Membranous Replication Factories Induced by Plus-Strand RNA Viruses

Abstract

In this review, we summarize the current knowledge about the membranous replication factories of members of plus-strand (+) RNA viruses. We discuss primarily the architecture of these complex membrane rearrangements, because this topic emerged in the last few years as electron tomography has become more widely available. A general denominator is that two "morphotypes" of membrane alterations can be found that are exemplified by flaviviruses and hepaciviruses: membrane invaginations towards the lumen of the endoplasmatic reticulum (ER) and double membrane vesicles, representing extrusions also originating from the ER, respectively. We hypothesize that either morphotype might reflect common pathways and principles that are used by these viruses to form their membranous replication compartments.

Full text

Viruses 2014, 6, 2826-2857; doi:10.3390/v6072826
viruses
ISSN 1999-4915
www.mdpi.com/journal/viruses
Review
Membranous Replication Factories Induced by Plus-Strand
RNA Viruses
Inés Romero-Brey * and Ralf Bartenschlager *
Department of Infectious Diseases, Molecular Virology, Heidelberg University, Im Neuenheimer Feld 345,
69120 Heidelberg, Germany
* Authors to whom correspondence should be addressed;
E-Mails: ines_romero-brey@med.uni-heidelberg.de (I.R.-B.);
ralf_bartenschlager@med.uni-heidelberg.de (R.B.); Tel.: +49-0-6221-566306 (I.R.-B.);
+49-0-6221-564225 (R.B.); Fax: +49-0-6221-564570 (I.R.-B. & R.B.).
Received: 29 April 2014; in revised form: 2 June 2014 / Accepted: 24 June 2014 /
Published: 22 July 2014
Abstract: In this review, we summarize the current knowledge about the membranous
replication factories of members of plus-strand (+) RNA viruses. We discuss primarily the
architecture of these complex membrane rearrangements, because this topic emerged in the
last few years as electron tomography has become more widely available. A general
denominator is that two ―morphotypes‖ of membrane alterations can be found that are
exemplified by flaviviruses and hepaciviruses: membrane invaginations towards the lumen
of the endoplasmatic reticulum (ER) and double membrane vesicles, representing extrusions
also originating from the ER, respectively. We hypothesize that either morphotype might
reflect common pathways and principles that are used by these viruses to form their
membranous replication compartments.
Keywords: membrane rearrangements; electron microscopy; electron tomography;
ultrastructure; double-membrane vesicles; membranous replication factories; hepatitis C
virus; flaviviruses; picornaviruses; coronaviruses
OPEN ACCESS

Viruses 2014, 6 2827
1. The Family Flaviviridae
Members of the family Flaviviridae are enveloped viruses with a single stranded RNA genome of
positive polarity. This family contains four different genera: Hepacivirus (from the Greek hepar,
hepatos, ―liver‖), Flavivirus (from the Latin flavus, ―yellow‖), Pestivirus (from the Latin pestis,
―plague‖) and the recently included genus Pegivirus [1,2] (Figure 1).
Figure 1. The family Flaviviridae. Phylogenetic tree based on the analysis (neighbour
joining) of aligned conserved motifs of the RNA dependent RNA polymerase (RdRp).
Shown are selected members of the family. A distance scale corresponding to amino
acid substitutions per position is shown. (Figure adapted from [1], with permission).
Hepatitis C Virus (HCV) is the prototype species of the genus Hepacivirus. It was first discovered
by Choo et al. [3] in the serum and tissues of a chimpanzee experimentally inoculated with serum from
an individual with chronic, non-A, non-B hepatitis. This virus was associated with a mild form of
chronic hepatitis frequently observed in recipients of blood transfusions [4] and was called HCV.
A second species within the genus Hepacivirus is GBV-B which was first identified in tamarins
that developed hepatitis following inoculation with the serum from a surgeon (initials G.B.) with
acute hepatitis. Additional GB-like viruses were discovered later on and have been assigned to
the new genus Pegivirus (an acronym derived from pe, persistent; g, GB or G) within the family
Flaviviridae [1].

Viruses 2014, 6 2828
The genus Flavivirus comprises 53 species and represents the largest genus within the family
Flaviviridae. Many of these viruses are transmitted by arthropods and therefore called ARBO viruses
(arthropod-borne). Flaviviruses such as Dengue Virus (DENV), Japanese Encephalitis Virus (JEV),
West Nile Virus (WNV) or Yellow Fever Virus (YFV) cause a variety of diseases, including fever,
encephalitis and hemorrhage [5,6].
Pestiviruses are important animal pathogens causing major losses in stock farming. The genus
comprises the species Bovine Viral Diarrhea Virus 1 and 2 (BVDV-1/-2), Border Disease Virus
(BDV), Classical Swine Fever Virus (CSFV), the tentative virus species ―Giraffe‖, as well as several
unassigned viruses [7]. Within the family Flaviviridae, pestiviruses show greater similarity in genome
structure and mechanism of translation initiation to the hepaciviruses than to flaviviruses. Pestivirus
infections can be subclinical or induce a range of clinical conditions, including acute diarrhea, acute
hemorrhagic syndrome, acute fatal disease, and a wasting disease [5].
These viruses have in common a single strand RNA genome containing one long open reading
frame flanked by 5’- and 3’-untranslated regions (UTRs) [5]. Viral proteins are produced by cleavage
of a single polyprotein that has a length of about 3000 amino acid residues. The N-terminal half of
the polyprotein contains the structural proteins forming the virus particle, while the C-terminal part
contains the nonstructural proteins (NS) involved in synthesis of the viral RNA genome. The NS
proteins comprise the enzymatic components of the RNA replicase, including a RNA helicase and
a RNA-dependent RNA polymerase (RdRp) [8].
Binding and uptake of viruses belonging to the Flaviviridae family into their host cells occur by
receptor-mediated endocytosis involving interactions between cellular receptors and viral envelope
proteins. Upon uncoating, the (+) RNA genome is released into the cytoplasm, where it serves as
messenger RNA (mRNA) for translation of all viral proteins, as template for RNA replication to
produce new (−) RNA copies and as genetic material packaged into new virus particles. During
RNA replication, the (+) RNA genome is converted into a (−) RNA copy that serves as template for
production of new (+) RNA genomes. Most of these steps occur in close association with intracellular
membranes that are rearranged upon viral infection giving rise to complex membranous replication
factories. These rearrangements correspond to invaginations or protrusions of membranes that are
frequently derived from the endoplasmic reticulum (ER). In this review, we summarize our current
knowledge about the 3D architecture of these complex membrane rearrangements, focusing on
members of the family Flaviviridae that generate either morphotype of replication compartment.
2. Architecture and Properties of the Replication Factories of Members of the Flaviviridae Family
2.1. Flaviviruses
The first publications on DENV-induced membrane modifications in mammalian cells (marked
cytoplasmic vacuolization and accumulation of virions in crystalloid arrays surrounded by membranes)
date from the seventies [9,10]. Several years later, studies on DENV-infected mosquito cells were
published in which the visualization of virions was reported [11,12]. In 1997, Grief and coworkers
described DENV-induced membrane alterations of various morphologies, including convoluted
membranes (CMs) and vesicle packets (VPs) [13]. Subsequent immuno-EM studies indicated that VPs

Viruses 2014, 6 2829
and CMs may represent the site of DENV replication and RNA translation/polyprotein processing,
respectively [14]. The most complete characterization of DENV-induced intracellular membrane
rearrangements elucidated their 3D architecture as well as their spatial connection with viral assembly
sites [15]. TEM of resin-embedded infected cells revealed a complex collection of convoluted and
vesicular structures, including CMs that were usually surrounded by multiple vesicles, often appearing
as longitudinal vesicle arrays. By using electron tomography (ET), the latter were found to correspond
to ER tubules containing 80–90 nm single-membrane vesicles (Ve) that result from the invagination
of the ER membrane into the ER lumen. By conventional EM, these vesicles appeared as double
membrane vesicles, likely corresponding to the VPs described earlier [13]. Immuno-EM confirmed
that the vesicles visible in resin-embedded cells were induced by DENV infection and contained all
NS proteins. However, only NS3 was detectable within the CMs, which could be due to lower affinity
of the antibodies or poor accessibility of the other NS proteins in the CMs. Double-stranded RNA
(dsRNA) detected by immunostaining appeared as discrete electron-dense structures inside or on the
cytosolic surface of a subset of vesicles, suggesting that dsRNA might be present only in some of the
vesicles at a given time point. Furthermore, the vesicles contain rather uniform pores of ~10 nm
diameter towards the cytosol (Figure 2A). Thus, both the topology of the vesicles and the
immunolabeling results support the idea that the vesicles might be the site of RNA replication.
Moreover, these results showed that replication factories are a continuous membrane network that
provides a platform for the transport of viral proteins and genomes between sites of RNA replication,
ribosome-containing compartments (RNA translation) and virus assembly sites. In fact, virus budding
sites were found in close proximity to the pores of the replication vesicles. This topological link may
ensure efficient production and delivery of viral RNA for the assembly of infectious virus progeny.
Consistent with these findings, a very recent publication using ET showed that these virally modified
structures were also observed in DENV-infected mosquito cells, with one exception: CMs were absent
from DENV-infected C6/36 mosquito cells [16]. In addition, after multiple rounds of virus replication,
tubular structures were also observed in the vicinity of VPs. These structures might represent a
hallmark of chronically infected insect cells, since these structures are also induced by TBEV in tick
cells (see below).
The first reports on WNV-infected cells described the visualization of virions [17]. An extensive
characterization of Kunjin Virus—the Australian variant of WNV (WNVKUN)—infected cells has been
carried out more recently [18–20]. Three well-defined structures were found, corresponding to large
CMs, paracrystalline arrays (PCs) and VPs that appeared as membrane sacs containing small vesicles
(Ve) [18,21]. Based on immunolocalization studies, a distinct redistribution of the trans-Golgi network
(TGN) and colocalization of TGN markers with dsRNA has been observed, suggesting that the
replication factories of WNVKUN were derived from the TGN [22]. Three-dimensional reconstructions
of the WNVKUN replication sites revealed an intimate association of the rough ER (rER) with the
bounding membrane of the VPs [20] (Figure 2B), resembling the vesicles observed in DENV-infected
cells. These results argue for an additional role of the rER in the formation of the WNVKUN replication
factories. Similar to DENV, individual necks were observed in the vesicles as well as the majority of
the viral RNA, as detected by immunolabeling with a dsRNA-specific antibody, resided within
these vesicles [18,20,22–24]. In most cases, viral RNA spanned the breadth of the vesicles and was
juxtaposed to the necks open to the cytoplasm [20].

Viruses 2014, 6 2830
Figure 2. Representative images of membrane rearrangements induced by different
members of the family Flaviviridae. (A) Dengue Virus (DENV); (B) West Nile Virus
(WNV); (C) Tick-borne Encephalitis Virus (TBEV); (D) Hepatitis C Virus (HCV). Slices
through tomograms of infected cells (on the left) and 3D top and lateral (90° rotation)
views of the same tomograms (on the right) are depicted, showing the characteristic
virus-induced structures. The replication vesicles (Ve) of DENV, WNV and TBEV (genus
Flavivirus) correspond to invaginations of ER membranes that remain connected to the
cytosol via 10 nm-pores (highlighted with white arrows in the 3D lateral views), forming
vesicle packets (VPs). The replication factory of HCV (genus Hepacivirus) is primarily
composed of double membrane vesicles (DMVs) that seem to be formed asER protusions
connected to ER membranes via neck-like structures (highlighted with white arrows in the
3D lateral view). The ER is shown in yellow (DENV, TBEV and HCV) or in red (WNV)
and the replication organelles in brown (DENV, TBEV and HCV) or in white (WNV). The
outer and inner membranes of DMVs are depicted in different shades of brown (outer
membrane in dark brown and inner membrane in light brown). Figure 2B is reproduced
with permission from [20].
In cells infected with TBEV, one of the most important tick-transmitted viruses in Europe and Asia,
virus particles and membrane-connected vesicles were also observed inside the ER [25], similar to
what was described for DENV and WNVKUN. The viral dsRNA was only detected inside the vesicular
structures within rER, suggesting that TBEV rearranges internal cell membranes to generate

Viruses 2014, 6 2831
a compartment that protects viral RNA from detection by cytoplasmic pathogen recognition
receptors (PRRs) [26–28]. This localization of dsRNA might suffice to delay the onset of the IFN
response [25]. For TBEV [25] and WNVKUN [19] it was shown that treatment with brefeldin A (BFA),
a drug which disrupts the Golgi apparatus, did not interfere with viral replication. However, this
treatment rendered WNVKUN sensitive to the antiviral action of the IFN-induced protein MxA,
indicating that BFA might have disrupted the membranous WNVKUN replication compartments, thus
leading to exposure of dsRNA and its detection by PRRs. In contrast, treatment of TBEV-infected cells
with BFA neither affected viral replication, nor the level of IFN production. These findings indicate
that TBEV dsRNA might be stored inside BFA-resistant membrane vesicles that robustly protect the
viral RNA from recognition by cellular sensors.
Vector-borne flaviviruses like DENV and TBEV must replicate in both mammalian and arthropod
cells. A few comparative studies have been published describing virus-induced structures such as
cytoplasmic membrane proliferations and vesicle formation, also in insect cells [11,16,21,29,30,].
A detailed comparative ultrastructural analysis of TBEV-induced modifications revealed that the
extent of membrane expansion and the abundance of vesicles were lower in insect cells [31].
Single-membrane vesicles, ranging in diameter from 60–100 nm were frequently found within
proliferated ER areas, often occurring in large groups contained within ER cisternae. Pore-like
openings connected these vesicles to the cytoplasm and to other vesicles. Apart from these vesicles,
in tick-infected cells elongated vesicles or tubules were found that were much more prevalent in
persistently than in acutely infected cells. These tubules were only occasionally noted in infected
mammalian cells, similar to what was found with DENV-infected cells [15]. The tubular structures had
a cross-sectional diameter of 60–100 nm, similar to the one of vesicles, reached up to 800 nm in
length, were closed at the ends and often arranged in fascicle-like bundles, shrouded with the ER
membrane. However, no pores between the tubules or towards the cytoplasm were observed [31]. The
function of these tubules is unclear and it is not known whether they represent bona fide features of
replication factories, aberrant structures as a result of incorrect membrane remodeling, or the result of
a cellular process to restrict infection [31]. In any case, the tubules might be a feature of persistent
infection, eventually linked to the high number of defective virus particles, because the lack of
pores could prevent proper replication or packaging of the viral genome [32–34]. Further studies
are required to shed light on the biogenesis and biological significance of these membranous
tubular structures.
A recent study identified 80 nm-diameter vesicles within the ER lumen of TBEV-infected BHK-21
cells and in cells transfected with a TBEV replicon [35]. ET revealed that these vesicles are invaginations
of the ER within a highly organized network of interconnected membranes with half of vesicles
containing pore-like connections to the cytoplasm (Figure 2C). However, no pore-like openings
were observed between adjacent/neighboring vesicles, in contrast to what has been described for
cells infected with Langat Virus (LGTV), a naturally attenuated tick-borne flavivirus [31] or in
WNVKUN-infected cells [20]. Interestingly, in TBEV replicon cells, the number of pore-containing
vesicles was slightly larger (~75%) and they were found in much more fragmented ER tubules as
compared to TBEV-infected cells. However, despite more extensive ER rearrangements in replicon
cells, they contained fewer vesicles, consistent with the lower level of viral replication [36–38].

Viruses 2014, 6 2832
Conventional EM analysis of neurons infected with Murray Valley Encephalitis Virus (MVEV)
revealed several ultrastructural features, including proliferation of ER and Golgi complex membranes
as well as the appearance of membrane-bound spherical vesicles (75–145 nm diameter) [39], similar to
those observed for the related flaviviruses Japanese Encephalitis Virus (JEV) [40,41] and St. Louis
Encephalitis Virus (SLEV) [42]. In the latter case, cylindrical membranous structures (or tubules) were
also observed [43]. The presence of vesicles was also detected in monkey liver cells infected with
Yellow Fever Virus (YFV) [44]. These findings indicate that all members of the genus Flavivirus
utilize the ER as a source of membranes for the formation of their replication factories, whereas
assembly of new virions seems to occur at ER sacs in close proximity to the replication sites [15,35],
thus creating an optimized membranous environment to support efficient viral replication and
assembly. Maturation of the newly synthesized virions takes place in the Golgi apparatus, where
flavirirus virions are often observed [15].
2.2. Hepaciviruses
In stark contrast to flaviviruses, HCV, the prototype of the genus Hepacivirus, provokes an
alternative rearrangement of intracellular membranes, originally designated ―membranous web‖
(MW). This term referred to compact vesicle accumulations embedded into a membranous matrix [45]
as detected in cells inducibly expressing the HCV polyprotein. By using different EM methods, we
and others have recently found that the MW is primarily composed of double membrane vesicles
(DMVs) [46–48]. The fact that the kinetics of their appearance correlates with HCV replication
suggests that these structures play an important role for viral RNA amplification [48]. Indeed,
immunolabeling of purified DMVs revealed an enrichment for viral proteins as well as dsRNA [46,49].
Importantly, DMVs contain enzymatically active viral replicase [49] and they originate from ER
membranes, similar to what has been found for other members of the family Flaviviridae. ET analysis
showed that most of the DMVs remain connected to the ER via their outer membrane [48]
(Figure 2D). Although DMVs are primarily closed structures, ~10% of them have an opening
towards the cytosol. Late in infection, multi-membrane vesicles (MMVs) with an average diameter
of 390 nm are generated, likely originating from DMVs by secondary enwrapping events [48].
By using Huh7.5 cells infected with the highly replicative HCV strain JFH-1, Ferraris and
coworkers observed three different types of membrane alterations: vesicles in clusters (ViCs),
contiguous vesicles (CVs) and DMVs [47]. The ViCs were small single-membrane vesicles of
variable size (100–200 nm), grouped together in well-delimited areas. Most of them had an internal
invagination. The CVs were small single-membrane vesicles, present in large numbers and widely
distributed throughout the cytoplasm, with a more homogeneous size (around 100 nm). They were
tightly associated to each other and tended to form a collar around lipid droplets (LDs). DMVs were
heterogeneous in size (150–1000 nm) and had a thick, electron-dense membrane consisting of
two closely apposed membranes. The increase of CVs’ number correlated with an increase of intracellular
HCV RNA levels, arguing for a possible role of CVs in the early stages of viral replication. The
presence of NS5A in CVs, as demonstrated by immunogold staining, is consistent with this hypothesis.
Alternatively, CVs might constitute the membranous platform for viral assembly. In fact, the core
protein is present in these structures (16%) as well as on the LD surface (81%). However, so far

Viruses 2014, 6 2833
visualization of virus particles in infected cells has not been possible, making this hypothesis difficult
to prove. While most of the dsRNA signal was located within DMVs or at DMV membranes, ViCs
were free of viral components and RNA and these structures as well as CVs were very rarely observed
in cells with a subgenomic JFH-1 replicon [46] or absent in cells infected with a JFH-1 variant
designated Jc1 [48]. The first 3D reconstruction of a complete HCV-infected cell revealed that all
these three membrane structures were tightly connected and closely associated with LD clusters [47].
Taken together, these findings indicate a fundamental role of DMVs in HCV replication.
An in-depth comparison of the study by Ferraris and coworkers [47] and our publication [48] suggests
that CVs might be also DMVs for several reasons: first, CVs have electron dense tightly apposed
membranes; second, by using correlative light and electron microscopy, we also detected DMV
accumulations around LDs, reminiscent of the CVs described by Ferraris and coworkers [47]; third,
taking into consideration the density of content and morphology, some of the structures described as
DMVs by Ferraris and colleagues might correspond to MMVs according to our nomenclature. This
might account for the differences in size between the DMVs reported in both studies (up to 1000 nm
versus 150 nm, respectively). Alternatively, the difference might be due to the use of distinct virus
strains (JFH-1 and Jc1) that differ in their capacity to produce infectious virus particles by ~3 orders of
magnitude [50], which might also explain the presence of ViCs only in JFH-1 infected cells.
2.3. Pestiviruses
Much less about membranous replication factories is known for pestiviruses. TEM-based studies
from the times in which the genus Pestivirus was still belonging to the family Togaviridae reported
that pestivirus-infected cells exhibited ultrastructural modifications of rER and contained small
numbers of virus-like particles (VLPs) [51,52]. Gray and Nettleton (1987) reported that Border
Disease Virus (BDV)-infected cells contained several profiles of ER and many dense lamellar bodies,
which when transversely sectioned appeared as multiple rows of tubules, 33 nm in diameter [52].
These lamellae were often found in association with rER and in one occasion VLPs appeared to be
budding within them. Bovine Viral Diarrhea Virus (BVDV)-infected cells contained rER modified into
tubules, in which electron-dense VLPs were present. More recent studies on BVDV-infected cells
revealed cytoplasmic vacuolization and VLPs in dilated ER cisternae [53,54]. In addition, membrane
structures consisting of vesicles of various sizes enclosed in much larger vesicles have been reported
[55]. These structures that morphologically resemble multivesicular bodies (MVBs) are distinct from
the HCV-induced membranous web and more reminiscent of the flavivirus-induced VPs.
Studies on the morphogenesis of pestiviral particles were hampered by a low rate of virion
production. In a recent study, Schmeiser and colleagues have overcome this problem by using high
multiplicity of infection in MDBK cells with a distinct virus strain, the Giraffe-1 strain [56]. Obtained
results define the ER as the site of pestivirus particle assembly, where budding of virions was
observed. Virus particles were also found inside the lumen of the Golgi and in vesicles associated with
the Golgi compartment, suggesting that virus egress occurs via the conventional secretory pathway.
Interestingly, replication kinetics of pestiviral RNA did not correlate with distinct membrane
rearrangements and only slight dilatation of the ER lumen was noticed. The absence of significant
membrane rearrangements argues for a major difference between pestiviruses and other members of

Viruses 2014, 6 2834
the Flaviviridae family. Interestingly, the authors detected the capsid protein and dsRNA, the marker
for viral replication intermediates, mainly in MVBs, indicating that pestiviruses are either using this
compartment for replication or that viral RNA and proteins are transferred to this compartment for
degradation. Similar assumptions have been made for HIV [57] and Marburg virus, a member of the
Filoviridae family [58,59]. Alternatively, pestiviral RNA and protein in MVBs might be intermediates
of the entry process, prior to fusion of the envelope with the endosomal membrane. Indeed, particles
inside MVBs matching the morphological criteria of pestivirus virions were detected [56]. However,
MVBs of non-infected cells also contain vesicles for lysosomal degradation termed intraluminal
vesicles (ILVs) that display a very similar morphology to pestiviral virions. Thus, unambiguous
discrimination between ILVs and pestivirus particles will require detailed immunolabeling approaches.
3. Architecture and Properties of the Replication Factories of Other (+) Strand RNA Viruses
3.1. Nodaviruses
The first visualization of the 3D architecture of a (+) strand RNA virus replication factory was
reported for Flock House Virus (FHV), a member of the family Nodaviridae [60]. This insect nodavirus
induces the formation of invaginations at the outer mitochondrial membrane (OMM) with an average
diameter of ~50 nm [60,61] (Figure 3A). The interior of these vesicles (called spherules) is connected
to the cytoplasm by a necked channel of ~10 nm diameter, which is wide enough to allow import of
ribonucleotides and export of synthesized RNA (diameter < 2 nm) [60]. Furthermore, metabolically
labeled FHV RNA localized between inner and outer mitochondrial membranes inside these spherules,
thus validating the spherules as bona fide FHV-induced compartments for viral RNA synthesis [60].
Figure 3. Positive strand RNA viruses usurp and modify cell membranes of different
origins to replicate their genomes. (A) Mitochondrial membranes are targeted by Flock
House Virus (FHV), which induces the formation of vesicles (white) at the outer
mitochondrial membrane (blue); (B) 3D surface-rendered model of SARS-CoV–infected
Vero E6 cells containing large double membrane vesicles (DMVs) (outer membrane,
gold; inner membrane, silver) that remain connected to their source, the ER (in bronze);
(C) Zippered ER, found in cells infected with the gammacoronavirus Infectious Bronchitis
Virus (IBV), is connected to spherules (red arrows). DMVs (red arrowheads) are also
found, but to a lesser extent; (D) Equine Arteriviruses (EAV) infection of HeLa cells
results in the formation of DMVs (brown), depicting a core (blue) that is associated with
the ER (beige) and close to ER tubules (green); (E) Closterovirus rearranges ER and
mitochondrial membranes to form DMVs (red arrowheads) and vesicles packets (VPs) (red
arrows); (F) Coxsackie B3 Virus (CVB3) usurps donor membranes most likely derived
from the Golgi, appearing early in infection as single-membrane tubules (green), open
(orange) and closed (yellow) DMVs and ER (blue); (G) 3D architecture of poliovirus
membranous replication factories at intermediate stages of development, originating from
cis-Golgi membranes. Single-membrane structures are depicted in different shades of blue.
Note that these single membrane vesicles undergo secondary invaginations giving rise to
DMVs at the late stages of infection; (H) Murine Norovirus 1 (MNV-1) infection results in

Viruses 2014, 6 2835
the formation of vesiculated areas (VAs, red arrowheads) within aggregates of MNV-1
particles (red arrows). These VAs seem to originate from ER, trans-Golgi and endosomes;
(I) 3D model of a cytopathic vacuole (CPV) found in Rubella Virus-infected cells. This
CPV (yellow) is surrounded by the rER (light green) and contains a number of vacuoles,
vesicles (white) and a rigid straight sheet (brown) that is connected with the periphery of
the CPV. Mitochondria are depicted in red; (J) Typical cytoplasmic vacuoles induced by
Semliki Forest Virus (SFV) in baby hamster kidney cells. Color code is as follows:
blue-framed images depict single-membrane vesicle inducers; green-framed images
correspond to double-membrane vesicle inducers. (The different parts were reproduced
with permission: see acknowledgement section)

Viruses 2014, 6 2836
3.2. Nidovirales: Corona- and Arteriviruses
Conventional TEM analyses of coronavirus-infected cells identified large numbers of isolated
DMVs [62]. At least in case of the Severe Acute Respiratory Syndrome (SARS)-Coronavirus, these
DMVs are part of an elaborate reticulovesicular network (RVN) of modified ER that consists of
convoluted membranes, numerous DMVs (diameter 200–300 nm) (Figure 3B) [63], and ―vesicle
packets‖ apparently arising from merging of DMVs. The CMs were most intensively immunolabeled
for viral replicase subunits whereas DMVs labeled abundantly for dsRNA. While this result argues
that DMVs might be the site of viral RNA synthesis, ET analyses failed to detect DMV connections to
the cytoplasm to allow transport of nascent RNA. Instead, DMVs are connected to each other, to
CMs and to the ER via their outer membranes.
Also, in case of another coronavirus, the Mouse Hepatitis Virus (MHV), ER membranes are thought
to be the lipid donor of the membranous replication compartment [64,65]. Qualitative and quantitative
analyses by (immuno)-electron microscopy of MHV-induced membrane rearrangements revealed the
appearance, in strict order, of DMVs (diameter 200–350 nm), CMs, large virion-containing vacuoles,
tubular bodies and cubic membrane structures [65].
The recently identified coronavirus Middle East Respiratory Syndrome Coronavirus (MERS-CoV)
induces extensive membrane rearrangements in the perinuclear region, including the formation of
DMVs and CMs [66]. The diameter of MERS-CoV induced DMVs ranged from 150–320 nm,
comparable to the SARS-CoV induced DMVs. In addition, CMs were always surrounded by DMV
clusters and were only observed in cells that appeared to be more advanced in infection. This
observation strengthens the notion that DMV formation precedes the development of CMs, as
postulated previously for SARS-CoV [63].
In addition to the betacoronaviruses (SARS-CoV, MHV and MERS-CoV), ER-derived DMVs
with a diameter of ~200 nm have been observed in primary avian and mammalian cells infected
with Infectious Bronchitis Virus (IBV), an important poultry pathogen belonging to the genus
Gammacoronavirus [67]. However, the most striking structures induced by IBV are zippered ER
membranes (Figure 3C). The zippered ER was associated to 60–80 nm diameter spherules, structures
that are not present in cells infected with betacoronaviruses. ET showed that these IBV-induced
spherules are tethered to the zippered ER and contain a 4.4 nm long channel connecting their interior
to the cytoplasm of the cell, making them the ideal candidates for the site of IBV RNA synthesis.
ER membranes are also targeted by nidovirales that belong to the family Arteriviridae. Cells
infected with the prototypic arterivirus, Equine Arterivirus (EAV), also contain DMVs associated to
ER tubules. These DMVs are ~2–3 times smaller (~95 nm) as compared to coronaviruses [68].
A recent in-depth ultrastructural analysis revealed that the outer membranes of EAV-induced DMVs
are interconnected with each other and with the ER (Figure 3D), thus forming a reticulovesicular
network (RVN) resembling the one previously described for the distantly related SARS-CoV [69].
Despite significant morphological differences, a striking parallel between the two virus groups,
and possibly all members of the order Nidovirales, is the accumulation of dsRNA, the presumed
intermediate of viral RNA synthesis, in the DMV interior. Along these lines, DMVs visualized by
means of electron spectroscopy imaging contained phosphorus in amounts corresponding on average
to a few dozen copies of the EAV RNA genome. Like in SARS-coronavirus infected cells, connections

Viruses 2014, 6 2837
between DMV interior and cytosol could not be unambiguously identified, suggesting that dsRNA is
compartmentalized by the DMV membranes. In addition, ET revealed a network of nucleocapsid
protein-containing protein tubules, intertwined with the RVN. This potential intermediate in
nucleocapsid formation, which was not observed in coronavirus-infected cells, suggests that arterivirus
RNA synthesis and assembly are spatially coordinated.
3.3. Picornaviruses
Membrane remodeling in picornavirus-infected cells has been studied for more than 50 years.
Massive virus-induced membrane modifications have been reported already in 1958 [70], but the
origin of these membranes is still a matter of controversy. Several lines of evidence, including
biochemical and structural data, suggest that the ER must play a major role in the formation of
those structures [71–73]. Early in infection membranous replication factories contain markers of
the Golgi [74,75], whereas markers of the ER, Golgi and lysosomes were all found to be associated
with poliovirus replication sites late in infection [76]. Initial reports identified membrane
rearrangements as U-bodies because of their horseshoe-like shape [70]. Later, Bienz et al. described
rearranged membranes as clusters of single-membrane vesicles [71,77], while other reports [76,78]
noticed the double-membrane morphology of picornavirus-induced vesicles. The single- versus
double-membrane morphology of the vesicles was first interpreted as two different models of their
formation. However, several recent publications suggest that picornavirus-induced membrane
rearrangements might occur in a consecutive manner. Thus, early in poliovirus infection small clusters
of single-membrane vesicles predominate that are transformed into bigger irregularly shaped
single-membrane structures (Figure 3G) and, late in infection, replaced by either round or irregularly
shaped DMVs [79]. Interestingly, the small clusters of single-membrane vesicles of ~100–200 nm
diameter contain GM130, a cis-Golgi marker, but did not stain positive for calnexin, an ER marker.
However, this does not exclude a role of the ER for biogenesis of these vesicles, because ER-resident
proteins might be sorted out as these membranes are transformed. Although dsRNA and metabolically
labeled viral RNA were detected in single-membrane vesicles and DMVs, the exponential phase
of viral RNA synthesis correlates with the appearance of single-membrane and intermediate
structures [79] arguing that these structures are most relevant for high level poliovirus RNA synthesis.
Similar results have been obtained with another member of the family Picornaviridae, coxsackie
B3 virus (CVB3) that also induces the formation of single- and double-membrane compartments,
whose relative abundance correlates with the stage of the replication cycle [80] (Figure 3F). Based on
the observation that the Golgi apparatus disappears in CVB3-infected cells, the membrane
rearrangements might originate from this organelle (Montserrat Bárcena, personal communication).
Similar to poliovirus, single-membrane tubular clusters occur predominantly early in infection,
whereas the number of DMVs increases as infection progresses. A budding event could account for
the formation of the tubules, depicting an average length of 654 ± 291 nm and an average diameter of
81 ± 7 nm. A subsequent enwrapping of these single-membrane tubules via an ―autophagy-like‖
mechanism could then lead to the formation of DMVs that have an average diameter of 159 nm ± 47 nm.
This transformation may require several steps: (i) membrane pairing; (ii) induction of curvature;
and (iii) membrane fusion [80]. This scenario would be consistent with the membrane surface of

Viruses 2014, 6 2838
DMVs as an average-sized DMV with a diameter of 160 nm would be equivalent to a tubule with
a length of 632 nm and a diameter of 81 nm. ER membranes were found near DMV clusters. However,
in contrast to previous observations in nidovirus-infected cells, these DMVs were not connected to
neighboring structures. In addition to these compartments, a third type of modification was detected in
CVB3-infected cells: multilamellar structures, which are typical for the late phase of infection and
that correspond to enwrapped DMVs: despite their various shapes and degrees of complexity, in all
instances they contained one DMV, surrounded by one or several layers of curved cisternae.
In conclusion, these results, and similar observations made for Foot and Mouth Disease Virus
(FMDV) (genus Aphthovirus) [81] suggest that members of the Picornaviridae family induce single-
and double-membrane vesicles. They appear in a time-dependent manner and seem to evolve from
each other, possibly in coordination with the progression of the viral replication cycle [80]. These
membrane rearrangements occur independently from the used virus strain and cell line.
3.4. Togaviruses
Rubella Virus (RUBV) is an important human teratogenic virus and the only member of the genus
Rubivirus (family Togaviridae) [82]. RUBV anchors its RNA synthesis machinery to membranes of
a cell organelle known as ―cytopathic vacuoles‖ (CPVs) that is derived from modified endosomes or
lysosomes and has an average diameter of 600–2000 nm [83–85]. Freeze-fracture and ET analysis of
RUBV-infected cells revealed a high complexity of CPVs that are composed of stacked membranes,
rigid sheets, small vesicles and large vacuoles (Figure 3I) [86]. The CPVs are interconnected and
linked to the endocytic pathway, as deduced from labeling experiments with endocytosed BSA-gold.
Furthermore, rER cisternae, mitochondria and Golgi stacks are recruited around CPVs to build up
RUBV factories. CPVs have several contacts with cellular organelles: they are coupled to the rER
through protein bridges of ~10–15 nm and closely apposed membranes and they are attached to Golgi
vesicles, whereas contacts with mitochondria were not detected [86]. It has been proposed that
RNA synthesis occurs on vesicular membranes within the CPVs, which are linked to the cytosol and
that the viral replicase molecules are associated with vesicles that transform with time into large
vacuoles and straight elements [86]. This is supported by immunogold labeling revealing replicase
components and dsRNA within the CPVs [83,84,87].
The modification of late endosomes and lysosomes is a feature that RUBV shares with
alphaviruses, the other genus of the family Togaviridae [88,89]. Cells infected with alphaviruses like
Semliki Forest Virus (SFV), Sindbis Virus and Western Equine Encephalitis Virus (WEEV) contain
large CPVs with a diameter ranging between 600 and 2000 nm. The inner surface of these CPVs is
covered with small invaginations or spherules that originate at the plasma membrane [90–92]
(Figure 3J). These spherules are comprised of a single membrane forming a vesicle with a diameter of
~50 nm. In addition, the spherules were shown to be the site of viral RNA synthesis as deduced from
metabolic labeling and detection by EM [89,91,92]. Importantly, the inside of the spherule is
connected to the cytoplasm by a pore with a diameter of 5–10 nm. The spherules are formed at
the plasma membrane by the concerted action of the viral nonstructural proteins (nsP1–nsP4) and
genomic viral RNA [89]. Furthermore, Froshauer et al. [88] showed that the CPVs that contain the
spherules possess endosomal and lysosomal markers.

Viruses 2014, 6 2839
Time course studies revealed that the spherules of SFV undergo an unprecedented large-scale
movement between cellular compartments [93]. The spherules first form as blebs (exvaginations) at
the plasma membrane. Then, they are internalized by an endocytic process requiring a functional
actin-myosin network. The spherules therefore represent an unusual type of endocytic cargo. After
endocytosis, spherule-containing vesicles, namely CPVs-I fuse with acidic endosomes and move along
microtubules. This leads to the formation of a very stable compartment, where the spherules
accumulate as invaginations on the outer surface of unusually large, acidic vacuoles localized in the
pericentriolar region [93].
3.5. Caliciviruses
Members of the genus Norovirus (NoVs, family Caliciviridae) are major agents of acute
gastroenteritis [94]. Ultrastructural examination of Murine Norovirus 1 (MNV-1)-infected cells
revealed a striking change in their overall morphology and intracellular organization [95]. Structures
resembling virus particles were observed within or next to single- or double-membrane vesicles in the
cytoplasm. The vesiculated areas increase in size with time and by 24 hpi, large numbers of these
vesicles and viral particles occupy most of the cytoplasm and displace the nucleus (Figure 3H).
In addition, a complete rearrangement of the ER and loss of intact Golgi apparatus was observed. Both
dsRNA and MNV-1 nonstructural protein 7, the RNA dependent RNA polymerase, localize to the
limiting membrane of individual vesicle clusters by immuno-EM [96]. Immunofluorescence-based
double-labeling showed that MNV-1 appears to recruit membranes derived from multiple cellular
organelles and/or compartments: the ER, trans-Golgi apparatus and endosomes. However, despite
extensive efforts, human norovirus cannot be grown in cultured cells [97]. Thus, detailed studies have
not been possible, but it is assumed that replication structures are similar to those of MNV-1.
Feline Calicivirus (FCV), a member of the genus Vesivirus within this family, is a major agent
of respiratory disease in cats, which replication originates also membranous rearrangements and
vesicles [98].
3.6. Plant Viruses
Brome Mosaic Virus (BMV, family Bromoviridae) generates its replication factory by hijacking ER
membranes, similar to what has been described for other plant viruses like Tobacco Mosaic Virus
(TMV, family Virgaviridae) [99], Tobacco Echt Virus (TEV, family Potyviridae) [100] and Red
Clover Necrosis Mosaic Virus (family Tombusviridae) [101]. However, other plant viruses such as
Alfalfa Mosaic Virus (AMV) [102] and Cucumber Mosaic Virus (CMV), both belonging to the family
Bromoviridae, and Turnip Yellow Mosaic Virus (TYMV, family Tymoviridae) [103] anchor their
replication sites on chloroplasts. Cucumber Necrosis Virus (CNV), family Tombusviridae, utilizes
peroxisomal membranes as replication platforms [104], while other plant viruses replicate on the
surface of mitochondria [105].
Although the 3D architecture of these membranous replication sites remains largely unknown, their
characteristics are strikingly similar to those for FHV (family Nodaviridae). Best studied is BMV that
induces spherules, of similar size as the insect nodavirus FHV, in the ER close to the nucleus, where
viral RNA synthesis and viral replication proteins are localized [106–108]. In the case of Beet Yellows

Viruses 2014, 6 2840
Closterovirus (BYV, family Closteroviridae), TEM of infected plant cells revealed the formation of
~100 nm-diameter DMVs and multivesicular complexes (single-membrane vesicles surrounded by
a common membrane) (Figure 3E) [109]. These multivesicular complexes often reside next to stacks
of aligned filamentous BYV particles [110,111] and resemble the DMVs and VPs produced by
nidoviruses and flaviviruses. Several BYV replication-associated proteins (L-PCP, MTR and HEL)
colocalize with DMV and VP membranes, supporting the role of these structures as replications
platforms [112,113]. The membranes in closterovirus DMVs and VPs are likely to be derived from ER
for members of the genus Crinivirus [114] or mitochondria in case of Ampelovirus [115,116]. Whether
these structures are ―closed‖ or ―necked‖ remain unknown.
4. Similarities and Differences between the Replication Factories of Flaviviruses and other (+)
Strand RNA Viruses
Based on amino acid sequence homologies of their RNA-dependent RNA polymerases, (+) RNA
viruses have been classified into three large supergroups [113,117,118]: supergroup I (Picornavirata
or picorna-like group), including picorna-, corona-, arteri- and nodaviruses; supergroup II (Flavivirata
or the flavi-like group), including tombus-, diantho-, pesti-, hepaci- and flaviviruses as well as
single-strand RNA bacteriophages; supergroup III (Rubivirata or the alpha-like group), including
tobamo-, hordei-, alpha- and rubiviruses as well as hepatitis E virus (HEV). These higher-order
taxonomic units encompass diverse viruses infecting different hosts from almost all kingdoms of life.
As discussed earlier [119], amongst these viruses two main architectures of remodeled membranes
(morphotypes) can be found that may reflect two alternative strategies to induce the membranous
microenvironments required to allow virus replication (summarized in Table 1).
The first morphotype involves the formation of negatively curved membranes, initiated by
invaginations of the pre-existing membrane bilayer and giving rise to spherules, vesicles or vacuoles
towards the lumen of the targeted cell organelle. These structures have been identified in a broad range
of mammalian, plant and insect cells infected with viruses belonging to supergroups II and III.
The second strategy involves the formation of membranes with positive curvature, giving rise to
double-membrane structures that are the predominant characteristic of the replication factories of the
picorna-like virus supergroup. The conservation of these two sorts of morphotypes in distantly related
viruses supports the assumption of an evolutionary conserved mechanism. A striking finding in this
regard was the observation that HCV, despite belonging to the flavi-like supergroup, induces DMVs
whereas flaviviruses induce the formation of negatively curved membranes. To our current knowledge,
HCV is the only member of the family Flaviviridae inducing the formation of membrane structures
with positive curvature, suggesting that HCV might share common host cell pathways to induce
membranous replication compartments with distantly related viruses such as corona-, arteri-, picorna-,
calivi- or closteroviruses (Table 1). However, it still remains to be elucidated whether other members
of the family Flaviviridae, belonging to the genera Pestivirus and Pegivirus, also utilize a picorna-like
membrane remodeling strategy.

Viruses 2014, 6 2841
Table 1. Main structures induced by different (+) strand RNA viruses.
Morphotype
Virus Group and
Representative Member
Name of
Membrane
Alteration
Size/Diameter
Membrane
Source
Single-membrane
vesicles
Nodaviruses
FHV
Spherules
~50 nm
Mitochondria
Bromoviruses
BMV
~60 nm
ER (adjacent to
the nucleus)
Togaviruses
RUBV
Cytoplasmic
vacuoles (CPVs)
600–2000 nm
Lysosomes
SFV
CPVs with
spherules
CPVs:
600–2000 nm
Spherules:
50 nm
Flaviviruses
DENV
Invaginated vesicles
80–120 nm
ER
WNVKUN
50–100 nm
TBEV
60–100 nm
MVEV, SLEV
and JEV
75–145 nm
YFV
unknown
Double-
membrane
vesicles
Hepaciviruses
HCV
Double membrane
vesicles (DMVs)
~150 nm
ER
Coronaviruses
SARS-CoV
200–300 nm
MHV
200–350 nm
MERS-CoV
150–320 nm
IBV
~200 nm
Arteriviruses
EAV
~95 nm
Picornaviruses
Poliovirus
100–300 nm
Golgi
CV3B
~160 nm
FMDV
~100 nm
unknown
(probably ER)
Calicivirus
MNV-1
~200 nm
ER/Golgi
Closterovirus
BYV
~100 nm
ER/Mitochondria
Abbreviations: FHV, Flock House Virus; BMV, Brome Mosaic Virus; RUBV, Rubella Virus; SFV, Semliki
Forest Virus; DENV, Dengue Virus; WNVKUN, West Nile Virus (Kunjin strain); TBEV, Tick Borne
Encephalitis Virus; MVEV, Murray Valley Encephalitis Virus; SLEV, St. Louis Encephalitis Virus; JEV,
Japanese Encephalitis Virus; YFV, Yellow Fever Virus; HCV, Hepatitis C Virus; SARS-CoV, Severe Acute
Respiratory Virus-Coronavirus; MHV, Mouse Hepatitis Virus; MERS-CoV (Middle East Respiratory Syndrome-
Coronavirus); IBV, Infectious Bronchitis Virus; EAV, Equine Arterivirus; CVB3, Coxsackie B3 Virus;
FMDV, Foot and Mouth Disease Virus; MNV-1, Murine Norovirus-1; BYV, Beet Yellows Virus.
A common feature associated with the spherule/vesicle/vacuole/-type of rearranged membranes is
their size (50–150 nm diameter) and the presence of a pore connecting the interior of the vesicle
with the cytoplasm [15,20, 60, 88]. Since RNA replication occurs in the vesicle interior, the pore
allows exchange of nucleotides and RNA products with the cytoplasm. The size of the pore is
variable, ranging from 4.4 nm in case of IBV-induced spherules to ~10 nm in case of membrane
invaginations induced by flaviviruses. In contrast, in the majority of DMVs no such channel or pore

Viruses 2014, 6 2842
has been detected. Nevertheless, as exemplified with nidoviruses, the inner compartments enclosed
by interconnected DMVs contain the bulk of dsRNA [63], and in some cases they depict
an electron-dense core assumed to correspond to viral RNA (IBV and EAV) [67,69]. Although
this represents a functional enigma in terms of RNA synthesis and transport, the presence of dsRNA in
the DMV interior does not necessarily indicate active RNA replication. Assuming a temporal
regulation, it is possible that DMVs might be sites of RNA synthesis as long as they are linked to the
cytoplasm, but replication would stop upon closure of the vesicles. Yet, another strategy appears to be
used by enteroviruses, where active RNA replication has been detected on the cytosolic side of the
membranous structures [71,120], consistent with the membrane topology of the nonstructural proteins
catalyzing RNA replication [121–123]. As described above, studies conducted with picornaviruses
revealed that the exponential phase of viral RNA synthesis coincides with the accumulation of
single-membrane tubules [79,80]. Importantly, pulse-radiolabeling experiments localized sites of
active RNA replication to the outer surface of single-membrane tubules [71] and isolation of the
membranous replication factories and their subsequent visualization by EM revealed that they form
rosette-like structures composed of virus-induced cytoplasmic vesicles [124]. RNA replication is
thought to occur at sites where the vesicles cluster, whereas RNA translation probably takes place on
the exposed periphery of the vesicles. This raises the question, what the role of DMVs in the
replication cycle of picornaviruses might be. It is possible that DMVs either support RNA synthesis or
serve as RNA storage sites (especially in case of closed DMVs). In this manner, DMVs might be
involved in regulating viral RNA replication: by complete sealing of the viral replicase inside the
vesicle, it would be inactive, thus regulating overall RNA copy number in the infected cell. Alternatively,
DMVs might be an epiphenomenon, resulting from the over-expression of membrane-active proteins
that accumulate especially during the late stages of infection.
The mechanism responsible for DMV formation is not clear. In case of picornaviruses, it is
thought that single-membrane structures are the precursors of DMVs [79,80]. Nevertheless, DMV
formation might also involve the autophagy machinery, or at least several components thereof, by
a process analogous to the formation of autophagic vacuoles [76]. This hypothesis is supported by
the morphological resemblance of DMVs and autophagosomes. It has been shown that the inhibition or
stimulation of autophagy results in a modest inhibition or stimulation of poliovirus and coxsackie B3
virus yield, respectively, and there are also data supporting the involvement of autophagy in
the replication of rhinovirus 2 and 14 [125,126]. However, Brabec-Zaruba et al. [127] reported
that replication of rhinovirus 2 was insensitive to pharmacological manipulation of autophagy and did
not induce detectable modification of LC3. This discrepancy might be due to the use of different cell
types and experimental conditions.
The mechanism of DMV formation in case of HCV and coronaviruses is also unclear. Biochemical
analysis of isolated host cell membranes associated with HCV RNA and proteins identified markers of
the autophagy machinery, including LC3-II, the lipidated form of LC3 (LC3-II) that is generated upon
activation of the autophagy machinery [46]. However, the role of autophagy in the HCV replication
cycle is also a matter of controversy. For instance, immunolabeling did not identify LC3-II at those
sites where nonstructural proteins accumulate [48]. Moreover, different roles of autophagy for
the HCV replication cycle have been proposed. These include a role of autophagy in HCV RNA
translation [128], initiation of RNA replication [129,130], production of infectious virus particles [131]

Viruses 2014, 6 2843
or suppression of the innate antiviral defense [132,133]. To clarify these discrepancies, future studies
should combine biochemical and cell biological approaches with ultrastructural analyses.
The autophagy machinery might also be involved in the formation of virus-induced membrane
invaginations/spherules. For instance, Lee and coworkers provided evidence that DENV infection
enhances autophagolysosome formation and that inhibition of the autophagy machinery by 3-methyladenine
(3-MA) reduces DENV particle production [134]. However, the effects were moderate, arguing that
this pathway may contribute to DENV replication to only a minor extent. Moreover, autophagy
includes membrane wrapping, leading to double-membrane compartments involved in lysosomal
degradation whereas DENV-induced vesicles are invaginations. Finally, immunolabeling experiments
failed to detect lamp-1 at these vesicles [15], arguing against the involvement of lysosomes in the
formation of the DENV replication vesicles. It remains to be determined whether autophagy is actively
induced by these viruses to provide a compartment favoring replication or induced as a bystander
defense against infection leading to degradation of the replicase proteins [135].
Another membrane compartment frequently induced by (+) RNA viruses are convoluted
membranes (CMs) that were observed e.g., in SARS-CoV-, MHV-, WNV- or DENV-infected
cells [13,15,18,63,65,66,136]. Morphologically, CMs resemble smooth ER membranes, lack
ribosomes and in case of DENV are induced by the sole expression of NS4A [137,138]. CMs are
often associated with late stages of infection, suggesting that DMV formation might precede the
development of CMs. In SARS-CoV-infected cells, DMVs appear to be connected with CMs [63],
while in MHV-infected cells no such connections have been observed [65]. The role of CMs for
the viral replication cycle is not well understood. In case of WNVKUN, CMs are supposed to be the site
of polyprotein processing [18,136]. This conclusion is based primarily on the strong immunolabeling
for NS2B and NS3 and the absence of NS1 and NS4B. Since polyprotein cleavage occurs
co-translationally and thus, should happen at the rER, this model would require the formation of rather
stable processing intermediates that are transferred from the rER to the CMs where further cleavage
would occur. Alternatively, at least in case of DENV, CMs might represent a storage site for proteins
and lipids involved in viral replication that can be recruited to vesicles upon demand. The fact that
CMs are physically linked with ER-containing invaginations and contain NS3 would be consistent
with this assumption [15]. Along these lines, the fact that insects are cholesterol auxotrophs and lack
several enzymes in the cholesterol biosynthesis pathway [139], suggests that cholesterol might be
a key component of CM structures, which would explain their absence in infected insect cells [16].
5. Role of Viral Proteins in the Formation of the Replication Organelles
Viral replication complexes are targeted to the respective membranous organelle primarily by
nonstructural (NS) proteins rather than viral RNA [140]. These NS proteins seem to have some
specificity in recognizing organelle subpopulations and often contain multiple hydrophobic domains
implicated in membrane targeting and rearrangement. The molecular mechanisms orchestrating the
formation of these complex structures are still poorly understood, but it is clear that NS proteins, often
working in a concerted action, are key players in replication factory biogenesis.

Viruses 2014, 6 2844
5.1. Single-Membrane Vesicle Inducers
One well-studied example among the single-membrane vesicle inducers is BMV where it was
shown that the sole expression of the NS protein 1a is sufficient to induce the formation of
single-membrane spherules resembling the ones observed in infected cells. These spherules had
a diameter of 50–70 nm, resided in the ER lumen and were shown to be the site of viral RNA
synthesis [108]. In case of the insect nodavirus FHV, protein A and replication competent
RNA were required for induction of the spherules. Expression of protein A alone induced only
―zippering‖ of the surfaces of adjacent mitochondria, but did not induce spherules. Thus, protein A
is necessary, but not sufficient for spherule formation. Moreover, spherules were not formed
when replication-competent FHV RNA templates were expressed with a protein A mutant lacking
polymerase activity or when wild-type protein A was expressed with a replication-incompetent
FHV RNA template. Thus, the membranous FHV replication compartment requires both a viral
protein and active RNA synthesis [141].
Feline Calicivirus (FCV) infection results in rearrangement of intracellular membranes and
production of numerous membrane-bound vesicular structures on which viral genome replication is
thought to occur. Expression of individual FCV nonstructural proteins revealed that p30 induces
significant reorganization of the ER into large, fenestrated membrane networks, resembling the
structures found in infected cells [142]. Moreover, expression of p39 and p32, two additional FCV NS
proteins, induced extensive reorganization of the ER and the nuclear envelope suggesting that the ER
is the primary source of the membranous replication factory [142].
Flavivirus membrane rearrangements are mainly induced by NS4A, as suggested by recent
studies with WNVKUN and DENV [137,138], but it is unknown whether the same applies to NS4A of
TBEV. In case of DENV, NS4A is thought to contain a central peripheral membrane domain
that intercalates into the luminal leaflet of the ER membrane [138]. It is tempting to speculate that
NS4A oligomers [143] might dilate the luminal leaflet, resulting in membrane invaginations towards
the ER lumen. However, the sole expression of NS4A is not sufficient to induce ER membrane
invaginations that have been detected in infected cells. Instead, expression of NS4A lacking the
C-terminal 2K fragment (corresponding to fully processed NS4A) induced ER membrane
rearrangements reminiscent of CMs, whereas unprocessed NS4A/2K did not induce membrane
alterations [138]. These results provide strong evidence that processing at the NS4A-2K site is
required for the induction of membrane alterations. The critical role of polyprotein cleavage for
induction of membrane rearrangements is supported by studies conducted with WNVKUN. There it was
shown that a regulated cleavage of a NS4A/2K/4B precursor by the viral NS2B/3 protease is needed
for induction of membrane rearrangements [137]. However, the same study reported that expression of
full-length (uncleaved) WNVKUN NS4A/2K led to membrane alterations similar to those induced in
infected cells whereas the 2K fragment impaired the ability of NS4A to induce membrane
rearrangements. Whether this reflects a biological difference between WNVKUN and DENV NS4A or
is due to the use of alternative experimental approaches remains to be determined.
One of the most fascinating mechanisms employed by (+) RNA viruses to induce their replication
factories is used by SFV. It was shown that the spherules of SFV arise by blebbing at the surface of the
plasma membrane [93]. These blebs are internalized and after fusion with lysosomes give rise to large

Viruses 2014, 6 2845
cytoplasmic vacuoles. Formation of these membrane alterations requires the viral protein nsP1, which
has several functions. It has guanine-7-methyltransferase and guanylyltransferase activities
and thus is critically involved in capping of the viral RNAs [144–146], but at the same time has
affinity to lipids [89]. In fact, of the four NS proteins of SFV, only nsP1 has affinity for membranes,
and when expressed alone, it is specifically targeted to the inner surface of the plasma membrane [147].
NsP1 is a monotopic membrane protein and its affinity for membranes is dictated by an amphipathic
α-helix, located in the central region of the protein [148,149]. NsP1 has a specific affinity for
negatively charged phospholipids, which might account for its prevalent localization to the plasma
membrane, where such lipids are enriched. Membrane binding of nsP1 via its amphipathic α-helix is
essential for alphavirus replication [93]. However, nsP1 is not sufficient for cytoplasmic vacuole
formation. For instance, it was found that nsP3 contributes to the transport of the replicase polyprotein
from the plasma membrane to the surface of endosomes [150]. These results indicate that nsp1 has to
cooperate with other viral and cellular factors to allow formation of the cytoplasmic vacuoles.
Furthermore, in a recent study Kallio et al. [151] have shown that the size of the spherule is
dependent on the length of the RNA template, in contrast to what has been observed for FHV, another
spherule-inducer [141]. These results indicate that in addition to the NS proteins, the viral RNA
template itself critically determines the morphology of the membranous vesicles.
5.2. Double-Membrane Vesicle Inducers
In order to induce the variety of membrane alterations observed in CVB3 -infected cells (Figure 3F),
several membrane-remodeling mechanisms are required: induction of membrane curvature, membrane
fusion and membrane-membrane interactions [80]. These rearrangements require the enteroviral
proteins 2BC and 3A; their coexpression generates ER membrane-derived structures mimicking
those observed during viral infection [78]. Importantly, 2B and 2C both contain an amphipathic
α-helix [152–154], a well-known curvature-inducing motif [155]. Along the same lines, FMDV 2B and
2BC locate to the ER when expressed on their own and cause a swelling of ER cisternae [156].
In case of HCV, we recently found that a concerted action of NS3/4A, NS4B, NS5A and NS5B is
required to generate the membranous web. Furthermore, all these replicase components were capable
of inducing membrane vesiculation with NS5A having the highest potential to trigger membrane
curvature. Importantly, some of these NS5A-induced structures corresponded to DMVs [48].
In addition, NS4B also plays an important role in triggering rearrangements of intracellular
membranes [45]. NS4B is an integral membrane protein containing two N-terminal amphipathic
α-helices, a highly hydrophobic central core domain composed of four putative transmembrane
segments, and a highly conserved C-terminal domain that is thought to harbor two α-helices (reviewed
in [157]). A recent study has demonstrated that NS4B oligomerizes through multiple conserved
determinants and that oligomerization appears to be required for membranous web induction [158].
Indeed, mutations affecting the highly conserved C-terminal domain impairing NS4B self-interaction
resulted in the formation of aberrant DMVs arguing for a central role of NS4B in formation of
functional replication compartments [159].
Studies on arteriviruses revealed that the sole expression of EAV nsp2 and nsp3 is sufficient
to induce membrane structures similar to those generated during EAV infection [160]. Mutations

Viruses 2014, 6 2846
within nsp3, which is a tetra-spanning integral membrane protein, alter membrane rearrangements,
highlighting the importance of this protein for the biogenesis of EAV-induced DMVs [161]. In case of
SARS-CoV, nsp3, nsp4 and nsp6 were found to be sufficient to induce the formation of DMVs that are
similar to those observed in SARS-CoV-infected cells [162]. These DMVs were, however, smaller in
diameter, suggesting a role for other viral proteins or the presence of viral RNA in determining
the DMV morphology. Importantly, EM analysis of nsp4 mutants that are impaired in RNA replication
and virus growth, revealed an aberrant morphology of DMVs as well as an increased prevalence of
CMs [163]. Another important viral protein involved in inducing the SARS-CoV membranous
replication factory is nsp6, which is predicted to contain seven transmembrane segments and a
hydrophilic cytoplasmic domain [164]. Nsp6 was shown to induce vesicles containing Atg5 and LC3-II
as well as phosphatidylinositol-3-phosphate, thus sharing many features with omegasomes, which are
omega-shaped membrane compartments that are formed during activation of autophagy [164]. This
result suggests that autophagy might contribute to the formation of the membranous replication site of
SARS-CoV.
6. Conclusions and Future Perspectives
In the last couple of years, our knowledge of the architecture of the replication factories induced
by (+) RNA viruses has increased substantially. This is primarily due to the more widespread use of
ET and other high-resolution imaging methods. Nevertheless, our knowledge is still rather restricted to
descriptions of the morphologies of these complex structures, whereas our understanding of their
biogenesis in most cases is very rudimentary. More efforts are required to elucidate the role of the viral
proteins in the formation of the replication vesicles, to identify the involved cellular components and
the mechanisms used by these proteins to subvert and exploit cellular pathways to establish
membranous replication factories. This includes determination of the 3D structure of involved (viral)
proteins as well as evaluation of host cell factors and lipids contributing to biogenesis and activity of
the replication compartment. In addition, further studies are needed to understand how viruses
utilize these compartments to coordinate the different steps of their life cycle (replication, assembly
and release) in space and time to achieve efficient replication.
Acknowledgments
The authors are very grateful to Erik J. Snijder (Leiden University Medical Center, Leiden, The
Netherlands) and Kèvin Knoops (European Molecular Biology Laboratory, Grenoble, France) for
providing the unpublished pictures depicted in Figure 3B and 3D and to Montserrat Bárcena (Leiden
University Medical Center, Leiden, The Netherlands) and Tero Ahola (University of Helsinki,
Helsinki, Finland) for providing the unpublished pictures depicted in Figure 3F and 3J, respectively.
Figures 3A, 3C, 3E, 3G, 3H and 3I are reproduced with permission from [60], [67], [109], [79], [95]
and [86], respectively.
We also would like to thank Jason M. Mackenzie (University of Queensland, Brisbane, Australia),
Paul Ahlquist (Howard Hughes Medical Institute, Madison, WI, USA), Cristina Risco (Centro
Nacional de Biotecnología, Madrid, Spain), Georg A. Belov (University of Maryland, College Park,
MD, USA), Christiane Wobus (Washington University School of Medicine, St Louis, MO, USA), Paul

Viruses 2014, 6 2847
Britton (The Pirbright Institute, Compton, UK) and Alexey A. Agranovsky (Moscow State University,
Moscow, Russia) for providing other electron micrographs and 3D reconstructions.
Work in the authors’ laboratory was supported by the Deutsche Forschungsgemeinschaft
(Sonder-forschungsbereich 638, TP5 and Transregional Collaborative Research Project TR83, TP13).
Conflicts of Interest
The authors declare no conflict of interest.
References and Notes
1. Stapleton, J.T.; Foung, S.; Muerhoff, A.S.; Bukh, J.; Simmonds, P. The GB viruses: A review
and proposed classification of GBV-A, GBV-C (HGV), and GBV-D in genus Pegivirus within
the family Flaviviridae. J. Gen. Virol. 2011, 92, 233–246.
2. Simmonds, P. The Origin of Hepatitis C Virus. In Hepatitis C Virus: From Molecular Virology
to Antiviral Therapy, 82nd ed.; Bartenschlager, R., Ed.; Springer Berlin Heidelberg: Berlin,
Heidelberg, Germany, 2013; Volume 369.
3. Choo, Q.L.; Kuo, G.; Weiner, A.J.; Overby, L.R.; Bradley, D.W.; Houghton, M. Isolation of
a cDNA clone derived from a blood-borne non-A, non-B viral hepatitis genome. Science 1989,
244, 359–362.
4. Feinstone, S.M.; Kapikian, A.Z.; Purcell, R.H.; Alter, H.J.; Holland, P.V. Transfusion-associated
hepatitis not due to viral hepatitis type A or B. N. Engl. J. Med. 1975, 292, 767–770.
5. Lindenbach, B.D.; Thiel, H.-J.; Rice, C.M. Flaviviridae: The Viruses and Their Replication.
In Fields Virology, 5th ed.; Fields, B.N., Knipe, D.M., Howley, P., Eds.; Raven Press:
New York, NY, USA, 2001; pp. 1001–1152.
6. Mackenzie, J.S.; Gubler, D.J.; Petersen, L.R. Emerging flaviviruses: The spread and resurgence
of Japanese encephalitis, West Nile and dengue viruses. Nat. Med. 2004, 10,
S98–S109.
7. Simmonds, P.; Becher, P.; Collett, M.S.; Gould, E.A.; Heinz, F.X.; Meyers, G.; Monath, A.;
Pletnev, A.; Rice, C.M.; Stiasny, K.; et al. Family Flaviviridae. In Virus taxonomy: Classification
and nomenclature of viruses. Ninth report of the International Committee on Taxonomy of
Viruses; King, A.M.Q., Adams, M.J., Carstens, E.B., Lefkowitz, E.J., Eds.; Academic Press:
San Diego, CA, USA, 2012; pp. 1003–1020.
8. Miller, R.H.; Purcell, R.H. Hepatitis C virus shares amino acid sequence similarity with pestiviruses
and flaviviruses as well as members of two plant virus supergroups. Proc. Natl. Acad. Sci. USA
1990, 87, 2057–2061.
9. Matsumura, T.; Stollar, V.; Schlesinger, R.W. Studies on the nature of dengue viruses. V.
Structure and development of dengue virus in Vero cells. Virology 1971, 46, 344–355.
10. Stohlman, S.A.; Wisseman, C.L., Jr.; Eylar, O.R.; Silverman, D.J. Dengue virus-induced
modifications of host cell membranes. J. Virol. 1975, 16, 1017–1026.
11. Hase, T.; Summers, P.L.; Eckels, K.H.; Baze, W.B. An electron and immunoelectron
microscopic study of dengue-2 virus infection of cultured mosquito cells: Maturation events.
Arch. Virol. 1987, 92, 273–291.

Viruses 2014, 6 2848
12. Tu, W.C.; Chen, C.C.; Hou, R.F. Ultrastructural studies on the reproductive system of male
Aedes aegypti (Diptera: Culicidae) infected with dengue 2 virus. J. Med. Entomol. 1998, 35,
71–76.
13. Grief, C.; Galler, R.; Cortes, L.M.; Barth, O.M. Intracellular localisation of dengue-2 RNA in
mosquito cell culture using electron microscopic in situ hybridisation. Arch. Virol. 1997, 142,
2347–2357.
14. Mackenzie, J.M.; Jones, M.K.; Young, P.R. Improved membrane preservation of
flavivirus-infected cells with cryosectioning. J. Virol. Methods 1996, 56, 67–75.
15. Welsch, S.; Miller, S.; Romero-Brey, I.; Merz, A.; Bleck, C.K.; Walther, P.; Fuller, S.D.;
Antony, C.; Krijnse-Locker, J.; Bartenschlager, R. Composition and three-dimensional
architecture of the dengue virus replication and assembly sites. Cell. Host. Microbe 2009, 5,
365–375.
16. Junjhon, J.; Pennington, J.G.; Edwards, T.J.; Perera, R.; Lanman, J.; Kuhn, R.J. Ultrastructural
characterization and three-dimensional architecture of replication sites in dengue virus-infected
mosquito cells. J. Virol. 2014, 88, 4687–4697.
17. Shahar, A.; Lustig, S.; Akov, Y.; David, Y.; Schneider, P.; Friedmann, A.; Levin, R. West Nile
virions aligned along myelin lamellae in organotypic spinal cord cultures. J. Neurosci. Res. 1990,
26, 495–500.
18. Westaway, E.G.; Mackenzie, J.M.; Kenney, M.T.; Jones, M.K.; Khromykh, A.A. Ultrastructure
of Kunjin virus-infected cells: Colocalization of NS1 and NS3 with double-stranded RNA, and of
NS2B with NS3, in virus-induced membrane structures. J. Virol. 1997, 71, 6650–6661.
19. Mackenzie, J.M.; Westaway, E.G. Assembly and maturation of the flavivirus Kunjin virus appear
to occur in the rough endoplasmic reticulum and along the secretory pathway, respectively.
J. Virol. 2001, 75, 10787–10799.
20. Gillespie, L.K.; Hoenen, A.; Morgan, G.; Mackenzie, J.M. The endoplasmic reticulum provides
the membrane platform for biogenesis of the flavivirus replication complex. J. Virol. 2010, 84,
10438–10447.
21. Mackenzie, J.M.; Jones, M.K.; Young, P.R. Immunolocalization of the dengue virus
nonstructural glycoprotein NS1 suggests a role in viral RNA replication. Virology 1996, 220,
232–240.
22. Mackenzie, J.M.; Jones, M.K.; Westaway, E.G. Markers for trans-Golgi membranes and the
intermediate compartment localize to induced membranes with distinct replication functions in
flavivirus-infected cells. J. Virol. 1999, 73, 9555–9567.
23. Mackenzie, J.M.; Kenney, M.T.; Westaway, E.G. West Nile virus strain Kunjin NS5 polymerase
is a phosphoprotein localized at the cytoplasmic site of viral RNA synthesis. J. Gen. Virol. 2007,
88, 1163–1168.
24. Mackenzie, J.M.; Khromykh, A.A.; Jones, M.K.; Westaway, E.G. Subcellular localization and
some biochemical properties of the flavivirus Kunjin nonstructural proteins NS2A and NS4A.
Virology 1998, 245, 203–215.
25. Overby, A.K.; Popov, V.L.; Niedrig, M.; Weber, F. Tick-borne encephalitis virus delays
interferon induction and hides its double-stranded RNA in intracellular membrane vesicles.
J. Virol. 2010, 84, 8470–8483.

Viruses 2014, 6 2849
26. Pichlmair, A.; Reis e Sousa, C. Innate recognition of viruses. Immunity 2007, 27, 370–383.
27. Pichlmair, A.; Schulz, O.; Tan, C.P.; Rehwinkel, J.; Kato, H.; Takeuchi, O.; Akira, S.; Way, M.;
Schiavo, G.; Reis e Sousa, C. Activation of MDA5 requires higher-order RNA structures
generated during virus infection. J. Virol. 2009, 83, 10761–10769.
28. Randall, R.E.; Goodbourn, S. Interferons and viruses: An interplay between induction, signalling,
antiviral responses and virus countermeasures. J. Gen. Virol. 2008, 89, 1–47.
29. Rahman, S.; Matsumura, T.; Masuda, K.; Kanemura, K.; Fukunaga, T. Maturation site of dengue
type 2 virus in cultured mosquito C6/36 cells and Vero cells. Kobe J. Med. Sci. 1998, 44, 65–79.
30. Senigl, F.; Grubhoffer, L.; Kopecky, J. Differences in maturation of tick-borne encephalitis virus
in mammalian and tick cell line. Intervirology 2006, 49, 239–248.
31. Offerdahl, D.K.; Dorward, D.W.; Hansen, B.T.; Bloom, M.E. A three-dimensional comparison of
tick-borne flavivirus infection in mammalian and tick cell lines. PLoS One 2012, 7, e47912.
32. Schmaljohn, C.; Blair, C.D. Persistent infection of cultured mammalian cells by Japanese
encephalitis virus. J. Virol. 1977, 24, 580–589.
33. Brinton, M.A.; Fernandez, A.V. A replication-efficient mutant of West Nile virus is insensitive to
DI particle interference. Virology 1983, 129, 107–115.
34. Poidinger, M.; Coelen, R.J.; Mackenzie, J.S. Persistent infection of Vero cells by the flavivirus
Murray Valley encephalitis virus. J. Gen. Virol. 1991, 72, 573–578.
35. Miorin, L.; Romero-Brey, I.; Maiuri, P.; Hoppe, S.; Krijnse-Locker, J.; Bartenschlager, R.;
Marcello, A. Three-dimensional architecture of tick-borne encephalitis virus replication sites and
trafficking of the replicated RNA. J. Virol. 2013, 87, 6469–6481.
36. Hoenninger, V.M.; Rouha, H.; Orlinger, K.K.; Miorin, L.; Marcello, A.; Kofler, R.M.;
Mandl, C.W. Analysis of the effects of alterations in the tick-borne encephalitis virus
3'-noncoding region on translation and RNA replication using reporter replicons. Virology 2008,
377, 419–430.
37. Khromykh, A.A.; Westaway, E.G. Subgenomic replicons of the flavivirus Kunjin: Construction
and applications. J. Virol. 1997, 71, 1497–1505.
38. Jones, C.T.; Patkar, C.G.; Kuhn, R.J. Construction and applications of yellow fever virus
replicons. Virology 2005, 331, 247–259.
39. Matthews, V.; Robertson, T.; Kendrick, T.; Abdo, M.; Papadimitriou, J.; McMinn, P.
Morphological features of Murray Valley encephalitis virus infection in the central nervous
system of Swiss mice. Int. J. Exp. Pathol. 2000, 81, 31–40.
40. Hase, T.; Summers, P.L.; Ray, P. Entry and replication of Japanese encephalitis virus in cultured
neurogenic cells. J. Virol. Methods 1990, 30, 205–214.
41. Hase, T.; Summers, P.L.; Dubois, D.R. Ultrastructural changes of mouse brain neurons infected
with Japanese encephalitis virus. Int. J. Exp. Pathol. 1990, 71, 493–505.
42. Murphy, F.A.; Harrison, A.K.; Gary, G.W., Jr.; Whitfield, S.G.; Forrester, F.T. St. Louis
encephalitis virus infection in mice. Electron microscopic studies of central nervous system.
Lab. Investig. 1968, 19, 652–662.
43. Whitfield, S.G.; Murphy, F.A.; Sudia, W.D. St. Louis encephalitis virus: An ultrastructural study
of infection in a mosquito vector. Virology 1973, 56, 70–87.

Viruses 2014, 6 2850
44. McGavran, M.H.; White, J.D. Electron microscopic and immunofluorescent observations on
monkey liver and tissue culture cells infected with the Asibi strain of yellow fever virus liver and
tissue culture cells infected with the asibi strain of yellow fever virus. Am. J. Pathol. 1964, 45,
501–517.
45. Egger, D.; Wolk, B.; Gosert, R.; Bianchi, L.; Blum, H.E.; Moradpour, D.; Bienz, K. Expression
of hepatitis C virus proteins induces distinct membrane alterations including a candidate viral
replication complex. J. Virol. 2002, 76, 5974–5984.
46. Ferraris, P.; Blanchard, E.; Roingeard, P. Ultrastructural and biochemical analyses of hepatitis C
virus-associated host cell membranes. J. Gen. Virol. 2010, 91, 2230–2237.
47. Ferraris, P.; Beaumont, E.; Uzbekov, R.; Brand, D.; Gaillard, J.; Blanchard, E.; Roingeard, P.
Sequential biogenesis of host cell membrane rearrangements induced by hepatitis C virus
infection. Cell. Mol. Life Sci. 2013, 70, 1297–1306.
48. Romero-Brey, I.; Merz, A.; Chiramel, A.; Lee, J.Y.; Chlanda, P.; Haselman, U.;
Santarella-Mellwig, R.; Habermann, A.; Hoppe, S.; Kallis, S.; et al. Three-dimensional
architecture and biogenesis of membrane structures associated with hepatitis C virus replication.
PLoS Pathog. 2012, 8, e1003056.
49. Paul, D.; Hoppe, S.; Saher, G.; Krijnse-Locker, J.; Bartenschlager, R. Morphological and
biochemical characterization of the membranous hepatitis C virus replication compartment.
J. Virol. 2013, 87, 10612–10627.
50. Pietschmann, T.; Kaul, A.; Koutsoudakis, G.; Shavinskaya, A.; Kallis, S.; Steinmann, E.;
Abid, K.; Negro, F.; Dreux, M.; Cosset, F.L.; et al. Construction and characterization of
infectious intragenotypic and intergenotypic hepatitis C virus chimeras. Proc. Natl. Acad. Sci. USA
2006, 103, 7408–7413.
51. Chasey, D.; Roeder, P.L. Virus-like particles in bovine turbinate cells infected with bovine virus
diarrhoea/mucosal disease virus. Arch. Virol. 1981, 67, 325–332.
52. Gray, E.W.; Nettleton, P.F. The ultrastructure of cell cultures infected with border disease and
bovine virus diarrhoea viruses. J. Gen. Virol. 1987, 68, 2339–2346.
53. Kubovicova, E.; Makarevich, A.V.; Pivko, J.; Chrenek, P.; Grafenau, P.; Riha, L.; Sirotkin, A.V.;
Louda, F. Alteration in ultrastructural morphology of bovine embryos following subzonal
microinjection of bovine viral diarrhea virus (BVDV). Zygote 2008, 16, 187–193.
54. Birk, A.V.; Dubovi, E.J.; Cohen-Gould, L.; Donis, R.; Szeto, H.H. Cytoplasmic vacuolization
responses to cytopathic bovine viral diarrhoea virus. Virus Res. 2008, 132, 76–85.
55. Weiskircher, E.; Aligo, J.; Ning, G.; Konan, K.V. Bovine viral diarrhea virus NS4B protein is
an integral membrane protein associated with Golgi markers and rearranged host membranes.
Virol. J. 2009, 6, 185.
56. Schmeiser, S.; Mast, J.; Thiel, H.J.; Konig, M. Morphogenesis of pestiviruses: New insights from
ultrastructural studies of strain Giraffe-1. J. Virol. 2014, 88, 2717–2724.
57. Welsch, S.; Keppler, O.T.; Habermann, A.; Allespach, I.; Krijnse-Locker, J.; Krausslich, H.G.
HIV-1 buds predominantly at the plasma membrane of primary human macrophages.
PLoS Pathog. 2007, 3, e36.
58. Kolesnikova, L.; Berghofer, B.; Bamberg, S.; Becker, S. Multivesicular bodies as a platform for
formation of the Marburg virus envelope. J. Virol. 2004, 78, 12277–12287.

Viruses 2014, 6 2851
59. Mittler, E.; Kolesnikova, L.; Strecker, T.; Garten, W.; Becker, S. Role of the transmembrane
domain of marburg virus surface protein GP in assembly of the viral envelope. J. Virol. 2007, 81,
3942–3948.
60. Kopek, B.G.; Perkins, G.; Miller, D.J.; Ellisman, M.H.; Ahlquist, P. Three-dimensional
analysis of a viral RNA replication complex reveals a virus-induced mini-organelle. PLoS Biol.
2007, 5, e220.
61. Lanman, J.; Crum, J.; Deerinck, T.J.; Gaietta, G.M.; Schneemann, A.; Sosinsky, G.E.;
Ellisman, M.H.; Johnson, J.E. Visualizing flock house virus infection in Drosophila cells with
correlated fluorescence and electron microscopy. J. Struct. Biol. 2008, 161, 439–446.
62. Snijder, E.J.; van der Meer, Y.; Zevenhoven-Dobbe, J.; Onderwater, J.J.; van der Meulen, J.;
Koerten, H.K.; Mommaas, A.M. Ultrastructure and origin of membrane vesicles associated
with the severe acute respiratory syndrome coronavirus replication complex. J. Virol.
2006, 80, 5927–5940.
63. Knoops, K.; Kikkert, M.; Worm, S.H.; Zevenhoven-Dobbe, J.C.; van der Meer, Y.; Koster, A.J.;
Mommaas, A.M.; Snijder, E.J. SARS-coronavirus replication is supported by a reticulovesicular
network of modified endoplasmic reticulum. PLoS Biol. 2008, 6, e226.
64. Gosert, R.; Kanjanahaluethai, A.; Egger, D.; Bienz, K.; Baker, S.C. RNA replication of mouse
hepatitis virus takes place at double-membrane vesicles. J. Virol. 2002, 76, 3697–3708.
65. Ulasli, M.; Verheije, M.H.; de Haan, C.A.; Reggiori, F. Qualitative and quantitative
ultrastructural analysis of the membrane rearrangements induced by coronavirus. Cell. Microbiol.
2010, 12, 844–861.
66. De Wilde, A.H.; Raj, V.S.; Oudshoorn, D.; Bestebroer, T.M.; van, N.S.; Limpens, R.W.;
Posthuma, C.C.; van der Meer, Y.; Barcena, M.; Haagmans, B.L.; et al. MERS-coronavirus
replication induces severe in vitro cytopathology and is strongly inhibited by cyclosporin A
or interferon-alpha treatment. J. Gen. Virol. 2013, 94, 1749–1760.
67. Maier, H.J.; Hawes, P.C.; Cottam, E.M.; Mantell, J.; Verkade, P.; Monaghan, P.; Wileman, T.;
Britton, P. Infectious bronchitis virus generates spherules from zippered endoplasmic reticulum
membranes. MBio 2013, 4, e00801–e00813.
68. Pedersen, K.W.; van der Meer, Y.; Roos, N.; Snijder, E.J. Open reading frame 1a-encoded
subunits of the arterivirus replicase induce endoplasmic reticulum-derived double-membrane
vesicles which carry the viral replication complex. J. Virol. 1999, 73, 2016–2026.
69. Knoops, K.; Barcena, M.; Limpens, R.W.; Koster, A.J.; Mommaas, A.M.; Snijder, E.J.
Ultrastructural characterization of arterivirus replication structures: Reshaping the endoplasmic
reticulum to accommodate viral RNA synthesis. J. Virol. 2012, 86, 2474–2487.
70. Kallman, F.; Williams, R.C.; Dulbecco, R.; Vogt, M. Fine structure of changes produced in
cultured cells sampled at specified intervals during a single growth cycle of polio virus.
J. Biophys. Biochem. Cytol. 1958, 4, 301–308.
71. Bienz, K.; Egger, D.; Pasamontes, L. Association of polioviral proteins of the P2 genomic region
with the viral replication complex and virus-induced membrane synthesis as visualized by
electron microscopic immunocytochemistry and autoradiography. Virology 1987, 160, 220–226.
72. Egger, D.; Bienz, K. Intracellular location and translocation of silent and active poliovirus
replication complexes. J. Gen. Virol. 2005, 86, 707–718.

Viruses 2014, 6 2852
73. Rust, R.C.; Landmann, L.; Gosert, R.; Tang, B.L.; Hong, W.; Hauri, H.P.; Egger, D.; Bienz, K.
Cellular COPII proteins are involved in production of the vesicles that form the poliovirus
replication complex. J. Virol. 2001, 75, 9808–9818.
74. Bienz, K.; Egger, D.; Rasser, Y.; Bossart, W. Intracellular distribution of poliovirus proteins and
the induction of virus-specific cytoplasmic structures. Virology 1983, 131, 39–48.
75. Hsu, N.Y.; Ilnytska, O.; Belov, G.; Santiana, M.; Chen, Y.H.; Takvorian, P.M.; Pau, C.;
van der Schaar, H.; Kaushik-Basu, N.; Balla, T.; et al. Viral reorganization of the secretory
pathway generates distinct organelles for RNA replication. Cell 2010, 141, 799–811.
76. Schlegel, A.; Giddings, T.H., Jr.; Ladinsky, M.S.; Kirkegaard, K. Cellular origin and ultrastructure
of membranes induced during poliovirus infection. J. Virol. 1996, 70, 6576–6588.
77. Bienz, K.; Egger, D.; Rasser, Y.; Bossart, W. Kinetics and location of poliovirus macromolecular
synthesis in correlation to virus-induced cytopathology. Virology 1980, 100, 390–399.
78. Suhy, D.A.; Giddings, T.H., Jr.; Kirkegaard, K. Remodeling the endoplasmic reticulum by
poliovirus infection and by individual viral proteins: An autophagy-like origin for virus-induced
vesicles. J. Virol. 2000, 74, 8953–8965.
79. Belov, G.A.; Nair, V.; Hansen, B.T.; Hoyt, F.H.; Fischer, E.R.; Ehrenfeld, E. Complex dynamic
development of poliovirus membranous replication complexes. J. Virol. 2012, 86, 302–312.
80. Limpens, R.W.; van der Schaar, H.M.; Kumar, D.; Koster, A.J.; Snijder, E.J.; van Kuppeveld, F.J.;
Barcena, M. The transformation of enterovirus replication structures: A three-dimensional study
of single- and double-membrane compartments. MBio 2011, 2, doi:10.1128/mBio.00166-11.
81. Monaghan, P.; Cook, H.; Jackson, T.; Ryan, M.; Wileman, T. The ultrastructure of the
developing replication site in foot-and-mouth disease virus-infected BHK-38 cells. J. Gen. Virol.
2004, 85, 933–946.
82. Frey, T.K. Molecular biology of rubella virus. Adv. Virus Res. 1994, 44, 69–160.
83. Fontana, J.; Tzeng, W.P.; Calderita, G.; Fraile-Ramos, A.; Frey, T.K.; Risco, C. Novel
replication complex architecture in rubella replicon-transfected cells. Cell. Microbiol. 2007, 9,
875–890.
84. Lee, J.Y.; Marshall, J.A.; Bowden, D.S. Characterization of rubella virus replication complexes
using antibodies to double-stranded RNA. Virology 1994, 200, 307–312.
85. Magliano, D.; Marshall, J.A.; Bowden, D.S.; Vardaxis, N.; Meanger, J.; Lee, J.Y. Rubella virus
replication complexes are virus-modified lysosomes. Virology 1998, 240, 57–63.
86. Fontana, J.; Lopez-Iglesias, C.; Tzeng, W.P.; Frey, T.K.; Fernandez, J.J.; Risco, C.
Three-dimensional structure of Rubella virus factories. Virology 2010, 405, 579–591.
87. Risco, C.; Carrascosa, J.L.; Frey, T.K. Structural maturation of rubella virus in the Golgi
complex. Virology 2003, 312, 261–269.
88. Froshauer, S.; Kartenbeck, J.; Helenius, A. Alphavirus RNA replicase is located on the
cytoplasmic surface of endosomes and lysosomes. J. Cell. Biol. 1988, 107, 2075–2086.
89. Kujala, P.; Ikaheimonen, A.; Ehsani, N.; Vihinen, H.; Auvinen, P.; Kaariainen, L. Biogenesis of
the Semliki Forest virus RNA replication complex. J. Virol. 2001, 75, 3873–3884.
90. Friedman, R.M.; Levin, J.G.; Grimley, P.M.; Berezesky, I.K. Membrane-associated replication
complex in arbovirus infection. J. Virol. 1972, 10, 504–515.

Viruses 2014, 6 2853
91. Grimley, P.M.; Berezesky, I.K.; Friedman, R.M. Cytoplasmic structures associated with an
arbovirus infection: Loci of viral ribonucleic acid synthesis. J. Virol. 1968, 2, 1326–1338.
92. Grimley, P.M.; Levin, J.G.; Berezesky, I.K.; Friedman, R.M. Specific membranous structures
associated with the replication of group A arboviruses. J. Virol. 1972, 10, 492–503.
93. Spuul, P.; Balistreri, G.; Kaariainen, L.; Ahola, T. Phosphatidylinositol 3-kinase-, actin-, and
microtubule-dependent transport of Semliki Forest Virus replication complexes from the plasma
membrane to modified lysosomes. J. Virol. 2010, 84, 7543–7557.
94. Green, K.Y.; Chanock, R.M.; Kapikian, A.Z. Human caliciviruses. In Fields Virology;
Knipe, D.M., Howley, P.M., Eds.; Lippincott Williams and Wilkins: Philadelphia, PA, USA,
2001; pp. 841–874.
95. Wobus, C.E.; Karst, S.M.; Thackray, L.B.; Chang, K.O.; Sosnovtsev, S.V.; Belliot, G.; Krug, A.;
Mackenzie, J.M.; Green, K.Y.; Virgin, H.W. Replication of Norovirus in cell culture reveals a
tropism for dendritic cells and macrophages. PLoS Biol. 2004, 2, e432.
96. Hyde, J.L.; Sosnovtsev, S.V.; Green, K.Y.; Wobus, C.; Virgin, H.W.; Mackenzie, J.M. Mouse
norovirus replication is associated with virus-induced vesicle clusters originating from
membranes derived from the secretory pathway. J. Virol. 2009, 83, 9709–9719.
97. Duizer, E.; Schwab, K.J.; Neill, F.H.; Atmar, R.L.; Koopmans, M.P.; Estes, M.K. Laboratory
efforts to cultivate noroviruses. J. Gen. Virol. 2004, 85, 79–87.
98. Green, K.Y.; Mory, A.; Fogg, M.H.; Weisberg, A.; Belliot, G.; Wagner, M.; Mitra, T.; Ehrenfeld, E.;
Cameron, C.E.; Sosnovtsev, S.V. Isolation of enzymatically active replication complexes from
feline calicivirus-infected cells. J. Virol. 2002, 76, 8582–8595.
99. Mas, P.; Beachy, R.N. Replication of tobacco mosaic virus on endoplasmic reticulum and
role of the cytoskeleton and virus movement protein in intracellular distribution of viral RNA.
J. Cell. Biol. 1999, 147, 945–958.
100. Schaad, M.C.; Jensen, P.E.; Carrington, J.C. Formation of plant RNA virus replication
complexes on membranes: Role of an endoplasmic reticulum-targeted viral protein. EMBO J.
1997, 16, 4049–4059.
101. Turner, K.A.; Sit, T.L.; Callaway, A.S.; Allen, N.S.; Lommel, S.A. Red clover necrotic
mosaic virus replication proteins accumulate at the endoplasmic reticulum. Virology 2004, 320,
276–290.
102. De, G.M.; Coscoy, L.; Jaspars, E.M. Localization and biochemical characterization of alfalfa
mosaic virus replication complexes. Virology 1993, 194, 878–881.
103. Hayes, R.J.; Buck, K.W. Complete replication of a eukaryotic virus RNA in vitro by a purified
RNA-dependent RNA polymerase. Cell 1990, 63, 363–368.
104. Panavas, T.; Hawkins, C.M.; Panaviene, Z.; Nagy, P.D. The role of the p33:p33/p92 interaction
domain in RNA replication and intracellular localization of p33 and p92 proteins of Cucumber
necrosis tombusvirus. Virology 2005, 338, 81–95.
105. Weber-Lotfi, F.; Dietrich, A.; Russo, M.; Rubino, L. Mitochondrial targeting and membrane
anchoring of a viral replicase in plant and yeast cells. J. Virol. 2002, 76, 10485–10496.
106. Restrepo-Hartwig, M.A.; Ahlquist, P. Brome mosaic virus helicase- and polymerase-like proteins
colocalize on the endoplasmic reticulum at sites of viral RNA synthesis. J. Virol. 1996, 70,
8908–8916.

Viruses 2014, 6 2854
107. Den Boon, J.A.; Chen, J.; Ahlquist, P. Identification of sequences in Brome mosaic virus
replicase protein 1a that mediate association with endoplasmic reticulum membranes. J. Virol.
2001, 75, 12370–12381.
108. Schwartz, M.; Chen, J.; Janda, M.; Sullivan, M.; den, B.J.; Ahlquist, P. A positive-strand RNA
virus replication complex parallels form and function of retrovirus capsids. Mol. Cell. 2002, 9,
505–514.
109. Gushchin, V.A.; Solovyev, A.G.; Erokhina, T.N.; Morozov, S.Y.; Agranovsky, A.A. Beet
yellows virus replicase and replicative compartments: Parallels with other RNA viruses.
Front. Microbiol. 2013, 4, 38.
110. Cronshaw, J.; Hoefert, L.; Esau, K. Ultrastructural features of Beta leaves infected with beet
yellows virus. J. Cell. Biol. 1966, 31, 429–443.
111. Esau, K.; Cronshaw, J.; Hoefert, L.L. Relation of beet yellows virus to the phloem and to
movement in the sieve tube. J. Cell. Biol. 1967, 32, 71–87.
112. Erokhina, T.N.; Vitushkina, M.V.; Zinovkin, R.A.; Lesemann, D.E.; Jelkmann, W.; Koonin, E.V.;
Agranovsky, A.A. Ultrastructural localization and epitope mapping of the methyltransferase-like
and helicase-like proteins of Beet yellows virus. J. Gen. Virol. 2001, 82, 1983–1994.
113. Zinovkin, R.A.; Erokhina, T.N.; Lesemann, D.E.; Jelkmann, W.; Agranovsky, A.A. Processing
and subcellular localization of the leader papain-like proteinase of Beet yellows closterovirus.
J. Gen. Virol. 2003, 84, 2265–2270.
114. Wang, J.; Stewart, L.R.; Kiss, Z.; Falk, B.W. Lettuce infectious yellows virus (LIYV) RNA
1-encoded P34 is an RNA-binding protein and exhibits perinuclear localization. Virology 2010,
403, 67–77.
115. Kim, K.S.; Gonsalves, D.; Teliz, D.; Lee, K.W. Ultrastructure and mitochondrial vesiculation
associated with closterovirus-like particles in leafroll-diseased grapevines. Phytopathology 1989,
79, 357–360.
116. Faoro, F.; Carzaniga, R. Cyotchemistry and immuno-cytochemistry of the inclusion
bodies induced by grapevine leafroll-associated closteroviruses GLRaV-1 and GLRaV-3.
Riv. Patol. Veg. 1995, 5, 85–94.
117. Koonin, E.V. The phylogeny of RNA-dependent RNA polymerases of positive-strand RNA
viruses. J. Gen. Virol. 1991, 72, 2197–2206.
118. Koonin, E.V.; Dolja, V.V. Evolution and taxonomy of positive-strand RNA viruses: Implications
of comparative analysis of amino acid sequences. Crit Rev. Biochem. Mol. Biol. 1993, 28,
375–430.
119. Paul, D.; Bartenschlager, R. Architecture and biogenesis of plus-strand RNA virus replication
factories. World J. Virol. 2013, 2, 32–48.
120. Takeda, N.; Kuhn, R.J.; Yang, C.F.; Takegami, T.; Wimmer, E. Initiation of poliovirus
plus-strand RNA synthesis in a membrane complex of infected HeLa cells. J. Virol. 1986, 60,
43–53.
121. Xiang, W.; Cuconati, A.; Hope, D.; Kirkegaard, K.; Wimmer, E. Complete protein linkage map
of poliovirus P3 proteins: Interaction of polymerase 3Dpol with VPg and with genetic variants of
3AB. J. Virol. 1998, 72, 6732–6741.

Viruses 2014, 6 2855
122. Wessels, E.; Notebaart, R.A.; Duijsings, D.; Lanke, K.; Vergeer, B.; Melchers, W.J.;
van Kuppeveld, F.J. Structure-function analysis of the coxsackievirus protein 3A: Identification
of residues important for dimerization, viral rna replication, and transport inhibition. J. Biol. Chem.
2006, 281, 28232–28243.
123. Fujita, K.; Krishnakumar, S.S.; Franco, D.; Paul, A.V.; London, E.; Wimmer, E. Membrane
topography of the hydrophobic anchor sequence of poliovirus 3A and 3AB proteins and the
functional effect of 3A/3AB membrane association upon RNA replication. Biochemistry 2007,
46, 5185–5199.
124. Bienz, K.; Egger, D.; Pfister, T.; Troxler, M. Structural and functional characterization of the
poliovirus replication complex. J. Virol. 1992, 66, 2740–2747.
125. Jackson, W.T.; Giddings, T.H., Jr.; Taylor, M.P.; Mulinyawe, S.; Rabinovitch, M.; Kopito, R.R.;
Kirkegaard, K. Subversion of cellular autophagosomal machinery by RNA viruses. PLoS Biol.
2005, 3, e156.
126. Wong, J.; Zhang, J.; Si, X.; Gao, G.; Mao, I.; McManus, B.M.; Luo, H. Autophagosome supports
coxsackievirus B3 replication in host cells. J. Virol. 2008, 82, 9143–9153.
127. Brabec-Zaruba, M.; Berka, U.; Blaas, D.; Fuchs, R. Induction of autophagy does not affect
human rhinovirus type 2 production. J. Virol. 2007, 81, 10815–10817.
128. Dreux, M.; Gastaminza, P.; Wieland, S.F.; Chisari, F.V. The autophagy machinery is required to
initiate hepatitis C virus replication. Proc. Natl. Acad. Sci. USA 2009, 106, 14046–14051.
129. Sir, D.; Chen, W.L.; Choi, J.; Wakita, T.; Yen, T.S.; Ou, J.H. Induction of incomplete
autophagic response by hepatitis C virus via the unfolded protein response. Hepatology 2008, 48,
1054–1061.
130. Guevin, C.; Manna, D.; Belanger, C.; Konan, K.V.; Mak, P.; Labonte, P. Autophagy protein
ATG5 interacts transiently with the hepatitis C virus RNA polymerase (NS5B) early during
infection. Virology 2010, 405, 1–7.
131. Tanida, I.; Fukasawa, M.; Ueno, T.; Kominami, E.; Wakita, T.; Hanada, K. Knockdown of
autophagy-related gene decreases the production of infectious hepatitis C virus particles.
Autophagy 2009, 5, 937–945.
132. Ke, P.Y.; Chen, S.S. Autophagy: A novel guardian of HCV against innate immune response.
Autophagy 2011, 7, 533–535.
133. Shrivastava, S.; Raychoudhuri, A.; Steele, R.; Ray, R.; Ray, R.B. Knockdown of autophagy
enhances the innate immune response in hepatitis C virus-infected hepatocytes. Hepatology
2011, 53, 406–414.
134. Lee, Y.R.; Lei, H.Y.; Liu, M.T.; Wang, J.R.; Chen, S.H.; Jiang-Shieh, Y.F.; Lin, Y.S.;
Yeh, T.M.; Liu, C.C.; Liu, H.S. Autophagic machinery activated by dengue virus enhances virus
replication. Virology 2008, 374, 240–248.
135. Wileman, T. Aggresomes and autophagy generate sites for virus replication. Science 2006, 312,
875–878.
136. Westaway, E.G.; Khromykh, A.A.; Kenney, M.T.; Mackenzie, J.M.; Jones, M.K. Proteins C and
NS4B of the flavivirus Kunjin translocate independently into the nucleus. Virology 1997, 234,
31–41.

Viruses 2014, 6 2856
137. Roosendaal, J.; Westaway, E.G.; Khromykh, A.; Mackenzie, J.M. Regulated cleavages at the
West Nile virus NS4A-2K-NS4B junctions play a major role in rearranging cytoplasmic
membranes and Golgi trafficking of the NS4A protein. J. Virol. 2006, 80, 4623–4632.
138. Miller, S.; Kastner, S.; Krijnse-Locker, J.; Buhler, S.; Bartenschlager, R. The non-structural
protein 4A of dengue virus is an integral membrane protein inducing membrane alterations in
a 2K-regulated manner. J. Biol. Chem. 2007, 282, 8873–8882.
139. Rawson, R.B. The SREBP pathway--insights from Insigs and insects. Nat. Rev. Mol. Cell. Biol.
2003, 4, 631–640.
140. Salonen, A.; Ahola, T.; Kaariainen, L. Viral RNA replication in association with cellular
membranes. Curr. Top. Microbiol. Immunol. 2005, 285, 139–173.
141. Kopek, B.G.; Settles, E.W.; Friesen, P.D.; Ahlquist, P. Nodavirus-induced membrane
rearrangement in replication complex assembly requires replicase protein a, RNA templates, and
polymerase activity. J. Virol. 2010, 84, 12492–12503.
142. Bailey, D.; Kaiser, W.J.; Hollinshead, M.; Moffat, K.; Chaudhry, Y.; Wileman, T.;
Sosnovtsev, S.V.; Goodfellow, I.G. Feline calicivirus p32, p39 and p30 proteins localize to
the endoplasmic reticulum to initiate replication complex formation. J. Gen. Virol. 2010, 91,
739–749.
143. Stern, O.; Hung, Y.F.; Valdau, O.; Yaffe, Y.; Harris, E.; Hoffmann, S.; Willbold, D.; Sklan, E.H.
An N-terminal amphipathic helix in dengue virus nonstructural protein 4A mediates
oligomerization and is essential for replication. J. Virol. 2013, 87, 4080–4085.
144. Ahola, T.; Kaariainen, L. Reaction in alphavirus mRNA capping: Formation of a covalent
complex of nonstructural protein nsP1 with 7-methyl-GMP. Proc. Natl. Acad. Sci. USA 1995, 92,
507–511.
145. Ahola, T.; Laakkonen, P.; Vihinen, H.; Kaariainen, L. Critical residues of Semliki Forest virus
RNA capping enzyme involved in methyltransferase and guanylyltransferase-like activities.
J. Virol. 1997, 71, 392–397.
146. Laakkonen, P.; Ahola, T.; Kaariainen, L. The effects of palmitoylation on membrane association
of Semliki forest virus RNA capping enzyme. J. Biol. Chem. 1996, 271, 28567–28571.
147. Peranen, J.; Laakkonen, P.; Hyvonen, M.; Kaariainen, L. The alphavirus replicase protein nsP1 is
membrane-associated and has affinity to endocytic organelles. Virology 1995, 208, 610–620.
148. Ahola, T.; Lampio, A.; Auvinen, P.; Kaariainen, L. Semliki Forest virus mRNA capping enzyme
requires association with anionic membrane phospholipids for activity. EMBO J. 1999, 18,
3164–3172.
149. Lampio, A.; Kilpelainen, I.; Pesonen, S.; Karhi, K.; Auvinen, P.; Somerharju, P.; Kaariainen, L.
Membrane binding mechanism of an RNA virus-capping enzyme. J. Biol. Chem. 2000, 275,
37853–37859.
150. Salonen, A.; Vasiljeva, L.; Merits, A.; Magden, J.; Jokitalo, E.; Kaariainen, L. Properly folded
nonstructural polyprotein directs the semliki forest virus replication complex to the endosomal
compartment. J. Virol. 2003, 77, 1691–1702.
151. Kallio, K.; Hellstrom, K.; Balistreri, G.; Spuul, P.; Jokitalo, E.; Ahola, T. Template RNA length
determines the size of replication complex spherules for Semliki Forest virus. J. Virol. 2013, 87,
9125–9134.

Viruses 2014, 6 2857
152. Paul, A.V.; Molla, A.; Wimmer, E. Studies of a putative amphipathic helix in the N-terminus of
poliovirus protein 2C. Virology 1994, 199, 188–199.
153. Echeverri, A.C.; Dasgupta, A. Amino terminal regions of poliovirus 2C protein mediate
membrane binding. Virology 1995, 208, 540–553.
154. Van Kuppeveld, F.J.; Melchers, W.J.; Kirkegaard, K.; Doedens, J.R. Structure-function analysis
of coxsackie B3 virus protein 2B. Virology 1997, 227, 111–118.
155. Miller, S.; Krijnse-Locker, J. Modification of intracellular membrane structures for virus
replication. Nat. Rev. Microbiol. 2008, 6, 363–374.
156. Moffat, K.; Knox, C.; Howell, G.; Clark, S.J.; Yang, H.; Belsham, G.J.; Ryan, M.; Wileman, T.
Inhibition of the secretory pathway by foot-and-mouth disease virus 2BC protein is reproduced
by coexpression of 2B with 2C, and the site of inhibition is determined by the subcellular
location of 2C. J. Virol. 2007, 81, 1129–1139.
157. Gouttenoire, J.; Penin, F.; Moradpour, D. Hepatitis C virus nonstructural protein 4B: A journey
into unexplored territory. Rev. Med. Virol. 2010, 20, 117–129.
158. Gouttenoire, J.; Roingeard, P.; Penin, F.; Moradpour, D. Amphipathic alpha-helix AH2 is a
major determinant for the oligomerization of hepatitis C virus nonstructural protein 4B. J. Virol.
2010, 84, 12529–12537.
159. Paul, D.; Romero-Brey, I.; Gouttenoire, J.; Stoitsova, S.; Krijnse-Locker, J.; Moradpour, D.;
Bartenschlager, R. NS4B self-interaction through conserved C-terminal elements is required
for the establishment of functional hepatitis C virus replication complexes. J. Virol. 2011, 85,
6963–6976.
160. Snijder, E.J.; van, T.H.; Roos, N.; Pedersen, K.W. Non-structural proteins 2 and 3 interact
to modify host cell membranes during the formation of the arterivirus replication complex.
J. Gen. Virol. 2001, 82, 985–994.
161. Posthuma, C.C.; Pedersen, K.W.; Lu, Z.; Joosten, R.G.; Roos, N.; Zevenhoven-Dobbe, J.C.;
Snijder, E.J. Formation of the arterivirus replication/transcription complex: A key role for
nonstructural protein 3 in the remodeling of intracellular membranes. J. Virol. 2008, 82,
4480–4491.
162. Angelini, M.M.; Akhlaghpour, M.; Neuman, B.W.; Buchmeier, M.J. Severe acute respiratory
syndrome coronavirus nonstructural proteins 3, 4, and 6 induce double-membrane vesicles. MBio
2013, 4, doi:10.1128/mBio.00524-13.
163. Gadlage, M.J.; Sparks, J.S.; Beachboard, D.C.; Cox, R.G.; Doyle, J.D.; Stobart, C.C.; Denison, M.R.
Murine hepatitis virus nonstructural protein 4 regulates virus-induced membrane modifications
and replication complex function. J. Virol. 2010, 84, 280–290.
164. Cottam, E.M.; Maier, H.J.; Manifava, M.; Vaux, L.C.; Chandra-Schoenfelder, P.; Gerner, W.;
Britton, P.; Ktistakis, N.T.; Wileman, T. Coronavirus nsp6 proteins generate autophagosomes
from the endoplasmic reticulum via an omegasome intermediate. Autophagy 2011, 7, 1335–1347.
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