The mystery remains: How do potyviruses move within and between cells?

Abstract

The genus Potyvirus is considered as the largest among plant single‐stranded (positive‐sense) RNA viruses, causing considerable economic damage to vegetable and fruit crops worldwide. Through the coordinated action of four viral proteins and a few identified host factors, potyviruses exploit the endomembrane system of infected cells for their replication and for their intra‐ and intercellular movement to and through plasmodesmata (PDs). Although a significant amount of data concerning potyvirus movement has been published, no synthetic review compiling and integrating all information relevant to our current understanding of potyvirus transport is available. In this review, we highlight the complexity of potyvirus movement pathways and present three potential nonexclusive mechanisms based on (1) the use of the host endomembrane system to produce membranous replication vesicles that are targeted to PDs and move from cell to cell, (2) the movement of extracellular viral vesicles in the apoplasm, and (3) the transport of virion particles or ribonucleoprotein complexes through PDs. We also present and discuss experimental data supporting these different models as well as the aspects that still remain mostly speculative.

This synthetic review aims to shed light on the mysteries surrounding the complexity of potyvirus movement pathways.

1. INTRODUCTION

To invade the whole plant after their replication, plant viruses must move intracellularly to reach plasmodesmata (PDs), the symplasmic tunnels between cells that are the gateways for this movement, cross them to enter neighbouring cells, and then enter sieve elements. Viruses are then transported with the source‐to‐sink flow of photoassimilates and are unloaded distantly from sieve elements into sink tissues (Schoelz et al., 2011). PDs are structures unique to plants, crucial for the communication between plant cells, that allow the regulated passage of signals, from small molecules to macromolecules involved in plant growth, development, and defence (Reagan & Burch‐Smith, 2020). PDs can be defined by three major structural components: the membrane‐bound tube that leads to plasma membrane and cytoplasmic continuity between adjacent cells, the central desmotubule, which is an appressed form of the endoplasmic reticulum (ER), and the cell wall surrounding the plasma membrane (Reagan & Burch‐Smith, 2020). Cell wall glucan deposition, insoluble glucans or callose, densifies at the neck regions of PDs, modulating PD permeability and affecting their size exclusion limit (SEL) (Benitez‐Alfonso et al., 2010; German et al., 2023). Despite the symplasm continuity between cells, the SEL of PDs constitutes a physical barrier that must be overcome by a virus to successfully move from cell to cell. Indeed, viral nucleic acids or ribonucleoprotein complexes are too large to move through PDs on their own, and virions are too large to passively cross PDs. According to Schoelz et al. (2011), “if PD are the doorway out of the cell, then plant viruses must possess the tools to find the door as well as the keys to unlock the door”. Indeed, plant viruses use active mechanisms to move from the site of replication within the cell to the PD for cell‐to‐cell movement. Plant virus genomes encode so‐called movement proteins (MPs), which interact with host proteins to modify PDs and permit cell‐to‐cell transportation of the viral genome (Heinlein, 2015; Kumar & Dasgupta, 2021). Viral MPs have been identified in nearly all plant viruses (Tilsner et al., 2014) and display a range of common functions such as the ability to (1) nonspecifically bind nucleic acids (Citovsky et al., 1992), (2) target themselves to PDs and mediate their own cell‐to‐cell movement (Oparka et al., 1997; Wei, Zhang, et al., 2010), and (3) increase the SEL of PDs, also known as gating (Howard et al., 2004; Oparka et al., 1997). Although a variety of models for how different plant viruses transport their genomes within and between cells have been proposed (Kumar & Dasgupta, 2021), two main categories have been described. In brief, the first one, exemplified by tobacco mosaic virus (TMV) (a tobamovirus), may be common to viruses that do not move as intact virions and are considered as “nontubule‐forming viruses” (because they do not morphologically drastically alter PDs). In the second category, the MPs form large tubules that insert into PDs to allow the transport of intact virions between cells. Those viruses are considered as “tubule‐forming viruses” and induce drastic PD alterations, with elimination of the desmotubule (Kumar & Dasgupta, 2021; Morozov & Solovyev, 2020).

However, the cell‐to‐cell movement mechanism of the largest family of plant RNA viruses, the Potyviridae, does not fall into either of the two previous categories. The Potyvirus genus comprises plant viruses impacting crops of economic interest, including important pathogens such as plum pox virus (PPV), potato virus Y (PVY), soybean mosaic virus (SMV), and turnip mosaic virus (TuMV) (Inoue‐Nagata et al., 2022). Potyviruses are nonenveloped, flexuous rod‐shaped particles of 680–900 nm in length and 11–15 nm in diameter. The virus has a single‐stranded, positive‐sense RNA genome of around 10 kb, which is linked to a viral protein (VPg, viral protein genome‐linked) at its 5′ end and polyadenylated at its 3′ end. When the virus enters host plant cells, it is uncoated and the viral RNA is translated into a large viral polyprotein, which is sequentially cleaved into 10 proteins (Figure 1). This cleavage is performed by three viral proteins, P1, the helper‐component proteinase (HC‐Pro), and NIa‐Pro. The 11th protein is called P3N‐PIPO, for “Pretty Interesting Potyviridae ORF”, and is embedded in the P3 cistron. During viral replication, RNA polymerase slippage leads to insertion of an additional adenine in some genomic RNA progeny molecules, bringing the PIPO‐encoding sequence in frame with the P3 protein gene. As a result, a fusion protein named P3N‐PIPO, which contains the N‐terminus of P3 fused to the PIPO sequence, is produced (Chung et al., 2008; Olspert et al., 2015; Rodamilans et al., 2015) (Figure 1). As for many other viral proteins, potyviral proteins have multiple functions and are involved in multiple processes such as replication, intracellular movement, cell‐to‐cell movement, and long‐distance systemic movement (Table 1).

FIGURE 1.

Genomic organization of potyviruses. The long open reading frame (ORF) encodes the polyprotein that is cleaved by three viral proteases (P1, HC‐Pro, and NIa‐Pro) into 10 mature viral proteins. The PIPO ORF embedded in the P3 cistron is indicated as a striped area. The viral genome‐linked protein (VPg) is represented as a black ellipse at the 5′ end of the genome. Arrows starting from the three proteases indicate their cleavage sites in the polyprotein. While P1 and HC‐Pro cleave themselves autoproteolytically, NIa‐Pro, in addition to itself, releases the remaining viral proteins from the polyprotein (adapted from Revers & García, 2015).

TABLE 1.

Relevant features of multifunctional potyviral proteins.

Protein	Molecular mass (kDa)	Protein functions	References	Reviews
P1	30–80 a	Serine protease; RNA binding; accessory factor for virus replication; virus adaptation to host	Brantley and Hunt (1993), Valli et al. (2007), Verchot and Carrington (1995), Verchot et al. (1991)	Revers and García (2015), Rohožková and Navrátil (2011)
HC‐Pro	c.56	Cysteine protease; aphid transmission; RNA silencing suppression	Anandalakshmi et al. (1998), Carrington et al. (1989, 1996), Govier and Kassanis (1974), Kasschau and Carrington (1998), Valli et al. (2014)	Gadhave et al. (2020), Valli et al. (2018)
P3	c.37	Virus adaptation to host; virus replication and movement	Cui et al. (2010), Klein et al. (1994), Salvador et al. (2008)	Revers and García (2015)
P3N‐PIPO	c.14	Virus cell‐to‐cell movement	Chai et al. (2020), Cui et al. (2017), Vijayapalani et al. (2012), Wei, Zhang, et al. (2010)	Wang (2021)
6K1	6	Virus replication; reduction of plant protease activity to increase virus accumulation	Bera et al. (2022), Cui and Wang (2016), Geng et al. (2017), Hu et al. (2023)
CI	c.79	Pinwheel formation; RNA helicase; virus replication; virus cell‐to‐cell and long‐distance movement	Calder and Ingerfeld (1990), Carrington et al. (1998), Deng et al. (2015), Edwardson (1966), Fernández et al. (1995), Lain et al. (1990), Otulak and Garbaczewska (2012), Wei, Zhang, et al. (2010)	Sorel et al. (2014)
6K2	6	Formation of viral replication complex vesicles; virus replication; virus intra‐/intercellular and long‐distance movement	Cotton et al. (2009), Grangeon et al. (2012, 2013), Schaad et al. (1997), Wan, Basu, et al. (2015), Wan, Cabanillas, et al., (2015)	Grangeon et al. (2010), He et al. (2023), Solovyev et al. (2022)
VPg	c.21	RNA replication and translation; virus cell‐to‐cell and long‐distance movement; RNA silencing suppression; virus phloem loading	Cheng and Wang (2017), Keller et al. (1998), Nicolas et al. (1997), Rajamäki and Valkonen (2002, 2009), Schaad et al. (1997), Siaw et al. (1985), Tavert‐Roudet et al. (2017)	Eruera et al. (2021), Jaramillo‐Mesa and Rakotondrafara (2023), Jiang and Laliberté (2011), Rantalainen et al. (2011)
NIa‐Pro	c.27	Cysteine protease; DNase	Anindya and Savithri (2004), Flint and Ryan (1997); Schaad et al. (1996)	Adams et al. (2005)
NIb	c.58	RNA replicase (RNA‐dependent RNA polymerase [RdRP]); assembly and activation of viral replication complexes (VRCs); suppression of host defence	Cheng et al. (2017), Hong and Hunt (1996), Li et al. (2018), Sanfaçon (2005)	Mäkinen and Hafrén (2014), Shen et al. (2020)
CP	30–40 a	Virion assembly; RNA binding; virus translation; virus cell‐to‐cell and long‐distance movement; aphid transmission	Atreya et al. (1991), Blanc et al. (1997), Dolja et al. (1995), Hafrén et al. (2010), Kendall et al. (2008), Seo et al. (2013), Shukla and Ward (1989)	Gadhave et al. (2020), Martínez‐Turiño and García (2020), Revers and García (2015)

Note: The column “References” corresponds to pioneering studies and the column “Reviews” to recent reviews focused on a particular potyviral protein.

^a Sweet potato feathery mottle virus (SPFMV) P1 protein is the largest among all potyviral P1 proteins, ranging from 74.1 to 80 kDa (Mingot et al., 2016); sweet potato virus G (SPVG) CP is 39.4 kDa, the largest of all known species of potyviruses (Li et al., 2012).

Although several recent models have been proposed (Martínez‐Turiño & García, 2020; Solovyev et al., 2022; Wang, 2021), the cell‐to‐cell movement of potyviruses still remains poorly understood. Two nonexclusive cell‐to‐cell transport pathways of potyviruses were recently reviewed: one mediated by the “second 6‐kDa membrane‐anchoring protein” (6K2), in which potyviruses use the host endomembrane system to produce membranous vesicles able to move between cells, in a fashion reminiscent of the way some animal viruses use membrane‐derived vesicles for exit from infected cells and entry into healthy ones (Solovyev et al., 2022), and another mechanism where virions or ribonucleoprotein complexes are transported through PDs (Wang, 2021). A third, very original mechanism was proposed based on the observation in the extracellular space and cell wall (apoplasm) of TuMV‐induced vesicles, illustrating a potential change of paradigm for virus cell‐to‐cell transport (Movahed, Cabanillas, et al., 2019; Wan & Laliberté, 2015). The aim of the present review is to compile and integrate all available information relevant to the current understanding of potyvirus transport, in one complete and synthetic review. We present and discuss experimental data supporting these different models as well as the aspects that still remain mostly speculative.

2. FOUR VIRAL MULTIFUNCTIONAL PROTEINS ARE INVOLVED IN POTYVIRUS MOVEMENT

For potyviruses, there is no dedicated MP per se. Reverse genetics and cell biology studies performed on a large number of potyvirus species have shown that at least four potyviral proteins are involved in potyvirus transport (Revers & García, 2015; Wang, 2021). These are (1) the cylindrical inclusion helicase (CI, a protein that forms the pinwheel‐shaped inclusion bodies in the cytoplasm of infected cells, a unique feature of the Potyviridae), (2) P3N‐PIPO, (3) the 6K2 protein, and (4) the capsid protein (CP). A potential role for HC‐Pro in movement has also been suggested as it was shown to increase the SEL of PDs (Rojas et al., 1997; Valli et al., 2018), but this could as well be associated with an ability to move autonomously from cell to cell and over long distances, needed to fulfil its silencing suppression function, rather than being involved in virus movement (Kasschau & Carrington, 2001; Kumar & Dasgupta, 2021).

In addition to replication, the CI protein is involved in cell‐to‐cell and long‐distance movement. Previous alanine‐scanning mutagenesis experiments reviewed by Sorel et al. (2014) and Deng et al. (2015) revealed that mutations in the N‐terminal part of the CI protein impair cell‐to‐cell spread of tobacco etch virus (TEV) and PPV (Carrington et al., 1998; Gomez de Cedron et al., 2006). The ability of the TuMV CI protein to target PDs and interact with CP seems to be associated with its role in viral cell‐to‐cell movement (Deng et al., 2015). It has since been reported that P3N‐PIPO recruits the CI protein to the cell wall near PDs and depends on the secretory pathway but not on actin or myosin motors for that function (Wei, Zhang, et al., 2010).

Before the discovery of P3N‐PIPO by Chung et al. (2008), it had already been noted that a silent mutation in the P3 cistron of wheat streak mosaic virus altered viral movement. This mutation is now known to prevent the polymerase slippage event and thus P3N‐PIPO production (Choi et al., 2005). A few studies have shown that P3N‐PIPO can be considered as an MP for potyviruses: green fluorescent protein (GFP)‐tagged P3N‐PIPO targets PDs and is able to spread between cells, thus being seemingly capable of increasing the SEL of PDs and promoting viral movement (Cui et al., 2017; Vijayapalani et al., 2012), even if P3N‐PIPO has not been shown to bind to nucleic acids. The N‐terminal domain of P3N‐PIPO conditions its localization to PDs in the case of sugarcane mosaic virus (Cheng et al., 2017) and stop codons introduced in the PIPO sequence decrease the intercellular spread of SMV while they do not affect virus accumulation (Wen & Hajimorad, 2010).

The 6K2 protein of potyviruses is a transmembrane protein involved in rearrangement of the ER, leading to the generation of membranous viral vesicles, important for replication (Beauchemin et al., 2007) as well as for intracellular and intercellular movement (Cotton et al., 2009; Grangeon et al., 2013). The 6K2 protein is predicted to contain a central hydrophobic transmembrane domain necessary for vesicle formation (Schaad et al., 1997), an N‐terminal tail exposed in the cytoplasm involved in vesicle export from the ER, and a C‐terminal tail exposed in the ER lumen or in vesicles (Jiang et al., 2015).

Besides the three nonstructural proteins listed above, the fourth protein important for potyviral transport is the CP (Martínez‐Turiño & García, 2020). The potyviral CP presents three structural domains, with both N‐ and C‐terminal regions exposed on the virion surface and a conserved core subunit structure (Dolja et al., 1994). A large number of reverse genetics experiments have shown that mutations in all domains of the CP may impact viral movement (Martínez‐Turiño & García, 2020).

3. 6K2‐INDUCED VESICLES: KEY PLAYERS IN COUPLING POTYVIRUS REPLICATION AND INTERCELLULAR MOVEMENT

3.1. The 6K2 protein induces the formation of ER‐derived vesicles

The first description of 6K2‐induced vesicles dates back 25 years ago, when Schaad et al. (1997) showed that the TEV 6K2 protein induces, alone or during TEV infection, some discrete 2–10 μm diameter vesicles derived from the ER. Since then, the dynamics and ultrastructure of TuMV 6K2‐induced vesicles have been extensively studied by confocal and transmission electron microscopy (TEM) (Grangeon et al., 2012; Li et al., 2020; Wan, Basu, et al., 2015). At an early stage of TuMV infection, the accumulation of convoluted membranes (CMs) connected to the rough ER is observed. Then, at the TuMV infection midstage, CMs turn into single‐membrane vesicle‐like structures (SMVLs), which are viral RNA replication sites according to immunogold labelling studies using anti‐RNA‐dependent RNA polymerase (RdRp) and anti‐double‐stranded RNA (dsRNA) polyclonal antibodies (Wan, Basu, et al., 2015). Late in infection, in addition to the SMVLs, double‐membrane vesicle‐like structures (DMVLs) are produced as well as virion bundles associated with vacuoles where encapsidation could occur (Li et al., 2020; Wan, Basu, et al., 2015). Wan, Basu, et al. (2015) suggested that DMVLs are probably underlying the perinuclear globular‐like structures previously observed by confocal microscopy and produced by amalgamation of ER, Golgi bodies, coat protein complex II (COPII) coatomers, and chloroplasts (Grangeon et al., 2012; Wei, Huang, et al., 2010). This perinuclear globular structure contains viral RNA and numerous 6K2‐induced vesicles, as well as viral proteins involved in replication (P3, 6K1, VPg, NIa‐Pro, NIb [RdRp], and CI helicase) (Cronin et al., 1995; Cui et al., 2010; Cui & Wang, 2016; Klein et al., 1994; Lõhmus et al., 2016; Wei, Huang, et al., 2010), together with host translation factors (eIF[iso]4E, poly[A]‐binding protein [PABP], and eukaryotic translation elongation factor 1A [eEF1A]) (Beauchemin & Laliberté, 2007; Dufresne et al., 2008; Lõhmus et al., 2016; Thivierge et al., 2008). Even though the ER and Golgi apparatus have lost their characteristic organization in this perinuclear globular structure, they remain connected to the host secretory pathway, which is probably important for the generation of peripheral 6K2‐induced vesicles, shown to exit from the globular structure (Grangeon et al., 2012). Potyvirus replication and probably viral translation occur at the level of those 6K2‐induced vesicles, each being derived from a single viral genome (Cotton et al., 2009). This is confirmed by experiments with TuMV‐infected protoplasts, in which VPg‐Pro, RdRp, and CI colocalize in vesicular punctate structures (probably 6K2‐induced vesicles) with immunofluorescence‐stained dsRNA replicative forms or neosynthesized 5‐bromouridine‐labelled RNA (Cotton et al., 2009).

In summary, the 6K2‐dependent reorganization of the ER in potyvirus‐infected cells leads to the formation of two types of structures that are both involved in replication: the large perinuclear globular structure and the mobile 6K2‐induced vesicles located at the cell periphery. This led to some confusion in the terminology used in the literature, where different terms are employed to refer to the perinuclear structure and 6K2‐induced vesicles, such as “viral replication complexes (VRCs)”, “vesicular structures”, “punctate structures”, “replication complex vesicles”, “membrane‐derived replication complexes”, “viral replication vesicles”, “viral replication organelles (VROs)”, “vesicular VRCs”, and “VRC‐associated membranous structures”. Therefore, in this review, for clarity, the term “viral replication compartment” (VRC) will be used to refer to the perinuclear structure (in reference to similar perinuclear replication compartments observed for different plant virus genera), whereas we will refer to the 6K2‐induced mobile vesicles as “6K2 vesicles”.

The formation of potyvirus VRCs and 6K2 vesicles induces drastic endomembrane rearrangements and relies on the early secretory pathway. Indeed, the detachment of 6K2 vesicles from the ER membrane involves the COPII‐mediated formation of transport vesicles, which bud from the ER membrane and deliver cargo to the Golgi cisternae (Brandizzi & Barlowe, 2013; He et al., 2023; Wei & Wang, 2008). In particular, the TuMV 6K2 protein interacts with the COPII coatomer Sec24a, the subunit interacting with cargo proteins for their packaging into COPII vesicles (Jiang et al., 2015). The N‐terminal cytoplasmic domain of the 6K2 protein is crucial for this interaction, and mutation of the conserved tryptophan residue at position 15 leads to partial retention of the 6K2 protein in the ER, alters TuMV replication, and inhibits TuMV cell‐to‐cell movement (Jiang et al., 2015). Furthermore, the atlastin‐like GTPase ROOT HAIR DEFECTIVE3 (RHD3), which plays an important role in the generation of the interconnected tubular ER network and in membrane shaping, interacts with 6K2 and is required for the maturation of TuMV 6K2 vesicles in replication‐competent vesicles (Movahed, Sun, et al., 2019). Another striking point highlighted by studies on TuMV is that the 6K2 vesicles are mobile, and their different fates, after they detach from the ER, are detailed below.

3.2. 6K2 vesicles are mobile and target multiple cellular compartments during viral replication and intercellular movement

Mobile 6K2 vesicles located at the cell periphery are observed, deriving from the VRC (Grangeon et al., 2012). The intracellular mobility of potyvirus 6K2 vesicles relies both on components of the early secretory pathway and on the actin network (but not on microtubules), while the vesicle trafficking along actin microfilaments depends on myosins XI‐2 and XI‐K (Agbeci et al., 2013; Cotton et al., 2009; Grangeon et al., 2012; Wei & Wang, 2008). Latrunculin B, a drug that disrupts actin microfilaments, impedes not only the intracellular movement of 6K2 vesicles (Cotton et al., 2009), but also TuMV cell‐to‐cell transport (Agbeci et al., 2013), suggesting that 6K2 vesicles could be involved in both intra‐ and intercellular movement of TuMV.

3.2.1. 6K2‐induced vesicles are targeted to chloroplasts

Some 6K2 vesicles traffic from the ER to chloroplasts, where they amalgamate and induce chloroplast membrane invaginations. Both the vesicular transport pathway and actomyosin motility systems are involved in this process (Wei, Huang, et al., 2010). Viral RNA, dsRNA, and viral replicase components are concentrated at the level of 6K2 vesicles associated with chloroplasts in infected cells, suggesting that these chloroplast‐bound 6K2 vesicles can also be the site for potyvirus replication (Wei, Huang, et al., 2010). This association of 6K2 vesicles at the periphery of chloroplasts is observed at 48 h post‐infection (hpi). Later in infection (96 hpi), 6K2 vesicles form an elongated tubular structure at the junctions between two adjacent chloroplasts (Wei et al., 2013). This occurs also when the 6K2 protein is expressed alone, independently of any other TuMV‐encoded proteins or viral RNA replication (Wei et al., 2013).

Some regulators of the secretory pathway, the soluble N‐ethylmaleimide‐sensitive factor attachment protein receptors (SNAREs), are crucial for the targeting of 6K2 vesicles to the chloroplast. In the case of TuMV, the ER SNARE protein Syp71 colocalizes with the chloroplast‐bound 6K2 vesicles that make elongated tubular structures (Wei et al., 2013). Down‐regulation of Syp71 inhibits the formation of such structures and reduces TuMV accumulation and virus systemic infection (Wei et al., 2013). However, Syp71 does not interact directly with the 6K2 protein, suggesting that the recruitment of Syp71 to the 6K2 vesicles associated with chloroplasts is not direct. Plant dynamin‐related protein 1 (DRP1) and DRP2 are large multidomain GTPases that play key roles in endocytosis and post‐Golgi trafficking in plants and are also co‐opted by TuMV for infection (Wu et al., 2020). Arabidopsis AtDRP1 and AtDRP2 interact with the TuMV 6K2 protein. AtDRP1‐labelled endosomes colocalize with the chloroplast‐associated TuMV 6K2 vesicles and VRCs, suggesting that AtDRP1 or the AtDRP1‐labelled endosomes are co‐opted by TuMV to support virus replication (Wu et al., 2020). AtDRP2 is also recruited to the VRCs in TuMV‐infected cells (Wu et al., 2018). As AtDRP2 plays an essential role in membrane remodelling and fusion, it is possible that TuMV co‐opts AtDRP2 for VRC assembly. However, the exact mechanical roles of AtDRP1 and AtDRP2 in VRC assembly remain to be further elucidated (Wu et al., 2018, 2020).

3.2.2. Some 6K2‐induced vesicles follow an unconventional trafficking route based on the endosomal/post‐Golgi multivesicular body pathway

Although a proportion of 6K2 vesicles does traffic to the Golgi apparatus, a large number of 6K2 vesicles bypass the Golgi apparatus through an unconventional trafficking route involving the late endosome (LE)/multivesicular body (MVB) pathway (Cabanillas et al., 2018). The central transmembrane domain of the TuMV 6K2 protein, which contains a GxxxG motif (x being any amino acid), is very important for vesicle production and this unconventional trafficking. Indeed, a mutation of glycine (G) residues in this motif leads to relocalization of 6K2 vesicles to Golgi bodies and the plasma membrane and severely affects virus replication (Cabanillas et al., 2018). Furthermore, the TuMV 6K2 protein interacts with Sec22, a SNARE protein that, when overexpressed, has a negative effect on the ER–Golgi equilibrium by impairing the fusion of transport vesicles between the ER and the Golgi apparatus. Overexpression of Sec22 impairs the fusion of 6K2 vesicles with the Golgi apparatus membrane and enhances TuMV intercellular movement, supporting the hypothesis that the Golgi compartment could be a dead end for TuMV 6K2 vesicles when it comes to viral movement (Cabanillas et al., 2018). The 6K2 protein is colocalized and copurifies with Vps10‐interacting protein 11 (Vti11), another SNARE protein localized to the trans‐Golgi network and LEs/MVBs, where dsRNAs are also detected by immunogold electron microscopy (Cabanillas et al., 2018). The biological role of the targeting of 6K2 vesicles to MVBs is strongly supported by the resistance of an Arabidopsis vti11 knockout line to TuMV infection (Cabanillas et al., 2018). Interestingly, Vti11 associates with VSR1, a member of the Vacuolar Sorting Receptor family of proteins, which localizes to prevacuolar compartments (PVCs)/LEs/MVBs (Song et al., 2006). Recently, it was shown that another member of the VSR family, VSR4, interacts with the TuMV 6K2 protein and plays a key role in targeting 6K2 vesicles to MVBs (Wu et al., 2022). In parallel, TuMV infection is impaired in Arabidopsis vsr4 knockout lines (Wu et al., 2022). Altogether, those results show that the intracellular transport of potyvirus 6K2 vesicles that are detached from the ER can bypass the Golgi apparatus, and for some of them, post‐Golgi membrane compartments are used to reach MVBs.

3.2.3. 6K2 vesicles are addressed to PDs and can move from cell to cell

Grangeon et al. (2013) showed that in TuMV‐infected cells, the 6K2 vesicles traffic towards the plasma membrane, where they can be observed in close vicinity to PDs, and have even been reported to move from cell to cell (Grangeon et al., 2013). This very intriguing conclusion is based on confocal microscopy observations performed with 6K2 fused to a photoactivable GFP (6K2:PAGFP) expressed in mock‐ or TuMV‐infected cells. After PAGFP activation, 6K2:PAGFP‐tagged vesicles can be observed moving to the cell periphery and across the cell wall to neighbouring cells. However, this was only observed in the infected condition (Grangeon et al., 2013). These results support the hypothesis that intercellular movement of TuMV may occur in the form of a membrane‐associated viral RNA vesicular complex induced by 6K2 (Grangeon et al., 2013).

Importantly, when expressed alone (in the absence of virus infection), the 6K2 protein can induce the formation of vesicles that are morphologically identical to those observed in infected cells (Figure 2), but these vesicles do not move from cell to cell (González et al., 2019; Grangeon et al., 2013; Lõhmus et al., 2016). However, when coexpressed with 6K2, P3N‐PIPO and CI are necessary and sufficient for the targeting of 6K2 vesicles to PDs and for their intercellular movement. Therefore, the CI and P3N‐PIPO proteins would form the minimal complex required for the intercellular movement of the 6K2 vesicles (Movahed et al., 2017). The current hypothesis is that P3N‐PIPO interacts with and recruits the CI protein to PDs via the PIPO domain (Wei, Zhang, et al., 2010). P3N‐PIPO does not interact directly with 6K2 but interacts with the P3 protein via their shared P3N domain, and this interaction has been suggested to allow the establishment of a connection between CI and 6K2 vesicles at the PD level (Chai et al., 2020). These results support the notion that the replication and cell‐to‐cell movement of potyviruses are coupled through the anchoring of 6K2 vesicles at the entrance of PDs.

FIGURE 2.

6K2‐induced vesicles in Nicotiana benthamiana epidermal cells imaged using confocal microscopy. (a) 6K2 vesicles observed when 6K2‐GFP is expressed alone at 2 days after agro‐infiltration. Chloroplast autofluorescence is shown in red. (b) 6K2 vesicles observed during infection by a recombinant TuMV clone expressing a 6K2‐mCherry fusion protein (TuMV‐6K2‐mCherry) 3 days after inoculation. The 6K2‐induced vesicles are indicated in magenta, while chloroplast autofluorescence is shown in blue. In both cases 6K2‐induced vesicles are observed in close association with chloroplasts. (c) 6K2‐induced vesicles close to the periphery of the cell during infection by a recombinant TuMV clone expressing a 6K2‐mCherry fusion (TuMV‐6K2mCherry) 3 days after inoculation. The 6K2‐induced vesicles are indicated in magenta merged. Scale bar: 10 μm.

3.2.4. 6K2 vesicles could move as extracellular vesicles

Extracellular vesicles have in the past few years emerged as a potential additional route for macromolecule transport in plants and in particular for delivering RNA molecules to distant tissues (Kehr & Kragler, 2018). Extracellular vesicles exist in plants and their proteome significantly overlaps with the PD proteome, suggesting that PD might be a hotspot for extracellular vesicles/exosome transfer between cells (Rutter & Innes, 2017, 2018). Extracellular vesicles can be derived from MVB intraluminal vesicles released upon fusion of the MVB boundary membrane with the plasma membrane (Rutter & Innes, 2018). Interestingly, the Vti11 SNARE, which plays a critical role in TuMV infection (Cabanillas et al., 2018), is also found in the proteome of extracellular vesicles isolated from Arabidopsis (Rutter & Innes, 2017). The functions of Vti11 in both TuMV infection and MVB–plasma membrane fusion suggest the possible existence of extracellular viral vesicles produced through the fusion of TuMV MVBs with the plasma membrane. This hypothesis is strongly supported by the confocal microscopy observations of 6K2‐induced aggregates present in the extracellular space of infected leaves, probably an amalgamation of 6K2 vesicles containing viral RNA and the viral RdRp as shown in xylem‐conducting tubes (Wan, Cabanillas, et al., 2015). This suggests that 6K2 vesicles can enter the extracellular space by exocytosis and possibly thus move between cells (Grangeon et al., 2013; Movahed, Cabanillas, et al., 2019). Indeed, by using TEM, both MVB–plasma membrane fusion events and vesicles present in the extracellular space can be observed in infected tissues, and these vesicles contain dsRNA (Movahed, Cabanillas, et al., 2019). Furthermore, the presence of 6K2 vesicles in xylem vessels suggests that viral components could also be found in other apoplastic regions of the plant (Wan, Cabanillas, et al., 2015; Wan & Laliberté, 2015). Altogether, these results highlight a potential novel pathway for intercellular movement of potyviruses that would represent an important change of paradigm for plant virus movement (Richardson, 2019).

In the previous part, we have highlighted the progress made in understanding how potyviruses recruit the host plant endomembrane system and cytoskeleton to move intracellularly to the cell wall and PDs, with a special focus on the role played by 6K2 vesicles in this process. A major question remains to be answered: What is the nature of the viral “entity” that crosses the PD? As discussed above, 6K2 vesicles are able to transit to noninfected cells; however, the exact composition of their viral load still remains to be determined, which is very technically challenging. At this point, it cannot be excluded that potyviral RNA moves from cell to cell through PDs, associated with still unknown plant factors and in a 6K2‐independent way. According to this hypothesis, the 6K2 vesicles could have the functions of allowing the synthesis and bringing the viral material (genomic RNA, viral proteins) close to PDs. It is also not known whether the encapsidation process precedes the transport of virions or whether the genomic RNA is transported in the form of a ribonucleoprotein complex. The demonstration that only replication‐competent RNA can be encapsidated into mature potyviral particles suggests that viral RNA replication, virion assembly, and viral movement are closely interconnected for potyviruses (Gallo et al., 2018).

In the next sections we will focus on the molecular and cellular events that take place at the PD level during potyvirus infection and address the following still open question: Do potyviruses move between cells through PDs as ribonucleoprotein complexes or as virions?

4. POTYVIRUSES COUNTERACT VIRAL‐INDUCED CALLOSE DEPOSITION AT PDS

Plant infection by viruses has been associated with callose regulation at the PD level. Callose is a 1,3‐β‐glucan that accumulates in cell wall microdomains surrounding PDs, regulating the cell‐to‐cell transport of macromolecules by changing the SEL (German et al., 2023). A negative role of callose accumulation in PD permeability has been confirmed experimentally, yet the roles of cytoskeletal elements and many PD‐associated proteins in this phenomenon still remain unclear. Many viruses have been reported to induce PD callose accumulation, including potyviruses (Zavaliev et al., 2013). Nevertheless, for efficient movement, potyviruses must somehow overcome the effect of callose deposition at PDs. Potyvirid infection is associated with the up‐regulation of genes involved in callose degradation, such as cell wall β‐1,3‐glucanases, as observed during cassava brown streak ipomovirus infection (Anjanappa et al., 2018) and zucchini yellow mosaic virus infection (Amoroso et al., 2022). In response to SMV infection, soybean endo‐1,3‐β‐glucanase (GmGLU) expression is also increased and GmGLU, among other enzymes, regulates callose deposition at the PD (Shi et al., 2020).

The roles of potyviral CI and P3N‐PIPO in regulating callose deposition at PDs have been highlighted in different studies. Ectopic expression of potato (Solanum tuberosum) REMORIN (StREM1.3), a plant protein that localizes at lipid rafts and PDs, in Nicotiana benthamiana increases PD callose deposition. However, ectopic expression of StREM1.3 in N. benthamiana affects virus spread differently: The movement of the potexvirus PVX is restricted, while the propagation of two potyviruses, TuMV and potato virus A (PVA), is enhanced (Perraki et al., 2018). The observation that the TuMV CI protein interacts with StREM1.3 at the plasma membrane and at the PD level suggests that the CI protein could counteract the role of StREM1.3 in promoting callose deposition at PDs to favour TuMV propagation, but the underlying mechanisms remain to be determined (Rocher et al., 2022). Furthermore, the presence of the CI protein of pea seedborne mosaic virus (PSbMV) at the PD level is linked with an apparent transient reduction in callose deposition in the vicinity of PDs (Roberts et al., 1998), suggesting that the CI protein might transiently increase the SEL to allow viral movement. However, this notion is not supported by microinjection experiments performed by Rojas et al. (1997), who showed that the CP, but not the CI protein, can increase the SEL of PDs. On the other hand, the SEL of PDs has been shown to be increased by P3N‐PIPO, which interacts with both CI and a divalent cation‐binding plasma membrane protein (PCaP1) via its C‐terminal PIPO domain (Chai et al., 2020; Vijayapalani et al., 2012). Therefore, it has been proposed that P3N‐PIPO could control the Ca²⁺ concentration around PDs, which could in turn affect the SEL by regulating callose deposition and severing the actin system (Cheng et al., 2017, 2020). Indeed, both actin and myosin localize in the PD pore and regulate not only the gating to but also the trafficking through it (Diao & Huang, 2021).

Besides callose regulation at the cell wall microdomains surrounding PDs, cell wall remodelling could also occur during potyvirus infection, involving a cell wall‐loosening protein, NbEXPA1 (Park et al., 2017). This α‐expansin is a protein involved in cell expansion, leaf development, and growth and has been shown to be a PD‐located protein in N. benthamiana (Park et al., 2017). When ectopically expressed, NbEXPA1 promotes TuMV replication and intercellular movement, suggesting that this PD‐specific cell wall‐loosening protein could be involved in PD modification and facilitate potyviral cell‐to‐cell movement (Park et al., 2017). A potential role of expansins in the regulation of PD gating was described in a very particular study system, the elongation of single‐celled cotton fibres (Ruan et al., 2001). The gating of the basal PDs connecting the fibres to the seed coat is highly coordinated with the expression of cell wall expansin during cotton fibre elongation (Ruan et al., 2001). Whether NbEXPA1 in leaves is also involved in the regulation of PD gating remains to be shown.

Altogether, those results show that potyviruses probably modulate callose deposition and possibly other cell wall parameters at the PD level in order to favour their cell‐to‐cell movement, although the exact underlying mechanism(s) still remain(s) to be determined.

5. DOCKING OF POTYVIRAL “MOVEMENT PROTEINS” AT THE PD LEVEL

Different ultrastructural analyses reviewed by Sorel et al. (2014) reveal that cylindrical inclusions are localized in close vicinity to the plasmalemma during early infection. In particular, the “step‐by‐step” ultrastructural analysis of PSbMV infection of pea cotyledons has shown that cytoplasmic inclusion bodies are detected at both sides of PD connecting two cells at the front edge of the viral infection (Roberts et al., 1998). Other studies with different potyviruses confirm that the CI is associated with pathognomonic cone‐shaped structures close to PDs and in the vicinity of the cytoplasmic inclusion bodies, where CP and viral RNA are also detected (Otulak & Garbaczewska, 2012; Roberts et al., 1998; Rodriguez‐Cerezo et al., 1997; Rojas et al., 1997). Furthermore, a TuMV CI protein mutated in N‐terminal charged residues failed to form conical inclusions and to interact with the virus CP at PDs, and the corresponding mutants are altered in their cell‐to‐cell movement, suggesting a functional link between CI localization at PDs and potyvirus movement (Deng et al., 2015).

The recruitment of the CI protein to PDs is mediated by its interaction with P3N‐PIPO, and the CI protein could then serve as a docking point for the intercellular movement of viral material or intact virions (Deng et al., 2015; Movahed et al., 2017; Wei, Zhang, et al., 2010). The anchoring of P3N‐PIPO to PDs depends on its interaction with plasma membrane‐associated PCaP1 (Vijayapalani et al., 2012). Indeed, knockout of PCaP1 in Arabidopsis thaliana induces a strong reduction in TuMV movement (Vijayapalani et al., 2012). Furthermore, the regulated plasma membrane protein NbDREPP in N. benthamiana, considered as the homologue of AtPCaP1, interacts with both P3N‐PIPO and the CI protein of tobacco vein banding mosaic virus (TVBMV) at the PD level. Its knockdown impedes the cell‐to‐cell movement of TVBMV, confirming the key role played by PCaP1 at the PD level for potyvirus transport (Geng et al., 2015). It is hypothesized that PCaP1 recruits the TuMV P3N‐PIPO protein to PDs and the actin filament‐severing activity of PCaP1 is required for TuMV intercellular movement (Cheng et al., 2020). However, Arabidopsis AtREM1.2 negatively regulates the cell‐to‐cell movement of TuMV via competition with PCaP1 for binding actin filaments (Cheng et al., 2020). This negative effect of AtREM1.2 on potyvirus movement contradicts the positive effect observed for StREM1.3 (Rocher et al., 2022), adding another level of complexity concerning the functions of REMORINs in viral infections.

Another protein involved in membrane trafficking, the synaptotagmin SYTA, has been shown to regulate P3N‐PIPO intercellular trafficking (Uchiyama et al., 2014). SYTA localizes to the sites where the ER makes contact sites with the plasma membrane, such as at the PD sites. TuMV propagation and the cell‐to‐cell movement of P3N‐PIPO‐GFP fusion proteins are delayed in Arabidopsis syta‐1 knockdown mutants. The authors suggest that SYTA could play a role in the way P3N‐PIPO, with its cargo of CI and viral material, could be directed to PDs. However, such hypothesis is not supported by the fact that no interaction between P3N‐PIPO and SYTA has been described so far (Uchiyama et al., 2014).

6. DO POTYVIRUS VIRIONS CROSS THE PD?

The structural protein CP is also part of the potyviral multicomponent transport system (Martínez‐Turiño & García, 2020). However, as detailed below, it is still not clear whether potyviruses are transported through PDs as virions or as nonvirion ribonucleoprotein complexes.

Studies on filamentous plant virus particles using atomic force microscopy (AFM) revealed the presence of protruding tip structures on PVY and PVA virions (Gabrenaite‐Verkhovskaya et al., 2008; Torrance et al., 2006). At the level of these protruding tips at the 5′ end of the PVA particles, the CI protein is detected, together with the VPg and HC‐Pro proteins (Gabrenaite‐Verkhovskaya et al., 2008). The role of those structural supplements at the virion end is still unknown; they may play roles at different viral infection stages, such as virus assembly/disassembly, movement, and vector transmission (Torrance et al., 2006). A possible hypothesis, based on the ATPase and RNA helicase activities of the CI protein, is that the CI protein could behave as a molecular motor to both disassemble virions and translocate the viral genome through PDs (Gabrenaite‐Verkhovskaya et al., 2008).

Pioneering studies such as that of Weintraub et al. (1974) suggest that PVY moves from cell to cell as particles, but due to limitations of imaging technologies, as well as the absence of specific labelling of CP in these experiments, this conclusion remains highly speculative. Even after immunolabelling experiments in other studies, it is difficult to conclude whether CP labelling within PDs corresponds to virions or to CP involved in nonvirion ribonucleoprotein complexes (Riedel et al., 1998; Roberts et al., 1998; Varrelmann & Maiss, 2000). Indeed, based on immuno‐electron microscopy observations, CP has been detected in the central core of the CI pinwheel inclusions, leading to the hypothesis that CI could position and even guide, like a funnel, viral particles across the cell wall (Roberts et al., 1998). Linear‐shaped CP “complexes” inside or attached to cylindrical inclusions near PD connections in tobacco leaves infected with tobacco vein mottling virus were also observed, but it was not possible to conclude whether these complexes are virions targeted for transport or nonvirion ribonucleoprotein complexes (Rodriguez‐Cerezo et al., 1997). Even though viral particles are observed near PDs, late in infection (Roberts et al., 1998), neither a “queue” of “waiting” virions nor “engaged” virions in PDs have been observed or reported in any electron microscopy studies.

To the best of our knowledge, there is no clear experimental evidence that links the RNA‐binding capacity of a potyvirus CP with virus cell‐to‐cell movement, except the data from Yan, Xu, Fang, Cheng, et al. (2021) showing that mutations of two residues in the RNA‐binding pocket of the CP from TVBMV reduced the RNA‐binding activity of CP and abolished virus cell‐to‐cell movement and replication.

As reviewed above, reverse genetics experiments showed that in addition to being the potyvirus structural protein, mutations or deletions of CP affect the intercellular movement of TEV and PPV, without impairing viral replication but hampering assembly of the particles (Roberts et al., 1998; Varrelmann & Maiss, 2000). This essential role of multiple conserved aromatic residues in the core CP in CP accumulation and virus movement was confirmed for TVBMV, watermelon mosaic virus, and PVY (Yan, Xu, Fang, Geng, et al., 2021). Mutations of three charged residues, arginine R245, histidine H246, and aspartic acid D250, in the C‐terminal domain also impair intercellular movement of SMV (Seo et al., 2013), while deletions in the CP core negatively impact TuMV particle assembly (Yuste‐Calvo et al., 2020) and impair TEV cell‐to‐cell movement and the production of virus particles (Dolja et al., 1994, 1995). Those results suggest that some potyviruses probably move as virions. On the other hand, for TuMV, intact virions are not needed for intercellular spread, because deletions in the N‐terminal part of CP induce the formation of aberrant long forms of virions without impairing cell‐to‐cell or systemic spread (Dai et al., 2020). In parallel, encapsidation is not sufficient for cell‐to‐cell or long‐distance movement of TEV and PVY as those potyviruses can form virions despite deletion of the C‐terminal part of their CP, abolishing movement (Dolja et al., 1995; Kežar et al., 2019). Other mutations in CP, leading to movement‐defective viruses, can be partially rescued by transcomplementation in CP transgenic plants, but without the formation of visible virions (Dolja et al., 1994, 1995), supporting the assumption that cell‐to‐cell and long‐distance transport of potyviruses are facilitated by their CP but without an absolute requirement for virion formation. Therefore, some conclusions about potyviruses moving as virions, based on reverse genetics experiments but without any experimental proof of virions crossing PDs (Seo et al., 2013), should probably be considered with caution. In summary, there is currently conflicting evidence about the need for encapsidation for potyvirus movement. In some cases, mutations have been identified that simultaneously prevent viral particle formation and potyviral movement, while other experimental results show that cell‐to‐cell movement occurs without accumulation of viral particles. In addition to particle encapsidation, mutations in CP could also modify its interactions with viral RNA or with other viral or cellular proteins, impacting vesicular transport and cell‐to‐cell spread. As a result, potyviruses could be transported through PDs as virions, as nonvirion ribonucleoprotein complexes involving CP, or even possibly as a combination of these two nonexclusive modes.

7. CONCLUSION

Taken together, data from this review including genetic, cellular, and ultrastructural studies highlight how potyvirus transport involves a large network of interactions between viral and cellular proteins. Despite now extensive investigations, whether potyviruses move between cells as virions or other movement complexes, such as ribonucleoproteins or 6K2 vesicles, still remains an unsolved conundrum, setting aside potyviruses from the majority of plant viruses. However, evidence obtained so far shows that viral replication, encapsidation, and movement are very closely interconnected in the case of potyviruses. Observations of 6K2 vesicle trafficking between cells and of viral extracellular vesicles containing dsRNA (potential replicative forms of viral RNA) penetrating and possibly even crossing the cell wall are highly innovative results that suggest the existence of a totally novel mechanism for plant virus cell‐to‐cell spread. Whether those results can be generalized to all potyvirus species besides TuMV and may apply to other plant virus genera remains to be determined.

Based on data gathered in this review, we propose an updated working model integrating all the identified pathways potentially involved in potyvirus movement (Figure 3). It must be noted that in this comprehensive model of the intracellular and intercellular movement of potyviruses, the chronological order shown in Figure 3 is provided for presentation clarity but probably does not reflect the biological reality, as some described events can occur simultaneously. Figure 3 shows that upon entry of the viral particle into the cell via aphid transmission or mechanical injury, the virion is uncoated (1). The positive‐sense RNA viral genome is translated and the synthesized viral proteins, including the 6K2 protein and co‐opted host factors, induce a drastic remodelling of the ER and Golgi endomembrane systems associated with the formation of a large perinuclear globular structure (VRC) (2) and the release of mobile 6K2 vesicles that move intracellularly following transvacuolar actin strands (3). These 6K2 vesicles contain viral RNA together with host and viral proteins involved in translation and replication. Some 6K2 vesicles target chloroplasts, where replication can occur (4). Others bypass the Golgi apparatus and become associated with PVCs/MVBs (5). MVBs can then fuse with the plasma membrane and release their vesicular content in the extracellular space, where the vesicles can move through pores in the cell wall to the neighbouring cell (5a). The 6K2 vesicles can also reach the plasma membrane and become associated with PDs, a process relying on the P3N‐PIPO and CI proteins, in association with host factors such as PCaP1 for PD gating, docking, and intercellular traffic of 6K2 vesicles (5b). The delivery of 6K2 vesicles to PDs probably increases the concentration of viral replication products nearby. After downloading of the content of the vesicles, newly synthesized genomic RNAs can be packaged by CP in virions in the vacuoles or be transported through PDs either as viral ribonucleoprotein complexes involving the CP (5c) or as nascent virions (5d). Investigations are still in dire need in order to sort out or to reconcile all these potential routes of intracellular and intercellular propagation of potyviruses and if possible to establish a chronology linking them.

FIGURE 3.

Schematic illustration of the different models for potyviral intercellular transport. After entry of the virus via aphid stylets or through mechanical injury, virions are uncoated (1) and genomic RNA is released in the cytoplasm, where translation occurs. The viral 6K2 protein induces the formation of a large perinuclear globular structure (VRC) associated with drastic remodelling of the endoplasmic reticulum (ER) and Golgi endomembrane systems (2) and of ER‐derived 6K2 vesicles involved in replication (3). A subpopulation of 6K2 vesicles targets the Golgi apparatus and chloroplasts, where replication occurs (4). The 6K2 vesicles can also bypass the Golgi apparatus and are associated with the prevacuolar compartments/late endosomes/multivesicular bodies (PVCs/LEs/MVBs) (5). MVBs can fuse with the plasma membrane and release their vesicular content in the extracellular space, where the vesicles can move through pores in the cell wall to neighbouring cells (5a). 6K2 vesicles can also reach the plasma membrane and associate with plasmodesmata (PDs), a process relying on the P3N‐PIPO and cylindrical inclusion (CI) proteins, in association with host factors such as PCaP1 for PD gating. The CI protein associated with 6K2 vesicles can serve as a docking point at the PDs for their intercellular movement (5b). The delivery of 6K2 vesicles to PDs probably increases the concentration of viral replication products nearby. After downloading of the content of the vesicles, newly synthesized genomic RNAs can be packaged by the coat protein (CP) in virions in the vacuole or be transported through PDs either as viral ribonucleoprotein complexes associated with the CP (5c) or as nascent virions (5d).

Xue, M. , Arvy, N. & German‐Retana, S. (2023) The mystery remains: How do potyviruses move within and between cells? Molecular Plant Pathology, 24, 1560–1574. Available from: 10.1111/mpp.13383

DATA AVAILABILITY STATEMENT

Not applicable.