TY - JOUR
AU1 - Tapken, Wiebke
AU2 - Murphy, Angus S.
AB - Abstract The plasma membrane is the interface between the cell and the external environment. Plasma membrane lipids provide scaffolds for proteins and protein complexes that are involved in cell to cell communication, signal transduction, immune responses, and transport of small molecules. In animals, fungi, and plants, a substantial subset of these plasma membrane proteins function within ordered sterol- and sphingolipid-rich nanodomains. High-resolution microscopy, lipid dyes, pharmacological inhibitors of lipid biosynthesis, and lipid biosynthetic mutants have been employed to examine the relationship between the lipid environment and protein activity in plants. They have also been used to identify proteins associated with nanodomains and the pathways by which nanodomain-associated proteins are trafficked to their plasma membrane destinations. These studies suggest that plant membrane nanodomains function in a context-specific manner, analogous to similar structures in animals and fungi. In addition to the highly conserved flotillin and remorin markers, some members of the B and G subclasses of ATP binding cassette transporters have emerged as functional markers for plant nanodomains. Further, the glycophosphatidylinositol-anchored fasciclin-like arabinogalactan proteins, that are often associated with detergent-resistant membranes, appear also to have a functional role in membrane nanodomains. Detergent-resistant membranes, membrane nanodomains, nanodomain function, ordered membrane domains, plasma membrane ABC proteins, sphingolipids, sterols. Introduction Lipid membranes define the boundary of a cell, its organelles, and secretory subcompartments. They spatially separate incompatible biochemical processes, provide structural support for transporters and receptors, function as exchange surfaces for mineral ions and signalling molecules, and are the initial point of contact for host–pathogen interactions. Glycerolipids [mainly phospholipids (PLs)], sphingolipids (SLs), and sterols are the three main lipid classes that constitute the plant plasma membrane (PM), and define membrane organization and function. Recent combinatorial studies using high-resolution microscopy and lipid biosynthesis inhibitors suggest that proteins form specialized laterally defined PM microenvironments characterized by SLs and sterols (Lasserre et al., 2008, Raffaele et al., 2009; Demir et al., 2014; Jarsch et al., 2014). These nanodomain (ND) structures have been shown to increase the stability and activity of embedded proteins and protein complexes: NDs have been associated with receptor complex function (Sun et al., 2002), transporter efficiency (Blakeslee et al., 2007; Titapiwatanakun et al., 2009), channel regulation (Demir et al., 2013), protein trafficking (Men et al., 2008; Yang et al., 2013), and plant–bacterial interactions (Bhat et al., 2005; Haney and Long, 2010; Lefebvre et al., 2010). The terms lipid rafts and detergent-resistant membranes (DRMs) are often used synonymously with NDs, largely for historical reasons. The concept of lipid rafts was introduced to describe membrane ‘patches’ in which discrete groups of proteins were localized in mammalian cells (Simons and Ikonen, 1997). DRMs are physically isolated fractions of denser sterol- and SL-enriched membranes, which are obtained through extraction at 4 °C with non-ionic detergents, such as Triton X-100 (Brown and Rose, 1992; Mongrand et al., 2004; Borner et al., 2005; Martin et al., 2005). The term ‘lipid raft’ emerged as a conceptualization of immiscible, sterol- and SL-rich ordered membrane domains with compositions similar to those of DRMs. Co-purification of certain proteins with DRMs suggested that SLs and sterols could form distinct sites of viral entry in mammalian cells (Nguyen and Hildreth, 2000; Samuel et al., 2001; Bavari et al., 2002; Chazal and Gerlier, 2003). This concept was later also applied to plants, and thus DRM analyses paved the way for studies that associated ordered membrane domains, or NDs, with physiological functions (Lasserre et al., 2008; Demir et al., 2013; Yang et al., 2013). Biochemical association of lipids and proteins with DRMs does not directly establish the presence of ordered protein and membrane complexes in plants any more than a sucrose density gradient fractionation of membranes conclusively demonstrates association of a particular protein with a specific cellular membrane structure. Interchangeable use of the terms ‘lipid raft’ and DRM is, therefore, inappropriate. Some authors suggest that the term ‘lipid raft’ should not be used to describe any type of plant-associated ordered membrane domain and question that these domains occur in plants (Tanner et al., 2011). This review does not in any way imply that this metaphorical term should be used in the plant literature. A number of artefacts are generated in DRM preparation. For instance, the strength of the non-ionic detergent used to isolate DRMs can alter the composition of enriched sterols, SLs, and associated proteins, and can also increase the membrane affinity of cytosolic proteins (Titapiwatanakun et al., 2008; Tanner et al., 2011). More importantly, solubilization with Triton X-100 results in the temperature-dependent restructuring of lipid domains, as observed in giant unilamellar vesicles (Heerklotz, 2002; Casadei et al., 2014). Thus, DRM extraction is a selective isolation method and it is possible that the composition of functional NDs differs in living cells. Due to its dependence on the isolation method used, the lipid and protein composition of isolated DRMs is not identical to that of NDs. However, there is increasing evidence that ordered membrane structures do exist in plants and that they are functionally relevant. Technological advances within the field of microscopy, including new probes for atomic force microscopy (AFM), super resolution confocal microscopy, high-resolution total internal reflection fluorescence (TIRF), the variable angle epifluorescence microscopy (VAEM) variant of TIRF used in plants (Konopka and Bednarek, 2008), and enhanced fluorescence recovery after photobleaching (FRAP), have enabled more precise definition of the size and composition of ND structures (Gutierrez et al., 2010; Truong-Quang and Lenne, 2014). The identification of SL and sterol biosynthesis mutants in Arabidopsis, in conjunction with these high-resolution imaging techniques, has provided a context for analyses of the contribution of these lipids to protein trafficking, membrane structure, and physiology (Willemsen et al., 2003; Men et al., 2008; Roudier et al., 2010; Markham et al., 2011; Yang et al., 2013). As a consequence, ND size estimations in plants have progressively decreased, from the micrometre scale to the order of 10–100nm, which is comparable with large protein complexes, such as the cellulose synthase complex (Kimura et al., 1999; Cacas et al., 2012; Truang-Quong and Lenne, 2013). The wide range of ND sizes has spawned the use of the ‘microdomain’ and ‘nanodomain’ nomenclature to better distinguish between differently sized complexes (Mongrand et al., 2010; Demir et al., 2013). However, the terms still lack concrete size delimitation. In summary, the current consensus is that the term DRM is utilized for lipids and proteins that have been biochemically associated with SL- and sterol-enriched membranes, and NDs as the functional assemblies of lipids and proteins, when determined by biophysical or microscopic techniques such as super resolution, TIRF, or AFM. In this review we will discuss the current understanding of what determines protein association with NDs, where these functional domains are formed, and how the lipid environment affects the trafficking of PM proteins. In particular, we focus on the role of NDs in regulating the trafficking, localization, stability, mechanistic function, and interactions of PM proteins, with an emphasis on ATP binding cassette (ABC) transporters from the B and G subgroups in Arabidopsis. Characteristics of the plant lipidome Cholesterol and ergosterol are the most abundant sterols in vertebrates and fungi, respectively (Shieh et al., 1977; Dupont et al., 2012). Yeast DRMs are enriched in ergosterol (Bagnat et al., 2000; Mongrand et al., 2004; Roche et al., 2008; Yang and Murphy, 2009), while plant DRMs are mostly enriched in the primary PM sterols β-sitosterol and campesterol. The sterols of the Arabidopsis PM and DRM fractions contain 80% β-sitosterol and 10% campesterol, with no measureable stigmasterol content (Kierszniowska et al., 2009). Stigmasterol is more highly abundant in the PM and DRMs from oat and maize roots (Grandmougin et al., 1989; Norberg and Liljenberg, 1991; Borner et al., 2005). Sterol concentrations in PM and DRM fractions from tobacco bright yellow 2 (BY2) cell cultures contain ~50% stigmasterol, ~20% 24-methyl cholesterol, ~14% cholesterol, and only ~14% sitosterol (Mongrand et al., 2004). Plant sterols can also be conjugated with sugars, which in turn can be fatty acylated to form steryl glycosides and acyl-steryl glycosides. Steryl glycosides and acyl-steryl glycosides are enriched in Arabidopsis DRMs (Laloi et al., 2007; Minami et al., 2008). Experiments with artificial membranes suggest that the lipid diversity in plants decreases the temperature sensitivity and enhances lipid affinity for lipid raft formation (Xu et al., 2001; Beck et al., 2007). Sphingosine and its derivative sphingomyelin are the most common backbones of animal SLs (Cacas et al., 2012), but are largely absent in plants. Instead, the hydroxysphinganine phytosphingosine is the primary plant SL backbone. In place of the glycosylceramides that are abundant in mammals, glycosyl inositol phosphoceramides (GIPCs) are the predominant SLs in plants (Sperling et al., 2005; Pata et al., 2010; Blaas and Humpf, 2013). GIPC formation requires the enzyme inositol phosphorylceramide glucuronyltransferase1 (IPUT1), which is localized to the Golgi (Rennie et al., 2014). GIPCs are highly abundant in membranes of Arabidopsis leaves (Markham et al., 2006) and are also enriched in DRMs (Borner et al., 2005). However, tobacco BY2 cell cultures are more enriched in glycosphingolipids (Mongrand et al., 2004). Hydrogen bonding between membrane-intercalated sterols and either long chain bases (fatty acids) or large hydrophobic head groups of SLs are energetically favoured over interactions with PL (Lingwood and Simons, 2010). This increases the potential for SL/sterol partitioning and lateral immiscibility, which forces them into a denser, liquid-ordered state. The long SL fatty acid chains confer a higher melting temperature (Brown and London, 1998) and are thought to increase the thickness of the PM in the presence of sterols. However, although largely accepted, the latter might in fact result from the presence of proteins in these lipid environments, rather than the presence of sterols (Mitra et al., 2003). Chemical stripping experiments have shown that NDs are dependent on the presence of sterols and SLs, and therefore the rigidity of the liquid-ordered state is assumed to be the basis of the structured nature of ND-associated proteins and their potential to associate into highly stable protein complexes. Proteins localized to NDs Although DRMs and NDs should not be thought of interchangeably in the context of their biochemical and biophysical properties, proteomic analyses of DRMs have provided important information about proteins that could potentially associate with NDs through their affinity for sterols and SLs (Borner et al., 2005; Morel et al., 2006; Minami et al., 2008; Fujiwara et al., 2009; Takahashi et al., 2012, 2013). These proteomic exercises have been the jumping-off point for studies seeking to relate NDs to specific functions such as signal transduction (Demir et al., 2013; Srivastava et al., 2013; Matsui et al., 2014; Zauber et al., 2014), substrate transport (Sutter et al., 2006; Blakeslee et al., 2007; Yang et al., 2013), stress responses (Beck et al., 2007; Minami et al., 2008; Li et al., 2011), and pathogen responses (Bhat et al., 2005; Raffaele et al., 2009; van der Meer-Janssen et al., 2010; Bozkurt et al., 2014). A non-exhaustive list of ND and/or DRM-localized proteins referenced herein can be found in Table 1. Table 1. Proteins associated with liquid-ordered domains Protein  Function  ND identified (microscopy)  SL/sterol dependence in DRMs  References  ABCB1  Auxin transport  –  Extracted with 1% Brij and Triton X-100  Demir et al. (2013)   ABCB4  Auxin transport  Yes  SL dependent  Borner et al. (2005); Titapiwatanakun et al. (2009); Yang et al. (2013)   ABCB19  Auxin transport  Yes  Mainly SL, but also sterol dependent  Titapiwatanakun et al. (2009); Yang et al. (2013)   ABCB21  Auxin transport  –  Extracted with 1% Brij and Triton X-100  Demir et al. (2013)   ABCG36/PEN3  Pathogen response  –  Not MβCD-responsive  Minami et al. (2008); Kierszniowska et al. (2009)   ABCG37/PIS1  Coumarin and auxinic compound transport  –  FB1 and FEN treatment induce no changes in PM localization  Yang et al. (2013); Fourcroy et al. (2014)   FLA2  Cell adhesion  –  Sterol dependent (MβCD); extracted with 1% Brij and Triton X-100  Kierszniowska et al. (2009); Demir et al. (2013)   FLA proteins  Cell adhesion    Sterol dependent (MβCD)  Kierszniowska et al. (2009)   Flotillin1 and flotillins  Pathogen response  Yes  Sterol dependent (MβCD)  Borner et al. (2005); Haney and Long (2010); Li et al. (2012); Jarsch et al. (2014)   KAT1  Potassium channel  Yes  Not sensitive to MβCD  Sutter et al. (2006); Reuff et al. (2010); Jarsch et al. (2014)   PIN1  Auxin transport  Yes  FB1 and tsc10a (SL) show more intracellular structures; cpi1-1 (sterol)  Blakeslee et al. (2007); Men et al. (2008); Yang et al. (2013)   PIN2  Auxin transport  Yes  cpi1-1, cvp1-3, and fackel-J79 (sterol); DRM preparation  Men et al. (2008); Pan et al. (2009); Titapiwatanakun et al. (2009)   PIP2;1  Aquaporin  Yes  Sterol dependent (MβCD); not sensitive to FB1 (SL) or FEN (SL); extracted with 1% Brij and Triton X-100  Li et al. (2011); Demir et al. (2013); Yang et al. (2013)   Remorin1.3  Pathogen response  Yes  Sterol dependent (MβCD); extracted with 1% Brij and Triton X-100  Raffaele et al. (2009); Demir et al. (2013)   Remorins  Pathogen response  Yes    Mongrand et al. (2004); Lefebvre et al. (2010); Jarsch et al. (2014)   SLAH3  Nitrate, ABA signalling  Yes  Sterol dependent  Demir et al. (2013)   SYP121/PEN1  Pathogen response  Yes  In structures marked by filipin (sterol); in DRMs of rice  Assaad et al. (2004); Bhat et al. (2005); Fujiwara et al. (2009)   Protein  Function  ND identified (microscopy)  SL/sterol dependence in DRMs  References  ABCB1  Auxin transport  –  Extracted with 1% Brij and Triton X-100  Demir et al. (2013)   ABCB4  Auxin transport  Yes  SL dependent  Borner et al. (2005); Titapiwatanakun et al. (2009); Yang et al. (2013)   ABCB19  Auxin transport  Yes  Mainly SL, but also sterol dependent  Titapiwatanakun et al. (2009); Yang et al. (2013)   ABCB21  Auxin transport  –  Extracted with 1% Brij and Triton X-100  Demir et al. (2013)   ABCG36/PEN3  Pathogen response  –  Not MβCD-responsive  Minami et al. (2008); Kierszniowska et al. (2009)   ABCG37/PIS1  Coumarin and auxinic compound transport  –  FB1 and FEN treatment induce no changes in PM localization  Yang et al. (2013); Fourcroy et al. (2014)   FLA2  Cell adhesion  –  Sterol dependent (MβCD); extracted with 1% Brij and Triton X-100  Kierszniowska et al. (2009); Demir et al. (2013)   FLA proteins  Cell adhesion    Sterol dependent (MβCD)  Kierszniowska et al. (2009)   Flotillin1 and flotillins  Pathogen response  Yes  Sterol dependent (MβCD)  Borner et al. (2005); Haney and Long (2010); Li et al. (2012); Jarsch et al. (2014)   KAT1  Potassium channel  Yes  Not sensitive to MβCD  Sutter et al. (2006); Reuff et al. (2010); Jarsch et al. (2014)   PIN1  Auxin transport  Yes  FB1 and tsc10a (SL) show more intracellular structures; cpi1-1 (sterol)  Blakeslee et al. (2007); Men et al. (2008); Yang et al. (2013)   PIN2  Auxin transport  Yes  cpi1-1, cvp1-3, and fackel-J79 (sterol); DRM preparation  Men et al. (2008); Pan et al. (2009); Titapiwatanakun et al. (2009)   PIP2;1  Aquaporin  Yes  Sterol dependent (MβCD); not sensitive to FB1 (SL) or FEN (SL); extracted with 1% Brij and Triton X-100  Li et al. (2011); Demir et al. (2013); Yang et al. (2013)   Remorin1.3  Pathogen response  Yes  Sterol dependent (MβCD); extracted with 1% Brij and Triton X-100  Raffaele et al. (2009); Demir et al. (2013)   Remorins  Pathogen response  Yes    Mongrand et al. (2004); Lefebvre et al. (2010); Jarsch et al. (2014)   SLAH3  Nitrate, ABA signalling  Yes  Sterol dependent  Demir et al. (2013)   SYP121/PEN1  Pathogen response  Yes  In structures marked by filipin (sterol); in DRMs of rice  Assaad et al. (2004); Bhat et al. (2005); Fujiwara et al. (2009)   View Large For example, the inward-rectifying potassium channel KAT1 is a guard cell-localized protein, which has been implicated in abscisic acid- (ABA) dependent stomatal movement (Sutter et al., 2007). KAT1 function at the guard cell PM is dependent on syntaxin of plants 121 (SYP121/PEN1), and KAT1 forms large protein aggregates of ~500nm in diameter, consisting of an estimated 50 homo-tetramers (Sutter et al., 2006; Reuff et al., 2010). Lateral movement of KAT1 at the PM is very slow, as observed through pulse–chase-like strategies in conjunction with kymographic analyses (Sutter et al., 2006). Another stomatal protein, slow anion channel homolog 3 (SLAH3), interacts with calcium dependent protein kinase 22 (CPK21) in guard cell NDs (Demir et al., 2013). ABA binding to the receptor regulatory components of ABA receptor1/PYR1 (pyrobactin resistance 1)-like proteinc9 (RCAR/PYL9) results in recruitment of the ABA insensitive 1 (ABI1) protein phosphatase 2C to the SLAH3/CPK21 complex and rapid activation of the anion channel. It appears that the ordered ND environment is in this case essential to complex formation and function. SYP121, also known as PENETRATION1 (PEN1), appears to function in rapid recruitment of proteins to NDs. SYP121 is recruited to sites of Blumeria graminis (powdery mildew) infection in Arabidopsis, where it is required for papillae formation on the apoplastic side of the PM (Assaad et al., 2004). In unchallenged cells, SYP121 is continuously cycled between the endomembrane system and the PM, but rapidly partitions into NDs after pathogen infection (Bhat et al., 2005; Nielsen et al., 2012), in a manner similar to what is observed with KAT1 in guard cells. Plasma membrane intrinsic protein2;1 (PIP2;1/PIP2A) is a member of the aquaporin family that facilitate water movement across the PM (Santoni et al., 2003; Katsuhara et al., 2008). PIP2;1 partitions into DRMs (Demir et al., 2013) and its association with NDs has been investigated in great detail (Li et al., 2011). In control conditions, PIP2;1 is more or less homogenously distributed at the PM. However, under salt stress (NaCl) it increasingly co-localizes with Flotillin1 in specific foci at the PM, which is accompanied by the reduction of its lateral mobility (Li et al., 2011). This restriction was suggested to lead ultimately to PIP2;1 internalization from the PM. PIP2;1 appears to be able to move in and out of NDs. As PIP2;1 functions as an oligomer, this flexibility has been suggested to facilitate protein–protein interactions at the PM (Li et al., 2011). Notably, the NDs with which PIP2;1 associates are distinct from ABCB19, as FEN (a sterol biosynthesis inhibitor) treatment does not alter PIP2;1 localization at the PM (whereas ABCB19 is affected), and salt treatment did not alter ABCB19 localization (Yang et al., 2013). Emergence of standard ND marker proteins One highly conserved ND protein family, which was initially identified in other kingdoms, has become a useful ND marker in plants. In animal cells, flotillins are proteins of unknown function that are associated with caveolae, which are sterol- and SL-rich membrane invaginations that mediate a variety of cell to cell communication processes (Bickel, 1997; Babuke and Tikkanen, 2007; Takeshita et al., 2012). Arabidopsis Flotillin1 forms highly dynamic punctate structures at the PM (Li et al., 2012). Plant flotillins appear to have a role in symbiotic interactions with nitrogen-fixing bacteria (Haney and Long, 2010). Another important marker for NDs are the remorins, which were originally identified in the Solanaceae (Farmer et al., 1989; Reymond et al., 1996). Remorins are plant-specific, peripheral membrane proteins that aggregate in clusters of ~70nm in diameter (Raffaele et al., 2009). One of the most commonly used ND markers, Remorin1.3, has been shown to function in immune responses upon infection with the Potato virus X and Phytophtora infestans (Perraki et al., 2012; Bozkurt et al., 2014), but remorins remain otherwise functionally uncharacterized. Flotillin and remorin isoforms associate with distinct ND types (Jarsch et al., 2014), and are therefore increasingly used to analyse functional relationships between proteins and specific NDs. Other identified ND markers are KAT1, SLAH3 (mentioned above), and multiple members of the ABC transporter family of the subclasses B and G. ABC transporters form a superfamily of membrane transporters that is ubiquitously found across kingdoms. Arabidopsis ABCs cluster into eight subclasses on the basis of sequence homology and domain organization (Verrier et al., 2008). Plant ABCs of the subclass B transport organic acid compounds such as indole-3-acetic acid (the principle auxin IAA; ABCB1, ABCB4, ABCB19, ABCB21; Noh et al., 2001; Geisler et al., 2005; Terasaka et al., 2005; Kamimoto et al., 2012), and malate and citrate (ABCB14; Kaneda et al., 2011). The full-length ABCGs ABCG36/PEN3 and ABCG37/PIS1 transport a greater range of substrates, including xenobiotics (ABCG37; Ito and Gray, 2006), compounds involved in pathogen responses (ABCG36; Stein et al., 2006), cadmium (ABCG36; Kim et al., 2007), and the artificial auxin 2,4-dichlorophenoxyacetic acid (2,4-D) (ABCG37; Ito and Gray, 2006) as well as the IAA precursor indole-3-butyric acid (ABCG36&ABCG37; Strader and Bartel, 2009; Růžička et al., 2010), and coumarin compounds (ABCG37; Fourcroy et al., 2014). Although all of these transporters have been associated with NDs and/or DRMs, ABCB4, ABCB19, and ABCG37 are most commonly identified in these structures (Minami et al., 2008; Kierszniowska et al., 2009; Demir et al., 2013; Yang et al., 2013). The glycosylphosphatidylinositol (GPI)-anchored fasciclin-like arabinogalactan (FLA) proteins and some jacalin lectin proteins are recurrent residents of DRMs (Borner et al., 2005; Kierszniowska et al., 2009; Demir et al., 2013) that are also used as ND markers. The potential of these proteins to mediate interactions of NDs with cell walls makes them prime targets for investigation. Determinants for ND association of proteins Protein association with DRMs depends on electrostatic (hydrogen bonds) and hydrophobic interactions with the lipids (Murray et al., 1997; Cacas et al., 2012). These interactions appear to contribute to the formation and stability of functional NDs and associated protein complexes. An example is the interaction of the two Arabidopsis auxin transporters ABCB19 and PINFORMED1 (PIN1; Fig. 2). ABCB19 is associated with a subset of DRMs characterized primarily by SLs, but also sterols (Titapiwatanakun et al., 2009; Yang et al., 2013). PIN1 is a highly polarized transporter and is the primary mediator of polar auxin streams required for organogenesis and shootward auxin transport (Gälweiler et al., 1998; Benková et al., 2003; Reinhardt et al., 2005; Marhavý et al., 2011). PIN1 polar localization in some cell types is destabilized in mutants that are deficient in sterol biosynthesis (Willemsen et al., 2003). ABCB19 and PIN1 have been shown to interact in DRMs (Titapiwatanakun et al., 2009), and ABCB19 stabilizes PIN1 in discrete domains when the two proteins co-occur. Loss of ABCB19 in cells where it otherwise would co-localize with PIN1 results in reduced abundance of PIN1 at the PM (Blakeslee et al., 2007; Titapiwatanakun et al., 2009; Yang et al., 2013). PIN1 was also observed to exhibit increased intracellular abundance in cells within the normal ABCB19 expression domain in both the abcb19 mutant and the wild type, treated with sterol/SL biosynthesis inhibitors and/or stripping reagents (Yang et al., 2013). Co-purification of FLA2 with ABCB19 suggests that the GPI-anchored FLA2 could be part of the ABCB19/PIN1 ND (Fig. 2). Finally, PM localization of ABCB19 itself was substantially reduced in SL- and sterol-deficient mutants. The sterol and SL dependence of ABCB19 transport function and PIN1 interaction has been demonstrated in heterologous systems and in planta. Auxin transport activity of ABCB19 in heterologous systems (Schizosaccharomyces pombe) is increased by addition of sterols (Titapiwatanakun et al., 2009) or when ergosterol-containing membrane domains are present (Yang and Murphy, 2009). When ABCB19 and PIN1 are co-expressed, a synergistic increase in auxin transport and substrate specificity is observed (Blakeslee et al., 2007), and loss of either ABCB19 or PIN1 increases polar transport of benzoic acid (Blakeslee et al., 2007), which is normally not transported in a polar manner. GPI and S-acylation modifications are hallmarks of ND-associated proteins in animals (Garner et al., 2007). GPI anchoring and some S-acylation modifications appear in some cases to be solely sufficient for conferring DRM partitioning in plant cells (Borner et al., 2005; Sorek et al., 2007). About 1% of the Arabidopsis proteome encodes GPI-anchored proteins (Arabidopsis Genome Initiative, 2000; Borner et al., 2003). GPI anchoring results in protein targeting to the exoplasmic leaflet in animals and plants (Ikonen and Simons, 1998). Failure to generate GPI anchors in the abnormal pollen tube guidance 1 (aptg1) mutant aptg1 results in embryo lethality (Dai et al., 2014), highlighting the importance of GPI-anchored proteins for plant development and reproduction. Many proteins involved in cell to cell communication during seed coat and pollen development are over-represented in the GPI proteome (Borner et al., 2003; Tsukamoto et al., 2010; Edstam and Edqvist, 2014). One of these is COBRA-LIKE 10 (COBL10), a polar-localized GPI-anchored pollen tube protein that interacts with female chemotropic signals to direct growth within the pistil, leading to successful fertilization (Li et al., 2013). In aptg1 heterozygotes, COBL10 fails to localize to the PM (Dai et al., 2014). Plasmodesmata (PDs) connect neighbouring plant cells and are crucial for facilitating the intercellular exchange of nutrients and hormones, and to provide passageways for pathogens. PDs are enriched in sterols and SLs, and many PD proteins are represented in the GPI proteome (Fernandez-Calvino et al., 2011). Among these are members of the PD callose binding protein (PDCB) family that adjusts PD pore size by regulating the amount of callus deposition. PDCBs have been experimentally verified to be GPI anchored (Simpson et al., 2009). S-Acylation is the post-translational addition of a palmitate or, less frequently, a stearate moiety to a cysteine residue of the target protein. Recent proteomic studies suggest that S-acylation is the most frequent lipid modification of proteins, with a total of 581 S-acylated proteins (~2% of the proteome) identified in Arabidopsis (Hemsley et al., 2013; Hemsley, 2014). For instance, some members of the remorin family are S-acylated (Hemsley et al., 2013; Konrad et al., 2014). Two acylating enzymes, protein S-acyltransferase 10 (PAT10), located in the Golgi and tonoplast, and TIP GROWTH DEFECTIVE 1 (TIP1), localized to the PM, acylate proteins in Arabidopsis (Hemsley et al., 2005; Zhou et al., 2013; Qi et al., 2013). S-Acylation is the only fully reversible lipid modification, making it particularly important in lipid signalling (Smotrys and Linder, 2004). Notable examples of S-acylated proteins in Arabidopsis are members of the large family of leucine-rich repeats (LRR) receptor-like kinases, soluble N-ethylmaleimide-sensitive factor attachment protein receptors (SNAREs) such as SYP121/PEN1, and members of the Rho-like GTPases in plants (ROPs; Sorek et al., 2007; Craddock et al., 2012). GTP-bound ROPs have been localized to DRMs and they regulate cell division, morphogenesis, and cell polarity (Yalovsky et al., 2008). S-Acylation within the G-domain in ROP6 (at which GTP binding and hydrolysis takes place) is vital for its association with DRMs and, thus, probaby its function in vivo (Sorek et al., 2007, 2010). Inactive, GDP-bound ROPs can exhibit other lipid modifications such as prenylation, but locate to the cytosol. N-Myristoylation is the addition of the 14-carbon saturated fatty acid myristate (myristoyl-CoA) through N-myristoyltransferases (NMT) to an N-terminal glycine of a target protein (Qi et al., 2000; Sharma, 2004; Martinez et al., 2008). The myristate moiety interacts with negatively charged phosphatidylserine or phosphatidylinositol phosphates in the endoplasmic leaflet to enhance protein affinity for the PM (Resh, 2013). N-Myristoylation alone is not sufficient for DRM association in animals and plants, but tandem N-myristoylation in conjunction with single S-acylation appears to provide a viable signal for DRM partitioning in animal cells (Zacharias et al., 2002). The virulence of some bacteria and viruses depends on this lipid modification, as it confers PM association of their effector proteins, such as members of the HopZ family from Pseudomonas syringae and AC4 of the East African cassava mosaic Cameroon virus (Fondong et al., 2007; Lewis et al., 2007). However, N-myristoylation of ND-associated proteins has not been unequivocally demonstrated in plants. Prenylation, like N-myristoylation, enhances the affinity of proteins for the PM. C-15 farnesyl or C-20 geranylgeranyl isoprenoid moieties are added by protein farnesyl transferase (PFT) and protein geranyl-geranyltransferase-1 (PGGT-1), respectively, to target proteins. Both enzymes consist of one α-and one β-subunit, with the β-subunit conferring substrate specificity (Maurer-Stroh et al., 2003). PFT and PGGT-1 recognize the motif cysteine-a1-a2-X, where ‘a’ corresponds to aliphatic amino acids and X can be cysteine, methionine, serine, glutamine, alanine, or leucine (Sorek et al., 2011). The amino acid at the ‘X’ position generally determines whether a protein is farnesylated (serine, glutamine, methionine, alanine) or geranylgeranylated (leucine) (Reid et al., 2004; Resh, 2006). However, to date, prenylated proteins have been shown to associate exclusively with phospholipid domains. A summary of software currently available to predict co-and post-translational modifications is given in Table 2. Table 2. Searchable databases for protein modifications Modification  Software  Source  Plant Specific  General  ScanProsite  de Castro et al. (2006)   No  ARAMEMNON  Schwacke et al. (2003)   Yes  ExPASy  Artimo et al. (2012)   No  GPI anchors  PredGPI  Pierleoni et al. (2008)   Yes  BIG_PI Plant Predictor  Eisenhaber et al. (2003)   Yes  GPI-SOM  Fankhauser and Mäser (2005)   Included  FragAnchor  Poisson et al. (2007)   Included  S-Acylation  CSS-Palm 2.0  Ren et al. (2008)   No  PalmPred  Kumari et al. (2014)   No  NBA-Palm  Xue et al. (2006)   No  N-Myristoylation  NMT-The Myr Predictor  http://mendel.imp.ac.at/myristate/SUPLpredictor.htm  No  ExPASy Myristoylator  http://web.expasy.org/myristoylator/  Included  PlantsP  Podell and Gribskov (2004)   Yes  TermiNator3  Martinez et al. (2008)   Included  Prenylation  PrePS  Maurer-Stroh and Eisenhaber (2005)   No  Modification  Software  Source  Plant Specific  General  ScanProsite  de Castro et al. (2006)   No  ARAMEMNON  Schwacke et al. (2003)   Yes  ExPASy  Artimo et al. (2012)   No  GPI anchors  PredGPI  Pierleoni et al. (2008)   Yes  BIG_PI Plant Predictor  Eisenhaber et al. (2003)   Yes  GPI-SOM  Fankhauser and Mäser (2005)   Included  FragAnchor  Poisson et al. (2007)   Included  S-Acylation  CSS-Palm 2.0  Ren et al. (2008)   No  PalmPred  Kumari et al. (2014)   No  NBA-Palm  Xue et al. (2006)   No  N-Myristoylation  NMT-The Myr Predictor  http://mendel.imp.ac.at/myristate/SUPLpredictor.htm  No  ExPASy Myristoylator  http://web.expasy.org/myristoylator/  Included  PlantsP  Podell and Gribskov (2004)   Yes  TermiNator3  Martinez et al. (2008)   Included  Prenylation  PrePS  Maurer-Stroh and Eisenhaber (2005)   No  View Large Recent evidence suggests that some protein structures could directly confer protein association with NDs. Experiments utilizing a fusion of yellow fluorescent protein (YFP) to the C-terminus of potato Remorin1.3 (RemCA) containing an α-helical structure that forms a tight amphipathic hairpin within the lipid bilayer showed that this structure alone was sufficient to target the protein to DRMs (Perraki et al., 2012). Homologous peptides were subsequently identified in publicly available databases (Raffaele et al., 2013), but have not been confirmed. Multiple members of the remorin family in Arabidopsis are predicted and/or verified to be S-acylated at C-terminal cysteine residues, which is not sufficient for their interaction with the PM, but rather appears to enhance their interaction with this lipid environment (Hemsley et al., 2013; Konrad et al., 2014). A putative S-acylation site was also predicted for Flotillin1 (Borner et al., 2005), but still awaits experimental confirmation. Sterol and SL inhibitors and dyes used to analyse ND function Sterol and SL biosynthetic inhibitors have been extensively used to dissect ND function (Laloi et al., 2007; Titapiwatanakun et al., 2009; Li et al., 2011; Markham et al., 2011; Yang et al., 2013). The effects of pharmacological inhibitors can be controlled in a spatiotemporal manner that increases the chances to pinpoint the involvement of specific lipid species in ND formation. In combination with microscopic techniques, dyes allow for the direct visualization of the effects of sterol and SL depletion in real-time. However, care must be taken to avoid indirect or solvent effects on membrane integrity that increase marker protein degradation. Some of these inhibitors have been adopted from animal and fungal studies (Baloch et al., 1984; Bagnat et al., 2000) and can be used to distinguish the effects of structural sterol depletion from those of the reduction of steroid hormones (brassinosteroids) in plants (Yang et al., 2013). For example, fenpropimorph (FEN) is a chemical inhibitor of sterol synthesis that targets the plant-specific, endoplasmic reticulum (ER)-localized cycloeucalenol-obtusifoliol isomerase, but also has some impact on glucosylceramide (SL) accumulation (Fig. 1; Rahier et al., 1989; Laloi et al., 2007; Men et al., 2008). FEN treatment decreased the amount of Δ5-sterols, glucosylceramide, and overall DRM content of PM fractions from leek seedlings (Laloi et al., 2007). Propiconazol and brassinolide are inhibitors of steroid hormone biosynthesis that can be used to distinguish the effects of structural sterol depletion from hormone signalling (Hartwig et al., 2012; Yang et al., 2013). Fig. 1. View largeDownload slide Sites of alteration of lipid biosynthesis/content resulting from genetic lesions or pharmacological inhibition. Sphingolipid biosynthetic inhibitors are marked in red, sterol inhibitors are blue, and trafficking inhibitors are in black. Short-term treatment with FEN and MβCD leads to the internalization of sterol-dependent PM proteins. FEN-sensitive PM proteins have been found to be internalized to TGN/EE compartments (marked by ‘1st’) and characterized by SYP61 (green spiked vesicle). After extended treatment with MβCD and FEN, internalized proteins can first be found in the PVC/MVB (marked by ‘2nd’). The SYP61 compartment functions primarily in anterograde secretion of PM proteins (thicker arrow) but also receives a subset of PM proteins after endocytosis. Nanodomain-associated ABCB proteins co-localize with so-called ‘endosin bodies’ marked by SYP61 that are formed after treatment with ES-1 (Robert et al., 2008). ConcA, which induces fusion of PVC/MVB vesicles with TGN/EE vesicles, is marked by SYP61 and SLs. The thiol protease inhibitor E-64d inhibits the fusion of SYP61 with the PVC/MVB in an SL-dependent manner. PVC/MVB, pre-vacuolar compartment/multivesicular body; TGN/EE, trans-Golgi network/early endosome; ConcA, concanamycin A; ES-1, endosidin1; FEN, fenpropimorph; FB1, fumonisin B1. Mutations: gcs, glycosylceramide synthase; pas1, pasticcino1; loh1,3: LAG1 homologue; tsc10a-2, temperature-sensitive CSG2 suppressor; cvp1-3, cotyledon vascular pattern1; cpi1-1, cyclopropylsterol isomerase1, erh1, enhancing RPW8-mediated HR-like cell death, codes for IPCS. Fig. 1. View largeDownload slide Sites of alteration of lipid biosynthesis/content resulting from genetic lesions or pharmacological inhibition. Sphingolipid biosynthetic inhibitors are marked in red, sterol inhibitors are blue, and trafficking inhibitors are in black. Short-term treatment with FEN and MβCD leads to the internalization of sterol-dependent PM proteins. FEN-sensitive PM proteins have been found to be internalized to TGN/EE compartments (marked by ‘1st’) and characterized by SYP61 (green spiked vesicle). After extended treatment with MβCD and FEN, internalized proteins can first be found in the PVC/MVB (marked by ‘2nd’). The SYP61 compartment functions primarily in anterograde secretion of PM proteins (thicker arrow) but also receives a subset of PM proteins after endocytosis. Nanodomain-associated ABCB proteins co-localize with so-called ‘endosin bodies’ marked by SYP61 that are formed after treatment with ES-1 (Robert et al., 2008). ConcA, which induces fusion of PVC/MVB vesicles with TGN/EE vesicles, is marked by SYP61 and SLs. The thiol protease inhibitor E-64d inhibits the fusion of SYP61 with the PVC/MVB in an SL-dependent manner. PVC/MVB, pre-vacuolar compartment/multivesicular body; TGN/EE, trans-Golgi network/early endosome; ConcA, concanamycin A; ES-1, endosidin1; FEN, fenpropimorph; FB1, fumonisin B1. Mutations: gcs, glycosylceramide synthase; pas1, pasticcino1; loh1,3: LAG1 homologue; tsc10a-2, temperature-sensitive CSG2 suppressor; cvp1-3, cotyledon vascular pattern1; cpi1-1, cyclopropylsterol isomerase1, erh1, enhancing RPW8-mediated HR-like cell death, codes for IPCS. Methyl-β-cyclodextrin (MβCD) can selectively bind the small sterols in its hydrophobic cavity (Zidovetzki and Levitan, 2007). Treatment of tissues with MβCD strips membrane sterols, and, in most cases, releases or destabilizes sterol-dependent membrane proteins (Ohvo and Slotte, 1996; Zidovetzki and Levitan, 2007; Kierszniowska et al., 2009). It is also possible that MβCD treatment induces endocytosis, as is seen with Flotillin1 (Li et al., 2012). MβCD will displace many ND proteins during transit through secretory compartments beginning at the smooth ER, the initial point of sterol incorporation. However, short-term treatments can be used to restrict MβCD stripping to the PM, and internalization of PM proteins resulting from sterol depletion can be observed within 30–60min of the treatment (Li et al., 2012; Yang et al., 2013). MβCD treatment has also been used as a tool in proteomic analyses to differentiate membrane proteins that partition into DRMs due to sterol association from other DRM proteins (Kierszniowska et al., 2009). This study showed that regulatory/signalling membrane proteins are less consistently found to be associated with DRMs compared with, for instance, the FLAs. Filipin is an antibiotic with sterol-binding fluorochrome properties. Its fluorescent property (excitation 360nm, emission 480nm) makes it a useful sterol visualization agent (Norman et al., 1972; Drabikowski et al., 1973). However, filipin also disrupts sterol interactions and has been used to interfere with internalization of PIN2 (Men et al., 2008). Filipin also binds to PL when used at higher concentrations (Drabikowski et al., 1973) and can introduce lesions in the lipid bilayer, which could lead to unspecific internalization or clustering of proteins (Santos et al., 1998). The primary pharmacological SL biosynthesis inhibitor used in plant ND studies is the mycotoxin fumonisin B1 (FB1). FB1 inhibits ceramide synthase activity and has been shown to interrupt SL biosynthesis in plants (Wang et al., 1991; Abbas et al., 1994). However, as SLs are highly stable, depletion of SLs from membranes after FB1 treatment is relatively slow (Yang et al., 2013). 1-Phenyl-2-palmitoylamino-3-morpholino-1-propanol, which is a glycosylceramide synthase inhibitor, is also used as an SL synthesis inhibitor (Yang et al., 2013), but is thought to have multiple targets in plants. An overview of where the pharmacological inhibitors and lipid stripping agents function in the plant cell is shown in Fig. 1. Use of sterol and SL biosynthetic mutants to analyse ND function An alternative to the use of sterol and SL biosynthetic inhibitors is the use of well-characterized sterol and SL biosynthesis mutants in Arabidopsis (Willemsen et al., 2003; Men et al., 2008; Pan et al., 2009; Carland et al., 2010; Yang et al., 2013). Biosynthetic mutants are particularly useful, as lesions in genes encoding enzymes in the biosynthetic pathway can eliminate a desired lipid species. For example, cotyledon vascular pattern 1 (cvp1) is defective in sterol methyl transferase 2 (SMT2) and contains low levels of structural sterols, but not low levels of steroid hormones (Carland et al., 2010). The cvp1-3 mutant develops in a relatively normal fashion and sets seed. It also expresses lower levels of SMT1 and SMT3, thus reducing structural sterol levels in the root epidermis where high- resolution light microscopy analyses of fluorescent proteins and membrane dyes are most effective. Studies of cvp1-3 have been used to identify defects in trafficking of sterol-associated proteins such as PIN2, ABCB4, and ABCB19 (Pan et al., 2009; Yang et al., 2013). However, the residual levels of structural sterols in cvp1-3 epidermal cells and their contribution to PM protein stability are difficult to determine. An alternative is the use of mutants with lesions in cyclopropylsterol isomerase1/cycloeucalenol-obtusifoliol isomerase (cpi1/coi) that is expressed in epidermal cells (Men et al., 2008), and has been used to analyse trafficking of sterol-associated proteins (Men et al., 2008; Pan et al., 2009; Yang et al., 2013). However, cpi1 mutants are severely stunted and are defective in brassinosteroid biosynthesis (Men et al., 2008). Further, levels of the minority structural sterols cycloeucalenol and 24-methylenecycloartanol are higher in cpi1 than in the wild type, and potentially substitute for stigmasterol and sitosterol, leading to false-negative results (Men et al., 2008; Yang et al., 2013). Other mutants that have been utilized in trafficking studies are fackel, a weak sterol C14 reductase allele that is defective in later steps of sterol biosynthesis and exhibits faulty stomatal precursor cell division (Jang et al., 2000; Qian et al., 2013), and sterol methyltransferase 1 (smt1orc) that is defective in conversion of 24-methylenelophenol into citrostadienol, a precursor of stigmasterol, and exhibits dwarfism and cotyledon defects (Willemsen et al., 2003). Both of these mutants exhibit faulty PM localization of PIN proteins (Willemsen et al., 2003; Pan et al., 2009). Reduction of SL levels results in a visible effect on localization of ND-associated proteins. Reductions of very long chain fatty acids that contribute to SL formation in pasticcino1 mutants and multiple SLs in lag1 homolog 1 and 3 result in reduced auxin transport and partial mislocalization of PIN1 (Roudier et al., 2010; Markham et al., 2011), similar to what is observed in the abcb19 mutant. The temperature-sensitive CSG2 suppressor gene (tsc10a-2) encodes a 3-ketodihydrosphinganine reductase, a SL biosynthesis mutant that exhibits increased tricotyledon formation (Chao et al., 2011; Yang et al., 2013) and phenotypes similar to abcb19 (Yang et al., 2013). ABCB19 was shown to aggregate partially in compartments marked by SYP61 and the V-SNARE VTI12 in tsc10a, and PIN1 exhibited mislocalization similar to that observed in abcb19 mutants. Trafficking of ND-associated proteins Plant sterols and SLs are initially synthesized in the smooth ER and then are elaborated by glycosylation, acylation, and lipidification during anterograde movement through the secretory system (Bowles et al., 1977; Moreau et al., 1998; Melkonian et al., 1999; Rennie et al., 2014). Analysis of membrane fractions from leek seedlings showed that DRMs are only present in post-ER membrane compartments (Laloi et al., 2007), and NDs appear first to assemble within the Golgi and trans-Golgi network (TGN) (Laloi et al., 2007; Klemm et al., 2009). However, the exact timing of formation of specific types of NDs, the extent to which ND-associated proteins help nucleate and maintain NDs, and the relative roles of lipid species in the process remain unresolved. These questions are increasingly being examined in live imaging studies using variants of laser scanning confocal and TIRF microscopy in combination with lipid dyes, pharmacological inhibitors, mutants, and subcellular trafficking markers. In these studies, ND–protein associations have been confirmed to occur primarily in post-Golgi compartments, especially at the TGN/early endosome (EE), where exocytotic and endocytotic events co-occur (Dettmer et al., 2006). For example, PM abundance of the auxin transporter PIN2 is regulated by both secretion and endocytosis from the PM. In the sterol biosynthesis mutant cpi1-1, an increase of PIN2 at the PM is detected (Men et al., 2008). FRAP analysis in the presence of the vesicle trafficking inhibitor brefeldin A (BFA) suggested that the increase of PIN2 at the PM is due to reduced endocytosis of the protein. This regulation is consistent with the observed regulation of PIN2 abundance by ROP GTPases (Chen et al., 2012; Lin et al., 2012) that have been independently shown to associate with NDs (Prior et al., 2001, Sorek et al., 2007, 2010). ABCB19 occurs in NDs that appear to be compositionally different from the most common forms, and has been shown to recruit PIN1 into, or maintain PIN1 within these domains (Titapiwatanakun et al., 2009). The ABCB19/PIN1-ND is formed during anterograde trafficking through the secretory pathway (Yang et al., 2013). Short-term treatment with FEN results in the initial accumulation of ABCB19 in a TGN/EE compartment labelled by SYP61 before ultimate diversion to the pre-vacuolar compartment (PVC) via vesicles marked by VTI12 (Yang et al., 2013). This SYP61 compartment is part of both exocytotic and endocytotic streams, and thus contains lipids and proteins from both pathways (Dettmer et al., 2006). FRAP analysis of ABCB19–green fluorescent protein (GFP) showed slow recovery of PM signals and enhanced fluorescence recovery in FEN-treated SYP61 compartments, indicating that indeed exit of ABCB19–GFP from the SYP61 compartment is impaired in the presence of FEN. Thus, only the last step in trafficking of ABCB19 through the secretory pathway is dependent on sterols. However, depletion of SLs resulted in the accumulation of ABCB19–GFP in a different compartment. Treatment with FB1 resulted in the accumulation of ABCB19–GFP in the Golgi, and only a fraction was able to reach the TGN and PM (Yang et al., 2013). Similar results were observed in the SL biosynthesis mutant tsc10a-2. These experiments indicate that SLs and sterols assemble with proteins associated with this class of NDs in discrete stages, with SL association preceding sterol packing. Although these results differ from what had been previously proposed for DRMs (Laloi et al., 2007), it is also more consistent with the more recent recognition of GIPC abundance in Arabidopsis PMs (Blaas and Humpf, 2013; Markham et al., 2013). ABCB4 is regularly detected in canonical DRMs (Borner et al., 2005), and recently in SYP61 proteomics (Drakakaki et al., 2012). ABCB4 is also reduced in tsc10a-2 and after FB1 treatment, though to a lesser extent than ABCB19 (Yang et al., 2013). ABCB4 localization to the PM is also sensitive to very low concentrations of dimethylsulphoxide (DMSO; 0.05%), which is a common solvent for pharmacological inhibitors, dyes, and growth regulators (Kubeš et al., 2012). DMSO introduces water pores into lipid bilayers (Notman et al., 2006), which could potentially disturb lipid–protein interactions in ND-associated proteins. Such dissociation is likely to contribute to the observed ND dissociation, subsequent internalization, and exposure of proteolytic cleavage sites in ABCB4. For example, mammalian protein kinase C is stable at the membrane when it interacts with acidic lipids. In the presence of phosphatidylserine and diacylglycerol it is destabilized, exposing a proteolytic site within the protein, which ultimately leads to its degradation by Arg-C (Orr et al., 1992). A clear example of ND-dependent recruitment is seen in ABA regulation of Arabidopsis SLAH3. SLAH3 is a PM/ND-localized anion channel that is involved in ABA signalling (Geiger et al., 2011). Without the indirect activation of SLAH3 by the presence of ABA, the protein is primarily found in detergent-soluble endomembranes (Demir et al., 2013). SLAH3 activity depends strongly on its interaction with CPK21 at the PM, but this interaction is inhibited in the absence of ABA through the binding of the signalling phosphatase ABI1 to CPK21. The presence of ABA prevents this interaction, as under these conditions ABI1 preferentially forms a complex with ABA-bound RCAR1/PYL9. In the presence of ABA, the protein complex SLAH3/CPK21 is associated with sterol- and SL-rich NDs (Fig. 2; Demir et al., 2013). CPK21 localization to the ND is sterol dependent, as treatment with MβCD releases it from the PM. SLAH3/CPK21-ND formation occurs at the PM and not during anterograde trafficking within the secretory pathway. This association has been described as fully reversible and thus appears to differ from ABCB19. Fig. 2. View largeDownload slide Schematic representation of the SLAH3/CPK21 and ABCB19/PIN1 membrane nanodomains. Left: SLAH3 is active in the presence of abscisic acid (ABA). Binding of ABA to the RCAR/PYL9 receptor recruits ABI1 to the complex and blocks CPK21 interaction. This allows for the formation of the SLAH3/CPK21 complex in sterol- and SL-rich nanodomains. Whether ABI1 shows preferential binding to either nanodomains (SL shown with yellow head groups, sterols as small grey structures) or phospholipids (blue head groups) of the plasma membrane is not known. Centre: in the absence of ABA, ABI1 binds to CPK21, and SLAH3/CPK21 complex formation is inhibited. Under these conditions, SLAH3 and CPK21 are mainly found in detergent-soluble membranes. Red arrows indicate movement of CPK21 out of the nanodomains. Right: ABCB19/PIN1 interact in nanodomains with the result of increased specificity and effectiveness of polarized auxin transport. Consistent identification of FLA2 and other GPI-anchored fasciclin-like arabinogalactan proteins in nanodomains, and co-purification of FLA2 with ABCB19 suggest that a subclass of FLAs function in nanodomain–cell wall interactions. Schematic representation of FLA2 interaction with the ND via the C-terminal GPI anchor is shown. Grey rectangle, ethanolamine; blue circles, mannose; green circle, glucosamine; red lipid head group, inositol. Fig. 2. View largeDownload slide Schematic representation of the SLAH3/CPK21 and ABCB19/PIN1 membrane nanodomains. Left: SLAH3 is active in the presence of abscisic acid (ABA). Binding of ABA to the RCAR/PYL9 receptor recruits ABI1 to the complex and blocks CPK21 interaction. This allows for the formation of the SLAH3/CPK21 complex in sterol- and SL-rich nanodomains. Whether ABI1 shows preferential binding to either nanodomains (SL shown with yellow head groups, sterols as small grey structures) or phospholipids (blue head groups) of the plasma membrane is not known. Centre: in the absence of ABA, ABI1 binds to CPK21, and SLAH3/CPK21 complex formation is inhibited. Under these conditions, SLAH3 and CPK21 are mainly found in detergent-soluble membranes. Red arrows indicate movement of CPK21 out of the nanodomains. Right: ABCB19/PIN1 interact in nanodomains with the result of increased specificity and effectiveness of polarized auxin transport. Consistent identification of FLA2 and other GPI-anchored fasciclin-like arabinogalactan proteins in nanodomains, and co-purification of FLA2 with ABCB19 suggest that a subclass of FLAs function in nanodomain–cell wall interactions. Schematic representation of FLA2 interaction with the ND via the C-terminal GPI anchor is shown. Grey rectangle, ethanolamine; blue circles, mannose; green circle, glucosamine; red lipid head group, inositol. ABCG36/PEN3 is associated with multiple stress responses including pathogen responses, suggesting that NDs may function in a manner analogous to pathogen interactions observed in other kingdoms (Triantafilou et al., 2002; Rogers et al., 2012). ABCG36/PEN3 is regarded as a component of pathogen-associated molecular patterns (PAMP)-triggered immune response in Arabidopsis, and is activated upon infection with the powdery mildew Blumeria graminis (Underwood and Somerville, 2013). ABCG36/PEN3 exhibits a uniform PM distribution in uninfected wild-type leaf cells (Underwood and Somerville, 2013), and is also present within the SYP61 compartment (Drakakaki et al., 2012). Upon attempted fungal penetration, it forms distinct puncta at the site of infection, where it accumulates in the extracellular space within the papillae, into which it facilitates the transport of antimicrobial metabolites. In the roots, ABCG36/PEN3 is polar localized to the basal PM, designated as the outer polar domain, which defines the interface between the plant and the surrounding soil (Langowski et al., 2010). Notably, neither FB1 (SL biosynthesis inhibitor) nor FEN (sterol biosynthesis inhibitor) treatment appear to have a specific effect on its PM localization (Yang et al., 2013). Inositol phosphoryl ceramide synthase (IPCS) converts ceramide (SL) into inositol phosphorylceramide, which can be subsequently converted to GIPC (Rennie et al., 2014). IPCS was originally identified in an enhancer screen for RESISTANCE TO POWDERY MILDEW8 (RPW8), which confers the hypersensitive response to multiple powdery mildew pathogens (Xiao et al., 2001; Wang et al., 2008). In IPCS-defective mutants, the ceramide concentration is increased, resulting in elevated salicylic acid accumulation and RPW8 expression, and ultimately to increased resistance to powdery mildew (Wang et al., 2008). IPCS, like ABCB19, ABCB4, and ABCG36/PEN3, was localized to the SYP61 compartment. Thus, it appears to have a dual role in resistance to powdery mildew, by controlling the amounts of the signalling lipid ceramide through IPCS, and shuttling ABCG36/PEN3 to the sites of infection. Together, this also focuses our attention on the SYP61 compartment as an important component in ND trafficking and formation. The presence of IPCS in this compartment raises the possibility that even after exit from the Golgi, ND formation and restructuring of proteins could still occur. Do cell wall interactions stabilize NDs? Although NDs are clearly shown to stabilize associated protein complexes at the PM, there is currently no evidence that they function directly in polar trafficking events. However, as is the case with polarized PIN1 auxin transporter (Feraru et al., 2011), enzymatic removal and mutational disruption of the cell wall results in a more diffuse and uniform distribution of ABCB auxin transporters at the PM. This suggests that ND interactions with cell walls may occur via embedded proteins with exoplasmic interaction structures. Interestingly, the cell wall itself was shown to be sufficient for constraining the movement of ND-partinioning minimal proteins (containing only the membrane-spanning domains; Martinière et al., 2012). Two groups of proteins that consistently occur in DRM fractions are the FLAs and jacalin lectin proteins. FLAs are considered primary candidates for this function, as they comprise a large family, are GPI anchored, and contain arabinogalactan moieties that would be likely to interact with cell wall carbohydrates (Elortza et al., 2003; Johnson et al., 2003; Huang et al., 2013). However, such a function has not yet been clearly demonstrated for specific NDs. Conclusions The role of ordered membrane domains in regulating the trafficking and function of plant membrane proteins has gradually moved from the realm of speculation to accepted science. Studies of ND function in plants have been greatly enhanced by elucidation of plant and mammalian pathogen responses and the development of lipid-selective isolation techniques. Recent elucidation of SL biosynthetic pathways, development of a wide array of tools to explore PM–cell wall interactions, and the adaptation of new techniques to live imaging of plants have improved the characterization of plant ND formation and function. Wholesale biochemical isolation of DRMs has now been largely replaced by more refined cell biological approaches that track candidate proteins as they are incorporated into and interact with other proteins in sterol- and SL-rich domains. The ability to target and modify ND-associated proteins selectively and visualize them in planta with high-resolution microscopy techniques is likely to clarify the precise functions of ND structures in the near future. Acknowledgements This work was supported by the Department of Energy, Basic Energy Sciences, grant no. 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TI - Membrane nanodomains in plants: capturing form, function, and movement
JF - Journal of Experimental Botany
DO - 10.1093/jxb/erv054
DA - 2015-02-27
UR - https://www.deepdyve.com/lp/oxford-university-press/membrane-nanodomains-in-plants-capturing-form-function-and-movement-7OnB4XyzqK
SP - 1573
EP - 1586
VL - 66
IS - 6
DP - DeepDyve
ER -