key: cord-259233-smmhhroe
authors: de Armas‐Rillo, Laura; Valera, María‐Soledad; Marrero‐Hernández, Sara; Valenzuela‐Fernández, Agustín
title: Membrane dynamics associated with viral infection
date: 2016-01-28
journal: Rev Med Virol
DOI: 10.1002/rmv.1872
sha: 
doc_id: 259233
cord_uid: smmhhroe

Viral replication and spreading are fundamental events in the viral life cycle, accounting for the assembly and egression of nascent virions, events that are directly associated with viral pathogenesis in target hosts. These processes occur in cellular compartments that are modified by specialized viral proteins, causing a rearrangement of different cell membranes in infected cells and affecting the ER, mitochondria, Golgi apparatus, vesicles and endosomes, as well as processes such as autophagic membrane flux. In fact, the activation or inhibition of membrane trafficking and other related activities are fundamental to ensure the adequate replication and spreading of certain viruses. In this review, data will be presented that support the key role of membrane dynamics in the viral cycle, especially in terms of the assembly, egression and infection processes. By defining how viruses orchestrate these events it will be possible to understand how they successfully complete their route of infection, establishing viral pathogenesis and provoking disease. © 2015 The Authors Reviews in Medical Virology Published by John Wiley & Sons, Ltd.

Viruses are small structures that lack the metabolic pathways and structures necessary to ensure their own survival, relying on their host's machinery to replicate their genome and spread their progeny. Accordingly, viruses have developed strategies to enter cells and exploit their structures to replicate. These strategies also serve to evade immune responses, such as those involving toll-like receptors and autophagic-mediated antigen presentation [1] [2] [3] [4] . Similarly, viruses use the target cell's main trafficking pathways to ensure their propagation, exploiting the endosome or vesicular compartments by recruiting the clathrin, coatomer protein complex (COPI) I and II (Figure 1 ), the endosomal-sorting complex required for transport (ESCRT) and their accessory proteins (reviewed by [1, 2, 5] : Figure 2 ), as well as small guanosine triphosphatases (GTPases) [2] . This is evident during neutrophil-mediated phagocytosis, where microorganisms can be cleared by granule and vesicle secretion [6] . Therefore, determining how viruses use and rearrange intracellular organelles during their biological cycle is an important goal that will aid the development of new antiviral strategies, and our understanding Figure 1 . Viral factories and virus-triggered autophagic membrane flux for replication and egression. Some viruses achieve replication by exploiting the cell's membrane transport pathways, thereby generating membrane organelles named Viral Factories (VFs). These VFs are organised by different viral proteins, and they represent specialized compartments for viral-gene replication, morphogenesis, export, maturation and release. Moreover, these compartments also serve to override or evade the immune responses directed against viral genomes. Viral proteins can enter secretory pathways by co-translational translocation into the ER in order for them to be further transported to the Golgi complex, either in vesicles or in a coatomer protein complex (COP) II-dependent manner. Viral complexes formed inside the VFs communicate with vesicles, mitochondria, Golgi cisternae and ER membranes. This interaction allows viral complexes to be transported through the Golgi network to the plasma membrane and it promotes their final release as viral particles. Alternatively, some viruses take advantage of the host's autophagic machinery for their own replication and pathogenesis. Viruses first initiate the formation of vesicles that bear key autophagic proteins, such as Beclin-1 and LC3, capturing portions of membranes from the ER and other cytoplasmic elements. This assembly evolves toward an immature double-membrane vesicle (DMV) that serves as an aggresome compartment to recruit viruses or newly formed viral replication complexes. Several RNA viruses induce the formation of these autophagosome-like vesicles (also referred to as DMVs) to enhance viral replication and non-lytic egression, such as poliovirus and CVB3, HIV-1 and HCV. How these viruses trigger the accumulation of autophagosome-like vesicles and DMVs remains unclear. Some theories involve blocking the fusion of nascent autophagosomes with late endosomes and lysosomes, as in the case of HIV-1 Nef, which appears to cause autophagosome accumulation by inhibiting their progression towards more mature stages. Indeed, autophagosome-like vesicles may represent a trafficking pathway for these viruses, connecting to multivesicular bodies (MVBs), and assuring virus assembly and budding at the cell surface while protecting them from intrinsic antiviral factors and immune responses. The morphogenesis and release of mature and infectious HBV particles also require Tsg101 and depend on the ESCRT-MVB system. Under standard conditions the lumen of autophagosomes acidifies after fusion with endosomes that carry vacuolar (H + )-ATPase (V-ATPase) to form amphisomes. The autophagic membrane flux progresses by fusing with lysosomes in order to form the autolysosome that contains the former's proteinases. Poliovirus inhibition of autophagosome formation attenuates viral replication while inhibiting autolysosome formation, and thus, catalytic activity does not affect the virus. However, degradation of cellular triglycerides by autophagy benefits DENV replication and autolysosome degradation dampens IFN activation following HCV infection of these pathologies. Indeed, there is growing evidence that cell's modify their membranes to defend themselves against pathogens and infection, altering their spatial reorganization and vesicle trafficking. In this review, we focus on the importance of the membrane flux triggered by viruses to achieve replication and egression, and to ensure their propagation.

Several of the cell's organelles and membrane structures are involved in viral replication and in fact, many viruses use specific cellular compartments to replicate, referred to as viral factories (VFs: Figure 1 ). These VFs provide a physical scaffold that brings together elements required for genome replication and morphogenesis [1, 7] . VFs are usually formed by rearranging the host's cell membranes, reorganizing the cytoskeleton and recruiting specific organelles, like mitochondria (reviewed in [8] ). These viral driven events involve the association of replication complexes (RCs) with ER derived membranes to form a VF. Hence, intracellular membrane dynamics appear to be crucial for viral replication and survival.

A well-known example of a VF is that used by vaccinia virus, an enveloped pathogen of the Poxvirus family that replicates in the cytoplasm by assembling small rough ER-derived cisternae into a microenvironment that resembles a cytoplasmic mini-nucleus for viral replication [9] . Similarly, the RCs of Togaviruses associate with endocytic membranes, while Nodavirus RCs associate with mitochondrial membranes (reviewed in [1] ). Thus, specific membrane compartments can be used as VFs by RNA viruses to concentrate viral replicases and key cofactors, and ensure efficient viral genome replication [10] . In this context, both rubella virus (RUBV), a relevant human teratogenic Togaviridae virus [11] , and Semliki forest virus (SFV), a member , some viruses attach structural polyproteins to PIP 2 -rich membrane regions of the infected cell for further budding and release into the intercellular space. PIP 2 confers fluidity to the cell membrane and favours virus-cell fusion. These virions then bind to specific receptors in order to infect the neighbouring target cell at the VS, fusing with its plasma membrane directly or after surfing on actin-structured filopodia, or being internalized by endocytosis as is believed to occur with HIV-1. The VS represents an efficient environment for viral budding. It typically arises in PIP 2 -enriched plasma membrane domains, where the membrane of the infected cell is polarized towards the synaptic junction through the movement of vesicles governed by the ESCRT/Alix-Tsg101 machinery or by MVBs coordinating the translocation of the MTOC. This scaffolding facilitates subsequent viral infection and spread from the infected to the nearby uninfected cell. In addition, long membrane nanotubes may also form between neighbouring cells, promoting viral protein trafficking. Other dynamic membrane events involved in viral infection and spreading are trogocytosis, Arf6/PIP 2 -mediated membrane dynamics and exosomal transport. Trogocytosis involves the exchange of cell surface membrane patches that may contain receptor clusters associated to viral particles, while exosomes are vesicles formed from MVBs that could participate in viral infection and spreading between cells of the Alphavirus group of this family [12] , couple their RNA synthesis to endosome and lysosome membranes modified by the association of virus specific components. The subsequent fusion of these late endosomes and lysosomes generates cytopathic vacuoles (CPVs) [13, 14] that are lined with small vesicular invaginations or spherules (viral RNA replication sites) [13, 14] . CPVs establish complex and reversible contact with endocytotic vesicles through internal membranes interconnected with transport endosomes [8] . For example RUBV forms VFs around CPVs via the recruitment of membrane structures from the ER cisternae, Golgi stacks and mitochondria [8] (Figure 1 ). The Golgi apparatus is a highly dynamic organelle with a sustained, functional flux of membrane proteins [15] , and it can serve as a morphogenic mould for Rubiviruses, Coronaviruses, Arteriviruses and Bunyaviruses [1, 8, 16, 17] . These RUBV factories connect viral replication with the assembly and maturation of nascent virions at Golgi membranes, contributing to the virus escaping from the host cell's defences [16] .

Some viruses induce the formation of doublemembrane vesicles (DMVs) and/or autophagosomes for replication [1] [2] [3] [4] 18, 19] , such as the positive RNA viruses of the Flaviviridae family and Nidovirales order [20, 21] (Figure 1 ). The RNA polymerase of the human poliovirus, a Picornaviridae family member responsible for poliomyelitis, can also assemble DMVs [22] . Infection by this virus triggers the modification of different intracellular membrane structures and organelles (but not mitochondria), converting them into virus replication vesicles. In fact, poliovirusassociated DMVs resemble autophagosomes, as also described for another Picornaviridae family member, Coxsackievirus B3 (CVB3 [23, 24] : Figure 1 ). Autophagosomes are DMVs generated by membrane trafficking and they are related to the catabolic process of autophagy, which involves the degradation of cytoplasmic components within lysosomes [25, 26] . Autophagy maintains the organism's homeostasis by sequestering undesired intracellular elements for lysosomal degradation and recycling [25, 26] . Viruses often use autophagy to complete their lifecycle and evade immune responses, even though it is based on catalytic pathways [3, 23, 24] . Poliovirus, like other positive RNA viruses, has evolved the capacity to convert autophagy into a key cellular motor for replication [3, 10, 23] . During autophagy, a cytosolic form of the microtubule-associated protein 1A/1B light chain 3 (LC3-I) conjugates with phosphatidylethanolamine to form LC3-II and associate with autophagosomal membranes, ultimately producing the degradation of LC3-II during the late steps of autophagy [27] . Conversely, the p62 protein (or sequestosome-1, SQSTM1) interacts with ubiquitinated proteins, LC3 and other proteins to ensure the correct degradation of undesired material. LC3-II augments during active autophagy when p62 is degraded [28, 29] . In this context, Poliovirus or CVB3 infection triggers the generation of autophagosomes with a higher LC3-II/LC3-I ratio and with LC3 foci, structures that support the RNA RC without promoting lysosome degradation (evident through p62 stabilization [23, 24] : Figure 1 ). However, it is unclear whether these viruses block autophagosome maturation into amphisomes, avoiding autophagosome fusion with endosomes [30] . Such events override the appearance of degradative autolysosomes [31] or they may provoke the formation of autophagosome-like structures disconnected from catalytic pathways. It is also thought that these autophagosomes may ultimately serve as a membrane scaffold to permit the egression of nascent virions from infected cells, preventing cell lysis [30] (Figure 1 ).

All HCV viral genotypes (1a, 1b and 2a), positive RNA flaviviruses that are a major cause of chronic liver disease [32], induce autophagosome accumulation [33, 34] . This involves regulation of the unfolded protein response (UPR), which relieves ER stress and prevents the formation of catalytic autolysosomes by suppressing their fusion with lysosomes [33,34] ( Figure 1 ). Apparently, the success of viral replication relies on the recruitment of membrane-trafficking proteins to ER-derived membrane scaffolds [35] [36] [37] [38] [39] [40] [41] . Hence, domain 1 of the non-structural 5A (NS5A) protein and the helicase domain of NS3 are sufficient to achieve efficient DMV formation, which also depends on tightly regulated cis cleavage of the HCV-polyprotein precursor [35] and requires cyclophilin A isomerase activity [36] . NS5A associates with NS5B, a RNAdependent RNA polymerase, a complex that interacts with VAMP (vesicle-associated membrane protein)-associated proteins (VAPs) [37,38] and recruits Ras-like small GTPases (e.g. Rab1, Rab5 and Rab7), enlarging the viral replication compartment by docking membrane vesicles [39] [40] [41] . This process also regulates autophagy [42], given that HCV-induced autophagosomes support viral replication and the delivery of incoming viral RNA to the translation apparatus, and/or the recruitment of cellular factors for translation. However, some controversy still surrounds this issue, autophagosomes can mature into acidic amphisomes in HCV-infected cells [43, 44] , and subsequently fuse with late endosomes or lysosomes [44] (Figure 1 ). Autophagic membrane flux appears to be necessary to translate the HCV genome, yet it appears to be dispensable once viral infection has begun [45] . Moreover, no changes in either p62 or the degradation of long-lived proteins are observed [33], despite the enhanced autolysosome formation in cells expressing HCV replicons [46] . While specific silencing of autophagy genes does not affect viral translation and RNA replication, it does apparently alter HCV morphogenesis [47] . However, the silencing of factors critical for autophagosomes formation, like LC3 or Atg7, appears to suppress HCV RNA replication [48], while HCV replication is apparently potentiated when the UPR promotes autophagy [49] . Conversely, HCV infection seems to promote autophagy without concomitant stimulation of the UPR and autophagy does not appear to be required as a platform for HCV RNA replication [50]. Thus, doubts remain about the role of autophagy and the UPR in HCV replication, although the distinct interactions between autophagy and HCV replication suggest that such membrane flux promotes viral replication.

The dengue virus (DENV) is a mosquito-borne single positive-stranded RNA virus of the Flaviviridae family that causes dengue fever [51] . There are five antigenically related but distinct DENV virus genotypes (DENV-1 to DENV-5) [51,52]. Like HCV, there is evidence that autophagy may be implicated in DENV replication. Following cell entry and nucleocapsid uncoating, DENV RNA is translated into a single polyprotein that passes into the ER lumen where the different viral proteins are processed. In fact, DENV-2 proteins involved in translation and replication are found in or in close proximity to autophagosomes during viral infection [53, 54] . Accordingly, inhibition of autophagosome formation dampens the production of infectious DENV-2 particles [53], while stabilizing autophagosomes and/or amphisomes by impeding their fusion with lysosomes enhances viral egression [55] . Indeed, DENV-3 seems to promote autophagy during early infection [54] , while inhibition of autophagosome formation also dampens the production of infectious DENV-3

[54]. Hence, DENV-2 and -3 appear to interact with the autophagy machinery in a different manner, and while it is conceivable that amphisomes or autophagosomes represent the site of DENV-2 translation/replication [54,55], autophagolysosomes could be the crucial site for DENV-3 viral replication [54] . The distribution of NS1 or DENV-2 and DENV-3 double-stranded RNA (dsRNA) in the different autophagy-associated membrane structures confirms these observations ( Figure 1 ). Moreover, nascent viral particles are formed and mature in these structures, then travelling through the trans-Golgi network (TGN) to egress [56, 57] . Remarkably, the precursor membrane (prM) protein of the DENV-1-4 genotypes behaves similarly and it is cleaved by the TGN-protease furin in the secretory pathway [58], assuring viral assembly and the infectivity of nascent viral particles [59] .

HIV is a single-stranded RNA virus (Lentivirus genus of the Retroviridae family) that causes AIDS. HIV type 1 (HIV-1) alters the autophagic membrane flux of the host cell's organelles, thereby modulating the intracellular milieu in favour of viral replication and propagation [3] (Figure 1 ). When CD4+ T cells, monocytes and dendritic cells (DCs) are infected with HIV-1, autophagic vacuole formation is blocked and the expression of autophagy proteins down-regulated (e.g. LC3 and Beclin1 [19, 60] : Figure 1 ). Remarkably, the HIV-1 protein Nef (negative factor) blocks the autophagic flux of membranes, especially during the autolysosome stage of autophagy, resulting in an accumulation of autophagosomes and LC3 in macrophages ( Figure 1 ). Thus, Nef prevents autophagic degradation of HIV-1 biosynthetic intermediates of virions by targeting the lipid class III phosphatidylinositol 3-kinase (C3-PI3K) complex and by associating with Beclin1 (Atg6-autophagy-related protein 6 -in yeast). Significantly, Beclin1 is actually part of the C3-PI3K complex, together with the vacuolar protein sorting-associated proteins 34 (Vps34) and 15 (p150). Nef therefore alters the sub-cellular distribution of Vps34, potentially ensuring the survival of the viral progeny [3, 61] . Indeed, Nef is thought to promote the appropriate HIV-1 Gag membrane localization and processing, thereby facilitating viral cell-to-cell transfer [62] . Although the catalytic activity of autophagy appears to be impeded by HIV-1, autophagosome formation or accumulation is still promoted. Hence, the HIV-1 Gag protein promotes early stages of autophagosome formation by directly interacting with LC3 in macrophages, enhancing HIV-1 yields and Gag processing, a critical step in virion assembly and release [61] (Figure 1) . Notably, newly identified components of the ubiquitin-like conjugation system all seem to be involved in HIV-1 replication (e.g. Atg7, Atg8-LC3 is its best characterized mammalian homologue-Atg12 and Atg16L2-responsible for vesicle elongation) [63] . However, it remains unclear how these factors actually affect HIV-1 replication, which occurs in the nucleus of infected cells. Moreover, while autophagic vacuoles would appear to be fundamental for HIV-1 morphogenesis and egression, how HIV-1 overrides or uses autophagy to persist remains poorly understood. Hence, the infectious capacity of nascent HIV-1 virions depends on the uptake of the viral infectivity factor (Vif) during viral budding, a process influenced by histone deacetylase 6 (HDAC6), which promotes autophagic clearance of Vif [64] . Other positive RNA viruses exploit the formation of ER-derived membrane scaffolds and membrane autophagic flux to replicate (e.g. the Norwalk virus), because the membranebound nsp48 protein also binds to VAP-A [65] .

RNA replication may occur in endosomes, lysosomes (Togaviruses), peroxisomes and chloroplasts (Tombusvirus), or mitochondria (Nodaviruses), shielded from immune responses. All positive RNA viruses transform cytoplasmic membranes into specialized viral replication sites [10] . The antiviral effect of Brefeldin A (BFA), an inhibitor of anterograde ER-Golgi network membrane dynamics, suggests that membrane trafficking must be active for enterovirus replication, as reported for Picornaviruses, poliovirus and Coxsackievirus [66, 67] . BFA prevents the membrane flux required to form replication compartments, blocking virus secretion from infected cells [68] by inhibiting ADP (adenosine diphosphate)ribosylation factor (Arf)-GTP exchange proteins (Arf-GEFs). This blockade negatively affects COPI coat generation at the Golgi by diminishing and sequestering Arf1-GTP [69] . For several Picornaviruses, COPII-coated vesicles may provide membranes suitable for replication [70] , although autophagosomes may also contribute at this point [23] (Figure 1 ). Reovirus and SFV also promote coated-pit formation [71] . Moreover, the small GTPase Rab7 is soon recruited for SFV internalization when associated to intermediate endosomes [72] , which in turn induces the formation of CPVs that is an important event for viral RNA synthesis in target cells [13] .

An important biological process common to the recently proposed Megavirales order is viral replication within cytoplasmic VFs [73] . Giant viruses (also called nucleocytoplasmic large DNA viruses-CLDVs) belonging to this order are double-stranded DNA (dsDNA) viruses with a genome and particle size comparable to those of small bacteria [74] . African swine fever virus (ASFV; from the Asfarviridae family), poxviruses and iridoviruses are the three families of NCLDVs that terminate or undergo their entire replication cycle in the cytoplasm [75] [76] [77] . This feature is not observed in herpes viruses or baculoviruses, other large DNA viruses of eukaryotes that undergo nuclear DNA replication and transcription [78] . Giant viruses provoke VF formation in the cytoplasm of infected cells to permit genome replication and morphogenesis [73, 79] . ASFV factories are similar to the aggresomes formed at the MTOC (microtubule organizing centre) [80] , and they provoke a rearrangement of the intermediate vimentin cytoskeleton at the MTOC into a star shaped structure that resembles the microtubule aster formed during mitosis, a structure required for late gene expression [81] . Together with an ASFV chaperone, the hsp70 cell chaperone is recruited to ASFV factories, along with mitochondria, facilitating the folding of viral structural proteins like the major capsid protein p72 [82, 83] . Nascent ASFV virions are formed from VF membranes through the assembly and recruitment of viral proteins in VFs. Thus, the viral membranes in VFs may be connected to cellular organelles, particularly given that resident ER markers are detected with the viral p17, p54 and pB318L proteins in new viral particles [84] [85] [86] [87] . ASFVs are thought to reorganize cell membranes through viral proteins that contain a KDE motif, inducing the redistribution of ERassociated proteins [88] and the viral p54 protein. The latter is required for the correct VF localization of the membranes and the collapse of the ER-derived cisternae [89] . ASFV infection is achieved by redistributing membranes from the secretory pathway and TGN [90] . Therefore, these common biological features of giant virus replication and virion architecture could reflect a common origin, and the sharing of a large set of ancestral genes [74, 91] .

Membrane dynamics during viral assembly and budding and infectiousness. During budding, successful infection is achieved by adjustment and distortion of the target cell's plasma membrane [4] . The structural Gag polyprotein is common to several retroviruses, like HIV-1 and the murine leukaemia virus (MLV), representing the minimal plasma membrane component required for viral assembly [92] . HIV-1 Gag localizes to phosphatidylinositol-4,5-bisphosphate (PIP 2 ) rich plasma membrane regions, where PIP 2 plays a critical role in HIV-1 virion assembly [93] (Figures 1 and 2) . In fact, the matrix viral protein (MA) within the unprocessed HIV-1 Gag polypeptide drives Gag towards these PIP 2 membrane domains [92, 94] in a myristoylation-dependent manner [95] , raft domains where HIV-1 buds [95] [96] [97] . Phosphate hydrolysis by polyphosphoinositide 5phosphatase IV (5ptaseIV) diminishes the plasma membrane PIP 2 [98] , causing the Gag polypeptide to translocate from HIV-1 budding sites at the membrane to CD63 rich compartments, thereby inhibiting viral release [93] . Similarly, Arf6/Q67L expression, a GTP-bound mutant of Arf6, alters the trafficking of Arf6/PIP 2 -associated vesicles, provoking their accumulation in the cytoplasm to where Gag is redirected. These complexes lie far from the budding sites at the membrane, thereby dampening virus release [93] . Although the assembly of HIV-1 at the cell surface is only partially understood, several key steps in the membrane trafficking of viral proteins have been defined, shedding light on both the viral assembly and budding processes [92, 99] .

Enveloped viruses like HIV-1, Vesicular stomatitis virus (VSV), Ebola virus (EBOV) and Hepatitis B virus (HBV), and other RNA and DNA viruses, mainly emerge from cells by co-opting the host's ESCRT machinery [100, 101] , which plays a vital role in cellular abscission and in multivesicular body (MVB) biogenesis (a process by which ubiquitinated misfolded or damaged proteins enter endosomes to be destroyed). In addition, MVBs are important intermediates in endolysosomal transport [102] (Figures 1 and 2) . Gag activity drives ESCRT-III complex formation at the budding site of HIV-1, which binds to and recruits the ESCRT-I complex and the ALG-2 (apoptosis-linked gene 2)interacting protein X (ALIX). This ESCRT-III complex promotes the excision of nascent virions at the cell surface, an event potentially equivalent to the cleavage of intraluminal vesicles from MVBs [103, 104] (Figures 1 and 2) . Moreover, the tumour susceptibility gene 101 (Tsg101) is a subunit of the ESCRT-I complex that drives viral RNA transport and envelope fusion to late endosomes, processes required for infection and RNA release [105] ( Figure 2) . However, the interaction of viral Gag protein with the ESCRT machinery appears not to be absolutely required for HIV-1 viral budding [106] [107] [108] . Nevertheless, interferon-stimulated gene 15 protein (ISG-15) inhibits HIV-1 egression by interfering with ESCRT-III protein membrane flux during budding [109, 110] .

Remarkably, morphogenesis and the release of HBV particles also require Tsg101 [111] , although this DNA virus lacks a viral protein bearing the late (L) domain necessary to interact with the ESCRTmachinery [101] . However, α-taxilin interacts directly with Tsg101 and with the large HBV surface protein (LHBs), thereby recruiting the viral capsids to ESCRT complexes, thus permitting correct viral formation and egression [111] . Therefore, HBV maturation and egression depends on the ESCRT-MVB system. Notably, HBV infected cells also produce large amounts of non-infectious spherical or filamentous envelope particles (SVPs). These SVPs are a mixture of lipids and viral surface proteins that accumulate in an ER-Golgi intermediate compartment (ERGIC), budding into the lumen and provoking release through the general secretory pathway [112] .

Many other enveloped RNA viruses bud in an ESCRT-dependent manner [5, 100, 113] , as do most negative-strand non-segmented single-stranded RNA (ssRNA) viruses, such as Rhabdoviruses, Filoviruses and most Paramyxoviruses, all of which recruit ESCRTs for viral egression [114, 115] . Even the budding of negative-strand segmented-ssRNA Arena viruses involves an ESCRT-dependent pathway [116, 117] . However, no evidence for the participation of ESCRTs has yet been reported in Nipah, Measles, HRSV or Bornaviridae budding ( [5] ). Indeed, the enveloped influenza virus buds in an ESCRT-independent manner as its matrix protein is devoid of an ESCRT-binding domain [118, 119] . Other viruses are also released from the host's plasma membrane through their MAs, such as the Newcastle disease virus or VSV. In these cases, bud formation and excision from the membrane are matrix-dependent processes [120, 121] , as for influenza virus. However, much work is still required to determine how membrane dynamics affect the trafficking and assembly of these viruses, particularly in terms of the cellular factors that control the [92, 122] . Given all of these findings, membrane dynamics has a crucial influence on the assembly and budding of numerous viruses, and it may represent an important and complex target to limit the viral life cycle.

Viruses use various cell communication pathways to achieve effective cell-to-cell dissemination [123] . First described for type 1 HTLV (HTLV-1) [124] , the virological synapse (VS) is a complex structure found at the interface of infected:uninfected cells. Viral receptors and the egression machinery accumulate at the VS [125] , making the infection and spread of HTLV-1 through T lymphocytes cell-cell dependent. Direct cell-to-cell transmission facilitated by the formation of stable cellular junctions has several advantages, including faster replication rates [126] , successful transmigration of infected cells across mucosal barriers [123] and viral protection from host responses. However, such transmission is still to be confirmed for HIV [127, 128] .

Cell-to-cell spreading of HIV-1 ( Figure 2 ) is considered to involve microtubule-mediated polarization and substantial budding, followed by the entry of free viral particles into target cells [129] . Thus, it involves pathways that regulate cell-free virus entry by modifying membrane dynamics. In this regard, most HIV-1-infected GALT (gut-associated lymphoid tissue) cells in intestinal crypts are infected by concentrated pools of free HIV-1 viral particles in HIV-1-infected humanized mice. Fewer infected cells are found in the mucosal regions and the lamina propria, where VS presumably occur [130] , explaining why infection of permissive cells by free viral particles is crucial for HIV-1 replication and pathogenesis in vivo. This is consistent with the recent identification of the key cell signals required for efficient early HIV-1 infection and the establishment of latency in CD4+ T cells [131] [132] [133] [134] [135] [136] . Interference with retroviral cell-tocell transmission is not only produced by blocking cytoskeletal motility [137] and depleting membranecholesterol [138] but also, by interfering with Arf6governed plasma membrane dynamics. Moreover, restricting plasma membrane fluidity caused by altering early HIV-1-triggered phosphatidylinositol-4-phosphate 5-kinase (PI4P5-K) Iα activation and the ensuing detrimental effects PIP 2 production on HIV-1 transmission [4, 133, 135] . In fact, Arf6-coordinated membrane trafficking is required for efficient HIV-1 fusion, entry and infection of CD4+ T lymphocytes [135] (Figures 1 and 2) . The flux and turnover of PIP 2 -enriched vesicles from the plasma membrane, driven and coordinated by the Arf6-GTP/GDP cycle, ensures cell surface membrane regeneration and it allows membrane exchange between the viral and target cell surfaces. This type of membrane trafficking, coupled with enhanced fluidity, is in strong synergy with the key HIV-1/receptor (CD4 and C-X-C or C-C chemokine receptor type 4 or 5-CXCR4 or CCR5), interactions that promote fusion pore formation in target cells. These interactions take place between the non-regenerative HIV-1 viral membrane and the dynamic cell surface membrane, favouring efficient virus-cell fusion, entry and infection, both for the free virus and in the context of the VS [4, 133, 135] (Figures 1 and 2) .

Despite these similarities, some contact-specific events that affect membrane flux should also be considered. During cell-cell HIV transmission, intense viral endocytosis drives entry into neighbouring cells even if they are in contact [139] (Figure 2 ). Remarkably, biofilm-like structures at the surface of infected cells concentrate HTLV-1 viruses for their efficient transmission to target cells [140] and cellular projection is used to transmit pseudo-rabies virus. Retroviruses also travel along membrane protrusions that contact adjacent cells and MLV surfs on the filopodia of fibroblasts before entering cells [141] (Figure 2 ). HIV-1 also takes advantages of filopodia for cell-tocell transmission [141] , similarly surfing on the narrower membranous nanotubes that connect cells separated no more than 100 μm [142] and facilitating the transfer of viral proteins to the inner side of the membrane. These actin structures extend from HIVinfected cells to target cells irrespective of receptorenvelope interactions [142] (Figure 2) .

The take up of larger membrane invaginations at the VS of connected cells [129] is known as trogocytosis, an event that may also control the extent and stability of the synapse, regulating its duration [143] . HIV particles, like CD4 molecules and other membrane components, are transferred by trogocytosis from uninfected to infected cells in a manner triggered by the HIV-1 envelope (Env)/ CD4 [128] (Figure 2 ). This mechanism could be very significant and render cells permissive to HIV infection, as recently proposed [144] .

The cell-cell contacts and signalling induced by the HIV-1 Env complex that occur at the VS can activate autophagic membrane flux, leading to apoptotic cell death of uninfected CD4+ T cells [60, 145] . This lethal autophagy may provoke or enhance immunodeficiency, as observed in vivo where the majority of CD4+ T cells undergoing apoptosis, as well as the peripheral blood and lymph nodes of HIV patients, remain uninfected [146] . Simultaneously, autophagy can be suppressed in infected CD4+ T cells [60] , thereby antagonizing Env-mediated apoptosis and allowing viral replication to occur in infected CD4+ T cells. In this context, HIV-1 evades immune responses in an HIV-1 Env-CD4dependent manner by efficiently impairing autophagy in DCs when early contacts are established [19] . Taking into account the role autophagy plays in viral replication, HIV-1 can enhance or suppress autophagy at different stages of its viral cell cycle, favouring persistence and the evasion of immune responses, and therefore, its pathogenesis [19] ( Figure 1 ). Finally, like other viruses (e.g. CMV), HIV-1 stably associates with professional APCs during infection (such as DCs) to further infect lymphocytes during T-cell scanning or antigen presentation [147] . In fact, HIV-1 enters DCs by exploiting exosomal trafficking during antigen presentation [148] (Figure 2 ).

This review examines the intracellular trafficking of viruses that occurs in association with cellmembrane structures, some of which may be newly assembled by viruses to ensure their replication and budding. Membranes derived from the ER, mitochondria, lysosomes and endosomes are sculpted by viruses to generate VFs, acquiring their own functional morphology. These structures help ensure RNA replication is accomplished without alerting the host's defence mechanisms.

Although the importance of membrane dynamics during viral infection has been established, several questions remain unanswered. It remains unclear how proteins from distinct viruses and host cells use the same intracellular membrane compartments or events (e.g. autophagy) to achieve viral replication, without affecting important cellular processes. Conversely, it is not clear why viruses replicate in different subcellular membrane compartments, how they move across membranes and which host factors are involved in these events. Similarly, we still do not know how these changes in membrane dynamics enable viruses to avoid immune responses. Indeed, it remains unclear whether rearranging intracellular organelles enables viruses to escape the anti-replicative activity of natural restriction factors, such as apolipoprotein B mRNA-editing enzyme-catalytic, polypeptide-like 3 (APOBEC3) proteins, Tetherin (BST-2/CD317/ HM1.24) or SAMHD1 (sterile alpha motif (SAM) and histidine-aspartate (HD) domain-containing protein 1) for HIV-1 [149] . Resolving these issues will help decipher how viruses rearrange membranes during their infection cycle, thereby aiding the design of new antiviral strategies that target these dynamic viral-cell interactions and combat viral infection. These findings may also produce innovations in non-viral gene delivery systems to tackle tumours and immune diseases.

New technical developments, such as more powerful microscopy systems [4, 8, 135, 150] , will allow dynamic viral trafficking and egression to be studied in cells with better spatial and temporal resolution. Such information will further our understanding of the viral infection process and of how viruses succeed in deceiving the host's immune responses. respectively, Spain), UNLL10-3E-783 (ERDF) and Fundación CajaCanarias, and by the Project RD12/0017/0034 integrated in the "Plan Nacional I + D + i" and co-funded by ISCIII-"Subdirección General de Evaluación" and ERDF (RIS-RETIC) grants. M-S.V and L.A-R are supported by RD12/0017/ 0034-(RIS-RETIC) and SAF2011-24671-FPI-associated grants and fellowships, respectively. We thank Dr Mark Sefton (Biomedred SL) for his linguistic revision of the manuscript. We apologize for all research studies and reviews that we have not discussed or cited in this Review. We have tried to avoid any and all such omissions but space limitations have surely made this an impossible endeavour. 

Modification of intracellular membrane structures for virus replication

Viruses and endosome membrane dynamics

When autophagy meets viruses: a double-edged sword with functions in defense and offense

Viral infection: moving through complex and dynamic cell-membrane structures

How to get out: ssRNA enveloped viruses and membrane fission

Neutrophil granules and secretory vesicles in inflammation

A guide to viral inclusions, membrane rearrangements, factories, and viroplasm produced during virus replication. Advances in

Ultrastructure of the replication sites of positive-strand RNA viruses

Vaccinia virus DNA replication occurs in endoplasmic reticulum-enclosed cytoplasmic mini-nuclei

Viral RNA replication in association with cellular membranes

Molecular biology of rubella virus

Biogenesis of the Semliki Forest virus RNA replication complex

Alphavirus RNA replicase is located on the cytoplasmic surface of endosomes and lysosomes

Rubella virus replication complexes are virus-modified lysosomes

Microscopic morphology and the origins of the membrane maturation model of Golgi apparatus function

Structural maturation of rubella virus in the Golgi complex

Polymorphism and structural maturation of bunyamwera virus in Golgi and post-Golgi compartments

Herpesviruses remodel host membranes for virus egress

Human immunodeficiency virus-1 inhibition of immunoamphisomes in dendritic cells impairs early innate and adaptive immune responses

ORF1a-encoded replicase subunits are involved in the membrane association of the arterivirus replication complex

Kunjin virus: an Australian variant of West Nile?

Oligomeric structures of poliovirus polymerase are important for function

Subversion of cellular autophagosomal machinery by RNA viruses

Autophagosome supports coxsackievirus B3 replication in host cells

Maturation of autophagic vacuoles in mammalian cells

Autophagosome formation: core machinery and adaptations

Autophagy: process and function

p62/ SQSTM1 forms protein aggregates degraded by autophagy and has a protective effect on huntingtin-induced cell death

p62/SQSTM1 binds directly to Atg8/LC3 to facilitate degradation of ubiquitinated protein aggregates by autophagy. The Journal of

That which does not degrade you makes you stronger: infectivity of poliovirus depends on vesicle acidification

Differential role of autophagy in CD4 T cells and macrophages during X4 and R5 HIV-1 infection

Autophagy pathway intersects with HIV-1 biosynthesis and regulates viral yields in macrophages

HIV-1 Nef promotes the localization of Gag to the cell membrane and facilitates viral cell-to-cell transfer

Identification of host proteins required for HIV infection through a functional genomic screen

The HDAC6/APOBEC3G complex regulates HIV-1 infectiveness by inducing Vif autophagic degradation

Norwalk virus nonstructural protein p48 forms a complex with the SNARE regulator VAP-A and prevents cell surface expression of vesicular stomatitis virus G protein

Inhibition of poliovirus RNA synthesis by brefeldin A

GBF1, a guanine nucleotide exchange factor for Arf, is crucial for coxsackievirus B3 RNA replication

Inhibition of cellular protein secretion by poliovirus proteins 2B and 3A

Involvement of cellular membrane traffic proteins in poliovirus replication

Cellular COPII proteins are involved in production of the vesicles that form the poliovirus replication complex

Endocytosis by random initiation and stabilization of clathrin-coated pits

Rab7 associates with early endosomes to mediate sorting and transport of Semliki forest virus to late endosomes

Vaccinia-like cytoplasmic replication of the giant Mimivirus

The origins of giant viruses, virophages and their relatives in host genomes

The biochemistry of icosahedral cytoplasmic deoxyviruses

Role of the host cell nucleus in the replication of African swine fever virus DNA

Frog virus 3 DNA replication occurs in two stages

Viral transcription during Autographa californica nuclear polyhedrosis virus infection: a novel RNA polymerase induced in infected Spodoptera frugiperda cells

Ultrastructural study of African swine fever virus replication in cultures of swine bone marrow cells

Aggresomes resemble sites specialized for virus assembly

Vimentin rearrangement during African swine fever vi-II

A virally encoded chaperone specialized for folding of the major capsid protein of African swine fever virus

Migration of mitochondria to viral assembly sites in African swine fever virus-infected cells

African swine fever virus is wrapped by the endoplasmic reticulum

African swine fever virus is enveloped by a two-membraned collapsed cisterna derived from the endoplasmic reticulum

Membrane rupture generates single open membrane sheets during vaccinia virus assembly

Vaccinia virus lacking A17 induces complex membrane structures composed of open membrane sheets

The subcellular distribution of multigene family 110 proteins of African swine fever virus is determined by differences in Cterminal KDEL endoplasmic reticulum retention motifs

Mechanism of collapse of endoplasmic reticulum cisternae during African swine fever virus infection

African swine fever virus organelle rearrangements. Virus

Megavirales", a proposed new order for eukaryotic nucleocytoplasmic large DNA viruses

Virus assembly and plasma membrane domains: which came first? Virus

Phosphatidylinositol (4,5) bisphosphate regulates HIV-1 Gag targeting to the plasma membrane. Proceedings of the National Academy of Sciences of the United States of

Gag localization and virus-like particle release mediated by the matrix domain of human T-lymphotropic virus type 1 Gag are less dependent on phosphatidylinositol-(4,5)-bisphosphate than those mediated by the matrix domain of HIV-1 Gag

Point mutations in the HIV-1 matrix protein turn off the myristyl switch

Evidence for budding of human immunodeficiency virus type 1 selectively from glycolipidenriched membrane lipid rafts

The HIV lipidome: a raft with an unusual composition

The isolation and characterization of a cDNA encoding phospholipid-specific inositol polyphosphate 5-phosphatase

M910119199 M910119199

HIV-1 assembly, budding, and maturation. Cold Spring Harbor Perspectives in

Late budding domains and host proteins in enveloped virus release

Mechanisms for enveloped virus budding: can some viruses do without an ESCRT?

Biogenesis and function of multivesicular bodies

The ESCRT machinery: new functions in viral and cellular biology

Membrane budding and scission by the ESCRT machinery: it's all in the neck

The ESCRT-I subunit TSG101 controls endosome-to-cytosol release of viral RNA

Analysis of the assembly function of the human immunodeficiency virus type 1 gag protein nucleocapsid domain

Functional role of Alix in HIV-1 replication

Human immunodeficiency virus type 1 nucleocapsid p1 confers ESCRT pathway dependence

The interferon-induced gene ISG15 blocks retrovirus release from cells late in the budding process

Budding of enveloped viruses: interferon-induced ISG15-antivirus mechanisms targeting the release process

Identification of alpha-taxilin as an essential factor for the life cycle of hepatitis B virus

Morphogenesis of hepatitis B virus and its subviral envelope particles

Viral late domains

Filovirus assembly and budding

Rabies virus assembly and budding

Efficient budding of the tacaribe virus matrix protein z requires the nucleoprotein

Arenavirus reverse genetics: new approaches for the investigation of arenavirus biology and 158 L. de Armas

development of antiviral strategies

Influenza virus hemagglutinin and neuraminidase, but not the matrix protein, are required for assembly and budding of plasmid-derived virus-like particles

Budding of filamentous and nonfilamentous influenza A virus occurs via a VPS4 and VPS28-independent pathway

Functional analysis of late-budding domain activity associated with the PSAP motif within the vesicular stomatitis virus M protein

Vesicle formation by self-assembly of membranebound matrix proteins into a fluidlike budding domain

Paramyxovirus glycoprotein incorporation, assembly and budding: a three way dance for infectious particle production

Avoiding the void: cell-to-cell spread of human viruses

Spread of HTLV-I between lymphocytes by virus-induced polarization of the cytoskeleton

Retroviral spread by induction of virological synapses

Quantitation of human immunodeficiency virus type 1 infection kinetics

Cell-to-cell HIV-1 spread and its implications for immune evasion

Antigp41 antibodies fail to block early events of virological synapses but inhibit HIV spread between T cells

Virological synapsemediated spread of human immunodeficiency virus type 1 between T cells is sensitive to entry inhibition

STINGdependent type I IFN production inhibits cell-mediated immunity to Listeria monocytogenes

Filamin-A regulates actindependent clustering of HIV receptors. Nature

PI4P5-kinase Ialpha is required for efficient HIV-1 entry and infection of T cells

Moesin is required for HIV-1-induced CD4-CXCR4 interaction, F-actin redistribution, membrane fusion and viral infection in lymphocytes

Arf6-mediated membrane dynamics to efficiently enter and infect T lymphocytes. Molecular

Gelsolin activity controls efficient early HIV-1 infection

Requirement for an intact T-cell actin and tubulin cytoskeleton for efficient assembly and spread of human immunodeficiency virus type 1

Human immunodeficiency virus type 1 virological synapse formation in T cells requires lipid raft integrity

On the steps of cellto-cell HIV transmission between CD4 T cells

Biofilm-like extracellular viral assemblies mediate HTLV-1 cell-to-cell transmission at virological synapses

Retroviruses can establish filopodial bridges for efficient cell-to-cell transmission

Membrane nanotubes physically connect T cells over long distances presenting a novel route for HIV-1 transmission

What is trogocytosis and what is its purpose?

Could CD4 capture by CD8+ T cells play a role in HIV spreading?

Autophagy is involved in T cell death after binding of HIV-1 envelope proteins to CXCR4

High expression of APO-1 (CD95) on T lymphocytes from human immunodeficiency virus-1-infected children

HIV and mature dendritic cells: Trojan exosomes riding the Trojan horse?

Exosomes and retroviruses: the chicken or the egg?

Host factors and HIV-1 replication: clinical evidence and potential therapeutic approaches

Virus trafficking-learning from single-virus tracking

This work and A.V.-F. are supported by the European Regional Development Fund (ERDF), SAF2008-01729 and SAF2011-24671 (MICINN and MINECO,

The authors have no competing interests to declare.