Virology 358 (2007) 448 – 458
www.elsevier.com/locate/yviro
Structures required for poly(A) tail-independent translation overlap with,
but are distinct from, cap-independent translation and RNA replication
signals at the 3′ end of Tobacco necrosis virus RNA
Ruizhong Shen a,1 , W. Allen Miller a,b,⁎
a
Interdepartmental Genetics Program and Department of Plant Pathology, Iowa State University, IA, USA
b
Department of Biochemistry, Biophysics and Molecular Biology, Iowa State University, IA, USA
Received 25 July 2006; returned to author for revision 14 August 2006; accepted 31 August 2006
Available online 4 October 2006
Abstract
Tobacco necrosis necrovirus (TNV) RNA lacks both a 5′ cap and a poly(A) tail but is translated efficiently, owing in part to a Barley yellow
dwarf virus (BYDV)-like cap-independent translation element (BTE) in its 3′ untranslated region (UTR). Here, we identify sequence downstream
of the BTE that is necessary for poly(A) tail-independent translation in vivo by using RNA encoding a luciferase reporter gene flanked by viral
UTRs. Deletions and point mutations caused loss of translation that was restored by adding a poly(A) tail, and not by adding a 5′ cap. The two 3′proximal stem–loops in the viral genome contribute to poly(A) tail-independent translation, as well as RNA replication. For all necroviruses, we
predict a conserved 3′ UTR secondary structure that includes the BTE at one end of a long helical axis and the stem–loops required for poly(A)
tail-independent translation and RNA replication at the other end. This work shows that a viral genome can harbor distinct cap- and poly(A) tailmimic sequences in the 3′ UTR.
© 2006 Elsevier Inc. All rights reserved.
Keywords: Barley yellow dwarf virus-like translation element; Stem-loop structure; 3′ untranslated region; Translational control; Tombusviridae; Necrovirus;
Poly(A) tail mimic
Introduction
Translational control is a major step of gene regulation for
RNA viruses, oocytes, and other systems with little or no
transcriptional control. Most translational control elements and
features in mRNAs exist in the 5′ and 3′ untranslated regions
(Hughes, 2006; Mignone et al., 2002; Pesole et al., 2001; Wilkie et
al., 2003). On average, 3′ UTRs are substantially longer than 5′
UTRs (Pesole et al., 2001). Consequently, the 3′ UTR is a region
with great regulatory potential. Many 3′ UTRs contain translational control elements and an array of diverse binding sites for
⁎ Corresponding author. Plant Pathology Department, 351 Bessey Hall, Iowa
State University, Ames, IA 50011, USA. Fax: +1 515 294 9420.
E-mail address: wamiller@iastate.edu (W.A. Miller).
1
Current address: Dept. of Medicine/Gastroenterology, 703 19th Street SouthZRB 633, University of Alabama, Birmingham, AL 35294, USA.
0042-6822/$ - see front matter © 2006 Elsevier Inc. All rights reserved.
doi:10.1016/j.virol.2006.08.054
regulatory proteins or RNAs (He and Hannon, 2004; Kuersten
and Goodwin, 2003; Wickens et al., 2000; Wilkie et al., 2003).
A ubiquitous 3′ UTR element is the poly(A) tail. It regulates
both stability and translational efficiency of mRNAs (Jacobson,
1996). The 5′ cap and poly(A) tail function synergistically to
facilitate efficient translation initiation via circularization of
mRNA-initiation factor complex (Gallie, 1991; Hentze, 1997;
Preiss and Hentze, 1999; Preiss and Hentze, 1998; Tarun and
Sachs, 1995; Wells et al., 1998). Eukaryotic initiation factor
(eIF) 4E binds the 5′ cap and is associated with eIF4G. eIF4G
also binds poly(A) binding protein (PABP), which binds to the
poly(A) tail (Kahvejian et al., 2005). Thus, mRNA is
circularized (Gallie, 1991; Hentze, 1997; Preiss and Hentze,
1999; Preiss and Hentze, 1998; Tarun and Sachs, 1995; Wells et
al., 1998). The mRNA circularization provides a framework to
understand how elements within 3′ UTRs control translation.
Many positive strand viral RNAs have a cap structure but
lack a poly(A) tail. Generally, specific sequences within the 3′
R. Shen, W.A. Miller / Virology 358 (2007) 448–458
UTR replace the function of a poly(A) tail in translation (Gallie
and Kobayashi, 1994; Gallie and Walbot, 1990). This region
also contains the promoter for negative strand synthesis, which
can be regulated by switchable repressor activity in many plant
viruses (Dreher, 1999; Dreher and Hall, 1988; Matsuda et al.,
2004; Olsthoorn et al., 1999; Petrillo et al., 2005; Pogany et al.,
2003; Zhang et al., 2004a).
The RNAs of Tobacco mosaic virus (TMV) (Gallie et al.,
1996; Gallie and Walbot, 1990), rotavirus (Piron et al., 1998;
Varani and Allain, 2002), Turnip yellow mosaic virus (Gallie
and Kobayashi, 1994; Matsuda and Dreher, 2004), and Brome
mosaic virus (Gallie and Kobayashi, 1994) have functional
alternatives for the poly(A) tail in the 3′ UTR. Histone mRNAs
of metazoans, but not plants, also lack a poly(A) tail and harbor
a stem–loop structure that functionally mimics a poly(A) tail
(Gallie et al., 1996; Gorgoni et al., 2005). The 3′ UTR of Alfalfa
mosaic virus (AMV) contains a series of stem–loops separated
by AUGC motifs that bind viral coat protein to facilitate
poly(A) tail-independent translation (Neeleman et al., 2001).
Multiple domains in the 3′ UTR each contribute to the poly(A)
tail-independent translation of dengue virus RNA (Chiu et al.,
2005; Holden and Harris, 2004). While poly(A) tail-independent translation has been studied in capped viral RNAs,
sequences that substitute for poly(A) tails have not been clearly
identified on uncapped viral RNAs.
To investigate poly(A) tail-independent translation of a
naturally uncapped viral RNA, we used Tobacco necrosis virus
strain D (TNV-D) as a model system. TNV is the type member
449
of genus Necrovirus in the Tombusviridae family. TNV-D has a
positive sense single-stranded RNA genome of 3762 nt that has
neither a 5′ cap (Lesnaw and Reichmann, 1970) nor a 3′
poly(A) tail (Coutts et al., 1991). TNV-D RNA encodes six
open reading frames (ORFs) (Fig. 1A). We reported previously
(Shen and Miller, 2004b) that TNV-D RNA has a BYDV-like
cap-independent translation element (BTE) within the 3′ UTR
that functionally mimics a 5′ cap (Fig. 1A). When located in the
3′ UTR (its natural location), the TNV BTE is predicted to base
pair to the viral 5′ UTR, via kissing stem–loops, to function.
The BTE also functions in the 5′ UTR when the AUG triplets
within the BTE are altered to prevent initiation before the
correct start codon. A 105 nt sequence (nts 3555–3659) is
sufficient for cap-independent translation in wheat germ extract,
but an undetermined amount of additional sequence downstream is required for efficient cap-independent translation in
vivo (Shen and Miller, 2004b). The extent and role of the extra
sequence are not clear.
Here, we report that a small portion of the extra sequence is
necessary for cap-independent translation in vivo, but most of
the extra sequence required for translation in vivo can be
replaced by a 60 nt poly(A) tail, and not by a 5′ cap. This
suggests that the extra sequence has a poly(A) tail-substitute
function. The extra sequence includes two stem–loops wellknown to serve as the promoter for initiation of negative strand
RNA synthesis. These results show that the poly(A) tailindependent and cap-independent translation require different,
but possibly overlapping, portions of the viral 3′ UTR.
Fig. 1. Deletion mapping of 3′ UTR translational control signals in oat protoplasts. (A) TNV genome organization. Open boxes represent open reading frames (ORFs)
with encoded protein names in or beside boxes. Shaded box indicates the in vitro-defined BYDV-like cap-independent translation element (BTE). Bold black lines
represent genomic RNA (gRNA) and subgenomic RNAs (sgRNAs). (B) Left side: maps of TLucT deletion mutants (not to scale). Positions of deletions are indicated
using the wild-type TNV-D genomic RNA numbering. fLuc: firefly luciferase ORF. 5′ and 3′ UTRs are indicated by black bold lines with blank areas indicating
deleted portions. Right side: relative luciferase activities of translation products of TLucT and deletion mutants obtained 4 h after electroporation of protoplasts.
Luciferase assays were performed in at least three independent experiments, each of which was in triplicate. Standard deviations are indicated.
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R. Shen, W.A. Miller / Virology 358 (2007) 448–458
Results
A sequence within the TNV-D 3′ UTR functionally replaces a
poly(A) tail
We showed previously that the in vitro-defined BTE of TNV
has a 3′ boundary immediately upstream of nt 3659, but an
unknown amount of additional downstream sequence is needed
for full expression of capped and uncapped RNAs in vivo (Shen
and Miller, 2004b). Because poly(A) tails are necessary for
translation of plant mRNAs in vivo, but not in wheat germ
extract (WGE), we speculated that it might be possible to
replace the additional sequence with a poly(A) tail. To test the
function of the 103 nucleotides downstream of the BTE needed
for translation in vivo, we made a series of deletions and
truncations in this region in reporter construct TLucT, which
encodes the firefly luciferase ORF flanked by TNV-D 5′ and 3′
UTRs (Shen and Miller, 2004b). We examined the translatability of these mutated RNAs containing (i) both cap and a 60 nt
poly(A) tail, (ii) cap only, (iii) poly(A) tail only, or (iv) neither a
cap nor a poly(A) tail.
Addition of a 60 nt poly(A) tail to TLucT with the complete
3′ UTR increased luciferase expression about 2.5-fold (Fig.
1B). While this is significantly higher than expression in the
absence of the tail, it is far less than the level of stimulation seen
on cellular mRNAs that require a poly(A) tail (Gallie, 1991;
Tarun and Sachs, 1995). Thus, significant translation of TLucT
occurs in the absence of the poly(A) tail.
Deletions of portions of the 3′ end between nt 3720 and 3762
(the 3′ terminus) reduced translation to about 40% of that from
TLucT (Fig. 1B, TLucT, D3720, D3748). Addition of a 5′ cap
had no notable effect on translation. However, addition of a 60 nt
poly(A) tail alone restored translation to a similar level as TLucT
with a poly(A) tail. Addition of a 5′ cap and a poly(A) tail had a
similar effect on translation as the poly(A) alone. We conclude
that sequences between nt 3720 and 3745, and downstream of nt
3748 are necessary for poly(A) tail-independent translation and
are not required for cap-independent translation.
Truncations further upstream, from the 3′ end up to nt 3700
or 3680, reduced luciferase expression 14- to 20-fold relative to
TLucT. Again, addition of a 5′ cap had no effect on translation
of transcripts with or without a poly(A) tail. Presence of the
60 nt poly(A) tail increased luciferase expression about 5- to 8fold (D3680, D3700, Fig. 1B), but only to about half of the level
obtained from non-polyadenylated TLucT. Thus, full translation
could not be restored on RNAs with these larger truncations,
even in the presence of a 5′ cap and poly(A) tail.
Truncating still further upstream, to nt 3661, caused a 17fold reduction in translation. Addition of a 60 nt poly(A) tail
increased translation 4.5-fold. Addition of a 5′ cap increased
translation 2.5-fold. Interestingly, a 37-fold increase in
translation was observed in D3661 RNA by adding both a 5′
cap and a poly(A) tail. The translation level of capped,
polyadenylated D3661 RNA was similar to capped, polyadenylated TLucT RNA (Fig. 1B, TLucT, D3661). Thus, when all
sequence downstream of 3661 was deleted, addition of a cap
and poly(A) had a strikingly synergistic effect.
Taken together, these data show that sequence between nt
3661 and 3680 is necessary for cap-independent translation in
vivo, even though it was not needed in vitro. Sequence
downstream of nt 3680 does not contribute to cap-independent
translation. Why the capped, polyadenylated forms of the two
RNAs with intermediate truncations (D3610 or D3700)
translated at lower levels than the capped, polyadenylated
RNA with even more sequence deleted (D3661) is unclear, but it
is obvious that addition of a poly(A) tail increased translation at
least five-fold. Thus, sequence downstream of nt 3681 contains
element(s) required only for poly(A) tail-independent translation. At least a portion of this poly(A) tail-independent
translation element (PITE) extends downstream of nt 3748.
Two stem–loops are required for poly(A) tail-independent
translation
To guide investigation of the structure and function of nts
3681–3762, we used the MFOLD program (Zuker, 2003) to
predict the secondary structure of this portion of the PITE.
Nucleotides 3681–3762 are predicted to form a structure with
two stem–loops separated by a 15 nt single-stranded pyrimidine-rich tract (nts 3725–3739) (Fig. 2A). Stem–loop II has two
internal bulges. Based on the results in Fig. 1B, deletions that
truncate or remove stem–loop I (D3720, D3748) reduced poly
(A) tail-independent translation by 50%, and deletions that
removed both stem–loops (D3661, D3680, D3700) abolished
translation in the absence of a poly(A) tail.
To determine the primary and secondary structures required
for the PITE function, we introduced mutations into the stems
and loops and deleted the pyrimidine-rich tract (Figs. 2 and 3).
Deletion of the pyrimidine-rich tract (D3726–3738) decreased
luciferase activity about 3-fold (Fig. 2B). The loop I mutation
(LI-mut) had little effect on translation. SL-I mutations C3753G
and G3744C, predicted (by mfold (Zuker, 2003)) to weaken or
disrupt the SL-I helix, reduced translation by 2- and 3-fold,
respectively. The combination of these two mutations (SI-re),
which is predicted to restore the stem, restored translation
efficiency to the wild-type level. Presence of the 60 nt poly(A)
tail caused all constructs to generate about 2- to 2.5-fold more
luciferase than non-polyadenylated “wild-type” TLucT RNA
(Fig. 2B). Thus, the pyrimidine tract and the secondary structure
of SL-I are important for poly(A) tail-independent translation.
To determine the biological relevance of these mutations,
we subcloned them into full-length TNV-D genomic RNA
and observed their effects on viral RNA accumulation in
tobacco NT-1 protoplasts (Fig. 2C). Because translation is
necessary for production of viral replicase, mutations that
reduce translation should reduce replication. The pyrimidine
tract deletion (Δ3762–3738) decreased the replication of
TNV-D to an undetectable level. Stem I mutations (C3753G
and G3744C) and loop I mutation (LI-mut) dramatically
reduced accumulation of TNV-D RNA. Remarkably, the
compensatory mutation (SI-re) that restored SL-I resulted in a
massive increase in TNV-D RNA accumulation, many fold
above that of wild-type TNV-D RNA. This is not due to a
major change in stability of SL-I (wt ΔG = − 9.8 kcal/mol, SI-
R. Shen, W.A. Miller / Virology 358 (2007) 448–458
451
luciferase accumulation in protoplasts by 2- to 3-fold (Fig. 3B).
The double mutant containing G3697C and C3704G, predicted
to re-form the SL-II helix (SII-re), restored translation to the
wild-type level. Addition of a poly(A) tail caused all mutants to
translate at the level of polyadenylated TLucT RNA, which is
about double that of non-polyadenylated TLucT. LII-mut and
stem II mutation G3697C abolished the replication of TNV-D.
Stem II mutation C3704G significantly decreased the replication of TNV-D. SII-re restored the accumulation of TNV-D to a
level higher than wild type. Thus, as for the mutations in SL-I,
effects of these mutations on replication were more extreme
than on translation (Fig. 3C). In summary, the distal SL-II helix
and the primary sequence of L-II are important for the function
of the TNV-D PITE, as well as for TNV-D replication.
The upper bulge in SL-II, GGGU, can potentially base pair to
the 3′ terminal four bases ACCC of the TNV genome (boxed
sequence connected by dashed arrow, Fig. 3A). A similar
interaction, but with five base pairs, is required for the Tomato
bushy stunt tombusvirus (TBSV) and Turnip crinkle virus
(TCV) replication silencer to down-regulate negative strand
RNA synthesis (Pogany et al., 2003; Zhang et al., 2004a). We
tested the effect of disrupting this interaction on translation by
introducing point mutations in either the GGGU or ACCC
sequence or both (Fig. 3D). (The predicted G–C pairs were not
altered because the terminal three C's comprise half of the SmaI
site required for plasmid linearization prior to in vitro
transcription.) In all cases, translation was reduced by only
36% to 44% in the disruptive mutants and remained at a similar
lower level in the double mutant predicted to restore the base
pairing (Fig. 3D). Thus, this pseudoknot interaction does not
play a major role in poly(A) tail-independent translation in
uninfected cells.
Mutations have little effect on stability of mRNA
Fig. 2. Effect of pyrimidine-rich tract deletion and SL-I mutations on TLucT
translation and TNV-D replication. (A) Predicted secondary structure of the 3′
end of the TNV genome showing mutated bases in bold, and boxed with
corresponding base on opposite strand. Dash between bases indicates Watson–
Crick complementarity. Adjacent mutated bases in loop L1 and pyrimidine-rich
tract are also boxed. Names of constructs are shown beside boxes. (B) Relative
luciferase activity expressed from TLucT, pyrimidine-rich tract deletion (d3726/
3738), and SL-I mutants in oat protoplasts. (C) Northern blot hybridization of
total RNA accumulated in NT-1 protoplasts 24 h after inoculation with TNV-D
transcripts containing the indicated mutations. Probed with negative sense
transcript complementary to the 3′ end of the TNV-D genomic RNA. Mobilities
of viral genomic and subgenomic RNA are indicated at right. The bottom panel
shows ethidium bromide staining of the gel prior to blotting to indicate the
amount of RNA loaded in each lane.
re ΔG = − 9.6 kcal/mol). Thus, RNA replication was more
sensitive to alterations in the 3′ UTR than was translation.
But, as expected, all mutations that reduced translation had a
negative impact on RNA accumulation.
We next examined the effects of disrupting and restoring SLII (Fig. 3A). Mutations in loop II (LII-mut) and those that
weakened or disrupted stem II (G3697C and C3704G) reduced
The expression differences of the mutants could be caused by
changes in translation efficiency and/or RNA stability. To
distinguish between these possibilities, we assessed the functional
stability of the lowest expressing mutant RNAs. Functional halflife, the time in which the rate of reporter protein accumulation
halves, was determined. Time course analyses showed that, while
total luciferase accumulation from actively translated RNAs
varied, all mutants had a similar functional half-life (Fig. 4). Thus,
the mutations that reduced luciferase accumulation did not
significantly affect the stability of the reporter mRNA. We
conclude that the mutations that reduced luciferase expression did
so by reducing the translation efficiency of the mRNA.
Full-length TNV-D 3′ UTR, but not the double stem–loop
structure alone, is sufficient to replace the poly(A) tail
Having established that stem–loop I and at least the upper
portion of SL-II contribute to poly(A) tail-independent translation, we next tested whether the 3′-terminal sequence harboring
these two stem–loops and extending to the genomic 3′ end is
sufficient to replace a poly(A) tail. Because the BTE is absent in
most of these constructs, all constructs in this experiment were
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R. Shen, W.A. Miller / Virology 358 (2007) 448–458
capped. We tested a luciferase reporter construct, VLucT122, in
which the 5′ UTR is from vector sequence and the 3′ UTR
consists of the 3′-terminal 122 nt of TNV-D RNA (nts 3641–
3762), which includes the double stem–loop sequence discussed
above (V indicates vector-derived sequence, T indicates TNV-D
sequence). We compared translation efficiencies of VLucT122,
Fig. 4. Functional stability assay of reporter constructs. (A) Time course of
luciferase activity accumulation from TLucT and selected mutant RNAs at
indicated times after electroporation into oat protoplasts. Data represent the
mean from three independent experiments. (B) mRNA functional half-live
calculated from data in panel A (see Materials and methods).
with and without a poly(A) tail, to those of constructs that
harbored only vector-derived UTRs, VLucVAn and VLucV294,
in oat protoplasts. The positive control, VLucVAn, has a 67 nt
vector-derived 3′ UTR followed by a 60 nt poly(A), whereas
VLucV294 has a 294 nt vector-derived sequence as its 3′ UTR.
VLucT122 and VLucV294 translated about 7% as efficiently as
VLucVAn (Fig. 5). Poor translation of VLucV294 shows that a
long, non-poly(A) sequence is insufficient to stimulate poly(A)
tail-independent translation, in contrast to a previous report that
Fig. 3. Effects of mutations in SL-II and the 3′ terminus on translation and TNVD replication. (A) Predicted secondary structure of the 3′ end of the TNV
genome. Mutated bases are in bold, and boxed with corresponding base on
opposite strand, or with adjacent bases if in a predicted loop or bulge. Names of
constructs are shown beside rectangles. Dashed arrow indicates potential base
pairing between the terminal four bases and the bulge in SL-I, that is disrupted or
weakened in constructs U3714A and A3759U and restored in the double mutant
AU. (B) Relative activity of luciferase translated from TLucT transcripts
harboring indicated mutations without (unshaded bars) or with (shaded bars) a
poly(A) tail. (C) Northern blot hybridization of TNV-D RNA accumulated in
NT-1 protoplasts 24 h after inoculation with TNV-D transcripts containing the
indicated mutations. Mobilities of viral genomic and subgenomic RNA are at the
right. Ethidium bromide staining of the gel prior to blotting is below the blot. (D)
Relative luciferase activity translated from TLucT transcripts harboring
indicated mutations at the 3′ end and in the four base bulge in SL-II.
R. Shen, W.A. Miller / Virology 358 (2007) 448–458
453
Fig. 5. Test of the ability of sequences from the TNV-D 3′ UTR to replace a poly(A) tail. Left side: maps of constructs beside their names on the left. The sizes and
sequence of TNV RNA fragments added to 3′ UTRs of reporter constructs are indicated. Right side: relative luciferase activity obtained from translation of each
construct in oat protoplasts. Relative luciferase activities were normalized to the capped, polyadenylated construct with vector-derived UTRs and a 60 nt poly(A) tail
(VLucVAn). Luciferase assays were performed in at least three independent experiments, each of which was in triplicate. Standard deviations are indicated.
any long 3′ UTR enhances translation in vivo (Tanguay and
Gallie, 1996). Addition of a poly(A) tail increased the translation
of VLucT122 to the level of VLucVAn (Fig. 5, VLucVAn,
VLucV294, and VLucT122). Thus, the TNV-D 122 nt sequence
containing the 3′ terminus of TNV-D RNA, including the two
stem–loops, is not sufficient to replace a poly(A) tail.
The inability of the 122 nt sequence to replace a poly(A) tail
in the above construct may be due to the absence of the TNV-D
5′ UTR or additional upstream 3′ UTR sequence. To test these
possibilities, we first replaced the 5′ UTR of VLucT122 and
VLucV294 with the 5′ UTR of TNV-D to create constructs
TLucT122 and TLucV294 and examined their translatability.
No statistically significant difference between luciferase activity
from constructs containing the vector 5′ UTR or the TNV-D 5′
UTR was detected (Fig. 5). Thus, the viral 5′ UTR in combination with the 3′-terminal 122 nt was insufficient to confer
poly(A) tail-independent translation. Secondly, we replaced the
122 nt 3′ UTR of TLucT122 with the 171 nt closest to the TNVD RNA 3′ end (nts 3592–3762) to construct TLucT171. We
also constructed TLucTBF which is the same as TLucT but with
a GAUC duplication in a BamHI site that destroys the capindependent translation function of the BTE (Shen and Miller,
2004b). The translation of TLucT171 was 4.2-fold higher than
that of TLucT122, 3.2-fold higher than that of TLucV294, and
was about 40% as efficiently translated as VLucVAn. Thus, the
3′-proximal 171 nt of TNV-D RNA contains significant but
incomplete poly(A) tail-independent translation activity. Nonpolyadenylated TLucTBF translated even more efficiently than
the polyadenylated control VLucVAn (Fig. 5). Thus, the fulllength 3′ UTR of TNV-D can functionally replace a poly(A) tail
and PITE is separable from the BTE.
predictions of secondary structure shared by all necroviruses.
Stem–loops I and II and the potential pairing of the four terminal
bases to a bulge in SL-II are phylogenetically conserved (Fig. 6).
This provides strong support of the experimental data that the
structures exist and serve an important function. Double stem–
loop structures at the 3′ termini of all necroviruses can be
grouped into two classes: TNV-D-like and TNV-A-like. In the
TNV-D-like viruses (TNV-D, Leek white stripe and Beet black
scorch), the bulge in stem–loop II that base pairs to the 3′
terminus consists only of the four bases complementary to the 3′
end, and there are no predicted single-stranded bases on the
opposite strand. In TNV-A-like necroviruses, TNV-A, Olive
mild mosaic virus (OMMV), and Olive latent virus-1 (OLV-1),
this bulge may be larger than for the other necroviruses as there
are unpaired bases on both sides of the bulge, but stem–loop II
may contain non-Watson–Crick pairing of a G.A/A.G tandem
(Fig. 6) (Na and White, 2006). Furthermore, the four terminal
bases involved in the pseudoknot interaction of TNV-D-like
necroviruses consist of RCCC (R = purine), whereas it is CCCC
in the TNV-A-like viruses, with an extra 3′-terminal A at the 3′
end of OLV-1 RNA.
It is noteworthy that sequence upstream of SL-II needed for
full poly(A) tail-independent translation forms a long bulged,
branched stem–loop structure that includes the BTE. In TNV-D
RNA, bases 3664–3679 that are downstream of the minimal in
vitro-defined BTE (circled in Fig. 6), which ends at nt 3663
(Shen and Miller, 2004b), and are upstream of stem–loop II, are
predicted to base pair to sequence upstream of the BTE to
extend helix IVof the BTE. This extended stem IVof the BTE is
conserved among all necroviruses (BTE S-IV, Fig. 6).
Discussion
Conserved secondary structure at the 3′ ends of necrovirus
genomes
To begin to understand the structural basis of the above
observations, we examined the secondary structure of the entire
translational control region of the 3′ UTR. We found MFOLD
The sequence that substitutes for a poly(A) tail differs from the
BTE
Here we report that the two 3′-proximal stem–loops and
upstream sequence of TNV-D RNA 3′ UTR are necessary for
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R. Shen, W.A. Miller / Virology 358 (2007) 448–458
Fig. 6. Secondary structures of 3′ ends of TNV-D and other necrovirus genomes predicted using mfold. Circled region in TNV-D RNA represents the BTE that is fully
functional in vitro. Arrows indicate the 3′-most boundary of sequence required for cap-independent translation in vivo (nt 3680), and the minimal region required for
poly(A) tail-independent translation (upstream of BTE to downstream of nt 3748). Dots connect possible G.A/A.G tandem pairs in TNV-A, OMMV, and OLV-1
genomes. Dashed lines indicate predicted, phylogenetically conserved pairing of the 3′ terminus with the SL-II bulge not demonstrated here, but known to affect
replication in other Tombusviridae. Genbank accession numbers of sequences: Tobacco necrosis virus strain D, UK isolate (D_00942), Leek white stripe virus
(NC_001822), Beet black scorch virus (NC_004452), Tobacco necrosis virus strain A (NC_001777), Olive mild mosaic virus (NC_006939), Olive latent virus-1
(NC_001721).
the 3′ end to functionally replace a poly(A) tail. Translation of
mRNAs with 3′ deletions up to nt 3720 was restored fully by
adding only a poly(A) tail, indicating that this region is
unnecessary for cap-independent translation. 3′ truncations to
nts 3700 or 3680 were somewhat puzzling because translation
remained low even in the presence of a cap and a poly(A) tail.
However, it is clear that the region lacks a cap-mimic sequence
because addition of a 5′ cap has no stimulatory effect. Either this
region contains a sequence that counteracts a negative element
further upstream or the deletion causes the RNA to misfold to
the extent that translation of even capped and polyadenylated
transcripts is low.
Sequence between nt 3661 and 3680 is required for capindependent translation in vivo. The cap and poly(A) tail
synergistically enhance translation of the RNA truncated at nt
3661 (Fig. 1B, D3661). Thus, 3′ boundary of in vivo-defined
TNV BTE is between nt 3662 and 3680, compared to the in
vitro-defined 3′ boundary at nt 3659 (Shen and Miller, 2004b).
The full-length 3′ UTR of TNV-D is not only sufficient to
replace a poly(A) tail, but it also allows 70% higher translation
than a 60 base poly(A) tail and increases translation more than
27-fold compared to a 294 nt vector-derived 3′ UTR sequence
(Fig. 5). In summary, it is clear that the BTE and the PITE have
different sequences, but the full PITE overlaps with the BTE.
The boundaries of these elements are difficult to define
because the deletions used to map them could alter important
secondary structures in unpredictable ways. Nevertheless, these
results reveal two important properties of the 3′ UTR. First, it
harbors both a PITE and a cap-independent translation element
sequence on a naturally uncapped, non-polyadenylated mRNA.
Many 3′ UTR elements have been identified that confer poly(A)
tail-independent translation on capped viral RNAs (Chiu et al.,
2005; Leathers et al., 1993; Neeleman et al., 2001; Piron et al.,
1998), and many cap-independent translation elements have
been characterized on polyadenylated mRNAs (Hellen and
Sarnow, 2001; Kneller et al., 2006), but to our knowledge, this
is the first poly(A) tail-independent translation element that
has been characterized on an RNA that lacks both a cap and a
R. Shen, W.A. Miller / Virology 358 (2007) 448–458
poly(A) tail. Secondly, stem–loops required for the TNV-D
PITE are also known to be required for polymerase recognition
and initiation of minus strand synthesis in related viruses (Fabian
et al., 2003; Panaviene et al., 2005; Stupina and Simon, 1997;
Zhang et al., 2004a). Thus, translational control signals overlap
with replication signals.
The location of both the cap- and poly(A) tail-substitute
sequences in the 3′ UTR poses interesting question about RNA
circularization. If translation factors and the 40S ribosomal
subunit bind these elements, one might predict “circularization”
within the 3′ UTR itself (i.e. two elements in the 3′ UTR
interacting directly or indirectly with eIF4G). The complex
would then require the BTE-mediated long-distance base
pairing with the 5′ UTR to deliver the translation factors to
the 5′ end. Alternatively to the above model, the PITE may
interact with the 5′ UTR directly or via host proteins to enhance
the interaction with the 5′ UTR mediated by the BTE.
Conserved secondary structure of 3′ end containing the
cap-independent and poly(A) tail-independent translation
elements
The conserved 3′ UTR secondary structures in Fig. 6 reveal
an extension of stem IV of the BTE (BTE S-IV) that is longer
than that on the minimal BTE necessary for function in vitro.
Interestingly, it is the extra sequence needed for capindependent translation in vivo (nts 3662–3680) that confers
the extension to BTE S-IV. This extended helical region ends
just two bases upstream of 3′ SL-II. Because helices have a
tendency to stack coaxially, we drew the 3′ end differently than
in the “standard” format for Tombusviridae (Fig. 3A). Drawn as
in Fig. 6 also allows placement of the 3′-terminal four bases in
proximity with their conserved complementary bases in the
bulge of 3′ SL-II. These base pairs are shown as dashed lines
because they do not seem to contribute substantially to
translation. Instead, this base pairing plays a role in replication,
based on experimental results and phylogenetic comparisons of
many other viruses in the Tombusviridae (Na and White, 2006;
Pogany et al., 2003; Zhang et al., 2004a, 2004b). The
asymmetry of the bulge is expected to form a kink in the 3′
SL-II helix. Base pairing of the four terminal bases may
reinforce the kink and stack coaxially with either the top or
bottom half of the SL-II helix. The terminal base pairing may
stack coaxially with (and between) both SL-I and the top of SLII. This was proposed for tombusviruses (Na and White, 2006),
but necroviruses lack unpaired bases in the SL-II that may be
needed to allow such a structure to form.
TNV-A, OMMV and OLV-1 differ slightly from the other
necroviruses in that there are unpaired bases on both sides of
the bulge in 3′ SL-II, and the four terminal bases to which
this bulge pairs are all cytosines. Na and White proposed that
the G.A/A.G tandem in this bulge may base pair, extending
SL-II (Na and White, 2006). Sheared non-Watson–Crick base
pairing in a G.A/A.G tandem occurs, for example, in the cisacting element required for incorporation of the amino acid
selenocysteine during translation of selenoproteins (Walczak
et al., 1998).
455
The pyrimidine-rich tract may be a spacer to provide length
to allow the 3′ terminal four bases to reach the SL-II bulge. A
sequence this long (15–16 nt) between the terminal stem–loops
is present in all Tombusviridae RNAs, except that it is
interrupted by a short stem–loop in genus Tombusvirus, and it
is not pyrimidine-rich in other Tombusviridae.
Presence of a cap-independent translation element immediately upstream of the 3′ terminal structure needed for replication
resembles the organization of other Tombusviridae, including
dianthoviruses which also harbor a BTE (Mizumoto et al.,
2003), as well as Tombusviridae that harbor non-BTE capindependent translation elements, such as TBSV (Fabian and
White, 2006), TCV (Qu and Morris, 2000), Panicum mosaic
virus (Batten et al., 2006), and Maize necrotic streak virus
(Scheets and Redinbaugh, 2006). These all differ from BYDV
in which the BTE is located 800 nt upstream from the 3′ end,
even though the 3′ structures required for replication bear some
resemblance to those of the Tombusviridae (Koev et al., 2002).
The poly(A) tail overlaps with replication signals
The primary and secondary structural requirements for
replication of viral RNAs in the Tombusviridae are wellcharacterized. Stem–loops of similar size and position as 3′ SLI and 3′ SL-II, referred to as Pr and H5, respectively, in the
carmoviruses and as gPR and SL3, respectively, in the
tombusviruses have been mutated exhaustively (Fabian et al.,
2003; Pogany et al., 2003; Zhang et al., 2004a; Zhang and
Simon, 2005). Stem–loop I is required for minus strand
synthesis and facilitates replicase assembly (Panaviene et al.,
2005). Furthermore, the pseudoknot base pairing between the 3′
end and the SL-II bulge is predicted in eight of the nine Tombusviridae genera (Na and White, 2006). In this interaction,
there are usually five, rather than the four, base pairs predicted
in the necroviruses. In the above viruses, the short helix formed
by the 3′ end-SL-II bulge base pairing is expected to stack with
the adjacent helical domain in SL-II. A similar coaxially stacked
helix interaction occurs at the BYDV RNA 3′ end (Koev et al.,
2002) which is a Tombusviridae-like luteovirus (Miller et al.,
2002). In all cases, this interaction is predicted to make the 3′
end inaccessible to proteins such as the viral replicase.
Consistent with this, the 3′ end-S-L II bulge base pairing
attenuates minus strand synthesis (Pogany et al., 2003; Zhang et
al., 2004a). This would allow preferential synthesis of plus
strands which is the goal of these positive strand RNA viruses.
The role of the above elements in translation had not been
investigated for Tombusviridae. The switch above may also
serve as a switch between minus strand synthesis and
translation. We show that the base pairing of the terminal four
bases to the bulge does not appear to enhance or facilitate
translation directly (Fig. 3). Due to the requirement for a 3′terminal CCC for in vitro transcription, we could change only
the fourth base from the 3′ end, A to U. It is possible that the
three C–G pairs are sufficient. The lack of any significant
change in activity when the disrupted U–A pair was replaced by
an A and a U, respectively, suggests that this terminal pairing
plays little role in translation in uninfected cells (Fig. 3D).
456
R. Shen, W.A. Miller / Virology 358 (2007) 448–458
Alternatively, an entirely different and unpredicted structure,
rather than the restored helix, may form in the presence of the A
and U substitutions expected to replace the U–A pair. Even if
this were the case, the modest decrease in translation in all of the
mutants at this site (> 50% of wild type, Fig. 3D) does not
suggest a major role in translation for this helix. Importantly, the
role in translation of this base pairing of the four terminal bases
to the SL-III bulge remains to be determined in infected cells. It
is possible that, by interacting with the unpaired conformation
of the 3′ end, replicase sequesters the RNA away from the
translational machinery. Thus, the base paired conformation of
the four 3′-terminal bases would enhance translation efficiency
only in infected cells.
Control of translation and replication by the 3′ UTR has been
investigated in other viruses. The tRNA-liked structures (TLS)
of the Bromoviridae, Tymoviruses, and Tobamoviruses both
facilitate translation in the absence of a poly(A) tail (Gallie and
Kobayashi, 1994; Matsuda and Dreher, 2004) and serve as the
key replicase recognition site for minus strand synthesis. Dreher
and colleagues showed that eIF1A binding blocked replicase
access to the initiation site (Matsuda et al., 2004). This provides
the same function as the molecular switch proposed above to
favor either translation or plus strand synthesis. Similarly, the 3′
terminus of AMV RNA consists of a series of stem–loops
which can refold into a pseudoknotted structure that resembles a
TLS. The AMV coat protein binds the stem–loop series
conformation, while the pseudoknotted version is proposed to
serve as the active minus strand template (Vlot et al., 2001).
Advantage of viral RNA with a poly(A) tail-independent
element instead of a poly(A) tail
Why have viral RNAs evolved a poly(A) tail-independent
translation element instead of having a poly(A)? By lacking a
poly(A) tail, non-polyadenylated viral RNAs may avoid host
translational regulation mechanisms. By obviating the need for
PABP, a PITE evades competition with cellular mRNAs for
PABP. For example, the Rotavirus NSP3 protein competes with
PABP for binding eIF4F (Groft and Burley, 2002), shutting off
translation of cellular polyadenylated mRNAs in favor of viral
translation. Secondly, poly(A) tail-independent translation may
help the viral RNA bypass the host's antiviral defenses that shut
down translation via negatively regulating translation factor
activity. Thirdly, absence of a poly(A) tail allows the viral RNA
to harbor key RNA replication control elements at the extreme
3′ terminus where minus strand synthesis initiates. It is possible
that absence of a poly(A) tail provides no advantage for
translation per se, but instead provides the replicative advantage
of switchable replication control signals at the extreme 3′
terminus.
Materials and methods
Plasmids and RNA constructs
All clones were verified by automated sequencing at the
Iowa State University DNA Sequencing and Synthesis Facility.
Plasmid pTNV-D is a full-length infectious clone of TNV-D,
kindly provided by R. H. A. Coutts, Imperial College, London
(Coutts et al., 1991). pTLucT is the template for TLucT, which
has a firefly luciferase ORF as a reporter flanked by the 5′ and
3′ UTRs of TNV-D (Shen and Miller, 2004b). D3720, D3700,
D3680, and d3726–3738 were constructed by replacing the 3′
UTR of TLucT with the respective shortened 3′ UTR of TNV-D
generated by PCR. In D3720, nts 3721–3744 were deleted. In
D3700, nts 3701–3759 were deleted. Nts 3681–3759 were
deleted in D3680. D3748 and D3661 were truncations of TLucT
at BglII3745 and SspI3659, respectively.
Mutants within the stem–loops were constructed by using
standard PCR-mediated, site-directed mutagenesis as in Guo et
al. (2000, 2001). VLucVAn was described in Guo et al. (2000),
in which a 60 base poly(A) tail was inserted into the StuI/SalI
site of pGEM-luc (Promega, Madison, WI) (Guo et al., 2000).
VLucT122 was constructed by replacing the 3′ UTR of
VLucVAn with a 122 nt sequence from TNV-D 3′ UTR (nts
3641–3762), which includes the PITE sequence. V indicates
sequence from vector, and T indicates sequence from TNV-D.
The template for in vitro transcription of VLucV294 was
pGEMLUC linearized with SspI. TLucT122 and TLucV294
were constructed by replacing the 5′ UTR of VLucT122 and
VLucV294 with the 5′ UTR of TNV-D. In all constructs with a
vector-derived 5′ UTR, the 5′ UTR sequence is the complement
of nts 1755–1803 in pGEM-luc (Genbank accession no.
X65316). The 3′ UTR of VlucVAn is the complement of
pGEM-luc nts 45–101, and the 3′ UTR of TLucV294 and
VlucV294 is the complement of nts 4739–101. TLucT171 was
constructed by replacing the 3′ UTR of TLucT122 with the
171 nt (nts 3592–3762) sequence from the 3′ UTR of TNV-D.
TLucTBF has the full-length 3′ UTR of TNV-D with a GUAC
duplication in a BamHI site. The cap-independent translation
function of TE is destroyed by this duplication (Shen and
Miller, 2004b).
In vitro transcription
Capped and uncapped RNAs were synthesized by in vitro
transcription using the T7 mMESSAGE mMACHINE® and
MegaScript® kits (Ambion, Austin, TX) as per manufacturer's
instructions, respectively. Templates for RNAs with a poly(A)
tail were linearized with VspI. Template for VLucV294 was
pGEMLUC linearized with SspI. Templates for D3661 and
D3748 were pTLucT linearized with SspI and BglII, respectively. All other templates were digested with SmaI.
In vivo translation
Oat (Avena sativa cv. Stout) protoplasts were prepared and
electroporated with RNA as described in Dinesh-Kumar and
Miller (1993). Luciferase assays were done as in Shen and Miller
(2004b). We included a capped and polyadenylated renilla
luciferase reporter as an internal control, and the Promega StopN-Glo™ (Madison, WI) system was used to assay both
luciferase activities. All luciferase assays were performed in
triplicate in at least three independent experiments. Firefly
R. Shen, W.A. Miller / Virology 358 (2007) 448–458
luciferase activities were first normalized with renilla luciferase
activity to minimize variation between samples. The luciferase
activities of all constructs were then compared to TLucT, whose
luciferase activity is defined as 100%.
Northern blot hybridization
For TNV-D replication assays, NT-1 protoplasts were used
and incubated for 24 h after electroporation. Total RNAs were
extracted from these cells by using the Trizol reagent
(Invitrogen, Carlsbad, CA) as per manufacturer's instructions
and analyzed by Northern blot analysis as described previously
(Shen and Miller, 2004a). A 32P-labeled probe, complementary
to the 107 nt TE of TNV-D, was used to detect TNV-D gRNA
and sgRNAs.
Stability assay
Functional stability assays were done as described in
Danthinne et al. (1993). Protein accumulation (A) as a function
of time (t) was analyzed by using the first order kinetics
equation: A(t) = A0e−kt. A function y = a ln(t) + b was achieved
from the logarithmic trend line of curve protein accumulation (A)
vs. time (t) by using Microsoft Excel. Constant k was calculated
by giving an arbitrary time t = 60 min and the function. We then
calculated the functional half time t1/2 = ln(1 / 2) * (1 / k). Data
shown were the mean from three independent experiments.
Acknowledgments
The authors thank R.H.A. Coutts for the kind gift of pTNVD, R. Sheldahl and A. Zakokar for technical assistance, and A.
Rakotondrafara for thoughtful reading of the manuscript. This
research was funded by NIH grant number RO1 GM067104 and
also supported by Hatch Act funds and the Iowa Agriculture and
Home Economics Experiment Station.
References
Batten, J.S., Desvoyes, B., Yamamura, Y., Scholthof, K.B., 2006. A translational
enhancer element on the 3′-proximal end of the Panicum mosaic virus
genome. FEBS Lett. 580, 2591–2597.
Chiu, W.W., Kinney, R.M., Dreher, T.W., 2005. Control of translation by the
5′- and 3′-terminal regions of the dengue virus genome. J. Virol. 79,
8015–8303.
Coutts, R.H.A., Rigden, J.E., Slabas, A.R., Lomonossoff, G.P., Wise, P.J., 1991.
The complete nucleotide sequence of tobacco necrosis virus Strain-D.
J. Gen. Virol. 72, 1521–1529.
Danthinne, X., Seurinck, J., Meulewaeter, F., Van Montagu, M., Cornelissen,
M., 1993. The 3′ untranslated region of satellite tobacco necrosis virus RNA
stimulates translation in vitro. Mol. Cell. Biol. 13, 3340–3349.
Dinesh-Kumar, S.P., Miller, W.A., 1993. Control of start codon choice on a plant
viral RNA encoding overlapping genes. Plant Cell 5, 679–692.
Dreher, T.W., 1999. Functions of the 3′-untranslated regions of positive strand
RNA viral genomes. Annu. Rev. Phytopathol. 37, 151–174.
Dreher, T.W., Hall, T.C., 1988. Mutational analysis of the sequence and
structural requirements in brome mosaic virus RNA for (−) strand promoter
activity. J. Mol. Biol. 201, 31–40.
Fabian, M.R., White, K.A., 2006. Analysis of a 3′-translation enhancer in a
457
tombusvirus: a dynamic model for RNA–RNA interactions of mRNA
termini. RNA 12, 1304–1314.
Fabian, M.R., Na, H., Ray, D., White, K.A., 2003. 3′-Terminal RNA secondary
structures are important for accumulation of tomato bushy stunt virus DI
RNAs. Virology 313, 567–580.
Gallie, D.R., 1991. The cap and poly(A) tail function synergistically to regulate
mRNA translational efficiency. Genes Dev. 5, 2108–2116.
Gallie, D.R., Kobayashi, M., 1994. The role of the 3′-untranslated region of nonpolyadenylated plant viral mRNAs in regulating translational efficiency.
Gene 142, 159–165.
Gallie, D.R., Walbot, V., 1990. RNA pseudoknot domain of tobacco mosaic
virus can functionally substitute for a poly(A) tail in plant and animal cells.
Genes Dev. 4, 1149–1157.
Gallie, D.R., Lewis, N.J., Marzluff, W.F., 1996. The histone 3′-terminal stem–
loop is necessary for translation in Chinese hamster ovary cells. Nucleic
Acids Res. 24, 1954–1962.
Gorgoni, B., Andrews, S., Schaller, A., Schumperli, D., Gray, N.K., Muller, B.,
2005. The stem–loop binding protein stimulates histone translation at an
early step in the initiation pathway. RNA 11, 1030–1042.
Groft, C.M., Burley, S.K., 2002. Recognition of eIF4G by rotavirus NSP3
reveals a basis for mRNA circularization. Mol. Cell 9, 1273–1283.
Guo, L., Allen, E., Miller, W.A., 2000. Structure and function of a capindependent translation element that functions in either the 3′ or the
5′ untranslated region. RNA 6, 1808–1820.
Guo, L., Allen, E., Miller, W.A., 2001. Base-pairing between untranslated
regions facilitates translation of uncapped, nonpolyadenylated viral RNA.
Mol. Cell 7, 1103–1109.
He, L., Hannon, G.J., 2004. MicroRNAs: small RNAs with a big role in gene
regulation. Nat. Rev., Genet. 5, 522–531.
Hellen, C.U., Sarnow, P., 2001. Internal ribosome entry sites in eukaryotic
mRNA molecules. Genes Dev. 15, 1593–1612.
Hentze, M.W., 1997. eIF4G: a multipurpose ribosome adapter? [published
erratum appears in Science 1997 Mar 14;275(5306):1553] Science 275,
500–501.
Holden, K.L., Harris, E., 2004. Enhancement of dengue virus translation: role of
the 3′ untranslated region and the terminal 3′ stem–loop domain. Virology
329, 119–133.
Hughes, T.A., 2006. Regulation of gene expression by alternative untranslated
regions. Trends Genet. 22, 119–122.
Jacobson, A., 1996. Poly(A) metabolism and translation: the closed-loop model.
In: Hershey, J.W.B., Mathews, M.B., Sonenberg, N. (Eds.), Translational
Control. Cold Spring Harbor Laboratory Press, Cold Spring Harbor,
pp. 451–480.
Kahvejian, A., Svitkin, Y.V., Sukarieh, R., M'Boutchou, M.N., Sonenberg, N.,
2005. Mammalian poly(A)-binding protein is a eukaryotic translation
initiation factor, which acts via multiple mechanisms. Genes Dev. 19,
104–113.
Kneller, E.L., Rakotondrafara, A.M., Miller, W.A., 2006. Cap-independent
translation of plant viral RNAs. Virus Res. 119, 63–75.
Koev, G., Liu, S., Beckett, R., Miller, W.A., 2002. The 3′-terminal structure
required for replication of barley yellow dwarf virus RNA contains an
embedded 3′ end. Virology 292, 114–126.
Kuersten, S., Goodwin, E.B., 2003. The power of the 3′ UTR: translational
control and development. Nat. Rev., Genet. 4, 626–637.
Leathers, V., Tanguay, R., Kobayashi, M., Gallie, D.R., 1993. A phylogenetically conserved sequence within viral 3′ untranslated RNA pseudoknots
regulates translation. Mol. Cell. Biol. 13, 5331–5347.
Lesnaw, J.A., Reichmann, M.E., 1970. Identity of the 5′-terminal RNA
nucleotide sequence of the satellite tobacco necrosis and its helper virus:
possible role of the 5′-terminus in the recognition by virus-specific RNA
replicase. Proc. Natl. Acad. Sci. U.S.A. 66, 140–145.
Matsuda, D., Dreher, T.W., 2004. The tRNA-like structure of Turnip yellow
mosaic virus RNA is a 3′-translational enhancer. Virology 321, 36–46.
Matsuda, D., Yoshinari, S., Dreher, T.W., 2004. eEF1A binding to aminoacylated viral RNA represses minus strand synthesis by TYMV RNAdependent RNA polymerase. Virology 321, 47–56.
Mignone, F., Gissi, C., Liuni, S., Pesole, G., 2002. Untranslated regions of
mRNAs. Genome Biol. 3 REVIEWS0004.
458
R. Shen, W.A. Miller / Virology 358 (2007) 448–458
Miller, W.A., Liu, S., Beckett, R., 2002. Barley yellow dwarf virus: Luteoviridae
or Tombusviridae? Mol. Plant Pathol. 3, 177–183.
Mizumoto, H., Tatsuta, M., Kaido, M., Mise, K., Okuno, T., 2003. Capindependent translational enhancement by the 3′ untranslated region of red
clover necrotic mosaic virus RNA1. J. Virol. 77, 12113–12121.
Na, H., White, K.A., 2006. Structure and prevalence of replication silencer–3′
terminus RNA interactions in Tombusviridae. Virology 345, 305–316.
Neeleman, L., Olsthoorn, R.C., Linthorst, H.J., Bol, J.F., 2001. Translation of
a nonpolyadenylated viral RNA is enhanced by binding of viral coat
protein or polyadenylation of the RNA. Proc. Natl. Acad. Sci. U.S.A. 98,
14286–14291.
Olsthoorn, R.C.L., Mertens, S., Brederode, F.T., Bol, J.F., 1999. A
conformational switch at the 3′ end of a plant virus RNA regulates viral
replication. EMBO J. 18, 4856–4864.
Panaviene, Z., Panavas, T., Nagy, P.D., 2005. Role of an internal and two
3′-terminal RNA elements in assembly of tombusvirus replicase. J. Virol. 79,
10608–10618.
Pesole, G., Mignone, F., Gissi, C., Grillo, G., Licciulli, F., Liuni, S., 2001.
Structural and functional features of eukaryotic mRNA untranslated regions.
Gene 276, 73–81.
Petrillo, J.E., Rocheleau, G., Kelley-Clarke, B., Gehrke, L., 2005. Evaluation of
the conformational switch model for alfalfa mosaic virus RNA replication.
J. Virol. 79, 5743–5751.
Piron, M., Vende, P., Cohen, J., Poncet, D., 1998. Rotavirus RNA-binding
protein NSP3 interacts with eIF4GI and evicts the poly(A) binding protein
from eIF4F. EMBO J. 17, 5811–5821.
Pogany, J., Fabian, M.R., White, K.A., Nagy, P.D., 2003. A replication silencer
element in a plus-strand RNA virus. EMBO J. 22, 5602–5611.
Preiss, T., Hentze, M.W., 1998. Dual function of the messenger RNA cap
structure in poly(A)-tail-promoted translation in yeast. Nature 392,
516–520.
Preiss, T., Hentze, M., 1999. From factors to mechanisms: translation and
translational control in eukaryotes. Curr. Opin. Genet. Dev. 9, 515–521.
Qu, F., Morris, T.J., 2000. Cap-independent translational enhancement of turnip
crinkle virus genomic and subgenomic RNAs. J. Virol. 74, 1085–1093.
Scheets, K., Redinbaugh, M.G., 2006. Infectious cDNA transcripts of Maize
necrotic streak virus: infectivity and translational characteristics. Virology
350, 171–183.
Shen, R., Miller, W.A., 2004a. Subgenomic RNA as a riboregulator: negative
regulation of RNA replication by Barley yellow dwarf virus subgenomic
RNA 2. Virology 327, 196–205.
Shen, R., Miller, W.A., 2004b. The 3′ untranslated region of tobacco necrosis
virus RNA contains a barley yellow dwarf virus-like cap-independent
translation element. J. Virol. 78, 4655–4664.
Stupina, V., Simon, A.E., 1997. Analysis in vivo of turnip crinkle virus satellite
RNA C variants with mutations in the 3′-terminal minus-strand promoter.
Virology 238, 470–4707.
Tanguay, R.L., Gallie, D.R., 1996. The effect of the length of the 3′-untranslated
region on expression in plants. FEBS Lett. 394, 285–288.
Tarun, S.J., Sachs, A.B., 1995. A common function for mRNA 5′ and 3′ ends in
translation initiation in yeast. Genes Dev. 9, 2997–3007.
Varani, G., Allain, F.H., 2002. How a rotavirus hijacks the human protein
synthesis machinery. Nat. Struct. Biol. 9, 158–160.
Vlot, A.C., Neeleman, L., Linthorst, H.J., Bol, J.F., 2001. Role of the 3′untranslated regions of alfalfa mosaic virus RNAs in the formation of a
transiently expressed replicase in plants and in the assembly of virions.
J. Virol. 75, 6440–6449.
Walczak, R., Carbon, P., Krol, A., 1998. An essential non-Watson–Crick base
pair motif in 3′UTR to mediate selenoprotein translation. RNA 4, 74–84.
Wells, S.E., Hillner, P.E., Vale, R.D., Sachs, A.B., 1998. Circularization of
mRNA by eukaryotic translation initiation factors. Mol. Cell 2, 135–140.
Wickens, M., Goodwin, E.B., Kimble, J., Strickland, S., Hentze, M., 2000.
Translational control of developmental decisions. In: Sonenberg, N.,
Hershey, J.W.B., Mathews, M.B. (Eds.), Translational Control of Gene
Expression. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY,
pp. 295–370.
Wilkie, G.S., Dickson, K.S., Gray, N.K., 2003. Regulation of mRNA translation
by 5′- and 3′-UTR-binding factors. Trends Biochem. Sci. 28, 182–188.
Zhang, J., Simon, A.E., 2005. Importance of sequence and structural elements
within a viral replication repressor. Virology 333, 301–315.
Zhang, G., Zhang, J., Simon, A.E., 2004a. Repression and derepression of
minus-strand synthesis in a plus-strand RNA virus replicon. J. Virol. 78,
7619–7633.
Zhang, J., Stuntz, R.M., Simon, A.E., 2004b. Analysis of a viral replication
repressor: sequence requirements for a large symmetrical internal loop.
Virology 326, 90–102.
Zuker, M., 2003. Mfold web server for nucleic acid folding and hybridization
prediction. Nucleic Acids Res. 31, 3406–3415.