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Dissertation presented at Uppsala University to be publicly examined in BMC B41,
Husargatan 3, Uppsala, Friday, May 8, 2009 at 13:15 for the degree of Doctor of Philosophy.
The examination will be conducted in English.
Abstract
Henriksson, N. 2009. Poly(A)-specific Ribonuclease (PARN). Structural and Functional
Studies of Poly(A) Recognition and Degradation. Acta Universitatis Upsaliensis. Digital
Comprehensive Summaries of Uppsala Dissertations from the Faculty of Science and
Technology 629. 65 pp. Uppsala. 978-91-554-7484-3.
Regulation of mRNA degradation is a powerful way for the cell to regulate gene expression.
A critical step in eukaryotic mRNA degradation is the removal of the poly(A) tail at the 3'end of the mRNA. Poly(A)-specific ribonuclease (PARN) is an oligomeric, processive and
capinteracting 3'-5' exoribonuclease that efficiently degrades eukaryotic mRNA poly(A) tails.
In addition to the exonuclease domain, PARN harbors two RNA-binding domains, a classical
RNA recognition motif (RRM) and an R3H-domain. In this project we have studied mechanisms by which PARN specifically recognizes and degrades poly(A).
We investigated the RNA binding properties of PARN by using electrophoretic mobility
shift assays and filter-binding analysis and we could show that PARN binds poly(A) with
high affinity and specificity. Furthermore, we showed that the RRM and R3H domains of
PARN separately could bind to poly(A).
To investigate specificity for and recognition of poly(A) in the active site of PARN, we performed a kinetic analysis on a repertoire of trinucleotide substrates in vitro. We showed that
PARN harbors affinity for adenosines in the active site and that both the penultimate and the
3' end located nucleotide play an important role for providing adenosine-specificity in the
active site of PARN.
Moreover, we solved a crystal structure of PARN in complex with m7GpppG cap analogue
and showed that the cap binding and active sites overlap both structurally and functionally. By
mutational analysis we identified residues in the active site that specifically recognize adenosines. Furthermore, biochemical data showed that the adenosine specificity in the active site is
lost when Mn2+ is used instead of Mg2+ as divalent metal ion.
Taken together, these results demonstrate that both RNA-binding properties of the RRM and
R3H-domains in addition to base recognition in the active site contributes to PARN poly(A)specificity.
Keywords: mRNA degradation, deadenylation, PARN, poly(A) specificity, RRM, R3Hdomain, active site, trinucleotides
Niklas Henriksson, Department of Cell and Molecular Biology, Box 596, Uppsala University,
SE-75124 Uppsala, Sweden
© Niklas Henriksson 2009
ISSN 1651-6214
ISBN 978-91-554-7484-3
urn:nbn:se:uu:diva-98710 (http://urn.kb.se/resolve?urn=urn:nbn:se:uu:diva-98710
In memory of my mother
List of Publications
This thesis is based on the following papers, which are referred to in the text
by their Roman numerals.
*
I
Nilsson, P., Henriksson, N., Niedzwiecka, A., Balatsos,
A.A.N., Kokkoris, K., Eriksson, J. and Virtanen, A. (2007) A
Multifunctional RNA Recognition Motif in Poly(A)-specific
Ribonuclease with Cap and Poly(A) Binding Properties.
J. Biol. Chem., 282, 32902-32911
II
*
Wu, M., *Nilsson, P., *Henriksson, N., Niedzwiecka, A., Lim,
M. K., Cheng, Z., Kokkoris, K., Virtanen, A. and Song, H.
(2009) Structural Basis of m7GpppG Binding to Poly(A)Specific Ribonuclease.
Structure, 17, 276-286
III
Henriksson, N., Nilsson, P., Wu, M., Song, H. and Virtanen, A.
Recognition of adenosine residues by the active site of poly(A)specific ribonuclease (PARN).
Manuscript 2009
IV
Henriksson, N., Nilsson, P., and Virtanen, A. Poly(A)-specific
ribonuclease (PARN) loses specificity for recognition of adenosine residues in the presence of Mn2+.
Manuscript 2009
V
Henriksson, N., Nilsson, P., and Virtanen, A. RNA binding
properties of the R3H domain of poly(A)-specific ribonuclease
(PARN).
Manuscript 2009
These authors contributed equally
Reprints were made with permission from the respective publishers.
Contents
General background ......................................................................................11
The life cycle of eukaryotic mRNAs........................................................11
mRNA processing ....................................................................................12
Capping and polyadenylation ..............................................................12
mRNA transport and initiation of translation ......................................13
mRNA degradation ..................................................................................14
Pathways of mRNA degradation .........................................................14
Enzymes involved in mRNA decay.....................................................15
mRNA surveillance .............................................................................19
mRNA degradation in prokaryotes ......................................................21
Regulation of mRNA decay .....................................................................21
AU-rich elements.................................................................................21
miRNAs ...............................................................................................22
PUF-proteins........................................................................................23
P-bodies ...............................................................................................23
RNA-binding domains .............................................................................23
RRM ....................................................................................................24
R3H......................................................................................................25
Other RNA-binding domains...............................................................26
Present investigations....................................................................................27
Introduction ..............................................................................................27
Aim...........................................................................................................29
Results ......................................................................................................31
The RRM of PARN binds both the cap and poly(A) (Paper I)............31
The R3H-domain of PARN binds RNA (Paper V)..............................33
The structure of PARN in complex with Cap (Paper II) .....................33
Recognition of adenosine residues by the active site of PARN (Paper
III)........................................................................................................35
Poly(A) specificity can be modulated when Mn2+ is used as divalent
metal ion (Paper IV) ............................................................................37
Discussion ................................................................................................38
Cap and poly(A) binding .....................................................................38
Processivity: A comparison with RNase II..........................................38
Specificity: The RNA-binding sites and the active site .......................39
Poly(A) recognition in poly(A)-binding proteins ................................40
Other DEDD exonucleases: A comparison with PARN......................41
PARN involved in degrading oligouridylated mRNAs? .....................43
Conclusions and future perspectives ........................................................43
Acknowledgements.......................................................................................45
Swedish summary .........................................................................................48
References.....................................................................................................50
”Det är den som går vilse som finner de nya vägarna.”
Nils Kjaer
Abbreviations
A
ARE
AMD
C
CBC
Ccr4
CPSF
Dcp1/2
DcpS
DNA
eIF4E
EMSA
G
Kd
Km
kDa
miRNA
mRNA
mRNP
NGD
NMD
NSD
PABP
PABP2
Pan
PAP
PARN
P-body
RNA
RNA pol
RNAse
RRM
RBD
U
UTR
Xrn1
adenosine
AU-rich element
ARE-mediated decay
cytidine
cap-binding complex
carbon catabolite repressor 4
cleavage and polyadenylation specificity factor
decapping protein 1/2
scavenger decapping enzyme
deoxyribonucleic acid
eukaryotic translational initiation factor 4E
electrophoretic mobility shift assay
guanosine
equilibrium dissociation constant
Michaelis-Menten constant
kilo Dalton
micro RNA
messenger RNA
messenger ribonucleoprotein
no-go dacay
nonsense-mediated decay
non-stop decay
cytoplasmic poly(A)-binding protein
nuclear poly(A)-binding protein
poly(A)-nuclease
poly(A)-polymerase
poly(A)-specific ribonulease
processing body
ribonucleic acid
RNA polymerase
ribonuclease
RNA-recognition motif
RNA-binding domain
uridine
untranslated region
exoribonuclease
General background
The life cycle of eukaryotic mRNAs
The central dogma in molecular biology describes the flow of genetic information from DNA via messenger RNA (mRNA) to protein. The genes encoded by DNA in the nucleus are transcribed into pre-mRNA in a process
called transcription. Transcription takes place in the nucleus and proteincoding genes are transcribed by RNA polymerase II (RNA pol II)
(Kornberg, 2001). The pre-mRNA is further processed in different ways
before transport to the cytoplasm. A cap consisting of two guanosine residues connected by a 5’-5’-triphosphate bridge (m7GpppG) is added to the 5’end of the mRNA and at the 3’-end, a poly(A)-tail consisting of 60-200
adenosine residues is added (Shatkin and Manley, 2000). In a third process,
called splicing, the non-coding introns present in pre-mRNA are removed
(Wahl et al., 2009). All processes are coupled and occur to a large extent
cotranscriptionally (Proudfoot et al., 2002; Zorio and Bentley, 2004). In the
cytoplasm, the mRNA is translated into proteins by the ribosome, a large
complex consisting of both RNA and proteins (Steitz, 2008). When the cell
does not need the protein that is translated from its coding mRNA, the
mRNA undergoes degradation. The half-lives of eukaryotic mRNAs vary
widely, from minutes to several hours. Interestingly, mRNAs encoding proteins that functions in the same process or act together in the same complex
seems to follow the same route of decay (Wang et al., 2002b).
Importantly, it is the balance between synthesis and degradation that sets
the steady-state level of the mRNA (GarcIa-MartInez et al., 2004). One can
think of the mRNA-pool as a water sink, where the incoming water from the
tap represents transcription and the water flushing down the drain represents
mRNA degradation. In order to regulate gene expression, the cell can either
control the synthesis of mRNA at the transcriptional level (adjust the flow
rate of the water-tap) or regulate the rate by which the mRNA is degraded
(adjust the flow rate down the drain). Historically, transcriptional control has
been well studied, but the post-transcriptional regulation at the mRNA-level
has been taken more and more into consideration (Molina-Navarro et al.,
2008). Therefore, the regulation of mRNA decay is a powerful way for the
cell to control gene expression (Wilusz and Wilusz, 2004). A large amount
of regulation is taking place at the first and rate-limiting step of mRNA degradation: deadenylation (Cao and Parker, 2001).
11
In this thesis I will describe one of the enzymes involved in the deadenylation step of mRNA degradation, poly(A)-specific ribunuclease (PARN).
I have focused on the mechanisms by which PARN specifically recognizes
and degrades poly(A).
mRNA processing
Capping and polyadenylation
When the nascent mRNA is approximately 20-35 nucleotides long and still
being transcribed by RNA pol II, the 5’ end undergoes capping. The
m7GpppG cap structure is linked to the 5’ end of the mRNA by a three step
reaction involving three enzymatic activities (Proudfoot et al., 2002; Shatkin
and Manley, 2000). The 5’ triphosphate of the first transcribed nucleotide
(most commonly a G, but sometimes an A) is first hydrolyzed to a diphosphate by an RNA 5’ triphosphatase (RTP). Then a GMP moiety is added
from GTP by guanylyltransferase (GT) via a 5’-5’ triphosphate linkage. Finally, the N7 position of the added GMP is methylated by a methyltransferase (MT). In yeast, three different enzymes are responsible for these enzymatic activities, while in mammals both the RTP and GT activities are part
of the same polypeptide. Importantly, all enzymes involved in capping bind
to the hyperphosphorylated C-terminal of RNA pol II, therefore allowing
capping to take place cotranscriptionally (Schroeder et al., 2000).
Polyadenylation is a two-step mechanism that involves both cis-acting
elements and trans-acting factors (reviewed in (Danckwardt et al., 2008;
Shatkin and Manley, 2000)). First, the poly(A)-signal AAUAAA in the 3’
untranslated region (UTR) is recognized by the cleavage and polyadenylation specificity factor (CPSF) consisting of at least four subunits. One component of this complex, CPSF 73 then cleaves the RNA at the poly(A)-site
located 10-30 nucleotides downstream of the AAUAAA sequence element
(Mandel et al., 2006; Ryan et al., 2004). Another important cis-element is the
GU-rich sequence located 30 nucleotides downstream of the poly(A)-site.
The heterotrimeric cleavage-stimulating factor (CstF) binds this downstream
element thereby promoting the efficiency of 3’ end processing. Second, a
poly(A)-tail consisting of 200-250 adenosines in mammals and 60-80 adenosines in yeast is added to the new 3’ terminus by Poly(A)-polymerase (PAP)
(Edmonds, 1990). Note that PAP adds the adenosines through a nontemplated reaction, in contrast to the DNA and RNA polymerases. As in the
case of capping, the C-terminal end of RNA polymerase interacts with components in both CPSF and CstF. Therefore, the C-terminal end of Pol II
12
works as a processing platform that enhances both capping and polyadenylation (Howe, 2002).
The histone mRNAs lack poly(A)-tails. Instead they possess a conserved
stem-loop at the 3’ end. The initial step in the 3’ processing of histone
mRNAs seems to work in the same way as with mRNAs with poly(A)-tails.
The stem-loop binding protein (SLBP) binds the stem-loop and recruits
CstF and CPSF that cleaves the histone mRNA (Dominski and Marzluff,
2007).
mRNA transport and initiation of translation
The cap and the poly(A)-tail are specific features that define the two ends of
the mRNA and help the different cellular machineries to distinguish mRNAs
from other RNAs (e.g. rRNAs, tRNAs, snRNAs and snoRNAs). Noteworthy, mRNAs usually do not exist as naked molecules. Instead they are found
in complex with a plethora of RNA-binding proteins, building up messenger
ribonucleoproteins (mRNPs) (Glisovic et al., 2008). The 5’ located cap and
the 3’ located poly(A)-tail are important docking-sites for various RNAbinding proteins that are important for both the transport of the mRNA from
the nucleus to the cytoplasm and for the correct initiation of translation
(Kohler and Hurt, 2007; Mendez and Richter, 2001).
In the nucleus, the cap is recognized and bound by the cap-binding complex (CBC). CBC consists of a large and a small subunit, CBP 80 and CBP
20 (Izaurralde et al., 1994). CBP 20 binds the cap (Calero et al., 2002;
Mazza et al., 2002) and CBP 80 plays a regulatory role by interacting with
various proteins. CBP assists in mRNA transport to the cytoplasm by a direct
interaction with the mRNA export machinery (Cheng et al., 2006). Studies
of export of the large Balbiani ring mRNP (Wieslander, 1979) shows that
ribosomes start translating this mRNA in the cytoplasm before the export is
fully accomplished (Mehlin et al., 1992). That is a good evidence that this
mRNP is transported out of the nucleus with the 5’ end first. It has been suggested that CBP 80 directs the transport in the 5’ to 3’ direction (Cheng et
al., 2006; Visa et al., 1996). In the nucleus, the newly synthesized poly(A)tail is coated by many copies of the nuclear poly(A) binding protein
(PABP2). The yeast homolog of PABP2 binds transportin, which implies an
active role for PABP2 in mRNA transport.
After transport to the cytoplasm, PABP2 dissociates and shuttles back to
the nucleus (Brune et al., 2005). Instead, the cytoplasmic poly(A)-binding
protein (PABP) binds to the poly(A)-tail (see (Kuhn and Wahle, 2004) for
review). At the 5’ end, CBC is exchanged for the eukaryotic translational
initiation factor 4E (eIF4E) that will bind the cap in the cytoplasm
(Marcotrigiano et al., 1997). eIF4E together with eIF4G and eIF4A forms
13
eIF4F that recruits the ribosome to the mRNA facilitating efficient initiation
of translation. Interestingly, eIF4E associates with eIF4G that simultaneously binds PABP. Therefore, this leads to a circularization of the mRNA
upon ongoing translation (Figure 1). This phenomenon has been visualized
by electron microscopy and will ensure fast recycling of the ribosome (Wells
et al., 1998). This circular structure must somehow be disrupted when the
mRNA is about to undergo degradation. A key step in the transition from
translation to mRNA decay is removal of the poly(A)-tail.
Figure 1. Pathways of translation and mRNA degradation.
mRNA degradation
Pathways of mRNA degradation
mRNA-degradation normally proceeds through one of two main decay
pathways (Figure 1) (Garneau et al., 2007) (Parker and Song, 2004) (Meyer
et al., 2004). Importantly, both pathways start with the removal of the 3’-end
located poly (A)-tail followed by degradation from the 5’ or 3’ end, respectively. In the deadenylation-dependent 3’5’ decay pathway the mRNA is
degraded from the 3’-end by the exosome, followed by cap-hydrolysis by the
14
scavenger decapping enzyme (DcpS). In the deadenylation-dependent 5’3’
decay pathway, the cap is removed by the decapping protein1/decapping
protein2 (Dcp1/Dcp2) complex and subsequently the mRNA is degraded
from the 5’-end by exoribonuclease1 (Xrn1). In yeast, the deadenylation
dependent 5’3’ pathway seems to be dominant, whereas in mammals the
deadenylation dependent 3’5’ pathway is regarded as the main pathway.
Some transcripts in yeast are degraded without the initial deadenylation
event. (Badis et al., 2004; Muhlrad and Parker, 2005). Instead, in the so
called deadenylation-independent decay pathway, these mRNAs first recruit
the decapping complex Dcp1/Dcp2. After decapping, the mRNA is degraded
5’3’ by Xrn1.
In the endonuclease-mediated decay pathway, degradation is initiated by
endonucleolytic cleavage of the mRNA. An interesting example is RNA
interference, where small interfering RNAs (siRNAs) induce mRNA decay
by endonucleolytic cleavage by argonaute 2 (Ago2) as part of the RNA induced silencing complex (RISC) (reviewed in(Hammond, 2005; McManus
and Sharp, 2002; Rana, 2007)). Thereafter, the generated 3’ and 5’ ends are
degraded by the exosome and Xrn1, respectively (Orban and Izaurralde,
2005).
Enzymes involved in mRNA decay
There are a number of different enzymes with different functions that participate in mRNA decay. A more detailed description of these enzymes follows.
Deadenylases
There are three main deadenylating activities that have been studied so far,
the Carbon catabolite repressor protein 4 (Ccr4)/Pop2-complex, the poly(A)nuclease2/poly(A)-nuclease3 (Pan2/Pan3)-complex and poly(A)-specific
ribonuclease (PARN) (reviewed in (Goldstrohm and Wickens, 2008).
The Ccr4/Pop2 complex is considered to be the major deadenylase in
yeast (Tucker et al., 2001). Ccr4 belongs to the exonuclease-endonucleasephosphatase (EEP) superfamily of ribonucleases and is the active 3’5’
deadenylase of the complex in Saccharomyces cerevisiae (Chen et al., 2002;
Tucker et al., 2002). Its activity is inhibited by PABP (Tucker et al., 2002).
Homologues of Ccr4 have been identified in higher eukaryotes (Dupressoir
et al., 2001), at least some of them harboring deadenylase activity (Baggs
and Green, 2003). Pop2 (also known as Ccr4 associating factor(Caf1)) is a
member of the DEDD superfamily of ribonucleases (Zuo and Deutscher,
15
2001) and shares homology with the Escherichia coli RNaseD 3’5’ exonuclease domain both in its amino acid sequence (Moser et al., 1997) as
well as structurally (Thore et al., 2003). Beacause Pop2 deadenylase activity
in the S. cerevisiae Ccr4/Pop2 complex is missing due to lack of catalytically
important amino acids in the active site, an alternative role for Pop2 might
be to enhance the activity of Ccr4. However, in higher eukaryotes Pop2 is an
active deadenylase (Bianchin et al., 2005; Schwede et al., 2008; Viswanathan et al., 2004) . Interestingly, Ccr4 and Pop2 are components of a large 1
mega Dalton (MDa) complex with the Not1-5 proteins and Caf40 and
Caf130 (Chen et al., 2001b). Because the Not1-5 proteins have been implicated as transcription factors (Liu et al., 1998), the Ccr4/Pop2/Not-complex
shows a coupling between transcription and mRNA decay.
The second deadenylating activity, the Pan2/Pan3 complex, is responsible
for the initial poly(A)-tail shortening both in yeast (Brown and Sachs, 1998)
and mammals (Yamashita et al., 2005). Pan2 belongs to the DEDD superfamily of ribonucleases and is the catalytically active subunit (Boeck et al.,
1996), whereas Pan3 binds to Pan2 and enhances the activity (Brown et al.,
1996). In contrast to Ccr4/Pop2, Pan2/Pan3 activity is stimulated by PABP
(Sachs and Deardorff, 1992). Pan3p interacts physically with PABP to
stimulate Pan2 activity (Mangus et al., 2004; Uchida et al., 2004).
Poly(A)-specific ribonuclease (PARN) differs from the other deadenylating activities in that it interacts with the 5’ cap-structure in addition to the 3’
poly(A)-tail (Dehlin et al., 2000; Gao et al., 2000; Martinez et al., 2000).
Even if the cap is not absolutely necessary for poly(A) hydrolysis by PARN,
it stimulates PARN activity (Dehlin et al., 2000; Gao et al., 2000; Martinez
et al., 2000) and thereby amplifies the processivity of PARN action
(Martinez et al., 2001). Therefore, the cap interaction with PARN leads to a
fast and efficient degradation of capped mRNAs. In turn, capped mRNAs
undergoing deadenylation by PARN are unavailable for the translational
initiation complex because their cap-structures are bound to PARN. The
involvement of PARN in mRNA decay is not fully understood at the moment. It has been argued that PARN is not involved in default mRNA degradation (Yamashita et al., 2005) while others have argued that PARN is the
major deadenylase involved in mRNA decay (Gao et al., 2000). PARN is
present in all higher eukaryotes, but is absent in S. cerevisiae and Drosophila
melanogater. PARN is involved in deadenylation of maternal mRNAs in
Xenopus laevis oocyte maturation and therefore plays a key role during development (Copeland and Wormington, 2001; Korner et al., 1998). PARN
was found in the same complex as the poly(A)-polymerase Gld2, cleavage
and polyadenylation specificity factor (CPSF) and cytoplasmic polyadenylation element binding protein (CPEB) (Kim and Richter, 2006). The two last
proteins are factors involved in the polyadenylation machinery and anchors
PARN and Gld to the 3’ end of the mRNA. Poly(A)-tail length remains short
by default because PARN activity overcomes the polyadenylating activity of
16
Gld2. Upon oocyte maturation, a signalling cascade starts that phosphorylates CPEB, causing PARN to dissociate from the complex. That allows
Gld2 to polyadenylate the mRNA and therefore induces translation of the
transcript.
3’5’ exonucleases: The exosome
The exosome, a multiprotein complex with 3’5’ exonuclease activity was
first discovered in yeast (Mitchell et al., 1997). It consists of a core of six
proteins: Rrp41, Rrp42, Rrp43, Rrp45, Rrp46 and Mtr3p forming a ring
structure. Three proteins, Rrp4, Csl4 and Rrp40 are located on top of the
barrel forming a “cap”. All six core proteins contain RNase PH –domains
and are homologous to the bacterial polynucleotide phosphorylases
(PNPases). The archaeal subunit Rrp 41 has phosphorolytic activity
(Lorentzen and Conti, 2005; Lorentzen et al., 2005), but none of the core
subunits of the eukaryotic exosome is active (Liu et al., 2006). Instead, it has
been shown that a tenth subunit, Rrp44 (also called Dis 3), is responsible for
the 3’5’ exonucleolytic activity of the eukaryotic exosome (Dziembowski
et al., 2007). Rrp44 belongs to the RNase II family of ribonucleases and the
hydrolytic activity has been mapped to its RNB-domain. Recently it has
been observed that Rrp 44 harbors additional endonucleolytic activity within
its PIN-domain (Lebreton et al., 2008; Schaeffer et al., 2009; Schneider et
al., 2009). The combined actions of endo and exo activities fit well with the
exosome being involved in processing of rRNAs, snoRNAs, snRNAs
(Allmang et al., 1999) and tRNAs, in addition to mRNA degradation from
the 3’-end. Electron microscopy (EM) pictures of the exosome (Wang et al.,
2007) has positioned Rrp44 at the bottom of the ring-structured core , but it
seems that Rrp44 is loosely attached to the core in yeast and human
exosomes. Therefore the inactive exosome core is believed to work like a
scaffold to which the other subunits can bind. Furthermore, the exosome is
involved in nuclear mRNA surveillance mechanisms outlined below. Another protein subunit, Rrp6 with hydrolytical exoribonucleolytic activity has
been found explicitly in nuclear exosomes (Midtgaard et al., 2006). Rrp6
belongs to the same DEDD superfamily of ribonucleases as Pop2, Pan2 and
PARN.
5’3’ exonucleases
There are two major 5’3’ exonucleases in yeast, the cytoplasmic Xrn1 and
the nuclear Rat1 (Johnson, 1997). In the cytoplasm, after decapping, mRNAs
are degraded by the 5’3’ exoribonuclease (Xrn1) (Hsu and Stevens, 1993).
The homolog of Xrn1 in D. melanogaster, which has been named Pacman, is
developmentally regulated (Till et al., 1998). However, little is known about
17
domain organization and catalytic mechanism of this enzyme. In the nucleus,
Rat1 (Xrn2 in human) (Amberg et al., 1992; Xiang et al., 2009; Zhang et al.,
1999) is responsible for the 5’3’ degradation of transcripts that have been
cleaved at the poly(A)-site. Rat1 then “chases” RNA Pol II along the downstream cleavage product leading to transcriptional termination in the so
called “torpedo model” (Luo et al., 2006; Proudfoot, 1989; West et al.,
2004).
Decapping enzymes
In eukaryotic cells, there are two types of decapping enzymes, the
Dcp1/Dcp2 complex and DcpS (reviewed in(Liu and Kiledjian, 2006)). The
Dcp1/Dcp2 complex was first discovered in yeast (Beelman et al., 1996;
Dunckley and Parker, 1999). The Dcp1 and Dcp2 proteins from both yeast
and human have been cloned, expressed and purified as recombinant proteins. It was shown that Dcp2 was the catalytically active protein of the
complex (Steiger et al., 2003; van Dijk et al., 2002; Wang et al., 2002c),
whereas Dcp1 enhanced the activity of Dcp2. The amino acids important for
decapping activity of Dcp2 lies within the Nudix-domain (She et al., 2006),
which is the catalytically active domain in the family of Nudix hydrolases to
which Dcp2 belongs (Mildvan et al., 2005). Dcp2 is a divalent metal cation
dependent hydrolase and is most efficient in decapping when Mn2+ is used
(Piccirillo et al., 2003). Dcp2 generates m7GDP and 5’ phosphorylated
mRNAs as products. The 5’ phosphorylated mRNA is then recognized and
degraded by the Xrn1 5’3’ exonuclease. Dcp2 binds RNA in addition to
the cap and is unable to hydrolyze the cap in absence of the RNA body
(Piccirillo et al., 2003). The structure of Dcp2 has been solved and the RNA
is proposed to lie in a channel formed by the C-terminal catalytical Nudix
domain and the N-terminal domain (Deshmukh et al., 2008). The N-terminal
domain (NTD) interacts with Dcp1 as visualized in the structure of the
Dcp1/Dcp2 complex (She et al., 2008). Dcp2 exists in an open and a closed
conformation, with the closed form being catalytically more active. This
structure also explains the activating role of Dcp1: It stabilizes the closed
conformation of Dcp2.
The Scavenger decapping enzyme (DcpS) is the other enzyme with decapping activity involved in mRNA decay. It decaps the short, capped
olionucleotides that results from 3’5’ exonucleolytic digestion by the
exosome in the 3’5 decay pathway. A direct association of DcpS with a
subset of exosomal proteins has also been shown in vivo (Wang and Kiledjian, 2001). DcpS can also cleave the m7GDP product of the Dcp2/Dcp1mediated decapping reaction of the 5’3’ decay pathway. In the former
case the products are m7GMP and an mRNA with a diphosphate in the 5’end. In the later case the products are m7GMP and phosphate. DcpS is unable to decap long substrates. DcpS is a member of the HIT family of pyro18
phosphatases (Liu et al., 2002) and the structure reveals a homodimeric
composition with a domain-swapped amino terminus (Chen et al., 2005; Gu
et al., 2004). When bound to m7GpppG, the dimer turns asymmetric with the
cap bound both to the closed productive site and to the open non-productive
site. A kinetic model of cap-hydrolysis by DcpS based on this structure has
been proposed, where substrate binding and cap hydrolysis are regulated by
negative cooperativity (Liu et al., 2008).
There are several ways that capping can be regulated by both cis-acting
elements and trans-acting factors. An interesting observation was that a 3’terminal oligo(U)-tract added post-transcriptionally can stimulate decapping
(Song and Kiledjian, 2007). More is known about the trans-acting factors
that stimulates decapping. The Lsm1-7 proteins (reviewed in (Khusial et al.,
2005; Wilusz and Wilusz, 2005)) interacts with Dcp1/Dcp2 and together
with the helicase Dhh1 enhances decapping (Fischer and Weis, 2002; Tharun
and Parker, 2001). Furthermore, two RNA-binding proteins, Enhancer of
decapping 1 (Edc1) and Edc2 have been shown to bind RNA and recruit the
decapping complex (Schwartz et al., 2003). PABP, which bind to the 3’ located poly(A)-tail, inhibits decapping (Caponigro and Parker, 1995; Coller et
al., 1998; Wilusz et al., 2001). PABP is proposed to do so in different ways.
First, the interaction with PABP and eIF4G (Tarun and Sachs, 1996), which
in turn binds to eIF4E that interacts with the cap circularizes the transcript
during translation and therefore prevents decapping enzymes from interacting with the cap. Second, it has been shown that PABP can directly interact
with the cap in addition to the poly(A)-tail, inhibiting decapping that way
(Khanna and Kiledjian, 2004).
mRNA surveillance
mRNAs containing errors may be deleterious for the cell if translated to proteins. Therefore different surveillance mechanisms exist that detects and
degrades aberrant transcripts (reviewed in (Garneau et al., 2007)).
Nonsense-mediated decay (NMD)
mRNAs containing premature termination codons (PTCs) would yield truncated proteins if translated. Such mRNAs are therefore degraded by a
mechanism called Nonsense-mediated decay (NMD). There are currently
two models describing how NMD works (reviewed in (Amrani et al., 2006;
Brogna and Wen, 2009; Maquat, 2004)) .
In the “EJC model", NMD is dependent on the Exon-junction complex
(EJC), a protein complex that “marks” Exon-exon boundaries after the splicing process in the nucleus (Zhang et al., 1998). After transport to the cytoplasm, the ribosome in the pioneer round of translation removes the EJC
19
complex from the mRNA (Ishigaki et al., 2001). However, if the ribosome
encounters a PTC, it stalls and the release factors 1 and 3 recruits the Upf1
protein important for NMD. In turn, Upf1 associates with Upf2 and Upf3 in
the EJC thereby forming a closed complex that triggers the degradation of
the PTC-containing mRNA.
In the “faux 3’ UTR model” the distinction between normal and premature termination codons lies in the distance between the PTC and the
poly(A)-tail at the 3’-end of the transcript (Amrani et al., 2004) (Buhler et
al., 2006). A normal stop codon close to the 3’ UTR cause the ribosomal
release factor 3 (eRF3) to interact with PABP bound to the poly(A)-tail
thereby causing normal translation termination. However, if the distance
from the termination codon to the 3’ mRNP is too long (which is the case of
a PTC), no interaction between the eRF3 and PABP can occur. Instead, the
NMD-factor Upf1 interacts with eRF3 thereby triggering NMD.
Both the deadenylation dependent 3’5 and 5’3’ decay pathways are
used in mammalian NMD. PARN coimmunopurifies with the NMD factors
Upf1, Upf2 and Upf3 (Lejeune et al., 2003) thereby implying that PARN
could be involved in NMD. In D. melanogaster it has been shown that NMD
is initiated by an endonucleolytic cleavage (Gatfield and Izaurralde, 2004)
and very recently the endonuclease responsible for this cleavage in human
cells was identified (Eberle et al., 2009). Therefore it is possible that initial
endonucleolytic cleavage is conserved in metazoan NMD.
Non-stop decay (NSD)
Transcripts lacking a stop codon are degraded by a pathway called Non-stop
decay (Frischmeyer et al., 2002). The stalled ribosome at the 3’ end of the
poly(A)-tail is released with help of the Ski7 protein and thereafter the
exosome is recruited and degrades both the poly(A)-tail and the mRNA from
the 3’ end (van Hoof et al., 2002).
No-go-decay (NGD)
When the ribosome is stalled due to for example a secondary structure in the
mRNA it undergoes degradation (Doma and Parker, 2006). In this pathway,
termed No-go-decay, the Dom34 and Hbs proteins bind the transcript close
to the stalled ribosome and initiate an endonucleolytic cleavage. The mRNA
fragments are then rapidly degraded 3’5’ by the exosome and 5’3’ by
the Xrn1 exonuclease.
Nuclear mRNA decay
In the nucleus, aberrant mRNAs are polyadenylated by the Trf4-Air2-Mtr4
(TRAMP) complex (LaCava et al., 2005; Vanacova et al., 2005). The resulting short stretches of oligo(A) makes these mRNAs substrates for the nu20
clear exosome (Vanacova and Stefl, 2007) causing them to be degraded before transported to the cytoplasm. This surveillance mechanism therefore
works in a similar way as mRNA degradation in prokaryotes (see below).
mRNA degradation in prokaryotes
In prokaryotes, poly(A)-tails are found on mRNAs, but the function of the
poly(A)-tail differs from eukaryotes (Sidney, 2004). In many cases, mRNAs
about to be degraded are polyadenylated by PAP and the poly(A)-tail functions as a signal for decay rather than a structure that protects the RNA from
degradation.
In E. coli, most mRNAs are degraded by a multienzyme complex called
the degradosome. It consists of RNaseE, PNPase, the helicase RhlB and
Enolase (reviewd in (Carpousis, 2007)). RNaseE is a homotetrameric endonuclease that performs the initial endonucleolytic attack on the mRNA.
Thereafter, the digested mRNA is polyadenylated and subsequently degraded by PNPase, a phosphorolytic 3’5’ exoribonuclease. The Helicase
RhlB assists in unwinding structured RNA. The function of Enolase is unknown, but it is attached to the unstructured C-terminal part of RNase E as
well as PNPase and RhlB thereby forming the degradosome.
Regulation of mRNA decay
The concept of a messenger ribonucleic particle (mRNP) is important in the
regulation of mRNA decay (Moore, 2005). As discussed earlier, mRNAs
usually do not exist as naked molecules. Rather, they are coated with a plethora of RNA-binding proteins bound to specific sequences (cis-acting elements). Most cis-acting elements that regulate mRNA decay are found in the
3’ untranslated region (3’-UTR) of the mRNA. There, trans-acting factors
like RNA-binding proteins can bind the RNA without disturbance from
translating ribosomes. The general action of these factors is to recruit the
RNA-decay machinery (reviewed in (Garneau et al., 2007; Goldstrohm and
Wickens, 2008; Wilusz et al., 2001)
AU-rich elements
AU-rich elements (AREs) are sequence elements based on the pentamer
AUUUA found in the 3’ UTR of mRNAs (Chen and Shyu, 1995). AREs
function as docking platforms /sequences for ARE-binding proteins that in
turn recruits the RNA degradation machinery. Therefore, AREs function as
destabilizing elements. Some examples of ARE-binding proteins are KH
splicing regulatory protein (KSRP) and RNA helicase associated with AU21
rich element (RHAU) that both bind to and recruit PARN and the exosome
(Gherzi et al., 2004; Tran et al., 2004). Two other examples are AU-rich
binding factor-1 (AUF1) and CUG-binding protein (CUG-BP) that recruit
the exosome and PARN respectively (Chen et al., 2001a; Moraes et al.,
2006) ,further exemplifying that the main decay pathway regulated by AREs
is the 3’5’ pathway. However, evidence for the involvement of the 5’3’
pathway in ARE-mediated decay (AMD) have also been found (Stoecklin et
al., 2006). Some ARE-binding proteins can modulate the activity of mRNA
decay elements without a direct physical interaction. One example is
tristetraprolin (TTP) that can promote deadenylation of ARE-containing
mRNAs by PARN (Lai et al., 2003). An important way for the cell to finetune the regulation of mRNA decay is by phosphorylation of ARE-binding
proteins. Phosphorylation can change the affinity of the protein for its substrate and facilitate an exchange of factors binding to the ARE. For example,
the phosphorylation of KSRP by a kinase makes KSRP to dissociate from
the ARE (Briata et al., 2005). Stabilizing factors such as HuR (Ford et al.,
1999) can then associate with the ARE. Proteins are not the only factors that
interact with AREs. It has been found that a micro RNA (miRNA) harboring
a sequence complementary to the ARE is involved in AMD (Jing et al.,
2005).
miRNAs
Micro RNAs (miRNAs) are ~21 nucleotides long RNAs that have been processed from stem-loop structures in primary transcripts. They form partially
complementary base-pairing to their target mRNAs, thereby downregulating
gene expression. The mechanism by which miRNAs downregulate gene
expression is either by inhibiting translation or accelerating mRNA decay
(reviewed in (Pillai et al., 2007)). It has been debated whether miRNA induced translational repression works through inhibition of translation initiation or at later translational steps. Evidence of the former was found in
(Kiriakidou et al., 2007) where the ability of Ago2 to bind the cap was reported. Ago2 might therefore compete with the translation initiation factor
4E for binding to the cap and therefore inhibit initiation of translation. Evidence of the latter has also been reported where the function of miRNAs
rather is to impede the association of the ribosomal subunits, trigger premature translational termination or promote cotranscriptional degradation of
nascent polypeptides. miRNAs can also trigger mRNA decay as reported in
(Giraldez et al., 2006; Wu et al., 2006) where it was shown that miRNAs
promotes deadenylation. The function by which miRNAs direct rapid deadenylation of mRNAs is by recruiting the Ccr4/Not complex (Behm-Ansmant
et al., 2006).
22
PUF-proteins
AU-rich elements are not the only sequence elements in the 3’-UTR recognized by RNA-binding proteins. The PUF proteins bind to a UGUR tetranucleotide in the 3’ UTR of mRNAs (reviewed in (Wickens et al., 2002)). PUF
proteins are named after the two first identified members of the family,
Pumilio in D. melanogaster (Murata and Wharton, 1995) and FBF (Fem-3
mRNA-Binding Factor) in C. elegans (Zhang et al., 1997). PUF-proteins
regulate a plethora of mRNA targets (Gerber et al., 2006) by accelerating
mRNA-decay. More precisely, PUF proteins enhance deadenylation by recruiting the Ccr4/Pop2/Not-daeadenylase complex through a direct interaction with Pop2, exemplified by studies with the S. cerevisiae PUF-protein
Mpt5 (Goldstrohm et al., 2006). The PUF protein-mediated deadenylation is
then catalyzed by Ccr4 (Goldstrohm et al., 2007).
P-bodies
Processing bodies (P-bodies) are granular cytoplasmic foci where enzymes
and factors involved in mRNA decay have been found (reviewed in (Eulalio
et al., 2007; Parker and Sheth, 2007). All proteins in the 5’3’ mRNA decay pathway have been shown to localize to P-bodies, including the
Ccr4/Pop2/Not-deadenylation complex (Cougot et al., 2004), the Dcp1/Dcp2
decapping complex (Sheth and Parker, 2003; van Dijk et al., 2002) and the
Xrn1 5’3’ exoribonuclease (Sheth and Parker, 2003). There is evidence
that mRNA degradation actively takes place inside P-bodies (Sheth and
Parker, 2003), but another view is that P-bodies are sites where mRNAs are
stored for return to translation (Brengues et al., 2005). Although PARN is
absent in P-bodies, the enzyme has been found in discrete cytoplasmic
granular structures distinct from P-bodies. An ARE-containing mRNA colocalized with PARN and exosome subunits, suggesting that AMD occur in
these cytoplasmic exosome granules.
RNA-binding domains
Proteins use conserved structural RNA-binding domains (RBDs) to interact
with RNA (see (Lunde et al., 2007; Yu Chen, 2005) for review). Often
RNA-binding proteins use multiple copies of the same RBD or a combination of different RBDs to obtain the wanted RNA-binding capability (Lunde
et al., 2007). For RNA-interacting enzymes, the catalytic domain in combination with one or several RBDs defines the targets of the enzymes and
regulates the catalytic activity. I will briefly go thorough some RBDs here.
23
RRM
The RNA rcognition motif (RRM), is the most common of all RNA-binding
domains (Cléry et al., 2008; Maris et al., 2005). ~0.5-1% of all human genes
contain one or several RRMs. The RRM is 80-90 amino acids long and folds
into a compact globular structure consisting of a four stranded anti-parallel
-sheet with two -helixes located behind with topology 112324
(Oubridge et al., 1994) (Figure 2). RNA recognition usually occurs on the
surface of the -sheet. Amino acids involved in RNA-binding reside in the
two central -strands, referred to as RNP1 (in 3-strand) and RNP2 (in 1strand). The consensus sequence for RNP1 is [RK]-G-[FY]-[GA]-[FY][ILV]-X-[FY] and for RNP2 [ILV]-[FY]-[ILV]-X-N-L. The two aromatic
amino acids that are involved in base-stacking of the RNA-bases are shown
in bold and the one involved in interaction with the phosphodiester backbone
of RNA is shown in italic. Not only the -sheet contributes to RNA-binding.
Also the loops connecting the -strands with the -helixes may contain amino acids that interact with RNA. An RRM can bind 2-8 nucleotides, but that
number can be increased by RRM oligomerization. Several examples are
known where two RRMs, located in tandem, will lead to an increased RNA
affinity (Handa et al., 1999; Perez-Canadillas, 2006; Wang and Tanaka Hall,
2001). Interestingly RRMs have also been found to mediate protein-protein
interactions in addition to RNA-protein interactions exemplified by spliceosomal proteins (Corsini et al., 2007; Rideau et al., 2006; Schellenberg et
al., 2006).
24
Figure 2. Structure of the RRM and R3H domains in PARN. (Adapted from the
PBD entries 3D45 and 2A1S respectively).
R3H
In contrast to the RRM, that have been extensively studied, less is known
about the R3H-domain. The three-dimensional structure of the R3H-domain
reveals a three-stranded antiparallel -sheet and two -helices located behind
(Liepinsh et al., 2003) with 1122 3 topology (Figure 2). The name R3H
refers to two highly conserved arginine and histidine residues in the second
-helix with three amino acids located in between. There are over 100 proteins from a diverse range of organisms that contains R3H-domains. Proteins
that contain R3H-motifs can be grouped into 8 families based on similarities
outside the R3H region (Grishin, 1998). The functions of other domains in
proteins containing an R3H-domain indicate that the R3H domain can be
involved in nucleic acid binding. The immunoglobulin μ chain switch region
(Sμ) binding protein 2 (Sμbp-2) contains an R3H-domain. The single
stranded DNA (ssDNA) binding ability of Sμbp-2 has been mapped to a
region harboring the R3H-domain (Fukita et al., 1993). Moreover, the R3Hdomain shows structural similarity to the small DNA-binding MutS-related
(Smr) domain of the Bcl3-binding protein (B3BP) (Diercks et al., 2008)
involved in DNA repair and to the C-terminal domain of translation initiation factor 3 (IF3) (Biou et al., 1995) that binds RNA (Pioletti et al., 2001).
Therefore, the R3H-domain is most likely involved in DNA and RNA binding, but no structure with DNA or RNA bound to R3H is yet solved.
25
Other RNA-binding domains
Zinc-binding domain (Zinc finger domain)
Zinc fingers are classical DNA-binding proteins containing zinc-binding
domains that can also bind to RNA (Lu et al., 2003). They are classified
according to the number of Cysteine and Histidine residues that are involved
in coordinating zinc, e.g. CCHH, CCCH and CCHC-motifs. Zinc-binding
domains are common in proteins that bind AU-rich elements, exemplified by
Tis11d (Hudson et al., 2004). The structure of a Tis11d-RNA complex
shows that the RNA bases points towards the Zinc-binding domains. Sequence-specific RNA-recognition is primarily achieved through hydrogenbonds between the RNA-bases and the main chain of the protein. Zinc finger
proteins that specifically binds to poly(A) has also been identified (Kelly et
al., 2007) although the exact mode of recognition is unknown.
KH-domain
The heterogeneous nuclear (hn)RNP K-homology domain (KH domain) can
bind both single stranded DNA (ssDNA) and ssRNA. The ~70 amino acids
long domain consists of a three-stranded -sheet packed against three helices. The KH domain is characterized by a [I/L/V]-I-G-X-X-G-X-X-[I/L/V]
motif near the centre of the domain (Grishin, 2001; Siomi et al., 1993). The
structure is therefore similar to the RRM, but aromatic amino acids are missing in the conserved motif. Instead of stacking interactions, recognition is
achieved by hydrogen bonding, electrostatic interactions and shape complementarity.
PUF-domain
The PUF-domain is the domain responsible for the RNA-binding property of
the PUF-proteins described earlier. The PUF-domain consists of 8 PUFrepeats with each repeat comprising ~40 amino acids. The PUF repeats, consisting of three -helices each, pack together in a curved, crescent-shaped
structure (Wang et al., 2001). The RNA is bound as an extended strand on
the inside of the structure, with the bases making the contacts to the proteinside chains (Wang et al., 2002a). The RNA is recognized by an alternating
pattern of stacking and base-specific hydrogen-bonds.
26
Present investigations
Introduction
In 1991, a 3'5 poly(A)-specific exoribonucleolytic activity was identified
in HeLa cell nuclear extracts (Astrom et al., 1991). The activity required a 3'
hydroxyl group, released 5'-AMP and left a 3’-hydroxylgroup on the last
adenosine residue of the remaining poly(A)-tail (Astrom et al., 1992). Further studies revealed that monovalent as well as divalent metal ions were
required for its activity. The deadenylase was later purified from calf thymus
and was found to be associated with a 74 kilo Dalton (kDa) polypeptide
(Korner and Wahle, 1997; Martinez et al., 2000). It was named PARN for
poly(A)-specific ribonuclease. PARN is not present in S. cerevisiae and D.
melanogaster, but found in X. laevis (Copeland and Wormington, 2001) and
higher eukaryotes. PARN showed both nuclear and cytoplasmic localization.
The human gene for PARN was cloned and the resulting recombinant protein expressed in E.coli showed specificity for poly(A) (Korner et al., 1998;
Martinez et al., 2000). The 639 amino acids long polypeptide showed sequence homology to bacterial RNaseD and was found to be a member of the
DEDD superfamily of exonucleases (Zuo and Deutscher, 2001). Four conserved acidic amino acids, D28, E30, D292 and D382 in the nuclease domain (Figure 4A) were essential for activity investigated by site-directed
mutagenesis (Ren et al., 2002b). By Fe2+-mediated hydroxyl radical cleavage
and metal ion switch experiments it was concluded that these four residues
coordinate the two catalytically important divalent metal ions in the active
site in a similar way as in the Klenow fragment of DNA pol I (Ren et al.,
2004; Ren et al., 2002b) (Figure 3). Furthermore, aminoglycosides were
found to inhibit PARN-activity with the suggested mechanism of binding in
the active site and thereby displacing the catalytically important divalent
metal ions (Ren et al., 2002a).
PARN is a processive enzyme and has among the ribonucleases the
unique property of interacting with the 5´-end cap structure in addition to the
poly (A)-tail (Dehlin et al., 2000; Gao et al., 2000; Martinez et al., 2000).
The interaction with the cap-structure enhances the rate of hydrolysis and
amplifies the processivity of PARN activity (Martinez et al., 2001). PARN is
a phosphoprotein and it has been suggested that phosphorylation of PARN
improves its binding to the cap (Seal et al., 2005). On the other hand, the
affinity of eIF4E to the cap is decreased upon phosphorylation implying the
27
opposing roles for eIF4E and PARN bound to the cap. PARN activity is
inhibited by the cap binding complex (CBC). CBP 80 inhibited PARN while
the cap-binding part of the complex, CBP 20 had no effect on PARN activity
(Balatsos et al., 2006). The inhibitory effect was mediated by the fact that
CBP 80 could directly interact with PARN. Moreover, PARN activity was
shown to be inhibited by PABP, most likely because the poly(A)-tail is
coated with PABP thereby blocking access for the nuclease (Körner 1997).
In addition to the nuclease domain, two RNA-binding domains have been
identified in PARN (Figure 4A). One is a classical RNA-recognition motif
(RRM) found in many proteins that bind RNA (Cléry et al., 2008; Maris et
al., 2005) and the other is an R3H-domain named by two conserved Arg and
His residues with three amino acids located in between (Grishin, 1998). However, in the R3H domain of PARN the histidine residue is exchanged by a tyrosine. Solution structures of both the RRM and R3H-domains in PARN have
been solved by NMR (PDB: 1WHV and 1UG8). Biophysical studies have suggested that the RRM stabilizes the overall structure of PARN (Zhang et al.,
2007) and that the R3H-domain acts as a protector or an intermolecular chaperone of the RRM (Liu et al., 2007). The cocrystal structure of a C-terminally
truncated version of human PARN, PARN (1-430), consisting of the nucleaseand R3H-domains, in complex with A10, revealed a dimeric composition of
PARN (Wu et al., 2005) (Figure 4B). Three adenosines were visible in the active site, however the structure could not explain how specificity for poly (A)
could be achieved since no base specific interactions were detected in the structure.
28
Figure 3. A schematic drawing for the active site of PARN and the proposed twometal ion catalytic mechanism. The model is based on the catalytic center of the
3’5’ exonuclease activity of E. coli DNA polymerase I (Brautigam and Steitz,
1998) and supported by biochemical (Ren et al., 2004; Ren et al., 2002b) and structural data (Wu et al., 2005).
Aim
The aim of this project was to understand the mechanisms by which PARN
specifically recognizes and degrades poly(A). In other words to figure out
what makes PARN preferring adenosine residues compared to other RNA
nucleotides. The first part covers adenosine specificity in the RNArecognition motif (RRM) (paper I) and the R3H domain (paper V) of PARN.
In the second part, we investigated adenosine specificity during the hydrolytic step in the active site of PARN (paper III) and how the cap and active
sites overlap each other both structurally and functionally (paper II). In addition we have investigated how divalent metal ions modulate poly(A)specificity (paper IV).
29
Figure 4. (A) Domain organization of human PARN showing the nuclease domain
(yellow), the R3H domain (blue) and the RRM domain (red). Residues important for
catalysis (in the nuclease domain) and cap-binding (in the RRM) are labeled. (B)
Structural model of homodimeric PARN containing all three functional domains
colored as in (A). The closed conformation of the nuclease domain and the RRM is
shown on the left and the open conformation is shown on the right. m7GpppG and
A3 ,superimposed in both monomers are shown in stick models with green carbons
for A3 and white carbons for m7GpppG respectively.
30
Results
We have established conditions for expression of recombinant human PARN
and mutants thereof in E. coli (Nilsson and Virtanen, 2006). By a 2-3 step
purification procedure, we were able to generate 90-95% pure PARNpreparations to be used in the biochemical and biophysical assays in the
studies described below.
The RRM of PARN binds both the cap and poly(A) (Paper I)
In this study our aim was to identify amino acids that were involved in
PARN-cap binding. It has previously been shown that several cap-binding
proteins e.g. CBP20 (Calero et al., 2002; Mazza et al., 2002), eIF4E
(Marcotrigiano et al., 1997), the DcpS scavenger decapping enzyme (Gu et
al., 2004) and the vaccinia virus cap modification enzyme VP39 (Hodel et
al., 1998) bind the cap by stacking of the 7-methylguansoine base of the cap
structure between aromatic amino acids. We investigated if this also was the
case for PARN by mixing either of the two cap analogs m7GpppG or m7GTP
and analyzed the mixture by fluorescence spectroscopy. When PARN is
excited with 280 nm light, a light peak at 342 nm is emitted (Figure 2A,
paper I). This is very close to the wavelength of free tryptophan, 340 nm.
Upon addition of increasing amount of free cap analogue, the intensity of the
tryptophan peak decreased, suggesting that either the cap quenches the emitted light from a tryptophan through a direct interaction between the cap and
tryptophans, or that conformational changes takes place in PARN upon cap
binding. From the fluorescence spectra the equilibrium dissociation constant
(Kd) of the PARN-cap interaction could be calculated and it was found to be
in the low micromolar range (Table 1, paper I). To further investigate which
tryptophan that could be involved in cap-binding, we systematically mutated
all six tryptophans in PARN. Mutation of tryptophan 475 to alanine drastically decreased cap binding, and no Kd –value could be obtained, suggesting
that this tryptophan is highly important for cap-binding. Another tryptophan
residue, W456, seemed to have some role in cap binding since we could
observe a 3-7 fold increase in the Kd-value when it was mutated to alanine.
Interestingly, W456 and W475 are situated within the RRM of PARN. To
investigate if the RRM alone is sufficient for cap-binding we cloned a
PARN-fragment, PARN (443-560) comprising the RRM. PARN(443-560)
bound the cap with a slightly higher Kd-value than PARN whereas the double mutant PARN(443-560, W456A, W475A) was unable to bind the cap.
Therefore we concluded that the RRM of PARN binds the cap and W475 is
especially important for cap-binding.
To elucidate if cap-binding is essential for PARN activity, we investigated the ability of PARN and PARN(E455A, W456A, W475A) (which is
severely defective in cap binding, Paper I, table 1) to degrade a capped or
31
non-capped L3(A30) RNA substrate respectively. The introduced mutations
did not affect PARN hydrolysis (Figure 3, paper I), suggesting that cap binding is not essential for PARN activity. Interestingly, recombinant PARN
showed a slightly higher activity on capped versus non-capped substrate,
even though it was not as prominent as with native PARN purified from calf
thymus (Martinez et al., 2001).
We next used electrophoretic mobility shift assays (EMSA) to test
whether the RRM of PARN binds RNA. By using oligo(A) of different
lengths we found that both PARN and the RRM of PARN form complexes
with A20 (Figure 5A and C, paper I), but not with A10 or A5. Moreover, cold
A20 was able to compete a PARN-A20 or PARN(443-560)-A20 complex,
whereas cold A10 competed to some extent and A5 not at all (Figure 4, paper
I). We used a filter-binding assay to measure Kd-values of the PARN-RNA
and the PARN(443-560)-RNA interactions. For A20, Kd-values around 5 nM
were found both for PARN and the RRM of PARN (Table 2, paper I). By
measuring Kd-values for the binding of shorter and shorter oligo(A)s to the
RRM of PARN we observed a drastic increase in Kd for oligos shorter than
10 adenosines (Figure 6, paper I). We conclude that more than 10 adenosine
residues are required for efficient PARN-RRM-Poly(A)-binding.
To test the specificity of the RRM in RNA-binding, we performed competition experiments with the PARN-A20 or PARN(443-560)-A20 complex.
We could show that the complexes were efficiently competed by poly(A), to
some extent by poly(G) and to a very little degree by poly(C) or heteropolymeric single stranded RNA. Therefore, we conclude that the RRM of PARN
binds specifically to poly(A).
Finally, we showed that the cap and RNA-binding sites on the RRM are
structurally and functionally separated from each other. We drew this conclusion primarily based on two observations. (i) The RNA-binding properties of the cap binding deficient PARN(E455A, W456A, W475A) and
PARN(443-560, W456A, W475A) did not differ from the wt counterparts
analyzed by EMSA and filter-binding (Figure 5 and Table 2 paper I). (ii)
The PARN-A20 or PARN-RRM-A20 complex was unaffected by the addition
of cap in excess in EMSAs and the corresponding Kd-values did not differ in
filter binding assays (Figure 7 and Table 2 paper I).
In conclusion, we showed that the RRM of PARN binds both poly(A) and
cap and that W475 is important for the cap-interaction. We further showed
that the cap and RNA-binding sites on the RRM are structurally and functionally separated from each other. These observations suggest that the RRM
is important for proper PARN activity.
32
The R3H-domain of PARN binds RNA (Paper V)
To investigate the RNA-binding properties of the R3H-domain, we cloned a
region in PARN comprising the R3H, PARN(170-245), expressed it recombinantly in E.coli and purified it to homogeneity. By using EMSA, we could
show that the R3H-domain forms a complex with A20 (Figure 1, paper V). To
further investigate the affinity of the R3H domain to A20, we used a Filter
binding assay. We showed that the R3H domain bound to A20 with a somewhat higher affinity (Equilibrium dissociation constant (Kd) 1.8 nM) compared to PARN and the RRM of PARN (Kd-values 5.0 nM and 8.0 nM respectively) (paper I). Next, we investigated the specificity for adenosines in
the R3H-domain of PARN using EMSA wherein the R3H/A20 complex was
competed with different RNA molecules. Figure 1 paper V shows that the
R3H-A20 complex was efficiently competed by poly(A), to some extent by
poly(G) or heteropolymeric RNA and very inefficiently by poly(C). Compared to the RRM (where the complex with A20 could not be competed with
heteroplymeric RNA) (paper I), the R3H-domain shows a more relaxed
specificity for poly(A). Moreover, we found that the R3H-domain bound U20
with a Kd of 4 nM. Competition experiments with the R3H/U20 complex
showed that the R3H harbors poly(U)-specificity (Figure 2 paper V). However, the specificity for poly(U) seems less prominent than for poly(A), indicated by the ability of cold heteroploymeric RNA to compete off the
R3H/U20-complex.
The structure of PARN in complex with Cap (Paper II)
In this paper, we have determined the crystal structure of mouse PARN containing the nuclease, R3H and RRM domains (mPARN 1-510) in complex
with the cap analog, m7GpppG at 3.0Å resolution. As previously reported
(Wu et al., 2005), PARN forms a homodimer with the nuclease domains
forming the dimer interface. The electron density for the R3H-domain was
not good enough for residue-building in our structure. The RRM is in close
contact with the nuclease domain through a hinge region (Figure 1B paper
II). The two subunits adopt different conformations with the orientation of
the RRM differing by 30° (Fig 1C paper II), referred to as open and closed
conformations.
Interestingly, amino acid residues located in the RRM and the nuclease
domains are involved in cap-binding in the closed conformation. In the RRM
the residue W468 (W475 in human PARN) stacks m7G (Figure 2 paper II).
This interaction explains why the PARN-mutant W475A, investigated in
paper I was unable to bind the cap. The other tryptophan that showed reduced affinity to the cap in paper I, W449 (W456 in human PARN) makes
additional contact with the m7G of the cap. In the nuclease domain, two resi33
dues from the active site, isoleucine 34 (I34) and asparagine 281 (N281)
(N288 in human PARN) were when mutated to alanines, defective and severely defective respectively in binding the cap as measured by fluorescence
spectroscopy (Table 3 paper II). In the structure, these residues recognize
atoms in the ribose and the base of the first transcribed guanosine. Furthermore, the first transcribed guanosine is clamped between one methionine and
one leucine, M418 and L284 (M425 and L291 in human PARN), respectively.
Interestingly, when the structure of mouse PARN with bound m7GpppG
in the closed conformation was superimposed on the structure of human
PARN with bound A3 in the active site (Wu et al., 2005), the penultimate
adenosine occupied the same position as the first transcribed G of the cap
(Figure 3A, paper II and Figure 5). Moreover, isothermal calorimetry (ITC)
experiments showed that A10 could be outcompeted by the m7GpppG capstructure and that cap-structure could be outcompeted by A10. Taken together, these data is strong evidence for the presence of overlapped cap and
poly(A) binding sites in the nuclease domain of PARN.
Next, we investigated the catalytic activity of the mutants defective in
cap-binding. We used three different substrates, A3, A20 and uncapped
L3(A30) to probe for activity in the active site, to elucidate the influence of
RNA binding in addition to hydrolytic activity in the active site and to investigate the catalysis of a substrate resembling a bonafide mRNA respectively.
Of the mutants defective in cap binding, at least two of them, I34 and N288
were clearly defective in deadenylating all three types of substrates (Figure 4
paper II). Moreover, mutations of residues N288, L291, S342 and M425
involved in the phosphate linkage and G base recognition showed defects in
the A2 to A1 conversion of an A3 substrate (Figure 4A paper II). Furthermore, we investigated the cap-stimulatory effect observed with native PARN
(Martinez 2001) by using a capped versus a non-capped L3(A30)-substrate.
When using a phosphate-buffered system we were able to reconstitute a 2.5fold cap stimulatory effect for PARN (Figure 4D, paper II). The
PARN(W475A) mutant did not reveal any cap-stimulatory effect, whereas
the PARN(N288A) mutant was slightly affected, as expected due to their
inability to bind the cap.
Finally, we combined all available structures of PARN to create a PARNmodel, where the RRM from one subunit and the R3H-domain from the
other subunit enclose the active site (Figure 6, paper II and Figure 4B). Most
importantly, we conclude from this study that the cap-binding site overlaps
with the active site in the nuclease domain of PARN.
34
Recognition of adenosine residues by the active site of PARN
(Paper III)
In this study, we investigated the molecular mechanisms behind the substrate
specificity of PARN. We started by comparing PARN hydrolysis of the
RNA substrates A20, U20, C20 and G20. We showed that, in keeping with previous results (Martinez et al., 2000) (Korner and Wahle, 1997), A20 was the
most efficient substrate and that U20 was surprisingly well hydrolyzed,
whereas C20 and G20 were poor substrates for PARN (Figure 1 paper III).
When using A20, reaction intermediates corresponding to A4 and A2 accumulated, implying that PARN degrades A20 through three phases, a fast phase
generates A4, a slower phase generates A2, and the last nucleotide is cleaved
with even slower kinetics. In the case of U20, three intermediates consisting
of U5, U4 and U2 were observed. In the case of C20, intermediates consisting
of C5 and C4 could be detected.
The preference for PARN hydrolyzing A20 and U20 could be due to a preference for A and U in PARN RNA-binding. We showed in paper I that the
RRM binds poly(A) and in paper V that the R3H domain binds poly(A) and
poly(U). Therefore it was not surprising that PARN could form a complex
with U20 using EMSA (Figure 2 paper III). However, as in the case of the
R3H-U20 complex, the PARN-U20 interaction seems less specific than the
PARN-A20 interaction, indicated by the ability of cold heteroploymeric RNA
to compete off the PARN/U20-complex. Anyhow, the ability of PARN to
bind A20 and U20 is in agreement with PARN being hydrolytically active on
U20 in addition to A20.
To investigate PARN specificity in the active site, we performed in vitro
RNA-degradation assays using a collection of trinucleotides as substrates.
We chose trinucleotides because in the structure of human PARN (Wu et al.,
2005) three adenosines were observed in the active site. In figure 3, paper III
we showed that PARN indeed harbored specificity in the active site by preferring A3 over G3, U3 and C3. Moreover, we could show that the preference
for adenosines was mainly Km-related. We conclude that PARN harbors
specificity for adenosines in its active site and that improved RNA-binding
to adenosines (in the active site) rather than a higher turnover number is the
reason for this specificity.
To further investigate the mechanisms behind adenosine-specificity in the
active site, we used trinucleotide substrates with different combinations of A
and C. C3 was the slowest hydrolyzed homotrinucleotide compared to A3
(Figure 3 and Table I, paper III), so we expected large kinetic differences
among these substrates. Changing the 5’ located nucleotide to an adenosine
did not improve the hydrolysis, as the kinetic data on ACC did not differ
from CCC (Figure 4 and Table I, paper III). This implies that the 5’-located
nucleotide is not a major factor in providing adenosine specificity in the
active site of PARN. Furthermore, all four substrates with a combination of
35
one adenosine and one cytosine in the penultimate or 3’ position e.g. CCA,
CAC, AAC and ACA were hydrolyzed with increased efficiency (Table 1
and Figure 4, paper III), due to an improved Vmax. Therefore, the penultimate
and 3’-position of a trinucleotide substrate are important for poly(A)specificity in the active site of PARN. As expected, CAA harboring adenosines both in the middle and last position was effectively hydrolysed by
PARN. Interestingly, this improvement in hydrolytic activity was due to an
approximately tenfold decrease in Km (Table 1, paper III). Taken together,
we conclude that the penultimate and 3’-position of a trinucleotide substrate
are important for poly(A)-specificity in the active site of PARN and that the
positional contributions to specificity are additive.
Our structural model of PARN implies that the penultimate adenosine
colocalizes with the first transcribed guanosine of the cap (paper II) that is
clamped by M425 and L291, respectively (Figure 5). We therefore investigated the activity of the mutant polypeptides PARN(M425A) and
PARN(L291A) on trinucleotide substrates. Both PARN(L291A) and
PARN(M425A) hydrolyzed A3 with less efficiency compared to wild-type
PARN (Table 1, paper III and Figure 4, paper II). The mutants hydrolyzed
ACA less efficiently than wt hydrolyzed ACA, but the difference by which
the mutants hydrolyzed ACA compared to AAA was smaller for the mutants
than for the wt. Therefore the specificity for an adenosine in the penultimate
position is reduced in the two mutants, implying that the residues M425 and
L291, at least to some extent are involved in recognizing the penultimate
adenosine.
Figure 5. (A) Superposition of the bound A3 and the m7GpppG cap structure in the
active site of PARN (closed conformation). (B) Residues making contacts with the
bases of cap and A3. Note that the amino acid numbering refers to human PARN.
36
Poly(A) specificity can be modulated when Mn2+ is used as
divalent metal ion (Paper IV)
PARN activity is dependent on divalent metal ions coordinating the amino
acids important for catalysis in the active site (Figure 3). In this work, we
investigated PARN activity in the presence of Mn2+ and compared to Mg2+induced PARN activity. Two interesting observations were made regarding
the specificity of PARN activity in the presence of Mn2+ using A20, U20, C20
and G20 RNA substrates (Figure 1, paper IV). (i) C20 was hydrolyzed much
more efficiently at a rate comparable with A20 and U20 and (ii) the short reaction intermediates that were observed when Mg2+ was used (Figure 1, paper
III) were not as pronounced in the presence of Mn2+. Therefore, PARN loses
specificity in the presence of Mn2+ and degrades the substrates with one reaction phase only. The specificity loss was not due to improved RNAbinding in Mn2+, since EMSA with increasing amount of Mn2+ or Mg2+ did
not alter complex formation (Figure 2, paper IV).
To investigate if the Mn2+-induced specificity-loss is due to altered hydrolytic properties in the active site, we performed RNA-degradation assays
using different homotrinucleotides as substrates in the presence of Mn2+. We
could show that PARN-adenosine specificity is indeed lost using A3, G3, C3
and U3 as substrates (Figure 3, paper IV). G3, U3 and C3 are approximately
10-fold more efficiently hydrolyzed in Mn2+ than in Mg2+ (Table 1, paper IV
and Table 1, paper III). However, the efficiency by which A3 is hydrolyzed
was not changed when the divalent metal cation was changed from Mg2+ to
Mn2+. This result was due to a 25 to 30-fold decrease in both Km and Vmax for
A3 in Mn2+ compared to Mg2+. For G3, U3 and C3, Km was decreased 2300,
40 and 90 times respectively when exchanging Mg2+ for Mn2+. However, the
corresponding change in Vmax was only 240, 4 and 7 times respectively giving rise to the 10-fold increase in catalytic efficiency (Vmax/Km) for G3, U3
and C3 in Mn2+ compared to Mg2+.
We conclude that PARN loses specificity in the active site when Mn2+ is
used as divalent metal ion and that specificity loss is due to changes in both
Km and Vmax. Therefore Mn2+ can be used as a probe to understand the
mechanisms behind poly(A) specificity in the active site of PARN.
37
Discussion
Cap and poly(A) binding
Our structural and functional analysis of PARN cap binding showed that Trp
475 in the RRM stacks the m7G of the cap. The same observation has been
made in two other structures of the PARN-RRM in complex with cap
(Monecke et al., 2008; Nagata et al., 2008). Trp 456 makes additional contacts with the m7G. Therefore, PARN utilizes the 1-1 loop and 2 in the
RRM to recognize the m7G moiety. This is a novel cap-binding mode that
differs from for example CBP 20, where the two tyrosines involved in capbinding are located on the -sheet normally used for RNA-binding (Calero et
al., 2002; Mazza et al., 2002) and Figure 1, paper I. Therefore, the canonical
RNA-binding site on the RRM domain -sheet is still available for poly(A)binding. We showed that the RRM binds poly(A) and that the cap and
poly(A) binding sites on the RRM are separate from each other. On the other
hand, in the nuclease domain, the cap-binding site and the active site overlap
to some extent. That explains why cap added in trans could inhibit PARN
activity (Martinez et al., 2001). Cap provided in cis stimulates PARN activity most likely because the cap can bind in one active site pocket and the
poly(A)-tail in the other active site of dimeric PARN.
Processivity: A comparison with RNase II
Another interesting observation is the three phases through which PARN
degrades an oligonucleotide consisting of 20 adenosine residues (Figure 1,
paper III). The fast phase that results in an intermediate product of four nucleotides can be due to an RNA-binding domain located five nucleotides
away from the active site. In our structural study we could observe that (paper II), the RRM is adjacent to the nuclease domain in the closed conformation, but because no structure containing both RNA and the RNA-binding
domains of PARN has been determined, it is also likely that the R3H-domain
might play this role. As long as an RNA-binding domain binds the substrate
with high affinity, hydrolysis is efficient. When the substrate becomes too
short, the affinity of the RNA to PARN is weaker as described in paper I and
subsequently the further hydrolysis continues with slower kinetics. The 3050 fold decrease in catalytic efficiency observed between long (Capped
L3(A30) and short substrates (A3) (paper III) was due to an increase in Km but
38
did not affect Vmax. The same result was obtained (Liu et al., 2007) with long
poly(A) substrates when the kinetic properties of full-length PARN (p74)
and a truncated version of PARN lacking both the RRM and the R3Hdomain (p54R3H) were compared. This implies that when either no RNAbinding domain is present at all or when the substrate is too short to contact
the RRM/R3H and the active site simultaneously, the only kinetic determinant is the properties of the active site itself.
Interesting parallels can be drawn with RNAse II that degrades olio(A)tails in bacteria (Marujo et al., 2000). RNAse II belongs to the RNB superfamily of exoribonucleases and is, just like PARN, a hydrolytic and processive exoribonuclease (Cannistraro and Kennell, 1994). It shares homology
with Rrp44, the only functional exoribonuclease in the eukaryotic exosome
(Lorentzen et al., 2008). RNase II consists of two N-terminal Cold-shock
domains, one central RNB catalytic domain and one C-terminal S1 domain.
The structure of RNaseII revealed that the RNA contacts the enzyme at two
different regions, at the anchor region made up by the CSDs and the S1 domain, and at the catalytic region within the RNB (Frazao et al., 2006). It has
been shown that a minimum of 10-15 nucleotides are required for enzyme
processivity. The structure indicates that an RNA consisting of 10 nucleotides is the shortest RNA that can contact the active site and the anchor region simultaneously. For shorter RNAs, no such interaction is possible,
hence making the enzyme more distributive. Moreover, in the active site the
last five nucleotides are stacked between two aromatic residues. This gave a
structural explanation to the observed four nucleotides long end-product of
RNA-hydrolysis by RNase II (Amblar et al., 2006).
Therefore, the first four nucleotides long intermediate observed in PARNA20-hydrolysis can be due to an RNA-binding domain located five nucleotides away from the active site. The second intermediate consisting of two
adenosines is more likely the result of properties in the active site itself. The
appearance of a dinucleotide product has previously been observed when the
coordination of divalent metal ions in the active site of PARN was investigated (Ren et al., 2004).
Specificity: The RNA-binding sites and the active site
We investigated the specificity of the RRM and the R3H-domains of PARN
in RNA-binding and found that both domains harbor specificity for poly(A).
Moreover, we showed that the active site itself could discriminate adenosines from other bases. Therefore, hydrolysis of A20 is efficient because A20
binds to PARN with high affinity and additional affinity for adenosines exists in the active site. U20 was surprisingly well hydrolyzed by PARN. That
could be explained by the fact that U20 binds to PARN with high affinity
even though the efficiency by which uridines are hydrolysed in the active
39
site is only 1/10 of the efficiency by which adenosines are hydrolysed. However, the molecular mechanisms of specificity in RNA-binding to the RRM
and R3H-domains are still unknown.
In the active site, it was shown that A3 was recognized by a combination
of hydrogen bonds, hydrophobic interactions and van der Waals contacts
(Wu et al., 2005). F115 is important for recognition of the A in the 3’-end by
a stacking interaction. In paper III, we suggest that M425 and L291 could be
involved in recognizing the penultimate A (Figure 5). An interesting property observed with the M425A-mutant was that an intermediate consisting of
A5 instead of the normal A4-intermediate appears when A20-substrate is being hydrolyzed (Figure 4B, paper II). In RNaseII, a similar phenomenon was
observed when one of the aromatic residues stacking the RNA in the active
site was mutated to an alanine (Barbas et al., 2008). In that case, mutation of
Y253 changed the smallest product of degradation from 4 nucleotides to 10
nucleotides. In PARN, M425 is located adjacent to the RRM and might
therefore, when mutated increase the distance between the nuclease domain
and the RRM by one nucleotide length. The only base-specific interaction
reported in the PARN-A3 structure was S112 that makes a hydrogen bond to
N3 of the 3’ located adenosine (Figure 5). Apart from that no base-specific
interactions was observed neither in that structure nor in our structural model
including the RRM and R3H-domains (paper II). However, in both structures, the C-terminal part of PARN, which has been reported to have a regulatory function (Balatsos et al., 2006), was missing. Therefore, we cannot
rule out that residues from the C-terminal part of PARN could play a pivotal
role in recognizing the penultimate and leaving base, respectively.
Poly(A) recognition in poly(A)-binding proteins
The cytoplasmic PABP (also called PABPC) contains four RRMs, with the
two N-teminally located RRMs (RRM 1 and 2) being sufficient for RNAbinding (Burd et al., 1991; Kuhn and Pieler, 1996). EMSA competition experiments showed that the minimal oligo(A) required for high affinity binding was 11 nucleotides (Deo et al., 1999). In the structure of human PABP
(RRM 1+2) in complex with A11, the two RRMs were arranged in tandem. 8
adenosines were visualized on the RNA recognition surface formed by the
two -sheets (Deo et al., 1999). Adenine recognition was mainly achieved by
van der Vaals contacts, hydrogen bonds and stacking interactions with conserved residues found in the RNP motifs of the RRMs .
The nuclear PABP2 (also called PABPN1) on the other hand, contains
only one RRM. Interestingly PABP2 is a dimer in absence of RNA with the
RRMs forming the dimer interface (Song et al., 2008). The canonical RNA
recognition site on the -sheet surface is occupied by a polyproline motif.
When poly(A) binds, PABP2 undergoes a transition from dimer to monomer
40
that removes the polyproline motif and allows the adenosine nucleotides to
interact with the RRM.
PARN and PARN-RRM binds oligo(A) in the low nanomolar range, just
like PABP (Gorlach et al., 1994) and PABP2 (Wahle et al., 1993). Interestingly, the shortest oligo(A) to bind to PARN-RRM with high affinity was 10
nucleotides (Figure 6, paper I), in accordance with the 11 nucleotides required for tight association with RRM 1+2 of PABP. It is tempting to speculate that the PARN-RRM either can dimerize or together with the R3Hdomain form a strong RNA recognition site upon RNA-binding.
Other DEDD exonucleases: A comparison with PARN
DNA polymerases
All known DNA polymerases share both structural and functional features
(Joyce and Steitz, 1995). Importantly, they have both 5’3’ polymerase and
3’5’ proofreading activities. Both activities are divalent-metal ion dependent. It has been shown that the DNA polymerases induce higher errorfrequencies causing mutations when Mn2+ is used instead of Mg2+ as divalent
metal ion. This fidelity-loss is due to a decrease in Km of the mismatched
deoxynucleotide for the polymerase-template complex and has been reported
for DNA polymerases from different organisms (Goodman et al., 1983)
(Beckman et al., 1985) (Copeland et al., 1993). It therefore seems that the
mutagenic effect of Mn2+ is due to an increase in the binding affinity of the
mismatched deoxynucleotides relative to the correctly paired deoxynucleotides to the polymerase-template complex. Moreover, the proofreading
3’5’ exonucleolytic activity seems to be more “sloppy” when Mn2+ is used
instead of Mg2+ as divalent metal cation (Goodman et al., 1983).
Interesting conclusions can be drawn when comparing the data for PARN
hydrolysis of trinucleotides in Mg2+ vs. Mn2+ (Paper III and IV) with what is
known for DNA polymerases. First, loss of adenosine specificity was observed for PARN when trinucleotide substrates were hydrolyzed in the presence of Mn2+ compared to when Mg2+ was the divalent metal ion source, in
line with loss of fidelity for DNA polymerases. Second, both kinetic parameters Km and Vmax were decreased in Mn2+. That was also the case when diaminopurine was used instead of dATP as incoming nucleotide in the polymerase reaction of T4 DNA polymerase (Goodman et al., 1983). Third, the
specificity-loss of PARN for adenosines in the active site was mostly due to
a decrease in Km for the other homotrinucleotides compared to A3 even if the
Vmax values also decreased. It therefore seems that the ability of DEDDnucleases to be unspecific and “sloppy” with Mn2+ might be evolutionary
conserved.
41
Pop2
Pop2 from Schizosaccaromyces pombe belongs to the same DEDDh superfamily as PARN. The structure of S. pombe Pop2 was recently solved to
1.4Å resolution (Jonstrup et al., 2007). Pop2, in contrast to PARN, exists as
a monomer consisting of only one nuclease-domain. However, the nuclease
domain in PARN is structurally very similar to Pop2 and superposition of A3
from the human PARN-structure (2A1R) (Wu et al., 2005) fits well in the
active site of Pop2 (Jonstrup et al., 2007) Because two Mg2+ ions are visualized in the active site of Pop2 it is highly likely that PARN also utilizes the
general two metal ion catalytic mechanism (Steitz and Steitz, 1993; Yang et
al., 2006) that also have been proposed by Ren et. al (Ren et al., 2004). In
the two metal ion mechanism, one metal ion (MeA2+) activates the attacking
water molecule whereas the second metal ion (MeB2+) coordinates and stabilizes the leaving group (Figure 3). Jonstrup investigated different combinations of Mg2+, Mn2+ and Zn2+ obtaining several interesting results. First, in
Mg2+, Pop2 stops after removing the poly(A)-tail of a generic-poly(A) RNA
substrate, whereas in Mn2+ it continues to degrade the whole substrate. This
loss of specificity is exactly the observation we see with PARN, and it seems
to be associated with one reaction phase only. Second, the addition of Zn2+
together with Mn2+ recovers the poly(A)-specificity of Pop2. The nature of
the divalent metal ions therefore seems to influence the finer architecture of
the active site, hence being important for the catalytic function of the protein. It would be interesting to see if this is the case with PARN as well.
Rrp6
The hydrolytic exoribonuclease found in nuclear exosomes, Rrp6, belongs to
the DEDDy superfamily of exoribonucleases. Rrp6 contains an N-terminal
domain, an exonuclease domain and a HRDC (helicase and RNase D Cterminal) domain. When solving the structure of Rrp6 cocrystallized with
AMP or UMP (Midtgaard et al., 2006), it was discovered that concomitant
binding of both Mn2+ and Zn2+ was required for formation of an enzymeproduct complex. Furthermore, upon substrate-binding a contraction of Rrp6
around the active site was observed. This contraction was due to a rotation
around a hinge-point that brings the HRDC domain closer to the exonuclease-domain. We are observing a similar phenomenon in PARN with closed
and open conformations (paper II). The RRM of PARN located immediately
C-terminal to the exonuclease domain (Figure 4) is attached to the exonuclease domain by a linker region and could play a similar role as the HRDCdomain in Rrp6. Moreover, the HRDC domain in Rrp6 is critical for proper
3’-end processing of stable RNAs (Midtgaard et al., 2006; Phillips and Butler, 2003) and has been implicated in nucleic acid binding. We show in paper
I that the RRM of PARN binds poly(A) and in (Wu et al., 2005) it is shown
that a truncated PARN missing the RRM is a much less efficient enzyme.
42
Finally, Midtgaard et al report a lysine (K342) in Rrp6 that is involved in
specific base-recognition by making hydrogen bonds to N6 and N7 of AMP
and to O4 of UMP. In PARN, that lysine seems to be conserved (K359), but
in the PARN-A3 structure (Wu et al., 2005) it is not found to be involved in
any base-specific interactions.
PARN involved in degrading oligouridylated mRNAs?
Recently, several poly(U)-polymerases have been discovered (Kwak and
Wickens, 2007) that adds short poly(U)-tails to the 3’-ends of histonemRNAs (Mullen and Marzluff, 2008) and polyadenylated actin mRNAs
(Rissland et al., 2007) (reviewed in (Rissland and Norbury, 2008; Wilusz
and Wilusz, 2008)). This polyuridylation resembles the prokaryotic mRNA
decay, in which polyadenylation is a signal for destruction of the mRNA
(reviewed in (Sidney, 2004)). In this thesis, I have shown that PARN degrades poly(U) in addition to poly(A). It is tempting to speculate that PARN
is involved in this novel mechanism of mRNA decay involving poly(U)tails.
Conclusions and future perspectives
In this thesis I have focused on understanding the mechanisms by which
PARN specifically recognizes and degrades poly(A). I have shown that both
the RRM and the R3H domains in PARN are involved in poly(A) binding. In
addition, I have shown that the active site itself harbors specificity for
poly(A) recognizing the last two nucleotides. Furthermore, adenosine specificity in the active site is lost when Mn2+ is used instead of Mg2+ as divalent
metal ion. In the crystal structure of a C-terminally truncated PARN dimer in
complex with m7GpppG I have shown that the cap-binding site overlaps with
the active site in the nuclease domain. In a modelled PARN, I have shown
that the RRM from one subunit and the R3H-domain from the other subunit
enclose the active site.
However, there are always more experiments to be performed before a
more complete view of the mechanisms for PARN poly(A)-binding and hydrolysis can be obtained. For example it would be beneficial to solve a crystal structure with full length PARN and a longer poly(A), interacting with
both the active site and the RRM/R3H-domains simultaneously. Moreover, it
would be interesting to see which amino acids in the RRM and/or R3H that
are involved in RNA-binding and identify the ones that define specificity
towards poly(A). Furthermore, such a structure would provide detailed insight into poly(A)-specificity in the active site and comparisons of structures
43
with Mg2+ vs. Mn2+ in the active site might give us a clue about the mechanisms for specificity-loss. It would also be beneficial to see how an mRNA
molecule with both a cap and a poly(A)-tail present simultaneously binds to
PARN. Finally it would be interesting to investigate PARN “in action”, for
example by a real-time technique.
My work on PARN ends here, but I’m sure that more interesting discoveries will be made on this enzyme in the future.
44
Acknowledgements
•I would like to thank my supervisor Anders Virtanen for accepting me as a
PhD-student and for being a part of a great research-team. You are always
full of ideas and willing to discuss them. You have taught me a lot about
how to interpret results, which conclusions to be drawn and how to present
data in a proper way. I’m truly grateful!
•Many thanks to Haiwei Song, Mousheng Wu and Anna Niedzwiecka for
nice and fruitful collaborations.
•Thank you all fellow members of the Virtanen group: Per, my “brother in
arms” in the PARN-factory. Over the years we have been struggling, sharing
joy and frustration. You are one of the most generous and kind persons I
know. Thank you for all help and good company! Also, thanks for proofreading my thesis. You finally won the pipette-rack-piling-competition, congratulations! Good luck in Japan! Pontus for always helping me with computer problems (no matter how frustrated I am). I admire your capability of
staying focused and working hard in the office sometimes in complete lack
of oxygen… Thanks for nice company on conferences and of course I’ll see
you around on the west coast! Lei for nice organization in the lab and good
company. Good luck with your little kid! You are next in line!
•All the project students over the years: Kyri and Assad for being really
good friends and colleagues. I really like the way you combined hard work
with fun times. Good luck with your PhD:s in Switzerland! Thank you Rickard, Hanna B, Robert, Jens E, Kalle, Felix and Albena. All the best!
•The ”old” people in the Virtanen lab: Thank you Javier for your enthusiasm, commitment and humour. You were one of the reasons why I started as
a PhD-student. Lotta and Helena for helping me in the lab when I was new
and confused. Yanguo for teaching me a lot about PARN. Christina, Nikos
and Jens S for creating a nice scientific atmosphere. Sussie for nice parties
and a great ski-week in Åre.
•The newcomers: Magnus L (well you are not really a newcomer)-Kom igen
Rögle! Harri-Sometimes I wish I could be more of a morning person like
you. Andrea-We should go out for a run soon!
45
•I would like to thank all the PIs at ICM: Måns Ehrenberg-I am deeply
impressed about your passion for science and knowledge in enzyme kinetics.
Leif Kirsebom-Thank you for believing in me and encouraging me. I will
never forget the ”Cumberland sausage ring” in Bath and your obsession with
Mg2+ ions. Gerhart Wagner, Lars Hellman, Nora Ausmees, Sandra
Kleinau, Suparna Sanyal, Santanu Dasgupta, Karin Carlson, Kurt
Nordström, Manfred Kilimann, Johan Elf, Jan Andersson and Diarmaid
Hughes-Thank you for nice times discussing both scientific and nonscientific topics. Maria Selmer-Thanks for valuable help with 3D protein
structure programs. Staffan Svärd-Did you finally beat Glenn Hysén in EMtipset?
I would like to thank all colleagues at ICM:
•Micro: Thanks Pernilla for fruitful scientific discussions and for being such
a nice friend. Linda and Mattias for planning my USA-vacation. Camp
Curry forever! I hope you are settled in San Diego now. Good luck over
there! Cia for nice travelling company in Chicago. Helene for nice (but
sometimes long) conversations about exercise and training. With your determination and fighting-spirit you will also get your PhD soon! Patricia for
chats and laughter, Disa, Marie, Hava, Fredrik P-you will be done just one
week after me! Shiying, Karin, Johan A-cool funk music in the cell lab,
Emma-fun parties, Jon-thank you for enzyme ordering, Johan R-pleasant
lunch conversations, Erik, Klas U, Aurelie, Nadja, Maiike, Amanda, Sonchita, Henrik, Mathias B, Jaydip, Mira, Ema, Tobias, Hanna S, Fredrik
A, Håkan and Janne.
•The Kilimann group: Nicole-my teaching mate, Greta-thanks for nice company in the lunch room watching big sport events, Marcus-thanks for nice
chats at parties, Siv-thanks for all sequencing, Agata, Christoph and Ben
•Molbio: Martin, thanks for nice scientific discussions and good company
in Banff, Canada. I hope everything is fine in Göteborg. Magnus J, Lamine,
Ray-Chelsea will win Champions Leauge this year! Michail-water master,
Doreen, Ikue, David F, Vasili, Kristin and Chandu
•The Elf’s: Brian, Petter, Prune and Mats. Keep the beer-club and the good
research going!
•The Susana Cristobal group: Jia and Itxaso
•The immunology girls: Jenny, Kajsa and Sofia
•The technical staff and administration: Thanks Eva & Ewa for preparing
endless amount of TB-media. Christer for technical support, Solan and
Ulrika for proper lab organization and ordering. Sigrid, Anders, Sofia, Åsa,
Christina F, Cristina M and Akiko for all help.
•The Dicty-group: Fredrik Söderbom, Andrea and Lotta-thanks for nice
company in Chicago.
•And everybody else I somehow forgot to mention!
46
Of course thank you all my great friends:
•Gopal ” the guru” for many fun “boys-night-out-nights”, interesting discussions and nice vacations in London and Paris!
•Stort tack till alla kompisar i OK Linné för proffsigt arrangerade träningar,
tävlingsresor och utlandsläger. Jag stod väl på toppen av min OL-karriär
genom segern i Madrid 2005, sedan har det bara gått utför. Nu när man är
klar med detta arbete kanske det är dags för comeback i skogarna!
•Tack Änglarna 018, Johan, Per och Patrik för trevliga Blåvita stunder
både live och i TV-soffan på ”Casa de la Brokk”.
•Stort tack till mina vänner på Västkusten: Magnus Andersson för sällskap
på långa löp-, skid-, skridsko- och kajakturer, samt gött tjöt! Peter ”Frankan” Andersson och Anders ”Taikon” Vikingsson för en oslagbar USAsemester 2006! Sorry, glömde visst att referera till ”Bruhn, G. & Ferm, W.
(1993) Molekylärbiologin i det 20e århundradet. Acta Östraboica, 23, 1012”
•Tack Johan W för många segeläventyr med ”Flax”. Edvin för grabbkvällar
med Rambo och Zlatan. Ville för all hjälp med mina tre cyklar. Henrik för
trevliga samtal om allt mellan himmel och jord.
Stort tack till min släkt och familj:
•Mårten, Holger, Majken, Peter, Maria, Dick, Eva, Marina och Robin
•Min bror Jonas som trots att vi är ganska olika har hittat på mycket kul
tillsammans genom åren. Pappa som alltid har ställt upp för mig och stöttat
mig i ur och skur. Jag älskar er båda väldigt mycket!
•Till sist skulle jag vilja tacka Kung Bore som med mycket snö och is i
2009 års version gjorde skrivandet av denna avhandling mer dräglig.
47
Swedish summary
Alla organismer är uppbyggda av ett stort antal celler. I våra celler finns en
mängd olika proteiner som är viktiga för uppbyggnaden av cellen och för att
cellen ska fungera och må bra. Våra gener (arvsanlag) fungerar som en mall
för hur cellens proteiner ska se ut. Varje gen utgör ritningen för ett protein
och finns lagrad som DNA. Bildandet av proteiner sker i två steg. 1) Genen
skrivs först av (transkriberas) till budbärar-RNA (mRNA). Informationen om
hur proteinet ska se ut har alltså överförts från DNA till mRNA. 2) Det maskineri (ribosomerna) som i sin tur bygger proteinet använder mRNA:t som
mall. Denna process kallas för translation.
Hur mycket protein som kan bildas i cellen beror på hur mycket mRNA
som finns tillgängligt för translation till protein. Ett sätt för cellen att reglera
mRNA-nivån är att stoppa transkriptionen, ett annat är att bryta ner mRNA:t.
Det första steget i mRNA nedbrytning är att ta bort en s.k. poly(A)-svans i
ena änden (3’ änden) av mRNA:t. Det sker genom att speciella enzymer
(exoribonukleaser) attackerar och bryter ner poly(A)-svansen från 3’ änden.
När väl den skyddande poly(A)-svansen är borta kommer mRNA:t snabbt att
brytas ned av andra exoribonukleaser.
I denna avhandling har jag studerat ett av de enzymer som är specialiserat
på att bryta ner poly(A)-svansen på mRNA: Poly(A)-specifikt ribonukleas
(PARN). Jag har fokuserat på de mekanismer som gör att PARN föredrar att
bryta ner poly(A) till skillnad från övriga RNA-sekvenser. PARN är ett 639
aminosyror stort protein uppbyggt av tre domäner: 1) exonukleasdomänen
där det aktiva sätet sitter, 2) ett RRM som är en klassisk RNA-bindande domän, samt 3) en R3H-domän som också kan binda RNA. PARN är beroende
av divalenta metalljoner för aktivitet. Me2+ koordinerar de negativt laddade
aminosyror i aktiva sätet som deltar i reaktionsmekanismen. Vidare är
PARN en dimer där nukleasdomänen från en subenhet interagerar med nukleasdomänen från den andra subenheten. PARN är unikt i och med att enzymet också känner igen och binder den i 5’-änden lokaliserade cap-strukturen
(m7GpppG) på mRNA:t.
Vi började med att undersöka PARNs förmåga att specifikt binda till
poly(A). Vi fann att PARN föredrar att binda till poly(A) framför poly(G),
poly(C) och ett heteropolymert RNA. Vidare klonade vi RRM, samt R3Hdomänerna i PARN och fann att de var för sig också binder specifikt till
poly(A). En intressant iakttagelse var att affiniteten för korta poly(A)svansar var mycket lägre än för långa. Vidare så bröts långa poly(A)-svansar
48
snabbt ner till en längd av 4 adenosiner, för att sedan brytas ned betydligt
långsammare. En tolkning av dessa resultat var att långa poly(A):n kan binda
både till de RNA-bindande domänerna och till det aktiva sätet samtidigt. I
och med att affiniteten för poly(A) är hög i de RNA-bindande domänerna
hålls en lång poly(A)-sekvens effektivt på plats så att katalysen blir effektiv.
För att undersöka om PARN har specificitet för adenosiner i det aktiva sätet använde vi oss av en repertoar av trinukleotidsubstrat. Vi resonerade att
trinukleotider var för korta substrat för att involvera bindning i de RNAbindande domänerna. Vi visade att PARN bröt ned A3 betydligt effektivare
än G3, U3 och C3 vilket pekar på en förmåga hos PARN att känna igen adenosiner även i det aktiva sätet. Genom att successivt byta ut alla tre adeosiner mot cytosiner visade vi att adenosinen i mitten och den 3’ lokaliserade
adenosinen känns igen av PARN i det aktiva sätet.
Vidare så löste vi kristallstrukturen på PARN i komplex med capstrukturen (m7GpppG). Intressant var att det cap-bindande sätet överlappar
med det aktiva sätet både strukturellt och funktionellt. Genom att jämföra
med en annan PARN-struktur innehållandes A3 i det aktiva sätet såg vi att
den första transkriberade guanosinen i cap-strukturen överlappade med mitten-adenosinen i A3. Genom att mutera de aminosyror som känner igen mitten adeosinen fick vi ett PARN som var mindre poly(A)-specifikt i det aktiva
sätet. Ett annat sätt att få PARN att tappa specificitet för adenosiner i det
aktiva sätet var att använda Mn2+ i stället för Mg2+ i valet av divalent metallkatjon.
Sammanfattningsvis visar denna studie att både RNA-bindande egenskaper i RRM och R3H domänerna samt igenkänning av adenosiner i det aktiva
sätet bidrar till poly(A)-specificiteten hos PARN.
49
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