosomal 8-oxo-G reduction
suggests that these exoribonucleases play a role in degrading oxidized RNA in the
ribosomes. One explanation is that some rRNA fragments are present in the ribosomes
and their degradation depends on these two exoribonucleases. Alternatively, PNPase and
RNase R may be involved in degrading intact rRNAs in ribosome. These possibilities
require further investigation.
3.4 Degradation of 16S and 23S ribosomal RNA under oxidative stress in
Escherichia coli
It is known that the majority of oxidative damage to cellular nucleic acids is
present in RNA, and only a small portion is in DNA (Liu et al., 2012). Ribosomal RNA
(rRNA) is the majority of cellular RNA, accounting for nearly 80% of total RNA in
actively dividing cells. It has been suggested that the ribosome structure protects rRNA
from degradation and chemical damage (Li et al., 2006). However, this is not the case for
RNA damage under oxidative stress conditions.
78
We have recently found that under normal growth conditions, E. coli rRNA
isolated from ribosomes contains much less oxidized RNA than RNA isolated from nonribosomal fractions. Levels of 8-hydroxyguanosine (8-oxo-G), an oxidized form of
guanosine, are three-fold lower in ribosomal RNA than in non-ribosomal RNA (Liu et al.,
2012). Surprisingly, after cells are exposed to an oxidant, ribosomal RNA can be quickly
oxidized to levels higher than non-ribosomal RNA, suggesting that neither ribosome nor
higher order RNA structure protects rRNA from attack by free radicals (Liu et al., 2012,
unpublished observations). These observations demonstrated that ribosomal RNA is
oxidized no less than non-ribosomal RNA. However, oxidized rRNA may be removed
quickly to maintain lower RNA oxidation levels in ribosomes under normal steady-state
conditions. This proposal is supported by our recent work showing ribosomal 8-oxo-G
level rise quickly in 15 min after addition of hydrogen peroxide (H2O2), followed by a
sharp decrease in a time course (Fig. 13). In contrast, 8-oxo-G in non-ribosomal RNA
decreased at a slower rate. Interestingly, both RNase R and PNPase appear to play a role
in the reduction of ribosomal 8-oxo-G levels, strongly suggesting that the decrease of
ribosomal 8-oxo-G is caused by selective degradation of oxidized rRNA.
To date, nothing about degradation of oxidized rRNA has been reported in
literature. However, rRNA degradation under other conditions has been documented in E.
coli previously. In general, rRNA in an intact ribosome is stable. However, rRNA that is
not associated with ribosomal proteins and rRNA that has been incorrectly assembled in
preribosome particles are degraded rapidly by quality control activities. PNPase and
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RNase R have been shown to participate in quality control of rRNA by degrading rRNA
fragments (Cheng and Deutscher, 2003; Basturea et al., 2011).
Under certain conditions, even rRNAs in intact ribosomes are completely
degraded. Under starvation of carbon source, rRNA is thought to be fragmented first by
endonucleolytic cleavages, followed by exonucleolytic activity which degrades rRNA
fragments into mononucleotides (Kaplan and Apirion, 1975). It is found that this
degradation is triggered by increased free ribosome subunits (Zundel et al., 2009),
suggesting that rRNA degradation under such conditions is a mechanism for eliminating
unused ribosomes.
It has been shown that free ribosome subunits are cleaved by
endoribonuclease at defined positions. The rRNA fragments are then degraded by
exoribonucleases, mainly RNase R and RNase II (Basturea et al., 2011). In the absence of
these exoribonucleases, the rRNA fragments accumulated to high level.
Although rRNA degradation under starvation and rRNA quality control both use
the similar exoribonucleases to clean up rRNA fragments, they differ in a number of
details. Firstly, the exact positions of the endonucleolytic cleavages on 23S rRNA are
different in the two degradative processes. Secondly, RNase R and II are most important
in removing rRNA fragments during starvation, whereas RNase R and PNPase are
important in the quality control of rRNA. Finally, rRNA degradation under starvation is
initiated by shortening of the 3’ end of 16S rRNA by RNase PH in the intact ribosome.
This will remove the rRNA sequence that interacts with the Shine-Dalgarno sequence of
mRNA and therefore inactivate the ribosome. The inactivated ribosome falls apart and
the ribosomal subunits are degraded by the endo- and exo- RNases mentioned above. In
80
contrast, quality control only degrades rRNA that may be trapped in incorrectly
assembled ribosome particles, and the process is independent of RNase PH (Basturea et
al., 2011).
In this work, we examined if any of the activities participating in rRNA
degradation under starvation or rRNA quality control are involved in the rRNA
degradation pathways under oxidative stress. Such activities are expected to play a major
role in eliminating the majority of oxidized RNA in E. coli cells.
3.4.1 rRNA fragments are detected in H2O2-challenged E. coli cells
In a recent study, we have shown that ribosomal RNA contains high levels of 8oxo-G shortly after cells were exposed to H2O2 (Liu et al., 2012). We have also observed
fast and selective elimination of 8-oxo-G containing ribosome RNA in a time course. To
examine whether oxidative stress induces rRNA degradation, we have carried out
Northern blotting to examine any rRNA fragment that are formed after exposure of
exponentially grown E. coli cultures to H2O2.
As shown in Figure 14A, treatment with 5 mM H2O2 for 30 min caused the
production of some new RNA bands in the wild type cell. At the 180 min time point, the
abundance of these products was reduced. These products can also be observed in trace
amounts when 1 mM H2O2 was applied. These products are fragments of rRNA as
shown by Northern blotting using oligonucleotide probes that are complementary to
different regions of 16S and 23S rRNA (Fig. 15A and 16A). Some products are shown in
smears, suggesting that they are shortened by exonucleolytic trimming. The results
clearly demonstrated oxidation dependent degradation of rRNA. The steady-state levels
81
of these RNA products are relatively low in the wild type cell due to the presence of the
full array of degradation activities. Nevertheless, accumulation of these products
indicates that cells have to handle rRNAs that are presumably present in the damaged
ribosomes under oxidative stress. At the higher H2O2 dosage, the degradation activity
becomes limited due to higher levels of ribosome damage and more rRNA fragments
accumulate.
A. wild-type (wt)
B. rnr pnpts
C. rnb rnr pnpts
Figure 14. Accumulation of rRNA fragments at different time courses with various
concentrations of H2O2. The cells were grown at 30 °C until OD550 ≈ 0.5, and then
shifted to 42 °C (0 min). H2O2 with indicated concentrations (0, 1, 5 mM) was added into
the culture, and the cells were collected at indicated time course (0, 30, 180 min). Total
cellular RNA from wild type (wt, CA244 rna) or mutants additionally lacking
exoribonucleases (rnr pnpts and rnb rnr pnpts) was extracted, separated on a 1.5% agarose
gel, and visualized by ethidium-bromide staining. A. rRNA fragments accumulated in wt.
B. rRNA fragments accumulated in rnr pnpts lacking RNase R and PNPase. C. rRNA
fragments accumulated in rnb rnr pnpts lacking RNase II, R, and PNPase.
82
3.4.2 The three processive exoribonucleases play a role in degrading rRNA
fragments under oxidative stress
Because PNPase, RNase R, and RNase II have been shown to degrade rRNA
fragments under other conditions, we have examined their role in rRNA degradation
under oxidative stress. E. coli mutants lacking PNPase and RNase R, or all three
enzymes, were treated with H2O2, and rRNA products were detected by agarose gel
electrophoresis and Northern blotting. Several fragments accumulated in the mutants,
depending on H2O2 dosage and increasing over time (Fig. 14B and 14C). As indicated in
Northern blotting, these fragments are generated from full-length 23S and 16S rRNA
(Fig. 15 and 16). The 16S fragments in the two mutants are essentially the same (Fig.
14B, 14C, 15B, and 15C), although the relative intensity may vary. These results suggest
that RNase R and PNPase are the main degrading enzymes of rRNA fragments under
oxidative stress. RNase II may help degrade some of the fragments.
3.4.3 Degradation of 16S rRNA under oxidative stress
To better characterize rRNA degradation intermediates under oxidative stress, we
carried out Northern blotting to analyze the identity and size of these fragments and their
relative positions in the full-length rRNA. Primer extension and 3’ RACE were also
performed to determine the 5’ and 3’ termini of some RNA fragments. To analyze 16S
rRNA degradation, oligonucleotide probes complementary to various regions of 16S
rRNA were used to detect the individual fragments generated by endo- and/or exonucleolytic digestion. For convenience, the oligonucleotide probes are named by the
complementary region in the rRNA. For instance, probe 16S15-36 is complementary to
the region of 15-36 nt in 16S rRNA. Altogether, 5 bands were detected with 16S probes
83
in the rnr pnpts and rnb rnr pnpts mutants, and 3 of them were also seen in wild type (Fig.
15). Three probes, 16S15-36, 16S611-630 and 16S838-857, detected products that are
marked as band 1, 2 and 3. Because the ~900 nt band 1 is the longest and most abundant
fragment, the results suggest that the 5’ half of 16S rRNA is a key degradation
intermediate. A shorter product, band 2, was detected by probe 16S611-630 and 16S838857, but not by 16S15-36, suggesting that band 2 is formed from band 1 by shortening at
the 5’ end. Band 3 is even shorter than band 2 at the 3’ end because it was not detected by
16S838-857. The accumulation of these intermediates suggests gradual degradation of
the 5’ half of 16S rRNA, presumably involving both endo- and exoribonucleases.
The downstream fragments, bands 4 and 5, are detected by probes 16S951-970
and/or 16S1448-1467 in wild type and at increased abundance in the RNase-deficient
mutants. Band 4, ~650 nt in length, was detected by both probes, as well as a probe
16S893-912 and 16S1520-1539 (data not shown), indicating that it contains the 3’ end of
the 1542 nt full-length 16S rRNA and it has overlapping region of band 1. The shorter
band 5 was only detected by probe 16S951-970, and not by 16S1448-1497, indicating
that it lacks the 3’ end of 16S rRNA. The 5’ ends of band 4 and band 5 RNA must have
been generated by endonucleolytic cleavages because 5’->3’ exoribonuclease activity has
not been identified in E. coli. The sizes and locations of these fragments in 16S rRNA
strongly suggest that the 3’ half was degraded independently from the 5’ half. Because in
all strains, band 4 is more abundant at 30 min than at 180 min after addition of H2O2, it is
likely that this fragment is shortened to band 5 RNA by activity(ies) independent of these
84
A. wild-type (wt)
B. rnr pnpts
C. rnb rnr pnpts
D.
Figure 15. Northern blot analysis of 16S rRNA fragments. Total cellular RNA was
extracted in the same way as in Figure 14, and transferred to a nylon membrane.
85
Oligonucleotide probes complementary to various regions of 16S rRNA were used to
detect the individual fragments generated by the endo- and/or exo-nucleolytic digestion,
which are named by the complementary region in the rRNA and indicated in each panel.
A. 16S rRNA in wide type (wt, CA244 rna) cells. B. 16S rRNA in rnr pnpts cells. C. 16S
rRNA in rnb rnr pnpts cells. D. Diagrams show the location of the probes and the main
endonucleolytic cleavage sites (arrows). The weight of the lines indicates the amount of
the fragments. endo: endonucleolytic cleavage
three exoribonuclease. Band 5 RNA may be degraded by these three RNases since it
accumulates over time after inactivation of these enzymes.
The non-overlapping pattern of the degradation intermediates at the 5’ and 3’
halves of 16S rRNA strongly suggests that an endonucleolytic cleavage at a position
around 912 nt is a key step that precedes the degradation of the fragments. The two
halves of 16S rRNA are subsequently degraded by a combination of endo- and
exonucleolytic activities.
The termini of some of the RNA products were determined. 3’ RACE detects 3’
ends of some RNA products at residues A696, G894, C1303, and A1413 etc, which could
be the ends of some major or minor degradation intermediates. Primer extension using
RNA from the rnb rnr pnpts mutant detected RNA containing 5’ ends at A559, U920,
U921, A923, and C924. Residue A919 was reported as the major cleavage site in 16S
rRNA during starvation and quality control (Basturea et al., 2011). Our result is
consistent with the existence of this cleavage site, and suggests that 16S rRNA
86
degradation under oxidative stress may involve the same endonucleolytic cleavage as
under starvation.
The results of various experiments were summarized in a diagram shown in
Figure 15. Additional probes were used to reveal 16S related products. The results from
these probes were not shown in the Northern images, and are summarized together with
other experiments in the diagram.
3.4.4 Degradation of 23S rRNA under oxidative stress
The degradation intermediates of 23S rRNA were also analyzed under oxidative
stress conditions. Because the results of 16S rRNA from the rnr pnpts strain are very
similar with those from the rnb rnr pnpts mutant, we have only included the triple mutant
in the analysis of 23S rRNA. The results and a summary diagram are shown in Figure 16.
In wild type cells, probe 23S7-26 detected three products, marked as band 6, 7,
and 8. These products are more abundant in the rnb rnr pnpts mutant, suggesting a role of
the RNases in degrading these products. The long RNA fragment, band 6, is about 1650
nt in length, while another fragment, band 6*, was slightly longer and detected in the
mutant by probe 16S1689-1708. Band 7 is an abundant product of about 1000 nt in
length, and band 8 is about 700 nt. Probes 23S1689-1708 and 23S1919-1938 detected a
range of products with sizes around 1200 and they are marked as band 9. These products
were not detected by probe 23S7-26, indicating that these fragments lack the 5’ end of
23S rRNA. Because band 9 is not a sharp band, it may be composed of products of
similar sizes with slightly different ends. Band 6, 7, 8 and 9 are all dependent to the
87
dosage of hydrogen peroxide. It is likely that band 6 is converted to band 7, and the latter
is in turn converted into band 8 during the degradation process. The conversion of the
longer RNA to shorter products appears to be independent of the exoribonulceases
because it happens in both wild type and the mutant lacking the RNases. The abundance
of bands 6 and 8 RNA increases over time in the RNase-deficient mutant, suggesting the
production of these products is faster than their removal by these enzymes. The intensity
of band 7 is lower at 180 min than at 30 min, probably due to more efficient conversion
of band 7 RNA to band 8 RNA during the time course. Band 9 RNA may be formed
independently from band 6 RNA because the two RNA products are different at both
ends. Small amount of band 9 was detected by downstream probes, 23S2131-2150,
23S2608-2627 and 23S2885-2904, in the mutant, which may convert to shorter
downstream fragments.
The downstream probes, 23S2131-2150, 23S2608-2627 and 23S2885-2904,
detected only trace amount of smearing products that may cover the 5’ half of 23S RNA.
In contrast, we have detected products that are 1000 nt or shorter (band 10, 11, 12 and 13)
in the mutant and they do not overlap with any of the 5’ products, strongly suggesting the
existence of an endonucleolytic cleavage(s) that separates the 5’ half from the 3’ region.
These 3’ products must be degradation products created by the exoribonucleases
that are missing in the mutant because they are not detected in the wild type cell. The
distribution of these 3’ products also suggests the involvement of multiple secondary
endonucleolytic cleavages in breaking down the 3’ half before or during exonucleolytic
degradation. Particularly, all the products in bands 10, 12 and 13 contain the 3’ end of
88
23S rRNA, since they are detected by probe 23S2885-2904, which is complementary to
the 3’ end of the RNA. Band 11 RNA is very short, and was detected only by probe
23S2131-2150. Bands 12 and 13 RNAs are longer than band 11, and were not detected
by probe 23S2131-2150, suggesting that they are separated from band 11 RNA by
endonucleolytic activity. In addition, some minor products corresponding to the 5’ end of
band 10 RNA were also detected. All the shorter RNAs in this region may be generated
from band10 by multiple endonucleolytic cleavages. An endonucleolytic cleavage in the
middle of band 10 RNA may generate band 12 RNA and an upstream RNA. The
upstream RNA maybe quickly degraded by endo- and exoribonucleases to produce band
11 RNA and the short RNAs that cover the 5’end of band 10 RNA. Band 12 RNA may
be an intermediate that is quickly transformed into band 13 RNA by another
endonucleolytic activity.
3’ RACE experiment revealed the 3’ ends of several RNA products at residues
A705, A1014, G1620, G1731, C1836, C2232, and C2301. Among them, A705 may be
the 3’ end of band 8 RNA. A1014 matches well with the 3’ end of band 7 RNA. G1620
can be the 3’ end of band 6 RNA. G1731 and C1836 could be some of the 3’ ends of
band 9 RNAs. C2232 or C2301 may be the 3’ end of band 11 RNA. Primer extension of
23S rRNA revealed several 5’ ends around residue C1942. This position is the major
endonucleolytic cleavage site of 23S rRNA during degradation under starvation and
quality control conditions (Basturea et al., 2011). The multiple 5’ ends of the downstream
RNA products suggest that multiple endonucleolytic cleavages occur in this region,
89
A. wild-type (wt)
B. rnb rnr pnpts
C.
Figure 16. Northern blot analysis of 23S rRNA fragments. Total cellular RNA was
processed as in Figure 14 and analyzed with 23S rRNA– specific probes as indicated.
90
Numbers are assigned to each fragment. A. 23S rRNA in wild-type (wt) cells. B. 23S
rRNA in mutant rnb rnr pnpts cells. C. Diagrams show the location of the probes and the
main endonucleolytic cleavage sites (arrows). The weight of the lines indicates the
amount of the fragments. endo: endonucleolytic cleavage
which would also generate multiple 3’ ends for the upstream products including the band
9 RNAs. Primer extension also revealed multiple 5’ ends from C2096 to U2109
corresponding to the 5’ ends of band 11 RNA, and 5’ ends at U2585, U2586, and C2591
corresponding to the 5’ ends of band 12 and/or 13 products. The results are summarized
in a detailed diagram in Figure 16.
3.4.5 Pre-existing rRNA is degraded under oxidative stress
It has been reported that rRNA in pre-existing ribosome was degraded under
starvation, while newly synthesized rRNA was the substrate of the quality control. To
investigate the substrate of degradation pathway under oxidative stress, cells were grown
with (+) or without (-) rifampicin and then collected to analysis the degradation.
Fragments were accumulated in wild type (wt) cells at 180 min in the absence of H2O2,
but the accumulation was increased under H2O2 challenge and dose-dependent on the
H2O2 (Fig. 17A, left panel). The 16S rRNA was shorter than the full-length one at 180
min with 5 mM H2O2, indicating that the pre-existing 16S rRNA was partly degraded.
The upstream fragment 1 and downstream fragments 4 and 5 were detected by the same
probes used in Figure 16 (Fig. 17A). A short band was detected by the probe 16S710729, but it was not detected by the probe 16S15-36, suggesting that it is produced by an
endo-nucleolytic cleavage. The 5’ half products of this endo-nucleolytic cleavage may be
91
A. wild-type (wt)
B. rnr pnpts
C. rnb rnr pnpts
Figure 17. Northern blot analysis of 16S rRNA in the wild type (wt) and mutant cells
with H2O2 and rifampicin treatments. The cells were grown at 30 °C until OD550 ≈ 0.5,
and then shifted to 42 °C. 200 µg/ml rifampicin and H2O2 of indicated concentrations
were added into the culture. Total cellular RNA was extracted as in Figure 14 and the
Northern blot analysis of 16S rRNA was performed as in Figure 15 with specific probes
92
indicated in each panel. A. 16S rRNA in wide type (wt) cells. B. 16S rRNA in mutant rnr
pnpts cells. C. 16S rRNA in mutant rnb rnr pnpts cells.
the short bands detected only by the probe 16S15-36, and they are shortened by exonucleolytic activities.
In the absence of RNase R and PNPase, the 16S rRNA fragments were
accumulated more than that in wt cells, and were dose-dependent on the H2O2 (Fig. 17B,
left panel). Comparing with the results of no rifampicin treatment in Figure 15, bands 1,
4, and 5 were also detected by the same probes used in Figure 15 (Fig. 17B). Moreover,
another upstream long band, close to full length 16S rRNA, was detected at 180 min with
1 mM and 5 mM H2O2. It is probably trimmed into band 1. Two upstream short bands
were detected only by probe 16S710-729, indicating they were produced by the
endonucleolytic cleavage (Fig. 17B). The accumulated fragments were decreased when
RNase II is further deleted (Fig. 17C, left panel). Bands 1, 4, and 5 were detected by the
same probes (Fig. 17C). The short bands detected by 16S710-729 in rnr pnpts were not
detected by 16S838-857 in rnb rnr pnpts. Although RNase II is known to cleave the 3’
end of RNA, which helps other exoribonucleases to bind the RNA fragments and
degrade, the role RNase II plays in this situation is not clear.
Upon rifampicin treatment, the major upstream and downstream bands of 16S
rRNA were detected in the wild type and two mutants, and they were dose-dependent on
the H2O2. Therefore, these bands were produced from the pre-existing rRNA under
oxidative stress. This degradation process under oxidative stress is similar with starvation
condition, but not with quality control.
93
3.4.6 RNase PH is not required for the initiation of 16S rRNA degradation under
oxidative stress
RNase PH removes the 3’ end of 16S rRNA and initiates the rRNA degradation
under starvation. In the absence of RNase PH, no degradation of 16S rRNA occurs under
starvation (Basturea et al., 2011). To examine if RNase PH also plays a role in 16S rRNA
degradation under oxidative stress, an E. coli mutant lacking PNPase, RNase II, RNase R
and RNase PH (rnb rnr rph pnp) was compared with the rnb rnr pnp mutant for rRNA
degradation analysis (Fig. 18A, 14C, and 15C). Importantly, the additional RNase PH
deficiency did not block H2O2-induced degradation. Instead, many more rRNA fragments
accumulated in the quadruple mutant than in the triple mutant upon exposure to H2O2
(Fig. 18A, left panel, and 14C). Northern blotting analysis further confirmed that the
quadruple mutant contains the same 16S rRNA fragments as the triple mutant (Fig. 18A
and 15C). Interestingly, bands 1, 4 and 5 RNAs increased to higher levels in the mutant
lacking RNase PH when the cells were treated with 5 mM H2O2 for 180 min, suggesting
a role for RNase PH in efficient degradation of these RNA fragments (Fig. 18A).
Altogether, unlike under starvation conditions, RNase PH does not remove the 3’ end of
16S rRNA to initiate the degradation under oxidative stress (Fig. 18A and 15C).
However, RNase PH may participate in the removal of certain degradation intermediates
of 16S rRNA under oxidative stress. The rRNA fragments accumulated in the mutant are
from pre-existing rRNA, since the same upstream and downstream bands are detected
when the cells were treated with rifampicin (Fig. 18B)
94
A.
B.
1448-1467
1520-1539
838-857
893-912
951-970
710-729
15-36
C.
1 ~ 900 nt
4 ~ 650 nt
endo
5
Figure 18. Northern blot analysis of 16S rRNA in the mutant rnb rnr rph pnpts. Total
cellular RNA was extracted as in Figure 15 and the Northern blot analysis of 16S rRNA
was performed as in Figure 16 with specific probes indicated in each panel. A. 16S rRNA
fragments accumulated at different time courses with various concentrations of H2O2 and
the related Northern blot analysis. B. 16S rRNA fragments accumulated at different time
courses with 200 µg/ml rifampicin and various concentrations of H2O2 and the related
Northern blot analysis. C. Diagram shows the location of the probes and the main
95
endonucleolytic cleavage sites (arrows). The weight of the lines indicates the amount of
the fragments. endo: endonucleolytic cleavage
3.4.7 RNase E and RNase G are responsible for the major endonucleolytic cleavage
of 23S rRNA under oxidative stress
To investigate the enzyme(s) responsible for the endonucleolytic cleavage of
rRNA, mutants lacking single endoribonuclease or double endoribonucleases were used
to analyze the 16S rRNA degradation. RNase I and several exoribonucleases are deleted
in the control strain, such as RNase II, RNase T, RNase BN, and RNase D. In the mutant
strains, PNPase and either one or two additional endoribonucleases, such as RNase E,
RNase G, RNase P, and/or RNase III, are missing. RNase E and RNase G are responsible
for the 5’ maturation of 16S rRNA. RNase P generates the mature 5’ end of all tRNAs.
RNase III cleaves the primary transcript to separate the individual rRNAs. The cells were
grown in the same conditions as above for mutant strains lacking exoribonucleases.
Probes of 16S and 23S rRNA specific for the endo-cleavage sites were used in Northern
blot.
To display the same bands, the RNA sample from the mutant rnb rnr pnpts
collected at 180 min with 5 mM H2O2 of Figure 15 was used, which is labeled as c in
Figure 19. The probe 16S951-970 detected two bands in the control strain which are the
same size with the bands of lane c, indicating they are the same products, bands 4 and 5
(Fig. 19A). In the mutant ams, only band 4 was detected. In the cafA, both bands 4 and 5
were detected, indicating RNase G did not play role in the endo-cleavage of band 5 (Fig.
19B). In the ams cafA, ams rnc, and ams rnpA49, only band 4 was detected, suggesting
96
that RNase E may be responsible for the endo-cleavage which produced band 5 (Fig. 19A
and 20A). However, in the rnpA49 rnc, band 5 was not detected, although RNase E was
present (Fig. 20B). In the rnpA49, both bands 4 and 5 were detected, suggesting that
RNase P was not responsible for the endo-cleavage of band 5 (Fig. 20B). Therefore, none
of these endoribonucleases, RNase E, RNase G, RNase P, are responsible for the endocleavage of band 5 in 16S rRNA.
A. the control strain and the mutant ams cafA
B. ams and cafA
Figure 19. Northern blot analysis of 16S rRNA in the control strain and mutants
lacking endoribonucleases RNase E and/or RNase G. Total cellular RNA was
analyzed as in Figure 15 with the probe 16S951-970. A. 16S rRNA in the control strain
(CA265 rna rnb rnt rbn rnd) lacking RNase I and exoribonucleases RNase II, RNase T,
RNase BN, and RNase D, and the mutant ams cafA additionally lacking PNPase, RNase
97
E, and RNase G. B. 16S rRNA in the ams mutant additionally lacking PNPase and
RNase E, and cafA mutant additionally lacking PNPase and RNase G. c: RNA extracted
from the mutant CA244 rna rnb rnr pnpts at 180 min with 5 mM H2O2 and used in
Figures 15.
A. ams rnc and ams rnpA49
B. rnpA49 and rnpA49 rnc
Figure 20. Northern blot analysis of 16S rRNA fragments in mutants lacking
PNPase and endoribonucleases. Total cellular RNA was analyzed as in Figure 15 with
the probe 16S951-970. A. 16S rRNA in the mutant ams rnc additionally lacking PNPase,
RNase E, and RNase III, and mutant ams rnpA49 additionally lacking PNPase, RNase E
and RNase P. B. 16S rRNA in the mutant rnpA49 additionally lacking PNPase and
RNase P, and mutant rnpA49 rnc additionally lacking PNPase, RNase P, and RNase III.
c: RNA extracted from the mutant CA244 rna rnb rnr pnpts at 180 min with 5 mM H2O2
and used in Figures 15.
98
Northern analysis was also performed on 23S rRNA of these mutants. The probe
23S1996-2015 detected band 10, in the control, ams, cafA strains, and also the lane c,
indicating the endo-cleavage responsible for the production of band 10 occurred in
mutants ams and cafA (Fig. 21). Band 9 in ams cafA was detected by the same probe (Fig.
21A). Both band 9 and band 10 contain the 3’ end of 23S rRNA, but the 5’ end of band 9
is more upstream than band 10, which suggests that band 10 is produced from band 9 by
endo-cleavage. Since band 9 is not detected in ams and cafA single mutant, either of the
absent enzymes, RNase E and RNase G plays the role in endo-cleavage to produce band
10. All other mutants, rnpA49, rnpA49 rnc, ams rnc, and ams rnpA49, detected the same
band 10, indicating that RNase P and RNase III are not responsible for the endo-cleavage
which produced the fragment 10 of 23S rRNA (Fig. 22).
3.4.8 Discussion
In this study, we have shown that oxidative stress induces rRNA degradation,
consistent with the notion that the highly damaged rRNA must be eliminated.
Degradation under oxidative stress employs similar endonucleolytic activities and the
processive exoribonucleases that are known to cleave rRNA and clean up the resulting
fragments under starvation and quality control conditions. However, oxidative stress
induced degradation differs from the other two processes in the following aspects. First,
oxidative stress induced rRNA degradation differs from rRNA degradation under
starvation in the role of RNase PH and RNase II in these two processes. RNase PH
initiates rRNA degradation by shortening the 3’ end of 16S rRNA in the intact ribosomes
99
A. the control strain and the mutant ams cafA
B. ams and cafA
Figure 21. Northern blot analysis of 23S rRNA fragments in mutants lacking
endoribonucleases RNase E and/or RNase G. Total cellular RNA was analyzed as in
Figure 16 with the probe 23S1996-2015. A. 23S rRNA in the control strain and the
mutant ams cafA additionally lacking PNPase, RNase E, and RNase G. B. 23S rRNA in
the ams mutant additionally lacking PNPase and RNase E, and mutant cafA additionally
lacking PNPase and RNase G. c: RNA extracted from the mutant CA244 rna rnb rnr
pnpts at 180 min with 5 mM H2O2 and used in Figures 16.
100
A. ams rnc and ams rnpA49
B. rnpA49 and rnpA49 rnc
Figure 22. Northern blot analysis of 23S rRNA fragments in mutants lacking
endoribonucleases. Total cellular RNA was analyzed as in Figure 16 with the probe
23S1996-2015. A. 23S rRNA in the mutant ams rnc additionally lacking PNPase, RNase
E, and RNase III, and mutant ams rnpA49 additionally lacking PNPase, RNase E, and
RNase P. B. 23S rRNA in the mutant rnpA49 additionally lacking PNPase and RNase P,
and mutant rnpA49 rnc additionally lacking PNPase, RNase P, and RNase III. c: RNA
extracted from the mutant CA244 rna rnb rnr pnpts at 180 min with 5 mM H2O2 and used
in Figures 16.
101
(Basturea et al., 2011), whereas it only affects the degradation efficiency of certain rRNA
fragments under oxidative stress (data presented in this report). RNase II is one of the
two exoribonucleases that play important roles in degrading the rRNA fragments under
starvation (Basturea et al., 2011). However, the role of RNase II in oxidative stress
induced rRNA degradation is minimal. Second, rRNA degradation under oxidative stress
uses similar exoribonucleases as rRNA quality control. However, the two processes differ
from each other by degrading rRNA of different status. The difference implies that cells
use these two processes to target different rRNA populations. Quality control is limited to
degrading newly synthesized rRNA that is not yet assembled into mature ribosomes
(Basturea et al., 2011). Under oxidative stress, rRNA in damaged ribosomes may be a
major target for degradation (data presented in this report).
Under oxidative stress, both 16S and 23S rRNAs are degraded by multiple endoand exonucleolytic activities. The cleavage around 919 nt of 16S rRNA may be a very
early step in the degradation of this RNA. Similarly, the cleavage around 1942 nt of 23S
rRNA may initiate the decay of this RNA. However, in the ribosome, whether one rRNA
is cleaved before the other is unknown. It was reported that rRNA degradation under
starvation or quality control conditions also involves similar cleavages on the two longer
rRNAs at the early stage (Kaplan and Apirion, 1975; Basturea et al., 2011). Therefore,
the three process may share the same endoribonuclese activity(ies) for these cleavages,
which remain to be elucidated.
The RNA fragments generated by the initial cleavages are either degraded by
secondary
endonucleolytic
activities
into
102
shorter
fragments
or
digested
by
exoribonucleases in the 3’->5’ direction. The fragments of 23S rRNA appears to be
subjected to more endonucleolytic cleavages than those of 16S rRNA, possibly due to the
fact that 23S rRNA, 2904 nt, is much longer than 16S rRNA, 1542 nt. Similar to the
quality control pathway, RNase R and PNPase play major roles in the degradation of
rRNA fragments by 3’->5’ digestion, and RNase II may also help degrade rRNA
fragments under oxidative stress (Basturea et al., 2011).
Our work also revealed an interesting role for RNase PH in the degradation of
certain 16S RNA intermediates under oxidative stress. One intermediate contains the 3’
end of 16S rRNA. Two other intermediates contain 3’ termini located internal of 16S
rRNA, representing newly observed substrates for RNase PH.
Under starvation
condition, RNase PH plays a more important role in rRNA degradation because it
initiates ribosome breakdown by shortening the 3’ end of 16S rRNA. In this regard, the
surveillance mechanism of rRNA under oxidative stress is closer to rRNA quality
control, and differs significantly from that under starvation.
The two fragments of 16S rRNA generated by the initial cleavage seem to be
degraded at a similar rate in the wild type cell, where formation of both depends on the
exoribonucleases and some unknown endonucleolytic activity(ies). In contrast, the initial
fragments of 23S rRNA are degraded at different rates. The upstream fragments, bands 69, accumulate in both mutant and wild type. The downstream fragments, bands 10-13,
accumulate only in the absence of RNase II, RNase R, and PNPase, and they are absent
in wild type. This pattern suggests that the downstream fragments of 23S rRNA are more
efficiently removed than the upstream fragments, probably due to the involvement of
103
multiple endonucleolytic cleavages in degrading the downstream products. The resulting
shorter products can be more rapidly degraded than the upstream products. The exact
features for the downstream half of 23S rRNA to receive more endonucleolytic cleavages
are unknown, but it may be related to the structure of this RNA.
It should be noted that the degradation intermediates identified under oxidative
stress are also found under conditions without oxidant-treatment. However, in most cases,
the amount of the intermediates is elevated after H2O2 treatment. This provides further
support to the idea that rRNA degradation under oxidative stress and quality control may
share the same pathway, although unique activities may be used in one of the processes.
It also suggests that rRNA degradation under oxidative stress demands high degradation
activities, causing accumulation of more intermediates than non-oxidative conditions. It
is also likely that some of the degradation activities are regulated by oxidative stress,
which deserves further study in the future.
104
4. Conclusions
Under oxidative stress conditions, such as upon treatment with hydrogen
peroxide, E. coli RNA is highly oxidized. H2O2-induced 8-oxo-G is mainly present in
ribosomal RNA. In fact, rRNA appears to contain higher normalized levels of 8-oxo-G
both in vivo and in vitro when treated with H2O2, likely caused by its high ability to bind
redox metals and/or by its high order structures. In contrast, ribosomal RNA contains
lower levels of 8-oxo-G than non-ribosomal RNA under normal conditions. Furthermore,
the levels of H2O2-induced 8-oxo-G are quickly reduced in ribosomes after removal of
the oxidant. These suggested that 8-oxo-G containing rRNA is selectively eliminated in
ribosomes. Importantly, the selective reduction of ribosomal 8-oxo-G depends on the
activities of PNPase and RNase R that are known to degrade structured RNA, suggesting
that RNA degradation is a major mechanism for controlling oxidized rRNA. In consistent
with this notion, rRNA undergoes degradation when E. coli cultures are exposed to H2O2.
The level of degradation of 16S and 23S rRNA depends on the dose of H2O2. H2O2induced rRNA degradation appears to be initiated by endonucleolytic cleavages. The
resulting rRNA fragments are digested exoribonucleases, mainly RNase R and PNPase.
The H2O2-induced rRNA degradation pathwayis similar to, and yet different from, those
for bulk rRNA decay under starvation and rRNA quality control (Basturea et al., 2011).
This work demonstrates that rRNA oxidative damage is an important problem, and
109
specific RNA degradation mechanisms are involved in the control of oxidized rRNA in
E. coli.
109
Appendices
I. Identification of RNA-related proteins that protect cells under oxidative stress
Our results presented in this dissertation as well as in published work suggest that
RNA oxidative damage is deleterious to cells, and cells may have developed additional
mechanisms to control damaged RNA. Based on these observations, we have searched
for other RNA-related activities in E. coli that may play protective roles under oxidative
stress. Our rational is that if a protein is involved in RNA metabolism, and if it is
important for cell survival under oxidative stress, the protein is likely involved in control
of oxidizedRNA.
We have identified such proteins by testing H2O2-sensitivity of mutants that each
lacks one of the protein candidates. Those mutants that were hypersensitive to H2O2
would suggest that the missing proteins are important for cell viability. The candidate
proteins were chosen based on either a proteomics screening for proteins that may bind
oxidized RNA with specificity, or a bioinformatics search for proteins with
known/predicted role in RNA metabolism. Results of cell viability analysis are
summarized below.
111
I.1 Selection of candidate proteins and construction of E. coli mutants lacking the
proteins
Our laboratory has performed a proteomic analysis to identify proteins that bind
oxidized RNA with higher affinity than binding normal control RNA (Zhe Jiang and
Zhongwei Li, unpublished data). Briefly, a 50-mer synthetic RNA was oxidized using
H2O2 (Wu et al., 2009). Both control RNA and oxidized RNA were covalently linked to
agarose beads. RNA-beads preparations were incubated with E. coli S100 cell extracts.
Proteins bound to the RNA were eluted and separated on a SDS-PAGE gel. The protein
bands that demonstrated increased amounts from oxidized RNA beads comparing to
those from control RNA beads were excised from the gel for MS-Spec analysis of protein
identity. Based on the MS-Spec data, we have selected 74 of the proteins to further
analyze their roles in cell protection under oxidative stress.
Sixty-four additional proteins were selected based on their known or predicted
functions in RNA metabolism. However, those proteins that are known to be essential for
normal cell growth were excluded in this study because H2O2-sensitivity data can not be
generated from the mutants without these activities.
The selected proteins were classified into several groups according to their
functions. These groups include ribonucleases, RNA helicases, RNA binding proteins,
ribosomal proteins, RNA modification enzymes, proteins that are induced under
oxidative stress, DNA repair enzymes, proteins playing roles in the metabolism of
carbohydrates, proteins, and fatty acids. There are also proteins responsible for
112
transportation, secretary, folding, etc. In addition, the function of 22 candidates is
unknown.
E.coli mutants lacking one of the non-essential genes have been systematically
constructed using strain K-12 BW25113 (Beba et al., 2006) by replacing the open reading
frame of the genes with a kanamycin-resistance gene cassette.The wild type BW25113
strain and mutants lacking one of the above 138 proteins were used in the initial
screening. Corresponding mutants were also constructed in E. coli K-12 CA244
background by transferring the mutant alleles via P1 transduction.
I.2 H2O2-sensitivity of E.coli mutants lacking RNA-related proteins
We have studied H2O2-sensitivity of the mutants of both BW25113 and CA244
backgrounds. Exponentially growing cultures were serially diluted and a small amount of
the diluted cultures were inoculated as spots on the surface of YT-agar plates with or
without H2O2. After incubation, growth was recorded. Usually, cells grow well in the
absence of H2O2, and separate colonies form only at the spots of the most diluted
cultures. In the presence of H2O2 at certain concentration, the growth was much inhibited,
resulting in the growth of fewer spots of less-diluted cultures depending on the sensitivity
of particular mutant strain.
The growth of mutant strains was compared with that of wild type (wt) (Fig. 23).
The results are summarized in Tables 3. Depending on the level of H2O2 sensitivity, the
mutations may cause severe, mild or no effect on cell growth under oxidative stress. The
113
growth defect of several mutants upon H2O2 treatment suggested that the absent proteins
protect the cell from oxidative stress.
A.
wt
fdrA
deoBB
ppK
-H2O2
+H2O2
B.
wt
fimD
degP
rhlB
truC
+H2O2
-H2O2
C.
wt
ycgG
truB
srmB
+H2O2
-H2O2
Figures 23. Hypersensitivity of E.coli cells to H2O2 treatment. Overnight culture of
E.coli wild type (wt) and mutant cells were diluted with fresh YT from and grown to
OD550 ≈ 0.5. Then the cells were serially diluted for 7 times with YT. The serially diluted
cultures were spotted onto the YT agar plate (+/- H2O2). A. wt and mutants fdrA, deoB,
and ppK, respectively were spotted on the same plate with H2O2 (+) or without (-). B. the
114
growth of wt and mutants fimD, degP, rhlB, and ruC with (+) or without (-) H2O2. C. the
growth of wt and mutants ycgG, truB, and srmB with (+) or without (-) H2O2.
Table 3: Summary of identified proteins that protect cells against oxidative stress.
Source of
protein
candidates
Proteomics
analysis
Functional
prediction
number of
proteins
74
64
Response of
mutants to
H2O2
Strain background
BW25113
only
CA244
only
Both
Hypersensitive
22
4
19
Not
hypersensitive
4
22
29
Hypersensitive
22
3
23
Not
hypersensitive
3
22
16
The results suggest that proteins of certain categories tend to have more profound
protective effect. Ten of thirteen ribonucleases, three of five RNA helicases, four proteins
binding with RNA, two ribosomal proteins, sixteen proteins modifying RNA, etc, showed
protection to the cell under oxidative stress. Overall, among the 74 proteins selected from
the proteomic analysis, 19 of them had protective effect in the BW25113 and CA244
backgrounds. Twenty-two were hypersensitive to H2O2 treatment in BW25113
background, and four were in CA244 background. Among the 64 proteins selected from
functional prediction, 23 of them were hypersensitive to H2O2 treatment in both
115
backgrounds. Twenty-two of them were hypersensitive to H2O2 challenge in BW25113,
and 3 were in CA244 background.
I.3 The protective roles of RNases, RNA helicases, or poly(A) polymerase under
oxidative stress
PNPase and RNA helicase RhlB are components of the RNA degradosome and
they are thought to work together to degrade a structured RNA with high efficiency. It
has also been postulated that PAP helps RNase R to degrade a structural region of RNA
by adding a poly(A) tail to the 3’ end of RNA. A mutant lacking both RNase R and
PNPase is unviable (Cheng et al., 2008). Therefore, the deletion of PNPase and PAP
would affect both activities known to degrade structured RNA. Mutation of both RNase
R and RhlB would have similar effect on degradation of structured RNA. Interestingly,
the double mutant, pnp pap and rnr rhlB, grew well under normal conditions, suggesting
that the remaining RNA degradation activities are sufficient for normal RNA metabolism.
We have shown that cells lacking either PNPase, RNase R or RNase II are
hypersensitive to H2O2 challenge (Wu et al., 2009; unpublished observations), suggesting
that both activities are important for controlling oxidized RNA. This prompted us to
analyze the sensitivity of the double mutant, pnp pap and rnr rhlB, under oxidative stress
conditions.
In the absence of PNPase and PAP, cells grew extremely poorly under oxidative
stress (Fig. 24A). The pnp pap mutant was much more sensitive than pnp to H2O2
treatment. A cell lacking PAP grew as well as wild type in the presence of H2O2.
116
Similarly, rnr rhlB cells were much more sensitive to H2O2 than the rnr or rhlB mutants
(Fig. 24B). These findings strongly suggest that blockage in structured RNA degradation
affects the ability to remove oxidized RNA, and the accumulation of damaged RNA
under oxidative stress may eventually cause cell death.
A.
wt
pap
pnp
pnp pap
+ H2O2
- H2O2
(A by Jinhua Wu)
B.
wt
rhlB
rnb
rnb rhlB
pnp
pnp rhlB
rnr
rnr rhlB
-H2O2
+H2O2
Figure 24. More sensitivity of combined mutation of RNase, RNA helicase RhlB, or
poly(A) polymerase under oxidative stress. This was done in the same way as in Figure
25. A. wt (wild-type) and mutants pnp, pap, and pnp pap lacking PNPase, PAP, or
PNPase and PAP, respectively. B. wt (wild-type) and mutants rhlB, rnb, rnb rhlB, pnp,
pnp rhlB, rnr, and rnr rhlB.
117
Moreover, we have also studied the H2O2 sensitivity of mutants lacking other
DEAD-box RNA helicases, RhlE, DbpA, SrmB, and DeaD, alone or in combination with
deficiencies of RNase R, PNPase, or RNase II (Fig. 25). It is known that these five RNA
helicases regulate RNA structure by unwinding RNA duplex, and are important for RNA
metabolism at low temperature (Jagessar and Jain, 2010). RhlE is a ribosome assembly
factor and plays a role in the rRNA maturation (Jain, 2008). DbpA is an ATP-dependent
3'-5' RNA helicase and interacts with 23S rRNA. Both SrmB and DeaD are also ribosome
assembly factors and work in ribosome biogenesis (Jagessar and Jain, 2010). DeaD also
facilitates translation of mRNAs with 5' secondary structures. SrmB is required for
efficient ribosomal 50S subunit assembly and stabilizes exposed mRNA, possibly by
protecting or destabilizing secondary structure. In the absence of SrmB or DeaD, rRNA
processing and ribosomal maturation are defective at low temperature and 37 °C
(Jagessar and Jain, 2010). When DbpA was combined with RNase II, PNPase, or RNase
R, the mutants were not more hypersensitive than the corresponding single mutant,
except rnr dbpA which grew worse than rnr, dbpA, and wt cells upon H2O2 challenge
(Fig. 25A). DeaD displayed a similar result with DbpA. Only rnr dead grew worse than
the corresponding single mutant (Fig. 25B). rnb srmB had the most serious defective
growth among all the single mutants and double mutants under oxidative stress (Fig.
25C). The deletion of RhlE did not affect the cell growth under oxidative stress, nor did
the combined absence with RNase II, PNPase, or RNase R (data not show).
118
A.
wt
dbpA
rnb
rnb dbpA
pnp
pnp dbpA
rnr
rnr dbpA
-H2O2
+H2O2
B.
wt
deaD
rnb
rnb deaD
pnp
pnp deaD
rnr
rnr deaD
-H2O2
+H2O2
C.
wt
srmB
rnb
rnb srmB
pnp
pnp srmB
rnr
rnr srmB
-H2O2
+H2O2
Figure 25. More sensitivity of combined mutation of RNase and RNA helicases
under oxidative stress. This was done in the same way as in Figure 24. A. wt (wild-type)
and mutants in the absent of RNA helicase DbpA, and one of the three exoribonucleases,
RNase II, PNPase, or RNase R. B. wt and mutants in the absent of RNA helicase DeaD,
119
and one of the three exoribonucleases, RNase II, PNPase, or RNase R. C. wt and mutants
in the absent of RNA helicase SrmB, and one of the three exoribonucleases, RNase II,
PNPase, or RNase R.
Altogether, the results suggest that some of the RNA helicases facilitate the
exoribonucleases in protecting cells under oxidative stress, probably by helping the decay
of oxidized RNA.
I.4 Discussion
This work determined enzymes protecting the cell from oxidative stress. It is
interesting that many proteins are important for the cell viability under oxidative stress. In
the absence of PPK, a component of RNA degradosome, the cell did not grow as well as
wild type cells upon H2O2 treatment. It is possible that the damaged RNA was removed
more slowly and accumulated in the cell, since the activity of the degradosome was
reduced due to the absence of PPK, which may contribute to the growth problem.
Moreover, PPK was selected by the proteomic analysis and bound with oxidized RNA
specifically, indicating that it may identify the oxidized RNA and signal the degradation
of damaged RNA. In the ppK mutant, both identification and degradation of the damaged
RNAs were affected by PPK mutation, and together play roles in the cell viability under
oxidative stress. However, the degradosome is not the only quality control pathway in the
cell, nor PPK is the the only protein specifically bound with oxidized RNA. Based on the
data, it is difficult to find out the connection between the absence of PPK and the growth
defect under oxidative stress. It is also hard to discover the relation in other mutants and
more works need to be done in the future.
120
II. Supplementary Data
Oligos and primers used in this work:
16S15-36: 5’-GCG TTC AAT CTG AGC CAT GAT C-3’
16S57-76: 5’-CCT GTT ACC GTT CGA CTT GC-3’
16S308-328: 5’-GTC TCA GTT CCA GTG TGG CT-3’
16S330-349: 5’-TCC CGT AGG AGT CTG GAC CG-3’
16S440-459: 5’-TAC TCC CTT CCT CCC CGC TG-3’
16S504-523: 5’-TGC TGG CAC GGA GTT AGC CG-3’
16S533-552: 5’-AAC GCT TGC ACC CTC CGT AT-3’
16S561-580: 5’-GTG CGC TTT ACG CCC AGT AA-3’
16S584-603: 5’-ATC TGA CTT AAC AAA CCG CC-3’
16S611-630: 5’-TTC CCA GGT TGA GCC CGG GG-3’
16S632-651: 5’-GCT TGC CAG TAT CAG ATG CA-3’
16S710-729: 5’-TTC GCC ACC GGT ATT CCT CC-3’
16S795-814: 5’-TAC GGC GTG GAC TAC CAG GG-3’
16S838-857: 5’-GGA AGC CAC GCC TCA AGG GC-3’
16S893-912: 5’-GAG TTT TAA CCT TGC GGC CG-3’
121
16S922-941: 5’-CGC TTG TGC GGG CCC CCG TC-3’
16S939-958: 5’- TAA ACC ACA TGC TCC ACC GC-3’
16S951-970: 5’-GTT GCA TCG AAT TAA ACC AC-3’
16S980-999: 5’-GGA TGT CAA GAC CAG GTA AG-3’
16S1010-1029: 5’-AGG CAC CAA TCC ATC TCT GG-3’
16S1037-1057: 5’-AGG GTT GCG CTC GTT GCG GG-3’
16S1065-1084: 5’-CAT TTC ACA ACA CGA GCT GA-3’
16S1269-1288: 5’-TTA TGA GGT CCG CTT GCT CT-3’
16S1382-1401: 5’-CGG TGT GTA CAA GGC CCG GG-3’
16S1411-1430: 5’-TTT GCA ACC CAC TCC CAT GG-3’
16S1448-1467: 5’-GGT AAG CGC CCT CCC GAA GG-3’
16S1477-1496: 5’-GAC TTC ACC CCA GTC ATG AA-3’
16S1520-1539: 5’-GGA GGT GAT CCA ACC GCA GG-3’
23S7-26: 5’-CAC CGT GTA CGC TTA GTC GC-3’
23S182-201: 5’-GAT GTT TCA GTT CCC CCG GT-3’
23S1689-1710: 5’-CAG CGT GCC TTC TCC CGA AG-3’
23S1799-1818: 5’-ACG TCC ACT TTC GTG TTT GC-3’
122
23S1855-1874: 5’-GCT TGC GCT AAC CCC ATC AA-3’
23S1880-1899: 5’-TAC CGG GGC TTC GAT CAA GA-3’
23S1919-1938: 5’-TTT CGC TAC CTT AGG ACC GT-3’
23S1942-1961: 5’-GTC GGA ACT TAC CCG ACA AG-3’
23S1974-1993: 5’-ACA GCC TGG CCA TCA TTA CG-3’
23S1996-2015: 5’-TTT CAC TGA GTC TCG GGT GG-3’
23S2019-2038: 5’-CTG CAT CTT CAC AGC GAG TT-3’
23S2061-2080: 5’-TAG TAA AGG TTC ACG GGG TC-3’
23S2131-2150: 5’-GAC TGG CGT CCA CAC TTC AA-3’
23S2608-2627: 5’-CGC CCA CGG CAG ATA GGG AC -3’
23S2885-2904: 5’-AAG GTT AAG CCT CAC GGT TC-3’
16S-sense-440-459: 5’-CAG CGG GGA GGA AGG GAG TA-3’
16S- sense-503-522: 5’-CCG GCT AAC TCC GTG CCA GC-3’
16S- sense-771-791: 5’-GTG GGG AGC AAA CAG GAT TA-3’
16S- sense-838-857: 5’-GCC CTT GAG GCG TGG CTT CC-3’
16S- sense-922-941: 5’-GAC GGG GGC CCG CAC AAG CG-3’
16S- sense-1059-1079: 5’-CTG TCG TCA GCT CGT GTT GT-3’
123
16S- sense-1127-1146: 5’-GCC AGC GGT CCG GCC GGG AA-3’
16S- sense-1342-1361: 5’-CGC TAG TAA TCG TGG ATC AG-3’
16S- sense-1382-1401: 5’-CCC GGG CCT TGT ACA CAC CG-3’
16S- sense-1448-1467: 5’-CCT TCG GGA GGG CGC TTA CC-3’
23S- sense-433-452: 5’-CTC CTG ACT GAC CGA TAG TG-3’
23S- sense-509-528: 5’-CCT GAA ACC GTG TAC GTA CA-3’
23S- sense-610-629: 5’-CCG AAT AGG GGA GCC GAA GG-3’
23S- sense-700-719: 5’-GGT TGA AGG TTG GGT AAC AC-3’
23S- sense-1200-1219: 5’-CTG TAA GCC TGT GAA GGT GT-3’
23S- sense-1350-1369: 5’-GGT TGA AGG TTG GGT AAC AC-3’
23S- sense-1652-1671: 5’-AGA ACT CGG GTG AAG GAA CT-3’
23S- sense-1752-1771: 5’-CGA AGA TAC CAG CTG GCT GC-3’
23S- sense-2131-2150: 5’-TTG AAG TGT GGA CGC CAG TC-3’
Primer of the Linker: 5’- TCT GCG TCG CTA CAA TGG AT- 3’
Linker: 5’- ATC CAT TGT AGC GAC GCA GA- 3’
Sense: same sequence with rRNA
124
5. References
Aas, P.A., Otterlei, M., Falnes, P.O., Vagbo, C.B., Skorpen, F., Akbari, M., Sundheim,
O., Bjoras, M., Slupphaug, G., Seeberg, E., and Krokan, H.E. (2003) Human and
bacterial oxidative demethylases repair alkylation damage in both RNA and DNA.
Nature 421:859-863.
Abe, T., Tohgi, H., Isobe, C., Murata, T., and Sato, C. (2002). Remarkable increase in the
concentration of 8-hydroxyguanosine in cerebrospinal fluid from patients with
Alzheimer’s disease. J. Neurosci. Res. 70:447 – 450.
Abelson, H. T., Johnson, L. F. S. Penman, S., and Green, H. (1974) Changes in RNA in
relation to growth of the fibroblast: II. The lifetime of mRNA, rRNA, and tRNA
in resting and growing cells. Cell 1:161-165.
Abernathy, J. W., Xu, D. H., Li, P., Klesius, P., Kucuktas, H., and Liu, Z. (2009)
Transcriptomic profiling of Ichthyophthirius multifiliis reveals polyadenylation of
the large subunit ribosomal RNA. Comp. Biochem. Physiol. Part D Genomics
Proteomics. 4:179-186.
Alexandrov, A., Chernyakov, I., Gu, W., Hiley, S. L., Hughes, T. R., Grayhack, E. J.,and
Phizicky, E. M. (2006) Rapid tRNA decay can result from lack of nonessential
modifications. Mol. Cell. 21:87-96.
125
Allmang, C., Mitchell, P., Petfalski, E., and Tollervey, D. (2000) Degradation of
ribosomal RNA precursors by the exosome. Nucleic Acids Res. 28:1684-1691.
Ames, B.N., and Gold, L.S. (1991) Endogenous mutagens and the causes of aging and
cancer. Mutat. Res. 250:3-16.
Baba, T., Ara, T., Hasegawa, M., Takai, Y., Okumura, Y., Baba, M., Datsenko, K.A.,
Tomita, M., Wanner, B.L., and Mori, H. (2006) Construction of Escherichia coli
K-12 in-frame, single-gene knockout mutants: the Keio collection. Mol. Syst.
Biol. 2:2006.0008.
Baker, K. E., and Parker, R. (2004) Nonsense-mediated mRNA decay: terminating
erroneous gene expression. Curr. Opin. Cell Biol. 16:293-299.
Bass, B. L. (2000) Double-stranded RNA as a template for gene silencing. Cell 101:235238.
Basturea, G. N., Zundel, M. A., and Deutscher, M. P. (2011) Degradation of ribosomal
RNA during starvation: comparison to quality control during steady-state growth
and a role for RNase PH. RNA 17:338-345.
Beelman, C. A., and Parker, R. (1995) Degradation of mRNA in eukaryotes. Cell 81:179183.
Bellacosa, A. and Moss, E.G. (2003) RNA repair: damage control. Curr. Biol. 13 R:482484.
126
Ben-Hamida, F., and Schlessinger, D. (1966) Synthesis and breakdown of ribonucleic
acid in Escherichia coli starving for nitrogen. Biochim. Biophys. Acta. 119:183191.
Bernstein, P., and Ross, J. (1989) Poly(A), poly(A) binding protein and the regulation of
mRNA stability. Trends Biochem. Sci. 14:373-377.
Bessarab, D. A., Kaberdin, V. R., Wei, C. L., Liou, G. G., and Lin-Chao, S. (1998) RNA
components of Escherichia coli degradosome: evidence for rRNA decay. Proc.
Natl. Acad. Sci. USA. 95:3157-3161.
Blum, E., Py, B., Carpousis, A. J., and Higgins, C. F. (1997) Polyphosphate kinase is a
component of the Escherichia coli RNA degradosome. Mol. Microbiol. 26:387398.
Borek, E., Baliga, B. S., Gehrke, C. W., Kuo, C. W., Belman, S., Troll, W., and Waalkes,
T. P. (1977) High turnover rate of transfer RNA in tumor tissue. Cancer Res.
37:3362-3366.
Broedbaek, K., Poulsen, H. E., Weimann, A., Kom, G. D., Schwedhelm, E., Nielsen, P.,
and Boger, R. H. (2009). Urinary excretion of biomarkers of oxidatively damaged
DNA and RNA in hereditary hemochromatosis. Free Radic. Biol. Med. 47:12301233.
Carpousis, A.J., Vanzo, N.F., and Raynal, L.C. (1999) mRNA degradation: a tale of poly
(A) and multiprotein machines. Trends Genet. 15:24-28.
127
Chang, Y., Kong, Q., Shan, X., Tian, G., Ilieva, H., Cleveland, D. W., Rothstein, J. D.,
Borchelt, D. R., Wong, P. C., and Lin, C. L. (2008) Messenger RNA oxidation
occurs early in disease pathogenesis and promotes motor neuron degeneration in
ALS. PLoS One. 3:2849-2867.
Che, Y., Wang, J.F., Shao, L., and Young, T. (2010). Oxidative damage to RNA but not
DNA in the hippocampus of patients with major mental illness. J. Psychiatry
Neurosci. 35:296-302.
Cheng, Z. F., and Deutscher, M. P. (2002) Purification and characterization of the
Escherichia coli exoribonuclease RNase R. Comparison with RNase II. J. Biol.
Chem. 277:21624-21629.
Cheng, Z. F., and Deutscher, M. P. (2003) Quality control of ribosomal RNA mediated
by polynucleotide phosphorylase and RNase R. Proc. Natl. Acad. Sci. USA.
100:6388-6393.
Cheng, Z.F., and Deutscher, M.P. (2005) An important role for RNase R in mRNA
decay. Mol. Cell. 17: 313-318.
Chernyakov, I., Baker, M. A, Grayhack, E. J, and Phizicky, E. M. (2008) Identification
and analysis of tRNAs that are degraded in Saccharomyces cerevisiae due to lack
of modifications. Methods Enzymol. 449:221-37.
Chernyakov, I., Whipple, J. M., Kotelawala, L., Grayhack, E. J., and Phizicky, E. M.
(2008) Degradation of several hypomodified mature tRNA species in
128
Saccharomyces cerevisiae is mediated by Met22 and the 5'-3' exonucleases Rat1
and Xrn1. Genes Dev. 22:1369-1380.
Cohen, S.N. (1995) Surprises at the 3′ end of prokaryotic RNA. Cell 80: 829-832.
Cole, S. E., and LaRiviere, F. J. (2008) Analysis of nonfunctional ribosomal RNA decay
in Saccharomyces cerevisiae. Methods Enzymol. 449:239-259.
Cole, S. E., LaRiviere, F. J, Merrikh, C. N, and Moore, M. J. (2009) A convergence of
rRNA and mRNA quality control pathways revealed by mechanistic analysis of
nonfunctional rRNA decay. Mol. Cell. 34:440-450.
Cooke, M. S., Evans, M. D., Dizdaroglu, M., and Lunec, J. (2003) Oxidative DNA
damage: mechanisms, mutation, and disease. FASEB J. 17:1195-1214.
Datsenko, K. A., and Wanner, B. L. (2000) One-step inactivation of chromosomal genes
in Escherichia coli K-12 using PCR products. Proc. Natl. Acad. Sci. 97:66406645.
Deana, A., Celesnik, H., and Belasco, J. G. (2008) The bacterial enzyme RppH triggers
messenger RNA degradation by 5' pyrophosphate removal. Nature 451:355-358.
de Souza-Pinto, N.C., and Bohr, V.A. (2002). The mitochondrial theory of aging:
involvement of mitochondrial DNA damage and repair. Int. Rev. Neurobiol.
53:519 – 534.
Deutscher, M. P. (2003) Degradation of stable RNA in bacteria. J. Biol. Chem.
278:45041-45044.
129
Deutscher, M. P. (2006) Degradation of RNA in bacteria: comparison of mRNA and
stable RNA. Nucleic Acids Res. 34:659-666.
Deutscher, M.P. and Hilderman, R.H. (1974). Isolation and partial characterization of
Escherichia coli mutants with low levels of transfer ribonucleic acid
nucleotidyltransferase. J. Bacteriol. 118: 621 – 627.
Ding, Q., Markesbery, W.R., Chen, Q., Li, F., and Keller, J.N. (2005) Ribosome
dysfunction is an early event in Alzheimer’s disease. J. Nuerosci. 25:9171-9175.
Doma, M. K. (2008) Methods to Study No‐Go mRNA Decay in Saccharomyces
cerevisiae. Methods Enzymol. 449: 3-21.
Dulebohn, D., Choy, J., Sundermeier, T., Okan, N., and Karzai, A. W. (2007) Transtranslation: the tmRNA-mediated surveillance mechanism for ribosome rescue,
directed protein degradation, and nonstop mRNA decay. Biochemistry 46:46814693.
ESCODD (European Standards Committee on Oxidative DNA Damage), Gedik, C.M.,
and Collins, A. (2005). Establishing the background level of base oxidation in
human lymphocyte DNA : results of an interlaboratory validation study. FASEB
J. 19:82 – 84.
Evans, M. D., Dizdaroglu, M., and Cooke, M. S. (2004) Oxidative DNA damage and
disease: induction, repair and significance. Mutat. Res. 567:1-61.
130
Fiala, E. S., Conaway, C. C., and Mathis, J. E. (1989) Oxidative DNA and RNA damage
in the livers of Sprague-Dawley rats treated with the hepatocarcinogen 2nitropropane. Cancer Res. 49:5518-5522.
Floyd, R. A., West, M. S., Eneff, K. L., and Schneider, J. E. (1989) Methylene blue plus
light mediates 8-hydroxyguanine formation in DNA. Arch. Biochem. Biophys.
273:106-111.
Gajewski, E., Rao, G., Nackerdien, Z., and Dizdaroglu, M. (1990). Modification of DNA
bases in mammalian chromatin by radiation- generated free radicals.
Biochemistry 29:7876 – 7882.
Ge, Z., Mehta, P., Richards, J., and Karzai, A. W. (2010) Non-stop mRNA decay initiates
at the ribosome. Mol Microbiol. 78:1159-1170.
Ghosh, S., and Deutscher, M. P. (1999) Oligoribonuclease is an essential component of
the mRNA decay pathway. Proc. Natl. Acad. Sci. 96: 4372-4377.
Gong, X., Tao, R., and Li, Z. (2006) Quantification of RNA damage by reverse
transcription polymerase chain reactions. Anal. Biochem. 357:58-67.
Görg, B., Qvartskhava, N., Bidmon, H.J., Palomero-Gallagher, N., Kircheis, G., Zilles,
K., and Häussinger, D. (2010). Oxidative stress markers in the Brain of patients
with cirrhosis and hepatic encephalopathy. Hepatology 52:256-265.
131
Haiser, H. J., Karginov, F. V., Hannon, G. J., and Elliot, M. A. (2008) Developmentally
regulated cleavage of tRNAs in the bacterium Streptomyces coelicolor. Nucleic
Acids Res. 36:732-741.
Hayakawa, H., Hofer, A., Thelander, L., Kitajima, S., Cai, Y., Oshiro, S., Yakushiji, H.,
Nakabeppu, Y., Kuwano, M., and Sekiguchi, M. (1999) Metabolic fate of
oxidized guanine ribonucleotides in mammalian cells. Biochemistry 38:36103614.
Hayakawa, H., Kuwano, M., and Sekiguchi, M. (2001) Specific binding of 8-oxoguaninecontaining RNA to polynucleotide phosphorylase protein. Biochemistry 40:99779982.
Hillebrand, J., Barbee, S. A., and Ramaswami, M. (2007) P-body components,
microRNA regulation, and synaptic plasticity. ScientificWorldJournal. 7:178-190.
Hofer, T., Badouard, C., Bajak, E., Ravanat, J.L., Mattsson, A. Cotgreave, I. (2005)
Hydrogen peroxide causes greater oxidation in cellular RNA than in DNA. Biol.
Chem. 386:333-337.
Hofer, T., Fontana, L., Anton, S.D., Weiss, E.P., Villareal, D., Malayappan, B., and
Leeuwenburgh, C. (2008a). Long-term effects of caloric restriction or exercise on
DNA and RNA oxidation levels in white blood cells and urine in humans.
Rejuvenation. Res. 11: 793-799.
Hofer, T., Marzetti, E., Xu, J., Seo, A.Y., Gulec, S., Knutson, M.D., Leeuwenburgh, C.,
and Dupont-Versteegden, E.E. (2008b). Increased iron content and RNA
132
oxidative damage in skeletal muscle with aging and disuse atrophy. Exp.
Gerontol. 43:563-570.
Hofer, T., Seo, A.Y., Prudencio, M., and Leeuwenburgh, C. (2006) A method to
determine RNA and DNA oxidation simultaneously by HPLC-ECD: greater RNA
than DNA oxidation in rat liver after doxorubicin administration. Biol Chem.
387:103-111.
Holley, C. L., and Topkara, V. K. (2011) An introduction to small non-coding RNAs:
miRNA and snoRNA. Cardiovasc. Drugs Ther. 25:151-159.
Honda, K., Smith, M.A., Zhu, X., Baus, D., Merrick, W.C., Tartakoff, A.M., Hattier, T.,
Harris, P.L., Siedlak, S.L., Fujioka, H., Liu, Q., Moreira, P.I., Miller, F.P.,
Nunomura, A., Shimohama, S., and Perry, G. (2005) Ribosomal RNA in
Alzheimer Disease Is Oxidized by Bound Redox-active Iron. J. Biol. Chem.
280:20978–20986.
Hsu, D., Shih, L. M., and Zee, Y. C. (1994) Degradation of rRNA in Salmonella strains: a
novel mechanism to regulate the concentrations of rRNA and ribosomes. J.
Bacteriol. 176:4761-4765.
Huang, H., Liao, J., and Cohen, S. N. (1998) Poly(A)- and poly(U)-specific RNA 3' tail
shortening by E. coli ribonuclease E. Nature. 391:99-102.
133
Ishibashi, T., Hayakawa, H., Ito, R., Miyazawa, M., Yamagata, Y., and Sekiguchi, M.
(2005). Mammalian enzymes for preventing transcriptional errors caused by
oxidative damage. Nucleic. Acids Res. 33:3779 – 3784.
Isken, O., and Maquat, L. E. (2007) Quality control of eukaryotic mRNA: safeguarding
cells from abnormal mRNA function. Genes Dev. 21:1833-1856.
Jacobson, A., and Gillespie, D. (1968) Metabolic events occurring during recovery from
prolonged glucose starvation in Escherichia coli. J. Bacteriol. 95:1030-1039.
Jagessar, K. L., and Jain, C. (2010) Functional and molecular analysis of Escherichia coli
strains lacking multiple DEAD-box helicases. RNA. 16:1386-1392.
Jain, C. (2002) Degradation of mRNA in Escherichia coli. IUBMB Life 54:315-321.
Jain, C. (2008) The E. coli RhlE RNA helicase regulates the function of related RNA
helicases during ribosome assembly. RNA 14:381-389.
Jöchl, C., Rederstorff, M., Hertel, J., Stadler, P. F., Hofacker, I. L., Schrettl, M., Haas, H.,
and Hüttenhofer, A. (2008) Small ncRNA transcriptome analysis from
Aspergillus fumigatus suggests a novel mechanism for regulation of protein
synthesis. Nucleic Acids Res. 36:2677-2689.
Kadaba, S., Krueger, A., Trice, T., Krecic, A. M., Hinnebusch, A. G., and Anderson, J.
(2004) Nuclear surveillance and degradation of hypomodified initiator tRNAMet
in S. cerevisiae. Genes Dev. 18:1227-1240.
134
Kadaba. S., Wang, X., and Anderson, J. T. (2006). Nuclear RNA surveillance in
Saccharomyces
cerevisiae:
Trf4p-dependent
polyadenylation
of
nascent
hypomethylated tRNA and an aberrant form of 5S rRNA. RNA 12:508–521.
Kaplan, R., and Apirion, D. (1975) Decay of ribosomal ribonucleic acid in Escherichia
coli cells starved for various nutrients. J. Biol. Chem. 250:3174-3178.
Kaplan, R., and Apirion, D. (1975) The fate of ribosomes in Escherichia coli cells starved
for a carbon source. J. Biol. Chem. 250:1854-1863.
Kawaji, H., Nakamura, M., Takahashi, Y., Sandelin, A., Katayama, S., Fukuda, S., Daub,
C. O., Kai, C., Kawai, J., Yasuda, J., Carninci, P., and Hayashizaki, Y. (2008)
Hidden layers of human small RNAs. BMC Genomics 9:157-177.
Keiler, K. C. (2008) Biology of trans-translation. Annu. Rev. Microbiol. 62:133-151.
Keiler, K. C., Waller, P. R., and Sauer, R. T. (1996) Role of a peptide tagging system in
degradation of proteins synthesized from damaged messenger RNA. Science
271:990-993.
Kevil, C. G., Walsh, L., Laroux, F. S., Kalogeris, T., Grisham, M. B., and Alexander, J.
S. (1997). An improved, rapid Northern protocol. Biochem. Biophys. Res.
Commun. 238: 277 – 279.
Khemici, V., and Carpousis, A. J. (2004) The RNA degradosome and poly (A)
polymerase of Escherichia coli are required in vivo for the degradation of small
135
mRNA decay intermediates containing REP-stabilizers. Mol. Microbiol. 51:777790.
Kikuchi, A., Takeda, A., Onodera, H., Kimpara, T., Hisanaga, K., Sato, N., Nunomura,
A., Castellani, R. J., Perry, G., Smith, M. A., and Itoyama, Y. (2002) Systemic
increase of oxidative nucleic acid damage in Parkinson's disease and multiple
system atrophy. Neurobiol. Dis. 9:244-248.
King, T. C., Sirdeskmukh, R., and Schlessinger, D. (1986) Nucleolytic processing of
ribonucleic acid transcripts in procaryotes. Microbiol. Rev. 50:428-451.
Komine, Y., Kitabatake, M., Yokogawa, T., Nishikawa, K., and Inokuchi, H. (1994) A
tRNA-like structure is present in 10Sa RNA, a small stable RNA from
Escherichia coli. Proc. Natl. Acad. Sci. USA. 91:9223-9227.
Kong, Q., and Lin, C.L. (2010). Oxidative damage to RNA: mechanisms, consequences,
and diseases. Cell. Mol. Life. Sci. 67:1817-1829.
Kong, Q., Shan, X., Chang, Y., Tashiro, H., and Lin, C. L. (2008) RNA oxidation: a
contributing factor or an epiphenomenon in the process of neurodegeneration.
Free Radic. Res. 42:773-777.
Kuai, L., Fang, F., Butler, J. S., and Sherman, F. (2004) Polyadenylation of rRNA in
Saccharomyces cerevisiae. Proc. Natl. Acad. Sci. USA. 101:8581-8586.
Kushner, S. R. (2002) mRNA Decay in Escherichia coli Comes of Age. J. Bacteriol. 184:
4658–4665.
136
LaRiviere, F. J., Cole, S. E., Ferullo, D. J and Moore, M. J (2006) A late-acting quality
control process for mature eukaryotic rRNAs. Mol. Cell. 24:619-626.
Le Hir, H., Moore, M. J., and Maquat, L.E. (2000) Pre-mRNA splicing alters mRNP
composition: evidence for stable association of proteins at exon-exon junctions.
Genes Dev. 14:1098-1108.
Le Hir, H., Izaurralde, E., Maquat, L. E., and Moore, M. J. (2000) The spliceosome
deposits multiple proteins 20-24 nucleotides upstream of mRNA exon-exon
junctions. EMBO J. 19:6860-6869.
Lee, S. R., and Collins, K. (2005) Starvation-induced cleavage of the tRNA anticodon
loop in Tetrahymena thermophila. J. Biol. Chem. 280:42744-42749.
Lengyel, J. A, and Penman, S. (1977) Differential stability of cytoplasmic RNA in a
Drosophila cell line. Dev. Biol. 57:243-253.
Lewicki, B. T., Margus, T., Remme, J., and Nierhaus, K. H. (1993) Coupling of rRNA
transcription and ribosomal assembly in vivo. Formation of active ribosomal
subunits in Escherichia coli requires transcription of rRNA genes by host RNA
polymerase which cannot be replaced by bacteriophage T7 RNA polymerase. J.
Mol. Biol. 231:581-593.
Li, Z., and Deutscher, M. P. (1994) The role of individual exoribonucleases in processing
at the 3' end of Escherichia coli tRNA precursors. J. Biol. Chem. 269:6064-6071.
137
Li, Z., and Deutscher, M. P. (2004) Exoribonucleases and endoribonucleases. In Curtiss,
R., III (Editor in Chief), Ecosal-Escherichia coli and Salmonella: Cellular and
Molecular Biology. Chapter 4.6.3 ([online] http://www.ecosal.org. ASM Press,
Washington, DC. )
Li, Z. and Deutscher, M.P. (2009). Analyzing the Decay of stable RNAs in E .coli.
Methods Enzymol. 447:31 – 45.
Li, Z., Pandit, S., and Deutscher, M. P. (1998) Polyadenylation of stable RNA precursors
in vivo. Proc. Natl. Acad. Sci. USA. 95:12158-12162.
Li, Z., Pandit, S., and Deutscher, M.P. (1999) RNase G (CafA protein) and RNase E are
both required for the 5' maturation of 16S ribosomal RNA. EMBO J. 18:28782885.
Li, Z., Reimers, S., Pandit, S., and Deutscher, M. P. (2002) RNA quality control:
degradation of defective transfer RNA. EMBO J. 21:1132-1138.
Li, Z., Wu, J., and Deleo, C. J. (2006) RNA Damage and Surveillance under Oxidative
Stress. IUBMB Life 58:581 – 588.
Liu, J., Head, E., Gharib, A.M., Yuan, W., Ingersoll, R.T., Hagen, T.M., Cotman, C.W.,
and Ames, B.N. (2002). Memory loss in old rats is associated with brain
mitochondrial decay and RNA/DNA oxidation: partial reversal by feeding acetylL-carnitine and/or R-alpha-lipoic acid. Proc. Natl. Acad. Sci. 99: 2356 – 2361.
138
Liu, M., Gong, X., Alluri, R. K., Wu, J., Sablo, T., and Li, Z. (2012) Characterization of
RNA damage under oxidative stress in Escherichia coli. Biol. Chem. 393:123–
132.
Loeb, J. N., Howell, R. R., and Tomkins, G. M. (1965) Turnover of ribosomal RNA in rat
liver. Science 149:1093-1095.
Maki, H., and Sekiguchi, M. (1992) MutT protein specifically hydrolyses a potent
mutagenic substrate for DNA synthesis. Nature 355:273-275.
Mangus, D. A., Evans, M. C., and Jacobson, A. (2003) Poly(A)-binding proteins:
multifunctional scaffolds for the post-transcriptional control of gene expression.
Genome Biol. 4:223.
Maquat, L. E. (2004) Nonsense-mediated mRNA decay: splicing, translation and mRNP
dynamics. Nat. Rev. Mol. Cell Biol. 5:89-99.
Maruyama, H., and Mizuno, D. (1970) Ribosome degradation and the degradation
products in starved Escherichia coli. I. Comparison of the degradation rate and of
the nucleotide pool between Escherichia coli B and Q-13 strains in phosphate
deficiency. Biochim. Biophys. Acta. 199:159-165.
Martinet, W., de Meyer, G.R., Herman, A.G., Kockx, M.M. (2004) Reactive oxygen
species induce RNA damage in human atherosclerosis. Eur J Clin Invest. 34: 323327.
139
Masaki, H., and Ogawa, T. (2002) The modes of action of colicins E5 and D, and related
cytotoxic tRNases. Biochimie. 84:433-438.
McKinlay, A., Gerard, W., and Fields, S. (2012) Global analysis of RNA oxidation in
Saccharomyces cerevisiae. Biotechniques 52:109-111.
Meselson, M., Nomura, M., Brenner, S., Davern, C., and Schlessinger, D. (1964)
Conservation of ribosomes during bacterial growth. Mol. Biol. 9: 696-711.
Miczak, A., Kaberdin, V. R., Wei, C. L., and Lin-Chao, S. (1996) Proteins associated
with RNase E in a multicomponent ribonucleolytic complex. Proc. Natl. Acad.
Sci. USA. 93:3865-3869.
Mitchell, P. and Tollervey, D. (2000) mRNA stability in eukaryotes. Curr. Opin. Genet.
Dev. 10:193-198.
Mohanty, B. K., and Kushner, S. R. (2003) Genomic analysis in Escherichia coli
demonstrates differential roles for polynucleotide phosphorylase and RNase II in
mRNA abundance and decay. Mol. Microbiol. 50:645-658.
Mroczek, S., and Kufel, J. (2008) Apoptotic signals induce specific degradation of
ribosomal RNA in yeast. Nucleic Acids Res. 36:2874-2888.
Mukhopadhyay, P., Gangopadhyay, T., and Ghosh, R. K. (1994) Unusual in vivo
turnover of transfer RNA in Vibrio cholerae. Microbiology 140:1775-1779.
140
Nunomura, A., Chiba, S., Kosaka, K., Takeda, A., Castellani, R.J., Smith, M.A., and
Perry, G. (2002) Neuronal RNA oxidation is a prominent feature of dementia with
lewy bodies. Neuroreport 13:2035-2039.
Nunomura, A., Hofer, T., Moreira, P.I., Castellani, R.J., Smith, M.A., and Perry, G.
(2009). RNA oxidation in Alzheimer disease and related neurodegenerative
disorders. Acta Neuropathol. 118: 151-166.
Nunomura, A., Honda, K., Takeda, A., Hirai, K., Zhu, X., Smith, M.A., and Perry, G.
(2006) Oxidative damage to RNA in neurodegenerative disease. J. Biomed.
Biotechnol. 2006:82323.
Nunomura, A., Perry, G., Pappolla, M. A., Wade, R., Hirai, K., Chiba, S., and Smith, M.
A. (1999) RNA oxidation is a prominent feature of vulnerable neurons in
Alzheimer's disease. J. Neurosci. 19:1959-1964.
Ogawa, T., Tomita, K., Ueda, T., Watanabe, K., Uozumi, T. and Masaki, H. (1999) A
cytotoxic ribonuclease targeting specific transfer RNA anticodons. Science
283:2097–2100.
Parker, R., and Sheth, U. (2007) P bodies and the control of mRNA translation and
degradation. Mol. Cell. 25:635-646.
Parker, R., and Song, H. (2004) The enzymes and control of eukaryotic mRNA turnover.
Nat. Struct. Mol. Biol. 11:121-127.
141
Passos, D. O., Doma, M. K, Shoemaker, C. J, Muhlrad, D., Green, R., Weissman, J.,
Hollien, J., and Parker, R. (2009) Analysis of Dom34 and its function in no-go
decay. Mol. Biol. Cell 20:3025-3032.
Phizicky, E. M., and Hopper, A. K. (2010) tRNA biology charges to the front. Genes
Dev. 24:1832-1860.
Poulsen, H. E., Specht, E., Broedbaek, K., Henriksen, T., Ellervik, C., Mandrup-Poulsen,
T., Tonnesen, M., Nielsen, P. E., Andersen, H. U., and Weimann, A. (2012) RNA
modifications by oxidation: A novel disease mechanism? Free Radic Biol Med.
Jan 28. [Epub ahead of print]
Radak, Z. and Boldogh, I. (2010). 8-Oxo-7,8-dihydroguanine: links to gene expression,
aging, and defense against oxidative stress. Free Radic. Biol. Med. 49:587 – 596.
Reinisch, K. M., and Wolin, S. L. (2007) Emerging themes in non-coding RNA quality
control. Curr. Opin. Struct. Biol. 17:209-214.
Rhee, Y., Valentine, M. R., and Termini, J. (1995) Oxidative base damage in RNA
detected by reverse transcriptase. Nucleic Acids Res. 23:3275-3282.
Richards, J., Liu, Q., Pellegrini, O., Celesnik, H., Yao, S., Bechhofer, D. H., Condon, C.,
and Belasco, J. G. (2011) An RNA pyrophosphohydrolase triggers 5'exonucleolytic degradation of mRNA in Bacillus subtilis. Mol. Cell 43:940-949.
142
Richards, J., Mehta, P., and Karzai, A. W. (2006) RNase R degrades non-stop mRNAs
selectively in an SmpB-tmRNA-dependent manner. Mol. Microbiol 62:17001712.
Ron, E., Kohler, R., and Davis, B. (1966). Polysomes extracted from Escherichia coli by
freeze-thaw-lysozyme lysis. Science 153: 1119 – 1120.
Schein, C.H. (2001) Nuclease methods and protocols. Methods in molecular biology. 160
Schlegel, R.A., Iversen, P., and Rechsteiner, M. (1978) The turnover of tRNAs
microinjected into animal cells. Nucleic Acids Res. 5:3715-3729.
Selinger, D.W., Saxena, R. M., Cheung, K. J., Church, G. M., and Rosenow, C. (2003)
Global RNA half-life analysis in Escherichia coli reveals positional patterns of
transcript degradation. Genome Res. 13: 216-223.
Seo, A.Y., Xu, J., Servais, S., Hofer, T., Marzetti, E., Wohlgemuth, S.E., Knutson, M.D.,
Chung, H.Y., and Leeuwenburgh, C. (2008). Mitochondrial iron accumulation
with age and functional consequences. Aging Cell 7: 706-716.
Shahbabian, K., Jamalli, A., Zig, L., and Putzer, H. (2009) RNase Y, a novel
endoribonuclease, initiates riboswitch turnover in Bacillus subtilis. EMBO J.
28:3523-3533.
Shan, X., Chang, Y., and Lin C.L. (2007) Messenger RNA oxidation is an early event
preceding cell death and causes reduced protein expression. FASEB J. 21:27532764.
143
Shan, X., and Lin, C. L. (2006) Quantification of oxidized RNAs in Alzheimer's disease.
Neurobiol. Aging 27:657-662.
Shan, X., Tashiro, H., and Lin, C. L. (2003) The identification and characterization of
oxidized RNAs in Alzheimer's disease. J. Neurosci. 23:4913-4921.
Shen, Z., Wu, W., and Hazen, S. L. (2000) Activated leukocytes oxidatively damage
DNA, RNA, and the nucleotide pool through halide-dependent formation of
hydroxyl radical. Biochemistry 39:5474-5482.
Siehnel, R. J., and Morgan, E. A. (1985) Unbalanced rRNA gene dosage and its effects
on rRNA and ribosomal-protein synthesis. J. Bacteriol. 163:476-486.
Slomovic, S., Laufer, D., Geiger, D., and Schuster, G. (2006) Polyadenylation of
ribosomal RNA in human cells. Nucleic Acids Res. 34:2966-2975.
Smith, J. D. (1975) Mutants which allow accumulation of tRNAtYr precursor molecules,
Brookhaven Symp. Biol. 26:1-11.
Spickler, C., and Mackie, G. A. (2000) Action of RNase II and polynucleotide
phosphorylase against RNAs containing stem-loops of defined structure. J.
Bacteriol. 182:2422-2427.
Steglich, C., Lindell, D., Futschik, M., Rector, T., Steen, R., Chisholm, S.W. (2010)
Short
RNA
half-lives
in
the
slow-growing
Prochlorococcus. Genome Biol. 11:R54.
144
marine
cyanobacterium
Szé kely, M., Brimacombe, R., and Morgan, J. (1973). A specifi c ribonucleoprotein
fragment from Escherichia coli 30-S ribosomes. Location of the RNA component
in 16-S RNA. Eur. J. Biochem. 35: 574 – 581.
Taddei, F., Hayakawa, H., Bouton, M., Cirinesi, A., Matic, I., Sekiguchi, M., and
Radman, M. (1997) Counteraction by MutT protein of transcriptional errors
caused by oxidative damage. Science 278:128-130.
Tanaka, M., Chock, P.B., and Stadtman, E.R. (2007). Oxidized messenger RNA induces
translation errors. Proc. Natl. Acad. Sci. USA. 104: 66 – 71.
Tanaka, M., Han, S., Küpfer, P. A., Leumann, C. J., and Sonntag, W. E. (2011a).
Quantification of oxidized levels of specific RNA species using an aldehyde
reactive probe. Anal.Biochem.417:142-148.
Tanaka, M., Han, S., Küpfer, P. A., Leumann, C. J., and Sonntag, W. E. (2011b). An
assay for RNA oxidation induced abasicsites using the Aldehyde Reactive Probe.
Free Radic. Res. 45: 237-247.
Temple, M. D., Perrone, G. G., and Dawes, I. W. (2005) Complex cellular responses to
reactive oxygen species. Trends Cell Biol. 15:319-326.
Thomason, L.C., Costantino, N., and Court, D.L. (2007) E. coli Genome Manipulation by
P1 Transduction. Curr. Protoc. Mol. Biol. Chapter 1:Unit 1.17.
Thompson, D. M., Lu, C., Green, P. J., and Parker, R. (2008) tRNA cleavage is a
conserved response to oxidative stress in eukaryotes. RNA 14:2095-2103.
145
Thompson, D. M., and Parker, R. (2009) The RNase Rny1p cleaves tRNAs and promotes
cell death during oxidative stress in Saccharomyces cerevisiae. J. Cell Biol.
185:43-50.
Thorp, H. H. (2000) The importance of being r: greater oxidative stability of RNA
compared with DNA. Chem. Biol. 7:R33-6.
Tomita, K., Ogawa, T., Uozumi, T., Watanabe, K., and Masaki, H. (2000) A cytotoxic
ribonuclease which specifically cleaves four isoaccepting arginine tRNAs at their
anticodon loops. Proc. Natl. Acad. Sci. USA. 97:8278-8283.
Tu, G. F., Reid, G. E., Zhang, J. G., Moritz, R. L., and Simpson, R. J. (1995) C-terminal
extension of truncated recombinant proteins in Escherichia coli with a 10Sa RNA
decapeptide. J. Biol. Chem. 270:9322-9326.
Vaidyanathan, P.P., Deutscher, M.P., and Malhotra, A. (2007). RluD, a highly conserved
pseudouridine synthase, modifi es 50S subunits more specifi cally and effi ciently
than free 23S rRNA. RNA 13: 1868 – 1876.
Valadkhan, S. (2005) snRNAs as the catalysts of pre-mRNA splicing. Curr. Opin. Chem.
Biol. 9:603-608.
van Hoof, A., Frischmeyer, P. A., Dietz, H.C., and Parker, R. (2002) Exosome-mediated
recognition and degradation of mRNAs lacking a termination codon. Science
295:2262-2264.
146
van Hoof, A., and Parker, R. (1999) The exosome: a proteasome for RNA? Cell 99:347350.
Vincent, H. A., and Deutscher, M. P. (2006) Substrate recognition and catalysis by the
exoribonuclease RNase R. J. Biol. Chem. 281:29769-29775.
Wamer, W. G., and Wei, R. R. (1997) In vitro photooxidation of nucleic acids by
ultraviolet A radiation. Photochem. Photobiol. 65:560-563.
Wardman, P. and Candeias, L. P. (1996). Fenton chemistry: an introduction. Radiat. Res.
145: 523 – 531.
Weidner, A. M., Bradley, M. A., Beckett, T. L., Niedowicz, D. M., Dowling, A. L.,
Matveev, S. V., Levine, H III., Lovell, M. A., and Murphy, M. P. (2011). RNA
Oxidation Adducts 8-OHG and 8-OHA Change with Aβ(42) Levels in Late-Stage
Alzheimer's Disease. PLoS One 6: e24930.
Wu, J., Jiang, Z., Liu, M., Gong, X., Wu, S., Burns, C. M., and Li, Z. (2009)
Polynucleotide phosphorylase protects Escherichia coli against oxidative stress.
Biochemistry 48:2012-2020.
Wu, J., and Li Z. (2008) Human polynucleotide phosphorylase reduces oxidative RNA
damage and protects HeLa cell against oxidative stress. Biochem. Biophys. Res.
Commun. 372:288-292.
Wurtmann, E. J., and Wolin, S. L. (2009). RNA Under Attack: Cellular Handling of
RNA Damage. Crit. Rev. Biochem. Mol. Biol. 44:34-49.
147
Yamamoto, Y., Sunohara, T., Jojima, K., Inada, T., and Aiba, H. (2003) SsrA-mediated
trans-translation plays a role in mRNA quality control by facilitating degradation
of truncated mRNAs. RNA 9:408-418.
Zhang, J., Perry, G., Smith, M.A., Robertson, D., Olson, S.J., Graham, D.G., and
Montine, T.J. (1999) Parkinson's disease is associated with oxidative damage to
cytoplasmic DNA and RNA in substantia nigra neurons. Am. J. Pathol. 154:14231429.
Zundel, M. A., Basturea, G. N, and Deutscher, M. P. (2009) Initiation of ribosome
degradation during starvation in Escherichia coli. RNA 15: 977–983.
148