Competing and noncompeting activities of miR-122
and the 5′ exonuclease Xrn1 in regulation of
hepatitis C virus replication
You Lia, Takahiro Masakia, Daisuke Yamanea, David R. McGiverna,b, and Stanley M. Lemona,b,c,1
aLineberger Comprehensive Cancer Center andbDivision of Infectious Diseases, Department of Medicine andcDepartment of Microbiology and Immunology,
University of North Carolina at Chapel Hill, Chapel Hill, NC 27599-7292
Edited by Francis V. Chisari, The Scripps Research Institute, La Jolla, CA, and approved November 19, 2012 (received for review August 3, 2012)
Hepatitis C virus (HCV) replication is dependent on microRNA 122
(miR-122), a liver-specific microRNA that recruits Argonaute 2 to the
cell-free reactions and in infected cells. Here we describe the RNA
degradation pathways against which miR-122 provides protection.
Transfected HCV RNA is degraded by both the 5′ exonuclease Xrn1
and 3′ exonuclease exosome complex, whereas replicating RNA
within infected cells is degraded primarily by Xrn1 with no contri-
bution from the exosome. Consistent with this, sequencing of the
5′ and 3′ ends of RNA degradation intermediates in infected cells
confirmed that 5′ decay is the primary pathway for HCV RNA deg-
radation. Xrn1 knockdown enhances HCV replication, indicating
that Xrn1 decay and the viral replicase compete to set RNA abun-
dance within infected cells. Xrn1 knockdown and miR-122 supple-
mentation have equal, redundant, and nonadditive effects on the
rate of viral RNA decay, indicating that miR-122 protects HCV RNA
from 5′ decay. Nevertheless, Xrn1 knockdown does not rescue rep-
lication of a viral mutant defective in miR-122 binding, indicating
that miR-122 has additional yet uncharacterized function(s) in the
viral life cycle.
host factor|RNA decay|translation|viral replicase
an important cause of human liver disease (1). Its replication is
uniquely dependent on miR-122, which is the most abundant
microRNA (miRNA) in the liver and accounts for >50% of ma-
ture miRNAs in human hepatocytes (2, 3). There are two con-
internal ribosome entry site (IRES) that mediates translation of
the viral polyprotein. Direct interactions between the miR-122
seed sequence (nts 2–8) and these sites in the5′ UTR areessential
for amplification of the HCV genome (4, 5). Additional supple-
mental base-pairing upstream of S1 and S2 has also been dem-
onstrated and is important for viral replication (6, 7).
binding within the 3′ UTR and promote translational repression
and/or destabilization of the target RNA (8), binding of miR-122
to the 5′ UTR of HCV genomic RNA stimulates viral protein ex-
stabilizes HCV RNA (10). The rate of decay of either transfected
synthetic genomic RNA or replicating viral RNA within infected
122, thereby supplementing its endogenous abundance. Con-
enhances the rate with which HCV RNA decays in either context.
HCV RNA is not thought to contain a 5′ cap structure, and the
stabilizing action of miR-122 on synthetic RNA can be function-
ally substituted by addition of a 5′ cap analog (10). These obser-
vations point to the importance of RNA decay pathways in HCV
replication and suggest that miR-122 is likely to prevent degra-
dation of viral RNA from its 5′ end. However, the specific
epatitis C virus (HCV) is a positive-strand RNA virus classi-
fiedin the family Flaviviridae. It is highly hepatotropic and
mechanisms involved in degradation of HCV RNA, or for that
matter most positive-strand viral RNAs, have not been well
studied and remain unclear. In addition, exactly how miR-122
stabilizes the viral RNA remains to be elucidated.
Because HCV genomicRNA is positivesense and servesdirectly
as the mRNA for viral protein translation, it is reasonable to
speculate that cellular mRNA decay pathways may also function in
degradation of the viral RNA. In eukaryotic cells, bulk mRNA
decay typically initiates with deadenylation, shortening the 3′ poly
(A) tail, followed by degradation of the RNA molecule in a 5′→3′
or 3′→5′ direction (11). In the 5′ decay pathway, the monomethyl
(12, 13), exposing the 5′ monophosphorylated product to pro-
gressive 5′→3′ exoribonucleolytic degradation by Xrn1 (14, 15). In
the 3′ decay pathway, deadenylated mRNA is degraded by the cy-
toplasmic RNA exosome complex, a multisubunit 3′→5′ exoribo-
nuclease (16). The residual cap structure resulting from 3′ decay is
HCV genomicRNA contains neithera 3′poly(A) tailnor5′cap,its
degradation requires neither deadenylation nor decapping. None-
theless, either or both of these two exoribonucleolytic pathways
may potentially contribute to decay of HCV RNA in infected cells.
Here, we describe the roles played by Xrn1 and the exosome
complex in HCV RNA decay. We show that viral RNA is de-
graded specificallyby Xrn1 within infectedcells. Ourresults reveal
that miR-122 protects the genome against Xrn1-mediated decay
but that it has additional functions beyond genome stabilization
that are essential for viral replication.
Xrn1 and the Exosome both Mediate Degradation of Transfected HCV
contribute to degradation of positive-strand HCV RNA, we ex-
amined the decay of HCV RNA transfected into HeLa cells that
do not express endogenous miR-122. To assess the role of the 5′
exonuclease Xrn1 (XRN1), we reduced its expression by prior
transfection of Xrn1-specific siRNA. We similarly knocked down
expression of Upf1 (UPF1), which is a key factor in nonsense-
mediated mRNA decay (18) and is also involved in degradation of
histone mRNA and HIV RNA metabolism (19, 20). Immunoblots
confirmed efficient knockdown of both RNA decay factors (Fig.
which contains a lethal mutation in its RNA polymerase) (10) was
Author contributions: Y.L., T.M., D.Y., and S.M.L. designed research; Y.L., T.M., and D.Y.
performed research; Y.L., T.M., D.Y., D.R.M., and S.M.L. analyzed data; D.R.M. contributed
new reagents/analytic tools; and Y.L. and S.M.L. wrote the paper.
The authors declare no conflict of interest.
This article is a PNAS Direct Submission.
See Commentary on page 1571.
1To whom correspondence should be addressed. E-mail: firstname.lastname@example.org.
This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.
| January 29, 2013
| vol. 110
| no. 5
electroporated into the cells with or without coelectroporation of
duplex miR-122. This strategy eliminates any potentially con-
founding effects of replication on measurements of RNA stability.
HCV RNA levels were determined by quantitative real-time RT-
PCR (qRT-PCR) and normalized to β-actin mRNA. As shown in
Fig. 1B, the transfected HCV RNA degraded rapidly, with a t1/2of
1.4 h in cells transfected with a scrambled control siRNA (siCtrl)
(Table S1). Knocking down Xrn1 significantly stabilized HCV
RNA, increasing the t1/2to 3.2 h, whereas Upf1 knockdown had
little effect on its decay (t1/2= 1.7 h) (Fig. 1B). As expected (10),
coelectroporation of miR-122 significantly stabilized the RNA,
leadingtoa t1/2of2.3and2.7hinthe siCtrlandsiUpf1-transfected
cells, respectively. miR-122 supplementation resulted in a pro-
portionately similar increase in RNA stability after Xrn1 knock-
down, boosting the t1/2from 3.2 to 4.9 h. Maximum stabilization of
HCV RNA was observed with combined Xrn1 knockdown and
miR-122 supplementation (Fig. 1B). Collectively, these results
indicate that Xrn1, but not Upf1, contributes to degradation of
transfected HCV RNA.
To determine whether the exosome is also involved in degrada-
tion of transfected RNA, we knocked down two exosome compo-
nents, Rrp41 (EXOSC4), a core structural component of the
exosome, and PM/Scl-100 (EXOSC10), a 3′→5′ exonuclease, using
with control lentivirus (shCtrl) than with siCtrl (t1/2= 2.1 vs. 1.4 h)
(Table S1), reflecting different conditions in these experiments
either exosome component substantially stabilized the RNA (Fig.
1C; Table S1), indicating that the exosome is also involved in decay
of transfected viral RNA. As with Xrn1, miR-122 supplementation
further increased the stability of the transfected HCV RNA in the
PM/Scl-100 and Rrp41 knockdown cells. We conclude from these
results that both Xrn1 and the exosome contribute to degrada-
tion of transfected HCV RNA.
Degradation of HCV RNA in Cell-Free S10 Lysate Is Primarily Mediated
by the Exosome. We previously demonstrated that miR-122 sta-
bilizes a chimeric poliovirus RNA that contains the HCV 5′ UTR
in lieu of the poliovirus 5′UTR when added with it to HeLa S10
lysate (10). Using SYTO 62 infrared fluorescence to quantify the
(H77S) (21) is similarly stabilized by miR-122 when incubated in
S10 lysate. HCV RNA degrades rapidly in HeLa S10 lysate (Fig.
2B). However, prior addition of duplex miR-122, but not miR-124
(a brain-specific miRNA), to the lysate doubled the half-life of the
RNA (t1/2increasing from 18.6 to 39 min; Table S2). In contrast, a
related viral RNA mutant with single base substitutions in S1 and
S2 that ablate miR-122 binding (H77S/S1-S2p6m; Fig. 2A) (4) was
not stabilized by miR-122, but was stabilized by the complemen-
tary miR-122 mutant (miR-122p6) (Fig. 2C; Table S2), thereby
confirming that HCV RNA is physically stabilized in S10 lysate as
of the capped RNA was further enhanced by miR-122 supple-
mentation (Fig. S1C).
To investigate how HCV RNA is degraded in this cell-free
system, we carried out similar assays using S10 lysates from Xrn1
and exosome knockdown cells. In contrast to what we observed
with transfected RNA, Xrn1 knockdown minimally affected the
HCV RNA decay rate (Fig. 2D; Table S3), whereas knocking
down either of the exosome components, Rrp41 and PM/Scl-100,
slowed degradation of HCV RNA substantially (Fig. 2E). Thus,
the 3′ exosome-mediated decay pathway is mainly responsible for
degradation of HCV RNA in HeLa S10 lysate. This finding is
5′ end (Fig. 2 A and B) and is considered further in Discussion.
Degradation of Replicating HCV RNA Is Mediated by Xrn1. The
experiments described above examined decay of synthetic HCV
RNA after transfectionintocellsorwhenadded tocell-free lysates
and may not recapitulate the RNA decay pathways acting on
replicating viral genomes. To assess this, we treated Huh-7.5 cells
infected with HCV (H77S.3 virus) with PSI-6130, a potent and
specific nucleoside inhibitor of the HCV NS5B RNA-dependent
RNA polymerase that blocks viral RNA synthesis (22). As ob-
served previously following the arrest of new viral RNA synthesis
(10), replicating RNA degraded much more slowly than trans-
fected RNA (t1/2of 10.6 h in siCtrl-transfected cells) (Fig. 3A, Left,
compare with Fig. 1A). Knocking down Xrn1 slowed this rate of
down the exosome component PM/Scl-100 had no effect (t1/2of
10.8 h) (Fig. 3 A, Right, and B). Thus, replicating HCV RNA is
degraded by the 5′ exonuclease Xrn1, but not the exosome com-
plex. The viral RNA was significantly stabilized by miR-122 sup-
plementation in siCtrl and PM/Scl-100 knockdown cells, reaching
a half-life approximating that in the Xrn1 knockdown cells (t1/2=
in no additional increase in RNA stability in the Xrn1 knockdown
cells (t1/2of 19.2 h). Thus, Xrn1 knockdown and miR-122 sup-
plementation have equal and redundant effects on stability of the
RNA, from which we infer that miR-122 enhances HCV RNA
stability by protecting it from Xrn1-mediated 5′→3′ degradation.
Xrn1 and Upf1 in HeLa cells following siRNA transfection and (Right) Rrp41
and PmScl100 in HeLa cells stably expressing the indicated shRNA. β-actin was
a loadingcontrol. (B) HeLa cells were transfected with theindicatedsiRNAsfor
48 h and electroporated with replication-deficient genotype 1a H77S-AAG
RNA with or without miR-122 (1 μM). The percentage of HCV RNA remaining
at each time point following electroporation without (Left) and with (Right)
miR-122 supplementation was determined by qRT-PCR, relative to the abun-
dance of β-actin mRNA. Results shown represent the means of three replicate
experiments ± SEM. (C) HeLa cells expressing the indicated shRNAs were
electroporated as in B. Percent HCV RNA remaining following electroporation
without (Left) and with (Right) miR-122 supplementation was quantified by
qRT-PCR relative to the abundance of β-actin mRNA. Results shown represent
the means of three replicate experiments ± SEM.
Decay of transfected HCVRNA in HeLacells. (A) Immunoblots of (Left)
| www.pnas.org/cgi/doi/10.1073/pnas.1213515110Li et al.
We also assessed the effect of miR-122 on HCV protein ex-
pression under these conditions. For this, we measured Gaussia
princeps luciferase (GLuc) secreted into the media by cells sup-
porting replication of a viral RNA (H77S.3/GLuc2A) that expresses
h intervals after the arrest of viral RNA synthesis with PSI-6130,
declined with a t1/2of about 8 h (Fig. 3C). miR-122 supplementation
resulted in a small but reproducible increase in GLuc production in
siCtrl-transfected cells (Fig.3C, Left),consistent with stabilization of
(Fig. 3C, Right), indicating that miR-122 supplementation does not
enhance viral protein translation under these conditions.
sites of mRNA storage and decay (24). We previously demon-
strated that double-stranded RNA (dsRNA), an intermediate in
the HCV replication cycle, is not localized to P bodies, the mor-
phology of which is well preserved within infected cells with only
a minimal reduction in their number (25). To ascertain whether
single-stranded positive-sense viral RNA colocalizes with Xrn1,
we used a sensitive FISH method to detect HCV RNA in infected
cells, counterstaining with antibody specific for Xrn1 (Fig. 3D).
FISH confirmed the absence of HCV RNA in P bodies. A
quantitative pixel analysis of confocal microscopic images
revealed0.67±0.35%SDoverlapof theRNA signalwith Xrn1 in
P bodies vs. 22.3 ± 5.4% SD overlap with Xrn1 in the cytosol.
Thus, although HCV RNA localizing to P bodies could be de-
graded by Xrn1 (and hence not detectable by FISH), it is possible
that HCV RNA is degraded by cytosolic Xrn1. Importantly, we
observed no differences in the cellular localization of Xrn1 in
infected vs. uninfected cells. Similar studies using an antibody
specific for the decapping factor DCP1a as a marker of P bodies
also showed no association of HCV RNA with P bodies or
changes in DCP1a localization (Fig. S2).
Identification of HCV RNA Degradation Intermediates. To directly
identifyHCVRNAsthathavebeenpartially degraded,we adapted
a circularization RT-PCR (cRT-PCR) strategy to capture HCV
RNAs with 5′ monophosphate (5′P). HCV RNAs with 5′P can be
ligated with the 3′OH to form a circular RNA, and the region
containing 5′ and 3′ ends is subsequently amplified by RT-PCR
(Fig. S3A). HCV RNAs with 5′ triphosphate are incompetent for
ligation and can only be detected if treated first with RNA poly-
phosphatase. We applied this method to total RNA isolated from
Huh-7 cells stably infected with HJ3-5 virus (Fig. S3B). Full-length
HCV RNA with intact 5′ and 3′ ends was identified only after
polyphosphatase treatment (Fig. S3C), confirming that the RNA
possesses a 5′ triphosphate RNA in cells. Importantly, without
pretreatment with RNA polyphosphatase, we could identify HCV
RNA degradation intermediates containing 5′P (Fig. S3D). When
amplified and sequenced, these intermediates all contained trun-
cated 5′ ends while retaining intact 3′ ends. This finding provides
degradation in cells and is consistent with Xrn1 being the major
contributor to degradation of replicating HCV RNA.
Xrn1 Knockdown Enhances HCV Replication. We next asked whether
the increased HCV RNA stability in Xrn1 knockdown cells would
decay pathway competes with viral RNA synthetic machinery to
set the level to which HCV RNA replicates in cells. To test this
hypothesis, we transfected Huh-7.5 cells with siRNAs specific for
Xrn1 or PM/Scl-100, resulting in efficient knockdown of these
proteins (Fig. 4A). The cells were then retransfected with H77S.3/
GLuc2A (23) RNA, and secreted GLuc activity was assessed
thereafter as a measure of its replication. Xrn1 knockdown re-
sulted in a twofold increase in GLuc activity at 48–72 h compared
with control siRNA, whereas PM/Scl-100 knockdown had no ef-
fect (Fig. 4B). Intracellular HCV RNA was similarly increased in
Xrn1 knockdown cells (Fig. 4C and Fig. S4), as was the abundance
of HCV core protein (Fig. 4A). Xrn1 knockdown also increased
the yield of infectious virus released into supernatant fluids (Fig.
4D). Collectively, these data show that Xrn1-mediated decay
the level of viral RNA in infected cells. Previous efforts to in-
vestigate how Xrn1 influences HCV replication have produced
conflicting results, with two studies showing no effects of Xrn1
depletion (26, 27). Our results are consistent with those of Jones
et al. (28) and Ruggieri et al. (29) and provide a mechanism for
why Xrn1 depletion promotes HCV replication.
We next determined whether miR-122 supplementation en-
hances HCV replication in Huh-7.5 cells following knockdown of
Xrn1. If miR-122 acts only to stabilize theviral RNA, we reasoned
that there shouldberelatively littleenhancementof replication, as
Xrn1 knockdown was as effective as miR-122 supplementation in
stabilizing a nonreplicating viral RNA (Fig. 3 A and B). We tested
this hypothesis by supplementing cells with miR-122 before
mutant miR-122p6 guide strand sequence; (Lower) 5′ terminal sequence of
HCV RNA (H77S.3/AAG) with S1 and S2 miR-122 seed sequence-binding sites
underlined. Point mutations (*) in the related S1-S2p6m mutant are shown
above. (B) H77S.3 RNA was incubated with HeLa S10 lysate containing the
indicated duplex miRNA (1 μM). RNAs were extracted at indicated time
intervals, stained with SYTO 62, and resolved in 1% agarose. Percent HCV
RNA remaining was quantified by the Odyssey Infrared Imaging System
relative to the 28S rRNA. Results are the means of three experiments ± SEM.
(C) Decay assays were carried out as in B with the H77S.3/S1-S2p6m mutant
RNA. (D and E) Decay assays were carried out as in B with lysates from (D)
HeLa cells transfected with control or Xrn1 siRNA (mean ± SEM from two
replicate experiments) or (E) lysates from HeLa cells stably expressing the
indicated shRNA (mean ± SEM from three replicate experiments).
Decay of HCV RNA in HeLa S10 lysate. (A) (Upper) miR-122 and
Li et al.PNAS
| January 29, 2013
| vol. 110
| no. 5
transfection of the H77S/GLuc2A RNA. Although Xrn1 knock-
down diminished the magnitude of the effect, miR-122 still effec-
tively boosted HCV replication, as determined by measurements of
GLuc expression (Fig. 4E, compare Left and Right) and viral
RNA abundance (Fig. 4F). To confirm that this was not due to
concentrations of siXrn1. This resulted in increasing depletion of
Xrn1, which was near maximal at 20 nM (Fig. 4G, Upper) and was
matched by increasing HCV RNA replication (GLuc expres-
sion), which plateaued at the same siXrn1 concentration (Fig.
4G, Lower). Despite this evidence for maximal knockdown of
Xrn1 expression, miR-122 supplementation resulted in a greater
increase in HCV replication (Fig. 4G, Lower). This increase was
associated with an increase in the number of large, NS5A-con-
taining replication complexes, which were visualized in micro-
scopic images of cells supporting replication of an HCV RNA in
which enhanced yellow fluorescent protein (EYFP) was fused in
frame with NS5A (30) (Fig. S5). Thus, whereas the enhanced level
of replication in Xrn1 knockdown cells confirms that Xrn1 medi-
ates viral RNA degradation in competition with RNA synthesis,
the continued ability of miR-122 to enhance replication in Xrn1-
depleted cells suggests that miR-122 plays a role in replication
beyond its ability to protect viral RNA from Xrn1 exonuclease.
Xrn1 Knockdown Does Not Rescue Replication of a miR-122 Binding
Mutant. To further assess this Xrn1-independent function of miR-
122 in HCV RNAreplication, we asked whether Xrn1 knockdown
could rescue replication of a viral RNA containing the p6m mu-
tation in both S1 and S2 sites (Fig. 2A) that ablates miR-122
binding (HJ3-5/GLuc2A-S1-S2p6m RNA) (7). As expected (4),
this mutant RNA did not replicate when transfected into Huh-7.5
cells, generating only diminishing GLuc activity over the ensuing
72 h (Fig. 4H, Left). Also as expected, cotransfection of the
complementary miR-122p6 mutant (Fig. 2A) rescued its replica-
tion (Fig. 4H, Right), confirming that the lack of replication is due
to its inability to bind miR-122 (4). Significantly, although GLuc
expression was increased about twofold (consistent with greater
RNA stability), there was no evidence of replication of the mutant
RNA when it was transfected into Xrn1 knockdown cells in the
absence of miR-122p6m (Fig. 4 H, Left, and I). Thus, although
Xrn1 knockdown is able to fully substitute for miR-122 in stabi-
lizing the viral RNA in infected Huh-7.5 cells (Fig. 3 A and B), it is
not able to rescue replication of a miR-122 binding mutant. Con-
sistent with the absence of an effect of miR-122 on viral translation
after PSI-6130 arrest of RNA synthesis (Fig. 3C, Right), polysome
analysis demonstrated no differences in the loading of the WT and
miR-122 binding mutant RNAs on ribosomes (Fig. S6). The
absence of a difference in ribosome loading is strong evidence
that miR-122 has additional function(s) in viral genome amplifi-
cation beyond stabilization and translation of HCV RNA.
Recognition that miR-122 slows the decay of HCV RNA through
with the importance of miR-122 in the overall viral life cycle (3, 4),
led us to characterize how HCV RNA is degraded in cells. We
to HeLa S10 lysate, and viral RNA replicating within infected cells
were substantially different (Figs. 1–3). These results suggest dif-
to decay pathways, despite the fact that miR-122 stabilizes the
is subject to both Xrn1 and exosome-mediated decay, with a t1/2of
∼1.8 h, whereas RNA added to S10 lysate has a t1/2of ∼17 min and
is mainly degraded by the exosome complex and not Xrn1. It is
interesting that miR-122 protects HCV RNA from decay in HeLa
lysate, as it does this by binding near the end of the 5′UTR (Fig.
2C). One possible explanation is that HCV RNA may be degraded
by an unknown 5′ exonuclease (not Xrn1) present in S10 lysate. A
second possibility isthat miR-122, ortheAgo2proteinitrecruitsto
the 5′ UTR (10, 31), may prevent 3′ exosome degradation by
promoting circularization of HCV RNA, similar to how poly(rC)
binding protein 2 (PCBP2) promotes circularization and stabili-
zation of the poliovirus genome (32, 33). Such a hypothesis could
also explain the increased stability conferred on capped HCV
RNA in S10 lysate by miR-122 (Fig. S1C).
Replicating viral RNA appears to be subject to very different
decay pathways than transfected RNA or RNA added to HeLa
lysate. siRNA-mediated depletion of Xrn1 could have unintended
consequences on cellular mRNA abundance, thereby potentially
cells following PSI-6130 arrest of viral RNA synthesis. (A) Huh-7.5 cells were
transfected with replication-competent genotype 1a H77S.3 RNA, and then
retransfected 48 h later with siRNAs specific for Xrn1 or PM/Scl-100 or
scrambled siCtrl. After an additional 48-h incubation (time = 0), the cells
were treated with 10 μM PSI-6130, with (Right) or without (Left) simulta-
neous miR-122 supplementation. Data shown represent percent HCV RNA
remaining following addition of PSI-6130. RNA was quantified by qRT-PCR
relative to the abundance of actin mRNA. Results are the means of three
experiments ± SEM. (B) Estimated half-life (t1/2) of HCV RNA ± 95% CI under
the conditions shown in A. The data were fit to a one-phase decay model
(R2= 0.946–0.983). While the decay constant, k, differed significantly be-
tween siXrn1-, siPmScl-100–, and siCtrl-transfected cells in the absence of
miR-122 supplementation (P < 0.0001 by the extra sum-of-squares F test),
there was no significant difference in cells supplemented with miR-122 (P =
0.19). (C) GLuc expression from cells transfected with H77S.3/GLuc2A RNA,
followed by transfection of siXrn1 or siCtrl and PSI-6130 arrest of new viral
RNA synthesis as in A. Cells were supplemented with miR-122 or miR-124 at
the time of addition of PSI-6130, and media were replaced at 4-h intervals
thereafter. Data shown are the mean GLuc activity ± SD from four replicate
cultures and are representative of multiple experiments. (D) Confocal mi-
croscopy demonstrating the absence of colocalization of HCV RNA with
Xrn1. Huh-7.5 cells were transfected with H77S.3 RNA for 4 d and then
subjected to FISH for detection of HCV RNA (red). Xrn1 was visualized by
subsequent immunostaining (green) and is concentrated in P bodies (arrow
in center panel). Nuclei (N) are marked by the absence of HCV RNA and Xrn1.
Inset represents an enlarged view of a portion of the merged image. An
uninfected cell in the lower right quadrant provides an internal control
Decay of replicating HCV RNA in Xrn1- and PM/Scl-100–depleted
| www.pnas.org/cgi/doi/10.1073/pnas.1213515110 Li et al.
altering the abundance of host cell proteins. Despite this caveat,
is mediated specifically by Xrn1 with no contribution from the 3′
exosome decay pathway (Fig. 3A). The slow rate of decay under
these conditions (t1/2∼11 h) is likely to reflect limited access of
Xrn1 to HCV RNA within the membranous web (34), a mem-
brane-associated complex containing multiple viral nonstructural
proteins that directs new viral RNA synthesis. If the membranous
web represents a sanctuary within which newly synthesized viral
RNA is protected from cytosolic Xrn1, it would explain the longer
t1/2observed for replicating viral RNA than transfected, replica-
constituting the membranous web are relatively static and stable
over many hours (35), consistent with the t1/2we observed for
replicating viral RNA. Viral RNA may become subject to Xrn1
degradation during movement from the membranous web to sites
The decay rate may thus be more indicative of the turnover and
release of viral RNAfrom these complexes, rather than therate of
Xrn1-mediated 5′→3′ exoribonucleolytic digestion per se.
Alternatively, the slow rate of decay of replicating RNA may
reflect the need for removal of a 5′ terminal triphosphate from
HCV RNA, as efficient degradation of RNA by Xrn1 requires a 5′
monophosphate (37). Proteins with pyrophosphatase activity on 5′
triphosphate RNA have been reported, including RppH in
Escherichia coli andRai1p in yeast (38, 39). Whether proteinswith
similar activity are necessary for HCV RNA degradation remains
to be determined. Other viruses in the family Flaviviridae express
subgenomic RNAs (sfRNAs) that are produced by Xrn1 degra-
dation of the viral genome and are important for pathogenicity
(40, 41). Unlike HCV, however, which does not express sfRNA,
theseflavivirusgenomes possess a 5′cap,and detailsoftheirdecay
pathway are thus likely to differ from HCV.
Although we show that miR-122 protects the viral RNA from
is an additional role for miR-122 in the viral life cycle beyond
stabilization of the RNA. The critical role that it plays as an HCV
host factor is demonstrated by the lack of replication of HCV
RNA with single base substitutions in S1 and S2 that ablate miR-
122 binding (Fig. 4H). We found that the binding of miR-122 to
the5′ UTRis essential forRNAreplication even afterknockdown
of Xrn1 (Fig. 4 H and I), conditions under which miR-122 sup-
plementation has no influence on the stability of the RNA (Fig. 3
A and B). Similarly, it does not appear that the requirement for
miR-122 is related to viral protein translation or the activity of the
protein expression (4, 9), but it seems likely that this effect is due
to its ability to stabilize HCV RNA and increase its abundance
(10). Importantly, we observed no increase in viral protein ex-
pression following miR-122 supplementation in Xrn1-depleted
cells (Fig. 3C, Right), and no defects in ribosome loading by a
mutant HCV RNA defective in miR-122 binding (Fig. S6). Col-
or viral protein expression.
What other role might miR-122 play in the viral life cycle? Al-
122 is also required for amplification of HCV RNA replicons (3)
and thus has an essential role independent of viral entry or as-
sembly and release. Absent an essential effect on viral protein
siCtrl and then 24 h later retransfected with H77S.3/GLuc2A RNA. (A) Immunoblots of Xrn1, PM/Scl-100, and HCV core protein 72 h after HCV RNA trans-
fection, with β-actin as a loading control. (B) GLuc activity in supernatant fluids from Huh-7.5 cells transfected with HCV RNA and the indicated siRNAs. (C)
HCV RNA was quantified by qRT-PCR 72 h after HCV RNA transfection relative to β-actin mRNA. (D) Infectious virus titer of supernatant fluids from Huh-7.5
cells transfected with HCV RNA for 48 or 72 h, determined by a fluorescent focus formation assay. (E) GLuc assays were carried out as in B with or without
cotransfection of miR-122 (50 nM) and the indicated siRNAs. (F) Cells in E at 72 h posttransfection were harvested, and HCV RNA was quantified by qRT-PCR
relative to β-actin mRNA. (G) Graded knockdown of Xrn1 (Upper) by transfection of increasing concentrations of siXrn1 results in proportionate increases in
GLuc expression (Lower) that plateaus at the highest siXrn1 concentrations significantly below the level of GLuc expression resulting from miR-122 sup-
plementation. Cells shown as transfected with 0 nM siXrn1 received 80 nM siCtrl. (H) Huh-7.5 cells were transfected with HCV RNA containing p6m mutations
in both miR-122 binding sites (HJ3-5/GLuc2A-S1-S2p6m) with cotransfection of the indicated miRNAs. GLuc activity in supernatant fluids was analyzed at
indicated time intervals. (I) Graded knockdown of Xrn1 (see G) results in increased transient expression of GLuc from HJ3-5/GLuc2A-S1-S2p6m (due to sta-
bilization of the transfected RNA) but fails to rescue replication of the miR-122 binding mutant. Results shown in G and I represent the mean ± range from
replicate cultures, whereas all other results represent the means of three replicate experiments ± SEM.
Xrn1 knockdown enhances HCV replication in Huh-7.5 cells. Huh-7.5 cells were transfected with siRNAs specific for Xrn1 or PM/Scl-100 or scrambled
Li et al.PNAS
| January 29, 2013
| vol. 110
| no. 5
translation or RNA stability in Xrn1-depleted cells, our data point Download full-text
strongly toward a primary requirement for miR-122 in HCV RNA
synthesis. Despite this, previous efforts to demonstrate direct in-
volvement of miR-122 in viral RNA synthesis have not succeeded.
Short-term pulse-labeling experiments failed to show a diminution
in HCV RNA synthesis in vivo following transfection of an anti-
sense RNA targeting miR-122 (43), and we found that altering the
RNA by isolated membrane-bound replicase complexes (44).
However, a requirement for miR-122 in initiation of viral RNA
synthesis, or for recruitment of host replication factors to the
replication complex, might be missed in short-term pulse-labeling
studies or in ex vivo studies of isolated replicase complexes (43,
effects of miR-122 on viral RNA stability and protein expression
versus viral genome amplification that we demonstrated in the
studies described herein and suggest new directions for future
efforts to elucidate the role of miR-122 in the HCV life cycle.
Materials and Methods
Viral RNA Stability in Transfected and Infected Cells. For transfection experi-
ments, RNA was transcribed in vitro from pH77S/GLuc2A-AAG (complete geno-
type 1a HCV sequence with GLuc2A placed in-frame within the polyprotein-
into cells together with duplex miRNA. Total RNA was harvested at intervals,
and HCV RNA was measured by qRT-PCR. Stability of replicating viral RNAs
was assessed following arrest of new viral RNA synthesis with PSI-6130 as
RNA Stability in HeLa S10 Lysate. ViralRNAswereincubatedinHeLaS10lysates,
prepared as previously described except for the deletion of RNase treatment
(32, 45), and preincubated with duplex miRNA as described previously. Reac-
tions were stopped at intervals, and RNA was extracted, labeled with SYTO 62
dye, and then resolved by electrophoresis through a 1% Tris Borate EDTA-
Statistical Methods. qRT-PCR data were fit to a one-phase decay model, and
decay constants were compared using the extra sum-of-squares F test.
Additional details can be found in SI Materials and Methods.
ACKNOWLEDGMENTS. We thank Dr. William Marzluff, Angela Lam, Phil
Furman, and Charles Rice for kindly providing reagents, and the Michael
Hooker Microscopy Facility and University of North Carolina In Situ Hybridiza-
tion Core Facility for technical support. This work was supported by National
Institutes of Health Grants R01-AI095690 and P20-CA150343 and the University
Cancer Research Fund.
1. Lemon SM, Walker C, Alter MJ, Yi M (2007) Hepatitis C viruses. Fields Virology, eds
Knipe DM, et al. (Lippincott Williams & Wilkins, Philadelphia), 5th Ed, pp 1253–1304.
2. Chang J, et al. (2004) miR-122, a mammalian liver-specific microRNA, is processed
from hcr mRNA and may downregulate the high affinity cationic amino acid trans-
porter CAT-1. RNA Biol 1(2):106–113.
3. Jopling CL, Yi M, Lancaster AM, Lemon SM, Sarnow P (2005) Modulation of hepatitis C
virus RNA abundance by a liver-specific MicroRNA. Science 309(5740):1577–1581.
4. Jangra RK, Yi M, Lemon SM (2010) miR-122 regulation of hepatitis C virus translation
and infectious virus production. J Virol 84:6615–6625.
5. Jopling CL, Schütz S, Sarnow P (2008) Position-dependent function for a tandem
microRNA miR-122-binding site located in the hepatitis C virus RNA genome. Cell Host
6. Machlin ES, Sarnow P, Sagan SM (2011) Masking the 5′ terminal nucleotides of the
hepatitis C virus genome by an unconventional microRNA-target RNA complex. Proc
Natl Acad Sci USA 108(8):3193–3198.
7. Shimakami T, et al. (2012) Base pairing between hepatitis C virus RNA and microRNA
122 3′ of its seed sequence is essential for genome stabilization and production of
infectious virus. J Virol 86(13):7372–7383.
8. Fabian MR, Sonenberg N (2012) The mechanics of miRNA-mediated gene silencing: A
look under the hood of miRISC. Nat Struct Mol Biol 19(6):586–593.
9. Henke JI, et al. (2008) microRNA-122 stimulates translation of hepatitis C virus RNA.
EMBO J 27(24):3300–3310.
10. Shimakami T, et al. (2012) Stabilization of hepatitis C virus RNA by an Ago2-miR-122
complex. Proc Natl Acad Sci USA 109(3):941–946.
11. Garneau NL, Wilusz J, Wilusz CJ (2007) The highways and byways of mRNA decay. Nat
Rev Mol Cell Biol 8(2):113–126.
12. Song M-G, Li Y, Kiledjian M (2010) Multiple mRNA decapping enzymes in mammalian
cells. Mol Cell 40(3):423–432.
13. Li Y, Kiledjian M (2010) Regulation of mRNA decapping. Wiley Interdiscip Rev RNA
14. Hsu CL, Stevens A (1993) Yeast cells lacking 5′→3′ exoribonuclease 1 contain mRNA
species that are poly(A) deficient and partially lack the 5′ cap structure. Mol Cell Biol
15. Jones CI, Zabolotskaya MV, Newbury SF (2012) The 5′ → 3′ exoribonuclease XRN1/
Pacman and its functions in cellular processes and development. Wiley Interdiscip Rev
16. Liu Q, Greimann JC, Lima CD (2006) Reconstitution, activities, and structure of the
eukaryotic RNA exosome. Cell 127(6):1223–1237.
17. Liu H, Rodgers ND, Jiao X, Kiledjian M (2002) The scavenger mRNA decapping enzyme
DcpS is a member of the HIT family of pyrophosphatases. EMBO J 21(17):4699–4708.
18. Lykke-Andersen J, Shu MD, Steitz JA (2000) Human Upf proteins target an mRNA for
nonsense-mediated decay when bound downstream of a termination codon. Cell
19. Kaygun H, Marzluff WF (2005) Regulated degradation of replication-dependent
histone mRNAs requires both ATR and Upf1. Nat Struct Mol Biol 12(9):794–800.
20. Ajamian L, et al. (2008) Unexpected roles for UPF1 in HIV-1 RNA metabolism and
translation. RNA 14(5):914–927.
21. Yi M, Villanueva RA, Thomas DL, Wakita T, Lemon SM (2006) Production of infectious
genotype 1a hepatitis C virus (Hutchinson strain) in cultured human hepatoma cells.
Proc Natl Acad Sci USA 103(7):2310–2315.
22. Stuyver LJ, et al. (2006) Inhibition of hepatitis C replicon RNA synthesis by β-D-2′-
deoxy-2′-fluoro-2′-C-methylcytidine: a specific inhibitor of hepatitis C virus replica-
tion. Antivir Chem Chemother 17(2):79–87.
23. Shimakami T, et al. (2011) Protease inhibitor-resistant hepatitis C virus mutants
with reduced fitness from impaired production of infectious virus. Gastroenterology
24. Balagopal V, Parker R (2009) Polysomes, P bodies and stress granules: States and fates
of eukaryotic mRNAs. Curr Opin Cell Biol 21(3):403–408.
25. Jangra RK, Yi M, Lemon SM (2010) DDX6 (Rck/p54) is required for efficient hepatitis C
virus replication but not IRES-directed translation. J Virol 84:6810–6824.
26. Scheller N, et al. (2009) Translation and replication of hepatitis C virus genomic RNA
depends on ancient cellular proteins that control mRNA fates. Proc Natl Acad Sci USA
27. Ariumi Y, et al. (2011) Hepatitis C virus hijacks P-body and stress granule components
around lipid droplets. J Virol 85(14):6882–6892.
28. Jones DM, Domingues P, Targett-Adams P, McLauchlan J (2010) Comparison of U2OS
and Huh-7 cells for identifying host factors that affect hepatitis C virus RNA replica-
tion. J Gen Virol 91(Pt 9):2238–2248.
29. Ruggieri A, et al. (2012) Dynamic oscillation of translation and stress granule for-
mation mark the cellular response to virus infection. Cell Host Microbe 12(1):71–85.
30. Ma Y, et al. (2011) Hepatitis C virus NS2 protein serves as a scaffold for virus assembly
by interacting with both structural and nonstructural proteins. J Virol 85(1):86–97.
31. Wilson JA, Zhang C, Huys A, Richardson CD (2011) Human Ago2 is required for effi-
cient miR-122 regulation of HCV RNA accumulation and translation. J Virol 85:
32. Murray KE, Roberts AW, Barton DJ (2001) Poly(rC) binding proteins mediate polio-
virus mRNA stability. RNA 7(8):1126–1141.
33. Herold J, Andino R (2001) Poliovirus RNA replication requires genome circularization
through a protein-protein bridge. Mol Cell 7(3):581–591.
34. Gosert R, et al. (2003) Identification of the hepatitis C virus RNA replication complex
in Huh-7 cells harboring subgenomic replicons. J Virol 77(9):5487–5492.
35. Wölk B, Büchele B, Moradpour D, Rice CM (2008) A dynamic view of hepatitis C virus
replication complexes. J Virol 82(21):10519–10531.
36. Miyanari Y, et al. (2007) The lipid droplet is an important organelle for hepatitis C
virus production. Nat Cell Biol 9(9):1089–1097.
37. Stevens A (1980) Purification and characterization of a Saccharomyces cerevisiae ex-
oribonuclease which yields 5′-mononucleotides by a 5′ leads to 3′ mode of hydrolysis.
J Biol Chem 255(7):3080–3085.
38. Xiang S, et al. (2009) Structure and function of the 5′→3′ exoribonuclease Rat1 and its
activating partner Rai1. Nature 458(7239):784–788.
39. Deana A, Celesnik H, Belasco JG (2008) The bacterial enzyme RppH triggers messen-
ger RNA degradation by 5′ pyrophosphate removal. Nature 451(7176):355–358.
40. Pijlman GP, et al. (2008) A highly structured, nuclease-resistant, noncoding RNA
produced by flaviviruses is required for pathogenicity. Cell Host Microbe 4(6):
41. Silva PA, Pereira CF, Dalebout TJ, Spaan WJ, Bredenbeek PJ (2010) An RNA pseu-
doknot is required for production of yellow fever virus subgenomic RNA by the host
nuclease XRN1. J Virol 84(21):11395–11406.
42. Randall G, et al. (2007) Cellular cofactors affecting hepatitis C virus infection and
replication. Proc Natl Acad Sci USA 104(31):12884–12889.
43. Norman KL, Sarnow P (2010) Modulation of hepatitis C virus RNA abundance and the
isoprenoid biosynthesis pathway by microRNA miR-122 involves distinct mechanisms.
J Virol 84(1):666–670.
44. Villanueva RA, et al. (2010) miR-122 does not modulate the elongation phase of
hepatitis C virus RNA synthesis in isolated replicase complexes. Antiviral Res 88(1):
45. Barton DJ, Morasco BJ, Flanegan JB (1996) Assays for poliovirus polymerase, 3D(Pol),
and authentic RNA replication in HeLa S10 extracts. Methods Enzymol 275:35–57.
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