tRNA biology charges to the front
Eric M. Phizicky1,4and Anita K. Hopper2,3
1Department of Biochemistry and Biophysics, Center for RNA Biology, University of Rochester School of Medicine, Rochester,
New York 14642, USA;2Department of Molecular Genetics, Center for RNA Biology, Ohio State University, Columbus, Ohio
tRNA biology has come of age, revealing an unprecedented
level of understanding and many unexpected discoveries
along the way. This review highlights new findings on the
diverse pathways of tRNA maturation, and on the forma-
tion and function of a number of modifications. Topics of
special focus include the regulation of tRNA biosynthesis,
quality control tRNA turnover mechanisms, widespread
tRNA cleavage pathways activated in response to stress
and other growth conditions, emerging evidence of signal-
ing pathways involving tRNA and cleavage fragments, and
the sophisticated intracellular tRNA trafficking that oc-
curs during and after biosynthesis.
tRNA biogenesis involves the synthesis of the initial
transcript, followed by processing to remove the 59 leader,
trim the 39 trailer, add CCA, splice introns that may be
present, modify multiple nucleoside residues (Fig. 1), and,
for eukaryotes, export the tRNA to the cytoplasm, before
its use in translation (Hopper and Phizicky 2003). Al-
though superficially a simple process, results in the last
several years have highlighted an unexpected complexity
and breadth in tRNA processing and trafficking path-
ways, and have uncovered multiple levels of regulation of
tRNA biosynthesis and function. For the first time, there
is a nearly complete working knowledge of the essential
toolkit of genes required for tRNA biogenesis in the yeast
Saccharomyces cerevisiae (Table 1), and the pace of
discovery in bacteria, archaea, and other eukaryotes is
accelerating rapidly. Exciting stories have unfolded that
describe unexpected layers of regulation in tRNA gene
transcription, new insights in end maturation and splic-
ing, and newly defined roles of tRNA modifications in
translation and tRNA quality control. Intriguing new
results demonstrate the widespread existence of tRNA
cleavage pathways activated by stress and other growth
conditions, the unexpected signaling roles of tRNA frag-
ments and tRNA molecules, and the surprisingly intricate
pathways of tRNA trafficking within the cell. In this
review, we discuss some of the highlights of these findings
in tRNA biology.
Multiple layers of regulation of tRNA transcription
tRNA and rRNA genes are highly transcribed, leading to
the production in yeast of ;3 million tRNAs per gener-
ation and 300,000 ribosomes (Waldron and Lacroute
1975), compared with about 60,000 mRNAs (Ares et al.
1999). Because of the energy devoted to tRNA and rRNA
transcription, and because of the required coordination of
tRNA and ribosome function, tRNA transcription via
RNA polymerase III (Pol III) and rRNA transcription via
Pol I need to be coordinated and regulated in response to
cellular nutrient availability and other environmental
information. The consequences of inappropriate regula-
tion of tRNA transcription have been underscored by the
results of Marshall et al. (2008) showing that elevated
tRNAiMettranscription can promote cell proliferation
and immortalization as well as tumors in mice. Within
the past decade, there has been much progress delineating
mechanisms by which Pol III transcription is regulated
and coordinated with environmental signals (for review,
see Willis and Moir 2007; Ciesla and Boguta 2008).
Pol III is negatively regulated by a single protein, Maf1,
first discovered in yeast by its effects on tRNA-mediated
nonsense suppression (Murawski et al. 1994; Moir et al.
2006). Maf1 is conserved throughout eukaryotes, al-
though mammalian Maf1 negatively regulates Pol I and
Pol II transcription in addition to Pol III transcription
(Pluta et al. 2001; Reina et al. 2006; Johnson et al. 2007).
Yeast and mammalian Maf1 interacts directly with Pol III
subunits (Pluta et al. 2001; Gavin et al. 2006; Oficjalska-
Pham et al. 2006; Reina et al. 2006) and components of
the TFIIIB transcription factor (Upadhya et al. 2002; Desai
et al. 2005; Reina et al. 2006; Rollins et al. 2007; for re-
view, see Ciesla and Boguta 2008).
Substantial evidence suggests that the PKA and TOR
pathways regulate Maf1. Under favorable growth condi-
tions, Maf1 is phosphorylated by both PKA and the TOR-
dependent kinase Sch9 (Huber et al. 2009; J Lee et al.
2009; Wei et al. 2009; for review, see Boguta 2009). Maf1
activity is also regulated by TOR via a Sch9-independent
mechanism (J Lee et al. 2009; Wei and Zheng 2009).
Phosphorylation of Maf1 prevents its negative regulation
of Pol III transcription.
Maf1 is dephosphorylated in response to conditions
that slow growth—nutrient deprivation, shift from fer-
mentation to respiration carbon sources, DNA damage,
and various other environmental stresses (Boisnard et al.
[Keywords: Cleavage; modifications; processing; splicing; tRNA; trafficking;
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Article is online at http://www.genesdev.org/cgi/doi/10.1101/gad.1956510.
1832 GENES & DEVELOPMENT 24:1832–1860 ? 2010 by Cold Spring Harbor Laboratory Press ISSN 0890-9369/10; www.genesdev.org
2009; for review, see Willis and Moir 2007; Ciesla and
Boguta 2008)—and unphosphorylated Maf1 is able to
negatively regulate Pol III transcription. Earlier studies
had implicated Tpd3, a regulatory subunit of the TOR-
dependent protein phosphatase PP2A, in tRNA transcrip-
tion (van Zyl et al. 1992), and it has since been learned
that Maf1 remains phosphorylated in PP2A mutants
(Boisnard et al. 2009), showing that PP2A is a phosphatase
acting on Maf1.
In some yeast strains, phosphorylated Maf1 is located
in the cytoplasm, and is therefore unable to access and
repress Pol III transcription. Maf1’s cytoplasmic location is
mediated by two mechanisms: phosphorylation-dependent
inactivation of the Maf1 nuclear location signals (NLSs)
(Moir et al. 2006), and nuclear export of phosphorylated
Maf1 by the exportin Msn5 (Towpik et al. 2008). Con-
versely, in nutrient-deprived and stress conditions, de-
phosphorylated Maf1 is located in the nucleus, and is
thereby able to access Pol III and down-regulate tRNA
Although the different distribution of Maf1 between
the nucleus and the cytoplasm could, in principle, ac-
count for Maf1’s regulation of Poll III transcription, two
lines of evidence suggest that the nuclear–cytoplasmic
dynamics of Maf1 may instead be a mechanism to fine-
tune regulation. First, Maf1 remains able to appropriately
regulate Pol III transcription in cells lacking Msn5, which
(Towpik et al. 2008). Second, although Maf1 is constitu-
tively located in the nucleus in the common W303 yeast
responsive to environmental signaling (Wei et al. 2009). As
phosphorylation of Maf1 also prevents the interaction of
Maf1 with Pol III (Oficjalska-Pham et al. 2006), it would
appear that this interaction may be the primary level of
negative regulation of Pol III transcription by Maf1.
tRNA end-processing and splicing reveal new insights
in biology and evolution
Surprises in 59 processing
The study of RNase P catalysis has revealed a series of
fascinating discoveries about the nature of catalysis and
the universal requirement for the reaction to occur. The
accepted dogma of tRNA processing has always involved
synthesis of a precursor tRNA containing a 59 leader se-
quence, which was subsequently removed by the endo-
nuclease RNase P (Walker and Engelke 2006). It has also
been accepted dogma since the early 1980s that RNase
P is a ribonucleoprotein (RNP), and that RNA is the
catalytic component in bacteria (Guerrier-Takada et al.
1983; Torres-Larios et al. 2005), which have one protein
component, and in archaea (Pannucci et al. 1999), which
have five protein components (Cho et al. 2010). However,
the source of RNase P catalysis was unknown in eukary-
otes such as yeast or humans, which havenine (Table 1) or
10protein subunits,respectively, inadditionto theirRNA
components (Walker and Engelke 2006), particularly since
their RNA components have substantial differences in
regions important for stability and catalysis (Marquez
et al. 2005, 2006). Nonetheless, results from Kirsebom
and coworkers (Kikovska et al. 2007) have demonstrated
that the RNA component of eukaryotic RNase P of
humans or of Giardia lamblia is, in fact, catalytic, albeit
with verypoorkinetics,likelydue inpart tothe absence in
eukaryotes of helices P15–P17 (Marquez et al. 2005, 2006).
This low level of catalytic activity supports the idea of
a prominent role of one or more of the eukaryotic protein
components in stimulating catalysis by the RNA compo-
nent(Marquez et al. 2006). Indeed, Gopalan and coworkers
(Tsai et al. 2006) have shown with reconstituted archaeal
RNase P holoenzyme that the protein subunits have
a 4000-fold stimulatory effect on catalytic efficiency.
tRNA in S. cerevisiae. tRNA is shown in its usual secondary
structure form, with circles representing nucleotides and lines
representing base pairs. (Green circles) Residues that are unmodi-
fied in all yeast tRNA species; (pink circles) residues that are
modified in some or all tRNA species; (white circles) additional
residues (20a and 20b) that are present in some, but not all, tRNAs
are modified in some tRNAs; (light-blue circles) the CCA end.
Conventional abbreviations are used; see the Modomics data-
base (http://modomics.genesilico.pl). (C) Pseudouridine; (Am)
29-O-methyladenosine; (Cm) 29-O-methylcytidine; (m1G) 1-meth-
ylguanosine; (m2G) 2-methylguanosine; (ac4C) 4-acetylcytidine;
(D) dihydrouridine; (Gm) 29-O-methylguanosine; (m2,2G) N2,
N2-dimethylguanosine; (m3C) 3-methylcytidine; (I) inosine;
(m5C) 5-methylcytidine; (mcm5U) 5-methoxycarbonylmethyl-
29-O-methyluridine; (m1I) 1-methylinosine; (i6A) N6-isopentenyl
adenosine; (yW) wybutosine; (t6A) N6-threonylcarbamoyladeno-
sine; (Um) 29-O-methyluridine; (m7G) 7-methylguanosine; (rT)
ribothymidine; ½Ar(p)? 29-O-ribosyladenosine (phosphate). The pic-
after residue 44, and some tRNAs have different numbers of
residues in the D-loop and the variable arm, but the anticodon is
always numbered residues 34, 35, and 36, and the CCA end is
always numbered residues 74, 75, and 76.
A schematic of modifications found in cytoplasmic
tRNA processing and trafficking dynamics
GENES & DEVELOPMENT1833
Table 1. S. cerevisiae genes implicated in tRNA processing and tRNA trafficking
Yeast gene Modification or functiona
Null mutant phenotypeReferences
Aebi et al. 1990
Bishop et al. 2002;
Xing et al. 2002
Xing et al. 2004
Xing et al. 2004
Xing et al. 2004
Jablonowski et al. 2004;
Huang et al. 2005, 2008
ELP1 (IKI3), ELP2, ELP3, ELP4,
ELP5 (IKI1), ELP6, KTI11
(DPH3), KTI12, KTI13,
KTI14, SIT4, SAP185, SAP190,
Export, re-export Not essential Hopper et al. 1980;
Murthi et al. 2010
Laten et al. 1978;
Dihanich et al. 1987
Eswara et al. 2009;
Murthi et al. 2010
Shaheen and Hopper 2005
Nakai et al. 2004, 2007;
Bjork et al. 2007;
Huang et al. 2008
See Chamberlain et al. 1998
Loss of suppression
NFS1, ISU1, ISU2, CFD1,
NBP35, CIA1, URM1, UBA4,
NCS2, NCS6 (TUC1), TUM1
POP1, POP3, POP4, POP5, POP6,
POP7, POP8, RPP1, RPR2, RPR1
RNase P Essential
C(35), C36, C65, C67
Not essentialSimos et al. 1996;
Motorin et al. 1998
Lecointe et al. 1998
Becker et al. 1997
Ansmant et al. 2001
Behm-Ansmant et al. 2003
Behm-Ansmant et al. 2004
Astrom and Bystrom 1994
Ho et al. 1990;
Trotta et al. 1997
El Yacoubi et al. 2009
Gerber et al. 1998
Gerber and Keller 1999
Johansson and Bystrom 2004
Gu et al. 2003
Culver et al. 1997
Phizicky et al. 1986
Ellis et al. 1986
Hopper et al. 1982;
Nordlund et al. 2000
Cavaille et al. 1999
Motorin and Grosjean 1999
SEN2, SEN15, SEN34, SEN54
m1G37, m1I37, yW37
Cm32, Cm34, Gm34,
Am4, Gm4, Cm4
TRM6 (GCD10), TRM61(GCD14)
Not essential; reading
Bjork et al. 2001
Anderson et al. 1998
Pintard et al. 2002
Alexandrov et al. 2002
Kalhor and Clarke 2003;
Studte et al. 2008
Jackman et al. 2003
Purushothaman et al. 2005
Wilkinson et al. 2007
Kotelawala et al. 2008
Takaku et al. 2003
Kalhor et al. 2005;
Noma et al. 2006
aFor complex modifications, the underlined portion indicates the part of the modification due to the corresponding gene(s).
Phizicky and Hopper
1834GENES & DEVELOPMENT
In light of this evidence that the RNA component of
RNase P is always the catalytic component, it was a
distinct surprise to find that human mitochondrial RNase
P has no RNA components at all (Holzmann et al. 2008).
Although there had been prior reports hinting that RNase
P activity in chloroplasts (Wang et al. 1988) and human
mitochondria (Rossmanith and Karwan 1998) did not re-
quire an RNA component, the chloroplast data had not
been confirmed in >20 years and the human mitochon-
drial data had been challenged (Puranam and Attardi
2001), leaving many convinced that active RNase P in-
variably required RNA. However, Rossmanith and col-
leagues (Holzmann et al. 2008) have shown convincingly
that human mitochondrial RNase P is instead comprised
of only three seemingly unrelated polypeptides: a protein
related to the tRNA m1G9methyltransferase, a member
of the short chain dehydrogenase/reductase complex,
and an uncharacterized ORF. Since active mitochondrial
RNase P was reconstituted after expression and purifica-
tion of these three protein subunits in Escherichia coli, it
is certain that the enzyme does not require RNA.
New results have also identified an exception to the
apparent universal requirement for RNase P activity dur-
ing tRNA biogenesis. Computational analysis suggested
that RNase P RNA was missing from Nanoarchaeum
(Li and Altman 2004). Subsequently, So ¨ll and coworkers
(Randau et al. 2008) provided experimental evidence that
N. equitans has tRNA genes that are transcribed without
a leader sequence, and that are apparently functional with-
out further processing. These tRNAs have triphosphory-
lated 59 ends, and, in several cases, also have an additional
purine base at the ?1 position, and are functional in vitro
RNase P is also now known to have other substrates in
bacteria, yeast, plants, and vertebrate cells (for review, see
Kirsebom 2007). In yeast, genome-wide studies to identify
authentic physiologically relevant RNase P substrates
revealed a role for RNase P in the processing of box C/D
small nucleolar RNAs (snoRNAs) (Coughlin et al. 2008)
and in the processing of the HRA1 noncoding RNA
(ncRNA) (Samanta et al. 2006; Yang and Altman 2007),
whereas, in vertebrates, RNase P has been shown to
process both the MALAT RNA and the Men b transcript
(Wilusz et al. 2008; Sunwoo et al. 2009). It remains to be
P, and whether, and to what extent, there is a biological
connection linking these activities with tRNA processing.
39 Processing and the rise of RNase Z
New data over the last several years have substantially
improved understanding of the process by which the 39
end of tRNAs is processed in different organisms. Matu-
ration of the 39 end of tRNA always requires removal of
the 39 trailer from the original transcript, and also often
requires the subsequent addition of CCA after N73, al-
though, in some bacteria and archaea, some or all tRNA
genes have encoded CCA (Vogel et al. 2005; Hartmann
et al. 2009).Earlier biochemical andin vivo workin E. coli
had established an important role for the endonuclease
RNase E in initial cleavage of most 39 trailers (Li and
Deutscher 2002; Ow and Kushner 2002), and for subse-
quent trimming by 39 exonucleases, primarily by RNase
PH and RNase T, but also by RNase II, PNPase, and
RNase BN (Li and Deutscher 1994, 1996). Similarly, prior
biochemical work in eukaryotes had implicated an endo-
nuclease activity (Castano et al. 1985; Furter et al. 1992)
and an exonuclease activity (Garber and Altman 1979;
Engelke et al. 1985), and fractionation in yeast suggested
more than one source of each activity (Papadimitriou and
Gross 1996). Subsequently, Wolin and colleagues (Yoo
and Wolin 1997) demonstrated that the 39 endonuclease
activity in yeast required the yeast La protein (Lhp1), and
provided evidence that, in the absence of La protein,
a number of tRNAs were processed by an alternative
pathway involving exonuclease activity.
A body of emerging data demonstrates that the endo-
nuclease RNase Z (also called tRNase Z) (Schiffer et al.
2002) plays a major role in removal of the 39 trailer from
tRNAs in several organisms (for review, see Vogel et al.
2005). Most RNase Z enzymes specifically cleave tRNAs
immediately after the discriminator base (N73) to remove
the 39 trailer, prior to the addition of CCA, and available
evidence suggests that RNase Z has this same role in vivo.
Thus, Bacillus subtilis strains depleted for RNase Z
accumulate a large number of tRNAs containing 39 exten-
sions from among the set of tRNAs lacking an encoded
of Drosophila S2 cells to knock down RNase Z expression
results in the accumulation of both nuclear and mitochon-
drial tRNAs that retain their 39 trailers (Dubrovsky et al.
One remarkable biochemical feature of many RNase Z
enzymes is their lack of cleavage of tRNA species with
a mature CCA end (Mohan et al. 1999). Thus, for the B.
subtilis RNase Z, substrates with a CCA sequence at the
beginning of the 39 trailer have a KMthat is 2.5-fold higher
and a Vmax that is 0.4 % of that of substrates with a UAA
sequence at this position, with most of the effect due to
the first C at position N74(Pellegrini et al. 2003). This
dramatic anti-determinant property of the CCA end pre-
vents mature tRNA from being subject to cleavage, and
a resulting futile cycle. This anti-determinant property
also means that other mechanisms must exist for 39 end
formation of tRNAs with encoded CCA sequences in
organisms such as B. subtilis, which have tRNAs both
with and without encoded CCAs. In this organism,
RNase PH and another RNase T-like exonuclease are
implicated in maturation of tRNAs with encoded CCA
sequences (Wen et al. 2005; Redko et al. 2007).
Understanding of the molecular basis for RNase Z
binding and activity and CCA inhibition has been pro-
pelled by extensive structural data (de la Sierra-Gallay
et al. 2005; Ishii et al. 2005, 2007; Kostelecky et al. 2006;
Li de la Sierra-Gallay et al. 2006), as well as by bio-
chemical data. From the cocrystal structure (Li de la
Sierra-Gallay et al. 2006), it is evident that the tRNA
substrate binds one subunit and is cleaved by the other
subunit, that the protein binds tRNA through numerous
tRNA processing and trafficking dynamics
GENES & DEVELOPMENT1835
contacts in the tRNA backbone, and that the D-stem and
anticodon stem are largely devoid of contacts, which is
consistent with biochemical data that these regions are
not important for activity (Nashimoto et al. 1999a). From
modeling and biochemical data, it is likely that the 39
trailer binds through an exit tunnel that precludes base-
pairing (Nashimoto et al. 1999b; de la Sierra-Gallay et al.
2005; Redko et al. 2007), and that the anti-determinant
function of the CCA end of RNase Z is due to the loop
between strands b1 and b2 and to the flexible arm, also
called the exosite (de la Sierra-Gallay et al. 2005; Redko
et al. 2007; Minagawa et al. 2008).
The CCA end does not behave universally as an anti-
determinant (Schiffer et al. 2003; Minagawa et al. 2004).
For example, the RNase Z enzyme from Thermotoga
maritima chooses its cleavage site based on the presence
or absence of a CCA sequence; the 45 tRNAs with an
encoded CCA sequence are cleaved after A76, whereas the
lone tRNA without an encoded CCA sequence is cleaved
at N75(Minagawa et al. 2004).
There are several additional fascinating aspects of
RNase Z function in vivo. While its prominent role in
tRNA 39 end maturation is well documented, it is still
unclear how the gene is linked to increasing the risk of
prostate cancer (Tavtigian et al. 2001). It is also unclear
which other substrates RNase Z has in vivo, since the
enzyme can act in the 59 processing of 5S RNA in vitro in
Halferax volcanii (Holzle et al. 2008) and has been
implicated in the turnover of several mRNAs in vivo in
E. coli (Perwez and Kushner 2006), consistent with the
known biochemical activity of the E. coli RNase Z
enzyme on unstructured RNAs (HS Shibata et al. 2006).
Multiple different substrates might partially explain the
presence of four Arabidopsis RNase Z homologs—includ-
ing one found in both the nucleus and mitochondria, one
found solely in the mitochondria, one found in the
cytoplasm—and an essential homolog found in the chlo-
roplast (Canino et al. 2009).
Splicing endonuclease structure and connection
Several results in the last several years have cast new
light on the components and mechanisms of tRNA
splicing, the medical importance of tRNA splicing, and
other uses and interactions of the tRNA splicing compo-
nents. It remains clear from analysis of multiple se-
quenced genomes that, although introns occur in only a
minority of tRNA genes, they are found in all sequenced
eukaryotes and archaea, and splicing is essential (or
nearly essential) in all of these organisms, based on the
presence of at least one family of tRNA genes in each
organism in which all or almost all of the tRNA genes
have introns (Genomic tRNA Database, http://gtrnadb.
ucsc.edu). While eukaryotic tRNA introns are invariably
found between nucleotides 37 and 38 of tRNAs, and are
most often found at that position in archaea, archaeal
tRNA introns are also found at 14 other positions around
the tRNA molecule (Marck and Grosjean 2003), all of
which feature versions of the bulge–helix–bulge motif
originally identified some years ago, comprised of a 4-base-
pair (bp) helix surrounded on each side by 3-nucleotide (nt)
bulges and a following helix (Thompson and Daniels 1990;
Marck and Grosjean 2003).
Splicing of tRNA is superficially much simpler than
the more common spliceosome-mediated mRNA splic-
ing, because it involves only a limited numberof proteins,
each carrying out a defined reaction (Fig. 2). In all or-
ganisms, tRNA splicing is initiated by an endonuclease
that excises the intron, leaving a 59 tRNA half-molecule
ending in a 29–39 cyclic phosphate, and a 39 tRNA half-
molecule beginning with a 59-OH group (Peebles et al.
1983). In yeast, the endonuclease has four different sub-
units (Table 1), including two related subunits (Sen2
and Sen34) with catalytic activity (Trotta et al. 1997),
whereas, in archaea, the endonuclease is often comprised
of fewer subunits, with a2, a2b2, or a4 configurations
(see Xue et al. 2006).
New structural information has provided a clear image
of a homodimeric endonuclease and its active site. The
structure of the endonuclease in complex with a model
substrate (Xue et al. 2006) demonstrates that all 3 nt of
each bulge of the RNA substrate are flipped out from their
stacking positions. Furthermore, each bulge of the RNA is
in its usual secondary structure, with the antidocodon indicated
by red circles, and the intron after residue 37 indicated by blue
circles. The endonuclease (comprised of Sen2, Sen15, Sen34, and
Sen54 in yeast) excises the intron by cleaving the pre-tRNA
at each exon/intron border, leaving tRNA half-molecules with
a 29–39 cyclic phosphate (indicated by a triangle with a white
circle containing the phosphate) and a 59-OH group at their ends.
In yeast and plants, the ligase (Trl1 in yeast) RNA 59 kinase
activity phosphorylates the 59-OH end of the 39 half-molecule
(black circle), and the ligase cyclic phosphodiesterase activity
opens the 29–39 cyclic phosphate to a 29 phosphate. Then ligase
joins the half-molecules (after activation of the 59 phosphate,
which is not shown), using the 59 phosphate (black circle) as the
junction phosphate, and leaving the 29 phosphate at the splice
junction (white circle). This 29 phosphate is subsequently trans-
ferred to NAD by the 29 phosphotransferase (Tpt1 in yeast). The
yeast-like ligation pathway is also found in vertebrates, but, in
vertebrates and some archaea, the predominant vertebrate ligase
directly joins the phosphate of the 29–39 cyclic phosphate (white
circle) to the 39 half-molecule.
tRNA splicing and ligation pathways. tRNA is shown
Phizicky and Hopper
1836 GENES & DEVELOPMENT
subunit, as well as by R280 and R302 of the other subunit,
as part of a composite active site (Xue et al. 2006). The
composite nature of the active site was also cleverly
documented biochemically with the yeast enzyme (Trotta
et al. 2006). The structure also clearly shows a catalytic
triad, with Y246 poised for attack of the 29-OH, H257
available to protonate the 59-O?leaving group, and K287
stabilizing the negative charge of the transition state,
although none of these residues appears to be absolutely
required for activity (Calvin et al. 2008).
Intriguingly, recent results demonstrate that mutations
in specific subunits of the tRNA splicing endonuclease
subunits are associated with two of the subtypes of the
neurodegenerative disease pontocerebellar hypoplasia. In
many afflicted individuals, the Sen54 endonuclease sub-
unit is mutated in a region that is conserved in mammals
(but not lower eukaryotes). In two other individuals, there
are alterations in either a strictly conserved amino acid of
the Sen2 subunit or a position in the Sen34 subunit that is
the finding that pontocerebellar hypoplasia is associated
with mutations in any of three different endonuclease sub-
units virtually proves a direct connection between the en-
donuclease complex and the disease, it remains to be deter-
mined exactly how these mutations exert their effect on
Partial resolution of the ligation pathway
One of the long-standing unsolved problems in tRNA
splicing has been the nature and identity of the ligase that
catalyzes the second step of tRNA splicing in vertebrates
(Fig. 2). In yeast and plants, ligation requires a cyclic
phosphodiesterase to hydrolyze the 29–39 cyclic phos-
phate of the 59 half-molecule and generate a 29 phosphate,
an RNA kinase to phosphorylate the 59-OH of the 39 half-
molecule, and a ligase to join the half-molecules after
adenylylation of the 59-P end (Greer et al. 1983). A single
yeast protein, Trl1 (Phizicky et al. 1986), which is es-
sential for tRNA splicing in vivo (Phizicky et al. 1992),
catalyzes all four of these activities. Trl1 also catalyzes
the ligation step of the nonconventional tRNA-like
splicing of yeast HAC1 mRNA, which is required for
the unfolded protein response, and is initiated by excision
of the HAC1 intron by the endonuclease Ire1 (Sidrauski
et al. 1996; Sidrauski and Walter 1997). The resulting
ligated RNA from either tRNA splicing or HAC1 mRNA
splicing bears a splice junction 29 phosphate that, for
tRNA, is subsequently transferred to NAD by the 29
phosphotransferase Tpt1 to form ADP-ribose 10–20 cyclic
phosphate (Culver et al. 1993; Spinelli et al. 1997).
In contrast to yeast and plants, vertebrates appear to
have two tRNA ligation pathways. The classical verte-
brate ligation pathway involves a ‘‘vertebrate’’ ligase
activity that directly joins the tRNA half-molecules in
vitro, using the phosphate of the 29–39 cyclic phosphate as
the junction phosphate (Nishikura and De Robertis 1981;
Filipowicz and Shatkin 1983; Laski et al. 1983). In
addition, vertebrates have a second yeast-like ligase/29
phosphotransferase pathway, since both of these activi-
ties have been detected in HeLa extracts (Zillmann et al.
1991, 1992), and the mammalian 29 phosphotransferase is
known to be functional in yeast (Spinelli et al. 1998).
Moreover, mammals have retained the nonconventional
tRNA-like splicing of their HAC1 mRNA ortholog (XBP1)
during the unfolded protein response (Yoshida et al. 2001;
Calfon et al. 2002). Therefore, it has been unclear which
ligation pathway is involved in which reaction.
Some recent results have favored the involvement of
the yeast-like ligation pathway in vertebrate tRNA splic-
ing. Thus, human Clp1 protein was shown to have an
RNA 59 kinase activity, and siRNA experiments in mam-
malian cells implicated Clp1 in tRNA splicing (Weitzer
and Martinez 2007). This finding is consistent with the
yeast-like ligase/29 phosphotransferase pathway, since
this pathway requires an RNA 59 kinase activity. In ad-
dition, prior results had shown that Clp1 copurifies with
the human tRNA splicing endonuclease (Paushkin et al.
2004), although it is known to be involved in cleavage and
polyadenylation of mRNA. Thus, it seemed likely that
splicing in vertebrate cells required the yeast-like ligation
However, it now seems more likely that the yeast-like
ligation pathway is not required in mice for either tRNA
splicing or the unfolded response, since a mouse strain
lacking the 29 phosphotransferase gene is fully functional
for translation of mRNAs requiring decoding by spliced
tRNA, and is normal for the unfolded protein response
(Harding et al. 2008). Thus, it seems likely that the classical
vertebrate ligation pathway is used for tRNA and XBP1
mRNA splicing in vertebrates, similar to the pathway used
for tRNA splicing in archaea (Zofallova et al. 2000; Salia
et al. 2003). The identity of the ligase awaits discovery.
Baroque tRNA gene organization and tRNA processing
An additional surprise is the discovery of two highly
unusual arrangements of tRNA genes that appear to use
tRNA processing enzymes in unexpected ways. In the
first example, So ¨ll and coworkers (Randau et al. 2005b)
have found that N. equitans generates functional tRNAs
by combining separate tRNA halves, which are located in
different parts of the chromosome and are individually
transcribed (Fig. 3). The 59 tRNA half-molecule tran-
scripts terminate in the anticodon loop, and are followed
by an additional sequence of ;12 nt, which pairs exactly
with a corresponding additional sequence at the 59 end
of the 39 tRNA half-molecule transcripts (Randau et al.
2005b). These paired halves are then processed by the
N. equitans splicing endonuclease and, presumably, by
a ligase (Randau et al. 2005a). In the second example,
Sekine and coworkers (Soma et al. 2007) found that the red
alga Cyanidioschyzon merolae had circularly permuted
tRNA genes in which the 59 end of the transcript began in
the D-loop, the anticodon loop, the T-stem, or the T-loop.
The resulting transcripts were subsequently processed to
remove linking sequences, presumably by a combination
of the splicing machinery, RNase P, and RNase Z (Soma
et al. 2007).
tRNA processing and trafficking dynamics
GENES & DEVELOPMENT1837
These unexpected mechanisms for engineering func-
tional RNAs from unusual use of tRNA processing
enzymes are as intriguing as the conserved use of the
tRNA ligation machinery for splicing of HAC1 (XBP1)
mRNA in the unfolded protein response (Sidrauski et al.
1996; Sidrauski and Walter 1997; Yoshida et al. 2001;
Calfon et al. 2002), and the use of RNase P to generate the
39 end of MALAT1 RNA in humans (Wilusz et al. 2008).
Another recent example of the flexibility of tRNA pro-
cessing machinery is the demonstration in yeast that
a designed STE2 mRNA containing an embedded intron-
containing pre-tRNA can be spliced by the tRNA splicing
machinery and RNase P to yield both a functional Ste2
protein and a functional tRNA (Di Segni et al. 2008).
Modification biology takes its place
at the translation table
One of the striking features of tRNA from all organisms is
their large number of post-transcriptional modifications.
A total of 92 different tRNA modifications are listed in
the RNA Modification Database (http://biochem.ncsu.
edu/RNAmods), and a survey of 561 sequenced tRNAs
from several different organisms (Sprinzl and Vassilenko
2005) shows that modifications are found on 11.9% of the
residues, with a median of eight modifications per tRNA
species (Phizicky and Alfonzo 2010). In the yeast S.
cerevisiae, 25 different modifications are found among
the 34 sequenced cytoplasmic tRNA species (Fig. 1).
These modifications occur at 36 different positions, with
an average of 12.6 modifications per species.
The last decade has witnessed great progress in the
identification of the genes and corresponding enzymes
that are required for modification, as well as in our un-
derstanding of their biochemistry and their biological roles.
Thus, in S. cerevisiae, the vast majority of the genes that
are responsible for modifications have now been identified
(Table 1), and a large number of modification genes and
enzymes have been identified in other organisms (see the
Modomics database, http://modomics.genesilico.pl).
Many of the functions of these modifications have
begun to be elucidated, and conform to three general
rules. First, many modifications in or around the antico-
don loop affect translation or growth. For example, yeast
strains lacking I34are inviable (Gerber and Keller 1999);
strains lacking m1G37or t6A are extremely sick (Bjork
et al. 2001; El Yacoubi et al. 2009); strains lacking Nm32
and Nm34, or C38and C39, grow poorly (Lecointe et al.
1998; Pintard et al. 2002); and strains lacking i6A37are
mildly defective for translation (Laten et al. 1978; Dihanich
et al. 1987). Other specific examples of the effects of mo-
difications around the anticodon loop are discussed below.
Second, many modifications in the main body of the tRNA
affect tRNA folding or stability. Thus, the lack of T54
lowers the Tm of tRNA by 2°C–6°C in tRNAPheand
tRNAfMet(Davanloo et al. 1979; Sengupta et al. 2000), the
lack of m1A9in human tRNALysleads to formation of
an alternative structure (Helm et al. 1999), the lack of
29-O-methylated nucleotides can destabilize the 39 endo
form of RNA (Kawai et al. 1992), and the lack of C can
destabilize helices (Durant and Davis 1999; Newby and
Greenbaum 2001). Although the specific biological conse-
quences of the lack of such body tRNA modifications are
not as apparent, emerging evidence elaborated later in this
review suggests that the lack of a number of different body
modifications can elicit tRNA degradation by two different
tRNA turnover pathways. Third, some modifications at
various positions specifically affect tRNA identity. Thus,
G?1 of tRNAHisis a positive determinant for HisRS
(Rudinger et al. 1994; Nameki et al. 1995), and Ar(p) at
position 64 is an identity element for tRNAiMetin yeast
(Astrom and Bystrom 1994), whereas m1G37of tRNAAsp
prevents misacylation by ArgRSin E. coli (Putz et al. 1994),
and lysidine at position 34 prevents misacylation of
tRNAIleby E. coli MetRS (Muramatsu et al. 1988).
The Elp complex is revealed as a modification enzyme
One major highlight has been the discovery that the
central role of the Elp complex in cell function is due to
its activity in modifying tRNA. The Elp complex had
been characterized previously as a complex that copuri-
fies with active elongating hyperphosphorylated Pol II
(Otero et al. 1999), acetylates histones H3 and H4
(Wittschieben et al. 1999; Winkler et al. 2002), and
interacts with Sec2 to regulate exocytosis (Rahl et al.
2005). This complex is comprised of a subcomplex con-
taining Elp1, Elp2, and Elp3, and another subcomplex
containing Elp4, Elp5, and Elp6 (Krogan and Greenblatt
2001; Winkler et al. 2001), and is associated with Kti11
and Kti12 (Fichtner et al. 2002, 2003).
N. equitans, split tRNA genes are transcribed from distant chro-
msomal loci (indicated by filled and open circles) as half-molecules,
each containing additional nucleotides at their 59 or 39 end (blue
circles). The additional sequences at the 39 end of the 59 half-
molecule occur after nucleotide 37, the usual position of an intron,
and pair with the additional 59 sequences of the 39 half-molecule,
forming a hybrid intron. Splicing by the endonuclease, followed
by ligation and CCA addition, results in formation of the mature
Formation of mature tRNA from split tRNA genes. In
Phizicky and Hopper
1838GENES & DEVELOPMENT
Remarkably, available evidence suggests that all of the
phenotypes associated with the lack of the Elp complex
derive from the lack of formation of mcm5s2U at position
34 of certain substrate tRNAs. First, it was found that
all six subunits of the Elp complex, as well as Kti11,
Kti12, and Kti13, were required for formation of mcm5U,
mcm5s2U, and ncm5U at position 34 (Huang et al. 2005).
Then it was shown that the Kluyveromyces lactis g-toxin
acts by cleaving tRNAGlu(UUC), tRNALys(UUU), and
tRNAGln(UUG)after U34(Lu et al. 2005), and that g-toxin
targets these tRNAs because of their mcm5s2U modifi-
cation, since g-toxin resistance is conferred by mutation
of the Elp complex, KTI11, KTI12, or KTI13 (Jablonowski
et al. 2001; Lu et al. 2005). Finally, it was shown that
overproduction of tRNAGln(UUG)and tRNALys(UUU)sup-
presses all of the known Elp complex phenotypes associ-
ated with transcription or exocytosis, and that mutation
of NCS2, which results in the lack of s2U modification of
these same tRNAs, results in the same phenotypes
ascribed to the lack of the Elp complex (Esberg et al.
2006). Thus, it appears that the effects on transcription
and on exocytosis are secondary consequences of the lack
of the mcm5s2U modifications on these tRNAs (Esberg
et al. 2006). It remains to be determined if the require-
ment of the Elp complex for filamentous growth signal-
ing is also directly due to effects on modified tRNAs
(Abdullah and Cullen 2009). It also remains to be deter-
mined why the activity of the Elp complex is defective in
mutants lacking Sit4 phosphatase (critical for cell cycle
progression and modification of Pkc1 function) (Sutton
et al. 1991; Torres et al. 2002), in mutants lacking both of
the Sit4-associated proteins (Sap185 and Sap190), and in
mutants lacking the Kti14 protein kinase (Mehlgarten
and Schaffrath 2003; Jablonowski et al. 2004; Huang et al.
The function of the Elp complex in tRNA modification
and the importance of the mcm5s2U modification are
conserved in metazoans. For example, in Caenorhabditis
elegans, the ELP1 ortholog is also required for formation
of the mcm5moiety of mcm5s2U and ncm5U (Chen et al.
2009), and the TUC1 ortholog (Bjork et al. 2007) is re-
quired for formation of the s2U moiety of mcm5s2U (Chen
et al. 2009). Furthermore, the metazoan Elp complex is
consistently associated with defects in neurological func-
tion. Mutation of the human ELP1 ortholog is associated
with the human neurodegenerative disease familial dys-
autonomia (Anderson et al. 2001; Slaugenhaupt et al.
2001). Similarly, mutation of the C. elegans ELP1 or ELP3
orthologs is associated with a defect in salt chemotaxis
learning, and mutation of both the ELP1 (or ELP3) and
TUC1 orthologs results in developmental defects (Chen
et al. 2009).
The decoding function of the mcm5s2U, mcm5U, and
ncm5U modifications has recently been investigated
in detail by examination of a series of multiple deletion
strains lacking specific tRNA genes and/or the genes
responsible for synthesis of the mcm5or s2moiety
(Johansson et al. 2008). These studies reveal that the
mcm5U modification is important for reading G in the
wobble position (wobble G) for both tRNAs with this
modification ½tRNAArg(UCU)and tRNAGly(UCC)?, based on
growth of the corresponding strain lacking tRNA with
C34. They also reveal that the mcm5s2U modification is
important for reading a wobble A and, in the context of
tRNAGln(UUG), also helps to read a wobble G (but not well
enough to allow growth of a knockout of the correspond-
ing tRNA with C34). Similar experiments show that the
ncm5moiety is important for reading a wobble G residue
for tRNAVal(UAC), tRNASer(UGA), and tRNAThr(UGU), and
that tRNAPro(UGG)can decode codons ending in all four
residues with or without its ncm5U moiety (Johansson
et al. 2008), a clear statement that this property does not
require an unmodified U.
A connection between TRM9 or its mcm5U product
and translation of DNA repair genes
Recent experiments have also revealed a striking trans-
lation role for Trm9 (Begley et al. 2007), which catalyzes
conversion of cm5U to mcm5U at position 34 of substrate
tRNAs (Kalhor and Clarke 2003). trm9-D mutants are
paromomycin-sensitive, implying some translation de-
fect thatis related to ribosome A site function (Kalhor and
Clarke 2003). Expression analysis (Begley et al. 2007) sug-
gests that trm9 mutants are defective in reading arginine
AGA codons as well as glutamate GAA codons, which are
read by tRNAs with the mcm5moiety, as well as arginine
AGG codons, which are read both by tRNA with an
exact match anticodon and tRNA with the mcm5moiety
(Johansson et al. 2008). Since trm9 mutants have reduced
amounts of Rnr1, Rnr3, and Yef3 proteins, each of which
has an overrepresentation of GAA and AGA codons, and
the sensitivity of trm9 mutants to DNA-damaging treat-
ments can be overcome by overexpression of RNR1 or
RNR3, these results suggest that lack of Trm9 and/or its
mcm5U product causes these phenotypes (Begley et al.
2007). Consistent with this explanation, recent results
show that depletion of the human Trm9 homolog ABH8
reduces the amount of mcm5U in tRNA, and leads to
increased sensitivity to DNA-damaging treatments (Fu
et al. 2010). Since Trm9 does not appear to have a major
effect on the growth of strains requiring tRNAArg(UCU)
to read AGG codons, or tRNAGly(UCC)to read GGG
codons, and each of these tRNAs has an mcm5U moiety
(Johansson et al. 2008), the precise effects of Trm9 on
translation may be specific for certain codons or tRNAs.
It is intriguing to note also that silencing of human ABH8
also leads to apoptosis of urothelial carcinoma lines and
down-regulation of NOX-1-dependent reactive oxygen
species, as well as suppression of angiogenesis and in-
vasion (Shimada et al. 2009). Since ABH8 does not have
demethylase activity (Fu et al. 2010), these effects may
also be due to the lack of the mcm5U moiety of tRNAs.
Formation and function of s2U revealed
Another crucial anticodon modification is s2U, which is
universally found at U34of tRNA species that also have
U35 (Bjork et al. 2007). Earlier experiments in E. coli
demonstrated that thiolation requires IscS to transfer
the sulfur from cysteine to form a persulfide, which was
tRNA processing and trafficking dynamics
GENES & DEVELOPMENT1839
transferred to MnmA, to catalyze 2-thiouridylation of
substrate tRNAs (Kambampati and Lauhon 2003). Sub-
sequent E. coli ribonucleome analysis by Suzuki and
coworkers (Ikeuchi et al. 2006) resulted in the identifica-
tion of five other genes involved in this pathway, desig-
nated tusA to tusE, and reconstitution experiments with
purified proteins showed that inclusion of all components
stimulates the biochemical reaction 100-fold. Thiolation
is accomplished by transfer of sulfur from IscS to TusA,
followed by sulfur transfer to TusD of the TusB–TusC–
TusD complex, and likely to TusE, which forms a com-
plex with MnmA and tRNA (Ikeuchi et al. 2006). The
yeast pathway for s2U modification appears to require
several similar components and a similar set of reactions
using Urm1 and Uba4, which also participate in urmyla-
tion of proteins (Nakai et al. 2004, 2007, 2008; Huang
et al. 2008; Schlieker et al. 2008; Leidel et al. 2009; Noma
et al. 2009). A crucial role for the s2U moiety is inferred,
since mutants lacking genes at the end of the pathway
grow poorly, which is consistent with the near universal
conservation of this modification in tRNAs with U34and
U35(Bjork et al. 2007).
The function of s2U is decoding, rather than prevention
of misreading, because overproduction of the unmodified
genes encoding tRNAs with s2U can overcome the
lethality caused by the total lack of mcm5s2U in an elp3
tuc1 double mutant (Bjork et al. 2007). Remarkably,
although tRNALys(UUU), tRNAGln(UUG), and tRNAGlu(UUC)
all have mcm5s2U, only tRNALys(UUU)overexpression is
required to suppress the lethal phenotype of the elp3 tuc1
to aid in the decoding of wobble G codons (Johansson et al.
2008). This is also true in mitochondria, since Tarassov
and coworkers (Kamenski et al. 2007) have shown that
reduced formation of the s2U moiety of cmnm5s2U34of
proteins with AAG codons in a strain unable to import
tRNALys(CUU), providing strong evidence that the long-
known import of tRNALys(CUU)(Martin et al. 1979) is
required at a high temperature for translation of these
Formation and function of wybutosine revealed
At long last, many of the particulars of the mysterious
wybutosine (Wye base, yW) modification have been
elucidated. Wybutosine (or its hydroxy derivative, perox-
ywybutosine) is a complex guanosine base modification
that is found exclusively at base 37 of tRNAPhein most,
but not all, eukaryotes, and other similar wyosine family
derivatives are widely found in archaeal tRNAPhespecies
(Waas et al. 2005; de Crecy-Lagard et al. 2010). Wybuto-
sine is of interest because of its unusual structure (which
contains an additional imidazole ring that is fused to
the guanosine ring, to which is attached the a-amino-
a-carboxy-propyl group of methionine, as well as several
appended groups), and because of reports that reduced
levels of Wye base are associated with tRNAPhein dif-
ferent tumor cells (Grunberger et al. 1975; Kuchino et al.
1982). It has been known for some time that the first step
in Wye base formation is catalyzed by the m1G methyl-
transferase Trm5 (Droogmans and Grosjean 1987; Bjork
et al. 2001). Recent work has shown that subsequent bio-
synthesis of wybutosine involves formation of the methyl
imidazole ring by Tyw1 (Waas et al. 2005; Noma et al.
2006), followed by addition of the a-amino-a-carboxy-
propyl group of methionine by Tyw2 (Kalhor et al. 2005;
Noma et al. 2006), and methylation of guanosine N3 by
Tyw3 (Noma et al. 2006). Intriguingly, the last step of
Wye base formation involves both methylation of the
a-carboxy end group and methoxycarboxylation of the
a-amino end group, with incorporation of carbon dioxide,
all seemingly catalyzed by Tyw4 (Noma et al. 2006;
Suzuki et al. 2009).
An analysis of translation has since shown that one
function of the Wye base is to prevent ?1 frameshifting
(Waas et al. 2007). This study was done using a derivative
of the slippery sequence used by the double-stranded
virus SCV-LA (GGGUUUA) in which the A site codon
was changed from UUA to UUU or UUC. Frameshifting
was most pronounced when the last codon was UUU
rather than UUC, and was progressively reduced as the
Wye base was increasingly formed, since frameshifting
was twofold higher than wild type in a tyw1 mutant,
which has tRNAPhewith m1G37, and 1.5-fold higher than
wild type in a tyw2 mutant, in which the Wye base is in
an intermediate state in which only the imidazole ring
is attached. This finding provides a highly satisfactory
explanation of the value, and perhaps the evolution, of
increased modification in the Wye base, and is consistent
with previous analysis of frameshifting in vitro (Carlson
et al. 2001). These results nicely complement earlier
work by Bjork and colleagues (Urbonavicius et al. 2001,
2003) demonstrating that several modifications in the
anticodon loop of tRNAs in E. coli, Salmonella typhimu-
rium, or S. cerevisiae affect +1 frameshifting, but not ?1
The conserved G?1of tRNAHisand novel activities
of Thg1, the tRNAHisguanylyltransferase
Recent experiments have revealed a number of intrigu-
ing results about the additional G?1residue of tRNAHis.
Decades of previous analysis had shown that this addi-
tional G?1was found uniquely in tRNAHis, that no other
tRNA species (with one exception) had any nucleotide
at this position, and that the additional G?1residue is
conserved by two very different mechanisms: In bacteria
and some archaea, the G?1is encoded (opposite C73) and
retained during processing (Orellana et al. 1986), whereas,
in eukaryotes and other archaea, the G?1residue is added
post-transcriptionally (across from A73and C73, respec-
tively) by tRNAHisguanylyltransferase (Cooley et al.
1982; Jahn and Pande 1991) encoded by the essential
THG1 gene in yeast (Gu et al. 2003). Depletion of Thg1
leads to the accumulation of uncharged tRNAHislacking
its G?1residue, which is consistent with biochemical data
that the G?1residue and its 59 phosphate are important for
HisRS activity (Rudinger et al. 1994; Nameki et al. 1995;
Rosen et al. 2006), as well as to activation of the GCN4
Phizicky and Hopper
1840GENES & DEVELOPMENT
pathway due to the accumulation of uncharged tRNAHis
(Gu et al. 2005). Unexpectedly, however, loss of Thg1 is
also associated with the delayed accumulation of m5C at
C48and C50, and with nuclear localization of a significant
fraction of the tRNAHis(Gu et al. 2005).
Recent results demonstrate four additional surprising
results about the G?1residue of tRNAHisand about Thg1.
First, Thg1 from both yeast and humans is somehow
associated with cell cycle progression. In yeast, thg1
mutants have a cell cycle defect at G2/M, and Thg1
interacts with Orc2 of the origin recognition complex
(Rice et al. 2005), while human Thg1 is cell cycle-
regulated, and its knockdown is associated with a defect
in cell proliferation and with the onset of polynucleate
cells (Guo et al. 2004).
Second, tRNAHisfrom a clade of a-proteobacteria in-
cluding Sinorhizobium meliloti has been found to lack a
G?1residue, and to have a HisRS species with variations
consistent with altered tRNAHisrecognition, thereby pro-
ving that the G?1residue of tRNAHiscannot be univer-
sally required (Wang et al. 2007). It is now known that,
despite its near universal conservation in eukaryotes, the
G?1residue of tRNAHisis also not absolutely required for
growth in eukaryotes, since yeast strains lacking Thg1
can survive, albeit poorly, without the G?1residue, pro-
vided that both tRNAHisand its synthetase are over-
produced (Preston and Phizicky 2010).
Third, it has been shown that Thg1 has several notable
mechanistic similarities to, but little obvious homology
synthetases, recognition of the anticodon is necessary and
sufficient for reaction at the acceptor end, and since the
chemical steps of adenylylation of the 59 phosphate of
tRNA and subsequent guanylyltransfer during G?1addi-
tion are mechanistically similar to formation of the amino-
acyl adenylate and the subsequent tRNA acyltransfer dur-
ing charging by tRNA synthetases (Jackman and Phizicky
Fourth, it has been shown that Thg1 has a distinct 39–59
template-dependent polymerase activity in which it adds
multiple guanine or cytidine nucleotides (or deoxynu-
cleotides) to the 59 ends of appropriate tRNA species, thus
forming multiple phosphodiester bonds in a direction
opposite to that of all known polymerases (Jackman and
Phizicky 2006b). Although the function of this unusual
39–59 template-dependent polymerase activity is not yet
known, it may have a repair function similar to that of the
mechanistically similar, but unidentified, editing activity
that repairs mismatches at the 59 end of tRNA species in
the mitochondria of Acanthamoeba castellani (Price and
Gray 1999). Recent results from Jackman and coworkers
(Abad et al. 2010) showthat archaealThg1 homologshave
a template-dependent uridine and guanidine nucleotide
addition activity across from A73and C73, respectively,
but no G?1addition activity across from A73. This result,
together with the observation that several archaea have
both an encoded G?1residue in their tRNAHisgenes and
a Thg1 homolog, suggests that template-dependent 39–59
nucleotide addition activity is the original function of the
Thg1 family of proteins, and that the known role of Thg1
in the G?1addition across from A73is a newer evolved
function (Abad et al. 2010). Nonetheless, the essential
role of Thg1 in yeast is its tRNAHisG?1addition activity,
although 39–59 template-dependent polymerization is ob-
served in vivo with substrate tRNAHisspecies bearing C73
(Preston and Phizicky 2010).
Unexpected deamination promiscuity in editing
Two other findings suggest expanded roles and functions
of RNA-editing proteins in tRNA function. First, recent
evidence suggests that the Trypanosoma brucei adenosine
deaminase complex ADAT2/ADAT3 can catalyze both
A-to-I editing of the wobble adenosine of tRNAThr(AGU), as
well as C-to-U editing of ssDNA (Rubio et al. 2007). Since
mutation of the active site of ADAT2 eliminates both
A-to-I editing of tRNA and C-to-U editing of DNA, while
not preventing complex formation, it seems highly likely
that the single active site contains both activities. Fur-
thermore, at least in vivo, this same protein appears to be
responsible for both the C-to-U editing that occurs at
residue 32 of tRNAThrand the A-to-I editing that occurs
at residue 34 (Rubio et al. 2006). These findings point to
a common evolutionary origin of both classes of these
editing activities, and suggest the possibility that both
editing activities in tRNA processing derive from one
activity (Rubio et al. 2007). Second, another recent result
demonstrates that tRNAs from Methanopyrus kandleri
have promiscuous editing of their tRNAs at C8, to form
U8at that position, which is crucial for maintenance of
the highly conserved U8:A14reverse Hoogsteen tertiary
base pair (Randau et al. 2009).
tRNA turnover as a quality control mechanism
Emerging results have challenged the widely held notion
that tRNA biosynthesis inevitably leads to product
tRNAs that are virtually infinitely stable. tRNA half-
lives are very long indeed, with estimates of 50 h in
chicken muscle (Nwagwu and Nana 1980), 3 d in avian
liver (Kanerva and Maenpaa 1981), and 44 h in Euglena
gracilis (Karnahl and Wasternack 1992), roughly compa-
rable with the half-life of rRNA (Nwagwu and Nana 1980;
Karnahl and Wasternack 1992). However, experimental
results in the last few years have led to the discovery and
characterization of two pathways by which tRNAs turn
over in the cell as part of quality control mechanisms that
monitor the integrity of tRNA during and after biogenesis
Nuclear surveillance of pre-tRNA and degradation
from the 39 end
Anderson and coworkers (Kadaba et al. 2004, 2006) doc-
umented the existence of a pathway that monitors the
quality of pre-tRNA during tRNA biogenesis. Previous
work had shown that Trm6/Trm61 (Gcd10/Gcd14) was
the methyltransferase that catalyzed formation of m1A58
in tRNA (Anderson et al. 2000), that the temperature
sensitivity of trm6tsmutants could be suppressed by
tRNA processing and trafficking dynamics
GENES & DEVELOPMENT1841
increased amounts of initiator tRNA on high-copy plas-
mids, and that trm6tsmutants had increased turnover of
newly synthesized tRNAiMet, but not elongator tRNAMet
(Anderson et al. 1998). The identification and analysis of
trm6tssuppressors led to the definition of the nuclear
surveillance turnover pathway in which nuclear pre-
tRNAiMetlacking m1A58is subject to polyadenylation
by Trf4 of the TRAMP complex, and then degradation by
Rrp6 and the nuclear exosome (Kadaba et al. 2004, 2006).
Other experiments have shown that overexpression of
the poly(A) polymerase Trf5 can substitute for the lack of
Trf4 in the nuclear surveillance pathway (Kadaba et al.
2006), and that a mutant with a defect in the ATP-
dependent RNA helicase activity of Mtr4 prevents deg-
radation of pre-tRNAiMet, but does not perturb its forma-
tion of the TRAMP complex or polyadenylation of the
pre-tRNA (Wang et al. 2008).
Two lines of evidence show that the 39–59 exonuclease
Rex1 plays a significant role in the nuclear surveillance
pathway. First, rex1trm6mutants have a synthetic growth
phenotypethat is associatedwith polyadenylationofthose
pre-tRNAiMetspecies and pre-tRNAVal(CAC)species that
have longer 39 trailers, and of dimeric tRNAArg–tRNAAsp
transcripts (Ozanick et al. 2009). Second, Wolin and co-
workers (Copela et al. 2008) have shown that unspliced
pre-tRNA species are subject to the nuclear surveillance
pathway, but not in rex1 mutants, suggesting that Rex1
function may generate these pre-tRNA degradation sub-
strates, and further experiments suggest that Rex1 is in
competition with La (Lhp1) protein for these 39 ends.
The full extent of the involvement of La protein in the
nuclear surveillance and/or other tRNA turnover path-
ways is not yet known, but it likely has some role, since
overexpression of La protein can also suppress trm6ts
mutants (Anderson et al. 1998). La protein acts by binding
the UUU-OH sequence at the end or pre-tRNA tran-
scripts (see Maraia and Bayfield 2006; Teplova et al. 2006
and references therein) through the use of two sites
(Bayfield and Maraia 2009), has an RNA chaperone ac-
tivity (Chakshusmathi et al. 2003), and appears to protect
RNA from Rrp6 of the nuclear surveillance pathway
in Schizosaccharomyces pombe (Huang et al. 2006).
However, La also protects RNA independently of Rrp6,
suggesting that it participates in another turnover path-
way (Huang et al. 2006).
Detailed and elegant biochemical experiments support
the claim that the TRAMP complex collaborates with the
nuclear exosome to polyadenylate substrate tRNAs and
et al. 2005), and show that this activity acts on a pre-tRNA
transcript (LaCava et al. 2005); on an unmodified tRNAiMet
transcript, but not its fully modified derivative (Vanacova
et al. 2005); on a tRNAAlavariant with a structural defect,
but not the unmodified wild-type tRNAAla(Vanacova et al.
2005); and on tRNAiMetlacking only m1A58, but not other
tRNA substrates (Schneider et al. 2007).
Rapid tRNA decay (RTD) of mature tRNA
from the 59 end
Another set of experiments has shown that mature
tRNAs are also subject to turnover. Thus, trm8 trm4
mutants, which lack m7G and m5C in their tRNAs, are
temperature-sensitive due to the specific degradation of
tRNAVal(AAC)by an RTD pathway that is distinct from the
nuclear surveillance pathway that acts on pre-tRNAiMet
lacking m1A58(Alexandrov et al. 2006). This RTD path-
way appears to be general, since several different tRNA
species lacking different combinations of modifications
are subject to degradation by this pathway (Chernyakov
et al. 2008). Moreover, degradation occurs at the level of
mature tRNA, rather than pre-tRNA, since cells treated
with the transcription inhibitor thiolutin undergo degra-
dation at the same rate and degrade all of the tRNA of
substrate species (Chernyakov et al. 2008). Genetic anal-
ysis suggests that tRNA degradation by the RTD pathway
ways in yeast. pre-tRNA transcribed in the nu-
cleus is processed (black arrows) in the nucleus
and the cytoplasm (steps 1) to remove the 59 leader
and 39 trailer (purple circles), to add CCA to the 39
end (blue circles), to remove the intron if present
(not shown), and to add modifications ½pink cir-
cles, as for tRNAVal(AAC)?, ultimately emerging in
the cytoplasm for translation (step 2). If m1A58is
not added to pre-tRNAiMet(absence of m1A in-
dicated by black circle), this pre-tRNA is degraded
by the nucelar surveillance pathway (step 3, red
arrow) in which the pre-tRNA is first polyadeny-
lated by the TRAMP complex, and then degraded
from the 39 end by the nuclear exosome. If m7G46
and m5C49are not added to tRNAVal(AAC)(black
circles), the hypomodified mature tRNA is at least
partially functional, but is degraded by the RTD
pathway (red arrows), by Xrn1 in the cytoplasm
(step 4), or by Rat1 in the nucleus (step 6), possibly
after nuclear import (step 5). (Step 7) The elevated
presence of pAp in met22 mutants inhibits the
RTD pathway by inhibiting both Xrn1 and Rat1.
Two different tRNA degradation path-
Phizicky and Hopper
1842GENES & DEVELOPMENT
is catalyzed by the 59–39 exonucleases Rat1 and Xrn1 and
requires the methionine biosynthetic enzyme Met22
(Chernyakov et al. 2008), likely because its substrate,
pAp, accumulates in met22 mutants (Murguia et al. 1996)
and inhibits Rat1 and Xrn1 (Dichtl et al. 1997). Intrigu-
ingly, the substrate for the RTD pathway may be amino-
acylated tRNA, since tRNA degradation is accompanied
by selective loss of the aminoacylated fraction of the
tRNAVal(AAC)population (Alexandrov et al. 2006), and
since mutations in MET22 or RAT1 and XRN1 that pre-
vent degradation also restore the fraction of charged
tRNA (Chernyakov et al. 2008). It is not yet known if
and to what extent any part of the translation machinery
participates in this pathway; why specific hypomodified
tRNA substrates are selected for degradation by this
pathway, whereas other tRNAs lacking the same modi-
fications are spared; and the extent to which this pathway
acts on other types of defective tRNAs.
tRNA cleavage pathways activated by stress and other
growth conditions, and signaling by tRNA fragments
A series of exciting studies in the last few years has de-
tailedthe existence of a previously unknown pathway that
occurs in a variety of organisms under specific growth
conditions, and generates tRNA cleavage in the region of
the anticodon loop (for review, see Thompson and Parker
2009b). Thus, Collins and coworkers (Lee and Collins
2005) have shown that starvation of Tetrahymena ther-
mophila induces cleavage of a large number of different
tRNAs that derive from the macronucleus, but not the
mitochondria. tRNA cleavage occurs at variable positions
in and around the anticodon loop and occasionally in the
variable arm, appears to target tRNAs that are modified
but lack their CCA end, is quantitatively minor, and ap-
pears to require translation (Lee and Collins 2005). Exper-
iments in yeast demonstrate that similar cleavage of mul-
tiple tRNA species occurs in cells undergoing oxidative
stress, methionine starvation, extended growth in station-
ary phase, and growth at high temperature, but not in cells
undergoing ultraviolet (UV) stress, nitrogen starvation, or
glucose starvation (Thompson et al. 2008). Furthermore,
similar cleavage is observed in Arabidopsis thaliana and
human cells undergoing oxidative stress (Thompson et al.
2008; Fu et al. 2009; Yamasaki et al. 2009), as well as
Streptomyces coelicolor (Haiser et al. 2008) and several
other organisms, generally under starvation conditions or
other conditions affecting developmental change (see
Thompson and Parker 2009b and references therein). Sub-
sequent experiments show that this pathway is mediated
by the RNase T2 family member Rny1, which relocalizes
from the vacuole to the cytoplasm during oxidative stress
and mediates cell death (Thompson and Parker 2009a).
However, the activity associated with Rny1 that mediates
cell death is demonstrably separate from its cleavage ac-
tivity (Thompson and Parker 2009a). Intriguingly, the hu-
man Rny1 ortholog RNASET2 has a set of activities sim-
ilar to those of yeast Rny1, since expression of RNASET2
in yeast restores the production of tRNA cleavage frag-
ments in an rny1 mutant strain and restores the reduced
viability of cells undergoing oxidative stress (Thompson
and Parker 2009a), while expression of RNASET2 in mam-
malian cells acts as a tumor suppressor independent of its
cleavage activity (Acquati et al. 2005; Smirnoff et al. 2006).
tRNA cleavage products are also generated by Dicer-
dependent cleavage. Deep sequencing has revealed the
existence of a prominent Dicer-generated tRNAIlefrag-
ment in mouse embryonic stem (ES) cells that derives
from the 39 end of the gene and part of the trailer se-
quence, and may arise because of alternative folding of
the tRNA transcript (Babiarz et al. 2008). In addition, deep
sequencing has identified a number of Dicer-generated
59 tRNA fragments in HeLa cells that tend to arise from
cleavage after residue 19 in the D-loop, and derive mostly
from four major tRNA species (Cole et al. 2009). Since
these RNA fragments appear to be modified at the 39 end
and bind Argonaute complexes weakly, they are not
likely to act as microRNAs (miRNAs); thus, their func-
tion remains unknown.
Possible signaling effects of tRNA fragments
Two recent reports have emerged suggesting that tRNA
cleavage products inhibit translation. First, Anderson and
coworkers (Yamasaki et al. 2009) have shown that each
of several different stress treatments of mammalian cells
activates the ribonuclease angiogenin to cleave tRNA to
produce tRNA-derived stress-induced RNAs (tiRNAs)
that inhibit translation. Angiogenin is a potent angio-
genic factor isolated from the medium of human cell lines
or from plasma (Shapiro et al. 1987), with ribonuclease
activity that is essential for its angiogenic activity (Shapiro
and Vallee 1987). Remarkably, the 59 tiRNAs produced by
angiogenin have a distinct inhibitory effect on cell growth,
since transfection of the 59 tiRNA fragments, but not the
39 tiRNA fragments, induces translation arrest indepen-
dent of phosphorylation of eIF2a (Yamasaki et al. 2009)
through formation of stress granules (Emara et al. 2010).
Second, pumpkin phloem sap has been shown to contain
tRNA fragments of several tRNAs, which are cleaved in
either the anticodon loop or the D-loop, and which appear
to inhibit translation based on reduced translation in the
presence of phloem sap RNA, or of RNase A-generated
fragments of yeast tRNA (Zhang et al. 2009). It remains to
be determined if translation is similarly inhibited in other
systems in which tRNA fragments are produced.
In another development, Dutta and colleagues (YS Lee
et al. 2009) have found by deep sequencing methods that
multiple different tRNA-derived fragments (tRFs) are
made in prostate cancer cell lines, at least one of which
has potent effects on promoting cell proliferation. These
tRFs are 13–26 nt long, comprise fully 40% of the non-
miRNA sequences in this size range, and derive from one
of three precise regions of the tRNA: the 59 end of mature
tRNA, the 39 end of the mature tRNA ending in the CCA
sequence, or the 39 trailer sequence beginning immedi-
ately after the 39 end of the tRNA. One of these tRF
species, tRF-1001, derives from the 39 trailer sequence of
a specific tRNASer(UGA)species and has profound effects
on cell growth. tRF-1001 is highly expressed in a number
tRNA processing and trafficking dynamics
GENES & DEVELOPMENT1843
of different proliferating cancer cell lines, and its expres-
sion is reduced after serum starvation or when cell
density is high. Furthermore, siRNA-mediated knock-
down of tRF-1001 expression is associated with decreased
cell proliferation, and cell proliferation is restored by trans-
fection of a synthetic 29-O-methylated tRF-1001 RNA
oligonucleotide that does not interfere with the siRNA
treatment. Further experiments show that tRF-1001 is
formed by the action of ELAC2, which encodes RNase Z
and was originally identified as a candidate prostate cancer
(and its pre-tRNA) is found almost exclusively in the
cytoplasm, leading to the suggestion that pre-tRNAs can
be processed either into tRNA in the nucleus, or in the
cytoplasm to form tRF-1001 and related species (YS Lee
et al. 2009). Since tRF-1001 does not affect expression of a
reporter that should be sensitive to siRNA or miRNA, the
to be determined. Similarly, it remains to be determined if
and to what extent other tRFs regulate cellular functions,
and how these tRFs are generated.
Distinct tRNA cleavage pathways as part of host
These widespread tRNA cleavage pathways are distinct
from a variety of tRNA cleaving toxins that act on
specific sets of tRNAs and at specific positions within
the tRNA anticodon loop to disable substrate tRNAs.
The most well known of these is the PrrC endonuclease,
which attacks tRNALysspecies during infection by phage
T4 (Amitsur et al. 1987), but several others have been
described, including the colicins and onconase (for re-
view, see Phizicky 2008). A notable recent addition to this
family is the g-toxin of K. lactis that targets specific
tRNAs in the yeast S. cerevisiae that have the mcm5s2U
modification at position 34 and cleave the tRNA by
formation of a 29–39 cyclic phosphate (Lu et al. 2005).
Since Shuman and coworkers (Nandakumar et al. 2008)
have shown that this cleavage pathway can be repaired by
a plant RNA ligase domain together with the yeast splicing
cyclic phosphodiesterase and kinase activities, it seems
plausible that any of the cleavage pathways described
above are also subject to RNA repair. Intriguingly, CCA
addition by nucleotidyl transferase may also act as part of
a quality control mechanism, since tRNAs that are nicked
are not efficient substrates for the enzyme (Dupasquier
et al. 2008). It is not clear what happens to such substrates
in vivo, but they could, in principle, also be repaired.
Nonconventional uses of tRNA and other signaling
pathways using tRNA
Although the conventional function of tRNAs is to de-
liver amino acids to the translation machinery as speci-
fied by mRNA codons, it has been well established in
both prokaryotic and eukaryotic cells that tRNAs serve
numerous other functions. In prokaryotes, aminoacylated
tRNAs serve as donors to deliver amino acids used for
diverse biochemical pathways, including peptidoglycan
biosynthesis, additions to lipids, production of certain anti-
biotics, and targeting of proteins for degradation via the
N-end rule pathway (for review, see Banerjee et al. 2010;
Francklyn and Minajigi 2010). Moreover, as prokaryotic
organisms lack a complete set of aminoacyl-tRNA synthe-
tases to generate all required aminoacylated tRNAs, mech-
anisms using noncognate aminoacyl-tRNAs function to
generate the full set of cognate aminoacyl-tRNAs. For
example, Glu-tRNAGlnand Asp-tRNAAsnare modified to
(for review, see Feng et al. 2004). Finally, in Gram-positive
bacteria, tRNAs also function as sensors to regulate gene
expression in response to nutrient availability, as demon-
strated clearly by the T-box riboswitch mechanism; this
tRNA-dependent process employs uncharged tRNAs,
which interact with mRNA leader sequences to generate
anti-termination elements and thereby allow the full tran-
scription of gene products involved in amino acid bio-
synthesis (for review, see Green et al. 2010).
tRNAs also serve nonconventional functions in eu-
karyotes. Most of these functions differ from nonconven-
tional functions for tRNAs in prokaryotes, but at least
two roles are similar among prokaryotes and eukaryotes.
The first isthe useof aminoacylated tRNAs as amino acid
donors for N-terminal conjugation of amino acids to
proteins, targeting the recipient proteins for degradation
(for review, see Varshavsky 1997; Mogk et al. 2007). The
second concerns the role of uncharged tRNAs in signal
transduction pathways responding to nutrient depriva-
tion. In yeast, the general amino acid response pathway
responds to amino acid deprivation using uncharged
tRNAs that interact with Gcn2, the protein kinase that
phosphorylates translation initiation factor eIF2. Phos-
phorylation of eIF2 by Gcn2 results in decreased levels
of general translation, but increased translation of the
transcription regulator Gcn4, which in turn results in
transcription of numerous genes involved in amino acid
and nucleotide biosynthesis (for review, see Dever and
Hinnebusch 2005). In mice, a similar Gcn2-dependent
process occurs in which binding of uncharged tRNAs to
Gcn2 results in an altered eating response (Hao et al.
2005; Maurin et al. 2005; for review, see Dever and
Hinnebusch 2005). Surprisingly, the Gcn4 pathway is
also turned on by DNA-damaging treatments in yeast
that act through Mec1 and Rad53 signaling to relocalize
Los1 to the cytoplasm, resulting in the accumulation of
unspliced tRNA in the nucleus (Ghavidel et al. 2007). It is
unknown which tRNA species activates this pathway,
but Gcn4 is also activated in los1-D cells independently of
Gcn2 (Qiu et al. 2000), so this DNA damage response
pathway likely occurs by the same pathway.
There are also nonconventional roles for tRNAs that
are not shared between eukaryotes and prokaryotes. For
example, tRNAs have been implicated recently in regu-
lation of apoptosis in mammalian cells (Mei et al. 2010).
These studies showed that tRNAs bind cytochrome c,
thereby preventing the interaction of cytochrome c with
the caspase activator Apaf-1 and preventing its activa-
tion. The results provide new ways to think about how
Phizicky and Hopper
1844GENES & DEVELOPMENT
tRNA cellular levels influence cell growth and oncogen-
esis. Another eukaryotic-specific nonconventional func-
tion for tRNAs is the employment of tRNAs in the
retroviral life cycles. Retroviruses have usurped tRNAs
to serve as primers for reverse transcription of their RNA
genomes(for review, see Marquet etal. 1995),and for HIV-
1 minus strand transfer (Piekna-Przybylska et al. 2010
and references therein). HIV has also usurped the retro-
grade tRNA pathway (see below) as one mechanism to
deliver the reverse-transcribed complex from the cyto-
plasm to the nucleus (Zaitseva et al. 2006).
The long and winding cellular road for tRNA biogenesis
The highly unexpected set of new mechanisms of regu-
lation of tRNA synthesis, new insights into modifica-
tions, new pathways of tRNA processing and turnover,
and new tRNA cleavage pathways is matched by equally
unexpected recent results describing the long and wind-
ing road by which tRNA biogenesis occurs. Indeed, as
described further below, the widespread cellular distribu-
tion of gene products functioning in transcription, pre-
cursor processing, and nuclear export events for tRNAs
differs substantially from the much more localized dis-
tribution of gene products functioning in analogous
events for biogenesis of other RNAs. For example, for
mRNA production, it appears that most of the biochem-
ical machinery is recruited to the site of transcription; the
capping, end processing, splicing, and components of the
export machinery are recruited to the loci or to mRNAs
cotranscriptionally (for review, see Iglesias and Stutz
2008). Similarly, as visualized by electron microscopy,
pre-rRNA processing was shown to be cotranscriptional
(Osheim et al. 2004), and recent kinetic studies of
metabolically labeled yeast cells concluded that, with
the prominent exception of cytoplasmic processing of 20S
pre-rRNA to 18S mature rRNA, processing excisions and
modification steps for rRNA production occur cotran-
scriptionally (Kos and Tollervey 2010). In stark contrast,
tRNA transcription and processing occur at several
distinct subcellular locations, including the nucleolus,
nucleoplasm, inner nuclear membrane (INM), cytoplasm,
and cytoplasmic surface of mitochondria (Fig. 5).
tRNA transcription in the nucleolus in yeast
Although best known for its prominent role in rRNA
transcription, pre-rRNA processing, and ribosome assem-
bly, the nucleolus is also the site for tRNA transcription
in yeast and pre-tRNA 59 end processing in numerous
organisms. The yeast genome contains 274 tRNA genes
distributed randomly among the 16 chromosomes. Yet, as
shown clearly by fluorescence in situ hybridization (FISH),
tRNA genes are recruited to a single subnuclear loca-
tion—the nucleolus—and are transcribed there (Thompson
et al. 2003; for review, see Hopper et al. 2010; Pai and
Engelke 2010). Location of the dispersed yeast tRNA genes
to the nucleolus is dependent on tRNA gene transcription,
as wild-type tRNA genes are located in the nucleolus
;50% of the time, whereas transcriptionally inactivated
tRNA genes are located in the nucleolus only ;10% of the
time (Thompson et al. 2003; Hopper et al. 2010). Localiza-
tion of tRNA genes to the nucleolus is also dependent on
intact nucleoli, as dispersion of the nucleolar structure
results in distribution of tRNA genes to the nucleoplasm
(Thompson et al. 2003; Wang et al. 2005).
synthesis and nuclear–cytoplasmic traf-
ficking for intron-containing tRNAs in
the yeast S. cerevisiae. tRNA transcription
and 59 end-processingoccurin the nucleolus.
Following 39 end-processing, CCA addition,
and various modification steps in the nucle-
oplasm and at the INM, intron-containing
pre-tRNAs are exported to the cytoplasm via
the Los1 exportin and at least one unknown
pathway. After pre-tRNA splicing on the
cytoplasmic surface of mitochondria, addi-
tional modifications in the cytoplasm, and
aminoacylation, mature charged tRNAs can
participate in protein synthesis. Cytoplasmic
tRNAs are constitutively imported into nu-
clei, directly or indirectly, via Mtr10. Re-
export of nuclear tRNAs to the cytoplasm is
mediated by Los1 and Msn5 and is regulated
by nutrient status; likely, Msn5-dependent
re-export requires that the tRNA be appro-
priately structured and aminoacylated in the
nucleus. (Green and red circles) Parts of the
The cell biology of tRNA bio-
tRNA that are maintained in the mature structure; (red circles) anticodon; (purple circles) transcribed 59 leader and 39 trailer sequences;
(dark-blue circles) intron sequence; (light-blue circles) CCA end; (yellow, orange, and pink circles) various modifications made in the
nucleoplasm, at the INM, and in the cytoplasm, respectively; (aa) amino acid. Processing steps are labeled, as are the b-importin members
that function in the nucleus–cytoplasm import and export steps.
tRNA processing and trafficking dynamics
GENES & DEVELOPMENT1845
The chromosome-condensing complex condensin is
involved in tRNA gene organization. Condensin associ-
ates physically with tRNA genes, and cells with condi-
tionally defective condensin subunits fail to cluster
tRNA genes in the nucleolus (D’Ambrosio et al. 2008;
Haeusler et al. 2008). Interestingly, condensin may func-
tion in clustering the tRNA genes together rather than in
locating them to the nucleolus per se. This idea is
supported by the observation that the nucleolar location
of tRNA genes can be disrupted by treatment with
nocodazole, but this treatment does not affect tRNA gene
clustering (Haeusler et al. 2008). Thus, clustering of tRNA
genes together and the location of the cluster to the
nucleolusappear tobeseparable processes. Although there
is little evidence for the localization of tRNA genes to the
nucleolus in other organisms, tRNA gene clustering may
be a conserved process, as it has been reported that dis-
persed tRNA genes in the fission yeast S. pombe cluster in
a nuclear region close to centromeres, and that clustering
at centromeres is dependent on condensin (Iwasaki et al.
2010). The roles for tRNA gene clustering and location
to the nucleolus are not completely understood, but this
located mRNA-encoding genes (Kendall et al. 2000; Wang
et al. 2005).
tRNA 59 end processing in the nucleolus
59 End processing is generally the first step of the tRNA
processing pathway. RNase P, the endonuclease respon-
sible for catalyzing removal of the 59 leader sequences
of pre-tRNAs (see above), is localized to the nucleolus
in yeast and vertebrate cells, although, in vertebrate
cells, some of the subunits of this complex RNP are also
located elsewhere(Bertrand et al. 1998; Jarrous et al. 1999).
Because RNase P shares subunits with RNase MRP
(Chamberlain et al. 1998), a complex RNP functioning
in pre-rRNA processing in the nucleolus (Schmitt and
Clayton 1993), the location of RNase P in the nucleolus
could merely facilitate sharing of the subunits with RNase
MRP. However, RNase P is more likely to have a more
direct nucleolar function, since it has been reported to
process yeast intron-encoded box C/D snoRNAs that are
required for appropriate pre-rRNA processing in the nu-
cleolus (Coughlin et al. 2008).
tRNA 39 end processing in the nucleoplasm?
39 End processing of pre-tRNAs generally follows 59 end
processing, and is catalyzed by both exonucleases (Rex1
in yeast) (Piper and Straby 1989; Copela et al. 2008;
Ozanick et al. 2009) and the RNase Z endonuclease (Trz1
in yeast) (for review, see Vogel et al. 2005). The balance
between endonucleolytic and exonucleolytic 39 end pro-
cessing depends, at least in part, on the tRNA-binding La
protein in both lower and higher eukaryotes (Huang et al.
2006; Copela et al. 2008; Zhao et al. 2009). Because there is
at least one yeast pre-tRNA for which 39 end processing
precedes 59 end processing (Kufel and Tollervey 2003), and
because some of the enzymes that function in 39 pre-tRNA
end processing also function in the pre-rRNA processing
pathway (e.g., Rex1) (van Hoof et al. 2000), one might have
predicted that pre-tRNA 39 end processing, like 59 end
processing, would occur in the nucleolus. However, there
appears to be no evidence for this, as the genome-wide
pools of Trz1 and Lhp1 (yeast La) are primarily nucleoplas-
mic, not nucleolar (Huh et al. 2003).
Nuclear tRNA modification enzymes reside at various
Several studies demonstrate that several different tRNA
modifications occur in the nucleus in various organisms.
Studies in Xenopus oocytes using a plasmid encoding
yeast tRNATyrinjected into oocyte nuclei showed that
some modifications were found on initial tRNA tran-
scripts, whereas others were not detected until the
tRNAs were end-matured (Melton et al. 1980; Nishikura
and De Robertis 1981). Since these initial transcripts and
processing intermediates were restricted to the nucleus,
the enzymes catalyzing modification must have resided
in the nucleus. Likewise, studies employing yeast with
defects in tRNA nuclear export showed that pre-tRNAs
contain some, but not all, modifications (Hopper et al.
1978; Knapp et al. 1978; Etcheverry et al. 1979). Consis-
tent with these findings, studies of individual tRNA mo-
dification proteins, as well as the genome-wide GFP-
tagging database in yeast, demonstrate that a subset of
the modification enzymes reside in the nucleus and lack
cytoplasmic pools (Simos et al. 1996; Huh et al. 2003; for
review, see Martin and Hopper 1994). For example, of the
eight pseudouridine synthases, two (Pus1 and Pus7) do
not appear to have cytoplasmic pools, and five of the 17
proteins involved with tRNA methylation (Trm1, Trm6,
Trm8, Trm61, and Trm82) appear to be nuclear-localized.
Surprisingly, modification enzymes residing primarily
in the nucleus have distinct subnuclear distributions, and
can be located in the nucleolus, in the nucleoplasm, or at
the INM. For example, Pus1 is rather evenly distributed
throughout the nucleoplasm (Simos et al. 1996; Huh et al.
2003). In contrast, MOD5-encoded isozymes are located
in the mitochondria, cytoplasm, and nucleus, and the
nuclear Mod5 is concentrated in the nucleolus (Tolerico
et al. 1999). Because the i6A modification catalyzed by
Mod5 (Laten et al. 1978; Martin and Hopper 1982) occurs
only after intron removal (Spinelli et al. 1997; for review,
see Grosjean et al. 1997), which happens in the cytoplasm
(Yoshihisa et al. 2003, 2007), the role of the Mod5 nu-
cleolar pool remains unknown. Trm1 is located at yet a
third intranuclear site, the INM. Although the cis-acting
and some of the trans-acting factors responsible for Trm1
distribution to the nucleus and the INM have been
characterized (Rose et al. 1992; Murthi and Hopper
2005; Lai et al. 2009), it remains unknown why Trm1 is
located in the nucleus and, specifically, at the INM, since
preventing Trm1 nuclear import does not prevent its
catalysis of tRNA modification (Rose et al. 1992). In sum,
reasons for the different subnuclear distributions of tRNA
modification enzymes remain to be resolved.
Phizicky and Hopper
1846GENES & DEVELOPMENT
tRNA modifications catalyzed in the cytoplasm
The Xenopus injection studies described above also
showed that some modifications, primarily those located
in the anticodon loop, appear only on mature tRNAs, and
therefore could be catalyzed in the cytoplasm (Melton
et al. 1980; Nishikura and De Robertis 1981); however,
there have been few reports regarding the distribution
of tRNA modification enzymes in higher eukaryotic
cells. In yeast, numerous tRNA modification proteins
have been reported to reside solely in the cytoplasm. For
some of these, cytoplasmic location may have evolved to
accommodate substrate specificity. For example, the
anticodon modifications of A37to i6A, and of C32and
G34to Cm32and Gm34, respectively, occur only after
intron removal (Jiang et al. 1997; Pintard et al. 2002; for
review, see Grosjean et al. 1997). Since, as described
below, intron removal in yeast is restricted to the cyto-
plasm (Yoshihisa et al. 2003, 2007), enzymes that catalyze
modifications that occur after splicing should be cyto-
plasmic, at least in yeast.
tRNA splicing on the mitochondrial cytoplasmic
surface in some, but not all, organisms
Perhaps the most curious partitioning of tRNA biogene-
sis enzymes concerns the location of the tRNA splicing
machinery in yeast. In stark contrast to the nuclear
location for tRNA splicing in vertebrates (Melton et al.
1980; Lund and Dahlberg 1998; Paushkin et al. 2004), pre-
tRNA splicing in yeast does not occur in the nucleus. As
described above, the yeast splicing machinery is com-
prised of three enzymes: tRNA splicing endonuclease,
tRNA ligase (Trl1), and 29 phosphotransferase (Tpt1).
Remarkably, the tRNA splicing endonuclease subunits
are located on the cytoplasmic surface of mitochondria
(Huh et al. 2003; Yoshihisa et al. 2003, 2007; Shaheen
and Hopper 2005). The other two proteins—Trl1 and
Tpt1—required for splicing are not excluded from the
nucleus (Huh et al. 2003; N. Dhungel and AK Hopper,
unpubl.). Nevertheless, since the pre-tRNAs that accu-
mulate in the cytoplasm are true processing intermedi-
ates (Yoshihisa et al. 2007), it is clear that, in yeast, tRNA
splicing occurs in the cytoplasm, not the nucleus.
The location of tRNA splicing in plants is not com-
pletely resolved. Expression of recombinant genes in
heterologous plants provided evidence for nuclear pools
of tRNA splicing endonuclease (Englert et al. 2007), con-
sistent with pre-tRNA splicing in plants occurring in the
nucleus as in vertebrates; however, other genetic studies
showed that inhibition of tRNA nuclear export results in
defects in intron removal (Park et al. 2005), most easily
explained by a cytoplasmic location of pre-tRNA splicing
in plants, as is the case for yeast. Although additional
in the plant kingdom, it is nevertheless clear that there are
different locations for pre-tRNA splicing among different
organisms. Possible reasons for the different subcellular
distributions of the tRNA splicing enzymes among eukary-
otes include selections based on other functions of the
splicing machinery (Sidrauski et al. 1996; Sidrauski and
Walter 1997; Paushkin et al. 2004) and/or coordination of
cellular metabolism and pre-tRNA processing in yeast, but
not in vertebrate cells (for review, see Hopper and Shaheen
2008; Hopper et al. 2010).
tRNA processes occurring in both the nucleus
and the cytoplasm
their 39-terminal CCA nucleotides, the E. coli genome
also has a CCA nucleotidyl transferase, and its deletion
causes slow growth, suggesting an important role of this
enzyme in tRNA 39 end repair in prokaryotes (Zhu and
Deutscher 1987). As described above, all eukaryotic
tRNA genes lack an encoded CCA and therefore require
CCA addition during processing, and available evidence
suggests that this step occurs in the nucleus. Injection
studies in Xenopus oocytes clearly document that CCA is
added to tRNAs before they are exported to the cyto-
plasm (Melton et al. 1980; Nishikura and De Robertis
1981; Lund and Dahlberg 1998), and it is known that the
CCA addition to tRNAs is required for efficient tRNA
nuclear export in vertebrate cells (Arts et al. 1998b;
Lipowsky et al. 1999). The evidence also supports a role
for CCA addition in the nucleus in yeast, as end-processed
intron-containing pre-tRNAs located primarily in the
nucleus possess CCA at their 39 ends (Wolfe et al. 1996).
Nevertheless, in addition to the nuclear pools, there are
also cytoplasmic pools of CCA nucleotidyl transferase in
both yeast and vertebrate cells (Solari and Deutscher
1982; Wolfe et al. 1996). These cytoplasmic pools likely
serve to repair tRNA 39 ends analogous to the function
of this activity in prokaryotes, since redistribution of cy-
toplasmic CCA nucleotidyl transferase to the nucleus in
yeast results in accumulation of mature tRNAs lacking
39 CCA nucleotides (Wolfe et al. 1994). Thus, in eukary-
otes, whereas nucleoplasmic CCA nucleotidyl transferase
functions in tRNA processing, cytoplasmic CCA nucleo-
tidyl transferase likely functions in tRNA end repair.
Although all E. coli tRNA genes encode
RTD turnover pathway
also appears to function in both the nucleus and the
cytoplasm. Evidence to support this is the fact that
hypomodified tRNAs that are subject to this pathway
are not completely stabilized if either the nuclear Rat1
59–39 exonuclease or the orthologous cytoplasmic Xrn1
exonuclease is deleted, but near-complete stabilization is
et al. 2008). In contrast to the RTD pathway, the TRAMP-
mediated tRNA turnover pathway is restricted to the
nucleus (for review, see Houseley et al. 2006).
The RTD turnover pathway
Enzymes shared by more than one
A common theme for tRNA processing enzymes is
sharing proteins encoded by single genes between the
cytoplasmic/nuclear compartments and in mitochondria.
The yeast tRNA modification enzyme Trm1 provided the
first example of eukaryotic genes encoding sorting iso-
zymes—multiple proteins encoded by single genes, each
with different subcellular distribution (Hopper et al.
tRNA processing and trafficking dynamics
GENES & DEVELOPMENT1847
1982). TRM1 encodes two proteins differing by possession
of a 16-amino-acid N-terminal peptide: Trm1-I, the long
form, is found exclusively in the mitochondria, and the
short form, Trm1-II, is distributed primarily to the
nuclear INM (Ellis et al. 1989; Li et al. 1989). Similar
mechanisms create different isozymes for Mod5 and
Cca1, resulting in the longest forms being located in
mitochondria and the shorter forms being located else-
where in cells (Wolfe et al. 1994; Tolerico et al. 1999; for
review, see Martin and Hopper 1994). Although the
information is still not complete, compartmental sharing
appears to be characteristic of tRNA processing enzymes
for yeast. For example, of the 17 known yeast genes
encoding subunits of nucleoside methyltransferases,
none appear to encode proteins solely targeted to mito-
chondria; thus, for those tRNA modifications that occur
on both mitochondrial and nuclear encoded tRNAs, the
mitochondrial enzymes must be encoded by the same
genes that encode the cytoplasmic- and/or nuclear-
located enzymes. Likewise, of the eight genes involved
in tRNA pseudouridylation, only two, PUS2 and PUS9,
encode activities that modify only tRNAs localized in
mitochondria (Behm-Ansmant et al. 2004, 2007). The
mechanisms by which the numerous dual-targeted activ-
ities are distributed to multiple subcellular compart-
ments have not been completely determined.
Compartmental sharing isalsofound amongsomeofthe
aminoacyl-tRNA synthetases. As first shown in yeast, a
single gene encodes both cytoplasmic and mitochondrial
histidyl-tRNA synthetase (Natsoulis et al. 1986). Simi-
larly, mitochondrial and cytoplasmic alanyl-tRNA, glycyl-
tRNA, and valyl-tRNA synthetases are encoded by single
genes (Chatton et al. 1988; Turner et al. 2000; Tang et al.
2004). The remainder of the yeast mitochondrial and
cytoplasmic tRNA synthetases appear to be encoded by
separate genes. In plants, a similar situation exists; mito-
chondrial and chloroplast tRNA synthetases are generally
encoded by the same gene, and there are multiple exam-
ples of single genes encoding both cytoplasmic and plastid
activities (Duchene et al. 2005).
Amazing nuclear–cytoplasmic tRNA dynamics
A long-standing view of the nuclear–cytoplasmic dynam-
ics for tRNAs was that tRNAs were transcribed and
processed in the nucleus and then delivered to the
cytoplasm for aminoacylation and function in transla-
tion. This view was challenged by the demonstration that
(1) tRNA aminoacylation is not restricted to the cyto-
plasm (Arts et al. 1998b; Lund and Dahlberg 1998; Sarkar
et al. 1999; Grosshans et al. 2000), (2) defects in tRNA
aminoacylation in the nucleus cause tRNA nuclear
accumulation (Lund and Dahlberg 1998; Sarkar et al.
1999; Grosshans et al. 2000; Azad et al. 2001; Steiner-
Mosonyi and Mangroo 2004), and, (3) in yeast, tRNA
splicing occurs not in the nucleoplasm, but rather in the
cytoplasm on the surface of mitochondria (Yoshihisa
et al. 2003, 2007). These observations and the fact that,
in yeast, spliced tRNAs (i.e., tRNAs that had ‘‘visited’’the
cytoplasm) accumulate in the nucleus when aminoacyl-
ation is defective (Grosshans et al. 2000; Feng and Hopper
2002) led to the proposal and subsequent proof that tRNA
subcellular traffic is not unidirectional, from the nucleus
to the cytoplasm. Rather, a retrograde pathway exists by
which cytoplasmic tRNAs are imported into the nucleus,
and they can be re-exported to the cytoplasm in response
to nutrient availability (Shaheen and Hopper 2005;
Takano et al. 2005; Hurto et al. 2007; Whitney et al.
2007). This retrograde process is conserved in vertebrate
cells, and HIVappears to have co-opted the process as one
mechanism to deliver retrotranscribed complexes to the
nucleus (Zaitseva et al. 2006; Shaheen et al. 2007). The
movement of tRNA between the nucleus and the cyto-
plasm thus occurs in three separate steps (Fig. 5). First,
newly transcribed and partially processed tRNAs are
exported from the nucleus to the cytoplasm (‘‘primary
tRNA nuclear export’’). Second, tRNAs are imported
from the cytoplasm into the nucleus (‘‘tRNA nuclear
import’’). Finally, tRNAs that were once in the cytoplasm
but reside in the nucleus can re-enter the cytoplasm by
‘‘tRNA re-export’’ (Whitney et al. 2007).
Primary tRNA nuclear export
There is a well-defined mechanism for the export of
tRNA from the nucleus to the cytoplasm that uses the
Ran-GTPase pathway and the vertebrate member of the
Ran-binding b-importin family, Xpo-t (Arts et al. 1998a;
Kutay et al. 1998). Xpo-t orthologs in fungi (Los1 or Xpot)
and plants (PSD or PAUSED) appear to function similarly
(Hellmuth et al. 1998; Sarkar and Hopper 1998; Hunter
et al. 2003). Xpo-t and its orthologs directly bind tRNA in
the presence of Ran-GTP, and the heterotrimeric Ran-
GTP•Xpo-t•tRNA complex moves through nuclear pores
to the cytoplasm, where, upon Ran-GAP-mediated en-
hancement of hydrolysis of Ran-GTP to Ran-GDP, the
complex dissociates and tRNA is released. Recently, a 3.2
A˚resolution structure of S. pombe Xpot in complex with
Ran-GTP and tRNA was obtained, as well as a 3.1 A˚
structure of the unbound Xpot (Cook et al. 2009). The
structures elegantly demonstrate that Xpot wraps around
tRNA, making contacts with the acceptor arm and the
TCC and D-loops, leaving the anticodon loop exposed.
These studies verify earlier work using structure probing
and the use of artificial substrates that demonstrated that
Xpo-t binds preferentially to tRNAs with mature 59 and
39 termini containing a CCA end, and that alterations of
tRNA sequences disrupting the cloverleaf or tertiary
tRNA structure result in reduced binding to Xpo-t, but
the presence of introns does not influence the binding
efficiency of Xpo-t to tRNA (Arts et al. 1998b; Lipowsky
et al. 1999). The interactions show that Xpo-t provides a
proofreading function so that tRNAs with inappropriate
structure or immature 59 and 39 termini fail to be de-
livered to the cytoplasm. The interactions also account
for how unspliced tRNAs containing introns in the
anticodon loop can interact with Los1 and be exported
to the cytoplasm. However, the data shed little light on
how aminoacylation affects tRNA nuclear export because
unspliced tRNAs cannot be aminoacylated (O’Farrell et al.
Phizicky and Hopper
1848 GENES & DEVELOPMENT
1978), but, in yeast, they are nevertheless exported from
the nucleus by Los1.
Two lines of evidence suggest that, at least in some
organisms, there is an additional pathway to deliver tRNA
to the cytoplasm that is independent of the Xpo-t ortho-
logs, First, LOS1 in budding and fission yeast, and its plant
ortholog, PSD, are each nonessential, even though tRNA
nuclear export is obviously essential for these cells.
Although plants missing PSD have developmental defects,
growth of los1D and LOS1 wild-type cells is very similar
(Hurt et al. 1987; Hunter et al. 2003; Li and Chen 2003).
Second, some organisms, such as arthropods, do not have
anXpo-torthologamongtheir b-importinfamily members
(S Shibata et al. 2006 and references therein).
In addition to its role in nuclear export of particular
phosphorylated proteins in budding yeast (Kaffman et al.
1998; DeVit and Johnston 1999; for review, see Hopper
1999) and in nuclear export of pre-miRNAs in metazoans
and plants (Lund et al. 2004; Zeng and Cullen 2004; Park
et al. 2005; S Shibata et al. 2006), the b-importin member
Exportin-5 likely serves as a tRNA nuclear exporter in
some organisms. Exportin-5 binds tRNA in the presence
of Ran-GTP (Bohnsack et al. 2002; Calado et al. 2002; S
Shibata et al. 2006). The structure of this complex has not
yet been reported, but there is a prediction of it based on
the recently reported 2.9 A˚structure of pre-miRNA in
complex with Exportin-5 and Ran-GTP (Okada et al.
2009). In yeast, biological experiments have documented
a role for the Exportin-5 ortholog Msn5 in tRNA nuclear
export. First, los1D msn5D double mutants accumulate
more tRNA in the nucleus than either single mutant
(Takano et al. 2005). (Since los1D msn5D double mutants
are hearty, there must be at least one other unidentified
exporter for tRNA nuclear export in yeast.) Second, as
described further below, Msn5 functions in the tRNA re-
export part of the retrograde process (Eswara et al. 2009;
Murthi et al. 2010). Likewise, in Drosophila, where the
Exportin-t ortholog is absent, the Exportin-5 ortholog
dmExp5 serves as a tRNA nuclear exporter in addition
to serving for pre-miRNA nuclear export (S Shibata et al.
2006). So, in yeast and flies, Exportin-5 homologs function
in tRNA nuclear export. This is in contrast to the biological
role in other organisms. For example, despite the ability of
vertebrate Exportin-5 to bind tRNA, it has not been con-
sidered a major contributor to tRNA nuclear–cytoplasmic
dynamics for vertebrates because Xpo-t exports the major-
ity of the tRNA nuclear pool to the cytoplasm in vertebrate
cells (Arts et al. 1998b; Lipowsky et al. 1999), and because
Exportin-5hasonly asmall effect onthetRNAnuclear pool
(Bohnsack et al. 2002; Calado et al. 2002). Similarly, the
plant Exportin-5 ortholog HST has been reported not to
function in tRNA nuclear export (Park et al. 2005). In sum,
Exportin-5 homologs appear to function in tRNA nuclear
export in some, but not all, organisms.
Retrograde tRNA nuclear import
One member of the yeast b-importin family, Mtr10
(vertebrate TNPO3 or TRN-SR2), is implicated in retro-
grade tRNA nuclear import. Deletion of MTR10 causes
defects in tRNA nuclear accumulation (Shaheen and
Hopper 2005; Hurto et al. 2007; Whitney et al. 2007).
Whether Mtr10 has a direct interaction with tRNA is
unknown, but its role may be indirect, based on well-
characterized interactions of Mtr10 and TNPO3 with
proteins, especially RNA-binding proteins (for review, see
Pemberton and Paschal 2005). tRNAs accumulate in the
nucleus upon nutrient deprivation (Shaheen and Hopper
2005; Hurto et al. 2007; Whitney et al. 2007). However,
the nuclear import process is not nutrient-sensitive, as it
has been shown to be a constitutive process (Shaheen and
Hopper 2005; Takano et al. 2005; Zaitseva et al. 2006;
Murthi et al. 2010).
Retrograde tRNA re-export
Since tRNA nuclear import is constitutive, but the
retrograde pathway is nutrient-sensitive, the re-export
step is likely to be regulated and nutrient-responsive.
Interestingly, two lines of evidence suggest that Los1
functions in both the tRNA primary and the re-export
processes. First, cytoplasmic tRNAs accumulate in nu-
clei in los1D heterokaryons (Shaheen and Hopper 2005;
Takano et al. 2005). Second, the nuclear pool of tRNAs
in los1D cells is markedly reduced when tRNA nuclear
import is reduced in mtr10D los1D cells (Murthi et al.
2010). Los1-mediated nuclear export is unlikely to be
responsible for nutrient-dependent or aminoacylation-
dependent tRNA re-export because Los1 can export pre-
tRNA that is not aminoacylated.
As described above, the yeast b-importin member
Msn5 functions in tRNA nuclear export. As deletion of
MSN5 has no effect on nuclear export of intron-contain-
ing pre-tRNAs, Msn5’s role in nuclear tRNA export is
restricted to the re-export pathway, at least for those
tRNAs encoded by intron-containing genes (Eswara et al.
2009; Murthi et al. 2010), and is the likely tRNA exporter
responding to nutrient availability. It is unknown how
to the cytoplasm, but not transcripts that contain introns.
It is also unknown how Msn5 could be nutrient-responsive
and/or export primarily aminoacylated tRNAs. One pro-
posal is that it would cooperate with another protein able to
distinguish mature tRNAs from intron-containing pre-
tRNAs, and aminoacylated from uncharged tRNAs. The
translation elongation factor Tef1/2 (vertebrate eEF1-a) is
a candidate for such a protein because it interacts only
with mature aminoacylated tRNA, has been implicated
in tRNA nuclear export (Grosshans et al. 2000; McGuire
and Mangroo 2007), and is able to move into and out of the
yeast nucleus (Murthi et al. 2010). Other yeast proteins
including Utp9 (Eswara et al. 2009), Utp8 (McGuire et al.
2009), and Sol1/2 (Shen et al. 1996; Stanford et al. 2004)
have also been implicated in the nuclear–cytoplasmic
to learn the roles of these proteins in tRNA re-export.
The pace of progress in our understanding of tRNA pro-
cessing, tRNA trafficking, and tRNA function is truly
tRNA processing and trafficking dynamics
GENES & DEVELOPMENT 1849
staggering, since almost all of the highlighted informa-
tion in this review is new since 2003 (Hopper and
Phizicky 2003), and since we omitted so many results
from the study of organisms other than yeast and from
organelles. The future promises to be equally revealing.
For example, we can expect that every tRNA modifica-
tion pathway will be worked out in yeast and in several
other model organisms, as well as in humans, and we will
know most of the functions in some detail. In yeast, the
recent finding that sua5 mutants lack the universally
conserved threonylcarbamoyladenosine modification
(t6A) has opened the door to study of this ancient modi-
fication (El Yacoubi et al. 2009). The increasing applica-
tion of synthetic genetic and chemical genetic arrays and
other similar methods for analyzing complex genetic
phenotypes promises to yield great insights into tRNA
biology (Gustavsson and Ronne 2008; Wilmes et al.
2008). Increased use of deep sequencing methods and
microarray-based analysis will undoubtedly contribute
significantly new information on tRNA expression, sig-
naling, and translation (Netzer et al. 2009). The increased
sophistication of crystallographic analysis (Murphy et al.
2004; Ogle and Ramakrishnan 2005) and in vitro trans-
lation assays (Kothe and Rodnina 2007; Zaher and Green
2009) promises to add great dimension to our understand-
ing of tRNA translation efficiency and fidelity, and ever
more sophisticated assay systems will allow an increas-
ingly clear picture of translation in vivo (Salas-Marco and
tRNA trafficking pathways. In addition, further study of
the involvement of tRNA and tRNA fragments in signal-
ing pathways and stress response pathways will undoubt-
edly lead to new links between tRNA function and other
global cellular response systems.
This work was supported by grants from the NIH: GM27930 to
A.K.H., and GM52347 to E.M.P. We thank Elizabeth Grayhack
for comments on the manuscript, and are grateful to the students
and post-doctoral fellows in the Hopper and Phizicky laborato-
ries for many stimulating conversations.
Note added in proof
Recent results demonstrate that the tRNA m5C methyltrans-
ferase Dnmt2, which modifies residue 38 of substrate tRNAs
(Goll et al. 2006), has a prominent role in the stress response
(Schaefer et al. 2010). Drosophila Dnmt2 mutants are signifi-
cantly less viable in response to oxidative or heat stress, and
substrate tRNAs in these mutants are more sensitive to stress-
induced cleavage and cleavage by angiogenin (Schaefer et al.
2010). Since this sensitivity to cleavage by angiogenin is also
observed in tRNA from mouse embyonic fibroblasts derived
from the corresponding mouse mutant (Schaefer et al. 2010),
these results suggest that tRNA modification may play a con-
served role in modulation of the stress response.
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