In this issue of New Phytologist (Deeks et al., pp. 529–540), Patrick Hussey and his coworkers report on the very distinct localization of Arabidopsis thaliana formins, AtFH4 and AtFH8, to cross-walls of roots, hypocotyl and shoot tissues. This is the first time that plant formins are reported to have such distinct domain-specific subcellular localizations. Here we discuss these pertinent findings from the broader perspective of plant cell polarity, cell wall–cytoskeleton adhesion domains, polar auxin transport, and the emerging unique status of these cross-walls in that they resemble neuronal and immunological synapses (Baluška et al., 2003a,b,c, 2005).
‘The recent finding by Deeks et al., that AtFH4 and AtFH8 are localized to the cross-walls of roots, hypocotyl and shoot tissues, is relevant for our understanding of the nongrowing but extremely dynamic actin-based cross-walls.’
In 2000, Fatima Cvrčková carried out a bioinformatic approach to search for plant formins; eight putative formin-coding genes were idenitified (Cvrčková, 2000). It was of great surprise to find that the majority of these plant formins contained potential transmembrane domains which are not present in the yeast and animal formins, and suggests that they are integral membrane proteins. Moreover, some of the plant formins also contained an exposed proline-rich domain which presumably inserts into the cell wall, exhibiting similarity to cell wall extensin. From these bioinformatic analyses, it was predicted that some plant formins might be involved in the direct cell wall–cytoskeleton communication (Cvrčková, 2000; Baluška et al., 2003c). During the last two years, our knowledge of plant formins has literally expanded. Bioinformatics data are now well advanced (Cvrčkováet al., 2004), and there are also first data on both cell biology (Cheung & Wu, 2004; Favery et al., 2004; Ingouff et al., 2005; Yi et al., 2005) and biochemistry (Michelot et al., 2005; Yi et al., 2005). This most recent study by Deeks et al. opens new avenues in our understanding of plant-specific formins as they report domain-specific enrichment of AtFH4 and AtFH8 at cross-walls of diverse plant organs. This raises the intriguing question as to what is so specific about these cross-walls.
Cross-walls: from auxin transport to vesicle recycling
Cross-walls of longitudinal cells have long been suspected to have special signaling properties, simply because it has been known for many years that developmental signals and cues are spread primarily along the longitudinal axis of organs such as roots and stems (Baluška et al., 2003a). Moreover, the cross-walls are known to harbour numerous plasmodesmata which interconnect adjacent cells of cell files into syncytium-like supercells (Baluška et al., 2003a,c). Recently, the interest into these subcellular domains increased further as a result of advances made concerning proteins that assist in the polar transport of auxin. Because this plant hormone is transported preferentially along cell files, the cross-walls inevitably represent domains which are transporting auxin from cell to cell (Baluška et al., 2003b, 2005).
For unknown reasons, auxin is not transported across plasmodesmata even though its small size should guarantee a free passage. In light of this, a theoretical model was proposed and followed which considers plasma membrane transporters (putative influx and efflux carriers) which should drive the polar transport of auxin across cross-walls (reviewed by Friml & Palme, 2002; Friml & Wiśniewska, 2005). This so-called chemiosmotic model is based on the chemical properties of auxin and acidic pH values found in cell walls vs basic pH values of the cytoplasm. In accordance with this model, putative auxin transporters are localized in a polar fashion (reviewed by Friml & Wiśniewska, 2005). Further new data are rather at variance with this model. For instance, although the early concept of polar auxin transport did not consider vesicle trafficking at all, this process slowly penetrated all the papers dealing with the auxin transport because putative efflux carriers turned out to perform rapid recycling between the plasma membrane and putative plant endosomes (Friml & Wiśniewska, 2005). The chemiosmotic model has difficulties to explain why plasma membrane transporters undergo such a rapid recycling rate. Nevertheless, one can easily explain this point by invoking impacts of developmental cues and signaling cascades on the flow of auxin via rapidly reshifting polar subcellular localizations of these putative auxin transporters.
However, there are more serious weak points of this model which claim the plasma membrane transporter should drive transcellular transport of auxin. First of all, despite the suggestive polar localization of putative transporters (including PIN1, PIN2, PIN3, PIN4 and AUX1) to the cross-walls (reviewed by Friml & Wiśniewska, 2005), all attempts to prove the plasma membrane transporter nature of these proteins has failed until now. Hence, the consensus of opinion has been slowly shifting to consider the carrier nature of these proteins as transport facilators or regulators. Secondly, brefeldin A (BFA; a potent inhibitor of vesicular secretion in plants, as in other eukaryotic cells) blocks polar auxin transport within a few minutes of application (Delbarre et al., 1998; Paciorek et al., 2005). Importantly, BFA not only exerts rapid inhibition of the auxin efflux but also causes the complete block of this process, whereas auxin influx is not affected (Delbarre et al., 1998). Because BFA blocks exocytosis while it stimulates endocytosis (Wang et al., 2005), these rapid effects of BFA on the polar auxin transport correspond well to the ‘neurotransmitter’ nature of auxin, being secreted out of exporting cells and perhaps taken up via endocytosis by a receiver cells (Baluška et al., 2003b; Friml & Wiśniewska, 2005). Finally, inhibitors of the polar auxin transport, irrespective of their chemical nature, turned out to act as inhibitors of endocytosis (Geldner et al., 2001, 2003). From the chemiosmotic model perspective, it is a mystery why inhibitors of endocytosis should block the polar transport of auxin, whereas the alternative ‘neurotransmitter’ model (Friml & Palme, 2002; Baluška et al., 2003b; Friml & Wiśniewska, 2005) can easily explain this conundrum.
Although all PINs as well as AUX1 are known to accomplish vesicular recycling at the cross-walls, and although they get trapped into the endocytic BFA-induced compartments in BFA exposed cells, this happens only after some 10–15 min, whereas the full size of BFA compartments is achieved only after 120 min (Geldner et al., 2001, 2003). However, BFA inhibits polar transport of auxin immediately after exposure, when most of the carriers are still localized to the plasma membrane (see fig. 2o in Paciorek et al., 2005). In addition, even after 90 min of BFA treatment, when BFA-induced compartments have reached large size, there is still a considerable portion of auxin carriers localized to the plasma membrane at cross-walls (Paciorek et al., 2005). Again, this is at variance with the chemiosmotic model but corresponds with the ‘neurotransmitter’ model of polar auxin transport.
The only known process which is blocked within a few minutes of BFA exposure is vesicular secretion, irrespective of whether it is the constitutive Golgi-apparatus-based secretion or the endocytosis-based and vesicle-recycling-driven regulated secretion (Šamaj et al., 2005). Therefore, the most plausible explanation of the very rapid blockage of auxin transport via BFA is that auxin is secreted via recycling-based regulated secretion. In support of this latter notion, we have recently localized auxin into vesicular structures as well as within endocytic BFA-induced compartments (our own unpublished data). Moreover, depolymerization of F-actin, which prevents endocytosis of auxin carriers (Geldner et al., 2001, 2003), and thus maintains them at the plasma membrane, inhibits polar auxin transport too (Sun et al., 2004). This latter finding is again at variance with the chemiosmotic model. Importantly, the F-actin-dependence of polar auxin transport implicates that F-actin nucleators will be critical for our mechanistic understanding of processes driving polar auxin transport across cellular boundaries.
Cross-walls: actin-, myosin VIII- and formin-enriched domains specialized for endocytosis, rapid vesicle recycling and signaling
Cross-walls are known to be actin-enriched domains (Baluška et al., 2003a). This is rather surprising because cross-walls are nongrowing domains in postmitotic root cells. Most eukaryotic cells, with the exception of neuronal synapses, assemble dense F-actin meshworks typically at growing domains (Baluška et al., 2003a). Recently, however, it is becoming clear that extensive endocytosis and vesicle recycling (Šamaj et al., 2005) is going on under nongrowing cross-walls, which balances exocytosis to such an extent that there is no net growth of cell periphery at these highly specialized subcellular domains (Baluška et al., 2003a,b,c). Thus, cross-walls resemble the neuronal synapses (Baluška et al., 2003b) and the imperative question that emerges concerns what molecules act as actin nucleators at the synaptic cross-walls.
The first obvious candidate was the ARP2/3 complex; indeed, knocking out this complex results in disassembly of cross-walls in epidermal cells (Basu et al., 2005; Mathur, 2005). However, cross-walls of nonepidermal cells remained intact in cells devoid of the ARP2/3 complex (reviewed by Mathur, 2005) and the overall phenotype of these mutants are mild, suggesting the existence of another powerful F-actin nucleator. Indeed, recent studies confirmed that plant formins are potent F-actin nucleators (Michelot et al., 2005; Yi et al., 2005). Therefore, the recent finding by Deeks et al., that AtFH4 and AtFH8 are localized to the cross-walls of roots, hypocotyl and shoot tissues, is relevant for our understanding of the nongrowing but extremely dynamic actin-based cross-walls. In addition to actin, cross-walls are enriched also with myosins of the class VIII type, and both actin and myosin VIII are known to be important for endocytosis (reviewed by Šamaj et al., 2005). Moreover, profilin was also localized to cross-walls (Baluška et al., 2001a) and, interestingly in this respect, AtFH4 binds to profilin and affects actin polymerization (Deeks et al.). Besides the putative auxin transporters, cell wall pectins are also internalized via the same recycling pathways and become trapped within the BFA compartments under the exposure of root apices to BFA (Šamaj et al., 2005). Pectins are well known to act as adhesive agents of plant cells (Lord & Mollet, 2002), and their recycling at the cell-cell adhesive cross-walls suggests that their function is tightly controlled via these recycling processes, which themselves are targets of developmental cues and signaling cascades. In fact, cross-walls in root apices are also enriched with plant Rho GTPases known as ROPs (Molendijk et al., 2001) and MAP kinases (J. Šamaj, University of Bonn, pers. comm.).
Cross-walls as actin- and pectin-based adhesion domains: do formins and myosins of the class VIII act as elusive adhesive molecules of plant cells?
Plants lack integrins and it still remains a mystery as to which molecules act as the linkers between cell wall components and the cytoskeleton. Recently, we surveyed all candidates and proposed that myosins of the class VIII and formins represent the best candidates for this role (Baluška et al., 2003c). Myosin VIII is enriched at subcellular cell periphery sites involved in callose synthesis and it is possible that it binds directly to one of the callose synthase subunits. This would interlink the cell wall with cytoskeleton via callosic cell periphery domains at plasmodesmata and pit-fields. On the other hand, plant formins of the group I are equipped with an extensin-like domain which is predicted to be inserted into the cell wall (Cvrčková, 2000; Cvrčkováet al., 2004). Again, this would provide plant cells with a direct linkage between the cell wall and cytoskeleton.
These two types of cell wall–cytoskeleton linkages would satisfy the plant-specific demands for a very dynamic cell periphery because plant cells often suffer from osmotic stress and respond with very rapid retraction of their plasma membrane/protoplast from the cell wall (Baluška et al., 2003c). Interestingly, this is associated with very rapid callose synthesis, especially to pit-fields at cross-walls, and recruitment of myosin VIII to these sites of callose synthesis (Wojtaszek et al., 2005). It would be interesting to test if formins, too, are recruited to these sites of enhanced adhesion sites.
Formins as synaptic proteins of plants?
More than 100 years ago, Bohumil Němec described in great detail very prominent longitudinal F-actin cables interacting at cross-walls, using the classical cytological methodology (Fig. 1a; Němec, 1901). Today, actin antibodies reveal two different types of F-actin arrays assembled at cross-walls. The first one is the very dense submembraneous meshwork (Fig. 1b) which is involved, together with myosin VIII, in the endocytosis and vesicle recycling. The second one is composed of distinct cables which traverse the cell longitudinally, interconnecting the opposite cross-walls and typically contacting the nuclear surface (Fig. 1b). Both the meshworks as well as longitudinal cables are essential for cell-to-cell communication and are sensitive to BFA (Fig. 1c). Formins are predicted to organize both these F-actin arrays: the group I formins which associate with cross-walls (Deeks et al.) may be relevant for the dense meshworks, whereas the group 2 formins can be expected to be important for assembly of thick cables (Fig. 2). The dynamic meshworks drive the endocytic recycling related to chemical neuronal synapses as well as immunological synapses. On the other hand, bundles interconnect the opposite cross-walls enriched with plasmodesmata, which might act as electrical neuronal synapses, and hence be involved in the rapid spread of signals, as proposed by Němec. Future studies will unveil how formins nucleate diverse arrays of F-actin at plant synapses in response to developmental cues and signaling cascades which ultimately impinge on the auxin transporting machinery that is based on BFA-sensitive vesicle recycling.