© 2000 Oxford University PressNucleic Acids Research, 2000, Vol. 28, No. 4
Influence of correct secondary and tertiary RNA folding
on the binding of cellular factors to the HCV IRES
F. E. Odreman-Macchioli, S. G. Tisminetzky, M. Zotti, F. E. Baralle* and E. Buratti
International Centre for Genetic Engineering and Biotechnology (ICGEB), Padriciano 99, 34012 Trieste, Italy
Received November 2, 1999; Revised and Accepted December 22, 1999
Structural integrity of the hepatitus C virus (HCV) 5′
UTR region that includes the internal ribosome entry
site (IRES) element is known to be essential for
efficient protein synthesis. The functional explanation
for this observation has been provided by the recent
evidence that binding of several cellular factors to
the HCV IRES is dependent on the conservation of its
secondary structure. In order to better define the
relationship between IRES activity, protein binding
and RNA folding of the HCV IRES, we have focused
our attention on its major stem–loop region (domain
III) and the binding of several cellular factors: two
subunits of eukaryotic initiation factor eIF3 and
ribosomal protein S9. Our results show that binding
of eIF3 p170 and p116/p110 subunits is dependent on
the ability of the domain III apical stem–loop region to
fold in the correct secondary structure whilst
secondary structure of hairpin IIId is important for
the binding of S9 ribosomal protein. In addition, we
show that binding of S9 ribosomal protein also
depends on the disposition of domain III on the HCV
5′ UTR, indicating the presence of necessary inter-
domain interactions required for the binding of this
protein (thus providing the first direct evidence that
tertiary folding of the HCV RNA does affect protein
Translation initiation in the hepatitis C virus (HCV) (1) is
under the control of an internal ribosome entry site (IRES)
starting at nt 20–45 of the 341 nt long 5′ untranslated region (5′
UTR) and extending ~30 nt in the core protein sequence (2–9;
for a recent review see 10). The 5′ UTR and the initial core
region fold upon themselves to form several characteristic
stem–loop domains (I, II, III and IV), held together by a helical
structure and a pseudoknot which are highly conserved in all
virus isolates (11,12). Mapping of the IRES boundaries has
shown that the sequence and structural integrity of domains II,
III and IV is important for driving efficient protein translation
(5,6,8,12–18) with domain I not participating directly in the
formation of the IRES itself but possibly playing a role in
inhibiting its activity (8). In addition, non-structured regions of
the HCV 5′ UTR (19) and long-range RNA–RNA interactions
(20) can also be important to preserve translation efficiency.
To date, the better characterized cellular factors which bind to
the IRES and may affect its activity are: p52 (La antigen) (21),
in regulation of HCV translation by acting outside the IRES
protein (25,26), hnNRP L (27), poly(C)-binding protein (28)
and eukaryotic translation initiation factor eIF3 (25,26,29).
of HCV IRES functioning and additional insight concerning
the general field of RNA–protein interactions. In fact, it has
become increasingly clear that in order to achieve correct
RNA–protein recognition two factors often have to be
conserved: nucleotide sequence integrity and correct RNA
secondary structure. Indeed, with regard to the secondary
structure, protein binding may also be needed to promote its
correct folding since it has recently been reported that binding
ofPTB to the EMCV 5′ UTR is essential for IRES activity only
when there is a necessity to stabilize the native secondary
Previously, we have described that the binding of two subunits
(p170 and p116/p110) of eIF3 to HCV RNA is important for
translation initiation (29). Using artificial mutants we identified
the binding region as the apical stem–loop region of domain III
and showed that direct disruption of its secondary structure
with a chemical and enzymatic footprinting study performed
on a binary eIF3–IRES complex which reported the binding of
four eIF3 subunits, p170, p116, p66 and p47, and identified the
binding site by footprinting analysis as consisting of the large
central helix of domain III and hairpins IIIa, IIIb and IIIc (26).
It was therefore interesting to further investigate the relationship
between IRES activity and correct folding of this domain,
evidently a complex issue since we had already described (29)
that inversion of stem–loop IIIb sequence resulted in a mutant
(5′ S/L) which had unexpectedly maintained IRES activity.
This mutant had retained the capacity to bind p170, although
not p116/p110, and the fact that one binding site was conserved
offered a likely explanation for the lack of significant variations
in the level of IRES activity (29).
In this study we have engineered several mutants which
carry selected portions of domain III and each mutant was
comprehensively analyzed for a variety of characteristics:
ability to drive IRES translation in vivo and in vitro, correct
folding of the apical stem–loop and ability to bind the eIF3
subunits previously identified (29). Our results show that only
*To whom correspondence should be addressed. Tel: +39 40 3757337; Fax: +39 40 3757361; Email: firstname.lastname@example.org
Nucleic Acids Research, 2000, Vol. 28, No. 4
one mutant comprising domain III sequence from nt 145 to nt
248 was capable of binding p170 and p116/p110 and, in
keeping with this result, the RNase digestion pattern of its
apical stem–loop showed that the RNA secondary structure
was identical to that of wild-type domain III. The fact that this
mutant was not capable of driving IRES translation prompted
us to look for additional cellular factors binding to the IIId
element and to investigate the importance of this stem–loop for
IRES activity. In particular, we focused our attention on the
interaction of our mutants with S9 ribosomal protein, whose
binding has already been seen to be affected by deletion of
hairpin IIa and IIIc (25). This second part of the study shows
that domain IIId structure is indeed implicated in S9 ribosomal
protein binding and that its binding can also be affected by the
relative disposition and orientation of the major HCV stem–loop
MATERIALS AND METHODS
Plasmid construction of HCV 5′ UTR mutants
The strategy used to swap the positions of different domains of
HCV 5′ UTR using a template sequence has already been
described in detail elsewhere (29). The domain II region
missing from 5′ MscStu (nt 23–102) was amplified and
inserted back in the MscI site of this plasmid (obtaining
plasmid 5′ domII) using two sense and antisense primers (5′-ggg-
cgacactccacc-3′ and 3′ oligo: 5′-aggctgcacgacact-3′). In the
StuI site we then inserted the amplified sequences of domain
III using the following 5′ and 3′ primers: 5′-accccccctcccg-
ggggtctgcgg-3′, 5′-cagtctcgcgggggca-3′ for mutant 5′ domIII
(145–248), 5′-accccccctcccgggaattgccagga-3′, 5′-caatctccag-
gcata-3′ for mutant 5′ domIII (172–227) and 5′-accccccctcccg-
gggaccgggtc-3′, 5′-cgagcgggttgatccaa-3′ for mutant 5′ domIII
(184–213) (see Fig. 1A for a schematic diagram of each
mutant).All these primers contained5′leader sequences aimed
at restoring the nucleotides deleted during the formation of the
MscI and StuI site. All these different mutants were inserted in
the pSV GH bicistronic expression system (31) for transfection
experiments in COS-1 cells as already described (29) and in
the XbaI/HindIII restriction enzymesites of pBluescript II KS+
(Stratagene). An in vitro transcription and translation assay
(Novagen) was performed according to manufacturer’s
instructions using 2 µg of each Bluescript plasmid. Mutant 5′
S/L, 5′ MscStu, and the wild-type 5′ UTR sequence (5′ wt) have
already been described in detail elsewhere (29). Mutants of
subdomain IIId were performed using the following 5′ and 3′sets
of primers: 5′-tagtgttgggtgtgatagccgcttgtggtac-3′, 5′-tcacac-
ccaacactactcggctagcagtctc-3′ for mutant IIId disrupted (5′ domIIId
disr.), 5′-gcgcttgggtgtgatagccgcttgtggtac-3′, 5′-atcacacccaagc-
gctcttccgtagcagtctc-3′ for mutant IIId repaired (5′ domIIId rep.)
and 5′-gcgctgggttgtgatgagccgcttgtggtac-3′, 5′-atcacaacccag-
cgctttccgtagcagtctc-3′ for mutant IIId S/L (5′ domIIId S/L).
These primers were used to amplify and insert back the
mutated domIIId in the wild-type HCV 5′ UTR as described
previously for the 5′ S/L mutant (29).
Transcription of the pBluescript II KS+ plasmids
All plasmids described in this study were linearized by digestion
with HindIII and transcription was performed with T7 RNA
polymerase (Stratagene) in the presence of [α-32P]UTP and
purified on a Nick column (Pharmacia) according to the
manufacturers’ instructions. The labeled RNAs were then
precipitated and resuspended in RNase-free water. The specific
activities oflabeledRNApreparationswere~4 × 106c.p.m./µg of
RNA. Ribosomal salt wash extract from COS-1 and
the binding conditions for the UV-crosslinking assay using
[α-32P]UTP labeled RNA probes (1 × 106c.p.m. per incuba-
tion) have already been described in detail in a previous work
(29). The samples were loaded on an 8 and 12% (as stated in
the figure legends) SDS–PAGE gel which was subsequently
dried and exposed to Kodak X-Omat AR films for 12–24 h.
Films were then scanned on a Macintosh G3 workstation using
Adobe Photoshop and printed using a Phaser 400 printer.
Enzymatic analysis of RNA secondary structure
In order to investigate the secondary structure of the domain III
mutants we used single- and double-strand specific RNases.
RNA was transcribed using T7 RNA polymerase (Stratagene)
from the Bls KS+ domIII (145–248), domIII (172–227),
domIII (184–213), 5′ wt and 5′ S/L mutants with HindIII.
Reaction mixes (100 µl final vol) contained 1 µg of RNA and
0.02 U of RNase V1 (Pharmacia Biotech), or 0.5 U of RNase
T1 (Sigma) in buffer A (10 mM Tris pH 7.5, 10 mM MgCl2,
50 mM KCl), or 20 U of S1 nuclease (Pharmacia Biotech) in
buffer B (buffer A plus 1 mM ZnSO4). The RNA was digested
at 30°C for15min in awaterbath and a control aliquot of RNA
without the addition of RNases was processed simultaneously
with the digested samples. Reactions were stopped by extraction
with phenol/chloroform and ethanol precipitated in 0.3 M
sodium acetate (pH 5.2). The pellet was resuspended in 3 µl of
water and RNase cleavage sites were identified by primer
extension with a domain IV specific end-labeled oligonucleotide
primer (5′-ggtgcacggtctacgaga-3′). In a total volume of 5 µl,
10 ng of32P-labelled primer was hybridized to the resuspended
RNA in RT buffer (50 mM Tris, pH 8.3, 3 mM MgCl2, 75 mM
KCl). The solution was heated at 65°C for 5 min and allowed
to cool for 5 min at room temperature. To each reaction we
then added 15 µl of a solution in RT buffer containing 0.2 U of
M-MLV reverse transcriptase (Gibco BRL), 2 µl of dNTP mix
5 mM and 2 µl of 0.1 M DTT. The mixture was held at 42°C
for 30 min and then 2 µg of RNase A were added and the
mixture incubated for a further 30 min at 37°C. It was then
phenol/chloroform extracted and ethanol precipitated in 0.3 M
sodium acetate (pH 5.2). The samples (enzymatically digested
RNA, control reaction and a sequencing reaction using the
same primer used for the RT reaction) were loaded on a 6%
polyacrylamide denaturing gel (8 M urea, 1× TBE) which was
subsequently run for 1–3 h at 1800 V, dried, and exposed to
Kodak X-Omat AR films for 12–24 h. Films were then
scanned on a Macintosh G3 workstation using Adobe
Photoshop and printed using a Phaser 400 printer.
Assembly and sucrose density gradients of binary IRES–40S
The purification of 40S ribosomal subunits and the assembly
of binary IRES–40S ribosomal complexes were performed
essentially as described by Pestova et al. (32). Briefly, 40S
ribosomal subunits were prepared from HeLa extracts by
precipitation for 4 h at 4°C and 30 000 r.p.m. in a Beckman
60Ti rotor and resuspended in buffer A (20 mM Tris–HCl
pH 7.6,2 mM DTTand6 mM MgCl2) with 0.25M sucrose and
Nucleic Acids Research, 2000, Vol. 28, No. 4
150 mM KCl to a concentrationof 40 A260U/ml. This suspension
was incubated with 1 mM puromycin (Sigma) for 10 min at
0°C and then for 10 min at 37°C before addition of KCl to
0.5 M final concentration. The 40S and 60S ribosomal subunits
were then separated by centrifugation of 2 ml aliquots of this
suspension through a 10–30% sucrose gradient in buffer A
with 0.5 M KCl for 17 h at 4°C and 22 000 r.p.m. using a
Beckman SW28 rotor. The 40S subunits were precipitated
from the pooled gradient fractions by centrifugation for 18 h at
4°C and50000r.p.m. ina 60Ti rotor. Pellets were resuspended
1 mM DTT, 1 mM MgCl2and 0.25 M sucrose) to a final
concentration of 60 A260U/ml. Ribosomal complexes were
assembled by incubating 1 µg of labeled RNAs for 10 min at
30°C in a 100 µl reaction volume that contained buffer E
(2 mM DTT, 100 mM potassium acetate, 20 mM Tris pH 7.6)
with 2.5 mM magnesium acetate, 100 U of RNasin (Promega),
1 mM ATP and 6 pmol of 40S subunits. The complexes were
resolved by centrifugation through a 10–30% sucrose gradient
in buffer E with 6 mM magnesium acetate for 16 h at 4°C and
24 000 r.p.m. using a Beckman SW41 rotor. The radioactivity
of gradient fractions was determined by Cerenkov counting.
Figure 1. (A) Schematic representation of the domain III mutants used in this study. The arrows indicate how each mutant was obtained from the original 5′ wt and
template sequence (5′ MscStu). The three novel mutants contain different regions of domain III: 145–248, 172–227 and 184–213. (B) The IRES activity of each
mutant as assayed in transfection in COS-1 cells. The arrows indicate the position of the processed (23 kDa) and unprocessed (25 kDa) HCV core protein. The
HCV core protein was recognized using MAb B12.F8 (35) and visualized by ECL staining. (C) In vitro transcription and translation assay of all three mutants
including 5′ wt and mutant 5′ S/L. The arrow indicates the unprocessed35S-labeled HCV core protein (visualized by autoradiography).
Nucleic Acids Research, 2000, Vol. 28, No. 4
We have previously shown that an isolated domain III
sequence RNA (nt 134–290) binds two eIF3subunits (29) even
when transcribed apart from the rest of the entire HCV 5′ UTR
sequence. This result is in agreement with a different study
showing that a binary complex between the entire eIF3 complex
and the full HCV IRES sequence protects from footprinting
analysis only stem–loops IIIa, IIIb and IIIc of this domain (26).
Taken together, these data were indicative that the apical
portion of domain III is the only HCV 5′ UTR region involved
with eIF3 binding to the IRES. Therefore, in order to better
define these observations we have prepared three additional
172–227 and 184–213 (Fig. 1A). First of all,consideringthatall
Figure 2. Enzymatic determination of the RNA secondary structure of HCV 5′ UTR domain III. In vitro transcribed RNA was enzymatically digested with S1
nuclease, T1 and V1 RNases and reversely transcribed. The RT products were then separated on a 6% polyacrylamide sequencing gel and a sequencing reaction
performed with the same primer used for the RT reaction was run in parallel to precisely determine the cleavage sites. Squares, circles and triangles indicate S1
nuclease, RNase T1 and V1 cleavage sites, respectively. Black, shaded and white symbols indicate high, medium and low cleavage intensities. The vertical bars
indicate the proposed loop regions of IIIa, IIIb, IIIc and IIId. No enzyme was added to the reaction mixture in lane N. The observed cleavage sites (panel on the
right) are reported on the proposed schematic diagram of domain III (panel on the left). Reported on this panel are also the portions of domain III inserted on the
template sequence to obtain the three novel mutants.
Nucleic Acids Research, 2000, Vol. 28, No. 4
these sequences were inserted back in the exact position of the
original wild-type domain III (nt 134–290) it was interesting to
measure if any of these mutants had retained some IRES
activity. Figure 1B shows that in an in vitro transcription and
translation assay and in a transfection assay in COS-1 cells
using a bicistronic system (Fig. 1C) all three mutants failed to
stimulate IRES activity as estimated by production of a
reporter HCV core protein. It should be noted that transfection
efficiencies in Figure 1C, as measured by growth hormone
production from the first cistron, were all similar,indicatingthat
all RNAs were equally stable. As positive controls we used both
the wild-type sequence (5′ wt) and the previously described
mutant (5′ S/L) in which the apical region sequence (IIIb) had
been inverted without occurring in a loss of IRES activity (29).
It should be noted that in the in vitro assay only the unpro-
cessed form of the core protein can be detected and that in both
assays the molecular weight of this protein is slightly higher
than the HCV native core protein (25 as opposed to 23 kDa) due
to the addition in our constructs of a C-terminal tag sequence
A necessary control to understand the inability of these
mutants todrive IRES translation is represented by the analysis
of their secondary structure (and comparing it with the wild-
type structure). Although the secondary structure ofHCV5′ UTR
validated based on computer predictions and confirmed by
mutational analysis/RNase probing (11,12), we decided to
repeat this analysis in order to obtain a reference pattern to
compare it with that of the mutants. As shown in Figure 2
(right panel) the cleavage data obtained in the domain III
region (nt 134–290) are in very good agreement with the
already existing model (Fig. 2, left panel). In particular, strong
T1 and S1 cleavages (indicating single-stranded regions) can
be detected in correspondence to the apical loop (IIIb) and the
lateral IIIa and IIId loops, with the exception of the proposed
loop IIIc. Interestingly, a prominent S1 cleavage can also be
detected in the lateral bulge which was described to be
protected in footprinting experiments using the entire eIF3
complex (26). Furthermore, all the V1 cleavages (indicating
double-stranded regions) detected in this analysis are entirely
consistent with the proposed stem structures.
The same procedure was then used to map the secondary
structure of each mutant and the results of this analysis are
shown in Figure 3. Only mutant 5′ domIII (145–248) shows an
apical cleavage pattern very similar to that of the wild-type
RNA. Indeed, not only are the single-strand cleavages in the
loop are conserved but also the V1 specific cleavages in the
stem immediately underneath. On the other hand, the apical
loop region in mutants 5′ domIII (172–227) and 5′ domIII
the structure of the 5′ S/L mutant shows that in the IIIb mutated
Figure 3. RNA secondary structure of the apical portion of each domain III mutants with respect to the wild-type and 5′ S/L sequence. In vitro transcribed RNAs
were enzymatically analyzed as described in the legend to Figure 2. Only the apical stem–loop region of domain III is shown and the putative IIIb loop portion is
indicated by the vertical bar. The structure of mutants 5′ domIII (172–227) and 5′ domIII (184–213) are crossed to indicate that the stem–loop configuration is not
consistent with the RNase analysis. In addition this figure shows the RNA secondary structure of domain III of the 5′ S/L mutant (its modified loop IIIb sequence
is indicated by an asterisk).
Nucleic Acids Research, 2000, Vol. 28, No. 4
region the two G residues in the loop IIIb (present in inverted
positions with respect to 5′ wt) were recognized by RNase T1
and that even the mutated stem region presented V1 cleavages,
confirming the maintenance of a stem–loop structure similar
(but not identical) to 5′ wt. Therefore, although the 5′ S/L
mutant presents a very extensive primary sequence mutation
its conformation resembles more the wild-type pattern than the
two 5′ domIII (172–227) and 5′ domIII (184–213) mutants.
It was then interesting to determine how the variation in
secondary structure of these mutants was reflected in their
ability to bind the previously described p170 and p116/p110
subunits. Figure 4A shows the UV-crosslinking assay of each
32P-labeled RNA with a ribosomal salt wash extract from COS-1
cells loaded on an 8% SDS–PAGE gel to better visualize high
molecular weight proteins. This analysis shows that, as
expected, mutant 5′ domIII (145–248) was capable of binding
both p170 and p116/p110 (although with lower efficiency than
5′ wt) whilst no binding could be detected for the other two
mutants 5′ domIII (172–227) and 5′ domIII (184–213). The
fact that mutant 5′ domain III (145–248) showed a structure
and binding pattern identical to that of the wild-type sequence
but was completely incapable of IRES activity made it an ideal
target to identify essential cellular factors that bind to (or
whose binding is affected by) the lower missing portion of
domain III (principally stem–loop IIId) and which could
account for this loss. This possibility was enhanced by the fact
that, in addition to eIF3, ribosomal protein S9 had also been
detected as binding todomainIII (25,26). In particular, binding
of this protein was reported to be reduced by deletions of
hairpins IIa and IIIc but not by deletion of hairpin IIIb (25).
Figure 4B shows a competition analysis performed on the 5′wt
labeled RNAusing aribosomal salt wash extract with5′ wt and
5′ domIII (145–248) cold RNA as competitor (in 2, 5 and 10
molar excess) and loaded on a 12% SDS–PAGE gel to visualize
low molecular weight proteins. In this analysis, a protein of
25 kDa apparent molecular weight, consistent with the
reported S9 ribosomal protein molecular weight observed in
previous studies (25), was found to bind the 5′ wt sequence and
could be competed away by cold 5′ wt RNA but not by cold 5′
domIII (145–248) RNA.
In order to better investigate the binding requirements for
this protein we then prepared a set of mutants (Fig. 5A) which
involved inversion of stem–loop IIId sequence (5′ IIId S/L),
disruption of the principal stem (5′ IIId disr.) and its compen-
mutants these mutants were analyzed for IRES activity both in
vivo (Fig. 5A) and in vitro (Fig. 5B). The results show that
none of these mutants displayed IRES activity, including the
one bearing the intended compensatory mutation.
Analysis of the secondary structure of each mutant is shown
in Figure 6 and in keeping with the loss in IRES activity both
repaired mutant fail to mimic that of the wild-type sequence or
the 5′ S/L mutant. The fact that even the supposedly repaired
mutant fails to fold again correctly highlights the fact that it is
very difficult to predict beforehand the effect of engineered
mutations. Figure 6 also shows the secondary structure of the
IIId stem–loop of mutant 5′ S/L and that it is identical to that of
5′ wt (although this was an expected result considering that the
engineered mutation was located in the apical IIIb stem–loop).
The results of the structural analysis were reflected in the
UV-crosslinkingassay,as showninFigure7:only5′wt and5′S/L
bind the 25 kDa protein, whilst none of the IIId mutants is
capable of doing so. Binding of S9 to the 5′ S/L (whose mutation
involved only hairpin IIIb) is in keeping with published data
that report a reduction of p25 UV crosslinking following deletion
of hairpins IIa and IIIc but not by deletion of hairpin IIIb (25).
Our results thus provide a likely reason for the loss in IRES
activity experienced by the correctly folded 5′ domIII (145–248)
mutant and also report for the first time that stem–loop IIId
affects the binding of S9 ribosomal protein to HCV 5′ UTR.
The fact that distant stem–loops (IIa, IIIc and in the present
work IIId) have been found to affect S9 ribosomal protein
binding suggests that, unlike p170 and p116/p110, the binding
requirements for this protein might not be present on a single
domain. A simple experiment to test this hypothesis was to see
if S9 ribosomal protein binding can still be detected on a HCV
5′ UTR in which the sequences of hairpins IIa, IIIc and IIId are
present (and unmutated) but in different orientations with
respect to the wild-type. Therefore, we used the mutants
already described in a previous paper in which the position of
domain II and domain III on the HCV 5′ UTR had been
inverted (Fig. 8A) and which did not possess any IRES
activity. The results ofthe UV-crosslinking analysis performed
on these mutants (Fig. 8B) shows that only the wild-type
sequence (5′ wt) can bind S9 whilst mutants 5′ MscStu domII,
5′ MscStu domIII and 5′MscStu domIII/domII do not bind this
protein. It is important to note that both 5′ MscStu domIII and
5′ MscStu domIII/domII have already been reported (29) to
bind both p170 and p116/p110, and that a secondary structure
analysis of their domain IIId structure showed that this hairpin
presented a cleavage pattern identical to that of the wild-type
structure (data not shown).
Figure 4. UV-crosslinking analysis used to identify proteins binding to the
wild-type HCV RNA and the domain III mutants. (A) UV-crosslinking assay
(loaded on an 8% SDS–PAGE gel) using a COS-1 ribosomal salt wash extract
with all the mutant UTRs carrying different portions of domain III. The arrow
indicates the position of p170 and p116/p110. To better visualize the binding
of these subunits we used 5 µg of cold 5′ MscStu template RNA as competitor
(which had already been reported not to bind p170 and p116/p110). (B) Com-
petition analysis (loaded on a 12% SDS–PAGE gel) of the proteins bound by
labeled 5′ wt with cold 5′ wt and 5′ domIII (145–248) RNA. The arrow
indicates the 25 kDa protein that binds labeled 5′ wt and can be competed by
cold 5′ wt but which cannot be competed by cold 5′ domIII (145–248) RNA.
Nucleic Acids Research, 2000, Vol. 28, No. 4
Finally, the failure to detect UV crosslinking of the S9
protein in our previous mutants could be due to two reasons.
does not actually bind to the mutant RNAs. Alternatively, the
mutant RNAs might be able to bind to the 40S ribosomal
subunit but in a different orientation compared to the wild-type
RNA. In order to differentiate between these two possibilities
we performed sucrose density gradient analysis of the wild-type
RNA and of representative mutants in the presence of purified
40S subunits. Previous studies have already shown that in
order to obtain binding of the 40S ribosomal subunit to the
HCV IRES no additional initiation factors are required (25).
The results shown in Figure 9 demonstrate that a ribosomal
complex can only be detected with the wild-type HCV IRES,
Figure 5. (A) Schematic representation of the stem–loop IIId mutants 5′ domIIId (disr.), 5′ domIIId (rep.) and 5′ domIIId S/L. (B) Transfection in COS-1 cells of
these mutant 5′ UTRs and the arrows indicate the position of the processed (23 kDa) and unprocessed (25 kDa) HCV core protein. The HCV core protein was
recognized using MAb B12.F8 (35) and visualized by ECL staining. (C) In vitro transcription and translation assay of all mutants. The arrow indicates the unprocessed
35S-labelled HCV core protein (visualized by autoradiography).
Nucleic Acids Research, 2000, Vol. 28, No. 4
whilst mutant 5′ domIII (145–248) and mutant 5′ IIId rep. (see
Figs 1 and 5, respectively) do not bind to the 40S ribosomal
Functional and mutational studies performed onthe HCV 5′UTR
region coupled with data obtained from UV crosslinking and
structural analyses have allowed us to gain substantial insight
concerning HCV IRES function, as recently reviewed in (10). In
particular, several cellular factors have been found to specifically
bind its structure and, in this respect, the HCV IRES has been
recently shown to share a similarity with the mechanism of
translation initiation in prokaryotes (25). In this study we have
further characterized the binding to the HCV 5′ UTR of two
subunits of eukaryotic initiation factor eIF3 and of the ribosomal
protein S9 with regards to conservation of RNA secondary
structure. In addition, we have investigated the possible
influence of RNA tertiary folding on their binding, providing
direct evidence that it plays a role in the binding of S9 ribo-
somal protein to the HCV IRES.
The effect of secondary structure on the binding of the eIF3
subunits has already been preliminarily analyzed in a previous
study in which we showed that binding could still occur in an
Figure 6. RNA secondary structure of the stem–loop IIId mutants with respect to the wild-type sequence 5′ wt and the 5′ S/L mutant. In vitro transcribed RNAs
were enzymatically analyzed as described in the legend to Figure 2. Only the lower region of domain III is shown and the putative IIId loop is indicated by a vertical
bar. The structure of mutants 5′ domIIId (disr.), 5′ domIIId (rep.) and 5′ domIIId S/L are crossed to indicate that the original stem–loop configuration is not
consistent with the RNase analysis.
Figure 7. UV-crosslinking assay (loaded on a 12% SDS–PAGE gel) using a
COS-1 ribosomal salt wash extract with all the mutant UTRs carrying different
mutations in stem–loop IIId. Binding of the 25 kDa protein (indicated by an
arrow) could be detected only for the wild-type sequence (5′ wt) and for mutant
and 5′ domIIId S/L mutant. To better visualize this binding we used 5 µg of
cold 5′ MscStu RNA as competitor.
Nucleic Acids Research, 2000, Vol. 28, No. 4
isolated domain III sequence and that it could be impaired by
mutations which were predicted to disrupt the RNA folding of
the apical stem–loop (29). In this study we report that binding of
can be achieved even when significant lower portions of
is provided by whether or not the apical stem–loop RNA
sequence is able to fold in the correct secondary structure. In
fact, RNase mapping of the 5′ UTR secondary structure shows
that nt 145–248 can fold autonomously in the correct RNA
conformation and can then bind both subunits: an observation
that definitively identifies this region as possessing the minimal
sequence and structural features necessary for recognition of
these subunits. This result is in agreement with the footprinting
data previously published concerning an eIF3–HCV IRES
binary complex which did not detect any protection in the
HCV IRES outside the apical domain III region (26). It is also
of particular interest the observation that mutants which
contain sizeable portions of the apical stem–loop sequences
previously identified as important for this interaction (nt 172–227
and 184–213) were not able to bind either p170 or p116/p110.
This observation was not totally expected if we consider the
fact that the previously described 5′ S/L mutant (in which the
sequence of the apical stem–loop was inverted) had retained
the ability to bind p170 but had lost the ability to bind p116/
p110 (29). Therefore, if the recognition of p170 and p116/p110
had been predominantly dependent on the primary sequence
some binding could still have taken place in the two 5′ domIII
(172–227) and 5′ domIII (184–213) mutants. The fact that no
binding occurred finds an explanation in the secondary structure
of these mutants which, as opposed to the one of mutant 5′ S/L,
revealed that in their case the structure of the apical stem–loop
had not been preserved. On the other hand, the structure of the
5′ S/L mutant hadconserved an unexpected similarity (considering
the extent of the primary nucleotide changes) to the wild-type
conformation. Taken together, these results show that conser-
vation of secondary structure is a fundamental necessity to
obtain protein binding and that it possesses just as great an
importance as the primary nucleotide sequence. In particular,
our results show that conserving one without the other
(irrespectively) is not sufficient to obtain correct binding.
The fact that the correctly folded 5′ domIII (145–248)
mutant was not capable of IRES translation (although its
binding and structural profile are most similar to those of 5′ wt,
even more so than those observed for the 5′ S/L mutant) was
also indicative that the missing lower sequence is probably
involved in the binding of some other essential cellular factor.
ribosomal protein S9 (25) binding to HCV domain III. In
accordance with this study we have identified a 25 kDa protein,
whose molecular weight is consistent with the molecular weight
ofthe S9ribosomal protein, that binds tothe 5′wtsequencebut
which cannot be competed away by the 5′ domIII (145–248)
Figure 8. (A) Mutated UTRs used in the part of the study concerning the effect of swapping the positions of domain II and III on the binding of protein S9. How
the different mutants were obtained from the original 5′ wt sequence is described in detail elsewhere (29). Major structural domains are indicated by Roman numerals
I–IV. (B) UV-crosslinking assay (loaded on a 12% gel SDS–PAGE) using a COS-1 ribosomal salt wash extract with all the mutant UTRs carrying different combinations of
domain II or III in swapped positions. The arrow indicates the 25 kDa protein observed to bind only to 5′ wt. No binding could be observed with 5′ MscStu domII,
5′ MscStu domIII and 5′ MscStu domIII/domII.
Nucleic Acids Research, 2000, Vol. 28, No. 4
An increasing body of evidence shows that the majority of
take place. Up to now, hairpins IIIb (29), IIIc (15,25), and
recently IIIe (18), have been demonstrated to be important for
efficient IRES activity. Therefore, we investigated whether the
integrity of hairpin IIId was also important for IRES activity
and whether its disruption affected S9 ribosomal protein
binding. Our results show that mutations of hairpin IIId which
cause both sequence and structural changes lead to undetectable
levels of IRES activity in conjunction with the loss of S9
protein binding. The analysis of IRES–40S binary ribosomal
complex showed that mutant 5′ domIII (145–248) and mutant
5′ IIId rep. do not bind to the 40S ribosomal subunit, thus
providing a functional explanation for the lack of S9 binding.
Notably, it is also indicative that mutant 5′ S/L, which was
observed to be capable of IRES translation, showed a IIId
cleavage pattern similar to that of the wild-type sequence and
can bind S9, providing an additional explanation of why this
mutation did not affect the translational activity.
The fact that bindingof the S9 ribosomal protein was already
reported to be affected by deletion of hairpins IIa, IIIc (25) and
IIId (this paper) suggested that the binding requirements for S9
are also extended to the tertiary folding of the RNA structure.
In this respect, using small angle X-ray crystallography Kieft
et al.haveshown that the HCV IRES possesses indeeda higher
order structure of spatially organized recognition domains in
the absence of cellular factors (34). Therefore, the S9 interaction
with the tertiary configuration was further analyzed by studying
the binding pattern in mutants already used in a previous work
where the position of wild-type domain II and domain III had
been inverted on a 5′ UTR template without introducing mutations
in their primary sequence (29). Our analysis shows that the
simple change of orientation of these domains with conservation
of sequence and RNA structure can abrogate binding of S9.
domain III on the 5′ UTR had no effect on p170 and p116/p110
binding (29), highlighting a clear difference in the binding
requirements between the eIF3 and S9 cellular factors. In
that of a highly organized 3D structure where the binding of
proteins required for translation initiation needs not only an
intact domain structure but also correct spatial configuration of
evolved by the HCV IRES to bind essential cellular factors.
1. Choo,Q.L., Kuo,G., Weiner,A.J., Overby,L.R., Bradley,D.W. and
Houghton,M. (1989) Science, 244, 359–362.
2. Kamoshita,N., Tsukiyama-Kohara,K., Kohara,M. and Nomoto,A. (1997)
Virology, 233, 9–18.
3. Tsukiyama-Kohara,K., Iizuka,N., Kohara,M. and Nomoto,A. (1992)
J. Virol., 66, 1476–1483.
4. Wang,C., Sarnow,P. and Siddiqui,A. (1993) J. Virol., 67, 3338–3344.
5. Wang,C., Sarnow,P. and Siddiqui,A. (1994) J. Virol., 68, 7301–7307.
6. Wang,C., Le,S.Y., Ali,N. and Siddiqui,A. (1995) RNA, 1, 526–537.
7. Fukushi,S., Katayama,K., Kurihara,C., Ishiyama,N., Hoshino,F.B.,
Ando,T. and Oya,A. (1994) Biochem. Biophys. Res. Commun., 199,
8. Rijnbrand,R., Bredenbeek,P., van der Straaten,T., Whetter,L.,
Inchauspe,G., Lemon,S. and Spaan,W. (1995) FEBS Lett., 365, 115–119.
9. Reynolds,J.E., Kaminski,A., Kettinen,H.J., Grace,K., Clarke,B.E.,
Carroll,A.R., Rowlands,D.J. and Jackson,R.J. (1995) EMBO J., 14,
10. Hellen,C.U.T. and Pestova,T.V. (1999) J. Viral Hepatitis, 6, 79–87.
11. Brown,E.A., Zhang,H., Ping,L.H. and Lemon,S.M. (1992)
Nucleic Acids Res., 20, 5041–5045.
12. Honda,M., Brown,E.A. and Lemon,S.M. (1996) RNA, 2, 955–968.
13. Buratti,E., Gerotto,M., Pontisso,P., Alberti,A., Tisminetzky,S.G. and
Baralle,F.E. (1997) FEBS Lett., 411, 275–280.
14. Fukushi,S., Kurihara,C., Ishiyama,N., Hoshino,F.B., Oya,A. and
Katayama,K. (1997) J. Virol., 71, 1662–1666.
15. Tang,S., Collier,A.J. and Elliott,R.M. (1999) J. Virol., 73, 2359–2364.
16. Honda,M., Beard,M.R., Ping,L.H. and Lemon,S.M. (1999) J. Virol., 73,
17. Hwang,L.H., Hsieh,C.L., Yen,A., Chung,Y.L. and Chen,D.S. (1998)
Biochem. Biophys. Res. Commun., 252, 455–460.
18. Psaridi,L., Georgopoulou,U., Varaklioti,A. and Mavromara,P. (1999)
FEBS Lett., 453, 49–53.
19. Varaklioti,A., Georgopoulou,U., Kakkanas,A., Psaridi,L., Serwe,M.,
Caselmann,W.H. and Mavromara,P. (1998) Biochem. Biophys. Res.
Commun., 253, 678–685.
Figure 9. Binary IRES–40S ribosomal complex formation on HCV wild-type
IRES and selected mutants. Assays were performed on a 10–30% sucrose
density gradient with labeled RNAs and purified 40S ribosomal subunits.
Assays were done using 5′ wt (A), 5′ domIII (145–248) (B) and 5′ IIId (rep)
(C) mutant RNAs. Sedimentation was from right to left. The position of the
binary complexes is indicated by an arrow.
Nucleic Acids Research, 2000, Vol. 28, No. 4 Download full-text
20. Honda,M., Rijnbrand,R., Abell,G., Kim,D. and Lemon,S.M. (1999)
J. Virol., 73, 4941–4951.
21. Ali,N. and Siddiqui,A. (1997) Proc. Natl Acad. Sci. USA, 94, 2249–2254.
22. Kaminski,A., Hunt,S.L., Patton,J.G. and Jackson,R.J. (1995) RNA, 1,
23. Ali,N. and Siddiqui,A. (1995) J. Virol., 69, 6367–6375.
24. Ito,T. and Lai,M.M. (1999) Virology, 254, 288–296.
25. Pestova,T.V., Shatsky,I.N., Fletcher,S.P., Jackson,R.J. and Hellen,C.U.
(1998) Genes Dev., 12, 67–83.
26. Sizova,D.V., Kolupaeva,V.G., Pestova,T.V., Shatsky,I.N. and
Hellen,C.U. (1998) J. Virol., 72, 4775–4782.
27. Hahm,B., Kim,Y.K., Kim,J.H., Kim,T.Y. and Jang,S.K. (1998) J. Virol.,
28. Spangberg,K. and Schwartz,S. (1999) J. Gen. Virol., 80, 1371–1376.
29. Buratti,E., Tisminetzky,S., Zotti,M. and Baralle,F.E. (1998)
Nucleic Acids Res., 26, 3179–3187.
30. Kaminski,A. and Jackson,R.J. (1998) RNA, 4, 626–638.
31. Nicholson,R., Pelletier,J., Le,S.Y. and Sonenberg,N. (1991) J. Virol., 65,
32. Pestova,T.V., Hellen,C.U. and Shatsky,I.N. (1996) Mol. Cell. Biol., 16,
33. Buratti,E., Baralle,F.E. and Tisminetzky,S.G. (1998) Cell. Mol. Biol.
(Noisy-le-grand), 44, 505–512.
34. Kieft,J.S., Zhou,K., Jubin,R., Murray,M.G., Lau,J.Y. and Doudna,J.A.
(1999) J. Mol. Biol., 292, 513–529.
35. Cerino,A., Boender,P., La Monica,N., Rosa,C., Habets,W. and
Mondelli,M.U. (1993) J. Immunol., 151, 7005–7015.